Beyond Induction of Histone Acetylation: the Multi-Facets of the Antineoplastic Effect of Hdac Inhibitors

BEYOND INDUCTION OF HISTONE ACETYLATION: THE MULTI-FACETS OF THE ANTINEOPLASTIC EFFECT OF HDAC INHIBITORS

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in the Graduate

School of The Ohio State University

Chang-Shi Chen, M.S.

* * * * *

The Ohio State University

2006

Dissertation Committee: Approved by Professor Ching-Shih Chen, Advisor

Professor Pui-Kai Li ______Adviser Professor Matthew D. Ringel Graduate Program in Pharmacy

ABSTRACT

Histone deacetylase (HDAC) is recognized as one of the promising targets for cancer therapy as aberrant regulation of this epigenetic regulatory system has been shown to cause the repression of tumor suppressor genes and promotion of tumorigenesis. To date, many HDAC inhibitors have entered human clinical trials in light of their high potency in inhibiting tumor cell growth in vivo without incurring significant toxicity.

Although the effect of HDAC inhibitors on transcriptional regulation is well understood, an increasing body of evidence suggests that modulation of gene expression through chromatin remodeling might not be exclusively responsible for the antiproliferative effects of these agents. In this dissertation research, we demonstrate that while histones still represent a primary target for the physiological function of these protein deacetylases, the antineoplastic effect of HDAC inhibitors might also be attributed to histone acetylation-independent and transcription-independent mechanisms by disrupting the dynamic macromolecular complexes involving HDACs and by modulating the acetylation status of a series of nonhistone targets.

In the first part of this study, we demonstrate a novel histone acetylation

-independent mechanism by which the HDAC inhibitor, trichostatin A (TSA), induces

Akt dephosphorylation in U87MG glioblastoma cells by disrupting HDAC-protein phosphatase 1 (PP1) complexes. Evidence from Western blotting analysis indicates that ii this Akt dephosphorylation is not mediated through deactivation of upstream kinases or activation of downstream phosphatases, including PP1 and protein phosphatase 2A

(PP2A). However, PP1 inhibition, but not that of PP2A, can rescue the effect of TSA on phospho-Akt. Immunochemical analyses reveal that TSA blocks specific interactions of

PP1 with HDACs 1 and 6, resulting in increased PP1-Akt association. Moreover, by using HDAC isozyme-specific siRNAs, the role of HDACs 1 and 6 were confirmed as key mediators in facilitating Akt dephosphorylation. This selective histone acetylation-independent action of HDAC inhibitors on HDAC-PP1 complexes represents the first example of modulating specific PP1 interactions by small-molecule agents.

The results of the second part of this study present the in vitro and in vivo efficacy of (S)-HDAC-42, a novel phenylbutyrate-derived HDAC inhibitor synthesized in our laboratory, with reference to suberoylanilide hydroxamic acid (SAHA) in prostate cancer cells. We demonstrate that the anti-tumor effects of (S)-HDAC-42 are attributed to both histone acetylation-dependent and -independent mechanisms by interfering with the activation or expression status of a number of signaling targets. Especially noteworthy are the effects of (S)-HDAC-42 on the dephosphorylation of Akt and the repression of

Bcl-xL and survivin in PC-3 cells, both of which were not as evident in SAHA-treated cells. Together, these mechanisms provide a molecular basis to account for the higher in vivo potency of (S)-HDAC-42 than SAHA in suppressing established PC-3 xenograft tumor growth.

The third part of this study reports another histone acetylation-independent mechanism by which HDAC inhibitors sensitize prostate cancer cells to DNA damaging agents by targeting Ku70 acetylation. Ku70 represents a crucial component of the

iii non-homologous end joining (NHEJ) repair machinery for DNA double-strand breaks

(DSBs). Our data indicate that pretreatment of the Bax-deficient DU-145 cells with

HDAC inhibitors, including TSA, SAHA, MS-275, and (S)-HDAC-42, led to increased

Ku70 acetylation. Although hyperacetylation of Ku70 does not affect the Ku heterodimer formation, it reduces Ku70’s DNA end-binding affinity, and diminishes the cellular capability to repair drug-induced DNA DSBs. Moreover, mimicking acetylation by replacing lysine residues with glutamine in Ku70’s DNA binding cradle via site-directed mutagenesis also achieves the same effect. The ability of HDAC inhibitors to regulate cellular ability to repair DNA damage by targeting Ku70 acetylation underlies the viability of their combination with DNA damaging agents as a therapeutic strategy for prostate cancer.

All together, through this dissertation research, two novel histone acetylation-/transcriptional regulation -independent anticancer effects of HDAC inhibitors have identified. Moreover, we also show that these histone acetylation-dependent and -independent mechanisms underscore the pleiotropic antineoplastic effects of HDAC inhibitors, at both epigenetic and cellular levels in the in vitro cell culture and in vivo xenograft tumor models.

Dedicated to my parents, sisters, and my wife

ACKNOWLEDGMENTS

I would like to thank my advisor, Dr. Ching-Shih Chen, for his guidance, constant encouragement, and all the supports through this dissertation research. He is one of my role models in terms of his work ethic and dedication to scientific investigations.

I am grateful to Dr. Woan-Ru Shieh for the assistance in the experiment of the detection of PIP3, Dr. Ping-Hui Tseng for the help in the PDK1 kinase assay, and Nicole for the aid in the immunocytochemical assay in the study of the effect of HDAC inhibitors on Akt dephosphorylation.

I appreciate Dr. Samuel Kulp for the help in the tumor xenograft study and critical review of this manuscript, and Dr. Da-Sheng Wang and Dr. Qiang Lu for the synthesis of the HDAC inhibitors used in this study.

I acknowledge Dr. Shigemi Matsuyama for providing the Ku70 expression plasmids and critical comments for the manuscript of the Ku70 paper, Dr. Chih-Cheng

Yang for the help in the site-directed mutagenesis, Jack for the assistance in the immunocytochemical assays, Dr. Hsiao-Ching Yang for the assistance in molecular modeling, and Dr. Yen-Shen Lu for the discussions in the study of Ku70.

I also thank all of the former and current members in Chen’s lab for your company through this journey.

VITA

Jan. 2, 1973 ...... Born - Hsin-Chu, Taiwan

1996...... B.S. Agricultural Chemistry National Taiwan University, Taiwan

1998...... M.S. Microbiology and Immunology National Yang-Ming University, Taiwan

2005...... M.S. Medicinal Chemistry and Pharmacognosy The Ohio State University

1996 - 1998 ...... Graduate Research Assistant Institute of Microbiology and Immunology National Yang-Ming University, Taiwan

2000 - 2002 ...... Teaching Assistant Institute of Microbiology and Immunology National Yang-Ming University, Taiwan

2002 - Present ...... Graduate Research Associate College of Pharmacy The Ohio State University

PUBLICATIONS

Research Publications

1. Ping-Hui Tseng, Shu-Chuan Weng, Yu-Chieh Wang, Jing-Ru Weng, Chang-Shi Chen, Robert W. Brueggemeier, Charles L. Shapiro, Ching-Yu Chen, Sandra E. Dunn, Michael Pollak, and Ching-Shih Chen. Overcoming Trastuzumab Resistance in HER2-Overexpressing Breast Cancer Cells by Using a Novel Celecoxib-Derived PDK-1 Inhibitor. Mol Pharmacol. 2006 Nov;70(5):1534-1541.

vii 2. Samuel K. Kulp, Chang-Shi Chen, Da-Sheng Wang, Ching-Yu Chen, and Ching-Shih Chen. Antitumor Effects of a Novel Phenylbutyrate-Based Histone Deacetylase Inhibitor, (S)-HDAC-42, in Prostate Cancer. Clin Cancer Res. 2006 Sep 1;12(17):5199-206.

3. Chih-Cheng Yang, Chia-Yu Ku, Shou Wei, Chung-Wai Shiau, Chang-Shi Chen, Joseph Pinzone, Mathew D. Ringel, and Ching-Shih Chen. Peroxisome Proliferator-Activated Receptor g-Independent Repression of Prostate-Specific Antigen Expression by Thiazolidinediones in Prostate Cancer Cells. Mol Pharmacol. 2006 May;69(5):1564-70.

4. Hsiang-Yu Lin*, Chang-Shi Chen*, Shuan-Pei Lin, Jing-Ru Weng, Ching-Shih Chen. Targeting histone deacetylase in cancer therapy. Med Res Rev. 2006 Jan 31;26(4):397-413. * equal contributions to this article

5. Chang-Shi Chen, Shu-Chuan Weng, Ping-Hui Tseng, Ho-Pi Lin, and Ching-Shih Chen. Histone acetylation-independent effect of histone deacetylase inhibitors on Akt through the reshuffling of protein phosphatase 1 complexes. J Biol Chem. 2005 Nov 18;280(46):38879-87.

6. Qiang Lu, Da-Sheng Wang, Chang-Shi Chen, Yuan-Dong Hu, and Ching-Shih Chen. Structure-Based Optimization of Phenylbutyrate-Derived Histone Deacetylase Inhibitors. J Med Chem. 2005 Aug 25;48(17):5530-5.

7. Chung-Wai Shiau, Chih-Cheng Yang, Samuel K. Kulp, Kuen-Feng Chen, Chang-Shi Chen, Jui-Wen Huang, and Ching-Shih Chen. Thiazolidenediones mediate apoptosis in prostate cancer cells in part through inhibition of Bcl-xL/Bcl-2 functions independently of PPARγ. Cancer Res. 2005 Feb 15;65(4):1561-9.

8. Ho-Pi Lin, Samuel K. Kulp, Ping-Hui Tseng, Ya-Ting Yang, Chi-Cheng Yang, Chang-Shi Chen, and Ching-Shih Chen. Growth inhibitory effects of celecoxib in human umbilical vein endothelial cells are mediated through G1 arrest via multiple signaling mechanisms. Mol Cancer Ther. 2004 Dec;3(12):1671-80.

9. Sameek Roychowdhury, Robert A. Baiocchi, Srinivas Vourganti, Darshna Bhatt, Bradley W. Blaser, Aharon G. Freud, Jason Chou, Chang-Shi Chen, Jim J. Xiao, Mark Parthun, Kenneth K. Chan, Charles F. Eisenbeis, Amy K. Ferketich, Michael R. Grever, Ching-Shih Chen, Michael A. Caligiuri. Selective efficacy of depsipeptide in a xenograft model of Epstein-Barr virus-positive lymphoproliferative disorder. J Natl Cancer Inst. 2004 Oct 6;96(19):1447-57.

10. Qiang Lu, Ya-Ting Yang, Chang-Shi Chen, Melanie Davis, John C. Byrd, Mark R. Etherton, Asad Umar, and Ching-Shih Chen. Zn2+-chelating motif-tethered

viii short-chain fatty acids as a novel class of histone deacetylase inhibitors. J Med Chem. 2004 Jan 15;47(2):467-74.

11. Chih-Cheng Yang, Ho-Pi Lin, Chang-Shi Chen, Ya-Ting Yang, Ping-Hui Tseng, Vivek M. Rangnekar, and Ching-Shih Chen. Bcl-xL mediates a survival mechanism independent of the phosphoinositide 3-kinase/Akt pathway in prostate cancer cells. J Biol Chem. 2003 Jul 11;278(28):25872-8.

FIELDS OF STUDY

Major Field: Pharmacy

TABLE OF CONTENTS

Page Abstract...... ii

Dedication...... v

Acknowledgments...... vi

Vita...... vii

List of Tables ...... xiii

List of Figures...... xiv

Chapters:

1. Introduction...... 1

2. Literature Review: Histone Deacetylases and Histone Deacetylase Inhibitors.... 6

2.1 Chromatin Remodeling and Histone Code ...... 6 2.2 Classification and Regulation of Histone Deacetylases...... 7 2.3 Nonhistone Substrates of HDACs ...... 9 2.4 Chemical Biology and Development of HDAC Inhibitors...... 10 2.5 Antitumor Mechanisms of HDAC Inhibitors ...... 12

3. Histone Acetylation-Independent Effect of Histone Deacetylase Inhibitors on Akt Dephosphorylation...... 22

3.1 Introduction...... 22 3.2 Materials and Methods...... 23 3.2.1 Cell Culture...... 23 3.2.2 Reagents ...... 24 3.2.3 Immunoblotting...... 24 3.2.4 Cell Viability Assay ...... 25 3.2.5 Flow Cytometry ...... 26 3.2.6 Affinity Purification of HDAC-PP1 Complexes...... 26 x 3.2.7 Co-immunoprecipitation of PP1-Akt complexes...... 26 3.2.8 Subcellular Fractionation ...... 27 3.2.9 Ser/Thr Phosphatase Activity...... 27 3.2.10 Kinase Assay for Phosphatidylinositol 3-kinase (PI3K)...... 28 3.2.11 Determination of Phosphoinositide Formation ...... 29 3.2.12 Kinase Assay for Phosphoinositide-dependent kinase-1 ...... 30 3.2.13 Isozyme-specific Knockdown of HDACs with siRNA ...... 30 3.2.14 Immunocytochemical Analysis for PP1 and Phospho-Akt... 31 3.3. Results...... 31 3.3.1 Differential Effects of HDAC Inhibitors on Akt Dephosphorylation...... 31 3.3.2 Selective Dephosphorylation of Signaling Kinases ...... 32 3.3.3 Differential Effects of HDAC Inhibitors on Cell Proliferation and Cell Cycle...... 33 3.3.4 TSA-mediated Akt Dephosphorylation Is Not Caused by Changes in the Expression Level of Proteins Involved in Phospho-Akt Regulation...... 34 3.3.5 Inhibition of PP1 Prevents TSA-mediated Akt Dephosphorylation...... 34 3.3.6 TSA Disrupts HDAC-PP1 Complexes, Resulting in Increased PP1-Akt Associations ...... 35 3.3.7 Validation of the Involvement of HDACs 1 and 6 in Akt Dephosphorylation...... 37 3.4. Conclusion ...... 37

4. Antitumor Effects of A Novel Phenylbutyrate-based Histone Deacetylase Inhibitor, (S)-HDAC-42, in Prostate Cancer...... 55

4.1 Introduction...... 55 4.2 Materials and Methods...... 56 4.2.1 Reagents ...... 56 4.2.2 Cell Culture...... 57 4.2.3 Cell Viability Assay ...... 57 4.2.4 Apoptosis Assay...... 58 4.2.5 Immunoblotting...... 58 4.2.6 Semiquantitative RT-PCR...... 59 4.2.7 In Vivo Studies ...... 59 4.2.8 Statistical Analysis...... 60 4.3 Result ...... 60 4.3.1 (S)-HDAC-42 Induces Apoptosis in Prostate Cancer Cells.... 60 4.3.2 (S)-HDAC-42 Facilitates the Dephosphorylation of Akt and Alters the Dynamics of Bcl-xL Expression...... 62 4.3.3 (S)-HDAC-42 Attenuates Protein Levels of IAP Family Members ...... 64 4.3.4 (S)-HDAC-42 Suppresses Prostate Tumor Xenograft Growth In Vivo...... 65 xi 4.4. Conclusion ...... 66

5. Evaluation of the DNA Double-Stranded Break Repair Function of Acetylated Ku70...... 78

5.1 Introduction...... 78 5.2 Materials and Methods...... 80 5.2.1 Cell Culture...... 80 5.2.2 Reagents ...... 80 5.2.3 Immunoblotting...... 81 5.2.4 Immunoprecipitation of Ku70...... 81 5.2.5 Transfection of Flag-tagged Ku70 Expression Plasmids ...... 82 5.2.6 DNA End-Binding Activity of Ku70 ...... 82 5.2.7 Cell Survival Assay...... 83 5.2.8 Apoptosis Assay...... 83 5.2.9 Immunocytochemical Detection of γH2AX Foci Formation.. 84 5.2.10 Molecular Modeling Analysis...... 84 5.2.11 Statistical Analysis ...... 85 5.3 Results...... 86 5.3.1 Pretreatment with HDAC Inhibitors Sensitizes Prostate Cancer Cells to Agents that Generate DNA DSBs ...... 86 5.3.2 HDAC Inhibition Leads to Increased Acetylation of Ku70.... 87 5.3.3 HDAC Inhibition Diminishes Cellular Ability to Repair DNA DSBs ...... 88 5.3.4 Constitutive Acetylation of Lysine Residues in DNA-Binding Domains Mimics HDAC Inhibitors in Suppressing Ku70’s End-Binding Affinity...... 89 5.3.5 Molecular Modeling Analysis of the Interaction of Ku70/Ku80 Heterodimer with DNA...... 90 5.3.6 Constitutive Acetylation of Ku70 Diminishes the Cellular Ability to Repair DNA DSBs...... 91 5.4 Conclusion ...... 91

6. Discussions and Perspectives...... 105

6.1 HDAC Inhibitors and Akt Dephosphorylation ...... 106 6.2 Anticancer Effects of (S)-HDAC-42...... 107 6.3 Acetylation of Ku70 Impairs Its DNA-Repair Function...... 108 6.4 Future Perspectives ...... 109

Bibliography ...... 113

xii

LIST OF TABLES

Table Page

2.1 HDAC Family ...... 15

2.2 Representative Nonhistone Substrates of HDACs ...... 16

2.3 Classification of HDAC Inhibitors ...... 17

2.4 Tumor-Associated Proteins Whose Transcriptional Expression is Altered in Response to HDAC Inhibitor Treatment of Cells ...... 18

3.1 Effects of HDAC Inhibitors on Cell Cycle Distribution in PTEN-Null U87MG Glioblastoma Cells ...... 39

xiii

LIST OF FIGURES

Figure Page

1.1 A Model for the Antitumor Action of HDAC Inhibitors ...... 4

1.2 Effects of Histone Deacetylase Inhibition on Nonhistone Proteins ...... 5

2.1 Space-Filling Representation of TSA in the Active Site of HDL ...... 19

2.2 Structural Basis for TSA Binding to HDLP Binding Site ...... 20

2.3 Molecular Docking of HTPB and (S)-HDAC-42 into the Active-Site Pocket of HDLP ...... 21

3.1 Effects of HDAC Inhibitors on Akt Dephosphorylation in PTEN-Null U87MG Glioblastoma Cells ...... 40

3.2 A Dose-Dependent Effects of TSA and SAHA on the Phosphorylation State of 308Thr- and 473Ser-Akt vis á vis ERK1/2, p38, and JNK MAP Kinases in U87MG Cells...... 41

3.3 Dose Dependence of the Growth Inhibitory Effects of HDAC Inhibitors in U87MG Cells...... 42

3.4 Akt Dephosphorylation Is Not Due to Alterations in the Expression Level of Proteins Involved in Phospho-Akt Regulationin U87MG Cells ...... 43

3.5 TSA Does Not Affect the Enzyme Activities of PI3K and PDK1 Kinases...... 44

3.6 TSA Does Not Affect the PIP3 Level in U87MG Cells...... 45

3.7 The Protein Phosphatase Inhibitors Exhibit Differential Effects on TSA-Mediated Dephosphorylation of Akt...... 46

3.8 TSA Does Not Affect the Enzyme Activities of PP1/2A Phosphatases...... 47

3.9 TSA Treatment Did Not Alter the Expression Level of HDACs...... 48 xiv

3.10 TSA Selectively Disrupts the Association of PP1 with HDAC1 and 6 in U87MG Cells...... 49

3.11 TSA Does Not Change the Relative Abundance of PP1 or PP2A in the Nucleus or Cytoplasm in Drug-Treated U87MG Cells ...... 50

3.12 TSA Treatment Leads to Increase PP1-Akt Association...... 51

3.13 TSA Treatment Leads to Increase PP1-Akt Association and Akt-Dephosphorylation...... 52

3.14 Validation of the Involvement of HDAC1 or 6 in PP1-Mediated Akt Dephosphorylation...... 53

3.15 Schematic Diagram Depicting the Effect of HDAC Inhibitors on the Dynamics of PP1 Complex Formation with HDACs and Phospho-Akt...... 54

4.1 Antiproliferative Effects of (S)-HDAC-42 and SAHA in Three Different Prostate Cancer Cell Lines...... 68

4.2 Antiproliferative Effects of (S)-HDAC-42 and SAHA in Normal Prostate Epithelial Cells, PrECs...... 69

4.3 Morphological changes in PC-3, DU-145 and LNCaP Cells Treated with Vehicle Control, 1 µM (S)-HDAC-42 or 2.5 µM SAHA for 24 Hours...... 70

4.4 Effects of (S)-HDAC-42 versus SAHA on Various Biomarkers Associated with HDAC Inhibition or Apoptosis in PC-3 Cells...... 71

4.5 Effects of (S)-HDAC-42 versus SAHA on Apoptosis in PC-3 Cells...... 72

4.6 Dose-Dependent Effects of (S)-HDAC-42 versus SAHA on the Phosphorylation State of 473Ser- and 308Thr-Akt and 136Ser-Bad in PC-3 Cells...... 73

4.7 Dose-Dependent Effects of (S)-HDAC-42 versus SAHA on the Expression Levels of Bad, Bcl-xL, Bcl-2, and Bax in PC-3 Cells...... 74

4.8 Dose-Dependent Effects of (S)-HDAC-42 versus SAHA on the Transcription of Survivin and Bcl-xl in PC-3 Cells...... 75

4.9 Dose-Dependent Effects of (S)-HDAC-42 versus SAHA on the Expression Levels of Members of The IAP Family of Proteins Including Survivin, cIAP-1, cIAP-2, and ILP, in PC-3 Cells...... 76

xv 4.10 Effects of Oral (S)-HDAC-42 at 25 mg/kg per day (q.d.) or 50 mg/kg every other day (q.o.d.) versus Oral SAHA at 50 mg/kg per day on the Growth of Established PC-3 Tumors in Nude Mice and the Expression of Intratumoral Biomarkers of Drug Activity...... 77

5.1 Expression Status of Ku70, Ku80, and Bax in Three Prostate Cancer Cell Lines, LNCaP, DU-145, and PC-3...... 94

5.2 Effect of (S)-HDAC42 on Sensitizing DU-145 Prostate Cancer Cells to DNA DSB-Inducing Agents ...... 95

5.3 Effect of 250 nM (S)-HDAC42 Pretreatment on Drug-Induced DNA Fragmentation in DU-145 Cells...... 96

5.4 Dose-Dependent Effect of Four Different HDAC Inhibitors, Including (S)-HDAC42, SAHA, MS-275, and TSA on Histone H3 Acetylation, p21WAF1/CIP1 Expression, α-tubulin Acetylation, and Expression of Ku70 and Ku80 in DU145 Cells...... 97

5.5 HDAC Inhibitor-Mediated Ku70 Acetylation in DU145 Cells ...... 98

5.6 (S)-HDAC42 Did Not Disrupt the Interaction between Ku70 and Ku80 ...... 99

5.7 Dose-Dependent Effect of (S)-HDAC42 and MS-275 on the DNA-End Binding Activity of Ku70 in DU145 Cells...... 100

5.8 Effect of (S)-HDAC42 on Enhancing Drug-Induced γ-H2AX Foci Formation...... 101

5.9 Effect of Constitutive Acetylation of Lysine Residues in the DNA-Binding Cradle of Ku70 on Its DNA-End Binding Activity...... 102

5.10 Molecular Modeling Analysis of the Mode of Interactions between the Ku Heterodimer and DNA ...... 103

5.11 Effects of the Constitutive Acetylation of K282 and K338 of Ku70 on Drug-Induced γH2AX Foci Formation ...... 104

xvi

CHAPTER 1

INTRODUCTION

The transformation of cancer cells requires a number of integrated etiologic events, including genetic, cytogenetic, and epigenetic processes. Among them, the genetic and cytogenetic routes to tumorigenesis have been extensively studied. Following the completion of the human genome project, epigenetics has emerged as an important area to understand how the genome translates its information in tumor formation. Epigenetic mechanisms, which involve DNA and histone modifications, affect the heritable changes in gene expression without altering the coding sequence of the underlying DNA. Moreover, these dynamic epigenetic events provide a mechanism for an organism to respond to environmental stimuli through changes in gene expression during cell growth and differentiation. Accumulating evidence indicates that dysregulation of these epigenetic processes can lead to abnormal expressions of a subset of genes and serious developmental defects, which underlie the pathogenesis of many diseases such as cancer (1-3). Histone modifications, including histone phosphorylation, methylation, sumoylation, ubiquitination, ADP-ribosylation, and especially acetylation, play essential roles in generating the dynamic state of chromatin structure (4) (Section 2.1 for discussion). Histone acetylation, which is predominantly linked to transcriptional regulation, is the most extensively studied posttranslational histone modification to date.

The functional balance of histone acetylation relies on its highly reversible nature that

1 depends on the accuracy and efficiency of its reverse reaction, histone deacetylation. The dynamic equilibrium of the acetylation status of histones is controlled by two specific groups of enzymes, named histone acetyltransferases (HATs) and histone deacetylases

(HDACs) (Section 2.2 for discussion). Many reports indicate that HDACs are critical regulators of fundamental cellular events and their dysregulation is involved in tumorigenesis (5,6). Importantly, HDAC inhibitors can selectively induce growth arrest, differentiation, and/or apoptosis of transformed cells and therefore are being explored as therapeutic agents for the treatment of certain forms of cancer (7,8) (Section 2.4 for discussion). The general concept for the epigenetic or transcriptional therapy of cancer by HDAC inhibition is that HDAC inhibitors can induce the re-expression of a specific set of tumor suppressor genes, differentiation genes, and cell cycle inhibitor genes which are down-regulated during neoplastic transformation (9) (Figure 1.1). However, recent studies have shown that regulation of transcription via the acetylation status of histones per se is not sufficient to explain the anticancer activity of HDAC inhibitors (10,11), although histones may still be their primary targets (12,13) (Section 2.5 for discussion). For example, expanding lists of acetylated proteins have been identified as the nonhistone substrates of HDACs (Section 2.3 for discussion). Targeting of the acetylation status of these signal mediators by HDAC inhibitors may also account for their anticancer activity (14-16) (Figure 1.2). The study of the biological functions of the acetylation status of histones and nonhistone proteins, called acetylation biology (17,18) or acetylomics (19), is becoming an important research area. The further characterization of HDACs and their substrates and a greater understanding of the dynamic function of these proteins may provide us new insights into tumorigenesis and new anticancer drug development.

The aim of this dissertation research is to evaluate the modes of action of the

HDAC inhibitors in the regulation of histone acetylation-/transcription

2 regulation-dependent and -independent antineoplastic effects in glioblastoma and prostate cancer cells. The present study particularly focuses on evaluation of the anticancer effects of (S)-(+)-N-hydroxy-4-(3-methyl-2-phenyl-butyrylamino)benzamide,

(S)-HDAC-42, which is a phenylbutyrate-derived histone deacetylase inhibitor developed in our laboratory and is currently undergoing preclinical evaluation through the Rapid Access to Intervention Development Program at the National Cancer Institute

(20,21). (Section 2.4 for discussion).

Figure 1.1: A Model for the Antitumor Action of HDAC Inhibitors (9)

Figure 1.2: Effects of Histone Deacetylase Inhibition on Nonhistone Proteins (16)

CHAPTER 2

LITERATURE REVIEW:

HISTONE DEACETYLASES AND HISTONE DEACETYLASE INHIBITORS

2.1 Chromatin Remodeling and Histone Code In eukaryotes, the packaging of genomic DNA into a higher ordered and dynamic structure provides a critical point for controlling gene expression by regulating the access of transcription machinery, such as transcriptional factors and RNA polymerases, to the promoter region. This highly ordered molecular structure is known as chromatin, which is composed of multiple repeating subunits called nucleosomes. A nucleosome consists of 146 base pairs of double-stranded DNA wrapped around a histone core octamer consisting of two copies of each histone 2A, H2B, H3 and H4 (22). Histones are small, basic proteins that consist of a globular domain and an N-terminal tail that protrudes from the nucleosome. Although histones are some of the most evolutionarily conserved proteins, they are also among the most variable in terms of posttranslational modifications (23). Based on current knowledge, histone modifications and DNA methylation are two of the major factors influencing chromatin architecture, which along with ATP-dependent chromatin remodeling, are the principle epigenetic mechanisms by which tissue-specific gene expression patterns and global silencing are established and maintained (24-26).

The lysine rich N-terminal histone tails extending from the nucleosome can serve as targets for diverse posttranslational modifications, including phosphorylation on serine

6 and threonine residues, ADP-ribosylation on aspartate and glutamate residues, methylation on lysine and arginine residues, and sumoylation, ubiquitination and acetylation on lysine residues (27-30). It has been proposed that the pattern of these modifications can act as an information code that regulates gene transcription. This pattern of modification has been termed the histone code (31,32). The particular pattern of histone modifications may play a role in determining the affinity for chromatin-associated proteins, which determine whether the chromatin takes on an active or silent state (32). For example, acetylation of histones can effectively mask their positively charged lysine residues, thereby interfering with the electrostatic interactions between histone tails and the negatively charged DNA backbone. This leads to relaxed nucleosomal structures and a permissive transcriptional state for targeted genes. By contrast, when histone proteins are in a hypoacetylated state, nucleosomes are tightly compacted, resulting in transcriptional repression due to the restricted access of transcriptional factors to the promoter region. It is noteworthy that aberrant regulation of this epigenetic marking system has been shown to cause inappropriate gene expression, a key event in the pathogenesis of many forms of cancer (1,4,33).

2.2 Classification and Regulation of Histone Deacetylases Histone deacetylases (HDACs) represent a family of enzymes that compete with the action of histone acetyltransferases (HATs) to modulate chromatin structure and regulate transcriptional activity via changing the acetylation status of nucleosomal histones. The opposing functions of HATs and HDACs in both activating and repressing transcription by controlling the tightness of nucleosomal integrity reflect the processes that are involved in transcriptional regulations (34). Since the identification of the first histone deacetylase, at least eighteen HDACs have been isolated in humans (35,36).

Based on their homology to yeast HDACs, mammalian HDACs can be categorized into

7 three classes. Moreover, each HDAC has been mapped to a chromosomal location (Table

2.1). Mechanistically, Class I and II HDACs are distinct from Class III HDACs in co-factor requirement; that is, the former require an active site zinc ion (Zn2+) to mediate deacetylation catalysis while the latter are dependent on nicotinamide adenine dinucleotide (NAD+) (37). Class I HDACs, including HDAC1, 2, 3 and 8, all share a certain degree of homology to the yRpd3 (yeast Reduced potassium dependency 3), are generally localized to the nucleus and ubiquitously expressed in many human cell lines and tissues (38).

Class II HDACs are homologous to yHda1 (yeast Histone deacetylase-A 1) and larger in size than the other two classes of HDACs (39). They can be further subdivided into two subclasses, IIa (HDAC4, 5, 7, 9) and IIb (HDAC6 and 10), based on their sequence homology and domain organization. Class IIa HDACs contain a highly conserved C-terminal deacetylase catalytic domain (~ 420 amino acids) homologous to yHda1, but have an N-terminal domain with no similarity to HDACs in other classes. Class IIb HDACs are characterized by presence of an extra deacetylase domain, although this duplication is partial in the case of HDAC10 (39). An analysis of the catalytic activity of these two separate catalytic domains of HDAC6 by site-directed mutagenesis suggests that these two domains might function independently (40). Class II HDACs are generally localized to cytoplasm and can be shuttled to the nucleus as they are needed, suggesting potential cellular functions by regulating the acetylation status of nonhistone substrates. Additionally, HDAC11 is the most recently cloned and characterized HDAC isoform and possesses properties common to both Class I and Class II HDACs (41). The third class of

HDACs is the sirtuins (SIRT1-7), which are homologous to the ySir2 (yeast Silent information regulator 2) family of proteins (42). As aforementioned, these enzymes require NAD+ for deacetylase activity in contrast to the Zn2+-catalyzed mechanism used

8 by class I and class II HDACs. Their biological functions have been linked to chromatin silencing, cellular metabolism and aging (42,43).

As vital regulators of many cellular process, the activities of HDACs are tightly regulated by multiple mechanisms (44). Many reports demonstrated that the activities of

HDACs can be regulated by posttranslational modifications, such as phosphorylation and sumoylation (45,46), as well as by subcellular localization which is controlled by 14-3-3 proteins (47,48). Less studied, but perhaps equally important, is that the activities of

HDACs can also be regulated by transcriptional regulation, availability of cofactors, and proteolytic processing (49-51). Thus, a complete understanding of how HDACs are regulated will contribute not only to our knowledge of chromatin structure and gene control, but will provide new insight into approaches for developing therapeutic HDAC inhibitors with improved specificity.

2.3 Nonhistone Substrates of HDACs Extensive phylogenetic analysis suggests that HDACs belong to an ancient enzyme family found in animals, plants, fungi and bacteria (52,53). It is believed that HDACs evolved in the absence of histone proteins. Indeed, eukaryotic HDACs are responsible for modifying diverse types of nonhistone proteins as well as histones, the most thoroughly studied protein substrates. It has been postulated that key HDAC substrates might not be histones, but instead belong to the expanding list of nonhistone proteins, including transcription factors, signal transduction mediators, a microtubule component, and a molecular chaperone (Table 2.2). Additionally, non-protein molecules such a polyamines or metabolic intermediates might also serve as valid substrates (19).

Acetylation, a key posttranslational modification of many proteins, is a dynamic, reversible, and highly regulated chemical modification responsible for regulating critical intracellular pathways (54). Acetylation of a protein can have many different effects.

9 Firstly, both acetylation and ubiquitination often occur on the same lysine residues, and there is a regulatory cross-talk between these two modification (55). Secondly, acetylation of the lysine residues can also compete with other posttranslational modifications, such as sumoylation, or influence other posttranslational modifications such as phosphorylation (56-58). Thirdly, the DNA-binding activity of nonhistone proteins, including transcriptional factors and enzymes involved in DNA metabolism and repair, can also be affected by acetylation modification (59,60). Consequently, this process may control the stability (61,62), localization (63,64), protein dimerization (65), and protein-protein (66-68) and protein-DNA interactions (59,60). It is noteworthy that acetylation of many proteins have been reported, however, the specific protein deacetylases that regulate the reverse process remain unclear (58,66,69-71).

2.4 Chemical Biology and Development of HDAC Inhibitors To date, several structurally distinct classes of HDAC inhibitors have advanced into phase I, II and/or III clinical trials in solid tumors and hematological malignancies (16). On the basis of their chemical structures, major HDAC inhibitors can be classified into four categories: short-chain fatty acids, hydroxamic acids, benzamide derivatives, and cyclic peptides (Table 2.3). In general, these inhibitors, especially the hydroxamates, do not exhibit isozyme specificity in HDAC inhibition. However, MS-275, a benzamide derivative, has been reported to be ineffective in inhibiting the α-tubulin deacetylase HDAC6 (72), while trichostatin A (TSA), a hydroxamate derivative, can induce robust hyperacetylation of α-tubulin at submicromolar concentrations (73). Conceivably, isozyme-specific HDAC inhibitors will be of both research and clinical interests. More recently, a small molecule, tubacin, has been identified, via high-throughput screening, to selectively inhibit HDAC6-mediated α-tubulin deacetylation without affecting the level of histone deacetylation (74). Tubacin selectively binds to one of the two catalytic

10 domains possessing tubulin deacetylase activity. However, rational design of isozyme-specific HDAC inhibitors proves to be challenging due to the shared mode of catalytic mechanism and the high degree of homology in the catalytic domain of class I and II HDACs.

X-ray crystallographic analysis of HDLP (histone deacetylase-like protein), a bacterial HDAC homologue, has revealed a distinct mode of protein-ligand interactions whereby inhibitors like suberoylanilide hydroxamic acid (SAHA) and TSA mediate enzyme inhibition (75) (Figure 2.1). The HDAC catalytic domain consists of a narrow, tube-like pocket of a length equivalent to that of 4-6 carbon straight chains. A Zn2+ cation is positioned near the bottom of this enzyme pocket, which, in cooperation with two His-Asp charge-relay systems, facilitates the deacetylation catalysis. The structure of TSA and many other HDAC inhibitors might be divided into three motifs, each of which interacts with a discrete region of the enzyme pocket. For TSA, these include a

Zn2+-chelating function (i.e., hydroxamic acid), a conjugated, aliphatic chain as linker, and a polar cap group (i.e., the dimethylaminophenyl moiety) (Figure 2.2). On the basis of this three-component working model (i.e., cap group—linker—Zn2+-chelating motif), we proposed that short-chain fatty acids mediated HDAC inhibition through nonspecific hydrophobic interactions with surface residues located at the enzyme pocket entrance and/or the hydrophobic region inside the tube-like pocket. Accordingly, we rationalized that the weak potency of short-chain fatty acids in

HDAC inhibition is attributable to their inability to access the Zn2+ cation in the active-site pocket, which plays a pivotal role in the deacetylation catalysis. This premise constitutes the basis for our hypothesis that tethering them with a Zn2+-chelating motif through a hydrophobic linker could enhance the HDAC-inhibitory potency of short-chain fatty acids. Consequently, we developed a novel strategy of tethering short-chain fatty acids (valproate, butyrate, phenylacetate, and phenylbutyrate) with hydroxamate through

11 aromatic amino acid linkers to generate a novel class of HDAC inhibitors (20). Among a series of derivatives with varying potency, HTPB (a hydroxamate-tethered phenylbutyrate) represented an optimal agent with IC50 of 44 nM, compared to 0.4 mM for the parent molecule phenylbutyrate. As HTPB is structurally distinct from other potent HDAC inhibitors such as TSA and SAHA, there exists a high degree of flexibility in the active-site pocket in accommodating cap groups with different stereoelectronic properties. Docking of HTPB into the HDLP binding domain reveals interaction with a high degree of similarity with that of TSA (Figure 2.3). Moreover, we proposed that the hydrophobic microenvironment encompassed by Phe-198 and Phe-200 could be exploited for enhancing the inhibitory potency. This premise was corroborated by a greater potency of (S)-HDAC-42 than that of HTPB (IC50, 16 nM versus 44 nM), of which the α-isopropyl moiety was favorable in interacting with this hydrophobic motif (21) (Figure 2.2). It is noteworthy that (S)-HDAC-42 is currently undergoing preclinical evaluation through the Rapid Access to Intervention Development Program at the National Cancer Institute.

2.5 Antitumor Mechanisms of HDAC Inhibitors The general paradigm for the antitumor action of HDAC inhibitors has been the induction of histone acetylation resulting in transcriptional regulation of critical genes that are involved in cytostasis (cell cycle arrest), differentiation, or apoptosis (76-79) (Table 2.4). However, several lines of evidence suggest that induction of growth arrest and apoptosis in a variety of human cancer cells by HDAC inhibitors cannot be solely attributed to the accumulation of hyperacetylated histones.

Firstly, histone hyperacetylation induced by histone deacetylase inhibitors, including sodium butyrate, phenylbutyrate and suberic bishydroxamate (SBHA), is not sufficient to cause growth inhibition in human dermal fibroblasts, although histone

12 acetylation remains a major target for HDAC inhibitors (10). It also has been reported that certain cell lines, including human dermal fibroblasts and erythroleukemia cells, are able to growth in the presence of HDAC inhibitors and the in the presence of hyperacetylated histones (80,81).

Secondly, various important survival signaling regulatory proteins, such as p53,

Ku70, and Hsp90, have been identified as the nonhistone substrates of HDACs, as mentioned previously. Targeting of the acetylation status of these signal mediators by

HDAC inhibitors may also account for their anticancer activity. For instance, induction of acetylation of the tumor suppressor p53 by HDAC inhibitors can inhibit its ubiquitination by Mdm2 and its degradation by proteasome-dependent proteolysis (82). Accordingly, it has been reported that the HDAC inhibitors, TSA and CG152, can stabilize acetylated p53 and induce cell cycle arrest or apoptosis in prostate cancer cells (14). TSA can also induce apoptosis through the acetylation of Ku70. Acetylation of Lys539 and Lys542 of Ku70 can disrupt the interaction between Ku70 and its binding protein, Bax. This can lead to the release of Bax from Ku70 and the increased mitochondrial localization of Bax, which triggers cytochrome c release, leading to caspase-dependent apoptosis in neuroblastoma cells (15). Furthermore, inhibition of HDAC6 can induce the acetylation of Hsp90 and lead to the lost of its chaperone activity of Hsp90 (67), which could serve as a possible antineoplastic target of HDAC inhibitors that can inhibit the action of HDAC6. Thirdly, it has been reported that HDACs can form protein complexes with many cellular proteins, including the transcription regulators, chaperone 14-3-3 proteins,

α-tubulin, polyubiquitin, and protein phosphatase 1 (PP1), which contribute to their complicated biological functions (83-86). Interestingly, the HDAC inhibitor TSA is able to disrupt the interaction of HDACs and PP1 (87). This suggests that HDAC inhibitors can elicit coordinate changes in cellular protein phosphorylation and acetylation, and that

13 changes in these protein modifications at multiple subcellular sites may contribute to the well documented ability of HDAC inhibitors to suppress cell growth and transformation.

Together these observations demonstrate that the biological effects of HDAC inhibitors can be mediated by events independent of the acetylation of histone. Therefore, the further characterization of HDAC-isoform specific functions and their substrates, as well as a greater understanding of the dynamic protein complexes involving HDAC may provide us new insights for novel anticancer drug development.

Class Enzymea Size Chromosome Subcellular References (amino localization acids) Class I HDAC1 482 1p34.1 Nucleus (35,88) (yRpd3-like) HDAC2 488 6q21 Nucleus (88-90) HDAC3 428 5q31.1-5q31.3 Nucleus/cytoplasm (88,91) HDAC8 377 Xq21.2-Xq21.3 Nucleus (92-94)

Class IIa HDAC4 1084 2q37.2 Nucleus/cytoplasm (95,96) (yHda1-like) HDAC5 1122 17q21 Nucleus/cytoplasm (47,97) HDAC7 855 12q13.1 Nucleus/cytoplasm (97-99) HDAC9 1011 7p15-p21 Nucleus/cytoplasm (88,97,100)

Class IIb HDAC6 1215 Xp11.23 Nucleus/cytoplasm (40,101) (yHda1-like) HDAC10 669 22q13.31-22q13.33 Nucleus/cytoplasm (102-105)

Class III SIRT1 747 10q22.2 Nucleus (106,107) (ySir2-like) SIRT2 373 19q13 Cytoplasm (108-110) SIRT3 399 11p15.5 Mitochondria (111,112) SIRT4 314 12q Unknown (113) SIRT5 310 6p22.3 Unknown (113,114) SIRT6 355 19p13.3 Nucleus (113,115) SIRT7 400 17q Unknown (113,116)

a HDAC11 is homologous to both class I and class II HDACs (41).

Table 2.1: HDAC Family

Protein Intracellular function Deacetylase References

Ku70 Suppresses apoptosis SIRT1, TSA-sensitive HDACs (15,50,68)

FOXO1 Transcription factor SIRT1 (117)

p300 Transcription factor SIRT1 (118)

p53 Tumor suppressor HDAC1, SIRT1 (82,107,11

9-122)

Androgen receptor Hormone receptor HDAC1 (123-125)

Smad7 Signal transducer of TGF-β HDAC1, 3 and 6 (126)

Stat3 Signal transducer of cytokines HDAC1, 2 and 3 (65,127)

NF-κB (RelA) Nuclear transcription factor HDAC3, STR1 (128-131)

SRY Y chromosome-encoded HDAC3 (132)

DNA- binding protein

α-tubulin Microtubule component HDAC6, SIRT2 (73,110,13

3,134)

Hsp90 Molecular chaperone HDAC6 (67,135)

Table 2.2: Representative Nonhistone Substrates of HDACs

Class Representative compounds Pharmacological profiles Potency (IC50) These HDAC inhibitors, alone or in COOH COOH combination with therapeutic agents, have entered different stages of clinical Short-chain Butyrate Phenylbutyrate evaluation. Although well tolerated in fatty acids COOH O O patients, these agents suffer from a short Low (mM) O O plasma half-life due to rapid metabolism, Valproate AN-9 nonspecific mode of action, and the high doses needed for therapeutic effects. H O N NHOH These HDAC inhibitors contain a broad set O SAHA of agents with nM potency against both OH O Class I and II enzymes. The natural product OH trichostatin A (TSA) was the first Hydroxamic N H hydroxamate identified with high potency, acids N HN but is not in clinical use due to toxicity. High (nM) LAQ-824 Synthetic hydroxamate such as O O suberoylanilide hydroxamic acid (SAHA) OH N and LAQ-824 are currently undergoing H N Phase I and/or II clinical trials. TSA O

O N H NH2 H N N These agents inhibit HDAC activity at µM Benzamides MS-275 O concentrations. Representatives include Moderately MS-275 and CI-944, both of which are in High (µM) O HN clinical trials. HN O CI-994 H2N Cyclic peptides constitute the most structurally complex class of HDAC inhibitors. Representatives include O O depsipeptide (FK-228), apicidin A, and N S NH cyclic hydroxamic acid containing peptide H (CHAP). Depsipeptide is metabolically Cyclic peptides NH S O activated via reduction of the High (nM) HN O intramolecular disulfide bond with a O decrease in IC50 from 36 nM to 1.6 nM. O Depsipeptide was in Phase I and II trials for various types of cancer, which, however, suffered a setback due to its cardiovascular toxicity.

Table 2.3: Classification of HDAC inhibitors (136)

Table 2.4: Tumor-Associated Proteins Whose Transcriptional Expression is Altered in Response to HDAC Inhibitor Treatment of Cells (79)

Figure 2.1: Space-Filling Representation of TSA in the Active Site of HDLP (75)

Figure 2.2: Structural Basis for TSA Binding to HDLP Binding Site (136). A, Structure of TSA. B, Molecular Docking of TSA into the Active-Site Pocket of HDLP

Figure 2.3: Molecular Docking of HTPB and (S)-HDAC-42 into the Active-Site Pocket of HDLP (136)

CHAPTER 3

HISTONE ACETYLATION-INDEPENDENT EFFECT OF HISTONE

DEACETYLASE INHIBITORS ON AKT DEPHOSPHORYLATION

3.1 Introduction The current paradigm underlying the antiproliferative effects of HDAC inhibitors has their regulation of the transcription of a defined set of genes through chromatin remodeling. However increasing evidence suggests that modifications of the epigenetic histone code may not represent the only mechanism for HDAC inhibitor-mediated growth inhibition and apoptosis in cancer cells (as reviewed in Chapter 2). Data from this and other laboratories show that HDAC inhibitors can facilitate the dephosphorylation of cell survival signaling proteins involved in cell proliferation and apoptosis, including protein kinase B (Akt) (137-143), extracellular signal-regulated kinase (Erk) (137-139,144), retinoblastoma protein (Rb) (145), and focal adhesion kinase (Fak) (146). However the molecular mechanism underlying this specific effect of HDAC inhibitors remains unclear. It is interesting that HDACs have been shown to form complexes with protein phosphatase 1 (PP1) (86,87), of which the combined deacetylase/phosphatase activities underlie the ability of HDAC1 to modulate transcriptional activity of cAMP-response-element-binding protein (CREB) (86) and that of HDAC6 to regulate microtubule dynamics (87). It is worth noting that the broad specificity HDAC inhibitor,

TSA, can specifically disrupt the interaction between HDACs and PP1, thereby releasing

22 PP1 from these protein complexes (87). Together these data suggest that HDAC inhibitors may elicit coordinate changes in cellular protein phosphorylation and acetylation at multiple cellular sites. In addition, a number of small molecules such as

C2-ceramide, 4-hydroxynonenal, FTY720, and N-ethylmaleimide have been reported to facilitate Akt dephosphorylation by activating PP1 or PP2A (147-151). Consequently, we attempted to establish a potential link between TSA-mediated Akt down-regulation and protein phosphatase activation by HDAC inhibitors in U87MG cells.

In this study, we demonstrate a novel histone acetylation-independent mechanism by which HDAC inhibitors induce Akt dephosphorylation in U87MG glioblastoma cells by disrupting HDAC- PP1 complexes resulting in increased association of PP1 with Akt. In light of the pivotal role of Akt in cell proliferation, the down-regulation of Akt activity through this non-conventional mechanism may contribute to the antitumor activities of HDAC inhibitors. This PP1-facilitated kinase dephosphorylation underscores the diverse functions of HDAC inhibitors in mediating antineoplastic activities at different cellular levels.

3.2 Materials and Methods 3.2.1 Cell Culture U87MG human glioblastoma cells were kindly provided by Dr. Ing-Ming Chiu (The Ohio State University, Columbus, OH), and PC-3 prostate cancer cells were purchased from the American Type Culture Collection (Manassas, VA). As both U87MG and PC-3 cells are PTEN null, they exhibit constitutively active Akt. These cancer cells were cultured in 10% fetal bovine serum (FBS)-supplemented RPMI 1640 medium containing 100 units/ml penicillin and 100 mg/ml streptomycin (Life Technologies, Inc.,

Grand Island, NY).

23 3.2.2 Reagents Trichostatin A (TSA), calyculin A, and okadaic acid were purchased from

Sigma-Aldrich (St. Louis, MO), and tautomycin was obtained from Calbiochem (La Jolla,

CA). The HDAC inhibitors suberoylanilide hydroxamic acid (SAHA) and HDAC-42

[N-hydroxy-4-(3-methyl-2-phenyl-butyrylamino)benzamide] were synthesized in our laboratory. Mouse antibodies against α-tubulin and acetylated α-tubulin were from Sigma-Aldrich (St. Louis, MO). Rabbit antibodies against Akt, phospho-473Ser-Akt, phospho-308Thr-Akt, PDK1, phospho-ERK1/2, phospho-p38, phosphor-JNK, and various HDAC isozymes were purchased from Cell Signaling Technology Inc. (Beverly, MA). Rabbit antibodies against PP1, PP2A, and nucleolin, and mouse antibodies against p21WAF1/CIP1 were from Santa Cruz Biotechnology Inc. (Santa Cruz, CA). Rabbit antibodies against p85, p110, H3, and acetyl-H3 were from Upstate Biotechnology (Lake

Placid, NY). Mouse monoclonal anti-β-actin was from ICN Biomedicals Inc. (Costa Mesa, CA). Rabbit anti-human CTMP antibodies and rabbit anti-TRB3 antibodies were from Alpha Diagnostic International (San Antonio, TX) and Oncogene Research Products (San Diego, CA), respectively. Goat anti-rabbit immunoglobulin G (IgG)-horseradish peroxidase conjugates and rabbit anti-mouse IgG horseradish peroxidase conjugates were from Jackson ImmunoResearch Laboratories (West Grove, PA). The siRNA Transfection Reagent, siRNA Transfection Medium, and siRNAs against HDAC1 and HDAC6, and a control, scrambled siRNA were from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA).

3.2.3 Immunoblotting U87MG cells treated with various concentrations of TSA in 10% FBS supplemented RPMI 1640 medium for 48 h were collected and lysed by NP-40 lysis buffer [50 mM Tris-HCl, pH 7.5, 120 mM NaCl, 1% (v/v) NP-40, 1 mM EDTA, 50 mM

24 NaF, 40 mM β-glycerophosphate, and 1 µg/ml each of aprotinin, pepstatin, and leupeptin]. Protein concentrations of the lysates were determined by using a Bradford protein assay kit (Bio-Rad, Hercules, CA). Equivalent amounts of proteins from each lysate were resolved by SDS-PAGE and then transferred onto Immobilon-nitrocellulose membranes (Millipore, Bellerica, MA) in a semidry transfer cell. The transblotted membrane was washed twice with Tris-buffered saline (TBS) containing 0.1% Tween 20

(TBST). After blocking with TBST containing 5% nonfat milk for 40 min, the membrane was incubated with the appropriate primary antibody in TBST containing 1% nonfat milk at 4 °C overnight. All primary antibodies were diluted in 1% nonfat milk-containing TBST. After treatment with the primary antibody, the membrane was washed two times with TBST for a total of 20 min, followed by goat anti-rabbit or anti-mouse IgG-horseradish peroxidase conjugates for 1 h at room temperature, and washed three times with TBST for a total of 1 h. The immunoblots were visualized by ECL chemiluminescence (Pharmacia, Uppsala, Sweden).

3.2.4 Cell Viability Assay The effect of the HDAC inhibitors on cell viability was assessed by the MTT {[3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide]} assay in 6 replicates. U87MG and PC-3 cells were seeded and incubated in 96-well, flat-bottomed plates in 10% FBS-supplemented RPMI-1640 medium 24 hours before drug treatment. Cells were exposed to individual HDAC inhibitors at the indicated concentrations in 10%

o FBS-supplemented RPMI-1640 medium at 37 C in 5% CO2 for 48 h. The medium was removed and replaced by 200 µl of 0.5 mg/ml of MTT in RPMI-1640 medium, and cells were incubated in the CO2 incubator at 37 oC for 2 h. Supernatants were removed from the wells, and the reduced MTT dye was solubilized with 200 µl DMSO. Absorbance was determined on a plate reader at 570 nm.

25 3.2.5 Flow Cytometry After drug treatment as described above, floating cells were collected, and adherent cells were harvested by scraping. The combined cells were fixed in ice-cold

70% ethanol and storage at –20 oC. Cells were centrifuged, washed with PBS, and incubated in 0.2 M phosphate citrate buffer, pH 7.8 at 37 oC to extract low-molecular-weight DNA. Samples are stained with propidium iodide solution (10 mg/ml) in the presence of RNase A (100 mg/ml) for 30 min at room temperature. Cell cycle phase distributions were determined on a FACScort flow cytometer and analyzed by ModFitLT V3.0 program.

3.2.6 Affinity Purification of HDAC-PP1 Complexes U87MG cells treated with TSA at the indicated concentrations in 10% FBS-supplemented RPMI 1640 medium for 48 h. Control cells received the vehicle treatment of 0.1% (v/v) DMSO. Cells were washed by phosphate-buffered saline, and lysed by the aforementioned NP-40 lysis buffer. Cell extract was incubated with microcystin-LR-Sepharose (Upstate Biotechnology, Lake Placid, NY) at 4 oC overnight on a rotator. After brief centrifugation, Sepharose beads were collected and washed with the lysis buffer four times. Bound proteins were eluted by 50 µl of SDS sample buffer and subjected to SDS-PAGE followed by immunoblotting with appropriate antibodies.

3.2.7 Co-immunoprecipitation of PP1-Akt Complexes U87MG cells were treated with various concentrations of TSA for 48 h, and lysed by the aforementioned NP-40 isotonic lysis buffer with a cocktail of protease inhibitors.

After centrifugation at 13000 x g for 15 min, the supernatants were collected, and incubated with protein A-Sepharose beads (Sigma, St. Louis, MO) for 15 min to eliminate nonspecific binding. The mixture was centrifuged at 1000 x g for 5 min, and

26 the supernatants were exposed to Akt or PP1 antibodies in the presence of protein

A-Sepharose beads at 4 oC for 2 h. After brief centrifugation, protein A-Sepharose beads were collected and washed with the aforementioned lysis buffer 4 times, suspended in 2 x

SDS sample buffer, and subjected to Western blot analysis with antibodies against PP1 or

Akt.

3.2.8 Subcellular Fractionation The nuclear and cytosolic fractions of U87MG cells were prepared by using a

Nuclear/Cytosol Fraction Kit (MBL Co., Watertown, MA) according to the manufacturer’s instruction. In brief, cells were cultured to 50% confluency in T-75 flasks, treated with TSA for 48 hr, and collected by centrifugation. The pelleted cells were resuspended in 0.2 ml of the Cytosol Extraction Buffer A Mix containing DTT and a cocktail of protease inhibitors, and mixed vigorously on a vortex mixer. The suspension was then incubated on ice for 10 minutes, mixed with 11 µl of the Cytosol Extraction Buffer B, and incubated on ice for 1 minute. The lysates were centrifuged at 16,000 x g for 5 min. The supernatant, representing the cytoplasmic fraction, was transferred to a pre-chilled tube, and the pellet was resuspended in 100 ml of the Nuclear Extraction Buffer Mix. The suspension was mixed vigorously on a vortex mixer for 15 s, incubated on ice for 40 minutes, and centrifuged at 16,000 x g for 10 min to collect the nuclear extract in the supernatant fraction.

3.2.9 Ser/Thr Phosphatase Activity The Ser/Thr phosphatase activity was determined by using a nonradioactive, malachite green-based Ser/Thr Phosphatase Assay kit (Upstate Biotechnology, Lake

Placid, NY) according to the manufacturer’s instruction. In brief, 5µg of U87MG cell extracts in the aforementioned lysis buffer were incubated with 175 µM of the

27 phosphopeptide substrate (K-R-pT-I-R-R) in the phosphatase buffer (20 mM MOPS, pH

7.5, 60 mM β-mercaptoethanol, 0.1 M NaCl and 0.1 mg/ml serum albumin) with a total volume of 25 µl. The reaction was incubated in a 96-well plate at room temperature for 10 min, terminated by adding the malachite green solution, and allowed to stand for 15 min to permit color development. Absorption at 650 nm was measured in a microplate reader, and the serine/threonine phosphatase activity was calculated using a standard curve based on free phosphate generated by a standard solution. All experiments were performed in triplicate.

3.2.10 Kinase Assay for Phosphatidylinositol 3-kinase (PI3K) PI3K activity was determined by the use of a PI3K ELISA Assay kit from Echelon Biosciences Inc. (Salt Lake City, UT) according to the manufacturer’s instruction. U87MG cells were cultured to 50% confluency in 10 cm culture dishes, and the medium was removed after TSA treatment. The drug-treated cells were washed with 10 ml per dish of ice-cold Buffer A (137 mM NaCl, 10 mM Tris-HCl, pH 7.4, 1 mM

CaCl2, 1 mM MgCl2, and 0.1 mM sodium orthovanadate) three times, incubated with 1 ml of ice cold Lysis Buffer (Buffer A plus 1% NP-40 and 1 mM PMSF) on ice for 20 min, scraped from the dish, transferred to microcentrifuge tubes, and centrifuged. The supernatant was collected, and protein concentrations were determined by the Bradford assay (Bio-Rad Lab.). Equal amounts of proteins were treated with 5 µl of anti-PI3K antibodies (Upstate Biotechnology, Lake Placid, NY) at 4 °C for 1 h, to which was added

60 µl of a 50% slurry of protein A-agarose beads in PBS, followed by incubation at 4 °C for 1 h with mixing. The immunocomplex was collected by brief centrifugation, washed with Lysis Buffer three times, and incubated for 2 h at room temperature with 10 µl of 10

µM L-α-phosphatidyl-D-myo-inositol 4,5-bisphosphate [PI(4,5)P2], 5 ml of 10X reaction buffer (40 mM MgCl2, 200mM Tris, pH 7.4, 100 mM NaCl, 250 µM ATP), and 35 µl of

28 distilled water. The reaction was terminated by the addition of 2.5 µl of 100 mM EDTA. After brief centrifugation, fifty ml of each reaction mixture was transferred to a 96 well plate, 50 µl of diluted L-α-phosphatidyl-D-myo-inositol 3,4,5-trisphosphate (PIP3) detector was added to each well, and incubated at room temperature for 1 h. 100 µl of reaction mixture from each well was transferred to the corresponding well in the detection plate, and incubated at room temperature for 1 h in the dark. After three washes with 300 µl of TBS-Tween (0.05% v/v), 100 µl of the secondary detection reagent was added to each well followed by incubation for 1 h in the dark, and three more washes with

300 µl of TBS-Tween (0.05% v/v). 100 µl of TMB solution was added to each well and incubated for approximately 20-30 min to allow color development. The reaction was stopped by the addition of 100 µl of stop solution (0.5 M H2SO4) to each well. Absorbance at 450 nm was read, and the PI3K activity in each sample was calculated using a standard curve generated by using a PIP3 standard solution. All experiments were performed in triplicate.

3.2.11 Determination of Phosphoinositide Formation U87MG cells, cultured in T25 flasks, were labeled with 1 mCi/ml of [32P]orthophosphate (HCl-free, DuPont NEN) in Dulbecco modified Eagle phosphate-free and serum-free medium (Gibco) for 4 h, washed three times with Buffer A

(30 mM Hepes, pH 7.2, 110 mM NaCl, 10 mM KCl, 1 mM MgCl2, and 10 mM glucose), and treated with various concentrations of TSA in 10% FBS-supplemented RMPI 1640 medium for 48 h. The cells were extracted with 3 ml of chloroform/methanol (1:2, v/v), followed by 4 ml of chloroform/2.4 M HCl (1:1, v/v), and 1 ml of chloroform four times.

The combined organic phase was dried under a stream of nitrogen, and resuspended in 90

µl of chloroform for thin layer chromatography (TLC) analysis by using a 20 x 20 cm Silica Gel 60 TLC plate (EM Science) impregnated with 1% potassium oxalate in 50 %

29 ethanol. The TLC plate was developed in chloroform/acetone/methanol/acetic acid/water

(80:30:26:24:14, v/v/v/v/v) (23). Radioactive spots were detected by autoradiography using Kodak X-Omat film, and total PIP3 quantified by both densitometry and an AMBIS B scanning system (San Diego, CA), both of which showed comparable results.

3.2.12 Kinase Assay for Phosphoinositide-dependent kinase-1 The phosphoinositide-dependent kinase-1 (PDK1) kinase activity was performed using a PDK-1 kinase assay kit (Upstate Biotechnology, Lake Placid, NY) according to a described procedure (152). This cell-free assay is based on the ability of recombinant PDK-1 to activate its downstream kinase serum- and glucocorticoid-regulated kinase which, in turn, phosphorylates the Akt/serum- and glucocorticoid-regulated kinase-specific peptide substrate RPRAATF with [γ-32P]ATP. The 32P-phosphorylated peptide was then separated from residual [γ-32P]-ATP by P81 phosphocellulose paper, and quantitated by a scintillation counter after three washes with 0.75% phosphoric acid.

3.2.13 Isozyme-specific Knockdown of HDACs with siRNA Isozyme-specific siRNAs were used to attenuate the expression of HDAC 1 and 6 in U87MG cells using reagents obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Transfection of U87MG cells with these siRNA was carried out according to the vendor's instructions. In brief, cells were cultured to 40 - 50% confluency in 10% FBS-supplemented antibiotics-free RPMI 1640 medium. The HDAC1 and HDAC6 siRNAs were transfected to the cells by the siRNA Transfection Reagent (Santa Cruz, CA) to the final concentration of 200 nM. A scramble siRNA was used in parallel experiments as a control. The transfected cells were incubated at 37 oC, and the extents of siRNA-mediated down-regulation of HDAC1 and HDAC6 expression were monitored by immunoblotting analysis at different time intervals. It was found that 30 h exposure to

30 siRNA caused more than 80% repression of HDAC expression, and thus was used through this study.

3.2.14 Immunocytochemical Analysis for PP1 and Phospho-Akt U87MG cells were treated with 0.5 µM TSA in 10% FBS-supplemented RPMI

1640 medium for different time intervals, washed with Dulbecco's PBS, fixed with 4% paraformaldehyde for 30 min at room temperature, and then washed with PBS. For double staining of phospho-Akt and PP1, the cells were permeabilized with 0.1% Triton

X-100 in 1% FBS-containing PBS, and treated with rabbit anti-phospho-473Ser-Akt at 4°C for 24 h, and washed with PBS. Subsequently, the cells were treated with mouse anti-PP1 for 8 h at room temperature, and washed with PBS. For fluorescent microscopy, Alexa Fluor 488 goat anti-mouse and anti-rabbit IgG (Molecular Probe, Inc) were used for conjugating PP1 and phospho-Akt, respectively. The nuclear counter staining was performed using a DAPI-containing mounting medium (Vector, CA) prior to examination. Images of immunocytochemically labeled samples were observed using a Zeiss confocal microscope (LSM510) with an Argon laser and a He-Ne laser, and appropriate filters (excitation wavelength: 488 nm for PP1, 633 nm for phospho -Akt, and 543 nm for DAPI).

3.3. Results 3.3.1 Differential Effects of HDAC Inhibitors on Akt Dephosphorylation To shed light onto the causative relationship between HDAC inhibition and Akt deactivation, we assessed the effect of four different HDAC inhibitors, including

HDAC-42, SAHA, MS-275, and TSA, on various HDAC-related biomarkers (histone H3 acetylation, p21WAF1/CIP1 expression, and tubulin acetylation) in relation to Akt phosphorylation state in U87MG gliobalstoma cells. Of the four HDAC inhibitors

31 examined, HDAC-42 belongs to a novel class of phenylbutyrate-based HDAC inhibitors

(20,21,153), and has an IC50 of 30 nM in inhibiting HDAC activity in nuclear extracts.

The reported IC50 values for TSA, SAHA, and MS-275 are 5 - 15 nM, 120 nM, and 4.8

µM, respectively (154).

Western blot analysis shows that exposure of U87MG cells to these inhibitors led to substantial increases in histone H3 acetylation and p21WAF/CIP1 expression (Figure 3.1).

However, these four inhibitors behaved differently with regard to α-tubulin acetylation, indicating differences in the respective activities in inhibiting the α-tubulin deacetylase HDAC6 (73,133). While HDAC-42 and TSA produced robust hyperacetylation of

α-tubulin at submicromolar concentrations, SAHA was effective at low micromolar concentrations, and MS-275 was totally ineffective, even at 5 µM, in inhibiting α-tubulin deacetylation. Moreover, examination of Akt phosphorylation at both 308Thr and 473Ser indicates that HDAC-42 and TSA at submicromolar concentrations were able to substantially reduce phospho-Akt levels. A similar effect on phospho-Akt was not achieved with SAHA until the concentration reached 5 µM. In contrast, the repressing effect of MS-275 on Akt phosphorylation, even at 5 µM, was marginal. It is noteworthy that there existed an inverse relationship between the levels of acetylated α-tubulin and phospho-Akt, providing a potential link between HDAC6 inhibition and Akt dephosphorylation. This premise was further supported by the differential effect of selective siRNA-mediated knockdown of individual HDAC isozymes on phospho-Akt levels (section 3.3.7).

3.3.2 Selective Dephosphorylation of Signaling Kinases Pursuant to the above finding, a question emerged with regard to the specificity of

HDAC inhibitor-mediated kinase dephosphorylation. Accordingly, we examined the impact of two representative HDAC inhibitors, TSA and SAHA, on the phosphorylation

32 status of ERKs, p38, and JNK MAP kinases versus Akt in U87MG cells (Figure 3.2). As noted, both TSA and SAHA caused a modest decrease in phospho-ERK1/2 levels despite their respective activities in Akt dephosphorylation, while those of phospho-p38 and phospho-JNK remained unaffected. These findings indicate that there existed a certain degree of specificity in HDAC inhibitor-mediated kinase dephosphorylation, suggesting the involvement of a unique signaling mechanism.

3.3.3 Differential Effects of HDAC Inhibitors on Cell Proliferation and Cell Cycle Together, these findings suggest that these four HDAC inhibitors exhibited distinct profiles regarding pharmacological targets, which might underlie differences in their antitumor activities. For example, although these agents were able to up-regulate p21WAF1/CIP1 expression and histone acetylation, two hallmark features in association with intracellular HDAC inhibition, at submicromolar concentrations, their ability to suppress cell proliferation varied by almost an order of magnitude (Figure 3.3). While TSA and HDAC-42 were effective in suppressing proliferation in U87MG cells at submicromolar concentrations, it would require at least 2.5 - 5 µM for SAHA and MS-275 to attain a similar antiproliferative effect. This differential potency paralleled the respective activities in causing Akt dephosphorylation, suggesting a putative link between Akt down-regulation and the antitumor activities of HDAC-42, TSA, and, to a lesser extent, SAHA. Moreover, flow cytometric analyses indicate that the effect of these HDAC inhibitors on growth inhibition was largely attributable to cell cycle arrest. Analyses of cell cycle distribution in drug-treated U87MG cells show that HDAC-42, TSA, and

SAHA induced dose-dependent accumulation of cells in the G2/M phase, accompanied by comparative decreases in the G0/G1 and, in particular, S fractions (Table 3.1). In contrast, MS-275 induced dose-dependent G1 cell cycle arrest in this cell line.

33 3.3.4 TSA-mediated Akt Dephosphorylation Is Not Caused by Changes in the

Expression Level of Proteins Involved in Phospho-Akt Regulation Mechanistically, this Akt dephosphorylation might be mediated through the deactivation of upstream kinases or the activation of downstream phosphatase. To discern the role of transcriptional activation in this drug action, we assessed the expression levels of a series of signaling proteins related to the regulation of Akt signaling pathways in

TSA-treated U87MG cells, which included the p85 regulatory and p110 catalytic subunits of PI3K, PDK-1, Akt, two negative Akt modulators CTMP (155) and TRB3

(156), PP1, and PP2A. As shown in Figure 3.4, TSA exposure did not alter the expression level of any of these signaling proteins, excluding the involvement of transcriptional activation in altering the status of Akt phosphorylation. Furthermore, three lines of evidence argued against the possibility that TSA-mediated Akt deactivation were caused by a decrease in PI3K and PDK-1 kinase activity. First, immunoprecipitated PI3K kinase activity in U87MG cells treated with different doses of TSA remained the same as that of the DMSO control (Figure 3.5; open bars). Second, TSA at different doses exhibited no appreciable inhibitory effects on the kinase activity of recombinant PDK-1 (Figure 3.5; gray bar). Third, autoradiographic

32 analysis of P-labeled phospholipids demonstrated that the level of PIP3, a PI3K lipid product, and other inositol lipids in U87MG cells was unaffected by 0.5 µM TSA treatment (Figure 3.6).

3.3.5 Inhibition of PP1 Prevents TSA-mediated Akt Dephosphorylation In the literature, a number of small molecules such as C2-ceramide,

4-hydroxynonenal, FTY720, and N-ethylmaleimide have been reported to facilitate Akt dephosphorylation by activating PP2A or PP2A-like phosphatase (147-151).

Consequently, we investigated a potential link between TSA-mediated Akt

34 down-regulation and protein phosphatase activation by examining the effect of tautomycin, calyculin A, and okadaic acid on phospho-Akt levels in TSA-treated

U87MG cells. These three compounds exhibit distinct specificity in protein phosphatase inhibition, i.e., tautomycin is a highly specific PP1 inhibitor (157), okadaic acid at low doses (<100 nM) is selective for PP2A(158,159), and calyculin A lacks selectivity between PP1 and PP2A. Figure 7 demonstrates that while none of these inhibitors affected the ability of TSA to inhibit histone deacetylation, the differential effects on the

TSA-mediated dephosphorylation of Akt was noteworthy. As shown, calyculin A (50 nM)

(Figure 3.7; panel A) and tautomycin (5 µM) could completely abrogate the effect of TSA on phospho-Akt, while okadaic acid lacked appreciable protective activity against the dephosphorylation (Figure 3.7; panel B). As PP1 represents a common target for calyculin A and tautomycin, this finding suggests the involvement of PP1 in TSA-facilitated kinase dephosphorylation.

3.3.6 TSA Disrupts HDAC-PP1 Complexes, Resulting in Increased PP1-Akt Associations The link between HDAC inhibition and PP1-mediated kinase dephosphorylation could be attributed to two plausible mechanisms. First, TSA treatment might lead to increased PP1 activity. Second, in light of the reported HDAC-PP1 complex formation (86,87), HDAC inhibitors might cause disruption of these complexes, which would free PP1 to interact with Akt. Of these two mechanisms, the former was refuted by lack of appreciable increases in either the PP1 expression level or the overall Ser/Thr protein phosphatase activity after TSA exposure (Figure 3.8). To evaluate the second possibility, we investigated the expression profile of HDAC isozymes in U87MG cells. Among the seven different isozymes examined, U87MG cells expressed HDACs 1, 3, 5, and 6, but the expression of HDACs 4 and 7 were undetectable. In addition, TSA treatment did not

35 alter the expression level of any of these HDACs (Figure 3.9). To assess the impact of

TSA on the dynamics of HDAC-PP1 complexes, we exposed the lysates of drug-treated

U87MG cells to microcystin affinity beads to purify PP1-associated complexes (86).

Western blot analysis of the affinity-purified PP1 complexes indicates that HDACs 1, 3, and 6 could be pulled-down by the affinity beads. Moreover, the levels of affinity bead-bound HDACs 1 and 6 decreased in response to TSA in a dose-dependent manner

(Figure 13.10). The level of PP1-associated HDAC3, however, remained unaltered by the

TSA treatment.

Based on the report that the PP1 binding domain of HDAC6 encompassed the catalytic motif (87), binding of the HDAC inhibitor to the catalytic domain might hinder the binding of PP1 to HDAC6. Presumably, HDAC inhibitors could also sequester HDAC1, but not HDAC3, from PP1 association through the same mode of mechanism. Also noteworthy is that TSA-mediated dissociation of HDAC-PP1 complexes did not significantly change the relative distribution of PP1 or PP2A in the nucleus or cytoplasm (Figure 3.11). In addition, two lines of evidence indicate that the TSA-mediated disruption of HDAC-PP1 complexes was accompanied by a dose-dependent increase in PP1-Akt associations. First, immunoprecipitation of PP1 in the lysates of TSA-treated cells followed by Western blotting of Akt, or vice versa, demonstrates a positive correlation between TSA doses and amounts of Akt co-immunoprecipitated with PP1 (Figure 3.12). Second, immunocytochemical examinations show that exposure of U87MG cells to TSA for 4 h led to co-localization of PP1 and phospho-Akt, which, however, was not noted with cells receiving DMSO vehicle (Figure 3.13). The physical interaction between PP1 and Akt led to partial and complete dephosphorylation at 8h and

24 h post-treatment, respectively.

36 3.3.7 Validation of the Involvement of HDACs 1 and 6 in Akt Dephosphorylation Together, the above findings suggest that TSA caused PP1-mediated Akt dephosphorylation by sequestering HDACs 1 and 6. To validate the role of these two isozymes in Akt deactivation, we used isozyme-specific siRNAs to selectively knockdown the expression of HDACs 1 and 6, and that of a scrambled siRNA as a negative control (Figure 14; panel A). As shown, repressed expression of HDACs 1 and 6 both led to decreased Akt phosphorylation, while control siRNA transfection had no appreciable effect on phospho-Akt levels (Figure 3.14; panel B).

3.4. Conclusion The present study demonstrates a novel histone acetylation-independent mechanism by which HDAC inhibitors mediate the dephosphorylation of Akt through the disruption of HDAC-PP1 complexes (Figure 3.15). Of the four inhibitors examined, HDAC-42 and TSA were most effective in facilitating Akt dephosphorylation, followed by SAHA, while MS-275 exhibited a marginal effect at therapeutically attainable concentrations (≤ 5 µM). In light of the pivotal role of Akt in cell proliferation, this differential activity in Akt down-regulation might represent the basis for the differences in the antitumor activities among these four HDAC inhibitors. TSA and HDAC-42 were potent in suppressing the proliferation of U87MG cells, in part, due to their ability to down-regulate Akt signaling. It is noteworthy that the activity of these agents to suppress

Akt phosphorylation paralleled the respective potency in inducing α-tubulin acetylation, a biomarker for HDAC6 (73,133). This finding suggests a role of HDAC6 inhibition in

Akt dephosphorylation. Consequently, MS-275 was ineffective in causing Akt deactivation as a result of its inability to inhibit HDAC6.

The present study further demonstrated a mechanistic link between PP1 and

TSA-mediated Akt dephosphorylation. Our immunochemical study showed that TSA

37 blocks specific associations of PP1 with HDACs 1 and 6, suggesting the involvement of both isozymes in regulating the dynamics of PP1 complexes. Subsequent co-immunoprecipitation and immunocytochemical assays revealed that disruption of

HDAC-PP1 complexes leads to increased association of PP1 with Akt, resulting in Akt deactivation. Moreover, we used isozyme-specific siRNAs to confirm the role of HDACs

1 and 6 as key mediators in facilitating Akt dephosphorylation. The effect of reduced abundance of HDACs 1 or 6 on Akt deactivation mimicked that of HDAC inhibitors. We reasoned that both siRNAs and HDAC inhibitors could destabilize PP1-HDAC complexes, resulting in increased association of PP1 with Akt. In conclusion, the ability of HDAC inhibitors to deactivate Akt through the reorganization of PP1 complexes underlines the complexity of the pharmacological functions of these agents. This study represents the first example of modulating specific PP1 interactions by small-molecule agents. From a clinical perspective, identification of this PP1-facilitated dephosphorylation mechanism underscores the potential use of HDAC inhibitors in lowering the apoptosis threshold for other therapeutic agents through Akt downregulation. In light of the clinical application of HDAC inhibitors, a better understanding of this novel histone-independent mechanism will allow the design of more effective strategies for optimizing the use of the agents in cancer treatment and/or prevention.

Table 3.1: Effects of HDAC Inhibitors on Cell Cycle Distribution in PTEN-Null U87MG Glioblastoma Cells

Figure 3.1: Effects of HDAC Inhibitors on Akt Dephosphorylation in PTEN-Null U87MG Glioblastoma Cells

Figure 3.2: A Dose-Dependent Effects of TSA and SAHA on the Phosphorylation State of 308Thr- and 473Ser-Akt vis á vis ERK1/2, p38, and JNK MAP Kinases in U87MG Cells

Figure 3.3: Dose Dependence of the Growth Inhibitory Effects of HDAC Inhibitors in U87MG Cells

Figure 3.4: Akt Dephosphorylation Is Not Due to Alterations in the Expression Level of Proteins Involved in Phospho-Akt Regulation in U87MG Cells

Figure 3.5: TSA Does Not Affect the Enzyme Activities of PI3K and PDK1 Kinases

Figure 3.6: TSA Does Not Affect the PIP3 Level in U87MG Cells

Calyculin A 50 nM 1 Hr

TSA 0 250 500 250 500 nM

p-Akt Thr308

p-Akt Ser473

Akt

p-ERK 1/2

Acetyl-H4

Acetyl-H4 H4 H4

p21 p-p21 p21

Actin

B. TC OA 5 µM 100 nM

TSA 0 0.25 0.5 0. 25 0.5 0.25 0.5 µM

p-Akt Ser473

Akt

p-ERK 1/2

ERK1/2

Actin

Figure 3.7: The Protein Phosphatase Inhibitors Exhibit Differential Effects on TSA-Mediated Dephosphorylation of Akt

PP1/2A

Figure 3.8: TSA Does Not Affect the Enzyme Activities of PP1/2A Phosphatases

TSA 0 1.0 2.5 5.0 µM

HDAC1

HDAC3

HDAC4

HDAC5

HDAC6

HDAC7

Figure 3.9: TSA Treatment Did Not Alter the Expression Level of HDACs

IP:PP1; IB: HDACs

Fig 3.10: TSA Selectively Disrupts the Association of PP1 with HDAC1 and 6 in U87MG Cells

Figure 3.11: TSA Does Not Change the Relative Abundance of PP1 or PP2A in the Nucleus or Cytoplasm in Drug-Treated U87MG Cells

Figure 3.12: TSA Treatment Leads to Increase PP1-Akt Association

Figure 3.13: TSA Treatment Leads to Increase PP1-Akt Association and Akt-Dephosphorylation

Figure 3.14: Validation of the Involvement of HDAC1 or 6 in PP1-Mediated Akt Dephosphorylation

Figure 3.15: Schematic Diagram Depicting the Effect of HDAC Inhibitors on the Dynamics of PP1 Complex Formation with HDACs and Phospho-Akt

CHAPTER 4

ANTITUMOR EFFECTS OF A NOVEL PHENYLBUTYRATE-BASED

HISTONE DEACETYLASE INHIBITOR, (S)-HDAC-42, IN PROSTATE

CANCER

4.1 Introduction HDAC inhibitors can induce growth arrest, differentiation and apoptosis in cancer cells, and suppress tumor growth in mouse models, based on substantial preclinical evidence. Among clinically relevant HDAC inhibitors, phenylbutyrate and other short-chain fatty acids are weak inhibitors of HDACs exhibiting millimolar potency in vitro (160-162). We recently reported our development of a novel class of potent phenylbutyrate-based HDAC inhibitors possessing submicromolar HDAC-inhibitory activity (20). Further structure-based lead optimization culminated in the generation of (S)-HDAC-42, a hydroxamate-tethered phenylbutyrate derivative with a low nanomolar

IC50 for HDAC inhibition (as reviewed in Chapter 2.4) (21). In light of the apparent role of HDACs in prostate carcinogenesis and progression (25,163-165), and the efficacy of HDAC inhibitors in preclinical models of prostate cancer (14,166-169), the antitumor potential of (S)-HDAC-42, in comparison to suberoylanilide hydroxamic acid (SAHA), a

HDAC inhibitor currently in clinical trials, was evaluated in in vitro and in vivo models of human prostate cancer.

We show here that (S)-HDAC-42 is a potent inhibitor of prostate cancer cell viability and induces a greater apoptotic response relative to SAHA. We also provide

55 evidence that this potent apoptogenic activity is associated with the greater ability of

(S)-HDAC-42 to influence multiple regulators of cell survival including the Bcl family proteins, the inhibitor of apoptosis protein (IAP) family proteins, and the histone-acetylation-independent target, Akt. Finally, these in vitro results were extended to an in vivo prostate cancer xenograft model in which orally administered (S)-HDAC-42 potently inhibited tumor growth in association with intratumoral histone hyperacetylation and reductions in phospho-Akt and Bcl-xL levels. Thus, (S)-HDAC-42, a potent orally bioavailable HDAC inhibitor with antitumor activity at multiple cellular levels, may have clinical value in prostate cancer chemotherapy and warrants further investigation in this regard.

4.2 Materials and Methods 4.2.1 Reagents The HDAC inhibitors suberoylanilide hydroxamic acid (SAHA) and (S)-(+)-Nhydroxy-4-(3-methyl-2-phenyl-butyrylamino)benzamide [(S)-HDAC-42] were synthesized in our laboratory with purities exceeding 99% as demonstrated by NMR spectroscopy (300 MHz). (S)-HDAC-42 (NSC 736012) is a novel hydroxamate-tethered phenylbutyrate derivative(20,21), and is currently undergoing preclinical evaluation through the Rapid Access to Intervention Development Program at the National Cancer Institute. For in vitro studies, stock solutions of inhibitors were prepared in DMSO and diluted in 10% serum-containing culture medium for treatment of cells (final concentration of DMSO, <0.1%). For in vivo studies, (S)-HDAC-42 and SAHA were prepared as suspensions in vehicle (0.5% methylcellulose w/v, 0.1% Tween 80 v/v, in sterile water) for oral administration to xenograft-bearing athymic nude mice. Rabbit antibodies against Akt, phospho-473Ser-Akt, phospho-308Thr-Akt, phospho-136Ser-Bad, Bad, Bax, Bcl-xL, Bcl-2, caspase 9 and poly (ADP-ribose) polymerase (PARP) were

56 purchased from Cell Signaling Technology, Inc. (Beverly, MA). Mouse antibodies against p21WAF1/CIP1 (clone F-5) and rabbit antibodies against cytochrome c (H-104), ILP

(H-202) and cIAP-2 (H-85) were obtained from Santa Cruz Biotechnology, Inc. (Santa

Cruz, CA). Rabbit antibodies against cIAP-1 and surviving were purchased from R&D

Systems Inc. (Minneapolis, MN) and against acetylated histone H3 from Upstate

Biotechnology, Inc. (Lake Placid, NY). Mouse antibodies against β-actin were obtained from ICN Biomedicals (Irvine, CA).

4.2.2 Cell Culture The androgen-insensitive PC-3 and DU-145, and androgen-sensitive LNCaP human prostate cancer cell lines were purchased from American Type Culture Collection (Manassas, VA) and cultured in RPMI-1640 medium (Gibco, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS; Gibco). Normal prostate epithelial cells (PrECs) were obtained from Cambrex Bioscience-Walkersville, Inc. (Walkersville, MD) and maintained in the vendor’s recommended defined prostate epithelial growth medium. All cell types were cultured at 37°C in a humidified incubator containing 5%

CO2. Cells in log phase growth were harvested by trypsinization for use in viability assays and in vivo studies.

4.2.3 Cell Viability Assay Cell viability was assessed by using the 3-(4,5-dimethylthiazol-2-yl)-2,5

-diphenyl-2H-tetrazolium bromide (MTT; Tokyo Chemical Industry, Inc, Tokyo, Japan) assay in six replicates (96-well format). Cancer cells and PrECs were seeded at 2,500 to

3,000 and 8,000 cells per well, respectively, in 96-well flat-bottomed plates. Twenty-four hours later cells were treated with HDAC inhibitors at the indicated concentrations in

10% FBS-supplemented RPMI-1640 medium or 10% FBS-prostate epithelial growth

57 medium. The controls received DMSO at a concentration equal to that in drug-treated cells. At the indicated intervals, 1/10 volume of 10X MTT (5 mg/mL) was added to each well and cells were incubated at 37 °C for 2 hours. Medium was removed and the reduced

MTT dye was solubilized in 200 µL/well DMSO. Absorbance was determined at 570 nm.

Concentrations of compounds that inhibited viability by 50% (IC50) were determined by the median-effect method of Chou and Talalay(170) using CalcuSyn software (Biosoft,

Ferguson, MO).

4.2.4 Apoptosis Assay Drug-induced apoptotic cell death was assessed using the Cell Death Detection ELISA kit (Roche Diagnostics, Mannheim, Germany), which quantitates cytoplasmic histone-associated DNA fragments in the form of mononucleosomes or oligonucleosomes. Cells were seeded and incubated at 50,000 cells/well in 12-well flat-bottomed plates in 10% FBS-supplemented RPMI-1640 medium. After 24 h, cells were treated with HDAC inhibitors at the indicated concentrations and duration. Both floating and adherent cells were collected and the assay was performed according to manufacturer’s instructions.

4.2.5 Immunoblotting Lysates of PC-3 cells treated with HDAC inhibitors at the indicated concentrations for 72 h were prepared for immunoblotting of acetylated histone H3, p21WAF1/CIP1, β-actin, cytochrome c, caspase 9, PARP, phosho-473Ser-Akt, Akt, phosho-136Ser-Bad, Bad, Bcl-xL, Bax, survivin, cIAP-1 and -2, and ILP. Western blot analysis was performed as previously reported (171). For immunoblotting of biomarkers in PC-3 tumor xenografts, tumor tissue homogenates were prepared and immunoblotting performed as previously described (152).

58 4.2.6 Semiquantitative RT-PCR Semiquantitative RT-PCR was performed on total RNA isolated from PC-3 cells treated for 72 hours with the indicated concentrations of (S)-HDAC-42 or SAHA using the RNeasy Mini Kit (Qiagen Inc., Valencia, CA). Following DNase I treatment, 1 µg RNA was reverse-transcribed using Omniscript RT (Qiagen) and oligo(dT). The minimum number of PCR cycles required to obtain a clear signal from the amplicon within the linear amplification phase was determined for each primer set. Following specific primer pairs were used: 5’-GCA TTG TTC CCA TAG AGT TCC-3’ and 5’-CAT

GGC AGC AGT AAA GCA AG-3’ for Bcl-xL; 5’-AAG AAC TGG CCC TTC TTG GA-3’ and 5’-CAA CCG GAC GAA TGC TTT TT-3’ for survivin; and 5’-TCT ACA

ATG AGCTGC GTG TG-3’, and 5’-GGT CAG GAT CTT CAT GAG GT-3’ for β-actin

4.2.7 In Vivo Studies Intact male NCr athymic nude mice (5-7 weeks of age) were obtained from the National Cancer Institute (Frederick, MD). The mice were group housed under conditions of constant photoperiod (12 hours light/12 hours dark) with ad libitum access to sterilized food and water. All experimental procedures utilizing these mice were performed in accordance with protocols approved by the Institutional Laboratory Animal Care and Use Committee of The Ohio State University.

Each mouse was inoculated s.c. with 5 x 105 PC-3 cells in a total volume of 0.1 ml serum-free medium containing 50% Matrigel (BD Biosciences, Bedford, MA) under isoflurane aneththesia. As tumors became established (mean starting tumor volume,

145.1 ± 22.1 mm3), mice were randomized to five groups (n = 7) that received the following treatments: (a) (S)-HDAC-42 at 25 mg/kg body weight q.d., (b) (S)-HDAC-42 at 50 mg/kg q.d., (c) (S)-HDAC-42 at 50 mg/kg q.o.d., (d) SAHA at 50 mg/kg q.d., and (e) the methylcellulose/Tween 80 vehicle. Mice received treatments by gavage (10 µl/g body

59 weight) for the duration of the study. Tumors were measured weekly using calipers and their volumes calculated using a standard formula: width2 × length × 0.52. Body weights were measured weekly. At the conclusion of the study, tumors were harvested, snap-frozen in liquid nitrogen, and stored at -80°C until needed for Western blot analysis of relevant biomarkers. The percent reductions in tumor growth were calculated using the formula: [1 - (Tf - Ti)/(Cf - Ci)] × 100, where Tf and Ti are the final and initial mean tumor volumes, respectively, of the group receiving a specified treatment, and Cf and Ci are the final and initial mean tumor volumes, respectively, of the control group. One mouse from each treatment group was submitted to The Ohio State University Mouse Phenotyping Shared Resource for evaluation of gross and histological pathology.

4.2.8 Statistical Analysis Tumor volume data met the assumptions of normality and homogeneity of variance for parametric analysis; thus, group means at 28 days of treatment were compared using one-way ANOVA followed by the Fisher’s least significant difference (LSD) method for multiple comparisons. Tumor growth data are expressed as mean tumor volumes ± SE. In vitro data are expressed as means ± SD and were assessed by one-way ANOVA and Fisher’s LSD method for post-hoc comparisons. Differences were considered significant at P < 0.05. Statistical analysis was performed using SPSS for Windows (SPSS, Inc., Chicago, IL)

4.3 Result

4.3.1 (S)-HDAC-42 Induces Apoptosis in Prostate Cancer Cells. The antitumor effects of (S)-HDAC42 with reference to SAHA were assessed in three different human prostate cancer cell lines, PC-3, DU-145 and LNCaP. These cells were exposed to individual agents within the dose range of 0.1 to 2.5 µM, and the cell

60 viability was determined by the MTT assay at 24, 48, and 72 h. Both agents showed a time-dependent reduction in cell viability. Neither (S)-HDAC-42 nor SAHA showed significant antiproliferative effect until 48 h, and (S)-HDAC42 clearly was more potent than SAHA in suppressing cell viability in all cell lines evaluated (Figure 4.1). These three cell lines were comparably susceptible to the antiproliferative effect of

(S)-HDAC-42 with submicromolar IC50 values after 72 h of treatment (PC-3, 0.48 ± 0.02; LNCaP, 0.39 ± 0.07; DU-145, 0.43 ± 0.05 µM; P > 0.8, ANOVA), irrespective of the difference in androgen responsiveness, the functional status of p53 and PTEN, and Bax and Bcl-xL expression levels that distinguish these cell lines. On the other hand, SAHA exhibited a statistically significant effect on growth inhibition among all the cell lines tested, with DU-145 cells being the most sensitive (IC50, 1.62 ± 0.16 µM), followed by

LNCaP and PC-3 cells for which IC50 values were extrapolated to be 2.20 ± 0.32 and 3.10 ± 0.16 µM, respectively (P < 0.05, ANOVA). This discrepancy in cellular sensitivity might reflect the differences in mechanisms underlying the antiproliferative effects of these two agents as described below. Assessment of effects on non-malignant cells revealed that PrECs exhibited a 7.5- to 9.5-fold lower sensitivity to the antiproliferative effects of (S)-HDAC-42 than the prostate cancer cells (IC50, 3.69 ± 0.96 µM; P < 0.001, ANOVA; Figure 4.2). It is noteworthy that although neither agent showed a sizeable inhibition of cell proliferation at 24 h of treatment, particularly in PC-3 cells, morphological changes were evident. As shown in Figure 4.3, 24 hours’ exposure of PC-3 cells to 1 µM (S)-HDAC-42 transformed the typical epithelial morphology to a more spindle-shaped morphology (top left and middle panels). These drug-treated cells appeared severely contracted with diminished cell-to-cell contacts and long spine-like processes. Similar morphological changes were also observed in PC-3 cells treated with 2.5 µM SAHA, though to a lesser extent (top right panel). These changes were also apparent in DU-145 (middle panels) and

61 LNCaP cells (bottom panels), though the typical spindle-like morphology of untreated

LNCaP cells made such changes in them more difficult to discern. Although the driving force underlying these morphological changes remain unclear, these findings indicate a clear impact of drug treatment on prostate cancer cells, which preceded the observable effects on cell viability and/or apoptosis.

Western blot analysis of the status of p21WAF1/CIP1 expression and histone H3 acetylation in drug-treated cells revealed dose-dependent up-regulation of these

HDAC-associated biomarkers (Figure 4.4A), which paralleled their activities in inhibiting cell proliferation. Of these two HDAC inhibitors, (S)-HDAC 42 exhibited higher potency in stimulating p21WAF1/CIP1 up-regulation and histone H3 hyperacetylation (Figure 4.4B). In addition, as evidenced by mitochondrial cytochrome c release, activation of caspase 9 and cleavage of PARP (Figure 4.4A), and DNA fragmentation (Figure 4.5), the antiproliferative activity of (S)-HDAC42 was, at least in part, attributable to apoptosis. Moreover, these changes in apoptotic markers were markedly higher than those observed in the SAHA-treated PC-3 cells, consistent with the greater potency of (S)-HDAC-42.

4.3.2 (S)-HDAC-42 Facilitates the Dephosphorylation of Akt and Alters the Dynamics of Bcl-xL Expression. Previously, we reported that (S)-HDAC-42 and, to a lesser extent, SAHA caused Akt dephosphorylation in U87MG and PC-3 cells through the disruption of

HDAC-protein phosphatase (PP1) complexes, leading to increased PP1-Akt association

(Chapter 3) (171). This finding indicates the complexity in the mechanism of

(S)-HDAC-42’s antitumor effects and suggests a basis for the difference in the apoptogenic potencies of these two compounds. Moreover, the modulation of the expression of Bcl-2 family members by several HDAC inhibitors has been reported in a

62 variety of cancer cell lines including mesothelioma, prostate cancer, melanoma, leukemia and lung cancer (172-176). To shed light onto the mechanism underlying the differential antiproliferative activities of (S)-HDAC-42 and SAHA, the effects of these agents on the status of Akt signaling and expression levels of Bcl-2 family members in PC-3 cells were compared.

Western blot analysis revealed that (S)-HDAC-42, even at 1 µM, caused substantial reductions in the levels of phospho-473Ser- and phospho-308Thr-Akt (64.4 ± 11.9% and 61.4 ± 14.8% reductions, respectively, compared with DMSO-treated controls) in PC-3 cells after 72 hours of exposure (Figure 4.6A and B). In contrast, SAHA required at least 5 µM to attain similar levels of Akt dephosphorylation (48.9 ± 12.7 and 58.0 ± 15.6%, respectively) (Figure 4.6A and B). To validate the functional consequence of diminished phospho-Akt, we examined the phosphorylation status of Bad, an apoptosis-relevant substrate of Akt. In parallel with phospho-Akt levels, the amount of phospho-Bad was significantly reduced by (S)-HDAC-42 without changes in the expression of total Bad protein (Figure 4.6A and B). Furthermore, (S)-HDAC-42 caused a pronounced, dose-dependent attenuation in Bcl-xL expression achieving a 75.4 ± 2.5% reduction at 2.5 µM (Figure 4.7A, B). In contrast, SAHA failed to reduce Bcl-xL protein levels below 82.8 ± 2.8% of the control level at the highest concentration tested (Figure 4.7A, B). This clear difference in the abilities of SAHA and (S)-HDAC-42 to suppress in Bcl-xL protein expression was mirrored by their respective effects on steady state levels Bcl-xL mRNA. Semi-quantitative RT-PCR revealed that SAHA reduced Bcl-xL mRNA levels by 9.5 ± 1.7, 9.3 ± 1.3 and 10.2 ± 6.0% at 0.5, 2.5 and 5.0 µM, respectively, compared to vehicle-treated controls, while (S)-HDAC-42 more potently suppressed mRNA levels by 16.7 ± 9.0, 38.9 ± 3.3 and 46.4 ± 4.4% at 0.25, 1.0 and 2.5 µM, respectively (Figure 4.8A, B). (S)-HDAC-42 also caused a nearly 2-fold increase in Bax expression level, while no detectable increase occurred in SAHA treated PC-3 cells

63 (Figure 4.7A). These changes in pro-apoptotic Bcl-2 family members were accompanied by increases in the expression levels of anti-apoptotic Bcl-2 protein in cells treated with either agent. (S)-HDAC-42 treatment induced a 1.6- to 4.9-fold increase in the amount of

Bcl-2 protein over the concentration range tested, while SAHA caused a 2.2-fold increase at 5 µM (Figure 4.7B).

4.3.3 (S)-HDAC-42 Attenuates Protein Levels of IAP Family Members. Members of the IAP family of proteins have been implicated in oncogenesis, cancer progression and therapeutic resistance, at least in part, through their well-known anti-apoptotic function of inhibiting caspase activity (177,178). Several reports have described the suppression of IAP family members by treatment with HDAC inhibitors in cancer cells (138,175,176,179-182), as well as tumor endothelium (181). Accordingly, we sought to determine the effects of (S)-HDAC-42 and SAHA on the suppression of IAP members, including survivin, cIAP-1, cIAP-2 and ILP in PC-3 cells. As shown in Figure 9A and 9B, (S)-HDAC-42 induced a profound, dose-dependent loss of survivin that ranged from a 64.6 ± 6.2% reduction at 0.25 µM up to 93.6 ± 2.5% at 2.5 µM. SAHA caused a similar, though less potent, effect on survivin expression, diminishing it at all concentrations tested up to a 64.1 ± 7.6% reduction after treatment with 5.0 µM. Although less robust, decrements in survivin were also observed at the level of mRNA. The highest concentrations of (S)-HDAC-42 and SAHA tested (2.5 and 5.0 µM, respectively) reduced steady-state levels of survivin mRNA to 67.1 ± 1.7 and 81.9 ± 1.1% of that of the vehicle-treated controls (Figure 8A and 8B). Of the other IAPs evaluated

(Figure 4.9A), cIAP-2 was the most responsive to either drug exhibiting 1.7- and 2.5-fold reductions after treatment with 5.0 and 2.5 µM of SAHA and (S)-HDAC-42, respectively.

(S)-HDAC-42 also induced a 2-fold inhibition of cIAP-1 at 2.5 µM; however, neither agent altered the expression of ILP.

64 4.3.4 (S)-HDAC-42 Suppresses Prostate Tumor Xenograft Growth In Vivo. Collectively, the in vitro data described above indicate that (S)-HDAC-42 is a potent inhibitor of prostate cancer cell proliferation and survival, and owes this antitumor efficacy to the modulation of apoptotic machinery at multiple levels, including Akt signaling, mitochondrial integrity and caspase activation. To further evaluate the antitumor potential of (S)-HDAC-42, athymic nude mice bearing established subcutaneous PC-3 tumor xenografts (mean tumor volume, 145.1 ± 22.1 mm3) were treated orally for 28 days with (S)-HDAC-42 at 25 mg/kg daily or 50 mg/kg every other day, with SAHA at 50 mg/kg daily, or with vehicle. As shown in Figure 4.10A, treatment of mice with SAHA, 25 mg/kg of (S)-HDAC-42 daily and 50 mg/kg of (S)-HDAC-42 every other day significantly inhibited PC-3 tumor growth by 31%, 52% and 67%, respectively, relative to vehicle-treated controls (P < 0.01). Moreover, at 50 mg/kg every other day, the antitumor effect of (S)-HDAC-42 was significantly greater than that of SAHA at 50 mg/kg daily (P < 0.001). This in vivo efficacy after oral administration confirmed (S)-HDAC-42’s oral bioavailability, which was previously determined to be 26%. Importantly, mice appeared to tolerate all of the agents without overt signs of toxicity, without significant changes in body weight compared to the vehicle-treated group, and without abnormalities in serum chemistry variables. The sole gross and histopathological findings associated with (S)-HDAC-42 treatment were testicular atrophy and a marked diffuse testicular degeneration with associated epididymal hypospermia or aspermia. These testicular changes were not observed in the

SAHA-treated mice.

To correlate this in vivo tumor-suppressive response to mechanisms identified in vitro, the effects of treatment with HDAC inhibitors on intratumoral biomarkers of drug activity were evaluated by immunoblotting of PC-3 tumor homogenates collected after

28 days of treatment. As shown in Figure 4.10B and 10C, treatment of mice with

65 (S)-HDAC-42 or SAHA increased levels of acetylated histone H3 in PC-3 tumor xenografts relative to the vehicle-treated controls, thus confirming HDAC inhibition in vivo. The effects of these agents on Akt dephosphorylation and Bcl-xL downregulation approximated those observed in vitro, which reflected their differential tumor suppressive activities in vivo. Treatment with (S)-HDAC-42 at 25 mg/kg per day and 50 mg/kg every other day induced marked reductions in intratumoral levels of both phospho-Akt (69.8 ± 22.1 and 76.8 ± 7.4% reductions, respectively, compared to vehicle-treated controls) and Bcl-xL (72.6 ± 13.2% and 90.9 ± 3.4% reductions, respectively). SAHA treatment (50 mg/kg/day), however, caused much smaller changes in these biomarkers, reducing phospho-Akt by 21.6 ± 15% and Bcl-xL by 23.6 ± 7.8%. In contrast to the marked suppression of survivin by (S)-HDAC-42 and SAHA in vitro, its in vivo suppression by these agents was less dramatic. (S)-HDAC-42 at 25 mg/kg per day and 50 mg/kg every other day reduced survivin expression by 42.8 ± 14.5% and 56.5 ± 25.6%, respectively, while SAHA diminished it by only 17.3 ± 20.3%.

4.4. Conclusion In this study, we describe the in vitro and in vivo efficacy of a novel phenylbutyrate-derived HDAC inhibitor, (S)-HDAC-42, in prostate cancer. Our findings show that (S)-HDAC-42 exhibits a broad spectrum of antitumor activities at low micromolar concentrations that involve not only histone acetylation, but also the modulation of apoptotic regulators at multiple levels, including Akt signaling, mitochondrial integrity and caspase activation.

In comparison to SAHA, a HDAC inhibitor currently in clinical trials,

(S)-HDAC-42 exhibited greater antiproliferative activity in multiple prostate cancer cell lines in vitro. The greater potency of (S)-HDAC-42 to induce apoptosis, as evidenced by cytochrome c release, activation of caspase 9, PARP cleavage and DNA fragmentation,

66 appeared to underlie this higher antitumor efficacy. Moreover, the marked differences in apoptosis induction between these two agents were paralleled by similarly differences in their relative abilities to decrease Akt phosphorylation, to stimulate Bax expression and to suppress expression levels of survivin and particularly Bcl-xL in vitro. This ability of

(S)-HDAC-42 to affect multiple regulators of cancer cell survival is reflected in its equipotent antiproliferative effects against three prostate cancer cells lines that differ with respect to the functional status of p53 and PTEN and Bcl-xL expression level, and suggests its potential clinical efficacy against molecularly heterogeneous tumors.

The correlation between the differential modulation of apoptotic signaling targets and in vitro antitumor activities of (S)-HDAC-42 and SAHA was mirrored in our in vivo study, in which (S)-HDAC-42 exhibited higher potency than SAHA in suppressing established PC-3 xenograft tumor growth. Western blot analysis of the tumor samples revealed that the greater tumor growth inhibition of (S)-HDAC-42 paralleled its greater ability to inhibit putative histone acetylation-independent biomarkers, particularly Akt phosphorylation and Bcl-xL expression. In conclusion, our results show that the novel orally bioavailable, phenylbutyrate-derived HDAC inhibitor, (S)-HDAC-42, is a potent inhibitor of HDAC, as well as targets regulating multiple aspects of cancer cell survival including Akt signaling, mitochondrial integrity and caspase activity. This broad spectrum of activity underlies the more potent apoptogenic and antitumor activities of (S)-HDAC-42 in vitro and in vivo relative to SAHA, and suggests its viability as part of a therapeutic strategy for prostate cancer.

Figure 4.1: Antiproliferative Effects of (S)-HDAC-42 and SAHA in Three Different Prostate Cancer Cell Lines

Figure 4.2: Antiproliferative Effects of (S)-HDAC-42 and SAHA in Normal Prostate Epithelial Cells, PrECs

Figure 4.3: Morphological changes in PC-3, DU-145 and LNCaP Cells Treated with Vehicle Control (left panels), 1 µM (S)-HDAC-42 (central panels) or 2.5 µM SAHA (right panels) for 24 Hours. Original magnification, 100X

Figure 4.4: Effects of (S)-HDAC-42 versus SAHA on Various Biomarkers Associated with HDAC Inhibition or Apoptosis in PC-3 Cells. A. western blot analysis of the dose-dependent effects of (S)-HDAC-42 and SAHA on p21WAF1/CIP1 expression, histone H3 acetylation, cytochrome c release, caspase 9 activation, and PARP cleavage in PC-3 cells. B. Signals for p21WAF1/CIP1 and acetylated histone H3 were quantitated by densitometry and normalized against that of β-actin.

Figure 4.5: Effects of (S)-HDAC-42 versus SAHA on Apoptosis in PC-3 Cells. DNA fragmentation was quantitatively measured by a cell death detection ELISA kit. Each bar represents a mean ± S.D. (n = 3)

Figure 4.6: Dose-Dependent Effects of (S)-HDAC-42 versus SAHA on the Phosphorylation State of 473Ser- and 308Thr-Akt and 136Ser-Bad in PC-3 Cells. A. representative immunoblots B. relative expression levels were quantitated by densitometry and normalized to those of β-actin. Each bar represents a mean ± S.D. (n = 3).

Figure 4.7: Dose-Dependent Effects of (S)-HDAC-42 versus SAHA on the Expression Levels of Bad, Bcl-xL, Bcl-2, and Bax in PC-3 Cells. A. representative immunoblots B. relative expression levels were quantitated by densitometry and normalized to those of β-actin. Each bar represents a mean ± S.D. (n = 3)

survivin

Bcl-xL

Actin

120 120

Survivin Bcl-xL / actin 100 100

80 80

60 60

40 40 % Relative ratio Relative % ratio Relative %

20 20

0 0 DMSO0.52.550.2512.5 DMSO 0.5 2.5 5 0.25 1 2.5 SAHA (s)-HDAC-42 SAHA (s)-HDAC-42

Figure 4.8: Dose-Dependent Effects of (S)-HDAC-42 versus SAHA on the Transcription of Survivin and Bcl-xl in PC-3 Cells. A. representative images B. relative expression levels were quantitated by densitometry and normalized to those of β-actin. Each bar represents a mean ± S.D. (n = 3)

Figure 4.9: Dose-Dependent Effects of (S)-HDAC-42 versus SAHA on the Expression Levels of Members of the IAP Family of Proteins Including Survivin, cIAP-1, cIAP-2, and ILP, in PC-3 Cells. A. representative immunoblots B. relative expression levels were quantitated by densitometry and normalized to those of β-actin. Each bar represents a mean ± S.D. (n = 3)

Fig 4.10: Effects of Oral (S)-HDAC-42 at 25 mg/kg per day (q.d.) or 50 mg/kg every other day (q.o.d.) versus Oral SAHA at 50 mg/kg per day on The Growth of Established PC-3 Tumors in Nude Mice and the Expression of Intratumoral Biomarkers of Drug Activity

CHAPTER 5

EVALUATION OF THE DNA DOUBLE-STRANDED BREAK REPAIR

FUNCTION OF ACETYLATED KU70

5.1 Introduction Ku70, an autoantigen recognized by the sera of patients with scleroderma polymyositis overlap syndrome (183), is the crucial component of the non-homologous end joining (NHEJ) machinery involved in DNA double-stranded break (DSB) repair (184,185). In cooperation with Ku80, Ku70 binds and bridges two proximal broken DNA ends, which facilitates DNA-end joining through a cascade of reactions that involve DNA-dependent protein kinase and DNA ligase IV. Ku70 contains two DNA-binding domains at N- and C-termini, both of which are required for the high binding affinity to DNA (186-189). Recent evidence indicates that the C-terminus of Ku70 also binds Bax, and suppresses its apoptotic translocation to mitochondria (190,191). Consequently, Ku70 mediates its cytoprotective function though two distinct mechanism, i.e., DNA repair and maintenance of mitochondrial integrity (192). Increased Ku70 activity, therefore, would simultaneously enhance the ability of cells to repair the DNA damage and reduce the tendency to initiate Bax-mediated apoptosis. The dual activity of Ku might be regulated at both transcriptional and post-translational levels in response to apoptotic stimuli.

It is noteworthy that eight lysine residues in Ku70 are targets for acetylation by the HATs and HDACs in vivo (50,68). Five of these, K539, K542, K544, K533 and K556,

78 lie in the C-terminal linker domain adjacent to the Bax interaction domain. Mimicking acetylation of K539 or K542, or treating cells with the HDAC inhibitors abolishes the ability of Ku70 to suppress Bax-mediated apoptosis (15,68). Although acetylation also occurs in Ku70’s DNA binding domain (68), defining the consequent effect on DNA

DSB repair remains elusive. According to the structure of human Ku70 (193,194), the positively charged lysine residues in the DNA-binding domains of Ku70 are crucial for the interaction between Ku70 and phosphate groups on the DNA backbone. It has been well demonstrated that acetylation at the lysine residues of DNA binding proteins can block the interaction of these proteins with DNA by interrupting their electrostatic interactions (59,60,195,196). Moreover, several recent reports have shown that HDAC inhibitors can sensitize cancer cells to DNA damaging agents by suppressing their DNA repair activity (197-201). However, the detailed molecular mechanism for this mode of action of HDAC inhibitors remains undefined. Here, we hypothesized that HDAC inhibitors mediate this radio- and chemo-sensitization through the modulation of the acetylation status of Ku70. By using the Bax-deficient DU-145 cells and site-directed mutagenesis targeted to the aforementioned those lysine residues on Ku70, we demonstrate a histone deacetylation-independent mechanism by which HDAC inhibitors sensitize prostate cancer cells to DNA damaging agents by targeting Ku70 acetylation. Our data indicate that pretreatment of the Bax-deficient DU-145 cells with HDAC inhibitors, including TSA, SAHA, MS-275, and (S)-HDAC-42, led to increased Ku70 acetylation irrespective of differences in their pharmacological profiles in HDAC inhibition. Although this hyperacetylation did not affect the Ku heterodimer formation, it reduced Ku70’s DNA end-binding affinity, and diminished the cellular capability to repair drug-induced DNA

DSBs, as evidenced by increased phosphorylation of histone H2AX (γH2AX).

79 5.2 Materials and Methods

5.2.1 Cell Culture DU145, LNCaP, and PC-3 prostate cancer cells were purchased from the

American Type Culture Collection (Manassas, VA). These cancer cells were cultured in

10% fetal bovine serum (FBS)-supplemented RPMI 1640 medium containing 100 units/ml penicillin and 100 mg/ml streptomycin (Life Technologies, Inc., Grand Island,

NY).

5.2.2 Reagents The HDAC inhibitors suberoylanilide hydroxamic acid (SAHA), MS-275, and (S)-HDAC-42 were synthesized in the authors’ laboratory. Trichostatin A (TSA) and bleomycin were purchased from Sigma-Aldrich (St. Louis, MO), and VP-16 (etoposide) and doxorubicin were purchased from Calbiochem (San Diego, CA). Antibodies against various proteins were obtained from the following sources. Mouse monoclonal antibodies: Ku70, Flag, α-tubulin and acetylated α-tubulin, Sigma-Aldrich (St. Louis, MO); p21WAF1/CIP1 and nucleolin (C23), Santa Cruz Biotechnology Inc. (Santa Cruz, CA); phosho-139Ser-histone H2AX (γ–H2AX), Upstate Biotechnology (Lake Placid, NY); β-actin, ICN Biomedicals Inc. (Costa Mesa, CA). Rabbit polyclonal antibodies: Ku80 and pan-acetylated-lysine, Santa Cruz; Bax and Bcl-xL, Cell Signaling Technology Inc. (Beverly, MA); histone H3 and acetyl-histone H3, Upstate Biotechnology. Agarose-conjugated goat polyclonal anti-Ku70 antibodies were purchased from Santa

Cruz, and goat anti-rabbit IgG-horseradish peroxidase conjugates and rabbit anti-mouse

IgG horseradish peroxidase conjugates were from Jackson ImmunoResearch

Laboratories (West Grove, PA). Anti-mouse IgG-Alexa Fluor 488 antibodies were purchased from Molecular Probes (Eugene, OR). Plasmids encoding various Ku70

80 mutants were generated by site-directed mutagenesis of pCMV2B Flag-tagged Ku70

(190) by using a QuickChange site-directed mutagenesis kit from Stratagene.

5.2.3 Immunoblotting Cells treated with various concentrations of HDAC inhibitors in 10% FBS supplemented RPMI 1640 medium for various time intervals were collected and lysed by

NP-40 lysis buffer [50 mM Tris-HCl, pH 7.5, 120 mM NaCl, 1% (v/v) NP-40, 1 mM

EDTA, 50 mM NaF, 40 mM β-glycerophosphate, and 1 µg/ml each of aprotinin, pepstatin, and leupeptin]. Protein concentrations of the lysates were determined by using a Bradford protein assay kit (Bio-Rad, Hercules, CA). Equivalent amounts of proteins from each lysate were resolved by SDS-PAGE and then transferred onto nitrocellulose membranes (Millipore, Bellerica, MA) in a semidry transfer cell. The transblotted membrane was washed twice with Tris-buffered saline (TBS) containing 0.1% Tween 20 (TBST). After blocking with TBST containing 5% nonfat milk for 40 min, the membrane was incubated with an appropriate primary antibody in TBST containing 1% nonfat milk at 4 °C overnight. All primary antibodies were diluted in 1% nonfat milk-containing TBST. After treatment with the primary antibody, the membrane was washed two times with TBST for a total of 20 min, followed by goat anti-rabbit or anti-mouse IgG-horseradish peroxidase conjugates for 1 h at room temperature, and washed three times with TBST for a total of 1 h. The immunoblots were visualized by ECL chemiluminescence (Pharmacia, Uppsala, Sweden).

5.2.4 Immunoprecipitation of Ku70 The immunoprecipitation was carried out according to a reported procedure (68).

In brief, DU145 cells were treated with various concentrations of HDAC inhibitors for 48 h, and lysed by Chaps lysis buffer (150 mM sodium chloride, 10 mM Hepes at pH 7.4 and

81 1.0% Chaps) with a cocktail of protease inhibitors (Sigma-Aldrich). After centrifugation at 13,000 x g for 15 min, the supernatants were collected, and incubated with protein

A-Sepharose beads for 15 min to eliminate nonspecific binding. The mixture was centrifuged at 1000 x g for 5 min, and the supernatants was incubated with agarose-conjugated goat polyclonal anti-Ku70 antibodies), followed by three washes in

1% triton in PBS. The immunocomplex was separated by SDS-PAGE and proteins were detected with mouse monoclonal anti-Ku70, Ku80, and rabbit polyclonal anti-pan-acetylated-lysine (pan-Ac-K).

5.2.5 Transfection of Flag-tagged Ku70 Expression Plasmids DU-145 cells were nucleofected with individual plasmids encoding Flag-tagged Ku70 and various Ku70 mutants by using a Nucleofactor kit L (Amaxa, Gaithersburg, MD) as described by the manufacturer. Transfection efficiency was at least 75% in DU-145 cells, as determined by pmaxGFP plasmid transfection.

5.2.6 DNA End-Binding Activity of Ku70 Preparation of nuclear extracts and assessment of DNA end-binding activity of Ku70 were carried out by using the Nuclear Extract Kit and Ku70/80 DNA Repair Kit, respectively (Active Motif, Carlsbad, CA). In brief, Ku70-containing nuclear proteins were extracted from drug-treated or nucleofected cells by a Nuclear Extract Kit. Protein concentrations of nuclear extracts were determined by using a Bradford protein assay kit.

Further quantification of protein loading amounts was determined by immunoblotting against Ku70 or Flag. Equivalent amounts of proteins from each lysate were loaded into a

96-well plate, of which each well contained immobilized linear oligonucleotides with blunt ends. Ku70 protein extracted from the nucleus could specifically bind to this oligonucleotide. The primary antibody used in the Ku70/86 DNA Repair Kit recognized

82 an epitope on Ku70 protein that was accessible upon DNA binding. Anti-Flag antibody was also used in the experiment for the transfection of Flag-tagged Ku70 plasmids.

Addition of a secondary HRP-conjugated antibody provided a sensitive colorimetric readout quantified by spectrophotometry.

5.2.7 Cell Survival Assay Survival curves for DU-145 cells were obtained by the clonogenic assay as previously described (198). Cells were plated at 800~1,000 cells/plate and exposed to 100

~ 250 nM (S)-HDAC-42 for 4 h or left untreated before exposure to the DSB-inducing drugs. After washing out the HDAC inhibitor, the cells were exposed to increasing amounts of bleomycin (0 - 100 mg/ml), doxorubicin (0 - 500 nM), and VP-16 (0 - 50 mM) for 1 h. The drugs were then washed away, and the cells were allowed to grow for 10 - 14 days. The colonies were fixed and stained with crystal violet (0.5% in 70% ethanol). Colonies containing more than 50 cells were scored. The survival fraction was calculated based on the number of colonies formed in drug-treated cells relative to that of the untreated control. Each dose was done in triplicate, and the experiments were repeated at least twice.

5.2.8 Apoptosis Assay Drug-induced apoptotic cell death was assessed using the Cell Death Detection ELISA kit (Roche Diagnostics, Mannheim, Germany), which quantitates cytoplasmic histone-associated DNA fragments in the form of mononucleosomes or oligonucleosomes. Cells were seeded and incubated at 8,000 cells/well in 12-well flat-bottomed plates in 10% FBS-supplemented RPMI-1640 medium. After 24 hours, cells were treated with (S)-HDAC-42 for 4 h, followed by bleomycin, doxorubicin, and

83 VP-16 at the indicated concentrations for 24. Both floating and adherent cells were collected and the assay was performed according to manufacturer’s instructions.

5.2.9 Immunocytochemical Detection of γH2AX Foci Formation All experiments were carried out in 10% FBS-supplemented RPMI 1640 medium.

Cells were seeded on cover slips at a density of 2 x 105 per slip, and incubated overnight.

These cells were treated with 250 nM (S)-HDAC-42 for 4 h, washed, and exposed to one of the following agents for 1 h: 10 µg/ml bleomycin, 100 nM doxorubicin, and 10 µM

VP-16. DMSO vehicle was used in the control. Drug-treated cells were fixed in 100% methanol at -20 °C for 30 min, rehydrated in PBS containing 0.1% triton X-100 for 10 min, and treated with mouse monoclonal anti-γH2AX antibody, followed by anti-mouse IgG-Alexa Fluor 488 antibody. The cover slips were mounted on slides with mounting solution containing 1 µg/ml DAPI, and observed under an Eclipse TE300 fluorescence microscope (Nikon, Melville, NY). Cell nuclei containing at least 5 condensed fluorescence dots were considered foci-positive, and 200 to 300 cells were counted in each slide to estimate the percentage of foci-positive cells in the whole cell population.

5.2.10 Molecular Modeling Analysis Protein sequence of human Ku70 (accession no. P23475) was retrieved from the National Center for Biotechnology Information Reference Sequence Collection. Structure of the Ku70/Ku80 complex was constructed by using the Modeller program with the published crystal structures of Ku heterodimer (RCSB entry code, 1JEY and

1JRR) as a modeling template. Subsequently, this Ku heterodimer structure was subject to the addition of polar hydrogens and the assignment of Kollman charges (202).

Three-dimensional affinity grids centered on the preformed channel with 0.375-Ǻ spacing were calculated for each of the following atom types: (a) Ku protein, A (aromatic

84 C), C, H, N, O; (b) DNA, C, A, N, O, P, H, e (electrostatic), and d (desolvation) using

Autogrid3 (203). AutoDock version 3.0.0 was used for the docking simulation. The DNA ligand structure was extracted from the published crystal structure of Ku heterodimer

(RCSB Protein Data Bank; PDB ID, 1JEY). We identified six Lysine residues in Ku 70 that are target for acetylation in docking simulation, of which K282, K317, K331, and K

338 are located within the ring structure, while K539, K542 lie within the C-terminal linker. The broken DNA was docked into the putative Ku preformed channel that exhibits a polarization of positive electrostatic charge focused on the inner surface of the channel and along the DNA-binding cradle (193). The Lamarckian genetic algorithm was used for DNA conformational searching because it has enhanced performance relative to simulated annealing or the simple genetic algorithm. Accordingly, docking simulations of the DNA-Ku binding were carried out by using molecular mechanics with the following parameters: trials of dockings, 100; population size, 150; random starting position and conformation, translation step ranges, 2.0 Ǻ; rotation step ranges, 50°; elitism, 1; mutation rate, 0.02; crossover rate, 0.8; local search rate, 0.06, and 100 million energy evaluations. Final docked conformations were clustered using a tolerance of 1.5 Ǻ root mean square deviation.

5.2.11 Statistical Analysis At least three independent experiments were performed for all clonogenic assays, Western blotting, and immunocytochemical analyses. Statistical differences in DNA binding affinity and γH2AX formation between drug-treated cells versus controls were determined by two-sided, unpaired Student’s t test. For statistical analysis, a statistically significant difference was defined as P < 0.05.

85 5.3 Results In light of the dual function of Ku70 in regulating DNA repair and Bax-mediated apoptosis (15,190,191), we chose DU-145 prostate cancer cells as a model based on the following considerations. First, DU-145 cells are deficient in Bax expression (Figure 5.1).

As the cytoprotective activity of Ku70 works, in part, through inhibition of Bax

(15,190,191), use of DU-145 cells would allow the assessment of the functional role of

Ku70 acetylation in the absence of Bax-mediated apoptosis. Second, DU-145 cells lack functional p53 (204), thereby eliminating the complication of p53 activation in response to apoptotic signals that produce DNA DSBs. Third, expression levels of Ku70 and Ku80 in DU-145 cells were similar to that of PC-3 and LNCaP cells (Figure 5.1), suggesting that these cells exhibit comparable levels of NHEJ DNA DSB repair activity.

5.3.1 Pretreatment with HDAC Inhibitors Sensitizes Prostate Cancer Cells to Agents that Generate DNA DSBs The ability of HDAC inhibitors to enhance the cell killing effect of drugs known to produce DNA DSBs, including bleomycin, doxorubicin, and VP-16, was demonstrated in Figure 5.2. The IC50 of (S)-HDAC-42 in suppressing DU-145 cell proliferation was 0.4 µM at 72 h (153). DU-145 cells were exposed to 0.25 µM (S)-HDAC-42 or DMSO vehicle in 10% FBS-supplemented medium for 4 h, washed with fresh medium, treated with different doses of individual agents for 1 h, and plated for clonogenic cell survival. As shown, a 4-h exposure to (S)-HDAC-42 alone over a dose range of 1 nM – 1 µM exhibited no appreciable effect on reducing cell survival (Figure 5.2). This is in line with our previous report that (S)-HDAC-42 and other HDAC inhibitors showed no significant antiproliferative effect in DU-145 cells within 24 h (Chapter 4) (153). In contrast, pretreatment of DU-145 cells with 0.25 µM (S)-HDAC-42 for 4 h shifted the dose response curves of bleomycin, doxorubicin, and VP-16 in inhibiting clonogenicity to the

86 left by almost an order of magnitude. This effect, however, was not noted in cells treated with 5-fluorouracil (5-FU) (Figure 5.2), an agent that causes minimum DNA damage

(205-207).

The chemosensitizing effect of (S)-HDAC-42 was, at least in part, attributable to its ability to augment cellular susceptibility to drug-induced apoptosis. DNA fragmentation ELISA indicates that pretreatment of cells with 250 nM (S)-HDAC-42 for

4 h significantly increased the extent of nucleosome formation in response to bleomycin, doxorubicin, and VP-16 in a dose-dependent manner (P < 0.05 compared with individual drug treatment alone) (Figure 5.3). Together, these findings suggest the chemosensitizing effect of HDAC inhibitors could be attributed to the ability to suppress DNA DSB repair activity. Among various components of the NHEJ DNA DSB repair pathway (185), Ku70 was especially noteworthy because it is targeted for deacetylation by HDACs.

5.3.2 HDAC Inhibition Leads to Increased Acetylation of Ku70 Pursuant to the above premise, we characterized the effect of four different HDAC inhibitors, including (S)-HDAC-42, SAHA, MS-275, and TSA, on Ku70 acetylation with reference to various HDAC-related biomarkers (histone H3 acetylation, p21WAF1/CIP1 expression, and α-tubulin acetylation) in DU-145 cells. These agents caused substantial increases in histone H3 acetylation and p21WAF1/CIP1 expression, but differed in their activity in inhibiting the α-tubulin deacetylase HDAC6 (Figure 5.4). (S)-HDAC-42 and TSA produced robust hyperacetylation of α-tubulin at submicromolar concentrations, while MS-275 was ineffective in inhibiting α-tubulin deacetylation. Moreover, we examined the effect of these inhibitors on the expression level of Ku70 and

Ku80 in light of a recent report that butyrate transcriptionally repressed these Ku proteins in melanoma cells (200). However, none of these agents showed an appreciable effect on suppressing the expression level of Ku70 or Ku80 (Figure 5.4).

87 To evaluate the drug effect on Ku70 acetylation, cell lysates of DU-145 cells treated with individual agents at the indicated concentrations for 24 h were immunoprecipitated with anti-Ku70, followed by immunoblotting with antibodies against acetyl-lysine, Ku80, and Ku70, respectively (Figure 5.5). As shown, these four inhibitors increased the acetylation level of Ku70 irrespectively differences in their pharmacological profiles. Moreover, acetylation of Ku70 did not affect its complex formation with Ku80 as the ratio of these two proteins in the immunoprecipitates remained unchanged (Figure 5.5 and 5.6).

5.3.3 HDAC Inhibition Diminishes Cellular Ability to Repair DNA DSBs Subsequently, the effect of acetylation on Ku70’s DNA-end binding activity in nuclear extracts was examined after exposing DU-145 cells to different doses of (S)-HDAC-42 and MS-275 for 24 h. As shown in Figure 5.7, these two HDAC inhibitors reduced Ku70’s binding affinity to broken DNA ends with IC50 less than 0.1 µM and 0.5 µM, respectively.

In the literature, phosphorylation of histone H2AX (γH2AX) has been widely used as a biomarker for drug-induced DNA damage in light of its occurrence at sites flanking DNA DSBs (208). Pursuant to the above finding, we assessed the impact of

(S)-HDAC-42 on the level of γH2AX in DU-145 cells treated with bleomycin, doxorubicin, or VP-16. Figure 5.8 depicts the immunochemical analysis of γH2AX foci formation in DU-145 cells pretreated with 250 nM (S)-HDAC-42 in 10%

FBS-supplemented medium for 4 h, followed by bleomycin (10 µg/ml), doxorubicin (100 nM), or VP-16 (10 µM) for 1 h. While (S)-HDAC-42 had no appreciable effect on

γH2AX phosphorylation levels, the sequential treatment significantly increased the number of γH2AX foci-positive cells as compared to individual chemotherapeutic agents

88 alone (P < 0.05) (Figure 5.8A and B), which was also confirmed by Western blot analysis of drug-treated DU-145 cell lysates (Figure 5.8C).

5.3.4 Constitutive Acetylation of Lysine Residues in DNA-Binding Domains Mimics

HDAC Inhibitors in Suppressing Ku70’s End-Binding Affinity Ku70 contains two DNA-binding domains. One is located at the N-terminus before residue 440, and the other resides in the C-terminus (residues 536 - 609) (186-189).

These two domains differ in their dependence on Ku80 for the DNA binding activity.

While the N-terminal domain requires the participation of Ku80 to bind broken DNA ends, the DNA binding activity of the C-terminal domain is independent of the heterodimer formation with Ku80. Moreover, the C-terminal DNA-binding domain encompasses the Bax-binding domain (68,190,191). It has been shown that mimicking a constitutively acetylated state of K539 or K542 by replacing it with glutamine (K to Q) abolished the Ku70-Bax interaction, thereby promoting Bax-mediated apoptosis (68). As the Ku heterodimer interacts with DNA through electrostatic interactions between its positively charged residues and DNA’s sugar-phosphate backbone (193,194), we hypothesized that acetylation of lysine residues would diminish Ku70’s DNA binding activity. This mode of regulation was reminiscent to that of human Flap endonuclease-1 (59) and the transcription factor interferon regulatory factor-7 (60) reported in the literature. To validate this hypothesis, we carried out site-directed mutagenesis to replace each of the following six lysine residues of Ku70 with glutamine: K282, K317, K331,

K338, K539, and K542, to mimic constitutive acetylation. Of these six residues, K282,

K317, K331, and K338 are located within the ring region, while K539 and K542 lie within the C-terminal linker of Ku70.

89 DU-145 cells were transfected with expression vectors encoding Flag-tagged wild type or each of these Ku70 mutants via nucleofection. Western blotting indicates that these ectopic Ku70s were expressed at levels several-fold higher than that of the endogenous protein (Figure 5.9A). Equal amounts of nuclear extracts of these transfected cells were analyzed for their ability to bind DNA broken ends. As shown in Figure 5.9B, constitutive acetylation at K282, K338, K539, and K542 suppressed the DNA end-binding activity by 40 - 60% (P < 0.005), while substitution of K317 or K331 with glutamine had no significant effect. It is interesting to note that the dual mutation at K282 and K338 (K282Q/K338Q) or replacement of all four lysine residues at the N-terminal DNA-binding domain (K282Q/K317Q/K331Q/K338Q) did not cause a higher degree of inhibition than that of K282Q or K338Q alone. This finding suggests that endogenous Ku70 contributed to the residual activity in the nuclear extracts of these transfected cells. In contrast, the double mutant K317Q/K331Q had no significant effect on suppressing the DNA end-binding activity, confirming that neither lysine residue played a role in interacting with DNA.

5.3.5 Molecular Modeling Analysis of the Interaction of Ku70/Ku80 Heterodimer with DNA To envisage the spatial arrangement of these six lysine residues relative to the sugar-phosphate backbone of DNA, we carried out molecular modeling analysis by docking a 14-bp duplex into the DNA-binding channel of the Ku heterodimer (Ku70,

Ku80, and broken DNA are colored in light blue, red, and yellow, respectively). Figure

5.10 depicts a consensus model of Ku-DNA complex formation on the basis of 50 docking simulations. As shown, these six lysine residues formed a positively charged lining of the DNA-binding cradle. However, the mean distance between the ε-amino moiety of individual lysine residue and the phosphate backbone of DNA varied to

90 different extents. The distance was approximately 3.5 Å for K282, K338, K539, and

K542, in contrast to 17.4 Å and 4.6 Å for K317 and K331, respectively. As both K317 and K331 were pointed outward the DNA binding ring, they were not within the effective range of electrostatic interactions with the DNA.

5.3.6 Constitutive Acetylation of Ku70 Diminishes the Cellular Ability to Repair

DNA DSBs We also examined the effect of the constitutive acetylation at K282 and K338 on augmenting drug-induced γH2AX foci formation. DU-145 cells were transiently transfected with a pCMV plasmid (mock) or individual plasmids encoding Flag-tagged wild type Ku70, and the mutants K282Q and K338Q. These cells were treated with 10 µg/ml bleomycin, 100 nM doxorubicin, 10 µM VP-16, or DMSO vehicle for 1 h, and the formation of γH2AX foci was assessed by immunocytochemistry (Figure 5.11). Immunostaining with anti-Flag antibodies indicates that ectopic Ku70 was expressed in nearly all cells examined. As compared to mock, overexpression of wild type Ku70 did not suppress drug-induced γH2AX formation. One plausible explanation for this lack of protection was that the Ku70-overexpressing cells might not have stoichiometric quantities of Ku80 to form heterodimeric Ku complexes. In contrast, overexpression of

K282Q or K338Q rendered cells more susceptible to the drug-induced γH2AX foci formation (P < 0.05).

5.4 Conclusion Here we report a unique histone acetylation-independent mechanism by which

HDAC inhibitors sensitize prostate cancer cells to DNA damaging agents through the regulation of the acetylation status of Ku70. It is well understood that Ku70 plays a dual role in repairing DNA DSBs and in suppressing Bax-mediated apoptosis by interacting

91 with Ku80 and Bax, respectively, in different cellular compartments (192). These cytoprotective functions of Ku70 are abrogated in HDAC inhibitor-treated cells due to hyperacetylation, thereby rendering cancer cells more susceptible to the killing effect of

DNA damage. This mechanistic finding has therapeutic relevance in terms of the combinatorial use of these two types of antitumor agents in cancer therapy. To achieve optimal therapeutic benefits, it might be desirable to use HDAC inhibitors preceding that of radiation/DNA-damaging agents.

Our data indicate that TSA, MS-275, SAHA, and (S)-HDAC-42 caused Ku70 hyperacetylation irrespective of differences in their pharmacological profiles in HDAC inhibition. For example, although MS-275 lacks the activity in α-tubulin acetylation, a biomarker for inhibiting HDAC6, it is equally efficacious in facilitating Ku70 acetylation as (S)-HDAC-42 and TSA. The DNA binding cradle of Ku70 contains multiple lysine residues to form a positively charged lining for interacting with broken DNA ends (68,193), many of which are acetylated in vivo (68). Our data also demonstrate that Ku70 hyperacetylation did not disrupt the complex formation with Ku80. However, not all of these lysine residues were involved in interacting with DNA. Of the six lysine residues examined (K282, K317, K331, K338, K539, and K542), constitutive acetylation of K317 and K331 had no appreciable effect on reducing binding affinity with DNA broken ends. Computer modeling data, indicating that K317 and K331 were not within an effective range to exert electrostatic interactions with DNA’s sugar-phosphate backbone, further confirming the bystander role of these two lysine residues. Previously, it was reported that substitution of either K539 or K542 in the C-terminal linker abolished Ku70’s ability to bind Bax, thereby facilitating Bax-mediated apoptosis. Here, we report that these two lysine residues also took part in binding DNA broken ends. The dual binding partners of the C-terminal linker underlines the crucial role of Ku70 in determining the fate of cells

92 that have undergone DNA damage, and provides a basis to design an effective strategy for the therapeutic combination of HDAC inhibitors with cytotoxic agents.

P a 45 C 1 3 N U C L D P

Ku80

Ku70

Bax

Bcl-xl

Actin

Figure 5.1: Expression Status of Ku70, Ku80, and Bax in Three Prostate Cancer Cell Lines, LNCaP, DU-145, and PC-3

Figure 5.2: Effect of (S)-HDAC-42 on Sensitizing DU-145 Prostate Cancer Cells to DNA DSB-Inducing Agents

Figure 5.3: Effect of 250 nM (S)-HDAC-42 Pretreatment on Drug-Induced DNA Fragmentation in DU-145 Cells

Figure 5.4: Dose-Dependent Effect of Four Different HDAC Inhibitors, Including (S)-HDAC-42, SAHA, MS-275, and TSA on Histone H3 Acetylation, p21WAF1/CIP1 Expression, α-tubulin Acetylation, and Expression of Ku70 and Ku80 in DU145 Cells

Figure 5.5: HDAC Inhibitor-Mediated Ku70 Acetylation in DU145 Cells

M DMSO TSA (S)-HDAC-42 1.0 1.0 µM

150 kDa

100 kDa

75 kDa Ku80 Ku70

Ku70, cleaved

50 kDa IgG

Figure 5.6: (S)-HDAC-42 Did Not Disrupt the Interaction between Ku70 and Ku80

DM 0.1 0.5 1.0 0.5 2.5 5.0 µM SO (S)-HDAC-42 MS-275

loading: Ku-70

Nucleolin

Figure 5.7: Dose-Dependent Effect of (S)-HDAC-42 and MS-275 on the DNA-End Binding Activity of Ku70 in DU145 Cells

100

Figure 5.8: Effect of (S)-HDAC-42 on Enhancing Drug-Induced γ-H2AX Foci Formation. A, immunocytochemical analysis. B, percentage γ-H2AX foci-positive cells in drug-treated versus DMSO-treated DU-145 cells. C, Western blot analysis of the effect of (S)-HDAC42 on drug-induced γ-H2AX phosphorylation.

101

Figure 5.9: Effect of Constitutive Acetylation of Lysine Residues in the DNA-Binding Cradle of Ku70 on Its DNA-End Binding Activity. A, Western blot analysis of the expression level of Flag-tagged Ku70 versus endogenous Ku70 in nuclear extracts. B, relative DNA end-binding activity of individual Ku70 mutants to wild type Ku70.

102

Figure 5.10: Molecular Modeling Analysis of the Mode of Interactions between the Ku Heterodimer and DNA. Ku70, Ku80, and broken DNA are colored in light blue, red, and yellow, respectively. The six lysine residues that underwent site-mutagenesis are indicated.

103

A. B.

Fig 5.11: Effects of the Constitutive Acetylation of K282 and K338 of Ku70 on Drug-Induced γ-H2AX Foci Formation

104

CHAPTER 6

DISCUSSIONS AND PERSPECTIVES

The biology of the various HDAC isozymes and their relationship to tumorigenesis is just beginning to be elucidated and is largely driven by the perceived clinical potential of HDAC inhibitors. However, a major challenge is lack of in-depth understanding of the biological effects of structurally diverse HDAC inhibitors. Moreover, the complexities of the histone code and the variety of other nonhistone proteins whose functions are modulated by acetylation/deacetylation underlie why the anticancer effects of HDAC inhibition remain ambiguous. In addition to modulating epigenetic events, emerging evidence indicates that HDAC inhibitors also target nonhistone substrates at different cellular levels, which constitute transcription -independent mechanisms to account for their anticancer activities. Consequently, these studies have indicated that the effects of HDAC inhibitors could be considerably broader and more complicated than originally understood. Through this dissertation research, we have identified two additional histone acetylation-independent mechanisms of HDAC inhibitors, i.e. targeting the phosphorylation status of Akt (Chapter 3), an important cell survival signal, and the DNA DSB-repair function of Ku70 (Chapter 5). We further demonstrated that, along with the transcriptional regulation of tumor suppressor genes and oncogenes, these two modes of action of HDAC inhibitors also provide a basis for their antineoplastic activities in single drug treatment or in combination with other anticancer drugs (Chapter 4 and 5).

105 6.1 HDAC Inhibitors and Akt Dephosphorylation In the first part of this study, we demonstrated a novel nonepigenetic mechanism by which the HDAC inhibitor, TSA, induces Akt dephosphorylation by disrupting

HDAC-PP1 complexes. Mechanistically, the HDAC inhibitor-mediated reorganization of PP1 complexes represents a unique mode of Akt regulation independent of PI3K signaling pathways. Previously, drug-facilitated PP1-dependent Akt dephosphorylation was reported in ErbB2-overexpressing breast cancer cells treated with the Hsp90 inhibitor geldanamycin or the ErbB inhibitor ZD1839 (209), however, through a distinct mechanism. In contrast to the reshuffling of PP1 complexes, the effect of geldanamycin and ZD1839 on Akt deactivation was attributable to the direct activation of PP1. In the present study, no increase in Ser/Thr protein phosphatase activity was observed in TSA-treated U87MG cells. In addition, it was reported that inhibition of HDAC6 resulted in the hyperacetylation and loss of the chaperone activity of Hsp90 (67). As Hsp90 forms intracellular complexes with Akt (210), inhibition of Hsp90 function provides a plausible mechanism for down-regulating Akt kinase activity through the promotion of Akt degradation (211,212). However, our finding that Akt levels remained unaltered refutes the involvement of Hsp90 in HDAC inhibitor-induced Akt dephosphorylation in U87MG cells. Together with the previous findings of the functional roles of HDAC-PP1 pairs in the modulation of CREB phosphorylation (86) and α-tubulin acetylation (87), this histone-independent mechanism provides a potential basis to account for the antineoplastic activities of HDAC inhibitors in growth inhibition and apoptosis induction.

Recently, PP1 has emerged as an important therapeutic target in light of its regulatory role in a plethora of cellular functions (213). However, targeting PP1 proves elusive because it forms complexes with over 50 established or putative regulatory subunits in different cellular compartments. The selective action of HDAC inhibitors on cellular

106 HDAC-PP1 complexes represents the first example of modulating specific PP1-protein interactions by small-molecule agents. From a clinical perspective, identification of this

PP1-facilitated dephosphorylation mechanism underlies the potential use of HDAC inhibitors in lowering the apoptosis threshold for other therapeutic agents through the down-regulation of Akt signaling. This therapeutic strategy is illustrated by the ability of

HDAC inhibitors to sensitize cancer cells to the apoptotic effects of the Bcl-Abl kinase inhibitor STI571 (Gleevec, imatinib mesylate) (142,144), the Her-2 antibody trastuzumab (140,214), the receptor tyrosine kinase FLT-3 inhibitor PKC412 (215), the purine analogue fludaribine(138), and the Hsp90 antagonist 17-allylamino-demethoxy geldanamycin (17-AAG) (139,216). In conclusion, the ability of HDAC inhibitors to deactivate Akt through the reorganization of PP1 complexes presents another example of the complexity of the pharmacological functions of these agents. In light of the clinical application of HDAC inhibitors, a better understanding of this novel histone-independent mechanism, for example characterizing the interaction domains of HDACs and PP1, will allow the design of more effective strategies for optimize the use of the agents in cancer treatment and/or prevention.

6.2 Anticancer Effects of (S)-HDAC-42 In the second part of this study, we examined the in vitro and in vivo efficacy of (S)-HDAC-42, a novel phenylbutyrate-derived HDAC inhibitor synthesized in our laboratory, with reference to SAHA in prostate cancer cells. We demonstrated that the antitumor effects of (S)-HDAC-42 and SAHA are attributed to both histone acetylation-dependent and -independent mechanisms by interfering with the activation or expression status of a number of signaling targets. Especially noteworthy are the effects of (S)-HDAC-42 on the dephosphorylation of Akt and the repression of Bcl-xL and

107 survivin in PC-3 cells, both of which were not as evident in SAHA-treated cells. Together, these mechanisms provide a molecular basis to account for the higher in vivo potency of

(S)-HDAC-42 than SAHA in suppressing established PC-3 xenograft tumor growth.

Moreover, since levels of phospho-Akt and survivin were also reduced, though to a lesser degree, by SAHA treatment, it is likewise unclear whether the differential effects of these agents on apoptotic regulators are merely a function of their different potencies and/or result from true differences in the spectrum of their respective pharmacological targets.

From the toxicological point of view; tumor growth suppression by (S)-HDAC-42 occurred in the absence of limiting toxicity, with the exception of the observed testicular pathology. While testicular toxicities are common adverse effects of cancer therapy (217), testicular pathology has not been reported as a consequence of HDAC inhibitor treatment in preclinical models of human cancer. A single study examining the antifertility effect of TSA in mice reported a completely reversible impairment of spermatogenesis resulting from increased apoptosis of spermatocytes (218). Although the cellular target mediating this toxicity of (S)-HDAC-42 remains unclear, the lack of pathological changes in the pituitary and accessory sex organs suggests a primary testicular toxicity. Supporting this supposition is the absence of histopathological changes in the gut and bone marrow, which are also common sites of adverse chemotherapeutic consequences.

6.3 Acetylation of Ku70 Impairs Its DNA-Repair Function The third part of this study reported another novel histone acetylation

-independent mechanism by which HDAC inhibitors sensitize prostate cancer cells to

DNA damaging agents by targeting Ku70 acetylation. Our data indicated that pretreatment of the Bax-deficient DU-145 cells with HDAC inhibitors led to Ku70 hyperacetylation. Although hyperacetylation of Ku70 does not affect the Ku heterodimer formation, it reduces Ku70’s DNA end-binding activity, and diminishes the cellular

108 capability to repair drug-induced DNA DSBs. Moreover, mimicking acetylation by replacing lysine residues with glutamine in Ku70’s DNA binding domains via site-directed mutagenesis also achieves the same effect. Previously, it was reported that substitution of either K539 or K542 in the C-terminal linker abolished the interaction between Ku70 and Bax, thereby facilitating Bax-mediated apoptosis. Here, we report that these two lysine residues also take part in binding DNA broken ends. The dual binding character of the C-terminal linker underlines the crucial role of Ku70 in determining the fate of cells that have undergone DNA damage, and provides a basis to design an effective strategy for the therapeutic combination of HDAC inhibitors with cytotoxic agents. In the literature, several different mechanisms have been proposed to account for the ability of HDAC inhibitors to sensitize cancer cells to DNA damaging agents. For example, sodium butyrate has been reported to reduce the gene expression of Ku70 and Ku80 in human melanoma cells (200), and to increase DNA topoisomerase IIa expression in human leukemic cells (219). Moreover, phenylbutyrate, but not sodium butyrate, was shown to enhance the cytotoxic effect of doxorubicin in multidrug-resistant cancer cells by suppressing the activity of antioxidant enzymes such as superoxide dismutase and glutathione reductase (220). These multiple mechanisms underscore the pleiotropic effects of HDAC inhibitors at both epigenetic and cellular levels. However, in this study, we did not find the repression of Ku70 or Ku80 in (S)-HDAC-42-treated DU-145 cells, which was in contrast to what was noted in sodium butyrate-treated melanoma cells. This discrepancy might be attributable to differences in the pharmacological properties of different HDAC inhibitors in the transcriptional regulation of Ku proteins, or to the genetic heterogeneity among different cancer cell lines.

In conclusion, data from this and other laboratories have demonstrated the in vivo efficacy of HDAC inhibitors as single-agent antitumor drugs, which target multiple facets

109 of cancer cell survival and cell cycle progression. The ability of HDAC inhibitors to regulate cellular capacity to repair DNA damage through the modulation of Ku70’s acetylation status underscores the viability of their synergistic combinations with DNA damaging agents as part of a therapeutic strategy for prostate cancer.

6.4 Future Perspectives HDAC inhibitors belong to a new category of anticancer agents with pleiotropic cellular effects that act cooperatively to mediate potent antitumor activities. However, due to the lack of complete insight into the biological functions and regulation of each HDAC isoform and the effects of HDACs on the diverse list of nonhistone substrates during tumorigenesis, the molecular processes underlying the antineoplastic or even the adverse effects of HDAC inhibitors remain to be fully elucidated. Currently, knowledge about the specific functions of each HDAC isoform is still limited. It remains unclear whether HDACs function redundantly in transcriptional regulation or other cellular events during development or tumor transformation. Nevertheless, progress in gaining and understating of the isoform-specific roles of Class II HDACs is being made. For example, it was recently reported that HDAC7 is specifically expressed in the vascular endothelium during early embryogenesis (221). Disruption of the HDAC7 gene in mice results in embryonic lethality due to a failure in endothelial cell-cell adhesion. These findings reveal, for the first time, a specific role for HDAC7 in the maintenance of vascular integrity and cardiovascular development. A report from the same group also demonstrated that HDAC5 and HDAC9 have redundant roles in cardiac development

(222). These findings suggest potential adverse cardiovascular effects of HDAC inhibitors that target these isozymes. Moreover, it is unclear whether structurally and pharmacologically diverse HDAC inhibitors have a common mechanism for inhibiting tumor growth across different cancer cell types, however, it seems that many of the

110 reported effects could be cell-type dependent. Consequently, defining the molecular events underlying the various antitumor activities of HDAC inhibitors will require a better understanding of the expression profiles of HDACs and the effects of HDAC inhibition on diverse cellular proteins and pathways in different cancer cell types.

Several HDAC inhibitors selectively inhibit tumor cell growth at concentrations up to one order of magnitude lower than those toxic to normal cells in vitro, which suggests that a favorable therapeutic window could be achieved. Although a number of clinical trials using different HDAC inhibitors have demonstrated promising antitumor responses, adverse side effects, including gastrointestinal symptoms (diarrhea, nausea, and anorexia), hematological abnormalities (thrombocytopenia, anemia, and myelosuppression), and cardiac abnormalities, have also been recorded (223,224). Whether these toxicities represent effects on histone acetylation-dependent or -independent targets of these compounds remains to be determined. One possible way to minimize toxicities associated with HDAC inhibition is the production of isoform-specific inhibitors. For instance, a HDAC inhibitor without activity against HDAC7 might have less cardiovascular toxicity. As mentioned previously, a HDAC6-specific inhibitor has been identified through high-throughput screening (74), and the solution of the crystal structures of other HDACs, such as HDAC8 (225), should pave the way for the development of other isoform-specific inhibitors. Therefore, a more detailed understanding of the specific roles played by various HDAC isoforms during human tumorigenesis might lead, not only to development of isoform-specific inhibitors, but also to more effective or less toxic antitumor therapeutics, as compared to the multiclass HDAC inhibitors that are currently undergoing clinical evaluation.

More general considerations for the use of HDAC inhibitors as cancer therapeutics also need to be addressed. Since the selection of a molecule for further drug development requires a balance of biological potency, safety and pharmacokinetics, it is

111 importance to elucidate the pharmacokinetic and toxicological properties of these HDAC inhibitors before they can be considered as potential new drugs. In addition, it has become clear that tumor cells are not homogeneous and that cancer stem cells do play some role in tumorigenesis and even in drug-resistance (226,227). It is reasonable to speculate that the lack of success of some current cancer therapeutics are, in part, due to the failure to target cancer stem cells (228). Therefore, using HDAC inhibitors to address this pivotal issue of tumor stem cell sensitivity should clarify the true potential of HDAC inhibitors in clinical usage. Finally, although a clinical role for HDAC inhibitors as novel cancer therapeutics seems inevitable, their general clinical utility will likely depend greatly on the future development of molecular or cellular predictors of their antitumor activity.

112

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