Cell Signaling & Molecular Targets in Cancer

Cell Signaling & Molecular Targets in Cancer

Malay Chatterjee • Khosrow Kashfi Editors

Cell Signaling & Molecular Targets in Cancer

Editors Malay Chatterjee Department of Pharmaceutical Technology Jadavpur University Kolkata, India [email protected]

Khosrow Kashfi Department of Physiology and Pharmacology City University of New York Medical School New York, USA [email protected]

ISBN 978-1-4614-0729-4 e-ISBN 978-1-4614-0730-0 DOI 10.1007/978-1-4614-0730-0 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011940810 © Springer Science+Business Media, LLC 2012 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Acknowledgement

I sincerely thank all my students, researchers and colleagues who initially put forth the idea of writing this book. The book is the result of their initiative and constant motivation. I wish to express special gratitude to Prof. Kho Kashfi, City University of New York, who collaborated with me as a co-editor in editing this book and sincerely acknowledge his valuable suggestions, comments and guidance in completing this book. I also like to thank our reviewers for their painstaking work. I would like to thank Mr. Subhadeep Das, research associate of the department who played a major role in assuring the consistency of the book. Many new approaches and topics in cancer research are included in this book and we have incorporated the skills, experience and perspective of a truly international complement of highly distinguished authors. The publication of this book could not have been possible but for ungrudging effort put in by all our authors. I would like to thank all of them for their valuable contribution. I especially thank our publisher Springer, USA, for their continuous support, enthusiasm and help in improving the usability of the text. Many individuals associated with our publishers need our special thanks. Outstanding among them are Florencia Schapiro, Cancer Research Editor at Springer Biomedicine, and her assistant Greg Baer who supervised the production of this book. I thank my family for their patience, love and support of this venture. Special thanks are to my wife Shipra. She has been the inspiration that has driven me onwards. Last but not the least I hope this book will provide useful and important information to researchers, clinicians, academics and pharmaceutical scientists engaged in the areas of cancer research. Malay Chatterjee

v

Contents

1

GRB2 Signaling as a Molecular Target for Cancer ........................... Alessio Giubellino

2

Human Arylamine N-acetyltransferase 1: From Drug Metabolism to Drug Target ................................................................... Fernando Rodrigues-Lima, Julien Dairou, Florent Busi, and Jean-Marie Dupret

1

23

3

Targeting Argininosuccinate Synthetase in Cancer Therapy ............ Niramol Savaraj, Min You, Chunjing Wu, Macus Tien Kuo, Vy Dinh, Medhi Wangpaichitr, and Lynn Feun

37

4

Parathyroid Hormone–Related Peptide Signaling in Cancer ............ Franco Oreste Ranelletti and Giovanni Monego

53

5

Signalling Molecules as Selective Targets for Therapeutic Strategies in Multiple Myeloma ............................................................ Francesco Piazza and Gianpietro Semenzato

6

Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract ......... Hiroshi Yasuda and Fumio Itoh

7

AIB1: A Transcriptional Coactivator Which Integrates Signaling Cross Talk in Cancer Cells ................................................... Macarena Ferrero and Jaime Font de Mora

8

9

Rational Design of DNA Anticancer Agent That Targets Signal Transducer and Activator of Transcription 3 (Stat3) for Cancer Therapy ............................................................................... Naijie Jing Estrogen Receptor Signaling in Lung Cancer ..................................... P.A. Hershberger and J.M. Siegfried

87 109

129

167 191

vii

viii

10

Contents

Microparticle Dissemination of Biological Activities: Implications for Cancer Biology ........................................................... Pauline P. Goh

11

Protein Kinase D Signaling in Cancer ................................................. Peter Storz

12

Cell Signaling and Cancer: Integrated, Fundamental Approach Involving Electron Transfer, Reactive Oxygen Species, and Antioxidants...................................................................... Peter Kovacic and Ratnasamy Somanathan

13

211 245

273

Targeting Signal Transducer and Activator of Transcription (STAT) for Anticancer Therapy............................................................ Fabio P.S. Santos, Inbal Hazan-Halevy, and Zeev Estrov

299

Index ................................................................................................................

323

Contributors

Florent Busi UFR des Sciences du Vivant, Unité de Biologie Fonctionnelle et Adaptative, Univ Paris Diderot-Paris 7, CNRS EAC 4413, 75013 Paris, France Julien Dairou UFR des Sciences du Vivant, Unité de Biologie Fonctionnelle et Adaptative, Univ Paris Diderot-Paris 7, CNRS EAC 4413, 75013 Paris, France Vy Dinh Miami V.A. Healthcare System, Miami, FL 33125, USA Jean-Marie Dupret UFR des Sciences du Vivant, Unité de Biologie Fonctionnelle et Adaptative, Univ Paris Diderot-Paris 7, CNRS EAC 4413, 75013 Paris, France Zeev Estrov Department of Leukemia, The University of Texas, Anderson Cancer Center, 1515 Holcombe Boulevard, Unit 0428, Houston, TX 77030, USA, [email protected] Macarena Ferrero Laboratory of Cellular and Molecular Biology, Centro de Investigación Príncipe Felipe, Avenida Autopista del Saler 16, 46012 Valencia, Spain Lynn Feun Sylvester Cancer Center, University of Miami School of Medicine, Miami, FL 33125, USA, [email protected] Alessio Giubellino, MD Urologic Oncology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD 20892, USA, [email protected] Pauline P. Goh Vascular Immunology Unit, Department of Pathology, Sydney Medical School, University of Sydney, Sydney, NSW 2006, Australia [email protected] Inbal Hazan-Halevy Department of Hematology and Bone Marrow Transplantation, Tel Aviv Sourasky Medical Center, Haifa, Israel ix

x

Contributors

P.A. Hershberger Department of Pharmacology, University of Pittsburgh, Pittsburgh, USA Fumio Itoh Division of Gastroenterology and Hepatology, St. Marianna University School of Medicine, 2-16-1 Sugao, 216-8511 Kawasaki, Japan Naijie Jing, Ph.D. Department of Medicine, Baylor College of Medicine, Houston, TX 77030, USA, [email protected] Peter Kovacic Department of Chemistry, San Diego State University, San Diego, CA 92182-1030, USA, [email protected] Macus Tien Kuo, M.D. Anderson Cancer Center, University of Texas, Houston, TX 77054, USA Giovanni Monego Institutes of Human Anatomy and Cell Biology, Universtà Cattolica del S. Cuore, Largo F. Vito 1, 00168 Rome, Italy Jaime Font de Mora Laboratory of Cellular and Molecular Biology, Centro de Investigación Príncipe Felipe, Avenida Autopista del Saler 16, 46012 Valencia, Spain, [email protected] Francesco Piazza Department of Clinical and Experimental Medicine, Haematology-Immunology Section and Venetian Institute of Molecular Medicine, Haematological Malignancies Unit, Padua University School of Medicine, Padua, Italy, Via Giustiniani 2, 35128 Padova, Italy Franco Oreste Ranelletti Institutes of Histology and Embryology, Universtà Cattolica del S. Cuore, Rome 00168, Italy, [email protected] Fernando Rodrigues-Lima Univ Paris Diderot-Paris 7, Unité de Biologie Fonctionnelle et Adaptative, 75013 Paris, France, [email protected] Fabio P.S. Santos Department of Leukemia, The University of Texas, M.D. Anderson Cancer Center, Houston, TX, USA Niramol Savaraj Miami V.A. Healthcare System, Miami, FL 33125, USA Gianpietro Semenzato Department of Clinical and Experimental Medicine, Haematology-Immunology Section and Venetian Institute of Molecular Medicine, Haematological Malignancies Unit, Padua University School of Medicine, Padua, 35128 Padova, Italy, [email protected]

Contributors

xi

J.M. Siegfried Co-Director, Thoracic Malignancies Program, Department of Pharmacology, University of Pittsburgh Cancer Institute, Pittsburgh, PA 15213, USA, [email protected] Ratnasamy Somanathan Department of Chemistry, San Diego State University, San Diego, CA, 92182-1030, USA Centro de Graduados e Investigación del Instituto Tecnológico de Tijuana, Apdo postal 1166, Tijuana, BC, Mexico Peter Storz, Ph.D. Department for Cancer Biology, Mayo Clinic, Jacksonville, FL 32224, USA, [email protected] Medhi Wangpaichitr Miami V.A. Healthcare System, Miami, FL 33125, USA Chunjing Wu Miami V.A. Healthcare System, Miami, FL 33125, USA Hiroshi Yasuda Division of Gastroenterology and Hepatology, St. Marianna University School of Medicine, 2-16-1 Sugao 216-8511 Kawasaki, Japan, [email protected] Min You Sylvester Cancer Center, University of Miami School of Medicine, Miami, FL 33136, USA

Chapter 1

GRB2 Signaling as a Molecular Target for Cancer Alessio Giubellino

Introduction Adaptor Proteins in Cell Signaling Eukaryotic cells have developed complex signaling processing circuits that operate inside the cellular cytoplasm and govern several cellular functions [1]. In an oversimplified model, upon stimulation by extracellular ligands (e.g., growth factors), cell surface receptors become activated. The activation is responsible for the transient posttranslational modification (e.g., phosphorylation) of specific residues inside a defined amino acid sequence, creating specific docking sites for intracellular signaling effectors [2, 3]. The specificity of such interactions is dictated by the presence inside the protein effector of “domains,” evolutionary conserved sequences with characteristic structures and functions [4, 5]. Structural analysis of the protein kinase Src uncovered the presence, besides the catalytic domain (named SH1 domain), of other sequences with peculiar and distinct structures [6]; those sequences where named consecutively Src-homology 2 (SH2) and Src-homology 3 (SH3) domains because of the proximity to the catalytic domain. Computational analysis has allowed the identification of similar domains in other proteins. In addition, several other protein-binding modules were discovered, each recognizing specific binding motifs on partner proteins [7]. Comprehensively these proteins go under the name of adaptor (or adapter) proteins. A more strict definition of this class of proteins includes only proteins containing two or more protein binding modules, and devoid of any catalytic activity [8].

A. Giubellino (*) Urologic Oncology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD 20892, USA e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_1, © Springer Science+Business Media, LLC 2012

1

2

A. Giubellino

Cell signaling has evolved as an intricate network of proteins organized into cascades where the function of one component is dependent upon interaction and activation by other proteins. In this context, adaptor proteins serve the role to connect and assemble the complex array of intercommunicating components. Increasing evidence, though, shows that they are not only simple connectors, but refined modulators of cell signaling; indeed posttranslational modifications (such as phosphorylation) of some of these proteins can finely tune the process, adding a further layer of complexity to the system. Protein–protein interactions are of paramount importance for virtually every process inside a cell, and define specificity in signal transduction. The presence of modular domains that specifically interact with a consensus sequence inside the structure of partner proteins (frequently recognizing specific posttranslational modifications) is at the base of the temporal and spatial specificity of signaling transduction networks. The versatility and the modular organization can target signaling proteins to the appropriate location within the cells, providing the correct wiring to control and integrate the flow of information. The family of adaptor proteins encompasses several members with a variety of modules and interaction specificity. For example, the phosphotyrosine-binding (PTB) domain, similarly to the SH2 domain, binds to phosphotyrosine-containing sequences [9]. Other domains can interact with other modified peptides (such as the 14-3-3 and WW domains with phosphoserine-containing sequences, or the Bromo domain with acetyl-lysine), with unmodified peptides (such as the EVH1 and SH3 domains with proline-rich sequences, or the VHS domain that bind to acidic di-leucine motifs in cytoplasmic domains of sorting receptors), with phospholipids (such as the PH or the FERM domain that bind to the phosphates in the headgroup of phosphoinositides) and even with nucleic acids (such as the PUM domain binding with RNA or the Tubby domain binding to DNA sequences). Several other modules recognize the identical module in other proteins (domain/domain interactions and formation of homotypic oligomers); examples of this category include the PDZ, SAM, and DD domains. It is remarkable that a limited number of modules can control the complex actions of many different cell types; since it is improbable that there are proteins for a single biological role, it is reasonable to consider that signaling proteins rather act in a combinatorial fashion. On this regard, a system biology approach will most likely give us a better opportunity to understand which combination of proteins is sufficient and necessary for a distinct biological function.

The Grb Family The discovery of the SH2 domain in the sequence of Fps and Src tyrosine kinases [10, 11] have sparked the detection of similar domains in other signaling molecules and the discovery of other regions of similarity in the sequence of other proteins. By screening bacterial expression libraries with the phosphorylated C-terminal tail of the EGF receptor, several other SH2 domain containing proteins were characterized [12].

1

GRB2 and Target Therapy

3

In particular, using the CORT (Cloning Of Receptor Targets) method, the first clone to be isolated was the p85 subunit of the previously identified PI-3 kinase; being the first protein to be discovered with this technique, it was named growth factor receptor-bound protein 1 (or Grb1) [13]. It was found that this 85-kDa protein had no intrinsic PI-3 kinase activity, but its structure was composed of two SH2 domains and one SH3 domain. The N-terminus of the p110 subunit (with catalytic activity) of PI3K is tightly associated with a region located between the two SH2 domains of p85. The PI3K family is then organized in three classes, each one with a unique preference for phosphoinositide substrates, producing specific lipid second messengers. The second clone, Grb2, was isolated from the same library used for Grb1/p85 and independently cloned by PCR [14, 15]. Grb2 is one of those proteins made exclusively of SH2 and SH3 domains (such as Crk and Nck). Developmental studies of the homologues in Caenorhabditis elegans (Sem-5) and in Drosophila melanogaster (Drk, then classified as Grb8) have elucidated the role of Grb2 in cell signaling as discussed below. Several other Grb clones that were isolated turned out to be proteins already discovered through other methods [16]. Grb3 was identified as the protein Crk and Grb4 turned out to be the protein Nck; both these proteins have a structural similarity to Grb2, in the fact they are exclusively composed of SH2 and SH3 domains. Grb5 was recognized as the tyrosine kinase Fyn, Grb6 as the phospholipase C-gamma (PLC-g) and Grb9 as the protein tyrosine phosphatase Syp. Grb7 and Grb10 were discovered as a novel family of SH3 domain proteins with a distinct structure [16]. Indeed the molecular architecture of Grb2 (the prototype of the superfamily of adaptor proteins including Grb10 and Grb14) includes an N-terminal proline-rich region, a C-terminal SH2 domain, and a central segment named GM region (for Grb and Mig) which includes a PH domain with sequence homology to the Caenorhabditis elegans protein Mig-10 [17, 18]. Despite the highly conserved structure homology, the members of the Grb7/10/14 superfamily exhibit a distinctive expression pattern among different tissue types. Interestingly, overexpression of these proteins has been observed in several cancers and correlate with tumor progression [19, 20]. A further level of complexity of this family of proteins (not completely understood and under investigation) is represented by potential different mechanisms of regulation, including phosphorylation and the occurrence of homo- and hetero-oligomerization (Table 1.1).

The Adaptor Protein GRB2 Grb2 in Cell Signaling The growth factor receptor-bound protein 2 (Grb2) is a prototype adaptor protein that has been extremely useful in understanding the function of other adaptor proteins in cell signaling. Grb2 is widely expressed and is essential for a variety of

4 Table 1.1 The growth factor receptor-bound protein family Common name Grb1 p85 subunit (PI-3 Kinase) Grb2 Grb2 Grb3 Crk Grb4 Nck Grb5 Fyn Grb6 Phospholipase C-g (PLC-g) Grb7 Grb7 Grb8 Drk (Grb2 homolog in Drosophila) Grb9 Syp (protein tyrosine phosphatase-2C) Grb10 Grb10 Grb14 Grb14

A. Giubellino

Domains SH2, SH3, Bcr SH2, SH3 SH2, SH3 SH2, SH3 SH2, SH3, TK SH2, SH3, PLC SH2, GM/PH SH2, SH3 SH2, TK SH2, GM/PH SH2, GM/PH

basic cellular functions, acting as a critical downstream intermediary in several oncogenic signaling pathways. The mature form has a mass of 25 kDa and a modular structure with one Src homology 2 (SH2) domain flanked by two SH3 domains [21]. Grb2 is also known under several names, but one name in particular (EGFRbinding protein) reflects the way it was originally isolated through screening for epidermal growth factor receptor (EGFR)–interacting proteins. Subsequent experiments have demonstrated that Grb2 interacts with several other proteins, becoming one of the adaptor proteins with the highest number of reported cellular interactions. In particular through its SH2 domain, which is a conserved sequence of 100 amino acids, Grb2 can interact directly with receptor tyrosine kinases (c-Met, PDGFR) and non-receptor tyrosine kinases, such as focal adhesion kinase (FAK) and Bcr/Abl [22], as well as substrates of tyrosine kinases, via preferential binding to the phosphopeptide motif pYXNX (where N is asparagine and X any residue). The two carbonyl and amino-terminal Src homology 3 (SH3) domains, which have a conserved sequence of around 50 amino acids, bind proline-rich regions within interacting proteins. The classical model of how Grb2 carries out its functions relies on the widely confirmed observation that Grb2 is constitutively associated with son of sevenless (Sos), a guanine-nucleotide exchange factor that promotes GDP–GTP exchange on Ras, with consequent activation of Ras and the mitogen-activating protein kinase (MAPK) cascade [23]. This important function is executed upon recruitment of Grb2 (after growth factor receptor activation) near the cell membrane, bringing Sos close to Ras. The Grb2 gene (in humans and located on chromosome 17) is highly conserved among species and Grb2 expression is critical for normal development [24]. Mouse embryos homozygous for a Grb2 null allele die at a very early embryological stage, precluding investigation of this gene later in development. However, using mutations originally identified in C. elegans, it has been possible to generate a hypomorphic allele of the mouse Grb2 gene and derive Grb2-/hypomorph mice that manifest morphogenic defects in neural crest cell migration into brachial arches and defects in

1

GRB2 and Target Therapy

5

cardiovascular development [25]. These findings reinforce the idea that Grb2 is critical for epithelial morphogenesis and for processes such as cell motility and vasculogenesis. A natural isoform known as Grb3-3, with a deletion within the SH2 domain, has been discovered [26]. Grb3-3 is unable to bind to the epidermal growth factor receptor, and inhibits EGF-initiated transactivation of a Ras-responsive element. This Grb3-3 acts as a dominant negative protein over Grb2 and experimental evidence points out a role of Grb3-3 in suppressing proliferative signals and activating apoptosis [27]. Together with Grb2, Grb3-3 has been reported to localize in the nucleus and bind to the heterogeneous nuclear ribonucleoprotein C (hnRNP C) [28]; the physiological implication of these interactions require further investigation, but it has been suggested to be part of a general mechanism involving Grb2 and Grb3-3 in the apoptotic process. The domain organization of Grb2 is similar to at least two other protein adaptors: Gads (Grb2-related adaptor protein 2) and Grap (Grb2-related adaptor protein). Gad is mainly involved in leukocyte-specific tyrosine kinase signaling, and interacts with several proteins, including Grb2-associated binding protein 1 (Gab1) and the SLP-76 leukocyte protein (LCP2) [29, 30]. Grap interacts with the stem cell factor receptor and the receptor for erythropoietin, and stabilizes the formation of a complex with the oncoprotein Bcr/Abl [31]. In analogy to other Grb2 family members, also Grb2 can be tyrosine phosphorylated; this posttranslational modification is orchestrated by the Bcr/Abl oncoprotein and the epidermal growth factor receptor [32]. The observation that Grb2 phosphorylation at tyrosine 209 disrupts its binding to Sos suggests a potential negative regulation by phosphorylation; further and more detailed investigation is required to better understand how this and other posttranslational modifications impact Grb2’s cellular functions.

Grb2 and Cancer Besides its role as a critical downstream intermediary in several oncogenic signaling pathways, Grb2 signaling has also been implicated directly in the pathogenesis of several specific human malignancies. The Grb2 gene is located in the chromosome region 17q22, which is known to be duplicated in solid tumors and in leukemias [33]. Interestingly, Grb2 and Her2 are both located on chromosome 17q, and amplifications in this region occur frequently in solid tumors such as neuroblastoma and ovarian and gastric cancers. In chronic myelogenous leukemia (CML), the chimeric Bcr/Abl tyrosine kinase oncoprotein is able to bind the Grb2 SH2 domain through Y177 in the BCR region, linking the fusion protein to the Ras pathway [34]. Inhibition of the ATP binding site of the Abl tyrosine kinase with small molecules such as Imatinib (STI-571) is currently the main therapy, but inhibition of Grb2 could become a valid adjuvant therapy or an alternative in patients resistant to other treatments.

6

A. Giubellino

Grb2 may also be overexpressed in solid tumors, like those found in breast cancer. Indeed, in addition to its role as a proximal mediator of ErbB2/Neu signaling, Grb2 itself was found to be overexpressed in several breast cancer cell lines and breast cancer tissue samples [35, 36], enhancing signaling through the MAPK pathway. Grb2 is also important in polyomavirus-induced mammary carcinoma, and Grb2 gene dosage is rate limiting for the onset and development of mammary carcinomas [25], highlighting its critical role in the transformation process. Grb2 is involved in keratinocyte growth factor (KGF)–induced motility in MCF-7 breast cancer cells [37] further suggesting that Grb2 can be a valid therapeutic target for pathological processes such as the spread of solid tumors through local invasion and metastasis. In bladder cancer cells where no EGFR overexpression or H-Ras mutations have been observed, Grb2 has been found overexpressed together with Sos1, as the only observed mechanism of oncogenesis [38]. In the highly metastatic cancer cell line 1-LN, Grb2 was one of the effector proteins significantly induced, together with Sos1, Shc, and Raf1, through activation of the [alpha]2-macroglobulin receptor [39]. Another group, using immunohistochemical analysis of tissue microarray from more than 1,000 specimens, has reported the overexpression of Grb2, together with overexpression of Her2, in gastric carcinoma, specifically with an increase of Grb2 expression in primary cancer and nodal metastasis when compared with normal gastric mucosa [40]; moreover, Grb2 overexpression was associated with poor survival rates, supporting a role for Grb2 in tumor cell aggressiveness. The same group has previously investigated Grb2 expression in colorectal cancers and found a significant increase in specimens derived from metastatic lesions [41]. Consistent with a role in tumor dissemination, several groups have reported specific direct and indirect interactions of Grb2 with molecules involved in cytoskeleton remodeling, motility, and other cellular processes recapitulated in the multistep cascade of cancer metastasis. Inhibitors of Grb2 have been demonstrated to reduce motility in vitro and decrease cancer metastasis in animal models [42–44]. Several studies, summarized below, elucidate the molecular mechanisms by which Grb2 contributes to cell motility and other processes characteristic of cancer metastasis.

Involvement of Grb2 in Invasion and Metastasis It is well known that the primary cause of mortality in cancer is due to the deleterious consequences of tumor dissemination and metastasis. Despite progressive advancement in our understanding and treatment of cancer over the last decade, the metastatic process remains the least understood at the molecular level [45]. To develop more effective therapies we need to better understand the molecular mechanisms underlying this multistep process and identify novel molecular targets [46]. Many current cancer treatments focus primarily on blocking the proliferation of tumor cells using cytostatic agents and targeted therapies, but these regimens offer limited success, with frequent relapse. Thus, to improve survival rates for most cancers, more effective ways of treating micrometastatic disease are required.

1

GRB2 and Target Therapy

7

Metastasis is a multistep process [47, 48] in which cells from the primary tumor migrate through the extracellular matrix, enter the circulation through newly formed blood vessels (tumor angiogenesis) and disseminate to distant sites (extravasation), where proliferation begins again. Blocking any stage of this process can potentially be an effective strategy to block the entire process of metastatic disease. The contribution of a protein such as Grb2 in oncogenesis goes beyond its original characterization in cell proliferation [49]; emerging evidence shows that Grb2 contributes to tumorigenesis in several other ways and to other stages of cancer development. Direct and indirect interactions between Grb2 and several intracellular proteins involved in the metastatic cascade have been the subject of numerous original studies that I will summarize in the following pages.

Grb2 Involvement in the Early Phases of the Metastatic Cascade Inside the primary tumor, several experimental evidences have demonstrated that changes occur in the adhesion properties of potentially metastatic cells, as an early event in the metastatic cascade [50]. Also the results of a number of clinical studies corroborate this notion, and the expression of several adhesion molecules is altered in different tumors and correlate with prognosis and biological behavior in primary tumor and metastasis [51]. Tumor cells lose their junctions to other cells and to the extracellular matrix (ECM), and display increased motility. In motile cells, new adhesion sites (focal complexes) located within cell edge protrusions (lamellipodia and filopodia) are transient and small compared to the more stable focal adhesions underlying the cell body and localized at the extremities of actin stress fibers [52]. The interaction of ECM components with the cell surface is mediated mainly by members of the integrin family of transmembrane receptors. Integrins do not have intrinsic catalytic activity but rely on the kinase activity of other nearby intracellular proteins. For example, focal adhesion kinase (FAK) co-localizes with integrin receptors upon engagement of the cell with the ECM and constitutes a nodal point in integrin signaling, and an important regulator of cell migration [53]. The overexpression of focal adhesion kinase in several types of tumors is associated with increased angiogenesis, metastasis, and poor prognosis [54]. FAK function is regulated through tyrosine phosphorylation at specific residues in its amino acid sequence. Several stimuli, such as those mediated by integrins, can induce FAK autophosphorylation, creating docking sites for proteins containing SH2 domains, including Src. Src can also activate FAK and promote phosphorylation on other tyrosine residues; one of these residues, Y925, occurs within a consensus sequence (pYXNX) for high-affinity binding to the SH2 domain of Grb2 [55]. Interestingly, elevated Src activity, as observed during colon cancer progression, specifically promotes phosphorylation on tyrosine Y925, inducing changes in integrin adhesion and deregulation of E-cadherin [56], leading to an E-cadherin/N-cadherin switch [57]. This event is part of an important hallmark of cell transformation and metastasis, namely epithelial-mesenchymal transition (EMT).

8

A. Giubellino

The interaction of Grb2 with FAK, Shc, and other proteins also leads to activation of the Ras and ERK2 pathways; integrin engagement of these pathways induces cell spreading through actin cytoskeleton rearrangement. Indeed, one of the differences between ERK2 pathway activation by growth factor receptors versus by integrin receptor is that the latter requires a functional actin cytoskeleton to signal, while growth factor receptors can signal even when the actin microfilaments are disrupted by cytochalasin D, a potent inhibitor of actin polymerization [58, 59]. The discovery of Grb2-mediated adhesion signaling raises questions as to its precise role in this context. In particular it is important to understand the signaling that permits migrating tumor cells to go through a series of continuous attachments and detachments, and how signaling proteins such as Grb2 direct these transitions and promote cell movement. After detaching from neighbor cells and from the ECM, tumor cells have to migrate through the surrounding tissue and reach blood vessels for dissemination. This event is made possible by several proteinases [60], such as the matrix metalloproteinases (MMPs), a family of secreted and transmembrane proteins capable of degrading virtually every component of the extracellular matrix. They function in physiologic processes such as tissue growth, morphogenesis, tissue repair, and angiogenesis, as well as in pathological conditions such as tumor invasion and metastasis [61]. The MMP family includes a class of membrane-anchored metalloproteinases, ADAMs (A Disintegrin And Metalloprotease), involved in proteolytical cleavage and release of membrane-bound growth factors, cytokines and receptors [62]. ADAMs overexpression and dysregulation have been implicated in angiogenesis and metastasis, especially in ErbB ligand cleavage and activation as well as in the processing of other proteins involved in oncogenesis [63]. Selective ADAMs inhibitors have been developed in recent years and are currently entering clinical trials as a therapeutic strategy in many cancers, reinvigorating the hopes placed earlier on MMP inhibitors [64, 65]. Grb2 is also involved in ECM remodeling; in fact the transmembrane metalloproteinase ADAM12 (Meltrin alpha), upregulated in several cancers, interacts directly with the SH3 domains of Grb2 through prolin-rich sequences in its cytoplasmic tail [66]. Moreover, Grb2 and ADAM12 co-localize at membrane ruffles, structures visible specifically during epithelial cell migration. Another protein in this family, ADAM15 (metargidin) also interacts with Src and Grb2 in vitro through prolinerich sequences in the cytoplasmic domain [67]. In addition to weakening the structural integrity of the ECM, these proteolytic activities can liberate matrix-bond chemokines and growth factors that further enhance the motility of cancer cells. Growth factors, upon binding to membrane receptors, initiate signaling leading to survival, increase specific metabolic cascades, and other complex activities such as motility and invasion. Thus, it is the complex downstream signaling triggered by growth factors that lead to these diverse effects. Most, if not all growth factors have motogenic effects. One of the best characterized growth factors involved in cell motility and invasion is the hepatocyte growth factor (HGF), also known as the Scatter Factor (SF), the ligand for the tyrosine kinase receptor Met [68]. Scattering is a spatially and temporally complex process in which

1

GRB2 and Target Therapy

9

a cluster of grouped cells lose apical/basal polarity and initiate membrane ruffling (centrifugal spreading phase, after 1–3 h), then dissociate from the ECM and neighboring cells through disruption of E-cadherin-mediated cell–cell adhesion (dissociation phase, after 3–6 h), and subsequently migrate (migration phase, after 6 h) [69]. All of these events are triggered upon HGF binding to the extracellular domain of c-Met; intracellularly, the c-Met kinase domain is activated and through a series of auto- and transphosphorylations on specific tyrosine residues, docking sites for effector proteins are created. Grb2 interacts directly with pY1356 in the multifunctional docking site of c-Met, or indirectly through the adaptor protein Gab1 (Grb2 Associated Binder 1). Several downstream pathways have been implicated in this process, such as activation of the GTPase Rac (Rho family) and the small GTPase Ras [70, 71]. Activation of the Ras–Rac1/Cdc42–PAK and Gab1–Crk–C3G—Rap1 effector cascade also regulates cytoskeletal and cell adhesion proteins such as cadherins, Arp2/3, N-Wiskott–Aldrich Syndrome protein (N-WASp), paxillin, integrins and FAK. Grb2 interacts directly with the actin filament machinery. The interaction with WASp [72], a regulator of actin cytoskeletal rearrangement, is a well documented example. Patients affected by a mutation in the gene encoding of this protein show functional defects in platelets, in T- and B-cell polarization, and in the ability of these cells to migrate to external stimuli, resulting in thrombocytopenia, immunodeficiency, and propensity to develop malignancies (mainly leukemia and lymphoma) [73]. The WASp contains proline-rich sequences that mediate interaction with the Grb2-SH3 domains. Binding to Grb2 translocates WASp from the cytosol to the plasma membrane, where it can interact with membrane-bound proteins such as Rac and Cdc42 [74]. Furthermore, Grb2 links the EGF receptor to WASp constitutively and this interaction is enhanced upon EGF stimulation. WASp at the membrane also interacts with Nck [75], and together with Grb2, cooperatively stabilizes the actin-nucleating complex. So Grb2 and Nck, both SH2 and SH3 domain–containing proteins, can link membrane receptors and membrane-bound proteins to intracellular cytoskeletal regulators, increasing their local concentrations at the membrane and facilitating enzymatic reactions and activation. Model organisms, such as Listeria monocytogenes and vaccinia virus, have been used to refine our understanding of the role of Grb2 in actin-based motility. These pathogens hijack the actin cytoskeletal machinery to navigate through the host cytoplasm [75, 76]. N-WASp is necessary for the actin-based motility of vaccinia virus, and in mammalian cells regulate actin polymerization through the Arp2/3 complex (the nucleation factor of newly formed actin filaments) and interaction with Cdc42 [77]. When N-WASp is not present, Grb2 triggers a weak activation of Arp2/3 with defective actin polymerization [78]. Several migratory signaling pathways from the cell surface converge on the p21activating kinases (PAKs), which consequently translocate to the leading edge of the cell and contributes to motility and invasion. Activation of PAKs and translocation to the plasma membrane are processes that involve the interaction with adaptor proteins such as Nck and Grb2. PAKs are serine/threonine kinases that regulate cancer signaling networks and are considered platforms that amplify and propagate oncogenic signals elicited by extracellular stimuli [79]. PAKs are important regulators of actin

10

A. Giubellino Table 1.2 List of the main proteins interacting with the SH2 or the SH3 domain of Grb2 SH2 domain interacting proteins SH3 domain interacting proteins EGFR SOS1 PDGFR WASP VEGFR PAK1 MET GAB1 RON p85/PI3K ERB-B2/B3 JAK1/2 KIT ELK1 LAT ADHALIN FGFR DYNAMIN NGFR SPROUTY1/2 RET ADAM12 EPH B2/B6/A2 MTA1/3 CSF1R SELECTIN-L SHP-1/2 VAV SRC DYSTROGLYCAN FAK FAS ligand CBL HUNTINGTIN CD72 MAP2 ZAP70 CRK CALDESMON CORTACTIN SHC MAGICIN IRS-1 DELTEX1 BCR/ABL DYNACTIN1 MUC1

cytoskeletal dynamics and the role of PAKs in cancer has been widely reported in literature. Specifically, PAK1, the best characterized member of the PAK family, was discovered in 1994 by Manser et al. [80] as a target for the Rho-GTPase related proteins CDC42 and Rac1, well-known regulators of the actin cytoskeleton and implicated in the formation of fingerlike protrusions (filopodia; CDC42) and sheetlike structures at the cell periphery (lamellipodia; Rac1). Interestingly, PAK can directly and specifically interact with Grb2 through a prolin-rich motif in the PAK sequence [81]. This interaction is independent of EGF stimulation, but it is increased after stimulation of the EGF receptor and EGFR-Grb2-PAK1 interaction is required for EGF induced lamellipodia formation. However, partially in contrast with this interaction, Yamaguchi et al. [82] have reported that Grb2 partial silencing using siRNA technology does not block invadopodia formation, at least in the rat mammary adenocarcinoma used by the authors. The complexity of Grb2 involvement in actin-based cell motility is highlighted by the growing list of interacting cytoskeletal proteins; we can schematically classify those connections based on the module of Grb2 mediating the interaction (Table 1.2). In a screening for Merlin-binding proteins, Wiederhold et al. [83] discovered a novel protein, Magicin (Merlin and Grb2 interacting cytoskeletal protein), with a consensus

1

GRB2 and Target Therapy

11

sequence (pYVNG) for the SH2 domain of Grb2. Magicin creates a multiprotein complex with Merlin, although the main interaction seems to be through the SH3 domain of Grb2. Merlin is the product of the Neurofibromatosis 2 (NF2) gene; NF2 mutations give rise to an autosomal dominant syndrome characterized by vestibular schwannomas and meningiomas. The sequence of Merlin shows similarities with ERM proteins (Ezrin, Radixin, and Moesin), that linker membrane proteins to the actin cytoskeleton; Merlin has the FERM domain for interaction with F-actin. Merlin localizes to the leading edge of cells (membrane ruffles), where it colocalizes with actin and contributes to actin assembly and interaction with the cortical cytoskeleton. In A431 human epidermoid carcinoma cells, HGF stimulates the phosphorylation of cortactin, a filamentous actin–binding protein, that translocates from the cytoplasm, where it’s located when inactive, to the cell periphery where it assists the Arp2/3 complex in nucleating actin [84], promoting the formation of lamellipodia; Grb2 can associate directly to cortactin, linking c-met to this molecule. When phosphorylated on serine or threonine by the extracellular-signal-regulated kinase (ERK), cortactin assumes a conformation that allows N-WASp and other nucleation-promoting factors to bind to the cortactin–Arp2/3–actin complex [85]. Cortactin is also overexpressed in certain cancers and is associated with an invasive phenotype, formation of invadopodia, and secretion of MMPs, favoring the spread of cancer cells through tissue [86, 87]. Interestingly, cortactin can bind to the actin and calmodulin-binding protein Caldesmon. Caldesmon, when tyrosine phosphorylated, can interact with the Shc–Grb2–Sos complex, enhancing the interaction between myosin and actin and driving transformation during sarcomagenesis [88]. Grb2 interacts also with POB1 (partner of RalBP1), a protein that shares a homology with Eps15, an epidermal growth factor (EGF) receptor substrate, and has two proline-rich motifs. POB1 interacts with RalBP1 that exhibits GAP activity for Rac and CDC42, controlling the cytoskeleton [89]. The importance of Grb2 in actin polymerization is further underscored in the formation of podosomes. Originally identified in cells of mesenchymal origin, podosomes have also emerged as adhesive structures in epithelial cells. These clusters of actin organized in rosette-like structures have components also found in focal contacts as well as dynamin, cortactin, Arp 2/3, N-WASp, and VASP [90]. Grb2, but not Nck, participates in podosome formation and overexpression of Grb2 interferes with the organization of these structures [91]. Characteristically, these structures disappear when cells become motile and re-form when cells are no longer moving. Not only the actin cytoskeleton seems to be affected by Grb2 mediated signaling, but also the function of another component of the cytoskeleton, the microtubule network. For example, in osteoclasts Grb2 is one of the major binding proteins for dynactin [92], a multisubunit activator of the microtubule motor protein dynein that contributes to cell polarization during migration. Hungtingtin, whose malfunction is associated with Huntington’s Disease, has several prolix-rich motifs and interacts specifically with the Grb2 SH3 domains, when Grb2 is dissociated from Sos (hungtingtin and Sos do not coexist in the same complex), linking hungtingtin to the EGF receptor [93]. Hungtingtin is mainly associated with microtubules and appears to function in cytoskeletal anchoring.

12

A. Giubellino

Another microtubule-associated protein, MAP2 (Microtubule associated protein2), a major cytoskeletal protein in neurons [94] that participates in neuronal morphogenesis, also binds Grb2, linking the microtubule network to multimeric signaling complexes in the cytosol. The number of Grb2 interacting proteins involved in the actin and tubulin cytoskeleton is continuously increasing, reinforcing the idea that Grb2 is playing a fundamental role in this context.

Role of Grb2 in Angiogenesis and Dissemination Detailed evaluation through experimental models and clinical observations has revealed that the metastatic process is extremely inefficient [95]; indeed, of the large number of cancer cells that can be shed into the circulation only few will effectively colonize a secondary site. In the first place, the primary tumors have to establish an effective blood supply in order to maintain its mass and grow. For this reason, tumor cells secrete several growth factors that induce the formation of new blood vessels (angiogenesis) [96]. Angiogenesis is an important step in the transition of the primary tumor to malignancy. In addition to nourishing the tumor, newly formed vessels provide a means to disseminate metastatic cells. Recent findings describe angiogenesis at the primary tumor as a mosaic of tumor cells and endothelial cells, and this structure facilitates the shedding of cancer cells into the systemic circulation. Many of the pathways in which Grb2 is involved are important in the formation of a novel vessel through angiogenesis and lymphangiogenesis. Several growth factors, such as VEGF, angiopoietin-1, FGF2, and HGF, contribute to the development of new blood vessels in physiologic and in pathological conditions, such as tumor angiogenesis. The vascular endothelial growth factor (VEGF) receptor 2 (VEGFR2/KDR) is phosphorylated in response to its ligand, VEGF-A which is secreted by many tumor cells, and this activation lead to the direct recruitment of Grb2, Shc, and Nck [97]. This occurs also for the VEGF receptor 3 (FLT4L). Besides the direct interaction, Grb2 can also bind indirectly through the intercession of Shc [98]. Consequently, Grb2 can lead to cell cycle progression via the Sos/Ras pathway and to cell motility through activation of the Rac1/ Rho pathway. The latter response appears to be sensitive to specific binding antagonists of the SH2 domain of Grb2 [99], with reduction of cell invasion and inhibition of more complex processes, such as endothelial cell tubulogenesis and formation of vessels in the chick chorioallantoic membrane (CAM). Other signaling pathways such as those driven by angiopoietin-1 and fibroblast growth factor-2 (FGF-2) also stimulate angiogenesis through direct and indirect interaction with Grb2, via intermediate proteins such as the FGF receptor substrate 2, Gab1, and Shc [100, 101]. Another mediator of angiogenic signaling is HGF, primarily through a direct effect on vascular endothelial cells. In particular, the activation of Met by HGF has been shown to enhance tumor angiogenesis and the interaction between tumor and endothelial cells, mostly increasing endothelial expression of CD44

1

GRB2 and Target Therapy

13

[102] or integrin expression in cancer cells [103]. These events require the activation of multiple signaling for which Grb2 has been demonstrated to be a key intermediate [104, 105]. Interestingly, NK4, a competitive antagonist of HGF-c-Met association, has been shown to be an angiogenesis inhibitor [106]. Several other angiogenic signaling pathways, including the ones driven by platelet-derived growth factor (PDGF), ALK, and Eph, require Grb2 as a critical downstream effector. Besides its role as a critical effector protein in focal adhesion platforms, FAK through Y925 and Grb2 binding is involved in tumor neo-vascularization. Indeed FAK activation and consequent binding to Grb2 within tumor cells induce the expression of VEGF; elevated VEGF production in tumor cells stimulate endothelial cell, facilitating cell motility and survival, without affecting cell proliferation [107]. Moreover, previous studies have demonstrated that hypoxia conditions, like those experienced inside tumors, is able to increase phosphorylation of FAK with consequent Grb2 binding [108]. Selective antagonists of the Grb2 SH2 domain have already shown promising effects on tumor angiogenesis [99] and may represent important tools to better understand how adaptor proteins are involved in the morphogenetic response of endothelial cells to angiogenic cues and in the formation of blood vessels. The final step in the metastatic cascade consists in the extravasation of circulating tumor cells and the colonization of the secondary site. One of the best characterized models of extravasation is represented by the movement of leukocytes from the circulation to the site of tissue damage or infection, in response to cytokines released by macrophages in the affected tissue. These cytokines induce the expression of a particular class of adhesion molecules, selectins, while specific chemokines act as chemoattractants on circulating leukocytes [109]. Selectins mediate the adhesion of lymphocytes on the endothelial cell luminal surface with a series of low-affinity binding sites, slowing down their speed to complete arrest. Studies on the intracellular signaling events upon L-selectin activation have identified Grb2 as an important adaptor: Grb2 binds directly to L-selectin and trigger activation of Rac2 [110]. L-selectin can enhance the formation of metastasis to lymph nodes in vivo [111]. Endothelial cells in different tissues are not alike and can express different patterns of adhesion molecules; this may explain in part why tumor cells prefer specific organs, skipping organs that are the most logical target based on vascular drainage from the primary tumor. Integrins are also important mediators of cancer cells rolling adhesion by circulating and subsequent extravasation [109]. Integrins expressed on the surface of circulating tumor cells engage the surface of endothelial cells and this binding triggers association of the integrin cytoplasmic tail to the intracellular cytoskeleton; consequently, cells can change their shape and form pseudopodia for facilitating passage between endothelial cells. After successful extravasation, tumor cells must again activate ECM degradation, migrate to the metastatic site and proliferate again. Thus, metastatic cells must switch from a migratory to a proliferative state, again modulating pathways in which Grb2 is a well-known mediator.

14

A. Giubellino

Development of Grb2 Inhibitors as Anticancer Drugs The recognition that Grb2 is important in several oncogenic signaling pathways is the origin of several efforts to selectively disrupt its intracellular network. Specifically, the peculiar structure of the SH2 domain of Grb2 has been the focus of intense study in an effort to synthesize selective antagonists. What is different about the SH2 domain of Grb2 (compared with other SH2 domains) is that it invariably prefers an asparagine at the second position downstream of the phosphotyrosine (pY + 2) and requires the ligand peptide to adopt a b-bend configuration [112]. The basis of these requirements became clear when the structure of Src and Grb2 proteins became available for comparison. For the Src kinase, the peptide that binds the SH2 domain has a linear configuration because it allocates in a broad pocket that is open and elongated, while the peptide sequence that binds to the Grb2 SH2 domain is forced into a much deeper pocket and this explains the sequence requirement at pY + 2 and the b-bend configuration. Medicinal chemists took advantage of this information and started the rational design and synthesis of sophisticated compounds. Several potent binding antagonists of the SH2 domain of Grb2 have been developed in the last decade. Short phosphotyrosine-containing peptides mimicking the motif within Grb2 target proteins were used as the starting point for the synthesis of this class of binding antagonists. A major challenge in designing SH2 domain antagonists is the presence of negative charges from phosphate and acidic side chains, which may present an obstacle to penetrating the cell membrane. Conferring phosphatase resistance through modification of the phosphotyrosine is also a big challenge because this can adversely affect SH2 domain–binding affinity. In addition, selectivity is essential to avoid random antagonism of SH2 domain interactions. Thus it is not surprising that the development of high affinity, stable, and cell-permeable SH2 inhibitors has been a difficult challenge; nonetheless, several advancements have been made. To confer phosphatase resistance the phosphotyrosyl residue has been substituted by phosphonomethyl phenylalanine (Pmp) residue, or similar structures [113]; another approach was to use non-phosphate containing ligands [114]. More recently, Song et al. [115] reported the synthesis of potent inhibitor free of phosphotyrosine or any phosphotyrosyl mimetic, demonstrating that complete replacement of the phosphotyrosyl residue can be accomplished without significant loss of binding affinity. High-affinity compounds, able to block EGFR-Grb2 interactions in intact cells [116] were derived through systematic and stepwise substitution of the backbone motif pYxN and mimicking the b-turn conformation. Starting from this platform, potent peptidomimetic inhibitors of SH2 domain interactions have been developed [117, 118]. Macrocyclization has been adopted to stabilize the b-turn conformation, obtaining potent peptidomimetic inhibitors. Other modifications have been identified that increase inhibitor affinity and the potential for cell membrane penetration, and it has been demonstrated that it is possible to design Grb2 Sh2 domain antagonists devoid of any peptidic character [119].

1

GRB2 and Target Therapy

15

Using synthetic binding antagonists of the Grb2 SH2 domain, Atabey et al. [120] demonstrated their potent blockade of HGF-stimulated cell motility, matrix invasion, and branching morphogenesis in epithelial and hematopoietic target cell models. The same compounds did not affect HGF-stimulated mitogenesis. Unpublished evidence suggests that these compounds can block mitogenesis in specific cell types, implying that dependence on Grb2 for mitogenic signaling may be cell-type specific. Interestingly, no evidence of toxicity, or loss of contractility required for cellular functions other than locomotion were observed. The same compounds were also tested for inhibition of the basic morphogenic events required for angiogenesis, such as the HGF, VEGF, and bFGF- driven endothelial cell migration and invasion [99], and inhibition of vasculogenesis in vivo in a chick chorioallantoic assay. Testing of a prototypical antagonist in two aggressive tumor models revealed inhibition of tumor metastasis without affecting primary tumor growth, thus highlighting the critical role of the Grb2 adapter function in motility, invasion, and the spreading of solid tumors [121]. Targeting the SH3 domain of Grb2 has been another important therapeutic strategy. The structure of these modules is known in great detail. The ligand binding surface of SH3 domains is relatively flat and hydrophobic and consists of three pockets characterized by conserved aromatic residues. The ligand typically occupies two of these pockets with two hydrophobic prolines, while the third pocket frequently interacts with basic residues, in a so called polyproline-2 (PPII) conformation [122]. The same conformation is recognized by other proline-recognizing modules such as WW and profilin domains [122], and this similarity has been the subject of several studies to better understand the selectivity of SH3 domains for target proteins. The importance of conformation over sequence for SH3 domain ligand recognition is also gaining acceptance. Although SH3 domains have long been thought to bind preferentially to proline rich sequences of the form PXXP in target proteins, several exceptions to this rule have been found [123]. The SH3 domains of Grb2 have been found to bind a RXXK core consensus motif [124]. The relatively low binding affinities reported for SH3/target protein interactions, together with the ability of SH3 domains from different proteins to recognize the same target, have raised questions regarding the basis for SH3 domain selectivities observed in intact cells as well as the feasibility of screening peptide libraries using SH3 domains to discover binding antagonists [125]. Nonetheless, progress has been made on both fronts [126]. For Grb2 in particular, SH3 domain target selectivity may be increased through two mechanisms, one intrinsic to the SH3 domains themselves and the other through their context in an SH2 domain containing protein. For example, both Grb2 SH3 domains may interact simultaneously with different sites on a single target molecule, e.g., SOS1, thereby increasing both the apparent affinity and selectivity of Grb2-target interaction [123]. Because the Grb2 SH2 domain restricts Grb2 subcellular localization, the pool of potential SH3 domain binding partners is also likely to be limited, further increasing their apparent selectivity. The SH3 domains from several proteins have been used to screen short peptide libraries in search of binding antagonists. Peptides binding the SH3 domain containing adaptor protein Mona/Gads with high affinity have been identified, and subsequent

16

A. Giubellino

structural studies using one of these peptides revealed a novel type of peptide-SH3 domain interaction [127]. Similarly, screening for peptides binding the SH3 domains of the C. elegans Grb2 homolog Sem-5 yielded a bivalent peptide ligand with nanomolar affinity [128]. Pak1-derived peptides encompassing proline-rich sequences in that protein were found to specifically disrupt Grb2 SH3 domain-Pak1 interactions with relevant impact on growth factor mediated migration and lamellipodia formation [81]. Non-natural amino acids analogs have been substituted at the proline-requiring site of Grb2 SH3 domain ligands [129]. These have provided peptides with nanomolar affinity and reinforced the concept that proline residues may be dispensable in the design of SH3 domain–binding antagonists. An adaptation of the target peptide screening approach in developing SH3 domain–binding antagonists has been to systematically introduce point mutations in target peptide sequences. High-affinity peptides capable of blocking the proliferation of primary blast cell cultures derived from patients with chronic myelogenous leukemia (CML) and Bcr/Abl positive cell lines have been developed using this strategy [130]. Subsequent modifications of these peptides to improve their ability to permeate cells yielded agents that more potently disrupted Grb2 signaling complexes in CML-derived cells [131]. These preclinical studies support the concept that Grb2 SH3 domain binding antagonists could provide a therapeutic alternative for CML patients developing resistance to standard treatments [132]. Enhancing the affinity and selectivity of artificial SH3 domain–binding antagonists by exploiting the existence of two SH3 domains in Grb2, dimeric peptides with high affinity binding to both SH3 domains of Grb2 have been designed with the goal of disrupting Grb2-SOS1 interactions [133]. These “peptidimers” inhibited cell growth in vitro and displayed anti-tumor effects in xenograft models, and thus represent the first examples of in vivo activity for this class of compounds [134]. Introducing N-alkylated residues into both monomers of the peptidimer and optimizing the linker improved the affinity for Grb2 to the subnanomolar range [135]. Finally, non-peptidic small molecule inhibitors have also been explored. The first example is the Src signal transduction inhibitor UCS15A that disrupts several SH3 domain mediated interactions, including those of Grb2 [136]. Although target selectivity remains to be improved, this and similar chemical structures may provide a platform for the development of small synthetic drugs that potently antagonize specific SH3 domain–binding interactions. As a final point, it is noteworthy to remember that other potential therapeutic approaches have been explored. Gene silencing using RNA interference (RNAi) has been used to explore the role of Grb2 in signal transduction [137] and in receptor downregulation [138]. Studies on leukemia and breast cancer using antisense oligonucleotide showed inhibition of proliferation, but more studies using RNAi technology to exploit its therapeutic potential in the inhibition of Grb2 are needed to understand if this is an effective strategy. Lentiviral-mediated silencing of Grb2 through RNAi can be envisioned as a potential approach that has been proved already to be effective for other therapeutic targets and in other diseases [139]. Further studies are necessary to refine our understanding of the complexity of Grb2 signaling and to better understand how Grb2 is linked to tumor metastasis in

1

GRB2 and Target Therapy

17

different tumor types. A promising and powerful strategy to explore target protein selectivity and mechanism of action of these compounds is the synthesis of tool compounds (e.g., modified with chemical tags, such as biotin) [140]. This and other strategies, such as the use of microarrays for the global analysis of gene expression profiles, will be extensively used to develop pharmacodynamic markers of drug action, a high priority in any modern drug development effort.

Conclusions and Prospectives The fundamental contribution of adaptor proteins to complex cell signaling cascades and the consequent appropriate cellular responses is outlined by many experimental observations. Several adaptor proteins have been involved directly or indirectly in the pathogenesis of numerous diseases, including cancer. To cite a few examples, the adaptor protein Shc is broadly involved in oncogenesis and in tumor progression [141]; Nedd9, an adaptor protein related to p130CAS, has been considered a melanoma metastasis gene, contributing to invasion and the spread of transformed cells [142]; the adaptors Nck and Crk, promoting the local state of actin polymerization, contribute to tumor progression and immune disorders [143]. Moreover, it has been observed that the role of adaptors may go beyond their function as mere effectors of kinase-initiated signaling. For example, the integrity of the SH2 and SH3 domains of tyrosine kinase c-Src are required for facilitating cell spreading, while its catalytic activity is dispensable [144]. The adaptor protein Grb2 has been the focus of extensive research and significant advances have been made in the understanding of its biology and the role it plays in specific signaling systems. Nonetheless, much more remains to be uncovered. A number of proteins that bind to Grb2 have been identified and their interaction characterized. The role of Grb2 as a signal-associated transducer of several cancerrelevant receptors and the broad involvement of Grb2 in several steps of tumorigenesis and tumor progression make it an excellent candidate for anti-tumor therapeutic strategies. Using Grb2 SH2 domain antagonists we have already learned important information about Grb2 and growth factor-induced signaling. Originally designed to block cell proliferation, we have discovered that compounds that selectively antagonize Grb2 potently inhibit cell migration, invasion, and angiogenesis; and promising evidence points to a potential use of these compounds as anti-metastatic therapeutics [145]. In vivo evidence of Grb2 dimers, and crystal structures of these dimers [146], may become the basis for development of dimeric small molecules as a possible strategy, such as has been explored for other therapeutic targets. However, more experimental evidence to prove that such dimeric form exists in solution, and in cells, plus modeling evidence that could finely estimate the distance between the two Grb2 SH2 domains in solution is needed to rationally undertake/attempt the synthesis of dimeric Grb2 inhibitors. Moreover, other experimental evidence is required to demonstrate the biological role of such dimerization.

18

A. Giubellino

Considering the fundamental role of adapter proteins such as Grb2, and other adaptors in cell signaling and in the development of human cancer, in the coming years we anticipate more work on developing specific drugs that selectively target these proteins, both to treat human disease and as a tool to deepen our knowledge of the role of adapter proteins in normal as well as diseased cell signaling. An interesting option is the use of these molecules in combination with conventional chemotherapeutics or with targeted therapies. Further studies are needed to determine the tumor-specific biology of Grb2 and other adaptor proteins in diverse human cancers in order to determine where and when these inhibitors may provide the best benefit. Considering the wide-spread presence of Grb2 in various tissues, it will be important to also address issues related to potential systemic toxicity of selective antagonists. Rationally designed drugs targeting specific molecules involved in oncogenesis and metastasis is still in its infancy. It will be important to define targets that are most likely to be uniquely involved in these processes. To address the problem of low efficacy for some single agent therapies, and to target a complex process such as metastasis, combinations of drugs acting at different signaling nodes will probably be needed. Drugs targeting protein–protein interactions hold the promise of fewer side effects and when used in combinations, may realize a better therapeutic efficacy than conventional medications. Disease-relevant intracellular protein–protein interactions are promising drug targets, especially where cancer cells are dependent on the activity of a specific oncogene. Indeed, oncogenic-relevant proteins (such as Bcr/Abl, Src, and Ras) are not acting alone as single proteins, but we need to consider them as complex protein networks that are perturbed and remodeled upon drug action [147]. The future will reserve a predominant importance of systems biology to the study of intracellular signaling complexes; a better understanding of the complexity of the interactome will allow for interpretation and understanding of pathological processes and help to determine therapeutic strategy to target this complexity. It is becoming progressively clear that the era of the “magic bullet,” as theorized by Paul Ehrlich [148], is leaving room for a new era where disciplines such as network medicine [149], network pharmacology [150] and system biology of cell signaling [151] bring the promise of a better understanding of how we will select targets and use drug combinations. A systems biology approach will most likely determine which molecular targets will be more useful to target in combination to reduce single drug dosages and achieve higher efficacy, reducing the potential of toxicity resulting from using higher doses of a single agent. In line with this view, adaptor proteins will be an important target for combination therapies.

References 1. T. Pawson, P. Nash, Science 300, 445 (2003). 2. T. Hunter, Cell 100, 113 (2000). 3. T. Hunter, Current Opinion in Cell Biology 21, 140 (2009).

1

GRB2 and Target Therapy 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52.

19

T. Pawson, P. Nash, Genes Dev. 14, 1027 (2000). T. Pawson, Cell 116, 191 (2004). J. Roskoski, Biochemical and Biophysical Research Communications 324, 1155 (2004). T. Pawson, Current Opinion in Cell Biology 19, 112 (2007). D. C. Flynn, Oncogene 20, 6270 (2001). M. T. Uhlik et al., Journal of Molecular Biology 345, 1 (2005). I. Sadowski, J. C. Stone, T. Pawson, Mol. Cell. Biol. 6, 4396 (1986). T. Pawson, Oncogene 3, 491 (1988). L. E. M. Marengere, T. Pawson, Journal of Cell Science 107, 97 (1994). E. Y. Skolnik et al., Cell 65, 83 (1991). E. J. Lowenstein, Cell 70, 431 (1992). K. Matuoka, M. Shibata, A. Yamakawa, T. Takenawa, Proceedings of the National Academy of Sciences of the United States of America 89, 9015 (1992). B. Margolis et al., Proceedings of the National Academy of Sciences of the United States of America 89, 8894 (1992). D. Stein et al., EMBO Journal 13, 1331 (1994). J. Manser, W. B. Wood, Developmental Genetics 11, 49 (1990). T. L. Shen, J. L. Guan, Frontiers in Bioscience 9, 192 (2004). M. A. Lim, H. Riedel, F. Liu, Frontiers in Bioscience 9, 387 (2004). E. J. Lowenstein et al., Cell 70, 431 (1992). P. G. Dharmawardana, B. Peruzzi, A. Giubellino, J. Burke, D. P. Bottaro, Anti-Cancer Drugs 17, 13 (2006). A. M. Tari, G. Lopez-Berestein, Seminars in Oncology 28, 142 (2001). A. M. Cheng et al., Cell 95, 793 (1998). T. M. Saxton et al., Current Biology 11, 662 (2001). I. Fath et al., Science 264, 971 (1994). I. Rey et al., Cell Death and Differentiation 2, 105 (1995). F. Romero, The Journal of biological chemistry 273, 7776 (1998). S. K. Liu, C. J. McGlade, Oncogene 17, 3073 (1998). N. Dimasi, International Journal of Biochemistry and Cell Biology 39, 109 (2007). G. S. Feng et al., J. Biol. Chem. 271, 12129 (1996). S. Li, A. D. Couvillon, B. B. Brasher, R. A. Van Etten, EMBO Journal 20, 6793 (2001). K. Huebner et al., Genomics 22, 281 (1994). A. M. Pendergast et al., Cell 75, 175 (1993). R. J. Daly, M. D. Binder, R. L. Sutherland, Oncogene 9, 2723 (1994). B. S. Verbeek, S. S. driaansen-Slot, G. Rijksen, T. M. Vroom, Journal of Pathology 183, 195 (1997). X. P. Zang, D. R. Siwak, T. X. Nguyen, A. M. Tari, J. T. Pento, Clinical and Experimental Metastasis 21, 437 (2004). T. Watanabe et al., IUBMB Life 49, 317 (2000). U. K. Misra, S. V. Pizzo, Cellular Signalling 16, 487 (2004). G. Z. Yu, Y. Chen, J. J. Wang, Journal of Cancer Research and Clinical Oncology, in press. G. Z. Yu et al., Oncology Reports 19, 1191 (2008). B. Gay et al., J. Biol. Chem. 274, 23311 (1999). N. Atabey et al., J. Biol. Chem. 276, 14308 (2001). A. Giubellino et al., Cancer Res 67, 6012 (2007). G. Christofori, Nature 441, 444 (2006). P. S. Steeg, D. Theodorescu, Nature Clinical Practice Oncology 5, 206 (2008). I. J. Fidler, Nat Rev Cancer 3, 453 (2003). P. S. Steeg, Nat Med 12, 895 (2006). J. Downward, FEBS Letters 338, 113 (1994). U. Cavallaro, G. Christofori, Nat Rev Cancer 4, 118 (2004). K. A. Paschos, D. Canovas, N. C. Bird, Cellular Signalling 21, 665 (2009). S. K. Mitra, D. A. Hanson, D. D. Schlaepfer, Nat Rev Mol Cell Biol 6, 56 (2005).

20

A. Giubellino

53. B. D. Cox, M. Natarajan, M. R. Stettner, C. L. Gladson, Journal of Cellular Biochemistry 99, 36 (2006). 54. G. W. McLean et al., Nat Rev Cancer 5, 505 (2005). 55. D. D. Schlaepfer, S. K. Hanks, T. Hunter, P. Van der Geer, Nature 372, 786 (1994). 56. E. Avizienyte et al., Nat Cell Biol 4, 632 (2002). 57. R. B. Hazan, R. Qiao, R. Keren, I. Badano, K. Suyama, Annals of the New York Academy of Sciences 1014, 155 (2004). 58. S. Miyamoto et al., Journal of Cell Biology 131, 791 (1995). 59. D. D. Schlaepfer, K. C. Jones, T. Hunter, Mol. Cell. Biol. 18, 2571 (1998). 60. C. J. Malemud, Frontiers in Bioscience 11, 1696 (2006). 61. M. D. Martin, L. M. Matrisian, Cancer and Metastasis Reviews 26, 717 (2007). 62. M. L. Moss, M. H. Lambert, Essays in Biochemistry 38, 141 (2002). 63. C. P. Blobel, Nat Rev Mol Cell Biol 6, 32 (2005). 64. B. B. Zhou et al., Expert Opinion on Investigational Drugs 14, 591 (2005). 65. J. S. Fridman et al., Clin Cancer Res 13, 1892 (2007). 66. A. Suzuki et al., Oncogene 19, 5842 (2000). 67. Z. Poghosyan et al., J. Biol. Chem. 277, 4999 (2002). 68. C. Birchmeier, W. Birchmeier, E. Gherardi, G. F. Vande Woude, Nat Rev Mol Cell Biol 4, 915 (2003). 69. J. F. Lai et al., J. Biol. Chem. 275, 7474 (2000). 70. A. J. Ridley, P. M. Comoglio, A. Hall, Mol. Cell. Biol. 15, 1110 (1995). 71. G. Hartmann, K. M. Weidner, H. Schwarz, W. Birchmeier, J. Biol. Chem. 269, 21936 (1994). 72. H. She et al., Mol. Biol. Cell 8, 1709 (1997). 73. H. D. Ochs, A. J. Thrasher, Journal of Allergy and Clinical Immunology 117, 725 (2006). 74. P. Aspenstrom, U. Lindberg, A. Hall, Current Biology 6, 70 (1996). 75. N. Scaplehorn et al., Current Biology 12, 740 (2002). 76. M. F. Carlier, C. Le Clainche, S. Wiesner, D. Pantaloni, Bioessays 25, 336 (2003). 77. H. N. Higgs, T. D. Pollard, Annual Review of Biochemistry 70, 649 (2001). 78. M. F. Carlier et al., J. Biol. Chem. 275, 21946 (2000). 79. R. Kumar, A. E. Gururaj, C. J. Barnes, Nat Rev Cancer 6, 459 (2006). 80. E. Manser, T. Leung, H. Salihuddin, Z. S. Zhao, L. Lim, Nature 367, 40 (1994). 81. L. A. Puto, K. Pestonjamasp, C. C. King, G. M. Bokoch, J. Biol. Chem. 278, 9388 (2003). 82. H. Yamaguchi et al., J. Cell Biol. 168, 441 (2005). 83. T. Wiederhold et al., Oncogene 23, 8815 (2004). 84. L. Crostella, S. Lidder, R. Williams, G. G. Skouteris, Oncogene 20, 3735 (2001). 85. N. Martinez-Quiles, H. Y. H. Ho, M. W. Kirschner, N. Ramesh, R. S. Geha, Mol. Cell. Biol. 24, 5269 (2004). 86. A. M. Weaver, Clinical and Experimental Metastasis 23, 97 (2006). 87. E. S. Clark, A. S. Whigham, W. G. Yarbrough, A. M. Weaver, Cancer Res 67, 4227 (2007). 88. J. L. Boerner et al., Oncogene 22, 6679 (0AD). 89. A. Hall, Annual Review of Cell Biology 10, 31 (1994). 90. L. Spinardi, P. C. Marchisio, European Journal of Cell Biology 85, 191 (2006). 91. L. Spinardi et al., Experimental Cell Research 295, 360 (2004). 92. M. Sahni et al., J. Biol. Chem. 271, 33141 (1996). 93. Y. F. Liu, R. C. Deth, D. Devys, J. Biol. Chem. 272, 8121 (1997). 94. R. W. L. Lim, S. Halpain, J. Biol. Chem. 275, 20578 (2000). 95. K. J. Luzzi et al., Am J Pathol 153, 865 (1998). 96. J. Folkman, Seminars in Oncology 29, 15 (2002). 97. J. Kroll, J. Waltenberger, J. Biol. Chem. 272, 32521 (1997). 98. E. Fournier et al., Oncogene 18, 507 (1999). 99. J. V. Soriano et al., Molecular Cancer Therapeutics 3, 1289 (2004). 100. T. Shono, H. Kanetake, S. Kanda, Experimental Cell Research 264, 275 (2001).

1

GRB2 and Target Therapy

21

101. K. V. Stoletov, K. E. Ratcliffe, B. I. Terman, The FASEB journal : official publication of the Federation of American Societies for Experimental Biology 16, 1283 (2002). 102. S. Hiscox, W. G. Jiang, Biochemical and Biophysical Research Communications 233, 1 (1997). 103. N. Kawakami-Kimura et al., British Journal of Cancer 75, 47 (1997). 104. E. M. Rosen et al., CIBA Foundation Symposia 215 (1997). 105. M. Rosario, W. Birchmeier, Trends in Cell Biology 13, 328 (2003). 106. K. Matsumoto, T. Nakamura, Cancer Science 94, 321 (2003). 107. S. K. Mitra et al., Oncogene 25, 5969 (2006). 108. Y. Seko et al., Biochemical and Biophysical Research Communications 262, 290 (1999). 109. A. E. Aplin, A. Howe, S. K. Alahari, R. L. Juliano, Pharmacological Reviews 50, 197 (1998). 110. B. Brenner et al., Proceedings of the National Academy of Sciences 93, 15376 (1996). 111. F. Qian, D. Hanahan, I. L. Weissman, Proceedings of the National Academy of Sciences of the United States of America 98, 3976 (2001). 112. H. W. H. G. Kessels, A. C. Ward, T. N. M. Schumacher, Proceedings of the National Academy of Sciences of the United States of America 99, 8524 (2002). 113. J. Burke et al., Biochemistry«6490 (1994). 114. Z. J. Yao et al., Journal of Medicinal Chemistry 42, 25 (1999). 115. Y. L. Song et al., Journal of Medicinal Chemistry 49, 1585 (2006). 116. B. Gay et al., International Journal of Cancer 83, 235 (1999). 117. K. Machida, B. J. Mayer, Biochimica et Biophysica Acta (BBA) - Proteins & Proteomics 1747, 1 (2005). 118. J. Burke, International Journal of Peptide Research and Therapeutics 12, 33 (2006). 119. G. Caravatti, J. Rahuel, G. Brigitte, P. Furet, Bioorganic and Medicinal Chemistry Letters 9, 1973 (1999). 120. N. Atabey et al., Journal of Biological Chemistry 276, 14308 (2001). 121. A. Giubellino et al., Cancer Research 67, 6012 (2007). 122. B. K. Kay, M. P. Williamson, M. Sudol, FASEB Journal 14, 231 (2000). 123. B. J. Mayer, Journal of Cell Science 114, 1253 (2001). 124. D. M. Berry, P. Nash, S. K. W. Liu, T. Pawson, C. J. McGlade, Current Biology 12, 1336 (2002). 125. A. R. Viguera, J. L. R. Arrondo, A. Musacchio, M. Saraste, L. Serrano, Biochemistry 33, 10925 (1994). 126. S. M. Feller, M. Lewitzky, Current Pharmaceutical Design 12, 529 (2006). 127. M. Harkiolaki et al., EMBO Journal 22, 2571 (2003). 128. M. R. Ferguson et al., Protein Sci 13, 626 (2004). 129. J. T. Nguyen, C. W. Turck, F. E. Cohen, R. N. Zuckermann, W. A. Lim, Science 282, 2088 (1998). 130. C. Kardinal et al., FASEB Journal 14, 1529 (2000). 131. C. Kardinal et al., Blood 98, 1773 (2001). 132. S. M. Feller, G. Tuchscherer, J. Voss, Leukemia and Lymphoma 44, 411 (2003). 133. D. Cussac et al., FASEB Journal 13, 31 (1999). 134. B. Gril et al., International Journal of Cancer 121, 407 (2007). 135. M. Vidal et al., Biochemistry 43, 7336 (2004). 136. C. Oneyama, H. Nakano, S. V. Sharma, Oncogene 21, 2037 (2002). 137. M. Di Fulvio, K. M. Henkels, J. Gomez-Cambronero, Biochemical and Biophysical Research Communications 357, 737 (2007). 138. F. Huang, A. Sorkin, Mol. Biol. Cell 16, 1268 (2005). 139. K. V. Morris, J. J. Rossi, Gene Therapy 13, 553 (2006). 140. A. Giubellino et al., Journal of Medicinal Chemistry 51, 7459 (2008). 141. J. Ursini-Siegel, W. J. Muller, Cell Cycle 7, 1936 (2008). 142. M. Kim et al., Cell 125, 1269 (2006). 143. S. Antoku, K. Saksela, G. M. Rivera, B. J. Mayer, Journal of Cell Science 121, 3071 (2008). 144. K. B. Kaplan, J. R. Swedlow, D. O. Morgan, H. E. Varmus, Genes Dev. 9, 1505 (1995). 145. A. Giubellino, J. Burke, D. P. Bottaro, Expert Opinion on Therapeutic Targets 12, 1021 (2008).

22

A. Giubellino

146. N. Schiering, E. Casale, P. Caccia, P. Giordano, C. Battistini, Biochemistry 39, 13376 (2000). 147. M. Brehme et al., Proceedings of the National Academy of Sciences of the United States of America 106, 7414 (2009). 148. K. Strebhardt, A. Ullrich, Nature Reviews Cancer 8, 473 (2008). 149. T. Pawson, R. Linding, FEBS Letters 582, 1266 (2008). 150. A. L. Hopkins, Nature Chemical Biology 4, 682 (2008). 151. L. Kuepfer, M. Peter, U. Sauer, J. Stelling, Nature Biotechnology 25, 1001 (2007).

Chapter 2

Human Arylamine N-acetyltransferase 1: From Drug Metabolism to Drug Target Fernando Rodrigues-Lima, Julien Dairou, Florent Busi, and Jean-Marie Dupret

Introduction Arylamine N-acetyltransferases (NAT) are xenobiotic-metabolizing enzymes (XME) that participate in the metabolism of numerous aromatic and heterocyclic amine compounds including drugs and carcinogens [1]. NAT enzymes catalyze the transfer of an acetyl moiety from acetyl-CoA (AcCoA) to the nitrogen or oxygen atom of aromatic amines and their N-hydroxylated metabolites (Fig. 2.1). These phase II XME play, thus, an important role in the detoxification and/or bioactivation of several therapeutic drugs and carcinogens [1]. NAT enzymes have been identified in several species including bacteria [2] and lower eukaryotes [3, 4]. In humans, there are two polymorphic NAT loci producing two functional enzymes, NAT1 and NAT2. These two isoenzymes are encoded adjacent to each other on chromosome 8. A third locus encodes a pseudogene [5, 6]. NAT1 and NAT2 are cytosolic enzymes (~31 kDa) that have very similar sequences (81% identity at the amino acid level). They have distinct but overlapping substrate preferences [7]. The two human isoenzymes have different tissue distributions and are differently expressed during development [8]. NAT2 is principally expressed in the liver and in colon epithelium, which is in agreement with its xenobiotic-metabolizing function. Conversely, NAT1 is distributed widely in adult tissues [8]. Identification of promoter regions suggests that NAT1 and NAT2 gene expression is independent and involves different transcription elements and factors [9, 10]. Polymorphisms affecting enzyme activity have been described for both genes and can lead to “slow” or “rapid” acetylator phenotypes. These interindividual variations have been associated with adverse drug reactions and several diseases such as cancer [11, 12]. In addition to the genetic processes which control their

F. Rodrigues-Lima (*) Univ Paris Diderot-Paris 7, Unité de Biologie Fonctionnelle et Adaptative, 75013 Paris, France e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_2, © Springer Science+Business Media, LLC 2012

23

24

F. Rodrigues-Lima et al. CoAS COCH3

R

CoA SH

R

NH2

Arylamine

Arylamide CoAS COCH3

R

NHOH

Hydroxyarylamine

NHCOCH3

CoA SH R

NHOCOCH3

Hydroxyarylamide

Fig. 2.1 Main reactions catalyzed by NAT enzymes. N-acetylation of arylamine from acetyl-CoA (CoASCOCH3) (upper part) and N-acetylation of hydroxyarylamines from acetyl-CoA (lower part)

expression, the activity of human NAT enzymes, particularly NAT1, can also be modulated by “nongenetic factors” such as substrate-dependent downregulation, drug inhibition, or biological oxidants [13–15]. The genetic polymorphism in NAT activity was first identified in patients treated with isoniazid for tuberculosis [16]. This drug is primarily excreted following acetylation by human NAT2. Since then, many therapeutic drugs such as hydralazine or sulfamethazine have been shown to be polymorphically acetylated in humans. It is now known that human NAT1 and NAT2 loci are highly polymorphic, with more than 26 and 53 alleles identified at the NAT1 and NAT2 locus, respectively. Several of these NAT1 and NAT2 alleles can lead to slow acetylator phenotypes due to altered enzyme activity [1]. Common consequences of the polymorphic acetylation of therapeutic drugs are drug-associated adverse effects either due to rapid or slow acetylation [17]. NAT-dependent acetylation plays an important role in the metabolic activation and detoxification of environmental carcinogens such as 4-aminobiphenyl (4-ABP) or benzidine and changes in the N- and/or O-acetylation of these aromatic chemicals have been linked to carcinogenesis [18]. In addition to its role in aromatic amine xenobiotic metabolism, there is strong evidence for an endogenous role of human NAT1 in folate catabolism through the NAT1-dependent N-acetylation of p-aminobenzoylglutamate (p-ABAGlu) (Fig. 2.2) [19]. More recently, several data have been published suggesting that NAT1 plays a biological role in breast cancer progression [20–22] and that this XME could be targeted in treatment [2, 23]. Proteomic and microarray studies have shown that in breast cancer there is overexpression of the human NAT1 gene in estrogen receptorpositive tumors [20, 22]. In addition, it has been reported that overexpression of human NAT1 in luminal epithelial cells leads to enhanced growth and drug resistance [20]. These recent data suggest that human NAT1 could be targeted in breast cancer treatment [2]. Very recently, the first human NAT1-specific inhibitors have been synthesized and tested in human breast cancer cell lines [23].

2

Human Arylamine N-acetyltransferase 1: From Drug Metabolism to Drug Target O

25

OH

O OH OH

N H N

O N H

N

NH2

N

N

Folic Acid

O

OH

O OH OH N

N H

N

O NH2

N

H2N

N

PABA-Glu

6 Methyl-pteridine

NAT1 O

OH

O OH N H

O

O H3C

N H

N-acetylPABA-Glu

Fig. 2.2 Potential role of NAT1 in folate catabolism through N-acetylation of p-ABAGlu

Human NAT1 Gene Structure and Polymorphism Human NAT1 gene is located on chromosome 8 at 8p22, a chromosomal region commonly deleted in certain human cancers [6]. NAT1 gene consists of at least nine exons spanning 52 kb [9]. The full coding sequence (870 bp) is contained within exon 9. Several NAT1 transcripts have been described which contain various combinations of the exons in the 5’ UTR [9, 24]. These mRNA originate mainly from two different promoters (Fig. 2.3) named NATa and NATb [25] which are, respectively, located 51.5 and 11.8 kb upstream of the single translated exon.

26 Fig. 2.3 Structure of the human NAT1 gene

F. Rodrigues-Lima et al. gene structure: NATa

1 2

NATb

3 4

5 6

7 8

9

open reading frame exon promoter

major mRNA:

4

8

9

Fig. 2.4 Structure of human NAT1 showing the positions of polymorphic variants. Backbone is shown as ribbons; wild-type residues are shown in ball-and-stick and correspond to the enzyme coded by NAT1*4 allele

A proximal promoter at −245 bp has also been described [26] but is unlikely to be of biological importance [25]. Transcription of NAT1 gene occurs in a wide variety of tissues and cell lines, and most mRNAs are initiated at the NATb promoter which is upstream of exon 4. This promoter gives rise to at least five different transcripts which have different translational efficiencies, although the biological significance of this is unknown [9, 24]. The most common transcript is the shortest and comprises exons 4, 8, and 9 only (Fig. 2.4). RT-PCR studies [27] showed that the alternative promoter, NATa, is most highly expressed in a few tissues, including lung, trachea, kidney, and liver suggesting that the activity of this promoter is tissue specific. Two major and four major transcripts arising from the NATa promoter and beginning with exon 1 have been reported [9]. So far, the biological significance of these different transcripts remains unknown [25]. The pattern of placental NAT1 expression during human development and the spatial and temporal expression of the murine ortholog of human NAT1 (Nat2) are likely due to transcriptional regulation [28]. Recently, androgens have been

2

Human Arylamine N-acetyltransferase 1: From Drug Metabolism to Drug Target

27

shown to increase NAT1 mRNA transcription from the NATb promoter in two prostate cancer cell lines [29]. In addition, high levels of NAT1 gene expression have been described in the estrogen receptor-positive human breast cancer ZR-75-I cell line correlated with high transcription level from the NATa promoter [22]. Expression of human NAT1 and NAT2 genes differs during development. Contrary to NAT2, NAT1 gene is expressed in placenta in the first trimester and at the blastocyst stage [8]. In addition to its expression in tissues possessing xenobiotic metabolism functions, NAT1 is also known to be expressed in tissues not normally associated with xenobiotic metabolism such as lymphocytes and red blood cells [8]. The large tissular expression of human NAT1 compared to NAT2 further supports the hypothesis that NAT1 could be involved in endogenous functions. The existence of polymorphic variants of the human NAT enzymes was first identified through population genetic studies of phenotypic variation [16]. NAT1 has long been thought to be genetically invariant. However, in 1993 several allelic variants at the NAT1 locus were first described [30]. To date, 26 different NAT1 alleles have been detected in human populations [31, 32]. Phenotypic differences are mainly due to single nucleotide polymorphisms (SNP) which are inherited effectively as haplotypes with up to six SNP per allele [33]. These SNPs and the NAT1 alleles are described in detail on the website http://louisville.edu/medschool/ pharmacology/NAT.html. Most SNPs (19 out of 21) occur in the coding region (exon 9). Certain SNPs have also been described in the 5¢ and 3¢UTR regions of NAT1; however, their functional effects are poorly understood [31]. In addition to SNPs, deletions and insertions within the 3’UTR region have also been reported. Several SNPs in the coding region lead to amino acid substitutions leading to enzyme variants with reduced or no activity [34]. Certain SNPs can also lead to truncated NAT1 enzyme [2]. As indicated above, NAT1 polymorphism can result in NAT1 protein variants with altered enzymatic activity or decreased abundance in human cells [2]. These genetic mechanisms are the basis for the existence of “slow” versus “rapid” NAT1 acetylator groups. Loss of function alleles that results in a slow acetylator phenotype include NAT1*14, NAT1*15, NAT1*17, NAT1*19, and NAT1*22 [31]. The effect of the human NAT1*10 allele, which in certain cases appears to increase activity and in other cases appears to have no effect [2], may be an example of the complex effects of polymorphisms in the noncoding region on transcription and the extent of translation of this transcript [32]. Contrary to NAT2, human NAT1 gene shows less variation in allelic frequencies among populations which may reflect a selective advantage in minimizing the persistence of various NAT1 alleles [35]. The reference NAT1*4 allele (~70% in populations of European descent) and NAT1*10 which contains two SNPs outside the coding region (~20% in populations of European descent) are common in most ethnic groups studied so far. Slow acetylator alleles account for less than 3% of the NAT1 alleles found in persons of European descent [11] but up to 25% in certain populations [36]. Decreased cellular activity of most NAT1 variants correlates well with reduced proteins levels. Most of these variants did not exhibit transcription deficiency [2]

28

F. Rodrigues-Lima et al.

indicating that the reduced cellular levels of these NAT1 alloenzymes are caused by mechanisms that act on NAT1 protein. Work from Hein’s group suggested that low intrinsic stability leading to intracellular degradation could account for the weak cellular activity of certain NAT1 variant [34]. Minchin and collaborators proposed a model for the rapid degradation of certain NAT1 variants by the proteasome. They hypothesized that NAT1 exists in the cell in either a stable acetylated state or an unstable non-acetylated state [14]. The variant alloenzymes may contain structural features that prevent catalytic formation of an acetylthioester and hence favor the formation of non-acetylated enzymes. These forms are more readily ubiquitinated and degraded by the protease, resulting in low amounts of NAT1 [14]. Recent work from Walters’s group also support the notion that low cellular NAT1 activity of certain variants is due to cellular degradation likely due to incorrect folding. Using green fluorescent protein-tagged NAT1 variants, they showed that contrary to the “wild-type,” active NAT1 enzyme which is distributed in the cytosol, certain variants are found clustered in aggresomes that are rapidly ubiquitinated and degraded [37]. It appears from all these studies that NAT1 slow acetylator phenotypes are associated with SNPs in the coding region that alter the NAT1 protein structure leading to its degradation in cells.

Human NAT1 Protein Structure The understanding of the structural characteristic of the NAT1 enzyme began with the identification of sulfhydryl-containing residues as being essential for acetylation of the substrates [38]. Site-directed mutagenesis confirmed this point by showing that a cysteine residue conserved in all NAT enzymes is crucial for catalytic activity [39]. In 2000, the crystal structure of the bacterial NAT from Salmonella typhimurium gave a clear understanding of the active site of NAT enzymes [40]. Unexpectedly, the structure showed a cysteine protease-like catalytic triad (CysHis-Asp) that appears to be conserved in all NAT enzymes [40]. Since then, the structure of several bacterial NAT enzymes have been reported [2, 41]. In 2007, the structures of the human NAT1 and NAT2 isoforms were published [42]. Overall, these structural studies indicate that all NAT enzymes share a conserved “NAT” fold that is related, at least in part (active site core), to cysteine protease enzymes [43]. The NAT fold consists of a three-domain structure of approximately equal length (~100 amino acids). The first domain is predominantly alpha helical, the second domain is predominantly beta sheets and the third domain is alpha–beta lid [2]. The residues of the active site are in domains one and two which are the most highly conserved. The C-terminus is less conserved and has a major role in substrate specificity. Indeed, the positioning of the C-terminal regions of human NAT1 and NAT2 proximal to the catalytic cavity allows for additional interactions between NAT amino acids and the substrates [2]. In addition, its length controls enzymatic activity [44]. Comparison of human and bacterial NAT structures identified the existence of a loop (between domains two and three) which is found only

2

Human Arylamine N-acetyltransferase 1: From Drug Metabolism to Drug Target

29

in eukaryotic NAT enzymes [2]. This “eukaryotic-specific” loop appears to be important for the differences between the CoA binding sites in eukaryotic and prokaryotic NAT enzymes [45]. The recent determination of the structures of the human NAT1 and NAT2 enzymes has rationalized previous comparative studies on the enzyme substrate specificity showing how variations at one position may contribute to selective binding of certain drugs or carcinogens to NAT1 or NAT2 [42, 46]. The crystal structures of NAT1 and NAT2 have contributed to understand the impact of polymorphisms on the enzyme structure and activity [2] and have also allowed the identification of the key bonds required to maintain both the active site and the NAT fold (Fig. 2.4). Comparison of the human and prokaryotic structures is likely to be of prime importance for the rational design of specific inhibitors of human and/or bacterial NAT isoforms [2].

Animal Models to Understand NAT1 Functions NAT enzymes have been identified in different eukaryotic species, in particular in rodents. Animal models have played an important role in understanding the pharmacogenetics variations and the molecular effects of the NAT polymorphisms [31]. Indeed, earlier studies reported variability in NAT activity among different mouse, Syrian hamster, and rat [2]. These animals were later found to possess three functional NAT genes, although one of these genes (Nat3) generates a relatively inactive enzyme. Nat3 is likely to be evolving to become a pseudogene, like the third locus (NATP1) in humans [33]. Mouse models have recently been very useful to understand the role of NAT enzymes and, in particular, of human NAT1. Indeed, mouse Nat2 and human NAT1 share close sequence identity and substrate specificity and have similar tissue distribution [47]. Nat2 is thus considered as the murine ortholog of NAT1, and several studies aimed at understanding the role of human NAT1 have used Nat2 as a model [8]. Knockout mouse strains for Nat2 gene showed that, in small population studies, the mice were devoid of any obvious developmental defects [48]. Inactivation of both Nat1 and Nat2 genes in a double-knockout strain was apparently aphenotypic. However, these mice were found to have compromised metabolic activities toward aromatic amine substrates [49] in agreement with their role in xenobiotic metabolism. In large population studies, strains devoid of Nat2 showed a bias in the sex ratio of live-born offsprings [50]. In addition, the frequency of neural tube defect was increased tenfold in heterozygous Nat2 null/ wt offsprings [51]. Interestingly, Nat2 has been reported to be involved in the susceptibility to teratogen-induced orofacial clefting [2]. It has been suggested that human NAT1 and Nat2 could play a role in the catabolism of folate. Indeed, both enzymes have been found to acetylate the folate catabolite, p-aminobenzoylglutamate (p-ABAGlu) [19, 47] (Fig. 2.2). Recent studies showed that acetylated p-ABAGlu was present in the urine of WT mice fed with a folate supplement,

30

F. Rodrigues-Lima et al.

but not in the urine of Nat2 null/null knockouts [52]. These data further support a putative endogenous role of NAT1/Nat2 in the metabolism of folate. Contrary to the Nat2 gene invalidation, overexpression of its human ortholog NAT1 in transgenic mice appears to have deleterious effects with many lethal malformations, including tail kinks resembling neural tube defects which could be associated with folate deficiency [53]. This is in agreement with a putative role of NAT1 in folate metabolism with excess NAT1 activity promoting folate deficiency [2]. As stated above, although the transgenic mice initially appeared to have no particular phenotype (except compromised aromatic amine metabolism), several studies suggest that deletion of Nat2 is associated with congenital abnormalities and imbalance in the sex ratios of offsprings. Most of these defects could be due to epigenetic effects through changes in DNA methylation [22]. Important studies are in progress to analyze the effects of carcinogen exposures on mice lacking Nat2.

Possible Relevance of NAT1 in Cancer Changes in the NAT-dependent N- and or O-acetylation of aromatic amine compounds have been linked to carcinogenesis [1, 54]. However, several studies strongly suggest that in addition to xenobiotic metabolism, NAT1 may have other functions [2, 32]. Recent data point to a role of NAT1 in cancer mechanisms and in particular in breast tumor biology [2]. In a comparative 2D proteomic study by Adams et al., NAT1 was found to be among the most consistently upregulated proteins in invasive ductal carcinoma and invasive lobular carcinoma when compared to normal human breast tissue [20]. Overexpression of NAT1 in normal breast luminal epithelial cells induced two of the hallmark traits of cancer, i.e., enhanced growth and resistance to etoposide, a therapeutic cytotoxic drug used in cancer treatment [20]. The proteomic data reported by Adams et al. were further supported by detailed analysis of a series of microarray-based studies that compared the expression profiles of thousands of human genes between malignant and normal breast tissue, or between estrogen receptor-positive (ER+) and ER-negative (ER−) breast tumors. The results of these microarray-based studies have been recently reviewed by Wakefield et al. [22] and are available through the oncomine website (http://www.oncomine.org/). NAT1 gene appears among the five most highly overexpressed genes in ER+ and in progesterone receptor-positive tumors. In addition, NAT1 was found to be among the genes that are downregulated in the late-stage breast tumors [22]. The molecular basis for the upregulation of NAT1 gene is not known. In their recent study, Wakefield et al. [22] showed that the levels of NAT1 in different ER+ breast cancer cell lines differ significantly (up to 40 times higher activity for the human ZR-75-1 cell line compared to the others). Elevated levels of NAT1 were not due to alteration in gene copy number in this cell line. Further analysis indicated that only the cell line with highest NAT1 activity (ZR-75-1) had NAT1 transcripts originating from the distal promoter NATa. The other cell lines tested had much less NAT1 activity, and NAT1

2

Human Arylamine N-acetyltransferase 1: From Drug Metabolism to Drug Target

31

transcripts were derived from the NATb promoter (Fig. 2.3). Interestingly, previous analysis of promoter use in normal human tissue indicates that the NATa promoter is tissue specific and active in liver, kidney, lung, and trachea [27]. In addition to the difference in the use of the promoters, there is variability in the 3¢ end of the human NAT1 gene transcript that may be associated with different levels of expression in breast cancer cell lines [22]. Moreover, both human NAT genes are encoded within a 200-kb region along with the human NAT pseudogene, and this region of the genome (8p22) [5] is unstable in several tumors, showing loss of heterozygosity or gene duplication [54]. So far, there is no clear evidence linking NAT1 expression and estrogens. Several other results add complexity to the involvement of NAT1 in breast cancer. For instance, Kim et al. [55] have reported differential methylation of the NAT1 gene in breast cancers compared to benign or normal tissue [55] suggesting that genetic and epigenetic mechanisms act on NAT1 expression. Clinical studies reported that high levels of NAT1 expression are associated with longer relapse-free survival of patients with ER+ breast cancer undergoing treatment with tamoxifen [21]. Interestingly, it has been shown that the activity of human NAT1 and its murine counterpart Nat2 could be inhibited by tamoxifen [47, 56]. This suggests that NAT1 may be influenced directly by estrogen agonists or antagonists. Recent data also suggest altered regulation of NAT1 in prostate cancer. Indeed, Butcher et al. [29] have shown the androgen induction of NAT1 in prostate cancer cells which is in agreement with the presence of androgen response elements upstream of the human NAT1 and murine Nat2 coding exons [29]. Androgens are known to be important for the prostate cancer development and progression [57]. In addition, a recent microarray study identified three separate tumor subtypes expressing different levels of NAT1 [58].

NAT1 as a Drug Target in Cancer The increasing evidence for an association of NAT1 with carcinogenesis, in particular with breast cancer, suggests that this XME could be targeted for breast cancer therapy [2]. Interestingly, recent studies have reported that certain well-known anticancer drugs are able to inhibit NAT1 enzyme in vitro and in vivo (Fig. 2.5). Tamoxifen, an antagonist of the estrogen receptor that is currently used to treat hormone-positive breast cancers, has been shown to inhibit NAT1 and its murine ortholog Nat2 [47, 56]. Recently, cisplatin, one of the most important chemotherapeutic compounds used in the treatment of various human cancers [59], has been found to irreversibly inhibit NAT1 [60]. Indeed, studies using recombinant NAT1 protein in human breast cancer cell lines and mice showed that cisplatin, at pharmacologically relevant concentrations, was able to impair the endogenous catalytic functions of NAT1. Mechanistic analysis of this inhibition indicated that NAT1 inactivation was rapid. The secondorder rate constant for the inhibition of NAT1 by cisplatin was found to be the highest reported for a reaction between cisplatin and a biological target [60]. Although these

32

F. Rodrigues-Lima et al. NHOH

N O

O

−

NO

O

COOH

peroxynitrite

Hydroxylamine para -aminobenzoic acid

2-nitrosofluorene

CH3 N O

HO CH3

O

O OH

Cl Cl CH3

tamoxifen

Pt

O

NH2 NH2

cisplatin

11a-hydroxycinnamosmolide

Fig. 2.5 Chemical structure of compounds known to inhibit NAT1 enzyme

data do not demonstrate that alteration of NAT1 functions in breast cancer cells contributes to the therapeutic effect of cisplatin, these results emphasize the putative link between NAT1 and breast cancer. Another recent study also reports that disulfiram also impairs NAT1 functions in human cancer cells [61]. Disulfiram has been used for decades to treat alcoholism [62]. However, several studies indicate that this drug could be used in cancer treatment [62]. Interestingly, disulfiram was found to inhibit enzymes that have been associated with cancer progression such as DNA topoisomerases, matrix metalloproteinases, and proteasome [61, 62]. Specific inhibitors for prokaryotic NAT enzymes have been identified/synthesized over the last years [63]. These studies have shown that inhibition of NAT activity in Mycobacterium bovis has effects similar to deleting the NAT gene [64]. More recently, Edith Sim’s team in Oxford has reported the identification, synthesis, and evaluation of a series of selective inhibitors (rhodanine and thiazolidin-2,4-dione derivatives) for human NAT1 and murine Nat2 [23]. The most potent inhibitor was found to act at submicromolar concentrations and to inhibit both the recombinant enzymes and human NAT1 in ZR-75 breast cancer cells in a competitive manner [23]. It is expected that rational drug design based both on the recently identified inhibitors and on the structure of human NAT1 will enable the synthesis of highly potent molecules. These compounds are expected to improve our understanding of the contribution of NAT1 to breast cancer mechanisms and putatively will be of therapeutic interest. Recent data from R. Minchin’s laboratory further support the

2

Human Arylamine N-acetyltransferase 1: From Drug Metabolism to Drug Target

33

contribution of NAT1 to cancer mechanisms and emphasize its potential role as a drug target. Using a specificic inhibitors or RNAi-mediated knock-down approaches, these authors show that inhibition of NAT1 in colon adenocarcinoma or breast cancer cells leads to growth inhibition [66, 67].

References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20.

21. 22. 23. 24. 25. 26. 27. 28. 29.

Hein, D. 2002, Mutat Res, 65, 65 Sim, E.., Walters, K. and Boukouvala, S. 2008, Drug Metab Rev, 40, 479 Rodrigues-Lima, F. and Dupret, J. M. 2002, Biochem Biophys Res Commun, 293, 783 Martins, M.., Rodrigues-Lima, F.., Dairou, J.., Lamouri, A.., Malagnac, F.., Silar, P. and Dupret, J. M. 2009, J Biol Chem, 284, 18726 Matas, N.., Thygesen, P.., Stacey, M.., Risch, A. and Sim, E. 1997, Cytogenet Cell Genet, 77, 290 Butcher, N.., Boukouvala, S.., Sim, E. and Minchin, R. F. 2002, Pharmacogenomics J, 2, 30 Grant, D. M.., Blum, M.., Beer, M. and Meyer, U. A. 1991, Mol Pharmacol, 39, 184 Sim, E.., Westwood, I. and Fullam, E. 2007, Expert Opin Drug Metab Toxicol, 3, 169 Butcher, N. J.., Arulpragasam, A.., Goh, H. L.., Davey, T. and Minchin, R. F. 2005, Biochem J, 387, 119 Husain, A.., Zhang, X.., Doll, M. A.., States, J. C.., Barker, D. F. and Hein, D. W. 2007, Drug Metab Dispos, 35, 721 Hein, D. W.., Doll, M. A.., Fretland, A. J.., Leff, M. A.., Webb, S. J.., Xiao, G. H.., Devanaboyina, U. S.., Nangju, N. A. and Feng, Y. 2000, Cancer Epidemiol Biomarkers Prev, 9, 29 Sim, E.., Lack, N.., Wang, C. J.., Long, H.., Westwood, I.., Fullam, E. and Kawamura, A. 2008, Toxicology, 254, 170 Atmane, N.., Dairou, J.., Paul, A.., Dupret, J. M. and Rodrigues-Lima, F. 2003, J Biol Chem, 278, 35086 Butcher, N. J.., Arulpragasam, A. and Minchin, R. F. 2004, J Biol Chem, 279, 22131 Rodrigues-Lima, F.., Dairou, J. and Dupret, J. M. 2008, Curr Drug Metab, 9, 505 Evans, D. A. P. and White, T. A. 1964, J. Lab. Clin. Med., 63, 394 Meisel, P. 2002, Pharmacogenomics, 3, 349 Hein, D. W. 2000, Toxicol Lett, 112-113, 349 Minchin, R. F. 1995, Biochem J, 307, 1 Adam, P. J.., Berry, J.., Loader, J. A.., Tyson, K. L.., Craggs, G.., Smith, P.., De Belin, J.., Steers, G.., Pezzella, F.., Sachsenmeir, K. F.., Stamps, A. C.., Herath, A.., Sim, E.., O’Hare, M. J.., Harris, A. L. and Terrett, J. A. 2003, Mol Cancer Res, 1, 826 Bieche, I.., Girault, I.., Urbain, E.., Tozlu, S. and Lidereau, R. 2004, Breast Cancer Res, 6, R252 Wakefield, L.., Robinson, J.., Long, H.., Ibbitt, J. C.., Cooke, S.., Hurst, H. C. and Sim, E. 2008, Genes Chromosomes Cancer, 47, 118 Russell, A. J.., Westwood, I. M.., Crawford, M. H.., Robinson, J.., Kawamura, A.., Redfield, C.., Laurieri, N.., Lowe, E. D.., Davies, S. G. and Sim, E. 2009, Bioorg Med Chem, 17, 905 Husain, A.., Barker, D. F.., States, J. C.., Doll, M. A. and Hein, D. W. 2004, Pharmacogenetics, 14, 397 Butcher, N. J.., Tiang, J. and Minchin, R. F. 2008, Curr Drug Metab, 9, 498 Butcher, N. J.., Arulpragasam, A.., Pope, C. and Minchin, R. F. 2003, Biochem J, 376, 441 Barker, D. F.., Husain, A.., Neale, J. R.., Martini, B. D.., Zhang, X.., Doll, M. A.., States, J. C. and Hein, D. W. 2006, Pharmacogenet Genomics, 16, 515 Wakefield, L.., Cornish, V.., Broackes-Carter, F. and Sim, E. 2005, J Histochem Cytochem, 53, 583 Butcher, N. J.., Tetlow, N. L.., Cheung, C.., Broadhurst, G. M. and Minchin, R. F. 2007, Cancer Res, 67, 85

34

F. Rodrigues-Lima et al.

30. Vatsis, K. P. and Weber, W. W. 1993, Arch Biochem Biophys, 301, 71 31. Boukouvala, S. and Fakis, G. 2005, Drug Metab Rev, 37, 511 32. Minchin, R. F.., Hanna, P. E.., Dupret, J. M.., Wagner, C. R.., Rodrigues-Lima, F. and Butcher, N. J. 2007, Int J Biochem Cell Biol, 39, 1999 33. Blum, M.., Grant, D. M.., McBride, W.., Heim, M. and Meyer, U. A. 1990, DNA Cell Biol, 9, 193 34. Fretland, A. J.., Doll, M. A.., Leff, M. A. and Hein, D. W. 2001, Pharmacogenetics, 11, 511 35. Upton, A.., Johnson, N.., Sandy, J. and Sim, E. 2001, Trends Pharmacol Sci, 22, 140 36. Dhaini, H. R. and Levy, G. N. 2000, Pharmacogenetics, 10, 79 37. Liu, F.., Zhang, N.., Zhou, X.., Hanna, P. E.., Wagner, C. R.., Koepp, D. M. and Walters, K. J. 2006, J Mol Biol, 361, 482 38. Riddle, B. and Jencks, W. P. 1971, J Biol Chem, 246, 3250 39. Dupret, J. M. and Grant, D. M. 1992, J Biol Chem, 267, 7381 40. Sinclair, J. C.., Sandy, J.., Delgoda, R.., Sim, E. and Noble, M. E. 2000, Nat Struct Biol, 7, 560 41. Martins, M.., Pluvinage, B.., de la Sierra-Gallay, I. L.., Barbault, F.., Dairou, J.., Dupret, J. M. and Rodrigues-Lima, F. 2008, J Mol Biol, 383, 549 42. Wu, H.., Dombrovsky, L.., Tempel, W.., Martin, F.., Loppnau, P.., Goodfellow, G. H.., Grant, D. M. and Plotnikov, A. N. 2007, J Biol Chem, 282, 30189 43. Rodrigues-Lima, F.., Deloménie, C.., Goodfellow, G. H.., Grant, D. M. and Dupret, J. M. 2001, Biochem J, 356, 327 44. Mushtaq, A.., Payton, M. and Sim, E. 2002, J Biol Chem, 17, 12175 45. Fullam, E.., Westwood, I. M.., Anderton, M. C.., Lowe, E. D.., Sim, E. and Noble, M. E. 2008, J Mol Biol, 375, 178 46. Liu, L.., Von Vett, A.., Zhang, N.., Walters, K. J.., Wagner, C. R. and Hanna, P. E. 2007, Chem Res Toxicol, 20, 1300 47. Kawamura, A.., Westwood, I.., Wakefield, L.., Long, H.., Zhang, N.., Walters, K.., Redfield, C. and Sim, E. 2008, Biochem Pharmacol, 75, 1550 48. Cornish, V.., Pinter, K.., Boukouvala, S.., Johnson, N.., Labrousse, C.., Payton, M.., Priddle, H.., Smith, A. J. H. and Sim, E. 2003, Pharmacogenomics J, 3, 169 49. Sugamori, K. S.., Wong, S.., Gaedigk, A.., Yu, V.., Abramovici, H.., Rozmahel, R. and Grant, D. M. 2003, Mol Pharmacol, 64, 170 50. Wakefield, L.., Long, H.., Lack, N. and Sim, E. 2007, Mamm Genome, 18, 270 51. Wakefield, L.., Cornish, V.., Long, H.., Griffiths, W. J. and Sim, E. 2007, Biochem Biophys Res Commun, 364, 556 52. Wakefield, L.., Cornish, V.., Long, H.., Griffiths, W. J. and Sim, E. 2007, Biochem Biophys Res Commun, 364, 556 53. Cao, W.., Chau, B.., Hunter, R.., Strnatka, D.., McQueen, C. A. and Erickson, R. P. 2005, Pharmacogenomics J, 5, 255 54. Hein, D. W. 2006, Oncogene, 25, 1649 55. Thygesen, P.., Risch, A.., Stacey, M.., Fakis, G.., Takle, L.., Knowles, M. and Sim, E. 1999, Pharmacogenetics, 9, 1 56. Kim, S. J.., Kang, H. S.., Chang, H. L.., Jung, Y. C.., Sim, H. B.., Lee, K. S.., Ro, J. and Lee, E. S. 2008, Oncol Rep, 19, 663 57. Lee, J. H.., Lu, H. F.., Wang, D. Y.., Chen, D. R.., Su, C. C.., Chen, Y. S.., Yang, J. H. and Chung, J. G. 2004, Res Commun Mol Pathol Pharmacol, 115-116, 217 58. Brooke, G. N.., Parker, M. G. and Bevan, C. L. 2008, Oncogene, 27, 2941 59. Lapointe, J.., Li, C.., Higgins, J. P.., van de Rijn, M.., Bair, E.., Montgomery, K.., Ferrari, M.., Egevad, L.., Rayford, W.., Bergerheim, U.., Ekman, P.., DeMarzo, A. M.., Tibshirani, R.., Botstein, D.., Brown, P. O.., Brooks, J. D. and Pollack, J. R. 2004, Proc Natl Acad Sci U S A, 101, 811 60. Ott, I. and Gust, R. 2007, Anticancer Agents Med Chem, 7, 95 61. Ragunathan, N.., Dairou, J.., Pluvinage, B.., Martins, M.., Petit, E.., Janel, N.., Dupret, J. M. and Rodrigues-Lima, F. 2008, Mol Pharmacol, 73, 1761 62. Malka, F.., Dairou, J.., Ragunathan, N.., Dupret, J. M. and Rodrigues-Lima, F. 2009, Febs J, 276, 4900 63. Sauna, Z. E.., Shukla, S. and Ambudkar, S. V. 2005, Mol Biosyst, 1, 127

2

Human Arylamine N-acetyltransferase 1: From Drug Metabolism to Drug Target

35

64. Brooke, E. W.., Davies, S. G.., Mulvaney, A. W.., Pompeo, F.., Sim, E. and Vickers, R. J. 2003, Bioorganic & Medicinal Chemistry, 11, 1227 65. Madikane, V. E.., Bhakta, S.., Russell, A. J.., Campbell, W. E.., Claridge, T. D.., Elisha, B. G.., Davies, S. G.., Smith, P. and Sim, E. 2007, Bioorg Med Chem, 15, 3579 66. Tiang, J. M.., Butcher, N. J.., Cullinane, C.., Humbert P. O. and Minchin, R. F. 2011, PLoS One 6:e17031 67. Tiang, J. M.., Butcher, N. J. and Minchin, R. F. 2010, Biochem Biophys Res Commun, 39, 95

Chapter 3

Targeting Argininosuccinate Synthetase in Cancer Therapy Niramol Savaraj, Min You, Chunjing Wu, Macus Tien Kuo, Vy Dinh, Medhi Wangpaichitr, and Lynn Feun

Introduction There has been renewed interest in the use of selective amino acid deprivation as a method to treat various cancers. It has been known for years that certain cancers are auxotrophic for particular nonessential amino acids. By exploiting the differences between normal human cells and cancer cells, one can target these cancers and potentially reduce the potential side effects of the drugs. The best example so far has been the use of asparaginase in the treatment of acute lymphoblastic leukemia (ALL). ALL cells have a requirement for L-asparagine, a nonessential amino acid in humans. By depleting this amino acid, ALL cells can be targeted and drug toxicity reduced. In this chapter, we will focus on arginine depletion as another amino acid deprivation target for cancers.

Background Reversible inhibition of mitosis in lymphocytes cultures by Mycoplasma was first noted by Copperman and Morton in 1966 [1]. The mechanism behind Mycoplasma inhibition of lymphocyte transformation induced by phytohaemagglutinin was described by Barile and Leventhal two years later [2]. They discovered that the depletion of arginine was the underlying mechanism. Gill and Pan in 1970 found that addition of arginine could reverse the inhibition of cell division in L5178Y cells by arginine-degrading Mycoplasma and the possible role of arginine

L. Feun (*) Sylvester Cancer Center, University of Miami school of Medicine, Miami, FL 33125, USA e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_3, © Springer Science+Business Media, LLC 2012

37

38

N. Savaraj et al.

deiminase [3]. Subsequently, Fenske and Kenny reported on the role of arginine deiminase in the growth of Mycoplasma [4], and Weickmann and Fahrney provided evidence of multiple forms of arginine deiminase from Mycoplasma [5]. It was not until 1990 that Sugimura identified arginine deiminase as the lymphocyte blastogenesis factor originating from Mycoplasma arginini [6–8]. In the same year, Miyazaki et al. reported on the potent growth inhibition of human tumor cells in culture by arginine deiminase purified from culture medium in cell lines infected with Mycoplasma [9]. Later, Takaku described the in vivo antitumor activity of arginine deiminase [10, 11]. Taken together, these reports suggest that arginine deprivation, such as after treatment with arginine deiminase isolated from Mycoplasma, represents a novel approach to the therapy of various malignancies which require exogenous arginine.

Why Arginine Deprivation Is Important The amino acid arginine is involved in several important cellular functions. These include polyamine synthesis, creatine production, and nitric oxide production [12–14]. Arginine is the only endogenous source of nitric acid in humans [12]. Humans derive arginine from several sources. These include the diet, body protein turnover, and de novo synthesis. In adult humans, arginine is considered a nonessential amino acid since it can be synthesized from citrulline. However, endogenous production of arginine may be insufficient under certain circumstances, such as when cells are under stress or need to proliferate during wound healing. As mentioned, certain tumor cells require exogenous arginine in order to grow and proliferate, and diet restriction of arginine may actually inhibit metastatic growth [15–17]. Arginine is synthesized from citrulline via the urea cycle (Fig. 3.1). There are two key enzymes involved: (1) argininosuccinate synthetase (ASS) which converts L-citrulline and aspartic acid to argininosuccinate and (2) argininosuccinate lyase (ASL) which then converts argininosuccinate to L-arginine and fumaric acid. L-arginine can also be degraded by the urea cycle enzyme arginase to L-ornithine. L-ornithine is converted back to L-citrulline by ornithine carbamoyl transferase (OCT) and then is recycled back to arginine. Wheatley has suggested that ASS and ASL are tightly coupled [18–20]. The sensitivity of tumor cells to arginine deprivation may depend on their ability to synthesize arginine from alternative intermediates in the urea cycle such as ornithine, citrulline, and argininosuccinate. There are three enzymes which can catabolize arginine: arginase, arginine decarboxlyase, and arginine deiminase (ADI). Since 1950, arginase has been tried in experimental animals as an antitumor agent. No major responses were observed. There are several potential problems with arginase as a targeted therapy for cancer. First of all, arginase has a low affinity for arginine, and larger amounts of the enzyme may be required to produce a response. Secondly, for arginase, the optimal pH is 9.5, which is high. On the other hand, Wheatley has shown that arginase may be effective at lower pH of 7.2–9 [15]. It is important to note that arginase catabolizes

3

Argininosuccinate Synthetase in Cancer Therapy

39

ARGININOSUCCINATE Argininosuccinate Lyase Fumarate ARGININE

Argininosuccinate (ASS) Synthetase

ADI

Putrescine

CITRULLINE + Ammonium Ornithine (OCT) Carbamoyl transferase

Arginase Urea

Aspartate

ORNITHINE

Carbamoyl phosphate

Detoxification

Fig. 3.1 Diagram for arginine metabolism in the urea cyle and the action of ADI and arginase

arginine to ornithine. The liver and the small bowel are known to be able to convert ornithine to citrulline due to the presence of OCT. It is not clear whether other normal tissues are able to do this since OCT gene is hypermethylated and not expressed in these tissues. Thus, there is the potential for normal tissue toxicity from the use of arginase. Despite these potential problems, pegylated arginase has been shown to have both in vitro and in vivo activity in hepatocellular carcinoma [21, 22]. Another novel approach involves transhepatic chemoembolization of the liver augmented with high-dose insulin to induce leakage of hepatic arginase from the liver into the circulation [23]. The high-dose insulin was administered to induce a state of hypoaminoacidemia to augment arginine depletion. Five of seven patients with hepatocellular carcinoma had response to this treatment. The two patients with no response had no significant reduction in plasma arginine levels. Since arginase has a short half-life, pegylation of arginase has been developed in attempt to produce sustained in vivo activity [24]. Further studies with pegylated arginase appear to be indicated. One of the important factors which make arginine deprivation an attractive approach to treat cancer is that certain tumors lack ASS expression. These tumors include melanoma, hepatocellular carcinoma, certain mesotheliomas, renal cell cancer and prostate cancers [25–28]. These ASS(−) tumors will require exogenous arginine for their growth. Thus, by depleting arginine, one can inhibit cell growth [16, 29–31]. Transfection of ASS also confers resistance to arginine deprivation treatment [30, 31]. The question remains which is the best enzyme to deplete arginine. ADI has several advantages for a targeted therapy. It degrades arginine to citrulline and ammonia. Tumor cells which lack ASS expression will not be able to synthesize arginine from citrulline, while normal cells are able to do so. Theoretically, normal tissue toxicity can be avoided by this approach. Secondly, ADI is active at physiologic pH. Thirdly, unlike arginase, ADI has high affinity for arginine [32]. The major drawback for ADI is that it is not a normal human enzyme, but made by Mycoplasma. Therefore, ADI is highly immunogenic and has a short half-life. One method to prolong the half-life of the enzyme and reduce immunogenecity

40

N. Savaraj et al.

is by pegylation. A number of pegylated forms of ADI have been tested. One pegylated form of ADI has been developed by Polaris, Inc (formerly Phoenix Pharmacologics) termed ADI-PEG20. ADI is a recombinant protein of approximately 46,000 daltons which has a covalent attachment to polyethylene glycol (PEG) of 20,000 daltons. Each molecule of ADI is attached to approximately 13–16 molecules of PEG. ADI-PEG20 incorporates the succinimidyl–succinate linker. ADI-PEG20 has undergone extensive in vitro and in vivo testing [30]. Subsequently, ADI-PEG20 has entered clinical trial in patients with malignant melanoma and hepatocellular carcinoma, and trials are planned for mesothelioma and possibly other tumor types [33–37].

Preclinical Antitumor Activity of ADI-PEG20 Takaku demonstrated that the addition of ADI to the culture media resulted in killing a number of tumors including melanoma and hepatocellular carcinoma [10]. Sugimura also showed that the growth inhibition of melanoma cells induced by arginine deprivation could not be reversed by the addition of the amino acid citrulline [38]. He hypothesized that melanoma cells lack argininosuccinate synthetase and hence cannot synthesize arginine from citrulline. Studies by Ensor et al. demonstrated that the human melanoma cell lines SK-mel2 and SK-mel28 were inhibited in a dose-dependent manner by adding ADI to the culture media [30]. These cells were found to express argininosuccinate lyase mRNA but not that of argininosuccinate synthetase by northern blot analysis. To confirm this hypothesis, SK-mel2 and SK-mel28 cells were transfected with a plasmid containing the human argininosuccinate synthetase cDNA using a cytomegalovirus promoter. The transfected cells were resistant to ADI. In the in vivo study, Takaku has shown that mice implanted with human melanomas and hepatocellular carcinomas and treated with ADI resulted in inhibition of tumor growth and extended life expectancy [10, 11]. These results were further confirmed with several other human melanomas including SK-mel2 and SK-mel28 [30]. Furthermore, when dogs with spontaneous melanoma of the oral cavity were treated with ADI-PEG20, clinical responses were observed. These results confirm that ADI-PEG20 has antitumor activity in melanoma. Similarly, hepatocellular carcinomas, SK-hep1, SK-hep2, and SΚ-hep3, implanted in mice were also inhibited when treated with ADI-PEG20.

ASS Expression and Arginine Deprivation in Melanoma As stated earlier, multiple studies have demonstrated that arginine deprivation using ADI showed antitumor activity both in vitro and in vivo. Furthermore, Ensor has tested 16 melanoma cell lines and found no ASS expression in all these melanoma cell lines. In human tumor samples, the majority of melanomas and hepatocellular carcinomas do not express ASS, while lung cancer and breast cancer cells

3

Argininosuccinate Synthetase in Cancer Therapy

41

were usually positive for ASS. In our laboratory, we evaluated the effect of degrading arginine in the culture media using ADI or exposing melanoma cells to arginine-free media. Our results demonstrated that ADI treatment results in growth inhibition which eventually leads to cell death [31]. Four ASS(−) melanoma cell lines (A375, SK-mel2, A2058, and Mel-1220), one normal human fibroblast cell line (BJ-1), and one ASS-positive NSCLC cell line were studied. The ID50 (growth inhibitory effect) for the four melanoma cell lines after 3-day treatment ranged from 0.05 to 0.1 ug/ml, and it was greater than 1 ug/ml in BJ-1 and NSCLC cell lines. It is of interest to note that at 72 h after treatment, the ASS protein can be detected by western blot analysis with A2058 expressing the highest level (2.76-fold increase), while in Mel-1220, it was not detectable, and A375 has negligible increased ASS protein. The cell lines (BJ-1 and NSCLC) which constitutively express ASS did not show changes in ASS protein after exposure to arginine-free media [31]. The changes in ASS expression upon arginine deprivation is also detected at the transcriptional levels using real-time RT-PCR. The ASS mRNA showed about 3-fold increase in A2058 followed by about twoto threefold increase in SK-mel2, while negligible changes occurred in A375 and Mel-1220. However, upon arginine replenishment, ASS mRNA and protein is back to normal [31]. Thus, it appears that arginine in the media controls ASS expression in certain melanoma cell lines. In our ongoing phase II trial in melanoma, we have studied ASS expression in tumor samples obtained prior to treatment with ADI-PEG20 and after progression or relapse. Two patients had tumor samples tested for ASS expression prior to therapy and were ASS(−). Both patients had a partial response to ADI-PEG20 but later had progression of disease. At the time of tumor progression, tumor samples were obtained again and were ASS positive. One patient’s tumor sample prior to treatment with ADI-PEG20 was found to be ASS positive, and this patient did not respond to therapy. Of 11 patients whose tumor samples were ASS(−), seven had evidence of antitumor activity (partial response, mixed response, minor response). While the data is still preliminary, it suggests that there may be a correlation between ASS tumor expression and clinical antitumor activity. This trial is continuing to accrue more patients.

Arginine Deprivation in Hepatocellular Carcinoma Dillon reported that all 51 hepatoma cell lines studied were ASS negative by immunohistochemical staining [25]. Ensor found that some hepatocellular cell lines which lack ASS expression were sensitive to arginine deprivation with ADI-PEG20 [30]. On the other hand, Cheng found that five hepatoma cell lines had ASS expression and were not sensitive to ADI [21, 22]. These cell lines lack OCT and were also sensitive to arginase. PEG-arginase also has been reported to have activity in vivo against hepatoma. Therefore, it is not clear how frequent ASS expression occurs in hepatoma and whether ADI or arginase is the better agent of choice for clinical trial.

42

N. Savaraj et al.

Arginine Deprivation in Other Tumor Cell Types Dillon has reported that some renal cell cancers and sarcomas as well as some prostate cancers lack ASS expression [25]. Other investigators also found lack of ASS expression in certain mesothelioma and neuroblastomas [26, 39]. Renal cell cancers which were negative for ASS expression by immunohistochemical staining were also found to be sensitive to ADI-PEG20 [25, 27]. This is of interest that the normal renal epithelium has high levels of ASS expression. Similarly, mesothelioma cell lines which lack ASS expression are also sensitive to arginine deprivation with ADI [26]. In contrast, mesothelioma cell lines which express ASS were resistant to ADI. In one study using immunohistochemical staining, 63% of mesothelioma cell lines express low levels of ASS. Thus, arginine deprivation may be a useful agent to treat certain mesotheliomas. A clinical trial using ADI-PEG20 is being planned for mesothelioma (Szlosarek, personal communication). Certain other tumor cell lines which may be sensitive to arginine deprivation include human T and B cell lymphoblastic cell lines [40, 41]. Interestingly, arginine deprivation with ADI-PEG20 showed better antitumor effect than L-asparaginase which is approved for lymphoblastic leukemia. Neuroblastoma cell lines have also been shown to be sensitive to arginine deprivation with ADI.

How Arginine Deprivation Affects Growth and Apoptotic Signaling Arginine Deprivation Affects mTOR Signaling Although arginine is a nonessential amino acid in normal cells which possess ASS, however, in ASS(–) tumor cells, it becomes an essential amino acid. Thus, arginine deprivation inhibits mTOR similar to those reported with nutritional deprivation such as amino acid deprivation. We have found that upon arginine deprivation, ASS(–) cells activate AMPK, which is known to regulate mTOR activity via energy/ nutrient sensing [42]. When cells are deprived of ATP (high AMP/ATP) ratio, AMPK is activated via phosphorylation by LKB or other upstream kinase(s) [43, 44]. Activated AMPK downregulates energetically demanding processes like translation by inhibiting mTOR [45]. In this regard, we have studied the effects of ADIPEG20 treatment on ATP levels and AMPK activation. Our data showed that the ATP decreased by 30%, 55%, 50%, and 60% in Mel-1220, A2058, SK-mel2, and A375, respectively, at 72 h corresponding to the activation of AMPK. Activation of AMPK also results in mTOR inhibition as evidenced by decreased 4E-BP phosphorylation, while p70S6K phosphorylation only decreased in two cell lines, which are ASS negative and cannot be induced. We have also studied three additional cell lines derived from patients whose tumors were ASS negative and cannot be induced. These cell lines also showed activation of AMPK upon arginine deprivation and

3

Argininosuccinate Synthetase in Cancer Therapy

43

inhibition of 4E-BP [42]. Thus, our data demonstrated that upon arginine deprivation in ASS(−) melanoma cells, activation of AMPK occurs which in turn has a negative impact on mTOR activity [42] . In this regard, Kim et al. also have shown that arginine deprivation results in decreased mTOR phosphorylation in prostate cancer cell lines which do not express ASS [28]. Thus, one can conclude that arginine deprivation in ASS(−) tumor cells results in mTOR inhibition which can ultimately lead to autophagy [46].

Arginine Deprivation on RAF/MEK/ERK1/2 Signaling It is known that approximately 60% of melanoma tumors possess BRAF mutation (V600E). Activation of BRAF has been shown to be one of the major growth signaling for melanoma. In fact, BRAF inhibitor has been developed to treat melanoma, with variable success depending on the specificity of the compound [47, 48]. We have found that upon treatment with ADI-PEG20, activated MEK and ERK, as detected by the phosphorylated form, increased, while no significant changes occurred in BRAF. Interestingly, the addition of MEK inhibitor (U0176 or PD98059) increased apoptosis when used in combination with ADI-PEG20 treatment. Thus, it appears activation of MEK and ERK may be an attempt for the cells to survive upon arginine deprivation through autophagy. To support this, recently it has been shown that activation of MEK and ERK are important in noncanonical autophagic process [49]. Inhibition of this process can result in apoptosis [49]. In addition, Wang et al. have shown that upon amino acid deprivation, activation of MEK and ERK occurs which results in protective autophagy, while sustained activation of MEK/ERK results in destructive autophagy [49]. They have also shown that activation of AMPK during nutritional deprivation activates MEK1 and leads to disassembling of mTORC1 complex via binding to and activation of TSC2. These findings confirm our results that arginine deprivation results in activation of MEK/ERK but has minor or no discernable effect on RAF. Activation of MEK/ERK leads to mTOR inhibition which results in autophagy.

Arginine Deprivation Leads to Autophagy in ASS(−) Cells Autophagy is a lysosomal degradation pathway which involves vesicular sequestration of proteins or organelles into autophagosomes which then fuse with lysosomes and become degraded [50, 51]. Low levels of autophagy also occur as a normal physiologic process to remove damaged organelles [52]. This process can be rapidly activated as an adaptive catabolic process in response to different forms of metabolic stress such as nutritional deprivation, hypoxia, or growth factor withdrawal [53–55]. The bulk forms of degradation generated are free amino acids and fatty acids which can be recycled to make essential proteins for adaptive survival. The signaling mechanisms of autophagy are complex and involve multiple pathways,

44

N. Savaraj et al.

depending upon the cellular contexts and type of inducers [56 –61]. Two evolutionally conserved nutrient sensors play a role in autophagy regulation: (1) mTOR kinase which turns off the energy consuming translational process during autophagy and (2) the eukaryotic kinase initiation factor 2a kinase (GCN2) which represses translation but upregulates transcriptional activator ATF4 which is required to transcribe stress related genes [62, 63]. Our initial data do not show alteration of GCN2 upon arginine deprivation. Downstream the TOR kinase, there are 17 genes which are involved in the autophagic process in yeast, termed ATG genes. The ATG gene is involved in the generation and maturation of autophagosome [64]. In cancer cells, the mechanism of how many ATG genes are involved in the autophagosome formation is not well understood. However, Vps34 (PI3 kinase class III) and Beclin 1(ATG6) have been shown to be involved in the vesicle nucleation process [51, 65]. On the other hand, autophagy can also lead to cell death (program type II or noncaspase-dependent cell death) which is characterized by degradation of organelles but preserved cytoskeletal process [63, 66]. Autophagic cell death is known to occur during the developmental process [67, 68] and recently with cancer chemotherapy and hormonal therapy [69, 70]. We have found that, upon arginine deprivation, ASS(−) melanoma cells undergo autophagy, most likely as part of the adaptive mechanisms to avoid cell death. These findings also have been shown by Kim et al. in prostate cancer cell lines which do not express ASS [28]. In fact, our preliminary data also suggest that knockdown of Beclin 1 increased cell death after ADI-PEG20 treatment by 20–30%. In this regard, Kim et al. also have shown that knockdown of Beclin 1 in ASS(−) prostate cancer cells increases cell death [28]. Thus, ADI-PEG20 or arginine deprivation treatment induces autophagy in ASS(−) melanoma cell lines, most likely representing a mechanism for survival. Inhibition of this process should be considered to increase the therapeutic effect of this treatment. One can conclude that arginine deprivation results in activation of AMPK which inhibits mTOR via excessive activation of MEK and ERK and possibly other pathways. This results in induction of autophagy in an attempt for the cell to survive (Fig. 3.2). However, prolonged arginine deprivation will ultimately result in caspase-dependent or independent apoptosis, depending on the cell type and the apoptotic machinery in the cells [28, 31, 41, 71].

How Arginine Deprivation Induces Cell Death in ASS(−) Melanoma Cells The ASS gene is located in chromosome 9q34.1. There are many pseudogenes (14 reported) located among other chromosomes [72]. The transcription and translation of ASS is not well understood. Two species of ASS mRNA which differ in their 5¢-UTR have been reported (Genbank Database). Previous published studies indicate that ASS regulation occurs at the pretranslational levels [12]. Multiple factors have been shown to positively and negatively influence ASS transcription, but are most likely tissue specific. Several investigators have demonstrated that glucocorticoids, glucagons, and cyclic

3

Argininosuccinate Synthetase in Cancer Therapy

Fig. 3.2 Possible pathways which can lead to autophagy upon arginine deprivation

45

Arginine deprivation

pAMPK

pMEK

TSC1/2

? pERK

mTOR

x AUTOPHAGY

AMP increase ASS expression, whereas fatty acids can repress ASS expression [73–81]. Both growth hormone and insulin have negative influence on ASS expression in liver tissue but do not affect ASS expression in other body tissues. Various cytokines, IL-Ib, interferon, and TGF-b, can affect ASS expression [82, 83]. Glutamine can stimulate ASS expression in rat hepatocytes and Caco-2 cells through O-glycosylation of SP-1 sites [84]. ASS expression can also be regulated by arginine [85]. We also have found that arginine in the media can regulate arginine expression in certain melanoma cells which possess very low levels of ASS not detected by western blot analysis but could be detected by real-time RT-PCR [31]. These cell lines can be induced to express ASS mRNA in the arginine free media. Our data in four different melanoma cells do not suggest any differences in the promoter region of these cell lines. Recently, we have found that ASS in melanoma cells is positively regulated by c-Myc but negatively by HIF1alpha. On the other hand, in other tumor types such as mesothelioma and ovarian cancer, aberrant methylation have been reported which results in epigenetic gene silencing [26]. Thus, it appears that ASS gene regulation in cancer cells is also tumor cell-type specific. From our data and the data available from other laboratories, ASS gene silencing occurs in certain tumor types such as melanoma, mesothelioma, and hepatoma. However, whether it is epigenetic or transcriptional factor-regulated or both are celltype specific requires further investigation.

ASS Expression and Cisplatin Resistance Downregulation of ASS has been found in cisplatin-resistant ovarian cancer cell lines [86, 87]. The exact mechanism(s) is not known. Recently, ASS methylation/ gene silencing at diagnosis was found to be associated with significantly reduced overall survival and relapse-free survival in ovarian cancer. Furthermore, in ovarian

46

N. Savaraj et al.

cancer cell lines, methylation of ASS promoter correlated with transcriptional silencing and selective resistance to platinum-based drug. The underlying mechanisms are not well understood. The author suggests that alteration of nitric oxide which affects apoptosis may be involved [86].

Other Effects of Arginine Deprivation Arginine deprivation can also have indirect effects on tumor growth and proliferation. For example, arginine deprivation may have anti-angiogenesis effects. Beloussow showed that arginine deprivation using recombinant ADI had anti-angiogenesis action in cultured human umbilical vein endothelial cells [88]. Park et al. used the same cell line to demonstrate that arginine deprivation with recombinant ADI inhibited angiogenesis in a dose-dependent manner [89]. The inhibition of angiogenesis was reversed by adding arginine back to the media. In addition, they showed that arginine deprivation with ADI produced inhibition of angiogenesis with chick embryo and mice. Thus, their in vivo studies confirmed the anti-angiogenesis results seen with their in vitro study demonstrating that ADI could inhibit tumor cell growth with CHO and HeLa cells. Therefore, ADI may affect tumor growth by its antiangiogenetic properties. Other investigators also have shown that ADI was effective in the inhibition of unfavorable neuroblastoma [39]. Thus, it appears that ADI could also exert its antitumor activity via inhibition of angiogenesis. ADI also has been used in combination with radiation therapy. The antitumor effect produced by radiation therapy was potentiated, suggesting a possible role for arginine deprivation with radiation therapy for certain tumors. Another effect of arginine deprivation may be on the immune system. It is well known that arginine is the only endogenous source for nitric oxide production. Arginine is the only substrate for nitric oxide synthetase [90], so arginine deprivation can inhibit nitric oxide production in macrophages challenged with interferon. Shen et al. showed that recombinant ADI could be a differential modulator of inducible and endothelial nitric oxide synthetase activity in cultured endothelial cells [91]. Furthermore, this effect was observed after mice were treated with TNF and endotoxin. They found that ADI-PEG20 did not inhibit nitric oxide production by eNOS, so ADI-PEG20 may be useful as a selective modulator for nitric oxide produced by iNOS. In addition, nitric oxide may affect ASS as Hao et al. found that ASS is reversibly inactivated by S-nitrosylation both in vitro and in vivo [92]. Further research into the complex interrelationships between these different parameters needs to be done to elucidate the role of arginine deprivation, nitric oxide production, and tumor cell survival. It is of interest that nitric oxide may promote tumor growth and metastasis by its stimulation of tumor cell migration, growth, and invasiveness. In this regard, Jadeski showed that nitric oxide promotes murine mammary cell tumor [93]. Some metastatic melanoma cells may also escape immunosurveillance through the novel mechanism of releasing nitric oxide which induces immunocyte dysfunction [94]. Other inhibitors of nitric oxide such

3

Argininosuccinate Synthetase in Cancer Therapy

47

as NG-methyl-L-arginine and NG-monomethyl-L-arginine have been shown to reverse or reduce the hypotensive effects in interleukin-2 therapy, enabling more drug to be delivered with less toxicity [95, 96]. Thus, arginine deprivation by ADI or arginase may be useful to reduce nitric oxide production and to enhance immunogenecity of tumors and lessen the side effect during interleukin-2 therapy [32].

Clinical Trials with ADI-PEG20 We have reported on a phase I/II trial of ADI-PEG20 in patients with advanced or metastatic melanoma [34]. This trial consisted of a phase I trial in the U.S. and a phase I/II trial in Italy. The phase I trial in the U.S. had 15 patients enrolled in four cohorts using different weekly doses of the drug. In Italy, there were 24 patients enrolled in the phase I trial, also using varying dose levels of ADI-PEG20. The dose of 160 IU/m² given weekly intramuscularly was felt to be the optimum biological dose (OBD). The most common side effect in both the U.S. and Italian part of the study was discomfort at the injection site. Other side effects were less common and included fatigue and mild elevation of the serum uric acid. In terms of response, none of the 15 patients enrolled in the phase I trial in the U.S. had a response. In Italy, 6 of 24 patients enrolled in the phase I/II had a response to treatment. This included one complete response and five partial responses, for a total response rate of 25%. In addition, 14 patients had stable disease for ³1 cycle of therapy, and six patients had stable disease for 3 months of the trial. All the patients with response or stable disease had received the OBD or higher dose of ADI-PEG20. We are currently conducting a phase II trial of ADI-PEG20 in patients with advanced or metastatic melanoma, and preliminary results have been published [34]. We have observed responses in at least four patients and stable disease in other patients. Toxicity has generally been mild, with discomfort at the injection site being the most common side effect. This trial is still ongoing. A phase I/II trial of ADI-PEG20 in melanoma is being conducted at New York University and Memorial Sloan-Kettering Cancer Center. This trial allowed heavily pretreated patients to be entered. While no major responses were observed, stable disease was seen in several patients. Arginine depletion with ADI-PEG20 has also been investigated in patients with hepatocellular carcinoma. A patient with unresectable hepatocellular carcinoma was treated with escalating doses of ADI-PEG20 in a single patient exemption trial. No drug toxicity was noted. The patient had a reduction in tumor size and a decrease in the serum alpha fetoprotein. The encouraging response in this single patient led to a series of phase I/II trials in hepatocellular carcinoma. In a study conducted at the Pascale Cancer Institute in Naples, Italy, patients were treated with various doses of ADI-PEG20. Nineteen patients were entered into the trial using one of four drug cohorts. The dose of 160 IU/m² given weekly intramuscularly was considered the OBD and lowered the serum arginine level to non-detectable levels (0.5 uM). In terms of toxicity, the side effects included discomfort at the injection site, elevation of the serum uric acid and fibrinogen, and occasional elevation of the serum lipase

48

N. Savaraj et al.

and amylase levels. No clinical evidence of pancreatitis was detected. No evidence of neutralizing antibody was found. In terms of response, two patients had complete response, and seven patients had partial response for a response rate of 47%. Duration of response was defined as the time from start of therapy to tumor progression. The median duration of response was >400 days (range 37– >680 days). Another phase I/II trial of weekly ADI-PEG20 was conducted at the M.D. Anderson Cancer Center in Houston, Texas. In this trial, 35 patients were enrolled. In terms of toxicity, 12 patients had grade 3 toxicity which consisted mainly of liver function or serum electrolyte abnormalities. In addition, three patients had grade 4 toxicity which consisted of liver function abnormalities or elevation of serum lipase. In terms of response, one patient had a partial response, and the patient’s disease became resectable after treatment. Furthermore, 16 patients had stable disease. Four patients did not complete the trial due to either allergic reaction or intercurrent disease. Twenty-eight patients had disease progression, with the mean time for progression of 3.4 months (range 1–13 months). We have treated nine patients with hepatocellular carcinoma with ADI-PEG20 as part of the phase 2B testing. Of the nine patients treated at the University of Miami, two patients had stable disease for at least 12 months. One of the two patients had stable disease for >2 years until he developed an allergic reaction, and the therapy was discontinued. Recently, a large phase II study in hepatoma treated with ADI-PEG20 showed a disease control rate of 31% and 7.3 months for overall survival. The patients with no detectable arginine appears to do better with an overall survival of 10 months [97].

Concluding Remarks and Future Direction The data from our laboratory and the data from other laboratories support the notion that arginine deprivation represents a novel targeted approach for antitumor therapy in humans. Tumors which cannot synthesize arginine from the urea cycle and thus are auxotrophic for arginine can be treated with drugs such as ADI-PEG20 or arginase. Furthermore, our in vitro laboratory data suggest that arginine deprivation combined with DNA-damaging agents such as temozolomide or cisplatin may be synergistic in certain tumor cell lines. Tumor cells which lack ASS expression may be unable to repair DNA damage without the presence of arginine. We are currently investigating the mechanism behind this. In addition, clinical trials combining ADIPEG20 to deplete arginine with DNA damaging agents should be explored for their synergistic activity. Melanoma and hepatocellular carcinoma are the most common tumor types that have been studied so far in clinical trials with arginine deprivation using ADI-PEG20. Arginine deprivation should also be investigated in certain mesotheliomas, renal cell cancers, and prostate cancers which lack ASS expression. Recently, a patient with pancreatic neuroendocrine tumor was treated with ADI-PEG20 and reportedly had a major response to therapy (Bomalaski, Polaris, Inc. unpublished data). Thus, ASS expression and arginine deprivation should be studied further in other tumor types.

3

Argininosuccinate Synthetase in Cancer Therapy

49

The development of drug resistance due to the induction of ASS gene expression represents a major problem. In our clinical trial with melanoma patients, biopsies of tumor after initial response to ADI showed that ASS expression was induced and correlated with drug resistance and relapse or tumor progression. Understanding the transcription/translational control of the ASS gene may aid in the development of ways to overcome the development of drug resistance. Pharmacological methods to control ASS expression and so prevent drug resistance to arginine depletion are currently being investigated in our laboratory.

References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28.

Copperman, R., and Morton, H. E. 1966, Proc Soc Exp Biol Med, 123, 790–795 Barile, M. F., and Leventhal, B. G. 1968, Nature, 219, 750–752 Gill, P., and Pan, J. 1970, Can J Microbiol, 16, 415–419 Fenske, J. D., and Kenny, G. E. 1976, J Bacteriol, 126, 501–510 Weickmann, J. L., and Fahrney, D. E. 1977, J Biol Chem, 252, 2615–2620 Sugimura, K., Fukuda, S., Wada, Y., Taniai, M., Suzuki, M., Kimura, T., Ohno, T., Yamamoto, K., and Azuma, I. 1990, Infect Immun, 58, 2510–2515 Sugimura, K., Kimura, T., Arakawa, H., Ohno, T., Wada, Y., Kimura, Y., Saheki, T., and Azuma, I. 1990, Leuk Res, 14, 931–934 Sugimura, K., Ohno, T., Fukuda, S., Wada, Y., Kimura, T., and Azuma, I. 1990, Cancer Res, 50, 345–349 Miyazaki, K., Takaku, H., Umeda, M., Fujita, T., Huang, W. D., Kimura, T., Yamashita, J., and Horio, T. 1990, Cancer Res, 50, 4522–4527 Takaku, H., Takase, M., Abe, S., Hayashi, H., and Miyazaki, K. 1992, Int J Cancer, 51, 244–249 Takaku, H., Matsumoto, M., Misawa, S., and Miyazaki, K. 1995, Jpn J Cancer Res, 86, 840–846 Husson, A., Brasse-Lagnel, C., Fairand, A., Renouf, S., and Lavoinne, A. 2003, Eur J Biochem, 270, 1887–1899 Lind, D. S. 2004, J Nutr, 134, 2837 S-2841 S; discussion 2853 S Morris, S. M., Jr. 2006, Am J Clin Nutr, 83, 508 S-512 S Wheatley, D. N., Philip, R., and Campbell, E. 2003, Mol Cell Biochem, 244, 177–185 Wheatley, D. N., and Campbell, E. 2003, Br J Cancer, 89, 573–576 Wheatley, D. N. 2004, Anticancer Drugs, 15, 825–833 Wheatley, D. N. 2005, Semin Cancer Biol, 15, 247–253 Wheatley, D. N., Campbell e, Lai P. B. S., Cheng, P. N. M. 2005, Gene Therapy and Molecular Biology, 9, 33–40 Wheatley, D. N., and Campbell, E. 2002, Pathol Oncol Res, 8, 18–25 Cheng, P., Lo, T., and Leung, T., et al. 2005, Proc Am Soc Clin Oncol, 23 Cheng, P. N., Lam, T. L., Lam, W. M., Tsui, S. M., Cheng, A. W., Lo, W. H., and Leung, Y. C. 2007, Cancer Res, 67, 309–317 Cheng, P. N., Leung, Y. C., Lo, W. H., Tsui, S. M., and Lam, K. C. 2005, Cancer Lett, 224, 67–80 Tsui, S. M., Lam, W. M., Lam, T. L., Chong, H. C., So, P. K., Kwok, S. Y., Arnold, S., Cheng, P. N., Wheatley, D. N., Lo, W. H., and Leung, Y. C. 2009, Cancer Cell Int, 9, 9 Dillon, B. J., Prieto, V. G., Curley, S. A., Ensor, C. M., Holtsberg, F. W., Bomalaski, J. S., and Clark, M. A. 2004, Cancer, 100, 826–833 Szlosarek, P. W., Klabatsa, A., Pallaska, A., Sheaff, M., Smith, P., Crook, T., Grimshaw, M. J., Steele, J. P., Rudd, R. M., Balkwill, F. R., and Fennell, D. A. 2006, Clin Cancer Res, 12, 7126–7131 Yoon, C. Y., Shim, Y. J., Kim, E. H., Lee, J. H., Won, N. H., Kim, J. H., Park, I. S., Yoon, D. K., and Min, B. H. 2007, Int J Cancer, 120, 897–905 Kim, R. H., Coates, J. M., Bowles, T. L., McNerney, G. P., Sutcliffe, J., Jung, J. U., GandourEdwards, R., Chuang, F. Y., Bold, R. J., and Kung, H. J. 2009, Cancer Res, 69, 700–708

50

N. Savaraj et al.

29. Shen, L. J., Beloussow, K., and Shen, W. C. 2006, Cancer Lett, 231, 30–35 30. Ensor, C. M., Holtsberg, F. W., Bomalaski, J. S., and Clark, M. A. 2002, Cancer Res, 62, 5443–5450 31. Savaraj, N., Wu, C., Kuo, M., You, M., Wangpaichitr, M., Robles, C., Spector, S., and Feun, L. 2007, Drug Target Insights, 2, 119–128 32. Dillon, B. J., Holtsberg, F. W., Ensor, C. M., Bomalaski, J. S., and Clark, M. A. 2002, Med Sci Monit, 8, BR248–253 33. Izzo, F., Marra, P., Beneduce, G., Castello, G., Vallone, P., De Rosa, V., Cremona, F., Ensor, C. M., Holtsberg, F. W., Bomalaski, J. S., Clark, M. A., Ng, C., and Curley, S. A. 2004, J Clin Oncol, 22, 1815–1822 34. Feun, L., Savaraj, N., Marini, A., and al, e. 2005, Proc Am Soc Clin Oncol, 23, 236s 35. Feun, L., and Savaraj, N. 2006, Expert Opin Investig Drugs, 15, 815–822 36. Curley, S. A., Bomalaski, J. S., Ensor, C. M., Holtsberg, F. W., and Clark, M. A. 2003, Hepatogastroenterology, 50, 1214–1216 37. Ascierto, P. A., Scala, S., Castello, G., Daponte, A., Simeone, E., Ottaiano, A., Beneduce, G., De Rosa, V., Izzo, F., Melucci, M. T., Ensor, C. M., Prestayko, A. W., Holtsberg, F. W., Bomalaski, J. S., Clark, M. A., Savaraj, N., Feun, L. G., and Logan, T. F. 2005, J Clin Oncol, 23, 7660–7668 38. Sugimura, K., Ohno, T., Kusuyama, T., and Azuma, I. 1992, Melanoma Res, 2, 191–196 39. Gong, H., Pottgen, C., Stuben, G., Havers, W., Stuschke, M., and Schweigerer, L. 2003, Int J Cancer, 106, 723–728 40. Gong, H., Zolzer, F., von Recklinghausen, G., Havers, W., and Schweigerer, L. 2000, Leukemia, 14, 826–829 41. Gong, H., Zolzer, F., von Recklinghausen, G., Rossler, J., Breit, S., Havers, W., Fotsis, T., and Schweigerer, L. 1999, Biochem Biophys Res Commun, 261, 10–14 42. Feun, L., Wu, C., Kuo, M., Wangpaichitr, M., Spector, S., and Savaraj, N. 2007, Current Phama Design 43. Hoyer-Hansen, M., and Jaattela, M. 2007, Autophagy, 3, 381–383 44. Hurley, R. L., Anderson, K. A., Franzone, J. M., Kemp, B. E., Means, A. R., and Witters, L. A. 2005, J Biol Chem, 280, 29060–29066 45. Sarbassov, D. D., Ali, S. M., and Sabatini, D. M. 2005, Curr Opin Cell Biol, 17, 596-603 46. Kim, R. H., Bold, R. J., and Kung, H. J. 2009, Autophagy, 5, 567–568 47. Smalley, K. S., and Herlyn, M. 2005, Ann N Y Acad Sci, 1059, 16–25 48. Fecher, L. A., Amaravadi, R. K., and Flaherty, K. T. 2008, Curr Opin Oncol, 20, 183–189 49. Wang, J., Whiteman, M. W., Lian, H., Wang, G., Singh, A., Huang, D., and Denmark, T. 2009, J Biol Chem 50. Maiuri, M. C., Zalckvar, E., Kimchi, A., and Kroemer, G. 2007, Nat Rev Mol Cell Biol, 8, 741–752 51. Cao, Y., and Klionsky, D. J. 2007, Cell Res, 17, 839–849 52. Levine, B., and Kroemer, G. 2008, Cell, 132, 27–42 53. Jin, S., and White, E. 2007, Autophagy, 3, 28–31 54. Pattingre, S., Espert, L., Biard-Piechaczyk, M., and Codogno, P. 2008, Biochimie, 90, 313–323 55. Zhang, H., Bosch-Marce, M., Shimoda, L. A., Tan, Y. S., Baek, J. H., Wesley, J. B., Gonzalez, F. J., and Semenza, G. L. 2008, J Biol Chem, 283, 10892–10903 56. Wang, Y., Singh, R., Massey, A. C., Kane, S. S., Kaushik, S., Grant, T., Xiang, Y., Cuervo, A. M., and Czaja, M. J. 2008, J Biol Chem, 283, 4766–4777 57. Shimizu, S., Kanaseki, T., Mizushima, N., Mizuta, T., Arakawa-Kobayashi, S., Thompson, C. B., and Tsujimoto, Y. 2004, Nat Cell Biol, 6, 1221–1228 58. Codogno, P., and Meijer, A. J. 2005, Cell Death Differ, 12 Suppl 2, 1509–1518 59. Kondo, Y., Kanzawa, T., Sawaya, R., and Kondo, S. 2005, Nat Rev Cancer, 5, 726–734 60. Amaravadi, R. K., Yu, D., Lum, J. J., Bui, T., Christophorou, M. A., Evan, G. I., ThomasTikhonenko, A., and Thompson, C. B. 2007, J Clin Invest, 117, 326–336 61. Lum, J. J., Bauer, D. E., Kong, M., Harris, M. H., Li, C., Lindsten, T., and Thompson, C. B. 2005, Cell, 120, 237–248 62. Kilberg, M. S., Pan, Y. X., Chen, H., and Leung-Pineda, V. 2005, Annu Rev Nutr, 25, 59–85 63. Levine, B., and Yuan, J. 2005, J Clin Invest, 115, 2679–2688

3 64. 65. 66. 67.

Argininosuccinate Synthetase in Cancer Therapy

51

Xie, Z., and Klionsky, D. J. 2007, Nat Cell Biol, 9, 1102–1109 Obara, K., Noda, T., Niimi, K., and Ohsumi, Y. 2008, Genes Cells, 13, 537–547 Scarlatti, F., Granata, R., Meijer, A. J., and Codogno, P. 2008, Cell Death Differ Bursch, W., Hochegger, K., Torok, L., Marian, B., Ellinger, A., and Hermann, R. S. 2000, J Cell Sci, 113 (Pt 7), 1189–1198 68. Mills, K. R., Reginato, M., Debnath, J., Queenan, B., and Brugge, J. S. 2004, Proc Natl Acad Sci U S A, 101, 3438–3443 69. Kanzawa, T., Germano, I. M., Komata, T., Ito, H., Kondo, Y., and Kondo, S. 2004, Cell Death Differ, 11, 448–457 70. Bilir, A., Altinoz, M. A., Erkan, M., Ozmen, V., and Aydiner, A. 2001, Pathobiology, 69, 120–126 71. Szlosarek, P., Klabatsa, A., Pallaska, A., Sheaf, f. M., Balkwell, F., and Fennell, D. 2006, Proc. Am Assoc Can Res, Washington D. C, USA 72. Carritt, B., and Povey, S. 1979, Cytogenet Cell Genet, 23, 171–181 73. Haggerty, D. F., Spector, E. B., Lynch, M., Kern, R., Frank, L. B., and Cederbaum, S. D. 1982, J Biol Chem, 257, 2246–2253 74. Gebhardt, R., and Mecke, D. 1979, Eur J Biochem, 97, 29–35 75. Lin, R. C., Snodgrass, P. J., and Rabier, D. 1982, J Biol Chem, 257, 5061–5067 76. Morris, S. M., Jr. 1987, Arch Biochem Biophys, 259, 144–148 77. Nebes, V. L., and Morris, S. M., Jr. 1988, Mol Endocrinol, 2, 444–451 78. Husson, A., Bouazza, M., Buquet, C., and Vaillant, R. 1983, Biochem J, 216, 281–285 79. Tomomura, M., Tomomura, A., Dewan, M. A., and Saheki, T. 1996, FEBS Lett, 399, 310–312 80. Grofte, T., Wolthers, T., Jensen, S. A., Moller, N., Jorgensen, J. O., Tygstrup, N., Orskov, H., and Vilstrup, H. 1997, Hepatology, 25, 964–969 81. Grofte, T., Jensen, D. S., Gronbaek, H., Wolthers, T., Jensen, S. A., Tygstrup, N., and Vilstrup, H. 1998, Am J Physiol, 275, E79–86 82. Hattori, Y., Shimoda, S., and Gross, S. S. 1995, Biochem Biophys Res Commun, 215, 148–153 83. Zhang, W. Y., Gotoh, T., Oyadomari, S., and Mori, M. 2000, Brain Res Mol Brain Res, 83, 1–8 84. Brasse-Lagnel, C., Fairand, A., Lavoinne, A., and Husson, A. 2003, J Biol Chem, 278, 52504–52510 85. Jackson, M. J., Allen, S. J., Beaudet, A. L., and O’Brien, W. E. 1988, J Biol Chem, 263 16388–16394 86. Nicholson, L. J., Smith, P. R., Hiller, L., Szlosarek, P. W., Kimberley, C., Sehouli, J., Koensgen, D., Mustea, A., Schmid, P., and Crook, T. 2009, Int J Cancer 87. Helleman, J., Jansen, M. P., Span, P. N., van Staveren, I. L., Massuger, L. F., Meijer–van Gelder, M. E., Sweep, F. C., Ewing, P. C., van der Burg, M. E., Stoter, G., Nooter, K., and Berns, E. M. 2006, Int J Cancer, 118, 1963–1971 88. Beloussow, K., Wang, L., Wu, J., Ann, D., and Shen, W. C. 2002, Cancer Lett, 183, 155–162 89. Park, I. S., Kang, S. W., Shin, Y. J., Chae, K. Y., Park, M. O., Kim, M. Y., Wheatley, D. N., and Min, B. H. 2003, Br J Cancer, 89, 907–914 90. Wu, G., and Morris, S. M., Jr. 1998, Biochem J, 336 (Pt 1), 1–17 91. Shen, L. J., Lin, W. C., Beloussow, K., Hosoya, K., Terasaki, T., Ann, D. K., and Shen, W. C. 2003, Biochem Pharmacol, 66, 1945–1952 92. Hao, G., Xie, L., and Gross, S. S. 2004, J Biol Chem, 279, 36192–36200 93. Jadeski, L. C., Hum, K. O., Chakraborty, C., and Lala, P. K. 2000, Int J Cancer, 86, 30–39 94. Zhang, X. M., and Xu, Q. 2001, Melanoma Res, 11, 559–567 95. Kilbourn, R. G., Fonseca, G. A., Griffith, O. W., Ewer, M., Price, K., Striegel, A., Jones, E., and Logothetis, C. J. 1995, Crit Care Med, 23, 1018–1024 96. Kilbourn, R. G., Fonseca, G. A., Trissel, L. A., and Griffith, O. W. 2000, Cancer J Sci Am, 6 Suppl 1, S21–30 97. Yang, T. -S., Lu, S. -N., Chao, Y., Sheen, I. -S., Lin, C. -C., Wang, T. -E., Chen, S. -C., Wang, J. -H., Liao, L. -Y., Thomson, J. A., Wang-Peng, J., Chen, P. -J., and Chen, L. -T. 2010, Br J Cancer, 103, 954–960

Chapter 4

Parathyroid Hormone–Related Peptide Signaling in Cancer* Franco Oreste Ranelletti and Giovanni Monego

Introduction Parathyroid hormone–related protein (PTHrP) was discovered in 1987 as a systemic humoral factor because of its association with the paraneoplastic syndrome of humoral hypercalcemia of malignancy (HHM) [1]. The hypercalcemic activity of PTHrP is based on its partial homology to parathyroid hormone (PTH) and on its ability to bind to the type 1 parathyroid (PTH)/PTH-related peptide receptor (PTH1R), with equal affinity as PTH, mimicking the stimulatory action of PTH on cAMP production in bone and kidney. This results in bone resorption and renal calcium retention, eventually leading to HHM. While PTHrP was discovered for its endocrine effects, malignant tumor, the secretory gland, and the bone tissue and kidneys being the target organs, it became clear that PTHrP is also expressed by nontransformed cells in almost all tissues where it functions locally in paracrine and/or autocrine manner. The most important regulatory roles of PTHrP relate to cell growth and differentiation, smooth muscle relaxation, and transepithelial calcium transport but not to serum calcium homeostasis [2]. Moreover, PTHrP behaves like an oncofetal protein, its expression being restricted to the period of fetal development or to malignancy. Although many cancers produce PTHrP, only a limited subset of them led to the development of HHM. About 50% of cases of HHM develop in the presence of squamous cell carcinomas, while the remaining cases are associated with kidney carcinoma, breast, colorectal, esophageal adenocarcinoma and hematologic neoplasia. However, despite the successful control of hypercalcemia, the development of HHM is associated with a poor prognosis [3], indicating that in addition to its

*

This work was supported in part by grant D1.1-2008 to F.O. Ranelletti from Università Cattolica del S. Cuore.

F.O. Ranelletti (*) Institute of Histology and Embryology, Università Cattolica del S. Cuore, Rome 00168, Italy e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_4, © Springer Science+Business Media, LLC 2012

53

54

F.O. Ranelletti and G. Monego

PTH-like effects, PTHrP has additional roles in cancer. Potential roles for PTHrP in the regulation of tumor growth and invasion are suggested by its growth factor– like properties together with the multifactorial regulation of its expression by growth and angiogenic factors, such as interleukins, epidermal growth factor (EGF), transforming growth factor-a (TGF-a), transforming growth factor-b (TGF-b), platelet-derived growth factor (PDGF), and vascular endothelial growth factor (VEGF) [2, 4]. Evidences have been accumulated, suggesting that the most important roles exerted by PTHrP in tumor development include: regulation of cancer cell growth, interference with apoptosis signaling pathways, acquisition of invasive phenotype by cancer cells, and induction and progression of bone metastasis. Moreover, the therapeutic potential of PTHrP-targeting strategies in human cancer has been highlighted by many studies, particularly in breast, prostate, lung, and kidney cancers [5–8]. This review article highlights some recent advances in the understanding of the signaling pathways involved in the PTHrP functions and of the factors that regulate PTHrP expression in cancer cells.

PTHrP Gene Structure and Regulation PTHrP owes the name to the structural homologies between the aminoterminal portion of the molecule and the corresponding aminoterminal portion of PTH. With regard to the primary structure, 8 of the first 13 amino acids (1–13) of N-terminal PTHrP are identical to PTH. Beyond the amino acidic sequence 1–13, the threedimensional structure of residues 13–34 of PTHrP presents strict similarities with PTH 13–34. PTHrP gene is localized on chromosome 12p11, whereas PTH, on 11p15, and it is considered that these two different chromosomes are derived from a common ancestor after tetraploidization events [9]. The human PTHrP gene presents a structure that spans more than 15 kilobases of genomic DNA and includes nine exons (Fig. 4.1). The 5¢-end of the gene is the region deputed to regulation of transcriptional processes and contains three distinct promoters named P1, P2, and P3, respectively [10–14]. P1 and P3 are promoters containing “TATA box”-like sequences, whereas P2 is a GC-rich promoter [14, 15]. P1 is located 25 bp 5¢ of exon 1, P2 is upstream of exon 3, and P3 is located in an intron 35 bp upstream to exon 4. The 5¢-untranslated regions in the mature PTHrP mRNA are referable to exons 1, 2, 3, and 4, while exon 5 encodes for the so-called prepro region. Exon 6 contains the major part of the coding region and, together with the product of exon 5, forms the sequence shared by all known PTHrP transcripts [16–18]. Exons 7, 8, and 9 are included in the 3¢-end of the gene and can be alternatively spliced in order to code three mRNA variants corresponding to three mature protein isoforms [11, 19]. These three PTHrP isoforms differ in length, being composed by 139 (PTHrP 1–139), 141 (PTHrP 1–141), and 173 (PTHrP 1–173) amino acids,

4

PTHrP Signaling in Cancer

55

Fig. 4.1 Structure of human parathyroid hormone–related peptide (PTHrP) gene. See text for details. The three different promoters are indicated as P1, P2, and P3; exons are shown as boxes numbered from 1 to 9. Alternative splice variants are shown by arrows. All the transcript variants are composed by a common part transcribed from exons 5 and 6 (red arrows) and variant specific parts transcribed by exon 7 (light blue arrow) for 1–139, by exon 8 (green arrow) for 1–173, and by exon 9 (yellow arrow) for 1–141 splice variants

respectively [1, 10, 11, 20]. Moreover, the three isoforms of PTHrP differ at the carboxyl termini, and at mRNA level, there are three different corresponding 3¢ transcripts. In fact, in exons 7, 8, and 9 coding sequences that are specific for PTHrP isoforms are located, and each of these exons also contains a specific 3¢-untranslated region (UTR). In detail, in addition to the common product of exons 5 and 6, PTHrP 1–139 is characterized by the product of exon 7; PTHrP 1–141 by the product of exon 8 and PTHrP 1–173 by the product of exon 9. The unique UTR contained in exons 7, 8, and 9 have strong influences on the stability of PTHrP mRNA splice variants, conditioning the half-life of the messengers. In fact, each of these 3¢-UTRs contains AUUUA regions, which are responsible for the rapid turnover of the mRNA with low steady-state levels of the transcripts, in spite of the high rate of PTHrP gene transcription [21–26]. These so-called AU-rich instability elements are present in the 3¢-UTR of PTHrP mRNA as well as in the sequences of the early response genes, such as c-fos, GM-CSF, and IL-2, which are also characterized by rapid turnover. The half-life of PTHrP mRNA ranges from 30 min to 4 h and can be increased by growth factors, such as TGF-b and EGF, probably by interfering with the binding of the 3¢-UTR with proteins that can promote rapid messenger degradation [27]. It has been reported that PTHrP mRNA 1–139 and 1–141 can be bound by proteins with molecular weights ranging from 37–40 kD to 80 kD, which are consistent with the molecular weights of p37AUF-1, p40AUF-1 (AUF-1 family members), and KSRP (K homology–type splicing regulatory protein), respectively [28, 29]. Moreover, IL-2 seems to play a role in stabilizing the PTHrP mRNA in HTLV-1 infected T cells [30]. PTHrP can theoretically start from all the three distinct promoters. Several studies were conducted on the expression of the three promoters of PTHrP, and after all, it is considered that P3 plays the major role in the regulation of the gene transcription in various cancers and tumor-derived cell lines [31]. As reported by some investigators,

56

F.O. Ranelletti and G. Monego

combined P2/P3 usage is associated with breast and bone tumor–derived cell lines, whereas P1 expression results detectable in squamous cancer and in cell lines of squamous origin [18, 32]. According to Dittmer [4], the PTHrP gene contains sequences for the binding of transcription factors. The authors described a Tax-responsive region upstream of the P3 promoter, and immediately upstream of P3 identified a 90-base region, which is probably inserted in the P3 core promoter and includes cis-acting elements involved in the recruitment of TATA binding protein–containing initiation complex. In this region, there are one inverted Sp1 DNA-binding consensus site and two overlapping Ets factor–binding sites (EBS I and EBS II). Ets factors are a family of nuclear transcription factors involved in the regulation of PTHrP gene expression, and Ets-1 is considered to play a pivotal role, particularly in PTHrP expression by adult-cell leukemia/lymphoma (ATLL) and during the differentiation of the T-cell lineage. [33– 37]. Furthermore, Ets-1 appears to exert a critical function in the TGF-b-mediated induction of PTHrP expression in MDA-MB-231 cells. Twenty-three bases downstream from EBS I site have been individuated a site for binding Smad complex. This Smadbinding site is located thirty-four bases upstream of the starting site of P3, which is considered the unique promoter to mediate the TGF-b-dependent activation of PTHrP. It is known that PTHrP expression can be increased by TGF-b through the dimerization of its type I and type II receptors and the phosphorylation of Smad proteins. The contiguity between EBS I and the Smad-binding site supports the hypothesis that the regulation of P3 could be based on strict interactions between Ets factors and Smad proteins. Experiments with mutated Smad, Sp1, or EBS I and with addition of Ets-1 confirmed the centrality of Ets-1 among the Ets family in PTHrP induction in breast cancer cell lines [38]. Nevertheless, the criticality of Ets-1’s role appears limited to the MDA-MB-231 cells. In fact, it has been reported that only Ets-1 is able to interact with Smad 3 to increase the transcriptional activity of PTHrP P3 promoter in MDA-MB-231 cells under TGF-b stimulation, whereas in MCF7 cells, Ets-2 mediates the increased PTHrP expression. Ets-1 is combined with the activation of PKCa second-messenger pathway, and Ets-2 is associated with the activation of PKCe second-messenger pathway, suggesting that TGF-b stimulation could be dependent on specific Ets factors combined with specific downstream signaling pathways [39, 40]. Moreover, Ets factors also appear to be involved in the regulation of PTHrP expression by the Ras–MAPK pathway [41]. Ets transcription factors represent the downstream target of Ras–MAPK signaling cascade, which articulates in three parallel arms terminating with ERK (extracellular signal–regulated kinase), p38, or JNK (Jun N-terminal kinase), respectively. Each of these three mitogen-activated kinases (MAPKs) converges to the phosphorylation of transcription factors and particularly of Ets family proteins [41]. In detail, ERK and JNK could interact with Ets-1 and Ets-2, which in turn could activate P3 through the EBS site upstream of the PTHrP promoter [42, 43]. The Ras–MAPK pathway consists of a sequence of signaling molecules that can work as common signal transducer for stimuli involved in the induction of PTHrP expression [20]. This pathway is composed by several arrays of kinases from the

4

PTHrP Signaling in Cancer

57

surface receptors on plasma membrane to the transcription factors in the nucleus, ranging from mitogen-activated kinase kinase kinase (MAPKKK) to mitogen-activated kinase kinase (MAPKK) and finally to mitogen-activated kinase (MAPK). Growth factor messages are generally passed on from Raf (MAPKKK) to MEK (MAPKK) to be transmitted to ERK (MAPK), whereas cytokines and stressors use ASK1 (MAPKKK), followed by MKK4 and MKK3 (MAPKK) terminating in JNK or p38, respectively. These two arms of Ras/MAPK cascade can be connected by Ras signaling through PI3 kinase and Rac/Cdc42 [44]. Moreover, Ras–MAPK pathway can also mediate the induction of PTHrP expression indirectly through stimuli that are recognized by the sensors connected with other transduction pathways that activate signaling molecules converging on Ras. For example, calcium is recognized by the calcium-sensing receptor (CaSR), which can promote the induction of matrix metalloproteinases through G protein activation, followed by mobilization of HB-EGF that can close the circuit activating EGFR and signaling through Ras–MAPK cascade [45–47]. Furthermore, the activation of PTHrP-P3 mediated by the ERK arm of the Ras–MAPK pathway could be triggered by the T-cell receptor, whereas the participation of the p38 arm to the TGF-b-dependent induction of PTHrP appears controversial [48, 49]. After all, it can be argued that different stimuli can converge on Ras–MAPK pathway in order to induce PTHrP gene expression. The PTHrP induction driven by P3 promoter activation is associated with the convergence on Ets proteins of the Ras–MAPK, TGF-b–Smad, and HTLV-1–Tax pathways and implies the cooperative interaction with the TATA box–associated basal transcription complex involving coactivators, such as CBP/p300 [33, 50]. The other TATA-containing PTHrP promoter P1 appears regulated by a cAMP-responsive element (CRE) positioned upstream of the starting point of exon 1 [51]. The position of the regulatory element and the P1 sensitivity to the induction by cAMP suggest that CRE is probably part of the core promoter and that P1 is the principal responsive target of the cAMP/PKA/CREB regulatory axis. It is considered that this pathway could be responsible for the PTHrP overexpression in squamous carcinoma cell lines associated with hypercalcemia, such as BEN lung cancer cells [32, 51, 52]. Little is known about the GC-rich promoter P2, results of which were expressed in several cell lines and tumors. It is possible to identify some regulatory elements upstream of the P2 starting site by screening the sequence with dedicated software. In detail, a vitamin D–responsive element (VDRE), three overlapping AP-2 sites, and two NF-kB sites were individuated [53]. An atypical VDRE is positioned 517–546 bp upstream of the P1 promoter of human PTHrP gene, supporting the participation of vitamin D to the regulation of PTHrP expression [54]. In this vitamin D–mediated regulatory mechanism, the vitamin D receptor (VDR), avoiding the recruitment of RXR (retinoid X receptor), is bound to the chromatinized atypical VDRE, and the stimulation with 1,25-(OH)2D3 causes the dissociation of the PTHrP VDRE/VDR complex, leading to the repression of gene expression from P1. The separation of VDR from the response element upstream of the promoter is caused by the phosphorylation of the receptor mediated by the catalytic subunit of the DNA-dependent protein kinase, which is recruited by 1,25-(OH)2D3 [55].

58

F.O. Ranelletti and G. Monego

1,25-(OH)2D3 administration can repress PTHrP gene expression in several cell lines, suggesting that this sterol could be able to interact with all the three promoters, as assumable based on a computerized analysis of gene sequence that identified three other potential VDR consensus sites upstream of the translation starting point. Another aspect of PTHrP gene structure is the presence of CpG dinucleotide sequences. The PTHrP gene presents a large CpG island located within the context of the GC-rich P2 promoter. Also, individual CpG sites may be positioned between P1 and P2. The functional spin-off for gene regulation of these CpG dinucleotide sequences appears still controversial. In fact, on the basis of the data reported by Broadus and Philbrick, it is assumable that methylation of residues upstream of the CpG island can repress PTHrP gene, whereas the data of Ganderton et al. suggest that methylation does not modulate gene expression [56–59]. After all, it is supposable that the methylation of upstream regions of PTHrP gene can modulate the expression in cell-type or promoter-dependent manners [31]. Also, PTHrP gene shows numerous potential glucocorticoid response elements within the region of P1 and P2 promoters, supporting the experimental evidences that suggest a strong inhibition of PTHrP expression determined by exposure to glucocorticoids in vitro [22, 60–64]. Nevertheless, these data were not confirmed by experiments on animal models [65, 66]. Similarly, controversy appears on the relationships between sex steroids and PTHrP expression, pointing out the need of further studies to investigate the structural bases and the mechanisms involved in the modulation of PTHrP expression by hormones [31].

Cancer Cell Growth There are evidences that PTHrP has tumor growth effects. In mice, specific overexpression of PTHrP in mammary gland produces a higher incidence of tumor formation [67]. A genetic polymorphism located in the osteostatin encoding region of the PTHrP gene, which produces Thyr/Pro PTHrP variants, was significantly associated with resistance or susceptibility to skin carcinogenesis in mice, suggesting a role of the PTHrP gene as a cancer modifier gene in skin carcinogenesis [68]. Treatment of rat with antisense oligonucleotides against PTHrP-RNA reduces the growth of pituitary cancer cells [69] and the size of tumor formed by implanted H-500 Leydig cells [70]. Furthermore, treatment of tumor-bearing mice with neutralizing PTHrP antibody inhibits growth of human MDA-MB-231 breast cancer cells metastasized to bone [71]. The effects of PTHrP on cancer cell growth are multifaceted. They reflect the fact that the native molecule can be processed into peptides that mediate pleiotropic actions through many molecular pathways. PTHrP should be considered as a polyhormone. In fact, a family of peptides arises from tissue-/cell-specific alternative splicing of the primary transcript. Moreover, alternative posttranslational endoproteolytic cleavage at dibasic amino acid regions forms N-terminal (PTHrP 1–36), midregion (PTHrP 38–94), and C-terminal (PTHrP 107–139) peptides with distinct pathophysiological

4

PTHrP Signaling in Cancer

59

Fig. 4.2 The functional domains of PTHrP protein. Signal peptide module serves to dock the nascent peptide to the secretory pathway. PTH1R indicates the receptor-binding region; midregion containing the bipartite NLS is essential for PTHrP nuclear import; osteostatin domain involved in bone turnover and b-arrestin binding. See text for details

functions (Fig. 4.2) and probably different cell surface receptors [72–76]. However, to date, only the PTH1R has been cloned, which is a G protein–coupled receptor (GPCR) belonging to the secretin-like GPCR family (class B), displaying a similar affinity for two distinct ligands, PTH and PTHrP N-terminal domains (PTHrP 1–36). The capability of PTH1R to respond in a specific manner to two different ligands can be attributed to various signal control mechanisms [77]. For example, unlike PTH, PTHrP has a second protein kinase C (PKC) activation domain, as indicated by the ability of picomolar concentrations of PTHrP to stimulate maximally membrane-associated protein kinase C (PKC) activity in the osteosarcoma cells [78]. PTHrP (1–36) binding to PTH1R activates adenylate cyclase (AC) through the action of stimulatory G-alpha proteins (Gas) coupled to the receptor. In turn, AC stimulates the formation of cyclic 3¢,5¢-adenosine monophosphate (cAMP) which binds to the regulatory subunit of protein kinase A (PKA) and releases the active catalytic subunits of the enzyme. Moreover, PTHrP (1–36) binding to PTH1R activates phospholipase Cb (PLCb) by Gaq, leading to the formation of diacylglycerol (DAG), which activates PKC and 1,4,5-inositol triphosphate (IP3), resulting in increased intracellular free Ca2+ [79]. It has been shown that high agonist concentrations (micromolar)

60

F.O. Ranelletti and G. Monego

and/or high receptor density are required for efficient activation of PLC; this is in contrast to PTH1R activation of AC, which occurs at physiologic (subnanomolar) agonist concentrations in the same cell host [80]. Interestingly, it has been observed that the Na+/H+ exchanger regulatory factor (NHERF), by binding to a PDZ domain within the carboxy terminal tail of PTH1R, shifts receptor signaling from AC to PLC through efficient coupling of the PTH1R–NHERF complex to Gq/11, which stimulates PLC and Gi, which inhibits AC [81]. Moreover, NHERF-mediated assembly of PTH1R and phospholipase C regulates PTHrP signaling in cells and membranes of polarized cells expressing NHERF and may account for many tissue- and cell-specific actions of PTH/PTHrP [81]. Desensitization of the receptor following agonist activation occurs mainly through phosphorylation of the C-terminal domain by second messenger–dependent protein kinases and GPCR kinases which is then followed by internalization of the receptor into clathrin-coated pits, a process primarily coordinated by the arrestins. Internalization of the receptor requires both ligand binding and receptor activation but does not involve stimulation of AC/PKA or PKC [82]. b-arrestins are recruited to agonist-occupied PTH1R that have been phosphorylated by GPCR kinases and sterically inhibit receptor-G protein coupling, which results in homologous receptor desensitization. Moreover, b-arrestins act as adaptors in clathrin-mediated receptor endocytosis [83]. In addition to their role as GPCR-specific endocytic adaptor proteins, b-arrestins also act as signal transducers via the formation of scaffolding complexes which control both the spatial and temporal activity of several MAPK modules, such as those of extracellular signal–regulated kinase (ERK1/2, ERK3/4, ERK5, ERK7/8), p38MAPK, and Jun N-terminal kinase (JNK)1/2/3 [84]. GPCR can stimulate ERK1/2 phosphorylation through an early G protein–dependent pathway mediated by PKA/PKC and a late G protein–independent pathway mediated by b-arrestin 1 and 2 [85]. Interestingly, it has been observed that b-arrestin and classical G protein–PKA/PKC pathways can be selectively activated by stimulating PTH1R with inverse agonist for cAMP or agonist ligands, respectively. This observation suggests that the receptor can exist in more active conformations, each one being able to initiate distinct signaling pathways [85, 86]. b-arrestin and classical G protein–PKA/ PKC pathways are not only mechanistically distinct but also perform different signaling functions. Thus, G protein–activated ERK1/2 generally translocate into the nucleus where they phosphorylate and activate many transcription factors, while b-arrestins-activated ERK1/2 compartimentalize in the cytoplasm where they can phosphorylate different sets of proteins [87, 88]. This mechanism could be of therapeutic impact as the two signaling pathways, leading to different cell responses, can be dissected. A recent advance in the understanding of PTH1R signaling is the discovery that several GPCR family members have been found to localize in the nucleus/nuclear membrane of cells. Similar to the propagation of signal through arrestins, nuclear GPCRs are providing a new avenue of investigation into the signaling potential of PTH1R [89]. PTH1R is the only member of the class B GPCR subfamily that translocates to the cell nucleus [90, 91]. It has been found that PTH1R contains a motif

4

PTHrP Signaling in Cancer

61

that fits the general consensus for a bipartite nuclear localization sequence (NLS). Moreover, the sequence of NLS is highly conserved across species, suggesting a key role for the nuclear import of PTH1R [92]. It has been suggested that NLS of PTH1R binds to importin a, which links the NLS-containing protein to importin b. Then, importin b docks the complex at the nuclear pore complex (NPC), facilitating entry of the PTH1R into the nucleus [91]. Nuclear PTH1R appears to be restricted to nucleoplasm and is not incorporated in the nuclear membrane [91]. PTH1R contains a nuclear export sequence (NES) which fits the generally accepted consensus for an NES and is highly conserved among several species. The association of PTH1R NES with chromosomal region maintenance 1 (CRM1) through a mechanism involving RanGTP results in the nucleus-to-cytoplasm translocation of the complex [93]. Nuclear–cytoplasmic shuttling of PTH1R appears to be regulated under normal physiologic conditions. In the mouse osteoblast-like cell line, MC3T3-E1, cultured in the presence of 10% fetal bovine serum, PTH1R displays both nuclear and cytoplasmic localization. On the contrary, in cells cultured under serum-starved conditions, PTH1R is predominantly localized in the nucleus. However, when cells are subsequently returned to normal serum conditions, PTH1R translocates from the nucleus to the cytoplasm where it appears predominantly localized. Interestingly, this effect of serum can be mimicked by the addition of the PTH1R ligand, PTHrP, which induces nuclear–cytoplasmic translocation in a time-dependent manner [93]. These observations suggest that nuclear import of PTH1R occurs constitutively through a ligand-independent mechanism, while its nuclear export depends on the presence of PTHrP. Moreover, it has been documented that, during interphase and telophase, PTH1R is in the nucleus when DNA is in the euchromatin state and transcriptionally active. In contrast, during stages in which DNA is compacted in heterochromatin states, such as prophase, metaphase, and anaphase, PTH1R nuclear localization is at a minimum [91]. Taken together, these observations suggest that PTH1R has a defined role in regulating cellular activities possibly by interacting directly with DNA and functioning as a transcription factor. PTH1R nuclear localization has been reported in association with ligand-independent proliferation [90]. PTH1R has a dual purpose in the cell; in addition to the classical membranebound signaling, it also has an intracellular signaling role that is largely unknown. Moreover, membrane PTH1R signaling is activated by ligand binding, while the nuclear PTH1R activity in the transcriptional control of genes is arrested by the ligand-dependent translocation of PTH1R out of the nucleus. However, if PTH1R deserves other functions, once that it is in the cytoplasm, is still unknown. Interestingly, overexpression of PTH1R confers a more aggressive phenotype to osteosarcoma cell line and results in increased proliferation, motility, and Matrigel invasion without addition of exogenous PTHrP, suggesting an autocrine mechanism. This cell behavior is nearly completely reversed by RNAi-mediated gene silencing [94], suggesting that the rate-limiting step in signal transduction might be at the level of the receptor that osteosarcoma cells possess rather than the availability of the ligand. In agreement with this observation, it has been reported that PTH1R is a better predictor than PTHrP for bone metastases and survival in breast cancer patients [95].

62

F.O. Ranelletti and G. Monego

Although many biological effects of PTHrP are mediated via signal transduction pathways initiated at membrane receptor(s), it has become apparent that PTHrP exerts cell surface receptor–independent biological actions mediated through an intracrine mechanism based on nuclear/nucleolar localization of the peptide. In fact, PTHrP encodes an NLS within the 87–107 residues of the mature protein (Fig. 4.2) and contains at least two translational initiation sites, one that generates a conventional signal peptide and one that, beginning at CUG codons downstream from the normal start codon, results in truncated PTHrP signal peptide which does not mediate entry into the secretory pathway [96]. Then, PTHrP can be either secreted or retained in the cytoplasm and translocated to the nucleus by binding to importin b [97, 98]. Alternative potential mechanisms of PTHrP entry into the cytoplasm have been proposed [99–101]. PTHrP can shuttle in both directions between cytoplasm and nucleus via the NPC, and it has been demonstrated that microtubule integrity plays an important role in PTHrP nuclear import [102]. PTHrP possesses a consensus motif for phosphorylation by p33cdk2 and p34cdc2 cyclin-dependent kinases immediately upstream of the NLS at Thr85 (Fig. 4.2), and phosphorylation at Thr85 prevents its nuclear import [98]. Moreover, it has been reported that CRM1/exportin 1 fulfills the task of PTHrP nuclear export receptor [103]. Although PTHrP nucleo-cytoplasmic flux occurs through distinct import and export receptors, the inhibition of nuclear export determines a concurrent reduction of PTHrP nuclear import, suggesting that the subcellular localization of PTHrP is regulated by an integrated operating system [103]. Moreover, it has been reported that nuclear translocation of PTHrP is cell cycle–dependent. However, in human keratinocyte cell line, HaCaT, PTHrP appears localized in the nucleolus in G1, but it relocalizes to the cytoplasm in mitotic cells [104], while in A10 vascular smooth muscle cells, PTHrP is localized in the nuclei of cells in the G2 or M phase of the cell cycle, suggesting that nuclear translocation is activated during cell division [97]. Since PTHrP NLS is homologous to that in other transcription factors, such as c-jun, c-fos, and p53, that directly bind DNA, it seems that PTHrP could also be a transcription factor that directly binds DNA. Interestingly, it has been recently reported that, in MDA-MB231 cells, PTHrP (38–94) amide can directly bind in vitro both discrete sites exposed by cellular chromatin and a purified 20-mer oligonucleotide and that it is involved in the transcriptional regulation of genes encoding for apoptosis factors and caspases [105]. As far as PTHrP proliferative effects are concerned, both positive and negative actions have been described depending on the combination of various factors, such as cell type, cell state, the effectual PTHrP domain (NH-terminal, midregion, COOH-terminal), paracrine/autocrine versus intracrine mechanisms, and the activity of signaling networks [4, 106, 107]. For example, a stimulatory effect on cell growth by cAMP/PKA has been described in PC12 pheochromocytoma cells [108], in PC-3 prostate carcinoma cells [109], and in rat enterocytes [110]. On the contrary, inhibition of growth has been reported in skin fibroblasts [106] and in PTHrP (1–34)-stimulated MCF-7 cells [111, 112]. The aberrations in cancer are pleiotropic, but mitogen-activated protein kinase (MAPK) pathways feature prominently. Mutations in MAPK pathways are a

4

PTHrP Signaling in Cancer

63

frequent occurrence in cancer and mostly affect Ras and B-Raf in the ERK pathways. PTH1R through AC–PKA and PLC–PKC pathways modulates the MAPK modules involved in many signaling pathways which regulate critical cell functions, such as proliferation, apoptosis, and differentiation [113]. The underlying mechanisms regulating the cross talk between PTH1R and MAPK pathways need to be elucidated in order to provide new therapeutic approach for cancer patients. The ERK pathway is the best studied of the mammalian MAPK pathways, since it is deregulated in approximately one third of all human cancers [114]. For example, it is interesting to note that in thyroid [115] and melanoma [116] cancer cells in which B-Raf mutations are common, cAMP is reported to activate ERK signaling rather than to inhibit it.

Regulation of ERK1/2 by Gs GPCRs through Gas-coupled subunit transduce signals to adenylyl cyclase, which converts ATP to cAMP. cAMP activates PKA, which plays a major role in Gasmediated stimulation as well as inhibition of MAPK modules, such as those of ERK1/2 and p38MAPK [117–125]. Gas-mediated stimulation of ERK1/2 appears to be mediated by a pathway involving Rap-1 and its activation of B-Raf, one of the isoforms of Raf [120, 124, 126–128]. It has been reported that Gas stimulation of ERK1/2 could be mediated at least by three different pathways: (1) Gas activates Rap-1 through cAMP-mediated, PKA-independent activation of EPAC (exchange protein directly activated by cAMP), a Rap-1-specific guanine nucleotide exchange factor [129–131]. (2) Gas exerts Rap-1-mediated activation of B-Raf–MEK–ERK module via a pathway involving cAMP–PKA–Src-mediated activation of C3G (Rap-1-GEF known as Crk SH3 domain-binding guanine nucleotide-releasing factor), which, in turn, stimulates Rap-1 [127, 128, 132, 133]; (3) Gas stimulates Ras signaling to ERK1/2 module via PKA-dependent or PKA-independent mechanism in a cell type–specific manner. Gas-mediated PKA-dependent stimulation of ERK1/2 involves a PKAdependent Ras-GEF (Ras-GRF1) [134, 135], whereas PKA-independent signaling involves a cAMP-responsive but PKA-independent Ras-GEF (CNrasGEF) as observed in melanoma cell lines [136] (Fig. 4.3). Gas-mediated inhibition of ERK pathways involves PKA-mediated phosphorylation at S43, S233, and S259 sites within N-terminal domain of a specific isoform of Raf, known as Raf-1 or C-Raf [117–119, 137–139]. Every one of the three phosphorylation sites blocks C-Raf activation and C-Raf interaction with Ras when cAMP levels are elevated (Fig. 4.3). PKA-phosphorylated S233 and S259 sites recruit 14-3-3 proteins to the N-terminus of C-Raf [140]. It has been suggested that, in the presence of elevated cAMP levels, PKA phosphorylates C-Raf on S43 and S233, creating a new 14-3-3 binding site, causing a rearrangement that switches 14-3-3 from the S621 site to the S233 site, but maintaining contact with the S259 site. C-Raf is then locked in a closed conformation that cannot

64

F.O. Ranelletti and G. Monego

Fig. 4.3 Gs regulation of ERK1/2 modules. See text for details. Gas stimulates the B-Raf-mediated activation of ERK modules via Rap-1 or Ras and inhibits Raf-1-mediated activation of ERK1/2 by phosphorylating Raf-1 through PKA. Also shown is the alternate PKA-independent pathway through which Gas activates Rap-1-mediated ERK1/2 activation via cAMP-activated EPAC

be activated because Ras cannot displace 14-3-3 from the N-region [124]. Interestingly, the C-terminal tail of PTH1R binds 14-3-3 proteins independently from ligand activation, and the phosphorylation of the PTH1R C-terminal tail by PKA reduces the interaction of the receptor with 14-3-3 [141], thus freeing 14-3-3 for interactions with other partners, such as C-Raf. Such a mechanism could potentiate the inhibition of Ras–C-Raf–ERK pathway mediated by the PKAdependent phosphorylation of 14-3-3 proteins. In fibroblast cell lines, such as NIH3T3 cells, it has been shown that the inhibition of C-Raf can also be determined by Gas-cAMP–PKA-mediated activation of Rap-1 [142, 143]. Once activated, Rap-1 binds and sequesters C-Raf from being activated by Ras as the effector domain of Rap-1 displays close sequence and structural similarities with that of Ras [142–144]. In addition to ERK1/2, Gas stimulates p38MAPK [121] module via cAMP–PKA pathway. Since it has been shown that Rap-1 can stimulate p38MAPK in various cell types [145, 146], a possible role of Rap-1 in this pathway can be hypothesized. Furthermore, Gas inhibits ERK5 module via cAMP–PKA pathway [125].

4

PTHrP Signaling in Cancer

65

Regulation of ERK1/2 by Gaq The Gq family of G proteins comprises Gaq, Ga11 and hematopoietic cell–specific Ga14, and Ga15/16 subunits [147, 148], which are able to activate ERK1/2, JNK, and p38MAPK modules. GPCRs transduce signals through the a-subunits of Gq family of G proteins to specific cellular responses via the activation of the effector PLCb [147, 149]. In turn, PLCb produces inositol triphosphate (IP3) and diacylglycerol (DAG) by hydrolyzing phosphatidylinositol 4,5-bisphosphate (PIP2). Both IP3 and DAG can directly activate protein kinase C (PKC). Moreover, they can indirectly activate PKC via the release of internally stored Ca2+. In addition to the Ga-subunit, also the bg-subunit released from Gq, activates PLCb [150, 151], signaling through both PLC–DAG–PKC- and PLC–IP3–Ca2+-mediated pathways. Gaq-activated PKC can stimulate ERK1/2 module either directly by phosphorylating and stimulating C-Raf [152–154] or indirectly by Ca2+–calmodulin-mediated activation of Pyk2, leading to the activation of Ras–ERK1/2 [155–157]. However, whether Gaq/11 activates ERK1/2 via the PKC–Raf signaling axis or IP3-Ca2+– calmodulin–Pyk2–Src–Ras pathway appears to be cell type–dependent [156–158]. In addition to these typical Gaq-mediated pathways, it has also been shown that Gaq can activate ERK through a novel mechanism involving a DAG- as well as a Ca2+-dependent Rap-1-GEF [159]. In summary, Gaq-mediated signaling to ERK1/2 is mediated by three different mechanisms: (1) activation of C-Raf mediated by PKC, (2) activation of Ras via Ca2+–Pyk2–Src-dependent pathway, and (3) activation of B-Raf mediated by Ca2+–DAG-stimulated Rap1(Fig. 4.4). It should be noted that all of these mechanisms need not be mutually exclusive as Gaq/11 activation results in the generation of both DAG and IP3. In addition, bg-subunits dissociate from GPCR coupled Gaq/11 and can contribute to Gaq/11 signaling to modules by stimulating PLCb. However, GPCR coupled to Gaq/11 activates ERK1/2 by utilizing preferentially PKC- and/or IP3–Ca2+ signaling pathways rather than bg-subunits [160]. The cell response to PKC activation can be either proliferative or antiproliferative in different cell types. Generally, PKC/C-Raf/Ras/ERK 1–2 pathway leads to cell proliferation. However, PKC activation can also result in inhibition of ERK1/2 modules and blocking of cell growth [161]. The type of cell response following PKC activation is PKC isoform–dependent as novel PKC isoforms (d, e, h, q, m), which are activated by DAG but not Ca2+-dependent, inhibit ERK1/2 activation by phosphorylating C-Raf and preventing its activation by Ras. Classical isoforms (a, bI, bII, g), which are both DAG- and Ca2+-dependent, and atypical isoforms (z, l, i), which are neither DAG- nor Ca2+-dependent, stimulate ERK1/2 module by phosphorylating Raf kinase inhibitory protein (RKIP) and determining its dissociation from C-Raf, which, in turn, activates downstream signaling to ERK1/2 module by associating with Ras [162, 163]. The cross talk between PTHrP and growth factor signaling pathways may result in additive or antagonistic effects on cancer cell proliferation. An example of additive effect comes from human oral cancer where PTHrP potentiates epidermal

66

F.O. Ranelletti and G. Monego

Fig. 4.4 Gq regulation of ERK1/2 modules. Gq activates ERK1/2 modules via PKC- as well as Ca2+-dependent pathways. Moreover, the bg subunits can activate ERK1/2 modules through CaMkinase–calmodulin–Pyk2–Src–Shc–Sos–Ras pathway. See text for details

growth factor (EGF) receptor (EGFR) signaling and promotes the malignancies of oral cancers [164]. EGF is secreted in the oral environment, and most oral cancer cells express abundant EGFR. Activation of EGFR upregulates the expression of PTHrP through ERK1/2 and p38 MAPK pathways. In turn, secreted PTHrP activates ERK1/2 MAPK and phosphatidylinositol 3-kinase (PI3K)/Akt pathways, potentiating the effects of EGF on tumor cell proliferation/survival, migration, and invasion and thus creating a vicious cycle. This observation suggests that targeting PTHrP may be beneficial to shut down oral squamous cell cancer progression, and the possibility that the combined application of PTHrP inhibition on dose reduction of EGFR-tyrosine kinase inhibitors may contribute to avoid the EGFR-TKI-derived adverse aspects, maintaining the antitumor potency [165]. Targeting PTHrP in preclinical settings has been attempted using a neutralizing antibody, antisense oligonucleotides, and RNAi, which resulted in prolonged survival of hypercalcemic mice

4

PTHrP Signaling in Cancer

67

[165], decreased cell proliferation of renal carcinoma [166], and increased apoptosis in human medulloblastomas [167], respectively. PTHrP and hedgehog (HH) signaling plays an important and coordinated role in regulating the differentiation of chondrocytes in the growth-plate cartilage through a negative-feedback loop [168]. PTHrP, secreted from growth-plate chondrocytes, acts on PTH1R on proliferating chondrocytes to keep them proliferating by inhibition of the cyclin-dependent kinase inhibitor p57Kip2 [169]. When chondrocytes are no longer sufficiently stimulated by PTHrP, they stop proliferating and synthesize Runx2 which induces the expression of Indian hedgehog (IHH) [170] which can then stimulate the production of PTHrP in growth-plate chondrocytes [171]. A similar mode of operation has been reported in breast cancer cells where PTHrP expression is driven by Gli2 [172], a mediator of HH signaling. Interestingly, cancer cells, when metastasize in the bone microenvironment, assume an osteomimicry property, responding to transforming growth factor b and taking advantage of gene regulatory mechanisms operative under the physiologic conditions. Moreover, metastatic cells use master transcription factors, such as Runx2/Cbfa1, to support their survival and to promote osteolytic disease through activation of the IHH–PTHrP pathway [173]. Recently, a direct regulation of HH-mediated transcriptional activation by PTHrP has been uncovered by studying the interaction of the two pathways in normal and neoplastic growth-plate chondrocytes [174]. HH-activated patched (PTCH) releases its inhibition on smoothened (Smo), which converts the transcription factor Gli3 from transcriptional repressor to transcriptional activator [175], facilitating its translocation to the nucleus. PTHrP-activated PTH1R promotes, via PKA activity, the cleavage of the full-length activator form of Gli3 to its repressor form, thus blocking HH signaling downstream of PTCH and Smo. Then, in selected tumor type, with mutations in PTCH or Smo, such as chondrosarcoma and medulloblastoma, PTHrP treatment could be used to decrease the HH signaling pathway. However, it has been reported that Gli2, but not Gli3, expression induces an increase of PTHrP production only in those breast cancer cell lines that cause osteolytic lesions, thus favoring the PTHrP role in the bone metastasis progression [172]. Then, it is still unclear whether the PTHrP inhibitory effect on HH signaling is Gli type- (Gli3 vs. Gli2) and/or context- and cell type–dependent.

Apoptosis Apoptosis is involved in the control of cell proliferation, and the interference with apoptosis signaling represents a potential mechanism to promote cancer growth and progression. PTHrP is generally considered as an antiapoptotic factor able to inhibit major apoptosis signaling pathways, but variable, and sometimes contradictory, data are reported in the literature. Actually, it is not possible to come to only one point of view on the role of PTHrP in the apoptotic process. The effects of PTHrP on apoptosis probably depend on tissue and cell type, cell line and clonal variation, physiological

68

F.O. Ranelletti and G. Monego

and pathological context, as well as on peptidic fragment. For example, in regard to lung cell physiopathology, Hastings reported that A549 cells and adult rat type II pneumocytes actively express PTHrP and PTH1R proteins, showing the possibility of producing peptide and responding to it through the cognate receptors, following autocrine/paracrine mechanisms. Hastings describes a sensitization of type II cells to apoptosis mediated by PTHrP 1–34 and PTHrP 67–86 [107, 176]. Nevertheless, Hastings et al. reported that PTHrP 1–34 and PTHrP 140–173 improve resistance of BEN cells to apoptosis induced by exposure to UV irradiation. This effect is reached through the reduction of the activity of caspase-3, -8, and -9. The modulation of caspases is associated with PKA activation [177] and cAMP increase. PTHrP 140–173 can regulate caspases also after stimulation of Fas (CD95) by activating antibody [178]. These data on the inhibitory influence of PTHrP on the apoptotic pathways are in accord with our data noticed from studying human medulloblastoma DAOY cells [167]. In fact, we observed that PTHrP gene silencing was associated with significant increase of apoptotic cell number in vitro. Capase-3 showed marked elevation of mRNA levels of about seven- to eightfold, involving the two major pathways converging on caspase-3 activation. Mitochondrial-mediated and death receptor–mediated signals showed different levels of activation. With respect to the intrinsic pathway, the Bax/Bcl-2 ratio mRNA levels rised to of about 160%, whereas Fas mRNA level reached a peak of about 1,700% in the context of the extrinsic pathway. Our data suggest that PTHrP can regulate apoptosis in DAOY cells prevalently by the modulation of death receptor–mediated signal and by the parallel minority involvement of mitochondrial-mediated signal. The correlations between PTHrP and Fas, as well as with antiapoptotic proteins, were confirmed by Gagiannis et al. [179], who developed the investigation on PTHrP interference with apoptosis pathways mediated by death receptors and mitochondria. Gagiannis and coworkers studied the role of PTHrP in the resistance of Saos-2 human osteosarcoma cells to the apoptosis induced by chemotherapeutic drugs. They noticed that both N-terminal and C-terminal PTHrP could inhibit significantly the apoptosis induced by doxorubicin and bleomycin, while the apoptotic effect of mitoxantrone is counteracted only by N-terminal PTHrP. This suggests that these antiapoptotic effects of PTHrP could be mediated by a PTH1R-dependent and cAMP/PKA-based mechanism as well as by a PTH1R-independent mechanism. At caspase level, the authors reported an inhibition of the activating effect of doxorubicin on caspase-2, -3, -6, -8, and -9 exerted by PTHrP 1–34 and PTHrP 107–139. These two peptides showed an ability to reduce the apoptosis produced by the stimulation with respective specific antibodies or ligands of CD95, TRAIL-R, and TNF-R, in addition with administration of cytostatic drugs, such as bleomycin and doxorubicin. The antagonistic influence of PTHrP on the extrinsic apoptosis signaling pathway was expressed also on the intrinsic pathway. In fact, the destabilizing proapoptotic effects of bleomycin and doxorubicin on mitochondrial membranes were counteracted by N-terminal and C-terminal PTHrP, whereas only N-terminal PTHrP was able to antagonize the effect of mitoxantrone. The exposition of Saos-2 cells to PTHrP 1–34 and PTHrP 107–139 interfered with the expression of proteins acting on mitochondria by producing accumulation

4

PTHrP Signaling in Cancer

69

of antiapoptotic Bcl-2 and Bcl-xl proteins while inducing downregulation of PUMA and Bax proapoptotic proteins. The consequent increase of the Bcl-2 and Bcl-xl to Bax ratio stops intrinsic apoptosis as well as the repression of the BH-3-only molecule PUMA, a direct transcriptional target of PTHrP and a promoter of oncogenic transformation [180]. Upstream to the extrinsic and intrinsic pathways, both N-terminal and C-terminal PTHrP exerted a significant inhibition of p53 family–mediated apoptosis. Moreover, this repression of transcriptional activation of p53 determined the consequent inhibition of the transactivation of target genes belonging to mitochondrial and death receptor–mediated apoptosis. In detail, N-terminal and C-terminal PTHrPs were able to inhibit p53-dependent transactivation of CD95 gene as well as the transactivation of Bax gene. This work reveals that PTHrP can inhibit apoptosis and confer chemoresistance by interfering with key regulator genes of apoptosis signaling cascade. Apoptosis modulation by PTHrP develops through p53 family–dependent as well as p53 family–independent signaling pathways and can affect both death receptor–mediated and mitochondria-mediated apoptosis. The results of PTHrP regulatory influence are critically linked to tumorigenesis because they consist in resistance to apoptosis induced by treatment with chemotherapeutic drugs and consequent promotion of tumor progression.

Invasiveness and Integrins Invasive phenotype is an integral part of the arsenal of cancer cells. Local invasiveness is the base of the metastatic potential of tumors, and it requires modifications of intercellular interactions as well as modification of interactions between cancer cell and extracellular matrix (ECM). It is reasonable to assume that the development of an invasive phenotype is associated with alterations in cancer cell adhesion and migration. Also, matrix proteolysis and degradation are considered key elements of the complex process of invasion–metastasis cascade. PTHrP intervenes in tumor invasiveness by regulating the expression of adhesion molecules, such as integrins. Integrins mediate cell–cell and cell–ECM adhesion and are considered able to influence cancer cell invasiveness and metastasizing capability. They belong to a family of transmembrane adhesion receptors that are heterodimers composed of a- and b-subunits. Different combinations of these subunits form receptors with different ligand specificities [181]. The ligands of integrins are the components of ECM: collagen type IV and laminin at level of basement membrane, and collagen type I and fibronectin at level of interstitial stroma. Adhesive interactions and attachment to ECM, plus local lyses and disruption of basement membrane, represent the essential steps for cancer cells to seep into stroma and to gain access to vascular compartment, spreading out through the circulation. Modifications in integrin expression and combination have been reported in association with cancer progression in vitro and in vivo [182–186]. Moreover, integrins

70

F.O. Ranelletti and G. Monego

can support tumor development by activating intracellular signaling pathways involved in the regulation of cell growth, differentiation, apoptosis, and motility [187]. After all, integrins are considered to play a major role in the development of the aggressive behavior of cancer cells [188, 189]. In PC-3 prostate cancer cells, Shen and Falzon observed that endogenous overexpression of wild-type PTHrP was selectively associated with increased expression of a1, a5, a6, and b4 integrin subunits, whereas in MDA-MB-231 and MCF-7 breast cancer cells, the overexpressed peptide upregulated only a6b4 integrin [187, 190–192]. Other studies on PTHrP overexpression highlighted that in MDA-MB-231 cells, the expression of a5, a6, b1, and b4 integrin subunits was enhanced, while in HT-29 colon cancer cells, only the expression of a5 and b1 subunits was increased [193]. The authors verified that the increased integrin subunit expression occurred both at transcriptional and cell surface levels, and they observed that the expression levels of the peptide directly correlated with those of the integrins. The involvement of a transcriptional mechanism in PTHrP-mediated effects on integrin expression, cell migration, and adhesion was confirmed by gene silencing experiments. The modulation of cell adhesion/migration is probably a PTHrP-mediated effect active in physiological conditions, exerted by the peptide at the transcriptional and/ or posttranscriptional level by increasing the gene transcription of the integrin subunits. PC-3 cells overexpressing wild-type PTHrP showed strong cytoplasmic and especially nuclear immunostaining, suggesting a nuclear accumulation of the peptide. This is in accord with the hypothesis that the intracrine pathway could mediate the regulatory effect of PTHrP. In fact, this upregulation was dependent on the presence of an intact NLS, particularly on the presence of amino acids 88–91, which are involved in importin b pathway (Fig. 4.2) [98]. The pivotal role of intracrine mechanism of action is supported by the observation that the administration of exogenous PTHrP 1–34, which lacks NLS and cannot enter the nucleus/nucleolus compartments, failed to induce significant modification of the expression of the integrins studied. The intracrine pathway is based on an influence exerted by PTHrP on gene transcription, either directly probably working as a transcription factor or indirectly by regulating the activities of other transcription factors. Integrin subunits selectively upregulated by endogenously overexpressed PTHrP are combined to assemble different heterodimeric adhesion receptors with specific adhesion targets. In detail, integrin a1b1 is deputed to mediate the adhesion to collagen type IV of basement membrane and to collagen type I of interstitial matrix. Also at the level of basement membrane, integrin a6b1 mediates the attachment to laminin, and the increased expression of this heterodimer can promote cancer cell attachment and the consequent transversion. Integrin a6b4 is considered an adhesion receptor of the laminins [194] and is also considered able to enhance migration and invasion of non-laminin ligands, such as collagen. The adhesion to stromal fibronectin appears mediated by a5b1 integrin [193]. Cell adhesion assays showed that PTHrP overexpression produced increased adhesion of MDA-MB-231 cells to collagen type I, laminin, and fibronectin, whereas

4

PTHrP Signaling in Cancer

71

in HT29 cells, the adhesion to collagen type I and fibronectin was enhanced. Transfection with NLS-mutant PTHrP confirmed the suggestion that the effects on cell adhesion and integrin expression induced by the peptide require its transport to the nucleus via an intracrine mechanism. Nevertheless, not all data obtained on the regulation of integrin expression and cell adhesion by PTHrP are concordant. For example, Anderson et al. [193] observed absence of correlation between integrin a6 and PTHrP expression in HT29 cells, while Shen and Falzon [195] reported that in LoVo colon cancer cells, the overexpression of PTHrP was associated with enhanced expression of a2, a5, a6, b1, and b4 integrin subunits. Ye et al. [196] did not find changes in adhesion to fibronectin and laminin in HT29 cells overexpressing PTHrP, while, consisting with previous data, Shen and Falzon [195] reported that LoVo cells overexpressing the peptide showed increased adhesion to collagen type I, laminin, and fibronectin. These conflicting evidences support the hypothesis that the regulation of integrin expression and cell adhesion exerted by PTHrP is tissue-specific and cell line–specific, and this specific regulation could influence the subsequent migration–metastasis sequence. In fact, basement membrane adhesion and transversion are the bases of tumor invasiveness and metastatic potential [197–202]. Another aspect that has to be considered in PTHrP-mediated contribution to cancer invasiveness is the upregulation of protein involved in matrix remodeling. The increased expression of matrix metalloproteases MMP2, MMP3, MMP9, and uPA (urokinase-type plasminogen activator) induced by exogenous peptides 1–141 and 1–84 in vitro could promote ECM degradation, favoring cancer cell invasion of the stroma [4]. In vitro studies highlighted that PTHrP overexpression is directly correlated with increased migration of MDA-MB-231 and MCF-7 cells. This enhancement in breast cancer cell migration induced by PTHrP is mediated by the upregulation of integrin a6b4, which represents a crucial point required for the development of PTHrP stimulatory effect [192]. Also, in PTHrP-overexpressing LoVo cells, migration and invasion were increased as well as the levels of integrin a6b4 subunits [187]. Expression/overexpression of integrin a6b4 has been associated with increased cancer aggressiveness [203–210]. Moreover, the progression of breast, thyroid, colorectal, gastric, and bladder tumors has been reported to correlate with the expression of integrin a6b4. Furthermore, integrin a6 can promote cancer cell metastasis to bone and engraftment in bone microenvironment by exerting its laminin-binding function on the laminins of bone extracellular matrix [211]. As reported above, in MDA-MB-231 cells, the overexpression of PTHrP induces enhanced adhesion to collagen type I, laminin, and fibronectin, the bone being composed of 95% of collagen type I, with laminin and fibronectin included in ECM [212]. This correspondence between upregulated integrin profile and presence of its ligands in the target tissue can help to explain the tendency of a primary tumor, such as breast cancer, to metastasize predominantly to a selected organ, such as bone. Another aspect of this integrinmediated tropism is based on the prevalence in the space of Disse of fibronectin and collagen type I, which are probably targeted by gastrointestinal cancer cell overexpressing a5b1 integrin subunits, as demonstrated by Anderson et al. [193].

72

F.O. Ranelletti and G. Monego

Besides modulating cell adhesion and migration, PTHrP-mediated modulation of integrin expression can support cancer development by other strategies involving cell proliferation and survival. In fact, integrins appear also able to influence intracellular signaling pathways involved in the regulation of cell growth and apoptosis [188, 189]. Overexpressing PTHrP in MDA-MB-231 and MCF-7 cells was associated with a significant increase in cell survival, which was reversed by cell transfection with PTHrP siRNA [192]. The PTHrP-overexpressing cells showed reduced apoptosis, while gene silencing produced an increase in apoptotic process [192]. These protective effects of the peptide against apoptosis were dependent on a6b4 subunit expression. Moreover, in PTHrP-overexpressing cells, augmented levels of phosphorylated Akt (p-Akt) and phosphorylated glycogen synthase kinase-3 (p-GSK-3) were detectable. In LoVo cells, Shen et al. [187] noted parallel increases in the expression of PTHrP, integrin a6b4, and PI3-K pathway components. Based on these data, it is assumable that the increased expression of PTHrP could increase the expression of integrin a6b4, which, in turn, could activate the pathway of phosphatidylinositol 3-kinase (PI3-K) [204]. PI3-K is a ubiquitous lipid kinase that can mediate the transduction of signals from tyrosine kinase receptors and GPCRs and is considered able to influence colon cancer progression [213–219]. PI3-K effects are mediated by its downstream effector Akt, which appears increased in several cancers, and especially in colon cancer, it correlates with clinicopathological parameters suggestive of cancer progression [220–222]. In LoVo colon cancer cells, the overexpression of PTHrP correlates with the levels of the phosphorylated Akt (p-Akt), which corresponds to the active form of Akt [187, 223, 224]. Similarly, the levels of p-Akt paralleled the expression of integrin a6b4 and PTHrP in MDA-MB-231 and MCF-7 cells, and also the levels of p-GSK-3 increased [192]. In fact, the phosphorylation of Akt leads to the phosphorylation of GSK-3 (glycogen synthase kinase-3). This glycogen synthase kinase-3 phosphorylates many proteins, such glycogen synthase, c-Myc, and cyclin D, keeping them inactivated or leading them to degradation [225]. So, the phosphorylation/inactivation of GSK-3 induces the derepression of pathways normally repressed, and this can be the strategy employed by PTHrP to indirectly promote cell survival and proliferation by triggering PI3-K/p-Akt pathway through integrin a6b4 upregulation. The a6b4 integrin works as a powerful activator of PI3-K in carcinoma cells [205, 211], but we cannot rule out the involvement of other survival pathways [192]. For example, Akt can be activated in a PI3-K-independent manner in the context of a pathway downstream to GPCRs, such as the PTH/PTHrP receptor PTH1R [226]. Moreover, upstream to PI3-K, a6b4 integrin can influence cell growth through the Ras-MAPK cascade [227, 228] and can also interact with some growth factor receptors, such as ErbB2 and Met [229, 230]. In particular, b4 subunit signaling is reported to promote cell proliferation via ErbB2, suggesting the possibility of a b4/ ErbB2 pathway for cancer cell growth and survival [231–233]. After all, the net effect of PTHrP-induced integrin a6b4 upregulation is a significant increase in cell proliferation, migration, invasion, and anchorage-independent cell

4

PTHrP Signaling in Cancer

73

growth, as reported by Shen et al. [187] in their work on LoVo cells. Moreover, in support of in vitro data, they described an increase of LoVo xenograft growth, with engrafted tumor cells overexpressing a6b4 subunits as well as PI3-K subunits (p85a regulatory and p110a catalytic) and p-Akt, plus total Akt. These experimental results highlight the pivotal role of the prosurvival and proinvasive integrin a6b4 in cancer progression [234], as confirmed by the experimental evidences that emerged from studies on the pharmacologic treatment of prostate cancer cell lines PC-3 and C4-2. Shen et al. [235] noted that the exposure to 1,25(OH)2D3 induced significant downregulation of PTHrP and integrin a6b4 expression in C4-2 cells. The decrease of integrin a6b4 expression followed the earlier decrease of PTHrP, supporting the mechanism based on the PTHrP-mediated control of integrin a6b4 expression. These effects resulted in cell proliferation reduction, as expected on the bases of the PTHrP/a6b4 control axis of cell proliferation described above. Moreover, recent data collected by Bhatia et al. demonstrated that the intracrine PTHrP signaling can protect C4-2 and PC-3 cells against the apoptosis induced by doxorubicin. This protective effect requires the a6b4-mediated activation of PI3-K/Akt pathway and results associated with enhanced level of c-Myc and increased ratio of antiapoptotic to proapoptotic members of Bcl-2 family. In this model, PTHrP is reported to increase the activity of nuclear factor kB as well as the anchorage-independent cell growth. The critical role played by integrin a6b4 in the induction of these survival/proliferation pathways is proved by the block of PTHrP-mediated activation of PI3-K/Akt pathway produced by a6b4 silencing [236]. Also a5b1 integrin results involved in an antiapoptotic pathway based on the selective activation of PI3-K [237] and probably PTHrP overexpression exerts its protective effect against apoptosis through the upregulation of a5 subunit [190].

PTHRP and Bone Metastases Based on literature data, there is a correlation between PTHrP expression by primary tumor and metastatic diffusion to bone tissue, and virtually all tumors that metastasize to bone are considered expressing PTHrP. Bone metastases are a frequent and serious complication of cancer and are associated with worsening of quality of life and considerable social costs. Bone metastases are often a “point of no return” in tumor clinical history because they are usually incurable and associated with devastating consequences, as severe bone pain, pathologic fractures, life-threatening hypercalcemia, and neural compression syndromes. The exact extent of skeletal metastatic involvement is not defined but, for example, it is estimated that in the United States, 350,000 patients affected by bone metastases die every year [238]. Bone metastases occur in up to 70% of patients with advanced cancer of breast or prostate [239], whereas in carcinoma of the lung, stomach, colon, rectum, uterus, kidney, bladder, or thyroid, the incidence is comprised from 15% to 30% approximately [240].

74

F.O. Ranelletti and G. Monego

Bone appears to be a preferred site of cancer metastasis, and this “predilection” is probably due to several structural and functional factors connected to bone microenvironment. First off, the high blood flow in red marrow areas can favor the localization of cancer cells in these sites [241]. In bone microenvironment, adhesive interactions can develop based on adhesive molecules, such as integrins, expressed by tumor cells. The adhesion of cancer cells to bone matrix and marrow stromal cells starts a bidirectional series of signals that gives rise to the development of a sort of fertile niche for metastatic cells. In fact, these adhesive interactions induce tumor cells to produce angiogenic factors and bone-resorbing factors that support tumor colony growth [242]. Moreover, bone resorption can mobilize and activate several growth factors previously embedded in bone matrix. The release of transforming growth factor b (TGF-b), insulin-like growth factors I and II (IGFs), fibroblast growth factors (FGFs), platelet-derived growth factors (PDGFs), bone morphogenic proteins (BMPs), and calcium can impact tumor growth, providing a suitable milieu for cancer cell proliferation [243, 244]. This is in accord with the “seed and soil” hypothesis suggested by Paget to describe the mechanism of bone metastasis [245]. However, when cancer cells metastasize to bone, a new altered microenvironment develops, and this causes the deregulation of bone remodeling. This dysregulated remodeling is a pathologic process that can range from osteolytic lesions to osteoblastic lesions, often showing both lesions coexisting in mixed forms of metastases. In fact, with the exception of the purely osteolytic lesions produced by multiple myeloma [246], it would be more correct to speak about predominantly osteolytic metastases as those generally associated with breast cancer or predominantly osteoblastic metastases as those in prostate cancer [247, 248]. Furthermore, the bone destruction caused by osteolytic metastasis is generally associated with an osteoblastic reaction finalized to bone neoformation, and so, there is also an increased component of osteosynthesis against the dominant osteolysis [249]. After all, the result is a poor quality of bone, which is exposed to the risk of pathologic fractures. In this picture, which role is played by the PTHrP/PTH1R system? In osteolytic metastases, the osteoclasts are liable of bone damage rather than cancer cells [250, 251]. However, tumor cells can act as pseudo-osteoblast by producing and releasing factors that can mediate the activation of osteoclasts: interleukin-1 (IL-1), interleukin-6 (IL-6), prostaglandin E2 (PGE2), tumor necrosis factor-a (TNFa), macrophage colony-stimulating factor (M-CSF), macrophage inflammatory protein-1a, and receptor activator of nuclear factor-kB ligand (RANKL) [240, 252]. Among tumoral osteoclastogenic factors, there is PTHrP, which is considered the primary stimulator of osteolytic destruction by bone metastases. PTHrP promotes bone resorption indirectly by binding PTH1R and inducing the expression of RANKL on osteoblastic and marrow stromal cells in bone microenvironment. The upregulated RANKL binds to RANK (receptor activator of nuclear factor-kB) on the surface of osteoclast precursors. RANKL signals can be transmitted downstream to RANK through the nuclear factor-kB and Jun N-terminal kinase pathways, inducing the maturation of osteoclasts and prolonging their survival by preventing apoptosis.

4

PTHrP Signaling in Cancer

75

Osteoclasts are the direct mediators of bone destruction through the release of acids and proteases at the level of the ruffled border of their plasma membrane, which is considered as a cellular resorbing organelle adhering to bone. As in the case of other bone-resorptive cytokines, the effect of PTHrP converges on preosteoclasts to promote the differentiation to mature and actively bone-resorbing osteoclasts. On the other hand, bone destruction results in release of IGFs, FGFs, BMPs, PDGF, and TGF-b from extracellular matrix. These growth factors in association with increased levels of extracellular calcium can feedback to cancer cells, stimulating proliferation and PTHrP production. Moreover, it has been reported that TGF-b is able to amplify PTHrP signaling by increasing the stability of PTHrP mRNA and prolonging its half-life [17]. So, a bidirectional interaction between cancer cells and bone microenvironment is set up. This reciprocal symbiotic interaction produces a vicious cycle that works as a support system for tumor localization and growth in bone, associated with progressive bone loss. Bone resorption is critical not only for osteolytic metastases development but also for tumor burden. In fact, experimental and clinical evidences suggest a close correlation between bone erosion and tumor growth since the pharmacologic block of bone destruction is associated with the decrease of tumor burden [253, 254]. Moreover, the inhibition of metastasization to skeleton can be associated with an improvement of clinical outcome [255]. The mechanism of osteolytic metastases from breast cancer is a paradigmatic example of the vicious circle structured around PTHrP expression by tumor cells. PTHrP is overexpressed in breast cancer cells that metastasize to bone, and it is not clear if this is due to an intrinsic functional property of tumor cell or is induced by the interaction with bone microenvironment. By the way, the metastasization to bone is associated with a marked increase in PTHrP expression, since only 50% of primary tumors produce the peptide, whereas 92% of bone metastases are verified to be PTHrP-expressing [256]. The production of PTHrP by bone metastatic cancer cells is upregulated by the increased expression of Runx2/CBFA1 (core-binding factor a1), a master transcription factor involved in the control of a cohort of genes required for bone development and turnover as well as for tumor growth and metastasis [173]. Breast cancer metastatic cells overexpress Runx2, and this activates Indian hedgehog (IHH) by binding to its promoter. In accord to physiologic sequence, the activation of IHH induces the downstream target gene PTHrP probably through Gli proteins, the mediators of hedgehog signaling. As mentioned before, PTHrP upregulation leads to RANKL expression and to osteoclastic activation, completing the bone-destructive mechanism operative in breast cancer cells. So, in breast cancer, the highly expressed Runx2 works through the physiological axis formed by the IHH–PTHrP pathway, and this supports the hypothesis that tumor cells in bone acquire osteomimicry property [257], gaining a growth boost by taking advantage of gene regulatory mechanisms active under physiologic conditions. Osteoclast-mediated resorption demineralizes bone matrix, increasing extracellular levels of calcium, which can promote cancer cell proliferation and PTHrP synthesis by signaling through CaSR (calcium-sensing receptor). The overproduced PTHrP can impact cancer proliferation by regulating the expression of its downstream

76

F.O. Ranelletti and G. Monego

target gene cyclin D1, a key cell cycle controller that can support tumor development [258]. Furthermore, metastatic cells actively synthesize several factors, such as M-CSF, IL-6, IL-1, TNF-a, and PGE2, which can contribute to osteoclast maturation and bone erosion. The osteolysis causes the release of growth factors and cytokines stored in mineralized extracellular matrix, resulting in a feedback support system for cancer growth. Among these growth factors mobilized from bone matrix, TGF-b can mediate an increase in PTHrP production via Smad and MAPK (mitogenactivated protein kinase) pathways. In this TGF-b-mediated induction of PTHrP, Runx2 plays an important role by directly activating IHH and stimulating the IHH downstream target gene PTHrP. This IHH-dependent induction of PTHrP requires a sort of activation complex that involves the synergistic interaction of Runx2 with coregulatory factors, such as Smad proteins and Ets proteins as well as the cooperation with hedgehog signaling molecules, such as Gli proteins, and the activation of MAPK signaling. Moreover, the signals of PTHrP and TGF-b converge on cyclin D1, which is an important target gene of both these factors for skeletal development [259, 260]. As mentioned above, cyclin D1 is a critical regulatory protein of cell cycle, and the signaling of PTHrP and TGF-b can contribute to the deregulation of cell proliferation by targeting cyclin D1 [258, 261]. This input to proliferation of cancer cells closes the vicious circle that links bone microenvironment to metastatic cells, improving progressively tumor burden and bone destruction. All these data support the concept that PTHrP plays a major role in osteolytic metastases from breast cancer and other solid tumors. Nevertheless, PTHrP, as well as PTH, can exert anabolic effects in bone through their common receptor. In detail, PTH inhibits collagen synthesis and osteoblast differentiation and, similarly to PTHrP, inhibits matrix mineralization probably through the matrix gla protein (MGP) [262, 263]. By contrast, PTHrP effect is more complex, depending on cell differentiation state. In fact, the peptide stops proliferation of confluent differentiated osteoblasts whereas increases the growth of osteoblastic cells proliferating under low-serum conditions by upregulating its target gene cyclin D1 [260, 264]. So, in bone microenvironment, the presence of different cellular pools at various differentiation stages makes the effect of PTHrP released by metastatic cells articulated. Moreover, posttranslational processing of PTHrP can produce multiple peptides characterized by different, and sometimes opposite, effects. The amino-terminal PTHrP is a potent enhancer of osteoclastogenesis through the binding with PTH1R, whereas the carboxyl-terminal is considered an inhibitor of osteoclast-mediated bone erosion and is defined osteostatin [265]. The complexity of PTHrP signaling accounts for the possibility that the interactions between cancer cells and bone microenvironment could give rise to a pathological structural rearrangement potentially oriented toward bone formation or toward bone destruction. In certain conditions, PTHrP could synergize with tumor-derived growth factors to support a predominant osteoblastic activity with osteoblast proliferation, whereas in other conditions, the peptide could work as an inhibitor of osteoblast development and as a powerful stimulator of osteoclastogenesis and bone

4

PTHrP Signaling in Cancer

77

catabolism by interacting cooperatively with matrix-derived factors and structuring the vicious circle of osteolytic metastasis. Osteoblastic metastasis is the opposite extreme of osteolytic metastasis in the pathologic bone remodeling process caused by cancer metastasization to skeleton. The histology of this type of metastasis is composed by increased newly formed woven bone with irregular structure of more abundant trabeculae and numerous osteoblasts. Osteoblastic lesions are rare and generally associated with cancer of the prostate. The mechanisms of osteoblastic metastases are still not definitively known, but some evidences suggest that a vicious circle could be involved also in this case. The vicious sequence could be based on an osteoblast-inducing activity performed by tumor cells followed by the production and the release by osteoblasts of growth factors that can improve tumor growth. The growth factors involved in the development of osteoblastic lesions are endothelin-1 and PDGF [266, 267]. The cancer of the prostate spreads metastases to the skeleton, and these lesions are predominantly osteoblastic. Prostate cancer cells release endothelin-1, PDGF, urokinase and urokinase-type plasminogen activator (u-PA), and prostate-specific antigen (PSA). Increased production of u-PA by prostate cancer cells results in an increase of bone metastases [268]. PSA is a kallikrein serine protease produced by prostate cancer cells. The release of PSA by tumor cells can indirectly impact on bone remodeling. In fact, PSA can cleave PTHrP at the amino-terminal part of peptide, cutting the site of binding to PTH1R and stopping the induction of bone resorption. PTHrP shows intense positive immunostaining in bone metastatic lesions and colocalizes with PSA. PSA can also take part in the activation of insulin-like growth factors I and II, and TGF-b. The release of these osteoblastic growth factors in bone microenvironment during metastases development represents the possible completion of the vicious cycle hypothesized above. As mentioned before, osteoblastic and osteolytic processes coexist in the deregulated bone remodeling induced by cancer metastases. In fact, in bone metastases of prostate cancer, the osteoblastic lesions are associated with increase of bone resorption markers. Probably, osteoclast-mediated bone resorption is an integral part in the pathogenesis of osteoblastic bone metastases and can represent an initial event followed by massive osteoblast activation with consequent extensive bone formation [266]. Another emerging aspect of PTHrP produced by the bone metastasizing cancer cells is the potential impact of the peptide on hematopoietic red marrow and, in particular, on stem cell niche. The hypothesis is that PTHrP could support bidirectional communication between tumor cells and hematopoietic cells. In detail, PTHrP, like PTH, could act on the PTH1R on osteoblast, inducing the expression of Jagged 1. Jagged 1 is a ligand of Notch, which is expressed on hematopoietic cells and is considered critical to support many steps of hematopoietic stem cell differentiation. So, PTHrP could promote hemopoiesis through the Jagged 1/Notch pathway. The increases in hematopoietic cells could, in turn, favor neoangiogenesis contributing to

78

F.O. Ranelletti and G. Monego

a fertile environment for cancer cells in bone. Moreover, PTHrP and PTH revealed ability to regulate hemopoiesis-relevant genes such as M-CSF (macrophage colonystimulating factor), MCP-1 (monocyte chemotactic protein-1), and VCAM-1 (vascular cell adhesion molecule 1) [269–271].

References 1. Suva, L. J., Winslow, G. A., Wettenhall R. E., Hammonds, R. G., Moseley, J. M., DiefenbachJagger, H., Rodda, C. P., Kemp, B. E., Rodriguez, H., Chen, E.Y. 1987, Science, 237, 893. 2. Philbrick, W. M., Wysolmerski, J. J., Galbraith, S., Holt, E., Orloff, J.J., Yang, K. H., Vasavada, R. C., Weir, E. C,, Broadus, A. E., and Stewart, A. F. 1996, Physiol. Rev., 76, 127. 3. Pecherstorfer, M., Schilling, T., Blind, E., Zimmer-Roth, I., Baumgartner, G., Ziegler, R., and Raue F. 1994, J. Clin. Endocrinol. Metab. 78, 1268. 4. Dittmer, J. 2004, Gene Ther. Mol. Biol., 8, 451. 5. Luparello, C., Romanotto, R., Tipa, A., Sirchia, R., Olmo, N., Lopez de Silanes, I., Turnay, J., Lizarbe, A., and Stewart, A. F. 2001, J. Bone Miner. Res., 16, 2173. 6. Dougherty, K. M., Blomme, E. A., Koh, A. J., Henderson, J. E., Pienta, K. J., Rosol, T..J., and Mc Cauley, L. K. 1999, Cancer Res., 59, 6015. 7. Hastings, R. H., Burton, D. W., Quintana, R. A., Biederman, E., Gujral, A., and Deftos, L. J. 2001, Cancer, 92, 1402. 8. Massfelder, T., Lang, H., Schordan, E., Lindner, V., Rothhut, S., Welsch, S., Simon-Assmann, P., Barthelmebs, M., Jacqmin, D., and Helwig, J. J. 2004, Cancer Res., 64, 180. 9. Comings, D. 1972, Nature, 238, 455. 10. Mangin, M., Ikeda, K., Dreyer, B.E., Milstone, L., and Broadus, A.E. 1988, Mol Endocrinol., 2, 1049. 11. Mangin, M., Ikeda, K., Dreyer, B.E., and Broadus, A.E. 1989, Proc. Nat. Acad. Sci. U.S.A., 86, 2408. 12. Suva, L.J., Mather, K.A., Gillespie, M.T., Webb, G.C., Ng, K.W., and Winslow, G.A. 1989, Gene, 1989; 77, 95. 13. Yasuda, T., Banville, D., Hendy, G.N., and Goltzman, D. 1993, Mol. Endocrinol., 7, 273. 14. Vasavada, R. C., Wysolmerski, J. J., Broadus, A. E., and Philbrick, W. M. 1993, Mol. Endocrinol., 7, 273. 15. Mangin, M., Ikeda, K., Dreyer, B.E., and Broadus, A.E. 1990, 4, 851. 16. Campos, R.V., Wang, C., and Drucker, D.J. 1992, Mol. Endocrinol., 6,1642. 17. Sellers, R.S., Luchin, A.I., Richard, Brena, R.M., Lima, D. and Rosol, T.J. 2004, J. Mol. Endocrinol., 33, 227. 18. Southby, J., O’Keeffe, L.M., Martin, T.J., and Gillespie, M.T. 1995, Br. J. Cancer, 72, 702. 19. Yasuda, T., Banville, D., Hendy, G. N., and Goltzman, D. 1989, J. Biol. Chem., 264, 7720. 20. Gillespie, M.T., and Martin, T.J. 1994, Mol. Cell. Endocrinol., 100, 143. 21. Ikeda, K., Arnold, A., Mangin, M., Kinder, B., Vydelingum, N.A., Brennan, M.F., and Broadus, A.E. 1989, J. Clin. Endocrinol. Metab., 69, 1240. 22. Ikeda, K., Lu, C., Weir, E.C., Mangin, M. and Broadus, A.E. 1989, J. Biol. Chem., 264, 15743. 23. Shaw, G., and Kamen, R. 1986, Cell, 46, 659. 24. Martin, T.J., Moseley, J.M., and Gillespie, M.T. 1991, Crit. Rev. Biochem. Mol. Biol., 26, 377. 25. Power, D.M., Ingleton, P.M., Flanagan, J., Canario, A.V., Danks, J., Elgar, G., and Clark, M.S. 2000, Gene, 250, 67. 26. Ikeda, K., Lu, C., Weir, E.C., Mangin, M., and Broadus, A.E. 1990, J. Biol. Chem., 265, 5398. 27. Sellers, R.S., LeRoy, B.E., Blomme, E.A., Tannehill-Gregg, S., Corn, S., and Rosol, T.J. 2004, Prostate, 58, 366. 28. Wilson, G.M., Sutphen, K., Chuang, K., and Brewer, G. 2001, J. Biol. Chem., 276, 8695.

4

PTHrP Signaling in Cancer

79

29. Chen, C.Y., Gherzi, R., Ong,S.E., Chan, E.L., Raijmakers, R., Pruijn, G.J., Stoecklin, G., Moroni, C., Mann, M., and Karin, M. 2001, Cell, 107, 451. 30. Ikeda, K., Okazaki, R., Inoue, D., Ogata, E., and Matsumoto, T. 1993, J. Biol. Chem., 268, 1174. 31. Richard, V., Rosol, T.J., and Foley, J. 2005, Crit. Rev. Eu. Gene Exp., 15, 115. 32. Southby, J., Murphy, L.M., Martin, T.J., and Gillespie, M.T. 1996, Endocrinology, 137, 1349. 33. Oikawa, T., and Yamada, T. 2003, Gene, 303, 11. 34. Dittmer, J., Gitlin, S.D., Reid, R.L., and Brady, J.N. 1993, J. Virol., 67, 6087. 35. Dittmer, J., Gegonne, A., Gitlin, S.D., Ghysdael, J., and Brady J.N. 1994, J. Biol. Chem. 269, 21428. 36. Dittmer, J., Pise-Masison, C.A., Clemens, K.E., Choi, K.S., and Brady, J.N. 1997, J. Biol. Chem. 272, 4953. 37. Levine, M., and Tjian, R. 2003, Nature, 424, 147. 38. Lindemann, R.K., Ballschmieter, P., Nordheim, A., and Dittmer, J. 2001, J. Biol. Chem., 276, 46661. 39. Lindemann, R.K., Braig, M., Ballschmieter, P., Guise, T.A., Nordheim, A., and Dittmer, J. 2003. Int. J. Oncol., 22, 799. 40. Lindemann, R.K., Braig, M., Hauser, C.A., Nordheim, A., and Dittmer, J. 2003, Biochem. J., 372, 787. 41. Wasylyk, B., Hagman, J., and Gutierrez-Hartmann, A. 1998. Trends Biochem. Sci., 23, 213. 42. Aklilu, F., Gladu, J., Goltzman, D., and Rabbani, S.A. 2000, Cancer Res., 60, 1753. 43. Cho, Y.M., Lewis, D.A., Koltz, P.F., Richard, V., Gocken, T.A., Rosol, T.J. Konger, R.L., Spandau, D.F., and Foley, J. 2004, J. Endocrinol., 181, 179. 44. Arbabi, S., and Maier, R. 2002, Crit. Care Med., Suppl 1: S74. 45. MacLeod, R.J,, Chattopadhyay, N., and Brown, E.M. 2003, Am. J. Physiol. Endocrinol. Metab., 284, E435. 46. Tfelt-Hansen, J., MacLeod, R.J., Chattopadhyay, N., Yano, S., Quinn, S., Ren, X. Terwilliger, E.F., Schwarz, P., and Brown, E.M. 2003, Am. J. Physiol. Endocrinol. Metab., 285, E329. 47. MacLeod, R.J., Yano, S., Chattopadhyay, N., and Brown, E.M. 2004, Biochem. Biophys. Res. Commun. 320, 455. 48. Kakonen, S.M., Selander, K.S., Chirgwin, J.M., Yin, J.J., Burns, S., Rankin, W.A., Grubbs, B.G., Dallas, M., Cui, Y., and Guise, T.A. 2002, J. Biol. Chem., 277, 24571. 49. Lindeman, R.K., Nordheim, A., and Dittmer, J. 2003, Mol. Cancer, 2, 20. 50. Yang, C., Shapiro, L.H., Rivera, M., Kumar, A., and Brindle, P.K. 1998, Mol. Cell. Biol., 18, 2218. 51. Chilco, P.J., Leopold, V., and Zajac, J.D. 1998, Mol. Cell. Endocrinol., 138, 173. 52. Wysolmerski, J.J., Vasavada, R., Foley, J., Weir, E.C., Burtis, W.J., Kukreja, S.C., Guise, T.A., Broadus, A.E., and Philbrick, W.M. 1996, Cancer Res., 56, 1043. 53. Quandt, K., Frech, K., Karas, H., Wigender, E., and Werner, T. 1995, Nucleic Acid. Res. 23, 4878. 54. Nishishita, T., Okazaki, T., Ishikawa, T., Igarashi, T., Hata, K., Ogata, E., and Fujita, T. 1998, J. Biol. Chem. 273, 10901. 55. Okazaki, T., Nishimori, S., Ogata, E., and Fujita, T. 2003, Biochem. Biophys. Res. Commun. 2003, 304, 632. 56. Holt, E.H., Vasavada, R.C., Bander, N.H., Broadus, A.E., and Philbrick, W.M. 1993, J. Biol. Chem., 268, 20639. 57. Ganderton, R.H., Day, I.N., and Briggs, R.S. 1995, Eur. J. Cancer, 31A, 1697. 58. Ganderton, R.H., and Briggs, R.S. 1997, Biochem. Biophys. Acta., 1352, 303. 59. Ganderton, R.H., and Briggs, R.S. 2000, Eur. J. Cancer., 36, 2128. 60. Liu, B., Goltzman, D., and Rabbani, S.A. 1993, Endocrinology,132, 1658. 61. Asadi, F., Faraj, M., Malakouti, S., and Kukreja, S.C. 2001, Int. Urol. Nephrol., 33, 47. 62. Merryman, J.I., Capen, C.C., McCauley, L.K., Werkmeister, J.R., Suter, M.M., and Rosol, T.J. 1993, Lab. Invest., 69, 347. 63. Glatz, J.A., Heath, J.K., Southby, J., O’Keeffe, L.M., Kiriyama, T., Moseley, J.M., Martin, T.J., and Gillespie, M.T. 1994, Mol. Cell. Endocrinol., 101, 295. 64. Kasono, K., Isozaki, O., Sato, K., Sato, Y., Shizume, K., Ohsumi, K., and Demura, H. 1991, Jpn. J. Cancer Res., 82, 1008.

80

F.O. Ranelletti and G. Monego

65. Schilling, T., Pecherstorfer, M., Blind, E., Kohl, B., Wagner, H., Ziegler, R., and Raue, F. 1996, Bone, 18, 315. 66. Cohen-Solal, M. E., Bouizar, Z., Denne, M. A., Graulet, A. M., Gueris, J., Bracq, S., Jullienne, A., and de Vernejoul, M. C. 1995, Horm. Metab. Res., 27, 403. 67. Wysolmerski, J .J., Dann, P.R., Zelazny, E., Dunbar, M. E., Insogna, K.L., Guise, T.A., and Perkins, A. S. 2002, J. Bone Miner. Res., 17, 1164. 68. Manenti, G., Peissel, B., Gariboldi, M., Falvella, F. S., Zaffaroni, D., Allaria, B., Pazzaglia, S., Rebessi, S., Covelli, V., Saran, A., and Dragani, T.A. 2000, Oncogene, 19, 5324. 69. Akino, K., Ohtsuru, A., Yano, H., Ozeki, S., Namba, H., Nakashima, M., Ito, M., Matsumoto, T. and Yamashita, S. 1996, Cancer Res., 56, 77. 70. Rabbani, S.A., Gladu, J., Liu, B., and Goltzman, D. 1995, Endocrinology, 136, 5416. 71. Guise, T. A., Yin, J. J., Taylor, S. D., Kumagai, Y., Dallas, M., Boyce, B. F., Yoneda, T, and Mundy, G. R. 1996, J. Clin. Invest., 98, 1544. 72. Orloff, J. J., Reddy, D., de Papp, A.E.,Yang, K., Soifer, N.E., Stewart, A.F. 1994, Endocrine Rev., 15, 40. 73. Plawner, L. L., Philbrick, W. M., Burtis, W. J., Broadus, A. E., and Stewart, A. F. 1995, J. Biol. Chem., 270, 14078. 74. Wu, T. L., Vasavada, R. C., Yang, K., Massfelder, T., Ganz, M., Abbas, S. K., Care, A. D., and Stewart, A. F. 1996, J. Biol. Chem., 271, 24371. 75. Deftos, L.J., Burton, D.W., Hastings, R.H., Terkeltaub, R., and Hook, V.Y. 2001, Endocrine, 15, 217. 76. Hook, V.Y., Burton, D., Yasothornsrikul, S., Hastings, R. H., and Deftos, L. J. 2001, Biochem. Biophys. Res. Commun., 285, 932. 77. Gensure, R. C., Gardella, T. J., and Juppner, H. 2005, Biochem. Biophys. Res. Commun., 328, 666. 78. Gagnon, L., Jouishomme, H., Whitfield, J. F., Durkin, J. P., MacLean, S., Neugebauer, W., Willick, G., Rixon, R. H., and Chakravarthy, B. 1993, J. Bone Miner. Res. 8: 497–503. 79. Abou-Samra, A. B., Juppner, H., Force, T., Freeman, M. W., Kong, X. F., Schipani, E., Urena, P., Richards, J., Bonventre, J. V., Potts, J. T. Jr., Kronenberg, H. M., and Segre, J. V. 1992, Proc. Natl. Acad. Sci. USA, 89, 2732. 80. Bringhurst, F.R., Juppner, H., Guo, J., Urena, P., Potts, J.T. Jr., Kronenberg, H.M., AbouSamra, A.B. and Segre, G.V. 1993, Endocrinology, 132, 2090. 81. Matthew, J., Mahon, M. J., Donowitz, M., Yun, C. C., and Segre, G. V. 2002, Nature 417, 858. 82. Tawfeek, H.A., Che, J., Qian, F., and Abou-Samra, A. B. 2001, 281, E545. 83. Ferguson, G. S. S. 2001, Pharmacol. Rev. 53, 1. 84. Luttrell, L.M. 2005, J Mol Neurosci., 26, 253. 85. Gesty-Palmer, D., Chen, M., Reiter, E., Ahn, S., Nelson, C. D., Wang, S., Eckhardt, A. E., Cowan, C. L., Spurney, R. F., Luttrell, L. M., and Lefkowitz, R. J. 2006, J. Biol. Chem., 281, 10856. 86. Azzi, M., Charest, P. G., Angers, S., Rousseau, G., Kohout, T., Bouvier, M., and Pineiro, G. 2003, Proc. Natl. Acad. Sci. USA, 100, 11406. 87. Pierce, K. L., Luttrell, L. M., and Lefkowitz, R. J. 2001, Oncogene, 20, 1532. 88. Tohgo, A., Pierce, K. L., Choy, E. W., Lefkowitz, R. J., and Luttrell, L. M. 2002, J. Biol. Chem., 277, 9429. 89. Watson, P. H., and Pickard, B. W. 2008, Crit. Rev. Eukaryot. Gene Expr., 18, 151. 90. Watson, P. H., Fraher, L. J., Natale, B. V., Kisiel, M., Hendy, G. N., and Hodsman, A. B. 2000, Bone, 26, 221. 91. Pickard, B. W., Hodsman, A. B., Fraher, L. J., and Watson, P. H. 2006, 147, 3326. 92. Watson, P. H., Fraher, L. J., Hendy, G. N., Chung, U. I., Kisiel, M., Natale, B. V., and Hodsman, A. B. 2000, J. Bone Miner. Res., 15, 1033. 93. Pickard, B. W., Hodsman, A. B., Fraher, L. J., and Watson, P. H. 2007, Endocrinology, 148, 2282. 94. Yang, R., Hoang, B. H., Kubo, T., Kawano, H., Chou, A., Sowers, R., Huvos, A. G., Meyers, P. A., Healey, J. H., and Gorlick, R. 2007, Int. J. Cancer, 121, 943.

4

PTHrP Signaling in Cancer

81

95. Linforth, R., Anderson, N., Hoey, R., Nolan, T., Downey, S., Brady, G., Ashcroft, L., and Bundred, N. 2002, Clin. Cancer. Res., 8, 3172. 96. Nguyen, M., He, B., and Karaplis, A. 2001, Endocrinology, 142, 694. 97. Massfelder, T., Dann, P., Wu, T. L., Vasavada, R., Helwig, J. J., and Stewart, A. F. 1997, Proc. Natl. Acad. Sci. U. S. A., 94, 13630. 98. Lam, M. H., Briggs, L. J., Hu, W., Martin, T. J., Gillespie, M. T., and Jans, D. A. 1999, J. Biol. Chem., 274, 7391. 99. Meerovitch, K., Wing, S., and Goltzman, D. 1998, J. Biol. Chem., 273, 21025. 100. Aarts, M.M., Rix, A., Guo, J., Bringhurst, R., and Henderson, J.E. 1999, J. Bone Miner. Res., 14, 1493. 101. Fiaschi-Taesch, N. M., Stewart, A. F. 2003, Endocrinology, 144, 407. 102. Lam, M. H., Thomas, R.J., Loveland, K. L., Schilders, S., Gu, M., Martin, T. J., Gillespie. M. T., and Jans, D.A. 2002, Mol. Endocrinol., 16, 390. 103. Lam, M. H., Henderson, B., Gillespie, M. T., and Jans, D. A. 2001, Traffic, 2, 812. 104. Lam, M. H., Olsen, S. L., Rankin, W. A., Ho, P. W., Martin, T. J., Gillespie, M. T., and Moseley, J. M. 1997, J. Cell. Physiol., 173, 433. 105. Luparello, C., Sirchia, R., and Lo Sasso, B. 2008, Breast Cancer Res. Treat., 111, 461. 106. Fortino, V., Torricelli, C., Gardi, C., Valacchi, G., Rossi Paccani, S., and Maioli, E. 2002, Cell.Mol. Life Sci., 59, 2165. 107. Hastings, R.H. 2004, Respiratory Physiology & Neurobiology, 142, 95. 108. Erhardt, P., Troppmair, J., Rapp, U. R., and Cooper, G. M. 1995, Mol. Cell. Biol., 15, 5524. 109. Tovar Sepulveda, V. A., and Falzon, M. 2002, Regul. Pept., 105, 109. 110. Gentili, C., Morelli, S., Boland, R., and de Boland, A. R. 2001, Biochim. Biophys. Acta, 1540, 201. 111. Falzon, M., and Du, P. 2000, Endocrinology, 141, 1882. 112. Maioli, E., and Fortino, V. Br. J. Cancer, 90, 1293. 113. Goldsmith, Z. G., and Dhanasekaran, D. N. 2007, Oncogene, 26, 3122. 114. Dhillon, A.S., Hagan, S., Rath, O., and Kolch, W. 2007, Oncogene, 26, 3279. 115. Iacovelli, L., Capobianco, L., Salvatore, L., Sallese, M., D’Ancona, G. M., and De Blasi, A. 2001, Mol. Pharmacol., 60, 924. 116. Busca, R., Abbe, P., Mantoux, F., EMBO J., 19, 2900. 117. Cook, S.J., and McCormick, F. 1993, Science, 262, 1069.. 118. Graves, L.M., Bornfeldt, K. E., Raines, E. W., Potts, B. C., Macdonald, S. G., Ross, R., and Krebs, E. G. 1993, Proc. Natl. Acad. Sci. U.S.A., 90, 10300. 119. Wu, J., Dent, P., Jelinek, T., Wolfman, A., Weber, M. J., and Sturgill, T. W. 1993, Science, 262,1065. 120. Wan, Y., and Huang, X, Y. 1998, J. Biol. Chem., 273, 14533. 121. Zheng, M., Zhang, S. J., Zhu, W.Z., Ziman, B., Kobilka, B. K., and Xiao, R. P. 2000, J. Biol. Chem., 275, 40635. 122. Laroche-Joubert, N., Marsy, S., Michelet, S., Imbert-Teboul, M., and Doucet, A. 2002, J. Biol. Chem., 277, 18598. 123. Stork, P. J., and Schmitt, J. M. 2002, Trends Cell. Biol., 12, 258. 124. Dumaz, N., and Marais, R. 2005, FEBS J., 272, 3491. 125. Pearson, G. W., Earnest, S., and Cobb, MH. 2006, Mol. Cell. Biol., 26, 3039. 126. Vossler, M. R., Yao, H., York, R. D., Pan, M. G., Rim, C. S., and Stork, P. J. 1997, Cell, 89, 73. 127. Weissman, J. T., Ma, J. N., Essex, A., Gao, Y., and Burstein, E. S. 2002, Oncogene, 23, 241. 128. Wang, Z., Dillon, T. J., Pokala, V., Mishra, S., Labudda, K., Hunter, B., and Stork, P. J. 2006, Mol. Cell. Biol., 26, 2130. 129. de Rooij, J., Zwartkruis, F.J., Verheijen, M.H., Cool, R.H., Nijman, S.M., Wittinghofer, A., and Bos, J.L. 1998, Nature, 396, 474. 130. Quilliam, L. A., Rebhun, J. F., and Castro, A. F. 2002; Prog. Nucleic Acid Res. Mol. Biol., 71, 391. 131. Bos, J.L. 2006, Trends Biochem. Sci., 31, 680. 132. Schmitt, J. M., and Stork, P. J. 2000, J. Biol. Chem., 275, 25342.

82

F.O. Ranelletti and G. Monego

133. Obara, Y., Labudda, K., Dillon, T. J., and Stork, P. J. 2004, J. Cell Sci., 117, 6085. 134. Norum, J. H., Methi, T., Mattingly, R. R., and Levy, FO. 2005, FEBS J., 272, 2304. 135. Norum, J. H., Dawood, H., Mattingly, R. R, Sandnes, D., and Levy, F. O. 2007, FEBS Lett., 581, 15. 136. Amsen, E.M., Pham, N., Pak, Y., and Rotin, D. 2006, J. Biol. Chem., 281, 121. 137. Sidovar, M.F., Kozlowski, P., Lee, J. W., Collins, M. A., He, Y., and Graves, L. M. 2000, J. Biol. Chem., 275, 28688. 138. Dhillon, A.S., Pollock, C., Steen, H., Shaw, P.E., Mischak, H., and Kolch, W. 2002, Mol. Cell. Biol., 22, 3237. 139. Dumaz, N., Light, Y., and Marais, R. 2002, Mol. Cell. Biol., 22, 3717. 140. Dumaz, N., Marais, R. 2003, J. Biol. Chem., 278, 29819. 141. Tazawa, H., Takahashia, S., and Zilliacus, J. 2003; Biochim. Biophys. Acta, 1620, 32. 142. Schmitt, J. M., and Stork, P. J. 2001, Mol. Cell Biol., 21, 3671. 143. Schmitt, J. M., and Stork, P. J. 2002, Mol. Cell, 9, 85. 144. Bos, J.L. 1998, EMBO J., 17, 6776. 145. Ahn, Y.H., Jung, J.M., and Hong, S.H. 2005, Cancer Res., 65, 4896. 146. Ahn, Y.H., Han, J.H., and Hong, S.H. 2006, J Cell. Physiol., 209, 1039. 147. Simon, M. I., Strathmann, M. P., and Gautam, N. 1991, Science, 252, 802. 148. Hubbard, K. B., and Hepler, J. R. 2006, Cell. Signal., 18, 135. 149. Johnson, G. L., and Dhanasekaran, N. 1989, Endocr. Rev., 10, 317. 150. Camps, M., Carozzi, A., Schnabel, P., Scheer, A., Parker, P.J., and Gierschik, P. 1992, Nature, 360, 684. 151. Katz, A., Wu, D., and Simon, M. I. 1992, Nature, 360, 686. 152. Kolch, W., Heidecker, G., Kochs, G., Hummel, R., Vahidi, H., Mischak, H. Finkenzeller, G., Marmé, D., and Rapp, U. R. 1993, Nature, 364, 249. 153. Ueda, Y., Hirai, S., Osada, S., Suzuki, A., Mizuno, K., and Ohno, S. 1996, J. Biol. Chem., 271, 23512. 154. Schonwasser, D. C., Marais, R. M., Marshall, C. J., and Parker, P. J. 1998, Mol. Cell Biol., 18, 790. 155. Lev, S., Moreno, H., Martinez, R., Canoll, P., Peles, E., Musacchio, J. M., Plowman, G. D., Rudy, B., and Schlessinger J. 1995, Nature, 376, 737. 156. Dikic, I., Tokiwa, G., Lev, S., Courtneidge, S.A., and Schlessinger J. 1996, Nature, 383, 547. 157. Della Rocca, G.J., van Biesen, T., Daaka, Y., Luttrell, D.K., Luttrell, L.M., and Lefkowitz, R.J. 1997, J. Biol. Chem., 272, 19125. 158. Hawes, B. E., van Biesen, T., Koch, W. J., Luttrell, L. M., and Lefkowitz R. J. 1995, J. Biol. Chem., 270, 17148. 159. Guo, F. F., Kumahara, E., and Saffen, D. 2001, J. Biol. Chem., 276, 25568. 160. Blaukat. A,, Barac, A., Cross, M.J., Offermanns, S., and Dikic, I. 2000, Mol. Cell. Biol., 20, 6837. 161. Vucenik, I., Ramakrishna, G., Tantivejkul, K., Anderson, L., and Mand Ramljak, D. 2005, Breast Cancer Res. Treat., 91, 35. 162. Corbit, K. C., Trakul, N., Eves, E. M., Diaz, B., Marshall, M., and Rosner, M. R. 2003, J. Biol. Chem., 278, 13061. 163. Maioli, E., Torricelli, C., and Fortino, V. 2006, Cell. Mol. Life Sci., 63, 637. 164. Yamada, T., Tsuda, M., Ohba, Y., Kawaguchi, H., Totsuka, Y., and Shindoh, M. 2008, Biochem. Biophys. Res. Commun., 368, 575. 165. Sato, K., Yamakawa, Y., Shizume, K., Satoh, T., Nohtomi, K., Demura, H., Akatsu, T., Nagata, N., Kasahara, T., and Ohkawa, H. 1993, J. Bone Miner. Res., 8, 849. 166. Talon, I., Lindner, V., Sourbier, C., Schordan, E., Rothhut, S., Barthelmebs, M., Lang, H., Helwig J.J., and Massfelder, T. 2006, Carcinogenesis, 27, 73. 167. Gessi, M., Monego, M., Calviello, G., Lanza, P., Giangaspero, F., Silvestrini, A., Lauriola, L., and Ranelletti, F. O. 2007, Acta Neuropathol., 114, 135. 168. Kronenberg, H. M. 2003, Nature, 423, 332. 169. MacLean, H.E., Guo, J., Knight, M.C., Zhang, P., Cobrinik, D., and Kronenberg, HM. 2004, J. Clin. Invest., 113, 1343.

4

PTHrP Signaling in Cancer

83

170. Lai, L. P., and Mitchell, J. 2005, J. Cell. Biochem. 96: 1173. 171. Vortkamp, A., Lee, K., Lanske, B., Segre, G. V., Kronenberg, H. M., and Tabin, C. J. 1996, Science, 273, 613. 172. Sterling, J. A., Oyajobi, B. O., Grubbs, B., Padalecki, S. S., Munoz, S. A., Gupta, A., Story, B., Zhao, M., and Mundy, G. R. 2006, Cancer Res., 66, 7548. 173. Pratap, J., Wixted, J. J., Gaur, T., Zaidi, S. K., Dobson, J., Gokul, K.D., Hussain, S., van Wijnen, A. J., Stein, J. L., Stein, G. S., and Lian, J. B. 2008, Cancer Res., 68, 7795. 174. Alman, B. A., and Wunder, J. S. 2008, Ann. N.Y. Acad. Sci., 1144, 36. 175. Ogden, S.K., Casso, D. J., Ascano, M. Jr., Yore, M. M., Kornberg, T. B., Robbins, D. J. 2006, J. Biol. Chem., 281, 7237. 176. Hastings, R.H., Summers-Torres, D., Cheung, T.C., Ditmer, L.S., Burton, D.W., Petrin, E.M., Spragg, R.G., Li, J. and Deftos, L.J. 1996, Am. J. Physiol. Cell. Physiol. 270, L353. 177. Hastings, R.H., Araiza, F., Burton, D.W., Bedley, M. and Deftos, L.J. 2004, Am. J. Physiol. Cell. Physiol., 287, 1616. 178. Hastings, R.H., Araiza, F., Burton, D.W., Zhang, L., Bedley, M. and Deftos, L.J. 2003, Am. J. Physiol. Cell. Physiol., 285, C1429. 179. Gagiannis, S., Müller, M., Uhlemann, S., Koch, A., Melino, G., Krammer, P.H., Nawroth, P.P., Brune, M. and Schilling, T. 2009, Int. J. Cancer. 180. Hemann, M.T., Zilfou, J.T., Zhao, Z., Burgess, D.J., Hannon, G.J. and Lowe, S.W. 2004, Proc. Nat. Acad. Sci. U.S.A., 101, 9333. 181. Ivaska, J., and Heino, J. 2000, Cell. Mol. Life Sci., 57,16. 182. Lang, S.H., Clarke, N.W., George, N.J., Testa, N.G. 1997, Clin. Exp. Metastasis, 15: 218. 183. Juliano, R.L., and Varner, J.A. 1993, Curr. Opin Cell. Biol., 5, 812. 184. Sato, M., Narita, T., Kawakami-Kimura, N., Higashiyama, S., Taniguchi, N., Akiyama, S., Hashimoto, T., Manabe, T., Kannagi, R. 1996, Cancer Lett., 102, 183. 185. Fishman, D.A., Kearns, A., Chilukuri, K., Bafetti, L.M., O’Toole, E.A., Georgapoulos, J. et al. 1998, Invasion Metastasis. 1998, 18: 15. 186. Lundstrom, A., Holmbom, J., Lindqvist, C., and Nordstrom, T. 1998, Biochem. Biophys. Res. Commun., 250: 735. 187. Shen, X., Rychahou, P.G., Evers, B.M., and Falzon, M. 2007, Cancer Letters, 258, 241. 188. Ruoslahti, E. 1991, J Clin Invest., 87, 1. 189. Ruoslahti, E., Noble, N.A., Kagami, S. and Border, W.A. 1994, Kidney Int., 45 (Suppl.44), S17. 190. Shen, X., and Falzon, M. 2003, Regul. Peptides, 113, 17. 191. Shen, X., Qian, L., and Falzon, M. 2004, Exp. Cell Res., 294, 420. 192. Shen, X., and Falzon, M. 2006, Exp. Cell., Nov 15, 312 (19), 3822. 193. Anderson, J.A., Grabowska, A.M. and Watson, S.A. 2007, Br. J. Cancer, 96, 1394. 194. Mercurio, A.M. 1995, Trends Cell. Biol., 5, 419. 195. Shen, X. and Falzon, M. 2005, Regul. Pept., 125, 17. 196. Ye, Y., Seitz, P.K. and Cooper, C.W. 2001, Regul. Pept., 101, 19. 197. Hemler, M.E., Crouse, C., Sonnenberg, A. 1989, J. Biol. Chem., 264, 6529. 198. Dedhar, S., and Saulnier, R. 1990, J. Cell. Biol. 1990, 110, 481. 199. Ramos, D.M., Cheng, Y.F., and Kramer, R.H. 1991, Invasion Metastasis, 11, 125. 200. Rabinovitz, I., Nagle, R.B. and Cress, A.E. 1995, Clin. Exp. Metastasis,13, 481. 201. Shaw, L.M., Chao, C., Wewer, U.M., and Mercurio, A.M. 1996, Cancer Res., 56, 959. 202. Mercurio, A.M., and Rabinovitz, I. 2001, Semin. Cancer Biol., 11, 129. 203. Chao, C., Lotz, M.M., Clarke, A.C., Mercurio, A.M. 1996, Cancer Res., 56, 4811. 204. Shaw, L.M., Rabinovitz, I., Wang, H.H., Toker, A., and Mercurio, A.M. 1997, Cell, 91: 949. 205. Rabinovitz, I., and Mercurio, A.M. 1997, J. Cell. Biol., 139, 1873. 206. O’Connor, K.L., Shaw, L.M., and Mercurio, A.M. 1998, J. Cell. Biol. 1997, 139, 1749. 207. Rabinovitz, I., Toker, A., and Mercurio, A.M. 1999, J. Cell. Biol., 146, 1147. 208. Rabinovitz, I., Gibson, I.K., and Mercurio, A.M. 2001, Mol. Cell. Biol., 12, 4030. 209. Chung, J., Bachelder, R.E., Lipscomb, E.A., Shaw, L.M., and Mercurio, A.M. 2002, j. Cell. Biol., 158, 165.

84

F.O. Ranelletti and G. Monego

210. Lipscomb, E.A. and Mercurio, A.M. 2005, Cancer Metastasis Rev., 24, 413. 211. King, T.E., Pawar, S.C., Majuta, L., Sroka, I.C., Wynn, D., Demetriou, M.C., Nagle, R.B., Porreca, F., and Cress, A.E. 2008, Plos One, 3, e3535. 212. Vinturi, C.L., Alvarez-Silva, M., Trentin, A.G. and Borelli, P. 2000, Braz. J. Med. Biol. Res., 33, 889. 213. Fruman, D.A., Meyers, R.E. and Cantley, L.C. 1998, Annu. Rev. Biochem., 67, 481. 214. Vanhaesebroeck, B. and Waterfield, M.D. 1999, Exp Cell Res., 253, 239. 215. Semba, S, Itoh, M., Itoh, E.M., Youssef, M., Harado, W., Kimura, M. and Yamakawa, M. 2002, Cancer Res., 8, 3824. 216. Shao, J., Evers, B.M. and Sheng, H. 2004, Cancer Res., 64, 229. 217. Rychahou, P.G., Murillo, C.A. and Evers, B.M. 2005, Surgery, 138, 391. 218. Ikenoue, T., Kanai, Y., Hibika, Y., Kunai, F., Obata, T., Tanaka, Y., Imamura, J., Ohta, M., Jazag, A., Guleng, B., Tateishi, K., Asaoka, Y., Matsumara, M., Kawabe, T. and Omata, M. 2005, Cancer Res., 65, 4562. 219. Rychahou, P.G., Jackson, L.N., Silva, S.R., Rajaraman, B.M. and Evers, B.M. 2006, Ann. Surg., 243, 833. 220. Fresno Vara, J.A., Casado, E., de Castro, E.J., Cejas, P., Belda-Iniesta, C., and GonzalezBarón, M. 2002, Cancer Treat. Res., 30, 193. 221. Roy, H.K., Olusola, B.F., Clemens, D.L., Karolski, W.J., Ratashak, A., Lynch, H.T., and Smyrk, T.C. 2002, Carcinogenesis, 23, 201. 222. Khaleghpour, K., Li, Y., Banville, D., Yu, Z., and Shen, S.H. 2004, Carcinogenesis, 25, 241. 223. Itoh, N., Semba, S., Ito, M., Takeda, H., Kawata, S. and Yamakawa, M. 2002, Cancer, 94, 3127. 224. Shen, X., Mula, R.V.R., Evers, B.M. and Falzon, M. 2007, Reg. Peptides, 141, 61. 225. Cross, D.A., Alessi, D.R., Cohen, P., Andjelkovich, M. and Hemmings, B.A. 1995, Nature, 378, 785. 226. Sable, C.L., Filippa, N., Hemmings, B. and Van Obberghen, E. 1997, FEBS Lett., 409, 253. 227. Maniero, F., Pepe, A., Wary, K.K., Spinardi, L., Mohammadi, M., Schlessinger, J. and Giancotti, F.G. 1995, EMBO J., 14, 4470. 228. Maniero, F., Murgia, C., Wary, K.K., Curatola, A.M., Pepe, A., Blumemberg, M., Westwick, J.K., Der, C.J. and Giancotti, F.G. 1997, EMBO J., 16, 2365. 229. Falcioni, R., Antonini, A., Nistico, P., DiStefano, S., Crescenzi, M., Natali, P.G. and Sacchi, A. 1997, Exp. Cell. Res., 236, 76. 230. Trusolino, L., Bertotti, A. and Comoglio, P.M. 2001, Cell, 107, 643. 231. Hu, Y.P., Venkateswarlu, S., Sergina, N., Howell, G., St Clair, P., Humphrey, L.E., Li, W., Hauser, J., Zborowska, E., Willson, J.K.V. and Brattain, G. 2005, J. Biol. Chem., 280, 27383. 232. Zhou, Y. and Brattain, M.G. 2005, Cancer Res., 65, 5848. 233. Guo, W., Pylayeva, Y., Pepe, A., Yoshioka, T., Muller, W.J., Inghirami, G. and Giancotti, F.G. 2006, Cell, 126, 489. 234. Rabinovitz, I., and Mercurio, A.M. 1996, Biochem. Cell. Biol., 74, 811. 235. Shen, X., Mula, R.V.R., Li, J., Weigel, N.L. and Falzon, M. 2007, Steroids, 72, 14, 930. 236. Bhatia, V., Mula, R.V., Weigel, N.L. and Falzon, M. 2009, Mol. Cancer Res., 7, 1119. 237. Lee, J.W., and Juliano, R.L. 2000, Mol. Cell. Biol., 11, 1973. 238. Mundy, G.R. 2002, Nat. Rev. Cancer, 2, 584. 239. Coleman, R.E., and Rubens, R.D. 1987, Br. J. Cancer, 55, 61. 240. Roodman, G.D. 2004, Mechanisms of bone metastases, N. Engl. J. Med., 350: 1655. 241. Kahn, D., Weiner, G.J., Ben-Haim, S., Ponto, L.L., Madsen, M.T., Bushnell, D.L., Watkins, G.L., Argenyi, E.A., and Hichwa, R.D. 1994, Blood, 84: 3602. 242. Van der Pluijm, G., Sijmons, B., Vloedgraven, H., Deckers, M., Papapoulos, S. and Lowik, C. 2001, J. Bone Miner. Res. 16, 1077. 243. Hauschka, P.V., Mavrakos, A.E., Iafrati, M.D., Doleman, S.E., and Klagsbrun, M. 1986, J. Biol. Chem., 261, 12665. 244. Pfeilschifter, J., and Mundy, G.R. 1987, Proc. Nat. Acad. Sci. U.S.A., 84, 2024. 245. Paget, S. 1889, Lancet, 1, 571.

4

PTHrP Signaling in Cancer

85

246. Taube, T., Beneton, M.N., McCloskey, E.V., Rogers, S., Greaves, M., and Kanis, J.A. 1992, Eur. J. Haematol., 49, 192. 247. Charhon, S.A., Chapuy, M.C., Delvin, E.E., Valentin-Opran, A., Edouard, C.M., and Meunier, P.J. 1983, Cancer, 51, 918. 248. Coleman, R.E., and Seaman, J.J. 2001, Semin. Oncol., 28, Suppl 6, 11. 249. Schneider, A., Kalikin, L.M., Mattos, A.C., Keller, E.T., Allen, M.J., and Pienta, K.J. 2005, Endocrinology, 146, 1727. 250. Boyde, A., Maconnachie, E., Reid, S.A., Delling, G., and Mundy, G.R. 1986, Scan. Electron. Microsc., 4, 1537. 251. Taube, T., Elomaa, I., Blomqvist, C., Beneton, M.N.C., and Kanis, J.A. 1994, Bone, 90, 161. 252. Roodman, G.D. 2001, J. Clin. Oncol., 19, 3562. 253. Croucher, P,I,, Shipman, C.M., Lippitt, J., Perry, M., Asosingh, K., Hijzen, A., Brabbs, A.C., van Beek, E.J., Holen, I., Skerry, T.M., Dunstan, C.R., Russell, G.R., Van Camp, B., and Vanderkerken, K. 2001, Blood, 98, 3534. 254. Oyajobi, B.O., and Mundy, G.R. 2003, Cancer, 97, Suppl. 813. 255. Powles, T., Paterson, S., Kanis, J.A., McCloskey, E., Ashley, S., Tidy, A., Rosenqvist, K., Smith, I., Ottestad, L., Legault, S., Pajunen, M., Nevantaus, A., Männistö, E., Suovuori, A., Atula, S., Nevalainen, J., and Pylkkänen, L. 2002, J. Clin. Oncol., 20, 3219. 256. Powell, G.J., Southby, J., and Danks, J.A. 1991, Cancer Res., 51, 3059. 257. Logothetis, C.J., and Lin, S.H. 2005, Nat. Rev. Cancer, 5, 21. 258. Alao, J.P. 2007, Mol. Cancer, 6, 24. 259. Beier, F., Ali, Z., and Mok, D. 2001, Mol. Biol. Cell, 12, 3852. 260. Datta, N.S., Pettway, G.J., Chen, C., Koh, A.J., and McCauley, L.K. 2007, J. Bone Miner. Res., 22, 951. 261. Guo, J., Chung, U.I., Yang, D., Karsenty, G., Bringhurst, F.R., and Kronemberg, H.M. 2006, Dev. Biol., 292, 116. 262. Koh, A.J., Beecher, C.A., Rosol, T.J., and McCauley, L.K. 1999, Endocrinology, 140, 3154. 263. Gopalakrishnan, R., Ouyang, H., Somerman, M.J., McCauley, L.K., and Franceschi, R.T. 2001, Endocrinology, 142, 4379. 264. Datta, N.S., Chen, C., Berry, J.E., and McCauley, L.K. 2005, J. Bone Miner. Res., 20, 1051. 265. Fenton, A.J., Kemp, B.E., Kent, G.N., Moseley, J.M., Zheng, M.H., Rowe, D.J., Britto, J.M., Martin, T.J., and Nicholson, G.C. 1991, Endocrinology, 129, 1762. 266. Yi, B., Williams, P.J., Niewolna, M., Wang, Y., and Yoneda, T. 2002, Cancer Res., 62, 917. 267. Guise, T.A., Yin, J.J., and Mohammad, K.S. 2003, Cancer, 97, Suppl. 779. 268. Achbarou, A., Kaiser, S., Tremplay, G., Ste-Marie, L.G., Brodt, P., Goltzman, D., and Rabbani, S.A. 1994, Cancer Res., 54, 2372. 269. Weir, E.C., Philbrick, W.M., Amling, M., Neff, L.A., Baron, R., and Broadus, A.E. 1996, Proc. Nat. Acad. Sci. U.S.A., 93, 10240. 270. Li, X., and Qin, L. and Partridge, N.C. A.S.B.M.R. 27th Annual Meeting. 271. Berry, J.E., Datta, N., Pettway, G.J., and McCauley, L. 2006, J. Dent. Res. 2006. Int. Ass. Dent. Res. Presented in Brisbane, Australia.

Chapter 5

Signalling Molecules as Selective Targets for Therapeutic Strategies in Multiple Myeloma Francesco Piazza and Gianpietro Semenzato

Epidemiology and Aetiology Multiple myeloma (MM) comprises approximately 10% of all haematological malignancies with an annual incidence of four to five cases per 105. Over the last years, the apparent rise in incidence is best attributable to a refinement of the diagnostic techniques rather than to a de facto increase of the MM cases. After nonHodgkin’s lymphomas, MM represents the second most common haematological malignancy. MM is a disease that affects the elderly, as the median age of patients is ranging from 65 to 70 years. There is a slight male predominance, and black people are more prone to the disease, while Asian population is less affected. The etiologic factors driving MM onset are largely unknown. Genotoxic agents have been involved in a fraction of cases, like radiation exposure and environmental toxics (herbicides and insecticides, organic solvents). Familiarity has been described in clusters of families. Chronic inflammation has also been invoked as a causative condition, and experimental evidence has supported this hypothesis, even though no specific infectious or other types of agents have been identified.

Pathobiology of MM MM pathogenesis is a multistep process [1]. MM plasma cells (MM PCs) are characterised by a low proliferation index and a strict dependence on the bone marrow microenvironment for their survival and growth; they are post-germinal centre

G. Semenzato (*) Department of Clinical and Experimental Medicine, Haematology-Immunology Section and Venetian Institute of Molecular Medicine, Haematological Malignancies Unit, Padua University School of Medicine, Padua, 35128 Padova, Italy e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_5, © Springer Science+Business Media, LLC 2012

87

88

F. Piazza and G. Semenzato

lymphocytes, harbouring class-switched IgH genes and extensive hypermutations of their Ig genes [2]. Phenotypically, MM PCs are distinguished from normal plasma cells for the loss of expression of CD19 and the gain of expression of CD56. Cytogenetically, almost all MGUS and MM tumours are characterised by primary and secondary chromosomal translocations and aberrations [3]. The former occur very likely in the early steps of the disease, affect the immunoglobulin (IgH) locus at 14q and are believed to be caused by errors during the heavy-chain switch recombination or, more rarely, during the somatic hypermutation in the germinal centre reaction; primary chromosomal translocations are present in: 40–50% of MGUS and smouldering MM (SMM), 50–80% of intramedullary MM, 85% of primary plasma cell leukaemia (PCL) and more than 90% of MM cell lines [1, 4]. The IgH gene locus has been described to fuse with at least five different partners, in 40% of MGUS and MM: (1) MMSET and FGFR3 at 4p16, (2) CyclinD3 at 6p21, (3) Cyclin D1 at 11q13, (4) c-MAF at 16q23, (5) MAFB at 20q11. Late stage-associated secondary translocations affecting the IgH locus involve infrequent but recurrent partner loci. The cMYC gene, located at 8q24, can be a fusion partner of the Ig enhancer, but it can also be involved in other complex translocations (with an overall prevalence of 15%) [5]. Other types of aberrations in the number and structure of chromosomes have been widely documented in MM tumours, some of which bear prognostic significance, such as loss of chromosome 13/13q/13q14 sequences (in 50–60% of MM and 45% of MGUS) that have been associated with a poor outcome [6] or other prognostically negative cytogenetic alterations, such as gains at 1q21 [7, 8] and hypodiploidy/pseudodiploidy or subtetraploidy [9, 10]. Recent gene expression profile studies have also provided insights into the pathogenesis and prognosis of plasma cell dyscrasias [11]. For instance, the expression levels of cyclin D1, D2, D3 and the presence of non-random chromosomal translocations in MM can identify distinct prognostic classes [12–15]. Further molecular alterations observed in MM affect H-Ras p21, which is increased in approximately 70% of MM cases, and specific mutations have been shown to portend a poor prognosis: the tumour suppressor TP53, which displays point mutations in a variable percent of cases, and the tumour suppressor Rb-1, which is deleted in approximately 50% of cases. The tumoural niche of MM in its intra-medullary phase is represented by the bone marrow (BM). Physiologically, the terminally differentiated PCs that have completed the maturation process reside in the BM where they receive survival inputs and are committed to expand if the immunological context requires doing so. This same environment is modified by the malignant transformation of the PCs and gains specific and critical features. MM cell survives, adheres to stromal cellular and extracellular components, gains the ability to stimulate neo-angiogenesis, as well as interferes with osteoblast and osteoclast development and function in the BM milieu. Here, several signalling pathways play critical roles in these processes, and the dissection of the mechanisms that control them have provided the framework for the development of novel therapeutic approaches to MM [16].

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

89

The acquisition of escaping drug-induced apoptosis is acquired by MM cell contact with fibronectin in the extracellular matrix. This phenotype is called also cell adhesion-mediated drug resistance (CAM-DR) [17]. Furthermore, autocrine and paracrine loops that lead to the production of cytokines, such as IL-6, TNFa, IL-1b, IGF-I, TGFb and VEGF, are instrumental for regulating MM growth in the BM [18]. Myeloma bone disease is also a pathological consequence of the interaction of MM cells with the BM mesenchymal components: an imbalance between the generation of osteoblasts and new bone synthesis and the activation of bone-resorbing cells, the osteoclasts, is linked with alterations of cell-adherent and soluble mediators, such as the RANKL–RANK receptors, osteoprotegerin [19, 20], MIP1a and MIP1b [21], SDF1 [22] and, as recently shown, Wnt-signalling inhibitors DKK1 [23] and sFRP2 [24]. The crucial importance of the MM tumoural environment has pushed the research efforts to identify novel anti-myeloma therapies towards “microenvironment”targeting approaches associated with strategies that hit aberrant signalling pathways intrinsic of the malignant PCs. This concept now represents a crucial starting point for the development of novel biologically based therapies for this disease (Figs. 5.1 and 5.2).

Targeting Growth- and Survival-Promoting Signalling Cascades in MM PCs MM cells are influenced for their growth by intrinsic, genetically determined alterations as well as by external, paracrine or autocrine stimuli. Many signalling pathways have been described to be constitutively active as a consequence of mutations affecting growth-controlling genes. The triggering of growth-promoting signalling cascades often results from the deregulation of the function of pivotal molecules. For instance, cyclin Ds-mediated signalling is altered as a consequence of translocations; similarly, Ras, p53, c-myc, NF-kB are targeted by somatic mutations that cause an aberrant function conditioning MM cell growth.

The IL-6 Signalling System Interleukin-6 (IL-6) represents one of the most important growth factor for MM plasma cells. IL-6 engagement of its receptor leads to the activation of three signalling pathways that have a relevant importance in MM plasma cell biology. IL-6 receptor forms a hexameric membrane complex (formed by IL-6, IL-6Ralpha and gp130 subunits), and upon its autophosphorylation, the JAK family of cytoplasmic tyrosine kinases are recruited to the complex, forming a scaffolding platform able to gather the STAT family of transcription factors. JAKs phosphorylate STAT on tyrosine residues (Tyr 705 on STAT3). This phosphorylation causes intramolecular modifications

90

F. Piazza and G. Semenzato

myeloma plasmacells

Chemokine receptors

Growth factor receptors associated with tyrosine kinase activity growth factors IGF-I HGF CSF TGFb

chemotactic factors IGF-I SDF1 chemokines

TGFbR

fibronectin/integrins

collagen fibers extracellular matrix

stromal cells VCAM VLA4 VEGF-R

VEGF VEGF-R

adhesion immunomodulation

TNFa

lymphocytes, NK cells

myeloma plasmacells

endothelial cells monocytes

angiogenesis myeloma plasmacells

Immune cells dendritic cells

Fig. 5.1 The bone marrow microenvironment in multiple myeloma pathogenesis. Schematic representation of the bone marrow milieu components playing a role in MM pathogenesis. The bone marrow microenvironment is composed of the extracellular matrix (ECM) and of the bone marrow stromal cells (fibroblasts, adipocytes, dendritic cells). Myeloma plasmacells contact the ECM (fibronectin) through surface integrins, interact with stromal cells through adhesion molecules (i.e. VCAM, ICAM, VLA-4), and express chemokine receptors and tyrosine kinase receptors for chemotactic agents and growth factors. Stromal cells may secrete growth factors (i.e. IGF-I, IL-6, HGF, CSF), chemotactic agents (i.e. IGF-I, SDF-I, chemokines), molecules that stimulate angiogenesis (VEGF) and molecules that modulate the immune response (i.e. TNFα, TGFβ). VCAM vascular cell adhesion molecule; ICAM intercellular adhesion molecule; VLA-4 very late antigen-4; IGF-I insulin-like growth factor-I; IL-6 interleukin-6; HGF hepatocyte growth factor; CSF colony stimulating factor; SDF-1 stromal cell derived factor-1; VEGF vascular endothelial cell growth factor; VEGF-R vascular endothelial cell growth factor-receptor; TNFα tumor necrosis factor α; TGFβ transforming growth factor β

that lead STAT3 translocation into the nucleus and activation of STAT3 target genes. IL-6 engagement of its receptor also activates the MAPK pathway through upstream of Ras mediators Grb2 and SOS1 and downstream molecules RAF1 and MAPKKs. Finally, the IL-6R-induced signalling activates also another crucial survival pathway in MM cells, the PI3K/PKB cascade. IL-6 can be secreted by MM PCs spontaneously, or upon engagement of CD40 thus acting as an autocrine factor [25]. It can also be secreted under the stimulation of other cytokines present in the bone marrow microenvironment (i.e. TNFa, VEGF and IL-1); however, the major source of IL-6 paracrine secretion in the MM microenvironment are the bone marrow stromal cells (BMSCs). BMSC secrete IL-6 when in contact with

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

91

Fig. 5.2 Signalling cascades involved in myeloma plasma cell biology. Schematic representation of some signalling pathways whose role in MM plasma cell biology has been established and of the levels of their possible molecular targeting with innovative therapeutic approaches (in particular, inhibitors of tyrosine kinase receptors, MAPK inhibitors, NF-kB inhibitors and proteasome inhibitors). The different cytokines and growth factors are produced either in a paracrine or in a autocrine manner. IL-6 signals to the nucleus upon engagement of its membrane-associated receptor and the activation of downstream tyrosine kinases (Tyk2 and Jak) and transcription factors STAT3; IGF-I, by means of its cognate receptor, triggers the activation of the PI3K/AKT as well as the MAPK signalling modules, which lead to the transmission of transcriptional inputs through ERK and FKHR regulated gene expression; similarly, TGFb-TGFbR interaction activates the SMAD transcription factors through phosphorylation and subsequent nuclear translocation; the cytokine TNFa stimulates the activation of the NF-kB pathway and nuclear shuttling by activation both the TRAF/RIP and the MEKK signalling modules. Note that more than one pathway can be activated by the same stimulus and different stimuli can trigger the same pathway. For instance, IL-6 activates the JAK/STAT and the PI3K/AKT module, TGFb is able to stimulate the SMAD as well as the MAPK signalling routes; similarly, the NF-kB pathway may be the point of convergence of several distinct stimuli. IGF-I insulin-like growth factor-I; IGF-IR insulin-like growth factor-I receptor; IL-6 interleukin-6; IL-6R interleukin-6 receptor; TNFα tumour necrosis factor α; TNFαR tumour necrosis factor α receptor; TGFb transforming growth factor b; TGFbR transforming growth factor b receptor; Tyk2 tyrosine kinase 2; JAK Janus protein kinase; STAT3 signal transducer and activator of transcription 3; MEKK MAP-ERK kinase protein kinase; MAPK mitogen activated protein kinase; MEK MAP-ERK protein kinase; ERK extracellular signal-regulated protein kinase; PI3K phospoinositide-3’ kinase; AKT/PKB AKR thymoma/protein kinase B; FKHR forkhead rhabdomyosarcoma; GSK3 glycogen synthase kinase 3; SMADs SMA and MAD related proteins; TRAF TNF receptor-associated factor; receptor-interacting kinase; IKK IkB kinase; IkB inhibitor of kB

92

F. Piazza and G. Semenzato

MM PCs [26] or when exposed to TNFa [27], VEGF [28] or TGFb [29]. The major effects of IL-6 on MM plasma cells are the stimulation of proliferation [30], survival [31] and resistance to drugs [32]. Interestingly, the levels of IL-6 and its receptor in the serum of MM patients, which are paralleled by the serum levels of C-reactive protein, are prognostic markers, and this might be particularly relevant in light of recent work that has shown that in most MM cases, STAT3 is constitutively active [33]. Molecular strategies targeting IL-6 signalling have been investigated. Antibodies against IL-6 have been developed and tested already in MM. Tocilizumab, a humanised anti-IL-6 receptor mAb, is currently being studied in MM. A recent study demonstrated in a murine MM model the effectiveness of tocilizumab in vitro and in vivo [34]. Another chimeric monoclonal anti-IL-6 antibody, CNTO 328, has shown anti-myeloma efficacy in preclinical studies [35]. This molecule has undergone phase I clinical trials whose results are still to be published. Moreover, Sant7, an IL-6 superagonist with high affinity and blocking activity for IL-6R is another anti-IL-6 strategy under investigation in MM, both in preclinical setting [36].

IGF-I Signalling Insulin-like growth factor-I is a molecule whose role in cancer cell biology has become increasingly clear over the last years. IGF-I signals through the binding to its receptor-activating signalling pathways that promote cell proliferation and survival, adhesion and migration. IGF-I mostly activates the Ras/Raf/MEK/ERK and the PI3K/PKB pathways. IGF-I also activates the NF-kappaB signalling pathway, resulting in up-regulation of anti-apoptotic signalling molecules. Therefore, targeting the IGF-I-dependent signalling pathways represents a rational anti-cancer therapeutic approach but is troublesome because of achieving selectivity. NVP-ADW742 is a selective IGF-IR tyrosine kinase inhibitor that has shown significant anti-myeloma activity in vitro in MM cell lines and in mouse tumour xenograft models [37, 38]. The human monoclonal antibody IMC-A12 directed against IGF-IR blocks IGF-IRmediated signalling and showed a remarkable anti-cancer activity. IMC-A12 is being tested in phase I clinical trials in multiple myeloma patients [39]. Very recently, other IGF-IR inhibitors (picropodophyllin or PPP) have been tested in a murine MM model [38]. These compounds were able to lead to cell cycle arrest of MM cells. In vivo treatment of MM-bearing mice led to a reduction of tumour burden, serum paraprotein and bone marrow angiogenesis [40]. Altogether, these studies suggest that the IGF-I axis represents a potentially important therapeutic target for MM.

The NF-kB Pathway NF-kB transcription factors are important survival molecules for cancer cells [41, 42]. The NF-kB pathway is a highly inducible signalling system that is kept in an “off” state by the binding of NF-kB members to the inhibitor IkBa. In the presence

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

93

of a stimulus (which can be manifold, like GFs, DNA damage, various cellular stress), the cascade is “switched on” by the phosphorylation, ubiquitination and proteasome-mediated degradation of IkBa. NF-kB transcription factors then migrate in the nucleus and activate the transcription of target genes. In a fraction of MM PCs from patients and MM cell lines, NF-kB is constitutively active due to extrinsic stimulation or gene mutations affecting members of its signalling pathway [33, 43, 44]. NF-kB has been shown to protect MM cells against dexamethasone-induced apoptosis [45]. Moreover, NF-kB has been shown to be downstream of several signalling pathways that regulate MM cell growth and survival, such as the TNFa, IL-1b, PI3K/ AKT and Ras/MAPK cascades. The usefulness of targeting NF-kB in MM was established through the use of proteasome inhibitors [46]. However, since activation of NF-kB may occur through the activity of the IkB kinase IKK, specific inhibitors of this kinase have been evaluated as potential therapeutic drugs [47–49]. Moreover, other work has recently shown that substances displaying an inhibitory effect on IKK, such as curcumin (diferuloylmethane), cause down-regulation of NF-kB activity, MM plasma cell growth arrest and apoptosis [50].

The PI3K/AKT Pathway This signal transduction pathway is activated by several extracellular signals, including IL-6 and IGF-I, and it regulates many cellular processes such as proliferation, survival, cell adhesiveness, protein synthesis and others. Phosphatidylinositol 3,4,5-triphosphate (PIP3) is generated by PI3K at the plasma membrane and triggers the activation of several downstream protein kinases, of which AKT/protein kinase B (PKB) critically regulates cell survival, proliferation and oncogenesis. The activity of PI3K is modulated by PTEN, a dual specificity phosphatase able to inactivate PIP3 by dephosphorylation and an essential tumour suppressor mutated or altered in a wide variety of cancers [51]. The PI3K/AKT signalling pathway is critical for MM cell survival [52]. Thus, it has attracted the attention of investigators as a therapeutic target in this disease. An inhibitor of AKT, perifosine, is a synthetic alkylophospholipid able to inhibit the phosphorylation occurring at the membrane and AKT activation dose dependently. Perifosine has shown efficacy in inhibiting MM cell growth and also in collaborating with other agents, such as proteasome inhibitors in causing MM cell death [47, 53, 54]. These preclinical results prompted a phase I and II clinical trials in association with the novel agent lenalidomide in patients with relapsed/refractory MM and relapsed/resistant to bortezomib MM, demonstrating encouraging and remarkable clinical activity (ASH meeting, 2008).

Bcl-2 Protein Family The Bcl-2 family of proteins is critical in the regulation of intrinsic apoptosis and cell death. It includes both proapoptotic and antiapoptotic members, subdivided

94

F. Piazza and G. Semenzato

into three main classes, based in part on sharing homology within the so-called Bcl-2 homology (BH) 1–4 domains. Bcl-2, Bcl-xL, Mcl-1, A1 and Bcl-w are the antiapoptotic members. The proapoptotic Bax and Bak have BH 1–3 domains; however, they need an activation event to bind Bcl-2 and Bcl-xL. The so-called BH3-only proapoptotic members, such as Bid, Bad, Bim, Noxa, Puma, are proximal sensors that transmit signals of death downstream diverse stimuli [ 55, 56]. In MM, Bcl-2 function is altered and contributes to malignant plasma cell resistance to apoptosis [57]. Bcl-2 can be targeted by anti-sense oligonucleotides (oblimersen, G3139) and inhibitors of Bcl-2/Bcl-xL. Oblimersen underwent already phase II clinical trials in association with thalidomide and dexamethasone or phase III clinical trials with high-dose dexamethasone [58, 59], encouraging further experimentation with other agents, such as IMiDs and bortezomib. Moreover, a small molecule inhibitor that binds strongly with Bcl-2 and Bcl-xL, ABT-737, is able to impend the sequestration of proapoptotic molecules, therefore shifting the balance towards cell apoptosis. Preclinical studies have proven that ABT-737 induces MM cell apoptosis and shows synergy with other anti-myeloma agents [60–62]. A structurally similar molecule, ABT-263, displayed a better oral bioavailability profile than that of ABT-737. This molecule has been investigated in xenograft mouse models of different human tumours. The treatment of xenotransplanted mice with the OPM-2 MM cell line with ABT-263 in association with bortezomib displayed a greater tumourkilling efficacy than single treatments. One proposed mechanism of synergy is the simultaneous reduction of Mcl-1 (by bortezomib) and Bcl-2/Bcl-xL (by ABT-263) [62].

Farnesyltransferase Inhibitors (FTIs) The rationale of employing agents that inhibit farnesyltransferase (FTIs) in MM is based on the relatively high frequency of Ras GDP/GTP-binding GTPase mutations found in this cancer [63]. Moreover, growth factors such as IL-6 and IGF-I may also activate the Ras/Raf/MAPK pathway in MM cells [64]. The covalent addition of a 15–carbon isoprenoid chain (farnesyl group) to the Ras molecule allows Ras to be subsequently attached to the intracellular membranes. MM cells were found to be sensitive to treatment with FTIs R11577 (tipifarnib®, Zarnestra), which caused cell growth arrest [65, 66], perillic acid [67] and FTI-277, which was active also on chemoresistant cell lines [68]. In the clinical setting, R11577 was tested in a phase I trial in chronic myelogenous leukaemia, myelofibrosis and MM patients. The compound displayed clinical activity, and one MM patient had a reduction of the monoclonal protein of about 34% [69]. In a phase II trial, R11577 was administered to 43 advanced MM patients: 64% of patients had stable disease, which did not correlate with the in vivo inhibition of farnesylation. This study also demonstrated that tipifarnib may suppress oncogenic pathways (PI3K/AKT and STAT3) in vivo in MM cells [70]. Recent studies have demonstrated a synergistic

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

95

effect of tipifarnib and the proteasome inhibitor bortezomib in MM and acute myeloid leukaemia cells [71] and of another farnesyltransferase inhibitor, lonafarnib and bortezimib in inducing MM cell death [72]. Interestingly, a role for geranylgeranylated proteins in MM cell growth has been demonstrated in a recent study [73], providing the rationale for associating FTIs with geranylgeranyltransferase inhibitors [74].

TRAIL TRAIL/Apo2L (tumour necrosis factor-related apoptosis-inducing ligand/apoptosis ligand 2) is a member of the TNF superfamily that has the peculiar property of inducing apoptosis in cancer cells while sparing non-cancer cells. When TRAIL binds to its cognate receptors DR4 or DR5, it activates a death-inducing signalling complex (DISC) next to the intracellular domain of the receptor, which leads to the triggering of the extrinsic apoptotic pathway [75]. TRAIL was shown to induce apoptosis in MM cell lines and freshly isolated cells from patients [76] and to kill a panel of drug-sensitive and drug-resistant MM cell lines and to cause a considerable reduction of tumour burden in myelomabearing mice in a xenotransplant MM model [77]. More recent studies have investigated the mechanisms underlying TRAIL–induced apoptosis in MM and will help in designing better strategies to use this molecule in the therapy of this disease [78, 79].

Targeting Protein Maturation, Degradation and Acetylation Hsp90 Inhibitors Molecular chaperones permit intracellular trafficking, ternary and quaternary structure conformations and correct folding of cellular proteins. Heat shock proteins (HSPs) are molecular chaperones whose levels are increased under certain protein-denaturing stressors. This family of proteins has gained growing importance in cancer pathogenesis as they can affect the levels and the activity of many oncoproteins [80]. One of the most abundant HSPs in the cell is Hsp90. A number of natural and synthetic compounds have been shown to inhibit Hsp90. Geldanamycin (GA) is one of such natural compounds that acts by inhibiting the ATPase activity of Hsp90, and it has been tested on MM cells. GA was shown to cause a decrease in IGF-I and IL-6 receptor expression on MM cells; it reduced the levels of AKT, Raf and IKK, of antiapoptotic proteins FLIP, XIAP, cIAP and inhibited the NF-kB pathway [81]. Another Hsp90 inhibitor, analogue of GA, 17-AAG (17-(demethoxy)-17-allylamino geldanamycin, tanespimycin or KOS-953)

96

F. Piazza and G. Semenzato

has recently been shown to exert an anti-myeloma activity on drug-resistant MM cells and to inhibit the growth of MM cells in an in vivo murine model [71] and is currently being tested in phase II/III clinical trials in relapsed MM in association with bortezomib. Other newly developed HSP inhibitors are currently undergoing phase I clinical evaluation in MM. Retaspimycin (IPI-504) is a new HSP inhibitor derivative of geldanamycin and 17-AAG. It is highly soluble in water and well tolerated. Early phase I/II trials have demonstrated activity in NSCLC and gastrointestinal stromal tumour. Phase I/II trials are currently underway to evaluate the dosing schedules and activity of IPI-504 in relapsed/refractory MM (information available on www.clinicaltrials.gov).

Proteasome Inhibitors The ubiquitin-proteasome degradation pathway may control the turnover of proteins involved in cancer pathogenesis [82]. Ubiquitin is a short chain of about 80 amino acids, which serves as a signal that instructs to target proteins to which it is covalently attached for proteolysis by the 26S proteasome, a macromolecular protease complex [83, 84].

Bortezomib Bortezomib (Velcade®, Millennium, Cambridge, MA; formerly known as PS-341) is a boronic acid compound that binds to and inhibits the portion of the 26S proteasome that displays a chymotrypsin-like activity [85]. This compound was demonstrated to have in vitro and in vivo cytotoxic activity against several tumours, including MM [85–88], and one pilot clinical study in relapsed/refractory haematological malignancies showed a peculiar activity (in terms of response rates) in MM [89]. Three additional trials demonstrated the usefulness of bortezomib in MM. Two phase II studies (SUMMIT and CREST trials) showed significant clinical activity in patients with relapsed/refractory MM [90, 91]. A subsequent phase III study (the APEX trial) compared bortezomib with high-dose dexamethasone in relapsed MM patients, showing an improved clinical outcome in the bortezomib-treated group as compared to the high-dose dexamethasone-treated group [92]. Proteasome inhibitors may favour apoptosis in more than one way [85, 93– 95]. One of these is through inhibition of IkBa degradation and sequestration of inactive NF-kB transcription factors in the cytoplasm [46]. In addition to apoptosis, proteasome inhibitors may target other signalling pathways [85, 93, 94]. Prolonged exposure to proteasome inhibitors may lead to the development of resistance, and several compounds have been used with bortezomib to overcome this drawback, such as the camptothecin analogue CPT-11 [96], the antagonist of the mitochondrial peripheral benzodiazepine receptor PK11195 [97], TRAIL [98–100] and other compounds [101]. Intriguingly, novel PIs are currently under development [102, 103].

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

97

NPI-0052 This molecule (salinosporamide A) represents a second generation proteasome inhibitor derived from marine bacteria Salinospora [104]. NPI-0052 displayed remarkable activity against MM cells in vitro, even when grown in the protective BM microenvironment. NPI-0052 inhibits all the three protease activity of the proteasome, and it induces apoptosis principally through the FADD/caspase-8 pathway. An important feature of this molecule is its oral bioavailability [102]. Phase I clinical trials are ongoing in relapsed/refractory MM.

Carfilzomib (PR-171) Carfilzomib is another new irreversible proteasome inhibitor. It inhibits the chymotrypsinlike as well as the immunoproteasome activity. Carfilzomib can induce apoptosis of bortezomib-resistant MM cells [105]. This drug is currently being evaluated in clinical trials in haematological malignancies including MM.

HDAC Inhibitors This novel group of molecules has recently gained interest for its ability to inhibit a particular class of enzymes, i.e. HDACs (histone deacetylases), inducing apoptosis and differentiation in cancer cells. Histone acetylation and deacetylation is believed to be instrumental for the generation of a “histone code” responsible for the overall transcriptional outcome in a cell [106]. Among haematopoietic malignancies, mainly acute myeloid leukaemias and myelodysplastic syndromes have been associated with deregulation of these processes and have been tested for their sensitivity to the treatment with HDACi [107, 108]. The mechanisms through which HDACi cause their effects on cancer cells very likely rely on different methods of action besides those directly exerted on the HDAC [109]. Early reports showed that HDACi were able to increase the levels of cell-cycle inhibitors, and they induce apoptosis in MM cells [110]. More recently, suberoylanilide hydroxaminic acid (SAHA), a prototypic HDACi, was shown to cause apoptosis and several molecular changes in MM cells and a reduction of IL-6 production by bone marrow stromal cells [111, 112]. Another HDACi, depsipeptide, was demonstrated to induce growth arrest of IL-6-dependent MM cells and to decrease the levels of antiapoptotic proteins [113]. Moreover, the proteasome inhibitor bortezomib was demonstrated to synergize with HDACi (SAHA and sodium butyrate) in inducing apoptosis [114]. Another study has recently shown that by inhibiting a particular HDAC, HDAC6, with tubacin, the aggresome-dependent protein catabolism is impaired. This resulted in a synergistic cytotoxic effect with bortezomib on MM cells [115], suggesting that other cellular processes regulated by HDACs act as relevant therapeutic targets.

98

F. Piazza and G. Semenzato

Targeting the Myeloma Malignant Bone Marrow Microenvironment Immunomodulatory Therapies Thalidomide (N-phthalimido-glutarimide and lenalidomide) is a synthetic glutamic acid, derivative originally employed as a sedative and anti-emetic in the 1950s. Because of teratogenic effects, its use was at that time discontinued. Subsequently, in the 1960s, the discovery of anti-inflammatory and anti-angiogenic properties of thalidomide prompted to study its mode of action, and in the early 1990s, experimental data demonstrated that it could inhibit TNFa synthesis by monocytes [116] and other cellular sources [117] and could hamper basic fibroblast growth factor (bFGF)-induced angiogenesis [118]. The rationale for the use of thalidomide in MM is based on the observation that in MM, the bone marrow microvessel density is increased, and this negatively correlates with prognosis (reviewed in [115]). In the first report of the clinical use of thalidomide in relapsed/refractory MM patients, the drug was administered daily from 200 mg up to 800 mg orally for a median of 80 days [119]. A clinical benefit of thalidomide was demonstrated in 30–40% of patients, of which 10% obtained a complete or near complete remission and 25% a partial remission, as evaluated by >50% reduction in serum or urine monoclonal paraprotein levels [119]. Thalidomide was then tested as a single agent or in combination with dexamethasone and other drugs in several clinical trials that have demonstrated its clinical efficacy [120–122] and is currently being analysed in several ongoing clinical studies. One recently published study have assessed the role of thalidomide in association with HDT and ASCT [123], demonstrating that its incorporation into aggressive protocols may increase the ORR without affecting, however, the OS. Besides the teratogenic potential, which precludes its use in pregnant women, the most common adverse side effects observed with thalidomide include sedation (which indicates its assumption at bedtime), constipation, cutaneous manifestations (dry and itchy skin) and sensorimotor peripheral neuropathy, especially of the limbs. Rarer side effects are hepatotoxicity, hypothyroidism and neutropenia [122]. It has been proposed that thalidomide-mediated reduction of TNFa, bFGF and VEGF production might contribute to MM PC cytotoxicity, together with the increased activation of myeloma-directed T lymphocytes [124]. More active thalidomide-derived immunomodulatory drugs (IMiDs) have shown similar effects in inhibiting MM PC growth. The mechanisms of these anti-myeloma effects include induction of apoptosis, inhibition of the MAPK pathway, inhibition of NF-kB activation and blockage of VEGF-induced cell migration [125]. Lenalidomide (Revlimid®, Celgene Corp) is a thalidomide derivative that is up to 30,000 times more potent a TNFa inhibitor than thalidomide and is also more active in stimulating T-cell proliferation, production of IL-2 and IFNg [126]. In the clinical setting, lenalidomide has been utilised in a phase I trial in relapsed/refractory MM

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

99

patients. There was an anti-myeloma clinical activity (decreases in serum paraprotein levels of >25%) in 66% and stabilisation or decrease in 79% of patients [127]. Importantly, lenalidomide treatment was associated to fewer side effects than thalidomide; in particular, less constipation, sedation and neuropathy were seen. Based on this, phase II and phase III trials with Revlimid® are currently under development [128]. One of these phase II trials has recently been completed [129]. Lenalidomide was associated with dexamethasone as first-line treatment in 34 MM patients. The administration schedule was 25 mg daily on days 1–21 of a 28-day cycle for lenalidomide and dexamethasone orally 40 mg daily on days 1–4, 9–12, 17–20 of each cycle. Six percent of patients achieved a complete response, and 32% a very good partial response or a near complete response, with an overall response rate of 91%. Fatigue and muscle weakness were the major adverse effects experienced [129]. Very recently, the results of a phase II trials testing the effectiveness of lenalidomide as monotherapy in relapsed/refractory MM, given at two different dosing schedules (15 mg twice daily or 30 mg once daily, oral) were published, demonstrating its activity and well tolerability [127]. Based on this, lenalidomide has been approved for the treatment of refractory/relapsed MM.

Anti-angiogenic Therapy VEGF Inhibitors Since vascular endothelial growth factor (VEGF) has been shown to regulate not only neo-angiogenesis in MM bone marrow but also proliferation and migration of MM cells which express high levels of VEGFR [130, 131], and increased microvessel density in MM bone marrow correlates with a worse clinical course [132], investigators have been prompted study the effects of the interruption of the VEGF–VEGFR axis as a potentially useful anti-MM therapy. Preclinical studies with a small synthetic specific VEGFR inhibitor, PTK787 (Novartis and Schering), showed a direct inhibition of MM cell proliferation and an inhibitory activity on IL-6 production in the bone marrow microenvironment [133]. In another study, a pan-VEGFR inhibitor, GW654652, displayed an antagonistic effect on MM PC migration and proliferation triggered by VEGF [134]. A tyrosine kinase inhibitor with selectivity for VEGFR2, SU5416 (Sugen), has been used in a phase II clinical trial on relapsed/ refractory MM patients. This study showed no clinical activity but confirmed biological activity in vivo (reduction of plasma VEGF levels) [135]. However, more recent clinical studies have demonstrated that the use of pazopanib (GW786034), an oral angiogenesis inhibitor targeting VEGFR, platelet-derived growth factor receptor (PDGFR) and c-kit, as a single agent had no clinical effectiveness in relapsed/ refractory MM [136]. Taken together, these studies suggest that the usefulness of hampering the VEGF–VEGFR in treating MM deserves further investigation.

100

F. Piazza and G. Semenzato

Fibroblast Growth Factor Receptor 3 (FGFR3) Inhibitors FGFs display pro-angiogenic activity and are secreted in the MM BM milieu by BMSCs. They also bind the receptor tyrosine kinase FGFR on MM cells and stimulate MM cell growth. The importance of this family of GFs in MM biology is highlighted by the rearrangement of FGFR3 in the t(4;14) (p16.3; q32.3) [137]. This translocation is present in approximately 15% of MM. Thus, FGFR3 has attracted the attention of researchers as a potential therapeutic target in MM. In one recent study, two FGFR selective inhibitors, SU5402 and SU10991 (Sugen), have been tested. These compounds inhibited the growth of FGFR3+ MM cell lines in vitro and in murine MM models [138]. Also the synthetic compound PD173074 caused cell growth arrest, increase of plasma cell differentiation markers and, eventually, apoptosis of FGFR3+ MM cells [139]. Recently, the small-molecule tyrosine kinase inhibitor PKC412 was found able to inhibit the growth of B cells expressing mutated FGFR3 and TELFGFR3 fusion protein and of MM cell lines harbouring FGFR3 rearrangements [140]. Moreover, another approach using an anti-FGFR3 neutralising antibody, PRO-001, has been demonstrated to be effective in inducing MM cell death [141].

Other Signalling Pathways P38 MAP Kinase Inhibitors The rationale of targeting p38 MAP kinase resides in its role upstream from the bortezomib-induced Hsp27 up-regulation, which could partly account for the resistance of MM cells to this agent. Indeed, p38 MAPK inhibition with a novel compound, SCIO-469 (Scios Inc., USA), while not responsible for significant MM cell toxicity, enhanced the bortezomib-induced apoptosis of MM cells together with a reduction of p38 MAPK phosphorylation and Hsp27 up-regulation [142]. Importantly, this compound rendered bortezomib-resistant MM cells sensitive to the proteasome inhibitor. Recently, a study demonstrated that the inhibition of p38 MAPK rescues the function of dendritic cells in MM by rendering them more able to elicit an anti-myeloma humoral and cellular immune response [143].

TGFb Inhibitors Initial studies demonstrated an increased production of TGFb by bone marrow stromal cells and malignant PCs in MM and showed that this cytokine stimulated the secretion of IL-6 by bone marrow stromal cells [144, 145]. Subsequently, different properties have been assigned to TGFb in MM, such as immunomodulation [146, 147], regulation of osteoblast biology and other processes [148]. Recently, the TGFbI receptor inhibitor SD-208 was used on MM cells. SD-208

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

101

determined a reduction in IL-6 and VEGF secretion from bone marrow stromal cells and in MM PC growth induced by the adhesion to stromal cells. At a molecular level, this compound blocked the accumulation of Smad2/3 and HIF1a downstream from TGFb signalling, thus altering IL-6 and VEGF transcription, respectively [149].

Summary The success of employing more targeted therapeutic strategies in acute promyelocytic leukaemia (all-trans retinoic acid) and chronic myeloid leukaemia (imatinib, inhibitor of ABL kinase), has opened the avenue to the research of the identification of signalling molecules suitable of pharmacological treatment also in other diseases, including MM. To this regard, in recent years, multiple myeloma has become a model to test novel therapeutic strategies targeting several signalling mechanisms that are deregulated in the malignant plasma cell. These molecular pathways are now better understood, and the expectations are for new agents that specifically target these mechanisms. The passage from the preclinical to the clinical setting has been faster in the last years, and novel, molecular-based treatments are currently under investigation in clinical trials. Collectively, the general strategy to follow is the identification of the best combination of drugs to induce clinically relevant responses. The goals are manifold, such as re-sensitise chemoresistant targets, capitalise on differences in mechanisms of action and gain an advantage in terms of systemic toxicity of conventional chemotherapeutics. Acknowledgements Supported by AIRC (Milan) and AIRC Regional Project with Fondazione CARIPARO e CARIVERONA. We are grateful to all the members of the Unit of Haematological Malignancies at VIMM and of the Haematology–Immunology Section at Padua University Hospital for support and suggestions.

References 1. Kuehl WM, Bergsagel PL. Multiple myeloma: evolving genetic events and host interactions. Nat Rev Cancer. 2002;2:175–187. 2. Bakkus MH, Heirman C, Van Riet I, Van Camp B, Thielemans K. Evidence that multiple myeloma Ig heavy chain VDJ genes contain somatic mutations but show no intraclonal variation. Blood. 1992;80:2326–2335. 3. Fonseca R, Barlogie B, Bataille R, et al. Genetics and cytogenetics of multiple myeloma: a workshop report. Cancer Res. 2004;64:1546–1558. 4. Fonseca R, Blood E, Rue M, et al. Clinical and biologic implications of recurrent genomic aberrations in myeloma. Blood. 2003;101:4569–4575. 5. Avet-Loiseau H, Gerson F, Magrangeas F, Minvielle S, Harousseau JL, Bataille R. Rearrangements of the c-myc oncogene are present in 15% of primary human multiple myeloma tumors. Blood. 2001;98:3082–3086.

102

F. Piazza and G. Semenzato

6. Tricot G, Barlogie B, Jagannath S, et al. Poor prognosis in multiple myeloma is associated only with partial or complete deletions of chromosome 13 or abnormalities involving 11q and not with other karyotype abnormalities. Blood. 1995;86:4250–4256. 7. Hanamura I, Stewart JP, Huang Y, et al. Frequent gain of chromosome band 1q21 in plasmacell dyscrasias detected by fluorescence in situ hybridization: incidence increases from MGUS to relapsed myeloma and is related to prognosis and disease progression following tandem stem-cell transplantation. Blood. 2006;108:1724–1732. 8. Carrasco DR, Tonon G, Huang Y, et al. High-resolution genomic profiles define distinct clinico-pathogenetic subgroups of multiple myeloma patients. Cancer Cell. 2006;9:313–325. 9. Debes-Marun CS, Dewald GW, Bryant S, et al. Chromosome abnormalities clustering and its implications for pathogenesis and prognosis in myeloma. Leukemia. 2003;17:427–436. 10. Smadja NV, Bastard C, Brigaudeau C, Leroux D, Fruchart C. Hypodiploidy is a major prognostic factor in multiple myeloma. Blood. 2001;98:2229–2238. 11. Zhan F, Huang Y, Colla S, et al. The molecular classification of multiple myeloma. Blood. 2006;108:2020–2028. 12. Bergsagel PL, Kuehl WM. Molecular pathogenesis and a consequent classification of multiple myeloma. J Clin Oncol. 2005;23:6333–6338. 13. Zhan F, Hardin J, Kordsmeier B, et al. Global gene expression profiling of multiple myeloma, monoclonal gammopathy of undetermined significance, and normal bone marrow plasma cells. Blood. 2002;99:1745–1757. 14. Davies FE, Dring AM, Li C, et al. Insights into the multistep transformation of MGUS to myeloma using microarray expression analysis. Blood. 2003;102:4504–4511. 15. Mattioli M, Agnelli L, Fabris S, et al. Gene expression profiling of plasma cell dyscrasias reveals molecular patterns associated with distinct IGH translocations in multiple myeloma. Oncogene. 2005;24:2461–2473. 16. Duhrsen U, Hossfeld DK. Stromal abnormalities in neoplastic bone marrow diseases. Ann Hematol. 1996;73:53–70. 17. Damiano JS, Cress AE, Hazlehurst LA, Shtil AA, Dalton WS. Cell adhesion mediated drug resistance (CAM-DR): role of integrins and resistance to apoptosis in human myeloma cell lines. Blood. 1999;93:1658–1667. 18. Mitsiades CS, Mitsiades N, Munshi NC, Anderson KC. Focus on multiple myeloma. Cancer Cell. 2004;6:439–444. 19. Pearse RN, Sordillo EM, Yaccoby S, et al. Multiple myeloma disrupts the TRANCE/ osteoprotegerin cytokine axis to trigger bone destruction and promote tumor progression. Proc Natl Acad Sci USA. 2001;98:11581–11586. 20. Giuliani N, Bataille R, Mancini C, Lazzaretti M, Barille S. Myeloma cells induce imbalance in the osteoprotegerin/osteoprotegerin ligand system in the human bone marrow environment. Blood. 2001;98:3527–3533. 21. Abe M, Hiura K, Wilde J, et al. Role for macrophage inflammatory protein (MIP)-1alpha and MIP-1beta in the development of osteolytic lesions in multiple myeloma. Blood. 2002;100:2195–2202. 22. Zannettino AC, Farrugia AN, Kortesidis A, et al. Elevated serum levels of stromal-derived factor-1alpha are associated with increased osteoclast activity and osteolytic bone disease in multiple myeloma patients. Cancer Res. 2005;65:1700–1709. 23. Tian E, Zhan F, Walker R, et al. The role of the Wnt-signaling antagonist DKK1 in the development of osteolytic lesions in multiple myeloma. N Engl J Med. 2003;349:2483–2494. 24. Oshima T, Abe M, Asano J, et al. Myeloma cells suppress bone formation by secreting a soluble Wnt inhibitor, sFRP-2. Blood. 2005;106:3160–3165. 25. Urashima M, Chauhan D, Uchiyama H, Freeman GJ, Anderson KC. CD40 ligand triggered interleukin-6 secretion in multiple myeloma. Blood. 1995;85:1903–1912. 26. Chauhan D, Uchiyama H, Akbarali Y, et al. Multiple myeloma cell adhesion-induced interleukin-6 expression in bone marrow stromal cells involves activation of NF-kappa B. Blood. 1996;87:1104–1112.

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

103

27. Hideshima T, Chauhan D, Schlossman R, Richardson P, Anderson KC. The role of tumor necrosis factor alpha in the pathophysiology of human multiple myeloma: therapeutic applications. Oncogene. 2001;20:4519–4527. 28. Dankbar B, Padro T, Leo R, et al. Vascular endothelial growth factor and interleukin-6 in paracrine tumor-stromal cell interactions in multiple myeloma. Blood. 2000;95: 2630–2636. 29. Franchimont N, Rydziel S, Canalis E. Transforming growth factor-beta increases interleukin-6 transcripts in osteoblasts. Bone. 2000;26:249–253. 30. Ogata A, Chauhan D, Teoh G, et al. IL-6 triggers cell growth via the Ras-dependent mitogenactivated protein kinase cascade. J Immunol. 1997;159:2212–2221. 31. Heinrich PC, Behrmann I, Haan S, Hermanns HM, Muller-Newen G, Schaper F. Principles of interleukin (IL)-6-type cytokine signalling and its regulation. Biochem J. 2003;374:1–20. 32. Hideshima T, Nakamura N, Chauhan D, Anderson KC. Biologic sequelae of interleukin-6 induced PI3-K/Akt signaling in multiple myeloma. Oncogene. 2001;20:5991–6000. 33. Bharti AC, Shishodia S, Reuben JM, et al. Nuclear factor-kappaB and STAT3 are constitutively active in CD138+ cells derived from multiple myeloma patients, and suppression of these transcription factors leads to apoptosis. Blood. 2004;103:3175–3184. 34. Yoshio-Hoshino N, Adachi Y, Aoki C, Pereboev A, Curiel DT, Nishimoto N. Establishment of a new interleukin-6 (IL-6) receptor inhibitor applicable to the gene therapy for IL-6dependent tumor. Cancer Res. 2007;67:871–875. 35. Voorhees PM, Chen Q, Small GW, et al. Targeted inhibition of interleukin-6 with CNTO 328 sensitizes pre-clinical models of multiple myeloma to dexamethasone-mediated cell death. Br J Haematol. 2009;145:481–490. 36. Tassone P, Neri P, Burger R, et al. Combination therapy with interleukin-6 receptor superantagonist Sant7 and dexamethasone induces antitumor effects in a novel SCID-hu In vivo model of human multiple myeloma. Clin Cancer Res. 2005;11:4251–4258. 37. Mitsiades CS, Mitsiades NS, McMullan CJ, et al. Inhibition of the insulin-like growth factor receptor-1 tyrosine kinase activity as a therapeutic strategy for multiple myeloma, other hematologic malignancies, and solid tumors. Cancer Cell. 2004;5:221–230. 38. Menu E, Jernberg-Wiklund H, Stromberg T, et al. Inhibiting the IGF-1 receptor tyrosine kinase with the cyclolignan PPP: an in vitro and in vivo study in the 5T33MM mouse model. Blood. 2006;107:655–660. 39. Rowinsky EK, Youssoufian H, Tonra JR, Solomon P, Burtrum D, Ludwig DL. IMC-A12, a human IgG1 monoclonal antibody to the insulin-like growth factor I receptor. Clin Cancer Res. 2007;13:5549 s–5555 s. 40. Stromberg T, Ekman S, Girnita L, et al. IGF-1 receptor tyrosine kinase inhibition by the cyclolignan PPP induces G2/M-phase accumulation and apoptosis in multiple myeloma cells. Blood. 2006;107:669–678. Epub 2005 Sep 2015. 41. Karin M, Cao Y, Greten FR, Li ZW. NF-kappaB in cancer: from innocent bystander to major culprit. Nat Rev Cancer. 2002;2:301–310. 42. Karin M, Greten FR. NF-kappaB: linking inflammation and immunity to cancer development and progression. Nat Rev Immunol. 2005;5:749–759. 43. Keats JJ, Fonseca R, Chesi M, et al. Promiscuous mutations activate the noncanonical NF-kappaB pathway in multiple myeloma. Cancer Cell. 2007;12:131–144. 44. Annunziata CM, Davis RE, Demchenko Y, et al. Frequent engagement of the classical and alternative NF-kappaB pathways by diverse genetic abnormalities in multiple myeloma. Cancer Cell. 2007;12:115–130. 45. Feinman R, Koury J, Thames M, Barlogie B, Epstein J, Siegel DS. Role of NF-kappaB in the rescue of multiple myeloma cells from glucocorticoid-induced apoptosis by bcl-2. Blood. 1999;93:3044–3052. 46. Hideshima T, Chauhan D, Richardson P, et al. NF-kappa B as a therapeutic target in multiple myeloma. J Biol Chem. 2002;277:16639–16647. 47. Hideshima T, Catley L, Yasui H, et al. Perifosine, an oral bioactive novel alkylphospholipid, inhibits Akt and induces in vitro and in vivo cytotoxicity in human multiple myeloma cells. Blood. 2006;107:4053–4062.

104

F. Piazza and G. Semenzato

48. Jourdan M, Moreaux J, Vos JD, et al. Targeting NF-kappaB pathway with an IKK2 inhibitor induces inhibition of multiple myeloma cell growth. Br J Haematol. 2007;138:160–168. 49. Hideshima T, Chauhan D, Kiziltepe T, et al. Biologic sequelae of I{kappa}B kinase (IKK) inhibition in multiple myeloma: therapeutic implications. Blood. 2009;113:5228–5236. 50. Bharti AC, Donato N, Singh S, Aggarwal BB. Curcumin (diferuloylmethane) down-regulates the constitutive activation of nuclear factor-kappa B and IkappaBalpha kinase in human multiple myeloma cells, leading to suppression of proliferation and induction of apoptosis. Blood. 2003;101:1053–1062. 51. Di Cristofano A, Pandolfi PP. The multiple roles of PTEN in tumor suppression. Cell. 2000;100:387–390. 52. Tu Y, Gardner A, Lichtenstein A. The phosphatidylinositol 3-kinase/AKT kinase pathway in multiple myeloma plasma cells: roles in cytokine-dependent survival and proliferative responses. Cancer Res. 2000;60:6763–6770. 53. Hideshima T, Catley L, Raje N, et al. Inhibition of Akt induces significant downregulation of survivin and cytotoxicity in human multiple myeloma cells. Br J Haematol. 2007;138:783–791. 54. Gajate C, Mollinedo F. Edelfosine and perifosine induce selective apoptosis in multiple myeloma by recruitment of death receptors and downstream signaling molecules into lipid rafts. Blood. 2007;109:711–719. 55. Danial NN, Korsmeyer SJ. Cell death: critical control points. Cell. 2004;116:205–219. 56. Danial NN. BCL-2 family proteins: critical checkpoints of apoptotic cell death. Clin Cancer Res. 2007;13:7254–7263. 57. Ludwig H. Advances in biology and treatment of multiple myeloma. Ann Oncol. 2005;16:ii106–112. 58. Badros AZ, Goloubeva O, Rapoport AP, et al. Phase II study of G3139, a Bcl-2 antisense oligonucleotide, in combination with dexamethasone and thalidomide in relapsed multiple myeloma patients. J Clin Oncol. 2005;23:4089–4099. 59. Chanan-Khan AA, Niesvizky R, Hohl RJ, et al. Phase III randomised study of dexamethasone with or without oblimersen sodium for patients with advanced multiple myeloma. Leuk Lymphoma. 2009;50:559–565. 60. Trudel S, Stewart AK, Li Z, et al. The Bcl-2 family protein inhibitor, ABT-737, has substantial antimyeloma activity and shows synergistic effect with dexamethasone and melphalan. Clin Cancer Res. 2007;13:621–629. 61. Kline MP, Rajkumar SV, Timm MM, et al. ABT-737, an inhibitor of Bcl-2 family proteins, is a potent inducer of apoptosis in multiple myeloma cells. Leukemia. 2007;21:1549–1560. 62. Tse C, Shoemaker AR, Adickes J, et al. ABT-263: a potent and orally bioavailable Bcl-2 family inhibitor. Cancer Res. 2008;68:3421–3428. 63. Rasmussen T, Kuehl M, Lodahl M, Johnsen HE, Dahl IM. Possible roles for activating RAS mutations in the MGUS to MM transition and in the intramedullary to extramedullary transition in some plasma cell tumors. Blood. 2005;105:317–323. 64. Hu L, Shi Y, Hsu JH, Gera J, Van Ness B, Lichtenstein A. Downstream effectors of oncogenic ras in multiple myeloma cells. Blood. 2003;101:3126–3135. 65. Le Gouill S, Pellat-Deceunynck C, Harousseau JL, et al. Farnesyl transferase inhibitor R115777 induces apoptosis of human myeloma cells. Leukemia. 2002;16:1664–1667. 66. Ochiai N, Uchida R, Fuchida S, et al. Effect of farnesyl transferase inhibitor R115777 on the growth of fresh and cloned myeloma cells in vitro. Blood. 2003;102:3349–3353. 67. Beaupre DM, McCafferty-Grad J, Bahlis NJ, Boise LH, Lichtenheld MG. Farnesyl transferase inhibitors enhance death receptor signals and induce apoptosis in multiple myeloma cells. Leuk Lymphoma. 2003;44:2123–2134. 68. Bolick SC, Landowski TH, Boulware D, et al. The farnesyl transferase inhibitor, FTI-277, inhibits growth and induces apoptosis in drug-resistant myeloma tumor cells. Leukemia. 2003;17:451–457. 69. Cortes J, Albitar M, Thomas D, et al. Efficacy of the farnesyl transferase inhibitor R115777 in chronic myeloid leukemia and other hematologic malignancies. Blood. 2003;101:1692–1697.

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

105

70. Alsina M, Fonseca R, Wilson EF, et al. Farnesyltransferase inhibitor tipifarnib is well tolerated, induces stabilization of disease, and inhibits farnesylation and oncogenic/tumor survival pathways in patients with advanced multiple myeloma. Blood. 2004;103:3271–3277. 71. Mitsiades CS, Mitsiades NS, McMullan CJ, et al. Anti-myeloma activity of heat shock protein-90 inhibition. Blood. 2005;107:1092–1100. 72. David E, Sun SY, Waller EK, Chen J, Khuri FR, Lonial S. The combination of the Farnesyl transferase inhibitor (Lonafarnib) and the proteasome inhibitor (Bortezomib) induces synergistic apoptosis in human myeloma cells that is associated with down-regulation of p-AKT. Blood. 2005;106:4322–4329. 73. van de Donk NW, Lokhorst HM, Nijhuis EH, Kamphuis MM, Bloem AC. Geranylgeranylated proteins are involved in the regulation of myeloma cell growth. Clin Cancer Res. 2005;11:429–439. 74. Morgan MA, Sebil T, Aydilek E, Peest D, Ganser A, Reuter CW. Combining prenylation inhibitors causes synergistic cytotoxicity, apoptosis and disruption of RAS-to-MAP kinase signalling in multiple myeloma cells. Br J Haematol. 2005;130:912–925. 75. Wang S, El-Deiry WS. TRAIL and apoptosis induction by TNF-family death receptors. Oncogene. 2003;22:8628–8633. 76. Gazitt Y. TRAIL is a potent inducer of apoptosis in myeloma cells derived from multiple myeloma patients and is not cytotoxic to hematopoietic stem cells. Leukemia. 1999;13:1817–1824. 77. Mitsiades CS, Treon SP, Mitsiades N, et al. TRAIL/Apo2L ligand selectively induces apoptosis and overcomes drug resistance in multiple myeloma: therapeutic applications. Blood. 2001;98:795–704. 78. Gomez-Benito M, Balsas P, Bosque A, Anel A, Marzo I, Naval J. Apo2L/TRAIL is an indirect mediator of apoptosis induced by interferon-alpha in human myeloma cells. FEBS Lett. 2005;579:6217–6222. 79. Crowder C, Dahle O, Davis RE, Gabrielsen OS, Rudikoff S. PML mediates IFN-alpha-induced apoptosis in myeloma by regulating TRAIL induction. Blood. 2005;105:1280–1287. 80. Whitesell L, Lindquist SL. HSP90 and the chaperoning of cancer. Nat Rev Cancer. 2005;5:761–772. 81. Mitsiades N, Mitsiades CS, Poulaki V, et al. Molecular sequelae of proteasome inhibition in human multiple myeloma cells. Proc Natl Acad Sci USA. 2002;99:14374–14379. 82. Mani A, Gelmann EP. The ubiquitin-proteasome pathway and its role in cancer. J Clin Oncol. 2005;23:4776–4789. 83. Ciechanover A, Orian A, Schwartz AL. Ubiquitin-mediated proteolysis: biological regulation via destruction. Bioessays. 2000;22:442–451. 84. Ciechanover A. Proteolysis: from the lysosome to ubiquitin and the proteasome. Nat Rev Mol Cell Biol. 2005;6:79–87. 85. Adams J. Proteasome inhibition in cancer: development of PS-341. Semin Oncol. 2001;28:613–619. 86. Elliott PJ, Ross JS. The proteasome: a new target for novel drug therapies. Am J Clin Pathol. 2001;116:637–646. 87. Hideshima T, Richardson P, Chauhan D, et al. The proteasome inhibitor PS-341 inhibits growth, induces apoptosis, and overcomes drug resistance in human multiple myeloma cells. Cancer Res. 2001;61:3071–3076. 88. LeBlanc R, Catley LP, Hideshima T, et al. Proteasome inhibitor PS-341 inhibits human myeloma cell growth in vivo and prolongs survival in a murine model. Cancer Res. 2002;62:4996–5000. 89. Orlowski RZ, Stinchcombe TE, Mitchell BS, et al. Phase I trial of the proteasome inhibitor PS-341 in patients with refractory hematologic malignancies. J Clin Oncol. 2002;20:4420–4427. 90. Richardson PG, Barlogie B, Berenson J, et al. A phase 2 study of bortezomib in relapsed, refractory myeloma. N Engl J Med. 2003;348:2609–2617. 91. Jagannath S, Barlogie B, Berenson J, et al. A phase 2 study of two doses of bortezomib in relapsed or refractory myeloma. Br J Haematol. 2004;127:165–172.

106

F. Piazza and G. Semenzato

92. Richardson PG, Sonneveld P, Schuster MW, et al. Bortezomib or high-dose dexamethasone for relapsed multiple myeloma. N Engl J Med. 2005;352:2487–2498. 93. Adams J, Palombella VJ, Sausville EA, et al. Proteasome inhibitors: a novel class of potent and effective antitumor agents. Cancer Res. 1999;59:2615–2622. 94. Rajkumar SV, Richardson PG, Hideshima T, Anderson KC. Proteasome inhibition as a novel therapeutic target in human cancer. J Clin Oncol. 2005;23:630–639. 95. Zhang HG, Wang J, Yang X, Hsu HC, Mountz JD. Regulation of apoptosis proteins in cancer cells by ubiquitin. Oncogene. 2004;23:2009–2015. 96. Cusack JC, Jr., Liu R, Houston M, et al. Enhanced chemosensitivity to CPT-11 with proteasome inhibitor PS-341: implications for systemic nuclear factor-kappaB inhibition. Cancer Res. 2001;61:3535–3540. 97. Chauhan D, Li G, Podar K, et al. Targeting mitochondria to overcome conventional and bortezomib/proteasome inhibitor PS-341 resistance in multiple myeloma (MM) cells. Blood. 2004;104:2458–2466. 98. Sayers TJ, Brooks AD, Koh CY, et al. The proteasome inhibitor PS-341 sensitizes neoplastic cells to TRAIL-mediated apoptosis by reducing levels of c-FLIP. Blood. 2003;102:303–310. 99. Sayers TJ, Murphy WJ. Combining proteasome inhibition with TNF-related apoptosis-inducing ligand (Apo2L/TRAIL) for cancer therapy. Cancer Immunol Immunother. 2005. 100. Nikrad M, Johnson T, Puthalalath H, Coultas L, Adams J, Kraft AS. The proteasome inhibitor bortezomib sensitizes cells to killing by death receptor ligand TRAIL via BH3-only proteins Bik and Bim. Mol Cancer Ther. 2005;4:443–449. 101. Chauhan D, Li G, Podar K, et al. A novel carbohydrate-based therapeutic GCS-100 overcomes bortezomib resistance and enhances dexamethasone-induced apoptosis in multiple myeloma cells. Cancer Res. 2005;65:8350–8358. 102. Chauhan D, Catley L, Li G, et al. A novel orally active proteasome inhibitor induces apoptosis in multiple myeloma cells with mechanisms distinct from Bortezomib. Cancer Cell. 2005;8:407–419. 103. Chauhan D, Catley L, Li G, et al. Preclinical evaluation of a novel and orally active proteasome inhibitor as a therapy in relapsed/refractory multiple myeloma. ASCO Annual Meeting Proceedings. 2005;23:3122. 104. Williams PG, Buchanan GO, Feling RH, Kauffman CA, Jensen PR, Fenical W. New cytotoxic salinosporamides from the marine Actinomycete Salinispora tropica. J Org Chem. 2005;70:6196–6203. 105. Kuhn DJ, Chen Q, Voorhees PM, et al. Potent activity of carfilzomib, a novel, irreversible inhibitor of the ubiquitin-proteasome pathway, against preclinical models of multiple myeloma. Blood. 2007;110:3281–3290. 106. Peterson CL, Laniel MA. Histones and histone modifications. Curr Biol. 2004;14:R546–551. 107. Piazza F, Semenzato G. Molecular therapeutic approaches to acute myeloid leukemia: targeting aberrant chromatin dynamics and signal transduction. Expert Rev Anticancer Ther. 2004;4:387–400. 108. Minucci S, Pelicci PG. Histone deacetylase inhibitors and the promise of epigenetic (and more) treatments for cancer. Nat Rev Cancer. 2006;6:38–51. 109. Marks PA, Richon VM, Miller T, Kelly WK. Histone deacetylase inhibitors. Adv Cancer Res. 2004;91:137–168. 110. Lavelle D, Chen YH, Hankewych M, DeSimone J. Histone deacetylase inhibitors increase p21(WAF1) and induce apoptosis of human myeloma cell lines independent of decreased IL-6 receptor expression. Am J Hematol. 2001;68:170–178. 111. Mitsiades N, Mitsiades CS, Richardson PG, et al. Molecular sequelae of histone deacetylase inhibition in human malignant B cells. Blood. 2003;101:4055–4062. 112. Mitsiades CS, Mitsiades NS, McMullan CJ, et al. Transcriptional signature of histone deacetylase inhibition in multiple myeloma: biological and clinical implications. Proc Natl Acad Sci USA. 2004;101:540–545. 113. Khan SB, Maududi T, Barton K, Ayers J, Alkan S. Analysis of histone deacetylase inhibitor, depsipeptide (FR901228), effect on multiple myeloma. Br J Haematol. 2004;125: 156–161.

5 Signalling Molecules as Selective Targets for Therapeutic Strategies…

107

114. Pei XY, Dai Y, Grant S. Synergistic induction of oxidative injury and apoptosis in human multiple myeloma cells by the proteasome inhibitor bortezomib and histone deacetylase inhibitors. Clin Cancer Res. 2004;10:3839–3852. 115. Hideshima T, Bradner JE, Wong J, et al. Small-molecule inhibition of proteasome and aggresome function induces synergistic antitumor activity in multiple myeloma. Proc Natl Acad Sci USA. 2005;102:8567–8572. 116. Sampaio EP, Sarno EN, Galilly R, Cohn ZA, Kaplan G. Thalidomide selectively inhibits tumor necrosis factor alpha production by stimulated human monocytes. J Exp Med. 1991;173:699–703. 117. Deng L, Ding W, Granstein RD. Thalidomide inhibits tumor necrosis factor-alpha production and antigen presentation by Langerhans cells. J Invest Dermatol. 2003;121:1060–1065. 118. D’Amato RJ, Loughnan MS, Flynn E, Folkman J. Thalidomide is an inhibitor of angiogenesis. Proc Natl Acad Sci USA. 1994;91:4082–4085. 119. Singhal S, Mehta J, Desikan R, et al. Antitumor activity of thalidomide in refractory multiple myeloma. N Engl J Med. 1999;341:1565–1571. 120. Barlogie B, Desikan R, Eddlemon P, et al. Extended survival in advanced and refractory multiple myeloma after single-agent thalidomide: identification of prognostic factors in a phase 2 study of 169 patients. Blood. 2001;98:492–494. 121. Alexanian R, Weber D, Anagnostopoulos A, Delasalle K, Wang M, Rankin K. Thalidomide with or without dexamethasone for refractory or relapsing multiple myeloma. Semin Hematol. 2003;40:3–7. 122. Barlogie B, Zangari M, Spencer T, et al. Thalidomide in the management of multiple myeloma. Semin Hematol. 2001;38:250–259. 123. Barlogie B, Tricot G, Anaissie E, et al. Thalidomide and hematopoietic-cell transplantation for multiple myeloma. N Engl J Med. 2006;354:1021–1030. 124. Vacca A, Scavelli C, Montefusco V, et al. Thalidomide downregulates angiogenic genes in bone marrow endothelial cells of patients with active multiple myeloma. J Clin Oncol. 2005;23:5334–5346. Epub 2005 Jun 5336. 125. Anderson KC. The role of immunomodulatory drugs in multiple myeloma. Semin Hematol. 2003;40:23–32. 126. Richardson P, Anderson K. Immunomodulatory analogs of thalidomide: an emerging new therapy in myeloma. J Clin Oncol. 2004;22:3212–3224. 127. Richardson PG, Blood E, Mitsiades CS, et al. A randomized phase 2 study of lenalidomide therapy for patients with relapsed or relapsed and refractory multiple myeloma. Blood. 2006. 128. Bartlett JB, Tozer A, Stirling D, Zeldis JB. Recent clinical studies of the immunomodulatory drug (IMiD) lenalidomide. Br J Cancer. 2005;93:613–619. 129. Rajkumar SV, Hayman SR, Lacy MQ, et al. Combination therapy with lenalidomide plus dexamethasone (Rev/Dex) for newly diagnosed myeloma. Blood . 2005;106:4050–4053. Epub 2005 Aug 4023. 130. Podar K, Tai YT, Davies FE, et al. Vascular endothelial growth factor triggers signaling cascades mediating multiple myeloma cell growth and migration. Blood. 2001;98:428–435. 131. Podar K, Tai YT, Lin BK, et al. Vascular endothelial growth factor-induced migration of multiple myeloma cells is associated with beta 1 integrin- and phosphatidylinositol 3-kinasedependent PKC alpha activation. J Biol Chem. 2002;277:7875–7881. 132. Vacca A, Ribatti D. Bone marrow angiogenesis in multiple myeloma. Leukemia. 2006;20:193–199. 133. Lin B, Podar K, Gupta D, et al. The vascular endothelial growth factor receptor tyrosine kinase inhibitor PTK787/ZK222584 inhibits growth and migration of multiple myeloma cells in the bone marrow microenvironment. Cancer Res. 2002;62:5019–5026. 134. Podar K, Catley LP, Tai YT, et al. GW654652, the pan-inhibitor of VEGF receptors, blocks the growth and migration of multiple myeloma cells in the bone marrow microenvironment. Blood. 2004;103:3474–3479. 135. Zangari M, Anaissie E, Stopeck A, et al. Phase II study of SU5416, a small molecule vascular endothelial growth factor tyrosine kinase receptor inhibitor, in patients with refractory multiple myeloma. Clin Cancer Res. 2004;10:88–95.

108

F. Piazza and G. Semenzato

136. Prince HM, Honemann D, Spencer A, et al. Vascular endothelial growth factor inhibition is not an effective therapeutic strategy for relapsed or refractory multiple myeloma: a phase 2 study of pazopanib (GW786034). Blood. 2009;113:4819–4820. 137. Chesi M, Nardini E, Brents LA, et al. Frequent translocation t(4;14)(p16.3;q32.3) in multiple myeloma is associated with increased expression and activating mutations of fibroblast growth factor receptor 3. Nat Genet. 1997;16:260–264. 138. Paterson JL, Li Z, Wen XY, et al. Preclinical studies of fibroblast growth factor receptor 3 as a therapeutic target in multiple myeloma. Br J Haematol. 2004;124:595–603. 139. Trudel S, Ely S, Farooqi Y, et al. Inhibition of fibroblast growth factor receptor 3 induces differentiation and apoptosis in t(4;14) myeloma. Blood. 2004;103:3521–3528. 140. Chen J, Lee BH, Williams IR, et al. FGFR3 as a therapeutic target of the small molecule inhibitor PKC412 in hematopoietic malignancies. Oncogene. 2005;24:8259–8267. 141. Trudel S, Stewart AK, Rom E, et al. The inhibitory anti-FGFR3 antibody, PRO-001 is cytotoxic to t(4;14) multiple myeloma cells. Blood. 2006;7:7. 142. Hideshima T, Podar K, Chauhan D, et al. p38 MAPK inhibition enhances PS-341 (bortezomib)induced cytotoxicity against multiple myeloma cells. Oncogene. 2004;23:8766–8776. 143. Wang S, Yang J, Qian J, Wezeman M, Kwak LW, Yi Q. Tumor evasion of the immune system: inhibiting p38 MAPK signaling restores the function of dendritic cells in multiple myeloma. Blood. 2006;107:2432–2439. 144. Urashima M, Ogata A, Chauhan D, et al. Transforming growth factor-beta1: differential effects on multiple myeloma versus normal B cells. Blood. 1996;87:1928–1938. 145. Kroning H, Tager M, Thiel U, et al. Overproduction of IL-7, IL-10 and TGF-beta 1 in multiple myeloma. Acta Haematol. 1997;98:116–118. 146. Cook G, Campbell JD, Carr CE, Boyd KS, Franklin IM. Transforming growth factor beta from multiple myeloma cells inhibits proliferation and IL-2 responsiveness in T lymphocytes. J Leukoc Biol. 1999;66:981–988. 147. Abildgaard N, Glerup H, Rungby J, et al. Biochemical markers of bone metabolism reflect osteoclastic and osteoblastic activity in multiple myeloma. Eur J Haematol. 2000;64:121–129. 148. Wright N, de Lera TL, Garcia-Moruja C, et al. Transforming growth factor-beta1 downregulates expression of chemokine stromal cell-derived factor-1: functional consequences in cell migration and adhesion. Blood. 2003;102:1978–1984. 149. Hayashi T, Hideshima T, Nguyen AN, et al. Transforming growth factor beta receptor I kinase inhibitor down-regulates cytokine secretion and multiple myeloma cell growth in the bone marrow microenvironment. Clin Cancer Res. 2004;10:7540–7546.

Chapter 6

Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract Hiroshi Yasuda and Fumio Itoh

Introduction Bile acids are synthesized in the liver through the oxidation of cholesterol, are excreted into the gallbladder, and are then released into the intestinal tract. Bile acids are required for cholesterol excretion and lipid absorption. They include the primary bile acids, cholic acid and chenodeoxycholic acid, their glycine and taurine conjugates, and the secondary bile acids, deoxycholic acid and lithocholic acid. Through the activity of the intestinal anaerobic bacterial flora, the primary bile acids undergo 7a-dehydroxylation to yield deoxycholic acid and lithocholic acid, respectively [1, Fig. 6.1]. Deoxycholic acid is partially reabsorbed in the large intestine and enters into the enterohepatic circulation. Lithocholic acid is fairly insoluble, and little is reabsorbed. Besides their roles in dietary lipid absorption and cholesterol homeostasis, it has become clear that bile acids also are signaling molecules with endocrine function. They affect lipid and glucose metabolism and also energy homeostasis. Bile acids activate the nuclear receptor, farnesoid X receptor (FXR, NR1H4) [2–4]. FXR regulates the transcription of genes encoding pivotal bile acid biosynthetic enzymes and transport proteins. These include CYP7A1, for 7a-cholesterol hydroxylase, which is the rate-limiting enzyme for bile acid synthesis, ileal bile acid-binding protein, and FGF15, an essential factor for filling the gallbladder [5, 6]. FXR is expressed in the adrenal glands, intestine, kidney, and liver. FXR binds to the FXR response element after heterodimerization with the retinoid X receptor (RXR).

H. Yasuda (*) Division of Gastroenterology and Hepatology, St. Marianna University School of Medicine, 2-16-1 Sugao, 216-8511 Kawasaki, Japan e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_6, © Springer Science+Business Media, LLC 2012

109

110 Fig. 6.1 Chemical structures of the major human bile acids

H. Yasuda and F. Itoh

cholic acid chenodeoxycholic acid deoxycholic acid lithocholic acid

R1

R2

OH OH H H

OH H OH H

Free acid, X= OH Glyco, X= NHCH2CO2H Tauro, X= NHCH2CH2SO3H

Bile acids activate several intracellular protein kinases, such as mitogen-activated protein kinase (MAPK) and protein kinase C (PKC). Recently, a membrane-type bile acid receptor TGR5, which has seven transmembrane domains and couples to trimeric G protein, has been identified [7, 8]. This review focuses on the role of bile acids in gastrointestinal carcinogenesis as a luminal toxin and endocrine modulator. We discuss our recent results on bile acid-induced activation of the EGFR–MAPK pathway via TGR5 activation and its role in gastrointestinal carcinogenesis.

Bile Acids and Carcinogenesis Cook et al. first reported that malignant tumors developed after repeated injection of deoxycholic acid into the flanks of mice [9]. This was the first experimental report of malignant growths stimulated by a compound that is present normally in the human body. Bile acids have been reported as tumor promoters of known mutagens in animal experiments, but no tumors were detected in the colons of rats given bile acid alone without a known mutagen. Bile acids have been studied predominantly in relation to colorectal carcinogenesis; however, bile acids now have been implicated in playing roles in carcinogenesis throughout the gastrointestinal tract [10]. Dietary habits may affect stool and serum concentrations of bile acids, which may affect colorectal carcinogenesis. Bile acids can induce mucosal injury, stimulate cell proliferation, and promote tumorigenesis. In addition, bile acids affect anatomical regions exposed abnormally for prolonged periods to high bile acid concentrations,

6 Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract

111

such as the resected stomach, gastric pylorus, and esophagogastric junction with reflux of bile acids from the duodenum and stomach.

Bile Acids and COX-2 Expression in Gastrointestinal Carcinogenesis Cyclooxygenase (COX) is a membrane-bound glycoprotein that functions as a ratelimiting enzyme in prostaglandin (PG) synthesis. COX catalyzes the conversion of arachidonic acid to the common precursor prostanoids, prostaglandins (PG) H2 and PGG2. PGH2 is subsequently converted to a variety of PGs, including PGE2, PGD1, PGF2a, PGI2, and thromboxane A2, by the respective PG synthetase. Each of these final metabolites binds to a specific GPCR to trigger intracellular responses. Several isoforms of COX enzymes have been reported. COX-1 is constitutively expressed and considered to be a housekeeping enzyme. Prostanoids synthesized via the COX-1 pathway are responsible for cytoprotection of the gastric mucosa, for the production of thromboxane by platelets, and for vasodilation of the kidney. In contrast, COX-2 is an immediate early gene induced by cytokines, growth factors, tumor promoters, and viral transformation. Its pathophysiological role has been connected to inflammation, the immune system, ovulation, and carcinogenesis [11–13]. Many different stimuli induce COX-2 expression. In 1971, Vane and colleagues demonstrated first that aspirin and indomethacin inhibited PG production by blocking COX activity [14]. Since then, it has been reported that nonsteroidal anti-inflammatory drugs (NSAIDs) affect COX activity directly. It is well documented that COX inhibitors reduce cancer growth. Prostanoids produced by COX-2, especially PGE2, enhance cell proliferation in normal and cancer cells through specific receptors [15, 16]. PGE2 receptors, namely EP1-4, are a family of GPCRs. PGE2 also is a potent immunosuppressor [17]. PGE2 inhibits the production of immune regulatory lymphokines, T-cell and B-cell proliferation, and the cytotoxic activity of NK cells, thus favoring tumor growth. In addition, COX-2 induces the production of vascular endothelial growth factor (VEGF), a proangiogenic growth factor, and may promote angiogenesis [18, 19]. Furthermore, prostanoids are involved in inhibition of cancer cell apoptosis [20]. Elevated expression of COX-2 has been reported in esophageal, gastric, and colon cancers. Elevated expression of COX-2 was associated with the reduced survival of patients undergoing surgery. COX-2 was involved in VEGF expression and angiogenesis and immunomodulation in these tumors. In the upper gastrointestinal tract, duodeno-gastro-esophageal reflux is one of the major factors for increased COX-2 expression. Bile acids have been reported to induce COX-2 expression in many cell culture systems, including cells from the gastrointestinal tract, which indicates a relationship with carcinogenesis [21–23]. Bile acids induced COX-2 by both transcriptional and post-transcriptional mechanisms [21]. Bile acids are potent activators of PKC isoenzymes in the colonic mucosa [24]. PKC is implicated as being important for bile acid-mediated induction of COX-2 [25, 26].

112

H. Yasuda and F. Itoh

Bile Acids and Human Gastrointestinal Cancer Esophagus Esophageal Squamous Cell Carcinoma The two major histological types of esophageal cancer are squamous cell carcinoma (SCC) and Barrett adenocarcinoma. In East Asia, SCC is the predominant histological type of esophageal cancer. Factors associated with increased risk of esophageal SCC include smoking and alcohol intake. Reflux of gastroduodenal contents after gastrectomy also has been reported as a risk factor of esophageal SCC [27, 28]. In an animal study, esophageal SCC or adenocarcinoma developed in rats with surgically induced gastro-duodeno-esophageal or duodeno-esophageal reflux [29].

Barrett’s Epithelium and Esophageal Adenocarcinoma Adenocarcinoma of the esophagus is one of most rapidly increasing cancers in Caucasian males in the United States, where the incidence of SCC of the esophagus has declined. Five-year survival rates rarely exceed 25%, even after intentionally curative resection, and early detection of esophageal adenocarcinoma is of great clinical significance. Barrett’s epithelium (BE) is a premalignant condition of the distal esophagus and is associated with a 30–125-fold higher risk and 0.5–1% conversion rate of developing esophageal adenocarcinoma, compared with individuals in the general population. BE is characterized by the replacement of the normal esophageal squamous epithelium with a metaplastic epithelium resembling a more distal intestine and is considered to be induced by chronic gastroesophageal reflux disease (GERD). Factors that increase gastroesophageal reflux are dietary components, increasing body mass index and eradicating Helicobacter pylori (H. pylori). BE could arise from two alternative cellular origins. One possibility is direct transdifferentiation from mature squamous cells. The other is that certain pluripotent stem cells from squamous epithelium or bone marrow might contribute to the development of BE [30]. The luminal environment of the distal esophagus may stimulate these cells to develop BE. Mixed reflux of the gastric and duodenal contents is common in GERD patients. Chronic exposure to duodeno-gastric refluxate, especially bile acids and gastric acid, is critical. These may affect intracellular signaling systems and genetic or epigenetic factors. Because treatment with proton pump inhibitors does not seem to reverse or halt the progression of Barrett adenocarcinoma, attention has turned to the role of bile acids in the genesis of BE and metaplasia. The bile acid compositions reported in esophageal aspirates include mostly glycine or taurine conjugated or sulfated forms. Duodenal refluxate into the esophagus of these patients can yield peak concentrations of bile acids up to 2 mM at levels as high as the maximum bile acid concentration reported in the fecal water of the colon after a high-fat diet [31, 32]. Moreover, the bile acid concentration of

6 Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract

113

esophageal aspirates was higher in patients with esophagitis and BE [33, 34]. Of note, conjugated bile acids stimulated cell proliferation in esophageal cell lines through the p38MAPK and p42/44MAPK pathways [35].

CDX-2 and Barrett’s Epithelium CDX-2 is a member of the caudal-related homeobox transcription factor gene family that plays an important role in the early differentiation and maintenance of the intestinal epithelium. CDX-2 is a direct transcriptional activator of many intestinespecific genes, such as MUC-2, sucrase-isomaltase, and guanylyl cyclase. While CDX-2 is not expressed in the normal esophageal epithelium, increased expression of CDX-2 protein and mRNA were observed in mucosa from BE [36, 37]. CDX-2 positive cells were observed in columnar epithelium of BE model rats with esophagojejunal anastomosis [38, 39]. An association between CDX-2 expression and intestinal metaplasia also has been reported in the stomach. While the normal gastric mucosa does not express CDX-2, intestinal metaplastic tissues in the human stomach stain intensively for the protein. In transgenic mice that overexpress CDX-2 in gastric parietal cells, the gastric fundic mucosa was changed into intestinal metaplastic mucosa [40, 41]. Of note, reflux of bile acids seems to be involved in CDX-2 expression and transdifferentiation from squamous to columnar epithelium. In cultured esophageal cells, bile acids induced CDX-2 expression via an NFkB-dependent pathway [37, 38]. Moreover, overexpression of CDX-2 in cultured rat esophageal keratinocytes induced MUC2. While CDX-2 is important for maintaining the intestinal phenotype, its expression was decreased in BE with dysplasia to Barrett adenocarcinoma [42]. In accordance with this, reduced expression of CDX-2 protein has been reported also in gastric and colon carcinomas and may be responsible for carcinogenesis [43]. Accordingly, exogenous CDX-2 expression in CRC cell lines inhibited cell proliferation [44]. Moreover, CDX-2+/− mice developed multiple intestinal adenomatous polyps in the proximal colon [45], and in addition, mice doubly heterozygous for APCD716 and Cdx-2+/− developed six times the number of colonic polyps than their APCD716 and CDX-2+/− littermates [46]. Interestingly, CDX-2 was shown to reduce COX-2 expression. In CRC cell lines, CDX-2 inhibited transcription of COX-2 by interfering with the binding of NF-kB to the NF-kB binding site [47, 48]. Collectively, CDX-2 seems to have a tumor-suppressive role in matured intestinal or metaplastic epithelium.

COX-2 Expression and Bile Acids in Esophageal Adenocarcinoma COX-2 has been implicated in the neoplastic progression of BE. COX-2 immunoreactivity was detected in most specimens of esophageal adenocarcinoma. Its expression was enhanced in the metaplastic and dysplastic epithelium of BE and in adenocarcinoma in a progressively increasing fashion [49]. Elevated expression of COX-2 was associated with reduced survival of patients undergoing surgery for

114

H. Yasuda and F. Itoh

esophageal adenocarcinoma [50]. Bile acids have been implicated in COX-2 expression in BE and Barrett adenocarcinoma. In in vitro studies, chenodeoxycholic acid and deoxycholic acid induced COX-2 mRNA expression and enhanced PGE2 production in human esophageal adenocarcinoma cell lines [21]. Unconjugated bile acids induce COX-2 expression in esophageal cell lines via the reactive oxygen speciesAKT mediated signaling pathway. Chenodeoxycholic acid induced COX-2 expression and VEGF production in esophageal adenocarcinoma and SCC cell lines [51]. In a BE model rat with esophagoduodenal anastomosis, COX-2 expression was induced in the BE and in esophageal adenocarcinoma formed in the distal esophagus during bile reflux [52]. Therapeutic strategies for Barrett adenocarcinoma targeting COX-2 have been investigated. Selective inhibition of COX-2 by NS-398 decreased cell growth and increased the rate of apoptosis in human esophageal adenocarcinoma cell lines [53]. A COX-2 inhibitor was effective in preventing cancer in an animal model of BE and esophageal adenocarcinoma [54].

Stomach Gastric cancer is one of the most common and lethal malignancies worldwide. Histologically, human gastric carcinoma may be divided into the intestinal type and diffuse type. Pathogenesis of the intestinal-type cancer has been connected to precursor changes, such as chronic atrophic gastritis, intestinal metaplasia, and dysplasia. Factors associated with an increased risk of gastric cancer include smoking, H. pylori infection, high salt intake, and low consumption of raw vegetables and fresh fruit. Among them, chronic H. pylori infection is most important and is associated with the development of both intestinal and diffuse types of gastric cancer [55]. Prophylactic eradication of H. pylori after endoscopic resection for early gastric cancer reduced the risk of the subsequent development of metachronous gastric cancer [56].

Gastric Carcinogenesis and Duodeno-gastric Reflux In addition to the factors mentioned above, duodeno-gastric reflux has been considered to be involved in gastric carcinogenesis. In particular, the Billroth II procedure resulted in a higher incidence of subsequent cancer than the Billroth I procedure [57]. This is likely because patients usually have a larger amount of duodenal reflux containing bile in the gastric stump after Billroth II gastrectomy than do those after Billroth I. Among the various components of the duodenal reflux, bile acids seem the most important in gastrointestinal carcinogenesis. Rats treated with NMMG (N-methyl-N¢-nitro-N-nitrosoguanidine) and taurocholic acids had a higher incidence of neoplastic lesions in the stomach [58]. Furthermore, duodenal reflux induced gastric adenocarcinoma in rats with gastrojejunal stomata [59]. No cancer was seen in rats with pancreaticoduodenal reflux, whereas rats with bile reflux and

6 Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract

115

Fig. 6.2 Early gastric intestinal-type carcinoma in the pyloric area

rats with combined reflux had gastric carcinoma. Thus, bile acids, and not pancreaticoduodenal secretions, were related to gastric carcinogenesis in rats with reflux through the pylorus [60].

Gastric Carcinogenesis and COX-2 COX-2 is an important factor in gastric carcinogenesis. Normal gastric mucosa expresses COX-1, but COX-2 expression is very low. COX-2 is overexpressed by the neoplastic cells in intestinal-type gastric cancer [61–63]. COX-2 expression increased gradually during the sequential development of metaplastic epithelium from dysplasia to carcinoma in the stomach [64]. COX-2 overexpression was correlated with tumor invasion into the lymphatic vessels in the gastric wall, lymph node metastasis, and TMN staging in patients with gastric cancer [65]. H. pylori has been implicated as an inducer of COX-2 in the stomach. COX-2 was expressed in H. pyloriassociated gastritis. H. pylori induced COX-2 expression and enhanced PGE2 production in a human gastric carcinoma cell line [66–68]. COX-2 expression was observed in stump cancers and conventional primary gastric cancer. In addition to H. pylori infection, physiological duodeno-gastric reflux in the non-resected stomach led to increased expression of COX-2. The gastric pylorus is a common site for the occurrence of gastric cancer (Fig. 6.2). COX-2 expression was higher in early intestinal-type gastric cancers located in the gastric pylorus compared to tumors occurring in other area of stomach [69, Fig. 6.3]. Exposure to high bile acid concentrations seems to be an additional mechanism for COX-2 induction in the pyloric area because of its anatomical location. In accordance with these observations, bile acids induced COX-2 expression and enhanced PGE2 production in human gastric adenocarcinoma cell lines via the EGFR-p42/44MAPK pathways. Thus, duodeno-gastric reflux in the non-resected stomach could cause gastric carcinogenesis.

116

H. Yasuda and F. Itoh

Fig. 6.3 Immunohistochemical staining of COX-2 in early gastric carcinoma

Colorectal Cancer Colorectal cancer (CRC) is one of the three leading causes of cancer mortality worldwide. CRC may be sporadic, hereditary, or on a background of inflammatory bowel disease (IBD) [70]. The majority of sporadic CRCs occur in adenomas due to local imbalance of growth regulation arising out of alteration of proto-oncogenes, loss of tumor suppressor gene activity, and abnormalities in genes involved in DNA mismatch repair. In hereditary CRC syndromes such as familial adenomatous polyposis (FAP) and hereditary non-polyposis colon cancer (HNPCC), there are inherited genetic mutations in either tumor suppressor genes (APC gene in FAP) or genes associated with DNA mismatch repair function (hMSH genes in HNPCC). IBDassociated CRC is thought to represent an inflammation-dysplasia-carcinoma sequence [71].

Colon Cancer and Bile Acids Environmental factors are also important in human colorectal carcinogenesis. Dietary habits constitute a major environmental factor. The consumption of animal fat is positively related to the incidence of colon cancer. A high-fat and low-fiber diet tends to increase secretion of bile acid. The predominant secondary bile acids

6 Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract

117

formed in the human colon are the 7a-dehydroxylated products of deoxycholic acid and lithocholic acid. They are the main bile acids in human fecal water [72] and have been implicated in CRC. In animal experiments, taurodeoxycholic acid, lithocholic acid, deoxycholic acid, and chenodeoxycholic acid had a promoting effect in colon carcinogenesis induced in rats by MNNG [73, 74]. Cholic acid had a promoting effect in colon carcinogenesis induced by intrarectal administration of MNU (N-methyl-N-nitrourea). This promoting effect might be exerted through conversion of cholic acid to deoxycholic acid by the intestinal bacterial flora. Dietary administration of chenodeoxycholic acid increased tumors in the duodenum of ApcMIN/+ mice without standard carcinogen such as MNNG [75]. In a clinical study, the serum concentration of unconjugated secondary bile acids such as deoxycholic acid and lithocholic acid was elevated in patients with colorectal adenomas [76]. Colonic mucosal proliferation was related to serum deoxycholic acid levels but not to lithocholic, cholic, or chenodeoxycholic acid levels [77]. Not all bile acids seem to act as carcinogens. Ursodeoxycholic acid (UDCA), a tertiary bile acid, is a “good bile acid” and dissolves cholesterol gallstones and shows cytoprotective effects in primary biliary cirrhosis [78]. UDCA, with its equatorial 7b-hydroxyl group, is a poor detergent and has been shown to be cytoprotective. UDCA inhibited proliferation of CRC cell lines in vitro and protected against deoxycholic acid-induced apoptosis by stimulating Akt-dependent survival signaling in a human colon cancer cell line [79]. UDCA feeding reduced the size and number of colon tumors induced by NMU or azoxymethane and reduced fecal levels of cytotoxic deoxycholic acid in rats [80]. Ulcerative colitis (UC) is associated with an increased risk of developing CRC. UC often is associated with primary sclerosing cholangitis (PSC). UDCA, which was used initially because of its hepatoprotective effect for PSC, caused a marked decrease in the risk for the development of CRC or dysplasia in patients with UC and PSC [81]. UDCA treatment for primary biliary cirrhosis (PBC) was associated with decreases in the probability of recurrences of colorectal adenoma following colonoscopic removal [82]. In a larger study, it was shown that UDCA treatment after removal of colorectal adenoma was associated with a reduction in recurrence of adenomas with high-grade dysplasia [83].

Colorectal Cancer and COX-2 COX-2 has been studied intensively in colon carcinogenesis [84]. COX-2 was undetectable in the normal intestine, but its expression was increased in CRC [85, 86]. COX-2 was overexpressed in 50% of benign polyps and 80–85% of CRCs. Human CRC patients with COX-2 positive tumors showed augmented neovascularization and had a poorer prognosis than those with tumors negative for COX-2. They had pathologically unfavorable findings, such as tumor size, differentiation, number of metastatic lymph nodes, and Duke’s stage [87]. Among the various prostanoids produced by COX-2, PGE2 seems important in colon carcinogenesis. PGE2 increased the growth and motility of the LS-174 human colorectal carcinoma cell line via the

118

H. Yasuda and F. Itoh

PI3 kinase/AKT pathway [88]. Disruption of the gene encoding COX-2 in the APCD716 heterozygote, a mouse model of human FAP, markedly reduced the number and size of intestinal polyps [12]. In addition, disruption of EP2 reduced the number and size of adenomas in APCD716 mice, indicating that EP2 was the major receptor mediating the PGE2 signal in intestinal polyposis [89]. Of note, bile acids induced cell proliferation [90] and COX-2 expression in colon cancer cells. A liquid extract of human fecal water and deoxycholic acid induced COX-2 promoter activity in the CRC cell line HCT166 [22]. Epidemiologic research indicates that there is a 40–50% reduction in mortality from colorectal cancer in individuals who take aspirin or other NSAIDs on a regular basis. Patients with FAP have an almost 100% risk of CRC. Chemoprevention of CRC with sulindac or the COX-2 selective inhibitor celecoxib caused regression of colorectal adenomatous polyps in patients with FAP and can be considered practical in a very high-risk population [91–93].

Bile Acid Receptors and Gastrointestinal Carcinogenesis GPCR and EGFR Crosstalk and Carcinogenesis The EGFR and its family members play a pivotal role in tumor development, and their expression strongly affects the clinical outcome of cancer patients. EGFR is overexpressed frequently in gastrointestinal tumors where the expression levels correlate with decreased 5-year survival rates [94]. EGFR activation has been shown in various cancer cell lines, including colon, non-small cell lung cancer, head and neck squamous cell carcinoma (HNSCC), pancreatic cancer, breast cancer, and prostate cancer. EGFR is emerging as an important therapeutic target for several epithelial tumors. The EGF ligand family consists of EGF, heparin-binding EGF-like growth factor (HB-EGF), transforming growth factor (TGF)-a, epiregulin, amphiregulin, neuregulins, and betacellulin [95]. The phosphorylation cascade stimulated by EGFR includes phosphatidylinositol 3-kinase, MAPK, including p38MAPK, p42/44MAPK, c-jun-N-terminal kinases (JNK), and PKC. Various extracellular stimuli unrelated to EGFR ligands, such as agonists of GPCR, cytokines, and chemical and environmental stressors [96–99], are able to utilize the EGFR as a downstream signaling partner in the generation of mitogenic signals. Among them, GPCR-mediated EGFR activation has drawn much attention. GPCR-mediated activation of EGFR was described originally in Rat-1 fibroblasts. Stimulation of these cells with the GPCR ligands of lysophosphatidic acid (LPA), endothelin-1 (ET-1), and thrombin induced phosphorylation of EGFR and the subsequent expression of p42/44MAPK or c-fos gene [96]. Two mechanisms have been proposed for EGFR transactivation by GPCRs: intracellular, ligand-independent and extracellular, ligand-dependent mechanisms. Several intracellular molecules have been identified in the ligand-independent mechanism. Non-receptor tyrosine kinases of the Src family have been suggested as both upstream and downstream

6 Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract

119

mediators of the ligand-independent EGFR transactivation. Angiotensin II was reported to induce ligand-independent and Src-dependent EGFR activation in vascular smooth muscle cells [100]. Gbg subunits have been reported to mediate Srcdependent EGFR phosphorylation [101]. PKC also has been shown to be involved in EGFR transactivation. ET-1 mediated EGFR activation was PKC-dependent in rat cardiomyocytes [102]. The Ca2+-regulated tyrosine kinase Pyk2 has been shown to activate EGFR transactivation in PC12 cells [103]. The activation of EGFR and Pyk2 by Src following GPCR stimulation of fibroblasts from Src-, Pyk2-, and EGFR-deficient mice did not require EGFR, Src, and Pyk2 kinases to link GPCR with the MAP kinases signaling cascade [104]. In terms of a ligand-independent mechanism, metalloprotease has been shown to be involved in GPCR-mediated EGFR transactivation in a series of experiments using metalloprotease inhibitors. EGFR transactivation requires activation of a disintegrin and metalloproteinase (ADAM), which induce the shedding of EGF ligand ectodomains during their transition to soluble bioactive factors [105]. Many GPCR agonists, cytokines, and stressors appear to mediate EGFR transactivation by activating metalloprotease-dependent HB-EGF shedding [106, 107]. GPCR-mediated EGFR transactivation has been reported in various cancer cells. For instance, LPA stimulated cell proliferation and EGFR transactivation in HNSCC cells in an MMP-dependent manner [108]. LPA stimulated EGFR in human colon cancer cells. LPA interacts with at least three GPCR, LPA-1, -2, and -3, although LPA-2 was overexpressed in colon cancer tissue [109]. In addition, PGE2 activated both EGFR and c-Met receptor and resulted in b-catenin and increased expression of the urokinase-type plasminogen activator receptor (uPAR) in human colon cancer cell lines [110]. ET-1 increased DNA replication in colorectal cancer cells via the ETA receptor. This action was mediated via the pertussis toxin-sensitive G-proteins, PI3K, PKC, and transactivation of EGFR [111]. Interleukin-8 (IL-8), a member of the chemokine superfamily that binds to GPCR IL-8RA and IL8RB, was shown to activate EGFR via the release of HB-EGF and to promote cell proliferation in the human colon cancer cell line Caco-2 [112]. On the other hand, the proteinase-activated receptor (PAR)-1 is a predominant receptor for thrombin. PAR-1 is absent from normal human colon epithelial cells but aberrantly expressed during colonic carcinogenesis [113]. Activation of PAR-1 by thrombin induced matrix metalloprotease (MMP)-dependent release of TGF-a, transactivation of EGFR, and subsequent activation of p42/p44MAPK in the human colon cancer cell line HT-29 [114].

Bile Acid Receptors and EGFR Transactivation Several reports have suggested that the cellular effects of bile acids involve activation of MAPK and EGFR. It was reported originally that deoxycholic acid induced the rapid activation of p42/44MAPK and EGFR in rat primary hepatocytes [115]. Moreover, it was reported that bile acids induced EGFR phosphorylation and enhanced COX-2 expression in a human cholangioma cell line. Bile acid-induced

120

H. Yasuda and F. Itoh

EGFR phosphorylation was associated with subsequent activation of p42/44MAPK, p38MAPK, and JNK. Src kinase activity was required for EGFR phosphorylation. Bile acid-induced COX-2 expression was dependent on the p42/44MAPK and p38MAPK pathways in these cells [116]. Although it was originally reported that bile acidinduced EGFR activation in hepatocytes was ligand independent [115], bile acids stimulate MMP-3-mediated release of TGF-a with consequent activation of EGFR in human cholangiocyte cell lines [117].

Bile Acids as Ligands for GPCR The nuclear receptor FXR was identified first as a bile acid receptor and has been studied widely. Recently, GPCR type bile acid receptors have been identified, and their role in gastrointestinal carcinogenesis has been investigated.

M3 Muscarinic Receptor and EGFR Activation It was reported originally that bile acids caused a rapid increase in cytosolic calcium in isolated rat hepatocytes [118]. Lithocholic acid and the conjugates taurolithocholic acid and taurolithocholic acid sulfate caused release of calcium from an Ins(1,4,5)P3-sensitive intracellular store [119]. Taurine- and glycine-conjugated bile acids, such as lithocholyltaurine and lithocholylglycine, have a similar molecular structure to acetylcholine. They mimic the action of acetylcholine and have been reported to activate the M3 muscarinic receptor and to transactivate EGFR [120– 122]. Of note, the M3 muscarinic receptor agonist carbachol stimulated EGFR transactivation and mediated MMP-mediated cleavage of TGF-a in T84 human colonic epithelial cells [123]. In studies using colon cancer cell lines, bile acids stimulated cell proliferation by a mechanism involving matrix metalloproteinase 7 (MMP-7)mediated release of an EGFR ligand and subsequent post-EGFR cell signaling [124]. In clinical studies, expression of MMP-7 (matrilysin) has been detected in SCC of the esophagus, neck, and lung and in adenocarcinomas of the breast, endometrium, prostate, stomach, and colon, which increased progressively with more advanced grades of colonic dysplasia and cancer [125]. Loss of matrilysin-deficiency in ApcMin/+ mice led to a 60% reduction in the number of tumors formed, along with a significant decrease in tumor size [126].

TGR5 and EGFR Transactivation A novel GPCR known as membrane-type bile acid receptor (M-BAR), or TGR5 [7, 8], was identified in a search of the GenBankTM data base for ligands for the orphan GPCRs. TGR5 has 30%, 29%, 26%, and 26% amino acid identity with EDG (sphingosine-1-phosphate receptor)-6, EDG-8, EDG-1, and EDG-7, respectively, and is a

6 Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract

121

member of the class 1 (rhodopsin-like) GPCR superfamily. Taurolithocholic acid increased [35S]GTPgS binding in membrane fractions from TGR5-transfected cells. Treatment of CHO cells expressing TGR-5 with bile acids induced intracellular cAMP production and activation of p42/44MAPK. The rank of order of potency of cAMP production in TGR5-transfected cells was lithocholic acid > deoxycholic acid > chenodeoxycholic acid > cholic acid. Their EC50 values were 0.035, 0.575, 4.0, >10 mM, respectively. Their taurine- or glycine-conjugated forms or UDCA also was active. The absence of a change in intracellular Ca2+ following stimulation of TGR5-transfected cells with bile acid suggested Gs, not to Gq or Gi, coupling to TGR5. Expression of TGR5 has been detected in most human tissues, including stomach, duodenum, ileum, jejunum, and colon, but not esophagus and rectum. TGR5 mRNA also is expressed in monocytes/macrophages. TGR5 also was expressed in sinusoidal endothelial cells in the rat liver [127]. Several metabolic roles of TGR5 have been explored. TGR5 was involved in bile acid-induced GLP secretion from intestinal neuroendocrine cells [128]. Recently, bile acids were shown to influence energy consumption in brown adipose tissue through the TGR5/cAMP pathway [129]. In addition, TGR5 seems to play a critical role in maintenance of bile acid homeostasis and cholesterol metabolism. The total bile acid pool size was decreased significantly in TGR5 disrupted mice. A high-fat diet led to fat accumulation with body weight gain in these mice [130]. They did not form cholesterol gallstones when fed a cholic acid-containing high-fat diet [131]. Recently, it was shown that TGR5 is involved in bile acid-induced transactivation of EGFR (Fig. 6.4). In AGS human gastric carcinoma cells, siRNA-mediated down-regulation of TGR5 was associated with suppression of deoxycholic acidinduced transactivation of EGFR [132]. In addition, siRNA-mediated down-regulation of ADAM-17 suppressed deoxycholic acid-induced transactivation of EGFR. Activation of ADAM-17 led to shedding of HB-EGF ectodomains, which activated EGFR. EGFR transactivation protected these cells partially from deoxycholic acidinduced apoptosis. Upregulated expression of ADAM-17 and EGFR was observed in human colon cancer tissue [133]. The TGR5–ADAM17–EGFR pathway seems to participate in the genesis and apoptosis of gastric carcinoma cells and thus may serve as a target for cancer prevention.

FXR and Colon Carcinogenesis In contrast to the tumorigenic roles of bile acid-induced GPCR activation, evidence from experiments using knockout mice [134] suggested a protective role of the bile acid nuclear receptor FXR in intestinal carcinogenesis. In a clinical study of FAP patients, FXR expression was decreased in tumors compared to normal mucosa. Increased colonic cell proliferation and increased prevalence and size of azoxymethane-induced adenocarcinoma were observed in the colons of FXRdeficient mice [135]. Moreover, loss of FXR in the ApcMin/+ mice resulted in early mortality and increased tumor progression by promotion of Wnt signaling [136]. FXR seems to protect against colon carcinogenesis in several ways. Firstly, mice

122

H. Yasuda and F. Itoh

Fig. 6.4 Effect of deoxycholic acid on the subcellular localization of EGFR in human gastric adenocarcinoma AGS cells. Cells were incubated with (lower) or without (upper) 50 mM deoxycholic acid for 5 min, and EGFR immunofluorescence was examined using a confocal laser microscope

lacking FXR expression have elevated levels of serum bile acids and an increase in the bile acid pool size [137]. Of note, FXR-deficient mice developed hepatocellular adenoma and carcinoma spontaneously after 12–15 months of age [138, 139]. A major role of FXR seems to protect the liver and intestine from the deleterious effect of bile acid overload by decreasing their endogenous production and by accelerating biotransformation and excretion. Secondly, FXR is important in protecting against colonic mucosal inflammation. Chronic inflammation is an important factor in colon carcinogenesis in patients with IBD. Mice lacking FXR had increased ileal levels of bacteria and a compromised epithelial barrier. Notably, FXR synthetic agonist GW4064 was able to reverse intestinal bacterial overgrowth following surgical

6 Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract

123

biliary obstruction in wild type but not in FXR-deficient mice. [140]. Thirdly, FXR is essential for glucose metabolism. Glucose was shown to regulate the expression of FXR, and streptozotocin-induced diabetes results in a down-regulation of hepatic FXR gene expression [141]. FXR-deficient mice developed severe fatty liver and elevated circulating free fatty acids, which was associated with impaired glucose tolerance and insulin resistance. In these mice, insulin signaling was impaired in peripheral insulin-sensitive tissue, such as skeletal muscle and white adipose tissue [142, 143]. Adiponectin is a key hormone responsible for insulin sensitization [144]. Reduced plasma levels of adiponectin have been reported to correlate with the risk of CRC [145]. When azoxymethane treated adiponectin-deficient mice were fed with a high-fat diet, increased numbers of colon polyps and reduced survival were observed [146]. It is likely that dysregulation of FXR expression in a diabetic and insulin resistant state may contribute to the increased incidence of CRC observed in the obese population. Collectively, FXR seems to protect against colon carcinogenesis by maintaining both bile acids and glucose homeostasis. There is a potential chemopreventive role for a selective FXR agonist.

Conclusions Several factors, including environmental, genetic, and epigenetic factors, are involved in gastrointestinal carcinogenesis. Among them, bile acids are important as luminal and endocrine factors through GPCRs and nuclear receptor pathways. In the upper gastrointestinal tract, bile acids in duodenal refluxate induce COX-2 expression by the esophageal and gastric epithelium during duodeno-gastric-esophageal reflux. A high-fat and low-fiber diet increases secretion of bile acids and is related to the incidence of CRC. Bile acids activate EGFR and MAPK, and induce COX-2 expression, in gastrointestinal cells via GPCR. GPCR may serve as a target for cancer prevention. In contrast to GPCRs, FXR protects against colon carcinogenesis by modulating glucose and bile acid homeostasis.

References 1. Russell, D.W., 2003, Ann. Rev. Biochem., 72, 137–174. 2. Makishima, M., Okamoto, A.Y., Repa, J.J., Tu, H., Learned, M., Luk, A., Hull, M.V., Lustig, K.D., Mangelsdorf, D.J., Shan, B., 1999, Science, 284, 1362–1365. 3. Park, D.J., Blanchard, S.G., Bledsoe, R.K., Chandra, G., Consler, T.G., Kliewer, S.A., Stimmel, J.B., Willson, T.M., Zavacki, A. M., Moore, D.D., Lehmann, J.M., 1999, Science, 284, 1365–1368. 4. Wang, H., Hung, J., Hollister, K., Sowers, L.C., Forman, B.M., 1999, Mol. Cell, 17, 683–694. 5. Lefebvre, P. Cariou, B., Lien, F., Kuipers, F., Staels, B., 2009, Physiol. Rev., 89, 147–191.

124

H. Yasuda and F. Itoh

6. Choi, M., Moschetta, A., Bookout A.L., Peng, L., Umetani, M., Holmstrom, S.R., Suino-Powell, K., Xu, H.E., Richardson, J.A., Gerard, R.D., Mangelsdorf, D.J., Kliewer, S.A., 2006, Nature Med., 12, 1253–1255. 7. Maruyama, T., Miyamoto, Y., Nakamura, T., Tamai, Y., Okada, H., Sugiyama, E., Nakamura, T., Itadani, H., Tanaka, K., 2002, Biochem. Biophys. Research Comm., 298, 714–719. 8. Kawamata, Y., Fujii, R., Hosoya, M., Harada, M., Yoshida, M., Miwa, M., Fukusumi, S., Habata, Y., Itoh, T., Shintani, Y., Hinuma, S., Fujisawa, Y., Fujino, M., 2003, J. Biol. Chem. 278, 9435–9440. 9. Cook, J.W., Kennaway, E.L., Kennaway, N.M., 1940, Nature, 3647, 627. 10. Bernstein, H., Bernstein, C., Payne, C.M., Dvorakova, K., Garewal, H., 2005, Mutation Research, 589, 47–65. 11. Williams, C.S., Mann, M., Dubois, R.N., 1999, Oncogene, 18, 7908–7916. 12. Oshima, M., Dinchuk, J.E., Kargman, S.L., 1996, Cell, 87, 803–809. 13. Iniguez, M.A., Punzon, C., Fresno, M., 1999, J. Immunol., 163, 111–119. 14. Vane, J.R., 1971, Nat. New Biol., 231, 232–235. 15. Narumiya, S., Sugimoto, Y., Ushikubi, F., 1999, Physiol. Rev., 79, 1193–1221. 16. Pai, R., Soreghan, B., Szabo, I.L., Pavelka, M., Baatar, D., S. Tarnawski, A.S., 2002, Nature Med., 8, 289–293. 17. Phipps, R.P., Stein, S.H., Roper, R.L., 1991, Immunol. Today, 12, 349–352. 18. Tsujii, M., Kawano, S., Tsuji, S,. Sawaoka, H., Hori, M., DuBois, R.N., 1998, Cell, 93, 705–716. 19. Pai, R., Szabo, I.L., Soreghan, B.A., Atay, S., Kawanaka, H., Tarnawski, A.S., 2001, Biochem. Biophys. Res. Commun., 286, 923–928. 20. Tsujii, M., Dubois, R.N., 1995, Cell, 83, 493–501. 21. Zhang, F., Subbaramaiah, K., Altorki, N., Dannenberg, A.J., 1998, J. Biol. Chem. 273, 2424–2428. 22. Glinghammar, B., Rafter, J., 2001, Gastrotenterol., 120, 401–410. 23. Zhang, F., Altorki, N.K., Wu, Y.-C., Soslow, R.A., Subbaramaiah, K., Dannenberg AJ., 2001, Gastroenterol., 121, 1391–1399. 24. Craven, P.A., Pfanstiel, J., DeRubertis, F.R., 1987, J. Clin. Invest., 79, 532–541. 25. Guillem, J.G., O’Brian, C.A., Fitzer, C.J., Johnson, M.D., Forde, K.A., LoGerfo, P., Weinstein, I.B., 1987, Arch. Surg., 122. 1475–1478. 26. Jones, B.A., Rao, Y.P., Stravitz, R.T., Gores, G.J., 1997, Am. J. Physiol., 272, G1109–1115. 27. Maeta, M., Koga, S., Shimizu, T., Matsui, K., 1990, J. Surgical Oncol., 44, 20–24. 28. Kitabayashi, K., Nakano, Y., Saito, H., Ueno, K., Kita, I., Takashima, S., Kurose, N., Nojima, T., 2001, Surgery Today, 31, 670–674. 29. Miwa, K., Hasegawa, H., Fujimura, T., Matsumoto, H., Miyata, R., Kosaka, T., Miyazaki, I., Hattori, T., 1996, Int. J. Cancer, 67, 269–274. 30. Fitzgerald, R.C., 2006, Gut, 55, 1810–1818. 31. Kauser, W.K.H., Peters, J.H., DeMeester, T.R., Feussner, A.P., Ireland, H.J., Stein, R.J., Siewert, R.J., 1997, Surgery, 122, 874–881. 32. Stadler J., Stern, H.S., Yeung, K.S., McGuire, V., Furrer, R., Marcon, N., Bruce, W.R., 1988, Gut, 29, 1326–1331. 33. Nehra, D., Howell, P., Williams, C.P., Pye, J.K., Beynon, J., 1999, Gut, 44, 598–602. 34. Menges, M., Müller, M., Zeitz, M., 2001, Am. J. Gastroenterol., 96, 331–337 35. Jaiswal, K., Lopez-Guzman, C., Souza, R.F., Spechler, S.J., Sarosiet, G.A., 2006, Am. J. Physiol., 290, G335–G342. 36. Eda, A., Osawa, H., Satoh, K., Yanaka, I., Kihira, K., Ishino, Y., Mutoh, H., Sugano, K., 2003, J. Gastroenterol., 38, 14–22. 37. Wong, N.S.C.S., Wilding, J., Bartlett, S., Liu, Y., Warren, B.F., Piris, J., Marynard, N., Marshall, R., Bodmer, W.F., 2005, Proc. Natl. Acad. Sci., 102, 7565–7570. 38. Kazumori, H., Ishihara, S., Rumi, M.A.K., Kadowaki, Y., Kinoshita, Y., 2006, Gut, 55, 16–25. 39. Kazumori, H., Ishihara, S., Kinoshita, Y., 2009, Gut, 58, 620–628.

6 Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract

125

40. Silberg, D.G., Sullivan, J.S., Kang, E., Swain, G.P., Moffett, J., Sund, N.J., Sackett, S.D., Kaestiner, K.H., 2002, Gastrotenterol., 122, 689–696. 41. Mutoh, H., Hakamata, Y., Sato, K., Eda, A., Yanaka, I., Honda, S., Osawa, H., Kaneko, Y., Sugano, K., 2002, Biochem. Biophys. Res. Commun., 294, 470–479. 42. Villanacci, V., Rossi, E., Zambelli, C., Galletti, A., Cestari, R., Missale, G., D. Della Casa, D., Bassott, G., 2007. Digestive Liver Disease, 39, 305–311. 43. Liu, Q., Teh, M., Ito, K., Shah, N., Ito, Y., Yeoh, K.G., 2007, Modern Pathology 20, 1286–1297. 44. Vidal Mallo, G., Soubeyran, P., Lissitzky, J.C., André, F., Farnarier, C., Marvaldi, J., Dagorn, J.C., Iovanna, J.L., 1998, J. Biol. Chem., 273: 14030–14036. 45. Chawengsaksophak, K., James, R., Hammond, V.E., Köntgen, F., Beck, F., 1997, Nature, 386, 84–87. 46. Aoki, K., Tamai, Y., Horiikem S., Oshima, M., Taketo, M.M., 2003, Nature Genet., 35, 323–330. 47. Kim, S.-P., Park, J.-W., Lee, S.-H., Lim, J.H., Jang, B.-C., Lee, S.-H., Jang, I.-H., Lee, S.-H., Jang, I.-H., Freund, J.-N., Suh, S.-I., Mun, H.C., Song, D.-K., Ha, E.-M., Lee, W.-J., Kwon, T.K., 2004, Biochem. Biophys. Re. Comm., 315, 93–99. 48. Mutoh, H., Hayakawa, H., Sakamoto, H., Sugano, K., 2007, J. Gastroenterol., 42, 719–729. 49. Shirvani, V.N., Ouatu-Lascar, R., Kaur, B.S., Omary, M.B., Triadafilopoulos, G., 2000, Gastroenterol., 118, 487–496. 50. Buskens, C.J., van Rees, B.P., Sivula, A., Reitsma, J.B., Haglund, C., Bosma, P.J., Offerhaus, G.J., van Lanschot, J.J.B., Ristimäki, A., 2002, Gastroenterol., 122, 1800–1807. 51. Soma, T., Kaganoi, J., Kawaba, A., Kondo, K., Tsunoda, S., Imamura, M., Shimada, Y., 2006, Int. J. Cancer, 119, 771–782. 52. Song, S., Guha, S., Liu, G., Buttar, N.S., Bresalier, R.S., 2007, Gut, 56, 1512–1521. 53. Souza, R.F., Shewmake, K., Beer, D.G., Cryer, B., Spechler, S.J., 2000, Cancer Res., 60, 5767–5772. 54. Buttar, N.S., Wang, K.K., Leontovich, O., Westcott, J.Y., Pacifico, R.J., Anderson, M.A., Krishnadath, K.K., Lutzke, L.S., Burgart, L.J., 2002, Gastrotenterol., 122, 1101–1112. 55. Uemura, N., Okamoto, S., Yamamoto, S., Matsumura, N., Yamaguchi, S., Yamakido, M., Taniyama, K., Sasaki, N., Schlemper, R., 2001, N. Engl. J. Med., 345, 784–789. 56. Fukase, K., Kato, M., Kikuchi, S., Inoue, K., Uemura, N., Okamoto, S., Terao, S., Amagai, K., Hayashi, S., Asaka, M., for the Japan Gast Study Group, 2008, Lancet, 372, 392–397. 57. Lundegardh, G., Adami, H.O., Helmick, C., Zack, M., Meirik, O., 1988, N. Engl. J. Med., 319, 195–200. 58. Kobori, O., Shimizu, T., Maeda, M., Atomi, Y., Watanabe, J., Shoji, M., Morioka, Y., 1984, J. Natl. Cancer Inst., 73, 853–861. 59. Miwa, K., Hasegawa, H., Fujimura, T., Matsumoto, H., Miyata, R., Kosaka, T., Miyazaki, I., Hattori, T., 1992, Carcinogenesis 13, 2313–2316. 60. Miwa, K., Fujimura, T., Hasegawa, H., Kosaka, T., Miyata, R., Miyazaki, I., Hattori, T., 1992, J. Cancer Res. Clin. Oncol., 118, 1570–4. 61. Ristimäki, A., Honkanen, N., Jänkälä, H., Sipponen, P., Härkönen, M., 1997, Cancer Res., 57, 1276–1280. 62. Saukkonen, K., Nieminen, O., van Rees, B., Vilkki, S., Härkönen, M., Juhola, M., Mecklin, J.P., Sipponen, P., Ristimäki, A., 2001, Clin. Cancer Res., 7, 1923–1931. 63. Saukkonen, K., Rintahaka, J., Sivula, A., Buskens, C.J., van Rees, B.P., Rio, M.C., Haglund, C., van Lanschot, J.B., Offerhaus, G.J.A., Ristimaki, A., 2003, APMIS 111, 915–925. 64. van Rees, B.P., Saukkonen, K., Ristimaki, A., Polkowski, W., Tytgat, G.N.J., Drillenburg, P., Offerhaus, J.A., 2002, J. Pathology, 196,171–179. 65. Murata, H., Kawano, S., Tsuji, S., Tsujii, M., Sawaoka, H., Kimura, Y., Shiozaki, H., Hori, M., 1999, Am. J. Gastrotenterol., 94, 451–455. 66. Romano, M., Vittorio, R,. Memoli, A., Tuccilli, C., Di Popolo, A., Sommi, P., Acquaviva, A.M., Blanco, C.D.V., Bruni, C.B., Zarrilli, R., 1998, J. Biol. Chem., 273, 28560–28563.

126

H. Yasuda and F. Itoh

67. Fu, S., Ramanujam, K.S., Wong, A., Wong, A., Fantry, G.T., Drachenberg, C.B., James, S.P., Meltzer, S.J., Wilson, K.T., 1999, Gastroenterol., 116, 1319–1329. 68. McCarthy, C.J., Crofford, L.J., Greenson. J., Scheiman, J.M., 1999, Am. J. Gastroenterol., 94, 1218–1223. 69. Yasuda, H., Yamada, M., Endo, Y., Inoue, K., Yoshiba, M. 2005, J. Gastroenterol., 40, 690–697. 70. Kinzler, K.W., Vogelstein, B., 1996, Cell, 87, 159–170. 71. Itzkowitz, S.H., Yio, X., 2004, Am. J. Pysiol., 287, G7-G17. 72. De Kok, T.M.C.M., van Faassen, A., Glinghammar, B., Pachen, D.M.F.A., Rafter, J.J., Baeten, C.G.M.I., Engels, L.G.J.B., Kleinjans, J.C.S., 1999, Digest. Dis. Sci., 44, 2218–2225. 73. Reddy, B.S., Narasawa, T., Weisburger, J.H., Wynder, E.L., 1976, J., Natl., Cancer Inst., 56, 441–442. 74. Reddy, B.S., Watanabe, K., Weisburger, J.H., Wynder, E.L., 1977, Cancer Research, 37, 3238–3242. 75. Mahmoud, N.N., Dannenberg, A.J., Bilinski, R.T., Mestre, J.R., Chadburn, A., Churchill, M., Martucci, C., Bertagnolli, M.M., 1999. Carcinogenesis, 20. 299–303. 76. Bayerdörffer, E., Mannes, G.A., Ochsenkühn, T., Dirschedl, P., Wiebecke, B., Paumgartner, G., 1995, Gut, 36, 268–273. 77. Ochsenkühen, T., Bayerdörffer, E., Meining, A., Schinkel, M., Thiede, C., Nüssler, V., Sackmann, M., Hatz, R., Neubauer, A., Paumgartner, G., 1999, Cancer, 85, 1664–1669. 78. Rubinm R.A., Kowalski, T.E., Khandelwal, M., Malet, P.F., 1994, Ann. Intern. Med. 121, 207–218. 79. Im, E., Akare, S., Powell, A.A., Martinez, J.D., 2005, Nutr. Cancer., 51, 110 – 116. 80. Batta, A.K., Salen, G., Holubec, H., Brasitus, T.A., Alberts, D., Earnest, D.L., 1998, Cancer Research, 58, 1684–1687. 81. Pardi, D.S., Loftus Jr., E.V., Kremers, W.K., Keach, J., Lindor, K.D., 2003, Gastroenterol., 124, 889–893. 82. Serfaty, L., De Leusse, A., Rosmordiuc, O., Desalint, B., Flejou, J.F., Chazoulleres, O., Poupon, R.E., Poupon, R., 2003, Hepatology, 38, 203–209. 83. Alberts, D.S., Martìnez, M.E., Hess, L.M., Einspahr, J.G., Green, S.B., Bhattacharyya, A.K., Guillen, J., Krutzsh, M., Batta, A.K., Salen, G., Fales, L., Koonce, K., Parish, D., Clouser, M., Roe, D., Lance, P. 2005, J. Natl. Cancer Inst., 97, 846–853. 84. Marnett, L.J., Dubois, R.N., 2002, Annu. Rev. Pharmacol. Toxicol.,42, 55–80. 85. Eberhart, C. E., Coffey, R. J., Radhika, A., Giardiello, F. M., Ferrebach, S., Dubois, R. N., 1994, Gastroenterol., 107: 1183–1188. 86. Sano, H., Kawahito, Y., Wilder, R.L., Hashiramoto, A., Mukai, S., Asai, K., Kimura, S., Kato, H., Kondo, M., Hla, T., 1995, Cancer Res., 55, 3785–3789. 87. Masunaga, R., Kohno, H., Dhar, D.K., Ohno, S., Shibakita, M., Kinugasa, S., Yoshimura, H., Tachibana, M., Kubota, H., Nagasue, N., 2000, Clin. Cancer Res., 6, 4064–4068. 88. Sheng, H., Shao, J., Washington, M.K., Dubois, R.N., 2001, J. Biol. Chem., 276, 18075–18081. 89. Sonoshita, M., Takazu, K., Sasaki, N., Sugimot, Y., Ushikubi, F., Narumiya, S., Oshima, M., Taketo, M.M., 2001, Nature Med., 7, 1048–1051. 90. Pai, R., Tarnawski, A.S., Tran, T., 2004, Mol. Biol. Cell, 15, 2156–2163. 91. Giardiello, F.M., Hamilton, S.R., Kruch, A.J., Piantadosi, S., Hylind, L.M., Celano, P., Booker, S.V., Robinson, C.R., Offerhaus, J.A., 1993, N. Engl. J. Med., 328, 1313–1316. 92. Steinbach, G., Lynch, P.M., Phillips, R.K.S., Wallace, M.H., Hawk, E., Gordon, G.B., Wakabayashi, N., Saunders, B., Fujimura, T., Su, L.K., Levin, B., 2000, N. Engl. J. Med., 342, 1946–1952. 93. Das, D., Arber, N., Jankowski, J.A., 2007, Digestion, 76, 51–67. 94. Sebastian, S., Settleman, J., Reshkin, S.J., Azzariti, A., Bellizzi, A., Paradiso, A., 2006, Biochim. Biophys. Acta, 1766, 120–139 95. Yarden, Y., MSliwkowski, M.X., 2001, Nat. Rev. Mol. Cell Biol. 2, 127–137. 96. Daub, H., Weuss, F.U., Wallasch, C., Ullrich, A., 1996, Nature 379, 557–560.

6 Role of Bile Acids in Carcinogenesis of Gastrointestinal Tract

127

97. Carpenter, G., J. Cell Biol., 146 (1999) 697–702. 98. Reardon, D.B., Contessa, J.N., Mikkelsen, R.B., Valerie, K., Amir, C., Dent, P., SchmidtUllrich, R.K., 1999, Oncogene,18, 4756–4766. 99. Warmuth, I., Harth, Y., Matsui, M.S., Wang, N., DeLeo, V.A., 1994, Cancer Res., 54, 374–376. 100. Bokemeyer, D., Schmitz, U., Kramer, H.J., 2000, Kidney Int., 58, 549–558. 101. Luttrell, L.M., Della Rocca, G.J., van Biesen, T., Luttrell, D.K., Lefkowitz, R.J., 1997, J. Biol. Chem., 272, 4637–4633. 102. Kodama, H., Fukuda, K., Takahashi, T., Sano, M., Kato, T., Tahara, S., Hakuno, D., Sato, T., Manabe, T., Konishi, F., Ogawa, S., 2002, J. Molecular Cellular Cardiol., 34, 139–150. 103. Soltoff, S.P., 1998, J. Biol. Chem., 273, 23110–23117. 104. Andreev, J., Galisteo, M.L., Kranenburg, O., Logan, S.K., Chiu, E.S., Okigaki, M., Cary, L.A., Moolenaar, W.H., Schlessinger, J., 2001, J. Biol. Chem., 276, 20130–20135. 105. Prenzel, M., Zwick, E., Daub, H., Leserer, M., Abraham, R., Wallasch, C., Ullrich, A., 1999, Nature, 402, 884–888. 106. Mifune, M., Ohtsu, H., Suzuki, H., Nakashima, H., Brailoiu, E., Dun, N.J., Frank, G.D., Inagami, Higashiyama, T.S., Thomas, W.G., Eckhart, A.D., Dempsey, P.J., Eguchi, S., 2005, J. Biol. Chem. 280, 26592–26599. 107. Higashiyama, S., Nanba, D., 2005, Biochim. Biophys. Acta., 1751, 110–117. 108. Gschwind, A., Prenzel, N., Ullrich, A., 2002, Cancer Research, 62, 6329–6336. 109. Shida, D., Watanabe, T., Aoki, J., Hama, K., Kitayama, J., Sonoda, H., Kishi, Y., Yamaguchi, H., Sasaki, S., Sako, A., Konishi, H., Nagawa, H., 2004, Lab. Invest., 84, 1352–1362. 110. Pai, R., Nakamura, T., Moon, W.S., Tarnawski, A.S., 2003, FASEB J., 17, 1640–1647. 111. Grant, K., Knowles, J., Dawas, K., Burnstock, G., Taylor, I., Loizidou, M. 2007, Br. J. Surgery, 94, 106–112. 112. Itoh, Y., Joh, T., Tanida, S., Sasaki, M., Kataoka, H., Itoh, K., Oshima, T., Ogasawara, N., Togawa, S., Wada, T., Kubota, H., Mori, Y., Ohara, H., Nomura, T., Higashiyama, S., Itoh, M., Cytokine, 29, 275–282. 113. Darmoul, D., Gratio, V., Devaud, H., Lehy, T., Laburthe, M., 2003, Am. J. Pathol., 162, 1503–1513. 114. Darmoul, D., Gratio, V., Devaud, H., Peiretti, F., Laburthe, M., 2004, Mol. Cancer Res., 2. 514–522. 115. Quao, L., Studer, E., Leach, K., McKinstry, R., Gupta, G., Decker, R., Kukreja, R., Valerie, K., Nagarkatti, P., Deiry, W.E.D., Molkentin, J., Schmidt-Ullrich, R., Fisher, P.B., Grant, S., Hylemon, P.B., Dent, P., 2001, Mol. Biol. Cell, 12, 2629–2645. 116. Yoon, J.H., Higuchi, H., Werneburg, N.W., Kaufman, S.H., Gores, G.J., 2002, Gastroenterol., 122, 985–993. 117. Werneburg, W.W., Yoon, J.H., Higuchi, H., Gores, G.J., 2003, Am. J. Physiol., 285, G31G36. 118. Anwer, M.S., Engelking, L.R., Nolan, K., Sullivan, D., Zimniak, P., Lester, R., 1988, Hepatology, 8, 887–891. 119. Combettes, L., Berthon, B., Doucet, E., erlinger, S., Claret, M., 1989, J. Biol., Chem., 264, 157–167. 120. Raufman, J.-P., Zimniak, P., Bartoszko-Malik, A., 1998, Am. J. Physiol., 274, G997-G1004. 121. Raufman, J.P., Cheng, K., Zimnial, P., 2003, Digestive Disease Sciences, 48,1431–1444. 122. Cheng, K., JRaufman, J.P., 2005, Biochem. Pharmacol., 70, 1035–1047. 123. McCole, D.F., Keely, S.J., Coffey, R.J., Barrett, K.E., 2002, J. Biol. Chem., 277, 42603–42612. 124. Cheng, K., Xie, G., Raufman, J.P., 2007, Biochem. Pharmacol., 73, 1001–1012. 125. Shattuck-Brandt, R.L., Lamps, L.W., Heppner Goss, K.J., DuBois, R.N., Matrisian, L.M., 1999, Mol. Carcinogenesis, 24, 177–187. 126. Wilson, C.L., Heppner, K.J., Labosky, P.A., Hogan, B.L., Matrisian, L.M., 1997, Proc. Natl. Acad. Sci. USA, 94, 1402–1407.

128

H. Yasuda and F. Itoh

127. Keitel, V., Reinehr, R., Gatsios, P., Rupprecht, C., Göerg, B., Häussinger, D., Kubitz, R., 2007, Hepatology, 45, 695–704. 128. Katsuma, S., Hirasawa, A., Tsujimoto, G., 2005, Biochem. Biophys. Res. Comm., 329, 386–390. 129. Watanabe, M., SHouten, S.M., Mataki, C., Christoffolete, M.A., Kim, B.W., Sato, H., Messaddeq, N., Harney, J.W., Ezaki, O., Kodama, T., Schoonjans, K., ABianco, A.C., Auwerx, J., 2006, Nature, 439, 484–489. 130. Maruyama, T., Tanaka, K., Suzuki, J., Miyoshi, H., Harada, N., Nakamura, T., Miyamoto, Y., Kanatani, A., Tamai, Y. 2006, J. Endocrinol., 191, 197. 131. Vassileva, G., Golovko, A., Markowitz, L., Abbondanzo, S.J., Zeng, M., Yang, S., Hoos, L., Tetzloff, G., Levitan, D., Murgolo, N.J., Keane, K., Davis, H.R., Hedrick, J., Gustafson, E.L., 2006, Biochem. J., 398, 423. 132. Yasuda, H., Hirata, S., Inoue, K., Mashima, H., Ohnishi, H., Yoshiba, M. 2007, Biochem. Biophys. Res. Commun., 354: 154–159. 133. Blanchot-Jossic, F., Jarry, A., Masson, D., Bach-Ngohou, K., Paineau, J., Denis, M., Laboisse, C., Mosnier, J.F., 2005, J. Pathology, 207, 156–163. 134. Sinal, C.J., Tohkin, M., Miyata, M., Ward, J.M., Lambert, G., Gonzalez, F.J., 2000, Cell, 102, 731–744. 135. Maran, R.R., Thomas, A., Roth, M., Sheng, Z., Esterly, N., Pinson, D., Gao, X., Zhang, Y., Ganapathy, V., Gonzalez, F.J., Guo, G.L., 2009, J. Pharmacol. Exp. Ther., 328, 469–477, 136. Modica, S., Murzilli, S., Salvatore, L., Schmidt, D.R., Moschetta, A., 2008, Cancer Res., 68, 9589–9594. 137. Kok, T., Hulsebos, C.V., Wolter, H., Havinga, R., Agellon, L.B., Stellaard, F., Shan, B., Schwarz, M., Kuipers, F., 2003, J. Biol. Chem. 278, 41930–41937. 138. Kim, I., Morimura, K., Shah, Y., Yang, Q., Ward, J.M., Gonzalez, F.J., 2007, Carcninogenesis, 28, 940–946. 139. Yang, F., Huang, X., Yi, T., Yen, Y., Moore D.D., Huang, W., 2007, Cancer Res, 67, 863–867. 140. Inagaki, T., Moschetta, A., Lee, Y.K., Peng, L., Zhao, G., Downes, M., Yu, R.T., Shelton, J.M., Richardson, J.A., Repa, J.J., Mangelsdorf, D.J., Kliewer, S.A., 2006, Proc. Natl. Acad. Sci., 103, 3920–3295. 141. Duran-Sandoval, D., Mautino, G., Martin, G., Percevault, F., Barbier, O., Fruchart, J.C., Kuipers, F., Staels, B., 2004, Diabetes, 53, 890–898. 142. Ma, K., Saha, P.K., Chan, L., Moore, D.D., 2006, J. Clin. Invest., 116, 1102–1109. 143. Cariu, B., van Harmelen, K., Duran-Sandoval, D., van Dijk, T.H., Grefhorst, A., Abdelkarim, M., Caron, S., Torpier, G., Fruchart, J.C., Gonzalez, F.J., Kuipers, F., Staels, B., 2006, J. Biol. Chem., 281, 11039–11049. 144. Yamauchi, T., Kamon, J., Wakim H., Terauchi, Y., Kubota, N., Hara, K., Mori, Y., Ide, T., Murakami, K., Tsuboyama-Kasaoka, N., Ezaki, O., Akanuma, Y., Gavrilova, O., Vinson, C., Reitman, M.L., H. Kagechika, Shudo, K., Yoda, M., Y. Nakano, Tobe, K., Nagai, R., Kimura, S., Tomita, M., Froguel, P., Kadowaki, T., 2001, Nature Med., 7, 941–946. 145. Wei, E.K., Giovanmucci, E., Fuchs, C.F., Wilett, W.C., Mantzoros, C.S., 2005, J.Natl.Cancer inst., 97, 1688–1694. 146. Fujisawa, T., Endo, h., Tomimoto, A., Sugiyama, M., Takahashi, H., Saito, S., Inamori, M., Nakajima, N., Watanabe, M., Kubota, N., Yamauchi, T., Kadowaki, T., Wada, K., Nakagama, H., Nakajima, A., 2008, Gut, 57, 1531–1538.

Chapter 7

AIB1: A Transcriptional Coactivator Which Integrates Signaling Cross Talk in Cancer Cells Macarena Ferrero and Jaime Font de Mora

Introduction: Cancer Etiology and Signal Transduction Cancer risk is determined by a combination of environmental factors and genetic predisposition. The etiology of cancer has a multi-factorial origin in which several elements impact tumor initiation and development with different degrees of relevance. Some of the factors contributing to cancer development have physiological implications and integrate disorders in signaling molecules that either in circulation or in a more localized environment directly affect cellular growth and proliferation. It is well-established that these factors, related to the expressing levels of hormones, growth factors, and cytokines, display a major role in cancer etiology [1, 2]. There are also alterations in transducers and effectors that mimic these signaling molecules. Hence, to counteract the excess of signaling, several therapeutic strategies have been developed against specific intracellular signaling networks for which inhibitory agents are currently undergoing clinical testing [3]. Another important factor that significantly predisposes for cancer is the genetic background inherited. The study of familial cancers has allowed the discovery of specific mutations in genes that strongly increase the chances to develop cancer in those relatives who inherited them. Most of these genes are tumor suppressors and their mutations are associated with lack of expression or function. Another characteristic of these tumor suppressors is that they are frequently associated to the risk of cancer in specific tissues such as BRCA1 and BRCA2 in breast and ovary [4], APC in colon [5], among others.

J. Font de Mora (*) Laboratory of Cellular and Molecular Biology, Centro de Investigación Príncipe Felipe, Avenida Autopista del Saler 16, 46010 Valencia, Spain e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_7, © Springer Science+Business Media, LLC 2012

129

130

M. Ferrero and J. Font de Mora

Fig. 7.1 Cancer etiology and signal transduction pathways. Cancer etiology is influenced by multiple diverse factors, all converging in chromosomal instability and alterations of signal transduction pathways. Two of the most frequently altered pathways in cancer are the phosphoinositide 3-kinase PI3K/AKT pathway and the Ras/ERK pathway that regulate key cellular processes and physiology

The level of contribution of environmental factors to cancer is more difficult to assess, although it is definitely very important, transuding with time in cancer development. Lifestyle, including diet and exercise [6], smoking and stress [7] are among others of significant influence that have come along with our current social habits. Finally, there are factors related to other diseases, such as viral infections [8], obesity [9, 10], and other genetic disorders such as ataxia telangiectasia [11] and others. It is very difficult to define how all of these factors contribute to cancer etiology, but what it is clear to researchers is that they all converge in chromosomal instability and eventually in alterations of the signal transduction pathways (see Fig. 7.1). An alteration of these pathways disables the cell to response adequately to other incoming signals. Cancer progression as a result of new modifications in the pathways that control transcriptional regulation, cell metabolism, proliferation, and survival is altered directing the cell to malignant transformation. Therefore, it is very important to understand these pathways and how they are altered in tumors in order to find new therapeutic approaches to more efficiently palliate each specific tumor.

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

131

The Steroid Receptor Coactivator or p160 Family The SRC proteins were firstly identified as factors that interacted with ligandbounded nuclear receptors and were able to enhance their dependent transcription. Nuclear receptors (NRs), such as thyroid hormone receptors, retinoid receptors, or estrogen receptors comprise a large family of ligand-dependent and DNA sequence-specific transcription factors which are implicated in many aspects of the animal physiology, involving reproduction, development, and energy homeostasis. They also play important roles in the initiation, progression, and treatment of numerous diseases including cancer. The activities of most NRs are regulated by small lipophilic molecules, including steroid/thyroid hormones, vitamin D3, and retinoids which finally control gene expression. Each type of ligand binds to and thereby activates one or a closely related set of a few nuclear receptor isoforms. Many NRs remain orphan receptors because they have no known ligand yet. On the other hand, it seems clear now that many receptors can influence transcription in the absence of ligand. Structurally, all members of the nuclear receptor family share a common structure with three major functional domains. The DNA-binding domain (DBD), in the middle region, is the most conserved one, and it is responsible for site-specific DNA binding at its response elements and for protein–protein interactions. The second most conserved domain is the transcriptional activation function 2 (AF-2), located at the carboxyl-terminal extreme. This region is composed of 12–13 alpha-helices folded into a globular structure and is responsible for ligand binding and aspects of homo-and/or hetero-dimerization. In some cases, it has the active conformation in the absence of ligand. Finally, within the amino-terminal extreme lies the transcriptional activation function 1 (AF-1). Unlike AF-2, AF-1 shows weak conservation and weaker transcriptional activity. This domain functions as a ligand-independent transcriptional activator and is responsible for interactions with proteins relevant to the transcription initiation such as TIFIIB and other accessory proteins. Although either AF-1 or AF-2 is able to regulate transcription alone, full transcriptional activity requires synergy between both domains. It is well understood that in order to orchestrate gene expression, NRs cooperate with several corregulators. Depending on the activating or suppressing effect of these corregulators upon gene transcription they are referred as coactivators or corepressors respectively [12, 13]. The initial step in transcriptional coactivation is the release of nuclear receptor corepressors. In general, after ligand binding, the receptor undergoes a conformational change that results in dissociation from these corepressors and recruitment of a diverse set of coactivators and regulatory molecules. Following this recruitment, the coactivator complex triggers chemical and structural modifications in the chromatin and communicates with the basal transcriptional machinery activating the gene transcription. Among the broad spectrum of identified coactivators, the proteins of the SRC family have been the most extensively studied. Three members comprise this family so far.

132

M. Ferrero and J. Font de Mora

Fig. 7.2 Structural domains of SRC proteins. The amino-terminal region contains the bHLH/PAS domain which is the most conserved domain; S/T, serine/threonine-rich regions; 1–9, LXXLL motifs; RID, nuclear receptor interaction domain; CID, CBP/p300 interaction domain; HAT, intrinsic histone acetyltransferase activity; squared yellow Q, glutamine-rich region; AD-1, activation domain 1; AD-2, activation domain 2

They are: SRC-1/nuclear receptor coactivator 1 (NCoA1) [14], SRC-2/nuclear receptor coactivator 2 (NCoA2)/transcriptional intermediary factor 2 (TIF2)/glucocorticoid receptor interacting protein 1 (GRIP1) [15] and SRC-3/nuclear receptor coactivator 3 (NCoA3)/amplified in breast cancer 1 (AIB1)/retinoid acid receptorinteracting protein 3 (RAC3)/acetyltransferase (ACTR)/thyroid hormone receptoractivated molecule 1 (TRAM-1) or p300/CBP-interacting protein (p/CIP) [16–21]. SRC proteins are encoded by different genes located in the human chromosomal regions 2p23, 8q21.1, and 20q12 [16, 22, 23]. These coactivators are about 160 KDa in size and share around 50–55% of sequence similarity and 43–48% of sequence identity. Structurally (see Fig. 7.2), at the amino-terminal region, all three members contain a basic helix-loop-helix (bHLH) followed by a double PAS domain (Per/ Ah receptor nuclear translocator/Sim domain) which is the most conserved region among the SRC proteins. This domain was initially identified in Drosophila [24] and it was implicated in DNA binding and protein–protein interactions. Although SRCs can interact with some transcription factors independently of the bHLH/ PAS domain, in other cases, this domain is required to interact and subsequently coactivate a few others, such as TEF-4, MEF-2C, and STAT5 [25–27]. In addition, this domain can also contribute to the coactivation function by recruitment of downstream coactivators, including GAC63, CoCoA, Flightless 1, and ANCO1 [28–31]. The central region, which is relatively conserved, contains several LXXLL motifs (L = Leucine and X = any amino acid) also called NR boxes because they are responsible for interaction with ligand-bounded NRs. Structural analysis has revealed that the LXXLL motif forms an amphipatic a-helix when bounded to a groove in the AF-2 domain of the nuclear receptors. Several studies have shown that the LXXLL

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

133

motif is necessary and sufficient to mediate binding with NRs [32–35]. However, not all the LXXLL motifs exhibit the same binding affinity for different NRs [36], suggesting that NRs show preferences over one coactivator or another. In fact, the LXXLL motif is necessary in the binding, but amino acids flanking this core motif are important in nuclear receptor recognition [37]. Within the carboxi-terminal region, there are two intrinsic transcriptional activation domains, AD-1 and AD-2. A transcriptional activation domain is defined in transcriptional assays with reporter genes and fusion proteins containing the Gal4 DNA-binding domain and fragments of SRC proteins. AD-1 is responsible for interaction with histone acetyltransferases (HATs), such as CBP/p300, pCAF, and more recently GCN5 [38]. Interestingly, in this region, there are several LXXLLlike motifs which are different from those recognizing NRs. It has been demonstrated that directed mutagenesis in key residues of these motifs or deletion of the motifs impairs the recruitment of CBP/p300 and subsequently, the coactivation function of SRC factors [35, 39, 40]. This fact suggests the importance of CBP/p300 interaction for the function of SRCs. Furthermore, a weak HAT activity has been attributed to SRC-1 and AIB1 [17, 41], raising the possibility that SRC family members may be directly involved in chromatin remodeling. Nonetheless, the importance of this activity still remains unclear. Recently, it has been observed that AIB1 and SRC-1 can acetylate ER81 in vitro (AIB1 also in vivo), but the degree is much lower than the acetylation by CBP/p300. Apparently, this acetylation does not contribute to the ER81 transcriptional activity, whereas acetylation by CBP/p300 is crucial [42]. The AD-2 region is responsible for interactions with histone methyltransferases such as CARM-1 and PRMT1 which also play an important role in chromatin remodeling and basal transcriptional machinery assembly. Within this domain, AIB1 presents a polyglutamine-rich region. Although the biological significance of this repeat is not established, it has been observed that short polyglutamine genotypes promote aggressive malignant phenotypes of ovarian carcinomas [43]. On the other hand, there is no evidence for an increased risk of breast cancer in BRCA1; BRCA2 mutation carriers associated with the length of this glutamine chain in AIB1 [44, 45]. Functionally, SRC coactivators have been proposed as bridge proteins among NRs and the basal transcriptional machinery. However, all three members are able to interact and coactivate not only NRs but also certain transcription factors, including AP-1 [46, 47], nuclear factor-кB (NF-кB) [48–50], signal transducers and activators of transcription (STATs) [27, 51–53], hepatocyte nuclear factors (HNFs) [54, 55], mitogen-activated protein kinase (MAPK)-dependent transcription factors (Ets) [42, 56], p53 [57], and E2F1 [58]. This means that p160 proteins play a much broader role in cell regulation than originally expected. In this way, changes in the levels and activities of these coactivators may greatly affect the expression levels of many genes and, as a consequence, influence a great number of cellular processes. The activities of SRC proteins can be regulated by a variety of post-translational modifications, including phosphorylations, ubiquitinations, sumoylations, methylations, and acetylations (reviewed in 59]. In Table 7.1, a summary of different AIB1interacting proteins responsible for these modifications as well as for other cellular

Peptidyl isomerase

Ubiquitin ligase Ubiquitin ligase

Ubiquitin ligase

26S proteasome ATPase

Promotes cell proliferation during S-phase Nuclear shuttling carrier Cell growth regulator

NR coactivators

Pin 1

CHIP E6AP

SCFFbw7a

SUG1

BRCA-IRIS

Importin a-3 SIP

NCoA proteins

Features G1/S cell cycle transition Facilitates the assembly of coregulator complexes to the local chromatin Modulates AIB1 coactivation function and regulates its turnover Reduces tumor growth and metastasis Mediates AIB1 proteasomal degradation

[166] [214]

[213]

References [211] [212]

Enhances ubiquitination and eventual transcrip- [62] tion-coupled degradation of AIB1 Interacts with AIB1 and coactivates RA target [215] genes Interacts with AIB1 and also with Activates cyclin D1 expression [178] SRC-1 Mediates nuclear import of AIB1 [216] Sequesters SRCs proteins in the cytosol and [217] inhibits estrogen-stimulated proliferation of mammary carcinoma cells NCoA proteins are able to Influences the formation and recruitment of the [53] interact each other through coactivator complex to the gene promoters PAS-B domain

Downregulates Smad and Twist Interacts with AIB1 carboxiterminal extreme in vitro Ubiquitinates K723 and K786

NR coactivator

Table 7.1 Non-transcriptional factors that interact with AIB1, regulating its function Interacting proteins Function Others GAS SRC-1 specific interaction ERa coactivator ANCAA AAA + ATPase ERa coactivator

134 M. Ferrero and J. Font de Mora

Function

Kinase

Kinase

Kinase

Kinase Kinase

Kinase Kinase

Phosphatase

Interacting proteins

Kinases and phosphatases p38

GSK3

IKK

ERK1/2 CK1d

Abl kinase PKCz

PP-1

Features

Coactivates RARa-mediated transcription and after, stimulates AIB1 degradation Switches ubiquitination and transcriptional coupled degradation of AIB1 Phosphorylates AIB1 Activates NF-кB-mediated transcription in response to TNFa and also ER-responsive genes in response to estrogens Phosphorylates in S857 and S860 Facilitates AIB1 interaction with ER Phosphorylates S601 Enhances interaction with ERa, coactivation capacity, and proteasomal stability Phosphorylates Y1357 Favors interaction with ERa and p300 Phosphorylates in the carboxiStabilizes AIB1 by inhibiting its interaction terminal region with the 20S proteasome Dephosphorylates AIB1 at S101 Stabilizes AIB1 and inhibits its activity and S102

Phosphorylates AIB1, at least in part, in S860 Phosphorylates S505

Others

[218]

[148] [65]

[147] [64]

[50, 157]

[62]

[63]

References 7 AIB1 is a Potential Therapeutical Target in Cancer Progression… 135

136

M. Ferrero and J. Font de Mora

functions is shown. Regarding degradation, it has been previously demonstrated that SRC family members are targets of the ubiquitin-proteasome system [60] and more recently that AIB1 can also be degraded in a ubiquitin and ATP-independent manner [61]. Furthermore, there are several factors that are involved in SRCs stability, such as subcellular distribution, phosphorylation by several kinases, including p38MAPK, GSK3, APKC, and CK1d, methylation by CARM-1, etc. [62–67]. The SRC family is well known for its oncogenic potential; hence, the study of all these factors affecting levels, activities, and subcellular distribution of p160 coactivators is of great importance as it may open new therapeutic approaches in cancer research.

Role of AIB1 in Physiology Most of our current knowledge on the role of AIB1 in physiology has been given by the study of animal models. AIB1 is mainly expressed in tissues of strong response to endocrine hormones, suggesting that AIB1 displays an important role in endocrine responsiveness. The study of the phenotypes developed by animal models re-enforces this hypothesis.

Role of AIB1 During Development Genetic disruption of AIB1 in mice results in a pleiotropic phenotype showing dwarfism, delayed puberty, reduced female reproductive function, and blunted mammary gland development [68]. Reduced somatic growth and body size in AIB1deficient mice is the result of both altered regulation of IGF-I gene expression in specific tissues and a defect of response to IGF-I [69]. In contrast, no alterations in somatic growth and fertility are observed in SRC-1 knockout mice [70]. All these results support that the physiological role of AIB1 is different from that of SRC-1 and proof the functional diversity among coactivator family members. In prostate cancer cells, it was observed that AIB1 overexpression resulted in increased AKT (or protein kinase B – PKB)/mTOR signaling activation and cell growth in a steroid-independent manner [71]. However, downregulation of AIB1 expression by small interfering RNA decreased cell growth, leading to a smaller cell size. Similarly, in AIB1 null mutant mice, AKT signaling is downregulated in those tissues that would normally express AIB1 in wild-type mice. Furthermore, AIB1 overexpression in the mammary epithelium increases mammary IGF-I mRNA and serum IGF-I protein levels together with the activation of the IGF-I receptor and downstream signaling molecules [72]. On the contrary, IGF-I circulating levels were reduced in AIB1−/− mice, although IGF-I mRNA levels were not affected [68]. Low IGF-I levels in AIB1−/− mice were the result of a rapid degradation of IGF-I. In the absence of AIB1, transcription of the vitamin D receptor (VDR)-target gene IGF-binding

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

137

protein 3 (IGFBP3) was reduced, resulting in the instability of IGF-I [73]. Both, acid-labile subunit (ALS) and IGF-binding proteins form complexes with IGF-I to maintain its circulating concentration and endocrine function. But in AIB1 knockout mice, IGFBP3 was specifically affected and not ALS or growth hormone (GH). Taken together, these results suggest that AIB1 is an important modulator for mammalian cell growth and that IGF-I signaling is an important effector of its physiological function.

Role of AIB1 in Reproductive Tissues Knockin mice expressing b-galactosidase in one of the AIB1 alleles demonstrated that it is specifically expressed in oocytes and its total ablation is very probably the cause of the low estrogen levels in females [68]. AIB1−/− females display delayed development of the reproductive functions. In contrast, mice lacking TIF2 have impaired fertility in both sexes [74]. Hypofertility in TIF2−/− males is due to defects in both spermiogenesis (teratozoospermia) and age-dependent testicular degeneration. TIF2 expression appears to be essential for adhesion of Sertoli cells to germ cells. Female hypofertility is due to a placental hypoplasia that most probably reflects a requirement for maternal TIF2 in decidua stromal cells that face the developing placenta. The progesterone receptor has shown to be an essential mediator of female fertility and development of reproductive tissues [75]. A transgenic mouse model that utilizes a progesterone receptor (PR) activity indicator (PRAI) system was very useful to define dynamic regulation of PR activity in vivo [76]. The study revealed the spatiotemporal activation of PR in female reproductive tissues. Importantly, PRAI mice under the genetic background of AIB1−/− showed that PR distinctly coordinates the p160 transcriptional coactivators in the different tissues [77]. Whereas AIB1 is the primary coactivator for PR in the breast, SRC-1 is the primary coactivator for PR in the uterus. Selective abrogation of TIF2 in PR expressing cells showed to be an indispensable coregulator for uterine and mammary gland responses that require progesterone [78]. In the absence of TIF2, uterine cells positive for PR were incapable of facilitating embryo implantation, a necessary first step toward the establishment of the materno-fetal interface. Importantly, such an implantation defect is not exhibited by knockouts for SRC-1 or AIB1, underscoring the unique coregulator importance of TIF2 in peri-implantation biology. Moreover, despite normal levels of PR, SRC-1, and AIB1, progesterone-dependent branching morphogenesis and alveologenesis fails to occur in the murine mammary gland in the absence of TIF2, thereby establishing a critical coregulator role for TIF2 in signaling cascades that mediate progesterone-induced mammary epithelial proliferation. Finally, the recent detection of TIF2 in the human endometrium and breast suggests that this coregulator may also represent a new clinical target for the future management of female reproductive health and/or breast cancer.

138

M. Ferrero and J. Font de Mora

Role of AIB1 in the Central Nervous System In the brain, AIB1 is expressed in the olfactory bulb and the hippocampus as well as SRC-1 and at lower levels TIF2. However, SRC-1 is also highly expressed in the piriform cortex, amygdala, hypothalamus, cerebellum, and brainstem. Importantly, disruption of SRC-1 specifically delays the Purkinje cells development and maturation in early stages and results in moderate motor dysfunction in adulthood [79]. In the nervous system, AIB1 and the other p160 coactivator family members display different functions depending on the cell type. In response to glucocorticoids, the glucocorticoid receptor (GR) differentially recruits p160 family members: In Schwann cells, GR recruits SRC-1a, SRC-1e (two distinct SRC-1 isoforms differing in their C-terminal region [80]), or AIB1, whereas in astrocytes, GR recruits SRC-1e and TIF2, and to a lesser extent AIB1 [81]. This can be explained by the different intracellular distribution of the p160 coactivators. For example, within astrocytes, SRC-1 and TIF2 were mainly nuclear, whereas AIB1 unexpectedly localized to the lumen of the Golgi apparatus. In contrast, in Schwann cells, SRC-1 showed a nucleocytoplasmic shuttling depending on hormonal stimulation, whereas TIF2 remained strictly nuclear and AIB1 remained predominantly cytoplasmic. Altogether, these results highlight the cell specificity and the time dependence of p160s recruitment by the activated GR in glial cells, revealing the complexity of GR-p160 assembly in the nervous system [82]. In aT3-1 pituitary cells, treatment with gonadotropin-releasing hormones induced progesterone receptor phosphorylation and translocation to the nucleus. Further transactivation of progesterone responsive genes required the recruitment of AIB1 to the activated PR transcriptional complexes, thus demonstrating the essential role of AIB1 in these pituitary cells [83]. Noteworthy, nuclear overexpression of AIB1 occurs in proliferative prolactinoma cells concomitantly with estrogen receptor a (ERa) and aromatase positivity [84], suggesting a novel mechanism whereby the abnormal high conversion of testosterone to estradiol is mediated by aromatase in these pituitary cells, upregulating AIB1 expression and or function, and leading to an increase in lactotroph proliferation.

Role of AIB1 in Energy Homeostasis AIB1 has been shown to control the metabolism resulting from increased mitochondrial function and energy expenditure as a consequence of activation of PGC-1a [85]. In this way, PGC-1a is an essential effector of AIB1 function, controlling mitochondrial function and energy homeostasis. Mechanistically, AIB1 regulates the expression of GCN5, the only acetyltransferase known to acetylate PGC-1a and consequently inhibiting its activity. Importantly, caloric ingestion induces AIB1 expression, resulting in the inhibition of PGC-1a and energy expenditure. On the

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

139

contrary, caloric restriction reduces AIB1 levels leading to enhanced PGC-1a activity and energy expenditure. Collectively, these data suggest that SRC-3 is a critical link in a cofactor network that uses PGC-1a as an effector to control mitochondrial function and energy homeostasis. In addition, AIB1 is a critical regulator of white adipocyte development [86]. In the absence of AIB1, adipocyte differentiation is impaired, reducing the transactivation capacity of the transcription factor CCAAT/enhancer-binding protein (C/EBP) and decreasing the expression levels of PPARg2. Similarly, gene ablation studies previously identified SRC-1 and TIF2 as being involved in the control of energy homeostasis [87]. However, whereas TIF2−/− mice are protected against obesity and display enhanced adaptive thermogenesis, SRC-1−/− mice are prone to obesity due to reduced energy expenditure. A genomic approach using microarray analysis demonstrated that the molecular targets of TIF2 in the liver stimulate fatty acid degradation and glycolytic pathway, whereas fatty acid, cholesterol, and steroid biosynthetic pathways were downregulated [88]. These findings strongly support the idea that gene expression changes are specific for each coactivator and non-overlapping in the liver.

Role of AIB1 in Other Tissues Another target of AIB1 is the thyroid hormone receptor beta (TRb). Mutations in the TRb develop thyroid hormone resistance and impaired growth. A novel mouse model harboring a mutation in TRb (TRbPV mice) previously found to cause resistance to thyroid hormone (RTH) in humans faithfully reproduces RTH of humans. Interestingly, depletion of AIB1 in this mouse model partially rescued dysfunction of the pituitary-thyroid axis, but growth impairment was worsened [89]. The lack of AIB1 reduced the growth of both the pituitary and the thyroid in TRbPV mice, thereby lessening the dysregulation of the pituitary-thyroid axis. In contrast, the lack of AIB1 exacerbated the growth impairment observed in TRbPV mice. The modulatory effect of AIB1 on growth is mediated via the IGF-1/PI3K/ AKT/mTOR signaling pathway. This dual action in the pituitary-thyroid axis and in growth can be explained by two mechanisms, one via its role as a receptor coregulator and the other via its growth regulatory role through the IGF-I/PI3K/ AKT/mTOR signaling. AIB1 is also expressed in vascular smooth muscle cells and endothelial cells, and its deficiency in knockout mice reveals its role in the inhibition of neointimal growth, suggesting its ER-dependent vasoprotective effects under conditions of vascular trauma [90]. An important physiological role of AIB1 has also being described in the inflammatory process. In macrophages, AIB1 has a novel cytoplasmic function by which activates the translational silencers TIA-1 and TIAR and thus inhibits the translation of proinflammatory cytokines [91]. The observation was originally made in AIB1−/− macrophages that produced significantly more proinflammatory cytokines, such as

140

M. Ferrero and J. Font de Mora

TNF-a, IL-6, and IL-1b, in response to induced endotoxicity, although the mRNA levels remained the same. Hence, AIB1 displays a protective action against the lethal endotoxic shock triggered by an acute inflammatory response. Based on the observations made in AIB1−/− mice, hematopoyesis is also affected. AIB1 regulates lymphopoiesis by inhibiting proliferation of lymphocytes. Absence of AIB1 decreases platelet and increases lymphocytes numbers, leading to the development of malignant B-cell lymphomas upon aging [92]. Lymphoid lineage derived from knockout mice displays constitutive activation of NFkB due to increased IkB kinase and consequently lower IkB levels, activations that can be reversed upon ectopic expression of AIB1. Therefore, taken all together, AIB1 exerts multiple functions depending on the cell type, causing proliferation and preventing from apoptosis in certain types like in the epithelial and inhibiting proliferation in others like in lymphoid cells.

Overexpression of AIB1 Affects Both Cancer Initiation and Progression Among the three coactivators of the SRC family, AIB1 has received more attention because compelling evidences demonstrate that this coactivator is a new oncogene intervening in both initiation and progression of cancer.

Evidences from Biopsies and Animal Models AIB1 has been implicated in the pathogenesis of a number of malignancies, including breast cancer, endometrial cancer, prostate cancer, ovarian cancer [16, 106–115], urothelial cancer [93], nasopharyngeal cancer [94], esophageal squamous cell carcinoma [95, 96], colorectal cancer [97–100], hepatocellular carcinoma [101], pancreatic carcinoma [102, 103], gastric cancer [104], and cutaneous melanoma [105]. The first indication that AIB1 might confer an advantage in cell growth came from Anzick et al. They examined AIB1 copy number by fluorescence in situ hybridization (FISH) in a series of ER-positive and ER-negative breast cancer cell lines and found that AIB1 was amplified in four of five ER-positive cell lines and none of six ER-negative [16]. Through Northern blotting assays, AIB1 was seen highly overexpressed in the cell lines which showed amplification but also in cell lines without amplification of the gene. In contrast, the levels of SRC-1 and GRIP1 remained constant. They also evaluated 105 specimens of breast cancer for FISH and found amplification of AIB1 in 9.5% of the cases. However, the amplification was not as high as in the cell lines. In these specimens, overexpression was observed in overall 64%, suggesting that overexpression without amplification of a gene was possible in breast cancers. They also demonstrated that AIB1 overexpression

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

141

increased ER-dependent transactivation in a dose-dependent manner. In a larger study, Bautista et al. observed amplification of AIB1 gene in 56 of 1,157 breast tumors (4.5%). They also analyzed 122 ovarian tumors finding a 7.4% of amplification [106]. Interestingly, a splice variant of AIB1 that lacks the bHLH domain and a proportion of the PAS domain (AIB1-Delta3 isoform) has also been observed overexpressed in breast cancer biopsies and in the breast cancer derived cell line MCF-7. This isoform has an enhanced efficacy activating nuclear receptors as well as potentiating EGF signaling compared to the full-length AIB1 [107]. There are more studies that show amplification and overexpression of AIB1 not only in breast tumors but also in endometrial, ovarian, and prostate carcinomas, including prostate cancer cell lines, such as LNCaP and prostate stromal cells where AIB1 is an important androgen receptor (AR) coactivator and may have a role in the progression of prostate cancer [108–112]. In general, malignant tumors are classified according to their size, grade of differentiation from their parental tissues, degree of spread to regional lymph nodes, and presence of metastasis. Tumors with high grade show little differentiation and poor prognosis. Interestingly, high expression of AIB1 has been correlated with poor patient outcome [113]. In line with this, breast tumors that show high levels of AIB1 have significantly shorter disease-free and overall survival times after surgery [114]. Similarly, it has been observed in endometrial carcinomas a correlation between AIB1 expression and clinical stage, myometrial invasion and dedifferentiation [115–118]. In ovarian cancer, Han et al. found a 36% of tumors with AIB1 overexpressed. They also observed high expression in poorly differentiated tumors at late stage and tumors with lymph node involvement [119]. These observations suggest that AIB1 may play a role in cancer progression. Amplification and overexpression of AIB1 is not restricted to endocrine tumors; there are also increased copy number and/or high levels of this coactivator in nonendocrine sensitive tumors, such as pancreatic and gastric cancer [103, 104], esophageal squamous cell carcinomas [95, 96], nasopharyngeal carcinomas [94], colorectal carcinomas [97–100], hepatocellular carcinomas [101], and cutaneous melanomas [105]. Henke et al. examined a series of pancreatic tissues and also observed high expression levels of AIB1 as malignancy staged. High levels of both mRNA and protein existed in pancreatic adenocarcinomas. Interestingly, AIB1 staining was located within the nucleus, whereas some cytoplasmic protein was also found in normal epithelia and low grade intraepithelial pancreatitis. Amplification of AIB1 gene was found in 37% of pancreatic adenocarcinomas [102]. In general, there is an association between AIB1 overexpression and tumor staging. All these findings suggest that AIB1 overexpression may be important in the acquisition of an invasive and/or metastatic phenotype, and hence, AIB1 could be used as a marker to predict prognosis in cancer. The development of animal models has improved our understanding of AIB1 and its role in cancer progression. Torres-Arzayus et al. established an AIB1 transgenic mouse under the control of the mouse mammary tumor virus (MMTV)/long terminal repeat (LTR) promoter/enhancer. They observed a more extensive degree of ductal branching, larger size of the cells, higher BrdU-positive cells, and less number

142

M. Ferrero and J. Font de Mora

of epithelial cells undergoing apoptosis in the transgenic (tg) mammary glands than in the wild-type (WT) glands. At 5 months of age, mammary glands of tg mice showed numerous hyperplastic lesions. These tg mice presented more tumor incidence than WTs. Tumors developed not only in breast but also in other organs, including lung, pituitary, and uterus, supporting the hypothesis that AIB1 was indeed involved in a wide range of cancers. Analysis of the mammary glands revealed that AIB1 activated PI3K/AKT/mTOR signaling pathway by increasing IGF-I circulating levels [72]. Therefore, it seemed that AIB1 was a potent oncogene and that the mechanism of its oncogenicity might involve the PI3K/AKT/mTOR signal transduction pathway. A transgenic mouse harboring the AIB1-Delta3 isoform was also generated [120]. Expression of the isoform in transgenic mice was driven by the human cytomegalovirus immediate early gene 1 (hCMVIE1) promoter. Although no tumors were seen, mice showed increased mammary proliferation and gland mass together with increased expression levels of cyclin D1, IGF-IR, and C/EBPb liver-enriched inhibitory protein (LIP) isoform. These results suggested that AIB1 worked through different pathways. Avivar et al. created a mouse model which expressed moderate levels of AIB1 to analyze its role during the first steps of cancer [121]. Similarly to the AIB1-Delta3 tg mice, these mice did not generate tumors. However, they displayed mammary hyperplasia at the onset of puberty. Interestingly, primary mammary epithelial cultures derived from these mice presented enhanced proliferation and augmented levels of cyclin D1 and E-cadherin. Therefore, overexpression of AIB1 may represent one of the pre-neoplastic changes in breast tissue. On the other hand, the lack of AIB1 in mice has been studied as well. The (MMTV)/v-Ha-ras (ras) transgenic mouse is a model with high frequency of mammary carcinomas. The lack of AIB1 in this transgenic mouse delays mammary tumor latency, reduces mammary tumor frequency, and suppresses primary tumor growth and metastasis to the lung [122]. Palpable breast tumors were first observed in AIB1+/+-ras and AIB1+/−-ras at ages 14 and 18 weeks, respectively, whereas the first tumor in AIB1−/−-ras was observed much later, at 40 weeks. Moreover, only 17% of AIB1−/−-ras mice presented focal lung tumors compared to 39% of AIB1+/−ras and 42% of AIB1+/+-ras mice. Cell lines generated from the mammary tumors of AIB1+/+-ras and AIB1−/−-ras mice showed less migration ability and reduced levels of IRS-1 and IRS-2 when AIB1 was deleted. The MMTV-Neu/AIB1+/− or AIB1−/− mice also present tumor latency delayed and protection from tumorigenesis [123]. Furthermore, in another study, it is reported that AIB1-knockout mice treated with the carcinogen 7,12-dimethylbenz[a]anthracene (DMBA) show mammary ductal morphogenesis and delayed onset of palpable tumors in comparison with WT rodents [124]. Hence, AIB1 deficiency protects against DMBA-induced tumorigenesis. Levels of IRS-1, IRS-2, p-AKT, and cyclin D1 were augmented in DMBAinduced WT tumors but not in DMBA-induced AIB1−/− tumors. The results revealed that AIB1 was required for IGF/PI3K/AKT signaling pathway-stimulated upregulation of cyclin D1 during DMBA-induced breast tumor initiation and progression. The role of AIB1 deficiency has also been studied in prostate and thyroid cancer

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

143

using TRAMP or TRbPV/PV mice models, respectively. In the case of TRAMP mice, loss of AIB1 significantly prolonged life and suppressed tumor progression. WT/ TRAMP mice presented poor differentiated prostate adenocarcinomas at 30 weeks, whereas AIB1−/−/TRAMP mice only formed early stage well-differentiated adenocarcinomas [125]. Unlike AIB1, loss of SRC-1 does not suppress either prostate cancer tumorigenesis or metastasis in the SRC-1−/−/TRAMP mice probably due to a compensation of AIB1 overexpression [126]. In TRbPV/PV rodents, the lack of AIB1 triggered a decrease in genes involved in cell cycle progression, including E2F1, Cdk2, Cdc6, and Cdc25a phosphatase. Due to this fact, thyroid carcinogenesis was delayed. Moreover, AIB1−/−/TRbPV/PV mice did not develop either vascular invasion or metastasis [127]. Cell models provide also useful information complementary to the one obtained with animal models. MCF10 cell line is an ER-negative, spontaneously immortalized, non-transformed human mammary epithelial cell line that has been extensively used to evaluate the activity of a great number of oncogenes. MCF10 infected with adenovirus expressing AIB1 displayed high frequency of colony formation in agar plates, while no colony was observed in the control infected with adenovirus expressing GFP [128]. Hence, this result is in accordance with animal model observations. Altogether, clinical studies, animal models, and cellular models reveal a clear oncogenic ability for AIB1. Therefore, factors which influence cellular levels and activity of this p160 family member could be of great importance in determining the potential for growth and transformation of a tissue.

Mechanisms of AIB1 Oncogenic Activity Hormone-Dependent Mechanisms: The Steroid Receptor Signaling Sex hormones or sex steroids, such as estrogen, androgen, and progesterone, affect the growth and function of reproductive organs, the development of sex characteristics, and the behavioral patterns of animals. However, they are also implicated in diseases, including breast, ovarian, endometrial, and prostate cancer. In line with this, Lathrop and Loeb found that an early ovariectomy prevented tumor incidence in a great number of mice because it eliminated sex steroids production [129]. Since sex steroids mediate their function through nuclear receptors and AIB1 is a NR coactivator, it is reasonable to think that overexpression of AIB1 together with a high exposition to steroids results in more NR activity (Fig. 7.3). Supporting this idea in breast cancer, AIB1 amplification and overexpression have been correlated with ER and PR positivity [106, 112, 130, 131]. It has been demonstrated that AIB1 and ER can be associated and, in fact, colocalize to the nuclei of ER-positive cells [132]. Furthermore, AIB1 is required for ER-binding to its target gene promoters. Treatment with 17-b-estradiol induces a dramatic increase in occupancy of both ERa

144

M. Ferrero and J. Font de Mora

Fig. 7.3 Schematic view of major signaling pathways converging on AIB1. AIB1 can stimulate PI3K/AKT signaling by increasing expression levels of several components of the pathway as well as enhance HER2/Neu pathway by augmenting its phosphorylated levels (not represented in the figure). Phosphorylation of AIB1 is necessary for its activity. Steroids, cytokines, and growth factors can induce AIB1 phosphorylation. Once activated, it will be able to interact and coactivate a variety of transcription factors. In response to estrogens, AIB1 can be recruited to the estrogenresponsive regions in target genes and stimulating cell cycle progression. Estrogen treatment facilitates the association of IKKa, ERa, and AIB1 to estrogen-responsive promoters and increases IKKa-mediated phosphorylation of ERa, AIB1, and histone H3. Alternatively, growth factor stimulation triggers ER and/or AIB1 phosphorylation, resulting in ER-mediated activation of cell proliferation independently of estrogens, developing resistance to endocrine therapy. Signal stimulation by cytokines activates the IKK complex and leads to phosphorylation of IkB as well as to nuclear localization of AIB1. In this sense, cytokines can enhance activities of both NF-kB and steroid receptor pathways. In addition, AIB1 also interacts and coactivates Ets and AP-1 transcription factors to stimulate the expression of MMPs which will subsequently allow the invasion of peripheral tissues. Noteworthy, HER2/Neu promoter is a target for Ets and AP-1; therefore, AIB1 might also increase its expression, promoting cell proliferation and generating again endocrine resistance. Finally, AIB1 is able to enhance transcription of E2F1-responsive genes, including cyclin E, cyclin A, Cdk2, AIB1, and E2F1 itself. AIB1 potentiates cell cycle progression and establishes a positive feedback loop with E2F1

and AIB1 at the CATD, pS2, and c-Myc gene promoters in MCF-7 cells [133], and in contrast, suppression of AIB1 with small interference RNA decreases the association between ER and promoters [134]. Hence, it is hypothesized that AIB1 is involved in cancer progression by overstimulating ER function. Accordingly, it has been observed that during breast cancer tumorigenesis there is a significant

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

145

upregulation of ER signaling in ER-positive tumors. This is due to an increased ER expression along with increased levels of factors that might activate ER function [135]. Interestingly, AIB1-Delta3 isoform has an enhanced efficacy coactivating ERa compared to AIB1 full-length probably because it is more available to NR when bHLH/PAS domain is lost. Tumors that overexpress this isoform might have more estrogenic stimulation of proliferation [136]. On the other hand, AIB1 depletion reduces estrogen-dependent proliferation of MCF-7 cells by decreasing the ability of estrogen to inhibit apoptosis [137]. Unlike AIB1, the role of SRC-1 in breast cancer seems to be less pronounced because SRC-1 is unable to compensate AIB1 loss [137]. Noteworthy, it has been demonstrated that SRC-1 interacts less with ER than AIB1 after estrogen treatment in MCF-7 cells [138]. Thus, AIB1 might have a specific role in mediating estrogen-dependent transcription in breast cancer. Of interest, another major ER downstream target gene to promote cell cycle in breast cancer cells is cyclin D1 [139]. AIB1 along with ER can be recruited to the estrogen-responsive cyclin D1 promoter to enhance cyclin D1 expression and hence, cell proliferation [140]. Taken all together, these observations suggest that overexpression of AIB1 might contribute to tumorigenesis by overstimulating steroid receptor-mediated transactivation. However, while it is likely that increased levels of AIB1 contribute to cancer progression through this mechanism in endocrine tumors, AIB1 overexpression has also been observed in ER-, PR-, and AR-negative tumors and cancer cell lines [141, 142], indicating that hormone-independent mechanisms exist in the process of cancer in response to AIB1. Hormone-Independent Mechanisms Many facts support the idea that AIB1 is involved in tumorigenesis via several signaling pathways: (1) AIB1 has the ability to interact and coactivate not only steroidresponsive nuclear receptors but also non-responsive nuclear receptors and other transcription factors, (2) its overexpression does not always correlate with ER or PR expression but instead correlates with HER2/Neu, and (3) AIB1 has been found amplified and overexpressed in a number of hormone-independent cancers. It is hence reasonable to believe that AIB1 might perturb signal integration by multiple transduction pathways other than steroid receptor signaling. During the last years, extensive efforts have been done to study the importance of AIB1 as a non-steroid receptor coactivator in cell growth, cell proliferation, and tumor development. So far, AIB1 has been observed to affect several signal transduction pathways, including IGF-I/Phosphatidylinositol 3-Kinase (PI3K)/AKT, HER2/Neu, NF-кB, and E2F1 signaling pathways.

The IGF/PI3K/AKT Signaling Pathway The binding of the insulin-like growth factor 1 (IGF-I) to its receptor (IGF-IR) initiates signaling via the insulin receptor substrates (IRS-1 and IRS-2) which associate

146

M. Ferrero and J. Font de Mora

to and activate several intracellular kinases including PI3K. Once activated, PI3K phosphorylates the membrane phospholipid phosphatidylinositol 4,5-bisphosphate (PIP2) to phosphatidylinositol 3,4,5-trisphosphate (PIP3) creating a binding site for the kinases PDK-1 and AKT. PDK-1 then triggers AKT phosphorylation, and in turn, activated AKT directly or indirectly phosphorylates a great number of substrates, including glycogen synthase kinase-3b (GSK3b), mammalian target of rapamycin (mTOR), p70S6K, caspase 9, BAD, and a few others to control processes, such as metabolism, cell survival, and cell proliferation [143]. Animal models, human tumors, and cell lines have demonstrated that AIB1 is involved in the regulation of AKT activity. There is a correlation between AIB1 and the expression levels of components of the IGF-I/PI3K/AKT signaling pathway. Mammary glands of AIB1 transgenic mice show high levels of phosphorylated IGF-IR and AKT indicating that PI3K/AKT pathway is more active in these animals [72]. The increased activity is due to an augment in the circulating levels of IGF-I. Interestingly, CMV-AIB1-Delta3 mice also display increased IGF-IR expression [120]. On the other hand, deletion of AIB1 in mice results in growth retardation as well as in overall reduction of IGF-I levels [68, 69, 71]. Wang et al. found a 30–50% decrease of IGF-I levels in AIB1 null rodents. Xu et al. registered a similar reduction (40%) with unaltered growth hormone levels. In addition, whereas no significant differences were found in IGF-IRß levels, levels of IGF-I, IRS-1, and IRS-2 were significantly reduced in MMTV-AIB1−/−-ras tumors compared to MMTV-AIB1+/+ -ras or MMTV-AIB1+/−-ras tumors [122]. In human prostate cancers, strong AIB1 expression correlated with significant higher rate of proliferation and high p-AKT levels [142]. Results in cell lines agree with those from biopsies and animal models. One study found a decrease in IGF-I-induced colony formation after AIB1 depletion in the human breast cancer cell line MCF-7 [144]. Cyclin D1, Bcl-2, ERK2, IGF-IR, and IRS-1 expression levels were also reduced, even in the presence of anti-estrogens, suggesting that AIB1 activity was important to maintain AKT signaling independently of its role in ER-dependent signaling in the breast cancer. In the prostate cancer cell lines LNCaP and PC3, AIB1 overexpression promotes cell growth by augmenting AKT activity [71]. Additional increase in AKT activity is not observed when LNCaP cells are treated with androgens indicating that AIB1 functions through steroid-independent pathways to regulate AKT activity. In contrast, AIB1 downregulation results in reduced cellular size and decreased AKT activity in these cells. Accordingly, overexpression of AIB1 in LNCaP cells reveals augmented expression levels of several components of the PI3K/AKT pathway, including IGF-I, IGF-II, IRS-1, IRS-2, PIK3CA, and AKT1, whereas AIB1 depletion produces a decrease of these proteins [47]. Noteworthy, AIB1 is recruited to the promoter regions of IGF-I, IGF-II, IRS-1, IRS-2, PIK3CA, AKT1, and Bcl-2, suggesting a direct function of AIB1 in the transcription of these genes. These observations demonstrate that AIB1 is required for IGF-I-dependent cell proliferation and survival in breast and prostate cancers. AIB1 opens another way of promoting cell growth independently of steroid hormones, suggesting that overexpression of AIB1 may play a role in endocrine resistance through enhancing growth factor signaling and reveals a new platform for cancer therapy.

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

147

The ErbB/HER Signaling Pathway In oncology, the HER family or ErbB family is one of the most important tyrosine kinase signaling networks. The HER family, also known as the epidermal growth factor receptor (EGFR) family, is composed of four main members. These receptors are well-known mediators of cell proliferation, differentiation, migration, and apoptosis [145]. The activation of HER receptors by the epidermal growth factors (EGFs) initiates a signal transduction cascade that ultimately leads to gene expression. A correlation between overexpression of AIB1 and HER2/Neu positivity in breast tumors has been observed [141, 146]. Interestingly, AIB1 is a target of the EGFR/HER-downstream activated kinases, such as ERK1/2 and c-Abl kinase [147–149]. The phosphorylation of AIB1 by these kinases promotes its nuclear localization and stimulates its transactivation by increasing the ability to interact with ERa and other transcription factors (Fig. 7.3), and recruit components of the basal transcriptional machinery such as CBP/p300. On the other hand, AIB1 and the AIB1-Delta isoform can also potentiate HER2/Neu signaling by increasing its phophorylation levels [107, 123, 150]. Therefore, it created a positive feedback loop in which AIB1 increases HER2/Neu signaling, and in turn, HER2/Neu signaling stimulates AIB1 function as a coactivator. The final outcome of this cross talk is not clear, but it could also constitute another mechanism for resistance to Herceptin (humanized monoclonal antibody which targets and inhibits HER2/Neu) [151]. Moreover, clinical studies have demonstrated that ER-positive breast tumors overexpressing both AIB1 and HER2/Neu frequently undergo resistance to endocrine therapies due to an increase of the agonistic activity of tamoxifen versus the antagonistic activity upon the ER [152].

The NF-кB Signaling Pathway Nuclear factor-кB (NF-кB) comprises a family of transcription factors that are implicated in the expression of genes controlling immune response and inflammation as well as cell cycle and apoptosis [153]. NF-кB consists in dimers of proteins that contain the Rel homology dimerization domain. Inactive NF-кB is sequestered to the cytosol by inhibitor proteins known as IkBs. Exposure of the cell to a variety of stimuli, including cytokines such as interleukin-1 (IL-1) or tumor necrosis factora (TNF-a), leads to the activation of the IkB kinase complex (IKK complex) which phophorylates IkB triggering its ubiquitination and subsequent degradation. Once released from the inhibitory IkB complex, NF-kB is translocated to the nucleus where it transactivates the expression of numerous target genes. AIB1 and SRC-1 can interact and enhance NF-кB-mediated transcription upon physiological stimulus in cell culture [48, 49]. However, only AIB1 interacts with IKK complex. In response to TNF-a, IKK complex phosphorylates AIB1 triggering its translocation to the nucleus where it encounters and coactivates NF-kB [50]. Estrogen treatment also augments AIB1 phosphorylation status to activate estrogenmediated transcription [154–156]. It has been recently shown that IKKa leads the

148

M. Ferrero and J. Font de Mora

phophorylation of both AIB1 and ER-a in response to estrogen, stimulating their nuclear localization. Within the nucleus, all three components form a complex to transactivate cyclin D1 expression [157]. However, the cyclin D1 promoter is not only regulated by ER-a but also by NF-kB [158]. Hence, it is likely that AIB1 can target more than one transcriptional pathway to activate cyclin D1 expression. Noteworthy, IKK can also be activated by AKT leading to activation of NF-kB [159, 160]. Since AIB1 overexpression can increase AKT activity, this would represent another possible mechanism through which AIB1 might promote cyclin D1 expression and cell cycle progression (Fig. 7.3). All these findings suggest that both AIB1 overexpression and exposure of tumor cells to estrogens and/or cytokines promote AIB1 oncogenic activity raising its phosphorylated levels in the nucleus and augmenting its transcriptional coactivation capacity.

The Rb/E2F1 Signaling Pathway The mammalian cell cycle is a process orchestrated by the expression and activation of a series of genes that form a highly organized and regulated network. The Rb/E2F1 pathway regulates G1/S transition and constitutes a key regulatory mechanism in the cell cycle [161]. Deregulation of any of the processes governing cell cycle is frequently associated with growth abnormalities including cancer. As a transcriptional coactivator, the discovery that AIB1 could also interact with E2F1 gave a novel role for AIB1 in cell cycle progression. AIB1 promoted cell proliferation by stimulating the expression of E2F1-responsive genes, such as cyclin E, cyclin A, cyclin B, Cdk2, p107, and E2F1 itself [58]. This increased proliferation occurred even in the presence of anti-estrogens, indicating again that AIB1 could function independently of steroids. Moreover, ChIP assays showed that the complex E2F1/AIB1 was recruited to the AIB1 promoter, suggesting a positive feedback regulatory loop in which high levels of both AIB1 and E2F1 were maintained [128, 162]. This result could explain a mechanism to achieve high levels of AIB1 independently of its gene amplification. Since it seems that AIB1 overexpression can bypass the growth-suppressive effects of anti-estrogens by activating E2F1- responsive gene expression, these findings are of great importance to explain how tumors acquire endocrine resistance.

Invasiveness Mechanisms AIB1 is also implicated in late stages of tumorigenesis: cell invasiveness and metastasis. The first observation suggesting a novel role for steroid hormones in cell motility and invasiveness was done in Drosophila [163]. It was reported that mutations in Taiman, the AIB1 orthologue in Drosophila, caused defects in migration of the border cells in the ovary. Another complementary study revealed that MCF-7 cells respond to 17-b-estradiol (E2) treatment by extending motile lamellipodia, but this behavior was suppressed in the presence of anti-estrogen [164]. These findings

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

149

suggested that steroids might stimulate invasive cell behavior. In the last few years, great efforts have been made to determine the mechanisms through which steroid receptor coactivators are affecting these processes. Clinical studies have correlated AIB1 not only with high cell proliferation and growth but also with metastatic stages of cancer [94, 99, 101, 102, 109], suggesting a role for AIB1 in tumor invasiveness and metastasis. The hallmark of cancer cells is to achieve the ability to migrate and invade surrounding tissues, reach blood vessels, and colonize distant organs. One important event in this process is the epithelial-mesenchymal transition or EMT. During this process, epithelial cells loose adhesion by repressing E-cadherin expression and acquire the capacity to migrate. Several transcription factors are known to suppress E-cadherin expression, including Twist and Snail [165]. Interestingly, a recent report correlated high levels of AIB1 with high levels of Twist [166]. The authors also observed an inverse correlation between the U-box-like ubiquitin ligase CHIP and AIB1 levels in breast cancer cells, indicating a role for CHIP in AIB1 degradation through the ubiquitin-proteasome system. CHIP levels were diminished with the concomitant increase of AIB1 levels in advanced tumor stages. That increase in AIB1 levels correlated with high levels of Smad-2 and EMT markers, including b-catenin, Twist, and vimentin. These results clearly indicate that high AIB1 levels might confer an invasive advantage through the stimulated expression of Twist. Although it has been demonstrated that SRC-1 does not affect either tumor initiation or growth, it seems to have importance in metastasis since its depletion in PyMT mice (SRC-1−/−/PyMT) drastically decreases the intravasation of mammary tumor cells and the frequency of metastasis [167]. Interestingly, SRC-1 correlates inversely with E-cadherin and potentiates PEA3-mediated Twist expression [168]. Hence, SRC-1 and AIB1 can both promote EMT and invasiveness. However, cells do not only have to lose adhesion but are also required to destruct biological barriers such as the basement membrane to make migration and invasion possible. That purpose can be achieved by the matrix metalloproteases (MMPs) that degrade the extracellular matrix proteins (ECMs). In line with this notion, AIB1 overexpression stimulates invasiveness and it functions as an AP-1 coactivator to upregulate the expression of MMP-7 and MMP-10 in breast cancer cell lines [169]. Similar results are obtained in prostate cancer cell lines in which AIB1 overexpression enhances coactivation of the transcription factors PEA3 (an Ets family member) and AP-1, thus increasing the levels of MMP-2 and MMP13 and resulting in greater ability to migrate [170]. These different expression patterns suggest that AIB1-induced MMP expression depends on the cellular context. In transgenic rodents for lung metastasis (MMTV-polyomavirus middle T antigen or PyMt mice), AIB1 deficiency extends tumor latency and suppresses lung metastasis [171]. Moreover, cells derived from these mice (AIB1−/−/PyMT and WT/ PyMT) expressed low levels of MMP-2 and MMP-9 when AIB1 was knockeddown. The expression of these two MMPs was possible through PEA3 transactivation activity. The same report correlates positively the expression of AIB1, PEA3, MMP-2, and MMP-9 in breast cancer specimens. AIB1 has also been associated with ER81, another member of the Ets family, to stimulate the expression of MMP-1 [42]. In this process, HER2/Neu is involved by activating ER81 together with its coactivators AIB1 and p300 and facilitating ER81-mediated transcription (Fig. 7.3).

150

M. Ferrero and J. Font de Mora

Of interest, the HER2/Neu promoter is a target for Ets (reviewed in 172]. Since AIB1 can interact and coactivate this transcription factor, AIB1 overexpression might be directly implicated in HER2/Neu overexpression. Again, the same molecules participate in a vicious cycle to favor proliferation and endocrine resistance. Therefore, it seems that high levels of AIB1 may play a role in tumor invasion and metastasis in addition to its function in cell proliferation and tumor growth. Blocking the PEA3/or ER81/AIB1 axis in cancer may represent a good approach to control invasiveness and cancer progression. A large body of compelling evidence reveals that AIB1 is an oncogene. AIB1 amplification and/or its overexpression occur in a wide range of human tumors, including breast, ovarian, prostate, pancreatic, gastric, hepatic, and colorectal carcinomas [16, 98, 100, 101, 103, 104, 106, 108, 110, 112]. Experiments in rodents have demonstrated that AIB1 alone is sufficient to initiate tumorigenesis [72]. In contrast, AIB1 deficiency protects from tumors [122–125, 127]. As described above, the mechanisms of AIB1 to promote cancer initiation, progression, and endocrine resistance involve a variety of signaling pathways, including ER, IGF/PI3K/AKT, HER2, NF-kB, and Ets as well as cell cycle regulation. However, the levels of AIB1 protein are not the unique determinants in AIB1 oncogenic function. Posttranslational modifications such as phosphorylation have also a great importance [155]. The phosphorylation of AIB1 is necessary for its activity and can be induced by steroids, cytokines, and growth factors [50, 147–149, 156]. Therefore, a correct understanding of the processes controlling AIB1 activity and half-life is necessary for future experimentation and drug discovery.

Cell Cycle Regulation by AIB1 The cell cycle is a highly regulated process by which cells duplicate genetic dosage and end up with cell division. It involves a complex dynamic transcriptional program through which cyclins, cyclin-dependent kinases, checkpoint regulators, DNA repair machinery, and replication proteins are timely expressed. Extracellular signals along with intracellular signals decide whether a cell will divide or not. The Rb/ E2F1 pathway plays the major role in the initiation of the cell cycle [173]. Firstly, E2F1 is associated to the tumor suppressor protein Rb, and this association prevents E2F1-mediated transcription. Upon growth factor-mediated stimulation, Ras is activated inducing an increase of cyclin D1 levels. In this way, association of cyclin D1 with Cdks is favored, promoting Rb phosphorylations and its consequently release from the E2F1 interaction and inhibition. Once released from Rb, E2F1 initiates the expression of molecules required for the entry and completion of the S-phase, such as cyclin E, cyclin A, and DNA replication proteins. The E2F family comprises 8 members referred to as E2F1-8. While E2F1-3 are positive regulators of the cell cycle, E2F4-8 function as transcriptional repressors. Because both cyclin D1 and E2F1 are important initiators of cell cycle, it is not surprising to find them deregulated in cancer [174, 175].

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

151

Cyclin D1 is one of the major ER-downstream gene targets [139]. It has been demonstrated that the functional coupling of ER at the cyclin D1 promoter is achieved through the actions of several intermediary proteins including AIB1. In fact, AIB1 is recruited to the cyclin D1 promoter, and moreover, its overexpression can enhance the ability of ER to interact with this promoter in an estrogen-dependent fashion [140, 157]. SRC-1 and TIF2 can also enhance cyclin D1 expression. However, in MCF-7 cells, ER preferentially interacts with AIB1 after estrogen treatment, highlighting a specific role for AIB1 in mediating estrogen-dependent gene transactivation in breast cancer cells. Nonetheless, there are ER-negative tumors that also overexpress cyclin D1 raising the possibility that its expression could also be regulated through other pathways. For instance, in certain mammary carcinoma cell lines, cyclin D1 expression was not as potently induced by estrogen as by serum, suggesting that other pathways are involved in its expression [176]. Furthermore, cyclin D1 promoter can be regulated by several transcription factors, including ER, STATs, AP-1, and NF-kB (reviewed in [177]). One possible way to activate cyclin D1 expression in a hormone-independent manner might involve AP-1. Nakuci et al. have demonstrated that c-Jun/AP-1 complex in the cyclin D1 promoter recruits BRCA-IRIS (a BRCA1 splice variant) alone or together with coactivators [178]. Since AIB1 is also a coactivator of AP-1 [47], it would be not surprising that AIB1 overexpression increases cyclin D1 levels through this pathway (Fig. 7.3). In addition, AIB1 can enhance NF-kB and STAT6 transcriptional activities [49, 179] increasing the possibilities. It seems clear that AIB1 plays an important function in activating cyclin D1 expression. Animals and AIB1 gene interfering in cancer cell lines also confirm that observation. When AIB1 is depleted in breast and prostate cancer cell lines the levels of cyclin D1 decrease [144, 180]. And in animals, knockdown of AIB1 results in lower levels of cyclin D1 in comparison to WTs [123, 124]. All these findings suggest that overexpression of AIB1 may confer a cell growth advantage by increasing the expression of cell cycle regulators such as cyclin D1. The overexpression of cyclin D1 might be one of the first steps in cancer progression induced by AIB1. AIB1 associates with the cell cycle positive regulator E2F1 but not with the cell cycle repressor E2F4 [58], participating in the core machinery for cell cycle progression. In fact, AIB1 depletion inhibited both the E2-dependent and-independent cell proliferation [58, 128]. This depletion decreased S-phase population with a concomitant increase in G0/G1-phase, suggesting a role of AIB1 in G1 to S-phase progression. In contrast, overexpression of AIB1 in breast cancer cell lines increased cell proliferation independently of E2, and more intriguingly, the elevated levels of AIB1 could promote the cell cycle even in the continuous presence of anti-estrogens [58]. In an attempt to find out why AIB1 overexpression could enhance cell proliferation under different conditions, these authors investigated the expression of cell cycle regulators when AIB1 levels were elevated. They found an increase in cyclin E, cyclin A, cyclin B, Cdk2, p107, and E2F1 expression levels; all of them are crucial genes for G1/S progression and well-characterized targets of the E2F1 transcriptional activity. Moreover, they observed that AIB1 was recruited independently of estrogen to the promoters of all those genes. This recruitment increased as cells

152

M. Ferrero and J. Font de Mora

entered S-phase [128], suggesting that AIB1 promotes cell cycle by facilitating the expression of genes essential for G1/S transition. In summary, all these findings demonstrate that AIB1 was an E2F1 coactivator and that its overexpression could override the anti-estrogen action of tamoxifen by functionally coactivating E2F1 allowing cell cycle progression. Importantly, AIB1 expression is cell cycle regulated with its protein levels peaking at G1–S-phase [128]. Moreover, Avivar et al. observed a correlation between BrdU incorporation and AIB1 nuclear staining in the mammary glands of both WT and AIB1 transgenic mice, whereas non-proliferative cells contained lower levels of AIB1 which were restricted to the cytoplasm [121]. This indicates a novel function of AIB1 during or after S-phase, in addition to its function during G1 as described previously. Interestingly, AIB1 promoter can also be regulated by the E2F1–AIB1 complex [128, 162], and the recruitment is increased at late G1 and decreased during S-phase, supporting the notion of regulated AIB1 expression during the cell cycle. These findings are important because it creates a positive feedback loop between high level of AIB1 and E2F1 and also might explain endocrine resistance in human cancers. AIB1 increases the expression of numerous cell cycle regulators, including cyclin D1, cyclin E, cyclin A, E2F1, and a few others. Hence, AIB1 plays a direct role in cell cycle regulation. One possible stepwise progress in ER-positive breast tumor evolution could be as follows: (1) during cancer initiation, AIB1 would coactivate ER function by interacting with ligand-bounded ER; (2) as tumor stages, amplification, and overexpression of AIB1 would increase the levels of PI3K/AKT signaling pathway components; (3) more active AKT and/or more activated HER2/Neu could stimulate ER and AIB1 phosphorylation resulting in increased hormone-independent ER-mediated transcription; (4) high AIB1 levels could also potentiate E2F1 function further increasing its levels and boosting cell cycle progression; (5) in addition, repression of E-cadherin starts to be effective and EMT initiates; and (6) PEA3-mediated expression of MMPs favors invasion of peripheral tissues and expression of metastasic genes such as TWIST mediates metastasis.

IGF-I Is a Mediator of AIB1 Signaling There are strong evidences, both in vitro and in vivo, that receptor tyrosine kinases play a key role in the formation and progression of human cancers. In particular, the insulin-like growth factor receptor (IGF-IR), a tyrosine kinase receptor for IGF-I and IGF-II, and to a lesser extent for insulin, has been well documented in cell culture, animal studies, and humans to play a role in malignant transformation, progression, and protection from apoptosis and metastasis [181]. Transgenic mouse models have shown that AIB1 can act as an oncogene [72, 121], giving rise to a premalignant hyperplastic mammary phenotype as well as to a high incidence of mammary tumors that are primarily ER-positive. In these models, the AIB1 transgene is responsible for continued activation of the insulin-like growth factor-I

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

153

receptor, suggesting a role for the stimulation of PI3K/AKT/mTOR pathway in the premalignant phenotype and tumor development. Importantly, treatment of AIB1 transgenic mice with the mTOR inhibitor RAD001 reverts the premalignant phenotype [182]. Moreover, combination of mTOR inhibition and ER-targeted endocrine therapy with 4-hydroxytamoxifen improves the outcome of the subset of ER-positive breast cancers overexpressing AIB1. This data provides preclinical support for the clinical development of RAD001 and suggests that AIB1 may be a predictive factor of RAD001 response. So far, IGF-I seems to be an important mediator of AIB1-induced tumor development. Although AIB1 is an important cofactor for estrogen and progesterone receptors, both animal models and human cancer cell line studies support a role for IGF-I in AIB1-dependent tumors. In human breast cancer cells, AIB1 is rate-limiting for IGF-I-dependent phenotypic changes and gene expression controlling the cell cycle and apoptosis [144]. These results reveal that AIB1 is required for IGF-Iinduced proliferation, signaling, cell survival, and gene expression in human breast cancer cells, independently of its role in estrogen receptor signaling. However, an N-terminal truncated isoform of AIB1 found being overexpressed in breast cancer can also induce mammary hyperplasia in mice without affecting serum IGF-I circulating levels [120], suggesting that perhaps other oncogenic mechanisms independent of IGF-I are also being activated by AIB1. Therefore, IGF-I could be just one of the several pathways activated by AIB1 that contribute together to the final outcome in cancer initiation and progression. Noteworthy, AIB1 upregulates the expression of multiple genes in the IGF-I/ AKT signaling pathway that are involved in cell proliferation and survival [47]. Of interest, prostate tumors of treated patients with the proteasome inhibitor Bortezomib and cell-derived cell lines underwent an unexpected higher proliferative expansion due to increased levels of AIB1 followed by increased AKT activation levels. Therefore, dual therapies targeting both the proteasome and AKT activation may improve efficacy [183]. In addition, AIB1 is recruited to the promoter of IRS2 and IGF-I by the transcription factor activator protein-1 AP-1, favoring proliferation and survival of cancer cells in a ligand-independent manner. These results strongly support the earliest observation connecting AIB1 and somatic growth through the regulation of IGF-I expression. Accordingly, the small size phenotype of AIB1 knockout mice reflects both altered regulation of IGF-1 gene expression in specific tissues and a cell-autonomous defect of response to IGF-1, including ineffective transcriptional activities by several classes of regulated transcription factors under specific conditions [69]. Regulation of IGF-I circulating levels has also been reported to be mediated by the regulation of IGFBP3 expression (see above in the role of AIB1 during development). Transcription of IGFBP3 is mediated by VDR and AIB1, resulting in increased IGF-I stability [73]. Novel insights in IGF-I-mediated oncogenesis by AIB1 were obtained by generating AIB1(+/+), AIB1(+/−), and AIB1(−/−) mice harboring the mouse mammary tumor virus/v-Ha-ras transgene that induces breast tumors [122]. This study revealed that mammary tumorigenesis was significantly suppressed in AIB1 deficient mice independently of ovarian hormone levels, suggesting that AIB1 and ovarian hormones contribute to mammary carcinogenesis through different pathways.

154

M. Ferrero and J. Font de Mora

Importantly, AIB1 deficiency also caused partial resistance to IGF-I because of a significant reduction in the insulin receptor substrates. The impaired IGF-I signaling was responsible in part for the suppression of mammary tumorigenesis and metastasis caused by inhibition of cell proliferation and migration. These interesting results support the notion for dual therapies to control breast cancer by targeting AIB1 together with antihormone therapy. Hence, further studies should be done focused in developing new strategies to override AIB1 oncogenic signals.

Overexpression of AIB1 and HER2/NEU Correlates with Tamoxifen Resistance Primary tumors are mostly classified as ER-positive or HER2/Neu-positive based in the staining results with specific antibodies. Therapy with the antiestrogenic compound tamoxifen has been frequently given to breast cancer patients with positive staining for ER. However, antiestrogen resistance is a major clinical problem in the treatment of breast cancer [184]. Noteworthy, development of resistance correlates with the novel expression of HER2/Neu [185]. Recent studies have provided important clues for the understanding on how this process is being triggered. Two independent mechanisms have been demonstrated so far, and curiously, AIB1 has been implicated in both of them. One mechanism is upstream of HER2/Neu controlling its transcription as a result of a competitive regulatory pathway. The second mechanism is downstream of HER2/Neu and depends on a bidirectional cross talk between HER2/Neu and the AIB1/ER transcriptional complex. Both mechanisms will be discussed further ahead in this section. Initially, broad studies with breast and ovarian tumors revealed a correlation between ER and PR positivity with AIB1 amplification [106, 112], suggesting a cooperative pathway for triggering cancer. Since AIB1 is an important cofactor for ER and PR transcriptional activity, it seemed reasonable at that time that AIB1 overexpression that ensued from amplification at 20q12-13 would enhance ER and PR signaling and result in the predicted pathological outcome. Subsequent studies demonstrated that AIB1 is overexpressed at the mRNA level in up to 60% of primary breast carcinomas; however, only 5% of these tumors show DNA amplification [110, 141]. In contrast with previous reports, one later study showed no correlation between AIB1 overexpression and ER and PR positivity, but instead it showed correlation with strong protein staining for HER2/Neu [141]. In addition, amplification of AIB1 and EGFR genes has been associated with lymph node metastasis of oral squamous cell carcinoma [186]. These additional findings suggested that AIB1 overexpression may promote breast cancer by different mechanisms involving either HER2/Neu overexpression or steroid receptor coexpression. In this way, AIB1 could be in the middle of the cross talk between different signals that modulate the pathological behavior of the tumor. Studies of endocrine therapy have revealed differential advantages depending on the association of AIB1 overexpression with ER or with HER2/Neu. Failure of the antitumor activity of tamoxifen is actually determined by both the levels of and

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

155

the interaction between AIB1 and HER2/Neu [187]. In fact, AIB1 overexpression correlates with increased levels of HER2/Neu and its ligand and resistance to tamoxifen therapy [141, 152, 188]. Based on the variety of tumors overexpressing AIB1 and the prognostic differences depending on the expression of other markers, it is clear that more data needs to be contrasted in order to have the big picture of all the signaling pathways leading to AIB1. For instance, it has been observed that ERb positive tumors do not express ERa protein, suggesting that a substantial fraction of ERb positive tumors could falsely be considered estrogen receptor negative [109]. Several molecular mechanisms leading to tamoxifen resistance have been postulated. Chromatin immunoprecipitation (ChIP)-on-chip analyses in ER-positive MCF-7 cells allowed the identification of an intronic ER-binding site within HER2/Neu gene. Noteworthy, treatment with either tamoxifen or estrogen induced the promoter recruitment of the paired box 2 gene product (PAX2), a crucial mediator of HER2/Neu repression by ER [189]. Conversely, expression of AIB1 competed with PAX2 for binding to the HER2/Neu-regulatory element, and this resulted in increased HER2/Neu transcription and cell proliferation in the presence of tamoxifen. Positive immunostaining for PAX2 corresponded to a significantly improved recurrence-free survival in patients relative to PAX2-negative tumors. Furthermore, within the PAX2-positive tumors, only those that were also positive for AIB1 had a worse clinical outcome than the tumors that were AIB1 negative. These results reveal the intrinsic role of AIB1-PAX2 expression in the development of tamoxifen resistance (Fig. 7.4). There are yet two mechanisms emanating from HER2/Neu cross talk that are also suggested to interplay an essential role in tamoxifen resistance. One mechanism promotes AIB1 transcriptional activity and the other inhibits its degradation, thus resulting in higher protein levels (Fig. 7.4). HER2/Neu is a cell surface tyrosine-kinase receptor that gets auto-phosphorylated upon activation, recruiting several transducer molecules that recognize and bind to the new phosphorylated sites. In this way, several signaling pathways are also becoming activated and transduce the signal to key regulatory molecules controlling metabolism, cell cycle, and survival and transcriptional regulation to generate the proper response to the incoming signal. Two of these pathways are the Ras/ERK pathway and the PI3K/AKT pathway (Figs. 7.1 and 7.4). Activation of both pathways results in the phosphorylation of ERa in its AF-1 domain. However, only induced phosphorylation by AKT at Ser167 is important for enhanced interaction with AIB1 and tamoxifen resistance [190]. In addition, activated ERK1/2 results in AIB1 phosphorylation allowing the recruitment of the transcriptional coactivators CBP/p300 [149]. As a result, activated macromolecular complexes with strong histone acetyltransferase activity are being formed around phosphorylated AIB1 and its bounded hormone receptor. Hence, in ER-positive tumors, ERK and AKT activation augments AIB1-ER transcriptional activity in a way that may even override antiestrogenic activity of tamoxifen (Fig. 7.4). Amplification of AIB1 gene at its genetic locus in 20q12 has been shown to be a mechanism leading to AIB1 overexpression [16]. However, such an amplification only occurs in only 5% of the tumors out of the approximately 60% that overexpress AIB1. Therefore, other mechanisms must be altered in tumors that increase AIB1 protein levels. One of these mechanisms is the activation of the PI3K/AKT pathway

156

M. Ferrero and J. Font de Mora

Fig. 7.4 Mechanisms for tamoxifen resistance. Three different mechanisms have been postulated so far for the development of resistance to tamoxifen. Tamoxifen-bound ER recruits PAX2 and represses Her2/Neu transcription [185]. Patients with PAX2 negative tumors develop resistance and display poorer recurrence-free survival. Activation of the PI3K/AKT pathway prevents proteasomal degradation of AIB1 [66], thus resulting in its overexpression and overriding antiestrogenic effect of tamoxifen. Finally, phosphorylation of AIB1 by ERK1 and 2 allows the recruitment of p300 to the coactivation complexes and enhancing their transcriptional activity [140]

that stabilizes AIB1 protein by inhibiting its ubiquitination and its proteasomal degradation [66]. As a result, AIB1 protein levels arise and may hypersensitize the ER-AIB1 transcriptional complex that even in the presence of tamoxifen may displace the equilibrium to the transcription of genes involved in tumor progression (Fig. 7.4). The result is again the resistance to the antiestrogenic therapy. Similar mechanisms could also explain the resistance to the aromatase inhibitor letrozole [191]. Taken all together, available data supports that AIB1 overexpression is involved in antiestrogenic resistance and, therefore, should be considered a new target for combined therapy.

Predictive and Prognostic Factors: Targeting AIB1 in Cancer Therapy Since the early discovery and characterization of AIB1 in 1997, efforts have been made to correlate the expression of AIB1 with either natural progression of the disease (“prognostic factor”) or with response to tamoxifen (“predictive factor”),

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

157

and to analyze potential correlations with tumor characteristics. As a prognostic factor, patients with breast tumor overexpressing AIB1 had significantly shorter disease-free and overall survival times after surgery [114]. Tamoxifen is one of the most successful treatments of breast cancer. However, resistance to tamoxifen is frequently developed [184]. Response to tamoxifen has been associated to the expression of AIB1 in tumor cells and varies depending on the coexpression of other molecules, such as HER2/Neu, ERa, and PAX2 [187, 189, 192]. Conversely, expression of ERa corepressors NCOR1 [193] or the scaffold attachment factors SAFB1 and SAFB2 (SAFB) [194] shows that low protein levels of these negative regulators of ERa predict poor prognosis of breast cancer patients. Special interest has been given in the scientific literature to the length of polyglutamine stretch, which has been studied as a risk factor for cancer. This stretch constitutes a polymorphism ranging from 26 to 32 glutamines with a heterozygosity of 54% [195]. Although AIB1 repeat genotype does not influence postmenopausal breast cancer risk [44], it showed a significative influence in BRCA1/2-associated breast cancer risk [196]. A more careful study revealed that longer repeat length correlated with elevated risk among BRCA1 mutation carriers, but not among BRCA2 mutation carriers [197]. However, contradictory results were obtained later showing no significant effect of AIB1 genetic variation on breast cancer risk, neither in BRCA1 mutation carriers nor in BRCA2 carriers [198, 199]. In contrast, ovarian cancer patients with short AIB1 genotype (28 glutamines or less] showed shorter time to disease recurrence and decreased overall survival [43]. To settle down all these discrepancies, a more recent study with a very large international sample size to detect previously reported effects concluded that the AIB1 glutamine repeat does not substantially modify risk of breast cancer in BRCA1 and BRCA2 mutation carriers [45]. Independently of all the controversy, it is still not clear enough the real function of this domain in AIB1 stability or binding capacity to other coactivators, implementing other non-studied partners for risk of cancer development and/or prognosis. Other predictive factors have been focused in the phosphorylation status of the ERa. Low phosphorylation of ERa at Ser118 was associated with significantly improved disease-free and overall survival, whereas AIB1 overexpression was significantly associated with activated ERK, AKT, HER2, and phosphorylation of ERa at Ser118 [200]. These results suggested that phosphorylation of ERa Ser118 affects survival in ER-positive breast cancer and could be helpful in distinguishing patients who are likely to benefit from endocrine therapy alone from those who are not. As discussed in the preceding section, association of AIB1 expression with HER2/Neu in tumors correlates with poor prognosis and tamoxifen resistance. In these cases, tamoxifen behaves like an estrogen agonist like in ER-positive breast cancer cells, resulting in de novo resistance. Based on crosses between AIB1 knockouts and HER2/Neu transgenic animals, HER2/Neu/AIB1(+/−) tumors display decreased phosphorylated HER2/Neu, cyclin D1, and cyclin E. Cellular proliferation is also reduced in these tumors, and AKT and JNK activation is barely detectable [123]. Similarly, v-ras-induced tumors are also reduced and less aggressive in the absence of AIB1 (also explained at the end of Sect. 7.7]. These data

158

M. Ferrero and J. Font de Mora

indicate that AIB1 is required for the oncogenic activity of HER2/Neu and v-ras. It is therefore predictable that reducing AIB1 levels or activity could potentiate therapies aimed at inhibiting HER2/Neu signaling in the mammary epithelium of breast cancers or Ras signaling in colon and pancreas cancers where AIB1 is also overexpressed. In fact, knockdown of AIB1 levels can eliminate the ER/HER2/ Neu crosstalk and restores the inhibitory effect of tamoxifen on cell proliferation [201, 202]. Knockdown of AIB1 in prostate cancer cells where it is also frequently overexpressed suppresses cell growth [203]. Resistance to the treatment against HER2/Neu activity with the antibody Herceptin has also been observed, but combination of this therapy with tamoxifen can improve the results [204]. As a factor with clinical significance, nuclear expression of AIB1 was initially correlated with the ERa status, and patients with AIB1 nuclear expression tended to be successfully treated by hormonal therapy [205]. Estrogen-bound ER produces a specific conformation enabling the recruitment of p160 class of coactivators [206]. This recruitment is sufficient for gene activation and for the growth stimulatory actions of estrogen in breast cancer supporting a model in which ER cofactors play unique roles in estrogen signaling [133]. Conversely, the tamoxifen-bound ER disables coactivator binding but recruits corepressors instead. In this regard, it would be of therapeutic interest to develop molecules that create steric hindrance or simply prevent ER interaction with coactivators such as AIB1 even in the presence of estrogen. Small peptide-like molecules mimicking LXXLL motifs with high specific affinity for estrogen-bounded ER could be an interesting approach for further investigation. Several preliminary approaches have been developed to target AIB1. Highthroughput screens have been used to discover small molecules that act as coactivator binding inhibitors that are able to disrupt this interaction of ER with AIB1 and ligands [207]. For this purpose, it was designed and optimized a set of time-resolved fluorescence resonance energy transfer (TR-FRET) assays to monitor the interaction of ER with SRC-3 and ligands. This system opens exciting expectancies for its high sensibility and its capacity to distinguish between conventional antagonists and coactivator binding inhibitors. Another approach used a combination of a mammalian two-hybrid screen to select compounds that disrupt the interaction between the ligand binding domain of ERa and AIB1 and a virtual screen to select compounds that fit onto the surface of ERa that interacts with LXXLL motifs, based on the X-ray crystal structure of the ERa complexed with a LXXLL peptide [208]. In contrast to classical ER antagonists, these novel inhibitors poorly displace estradiol in the ER-ligand competition assay. These small molecules may represent new classes of ER antagonists and may have the potential to provide an alternative for the current anti-estrogen therapy. MicroRNAs (miRNA) are single-stranded RNA molecules of 21–23 nucleotides in length, partially complementary to one or more messenger RNA (mRNA) molecules, and their main function is to downregulate gene expression. The microRNA Mir-17-5p has extensive complementarity to AIB1 mRNA and can downregulate AIB1 expression in cell culture experiments through translational inhibition [209]. Expression levels of Mir-17-5p have pathological consequences since they are low

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

159

in breast cancer cell lines, and downregulation of AIB1 by Mir-17-5p decreased proliferation of breast cancer cells. Importantly, miRNAs are on the scope for diagnostic potential in human cancer and even miRNA-based cancer therapies may be on the horizon [210]. The development of these novel inhibitors for cancer therapy requires further clinical studies. Undoubtedly, the next decade will see new insights into the basic biology of AIB1 which will facilitate the potential clinical applications which we have discussed in the context of this chapter.

References 1. Foster, J.R. 2001, Int J Exp Pathol, 82, 171. 2. Lupulescu, A.P. 1996, Cancer, 78, 2264. 3. Adjei, A.A., and Hidalgo, M. 2005, J Clin Oncol, 23, 5386. 4. Radice, P. 2002, J Exp Clin Cancer Res, 21, 9. 5. Fodde, R. 2002, Eur J Cancer, 38, 867. 6. Pan, S.Y., and DesMeules, M. 2009, Methods Mol Biol, 472, 191. 7. Chida, Y., Hamer, M., Wardle, J., and Steptoe, A. 2008, Nat Clin Pract Oncol, 5, 466. 8. Ziegler, J.L., and Buonaguro, F.M. 2009, Front Biosci, 14, 3455. 9. LeRoith, D., Novosyadlyy, R., Gallagher, E.J., Lann, D., Vijayakumar, A., and Yakar, S. 2008, Exp Clin Endocrinol Diabetes, 116 Suppl 1, S4. 10. Pischon, T., Nothlings, U., and Boeing, H. 2008, Proc Nutr Soc, 67, 128. 11. Mavrou, A., Tsangaris, G.T., Roma, E., and Kolialexi, A. 2008, Anticancer Res, 28, 401. 12. Hermanson, O., Glass, C.K., and Rosenfeld, M.G. 2002, Trends Endocrinol Metab, 13, 55. 13. Lonard, D.M., and O’Malley, B.W. 2006, Cell, 125, 411. 14. Onate, S.A., Tsai, S.Y., Tsai, M.J., and O’Malley, B.W. 1995, Science, 270, 1354. 15. Hong, H., Kohli, K., Garabedian, M.J., and Stallcup, M.R. 1997, Mol Cell Biol, 17, 2735. 16. Anzick, S.L., Kononen, J., Walker, R.L., Azorsa, D.O., Tanner, M.M., Guan, X.Y., Sauter, G., Kallioniemi, O.P., Trent, J.M., and Meltzer, P.S. 1997, Science, 277, 965. 17. Chen, H., Lin, R.J., Schiltz, R.L., Chakravarti, D., Nash, A., Nagy, L., Privalsky, M.L., Nakatani, Y., and Evans, R.M. 1997, Cell, 90, 569. 18. Li, H., Gomes, P.J., and Chen, J.D. 1997, Proc Natl Acad Sci U S A, 94, 8479. 19. Suen, C.S., Berrodin, T.J., Mastroeni, R., Cheskis, B.J., Lyttle, C.R., and Frail, D.E. 1998, J Biol Chem, 273, 27645. 20. Takeshita, A., Cardona, G.R., Koibuchi, N., Suen, C.S., and Chin, W.W. 1997, J Biol Chem, 272, 27629. 21. Torchia, J., Rose, D.W., Inostroza, J., Kamei, Y., Westin, S., Glass, C.K., and Rosenfeld, M.G. 1997, Nature, 387, 677. 22. Carapeti, M., Aguiar, R.C., Chase, A., Goldman, J.M., and Cross, N.C. 1998, Genomics, 52, 242. 23. Kalkhoven, E., Valentine, J.E., Heery, D.M., and Parker, M.G. 1998, Embo J, 17, 232. 24. Huang, Z.J., Edery, I., and Rosbash, M. 1993, Nature, 364, 259. 25. Belandia, B., and Parker, M.G. 2000, J Biol Chem, 275, 30801. 26. Chen, S.L., Dowhan, D.H., Hosking, B.M., and Muscat, G.E. 2000, Genes Dev, 14, 1209. 27. Litterst, C.M., Kliem, S., Marilley, D., and Pfitzner, E. 2003, J Biol Chem, 278, 45340. 28. Chen, Y.H., Kim, J.H., and Stallcup, M.R. 2005, Mol Cell Biol, 25, 5965. 29. Kim, J.H., Li, H., and Stallcup, M.R. 2003, Mol Cell, 12, 1537. 30. Lee, Y.H., Campbell, H.D., and Stallcup, M.R. 2004, Mol Cell Biol, 24, 2103. 31. Zhang, A., Yeung, P.L., Li, C.W., Tsai, S.C., Dinh, G.K., Wu, X., Li, H., and Chen, J.D. 2004, J Biol Chem, 279, 33799.

160

M. Ferrero and J. Font de Mora

32. Ding, X.F., Anderson, C.M., Ma, H., Hong, H., Uht, R.M., Kushner, P.J., and Stallcup, M.R. 1998, Mol Endocrinol, 12, 302. 33. Heery, D.M., Kalkhoven, E., Hoare, S., and Parker, M.G. 1997, Nature, 387, 733. 34. Li, H., and Chen, J.D. 1998, J Biol Chem, 273, 5948. 35. Voegel, J.J., Heine, M.J., Tini, M., Vivat, V., Chambon, P., and Gronemeyer, H. 1998, Embo J, 17, 507. 36. Leo, C., Li, H., and Chen, J.D. 2000, J Biol Chem, 275, 5976. 37. Savkur, R.S., and Burris, T.P. 2004, J Pept Res, 63, 207. 38. Brown, K., Chen, Y., Underhill, T.M., Mymryk, J.S., and Torchia, J. 2003, J Biol Chem, 278, 39402. 39. Demarest, S.J., Deechongkit, S., Dyson, H.J., Evans, R.M., and Wright, P.E. 2004, Protein Sci, 13, 203. 40. McInerney, E.M., Rose, D.W., Flynn, S.E., Westin, S., Mullen, T.M., Krones, A., Inostroza, J., Torchia, J., Nolte, R.T., Assa-Munt, N., Milburn, M.V., Glass, C.K., and Rosenfeld, M.G. 1998, Genes Dev, 12, 3357. 41. Spencer, T.E., Jenster, G., Burcin, M.M., Allis, C.D., Zhou, J., Mizzen, C.A., McKenna, N.J., Onate, S.A., Tsai, S.Y., Tsai, M.J., and O’Malley, B.W. 1997, Nature, 389, 194. 42. Goel, A., and Janknecht, R. 2004, J Biol Chem, 279, 14909. 43. Li, A.J., Lerner, D.L., Gapuzan, M.E., and Karlan, B.Y. 2005, Cancer Epidemiol Biomarkers Prev, 14, 2919. 44. Haiman, C.A., Hankinson, S.E., Spiegelman, D., Colditz, G.A., Willett, W.C., Speizer, F.E., Brown, M., and Hunter, D.J. 2000, Breast Cancer Res, 2, 378. 45. Spurdle, A.B., Antoniou, A.C., Kelemen, L., Holland, H., Peock, S., Cook, M.R., Smith, P.L., Greene, M.H., Simard, J., Plourde, M., Southey, M.C., Godwin, A.K., Beck, J., Miron, A., Daly, M.B., Santella, R.M., Hopper, J.L., John, E.M., Andrulis, I.L., Durocher, F., Struewing, J.P., Easton, D.F., and Chenevix-Trench, G. 2006, Cancer Epidemiol Biomarkers Prev, 15, 76. 46. Lee, S.K., Kim, H.J., Na, S.Y., Kim, T.S., Choi, H.S., Im, S.Y., and Lee, J.W. 1998, J Biol Chem, 273, 16651. 47. Yan, J., Yu, C.T., Ozen, M., Ittmann, M., Tsai, S.Y., and Tsai, M.J. 2006, Cancer Res, 66, 11039. 48. Na, S.Y., Lee, S.K., Han, S.J., Choi, H.S., Im, S.Y., and Lee, J.W. 1998, J Biol Chem, 273, 10831. 49. Werbajh, S., Nojek, I., Lanz, R., and Costas, M.A. 2000, FEBS Lett, 485, 195. 50. Wu, R.C., Qin, J., Hashimoto, Y., Wong, J., Xu, J., Tsai, S.Y., Tsai, M.J., and O’Malley, B.W. 2002, Mol Cell Biol, 22, 3549. 51. Giraud, S., Bienvenu, F., Avril, S., Gascan, H., Heery, D.M., and Coqueret, O. 2002, J Biol Chem, 277, 8004. 52. Litterst, C.M., and Pfitzner, E. 2002, J Biol Chem, 277, 36052. 53. Razeto, A., Ramakrishnan, V., Litterst, C.M., Giller, K., Griesinger, C., Carlomagno, T., Lakomek, N., Heimburg, T., Lodrini, M., Pfitzner, E., and Becker, S. 2004, J Mol Biol, 336, 319. 54. Iordanidou, P., Aggelidou, E., Demetriades, C., and Hadzopoulou-Cladaras, M. 2005, J Biol Chem, 280, 21810. 55. Soutoglou, E., Papafotiou, G., Katrakili, N., and Talianidis, I. 2000, J Biol Chem, 275, 12515. 56. Myers, E., Hill, A.D., Kelly, G., McDermott, E.W., O’Higgins, N.J., Buggy, Y., and Young, L.S. 2005, Clin Cancer Res, 11, 2111. 57. Lee, S.K., Kim, H.J., Kim, J.W., and Lee, J.W. 1999, Mol Endocrinol, 13, 1924. 58. Louie, M.C., Zou, J.X., Rabinovich, A., and Chen, H.W. 2004, Mol Cell Biol, 24, 5157. 59. Li, S., and Shang, Y. 2007, Cell Signal, 19, 1101. 60. Yan, F., Gao, X., Lonard, D.M., and Nawaz, Z. 2003, Mol Endocrinol, 17, 1315. 61. Li, X., Lonard, D.M., Jung, S.Y., Malovannaya, A., Feng, Q., Qin, J., Tsai, S.Y., Tsai, M.J., and O’Malley, B.W. 2006, Cell, 124, 381. 62. Wu, R.C., Feng, Q., Lonard, D.M., and O’Malley, B.W. 2007, Cell, 129, 1125.

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

161

63. Gianni, M., Parrella, E., Raska, I., Jr., Gaillard, E., Nigro, E.A., Gaudon, C., Garattini, E., and Rochette-Egly, C. 2006, Embo J, 25, 739. 64. Giamas, G., Castellano, L., Feng, Q., Knippschild, U., Jacob, J., Thomas, R.S., Coombes, R.C., Smith, C.L., Jiao, L.R., and Stebbing, J. 2009, Nucleic Acids Res, 65. Yi, P., Feng, Q., Amazit, L., Lonard, D.M., Tsai, S.Y., Tsai, M.J., and O’Malley, B.W. 2008, Mol Cell, 29, 465. 66. Ferrero, M., Avivar, A., Garcia-Macias, M.C., and de Mora, J.F. 2008, Cancer Res, 68, 5450. 67. Naeem, H., Cheng, D., Zhao, Q., Underhill, C., Tini, M., Bedford, M.T., and Torchia, J. 2007, Mol Cell Biol, 27, 120. 68. Xu, J., Liao, L., Ning, G., Yoshida-Komiya, H., Deng, C., and O’Malley, B.W. 2000, Proc Natl Acad Sci U S A, 97, 6379. 69. Wang, Z., Rose, D.W., Hermanson, O., Liu, F., Herman, T., Wu, W., Szeto, D., Gleiberman, A., Krones, A., Pratt, K., Rosenfeld, R., Glass, C.K., and Rosenfeld, M.G. 2000, Proc Natl Acad Sci U S A, 97, 13549. 70. Xu, J., Qiu, Y., DeMayo, F.J., Tsai, S.Y., Tsai, M.J., and O’Malley, B.W. 1998, Science, 279, 1922. 71. Zhou, G., Hashimoto, Y., Kwak, I., Tsai, S.Y., and Tsai, M.J. 2003, Mol Cell Biol, 23, 7742. 72. Torres-Arzayus, M.I., Font de Mora, J., Yuan, J., Vazquez, F., Bronson, R., Rue, M., Sellers, W.R., and Brown, M. 2004, Cancer Cell, 6, 263. 73. Liao, L., Chen, X., Wang, S., Parlow, A.F., and Xu, J. 2008, Mol Cell Biol, 28, 2460. 74. Gehin, M., Mark, M., Dennefeld, C., Dierich, A., Gronemeyer, H., and Chambon, P. 2002, Mol Cell Biol, 22, 5923. 75. Conneely, O.M., Mulac-Jericevic, B., Lydon, J.P., and De Mayo, F.J. 2001, Mol Cell Endocrinol, 179, 97. 76. Han, S.J., Tsai, S.Y., Tsai, M.J., and O’Malley, B.W. 2007, Endocrinology, 148, 2471. 77. Han, S.J., DeMayo, F.J., and O’Malley, B.W. 2007, Ernst Schering Found Symp Proc, 25. 78. Mukherjee, A., Amato, P., Allred, D.C., DeMayo, F.J., and Lydon, J.P. 2007, Nucl Recept Signal, 5, e011. 79. Nishihara, E., Yoshida-Komiya, H., Chan, C.S., Liao, L., Davis, R.L., O’Malley, B.W., and Xu, J. 2003, J Neurosci, 23, 213. 80. Hayashi, Y., Ohmori, S., Ito, T., and Seo, H. 1997, Biochem Biophys Res Commun, 236, 83. 81. Grenier, J., Trousson, A., Chauchereau, A., Amazit, L., Lamirand, A., Leclerc, P., GuiochonMantel, A., Schumacher, M., and Massaad, C. 2004, Mol Endocrinol, 18, 2866. 82. Grenier, J., Trousson, A., Chauchereau, A., Cartaud, J., Schumacher, M., and Massaad, C. 2006, Mol Endocrinol, 20, 254. 83. An, B.S., Selva, D.M., Hammond, G.L., Rivero-Muller, A., Rahman, N., and Leung, P.C. 2006, J Biol Chem, 281, 20817. 84. Blanco, E.J., Hernandez, C.M., Rubio, M., Riesco, J.M., Carretero, M., Perez, E., Herrero, J.J., de Mora, J.F., Burks, D.J., and Carretero, J. 2007, Journal of Anatomy, 210, 632. 85. Coste, A., Louet, J.F., Lagouge, M., Lerin, C., Antal, M.C., Meziane, H., Schoonjans, K., Puigserver, P., O’Malley, B.W., and Auwerx, J. 2008, Proc Natl Acad Sci U S A, 105, 17187. 86. Louet, J.F., Coste, A., Amazit, L., Tannour-Louet, M., Wu, R.C., Tsai, S.Y., Tsai, M.J., Auwerx, J., and O’Malley, B.W. 2006, Proc Natl Acad Sci U S A, 103, 17868. 87. Picard, F., Gehin, M., Annicotte, J., Rocchi, S., Champy, M.F., O’Malley, B.W., Chambon, P., and Auwerx, J. 2002, Cell, 111, 931. 88. Jeong, J.W., Kwak, I., Lee, K.Y., White, L.D., Wang, X.P., Brunicardi, F.C., O’Malley, B.W., and DeMayo, F.J. 2006, Mol Endocrinol, 20, 1138. 89. Ying, H., Furuya, F., Willingham, M.C., Xu, J., O’Malley, B.W., and Cheng, S.Y. 2005, Mol Cell Biol, 25, 7687. 90. Yuan, Y., Liao, L., Tulis, D.A., and Xu, J. 2002, Circulation, 105, 2653. 91. Yu, C., York, B., Wang, S., Feng, Q., Xu, J., and O’Malley, B.W. 2007, Mol Cell, 25, 765.

162

M. Ferrero and J. Font de Mora

92. Coste, A., Antal, M.C., Chan, S., Kastner, P., Mark, M., O’Malley, B.W., and Auwerx, J. 2006, Embo J, 25, 2453. 93. Luo, J.H., Xie, D., Liu, M.Z., Chen, W., Liu, Y.D., Wu, G.Q., Kung, H.F., Zeng, Y.X., and Guan, X.Y. 2008, Int J Cancer, 122, 2554. 94. Liu, M.Z., Xie, D., Mai, S.J., Tong, Z.T., Shao, J.Y., Fu, Y.S., Xia, W.J., Kung, H.F., Guan, X.Y., and Zeng, Y.X. 2008, Am J Clin Pathol, 129, 728. 95. Xu, F.P., Liu, Y.H., Zhuang, H.G., Li, L., Luo, X.L., and Wen, J.M. 2008, Zhonghua Bing Li Xue Za Zhi, 37, 250. 96. Xu, F.P., Xie, D., Wen, J.M., Wu, H.X., Liu, Y.D., Bi, J., Lv, Z.L., Zeng, Y.X., and Guan, X.Y. 2007, Cancer Lett, 245, 69. 97. Grivas, P.D., Tzelepi, V., Sotiropoulou-Bonikou, G., Kefalopoulou, Z., Papavassiliou, A.G., and Kalofonos, H. 2009, Int J Colorectal Dis. 98. Lassmann, S., Weis, R., Makowiec, F., Roth, J., Danciu, M., Hopt, U., and Werner, M. 2007, J Mol Med, 85, 293. 99. Xie, D., Sham, J.S., Zeng, W.F., Lin, H.L., Bi, J., Che, L.H., Hu, L., Zeng, Y.X., and Guan, X.Y. 2005, Hum Pathol, 36, 777. 100. Zeng, S.D., Xie, D., Lin, F., Wang, C.X., Tao, Y., and Zhang, M. 2005, Zhonghua Wei Chang Wai Ke Za Zhi, 8, 67. 101. Wang, Y., Wu, M.C., Sham, J.S., Zhang, W., Wu, W.Q., and Guan, X.Y. 2002, Cancer, 95, 2346. 102. Henke, R.T., Haddad, B.R., Kim, S.E., Rone, J.D., Mani, A., Jessup, J.M., Wellstein, A., Maitra, A., and Riegel, A.T. 2004, Clin Cancer Res, 10, 6134. 103. Ghadimi, B.M., Schrock, E., Walker, R.L., Wangsa, D., Jauho, A., Meltzer, P.S., and Ried, T. 1999, Am J Pathol, 154, 525. 104. Sakakura, C., Hagiwara, A., Yasuoka, R., Fujita, Y., Nakanishi, M., Masuda, K., Kimura, A., Nakamura, Y., Inazawa, J., Abe, T., and Yamagishi, H. 2000, Int J Cancer, 89, 217. 105. Rangel, J., Torabian, S., Shaikh, L., Nosrati, M., Baehner, F.L., Haqq, C., Leong, S.P., Miller, J.R., 3rd, Sagebiel, R.W., and Kashani-Sabet, M. 2006, J Clin Oncol, 24, 4565. 106. Bautista, S., Valles, H., Walker, R.L., Anzick, S., Zeillinger, R., Meltzer, P., and Theillet, C. 1998, Clin Cancer Res, 4, 2925. 107. Reiter, R., Wellstein, A., and Riegel, A.T. 2001, J Biol Chem, 276, 39736. 108. Glaeser, M., Floetotto, T., Hanstein, B., Beckmann, M.W., and Niederacher, D. 2001, Horm Metab Res, 33, 121. 109. Hudelist, G., Czerwenka, K., Kubista, E., Marton, E., Pischinger, K., and Singer, C.F. 2003, Breast Cancer Res Treat, 78, 193. 110. List, H.J., Reiter, R., Singh, B., Wellstein, A., and Riegel, A.T. 2001, Breast Cancer Res Treat, 68, 21. 111. Nessler-Menardi, C., Jotova, I., Culig, Z., Eder, I.E., Putz, T., Bartsch, G., and Klocker, H. 2000, Prostate, 45, 124. 112. Tanner, M.M., Grenman, S., Koul, A., Johannsson, O., Meltzer, P., Pejovic, T., Borg, A., and Isola, J.J. 2000, Clin Cancer Res, 6, 1833. 113. Harigopal, M., Heymann, J., Ghosh, S., Anagnostou, V., Camp, R.L., and Rimm, D.L. 2008, Breast Cancer Res Treat, 114. Zhao, C., Yasui, K., Lee, C.J., Kurioka, H., Hosokawa, Y., Oka, T., and Inazawa, J. 2003, Cancer, 98, 18. 115. Han, X.Y., Zhang, X.M., Chen, Y., Yang, K.X., and Qie, M.R. 2007, Sichuan Da Xue Xue Bao Yi Xue Ban, 38, 246. 116. Hirai, Y., Utsugi, K., Takeshima, N., Kawamata, Y., Furuta, R., Kitagawa, T., Kawaguchi, T., Hasumi, K., and Noda, T. 2004, Am J Obstet Gynecol, 191, 1173. 117. Sakaguchi, H., Fujimoto, J., Sun, W.S., and Tamaya, T. 2007, J Steroid Biochem Mol Biol, 104, 237. 118. Balmer, N.N., Richer, J.K., Spoelstra, N.S., Torkko, K.C., Lyle, P.L., and Singh, M. 2006, Mod Pathol, 19, 1593.

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

163

119. Han, X.Y., Chen, Y., Hou, M.M., Zhang, J., Yang, K.X., Chen, Y.Y., and Qie, M.R. 2008, Sichuan Da Xue Xue Bao Yi Xue Ban, 39, 619. 120. Tilli, M.T., Reiter, R., Oh, A.S., Henke, R.T., McDonnell, K., Gallicano, G.I., Furth, P.A., and Riegel, A.T. 2005, Mol Endocrinol, 19, 644. 121. Avivar, A., Garcia-Macias, M.C., Ascaso, E., Herrera, G., O’Connor, J.E., and de Mora, J.F. 2006, FEBS Lett, 580, 5222. 122. Kuang, S.Q., Liao, L., Zhang, H., Lee, A.V., O’Malley, B.W., and Xu, J. 2004, Cancer Res, 64, 1875. 123. Fereshteh, M.P., Tilli, M.T., Kim, S.E., Xu, J., O’Malley, B.W., Wellstein, A., Furth, P.A., and Riegel, A.T. 2008, Cancer Res, 68, 3697. 124. Kuang, S.Q., Liao, L., Wang, S., Medina, D., O’Malley, B.W., and Xu, J. 2005, Cancer Res, 65, 7993. 125. Chung, A.C., Zhou, S., Liao, L., Tien, J.C., Greenberg, N.M., and Xu, J. 2007, Cancer Res, 67, 5965. 126. Tien, J.C., Zhou, S., and Xu, J. 2009, Int J Biol Sci, 5, 256. 127. Ying, H., Willingham, M.C., and Cheng, S.Y. 2008, Oncogene, 27, 823. 128. Louie, M.C., Revenko, A.S., Zou, J.X., Yao, J., and Chen, H.W. 2006, Mol Cell Biol, 26, 3810. 129. Lathrop, A.E., and Loeb, L. 1916, J Cancer Res, 1, 1. 130. Kurebayashi, J., Otsuki, T., Kunisue, H., Tanaka, K., Yamamoto, S., and Sonoo, H. 2000, Clin Cancer Res, 6, 512. 131. Rey, J.M., Pujol, P., Callier, P., Cavailles, V., Freiss, G., Maudelonde, T., and Brouillet, J.P. 2000, J Mol Endocrinol, 24, 433. 132. Azorsa, D.O., Cunliffe, H.E., and Meltzer, P.S. 2001, Breast Cancer Res Treat, 70, 89. 133. Shang, Y., Hu, X., DiRenzo, J., Lazar, M.A., and Brown, M. 2000, Cell, 103, 843. 134. Shao, W., Keeton, E.K., McDonnell, D.P., and Brown, M. 2004, Proc Natl Acad Sci U S A, 101, 11599. 135. Murphy, L.C., Simon, S.L., Parkes, A., Leygue, E., Dotzlaw, H., Snell, L., Troup, S., Adeyinka, A., and Watson, P.H. 2000, Cancer Res, 60, 6266. 136. Reiter, R., Oh, A.S., Wellstein, A., and Riegel, A.T. 2004, Oncogene, 23, 403. 137. List, H.J., Lauritsen, K.J., Reiter, R., Powers, C., Wellstein, A., and Riegel, A.T. 2001, J Biol Chem, 276, 23763. 138. Tikkanen, M.K., Carter, D.J., Harris, A.M., Le, H.M., Azorsa, D.O., Meltzer, P.S., and Murdoch, F.E. 2000, Proc Natl Acad Sci U S A, 97, 12536. 139. Doisneau-Sixou, S.F., Sergio, C.M., Carroll, J.S., Hui, R., Musgrove, E.A., and Sutherland, R.L. 2003, Endocr Relat Cancer, 10, 179. 140. Planas-Silva, M.D., Shang, Y., Donaher, J.L., Brown, M., and Weinberg, R.A. 2001, Cancer Res, 61, 3858. 141. Bouras, T., Southey, M.C., and Venter, D.J. 2001, Cancer Res, 61, 903. 142. Zhou, H.J., Yan, J., Luo, W., Ayala, G., Lin, S.H., Erdem, H., Ittmann, M., Tsai, S.Y., and Tsai, M.J. 2005, Cancer Res, 65, 7976. 143. Franke, T.F. 2008, Oncogene, 27, 6473. 144. Oh, A., List, H.J., Reiter, R., Mani, A., Zhang, Y., Gehan, E., Wellstein, A., and Riegel, A.T. 2004, Cancer Res, 64, 8299. 145. Holbro, T., Civenni, G., and Hynes, N.E. 2003, Exp Cell Res, 284, 99. 146. Thorat, M.A., Turbin, D., Morimiya, A., Leung, S., Zhang, Q., Jeng, M.H., Huntsman, D.G., Nakshatri, H., and Badve, S. 2008, Histopathology, 53, 634. 147. Amazit, L., Pasini, L., Szafran, A.T., Berno, V., Wu, R.C., Mielke, M., Jones, E.D., Mancini, M.G., Hinojos, C.A., O’Malley, B.W., and Mancini, M.A. 2007, Mol Cell Biol, 27, 6913. 148. Oh, A.S., Lahusen, J.T., Chien, C.D., Fereshteh, M.P., Zhang, X., Dakshanamurthy, S., Xu, J., Kagan, B.L., Wellstein, A., and Riegel, A.T. 2008, Mol Cell Biol, 28, 6580. 149. Font de Mora, J., and Brown, M. 2000, Mol Cell Biol, 20, 5041. 150. Lahusen, T., Fereshteh, M., Oh, A., Wellstein, A., and Riegel, A.T. 2007, Cancer Res, 67, 7256. 151. Nahta, R., and Esteva, F.J. 2006, Breast Cancer Res, 8, 215.

164

M. Ferrero and J. Font de Mora

152. Osborne, C.K., Bardou, V., Hopp, T.A., Chamness, G.C., Hilsenbeck, S.G., Fuqua, S.A., Wong, J., Allred, D.C., Clark, G.M., and Schiff, R. 2003, J Natl Cancer Inst, 95, 353. 153. Perkins, N.D. 2007, Nat Rev Mol Cell Biol, 8, 49. 154. Marquez, D.C., Chen, H.W., Curran, E.M., Welshons, W.V., and Pietras, R.J. 2006, Mol Cell Endocrinol, 246, 91. 155. Wu, R.C., Qin, J., Yi, P., Wong, J., Tsai, S.Y., Tsai, M.J., and O’Malley, B.W. 2004, Mol Cell, 15, 937. 156. Zheng, F.F., Wu, R.C., Smith, C.L., and O’Malley, B.W. 2005, Mol Cell Biol, 25, 8273. 157. Park, K.J., Krishnan, V., O’Malley, B.W., Yamamoto, Y., and Gaynor, R.B. 2005, Mol Cell, 18, 71. 158. Cao, Y., Bonizzi, G., Seagroves, T.N., Greten, F.R., Johnson, R., Schmidt, E.V., and Karin, M. 2001, Cell, 107, 763. 159. Ozes, O.N., Mayo, L.D., Gustin, J.A., Pfeffer, S.R., Pfeffer, L.M., and Donner, D.B. 1999, Nature, 401, 82. 160. Romashkova, J.A., and Makarov, S.S. 1999, Nature, 401, 86. 161. Fang, Z.H., and Han, Z.C. 2006, Histol Histopathol, 21, 403. 162. Mussi, P., Yu, C., O’Malley, B.W., and Xu, J. 2006, Mol Endocrinol, 20, 3105. 163. Bai, J., Uehara, Y., and Montell, D.J. 2000, Cell, 103, 1047. 164. DePasquale, J.A. 1999, Histochem Cell Biol, 112, 341. 165. Yang, J., Mani, S.A., Donaher, J.L., Ramaswamy, S., Itzykson, R.A., Come, C., Savagner, P., Gitelman, I., Richardson, A., and Weinberg, R.A. 2004, Cell, 117, 927. 166. Kajiro, M., Hirota, R., Nakajima, Y., Kawanowa, K., So-Ma, K., Ito, I., Yamaguchi, Y., Ohie, S.H., Kobayashi, Y., Seino, Y., Kawano, M., Kawabe, Y.I., Takei, H., Hayashi, S.I., Kurosumi, M., Murayama, A., Kimura, K., and Yanagisawa, J. 2009, Nat Cell Biol, 167. Wang, S., Yuan, Y., Liao, L., Kuang, S.Q., Tien, J.C., O’Malley, B.W., and Xu, J. 2009, Proc Natl Acad Sci U S A, 106, 151. 168. Qin, L., Liu, Z., Chen, H., and Xu, J. 2009, Cancer Res, 69, 3819. 169. Li, L.B., Louie, M.C., Chen, H.W., and Zou, J.X. 2008, Cancer Lett, 261, 64. 170. Yan, J., Erdem, H., Li, R., Cai, Y., Ayala, G., Ittmann, M., Yu-Lee, L.Y., Tsai, S.Y., and Tsai, M.J. 2008, Cancer Res, 68, 5460. 171. Qin, L., Liao, L., Redmond, A., Young, L., Yuan, Y., Chen, H., O’Malley, B.W., and Xu, J. 2008, Mol Cell Biol, 28, 5937. 172. Hurst, H.C. 2001, Breast Cancer Res, 3, 395. 173. Sun, A., Bagella, L., Tutton, S., Romano, G., and Giordano, A. 2007, J Cell Biochem, 102, 1400. 174. Gillett, C., Fantl, V., Smith, R., Fisher, C., Bartek, J., Dickson, C., Barnes, D., and Peters, G. 1994, Cancer Res, 54, 1812. 175. Zhang, S.Y., Liu, S.C., Al-Saleem, L.F., Holloran, D., Babb, J., Guo, X., and Klein-Szanto, A.J. 2000, Cancer Epidemiol Biomarkers Prev, 9, 395. 176. Planas-Silva, M.D., Donaher, J.L., and Weinberg, R.A. 1999, Cancer Res, 59, 4788. 177. Klein, E.A., and Assoian, R.K. 2008, J Cell Sci, 121, 3853. 178. Nakuci, E., Mahner, S., Direnzo, J., and ElShamy, W.M. 2006, Exp Cell Res, 312, 3120. 179. Arimura, A., vn Peer, M., Schroder, A.J., and Rothman, P.B. 2004, J Biol Chem, 279, 31105. 180. Zou, J.X., Zhong, Z., Shi, X.B., Tepper, C.G., deVere White, R.W., Kung, H.J., and Chen, H. 2006, Prostate, 66, 1474. 181. Samani, A.A., Yakar, S., LeRoith, D., and Brodt, P. 2007, Endocr Rev, 28, 20. 182. Torres-Arzayus, M.I., Yuan, J., DellaGatta, J.L., Lane, H., Kung, A.L., and Brown, M. 2006, Cancer Res, 66, 11381. 183. Ayala, G., Yan, J., Li, R., Ding, Y., Thompson, T.C., Mims, M.P., Hayes, T.G., MacDonnell, V., Lynch, R.G., Frolov, A., Miles, B.J., Wheeler, T.M., Harper, J.W., Tsai, M.J., Ittmann, M.M., and Kadmon, D. 2008, Clin Cancer Res, 14, 7511. 184. Clarke, R., Leonessa, F., Welch, J.N., and Skaar, T.C. 2001, Pharmacol Rev, 53, 25. 185. Osborne, C.K., and Schiff, R. 2003, Breast, 12, 362.

7

AIB1 is a Potential Therapeutical Target in Cancer Progression…

165

186. Chen, Y.J., Lin, S.C., Kao, T., Chang, C.S., Hong, P.S., Shieh, T.M., and Chang, K.W. 2004, J Pathol, 204, 326. 187. Schiff, R., Massarweh, S., Shou, J., and Osborne, C.K. 2003, Clin Cancer Res, 9, 447S. 188. Kirkegaard, T., McGlynn, L.M., Campbell, F.M., Muller, S., Tovey, S.M., Dunne, B., Nielsen, K.V., Cooke, T.G., and Bartlett, J.M. 2007, Clin Cancer Res, 13, 1405. 189. Hurtado, A., Holmes, K.A., Geistlinger, T.R., Hutcheson, I.R., Nicholson, R.I., Brown, M., Jiang, J., Howat, W.J., Ali, S., and Carroll, J.S. 2008, Nature, 456, 663. 190. Glaros, S., Atanaskova, N., Zhao, C., Skafar, D.F., and Reddy, K.B. 2006, Mol Endocrinol, 20, 996. 191. Shin, I., Miller, T., and Arteaga, C.L. 2006, Clin Cancer Res, 12, 1008s. 192. Iwase, H. 2006, Nippon Rinsho, 64, 555. 193. Girault, I., Lerebours, F., Amarir, S., Tozlu, S., Tubiana-Hulin, M., Lidereau, R., and Bieche, I. 2003, Clin Cancer Res, 9, 1259. 194. Hammerich-Hille, S., Bardout, V.J., Hilsenbeck, S.G., Osborne, C.K., and Oesterreich, S. 2009, Breast Cancer Res Treat, 195. Hayashi, Y., Yamamoto, M., Ohmori, S., Kikumori, T., Imai, T., Funahashi, H., and Seo, H. 1999, Endocr J, 46, 279. 196. Rebbeck, T.R., Wang, Y., Kantoff, P.W., Krithivas, K., Neuhausen, S.L., Godwin, A.K., Daly, M.B., Narod, S.A., Brunet, J.S., Vesprini, D., Garber, J.E., Lynch, H.T., Weber, B.L., and Brown, M. 2001, Cancer Res, 61, 5420. 197. Kadouri, L., Kote-Jarai, Z., Easton, D.F., Hubert, A., Hamoudi, R., Glaser, B., Abeliovich, D., Peretz, T., and Eeles, R.A. 2004, Int J Cancer, 108, 399. 198. Montgomery, K.G., Chang, J.H., Gertig, D.M., Dite, G.S., McCredie, M.R., Giles, G.G., Southey, M.C., Hopper, J.L., and Campbell, I.G. 2005, Breast Cancer Res, 7, R353. 199. Hughes, D.J., Ginolhac, S.M., Coupier, I., Barjhoux, L., Gaborieau, V., Bressac-de-Paillerets, B., Chompret, A., Bignon, Y.J., Uhrhammer, N., Lasset, C., Giraud, S., Sobol, H., Hardouin, A., Berthet, P., Peyrat, J.P., Fournier, J., Nogues, C., Lidereau, R., Muller, D., Fricker, J.P., Longy, M., Toulas, C., Guimbaud, R., Yannoukakos, D., Mazoyer, S., Lynch, H.T., Lenoir, G.M., Goldgar, D.E., Stoppa-Lyonnet, D., and Sinilnikova, O.M. 2005, Int J Cancer, 117, 230. 200. Yamashita, H., Nishio, M., Toyama, T., Sugiura, H., Kondo, N., Kobayashi, S., Fujii, Y., and Iwase, H. 2008, Endocr Relat Cancer, 15, 755. 201. Su, Q., Hu, S., Gao, H., Ma, R., Yang, Q., Pan, Z., Wang, T., and Li, F. 2008, Oncology, 75, 159. 202. Zhao, W., Zhang, Q., Kang, X., Jin, S., and Lou, C. 2009, Biochem Biophys Res Commun, 380, 699. 203. Hashimoto, Y., Naruyama, H., Ando, R., Okada, S., Tozawa, K., and Kohri, K. 2008, Hinyokika Kiyo, 54, 57. 204. Chen, B., Wang, Y., Kane, S.E., and Chen, S. 2008, J Mol Endocrinol, 41, 367. 205. Iwase, H., Omoto, Y., Toyama, T., Yamashita, H., Hara, Y., Sugiura, H., and Zhang, Z. 2003, Breast Cancer Res Treat, 80, 339. 206. Shiau, A.K., Barstad, D., Loria, P.M., Cheng, L., Kushner, P.J., Agard, D.A., and Greene, G.L. 1998, Cell, 95, 927. 207. Gunther, J.R., Du, Y., Rhoden, E., Lewis, I., Revennaugh, B., Moore, T.W., Kim, S.H., Dingledine, R., Fu, H., and Katzenellenbogen, J.A. 2009, J Biomol Screen, 14, 181. 208. Shao, D., Berrodin, T.J., Manas, E., Hauze, D., Powers, R., Bapat, A., Gonder, D., Winneker, R.C., and Frail, D.E. 2004, J Steroid Biochem Mol Biol, 88, 351. 209. Hossain, A., Kuo, M.T., and Saunders, G.F. 2006, Mol Cell Biol, 26, 8191. 210. Blenkiron, C., and Miska, E.A. 2007, Hum Mol Genet, 16 Spec No 1, R106. 211. Liang, J., Zhang, H., Zhang, Y., Zhang, Y., and Shang, Y. 2009, EMBO Rep, 10, 51. 212. Zou, J.X., Revenko, A.S., Li, L.B., Gemo, A.T., and Chen, H.W. 2007, Proc Natl Acad Sci U S A, 104, 18067. 213. Yi, P., Wu, R.C., Sandquist, J., Wong, J., Tsai, S.Y., Tsai, M.J., Means, A.R., and O’Malley, B.W. 2005, Mol Cell Biol, 25, 9687.

166

M. Ferrero and J. Font de Mora

214. Mani, A., Oh, A.S., Bowden, E.T., Lahusen, T., Lorick, K.L., Weissman, A.M., Schlegel, R., Wellstein, A., and Riegel, A.T. 2006, Cancer Res, 66, 8680. 215. Ferry, C., Gianni, M., Lalevee, S., Bruck, N., Plassat, J.L., Raska, I., Jr., Garattini, E., and Rochette-Egly, C. 2009, J Biol Chem, 216. Yeung, P.L., Zhang, A., and Chen, J.D. 2006, Biochem Biophys Res Commun, 348, 13. 217. Zhang, Y., Zhang, H., Liang, J., Yu, W., and Shang, Y. 2007, Embo J, 26, 2645. 218. Li, C., Liang, Y.Y., Feng, X.H., Tsai, S.Y., Tsai, M.J., and O’Malley, B.W. 2008, Mol Cell, 31, 835.

Chapter 8

Rational Design of DNA Anticancer Agent That Targets Signal Transducer and Activator of Transcription 3 (Stat3) for Cancer Therapy Naijie Jing

Introduction A decade ago, signal transducer and activator of transcription (STAT) proteins were discovered as latent cytoplasmic transcription factors [1]. Seven known mammalian STAT proteins – Stat1, 2, 3, 4, 5a, 5b, and 6 – are involved in cell proliferation, differentiation, and apoptosis [2–12]. These proteins contain several domains: a tetramerization domain, a coil-coil domain, a DNA-binding domain, a linker domain, an Src-homology 2 (SH2) domain, a critical tyrosine residing near the C-terminal end (position 705 in Stat3), and a C-terminal transactivation domain [13]. STAT proteins are activated in response to the binding of a number of ligands to their cognate cell surface receptors, especially cytokines (e.g., IL-6) and growth factors (e.g., EGF). STAT proteins exist as monomers or N-terminal head-to-head dimers in the cytoplasm of non-stimulated cells [14]. Ligand binding to cytokine or growth factor surface receptors results in receptor dimerization and transphosphorylation by receptor-intrinsic or receptor-associated tyrosine kinases. The kinases phosphorylate the cytoplasmic tails of the receptor to provide a docking site for the recruitment of monomeric STATs. STAT proteins are recruited to specific phosphotyrosine residues within receptor complexes through their SH2 domains; they subsequently become phosphorylated on the tyrosine residue within their C-terminus, and dimerize through reciprocal interactions between the SH2 domain of one monomer and the phosphorylated tyrosine of the other. The activated dimers translocate to the nucleus, where they bind to DNA-response elements in the promoters of target genes and activate specific gene expression programs (Fig. 8.1) [15–17]. Previously reported X-ray crystal structures of Stat1 and Stat3 homodimers bound to DNA reveal many details involving dimerization and DNA binding [18, 19].

N. Jing (*) Department of Medicine, Baylor College of Medicine, Houston, TX 77030, USA e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_8, © Springer Science+Business Media, LLC 2012

167

168

N. Jing

Fig. 8.1 STAT signaling pathway

Stat3 was originally termed acute-phase response factor (APRF) because it was first identified as a DNA-binding activity within IL-6-stimulated hepatocytes, and was capable of selectively interacting with an enhancer element in the promoter of acute-phase response genes [12, 20, 21]. The protein derived from the livers of IL-6treated mice was first purified to homogeneity, and its cDNA isolated and sequenced [22]. Receptors linked to Stat3 activation include: receptors for G-CSF (granulocyte colony-stimulating factor) and the IL-6 cytokine family members, as well as other type I and type II cytokine receptors; receptor tyrosine kinases; and G protein-coupled receptors [23]. In various in vitro systems, Stat3 activation downstream of these receptors has been demonstrated to influence multiple cell fate decisions, including proliferation [24], differentiation [25–27] , and apoptosis [5, 28]. Targeted disruption of the mouse Stat3 gene leads to embryonic lethality at 6.5–7.5 days [11], indicating that Stat3 – which is essential to early embryonic development – is potentially involved in gastrulation or visceral endoderm function [2]. Moreover, the tissue-specific deletion of Stat3 has confirmed in vitro results by demonstrating that Stat3 plays a crucial role in a variety of biological functions, including cell growth, the suppression and induction of apoptosis, and cell motility [2]. Stat3 has been identified as an important target for cancer therapy, since it participates in oncogenesis through the upregulation of genes encoding apoptosis inhibitors (Bcl-xL, Bcl-2, Mcl-1, and survivin), cell-cycle regulators (cyclin D1 and c-myc), and inducers of angiogenesis (VEGF) [17]. Mounting evidence shows that Stat3 is constitutively activated in many human cancers [15–17], including: 82% of prostate cancers, 70% of breast cancers, more than 90% of head and neck cancers, and more than 50% of lung cancers [29–32]. Recent studies have demonstrated that Stat3 is constitutively activated both in tumor cells and in immune cells in the tumor

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

169

microenvironment [33]. Constitutively activated Stat3 in tumor cells inhibits the expression of mediators necessary for immune activation against tumor cells and results in production of numerous mediators such as VEGF which inhibit dendritic cell function. In addition, activation of Stat3 in hematopoietic cells restrains natural tumor surveillance. Thus, targeting Stat3 is expected to decrease survival of tumor cells, inhibit tumor angiogenesis, and activate immune cells in the tumor stroma.

Developed Inhibitors of Stat3 Signaling The design of Stat3 inhibitor is based on the mechanism of Stat3 activation. When stimulated by cytokines [34], Janus kinases (Jak), growth factors (EGF, PDGF) or tyrosine kinases (Src) [35], Stat3 is activated by phosphorylating the tyrosine residue Y705 for Stat3 dimerization. Phosphorylated Stat3 dimer translocates into the nucleus to bind with enhancer sequences of target genes. The crystal structure of p-Stat3 dimer with bound DNA duplex has been determined [18], which makes it possible to discover Stat3 inhibitor by employing structure-based drug design technology. Several strategies can be used to design a Stat3 inhibitor: (a) suppressing the expression of phosphorylated Stat3, (b) inhibiting upstream kinase activation such as Jak and Src to reduce the phosphorylation of Stat3, (c) blocking Stat3 dimerization, and (d) inhibiting the DNA elements to bind within Stat3 dimer. Currently, many inhibitors of Stat3 signaling have been developed. However, no anti-tumor drug directly targeting Stat3 has yet reached the clinic.

Organic Compounds The following Stat3 inhibitors mainly target upstream regulators, such as Jak-Stat3 and Src/Stat3 signaling, to compete with associated receptor for Stat3 activation or to inhibit Stat3 dimerization: AG-490 is a potent inhibitor of protein tyrosine kinase that selectively inhibits Jak kinase, especially Jak2 kinase [36, 37]; WP-1034 is a novel Jak-Stat inhibitor with a similar structure with AG-490 [38]; and Cucurbitacin I (JSI-124), which was identified as a novel Jak-Stat3 inhibitor, is a natural product, isolated from plants with the Cucurbitaceae and Cruciferae families, and has been used as a folk remedy in China and India for centuries [39]. Recently, Cryptotanshinone was demonstrated to inhibit constitutive Stat3 by blocking its dimerization and also to suppress JAK2 as a secondary effect [40].

Peptide Inhibitors PY*LKTK (Y* means phosphotyrosine) was exactly the same as the segment of monomer Stat3 starting from residue P704 to residue K709 to mimic the interaction between two Stat3 monomers [41]; Y1068 (1063LPVPEY*INQSVP1074) inhibited

170

N. Jing

EGF-stimulated Stat3 activity and transforming growth factor (TGF)-a/EGFRmediated autocrine growth when delivered into cancer cells [42]; and 904Y*LPQTV909 was a Stat3 inhibitor by mapping gp130 residues starting from Y904 to V909 [43].

DNA Inhibitors Decoy oligonucleotide was composed of a 15-mer double-stranded oligonucleotide, and the sequences were close to the Stat3 response element within the c-fos promoter [44]; Antisense oligos, which target Stat3 or a dominant-negative Stat3 (DN-Stat3) plasmid, reduce Stat3 protein level in cytoplasm [45–47].

Protein Inhibitors PIAS are able to disrupt STAT DNA-binding activity by direct association with STAT proteins [48, 49]; SOCS (suppressors of cytokine signaling) family, which were also referred to as JAB (Jak-binding protein) or SSI (Stat-induced Stat inhibitors) [50–52], and phosphatases T cell PTP (TC-PTP), PTP-1B, SHP-1, and SHP-2 have been characterized as another class of inhibitors to Jak-STAT signaling [53]. Receptor-related phosphatases consist of three major regions: phosphotyrosinebinding SH2 motifs, a phosphatase domain, and regulatory tyrosine residues which are responsible for specific phosphorylation.

G-Quartet Oligodeoxynucleotides (GQ-ODNs) as a Potential Anticancer Drug G-quartet oligonucleotides (ODNs) were originally examined as modified substrate drug candidates based on their ability to inhibit telomerase activity, c-myc transcription, HIV infection, and DNA topoisomerase I activity in cells [54, 55]. We suggested that this new class of drugs could be engineered to potently and selectively inhibit another key oncogenic protein – Stat3 [54]. GQ-ODN is composed of G-rich DNA sequences, which have been identified, cloned, and characterized in the telomeres of many organisms (e.g., fungi, ciliates, vertebrates, and insects) [56]. G-quartet structures, which were initially observed within telomeres, can be formed by G-rich oligonucleotides [57]. In addition to telomeric sequences [58–60], G-quartet structures have been shown in vitro in: fragile X syndrome nucleotide repeats [61], HIV-1 RNA sequences [62, 63], and the immunoglobulin switch region [64]. G-quartets emerge from the association of four G-bases into a cyclic Hoogsteen H-bonding arrangement. Each G-base makes two H-bonds with its neighbor G-base (N1 to O6 and N2 to N7) (Fig. 8.2). G-quartets stack on top of one another, creating tetrad-helical structures,

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

171

Fig. 8.2 H-bond formation of G-quartet bases

and forming a family of structures that includes folded G-quartets produced by intramolecular interactions, as well as hairpin dimers and parallel-stranded tetramers resulting from intermolecular interactions. The formation of G-quartet structures depends on the DNA sequence, loop geometry, base composition of the nucleic acids, and the presence of metal ions [65–68]. It has been observed that K+ prefers to induce a folded G-quartet structure, while Na+ preferentially forms a linear four-strand G-quartet structure [69]. The preference for inducing a folded G-quartet structure by metal ions has been proposed as K+ > Rb+ > Na+ > Li+ or Cs+ [69, 70]. In general, G-quartet structures are very stable; however, their stability depends on the presence of monovalent cations and the concentration of G-rich oligonucleotides [71–74]. Because of its optimal size, the interaction of potassium within a G-octamer greatly increases structural stability. Furthermore, the high intracellular K+ concentration (140 mM) significantly promotes formation of folded G-quartet structures within the cytoplasm, greatly increasing their biological activity [75]. We, and others, have reported that G-rich ODNs – in our case, T30923 (also called T30695) and T30177 (also called AR177), which are both inhibitors of HIV-1 integrase – form an intramolecular G-quartet. These structures have been defined by nuclear magnetic resonance (NMR), circular dichroism, modeling, kinetics, and other biophysical methods [76–78]. The findings demonstrate that T30923 forms two kinds of G-quartet structures, those with unfolded and folded loop domains. Potassium ions induce a conformational transition from an unfolded G-quartet to a folded G-quartet in which K+ ions are bound into the structure [75]. Only the structure with folded loop domains demonstrates pharmacologic activity. The principal difficulty in developing GQ-ODN as a pharmacologic agent arises from the physical and structural properties of GQ-ODN. With a large molecular size, GQ-ODN cannot directly penetrate cell membranes. In order to address this issue, we have developed a novel delivery system for GQ-ODN that increases drug activity in cells and in vivo [79, 80]. This quartet structure enables interaction with the molecular target and resists endonuclease digestion.

172

N. Jing

Rational Design of G-Rich ODNs That Target Stat3 Signaling Structure-Based Drug Design Computer-assisted rational drug design has been wildly used to search or screen a novel drug in many medicinal areas. The structure-based drug design is based on two critical conditions: (a) The molecular structure of the target molecule has been well determined or established, which contains a potential binding site to inhibit its activation. (b) The complex of target-molecular/inhibitor obtained from computational docking is a reasonable starting point for predicting effects of designed inhibitors. Establishing a structure–activity relationship (SAR) is a critical step to direct the search of a lead compound for drug development. A panel of G-rich ODNs that are capable of forming G-quartet structures was designed as potential inhibitors of Stat3 candidates [79]. Each of the GQ-ODNs was randomly docked onto the known structure of the Stat3 dimer [18] 1,000 times using GRRAM docking program. H-bonds play a very important role in governing the interaction between proteins and DNA. The distribution of H-bonds that formed between Stat3 and each GQ-ODN demonstrated a potential binding site in Stat3 dimer for the GQ-ODN [79]. Analysis of the distribution can predict the ability of each GQ-ODN to inhibit Stat3 activation. Then electrophoretic mobility shift assay (EMSA) was employed to determine activity of the inhibition of Stat3 activation for each GQ-ODN. Combining the percentage of H-bonds of each GQ-ODN binding within the site, which is composed of residues 638–652 of Stat3 dimer, with its IC50 of the inhibition of Stat3 activation together, a linear structure-activity relationship (SAR) between Stat3 and GQ-ODNs was established (Fig. 8.3a and Table 8.1) [79]. The SAR showed that T40214 with the highest possibility (40%) to interact with Stat3 dimer corresponds to the strongest activity to inhibit Stat3 activation (IC50 = 5 mM). Thus, T40214 with sequence of (GGGC)4 was selected as a lead compound, which forms an intramolecular G-quartet structure with two G-quartets in the middle and a G-C-G-C loop on the top and the bottom determined by NMR (Fig. 8.3b) [77, 81].

GQ-ODN T40214 Selectively Inhibiting Stat3 Activation Selectively targeting Stat3 is not only a key factor but also a challenge issue in the design of a potent and specific anti-cancer agent. Besides Stat3, other STAT family members have important roles in intracellular signaling. Stat1 acts in a pro-apoptotic and anti-proliferative manner and appears to be a tumor suppressor whose action is contrary to Stat3 [17]. Also, the INF-g activated Stat1 results in transcription of pro-inflammatory cytokine and chemokine genes, which induce migration of effective T cells to the tumor site and then penetration of the tumors [82]. Thus, selective targeting of Stat3, rather than Stat1, is a critical issue in the development of an effective

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

173

Fig. 8.3 (a) Plot demonstrating a linear relationship of the percentage of GQ-ODN H-bonding localized in residues 638–652 of Stat3 dimer versus its IC50 against Stat3 (R2 = 0.9) [79]. (b) Structure of T40214 [80]

Table 8.1 IC50s and binding possibility of the designed G-rich ODNs [79] ÿ T40214 T40212 T40215 T40217 T40229 T40230 T40231 T40233

ÿ GGGCGGGCGGGCGGGC GGGCGGGTGGGCGGGT (GGGGT)4 GGGGTGGGTGGGTGGGTT TAGGGTGGGTGGGTGGGTAT GTGGGTGGGTGGGTGGGTTG GGTGGGTGGGTGGG GGGCGGGTGGGCGG

IC50 (mM) 5.0 35 49 64 41 43 10 34

% of H-bonds in the site of Stat3 40 30 28 27 29 28 37 29

174

N. Jing

Stat3 inhibitor. However, selectively targeting Stat3 rather than Stat1 is difficult since (a) Stat3 and Stat1 have similar structures [18, 19]. Protein sequences demonstrate 51%, 60%, and 66% identity in the Stat1 and Stat3 full length, SH2 domain, and DNA-binding domain, respectively. (b) Both are phosphorylated on tyrosine residues, and both form dimers through SH2 domain interactions. (c) Stat3 and Stat1 can also form a heterodimer in activation. To confirm whether the GQ-ODN selectively inhibits Stat3, T40214 was examined to inhibit the DNA-binding activity of IL-6-activated Stat3 and IFN-g-activated Stat1 via EMSA. T40214 strongly inhibited the DNA binding of Stat3 (IC50 = 5 mM), while 50% inhibition of Stat1 DNA binding was not achieved under the same conditions, using concentrations of T40214 up to 142 mM (Fig. 8.4a) [83]. Also, T40214 was demonstrated to not inhibit other Stats, e.g., Stat5 [84]. The mechanism responsible for T40214 selectively targeting Stat3 was determined by computational studies. First, GQ-ODN T40214 was randomly docked onto the dimer structures of both Stat3 and Stat1 (1,000 times), without setting any constraints, and the distributions of H-bonds formed between GQ-ODN and Stat3 dimers and between GQ-ODN and Stat1 dimers were analyzed. The histograms of H-bond distributions show clearly that the interaction between GQ-ODN T40214 and the Stat3 dimer was highly concentrated on the binding site composed of amino acids 638–652 (40%) – especially in the residues Q643, N646, and N647. However, specific binding interaction did not occur within the Stat1 dimer (Fig. 8.4b) [83]. Then, a high-resolution GRAMM docking program was employed to predict how GQ-ODN selectively targets the Stat3 dimer. The lowest energy complexes obtained from the dockings show that in the Stat3 dimer, residue Q643 of one monomer repels N646 of another due to the presence of similarly charged polar side chains. This interaction opens a channel (~9 Å) in the SH2 domains. In contrast, in Stat1, K637 of one monomer interacts with S640 of another, which pulls two domains together and blocks GQ-ODN interaction (Fig. 8.4c) [83, 85]. The docking results also demonstrate that residues in the loop domains of GQ-ODN form seven H-bonds with residues Q643 to N647 and tightly bind into the Stat3 site, thereby blocking Stat3 DNA binding and destabilizing dimer formation. In contrast, GQ-ODN cannot bind within the Stat1 dimer to block Stat1 DNA binding.

An Effective Drug Delivery System for GQ-ODNs An effective intracellular delivery system is essential for successful development of DNA anti-cancer agent. The principal difficulty of delivering GQ-ODNs into cells arises from the physical and structural properties of GQ-ODNs. GQ-ODN with a large molecular size cannot directly penetrate through cell membranes. Recently, we have developed an effective intracellular delivery system for GQ-ODNs (Fig. 8.5a) [75, 86]. The important development in this system is that based upon the property of potassium-dependent formation of G-quartet structure. In solution, the complex of GQ-ODN/K+ has a neutral surface charge and cannot effectively bind cationic vehicles. When heated, however, G-quartet oligos denature into random coils which permit the

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

175

Fig. 8.4 GQ-ODN T40214 specifically targets p-Stat3 dimer. (a) Data from EMSA assays. (b) Histograms of the number of H-bonds versus amino acids of Stat1 (top panel) and Stat3 (lower panel). (c) Docking complexes of T40214/Stat1 (left panel) and T40214/Stat3 (right panel) [83]

176

N. Jing

Fig. 8.5 (a) A novel delivery system built for GQ-ODN T40214. (b) Non-denatured DNA gel demonstrating drug delivery of T40214 into cells. T40214/PEI (Lane 2) appears as two bands: The higher band corresponds to ODNs adhering on cell membranes (a large-sized molecule with slower migration), while the lower band corresponds to the ODNs released inside cells. T40214 in the absence of PEI (Lane 3) appears as a single, higher band, indicating that T40214 cannot penetrate cells directly [86]

electrostatic interactions necessary for forming the GQ-ODN/vehicle complexes. The T40214/PEI complex was formed by mixing the denatured T40214 (possessing negative charges) with PEI (polyethylenimine), which is positively charged. The positive charges on the surface of the T40214/PEI complexes enhance their cellular uptake. Formation of the active G-quartet structure depends on the presence of cations, especially potassium. The intracellular potassium concentration of 140 mM (compared to the 4-mM extracellular concentration) is well above the 50-mM KCl concentration required for T40214 G-quartet formation. Thus, T40214 molecules maintain their unfolded structure before entering cells. Once delivered, T40214 forms a G-quartet structure in the high intracellular K+ concentration to bind within p-Stat3 dimers, and penetrates into the nucleus through nuclear pores to block Stat3 transcription. PEI facilitates delivery of T40214 to target cells for endocytosis, but PEI itself does not enter cells due to its positive surface charges [86].

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

177

To determine whether T40214 was delivered into the cells and whether T40214 forms a G-quartet structure inside cells, the experiments were performed as described previously [86]. In the non-denatured polyacrylamide gel (Fig. 8.5b), the free T40214 (Lane 1) corresponds to a G-quartet structure. T40214 with PEI (Lane 2) has two bands, indicating that a portion of the ODN adhered to cell membranes (the higher band), and a portion of ODNs entered into the cells (the lower band). Based on the same migration with free T40214, the GQ-ODN inside the cells formed the G-quartet structure. Analysis of the band intensities indicated that the ratio of delivery of T40214 is about 72%. T40214 without PEI (Lane 3) has only a single higher band in the gel, demonstrating that T40214 did not directly penetrate into cells.

The Drug Activity of GQ-ODN Was Greatly Increased by the Effective Delivery Drug Activity in Cancer Cells Western blots (Fig. 8.6a) showed that pre-incubation of cancer cells with the T40214/PEI complex (upper panel) completely inhibited IL-6-mediated activation of Stat3 while Stat1 activation was minimally affected, confirming the selectivity of T40214 for Stat3 within cells. Pre-incubation of cancer cells with T40214 only (no PEI) (lower panel) showed that neither Stat3 nor Stat1 activation was inhibited, confirming that PEI is required for delivery of GQ-ODN into cells. To determine if GQ-ODN interferes with Stat3 activation and translocation into the nucleus, we examined levels of p-Stat3 within the nuclei of cells stimulated with IL-6 following GQ-ODN pre-treatment. The nuclear p-Stat3 level was reduced 50–70% when the T40214 concentration was increased from 3.5 to 142 mM (Lanes 4–7) (Fig. 8.6b) [79]. Also, the result shows that T40214 only inhibits the activation of phosphorylated Stat3 (p-Stat3) but not that of unphosphorylated Stat3 (total Stat3 or T-Stat3).

Drug Delivery and Activity In Vivo 5’-Fluorescently labeled T40214 (10 mg/kg) plus PEI (2.5 mg/kg) was administered via intraperitoneal (IP) injection in xenograft tumors of nude mice. After injections, tumors were harvested at 24, 48, and 72 h and examined by fluorescence microscopy. At 48 and 72 h, the level of GQ-ODN in tumors was roughly 60% and 20%, respectively, of that at 24 h (Fig. 8.6c) [83]. GQ-ODN is effectively delivered to and has prolonged presence within tumor cells. Then, T40214 with and without PEI was injected via IP (intraperitoneal) route into nude mice with prostate tumors every other day for 3 weeks. Compared with tumor growth in control and PEItreated mice, tumor growth in T40214-treated mice was only minimally inhibited (p = 0.021 to untreated tumors & p = 0.23 to PEI-treated tumors); however, tumor

178

N. Jing

Fig. 8.6 (a) EMSA of extracts of cancer cells stimulated with IL-6 (25 ng/ml) after pre-incubation of cells with T40214 complexed with (top panel) or without (lower panel) PEI. Extracts were incubated with antibodies against Stat1 (Ab1; lane 1) or Stat3 (Ab3; lane 2) to confirm the composition of the homodimer bands [79]. (b) Immunoblot of nuclear extracts of cancer cells pre-incubated with media, PEI, or T40214 demonstrated that T40214 suppresses the level of phosphorylated Stat3. Equal amount of total Stat3 protein was loaded in each lane as control [79]. (c) Fluorescent micrographs show the distribution of labeled T40214 in HNSCC tumors of nude mice after 24, 48, and 72 h of drug administration. The control panel indicates the tumor without drug injection [83]. (d) PEI as vehicle significantly increases T40214 drug activity in vivo [86]

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

179

growth was significantly suppressed by the T40214/PEI complex (p < 0.0005 and p < 0.004) (Fig. 8.6d) [86]. Therefore, an effective delivery system significantly increases T40214 drug activity and is a key factor for T40214 in clinical testing.

T40214 Suppresses the Growth of Prostate Tumor Xenografts in Nude Mice Prostate Cancer Therapy Prostate cancer is the most frequently diagnosed cancer among men in the United States and is second only to lung cancer as a cause of cancer death [87]. Prostate cancer initially occurs as an androgen-dependent tumor. Patients with metastatic disease respond initially to androgen-ablation therapy; however, when the disease recurs, it is often insensitive to hormonal manipulation [88, 89]. Salvage therapy for hormone refractory metastatic disease includes chemotherapeutic agents which can have undesirable side effects and produce relatively short survival periods. In fact, median survival is only 18.9 months even with the most active chemotherapeutic agents [90]. Therefore, novel therapies, including agents with different therapeutic mechanisms of action and novel molecular targets, are urgently needed to treat prostate cancer patients. Stat3 is constitutively activated in many human cancers, including 82% of prostate cancers [30]. Therefore, Stat3 signaling was proposed to strongly influence progression of prostate cancer, and targeting Stat3 with T40214 could produce a novel and effective treatment for prostate cancer. Approximately 2.5–4.0 million cancer cells in 200 ml of PBS were injected subcutaneously into the right flank of athymic nude mice. After tumors were established (50–150 mm3), mice were randomly assigned to several groups: Group 1, untreated controls; Group 2 received treatment with PEI alone; Group 3 was treated with non-specific ODN/ PEI (a control ODN); Group 4 was treated with T40214 without PEI; Group 5 was treated with paclitaxel, a chemotherapeutic agent, as positive control; and Groups 6 and 7 were treated with T40214/PEI at doses of 5 and 10 mg/kg, respectively. Each group was composed of five mice. The nude mice received IP injection every other day for 24 days. Body weight and tumor size were measured every other day, and tumor size was calculated using the function (a × 0.5b2), where a equals the length, and b equals the width of tumors. The results (Fig. 8.7 and Table 8.2) [86] demonstrated that GQ-ODN T40214 caused significant suppression of tumor growth and increased survival time in a nude mouse xenograft model of prostate cancer.

Other Solid Tumor Therapies Innovative treatment approaches, including new agents with different therapeutic mechanisms of action and novel molecular targets, are urgently needed for many human cancers (e.g., breast cancer, non-small cell lung cancer [NSCLC], squamous

180

N. Jing

Fig. 8.7 (a) Prostate tumors treated by paclitaxel and T40214 in 16 and 24 days. (b) Plot of tumor volume (mm3) versus days of treatment. (c) The effects of the treatments on animal survival show that 50% of the mice remained alive in the untreated, placebo-treated, and ns-ODN-treated mice in 7, 12, and 10 days, respectively. Paclitaxel treatment extended the 50% survival point to 18 days. The 50% survival point for both T40214/PEI-treated mice was ³24 days with 100% of the mice in the 10-mg/kg group surviving to 24 days [86]

Table 8.2 In vivo effects of T40214/PEI against PC-3 xenograft tumors in nude mice [86] No. of mice Weight of mice (g) Tumor (mm3) Group Drug dose Start End Start End Start Control Untreated 5 5 27.2 ± 1.5 25.2 ± 1.7 93.1 ± 25 PEI alone 3.75 mg/kg 5 5 24.9 ± 1.7 23.7 ± 1.2 58.3 ± 7.9 ns-ODN/PEI 5 + 1.25 mg/kg 5 5 27.2 ± 1.5 22.3 ± 1.6 67.9 ± 12 T40214 10 mg/kg 5 5 25.1 ± 0.5 22.2 ± 1.9 55.7 ± 4.9 Paclitaxel 10 mg/kg 5 5 18.6 ± 1.0 20.6 ± 1.0 70.7 ± 7.3 T40214/PEI 5 + 1.25 mg/kg 5 5 25.5 ± 0.7 26.8 ± 1.5 72.9 ± 8.3 T40214/PEI 10 + 2.5 mg/kg 5 5 25.3 ± 1.1 26.5 ± 1.6 91.0 ± 16 End 2,633 ± 657 1,924 ± 374 2,362 ± 565 1,327 ± 333 672 ± 300 329 ± 147 55 ± 38

Fold of tumor growth 41.20 ± 9.30 40.85 ± 7.95 40.79 ± 9.75 23.82 ± 5.97 9.50 ± 5.67 5.15 ± 2.31 0.61 ± 0.41

Mean tumor weight (g) 1.37 ± 0.37 1.32 ± 0.58 1.61 ± 0.38 1.50 ± 0.40 0.47 ± 0.22 0.38 ± 0.14 0.18 ± 0.10

P value – – – – < 0.005 < 0.001 < 0.0005

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor 181

182

N. Jing

cell carcinoma of head and neck [SCCHN], and other cancers). In recent years, mounting evidence has indicated that Stat3 is an important target for cancer therapy, since Stat3 participates in oncogenesis through the upregulation of genes encoding apoptosis inhibitors, cell-cycle regulators, and inducers of angiogenesis. Stat3 is constitutively activated in many human cancers – including SCCHN, NSCLC, breast, and other cancer cells. T40214 has also been demonstrated to significantly suppress the growth in nude mice of other tumors, including breast cancer [79], head and neck squamous cell carcinoma (HNSCC) [83], and non-small cell lung cancer (NSCLC) [84] (Fig. 8.8). The in vivo results provided solid evidence that T40214 is a promising candidate of anti-cancer agent that targets Stat3 for treatment of human cancers.

The Mechanism of GQ-ODN T40214 as a Potent Anticancer Agent The Mechanism of T40214 Suppression of Tumor Growth The tumor tissues which were harvested from mice at the end of drug treatment periods were examined by western blot and TUNEL assays [86]. The results demonstrate that Stat3 proteins were constitutively phosphorylated in androgen-independent prostate tumors (PC-3) and that activation of p-Stat3 in the tumors was totally inhibited by the T40214/PEI complex (Fig. 8.9a). In addition, delivery of T40214 completely inhibited the expression of p-Stat3 downstream proteins, including Bcl-2, VEGF, and cyclin D1, in the prostate tumors. Finally, TUNEL assays demonstrated significant apoptosis in the T40214/PEI-treated tumor. The mean percentage of apoptotic cells in placebo-treated tumors was 10% ± 5% and that in T40214-treated mice was 80% ± 5% [86]. Moreover, it has been demonstrated that GQ-ODN T40214 significantly inhibited Stat3 activation, but did not inhibit the activation of Stat1 and Stat5 in tumors, showing T40214 selectively targets phosphosrylated Stat3 in vivo as well [84].

T40214 as an Anticancer Agent Targeting Stat3 Signaling for Cancer Therapy Putting all together, we established a system that targets the Stat3 signaling pathway, via GQ-ODN T40214, for cancer therapy (Fig. 8.9b). Monomers of Stat3 in the cytoplasm of cancer cells become activated through the stimulation by IL-6 or EGF. Stat3 is also recruited by intrinsic or receptor-associated tyrosine kinases, such as JAK or Src. Tyrosine phosphorylation of Stat3 induces the formation of active Stat3 dimers through the SH2 domains, and the activated Stat3 dimers translocate to the nucleus, where they bind to DNA-response elements in the promoters of target genes and activate specific gene expression programs [16–18]. GQ-ODN T40214 is

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

183

Fig. 8.8 The photos of in vivo drug tests of T40214/PEI in NSCLC tumors (I) and in SCCHN tumors (II) [83, 84]

delivered into the cytoplasm by PEI. In cytoplasmic K+ concentrations, T40214 forms G-quartet structures, binds to the SH2 domains of p-Stat3 dimers, and blocks DNA-binding activity. T40214 binding induces p-Stat3 dephosphorylation and may retard Stat3 rephosphorylation in cancer cells by inhibiting Stat3 recirculation.

184

N. Jing

Fig. 8.9 (a) Western blots were performed in prostate tumors that were harvested from the mice at the end period of drug treatments, including one sample of untreated tumors as a control and two samples of PEI-, ns-ODN (control ODN), and T40214-treated tumors, respectively [ 86 ] . ( b ) The pathway of inhibition of Stat3 by GQ-ODN T40214 in cancer therapy

The inhibition of Stat3 activation disrupts the translation of Stat3-regulated genes, including those encoding anti-apoptotic proteins (Bcl-2, Bcl-xL, Mcl-1, and survivin), cell-cycle regulators (cyclin D1 and c-myc), and inducers of angiogenesis (VEGF) [84, 86]. Consequently, T40214 – with the inhibition of Stat3 activation – significantly promotes apoptosis, reduces angiogenesis and cell proliferation, and strongly suppresses tumor growth.

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

185

Toxicity of GQ-ODN T40214/PEI Complex Toxicity of GQ-ODNs: G-Quartet ODNs Are Low-Toxicity Agents Toxicity studies have been reported for AR177, an analogue of GQ-ODN T40214 [91]. GQ-ODN exhibited no genetic toxicity in three different mutagenic assays: Ames/Salmonella mutagenesis assay, CHO/HGPRT mammalian cell mutagenesis assay, and mouse micronucleus assay. Acute toxicity studies in mice showed that GQ-ODN had an LD50 (the lethal dose to half of the animals) of 1.5 g/kg body weight, which is more than 150-fold higher than our in vivo therapeutic dose. Also, the multiple-dose-toxicity studies in mice showed that GQ-ODN did not cause specific mortality and changes in serum chemistry, hematology, and histology until doses reached 600 mg/kg, which are >60-fold that of therapeutic level. Clinical chemistry findings included changes in liver function and decreased erythrocyte values at 250 and 600 mg/kg.

Toxicity of PEI PEI (polyethylenimine, 25 K) as a DNA carrier has been widely used due to its high delivery efficiency and low toxicity [92, 93]. The ratio of PEI to DNA is critical for complex formation. PEI as a DNA carrier has a positively charged molecular surface that improves binding with DNA oligos and increases delivery efficiency since the positive charges enhance cellular uptake [94, 95]. PEI also protects DNA against nuclease cleavage; however, excessive PEI (or using it alone) could cause toxicity. The N/P ratio (nitrogen atoms of PEI to DNA phosphates) determines the positive net charge of the PEI/DNA complex, which dramatically influences the efficiency of the DNA delivery system. Previous studies suggest that when an N/P ratio is lower than 3.0, PEI has the highest efficacy and lowest toxicity in animal studies; decreasing the N/P ratio by decreasing the cationic charge density is beneficial for the in vivo microenvironment [95].

T40214/PEI Complex Shows No Toxicity Based on the structure and sequence of T40214, the N/P ratio of T40214/PEI complex in our studies was adjusted to lower than 2.7. At this ratio, we did not find any toxicity induced by T40214/PEI complex in normal epithelial cells and in normal tissues. TUNEL assay was used to determine the effect of T40214/PEI on apoptosis in normal epithelial cells compared to malignant epithelial cells. As shown in Fig. 8.10a, the cancer cells (HepG2) (lower panels) and normal epithelial cells (NRK-52E) (top panels) were pre-incubated with PEI alone (middle panels) or T40214/PEI complex (right panels) under the same conditions for 24 h. Apoptotic cells stained in dark brown in photomicrographs. Compared with control cells

186

N. Jing

Fig. 8.10 (a) TUNEL of T40214/PEI complex in cancer and normal cells. Apoptosis cells stained in dark brown. (b) Western blots showed that Stat3 was not constitutively phosphorylated in normal tissues, and T40214 does not inhibit the unphosphorylated Stat3 (T-Stat3) in normal tissues. The control sample was harvested from a tumor in an untreated mouse. (c) TUNEL of T40214/PEI in NSCLC tumors and normal tissues, which were harvested from the tissues around the NSCLC tumors in mice

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

187

(left panels), PEI did not induce apoptosis in cancer or normal epithelial cells (middle panels). In contrast, T40214 (right panels) significantly increased apoptosis in cancer cells, where p-Stat3 is activated, but not in normal epithelial cells, demonstrating that T40214/PEI is not toxic to normal cells. To determine whether T4021/PEI induces toxicity in normal animal tissues by inhibiting p-Stat3 activation, we harvested the normal tissues (e.g., kidney and liver) from the animals grown with prostate tumors (PC3): two samples from untreated and two samples from T40214/PEI-treated (10 + 2.5 mg/kg) nude mice. Western blots (Fig. 8.10b) demonstrated no phosphorylated Stat3 (p-Stat3) in normal tissues. This may be a result of the rapid dephosphorylation of Stat3 in normal cells in contrast to tumors cells [96]. T40214/PEI did not decrease expression of total Stat3 (T-Stat3) in normal tissues, consistent that T40214/PEI only reduced levels of phosphorylated Stat3, but not total Stat3, in tumors (Fig. 8.9a). Also, we performed TUNEL staining to determine if T40214/PEI induces apoptosis of normal tissues. The samples were harvested from NSCLC tumors (Fig. 8.10c, lower panels) and the normal tissues grown around the NSCLC tumors (Fig. 8.10c upper panels) in nude mice xenografts. PEI alone did not induce apoptosis in either normal or tumor cells assessed by TUNEL staining; however, T40214/PEI significantly promoted apoptosis in tumor cells but not in normal cells. Thus, the T40214/PEI complex would be expected to demonstrate low toxicity in clinical trials.

Summary Stat3: An Important Target in Cancer Therapy Stat3 has been identified as an important target for cancer therapy since Stat3, which is constitutively activated in many human cancers, participates in oncogenesis through the upregulation of genes encoding apoptosis inhibitors, cell-cycle regulators, and inducers of angiogenesis. Also, Stat3 is constitutively activated both in tumor cells and in immune cells in the tumor microenvironment. Constitutively activated Stat3 in tumor cells inhibits the expression of mediators necessary for immune activation against tumor cells and results in production of numerous mediators such as VEGF which inhibit dendritic cell function. Therefore, targeting Stat3 can produce a novel and effective treatment for human cancers.

Potential Impact of T40214 on Cancer Therapy We have laid the initial groundwork in the development of the G-quartet oligodeoxynucleotide (GQ-ODN) T40214 as a potent inhibitor of Stat3, which has several unique features. First, T40214 selectively targets phosphorylated Stat3 in vitro and

188

N. Jing

in vivo. The phosphorylated Stat3 participates in oncogenesis through the upregulation of genes encoding apoptosis inhibitors, cell-cycle regulators, and inducers of angiogenesis. Activation of Stat3 also has a role in protecting cancer cells from the immune system. Second, by inhibition of Stat3 activation, T40214 promotes apoptosis, reduces angiogenesis and cell proliferation, and significantly suppresses tumor growth in animal models, including prostate, breast, head and neck, and lung cancers. Third, T40124 has been showed to have a prolonged in vivo effect and significantly increased the survival of mice with the androgen-independent prostate tumors. Finally, the animal studies showed that T40214 appears to be nontoxic with fewer side effects than chemotherapeutic agents that are now used to treat patients. Therefore, T40214 could provide a novel strategy for cancer therapy and is expected to have potential benefits for the patients with cancers in clinical treatments. Acknowledgments Author greatly appreciates all the post doctors, staffs, and collaborators who made contributions to develop GQ-ODN T40214 as an anticancer agent. This project was mainly supported by grants DOD PC020407 and R01 CA104035.

References 1. Darnell JE. 1997, Science, 277, 1630–5. 2. Akira S. 2000, Oncogene, 19, 2607–11. 3. Bromberg JF, Horvath CM, Wen Z, Schreiber RD, Darnell JE. 1996, Proc Natl Acad Sci USA, 93, 7673–8. 4. Cressman DE, Greenbaum LE, DeAngelis RA, Ciliberto G, Furth EE, Poli V, Taub R. 1996, Science, 274, 1379–83. 5. Fukada T, Hibi M, Yamanaka Y, Takahashi-Tezuka M, Fujitani Y, Yamaguchi T, Sugiyama K, Yamanaka Y, Nakajima K, Hirano T. 1996, Immunity, 5, 449–60. 6. Hirano T, Ishihara K, Hibi M. 2000, Oncogene, 19, 2548–56. 7. Ihle JN. Cytokine receptor signalling. 1995, Nature, 377, 591–4. 8. Kaplan MH, Sun YL, Hoey T, Grusby MJ. 1996, Nature, 382, 174–7. 9. Smithgall TE, Briggs SD, Schreiner S, Lerner EC, Cheng H, Wilson MB. 2000, Oncogene, 19:2612–8. 10. Stephens JM, Morrison RF, Pilch PF. 1996, J Biol Chem. 271, 10441–4. 11. Takeda K, Noguchi K, Shi W, Tanaka T, Matsumoto M, Yoshida N, Kishimoto T, Akira S. 1997, Proc Natl Acad Sci USA. 94, 3801–4. 12. Zhong Z, Wen Z, Darnell JE. 1994, Science, 264, 95–8. 13. Zhang X, Blenis J, Li HC, Schindler C, Chen-Kiang S. 1995, Science, 267, 1990–4. 14. Braunstein J, Brutsaert S, Olson R, Schindler C. 2003, J Biol Chem. 278, 34133–40. 15. Bowman T, Garcia R, Turkson J, Jove R. 2000, Oncogene, 19, 2474–88. 16. Buettner R, Mora LB, Jove R. 2002, Clin Cancer Res. 8, 945–54. 17. Yu H, Jove R. 2004, Nat Rev Cancer, 4, 97–105. 18. Becker S, Groner B, Muller CW. 1998, Nature, 394, 145–51. 19. Chen X, Vinkemeier U, Zhao Y, Jeruzalmi D, Darnell JE, Jr., Kuriyan J. 1998, Cell, 93, 827–39. 20. Lutticken C, Wegenka UM, Yuan J, Buschmann J, Schindler C, Ziemiecki A, et al. 1994, Science, 263, 89–92. 21. Wegenka UM, Lutticken C, Buschmann J, Yuan J, Lottspeich F, Muller-Esterl W, et al. 1994, Mol Cell Biol. 14, 3186–96.

8 GQ-ODN T40214, a Rational Designed Stat3 Inhibitor

189

22. Akira S, Nishio Y, Inoue M, Wang XJ, Wei S, Matsusaka T, et al. 1994, Cell, 77, 63–71. 23. Schindler C, Darnell JE. 1995, Annu Rev Biochem. 64, 621–51. 24. Fukada T, Ohtani T, Yoshida Y, Shirogane T, Nishida K, Nakajima K, et al. 1998, Embo J., 17, 6670–7. 25. Chakraborty A, Tweardy DJ. 1998, Leuk Lymphoma, 30, 433–42. 26. de Koning JP, Soede-Bobok AA, Ward AC, Schelen AM, Antonissen C, van Leeuwen D, et al. 2000, Oncogene, 19, 3290–8. 27. Shimozaki K, Nakajima K, Hirano T, Nagata S. 1997, J Biol Chem., 272, 25184–9. 28. Shirogane T, Fukada T, Muller JM, Shima DT, Hibi M, Hirano T. 1999, Immunity, 11, 709–19. 29. Dolled-Filhart M, Camp RL, Kowalski DP, Smith BL, Rimm DL. 2003, Clin Cancer Res. 9, 594–600. 30. Mora LB, Buettner R, Seigne J, Diaz J, Ahmad N, Garcia R, et al. 2002, Cancer Res. 62, 6659–66. 31. Nagpal JK, Mishra R, Das BR. 2002 Cancer, 94, 2393–400. 32. Song L, Turkson J, Karras JG, Jove R, Haura EB. 2003, Oncogene, 22, 4150–65. 33. Yu H, Kortylewski M, Pardoll D. 2007, Nat Rev Immunol., 7, 41–51. 34. Darnell JE, Jr., Kerr IM, Stark GR.1994 Science, 264, 1415–21. 35. Silvennoinen O, Schindler C, Schlessinger J, Levy DE. 1993, Science, 261, 1736–9. 36. Meydan N, Grunberger T, Dadi H, Shahar M, Arpaia E, Lapidot Z, et al. 1996, Nature, 379, 645–8. 37. Nielsen M, Kaltoft K, Nordahl M, Ropke C, Geisler C, Mustelin T, et al. 1997, Proc Natl Acad Sci USA. 94, 6764–9. 38. Faderl S, Ferrajoli A, Harris D, Van Q, Priebe W, Estrov Z. 2005, Anticancer Res., 25, 1841–50. 39. Blaskovich MA, Sun J, Cantor A, Turkson J, Jove R, Sebti SM. 2003, Cancer Res., 63, 1270–9. 40. Shin DS, Kim HN, Shin KD, Yoon YJ, Kim SJ, Han DC, Kown BM. 2009, Cancer Res., 69, 193–202. 41. Turkson J, Ryan D, Kim JS, Zhang Y, Chen Z, Haura E, et al. 2001, J Biol Chem., 276, 45443–55. 42. Shao H, Cheng HY, Cook RG, Tweardy DJ. 2003, Cancer Res., 63, 3923–30. 43. Ren Z, Cabell LA, Schaefer TS, McMurray JS. 2003, Bioorg Med Chem Lett., 13, 633–6. 44. Leong PL, Andrews GA, Johnson DE, Dyer KF, Xi S, Mai JC, et al. 2003, Proc Natl Acad Sci USA. 100, 4138–43. 45. Barton BE, Karras JG, Murphy TF, Barton A, Huang HF. 2004, Mol Cancer Ther. 3, 11–20. 46. Grandis JR, Drenning SD, Zeng Q, Watkins SC, Melhem MF, Endo S, et al. 2000, Proc Natl Acad Sci USA. 97, 4227–32. 47. Karras JG, McKay RA, Lu T, Pych J, Frank DA, Rothstein TL, et al. 2000, Cell Immunol. 202, 124–35. 48. Chung CD, Liao J, Liu B, Rao X, Jay P, Berta P, et al. 1997, Science, 278, 1803–5. 49. Liu B, Liao J, Rao X, Kushner SA, Chung CD, Chang DD, et al. 1998, Proc Natl Acad Sci USA. 95, 10626–31. 50. Endo TA, Masuhara M, Yokouchi M, Suzuki R, Sakamoto H, Mitsui K, et al. 1997, Nature, 387, 921–4. 51. Naka T, Narazaki M, Hirata M, Matsumoto T, Minamoto S, Aono A, et al. 1997, Nature, 387, 924–9. 52. Starr R, Willson TA, Viney EM, Murray LJ, Rayner JR, Jenkins BJ, et al. 1997, Nature, 387, 917–21. 53. Shuai K, Liu B. 2003, Nat Rev Immunol. 3, 900–11. 54. Jing N, Sha W, Li Y, Xiong W, Tweardy DJ. 2005, Curr Pharm Des. 11, 2841–54. 55. Jing N, Tweardy DJ. 2005, Anticancer Drugs. 16, 601–7. 56. Henderson E. 1995, Telomeres. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory p. 11–34.

190

N. Jing

57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76.

Gellert M, Lipsett MN, Davies DR. 1962, Proc Natl Acad Sci USA. 48, 2013–8. Kang C, Zhang X, Ratliff R, Moyzis R, Rich A. 1992, Nature, 356, 126–31. Smith FW, Feigon J. 1992, Nature, 356, 164–8. Sundquist WI, Klug A. 1989, Nature, 342, 825–9. Fry M, Loeb LA. 1994, Proc Natl Acad Sci USA. 91, 4950–4. Awang G, Sen D. 1993, Biochemistry, 32, 11453–7. Sundquist WI, Heaphy S. 1993, Proc Natl Acad Sci USA.90, 3393–7. Sen D, Gilbert W. 1988, Nature, 334, 364–6. Gilbert DE, Feigon J. 1999, Curr Opin Struct Biol. 9, 305–14. Rhodes D, Giraldo R. 1995, Curr Opin Struct Biol. 5, 311–22. Shafer RH, Smirnov I. 2000, Biopolymers, 56, 209–27. Williamson JR. 1994, Annu Rev Biophys Biomol Struct. 23, 703–30. Sen D, Gilbert W. 1990, Nature, 344, 410–4. Jing N, Rando RF, Pommier Y, Hogan ME. 1997, Biochemistry, 36, 12498–505. Guo Q, Lu M, Kallenbach NR. 1993, Biochemistry, 32, 3596–603. Han H, Hurley LH. 2000, Trends Pharmacol Sci. 21, 136–42. Mergny JL, Phan AT, Lacroix L. 1998, FEBS Lett. 435, 74–8. Pilch DS, Plum GE, Breslauer KJ. 1995, Curr Opin Struct Biol. 5, 334–42. Jing N, Xiong W, Guan Y, Pallansch L, Wang S. 2002, Biochemistry, 41, 5397–403. Jing N, De Clercq E, Rando RF, Pallansch L, Lackman-Smith C, Lee S, et al. 2000, J Biol Chem. 275, 3421–30. Jing N, Hogan ME.1998, J Biol Chem. 273, 34992–9. Jing N, Marchand C, Liu J, Mitra R, Hogan ME, Pommier Y. 2000, J Biol Chem. 27, 21460–7. Jing N, Li Y, Xiong W, Sha W, Jing L, Tweardy DJ. 2004, Cancer Res. 64, 6603–9. Jing N, Li Y, Xu X, Sha W, Li P, Feng L, et al. 2003, DNA Cell Biol. 22, 685–96. Jing N, Gao X, Rando RF, Hogan ME. 1997, J Biomol Struct Dyn. 15, 573–85. Neeson P, Paterson Y. 2006, Immunol Invest. 35, 359–94. Jing N, Zhu Q, Yuan P, Li Y, Mao L, Tweardy DJ. 2006, Mol Cancer Ther. 5, 279–86. Weerasinghe P, Garcia GE, Zhu Q, Yuan P, Feng L, Mao L, Jing N. 2007, Int J Oncol. 31, 129–36. Zhu Q, Jing N. 2007, J Comput Aided Mol Des. 21, 641–8. Weerasinghe P, Li Y, Guan Y, Zhang R, Tweardy DJ, Jing N. 2008, Prostate, 68, 1430–42. Jemal A, Siegel R, Ward E, Murray T, Xu J, Thun MJ. 2007, Cancer J Clin. 57, 43–66. Edwards J, Bartlett JM. 2005, BJU Int. 95, 1320–6. Edwards J, Bartlett JM. 2005, BJU Int. 95, 1327–35. Tannock IF, de Wit R, Berry WR, Horti J, Pluzanska A, Chi KN, 2004, N Engl J Med. 351, 1502–12. Wallace TL, Gamba-Vitalo C, Loveday KS, Cossum PA. 2000, Toxicol Sci. 53, 63–70. Lungwitz U, Breunig M, Blunk T, Gopferich A. 2005, Eur J Pharm Biopharm. 60, 247–66. Neu M, Fischer D, Kissel T. 2005, J Gene Med. 7, 992–1009. Boussif O, Lezoualc’h F, Zanta MA, Mergny MD, Scherman D, Demeneix B, et al. 1995, Proc Natl Acad Sci USA. 92, 7297–301. Fahrmeir J, Gunther M, Tietze N, Wagner E, Ogris M. 2007, J Control Release. 122, 236–45. Darnell JF. 2005, Nat. Med. 11:595–596.

77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 87. 88. 89. 90. 91. 92. 93. 94. 95. 96.

Chapter 9

Estrogen Receptor Signaling in Lung Cancer P.A. Hershberger and J.M. Siegfried

Introduction Eighty-five percent of lung cancers are classified as non-small cell lung cancers (NSCLC), of which there are two major subtypes: adenocarcinoma and squamous cell carcinoma. Adenocarcinomas arise in the glandular tissues of peripheral airways. Squamous cell carcinomas arise in the epithelial cells of the central airways and show a greater association with smoking than do adenocarcinomas [1]. Eightyseven percent of lung cancers are attributable to smoking. Smoking-related lung cancers develop when tobacco pro-carcinogens (such as benzo[a]pyrene) become activated and form DNA adducts in lung epithelial cells. If these adducts are not repaired correctly, they can lead to DNA mutations that increase cancer risk. It has been proposed that women may be more susceptible to the harmful effects of tobacco than are males. This is based on the findings that female lung cancer patients have a higher level of smoking-induced DNA adducts [2, 3] and decreased DNA repair capacity [4] compared to male patients. However, inconsistent results have been obtained in epidemiological studies designed to test this hypothesis formally (reviewed in [5]). Despite the debate that surrounds this issue, it is now well accepted that women are more susceptible to those lung cancers that arise in the absence of a smoking history: Females who are lifetime nonsmokers are three times more likely than their male counterparts to develop lung cancer [6]. This striking sex difference in lung cancer presentation provides a strong rationale for considering the contribution of hormonal factors, such as estrogen, in the development and/ or progression of lung cancer.

J.M. Siegfried (*) Co-Director, Thoracic Malignancies Program, Department of Pharmacology, University of Pittsburgh Cancer Institute, Pittsburgh, PA 15213, USA e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_9, © Springer Science+Business Media, LLC 2012

191

192

P.A. Hershberger and J.M. Siegfried

Estrogen Receptors Mediating Estrogen Action The cellular response to estrogens is mediated by estrogen receptors ERa and ERb, which belong to the nuclear steroid hormone receptor superfamily. These receptor proteins function as sequence-specific, ligand-dependent, transcription factors and regulate the expression of genes implicated in cell-cycle control, signal transduction, and cell survival (as an example, see [7]). Analogous to other nuclear hormone receptors, the ERa protein contains a number of functional domains including (1) a sequence located near the NH2-terminus that functions as a ligand-independent transcriptional activation domain termed AF-1, (2) a central DNA-binding domain, and (3) a COOH-terminal domain that is responsible for ligand binding. The COOH terminus also contains a second transcriptional activation function, termed AF-2, which is responsible for ligand-dependent activation of the receptor (Fig. 9.1). A second high-affinity estrogen receptor, termed ERb, was cloned independently by two laboratories in 1996 [8, 9]. Although the structure of the ERb protein is similar to ERa, the AF-1 domain of ERb is shorter and has a lower transcriptional activation capacity than the corresponding domain in ERa [10]. Both ERa and ERb bind with high affinity to the natural ligand 17b-estradiol (E2). According to a classical model for estrogen induction of gene expression, the ligand-bound transcription factors are released from inhibitory heat shock proteins and recruited to chromatin through an interaction with specific DNA elements termed estrogen-response elements (EREs) located in the promoters of estrogenresponsive genes (Fig. 9.2). When bound by an agonist such as E2, the AF-2 domains of the receptors undergo a conformational change that creates a surface favorable for interaction with coactivator proteins such as AIB1, SRC-1, and GRIP1/TIF2 [11, 12]. The coactivators facilitate chromatin unwinding and the recruitment of the basal transcription machinery to the promoter, leading to an increase in gene transcription. The receptors may also be indirectly recruited to genomic regions through a tethering mechanism involving interactions with other transcription factors such as AP-1 and Sp1 [13, 14]. For a more detailed mechanistic description of estrogen-regulated transcription, readers are referred to the recent work of Kininis et al. [15].

Fig. 9.1 Structure of ERa. The functional domains which are described in the text above are indicated. The designations (A–F) refer to structural domains that are characteristic of nuclear hormone receptors. The percentage of sequence identity between ERa and ERb is shown for the transactivation and DNA-binding domains

9

Estrogen Receptor Signaling in Lung Cancer

193

Fig. 9.2 Schematic representation of the mechanism by which the ER activates transcription. Upon ligand binding, the ER is released from an inhibitory association with heat shock protein 90 (hsp90). The receptor dimerizes and binds to EREs located in the promoters of estrogen-responsive genes. When bound by an agonist, the receptor recruits coregulatory proteins belonging to the p160 and p300 families. These proteins facilitate chromatin unwinding, interact with the basal transcription machinery and increase gene transcription

Due to the ability of estrogens to promote breast cancer growth, pharmacologic agents aimed at inhibiting estrogen signaling have been developed. These agents fall into two classes: (1) Antiestrogens which inhibit E2 from binding to its receptor, and (2) aromatase inhibitors which suppress endogenous estrogen biosynthesis. Clinically utilized antiestrogens include the selective estrogen response modifiers (SERMs) tamoxifen and raloxifene, and the pure ER antagonist, ICI 182,780 (fulvestrant). The mechanisms of action of these agents are described in more detail in the paragraphs immediately below. Clinically utilized aromatase inhibitors include letrozole, anastrazole, and exemestane. These agents are described in more detail later in this chapter. SERMs bind to the ER and regulate its function in a tissue-selective manner. For example, tamoxifen functions as an ER antagonist in breast cancer but as an ER agonist in the uterus. Therefore, while tamoxifen is an effective agent for the prevention and treatment of hormone-responsive breast cancers, it increases the risk of developing endometrial cancer [16]. The work by Shang and Brown demonstrates that the tissue specificity of SERMs is controlled, at least in part, by the cellular balance in ER coactivator and corepressor expression [17]. In breast cancer cells that lack expression of the coactivator SRC-1, tamoxifen mediates the preferential recruitment of corepressor proteins NCoR and SMRT to the promoters of E2-regulated genes. No gene induction is observed. In contrast, tamoxifen stimulates coactivator recruitment and induces the expression of E2-regulated genes in endometrial cells that endogenously express SRC-1. In preliminary studies, we

194

P.A. Hershberger and J.M. Siegfried

observed that tamoxifen increased the growth of human lung tumors (Stabile and Siegfried, unpublished observations). Therefore, the use of tamoxifen in lung cancer has not been widely pursued by us due to concerns about its potential agonist actions. Fulvestrant is an analogue of estrogen that functions as a pure receptor antagonist [18]. When bound to ERa, fulvestrant disrupts nuclear localization of the receptor and promotes receptor degradation [19, 20]. In preclinical studies, fulvestrant was demonstrated to block E2 regulation of gene expression and the growth of estrogensensitive breast tumors in vitro and in vivo [21, 22]. The mechanism by which fulvestrant inhibits ERb signaling may be distinctive. Studies indicate that ERb is stabilized in vitro and in vivo and retains its mobility in the presence of fulvestrant [23–25]. Nonetheless, we and others have observed that in tumor cells that express ERb, fulvestrant decreases the activation of a synthetic promoter construct that contains an ERE [26, 27] and decreases cell proliferation [26, 28, 29].

Lung: An Estrogen-Responsive Tissue Reproductive organs and bone are classically recognized as being estrogen-responsive. To identify other organs that respond to estrogen exposure in vivo, Ciana et al. developed transgenic mice in which an ERE-luciferase reporter construct was expressed in 26 different tissues [30]. Estrogen-responsive tissues were identified by treating ovarectomized female transgenic mice with vehicle (controls) or a single subcutaneous injection of 50 mg/kg 17b-estradiol. Tissues were harvested after 16 h and analyzed for luciferase activity ex vivo. The lungs displayed 15-fold induction of reporter gene activity upon estradiol exposure, which was greater the effect observed in tissues that are considered to be hormone responsive including the bone, uterus, and mammary gland. Using a different ERE-luciferase reporter transgenic mouse model, Lemmen et al. showed that 17b-estradiol also significantly induced luciferase activity in the lungs of male mice [31]. Based on the induction of a transcriptional response at a classical ERE, the lungs of both male and female mice can be classified as estrogen-responsive tissues. Consistent with the notion that estrogen signaling through ERb plays an important role in normal lung development; targeted inactivation of ERb results in lung abnormalities. At 3 months of age, the number of alveoli per field is significantly decreased in lungs from female ERb−/− mice compared to WT controls (309 versus 433, respectively). ERb−/− females also display altered surfactant homeostasis [32]. By 5 months of age, both male and female ERb-deficient mice exhibit signs of significant lung dysfunction (abnormal extracellular matrix deposition and alveolar collapse), which leads to systemic hypoxia [33]. It is intriguing that this phenotype occurs in both adult female and male mice, which are predicted to differ in circulating estrogen levels. This may suggest that the regulation of extracellular matrix activity by ERb occurs in a ligand-independent manner or that estrogens are produced and act locally to regulate lung biology in both male and female mice.

9

Estrogen Receptor Signaling in Lung Cancer

195

Effects of Endogenous and Exogenous Estrogens (Hormone Replacement Therapy) on Lung Cancer Risk and Lung Cancer Outcomes It is well established that estrogens are required for mammary gland development, but that aberrant estrogen signaling leads to breast cancer. Perhaps an analogous situation exists in the lung, where estrogens are required for the development and maintenance of normal tissue architecture, while excessive signaling increases the risk of developing cancer. Epidemiologic studies have been conducted to explore both the relationship between estrogen exposure and risk of developing lung cancer and the relationship between estrogen exposure and survival following a lung cancer diagnosis. If estrogen plays a role in lung cancer initiation and/or promotes the growth of established tumors, it is predicted that lung cancer risk would increase and/or that survival would decrease with increased exposure to estrogens. Three recent reports are consistent with a detrimental effect of endogenous estrogens on lung cancer outcomes. In an analysis of the SEER database, we found that women diagnosed with either squamous cell carcinoma or bronchiolalveolar carcinoma who were between 55 and 59 years old (and were presumed to be postmenopausal) had significantly better survival than women who were 40–49 years old (and were presumed to be premenopausal) [34]. More specifically, we determined that 5-year adjusted survival rates were 19% for premenopausal women with squamous cell carcinoma versus 23% for postmenopausal women (p = 0.03) and 40% for premenopausal women with bronchiolalveolar carcinoma versus 54% for postmenopausal women (p = 0.04). No age-related survival differences were observed in males. In a retrospective analysis of SWOG clinical trials, Albain et al. identified a survival benefit for women diagnosed with advanced non-small cell lung cancer (NSCLC) who were treated with a platinum-containing regimen compared to comparably treated men (11 months versus 8 months) [35]. However, the survival advantage was observed only in women who were age 60 or older. In a third study, Ross et al. quantified free estradiol concentrations in blood samples collected from males enrolled in phase III clinical trials in advanced NSCLC [36]. It was discovered that men with high free estradiol levels had significantly poorer survival than men with lower estradiol levels. With regard to exogenous estrogens (i.e., hormone replacement therapy (HRT)), epidemiologic studies have not shown a consistent association between estrogen use and lung cancer risk (reviewed in [37]). However, in two of three recent studies, an association was established between HRT use and poor survival following a lung cancer diagnosis [38–40]. Ganti et al. conducted a retrospective chart review of approximately 500 women diagnosed with lung cancer between 1994 and 1999 at a community-based hospital. HRT was defined as the use of estrogen plus progestin or estrogen only for at least 6 weeks of continuous use prior to the diagnosis of lung cancer. These investigators found that women with lung cancer who used HRT were significantly younger (63 years old) than women with lung cancer who never used

196

P.A. Hershberger and J.M. Siegfried

HRT (68 years old). Furthermore, women who were diagnosed with lung cancer and used HRT had significantly worse overall survival (39 months) compared to women who did not use HRT (79 months). Ayeni et al also conducted a retrospective chart review of nearly 400 women diagnosed with lung cancer between 1999 and 2003 in Ontario to test the hypothesis that HRT use impacted lung cancer survival. These investigators found no effect of HRT on overall survival, with reported median survivals of 13 months in the nonHRT group and 14 months in the HRT group. Of note, the cohort analyzed by Ayeni reported greater HRT use and included a greater percentage of cases of advanced NSCLC than the population studied by Ganti. The dramatic difference in overall survival between the non-HRT populations in the two studies (79 months versus 13 months) provides evidence of non-equivalence in the population of patients analyzed. Potential confounders of the above-described studies included the fact that the precise duration of HRT use was not specified, nor was a distinction made between the use of estrogen only versus the use of estrogen plus progestin. Both of these limitations were overcome in the study by Chlebowski et al., in which lung cancer outcomes were assessed in a secondary analysis of the Womens’ Health Initiative (WHI) randomized clinical trial [40]. In the WHI trial, more than 16,000 postmenopausal women were randomized to receive placebo or daily HRT (comprised of 0.625 mg estrogen plus 2.5 mg medroxyprogesterone acetate) for a period of 5.6 years. Factors known to influence lung cancer outcomes such as smoking history and age were balanced between the two groups. The investigators found a trend toward more NSCLC diagnoses in the HRT group compared to the placebo group, although this did not reach statistical significance. Although lung cancer risk may not have increased, the HRT group experienced a significantly greater likelihood of dying from NSCLC than did the placebo group (46% mortality versus 27%, respectively). These data support the conclusion that HRT use (as defined above) for a period of greater than 5 years significantly increases a woman’s chance of dying from lung cancer. This conclusion is consistent with the hypothesis that HRT provides a tumor growth advantage in lung cancer, as it does in breast cancer.

Preclinical Studies Indicating that Estrogen Supports the Development or Growth of NSCLC To evaluate the role of estrogen in lung cancer further, the effects of E2 have been studied using preclinical models. In an animal model in which lung adenocarcinomas were induced by K-ras activation and p53 deletion, E2 promoted tumor progression: Compared to untreated controls, E2 administration doubled the number of tumor foci evident on the lung surface in both males and ovariectomized females [41]. In established lung cancer cell lines, E2 significantly increased cell proliferation in vitro and in vivo, increased gene expression, and promoted VEGF secretion, and induced rapid phosphorylation/activation of p44/p42 mitogen activated

9

Estrogen Receptor Signaling in Lung Cancer

197

protein kinase (MAPK) [26, 42–45]. The induction of pro-proliferative/pro-survival responses by E2 provides an explanation for the poorer clinical outcomes observed in NSCLC patients who are either predicted or documented to have high estrogen levels. Given the ability of estrogens to increase gene expression in and stimulate the growth of NSCLC cells, preclinical investigations into the potential use of antiestrogens in lung cancer therapy have been initiated by us and others. To date, most efforts have focused on use of the pure ER antagonist, fulvestrant. Fulvestrant significantly inhibits E2-mediated stimulation of gene expression (measured by EREluciferase reporter assays), MAPK activation (measured by immunofluorescence), and proliferation of NSCLC cells in vitro [26, 44]. Fulvestrant also significantly inhibits the growth of 201 T, H23, and A549 lung tumor xenografts [26, 44, 46], providing strong rationale for the evaluation of this antiestrogen in lung cancer therapy.

Evaluation of the Functionality of the Classical Genomic Estrogen-Signaling Pathway in NSCLC Cells According to a classical model for estrogen regulation of gene expression, transcription is increased when ligand-bound ERs are recruited to chromatin through an interaction with EREs, which are located in the promoters of estrogen-responsive genes. To evaluate the functionality of this pathway in NSCLC cell lines in a facile manner, we have transfected cells that express endogenous ERs with an ERE-tkluciferase reporter construct. Following transfection, the cells received vehicle (controls) or were treated with increasing concentrations of E2. Luciferase reporter gene activity, which serves as a measure of ER-mediated transcription, was quantified the next day. In H23 and 201 T cells, nanomolar concentrations of E2 stimulate reporter gene activity approximately twofold [26, 27]. Indicative of involvement of the ER, E2-mediated activation of reporter gene expression in these cells was significantly blocked by fulvestrant.

Membrane-Initiated Steroid Signaling (MISS) in Lung Epithelial Cells and NSCLC Steroid hormones, including estrogen, rapidly activate second messenger–signaling pathways, leading to intracellular increases in cAMP, calcium fluxes, and activation of PI3K and MAPK. This process, which is initiated at the plasma membrane, leads to changes in protein structure/function and gene transcription and is imperative in eliciting full responses to estrogen. For example, pharmacologic inhibition of PI3K or MAPK prevents 17b-estradiol from stimulating the proliferation of endothelial cells [47]. The identity of the estrogen-binding protein responsible for plasma

198

P.A. Hershberger and J.M. Siegfried

membrane signaling has been the subject of much discussion, and it has been hypothesized that alternative receptors (other than the classical nuclear ER) are responsible. However, elegant studies by Pedram et al. have shown that, in breast cancer cells, the membrane-signaling protein is similar to or the same as the nuclear ERa [47]. In these studies, E2-binding proteins were purified from the nucleus and plasma membrane fractions of MCF-7 cells and identified by mass spectrometry. The plasma membrane–associated protein had a molecular weight of 67 kDa and had a sequence corresponding to the nuclear ERa protein. Consistent with a common identification for the membrane receptor and the nuclear receptor, transfection of siRNA directed against classical ERa in MCF-7 cells eliminates membraneinitiated estrogen signaling [47]. Studies by a number of different investigators show that 17b-estradiol initiates membrane signaling in NSCLC cells, as evidenced by rapid increases in phosphorylation of p42/p44 MAPK and/or Akt [27, 44, 45, 48]. Plasma membrane fractions from NSCLC cells contain estradiol-binding activity, and isolated caveolae from NSCLC cells contain proteins that react with antibodies specific for nuclear ERa and ERb [44]. These data support the hypothesis that a subset of the classical ER pool associates with the plasma membrane and triggers membrane signaling in NSCLC cells, as it does in breast cancer cells.

Estrogen Receptors and Mitochondria It has long been known that estrogens increase mitochondrial function. In recent years, interest in understanding the mechanistic basis for estrogen action at the mitochondria has re-emerged and resulted in the conduct of a series of elegant studies which show that: (a) nuclear ERa activates transcription of nuclear respiratory factor-1 (NRF-1) in breast cancer cells [49], (b) ERb modulates mitochondrial sensitivity to oxidative stressors in HT-22 hippocampal cells [50], and (c) ERa and ERb bind to human mitochondrial DNA EREs and may thus regulate the expression of genes encoded by the mitochondrial genome [51]. For a comprehensive consideration of this topic area, readers are directed to two recent reviews [52, 53]. The Nuclear Respiratory Factor-1 (NRF-1) gene encodes a transcription factor that activates the expression of genes required for mitochondrial DNA transcription and replication [54]. Mattingly et al. demonstrated that NRF-1 mRNA expression is increased by E2 treatment both in MCF-7 breast cancer cells via an ERa dependent mechanism and in H1793 lung cancer cells by an ERb-dependent mechanism [49]. NRF-1 induction by E2 was attributed to direct binding of the ligand-activated nuclear ER to the NRF-1 promoter based on the results of chromatin-immunoprecipitation studies and transient transfection assays which employed a NRF-1 promoter-reporter plasmid. E2 treatment of MCF-7 cells also led to (1) an increase in the expression of the NRF-1 target gene, TFAM; (2) the subsequent transcription

9

Estrogen Receptor Signaling in Lung Cancer

199

of TFAM-regulated mitochondrial DNA encoded genes; and (3) increased mitochondrial biogenesis. Based on this evidence, the authors conclude that E2 stimulates mitochondrial function/biogenesis through a genomic pathway that is initiated by the binding of nuclear ERs to the NRF-1 promoter. The observation that ERb localizes to the mitochondria in multiple cell types (including but not limited to neurons, breast cancer cells, skeletal muscle cells, and human lens cells) is consistent with a local role for this protein in the direct regulation of mitochondrial function [55–58]. Mitochondrial ER (mtER) is absent in endothelial cells isolated from ERa/ERb knockout mice, indicating that the mtER is derived from the same gene that encodes the nuclear ER [59]. In MCF-7 cells, E2 treatment caused a dose- and time-dependent increase in both the amount of ERb associated with the mitochondria and the abundance of mRNAs transcribed from genes encoded by mitochondrial DNA [56]. The concordance of these events raises the possibility that mitochondrial localized ERb directly affects transcription of the mitochondrial genome. Human mitochondrial DNA contains putative EREs (mtEREs). When oligonucleotides containing mtERE sites were used in electrophoretic mobility assays, mitochondrial extracts prepared from MCF-7 cells (that contained both ERa and ERb) displayed specific binding [51]. Protein binding to the mtEREs was increased in a dose- and time-dependent manner by E2 and decreased by ERb-specific (but not ERa-specific) antibodies. These data support a model in which E2 enhances mitochondrial function by a direct mechanism involving the binding of ERb in the mitochondria to EREs within mtDNA. ERb specific siRNA depletes the mitochondrial pool of estrogen receptors in transformed HT-22 hippocampal cells. Interestingly, HT-22 cells with engineered loss of ERb are relatively refractory to cell death induced by mitochondrial stressors [50]. Compared to WT controls, ERb knock-down cells also displayed reduced superoxide production after oxidative insult, had reduced resting mitochondrial membrane potential, and displayed increased resistance to hydrogen peroxideinduced membrane depolarization. Based on these findings, the authors conclude that ERb is a factor that renders mitochondria/cells vulnerable to oxidative stress. Because E2-treated WT HT-22 cells have a phenotype similar to ERb knock-down cells, the authors surmise that estrogen signaling through ERb nullifies the ability of ERb to promote mitochondrial vulnerability. According to such a model, an increase in estrogen signaling through ERb in the lung would be expected to promote lung cancer development or progression by reducing cellular susceptibility to oxidative insults. Conversely, it is predicted that lung cancer cells would become more susceptible to stress/apoptosis inducing agents under conditions of estrogen deficiency. Although mitochondrial localization of ERb in NSCLC cell lines has been reported [49], the majority of primary NSCLC specimens analyzed to date display primarily a nuclear pool of receptors (see section immediately below). Given this discrepancy, it is too early to comment on the contribution of mitochondrial ERb to the development or progression of NSCLC. Studies are required to determine whether ERb is expressed in the mitochondria of primary lung cancer cells and whether estradiol regulates the function of ERb from this subcellular location.

200

P.A. Hershberger and J.M. Siegfried

Estrogen Receptors Expressed in NSCLC Cells and Human Primary Lung Tumors As evidence accumulates indicating that estrogen promotes lung cancer growth, interest in identifying the receptor that is responsible has increased. Our initial approach to identify the biologically relevant receptor(s) involved assessing the expression of ERa and ERb mRNA and protein in NSCLC cell lines. In the mRNA analysis of our cell lines, we found that ERa transcripts were detected but were comprised of mostly alternatively spliced variants [26]. Consistent with this observation, we were unable to detect the predicted full-length 66-kD ERa protein in whole-cell extracts prepared from NSCLC cell lines despite the use of several different antibodies [26, 27, 42]. However, we did observe immunoreactive bands of »42 kD and »54 kD using antibodies against the hinge region and COOH terminus of ERa, respectively. Although we cannot exclude the possibility that these smaller proteins represent variant forms of ERa, they do not share common epitopes or subcellular localization and may represent non-specific cross-reactive bands [27]. In contrast to ERa, ERb transcripts and a full-length protein are consistently detected in the cell lines we have analyzed to date (including but not limited to 201 T, A549, H23, and 128.88 T). We conclude from our studies that ERb is the primary estrogen receptor expressed in NSCLC cells. In a complementary approach, investigators have sought to identify the biologically relevant receptor in lung cancer by using immunohistochemistry (IHC) to establish the relationship between ER expression in lung tumor tissue and disease outcomes. In 4 recent studies that utilized different ERb antibodies and distinctive scoring criteria to establish a NSCLC case as receptor positive, 45–69% of specimens were characterized as expressing a nuclear pool of ERb proteins [60–63]. In each of these, high nuclear ERb was found to be a favorable prognostic indicator, although this was observed only in males in two of the studies [60–63]. We recently completed an immunohistochemical analysis of approximately 160 NSCLC cases in which the relationship between ERb expression in different cellular compartments and disease outcome was analyzed. In contrast to other studies, we found that the cytoplasmic form of ERb was a negative prognostic factor for overall survival in both men and women, while nuclear ERb levels did not correlate with survival (unpublished observations). Our finding may indicate that induction of non-genomic signaling is the more important biological function of ERb. We also found that expression of progesterone receptor was protective (unpublished observation). Possibly, the interplay of both receptors is important in outcome. The analysis of ERa-staining patterns in NSCLC has proven to be difficult. Nuclear ERa immunostaining is either never, or only rarely, detected in primary lung tumors [60–63]. ERa is detected in the cytoplasm of some NSCLC cases, but the percentage of positive specimens varies considerably between studies (from approximately 3% [60] to 73% [62]). Even within the same study, the ability to detect a cytoplasmic pool of ERa was found to depend upon the antibody employed: An immunoreactive pool of receptors was detected when an antibody specific for a

9

Estrogen Receptor Signaling in Lung Cancer

201

COOH-terminal epitope of ERa was used, but not when an antibody specific for an NH2-terminal epitope was used [62]. This discrepancy may indicate that the COOH-terminal antibody reacts non-specifically or that the immunoreactive protein that is detected in the cytoplasm of lung tumors represents an ERa variant lacking the NH2-terminus. In studies where the prognostic significance of ERa expression was reported, it had either no correlation with survival [63] or correlated with poor prognosis [62].

Using Subtype Selective Ligands to Assess the Relative Contributions of ERb and ERa in NSCLC Cell Lines The sequences of ERa and ERb show only moderate conservation (approximately 53%) within the ligand-binding domain (Fig. 9.1). This has permitted the development of subtype-selective ligands which have been extensively used to distinguish ERa and ERb functions in cells/tissues that may express both receptors. Ligands that are commonly employed include the ERb subtype-selective agonists genistein (GEN) and 2,3-bis(4-hydroxyphenyl)propionitrile (DPN) and the ERa-selective agonist 4,4¢,4²-(4-propyl-[1 H]-pyrazole-1,3,5-triyl)trisphenol (PPT). GEN has a 26-fold binding affinity preference for ERb versus ERa [64]. DPN has a 70-fold binding affinity preference for ERb versus ERa [65]. PPT has a >400-fold binding affinity preference for ERa versus ERb [66]. As indicated above, E2 induces a variety of responses in NSCLC cells including cell proliferation, modulation of gene expression, and rapid MAPK activation. Because a majority of primary lung cancers express only ERb, we sought to determine whether ERb was sufficient to generate these responses. To do this, we determined the effects of the ERb-selective agonists GEN and DPN and the ERa-selective agonist PPT in 201 T cells that endogenously express ERb but not ERa [27]. GEN, but not PPT, mediated a significant 1.5-fold increase in reporter activity in cells transiently transfected with ERE-tk-luciferase, demonstrating that endogenous levels of ERb (but not those of ERa) are sufficient for transcription. We also studied membrane-initiated estrogen signaling in 201 T cells using MAPK phosphorylation as a read-out. PPT and DPN induced activation of MAPK with similar kinetics. Phosphorylation of MAPK was maximal within 5 min and returned to near-baseline levels within 30 min of treatment. However, only DPN mediated a statistically significant increase in 201 T cell growth in vitro and in vivo. We interpreted these results as evidence that endogenous levels of ERb are sufficient to generate genomic and non-genomic responses to estrogen in NSCLC cells, and that activation of both pathways is necessary for increased proliferation. We were unable to make any conclusions regarding the role of ERa in mediating estrogenic responses in lung cancer because our cell lines do not endogenously express full-length ERa protein. However, others have assessed the ability of endogenous ERa and ERb to increase the proliferation of NSCLC cells by transiently transfecting siRNA duplexes targeting each receptor into H23 cells that express

202

P.A. Hershberger and J.M. Siegfried

both ERa and ERb [46]. siRNA-mediated suppression of mRNA corresponding to either receptor resulted in approximately 20% inhibition of in vitro growth. This result suggests that ERa and ERb may contribute equally to the growth of lung cancer cells that express both receptors.

E2 Cooperates with Growth Factor Receptor Signaling to Increase the Growth of NSCLC Cells The epidermal growth factor receptor (EGFR) is a receptor tyrosine kinase that signals from the plasma membrane to regulate a variety of biological responses including proliferation, differentiation, migration, and apoptosis. The EGFR is recognized as a major therapeutic target in lung cancer, and a variety of agents to inhibit its activity have been developed. Most notable among these are the small molecule tyrosine kinase inhibitors gefitinib (Iressa) and erlotinib (Tarceva) and the EGFR blocking antibody, cetuximab (Erbitux). Currently, erlotinib is FDA-approved for use in patients with metastatic or locally advanced NSCLC who have failed at least one previous round of chemotherapy. Upon ligand binding, the EGFR dimerizes and undergoes autophosphorylation at tyrosine residues within its intracellular domain. These phosphotyrosines serve as binding sites for adapter molecules, which mediate the recruitment and activation of intracellular-signaling cascades. For example, phosphorylated EGFR binds to Grb2:SOS, which activates the p21Ras pathway leading to MAPK activation. Once activated, MAPK can translocate to the nucleus where it phosphorylates/activates nuclear transcription factors, including the ER. Phosphorylation of Ser118 in the A/B domain of ERa by MAPK leads to activation of the nuclear receptor [67], and provides a mechanism by which signaling through the EGFR can increase the activity of the steroid receptor-signaling pathway (Fig. 9.3). A reciprocal pathway for cross-talk between the ER and EGFR also exists, in which estrogen activation of the ER leads to enhanced signaling through EGFR. Evidence supportive of this mechanism includes the findings that EGF-induced DNA synthesis and transcription are absent in reproductive organs from ERa knockout mice [68], and that the ER antagonist, fulvestrant, blocks the ability of EGF to stimulate the proliferation of MCF-7 breast cancer cells [69]. Available data indicate that E2 mediates EGF signaling by activating cell surface-associated matrix metalloproteinases (MMP-2 and MMP-9) via a Src-dependent process. The proteinases then stimulate the release of EGFR ligands from the surface of the MCF-7 cells [70]. Because E2 and EGF stimulate the proliferation of NSCLC cells and the potential for cross-talk between the ER and EGFR pathways exists, Stabile et al. and Pietras et al independently launched a series of studies to explore the effect of combined targeting of the ER and EGFR pathways in NSCLC cells. Several observations were reported that provide provocative evidence for receptor cross-talk: (1) E2 induced rapid activation of MAPK in NSCLC cells in vitro, which was blocked by

9

Estrogen Receptor Signaling in Lung Cancer

203

Fig. 9.3 Cross-regulation between the ER and EGFR signaling pathways. E2 binding to ERs localized at the plasma membrane leads to the activation of matrix metalloproteinases (MMPs). MMPs catalyze the release of EGFR ligands (such as HB-EGF) from the cell surface. The free ligands bind to the EGFR and trigger the activation of downstream signaling pathways. ERK translocates to the nucleus upon its activation, where it can phosphorylate and activate the ER in a ligand-independent manner. Available data indicates that the plasma membrane ER and the nuclear ER are encoded by the same gene

the EGFR blocking antibody M225 [45], (2) the ER and the EGFR co-localized in caveolae purified from NSCLC cells [46], and (3) combination of E2 with EGF resulted in a greater than additive increase in MAPK activation in two different NSCLC cell lines. Stabile and Pietras then explored the therapeutic value of co-targeting the ER and EGFR pathways in NSCLC models. The combined use of fulvestrant plus the EGFR tyrosine kinase inhibitor (TKI) gefitinib resulted in greater growth inhibition than the TKI alone in 4 different NSCLC cell lines in vitro. In corresponding in vivo models, the growth of A549 human lung tumor xenografts was maximally suppressed by combination of fulvestrant with erlotinib [46], while the growth of 201 T human lung tumor xenografts was maximally inhibited by the combination of fulvestrant with gefitinib [45]. When compared with either single agent, the increased efficacy of the fulvestrant plus gefitinib combination in the 201 T xenograft model was associated with an increase in tumor cell apoptosis and a decrease in the proliferation marker, Ki67. Based on these compelling preclinical findings, a pilot clinical trial of gefitinib plus fulvestrant in the treatment of postmenopausal women with advanced NSCLC was conducted [71]. Twenty-two women (median age 66 year old) with pathologically confirmed advanced NSCLC were enrolled in the study, 14 of whom had

204

P.A. Hershberger and J.M. Siegfried

received at least 1 prior chemotherapy regimen. Eighteen women received gefitinib (250 mg orally once daily) plus fulvestrant (250 mg IM, on day 1 of each 28 day cycle). The remaining four women received gefitinib as above, but the fulvestrant dosing was modified to reduce the time required to achieve steady-state plasma concentrations. To do this, fulvestrant was administered at a loading dose of 500 mg IM on day 1, followed by 250 mg IM on days 15 and 29, and then administered every 28 days thereafter. Overall, the therapy was well tolerated. In 20 patients evaluable for response, three partial responses were observed. The median progressionfree survival for the population was 12 weeks, the median overall survival was 38.5 weeks, and the 1-year survival was 41%. The safety of the combination supported our subsequent design of a randomized phase II clinical trial in which the efficacy of erlotinib (i.e., objective tumor response) is being compared with erlotinib plus fulvestrant in patients with advanced NSCLC. This trial (ClinicalTrials.gov identifier NCT00100854) is currently open and accruing patients. The combination of fulvestrant plus erlotinib is also being evaluated in a separate Phase II trial led by Dr. Bazhenova at the University of California, San Diego (ClinicalTrials.gov identifier NCT00592007). This latter trial is designed to determine whether the addition of fulvestrant to erlotinib extends progression free survival in patients with advanced NSCLC. According to the design of this study, a patient must have a tumor that expresses the ER and have stable disease while on erlotinib monotherapy for a period of at least 2 months in order to be eligible to receive fulvestrant. Progression-free survival for the combination will be compared to historical controls of erlotinib monotherapy. Interestingly, a recent case report suggests that the combination of an EGFR TKI with hormone therapy will have efficacy in NSCLC [72]. A 57-year-old female with advanced NSCLC who was previously treated with cisplatin and vinrelobine presented with skin metastasis of lung adenocarcinoma. The skin lesions were positive for expression of both EGFR and ER. The patient received gefitinib which resulted in partial healing of the skin lesions and disease stabilization. Approximately 8 months later the aromatase inhibitor, letrozole, was added to her therapy. There was complete resolution of cutaneous lesions after 3 months of therapy with letrozole and gefitinib. At the time at which the case report was published (nearly 2 years after initiation of combination therapy), the patient showed no clinical or radiologic disease progression.

Aromatase Aromatase is a cytochrome P450 enzyme that catalyzes the synthesis of estrogens from androgens. Aromatase is over-expressed in breast cancers, where it supports the growth of estrogen-responsive tumors [73–75]. Steroidal (exemestane) and nonsteroidal (anastrazole or letrozole) inhibitors have been developed to circumvent the action of aromatase. The superior efficacy (i.e., disease free survival) of these agents over tamoxifen in the treatment of postmenopausal women with estrogen receptor– positive breast cancer was established in multiple, large clinical trials (reviewed

9

Estrogen Receptor Signaling in Lung Cancer

205

recently in [76, 77]). Consequently, treatment recommendations for adjuvant therapy of postmenopausal breast cancer patients with receptor-positive disease now include an aromatase inhibitor as part of initial treatment or following a course of treatment with tamoxifen. The observation that aromatase is expressed in human lung tissue [78, 79] raises the possibility that local tissue production of estrogen contributes to lung cancer growth, as it does in breast cancer, and that aromatase inhibitors may be useful in lung cancer therapy. To determine whether aromatase is expressed and active in NSCLC cells, Weinberg et al. quantified aromatase transcripts, protein, and activity in a panel of human NSCLC cell lines [80]. The authors observed that a majority of the lines expressed aromatase, and that protein expression level correlated with enzymatic activity. For example, A549 cells that expressed aromatase were capable of synthesizing estradiol from testosterone in vitro. Moreover, when A549 cells were implanted into ovariectomized mice, daily administration of the aromatase substrate androstenedione resulted in a significant increase in tumor growth compared to vehicle controls [81]. The stimulatory effect of androstenedione on the growth of aromatase-positive lung tumor xenografts was significantly inhibited by either the non-steroidal aromatase inhibitor, anastrazole [80] or the steroidal aromatase inhibitor, exemestane [82]. Together, these preclinical results support the notion that aromatase-positive NSCLC cells locally synthesize pro-proliferative estrogens from circulating androgen precursors and that blockade of estrogen production is a useful strategy to suppress tumor growth. To establish the clinical relevance of aromatase expression in lung cancer, Weinberg et al. subjected 53 primary NSCLC specimens to immunhistochemical staining with an aromatase-specific antibody. When a cut-off of 15% cells with specific staining was used as the definition of a positive case, 86% of primary NSCLC cases were determined to express aromatase [80]. Staining was observed in the tumor epithelial cells themselves, and no significant differences in expression were observed between adenocarcinomas and squamous cell carcinomas or between males and females. The aromatase enzyme detected by immunohistochemistry is likely to represent functional protein because archival tumor specimens displayed in vitro enzymatic activity. In a separate study that utilized 59 primary NSCLC specimens, Niikawa et al. found that a positive correlation exists between lung tumor expression of aromatase and intratumoral estradiol levels measured by liquid chromatography/electrospray tandem mass spectrometry [83]. These authors also observed that intratumoral concentrations of estradiol were positively associated with tumor size and Ki-67 labeling index only if the lung tumor cells expressed either ERa or ERb, consistent with the hypothesis that the locally produced estrogens support tumor growth via an ER-dependent pathway. Based on the observed expression of aromatase in clinical lung cancer specimens and the significant antitumor activity of aromatase inhibitors in preclinical xenograft models, clinical trials of aromatase inhibitors in lung cancer have been designed by us. While results from these trials will not be available for several years, results from a randomized trial of exemestane after 2–3 years of tamoxifen therapy in postmenopausal women with primary breast cancer provide provocative evidence of a

206

P.A. Hershberger and J.M. Siegfried

beneficial effect of aromatase inhibitors in lung cancer therapy [84]. In the exemestane treatment arm, 4/2,362 women (0.17%) developed primary lung cancer versus 12/2,380 women (0.5%) in the tamoxifen treatment arm.

Summary and Future Directions Preclinical studies from multiple laboratories provide consistent evidence that proliferative estrogen receptor (ER)–signaling pathways exist in NSCLC cell lines and that suppression of these pathways (via the use of ER antagonists or aromatase inhibitors) suppresses lung tumor growth in vitro and in vivo. It is also clear from preclinical studies that cross-regulation between the ER pathway and EGFR pathway occurs and results in enhanced growth. These latter findings provided rationale for the design and conduct of clinical trials combining an EGFR TKI plus fulvestrant in NSCLC patients. Results from these ongoing trials, and planned trials of novel regimens containing aromatase inhibitors, will be important in establishing the extent to which modulation of estrogen signaling provides therapeutic benefit in both males and females with lung cancer. Alternative approaches to suppressing estrogen signaling in lung cancer may follow the identification of the receptor that is responsible. Although most studies to date point to a prominent role for ERb, ERa and potential variants are observed in some cell lines and primary lung tumors. Additional experiments are warranted to establish the relative contribution of ERa and ERb in mediating genomic, nongenomic, and proliferative responses to estrogen in NSCLC cells that may express both receptors. As this field evolves, consideration should also be given to the potential impact of non-classical estrogen receptors, such as GPR30, on the response of NSCLC cells to estrogen and antiestrogens. Summarized within this chapter are preclinical studies which demonstrate that ERs can signal from multiple locations within the cell, including the nucleus, plasma membrane, and mitochondria. Ongoing efforts in our laboratories are aimed at elucidating how these pathways are interconnected in lung cancer cells and whether each must be properly engaged in order to support cell proliferation and survival.

References 1. Muscat JE, Wynder EL. Lung cancer pathology in smokers, ex-smokers and never smokers. Cancer Lett 1995;88(1):1–5. 2. Ryberg D, Hewer A, Phillips DH, Haugen A. Different susceptibility to smoking-induced DNA damage among male and female lung cancer patients. Cancer Res 1994;54(22):5801–3. 3. Mollerup S, Ryberg D, Hewer A, Phillips DH, Haugen A. Sex differences in lung CYP1A1 expression and DNA adduct levels among lung cancer patients. Cancer Res 1999;59(14): 3317–20. 4. Wei Q, Cheng L, Amos CI, et al. Repair of tobacco carcinogen-induced DNA adducts and lung cancer risk: a molecular epidemiologic study. J Natl Cancer Inst 2000;92(21):1764–72.

9

Estrogen Receptor Signaling in Lung Cancer

207

5. Patel JD, Bach PB, Kris MG. Lung cancer in US women: a contemporary epidemic. JAMA 2004;291(14):1763–8. 6. Parkin DM, Bray F, Ferlay J, Pisani P. Global cancer statistics, 2002. CA Cancer J Clin 2005; 55(2):74–108. 7. Frasor J, Danes JM, Komm B, Chang KCN, Lyttle CR, Katzenellenbogen BS. Profiling of Estrogen-Up- and Down-Regulated Gene Expression in Human Breast Cancer Cells: Insights into Gene Networks and Pathways Underlying Estrogenic Control of Proliferation and Cell Phenotype. Endocrinology 2003;144:4562–74. 8. Kuiper GG, Enmark E, Pelto-Huikko M, Nilsson S, Gustafsson JA. Cloning of a novel receptor expressed in rat prostate and ovary. Proc Natl Acad Sci USA 1996;93(12):5925–30. 9. Mosselman S, Polman J, Dijkema R. ER beta: identification and characterization of a novel human estrogen receptor. FEBS Lett 1996;392(1):49–53. 10. Delaunay F, Pettersson K, Tujague M, Gustafsson JA. Functional differences between the amino-terminal domains of estrogen receptors alpha and beta. Mol Pharmacol 2000;58(3): 584–90. 11. Brzozowski AM, Pike AC, Dauter Z, et al. Molecular basis of agonism and antagonism in the oestrogen receptor. Nature 1997;389(6652):753–8. 12. Shiau AK, Barstad D, Loria PM, et al. The structural basis of estrogen receptor/coactivator recognition and the antagonism of this interaction by tamoxifen. Cell 1998;95(7):927–37. 13. Kushner PJ, Agard, D.A., Greene, G.L., Scanlan, T.S., Shiau, A.K., Uht, R.M., Webb, P. Estrogen receptor pathways to AP-1. Journal Steroid Biochemistry Molecular Biology 2000;74:311–7. 14. Safe S, Kim K. Non-classical genomic estrogen receptor (ER)/specificity protein and ER/ activating protein-1 signaling pathways. J Mol Endocrinol 2008;41(5):263–75. 15. Kininis M, Chen BS, Diehl AG, et al. Genomic analyses of transcription factor binding, histone acetylation, and gene expression reveal mechanistically distinct classes of estrogen-regulated promoters. Mol Cell Biol 2007;27(14):5090–104. 16. Fisher B, Costantino JP, Wickerham DL, et al. Tamoxifen for prevention of breast cancer: report of the National Surgical Adjuvant Breast and Bowel Project P-1 Study. J Natl Cancer Inst 1998;90(18):1371–88. 17. Shang Y, Brown, M. Molecular Determinants for the Tissue Specificity of SERMs. Science 2002;295:2465–8. 18. Osborne CK, Wakeling A, Nicholson RI. Fulvestrant: an oestrogen receptor antagonist with a novel mechanism of action. Br J Cancer 2004;90 Suppl 1:S2-6. 19. Dauvois S, Danielian PS, White R, Parker MG. Antiestrogen ICI 164,384 reduces cellular estrogen receptor content by increasing its turnover. Proc Natl Acad Sci USA 1992;89(9): 4037–41. 20. Dauvois S, White R, Parker MG. The antiestrogen ICI 182780 disrupts estrogen receptor nucleocytoplasmic shuttling. J Cell Sci 1993;106 ( Pt 4):1377–88. 21. DeFriend DJ, Anderson E, Bell J, et al. Effects of 4-hydroxytamoxifen and a novel pure antioestrogen (ICI 182780) on the clonogenic growth of human breast cancer cells in vitro. Br J Cancer 1994;70(2):204–11. 22. Osborne CK, Coronado-Heinsohn EB, Hilsenbeck SG, et al. Comparison of the effects of a pure steroidal antiestrogen with those of tamoxifen in a model of human breast cancer. J Natl Cancer Inst 1995;87(10):746–50. 23. Van Den Bemd GJ, Kuiper GG, Pols HA, Van Leeuwen JP. Distinct effects on the conformation of estrogen receptor alpha and beta by both the antiestrogens ICI 164,384 and ICI 182,780 leading to opposite effects on receptor stability. Biochem Biophys Res Commun 1999; 261(1):1–5. 24. Peekhaus NT, Chang T, Hayes EC, et al. Distinct effects of the antiestrogen Faslodex on the stability of estrogen receptors-alpha and -beta in the breast cancer cell line MCF-7. J Mol Endocrinol 2004;32(3):987–95. 25. Damdimopoulos AE, Spyrou G, Gustafsson JA. Ligands differentially modify the nuclear mobility of estrogen receptors alpha and beta. Endocrinology 2008;149(1):339–45.

208

P.A. Hershberger and J.M. Siegfried

26. Stabile LP, Gaither Davis AL, Gubish CT, Hopkins TM, Luketich JD, Christie N, Finkelstein S, and Siegfried JM. Human Non-Small Cell Lung Tumors and Cells Derived from Normal Lung Express Both Estrogen Receptor a and b and Show Biological Responses to Estrogen. Cancer Research 2002;62:2141–50. 27. Hershberger PA, Stabile LP, Kanterewicz B, et al. Estrogen receptor beta (ERbeta) subtypespecific ligands increase transcription, p44/p42 mitogen activated protein kinase (MAPK) activation and growth in human non-small cell lung cancer cells. J Steroid Biochem Mol Biol 2009. 28. Lau KM, LaSpina M, Long J, Ho SM. Expression of estrogen receptor (ER)-alpha and ER-beta in normal and malignant prostatic epithelial cells: regulation by methylation and involvement in growth regulation. Cancer Res 2000;60(12):3175–82. 29. Belcher SM, Ma X, Le HH. Blockade of estrogen receptor signaling inhibits growth and migration of medulloblastoma. Endocrinology 2009;150(3):1112–21. 30. Ciana P, DiLuccio G, Belcredito S, Pollio G, Vegeto E, Tatangelo L, Tiveron C, Maggi A. Engineering of a Mouse for the in Vivo Profiling of Estrogen Receptor Activity. Molecular Endocrinology 2001;15:1104–13. 31. Lemmen JG, Arends RJ, van Boxtel AL, van der Saag PT, van der Burg B. Tissue- and timedependent estrogen receptor activation in estrogen reporter mice. J Mol Endocrinol 2004; 32(3):689–701. 32. Patrone C, Cassel TN, Pettersson K, Piao Y-S, Cheng G, Ciana P, Maggi A, Warner M, Gustafsson J-A, Nord M. Regulation of Postnatal Lung Development and Homeostasis by Estrogen Receptor b. Molecular and Cellular Biology 2003;23:8542–52. 33. Morani A, Barros RP, Imamov O, et al. Lung dysfunction causes systemic hypoxia in estrogen receptor beta knockout (ERbeta−/−) mice. Proc Natl Acad Sci USA 2006;103(18):7165–9. 34. Oton AB BC, Cai C, Owonikoko T, Gooding W, Siegfried J, Ramalingam R. Comparison of survival for non-small cell lung cancer between premenopausal and postmenopausal women: An analysis of the National Surveillance, Epidemiology and End Results (SEER) Database. Journal of Clinical Oncology, ASCO Annual Meeting Proceedings 2006;24(18 S):7038. 35. Albain KS UJ, Gotay CC, Davies AM, Edelman M, Herbst RS, Kelly K, Williamson S, Wozniak AJ, Gandara DR. Toxicity and survival by sex in patients with advanced non-small cell lung carcinoma on modern Southwest Oncology Group (SWOG) trials. Journal of Clinical Oncology ASCO Annual Meeting Proceedings 2007;25(18 S):7549. 36. Ross H OF, Bandstra B, Sandalic L, Bianco J, Bonomi P, Singer JW. Serum-free estradiol (E2) levels are prognostic in men with chemotherapy-naive advanced non-small cell lung cancer and performance status (PS) 2. Journal of Clinical Oncology ASCO Annual Meeting Proceedings 2007;25(18 S):7683. 37. La Vecchia C. Hormone replacement therapy in menopause and lung cancer: an update. Eur J Cancer Prev 2006;15(3):189–90. 38. Ganti AK, Sahmoun AE, Panwalkar AW, Tendulkar KK, Potti A. Hormone replacement therapy is associated with decreased survival in women with lung cancer. J Clin Oncol 2006;24(1):59–63. 39. Ayeni O, Robinson A. Hormone replacement therapy and outcomes for women with nonsmall-cell lung cancer: can an association be confirmed? Curr Oncol 2009;16(3):21–5. 40. Chlebowski R.T. SA, Wakelee H., Anderson G.L., Stefanick M.L., Manson J.E., Chien J.W., Chen C., Wactawski-Wende J., Gass M. Non-small cell lung cancer and estrogen plus progestin use in postmenopausal women in the Women’s Health Initiative randomized clinical trial. J Clin Oncol 2009;27:18 s. 41. Hammoud Z, Tan B, Badve S, Bigsby RM. Estrogen promotes tumor progression in a genetically defined mouse model of lung adenocarcinoma. Endocr Relat Cancer 2008;15(2):475–83. 42. Hershberger PA, Vasquez AC, Kanterewicz B, Land S, Siegfried JM, Nichols M. Regulation of endogenous gene expression in human non-small cell lung cancer cells by estrogen receptor ligands. Cancer Res 2005;65(4):1598–605. 43. Jarzynka MJ, Guo P, Bar-Joseph I, Hu B, Cheng SY. Estradiol and nicotine exposure enhances A549 bronchioloalveolar carcinoma xenograft growth in mice through the stimulation of angiogenesis. Int J Oncol 2006;28(2):337–44.

9

Estrogen Receptor Signaling in Lung Cancer

209

44. Pietras RJ, Marquez DC, Chen HW, Tsai E, Weinberg O, Fishbein M. Estrogen and growth factor receptor interactions in human breast and non-small cell lung cancer cells. Steroids 2005;70(5–7):372–81. 45. Stabile LP, Lyker JS, Gubish CT, Zhang W, Grandis JR, Siegfried JM. Combined Targeting of the Estrogen Receptor and the Epidermal Growth Factor Receptor in Non-Small Cell Lung Cancer Shows Enhanced Anti-Proliferative Effects. Cancer Research 2005; 65(4):1459–70. 46. Marquez-Garban DC, Chen HW, Fishbein MC, Goodglick L, Pietras RJ. Estrogen receptor signaling pathways in human non-small cell lung cancer. Steroids 2007;72(2):135–43. 47. Pedram A, Razandi M, Levin ER. Nature of functional estrogen receptors at the plasma membrane. Mol Endocrinol 2006;20(9):1996–2009. 48. Zhang G, Liu X, Farkas AM, et al. Estrogen receptor beta functions through nongenomic mechanisms in lung cancer cells. Mol Endocrinol 2009;23(2):146–56. 49. Mattingly KA, Ivanova MM, Riggs KA, Wickramasinghe NS, Barch MJ, Klinge CM. Estradiol stimulates transcription of nuclear respiratory factor-1 and increases mitochondrial biogenesis. Mol Endocrinol 2008;22(3):609–22. 50. Yang SH, Sarkar SN, Liu R, et al. Estrogen receptor beta as a mitochondrial vulnerability factor. J Biol Chem 2009;284(14):9540–8. 51. Chen JQ, Eshete M, Alworth WL, Yager JD. Binding of MCF-7 cell mitochondrial proteins and recombinant human estrogen receptors alpha and beta to human mitochondrial DNA estrogen response elements. J Cell Biochem 2004;93(2):358–73. 52. Klinge CM. Estrogenic control of mitochondrial function and biogenesis. J Cell Biochem 2008;105(6):1342–51. 53. Simpkins JW, Yang SH, Sarkar SN, Pearce V. Estrogen actions on mitochondria--physiological and pathological implications. Mol Cell Endocrinol 2008;290(1–2):51–9. 54. Virbasius CA, Virbasius JV, Scarpulla RC. NRF-1, an activator involved in nuclear-mitochondrial interactions, utilizes a new DNA-binding domain conserved in a family of developmental regulators. Genes Dev 1993;7(12A):2431–45. 55. Yang SH, Liu R, Perez EJ, et al. Mitochondrial localization of estrogen receptor beta. Proc Natl Acad Sci USA 2004;101(12):4130–5. 56. Chen JQ, Delannoy M, Cooke C, Yager JD. Mitochondrial localization of ERalpha and ERbeta in human MCF7 cells. Am J Physiol Endocrinol Metab 2004;286(6):E1011-22. 57. Milanesi L, Vasconsuelo A, de Boland AR, Boland R. Expression and subcellular distribution of native estrogen receptor beta in murine C2C12 cells and skeletal muscle tissue. Steroids 2009;74(6):489–97. 58. Cammarata PR, Chu S, Moor A, Wang Z, Yang SH, Simpkins JW. Subcellular distribution of native estrogen receptor alpha and beta subtypes in cultured human lens epithelial cells. Exp Eye Res 2004;78(4):861–71. 59. Pedram A, Razandi M, Wallace DC, Levin ER. Functional estrogen receptors in the mitochondria of breast cancer cells. Mol Biol Cell 2006;17(5):2125–37. 60. Wu CT, Chang YL, Shih JY, Lee YC. The significance of estrogen receptor beta in 301 surgically treated non-small cell lung cancers. J Thorac Cardiovasc Surg 2005;130(4):979–86. 61. Schwartz AG, Prysak GM, Murphy V, et al. Nuclear estrogen receptor beta in lung cancer: expression and survival differences by sex. Clin Cancer Res 2005;11(20):7280–7. 62. Kawai H, Ishii A, Washiya K, et al. Estrogen receptor alpha and beta are prognostic factors in non-small cell lung cancer. Clin Cancer Res 2005;11(14):5084–9. 63. Skov BG, Fischer BM, Pappot H. Oestrogen receptor beta over expression in males with nonsmall cell lung cancer is associated with better survival. Lung Cancer 2007. 64. Escande A, Pillon A, Servant N, et al. Evaluation of ligand selectivity using reporter cell lines stably expressing estrogen receptor alpha or beta. Biochem Pharmacol 2006;71(10):1459–69. 65. Meyers MJ, Sun J, Carlson KE, Marriner GA, Katzenellenbogen BS, Katzenellenbogen JA. Estrogen receptor-beta potency-selective ligands: structure-activity relationship studies of diarylpropionitriles and their acetylene and polar analogues. J Med Chem 2001; 44(24):4230–51.

210

P.A. Hershberger and J.M. Siegfried

66. Stauffer SR, Coletta CJ, Tedesco R, Nishiguchi G, Carlson K, Sun J, Katzenellenbogen BS, Katzenellenbogen JA. Pyrazole ligands: structure-affinity/activity relationships and estrogen receptor-alpha-selective agonists. J Medicinal Chemistry 2000;43(26):4934–47. 67. Kato S, Endoh H, Masuhiro Y, et al. Activation of the estrogen receptor through phosphorylation by mitogen-activated protein kinase. Science 1995;270(5241):1491–4. 68. Curtis SW, Washburn T, Sewall C, et al. Physiological coupling of growth factor and steroid receptor signaling pathways: estrogen receptor knockout mice lack estrogen-like response to epidermal growth factor. Proc Natl Acad Sci USA 1996;93(22):12626–30. 69. Migliaccio A, Di Domenico M, Castoria G, et al. Steroid receptor regulation of epidermal growth factor signaling through Src in breast and prostate cancer cells: steroid antagonist action. Cancer Res 2005;65(22):10585–93. 70. Razandi M, Pedram A, Park ST, Levin ER. Proximal events in signaling by plasma membrane estrogen receptors. J Biol Chem 2003;278(4):2701–12. 71. Traynor AM, Schiller JH, Stabile LP, et al. Pilot study of gefitinib and fulvestrant in the treatment of post-menopausal women with advanced non-small cell lung cancer. Lung Cancer 2009;64(1):51–9. 72. Giovannini M, Belli C, Villa E, Gregorc V. Estrogen receptor (ER) and epidermal growth factor receptor (EGFR) as targets for dual lung cancer therapy: not just a case? J Thorac Oncol 2008;3(6):684–5. 73. Miller WR, Hawkins RA, Forrest AP. Significance of aromatase activity in human breast cancer. Cancer Res 1982;42(8 Suppl):3365 s-8 s. 74. Bulun SE, Simpson ER. Breast cancer and expression of aromatase in breast adipose tissue. Trends Endocrinol Metab 1994;5(3):113–20. 75. Yue W, Wang J, Savinov A, Brodie A. Effect of aromatase inhibitors on growth of mammary tumors in a nude mouse model. Cancer Res 1995;55(14):3073–7. 76. Buzdar AU, Coombes RC, Goss PE, Winer EP. Summary of aromatase inhibitor clinical trials in postmenopausal women with early breast cancer. Cancer 2008;112(3 Suppl):700–9. 77. Eisen A, Trudeau M, Shelley W, Messersmith H, Pritchard KI. Aromatase inhibitors in adjuvant therapy for hormone receptor positive breast cancer: a systematic review. Cancer Treat Rev 2008;34(2):157–74. 78. Price T, Aitken J, Simpson ER. Relative expression of aromatase cytochrome P450 in human fetal tissues as determined by competitive polymerase chain reaction amplification. J Clin Endocrinol Metab 1992;74(4):879–83. 79. Pezzi V, Mathis JM, Rainey WE, Carr BR. Profiling transcript levels for steroidogenic enzymes in fetal tissues. J Steroid Biochem Mol Biol 2003;87(2–3):181–9. 80. Weinberg OK, Marquez-Garban DC, Fishbein MC, et al. Aromatase inhibitors in human lung cancer therapy. Cancer Res 2005;65(24):11287–91. 81. Mah V, Seligson DB, Li A, et al. Aromatase expression predicts survival in women with earlystage non small cell lung cancer. Cancer Res 2007;67(21):10484–90. 82. Marquez-Garban DC, Chen HW, Goodglick L, Fishbein MC, Pietras RJ. Targeting aromatase and estrogen signaling in human non-small cell lung cancer. Ann N Y Acad Sci 2009;1155: 194–205. 83. Niikawa H, Suzuki T, Miki Y, et al. Intratumoral estrogens and estrogen receptors in human non-small cell lung carcinoma. Clin Cancer Res 2008;14(14):4417–26. 84. Coombes RC, Hall E, Gibson LJ, et al. A randomized trial of exemestane after two to three years of tamoxifen therapy in postmenopausal women with primary breast cancer. N Engl J Med 2004;350(11):1081–92.

Chapter 10

Microparticle Dissemination of Biological Activities: Implications for Cancer Biology Pauline P. Goh

Introduction Extensive reviews in the current literature indicate that tumor growth and metastasis are dependent on reciprocal communication between cancer cells and cells of the tumor microenvironment. However, the recognition that the tumor microenvironment is enriched not only in activated stromal cells but also in tumor and stromal cell-derived microparticles (MPs) with biological function has added a new level of complexity to the tumor stroma. Previously regarded as passive components of the cell environment, accumulating evidence definitively demonstrate that shed MPs from tumor cells and stromal cells engage in cell–cell communication, playing multiple roles in various facets of tumor biology including tumor growth, angiogenesis, and metastasis, as well as cancer-associated thrombosis [1–5]. Subcellular procoagulant particles were first described by Wolf in 1967 [6], who found that minute particles in normal human plasma and serum, which he termed “platelet dust,” were responsible for the platelet-related coagulant activity of whole plasma and serum. In 1971, Crawford [7] extended these observations and concluded that these microparticles were formed by in vivo fracture of membrane buds from extended platelet pseudopods. Many studies have now reported the in vitro release of vesicles from apoptotic or activated cells and their presence in human plasma. The membrane vesicles that cells release are heterogeneous in size and have been designated as exosomes when they constitute smaller vesicles (40–100 mm) originating from the endoplasmic membrane, microparticles (MP), or microvesicles (0.1–1 mm) derived from the plasma membrane and apoptotic bodies (>1.5 mm) containing nuclear material [8–10]. Apoptotic or activated cells release MPs which expose the anionic phospholipid phosphatidylserine (PS) and express membrane antigens that

P.P. Goh (*) Vascular Immunology Unit, Department of Pathology, Sydney Medical School, University of Sydney, Sydney, NSW 2006, Australia e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_10, © Springer Science+Business Media, LLC 2012

211

212

P.P. Goh

reflect their cellular origin [11, 12] The most abundant MPs in the blood arise from platelets, although MPs in the periphery can also arise from lymphocytes, monocytes, and endothelial cells. While MPs are present in peripheral blood of healthy individuals, elevated numbers of circulating procoagulant MPs derived from leukocytes, platelets, and endothelial cells have been described in various pathological states. These conditions include cancer, acute coronary syndromes, antiphospholipid antibody syndrome, sickle cell disease, sepsis, and diabetes [8, 11, 13]. MPs secreted by tumor cells and vascular cells in both solid tumors and hematological malignancies have been reported to circulate at high levels in venous blood of cancer patients, and in some studies, have been shown to correlate with disease activity [14–18]. There is still a paucity of experimental and clinical data relating to the contribution of MPs to various aspects of tumor biology. The available literature, however, has provided some valuable insights into the repertoire of potential roles that circulating or blood-borne MPs can assume. Some tumor MPs expressing FasL appear to participate in immune escape [18, 19]. Furthermore, the high levels of tissue factor (TF)-expressing MPs demonstrated in cancer patients, and their positive correlation with coagulation activation markers, suggest that TF+MP may contribute to hypercoagulability in cancer patients [17, 20–22]. Some researchers have also postulated that elevated levels of circulating tumor – as well as plateletderived MPs which correlate with disease activity – may serve as useful predictors of cancer metastasis [14, 18, 23]. This chapter will firstly provide an overview of how MPs are generated, what stimuli (both in vitro and in vivo) trigger vesiculation, and what are the current technologies used for measuring MPs. Next, the focus will be on how MPs derived from cancer cells or platelets mediate tumor growth, angiogenesis, and metastasis. Finally, the potential role of tissue factor-positive MPs in facilitating cancer-associated thrombosis will be examined.

Cell-Derived Microparticles Mechanism of MP Generation, Release, and Action Resting mammalian cells are characterized by membrane phospholipid asymmetry, with the cytoplasmic leaflet predominantly formed by phosphatidylserine (PS) and phosphatidylethanolamine (PE), whereas the external leaflet is composed of phosphatidylcholine and sphingomyelin [24]. The transbilayer lipid distribution is under the control of three major enzymes: an inward-directed pump, termed flippase with specificity for PS and PE, an outward-directed pump referred as floppase, and a lipid scramblase promoting nonspecific, bidirectional redistribution across the bilayer [25, 26]. A significant and sustained increase of cytosolic Ca2+ accompanying cell stimulation may lead to the loss of phospholipid asymmetry, exposure of PS

10

Microparticles and Cancer Biology

213

on the outer cell surface, and cell membrane vesiculation [25, 26]. Furthermore, since MPs generated from apoptotic endothelial cells (ECs) have been shown to have higher levels of PS on their surface compared with MPs generated from activated ECs, it has been suggested that there are distinct mechanisms for the formation of MPs in apoptotic and activated cells [27]. To date, the exact mechanisms governing MP formation are not completely understood. An earlier study, however, showed that fluvastatin significantly suppressed the generation of MPs from TNFa-stimulated human coronary artery ECs (HCAECs), as well as TNF-a-induced Rho activation. In addition, a Rho-kinase inhibitor was shown to produce a similar suppression of MP release, implicating this kinase in the formation of MPs [28].

Stimuli Triggering MP Generation Data from in vitro studies have shown that MPs are shed from the surface of cells following exposure to a variety of chemical stimuli such as cytokines, thrombin, and endotoxin, or physical stimuli such as shear stress or hypoxia [10]. Depending on the stimuli used to generate them, MPs may differ in their content of exposed membrane anionic phospholipids and therefore their procoagulant potential [29]. In general, microparticle release, occurring minutes after addition of agonist, is dependent on the increase in cytosolic calcium [30]. Platelets are activated by thrombin, calcium ionophore A23187, ADP plus collagen, the terminal complement complex C5b-9, or shear stress [10]. Monocytes, EC, hepatocytes, and arterial smooth muscle cells release MPs upon activation by bacterial lipopolysaccharides (LPS) and cytokines such as tumor necrosis factor-a or interleukin-1b [10]. In an earlier study, the C5b-9 complex was reported to induce vesiculation of HUVEC plasma membrane and express binding sites for factor Va, perhaps contributing to fibrin deposition associated with immune endothelial injury [31]. Similarly, in response to apoptotic signals or cellular activation that may accompany inflammatory or hypercoagulable states present in malignancy, tumor cells and the cells which constitute the tumor microenvironment release MPs. Presently, the different endogenous agonists or stimuli which trigger cell vesiculation in vivo have not been thoroughly investigated. Of significance to cancer biology are observations that many of the stimuli or agonists demonstrated to trigger cell vesiculation in vitro are either hemostatic proteins expressed by tumor cells for activation of the coagulation system or physical stimuli like hypoxia which promote tumor angiogenesis critical for tumor growth and metastasis. Hemostatic factors: Tumor cells can express procoagulant proteins such as tissue factor (TF), the primary initiator of blood coagulation, which result in thrombin generation and ultimately clot formation [32]. Tumor cells also express fibrinolysis proteins like urokinase-type plasminogen (uPA) and tissue-type plasminogen activators and their inhibitors, plasminogen activator inhibitors 1 and 2 (PAI-1 and PAI-2) [32, 33]. Increased production of PAI-1 resulting in a defect in the generation of

214

P.P. Goh

normal plasma fibrinolytic activity may represent another mechanism for the development of venous thromboembolism (VTE) in patients with solid tumors [34]. In an in vitro study, PAI-1 has been shown to rapidly and dose-dependently promote formation of procoagulant endothelial MP (EMP) from EC. Interestingly, PAI-1 knockout mice showed a significantly decreased numbers of EMP than wildtype mice. Moreover, PAI-1-deficient mice responded to an infusion of PAI-1 with a more pronounced rise in the number of MP, and the number of MP stained with Annexin V, evidence for the expression of anionic phospholipid. Increased procoagulant potential was indicated by accelerated thrombin formation by ECs [35]. Platelet activation factors: By the release of soluble factors, such as adenosine diphosphate (ADP), or by thrombin generation via activation of the coagulation cascade, tumor cells also induce platelet activation and aggregation [36, 37]. Inflammatory cytokines: Tumor cells synthesize and release a variety of pro-inflammatory cytokines like TNF-a and IL-1b. In response to the tumor insult, normal inflammatory host tissue also engages in cytokine overproduction [32, 37]. In addition to inducing TF expression in tumor-associated macrophages and ECs, TNF-a and IL-1b also stimulate ECs to increase the production of the fibrinolytic inhibitor PAI-1 [32]. Hypoxia: Clinical investigations carried out over the last two decades have clearly demonstrated that the prevalence of hypoxic tissue is a characteristic pathophysiological property of locally advanced solid tumors, promoting tumor progression and resistance to therapy [38]. It is now well established that hypoxia is a key regulator of tumor angiogenesis, a process crucial to tumor growth and metastasis. In response to hypoxia, cancer cells and normal cells have been reported to secrete vascular endothelial growth factor (VEGF), a key angiogenic factor [39]. Recently, Wysoczynski and Ratjczak [40] reported that human and murine lung cancer cells treated with non-apoptotic doses of either hypoxia (1% O2) or g-irradiation (1,000 cGy) secreted up to 4 times more MPs compared to controls. These tumor-derived MPs (TMPs) activated and chemoattracted stroma fibroblasts and ECs. Significantly, these TMPs also induce expression of angiopoietic factors like interleukin-8 (IL-8), VEGF, and matrix metalloproteinase-9 (MMP-9) in these cells. Other studies have highlighted the ability of chemotherapy and procedures adopted for blood storage and processing to induce cell vesiculation. Since chemotherapy and blood transfusion form part of the treatment modalities for cancer patients, these in vitro data, though preliminary, merit our attention. Effect of cisplatin on cell vesiculation: Cisplatin-based chemotherapy has been reported to predispose cancer patients to thromboembolic events [41]. A recent study demonstrated that cisplatin dose- and time-dependently increased the formation of procoagulant EMPs from human umbilical vein endothelial cells (HUVECs) and human pulmonary microvesicular ECs. Blocking of phospholipids by Annexin V markedly diminished EMP-associated procoagulant activity which is TF independent [42]. Effect of blood product storage and processing on cell vesiculation: Of significance to hematologists/oncologists are preliminary results reported by Gelderman’s lab [43]

10

Microparticles and Cancer Biology

215

which indicated that apheresis platelet units (APU) analyzed on day 6 of the platelet storage period contained significantly elevated counts of MPs derived from platelets, WBC, RBC, and ECs when compared to plasma from healthy volunteers. Importantly, overnight incubation of APU MPs with cultured ECs induced a significant increase in cell apoptosis and endothelial expression of CD154 (ICAM-1) and CD142 (TF), markers of inflammation and coagulation, respectively. The potential impact of collection, processing, and storage steps on MP generation in blood and blood products has been reviewed by Simak and Gelderman [44].

Measurement of MP Microplate affinity assays and flow cytometry: In general, most investigators use either microplate affinity assays or flow cytometry assays for MP analysis [45]. Both types of assays depend on the antigenic composition of MPs and allow them to be enumerated according to their cellular origin. There are two types of microplate affinity assays. One frequently used assay, the enzyme-linked immunosorbent assay (ELISA) can be designed using antibodies to MP antigens for capture, detection, or both, whereas the other assays use Annexin V in the presence of Ca2+ to capture or detect PS+MP [44]. Conventional flow cytometry (FACS) is the most commonly used method for analyzing the number, size, and properties of MPs. By definition, MPs are small structures (0.1–1 mm) which display characteristic forward and side scatter patterns. As in the case of FACS analysis of cells, antibodies to cell surface molecules allow the identification of specific MP subpopulations. While ELISAs can assess the total amount of MP-related material in a specimen, they cannot provide information on the number or size of MPs. A critical review of the technical aspects of MP assays has been published recently [46]. MP analysis can be impacted not only by the type of assay but also by the manner in which blood is collected and processed, including sampling site, needle diameter, centrifugation, and freeze-thaw methods [44]. At present, MP analysis is constrained by the lack of standardized MP assays, a challenge now being addressed by the Working Group on Vascular Biology of the Scientific and Standardization Committee of the International Society on Thrombosis and Hemostasis. Proteomic analysis of proteins in MPs: Because of their critical pathophysiological role, the characterization of which cellular proteins are represented in MP is pivotal to the understanding of their function. Proteomics has been regarded by many researchers as the technology of choice for this kind of research, and some investigations into the microparticle proteome have appeared in the literature [47]. The current view is that protein-based microarray technologies using 2-dimensional electrophoresis (2-DE) and mass spectrometry represent an advancement in highthroughput analysis compared to standard ELISA techniques.

216

P.P. Goh

Role of Cell-Derived MPS in Tumor Growth, Angiogenesis, and Metastasis Tumor Progression and Metastasis Cancer metastasis, a hallmark of cancer, is a highly coordinated and dynamic multistep process and is recognized as the product of an evolving and extensive cross talk between genetically abnormal cells and stromal cells within the tumor microenvironment [48]. Briefly, the metastatic cascade begins with tumor invasion enabling cancer cells to break free from the primary tumor, intravasation into the blood circulation, survival of cells in the circulation, arrest in a new organ, extravasation into the surrounding tissue, initiation and maintenance of growth in metastatic foci, and vascularization of the metastatic tumor [49]. A great deal of experimental evidence has provided support for the observation that cancer cells can alter their adjacent stroma to form a permissive and supportive environment for tumor progression by producing a range of stroma-modulating growth factors and proteolytic enzymes [50]. These stromal modulators disrupt normal tissue homeostasis and act in a paracrine manner to induce angiogenesis and inflammation, as well as activate stromal fibroblasts and other stromal cells in the tumor microenvironment. Thus, the reactive tumor stroma is characterized by the presence of proliferating ECs, resulting in a high number of leaky tumor vessels, increased numbers of inflammatory cells, fragments of extracellular matrix (ECM) molecules, active proteases, and activated fibroblasts called myofibroblasts [50]. It is now widely appreciated that cancer cells constitutively shed membrane vesicles into the extracellular environment both in vitro and in vivo [51]. By actively shedding MPs which can modify the phenotype as well as influence diverse biological functions of a range of target cells that constitute the tumor stroma, cancer cells have developed an additional strategy to support its growth and progression. In a reciprocal fashion, MPs derived from activated stromal cells are able to modify the phenotype of tumor cells, increasing their invasiveness and metastatic potential.

Tumor Cell-Derived MPs (TMPs) and Tumor Growth, Angiogenesis, and Metastasis Of significance to tumor biology is the demonstration that tumor cell lines with highly metastatic potential release a greater amount of microparticles (TMPs) than cells with low metastatic ability [52]. Phenotypic characterization of TMPs has indicated that several, but not all, determinants expressed by tumor cells are present on TMPs. Furthermore, their level of expression on TMPs does not always correlate with that on tumor cells [53]. As summarized in Fig. 10.1 and reviewed in the subsequent sections, TMPs assume many roles in tumor biology. Since TMPs are rich in matrix-degrading proteinase activities, they may play critical roles in ECM digestion, facilitating invasion

10

Microparticles and Cancer Biology

Trafficking to Metastatic foci CCL20

217

ECM degradation

CXCL12

CXCR4 MMPs, TIMP1, uPA,MT-1 MMP

CCR6 TMP TMP

Primary Tumor Cell TMP

TMP

β1 integrin, VEGF, HGF, angiogenin,EGFR IL-6, IL-8, MMPS, CD147

FasL, TGF-β, HLA 1

TMP-Induced Angiogenesis

Immune Escape Fig. 10.1 Schematic diagram of some phenotypes of tumor cell–derived microparticles implicated in tumor growth, angiogenesis, and metastasis

of tumor cells during metastasis and migration of endothelial cells during angiogenesis. TMPs can also mediate tumor angiogenesis by transferring proangiogenic mRNA and proteins to ECs. Furthermore, their ability to induce apoptosis, or inhibit proliferation of immune cells, forms part of their arsenal to escape immune surveillance. Finally, signaling through chemokine receptors acquired from TMPs also enhances chemotactic and invasive responses of target cells, with ultimate homing to specific metastatic niches.

TMPs and Tumor Invasion and Metastasis In many pathological situations, cells cause excessive degradation of their own matrix. One of the most striking examples of unbridled attack upon the ECM is seen in the process of tumor invasion and metastasis, where cells of the primary tumor acquire an invasive phenotype. Invasion of the ECM by primary tumor cells during metastasis requires the concerted effort of ECM-degrading proteinases produced by many tumors with a migratory and invasive phenotype [54]. Research data accumulated over the last decade have shown that TMPs released from cultured cancer cell lines are enriched in proteolytic enzymes like matrix metalloproteinases (MMPs) which are free or complexed with their tissue inhibitors (TIMPs), zymogen pro-MMPs, membrane-type MMPs (MT-MMP), and urokinase-type

218

P.P. Goh

plasminogen activator (uPA) [55–60]. MMPs degrade basement membrane collagens, whereas uPA catalyzes the conversion of plasminogen to plasmin, a broad-spectrum serine protease that degrades numerous components of the ECM including fibrin, laminin, fibronectin, and vitronectin [57]. That cellular membrane vesicle shedding may represent a possible pathway of MMP enzyme release was suggested by early electron microscopic observations of invasive carcinoma tissue that showed destruction of connective stroma by membrane vesicles close to the invading epithelial cells [61]. Phenotyping of TMPs shed by human HT1080 fibrosarcoma cells indicate that they are rich in zymogen (pro-MMP-2 and pro-MMP-9) and active forms of MMP-2 and MMP-9, as well as MMP-9–TIMP 1 and uPA–uPA receptor complexes [56]. Interestingly, the TMPs also exhibited a strong plasminogen-dependent fibrinolytic activity due to TMP-associated uPA. Furthermore, addition of exogenous plasminogen resulted in activation of TMP-associated pro-MMPs to active MMPs, indicating a role for the urokinase–plasmin system in MMP-2 and MMP-9 activation. Based on these observations, the authors postulated that TMPs may provide a large membrane surface for the activation of membrane-associated proteinases involved in ECM degradation and tissue invasion [56]. In ovarian cancer, malignant ovarian ascites-derived membrane vesicles from stages III and IV ascites also contain activated MMP-2, MMP-9, and uPA and are potent inducers of proteinase-dependent invasion in cultured malignant ovarian epithelium [57]. Furthermore, TMPs shed by 8701-BC breast carcinoma cells were found to be densely packed with the b1 integrin subunit [55]. Since integrins are the major class of receptors that regulate cell adhesion to ECM macromolecules [62, 63], the researchers hypothesized that compacting of integrins on the membranes of shed TMPs may facilitate the binding of these TMPs to ECM components, thus permitting proteolytic enzymes to act focally on their substrates [55].

TMPs and Tumor-Induced Angiogenesis During adulthood, most blood vessels remain quiescent, and angiogenesis or the formation of new blood vessels from preexisting ones occurs only under tight regulation in the cycling ovary, in the placenta during pregnancy, and during wound healing [64–66]. However, in response to a hypoxic tumor microenvironment, the balance between angiogenesis stimulators and inhibitors is tilted, resulting in an angiogenic switch with resultant dysregulated tumor neovascularization [67]. Pioneering work by Folkman in the early 1970s showing that tumors cannot grow beyond a critical size or metastasize to another organ without blood vessels has led to the recognition that tumor growth and metastasis are angiogenesis dependent [68]. Thus, as a primary tumor grows, it needs to develop a blood supply that can support its metabolic needs, as well as provide an escape route by which tumor cells can leave the primary tumor mass and enter the body’s circulatory blood system to metastasize to secondary sites. Furthermore, once the metastatic cell has reached its target organ, angiogenesis is again required for the metastases to grow [69–71].

10

Microparticles and Cancer Biology

219

Angiogenesis consists of several processes like vascular basement membrane degradation, endothelial cell migration and proliferation, capillary tube formation, and finally, differentiation into mature vessels. All these processes are mediated by angiogenic growth factors and their receptors, proteolytic enzymes, as well as by cell adhesion molecules [72, 73]. As depicted in Fig. 10.1, TMPs from many cultured tumor cells with the ability to induce an invasive and angiogenic phenotype in stromal cells express proangiogenic molecules like VEGF, angiogenin, interleukin-6 (IL-6), interleukin-8 (IL-8), hepatocyte growth factor (HGF), MMP-2, MMP-9, membrane type 1 MMP (MT1-MMP), CD147/extracellular matrix metalloproteinase inducer (CD 147/EMMPRIN), epidermal growth factor receptor (EGFR), or tissue factor (TF). Human ovarian carcinoma cell-derived TMPs expressing MMP-2, MMP-9, and VEGF have been reported to stimulate the motility, invasiveness, and proliferation of HUVEC in vitro [59]. Glioblastoma tumor cell–derived TMPs, with the ability to stimulate tubule formation and initiate angiogenesis in brain ECs, are enriched in the angiogenic proteins VEGF, angiogenin, IL-6, and IL-8 [60], all of which have been implicated in glioma angiogenesis and increased malignancy [74–76]. Additionally, IL-8 has been shown to directly enhance HUVEC proliferation, survival, and MMP-2 and MMP-9 expression in CXCR1- and CXCR2-expressing EC, and to regulate angiogenesis [77]. Furthermore, these glioma TMPs stimulated proliferation of a human glioma cell line, perhaps via IL-6-mediated glial tumor cell growth by an auto/paracrine mechanism. In gliomas, IL-6 has been reported to promote tumor growth [78], with the level of IL-6 gene expression shown to increase with the grade of malignancy [76]. Angiogenesis shares many functional similarities with tumor invasion, including the requirement for expression of the invasive phenotype and associated proteolytic activity. Hence, many studies have provided compelling evidence for the involvement of MMPs in tumor-induced angiogenesis, with several MMPs exerting both positive and negative effects. MMPs act as proangiogenic factors by degrading the ECM, and providing a pathway for invading ECs , and by releasing ECM-bound growth factors and cytokines [54]. Thus, MMP-9, through its release of ECMassociated proangiogenic factors like VEGF, has been identified as a major contributor and a crucial factor triggering activation of quiescent vasculature during the tumor-induced angiogenic switch [79]. Although the proangiogenic role of MMP-9 in tumor angiogenesis has been definitively established, it became evident that this MMP as well as other MMPs could also inhibit angiogenesis. This contrasting function of MMPs is based on the ability of MMPs to generate endogenous molecules with strong antiangiogenic activity derived from either ECM proteins (e.g., tumstatin, endostatin) or non-matrix-derived extracellular molecules (e.g., angiostatin from plasminogen). Based on these considerations, MMP inhibition may result in the stimulation, rather than inhibition, of angiogenesis [80]. In addition to MMPs, MT1-MMP was detected on malignant ovarian ascitesderived membrane particles [57]. Overexpression of MT1-MMP in melanoma cells has been observed to increase tumor vascularization and tumor growth [81]. Furthermore, MT1-MMP has been reported to contribute to tumor angiogenesis via direct stimulation of VEGF secretion [81, 82].

220

P.P. Goh

Recent studies have demonstrated that TMPs may induce proangiogenic activities in stromal cells by a CD147-mediated mechanism [53, 83–85]. CD 147/EMMPRIN or basigin, a member of the immunoglobulin superfamily, is a plasma membrane glycoprotein enriched on the surface of many malignant tumor cells [86, 87]. Cumulative data from several studies have demonstrated that CD147/EMMPRIN could play a crucial role in tumor angiogenesis by regulating expression of VEGF and MMPs in stromal cells [86, 88, 89]. Thus, some researchers have shown that CD147/EMMPRIN can interact with fibroblasts and stimulate their production of MMP-1, MMP-2, MMP-3, and membrane type 1 MMP (MT1-MMP) [90, 91]. Furthermore, CD147 is also known to upregulate the production of MMPs in both tumor and endothelial cells [92]. Analogous studies have provided evidence that microvesicular release of CD147/ EMMPRIN from tumor cells may also play a role in tumor–stromal interactions through upregulation of fibroblast or endothelial cell MMP production. Thus, CD147/EMMPRIN released from the surface of NCI-H460 lung carcinoma cells via microparticle shedding was reported to stimulate MMP-1 transcription in fibroblasts and hence facilitate tumor invasion and metastasis [83]. Similarly, CD147/ EMMPRIN-positive TMPs obtained from PC3, a prostate carcinoma cell line with high metastatic potential, were shown to induce the activation of fibroblasts. Activated fibroblasts displayed extracellular signal-regulated kinase 1/2 (ERK 1/2) phosphorylation, MMP-9 upregulation, increased motility, and resistance to apoptosis. Subsequently, these activated fibroblasts, in turn, shed CX3CL1/fractalkinepositive ligand microparticles that were able to promote chemotaxis of highly metastatic PC3 cells, dependent partially on expression of the chemokine receptor CX3CR1 by PC3 cells [93]. Additional evidence presented by Millimaggi’s team [84] also implicated CD147/EMMPRIN-positive TMP shed by ovarian cancer cell lines in mediating in vitro angiogenesis, attributed in part to transcriptional upregulation of endothelial cell MMPs. Importantly, there was a direct correlation between CD147/EMMPRIN expression on TMPs and the ability of these TMPs to enhance EC invasiveness. To validate their initial observations, these researchers prepared a CD147 transfectant using the noninvasive CABA 1 human ovarian cell line that had nondetectable CD147 expression. As expected, these CD147-positive TMPs shed by the CD147 transfectant exhibited a greater capacity to induce both migration of HUVECs and the transcription of MT1-MMP, MMP-1, and MMP-2 mRNA compared with microparticles generated from vector-transfected cells. Moreover, TMPs derived from OVCAR3 cells with silenced CD147 expression were less effective at inducing migration and upregulating MT1-MMP, MMP-1, and MMP-2 mRNA in HUVECs compared with untreated control cell-derived TMPs [84]. Because stromal cells are the major source of MMPs in most of the human cancers analyzed to date, CD147/EMMPRIN is deemed to be a critical determinant of cancer growth and dissemination [88, 92, 94, 95]. The functional repertoire of TMPs has recently been expanded with exciting new in vitro and in vivo data highlighting the pivotal role that microvesicular transfer of cell surface receptors like TF and EGFR may play in tumor-associated angiogenesis. A recent study by Yu’s lab [96] demonstrated that transfer of TF from TMPs

10

Microparticles and Cancer Biology

221

A431 cancer cell

EGFR TMP

EGFR

HUVEC

Activates MAPK & Aktpathways Triggers endogenous expression of VEGF Stimulates VEGFR2

TMP-Induced angiogenesis

Transfers intact & functional EGFR mediated by PS

Fig. 10.2 Microvesicular transfer of oncogenic receptor EGFR from A431-derived microparticles mediates induction of the angiogenic phenotype in endothelial cells

derived from A431 human squamous carcinoma cells to TF-negative mouse brain EC in vitro initiated angiogenesis. Moreover, host blood vessels in A431 xenografts in SCID mice were positive for both human TF and EC markers (CD105 or endoglin), suggesting that the intercellular transfer of TF also occurred in vivo. TF is frequently overexpressed in human tumors [97]. Although implicated in various aspects of tumorigenesis, the precise mechanisms of TF involvement in tumor growth, angiogenesis, and metastasis still require additional fundamental studies [98, 99]. Based on insights gained from a series of elegant studies, Al-Nedawi and coworkers [100] hypothesized that one source of EGFR expression by tumor-associated ECs may be through the transfer of TMPs from adjacent cancer cells harboring this oncogenic receptor. These researchers demonstrated that this form of EGFR acquisition triggered activation of the autocrine VEGF/VEGFR-2 pathway in a manner dependent on phosphatidylserine (PS)-mediated microparticle exchange between tumor and ECs. As depicted in Fig. 10.2, uptake of intact and functional EGFR from TMPs derived from A431 human squamous cell carcinoma triggered phosphorylation of MAPK (Erk1/2) in ECs. This was accompanied by the onset of endothelial VEGF expression, followed by autocrine activation of its key signaling receptor VEGFR2. These effects were obliterated by preincubation of TMPs with PS blocking agents Annexin V and Diannexin or with an irreversible EGFR inhibitor CI-1033. These findings were corroborated by an in vivo study where A431 human tumor xenografts in mice were found to stain positively for EGFR and phospho-EGFR with Diannexin treatment noted to inhibit growth and angiogenesis of A431 tumors. Collectively, these new data concur with earlier studies investigating molecular profiles of tumor-associated endothelium which indicated that one distinguishing feature of tumor EC is their propensity to express the EGF receptor (EGFR) [101]. Moreover, activation of EGFR on ECs of tumor blood vessels appears to play a crucial role in tumor progression inasmuch as pharmacological

222

P.P. Goh

suppression of this signaling cascade in experimental tumors was shown to inhibit the growth of primary lesions and, more importantly, to reduce the frequency of metastasis [102–104]. In a similar context, results obtained using an in vivo transplantation mouse model showed that angiogenesis and more specifically, persistent expression of VEGFR2 are prerequisites for tumor invasion and malignancy [50]. This finding has been substantiated by other data indicating that intraperitoneal injection of an anti-VEGFR2 antibody (DC-101) into mice after transplantation of both well-differentiated, as well as highly malignant, metastasizing keratinocytes resulted in inhibition of both angiogenesis and tumor invasion. Also noteworthy from the same study is the observation that VEGFR2 blockade precipitated a phenotypic shift from a highly malignant to a premalignant, noninvasive tumor phenotype [105, 106].

TMPs and Immune Escape Patients with advanced malignancies have progressively impaired immune responses, indicating that tumor cells have developed various mechanisms to subvert the immune system [107]. Upregulation of Fas ligand (FasL/CD95L) expression on various tumors including melanoma, lymphoma, esophageal carcinoma, gastric carcinoma, colon carcinoma, and breast carcinoma appears to represent one such mechanism [108]. FasL is a transmembrane protein belonging to the tumor necrosis factor (TNF) superfamily that can trigger apoptotic cell death following ligation to its receptor Fas (CD95/APO-1) [109]. FasL confers immune privilege on tumors by inducing apoptosis of infiltrating lymphocytes that express Fas [110]. Accumulating evidence suggests that, in addition to local defense, FasL overexpression may also play an important role in tumor progression and in the establishment of tumor metastases. Hence, FasL expression was found to be higher in metastatic tumors than in primary ones. In breast and cervical tumors, high FasL expression was significantly associated with lymph node metastases [111, 112], whereas stronger FasL expression was found in liver metastases of colon cancer relative to the primary tumor [113]. Previous studies have documented that alterations in expression and function of the TcR2-associated signal transduction z chain may be responsible for deficient immune responsiveness of T cells in patients with cancer [114]. Significantly, tumor cell expression of FasL has been found to correlate with loss of CD3 z chain expression by tumor infiltrating lymphocytes (TILs) [110]. It is well documented that the CD3 z chain, by transducing activation signals from the antigen-binding T cell receptor to the T cell nucleus, is crucial for T cell receptor signaling. Therefore, FasL expressed on tumors may contribute to the functional abnormalities frequently found in lymphocytes isolated from the tumor microenvironment. Like the parent tumor cells from which they were derived, FasL-expressing TMPs shed by human colorectal and melanoma cancer cells [115, 116], or isolated from sera of patients with human colorectal or oral squamous cancer cells [19, 115], were capable of inducing apoptosis of T lymphocytes. Furthermore, ovarian cancer

10

Microparticles and Cancer Biology

223

cell-derived FasL-positive TMPs also exhibited dose-dependent suppression of T cell receptor/CD3 z expression [117]. These researchers also reported a strong correlation between the decrease in z expression and the level of FasL expression in TMPs. In this instance, the suppression of CD3 z by TMPs appeared to be linked to the induction of apoptosis and caspase-3 within T cells [117]. Taken together, these results showed that, in addition to apoptosis of TILs, FasL expressed by TMPs may also contribute to the functional abnormalities seen in patients with advanced malignancies. Another mechanism which tumors may develop to evade immune surveillance appears to be the inhibition of lymphocyte proliferation by shed TMPs. The observation that anti-TGF-b antibodies could completely neutralize the inhibition of lymphocyte proliferation by TMPs from human breast carcinoma cells led Dolo’s group [118] to conclude that TGF-b+ TMP may play a role in the escape of breast carcinoma cell from immune response. Their results concur with earlier experimental data highlighting the role of TGF-b in suppression of T cell-mediated antitumor immunity [119]. Many tumors including breast cancer, colorectal cancer, hepatocellular carcinoma, and malignant melanoma overexpress TGF-b [120–123]. Compelling evidence indicate that TGF-b does not inhibit tumor growth but dampens the immune response toward the tumor via its potent effects on tumor infiltrating immune cells. Hence, T cell-specific blockade of TGF-b signaling has been shown to induce an enhanced antitumor immune response, with consequent eradication of tumors in mice challenged with live tumor cells [124]. Similarly, treatment with anti-TGF-b antibodies has been reported to suppress the development of primary tumors and metastasis in a human xenotransplant tumor model in nude mice [125]. Overall, these studies suggest that TGF-b has potent effects on the antitumor immune response by suppressing the effector response of tumor infiltrating immune cells. Vesicle shedding is now recognized as a mechanism for tumor cells to remove antigens that could elicit an immune response against them [126]. Indeed, the observation that breast cancer- and colorectal cancer-derived TMPs carry both tumorassociated antigens and HLA class I molecules [55, 115] indicates that these structures could, in principle, present antigens to the immune system [118], providing another mechanism for tumor cells to escape from immune surveillance. Supporting this concept are immunohistological and flow cytometric data featuring total or selective losses of HLA class I antigens in different human tumor samples [127].

TMPs and Trafficking to Metastatic Foci Metastasis represents a highly organized, nonrandom, and organ-selective process [128], sharing many similarities with leukocyte trafficking, which is critically regulated by chemokines and their receptors [129]. Chemokines are a superfamily of small, cytokine-like proteins that induce, through their interaction with G-proteincoupled receptors, directional cell migration and homing to specific anatomical sites [130]. Tumor cells from both solid cancers and hematological malignancies have been reported to express a distinct, nonrandom pattern of functionally active

224

P.P. Goh

Main Metastatic foci

Regional lymph nodes, Bone lungs, liver, & bonemarrow marrow

Skin

Liver

Bone marrow

Chemokine

CXCL12

CXCL12

CCL21

CXCL12

CCL21 CCL27,28

CCL20

CX3CL1

Chemokine receptor

CXCR4

CXCR4

CCR7

CXCR4

CCR7

CCL21

CX3CR1

Type of Cancer

AML

Breast

Melanoma

CRC

Prostate

Reference

[16]

[131]

[131]

[132] & [134]

[133]

CCR10

Fig. 10.3 Selected chemokines expressed by different cancer cells and their corresponding chemokine ligands in preferred metastatic foci CRC (colorectal cancers)

chemokine receptors [130]. Signaling through these chemokine receptors induced chemotactic and invasive responses, with ultimate homing to specific metastatic niches. Significantly, organ-specific metastasis was governed, in part, by interactions between chemokine receptors on cancer cells and matching chemokines in target organs. As summarized in Fig. 10.3, this observation has been demonstrated in numerous studies implicating various chemokine receptors, namely, CXCR4, CCR6, and CCR7, and their corresponding ligands, in the regulation of metastasis in malignant tumors [16, 131–134]. Previous data presented by Muller and coworkers [131] clearly showed that signaling through CXCR4 or CCR7 mediated actin polymerization, pseudopodia formation, and subsequently, chemotactic and invasive responses in breast cancer cells expressing high levels of functionally active CXCR4 and CCR7. Furthermore, neutralizing the interactions of CXCL12/CXCR4 in severe combined immunodeficient (SCID) mice injected with CXCR4-positive, human breast carcinoma cells significantly impaired metastasis of breast cancer cells to CXCL12/SDF-1-rich regional lymph nodes and lungs, preferential sites for the metastasis of breast cancer tumors. Similar to breast cancer metastasis, CXCL12 found in the ascitic fluid of patients with ovarian cancer may have a potential role in the spread of CXCR4-positive ovarian cancer cells to the peritoneum [135]. Other studies implicating the chemokine–chemokine receptor axis in metastasis have shown that CCR6 is significantly upregulated in primary colorectal cancers (CRC) and colorectal liver metastases (CRLM), with peak levels of its related ligand CCL20 in the liver attracting and arresting these CCR6-positive migrating CRC cells [132, 134]. In hematological malignancies, functional CXCR4 expressed by acute myeloid leukemia (AML) cells has been shown to induce leukemia cell chemotaxis and migration beneath marrow stromal cells [136]. Interestingly, researchers from Lapidot’s lab [16] detected increased CXCR4+MPs in the peripheral blood and bone marrow plasma samples of newly diagnosed adult AML patients. These CXCR4+MPs exhibited a strong correlation with WBC counts, a commonly used prognostic

10

Microparticles and Cancer Biology

225

marker in AML patients. Also, the majority of these CXCR4+MPs were CD45+, suggestive of a leukocyte origin. Moreover, transfer of functionally active CXCR4 from these MPs to AML-derived HL-60 cells elicited enhanced cellular migration to CXCL12/SDF-1 in vitro and increased homing/retention to the CXCL12/SDF-1rich bone marrow of irradiated NOD/SCID/b2m null mice. The CXCR4 antagonist AMD3100 reduced these effects. Based on their results, the authors postulated that the CXCR4–CXCL12 axis may play a significant role in trafficking and tissue dissemination of AML cells in vivo [16]. On co-incubation with CCR6-expressing TMPs derived from pancreatic, colorectal, and lung adenocarcinoma, approximately 20% of normal monocytes were reported to internalize these TMPs, concomitantly acquiring CCR6 and CD44v7/8, two antigenic determinants not expressed by monocytes [137]. Moreover, CCR6 was biologically active only in 6% of CCR6+ monocytes determined in the presence of its chemokine CCL20/MIP-3a. Upregulated expression of CD44v on monocytes cultured in vitro is associated with an increased binding of hyaluronan (HA) [138]. Hence, the transfer of CD44v to monocytes by TMP may enhance their ability to react with HA and other ECM molecules. The overall significance of these findings to tumor biology is presently unclear.

Platelet-Derived MP (PMP) and Tumor Growth, Angiogenesis, and Metastasis Substantial experimental and clinical data have validated the view that platelets may aggravate tumor progression/metastasis and angiogenesis [139–143]. Hence, induction of thrombocytopenia has been observed to reduce experimental metastases, whereas platelet transfusions promote them [143]. Moreover, a high platelet count has been associated with poor survival in cancer patients suffering from malignant mesothelioma, gynecological malignancies, lung, renal, gastric, colorectal, or breast cancers [144]. Since the ability of cancer cells to aggregate platelets has been documented to correlate with their metastatic potential [145], tumor cell-induced platelet aggregation (TCIPA) has assumed increased significance in tumor progression and metastasis. TCIPA, perceived as a critical mechanism enabling tumor cells to escape from immune surveillance, confers a number of advantages to the tumor cell, ensuring its survival in the vasculature, and, ultimately, successful migration to distant sites. For instance, when covered with a coat of platelets, tumor cells acquire the ability to evade the body’s immune system [146]. Furthermore, activated platelets expressing adhesion receptors facilitate the adhesion of tumor cells to the vascular endothelium during hematogenous metastasis [147]. Like activated platelets, PMPs have been reported to express several platelet– endothelium attachment receptors like glycoprotein IIb/IIIa (CD41), Ib, and Ia IIa on their surface [148, 149],as well as harbor bioactive lipids like sphingosine-1phosphate (S1P) and arachidonic acid (AA) [150]. In two similar studies investigating PMP–tumor cell interactions (Fig. 10.4), PMPs derived from normal platelet concentrates were shown to transfer platelet-derived integrin CD41 to human lung

226

P.P. Goh

Transfer of intact & functional surface receptors CD 41 & CXCR4 CXCR4 PMP

CD 62P CD 41

M

Lung or Breast cancer cell

Increased production of MMPs Stimulation of cyclin D2 and MT1-MMP mRNA Activation of MAPK p42/44 & Akt signalling pathways Upregulation of VEGF, IL-8, HGF gene expression

M =mucin

PMP-induced invasive, migratory & angiogenic phenotype

Fig. 10.4 Schematic representation of the induction of the invasive, migratory, and angiogenic phenotype in lung or breast cancer cells by platelet-derived microparticles (PMPs)

and breast cancer cells, thus enhancing their adhesion to endothelial cells. Furthermore, PMPs also stimulated the production of MMPs [151, 152]. The authors postulated that increased MMP activity and greater adhesiveness of these cancer cells could potentially promote transendothelial migration of cancer cells into the tissue during hematogenous metastasis. Furthermore, PMPs could act as signaling molecules, interacting directly with both breast and lung cancer cells via plateletderived P-selectin/CD62P and mucins on target cancer cells to increase cell proliferation by promoting activation of MAPK p42/44 and AKT signaling pathways. More specifically, PMP may also facilitate tumor vascularization via upregulation of proangiogenic VEGF, IL-8, and HGF gene expression in lung cancer cells. Additionally, PMPs may also induce a more migratory and invasive phenotype in lung cancer cells by augmenting their expression of both cyclin D2 and MT1-MMP mRNA [151]. Overexpression of cyclin D2 has been associated with increased invasiveness in vivo and progression of various tumors such as testicular germ tumors and gastric cancer cells [153, 154]. The MT-MMPs are highly expressed in almost all types of human cancers [155, 156], with MT1-MMP localized predominantly to specialized membrane extensions; this localization appears to be essential for cancer cell invasion [156]. PMPs also strongly chemoattracted the highly metastatic human A549 lung cancer cell line and the murine Lewis lung carcinoma (LLC) cells. This phenomenon could be attributed to the presence of potent platelet-derived chemoattractants like S1P and AA in PMPs [150, 157–161]. The role of PMPs in mediating the increase in metastatic potential of murine LLC cells was confirmed by in vivo evidence showing that mice injected with PMP-covered LLC cells developed significantly more metastatic foci and LLC cells in murine lungs and bone marrow cavities, respectively, compared to mice injected with control LLC cells without PMP [151]. Since PMPs have been documented to transfer platelet-derived CXCR4 onto the

10

Microparticles and Cancer Biology

227

surface of target cells, [148, 162, 163], these researchers surmised that LLC cells preincubated with PMPs may have acquired CXCR4 on their surface, enabling them to respond to the CXCL12/SDF-1-rich environment found in the murine lung and bone marrow. Similarly, PMPs were shown to increase CXCR4 expression in invasive breast cancer cells and their chemotaxis toward a CXCL12/SDF-1 gradient [152]. Like prostate cancer cell-derived MPs discussed earlier (Section 2.1), PMPs also induced marrow fibroblasts to secrete MMP-9. These experimental data showing the ability of PMPs to mediate tumor progression via enhancement of the in vitro invasive potential of metastatic lung and breast cancer cell lines warrant further evaluation due to the implications of platelet transfusions in cancer patients.

Tissue Factor-Positive Microparticles and Cancer-Associated Thrombosis Cancer-Associated Thrombosis Thromboembolism is a well-recognized complication in patients with solid tumors or hematological malignancies, contributing significantly to their morbidity and mortality [1, 37, 164–166]. Clinical manifestations vary from venous thromboembolism (VTE) to disseminated intravascular coagulation (DIC) and arterial embolism [164]. Furthermore, the risk of VTE is further accentuated in patients on cytotoxic chemotherapy. In addition, antiangiogenic therapies (e.g., thalidomide) as well as hormonal therapies (e.g., tamoxifen) have also been associated with an increased risk of thrombosis [167–169]. Multimodal therapeutic approaches which combine chemotherapy and an angiogenesis inhibitor are also associated with an increased risk of VTE that may exceed the risk associated with the use of either modality [167, 170]. For instance, the incidence of deep vein thrombosis (DVT) has been reported to be about 1–3% for thalidomide alone and 10–30% when thalidomide is used in combination with dexamethasone [171]. Despite abundant experimental and clinical evidence in this rapidly expanding field of cancer-associated thrombosis, the mechanisms for the production of the hypercoagulable state characteristic of cancer has remained speculative. To date, possible underlying mechanisms postulated to contribute to the development of thrombosis include pro-inflammatory cytokines, circulating cell- or microparticle-bound tissue factor (TF), TF-independent procoagulant activity (including prothrombinase activity) and tumor-associated cysteine proteinase (cancer procoagulant) associated with cells and/or microparticles (MPs), and carcinoma-derived mucins [172].

Tissue Factor in Cancer-Associated Thrombosis Although abnormal coagulation profiles have been found in cancer patients, such abnormalities did not correlate with the development of thrombosis [173].

228

P.P. Goh

Tissue factor (TF), the physiological initiator of blood coagulation, is overexpressed in tumor cells and cells resident in the tumor microenvironment and is thought to significantly contribute to the hypercoagulable state in patients with malignancy [174]. Aberrant TF expression correlating with grade and tumor progression has been reported in association with solid tumors and hematological malignancies [98]. As reviewed by Lopez-Pedrera and coworkers [175], hypercoagulability in patients with hematological malignancies could be attributed to the activation of TF, which may be derived from tumor cells or cells of the tumor microenvironment. In the early 1990s, it was established that monocytes from lymphoma patients are endowed with functional abnormalities with simultaneous expression of TF and antifibrinolytic activity [175]. Recent studies have further demonstrated that DIC in malignant lymphoma may be due to elevated cytokine expression by lymphoma cells stimulating the expression of TF in blood cells or surrounding tissue [175]. Overexpression of TF in AML cells, particularly in leukemic blasts from subtypes M3 and M5, may account for an increased risk of thrombohemorrhagic alterations in these patients [175].

Studies Implicating Tissue Factor-Positive MPs in Cancer-Associated Thrombosis TF is constitutively expressed in the adventitial and medial layers of the vessel wall. This has led to the suggestion that TF is part of a hemostatic envelope surrounding the vessel lumen which triggers blood coagulation only in the case of disrupted vascular integrity [176, 177]. Subsequent findings indicate, however, that preformed TF can also be associated with blood cells and circulating microparticles [178]. Microparticles are considered to have pathophysiological importance in relation with the hemostatic system because like activated cells, chiefly platelets, they expose procoagulant anionic phospholipids such as phosphatidylserine (PS). PS expressed on the microparticle membrane surface can provide the catalytic surface for the assembly and activation of the tenase and prothrombinase complexes [1]. TF antigen and TF activity associated with microparticles in plasma have been measured in patients with various types of cancer. Although measurement of TF activity is considered more relevant, because TF activity more closely reflects the ability of TF to initiate coagulation, many clinical studies have measured TF antigen. TF antigen in clinical studies: Many researchers have evaluated the hypothesis that elevated numbers of TF-bearing MPs may contribute to cancer-associated thrombosis [4, 179, 180]. Furie and coworkers [179] have established that TF-bearing MPs are found in about one third of patients with advanced cancer, including patients with pancreatic carcinoma, ovarian carcinoma, colon cancer, and breast cancer. Whether TF associated with leukocyte (like monocyte) MPs or TF associated with tumor MPs is important in the hypercoagulable state associated with malignant diseases remains unknown [179]. Recently, TF-positive MPs have been demonstrated in the blood of cancer patients diagnosed with thrombotic syndromes [4, 17, 180–184].

10

Microparticles and Cancer Biology

229

In colorectal cancer patients, the amount of plasma TF-positive MPs was significantly correlated with levels of plasma d-Dimer, a very sensitive marker of coagulation activation which strongly predicts venous thromboembolic events in cancer patients [20]. Interestingly, the increase in circulating TF-positive microparticles was attributed to MPs derived from platelets. The positive correlation between TF-positive MPs and d-Dimer levels has led Rauch [165] to the challenging assumption that TF-positive MPs, especially when derived from platelets, may have a role in the hypercoagulopathy of cancer patients. TF procoagulant activity in clinical studies: The above data presented by Hron [20] concur with those of Tesselaar and coworkers [4] who reported that metastatic breast and pancreatic cancer patients, who presented with acute VTE, had higher numbers of circulating MPs (by a factor of 2–4), with greater than 90% of Annexin V+ MP expressing the platelet antigen CD 61. These patients also exhibited higher levels of MP–TF activity compared to healthy subjects, cancer patients without VTE, and subjects with idiopathic VTE. In addition, MP–TF activity correlated with circulating MPs expressing the epithelial antigen MUC1. Metastatic breast and pancreatic cancer patients with elevated MP–TF activity and detectable MUC1+-MP had a lower survival rate at follow-up than those with normal MP–TF activity and absence of MUC1+-MP. Confocal immunofluorescence microscopy studies of the co-expression of MUC1 and platelet antigen CD61 on MP also indicate that a small part of circulating MP seemed to result from fusion of cellular vesicles originating from malignant epithelial cells and platelets in patients with disseminated breast and pancreatic adenocarcinoma. Recently, Langer and coworkers [185] in a small but related study demonstrated significantly increased levels of TF-specific procoagulant activity (PCA) of plasma MPs in five patients presenting with overt disseminated intravascular coagulation (DIC) due to non-small cell lung cancer (n = 1), melanoma (n = 1), prostate cancer (n = 2), and acute promyelocytic leukemia (n = 1). Clotting experiments on available tumor cell samples suggested that cancer cells were a potential source of circulating TF-positive MPs. Significantly, follow-up plasma samples from two surviving patients revealed that the response of their malignancies to specific anticancer therapy was paralleled by resolution of overt DIC and a significant decline in MP-associated TF PCA. Although TF on circulating MPs have been implicated in the induction of a prothrombotic state in cancer patients, MPs have also been demonstrated to support the assembly of blood coagulation protein complexes in the absence of TF. In a recent study, ECs treated with cisplatin in vitro were shown to undergo apoptosis, concomitantly releasing MPs that had TF-independent procoagulant activity that could be inhibited by neutralization of anionic phospholipids [42]. TF in in vivo studies using mouse models: Using laser injury in a mouse model of thrombosis, Falati and coworkers [186] demonstrated that MPs expressing TF and PSGL-1 accumulate in the developing thrombus. Intriguingly, capture of TF+-MPs by activated platelets was mediated via its ligand PSGL-1 and P-selectin expressed on activated platelets in the platelet thrombus.

230

P.P. Goh

Tissue Factor-Positive MPs – Cellular Origin and Procoagulant Activity In studies centered on the pathophysiology of tumor-associated hypercoagulability, two of the most intriguing questions which have fueled intense debate are as follows: (a) What is the cellular origin of TF-positive MPs? (b) Do specific MPs express functionally active TF, that is, are TF-positive MPs procoagulant, able to mediate coagulation, and participate in thrombus formation? Cellular origin of TF-positive MPs: The elevated number of TF-positive MP and the MP-bound TF activity found in colorectal, breast, and pancreatic cancer patients were found to be mainly derived from platelets [4, 20]. As noted by Hron [20], the increased level of platelet-derived TF-positive MPs may stem from TF originally synthesized by other cells and subsequently transferred to activated platelets, which then shed TF-positive MPs into the blood of cancer patients. This possibility is supported by previous studies which indicated that TF-positive MPs obtained from monocytic cells expressing P-selectin ligands (either CD15 or PSGL-1) have the propensity to adhere and bind to P-selectin on activated platelets, concomitantly transferring TF [187, 188]. Thus, TF present on activated platelets might be acquired from other primary cell sources by fusion of MPs with the platelet surface membrane and possibly, be released as TF-positive platelet MPs upon vesiculation in response to further platelet activation. On the other hand, platelets themselves have been demonstrated to store small amounts of TF in a-granules and to release TF-positive MPs, which can increase the blood TF activity [189]. It is now widely appreciated that constitutive microvesicle shedding is a common feature of cancerous cells [51, 190]. Many studies have described shedding of procoagulant microvesicular structures from tumor cells. As early as 1985, membrane vesicles were found in certain types of leukemia, including acute promyelocytic leukemia, acute myelogenous leukemia, and acute monocytic leukemia [190]. These vesicles shared antigens with leukemic cells. Some of these vesicles expressed procoagulant activity, but it was not possible to correlate procoagulant activity of these vesicles with the clinical coagulation disorders in these patients. To date, there are very few clinical studies which focus specifically on tumor cell-derived MP which express TF. Recently, Langer’s lab [185] showed that cancer cells were a potential source of TF-positive MPs. Due to the small number of patients evaluated, the results, albeit significant, need to be confirmed. In a recent case study of lung cancer-associated Trousseau’s syndrome, tumor cells were proposed as the main source of TF-positive MPs [17]. Colorectal carcinoma, gastric carcinoma, melanoma, and squamous cell carcinoma cell lines also shed procoagulant TF-positive MPs, with TF activity shown to correlate with intracellular TF mRNA levels [191]. Other studies have indicated that TF shed from human glioblastoma cells was able to induce coagulation, platelet aggregation, and thrombus formation [192]. Also noteworthy is the report that in vitro-generated TF expressing MPs from human pancreatic cancer cells when injected into an orthotopic murine model of pancreatic cancer were capable of activating coagulation in vivo as shown by the presence of TF-dependent procoagulant activity and thrombin–antithrombin complex in cell-free murine plasma [182].

10

Microparticles and Cancer Biology

231

Fig. 10.5 Modulators of tissue factor procoagulant activity (TF PCA)

Procoagulant activity of TF-positive MPs: It is not yet known whether TF on MPs is constitutively active or needs to be decrypted from an inactive form or location for binding to FVII/FVIIa to initiate coagulation. A few studies support the notion that TF on MPs originally have a rather low procoagulant activity that becomes markedly increased when MPs interact with platelets [187, 188, 193]. In addition, the monocyte-derived MPs bind to activated platelets in an interaction mediated by platelet P-selectin and microparticle P-selectin glycoprotein ligand (PSGL-1) or CD15 (an epitope also found on PSGL-1) [187, 188]. Further support for the involvement of the PSGL-1/P-selectin ligand-receptor mechanism in facilitating binding of TF-positive MPs to activated platelets stems from recent data published by Furie and Furie [194]. Fluorescently labeled mouse MPs derived from WEHI cells, a monocytoid cell line, were infused into a recipient wild type or P-selectinnull mouse. Microparticles localized within the developing thrombus of the wildtype mouse, but not the P-selectin-null mouse, indicating that one pathway for the initiation of blood coagulation in vivo involves the accumulation of tissue factorand PSGL-1-containing microparticles in the platelet-rich thrombus expressing P-selectin [194].

Protein Disulfide Isomerase – Potential Modulator of TF Activity Cryptic and coagulant TF: It is now well documented that TF on unperturbed cells are encrypted and reside in a cryptic configuration requiring activation to initiate coagulation [195]. As shown in Fig 10.5, TF activation may be regulated at multiple levels, including increase in cytosolic calcium, rapid feedback inhibition by TF pathway inhibitor, and exposure of PS on the outer leaflet of the plasma membrane of activated/apoptotic cells and microparticles. Coagulant activity of TF may also be attenuated by TF dimerization and/or compartmentalization in lipid rafts such as caveolae [195]. Activated platelets may also promote TF activity in monocyte

232

P.P. Goh

Fig. 10.6 TF disulfide bond redox status and protein disulfide isomerase (PDI) regulate TF activity

membranes in a process entailing transfer of TF to activated platelets [193]. In addition to cellular surfaces, TF present on monocyte-derived microvesicles has also been shown to require activation that is dependent on fusion of microvesicles with activated platelets [188]. Modulation of TF activity by protein disulfide isomerase: Novel and exciting new data from Ahamed’s lab [196] indicate that the oxidoreductase protein disulfide isomerase (PDI) may have a role in the modulation of TF procoagulant activity. Their findings suggest that active coagulant and inactive encryptic TF may, in fact, be two structural entities, with decryption and subsequent procoagulant activity involving formation of an allosteric surface-accessible, extracellular disulfide bond (Cys186-Cys209) (Fig. 10.6). The redox state of the bond appears to be controlled by the oxidoreductase PDI with TF coagulant activity suppressed during association of cell surface PDI with inactive TF via a mixed disulfide bond. They further showed that PDI suppressed TF coagulant activity in a nitric oxide (NO)-dependent pathway. Cryptic, but not coagulant, TF is S-nitrosylated. S-Nitrosylation of TF resulting from reaction of nitric oxide (NO) with the unpaired TF cysteine thiol therefore appears to play a role in maintaining the cofactor in an encrypted state. Denitrosylation of TF leads to cleavage of the disulfide bond holding the TF–PDI complex together, with formation of the TF allosteric disulfide bond and coagulant TF [196]. Cellular sources of PDI: Although primarily an endoplasmic reticulum resident protein, PDI has been demonstrated on the surface of numerous cell types including endothelial cells and activated platelets [197, 198]. In addition to the archetypal PDI, a novel PDI named endothelial PDI (EndoPDI) because of its high expression in endothelial cells has been identified by Sullivan’s group [199]. Significantly, EndoPDI expression is rare in normal tissues but was present in the endothelium of tumors, and in other hypoxic lesions such as atherosclerotic plaques. In addition, other researchers have shown that upon activation, platelets secrete up to 20% of total platelet PDI. These researches also proposed that upregulation of TF activity in association with activated platelets may depend on secreted PDI [200]. Interestingly, data from Claudio’s lab showed PDI to be the most significantly upregulated gene (MGC3178) in 100% of the myeloma cell lines tested [201].

10

Microparticles and Cancer Biology

233

Fig. 10.7 The multifaceted roles of PDI [208]

Microparticle-associated PDI: Activation of platelets with calcium ionophore A23187 has been shown to induce release of PDI-positive vesicles [200]. Recently, Raturi and coworkers [202] detected increased levels of PDI-containing platelet MPs (PMPs) from patients with Type II diabetes (T2D). These PMP membranes demonstrated 50- to 100-fold higher specific procoagulant activity compared to activated platelets [203]. PDI precursors were also found to be associated with procoagulant MPs released from TNF-activated HUVEC [204]. In a recent study, Versteeg and Ruf [205] showed that purified PDI can enhance TF procoagulant activity on microvesicles shed from HUVECs. TF and PDI in thrombus formation using mouse models: Following laser-induced arteriolar injury, extracellular PDI [206] and both PDI and TF [207] were demonstrated to stimulate fibrin generation and thrombus formation in mice models. As shown in Fig. 10.7, PDI has been reported to catalyze oxidation, reduction, and isomerization of disulfide bonds [208]. The active site of PDI contains a reactive dithiol/disulfide in a CysXXCys motif that has redox potential. PDI-catalyzed disulfide formation occurs when oxidizing equivalents are transferred from the disulfide of the oxidized PDI active site to the reduced substrate (e.g., TF), forming reduced PDI. Conversely, the reduction of a substrate disulfide oxidizes the PDI active site. Disulfide isomerization or rearrangement of incorrect disulfides, which requires PDI in the sulfhydryl or reduced form, does not result in any net oxidation of PDI [208]. As depicted in Fig. 10.8, the reaction that PDI catalyzes (oxidation, reduction, or isomerization) will depend on the equilibrium position of the overall reaction and the redox state of the PDI active site. The reduced active-site cysteines of PDI can react with cellular glutathione disulfide (GSSG) to form oxidized PDI and two molecules of reduced glutathione (GSH) [208].

234

P.P. Goh

Fig. 10.8 PDI and cellular glutathione/glutathione disulfide [208]

Proposed Model for Promotion of Thrombus Development by TF-Expressing MP and Platelet-Derived PDI The current scientific evidence gleaned from in vitro studies using cancer cell lines and in vivo studies using mouse models, as well as clinical data obtained from patients with clinically overt thrombotic syndromes, support a role for TF-positive MPs and activated platelets in promoting TF procoagulant activity, fibrin generation, and thrombus formation. Moreover, binding of TF-MPs to activated platelets was mediated by platelet P-selectin and microparticle PSGL-1. Activated platelets have been shown to enhance the procoagulant activity of TF-positive MP. How activated platelets facilitate this process is presently unknown. The seminal study by Ahamed’s group [196] has provided valuable insights into how PDI, perhaps derived from activated platelets and/or their shed microparticles, may promote TF-mediated blood coagulation, fibrin formation, and thrombus development (Fig. 10.9). In concluding, we would like to propose a model implicating monocyte-derived or tumor cell-derived TF-positive MP and PDI expressed on activated platelets or their shed MPs in cancer-associated thrombosis. Therefore, we postulate that cryptic noncoagulant TF-positive MPs derived from activated leukocytes (like monocytes) or tumor cells are zoom in to activated PDI-positive platelets or PDI-positive platelet MPs in the developing thrombus. Accumulation of TF-MPs in the thrombus is facilitated by P-selectin expressed on activated platelets/platelet MP which capture these TF-positive MPs via its ligand PSGL-1 expressed on leukocyte MP or tumor mucins expressed on tumor cells. P-selectin, an integral membrane protein receptor that binds monocytes and neutrophils, is expressed on activated platelets and activated endothelium. P-selectin mediates adhesion to glycoproteins like mucins with carbohydrate structures containing sialyl-Lewis X [209]. The epithelial mucin MUC1, overexpressed on the cell surface of many epithelial malignancies as well as on some B cell lymphomas and multiple myelomas [210], present ligands for adhesion receptors, such as the selectins, which promote the ability of tumor cells to interact with host platelets and endothelial cells [209]. The work of del Conde [188] has shown that TF present on monocyte-derived microvesicles is poorly coagulant, and fusion of microvesicles with activated platelets is required for TF activity. We also hypothesize that once incorporated into the platelet, platelet-PDI catalyzes the conversion of TF into its procoagulant form which contains the Cys 186–Cys 209 disulfide bond, critical for coagulation.

10

Microparticles and Cancer Biology

P-selectin

235

GSH/GSSG on account of elevated oxidative stress in cancer, platelet PDI probably has oxidase activity

activated platelet

PDI ox P-selectin

2GSH

GSSG

−S

SH

MP

PDI ox S

PDI oxidase SH SH

Monocyte MP

SH

PDI reductase S -S

TF PSGL-1

Cancer Cell MP

TF Mucin

sulfhydryl Cryptic TF

disulfide Procoagulant TF

Fig. 10.9 A hypothetical model depicting potential roles for TF-positive microparticles and platelet-derived PDI in thrombus development

As documented in the existing literature, blood glutathione redox status may modulate PDI function [208]. In studies involving mouse models and cancer patients, the glutathione redox status (glutathione/glutathione disulfide or GSH/ GSSG) has been observed to decrease in blood of Ehrlich ascites tumor-bearing mice, as well as in patients with breast or colon cancers [211]. This effect is mainly due to an increase in GSSG levels attributed to an increase in peroxide production by the tumor, resulting in GSH oxidation. On account of elevated oxidative stress in cancer, culminating in a reduced GSH/GSSG ratio [211], we surmise that plateletPDI would probably have oxidase activity and be capable of catalyzing the formation of procoagulant TF. The latter is characterized by the presence of the disulfide (Cys 186–Cys 209) bond. Even though blood coagulation is initiated by TF, PS on activated platelets and platelet MPs may propagate the coagulation process by providing a surface for assembly and activation of the prothrombinase and/or tenase complexes in TF-mediated fibrin generation and thrombus development.

Conclusion and Future Perspectives The present review of the available literature, albeit not comprehensive, has lend credence to the view that cancer cells as well as their shed microparticles can modify the adjacent matrix and stroma cells to create a permissive and supportive

236

P.P. Goh

environment for tumor growth, angiogenesis, invasion, and metastasis. Armed with proteolytic MMPs and uPA, TMPs degrade the ECM, as well as induce the invasive phenotype in stromal cells. By stimulating MMP expression in fibroblasts, CD147/ EMMPRIN-positive TMPs mediate a novel form of tumor-fibroblast cross talk enabling cancer cells to harness the support of fibroblasts, thus facilitating tumor invasion and metastasis. Although it is clear that CD147/EMMPRIN can influence MMP production in stromal fibroblasts, many aspects of the phenomenon, for example, the identity of the CD147/EMMPRIN receptor on fibroblasts, remain to be elucidated. The expression of the b1 integrin subunit and proteolytic MMPs on membrane MPs have led some researchers to postulate that they may promote TMP adhesion to matrix components and local degradation of ECM, respectively. Additional investigations are needed, however, to establish which integrin dimers are clustered on membrane microvesicles and also identify their possible interaction with MMP-9 [55]. Over the last 10 years, accumulating evidence has established TMPs as carriers of proangiogenic factors, with the propensity to induce the invasive and angiogenic phenotype in endothelial cells. In two independent studies, microvesicular transfer of surface receptors TF or oncogenic receptor EGFR from A431 human squamous carcinoma cell-derived MPs has been reported to induce the angiogenic phenotype in endothelial cells. Of relevance are novel findings presented by [212] which implied that oncogenic changes may impact the level of tumor cell-derived TF and thereby affect cancer coagulopathy, angiogenesis, and other vascular effects associated with cancer. Supporting this notion is the observation that TF upregulation in cancer cells parallels their expression of several mutant oncogenes, including K-ras, EGFR, EGFRvIII, and HER2 [213]. To escape from immune surveillance and engage in successful hematogeneous metastasis, tumor cells have developed different mechanisms like shedding of FasL, HLA class I antigens, or TGF-b-positive TMPs to counter the host immune response. To date, the functional repertoire of MPs also includes microvesicular transfer of functional CXCR4 from MPs isolated from AML patient plasma or PMPs from activated normal platelets to cancer cells, enhancing their migration and homing to CXCL12/SDF-1-rich niches in vitro and in vivo. Furthermore, the preliminary findings that PMPs have the ability to enhance the invasive and metastatic potential of breast cancer cells warrant further clinical evaluation, since platelet transfusions form part of the treatment modalities undertaken by many cancer patients. Thrombosis is one of the major complications of malignant disease, but the underlying molecular and cellular basis remains elusive. Exploration of a potential role of MPs in cancer-associated thrombosis showed that increased numbers and thrombogenic activity of TF-expressing MPs are present in a spectrum of cancer patients known to have a high incidence of thromboembolic complications. Furthermore, the positive correlation of these TF-expressing MPs with coagulation activation markers like d-Dimer,indicate an important role for TF-expressing MPs in the pathogenesis of cancer-associated thrombosis. Before TF-positive MP can be considered as a biomarker for VTE, methodological issues regarding isolation, quantification, and antigenic characterization of TF-positive MPs need to be

10

Microparticles and Cancer Biology

237

addressed in order to obtain a reliable method for determining the cellular source of MP in vivo. Also, standardization of preanalytical procedures and development of more sensitive technologies are needed to improve our current understanding of the role of circulating TF-positive MPs in thrombosis. Similarly, findings derived from studies regarding issues concerning methodologies and standardization of TF-positive MPs will also have application in studies investigating the role of MPs in tumor growth, angiogenesis, and metastasis. In view of the pathophysiological implications of MPs in cancer, and to pave the way for novel and beneficial therapeutic interventions and strategies targeting cellular microparticles implicated in tumor growth and metastasis, as well as cancerassociated thrombosis, more intense and focused multidisciplinary research efforts are urgently needed to: (a) Standardize preanalytical procedures and assay methods and to develop more sensitive and high throughput technologies (b) Unravel the mechanisms of MP production and action (c) Characterize the cellular proteins and lipids represented in MP that is so pivotal to the understanding of their function (d) Compare the protein and lipid composition of MPs formed by exposure of cells to different agonists/stimuli and those formed in vivo in different cancers at different stages of disease as well as pre- and post-cancer therapy (e) To elucidate mechanisms (like MP-cell fusion or ligand–receptor interactions) by which MP interact with their target cells

References 1. Furie, B., and Furie, B. C. 2006 Blood Cells Mol Dis 36, 177 2. Ratajczak, J., Wysoczynski, M., Hayek, F., Janowska-Wieczorek, A., and Ratajczak, M. Z. 2006 Leukemia 20, 1487 3. Amin, C., Mackman, N., and Key, N. S. 2008 Pathophysiol Haemost Thromb 36, 177 4. Tesselaar, M. E., Romijn, F. P., Van Der Linden, I. K., Prins, F. A., Bertina, R. M., and Osanto, S. 2007 J Thromb Haemost 5, 520 5. Castellana, D., Kunzelmann, C., and Freyssinet, J. M. 2009 Hamostaseologie 29, 51 6. Wolf, P. 1967 Br J Haematol 13, 269 7. Crawford, N. 1971 Br J Haematol 21, 53 8. Hugel, B., Martinez, M. C., Kunzelmann, C., and Freyssinet, J. M. 2005 Physiology (Bethesda) 20, 22 9. Thery, C., Zitvogel, L., and Amigorena, S. 2002 Nat Rev Immunol 2, 569 10. VanWijk, M. J., VanBavel, E., Sturk, A., and Nieuwland, R. 2003 Cardiovasc Res 59, 277 11. Morel, O., Toti, F., Hugel, B., Bakouboula, B., Camoin-Jau, L., Dignat-George, F., and Freyssinet, J.-M. 2006 Arterioscler Thromb Vasc Biol 26, 2594 12. Burnier, L., Fontana, P., Kwak, B., and Angelillo-Scherrer, A. 2009 Thromb Haemost. 101, 439 13. Freyssinet, J. M. 2003 J Thromb Haemost 1, 1655 14. Kim, H. K., Song, K. S., Park, Y. S., Kang, Y. H., Lee, Y. J., Lee, K. R., Kim, H. K., Ryu, K. W., Bae, J. M., and Kim, S. 2003 Eur J Cancer 39, 184 15. Savasan, S., Buyukavci, M., Buck, S., and Ravindranath, Y. 2004 J Clin Pathol 57, 651

238

P.P. Goh

16. Kalinkovich, A., Tavor, S., Avigdor, A., Kahn, J., Brill, A., Petit, I., Goichberg, P., Tesio, M., Netzer, N., Naparstek, E., Hardan, I., Nagler, A., Resnick, I., Tsimanis, A., and Lapidot, T. 2006 Cancer Res 66, 11013 17. Del Conde, I., Bharwani, L. D., Dietzen, D. J., Pendurthi, U., Thiagarajan, P., and Lopez, J. A. 2007 Journal of Thrombosis and Haemostasis 5, 70 18. Bergmann, C., Strauss, L., Wieckowski, E., Czystowska, M., Albers, A., Wang, Y., Zeidler, R., Lang, S., and Whiteside, T. L. 2009 Head Neck 31, 371 19. Kim, J. W., Wieckowski, E., Taylor, D. D., Reichert, T. E., Watkins, S., and Whiteside, T. L. 2005 Clin Cancer Res 11, 1010 20. Hron, G., Kollars, M., Weber, H., Sagaster, V., Quehenberger, P., Eichinger, S., Kyrle, P. A., and Weltermann, A. 2007 Thromb Haemost 97, 119 21. Amabile, N., Guerin, A. P., Leroyer, A., Mallat, Z., Nguyen, C., Boddaert, J., London, G. M., Tedgui, A., and Boulanger, C. M. 2005 J. Am. Soc. Nephrol. 16, 3381 22. Haubold, K., Rink, M., Spath, B., Friedrich, M., Chun, F. K., Marx, G., Amirkhosravi, A., Francis, J. L., Bokemeyer, C., Eifrig, B., and Langer, F. 2009 Thromb Haemost 101, 1147 23. Kim, H. K., Song, K. S., Park, Y. S., Kang, Y. H., Lee, Y. J., Lee, K. R., Ryu, K. W., Bae, J. M., and Kim, S. 2003 Eur J Cancer 39, 184 24. Zwaal, R. F. A., and Schroit, A. J. 1997 Blood 89, 1121 25. Bevers, E. M., Comfurius, P., Dekkers, D. W. C., and Zwaal, R. F. A. 1999 Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids 1439, 317 26. Bevers, E. M., Comfurius, P., and Zwaal, R. F. A. 2003 Membrane phospholipid asymmetry: biochemical and pathophysiological perspectives. in Advances in Molecular and Cell Biology E. E. Bittar (Ed.), Elsevier, 387 27. Jimenez, J., Jy W, Mauro LM, Soderland, C., Horstmann, L., and Ahn, Y. 2003 109, 175 28. Tramontano, A. F., O’Leary, J., Black, A. D., Muniyappa, R., Cutaia, M. V., and El-Sherif, N. 2004 Biochemical and Biophysical Research Communications 320, 34 29. Perez-Pujol, S., Marker, P. P., and Key, N. S. 2007 Cytometry Part A 71A, 38 30. Wiedmer, T., Shattil, S. J., Cunningham, M., and Sims, P. J. 1990 Biochemistry 29, 623 31. Hamilton, K. K., Hattori, R., Esmon, C. T., and Sims, P. J. 1990 J. Biol. Chem. 265, 3809 32. Falanga, A., Panova-Noeva, M., and Russo, L. 2009 Best Practice & Research Clinical Haematology 22, 49 33. Kwaan, H. C., and Keer, H. N. 1990 Semin Thromb Hemost 16, 230 34. Rickles, F. R., and Falanga, A. 2001 Thromb Res 102, V215 35. Brodsky, S. V., Malinowski, K., Golightly, M., Jesty, J., and Goligorsky, M. S. 2002 Circulation 106, 2372 36. Varon, D., and Brill, A. 2001 Haemostasis 31 Suppl 1, 64 37. Prandoni, P., Falanga, A., and Piccioli, A. 2005 The Lancet Oncology 6, 401 38. Vaupel, P., and Mayer, A. 2007 Cancer and Metastasis Reviews 26, 225 39. Harris, A. L. 2002 Nat Rev Cancer 2, 38 40. Wysoczynski, M., and Ratajczak, M. Z. 2009 Int J Cancer 125, 1595 41. Grenader, T., Shavit, L., Ospovat, I., Gutfeld, O., and Peretz, T. 2006 Mt Sinai J Med 73, 810 42. Lechner, D., and Weltermann, A. 2008 Semin Thromb Hemost 34, 199 43. Gelderman, M. P., Carter, L. B., and Simak, J. 2004 ASH Annual Meeting Abstracts 104, 3635 44. Simak, J., and Gelderman, M. P. 2006 Transfus Med Rev 20, 1 45. Jy, W., Horstman, L. L., Jimenez, J. J., Ahn, Y. S., Biro, E., Nieuwland, R., Sturk, A., DignatGeorge, F., Sabatier, F., Camoin-Jau, L., Sampol, J., Hugel, B., Zobairi, F., Freyssinet, J. M., Nomura, S., Shet, A. S., Key, N. S., and Hebbel, R. P. 2004 J Thromb Haemost 2, 1842 46. Horstman, L. L., Jy, W., Jimenez, J. J., Bidot, C., and Ahn, Y. S. 2004 Keio J Med 53, 210 47. Pula, G., Perera, S., Prokopi, M., Sidibe, A., Boulanger, C., and Mayr, M. 2008 PROTEOMICS CLINICAL APPLICATIONS 2, 882 48. Liotta, L. A., and Kohn, E. C. 2001 Nature 411, 375 49. Chambers, A. F., Groom, A. C., and MacDonald, I. C. 2002 Nat Rev Cancer 2, 563 50. Mueller, M. M., and Fusenig, N. E. 2004 Nat Rev Cancer 4, 839

10

Microparticles and Cancer Biology

239

51. Taylor, D., and Black, P. 1986 Shedding of plasma membrane fragments. Neoplastic and developmental importance. in Developmental Biology, M.S.Steinberg (Ed.), Plenum Press, New York, 33 52. Barz, D., Goppelt, M., Szamel, M., Schirrmacher, V., and Resch, K. 1985 Biochim Biophys Acta 814, 77 53. Baj-Krzyworzeka, M., Szatanek, R., Weglarczyk, K., Baran, J., Urbanowicz, B., Branski, P., Ratajczak, M. Z., and Zembala, M. 2006 Cancer Immunol Immunother 55, 808 54. Deryugina, E. I., and Quigley, J. P. 2006 Cancer Metastasis Rev 25, 9 55. Dolo, V., Ginestra, A., Cassara, D., Violini, S., Lucania, G., Torrisi, M. R., Nagase, H., Canevari, S., Pavan, A., and Vittorelli, M. L. 1998 Cancer Res 58, 4468 56. Ginestra, A., Monea, S., Seghezzi, G., Dolo, V., Nagase, H., Mignatti, P., and Vittorelli, M. L. 1997 J Biol Chem 272, 17216 57. Graves, L. E., Ariztia, E. V., Navari, J. R., Matzel, H. J., Stack, M. S., and Fishman, D. A. 2004 Cancer Res 64, 7045 58. Taraboletti, G., D’Ascenzo, S., Borsotti, P., Giavazzi, R., Pavan, A., and Dolo, V. 2002 Am J Pathol 160, 673 59. Taraboletti, G., D’Ascenzo, S., Giusti, I., Marchetti, D., Borsotti, P., Millimaggi, D., Giavazzi, R., Pavan, A., and Dolo, V. 2006 Neoplasia 8, 96 60. Skog, J., Wurdinger, T., van Rijn, S., Meijer, D. H., Gainche, L., Sena-Esteves, M., Curry, W. T., Jr., Carter, B. S., Krichevsky, A. M., and Breakefield, X. O. 2008 Nat Cell Biol 10, 1470 61. Tarin, D. 1972 Epithelial-Mesenchymal Interactions in Carcinogenesis Academic Press, London 62. Hood, J. D., and Cheresh, D. A. 2002 Nat Rev Cancer 2, 91 63. Ahmed, N., Riley, C., Rice, G., and Quinn, M. 2005 Clin Exp Metastasis 22, 391 64. Augustin, H. G. 2000 Baillieres Best Pract.Res. Clin. Obstet. Gynaecol. 14, 867 65. Reynolds, L. P., Grazul-Bilska, A. T., and Redmer, D. A. 2002 Intl. J. Exptl. Pathol 83, 151 66. Arnold, F., and West, D. C. 1991 Pharmacology & Therapeutics 52, 407 67. Carmeliet, P. 2003 Nat. Med. 9, 653 68. Folkman, J. 2002 Semin Oncol 29, 15 69. Folkman, J. 1995 Nature Med. 1, 27 70. Folkman, J. 1995 N. Engl. J. Med. 333, 1757 71. Folkman, J. 1971 N Engl J Med 285, 1182 72. Papetti, M., and Herman, I.M. 2002 Am J Physiol Cell Physiol 282, 947 73. Carmeliet, P., and Jain, R. K. 2000 Nature 407, 249 74. Brat, D. J., Bellail, A. C., and Van Meir, E. G. 2005 Neuro-oncology 7, 122 75. Eberle, K., Oberpichler, A., Trantakis, C., Krupp, W., Knupfer, M., Tschesche, H., and Seifert, V. 2000 Anticancer Res 20, 1679 76. Rolhion, C., Penault-Llorca, F., Kemeny, J.-L., Lemaire, J.-J., Jullien, C., Labit-Bouvier, C., Finat-Duclos, F., and Verrelle, P. 2001 Journal of Neurosurgery 94, 97 77. Li, A., Dubey, S., Varney, M. L., Dave, B. J., and Singh, R. K. 2003 J Immunol 170, 3369 78. Goswami, S., and Gupta, A. 1998 Journal of Neurochemistry 71, 1837 79. Bergers, G., Brekken, R., McMahon, G., Vu, T., Itoh, T., Tamaki, K., Tanzawa, K., Thorpe, P., Itohara, S., Z. &, W., and Hanahan, D. 2000 Nature Cell Biology, 2, 737 80. Folkman, J. 2008 Angiostatin and Endostatin: Angiogenesis Inhibitors in Blood and Stroma. in Angiogenesis: An Integrative Approach from Science to Medicine, W.D. Figg and J. Folkman (Eds.), Springer US., 129 81. Sounni, N. E., Devy, L., Hajitou, A., Frankenne, F., Munaut, C., Gilles, C., Deroanne, C., Thompson, E. W., Foidart, J. M., and Noel, A. 2002 FASEB J. 16, 555 82. Deryugina, E. I., Soroceanu, L., and Strongin, A. Y. 2002 Cancer Res 62, 580 83. Sidhu, S. S., Mengistab, A. T., Tauscher, A. N., LaVail, J., and Basbaum, C. 2004 Oncogene 23, 956 84. Millimaggi, D., Mari, M., D’Ascenzo, S., Carosa, E., Jannini, E. A., Zucker, S., Carta, G., Pavan, A., and Dolo, V. 2007 Neoplasia 9, 349

240

P.P. Goh

85. Castellana, D., Zobairi, F., Martinez, M. C., Panaro, M. A., Mitolo, V., Freyssinet, J.-M., and Kunzelmann, C. 2009 Cancer Res 69, 785 86. Biswas, C., Zhang, Y., DeCastro, R., Guo, H., Nakamura, T., Kataoka, H., and Nabeshima, K. 1995 Cancer Res 55, 434–439 87. Toole, B. 2003 Curr Top Dev Biol 54, 371–389 88. Tang, Y., Nakada, M. T., Kesavan, P., McCabe, F., Millar, H., Rafferty, P., Bugelski, P., and Yan, L. 2005 Cancer Res 65, 3193 89. Tang, Y., Kesavan, P., Nakada, M. T., and Yan, L. 2004 Mol Cancer Res 2, 73 90. Sun, J., and ME, H. 2001 Cancer Res 61, 2276–2281 91. Guo, H., Zucker, S., Gordon, M. K., Toole, B. P., and Biswas, C. 1997 J Biol Chem 272, 24 92. Caudroy, S., Polette, M. N.-R., B., Cao, J., Toole, B. P., Zucker, S., and Birembaut, P. 2002 Clin Exp Metastasis 19, 697 93. Kanekura, T., and Chen, X. 2010 J Dermatol Sci 57, 149 94. Stetler-Stevenson, W. G., Aznavoorian, S., and Liotta, L. A. 1993 Annu Rev Cell Biol 9, 541 95. Stetler-Stevenson, W. G. 2001 Surg Oncol Clin N Am 10, 383 96. Yu, J., May, L., Milsom, C., Anderson, G. M., Weitz, J. I., Luyendyk, J. P., Broze, G., Mackman, N., and Rak, J. 2008 Arterioscler Thromb Vasc Biol 28, 1975 97. Forster, Y., Meye, A., Albrecht, S., and Schwenzer, B. 2006 Clin Chim Acta 364, 12 98. Rak, J., Milsom, C., Magnus, N., and Yu, J. 2009 Best Pract Res Clin Haematol 22, 71 99. Schaffner, F., and Ruf, W. 2009 Arterioscler Thromb Vasc Biol, Aug 6. [Epub ahead of print] 100. Al-Nedawi, K., Meehan, B., Kerbel, R. S., Allison, A. C., and Rak, J. 2009 Proc Natl Acad Sci USA 106, 3794 101. Amin, D. N., Hida, K., Bielenberg, D. R., and Klagsbrun, M. 2006 Cancer Res 66, 2173 102. Baker, C. H., Kedar, D., McCarty, M. F., Tsan, R., Weber, K. L., Bucana, C. D., and Fidler, I. J. 2002 Am J Pathol 161, 929 103. Yokoi, K., Thaker, P. H., Yazici, S., Rebhun, R. R., Nam, D. H., He, J., Kim, S. J., Abbruzzese, J. L., Hamilton, S. R., and Fidler, I. J. 2005 Cancer Res 65, 3716 104. Cheng, H., Langley, R. R., Wu, Q., Wu, W., Feng, J., Tsan, R., Fan, D., and Fidler, I. J. 2005 Neoplasia 7, 1065 105. Vosseler, S., Mirancea, N., Bohlen, P., Mueller, M. M., and Fusenig, N. E. 2005 Cancer Res 65, 1294 106. Skobe, M., Rockwell, P., Goldstein, N., Vosseler, S., and Fusenig, N. E. 1997 Nature Med. 3, 1222 107. Hellstrom, K. E., and, and Hellstrom, I. 1991 Principles of tumor immunity: tumor antigens. in Biologic Therapy of Cancer V. T. DeVita, S. Hellman, and Rosenberg, S. A. (Eds.), Lippincott, New York, 35 108. Kim, R., Emi, M., Tanabe, K., Uchida, Y., and Toge, T. 2004 Cancer 100, 2281 109. Maher, S., Toomey, D., Condron, C., and Bouchier-Hayes, D. 2002 Immunol Cell Biol 80, 131 110. Rabinowich, H., Reichert, T. E., Kashii, Y., Gastman, B. R., Bell, M. C., and Whiteside, T. L. 1998 J Clin Invest 101, 2579 111. Mottolese, M., Buglioni, S., Bracalenti, C., Cardarell, M. A., i, Ciabocco, L., Giannarelli, D., Botti, C., Natali, P. G., Concetti, A., and Venanzi, F. M. 2000 International Journal of Cancer 89, 127 112. Chopin, D., Barei-Moniri, R., Maille, P., Le Frere-Belda, M.-A., Muscatelli-Groux, B., Merendino, N., Lecerf, L., Stoppacciaro, A., and Velotti, F. 2003 Am J Pathol 162, 1139 113. Mann, B., Gratchev, A., Bohm, C., Hanski, M. L., Foss, H. D., Demel, G., Trojanek, B., SchmidtWolf, I., Stein, H., Riecken, E. O., Buhr, H. J., and Hanski, C. 1999 Br J Cancer 79, 1262 114. Lai, P., Rabinowich, H., Crowley-Nowick, P. A., Bell, M. C., Mantovani, G., and Whiteside, T. L. 1996 Clin Cancer Res 2, 161 115. Huber, V., Fais, S., Iero, M., Lugini, L., Canese, P., Squarcina, P., Zaccheddu, A., Colone, M., Arancia, G., Gentile, M., Seregni, E., Valenti, R., Ballabio, G., Belli, F., Leo, E., Parmiani, G., and Rivoltini, L. 2005 Gastroenterology 128, 1796

10

Microparticles and Cancer Biology

241

116. Andreola, G., Rivoltini, L., Castelli, C., Huber, V., Perego, P., Deho, P., Squarcina, P., Accornero, P., Lozupone, F., Lugini, L., Stringaro, A., Molinari, A., Arancia, G., Gentile, M., Parmiani, G., and Fais, S. 2002 J Exp Med 195, 1303 117. Taylor, D. D., Gercel-Taylor, C., Lyons, K. S., Stanson, J., and Whiteside, T. L. 2003 Clin Cancer Res 9, 5113 118. Dolo, V., Pizzurro, P., Ginestra, A., and Vittorelli, M. L. 1995 J Submicrosc Cytol Pathol 27, 535 119. Becker, C., Fantini, M. C., and Neurath, M. F. 2006 Cytokine & Growth Factor Reviews 17, 97 120. Reiss, M., and Barcellos-Hoff, M. H. 1997 Breast Cancer Research and Treatment 45, 81 121. Friedman, E., Gold, L. I., Klimstra, D., Zeng, Z. S., Winawer, S., and Cohen, A. 1995 Cancer Epidemiol Biomarkers Prev 4, 549 122. Matsuzaki, K., Date, M., Furukawa, F., Tahashi, Y., Matsushita, M., Sakitani, K., Yamashiki, N., Seki, T., Saito, H., Nishizawa, M., Fujisawa, J., and Inoue, K. 2000 Cancer Res 60, 1394 123. Reed, J. A., McNutt, N. S., Prieto, V. G., and Albino, A. P. 1994 Am J Pathol 145, 97 124. Gorelik, L., and Flavell, R. A. 2001 Nat Med 7, 1118 125. Hoefer, M., and Anderer, F. A. 1995 Cancer Immunology, Immunotherapy 41, 302 126. Whiteside, T. L. 2005 Br J Cancer 92, 209 127. Garcia-Lora, A., Algarra, I., and Garrido, F. 2003 Journal of Cellular Physiology 195, 346 128. Nicolson, G. L. 1993 Cancer and Metastasis Reviews 12, 325 129. Baggiolini, M. 1998 Nature 392, 565 130. Balkwill, F. 2004 Nat Rev Cancer 4, 540 131. Muller, A., Homey, B., Soto, H., Ge, N., Catron, D., Buchanan, M. E., McClanahan, T., Murphy, E., Yuan, W., Wagner, S. N., Barrera, J. L., Mohar, A., Verastegui, E., and Zlotnik, A. 2001 Nature 410, 50 132. Rubie, C., Oliveira-Frick, V., Rau, B., Schilling, M., and Wagner, M. 2006 J Clin Oncol 24, 5173 133. Shulby, S. A., Dolloff, N. G., Stearns, M. E., Meucci, O., and Fatatis, A. 2004 Cancer Res 64, 4693 134. Ghadjar, P., Rubie, C., Aebersold, D. M., and Keilholz, U. 2009 Int J Cancer 125, 741 135. Scotton, C. J., Wilson, J. L., Milliken, D., Stamp, G., and Balkwill, F. R. 2001 Cancer Res 61, 4961 136. Burger, J., A., and Bürkle, A. 2007 British Journal of Haematology 137, 288 137. Baj-Krzyworzeka, M., Szatanek, R., Weglarczyk, K., Baran, J., and Zembala, M. 2007 Immunol Lett 113, 76 138. Levesque, M. C., and Haynes, B. F. 1996 J Immunol 156, 1557 139. Honn, K. V., Tang, D. G., and Crissman, J. D. 1992 Cancer Metastasis Rev 11, 325 140. Honn, K. V., Tang, D. G., and Chen, Y. Q. 1992 Semin Thromb Hemost 18, 392 141. Taucher S, Salat A, G. M., Kwasny W, Mlineritsch B, Menzel R, Schmid M, Smola MG, Stierer M, Tausch C, Galid A, Steger G, e. a., and . 2003 Thromb Haemost 89, 1098 –1106 142. McCarty, O. J. T., Mousa, S. A., Bray, P. F., and Konstantopoulos, K. 2000 Blood 96, 1789 143. Nash, G., Turner, L., Scully, M., and Kakkar, A. 2002 Lancet Oncol. 3, 425 144. Jurasz, P., Alonso-Escolano, D., and Radomski, M. W. 2004 Br J Pharmacol 143, 819 145. Radomski, M. W., Jenkins, D. C., Holmes, L., and Moncada, S. 1991 Cancer Res 51, 6073 146. Philippe, C., Philippe, B., Fouqeray, B., Perez, J., Lebret, M., and Baud, L. 1993 Am. J. Pathol. 143, 1713–1723 147. Konstantopoulos, K., and Thomas, S. N. 2009 Annual Review of Biomedical Engineering 11, 177 148. Janowska-Wieczorek, A., Majka, M., Kijowski, J., Baj-Krzyworzeka, M., Reca, R., Turner, A. R., Ratajczak, J., Kowalska, M. A., and Ratajczak, M. Z. 2001 Blood 98, 3143–3149. 149. Baj-Krzyworzeka M, Majka M, Pratico D, Ratajczak J, Vilaire G, Kijowski J, Reca R, Janowska-Wieczorek A, and MZ., R. 2002 Exp Hematol 30, 450 –459 150. English, D., Garcia, J. G. N., and Brindley, D. N. 2001 Cardiovasc Res 49, 588 151. Janowska-Wieczorek, A., Marcin, W., Jacek, K., Leah, M.-C., Bogdan, M., Janina, R., and Mariusz, Z. R. 2005 International Journal of Cancer 113, 752 152. Janowska-Wieczorek, A., Marquez-Curtis, L., Wysoczynski, M., and Ratajczak, M. 2006 Transfusion. 46, 1199

242

P.P. Goh

153. Houldsworth, J., Reuter, V., Bosl, G. J., and Chaganti, R. S. 1997 Cell Growth Differ 8, 293 154. Takano, Y., Kato, Y., Masuda, M., Ohshima, Y., and Okayasu, I. 1999 The Journal of Pathology 189, 194 155. Seiki, M., Koshikawa, N., and Yana, I. 2003 Cancer Metastasis Rev 22, 129 156. Zucker, S., Pei, D., Cao, J., and Lopez-Otin, C. 2003 Membrane type-matrix metalloproteinases (MT-MMP). in Current topics in developmental biology: Cell surface proteases (Zucker S, and WT, C. eds.), Academic Press, New York. pp 1 157. English, D., Welch, Z., Kovala, A. T., Harvey, K., Volpert, O. V., Brindley, D. N., and Garcia, J. G. 2000 Faseb J 14, 2255 158. Barry OP, Pratico D, Lawson JA, and GA., F. 1997 J Clin Invest 99, 2118 –2127 159. Barry, O. P., Pratico, D., Lawson, J. A., and FitzGerald, G. A. 1997 J Clin Invest 99, 2118 160. Barry, O. P., Kazanietz, M. G., Pratico, D., and FitzGerald, G. A. 1999 J Biol Chem 274, 7545 161. Kim, C. W., Lee, H. M., Lee, T. H., Kang, C., Kleinman, H. K., and Gho, Y. S. 2002 Cancer Res 62, 6312 162. Baj-Krzyworzeka, M., Majka, M., Pratico, D., Ratajczak, J., Vilaire, G., Kijowski, J., Reca, R., Janowska-Wieczorek, A., and Ratajczak, M. Z. 2002 Experimental Hematology 30, 450 163. Rozmyslowicz, T., Majka, M., Kijowski, J., Murphy, S. L., Conover, D. O., Poncz, M., Ratajczak, J., Gaulton, G. N., and Ratajczak, M. Z. 2003 AIDS 17, 33 164. Prandoni, P., Falanga, A., and Piccioli, A. 2007 Thrombosis Research 120, S137–S140 165. Rauch, U., and Antoniak, S. 2007 Thromb Haemost 97, 9 166. Falanga, A., Barbui, T., and Rickles, F. R. 2008 Semin Thromb Hemost 34, 204 167. Zangari, M., Anaissie, E., Barlogie, B., Badros, A., Desikan, R., Gopal, A. V., Morris, C., Toor, A., Siegel, E., Fink, L., and Tricot, G. 2001 Blood 98, 1614 168. Zangari, M., Elice, F., Fink, L., and Tricot, G. 2007 Expert Rev Anticancer Ther 7, 307 169. Pritchard, K. I., Paterson, A. H., Paul, N. A., Zee, B., Fine, S., and Pater, J. 1996 J Clin Oncol 14, 2731 170. Haddad, T. C., and Greeno, E. W. 2006 Thromb Res 118, 555 171. Petropoulou, A. D., Gerotziafas, G. T., Samama, M. M., Hatmi, M., Rendu, F., and Elalamy, I. 2008 Thrombosis Research 121, 493 172. Varki, A. 2007 Blood 110, 1723 173. Falanga, A. 2005 Semin Thromb Hemost 31, 104 174. Rickles, F. R., and Brenner, B. 2008 Semin Thromb Hemost 34, 143 175. Lopez-Pedrera, C., Barbarroja, N., Dorado, G., Siendones, E., and Velasco, F. 2006 Leukemia 20, 1331 176. Drake, T. A., Morrissey, J. H., and Edgington, T. S. 1989 Am. J. Pathol. 134, 1087 177. Mackman, N. 2004 Arterioscler. Thromb. Vasc. Biol. 24, 1015 178. Giesen, P. L., Rauch, U., Bohrmann, B., Kling, D., Roque, M., Fallon, J. T., Badimon, J. J., Himber, J., Riederer, M. A., and Nemerson, Y. 1999 Proc Natl Acad Sci USA 96, 2311 179. Furie, B., Zwicker, J., LaRocca, T., Kos, C., Bauer, B., and Furie, B. C. 2005 Haematologica reports 1, 5 180. Zwicker, J. I. 2008 Semin Thromb Hemost 34, 195 181. Tilley, R. E., Holscher, T., Belani, R., Nieva, J., and Mackman, N. 2008 Thromb Res 122, 604 182. Davila, M., Amirkhosravi, A., Coll, E., Desai, H., Robles, L., Colon, J., Baker, C. H., and Francis, J. L. 2008 J Thromb Haemost 6, 1517 183. Lechner, D., and Weltermann, A. 2008 Thrombosis Research 122, S47–S54 184. Khorana, A. A., Francis, C. W., Menzies, K. E., Wang, J. G., Hyrien, O., Hathcock, J., Mackman, N., and Taubman, M. B. 2008 J Thromb Haemost 6, 1983 185. Langer, F., Spath, B., Haubold, K., Holstein, K., Marx, G., Wierecky, J., Brummendorf, T. H., Dierlamm, J., Bokemeyer, C., and Eifrig, B. 2008 Ann Hematol 87, 451 186. Falati, S., Liu, Q., Gross, P., Merrill-Skoloff, G., Chou, J., Vandendries, E., Celi, A., Croce, K., Furie, B. C., and Furie, B. 2003 J Exp Med 197, 1585 187. Rauch, U., Bonderman, D., Bohrmann, B., Badimon, J. J., Himber, J., Riederer, M. A., and Nemerson, Y. 2000 Blood 96, 170

10

Microparticles and Cancer Biology

243

188. del Conde, I., Shrimpton, C. N., Thiagarajan, P., and Lopez, J. A. 2005 Blood 106, 1604 189. Muller, I., Klocke, A., Alex, M., Kotzsch, M., Luther, T., Morgenstern, E., Zieseniss, S., Zahler, S., Preissner, K., and Engelmann, B. 2003 Faseb J 17, 476 190. Carr, J. M., Dvorak, A. M., and Dvorak, H. F. 1985 Cancer Res 45, 5944 191. Yu, J. L., and Rak, J. W. 2004 Journal of Thrombosis and Haemostasis 2, 2065 192. Bastida, E., Ordinas, A., Escolar, G., and Jamieson, G. A. 1984 Blood 64, 177 193. Osterud, B. 2003 Dis Mon 49, 7 194. Furie, B., and Furie, B. C. 2004 Trends in Molecular Medicine 10, 171 195. Bach, R. R. 2006 Arterioscler. Thromb. Vasc. Biol. 26, 456 196. Ahamed, J., Versteeg, H. H., Kerver, M., Chen, V. M., Mueller, B. M., Hogg, P. J., and Ruf, W. 2006 Proc Natl Acad Sci USA 103, 13932 197. Graven, K. K., Molvar, C., Roncarati, J. S., Klahn, B. D., Lowrey, S., and Farber, H. W. 2002 Am J Physiol Lung Cell Mol Physiol 282, L996 198. Essex, D. W., Chen, K., and Swiatkowska, M. 1995 Blood 86, 2168 199. Sullivan, D. C., Huminiecki, L., Moore, J. W., Boyle, J. J., Poulsom, R., Creamer, D., Barker, J., and Bicknell, R. 2003 J. Biol. Chem. 278, 47079 200. Chen, K., Detwiler, T. C., and Essex, D. W. 1995 Br J Haematol 90, 425 201. Claudio, J. O., Masih-Khan, E., Tang, H., Goncalves, J., Voralia, M., Li, Z. H., Nadeem, V., Cukerman, E., Francisco-Pabalan, O., Liew, C. C., Woodgett, J. R., and Stewart, A. K. 2002 Blood 100, 2175 202. Raturi, A., Miersch, S., Hudson, J. W., and Mutus, B. 2008 Biochim Biophys Acta 1778, 2790 203. Sinauridze, E. I., Kireev, D. A., Popenko, N. Y., Pichugin, A. V., Panteleev, M. A., Krymskaya, O. V., and Ataullakhanov, F. I. 2007 Thromb Haemost 97, 425 204. Banfi, C., Brioschi, M., Wait, R., Begum, S., Gianazza, E., Pirillo, A., Mussoni, L., and Tremoli, E. 2005 PROTEOMICS 5, 4443 205. Versteeg, H. H., and Ruf, W. 2007 J. Biol. Chem. 282, 25416 206. Cho, J., Furie, B. C., Coughlin, S. R., and Furie, B. 2008 J Clin Invest 118, 1123 207. Reinhardt, C., von Bruhl, M. L., Manukyan, D., Grahl, L., Lorenz, M., Altmann, B., Dlugai, S., Hess, S., Konrad, I., Orschiedt, L., Mackman, N., Ruddock, L., Massberg, S., and Engelmann, B. 2008 J Clin Invest 118, 1110 208. Wilkinson, B., and Gilbert, H. F. 2004 Biochimica et Biophysica Acta (BBA) - Proteins & Proteomics 1699, 35 209. Stone, J. P., and Wagner, D. D. 1993 J Clin Invest 92, 804 210. Brossart, P., Schneider, A., Dill, P., Schammann, T., Grunebach, F., Wirths, S., Kanz, L., Buhring, H.-J., and Brugger, W. 2001 Cancer Res 61, 6846 211. Navarro, J., Obrador, E., Carretero, J., Petschen, I., Avino, J., Perez, P., and Estrela, J. M. 1999 Free Radic Biol Med 26, 410 212. Yu, J. L., May, L., Lhotak, V., Shahrzad, S., Shirasawa, S., Weitz, J. I., Coomber, B. L., Mackman, N., and Rak, J. W. 2005 Blood 105, 1734 213. Rak, J., Milsom, C., and Yu, J. 2008 Curr Opin Hematol 15, 522

Chapter 11

Protein Kinase D Signaling in Cancer Peter Storz

Introduction The protein kinase D (PKD) family of serine/threonine kinases consists of three isoenzymes, PKD1/PKCm, PKD2, and PKD3/PKCn [1–4]. Due to similarities in some domain structures, PKD was originally classified as a member of the protein kinase C (PKC) family [2]. With the description of the relations of kinases in the whole human kinome, it became clear that PKD represents an own group more closely related to the calcium/calmodulin-regulated (Cam) kinases [5]. Like protein kinase C, PKD enzymes are phorbol ester- and diacylglycerol (DAG)-responsive kinases and are further regulated by phosphorylation events to gain full activity [6]. Activating phosphorylations are mediated by members of the novel PKC (nPKC) group and tyrosine kinases such as Src and Abl, all of which have been implicated in various functions in cancer before (Fig. 11.1). In additional to the regulation of PKD kinase activity, epigenetic regulation of PKD1 occurs in multiple cancers at the transcriptional levels through methylation of a CpG island in its promoter [7, 8]. PKD molecules consist of N-terminal regulatory and C-terminal kinase domains (Fig. 11.1). The regulatory region comprises the CRD domain (two cysteine-rich C1 domains C1a and C1b), an acidic region (AR), and the pleckstrin homology (PH) domain. The deletion of either of these regions or domains leads to increased PKD1 activity, suggesting that the N-terminal region is negative-regulatory for the kinase domain [9, 10]. As a potential autoinhibitory mechanism, it was suggested that the regulatory domain folds back on the kinase domain. Activation of the kinase may occur through binding to DAG and subsequent phosphorylation of PKD at tyrosine residues in its PH domain and/or at the activation loop serines in the kinase domain [11, 12]. The C1 domains localize the kinase to the plasma membrane in response to

P. Storz (*) Department for Cancer Biology, Mayo Clinic, Jacksonville, FL 32224, USA e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_11, © Springer Science+Business Media, LLC 2012

245

246

P. Storz

Fig. 11.1 Domain structure and phosphorylation-dependent regulation of PKD. Shown are the domain structure of PKD1, the activity-related phosphorylation sites, the respective function of their phosphorylation, and the involved upstream kinases. PKD can be divided into a catalytic domain and a regulatory region that consists of multiple lipid and protein binding domains as well as regions of unknown function such as AR (C1 C1 domains C1a and C1b, AR acidic region, PH pleckstrin homology domain, kinase kinase domain, PDZ PDZ protein binding motif). PKD1 phosphorylation occurs by upstream kinases or by auto-/transphosphorylation. Activity-relevant phosphorylation sites are included in the PKD1 domain structure, and kinases regulating these are indicated (proteins that interact directly with PKD1 are labeled blue). CQND labels the caspase cleavage site

the generation of DAG or treatment with phorbol esters or to the mitochondria in response to oxidative stress [13, 14]. Both C1 domains have distinct functional roles in targeting and maintaining PKD1 at the plasma membrane. C1a achieves fast maximal and reversible translocation, while C1b translocates the enzyme partially, but persistently. Further, persistent localization requires the binding of the C1b domain to Gaq protein [15]. C1a is also required for PKD1 localization to the golgi apparatus [16, 17] and C1b for nuclear import [18]. While the function of the AR is unknown, the PH domain does not bind any particular phosphoinositide with high specificity but serves as a protein–protein interface by binding to upstream kinases such as PKCh [19] and the G-protein bg-subunit [20], but also is required for nuclear export [18]. PKD1 has multiple functions within cells ranging from regulating golgi organization and vesicle transport, cell proliferation and cell survival, to mitochondrial signaling to the nucleus (for reviews, see [21, 22]). Recently, a role in regulating the actin cytoskeleton was added to this list, since it was shown that PKD1 interacts with F-actin structures [23], is regulated by RhoGTPases and kinases that are involved in regulating cell migration and adhesion [24], and once active, inhibits F-actin reorganization at the lamellipodium [25]. While some of these functions are mediated redundantly by all three enzymes, others are specifically regulated by individual PKD isoforms. For example, in oxidative stress signaling in tumor cells, PKD1 and PKD2 may have overlapping roles, but PKD3 may have functions distinct from PKD1/2.

11

Protein Kinase D Signaling in Cancer

247

Fig. 11.2 Upstream signaling events leading to PKD activation. Due to different localization within cells, the activation mechanisms for PKD vary with the upstream stimuli. Generally, PKD1 is localized to membranes by binding to DAG (diacylglycerol) and then activated by phosphorylations. Shown are three variant mechanisms that have been described as leading to PKD1 activity (yellow, lipid-generating protein; blue, protein directly interacting with PKD1; green, PKD; phosphorylations occurring in the respective signaling pathway are indicated). (a) In order to facilitate vesicle transport to the plasma membrane, PKD1 may be constitutively active at the golgi compartment. This is mediated by binding of PKD1 to DAG and Gbg, anchoring the protein to the golgi, as well as PKCh-mediated activation loop phosphorylation at S738 and S742 (for human PKD1). (b) PKD1 is activated by a similar mechanism at the plasma membrane that uses PKCe and may require binding to Ga. Active PKD1 is then released into the cytosol or translocates to the nucleus. (c) Tyrosine phosphorylations of PKD1 are mediated by Abl (Y463 in human PKD1) and Src (Y463 in human PKD1) and induce mitochondrial localization and interaction with PKCd. The generation of DAG is important for PKD1 and PKCd localization to the mitochondria

With the identification of LXRXXS (with X as any amino acid, L, leucine, R, arginine, and S, serine) as the PKD minimal substrate phosphorylation motif [26] and the subsequent generation of a substrate-specific antibody [27], optimal conditions were provided for discovering PKD1 substrates. This led to the description of an increasing wealth of PKD1 substrates and further defined the role of PKD in various processes in cancer (Fig. 11.2). The functions of PKD enzymes in cancer range from tumor-promoting events including increased proliferation and survival signaling to tumor cell progression, including the regulation of cell motility and angiogenesis (reviewed in [22]). However, with respect to cell signaling and function in tumor biology, PKD3 seems to be very distinct in its functions from the two other PKDs. For example, PKD3 rather seems to increase cell migration and invasion of some tumors [28–30], while PKD1 and PKD2 exhibit an inhibitory role on cell migration [25, 31]. Currently, a major focus in the field is to further determine the exact functions of each PKD family member.

248

P. Storz

Activation of PKD Upstream Activators for PKD Upstream activators for PKD in cancer cells include tumor-promoting phorbol esters (e.g., PdBu), growth factor receptors (GF-Rs), heterotrimeric G-proteincoupled receptors (GPC-Rs), RhoGTPases, inducers of reactive oxygen species (ROS), as well as inducers of apoptosis (e.g., chemotherapeutics). Activation of PKD by phorbol esters – Phorbol esters are natural products from plants and were initially characterized as potent tumor promoters in mouse skin. Phorbol esters exert divergent cellular responses, and multiple effectors are accountable for the various responses in cells and animal models for cancer [32]. Phorbol esters bind to C1 domains of DAG receptors and mimic DAG action by affecting cell proliferation, differentiation, survival, and transformation (for a review on this topic: [33]). Besides protein kinase C as one of the primary targets, PKD is also a high-affinity receptor for phorbol ester [32], and PKD activation by phorbol esters has been demonstrated for multiple tumor cell lines [34]. Phorbol ester binding of PKD1 occurs via C1b and localizes the kinase to the plasma membrane, where it is further activated [35]. Activation of PKD by growth factor receptors – Protein kinase D is activated downstream of growth factors such as platelet-derived growth factor (PDGF), epidermal growth factor (EGF), and vascular endothelial growth factor (VEGF) [36, 37]. For example in NIH-3T3 cells, the activation of PKD1 through PDGF is mediated by phospholipase Cg (PLCg) and PKC, but not PI 3-kinase (PI3K) [37]. EGF-mediated activation of PKD1 was shown in A431 cells [38]; however in other cell systems, PKD1 is not activated by EGF. Moreover, PDGF-mediated activation of PKD1 leads to a modulation of EGF signaling through direct phosphorylation of the EGF receptor [39]. The activation of PKD by growth factors seems to be cell-type specific and only occurs when the respective growth factor receptor is expressed in very high copy numbers such as PDGF-R in NIH-3T3 or EGF-R in A431 cells and may also be linked to integrin signaling. The regulation of PKD by VEGF occurs in endothelial cells and was shown to contribute to multiple aspects involved in angiogenesis (see chapter below). Activation of PKD by G proteins – PKD is activated by a variety of heterotrimeric guanine nucleotide-binding regulatory protein (G-protein)-coupled receptors including receptors for mitogenic peptides such as bombesin, vasopressin, endothelin, and bradykinin, or receptors for lysophosphatidic acid (LPA) [36]. Heterotrimeric G proteins are composed of a-, b-, and g-subunits, and G-protein a-subunits are classified into four subgroups, Gas, Gai, Gaq, and Ga12/Ga13. It was shown for PKD1 and PKD3 that activity is increased by G-protein a-subunits of the Ga12/Ga13, Gaq, and Gai subgroups [14, 40–43]. For example, Gaq and, to some extent, Gai contribute to LPA- and angiotensin-mediated activation of PKD1 [44]. In addition to its regulation by a-subunits, G-protein bg-subunits also contribute to PKD localization and activation [20].

11

Protein Kinase D Signaling in Cancer

249

Dependent on the stimulus and a-subunit involved, G-protein-coupled receptors activate PKD1 via DAG generation through phospholipase Cb or g enzymes (PLCb, PLCg) or phospholipase D (PLD) [45, 46]. For example, Gaq stimulates DAG formation through PLCb. Lysophosphatidic acid (LPA)-mediated activation of PKD1 occurs through PLD-mediated generation of phosphatidic acid (PA) and subsequent formation of DAG [46]. One functional consequence of PKD activation by LPA is the activation of the transcription factor NF-kB, and it was recently shown that PKD2 mediates LPA-induced interleukin 8 expression via NF-kB [47]. As another example, the peptide gastrin has been implicated in the development of colorectal cancers that express CCKB/gastrin receptors and is involved in a wide range of functions including neoplastic transformation [48]. PKD2 is activated by the CCKB/ gastrin receptor involving a heterotrimeric Gaq protein as well as PLC [49]. Activation by RhoGTPases – PKD isoenzymes are also activated by small RhoGTPases including RhoA, Rac, and Cdc42. For example, it was shown that ectopic expression of constitutively active RhoA mainly activates PKD1, and constitutively active Rac activates PKD3 [14, 40–43]. RhoA has a major role in regulating responses that lead to cytoskeletal reorganization involved in the formation of actin stress fibers, focal adhesions, and regulation of cell motility [50], and it was shown that PKD1 regulates some of the RhoA-mediated cytoskeletal changes [14, 24, 25]. There is also a crosstalk between heterotrimeric G proteins and RhoGTPases. For example, members of the Ga12 family mediate activation of RhoA via guanine nucleotide exchange factors that link the a-subunits to RhoA [51, 52]. Further, interactions of Gaq and Ga12/13 with the RhoGTPase Rac lead to the activation of PKD3 in bombesin receptor signaling [42, 43]. Activation of PKD by oxidative stress – Due to altered or increased metabolism or mitochondrial dysfunction, tumor cells often show increased levels of reactive oxygen species (ROS). ROS can have multiple functions in cancer including the regulation of cell proliferation, survival, motility, and angiogenesis (reviewed in [53, 54]). Protein kinase D1 is a sensor for oxidative stress in tumor cells [55]. The elevation of cellular ROS through treatment with hydrogen peroxide or increase of intracellular oxidative stress levels through L-S,R-buthionine sulfoximine (BSO)-mediated pharmacological depletion of the ROS scavenger glutathione (GSH) leads to tyrosine phosphorylation-mediated activation of PKD [12, 38, 56, 57]. Further, PKD1 activity is increased by a variety of agents such as rotenone that induce the mitochondrial generation of superoxide [58]. Dependent on the cellular system, growth factor receptors also elevate intracellular superoxide levels through NADPH oxidase or the mitochondria [54]. NADPH oxidase is activated via the small GTPase Rac-1 [59], which was described as a regulator of PKD3 [42, 60]. Moreover, production of ROS by neutrophils and macrophages as a mechanism to kill tumor cells is well established [54]. In these cells, a rapid burst of superoxide formation primarily mediated by NAPDH oxidase leads to subsequent production of hydrogen peroxide [61, 62]. However, the activation of PKD enzymes through NADPH oxidase-mediated ROS has not been described so far.

250

P. Storz

Activation of PKD by cleavage – In response to apoptotic stimuli, PKD1 can be activated by cleavage through caspases. Caspase cleavage sites are located in the regulatory region of the enzyme located between the acidic region and the pleckstrin homology domain [63]. Caspase-mediated cleavage of PKD1 was described after treatment of cells with chemotherapeutically used genotoxic agents including 1-b-D-arabinofuranosylcytosine (ara-C), etoposide, cisplatin, and doxorubicin, and upon activation of death receptors [63–65]. Caspase-mediated cleavage generates an unregulated kinase fragment that sensitizes cells to apoptotic cell death, but is not apoptotic per se [64].

Signaling Mechanisms Regulating PKD Activity Protein kinase D activity is regulated by the binding to lipid or protein cofactors, regulating its membrane localization, and by covalent modifications through phosphorylation by upstream kinases [66]. The activation mechanisms as well as the activating and interacting proteins slightly vary between the intracellular localization of PKD (Fig. 11.2). Membrane targeting as an initial activation step – An initial step in PKD activation is the translocation of the enzyme from the cytosol to intracellular membranes. This can occur either through binding to DAG via its CRD domain or through binding to G-protein a- or bg-subunits. The CRD domain of PKD includes two C1 domains, C1a and C1b, which both have slightly different specificities to DAG [35]. Dependent on the PKD-activating pathway, DAG is either directly generated through PLCb or PLCg enzymes, or indirectly formed via PA generated by PLD1. The later activation mechanism has been implicated in oxidative stress-mediated activation of PKD1 at the mitochondria, as well as LPA-mediated activation of PKD1 [13, 46]. All other PKD-activating pathways described so far seem to utilize PLCg or PLCb to generate DAG. Regulation of PKD by activation loop phosphorylation – The function of DAG binding of PKD is to release the inhibitory regulatory domain from the kinase domain and to bring the enzyme to intracellular membranes and into proximity to its upstream activators, allowing activation by phosphorylation. PKD enzymes are direct targets for the nPKC isoforms PKCe, PKCd, PKCq, and PKCh, which are also located to cellular membranes upon binding to DAG [11, 19, 56, 67–71]. Additionally, PKD and nPKC enzymes seem to form complexes. For example, PKCh binds directly to the PH domain of PKD1 [19], and PKCd binds with its C2 domain to the N-terminus of PKD1 through a phosphorylated tyrosine residue [69]. With recent work, it becomes evident that different stimuli utilize different nPKC isoforms to activate PKD at the membranes of various intracellular compartments. Activation of PKD by nPKCs occurs through phosphorylation of the activation loop serines S738 and S742 (in human PKD1) [72]. The phosphorylated residues in the activation loop most likely fork an ionic bridge with amino-acid residues in the

11

Protein Kinase D Signaling in Cancer

251

catalytic loop, thereby stabilizing the active catalytic structure of the enzyme [66, 73]. Phosphorylation of the activation loop serines has been shown to be an absolute requirement for the kinase to achieve full catalytic competence. Regulation of PKD by tyrosine phosphorylations – Tyrosine phosphorylation of PKD was described in response to increased oxidative stress [12, 38], expression of BCR–Abl [74], or activation of the RhoGTPase RhoA [24], and does not occur in response to GPC-R activation [38]. For oxidative stress, a sequential order of activation steps has been described initiated by DAG, which contributes to PKD1 localization to the mitochondria [13]. A key kinase in this activation is Src, which can be activated by hydrogen peroxide through direct oxidation or through reversible oxidation and inactivation of its negative-regulatory phosphatases [75, 76]. In response to ROS, Src initiates the phosphorylation of PKD1 at several sites including Y95 and Y463 [12, 69]. While Y95 is directly phosphorylated by Src [69], Y463 phosphorylation is a consequence of Src-mediated activation of Abl [38]. The phosphorylation of PKD1 at this residue is directly mediated by Abl [12] and leads to a conformational change and to mitochondrial localization [13, 38]. The phosphorylation of PKD1 at Y95 facilitates binding of PKCd through its C2 domain and subsequent phosphorylation of the activation loop of PKD1, leading to full activity [56, 57, 69]. This PKD1 activation pathway originally was described in cervix carcinoma cells, but meanwhile has been shown to be active in a multitude of tumor cell lines including breast cancer and pancreatic cancer cells. It is required for PKD1 and PKD2 activation by oxidative stress. PKD3 lacks a residue homologue to Y95 and is not activated via this mechanism [69]. Tyrosine phosphorylation-mediated PKD1 activation induces activation of the transcription factor nuclear factor k-B (NF-kB) and increases cell survival [38]. In BCR–Abl positive myeloid leukemia cells, PKD2 can be activated by its phosphorylation at a tyrosine residue homologue to Y463 but does not need the activation loop phosphorylations to mediate the activation of NF-kB [74]. Regulation of PKD by phosphorylation and protein interactions – Additional in vivo phosphorylation sites in PKD1 leading to its regulation by facilitating the binding of regulatory proteins were described [77–80]. These include residues S203, S208, S219, and S223 whose phosphorylation regulates binding of PKD1 to 14-3-3, a mechanism that decreases the activity of the kinase [78]. Although it was suggested that phosphorylation of these residues may occur through auto- or transphosphorylation, the amino acids surrounding these serines are not generating PKD phosphorylation motifs. Another in vitro PKD autophosphorylation site is serine residue 910 (in human PKD1) [79]. The amino acids surrounding S910 also do not represent an ideal phosphorylation motif for the kinase, and the role of PKD phosphorylation at S910 in vivo is not well understood. Furthermore, it is unclear if the phosphorylation of this site is a consequence of PKD1 activation, since it was shown that it occurs independent or prior to PKD1 activation loop phosphorylation, suggesting that in vivo, also other kinases may be involved in the phosphorylation of this residue [57, 81]. For example, in response to PLD1 activation, DAG generation from PA

252

P. Storz

induces PKD1 autophosphorylation at S910, and this occurs prior to PKC-mediated activation loop phosphorylation [46]. S910 is localized within a PDZ (Postsynaptic density-95/Disks large/Zonula occludens-1-binding motif) protein binding motif [80]. PKD1 controls the fission of golgi transport carriers specifically destined to the cell surface [17]. As an example for such mechanism, PKD1 and PKD2 regulate neurotensin secretion in pancreatic cancer cells [82]. It was shown that phosphorylation of S910 leads to a release of a PDZ-binding protein from the PDZ motif, resulting in the release of vesicles from the trans-golgi network [80]. Phosphorylation of PKD1 at S910 is also necessary for the recycling of avb3-integrin [83]. Moreover, it was shown that death-associated protein kinase (DAPK) leads to S910 but not activation loop phosphorylation of PKD1 and contributes to ROS-mediated activation of JNK [84]. Since PKD1 inhibits JNK activity [70], DAPK-mediated phosphorylation of PKD at S910 may also lead to a modulation of PDZ motif/PDZ binding protein interactions. Taken together, it is very likely that S910 phosphorylation represents a yet not well-defined additional regulation mechanism for PKD.

Role of PKD in Tumor Cell Proliferation Highly proliferative cancers such as mouse skin carcinomas show increased PKD expression. Moreover, increased levels of PKD1 in mouse keratinocytes were correlated with increased DNA synthesis and proliferation [85]. Furthermore, expression of PKD1 in human pancreatic ductal adenocarcinoma (PDAC) cell lines leads to enhanced proliferation and significant increases in telomerase activity [86]. Similarly, stable expression of PKD2 increases the proliferation of cell lines of human carcinoid tumors, rare neuroendocrine cancers with a predilection for the gastrointestinal tract [87, 88]. In androgen-dependent prostate cancer cells, a forced expression of PKD3 promoted S-phase entry. Further, depletion of these cells from PKD3 led to G0–G1 phase arrest and inhibition of androgen-independent cell proliferation [28]. Proliferative signaling by PKD seems to be mediated in response to triggering of growth factor receptors (GFRs) or G-protein-coupled receptors (GPCRs) [89]. For example, neuropeptides such as neurotensin (NT) and bombesin-like peptides act as autocrine or paracrine growth factors and potent cellular mitogens for a variety human cancers [90, 91]. Neuropeptides signal through Gaq subunits of G-proteincoupled receptors. All three PKD isoforms, PKD1, PKD2, and PKD3, have been shown to be activated and redistributed within cells by GPCRs. For example, PKD1 is shuttled to the nucleus after its activation [18, 92], and similar redistribution was shown for PKD2 and PKD3 in epithelial cell lines [60]. However, the cellular targets for active, nuclear PKD in response to these signals is so far unknown. One possibility is that nuclear PKD targets histone deacetylases (HDACs) to modulate gene expression. Histone-dependent packaging of genomic DNA into chromatin is a fundamental mechanism to regulate gene expression. Histone acetyltransferases

11

Protein Kinase D Signaling in Cancer

253

(HATs) mediate relaxation of nucleosomal structures through acetylation of histones, allowing transcriptional activators to access genes and activate gene transcription. Conversely, histone deacetylases condense the chromatin through deacetylation of histones, leading to transcriptional repression [93]. PKD phosphorylates the class II histone deacetylases HDAC5 and HDAC7 directly [94, 95]. PKD isoenzymes seem to have redundant roles in regulating HDACs [96]. Signaling through G-protein-coupled receptors induces the phosphorylation of HDAC5 at two serine residues, S259 and S498, of which all three PKD isoforms catalyze the phosphorylation of S498 [97]. Direct phosphorylation of HDAC5 through PKD drives its nuclear export [95]. A-kinase anchoring protein (AKAP)-Lbc coordinates activation and movement of signaling proteins and couples activation of PKD1 with the phosphorylation-dependent nuclear export of HDAC5 [98]. Neurotensin is a gut peptide with an important role in gastrointestinal cell motility and stimulates growth of normal gut mucosa. Additionally, NT also stimulates proliferation of NT receptor-positive cancers, including prostate, pancreatic, and colon cancer. Neurotensin and EGF induce synergistic stimulation of DNA synthesis in PDAC cells lines [91]. This is mediated by an increase in the duration of Erk1/2 signaling [99]. Both PKD1 and PKD2 have been shown to increase the duration of Erk signaling and potentiate DNA synthesis [100, 101]. Further, expression of constitutively active PKD1 leads to an activation of Raf-1 and subsequent activation of Erk1/2 [102]. PKD1-mediated regulation of the MAPK pathway also enhanced the transcriptional activity of SRE (serum response element)-driven genes through the Elk-1 ternary complex factor [102]. Similarly, PKD3 contributes to prostate cancer cell proliferation through regulation of Erk1/2 activity [28]. The expression of constitutively active Ras mutants has been identified as an early event in multiple cancers. For example, in pancreatic cancer, K-ras-activating mutations have been detected in over 95% of all cases [103]. Ras activates cell proliferation via the Raf1-Erk1/2 signaling cascade. As a potential mechanism, PKD regulates the Ras effector RIN1 (Ras and Rab interactor 1) through phosphorylation [104]. RIN1 competes with Raf1 for binding to Ras, whereby RIN1 seems to be a negative regulator of Ras signaling. For example, RIN1 efficiently blocks Ras-mediated transformation. Upon phosphorylation of RIN1 by PKD at S351, RIN binds to 14-3-3 proteins and loses its affinity to Ras and membrane localization [104]. Another potential mechanism of how PKD may contribute to increased cell proliferation is through regulation of sphingosine kinase 2 (SPHK2). PKD1 phosphorylates SPHK2 at its nuclear export sequence and induces its nuclear export. SPHK2 in the nucleus inhibits DNA synthesis under stress conditions [105, 106]. After its nuclear export, SPHK2 is active and generates the second messenger sphingosine 1-phosphate [107]. Sphingosine 1-phosphate is a bioactive lipid that regulates cell proliferation, cell survival, cell differentiation, and cell motility, dependent on its locus of generation [108]. Some of these processes are mediated by sphingosine 1-phosphate-specific G-protein-coupled receptors [108]. Expression of SPHK enhanced tumor cell proliferation and promoted G1/S transition, protected cells from apoptosis, and induced tumor formation in mice [109–112].

254

P. Storz

Role of PKD in Tumor Cell Survival Signaling PKD1 promotes cell survival in cancer cells in response to different inducers of cell death. For example, PKD1 activation occurs after DNA damage by genotoxic stress and is critically involved in survival signaling [113]. Further, in PDAC (pancreatic ductal adenocarcinoma) cell lines, the increased expression of PKD1 correlates with their resistance to CD95-mediated apoptosis [86]. Loss of growth control and resistance to apoptosis are major mechanisms that drive the progression of pancreatic tumors. PKD1 may protect pancreatic cancer cells by upregulating anti-apoptotic proteins such as c-FLIPL and survivin [86]. PKD1 also blocks TNFa-mediated apoptosis since it upregulates NF-kB-dependent protective genes such as TRAF-1 (TNF receptor-associated protein 1) or cIAP2 (inhibitor of apoptosis protein 2) [114]. Furthermore, PKD1 protects from ROS-mediated cell death in multiple cancer cell types through upregulation of antioxidant and anti-apoptotic genes, including SOD2 (encoding MnSOD) and A20 [58, 115]. This is mediated through a mitochondria-to-nucleus signaling pathway involving the activation of PKD1 through PLD1, Src, and PKCd that leads to the induction of NF-kB through its canonical activation pathway (IKKa/IKKb/NEMO complex) [38]. Tyrosine phosphorylation-mediated activation of PKD also occurs in chronic myelogenous leukemia (CML). CML is characterized by the expression of the BCR–Abl fusion protein (detected in 95% of all CML patients). The BCR–Abl oncogene renders the Abl kinase constitutively active and induces cell proliferation, transformation, and resistance to apoptosis in myeloid leukemia cells. In CML cell lines, PKD2 is tyrosinephosphorylated by BCR–Abl at a residue homologue to the Abl site in PKD1 for ROS signaling. Furthermore, similar to PKD1, tyrosine phosphorylation of PKD2 by BCR–Abl leads to its activation and subsequent induction of NF-kB [74]. The seine/threonine kinase Akt is a key effector of PI3K-mediated survival signaling [116, 117]. Although it was shown that PKD1 signaling to NF-kB is uncoupled from the PI3K–Akt signaling pathway [38], in prostate cancer ectopical expression of PKD3 increased Akt activity, and this required PI3K and p38 kinase [28]. Akt targets multiple pro-apoptotic substrates such as Bad and FOXO3a with the net effect to protect tumor cells from cell death [118–120]. This suggests that the PKD3 isoform may protect tumor cells from cell death via Akt. Besides actively inducing cell survival pathways, PKD1 also inhibits pro-apoptotic signaling through c-Jun N-terminal kinases (JNKs). JNK contributes to cell death by negatively regulating Bcl-2 [121]. Inhibition of PKD leads to increased activity of JNK and its target, the transcription factor c-Jun [122]. Resistance from cell death requires nPKC/PKD1 and Erk1/2 activation to switch off pro-apoptotic JNK/c-Jun signaling [122]. For example, PKCh-mediated activation of PKD1 led to an increase of Erk1/2 activity and induction of the serum response element. At the same time, PKD1 activation reduced JNK activity, suggesting that PKD1 in this pathway switches from apoptotic JNK signaling to proliferative signaling [70]. Similar inhibitory effects of PKD1 on JNK activity were obtained when PKCe was activated via PDK-1 (phosphoinositide-dependent kinase-1) [68]. It is unclear if

11

Protein Kinase D Signaling in Cancer

255

PKD1 negatively regulates JNK by direct phosphorylation; however, it was demonstrated that activation loop phosphorylation of PKD1 leads to a complex formation with JNK [123]. PKD1 may also modulate upstream activators for JNK. For example, in some cell types PKD1 has been implicated in the attenuation of EGF-induced activation of JNK [39]. PKD1 directly phosphorylates the EGF receptor at threonine residues T654 ad T669, which reduces the signaling capacity of EGF-R toward JNK. Interestingly, in lung and pancreatic cell lines, where the EGF–JNK pathway has also been implicated in sustaining the cancerous phenotypes, EGF-stimulated JNK activity is uncoupled from PKD1 suppression [124]. Further, PKD1 also phosphorylates the JNK target c-Jun directly in vitro at serine residue S58 and suppresses its phosphorylation by JNK at S63 and S73 [123, 125]. The function of this phosphorylation is not further characterized, but it may be speculated that it leads to an inhibition of c-Jun function. In some cell systems, PKD signaling toward JNK may also be inactivated to promote cell death. For example, in endothelial cells in response to some apoptotic stimuli, PKD1 interacts with the JNK upstream kinase ASK1 (apoptosis signalregulated kinase 1). This association is mediated through the PH domain of PKD1 and the C-terminal region of ASK1 but also involves the binding of 14-3-3 proteins to PKD1 [126]. Since the binding of 14-3-3 to PKD1 has been shown to be a negative-regulatory event [78], it is very likely that such signaling negatively regulates PKD1 but also increases ASK1 activity, leading to JNK activation and promoting apoptosis in endothelial cells. As another example, death-associated protein kinase (DAPK) is a cell death–promoting S/T kinase that is activated by ROS and induces JNK signaling. PKD1 is a target for DAPK, and both interact in response to ROS. DAPK leads to phosphorylation of PKD1 at S910 and contributes to ROS-mediated activation of JNK [84]. It is unclear if this S910 phosphorylation represents PKD1 activation or, since DAPK does not induce PKD1 activation loop phosphorylation, leads to a modulation of PDZ protein/PDZ binding protein interactions and inactivation of PKD1 toward JNK. However, proteolytic cleavage of PKD1 may lead to an increased sensitivity of cancer cells to apoptotic stimuli. PKD1 is cleaved and activated by caspase-3 in the apoptotic response of cancer cells to genotoxic agents including 1-b-D-arabinofuranosylcytosine, etoposide, cisplatin, or doxorubicin, and to inducers of cell death such as TNFa [63–65]. Such cleavage is blocked in cells that express anti-apoptotic proteins such as Bcl-XL. In apoptotic signaling, PKD1 cleavage is associated with the generation of a constitutively active (unregulated) catalytic fragment. Cells stably expressing this kinase fragment are sensitized to genotoxic stress-induced apoptosis [64].

Functions of PKD in Chemoresistance RNA interference (RNAi) screens of human kinase RNAi libraries identified PKD as a regulator of chemoresistance and a survival kinase [127]. Mechanisms of how PKD mediates chemoresistance however remain unclear. Recent data suggest the

256

P. Storz

contribution of PKD to several signaling events that previously have been shown to mediate chemoresistance of tumor cells. ROS in tumor cells can contribute to the oncogenic phenotype. However exuberant high levels also can lead to increased macromolecule damage that eventually kills tumor cells, a mechanism that is used by multiple chemotherapeutics (reviewed in [54]). Tumor cells have developed multiple protective mechanisms, which are activated in response to such threats, and their activation results in the upregulation of survival and repair of genes [128, 129]. For example, the activation of the transcription factors FOXO3a and NF-kB has been linked to upregulation of antioxidant genes such as SOD and catalase in response to ROS or radiation [58, 128, 130–134]. PKD1 is a sensor for oxidative stress that relays an increase in cellular ROS to the NF-kB-mediated expression of antioxidant genes. In pancreatic cancer, which is known for its increased ROS levels in tumor cells, NF-kB has been shown to mediate tumor cell survival and to induce resistance to chemotherapeutic agents [135, 136]. Further, it was recently shown that metallothionein 2A (MT 2A) interacts with the kinase domain of PKD1 in prostate cancer [137]. Metallothioneins are smallmolecular-weight trace metal scavenging proteins that also bind to p50 NF-kB [138]. Like NF-kB, metallothioneins are known to cause resistance to chemotherapy in several cancers including PC [139]. Hsp27 is a ubiquitous expressed member of the small heat shock protein family, with major roles in regulating apoptosis, actin reorganization processes, and cell migration. Like NF-kB, Hsp27 has been implicated in chemoresistance of pancreatic cancer cells [140]. Hsp27 is a PKD1 substrate, and PKD1 phosphorylates Hsp27 at S82 [27]. Of note, Hsp27 is also in vitro phosphorylated at S82 by MAPKAPK [141], but PKD1 is the relevant physiological kinase for this site in response to oxidative stress [27]. Phosphorylation of Hsp27 at this site has previously been demonstrated in response to inducers of stresses, including inflammatory cytokines, osmotic stress, ROS, and heat shock [141, 142]. In Panc-1 cells, neurotensin-induced PKD1 and PKD2 activity also mediated the phosphorylation of Hsp27 at S82 [143]. Unphosphorylated Hsp27 forms large multimers, and its phosphorylation at Ser 82 in particular promotes their dissociation [142]. Phosphorylation of Hsp27 modulates its oligomerization and chaperone function, resulting in the protection of tumor cells from injury due to stress [144]. Therefore, PKD-mediated phosphorylation of S82 may protect tumor cells from cellular stresses and thus contribute to chemoresistance in cancer.

Role of PKD in Tumor Cell Migration and Invasion The roles of the PKD isoforms in processes regulating tumor cell migration and invasion are still ill-defined. Moreover, converse results were obtained regarding the function of PKD1 in regulating cell migration in epithelial or endothelial cells. It is also unclear if different PKD isoforms may have opposing effects on cell motility. Most studies were performed with PKD1, and an emerging picture for this

11

Protein Kinase D Signaling in Cancer

257

kinase is that it may inhibit the migration and invasion of epithelial cancer cells at multiple levels. Role of PKD in cell–cell and cell–matrix aggregation – Recent data suggest that PKD1 may be crucial for cell–cell adhesion and linking adhesion complexes to the actin cytoskeleton [24, 145, 146]. The formation of E-cadherin/b-catenin complexes at the cell junctions plays an important role in maintaining cell integrity, polarity, and morphogenesis [145], and an aberrant expression or localization of the components of this complex has been associated with cancer progression and metastasis. Phosphorylation of E-cadherin or b-catenin results in increased stabilization of the cadherin–catenin complex and increases cell–cell adhesion in adherens junctions [147, 148]. In prostate cancer cells, PKD1 is localized to such cell junctions, where it phosphorylates E-cadherin [149]. This was associated with increased cell aggregation and decreased motility [150]. Furthermore, downregulation of E-cadherin or PKD1 increased cell migration and invasion [151]. b-catenin is essential for E-cadherin-mediated cell adhesions in epithelial cells and also acts as a key cofactor for transcriptional activity. PKD1 interacts with b-catenin and phosphorylates it at two threonine residues [152]. Blunting mutations of these residues abolished its interaction with a-catenin and resulted in increased nuclear localization of b-catenin, where it shows transcriptional activity. Expression of active PKD1 in prostate cancer cells stabilizes its interaction with E-cadherin and represses b-catenin-induced transcriptional activity [152]. Moreover, the activation of PKD1 with bryostatin 1, a natural macrocyclic lactone with antineoplastic properties in many tumors, induces a co-localization of the cytoplasmatic pool of b-catenin with PKD1 and proteins involved in vesicular trafficking [145]. PKD1 was also linked to an integrin-mediated cell adhesion to extracellular matrix. Such cell–matrix interactions are important for many cellular processes, including the maintenance of tissue integrity and tumor cell metastasis. Cell adhesion to collagen confers motility and invasive properties in vitro as well as the metastatic potential of tumor cells in vivo [153]. It was shown that cis-polyunsaturated fatty acids or arachidonic acid enhances b1-integrin-mediated adhesion of human breast carcinoma cells to type IV collagen by activating PKD1 and its upstream kinase PKCe [154]. PKD1 also associates with the b3 of the avb3-integrin and promotes avb3-integrin recycling. Furthermore, incorporation of avb3-integrin in newly forming focal adhesions requires its association with active PKD1 [155]. PKD functions at the leading edge of migrating cells – Cancer cell migration toward a chemotactic stimulus requires rapid actin severing, branching, and polymerization at the leading edge to mediate membrane protrusion [156, 157]. This is concerted by a complex interplay of proteins such as Wiskott–Aldrich syndrome protein (WASp), Arp2/3 complex, capping proteins, Lin-11/Isl-1/Mec-3 kinases (LIMK), slingshot (SSH) phosphatases, ADF (actin depolymerizing factor)/cofilin, and profilin [156]. In cervix carcinoma, pancreatic cancer, and breast cancer cells, PKD1 is recruited to sites of such actin reorganization at the leading edge, where it negatively regulates directed cell migration [23, 25]. Stimulus-induced lamellipodium protrusion and

258

P. Storz

directed cell migration is initiated by actin severing through ADF/cofilin. ADF/ cofilin is negatively regulated by phosphorylation through LIMK, and this is reversed by the phosphatase slingshot (SSH) [158]. Slingshot activity is regulated through two kinases, PAK4 (p21-activated kinase) and PKD1 [25, 159]. While PAK4 phosphorylates slingshot at the N-terminus [159], PKD1 phosphorylates the slingshot isoform SSH1L at a serine residue located within its actin binding motif, releasing it from F-actin through facilitating the binding of 14-3-3 [25]. Binding to 14-3-3 proteins sequesters SSH1L in the cytoplasm, rendering it inactive toward cofilin [160]. This results in enhanced phospho-cofilin levels in invasive tumor cells correlating with decreased formation of free actin filament barbed ends and effective inhibition of directed cell migration [25]. A similar mechanism was confirmed for PKD2 [31]. PKD1 also interacts with and phosphorylates the Ena/VASP protein family member EVL-1 in its EVH2 domain. Since EVL-1 is also localized at the leading edge of migrating cells, this may have a role in filopodia formation or the regulation of lamellipodia and cell–cell contacts [161]. Moreover, cortactin, an actin binding protein that accumulates in lamellipodia and membrane ruffles, recently has been characterized as an in vivo substrate for PKD1 [162]. However, PKD1mediated phosphorylations did not affect cortactin localization, and the cellular function of this signaling event remains unknown. PKD functions in cargo transport as a mechanism to impact migration – A subcellular pool of PKD is localized to the trans-golgi network (TGN) and regulates the fission of transport carriers specifically destined to the plasma membrane [163, 164] (reviewed in [165]), a process that also needs actin nucleation and reorganization. Therefore it is possible that PKD1 at the golgi has similar targets and functions as at the leading edge [158]. Further, an inactive PKD1 mutant inhibits golgi cargo transport from the TGN to the plasma membrane and the retrograde flow of surface markers and filamentous actin [166]. Role of PKD in cell invasion – Recent data indicate that in some epithelial cancers, PKD1 expression and activity negatively regulate cell invasion [7]. For example, PKD1 expression is lost at the mRNA level in highly invasive breast cancer cell lines including MDA-MB-231 cells, whereas PKD1 is still expressed in low- or nonmotile cell lines such as MCF-7. The knockdown of PKD1 from such lowmotile cells, however, increases their motility and invasion. Moreover, the reintroduction of a constitutively active PKD1 in MDA-MB-231 cells completely abolishes the ability of these cells to invade [7]. As a potential mechanism for such effects on the invasive phenotype, it was shown that PKD1 negatively regulates the expression of multiple matrix metalloproteases (MMPs). These enzymes previously have been shown to be involved in the degradation of extracellular matrix. When ectopically expressed, PKD1 negatively regulates the expression of MMP-2, MMP-7, MMP-9, MMP-10, MMP-11, MMP-13, and MMP-14 [7], all known to enhance the invasion of MDA-MB-231 cells in Matrigel [167–169]. The mechanisms of how PKD1 regulates the expression of these molecules are unknown. However, HDACs have been shown to regulate the expression of MMPs. Since PKD is a negative regulator of multiple HDACs [94–97], it may be speculated that PKD1 regulates the expression

11

Protein Kinase D Signaling in Cancer

259

of MMPs by inhibiting the function of HDACs. In contrast to downregulating the expression of multiple MMPs, PKD1 expression also increased the expression of MMP-3. This was observed in human fibroblasts and breast cancer cell lines [7, 170]. This is interesting since in breast cancer, MMP-3 expression was associated with benign early stage tumors and is frequently lost in advanced stage, aggressive cancers [171]. Therefore, it was speculated that loss of PKD1 expression and the resulting altered MMP expression may be linked to a switch driving the progression from benign to malignant tumors [7]. Notably, the increased expression of the PKD isoform PKD3 was linked to an increased invasiveness. Therefore, it may also be speculated that an isoform switch from PKD1 to PKD3 may promote tumor cell invasion. Finally, it was also shown that PKD forms complexes with cortactin and paxillin at sites of extracellular matrix degradation, but functions for this complex have not been assigned [172]. Multiple myeloma is an incurable B-cell neoplasia characterized by accumulation of malignant plasma cells in the bone marrow after crossing of endothelial barriers. The growth factor IGF-1 acts as a chemotactic factor for multiple myeloma cells and promotes migration and tissue invasion. In later stages of the disease, these cells extravasate through blood vessels and are responsible for metastasis. In this context, IGF-1 promotes transmigration through vascular endothelial cells and bone marrow stromal cells. With an inhibitor study, PKD1 and ROCK (Rho-associated kinase) were identified as mediators of such migration, indicating that PKD1 may have a converse function in tumor cells other than epithelial origin [173].

Regulation of PKD Expression in Cancer In normal tissues, PKD2 and PKD3 are expressed ubiquitously, whereas PKD1 expression seems to be restricted to immune cells, endothelial and epithelial cells. In tumor tissues, depending on the PKD isoenzyme, as well as the subtype and stage of the respective cancer, up- or downregulation of PKD expression has been described. While little is known for the two PKD isoforms, PKD2 and PKD3, recent data suggest that in epithelial cancers, PKD1 expression or activity may be needed at early stages of tumor formation but may be gradually lost with acquisition of increased cell motility at later more-aggressive stages. Gastric cancer – In gastric cancer, the PKD1 promoter is epigenetically inactivated [8]. Analysis of paired clinical primary gastric cancer samples indicated that approximately 60% of the tumors showed a decrease in PKD1 expression as compared to normal tissue, and this downregulation was correlated with methylation of the PKD promoter [8]. Methylation is also observed when cells of normal appearing mucosal tissue age, predicting that PKD1 methylation may be one of the earliest events that predispose an individual to gastric cancer. Moreover, approximately 70% of examined gastric cancer cell lines showed CpG hypermethylation of the PKD1 promoter region. In gastric cancer, PKD1 expression was further correlated with a negative regulation of motility and invasion, and the knockdown of PKD1 from gastric cancer cell lines increased cell invasion [8].

260

P. Storz

Breast cancer – Human tissue samples from invasive breast cancer and normal breast tissue were evaluated for PKD expression. PKD1 was highly expressed in ductal epithelial cells of normal human breast tissue but was approx. 50% reduced in its expression in more than 95% of the analyzed samples of human invasive breast tumors. PKD2 and PKD3 showed no significant differences in their expression levels [7]. Furthermore, PKD1 is not expressed in invasive breast cancer cell lines, whereas noninvasive or low-invasive breast cancer cell lines express PKD1 [7]. For cell lines, it was shown that the decrease in PKD1 expression is probably mediated through DNA methylation [7]. Similarly as shown for gastric cancer, a decreased PKD1 expression or activity led to an increased motility and invasion of breast cancer cells [7]. Prostate cancer – Prostate cancer is the second leading cause of death in men in the United States [174]. Progression of this cancer to androgen independence and chemoresistance are the main causes of death in patients [137]. PKD1 expression at transcription and translational levels is decreased in 100% of androgen-independent (AI) human prostate cancers [150]. Androgen-dependent (AD) prostate cancer cell lines show a 4- to 16-fold higher PKD1 expression as androgen-independent (AI) cell lines [175]. This suggests a potential role for a loss of PKD expression in progression to a more-aggressive stage. PKD1 complexes with the androgen receptor and affects its transcriptional activity [176]. It was also shown that PKD1-mediated phosphorylation of Hsp27 at S82 is necessary for the repression of the transcriptional activity of the androgen receptor and androgen-dependent proliferation of prostate cancer cells [177]. Only little is known about the involved mechanism by which PKD1 is silenced in prostate cancer, but it may be speculated that a similar epigenetic silencing occurs than observed for gastric and breast cancer. In contrast to PKD1, PKD3 seems to be overexpressed in prostate cancer [28] and may contribute to cancer progression [30]. Other cancers – In contrast to gastric, breast, and prostate cancer, immunohistochemical analysis of pancreatic tumor tissue revealed marked overexpression of PKD1/2 in tumors [86]. Furthermore, a majority of human hyperproliferative skin disorders such as psoriasis and basal cell carcinomas (BCC) show significantly elevated PKD1 protein levels when compared to normal epidermis [178]. PKD1 levels and activation status were also increased in a neoplastic mouse keratinocyte cell line [178].

PKD as a Regulator of Angiogenesis Angiogenesis, the formation of new blood capillaries, is an important component of tumor progression [179]. Vascular endothelial growth factor is essential for many angiogenic processes in normal and pathological conditions [180]. For example, the genetic inactivation of the VEGF receptor leads to a complete lack of development of blood vessels in the mouse embryo [181], and inactivation of the VEGF receptor function dramatically impairs the proliferation of endothelial cells in vivo [182].

11

Protein Kinase D Signaling in Cancer

261

PKD is required for VEGF-induced angiogenesis, and PKC and PKD regulate endothelial function at multiple levels. PKC enzymes are involved in the monolayer integrity of the endothelium, and PKCd, for example, protects against barrier dysfunction [183]. Further, PMA- and DAG-induced activation of PKCd and PKD leads to pulmonary microvascular hyperpermeability response across the endothelial monolayer [184]. PKD1 and PKD2 also are required for VEGF-mediated tubulogenesis and endothelial cell migration [185]. This is in contrast to epithelial tumor cell migration, which is inhibited by PKD1 and PKD2. Finally, PKD is also rapidly activated in vascular smooth muscle cells by phorbol esters, angiotensin II, and PDGF, also with a potential function in cell proliferation [186]. PKD activation by VEGF in endothelial cells is mediated through PKCa and PLCg [182]. In HUVECs (human umbilical vein endothelial cells), VEGF stimulates the tyrosine phosphorylation of PKD1 by the VEGF-R2 (KDR), and the knockdown of PKD1 or the expression of a kinase-inactive PKD1 or a mutant, which cannot be phosphorylated at Y463, inhibits VEGF-mediated cell proliferation and migration [187]. In response to VEGF-mediated activation, PKD1 and PKD2 phosphorylate the small heat shock protein Hsp27 at S82. This is independent of the kinase MAPKAPK2, which also can mediate the phosphorylation of Hsp27 at this residue [185]. PKD and its target Hsp27 seem to play major roles in the angiogenic response to VEGF [185].

Perspective Since its first description in 1994, protein kinase D was known to be expressed in a variety of different tumor cell lines. However, till date the roles of PKD isoforms in the different aspects of tumor formation, progression, and metastasis are still illdefined. Most data available focus on the isoform PKD1 and suggest that PKD1 contributes to early events leading to epithelial tumor formation and growth but is also a negative regulator of cell invasion that facilitates metastatic progression. This suggests that PKD1 may act as a switch from an invasive phenotype to the proliferative phenotype. While recently the benzoxoloazepinolone CID755673 was extensively characterized as the first PKD-specific inhibitor [30], a screening for PKD activators to inhibit tumor cell invasion was not advanced so far. Only little is known for the role of the two other PKD isoforms, PKD2 and PKD3, in cancer, and it will be exciting to see if they have similar or converse functions in tumor progression. Due to the fact that PKD1 may promote events leading to tumor formation and tumor growth, but also as a suppressor of metastasis, the treatment of cancer patients with compounds modulating PKD activity needs to be well judged. Patients with tumors at an early stage may be treated with PKD inhibitors, whereas an activation strategy for PKD1 may be more suitable to block metastasis in advanced tumors. An additional problem that may occur is that PKD isoform-specific activators or inhibitors may be needed.

262

P. Storz

Fig. 11.3 PKD1 substrates and their contribution to oncogenic signaling. A subset of so far characterized PKD1 substrates (red), some of the proteins involved in their signaling, as well as the known inducers of PKD1-mediated phosphorylation of these substrates are depicted (PdBu phorbol ester, GF-R growth factor receptor, GPC-R G-protein-coupled receptor). The NF-kB pathway is activated by PKD1 through a so far unidentified substrate or mechanism. Further indicated is the potential role of the PKD1-phosphorylated substrates in tumorigenesis. A role for PKD in mediating tumor cell survival and proliferation is well established; the role of PKD enzymes in cancer cell migration, however, varies with the isoform or cell type studied. Best studied in this context is PKD1 in epithelial cancers, where it may act as a molecular switch that alters the transition from a motile to a proliferative and stress-resistant phenotype

Inhibition of PKD by a variety of inhibitors including the indolocarbazole Gö6976, staurosporine, and K252 was described; however, they all are rather unspecific. For example, Gö6976 inhibits PKD1 with an IC50 of 20 nM [188] but is also a potent inhibitor of cPKC enzymes. The use of such “nonspecific” inhibitors in the PKD literature led to a variety of contradictory results, also due to a lack of some studies to verify the obtained data with more-specific methods, such as a knockdown of PKD. Nevertheless, there are examples for the successful use of broadspectrum agents as inhibitors of PKD-regulated signaling pathways in cancer cells (Fig.11.3). For example, PKD1-mediated signaling to NF-kB, a key event in PKD1 survival signaling, can be blocked by antioxidants such as NAC or resveratrol (trans3,4¢,5-trihydroxystilbene). Resveratrol is a naturally occurring antioxidant that has been shown to function in the prevention of several human pathological processes including carcinogenesis [189–193]. Resveratrol directly inhibits both PKD and its upstream activators nPKC and blocks NF-kB induction [57, 194–197]. In summary, it becomes clear that, to fully understand the functions of PKD enzymes and to further evaluate if they can serve as therapeutic targets for tumor therapy, more knowledge needs to be accumulated regarding their functions in all aspects of tumor biology.

11

Protein Kinase D Signaling in Cancer

263

Acknowledgments Research in the Storz laboratory is supported by grants from the Mayo Clinic SPORE for Pancreatic Cancer (P50 CA102701), the Mayo Clinic Breast Cancer SPORE (CA116201-03DR4), the NIH (GM86435 and CA135102), as well as a Bankhead–Coley grant (10BG11) from the Florida Department of Health.

References 1. Sturany, S., et al., Molecular cloning and characterization of the human protein kinase D2. A novel member of the protein kinase D family of serine threonine kinases. J Biol Chem, 2001. 276(5): p. 3310–8. 2. Johannes, F.J., et al., PKCu is a novel, atypical member of the protein kinase C family. J Biol Chem, 1994. 269(8): p. 6140–8. 3. Hayashi, A., et al., PKCnu, a new member of the protein kinase C family, composes a fourth subfamily with PKCmu. Biochim Biophys Acta, 1999. 1450(1): p. 99–106. 4. Valverde, A.M., et al., Molecular cloning and characterization of protein kinase D: a target for diacylglycerol and phorbol esters with a distinctive catalytic domain. Proc Natl Acad Sci USA, 1994. 91(18): p. 8572–6. 5. Manning, G., et al., The protein kinase complement of the human genome. Science, 2002. 298(5600): p. 1912–34. 6. Dieterich, S., et al., In vitro activation and substrates of recombinant, baculovirus expressed human protein kinase C mu. FEBS Lett, 1996. 381(3): p. 183–7. 7. Eiseler, T., et al., Protein kinase D1 regulates matrix metalloproteinase expression and inhibits breast cancer cell invasion. Breast Cancer Res, 2009. 11(1): p. R13. 8. Kim, M., et al., Epigenetic inactivation of protein kinase D1 in gastric cancer and its role in gastric cancer cell migration and invasion. Carcinogenesis, 2008. 29(3): p. 629–37. 9. Iglesias, T. and E. Rozengurt, Protein kinase D activation by mutations within its pleckstrin homology domain. J Biol Chem, 1998. 273(1): p. 410–6. 10. Iglesias, T. and E. Rozengurt, Protein kinase D activation by deletion of its cysteine-rich motifs. FEBS Lett, 1999. 454(1–2): p. 53–6. 11. Waldron, R.T. and E. Rozengurt, Protein kinase C phosphorylates protein kinase D activation loop Ser744 and Ser748 and releases autoinhibition by the pleckstrin homology domain. J Biol Chem, 2003. 278(1): p. 154–63. 12. Storz, P., et al., Tyrosine phosphorylation of protein kinase D in the pleckstrin homology domain leads to activation. J Biol Chem, 2003. 278(20): p. 17969–76. 13. Cowell, C.F., et al., Mitochondrial diacylglycerol initiates protein-kinase D1-mediated ROS signaling. J Cell Sci, 2009. 122(Pt 7): p. 919–28. 14. Yuan, J., L.W. Slice, and E. Rozengurt, Activation of protein kinase D by signaling through Rho and the alpha subunit of the heterotrimeric G protein G13. J Biol Chem, 2001. 276(42): p. 38619–27. 15. Oancea, E., et al., Mechanism of persistent protein kinase D1 translocation and activation. Dev Cell, 2003. 4(4): p. 561–74. 16. Baron, C.L. and V. Malhotra, Role of diacylglycerol in PKD recruitment to the TGN and protein transport to the plasma membrane. Science, 2002. 295(5553): p. 325–8. 17. Maeda, Y., et al., Recruitment of protein kinase D to the trans-Golgi network via the first cysteine-rich domain. Embo J, 2001. 20(21): p. 5982–90. 18. Rey, O., et al., Regulated nucleocytoplasmic transport of protein kinase D in response to G protein-coupled receptor activation. J Biol Chem, 2001. 276(52): p. 49228–35. 19. Waldron, R.T., T. Iglesias, and E. Rozengurt, The pleckstrin homology domain of protein kinase D interacts preferentially with the eta isoform of protein kinase C. J Biol Chem, 1999. 274(14): p. 9224–30.

264

P. Storz

20. Jamora, C., et al., Gbetagamma-mediated regulation of Golgi organization is through the direct activation of protein kinase D. Cell, 1999. 98(1): p. 59–68. 21. Van Lint, J., et al., Protein kinase D: an intracellular traffic regulator on the move. Trends Cell Biol, 2002. 12(4): p. 193–200. 22. Wang, Q.J., PKD at the crossroads of DAG and PKC signaling. Trends Pharmacol Sci, 2006. 27(6): p. 317–23. 23. Eiseler, T., et al., PKD is recruited to sites of actin remodelling at the leading edge and negatively regulates cell migration. FEBS Lett, 2007. 581(22): p. 4279–87. 24. Cowell, C.F., et al., Loss of cell-cell contacts induces NF-kappaB via RhoA-mediated activation of protein kinase D1. J Cell Biochem, 2009. 106(4): p. 714–28. 25. Eiseler, T., et al., Protein kinase D1 regulates cofilin-mediated F-actin reorganization and cell motility through slingshot. Nat Cell Biol, 2009. 11(5): p. 545–56. 26. Hutti, J.E., et al., A rapid method for determining protein kinase phosphorylation specificity. Nat Methods, 2004. 1(1): p. 27–9. 27. Doppler, H., et al., A phosphorylation state-specific antibody recognizes Hsp27, a novel substrate of protein kinase D. J Biol Chem, 2005. 280(15): p. 15013–9. 28. Chen, J., et al., Protein kinase D3 (PKD3) contributes to prostate cancer cell growth and survival through a PKCepsilon/PKD3 pathway downstream of Akt and ERK 1/2. Cancer Res, 2008. 68(10): p. 3844–53. 29. Chen, J., G. Lu, and Q.J. Wang, Protein kinase C-independent effects of protein kinase D3 in glucose transport in L6 myotubes. Mol Pharmacol, 2005. 67(1): p. 152–62. 30. Sharlow, E.R., et al., Potent and selective disruption of protein kinase D functionality by a benzoxoloazepinolone. J Biol Chem, 2008. 283(48): p. 33516–26. 31. Peterburs, P., et al., Protein kinase D regulates cell migration by direct phosphorylation of the cofilin phosphatase slingshot 1 like. Cancer Res, 2009. 69(14): p. 5634–8. 32. Wang, Q.J., et al., Ligand structure-activity requirements and phospholipid dependence for the binding of phorbol esters to protein kinase D. Mol Pharmacol, 2003. 64(6): p. 1342–8. 33. Yang, C. and M.G. Kazanietz, Divergence and complexities in DAG signaling: looking beyond PKC. Trends Pharmacol Sci, 2003. 24(11): p. 602–8. 34. Paolucci, L. and E. Rozengurt, Protein kinase D in small cell lung cancer cells: rapid activation through protein kinase C. Cancer Res, 1999. 59(3): p. 572–7. 35. Iglesias, T., S. Matthews, and E. Rozengurt, Dissimilar phorbol ester binding properties of the individual cysteine-rich motifs of protein kinase D. FEBS Lett, 1998. 437(1–2): p. 19–23. 36. Zugaza, J.L., et al., Bombesin, vasopressin, endothelin, bradykinin, and platelet-derived growth factor rapidly activate protein kinase D through a protein kinase C-dependent signal transduction pathway. J Biol Chem, 1997. 272(38): p. 23952–60. 37. Van Lint, J., et al., Platelet-derived growth factor stimulates protein kinase D through the activation of phospholipase Cgamma and protein kinase C. J Biol Chem, 1998. 273(12): p. 7038–43. 38. Storz, P. and A. Toker, Protein kinase D mediates a stress-induced NF-kappaB activation and survival pathway. Embo J, 2003. 22(1): p. 109–20. 39. Bagowski, C.P., et al., Cell-type specific phosphorylation of threonines T654 and T669 by PKD defines the signal capacity of the EGF receptor. Embo J, 1999. 18(20): p. 5567–76. 40. Yuan, J., et al., Activation of protein kinase D by signaling through the alpha subunit of the heterotrimeric G protein G(q). J Biol Chem, 2000. 275(3): p. 2157–64. 41. Yuan, J., et al., Cooperation of Gq, Gi, and G12/13 in protein kinase D activation and phosphorylation induced by lysophosphatidic acid. J Biol Chem, 2003. 278(7): p. 4882–91. 42. Yuan, J., O. Rey, and E. Rozengurt, Activation of protein kinase D3 by signaling through Rac and the alpha subunits of the heterotrimeric G proteins G12 and G13. Cell Signal, 2006. 18(7): p. 1051–62. 43. Yuan, J., O. Rey, and E. Rozengurt, Protein kinase D3 activation and phosphorylation by signaling through G alpha q. Biochem Biophys Res Commun, 2005. 335(2): p. 270–6.

11

Protein Kinase D Signaling in Cancer

265

44. Chiu, T. and E. Rozengurt, PKD in intestinal epithelial cells: rapid activation by phorbol esters, LPA, and angiotensin through PKC. Am J Physiol Cell Physiol, 2001. 280(4): p. C929-42. 45. Rey, O., et al., G protein-coupled receptor-mediated phosphorylation of the activation loop of protein kinase D: dependence on plasma membrane translocation and protein kinase Cepsilon. J Biol Chem, 2004. 279(33): p. 34361–72. 46. Kam, Y. and J.H. Exton, Role of phospholipase D in the activation of protein kinase D by lysophosphatidic acid. Biochem Biophys Res Commun, 2004. 315(1): p. 139–43. 47. Chiu, T.T., et al., Protein kinase D2 mediates lysophosphatidic acid-induced interleukin 8 production in nontransformed human colonic epithelial cells through NF-kappaB. Am J Physiol Cell Physiol, 2007. 292(2): p. C767–77. 48. Rozengurt, E. and J.H. Walsh, Gastrin, CCK, signaling, and cancer. Annu Rev Physiol, 2001. 63: p. 49–76. 49. Sturany, S., et al., Mechanism of activation of protein kinase D2(PKD2) by the CCK(B)/gastrin receptor. J Biol Chem, 2002. 277(33): p. 29431–6. 50. Hall, A., Rho GTPases and the actin cytoskeleton. Science, 1998. 279(5350): p. 509–14. 51. Hart, M.J., et al., Direct stimulation of the guanine nucleotide exchange activity of p115 RhoGEF by Galpha13. Science, 1998. 280(5372): p. 2112–4. 52. Kozasa, T., et al., p115 RhoGEF, a GTPase activating protein for Galpha12 and Galpha13. Science, 1998. 280(5372): p. 2109–11. 53. Halliwell, B., Oxidative stress and cancer: have we moved forward? Biochem J, 2007. 401(1): p. 1–11. 54. Storz, P., Reactive oxygen species in tumor progression. Front Biosci, 2005. 10: p. 1881–96. 55. Storz, P., Reactive oxygen species-mediated mitochondria-to-nucleus signaling: a key to aging and radical-caused diseases. Sci STKE, 2006. 2006(332): p. re3. 56. Storz, P., H. Doppler, and A. Toker, Protein kinase Cdelta selectively regulates protein kinase D-dependent activation of NF-kappaB in oxidative stress signaling. Mol Cell Biol, 2004. 24(7): p. 2614–26. 57. Storz, P., H. Doppler, and A. Toker, Activation loop phosphorylation controls protein kinase D-dependent activation of nuclear factor kappaB. Mol Pharmacol, 2004. 66(4): p. 870–9. 58. Storz, P., H. Doppler, and A. Toker, Protein kinase D mediates mitochondrion-to-nucleus signaling and detoxification from mitochondrial reactive oxygen species. Mol Cell Biol, 2005. 25(19): p. 8520–30. 59. Chiarugi, P. and T. Fiaschi, Redox signalling in anchorage-dependent cell growth. Cell Signal, 2007. 19(4): p. 672–82. 60. Rey, O., et al., Protein kinase C nu/protein kinase D3 nuclear localization, catalytic activation, and intracellular redistribution in response to G protein-coupled receptor agonists. J Biol Chem, 2003. 278(26): p. 23773–85. 61. Babior, B.M., The respiratory burst oxidase. Curr Opin Hematol, 1995. 2(1): p. 55–60. 62. Segal, A.W. and K.P. Shatwell, The NADPH oxidase of phagocytic leukocytes. Ann N Y Acad Sci, 1997. 832: p. 215–22. 63. Vantus, T., et al., Doxorubicin-induced activation of protein kinase D1 through caspase-mediated proteolytic cleavage: identification of two cleavage sites by microsequencing. Cell Signal, 2004. 16(6): p. 703–9. 64. Endo, K., et al., Proteolytic cleavage and activation of protein kinase C [micro] by caspase-3 in the apoptotic response of cells to 1-beta -D-arabinofuranosylcytosine and other genotoxic agents. J Biol Chem, 2000. 275(24): p. 18476–81. 65. Haussermann, S., et al., Proteolytic cleavage of protein kinase Cmu upon induction of apoptosis in U937 cells. FEBS Lett, 1999. 462(3): p. 442–6. 66. Waldron, R.T., T. Iglesias, and E. Rozengurt, Phosphorylation-dependent protein kinase D activation. Electrophoresis, 1999. 20(2): p. 382–90. 67. Yuan, J., et al., Protein kinase D is a downstream target of protein kinase Ctheta. Biochem Biophys Res Commun, 2002. 291(3): p. 444–52.

266

P. Storz

68. Brandlin, I., et al., Protein kinase C(mu) regulation of the JNK pathway is triggered via phosphoinositide-dependent kinase 1 and protein kinase C(epsilon). J Biol Chem, 2002. 277(47): p. 45451–7. 69. Doppler, H. and P. Storz, A novel tyrosine phosphorylation site in protein kinase D contributes to oxidative stress-mediated activation. J Biol Chem, 2007. 282(44): p. 31873–81. 70. Brandlin, I., et al., Protein kinase C (PKC)eta-mediated PKC mu activation modulates ERK and JNK signal pathways. J Biol Chem, 2002. 277(8): p. 6490–6. 71. Tan, M., et al., Thrombin rapidly induces protein kinase D phosphorylation, and protein kinase C delta mediates the activation. J Biol Chem, 2003. 278(5): p. 2824–8. 72. Iglesias, T., R.T. Waldron, and E. Rozengurt, Identification of in vivo phosphorylation sites required for protein kinase D activation. J Biol Chem, 1998. 273(42): p. 27662–7. 73. Johnson, L.N., M.E. Noble, and D.J. Owen, Active and inactive protein kinases: structural basis for regulation. Cell, 1996. 85(2): p. 149–58. 74. Mihailovic, T., et al., Protein kinase D2 mediates activation of nuclear factor kappaB by BcrAbl in Bcr-Abl + human myeloid leukemia cells. Cancer Res, 2004. 64(24): p. 8939–44. 75. Rhee, S.G., et al., Hydrogen peroxide: a key messenger that modulates protein phosphorylation through cysteine oxidation. Sci STKE, 2000. 2000(53): p. pe1. 76. Meng, T.C., T. Fukada, and N.K. Tonks, Reversible oxidation and inactivation of protein tyrosine phosphatases in vivo. Mol Cell, 2002. 9(2): p. 387–99. 77. Vertommen, D., et al., Regulation of protein kinase D by multisite phosphorylation. Identification of phosphorylation sites by mass spectrometry and characterization by site-directed mutagenesis. J Biol Chem, 2000. 275(26): p. 19567–76. 78. Hausser, A., et al., Protein kinase C mu is negatively regulated by 14-3-3 signal transduction proteins. J Biol Chem, 1999. 274(14): p. 9258–64. 79. Matthews, S.A., E. Rozengurt, and D. Cantrell, Characterization of serine 916 as an in vivo autophosphorylation site for protein kinase D/Protein kinase Cmu. J Biol Chem, 1999. 274(37): p. 26543–9. 80. Sanchez-Ruiloba, L., et al., Protein kinase D intracellular localization and activity control kinase D-interacting substrate of 220-kDa traffic through a postsynaptic density-95/discs large/zonula occludens-1-binding motif. J Biol Chem, 2006. 281(27): p. 18888–900. 81. Rybin, V.O., J. Guo, and S.F. Steinberg, Protein kinase D1 autophosphorylation via distinct mechanisms at Ser744/Ser748 and Ser916. J Biol Chem, 2009. 284(4): p. 2332–43. 82. Li, J., et al., PKD1, PKD2, and their substrate Kidins220 regulate neurotensin secretion in the BON human endocrine cell line. J Biol Chem, 2008. 283(5): p. 2614–21. 83. White, D.P., P.T. Caswell, and J.C. Norman, alpha v beta3 and alpha5beta1 integrin recycling pathways dictate downstream Rho kinase signaling to regulate persistent cell migration. J Cell Biol, 2007. 177(3): p. 515–25. 84. Eisenberg-Lerner, A. and A. Kimchi, DAP kinase regulates JNK signaling by binding and activating protein kinase D under oxidative stress. Cell Death Differ, 2007. 14(11): p. 1908–15. 85. Rennecke, J., et al., Protein-kinase-Cmu expression correlates with enhanced keratinocyte proliferation in normal and neoplastic mouse epidermis and in cell culture. Int J Cancer, 1999. 80(1): p. 98–103. 86. Trauzold, A., et al., PKCmu prevents CD95-mediated apoptosis and enhances proliferation in pancreatic tumour cells. Oncogene, 2003. 22(55): p. 8939–47. 87. Woodside, K.J., C.M. Townsend, Jr., and B. Mark Evers, Current management of gastrointestinal carcinoid tumors. J Gastrointest Surg, 2004. 8(6): p. 742–56. 88. Jackson, L.N., et al., Overexpression of wild-type PKD2 leads to increased proliferation and invasion of BON endocrine cells. Biochem Biophys Res Commun, 2006. 348(3): p. 945–9. 89. Romero, D.G., et al., Angiotensin II-mediated protein kinase D activation stimulates aldosterone and cortisol secretion in H295R human adrenocortical cells. Endocrinology, 2006. 147(12): p. 6046–55. 90. Rozengurt, E., Early signals in the mitogenic response. Science, 1986. 234(4773): p. 161–6.

11

Protein Kinase D Signaling in Cancer

267

91. Guha, S., O. Rey, and E. Rozengurt, Neurotensin induces protein kinase C-dependent protein kinase D activation and DNA synthesis in human pancreatic carcinoma cell line PANC-1. Cancer Res, 2002. 62(6): p. 1632–40. 92. Rey, O., et al., Rapid protein kinase D translocation in response to G protein-coupled receptor activation. Dependence on protein kinase C. J Biol Chem, 2001. 276(35): p. 32616–26. 93. Haberland, M., R.L. Montgomery, and E.N. Olson, The many roles of histone deacetylases in development and physiology: implications for disease and therapy. Nat Rev Genet, 2009. 10(1): p. 32–42. 94. Xu, X., et al., Angiotensin II stimulates protein kinase D-dependent histone deacetylase 5 phosphorylation and nuclear export leading to vascular smooth muscle cell hypertrophy. Arterioscler Thromb Vasc Biol, 2007. 27(11): p. 2355–62. 95. Vega, R.B., et al., Protein kinases C and D mediate agonist-dependent cardiac hypertrophy through nuclear export of histone deacetylase 5. Mol Cell Biol, 2004. 24(19): p. 8374–85. 96. Matthews, S.A., et al., Essential role for protein kinase D family kinases in the regulation of class II histone deacetylases in B lymphocytes. Mol Cell Biol, 2006. 26(4): p. 1569–77. 97. Huynh, Q.K. and T.A. McKinsey, Protein kinase D directly phosphorylates histone deacetylase 5 via a random sequential kinetic mechanism. Arch Biochem Biophys, 2006. 450(2): p. 141–8. 98. Carnegie, G.K., et al., AKAP-Lbc mobilizes a cardiac hypertrophy signaling pathway. Mol Cell, 2008. 32(2): p. 169–79. 99. Kisfalvi, K., S. Guha, and E. Rozengurt, Neurotensin and EGF induce synergistic stimulation of DNA synthesis by increasing the duration of ERK signaling in ductal pancreatic cancer cells. J Cell Physiol, 2005. 202(3): p. 880–90. 100. Sinnett-Smith, J., et al., Protein kinase D2 potentiates MEK/ERK/RSK signaling, c-Fos accumulation and DNA synthesis induced by bombesin in Swiss 3 T3 cells. J Cell Physiol, 2007. 211(3): p. 781–90. 101. Sinnett-Smith, J., et al., Protein kinase D potentiates DNA synthesis induced by Gq-coupled receptors by increasing the duration of ERK signaling in Swiss 3 T3 cells. J Biol Chem, 2004. 279(16): p. 16883–93. 102. Hausser, A., et al., Protein kinase C mu selectively activates the mitogen-activated protein kinase (MAPK) p42 pathway. FEBS Lett, 2001. 492(1–2): p. 39–44. 103. Perez-Mancera, P.A. and D.A. Tuveson, Physiological analysis of oncogenic K-ras. Methods Enzymol, 2006. 407: p. 676–90. 104. Wang, Y., et al., The RAS effector RIN1 directly competes with RAF and is regulated by 14-3-3 proteins. Mol Cell Biol, 2002. 22(3): p. 916–26. 105. Okada, T., et al., Involvement of N-terminal-extended form of sphingosine kinase 2 in serumdependent regulation of cell proliferation and apoptosis. J Biol Chem, 2005. 280(43): p. 36318–25. 106. Igarashi, N., et al., Sphingosine kinase 2 is a nuclear protein and inhibits DNA synthesis. J Biol Chem, 2003. 278(47): p. 46832–9. 107. Ding, G., et al., Protein kinase D-mediated phosphorylation and nuclear export of sphingosine kinase 2. J Biol Chem, 2007. 282(37): p. 27493–502. 108. Spiegel, S. and S. Milstien, Sphingosine 1-phosphate, a key cell signaling molecule. J Biol Chem, 2002. 277(29): p. 25851–4. 109. Nava, V.E., et al., Sphingosine kinase type 1 promotes estrogen-dependent tumorigenesis of breast cancer MCF-7 cells. Exp Cell Res, 2002. 281(1): p. 115–27. 110. Olivera, A., et al., Sphingosine kinase expression increases intracellular sphingosine-1-phosphate and promotes cell growth and survival. J Cell Biol, 1999. 147(3): p. 545–58. 111. Olivera, A., et al., Sphingosine kinase type 1 induces G12/13-mediated stress fiber formation, yet promotes growth and survival independent of G protein-coupled receptors. J Biol Chem, 2003. 278(47): p. 46452–60. 112. Xia, P., et al., An oncogenic role of sphingosine kinase. Curr Biol, 2000. 10(23): p. 1527–30. 113. Besirli, C.G. and E.M. Johnson, Jr., The activation loop phosphorylation of protein kinase D is an early marker of neuronal DNA damage. J Neurochem, 2006. 99(1): p. 218–25.

268

P. Storz

114. Johannes, F.J., et al., Protein kinase Cmu downregulation of tumor-necrosis-factor-induced apoptosis correlates with enhanced expression of nuclear-factor-kappaB-dependent protective genes. Eur J Biochem, 1998. 257(1): p. 47–54. 115. Storz, P., et al., Functional dichotomy of A20 in apoptotic and necrotic cell death. Biochem J, 2005. 387(Pt 1): p. 47–55. 116. Toker, A., Protein kinases as mediators of phosphoinositide 3-kinase signaling. Mol Pharmacol, 2000. 57(4): p. 652–8. 117. Toker, A. and M. Yoeli-Lerner, Akt signaling and cancer: surviving but not moving on. Cancer Res, 2006. 66(8): p. 3963–6. 118. Brunet, A., et al., Akt promotes cell survival by phosphorylating and inhibiting a Forkhead transcription factor. Cell, 1999. 96(6): p. 857–68. 119. Franke, T.F. and L.C. Cantley, Apoptosis. A Bad kinase makes good. Nature, 1997. 390(6656): p. 116–7. 120. Datta, S.R., A. Brunet, and M.E. Greenberg, Cellular survival: a play in three Akts. Genes Dev, 1999. 13(22): p. 2905–27. 121. Storz, P., Mitochondrial ROS--radical detoxification, mediated by protein kinase D. Trends Cell Biol, 2007. 17(1): p. 13–8. 122. Wang, Y., et al., Hepatocyte resistance to oxidative stress is dependent on protein kinase C-mediated down-regulation of c-Jun/AP-1. J Biol Chem, 2004. 279(30): p. 31089–97. 123. Hurd, C., R.T. Waldron, and E. Rozengurt, Protein kinase D complexes with C-Jun N-terminal kinase via activation loop phosphorylation and phosphorylates the C-Jun N-terminus. Oncogene, 2002. 21(14): p. 2154–60. 124. Hurd, C. and E. Rozengurt, Uncoupling of protein kinase D from suppression of EGFdependent c-Jun phosphorylation in cancer cells. Biochem Biophys Res Commun, 2003. 302(4): p. 800–4. 125. Waldron, R.T., et al., Identification of a novel phosphorylation site in c-Jun directly targeted in vitro by protein kinase D. Biochem Biophys Res Commun, 2007. 356(2): p. 361–7. 126. Zhang, W., et al., Protein kinase D specifically mediates apoptosis signal-regulating kinase 1-JNK signaling induced by H2O2 but not tumor necrosis factor. J Biol Chem, 2005. 280(19): p. 19036–44. 127. MacKeigan, J.P., L.O. Murphy, and J. Blenis, Sensitized RNAi screen of human kinases and phosphatases identifies new regulators of apoptosis and chemoresistance. Nat Cell Biol, 2005. 7(6): p. 591–600. 128. Kops, G.J., et al., Forkhead transcription factor FOXO3a protects quiescent cells from oxidative stress. Nature, 2002. 419(6904): p. 316–21. 129. Mercurio, F. and A.M. Manning, NF-kappaB as a primary regulator of the stress response. Oncogene, 1999. 18(45): p. 6163–71. 130. Li, N. and M. Karin, Is NF-kappaB the sensor of oxidative stress? Faseb J, 1999. 13(10): p. 1137–43. 131. Das, K.C., Y. Lewis-Molock, and C.W. White, Activation of NF-kappa B and elevation of MnSOD gene expression by thiol reducing agents in lung adenocarcinoma (A549) cells. Am J Physiol, 1995. 269(5 Pt 1): p. L588-602. 132. Guo, G., et al., Manganese superoxide dismutase-mediated gene expression in radiation-induced adaptive responses. Mol Cell Biol, 2003. 23(7): p. 2362–78. 133. Rojo, A.I., et al., Regulation of Cu/Zn-superoxide dismutase expression via the phosphatidylinositol 3 kinase/Akt pathway and nuclear factor-kappaB. J Neurosci, 2004. 24(33): p. 7324–34. 134. Olmos, Y., et al., Mutual dependence of Foxo3a and PGC-1alpha in the induction of oxidative stress genes. J Biol Chem, 2009. 284(21): p. 14476–84. 135. Huang, Z., et al., Effects of 5-fluouracil combined with sulfasalazine on human pancreatic carcinoma cell line BxPC-3 proliferation and apoptosis in vitro. Hepatobiliary Pancreat Dis Int, 2007. 6(3): p. 312–20. 136. Sun, C., et al., Aurora kinase inhibition downregulates NF-kappaB and sensitises tumour cells to chemotherapeutic agents. Biochem Biophys Res Commun, 2007. 352(1): p. 220–5.

11

Protein Kinase D Signaling in Cancer

269

137. Rao, P.S., et al., Metallothionein 2A interacts with the kinase domain of PKCmu in prostate cancer. Biochem Biophys Res Commun, 2003. 310(3): p. 1032–8. 138. Abdel-Mageed, A.B. and K.C. Agrawal, Activation of nuclear factor kappaB: potential role in metallothionein-mediated mitogenic response. Cancer Res, 1998. 58(11): p. 2335–8. 139. Vasak, M. and D.W. Hasler, Metallothioneins: new functional and structural insights. Curr Opin Chem Biol, 2000. 4(2): p. 177–83. 140. Mori-Iwamoto, S., et al., Proteomics finding heat shock protein 27 as a biomarker for resistance of pancreatic cancer cells to gemcitabine. Int J Oncol, 2007. 31(6): p. 1345–50. 141. Landry, J., et al., Human HSP27 is phosphorylated at serines 78 and 82 by heat shock and mitogen-activated kinases that recognize the same amino acid motif as S6 kinase II. J Biol Chem, 1992. 267(2): p. 794–803. 142. Lambert, H., et al., HSP27 multimerization mediated by phosphorylation-sensitive intermolecular interactions at the amino terminus. J Biol Chem, 1999. 274(14): p. 9378–85. 143. Yuan, J. and E. Rozengurt, PKD, PKD2, and p38 MAPK mediate Hsp27 serine-82 phosphorylation induced by neurotensin in pancreatic cancer PANC-1 cells. J Cell Biochem, 2008. 103(2): p. 648–62. 144. Rogalla, T., et al., Regulation of Hsp27 oligomerization, chaperone function, and protective activity against oxidative stress/tumor necrosis factor alpha by phosphorylation. J Biol Chem, 1999. 274(27): p. 18947–56. 145. Jaggi, M., et al., Bryostatin 1 modulates beta-catenin subcellular localization and transcription activity through protein kinase D1 activation. Mol Cancer Ther, 2008. 7(9): p. 2703–12. 146. Medeiros, R.B., et al., Protein kinase D1 and the beta 1 integrin cytoplasmic domain control beta 1 integrin function via regulation of Rap1 activation. Immunity, 2005. 23(2): p. 213–26. 147. Lickert, H., et al., Casein kinase II phosphorylation of E-cadherin increases E-cadherin/ beta-catenin interaction and strengthens cell-cell adhesion. J Biol Chem, 2000. 275(7): p. 5090–5. 148. Serres, M., et al., The disruption of adherens junctions is associated with a decrease of E-cadherin phosphorylation by protein kinase CK2. Exp Cell Res, 2000. 257(2): p. 255–64. 149. Jaggi, M., et al., E-cadherin phosphorylation by protein kinase D1/protein kinase C{mu} is associated with altered cellular aggregation and motility in prostate cancer. Cancer Res, 2005. 65(2): p. 483–92. 150. Jaggi, M., et al., Protein kinase C mu is down-regulated in androgen-independent prostate cancer. Biochem Biophys Res Commun, 2003. 307(2): p. 254–60. 151. Syed, V., et al., Beta-catenin mediates alteration in cell proliferation, motility and invasion of prostate cancer cells by differential expression of E-cadherin and protein kinase D1. J Cell Biochem, 2008. 104(1): p. 82–95. 152. Du, C., et al., Protein kinase D1-mediated phosphorylation and subcellular localization of beta-catenin. Cancer Res, 2009. 69(3): p. 1117–24. 153. Vihinen, P., et al., Integrin alpha 2 beta 1 in tumorigenic human osteosarcoma cell lines regulates cell adhesion, migration, and invasion by interaction with type I collagen. Cell Growth Differ, 1996. 7(4): p. 439–47. 154. Palmantier, R., et al., Cis-polyunsaturated fatty acids stimulate beta1 integrin-mediated adhesion of human breast carcinoma cells to type IV collagen by activating protein kinases C-epsilon and -mu. Cancer Res, 2001. 61(6): p. 2445–52. 155. Woods, A.J., et al., PKD1/PKCmu promotes alphavbeta3 integrin recycling and delivery to nascent focal adhesions. Embo J, 2004. 23(13): p. 2531–43. 156. Pollard, T.D. and G.G. Borisy, Cellular motility driven by assembly and disassembly of actin filaments. Cell, 2003. 112(4): p. 453–65. 157. Wang, W., R. Eddy, and J. Condeelis, The cofilin pathway in breast cancer invasion and metastasis. Nat Rev Cancer, 2007. 7(6): p. 429–40. 158. Storz, P., Protein kinase D1: a novel regulator of actin-driven directed cell migration. Cell Cycle, 2009. 8(13): p. 1975–6. 159. Soosairajah, J., et al., Interplay between components of a novel LIM kinase-slingshot phosphatase complex regulates cofilin. Embo J, 2005. 24(3): p. 473–86.

270

P. Storz

160. Nagata-Ohashi, K., et al., A pathway of neuregulin-induced activation of cofilin-phosphatase Slingshot and cofilin in lamellipodia. J Cell Biol, 2004. 165(4): p. 465–71. 161. Janssens, K., et al., Characterization of EVL-I as a protein kinase D substrate. Cell Signal, 2009. 21(2): p. 282–92. 162. De Kimpe, L., et al., Characterization of cortactin as an in vivo protein kinase D substrate: interdependence of sites and potentiation by Src. Cell Signal, 2009. 21(2): p. 253–63. 163. Liljedahl, M., et al., Protein kinase D regulates the fission of cell surface destined transport carriers from the trans-Golgi network. Cell, 2001. 104(3): p. 409–20. 164. Hausser, A., et al., Protein kinase D regulates vesicular transport by phosphorylating and activating phosphatidylinositol-4 kinase IIIbeta at the Golgi complex. Nat Cell Biol, 2005. 7(9): p. 880–6. 165. Ghanekar, Y. and M. Lowe, Protein kinase D: activation for Golgi carrier formation. Trends Cell Biol, 2005. 15(10): p. 511–4. 166. Prigozhina, N.L. and C.M. Waterman-Storer, Protein kinase D-mediated anterograde membrane trafficking is required for fibroblast motility. Curr Biol, 2004. 14(2): p. 88–98. 167. Hegedus, L., et al., Additional MDA-MB-231 breast cancer cell matrix metalloproteinases promote invasiveness. J Cell Physiol, 2008. 216(2): p. 480–5. 168. Hotary, K., et al., A cancer cell metalloprotease triad regulates the basement membrane transmigration program. Genes Dev, 2006. 20(19): p. 2673–86. 169. Ramos-DeSimone, N., et al., Activation of matrix metalloproteinase-9 (MMP-9) via a converging plasmin/stromelysin-1 cascade enhances tumor cell invasion. J Biol Chem, 1999. 274(19): p. 13066–76. 170. Reuben, P.M., Y. Sun, and H.S. Cheung, Basic calcium phosphate crystals activate p44/42 MAPK signal transduction pathway via protein kinase Cmicro in human fibroblasts. J Biol Chem, 2004. 279(34): p. 35719–25. 171. Farina, A.R., et al., Inhibition of human MDA-MB-231 breast cancer cell invasion by matrix metalloproteinase 3 involves degradation of plasminogen. Eur J Biochem, 2002. 269(18): p. 4476–83. 172. Bowden, E.T., et al., An invasion-related complex of cortactin, paxillin and PKCmu associates with invadopodia at sites of extracellular matrix degradation. Oncogene, 1999. 18(31): p. 4440–9. 173. Qiang, Y.W., et al., Insulin-like growth factor I induces migration and invasion of human multiple myeloma cells. Blood, 2004. 103(1): p. 301–8. 174. Hellerstedt, B.A. and K.J. Pienta, The current state of hormonal therapy for prostate cancer. CA Cancer J Clin, 2002. 52(3): p. 154–79. 175. Powell, C.T., et al., Persistent membrane translocation of protein kinase C alpha during 12-0tetradecanoylphorbol-13-acetate-induced apoptosis of LNCaP human prostate cancer cells. Cell Growth Differ, 1996. 7(4): p. 419–28. 176. Mak, P., et al., Protein kinase D1 (PKD1) influences androgen receptor (AR) function in prostate cancer cells. Biochem Biophys Res Commun, 2008. 373(4): p. 618–23. 177. Hassan, S., et al., Heat shock protein 27 mediates repression of androgen receptor function by protein kinase D1 in prostate cancer cells. Oncogene, 2009. 28(49): p. 4386–96. 178. Ristich, V.L., et al., Protein kinase D distribution in normal human epidermis, basal cell carcinoma and psoriasis. Br J Dermatol, 2006. 154(4): p. 586–93. 179. Folkman, J., Angiogenesis in cancer, vascular, rheumatoid and other disease. Nat Med, 1995. 1(1): p. 27–31. 180. Yancopoulos, G.D., et al., Vascular-specific growth factors and blood vessel formation. Nature, 2000. 407(6801): p. 242–8. 181. Carmeliet, P., et al., Abnormal blood vessel development and lethality in embryos lacking a single VEGF allele. Nature, 1996. 380(6573): p. 435–9. 182. Wong, C. and Z.G. Jin, Protein kinase C-dependent protein kinase D activation modulates ERK signal pathway and endothelial cell proliferation by vascular endothelial growth factor. J Biol Chem, 2005. 280(39): p. 33262–9. 183. Harrington, E.O., et al., Role of protein kinase C isoforms in rat epididymal microvascular endothelial barrier function. Am J Respir Cell Mol Biol, 2003. 28(5): p. 626–36.

11

Protein Kinase D Signaling in Cancer

271

184. Tinsley, J.H., N.R. Teasdale, and S.Y. Yuan, Involvement of PKCdelta and PKD in pulmonary microvascular endothelial cell hyperpermeability. Am J Physiol Cell Physiol, 2004. 286(1): p. C105-11. 185. Evans, I.M., G. Britton, and I.C. Zachary, Vascular endothelial growth factor induces heat shock protein (HSP) 27 serine 82 phosphorylation and endothelial tubulogenesis via protein kinase D and independent of p38 kinase. Cell Signal, 2008. 20(7): p. 1375–84. 186. Abedi, H., E. Rozengurt, and I. Zachary, Rapid activation of the novel serine/threonine protein kinase, protein kinase D by phorbol esters, angiotensin II and PDGF-BB in vascular smooth muscle cells. FEBS Lett, 1998. 427(2): p. 209–12. 187. Qin, L., H. Zeng, and D. Zhao, Requirement of protein kinase D tyrosine phosphorylation for VEGF-A165-induced angiogenesis through its interaction and regulation of phospholipase Cgamma phosphorylation. J Biol Chem, 2006. 281(43): p. 32550–8. 188. Gschwendt, M., et al., Inhibition of protein kinase C mu by various inhibitors. Differentiation from protein kinase c isoenzymes. FEBS Lett, 1996. 392(2): p. 77–80. 189. Tinhofer, I., et al., Resveratrol, a tumor-suppressive compound from grapes, induces apoptosis via a novel mitochondrial pathway controlled by Bcl-2. Faseb J, 2001. 15(9): p. 1613–5. 190. Cal, C., et al., Resveratrol and cancer: chemoprevention, apoptosis, and chemo-immunosensitizing activities. Curr Med Chem Anticancer Agents, 2003. 3(2): p. 77–93. 191. Udenigwe, C.C., et al., Potential of resveratrol in anticancer and anti-inflammatory therapy. Nutr Rev, 2008. 66(8): p. 445–54. 192. Kundu, J.K. and Y.J. Surh, Cancer chemopreventive and therapeutic potential of resveratrol: mechanistic perspectives. Cancer Lett, 2008. 269(2): p. 243–61. 193. Goswami, S.K. and D.K. Das, Resveratrol and chemoprevention. Cancer Lett, 2009. 284(1): p. 1–6. 194. Manna, S.K., A. Mukhopadhyay, and B.B. Aggarwal, Resveratrol suppresses TNF-induced activation of nuclear transcription factors NF-kappa B, activator protein-1, and apoptosis: potential role of reactive oxygen intermediates and lipid peroxidation. J Immunol, 2000. 164(12): p. 6509–19. 195. Slater, S.J., et al., Inhibition of protein kinase C by resveratrol. Biochim Biophys Acta, 2003. 1637(1): p. 59–69. 196. Haworth, R.S. and M. Avkiran, Inhibition of protein kinase D by resveratrol. Biochem Pharmacol, 2001. 62(12): p. 1647–51. 197. Stewart, J.R., et al., Resveratrol preferentially inhibits protein kinase C-catalyzed phosphorylation of a cofactor-independent, arginine-rich protein substrate by a novel mechanism. Biochemistry, 1999. 38(40): p. 13244–51.

Chapter 12

Cell Signaling and Cancer: Integrated, Fundamental Approach Involving Electron Transfer, Reactive Oxygen Species, and Antioxidants Peter Kovacic and Ratnasamy Somanathan

Abbreviations AA AO DADs ERK ET GSH HO-1 IL IOA MARK OS PAHs PDT ROS SO SOD UV

Amino acid Antioxidant Dialkyl disulfide Extracellular signal-related kinase Electron transfer Glutathione Heme oxygenase Interleukin Isoobtusilactone Nitrogen-activated protein kinase Oxidative stress Polycyclic aromatic hydrocarbons Photodynamic therapy Reactive oxygen species Superoxide Superoxide dismutase Ultraviolet

Introduction Cell signaling has played an important role in the life sciences. Therefore, it is not surprising to find extensive literature devoted to the cancer area. The present review presents an integrated approach to the topic involving electron transfer (ET),

P. Kovacic (*) Department of Chemistry, San Diego State University, San Diego, CA 92182-1030, USA e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_12, © Springer Science+Business Media, LLC 2012

273

274

P. Kovacic and R. Somanathan

reactive oxygen species (ROS), and antioxidants (AOs). Since carcinogenesis and cancer biochemistry involve a complex area, a broad, encompassing approach dealing with fundamental aspects should provide important insight. Our presentation includes both endogenous and exogenous agents, as well as participation of AOs as preventive agents and chemotherapeutics as curatives. It should be emphasized that our mechanistic approach is just one aspect of a multifaceted subject. Our objective does not include other modes of action. Some original references can be found in cited reviews and books.

Tenets of OS Theory Of the numerous theories that have been advanced, oxidative stress (OS) is the most comprehensive, and has stood the test of time. It can rationalize and correlate most aspects associated with carcinogenesis [1, 2]: (1) generation of ROS from the main classes of carcinogens, in addition to miscellaneous ones; (2) activated oxygen as a universal factor in oncology, (3) DNA cleavage and base oxidation, (4) association with the three major stages of carcinogenesis, (5) relationship to mutagenesis, (6) oncogene activation, (7) tobacco effect, (8) dietary impact, (9) aging factor, (10) association with illness, such as inflammation, (11) correlation with certain hereditary illnesses, (12) estrogen involvement, (13) probability of second cancers, (14) Haddow paradox, (15) beneficial influence of AOs, and (16) application to a wide range of other physiologically active substances. ET and OS have been implicated in the actions of drugs and toxins, such as, carcinogens [2], anti-infective agents [3], anticancer drugs [4], reproductive toxins [5], nephrotoxins [6], hepatotoxins [7], nerve toxins [8], cardiovascular toxins [9], mitochondrial toxins [10], abused drugs [11], ototoxins [12], immune system toxins [13], and various others, in addition to human illnesses [14]. The preponderance of bioactive substances or their metabolites incorporate ET functionalities, which, we believe, play important roles in physiological responses [1]. These main groups include quinones (or phenolic precursors), metal complexes (or complexors), aromatic nitro compounds (or reduced derivatives), and conjugated imines (or iminium species). In vivo redox cycling with oxygen can occur, giving rise to OS through generation of ROS, as discussed in the Metabolism section. In some cases, ET results in interference with normal electron transport chains, e.g., respiration. Alternatively, ROS can arise in some instances by non-ET avenues. Generally, active entities possessing ET groups display reduction potentials in the physiologically responsive range, i.e., more positive than −0.5 V. However, a correlation between reduction potential and activity is not always observed since important roles are played by other factors, such as, solubility, metabolism, diffusion, adsorption, site binding, cell permeability, and stereochemistry. Reduction potential is influenced by various factors including conformation which can differ in vitro vs. in vivo. Hence, electrochemistry, which has enjoyed relatively little

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

275

attention, can provide valuable insight into the mode of action. Our theoretical framework incorporates several features common to most carcinogens: 1. Binding to DNA by alkylation or complexation 2. Existence of an ET entity present in the parent carcinogen or frequently in a metabolite 3. Formation of ROS usually by ET involving oxygen 4. ROS generation in close proximity to DNA giving rise to mutation, apparently by strand cleavage and base oxidation There is a plethora of experimental evidence supporting the OS theoretical framework, including generation of the common ROS, lipoperoxidation, degradation products of oxidation, depletion of AOs, effect of exogenous AOs, DNA oxidation and cleavage products, as well as electrochemical data. This comprehensive, unifying mechanism is in keeping with the frequent observations that many ET substances display a variety of activities, e.g., multiple drug properties, as well as toxic effects. Although our focus is on this theory, it should be emphasized that bioactivity is quite complicated. Other well-supported, general proposals include the action of oncogenes. Evidence indicates that OS plays a role in a number of the alternative hypotheses. The most likely scenario is complementarity entailing a multifaceted approach.

Chronology The fundamental framework of the OS theory was laid down by four groups in the 1950s [1]. It is remarkable that the essential features still provide a useful base even though the experimental, supporting evidence at the time was quite limited. The unifying theme is in marked contrast to the approach, still prevalent, that each type of cancer has a different cause. Several corollaries were advanced, including protection by antioxidants and application to anticancer agents and aging. Though little attention was paid to the theory in the ensuing 10–15 years, a surge of activity began in the 1970s. Despite the mounting volume of research, the OS theory was largely ignored by most oncologists until recent years.

Stages in Carcinogenesis Carcinogenesis is a complex process characterized by three major stages: initiation, promotion, and progression [1]. Complete carcinogens are both initiators and promoters, whereas incomplete carcinogens require promoters to produce malignant transformations. Most carcinogens must be converted metabolically into an active form, i.e., transformation from a pro-carcinogen into an ultimate carcinogen. Our review is primarily concerned with initiation, the first stage, which is the most studied and best understood. Interaction of the ultimate carcinogen with DNA results in

276

P. Kovacic and R. Somanathan

irreversible alterations, i.e., mutations. Many investigators have pointed out an unmistakable correlation between carcinogenesis and mutagenesis, indicating that the former entails a type of mutation. Extensive evidence is documented which indicates that ROS species are pervasive mutagens. Numerous antecedent studies have demonstrated that ROS participate in all three stages. In addition to the modes addressed in this review, ROS can also interfere with cell signaling by altering protein kinase cascades and transcription factors, ultimately leading to tumor development [13]. Promotion, which follows initiation, is a multistage, often reversible, process of gene activation. There is convincing evidence that cellular oxidation states, i.e., the relative levels of ROS, AO defense entities, and radical scavengers, can promote initiated cells to neoplastic growth. Investigations dealing with involvement of ROS have continued into more recent years. One of the most powerful tumor promoters is phorbol myristate acetate. It is well established that its activation is accompanied by appearance of ROS, including superoxide (SO) and hydrogen peroxide, as well as hydroxyl radicals and lipid peroxidation. Phagocytic stimulation also plays a role. It is not surprising that cancer cells, which display no specialized function within the body, appear to represent a simpler type. Reversion to unicellular behavior and to a fetal phenotype have been discussed. Another major difference with normal cells concerns telomerase. Cellular immortality, a hallmark of cancer, involves this enzyme at various stages along the continuum of multistage carcinogenesis. The catalyst rebuilds telomeres, the termini on chromosomes. Telomerase may have potential as an early biomarker or, perhaps, as a player in the control of cancer.

Cell Signaling Cell signaling is known to be importantly involved in various aspects of biological function, including normal processes, therapeutic drug action, and toxicology. More than 10 years ago, ROS attracted attention in relation to cell signaling. Since then several books [15, 16] and a book chapter [17] have addressed this aspect. A recent review has provided further insight [18]. Evidence has accumulated that ROS, such as hydrogen peroxide, SO, and the hydroxyl radical, are important chemical mediators that regulate the transduction of signals by modulating protein activity via redox chemistry. Authors have proposed that ROS have been conserved throughout evolution as universal second messengers. Nearly every step in signal transfer is sensitive to ROS, which can function as second messengers in the activation of transcription factors. Various types of radiation, which are generators of ROS, also influence cell communication. Since the messengers must possess appreciable lifetimes in order to migrate, a certain degree of stability is required. For example, the hydroxyl radical would not be a messenger due to its extremely high reactivity with its resultant very short time of existence, although it would generate messenger radicals. Likely candidates include superoxide and resonance-stabilized peroxyl radicals. Others would be stable ROS arising from AOs, such as vitamin C, vitamin E, and flavonoids. Members can be envisioned from the reactive nitrogen species (RNS)

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

277

category. NO is a well-known radical that plays an important role in cell signaling. In relation to theory, a puzzling aspect is its relatively short life time. Perhaps more stable complexes are involved. Other nitrogen radicals can play a role. Important candidates comprise small proteins possessing redox groups. In effect, cell signaling can be regarded as proceeding via a long redox chain in which the standard parameters of initiation, propagation, and termination pertain, involving omnipresent conduit species with unshared electrons. A series of relay stations may be operative. Based on the redox chain framework, the second messenger might be superoxide formed by a redox process involving oxygen and a second messenger electron from an ET functionality in the receptor site. At the termination of the initial journey, radical character would be transmitted to a site, mobile or stationary. For example, a redox amino acid (AA) side chain acting as a relay (transfer station), that could then pass on (initiate) radical character to a third messenger. These types of interactions, widespread in AA chemistry, usually involving electron and/or hydrogen abstraction to generate radical species, are treated in a review [19]. The numerous redox moieties in anchored proteins might fall in the relay category. There has been dramatic increase in attention devoted to free-radical species in cell signaling, although the bulk of the signal transduction literature pays no attention to this aspect. Certain second messengers, most notably calcium ions and phosphoinositides, are also charged [20]. Therefore, electrostatic interactions may play a role in their biological activities.

Integrated Approach Endogenous Agents More recent years have shown an escalation in research dealing with an integrated approach to cancer in relation to ROS, OS, AOs, receptors, and cell signaling. Various other aspects of oncology are also addressed. There are articles that deal generally with the integrated approach of the present review [21–26]. There are many reports that address more specific aspects as presented in the ensuing material. ROS generation is a tightly controlled process that plays a central role in cell signaling [27]. Regulated changes in intracellular ROS levels can induce signaling events that control cellular functions, such as proliferation and apoptosis that are involved in carcinogenesis. The review summarizes ROS-mediated cell signaling and its relation to cancer induction. Special attention is given to redox-active iron in relation to ROS-mediated signaling in cancer development. Elucidation of molecular mechanisms that govern ROS-mediated regulation of cell signaling may lead to therapeutic strategies for cancer treatment and prevention. When redox homeostasis is disturbed, OS may contribute to disease development, including cancer [28]. The review focuses on the role of key transcription

278

P. Kovacic and R. Somanathan

factors and signal-transduction pathways in affecting cell survival, as well as the redox systems that regulate functions. How redox systems contribute to the development of cancer is discussed, including strategies for treatment. Nox and Duox enzymes generate ROS as part of normal and abnormal processes in vivo, including signal transduction [29]. The most common conditions associated with Nox-derived ROS are chronic diseases that commonly occur late in life, such as cancer. The pathological role of Nox enzymes might be understood in terms of negative pleiotropy, involving genes that are beneficial early in life, but have harmful effects later. Another report also deals with the Nox category [30]. When cells are stimulated by ROS, cell-signaling cascades are activated [31]. The cellular redox potential is an important determinant of cell function, and interruption of redox balance may have adverse effects. AOs may intercept critical ROS involved in signaling, and thus both protect against and foster pathogenesis. Further study is needed to unravel the role of ROS in redox regulation, and the potential outcome of AO use on cellular responses. Gamma-glutamyltransferase (GGT) plays an important role in the homeostasis of GSH [32]. Expression of GGT is upregulated after OS. In a study of signal transduction effects, OS-induced activation of GGT was found to involve Eas and several downstream signaling pathways in rat colon carcinoma cells. Glucose deprivation-induced cell death is associated with apoptosis which is characterized by membrane blebbing in human breast cancer cells [33]. Results suggest that the alteration of phosphorylation/dephosphorylation by paxillin may be related to events mediated by OS and the stress-activated protein kinase signaling pathway. Overexpression of catalase, but not Mn SOD, inhibited glucose deprivation-induced cytotoxicity and c-Jun-N-terminal kinase activation in human adrenocarcinoma cells [34]. Hydrogen peroxide, rather than superoxide, acts as a second messenger of metabolic OS in activation of certain signal transduction pathways. Production of ROS by oncogenic Ras is thought to be crucial, but little is known about the signaling mechanism involved [35]. Overexpression of a leukotriene receptor caused increased ROS production. An associated cascade appears to be involved. In a study of signaling mechanism by ROS in tumor progression, ROS, as a signaling messenger, oxidizes critical target molecules, such as PKC and protein tyrosine phosphates [36]. Two of the downstream entities regulated by ROS are MAPK and PAK. MAPK cascades comprise a major signal pathway for metastasis, which are mediated by PKC. Hydrogen peroxide plays a role in upregulation via an AO-response element. Understanding the critical role of ROS-triggered signaling transduction, transcriptional activation and regulation of gene ROS in tumor progression is helpful. Redox modification of proteins permits regulation of signaling pathways as in cancer cells [37]. ROS can modify signaling proteins through thiol oxidation and nitrosylation. These modifications modulate protein activity and are important in signaling events, especially those pertaining to cell death or survival. NF-kB has been implicated in various events leading to cancer promotion and this family also regulates the expression of many genes [38]. The review summarizes features of regulation, including signaling pathways by kinases. Focus is also on the role of NF-kB in cell survival and OS. There are related reports [39, 40].

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

279

Identifying target proteins undergoing redox modifications is key to understanding how oxidants mediate pathological processes, such as tumor promotion [41]. These proteins may also be targets for chemopreventive AOs which can block signaling induced by oxidants. PKS is a logical candidate for redox modification by ROS and AOs that may play a role in cancer-promoting and anticancer activity, respectively. PKCs contain structural features readily susceptible to oxidative modification, namely thiol residues. Mitochondria play roles in many cellular functions, including energy production and cell signaling [42]. The report addresses the role of mitochondrial proteins in signaling pathways, such as those dealing with OS and apoptosis. These proteins are modified through redox processes in various age-related diseases, e.g., cancer and Parkinson’s disease. There are related reviews that deal with ROS, cell signaling, and mitochondria [43–45]. A review discusses interplay between integrin-mediated ROS production and matrix metalloprotein expression in relation to cancer cell invasion and metastasis [46]. Increased levels of ROS from mitochondria have been observed in many cancer cells [47]. ROS derived from a novel protein (Romo 1) are indispensable for cancer cell proliferation and many play an important role in redox signaling in cancer cells. A comprehensive review addresses redox-dependent regulation of cell signaling through Cys oxidation in protein tyrosine phosphatase [48, 49]. ROS can stimulate cancer cell growth by regulating AMPK and COX-2 [50]. There is a correlation between H. pylori infection and gastric carcinogenesis [51–53]. Signal transduction is involved, as well as cell proliferation, apoptosis and production of ROS. As is well characterized, ROS are associated with tumor formation, with involvement as the second messenger in signaling cascades [54]. The study observed increased ROS formation in cancer cells in comparison with normal ones. Kupffer cells appear to play a central role in the hepatic response to carcinogenic agents [55]. Activation results in the release of cytokines and/or ROS that induce hepatocyte cell proliferation. ROS and RNS are mediators of cell signaling in pulmonary epithelial cells by asbestos, silica airborne particulates, diesel exhaust, ozone, and cigarette smoke [56]. Targeting of MARK and related signaling cascades may be critical to the prevention of lung cancer and control of a number of lung diseases. Interleukin (IL) plays an important role in modulating expression of various growth and angiogenic factors in tumor cells [57]. Exogenous hydrogen peroxide enhanced IL secretion and thiol prevented IL-induced ROS production. Various signaling pathways are involved in IL-induced secretion which enhance endothelial cell proliferation. The signaling adapter p62 is induced by Ras with increased levels in human tumors [58]. Its deficiency produces increased ROS which accounts for enhanced cell death. High levels of ROS are spontaneously produced by ovarian and prostate cancer cells [59], and the elevated levels were inhibited by NADPH oxidase inhibitor and mitochondrial ET chain inhibition. Evidence indicates that ROS play an important role in inducing angiogenesis and tumor growth. JNK was found to potentiate tumor necrosis factor (TNF)-stimulated necrosis [60]. Data indicate that JNK can shift the balance of TNF-stimulated cell

280

P. Kovacic and R. Somanathan

death from apoptosis to necrosis. TNF is an important regulator of immune responses, including signal pathways. ROS are generated in response to inhibition of protein phosphatase 2A (PP2A) in leukemia cells [61]. A differential role is indicated for ROS in apoptosis caused by PP2A inhibition. ROS generated by tumor promoter TPA play an important role in mediating cell signaling for regulation of gene expression [62]. Scavengers of ROS, such as SOD, catalase and thiol, suppressed TPA-triggered migration of human hepatoma cells. TPA can induce gene expression in a ROS-dependent manner. ROS produced by NADPH mediate the antiapoptotic effect of growth factors [63]. Results suggest a novel signaling pathway in which NADPH oxidase activation results in inhibition of protein tyrosine phosphatases, leading to enhanced phosphorylation of kinases and suppression of apoptosis. Bile reflux contributes to esophagal injury and neoplasia, in which COX-2 is involved [64]. Bile acids stimulated COX-2 expression and induced phosphorylation. ROS-mediated activation of signaling pathways occurred. Hypoxia is closely related to pathophysiological conditions, such as cancer, in which ROS have been implicated [65]. In this connection, the role of ROS in cell signaling was investigated. A novel role was found for adenosine monophosphate-activated protein kinase (AMPK) and the upstream signaling components. Regulation of hypoxiainducible factor is under the control of c-Jun N-terminal kinase and Janus kinase 2 pathways. Thus, AMPK is a key determinant of HIF-1 functions in response to ROS. Bilirubin produces major biological effects, such as eliciting cell toxicity and inhibiting proliferation in cancer cells [66]. It induces OS by promoting an increase in ROS. The ROS increase activated the AO cell response through a master redox regulator in eukaryotic cells. The activation was followed by concomitant activation of transcriptional factor and upregulation of tumor suppressor. Blocking ROS generation with N-acetylcysteine pretreatment avoided the adverse effects. In a related study, bile acids initiated signaling pathways via COX-2 induction with participation of ROS [67]. Understanding is increased of the mechanisms by which bile acids promote development of esophageal adenocarcinoma. Tumor suppressor gene PTEN functions as a lipid phosphatase, thereby regulating certain signaling pathways [64]. OS regulates accumulation of PTEN which reduces tumor progression and protects cells upon oxidative damage. TNFa induces autophagy which requires ROS production with participation in the apoptotic signaling pathway [68]. NF-kB activation mediates the repression of autophagy in response to TNFa. Investigations address the role of NOX in cancer. A review addresses involvement of ROS and NOX in tumor angiogenesis through the regulation of different biological systems [69]. Signaling pathways influenced by ROS and NOX involve proliferation and angiogenesis. ROS targets regulating proliferation include phosphatases, AP1 and NF-kB, as well as cell cycle targets. There is a mediating role for NOX redox signaling for Ras oncogene-induced changes [70]. Hypoxia-inducible factor-1 (HIF-1) represents the key mediator of hypoxia response [71]. HIF-1 overexpression is linked to tumor initiation and progression. Elevated ROS, observed in the tumors, have been implicated in HIF-1 signaling.

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

281

Various reports deal with the effects of photodynamic therapy (PDT). The procedure entails binding of a photosensitizer to the target tumor tissue, which upon light exposure generates ROS including singlet oxygen [72]: cancer cells reacted to PDT by covalent cross-linking of signal transducer and activator of transcriptor 3 (STAT3). Multiple signaling cascades are activated in cancer cells exposed to PDT [73]. These signals are transduced into cell death responses involving ROS. The killing can be direct by induction of apoptotic, as well non-apoptotic, pathways. A study deals with mitogen-activated protein kinase (MARK) signaling after PDT [74]. Both extracellular signal-related kinase (ERK) and p38 were activated. There was photochemically induction of ERK phosphorylation. Also, with PDT treated cancer cells, signaling pathways and mechanisms were characterized that lead to up-regulation of the AO enzyme heme oxygenase (HO-1) [75]. The p38MARK and P13K signaling cascade is required for HO-1 induction. Stimulation of HO-1 expression is a cytoprotective mechanism governed by the signaling pathways. Several investigations dealt with the role of thiols. ROS are able to modify signaling proteins through thiol oxidation [76]. Protein thiol modifications can control signal transduction effectors that include protein kinases, phosphatases and transcription factors. Related studies involve redox regulation of thiol dependent signaling pathways in cancer [77, 78]. MEK inhibitors markedly enhanced the efficacy of histone deacetylase inhibitors to induce generation of ROS and apoptosis in certain tumor cells. [79]. Results suggest that ROS induced by luteolin, a flavonoid, play a pivotal role in suppression of NF-kB and potentiation pf JNK in sensitization of cancer cells to TNF-induced apoptosis [80]. ROS promote squamous cell carcinoma by the ability to modulate intracellular signaling and to affect gene expression [81]. Both ROS and RNS activated NF-kB. Cancer may be regarded as the end result of defects in cellular signaling processes that play a key role. A report identifies signal transduction molecules as targets for cancer prevention [82]. The fusion gene BCR-ABL is known to activate several signaling pathways that promote cell proliferation in chronic myeloid leukemia [83].

Exogenous Agents The integrated approach is also applicable to a wide variety of exogenous agents. Metals and Particulates This class is known for toxicity and carcinogenesis as shown in the various reviews cited in the Introduction. The theme of ET-OS-ROS has enjoyed widespread mechanistic support. With regard to electrochemistry, the reduction potentials of heavier metals are generally in the range amenable to ET in biosystems. Metal ions influence

282

P. Kovacic and R. Somanathan

a variety of signaling processes [84]. Examples of metals relevant to the present approach are presented. Numerous metals are reported as carcinogens [85]. Underlying mechanisms have been reviewed including involvement of free radicals and signaling pathways. The following articles deal with participation of ROS and signal transduction for a number of carcinogenic metal compounds, including those of As [86–89], Cr [90–92], Ca [93, 94], Cd [95, 96], Se [95] and Ni [90]. In relation to carcinogenesis, an important role of free radicals from fibers is discussed, as well as modulation of genes and transcription factors caused by OS [97]. Cell signaling pathways elicited by asbestos are addressed involving stimulation of signaling by ROS generated via phagocytosis or redox reactions on the mineral surface [98]. Metals also play important roles in the physiological effects of particulate matter, such as asbestos and silica as discussed in the reviews on carcinogenesis and toxicity in the Introduction. Inhalation of silica is associated with various pathologies, including cancer, inflammation, and autoimmune disorders [99]. Several rationales have been developed in relation to toxicity, e.g., production of ROS, either directly or indirectly, triggering cell signaling pathways that initiate cytokine release and apoptosis. These processes are described as a function of receptor-mediated signaling. With murine macrophages, RNS are also generated in the initial respiratory burst. Similar results were obtained in another study in relation to ROS and signaling pathways [100].

Ultraviolet (UV) Radiation Increased UV irradiation resulting from thinner ozone layers results in enhanced rate of skin cancer [101]. An essential signal pathway entails activation of several growth factor receptors, including that for tyrosine phosphorylation. The radiation can generate hydroxyl radicals by hemolytic cleavage with subsequent attack of DNA, and dimerization of pyrimidine bases resulting in DNA damage [1]. Protection is provided by various skin and exogeneous AOs. UV radiation is responsible for many skin pathologies, including cancer, inflammation and photoaging [102]. These effects are associated with activation of many signal transduction pathways, e.g., MAPK and NF-kB, and also with alteration of gene expression. UV exposure also activates various receptors through the production of ROS. The role of a protooncogene is treated.

Hydrogen Peroxide Cervical cancer, a common one among females, is associated with ROS and inflammation [103, 104]. ROS in turn affects the expression of pro- and anti- apoptotic proteins. Apoptosis by H2O2 is triggered via the mitochondrial pathway involving upregulation and downregulation of various signal pathways. An overview deals with hydrogen peroxide production and signaling actions in relation to cancer [105].

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

283

Alcohol Alcohol metabolism in the liver can lead to cirrhosis, the common precursor of hepatic cancer [106]. The metabolism generates acetaldehyde and ROS that react readily with numerous cell targets, including components of cell signaling pathways and DNA. In the process, glutathione (GSH) is depleted. Induction of cytochrome P450 causes further free radical production. Involvement of OS has led to renewed interest in the use of AOs for prevention. The topic has been addressed in detail in a recent review [107].

Ginkgo Biloba Ginkgolide B, the major active component of Ginkgo biloba extracts, can both stimulate and inhibit apoptotic signaling in cancer cells [108]. The agent can increase the level of ROS, leading to elevations of Ca and NO and to signaling events. A model was produced for the induced cell signaling cascade in the tumor cells.

Curcumin A report shows gene upregulation by curcumin in colon cancer cells [109]. There is involvement of GSH and sulfhydryl enzymes associated with signal transduction in mediating the effect. The gene upregulation was attenuated by GSH or N-acetylcysteine. Curcumin increased ROS, decreased GSH, and activated sulfhydryl enzymes involved in signal transduction linked to gene expression. The findings suggest that a regulatory thiol redox-sensitive signaling cascade exists leading to induction of gene expression.

Endosulfan The organochlorine insecticide is a potential carcinogen in humans, which alters MAK kinase signaling pathways and may affect cell growth and differentiation in human keratinocytes [110]. The toxicant induces ROS generation leading to sustained kinase phosphorylation.

Benzene The pollutant is a leukemogen that forms metabolites capable of generating ROS [111]. Active metabolites are benzoquinone and hydroquinone which can undergo redox cycling. The toxicity may be mediated through alterations by ROS on the c-MYB signaling pathway.

284

P. Kovacic and R. Somanathan

Polycyclic Aromatic Hydrocarbons (PAHs) PAHs are known mammary carcinogens that can mimic growth factor signaling and increase cell proliferation [112]. Quinones are metabolites that are associated with production of ROS, such as superoxide and hydrogen peroxide. Data indicate that the quinones, through the generation of ROS, activate the epidermal growth factor receptor (EGFR) leading to cell proliferation. The signaling mechanism of EGFR involves generation of hydrogen peroxide. ROS signaling often results in phosphorylation and activation of growth factor receptor pathways.

Estrogen Quinone The highly redox active catechol quinone metabolites affect estrogen-sensitive genes where large amounts of ROS are generated causing DNA damage [113]. A major pathway is the hormonal one by which estrogen stimulates cell proliferation through receptor-mediated signaling, thus resulting in an increased risk of genomic mutations during DNA replication. Extranuclear estrogen signaling pathways also appear to be involved.

Antioxidants There is a large body of evidence that supports participation of ROS in the action of carcinogens [1]. Hence, it is reasonable to consider AOs as preventive agents which also appears plausible based on numerous literature reports [2]. Herein are provided additional examples, including involvement of cell signaling. OS is increasingly associated with pathogenesis of various disorders, including cancer [114]. Fruits and vegetables have the capacity to reduce the incidence of cancer, apparently through the AO properties of constituents. However, reports suggest that the chemopreventive effects may also be ascribed to their ability to modulate components of cell signaling pathways. Focus is on signal transduction mechanisms and prevention of gap junction intercellular communication. Apple peels are known to inhibit tumor cells proliferation [115]. Peel extract may prevent carcinogenesis and associated cell signaling, suggesting that the favorable effects may operate through the AO properties by blocking ROS-mediated cell signaling activation. Carnosol, a phytopolyphenol in rosemary, functions as an AO and as a chemopreventive agent against cancer [116]. The compound scavenged free radicals and protected DNA against oxidative attack. Carnosol inhibited NF-kB activation and exhibited various influences on cell signaling. Oxidative stress and inflammation are features of smoking-related disorders, such as cancer [117]. Flavonoids may be suitable prophylactic agents due to their AO properties. Thus, OS from smoking could be attenuated by the AO compounds, perhaps linked to a genetic polymorphism and modulation of complex cell signaling cascades.

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

285

Blackberries are a rich source of flavonoids which can inhibit carcinogenesis [118]. The extract is an effective scavenger of free radicals and inhibitor of 8-OH-dG formation. The chemopreventive effects may function through the AO properties by blocking ROS-mediated protein kinase activation. There are related reports involving flavonoids [103, 119–121]. Other compounds in the category are the following: tocotrienol [122] and general [123–130]. Various thiol containing AOs also play roles in prevention of toxicity. An important in vivo member is GSH which targets harmful ROS [131]. Links between the favorable AO properties of GSH and diseases, such as cancer, neurodegeneration, cystic fibrosis, HIV and aging have been demonstrated. Expression of enzymes involved in GSH homeostasis influences susceptibility and progression of these conditions. The review provides an overview of the importance of GSH. N-acetylcysteine (NAC) decreases glioma cell proliferation [132], as well as intracellular oxidants was observed. NAC also lowers Akt activity, signal-regulated kinase 1/2 are the redox-sensitive transcriptor factor NF-kB, with relationship to protein kinase C activity. There is a related study involving NAC [133]. Superoxide, produced in proper amounts, is a normal and useful metabolite, serving important roles, e.g., as a signaling agent [134]. However, in excess, the radical anion can initiate harmful oxidation of lipids, protein, and DNA, leading to cell death. The role of SOD is to help correct this imbalance in superoxide. Numerous studies in animals and humans demonstrate the chemopreventive effects of Se, presumably acting as an AO [135]. Evidence indicates that the methylated metabolite plays an important role. A mechanism is proposed entailing Se catalysis of reversible Cys/disulfide transformations that occur in a number of redoxregulated proteins, including transcription factors. Thioredoxin reductase is a selenoprotein that exhibits increased activity with Se supplementation. Enhanced levels of the enzyme could have beneficial effects in lowering OS and in regulating cell signal pathways. There is another relevant report on beneficial effects of Se [136].

Chemotherapeutic Agents and Preventions Because of the serious and resistant nature of the cancer problem, much research has been devoted to curative agents, including the following examples. These substances can also be included into the mechanistic framework of ET-ROS-OS. One should distinguish between agents that prevent cancer and those that destroy malignant cells.

Aminoflavone Aminoflavone, a novel anticancer agent, induces DNA-protein cross-links, phosphorylation, aryl hydrocarbon receptor signaling, and apoptosis [137]. The drug caused oxidative DNA damage. The AO N-acetyl-L-cysteine attenuated the

286

P. Kovacic and R. Somanathan

cytotoxic effects. Caspase activation was brought about. Inhibition of cancer cells apparently entails ROS production, oxidative DNA damage and apoptosis.

Curcumin Curcumin exhibits anticancer activity and triggers tumor cell apoptosis which is associated with ROS production and OS [138–140]. Mitochondrial respiration and redox tone are pivotal determinants in the apoptosis signaling.

Radiation Exposure of tumor cells to ionizing radiation causes DNA damage and ROS generation [141]. Radiation induces ROS-dependent activation of tyrosine kinases leading to activation of multiple downstream signaling pathways. There are other related investigations [142, 143].

Nitric Oxide Nitric oxide generates a state of OS which affects several redox-sensitive signaling pathways, leading to the elimination of neoplastic cells via apoptosis [144]. The gas generates ROS and modulates several signaling cascades [78, 145]. Insight into mechanisms involved in ROS signaling may offer novel avenues to facilitate discovery of cancer-specific therapies [125].

Resveratrol Resveratrol possesses therapeutic effects for various cancers [146]. The mode of action entails the ability to control intracellular signaling cascades of protein kinase responsible for inducing apoptosis, as well as induction of ROS.

Polyphenols Although polyphenols are reported to prevent cancer via AO action (see AO section), they are also reported to be chemotherapeutic agents [147]. The anti-tumor effects are mediated, at least in part, by protein kinases and downstream pathways of signal transduction. Apoptosis induction involves ROS, particularly hydrogen peroxide, and inactivation of the telomerase enzyme. ROS modulate the activity of regulating kinases and transcription factors. Epigallocatechin-3-gallate, a tea phenol, suppresses cancer cell proliferation, induces apoptosis, may increase the efficacy of chemotherapy and is involved with

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

287

several signaling pathways [148]. The drug may exert cell cytotoxicity through modulating AMPK followed by decrease in COX-2 expression. ROS are an upstream signal of AMPK. As discussed elsewhere in this review, polyphenols can generate ET quinones under the appropriate conditions. A related study also involves tea polyphenols [149]. Action of classical AOs as pro-oxidants, including mechanism, has been addressed [107]. Formaldehyde Formaldehyde releasing prodrugs diminish the level of GSH resulting in increase of ROS followed by signaling events that lead to cancer cell death [150]. Thiols, such as GSH and N-acetlycysteine, protected cancer cells from death induced by formaldehyde-releasing prodrugs. Isoobtusilactone (IOA) IOA exhibited effective cell growth inhibition by inducing cancer cells to undergo phase arrest and apoptosis [151]. Generation of ROS is a critical mediator of cell growth inhibition. Enhancement of ROS activated apoptosis signal-regulating kinase, which in turn resulted in activation of other cell signaling events. AOs significantly decreased apoptosis. IOA enhanced phosphorylation, triggered the mitochondrial apoptotic pathway, and induced mitochondrial membrane potential loss. Genipin This aglycon of geniposide, induces apoptotic cell death in human hepatoma cells [152]. Apparently, the mode of action entails activation of the mitochondrial pathway via NADPH oxidase generated ROS and JNK dependent cell signaling. Decursin Decursin, an herbal pyranocoumarin, exerts a long-lasting inhibition of androgen receptor-mediated signaling which is crucial for prostate cancer development [153]. Also, there is growth arrest, apoptotic effects, and generation of ROS. Quinoline Quinone A quinolinequinone induced growth arrest and apoptosis of renal carcinoma cells [154]. Apoptosis occurred via dephosphorylation of kinases. Various signaling events were associated with ROS formation. Thiol AO opposed apoptosis. The quinone functionality is a well-known generator of ROS by ET.

288

P. Kovacic and R. Somanathan

Acacetin Acacetin induces apoptosis in human breast cancer and also causes growth inhibition [155]. There was activation of caspase-7 and enhanced ROS production which was reduced by thiol AO. Induction of signaling pathways occurred. The compound is a flavonoid which is commonly regarded as an AO. However, under the appropriate conditions, pro-oxidant effects can occur, e.g., by oxidation to quinone [106, 107].

Berberine Berberine displays multiple activities, including promotion of apoptosis and anticancer effect [156]. The drug may play a role in an apoptotic cascade in cancer cells by activation of the JNK/p38 pathway and by induction of ROS production. Phosphorylation may be involved. The alkaloid is a conjugated iminium species and a catechol ether which can serve as precursor to ET o-quinone.

PAC-1 and UCS1025A Several recent anticancer agents, PAC-1 and UCS 1025A have attracted attention due to considerable potential based on new approaches [84]. PAC-1 activates procaspase-3 to caspase-3, resulting in induction of apoptosis in tumor cells. UCS1025A is an inhibitor of the enzyme telomerase, present in cancer cells, which is crucially involved in tumor cell immortality. The agents possess chelating sites for metal binding, which has not been appreciated. Complexes of heavier metals are well known ET functionalities that can generate ROS. The literature addresses involvement of signal transduction for the agents.

Selenium The anticancer effect of Se is mediated by AMPK via a signaling pathway involving COX-2 [157]. ROS play a role as an AMPK activation signal in Se-treated cells. Evidence shows that Se can act both as an AO and a pro-oxidant [107]. Earlier, it was proposed that ROS are importantly involved in the action of chemotherapeutic drugs, analogous to the mode of action of phagocytes against foreign cells [3, 4].

Dialkyl Disulfide (DADS) An early event in treatment of neuroblastoma cells with DADS is an increase in ROS resulting in oxidation of lipid and protein [158]. The treatment induces JNK activation by phosphorylation. There is a pivotal role for ROS in apoptosis. DADS may be useful as an antiproliferative agent in cancer therapy.

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

289

Onion This vegetable reduces the incidence of cancer in several tissues and exerts an anticarcinogenic effect in multiple organs of animals [159]. The chemopreventive effects have been ascribed to various mechanisms: enhancement of the activity of MFOs, induction of phase II enzymes, increased synthesis of the AO GSH, and induction of cell cycle arrest and apoptosis. SOD Mn SOD and Cu Zn SOD are AOs that are tumor suppressive [160]. Data indicate that these AOs are regulated by the protein kinase C/ERK1/2 signaling pathway. Phytochemicals Many chemopreventive and chemoprotective phytochemicals enhance cellular AO capacity through activation of Nrf2 therapy-blocking carcinogenesis [161]. The transcription factor may activate gene encoding detoxifying and AO enzymes. Components of the cell signaling pathways include NF-kB and activator protein 1 (AP-1) which have been implicated in carcinogenesis, including an inflammatory component.

References 1. Kovacic P, Jacintho JD. Mechanisms of carcinogenesis: focus on oxidative stress and electron transfer. Curr Med Chem 2001; 8: 773–796. 2. Kovacic P, Somanathan R. Mechanism of tumorigenesis: focus on oxidative stress, electron transfer and antioxidants. In: Wong DK, ed., Tumorigenesis Research Advances, Hauppauge, New York: Nova Science Publishers; 2007: 23–65. 3. Kovacic P, Becvar LE. Mode of action of anti-infective agents: emphasis on oxidative stress and electron transfer. Curr Pharmaceut Des. 2000; 6:143–167. 4. Kovacic P, Osuna JA. Mechanisms of anticancer agents: emphasis on oxidative stress and electron transfer. Curr Pharmaceut Des. 2000; 6:277–309. 5. Kovacic P, Jacintho JD. Reproductive toxins: pervasive theme of oxidative stress and electron transfer. Curr Med Chem. 2001: 8: 863–892. 6. Kovacic P, Sacman A, Wu-Weis M. Nephrotoxins: widespread role of oxidative stress and electron transfer. Curr Med Chem. 2002; 9: 823–892. 7. Poli G, Cheeseman KH, Dianzani MU, Slater TF. Free radicals in the Pathogénesis of Liver Injury, NY: Pergmon Press: 1989. 8. Kovacic P, Somanathan R. Neurotoxicity: The broad framework of electron transfer, oxidative stress and protection by antioxidants. Curr Med Chem-CNS Agents, 2005: 5: 249–258. 9. Kovacic P, Thurn LA. Cardiovascular toxicity from the perspective of oxidative stress, electron transfer, and prevention by antioxidants. Curr Vasc Pharmacol. 2005: 3: 107–118. 10. Kovacic P, Pozos RS, Somanathan R, Shangari N, O’Brien PJ. Mechanism of mitochondrial uncouplers, inhibitors, and toxins: focus on electron transfer, free radicala, and structure-activity relationships. Curr Med Chem. 2005: 12; 2601–2623.

290

P. Kovacic and R. Somanathan

11. Kovacic P, Cooksy AL. Unifying mechanism for toxicity and addiction by abused drugs: electron transfer and reactive oxygen species. Med Hypotheses 2005; 64: 357–366. 12. Kovacic P, Somanathan R. Ototoxicity and noise trauma: electron transfer, reactive oxygen species, cell siganaling, electrical effects and protection by antioxidants; practical medical aspects. Med Hypotheses 2008; 70: 914–923. 13. Kovacic P, Somanathan R. Integrated approach to immunotoxicity: electron transfer, reactive oxygen species, antioxidants, cell signaling and receptors. J Recept Signal Transduct. 2008; 28: 323–346. 14. Halliwell B, Gutteridge JMC. Free Radicals in Biology and Medicine, NY: Oxford University Press; 1999. 15. Forman HG, Cadenas E. Oxidative Stress and Signal Transduction. Chapman and Hall, New York, 1997, pp. 1–475. 16. Hancock GT. Cell Signaling. Oxford University Press, New York, 2005, 1–296. 17. Demple B. Oxidative stress and signal transduction. In: Handbook of Cell Signaling, Bradshaw RA, Dennis EA, Eds., Academic Press, Boston. 2004, 78, 293–307. 18. Kovacic P, Pozos RS. Cell signaling (mechanism and reproductive toxicity): redox chains, electrons, relays, conduit, electrochemistry, and other medical implications. Birth Defect Res. Part C 2006, 78, 333–344. 19. Kovacic P. Protein electron transfer (mechanism and reproductive toxicity); iminium, hydrogen bonding, homoconjugation, amino acid side chains, and cell signaling. Birth Defects Res. Part C 2007. 20. Cevc G. Membrane electrostatics. Biochim Biophys Acta 1990, 1031–3, 311–382. 21. Clerkin JS, Naughton R, Quiney C, Cotter TG. Mechanisms of ROS modulated cell survival during carcinogenesis. Cancer Lett. 2008; 266: 30–36. 22. Kim E-H, Na H-K, Kim D-H, Park S-A, Kim H-N, Song N-Y, Surh Y-J. 15-Deoxy-D12,14prostaglandin J2 induces COX-2 expression through Akt-driven AP-1 activation in human breast cancer cells: a potential role of ROS. Carcinogenesis 2008; 29: 688–695. 23. Klaunig JE, Kamendulis LM. The role of oxidative stress in carcinogenesis. Annu Rev Pharmacol Toxicol. 2004; 44: 239–267. 24. Maher P, Schubert D. Signaling by reactive oxygen species in the nervous system. Cell Mol Life Sci. 2000; 57: 1287–1305. 25. Matés JM, Segura JA, Alonso FJ, Mrquez J. Intracellular redox status and oxidative stress: implications for cell proliferation, apoptosis, and carcinogenesis. Arch. Toxicol. 2008; 82: 273–299. 26. Waris G, Ahsan H. Reactive oxygen species: role in the development of cancer and various chronic conditions. J Carcinogen. 2006; 5:14. 27. Galaris D, Skiada V, Barbouti A. Redox signaling and cancer : the roll of “labile” iron. Cancer Lett. 2008; 266: 21–29. 28. Trachootham D, Ogasawara MA, Nilsa RD, Huang P. Redox regulation of cell survival. Antioxid Redox Signal 2008; 10: 1343–1374. 29. Lambeth JD. Mox enzymes, ROS, and chronic disease: an example of antagonistic pleiotropy. Free Rad Biol Med. 2007; 43: 332–347. 30. Arnold RS, He J, Remo A, Ritsick D, Yin-Goen Q, Lambeth J. D, Datta MW, Young AN, Petros JA. Nox1 expression determines cellular reactive oxygen and modulates c-fos-induced growth factor, interleukin-8, and Cav-1. Am J Pathol. 2007; 171: 2021–2032. 31. Martin KR, Barrett JC. Reactive oxygen species as double-edged swords in cellular processes: low-dose cell signaling versus high-dose toxicity. Hum Exp Toxicol. 2002; 21: 71–75. 32. Pandur S, Pankiv S, Johannessen M, Moens U, Huseby NE. Gamma-glutamyltransferase is upregulated after oxidative stress through the Ras signal transduction pathway in rat colon carcinoma cells. Free Rad Res. 2007; 41: 1376–1384. 33. Song SH, Lee KH, Kang MS, Lee YJ.. Role of paxillin in metabolic oxidative stress-induced cytoskeletal reorganization: involvement of SAPK signal transduction pathway and PTOOEST expression. Free Rad Biol Med. 2000; 29: 61–70.

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

291

34. Song JJ, Lee YJ. Catalase, but not MnSOD, inhibits glucose deprivation-activated ASK1MEK-MAPK signal transduction pathway and prevents relocalization of Daxx: hydrogen peroxide as a messenger of metabolic oxidative stress. J Cell Biochem. 2003; 90: 304–314. 35. Choi J-A, Kim E-Y, Song H, Kim C, Kim J-H.. Reactive oxygen species are generated through a BLT2-linked cascade in Ras-transformed cells. Free Rad Biol Med. 2008; 44: 624–634. 36. Wu W-S. The signal mechanism of ROS in tumor progression. Cancer Metastasis Rev. 2006; 25: 695–705. 37. Englnad K, Cotter TG. Direct oxidative modifications of signaling proteins in mammalian cells and their effects on apoptosis. Redox Rep. 2005; 10: 237–245. 38. Holmes-McNary M. Nuclear factor kappa B signaling in catabolic disorders. Curr. Opin Clin Metab Care 2002; 5: 255–263. 39. Janssen-Heininger YMW, Macara I, Mossman BT. Cooperativity between oxidants and tumor necrosis factor in the activation of nuclear factor (NF)-kB. Requirement of Ras/mitogen-activated protein kinase in the activation of NF-kB by oxidants. Am J Respir Cell Mol Biol. 1999; 20: 942–952. 40. Wang Y, Huang X, Cang H, Gao F, Yamamoto T, Osaki T, Yi J.. The endogenous reactive species promote NF-kB activation by targeting on activation of NF-kB-inducing kinase in oral squamous carcinoma cells. Free Rad Res. 2007; 41: 963–971. 41. Gopalakrishna R, Jaken S. Protein kinase C signaling and oxidative stress. Free Rad Biol Med. 2000; 28: 1349–1361. 42. Gibson BW. The human mitochondrial proteome: oxidative stress, protein modifications and oxidative phosphorylation. Int Biochem Cell Biol. 2005; 37: 927–934. 43. Herrera B, Álvarez AM, Sánxhez A, Fernández M, Roncero C, Benito M, Fabregat I.. Reactive oxygen species (ROS) mediates the mitochondrial-dependent apoptosis induced by transforming growth factor b in fetal hepatocytes. FASEB J. 2001;15: 741–751. 44. Lluis JM, Buricchi F, Chiarugi P, Morales A, Fernandez-Checa JC. Dual role of mitochondrial reactive oxygen species in hypoxia signaling: activation of nuclear factor-kB via c-SRCand oxidant-dependent cell death. Cancer Res. 2007; 67: 7368–7377. 45. Thannickal VJ, Fanburg BL. Reactive oxygen species in cell signaling. Am J Physiol Lung Cell Mol Physiol. 2000; 279: L1005-L1028. 46. Svineng G, Ravuri C, Rikardsen O, Huseby N-K, Winberg J-O. The role of reactive oxygen species in integrin and matrix metalloproteinase expression and function. Connect Tissue Res. 2008; 49: 197–202. 47. Na A. R, Chung Y. M, Lee S. B, Park S. H, Lee M-S, Yoo Y. D. A critical role of Romo1derived ROS in cell proliferation. Biochem Biophy Res Commun. 2008; 369: 672–678. 48. Meng T-C, Chen Y-Y. Insights into cysteine oxidation of protein tyrosine phosphatases that involves regulation of signal transduction and development of human diseases. Protein Oxid Dis. 2008; 55–71. 49. Meng T-C, Lou Y-W, Chen Y-Y, Hsu S-F, Huang Y-F. Cys-oxidation of protein tyrosine phosphatases; its role in regulation of signal transduction and its involvement in human cancer. J Cancer Mol. 2006; 2: 9–16. 50. Park I-J, Hwang J-T, Kim Y. M, Ha J, Park O. J. Differential modulation of AMPK signaling pathways by low or high levels of exogenous reactive oxygen species in colon cancer cells. Ann New York Acad Sci. 2006; 1091(Part B): 102–109. 51. Cho S. O, Kim K. H, Yoon J-H, Kim H. Signaling for integrin a5/b1 expression in Heliobacter pylori-infected gastric epithelial AGS cells. Ann New York Acad Sci. 2006; 1090: 298–304. 52. Handa O, Naito Y, Yoshikawa T. CagA protein of Helicobacter pylori; a hijacker of gastric epithelial cell signaling. Biochem Pharmacol. 2007; 73: 1697–1702. 53. Kim MH, Yoo H. S, Kim M. Y, Jang H. J, Baek M. K, Kim H. R, Kim K. K, Shin B. A, Ahn B. W, Jung Y. D. Helicobacter pylori stimulate urokinase plasminogen activator receptor expression and cell invasiveness through reactive oxygen species and NF-kB signaling in human gastric carcinoma cells. Int J Mol Med. 2007; 19: 689–697. 54. Kumar B, Koul S, Khandrika L, Meacham R. B, Koul H. K. Oxidative stress is inherent in prostate cancer cells and is required for aggressive phenotype. Cancer Res. 2008; 68: 1777–1785.

292

P. Kovacic and R. Somanathan

55. Roberts R. A, Ganey P. E, Ju C, Kamendulis I, Rusyn I, Klaunig J. E. Role of the Kupffer cell in mediating hepatic toxicity and carcinogenesis. Toxicol Sci. 2007; 96: 2–15. 56. Mossman B. T, Lounsbury K. M, Reddy S. P. Oxidants and signaling by mitogen-activated protein kinase in lung epithelium. Am J Respir Cell Biol. 2006; 34: 666–669. 57. Sawai H, Funahashi H, Okada Y, Matsu Y, Sakamoto M, Yamamoto M, Takeyama H, Manabe T. Imterleukin-1a enhances IL-secretion through p38 mitogen-activated protein kinase and reactive oxygen species signaling in human pancreatic cancer cells. Med Sci Moni. 2005; 11: BR343-BR350. 58. Duran A, Linares JF, Galvez AS, Wikenheiser K, Flores JM, Diaz-Meco MT, Moscat J. The signaling adaptor p62 is an important NF-kB mediator in tumorigenesis. Cancer Cell 2008; 13: 343–354. 59. Xia C, Meng Q, Liu L-Z, Rojanasakul Y, Wang X-R, Jiang B-H. Reactive oxygen species regulate angiogenesis and tumor growth through vascular endothelial growth factor. Cancer Res. 2007; 67: 10823–10830. 60. Ventura J-J, Cogswell P, Flavell RA, Baldwin AS. JNK potentiates TNF-stimulated necrosis by increasing the production of cytotoxic reactive oxygen species. Genes Develop. 2004; 18: 2905–2915. 61. Boudreau RT, Conrad DM, Hoskin DW. Differential involvement of reactive oxygen species in apoptosis caused by the inhibition of protein phosphatase 2A in Jurkat and CCRF-CEM human T-leukemia cells. Exp Mol Pathol. 2007; 83: 347–356. 62. Wu W-S, Tsai RK, Chang CH, Wang S, Wu J-R, Chang Y-X. Reactive oxygen species mediated sustained activation of protein kinase C a and extracellular signal-regulated kinase for migration of human hepotoma cell Hepg2. Mol Cancer Res. 2006; 4: 747–758. 63. Lee JK, Edderkaoui M, Truong P, Ohno I, Jang K-T, Berti A, Pandol SJ, Gukovskaya AS. NADPH oxidase promotes pancreatic cancer cell survival via JAK2 dephosphorylation by tyrosine phosphatases. Gastroenterology 2007; 133: 1637–1648. 64. Song S, Guha S, Liu K, Buttar N, Bresalier RS. COX-2 induction by unconjugated bile acids involves reactive oxygen species-mediated signaling pathways in Barrett’s oesophagus and oesophagaeal adenocarcinoma. Gut 2007; 56: 1512–1527. 65. Jung S-N, Yang WK, Kim J, Kim HS, Kim EJ, Yun H, Park H, Kim SO, Choe W, Kang I, Ha J. Reactive oxygen species stabilize hypoxia-inducible factor-1 alpha protein and stimulate transcriptional activity via AMP-activated protein kinase in DU145 human prostate cancer cells. Carcinogenesis 2008; 29: 713–721. 66. Cesaratto L, Calligaris SD, Vascotto C, Deganuto M, Bellarosa C, Quadrifoglio F, Ostrow JD, Tiribelli C, Tell G. Bilirubin-induced cell toxicity involves PTEN activation through an APE1/ ref-1-dependent pathway. J Mol Med. 2007; 85: 1099–1112. 67. Chang C-J, Mulholland DJ, Valamehr B, Mosessian S, Sellers WR, Wu H. PTEN nuclear localization is regulated by oxidative stress and mediates p53-dependent tumor suppression. Mol Cell Biol. 2008; 28: 3281–3289. 68. Djavaheri-Mergny M, Amelotti M, Mathieu J, Besancon F, Bauvy C, Codogno P. Regulation of autophagy by NFkB transcription factor and reactive oxygen species. Autophagy 2007; 3: 390–392. 69. Blanchetot C, Boonstra J. The ROS-NOX connection in cancer and angiogenesis. Crit Rev Eukary Gene Expr. 2008; 18: 34–45. 70. Shinohara M, Shang W-H, Kubodera M, Harada S, Mitsushita J, Kato M, Miyazaki H, Sumimoto H, Kamata T. Nox1 redox signaling mediates oncogenic Ras-induced disruption of stress fibers and focal adhesions by down-regulating Rho. J Biol Biochem. 2007; 282: 17640–17648. 71. Galanis A, Pappa A, Giannakakis A, Lantis E, Dangaj D, Sandaltzopoulos R. Reactive oxygen species and HIF-1 signaling in cancer. Cancer Lett. 2008; 266: 12–20. 72. Henderson BW, Daroqui C, Tracy E, Vaughan LA, Loewen GM, Cooper MT, Baumann H. Cross-linking of signal transducer and activator of transcription 3-A molecular marker for photodynamic reaction in cells and tumors. Clin Cancer Res. 2007; 13: 3156–3163.

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

293

73. Buytaert E, Dewaele M, Agostinis P. Molecular effectors of multiple cell death pathways initiated by photodynamic therapy. Biochim Biophys Acta, Review on Cancer 2007; 1776: 86–107. 74. Weyergang A, Kaalhus O, Berg K. Photodynamic therapy with an endocytically located photosensitizer cause a rapid activation of the mitogen-activated protein kinase extracellular signal-regulated kinase, p38, and c-Jun NH2 terminal kinase with opposing effects on cell survival. Mol Cancer Therapeut. 2008; 7: 1740–1750. 75. Kocanova S, Buytaert E, Matroule J-Y, Piette J, Golab J, de Witte P, Agostine P. Induction of heme-oxygenase 1 requires the p38MAPK and PI3K pathways and suppresses apoptotic cell death following hypericin-mediated photodynamic therapy. Apoptosis 2007; 12: 731–741. 76. Cross JV, Templeton DJ. Regulation of signal transduction through protein cysteine oxidation. Antioxid Redox Signal. 2006; 8: 1819–1827. 77. Davis W, Ronai Z, Tew KD. Cellular thiols and reactive oxygen species in drug-induced apoptosis. J Pharmcol Exp Therapeut. 2001; 296: 1–6. 78. Giles GI. The redox regulation of thiol dependent signaling pathways in cancer. Curr Pharm Des. 2006; 12: 4427–4443. 79. Ozaki K-I. Targeted molecular strategies for cancer therapy based on the blockade of oncogenic pathways in human tumor cells. Yakugaku Zasshi 2007; 127: 983–991. 80. Ju W, Wang X, Shi H, Chen W, Belinsky SA, Lin Y. A critical role of luteolin-induced reactive oxygen species in blockage of tumor necrosis factor-activated nuclear factor-kB pathway and sensitization of apoptosis in lung cancer cells. Mol Pharmacol. 2007; 71: 1381–1388. 81. Bradburn JE, Pei P, Kresty LA, Lang JC, Yates AJ, McCormick AP, Mallery SR. The effects of reactive oxygen species on the tumorigenic phenotype of human head and neck squamous cell carcinoma (HNSCC) cells. Anticancer Res. 2007; 27: 3819–3829. 82. Bode AM, Dong Z. Signal transduction molecules as targets for cancer prevention. Sci Signal. 2009; 2: mr2. 83. Wan Y, Li B, BCR-ABL. BCR-ABL in hematopoietic stem cell renewal and chronic myeloid leukemia. In:Wong, DK, ed. Tumorigenesis Research Advances. Hauppauge, New York, Nova Science Publ. 2007. 159–178. 84. Kovacic P. Does structural commonality of metal complex formation by Pac-1 (anticancer), DHBNH (anti-HIV), AHL (autoinducer), and UCS1025A (anticancer) denote mechanistic similarity? Signal transduction and medical aspects. J Recept Signal Transd. 2008; 28: 141–152. 85. Leonard SS, Bower JJ, Shi X. Metal-induced toxicity, carcinogenesis, mechanisms and cellular responses. Mol Cell Biochem 2004; 255: 3–10. 86. Chen Y, Yang J, Hu Q, Guo Q, Yi J. Emodin-enhanced arsenic-induced apoptosis on esophageal tumor cells. Shanghai Jiaotong Daxue Yixueban 2006; 26: 1227–1232. 87. Kim DW, Ahan SH, Kim TY. Enhancement of arsenic trioxide (As2O3)-mediated apoptosis using berberine in human neuroblastoma SH-SY5Y cells. J Korean Neurosurg Soc. 2007; 42: 392–399. 88. Rodríguez-Gabriel MA, Russell P. Distinct signaling pathway to arsenite and reactive oxygen species in Schzosacchromyces pombe. Eukary Cell 2005; 4: 1396–1402. 89. Zhou J, Ye J, Zhao X, Li A, Zhou J. JWA is required for arsenic trioxide induced apoptosis in HeLa and MCF-7 cells via reactive oxygen species and mitochondria linked signal pathway. Toxicol App Pharmacol. 2008; 230: 33–40. 90. Barchowsky A, O’Hara KA. Metal-induced cell signaling and gene activation in lung diseases. Free Radic Biol Med. 2003; 34: 1130–1135. 91. Wang S, Leonard SS, Ye J, Ding M, Shi X. The role of hydroxyl radical as a messenger in Cr(VI)-induced p53 activation. Am J Physiol Cell Physiol. 2000; 279: C868-C875. 92. Ye J, Wang S, Leonard SS, Sun Y, Butterworth L, Antonini J, Ding M, Rojanasakul Y, Vallyathan V, Castranova V, Shi X. role of reactive oxygen species and p53 in chromium(VI)induced apoptosis. J Biol Chem. 1999; 274: 34974–34980. 93. Benedyk M, Sopalla C, Nacken W, Bode G, Melkonyan H, Banfi B, Kerkhoff C. HaCaT keratinocytes overexpressing the S100 proteins S100A8 and Sa00A9 show increased NADPH oxidase and NF-kB activities. J Invest Dermatol. 2007; 127: 2001–2011.

294

P. Kovacic and R. Somanathan

94. Lee J-H, Li Y-C, Ip S-W, Hsu S-C, Chang N-W, Tang N-Y, Yu C-S, Chou S-T, Lin SS, Lin C-C, Yang J-S, Chung J-G. The role of Ca2+ in baicalein-induced apoptosis in human MDAMB-231 cancer cells through mitochondria and caspase-3-dependent pathway. Anticancer Res. 2008; 28: 1701–1711. 95. He X, Chen MG, Ma Q. Activation of Nrif2 in defense against cadmium-induced oxidative stress. Chem Res Toxicol. 2008; 21: 1375–1383. 96. Waisberg M, Joseph P, Hale B, Beyersmann D. Molecular and cellular mechanisms of cadmium carcinogenesis. Toxicol. 2003; 192: 95–117. 97. Bhattacharya K, Alink G, Dopp E. Oxidative stress and changed gene expression profiles in fiber-/particle-induced carcinogenesis. Int J Hum Genet. 2007; 7: 1–21. 98. Mossman BT, Faux S, Janssen Y, Jimenez, LA, Timblin C, Zanella C, Goldberg E, Walsh E, Barchowsky A, Driscoll K. Cell signaling pathways elicited by asbestos. Environ Health Perspect. 1997; 105: 1121–1125. 99. Hamilton RF, Thakur SA, Holian A. Silica binding and toxicity in alveolar macrophage. Free Radic Biol Med. 2008; 44: 1246–1258. 100. Castranova V. Signaling pathways controlling the production of inflammatory mediators in response to crystalline silica exposure: role of reactive oxygen/nitrogen species. Free Rad Biol Med. 2004; 37: 916–925. 101. Huang R-P, Wu J-X, fan Y, Adamson ED. UV activates growth factor receptors via reactive oxygen intermediates. J Cell Biol. 1996; 133: 211–220. 102. Madson JG, Hansen LA. Multiple mechanisms of Erbb2 activation after ultraviolet irradiation of the skin. Molec Carc. 2007; 46: 624–628. 103. Singh S, Malik B K, Sharma DK. Protein kinase C in prostate cancer and herbal products: a bioinformatics approach. Inten J Integrat Biol. 2007; 1: 71–87. 104. Singh M, Sharma H, Singh N. Hydrogen peroxide induces apoptosis in HeLa cells through a mitochondrial pathway. Mitochondrion 2007; 7: 367–373. 105. Ree SG. Cell signaling: H2O2, a necessary evil for cell signaling. Science 2007; 312: 1882–1883. 106. Kovacic P, Somanathan R. Alcohol mechanisms, Cell Signaling, Toxicity, Addiction, Prevention, Therapy and Beneficial Effects. In: Brozner EY. Ed. New research on Alcohol Abuse and Alcoholism. NY: Nova Science Publishers Inc., 2006: 40–101. 107. Kovacic P, Somanathan R. Beneficial Effects of Antioxidants in Relation to Carcinogens, Toxins and Various Illnesses In: Panglossi HV ed. Antioxidants: New Research. NY Nova Science Publishers Inc. 2006: 1–37. 108. Chan W-H. The signaling cascade of ginkgolide B-induced apoptosis in MCF-7 breast cancer cells. Int J Mol Sci. 2007; 8: 1177–1195. 109. Scott DW, Loo G. Curcumin-induced GADD153 upregulation: modulation by glutathione. J Cell Biochem. 2007; 101: 307–320. 110. Antherieu S, Ledirac N, Luzy A-P, Lenormand P, Caron J-C, Rahmani R. Endosulfan decreases cell growth and apoptosis in human HaCaT keratinocytes; partial ROS-dependent ERK1/2 mechanism. J Cellular Physiol. 2007; 213: 177–186. 111. Wan J, Winn LM. Benzene’s metabolites alter c-MYB activity via reactive oxygen species in HD3 cells. Toxicol Appl Pharmacol. 2007; 222: 180–189. 112. Burdick AD, Davis JW, Liu KJ, Hudson LG, Shi H, Monske ML, Burchiel SW. Benzo(a)pyrene quinones increase cell proliferation, generate reactive oxygen species, and transactivate the epidermal growth factor receptor in breast epithelial cells. Cancer Res. 2003; 63: 7825–7833. 113. Bolton JL, Thatcher GRJ. Potential mechanisms of estrogen quinone carcinogenesis. Chem Res Toxicol. 2008; 21: 93–101. 114. Soobrattee MA, Bahorun T, Aruoma OI. Chemopreventive actions of polyphenolic compounds in cancer. Biofactors 2006; 27: 19–35. 115. Ding M, Lu Y, Bowman L, Huang C, Leonard S, Wang L, Vallyathan V, Castranova V, Shi X.. Inhibition of AO-1 and neoplastic transformation by fresh apple peel extract. J Biol Chem. 2004; 279: 10670–10676.

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

295

116. Lo A-H, Liang Y-C, Lin-Shiau S-Y, Ho C-T, Lin J-K. Carnosol, an antioxidant in rosemary, suppresses inducible nitric oxide synthase through down-regulating nuclear factor-[kapp]b in mouse macrophages. Carcinogenesis 2002; 23: 983–991. 117. Aruoma OI, Kang K-S, Bahorun T, Sung M-K, Rahman I. Oxidative damage and chronic inflammation induced by smoking; potential antioxidant and peripheral biomarker considerations. Cancer Prevent Res. 2005; 10: 149–158. 118. Feng R, Bowman LL, Lu Y, Leonard SS, Shi X, Jiang BH, Castronova V, Vallyathan V, Ding M. Blacberry extracts inhibit activating protein 1 activation and cell transformation by perturbing the mitogenic signaling pathway. Nutr Cancer 2004; 50: 80–89. 119. Ahamed N, Gali H, Javed S, Agarwal R. Skin cancer chemopreventive effects of a flavonoid antioxidant silymarin are mediated via impairment of receptor tyrosine kinase signaling and perturbation in cell cycle progression. Biochem Biophys Res Commun. 1998; 247: 294–301. 120. McLoughlin P, Roengvoraphoj M, Gissel C, Hescheler J, Certa U, Sachinidis A.. Transcriptional response to epigallocatechins-3 gallate in HT 29 colon carcinoma spheroids. Genes Cells 2004; 9: 661–669. 121. Sarkar FH. The role of isoflavones in cancer chemoprevention. Front Biosci. 2004; 9: 2714–2724. 122. Shibata A, Nakagawa K, Sookwong P, Tsuzuki T, Oikawa S, Miyazawa T.Tumor anti-angiogenic effect and mechanism of action of d-tocotrienol. Biochem. Pharmacol. 2008; 76: 330–339. 123. Cerimele F, Battle T, Lynch R, Frank DA, Murad E, Cohen C, Macaron N, Sixbey J, Smith K, Watnik RS, Ellopoulos A, Shehata B, Arbiser JL.. Reactive oxygen signaling and MAPK activation distinguish Epstein-Barr virus (EBV)-positive versus EBV-negative Burkitt’s lymphoma. PNAS 2005; 102: 175–179. 124. Eskin NAM, Bird R. Antioxidants and cancer therapy: to take or not to take: that is the question? ACS Symposium Series 2007; 956 (Antioxidant Measurement and Application): 8–14. 125. Fruehauf, J. P., Meyskens, F. L. Reactive oxygen species: a breath of life or death? Clin Cancer Res. 2007; 13, 789–794. 126. Hensley K, Robinson KA, Gabbita SP, Salsman S, Floyd R A. Reactive oxygen species, cell signaling, and cell injury. Free Rad Biol Med. 2000; 28: 1456–1462. 127. Nair S, Wenge K, Ah-Ng T. Natural dietary anti-cancer chemopreventive compounds: redoxmediated differential signaling mechanisms in cytoprotection of normal cells versus cytotoxicity in tumor cells. Acta Pharmacol Sinica 2007; 28: 459–472. 128. Palović D, Dordevic V, Kocić GA. “Cross-talk” between oxidative stress and redox cell signaling. Med Biol. 2002; 9: 131–137. 129. Valko M, Leibfritz D, Moncol J, Cronin MT, Mazur M, Telser J.. Free radicals and antioxidants in normal physiological functions and human disease. Int J Biochem Cell Biol. 2007; 39: 44–84. 130. Valko M, Rhodes CJ, Moncol J, Izakovic M, Mazur M. Free radicals, metals and antioxidants in oxidative stress-induced cancer. Chem Biol Interact. 2006; 160: 1–40. 131. Townsend DM, Tew KD, Tapiero H. The importance of glutathione in human disease. Biomed Pharmacother 2003; 57: 145–155. 132. Martin V, Herrera F, Garcia-Santos G, Antolin I, Rodriguez-Blanco J, Rodriguez C. Signaling pathways involved in antioxidant control of glioma cell proliferation. Free Rad Biol Med. 2007; 42: 1715–1722. 133. Wu S-J, Ng LT, Lin C-C. Effects of antioxidants and caspase-3 inhibitor on the phenylethyl isothiocyanate-induced apoptic signaling pathways in human PLC/PRF/cells. Eur J Pharmacol. 2005; 518: 96–106. 134. McCord JM. SOD, oxidative stress and human pathologies: a brief history and future vision. Biomed Pharmacother. 2005; 59: 139–142. 135. Ganther HE. Selenium metabolism, selenoproteins and mechanism of cancer prevention: complexities with thioredoxin reductase. Carcinogenesis 1999; 20: 1657–1666.

296

P. Kovacic and R. Somanathan

136. Verma A, Atten BM, Holian O. Selenomethionine stimulates MAPK (ERK) phosphorylation, protein oxidation, and DNA synthesis in gastric cancer cells. Nutr Cancer 2004; 49: 184–190. 137. McLean L, Soto U, Agama K, Francis J, Jimenez R, Pommier Y, Sowers L, Brantley E. Aminoflavin induces oxidative DNA damage and reactive oxidative species-mediated apoptosis in breast cancer. Int J Cancer 2008; 122: 1665–1674. 138. Chathoth S, Thayyullathil F, Galadari S. Curcumin cell signaling: a possible target for chemotherapy. Curr Trends Biotech Pharmacy 2008; 2: 226–238. 139. Jawadi P, Segan AT, Tuttle SW, Koumenis C. The chemopreventive agent cucumin is a potent radiosensitizer of human cervical tumor cells via increased reactive oxygen species production and over activation of the mitogen-activated protein kinase pathway. Mol Pharmacol. 2008; 73: 1491–1501. 140. Numsen H. Mitochondrial reactive oxygen species affect sensitivity to curcumin-induced apoptosis. Free Rad Biol Med. 2008; 44: 1382–1393. 141. Valerie K, Yacoub A, Hagan MP, Curiel DT, Fisher PB, Grant S, Dent P. Radiation-induced cell signaling: inside-out and outside-in. Mol Cancer Ther. 2007; 6: 789–801. 142. Bauer G, Abteilung V. Low dose radiation and intercellular induction of apoptosis: potential implications for the control of oncogenesis. Int J Rad Biol. 2007; 83: 873–888. 143. Shao C, Folkard M, Held KD, Prise KM.. Estrogen enhanced cell-cell signaling in breast cancer exposed to targeted irradiation. BMC Cancer 2008; 8: 184. 144. Rigas B, Sun Y. Induction of oxidative stress as a mechanism of action of chemopreventive agents against cancer. Brit J Cancer 2008; 98: 1157–1160. 145. Rigas B, Williams JL. NO-donating NSAIDs and cancer: overview with a note on whether NO is required for their action. Nitric Oxide 2008; 19: 199–204. 146. Hwang J-T, Dong W, Lin SK, Kim HM, Kim YM, Park OJ. Resveratrol induces apoptosis in chemoresistant cancer cells via modulation of AMPK signaling pathway. Ann New York Acad Sci. 2007; 1095: 441–448. 147. Al-Ayyoubi S, Gali-Muhtasib H. Anti-tumor signaling pathways modulated by plant polyphenols.In: Valentine RG, ed. New Cell Apoptosis Research. Hauppauge, N. Y: Nova Science Publishers; 2007: 85–105. 148. Hwang J-T, Ha J, Park I-J, Lee S-K, Baik HW, Youg M, Park OJ. Apoptotic effect of EGCG in HT-colon cancer cells via AMPK signal pathway. Cancer Lett. 2007; 247: 115–121. 149. Halder B, Bhattacharya U, Mukhpadhyay S, Giri AK. Molecular mechanism of black tea polyphenols induced apoptosis in human skin cancer cells: involvement of bax translocation and mitochondria mediated death cascade. Carcinogenesis 2008; 29: 129–138. 150. Levovich I, Nudelman A, Berkovitch G, Swift LP, Cutts SM, Phillips DR, Rephaeli A. Formaldehyde-releasing prodrugs specifically affect cells by depletion of intracellular glutathione and augmentation of reactive oxygen species. Cancer Chemo Phamacol. 2008; 62: 471–482. 151. Kuo P-L, Chen C-Y, Hsu Y-L. Isoobtusilactone A induces cell cycle arrest and apoptosis through reactive oxygen species/apoptosis signal-regulating kinase 1 signal pathway in human breast cancer cells. Cancer Res. 2007; 67: 7406–7420. 152. Hong H-Y, Kim B-C. Mixed lineage kinase 3 connects reactive oxygen species to c-Jun NH2terminal kinase-induced mitochondrial apoptosis in genipin-treated prostate cancer cells. Biochem Biophys Res Commun. 2007; 362: 307–312. 153. Guo J, Jiang C, wang Z, Lee H-J, Hu H, Malewicz B, lee H-J, Lee J-H, Baek N-I, Jeong J-H, Kim D-K, Kang K-S, Kim, S-H, Lu J. A novel class of pyranocoumarin anti-androgen receptor signaling compounds. Mol Cancer Therap. 2007; 6: 907–917. 154. Ambrose M, Ryan A, O’Sullivan GC, Dunne, C, Barry OP. Induction of apoptosis in renal cell carcinoma by reactive oxygen species: involvement of extracellular signal-regulated kinase ½, p38d/g, cyclooxygenase-2-down-regulation, and translocation of apoptosis-inducing factor. Mol Pharmacol. 2006; 69: 1879–1890. 155. Shim H-Y, Park J-H, Paik H-D, Nah S-Y, Kim DSHL, Han YS. Acacetin-induced apoptosis of human cancer MCF-7 cells involves caspase cascade, mitochondria-mediated death signaling and SAPK/JNK1/2-cjun activation. Mol Cells 2007; 24: 95–104.

12 Cell Signaling and Cancer: Integrated, Fundamental Approach…

297

156. Hsu W-H, Hsieh Y-S, Kuo H-C, Teng C-Y, Huang H-I, Wang C-J, Yang S-F, Liou Y-S, Kuo W-H. Berberine induces apoptosis in SW620 human colonic cells through generation of reactive oxygen species and activation of JNK/p38 MAPK and FasL. Arch Toxicol. 2007; 81:719–728. 157. Hwang J-T, Kim YM, Surh Y-J, Baik HW, Lee S-K, Ha, J, Park OJ. Selenium regulates cyclooxygenase-2 and extracellular signal-regulated kinase signaling pathways by activating AMP-activated protein kinase in colon cancer cells. Cancer Res. 2006; 66: 10057–10063. 158. Filmeni G, Aquilano K, Rotillo G, Ciriolo MR. Reactive oxygen species-dependent c-Jun NH2-terminal kinase/c-jun signaling cascade mediates neuroblastoma cell death induces by diallyl disulfide. Cancer Res. 2003; 63; 5940–5949. 159. Wu X-J, Stahl T, Hu Y, Kassie F, Mersch-Sundermann V. The production of reactive oxygen species and the mitochondrial membrane potential are modulated during onion oil-induced cell cycle arrest and apoptosis in A549 cells. J Nutr. 2006; 136: 608–613. 160. Kim C, Hee H, Song L, Lee SY, Youk HS, Moon JY, Duong HQ, Park MJ, Joo YM, Park HG, Kim YJ, Yoo MA, Lim S-C, Kang HS. Protein kinase c-ERK1/2 signal pathway switches glucose depletion-induced necrosis to apoptosis by regulating superoxide dismutase and suppressing reactive oxygen species production in A549 lung cancer cells. J Cellular Physiol. 2007; 211: 371–385. 161. Sur YJ, Na HK, Lee JS. Redox-sensitive transcription factors as prime targets for chemoprevention with anti-inflammatory and antioxidative phytochemicals. J Nutr. 2005; 135: 2993S–3001S.

Chapter 13

Targeting Signal Transducer and Activator of Transcription (STAT) for Anticancer Therapy* Fabio P.S. Santos, Inbal Hazan-Halevy, and Zeev Estrov

Introduction In the 1980s, studies of the mechanism of action of cytokines, their corresponding receptors, and the effects of activated receptors on intracellular molecules identified a series of intracellular pathways responsible for activation of gene transcription. One of the first pathways studied in detail was the pathway activated by interferons (IFNs). These elaborate studies of IFN signaling led to the discovery of the signal transducer and activator of transcription (STAT) molecule [1–3]. STATs are latent cytoplasmic transcription factors. They are activated by tyrosine phosphorylation, dimerize, and translocate to the nucleus where they bind to DNA and act as transcription factors activating gene expression [4]. Their dual role as both transducers of cell signaling and activators of transcription inspired their name [3]. STATs are vital molecules in the transduction of almost all growth factors and cytokine signaling, having important roles in hematopoiesis, immunology, and tumor biology [5–7]. Today, it is well known that STAT molecules play an important role in cancer biology, being constitutively activated in different cancer types [8–13]. Because STATs play a key role in cancer biology, several investigators have been developing compounds capable of inhibiting STATs and/or their signaling pathways.

*

Conflicts of interest: none to declare.

Z. Estrov (*) Department of Leukemia, The University of Texas, Anderson Cancer Center, 1515 Holcombe Boulevard, Unit 0428, Houston, TX 77030, USA e-mail: [email protected] M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0_13, © Springer Science+Business Media, LLC 2012

299

300

F.P.S. Santos et al.

Structure and Function of STATs STAT Activation–Inactivation Cycle A classical pathway for STAT activation–inactivation has been described, even though there are many variations to this scheme [3, 14] (Fig. 13.1). STAT molecules exist in a monomeric, latent state in the cytoplasm. In order to be activated, they need to be phosphorylated on a tyrosine residue located in the C-terminal portion of the protein. Once the STAT monomers are phosphorylated, they form dimers by means of a reciprocal interaction between the phosphorylated tyrosine residue and the Src-homology 2 (SH2) domain of each molecule. The activated dimer translocates to the nucleus, where it binds to specific DNA sequences in the promoter of target genes according to the type of dimer that was formed.

Fig. 13.1 STAT activation pathways. STATs are tyrosine phosphorylated, dimerize through reciprocal association of the phosphotyrosine residue with the SH2 domain, and translocate to the nucleus where they bind to DNA and increase transcription of target genes. Several kinases are responsible for tyrosine phosphorylation of STATs, including non-receptor tyrosine kinases (Jaks [which can be bound to cytokine receptors or G-protein receptors] and Src) and receptor tyrosine kinases (such as EGFR and PDGFRa/b)

13

Targeting Stats in Cancer

Table 13.1 Mouse knockout gene models STAT Knockout phenotype STAT1 • Viable • Increased risk of bacterial and viral infections • Increased risk of tumors • Diminished response to IFN type I and type II STAT2 • Viable • Increased risk of bacterial infections • Diminished response to IFN type I STAT3 • Embryonically lethal • Conditional knockout revealed: ° Impaired cell survival ° Impairment of wound healing and keratinocyte migration ° Thymus hypoplasia ° Diminished T-lymphocyte survival ° Increase in granulocytes ° Enhanced inflammatory responses to pathogens, leading to chronic colitis ° Impaired breast development ° Impaired neuronal migration ° Heart: decrease in resistance to inflammation and increase in fibrosis ° Multiple other effects STAT4 • Viable • Diminished Th1 CD4+ T-lymphocyte polarization • Diminished risk of autoimmune disorders • Increased risk of intracellular bacterial infections STAT5a/b • Viable • Impaired development of mammary gland (STAT5a) • Impaired growth (mSTAT5b) • Diminished T-lymphocyte survival • Diminished responsiveness to EPO • Diminished stem cell repopulation ability STAT6 • Viable • Diminished Th2 CD4+ T-lymphocyte polarization • B-lymphocytes cannot class-switch and produce IgE

301

References [50, 52, 53]

[60]

[ 71–78, 177–191]

[84, 85]

[94–97, 99–101]

[106, 107]

STAT Function As multifaceted regulators of transcription, with many different activation pathways, it is no wonder that STATs perform a myriad of functions in diverse tissues. Mouse knockout models of STAT genes provide insight into the primary roles of these proteins (Table 13.1).

302

F.P.S. Santos et al. Table 13.2 STAT subtypes – gene locus and phosphorylation residues Phosphorylation site STAT Gene locus Tyrosine Serine References STAT1 2q32.2-q32.3 701 727 [47, 48] STAT2 12q13.2 690 – [58] STAT3 17q21 705 727 [70] STAT4 2q32.2-q32.3 694 721 [48] STAT5a 17q11.2 694 725/779 [88] STAT5b 17q11.2 699 730 [88] STAT6 12q13 641 756 [102]

Fig. 13.2 STAT protein structure and domains

STAT Phosphorylation and Activation In order to be activated, STATs need to be phosphorylated on a tyrosine residue located in the TAD (~position 700) (Table 13.2, Figs. 13.1 and 13.2). Upon phosphorylation on tyrosine residues, two STAT molecules dimerize, forming homodimers or heterodimers [3, 14]. All STATs (with the exception of STAT2) can form homodimers, and STAT1/STAT2 and STAT1/STAT3 can form heterodimers. STATs can be phosphorylated on tyrosine residues by several mechanisms (Fig. 13.1). The first one to be described was the cytokine receptor pathway [3]. Cytokine receptors usually do not have intrinsic tyrosine kinase (TK) activity. They associate with Jaks, which are cytoplasmic TKs first described in 1989 [15]. There are four members of the Jak family: Jak1, Jak2, Jak3, and Tyk [16]. Jaks bind to the cytoplasmic portion of cytokine receptors. Once the cytokine binds to its receptor, it induces dimerization or oligomerization of the receptor complex. This approximates the Jak kinases bound to each receptor chain and allows for transphosphorylation and activation of each Jak kinase. Once activated, the Jak kinase phosphorylates the receptor chain. This allows

13

Targeting Stats in Cancer

303

binding to the receptor of molecules with SH2 domains, such as STAT molecules. Once they bind to the receptor, the Jak kinase phosphorylates and activates STAT [6, 17].

STAT Serine Phosphorylation and Other STAT Modifications Besides tyrosine phosphorylation, STAT molecules can undergo other modifications with important biological effects. The most relevant ones are serine phosphorylation, lysine acetylation, and threonine O-glycosylation [18]. For a review of STAT serine phosphorylation [19]. Whereas STAT tyrosine phosphorylation has a clear function, leading to STAT dimerization, nuclear translocation, and activation of transcription, STAT serine phosphorylation has different consequences, depending on the stimulus that leads to serine phosphorylation, the kinases involved, and the subtype of STAT molecule phosphorylated. In the context of STAT tyrosine phosphorylation, serine phosphorylation is ligand induced and is necessary for full transcriptional activity [19, 20].

Activation of Transcription As transcription factors, the main function of STATs is to bind to DNA and activate transcription of target genes. Once they reach the nucleus, STATs bind to the promoter regions of specific genes and induce transcription. The TAD domain is essential for the transcriptional activity of STATs, and their ability to increase transcription depends on the recruitment and interaction between the TAD and coactivator proteins and other transcription factors [21, 22]. Protein Inhibitor of Activated STAT PIAS is another constitutive mechanism of suppressing STAT activity. These were originally described as inhibitors of STATs but now have been found to inhibit other transcription factors [23]. There are four proteins in the PIAS family: PIAS1, PIAS3 (isoforms a and b), PIASx (also known as PIAS2), and PIASy (also known as PIAS4) [23]. All PIAS proteins bind to STAT dimers. PIAS1 and PIASx bind to STAT1 dimers. PIAS3 binds to STAT3, and PIASy binds to STAT4 [23]. PIAS1 and PIAS3 act by blocking and impeding DNA binding of the STAT dimer [24, 25]. PIASx and PIASy act by recruiting corepressors such as histone deacetylases and blocking transcription of target genes [26, 27]. Furthermore, all PIAS proteins have SUMO E3 ligase activity, which mediates the SUMOylation of STATs [28–31]. As already mentioned, SUMOylation of STAT1 at lysine residue 703 has a negative regulatory function and inhibits STAT1 activity [28].

304

F.P.S. Santos et al.

STAT and Tumor Biology Overview Neoplasms are thought to be induced by deregulation of gene function caused by chromosomal deletions, translocations, point mutations, or epigenetic DNA modifications. Several molecular abnormalities in cancer cells constitutively activate intracellular signaling pathways, thereby enhancing proliferation, resistance to apoptosis, impaired differentiation, angiogenesis, and tumor cell dissemination. Transcription factors are unique mediators of cellular function, as each transcription factor regulates several genes. Therefore, deregulation of transcription factor function might induce cellular transformation. In many neoplastic diseases, STATs, particularly STAT3 and STAT5, are constitutively activated and, as a result, transcribe genes known to induce a malignant phenotype [5] (Fig. 13.3). Therefore, STATs are an attractive target for anticancer therapy.

Constitutive Activation of STATs in Cancer The first studies that demonstrated the role of STAT3 in oncogenesis were conducted in virally transformed cells. STAT3 was found to be activated in Rous sarcoma virus (v-Src) transformed cells [32], and inhibition of STAT3 signaling impeded transformation of mouse fibroblasts infected by v-Src [33, 34], suggesting that the ability of v-Src to transform cells depends on STAT3. A mutant form

Fig. 13.3 STAT3 activation and cancer pathogenesis. Several different mechanisms can lead to STAT3 constitutive activation in cancer cells, and the consequences of STAT3 activation include increased cellular proliferation, resistance to apoptosis, increased angiogenesis, and suppression of antitumor immune responses

13

Targeting Stats in Cancer

305

of STAT3, which dimerizes spontaneously due to the substitution of two cysteine residues in the SH2 domain, transformed fibroblasts and induced tumor formation in nude mice, suggesting that STAT3 is a potential oncogene [34]. These initial studies were followed by several reports that indicated the importance of STATs in several tumor types. However, the mechanisms of STAT activation in cancer cells are not completely understood. Besides tyrosine phosphorylation, serine phosphorylation of STATs has also been demonstrated in cancer cells. STAT3 is constitutively phosphorylated at S727 in chronic lymphoid leukemia (CLL) cells, and phospho-serine STAT3 binds DNA and activates transcription in the absence of tyrosine phosphorylation in CLL [35, 36]. Transfection of CLL cells with a STAT3 small hairpin RNA (STAT3-shRNA) leads to a reduction in the expression of several STAT3 target genes and subsequent apoptosis, indicating that serine phosphorylation of STAT3 in CLL has biological significance and leads to gene transcription [36]. Constitutive serine phosphorylation of STAT3 has also been described in prostate cancer cells, where it drives prostate tumorigenesis independent of tyrosine phosphorylation [37]. STAT inhibitors also seem to play a role in cancer pathogenesis. Silencing of STAT inhibitors can lead to hyperactivation of STAT3 and/or STAT5, and this could possibly lead to the same outcome as increasing the rate of STAT phosphorylation and activation. DNA methylation is the addition of a methyl group to the C5 position of the cytosine (C) pyrimidine ring [38]. This occurs when C is followed by a guanine (G) in the so-called CpG islands, which are clustered in gene regulatory regions, such as promoters (promoter CpG islands) [38]. DNA methylation of promoter CpG islands leads to gene silencing, similar to what would occur if the gene was deleted [38]. Cancer cells have abnormal patterns of DNA methylation, and this contributes to the malignant phenotype [38]. Silencing of STAT3 inhibitors by gene promoter hypermethylation has been described in solid tumors and hematologic malignancies and was recently reviewed [39].

STATs and Tumor Cell Proliferation and Resistance to Apoptosis It can be seen that STATs (especially STAT3 and STAT5) are activated by several oncogenic signaling pathways, which is consistent with their role as transcription factors that are active in malignancies. STAT3 increases the transcription rate of several genes that contribute to tumorigenesis. Using a gene expression array, one study showed that activation of STAT3 in fibroblasts enhanced the expression of about 100 target genes, using a threshold of a 1.5-fold increase in expression [40]. Subsequent analysis of the data revealed that 12 STAT3 target genes were highly expressed in several malignancies, consistent with a STAT3 gene signature that was found in cancer [40]. These 12 target genes include genes involved in increased proliferation (early growth response-1 [EGR-1]), resistance to apoptosis (myeloid cell leukemia-1 [MCL-1]), and increased angiogenesis (vascular endothelial growth factor [VEGF]) [40]. It is thus clear that STAT3 activates the transcription of several genes that contribute to the phenotype of the cancer cell.

306

F.P.S. Santos et al.

The BCL-2 family of genes is one of the main regulators of apoptosis, and it comprises both anti-apoptotic and pro-apoptotic genes [41]. A mutant and constitutively active form of STAT3 has been demonstrated to increase the expression of the anti-apoptotic gene BCL-XL [42]. In multiple myeloma (MM) cell lines dependent on IL-6, activation of STAT3 has been demonstrated to increase BCL-XL, and blockage of this signaling pathway leads to a decrease in BCL-XL levels [43, 44]. Genes that are responsible for cell cycle regulation are also a target of STAT3 and STAT5. Cyclin D levels increase during the initial portion of the G1 phase of the cell cycle [45]. Cyclin D binds to cyclin-dependent kinases 4 and 6 (CDK 4/6) and increases their catalytic activity [45]. CDK 4/6 are serine–threonine kinases that are responsible for initiating the process that leads the cell to division [45]. Among the many targets of CDK 4/6, the retinoblastoma protein (pRb) stands as one of the most important [45]. Phosphorylation of pRb permits the cell to transverse the G1 phase restriction point and proceed to the next phase in the cell cycle, DNA synthesis (S phase) [45]. Normally, cyclin D levels are regulated by extracellular mitogenic signals [45]. It is no wonder then that STAT3 and STAT5 are associated with upregulation of cyclin D. C-myc is another gene responsible for cell cycle control that is also regulated by STATs. C-myc is a transcription factor, and it increases the expression of several genes that lead to cell proliferation and blockade of cellular differentiation [46]. Several genes have their transcription levels enhanced by c-myc, including cyclin D, cyclin E, CDK 4/6, and CUL1 (Cullin 1; degrades the CDK inhibitor p27Kip1), as well as the transcription factors of the E2F family [47–52]. Cells transfected with the STAT3C mutant have increased levels of c-myc mRNA [42]. STAT5 has been shown to regulate c-myc in CML cell lines [53]. IL-6 induces an increase in c-myc levels by signaling through STAT3 [54]. The p53 gene is a key tumor suppressor gene that is induced by several signals, especially DNA damage, and activates genes responsible for blocking cell proliferation, repairing DNA, and activating apoptosis [55]. One important study demonstrated that activated STAT3 binds to the promoter of wild-type p53 and inhibits its transcription [56]. As STAT3 is upregulated in many cancers, targeting STAT3 might lead to an increase in the transcription level of p53 in tumors without p53 mutations.

STATs and Tumor Angiogenesis Angiogenesis is the biological process of formation of new blood vessels from preexisting vasculature, and it is essential for uncontrolled tumor growth, since hypoxia and diminished blood supply can limit the size of tumors [57]. Angiogenesis is a complex process and depends on the interplay between angiogenic and angiostatic molecules [57]. VEGF (also known as VEGF-A) is one of the most important mediators of angiogenesis [57].

13

Targeting Stats in Cancer

307

VEGF is a target gene of STAT3, and STAT3-mediated increase in VEGF plays an important role in tumor angiogenesis [58, 59]. It has been demonstrated that there is a correlation between expression of VEGF and elevated STAT3 activity in several tumor types, including pancreatic cancer, melanoma, head and neck cancer, breast cancer, and gastric cancer [59–65]. A mutant, constitutively active form of STAT3 (STAT3C) leads to increased expression of VEGF in NIH3T3 fibroblasts [42]. STAT3 binding sequences were found in the promoter region of VEGF. It has been shown that cytokines (e.g., IL-6) and TKs (e.g., Src) have an important role in promoting tumor angiogenesis and VEGF expression [58]. There is now ample evidence to indicate that the angiogenic activity of cytokines and growth factors is actually mediated by STATs, especially STAT3 [66–68]. IL-6 increases the expression of VEGF in C33A cervical cancer cells, and blockade of STAT3 by a dominant-negative mutant abolishes VEGF expression [66]. IL-6 also has been shown to increase VEGF levels by STAT3-dependent pathways in gastric cancer [69]. Luciferase assay indicated that only the Jak2 inhibitor AG490 was capable of suppressing VEGF production induced by IL-6, implicating Jak/STAT3 in the signaling pathway responsible for IL-6-induced angiogenesis [69]. The VEGF receptor (VEGFR) signals through STAT3 pathways in endothelial cells [70]. Blocking STAT3 inhibits endothelial cell migration and angiogenesis [71]. STAT3 also upregulates matrix metalloproteinases (MMP-2 and MMP-9), which are proteolytic enzymes that promote angiogenesis, tumor invasion, and metastasis by degrading the extracellular matrix and the basement membrane [72, 73].

Role of STATs in Evading Immune Surveillance Mechanisms The immune system has developed innate and adaptive immune responses to prevent and control the development of tumors. Evading these immune surveillance mechanisms is crucial for the progression of tumors [74, 75]. It is known that tumor cells induce an immunosuppressive microenvironment that abolishes tumor-specific immune responses [76, 77]. Recently, several studies have indicated that STAT3 plays a major role in the genesis of tumor-induced immunosuppression by inhibiting Th1 immune response, blocking the maturation of dendritic cells, inducing regulatory T-cells (TRegs), and shifting the balance to the production of pro-inflammatory and oncogenic cytokines such as IL-23 [78–80]. In this crosstalk between tumor cells and immune cells, STAT3 is a central molecule, linking oncogenic pathways with immunosuppression. The observation that STAT3 might be involved in generating an immunosuppressive environment came from a gene therapy study of the melanoma B16 tumor in mice. Another study showed an inverse correlation between expression of STAT3 and immune cells infiltrating melanoma tumors [80]. Neutrophils isolated from tumors show constitutively active STAT3, and specific ablation of STAT3 in the hematopoietic system leads to increased cytotoxicity of neutrophils and NK cells

308

F.P.S. Santos et al.

against tumor targets [78]. All these results indicate that STAT3 acts to suppress the expression of pro-inflammatory cytokines that might act as danger signals and attract immune cells to the tumor and to inhibit the function of immune cells that are present in the tumor microenvironment. STAT3 also downregulates Th1 immune responses, not only by stimulating the production by tumor cells of immunosuppressive factors (VEGF, IL-10) but also by counteracting the immunostimulatory effects of NFkB and STAT1 [81, 82] and inducing the accumulation of TRegs [78]. Because many of the immunosuppressive factors are regulated by STAT3, and they signal STAT3-dependent pathways, the maintenance of tumor-induced immunosuppression seems to be dependent on this specific molecule, which makes it an attractive target for cancer therapy.

Targeting STATS Peptidic Inhibitors and Peptidomimetic Inhibitors Peptidic inhibitors targeting the STAT3 SH2 domain could be developed based on peptide sequences located on STAT3 itself (around Y705) or located on the receptors that activate STAT3. STAT3 binds via its SH2 domain to phosphorylated peptides based on receptor motifs that contain the sequence pYXXQ (pY is the phosphorylated tyrosine) [83–85] and also to phosphorylated peptides based on the STAT3 Y705 sequence pYLKT [86]. The main challenges with the use of peptides as drugs are their low stability, being subject to degradation by peptidases, and polar nature, which decreases membrane permeability and intracellular concentrations. The requirement of a phosphotyrosine adds another level of difficulty, since it can be targeted by phosphatases. Peptidic inhibitors, however, could be used as a framework upon which to develop peptidomimetic drugs with greater membrane permeability and increased stability.

Small-Molecule Non-peptidic Inhibitors Small-molecule non-peptidic inhibitors represent a class of drugs with greater potential for being developed into clinical use because of their greater stability, increased cell membrane permeability, and non-requirement for a pY residue. Based on computational modeling of the interaction between the SH2 domain and the phosphopeptide, several compounds have been identified by screening of virtual databases. After screening 429,000 compounds in the National Cancer Institute (NCI) Chemical Library, the small molecule inhibitor STA-21 was found to inhibit STAT3 dimerization in vitro at concentrations of 20 mM, and at concentrations of 20–30 mM, it inhibited STAT3 gene transcription and induced apoptosis of human breast carcinoma cells

13

Targeting Stats in Cancer

309

(MDA-MB-231, MDA-MB-435, and MDA-MB-468) with constitutively active STAT3 [87]. STA-21 exerted minimal effects upon cells without constitutively active STAT3. Another virtual screening of the NCI Chemical Library identified S3I-201 (also known as NSC 74859) as another potential STAT3 inhibitor [88]. It inhibits STAT3 dimerization, DNA binding, and transcriptional activity, with an IC50 value of 86 mM by EMSA assay [88]. S3I-201 induced growth inhibition and apoptosis in human breast carcinoma cell lines and suppressed the expression of the STAT3 target genes cyclin D1, BCL-xL, and survivin [88]. Furthermore, intravenous injection of S3I-201 at a dose of 5 mg/kg every 2 days led to the growth inhibition of human breast carcinoma tumors xenografted into nude mice [88]. Another study identified by virtual screening of another chemical library the compound Stattic (STAT three-inhibitory compound). Stattic was found to inhibit the binding of the phosphopeptide GpYLPQTV to the STAT3 SH2 domain [89]. The effect of Stattic is temperature dependent, exhibiting weak activity at 22°C and strong inhibition at 37°C (IC50 value of 5.1 mM). Stattic was shown to abrogate IL-6-induced STAT3 activation in human HepG2 hepatoma cell lines at a concentration of 20 mM and induce apoptosis of human breast carcinoma cells with constitutively active STAT3 [89]. Recently, the catechol (1,2 dihydroxybenzene) moiety was found to be a STAT3 SH2 inhibitor by virtual screening [90]. Catechol compounds dock to the SH2 domain, and computational modeling showed that they form hydrogen bonds with the pY-interacting amino acids [90]. A series of catechol compounds was synthesized and evaluated for activity in vitro as STAT3 inhibitors [90]. One compound (no. 8) inhibited STAT3 activity with an IC50 value of 106 mM. Further studies of these compounds to evaluate their in vitro and in vivo activities are awaited [90].

Inhibitors of STAT DNA Binding Domains Since STAT is a transcription factor and its action is dependent on DNA binding and activation of transcription, several compounds have been developed that target the DNA binding domain of STAT3 with hopes of blocking its transcriptional activity. Several compounds derived from platinum (IV) were found to have activity as STAT3 inhibitors. Their mechanism of action is not completely understood, but it involves interacting with STAT3 and blocking its ability to bind DNA. The platinum (IV) derivative IS3 295 was identified on screening of the NCI 2000 diversity set [91]. IS3 295 was shown to block STAT3 DNA binding activity with an IC50 of 1.4 mM [91]. IS3 295 directly inhibits STAT3 DNA binding ability, and it interacts with cysteine residues within STAT3 [91]. IS3 295 abrogated the expression of STAT3 target genes such as cyclin D1 and BCL-xL, and it induced cell growth arrest at G0/G1 and apoptosis of v-Src-transformed NIH3T3 fibroblasts and human breast carcinoma cells at a 10-mM concentration [91]. Another compound that targets the DNA binding domain of STAT3 is a tetrahydro-isobenzofuranone derivative isolated from fermentation broth of the ascomycete strain A111-95 [92]. Galiellalactone was discovered during a search for

310

F.P.S. Santos et al.

compounds that inhibit IL-6-mediated signal transduction in HepG2 cells [534]. It blocks IL-6-induced signaling, with IC50 values of 0.25–0.5 mM [92], and blocks STAT3 DNA binding without affecting its phosphorylation status. Galiellalactone loses its activity after being incubated with cysteine, and it has been proposed that it acts by binding to a cysteine residue located on the DNA binding domain (C468) of STAT3 and forming an inactive complex [92]. Galiellalactone at a concentration of 25 mM was demonstrated to downregulate STAT3 gene expression and induce apoptosis of the androgen-insensitive human prostate cancer cell lines DU145 and PC-3 that express constitutive STAT3 [93]. In nude mice with tumor xenografts of human prostate cancer (DU145 cells), galiellalactone administered at a dose of 3 mg/kg intraperitoneally induced tumor regression, demonstrating in vivo activity [93]. Another approach to block DNA binding of STAT3 would be to target it utilizing peptide aptamers. Peptide aptamers are short-chain peptides that target a specific molecule and are bound at each end to a protein scaffold [94]. Recently, 20-mer peptide aptamers have been identified that target the STAT3 DNA binding domain and block its transcriptional activity [95]. The peptides were selected from a peptide library of high complexity by an adaptation of the yeast two-hybrid procedure [95]. Peptide aptamers inhibited DNA binding and transactivation by STAT3 following EGF stimulation of NIH3T3/EGFR (Herc) cells [95]. Peptide aptamers tagged with a protein transduction motif of nine arginines and fused to thioredoxin as a scaffold protein were cell permeable and induced apoptosis of human multiple myeloma U266 cells at a concentration of 0.27 mM [95].

Inhibitors of the N-Terminal Domain The N-terminal domain of STAT3 is composed of eight a-helices, and it is involved in several protein–protein interactions, such as the formation of STAT3 tetramers and the interaction between STAT3 and other transcription factors [96]. It was hypothesized that targeting the N-terminal domain of STAT3 might interfere with its transcriptional activity. One recent report demonstrated for the first time the efficacy of STAT3 aminoterminal domain inhibitors in abrogating STAT3 activity [97]. Based on STAT4 data and sequence homology, a library of synthetic peptides derived from the second a-helix of the amino-terminal domain of STAT3 was constructed [97]. The peptides were fused to penetratin (a 16-amino acid peptide fragment that is internalized by cells) to enable membrane penetration. The fused peptide was found to inhibit STAT3 DNA transcription activity and induce growth inhibition and apoptosis in the human breast cancer cell lines MDA-MB-231, MDA-MB-435, and MCF-7 [97]. Preliminary data suggest that the peptides interfere with the binding of the histone deacetylase HDAC1 and the DNA methyltransferase DNMT1 to STAT3, but the precise mechanism of action is still unknown [97]. The same problems described above, with the utilization of peptides as drugs for clinical use, also have to be overcome before this inhibitor can be evaluated in the clinical setting. However, the

13

Targeting Stats in Cancer

311

study is important because it demonstrates the proof-of-concept that inhibiting the STAT3 amino-terminal domain blocks its transcriptional activity and represents another approach for targeting it.

Oligonucleotides Targeting STAT3 Oligonucleotides can be used to target STAT3, leading to degradation of STAT3 mRNA, inhibition of DNA binding activity, and disruption of dimerization. Oligonucleotides utilized include single-stranded antisense oligodeoxynucleotides (ODNs), small interfering RNA (siRNA), decoy ODN, and G-quartet ODN. Single-stranded antisense ODNs (ASO) have a sequence that is complementary to the mRNA of the targeted gene. ASO hybridize with the target mRNA in a sequence-specific manner and lead to mRNA degradation, inhibition of mRNA maturation by interfering with the splicing mechanism, and inhibition of mRNA translation and protein synthesis [98]. It has been demonstrated that ASO targeting STAT3 mRNA induce growth inhibition and apoptosis in human hepatocarcinoma and prostate cancer cell lines [99, 100]. A second-generation 2’-O-methoxyethyl (2’-MOE) modified ASO (for increased stability) targeting STAT3 (ISIS 345794) is currently in preclinical development and is expected to enter clinical trials soon [101]. siRNA is a class of 20–25-nucleotide-long double-stranded RNA molecules that interfere with the expression of a particular gene. Therapy with siRNA has been utilized to decrease the activity of STAT3 both in vitro and in vivo. Knockdown of STAT3 by siRNA leads to apoptosis in human astrocytoma cell lines A172 and T98G, with decreases in BCL-XL and survivin [102]. In another study, STAT3 mRNA was targeted using a dicistronic lentivirus small hairpin RNA (shRNA) [103]. 4T1 breast cancer cells transduced with the shRNA had decreased levels of STAT3 and did not induce tumor formation in BALB/c mice, whereas 4T1 cells that were not transduced with the shRNA efficiently formed tumors [103]. Decoy ODNs inhibit STAT3 DNA binding activity and mimic the consensus sequence of cis-elements that have a high specificity for the transcription factor [104]. Exogenous double-stranded decoy ODN essentially competes with the endogenous cis-elements for binding the transcription factor, resulting in downregulation of the expression of target genes [104]. A 15-mer STAT3 decoy ODN with the sequence 5¢-CATTTCCCGTAAATC-3¢ was shown to bind specifically to activated STAT3 and block its binding to DNA sequences found in target genes [105]. It inhibited proliferation and decreased expression of STAT3 target genes in squamous cell carcinoma of the head and neck cell lines [105]. Other studies followed and confirmed the potential of decoy ODN in inhibiting STAT3 signaling in several cancer cell lines with aberrant STAT3 activity [106–108]. Furthermore, decoy ODN injected intratumorally at a dose of 25 mg/day led to regression of tumor xenografts of A459 human lung cancer cells, confirming in vivo activity [109]. Another study evaluated the toxicity of an intramuscular injection of a low (0.8 mg) and high (3.2 mg) dose of STAT3 decoy ODN in cynomolgus monkeys [110]. There was no toxicity observed, and in the monkeys who received the high-dose injection,

312

F.P.S. Santos et al.

there was decreased expression of cyclin D1 and BCL-XL [110]. A phase I trial evaluating the safety and biological activity of a single intratumoral injection of STAT3 decoy ODN in patients with head and neck cancer is expected to start soon. Another class of ODN agents is G-quartet ODNs. G-quartet ODNs are guanine (G)-rich ODNs that have the ability to form inter- and intramolecular four-stranded structures known as G-quartets [111]. The G-quartet ODN T40214 was identified as a potent inhibitor of STAT3 DNA binding activity, with an IC50 value of 7 mM [112]. Computational modeling studies revealed that T40214 binds to the SH2 domain of STAT3 and forms hydrogen bonds with residues Q643, Q644, N646, and N647, thus disrupting STAT3 dimerization and DNA binding activity [113]. In a xenograft model of prostate and breast tumors, T40214 demonstrated in vivo activity leading to growth inhibition and tumor regression when given intravenously at a dose of 5.0 mg/kg [114].

Tyrosine Kinase Inhibitors The class of drugs known as TKIs includes several agents that are currently approved for use in cancer, including the Bcr-Abl/C-Kit inhibitor imatinib (used to treat CML and gastrointestinal stromal tumor [GIST]) [115, 116], the EGFR inhibitor erlotinib (used in non-small cell lung cancer) [117], the EGFR/Her-2 inhibitor lapatinib (used in breast cancer) [118], and the Bcr-Abl/Src inhibitor dasatinib (used in CML) [119]. TKIs are small-molecule inhibitors that compete with ATP for binding in the ATP pocket located in the TK [120]. Since the main mechanism of activation of STATs is tyrosine phosphorylation, it is logical that these compounds are indirect STAT inhibitors. However, TKIs also affect several other signaling pathways that also depend on tyrosine phosphorylation, and it is difficult to delineate how much of their effect is due to STAT inhibition.

Natural Compounds There are several compounds derived from natural sources that can inhibit the STAT3 pathway [121]. Screening of the NCI diversity set using a cytoblot assay to search for STAT3 phosphorylation inhibitors led to the identification of JSI-124 (cucurbitacin-I) [122]. JSI-124 is a member of the cucurbitacin family of compounds, isolated from the Cucurbitaceae plant family, which is used in herbal medicine in China and India [123]. Cucurbitacins are a group of triterpenoid substances structurally characterized by a tetracyclic cucurbitane nucleus skeleton (9b-methylnorlanosta-5-ene) with a variety of oxygen substitutions at different positions [123]. Cucurbitacin-I was first isolated from Ecballium elaterium [124], and it has been shown to inhibit DNA, RNA, and protein synthesis and proliferation in HeLa cells and to inhibit proliferation of Ehrlich ascites carcinoma cells [125, 126].

13

Targeting Stats in Cancer

313

JSI-124 selectively inhibits STAT3 phosphorylation in vitro at a concentration of 10 mM [122]. The precise mechanism of action remains unknown, however, since Src, Jak, Erk, and Akt kinases are not inhibited [122]. JSI-124 decreased the levels of phosphorylated STAT3 in NIH3T3/v-Src transformed fibroblasts and in human lung, breast, and pancreatic cancer, glioma, and lymphoma cells [30, 122]. JSI-124 administered intraperitoneally at a dose of 1 mg/kg inhibited the growth of human breast (MDA-MD-468) and lung (A459) tumor xenografts in nude mice [122]. JSI124 prolonged the survival of mice with murine melanoma and induced antitumor immune responses of dendritic cells, presumably by inhibition of STAT3 activation [127]. Also, CpG deoxynucleotide (a dendritic cell activator) was combined with JSI-124 in the treatment of melanoma in mice [128]. The combination led to a greater antitumor effect and correlated with an increase in the level of inflammatory Th-1 cytokines (including IL-12, IFN-g, TNF-a, and IL-2) and an increase in intratumoral CD8+ and CD4+ T-cells expressing activation/memory markers and NK cells [128]. There was also a significant decrease in immunosuppressive cytokines, as evidenced by lower intratumoral levels of VEGF and TGF-b, and a decreased number of CD4+CD25+Foxp3+ TReg cells in lymph nodes [128]. JSI-124 is poorly water soluble, and polymeric micelles were developed that increase the water solubility and improve the delivery of the drug [129]. Other cucurbitacins have been shown to have activity as STAT3 inhibitors [35]. Withacnistin, originally identified erroneously as cucurbitacin-Q, is a potent inhibitor of STAT3 [130]. It leads to tumor cell apoptosis and suppresses the growth of human lung and breast tumor xenografts [130]. It also does not inhibit Jak2, Src, Akt, Erk, or JNK. Cucurbitacin-B is another STAT3 inhibitor that has activity against hepatocellular carcinoma cells and was recently shown to synergize with docetaxel in inducing apoptosis in human laryngeal cancer cells in vitro [131, 132]. Curcumin (diferuloylmethane) is the active ingredient of the herbal remedy and dietary spice turmeric [133]. Turmeric is obtained from the rhizome of the plant Curcuma long and has been used for a long time in traditional medicine in India and China [133]. Recently, curcumin has been demonstrated to be active as a chemotherapeutic and chemopreventive agent. Curcumin inhibits STAT3 and several other signaling pathways [121, 134]. The precise mechanism is unknown, but it has been demonstrated that curcumin inhibits several TKs that are responsible for the activation and phosphorylation of STAT3, including Jak1, Jak2, Src, Erb-b2, and EGFR [44]. Curcumin can also activate a SHP-2 phosphatase, which leads to the dephosphorylation of Jak1/2 [135]. Curcumin suppresses STAT3 phosphorylation after only 30 min at a concentration of 10 mM and downregulates the expression of STAT3 target genes such as BCL-XL and cyclin D1 [136, 137]. It was shown that curcumin inhibited IL-6 signaling and STAT3 activation in multiple myeloma U266 cells, inducing growth inhibition and apoptosis [136]. Curcumin was more effective than another Jak2 inhibitor, AG490, in blocking STAT3 phosphorylation in myeloma cells (10 vs. 100 mM) [136]. Curcumin was shown to inhibit both NFkB and STAT3 activity in myeloma CD138+ cells obtained from patients with myeloma, and it was more potent than dexamethasone in this regard [138]. Curcumin was also shown to

314

F.P.S. Santos et al.

inhibit STAT3 in other human tumor cell lines, including Hodgkin’s lymphoma and head and neck squamous cell carcinoma cells with constitutive STAT3 activation [51]. Synthetic analogues of curcumin have been developed and have demonstrated activity in a xenograft model of breast cancer [139, 140]. Curcumin is safe in humans, and clinical trials did not reveal any dose-limiting toxicities when administered in doses up to 8 g/day [141]. Currently, there are several clinical trials evaluating the role of curcumin in the treatment of a broad range of malignancies. Another natural compound derived from spices is capsaicin, a constituent of green and red peppers from the plant genus Capsicum. Capsaicin blocks IL-6induced STAT3 activation [142]. The mechanism involves depletion of intracellular gp130 pools through induction of endoplasmatic reticulum stress [143]. Indirubin is the active component of a traditional Chinese herbal medicine, Danggui Longhui Wan, and is used for treatment of CML in China [144]. Indirubin and its derivatives have activity as cyclin-dependent kinase inhibitors (CDK), competing with ATP for binding in the catalytic domain of CDK [145]. Indirubin induces growth arrest and apoptosis in several tumor cell lines [60]. Recently, it was shown that the indirubin derivatives E564, E728, and E804 have activity as Src inhibitors, blocking constitutive STAT3 signaling and inducing apoptosis in human breast and prostate cancer cells, with concomitant decreases in the levels of the anti-apoptotic proteins Mcl-1 and survivin [146]. Two other compounds with activity as CDK inhibitors were also found to be STAT3 or STAT5 inhibitors [147, 148]. Flavopiridol is a flavonoid derivative of the natural product rohitukine isolated from the bark of the tree Dysoxylum binectariferum, and it is currently being evaluated in clinical trials for a variety of malignancies [149, 150]. Flavopiridol has activity as a CDK inhibitor [151], but it was also reported that it can inhibit STAT3–DNA interactions and disrupt STAT3directed gene transcription, leading to downregulation of Mcl-1 [147]. Roscovitine is not a natural compound, but it is a purine analogue and CDK inhibitor that was recently reported to inhibit STAT5 phosphorylation and induce apoptosis in MT-2 T-cells transformed by the human T-cell leukemia virus-1 (HTLV-1) [148]. Magnolol is the active component isolated from the Chinese herb Hou p’u (Magnolia officinalis). Magnolol has potential therapeutic benefit as an anti-inflammatory and anticancer agent [152, 153]. In one study, magnolol was shown to inhibit IL-6-induced STAT3 phosphorylation, DNA binding, and transcriptional activity in endothelial cells, demonstrating that magnolol possesses anti-STAT3 activity [154]. Magnolol did not affect the phosphorylation status of Jak1/2 or Erk 1/2 [154]. Other compounds derived from natural sources that were demonstrated to have STAT3 inhibitory activity are resveratrol and silibinin [155, 156]. Resveratrol is a phytoalexin with antioxidant and anti-inflammatory properties that inhibits Src TK activity and blocks constitutive STAT3 protein activation in malignant cells, leading to growth arrest and apoptosis [156]. Silibinin is a flavanone isolated from the milk thistle (Silybum marianum) that inhibits constitutively active STAT3 and has been shown to induce apoptosis in vitro in DU145 prostate cancer cells [155].

13

Targeting Stats in Cancer

315

Other Compounds with STAT3 Inhibitory Activity Sulindac is a nonsteroidal anti-inflammatory agent currently being studied for chemoprevention of colon cancer. Some of its antineoplastic activity may be due to inhibition of STAT3 [157]. Sulindac decreases the level of phosphorylated and unphosphorylated STAT3 and induces growth arrest and apoptosis in head and neck squamous cell carcinoma cell lines [157, 158]. Survivin is one of the downstream mediators of STAT3 that is targeted by sulindac, as forced reexpression of survivin partially reversed the effects of STAT3 inhibition [159]. Arsenic trioxide (ATO) is currently used in the treatment of relapsed APL [160]. In vitro, ATO induces growth arrest and apoptosis of non-APL AML cells from patient samples and from the AML cell line HEL, which has constitutive STAT3 signaling [161]. This effect was preceded by the downregulation of activated STAT proteins as early as 6 h after treatment with ATO [161]. Blockage of STAT activity was due to the inhibition of several TKs that are responsible for STAT3 and STAT5 phosphorylation, including Jaks, ZNF198-fibroblast growth factor receptor-1 (FGFR1), FLT3, and Bcr-Abl [161]. Another study showed that sodium arsenite inhibited the Jak–STAT pathway via direct inhibition of Jak by arsenite [162]. This effect was independent of arsenite activation of MAPK pathways [162]. ATO may also induce demethylation of the Jak–STAT pathway inhibitor SOCS1 [163]. Even though ATO is already being used for the treatment of APL, more studies are necessary to confirm its activity in other neoplasms [164, 165]. Triterpenoids are natural compounds synthesized in plants by the cyclization of squalene [166]. More than 20,000 triterpenoids are known to occur in nature, and two compounds have known anti-inflammatory properties, oleanoic acid and ursocholic acid [167, 168]. In an effort to improve their potency, modifications were made to the basic triterpenoid structure to obtain synthetic triterpenoids, a new class of noncytotoxic and multifunctional drugs that have several potential applications for anticancer and anti-inflammatory therapy [169]. The synthetic triterpenoid CDDO-imidazole (1-[2-cyano-3,12-dioxooleana-1,9-dien-28-oyl]imidazole) rapidly (within 30–60 min) blocks constitutive or IL-6-induced STAT3 and STAT5 activity in human multiple myeloma and lung cancer cells [170]. Induction of apoptosis occurred after brief (4 h) exposure to CDDO-imidazole [170]. The level of phosphorylated STAT3 in myeloma cells was almost undetectable after only 1 h of treatment with low concentrations (0.25 mM) of CDDOimidazole, indicating the high potency of this compound [170]. CDDO-imidazole induced the expression of the Jak–STAT pathway inhibitors SOCS1 and SHP-1, which may contribute to its activity [170]. Another study demonstrated that the similar compound CDDO-methyl ester blocked the Jak1–STAT3 pathway by forming adducts with Jak1 at C1077 in the kinase domain and inhibiting Jak1 activity and by binding directly to STAT3 and inhibiting the formation of STAT3 dimers [171]. Clinical studies with CDDO and other synthetic triterpenoids are ongoing.

316

F.P.S. Santos et al.

Activators of STAT Inhibitors Activation of STAT3 and STAT5 inhibitors is another therapeutic approach that could be exploited. The compound FTY720 (2-amino-2-[2-(4-octylphenyl)ethyl]1,3-propanediol hydrochloride; also known as fingolimod) is an immunomodulator in clinical trials for patients with multiple sclerosis that was shown to activate the phosphatase PP2A and to inhibit the activity of both Bcr-Abl and STAT5 in vitro and in vivo in CML cell lines that are sensitive or resistant to imatinib/dasatinib [172]. Peroxisome proliferator-activated receptor g (PPARg) is a member of the nuclear receptor superfamily. There are several PPARg ligands, including the prostanoid 15-deoxy-D-12,14-prostaglandin J2 (15d-PGJ2) and the antidiabetic thiazolidinedione rosiglitazone [173, 174]. Both compounds were shown to inhibit the phosphorylation and activity of Jak1 and Jak2 as well as STAT1 and STAT3 in glial cells [175]. Rosiglitazone induces transcription of the Jak–STAT pathway inhibitors SOCS1 and SOCS3. 15d-PGJ2 has the same effect and also induces activation of SHP-2, another negative regulator of the Jak–STAT pathway [175]. Of note, these effects were independent of PPARg activation [175]. In vitro, both rosiglitazone and 15d-PGJ2 were shown to induce differentiation and apoptosis of myeloid (U937 and HL-60) and lymphoid (Su-DHL, Sup-M2, Ramos, Raji, Hodgkin’s cell lines, and primary chronic lymphocytic leukemia) leukemic cells [176].

Conclusion It is clear that STATs are not mere transcription factors for cytokine receptors but rather central molecules in several intracellular signaling pathways. Constitutive activation of STATs contributes to the generation of the malignant phenotype in cancer cells. There are currently no approved agents for targeting STATs, but it is expected that with further development, new compounds will become available that will hopefully improve the outcomes for cancer patients.

References 1. Fu, X.Y., Schindler, C., Improta, T., Aebersold, R., and Darnell, J.E., Jr. 1992, Proc Natl Acad Sci USA, 89, 7840–3. 2. Schindler, C., Fu, X.Y., Improta, T., Aebersold, R., and Darnell, J.E., Jr. 1992, Proc Natl Acad Sci USA, 89, 7836–9. 3. Darnell, J.E., Jr., Kerr, I.M., and Stark, G.R. 1994, Science, 264, 1415–21. 4. Levy, D.E. and Darnell, J.E., Jr. 2002, Nat Rev Mol Cell Biol, 3, 651–62. 5. Yu, H. and Jove, R. 2004, Nat Rev Cancer, 4, 97–105. 6. Ihle, J.N. 2001, Curr Opin Cell Biol, 13, 211–7. 7. Ward, A.C., Touw, I., and Yoshimura, A. 2000, Blood, 95, 19–29. 8. Bromberg, J. 2002, J Clin Invest, 109, 1139–42.

13

Targeting Stats in Cancer

317

9. Bromberg, J. and Darnell, J.E., Jr. 2000, Oncogene, 19, 2468–73. 10. Weber-Nordt, R.M., Egen, C., Wehinger, J., Ludwig, W., Gouilleux-Gruart, V., Mertelsmann, R., et al. 1996, Blood, 88, 809–16. 11. Yu, C.L., Jove, R., and Burakoff, S.J. 1997, J Immunol, 159, 5206–10. 12. Zhang, Q., Nowak, I., Vonderheid, E.C., Rook, A.H., Kadin, M.E., Nowell, P.C., et al. 1996, Proc Natl Acad Sci USA, 93, 9148–53. 13. Zhang, Q., Raghunath, P.N., Xue, L., Majewski, M., Carpentieri, D.F., Odum, N., et al. 2002, J Immunol, 168, 466–74. 14. Darnell, J.E., Jr. 1997, Science, 277, 1630–5. 15. Wilks, A.F. 1989, Proc Natl Acad Sci USA, 86, 1603–7. 16. Leonard, W.J. and O’Shea, J. 1998, Annu Rev Immunol, 16, 293–322. 17. Shuai, K., Ziemiecki, A., Wilks, A.F., Harpur, A.G., Sadowski, H.B., Gilman, M.Z., et al. 1993, Nature, 366, 580–3. 18. Schindler, C., Levy, D.E., and Decker, T. 2007, J Biol Chem, 282, 20059–63. 19. Decker, T. and Kovarik, P. 2000, Oncogene, 19, 2628–37. 20. Wen, Z., Zhong, Z., and Darnell, J.E., Jr. 1995, Cell, 82, 241–50. 21. Horvath, C.M. 2000, Trends Biochem Sci, 25, 496–502. 22. Shuai, K. 2000, Oncogene, 19, 2638–44. 23. Shuai, K. and Liu, B. 2005, Nat Rev Immunol, 5, 593–605. 24. Chung, C.D., Liao, J., Liu, B., Rao, X., Jay, P., Berta, P., et al. 1997, Science, 278, 1803–5. 25. Liu, B., Liao, J., Rao, X., Kushner, S.A., Chung, C.D., Chang, D.D., et al. 1998, Proc Natl Acad Sci USA, 95, 10626–31. 26. Arora, T., Liu, B., He, H., Kim, J., Murphy, T.L., Murphy, K.M., et al. 2003, J Biol Chem, 278, 21327–30. 27. Liu, B., Gross, M., ten Hoeve, J., and Shuai, K. 2001, Proc Natl Acad Sci USA, 98, 3203–7. 28. Ungureanu, D., Vanhatupa, S., Kotaja, N., Yang, J., Aittomaki, S., Janne, O.A., et al. 2003, Blood, 102, 3311–3. 29. Ungureanu, D., Vanhatupa, S., Gronholm, J., Palvimo, J.J., and Silvennoinen, O. 2005, Blood, 106, 224–6. 30. Kotaja, N., Karvonen, U., Janne, O.A., and Palvimo, J.J. 2002, Mol Cell Biol, 22, 5222–34. 31. Rogers, R.S., Horvath, C.M., and Matunis, M.J. 2003, J Biol Chem, 278, 30091–7. 32. Yu, C.L., Meyer, D.J., Campbell, G.S., Larner, A.C., Carter-Su, C., Schwartz, J., et al. 1995, Science, 269, 81–3. 33. Bromberg, J.F., Horvath, C.M., Besser, D., Lathem, W.W., and Darnell, J.E., Jr. 1998, Mol Cell Biol, 18, 2553–8. 34. Turkson, J., Bowman, T., Garcia, R., Caldenhoven, E., De Groot, R.P., and Jove, R. 1998, Mol Cell Biol, 18, 2545–52. 35. Frank, D.A., Mahajan, S., and Ritz, J. 1997, J Clin Invest, 100, 3140–8. 36. Hazan-Halevy, I., Harris, D., Liu, Z., Ferrajoli, A., Keating, M.J., and Estrov, Z. 2008, Blood, 112, Abstract 1058. 37. Qin, H.R., Kim, H.J., Kim, J.Y., Hurt, E.M., Klarmann, G.J., Kawasaki, B.T., et al. 2008, Cancer Res, 68, 7736–41. 38. Herman, J.G. and Baylin, S.B. 2003, N Engl J Med, 349, 2042–54. 39. Ghoshal Gupta, S., Baumann, H., and Wetzler, M. 2008, Leuk Res, 32, 1005–14. 40. Alvarez, J.V., Febbo, P.G., Ramaswamy, S., Loda, M., Richardson, A., and Frank, D.A. 2005, Cancer Res, 65, 5054–62. 41. Cory, S. and Adams, J.M. 2002, Nat Rev Cancer, 2, 647–56. 42. Bromberg, J.F., Wrzeszczynska, M.H., Devgan, G., Zhao, Y., Pestell, R.G., Albanese, C., et al. 1999, Cell, 98, 295–303. 43. Catlett-Falcone, R., Landowski, T.H., Oshiro, M.M., Turkson, J., Levitzki, A., Savino, R., et al. 1999, Immunity, 10, 105–15. 44. Catlett-Falcone, R., Dalton, W.S., and Jove, R. 1999, Curr Opin Oncol, 11, 490–6. 45. Malumbres, M. and Barbacid, M. 2001, Nat Rev Cancer, 1, 222–31.

318

F.P.S. Santos et al.

46. Pelengaris, S., Khan, M., and Evan, G. 2002, Nat Rev Cancer, 2, 764–76. 47. Leone, G., Sears, R., Huang, E., Rempel, R., Nuckolls, F., Park, C.H., et al. 2001, Mol Cell, 8, 105–13. 48. Mateyak, M.K., Obaya, A.J., and Sedivy, J.M. 1999, Mol Cell Biol, 19, 4672–83. 49. O’Hagan, R.C., Ohh, M., David, G., de Alboran, I.M., Alt, F.W., Kaelin, W.G., Jr., et al. 2000, Genes Dev, 14, 2185–91. 50. Bouchard, C., Thieke, K., Maier, A., Saffrich, R., Hanley-Hyde, J., Ansorge, W., et al. 1999, Embo J, 18, 5321–33. 51. Coller, H.A., Grandori, C., Tamayo, P., Colbert, T., Lander, E.S., Eisenman, R.N., et al. 2000, Proc Natl Acad Sci USA, 97, 3260–5. 52. Hermeking, H., Rago, C., Schuhmacher, M., Li, Q., Barrett, J.F., Obaya, A.J., et al. 2000, Proc Natl Acad Sci USA, 97, 2229–34. 53. Huang, M., Dorsey, J.F., Epling-Burnette, P.K., Nimmanapalli, R., Landowski, T.H., Mora, L.B., et al. 2002, Oncogene, 21, 8804–16. 54. Kiuchi, N., Nakajima, K., Ichiba, M., Fukada, T., Narimatsu, M., Mizuno, K., et al. 1999, J Exp Med, 189, 63–73. 55. Vogelstein, B., Lane, D., and Levine, A.J. 2000, Nature, 408, 307–10. 56. Niu, G., Wright, K.L., Ma, Y., Wright, G.M., Huang, M., Irby, R., et al. 2005, Mol Cell Biol, 25, 7432–40. 57. Bergers, G. and Benjamin, L.E. 2003, Nat Rev Cancer, 3, 401–10. 58. Chen, Z. and Han, Z.C. 2008, Med Res Rev, 28, 185–200. 59. Niu, G., Wright, K.L., Huang, M., Song, L., Haura, E., Turkson, J., et al. 2002, Oncogene, 21, 2000–8. 60. Choi, J.H., Ahn, M.J., Park, C.K., Han, H.X., Kwon, S.J., Lee, Y.Y., et al. 2006, Apmis, 114, 619–25. 61. Gong, W., Wang, L., Yao, J.C., Ajani, J.A., Wei, D., Aldape, K.D., et al. 2005, Clin Cancer Res, 11, 1386–93. 62. Hsieh, F.C., Cheng, G., and Lin, J. 2005, Biochem Biophys Res Commun, 335, 292–9. 63. Wei, D., Le, X., Zheng, L., Wang, L., Frey, J.A., Gao, A.C., et al. 2003, Oncogene, 22, 319–29. 64. Xie, T.X., Huang, F.J., Aldape, K.D., Kang, S.H., Liu, M., Gershenwald, J.E., et al. 2006, Cancer Res, 66, 3188–96. 65. Masuda, M., Ruan, H.Y., Ito, A., Nakashima, T., Toh, S., Wakasaki, T., et al. 2007, Oral Oncol, 43, 785–90. 66. Wei, L.H., Kuo, M.L., Chen, C.A., Chou, C.H., Lai, K.B., Lee, C.N., et al. 2003, Oncogene, 22, 1517–27. 67. Suganami, E., Takagi, H., Ohashi, H., Suzuma, K., Suzuma, I., Oh, H., et al. 2004, Diabetes, 53, 2443–8. 68. Valdembri, D., Serini, G., Vacca, A., Ribatti, D., and Bussolino, F. 2002, Faseb J, 16, 225–7. 69. Huang, S.P., Wu, M.S., Shun, C.T., Wang, H.P., Lin, M.T., Kuo, M.L., et al. 2004, J Biomed Sci, 11, 517–27. 70. Bartoli, M., Platt, D., Lemtalsi, T., Gu, X., Brooks, S.E., Marrero, M.B., et al. 2003, Faseb J, 17, 1562–4. 71. Yahata, Y., Shirakata, Y., Tokumaru, S., Yamasaki, K., Sayama, K., Hanakawa, Y., et al. 2003, J Biol Chem, 278, 40026–31. 72. Dechow, T.N., Pedranzini, L., Leitch, A., Leslie, K., Gerald, W.L., Linkov, I., et al. 2004, Proc Natl Acad Sci USA, 101, 10602–7. 73. Xie, T.X., Wei, D., Liu, M., Gao, A.C., Ali-Osman, F., Sawaya, R., et al. 2004, Oncogene, 23, 3550–60. 74. Dunn, G.P., Bruce, A.T., Ikeda, H., Old, L.J., and Schreiber, R.D. 2002, Nat Immunol, 3, 991–8. 75. Pardoll, D. 2003, Annu Rev Immunol, 21, 807–39. 76. Gabrilovich, D. 2004, Nat Rev Immunol, 4, 941–52. 77. Zou, W. 2005, Nat Rev Cancer, 5, 263–74.

13

Targeting Stats in Cancer

319

78. Kortylewski, M., Kujawski, M., Wang, T., Wei, S., Zhang, S., Pilon-Thomas, S., et al. 2005, Nat Med, 11, 1314–21. 79. Kortylewski, M., Xin, H., Kujawski, M., Lee, H., Liu, Y., Harris, T., et al. 2009, Cancer Cell, 15, 114–23. 80. Burdelya, L., Kujawski, M., Niu, G., Zhong, B., Wang, T., Zhang, S., et al. 2005, J Immunol, 174, 3925–31. 81. Welte, T., Zhang, S.S., Wang, T., Zhang, Z., Hesslein, D.G., Yin, Z., et al. 2003, Proc Natl Acad Sci USA, 100, 1879–84. 82. Hoentjen, F., Sartor, R.B., Ozaki, M., and Jobin, C. 2005, Blood, 105, 689–96. 83. Stahl, N., Farruggella, T.J., Boulton, T.G., Zhong, Z., Darnell, J.E., Jr., and Yancopoulos, G.D. 1995, Science, 267, 1349–53. 84. Gerhartz, C., Heesel, B., Sasse, J., Hemmann, U., Landgraf, C., Schneider-Mergener, J., et al. 1996, J Biol Chem, 271, 12991–8. 85. Wiederkehr-Adam, M., Ernst, P., Muller, K., Bieck, E., Gombert, F.O., Ottl, J., et al. 2003, J Biol Chem, 278, 16117–28. 86. Wang, T., Niu, G., Kortylewski, M., Burdelya, L., Shain, K., Zhang, S., et al. 2004, Nat Med, 10, 48–54. 87. Song, H., Wang, R., Wang, S., and Lin, J. 2005, Proc Natl Acad Sci USA, 102, 4700–5. 88. Siddiquee, K., Zhang, S., Guida, W.C., Blaskovich, M.A., Greedy, B., Lawrence, H.R., et al. 2007, Proc Natl Acad Sci USA, 104, 7391–6. 89. Schust, J., Sperl, B., Hollis, A., Mayer, T.U., and Berg, T. 2006, Chem Biol, 13, 1235–42. 90. Hao, W., Hu, Y., Niu, C., Huang, X., Chang, C.P., Gibbons, J., et al. 2008, Bioorg Med Chem Lett, 18, 4988–92. 91. Turkson, J., Zhang, S., Mora, L.B., Burns, A., Sebti, S., and Jove, R. 2005, J Biol Chem, 280, 32979–88. 92. Weidler, M., Rether, J., Anke, T., and Erkel, G. 2000, FEBS Lett, 484, 1–6. 93. Hellsten, R., Johansson, M., Dahlman, A., Dizeyi, N., Sterner, O., and Bjartell, A. 2008, Prostate, 68, 269–80. 94. Baines, I.C. and Colas, P. 2006, Drug Discov Today, 11, 334–41. 95. Nagel-Wolfrum, K., Buerger, C., Wittig, I., Butz, K., Hoppe-Seyler, F., and Groner, B. 2004, Mol Cancer Res, 2, 170–82. 96. Zhang, X. and Darnell, J.E., Jr. 2001, J Biol Chem, 276, 33576–81. 97. Timofeeva, O.A., Gaponenko, V., Lockett, S.J., Tarasov, S.G., Jiang, S., Michejda, C.J., et al. 2007, ACS Chem Biol, 2, 799–809. 98. Chan, J.H., Lim, S., and Wong, W.S. 2006, Clin Exp Pharmacol Physiol, 33, 533–40. 99. Li, W.C., Ye, S.L., Sun, R.X., Liu, Y.K., Tang, Z.Y., Kim, Y., et al. 2006, Clin Cancer Res, 12, 7140–8. 100. Barton, B.E., Karras, J.G., Murphy, T.F., Barton, A., and Huang, H.F. 2004, Mol Cancer Ther, 3, 11–20. 101. Gleave, M.E. and Monia, B.P. 2005, Nat Rev Cancer, 5, 468–79. 102. Konnikova, L., Kotecki, M., Kruger, M.M., and Cochran, B.H. 2003, BMC Cancer, 3, 23. 103. Ling, X. and Arlinghaus, R.B. 2005, Cancer Res, 65, 2532–6. 104. Tomita, N., Ogihara, T., and Morishita, R. 2003, Curr Drug Targets, 4, 603–8. 105. Leong, P.L., Andrews, G.A., Johnson, D.E., Dyer, K.F., Xi, S., Mai, J.C., et al. 2003, Proc Natl Acad Sci USA, 100, 4138–43. 106. Zhang, X., Zhang, J., Wei, H., and Tian, Z. 2007, Oncol Rep, 17, 1377–82. 107. Sun, X., Zhang, J., Wang, L., and Tian, Z. 2008, Cancer Lett, 262, 201–13. 108. Gu, J., Li, G., Sun, T., Su, Y., Zhang, X., Shen, J., et al. 2008, J Neurooncol, 89, 9–17. 109. Zhang, X., Zhang, J., Wang, L., Wei, H., and Tian, Z. 2007, BMC Cancer, 7, 149. 110. Sen, M., Tosca, P.J., Zwayer, C., Ryan, M.J., Johnson, J.D., Knostman, K.A., et al. 2008, Cancer Chemother Pharmacol. 111. Williamson, J.R. 1994, Annu Rev Biophys Biomol Struct, 23, 703–30. 112. Jing, N., Li, Y., Xu, X., Sha, W., Li, P., Feng, L., et al. 2003, DNA Cell Biol, 22, 685–96. 113. Zhu, Q. and Jing, N. 2007, J Comput Aided Mol Des, 21, 641–8.

320

F.P.S. Santos et al.

114. Jing, N., Li, Y., Xiong, W., Sha, W., Jing, L., and Tweardy, D.J. 2004, Cancer Res, 64, 6603–9. 115. Demetri, G.D., von Mehren, M., Blanke, C.D., Van den Abbeele, A.D., Eisenberg, B., Roberts, P.J., et al. 2002, N Engl J Med, 347, 472–80. 116. O’Brien, S.G., Guilhot, F., Larson, R.A., Gathmann, I., Baccarani, M., Cervantes, F., et al. 2003, N Engl J Med, 348, 994–1004. 117. Shepherd, F.A., Rodrigues Pereira, J., Ciuleanu, T., Tan, E.H., Hirsh, V., Thongprasert, S., et al. 2005, N Engl J Med, 353, 123–32. 118. Geyer, C.E., Forster, J., Lindquist, D., Chan, S., Romieu, C.G., Pienkowski, T., et al. 2006, N Engl J Med, 355, 2733–43. 119. Talpaz, M., Shah, N.P., Kantarjian, H., Donato, N., Nicoll, J., Paquette, R., et al. 2006, N Engl J Med, 354, 2531–41. 120. Krause, D.S. and Van Etten, R.A. 2005, N Engl J Med, 353, 172–87. 121. Aggarwal, B.B., Sethi, G., Ahn, K.S., Sandur, S.K., Pandey, M.K., Kunnumakkara, A.B., et al. 2006, Ann N Y Acad Sci, 1091, 151–69. 122. Blaskovich, M.A., Sun, J., Cantor, A., Turkson, J., Jove, R., and Sebti, S.M. 2003, Cancer Res, 63, 1270–9. 123. Chen, J.C., Chiu, M.H., Nie, R.L., Cordell, G.A., and Qiu, S.X. 2005, Nat Prod Rep, 22, 386–99. 124. Rao, M.M., Meshulam, H., and Lavie, D. 1974, J. Chem. Soc. Perkin Trans. I 22, 2552–2556. 125. Witkowski, A., Woynarowska, B., and Konopa, J. 1984, Biochem Pharmacol, 33, 995–1004. 126. Gitter, S., Gallily, R., Shohat, B., and Lavie, D. 1961, Cancer Res, 21, 516–21. 127. Nefedova, Y., Nagaraj, S., Rosenbauer, A., Muro-Cacho, C., Sebti, S.M., and Gabrilovich, D.I. 2005, Cancer Res, 65, 9525–35. 128. Molavi, O., Ma, Z., Hamdy, S., Lai, R., Lavasanifar, A., and Samuel, J. 2008, Immunol Cell Biol, 86, 506–14. 129. Molavi, O., Ma, Z., Mahmud, A., Alshamsan, A., Samuel, J., Lai, R., et al. 2008, Int J Pharm, 347, 118–27. 130. Sun, J., Blaskovich, M.A., Jove, R., Livingston, S.K., Coppola, D., and Sebti, S.M. 2005, Oncogene, 24, 3236–45. 131. Liu, T., Zhang, M., Zhang, H., Sun, C., Yang, X., Deng, Y., et al. 2008, Eur J Pharmacol, 587, 78–84. 132. Zhang, M., Zhang, H., Sun, C., Shan, X., Yang, X., Li-Ling, J., et al. 2009, Cancer Chemother Pharmacol, 63, 635–42. 133. Ammon, H.P. and Wahl, M.A. 1991, Planta Med, 57, 1–7. 134. Kunnumakkara, A.B., Anand, P., and Aggarwal, B.B. 2008, Cancer Lett, 269, 199–225. 135. Kim, H.Y., Park, E.J., Joe, E.H., and Jou, I. 2003, J Immunol, 171, 6072–9. 136. Bharti, A.C., Donato, N., and Aggarwal, B.B. 2003, J Immunol, 171, 3863–71. 137. Bharti, A.C., Donato, N., Singh, S., and Aggarwal, B.B. 2003, Blood, 101, 1053–62. 138. Bharti, A.C., Shishodia, S., Reuben, J.M., Weber, D., Alexanian, R., Raj-Vadhan, S., et al. 2004, Blood, 103, 3175–84. 139. Adams, B.K., Cai, J., Armstrong, J., Herold, M., Lu, Y.J., Sun, A., et al. 2005, Anticancer Drugs, 16, 263–75. 140. Adams, B.K., Ferstl, E.M., Davis, M.C., Herold, M., Kurtkaya, S., Camalier, R.F., et al. 2004, Bioorg Med Chem, 12, 3871–83. 141. Hsu, C.H., Chuang, S.E., Hergenhahn, M., Kuo, M.L., Lin, J.K., Hsieh, C.Y., et al. 2002, Gan To Kagaku Ryoho, 29 Suppl 1, 194–200. 142. Bhutani, M., Pathak, A.K., Nair, A.S., Kunnumakkara, A.B., Guha, S., Sethi, G., et al. 2007, Clin Cancer Res, 13, 3024–32. 143. Lee, H.K., Seo, I.A., Shin, Y.K., Park, J.W., Suh, D.J., and Park, H.T. 2009, Biochem Biophys Res Commun. 144. Xiao, Z., Hao, Y., Liu, B., and Qian, L. 2002, Leuk Lymphoma, 43, 1763–8. 145. Hoessel, R., Leclerc, S., Endicott, J.A., Nobel, M.E., Lawrie, A., Tunnah, P., et al. 1999, Nat Cell Biol, 1, 60–7.

13

Targeting Stats in Cancer

321

146. Nam, S., Buettner, R., Turkson, J., Kim, D., Cheng, J.Q., Muehlbeyer, S., et al. 2005, Proc Natl Acad Sci USA, 102, 5998–6003. 147. Lee, Y.K., Isham, C.R., Kaufman, S.H., and Bible, K.C. 2006, Mol Cancer Ther, 5, 138–48. 148. Mohapatra, S., Chu, B., Wei, S., Djeu, J., Epling-Burnette, P.K., Loughran, T., et al. 2003, Cancer Res, 63, 8523–30. 149. Byrd, J.C., Lin, T.S., Dalton, J.T., Wu, D., Phelps, M.A., Fischer, B., et al. 2007, Blood, 109, 399–404. 150. Shah, M.A., Kortmansky, J., Motwani, M., Drobnjak, M., Gonen, M., Yi, S., et al. 2005, Clin Cancer Res, 11, 3836–45. 151. Kim, K.S., Sack, J.S., Tokarski, J.S., Qian, L., Chao, S.T., Leith, L., et al. 2000, J Med Chem, 43, 4126–34. 152. Shen, Y.C., Sung, Y.J., and Chen, C.F. 1998, Eur J Pharmacol, 343, 79–86. 153. Kong, C.W., Tsai, K., Chin, J.H., Chan, W.L., and Hong, C.Y. 2000, Shock, 13, 24–8. 154. Chen, S.C., Chang, Y.L., Wang, D.L., and Cheng, J.J. 2006, Br J Pharmacol, 148, 226–32. 155. Agarwal, C., Tyagi, A., Kaur, M., and Agarwal, R. 2007, Carcinogenesis, 28, 1463–70. 156. Kotha, A., Sekharam, M., Cilenti, L., Siddiquee, K., Khaled, A., Zervos, A.S., et al. 2006, Mol Cancer Ther, 5, 621–9. 157. Nikitakis, N.G., Hamburger, A.W., and Sauk, J.J. 2002, Cancer Res, 62, 1004–7. 158. Scheper, M.A., Nikitakis, N.G., Chaisuparat, R., Montaner, S., and Sauk, J.J. 2007, Neoplasia, 9, 192–9. 159. Scheper, M.A., Nikitakis, N.G., and Sauk, J.J. 2007, Int J Oral Maxillofac Surg, 36, 632–9. 160. Sanz, M.A., Grimwade, D., Tallman, M.S., Lowenberg, B., Fenaux, P., Estey, E.H., et al. 2009, Blood, 113, 1875–91. 161. Wetzler, M., Brady, M.T., Tracy, E., Li, Z.R., Donohue, K.A., O’Loughlin, K.L., et al. 2006, Clin Cancer Res, 12, 6817–25. 162. Cheng, H.Y., Li, P., David, M., Smithgall, T.E., Feng, L., and Lieberman, M.W. 2004, Oncogene, 23, 3603–12. 163. Wang, M.M., Zhu, Q., Ren, Z.H., Zou, L.F., Dou, H.J., and Hu, J.P. 2008, Zhongguo Shi Yan Xue Ye Xue Za Zhi, 16, 1064–8. 164. Parmar, S., Rundhaugen, L.M., Boehlke, L., Riley, M., Nabhan, C., Raji, A., et al. 2004, Leuk Res, 28, 909–19. 165. Hussein, M.A., Saleh, M., Ravandi, F., Mason, J., Rifkin, R.M., and Ellison, R. 2004, Br J Haematol, 125, 470–6. 166. Phillips, D.R., Rasbery, J.M., Bartel, B., and Matsuda, S.P. 2006, Curr Opin Plant Biol, 9, 305–14. 167. Huang, M.T., Ho, C.T., Wang, Z.Y., Ferraro, T., Lou, Y.R., Stauber, K., et al. 1994, Cancer Res, 54, 701–8. 168. Nishino, H., Nishino, A., Takayasu, J., Hasegawa, T., Iwashima, A., Hirabayashi, K., et al. 1988, Cancer Res, 48, 5210–5. 169. Liby, K.T., Yore, M.M., and Sporn, M.B. 2007, Nat Rev Cancer, 7, 357–69. 170. Liby, K., Voong, N., Williams, C.R., Risingsong, R., Royce, D.B., Honda, T., et al. 2006, Clin Cancer Res, 12, 4288–93. 171. Ahmad, R., Raina, D., Meyer, C., and Kufe, D. 2008, Cancer Res, 68, 2920–6. 172. Neviani, P., Santhanam, R., Oaks, J.J., Eiring, A.M., Notari, M., Blaser, B.W., et al. 2007, J Clin Invest, 117, 2408–21. 173. Forman, B.M., Tontonoz, P., Chen, J., Brun, R.P., Spiegelman, B.M., and Evans, R.M. 1995, Cell, 83, 803–12. 174. Lehmann, J.M., Moore, L.B., Smith-Oliver, T.A., Wilkison, W.O., Willson, T.M., and Kliewer, S.A. 1995, J Biol Chem, 270, 12953–6. 175. Park, E.J., Park, S.Y., Joe, E.H., and Jou, I. 2003, J Biol Chem, 278, 14747–52. 176. Konopleva, M., Elstner, E., McQueen, T.J., Tsao, T., Sudarikov, A., Hu, W., et al. 2004, Mol Cancer Ther, 3, 1249–62.

Index

A Adaptor protein GRB2 anticancer drugs b-bend configuration, 14 chronic myelogenous leukemia, 16 phosphatase resistance, 14 SH3 domain target selectivity, 15 cancer, 5–6 cell signaling epithelial morphogenesis, 5 Src homology 2 (SH2) domain, 4 invasion and metastasis actin-based cell motility, 10 ADAMs inhibitors, 8 angiogenesis and dissemination, 12–13 Caldesmon, 11 ECM, 7 FAK autophosphorylation, 7 Hungtingtin, 11 Merlin-binding proteins, 10 podosomes, 11 scatter factor, 8 ADI. See Arginine deiminase (ADI) A Disintegrin And Metalloprotease (ADAMs), 8 Amplified in breast cancer 1 (AIB1) biopsies and animal models androgen receptor (AR) coactivator, 141 cancer progression, 141 colony formation, 143 FISH, 140 mammary tumors, 142 pancreatic adenocarcinomas, 141 cancer etiology and signal transduction, 129–130

cell cycle regulation anti-estrogens, 151 cyclin D1, 151 endocrine resistance, 152 Rb, E2F1, 150 hormone-dependent mechanisms endocrine tumors, 145 ER function, 144 estrogen-dependent transcription, 145 sex steroids, 143 signaling pathways, 144 hormone-independent mechanisms ErbB/HER signaling pathway, 147 IGF/PI3K/AKT signaling pathway, 145–146 NF-kB signaling pathway, 147–148 Rb/E2F1, 148 IGF-I, 152–154 invasiveness mechanisms Drosophila, 148 E-cadherin, 149 epithelial-mesenchymal transition, 149 phosphorylation, 150 physiology central nervous system, 138 development, 136–137 energy homeostasis, 138–139 inflammatory process, 139 lymphocytes, 140 reproductive tissues, 137 TRb, 139 predictive and prognostic factors ERa Ser118, 157 HER2/Neu activity, 158 microRNA, 158

M. Chatterjee and K. Kashfi (eds.), Cell Signaling & Molecular Targets in Cancer, DOI 10.1007/978-1-4614-0730-0, © Springer Science+Business Media, LLC 2012

323

324 Amplified in breast cancer 1 (AIB1) (cont.) steroid receptor coactivator/p160 family, 131–136 tamoxifen resistance antiestrogen resistance, 154 endocrine therapy, 154 HER2/Neu, 154 histone acetyltransferase activity, 155 mechanisms, 156 PI3K/AKT pathway, 155 Anti-angiogenic therapy FGFR3 inhibitors, 100 VEGF inhibitors, 99 Arginine ADI-PEG20, 40 antitumor agent, 38 ASS(-) melanoma cells ADI-PEG20, 47–48 anti-angiogenesis effects, 46 ASS expression and cisplatin resistance, 45–46 radiation therapy, 46 citrulline, 38 growth and apoptotic signaling autophagy, ASS(-) cells, 43–44 mTOR signaling, 42–43 RAF/MEK/ERK1/2 signaling, 43 hepatocellular carcinoma, 41 mesothelioma cell lines, 42 mycoplasma, 37 OCT, 38 urea cyle, 39 Arginine deiminase (ADI), 38 Argininosuccinate synthetase (ASS) ASS(-) melanoma cells ADI-PEG20, 47–48 anti-angiogenesis effects, 46 ASS expression and cisplatin resistance, 45–46 radiation therapy, 46 transfection, 39 Aromatase inhibitors. See Lung cancer Arsenic trioxide (ATO), 315 Arylamine N-acetyltransferase 1 acetyl-CoA, 24 androgens, 26 animal models, 29–30 breast cancer progression, 24 cancer, 30–31 drug target, 31–33 gene structure and polymorphism, 25–28 human and prokaryotic structure, 28–29 non-acetylated enzymes, 28 p-aminobenzoylglutamate, 25 phenotypic variation, 27

Index polymorphisms, 23 SNP, 27 therapeutic drugs, 24 transcriptional regulation, 26 xenobiotic-metabolizing function, 23

B Barrett’s epithelium (BE), 112 Bile acids carcinogenesis, 110–111 chemical structures, 110 CRC colon cancer and bile acids, 116–117 COX–2, 117–118 esophagus BE and esophageal adenocarcinoma, 112–113 CDX–2 and Barrett’s epithelium, 113 COX–2 expression and bile acids, 113–114 squamous cell carcinoma, 112 gastric carcinogenesis COX–2, 115–116 duodeno-gastric reflux, 114–115 GPCR and EGFR EGFR transactivation, 119–120 gastrointestinal tumors, 118 metalloprotease, 119

C Caenorhabditis elegans, 3 cAMP-responsive element (CRE), 57 Colorectal cancer (CRC) colon cancer and bile acids, 116–117 COX–2, 117–118 Cortactin, 11 Cyclooxygenase (COX), 111

D Dialkyl disulfide (DADS), 288 Drug target, NAT1 acetyl-CoA, 24 androgens, 26 animal models, 29–30 breast cancer progression, 24 cancer, 30–31 gene structure and polymorphism, 25–28 human and prokaryotic structure, 28–29 human isoenzymes, 23 non-acetylated enzymes, 28 p-aminobenzoylglutamate, 25 phenotypic variation, 27

Index polymorphic acetylation, 24 SNP, 27 therapeutic drugs, 24 transcriptional regulation, 26 xenobiotic-metabolizing function, 23

E Endothelial cells (ECs), 213 Esophagus BE and esophageal adenocarcinoma, 112–113 CDX–2 and Barrett’s epithelium, 113 COX–2 expression and bile acids, 113–114 squamous cell carcinoma, 112 Estrogen receptors ERa, structure of, 192 fulvestrant, 194 gene transcription, 192 human primary lung tumors, 200–201 lung biology, 194 mitochondria, 198–199 SERMs, 193 Extracellular matrix (ECM), 7, 69

F Farnesoid X receptor (FXR), 109 Farnesyltransferase inhibitors (FTIs), 94–95 Fibroblast growth factor receptor 3 (FGFR3), 100 Fluorescence in situ hybridization (FISH), 140 Focal adhesion kinase (FAK), 7

G Gastroesophageal reflux disease (GERD), 112 G protein–coupled receptor (GPCR) EGFR transactivation, 119–120 FXR and colon carcinogenesis chronic inflammation, 122 deoxycholic acid, 122 glucose metabolism, 123 gastrointestinal tumors, 118 metalloprotease, 119 M3 muscarinic receptor and EGFR activation, 120 TGR5 and EGFR transactivation, 120–121 G-Quartet oligodeoxynucleotides (GQ-ODNs) cancer cells, 177 drug delivery and activity in vivo, 177–179 intracellular delivery system DNA anti-cancer agent, 174 endocytosis, 176 potassium-dependent formation, 174 structure-based drug design, 172

325 T40214, Stat3 activation anti-cancer agent, 172 cancer therapy, 182–184 DNA-binding activity, 174 IC50, 173 low-toxicity agents, 185 NSCLC tumors, 187 polyethylenimine, 185 prostate cancer therapy, 179 tumor growth, 182 TUNEL assay, 185 in vivo effects, 181 Growth factor receptor-bound protein 2 (Grb2) signaling adaptor proteins, 1–2 angiogenesis and dissemination, 12–13 Grb family, 2–3 metastatic cascade actin-based cell motility, 10 ADAMs inhibitors, 8 caldesmon, 11 ECM, 7 FAK autophosphorylation, 7 Hungtingtin, 11 Merlin-binding proteins, 10 podosomes, 11 scatter factor, 8

H Heat shock proteins (HSPs), 95 Hedgehog (HH), 67 Humoral hypercalcemia of malignancy (HHM), 53 Huntington’s Disease, 11

I Insulin-like growth factor-I (IGF-I), 92 Interleukin–6 (IL–6), 89 Isoobtusilactone (IOA), 287

L L-asparagine, 37 Listeria monocytogenes, 9 Lung cancer aromatase immunohistochemistry, 205 postmenopausal breast cancer patients, 205 tamoxifen, 206 estrogens endogenous and exogenous, 195–196 estrogen-responsive tissue, 194 mitochondria, 198–199

326 Lung cancer (cont.) membrane-initiated steroid signaling, 197–198 NSCLC, 196–197

M Microparticles (MP) dissemination blood product storage and processing, 214–215 cancer-associated thrombosis, TF antigen, 228–229 hypercoagulability, 228 mouse models, 229 procoagulant activity, 229 cisplatin, 214 hemostatic factors, 213–214 hypoxia, 214 inflammatory cytokines, 214 measurement of, 215 mechanism of, 212–213 platelet activation factors, 214 platelet-derived MP, 225–227 thrombus development, 234–235 tumor cell-derived MPs angiogenesis, 218–222 immune escape, 222–223 metastatic foci, 223–225 tumor invasion and metastasis, 217–218 Mitogen-activated protein kinase (MAPK), 62 Multiple myeloma (MM) bone marrow microenvironment immunomodulatory therapies, 98–99 epidemiology and aetiology, 87 FGFR3 inhibitors, 100 growth-promoting signalling cascades Bcl–2 protein family, 93–94 FTIs, 94–95 IGF-I signalling, 92 IL–6 signalling system, 89–92 NF-kB pathway, 92–93 PI3K/AKT pathway, 93 HDAC inhibitors, 97 Hsp90 inhibitors, 95–96 pathobiology, 87–89 P38 MAP kinase inhibitors, 100 proteasome inhibitors bortezomib, 96 carfilzomib (PR–171), 97 NPI–0052, 97 TGFb inhibitors, 100–101 TRAIL, 95 VEGF inhibitors, 99 Mycoplasma, 37

Index N Non-small cell lung cancers (NSCLC) classical genomic estrogen-signaling pathway, 197 EGFR, 202–204 estrogen receptors, 200–201 membrane-initiated steroid signaling, 197–198 preclinical models, 196–197 subtype-selective ligands, 201–202 N-Wiskott–Aldrich Syndrome protein (N-WASp), 9

O Ornithine carbamoyl transferase (OCT), 38 Oxidative stress (OS) antioxidants, 284–285 carcinogenesis, stages, 275–277 chemotherapeutic agents and preventions acacetin, 288 aminoflavone, 285–286 berberine, 288 curcumin, 286 DADS, 288 decursin, 287 formaldehyde, 287 genipin, 287 IOA, 287 nitric oxide, 286 onion, 289 PAC–1 and UCS1025A, 288 phytochemicals, 289 polyphenols, 286–287 quinoline quinone, 287 radiation, 286 resveratrol, 286 selenium, 288 SOD, 289 chronology, 275 endogenous agents bile acids, 280 interleukin, 279 leukotriene receptor, 278 Nox and Duox enzymes, 278 redox modifications, 279 ROS-mediated cell signaling, 277 exogenous agents alcohol, 283 benzene, 283 curcumin, 283 endosulfan, 283 estrogen quinone, 284 ginkgo biloba, 283 hydrogen peroxide, 282

Index metals and particulates, 281–282 PAHs, 284 ultraviolet radiation, 282 tenets of, 274–275

P Parathyroid hormone–related protein (PTHrP) apoptosis caspase–3 activation, 68 cell proliferation, 67 death receptors and mitochondria, 68 transcriptional activation, p53, 69 bone metastases adhesive interactions, 74 anabolic effects, 76 bone-resorptive cytokines, 75 cancer proliferation, 75 osteolytic metastases, 74 PSA, 77 skeletal metastatic involvement, 73 TGF-b, 76 cancer cell growth adenylate cyclase, 59 b-arrestins, 60 ERK1/2 by Gaq, 65–67 ERK1/2 by Gs, 63–64 functional domains of, 59 GPCR kinases, 59 ligand-independent mechanism, 61 microtubule integrity, 62 NHERF, 60 nuclear translocation, 62 osteosarcoma cells, 61 polyhormone, 58 skin carcinogenesis, 58 gene structure and regulation cancers and tumor-derived cell lines, 55 CpG dinucleotide sequences, 58 CRE, 57 Ets factors, 56 methylation, 58 protein isoforms, 54 Ras–MAPK pathway, 56 transcriptional process, 54 3’-untranslated region, 55 VDRE, 57 invasiveness and integrins basement membrane adhesion and transversion, 71 a6b4 integrin, 72 cell adhesion/migration, 70 ECM, 69 intracrine mechanism, 71

327 LoVo colon cancer cells, 72 matrix proteolysis and degradation, 69 transcriptional mechanism, 70 Phosphatidylethanolamine (PE), 212 Photodynamic therapy (PDT), 280 Platelet-derived MP (PMP), 225 Polycyclic aromatic hydrocarbons (PAHs), 284 Polyethylene glycol (PEG), 40 Prostate-specific antigen (PSA), 77 Proteasome inhibitors bortezomib, 96 carfilzomib (PR–171), 97 NPI–0052, 97 Protein disulfide isomerase (PDI) cellular sources, 232 cryptic and coagulant TF, 231–232 TF, modulation of, 232 thrombus formation, 233 Protein kinase D (PKD) actin cytoskeleton, 246 angiogenesis, 260–261 autoinhibitory mechanism, 245 breast cancer, 260 chemoresistance, 255–256 domain structure and phosphorylationdependent regulation, 246 gastric cancer, 259 oncogenic signaling, 262 PKD3, 247 prostate cancer, 260 signaling mechanisms activation loop phosphorylation, 250–251 initial activation step, 250 phosphorylation and protein interactions, 251–252 tyrosine phosphorylation, 251 tumor cell migration and invasion cargo transport, 258 cell–cell and cell–matrix aggregation, 257 chemotactic stimulus, 257 cortactin, 257 filopodia formation, 257 multiple matrix metalloproteases, 258 multiple myeloma, 259 tumor cell proliferation, 252–253 tumor cell survival signaling, 254–255 upstream activators cleavage, 250 G proteins, 248–249 growth factor receptors, 248 oxidative stress, 249 RhoGTPases, 249 upstream signaling events phorbol esters, 248

328 S Selective estrogen response modifiers (SERMs), 193 Signal transducer and activator of transcription (STAT) activation–inactivation cycle, 300–301 ATO, 315 function, 301–302 natural compounds cucurbitaceae, 312 curcumin, 313 indirubin, 314 magnolol, 314 peptidic inhibitors and peptidomimetic inhibitors, 308 phosphorylation and activation, 302–303 protein inhibitor of activated STAT, 303 serine phosphorylation, 303 small-molecule non-peptidic inhibitors DNA binding domain, 309–310 N-terminal domain, 310–311 oligonucleotides, 311–312 sulindac, 315 transcription factors, 303 triterpenoids, 315 tumor biology angiogenesis, 306–307 constitutive activation, 304–305 immune surveillance mechanisms, 307–308 neoplasms, 304 tumor cell proliferation, 305–306 tyrosine kinase inhibitors, 312 Signal transducer and activator of transcription 3 (STAT3) biological functions, 168 cancer therapy, 168 cytokines and growth factors, 167 DNA inhibitors, 170 G-quartet oligonucleotides, 170–171 immune system, 188 organic compounds, 169 peptide inhibitors, 169–170 protein inhibitors, 170 signaling pathway, 168 T40214 anti-cancer agent, 172 cancer therapy, 182–184 DNA-binding activity, 174 IC50, 173 low-toxicity agents, 185 NSCLC tumors, 187 polyethylenimine, 185 prostate cancer therapy, 179

Index tumor growth, 182 TUNEL assay, 185 in vivo effects, 181 tumor microenvironment, 187 tyrosine residue, 167 Single nucleotide polymorphisms (SNP), 27 Steroid receptor coactivator (SRC) basic helix-loop-helix, 132 DNA-binding domain, 131 HAT activity, 133 non-transcriptional factors, 134–135 nuclear receptors, 131 structural domains, 132 transcriptional activation domain, 133 ubiquitin-proteasome system, 136

T Tamoxifen resistance antiestrogen resistance, 154 endocrine therapy, 154 HER2/Neu, 154 histone acetyltransferase activity, 155 mechanisms, 156 PI3K/AKT pathway, 155 TGN. See Trans-golgi network (TGN) Thalidomide, 98 Thyroid hormone receptor beta (TRb), 139 Tissue factor (TF) antigen, 228–229 cellular origin, 230 cellular sources, 232 cryptic and coagulant, 231–232 hypercoagulability, 228 modulation of, 232 mouse models, 229 procoagulant activity, 229, 231 thrombus formation, 233 Trans-golgi network (TGN), 258 Tumor-derived MPs (TMPs) angiogenesis, 218–222 immune escape, 222–223 metastatic foci, 223–225 tumor invasion and metastasis, 217–218

V Vaccinia virus, 9 Vascular endothelial growth factor (VEGF), 99 Vitamin D–responsive element (VDRE), 57

W Wiskott–Aldrich syndrome protein (WASp), 257