The Pennsylvania State University. The Graduate School. The Huck Institutes of the Life Sciences

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1 , The Pennsylvania State University The Graduate School The Huck Institutes of the Life Sciences ROLE OF THE ENDOPLASMIC RETICULUM STRESS SENSOR IRE1 ALPHA IN ONCOGENE-INDUCED SENECENCE A Dissertation in Integrative Biosciences by Nicholas Blazanin 2015 Nicholas Blazanin Submitted in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy May 2015

2 The dissertation of Nicholas Blazanin was reviewed and approved* by the following: Adam B. Glick Associate Professor of Veterinary and Biomedical Sciences Director of the Center of Molecular Toxicology Chair of Molecular Medicine program Dissertation Advisor Chair of Committee Gary H. Perdew John T. and Paige S. Smith Professor in Agricultural Sciences Curtis J. Omiecinski Professor of Veterinary and Biomedical Sciences H. Thomas and Dorothy Willits Hallowell Chair Wendy Hanna-Rose Associate Professor of Biochemistry and Molecular Biology Peter J. Hudson Willaman Professor of Biology Director of the Huck Institutes of Life Sciences *Signatures are on file in the Graduate School

3 iii Abstract Inositol requiring enzyme 1α (IRE1α) activation during chemically-induced endoplasmic reticulum (ER) stress regulates divergent cell fate responses-adaptation versus cell death- through mechanisms that depend on its cytosolic endoribonuclease (RNase) domain. Adaptive responses are mediated by X-box binding protein 1 (XBP1), where IRE1α promotes cleavage and splicing of unspliced Xbp1 mrna to generate an active transcription factor, XBP1S. In contrast, cell death is mediated through preferential cleavage and degradation of mrnas important in survival through a process called regulated IRE1α dependent decay (RIDD). However, it is not clear if IRE1α regulates similar divergent responses during cancer progression. To model and study the contribution of both IRE1α RNase outputs to oncogene activation, we transduced oncogenic v-ha-ras (v-ras Ha ) into primary keratinocytes. During the initial proliferative response, oncogenic v-ras Ha simultaneously activates IRE1α-mediated splicing of Xbp1 mrna as well as promotes mrna decay through RIDD. ShRNA knockdown and pharmacological approaches implicate ER stressdependent Xbp1 mrna splicing as part of the proliferation response to oncogenic v- RAS Ha, while RIDD mediates cell cycle growth arrest in the absence of XBP1. The requirement of Xbp1 mrna splicing for proliferation is linked to suppression of v-ras Ha - induced senescence. In contrast, IRE1α promotes v-ras Ha -induced senescence in the absence of XBP1 and is required for suppression of malignant transformation. Gene expression profiling of v-ras Ha keratinocytes deficient in IRE1α identified a number of RIDD candidate mrna substrates that are linked to senescence and malignant transformation. Among those identified was the ID family, of which Id1 mrna was

4 confirmed as a direct target of IRE1α RNase activity using cell-free assays. Furthermore, shrna knockdown of Id1 mrna rescued the senescence phenotype in v- iv RAS Ha keratinocytes with inactive IRE1α. Together, these results suggest that the balance between IRE1α-XBP1S and IRE1α-RIDD determines whether a premalignant tumor cell undergoes proliferation and tumor progression or senescence and tumor suppression. Next, we showed that regulation of IRE1α RNase outputs in v-ras Ha keratinocytes can be modulated by the potent TGFβ1 tumor suppressor pathway. TGFβ1 treatment significantly enhanced v-ras Ha - induced senescence which was associated with dampened ER stress and Xbp1 mrna splicing but caused greater downregulation of RIDD substrates. Treatment of v-ras Ha keratinocytes with the TGFβ1 type I receptor inhibitor SB , previously shown to inhibit v-ras Ha -induced senescence, enhanced Xbp1 mrna splicing and attenuated RIDD. Pharmacological inactivation or depletion of IRE1α prevented TGFβ1-induced senescence independent of XBP1 and this was not due inhibition of TGFβ1-induced transcriptional activation. Conversely, pharmacologically-induced ER stress inhibited TGFβ1-induced senescence and growth arrest in v-ras Ha keratinocytes. This inhibition resulted from inactivation of downstream TGFβ1 signaling through a mechanism that involved impairment of ligandinduced receptor internalization. Together, selective modulation of v-ras Ha -induced IRE1α RNase activity toward an mrna decay pathway by TGFβ1 may suppress tumorigenesis by promoting senescence. Furthermore, ER stress, which promotes the adaptive functions of the UPR, represents a new mechanism to inactivate TGFβ1 signaling and inhibit the senescence response.

5 v Table of Contents List of Figures... viii List of Tables... x Abbreviations... xi Acknowledgements... xiv Chapter 1 Introduction General overview of ER stress and the UPR ER structure and function ER stress and UPR signaling Role of IRE1α in regulating cell fate responses Role of IRE1α signaling in cancer RAS oncogene RAS signaling RAS and cancer RAS and senescence Skin Carcinogenesis Structure and physiology of skin Two-Stage skin chemical carcinogenesis TGFβ1 signaling Overview of TGFβ1 signaling pathway Dual role of TGFβ1 signaling in cancer Interactions between TGFβ1 and RAS signaling Hypothesis and Aims Bibliography Chapter 2 Methods Cell culture and reagents Plasmids Virus production and infection BrdU incorporation assay Senescence associated β-glactosidase assay In vitro malignant conversion assay RNA isolation and quantitative PCR (q-pcr) Western blot analysis Phos-Tag SDS-PAGE In vitro RNA cleavage assays Luciferase reporter assays Microarray analysis Cell surface biotinylation assays... 93

6 vi 2.14 TGFβ1 receptor internalization assays Statistical analysis Bibliography Chapter 3 IRE1α regulates divergent RNase outputs that dicate senescence and malignant conversion induced by oncogenic HRAS Abstract Introducton Results Oncogenic v-ras Ha activates IRE1α-mediated Xbp1 splicing and RIDD during proliferation ER stress dependent activation of IRE1α by MEK-ERK signaling Opposing roles of IRE1α and XBP1 during proliferation and senescence XBP1 depletion causes hyperactive IRE1α and MEK-ERK signaling to promote senescence in v-ras Ha keratinocytes Expression profiling and identification of IRE1α-RIDD substrates linked to v-ras Ha -induced senescence and suppression of malignant conversion Id1 mrna is a direct IRE1α-RIDD cleavage substrate Downregulation of Id1 mrna is required for IRE1α-induced senescence in v-ras Ha keratinocytes Discussion Bibliography Chapter 4 IRE1α and ER stress dictates TGFβ1 signaling and senescence Abstract Introducton Results TGFβ1-induced senescence is associated with dampened IRE1αmediated Xbp1 mrna splicing and enhanced RIDD Inactivation of TGFβ1 signaling in v-ras Ha keratinocytes enhances Xbp1 mrna splicing and inhibits RIDD IRE1α inactivation blocks TGFβ1-induced senescence without inhibiting Smad activation ER stress inhibits TGFβ1-induced senescence ER stress inhibits Smad activation through impairment of TGFβ1 receptor internalization Discussion Bibliography

7 vii Chapter 5 Discussion Dual and opposing role of IRE1α RNase outputs during v-ras Ha -induced senescence Future studies Stage specific effects of IRE1α RNase activites during cancer progression Characterization of IRE1α-specific cancer mutants during v-ras Ha - induced senescence and malignant conversion Allosteric modulation of IRE1α as a therapeutic option for the treatment of cancer Bibliography

8 viii List of Figures Figure 1.1: General overview of ER functions under homeostatic conditions... 4 Figure 1.2: The Unfolded protein response (UPR) signaling pathway... 6 Figure 1.3: ER stress and the UPR triggers apoptosis through the intrinsic mitochondrial pathway Figure 1.4: Mechanism of IRE1α activation and regulation of cell fate during ER stress Figure 1.5: RAS signaling Figure 1.6: Modeling of RAS-driven multistage epithelial carcinogenesis in vivo and in vitro Figure 1.7: TGFβ1 signaling pathway Figure 1.8: TGFβ1-mediated growth inhibition in normal epithelial cells Figure 1.9: General schematic of antagonistic and synergistic interactions between RAS and TGFβ1 signaling pathways Figure 3.1: Oncogenic v-ras Ha promotes ER stress during proliferation Figure 3.2: Oncogenic v-ras Ha activates IRE1α but not PERK or ATF Figure 3.3: Activation of IRE1α-mediated RIDD in v-ras Ha keratinocytes Figure 3.4: Oncogenic v-ras Ha -induced ER stress and MEK-ERK signaling regulates IRE1α activation Figure 3.5: IRE1α RNase activity promotes growth arrest in the absence of XBP1 in v-ras Ha keratinocytes Figure 3.6: Divergent IRE1α RNase outputs is associated with v-ras Ha -induced senescence Figure 3.7: Opposing roles of IRE1α and XBP1 during v-ras Ha -induced senescence Figure 3.8: XBP1 depletion causes hyperactive IRE1α and MEK-ERK signaling to promote senescence

9 ix Figure 3.9: Identification of IRE1α-RIDD candidate genes linked to senescence and malignant conversion Figure 3.10: Validation of IRE1α-RIDD candidate genes Figure 3.11: Id1 mrna is a direct cleavage substrate of IRE1α Figure 3.12: IRE1α-mediated cleavage of Id1 mrna controls v-ras Ha -induced senescence response Figure 3.13: Model for opposing IRE1α RNase outputs during v-ras Ha -induced proliferation and senescence Figure 4.1: TGFβ1 promotes senescence in v-ras Ha keratinocytes Figure 4.2: TGFβ1 inhibits IRE1α-mediated Xbp1 mrna splicing but enhances downregulation of RIDD substrates during senescence Figure 4.3: SB enhances Xbp1 mrna splicing and inhibits downregulation of RIDD mrna substrates in v-ras Ha keratinocytes Figure 4.4: IRE1α is required for TGFβ1-induced senescence but does not affect Smad activation Figure 4.5: Pharmacological ER stress inhibits TGFβ1-induced senescence Figure 4.6: Pharmacological ER stress inhibits TGFβ1 signaling Figure 4.7: Pharmacological ER stress blocks TGFβ1-induced receptor internalization Figure 4.8: Model for regulation of TGFβ1-induced senescence dependent on ER stress and modulation of IRE1α RNase activity

10 x List of Tables Table 3.1: Candidate IRE1α-RIDD targets linked to senescence and supppression of malignant conversion in v-ras Ha keratinocytes

11 xi Abbreviations S1P site 1 protease S2P site 2 protease BiP binding immunoglobulin protein PDI protein disulphide isomerase GRP94 glucose-regulated protein 94 eif2α elongation intiation factor 2 alpha ATF4 activating transcription factor 4 XBP1 xbox binding protein 1 XBP1U unspliced xbox binding protein 1 XBP1S spliced xbox binding protein 1 BCL-2 B cell lymphoma 2 BAX BCL-2-associated x proein BAK BCL-2 homologous antagonist/killer PUMA p53 upregulated modulator of apoptosis NOXA phorbol-12-myristate-13-acetate induced protein 1 BIM BCL-2 interacting mediator of cell death ASK1 apoptosis signal regulating kinase 1 JNK JUN N-terminal kinase TRAF2 TNF receptor-associated factor 2 CHOP CCAAT/Enhancer binding protein GADD34 growth arrest and DNA damage inducible protein 34 HER2 human epidermal growth factor receptor 2 HIF1α hypoxia inducible factor 1 alpha VEGF vascular endothelial growth factor IL-6 interleukin 6 PER1 period circadian clock 1 APC min adenomatous polyposis coli multiple intestinal neoplasia RAS rat sarcoma HRAS harvey rat sarcoma viral oncogene homolog NRAS neuroblastoma viral oncogene homolog KRAS kirsten rat sarcoma viral oncogene homolog v-ras Ha Harvey rat sarcoma viral oncogene homolog SPARC secreted protein, acidic, cysteine rich GTPase guanosine triphosphate hydrolase enzyme GTP guanosine triphosphate GDP guanosine diphosphate GRB2 growth factor receptor bound protein 2 SOS son of sevenless EGF-epidermal growth factor RASGAP RAS GTPase activating protein NF1 neurofibromatosis type 1 RAF v-raf murine sarcoma 3611 viral oncogene homolog PI3K phosphatidylinositol-4,5-bisphosphate 3-kinase

12 RalGDS ral guanine nucleotide dissociation stimulator MEK mitogen activated protein kinase kinase ERK extracellular signal-regulated kinase JUN v-jun sarcoma virus 17 oncogene homolog ELK1 ETS-like gene 1 FOS FBJ murine osteosarcoma viral oncogene homolog AKT v-akt murine thymoma vival oncogene PDK1 pyruvate dehydrogenase kinase 1 FOXO forkhead box O NFkB nuclear factor of kappa light polypeptide gene enhancer in B cells RGL ral guanine nucleotide dissociation factor Rlf/RGL2 ral guanine nucleotide dissociation factor 2 RhoA RAS homolog gene family, member A Rac1 RAS related C3 botulinum toxin substrate 1 Cdc42 cell division cycle 42 RLIP1 rala binding protein 1 TGF-α-transforming growth factor alpha AREG - amphiregulin HBEGF heparin-binding EGF-like growth factor TGFβ1-transforming growth factor beta 1 GSK3β glycogen synthase kinase 3 beta IAP inhibitors of apoptosis PAR4 prostate apoptosis response 4 BCL-XL apoptosis regulator BCL-X BAD BCL-2-associated agonist of cell death ARC activity regulated cytoskeleton-associated protein ARNT aryl hydrocarbon receptor nuclear translocator mtor mechanistic targets of rapamycin GLUT1 glucose transporter 1 ATG5 autophagy related 5 ATG7 autophagy related 7 Mcl-1 myeloid cell leukemia 1 bfgf basic fibroblast growth factor PDGF platelet derived growth factor IL-8 interleukin 8 COX2 cyclooxygenase 2 MMP-2 matrix metalloproteinase 2 MMP-9 matrix metalloproteinase 9 upa urokinase plasminogen activator TSP-1 thrombospondin-1 SNAIL zinc finger protein SNAI1 SLUG SNAIL homolog 2 ROS-reactive oxygen species prb-retinoblatoma protein MDM2 xii

13 PIRH2 P53 induced protein with a RING H2 domain COP1 constitutive photomorphogenesis protein 1 ARF alternate reading frame protein CDK2-cyclin dependent kinase 2 CDK4/6-cyclin dependent kinase 4/6 Bmi-1 Bmi1 polycome ring finger protein CBX7 chromobox homolog 7 ID1-inhibitor of differentiation 1 Ets-1 v-ets avian erythroblastosis virus E26 oncogene homolog 1 MKK3 mitogen activated protein kinase kinase 3 MKK6 mitogen activated protein kinase kinase 6 E2F E2F transcription factor PLC phospholipase C PKC protein kinase C AP1 activated protein 1 RAR retinoic acid receptor RXR retinoid x receptor BMP-bone morphogenic protein GDF growth differentiation factor SARA smad anchor for receptor activation TAK1-tgfbeta activated kinase 1 Ski v-ski avian sarcoma viral homolog 1 Sno strawberry notch homolog 1 TGIF TGFbeta induced factor homeobox 1 PPARβ/δ peroxisome proliferator activated receptor beta/gamma PMP22 peripheral myelin protein 22 HGSNAT heparin-alpha-glucosminide N-acetyltransferase TIMP3 tissue inhibitor of metalloproteinase 3 IGFBP2 insulin growth factor binding protein 2 PP2Ce protein phosphatase 2C episilon PTP-1B protein tyrosine phosphtase 1B ATF3 activating transcription factor 3 IL-1 interleukin 1 xiii

14 xiv Acknowledgements I would first like to thank my advisor, Adam Glick, for giving me the opportunity to grow as an independent scientist, provide the necessary resources and reagents for my graduate studies, and for putting up with my emotional highs and lows as I tried to blaze a meaningful path in the cancer research field. Thank you for tolerating my ways for so many years, I would like to thank my committee for their patience, encouragemet and criticisms as I turned negative situation into a positive and productive one. I would especially like to thank Dr. Omiecinski for his willingness to be a part of my thesis defense on short notice. I would like to thank my current and former lab members Michael Podolsky, Kyle Breech, and Jeongin Son for meaningful input on my studies and entertaining discussions on politics, pop culture, and video games. I would especially like to thank my most recent undergraduate students Alayna-Craig Lucas, Christian John, Xiao Cui and Deirdre DeFranco that I have had the privilege to know and teach. It was always a joy for me despite my occasional grumpiness. I also would like to thank Michael Borland, Tajas Lohti, Ian Murray, and Kayla Smith for the many scientific discussions over the years and also for kindly providing valuable reagents for my graduate studies when in dire need. Finally, I want to thank my wonderful family. Without them I would not be the person I am today. To my mom thank you so very much for guiding me and encouraging me not to give up. Thank you for being there at my weakest and providing the emotional support I desperately needed many times. To my brother Vlad I will

15 xv always admire your drive and unflinching work ethic. I hope to emulate that one day. To my brother Tim, you are incredibly smart more so than me. Don t let it get to your head. To my sister Kim, thank you for getting a new dog without permission and making him my responsibility. To my dog Scarlett I miss you and to my dog Louie thank you for keeping things interesting while I was writing. I love you all. Finally to my dad, I wish you were here in person to see me finish what I started but I know you are looking and smiling down on me. I miss you terribly and love you. This is for you.

16 1 Chapter 1: Introduction This review provides an overview of endoplasmic reticulum (ER) stress and the signaling pathways involved in the unfolded protein response (UPR) as well as mechanisms employed in promoting adaptation or cell death. Emphasis is given to IRE1α, which is the most evolutionarily conserved among the three UPR transducers and the subject of the studies presented herein, where emerging evidence shows that it is an important mediator of controlling cell fate in chemically-induced ER stress models (1-3). A brief overview of the role of ER stress and the UPR in cancer and an in depth analysis of the critical role of IRE1α during cancer progression is presented. This will be followed by a general discussion of the role of oncogenic RAS in transformation and senescence and its relevance in the two-stage skin carcinogenesis model. Additionally, the role of TGFβ1 signaling in cancer is examined as well as its interactions with oncogenic RAS in this context. Here, I will show that both pathways regulate IRE1α to determine cell fate during early stages of cancer progression. 1.1 General overview of ER stress and the UPR All eukaryotic cells have developed elaborate quality control mechanisms to ensure proper maturation and folding of proteins within the ER before being exported to the cell surface or secreted extracellularly (4). Signaling receptors and growth factors that are essential for communication with the environment are synthesized in the ER. Thus, proper quality control of these proteins is needed to regulate cellular processes such as proliferation, differentiation, migration, and survival (4). Environmental insults that disrupt the fidelity of the protein maturation process can lead to an accumulation of

17 2 unfolded proteins within the ER and trigger a condition known as ER stress. To abate this imbalance, a set of overlapping evolutionarily conserved signaling pathways, termed the UPR, monitors conditions within the ER and transmit signals to the nucleus to help restore ER homeostasis(4). Paradoxically, studies in cell culture systems have shown that chronic ER stress promotes cell death through yet still partially undefined UPR-dependent mechanisms (5, 6). Thus, the underlying paradigm is that the UPR functions as a rheostat that senses the level of ER stress and controls cell fate decisions. Chronic ER stress-induced apoptosis has a now well documented role in several debilitating diseases, including neurodegenerative diseases such as Alzheimer s, diabetes, atherosclerosis, and renal disease (7). In contrast in diseases such as cancer ER stress is high and the UPR is activated but subverts the apoptosisinducing role and promotes the cytoprotective features of this response (8, 9). Due to the overlapping nature and dualism of the UPR and the disease context therapeutic intervention in the UPR is difficult. Thus, it is important to understand the molecular mechanisms of the UPR at the cellular, tissue and organismal level so that small molecule inhibitors or activators can be developed that selectively modulate discreet steps of the pathway ER structure and function The ER is a large, membranous network of cisternae and the first compartment in the secretory pathway where proteins destined for the cell surface or secreted into the extracellular milieu undergo synthesis, folding and posttranslational modifications that ensure proper functioning (4). While the general features of protein folding are similar in both the cytosol and ER compartments, the ER is much more complex as it is

18 3 a highly oxidized and Ca 2+ rich environment, where the addition of posttranslational modifications such as N-linked glycosylation and disulphide bond formation require specialized machinery to ensure proper folding. As such, the ER contains a number of unique resident protein chaperones and folding enzymes that associate with newly synthesized proteins to prevent their aggregation and ensure proper assembly before exiting the ER through transport vesicles. Furthermore, quality control mechanisms exist where only properly folded proteins are packaged into transport vesicles and secreted into the extracellular space or displayed on the cell surface (10, 11). Misfolded proteins are retained in the ER and retrotranslocated to the cytosol where they are degraded by the 26S proteasome (12). This process, called ER-associated degradation (ERAD) is required for continued removal of misfolded proteins (Figure 1.1) ER stress and UPR signaling When the influx of newly synthesized unfolded proteins entering the ER exceeds the folding capacity, unfolded proteins accumulate and ER homeostasis is perturbed leading to ER stress. Perturbations that cause ER stress can be induced by any number of physiological changes and in some cases occurs during normal processes. For example, B cells undergoing immunoglobulin secretion or differentiation into plasma cells, viral infections, and host response to microbial infection in plants have higher secretory activity which in turn causes ER stress (13-15). To combat ER stress, the UPR is activated to attenuate ER stress and restore ER homeostasis. Therefore, the UPR is an adaptive response to ER perturbations. The UPR mitigates ER stress by decreasing the folding demand and increasing the folding capacity of the ER. The folding demand is decreased by reducing the transcription and translation of proteins

19 4 Figure 1.1. General overview of ER functions under homeostatic conditions. (A) Proteins destined for the secretory pathway are first translated on ER-bound polysomes and simultaneously translocated into the ER through a protein channel called a translocon. (B) Resident chaperones and folding enzymes such as BiP, protein disulphide isomerase (PDI), and ERO1 facilitate proper folding of newly synthesized proteins and prevent aggregation, ensure proper processing and addition of branced glycans, and aid in disulphide bond formation for protein stabilization. (C) Correctly folded and matured proteins exit the ER through formation of transport vesicles and continue through the secretory pathway. (D) Existing protein quality control mechanisms which deem proteins misfolded are retrotranslocated to the cytosol and degraded by the 26S proteasome.

20 5 entering the ER or by increasing the clearance of unfolded proteins by ERAD (4). To increase ER capacity, the UPR increases the synthesis of chaperones and folding enzymes as well as promotes an increase in ER size, which attenuates the incoming protein load (4). To date, three major branches of the UPR have been characterized (Figure 1.2). Each branch consists of a stress sensing ER-resident transmembrane protein that signals downstream to the nucleus through unique mechanisms that differs from traditional signaling from the cell surface to the nucleus. The three branches of the UPR include activating transcription factor 6 (ATF6), double-stranded RNA-activating protein kinase (PKR)-like ER kinase (PERK), and inositol requiring enzyme 1 (IRE1). Of these three, IRE1 is the most evolutionarily conserved among eukaryotes (16). ATF6 is initially synthesized as an ER-resident transmembrane protein which contains a large ER-luminal domain. Upon ER stress ATF6 is packaged into transport vesicles and delivered to the Golgi apparatus where cleavage of the luminal domain by two proteases, S1P and S2P, which releases the N-terminal cytoplasmic portion ATF6 (N). ATF6(N) is a transcription factor that localizes to the nucleus to regulate UPR target genes (4). Mechanisms by which ATF6 senses accumulation of unfolded proteins in the ER are not well characterized. One likely mechanism of activation is that the ER chaperone BiP associates with the ATF6 luminal domain during homeostatic conditions, and upon ER stress, is released leading to ATF6 transport to the Golgi and subsequent cleavage (17). This mechanism of BiP association to regulate UPR activation is shared with both PERK and IRE1 as well (4). Once activated, ATF6 regulates expression of

21 6 Figure 1.2. The Unfolded protein response (UPR) signaling pathway. The UPR consists of three stress sensors, ATF6, PERK, and IRE1, which become activated due to accumulation of unfolded proteins within the ER and transduce signals about ER status to the cytosol and nucleus to restore ER homeostasis. ATF6 is proteolytically processed to produce a transcription factor that encodes genes involved in protein folding and ER biogenesis. PERK activation phosphorylates eif2α to dampen protein synthesis but also leads to specific translation of ATF4, which encodes genes involved in protein folding as well as amino acid synthesis and antioxidants. IRE1 activation leads to dimerization and autophosphorylation triggering its cytosolic RNase activity, which splices unspliced XBP1 (XBP1U) into an active transcription factor, XBP1S. XBP1S translocates to the nucleus where it encodes genes involved in protein folding, ER biogenesis, and protein degradation. IRE1 also degrades select mrnas through a process called regulated IRE1 dependent decay or RIDD that serves to reduce protein load.

22 7 genes involved in protein folding including BiP, PDI, and GRP94 as well as those involved in ERAD (18, 19). PERK is an ER-resident transmembrane kinase. In unstressed cells PERK exists as an inactive monomer bound to BiP but upon ER stress PERK undergoes oligomerization followed by autophosphorylation to activate its kinase domain (4). This process is required to phosphorylate and inactivate eif2α which reduces mrna translation, thereby limiting the protein load entering the ER and reducing ER stress (4). Interestingly, despite a general reduction in mrna translation, PERK activation leads to selective translation of the transcription factor ATF4. This occurs due to two short upstream open reading frames in the Atf4 mrna that are preferentially translated when eif2α levels are sufficiently low (20). Activation of ATF4 regulates a subset of genes involved in amino acid metabolism, oxidative stress and protein folding (4, 21). IRE1 is the third UPR signaling transducer and consists of two isoforms, IRE1α and IRE1β (4). IRE1 is unique in that it is a bifunctional ER transmembrane kinase and endoribonuclease (RNase) that catalyzes the non-conventional cleavage and splicing of XBP1 mrna to transmit downstream UPR signaling (4). Similar to PERK, IRE1 exists as an inactive monomer in non-stressed cells bound to BiP but forms dimers or highorder oligomers during ER stress depending on the intensity or duration (22). IRE1 oligomerization promotes transautophosphorylation leading to conformational changes that activates the RNase domain. Xbp1 mrna cleavage by IRE1 removes a 26 base intron at two sequence specific sites that form stem loop structures (4, 23). The two exons are joined by a trna ligase and Xbp1 mrna is translated into a stable and

23 8 active transcription factor called XBP1S. XBP1S translocates to the nucleus to regulate UPR target genes such as those involved in ERAD and protein folding (24) as well as components involved in ER biogenesis (25). IRE1 also promotes degradation of mrnas through a process called regulated IRE1 dependent decay (RIDD), and reduces protein load into the ER lumen during ER stress (22). RIDD cleavage also is site specific as validated target substrates contain consensus sequence 5 -CUGCAG-3 and stem loop structures similar to Xbp1 mrna(26). Initial discoveries of RIDD substrates suggested that IRE1 would only target mrnas associated with ER- bound polysomes that were part of the secretory pathway since that is the primary site of IRE1 localization (27). However, IRE1 can induce the decay of pre-mirnas as well as cytosolic mrnas that encode cytosolic or nuclear proteins (2, 28). One possible mechanism for localization of cytosolic mrnas to the ER come from recent discoveries involving unspliced Xbp1 mrna, which itself is localized in the cytosol. XBP1U, a protein translated from unspliced Xbp1 mrna, can act as a chaperone for unspliced Xbp1 mrna and facilitate association to ER-bound polysomes suggesting perhaps a similar mechanism could occur with presenting other cytosolic mrnas to IRE1(29). 1.2 Role of IRE1α in regulating cell fate responses Activation of the UPR in response to ER stress first serves as an adaptive mechanism by reducing unfolded protein load within the ER through several pro-survival mechanisms. These include ER expansion/biogenesis, selective induction of key protein folding enzymes and quality control machinery, and dampening of incoming proteins into the ER. However, when ER stress is persistent or at high levels and ER

24 9 homeostasis cannot be restored, the UPR triggers apoptosis (22). Apoptosis triggered by chronic ER stress is dependent on the intrinsic mitochondrial apoptosis pathway regulated by both pro- and anti-apoptotic BCL-2 family members. BCL-2 family members are characterized by presence of at least four BCL-2 (BH) homology domains. Intrinsic apoptosis is mediated by the activation of pro-apoptotic BCL-2 members, BAX and BAK, at the mitochondria leading to cytochrome c release, and activation of the caspase cascade(30). BAX and BAK are regulated upstream by BH3 domain-only proteins such as PUMA and NOXA (30). ER stress-induced apoptosis promotes transcriptional induction of both PUMA and NOXA (31). In addition, the pro-apoptotic BH3-only protein, BIM, is also upregulated at both the transcriptional and posttranslational level during chronic ER stress (32). Current evidence implicates both IRE1α and PERK as having bifunctional roles in triggering adaptation to ER stress as well as apoptosis. IRE1α can activate ASK1 and JNK signaling pathways through an association with TRAF2, which are mediators of the apoptotic response in cells undergoing ER stress (33). In addition, recent studies suggest that the RIDD function of IRE1α has a significant role in triggering the apoptotic response through degradation of mirnas that repress Caspase 2 (2, 3). Furthermore, RIDD may trigger apoptosis by promoting decay of mrnas of key growth-promoting proteins (22, 34-36). ATF4 induction upon PERK activation controls the induction of the pro-apoptotic protein CHOP. Although not well understood, CHOP can induced apoptosis through downregulation of anti-apoptotic BCL-2, or the transcriptional upregulation of pro-apoptotic proteins BIM, PUMA, and GADD34 (37-40) (Figure 1.3).

25 Figure 1.3. ER stress and the UPR triggers apoptosis through the intrinsic mitochondrial pathway. Under irremediable levels of ER stress, IRE1α and PERK send signals to BCL-2 proteins to activate apoptosis through the mitochondria. IRE1α induces endonucleolytic decay of ERlocalized mrnas, reducing proteins important in growth control or ER homeostasis which may lead to more severe ER stress. IRE1α may also degrade micrornas that derepress caspase 2, leading to activation of proapoptotic BID. IRE1α also recruits TRAF2 and activates ASK1 and JNK. JNK then activates pro-apoptotic BIM and inhibits anti-apoptotic BCL-2. High ER stress also leads to PERK activation, which causes selective translation of ATF4. ATF4 upregulates CHOP, which inhibits expression of anti-apoptotic BCL-2 while inducing expression of pro-apoptotic BIM. ER stress also promotes p53-dependent transcriptional upregulation of NOXA and PUMA through unknown mechanisms. Together, these proteins activate BAX and BAK on the mitochondria, leading to oligomerization that then drives permeabilization of the outer mitochondrial membrane, cytochrome c release, and activation of caspase-3-mediated apoptosis. 10

26 A critical question is what mechanisms trigger apoptotic cell death in response to ER stress? Emerging evidence suggests that the intensity and kinetics of different UPR 11 signal transducers are tightly regulated, which impact cell fate responses. Frequently, chemicals used to induce ER stress, such as thapsigargin or DTT, are used at high doses which lead to chronic ER stress and apoptosis. In contrast, certain specialized cell types that display high secretory capacity, such as plasma cells or pancreatic beta cells, undergo ER stress that is beneficial (41). Thus, ER stress is critically important to inducing the adaptive and apoptotic arms of the UPR. However, it is often difficult to delineate the adaptive versus apoptotic functions of individual UPR transducers. For example, chemically-induced ER stress often exhibits induction of adaptive proteins such as chaperones and ERAD components simultaneously with induction of apoptoticrelated factors (22). Previous studies have shown that PERK and IRE1α have similar stress-sensing luminal domains but their signaling dynamics are drastically different (22, 42). In some cases, IRE1α signaling is turned off upon chronic ER stress whereas PERK signaling is persistent (43, 44). This suggests that inactivation of IRE1α reduces the pro-survival arm through decreased XBP1S, and that activation of PERK during chronic ER stress may trigger its pro-apoptotic functions through induction of CHOP (22). Alternatively, PERK signaling is a transient response and IRE1α signaling is sustained (2, 3, 22, 45) suggesting an important role for IRE1α in the ER stressmediated apoptotic response. Indeed, examination of IRE1α-mediated XBP1S and RIDD during ER stress revealed that when XBP1S is at its maximum, RIDD target substrates are just beginning to decrease and continue to do so until cell death occurs (2, 3, 45, 46). In agreement, it appears RIDD activity increases proportionally as ER

27 12 stress levels increase while XBP1S is dampened at these later timepoints as ER stress damage becomes irreversible (46). Together this suggests that RIDD has a causal role in initiating ER stress-induced cell death and that the balance of IRE1α RNase outputs, XBP1S and RIDD, dictates this cell fate response. Perhaps the most relevant study to date to demonstrate that IRE1α regulates divergent cell fates comes from Han et al., which show through chemical-genetic studies that IRE1α regulates Xbp1 mrna splicing and RIDD through two distinct activation modes: called pseudokinase and phosphotransfer activation (3). By using an IRE1α mutant (I642G) that contains an engineered nucleotide binding pocket, they were able to use an ATP-competitive inhibitor (NM-PP1) to allosterically regulate IRE1α activation in the absence of ER stress phosphotransfer activation (3). In this system, NM-PP1 treatment caused Xbp1 mrna splicing but not RIDD and promoted only the cytoprotective functions (3). This suggests that NM-PP1 treatment, by acting as a pseduokinase, can promote IRE1α dimerization in the absence of phosphotransfer activation leading to Xbp1 mrna splicing. In contrast, phosphotransfer activation and high-order oligomer formation, which relaxes specificity of the IRE1α RNase pocket, promotes RIDD and apoptosis. Another study showed that quercetin, a naturally occurring flavonol, can activate the IRE1α RNase through potentiating activation by ADP, which engages the nucleotide binding cleft of IRE1α (47). Thus, this suggests that endogenous ligands, in corroboration with ER stress signals may function to modulate IRE1α activity to determine cellular functions. It is also now well-appreciated that IRE1α undergoes dynamic clustering into high-order oligomers in vivo (48, 49) and other studies suggest that formation of these high-order oligomers could be associated with RIDD activity in

28 13 vitro (3, 50). It is currently not clear whether clustering is associated with Xbp1 mrna splicing, RIDD, or both in vivo as oligomerization has only been observed in overexpression studies and needs to be further examined in an unperturbed setting (49). Taken together, the current proposed model of IRE1α activation to determine divergent cell fates involves a series of sequential steps: 1) Mild levels of ER stress promote formation of IRE1α dimers, which preferentially juxtaposes IRE1α monomers to form a RNase binding pocket preferential for Xbp1 mrna splicing although basal levels of RIDD may occur. Ultimately this leads to the adaptive features of IRE1α. 2) Chronic or high levels of ER stress further increase IRE1α transautophosphorylation and promote formation of high-order oligomers. Oligomer formation relaxes the RNase pocket thereby reducing preference for Xbp1 mrna splicing but enhancing degradation of mrnas involved in survival and apoptosis induction (Figure 1.4). 1.3 Role of IRE1α in cancer There is a strong association between UPR activation and cancer, due to conditions such as hypoxia, nutrient deprivation, metabolic stress, altered ph, or increased protein synthesis within the growing tumor (8, 51-53). It is clear that many features of the adaptive UPR have an essential role in tumor growth. However, since the UPR can also trigger apoptosis, it may serve to prevent tumor development. Currently, it remains to be elucidated where the balance of the UPR ultimately lies during cancer progression. Many aspects of the UPR have been studied extensively to understand its role in cell proliferation, angiogenesis, inflammation, and tumor dormancybut only IRE1α will be the focus of this section. There are several other

29 Figure 1.4. Mechanism of IRE1α activation and regulation of cell fate during ER stress. (Left) Under basal conditions IRE1α exists as an inactive monomer bound to the ER chaperone BiP. (Middle) Mild ER stress levels activate the adaptive arm of IRE1α with the formation of dimers. Dimer formation produces an RNase pocket that preferentially splices XBP1 mrna leading to nuclear translocation and transcription of genes that ameliorate ER stress. (Right) If the levels of ER stress become chronic or too high, IRE1α phosphorylation levels increase as well as formation of high order oligomers occur. Oligomerization relaxes the IRE1α RNase pocket facilitating degradation of ER-localized and cytosolic mrnas. MRNAs critical for survival, ER chaperones that aid in protein folding, and mirnas that suppress caspasemediated apoptosis are degraded leading to apoptosis. 14

30 15 excellent reviews detailing the overall role of the UPR during tumor development (8, 51, 54). The role of the IRE1α pathway and its downstream RNase functions in cancer development remains controversial. While there is strong support for a pro-oncogenic function of the IRE1α substrate XBP1 in most cancer types, the role of RIDD is currently unknown due to limited studies and has no clear role in malignant progression. XBP1 is overexpressed in several human cancers including breast (55) liver (56), and colon (57) and high levels of XBP1S are associated with poor clinical prognosis in breast cancer patients (58). Mechanistically, a recent study by Chen et al. demonstrated a critical role for XBP1 in triple negative breast cancer (TNBC)- a highly aggressive form of breast cancer with limited treatment options that lacks expression of estrogen, progesterone, and HER2 receptors (59). In breast cancer cell line models, XBP1 acted as a coregulator of HIF1α target genes to regulate angiogenesis and invasion. Furthermore, XBP1 deficiency in these cell lines reduced tumor growth and tumor relapse in mice. Another study showed that constitutive overexpression of XBP1S is sufficient to promote multiple myeloma in vivo (60) while XBP1 deletion in a fibrosarcoma model reduced tumor growth and survival independent of VEGF expression (61). In addition, dominant-negative IRE1α inhibited XBP1S induction of cyclin A, thereby reducing prostate cancer cell line proliferation (62) Similar to studies examining the role of XBP1, lung adenocarcinoma cells expressing a dominant negative form of IRE1α displayed reduced levels of VEGFA under conditions of hypoxia, and this correlated with reduced angiogenesis and tumor growth in vivo (63). Similarly, malignant glioblastoma cells expressing a dominant negative IRE1α displayed reduced angiogenesis and tumor

31 16 growth that was dependent on IL-6 expression (64). However, in these studies with a dominant negative IRE1α the direct contribution of RIDD to malignant progression was not established. One recent study suggests a pro-oncogenic role for RIDD. IRE1α mediated decay of the mrna to the core circadian clock gene, PER1, enhanced tumor growth and angiogenesis of malignant glioblastoma cells and reduced shorter survival times in mice (65). Paradoxically, there are a few reports implicating IRE1α RNase activities as having an anti-oncogenic role during tumor development. For example, Neiderreiter et al. show that reduced expression of XBP1 confers a genetic risk for inflammatory bowel disease characterized by a regenerative response and expansion of intestinal stem cells (66). Long-term effects from this are colitis-associated and spontaneous development of intestinal tumors on an APC min background. Another study demonstrates that oncogenic HRAS activates IRE1α-mediated Xbp1 mrna splicing to promote premature senescence in primary melanocytes suggesting a tumor suppressor role during early stages of cancer development (67). Lastly, while dominant negative IRE1α malignant glioblastoma cells displayed reduced tumor growth and angiogenesis, this was associated with a higher migratory and invasive capacity (34). By extension, Dejeans et al. showed that increased migration and invasion in IRE1α deficient cells was due to impaired mrna cleavage of the extracellular matrix gene, SPARC, indicating RIDD may have a anti-onogenic role in certain cellular and biological contexts (34). The role of IRE1α and its RNase outputs is further obscured in that a comprehensive proteomic study of human cancers identified IRE1α as the fifth most protein kinase containing somatic mutations that is also associated with at least one

32 17 driver mutation (68). These IRE1α mutations are found in glioblastoma, lung adenocarcinoma, and ovarian cancers although at low frequencies (50, 69). A total of five IRE1α mutants have been characterized. Four are missense mutations while one is a nonsense mutation that deletes the RNase domain of IRE1α completely (50, 69). While none of these mutants have been tested in a cancer model, preliminary studies have demonstrated that all are defective in RIDD and cannot induce apoptosis during ER stress. Moreover, three of these mutants retain the ability cause Xbp1 mrna splicing (50), suggesting that disabling RIDD functions is paramount to loss of Xbp1 mrna splicing for tumor progression. Taken together, the role of IRE1α during cancer progression is complex. While studies suggest oncogenic and tumor suppressor roles for both XBP1 splicing and RIDD the consensus is unclear due to use of different model systems and methodologies that either completely inactivate or artificially activate IRE1α, or overlook potential complexity caused by divergent effects of IRE1α RNase targets on cancer development. Future studies should be geared towards complete and dual assessment of both RNase functions within a singular model system to accurately determine the role of IRE1α in cancer. This approach should in essence guide rational therapeutic and prevention strategies to develop pharmacological kinase inhibitors and activators to modulate IRE1α RNase activity. 1.4 RAS oncogene RAS signaling

33 RAS and its downstream effector pathways are central regulators to many biological processes and are involved in numerous diseases including cancer. RAS 18 Figure 1.5. RAS signaling. RAS proteins function as switches that transmit extracellular signals to intracellular signalling. The binding of GTP to RAS causes activation, which promotes interactions with downstream effector pathways such as MEK and PI3K pathways. Termination of RAS signaling occurs when its intrinsic GTPase activity cleaves the γ-phosphate of GTP, leading to RAS inactivation. This cycle of GTP/GDP is modulated by RASGAPs and RASGEFs. RASGAPs, such as NF1, enhances the intrinsic GTPase activity of RAS, negatively regulating RAS function. In contrast, RASGEFs such as SOS, promote release of GDP to facilitate GTP binding and RAS activation. Activated RAS engages effector molecules that initiate several pathways that can promote survival, cell growth and proliferation, and migration

34 19 proteins were first discovered as transduced oncogenes from the Harvey and Kirstin strains of rat sarcoma viruses (70, 71). Cloning of the transforming viral sequences using transfection based methodologies revealed mutationally activated forms of RAS, HRAS and KRAS (72, 73). A third member, NRAS, was later discovered from neuroblastoma cell lines (74). RAS proteins are small GTPases associated with the plasma membrane and transmit downstream signaling through a wide range of effector pathways (Figure 1.5). These pathways are involved in a number of important intrinsic and extrinsic biological functions including cell proliferation, differentiation, survival, apoptosis, angiogenesis and immune regulation (75). RAS proteins signal by binding to GTP, which causes a conformational shift that displays a binding surface with high affinity for downstream effector proteins (76). In contrast, GTP hydrolysis restores RAS proteins to their original conformational state, reduces binding affinity for effector proteins and leads to attenuation of downstream signaling. In normal somatic cells, activation of RAS proteins is facilitated by guanine nucleotide-exchange factors (GEFs) of which SOS is the prototypical member of this family. SOS accelerates the removal of GDP and permits higher binding affinity of GTP. Preference for GTP binding occurs because of 10-fold higher intracellular concentrations of this molecule and preferred binding affinity over GDP (77). SOS in association with GRB2 serves as a bridge from EGF receptor activation at the cell surface to RAS activation (78, 79), linking extracellular signals to activation of intracellular RAS proteins. Attenuation of RAS signaling is modulated by GTPase activating proteins (GAPs) which stabilizes GTPase activity of RAS. Members of this family include RASGAP (80-82) and NF1, which was

35 20 found to be disrupted in neurofibromatosis patients (83-85). This provided the first indicator of deregulated RAS signaling in a developmental disorder. The best characterized effector pathways downstream of activated RAS signaling include RAF, PI3K, and RalGDS (86). RAF is a serine/threonine kinase and was the first effector pathway implicated in transducing activated RAS signaling. RAF directly interacts with RAS only in the activated GTP-bound form (87-89) and this association is required for stimulation of the mitogen-activated protein kinase (MAPK) pathway (90). RAF directly activates the MAPK pathway through phosphorylation of MEK which in turn phosphorylates ERK. ERK phosphorylates both cytosolic and nuclear protein substrates including transcription factors such as JUN and ELK1. ELK1 is an E26 transformation-specific sequence (ETS) family member that can regulate FOS expression (91-93). In turn, JUN and FOS proteins form the AP1 transcription factor complex which can regulate proteins that control cell cycle entry such as cyclin D1 (94). Activated RAS can also bind directly to the effector PI3K. This binding occurs on the PI3K catalytic domain and causes allosteric conformational changes leading to activation (95). PI3K activation causes translocation to the plasma membrane where it phosphorylates phosphatidylinositol-4, 5-bisphosphate (PIP2) to generate phosphatidylinositol-4, 5-triphosphate (PIP3). PIP3 then recruits proteins containing a pleckstrin homology domain (PHD) including the serine/threonine kinase AKT as well as its activating kinase PDK1 (96). AKT, which is important in cell survival, can regulate several transcription factors such as FOXO and NFκB (86). FOXO proteins negatively regulate cell survival and AKT inhibits FOXO function by phosphorylating three conserved amino acid residues which conceals the nuclear localization sequence and

36 21 impairs DNA binding (97). This prompts nuclear export and subsequent degradation of FOXO proteins by the ubiqutin-proteasomal pathway (98). RAS also signals through a class of GEFs that include RalGDS, RGL, and Rlf/RGL2 although their function is the least well understood of the three effector pathways described here. These proteins link RAS activation to Ral small GTPases (99-101). Ral activation by RalGDS can modulate RhoA/Rac1/Cdc42 signaling through RLIP1 activation (102). RhoA and other related family members are small GTPases that can interact with a wide range of cell surface receptors including G-protein coupled receptors, receptor tyrosine kinases, and integrins which dramatically impacts downstream signaling pathways (103). The importance of RhoA GTPase family members is highlighted by the fact that oncogenic RAS requires these proteins for cell proliferation and transformation (103, 104) RAS and cancer Activating RAS mutations occur in approximately 30% of all human cancers although the distribution and frequency of these mutations varies for each RAS isoform (105). HRAS mutations are frequently associated with squamous skin tumors and head and neck squamous cell carcinomas (HNSCC), KRAS mutations are associated with colorectal, lung, and pancreatic cancers, and NRAS mutations with hematopoietic cancers. The diversity of RAS mutations in different cancer pathologies suggests nonredundant functions of each isoform and perhaps different oncogenic outputs. In support of this, Haigis et al. engineered knock-in mice to express endogenous KRAS or NRAS mutants in colonic epithelium (106). In this model, only KRAS was able to trigger hyperproliferation and enhance tumorigenesis while NRAS did not. Another study demonstrated that endogenously expressed mutant NRAS drove a mild

37 22 myeloproliferative disorder (MPD) in mice as well as promoted acute myeloid leukemia that is similar to human NRAS-driven leukemias. KRAS mutant expression on the other hand caused a more aggressive MPD phenotype resulting in rapid death and was likely due to higher expression of RAS-GTP levels than mutant NRAS (107). Despite studies suggesting specific roles of different RAS isoforms impacting cancer progression other studies have shown that RAS isoforms can be interchangeable. This is best demonstrated in a model of lung tumorigenesis where endogenously expressed oncogenic HRAS replaced oncogenic KRAS at the Kras promoter (108) and was able to recapitulate many of the same features of oncogenic KRAS in urethane-induced lung tumorigenesis. This suggests that local regulatory elements surrounding the promoter region determine the cancer specificity of each isoform. Oncogenic mutations of RAS found in tumors can impair GTPase activity, reduce responsiveness to RASGAPs, or enhance guanine nucleotide exchange of GDP to GTP (109). This leads to an abundance of GTP-bound RAS and constitutive activation of the many downstream effector pathways. Here, a general overview of the role of oncogenic RAS in modulating these functions is discussed. Proliferation Perhaps the most obvious role of oncogenic RAS is in regulation of cell proliferation due to stimulation of mitogenic pathways such as RAF-MEK-ERK. Initial studies revealed that oncogenic RAS can drive quiescent cells into the cell cycle (110). In addition, oncogenic RAS upregulates transcription of a number of growth factor genes such as TGF-α, AREG, and HBEGF to facilitate maximal proliferative potential ( ). In some cases, oncogenic RAS can also promote cell proliferation by

38 23 inhibition of anti-proliferative signals such as those induced by the cytostatic growth factor TGFβ1 and its downstream intracellular proteins, the Smads (114). Oncogenic RAS principally stimulates proliferation through the regulation of cyclin D1, which can be induced by all three major effector pathways as well as several downstream transcription factors including JUN, ELK1, and NFkB (115, 116). Furthermore, cyclin D1 protein stability can be increased through PI3K-dependent inhibition of GSK3β kinase, which phosphorylates and targets cyclin D1 for ubiquitination and proteasomal degradation (117). Oncogenic RAS can also promote cell cycle progression through inhibition of cyclin dependent kinase inhibitors (CKIs) p21 and p27, which inhibit cyclin dependent kinases (CDKs) with cyclin complexes ( ). Indeed, the importance of cyclin D1 as a growth promoting factor during RAS-induced transformation is further substantiated in mice lacking cyclin D1 (121). These mice are resistant to tumor development induced by oncogenic H-RAS in mammary and squamous tissues (121, 122). Apoptosis Evasion of the apoptotic machinery is one of the hallmarks of cancer progression (123). Apoptosis can be triggered by extrinsic factors such as growth factor withdrawal or detachment from extracellular matrix which activates cell surface death receptors or through intrinsic mechanisms mediated by the mitochondria such as DNA damage (123). Both extrinsic and intrinsic pathways converge on caspase 3 leading to activation and cell death. The maintenance and development of tumors depends on continued oncogenic RAS expression to suppress apoptosis. In transgenic mice expressing an inducible melanocyte targeted HRAS V12G withdrawal lead to melanoma

39 24 regression that was associated with significant apoptosis in both tumor cells and endothelial cells (124). Similarly, another study showed that withdrawal of inducible oncogenic K-RAS expression in type II pneumocytes caused apoptosis of both highly proliferative focal lesions and lung adenocarcinomas (125). From a mechanistic standpoint, oncogenic RAS suppresses apoptosis through regulation of various pro- and anti-apoptotic molecules mediated by the PI3K and RAF effector pathways. For example, PI3K can downregulate the pro-apoptotic BCL-2 family member BAK (126) or upregulate anti-apoptotic IAP family members through NFkB activation ( ). On the other hand, activation of RAF signaling by oncogenic RAS suppresses apoptosis through downregulation of pro-apoptotic transcriptional repressor PAR4 or upregulation of anti-apoptotic BCL-2 and ARC proteins (129, 130). Furthermore, RAF and PI3K pathways can cooperate in phosphorylation of proapoptotic BAD proteins on serine 136 and serine 122, which causes BAD to form an inactive complex with This ultimately prevents association of BAD with antiapoptotic proteins BCL-2 and BCL-XL, leading to their inactivation (131, 132). Paradoxically, oncogenic RAS can also trigger apoptosis depending on the cellular context and signals emitted (133). One study showed that while PI3K signaling suppresses apoptosis through an AKT dependent manner, preferential activation of RAF signaling triggers this response (134). Furthermore, whether oncogenic RAS suppresses or activates apoptotic programmes may be linked to JNK activation (135). Taken together, oncogenic RAS can both induce or inhibit apoptosis. However, given that oncogenic RAS mutations are frequently detected in cancer as well as frequent

40 25 activation of its effector pathways indicates that the inhibitory aspects of oncogenic RAS on apoptosis play a dominant role in cancer progression. Energy metabolism Cancer cells typically have increased metabolic demands due to increased proliferation rates and produce important metabolites to sustain growth and survival (136). Initial characterizations by Otto Warburg in the 1920s suggested that increased glucose uptake, and a shift from mitochondrial oxidative phosphorylation to aerobic glycolysis are typical of cancer cells, the so-called Warburg effect (137, 138). Although a rather inefficient process as less ATP is generated, the catabolism of glucose through glycolysis is efficient in producing macromolecules such as nucleotides, amino acids, and lipids that are required for the replication and growth of cancer cells (137). Oncogenic RAS regulates metabolic processes through two mechanisms. First, oncogenic RAS regulates HIF1α, which associates with ARNT to form a transcription factor complex that aids in the shift toward glycolysis. Second, oncogenic RAS regulates autophagy a process of self-eating that generates energy, produces macromolecules important in survival, and sustains organelle homeostasis (136). Both RAF and PI3K effector pathways cause increased mtor activity (139, 140) which. in turn induces HIF1α through post-transcription mechanisms that increase protein translation and stability (141). HIF1α induction increases glucose transport in cancer cells as well as processing of glucose into biological intermediates. For example, increased transcription of the glucose transporter GLUT1 enhances a cells ability to take up glucose and increased transcription of several glycolytic enzymes such as hexokinase and phosphofructokinase that aid in glucose breakdown ( ). Several

41 26 studies have shown that basal induction of autophagy by oncogenic RAS supports tumor growth and survival. Oncogenic RAS does this by maintaining mitochondrial oxidative phosphorylation and promoting glycolysis through the transcriptional induction of several autophagy related genes such as ATG5 and ATG7 ( ). Paradoxically, autophagy also has a documented role in preventing oncogenic RAS-induced transformation (149). Egelndy et al. demonstrated that non-tumorigenic HOSE cells transduced with oncogenic RAS underwent autophagy-mediated cell death that was dependent on an association between NOXA and Beclin1 and displacement of Mcl-1 (150). Furthermore, inhibition of NOXA or Beclin1 reduced autophagy leading to enhanced clonogenic survival of these cells. Another study showed that oncogenic RAS expression induced autophagy-mediated cell death through JNK activation and upregulation of ATG5 in rat fibroblasts (151). Taken together, evidence suggests that autophagy can either support tumor growth and survival or promotecell death in cells expressing oncogenic RAS. These discrepancies likely are cell context dependent as well as the type of effector pathways triggered by oncogenic RAS in these systems. Angiogenesis In addition to having cell autonomous effects, oncogenic RAS can promote changes within the tissue microenvironment that aid in tumor progression. The development of new blood vessels through the process of angiogenesis allows tumor cells as well as other cell types within the growing tumor access to an adequate supply of oxygen and nutrients (152, 153). While a complex process, oncogenic RAS can regulate several key pro-angiogenic growth factors that lead to endothelial cell proliferation and stromal remodeling allowing formation of new blood vessels. Major

42 27 pro-angiogenesis targets regulated by oncogenic RAS include VEGFA, bfgf, and PDGF as well as inflammatory cytokines such as IL-8 and IL-6 ( ). Both HIF1α and COX2 can mediate VEGF induction by oncogenic RAS. Elevated COX2 increases prostaglandin synthesis causing cyclin AMP dependent transcription at the VEGFA promoter (157, 158). COX2 can also induce bfgf and PDGF expression both of which are required for endothelial cell spreading and migration (159). Increased expression of IL-8 and IL-6 recruits immune cells such as neutrophils and macrophages to the tumor, which in turn secrete pro-angiogenic growth factors (160, 161). Cancer cells are surrounded by extracellular matrix (ECM), comprised mostly of collagen and laminin. The ECM is a reservoir for secreted growth factors and cytokines and restricts their access to endothelial and immune cells. Remodeling of the ECM is necessary for secreted growth factors and cytokines to reach endothelial cells and immune cells or reciprocally for infiltration of endothelial and immune cells into the growing tumor (162). Oncogenic RAS regulates several proteases such as MMP-2, MMP-9 and upa that can degrade the ECM or remove the barrier function of the basement membrane that surround cancer cells (159, 163). In particular, a number of studies have documented a crucial role for MMPs and upa in facilitating endothelial cell migration, adhesion and vessel sprouting (164, 165). Furthermore, infiltrating immune cells such as neutrophils or macrophages secrete MMPs and are important in driving epithelial-derived malignancies (166). Oncogenic RAS can also promote angiogenesis by negatively regulating inhibitors of the angiogenic switch such as Thrombospondin-1 (TSP1) (167). TSP1 prevents new blood vessel formation by inhibiting endothelial cell migration, survival and growth factor mobilization (168).

43 28 Metastasis One of the most detrimental aspects of a developing tumor is the acquisition of metastatic properties where cancer cells spread to the surrounding tissue and distant organs. Metastasis is a multi-step process that includes the establishment of local tumor cell invasion, transendothelial migration of cancer cells into the vasculature (intravasation), survival in the blood stream, reentry into the distant organ site (extravasation), and subsequent proliferation and colonization (169). Mutations in oncogenic RAS are prevalent in metastatic tumors of the lung, pancreas, and colon (170). Oncogenic RAS plays an important role in facilitating the metastatic potential of cancer cells at almost every step in the metastatic cascade. The first step, the establishment of local tumor cell invasion and destruction of the basement membrane at the primary organ site has been extensively studied of which oncogenic RAS has an important role. Oncogenic RAS promotes epithelial-mesenchymal transition (EMT), an invasive/migratory phenotype that is driven by alterations in cell-cell and cell-ecm interactions (171). E-cadherin and the cytoplasmic protein β-catenin, are part of adhesion junctions. Loss of E-cadherin and adherins junctions are hallmarks of EMT that are lost during cancer progression (172). Oncogenic RAS downregulates E- cadherin levels through multiple mechanisms including induction of E-cadherin transcriptional repressors SNAIL and SLUG, proteolytic degradation of the E-cadherin protein, or methylation of the E-cadherin promoter ( ). Oncogenic RAS can also disrupt the E-cadherin-β-catenin complex by facilitating β-catenin re-localization (174, 176). In addition, cell-cell and cell-ecm interactions can be disrupted by oncogenic RAS by reducing the levels of integrin receptors ( ). In addition to alterations in cell

44 29 contacts, oncogenic RAS protects cancer cells from anoikis, which is a form of apoptosis that occurs when detached from surrounding ECM (180, 181). This is especially important when cancer cells intravasate into the circulatory system and extravasate to distant organ sites for colonization. All three major effector pathways induced by oncogenic RAS promote metastatic spread. These include the RAF-MEK-ERK, PI3K, and RALGDS pathways. In some cases cooperation of oncogenic RAS with other pathways such as TGFβ1 signaling enhances metastasis (182). However, despite understanding the general nature of the pathways involved each can differentially regulate metastasis at distinct steps and also depends on the tissue involved and genetic background RAS and senescence The work of Heyflick and Moorhead first demonstrated that primary cells have a limited number of population doublings and eventually stop proliferating in vitro, even in optimal culturing conditions. This phenomenon was later defined as replicative senescence and is a process strongly linked to organismal aging (183). Even though senescent cells are metabolically active they are unable to respond to mitogenic stimuli or undergo further cell cycle divisions and are considered irreversibly arrested. Replicative senescence of primary cells is largely attributed to the shortening of telomeres, which are maintained by telomerase; however because primary cells express low levels of this enzyme they are often susceptible to the progressive chromosomal erosion of their telomeres with each round of cell division (184). In addition to telomere shortening, it is now clear that various stress-inducing stimuli that alter DNA integrity can trigger a senescence response such as DNA double strand

45 30 breaks or oxidative lesions caused by environmental insults, genetic defects or endogenous processes such as the production of ROS due to mitochondrial respiration ( ). While initial discoveries have shown that oncogenic RAS can transform immortalized cell lines or collaborate with other oncogenes to transform primary cells, introduction of oncogenic RAS alone in primary cells causes growth arrest followed by a phenotypically similar form of replicative senescence, called oncogene-induced senescence (OIS) (188). OIS is a state of premature senescence which occurs prior to any reduction in telomere length (189). Premature senescence is a powerful tumor suppressor response to oncogene activation. Moreover, premature senescence is likely the dominant mechanism by which premalignant or benign tumors are suppressed from development into malignant neoplasms (190). Hyperactivation of RAS-MEK-ERK pathway and robust mitogenic signals from in vitro models of oncogenic RAS overexpression or other oncogenic stimuli triggers the senescence response (191). Expression of oncogenic RAS at physiological levels from endogenous promoters without the deleterious effects of overexpression promotes cell proliferation and transformation but not senescence without the aid other cooperating oncogenes (192, 193) This suggests that low levels of oncogene stimulation enable evasion of the senescence response. As mentioned previously, senescence can be induced by a plethora of insults, but they all appear to converge on two tumor suppressor pathways that maintain the growth arrest associated senescent state. These two pathways include the gatekeeper proteins p53 and prb (183). p53 mediates responses to DNA damage and it has been shown that p53 loss abrogates replicative senescence of human and mouse cells (194, 195). In the context of RAS, the production of ROS due

46 31 to RAS-induced mitogenic effects causes a DNA damage response and is likely the causative factor that activates p53 ( ). p53 is negatively regulated by E3 ubiquitin ligases that target it for proteosomal degradation such as MDM2, PIRH2, and COP1 (199, 200). In contrast, positive regulation occurs through posttranslational processes that stabilize p53 such as ARF proteins in human (p14) or mouse (p19) (201). The level of p53 activation mediated by these negative and positive regulators can be an essential determinant of the senescence response. Increased p53 levels result in cell cycle growth arrest through transcriptional regulation of p21 which can inhibit the activity of Cyclin E/CDK2 and Cyclin D/CDK4/6 complexes. This promotes hypophosphorylation of prb, which prevents transcription of a number of cell cycle regulatory proteins and causes induction of senescence (202). While p53 inactivation in some cell types reverses senescence-associated growth arrest, other studies have shown that p53 loss is dispensable in other cell types (203). The causative factor that distinguishes cells that are still senescent despite p53 inactivation may depend on the cell cycle inhibitor p16 (203). p16 like the p53-p21 pathway prevents Rb phosphorylation via inhibition of Cyclin D/CDK4/6 activity and is induced by a number of stimuli including overexpression of oncogenic RAS (204). Furthermore, a number of positive and negative regulators of p16, including Bmi-1, CBX7, ID1, and Ets-1 regulate senescence that is dependent on prb ( ). However, p16 and prb may also function independently of one another during the senescence response. For example, deletion of p16, but not prb, did not impede senescence suggesting a compensatory role (209, 210). Together, p16 and p21 likely cooperate to maintain prb in the hypophosphorylated state during senescence or alternatively p21 first transiently inhibits

47 prb while later induction of p16 causes permanent prb hypophosphorylation and 32 subsequent irreversible senescence (211). Indeed, oncogenic RAS promotes senescence through induction of both p53 and p16 further supporting the idea that multiple tumor suppressor pathways contribute to fulfillment of this phenotype (198). In some cases, oncogenic RAS triggers an initial proliferation response that is immediately followed by premature senescence. These mitogenic signals induced by RAS in primary cells may serve as a failsafe mechanism from oncogenic transformation. Since RAS activates the RAF-MEK-ERK pathway it is now apparent that this pathway is paradoxically involved in the senescence response. When RAF was expressed in nonimmortal human fibroblasts it induced premature senescence similar to RAS and was largely dependent on p16 but not p53 (212). Similarly, constitutive MEK expression in primary human and mouse fibroblasts induced premature senescence; however this response was dependent on both p53 and p16 (212). One possible mechanism of the involvement of the RAF-MEK-ERK pathway during oncogenic RAS-induced senescence is through activation of stress-activated MAPK p38 (213). Wang et al. showed that constitutive activation of MKK3 or MKK6, which directly activates p38, prompted induction of senescence, that p38 inhibition blocked oncogenic RAS-induced senescence and that inhibition of the RAF-MEK-ERK pathway inhibited p38 induction (213). Deng et al. demonstrated that high intensity RAS signaling activated the p38 pathway via RAF-MEK-ERK but moderate levels did not and this led to cell growth and proliferation (214). Additionally, oncogenic RAS expressed at low levels maintained its transformative potential when expressed with other oncogenes, suggesting that RAS oncogenic activity is not sacrificed at the expense of gene dosage levels.

48 33 In addition to activation of tumor suppressor and aberrant mitogenic signaling pathways, there are a number of other characteristics of cells undergoing senescence induced by oncogenic RAS and other insults. One of the most easily distinguishable features of senescent cells is they become extremely large, flat and multinucleated irrespective of the senescent stimuli (183) In some cases, particularly primary melanocytes (183) there is extensive cellular vacuolization that is associated with this flattening morphology. One of the classical biomarkers of senescent cells is expression of senescence associated-β-galactosidase activity (SA-β-Gal). This increased activity is derived from the lysosomal β-d galactosidase enzyme and is likely due to expansion of the lysosomal compartment within the cell (215). However, there currently is no evidence indicating an actual role of SA-β-Gal in the senescence response (216). In some cases, senescent cells are also associated with the development of senescence associated heterochromatin foci (SAHF), which causes a redistribution of chromatin into localized punctate bodies and can be easily visualized with common DNA dyes (217). SAHF can further be characterized by histone modifications such as the presence or absence of methylation and/or acetylation on lysine residues of histone H3 (217). Functionally, these SAHF in senescent cells directly interfere with promoter regions of several E2F target genes and prevents their transcription and can be bypassed when the p16-rb pathway is abrogated (218) as well as by other tumor suppressor pathways important in the senescence response ( ). 1.5 Skin carcinogenesis Structure and physiology of skin

49 34 The skin is the largest organ of the body and forms a barrier against harmful pathogens, environmental insults such as radiation, temperature fluctuations and is a protective barrier to water loss (222). The skin is made up of three layers consisting of the epidermis, dermis and hypodermis (adipose tissue). Keratinocytes are the most abundant cell type within the epidermis. These cells express structural proteins that produce the epidermal barrier through a tightly regulated process of terminal differentiation (222, 223). The innermost layer of the epidermis or basal layer consists of proliferating cells. Some of these cells remain in the basal layer as self-renewing stems cells while their progeny migrate into the upper differentiating layers termed the spinous and granular layer. These migrating cells ultimately transition from spinous to granular layers and finally to the surface into a layer of dead cells, the stratum corneum (222, 223), which forms the water impermeable barrier. The transition from the basal layer to the stratum corneum in the epidermis is characterized by several biochemical changes. Keratins which are intermediate filament proteins constitutes approximately 30% of the protein in basal keratinocytes but increases to 85% in differentiated keratinocytes (224). Keratins can be divided into two groups (type I or type II) based on amino acid sequence, molecular mass, or isoelectric ph. Basal keratinocytes express heterodimer pairs of type II keratin K5 and type I keratin K14 (225). As the basal keratinocyte transitions into the spinous layer, the expression of K5 and K14 is turned off while K1 and K10 expression are induced (225, 226). K1 and K10 are only expressed in the suprabasal layers and represent one of the earliest biochemical markers of terminal differentiation (226). In addition to keratins, epidermal keratinocytes produce other structural components that are restricted to the

50 35 differentiating layers and provide rigidity and stability to these cells. Involucrin is a soluble cytosolic protein that is gradually synthesized at much higher levels in spinous and granular layers and eventually becomes covalently cross-linked into the plasma membrane of cells within the stratum corneum (227). Crosslinking is mediated by transglutaminase, which catalyzes the formation of an isopeptide bond between involucrin and a membrane-bound precursor protein (228). When keratin filaments move from the granular layer to the stratum corneum they form dense protein aggregates within cornified cells which is mediated by filaggrin (229). Filaggrin is synthesized as a precursor protein in the granular layer that is unable to aid in keratin aggregation due to being heavily phosphorylated. However, in the cornified layer, filaggrin becomes dephosphorylated and is proteolytically cleaved to allow interaction with keratin filaments (230). Regulation of keratinocyte growth and differentiation is a highly coordinated process that is mediated by several extracellular signals and growth factors to maintain homeostasis. Therefore, determining the molecular mechanism of growth and differentiation in the epidermis is of utmost importance for understanding relevant changes associated with different skin diseases and cancer. Calcium is perhaps the most important and best studied regulator of keratinocyte differentiation. Based on studies performed in vitro and in vivo, a calcium gradient exists within the epidermis with low concentrations in basal keratinocytes but higher levels in differentiated keratinocytes (231). When keratinocytes isolated from newborn mice are cultured in media containing 0.05mM Ca 2 + they exhibit a proliferative basal cell phenotype (232). However, when switched to 0.12mM Ca 2+, these cells undergo an irreversible growth

51 36 arrest and display features of stratification and cornification (232). Increased extracellular Ca 2+ in vitro promotes many of the same markers observed in vivo including the appearance of desmosomes, increased expression of K1 and K10, loss of K5 and K14, and increased expression of involucrin and filaggrin (223, 233). Increasing extracellular Ca 2+ levels induces keratinocyte differentiation by triggering PLC activity, which causes an increase in DAG leading to PKC activation ( ). PKC family members, which are serine threonine protein kinases, are important in the regulation of terminal differentiation (237). PKCα, the prototypical member of this family, is localized to the membranes of differentiating keratinocytes and can be found associated with desmosomes (238, 239). PKCα activation induces loricrin and filaggrin (240) possibly through the activation of AP1 transcription factor proteins (241). In contrast to calcium, vitamin A and other related retinoid compounds suppress keratinocyte differentiation ( ). Previous studies have shown retinoids inhibit expression of early differentiation markers K1 and K10 (244), suppress involucrin and filaggrin expression ( ), inhibit synthesis of desmosomes (248) and impede cornified envelope formation (249). Retinoids suppress differentiation by transcriptional mechanisms involving activation of intracellular RAR steroid hormone receptors (246, 247, 250) which bind to consensus DNA sequences of target genes. RARγ is the major isoform expressed in skin and in cultured keratinocytes with RARα having minor contributions (251, 252). RARs are able to heterodimerize with each other, RXRs, and with thyroid receptors and can interact with AP1 proteins to negatively regulate differentiation-associated genes (253, 254). Interestingly, in contrast to activation of RARs with retinoids, dominant negative RAR mutants expressed in the epidermis inhibit

52 37 differentiation suggesting complex interactions exist between RAR and other nuclear proteins during terminal differentiation (255, 256). Epidermal keratinocytes that undergo terminal differentiation must be able to exit the cell cycle in an irreversible manner. As such, negative regulation of keratinocyte cell growth is controlled via autocrine effects of the growth inhibitor TGFβ1 in basal keratinocytes and TGFβ2 in differentiated keratinocytes (257). However, TGFβ1 and TGFβ2 can also maintain the basal keratinocyte phenotype through increased expression of K5 and K14 (258). This suggests that while TGFβs can suppress growth of basal keratinocytes and set the stage to differentiation; they likely work in concert with other signaling mediators such as those mentioned earlier to fully commit to the maturation process. In contrast to TGFβs, stimulation of keratinocyte cell growth is mediated by TGFα and other EGF receptor ligands (259). EGF ligand-mediated cell growth is restricted to basal keratinocytes as EGF receptors are only expressed on these cells (259). Indeed, EGF ligands are up-regulated in hyperproliferative skin disorders and overexpression in the epidermis can produce similar phenotypes (260) Two stage skin chemical carcinogenesis Most human cancers originate from epithelial-derived tissues including but not limited to breast, prostate, lung, and skin. The two-stage skin chemical carcinogenesis model represents a biologically relevant model that mimics the natural progression of epithelial malignancies that occurs in humans. This model is characterized by initiation, promotion, and malignant conversion that reflects the multi-stage development of cancer (261). Initiation in this model is caused by a single topical dose of the carcinogen 7, 12 dimethylbenz [a] athracene (DMBA), which undergoes bioactivation

53 38 into highly reactive diol epoxide metabolites and causes irreversible DNA damage and mutation (262). Amoung many mutations, DMBA causes a point mutation at codon 61 in the HRas gene of epidermal keratinocytes and produces a constitutively active, oncogenic GTPase (263, 264). This mutation is present in nearly all benign and malignant skin tumors in this model and can be detected by PCR as early as 4-5 weeks suggesting it is the cancer intiating mutation. The most likely cellular targets for DMBAinduced HRAS mutations are proliferating basal keratinocytes in the interfollicular epidermis or in the bulge region of the hair follicle, which contain stem cells (265, 266). Initiation alone with DMBA does not cause tumor formation, however repeated applications of a tumor promoter for many weeks, such as 2-O-tetradecanoylphorbol- 13-acetate (TPA), can promote clonal expansion of RAS-initiated keratinocytes that leads to development of benign papillomas. TPA is a non-genotoxic phorbal ester and a primary activator of PKC signaling (261). Repeated TPA applications following DMBA exposure alters the balance between cells that will terminally differentiate versus those that undergo proliferation (267), and eventually leads to epidermal hyperplasia and focal development of tumors. TPA provides a selective advantage for outgrowth of keratinocytes with RAS mutations since these cells are resistant to the accelerated differentiation ( ). A significant percentage of benign papillomas are dependent on repeated TPA applications for continued outgrowth. If TPA treatment is stopped these tumors frequently regress, or persist in a permenant benign state. In most inbred mouse strains very few squamous benign papillomas convert to invasive SCC. Several studies have identified two subclasses of benign papillomas with different propensities to undergo malignant progression to invasive SCC. These are characterized by several

54 39 biomarkers that allow discrimination at early premalignant stages ( ). Low-risk papillomas are terminally benign and do not progress to SCC while high-risk papillomas progress to SCC at high frequency (271). High-risk papillomas exhibit a gene signature similar to SCC while low-risk benign lesions do not (275). Tumor promoters such as TPA do not further increase the frequency of malignant progression but rather is a spontaneous process brought on by accumulation of additional mutations, chromosomal abnormalities within papillomas such as trisomies on chromosome 6 and 7 and aneuploidy (276). Benign papillomas are heterozygous for the mutant HRas allele but become homozygous in SCC (277, 278). In addition, when oncogenic HRAS is introduced into papilloma cell lines which harbor heterozygous HRAS mutations, they progress to carcinomas suggesting gene dosage is a critical event to malignant progression (279, 280). Mutations in the tumor suppressor p53 also play a role in malignant progression. p53 mutations are rare in chemically-induced benign papillomas but are frequently present in carcinomas (281, 282). Furthermore, p53-deficent mice have no effect on the number and size of benign papillomas when compared to wild type mice but of those tumors that do develop there is a higher percentage that progress to invasive SCC (283). An in vitro surrogate to study the effects of oncogenic RAS in skin carcinogenesis two stage is transduction of primary keratinocytes with a replication defective retrovirus containing the v-ras Ha gene (284). This method recapitulates the phenotype of the initiated preneoplastic keratinocyte caused by DMBA exposure in vivo. When v-ras Ha transduced keratinocytes are mixed with fibroblasts and transplanted onto athymic mice, they form benign papillomas that are histologically similar to those generated from

55 40 A B Figure 1.6. Modeling of RAS-driven multistage epithelial carcinogenesis in vivo and in vitro. (A) Treatment of mouse skin with the carcinogen DMBA followed by repeated applications of the tumor promoter TPA causes clonal expansion and development of benign papillomas that contain an activating HRAS mutation at codon 61. Eventually, some papillomas progress into invasive SCC and are characterized by tumor suppressor loss such as p53, aneuploidy, and amplification of mutant HRAS allele and loss of HRAS wild-type allele. (B) Transduction of newborn primary keratinocytes with v-ras Ha in vitro undergo proliferation followed by growth and senescence and mimic the benign papilloma phenotype when grafted onto nude mice. Perturbations of various tumor suppressor pathways such as TGFβ1, p53, and the INK4a/ARF locus leads to bypass of senescence, resistance to differentiation, and form invasive SCC when grafted on nude mice or undergo malignant conversion in vitro.

56 41 two-stage chemical carcinogenesis (264). In vitro v-ras Ha keratinocytes undergo rapid proliferation followed by growth arrest and premature senescence expressing similar senescence markers as discussed in section (285). A small percentage of v- RAS Ha keratinocytes cultured in vitro acquire resistance to growth arrest and differentiation signals induced by elevated calcium (286). These cells have properties similar to SCC derived from two-stage chemical carcinogenesis and form SCC in vivo when grafted on nude mice(286). Conversely, SCC derived from tumors can proliferate in elevated calcium ( ). Perturbation of various signaling molecules can lead to escape from senescence and increase malignant conversion in this in vitro model. For example, inactivation of TGFβ1 signaling (285, 289), loss of p16 and p19 (290), and p53 (291) all bypass senescence and promote malignant conversion (Figure 1.6). 1.6 TGFβ1 signaling Overview of TGFβ1 signaling pathway TGFβ1 is the best characterized member of a large family of structurally related growth factors with a multitude of functions in virtually every cell type and across a myriad of species (292, 293). TGFβ1 was first discovered from supernatants of sarcoma virus transduced cell lines that when added to culture media of untransformed cells caused a phenotype similar to the original transformed cell. Through subsequent purification and further characterization two major transforming factors were isolated termed TGFα, later determined to be a ligand for the EGF receptor and TGFβ, later named TGFβ1 (294).

57 Over the past 30 years other growth factors structurally related to TGFβ gave rise to the TGFβ superfamily with approximately 27 members. The TGFβ superfamily has 42 Figure 1.7. TGFβ1 signaling pathway. Receptor complex formation induced by TGFβ1 binding phosphorylates a member of the R-Smads (Smad2 or Smad3), enabling its association with the Co-Smad, Smad4 and translocation into the nucleus. In the nucleus, the activated smad complex associates with various DNA binding cofactors, transcriptional coactivators or repressors that determines transcriptional status of target genes. In the unphosphorylated state, R-Smads are retained in the cytoplasm by binding to SARA.

58 43 been categorized into two subfamilies 1) TGFβ/Activin/Nodal subfamily and 2) BMP/GDF subfamily. Some members are expressed in many cell types and throughout development and adulthood while others are restricted to certain cell types and expressed only during a specific developmental stage (292). Together, TGFβ1 and other related members have significant roles in nearly all phases of development and tissue homeostasis, and in pathological diseases such as cancer making them important regulators of processes such as cell growth, differentiation, migration, invasion, and apoptosis (292, 295). TGFβ1 activates downstream signaling by binding and assembling a cell surface receptor complex within the plasma membrane, in turn activates receptor regulated intracellular transcription factors called R-Smads (Figure 1.7). TGFβ1 binding promotes association of two types of serine/threonine receptor kinases, the type I (also called ALK5) and type II receptors. First, TGFβ1 binds to the basally active type II receptor which then recruits and phosphorylates serine and threonine residues within the conserved GS domain of ALK5 (296). Phosphorylated ALK5 in turn phosphorylates serine residues in the C-terminal SSXS motif of Smad2 and Smad3. These R-Smads are in an inactive state in the cytoplasm bound to SARA (297), and phosphorylation of Smad2 and Smad3 diminishes their affinity to SARA and exposes a nuclear import signal (298). Phosphorylation of R-Smads also increases their affinity for Smad4 (298). This R-Smad/Smad4 heteromeric complex then translocates to the nucleus to regulate target genes (292). Once in the nucleus, the R-Smad/Smad4 complex binds to Smad binding element (SBE) consensus sequences and associates with various DNA-binding cofactors and allowing transcriptional activation. Negative regulation of this pathway

59 44 comes in the form of inhibitory Smads (I-Smads), Smad6 and Smad7. Smad6 is specific for the BMP/GDF subfamily and prevents Smad1, 5, and 8 from interacting with Smad4 (299). On the other hand, Smad7 is specific for the TGFβ/Activin/Nodal subfamily and blocks phosphorylation of Smad2 and Smad3 by the TGFβ receptor complex (300). Both inhibitory Smads are regulated through R-Smad/Smad4 complex binding to their gene promoters thus forming a negative feedback loop (300). In addition to Smad-dependent transcriptional activation, TGFβ1 regulates other signaling pathways that are Smad independent such as the ERK, p38, and JNK signaling pathways (301). Evidence that TGFβ1 regulates these pathways through Smad-independent mechanisms comes from studies using Smad4-deficent cells or Smad dominant-negative approaches (302, 303). While currently unclear, the mechanism by which TGFβ1 activates these signaling pathways may be dependent on RAS activation (304) or through activation TAK1 (305). Regardless, activation of these pathways may serve to amplify or dampen Smad-dependent transcriptional responses as ERK and JNK pathways can regulate Smads through a variety of mechanisms ( ). TGFβ1 is best known as a potent inhibitor of cell growth and proliferation in multiple cell types including epithelial, endothelial, hematopoietic, and mesenchymal cells (309). TGFβ1-mediated inhibition of cell proliferation is primarily through upregulation of CDK inhibitors to cause a G1 cycle growth arrest (Figure 1.8). Perhaps the best known transcriptional target by TGFβ1 is the CDK inhibitor p15 (310). p15 inhibits cyclin D-dependent kinases, CDK4 and CDK6, by binding to the CDK and

60 interfering with the cataylic activity and thus preventing assembly of cyclin D-CDK4/6 complexes. Cyclin D-CDK4/6 promote proliferation by serving as a mitogen sensor, 45 Figure 1.8. TGFβ1-mediated growth inhibition in normal epithelial cells. TGFβ1 induces two distinct anti-proliferative responses through either c-myc downregulation or modulation of CDK inhibitory responses. CDK inhibition occurs through upregulation of p15 and p21 and downregulation of cdc25a. C-myc antagonizes TGFβ1 signaling by repressing CDK inhibition. Downregulation of cdc25a and upregulation of p15 and p21 leads to inhibition of cyclind-cdk4/6 and cyclin E-CDK2 and fullfillment of cell cycle growth arrest.

61 46 thus inhibiting this association impedes G1 progression. Other studies have shown that p15 upregulation by TGFβ1 can impede G1 progression through other ways, including prevention of sequestration of CDK inhibitor p27 by cyclin D-CDK4/6 (311, 312). Here, p27 normally inhibits G1 progression through association with cyclin E-CDK2 complexes, however when cells are proliferative p27 remains inactive through association with cyclin D-CDK4/6 complexes. The association of p27 with cyclind- CDK4/6 ends once TGFβ1 induces p15 leading to displacement of p27 and association with cyclin-cdk2 and G1 growth arrest. TGFβ1 also can induce the CKI p21 in multiple cell types which can associate and inactivate cyclin D-CDK4/6 and cyclin E-CDK2 complexes (313). In some cases, TGFβ1 can also induce p16 and p19 although their role seems to be associated with irreversible premature senescence and not transient G1 growth arrest (314). TGFβ1 can also inhibit CDK activity indirectly through downregulation of cdc25a (315). Cdc25A is a tyrosine protein phosphatase which removes the inhibitory effects of tyrosine phosphorylation on CDK proteins. TGFβ1 downregulation of cdc25a in mammary epithelial cells leads to increased tyrosine phosphorylation of CDK4 and CDK6 and subsequent kinase inhibition (316). TGFβ1 can also cause transcriptional downregulation of c-myc, which occurs upstream of CDK inhibition (317, 318). C-myc is a member of the basic helix-loop-helix leucine zipper family of transcription factors and is a promoter of cell growth and proliferation and is also a potent oncogene. C-myc can act as an activator and repressor of transcription through a number of mechanisms, and transcriptional activation usually occurs in a complex with Max, which binds to E-box enhancer elements on DNA. C-Myc transcriptional downregulation is a rapid and transient response to TGFβ1 which results

62 in rapid reduction in protein levels due to a short protein half-life (309). C-myc 47 downregulation is also critical for induction of p15 by TGFβ1. C-myc acts as a repressor at the p15 promoter through association with Miz1. TGFβ1 treatment prevents interaction of c-myc to Miz1 that relieves repression of p15 allowing activation by Smads (319). Taken together, TGFβ1 signaling triggers a coordinated program that upregulates multiple inhibitors of cell cycle progression as well inhibits key mitogens that ultimately leads to a potent G1 cell cycle arrest in normal epithelial cells Dual role of TGFβ1 signaling in cancer Since TGFβ1 is a potent growth inhibitor of many epithelial cell types it comes as no surprise that it functions as a suppressor of tumorigenesis. As such, the antiproliferative response induced by TGFβ1 is normal epithelial cells is frequently lost in tumor-derived cell lines (320) and nearly all pancreatic and colon cancers have mutations that inactivate critical components of the TGFβ1 pathway (321, 322). TβRII inactivating mutations occur in most human colorectal and gastric carcinomas with microsatellite instability (MSI) (323). Introduction of wild-type TβRII into colon or gastric cell lines with MSI restores TGFβ1-mediated growth arrest and mitigates tumorigenic potential when introduced into athymic mice suggesting that loss of this pathway is a required early tumorigenic event (324, 325). Mutational inactivation in TβRI also occurs in human cancer as well. About one third of ovarian cancers contain TβRI mutations and this occurs with no apparent TBRII mutations within the same sample set (326). In addition, deletions of TβRI have also been observed in pancreatic and billary carcinomas as well as in T-cell lymphomas at low frequencies (321, 327). The downstream intracellular mediator proteins of TGFβ1 signaling, the Smad proteins are

63 48 also disrupted in human cancer. Smad4 is deleted in one half of all pancreatic cancers and one third of colon cancers, and mutations have been found at lower frequencies in a variety of other carcinomas (328, 329). Interestingly, Smad4 mutations have been found associated with TβRI and TβRII mutations in biliary and colon cancer, respectively (321). Smad2 mutations have been found in a small percentage of colon carcinomas (330). No Smad3 mutations have been found in human cancer; however studies in mouse models suggest a critical role for this protein. For example, Smad3 null mice develop metastatic colon cancer at an early age (331), although another study shows that Smad3 loss in mice promotes resistance to chemically-induced tumor development (332). Thus, Smad3 may have both tumor suppressor and oncogenic roles during cancer development. Paradoxically, TGFβ1 signaling exacerbates tumor progression and the malignant phenotype in experimental systems and evidence also suggests this occurs in human cancer. High levels of TGFβ1 correlate with advanced clinical stage of the tumor in breast (333), lung (334), and colon cancer (335) and this is associated with a shorter time to relapse, disease progression, and metastasis. In vitro systems using human cancer cell lines that overexpress TGFβ1 or that inhibit endogenous TGFβ1 signaling support the idea that this pathway enhances the malignant phenotype ( ). Tumor-derived TGFβ1 could contribute to the malignant phenotype by suppressing immune surveillance, enhance angiogenesis, and alter the balance of extracellular matrix production and degradation, or promotion of a mesenchymal phenotype by EMT (339). Many of these functions are due to autocrine or paracrine

64 effects in which cancer cell lines have selectively lost their anti-proliferative responses to TGFβ1 (180, 340, 341) Interactions between TGFβ1 and RAS signaling The switch from tumor suppressor to pro-oncogenic factor may depend on context and stage specific effects of the growing tumor on TGFβ1 signaling. One idea is genetic changes within a tumor alters TGFβ1 signaling in such a way to subvert its growth inhibitory functions but promote other biological responses that leads to cancer progression. Indeed, antagonistic and synergistic interactions exist between oncogenic RAS and TGFβ1 signaling that dictate the type response the cancer cell elicits (Figure 1.9). In normal epithelial cells, RAS signaling may be important to TGFβ1 signaling as blockade of RAS activation or downstream signaling inhibits TGFβ1 signaling (342). Reciprocally, activation of TGFβ1 signaling in these cells can lead to RAS activation (343). However, overexpression of oncogenic RAS in cancer cell lines or activation of MEK-ERK signaling blocks TGFβ1 signaling through phosphorylation of Smad proteins within the linker region by ERK (308). In addition, further investigations by Sekimoto et al. have shown that Smad linker phosphorylation by oncogenic RAS is required for oncogenesis since this fosters tumor cell invasion through upregulation of matrix degrading proteases (344). In some cases oncogenic RAS can also inhibit TGFβ1 signaling through degradation of Smad4 (345). Furthermore, other studies have shown that activated RAS signaling leads to activation of TGFβ1 transcriptional corepressors such as Ski/SnoN and TGIF which may selectively inhibit critical TGFβ1 target genes ( ). However, in in vitro models of early stages of carcinogenesis TGFβ1 still

65 Figure 1.9. General schematic of antagonistic and synergistic interactions between RAS and TGFβ1 signaling pathways. 1) Inhibition of Smads by activated ERK can prevent translocation and target gene expression 2) TGFβ1 can inhibit proliferation and promote senescence of RAS transformed epithelial cells and prevent RAS-driven tumor development. 3) RAS and TGFβ1 can synergize in vitro and in vivo to promote EMT and invasion. 50

66 51 retains significant abilities to inhibit oncogenic RAS-induced proliferation (314, 349). In addition, this is supported by many in vivo transgenic mouse models where TGFβ1 overexpression can inhibit RAS-induced tumor formation or loss of this pathway accelerates early stages of cancer progression (340, 350, 351) While studies have shown that oncogenic RAS and downstream pathways can inhibit TGFβ1 signaling or in other cases TGFβ1 signaling remains intact to suppress RAS-induced transformation, there are numerous reports demonstrating synergistic and cooperative actions of both pathways to promote malignant progression. These actions have largely been documented with the promotion of EMT, a migratory and invasive phenotype associated with later stages of cancer and characterized by loss of E- cadherin and expression of mesenchymal markers such as vimentin (352). Oncogenic RAS and TGFβ1 cooperate to induce EMT in both human and mouse breast squamous cell carcinoma cell lines. When normal EpH4 mammary epithelial cell line is treated with TGFβ1 they undergo growth arrest and apoptosis, however when this cell line is transduced with RAS it blocks the growth inhibitory response by TGFβ1 and induces EMT and invasion (353, 354). This cooperative interaction requires ERK and Smad signaling and is dependent on a SNAIL-SMAD3/4 repressor complex on the gene promoters of E-cadherin and CAR to facilitate EMT ( ). Another study showed that Smad2 synergizes with RAS to drive EMT and metastasis in multistage chemical carcinogenesis and this is dependent on increasing levels of nuclear Smad2 driven by oncogenic RAS to promote this phenotype (356). Furthermore, overexpression of TGFβ1 can suppress clonal outgrowth of benign tumors in which oncogenic RAS is the initiated oncogene, but of those tumors that do develop form highly invasive, local

67 52 spindle cell carcinomas (340). In addition, inducible overexpression of TGFβ1 for 15 weeks in benign papillomas induced by chemical carcinogens promotes a metastatic phenotype with similar features to EMT (357). 1.7 Hypothesis and Aims As discussed above, previous studies of the UPR using models that involve pharmacological-induced ER stress have delineated many of the important parameters that lead to adaptation or apoptosis of cells in response to ER stress. However, an understanding of the role of the UPR and physiologically-induced ER stress in cancer development is lacking. IRE1α, one the most evolutionarily conserved and important members of the UPR, has an emerging role as an executioner of different cell fate responses (3). Due to this complexity and the multiple functions regulated by its intrinsic RNase activity, it is not surprising that a clear understanding of IRE1α function in cancer progression is lacking. To push this research field forward, we have used our wellestablished in vitro model of skin carcinogenesis that similarly has divergent cell fate responses to various perturbations driven by oncogenic HRAS. We have used this model to characterize both adaptive and destructive functions of IRE1α. We hypothesize that IRE1α provides contrasting RNase activities, similar to the chemical ER stress models, to dictate the phenotype of the cancer cell. In chapter 3 I present data which reveal a novel and paradoxical role for two distinct IRE1α RNase outputs in RAS-driven epithelial carcinogenesis. I show that IRE1α-mediated Xbp1 mrna splicing promotes cell proliferation that is ER stress dependent. In contrast, IRE1α-mediated RIDD promotes growth arrest and senescence

68 53 independent of XBP1 and blockade of this response enhances malignant conversion. Furthermore, I have identified Id1 as a novel mrna cleavage target of IRE1α and its downregulation is responsible for promoting senescence induced by oncogenic v- RAS Ha. With the development of kinase inhibitors that can selectively inhibit or activate the RNase of IRE1α (50, 358), these data have significant ramifications for the rational use of these drugs as a therapeutic option for the treatment of cancer and possibly other pathophysiological diseases. In chapter 4 I show that the potent tumor suppressor, TGFβ1, can modulate IRE1α RNase functions in v-ras Ha expressing keratinocytes. TGFβ1 suppresses Xbp1 mrna splicing and enhances RIDD to promote premature senescence. This is a novel mechanism of tumor suppression by TGFβ1. Furthermore, RIDD is required for TGFβ1- induced senescence through a mechanism that does not immediately alter Smad transactivation. Lastly, I provide evidence that mild levels of ER stress, which activates the adaptive arm of the UPR, can inhibit TGFβ1 signaling and senescence thus providing a new mechanism to dampen this tumor suppressor pathway and promote oncogenic RAS-induced transformation during epithelial carcinogenesis. Together, these studies highlight a previously underappreciated role for the physiological significance of the ER and its stress sensor, IRE1α, in epithelial cancer development harboring oncogenic RAS mutations and could be a desirable therapeutic target outside traditional targeting of classical signaling pathways. These data contribute new and meaningful insight into how IRE1α affects the carcinogenic process through modulation of its RNase activity and governs cell fate through either oncogenesis or tumor suppression.

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100 85 Chapter 2: Materials and Methods 2.1 Cell culture and reagents Primary mouse keratinocytes from either FVB/n or C57/Bl7 mouse strains were isolated from 1-3 day old newborn littermates ccording to standard protocols (1). K5rTA x tetoras V12G (2) mice that express human HRAS V12G when induced were obtained from the NCI mouse repository. Newborn mice were obtained from crosses of heterozygous K5rtTA (K5) and homozygous tetoras V12G (tetoras) mice and genotyped by PCR. Primary keratinocytes were plated in Eagle's minimal essential medium (Formula # G, Lonza) containing 8% chelexed FBS, 18.3 I.U/ml penicillin, 18.3 μg/ml streptomycin and 0.2 mm Ca 2+. The following day, media was changed to 0.05 mm Ca 2+. K5rTA x tetoras V12G keratinocytes was induced by addition of 1μg/ml doxycyline (Dox) to culture media. Short-term inhibition of ER stress with 4- phenyl butyrate (4-PBA, Sigma) began on day 4 after v-ras Ha transduction for 24 hours and was achieved using varying doses as indicated (1-5mM). Short-term inhibition of MEK signaling with 5µM U0126 (Millipore) in v-ras Ha keratinocytes began on day 2 after transduction for 48 hours. Short-term inhibition of IRE1α RNase activity (both Xbp1 mrna splicing and RIDD) with 25µM 4µ8C (Millipore) in v-ras Ha keratinocytes began on day 2 after transduction for 24 or 48 hours. Inhibition of ALK5 in v-ras Ha keratinocytes with 10µM SB (Sigma) began on day 2 after transduction and refreshed every 2 days. TGFβ1(R & D systems) was used at 1ng/mL and refreshed every 2 days. Long term treatment of v-ras Ha keratinocytes were dosed with U0126 (5µM), and 4µ8C (25µM) which began on day 2 after transduction and refreshed every

101 86 2 days until harvest at indicated times. DMSO was used as vehicle control where indicated. 2.2 Plasmids The plasmids ppax2, pmd2g, and pwpi empty vector were obtained from Addgene. Plasmid psbe4-luc was a gift from Joan Massague (Memorial Sloan- Kettering, New York). Plasmid plko.1puro constructs containing the target gene shrna sequences were obtained from Sigma: XBP1 shrna was 5 - CCGGCCCAGCTGATTAGTGTCTAAACTCGA GTTTAGACACTAATCAGCT GGGTTTTT-3 ; IRE1α shrna was 5 -CCGGCCCACTTCTCTTTCTTTCT AACTCGAGTTAGAAAGAAAGAGAA GTGGGTTTTT-3. Non-target control shrna was obtained from Sigma. Plasmid pwpi-xbp1s overexpression construct was made using In-Fusion HD cloning kit following the manufacturer s instructions (Clontech). 2.3 Virus production and infection The v-ras HA retrovirus was generated from ψ2 producer cells as described previously (3). Virus titer was determined using a NIH3T3 focus-forming assay and was routinely 1 x 10 7 virus/ml. Primary keratinocytes were infected with v-ras HA retrovirus in the presence of 4µg/mL polybrene on day 3 of culture at an MOI 2-3 to ensure nearly 100% of cells were infected. HEK293T/N cells were used for shrna lentivirus production. Transient transfections were performed using Lipofectamine 3000 (Invitrogen) at a 1:1 ratio using a combination of ppax2 and pmd2g, and appropriate plko.1puro shrna vector in serum free and antibiotic free high glucose DMEM media supplemented with 10mM HEPES, 1X non-essential amino acids, 1mM sodium pyruvate, and 0.75mg/mL sodium bicarbonate. Lentiviral supernatants were collected

102 87 twice over a 96 hour period. Cell debris was pelleted and supernatant was stored at 4ºC in ice-cold PEG solution (10% PEG 6000, 0.5M NaCl) for up to 3 days. Lentivirus was spun 3000xg using a swinging bucket centrifuge for 30 minutes and supernatant aspirated. After another spin for 5 minutes and residual media removed, lentivirus pellets were suspended in ice-cold 1X PBS (ph 7.4) concentrated by fold. Aliquots were made and stored at -80ºC. Lentiviral titers were determined using a quantitative PCR-based assay kit following the manufacturer s instructions (ABMgood). Primary keratinocytes were infected on day 3, along with v-ras Ha as described above, with shrna lentivirus at an MOI 20. MOI was chosen after doing titrations of increasing amounts of lentivirus that yielded effective knockdown of target mrnas. After 2 days infection, cells were selected with 1µg/mL puromycin for 2 days and subsequent analysis at different timepoints examined. This protocol yielded nearly 100% survival and knockdown to near undetectable levels as determined by western blot. 2.4 BrdU incorporation assay At indicated times, cells were pulsed with 10µM 5-bromo-2 -deoxyuridine (BrdU) for 45 minutes, washed 2X with 1X PBS (ph 7.4), and stored at 4ºC in 70% ethanol until staining. For staining, cells were treated with 0.7N NaOH for 2 minutes, washed with 1X PBS, and blocked with 5% normal goat serum for 30 minutes at room temperature and subsequently incubated with primary mouse anti-brdu antibody (1: 50, Becton Dickinson) and incubated for 1 hr at room temperature. Cells were then incubated with secondary biotinylated goat anti-mouse IgG (1:1000, Jackson ImmunoResearch) for 30 minutes at room temperature followed by incubation in avidin biotin horseradish peroxidase (ABC kit, Vector Laboratories) for 30 minutes at room temperature. 3,3 -

103 88 diaminobenzidine (DAB) substrate kit for peroxidase (Vector Laboratories) was used following the manufacturer s recommended procedures for detection of positively labeled cells. The percentage of BrdU-positive cells was quantified using a Nikon inverted microscope (20x microscope frame) and were expressed as a percentage of total cells for each treatment group. Three different fields from each well were counted (at least 100 cells per field) and triplicate samples were analyzed for each treatment group. 2.5 Senescence associated β-galactosidase (SA-β-Gal) assay At indicated timepoints, cells were washed 2 times with PBS (ph 7.4) and fixed for 5 minutes in 0.5% glutaraldehyde in PBS (ph 7.4). The fixative was aspirated and cells were washed twice with PBS (ph 7.4). The PBS solution was removed and 500μl of staining solution (0.5 mm K3Fe[CN]5, 0.5mM K4Fe[CN]6, 1mM MgCl2, 1mg/mL X- gal (Gold Biotechnology) in PBS, ph 6.0) was added to each well. Cells were incubated with staining solution for 24 hours at 37 C in a non-co2 incubator. Positive cells stained blue. The percentage of β-galactosidase positive cells was quantified using a Nikon inverted microscope (20x microscope frame) and were expressed as a percentage of total cells for each treatment group. Three different fields from each well were counted (at least 100 cells per field) and triplicate samples were analyzed for each treatment group. 2.6 In vitro malignant conversion assay Briefly, primary keratinocytes were seeded in 6-well culture dishes, cotransduced with v-ras Ha or corresponding shrna for 2 days, selected with puromycin (1ug/mL) for 2 days and then cultured for 10 days further in Eagle's minimal essential

104 89 medium containing 0.05 mm Ca 2++ and chelexed 8% FBS. The cells were then switched to 0.5 mm Ca 2++ medium for 6 weeks, and colonies of calcium-resistant keratinocytes were identified by staining the dishes with 0.35% rhodamine/10% formalin and counted with a dissecting microscope. Foci isolated from this assay produce carcinomas when tested in vivo in a skin graft (4). 2.7 RNA isolation and quantitative RT-PCR (q-pcr) Total RNA was isolated using Ribozol (Amresco) according to the manufacturer s instructions. Reverse transcription and quantitative PCR (qpcr) was performed as previously described except intron-spanning primers were used (5). The relative level of mrna was normalized to either glyceraldehyde 3-phosphate (Gapdh) or 18s RNA levels. The following primers were used for qpcr analysis: Xbp1u forward primer: 5 - AGTCCGCAGCACTCAGACTAT-3 ; Xbp1u reverse primer: 5 -TGAAGAGGCAACAGT GTCAGA-3 ; Xbp1s forward primer: 5 -CTGAGTCCGCAGCAGGTG-3 ; Xbp1s reverse primer: 5 -TCTGAAGAGGCAACAGTGTCA-3 ; Ire1α forward primer: 5 -TGTTTGTCTC GACCCTGGATG-3 ; Ire1α reverse primer: 5 -CGTTGTTCTTGCCTCCAAGTG-3 ; Hgsnat forward primer: 5 -AGCGCTGATTACCAACCAGAA-3 ; Hgsnat reverse primer: 5 -AAACCATGGGAAGACGAGGTC-3 ; Pmp22 forward primer: 5 -TGGCAGAA CTGTACCACATCC-3 ; Pmp22 reverse primer: 5 -ACGCTG AAGATGACAGACAGG- 3 ; Id1 forward primer: 5 -TACGACATGAACGGC TGCTACTCA-3 ; Id1 reverse primer: 5 -TTACATGCTGCAGGATCTCCACCT-3 ; Id2 forward primer: 5 - TGAACGACTGCTACT CCAAGCTCA-3 ; Id2 reverse primer: 5 - GTGCTGCAGGATTTCCATCTTGGT-3 ; Id3 forward primer: 5 -ACAA GAGGAGCTTTTGCCACT-3 ; Id3 reverse primer: 5 -GAGGCGTTGA

105 90 GTTCAGGGTAA-3 ; Timp3 forward primer: 5 -GGGAAAGAAGCTGGTGAA GGA-3 ; Timp3 reverse primer: 5 -AGACTTTCAGAGGCTTCCGTG-3 ; Igfbp2 forward primer: 5 -GCGGGTACCTGTGAAAAGAGA-3 ; Igfbp2 reverse primer: 5 - ATTGACCTTCTCCCGGAACAC Western blot analysis Total cell lysates were made in RIPA lysis buffer (50mM Tris-HCl, ph 7.4, 150mM NaCl, 1% IGEPAL, 0.5% Sodium Deoxycholate, 0.1% SDS, 2mM EDTA, 1µg/mL Aprotinin, 1µg/mL Pepstatin, 1µg/mL Leupeptin, 5mM NaF, 1mM PMSF, 2mM β-glycerophosphate, 2mM Sodium orthovanadate ), rotated for 1 hr at 4 o C and centrifuged to clear cellular debris. Protein concentrations were determined using BCA protein assay (Pierce). 20µg of protein were separated on 8%, 10%, or 15% SDS- PAGE gels depending on antibody used and transferred to nitrocellulose using transblot TURBO transfer system (Bio-Rad) according to manufacturer s instructions. The primary antibodies used were from Cell Signaling (BiP, phospho-erk, ERK, phospho- MEK, MEK, IRE1α, phospho-perk, PERK, phospho-eif2α, eif2α, p15), Santa Cruz Biotechnology (p21 Ras, ID1, p16), Biolegend (XBP1S), and Abcam (ATF6). ACTIN (Millipore) was used as a loading control. Proteins were detected using ECL reagent (Pierce). 2.9 Phos-Tag SDS-PAGE Phos-Tag (Waco) SDS-PAGE was performed as described with slight modifications (6). Briefly, 5% Phos-tag copolymerized SDS-PAGE gels for IRE1α (Cell Signaling) were done with the following running conditions: 100 V for 3 hr using 25 µm

106 91 Phos-tag. The IRE1α blot was routinely reprobed with anti-vinculin (Cell Signaling) as a loading control In vitro RNA cleavage assays The cytoplasmic domain of human IRE1α was obtained from Sino Biologicals. Increasing amounts of human IRE1α was added to a reaction mixture containing total RNA purified from primary keratinocytes in a buffer containing 50 mm Tris (ph 7.5), 150 mm NaCl, 1 mm MgCl2, 1 mm MnCl2, 5 mm β-mercaptoethanol, 4 units RNasin and 2 mm ATP, as described previously (6). Recombinant human IRE1α boiled at 100ºC for 10 minutes was used as a negative control. After incubation at 37 C for 2 hr, the substrate RNAs were purified with an RNeasy kit (Qiagen) and reverse transcription performed to make cdna. Quantitative PCR was performed using primers flanking regions with or without potential cleavage sites for Id1 mrna. Xbp1 mrna was used as a positive and negative control for specificity of IRE1α cleavage reaction. The relative mrna level for each cleavage reaction was normalized with Gapdh (see RNA isolation and qpcr section for sequences). The primers used are as follows: Id1 (43-292) forward primer: 5 -CACTC TGTTCTCAGCCTCCTC-3 ; Id1 (43-292) reverse primer: 5 - GTGAGTAGCAGCCGTTC ATGT; Id1 ( ) forward primer: 5 -CAAAGTGAGCAA GGTGGAGAT-3 ; Id1 ( ) reverse primer: 5 -CAAACCCTCTACCCACTGGAC-3 ; Id1 ( ) forward primer: 5 -CCAGTGGGTAGAGGGTTTGAT-3 ; Id1( ) reverse primer: 5 - TTCC TCAGAAATCCGAGAAGC-3 ; Xbp1( ) forward primer: 5 AGTCCGCAGCAC TCAGACTAT-3 ; Xbp1( ) reverse primer: 5 -TGAAGAGG CAACAGTGTCAGA-3 ; Xbp1 ( ) forward primer: 5 -CTCCTGGGAGGATAC TTTTGC-3 ; Xbp1 ( ) reverse primer: 5 -CAATGTGATGGTCAGGGAAAG-3.

107 Luciferase reporter assays Primary keratinocytes were co-infected with v-ras Ha and non-target control shrna, IRE1α shrna, or XBP1 shrna for 2 days, selected with puromycin (1µg/mL) for 2 days and then transiently transfected with psbe4-luc and pcmv-renilla. Twenty four hours after transfection, cells were treated with TGFβ1 (1ng/mL) for 24 hrs. Cells were lysed with 1X passive lysis buffer (Promega) and luciferase activity measured with Dual-Luciferase Assay reagent (Promega) and a 96-well luminometer (Promega) DNA microarray analysis Primary keratinocytes were co-transduced with v-ras Ha and control, IRE1α, or XBP1 shrna for 2 days, selected with puromycin (1µg/mL) for 2 days, and harvested on day 5. At the same time, primary keratinocytes were transduced with control shrna, selected with puromycin (1µg/mL) for 2 days, and harvested on day 5. Total RNA was isolated from 3 biological replicates for each group with RiboZOL (Amresco) and purified with RNeasy Mini Kit (Qiagen). 100ng RNA was prepared, reverse transcribed and hybridized to GeneChip Mouse Gene ST 2.0 arrays (Affymetrix) in the Penn State Genomics Core Facility. Labeling of cdna, hybridization of arrays was done according to manufacturer s instructions and arrays were scanned using a GeneChip Scanner 30007G. Arrays were analyzed using ArrayStar 11 Software (DNASTAR, Madison WI) with RMA background correction and quantile normalization. Mean log2 signal was used to compare gene expression between groups and significantly different genesidentified using a 1.5-fold cut off and 10% FDR using the method of Benjamini and Hochberg (7). Hierarchal clustering and heat map were generated using Gene Cluster 3.0 and Treeview software (8). Functional annotation was done using DAVID

108 93 Bioinformatics Software (9, 10). Genes whose protein products are localized to the lysosome, integral to the plasma membrane or localized to extracellular regions are designated as genes encoding ER cargo; genes whose protein products are localized in either the ER or Golgi are designated genes encoding ER resident; and genes localized to the cytosol or nucleus are designated as Cytosol Cell surface biotinylation assay At end of treatment, cells were placed on ice and washed three times with icecold PBS (ph 8.0) to remove amine-containing media and proteins from cells. 3mL of 1mg/mL ice-cold Sulfo-NHS-SS-Biotin (Pierce, diluted in 1X PBS ph 8.0) to each dish and gently rock for 30 minutes at 4ºC. Sulfo-NHS-SS-Biotin does not penetrate plasma membrane thus only proteins on cell surface become labeled. Cells were washed three times with ice-cold 100mM Glycine + PBS ph 8.0 to quench and remove excess biotin reagent and byproducts and kept on ice. 0.5mL 1% RIPA lysis buffer plus protease and phosphastase inhibitors was added to each dish and lysates scraped and transferred to a microcentrifuge tube and rotated for 1 hr to solubilize proteins. Cell debris was pelleted and supernatant transferred to new tube and protein quantified using BCA protein assay kit (Pierce). 500ug of protein was immunoprecipitated using streptavidinconjugated agarose beads (Pierce) according to manufacturer s instructions and cell surface proteins against TβRII and ALK5 (Santa Cruz) examined by western blot. An extra dish without biotinylation was used as a negative control TGFβ1 receptor internalization assay V-RAS Ha keratinocytes were pretreated with Thapsigargin (5nM) for 24 hours. To examine TGFβ1 receptor internalization, cells were place on ice and washed three

109 94 times with ice-cold PBS ph 8.0 to cease active endocytosis of cell surface proteins. Cells were labeled with 3mL Sulfo-NHS-SS- Biotin (1mg/mL) made in ice-cold PBS ph 8.0 for 30 minutes at 4ºC with gentle rocking and washed three times with ice-cold 100mM glycine + PBS ph 8.0 to quench and remove excess biotin. Cells were placed back at 37ºC in prewarmed media containing TGFβ1 (1ng/mL) to re-initiate endocytosis and let go for 1 hour. At end of 1 hr incubation, cells were placed back at 4ºC on ice and washed three times with ice-cold PBS ph 7.4 and incubated two times with a 50mM Glutathione-reduced solution for 15 minutes each to strip any remaining biotin on cell surface. Glutathione is unable to penetrate plasma membrane hence internalized biotin labeled proteins are protected. After three more washes with ice-cold PBS ph 7.4 to remove any excess glutathione and byproducts cells were lysed with 1% RIPA lysis buffer with protease and phosphatase inhibitors, solubilized for 1 hour with rotating and cell debris pelleted. Supernatants were transferred to new microfuge tube and protein quantified using BCA protein assay kit (Pierce). 500ug of protein was immunoprecipitated using streptavidin-conjugated agarose beads (Pierce) according to manufacturer s instructions and internalized proteins against TβRII (Santa Cruz) was examined by western blot. As controls, one dish was included to show total biotinylated proteins and another that was stripped with glutathione to ensure complete removal of biotinylated proteins Statistical analysis Values are expressed as the mean ± SEM. Student s t test was used to compare the indicated groups. P-values of 0.05 were regarded as statistically significant.

110 Bibliography 1 Dlugosz,A.A., Glick,A.B., Tennenbaum,T., Weinberg,W.C. and Yuspa,S.H. Isolation and utilization of epidermal keratinocytes for oncogene research, Methods Enzymol., 254:3-20.: 3-20, Chin,L., Tam,A., Pomerantz,J., Wong,M., Holash,J., Bardeesy,N., Shen,Q., O'Hagan,R., Pantginis,J., Zhou,H., Horner,J.W., Cordon-Cardo,C., Yancopoulos,G.D. and DePinho,R.A. Essential role for oncogenic Ras in tumour maintenance, Nature, 400: , Roop,D.R., Lowy,D.R., Tambourin,P.E., Strickland,J., Harper,J.R., Balaschak,M., Spangler,E.F. and Yuspa,S.H. An activated Harvey ras oncogene produces benign tumours on mouse epidermal tissue, Nature, 323: , Yuspa,S.H. and Morgan,D.L. Mouse skin cells resistant to terminal differentiation associated with initiation of carcinogenesis, Nature (London), 293: 72-74, Bae,D.S., Blazanin,N., Licata,M., Lee,J. and Glick,A.B. Tumor suppressor and oncogene actions of TGFbeta1 occur early in skin carcinogenesis and are mediated by Smad3, Mol.Carcinog., 48: , Lee,A.H., Heidtman,K., Hotamisligil,G.S. and Glimcher,L.H. Dual and opposing roles of the unfolded protein response regulated by IRE1alpha and XBP1 in proinsulin processing and insulin secretion, Proc.Natl.Acad.Sci.U.S.A, 108: , Reiner,A., Yekutieli,D. and Benjamini,Y. Identifying differentially expressed genes using false discovery rate controlling procedures, Bioinformatics., 19: , Eisen,M.B., Spellman,P.T., Brown,P.O. and Botstein,D. Cluster analysis and display of genome-wide expression patterns, Proc.Natl.Acad.Sci.U.S.A., 95: , Bickel,P.J., Brown,J.B., Huang,H. and Li,Q. An overview of recent developments in genomics and associated statistical methods, Philos.Trans.A Math.Phys.Eng Sci., 367: , Huang,d.W., Sherman,B.T. and Lempicki,R.A. Systematic and integrative analysis of large gene lists using DAVID bioinformatics resources, Nat.Protoc., 4: 44-57, 2009.

111 Chapter 3: IRE1α regulates divergent RNase outputs that dictate senescence and malignant conversion induced by oncogenic HRAS Abstract IRE1α is a key player in the UPR and is a unique bifunctional kinase/endoribonuclease that regulates adaptive or destructive responses to ER stress. However, whether IRE1α regulates a similar divergent response at early stages of cancer development is currently unknown. Here we show for the first time that the two actions of the IRE1α endoribonuclease (RNase) regulate opposing responses to oncogene activation. Transduction of primary keratinocytes with oncogenic v-ras Ha causes rapid hyperproliferation followed by irreversible growth arrest and senescence, a potent barrier to tumor development. During the initial proliferative response, oncogenic v-ras Ha simultaneously activates IRE1α-mediated mrna cleavage and splicing of XBP1 mrna to produce the adaptive transcription factor XBP1S as well as promotes mrna endoribonucleolytic decay through RIDD. Utilization of shrna and pharmacological approaches implicates ER stress-dependent XBP1S as part of the proliferation response to oncogenic v-ras Ha, while RIDD mediates cell cycle growth arrest in the absence of XBP1. The requirement of XBP1S for proliferation is linked to suppression of v-ras Ha -induced senescence. In contrast, IRE1α promotes v-ras Ha - induced senescence in the absence of XBP1 and is required for suppression of malignant transformation. Gene expression profiling of v-ras Ha keratinocytes deficient in IRE1α identified a number of RIDD candidate mrna substrates that are linked to senescence and malignant transformation. Among those identified was the ID family, of which Id1 mrna was confirmed as a direct target of IRE1α endoribonucleolytic activity

112 97 using cell-free assays. Furthermore, shrna knockdown of Id1 mrna rescued the senescence phenotype in v-ras Ha keratinocytes with inactive IRE1α RNase activity. Together, these results suggest that the balance between IRE1α-XBP1S and IRE1α- RIDD determines whether a premalignant tumor cell undergoes proliferation and tumor progression or senescence and tumor suppression. 3.2 Introduction The endoplasmic reticulum (ER) is an intracellular organelle responsible for protein folding and assembly, lipid and sterol biosynthesis, and calcium storage (1, 2). Pathophysiological states that disrupt ER homeostasis by increased protein folding demand or altered protein folding create an imbalance between protein load and capacity of the ER, causing the accumulation of unfolded proteins and a condition termed ER stress (3). To adapt to ER stress, the ER has evolved the unfolded protein response (UPR) signaling pathway which is mediated by three major ER transmembrane proteins, including PKR-like ER kinase (PERK), inositol-requiring enzyme 1α (IRE1α), and activating transcription factor 6 (ATF6) (1). IRE1α is the most evolutionarily conserved UPR transducer that functions as a dual protein kinase and endoribonuclease (RNase). During ER stress IRE1α homodimerizes followed by transautophosphorylation of its kinase domain in the ER membrane, causing conformational changes which activate its RNase (4). IRE1α activated RNase causes the site-specific cleavage of the Xbp1 mrna. Religation and translation of XBP1 mrna at an alternate open reading frame encodes a potent transcription factor, XBP1S, which regulates the expression of genes involved in protein folding and

113 98 degradation leading to resolution of ER stress (4). IRE1α RNase activity can also promote endonucleolytic cleavage and degradation of mrnas through a process called Regulated IRE1 Dependent Decay (RIDD), which may serve to reduce protein load by targeting mrnas being co-translated on the ER membrane (4-6). However, recent studies indicate that RIDD plays a role in promoting cell death during conditions of chronic ER stress (7) either through 1) cleavage of several cytosolic micrornas that inhibit apoptotic machinery (8) or 2) cleavage of cell surface and secretory pathway mrnas that encode proteins that may have growth promoting roles (6). Thus, the emerging paradigm suggests that IRE1α can regulate differing cell fate responses through two distinct RNase outputs. In the mouse two-stage skin carcinogenesis model, treatment with the carcinogen DMBA causes mutations in the c-ha-ras gene, and repeated applications of the tumor promoter TPA causes papilloma outgrowths with a small percentage that eventually convert to malignant squamous cell carcinoma (SCC) (9). During two-stage skin carcinogenesis, overexpression, gene amplification, and homozygousity of the mutant HRAS allele occurs as tumors undergo malignant progression (10). In an in vitro model, transduction of primary mouse keratinocytes with an oncogenic v-ras Ha retrovirus initially causes hyperproliferation but this is followed by irreversible growth arrest and senescence (11). Oncogene-induced senescence is a potent tumor suppressor pathway that prevents malignant progression and is linked to the benign tumor phenotype in mouse cancer models and human cancer (12, 13). Oncogeneinduced senescence in primary cells triggered by HRAS is characterized by induction of tumor suppressors such as p53 and p16 (14) and inactivation of these proteins is linked

114 99 to malignant progression (15). Inactivation of other tumor suppressor pathways, such as TGFβ1 and PPARβ/δ also lead to escape from senescence and malignant progression (11, 16). Here, we examined ER stress and activation of the UPR in mouse keratinocytes expressing v-ras Ha. We found that only the IRE1α pathway was potently activated by v-ras Ha. In this chapter we show that the two outputs of the IRE1α RNase, Xbp1 mrna splicing and RIDD have opposing roles in the proliferative and senescence response to oncogenic v-ras Ha. Given that oncogenic RAS mutations are the initiating driver mutations in skin cancer and prevalent in multiple human cancers, these results have important implications for understanding the role of IRE1α in RAS-driven human cancer. 3.3 Results Oncogenic v-ras Ha activates IRE1α-mediated Xbp1 mrna splicing and RIDD during proliferation. To determine the effect of oncogenic RAS on ER stress and the UPR we transduced freshly isolated mouse neonatal keratinocytes with a high titer v-ras Ha retrovirus at an MOI of 1-3 (17). As expected, v-ras Ha keratinocytes displayed robust activation of MEK and ERK kinases at 2, 4, and 5 days after transduction as indicated by increased protein levels of phosphorylated MEK and ERK (Figure 3.1A). In agreement with previous studies (11), v-ras Ha caused rapid cell proliferation as early

115 100 Figure 3.1. Oncogenic v-ras Ha promotes ER stress during proliferation. Primary and v-ras Ha keratinocytes were compared after 2, 4, and 5 days after transduction. (A) Western blot analysis of p21 Ras, p-mek, MEK, p-erk, ERK and ACTIN as a loading control. (B) Quantification of percent BrdU positive cells at each timepoint. (C) Western blot analysis of the ER chaperone BiP and ACTIN as a loading control. (D) Mean fluorescence intensity of primary and v-ras Ha keratinocytes stained with ER-Tracker Green and divided by the forward scatter to normalize for cell size at each timepoint as determined by flow cytometry. Values = mean ± SEM. *indicates significantly different values (P 0.05) from primary keratinocytes without v-ras Ha.

116 101 as 4 days after transduction and this was maximal by 5 days as determined by pulsing cells with BrdU and in situ immunohistochemical analysis (Figure 3.1B). At 5 days after transduction approximately 50% of the v-ras Ha keratinocytes were BrdU positive while only 10% were positive in mock-transduced primary keratinocytes. To determine if v- RAS Ha keratinocytes undergoing cell proliferation exhibited ER stress, we examined protein levels of the ER chaperone BiP as well as expansion of the ER, two well accepted markers of ER stress (1). In primary keratinocytes BiP protein levels were undetectable at all time points, while BiP protein levels increased 4 days after v-ras Ha transduction correlating with the onset of cell proliferation (Figure 3.1C). Consistent with increased BiP levels, there was approximately a 2-fold on day 2 and a 4-fold increase on day 5 in ER content, respectively, after v-ras Ha transduction as determined by flow cytometry analysis using ER-Tracker Green ( Figure 3.1D). Since the mean fluorescence intensity (MFI) of ER tracker staining was normalized to cell size based on forward scatter properties of primary and v-ras Ha keratinocytes, this increase was likely an active response to v-ras Ha transduction. Taken together, transduction of primary keratinocytes with v-ras Ha causes ER stress. In unstressed cells, BiP associates with and maintains PERK, ATF6, and IRE1α in an inactive state but under ER stress dissociates from these proteins and binds to misfolded proteins leading to UPR activation (18, 19). However, we did not observe an increase in phosphorylated PERK or total PERK levels, or phosphorylation of the PERK downstream target eif2α in v-ras Ha keratinocytes or increased levels of cleaved ATF6 (Figure 3.2A). To examine IRE1α phosphorylation, we initially used a phospho-specific antibody that detects phosphorylation at serine 724 within the activation loop of IRE1α.

117 102 Figure 3.2. Oncogenic v-ras Ha activates IRE1α but not PERK or ATF6. Primary and v-ras Ha keratinocytes were examined after 2, 4, and 5 days after transduction. (A) Western blot analysis of cleaved ATF6 and PERK-eIF2α UPR signaling pathways. (B) Western blot analysis of phosphorylated and total IRE1α. Horizontal bar denotes Phos-Tag reagent was used for SDS-PAGE. ACTIN and VINCULIN were used as loading controls (C) λppase treatment of day 5 whole cell protein lysates and western blot analysis of phosphorylated and total IRE1α (D) Q-PCR of Xbp1s mrna levels at each timepoint. (D) Western blot analysis of XBP1S at each timepoint. Values = mean ± SEM. *indicates significantly different vales (P 0.05) from primary keratinocytes without v-ras Ha.

118 103 However, we had difficulty detecting differences in IRE1α phosphorylation when comparing primary keratinocytes to those transduced with v-ras Ha. Furthermore, when we depleted IRE1α protein levels using shrna no effect in IRE1α phosphorylation was observed at this residue (data not shown). However, we detected increased IRE1α phosphorylation in v-ras Ha keratinocytes when compared to primary keratinocytes using a Phos-Tag-based approach (Figure 3.2B). Phos-Tag is a functional molecule that binds specifically to phosphorylated ions when in the presence of manganese or zinc ions (20). When Phos-Tag is copolymerized with SDS-PAGE gels, phosphorylated proteins migrate slower compared to the unphosphorylated forms and can be separated. Furthermore, this reagent is suitable for detection of physiological levels of IRE1α phosphorylation under ER stress (21). IRE1α phosphorylation was not detected when western blots were overexposed using gels without Phos-Tag (Figure 3.2B). Furthermore, lambda phosphatase treatment of cell lysates removed the phosphorylated form of IRE1α when separated on Phos-Tag gels (Figure 3.2C). In addition to an increase in IRE1α phosphorylation this was accompanied by an increase in total IRE1α protein levels at each timepoint examined (Figure 3.2B). ATF6 and IRE1α mediate the induction of Xbp1 mrna as well as processing into an active transcription factor, respectively (22). In agreement with lack of ATF6 activation, unspliced Xbp1 mrna remained unchanged (data not shown). Rather, there was a significant increase in Xbp1 mrna splicing as detected by quantitative RT-PCR as well as XBP1S protein levels at each timepoint examined, specifically linking IRE1α activation to the adaptive functions of the UPR in v-ras Ha keratinocytes undergoing proliferation (Fig. 3.2D and E).

119 104 In addition to Xbp1 mrna splicing, IRE1α also promotes mrna cleavage and degradation through Regulated IRE1 Dependent Decay (RIDD) during both mild and chronic ER stress (5, 23). To test if RIDD is activated in v-ras Ha keratinocytes we examined mrna levels of Hgsnat and Pmp22, which are ER-localized mrnas that have been previously characterized as IRE1α-RIDD cleavage substrates (5). HGSNAT is a membrane bound lysosomal transferase involved in the degradation of heparin sulfate and PMP22 is an ECM protein and major component of myelin in the peripheral nervous system (5). When compared to mock transduced primary keratinocytes, oncogenic v-ras Ha rapidly downregulated Hgsnat and Pmp22 mrna levels by 2 days after transduction and this was sustained up to 5 days (Figure 3.3A). This is consistent with increased IRE1α activation and RNase activity observed at each timepoint. To implicate IRE1α directly in regulating Hgsnat and Pmp22 mrna levels, we treated v- RAS Ha keratinocytes with the IRE1α RNase inhibitor 4µ8C which has previously been demonstrated to block Xbp1 mrna splicing and RIDD (24). 4µ8C is a specific coumarin based inhibitor that forms a stable Schiff base within lysine 907 of the IRE1α RNase domain. This prevents mrna substrate access and inhibits RNase activity without altering the IRE1α kinase (24). Preliminary dose-response experiments determined that treatment with 25µM 4µ8C for 24 hours in v-ras Ha keratinocytes as a suitable dose to significantly inhibit Xbp1 mrna splicing and XBP1S protein levels without any overt cytotoxicity (Figure 3.2B and data not shown). Importantly, treatment with 25µM 4µ8C also blocked mrna downregulation of Hgsnat and Pmp22 after 24 hours in v-ras Ha keratinocytes to levels close to that observed in primary keratinocytes (Figure 3.3C). This suggests that these RIDD targets are predominately dependent on

120 105 Figure 3.3. Activation of IRE1α-mediated RIDD in v-ras Ha keratinocytes. (A) Q-PCR of Hgsnat and Pmp22 mrna levels 2, 4, and 5 days after v-ras Ha transduction. (B) Q-PCR (left) of Xbp1s mrna levels and western blot analysis (right) of XBP1S protein levels in v- RAS Ha keratinocytes treated with 4µ8C (25µM) for 24 hours. (C) Q-PCR of Hgsnat and Pmp22 mrna levels in v-ras Ha keratinocytes treated with 4µ8C (25µM) for 24 hours. (D) Western blot analysis of IRE1α and XBP1S protein levels after co-transduction of v-ras Ha and lentiviral shrna against IRE1α or XBP1 after 5 days. ACTIN was used as a loading control. (D) Q-PCR of Hgsnat and Pmp22 mrna levels in v-ras Ha keratinocytes deficient in IRE1α or XBP1 after 5 days. Values = mean ± SEM. *indicates significantly different values (P 0.05). All expression values were normalized to primary keratinocytes without v-ras Ha and represent fold-change.

121 IRE1α RNase activation for mrna downregulation by v-ras Ha. We next determined if IRE1α functionally degrades RIDD targets in the absence of XBP1. We co-transduced 106 v-ras Ha keratinocytes with lentivirus containing shrna against either a non-target control, IRE1α or XBP1 at an MOI of 20 to ensure close to 100% survival when selected with puromycin (see Materials and Methods), and examined RIDD substrate mrna levels 5 days after transduction. IRE1α shrna knockdown significantly reduced IRE1α and XBP1S protein levels to nearly undetectable levels when compared to v-ras Ha keratinocytes infected with a non-targeting control shrna (Figure 3.3D). In contrast, XBP1 knockdown reduced XBP1S protein levels similar to that of IRE1α knockdown but IRE1α protein levels remained intact (Figure 3.3D). As expected, IRE1α depletion blocked Hgsnat and Pmp22 mrna downregulation by v-ras Ha similar to 4µ8C (Figure 3.3E). However and more importantly, this inhibition was solely dependent on IRE1α RNase activity separate from Xbp1 mrna splicing as XBP1 depletion did not block downregulation of Hgsnat and Pmp22 mrna levels (Figure 3.3F). As expected, this indicates that IRE1α was still active in the absence of XBP1. Taken together, these results indicate that hyperproliferation of v-ras Ha keratinocytes correlates with ER stress and selective activation of IRE1α but not PERK or ATF6. Furthermore, both IRE1α RNase functions, Xbp1 mrna splicing and RIDD, are activated in response to v- RAS Ha in primary keratinocytes ER stress dependent activation of IRE1α by MEK-ERK signaling It is not clear if v-ras Ha -induced ER stress regulates IRE1α RNase activation or if activation is due to IRE1α overexpression. Malfolded proteins in the ER promote oligomerization of IRE1α luminal domains, which in turn causes IRE1α to

122 107 transautophosphorylate, leading to RNase activation (7, 25). However, IRE1α oligomerization can also be driven through mass action on the ER membrane independent of ER stress (26). Acute overexpression of IRE1α causes the ER luminal domains to spontaneously undergo oligomerization, followed by transautophosphorylation and subsequent RNase activation (6, 26).To determine whether IRE1α RNase activation is ER stress dependent or independent, primary and v- RAS Ha keratinocytes were exposed to increasing doses of 4-phenyl butyrate (4-PBA) for 24 hours. 4-PBA is a low-molecular weight chemical chaperone that has been shown to increase ER function and reduce misfolded/aggregated proteins in the ER lumen and, in turn, ameliorate ER stress (27). 4-PBA treatment of v-ras Ha keratinocytes led to a dose dependent decrease in IRE1α phosphorylation and this correlated with similar decreases in BiP and XBP1S protein levels (Figure 3.4A). Similarly, downregulation of Hgsnat and Pmp22 mrna levels was inhibited with 4-PBA indicating both arms of IRE1α are deactivated (Figure 3.4B). Interestingly, despite complete inhibition of IRE1α activation at high 4-PBA doses (5mM), induction of total IRE1α levels by v-ras Ha remained unaffected. Similarly, 4-PBA treatment did not affect total RAS levels or activation of its downstream effector pathway, MEK-ERK (Figure 3.4A). Activation of the MEK-ERK pathway by oncogenic RAS is central to many downstream biological processes including cell proliferation and survival (28). Furthermore, a previous study demonstrated that MEK-ERK signaling was required for adaptation to ER stress and that IRE1α-mediated Xbp1 mrna splicing regulated proliferation of melanoma cells expressing mutant BRAF V600E (29). To determine whether v-ras Ha -induced IRE1α activation is similarly MEK-ERK dependent, we treated

123 108 Figure 3.4. Oncogenic v-ras Ha -induced ER stress and MEK-ERK signaling regulates IRE1α activation (A) Inhibition of ER stress in primary and v-ras Ha keratinocytes treated on day 4 after transduction with increasing doses of 4-PBA for 24 hours. Western blot analysis of phosphorylated IRE1α, total IRE1α, BiP, XBP1S, p21 Ras, p-erk, and ERK were examined. ACTIN and VINCULIN were used as loading controls. (B) Q-PCR of Hgsnat and Pmp22 mrna levels in primary and v-ras Ha keratinocytes treated on day 4 with 4-PBA (2.5mM) for 24 hours. (C) Primary or v-ras Ha keratinocytes were treated on day 2 after transduction with U0126 (5uM) for 48 hours and IRE1α activation determined by western blot analysis. (D) Q-PCR of Ire1α and Xbp1s mrna levels in primary and v-ras Ha keratinocytes similarly treated as in (C). (E) Q-PCR of Hgsnat and Pmp22 mrna levels similarly treated as in (C). Values = mean ± SEM. *indicates significantly different values (P 0.05). All expression values were normalized to primary keratinocytes and represent foldchange.

124 109 primary and v-ras Ha keratinocytes with the MEK kinase inhibitor U0126 and examined IRE1α signaling after 48 hours. As expected, U0126 treatment potently blocked ERK phosphorylation (Figure 3.4C) and resulted in decreased cell proliferation with no noticeable cytotoxicity (data not shown). U0126 treatment reduced Ire1α mrna levels as well as total IRE1α protein levels (Figure 3.4C, D) and this also resulted in a significant decrease in Xbp1 mrna splicing and BiP protein levels (Figure 3.4C, D). Furthermore, downregulation of Hgsnat and Pmp22 mrna levels was inhibited by U0126 in v-ras Ha keratinocytes (Figure 3.4E). There was no effect on IRE1α RNase function in primary keratinocytes treated with U0126 further confirming this as a specific effect to v-ras Ha keratinocytes. Collectively, these data suggest that IRE1α activation is ER stress dependent and IRE1α expression is driven by MEK-ERK signaling Opposing roles of IRE1α and XBP1 during proliferation and senescence 4-PBA treatment of v-ras Ha keratinocytes caused a dose-dependent decrease in proliferation despite intact MEK-ERK signaling (Figure 3.4A and data not shown). However, it is not clear if this effect on cell proliferation was due to inhibition of ER stress or specific inhibition of IRE1α RNase activity. We treated v-ras Ha keratinocytes 2 days after transduction with 4µ8C and examined BrdU incorporation after 24 and 48 hours. 4µ8C treatment significantly reduced v-ras Ha -induced cell proliferation at both timepoints indicating that intact IRE1α RNase activity is required for maximal proliferative response to v-ras Ha (Figure 3.5A). Since 4µ8C suppresses both Xbp1 mrna splicing and RIDD, we generated a lentivirus expressing mouse XBP1S (See Methods section) to determine the role of this arm of the IRE1α RNase in v-ras Ha - induced proliferation. Increasing MOI of empty and XBP1S lentivirus were co-

125 110 Figure 3.5. IRE1α RNase activity promotes growth arrest in the absence of XBP1 in v-ras Ha keratinocytes. (A) Primary and v-ras Ha keratinocytes were treated 2 days after transduction with 4µ8C (25µM) for 24 and 48 hours and BrdU positive cells quantified. (B) v-ras Ha keratinocytes were co-transduced with increasing amounts (MOI 2, 5, 10) of Empty or XBP1S lentivirus and XBP1S protein levels examined 4 days after transduction by western blot analysis. (C) v-ras Ha keratinocytes were transduced as in (B) and BrdU positive cells quantified on 4 days after transduction. (D) v-ras Ha keratinocytes were co-transduced with control, IRE1α or XBP1 lentivirus and 2 days after transduction treated with 4µ8C (25µM) for 72 hours. BrdU positive cells were then quantified. Values = mean ± SEM. *indicates significantly different values (P 0.05). ns = not significant.

126 transduced with v-ras Ha in primary keratinocytes and XBP1S protein levels and cell proliferation measured 4 days after transduction by BrdU incorporation and 111 immunohistochemical analysis. Nearly 100% of v-ras Ha keratinocytes were GFPpositive with increasing MOI (data not shown) and increasing protein levels of XBP1S confirmed by western blot analysis (Figure 3.5B). Furthermore, XBP1S overexpression in v-ras Ha keratinocytes increased cell proliferation in a dose-dependent manner (Figure 3.5C). Several studies have demonstrated opposing roles of XBP1 and IRE1α- RIDD in physiological and pathological settings. For example, mice deficient in XBP1 are protected against acetaminophen (APAP)-induced liver toxicity. This is due to IRE1α activation and mrna decay of Cyp1a2 and Cyp2e1, two enzymes responsible for conversion of APAP into hepatotoxic metabolites (30). Similarly, another study demonstrated that XBP1-deficient mice caused hypolipidemia, indicating an important role in lipid metabolism, and that silencing of IRE1α reversed this phenotype. Moreover, IRE1α mediated hypolipidemia through mrna decay of enzymes involved in lipogenesis and lipoprotein metabolism (31). To determine if IRE1α and XBP1 display a similar opposing response to cell proliferation, we compared v-ras Ha keratinocytes cotransduced with control, IRE1α and XBP1 shrna for 2 days and then treated with 4µ8C for 72 hours. As expected, on day 5 after v-ras Ha transduction IRE1α and XBP1- depleted cells exhibited a decrease in cell proliferation to similar levels (Figure 3.5D). Furthermore, 4u8C treatment of control and IRE1α-depleted v-ras Ha keratinocytes exhibited a cooperative decrease in cell proliferation (Figure 3.5D). In contrast, when XBP1-depleted v-ras Ha keratinocytes were treated with 4µ8C, there was no decrease in cell proliferation when compared to control and IRE1α-depleted v-ras Ha

127 keratinocytes (Figure 3.5D). However, although not statistically significant, 4µ8C 112 treatment in XBP1-depleted v-ras Ha keratinocytes increased the number of BrdUpositive cells to levels comparable to control v-ras Ha keratinocytes, suggesting that IRE1α RNase activity in the absence of XBP1 functions to inhibit cell proliferation. Next, we examined the kinetics of IRE1α-mediated Xbp1 mrna splicing and RIDD during v- RAS Ha -induced senescence. Primary keratinocytes transduced with v-ras Ha undergo senescence after about 8-10 days in culture (11). In agreement with this, we observed increased senescence-associated β galactosidase (SA-β-Gal) by 11 days as well as a concomitant decrease in the percentage of BrdU positive cells (Figure 3.6A, C). V- RAS Ha -induced senescence was also associated with extensive cytoplasmic vacuolization (Figure 3.6B), a response that is associated with HRAS-induced senescence in keratinocytes and melanocytes (32). As expected, v-ras Ha -induced senescence was associated with a further increase in p-erk, p-p38 activation, as well as increases in CDKi senescence markers p15 and p16 (Figure 3.6D). IRE1α phosphorylation transiently increased at day 8 and 11 after v-ras Ha transduction when compared to day 5 but returned to similar levels by day 14 (Figure 3.6D). Moreover, there was an increase in total IRE1α protein levels when compared to v-ras Ha keratinocytes on day 5. Surprisingly, despite an increase in IRE1α phosphorylation and total IRE1α protein levels during v-ras Ha -induced senescence, Xbp1 mrna splicing and XBP1S protein levels continually decreased between 5 to 14 days after v-ras Ha transduction ( Figure 3.6D,E). Furthermore, the change in Xbp1 mrna splicing was not due to alterations in unspliced Xbp1 mrna (Figure 3.6E). Similarly, there was a decrease in BiP protein levels indicating that, in conjunction with reduced spliced Xbp1

128 113 Figure 3.6. Divergent IRE1α RNase outputs is associated with v-ras Ha -induced senescence. V-RAS Ha keratinocytes were examined at 5, 8, 11, and 14 days after transduction. (A) Quantification of percent SA-β-Gal positive cells, (B) percent vacuolated positive cells, and (C) percent BrdU positive cells at each timepoint. (D) Western blot analysis of phosphorylated IRE1α, total IRE1α, BiP, XBP1S, p21 Ras, p-erk, total ERK, p-p38, total p38, p15 and p16. VINCULIN and ACTIN were used as loading controls. (E) Q-PCR of Xbp1u and Xbp1s mrna levels at each timepoint. (F) Q-PCR of Hgsnat and Pmp22 mrna levels at each timepoint, as indicated. Values = mean ± SEM. *indicates significantly different values (P 0.05) from v-ras Ha values on day 5. All expression values were normalized to primary keratinocytes without v-ras Ha on day 5 and represent fold-change.

129 114 mrna and XBP1S protein levels, suggests ER stress is reduced during v-ras Ha - induced senescence (Figure 3.6D). In contrast, downregulation of Pmp22 and Hgsnat mrna levels was enhanced during v-ras Ha -induced senescence in agreement with increased IRE1α phosphorylation and total IRE1α protein levels (Figure 3.6F). This evidence suggests that both decreased Xbp1 mrna splicing and increased RIDD are associated with induction of senescence. To investigate a divergent role for both Xbp1 mrna splicing and RIDD during senescence, we depleted IRE1α and XBP1 using lentiviral shrna in v-ras Ha keratinocytes. IRE1α depletion significantly inhibited senescence and cellular vacuolization and this was associated with a higher number of BrdU positive cells when compare to control v-ras Ha keratinocytes (Figure 3.7A-D). Furthermore, IRE1α depleted v-ras Ha keratinocytes did not exhibit characteristic cell flattening associated with senescence and growth arrest (Figure 3.7A). In contrast, as early as 5 days after v-ras Ha transduction, XBP1 depletion caused a significant increase in the percentage of SA-β-Gal positive cells and cytoplasmic vacuolization, and decrease in cell proliferation. Taken together, these results suggest IRE1α RNase functions act opposite of one another to regulate v-ras Ha -induced proliferation and senescence XBP1 depletion causes hyperactive IRE1α and MEK-ERK signaling to promote senescence in v-ras Ha keratinocytes To define the differential senescence response between IRE1 and XBP1, we examined several classical markers of oncogene-induced senescence (33), including hyperactivation of RAS and ERK, activation of the stress kinase p38, and tumor suppressors p15 and p16 (34, 35). Consistent with a role in promoting senescence

130 115 Figure 3.7. Opposing roles of IRE1α and XBP1 during v-ras Ha -induced senescence. Primary keratinocytes were transduced with v-ras Ha or control, IRE1α, and XBP1 shrna and examined after 5, 8, 11, and 14 days after transduction(a) Bright field (Top) and SA-β-Gal (Bottom) images of v-ras Ha keratinocytes 11 days after transduction of indicated groups. Red arrows denotes cells that are vacuolated or positive for SA-β-Gal. (B) Quantification of percent SA-β-Gal positive cells (C) percent vacuolated positive cells (D) and percent BrdU positive cells at each time point. Values = mean ± SEM. *indicates significantly different values (P 0.05) from v-ras Ha values on day 5.

131 116 (Figure 3.7A-D), IRE1α depletion in v-ras Ha keratinocytes dampened hyperactive RAS and phosphorylated ERK protein levels and completely abolished phosphorylation of the stress kinase p38 (Figure 3.8A). However, IRE1α depletion did not significantly effect p15 and p16 protein levels (Figure 3.8A). Surprisingly, there was also a decrease in BiP protein levels in v-ras Ha keratinocytes deficient in IRE1α. This is in contrast to senescence of control v-ras Ha keratinocytes where IRE1α protein levels increase and BiP protein levels decrease (Figure 3.6A). In contrast to IRE1α depletion, XBP1 depletion in v-ras Ha keratinocytes caused pronounced and rapid activation of some of the senescence markers examined including phosphorylation of ERK and p38 (Figure 3.8A). Interestingly, XBP1-depletion increased both phosphorylated and total IRE1α protein levels suggesting that XBP1 negatively regulates IRE1α activation and expression. This observation is consistent with other model systems using XBP1- deficient mice in which XBP1 deficiency leads to hyperactive IRE1α (31). Previous studies have shown that blockade of MEK signaling using dominant negative or pharmacological approaches inhibits the senescence response induced by oncogenic RAS (36). In agreement, XBP1-depleted v-ras Ha keratinocytes treated with U0126 significantly inhibits senescence (Figure 3.8B). This indicates that MEK-ERK signaling mediates senescence in v-ras Ha keratinocytes deficient in XBP1. Furthermore, inhibition of MEK-ERK signaling in XBP1-depleted v-ras Ha keratinocytes with U0126 inhibited IRE1α phosphoarylation and total IRE1α protein levels (Figure 3.8C). Therefore, we next reasoned that IRE1α RNase activity mediates senescence induced by XBP1 depletion in v-ras Ha keratinocytes. Indeed, treatment of XBP1-depleted v- RAS Ha keratinocytes with 4µ8C for 3 days inhibited senescence as well as cellular

132 117 Figure 3.8. XBP1 depletion causes hyperactive IRE1α and MEK-ERK signaling to promote senescence. (A) Western blot analysis of p-ire1α, BiP, p21 RAS, p-erk, p-p38, p15, p16 and total levels of v-ras Ha keratinocytes transduced with control, IRE1α, or XBP1 shrna and examined at indicated timepoints. ACTIN and VINCULIN were used as loading controls. (B) Quantification of percent SA-β-gal positive cells of v-ras Ha keratinocytes transduced with control, IRE1αα, or XBP1 shrna and treated with U0126 (5µM) on day 2 after transduction and examined on day 5. (C) Western blot analysis of p-ire1α, IRE1α, p- ERK, and ERK in XBP1-depleted v-ras Ha keratinocytes treated with U0126 (5µM) on day 2 after transduction and examined on day 8 and 11. ACTIN and VINCULIN were used as loading controls. (D) Quantification of percent SA-β-Gal positive cells in v-ras Ha keratinocytes transduced with control, IRE1α, and XBP1 shrna treated with 4µ8C (25uM) and examined on day 5. (E) Photomicrographs of Bright field (Left) and SA-β-Gal (Right) staining of v-ras Ha keratinocytes co-transduced with XBP1 shrna, treated 4µ8C(25µM). Red arrows indicate positive cells. (F) Western blot analysis of p21 Ras, p-p38, p38 and ACTIN of XBP1-depleted v-ras Ha keratinocytes treated with 4µ8C (25µM). Values = mean ± SEM. *indicates significantly different values (P 0.05) from v-ras Ha values on day 5.

133 vacuolization while little effect was observed in control and IRE1α depleted cells (Figure D, E). Moreover, inhibition of IRE1α RNase activity with 4µ8C prevented the increase in RAS and phosphorylated p38 protein levels in XBP1-depleted cells consistent with inhibition of senescence (Figure 3.8F). Collectively, these results provide the first evidence that IRE1α and XBP1 can have opposing responses to v-ras Ha - induced senescence Expression profiling and identification of IRE1α-RIDD substrates linked to v- RAS Ha -induced senescence and suppression of malignant conversion Our data suggests that IRE1α RNase activity independent of Xbp1 mrna splicing promotes v-ras Ha -induced senescence. Thus, we sought to identify RIDD substrates critical to the senescence response in v-ras Ha keratinocytes by using a microarray-based approach. We co-transduced primary keratinocytes with v-ras Ha and non-target control, IRE1α, or XBP1 shrna, selected with puromycin and purified total RNA after 5 days, amplified and labeled these samples, and hybridized to Mouse Gene 2.0 ST arrays. Primary keratinocytes transduced with non-target shrna was used as a control (see Materials and Methods). To select the most likely RIDD candidates important in the senescence response, the following data filtering criteria were used: 1) Genes whose expression was significantly downregulated by v-ras Ha 2) and were significantly induced by IRE1α-depletion when compared to control v-ras Ha keratinocytes 3) and exhibited no difference/enhanced downregulation by XBP1 depletion when compared to control v-ras Ha keratinocytes. This was to account for the IRE1α hyperactivation observed in XBP1 depleted cells. Using this approach, we identified a total of 71 genes that fit these criteria and represent potential RIDD mrna

134 119 Figure 3.9. Identification of IRE1α-RIDD candidate genes linked to senescence and malignant conversion. (A) Hierarchical clustering analysis of gene expression changes from DNA microarrays. Genes whose expression is 1.5 fold or more increased in IRE1α depleted v-ras Ha keratinocytes compared to control cells and no different/downregulated in XBP1-depleted v- RAS Ha keratinocytes compared to control cells on day 5 after transduction were considered RIDD candidates. Gene expression changes were normalized to primary keratinocytes transduced with control shrna. (B) Disease analysis of significant RIDD candidates altered by IRE1α. (C) Functional role of RIDD candidates during cancer progression. (D) In vitro malignant conversion assay of v-ras Ha keratinocytes co-transduced with control, IRE1α, and XBP1 shrna. Values = mean ± SEM. *indicates significantly different values (P 0.05)

135 120 substrates for IRE1α in v-ras Ha keratinocytes (Figure 3.9A and Table 3.1). Using the annotation database DAVID (37), we find that 49.2% of the mrnas identified are predicted to encode secretory pathway proteins. As expected, many of these mrnas are predicted or are localized on the ER membrane in close proximity to IRE1α as they are translated on ER-bound ribosomes and enter the ER lumen for folding and maturation. These secretory pathway proteins include those that are secreted into the extracellular matrix and include the matrix metalloproteinase inhibitor Timp3, the potent vasoconstrictor Endothelin 1, the disintegrin Adamts1, the mitogen and cell adhesion protein Ctgf, the binding protein Igfbp2 and Bmp4, a growth factor that mediates differentiation. Endothelin 1, for instance, is induced in IRE1α-depleted v-ras Ha keratinocytes with a log2 ratio of when compared to in control v-ras Ha keratinocytes, while in XBP1 depleted v-ras Ha keratinocytes the log2 ratio was (Table 3.1). In addition, 45% of those RIDD mrna substrates identified were predicted as putative cytosolic mrnas that encode cytoplasmic or nuclear proteins. These include the zinc finger transcription factor Plagl1, the GTPase activating protein Dlc1, and Calponin 1, a filament-associated protein implicating in regulated smooth muscle contraction. Next, we reasoned that since inactivation of IRE1α leads to inhibition of v- RAS Ha -induced senescence, RIDD candidate genes may facilitate cancer progression. Indeed, INGENUITY pathway analysis revealed that 38 of the 71 RIDD candidate genes identified were strongly associated with cancer (Figure 3.9B). Moreover, approximately 63% had positive roles in cell growth and proliferation, 24% in metastasis, and 7% in invasive processes (Figure 3.9C). These results are consistent with the role of IRE1α promoting RIDD to facilitate senescence and growth arrest and that this arm of the

136 121

137 122

138 123

139 124 IRE1α RNase may be disabled to promote malignant progression. To investigate a role of IRE1a-RIDD during malignant progression, we performed an in vitro malignant conversion assay comparing v-ras Ha keratinocytes lacking IRE1α or XBP1. Here, v- RAS Ha keratinocytes transduced with control, IRE1α, or XBP1 shrna were cultured in low Ca ++ medium for 14 days and then switched to high Ca ++ medium to trigger terminal differentiation and maintained in this medium for 6 weeks. Colonies that are resistant to Ca ++ -induced differentiation form malignant SCC when grafted onto athymic mice and recapitulate many of the features associated with SCC generated by two-stage chemical carcinogenesis (38). As expected, there was approximately a 20-fold increase in the number of calcium-resistant foci in IRE1α-depleted v-ras Ha keratinocytes when compared to control cells (Figure 3.9C). Furthermore, these colonies were highly proliferative (data not shown). In contrast, no significant increase in the number of Ca ++ resistant foci was observed in v-ras Ha keratinocytes lacking XBP1. Further analysis of the list of potential RIDD mrna substrates identified by gene expression profiling, we were surprised to find that IRE1α regulates mrna expression of three members of the Id family, Id1, Id2, and Id3 (Table 3.1). ID proteins belong to the helix-loop-helix (HLH) family of transcription factors. Unlike most members of this family, ID proteins lack a DNA binding domain but instead associate with other family members and prevent their DNA binding thereby acting as a negative regulator of transcription (39). ID proteins contribute to tumorigenesis by inhibiting differentiation, stimulating proliferation, and regulating cell fate determination (39). Moreover, ID proteins have a role in negatively regulating RAS-induced senescence as well as cooperating with RAS to promote malignancy and metastasis (40, 41). Using

140 125 quantitative RT-PCR we confirmed increased mrna levels of Id1, Id2, and Id3 levels in IRE1α-depleted v-ras Ha keratinocytes at day 5 and day 11 after transduction when compared with control cells (Figure 3.10A). In contrast, decreased mrna levels were observed in XBP1-depleted v-ras Ha keratinocytes at similar time points highlighting the divergent regulation of Id mrnas by IRE1α and XBP1 in these cells. We also validated mrna levels of two secretory pathway proteins, Igfbp2 and Timp3 (Figure 3.10B). Consistent with the changes in Id1 mrna levels, ID1 protein levels were similarly increased or decreased in IRE1α or XBP1-depleted v-ras Ha keratinocytes at day 5 and day 11, respectively (Figure 3.10C). Next, we treated doxycycline-inducible bitransgenic keratinocytes that express human RAS V12G with 1µg/mL doxycycline and 25µM 4µ8C for 48 hours (Figure 3.10D). We confirmed downregulation of Id1 and Igfbp2 mrna levels is mediated by the IRE1α RNase as well as ID1 protein levels (Figure 3.10E-F). This indicates that IRE1α has similar functions in response to a human RAS oncogene Id1 mrna is a direct IRE1α-RIDD cleavage substrate We next chose to further characterize Id1 mrna as an IRE1α cleavage target given its critical role in senescence and malignant progression. Although Id1 is a cytosolic mrna evidence indicates that mrnas that encode soluble proteins can be targeted to IRE1α for degradation. For example, unspliced Xbp1 mrna is localized in the cytosol but is targeted by IRE1α for cleavage and splicing to generate XBP1S protein. Moreover, while it is generally accepted that protein synthesis is compartmentalized, fractionation studies examining mrnas bound to free and ERbound ribosomes have shown that a broad representation of mrnas encoding soluble

141 126 Figure Validation of IRE1α-RIDD candidate genes. (A) Q-PCR of Id1, Id2, and Id3 mrna levels in v-ras Ha keratinocytes transduced with control, IRE1α, and XBP1 shrna and examined on day 5 and day 11 after transduction. (B) QPCR of Igfbp2 and Timp3 mrna levels similar as in (A). (C) Western blot analysis of ID1 protein levels at similar timepoints as in (A). (D) Schematic of K5rTA x tetoras V12G doxycycline-inducible system to express human RAS V12G in primary keratinocytes. (E) Human RAS V12G was induced in primary keratinocytes by addition of doxycycline (1µg/mL) in the presence of 4µ8C(25µM) for 48 hours. Q-PCR of Id1 and Igfbp2 mrna levels was examined and expression normalized to primary keratinocytes lacking K5rTA in the presence of doxycycline. (F) Western blot analysis of XBP1S and ID1 in human RAS V12G keratinocytes treated similarly as in (E). Values = mean ± SEM. *indicates significantly different values (P 0.05) from respective controls.

142 127 proteins are localized on the ER. Furthermore, these ER-bound ribosomes actively engage mrnas encoding soluble proteins for protein synthesis (42). Indeed, although not proven in our studies Id1 mrna was identified as localized to the ER in one fractionation study (43). To determine if Id1 mrna is an IRE1α mrna substrate, we treated primary keratinocytes with 4µ8C and monitored mrna levels in the absence and presence of ER stress. Consistent with being an RIDD mrna substrate, Id1 mrna levels significantly decreased after 6 hours when treated with Thapsigargin (Figure 3.11A), which promotes ER stress by depleted ER Ca 2++ stores, and this decrease was blocked in IRE1α-depleted primary keratinocytes. Sequence analysis of Id1 mrna revealed five IRE1α consensus cleavage sites similar to unspliced Xbp1 mrna and in some cases these were associated with predicted stem loop structures using M-fold software (Figure 3.11B and data not shown)(44). We next investigated whether Id1 mrna serves as a direct IRE1α cleavage substrate at these consensus sites by using an in vitro RNA cleavage assay. Total RNA from primary keratinocytes was subject to in vitro RNA cleavage assays using the cytosolic portion of GST-IRE1α, which includes the kinase and RNase domains. Quantitative RT-PCR was performed to determine the effect of GST-IRE1α on Id1 mrna levels using primers that flank these putative cleavage sites identified by sequence analysis (Figure 3.11C). Id1 mrna levels were significantly decreased following addition of increasing amounts of GST-IRE1α in regions corresponding to stem loop sites ( , ) (Figure 3.11E). In addition, as a control heat denatured GST-IRE1α did not decrease Id1 mrna levels at these cleavage sites. Moreover, Id1 mrna levels were not cleaved within regions that did not contain putative stem loop sites indicating that IRE1α cleavage was specific within Id1

143 Figure Id1 mrna is a direct cleavage substrate of IRE1α. (A) Id1 mrna levels were measured by q-pcr upon treatment with ER stress inducer Thapsigargin (500nM) for 5 hours in primary keratinocytes (B) Sequence alignment of Xbp1 mrna cleavage sites with similar regions in Id1 mrna. (C) Diagram of XBP1 and ID1 primers used that flank putative IRE1α-mediated cleavage sites or flank regions lacking putative sites (D,E) In vitro RNA cleavage assay. Total RNA extracted from primary keratinocytes was incubated with increasing amounts of GST-IRE1α-cyto in the presence of ATP for 2 hours at 37 C. Q-RT-PCR was then conducted to determine Xbp1 and Id1 mrna levels. Gapdh mrna levels was used for normalization. Values = mean ± SEM. *indicates significantly different values (P 0.05). 128

144 129 mrna (Figure 3.11E). Lastly, as a positive control, mrna levels decreased when using primers that flanked regions of well characterized stem loop sites in Xbp1 mrna. In contrast, no effect as observed within regions that did not contain these sites confirming that GST-IRE1α-mediated mrna cleavage was sequence specific (Figure 3.11D). Taken together, these data indicate that IRE1α mediates Id1 mrna cleavage, which will eventually lead to its degradation in a cellular environment such as in our model of v-ras Ha activation Downregulation of Id1 mrna is required for IRE1α-induced senescence in v- RAS Ha keratinocytes Since we identified Id1 as a direct RIDD mrna cleavage substrate of IRE1α, we determined if induction of ID1 due to inhibition of IRE1α RNase activity mediated the senescence response in v-ras Ha keratinocytes. To that end, we utilized ID1 shrna lentivirus and also an empty vector shrna lentivirus as a control. Co-transduction of primary keratinocytes with each shrna lentivirus in the presence of v-ras Ha followed by treatment with DMSO or 25µM 4µ8C until day 11 after transduction. ID1 shrna knockdown decreased the protein levels of ID1 when compared to v-ras Ha keratinocytes transduced with empty vector shrna (Figure 3.12A). Importantly, ID1 shrna knockdown suppressed ID1 protein levels induced by 4µ8C treatment (Figure 3.12A). We next examined if ID1 depletion rescued the senescence response in the presence of 4µ8C treatment in v-ras Ha keratinocytes. As expected, 4µ8C treatment significantly decreased the percentage of SA-β-Gal positive cells 8 and 11 days after transduction. In contrast, ID1 depletion restored the percent SA-β-Gal positive cells treated with 4µ8C to levels comparable to DMSO treated v-ras Ha keratinocytes (Figure

145 B). This suggests that inhibition of IRE1α RNase activity and subsequent inhibition of v-ras Ha induced senescence is mediated by induction of ID1. Together, these data suggest Id1 mrna cleavage and degradation by IRE1α precedes and ultimately promotes v-ras Ha -induced senescence. This IRE1α-Id1 mrna decay axis in combination with reduced Xbp1 mrna splicing during senescence contributes to inhibit tumor development and malignant progression. 3.4 Discussion Oncogene-induced senescence induced by oncogenic RAS or other pertinent oncogenes is a potent tumor suppressive pathway serving as a fail-safe mechanism to development of malignant cancer (13, 45). While initially thought to be an artifact of culturing conditions in vitro, recent studies have validated the importance of this pathway in vivo with the detection of senescent cells and senescence-associated signaling programmes in premalignant tumors (46, 47). While the senescence phenotype is conserved among different cell types and tissues, the genetic requirements involved remain debatable (48). Similarly, activation of the UPR has been reported in a variety of tumors (49, 50). However, pharmacological-induced ER stress models have established that the UPR can have both cytoprotective and cytotoxic outcomes, depending on levels of ER stress. Thus, the role of ER stress and the UPR in tumor development remains unclear. Previous reports have shown that ER stress and the UPR can promote or impede oncogene-induced senescence (32, 51). In some cases, the UPR has no role altogether in the senescence response (32). The causative factors that determine a pro- or anti-oncogenic role for ER stress and the

146 131 UPR during senescence are likely very complex and depend on cell type, oncogenic insult, and the signaling pathways involved. Our studies presented herein are the first to directly demonstrate that IRE1α can mediate both tumor suppressor and oncogenic functions in cancer through opposing RNase outputs. Evidence of these divergent functions by IRE1α come from chemical-genetic studies, where Xbp1 mrna splicing serves as an adaptive pro-survival response while mrna decay through RIDD has a Figure IRE1α-mediated cleavage of Id1 mrna controls v- RAS Ha -induced senescence response. V-RAS Ha keratinocytes were transduced with shcontrol or shid1 and treated with 4µ8C (25µM) on day 2 after transduction and examined on day 8 and day 11. (A) Western blot analysis of ID1 protein levels on Day 11 after transduction. ACTIN was used as a loading control. (B) Quantification of SA-β-Gal positive cells. Values = mean ± SEM. *indicates significantly different values (P 0.05).

147 132 pro-apoptotic role (6). Moreover, a role for RIDD has been illustrated in other disease models and in some cases this function opposes XBP1 (52), lending credence to the idea that both IRE1α RNase functions are relevant in certain physiological contexts. In our model, transduction of primary keratinocytes with oncogenic v-ras Ha rapidly activates IRE1α-mediated Xbp1 mrna splicing as well as RIDD. In contrast, we did not observe activation of ATF6 and PERK UPR signaling pathways. Selective activation of distinct UPR signaling pathways as highlighted here with v-ras Ha is contrasted against pharmacological-induced ER stress, where all three arms of the UPR are activated(25). Under certain physiological conditions, selective activation of distinct arms of the UPR may be required for malignant progression. For example, KRAS transformed mouse embryonic fibroblasts require PERK activation under hypoxic stress conditions, ensuring cancer cells receive an adequate supply of oxygen and nutrients by regulating angiogenesis and tumor growth (53). In contrast, primary melanocytes transduced with HRAS activate all three branches of the UPR and each transducer has a causal role in promoting premature senescence (32). One question that needs to be addressed is what can account for selectivity of IRE1α activation in our model? Evidence suggests that all three UPR transducers bind BiP when in an inactive state on their ER-luminal domains (18, 19). While all three ER-luminal domains share some homology, with PERK and IRE1α being more similar than ATF6 (1), it is possible that upon accumulation of unfolded proteins BiP remains associated with PERK and ATF6, but selectively dissociates from IRE1α leading to activation. In addition, it has been shown that the ER-luminal domain of IRE1α has a similar structure and groove to major histocompatibility complex class 1 (MHC-1) and can recognize and bind unfolded

148 133 proteins upon dissociation of BiP leading to oligomerization (54). Exploring these ideas should lead to a better understanding of selective activation of IRE1α versus complete activation of all three UPR transducers in physiological disease settings. Here, we show that activation of the IRE1α-XBP1S pathway is linked to cell proliferation as our results demonstrate that XBP1S and not RIDD is required for the initial proliferative response by v-ras Ha and is dependent on ER stress. This is in agreement with previous studies that show XBP1S is overexpressed in many human cancers and is critical for cell proliferation and tumor growth (55-57). While it seems counterintuitive that RIDD would be activated during v-ras Ha -induced proliferation given a role in apoptosis, basal activation of this pathway occurs even before onset of apoptosis during ER stress (5, 6). This suggests that mrna decay in some capacity can be cytoprotective by modulating incoming protein load in the ER. Indeed, basal activation of RIDD is required to maintain proper ER membrane homeostasis (5, 23) although it is not clear from our studies if a similar mechanism occurs in v-ras Ha keratinocytes. The balance between IRE1α-mediated Xbp1 mrna splicing and mrna decay through RIDD seems critical to the type of cellular response triggered. In pharmacological-induced ER stress, Xbp1 mrna splicing occurs in a rapid but transient manner, but it is not until these levels are reduced that RIDD activation increases and can promote apoptosis (23). Here it seems that the overt effects of RIDD only take place when the XBP1S-mediated adaptive functions of IRE1α are reduced and mrna decay becomes sustained. We observed a similar response as Xbp1 mrna splicing is highest during v-ras Ha -induced cell proliferation but decreases during senescence. In

149 134 contrast, RIDD mrna levels are further downregulated during v-ras Ha induced senescence suggesting activating of this function of IRE1α. However, ER stress levels seem to be dampened during senescence and previously, we have shown that pharmacological-induced ER stress inhibits v-ras Ha -induced senescence (51), suggesting that RIDD may function independent of ER stress. The opposing function of IRE1α and XBP1 was validated as shrna knockdown of each protein inhibited or promoted senescence, respectively. Furthermore, we demonstrated that XBP1 depletion promotes senescence through feedback activation of IRE1α RNase activity. Hyperactive IRE1α has been observed in other XBP1 deficient models. In one study, genetic loss of XBP1 facilitates cancer progression in intestinal epithelium through, in part, intestinal stem cell (ISC) expansion, which is mediated by IRE1α hyperactivation (58). In contrast to other studies and our result presented here, this suggests an antioncogenic role for XBP1 but a pro-tumorigenic role for IRE1α. In other cases, XBP1 deficiency does not overexpress IRE1α such as in triple negative breast cancer (estrogen -, progesterone -, HER2 -), a highly aggressive form of malignant cancer with limited treatment options (56). However, it is not clear from these studies if the RIDD function is activated regardless of IRE1α activation status in the absence of XBP1. Our studies do not explain how IRE1α RNase functions are selectively modulated during v-ras Ha -induced senescence. Previous studies have shown that under ER stress IRE1α forms discrete foci of high order oligomers (59). Furthermore, another study proposes that formation of high-order IRE1α oligomers and subsequent increases in IRE1α phosphorylation have reduced specificity for unspliced Xbp1 mrna but becomes more preferential for cleavage of RIDD substrates due to a relaxed RNase

150 135 pocket (6). One idea is that as v-ras Ha keratinocytes undergo senescence this promotes the formation of IRE1α oligomers that 1) leads to reduced Xbp1 mrna specificity or 2) and causes sustained activation of RIDD. However, further experiments are needed to investigate this mechanism of selectively in v-ras Ha keratinocytes by examination of oligomer formation using protein cross-linking techniques or visually using a GFP-tagged IRE1α (59). Initially, IRE1α-mediated Xbp1 mrna splicing and RIDD is regulated by the MEK-ERK pathway during cell proliferation. However, IRE1α or XBP1-depleted v-ras Ha keratinocytes differentially regulate the MEK-ERK pathway during senescence paralleling the divergent senescence response. Several studies have demonstrated that hyperactive RAS as well as MEK-ERK signaling can paradoxically trigger senescence (36, 60, 61). Threshold levels of this pathway may serve as a potent fail-safe mechanism triggering activation of stress pathways that fundamentally alters the signaling to induce expression of p53 as well as cell cycle inhibitors such as p16 and p21(45). Not only did IRE1α-depleted v-ras Ha keratinocytes evade senescence by reducing MEK-ERK signaling and p38 activation, we observed opposite regulation in XBP1-depleted v-ras Ha and this, in turn, was mediated by IRE1α. This suggests that modulation of v-ras Ha -induced senescence by IRE1α occurs through both positive and negative feedback mechanisms converging on the MEK-ERK pathway and is dependent on the level of Xbp1 mrna splicing as well as RIDD (Figure 3.13). We provided evidence that Id1 mrna downregulation mediates senescence induction by IRE1α. ID proteins have a well characterized role in promoting tumorigenesis(62). Moreover, ID1 overexpression suppresses RAS-induced

151 136 senescence (41) suggesting that at least in early stages of cancer progression with activated RAS mutations ID1 is downregulated. Furthermore additional studies are needed to compare regulation Id1 mrna in primary, premalignant and malignant cancer cell lines to determine if the RIDD function remains intact through cancer progression or is just obscured due to transcriptional activation at the ID1 promoter by activation of other oncogenic signaling pathways. Although Id1 is a cytosolic mrna, a previous study that fractionated ER-bound and soluble polysomes identified this mrna as enriched on the ER membrane (42). Thus, it is possible that Id1 mrna is in close proximity to IRE1α for mrna degradation. Regardless, further studies are needed to examine if Id1 mrna is enriched in ER-bound polysome fractions in our v-ras Ha model and if IRE1α depletion increases Id1 mrna levels on the ER. In addition to Id1 mrna, we identified a number of other potential RIDD mrna substrates including Igfbp2 and Timp3 (see Table 3.1). Approximately 50% of those mrnas identified are secreted into the ECM or associate with the plasma membrane. Thus, the potential of these IRE1α-regulated mrnas to impact other signaling pathways could be critically important to determining the senescence response induced by v- RAS Ha. For example, IGFBP2 is one of six binding proteins that associates with insulinlike growth factor (IGF) and regulates its access to the IGF receptor (63). IGFBP2 is associated with several different cancers including ovarian (64), prostate (65), and glioblastoma (66). Furthermore, IGFBP2 can promote cell proliferation, invasion, and facilitate chemoresistance in several cancer cell lines through activation of signaling pathways including MEK-ERK (64, 67). Furthermore, IGFBP2 can interact with α5 integrin, and modulate migration and invasion in glioblastoma cell lines (65). Based on

152 137 Figure Model for opposing IRE1α RNase outputs during v- RAS Ha -induced proliferation and senescence. (A) Transduction of v- RAS Ha into primary keratinocytes causes ER stress-dependent activation of IRE1α and subsequent Xbp1 mrna splicing and RIDD. XBP1S dampens IRE1α oligomerzation formation through as of yet undefined mechanisms, retaining RIDD activation in a basal state and preventing MEK-ERK hyperactivation and senescence. ER stress and XBP1S in turn promote proliferation. (B) During v-ras Ha -induced senescence, ER stress levels are reduced leading to downregulation of XBP1S. Reduced XBP1S leads to IRE1α hyperactivation and subsequent high-order oligomerization formation, allowing increased mrna decay of critical senescence regulators such as ID1. This cuases amplification of RAS signals including ERK and stress kinase p38.

153 138 this evidence, further studies need to be carried out to determine if Igfbp2 mrna or other targets identified from our studies are critical to malignant progression, is a direct IRE1α cleavage substrate and can modulate the cellular response to v-ras HA. One of the outstanding questions from our studies is if these divergent responses can be recapitulated in an in vivo environment. Thus, studies need to be performed where v-ras Ha keratinocytes deficient in IRE1α or XBP1 are mixed with normal fibroblasts and seeded onto a prepared graft site to allow tumor formation. This assay is well-established and after several weeks benign papillomas form that are histologically similar to DMBA-TPA papillomas (17). Furthermore, a small percentage of these grafts convert to carcinomas. Therefore, by comparing IRE1α and XBP1 deficient v-ras Ha keratinocytes, we can determine rate of tumor growth and frequency of malignant conversion (68). Based on our current evidence in vitro, we would expect XBP1 deficiency to attenuate tumor growth and prevent formation of tumors. In contrast, IRE1α deficiency should at the very least show a significant increase in malignant conversion. Other studies would be designed to determine if RIDD mrna substrates such as Id1 is responsible for these changes. Similarly, tumor induction studies with DMBA-TPA protocols should be performed with IRE1α fl/fl (69) and XBP1 fl/fl (70) mice both crossed with K14-Cre mice to allow keratinocyte specific deletion of these proteins in the skin. Tumor number, size, and histology from each of these groups should be determined. Furthermore, examination of IRE1α signaling and RIDD mrna substrate expression should be performed. Moreover, isolation and propagation of benign and malignant cell lines from these genotypes at different tumor stages should be done so

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161 146 Chapter 4: IRE1α and ER stress dictates TGFβ1 signaling and senescence 4.1 Abstract We have previously shown that the UPR transducer IRE1α regulates divergent cell fate responses dependent on two distinct RNase activities- Xbp1 mrna splicing and mrna decay of ER localized and cytosolic mrnas through a process called RIDD using an in vitro model of epithelial carcinogenesis driven by oncogenic v-ras Ha. In this model, Xbp1 mrna splicing serves as an adaptive response causing cell proliferation and inhibiting v-ras Ha -induced senescence, while RIDD promotes growth arrest and senescence in the absence of XBP1. Here we further investigated the regulation of IRE1α RNase outputs in v-ras Ha keratinocytes by the TGFβ1 tumor suppressor pathway. TGFβ1 treatment significantly enhanced v-ras Ha - induced senescence which was associated with dampened ER stress and Xbp1 mrna splicing but caused greater downregulation of RIDD substrates. Treatment of v-ras Ha keratinocytes with the TGFb1 type I receptor inhibitor SB , previously shown inhibit v-ras Ha -induced senescence, enhanced Xbp1 mrna splicing and attenuated RIDD. Furthermore, pharmacological inactivation or depletion of IRE1α prevented TGFβ1-induced senescence independent of XBP1 and this was not due inhibition of TGFβ1-induced transcriptional activation. Conversely, pharmacologically induced ER stress inhibited TGFβ1-induced senescence and growth arrest in v-ras Ha keratinocytes. This inhibition was due to inactivation of downstream TGFβ1 signaling through a mechanism that involved impairment of ligand-induced receptor internalization. Together, selective modulation of v-ras Ha -induced IRE1α RNase activity toward an mrna decay pathway by TGFβ1 may suppress tumorigenesis by promoting senescence. Furthermore, mild

162 levels of ER stress, which promotes the adaptive functions of the UPR, represents a new mechanism to inactivate TGFβ1 signaling and inhibit the senescence response Introduction IRE1α is a bifunctional kinase/endoribonuclease (RNase) stress sensor, and part of the unfolded protein response (UPR), that becomes activated in response to misfolded protein accumulation in the ER (1). Under ER stress, IRE1α activation initiates the cytosolic splicing of the Xbp1 mrna whose mature transcript encodes an active transcription factor, called XBP1S. In turn, XBP1S regulates the expression of genes that restore homeostatic conditions within the ER (1). However, recent studies suggests that when ER stress is at high levels IRE1α undergoes a series of further activation steps, resulting in higher levels of phosphorylation and oligomerization leading to increased activation of a potent mrna degradation pathway, called RIDD, that cleaves mrnas critical for survival and leading to apoptosis (1-3). Thus, IRE1α is a critical executioner of cell fate responses in response to varying levels of ER stress. While most of these studies have elucidated the role of IRE1α using chemical genetic and pharmacological ER stress models, the role of IRE1α in pathological conditions such as cancer remains unresolved due to different model systems or incomplete examination of both IRE1α RNase functions. Oncogenic HRAS triggers irreversible cellular senescence and growth arrest in normal cells and is considered a fail-safe mechanism to suppress tumor development by preventing the progression of benign lesions to malignancies in the absence of additional oncogenic mutations (4). Senescence induced by oncogenic HRAS is

163 148 associated with concomitant induction of various tumor suppressor pathways including the expression of p16 and p53 (5), induction of stress pathways such as p38 (6), and paradoxical activation of the mitogenic MEK-ERK pathway (7). Furthermore, recent evidence implicates a role for ER stress during oncogenic HRAS-induced senescence although it remains controversial (8, 9). Primary melanocytes expressing oncogenic HRAS undergo senescence and growth arrest that is dependent on IRE1α-XBP1S, ATF6, and PERK UPR members suggesting an anti-oncogenic role of the UPR in this response. However, this occurs in the absence of induction of classical senescence markers such as p16 and p53 (8). In contrast, pharmacological ER stress or knockdown of specific UPR members such as XBP1 and ATF4, suppresses or enhances senescence, respectively (9). Data in Chapter 3 shed new light on these discrepancies, as IRE1α can inhibit or promote oncogenic HRAS-induced senescence depending on the levels of its RNase outputs. IRE1α-mediated Xbp1 mrna splicing primarily serves as an adaptive response by promoting cell proliferation in response to ER stress. However, as cells undergo senescence, XBP1S levels are downregulated while IRE1α-mediated mrna decay through RIDD promotes senescence. These data further suggest that inactivation of RIDD is required for malignant progression and is supported by the fact that a recent proteomic study identified several IRE1α mutations in human cancers that may be defective in its apoptotic functions under ER stress (10, 11). Furthermore, we identified the oncogenic factor ID1 as a target of IRE1α RNase activity further substantiating the role of RIDD as a potent tumor suppressor pathway. TGFβ1 is a potent cytostatic growth factor and tumor suppressor during the earliest stages of cancer progression (12, 13). Furthermore, several studies have

164 149 implicated TGFβ1 signaling as a critical mediator of oncogenic HRAS-induced senescence. High levels of TGFβ1 occur as cells expressing oncogenic HRAS undergo senescence and this is associated with increased levels of Smad3 (14, 15). Indeed, genetic inactivation of TGFβ1 (15), Smad3 (14), or overexpression of Smad7 (16) inhibits senescence and promotes malignant progression. Furthermore, pharmacological blockade of the TGFβ type I receptor, ALK5, inhibits senescence (17). Together, TGFβ1 signaling is an essential determinant of whether oncogenic HRAS undergoes senescence and growth arrest or oncogenic transformation. Thus, the present studies were implemented to determine if a potent tumor suppressor pathway can modulate IRE1α RNase functions to favor senescence using genetic and pharmacological approaches and also to determine if pharmacological ER stress can inhibit senescence induced by TGFβ Results TGFβ1-induced senescence is associated with dampened IRE1α-mediated Xbp1 mrna splicing and enhanced RIDD. Previous studies implicate components of TGFβ1 signaling as a critical mediators of v-ras Ha -induced senescence and inactivation of this pathway leads to malignant conversion(14-16). Given the results in Chapter 3, and that v-ras Ha keratinocytes secrete high levels of TGFβ1 during senescence (15), we hypothesized that this pathway may modulate IRE1α-mediated Xbp1 mrna splicing and RIDD to favor development of the senescence phenotype in v-ras Ha keratinocytes. To test this hypothesis, we treated v-ras Ha keratinocytes 2 days after transduction with 1ng/mL

165 150 TGFβ1 every other day for 5 days during a time period when there are few senescent cells in untreated v-ras Ha keratinocytes. As expected, there was a significant increase in SA-β-Gal positive cells as early as 1 day after TGFβ1 treatment and this continued to increase over the course of subsequent treatments (Figure 4.1A, C). After 5 days treatment, the percent SA-β-Gal positive cells was approximately 60% in the TGFβ1 treated group and 5% in untreated v-ras Ha keratinocytes. These cells displayed a similar morphology to v-ras Ha -induced senescence as well as previous studies where TGFβ1 caused cytoplasmic vacuolization and cell flattening (8, 15) (Figure 4.1C). Furthermore, as early as 1 day after TGFβ1 treatment there was a significant reduction in cell proliferation, from 40% BrdU positive cells in untreated to 5% BrdU positive cells in TGFβ1 treated v-ras Ha keratinocytes (Figure 4.1B). This is in agreement with our previous studies that oncogenic v-ras Ha does not block immediate cytostatic effects of TGFβ1 in primary keratinocytes (18). In addition, TGFβ1 treatment of v-ras Ha keratinocytes induced many of the same senescence markers associated with v-ras Ha - induced senescence including phosphorylation of p38, and induction p16 senescence markers (Figure 4.1D). TGFβ1 treatment of v-ras Ha keratinocytes reduced IRE1α phosphorylation but increased total IRE1α levels at each timepoint (Figure 4.2A). Reduced IRE1α phosphorylation correlated with decreased ER stress markers XBP1S and BiP protein levels as early as 1 day after TGFβ1 treatment (Figure 4.2A), suggesting that TGFβ1 dampens ER stress in v-ras Ha keratinocytes. To determine the effect of TGFβ1 on the RIDD function of IRE1α, we examined mrna levels of Igfbp2, Timp3, and Id1. All three targets were previously identified by microarray analysis as candidate RIDD substrates and Id1 was validated using in vitro RNA cleavage assays

166 151 Figure 4.1. TGFβ1 promotes senescence in v-ras Ha keratinocytes. V- RAS Ha keratinocytes were treated on day 2 after transduction with TGFβ1 (1ng/mL) and examined 1, 3, and 5 days after treatment. (A) Quantification of percent SA-β-Gal positive cells and (B) Quantification of percent BrdU positive cells in v-ras Ha keratinocytes treated with TGFβ1. (C) Photomicrographs of SA-β-Gal staining (Top) and bright field images (Bottom) of cytoplasmic vacuolization. Red arrows point to positive cells. (D) Western blot analysis of p-p38, total p38, p16 and ACTIN in v-ras Ha keratinocytes treated with TGFβ1. Values = mean ± SEM. *indicates significantly different values (P 0.05) from v-ras Ha values left untreated at similar timepoints.

167 (see Chapter 3). As expected, compared to primary keratinocytes all three were 152 downregulated by v-ras Ha (Figure 4.2B). TGFβ1 treatment further downregulaion mrna levels of Igfbp2, Timp3, and Id1 at each timepoint (Figure 4.2B) consistent with a role for RIDD in promoting v-ras Ha induced senescence. Taken together, these data demonstrate that exogenous TGFβ1 can dampen ER stress and modulate IRE1α RNase activity to promote senescence in v-ras Ha keratinocytes Inactivation of TGFβ1 signaling in v-ras Ha keratinocytes enhances Xbp1 mrna splicing and inhibits RIDD. Since treatment of v-ras Ha keratinocytes with TGFβ1 inhibits IRE1α-mediated Xbp1 mrna splicing and enhances mrna downregulation of RIDD substrates, we next tested if basal TGFβ1 signaling in v-ras Ha keratinocytes also regulates IRE1αmediated Xbp1 mrna splicing and RIDD since both RNase functions are activated early upon v-ras Ha transduction. To inhibit TGFβ1 signaling, we treated v-ras Ha keratinocytes with the TGFβ1 type I receptor inhibitor (ALK5) SB , which acts as a competitive ATP binding site kinase inhibitor and prevents Smad phosphorylation (19). Furthermore, we have previously shown that treatment with SB inhibits v- RAS Ha induced senescence (17). Figure 4.3A shows that treatment of v-ras Ha keratinocytes with 10µM SB for 1 day increased IRE1α phosphorylation but had no effect on total IRE1α protein levels. This increase in IRE1α phosphorylation was associated with increased spliced Xbp1 mrna as well as XBP1S protein levels. There was also an increase BiP protein levels suggesting that SB treatment increases ER stress (Figure 4.3A, B). However, after SB treatment for 4 days the increase in IRE1α phosphorylation, XBP1S and BiP protein levels had returned to control levels

168 153 Figure 4.2. TGFβ1 inhibits IRE1α-mediated Xbp1 mrna splicing but enhances downregulation of RIDD substrates during senescence. (A) Western blot analysis of p-ire1α, total IRE1α, BiP, and XBP1S in v-ras Ha keratinocytes treated with TGFβ1 (1ng/mL) on day 2 after transduction and examined at 1, 3, and 5 days after treatment. ACTIN and VINCULIN were used as loading controls. (B) Q-PCR of Timp3, Igfbp2, and Id1 of v-ras Ha keratinocytes treated with TGFβ1 (1ng/mL) and examined at similar timepoints as in (A). Values = mean ± SEM. *indicates significantly different values (P 0.05). All expression values were normalized to primary keratinocytes without v-ras Ha on day 3 and represent fold-change.

169 and total IRE1α protein levels were reduced when compared to v-ras Ha keratinocytes treated with DMSO (Figure 4.3A). Thus, these effects of SB were transient in v- 154 RAS Ha keratinocytes. Next, we determined the impact of SB treatment on IRE1α-mediated RIDD by examining mrna levels of substrates identified in Chapter 3. Treatment with SB lead to a significant inhibition in downregulation of mrna levels of RIDD targets Id1, Igfbp2, and Timp3 by 2 days in v-ras Ha keratinocytes and this inhibition remained persistent and was even enhanced by 4 days (Figure 4.3B), suggesting that inhibition of TGFβ1 signaling reduces cleavage of RIDD substrates in v-ras Ha keratinocytes. Taken together, these results are consistent with endogenous TGFβ1 signaling promoting the senescence response through modulation of IRE1αmediated Xbp1 mrna splicing and RIDD IRE1α inactivation blocks TGFβ1-induced senescence without inhibiting Smad activation. TGFβ1 signaling is a potent inhibitor of cell growth and promotes senescence in v-ras Ha keratinocytes presumably through activation of various inhibitors of cell cycle progression including induction of p16 and p19 (20), or downregulation of proteins that induce proliferation such as c-myc (14). To determine if IRE1α mediates TGFβ1- induced senescence, v-ras Ha keratinocytes were co-transduced with control, IRE1α, and XBP1 shrna, or pretreated on day 2 after v-ras Ha transduction with 4µ8C for 2 days followed by treatment with TGFβ1 for 3 days to induce senescence (Figure 4.4). As expected, TGFβ1 treatment induced senescence with approximately 40% SA-β-Gal positive cells detected in v-ras Ha keratinocytes transduced with control shrna or treated with DMSO (Figure 4.4A). Similar to studies in Chapter 3, IRE1α depletion

170 155 Figure 4.3. SB enhances Xbp1 mrna splicing and inhibits downregulation of RIDD mrna substrates in v-ras Ha keratinocytes. V-RAS Ha keratinocytes were treated with SB (10µM) on day 2 after transduction and examined 1 and 4 days after treatment. (A) Western blot analysis of p-ire1α, total IRE1α, BiP, and XBP1S. VINCULIN and ACTIN were used as loading controls. (B) Q-PCR of Xbp1s, Timp3, Igfbp2, and Id1 mrna levels at indicated timepoints. Values = mean ± SEM. *indicates significantly different values (P 0.05) from v-ras Ha values treated with DMSO at similar timepoint. All expression values were normalized to primary keratinocytes wthout v-ras Ha and treated with DMSO on day 1 and represent fold-change.

171 156 reduced the number of SA-β-Gal positive cells detected upon induction by TGFβ1 by about 4-fold to approximately 10% (Figure 4.4A). Similarly, treatment with 4µ8C inhibited senescence to similar levels as IRE1α depletion suggesting that TGFβ1- induced senescence in v-ras Ha keratinocytes is dependent on intact IRE1α RNase activity. In contrast, XBP1 depletion enhanced TGFβ1-induced senescence with about 60% SA-β-Gal positive cells detected (Figure 4.4A). Our previous studies showed that Smad3 inactivation or Smad7 overexpression inhibited v-ras Ha -induced senescence and promoted malignant conversion indicating that Smad signaling is critical in mediating senescence (14, 16). Therefore, we tested if lack of functional IRE1α inhibits senescence by impacting TGFβ1-induced Smad activation. Control, IRE1α, and XBP1 shrna were co-transduced with v-ras Ha in primary keratinocytes and on day 4 after transduction transfected with a Smad binding element (SBE) reporter plasmid to measure Smad transcriptional activity in the nucleus. Figure 4.4B shows that following TGFβ1 treatment for 48 hours, there was slight decrease in Smad transcriptional activity when IRE1α depleted v-ras Ha keratinocytes were compared to control cells. However, this decrease was not significant and does not account for the nearly complete inhibition of senescence induced by TGFβ1 (Figure 4.4A). Similarly, no difference was observed when XBP1 depleted v-ras Ha keratinocytes were compared to control cells (Figure 4.4B). Taken together, this indicates that despite intact TGFβ1 signaling, the senescence response can be inhibited through ER perturbations dependent on IRE1α RNase activity independent of spliced Xbp1 mrna.

172 157 Figure 4.4. IRE1α is required for TGFβ1-induced senescence but does not affect Smad activation. (A) v-ras Ha keratinocytes were transduced with control, IRE1α, or XBP1 shrna or pretreated with 4µ8C (25µM) for 48 hours on day 2. On day 4, TGFβ1 (1ng/mL) was added in the presence or absence of 4µ8C where appropriate and refreshed after 48 hours. Percent SA-β-Gal positive cells were determined on Day 7. (B) v-ras Ha keratinocytes were transduced with indicated shrna as in (A) and on day 4 after transduction transfected with psbe-luc Smad reporter and prl-tk control plasmids for 24 hours followed by treatment with TGFβ1 (1ng/mL) for 48 hours. Relative firefly luciferase was measured with a illuminometer and normalized to that of Renilla luciferase. Values = mean ± SEM. *indicates significantly different values (P 0.05), ns indicates not significant.

173 ER stress inhibits TGFβ1-induced senescence Several lines of evidence implicate ER stress as an adaptive response to oncogenic v-ras Ha. First, data in Chapter 3 implicate ER stress as part of the proliferation response to oncogenic v-ras Ha. Furthermore, our previous study has shown that treatment of v-ras Ha keratinocytes with pharmacological drugs that induce ER stress can block v-ras Ha -induced senescence (9). To determine if ER stress inhibits senescence induced by TGFβ1, we used two different pharmacological ER stress agents that lead to accumulation of unfolded proteins, thapsigargin and tunicamycin. Thapsigargin treatment leads to ER Ca 2+ depletion due to inhibition of an ER ATPase and tunicamycin inhibits N-linked glycosylation. Thapsigargin and tunicamycin were pretreated at 5nM and 50ng/mL, respectively, for 24 hours in v-ras Ha keratinocytes and then treated with TGFβ1 in the presence of absence of these drugs for 5 days. Pharmacological-induced ER stress significantly inhibited TGFβ1-induced senescence although thapsigargin treatment had a more pronounced effect when compared to tunicamycin treatment (Figure 4.5B). Little effect was observed by thapsigargin or tunicamycin in untreated v-ras Ha keratinocytes. In addition, characteristic cytoplasmic vacuolization and cell flattening observed upon TGFβ1- induced senescence was suppressed in the presence of thapsigargin or tunicamycin (Figure 4.5C and data not shown). Similarly, treatment of v-ras Ha keratinocytes with thapsigargin and tunicamycin in the presence of TGFβ1 had a significant effect in blocking growth arrest in agreement with inhibition of senescence in these cells (Figure 4.5D).

174 159 Figure 4.5. Pharmacological ER stress inhibits TGFβ1-induced senescence. (A) V-RAS Ha keratinocytes were treated with DMSO, Thapsigargin(5nM), or Tunicamycin (50ng/mL) in the presence of absence of TGFβ1 (1ng/mL) and percent SA-β-Gal positive cells determined on day 1, 3, and 5 days after treatment. (B) V-RAS Ha keratinocytes were treated as in (A) and percent BrdU positive cells quantified after 48 hours. Values = mean ± SEM. *indicates significantly different values (P 0.05) from respective controls ER stress inhibits Smad activation through impairment of TGFβ1 receptor internalization. We next examined if pharmacological-induced ER stress inhibited senescence and growth arrest through changes in TGFβ1 signaling. In contrast to the effect of IRE1α inactivation (see Figure 4.4C), when we treated v-ras Ha keratinocytes with thapsigargin or tunicamycin in the presence or absence of TGFβ1, there was a dose dependent inhibition in Smad transcriptional activity even at the lowest doses as determined by SBE reporter (Figure 4.6A, B). Similarly, examination of Smad2 phosphorylation by western blot analysis verified this dose dependent inhibition of

175 160 TGFβ1 activation although thapsigargin was more potent than tunicamycin in agreement with inhibition of TGFβ1-induced senescence (Figure 4.6C, D). Next, we sought to determine the mechanism of inhibition Smad activation by pharmacological ER stress. When we transduced v-ras Ha keratinocytes with increasing amounts of a constitutively active type I ALK5 receptor adenovirus for 24 hours (14), we observed that increasing MOI caused a dose dependent increase in Smad activation (Figure 4.7A). However, when v-ras Ha keratinocytes transduced with the ALK5 adenovirus was treated in the presence of 5nM thapsigargin, pharmacological ER stress was still able to inhibit Smad transcriptional activity at each MOI (Figure 4.7A). This suggests that despite an active type I receptor, Smad phosphorylation is still impaired and this blocked responses to TGFβ1. We next reasoned that since the type II and type I TGFβ1 receptors undergo maturation in the ER before being displayed on the cell surface, ER stress could alter expression of these proteins. We treated v-ras Ha keratinocytes in the presence or absence of both ER stress inducers and labeled cell surface proteins with a cell impermeable Sulfo-NHS-biotin reagent. Following lysis, cells were immunoprecipitated with streptavidin agarose beads and western blot analysis performed for TβRII and TβRI (Figure 4.7B). Surprisingly, thapsigargin or tunicamycin treatment caused a significant increase in the expression of both receptors with treatment of thapsigargin having a more dramatic effect consistent with the impact of this agent on senescence and Smad phosphorylation (Figure 4.7B). TGFβ1 receptors constitutively undergo endocytosis in the absence of TGFβ1. After undergoing endocytosis, they are sorted and recycled back to the plasma membrane. However, upon TGFβ1 treatment, TGFβ1-bound receptors can continue and even maximize their

176 161 Figure 4.6. Pharmacological ER stress inhibits TGFβ1 signaling. V-RAS Ha keratinocytes were transfected with psbe-luc Smad reporter and prl-tk control plasmids for 24 hours followed by pretreatment of increasing doses of Thapsigargin (A) or Tunicamycin (B) for 24 hours followed by treatment of TGFβ1 (1ng/mL) for 24 hours. Relative firefly luciferase was measured with a illuminometer and normalized to that of Renilla luciferase. (C) Western blot analysis of p-smad2 and total SMAD2 of v-ras Ha keratinocytes pretreated with Thapsigargin at increasing doses as indicated for 24 hours followed by treatment with TGFβ1(1ng/m) for 6 hours. (D) Western blot analysis of p-smad2 and total SMAD2 of v-ras Ha keratinocytes pretreated with Tunicamycin with increasing doses as indicated similar to that in (C). Values = mean ± SEM. *indicates significantly different values (P 0.05) when compared to v-ras Ha keratinocytes treated with TGFβ1.