Regulation of Erk Signal Transduction by Signal-Induced Cysteine Oxidation

REGULATION OF ERK SIGNAL TRANSDUCTION BY SIGNAL-INDUCED CYSTEINE OXIDATION

JEREMIAH D. KEYES

A Dissertation submitted to the Graduate Faculty of WAKE FOREST UNIVERSITY GRADUATE SCHOOL OF ARTS AND SCIENCES in Partial Fulfillment of the Requirements for the Degree of DOCTOR OF PHILOSOPHY In Biochemistry and Molecular Biology August 2016 Winston-Salem, North Carolina

Approved By: Leslie Poole, Ph.D., Advisor Cristina Furdui, Ph.D., Chair Doug Lyles, Ph.D. Larry Daniel, Ph.D. W. Todd Lowther, Ph.D. ACKNOWLEDGMENTS

This has been an extremely challenging and rewarding project, and I could not have brought it to this point without the help of several individuals. I’d like to first thank my advisor, Dr. Leslie Poole, for encouraging me to pursue my interest in this project despite the fact that it diverged from her normal areas of study. Dr.

Poole fostered a rich atmosphere for study and collaboration and helped me to get to an unusual number of conferences to push my project forward. The last couple months she has especially put forth a heroic effort to see me to this point.

I would also like to thank Dr. Kimberly Nelson, who taught critical thinking through every day discussions over experimental design and analysis. Dr. Derek

Parsonage was an essential element in my study by teaching and performing molecular biology and protein purification. I need to especially thank LeAnn

Rogers, who taught me cell culture and, along with Rama Yammani who came in at just the right time, heroically helped in key discoveries to enable me to finish my project. I would also like to thank Gao, Laura Soito, and Chelsea Kesty who all contributed to my training and progress in this project. I thank Dr. Julie Reisz and Dr. Cristina Furdui for advice and analysis on mass spectrometry experiments. I need to give a special thank you to Dr. Rob Newman, who not only aided in his kinase signaling expertise, but encouraged me at a time when I was discouraged, which led me to an exciting post-doctoral opportunity. I also could not have made the progress I have without the help of Dr. Rony Seger, who gave critical and encouraging feedback on my studies, and Dr. Natalie Ahn who provided several established plasmids for ERK and MEK purification. I

i would also like to acknowledge Dr. Richard Watt, who introduced me to a love of research as an undergraduate student at Brigham Young University. I would like to thank each member of my committee, Dr. Doug Lyles, Dr. Todd Lowther, Dr.

Larry Daniel, and Dr. Cristina Furdui. Not only did they guide me throughout my graduate school tenure, but they helped me learn that consistent results are the truth – even if those results don’t match initial results.

I would like to especially thank members of my family. My mom, who helped me with my homework every day when I was young. My dad, who took me to science centers and taught me to cherish learning and always keep my doors open. My children, Porter, Hazel, and Nora, who have kept my imagination young by turning Eppendorf tubes into spaceships and allowing me to play with them almost every day. My brother Nathan, who reminded me at critical times of disappointment that science is more than failed experiments; it is enrapturing, fun, and wondrous. But most especially, my wife Samantha, who has by far been my greatest support and believer. She reminded me of my potential during my most difficult times. Her strength the past several years is unmatched as my attention has been siphoned into completing this project.

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS………………………………………………… i

LIST OF ABBREVIATIONS………………………………………………. iv

LIST OF ILLUSTRATIONS………………………………………..……… vii

ABSTRACT……………………………………………………………..….. x

1. INTRODUCTION…………………………………………………….... 1

2. ENDOGENOUS SULFENYLATION OF ERK IN RESPONSE

TO PROLIFERATIVE SIGNALS…………………………………….. 47

3. MODIFICATION OF rERK2 ACTIVITY IN RESPONSE TO IN

VITRO OXIDATION BY H2O2………………………………………… 90

4. CONCLUSIONS……………………………………………………….. 121

SCHOLASTIC VITAE……………………………………………………… 137

iii

LIST OF ABBREVIATIONS

ATP adenosine triphosphate

BCA bicinchoninic acid

CDK Cyclin Dependent Kinase

CK2 Casein Kinase 2

Clk Cell Division Cycle-like-kinase

DMEM Dulbecco's Modified Eagle Medium

DRS D-recruitment site

DTT Dithiothreitol

EDTA Ethylenediaminetetraacetic acid

EGF Epidermal Growth Factor

ELISA Enzyme-linked immunosorbent assay

EMEM Eagle's minimal essential medium

ERK Extracellular signal-regulated kinase

FBS Fetal Bovine Serum

FRET fluorescence resonance energy transfer

FRS F-recruitment site

GDP Guanosine diphosphate

GPCR G-protein couple receptor

GSK3 glycogen synthase kinase 3

GSNO S-Nitrosoglutathione

GTP Guanosine triphosphate

HR Hormone receptor

IAM iodoacetamide

JNK c-Jun N-terminal kinase

KO knock-out

LPA Lysophosphatidic acid

MAPK Mitogen Activated Protein Kinase

MEF Mouse Embryonic Fibroblast

MEK MAPK/ERK kinase

MKP Mitogen activated protein kinase phosphatase

MW Molecular Weight

NEM N-ethylmaleimide

NO Nitric Oxide

NTS Nuclear Translocation Signal

PDGF Platelet Derived Growth Factor

PDGFR platelet-derived growth factor receptor

PEG-Catalase Polyethylene Glycol-Catalase

PKC protein kinase C

PTP Protein Tyrosine Phosphatase

RNS Reactive Nitrogen Species

ROS Reactive Oxygen Species

RSS Reactive Sulfur Species

RTK Receptor Tyrosine Kinase

SDS-PAGE Sodium dodecyl sulfate – polyacrylamide gel electrophoresis

SNP Sodium Nitroprusside

SOS Son of Sevenless

TIRF Total internal reflection fluorescence microscopy

VEGF Vascular Endothelial Growth Factor

VSMC Vascular Smooth Muscle Cells

LIST OF ILLUSTRATIONS AND TABLES

TABLES PAGE

1-I Redox Sensitive Kinase 21

1-II MAPK oxidation in literature 22-23

3-I Cysteine Modifications Observed on rERK2 103

3-II List of proteins on function protein array 108

FIGURES PAGE

1-1 MAPK Signaling Cascades 4

1-2 MAPK Alignment 5

1-3 Signal-specific context to induce distinct cellular

responses through ERK1/2 signal cascade 8

1-4 Rat ERK2 and human ERK1/2 alignment with

secondary structure assignments 10

1-5 Structural analysis of ERK1/2 docking sites and NTS 12

1-6 Cysteine oxidation chemistry 27

2-1 Sulfenic acid trapping of cellular proteins during lysis 51

2-2 ERK1/2 cysteines are oxidized to sulfenic acid in

response to PDGF in NIH-3T3 cells 61

vii

2-3 ERK1/2 oxidation is caused by H2O2 generated in

response to PDGF in NIH-3T3 cells 62

2-4 Serum-depleted WI38 fibroblasts exhibit distinct

temporal patterns of sulfenic acid formation in response

to PDGF compared to serum-replete WI38 cells 64

2-5 Endogenous ERK1/2 oxidation inhibits kinase activity

towards Elk1 67

2-6 Potential modes of ERK regulation 76

2-S1 ERK1/2 biotinylation is due to labeling by DCP-Bio1 84

2-S2 Oxidation of ERK1/2 in prostate cancer-derived PC3

cells in response to LPA 85

2-S3 ERK1/2 oxidation in ovarian-cancer derived SKOV3 cells 86

2-S4 Observed oxidation of ERK in HeLa cells treated with EGF 87

2-S5 Observed oxidation of total and TEY-phosphorylated

ERK1/2 in response to Androgen agonist R1881 in

prostate-epithelium derived RWPE-1 cells 88

2-S6 Endogenous ERK1/2 oxidation from SKOV3 cells

inhibits activity towards Elk1 89

viii

3-1 rERK2 activity towards Elk1 is inhibited by adventitious

and H2O2-dependent oxidation 99

3-2 ERK1/2 oxidized during cell signaling events does not

make inter- or intramolecular disulfide bonds with itself

or other proteins 101

3-3 Identification of rERK2 cysteine sensitivity to sulfenylation

by H2O2 102

3-4 C159/252S (C2xS) double mutant rERK2 responds to

H2O2 differently than WT rERK2 104

3-5 C159S responsible for observed inhibition of ERK2 kinase activity

towards DRS-binding substrates 106

3-6 Dose-dependent, global changes in ERK2 kinase activity 113

ABSTRACT

The ERK1/2 pathway plays roles across eukaryotic biology by transducing extracellular signals into cell-fate decisions. One conundrum is in understanding how disparate signals induce specific responses through a common, ERK- dependent kinase cascade. While studies have revealed intricate modes to control ERK1/2 through spatiotemporal localization and phosphorylation dynamics, additional details of ERK1/2 signaling remain elusive. We hypothesized that fine-tuning of ERK1/2 signaling could occur by cysteine oxidation. We report that ERK1/2 is actively and directly oxidized by signal- generated H2O2 during proliferative signaling, and that oxidation occurs downstream of a variety of receptor classes tested in six cell lines. Furthermore, within the tested cell lines and proliferative signals, we observed that both phosphorylated and non-phosphorylated ERK1/2 undergo sulfenylation in cells, that there is a difference in the consistency of ERK1/2 oxidation between transformed and non-transformed cells, and that dynamics of ERK sulfenylation is dependent on whether or not cells are serum starved prior to stimulus. We also tested the effect of endogenous ERK1/2 oxidation on kinase activity and report that phospho-transfer reactions are drastically inhibited by oxidation, underscoring the importance of considering this alternative modification when assessing ERK1/2 activation. In light of these paradoxical results, we undertook a series of in vitro analyses to elucidate how oxidation could modulate ERK2 activity in a cellular environment. We identified one cysteine residue that is sensitive to in vitro oxidation by H2O2. C159 (Rat ERK2 numbering) was found to

x be highly sensitive to sulfenylation. We also have evidence that C252 undergoes sulfenylation at moderately higher concentrations of H2O2. Interestingly, neither of these cysteines are near the active site, indicating that observed inhibition by oxidation must occur by another mechanism than by direct inhibition of the kinase’s phospho-transfer activity. Indeed, both of these cysteines are part of important regulatory regions of ERK1/2, and their oxidation will likely affect protein-protein interactions with regulatory proteins and substrates and alter the spatiotemporal dynamics of ERK1/2 within cells. Finally, preliminary results indicate that oxidation of ERK1/2 actually activates kinase activity towards some substrates while inhibiting activity towards others, bolstering our hypothesis that oxidation is a more complex mode of ERK1/2 regulation than a simple switch to control ERK1/2 activity.

Chapter 1. Introduction

1.1 ERK is a key signal transducer

At the heart of a multitude of signaling pathways lies Extracellular signaling-regulated kinases 1 and 2, or ERK1/2. ERK2 was first identified due to its robust tyrosine phosphorylation in response to a variety of extracellular signals and in transformed cell lines; it was designated as p42 in the early 1980s (1-9).

Later, what was initially thought to be a separate but important kinase known as

Microtubule-associated protein kinase was shown to be one and the same as p42 (12). It wasn’t until it was first cloned by Melanie Cobb in 1990 that it was designated as ERK and shortly thereafter the MAPK acronym was modified to stand for “Mitogen Activated Protein Kinases” (14). It was clear in even the earliest work that ERK was fundamental to the signal transduction of growth factors. However, despite intensive study of ERK over the past 30+ years, there are still numerous unanswered questions about the regulation and functional consequences of ERK activation and its relation to human health and disease.

There are a variety of signal transduction molecules in cells. These include receptors such as Receptor Tyrosine Kinases (RTKs), G-protein coupled receptors (GPCRs), and hormone receptors (HRs), small GTPases such as Ras, small-molecule second messengers such as Ca2+ or diacylglycerol, and kinases and phosphatases that catalyze either the addition or removal of phosphate groups from biomolecules such as nucleotides, lipids, or proteins. Within this array of signaling molecules, ERK1 (p44) and ERK2 (p42) are Ser/Thr protein kinases that are members of the MAPK group within the CMGC subfamily of kinases consisting of cyclin-dependent kinases (CDKs), MAPKs, glycogen

2 synthase kinase 3 family of kinases (GSK3), and the CDC-like kinase family (Clk)

(15). MAPKs are known for their 3-5 tiered kinase cascade (Figure 1-1), in which a series of activations by phosphorylation occur down the cascade, serving to amplify the original signal. Within the ERK1/2 kinase cascade, Raf’s only known substrates are MEK1/2, and MEK1/2s’ only known substrates are ERK1/2.

However, ERK1/2 proteins have over 300 known substrates (16), indicating that these kinases are the primary workhorses within their respective MAPK cascade to elicit cellular responses to stimuli.

As shown in Figure 1-1, the ERK1/2 pathway is one of four established

MAPK cascades. The other known pathways are the p38, JNK, and ERK5.

Whereas p38 and JNK are primarily activated in response to stressors, ERK1/2 are primarily activated in response to extracellular signals to initiate proliferation, differentiation, survival, and a host of other cellular responses such as migration

(17,18). Alignment among the common MAPK homologues (ERK1/2, p38α,

JNK1, and ERK5) reveals surprising divergence among the sequences of these enzymes (Figure 1-2). However, several key motifs, with known and unknown functions, are shared among these MAPK family members, including several conserved cysteine residues.

Figure 1-1: MAPK signaling cascades. ERK1/2 signaling pathway is one of four currently identified Mitogen-activated protein kinase (MAPK) pathways. The other pathways include p38 and JNK, which are primarily activated in response to stresses, and ERK5, which appears to respond to both mitogens and stress (10). Each cascade has a 3-tier core that consists of a Mitogen-activated protein kinase kinase kinase (MAP3K), which activates by phosphorylation the Mitogen-activated protein kinase kinases (MAPKK). MAPKK are dual-specificity kinases which activate the MAPKs by phosphorylation on Thr and Tyr residues on the L12 activation loop (sometimes referred to as the activation lip (11)). MAPKs then go on to phosphorylate any number of substrates in both the cytosol and nucleus to induce specific cell responses. Most MAPKs are also linked to a set of kinases, referred to as Mitogen- activated protein kinase-activated protein kinases (MAPKAPK), that have their own set of substrates. Select MAPK cascades also have a set of MAP4Ks that are necessary to activate, by phosphorylation, the MAP3Ks. The ERK1/2 pathway, however, is primarily initiated by activation of the small GTPase Ras, although under certain signaling conditions Protein Kinase C (PKC) can be considered a MAP4K and can directly activate the MAP3K Raf proteins, bypassing the need for Ras activation.

ERK2 MAAAAA------AGAGPEM----VRGQVFDVGPRYTNLSY 30 ERK1 MAAAAAQG-G--GGGEPRRTEGVGPG------VPGEVEM----VKGQPFDVGPRYTQLQY 47 P38α ------MSQERPTFYRQELNKTIWEVPERYQNLSP 29 JNK1 ------MSRSKRDNNFYSVEIGDSTFTVLKRYQNLKP 31 ERK5 MAEPLKEEDGEDGSAEPPGPVKAEPAHTAASVAAKNLALLKARSFDVTFDVGDEYEIIET 60 : : * .* :.

ERK2 IGEGAYGMVCSAYDNVNKVRVAIKKIS-PFEHQTYCQRTLREIKILLRFRHENIIGINDI 89 ERK1 IGEGAYGMVSSAYDHVRKTRVAIKKIS-PFEHQTYCQRTLREIQILLRFRHENVIGIRDI 106 P38α VGSGAYGSVCAAFDTKTGLRVAVKKLSRPFQSIIHAKRTYRELRLLKHMKHENVIGLLDV 89 JNK1 IGSGAQGIVCAAYDAILERNVAIKKLSRPFQNQTHAKRAYRELVLMKCVNHKNIIGLLNV 91 ERK5 IGNGAYGVVSSARRRLTGQQVAIKKIPNAFDVVTNAKRTLRELKILKHFKHDNIIAIKDI 120 :*.** * *.:* .**:**: *: .:*: **: :: ..*.*:*.: ::

ERK2 IRAPT-IEQMKDVYIVQDLMETDLYKLLK-TQHLSNDHICYFLYQILRGLKYIHSANVLH 147 ERK1 LRAST-LEAMRDVYIVQDLMETDLYKLLK-SQQLSNDHICYFLYQILRGLKYIHSANVLH 164 p38α FTPARSLEEFNDVYLVTHLMGADLNNIVK-CQKLTDDHVQFLIYQILRGLKYIHSADIIH 148 JNK1 FTPQKSLEEFQDVYIVMELMDANLCQVIQ--MELDHERMSYLLYQMLCGIKHLHSAGIIH 149 ERK5 LRPTVPYGEFKSVYVVLDLMESDLHQIIHSSQPLTLEHVRYFLYQLLRGLKYMHSAQVIH 180 : :..**:* .** ::* :::: * ::: :::**:* *:*::*** ::*

ERK2 RDLKPSNLLLNTTCDLKICDFGLARVA-DPDHDHTGFLTEYVATRWYRAPEIMLNSKGYT 206 ERK1 RDLKPSNLLINTTCDLKICDFGLARIA-DPEHDHTGFLTEYVATRWYRAPEIMLNSKGYT 223 P38α RDLKPSNLAVNEDCELKILDFGLARHTD------DEMTGYVATRWYRAPEIMLNWMHYN 201 JNK1 RDLKPSNIVVKSDCTLKILDFGLARTAG-----TSFMMTPYVVTRYYRAPEVILG-MGYK 203 ERK5 RDLKPSNLLVNENCELKIGDFGMARGLCTSPAEHQYFMTEYVATRWYRAPELMLSLHEYT 240 *******: :: * *** ***:** :* **.**:*****::* *.

ERK2 KSIDIWSVGCILAEMLSNRPIFPGKHYLDQLNHILGILGSPSQEDLNCIINLKARNYLLS 266 ERK1 KSIDIWSVGCILAEMLSNRPIFPGKHYLDQLNHILGILGSPSQEDLNCIINMKARNYLQS 283 P38α QTVDIWSVGCIMAELLTGRTLFPGTDHIDQLKLILRLVGTPGAELLKKISSESARNYIQS 261 JNK1 ENVDLWSVGCIMGEMVCHKILFPGRDYIDQWNKVIEQLGTPCPEFMKKLQ-PTVRTYVEN 262 ERK5 QAIDLWSVGCIFGEMLARRQLFPGKNYVHQLQLIMMVLGTPSPAVIQAVGAERVRAYIQS 300 : :*:******:.*:: : :*** .::.* : :: :*:* :: : .* *: .

ERK2 LPHKNKVPWNRLFPNA------DSKALDLLDKMLTFNPHKRIEVEQALAHPYLE 314 ERK1 LPSKTKVAWAKLFPKS------DSKALDLLDRMLTFNPNKRITVEEALAHPYLE 331 P38α LTQMPKMNFANVFIGA------NPLAVDLLEKMLVLDSDKRITAAQALAHAYFA 309 JNK1 RPKYAGYSFEKLFPDVLFPADSEHNKLKASQARDLLSKMLVIDASKRISVDEALQHPYIN 322 ERK5 LPPRQPVPWETVYPGA------DRQALSLLGRMLRFEPSARISAAAALRHPFLA 348 : :: * .** :** :: ** . ** * ::

ERK2 QYYDPSDEPIAEAP-FKFDMELDDLPKEKLKELIFEETARFQPGYRS------360 ERK1 QYYDPTDEPVAEEP-FTFAMELDDLPKERLKELIFQETARFQPGVLEAP------379 P38α QYHDPDDEPVAD-P-YDQSFESRDLLIDEWKSLTYDEVISFVPPPLDQEEMES------360 JNK1 VWYDPSEAEAPPPKIPDKQLDEREHTIEEWKELIYKEVMDLEERTKNGV--IRGQPSPLG 380 ERK5 KYHDPDDEPDCAPP-FDFAFDREALTRERIKEAIVAEIEDFHARREGIRQQIRFQPSLQP 407 ::** : :: :. *. * :

ERK2 ------ERK1 ------P38α ------JNK1 AAVINGSQHPSSSSSVNDVSSMSTDPTLASDTDSSLEAAAGPLGCCR------427 ERK5 VASEPGCP------DVEMPSPWAPSGDCAMESPPPAPPPCPG 443

Figure 1-2: MAPK Alignment. Human ERK1/2, p38α, JNK1, and ERK5 sequences were aligned on the Uniprot server using ClustalO algorithm. “*” indicates conserved residues, “:” indicates highly conserved substitutions, and “.” indicates residues that are conserved among most homologues. The T and Y phosphorylated by upstream MAPKK are conserved (bold and underlined). The SPS motif (below the TxY motif) is conserved among ERK1/2 and ERK5. Several cysteine residues, including C161 (human ERK2 numbering) and C216 are also conserved.

Within the sphere of proliferation, ERK1/2 have been shown to be vital

components for healing, for organ homeostasis, and for the growth and

development of an organism (19). For example, lung tissue-specific deletion of

MEK1 in MEK2 null mice led to several lethal abnormalities in lung growth and

development (20), and the utilization of a fluorescence resonance energy transfer

(FRET)-based ERK1/2 kinase activity reporter expressed in the skin of living

mice revealed bursts of ERK1/2 activity in areas of epidermal growth and healing

(21). However, ERK1/2’s role in proliferation renders it a frequent contributor to

cancer initiation and growth (16,22). Although ERK1/2 is rarely identified as an

oncogene itself, oncogenic forms of its upstream activators Ras and Raf are

found in up to 35% of all human cancers, and the ERK1/2 cascade pathway is

found to be upregulated and hyperactivated in up to 90% of human cancers (16).

Thus, the ERK1/2 cascade is included in highly sought-after targets for the

development of cancer therapeutics.

ERK knockout (KO) mice have shed light on the fundamental role that

ERK1/2 proteins play not only in proliferation, but also in differentiation. Ablation

of ERK2, which is normally expressed in higher abundance than ERK1 (23), is

embryonically lethal; the mesoderm never develops into the early embryo (24).

ERK1 KO, however, is not embryonically lethal, although the mice have impaired

T-cell differentiation that originates with altered immune stem cells in the thymus

(25), and hindered adipogenesis in high-fat diet fed mice (26). These initial studies suggested non-redundant roles between ERK1 and ERK2, and this hypothesis was corroborated by reports that the two different isoforms exhibit different substrate specificities in vitro (27,28). However, a recent elegant study in which ERK1 was over-expressed in ERK2 KO mice reported the normal and healthy development of these mice (29), suggesting ERK1/2 dosage is more important in the development of an organism than any differences between

ERK1 and ERK2 isoforms. However, it should be noted that this study was done with mice living in ideal, non-stress conditions, and differences between WT and

ERK1-overexpressed mice may be manifested under various stress conditions.

Furthermore, it has not yet been tested if over-expressing ERK2 in ERK1 KO mice would restore adipogenesis in high-fat diet mouse models of for the ability of the thymus to produce mature T-cells.

While ERK1/2 have been primarily studied for their vital roles in proliferation and differentiation, there are other cell responses to stimuli that require or are facilitated by ERK1/2, including migration (30,31), progression throughout various stages of the cell cycle (32-34), autophagy (35,36), metabolism (37-39), insulin secretion and response (40,41), and even apoptosis

(42,43). These wide-ranging cell responses to a variety of signals brings up a conundrum: how is signaling specificity achieved between these varied signals when sharing a core signaling pathway (Figure 1-3A)? Over the years, this conundrum has led researchers to earnestly seek a better understanding of how

ERK1/2 is differentially regulated between these various ERK-dependent cell responses.

Figure 1-3: Signal-specific context to induce distinct cellular responses through ERK1/2 signal cascade. A) ERK1/2 signal transduction is induced by various different extracellular signals to elicit varied cellular responses to stimuli. Research has identified several alternate behaviors and mechanisms by which ERK1/2 is differentially regulated between signals. These include differences in the temporal dynamics of ERK1/2 TEY-phosphorylation and SPS-phosphorylation to induce nuclear translocation. However, the mechanistic details of how these regulations occur in different contexts has yet to be resolved. B) A complete understanding of how ERK1/2 is differentially regulated will enable to development of context-specific therapeutics. This is necessary to be able to inhibit ERK1/2 pathway signaling in unhealthy tissue (i.e. cancer) without inhibiting ERK1/2 necessary for the homeostasis of healthy tissue (i.e. nervous system, immune system, etc.)

Because of the ERK1/2 proteins’ role in cancer and other diseases, researchers have long sought to target components of the ERK1/2 pathway. Yet, because of its ubiquitous expression and vital role in multiple signaling pathways, total inhibition of ERK1/2 signaling can lead to drastic consequences such as depression (44,45), altered glucose metabolism (41,46,47), and impairments in memory (48,49), healing (50), and the immune system (25,51,52). It is therefore absolutely vital that ERK1/2 differential regulation between different signals is

8 fully understood (Figure 1-3A). If distinct modes of ERK1/2 regulation that are specific to a subset of signals are identified, then it is possible to therapeutically target ERK1/2 under those specific circumstances, thereby bypassing total-ERK inhibition and its side-effects (Figure 1-3B)

An excellent example of signal-specific inhibition is found in Rony Seger’s recent work, where they developed peptide-based inhibitors to ERK1/2 nuclear translocation (53). These inhibitors stopped the proliferation of melanoma cell lines and xenografts in animal models, without affecting the viability of non- transformed cell lines. Thus, it is clear that the more that is understood about the context of signal-specific ERK1/2 regulation, there more power scientists have in targeting the ERK1/2 pathway.

1.2 Established modes of ERK1/2 regulation

1.2.1 Activation-loop phosphorylation on TEY-motif as an activity switch

The primary established mode of regulation is through phosphorylation;

ERK1/2 in its native, unmodified state is unable to catalyze the phospho-transfer reaction from ATP to a protein substrate (18). ERK1/2 is the final stage in its respective MAPK cascade; it is phosphorylated on T183 and Y185 (rat ERK2 numbering, see Figure 1-4 for rat ERK2 alignment with human ERK1/2) by the dual-specificity kinases MEK1/2. MEK1/2, too, are activated by phosphorylation by Raf kinases, and Raf kinase activation is initiated either by PKC (54) or, more canonically, through its interactions with the small GTPase Ras (17). Ras is a small, membrane bound GTPase that is recruited to activated receptors, most

Figure 1-4: Rat ERK2 and human ERK1/2 alignment with secondary structure assignments. Secondary structure assignments are based crystallographic structures of Rat ERK2 in Zhang et al. (10) and mouse PKA solved by Knighton et al (13). Blue secondary structure elements indicate regions in the N-terminal lobe, with lighter blue coloring for β-strands and darker blue for α-helices. Peach coloring indicates elements in the C-terminal lobe with lighter shades for β- strands and darker shades for α-helices. Each loop region is numbered (L0-L16) which include secondary structure elements that are not in the canonical protein kinase structure. TEY and SPS phosphorylation motifs are underlined with modifiable residues in bold. Cysteine residues of particular interest that are mentioned in the text are double underlined, solvent-exposed cysteines are blue, and all cysteines are bolded. Rat ERK2 is the common species homologue used in structural and biochemical studies, and residue numbering throughout the text will refer to Rat ERK2 numbering. Rat ERK2 and Human ERK2 share 98.6% sequence identity as calculated by ClustalO algorithm on UniProt server; Human ERK1 and ERK2 share 82.1% sequence identity. often by GRB2, and the Guanine-nucleotide exchange factor SOS replaces GDP with GTP in the Ras protein, thereby activating Ras and initiating the ERK1/2 signaling cascade (17) (Figure 1-1).

The dual-phosphorylation of ERK1/2 on T183 and Y185, hereafter referred to as the TEY motif, induces a conformational change in the activation loop, opening up the active site of ERK and increasing the overall flexibility and dynamics of the kinase, particularly allowing movement between the N and C lobes of the structure (55-57) (Figure 1-3), an area which contains the ATP- binding pocket and active site residues (Figure 1-5). These changes induce a 3- order of magnitude increase in the catalytic activity (58), essentially transitioning the kinase from an “off” to an “on” state.

In general, researchers consider that the TEY-phosphorylated form of

ERK1/2 is active, and many will report that a certain stimulus or treatment changes the activity of ERK1/2 simply by measuring, via western blot, the

Figure 1-5: Structural analysis of ERK1/2 docking sites and NTS. A) Cartoon representing positioning of the docking motifs (dark blue), active site (light tan), ATP binding region (light orange), activation loop, Nuclear Translocation Site (NTS – Dark Brown). Yellow dots represent C159 located in the DRS docking motif, C164 facing ATP binding pocket, and C252 near the FRS and NTS. B) Ribbon representation of active Erk structure (PDB 2ERK). Erk structure oriented and colored to match reader’s view in A. All cysteine residues are shown as yellow ball-and-stick representations. Phosphate groups on Y183 and T185 are also shown. Only D147 is colored orange for the active site because this is the primary catalytic residue necessary for phospho-transfer to a serine/threonine on substrate. C) Surface representation of DRS docking motif over ribbon diagram of ERK. Rotated about 90ᵒ from viewing angle in B. Carbons are grey, negative charges are red and positive charges are blue. The hydrophobic residues from the consensus sequence ((Arg/Lys)2-3-X2-6--X-, Φ representing hydrophobic residues) of the insert into the grooves separated by C159. Asp 165 and 169, located on a short helical segment left of the hydrophobic grooves facilitate electrostatic interactions with the basic residues on the DRS recognition sequence. D) Surface representation of FRS docking motif, and with same colorings for the surface as in C. E) Nuclear Translocation Signal – consisting of L14 and α1L14. The two serine residues on L14 (shown) are phosphorylated by Casein Kinase 2 (CK2) which is necessary and sufficient for translocation to the nucleus. The asp residues on the α-helix are combine with the negative charges introduced by the phosphate groups to elicit ERK1/2 interaction with importin 7/9. C252 residue on the NTS structure is highlighted.

12 phosphorylation state of the TEY motif without reporting an actual measurement of kinase activity (see references (37,59,60) as select examples of where

ERK1/2 “activation” is primarily reported by TEY-phosphorylation status). It should be noted, however, that Templeton et al. reported that the oxidation of p38, another MAPK, inhibits p38 kinase activity even when phosphorylated on its homologous activation loop (61). Furthermore, a recent study of ERK1/2 activity of induced-ischemic rat brains reported that the extent of phosphorylation did not match the measured kinase activity (62), indicating that TEY-phosphorylation should not be considered as the sole factor in determining ERK1/2 kinase activation in response to stimuli.

As described in section 1.1, ERK1/2 TEY-phosphorylation occurs downstream of a plethora of various signals that have distinct cellular responses

(Figure 1-3). Thus, regulation of ERK1/2 by a simple “on” and “off” mechanism is insufficient to properly direct ERK1/2 activity to transduce signals into appropriate responses. Extensive research has gone into elucidating these intricate modes of

ERK1/2 regulation, and while many discoveries have helped clarify signal- specific modes of ERK1/2 regulation, details of how these modes work are still elusive. Many key aspects of what is known and not known will be detailed in the remainder of section 1.2.

1.2.2 Temporal regulation of TEY-phosphorylation of ERK1/2

One established mode of regulation is through modulation of temporal dynamics of ERK1/2 TEY-phosphorylation. Murphy et al. demonstrated that while

Epidermal Growth Factor (EGF) and Platelet-derived Growth Factor (PDGF) both

13 induce initial TEY-phosphorylation of ERK1/2 in Swiss-3T3 cells, PDGF elicited

S-phase entry but EGF did not (63). The authors then analyzed the extent of

TEY-phosphorylation over time and found that PDGF induced a sustained phosphorylation whereas EGF induced a transient phosphorylation. This difference in temporal dynamics of TEY-phosphorylation was responsible for the different cell response, which was mediated by differences in ERK1/2’s ability to phosphorylate one particular transcription factor, c-Fos. In these cells, c-Fos was not constitutively expressed under serum-depleted conditions. Upon ERK1/2

TEY-phosphorylation resulting from either EGF or PDGF, transcription and translation of c-Fos protein was induced. However, without phosphorylation by

ERK1/2, c-Fos is unstable and quickly degraded. When ERK1/2 exhibited sustained TEY-phosphorylation, c-Fos was subsequently stabilized and able to modify transcription and induce S-phase entry. When ERK1/2 TEY- phosphorylation was transient, however, c-Fos could not be stabilized by

ERK1/2-dependent phosphorylation and was quickly degraded, leading to an alternate cellular response. However, it was not shown how the same cell line can facilitate both a transient and sustained TEY-phosphorylation of ERK1/2; it likely is not due to differences between the expression of phosphatases because of the short time frame in which ERK1/2 is dephosphorylated in response to EGF compared to the induced expression of the ERK1/2-specific phosphatase MKP3

(64).

1.2.3 Nuclear translocation of ERK1/2

Another well-studied mode of regulation is the translocation of ERK1/2 in response to proliferative signals, where ERK1/2 translocates to the nucleus within 5-20 minutes of stimulus (65,66) and may be retained in the nucleus from minutes to hours after stimulation. This translocation is a way to control substrate specificity by sequestering TEY-phosphorylated ERK1/2 away from cytoplasmic substrates while enabling kinase activity towards nuclear substrates. Although this translocation was evident for many years, it wasn’t until the past 5-8 years that researchers were able to elucidate the molecular details of how translocation is facilitated (66-72).

Instead of a canonical nuclear localization sequence, Chuderland et al. discovered what they termed the “Nuclear Translocation Signal” (NTS) located within the MAPK insert region of ERK1/2 (Figures 1-4, 1-5) (71). Within this sequence are two serine residues separated by a proline residue, hereafter referred to as the SPS-motif, that get phosphorylated by Casein Kinase 2 (CK2)

(68) after ERK1/2 is released from anchor proteins via TEY-phosphorylation. The negative charges introduced by the phosphate groups on the NTS work with aspartate residues in the same region to enable ERK1/2 binding with importin

7/9, which transports ERK1/2 into the nucleus. What is not yet understood, however, is exactly why some signals induce this SPS-phosphorylation and others do not.

There are other aspects of controlling ERK spatial dynamics that are important for ERK1/2 regulation besides translocation to the nucleus. For

15 example, TEY-phosphorylated ERK is found at the leading edge of migrating cells and this localization is necessary for migration (30,31), although it is yet not known what anchors the TEY-phosphorylated ERK to the leading edge. There are also several scaffold proteins, including IQGAP, β-arrestin, and Pea-15, that anchor ERK1/2 near receptors, essentially priming the cell to quickly respond to extracellular signals by keeping a population of ERK1/2 near the receptors and upstream activators. Even MEK1/2 proteins act as anchors to sequester ERK1/2 in the cytoplasm; when ERK1/2 is overexpressed, stoichiometry of ERK: MEK leads to erroneous ERK1/2 relocation to the nucleus (66,68-70). It should be noted that scaffold proteins can have profound and varying effects on ERK signaling. For example, the nature of the interaction between β-arrestin and its coupled GPCR determines whether or not β-arrestin-associated ERK1/2 remains localized by the receptor during the course of the signal or is released from β- arrestin to freely diffuse throughout the cytosol and translocate to the nucleus

(73).

While TEY-phosphorylation usually disrupts ERK1/2 interactions with scaffold proteins, this is not always the case. For example, Pea-15 can still interact and anchor ERK1/2 independent of ERK1/2 proteins’ phosphorylation state (74-76) However, it is not yet known why TEY-phosphorylation facilitates

ERK1/2 release from scaffolds, or how ERK is recruited to the right scaffolds at the right time.

1.2.4 ERK1/2 regulation through protein-protein interactions

Much of the regulation of ERK1/2 in cells roots back to its protein-protein interactions, particularly through two distinct docking motifs. These motifs are known as the D-recruitment site (DRS) and the F-recruitment site (FRS) (Figure

1-5). The DRS is found near the hinge region between the N and C lobes of

ERK1/2, comprising parts of αE, L11 and L16, and β7 and 8, whereas the FRS is made up of residues from αG, α1L14 and α2L14 from within the MAPK insert

(Figure 1-4) (77-86).

The consensus sequence that binds to the DRS is (Arg/Lys)2-3-X2-6-Φ-X-Φ, where the R/K interacts with Asp 316 and Asp 319 (rat ERK2 numbering) in L16 of ERK1/2, and the Φ designate hydrophobic residues that interact with hydrophobic pockets on either side of a ridge created by C159 and H123 (78,86-

89). In spite of the discovery of this loose consensus sequence, it should be noted that some proteins, such as RSK, that are known to interact with ERK1/2 via the DRS, do not perfectly conform to this consensus sequence (87), but this has not been addressed in the literature. The DRS enables interactions with

MEK1/2 (90-93), MAPK-specific phosphatases (85,94-99), scaffold proteins and anchors, and many substrates (77).

The sequence that binds to the FRS is FXFP, although it has been shown that either of the phenylalanines can be replaced with another large hydrophobic amino acid (100). These large hydrophobic residues insert into a hydrophobic cleft on the MAPK insert. It appears that this docking domain is primarily used for interaction with substrates, and particularly transcription factors, although it has

17 also been implicated in interactions with other regulatory proteins such as

MEK1/2 (77).

Both of these motifs help determine substrate specificity. While ERK1/2 can technically phosphorylate any serine or threonine followed by a proline, with a slight preference for PXS/TP sequence (101,102), the rate of ERK phosphorylation is significantly enhanced when a substrate contains one of these docking sites (77,102,103) by increasing the successful sampling of phosphorylation motifs into the ERK active site (79). Furthermore, the anchor

Pea-15 has been shown to bind to and hide the DRS (75,76), essentially rendering ERK unable to phosphorylate DRS-interacting substrates, although it has yet to be tested if ERK is still able to phosphorylate FRS-interacting substrates while bound to Pea-15.

Beyond modulating substrate specificity, the DRS has a significant impact on ERK1/2 interactions with regulatory proteins (77). For example, MPK3 is a dual-specificity phosphatase that has shown to be one of the primary ERK1/2- specific phosphatases in vivo. In vitro studies have shown that when the DRS- binding consensus sequence is mutated on MKP3, it is unable to dephosphorylate ERK2 (99) despite retaining general phosphatase activity. Thus, modulation of the DRS on ERK1/2 could be one of the mechanisms by which cells could protect ERK1/2 from dephosphorylation during signal transduction.

1.2.5 Remaining questions regarding ERK1/2 regulation

The above examples, along with others not reviewed here, illustrate that evolution has led to the development of sophisticated and intricate mechanisms to control ERK1/2 activity to induce distinct responses, but there are still detailed questions remaining to fully understand how these modes of regulation take place. In particular, why do some signals induce the SPS-phosphorylation, resulting in nuclear translocation, and other signals do not? What are the exact mechanisms that recruit and release ERK1/2 from sequestering proteins? Why is

ERK1/2 protected from dephosphorylation during one signal while rapidly undergoing dephosphorylation during another signal? Despite detailed molecular knowledge about different docking sites, it is still unclear how TEY- phosphorylated ERK1/2 manages to discriminate between a mixed pool of substrates. These are all questions that are vital to answer in order to develop therapeutics that are targeted towards the ERK1/2 pathway under specific signaling conditions.

An alternative mode of molecular regulation that would induce sufficient changes in ERK1/2 surfaces and structure to elicit changes in protein-protein interactions would be through additional post-translational modifications (PTMs).

A perfect example of a recently discovered PTM is described above in the SPS- motif phosphorylation. Without this phosphorylation, ERK1/2 is unable to bind to importin7/9 and therefore is unable to translocate to the nucleus (68,71). It is likely that there are alternate, yet-to-be described PTMs that modulate ERK protein-protein interactions and activity. Of particular note is the possibility of

19 cysteine modifications. Although protein-tyrosine phosphatases (PTPs) were the initial signaling protein family identified to be sensitive to oxidation, there are a growing number of examples in the literature of direct oxidation of kinases (Table

1-I). There are a handful of examples of regulatory MAPK oxidation, particularly of ERK1/2, in the literature (Table 1-II) describing modulation of ERK1/2 activity, either directly or indirectly, by Reactive Oxygen Species (ROS) or Reactive

Nitrogen Species (RNS). These data, taken into consideration with the substantial body of work detailing the important role of ROS in signal transduction, bolsters the hypothesis that a vital missing component to understanding ERK1/2 regulation could be signal-dependent cysteine modification.

Table 1-I: Redox sensitive kinases Kinase Mod Type Modified Site Effect Ref MEKK1 PS-SG C1238 - (104) PKA C PS-SG C199 +/- (105) PKC PS-SG ND - (106,107) PKC1 PS-SG ND - (106) PKC2 PS-SG ND - (106) PKC PS-SG ND - (106) PKC PS-SG ND +/- (106) PKC PS-SG ND - (106) PKC PS-SG ND - (106) PKC PS-SG ND - (106) RPS6KA1 PS-SG C223 - (108) CAMK1 PS-SG C179 - (109) c-ABL PS-SG ND - (110) EGFR SOH C797 + (111) MAPK1 SOH; SO2H C38,159,214 -/NC (112-114) MAPK9 SOH; SO2H C41,137,177,222 NC (112) MAPK14 SOH; SO2H C162 NC (112) SRC S-S C245,277,487 +/- (115-117) LYN S-S C466 + (118) c-RET S-S C376 + (119) TYK1 S-S ND + (120) VEGFR2 S-S ND + (121) FGFR1 S-S C488 - (116) AKT1 S-S C297,311 NC (122) AKT2 S-S C124 - (123,124) PKG1 S-S C42 + (125) ATM S-S C2991 +/NC (126,127) PS-SG: glutathionylation; SOH: sulfenic acid; SO2H: sulfinic acid; S-S: disulfide (either intra- or intermolecular); ND: not determined; +, -, NC: increase, decrease, or no change in kinase activity, respectively. Table adapted with permission from Dr. Robert Newman.

Table 1-II: MAPK oxidation in the literature Modification: Effect On: Reference Kinase Context Treatment Type Site Kinase Cell response to stimulus Increases ERK phosphorylation a c ERK1/2 In vitro 0.1-1 μM H2O2 R-SO2/3 C38, 214 by MEK; no ND change in ERK activityb Decreases ERK phosphorylation Galli et al. ERK1/2 In vitro 10 μM H2O2 ND ND by MEK; no ND (128) change in ERK activity Changes in Cells increase rate of proliferation ERK1/2 spatiotemporal Transfecte in response to 1 μM H2O2 and WT, subcellular d LP07 1-50 μM H2O2 ND ND undergo apoptosis in response to C38A, locations, cells 50 μM H2O2. Cell response link to C214A disrupted MEK ERK1/2 oxidation was not tested. interactions Brain ERK2 cytosolic 1 mM GSNO R-SNO C161d ND ND Paige et al. lysates (129) sEnd.1 Constitutive NOS ERK2 R-SNO ND ND ND cells activity Kaplan et HUVEC ERK1/2 VEGF R-SOH ND ND ND al. (130) cells Cells undergo apoptosis in Luanpitong response to doxorubicin. The link ERK1/2 HaCat cells Doxorubicin R-SOH ND ND et al. (131) between ERK oxidation and cell response was not tested Note: This publication does not identify ERK1/2 oxidation in this ischemia model. However, their Rat focal +/- preconditioning Takahashi finding that ERK activity did not ERK1/2 ischemia before permanent ND ND ND et al. (62) always match ERK TEY- model brain ischemia phosphorylation supports the notion that oxidation could happen and affect kinase activity in vivo

Modification: Effect On: Reference Kinase Context Treatment Type Site Kinase Cell Response Note: Researchers found that C159, other crystals with ERK2 and ATP would βME cysteine unreported Facilitated Zhang et al Protein β-mercaptoethanol only form in presence of βME, but ERK2 adducts, R- cysteinese, packing in (132) crystals (βME) that initial crystals had “several SOH C63 (R- presence of ATP solvent-exposed cysteines formed SOH) disulfide bonds to βME.” C183 S-nitrosylation on ERK1 MCF-7 Decreases TEY- ERK1 1-2 mM SNP R-SNO C183f appears to lead to MCF-7 cell cells phosphorylation apoptosis in response to SNP Feng et al. 293T cell ERK1/2 Cys-NO or SNP R-SNO ND ND ND (133) lysates 6 hours post 500 MCF-7 ERK1/2 μM H2O2 or 50 R-SNO ND ND ND cells ng/mL TNF-α Inhibits kinase 10 mM H2O2, 100- activity as Templeton 250 μM ND, but C119, p38 HeLa cells measured by in ND et al. (61) prostaglandin J2 reversible C162g vitro kinase assay (PGJ2) towards ATF2 RKO colon C161 Yang et al ERK2 adenocarci 500 μM H2O2 R-SOH (Human ND ND (134) noma cells ERK2) a For Identification of cysteines, authors treated recombinant GST-ERK2 with H2O2, followed by protein drying, ZipTip, and ionization into the LCQ mass spectrometer. Authors never blocked free thiols by alkylation, thus observed oxidations could be a result of artificial oxidation during drying or ionization processes. b Activity was measured towards general substrate Myelin Basic Protein. c ND = Not Determined d Human ERK2 numbering (See Figure 1-3) e When authors first observed several βME adducts, they soaked the crystal with DTT which removed all adducts except for C159, which was involved in crystal packing. f Human ERK1 numbering (See Figure 1-3) g Cysteine IDs were solved by making point mutations in p38α and testing if mutants were captured by PROP method developed in publication. Although mutants containing only C119 restored p38 pulldown approximately 55% of WT, it appears that other cysteines make contributions to the pulldown of p38α in PGJ2-treated HeLa cells.

1.3 ROS in signal transduction

Historically, the role of ROS in biology was viewed as responsible for the detrimental side effects of aerobic life, their presence damaging lipids, proteins, and DNA (135,136). While oxidative stress is a major factor in a host of diseases and increases with age (135,137-139), research over the past twenty years has shown that ROS play an essential role in normal physiology by acting as second messengers during signal transduction (136,140-144). As second messengers,

ROS react with protein thiols to modify the protein’s surface and structure

(11,145), leading to changes in protein function, thereby affecting signal transduction and the eventual biological response.

The landmark study that first demonstrated a vital role for ROS in signal transduction was by Sundaresan et al. in 1995 (146), where they determined that

H2O2 was produced in response to PDGF in primary rat vascular smooth muscle cells (VSMCs). Not only did catalase loading of cells lead to a decrease in fluorescence of the ROS-sensitive fluorophore 2',7'-dichlorofluorescin (DCF), indicating near-complete removal of intracellular H2O2, but it also resulted in an overall decrease in tyrosine phosphorylation, DNA synthesis, and migration.

Thus, H2O2 production was necessary for the cells’ normal physiological response to PDGF: proliferation. Similar results have been reported in a host of other cell types and in response to a variety of extracellular signals, including

LPA in ovarian and prostate cancer cells (147-149), VEGF on the migration of

HUVECs (130), 17-beta-Estradiol response in ovarian adenocarcinoma cells

(150), chondrocyte response to fibronectin fragments (151), and the nuclear reprogramming of MEFs into pluripotent stem cells (152).

These ROS are generated by the mitochondria (153) or by membrane-

•- bound NADPH oxidase (NOX) complexes that generate superoxide (O2 ) or, in

•- the case of Nox4, may generate H2O2 directly (154-156). O2 generated by NOX dismutates either spontaneously or through superoxide dismutase (SOD) to

H2O2, which is considered to be one of the primary ROS that is able to react with protein cysteines to act as a second messenger (141,145,154,156). However, in order for ROS, such as H2O2, to act as a second messenger, it needs to be able to exist in the cell long enough to react with protein thiols. Evolution in an aerobic environment has led to substantial antioxidant defenses, including small molecule-thiols such as glutathione, along with enzymes like catalase, peroxiredoxins (Prxs), and glutathione peroxidases (GPXs) that catalyze the removal of H2O2. In recent years, research is bringing to light how these antioxidant defenses are bypassed, or used, to potentiate ROS in signal transduction. In particular, mammalian Prxs are prone to inactivation by hyperoxidation when local H2O2 rise to certain thresholds (157-161). Although hyperoxidation is normally irreversible, Prx hyperoxidation at the level of sulfinic acid (see below) can be reversed by the ATP-dependent reduction via the enzyme sulfiredoxin (162). Thus, hyperoxidation of Prxs is a signal-induced switch which may allow for the accumulation of localized H2O2 to persist long enough to react with cysteine residues of signal transduction proteins such as phosphatases and kinases (157). Indeed, there are a handful of studies to date

25 that have established that signal-dependent, localized pools of ROS and protein oxidation occur near endosomes which have been termed “redoxosomes” (163).

Alternatively, recent evidence suggests that Prxs can also be rendered inactive by phosphorylation, thereby allowing for the accumulation of H2O2 for their use in signaling (160,164). Additionally, there is also a recent study demonstrating that

Prxs themselves can act as oxidants to modify the signaling protein Stat3 (165).

An important aspect in elucidating the impact of ROS second messengers on signal transduction is to identify proteins that undergo cysteine oxidation during signaling events. There are various methods that have been developed to probe for cysteine-oxidized proteins (166). Two of particular note utilize the labile nature of cysteine redox chemistry (Figure 1-6) (11,145). When cysteines react with peroxides, or a narrow set of other ROS (145), they first form a metastable species known as sulfenic acid (R-SOH). The sulfenic acid will quickly react with reduced thiols (R-SH) to form disulfide bonds with small molecule thiols such as glutathione, with other cysteine residues on other proteins (intermolecular disulfide), or with cysteine residues within itself (intramolecular disulfide) (11).

Sulfenic acids can also react with the amide nitrogen of the protein backbone to form what is known as a sulfenamide, or they can react with additional ROS molecules to form the hyperoxidized, and irreversible, modification to sulfinic (R-

SO2) or sulfonic (R-SO3) acid, as discussed with Prxs above. It should also be noted that cysteine residues can react with other oxidant molecules, such as

Figure 1-6: Cysteine oxidation chemistry. Protein cysteines, when ionized to the thiolate form, are able to react with ROS such as H2O2, organic hydroperoxides, hypochlorous acid, and even peroxynitrite to form the meta-stable sulfenic acid. Sulfenic acid subsequently reacts with other thiols, such as glutathione or other cysteine thiols, to form disulfides. Alternatively, the neighboring amide nitrogen from the peptide backbone can attack the sulfenic acid sulfur to form sulfenamide (bottom), or an excess of ROS can react with the sulfenic acid to form the irreversible oxidations of sulfinic or sulfonic acid (red box). Reprinted with permission from (11)

RNS or reactive sulfur species (RSS), but these are not the focus of the current study and go through a different oxidation pathway as outlined in Figure 1-6.

Two key techniques for labeling oxidized cysteines are to target either the intermediate sulfenic acid or the reducible endpoint oxidation state, such as the disulfide or sulfenamide species. Several probes have been developed that specifically alkylate sulfenic acids (145,166,167), labeling them with biotin, with a fluorescent reporter, or with a click-chemistry molecule adaptable to the needs of

27 the researcher. The advantage with targeting the sulfenic acid intermediate is that, when a protein is identified using this technique, the researcher can know exactly at what time points active oxidation with peroxides were taking place on that protein in situ. However, the transient nature of sulfenic acids and the rate of reaction of sulfenic acids with current probes make it difficult to catch all active protein oxidations via proteomic analyses or to quantitate total amount of protein undergoing active oxidation, although relative changes in active oxidation can be assessed (168).

The second common technique is to target what can be termed an

“endpoint” oxidation, such as disulfide linkages, by what is commonly called a

“biotin switch” or “thiol switch” assay (169). First, all reduced thiols in a given sample are blocked with thiol-alkylating chemicals such N-ethylmaleimide (NEM) or Iodoacetamide (IAM). After this initial step of alkylation, oxidized thiols can be reduced and subsequently alkylated with various versions of IAM or maleimide that are linked to reporters such as biotin, fluorescent markers, or even ~15kDa oligonucleotides to promote dramatic shifts on protein gels (170). One advantage to this technique is that it is possible to selectively reduce certain types of oxidations while leaving others intact. Some examples include ascorbate, which has been reported to specifically reduce nitrosothiols (171,172), arsenite for the reduction of sulfenic acids (172), or general reductants such as dithiothreitol

(DTT) or tris(2-carboxyethyl)phosphine (TCEP) that will reduce all cysteine oxidations except for sulfinic or sulfonic acids or other irreversible oxidized crosslinks such as sulfonamides (169). Another advantage of this technique is

28 that it allows researchers to potentially quantitate total levels of protein oxidation in cells at a given time period. However, these advantages are coupled with several disadvantages. First, commonly used thiol alkylators such as NEM and

IAM are not completely specific for reduced thiols (173), leading to difficulties in quantitating total protein oxidation and potentially missing modifications in proteomic analyses. Additionally, many proteins, including ERK1/2, have multiple cysteine residues that can be oxidized, thus leading to labeling of the same protein molecule with the reporting alkylator multiple times. This alkylating multiplicity can result in unresolved total oxidation of a particular protein.

Furthermore, it is still debated whether or not particular reductants are fully specific for a particular type of oxidation (173), so researchers cannot definitively conclude if the oxidation is a result of reversal of a disulfide, a sulfenic acid, a sulfenamide, a S-nitrosothiol, or another type of oxidative modification.

Despite these caveats with labeling and identifying proteins oxidized in situ, current techniques have still revolutionized our ability to investigate exciting avenues of research that were unavailable even just 5-10 years ago. Many different classes of proteins important in signal transduction have been identified as targets of ROS and RNS, including phosphatases (174,175), transcription factors (176-179), G-proteins (180,181), receptors (182,183), and kinases (Table

1-I) (184,185). Phosphatases, specifically members of the PTP superfamily, were one of the first classes of signaling enzymes shown to be oxidized in response to physiologically-relevant signals (175). Members of this family of phosphatases have an active site cysteine whose pKa is lowered by coordination with an

29 arginine residue. This cysteine-arginine pairing assists in activating the cysteine to remove phosphate groups from tyrosines through a nucleophilic attack by the thiolate. The lowered pKa also renders this cysteine sensitive to direct oxidation by ROS, allowing the phosphatase to be inactivated by oxidation (175,186). This inactivation of PTPs allows for a sustained phosphorylation of kinases and their targets during signaling.

Over the past several years, a number of kinases have also been shown to be sensitive to oxidation, in response to either oxidative stress or extracellular signals (Table 1-I). Two notable examples of these are Protein Kinase A (PKA) and Protein Kinase B β (PKBβ), also known as Akt2, both of which are members of the AGC subfamily of kinases (so named for Protein kinases A, G, and C) (15).

There have been multiple reports of PKA oxidation, both in the catalytic subunit

(187,188) and on the regulatory subunit R1α (189). In the catalytic subunit of

PKA, the pKa of C199 is dramatically lowered by R165 within the active site

(188). Although oxidation of this site is yet to be identified in situ, in vitro studies reveal that this cysteine can be readily oxidized and glutathionylated, which appears to promote the dephosphorylation of S388, rendering PKA inactive

(188). An alternative, yet still direct, regulation of PKA by oxidation was recently shown in one version of the regulatory subunits, R1α, which is sensitive to oxidation to stimulate PKA signaling necessary for angiogenesis (189).

Akt2 oxidation was identified in NIH-3T3 mouse fibroblast cells and in

Mouse Embryonic Fibroblasts (MEFs) treated with PDGF (190,191) and in ovarian and prostate cancer cells treated with lysophosphatidic acid (LPA) (148).

This oxidation is specific for Akt2 on C124 (190), which is not conserved between the three Akt isoforms, indicating differential modulation of Akt activity between its different isoforms that, until recently, has been difficult to assess. Oxidation of this cysteine is in a linker region between the catalytic domain and pleckstrin homology (PH) domain and appears to inhibit Akt2 kinase activity as measured in vitro (190). Furthermore, mutating this cysteine disrupts the normal cell cycle distribution and migration pattern of MEFs (191).

These evidences that kinases are also susceptible to oxidation during oxidative stress or exposure to extracellular signals strongly encourages the hypothesis that the PTM of cysteine modification could be a missing piece in understanding the regulation of several types of kinases, including ERK1/2.

There are quite a few reports suggesting that MAPK enzymes, particularly

ERK1/2, are sensitive to cysteine modifications (Table 1-II). While most of the literature for ERK1/2 oxidation is in the context of bolus addition of oxidants to cells such as H2O2 (128), nitric oxide donors (129,133) or other oxidants such as doxorubicin (131), there are two select papers whose data suggest ERK1/2 can be oxidized in response to the extracellular signal VEGF (130) or nitrosylated in response to TNFα (133). Some of these papers went on to identify specific cysteine residues that are modified, whereas others simply showed that ERK1/2 does have an oxidative modification as measured by either the thiol-switch assay as described above (129) or via the use of a dimedone-recognizing antibody

(131).

Oxidation of ERK1/2 could be a key aspect regulating ERK1/2 activity.

Several of the cysteine residues found in ERK1/2 are in important regulatory regions of the enzyme. Of particular note are C38, C164, C159, and C252

(10,55). C38, although in a buried, hydrophobic region, is a serine in ERK1 and therefore could be a way to distinctly modulate ERK2 activity differently from

ERK1 (Figure 1-4). C164 is in the ATP binding pocket, so oxidation of this cysteine could sterically impede ATP binding, rendering the kinase inactive toward all substrates. C159, as mentioned in section 1.2, makes up a significant part of the surface of the DRS, so oxidation of this cysteine could preclude or modify which DRS-binding proteins are able to interact with ERK1/2 (Figure 1-5).

C252 is near the FRS, and immediately adjacent to the SPS-phosphorylation motif, so oxidation of this cysteine could either modify ERK1/2 protein-protein interactions with FRS-interacting partners or modulate ERK1/2 interactions that affect its translocation to the nucleus (Figure 1-5). In truth, oxidation of any of these cysteines could be a key mode of ERK1/2 regulation. Thus, elucidating the role of specific cysteine oxidation on ERK1/2 activity will enable a clearer understanding of how cells control ERK1/2 to elicit signal-specific cell responses to distinct stimuli.

1.4 Statement of purpose

There is clear evidence in the literature that ROS, particularly H2O2, can act as second messengers and are able to target several classes of key signaling proteins including kinases (184,185). In particular, evidence of ERK1/2 oxidation is sufficient (Table 1-II) to propose the hypothesis that these two central kinases

32 are cysteine-oxidized in response to extracellular signals. Furthermore, preliminary studies done by the Poole and Daniel laboratories suggested that

ERK1/2 may be oxidized in response to LPA in SKOV3 cells. My graduate research has focused on two key projects to better reveal the role that oxidative modifications of ERK1/2 play in regulating the response specificity of ERK1/2 signal transduction:

1. I undertook an analysis of ERK1/2 sulfenic acid formation in response

to a variety of proliferative signals in both non-transformed and cancer-

derived cell lines. I have identified that oxidation of ERK1/2 occurs

downstream of RTKs, GPCRs, and HRs. I have analyzed the temporal

dynamics of oxidation among total and TEY-phosphorylated

populations of ERK1/2 in each of these cell lines stimulated with cell-

specific proliferative signals. In these analyses, I have identified

differences in temporal dynamics of TEY-phosphorylated ERK1/2

oxidation between the same signal in two closely-related cell lines and

between the same cell line when stimulated under different resting

conditions. This and supporting data covered in Chapter 2 of the main

body of my thesis strongly support the hypothesis that ERK regulation

by cysteine oxidation occurs in situ.

2. I undertook an analysis of recombinant rat ERK2 (rERK2) reaction with

H2O2 in vitro and sought to understand how reaction with H2O2 affects

kinase activity towards a variety of substrates. We were able to identify

oxidant-sensitive cysteines and analyze changes in substrate

specificity when ERK2 is oxidized by H2O2 in vitro. Furthermore, I was

able to identify that, while endogenous ERK1/2 oxidation in cells is

H2O2-dependent, oxidation of ERK2 in vitro with H2O2 does not

completely mimic observations of how endogenous ERK oxidation

affects kinase activity, prompting the need for further cellular and in

vitro analyses to better understand how ERK1/2 is oxidized in cells and

how this modification regulates ERK signaling.

Due to the clinical need to find effective ERK1/2 pathway inhibitors for the treatment of cancer and other diseases, it is imperative that a more complete understanding of signal-specific ERK1/2 regulation is achieved in order to enable targeting of the ERK1/2 pathway in the context of specific signals to prevent the drastic potential side effects of ubiquitous ERK1/2 inhibition (16,192,193). My graduate work has made a significant contribution to this need by identifying that

ERK1/2 is indeed oxidized in response to extracellular signals and that this oxidation significantly affects ERK1/2 kinase activity and substrate specificity.

References

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Chapter 2: Endogenous Sulfenylation of ERK1/2 in Response to

Proliferative Signals

Jeremiah Keyes, Derek Parsonage, Rama Yammani, LeAnn Rogers, Chelsea

Kesty, Kimberly Nelson and Leslie Poole

This chapter will be submitted to Free Radical Biology and Medicine. Keyes performed cell harvests, DCP-Bio1 labeling, affinity capture, immunoblots, ppERK1/2 immunoprecipitations, GST-Elk1-His6 cloning, expression, and purification, and endogenous ERK1/2 activity assays. Yammani and Rogers aided in cell culture, DCP-Bio1 labeling and affinity captures; Yammani also aided in

Western Blotting. Kesty aided in ppERK1/2 immunoprecipitations. Dr. Nelson aided experiment design and analysis. Dr. Parsonage aided in GST-Elk1-His6 cloning, expression, and purification. Dr. Poole served in an advisory and editorial capacity.

Abstract

The ERK pathway plays roles across eukaryotic biology by transducing extracellular signals into cell-fate decisions. One conundrum is in understanding how disparate signals induce specific responses through a common, ERK- dependent kinase cascade. While studies have revealed intricate modes to control ERK through spatiotemporal localization and phosphorylation dynamics, additional details of ERK signaling remain elusive. We hypothesized that fine- tuning of ERK signaling could occur by cysteine oxidation. We report that ERK is actively and directly oxidized by signal-generated H2O2 during proliferative signaling, and that oxidation occurs downstream of a variety of receptor classes tested in six cell lines. Furthermore, within the tested cell lines and proliferative signals, we observed that both TEY-phosphorylated and non-phosphorylated

ERK undergo sulfenylation in cells, that there is a difference in the consistency of

ERK oxidation between transformed and non-transformed cells, and that dynamics of ERK sulfenylation is dependent on whether or not cells are serum starved prior to stimulus. We also tested the effect of endogenous ERK oxidation on kinase activity and report that phospho-transfer reactions are drastically inhibited by oxidation, underscoring the importance of considering this alternative modification when assessing ERK activation.

Introduction

Reactive oxygen species (ROS), while previously considered little more than detrimental byproducts of respiration (1,2), are now recognized as significant contributors to cellular signaling processes(2-7). Intensive study of

ROS in signaling began to garner attention when Sundaresan et al. demonstrated that H2O2 was formed in response to PDGF in primary rat vascular smooth muscle cells, and that the cells’ proliferative response to PDGF depended upon sufficient generation of hydrogen peroxide (8). This has now been demonstrated in many cell types and in vivo models (9-12), including recent work by our research team establishing the important role of ROS in the proliferative responses to lysophosphatidic acid (LPA), a bioactive lipid, of ovarian and prostate cancer cell lines (13,14). ROS-dependent signaling is not limited to proliferative signaling, as demonstrated by several recent examples: sulfenic acid formation found on the leading edge of HUVEC cells was found to be necessary for migration(15), stem cell pluripotency and differentiation was dependent on subtle changes in the delicate redox balance of cells(16), the redox landscape of a cell contributes nuclear reprogramming (17), and ROS formation is a vital component of chondrocyte response to fibronectin fragments.(18,19)

Much work has been done by our research team along with others in the field of redox signaling to identify targets of signal-produced ROS (20-28). Many

ROS and reactive nitrogen species (RNS) can react with cysteine residues of proteins, modifying the protein’s structure, dynamics, and/or function. Most often,

49 the oxidation of a cysteine residue is reversible (29-31), which sets up the oxidation of a target cysteine on a protein to be a molecular switch. This switch- like property of cysteines allows them to have evolved to serve specific functions as redox sensors on proteins (30,32-35). For example, a well-established example of a binary cysteine switch is in the protein tyrosine phosphatase (PTP) family of proteins (36). These enzymes contain an active site cysteine whose pKa is lowered by interaction with an arginine residue. This cysteine-arginine pairing assists in activating the cysteine to remove phosphate groups from tyrosines through a nucleophilic attack by the thiolate. However, it also renders this cysteine sensitive to direct oxidation by ROS. It has been shown in many signaling systems that PTPs are inactivated by oxidation on this active site cysteine during signaling, allowing for a sustained phosphorylation response to stimulus (37,38).

Identifying novel proteins whose cysteines are oxidized in response to extracellular signals has been historically difficult due to the labile chemistry of cysteines. One class of cysteine probes developed by our research team is based on dimedone (figure 2-1A) (39,40). Dimedone alkylates sulfenic acid (R-

SOH)(41), which is the intermediate oxidized form of a cysteine that has reacted with ROS molecules such as H2O2. Sulfenic acids are reactive toward other thiols, forming disulfide bridges which are likely a less transient product of the initial oxidation, and dimedone-like compounds can interfere with native disulfide bond formation (42). Using dimedone-based probes to capture the sulfenic acid- containing proteins undergoing active oxidation in cells responding to external

Figure 2-1. Trapping of sulfenic acids within cellular proteins during lysis. A) Labeling of oxidized proteins with DCP-Bio1. The reactive carbon of DCP-Bio1 selectively reacts with sulfenic acids to form a covalent adduct, thereby biotinylating proteins containing sulfenic acids at the time of lysis. B) Overall experimental approach for labeling of proteins undergoing active oxidation to sulfenic acid. Cells are cultured to ~80% confluency, treated with an exogenous signal of interest such as platelet-derived growth factor (PDGF), and harvested in lysis buffer containing DCP-Bio1 reagent, N-ethylmaleimide (NEM) and iodoacetamide (IAM) to alkylate free, reduced thiols (to prevent thiol oxidation during lysis), and catalase to remove any H2O2 formed during lysis. After incubation for 30 min on ice, lysates are clarified and excess DCP-Bio1 is removed with a molecular sieve (BioGel P6) spin column. Clarified lysates are subjected to avidin-based affinity capture, then beads are stringently washed to remove any unlabeled proteins carried over during capture. Captured proteins are eluted by incubation in 2% sodium dodecyl sulfate (SDS) at 100 ᵒC for 10 minutes and then subjected to an immunoblot for the protein of interest. To control for differences in capture efficiency and gel loading, a pre-biotinylated bacterial protein, AhpC, is added to lysates before affinity capture.

51 signals enable their identification and, combined with mass spectrometry, can reveal the identity of the reactive cysteine within these proteins (43,44).

One of the probes developed by our collaborative team of researchers,

DCP-Bio1 (Figure 2-1A), contains the reactive core of dimedone linked to biotin, allowing for sulfenic acid labeling and then affinity capture of cellular proteins undergoing active oxidation in the cell (figure 2-1B) (21,22). This probe has been used to identify many novel targets of oxidation as a result of extracellular signaling (14,15,22,45-47). One of the notable proteins identified is the signaling kinase Akt2, which is oxidized on an isoform-specific cysteine during PDGF signaling in NIH-3T3 cells (47), and as a result of LPA signaling in SKOV3 and

PC-3 cells (14). Oxidation of this cysteine appears to be important for normal cell cycle distribution and cell migration of MEFs (48).

Based on preliminary data from early experiments with DCP-Bio1 labeling of proliferating cells, one of the signaling enzymes further investigated for redox regulation was the Mitogen-activated protein kinases ERK1 and ERK2

(designated ERK1/2). These proteins are at the heart of a variety of signaling pathways leading to such responses as proliferation, differentiation, survival, migration(49-51), and even apoptosis (52-54). This presents a conundrum of specificity – how do disparate signals induce specific responses while sharing the

ERK signaling cascade? Evaluating ERK1/2 regulation is vital to understanding a variety of diseases from cancer to diabetes to neurological disorders. In fact, the various components of the ERK1/2 signaling cascade (particularly Ras and Raf) are highly upregulated through various oncogenic mutations, leading to an

52 overall reported hyperactivation of the ERK1/2 pathway in up to 90% of all human cancers (55).

Extensive research to date has revealed much about the ERK1/2 pathway and its regulation. Upon binding of an appropriate ligand to its receptor, a cascade of signaling protein activation occurs through the GTPase Ras, which activates the Raf kinases, which in turn activate the MAP kinase kinases MEK1 and MEK2 (referred to as MEK1/2). MEK1/2 proteins doubly phosphorylate

ERK1/2 on T183 and Y185 (51), hereafter designated as the TEY motif, which induces a conformational change that opens up the active site of ERK1/2 (56,57) and enhances structural flexibility (58,59), allowing ERK1/2 to phosphorylate substrates. However, with over 300 known substrates split between the cytosol and nucleus (55), evolution has created multiple layers of regulation. These additional layers include temporal and spatial control of TEY phosphorylation and dephosphorylation (60,61). Yet, while we know that this dual phosphorylation is a prerequisite for ERK1/2 activation, it is not yet fully clear how cells orchestrate

ERK1/2 localization, discriminate between substrates, or regulate the dephosphorylation of ERK1/2 by protein phosphatases (62).

Our knowledge of cysteine switches, including the tentative identification of ERK1/2 oxidation resulting from vascular endothelial growth factor (VEGF) signaling (15), led us to hypothesize that ERK1/2 cysteine oxidation could be a mode to control cell decisions regarding ERK1/2 regulation. Human ERK1 and

ERK2 contain 6 and 7 cysteines, respectively, half of which are solvent exposed and conserved in the MAPK family (56,63). Analysis of the literature gives ample

53 evidence that a few of these cysteines could be redox sensitive (Table 1-II).

Indeed, others have shown that under oxidative stress conditions, as mimicked by the bolus addition of H2O2 or nitric oxide donors, ERK1/2 cysteines respond to these stresses (15,64-67). While there are evidences in the literature that

ERK1/2 cysteine modification can occur downstream of the physiological signals of VEGF (15) and TNF-α (67), it has not definitively been shown that ERK1/2 oxidation commonly occurs as a result of a variety of extracellular signals. We thus used the dimedone-based DCP-Bio1 to test this hypothesis; this is the first report to document the modification of ERK1/2 cysteines in response non-stress extracellular signals and to relate any observed sulfenic acid formation to endogenous ERK1/2 kinase activity. We report that ERK1/2 cysteine sulfenylation occurs in response to a variety of proliferative signals and that endogenous oxidation affects ERK1/2 kinase activity.

Materials and Methods

Reagents and antibodies

Primary antibodies for Western blots to ERK (recognizing both ERK1 and

ERK2, p44 and p42, respectively), TEY-phosphorylated ERK1/2, Elk1 and phospho-Elk1, along with anti-rabbit, anti-mouse HRP-conjugated secondary antibodies and Platelet Derived Growth Factor BB (PDGF-BB) were from Cell

Signaling Technology. AhpC was expressed and purified as described previously(68,69), and anti-AhpC antibody was purified from rabbit serum (22).

DCP-Bio1 was synthesized as described previously (40). PEG-Catalase was from Sigma. DMEM and Fetal Bovine Serum were from Lonza; EMEM was from

Gibco. Chemiluminescence reagents for Western Blotting (SuperSignal Dura) were from Thermo Scientific. Nitrocellulose was from GE Healthcare -

Amersham. Magnetic A beads (Dynabead) for phospho-ERK1/2 immunoprecipitations were from Novex by Life Technologies, and ATP for in vitro kinase assay was acquired from Promega. Iodoacetamide (IAM) was from

Sigma-Aldrich; Dithiothreitol (DTT) and N-ethylmaleimide (NEM) was from

Fisher-scientific.

Elk1 for ERK1/2 in vitro activity assays was cloned, expressed, and purified as described below. GST-Elk1-Hisx6 was expressed from a pGex-6P-1- derived bacterial expression plasmid. First, the Elk1 DNA sequence corresponding to residues 307-428 of human Elk1 was synthesized by

GenScript, with the codon usage modified for expression in E. coli. The gene was sub cloned into pGex-6p-1 between BamH1 and XhoI restriction sites. Due to issues with truncated versions of the protein being expressed in E. coli, a 6-His tag was added to the C-terminal end of Elk1 using PCR to reengineer the DNA insert, with a forward primer (5’-CTGGGATCCATTTCGCAACCGC-3’) and a reverse primer encoding the extended His tag at the C-terminus (5’-

GCGCTCGAGTTAATGGTGATGGTGATGGTGCGGTTTTTGCGGACCCGGCG

AC-3’). After PCR product purification, the DNA fragment was ligated into pGex-

6p-1. Successful cloning was verified by restriction digest followed by gel analysis and was verified in its entirety by sequencing (Eurofins Genomics).

GST-Elk1-Hisx6 was expressed in BL21 Gold cells with growth in TYP media supplemented with 30 mM glucose and 100 μg/mL ampicillin at 37 °C to

55 an optical density of 1 at 600 nm. Expression was then induced by addition of 1 mM IPTG and incubation continued at 24 °C for 4 hours, after which cells were centrifuged and stored at -80 °C until used for purification. Cells were thawed and resuspended in 25 mM potassium phosphate at pH 7.0, with 150 mM NaCl and

10% glycerol, and disrupted using an Avestin C5 emulsifier. After centrifugation, the supernatant was applied to a cobalt-NTA column (GE Healthcare) preequilibrated in 50 mM sodium phosphate at pH 8.0, with 0.3 mM NaCl and 20 mM imidazole, washed with 4 column volumes of the same buffer, then eluted by increasing the imidazole to 500 mM. Elk1-containing fractions were pooled, bound to Glutathione Sepharose (GE Healthcare) preequilibrated and subsequently washed with 50 mM Tris-HCl at pH 8.0 (4 °C), with 150 mM NaCl,

1 mM EDTA and 10% glycerol. Bound protein was eluted using 50 mM Tris-HCl at pH 8.0 containing 10 mM reduced glutathione. GST-Elk1-Hisx6 was further purified on a 15 mL Source Q column using a gradient of 50 mM to 1 M NaCl 50 mM Tris-HCl pH 7.5, 10% glycerol. Elk1-containing fractions were then pooled, concentrated, exchanged by ultrafiltration into 20 mM Tris-HCl at pH 8.0, with 1 mM EDTA and 10% glycerol, aliquoted, and stored at -80 °C until use.

Cell culture and treatments

NIH-3T3 and WI-38 cells (from ATCC stocks) were cultured at 37 °C with

5% CO2 in either DMEM (NIH-3T3) or EMEM (WI-38) supplemented with 10% fetal bovine serum, L-glutamine, penicillin, and streptomycin. Except where indicated, cells were transferred into fresh, serum-free media for 18 hours before treatment with PDGF. PDGF-BB was added to the cells to a final concentration of

20 ng/mL and then cells were harvested at indicated time points. Where indicated, PEG-Catalase was added at 400 U/mL 18 hours before PDGF stimulation.

DCP-Bio1 labeling and affinity capture

NIH-3T3 or WI-38 cells (~5 x 105) were grown in FBS-supplemented media in 100-mm plates for 24 hours before serum-starvation pretreatment, except for when cells were stimulated in serum-replete conditions as indicated.

Cells grown to ~80% confluency were treated with 20 ng/mL PDGF and harvested at indicated time points with lysis buffer supplemented with DCP-

Bio1essentially as described previously (21). Briefly, lysis buffer (50 mM Tris-HCl at pH 8.0, with 100 mM NaCl, 100 μM diethylene triamine pentaacetic acid

(DTPA), 20 mM β-glycerophosphate, 0.1% SDS, 0.5% Na Desoxycholate, 0.5%

NP-40, and 0.5% Triton-X-100) was freshly prepared with 1 mM PMSF, 10 μg/mL aprotinin, 1 mM Na3VO4, 10 mM NaF, 1 mM DCP-Bio1, 10 mM NEM, 10 mM iodoacetamide (IAM), and 200 U/mL Catalase. Seventy-five μL of lysis buffer was added to each dish and cells were scraped from the plates, transferred to micro- centrifuge tubes, incubated on ice for 30 minutes, and then stored at -80 °C.

Before affinity capture and elution of labeled proteins (22), samples were thawed and clarified by centrifugation, then excess DCP-Bio1 was removed via a BioGel

P6 spin column. Samples were then assayed for protein concentration, diluted into 2 M urea and supplemented with 0.5 μg prebiotinylated AhpC to control for the efficiency of the affinity capture, elution, and gel loading steps. After samples were precleared with Sepharose CL-4B beads (Sigma), they were applied to

57 plugged spin columns containing high capacity streptavidin-agarose beads

(Thermo scientific) and incubated overnight at 4 °C with constant rotation.

Samples then underwent a series of stringent washes to remove any contaminating, non-labelled proteins that were co-captured with DCP-Bio1 conjugated proteins. Washes consisted of 2 washes each with 4 column volumes of, (in order) 1% SDS, 4 M Urea, 1 M NaCl, 10 mM DTT, 50 mM ammonium bicarbonate and water. Labeled proteins were eluted with 25 mM Tris-HCl at pH

6.8, containing 2% SDS, 50 mM DTT, 0.02% bromophenol blue, and 5% glycerol, incubated at 100 °C for 10 minutes, then centrifuged to separate protein from beads. Samples were then stored at -80 °C as needed and analyzed by

Western blot as described below.

Western Blotting

For immunoblots of proteins labeled with DCP-Bio1, 40-80 μg of protein was separated on 10% SDS-polyacrylamide gels and transferred to nitrocellulose membranes for 1 hour at 100 V at 4 ˚C (or overnight at 100 mAmps in the cold).

For immunoblots of cell lysates, typically 10 μg of protein was loaded per lane.

For phospho-Elk1 immunoblots, 1.2-2.4 μg of ERK1/2-treated GST-Elk1-Hisx6 was added to each lane. After transfer to nitrocellulose, membranes were blocked with 5% BSA, probed with protein-specific antibodies, and visualized using SuperSignal Dura chemiluminescence reagent. Exposed film was then scanned on an Epson Perfect V30, where original image integrity was maintained by adjusting scanner settings to prevent auto-adjustments in contrast and gamma. Band density was measured using ImageJ software. To analyze data

58 from DCP-Bio1 experiments, each sample was first normalized to AhpC band density to control for variations in affinity-capture efficiency, and then each replicate was normalized to the most intense ERK1/2 band, which was set to 1.

For quantitation of Elk1 phosphorylation after in vitro kinase assays of immunoprecipitated ERK1/2, each lysate sample was normalized to phospho-

Elk1 intensity from kinase reactions including DTT. For DCP-Bio1 labeling experiments, p-values were calculated using Student t-tests assuming 2-tailed distributions with two-sample equal variance. For activity assays, p-values were calculated using Student’s t-test for paired samples, 2-tailed distribution.

In vitro kinase assays of ERK1/2 immunoprecipitated from cells grown in culture

NIH-3T3 cells were treated and harvested as described above, but without the addition of DCP-Bio1, IAM, or NEM to the lysis buffer. Lysates were subsequently clarified by centrifugation and protein content of the supernatants was measured via BCA assay. Samples containing 50 μg of protein were added to 4 μL of anti-phospho-p42/44 rabbit antibody into a total volume 200 μL of lysis buffer supplemented with protease and phosphatase inhibitors and 200 U/mL of catalase. Twenty-five μL of magnetic A beads were suspended in the lysate plus antibody mix and incubated overnight at 4 °C with constant rotation. The following morning beads were washed twice with 5 column volumes of lysis buffer with phosphatase and protease inhibitors, followed by 2 washes with 5 volumes of kinase buffer (25 mM Tris-HCl at pH 7.5, with 5 mM β- glycerophosphate, 0.1 mM Na3VO4, and 10 mM MgCl2). After washes, beads were resuspended in 100 μL of 40 mM Tris-HCl at pH 7.5, with 0.1 mM Na3VO4;

59 samples were diluted five-fold with either DTT (to give a final concentration of 2 mM) or HPLC water, then incubated with 2.21 μM GST-Elk1-Hisx6 fusion protein substrate, 200 μM ATP, and Kinase buffer at 30 °C for 30 minutes with constant shaking. Reactions were quenched by addition of protein gel loading buffer (25 mM Tris-HCl at pH 6.8, with 2% SDS, 50 mM DTT, 0.2 % bromophenol blue, and

5% glycerol) and incubated at 100 °C for 5 minutes. Samples were either stored at -80 °C or directly loaded onto a 10% SDS-polyacrylamide gel, transferred to nitrocellulose, and probed for phospho-Elk1 as described above.

Results

To test whether or not ERK1/2 is a target of direct oxidation in cells responding to proliferative signals, we used the dimedone based probe DCP-

Bio1, as described in figure 1-1 and Materials and Methods. Various cell lines were treated with cell-specific growth factors over 5 to 30 minutes before harvest.

Cells were harvested in the presence of DCP-Bio1, catalase, and the cysteine alkylators (NEM and IAM) to minimize artifactual oxidation of cysteines during lysis (21). After clarification, lysates were subjected to biotin-dependent affinity capture as described in Materials and Methods. Captured proteins were resolved by SDS-PAGE and immunoblotted for total and doubly (TEY) phosphorylated

ERK1/2 (22). We observed that ERK1/2 was labeled by DCP-Bio1 in response to

PDGF in NIH-3T3 mouse fibroblasts (figure 2-2) as well as other treated cell types (vide infra). Considering that ERK1/2 was not biotinylated by endogenous means (supplemental figure 1), we can conclude that the biotinylation of ERK1/2 when DCP-Bio1 is added to lysis buffer indicates active sulfenic acid formation

occurring on ERK1/2 at indicated time points after stimulus. In NIH-3T3 cells, we

observed a consistent increase in sulfenic acid formation of ERK1/2 10 minutes

after stimulation with a return to basal levels of sulfenic acids 30 minutes after

stimulation (figures 2-2), confirming our hypothesis that ERK1/2 oxidation occurs

during physiologically-relevant extracellular signals.

Figure 2-2. ERK1/2 cysteines are oxidized to sulfenic acid in response to PDGF in NIH-3T3 cells. ERK oxidation was monitored during proliferative signaling as described in Materials and Methods. Briefly, cells were treated with 20 ng/mL PDGF after 18 hours in serum-free media and lysed in the presence of the sulfenic-acid trap DCP-Bio1, thereby biotinylating proteins undergoing cysteine oxidation. After addition of prebiotinylated AhpC as an internal control, DCP-Bio1 labeled proteins were captured via streptavidin-agarose beads, resolved by SDS-PAGE, and subjected to western blot for total and doubly- (TEY) phosphorylated ERK (ppERK). Upper panels of (A) and (B) show total (A) and doubly (TEY) phosphorylated (B) ERK oxidation detected in NIH-3T3 cells treated with PDGF. Lower panels summarize ERK band intensity from multiple experiments (n=4) of captured proteins in (A) and (B), respectively, following normalization to AhpC and then to the most intense band (10 min time point) for each replicate (set to 1). ** p < 0.0005, *** p<0.0001, ^ p<0.05, ^^ p<0.001, ^^^p<2x10-19.

To further investigate the source and nature of the ROS responsible for

ERK1/2 oxidation, we utilized the ROS modulator PEG-Catalase, which

scavenges H2O2 and is cell permeable. NIH-3T3 cells were treated with PDGF

for 10 minutes with or without PEG-Catalase. We observed that PEG-Catalase

significantly dampened the amount of ERK1/2 labeled by DCP-Bio1 (figure 2-3).

Thus, intracellular H2O2, or a derivative thereof, is implicated as the ROS

molecule responsible for ERK1/2 oxidation in cells.

Figure 2-3. ERK1/2 oxidation is caused by H2O2 generated in response to PDGF in NIH-3T3 cells. NIH- 3T3 cells were grown as in Fig. 2 and treated or not with PDGF for 10 minutes with or without the addition of PEG-Catalase as described in Materials and Methods. PEG-Catalase significantly reduced the amount of sulfenic acid formation on total-ERK (A) and phosphorylated ERK (B) in response to PDGF, indicating that sulfenic acid formation on ERK is a result of cysteine reaction with H2O2 (or a derivative of H2O2). Lower panels summarize ERK band intensity from multiple experiments (n=4) normalized as described in Figure 2. Images are representative of four biological replicates. * p< 0.01, ** p< 0.0005, ^p<0.05, ^^ p<0.1x10-8.

We also investigated the temporal dynamics of ERK1/2 sulfenic acid formation in another fibroblast cell line, WI-38, in response to PDGF stimulation.

WI-38 cells were serum-starved for 18 hours before PDGF stimulation, as was done with NIH-3T3 cells in Figure 2-2. Upon PDGF stimulation we observed a very similar pattern in sulfenic acid formation among doubly TEY-phosphorylated

ERK1/2, where there was an initial abrupt increase followed by a decline in observed sulfenic acids on ERK1/2 (Figure 2-4B). However, when probing DCP-

Bio1 labeled proteins with an antibody expected to measure total-ERK1/2 in a sample; we observed a relatively modest amount of total-ERK1/2 undergoing active oxidation before PDGF treatment. Upon PDGF stimulation, sulfenylation of the total population of ERK1/2 decreased and then considerably increased 30 minutes after treatment (figure 2-4A).

We also designed experiments to test the hypothesis that oxidation patterns vary with serum presence or absence during cell culture and treatment.

To test this hypothesis, we repeated the experiments described in figure 4A and

B in WI-38 cells but under serum replete conditions. In contrast to the PDGF- induced oxidation pattern observed in serum-depleted cells, serum-replete WI-38 cells exhibited very different dynamics of ERK1/2 oxidation (figure 4C, D). We observed relatively very high levels of total-ERK1/2 oxidation occurring before

PDGF treatment that significantly decreased 10 minutes after stimulation and did not increase to high levels of sulfenic acid formation by 30 minutes. Doubly-TEY phosphorylated ERK1/2, however, had a delayed oxidation response when

Figure 2-4. Serum-depleted WI38 fibroblasts exhibit distinct temporal patterns of sulfenic acid formation in response to PDGF compared to serum-replete WI38 cells. Cells were either serum-starved for 18 hours (A) and (B) or maintained in 10% fetal bovine serum (C) and (D) before treatment with PDGF. After indicated times with PDGF, cells were lysed in the presence of DCP-Bio1 as described in Figures 1 and 2. Affinity-captured proteins were immunoblotted for total (A) and (C) or for doubly (TEY) phosphorylated (B) and (D) ERK1/2. Lower panels represent multiple replicates normalized as in Fig. 2 (n=4 for serum-depleted; n=3 for serum-replete). * p< 0.05, ** p< 0.01, *** p< 0.005, ^ p< 0.0005, ^^ p< 0.00005, ^^^ p< 5x10-10.

compared to the response observed in serum-depleted cells, with a maximal

increase in sulfenic acid formation in serum-replete cells occurring 30 minutes

after PDGF stimulation. These data confirm that the basal growth conditions of

cells modulate the temporal dynamics of protein oxidation in response to a

stimulus.

Using this same sulfenic-acid labeling technique, we also investigated

ERK1/2 cysteine oxidation in HeLa, SKOV3, PC3, and RWPE-1 cells during

responses to the respective natural growth factors of EGF, LPA (for both SKOV3

and PC3), and the synthetic androgen-agonist R1881, respectively. Although the

patterns of total and phosphorylated ERK1/2 oxidation were typically less

consistent in these cells lines, we observed ERK1/2 oxidation occurring in each

(Supplemental figures 2-5). It is interesting to note that SKOV3, PC3, and HeLa

cell lines are derived from human tumors, whereas the non-transformed NIH-3T3

embryonic mouse fibroblasts, WI-38 embryonic human fibroblasts, and RWPE-1

human prostate epithelial cell lines are derived from normal, non-diseased

tissues. This suggests that the regulation of ERK1/2 oxidation is aberrant in

cancers.

To determine the effect of cysteine oxidation on ERK1/2 kinase activity,

NIH-3T3 cells treated with PDGF (figure 2-5) and SKOV3 cells treated with LPA

(Figure 2-S6) were harvested at various time points after stimulus and TEY- phosphorylated ERK1/2 was immunoprecipitated. Each sample of immunoprecipitated ERK1/2 was split, with one part incubated with 2 mM DTT to reverse endogenous oxidation, and the other half lacking DTT (to preserve the endogenous oxidation state from cells). Each sample was assayed in vitro by the addition of recombinant Elk1 and ATP for 30 minutes at 30ᵒC as described in

Materials and Methods. Samples were then immunoblotted for phosphorylation of

S383 of Elk1 (Figure 2-5, C and Figure 2-S6). At each time point tested, ERK1/2 kinase activity towards Elk1 was dramatically increased after reduction, revealing that ERK1/2 kinase activity, at least towards the substrate Elk1, is inhibited by endogenous ERK1/2 oxidation. These data strongly indicate that the activity of

ERK1/2 is modulated by endogenous oxidative cysteine modifications, revealing that the observed increase in sulfenic acid formation, and subsequent oxidation products formed, have an effect on ERK1/2 kinase activity.

Figure 2-5. Endogenous ERK oxidation inhibits kinase activity towards Elk1. Doubly (TEY) phosphorylated ERK1/2 was immunoprecipitated from NIH-3T3 cells treated with PDGF for various time points as in Fig. 2 and as described in Materials and Methods. Each sample of immunoprecipitated ERK1/2 was then split into two: one sample was diluted into dithiothreitol (DTT)-containing buffer while the other was left in its native redox state (diluted into buffer lacking DTT). Recombinant Elk1 and ATP were added and samples were incubated at 30ᵒ C for 30 minutes. An immunoblot of S383- phosphorylated Elk1 was conducted to measure relative activities of immunoprecipitated ERK1/2. (A) Representative immunoblot of phosphorylated-Elk1. (B) Normalized data as in (A) from 5 independent replicates. DTT- treated samples are more than 5-fold more active toward Elk1 than non-treated at all time points tested after PDGF addition. *P< 0.001 ** p< 0.000005 (C) Representative immunoblot for doubly (TEY) phosphorylated (top) and total (bottom) ERK1/2 from cell lysates.

Discussion

ERK1/2 are extensively studied serine/threonine kinases central to the signal transduction of a variety of pathways. Due to its ubiquitous nature, understanding ERK1/2 regulation is key to identifying how ERK1/2 signaling induces varying, and even contradictory, responses to particular signals (62). To date, several modes of regulation have been identified, including the activating dual phosphorylation on the TEY motif (56), interactions with anchoring proteins and scaffolds (70,71), control over localization via phosphorylation on the SPS motif (72), crosstalk with other signaling cascades (62), and signal duration as controlled by phosphatases (73-76). This current work identifies an additional layer of regulation through the post-translational modification of cysteine oxidation, which we demonstrate to be a modification not restricted to a particular class of receptors or cell types.

First, it is evident in several cell lines that ERK1/2 is labeled by DCP-Bio1 upon activation of RTKs, GPCRs, and nuclear hormone receptors (Figures 2-2,

2-4, 2-S2-S5). DCP-Bio1 labeling of ERK1/2 is indicative that at least one of the cysteine residues on ERK1/2 is reacting with oxidants to form sulfenic acid in response to the extracellular signal. Although this does not rule out the possibility that ERK1/2 oxidation can occur through a disulfide-transfer mediated by an oxidized protein such as a peroxiredoxin (77), the fact that dimedone-based probes are able to react directly with ERK1/2 reveals that ERK1/2 can react directly with oxidants produced by the cell. Furthermore, no biotinylation of

ERK1/2 is observed without the addition of DCP-Bio1 to lysis buffer, indicating

68 that biotinylation is due to sulfenic acid formation on ERK1/2 and not from endogenous biotinylation (Figure 2-S1). However, it should be noted that DCP-

Bio1 detects only the sulfenic acid intermediate and not the total pool of oxidized protein.

In NIH-3T3 mouse fibroblasts, there is very low basal sulfenic acid formation on ERK1/2 in the absence of serum (Figure 2-2). Upon PDGF treatment, sulfenic acid formation dramatically increases 5-10 minutes after stimulation (Figure 2-2). Thirty minutes after PDGF stimulation, sulfenic acid formation decreased to basal levels (although much protein remains oxidized, as detected by activity assays) even though the relative extent of TEY- phosphorylation did not decrease, indicating that active oxidation of ERK1/2 and

TEY-phosphorylation are not coupled. This conclusion is further corroborated in

Figure 2-3, where scavenging of H2O2 by PEG-Catalase abrogated sulfenic acid formation on ERK1/2 but did not decrease TEY-phosphorylation. While DCP-

Bio1 labeling and subsequent immunoblotting is unable to quantitate absolute ratios of sulfenic acid-oxidized ERK1/2 to total or phosphorylated forms of

ERK1/2, the temporal pattern of oxidation of total ERK1/2 and TEY- phosphorylated ERK1/2 are identical. This suggests, in NIH-3T3 cells’ response to PDGF, that only TEY-phosphorylated ERK1/2 undergoes active reaction with

H2O2 to form the sulfenic acids, or that phosphorylation and oxidation are co- temporal events.

Serum-depleted NIH-3T3 and WI-38 cells, both being embryonic, mesenchymal-derived fibroblast cells lines (78,79), exhibited very similar

69 responses of TEY-phosphorylated ERK1/2 oxidation to sulfenic acid upon PDGF stimulation; both exhibited rapid increases in oxidation 5-10 minutes after stimulation followed by a precipitous drop in active oxidation. Although a direct comparison has not been made in the kinetics of cell responses between these two cell lines, it is not surprising that they exhibit a similar response in ERK1/2 cysteine oxidation to sulfenic acid given the fact that both cell lines require PDGF for sustained growth and survival along with a chemotactic response (80,81) .

However, it is interesting that the patterns of oxidation in WI-38 cells detected using antibodies against total ERK1/2 differ from those observed using antibodies against TEY-phosphorylated ERK1/2. Before PDGF stimulation, serum-depleted cells have almost no observable TEY-phosphorylated ERK1/2

(Figure 2-4B), indicating that the oxidized ERK1/2 observed before PDGF treatment in figure 2-4A is largely non-phosphorylated ERK1/2. To our knowledge, there is no direct comparison between NIH-3T3 and WI-38 cell lines in the literature, therefore it is difficult to speculate why WI-38 cells exhibit this basal level of ERK1/2 oxidation and NIH-3T3 cells do not. Furthermore, it is unclear why this basal oxidation is occurring; further investigation into the role of unphosphorylated ERK1/2 oxidation, along with clear quantitation of the ratios of oxidation to the different phosphorylation-state subpopulations of ERK1/2, is needed to understand the role this basal oxidation contributes to cell homeostasis.

Interestingly, serum-replete WI-38 cells treated with PDGF exhibited a very different pattern of ERK1/2 sulfenic acid formation (Figure2-4C, D) than

70 serum-starved WI-38 cells treated with PDGF (Figure 2-4A, B). The serum- replete fibroblast cells exhibited high levels of basal sulfenic acid oxidation of unphosphorylated ERK1/2; upon PDGF treatment there was a considerable decrease in active oxidation of total-ERK1/2 by 10 min. TEY-phosphorylated

ERK1/2, on the other hand, exhibited a delayed response in ERK1/2 oxidation compared to serum-free PDGF-stimulated WI-38 cells, with little immediate increase in active oxidation until 30 minutes after stimulus. The fact that the same cell line exhibited such a remarkable difference in oxidative response of

ERK1/2 indicates that the basal state of the cells has a significant impact on how the cells utilize ROS as second messengers upon stimulation. It is possible that this observation is a result to changes in overall expression of the PDGFR (82), indicating that changes in redox signaling can originate from the level of receptor regulation. Once again, further analysis comparing WI-38 cell responses to

PDGF under these two different conditions, and an investigation on the role that

ERK1/2 cysteine oxidation plays in this biological response, are needed to better understand how cells are primed and utilize ROS second messengers during signal transduction to induce biological responses to stimuli.

ERK1/2 oxidation also occurred in various other cell lines, including the prostate-epithelial cells RWPE-1 and the cancer-derived SKOV3, PC3, and HeLa cell lines (figures 2-S2-5). Although the patterns of relative changes in ERK1/2 sulfenic acid formation in these cell lines were not always consistent based on acquired replicates, we can conclude that oxidation of ERK1/2 does not only occur in a small subset of cell types or in response to only a certain class of

71 receptors. Indeed, our data demonstrate that ERK1/2 oxidation can occur downstream of not only RTKs, but also GPCRs and hormone receptors. It is also interesting to note that ERK1/2 was constitutively oxidized in SKOV3 cells

(Figure 2-S3), whereas the same signal in PC3 cells elicited a dramatic increase in ERK1/2 oxidation compared to untreated PC3 cells (Figure 2-S2).

Furthermore, it is notable that the cancer-derived cell lines exhibited either constitutive, as in SKOV3, or erratic patterns of ERK1/2 oxidation compared to the non-transformed cell lines used in this study. This observation suggests that the regulation of ERK1/2 oxidation could be mishandled in cancer cells.

While there are a number of functions of ERK1/2 independent of its kinase activity (83-85), its primary mode of action is through phosphorylation of over 300 known substrates (55). One of the early response transcription factors phosphorylated by ERK1/2 during proliferative signaling is Elk1 (86). Figures 2-5 and 2-S6 demonstrate that endogenous ERK1/2 kinase activity towards Elk1 is severely dampened in the absence of reductant, despite the fact that ERK1/2 is phosphorylated on the activation loop. These results seem to belie the known role of ERK1/2 in proliferative signaling, where ERK1/2 kinase activity is required for the cell response to the proliferative stimuli (55). However, ERK1/2 regulation by cysteine oxidation could be a mode of regulation more complex than simply switching the enzyme activity between on and off states. For example, with its vast array of known substrates, spanning both the cytosol and nucleus, it is possible that ERK1/2 cysteine oxidation occurs to prevent phosphorylation of substrates in the cytosol while ERK1/2 is in route to the nucleus; ERK1/2 in the

72 nucleus would then be subjected to a more reducing environment than in the cytosol (87).

Alternatively, considering of the location of several cysteine residues in

ERK1/2, it becomes apparent that oxidation could control more than just ERK1/2 kinase activity (15,64-67,88-91). Of particular note are cysteines 159, 164 and

252 (rat ERK2 numbering). C252 is in close proximity to the F-recruitment site

(FRS) (92,93), a vital region facilitating interactions between ERK1/2 and many of its substrates, including Elk1. Thus, C252 oxidation could alter the functionality of this protein recruitment site. Additionally, C252 is physically near S246 and

S248, residues that are phosphorylated by casein kinase 2 (CK2) in response to proliferative signals (50,94,95). Phosphorylation of these serines is sufficient and necessary for ERK1/2 translocation to the nucleus. Plotnikov et al. created a model of interaction between ERK2 and CK2, and C252 points directly into the active site of CK2. Thus, oxidation of C252 would very likely affect ERK1/2 localization through its nuclear translocation signal.

Another interesting cysteine, C164, has been shown to be nitrosylated in cells treated with GSNO, a NO donor (67). Nitrosylation of this cysteine appears to lead cells into apoptosis. Thus, oxidation or nitrosylation of this cysteine could prevent, or slow, ATP binding and exchange, thereby inhibiting kinase activity.

C159 makes a major contribution to the D-recruitment site (96-102), another important region facilitating protein-protein interactions between ERK1/2 and its regulatory proteins such as MKP3 (103-105), MEK1/2 (106,107), and anchors and scaffolds (108). Many substrates also interact with ERK1/2 in this

73 binding region. Structural studies reveal that hydrophobic residues of interacting proteins insert into hydrophobic pockets on the surface of ERK1/2 on either side of C159 (93,97,100,109-111). Thus, oxidation of this residue could sterically prevent ERK1/2 from interacting with DRS partners. Indeed, DRS-binding peptides protect C159 from alkylation (102,112). Furthermore, analysis of known

ERK2 structures reveals that C159 could hydrogen-bond with H123, potentially lowering the pKa of C159 and making it more reactive with oxidants (56,63).

Similar to how phosphorylation of ERK1/2 on key residues affects ERK1/2 structure and function, oxidation of any of these cysteines would affect the surface, structure, and dynamics of ERK1/2 in key regions of ERK1/2 functionality (figure 2-6A.) Moreover, the observation that sulfenic acid formation occurs with different temporal dynamics between distinct signals and cell lines is reminiscent of how the temporal dynamics of ERK1/2 TEY-phosphorylation profoundly impacts on cell response to stimulus(62) (Figure 2-6B).

To be able to develop therapeutics that target ERK1/2 in specific signaling conditions, there is still a need for a more solid understanding of the molecular details that govern how ERK1/2 signal transduction induces distinct biological responses. By identifying that ERK1/2 cysteine residues are directly oxidized as a result of extracellular signals, our experiments have revealed cysteine oxidation of ERK1/2 to be a realistic way for cells to regulate ERK1/2 activity. Several cysteine residues on ERK1/2 are candidates as sites of oxidation which would regulate of ERK1/2 activity through modulation of subcellular localization, protein- protein interactions, and substrate specificity. Additionally, we have

74 demonstrated that endogenous ERK1/2 oxidation dramatically decreases kinase activity. This observation underscores the fact that TEY-phosphorylation is not the sole factor determining ERK1/2 kinase activity, as it is often assumed in the literature (88). Further studies to understand how ERK1/2 is regulated by oxidation are absolutely essential to illuminate modes of ERK1/2 signaling specificity, potentially leading to the development of novel therapeutics to target

ERK1/2 in signal-specific redox states.

Figure 2-6. Potential modes of ERK regulation. A) ERK regulation by PTMs. 1) Without any phosphorylations, the activation loop conceals the active site and ERK is essentially turned “off.” 2) Upon double phosphorylation on T183 and Y185 (TEY motif) by MEK1/2, a conformational change in the activation loop opens up the active site and renders the kinase activity “on.” 3) Double phosphorylation on the SPS-motif in the NTS by CK2is necessary and sufficient for ERK translocation to the nucleus. 4) As evidenced in various cell types within this chapter, unphosphorylated (4) and TEY- phosphorylated (5) ERK undergoes active cysteine sulfenylation in response to physiologically-relevant proliferative signals. Like phosphorylations that modify the charge, structure, and dynamics of specific regions of ERK, cysteine modification could lead to significant changes in ERK regulation. Of particular interest are cysteines 159, 164, and 252, as described in Conclusions. It is not yet known how these cysteine modifications crosstalk with phosphorylation PTMs or how these oxidations affect ERK activity in cells.

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Supplementary data (Keyes et al.)

Results

Figure 2-S1: ERK1/2 biotinylation is due to labeling by DCP-Bio1 and not endogenous biotinylation. To verify that biotinylation of ERK resulted from the sulfenic acid trap DCP-Bio1 and not from endogenous biotinylation in the cells, NIH-3T3 cells were treated with PDGF for various time points and harvested either in the presence or absence of DCP-Bio1. Samples from all three time points without DCP-Bio1 were combined, and all samples were then subjected to biotin- affinity capture with streptavidin-agarose beads as described in Figure 1 and Materials and Methods. Images are representative of 2 biological replicates. Parallel control experiments were also performed in SKOV3 and Hela cells (data not shown).

Result: Immunoblot for total-ERK1/2 reveals biotinylation of ERK only when DCP-Bio1 is included in lysis buffer.

Figure 2-S2: Oxidation of ERK1/2 in prostate cancer-derived PC3 cells in response to LPA. PC3 cells grown in 10% serum were treated with 100 nM LPA for 5, 10, or 30 minutes before lysis in presence of DCP-Bio1, as described in Fig. 2-1. Shown in (A) and (B) are Western blots of biotinylated proteins after affinity capture, blotted for total (left panels) and doubly (TEY) phosphorylated (right panels) ERK1/2. Lower panels of each show quantitation of ERK1/2 Western blots as described in Materials and Methods. * p<0.05, ** p<0.005, *** p<0.001.

Result: Sulfenylation of total-ERK1/2 is highly variable in PC3 cells, but it appears to be relatively low at 10 minutes post-stimulus and peak 30 minutes post-stimulus. Due to variability, pre- stimulation sulfenylation of ERK1/2 may or may not occur similar to WI-38 cells (Figure 2-4C, D) under serum-replete conditions. The pool of doubly phosphorylated ERK exhibits greater oxidation at later time points (10 and 30 min).

Figure 2-S3: ERK1/2 oxidation in ovarian-cancer derived SKOV3 cells. SKOV3 cells depleted of serum for 18 hr were treated with 100 nM LPA and harvested in the presence of DCP-Bio1 as described in Figure 2-1. The upper panel shows representative immunoblots of DCP-Bio1 labeled proteins for total ERK (A) and TEY-phosphorylated ERK1/2 (B). The lower panels depict averaged relative intensity for each sample after LPA treatment (n=6 total-ERK, n=3 for phospho- ERK). * p<0.001, ** p<0.0005

Result: There is no detectable pattern for sulfenic acid formation among the total population of ERK1/2 in response to LPA, but it appears that total-ERK1/2 is constitutively oxidized in SKOV3 cells. Among the population of TEY-phosphorylated ERK1/2, however, there is a significant increase in the amount of active sulfenylation of ERK1/2 5 minutes post-stimulus that then drops off to basal levels of sulfenylation 10 minutes post-stimulus.

Figure 2-S4: Observed oxidation of ERK in HeLa cells treated with EGF. HeLa cells were treated with EGF without serum and harvested in the presence of DCP-Bio1 as described in Figure 2-1. Panels (A) through (C) show immunoblots for total ERK (upper panels) and the internal AhpC controls (lower panels) for three different biological replicates (D) Representative total ERK1/2 immunoblot from whole cell lysates.

Result: While ERK does undergo active oxidation in HeLa cells, timing and relative extent of ERK oxidation is highly variable and may not occur in response to EGF.

Figure 2-S5: Observed oxidation of total and TEY-phosphorylated ERK1/2 in response to Androgen agonist R1881 in prostate-epithelium derived RWPE-1 cells. RWPE-1 cells were cultured as described in Materials and Methods and were treated with the androgen agonist R1881, then cells were lysed in presence of DCP-Bio1 as described in Figure 2. (A) First replicate showing that both total and TEY-phosphorylated ERK1/2 were maximally labeled 5 minutes after stimulation by DCP-Bio1. Due to technical issues, AhpC control band was unable to be visualized for this set of samples. Thus, quantitation for relative changes in DCP-Bio1 cannot be assumed. (B) A second replicate showing total-ERK oxidation decreasing in response to R1881 and (C) TEY-phosphorylated ERK oxidation increasing in response to R1881.

Result: Although variability between replicates makes the temporal response of ERK oxidation unclear, we are able to conclude that ERK does undergo oxidation in R1881-treated RWPE-1 prostate epithelium cells.

Figure 2-S6: Endogenous ERK oxidation inhibits kinase activity towards Elk1 in SKOV3 cells treated with LPA. TEY-phosphorylated ERK1/2 was immunoprecipitated from SKOV3 cells treated for various times with LPA and assessed for kinase activity with Elk1 in the presence or absence of dithiothreitol as described in figure 5. (A) Representative immunoblot of Elk1 (upper panel) and band density quantitation (lower panel) from 5 independent replicates. (B) Aggregated and normalized data from 5 independent replicates. At 5 and 10 minutes after LPA treatment, Elk1 phosphorylation levels are ~50% of those obtained in the presence of DTT when ppERK1/2 is left in endogenous oxidation state after immunoprecipitation, indicating that the endogenous ERK oxidation dramatically decreases ERK kinase activity. *P< 0.002 ** p< 0.000001. (C) Immunoblot for TEY-phosphorylated (top) and total (bottom) ERK1/2 from cell lysates. Result: Endogenous ERK oxidation inhibits kinase activity towards Elk1 in SKOV3 cells treated with LPA.

Chapter 3: Modification of rERK2 activity in response to in vitro oxidation by H2O2

Jeremiah Keyes, Derek Parsonage, Kimberly Nelson, Julie Reisz, Robert

Newman, and Leslie Poole

This chapter details preliminary studies investigating the effect of H2O2 in modulating rERK2 activity towards different substrates. Keyes performed protein cloning and purification, rERK2 activity assays using Elk1 as a substrate, protein mobility experiments, and prepared rERK2 for mass spectrometry and protein array experiments. Dr. Parsonage assisted in protein cloning and purification. Dr.

Nelson assisted in experimental design and analysis and helped prepare mass spectrometry figure. Dr. Reisz analyzed rERK2 peptides. Dr. Newman performed and analyzed protein array assays. Dr. Poole performed rERK2 activity assays towards Sub-D and served in an advisory and editorial capacity.

Introduction

ERK1/2 are central to cellular responses to extracellular signals. While the upstream kinases within the ERK MAPK cascade have very narrow substrate profiles, ERK1/2 enzymes have over 300 known and verified substrates – many of which induce opposite cell responses. Furthermore, ERK1/2 shares the same recognition motif as other MAPK enzymes: S/T-P. Given the simplicity of this motif, all MAPK should be able to phosphorylate thousands of proteins without any discrimination between the different kinases. However, the different MAPK enzymes do not share the same set of substrates in vivo. This has been traced to differences in docking interactions outside of the active site.

ERK1/2 enzymes have two primary docking sites facilitating protein- protein interactions, the D-recruitment site (DRS) and the F-recruitment site

(FRS) (1). As discussed in chapter 1, the DRS is important for interactions with scaffold proteins, the upstream kinases MEK1/2, and the phosphatase MKP3.

Several substrates also interact with ERK1/2 via the DRS, and although there is a general consensus sequence among these DRS-docking proteins, not all substrates that have been shown to interact in this region perfectly match the consensus sequence. The FRS has been shown to primarily be utilized for interactions with substrates, primarily transcription factors. Despite the knowledge of these docking sites, it is still not fully clear how ERK1/2 achieves substrate specificity to induce distinct cellular responses.

One of the potential mechanisms to regulate substrate specificity is through the alternate PTM of cysteine modification. The DRS, in particular, has a

90 notable cysteine contributing to the structure of the docking site (2-6). Indeed, it has been shown that when a peptide is docked at the DRS, this cysteine is protected from alkylation (7). Based on crystal structures of peptides docked to the DRS, it is reasonable to assume that alkylation, or cysteine modification, of

C159 would sterically preclude the interaction of DRS-docking proteins with

ERK1/2.

Because of our finding detailed in Chapter 2 that ERK1/2 undergoes cysteine sulfenylation in response to proliferative signals, and that endogenous oxidation inhibits kinase activity, we hypothesized that cysteine oxidation in vitro could modify substrate specificity. To test this hypothesis, we set out to analyze the effect of in vitro oxidation by H2O2 on ERK1/2 activity towards Elk1, towards the peptide substrates specific for the DRS (Sub-D), and towards functional protein arrays in order to analyze global positive or negative specificity changes in ERK1/2 kinase activity (8). Although many of the results in this chapter are preliminary, we have found exciting evidence that ERK1/2 oxidation by H2O2 is not a simple switch between “on” and “off” kinase activity, but is likely a mode to modulate ERK1/2 kinase activity between different classes of substrates.

Materials and Methods

Reagents

γ-32P ATP was purchased from Perkin-Elmer, non-radioactive ATP was from Promega. Catalase was purchased from Sigma. Anti-phospho (S383) Elk1, anti-phospho (T183/Y185) ERK1/2 and anti-ERK1/2 antibodies were from Cell

Signaling Technology, Mouse anti-glutathione antibody was from Virogen. H2O2 was from Fluka. Dimedone and iodoacetamide (IAM) were from Sigma-Aldrich.

Trypsin was from Worthington Enzymes. Functional protein arrays were provided by Rob Newman (North Carolina Agricultural and Technical State University) who purchased them from Heng Zhu at John Hopkins University. Sub-D peptide was synthesized by Dr. Andrew Wommack and Ms. Olivia Tornow essentially as described previously (5).

Protein cloning and expression

Recombinant rat ERK2 (rERK2) was expressed from NpT7 vector obtained from Addgene (Plasmid #39230) essentially as described previously (9-

11). Briefly, proteins from bacterial lysates were applied to a cobalt-NTA column

(GE Healthcare) preequilibrated in 50 mM sodium phosphate at pH 8.0, with 0.3 mM NaCl and 20 mM imidazole, washed with 4 column volumes of the same buffer, then eluted by increasing the imidazole to 500 mM. Fractions containing rERK2 were verified by immunoblot against ERK2 and pooled together, exchanged and concentrated into storage buffer consisting of 20 mM Tris at pH

7.5, pH adjusted at 4 °C with 1 mM EDTA, 1 mM DTT, and 10% glycerol. In preparation for activity assays, rERK2 was phosphorylated in vitro by MKK-G7B, a constitutively-active form of MKK1, as described previously (12). MKK-G7B plasmid (pRSETa) was a kind gift from Natalie Ahn (University of Colorado at

Boulder). MKK-G7B was expressed and purified as described previously (12,13).

After in vitro TEY-phosphorylation by MKK-G7B, rERK2 was applied to a 15 mL

Source Q column to separate mono- and diphosphorylated rERK2 from each

92 other and from unphosphorylated rERK2. Doubly-phosphorylated rERK2 was verified by immunoblot against TEY-phosphorylated ERK2 and by mass spectrometry. GST-Elk1-His6 was expressed and purified as described in

Chapter 2 Materials and Methods. The C2xS double mutant of rERK2 with both

Cys159 and Cys252 mutated to Ser was generated from a DNA sequence synthesized by Genscript using the same nucleotide sequence as for WT (in the

NpT7 vector) except at these two codons. A DNA fragment from the synthesized construct was generated by restriction digests with ApaI and NotI and ligated into the same sites of the ERK2-expressing NpT7 vector. The resulting plasmid was verified by restriction digests and sequencing of the entire coding region of the construct. rERK2 kinetic analyses

For all rERK2 activity assays, 0.5 μM rERK2 was treated with water or

H2O2 in 50 mM Tris-HCl at pH 7.5 for 10 min at 30 ˚C (15 min for assays conducted by immunoblotting for pElk1). Excess H2O2 was then scavenged by the addition of 1 U of Catalase. Samples were further diluted into Kinase Buffer A

(50 mM Tris-HCl at pH 7.5 with 10 mM MgCl2 for Elk1 activity assays,), Kinase

Buffer B (50 mM HEPES at pH 7.5 with 20 mM MgCl2, 40 ug/ml BSA, and 100 mM KCl for Sub-D activity assays), or Kinase Buffer C (50 mM Tris-HCl, pH 7.5,

100 mM NaCl, 10 mM MgCl2, 1 mM MnCl2, 1 mM EGTA, 25 mM HEPES-KOH, pH 7.5) for functional protein array activity assays. In some assays, one half of the peroxide-treated sample was diluted into buffer containing 2 mM DTT while the other half was diluted in Kinase Buffer without DTT (Figure 1A).

For assays with GST-Elk1-His6 using immunoblots to detect phospho-

Elk1, following H2O2 and catalase treatments and addition of DTT to half of each sample, 2 μM recombinant GST-Elk1-His6 and 200 μM ATP were added and incubated for 30 minutes at 30 ˚C. Reactions were quenched by the addition of

2% SDS and 10 mM DTT, and boiled at 100 ˚C for 5 minutes. To detect the activity of rERK2, an immunoblot to detect phosphorylation at S383 of Elk1 was performed. For quantitative assays with radioactive ATP, 25 nM rERK2 was incubated with 2 μM GST-Elk1-His6 and 50 μM ATP (500 counts per minute

(CPM) per pmol ATP) at 30 ˚C. For Sub-D assays, peptides were included at 100

μM and ATP concentration was 300 μM. For time-dependent kinase assays with

Elk1 or the peptide substrates, aliquots of each reaction mixture were removed and quenched at various time points by adding 10 μL of the reaction mixture to

200 μL 0.5 M EDTA at pH 7.0. Each sample was then applied to a Bio-Rad Dot-

Blot apparatus and proteins or peptides were captured on a nitrocellulose membrane while excess ATP was removed by washing each spot on the Dot Blot

4 times with 50 mM Tris-buffered saline (TBS, containing 0.9% w/v NaCl). After washes, the nitrocellulose was removed and dried for 10 minutes, then individual excised samples were added to 3 mL of ECOLUME ES scintillation fluid from

MPBio in vials for counting on a Beckman-Coulter LS 6500 multi-purpose scintillation counter. Data were analyzed using Microsoft Excel and Kaleidagraph

4.5. For H2O2-treated and untreated pairs of samples, rates were normalized to that of fully-reduced, H2O2-untreated rERK2. A general workflow for all activity assays is described in Figure 3-1A.

Protein chip assays were conducted essentially as described previously

(14). Briefly, rERK2 in Kinase Buffer C, treated or not with H2O2 as described above, was applied to slides arrayed with sets of 400 proteins and covered with a glass cover slip. Slides were placed in a humidity chamber and incubated at 30

˚C. After 30 minutes, reactions were terminated by dipping each slide into 500 mL TBS/T (TBS containing 0.1% Tween-20). Slides were then washed three times for 10 minutes each with TBS/T, and then with 0.5% SDS, followed by a quick wash in warm deionized water. Slides were dried and then exposed to high-resolution BIOMAX MR film from Carestream for various times. Film was scanned and analyzed using GenePix software from Molecular Devices.

Endogenous ERK1/2 oxidation detected by gel electrophoresis

SKOV3, HeLa, and NIH-3T3 cells were cultured and treated with LPA,

EGF, and PDGF, respectively, as described in Chapter 2 Materials and Methods.

Protein content for cell lysates was quantified using a BCA protein assay kit from

Thermo Scientific. For resolution of proteins on gels, equal quantities of protein were denatured either in non-reducing or reducing (10 mM DTT) sample buffer

(50 mM Tris-HCl at pH 6.8, with 2% SDS, 0.04% bromophenol blue, and 10% glycerol), incubated at 100 °C for 5 min, then loaded onto 10% SDS- polyacrylamide gels and electrophoresed. Proteins were then transferred to nitrocellulose membrane, which was blocked with 5% powdered milk in TBS/T and then immunoblotted for total-ERK (recognizing both ERK1 and ERK2).

Identifying redox-sensitive cysteines

rERK2 was treated with 0, 100, 500, or 1000 μM H2O2 in the presence of 7 mM dimedone in 50 mM Tris pH 7.5 for 1 hour at room temperature. Reactions were quenched by addition 200 mM IAM at room temperature in the dark for 10 minutes, followed by dilution into 6M Guanidine HCl. Denatured protein was incubated in these conditions with 50 mM IAM for 1 hour at room temperature in the dark. Samples were then precipitated in cold methanol overnight at -20 ˚C.

After washing samples, they were then resuspended in 40 mM ammonium bicarbonate, 1 mM CaCl2, and 10% acetonitrile and digested by trypsin at 37 ˚C overnight. Peptides were then analyzed on a Thermo Scientific LTQ Orbitrap XL.

For gel analysis of WT and C2xS rERK2 under reducing and non-reducing gels, proteins were treated with varying concentrations of peroxide and prepared for electrophoresis as described above. Gels were rinsed with deionized water and stained with Gel Code Blue from Fisher Scientific.

Results

rERK2 activity towards Elk1 is modestly inhibited by H2O2

To test the effect of H2O2 on the activity of recombinant ERK2 kinase, purified rERK2 was treated with varying concentrations of H2O2 for 10 or 15 min at 30 ˚C, then treated with catalase and assessed for activity in the presence or absence of DTT using immunoblots for phospho-Elk1 or radioactivity assays for incorporation of 32P into Elk1 protein (Figure 3-1A). For immunoblot analysis,

H2O2-treated or untreated rERK2 samples were incubated with recombinant

GST-Elk1-His6 and ATP in the presence or absence of DTT for 30 minutes, then

96 immunoblotted to detect phosphorylation at S383 of Elk1 (Figure 3-1B). Modest inhibition of rERK2 activity towards Elk1 was detected at 10, 100, and 1000 μM

H2O2 concentrations; we occasionally observed a slight increase in the activity of

0 or 1 μM H2O2-treated rERK2 when DTT was not present in reaction. However, these results were quantitatively inconsistent, therefore we were not able to determine percent changes in kinase activity between the differentially treated rERK2 conditions using this technique.

To further quantitate the effect of H2O2 on rERK2 kinase activity towards

Elk1, we measured activity using γ-32P ATP to track the extent of phospho- transfer to substrate, as described in Materials and Methods. Initially, significant inhibition of rERK2 activity was observed in reactions without DTT, and there was evidence of partial irreversible inhibition in the H2O2-treated samples (Figure 3-

1C). The reversible, diminished activity observed in H2O2-untreated rERK2 indicates that adventitious oxidation is occurring. (While only one replicate is shown in Figure 3-2C, very similar results were observed multiple times using other techniques.) In subsequent experiments, atmospheric O2 was removed from buffers by bubbling all buffers with argon and flushing tubes with argon each time they were opened. Under these O2-depleted conditions, we observed abrogation of adventitious inhibition by oxidation and a minimal effect of H2O2 on rERK2 kinase activity (data not shown). These data indicate that the adventitious oxidation, and the H2O2-induced irreversible inhibition on rERK2 kinase activity is dependent largely on atmospheric oxygen. To minimize adventitious and irreversible oxidation, subsequent experiments were conducted with activity

97 assays for non-DTT reactions performed as soon as possible after H2O2 treatment. Under these conditions, we observed a modest, dose-dependent effect of H2O2 on rERK2 kinase activity towards Elk1 (Figure 3-2D).

Figure 3-1: rERK2 activity towards Elk1 is inhibited by adventitious and H2O2-dependent oxidation. A) General scheme for rERK2 activity assays used throughout Chapter 3 (See figures 3-1, 3-5, 3-6). rERK2 treatment with H2O2 is followed by addition of Catalase for removal of excess H2O2, and splitting of sample into reduced or non-reduced pairs. rERK2 activity is measured by addition of substrate (GST-Elk1-His6 in Figure 3-1, Sub-D in Figure 3-5, or functional protein arrays in Figure 3-6) and ATP. All assays except for Figure 3-1B are performed using γ-32P ATP to measure phospho-transfer to substrate. B) Initial measurement of rERK2 activity towards Elk1 as measured by immunoblot against S383 phosphorylation on Elk1. C) Observation of adventitious and irreversible oxidation inhibiting rERK2 activity towards Elk1. Rates of rERK2 phospho-transfer activity were measured, and we observed that rERK2 activity, without peroxide treatment or reductant, was consistently ~50% of non-peroxide treated rERK2 in the presence of the reductant DTT. We also observed irreversible inhibition in peroxide-treated samples. D) Techniques were developed to minimize irreversible and adventitious oxidation of rERK2. Under these conditions, we observed modest, dose-dependent inhibition of rERK2 activity towards Elk1. * indicates statistically significant differences in the rate of rERK2 activity as compared to 0 H2O2 + DTT; ^ indicates statistically significant differences in the rate of rERK2 activity as compared to 0 H2O2 –DTT. * p < 0.005, ** p < 0.00002, *** p < 1x108, ^ p < 0.01, ^^ p < 0.005, ^^^ p < 0.00001.

ERK1/2 oxidation does not result in detectable formation of intra- or inter-subunit disulfide bonds in the presence of other cellular proteins

We then sought to identify the nature of endogenous oxidation products of

ERK1/2 in situ in order to ascertain whether or not the cell-based oxidation of

ERK1/2 is recapitulated with in vitro oxidation of recombinant ERK2 by H2O2.

SKOV3, NIH-3T3, or HeLa cells were treated with LPA, PDGF, or EGF, respectively, as described in Chapter 2. Cell lysates were resolved by SDS-

PAGE under non-reducing and reducing conditions, and then transferred to nitrocellulose. Western blots for total-ERK1/2 reveal no evidence that ERK1/2 makes intermolecular or intramolecular disulfide bonds given the lack of an apparent molecular weight shift in either direction (Figure 3-2), although this does not rule other the possibility that ERK1/2 makes a disulfide bond with a small- molecule thiol such as glutathione. Western blots of similarly-treated samples probed with anti-glutathione antibodies were conducted and hinted at the potential presence of glutathionylation on ERK proteins from stimulated cells, but

99 these experiments were largely inconclusive without further experimentation and better quality antibodies.

Figure 3-2: ERK1/2 oxidized during cell signaling events does not make inter- or intramolecular disulfide bonds with itself or other proteins. SKOV3, NIH-3T3, and HeLa cells were treated with LPA, PDGF, and EGF, respectively, for various time points and cells were alkylated and harvested as described in Materials and Methods. Cell lysates were subjected to non-reducing (left) and reducing (right) SDS-PAGE and subsequent immunoblot for total-ERK1/2.

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C159, C214, and C252 are sensitive to in vitro oxidation

To identify cysteines sensitive to oxidation, rERK2 was treated with varying concentrations of H2O2 in the presence of the sulfenic acid trap dimedone as described in Materials and Methods. Mass spectrometry analysis of rERK2 peptides identified C159-dimedone adducts in each sample, including untreated

(Figure 3-3 and Table 3-I). At 500 and 1000 μM H2O2, C159 formed sulfinic acid, a modification that is not reversed by DTT. Although spectral quality was not as high quality as for C159, C214 appeared to yield a dimedone adduct at 500 μM

H2O2 and C252 a dimedone adduct at 1000 μM H2O2 (Table 3-I).

Figure 3-3: Identification of rERK2 cysteine sensitivity to sulfenylation by H2O2. rERK2 was treated with 0, 100, 500, and 1000 μM H2O2 for one hour in the presence of 7 mM dimedone at room temperature. Free thiols were then alkylated by iodoacetamide (IAM) and protein was precipitated and resuspended in 10% ACN, 40 mM ammonium bicarbonate then digested overnight with trypsin at 37 ˚C. Peptides were analyzed on a Thermo Scientific LTQ Orbitrap XL.

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Table 3-I: Cysteine Modifications Observed on rERK2 Mock 100 M H2O2 500 M H2O2 1000 M H2O2 C38 IAM IAM IAM IAM C63 IAM IAM IAM IAM C125 IAM IAM IAM IAM IAM IAM IAM IAM C159 dimedone dimedone dimedone dimedone SO2 SO2 C164 IAM IAM IAM IAM IAM C214 IAM IAM IAM dimedone IAM C252 IAM IAM IAM dimedone

Given these data, a mutant rERK2 with both C159 and C252 substituted by serine, designated C2xS, was then tested for its response to H2O2 on non- reducing gels. Recombinant WT and C2xS rERK2 were treated with varying concentrations of H2O2 and subjected to SDS-PAGE gel electrophoresis under non-reducing and reducing conditions. Under these conditions, WT rERK2 formed a small amount of a high MW species in a dose-dependent manner

(migrating at a much larger apparent molecular weight than would be expected for dimers), but C2xS rERK2 did not, indicating that C159 or C252 was responsible for this observed MW shift.

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Figure 3-4: C159/252S (C2xS) double mutant rERK2 responds to H2O2 differently than WT rERK2. WT and C2xS rERK2 were treated with various concentrations of H2O2 and subjected to Non-reducing (left) and reducing (right) SDS-PAGE. Gels were rinsed with deionized water and stained with Gel Code Blue.

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C159S responsible for observed inhibition of ERK2 kinase activity towards DRS- specific substrate

Due to the finding that C159 is extremely sensitive to oxidation, we hypothesized that oxidation of this cysteine is responsible for the observed inhibition of kinase activity. To test this hypothesis, we obtained a peptide-based substrate Sub-D that specifically recognizes the DRS as opposed to Elk1 that binds to ERK via the DRS and the FRS interaction regions (5). We tested the effect of 1mM H2O2 on both WT and C159S activity towards Sub-D. We observed that H2O2 decreased the rate of ERK2 activity towards Sub-D by 40%, whereas

C159S ERK2 was not inhibited by H2O2 (Figure 3-5). Thus, we can conclude that

C159 is at least in part responsible for observed changes in ERK kinase activity as a result of H2O2 treatment. However, further analysis using other cysteine mutants and peptide substrates that are specific for only the FRS are required to fully assess the specific role of other cysteines in modulating ERK activity towards specific substrates. Indeed, it is likely that C159 oxidation would inhibit kinase activity towards DRS-specific substrates but not FRS substrates, thus acting as a switch to modify rERK2 substrate specificity rather than simply inhibiting kinase activity.

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Figure 3-5: C159S responsible for observed inhibition of ERK2 kinase activity towards DRS-binding substrates. WT and C159S rERK2 kinase activity after 1 mM H2O2 treatment was measured by tracking phospho-transfer of 32P onto substrates, as described in Materials and Methods and Figure 3-1A. For WT, n=6; for C159S n=8. * p < 0.005 for comparing difference in activity ratio of H2O2-treated to non-treated of C159S and WT ERK2.

Global changes in rERK2 activity and specificity resulting from H2O2 oxidation

To test the hypothesis that there are global changes in ERK substrate specificity in response to cysteine oxidation by H2O2, we utilized functional- protein microarrays composed of 400 full-length human proteins immobilized on a functionalized microscope slide (Table 3-II). To this end, rERK2 was treated with varying concentrations of H2O2 for 10 minutes at 30 ˚C, excess peroxide was removed by addition of catalase, and samples were diluted into Kinase Buffer 3

105 containing γ-32P-ATP. After incubation for 30 minutes at 30 ˚C with the protein microarrays, the reactions were quenched and the microarrays were exposed to high-resolution film as described in Materials and Methods. To date, these experiments have been conducted on two separate days, with two independent microarrays being used for each condition, each day. Preliminary results from these experiments suggest that rERK2 undergoes dose-dependent changes in activity toward select substrates. Overall, we observed changes in ERK activity towards 21 proteins among the two sets of replicates. ERK2 activity decreased for some substrates, increased for others, and is unchanged for yet others in response to H2O2 (see Fig 3-6 for an example of raw data). Although the results were reproducible using two independent microarrays on each of the days that an experiment was performed, phosphorylation of only one substrate was reproducible across the two different days indicating that further refinement of the experimental conditions are needed to fully test the hypothesis that ERK oxidation modulates substrate specificity and to determine which substrates are affected with higher confidence. However, this preliminary, but exciting, result indicates that, for rERK2, cysteine oxidation is likely a mode for controlling substrate specificity.

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Table 3-II: List of proteins on functional protein array Clone ID Gene symbol Clone ID Gene symbol IOH27760 CSNK1G2 IOH21006 TBK1 IOH21981 DCAMKL2 IOH21855 PNCK IOH26823 PRKCH IOH3648 RPS6KA2 IOH26839 TBK1 IOH54085 ITK IOH26143 RIPK3 IOH63245 NPR2 IOH62118 TNNI3K IOH60533 FLJ25006 IOH26432 NEK10 IOH14340 TRIB3 IOH60102 AURKC IOH22419 BMPR1A IOH26729 BMPR1B IOH22636 ADCK5 IOH27376 TSSK3 IOH3692 AKT1 IOH26033 CAMK1D IOH41408 VRK1 IOH27746 TSSK2 IOH13030 STK38 IOH23113 WNK1 IOH12327 MAPK1 IOH25726 PDK4 IOH61637 SNRK IOH27738 TTBK2 IOH2475 PAK4 IOH27515 STK33 IOH21137 ARAF IOH26807 CAMK2A IOH12032 PDIK1L IOH27465 ZAP70 IOH63228 ACVR2B IOH13704 CSNK2A1 IOH21132 CAMKV IOH9712 PCTK3 IOH62403 GSG2 IOH3435 MAPK13 IOH2412 DYRK2 IOH13493 SCYL3 IOH14023 SGK2 IOH61176 SCYL2 IOH21149 TRIB2 IOH21715 MAP2K3 IOH21117 TYK2 IOH11272 PIM2 IOH11645 BMX IOH12184 CDK4 IOH21152 FGR IOH14583 CDC2 IOH21127 MAPK12 IOH4873 CDK5 IOH63248 RPS6KA3 IOH22804 CAMKK1 IOH21155 OXSR1 IOH59135 PRKY IOH14775 BRSK2 IOH4008 CLK3 IOH2136 CDK9 IOH21860 ACVR1 IOH14307 PXK IOH21077 MAP3K7 IOH20961 PAK6 IOH21077 MAP3K7 IOH20970 DCAMKL1 IOH21160 CSNK1E IOH21081 FYN IOH21048 ACVRL1 IOH63235 ACVR2B IOH21042 BUB1 IOH6643 CLK3 IOH21026 CSNK1G1 IOH4507 GSK3B IOH21007 CLK1 IOH6368 RIPK2

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Clone ID Gene symbol Clone ID Gene symbol IOH5845 MAPK7 IOH3969 LOC51035 IOH5262 AAK1 IOH4079 S100A11 IOH4506 MATK IOH4113 ZNF593 IOH10843 ULK4 IOH4157 ITM2C IOH10145 PDK3 IOH6323 NR1H3 IOH5257 HSPB8 IOH9779 IHPK1 IOH10103 PRKACB IOH13601 LPXN IOH7002 ALS2CR2 IOH13735 ABI1 IOH6284 IKBKB IOH21137 ARAF IOH4032 PHKG2 IOH3908 ITGB4BP IOH3889 MAPKAPK3 IOH3935 CIAO1 IOH6258 PCTK1 IOH3957 SEC22B IOH10813 ACVR1C IOH3970 SERBP1 IOH10417 CSNK1G2 IOH4081 NDE1 IOH27916 STK25 IOH4117 TAGLN2 IOH6107 ARAF IOH4161 C16orf67 IOH5352 MKNK1 IOH7243 SH3YL1 IOH3192 SGK IOH9633 KCNIP3 IOH5866 PDXK IOH14010 CA12 IOH6848 DBN1 IOH13974 BCKDK IOH13374 OSGEPL1 IOH21104 LYPLA1 IOH12032 PDIK1L IOH3910 CBARA1 IOH14139 ERRFI1 IOH3937 TMEM14C IOH3903 ATP5E IOH3958 SAT1 IOH3930 IFI35 IOH3973 C9orf95 IOH3952 CCDC86 IOH4089 SOD1 IOH3968 PHF23 IOH4119 CNOT2 IOH4078 HN1 IOH4167 TARS2 IOH4112 NDUFB7 IOH7249 MYOT IOH4140 PSMC2 IOH10705 PANK3 IOH5913 TRIM39 IOH14516 TRIML1 IOH6899 FN3KRP IOH13976 NRBF2 IOH10757 FGF12 IOH21110 TOE1 IOH13899 FAM62B IOH3912 RAP1GDS1 IOH2268 VPS4B IOH3939 C9orf32 IOH3907 TMEM147 IOH3959 C14orf122 IOH3933 SCAMP2 IOH4068 PSME3 IOH3953 C1orf43

108

Clone ID Gene symbol Clone ID Gene symbol IOH4090 ATP6V1D IOH4138 NFKBIA IOH4125 DDOST IOH4181 CNIH4 IOH4169 GPSN2 IOH6787 RXRA IOH7464 ATPBD1C IOH29276 MAP2K4 IOH12929 NR4A2 IOH26280 CDCA7L IOH13607 NR3C1 IOH27006 VASH1 IOH14772 BTN2A2 IOH7356 ZMYM6 IOH21106 PLCD4 IOH12318 KLRG1 IOH3918 RPL27 IOH40810 IGKC IOH3941 RPS27A IOH40822 C16orf14 IOH3961 TM4SF4 IOH40905 C3orf54 IOH4069 RTN4 IOH27327 CYP4F12 IOH4105 WDR8 IOH10863 PPWD1 IOH4127 MTCP1 IOH10022 TMEM66 IOH4171 NUDT9 IOH6905 HLX1 IOH5756 STAC3 IOH27629 ZNF785 IOH13481 HPCAL1 IOH26668 COMTD1 IOH11681 SFN IOH9910 CCNDBP1 IOH14670 TRIM5 IOH6644 TMEM79 IOH3882 SAE1 IOH13069 NDRG4 IOH3920 LGALS3 IOH40811 SPHAR IOH3944 CBR1 IOH40826 LATS1 IOH3962 HRASLS3 IOH40907 C6orf89 IOH4071 MECR IOH27387 PACSIN1 IOH4108 NPC2 IOH21706 KLHL6 IOH4136 UROS IOH10530 HSPB1 IOH4176 LRRC61 IOH6923 IGSF21 IOH5938 CAPN1 IOH25891 SLC27A6 IOH11024 PMS2L5 IOH26770 C12orf40 IOH13806 MYF6 IOH10044 COG8 IOH14570 CAPN2 IOH12781 F11R IOH3886 PLEKHJ1 IOH4613 BUB3 IOH3926 FGFBP1 IOH40813 SLC2A12 IOH3948 IFIT3 IOH40830 BCDIN3 IOH3967 TCF19 IOH40910 C11orf48 IOH4076 PIR IOH27443 COL23A1 IOH4109 HLA-E IOH21818 GABRD

109

Clone ID Gene symbol Clone ID Gene symbol IOH3160 THOC3 IOH26021 CTDSPL2 IOH7216 NDUFA2 IOH22169 MRPL50 IOH26614 CHST14 IOH6177 OTUB1 IOH27238 GLRA3 IOH14187 NCLN IOH3008 MOBK1B IOH22453 RNF182 IOH5127 MALL IOH40819 SFXN3 IOH5300 ANXA7 IOH40898 RTN1 IOH40815 S100A12 IOH26713 TBC1D7 IOH40841 ZNF84 IOH11561 KCNS2 IOH40915 GAS7 IOH9803 LOC146325 IOH27520 CPNE3 IOH26573 CLCNKA IOH21864 SLC7A3 IOH3868 PLEKHA1 IOH3083 ARL5A IOH26279 SURF5 IOH7339 NME7 IOH12619 C19orf23 IOH39582 PPM1B IOH6341 FLJ20105 IOH27406 PDPK1 IOH14548 ETFDH IOH26105 GTPBP4 IOH44362 EPO IOH28188 OR5F1 IOH40820 KLHL23 IOH7399 SLC25A17 IOH40899 SEC13 IOH40817 CD3D IOH27132 EIF5 IOH40845 PKN3 IOH11603 TBX5 IOH40917 PTCD3 IOH9983 PPM1M IOH11702 UCHL5 IOH6191 ELOF1 IOH13344 DNTTIP1 IOH4503 GPBP1 IOH22415 PPP2R2C IOH4523 TMEM59 IOH7401 BLVRA IOH3147 KRT8 IOH26986 AOC3 IOH3983 SLC29A1 IOH21800 TBN IOH4504 DDX50 IOH3538 UBE2N IOH3995 RPL4 IOH22545 TMPIT IOH4505 AKR1B1 IOH12766 ADAM15 IOH3996 C16orf33 IOH40818 FUNDC2 IOH3706 MEST IOH40849 GOLGA2 IOH4513 FAM53C IOH26505 LOC124446 IOH4606 ATG4B IOH11698 DMD IOH3762 ANAPC5 IOH9777 MLX IOH3713 MRPL4 IOH26225 DDX6 IOH4030 ADI1 IOH7437 BZW2

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Clone ID Gene symbol Clone ID Gene symbol IOH4562 GINS2 IOH13576 MGST2 IOH3438 PHGDH IOH14713 ZNF331 IOH3717 PRNPIP IOH13846 ADM IOH3978 SERPINB6 IOH14206 EIF2A IOH4036 ALDH3B1 IOH13883 TNIP1 IOH14407 RNF167 IOH13915 TMEM39A IOH13517 -- IOH14024 FLOT2 IOH14612 HPGD IOH14231 NSMCE1 IOH14566 DNAJC7 IOH14757 TBCC IOH14158 ZNF266 IOH14847 GNA12 IOH14156 ETS1 IOH14460 PLEKHO1 IOH14208 WDR85 IOH14544 DHRS1 IOH14097 FUBP1 IOH14207 DMKN IOH14697 LAMP1 IOH14724 C1orf166 IOH13553 C7orf42 IOH10227 ACAD11 IOH14434 CD36 IOH13986 FAM189B IOH14290 TFG IOH14098 CUTL1 IOH13870 GGCX IOH14052 RARG IOH14625 PFDN1 IOH14347 ELK3 IOH14811 TSR2 IOH14255 SELI IOH14526 ARHGEF5 IOH14812 HNRPLL IOH13653 IGHM IOH14569 CDC25C IOH14720 LMAN2 IOH14463 APOBEC3C IOH14047 VCAM1 IOH14082 WDR33 IOH14051 RPL31 IOH13969 PARL IOH14046 KPNA3 IOH13857 ANKS6 IOH14457 SLAMF1 IOH13983 UBE2A IOH14505 POLR2D IOH13943 ATP1B3 IOH14687 TMEM59 IOH14819 ANKRD1 IOH14168 NY-SAR-48 IOH13910 DOM3Z IOH14513 SLC35B1 IOH13844 ERGIC1 IOH13651 LOC492311 HeLa lysate HeLa lysate IOH14642 RAB24 IOH13914 MGC29506

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0 μM H2O2

20 μM H2O2

100 μM H2O2

500 μM H2O2

2 mM GSSG

Figure 3-6: Dose-dependent, global changes in ERK2 kinase activity. As described in Materials 32 and Methods, H2O2-treated ERK2 is incubated with P-γATP on functional protein microarrays to test for overall changes in kinase activity resulting from oxidation. Shown above is an example of a pilot array (containing ~ 400 distinct proteins) where protein phosphorylation is measured by assessment of 32P incorporation on proteins. Blue boxes indicate a decrease in ERK2 activity at those locations on the array, yellow boxes indicate an increase in ERK2 activity, and white boxes indicate no change.

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Discussion

Treating rErk2 with H2O2 in vitro results in changes in kinase activity

(Figures 3-1, 3-5, and 3-6). We initially chose Elk1 as a substrate because of its importance as an early-response transcription factor that is activated by ERK1/2 in vivo (1). We observed that rERK2 reacts with H2O2, along with an atmospheric

O2-dependent ROS, to reversibly and irreversibly inhibit its kinase activity (Figure

3-1C). Removing dissolved O2 from buffers, along with keeping argon in the tops of all tubes, led to protection of inhibition by H2O2 and adventitious ROS (data not shown). This finding could mean that H2O2 reacts with rERK2 cysteines, particularly C159 as evidenced by Figure 3-4A and 3-5, to form sulfenic acid.

However, it is possible that sulfenic acid formation isn’t sufficient to inhibit rERK2 activity towards Elk1. However, sulfenic acid could react with dissolved O2 to form the irreversible sulfinic or sulfonic acids, accounting for our observation in

Figure 3-1C). Thus, considering the observation that C159 is sulfenylated before

H2O2 treatment on at least some portion of rERK2 (Figure 3-3, Table 3-I), and sulfinic acid formation was observed at 500 μM H2O2, and that mutation of C159 leads to insensitivity towards inhibition by H2O2 towards the DRS-specific substrate Sub-D, indicates C159 is the cysteine responsible for adventitious and irreversible inhibition of rERK2 activity in vitro. This is co

In testing rERK2 activity towards Elk1 under conditions to minimize this adventitious oxidation, we observed that 100 and 1000 μM H2O2 did lead to slight, but statistically significant, inhibition by partially reversible and partially irreversible oxidation. In particular, 100 and 1000 μM H2O2-treated rERK2 was

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88.4% and 77.2 % as active in the presence of DTT as untreated rERK2. The kinase activity in untreated rERK2 without DTT was 76% that of untreated with

DTT, and activity of 1000 μM H2O2 treated rERK2 without DTT was 58.9% that of fully reduced rERK2 (Figure 3-2C). We also observed that all kinase reactions performed without reductant yielded rates that were statistically different than those of fully reduced rERK2. However, among the DTT-lacking group of assays, only 1000 μM H2O2-treated rERK2 without reductant yielded a rate that was statistically different from that obtained with non-peroxide treated rERK2. This indicates that, under these reaction conditions, in vitro oxidation by H2O2 has a very modest effect on rERK2 activity towards Elk1.

This finding underscored the importance of identifying the in situ oxidation status of ERK1/2. Thus, we sought to determine if ERK1/2 forms intermolecular or intramolecular disulfides by resolving cell lysates on non-reducing polyacrylamide gels. There were no apparent molecular weight shifts in ERK1/2, indicating that ERK1/2 does not make intermolecular disulfides with other proteins at the time points tested in SKOV3, NIH-3T3, or HeLa cells. Nor does it appear that ERK1/2 proteins make any intramolecular disulfide bonds based on the lack of downward shifts in mobility on non-reducing gels. However, this finding does not rule out the possibility that ERK1/2 makes a disulfide with a small-molecule thiol such as glutathione. We sought to test this hypothesis in different ways, including conducting immunoblots against glutathione in ERK1/2- immunoprecipitated samples and assaying rERK2 treated at various stages with

114 reduced and/or oxidized glutathione, but results were largely inconclusive (data not shown).

To better understand how H2O2 affects ERK function, we used mass spectrometry to identify rERK2 cysteine residues that are sensitive to sulfenylation by H2O2. C159 was strongly labeled in all samples (Table 3-I), including the non-peroxide treated sample (only 100 μM H2O2-treated sample is shown in Figure 3-3). As discussed above, observations of adventitious sulfenylation and sulfinic acid oxidation on C159 match activity results in Figure

3-1C. We also observed that C214 was sulfenylated at 500 μM H2O2, although this spectrum was weak and contained many unidentified peaks. Furthermore, this cysteine is not solvent exposed in either the non-phosphorylated or TEY- phosphorylated rERK2 structures (15,16), so it is currently unclear if this oxidation would happen in vivo. C252 was sulfenylated in the 1000 μM H2O2- treated sample, but this was also not the strongest-quality spectrum. C252 is solvent-exposed, however, and it is more likely to be able to react with ROS in vivo. While it is not solidly determined at this time whether or not C214 or C252 are sensitive to oxidation, their oxidation at higher concentrations of peroxide would imply that ERK1/2 is able to sense and respond to different levels of ROS accumulation in cells.

WT rERK2, treated with H2O2 in vitro, induces generation of a minor band with a high apparent molecular weight on 10% SDS-PAGE gels run under non- reducing conditions, indicating that in vitro oxidation elicits some intermolecular disulfide bond formation. Interestingly, the shifted band corresponds to just about

115

116 kDa according to the ladder, suggesting the formation of a trimer of rERK2, which has never been reported in the literature. The origin and structural nature of this apparent high molecular weight species is not yet clear. However, we can conclude from Figure 3-4 that the MW shift is due to contributions from either

C159 or C252, because the C2xS rERK2 double-mutant does not form this band.

Thus, WT rERK2 and C2xS do not respond to in vitro incubation with H2O2 in the same fashion, suggesting that the observed inhibition in Figures 3-1B and 3-2C is due to either C159 or C252.

Because of the proximity of C159 to the DRS, respectively, we sought to test the effect of peroxide on rERK2 activity towards the DRS specific substrate

Sub-D. We found that, like with Elk1, treating WT ERK2 with 1 mM H2O2 led to a

60% reduction in activity towards Sub-D. (Figure 3-5). However, C159S mutant was not inhibited by H2O2, indicating that C159 oxidation is responsible for observed inhibition of activity.

We also sought to test if H2O2-induced oxidation could have a widespread effect on rERK2 activity by incubating reduced and oxidized rERK2 with functional protein array chips, in which there are approximately 400 unique proteins on the slide. These particular chips are pilot arrays designed to prepare for a more rigorous assay with chips containing over 4000 unique human proteins. Our preliminary results with the pilot arrays indicate that H2O2 has a differential effect on rERK2 activity between substrates; activity was decreased towards some substrates and increased towards others in a dose-dependent manner (Figure 3-6). This indicates that oxidation of ERK is not a simple mode to

116 switch ERK activity between “on” and “off” modes. Rather, oxidation has the power to change substrate specificity, which would be an important way to regulate ERK1/2 activity in vivo, given its many known substrates (17).

In conclusion, it is clear that H2O2 is able to react directly with rERK2 cysteines, particularly C159, which can have a major effect on ERK1/2 protein- protein interactions. However, it appears that ERK oxidation is complex and we have yet to identify all possible oxidations or what the endogenous oxidation state(s) is/are and how to replicate that in vitro. However, the work outlined in this chapter underscores the need to elucidate how cysteine oxidation affects ERK structure, function, and kinase activity among various substrates.

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References

1. Chuderland, D., and Seger, R. (2005) Protein-protein interactions in the regulation of the extracellular signal-regulated kinase. Molecular Biotechnology 29, 57-74 2. Piserchio, A., Warthaka, M., Devkota, A. K., Kaoud, T. S., Lee, S., Abramczyk, O., Ren, P. Y., Dalby, K. N., and Ghose, R. (2011) Solution NMR Insights into Docking Interactions Involving Inactive ERK2. Biochemistry 50, 3660-3672 3. Burkhard, K. A., Chen, F., and Shapiro, P. (2011) Quantitative Analysis of ERK2 Interactions with Substrate Proteins ROLES FOR KINASE DOCKING DOMAINS AND ACTIVITY IN DETERMINING BINDING AFFINITY. J. Biol. Chem. 286, 2477-2485 4. Ma, W. F., Shang, Y. A., Wei, Z. Y., Wen, W. Y., Wang, W. N., and Zhang, M. J. (2010) Phosphorylation of DCC by ERK2 Is Facilitated by Direct Docking of the Receptor P1 Domain to the Kinase. Structure 18, 1502-1511 5. Lee, S., Warthaka, M., Yan, C., Kaoud, T. S., Ren, P., and Dalby, K. N. (2011) Examining Docking Interactions on ERK2 with Modular Peptide Substrates. Biochemistry 50, 9500- 9510 6. Critton, D. A. y., Tortajada, A., Stetson, G., Peti, W., and Page, R. (2008) Structural Basis of Substrate Recognition by Hematopoietic Tyrosine Phosphatase. Biochemistry 47, 13336-13345 7. Abramczyk, O., Rainey, M. A., Barnes, R., Martin, L., and Dalby, K. N. (2007) Expanding the repertoire of an ERK2 recruitment site: Cysteine Footprinting identifies the D- recruitment site as a mediator of Ets-1 binding. Biochemistry 46, 9174-9186 8. Newman, R. H., Hu, J. F., Rho, H. S., Xie, Z., Woodard, C., Neiswinger, J., Cooper, C., Shirley, M., Clark, H. M., Hu, S. H., Hwang, W., Jeong, J. S., Wu, G., Lin, J., Gao, X. X., Ni, Q., Goel, R., Xia, S. L., Ji, H. K., Dalby, K. N., Birnbaum, M. J., Cole, P. A., Knapp, S., Ryazanov, A. G., Zack, D. J., Blackshaw, S., Pawson, T., Gingras, A. C., Desiderio, S., Pandey, A., Turk, B. E., Zhang, J., Zhu, H., and Qian, J. (2013) Construction of human activity-based phosphorylation networks. Mol. Syst. Biol. 9 9. Robbins, D. J., Zhen, E. Z., Owaki, H., Vanderbilt, C. A., Ebert, D., Geppert, T. D., and Cobb, M. H. (1993) REGULATION AND PROPERTIES OF EXTRACELLULAR SIGNAL- REGULATED PROTEIN KINASE-1 AND KINASE-2 INVITRO. J. Biol. Chem. 268, 5097-5106 10. Wilsbacher, J. L., and Cobb, M. H. (2001) Bacterial expression of activated mitogen- activated protein kinases. Methods Enzymol. 332, 387-400 11. Heise, C. J., and Cobb, M. H. (2006) Expression and characterization of MAP kinases in bacteria. Methods 40, 209-212 12. Shapiro, P. S., Vaisberg, E., Hunt, A. J., Tolwinski, N. S., Whalen, A. M., McIntosh, J. R., and Ahn, N. G. (1998) Activation of the MKK/ERK pathway during somatic cell mitosis: Direct interactions of active ERK with kinetochores and regulation of the mitotic 3F3/2 phosphoantigen. Journal of Cell Biology 142, 1533-1545 13. Mansour, S. J., Candia, J. M., Matsuura, J. E., Manning, M. C., and Ahn, N. G. (1996) Interdependent domains controlling the enzymatic activity of mitogen-activated protein kinase kinase 1. Biochemistry 35, 15529-15536 14. Zhu, J., Liao, G. L., Shan, L., Zhang, J., Chen, M. R., Hayward, G. S., Hayward, S. D., Desai, P., and Zhu, H. (2009) Protein Array Identification of Substrates of the Epstein-Barr Virus Protein Kinase BGLF4. Journal of Virology 83, 5219-5231

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15. Zhang, F. M., Strand, A., Robbins, D., Cobb, M. H., and Goldsmith, E. J. (1994) ATOMIC- STRUCTURE OF THE MAP KINASE ERK2 AT 2.3-ANGSTROM RESOLUTION. Nature 367, 704-711 16. Canagarajah, B. J., Khokhlatchev, A., Cobb, M. H., and Goldsmith, E. J. (1997) Activation mechanism of the MAP kinase ERK2 by dual phosphorylation. Cell 90, 859-869 17. Maik-Rachline, G., and Seger, R. (2016) The ERK cascade inhibitors: Towards overcoming resistance. Drug Resistance Updates 25, 1-12

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Chapter 4: Discussion and Conclusions

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The vital role that ERK1/2 proteins play in signal transduction is absolutely vital for the growth, development, and function of all eukaryotic organisms (1).

Furthermore, their misregulation makes a well-established contribution to a host of human diseases including cancer, diabetes, and immunological disorders

(2,3). Due to the ubiquitous expression of activity of ERK1/2 throughout the body and its common use as a signal transducer for a variety of contrasting extracellular signals, it is necessary to better understand how signals differentially regulate ERK1/2 signaling to elicit specific cell responses. Research and discoveries outlined in this dissertation provide evidence that ERK1/2 is sulfenylated in response to a variety of different proliferative signals and that endogenous oxidation of ERK1/2 affects phospho-transfer activity, bolstering the hypothesis that an important mode of ERK1/2 regulation is through signal- induced cysteine oxidation. Furthermore, my research provides evidence that

ERK1/2 cysteine oxidation affects ERK1/2 activity by a more intricate mechanism than a simple switch between “on” and “off” modes for kinase activity.

4.1 ERK1/2 signal-dependent sulfenylation as a regulatory PTM

Results outlined in chapter 2 clearly demonstrate that ERK1/2 sulfenylation occurs downstream of RTKs, GPCRs, and HRs during proliferative signaling in both non-transformed and transformed cell lines. Interestingly,

ERK1/2 oxidation in the non-transformed cell lines had very consistent oxidation dynamics (Figures 2-2, 2-4, 2-S5) compared to the more sporadic response in transformed cell lines (Figures 2-S2-S4). While a careful and dedicated analysis between matched transformed and non-transformed cell lines is required to

122 substantiate this observation, the lack of tight control over the dynamics of

ERK1/2 sulfenic acid formation indicates that the mechanisms controlling H2O2 signaling are aberrantly regulated in cancer cells.

The dynamics of ERK1/2 active oxidation to sulfenic acid is significantly dependent on the basal conditions of the cells prior to stimulus (Figure 2-4).

When the human embryonic lung fibroblast cell line, WI-38, was serum-starved for 18 hours prior to PDGF treatment, TEY-phosphorylated ERK1/2 was maximally sulfenylated 5 minutes after treatment, and relative levels of sulfenylation on the total-ERK1/2 population occurred 30 minutes after stimulation. However, when WI-38 cells were treated with PDGF in serum-replete media (with 10% FBS), maximal TEY-phosphorylated ERK1/2 was delayed until

30 minutes after stimulation and total levels of ERK1/2 were already undergoing active oxidation prior to PDGF treatment. Because there is relatively little TEY- phosphorylated ERK1/2 prior to PDGF treatment, this indicates that, under the basal conditions of 10% serum added to the media (which provides a number of unspecified growth factors), non-phosphorylated ERK1/2 is undergoing constitutive oxidation, similar to what is observed in some of the cancer-derived

PC3 and SKOV3 cell lines (Figures 2-S2-3).

The observed differences between total-ERK1/2 and TEY-phosphorylated

ERK1/2 oxidation levels emphasize the need to develop truly quantitative methods to measure ratios of various PTMs and their relationship to each other.

One particular method that can be adapted for this purpose is a recently developed technique that combine sandwich-ELISA (Enzyme-linked

123 immunosorbent assay) with Total internal reflection fluorescence microscopy

(TIRF) for single-molecule counting on a fluorescent confocal microscope (4).

Using this technique, researchers could either measure sulfenic acids using fluorescent conjugated sulfenic-acid traps or measure total levels of a reversibly oxidized protein of interest utilizing the thiol-switch technique discussed in chapter 1. After fluorescently tagging oxidized cysteines in cell lysate by either method described above, the protein of interest is selectively immunoprecipitated onto a microscope slide that has been covalently linked to antibodies recognizing total levels of the protein. Alternatively, the protein of interest could be selectively biotinylated and captured onto a streptavidin-plated microscope slide. After capture of the protein of interest to the slide, the slide would be incubated with additional antibodies that recognize either total levels or phosphorylated forms of the protein of interest and subsequently analyzed to measure ratios of oxidized protein to phosphorylated protein of interest.

In addition to identifying ERK1/2 sulfenylation occurring in response to proliferative signals, Figures 2-5 and 2-S6 reveal that endogenous oxidation of

ERK1/2 significantly inhibits in vitro kinase activity towards the important transcription factor Elk1. However, in vitro oxidation of rERK2 with H2O2 was not as inhibitory as endogenous oxidation assessed by Elk1 phosphorylation

(Figures 2-5, 2-S6, 3-1), despite the confirmation that recombinant rat ERK2 reacts with H2O2 in vitro (Figure 3-3) and the evidence that H2O2 is the ROS molecule necessary for ERK1/2 sulfenylation in NIH-3T3 cells (Figure 2-3). This disparity could result from a number of factors; one primary hypothesis is that the

124 in vitro oxidation of ERK2 by H2O2 does not match the endogenous oxidation because of a missing factor found in cell lysates, such as glutathione. While preliminary data suggest that glutathionylation of rERK2 can affect kinase activity, we were not able to identify whether or not glutathionylation happens in situ (data not shown). We further sought to identify if the endogenous oxidation formed inter- or intramolecular disulfides, but these studies revealed that these types of disulfides do not form within cells, although disulfides with small molecule thiols have yet to be ruled out (Figure 3-2). However, in vitro oxidation by H2O2 does cause a shift in apparent molecular weight on non-reducing SDS polyacrylamide gels for rERK2 (Figure 3-4), substantiating the hypothesis that in vitro oxidation is not fully replicating endogenous oxidation, highlighting the need for further study to identify the nature of endogenous oxidation.

Identification of ERK1/2 sulfenylation occurring in response to a variety of proliferative signals has opened the door to recognizing a novel mode of ERK1/2 regulation, but it has also given rise to many new questions. In particular, why do proliferative signals induce both activation of ERK1/2 via TEY-phosphorylation and inhibition of activity via oxidation? This apparent paradox, along with other new hypotheses resulting from this work, will be addressed further in section 4.4.

4.2 Identification of H2O2-sensitive cysteines on rERK2

ERK2 has 7 cysteine residues, 4 of which are solvent exposed and 1 of which is not conserved between ERK2 and ERK1 (Figure 1-4). To assess whether or not there could be a difference between ERK1 and 2 oxidation and to better understand how oxidation could affect ERK1/2 function, we identified

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H2O2-sensitive cysteines on recombinant, rat ERK2 (Figure 3-3 and Table 3-I).

We found C159 to be extremely sensitive to sulfenylation, forming sulfenic acid even without exogenous addition of H2O2, matching the inhibitory adventitious oxidation we observe in rERK2 activity assays (Figure 3-1C). We also observed the formation of sulfinic acid on C159 when rERK2 was treated with 500 μM and

1000 μM H2O2, also matching the partial irreversible inhibition observed in rERK2 activity assays (Figure 3-1C).

We also observed C214 and C252 sulfenylation starting at 500 and 1000

μM H2O2, respectively, but we did not observe sulfinic or sulfonic oxidation on these cysteines (Table 3-I). C252 is solvent exposed, whereas C214 is buried in a hydrophobic pocket and thus seems unlikely to be able to react with ROS (5,6).

However, if one or both of these cysteines are able to react with ROS in vivo, this differential oxidation susceptibility in vitro could indicate that ERK1/2 is capable of sensing and responding to varying levels of ROS in cells. For example, C159 is more likely to pick up low levels of H2O2 generated during signaling, whereas

C214 or C252 may only undergo oxidation at higher concentrations of ROS formed during oxidative stress. Detailed analysis of ERK1/2 oxidation in cysteine mutant- expressing cells are needed to test this particular hypothesis generated from this body of work. Although the role of these particular cysteines in vivo is not yet established, this work demonstrates that not only are these cysteines able to be sulfenylated in vitro, rERK2 double cysteine mutants do not respond to

H2O2 in the same way WT rERK2 does (Figure 3-4).

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To date, there are other publications that have reported identifying redox- active cysteines on ERK1/2. Galli et al. reported the oxidation of C38 and C214 to sulfinic and sulfonic acids when recombinant rat ERK2 was treated with 0.1

μM H2O2 in vitro (7) (Table 1-II). Although they found that mutating these cysteines negatively impacted the ability of MEK to phosphorylate ERK1/2 in both in situ and in vitro contexts, there are several considerations to be taken into account when comparing their results to the findings in this dissertation. In assessing the oxidation of ERK2, the authors did not alkylate reduced thiols prior to ionization and mass spectrometry analysis, likely leading to artifactual oxidation of cysteine residues during analysis. Furthermore, a structural analysis of the location of these cysteines reveals that both of them are in buried, hydrophobic regions of ERK1/2 structure (5,6). Oxidation of these cysteines to charged species such as sulfinic or sulfonic acids likely would completely alter protein folding; this fact and the inaccessibility of these residues in the folded protein make these modifications unlikely to happen in vivo. Finally, their studies were considering the response of ERK1/2 to the bolus addition of H2O2 to cells, whereas the work focused on in this dissertation has analyzed the oxidative regulation of ERK1/2 in response to physiologically-relevant proliferative signals.

While Galli et al. found many interesting patterns in the H2O2 response of LP07 mouse cancer cells, such as changes in ERK1/2 translocation and perturbed interactions with MEK, it can be concluded that their observations are more relevant to their specific system than to proliferative signaling as a whole.

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Another report by Feng et al. described the nitrosylation of C183 on ERK1

(C164 on rat ERK2, see Figure 1-4 and Table 1-II) in MCF-7 breast cancer cells treated with 1 – 2 mM Sodium Nitroprusside (SNP) (8). Identification of this cysteine was done by expressing point mutations in cells and identifying whether or not each point mutation could be captured by the S-nitrosylation specific biotin-switch technique developed by Jaffrey et al. (9). The authors noted that

C183A did not fully reduce levels of ERK1/2 captured by this technique, but they concluded that C183A is the primary cysteine undergoing nitrosylation in their study. Expression of the C183A ERK1 mutant in MCF-7 cells abrogated SNP- induced apoptosis. This cysteine is found in the ATP binding pocket of ERK2, therefore oxidation of this cysteine could affect ERK1/2 activity by disrupting ATP binding. The authors did not investigate this hypothesis, but were able to show that WT ERK1/2 was dephosphorylated at the TEY-motif in response to SNP, whereas C183A remained phosphorylated. Thus, SNP appears to affect either

MEK or MKP3 activity and C183 on ERK1 is important for the action of one or both of these regulatory proteins on ERK1.

Paige et al. found that C161 (C159 in rat, see Figure 1-4 and Table 1-II), was stably nitrosylated when cell lysates were treated with 1 mM GSNO, or when in cells expressed constitutively-active nitric oxide synthase (10). Identification of this cysteine was done by mass spectrometry following the same biotin-switch method used by Feng et al. While no functional outcome of this nitrosylation was tested, the fact that the authors identified that this cysteine is able to be oxidatively-modified matches the results presented in this thesis. Thus, C159 is

128 potentially able to be modified by multiple types of oxidative species. Full functionality of this cysteine and its modified forms in a cellular environment needs to be assessed to help elucidate the redox regulation of ERK1/2.

4.3 Effect of H2O2 on rERK2 substrate specificity

Due to the sensitivity of C159 towards oxidation, and because of its location within the DRS, we hypothesized that the inhibition of ERK1/2 activity we observed in chapters 2 and 3 may result from changes in protein-protein interactions rather than a direct inhibition of phospho-transfer activity. To address this, experiments using, as substrates, peptides recognized by only one of these motifs (11) are underway (Figure 3-5). Using the peptide substrate Sub-D, we have identified as C159 as the cysteine responsible for observed inhibition of

ERK2 activity towards DRS-interacting substrates. In addition, we have begun testing rERK2 activity towards a large array of proteins using functional protein array chips (See Table 3-II for a list of proteins on pilot arrays). In preliminary experiments, we have observed that rERK2 activity is inhibited in a dose- dependent manner towards some substrates, while activity increases upon oxidation towards other substrates. Currently, these results were obtained using pilot arrays that have ~300 proteins on the chip; these initial results will be validated and extended by repetition on protein chips containing ~4000 proteins.

An analysis on this scale would substantially increase the field’s understanding of how oxidation changes ERK2 substrate specificity.

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4.4 Future Directions

There are many new and unanswered questions and hypotheses resulting from this body of work, many of which are touched on above. However, this section will highlight central hypotheses of how signal-induced oxidation of

ERK1/2 could regulate ERK1/2 activity to elicit distinct, signal-specific responses.

First, what are the roles of C159, C214, and C252 oxidation in cellular responses to stimuli? As mentioned in section 4.2, C159 is much more sensitive to oxidation by H2O2 than C214 or C252. This observation leads to the hypothesis that these cysteines are oxidized for different purposes in a cellular context. Endogenous knockdown of ERK1/2 using CRISPR techniques can be followed by generation of various ERK-expressing cells (WT, C159S, or C252S, for example) to test this hypothesis and to learn the roles of these cysteines in

ERK1/2 signal transduction. In particular, rates of cell proliferation, migration, differentiation, along with analyses of cell cycle distribution, targeted or global mRNA levels, and the phosphoproteome can be assessed for differences between the different ERK cell lines in response to a variety of signals. This in- depth analysis of cellular responses to stimuli would be a powerful addition to the field’s understanding of ERK1/2 regulation.

Second, how does ERK1/2 oxidation affect its spatiotemporal activity in cells? Does oxidation affect substrate specificity in situ? These closely related questions, when answered, will elucidate the mechanism by which particular

ERK cysteine oxidations lead to specific cellular responses. Using the same cell lines expressing WT, C159S, or C252S ERK, ERK-specific biosensors can be

130 utilized to assess where and when the various forms of ERK are active in cells and what type of substrates are being targeted. These biosensors can be designed to interact with ERK1/2 via either the DRS or the FRS site and they can be localized to different regions of the cell including the nucleus, the cytosol, or the leading edge of migrating cells. With these biosensor tools and with the use of live cell and super-resolution imaging, details of how these cysteines control the spatiotemporal dynamics of ERK1/2 activity to induce specific cellular responses to stimuli will emerge.

Third, what role do C159 and C252 play on the interaction of ERK1/2 with regulatory proteins such as MEK1/2, MKP3, CK2, and scaffolds? MEK1/2 are known to interact with ERK1/2 via the DRS, thus C159 oxidation could affect dynamics of ERK1/2 TEY-phosphorylation. Furthermore, MEK acts as an anchor to keep ERK1/2 in the cytosol and can even pull ERK1/2 out of the nucleus.

Thus, cysteine-dependent interactions with ERK1/2 could be important for sequestering ERK1/2 away from the nucleus.

MKP3 activity towards ERK1/2 has been shown to rely on interactions at the DRS (12,13), thus oxidation of C159 could protect ERK1/2 from phosphate removal from the TEY-motif, sustaining its activity. Thus, C159 oxidation could also be a mechanism by which cells differentially regulate the dynamics of TEY- phosphorylation as discussed in section 1.2.2.

Recent work by Rony Seger’s group, as discussed in chapter 1 (14,15), revealed the mechanism of ERK1/2 translocation to the nucleus, which occurs through phosphorylation on a novel Nuclear Translocation Sequence (NTS). The

131 charges introduced by the phosphates on the NTS enables ERK1/2 interactions with importin7/9, which facilitate import into the nucleus. C252 is 4 residues away

(Figures 1-4, 1-5) from the SPS-motif and facing in the right direction to affect these regulatory interactions, thus oxidation of C252 could influence spatiotemporal dynamics via the NTS.

Finally, scaffold proteins have been shown to interact by various means with ERK1/2, primarily via DRS docking (16). Thus C159 oxidation could disrupt binding to scaffolds and therefore be the mechanism by which ERK1/2 is released from scaffolds to interact with substrates and other regulatory proteins throughout the cell. Each of these questions and hypotheses can be tested both by in vitro and cell-based assays with WT and cysteine mutants.

Fourth, it will be important to better understand the spatiotemporal context of where and when ERK1/2 oxidation happens. In redox-mediated signaling,

ROS can be generated by NOX enzymes at the cell membrane (17), within endosome signaling vesicles (Redoxosome model) (18), or by the Mitochondria

(19,20). Using established proximity-ligation techniques in conjunction with sulfenic acid labeling of proteins, researchers can test where ERK1/2 oxidation occurs in relation to other signaling components within the cell. Furthermore, the mechanism and timing of ERK1/2 reduction need to be established – is ERK1/2 reduced after translocation to the nucleus? What mechanisms lead to its reduction? Finally, what subpopulations of ERK1/2 are undergoing oxidation and reduction? Understanding the context of ERK1/2 oxidation could enable the production of highly context-specific therapeutics (21). For example, if it becomes

132 apparent that ERK1/2 is reduced and reactivated by a particular reductase in the nucleus, perhaps targeting that specific reductase will leave ERK1/2 in an oxidized, and inhibited, state, thereby leading to cancer cell apoptosis while leaving healthy cells intact.

4.5 Summary of Contribution to ERK1/2 and Redox Signaling

The findings outlined in this dissertation are of particular importance to the field of MAPK signaling, where there is a current heavy emphasis on overcoming the side effects and developed resistance to ERK1/2 pathway inhibitors for cancer treatment (21-23). The field is now one step closer to understanding the conundrum of signaling specificity, thereby enabling advances in targeted therapeutics to treat not only cancer, but insulin resistance and others.

Furthermore, by identifying that endogenous ERK1/2 oxidation inhibits activity, this work underscores the importance that researchers in the signaling field should not consider the detection of phosphorylation on the activation loop of kinases to be the final determinant of kinase activity, as is often done with

MAPKs.

The field of redox biology and signaling has made vast advancements since Toren Finkel’s landmark study identifying H2O2 as a necessary second messenger in the response of VSMCs to PDGF (24). This body of work further moves this field forward by adding ERK1/2 kinases to the list of proteins identified as direct targets of signal-induced ROS.

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Finally, this body of work provides a new example of the intriguing and sophisticated way that redox modifications can modulate protein activity. Many oxidative modifications of signaling proteins were found to be a simple switch between “on” and “off” modes (notable examples include PTPs and PKA) (25-

27). However, ERK1/2 oxidation may modulate protein-protein interactions without directly affecting phopsho-transfer activity. This is evidenced by the discovery that there are likely at least two ROS-sensitive cysteines on ERK2 with different susceptibilities to sulfenylation, located in non-active site regulatory regions of ERK1/2 structure. Additionally, the preliminary result that oxidation increases activity towards some substrates while decreasing it towards others bolsters this hypothesis and warrants future study.

In conclusion, researchers in the fields of signal transduction and redox biology are now in a better position to understand how signal specificity is achieved through the redox-regulation of ERK1/2. These discoveries will pave the way for the development of new avenues for the treatment of diseases resulting from misregulated ERK1/2-dependent pathways.

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Jeremiah David Keyes 3616 Konnoak Drive • Winston Salem, NC 27127 • Phone: (336) 210-0836 Email: [email protected] or [email protected]

EDUCATION Wake Forest School of Medicine, Winston Salem, NC PhD Candidate, Biochemistry and Molecular Biology, Fall 2010 - Present Expected Graduation: July 2016 Dissertation Title: Signal-dependent modulation of Extracellular signal-regulated kinase activity by reversible cysteine modification

The goal of my project is to discover the mechanisms underlying the redox regulation of Extracellular signal-regulated kinase (ERK) activity and to establish the specific role of ERK oxidation in proliferative signaling. The primary established mode regulating ERK activity is through the Post-Translation Modification (PTM) of phosphorylation, in which ERK is activated by phosphorylation. My work shows that an additional PTM regulating ERK activity is cysteine oxidation by signal-generated reactive oxygen species. I hypothesize that ERK cysteine oxidation regulates ERK function by altering kinase activity, protein-protein interactions, and modification of spatial-temporal localization within cells. This project underscores the importance in considering a variety of PTMs when assessing the regulation ofkinase signal transduction.

Brigham Young University, Provo, Utah B.S. Biochemistry, minor in Theatre Studies - June 2010

TEACHING INTERESTS General Chemistry, Biochemistry, Molecular Biology, Cell Biology, Biophysical chemistry, Scientific Communication, General Biology

TEACHING EXPERIENCE AND TRAINING Subject Matter Expert for Digital Learning Experience – Carbohydrate Chemistry, Spring 2016  Prepared curriculum for a digital resource to replace the traditional textbook for Chemistry 132 (Survey of Organic and Biochemistry) at Forsyth Technical Community College  Curriculum included interactive text, figures, and questions to guide students through understanding the nomenclature, chemistry, and the role of carbohydrates in life.

Workshop: Writing Effective Learning Objectives, Wake Forest Teaching and Learning Center, Jan 2016  Instructed in Bloom’s Taxonomy and its utilization in writing clear learning objectives  Practiced writing learning objectives for recent teaching experiences under mentorship from workshop director

Lecturer: Biochemistry 704 – “Preparatory Biochemistry” – Fall 2015  Team-taught biochemistry course for a pre-medical Master Program in Biomedical Sciences  Prepared and taught three lectures covering amino acids, protein structure, and enzymology  Prepared, administered, and graded exam covering lecture topics  Prepared two practice exams for students to self-test their knowledge

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Facilitator: exam-preparation review session - Fall 2014  Reviewed basic biochemistry principles with 1st year physiology-pharmacology graduate students  Session goals were to prepare students with minimal background in biochemistry for upcoming examinations

Laboratory mentor - Fall 2014 – present  Trained new technician and graduate student in laboratory techniques, including: o SDS-PAGE gel electrophoresis, immunoblotting, protein quantitation, cell culture, molecular cloning, expression of recombinant protein in bacterial and mammalian cells

Guest Lecturer for multiple graduate level courses:  Biological Spectroscopy (MCB 712): Applications of Biological Spectroscopy - Spring 2014 o Prepared and presented concluding lecture in biological spectroscopy course, focused on applying principles to students’ specific research projects o Prepared lecture based on students’ interests in using particular techniques in their research o Prepared and graded exam questions specific to each student’s chosen research project and spectroscopy technique  Molecular and Cellular Biosciences Core Course I: Macromolecular synthesis, structure and function, gene expression and genetics (MCB701) - Fall 2013 o Prepared and presented lecture covering basics of enzyme kinetics o Prepared review questions and held review session to help students prepare for exam  Biological Systems and Structures. (MCB 711) - Fall 2013 o Prepared and presented lecture on protein mutagenesis and engineering o Prepared and graded exam questions covering material covered in lecture

Class: Special Topics in Biochemistry - Fall 2011, 2012, 2014, 2015, 2016; Spring 2012, 2013, 2015, 2016  Participated as a student in discussion-focused class centered on student presentation and examination of primary scientific literature in the fields of biochemistry, molecular biology, and cell biology  Presented several lectures reviewing recent scientific articles  Directed discussion about experiments, techniques, and results of articles with other students and one faculty facilitator

Private Tutor, General Chemistry - Fall 2013  Aided freshman college student in general chemistry  Reviewed concepts discussed in class lectures, including basic chemical principles such as stoichiometry, balancing equations, acid-base and redox reaction chemistry, thermodynamics, gas phase chemistry, atomic theory, and molecular geometry  Prepared problems to practice together during review sessions, reviewed exam problems and gave regular feedback and assessments on student’s progress

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Teaching Assistant, Organic Chemistry Laboratory - Brigham Young University - Summer 2009  Taught basic organic chemistry laboratory procedures to synthesize, isolate, and identify organic compounds  Instructed students in safe laboratory practices  Assisted in laboratory classroom preparation and clean up  Assisted students with homework and in preparation for exams and quizzes  Graded weekly homework and laboratory reports

RESEARCH EXPERIENCE Graduate Research Assistant, Department of Biochemistry, Wake Forest School of Medicine, Fall 2010 – Present Research Advisor: Leslie Poole, PhD  Independently investigating regulation of Extracellular regulated kinase (ERK) by reversible cysteine modification during proliferative signaling  Using both in vitro and in situ methods, determining molecular mechanisms by which oxidation affects ERK function

Laboratory Research Assistant, Department of Chemistry and Biochemistry, Brigham Young University, April 2007 – June 2010 Research Advisor: Richard Watt, PhD  Investigated photocatalytic properties of ferrihydrite iron core within ferritin  Developed technique to utilize ferritin’s photocatalytic capabilities to synthesize stable, soluble gold nanoparticles

LABORATORY SKILLS Protein chemistry, immunoblotting, molecular cloning, protein expression and purification, cell culture, enzyme kinetics, immunoprecipitation, affinity capture, cysteine chemistry, MALDI-TOF mass spectroscopy, radioactive kinase assays, UV-Vis spectroscopy, Proximity ligation, immunofluorescence

PUBLICATIONS Keyes, J. D., Nelson, K. J., Rogers, L., Kesty, C. Parsonage, D., Poole, L. Endogenous Sulfenylation of ERK in Response to Proliferative Signals. (Manuscript to be submitted July 2016.)

Keyes, J. D., Hilton, R. J., Farrer, J., and Watt, R. K. (2011) Ferritin as a photocatalyst and scaffold for gold nanoparticle synthesis, Journal of Nanoparticle Research 13, 2563-2575.

Hilton, R. J., Keyes, J. D., and Watt, R. K. (2010) Maximizing the efficiency of ferritin as a photocatalyst for applications in an artificial photosynthesis system, Proceedings of SPIE 76460J-76460J.

Hilton, R. J., Keyes, J. D., and Watt, R. K. (2010) Photoreduction of Au(III) to form Au(0) nanoparticles using ferritin as a photocatalyst, Proceedings of SPIE 764607-764607.

INVITED TALKS Keyes, J.D. “Redox Biology, Cell Signaling, and Graduate School.” Anderson University Science Club, Invited Seminar Speaker – November 2015

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Keyes, J.D. “Modulation of ERK Activity by Reversible Cysteine Modification.” Gordon Research Seminar: Thiol-Based Redox Regulation and Signaling, Invited Speaker - July 2014

Keyes, J.D. “Modulation of ERK Activity by Reversible Cysteine Modification.” Gordon Research Seminar: Phosphorylation & G-Protein Mediated Signaling Networks, Invited Speaker - June 2014

PRESENTATIONS Keyes, J.D., Nelson, K. J., Kesty, C., Parsonage, D., King, B., Poole, L. B. Modulation of ERK Activity by Reversible Cysteine Modification. ASBMB Special Symposia: Kinases and Pseudokinases: Spines, Scaffolds and Molecular Switches, Poster – December 2015

Keyes, J.D., Nelson, K. J., Kesty, C., Parsonage, D., King, B., Poole, L. B. Modulation of ERK Activity by Reversible Cysteine Modification. ESF/EMBO Thiol Switches in Life Sciences, Poster – September 2015

Keyes, J.D., Nelson, K. J., Kesty, C., Parsonage, D., King, B., Poole, L. B. Modulation of ERK Activity by Reversible Cysteine Modification. FASEB Science Research Conference: Protein Kinases and Protein Phosphorylation, Poster – July 2015

Keyes, J.D., Nelson, K. J., Kesty, C., Parsonage, D., King, B., Poole, L. B. Modulation of ERK Activity by Reversible Cysteine Modification. Future Fellow Research Conference: St. Jude’s Children Research Hospital, Poster – June 2015

Keyes, J.D., Nelson, K. J., Klomsiri, C., Parsonage, D., Poole, L. B. Modulation of ERK Activity by Reversible Cysteine Modification. Gordon Research Conference: Thiol-Based Redox Regulation and Signaling, Poster - July 2014

Keyes, J.D., Nelson, K. J., Parsonage, D., Poole, L. B. Modulation of ERK Activity by Reversible Cysteine Modification. Gordon Research Conference: Phosphorylation and G-protein mediated networks, Poster - June 2014

Keyes, J.D. Modulation of ERK Activity by Reversible Cysteine Modification. Center for Molecular Communication and Signaling of Wake Forest University, Oral Presentation - October 2013, 2014

Keyes, J.D., Klomsiri, C. Nelson, K. J. Parsonage, D., Poole, L. B. Modulation of ERK Activity by Reversible Cysteine Modification. Society for Free Radical Biology and Medicine Conference, Poster - Nov 2013

Keyes, J.D., Klomsiri, C. Nelson, K. J. Parsonage, D., Poole, L. B. Modulation of Kinase Activity by Reversible Cysteine Modification. American Society for Biochemistry and Molecular Biology Conference, Poster - April 2013

Keyes, J.D., Rogers L. C., Klomsiri, C., Nelson, K. J., Daniel, L. W., Poole, L. B. Modulation of lysophosphatidic acid-dependant proliferative and survival signals by localized thiol oxidation. Glutathione and Related Thiols in Living Cells, Poster - September, 2011

Keyes, J.D. Ferritin as a photocatalyst and scaffold for gold nanoparticle synthesis. Utah Conference of Undergraduate Research, Oral Presentation - February 2010

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Keyes, J.D. Ferritin as a photocatalyst and scaffold for gold nanoparticle synthesis. Brigham Young University Physical and Mathematical Sciences Spring Research Conference, Oral Presentation - Spring 2009, 2010

Keyes, J.D. Hilton, R.J. and Watt, R. K. Ferritin as a photocatalyst and scaffold for gold nanoparticle synthesis. Nano Utah Conference, Poster - October, 2009

GRANTS/AWARDS  Research Grants o Mini-grant, Center for Molecular Communication and Signaling, Wake Forest University. Applied for a mini-grant to fund the purchase of materials for the knock out endogenous expression of ERK1/2 in cells and induce expression of mutant forms of ERK using CRISPR/Cas9 technology, $2500 awarded January 2016 o Recipient of T32 Training Grant from Department of Immunology at Wake Forest School of Medicine. Applied for and received competitive support to test for ERK oxidation occurring in a sepsis-signaling cell model. o Student fellow, Center for Molecular Communication and Signaling at Wake Forest University. This is a competitive fellowship that is “intended to foster interdisciplinary collaboration, to catalyze and support research, to promote the training of new researchers in the area of molecular communication and signaling, and to develop new approaches and research directions.” Fellowship paid student stipend from May 2014 – May 2015. o Undergraduate Research Award, BYU Department of Chemistry These awards are competitive proposals to participate as an undergraduate student in a research laboratory at BYU’s Department of Chemistry and Biochemistry - Summer 2007, Summer 2008-Spring 2010  Travel Awards and fellowships o Cowgill Fellow of the Department of Biochemistry and Wake Forest School of Medicine: This is a competitive fellowship awarding $2500 for conference travel expenses for attending conference of student’s choice and - 2014-2015, 2015-2016 Academic Years o Camillo Artom Fellow of the Department of Biochemistry and Wake Forest School of Medicine. This is a competitive fellowship awarding $2500 for conference travel expenses for attending conference of student’s choice - 2013-2014 Academic Year o Graduate Student Travel Award from the American Society for Biochemistry and Molecular Biology to attend the 2013 ASBMB annual meeting. $1000 - April 2013 o Student Travel Grant from European Science Foundation to attend Glutathione and Related Thiols Conference, September 2011, 2015

PROFESSIONAL MEMBERSHIPS ACS, student member AAAS, student member SFRBM, student member

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VOLUNTEER WORK Guest Instructor, Backyard Shakespeare Festival. Summer 2012, 2013  Taught basic presentation and acting skills to 10-18 year-old students at a local one-week summer camp  Mentored students as they practiced incorporating lessons into the scenes they would perform at end of camp

Science Fair Judge, Downtown Arts Based School, Spring 2011, 2012, 2013, 2014, 2015

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REFERENCES Leslie Poole, PhD Professor, Department of Biochemistry Wake Forest School of Medicine Medical Center Blvd. Winston-Salem, NC, 27157 Phone: (336) 716-6711 Email: [email protected]

Robert Newman Assistant Professor, Department of Biology North Carolina A&T State University 1601 E Market Street Greensboro, NC 27411 Phone: 336-285-2189 Email: [email protected]

Allyn Howlett, PhD Professor, Department of Physiology and Pharmacology Director, Integrative Physiology & Pharmacology Graduate Program Director, Office of Postdoctoral Affairs, WFU Graduate School Co-Chair, Health Equities Research Opportunities Fellowship Maya Angelou Center for Health Equities Wake Forest University Health Sciences One Medical Center Blvd. Winston-Salem, NC 27157 Phone 336-716-8545 Email: [email protected]

Richard K. Watt, PhD Associate Professor, Department of Chemistry & Biochemistry C-210 Brigham Young University Benson Building Provo, UT 84604 Phone: (801) 422-1923 Email: [email protected]

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