Regulation of Caspase-9 by Natural and Synthetic Inhibitors Kristen L

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5-2012 Regulation of Caspase-9 by Natural and Synthetic Inhibitors Kristen L. Huber University of Massachusetts Amherst, [email protected]

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REGULATION OF CASPASE-9 BY NATURAL AND SYNTHETIC INHIBITORS

A Dissertation Presented

KRISTEN L. HUBER

Submitted to the Graduate School of the University of Massachusetts Amherst in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

MAY 2012

Chemistry

REGULATION OF CASPASE-9 BY NATURAL AND SYNTHETIC INHIBITORS

A Dissertation Presented

KRISTEN L. HUBER

Approved as to style and content by:

______Jeanne A. Hardy, Chair

______Lila M. Gierasch, Member

______Robert M. Weis, Member

______Peter Chien, Member

______Craig T. Martin, Department Head Department of Chemistry

DEDICATION

For my mother, Elizabeth Huber, who taught me perseverance and to always follow my dreams, no matter how big or small

For my father, Kenneth Huber, who taught me to believe in myself and to remember you can always find a new beginning in tomorrow

For my sister, Elyse Huber, for being the person who inspires me every single day and who taught me to be proud of who I am

ACKNOWLEDGMENTS

To my mentor, Professor Jeanne Hardy, I thank you for guidance and support over this long and somewhat bumpy road. We have both grown in many ways since our first year here at UMass; you as a mentor, me as a budding scientist and both as individuals. Between the memories of our scientific travels and your enthusiastic high fives, you have truly made this an experience I will never forget.

To my committee, Dr. Lila Gierasch, Dr. Bob Weis and Dr. Peter Chein, you have pushed me to be a better scientist by teaching me to think outside the box and appreciate the meaning of testing a hypothesis. I thank you.

To the Hardy Lab members, my scientific family, both past and present, you will always be held near and dear to my heart. From the Monday morning sports talk to the mid week frustrations followed up by the Friday fireside chats and all of the fun times, you filled my days with laughter and enjoyment. Thank you for being the everyday rock I could always count on.

To my all my friends, you have been a shoulder to lean on and a breath of fresh air when I needed it most. To Mike Wilson Jr. and Shannon Coates Flagg, thank you for always listening, understanding, and making me smile. This road would have been difficult without all of your support so let the good times roll!

And last but certainly not least, to my family, I would not be where I am today or the person I am today without your unconditional love, support, and understanding. Thank you, not only for being my biggest fans but for supporting me through the tough times and keeping me grounded as a person. Mama, thank you for encouraging me to not only dance to the beat of a different drummer but to polka whenever I had the chance. Pappy, thank you for always finding the right words to make me feel at ease and giving me the encouragement to always stand up for what I believe in. Elyse, my partner in crime, thank you for teaching me all of the important life skills I needed to accomplish my goals. You have taught me how to think on my feet, especially when getting framed for taking cookies out of the kitchen, how to accept constructive criticism by convincing me there is always room for improvement, particularly when it comes to my touchdown dance moves, and patience by forgetting to find me when playing hide-n-go-seek. We are truly two peas in a pod, SBN! I love you all!

“You have brains in your head. You have feet in your shoes You can steer yourself any direction you choose. You're on your own. And you know what you know. And YOU are the guy who'll decide where to go.” Oh, the Places You’ll Go! -Dr. Seuss

ABSTRACT

REGULATION OF CASPASE-9 BY NATURAL AND SYNTHETIC INHIBITORS

MAY 2012

KRISTEN L. HUBER, B.S., QUINNIPIAC UNIVERSITY

Ph.D., UNIVERSITY OF MASSACHUSETTS AMHERST

Directed by: Professor Jeanne A. Hardy

Tight regulation of caspase-9, a key initiator of apoptosis, is required to uphold cellular homeostasis. Although it is controlled on a multifactorial level, misregulation of this process does occur, which is a characteristic of a variety of diseases from ischemic injury to cancer. Therefore it remains important to gain a detailed understanding of the mechanisms behind native caspase-9 regulatory pathways and harness these mechanisms for therapeutic purposes.

Based on known mechanisms, such as the unique inhibitory complex of caspase-

9 and XIAP-BIR3, development of synthetic regulators can be envisioned, while other mechanisms such as zinc-mediated inhibition and CARD activation of caspse-9 remain undefined. Intrigued by the multiple ways to control caspase-9’s activity, we sought after designing synthetic caspase-9 inhibitors in addition to defining the mechanistic details metal regulation and CARD domain activation.

We report the first stabilized α-helical peptides that harness the native regulatory mechanism of caspase-9 and the BIR3 domain which lead to the understanding of the importance of exosites in inhibitory complexes. Our studies also revealed that there are two distinct zinc binding sites, one at the active site and another at a novel zinc binding site of yet unknown function in caspase-9 however this site may have the potential to

vi control caspase-6 based on its regulatory mechanism. Furthermore, an interaction was discovered between CARD and the catalytic core of caspase-9 in the presence of a properly formed substrate binding groove, a potential mechanism utilized by the apoptosome for activation of the enzyme.

All in all, the regulation of caspase-9 occurs on a variety of levels that requires almost every surface of the enzyme. Through exploring these underlying molecular details behind the various mechanisms, not only has the field of caspase-9 regulation mechanisms been extended, essential information was gained for further pursuit in an advancement towards the design of caspase-9 activators and inhibitors.

vii

TABLE OF CONTENTS

Page

ACKNOWLEDGMENTS ...... v

ABSTRACT ...... vi

LIST OF TABLES ...... xiii

LIST OF FIGURES ...... xiv

CHAPTER

I. APOPTOSIS, DISEASE AND THE REGULATION OF CASPASE-9 ...... 1

1.1. The Roles of Apoptosis in Disease ...... 1

1.2. Caspases: Facilatators of Apoptosis...... 4

1.3. Caspase Active Site and Catalytic Mechanism ...... 6

1.4. Apoptotic Pathways ...... 9

1.5. Natural Regulation of Caspase-9 ...... 11

1.6. Synthetic Regulation of Caspase-9 ...... 15

1.7. Caspase-9 and Its Role in Disease ...... 19

1.8. Refrences...... 21

II. ROBUST PRODUCTION OF A LIBRARY OF CASPASE-9 INHIBITOR PEPTIDES USING METHODOLOGICAL SYNCHRONIZATION ...... 30

2.1. Introduction ...... 31

2.2. Results ...... 34

2.2.1. Construction of Peptide-Fusion Expression Vector and aPP Variants ...... 34

2.2.2. Expression and Purification of a Recombinant Peptide Library ...... 36

2.3. Discussion ...... 41

2.4. Materials and Methods ...... 45

viii

2.4.1. Cloning of Recombinant Peptide-Fusion Variants ...... 45

2.4.2. Expression and Purification of Recombinant Peptide-Fusion Variants ...... 46

2.4.3. Expression and Purification of Human Rhinovirus 3C Protease .....47

2.4.4. Peptide-Fusion Cleavage by Human Rhinovirus 3C Protease and Separation from Fusion Partners ...... 48

2.4.5. Mass Spectromerty...... 49

2.5. References ...... 49

III. INHIBITION OF CASPASE-9 BY STABILIZED PEPTIDES TARGETING THE DIMERIZATION INTERFACE ...... 54

3.1. Introduction ...... 55

3.2. Results ...... 61

3.2.1. Native and Aib-Stabilized Peptides ...... 61

3.2.2. aPP-Scaffolded Peptides ...... 66

3.2.3. Aliphatic Stapled Peptides ...... 71

3.3. Discussion ...... 73

3.4. Materials and Methods ...... 77

3.4.1. Caspase-9 Expression and Purification ...... 77

3.4.2. Caspase-7 WT and Caspase-7 C186A Expression and Purification ...... 79

3.4.3. Peptide Production ...... 79

3.4.3.1. Synthesis of N-Fmoc-S-2-(2'-pentyl) alanine...... 79

3.4.3.2. Synthesis of N-Fmoc-S-2-(2'-octyl) alanine...... 82

3.4.4 Activity Assays ...... 84

3.4.5. Mass Spectromerty...... 84

3.4.6. Secondary Structure Analysis by Circular Dichroism ...... 85

3.4.7. Computational Structure Prediction ...... 85

3.5. References ...... 86

IV. MECHANISM OF ZINC-MEDIATED INHIBITION OF CASPASE-9 ...... 94

4.1. Introduction ...... 94

4.2. Results ...... 95

4.2.1. Metal Affects on the Properties of Caspase-9 ...... 95

4.2.2. Determining the Location of Zinc Binding Sites ...... 99

4.3. Discussion ...... 102

4.4. Materials and Methods ...... 104

4.4.1. Caspase-9 Expression and Purification ...... 104

4.4.2. Prediction of Metal Lingands and Construction of Ligand Substitution Variants ...... 106

4.4.3. Caspase-7 C186A Expression and Purification ...... 107

4.4.4 Activity Assays ...... 107

4.4.5. Oligomeric-State Determination ...... 109

4.4.6. Secondary Structure Analysis ...... 110

4.4.7. Zinc Binding Analysis by ICP-OES ...... 110

4.4.8. Model of Zinc Binding to Caspase-9 ...... 111

4.5. Structure Determination trials of Caspase-9 and Zinc ...... 112

4.5.1. Crystallization of Caspase-9 in the Presence and Absence of Zinc ...... 112

4.5.2. Data Collection on Crystals of Caspase-9 ...... 114

4.5.3. Zinc Soaks of Caspase-9 Crystals ...... 115

4.6. References ...... 117

V. CASPASE-9 CARD:CORE DOMAIN INTERACTIONS REQUIRE A PROPERLY-FORMED ACTIVE SITE ...... 121

5.1. Introduction ...... 121

5.2. Results ...... 124

5.2.1. The Influence of CARD on the Oligomeric State and Stability of Caspase-9 ...... 124

5.2.2. Determining an Interaction Site Between Caspase-9 Catalytic Core and CARD Domains ...... 130

5.3. Discussion ...... 134

5.4. Materials and Methods ...... 137

5.4.1. Caspase-9 Expression and Purification ...... 137

5.4.2. Oligomeric State Determination ...... 139

5.4.3. CARD Expression and Purification ...... 140

5.4.4 Thermal Stability and Secondary Structure Analysis by Circular Dichroism...... 141

5.4.5. Caspase-3 Expression and Purification ...... 142

5.4.6. Native Gel Analysis and Ni-NTA Pull Down Assay to Determine in trans Interactions ...... 142

5.4.7. Activity Assays ...... 143

5.5. References ...... 145

VI. A SURFACE-WIDE VIEW OF THE NATIVE REGULATORY MECHANISMS OF CASPASE-9 ...... 147

6.1. Regulation of Caspase-9 Occurs at a Variety of Locations on its Surface...... 147

6.2. References ...... 153

APPENDICES

A. SMALL MOLECULE ACTIVATION OF CASPASE-9 ...... 157

B. DESIGN OF AN ACTIVATIABLE INITIATOR CASPASE ...... 170

C. EXPRESSION AND PURIFICATION OF YEAST METACASPASE YCA1 ...... 180

BIBLIOGRAPHY ...... 188

xii

LIST OF TABLES

Table Page

2.1. Peptide Expression, Yield, and Chemical Characteristics ...... 40 3.1. Overall Inhibition of Caspsae-9 Full-Length by Peptides ...... 65 4.1. Zinc Binding Analysis of Caspase-9 Variants ...... 100 5.1. Kinetic Parameters for Full-Length and ΔCARD variants of Caspase-9 ...... 125 5.2. Kinetic Parameters for Full-Length Caspase-9 Charge Swap Variants...... 134

xiii

LIST OF FIGURES

Figure Page

1.1. Caspase Structure ...... 5 1.2. Caspsae Active Site Structure and Mechanism ...... 7 1.3. Intrinsic and Extrinsic Pathways of Apoptosis ...... 9 1.4. Inhibitory Complex of Caspase-9 with XIAP-BIR3 Domain ...... 14 1.5. Synthetic Inhibitors of Caspase-9 Activity ...... 16 2.1. aPP Structure ...... 33 2.2. Schematic Representation of Expression Vector, pET32-Peptide ...... 34 2.3. Structure of Peptide Library...... 35 2.4. Peptide-Fusion Purifications ...... 37 2.5. Peptide-Thioredoxin Fusion Protein Cleavage ...... 38 2.6. HPLC Purification Chromatograms of Cleaved Peptide-Fusion Samples ...... 39 2.7. Mass Spectra of Peptides ...... 39 2.8. Methodological Synchronization ...... 44 3.1. Interactions Required for Inhibition of Caspase-9 are Clustered in the α5 Helix...... 58 3.2. Sequences of the α5 Region of XIAP-BIR3 and Peptides 1-9 ...... 62 3.3. Circular Dichroism Spectra of Native Peptide 5 and the Aib-Stabilized Peptide 8 ...... 63 3.4. Native and Aib-Stabilized Peptides Show Some Inhibition of Caspase-9 Activity ...... 63 3.5. Peptide 2 Non-Specifically Activates Caspase-9 ...... 66 3.6. BIR3 Interactions Grafted onto Stabilized Miniature Protein aPP ...... 67 3.7. aPP and Peptide Structures Predicted Computationally by Rosetta ...... 69 3.8. Properties of aPP Based Peptides ...... 69 3.9. Aliphatic Stapled Peptide Design ...... 71 3.10. Properties of Aliphatically Stapled Peptides...... 72 3.11. Analysis of Aliphatically Stapled Peptides ...... 73 3.12. Full-Length BIR3 is Required for High-Affinity Caspase-9 Inhibition ...... 76 4.1. Zinc Exclusively Inhibits Caspase-9 Activity...... 96 4.2. Kinetics of Full-Length Caspase-9 Wild-Type in the Presence of 0-50 μM ZnCl2 ...... 97 4.3. Zinc Does Not Alter the Biophysical Properties of Caspase-9 ...... 98 4.4. Zinc Binding in Caspase-9 ...... 99 4.5. Kinetics of Full-Length Caspase-9 C272A Variant in the Presence of 0-50 μM ZnCl2 ...... 102 4.6. A Model of Caspase-9 Active-Site Ligand Interactions with a Modeled Zinc Ion ...... 103 4.7. Crystals of Wild-Type Caspase-9 Full-Length Pre and Post Optimization ...... 114 4.8. Diffraction Image of Apo Wild Type Caspase-9 Crystals Soaked in a 20% PEG 400 Cryoprotectant for One Hour ...... 115 4.9. Diffraction Image of Apo Wild Type Caspase-9 Crystals Soaked in ZnCl2 .... 117

xiv

5.1. Model Depicting the Increase in Enzymatic Activity of Caspase-9 ...... 123 5.2. Size Exclusion Chromatography of Caspase-9 Full-Length and ΔCARD in the Presence and Absence of Active Site Ligand z-VAD-FMK ...... 125 5.3. Thermal Denaturation Analysis and Circular Dichroism Spectrum of Monomeric, Cleaved Caspase-9 Full-Length, ΔCARD, and CARD Only ...... 126 5.4. Thermal Denaturation Analysis and Circular Dichroism Spectrum of Dimeric, Cleaved Caspase-9 Full-Length and ΔCARD ...... 127 5.5. Overlay of the Circular Dichroism Spectrum for Full-Length Caspase-9 in the Presence and Absence of Active Site Ligand z-VAD-FMK ...... 128 5.6. Thermal Denaturation Analysis and Circular Dichroism Spectrum of Monomeric, Uncleaved caspase-9 and Monomeric, Cleaved Caspase-9 Full-Length ...... 129 5.7. SDS-PAGE Analysis of Full-Length Uncleaved Caspase-9 in the Presence and Absence of 3% Active Caspase-3 Enzyme ...... 129 5.8. Analysis of CARD:Caspse-9 Core Domain Interactions in trans ...... 130 5.9. Characteristics of the Ser-Gly Linker Extension Caspase-9 Variant ...... 131 5.10. Top RosettaDock Models of the Potential Interaction Site Between CARD and the Catalytic Core of Caspase-9 ...... 133 5.11. Model of Caspase-9 Activation States in the Presence of CARD ...... 136 6.1. Structure of Caspase-9 with Mapped Regulatory Surfaces ...... 151 A.1. Molecular Structure of FlAsH-EDT2 ...... 157 A.2. Models of the Caspase-9 FlAsH Binding Variants ...... 159 A.3. Analysis of FlAsH Binding to Caspase-9 ΔCARD ...... 163 A.4. Analysis of FlAsH Binding to Full-Length Caspase-9 ...... 165 A.5. Western Blot Analysis of FlAsH Binding to Full-Length Caspase-9 ...... 165 B.1. Model of the Designed Caspase-9 Disulfide Cross-Linked Variant ...... 172 B.2. Disulfide Cross-Linking of Wild Type Caspase-9 Full-Length and Disulfide Variant ...... 174 B.3. Chemical Cross-Linking of Wild Type Caspase-9 Full-Length and Disulfide Variant ...... 176 C.1. Schematic Representation of Expression Vector, pYCA1 ...... 182 C.2. Purification of YCA1 ...... 184

CHAPTER I

APOPTOSIS, DISEASE, AND THE REGULATION OF CASPASE-9

Apoptosis, otherwise known as programmed cell, has been established as a key component for a variety of disease. Pivotal roles are played by the facilitators of this process, the caspases, with caspase-9 (ICE-LAP6, Mch6) being of particular interest due to its role in initiating the cascade. Its activation mechanism and regulatory checkpoints are targets for treatment of a variety of apoptosis-related diseases. Thus, an improved understanding of these processes would improve our ability to create new therapies to treat diseases in which apoptosis has gone awry.

1.1. The Roles of Apoptosis in Disease

Cells go through a highly regulated process called apoptosis. Apoptosis is critical for the normal development and stability or homeostasis of all multi-cellular organisms.

Apoptosis plays a role in sculpting and structuring developing tissue and organs throughout the all stages of an organism‟s development. It is responsible for eliminating tissue or appendages that were once required in early development but are no longer necessary for the mature stages of life, for controlling the number of cells during the developmental process to achieve proper organ function, as well as, for eliminating defective or damaged cells from the developing system1,2. Therefore, any disturbance in the apoptotic process is harmful to the system.

Apoptosis has been found to play a role in a variety of diseases. Upregulation of apoptosis has been observed in a variety of cardiovascular diseases such as myocardial infarction, dilated cardiomyopathy and end-stage heart failure3-6. A build-up of apoptotic macrophages and smooth muscle cells have been observed in rupturing atherosclerotic

1 plaques resulting in macrophage build-up, clogging and weakening of the arterial cell wall. Furthermore, increased levels of cardiac-myocyte apoptosis is observed in patients suffering from chronic heart failure, which reduces the ability of the heart to maintain contractile function, providing a strong impetus for further investigating the role of apoptosis in heart disease. In addition, diabetes, a disease characterized by inappropriate blood glucose levels and insulin imbalance, has also been linked to an increase in apoptosis7-10. For both Type 1 and Type 2 diabetic patients, apoptotic beta cells in the islets of the pancreas have been observed. Although not clearly understood, beta-cell apoptosis in Type 1 diabetes is thought to be the main cause of the disease whereas the beta-cell death found in Type 2 diabetes is thought to be triggered by increased toxins in the beta cells themselves, such as glucose, saturated fatty acids, and islet amyloid polypeptides, which together induce oxidative stress and thus apoptosis. Given that 25.8 million children and adults in America have diabetes11, an improved understanding of the involvement of apoptosis is strongly warranted.

Further evidence of the harmful effects of a apoptotic imbalance can be observed in chronic airway inflammatory diseases such as bronchial asthma. This condition affects

34 million Americans12 and is caused by damage to the epithelium lining of the airways also caused by an increase in apoptotic cells. Although apoptosis is a normal process for clearing damaged cells from the epithelium layer of the nasal passage, trachea, and bronchia, an excessive amount of apoptosis is undesirable and thought to highly contribute to the pathogenesis of this class of disease13,14. Hepatitis C, a viral disease of the liver has been shown to cause an increase in apoptosis of infected cells, a process thought to provide a mechanism for viral shedding15 in addition to the host‟s own

2 mechanism of eliminating the virus from liver cells16. It has also been shown that during a systematic inflammatory response caused by a bacterial infection, otherwise known as sepsis, the body tries to regain homeostasis via initiating a compensatory anti- inflammatory response. Although this process is not highly understood, one area of focus has been lymphocyte apoptosis, characterized by reduced levels of CD4, B-, T- and dendritic cells17-19. Treatment of these cells with apoptotic inhibitors has shown increased survival rates in sepsis models17,20. Furthermore, neuronal cell death has been shown as a characteristic of neurodegenerative diseases such as amyotrophic lateral sclerosis (ALS),

Huntington's disease, Alzheimer's disease, and Parkinson's disease. Upregulation of the apoptotic machinery is evident in early and late stages of disease progression, characterized by increased activation of pro-apoptotic proteins. Treatment with apoptotic inhibitors, however, provides protection from additional cell death in mouse models, further highlighting apoptosis as an important area of study.

Down regulation of apoptosis is also prevalent in a variety of diseases, such as rheumatoid arthritis and cancer. Rheumatoid arthritis is a chronic inflammatory disorder affecting 2.1 million adults and approximately 1 in every 250 children under the age of

18 in America. Rheumatoid arthritis is caused by an imbalance between proliferating and apoptotic cells resulting in synovial hyperplasia and angiogenesis21. A defective apoptotic pathway by means of increase apoptotic inhibitors in synovial tissue has been discovered and prevention of apoptotic cell death is thought to restore the synovial membrane tissue22. In a similar respect, the apoptotic machinery is found to be absent or in low cellular levels in breast, gastric, colorectal and lung cancers, which also show an increased level of apoptotic inhibitors23-25. Not only is there a change in oncogenic and

3 pro-apoptotic-protein expression levels in cancer cells, mutations in other pro-apoptotic proteins26-31 have also been observed.

Because it is an ongoing area of research interest, irregularities within apoptosis are continually being discovered in a wide array of diseases. It is thought that irregularities in apoptosis may account for up to 50% of all diseases in which there are no suitable therapies32. Therefore, a further understanding of the apoptotic process and its regulators is still necessary in order to understand how diseases are able to evade this important cellular process.

1.2. Caspases: Facilitators of Apoptosis

Facilitators of this lethal cellular mechanism of apoptotic cell death are the cysteine aspartate proteases, or caspases. Caspases are classified into two subgroups based on their function in the cascade, structural characteristics, and substrate specificity33. The apoptotic caspases are commonly classified as in the initiator or executioner subgroups. Initiator caspases, caspase-2, -8, -9, and -10 emerge in the initial stages of the apoptotoic cascade and are known for their ability to be involved in large oligomeric activation assemblies. The executioner caspases-3, -6, and -7 are located in the latter half of the cascade and are responsible for the majority of proteolytic events resulting in the orderly demise of the cell. Caspases-1, -4, -5, -11, -12 and -14 are involved in the inflammatory response or other cell processes, but play no known role in apoptosis.

Caspases are synthesized as three-domain polypeptide chains (Fig. 1.1 A). The N- terminal domain, called the prodomain, is unique to each caspase, ranging from 16 to 220

Figure 1.1 Caspase Structure. (A) Caspase polypeptide chain consists of a pro or CARD domain (yellow), large subunit (dark purple), and small subunit (light purple). (B) Caspase-9 CARD domain (PDB ID: 3YGS). (C) Caspase-9 catalytic core (PDB ID: 1JXQ). Individual monomers represented in purple and green about the two fold axis () with position of catalytic cysteine (cyan) and histidine (orange) indicated. (D-E) Caspase-9 active site loop bundle. The active loop bundle conformation is disordered in the (D) absence of substrate and becomes ordered (E) upon binding substrate (gray). Images generated in the PyMOL Molecular Graphics System, Version 1.3, Schrödinger, LLC. amino acids in length. Prodomains shorter in length, found on the executioner caspases, are yet to have a specific function identified, although they have been hypothesized to

be involved in subcellular targeting34,35 or chaperoning folding of the core domain36. The longer prodomains observed in the initiator caspases act as recruitment or regulatory domains. Examples include the Caspase Activation and Recruitment Domain (CARD) found in caspase-2 and -9 and the Death Effector Domain (DED) of caspase-8 and -10.

Although CARD and DED domains are highly dissimilar, both fold into an anti-parallel six-α-helix Greek key structure and facilitate the formation of large protein complexes

(Fig, 1.1 B). The two domains C-terminal to this prodomain region are the large and small subunits connected via the intersubunit linker. Together, these domains fold into a heterodimer structure, common to all caspases. Homodimers of the heterodimer large and small subunits are formed about a two-fold axis which in turn makes up the catalytic core of the enzyme. The large subunit, averaging 17-20 kDa in size, houses the catalytic Cys-

His dyad. The small subunit, averaging 10-12 kDa, comprises the main portion of the homodimer interface. This catalytic core region of the caspases folds into a canonical 12- strand β-sheet core packed between two α-helical layers (Fig 1.1 C). Specific loops which connect portions of the catalytic core secondary structure known as L1, L2 and L2„(the product of cleavage of the intersubunit linker), L3 and L4 also form the catalytic loop bundle. These loops are known to alter their structural position based upon the activation state of the protease. In the apo unliganded, state, these loops are disordered (Fig. 1.1 D) however upon binding of substrate, the L2 loop from one of the monomers interacts with the L2` loop from the opposing monomer (Fig. 1.1 E). Furthermore, L1, L3, and L4 loops are in an ordered state around the substrate binding groove to complete the formation of the catalytic loop bundle in the active state.

1.3. Caspase Active Site and Catalytic Mechanism

Caspases have evolved to be highly specific enzymes with the moderate catalytic

-1 37 properties: Km in the range of 4 to 408 μM and a kcat of 0.2 to 9.1 s . Caspase-3 is the fastest in its class while caspase-9 possesses the slowest catalytic rates overall. The differences in catalytic properties most likely stem from their active site architecture and

6 target specificity. The active site structure of a caspase consists of a substrate binding groove, the catalytic Cys-His dyad, and four mobile loops. The substrate-binding groove

(Fig 1.2 A) provides the specificity and binding determinants for a particular substrate sequence via four interaction interfaces or specificity subsites (S1 – S4). Closest to the catalytic cysteine lies the S1 pocket. This region provides the highest specificity for substrate recognition, essentially always accommodating an aspartate residue, although caspase-9 occasionally recognizes glutamate residues. The S2 substrate-binding site is

38-40 thought to be involved in substrate differentiation . S3 provides an anchoring function for substrate while S4 provides specificity between the subclasses of caspases.

Figure 1.2 Caspase Active Site Structure and Mechanism. (A) Model of the caspase substrate binding groove with peptide z-EVD bound (gray). Catalytic Cys (blue) and His (orange) are in close proximity

to the Asp recognition element found in the S1 pocket (white). S2 (yellow), S3 (tan), and S4 (green), accommodates the remainder of the peptide. (B) Catalytic mechanism of caspases adapted from Fuentes-Prior and Salvesen43.

41 Executioner caspases prefer to bind acidic amino acids in their S4 pocket while initiator caspase-9 prefers bulkier hydrophobic residues (reviewed in42).

The cysteine and histidine catalytic residues responsible for cleavage of the substrate peptide bond are located on the C-terminal end of the β4 strand and the N- terminal end of the βI strand respectively. Caspases are thought to have similar catalytic

43 mechanisms (Fig. 1.2 B) as that of other cysteine proteases (reviewed in ) where the P1 recognition element is anchored through hydrogen bonding to the oxyanion hole, thus polarizing the carbonyl carbon of the peptide bond. Deprotonation of the catalytic cysteine thiol via the catalytic acid/base, a histidine residue, occurs as the initial step of the cleavage mechanism. The anionic sulfur of the now deprotonated catalytic cysteine then performs a nucleophilic attack on the substrates carbonyl carbon, C-terminal to the

P1 aspartate recognition element in the case of the caspases. The catalytic histidine then protonates the α-amino group of the peptide leaving group. After release of this C- terminal peptide of the cleaved substrate, the deprotonated histidine abstracts a proton from water to restore its native state. Activated water then hydrolizes the thioester bond that links the substrates carboxy-terminus to the cysteine thiol and thus restores the enzyme to its native, unliganded state.

Alternative mechanisms to the classical cysteine protease reaction mechanism have also been proposed. For example, the catalytic histidine Nγ is proposed to stabilize the developing charge on the leaving group while a water is utilized as the proton donor for the peptide leaving group instead of a protonated histidine residue44. Additionally, based upon known crystal structures of the caspase family of proteases, the distance of the caspase catalytic residues are observed to be approximately 5Å apart which would be

8 too far for hydrogen abstraction of the cysteine thiol by the catalytic acid/base, making the common cysteine protease reaction mechanism difficult to achieve. Therefore, an alternative mechanism has been proposed by having the active site cysteine nucleophile develop along the reaction coordinate instead of being polarized by the catalytic histidine45. This particular part of the mechanism would be governed by the pH optimum requirement for these enzymes where the executioner caspases lie more toward the neutral to slightly basic pH range of 7.0-8.0 while initiator caspases-8 and -9 have a lower pH optimum of 6.5-7.037. Further experimental data is required however, in order to discern which reaction mechanism is proper for caspase catalytic function.

1.4. Apoptotic Pathways

Although apoptosis‟ final phenotype is cell death, the cascade can be initiated via different mechanisms which would activate either the extrinsic and intrinsic pathways

(Fig 1.3). The decision of which pathway to initiate depends on the type of cellular stress signal received.

Figure 1.3 Intrinsic and extrinsic pathways of apoptosis. Caspases and other important proteins involved in apoptosis are shown. Figure by Kristin Paczkowski.

Activation of the extrinsic pathway through triggers such as cytokine release induces signaling of the pro-apoptotic receptors associated with the TNF or TRAIL family of proteins that reside on the cell surface. These membrane bound receptors bind ligands, such as FasL, TNF-α, or TRAIL, triggering the recruitment and clustering of adapter molecules, such as Fas-associated death domain (FADD), to the cytoplasmic death domain region of the receptor. Additional molecules, caspase-8 or -10 are also recruited ultimately forming the death inducing signaling complex (DISC)46-49. Once bound, the caspases are thought to undergo autoproteolysis and thus activation. As activated caspases, the initiators -8 and -10 can continue the cascade by cleaving their downstream targets, caspases-3, -6, and -7.

If a stress signal, such as DNA damage or build up of reactive oxygen species, occurs from within the boundaries of the cellular membrane, the intrinsic pathway is initiated. This type of stress induces recruitment of the pro-apoptotic Bcl-2 proteins to invade the outer mitochondrial membrane. This event, which controls mitochondrial integrity, induces translocation of other stress dependent proteins known as Bax50, Bad51, or Bid52 to the mitochondrial membrane, resulting in pore formation. As a result, a release of cytochrome c from the mitochondria into the cytosol occurs. Within the cytosol an inactive form of apoptotic protease activating factor-1 (Apaf-1) resides. Upon binding of cytochrome c, Apaf-1 undergoes a conformational change which exposes the Apaf-1

CARD53,54 resulting in the oligomerization of the molecule in an ATP/dATP dependent manner55-57. The now heptameric Apaf-1 complex with exposed CARD domains is able to recruit the zymogen form of caspase-9 through an interaction between the caspase recruiting domains (CARD) on both Apaf-1 and the caspase-9 full length protein. This

10 allows for multiple procaspase-9 monomers to be within close distance for activation and for intermolecular self-processing. This large oligomeric complex formed of cytochrome c, Apaf-1 and pro-caspase-9 is known as the apoptosome. Cryo electron microscopy of the holo form of the apoptosome58,59 indicate the seven Apaf-1 monomers oligomerize into a wheel-shape with each monomer representing one spoke. There is a central hub which houses Apaf-1 CARD which associates with caspase-9. Once bound to the apoptosome, procaspase-9 can become activated and cleave the executioner caspases-3, -

6, and -7 while still associated to the apoptosome.

1.5. Natural Regulation of Caspase-9

Regulating the caspases is a crucial step in controlling unwanted cell death. For this reason, a variety of regulatory check points have evolved to make sure caspase activation does not go awry. These precautions throughout the apoptotic pathway ensure that caspase activation occurs only upon the appropriate cellular signals. In particular, multiple levels of regulation exist for the initiator caspase of the intrinsic pathway, caspase-9. Caspase-9 is a 416 amino acid protein consisting of a 15 kDa CARD domain which facilitates its binding to the apoptosome, an 18 kDa large subunit which houses the catalytic Cys287 and His 237 residues and a 10 kDa small subunit. Caspase-9 not only begins the apoptotic cleavage cascade of the intrinsic pathway, it is a focal point for regulation.

In a natural non-apoptotic state, caspase-9 exists as an inactive, monomeric zymogen. In order to become active, the CARD domain of caspase-9 becomes involved in a protein-protein interaction with the Apaf-1 CARD resulting in a profound increase in caspase-9 activity55,57,60. This is further supported by the increase in activity gained by a

11 stepwise addition of the domains involved in this interaction to the caspase-9 catalytic core, the simplest active unit which possesses the least amount of enzymatic activity.

Addition of caspase-9 CARD increases enzymatic activity by 20 % which is five-fold further enhanced by the addition of Apaf-1 CARD61. Maximal activity is only achieved upon association with the entire apoptosome. Furthermore, caspases are known to be

62,63 active as dimers. The dimerization constant, KD, for caspase-9 is in the μM range , enforcing a fairly weak complex considering cellular concentrations of ~20 nM54. In

64 comparison, KD of the executioner caspases is <50 nM . Initial models of the apoptosome predict a monomeric caspase-9 which dimerizes via increasing the local concentration of monomeric caspase-9 resulting in recruitment of the dimeric partner65 or dimerization amongst the apoptosome bound monomers66 or induced conformations around the active site region of the enzyme67. However, evidence is emerging from high- resolution cryo electron microscopy that caspase-9 monomers are active bound to the apoptosome58.

In general, for caspases to reach their full activity, they need to be proteolytically processed within their intersubunit linker, which connects the large and small subunits of the enzyme. Full activation upon this cleavage serves as a secondary form of regulation.

In the case of caspase-9 this occurs either through autoprocessing or aid from an additional caspase or other protease such a granzyme B. As a cleaved dimer, the L2 loop from one half of the dimer is able to interact with the L2` loop from the other half in the presence of substrate, thus positioning the active site loop bundle62 for catalysis. Caspase-

9, which becomes cleaved at Asp315 has a ten-fold increase in activity upon cleavage in vitro45. This value rises to 2000-fold54 when caspase-9 is associated as part of the

12 apoptosome. However, it should be noted that caspase-9 does not require cleavage of its intersubunit linker to have some minimal activity54. Furthermore, proteolytic removal of the CARD domain from the large subunit, which results in decreased activity, has also been observed in addition to processed caspase-9 being displaced from the apoptosome by additional molecules of procaspase-968. Without these early checks and balances of caspase-9 activation in place, caspase-9 would prematurely activate the caspase cleavage cascade resulting in unwanted cell death.

As a failsafe, protein-based inhibitors also exist in the cell to control caspase activity. The most potent inhibitors69 of both the intrinsic and extrinsic pathways of apoptosis70 are known as the inhibitors of apoptosis or IAP‟s. These multidomain inhibitors consist of baculoviral IAP zinc-binding repeat (BIR) domains which are approximately 70 to 80 residues in length and contain a characteristic zinc-binding motif of -CX2CX16HX6C-. A single IAP consists of one to three BIR domains. Typically a carboxy-terminal RING domain is also associated, which allows targeting for ubiquitination and thus proteosomal degredation. Mammalian IAPs including X- chromosome-linked inhibitors of apoptosis (XIAP), cIAP1, cIAP2, Livin/ML-IAP, and neuronal apoptosis inhibitor protein also share this same baculoviral zinc motif.

XIAPs have been shown to inhibit both the initiator and executioner caspases in human cells. They consist of three tandem BIR domains named BIR1, BIR2 and BIR3 followed by a ring domain consisting of one zinc ion chelated to three cysteines and one histidine and an additional zinc ion bonded to four cysteines. Although similar in structure and sequence with approximately 30% sequence identity, each BIR domain possesses its own individual functions. Caspases-3 and -7 can be regulated by blocking of

13 the active site through binding of a small linker region N-terminal to the BIR2 domains of

IAPs (residues 164–235). Binding this linker prevents substrate from entering the active site71-73. In addition, a BIR2 domain interaction with the caspase interaction binding motif

74 73,75 distal from the active site of the enzyme with a Ki of 0.2-5 nM is present . In the case of caspase-9, the BIR3 domain, residues 251-350, causes inhibition of the enzyme73,76

76-78 with a Ki of 10-20 nM (Fig 1.4). BIR3 binds to the dimerization interface of the

Figure 1.4 Inhibitory complex of caspase-9 monomer (purple) with XIAP-BIR3 domain (blue). A structural zinc molecule in BIR3 is shown in red. inactive caspase-9 monomer and sequesters the N-terminus of the small subunit, thus controlling its activity on multiple levels by preventing dimerization and formation of the active site loop bundle79. Interestingly, not only has the cell adapted ways to control unwanted apoptosis, viruses have adapted means of survival by mimicking these natural inhibitory mechanisms. For example, viral IAPs are found in insects80,81 and animal viruses such as African swine fever82.

In addition to enzymatic cleavage, binding platforms and protein based inhibitors, binding of metal ions and post-translational modifications, such as phosphorylation, can alter the function of the caspase class of enzymes. Metal ions commonly used for both biological structure and cellular processes, such as zinc and copper, have been observed as specific regulators of apoptosis by targeting the caspases83. HeLa cells depleted of zinc for nine hours resulted in the processing of caspase-984 while the presence of zinc protected caspase-9 activation from manganese mediated apoptosis85. Further correlations linking zinc and caspases have been observed via the cellular localization of zinc and caspases under apoptotic and zinc deficient conditions13, however, the cellular processes which trigger metal based regulation in addition to how metal ions regulate caspase activity still remain unclear. Kinases, on the other hand, such as Akt, ERK MAPK‟s, and c-Abl regulate caspase-9 activity, both through activation and inhibition of the enzyme.

This occurs based on the position of the phosphorylation. For example, phosphorylation at position T153 increases the activity of the enzyme86 whereas T125 phosphorylation prevents processing of caspase-9 and subsequent executioner caspase activation87. These alternate modes of caspase regulation further enhance the idea of a multi-factorial requirement for controlling caspase activity and unwanted cell death.

1.6. Synthetic Regulation of Caspase-9

Discovery of the caspases and their crucial role in apoptosis opened a new area of therapeutic interest and thus the discovery and design of small synthetic inhibitors. Initial inhibitor designs were inspired by the substrate specificity elements of the enzymes themselves. Designed amino acid analogues (Fig 1.5 A) utilized the caspase substrate recognition element aspartate to target all caspases, including caspase-9. Additional

Figure 1.5 Synthetic Inhibitors of Caspase-9 Activity. (A) Protected amino acid monomers to tetrapeptide substrate mimics are tuned for caspase-9 active site specificity. The recognition element Asp is utilized for capped amino acids while LEHD is utilized for tetrapeptide sequences which can be functionalized on the N- and C- terminus. (B) N-terminal protective groups and C-terminal electrophilic warheads provide variation and increase potency of the peptide-based inhibitors. (C-E) Small molecule and α-helical peptide inhibitors are designed to disrupt caspase-9 activation via binding to Apaf-1 and preventing the CARD-CARD interactions. Images generated in ChemBioDraw Ultra 12.0, CambridgeSoft Corporation and the PyMOL Molecular Graphics System, Version 1.3, Schrödinger, LLC. analogues branched out into di-, tri-, and tetrapeptides that incorporated the sequence requirements for the remainder of the substrate binding groove. Sequences such as Val-

Asp and Val-Ala-Asp targeted a broad spectrum of caspases and are classified as pan- caspase inhibitors88. As sequence specificities for individual caspases were determined, specific peptide inhibitors were designed such as Leu-Glu-His-Asp (LEHD) in case of caspase-933 (Fig 1.5 A).

Caspase targeting and potency of peptide-based inhibitors can be altered through addition of protective groups, fluorophores, and reactive electrophilic warheads (Fig. 1.5

B). Functionalization on the N-terminus of the peptide sequence has included protective groups such as the N-acetyl (ac), benzyloxycarbonyl (Z), or quinolyl (Q) moieties.

Fluorophores such as FITC have also been included on the amino terminal end of this class of peptides to enable detection. The C-terminal portion of the peptide inhibitors typically comprise electrophilic warheads that react with the catalytic nucleophile.

Aldehydes, nitriles and ketones are used for reversibility, while halomethylketones (e.g.

FMK, fluoromethyketone; CMK, chloromethylketone), diazomethylketones and acylomethylketones serve as irreversible warheads33,89,90. The C-terminal O-phenoxy group was also incorporated into the warhead class of substrate based inhibitors due to its improved leaving properties and reactivity compared to fluoromethylketones. Peptide inhibitor Q-VD-Oph was determined to have an IC50 of 25-430 nM against a broad spectrum of caspases, with caspase-9 being on the highest end of the spectrum91. Potency and specificity vary greatly amongst these inhibitors with dissociation constants ranging anywhere from pM to μM. Caspase-9-specific, warhead driven inhibitors such as LEHD-

FMK, are potent against enzymatic activity. Compound IDN-6556 ((3 2-[(2-tert-butyl- phenylaminooxalyl)-amino]-propionylamino 4-oxo-5-(2,3,5,6-tetrafluoro-phenoxy)- pentanoic acid)) (Fig 1.5 C), a broad-spectrum irreversible inhibitor of the caspases with

92 an IC50 of 25 nM , encompasses a similar premise with incorporation of a peptide backbone and the aspartate recognition element however, IDN-6566 utilizes an N- terminal oxamide and C-terminal tetrafluorophenoxy warhead. This class of substrate mimicking inhibitors have been studied in the treatment of acute liver failure93,

17 myocardial ischemia-reperfusion injury94, and traumatic brain injury95. In this thesis, we have explored an entirely new class of inhibitors (Chapter III).

Caspase-9‟s mechanism for activation via binding to the apoptosome complex has also been targeted to control its enzymatic activity. Small molecule and α-helical peptide inhibitors have been designed to either disrupt oligomerization of Apaf-1, which creates the bulk of the apoptosome platform, or caspase-9‟s association with the apoptosome. N- alkylglycine peptoid inhibitors (Fig 1.5 D) and their solubility-enhancing analogues96 bind to Apaf-1 (KD = 57 ± 12 nM) and prevent recruitment of caspase-9 to the apoptosome. Fusion of cell penetrating peptides and a polymeric carrier to the N- alkylglycine peptoid inhibitors have also been explored to enhance membrane permeability and efficacy respectively for their ability to decrease apoptosis in neonatal rat cardiomyoctyes under hypoxic conditions. Diarylurea based compounds also inhibit formation of the apoptosome97 however the mechanism is slightly less clear. This class of inhibitor is determined to either prevent recruitment cytochrome c, dATP, or caspase-9 to

Apaf-1. α-helical polypeptide modulators on the other hand, specifically target the

CARD-CARD mediated interaction of caspase-9 and Apaf-198. Synthetic peptides were derived from the acidic patch region of helices 2 and 3 of Apaf-1 CARD domain (Fig 1.5

D) and the basic patch of helices 1 and 3 of the caspase-9 CARD. This class of inhibitors successfully disrupted the interaction of Apaf-1 and caspase-9, thus preventing caspase-9 activation both in vitro and in cell extracts with an IC50 of 68-102 μM for apoptosome activity as judged by cleavage of caspase-3 and 63-113 μM in human embryonic kidney

293 (HEK 293) cellular extracts.

1.7. Caspase-9 and its Role in Disease

Caspase-9 is of high interest due to its entry point into the apoptotic pathway and its involvement in disease. As an initiator caspase in the intrinsic pathway of apoptosis, targeting caspase-9 provides flexibility between the upstream apoptotic signals and the cleavage cascade, making this a critical target point for cell survival or cell death. This role is reflected by its specific involvement in a variety of diseases.

Down regulation of caspase-9 has been observed in cancers as well as viral infections. In colonic carcinoma cells, expression levels are found to be low or even absent when compared to normal cells99 suggesting caspase-9 as the critical apoptotic checkpoint within this cell line. In the case of testicular cancer, caspase-9 fails to activate100. Through the targeting caspase-9 in this manner, the testicular cancer cells are also able to resist treatment with cisplatin, a treatment in which cell death is a priority, further confirming the importance of this cell death protease in cancer. The viral infection caused by the vaccinia virus, has also adopted means of controlling caspase-9. Viral protein F1L has been found not only to mimic apoptosis-like proteins upstream of the caspase cascade101,102 but to directly affect caspase-9 function103. F1L inhibits caspase-9 activity by preventing zymogen caspase-9‟s association with the apoptosome in addition to blocking the enzymes protease function, driving home the significance of caspase-9 in controlling apoptosis.

Up regulation of caspase-9 has also been observed in a variety of neurological diseases and immune disorders. Amyotrophic lateral sclerosis (ALS) exhibits increased caspase-9 activity in the spinal motor neurons of ALS mouse models104 and has been found to be a key player in the progression of the disease. Increased caspase-9 expression

19 levels and activity are also observed in the severe neuropathological cases of

Huntington‟s disease patients and in a Huntington mouse model, implicating caspase-9‟s involvement in neuronal death at the end stage of the disease105. The same trend is also observed in the endothelial cells of patients with a rare multisystemic immune disorder known as Behçet's disease106, further confirming caspase-9‟s pivotal role in the progression of a disease.

In addition to direct action of caspase-9, diseases also co-opt the regulators of caspase-9 in order to ensure activation of the apoptotic pathway. A characteristic of renal cell carcinoma107,108, lung cancer109, testicular germ cell tumors110 and hepatocellular cancer111 is the release of caspase inhibitor antagonist, Smac/Diablo, which is a direct antagonists of BIR3 resulting in the release its inhibition over caspase-9. This allows caspase-9 to activate and initiate the caspase cascade. In a similar fashion, while experiencing ischemia-reperfusion injury of the heart, the BIR3 antagonist, Omi/HtrA2, is released thus causing activation of caspase-9 as well112.

Given these unique roles of caspase-9 in a wide variety of diseases, fully understanding the process of apoptosis is vital to understanding the normal homeostasis of a cell and to controlling a variety of diseases in which the cellular death process has been altered. By focusing on the caspases, the ultimate “hit men” of the cell, control can be achieved, specifically when targeting the initiator caspase-9. By controlling caspase-9 activity, the transmission of the cell death signal can be controlled as well. For this reason, the focus of this thesis is to harness and further understand the natural regulatory events inhibition of caspase-9 by BIR3, CARD domain activation and metal inhibition via biochemical and biophysical means.

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CHAPTER II

ROBUST PRODUCTION OF A LIBRARY OF CASPASE-9-INHIBITOR PEPTIDES USING METHODOLOGICAL SYNCHRONIZATION

This chapter was published as: Huber, K. L., Olsen, K. D.* and Hardy, J. A., 2009. “Robust Production of a Peptide Library using Methodological Synchronization.” Protein Expression and Purification, 67,139-147. KLH designed initial parent construct and mutational sites, in addition to performing metal ion affinity and reverse HPLC purifications, ESI-MS, and peptide expression analysis. KDO performed site directed mutagenesis and affinity purification steps.

Abstract

Peptide libraries have proven to be useful in applications such as substrate profiling, drug candidate screening and identifying protein-protein interaction partners. However, issues of fidelity, peptide length and purity have been encountered when peptide libraries are chemically synthesized. Biochemically produced libraries, on the other hand, circumvent many of these issues due to the fidelity of the protein synthesis machinery. Using thioredoxin as an expression partner, a stably folded peptide scaffold (avian pancreatic polypeptide) and a compatible cleavage site for human rhinovirus 3C protease, we report a method that allows robust expression of a genetically-encoded XIAP-Bir3 inspired peptide library, which yields peptides of high purity. In addition, we report the use of methodological synchronization, an experimental design created for the production of a library, from initial cloning to peptide characterization, within a five-week period of time.

Total peptide yields ranged from 0.8 to 16%, which corresponds to 2-70 milligrams of pure peptide per one liter of cell culture. Additionally, no correlation was observed between the ability to be expressed or overall yield of peptide-fusions and the intrinsic chemical characteristics of the peptides, indicating that this system can be used for a wide variety of peptide sequences with a range of chemical characteristics.

2.1. Introduction

Peptide libraries are commonly used in a variety of endeavors including identifying peptide-DNA interactions 1, as surrogates for screening protein-protein interactions2-5, and as a basis for finding potential peptidic drug molecules. Peptide libraries have also been used for profiling substrate specificities of proteases6-11, phosphatases12-14, kinases15-

18 and other drug targets19-22. Thus, the production of high quality peptide libraries is of wide interest for many applications.

Typically, peptides for libraries have been produced via solid-phase synthesis, which involves sequential coupling of amine-protected amino acids to resin-bound amino acids.

The resulting libraries generally contain peptides no longer than 15-20 amino acids, which can prove limiting in applications requiring longer peptides. The length constraints for chemically synthesized peptides are the result of coupling and deprotection efficiencies at each step, such that an exponential decrease in sequence fidelity is observed as a function of length. In addition, enantiomerization of nineteen of the twenty naturally-occurring amino acids and the associated difficulties in purification of the D- isomer-containing peptides from the L-isomers set practical limits to peptide lengths in library synthesis23,24. Sometimes peptide sequences are restricted due to steric clash of adjacent amino acids with bulky chemical catalysts or intermolecular aggregation which results in a low efficiency of the chemical coupling reaction25-27. Overall, chemically synthesized peptide libraries of longer lengths tend to have decreased sample quality, be costly and have increased production time.

Genetically-encoded peptide libraries offer several advantages in library design. The

resulting libraries can contain longer peptide lengths and significantly increased yields while avoiding the more common limitations of chemical synthesis. Biological synthesis of peptides of longer lengths can be particularly important for production of isotopically- labeled peptides for use in heteronuclear Nuclear Magnetic Resonance (NMR) spectroscopy. The principle advantages of biological production result from the fidelity of protein machinery, particularly because the ribosome is not limited in sequence or length of synthesized peptides and has an error rate of only 0.01% error per amino acid added28. Genetically-encoded peptide libraries have not been widely used due to difficulties in the expression and purification of small peptides. This problem has been overcome by the use of protein-expression fusion partners such as glutathione-s- transferase29, maltose binding protein30 and thioredoxin31 being the three most widely- used fusions. While fusion proteins often promote production, this method can be time consuming and requires post expression protease processing and purification of the peptide from the protease. However, the addition of fusion partners has resulted in the improvement of the overall yields by enhancing solubility of the peptide of choice.

We report a method that allows robust expression of a genetically-encoded XIAP-

Bir3 inspired peptide library that addresses the issues of purity, yield, length of the purified peptide, batch-to-batch variability which we have observed with chemically synthesized peptides, in addition to cost and time of production. This genetically-encoded system features thioredoxin as a fusion tag for the stably-folding peptide scaffold avian pancreatic polypeptide (aPP). aPP is a 36 amino acid peptide that contains a hydrophobic core and hydrogen bond network between the α-helix and polyproline helix of the

peptide32,33 (Fig. 2.1 A and B). These properties are responsible for its fold and stability. Previous work has demonstrated that the presence of the hydrophobic core and hydrogen bond network render aPP insensitive to mutations over much of the amino acid Figure 2.1 aPP structure. (A) aPP backbone (lightest gray) has a hydrophobic core created by three 34,35 prolines located on the poly-proline helix (medium sequence or to a c-terminal gray) and various residues on the α-helix (dark gray). (B) aPP’s internal hydrogen bond network, truncation33,36. highlighted in dashed lines, is responsible for the impressive structural stabilization of a peptide that is A common difficulty with peptide just 36 amino acids long. expression is residual amino acid overhangs following the protease cleavage event. The native sequence of aPP allows inclusion of an N-terminal cleavage site for human rhinovirus 3C protease, a highly specific protease, without any unwanted amino acid additions to the peptides. Human rhinovirus 3C protease cleaves the sequence LEVLFQ-

GP, generating a gly-pro overhang upon cleavage. In aPP the first two amino acids are gly-pro, so no residual amino acids are left upon cleavage. Combining this optimized expression system with methodological synchronization of site-directed mutagenesis, protein expression and purification, a twenty-member aPP variant library of peptides 27 amino acids in length was generated. This method allowed production of the desired library with overall improved yields, purity, and cost, all on a time frame that is comparable to synthetic peptide library methods on the same production scale.

2.2. Results

2.2.1. Construction of Peptide-Fusion Expression Vector and aPP Variants

A peptide-expression system for the library of aPP-based peptides was generated that expressed thioredoxin fusion proteins. A goal of this project was to streamline all steps in the process of cloning, expression and purification. In order to limit subsequent subcloning events, the parent vector (Fig. 2.2) was created through a single PCR reaction that resulted in the gene for a Figure 2.2 Schematic representation of expression vector, human rhinovirus 3C protease pET32-Peptide. The T7 promoter, a fusion partner thioredoxin (TrxA), 6xHis tag, human rhinovirus 3C cleavage site (LEVLFQ) and cleavable linker, gene for peptide, and ampicillin resistance are shown. the truncated aPP sequence

(GPSQPTYPGDDAPVE-DLIRFYNDLQQY). This gene product was then ligated into the thioredoxin fusion gene and 6xHis purification tag containing vector, pET32b via restriction sites NcoI and XhoI. The resultant thioredoxin-peptide fusion construct then served as the first base sequence for further mutagenesis.

aPP variant sequences were created by mutating codons for six to twelve of the amino acids in the truncated 27 amino acid peptide scaffold (Fig. 2.3 A) via a

QuikChange (Stratagene) mutagenesis strategy. This limited library was designed as aPP mimics of the natural caspase inhibitor, XIAP-BIR3. Mutations included the desired variations in the original peptide sequences and took into consideration amino acids that 34

Figure 2.3 Structure of peptide library. (A) Amino acid sequences of the twenty-member peptide library. Mutated residues are bold. (B) Cartoon representation of truncated aPP peptide scaffold with sites of mutations shown in stick representation and highly interrogated site D11 represented in spheres. are known to be structurally important for scaffold stability. Sites of mutagenesis are throughout the truncated aPP sequence and are spread across the aPP structural elements

(Fig. 2.3 B). Mutational sites include Q4, T6, D10 and D11 on the polyproline helix and

D16, D17, I18, Y21, D23, L24 Q25, and Y26 on the α-helix. In addition D11 was selected as a site to be extensively interrogated. By focusing our investigation in one region, we were able to produce many variants in a limited number of mutagenic steps.

Moreover, all mutational oligonucleotides were designed to limit cost and allow rapid production of the new peptide encoding DNA constructs. These designs focused on producing an array of peptide sequences using the minimum number of rounds of

mutagenesis. If mutagenesis was performed not by sequential means, 123 rounds of mutagenesis would have been performed to complete the library. Mutations were quickly assessed by growing the transformed E. coli cells for eight hours and plasmid prepping the DNA in time for same day sequencing by an outsourced company (Genewiz Inc).

Upon sequence analysis, one out of two or more clones tested obtained the desired mutations. The success rate was directly related to the thoroughness of the DpnI digestion step during the QuikChange protocol (data not shown).

2.2.2. Expression and Purification of a Recombinant Peptide Library

Once the desired genetic mutations were verified, the vectors were transformed into the BL21(DE3) strain of E. coli, expressed and harvested. The resulting cell pellets were lysed and prepared for purification. Various avenues were tested to optimize the purification of the peptide-fusion from other bacterial proteins. This was done in order to minimize the potential additive loss of protein during each purification step. Therefore, a nickel affinity step gradient, alone or in combination with ion exchange methods as well as nickel affinity linear gradients were explored to obtain a peptide-fusion sample of adequate purity without sacrificing yield. For example, peptide B3NK was purified via a

Ni-affinity step gradient and ion exchange methods while peptide B3NR was purified by a Ni-affinity linear gradient only (Fig. 2.4). It is clear that a two-step purification yields protein of higher purity compared to Ni-gradient alone. However, since only a few contaminants are removed by the ion-exchange chromatography step in spite of a significant loss of sample, the single-step linear Ni-gradient became the preferred method of purification.

Figure 2.4 Peptide-fusion purifications. (A) SDS-PAGE gels of peptide-fusion B3NK purified by a Ni-Affinity Step gradient (Load-L, Flow through 1-F1, Flow through 2-F2, Flow through 3-F3, Wash 1-W1, Wash 2-W2, Wash3 W3, and Elution-E) and ion exchange (fractions A-G) are indicated. (B) SDS-PAGE gel of peptide-fusion B3NR purified by Ni-affinity linear gradient (Load-L, Wash1-W1, Wash 2-W2 and elution fractions A-C 250 mM Imidazole. Fraction D is the elution from 1M imidazole and shows tightly bound B3NK and contaminants.) Uncleaved peptide-fusion MW = 24 kDa.

Once the peptide-fusions were purified to a satisfactory degree, cleavage of the peptide from its fusion partner and linkers was accomplished by the highly specific human rhinovirus 3C protease (Fig. 2.5). Various proteases to peptide fusion ratios were examined for optimal cleavage in minimal time. As a 5-hour digestion with a 1:25 protease to protein ratio was sufficient for full cleavage of peptide from the fusion tag, this method emerged as the time-optimized protocol. To separate the liberated peptide from the other components in solution, reverse phase HPLC was employed. By employing a three phase gradient which changes in organic phase from steep for buffer component elution, to shallow for peptide elution, to a second steep phase to regenerate

Figure 2.5 Peptide-thioredoxin fusion protein cleavage. SDS-PAGE gel of the purified thioredoxin- peptide fusion (Lane 1). Purified human rhinovirus 3C protease (Lane 2) and cleavage reaction including human rhinovirus 3C protease, cleaved thioredoxin and free aPP peptide (Lane 3). Peptide MW = 3176 Da. Human rhinovirus 3C protease MW = 46,000 Da. Thioredoxin with tags MW = 27,912 Da. the column and elute larger proteins such as thioredoxin and human rhinovirus 3C protease, the peptide was separated during a 21.5 minute gradient. For example, cleaved peptides B3DE, B3II and B3NR, all were successfully separated from the contaminating proteins and cleaved thioredoxin fusion partners via the three phase gradient (Fig. 2.6).

Major peaks in the chromatogram correspond to buffer components (peak 1) and larger proteins such as the thioredoxin tag and protease (peak 3) as verified by SDS-Page analysis. To verify the peptide identity and purity from the expected peptide samples

(peak 2) on the reverse phase HPLC chromatogram, electrospray ionization mass spectrometry (ESI-MS) was performed. The spectra indicate peptides are of the expected molecular weights (3,119.1 – 3,201.3 Da) (Fig. 2.7). In addition, the chromatogram for the liquid chromatography that is in line with the mass spectrometer showed only a single peak. Averaging of spectra from all regions of the peak resulted in uniform mass spectra indicating only one molecular species was present. These data suggest that the purified

Figure 2.6 HPLC purification chromatograms of cleaved peptide-fusion samples. (A-C) SDS-PAGE gel of uncleaved and cleaved peptide-fusion with corresponding HPLC separation chromatograms. Peak 1 consists of buffer components, peak 2 is eluted peptide indicated by * and peak 3 is cleaved thioredoxin and human rhinovirus 3C protease. (A) B3II (B) B3DE (C) B3NR (D) SDS-PAGE gel of peaks 1 and 3 from HPLC separation. (E) Mass spectra of HPLC peak 2, the purified peptide B3II, MW = 3119.1 Da.

Figure 2.7 Mass spectra of peptides. (A) B3NQ MW=3143.2 Da (B) B3II MW=3119.1 Da (C) B3DE MW=3174.2 Da (D) B3DR MW=3201.3 Spectra show double (+2) triply (+3), and quadruply (+4) charged ions.

peptides are of extremely high purity.

To assess the overall utility of the peptide library construction system, the total amount of peptide fusion that was expressed in E. coli lysates is compared to the overall yield of each of the purification methods (Table 2.1). Total peptide yield ranged from 0.8 to 16% from the overall expression. A major contributor to low peptide yield is

Table 2.1 Peptide Expression, Yield and Chemical Characteristics. Total peptide fusion expressed is compared to the overall yield of the peptide as a function of purification method used. Peptide characteristics such as charge, isoelectric point (IEP), the number of acidic, basic, polar, and non- polar amino acid residues are also tabulated.

Percent peptide in raw E. coli lystates was calculated using GeneTools (SynGene). * Indicates % peptide calculated was estimated by inspection of the stained SDS-PAGE gels. Ni: Ni-Affinity chromatography. IE: ionic exchange chromatography. Acidic, Basic, Polar and Non-polar represent the number of amino acids in the 27-residue peptide that have the particular chemical characteristic.

likely loss of protein at the step of column loading in the flow through during initial purification due to overloaded resin. In addition, other contributing losses are observed at the ionic exchange and reverse phase HPLC portions of the purification. For each

peptide, the ability to be expressed and the overall yield after purification was compared to different peptide features including charge, isoelectric point, and amino acid characteristics. Peptide charge ranged from -0.06 to -4.06 and calculated isoelectric point from 3.77 to 6.92. The number of acidic residues in the peptide sequences ranged from two to five while one to three basic residues, six to twelve polar residues and seven to nine hydrophobic residues were present in the various sequences. Favorably, no correlation between the expressability of the peptide-fusion and intrinsic chemical characteristics of the peptides was found. Likewise, no correlation between peptidic character and overall yield of pure peptide was observed. This suggests that our system can be used for a wide variety of peptide sequences with a range of chemical characteristics.

2.3.Discussion

During chemical synthesis of peptides, sequence errors are a likely result due to the imperfect coupling efficiency at each step. For chemically synthesized peptides, the contaminants are extremely similar to the desired product in size and chemical characteristics and are therefore difficult to separate based on chromatography. Peptides produced by biochemical synthesis by the E. coli ribosome are expected to be very homogeneous due to the ribosome’s low error rate. In the system described here, any contaminants that remain after affinity purification and protease cleavage of the biochemically synthesized peptide fusions are unlikely to be chemically similar to the peptides. Our method for production of peptides uses reverse-phase HPLC purification as the final step. Because the non-peptide components are not chemically related to the

peptides themselves, HPLC purification allows for simultaneous removal of the fusion partner, the protease, and any residual components of the buffer solution. This makes a simple, two-step purification possible. In addition, because of the high fidelity of biochemical synthesis, the homogeneity of our peptides is very impressive. This results in a library of extremely high purity.

All of the peptide-fusion proteins described here were purified using a six- histidine affinity tag. Following a Ni-affinity step gradient purification the peptide fusions were approximately 50% pure. We probed the effect of more extensively purifying the peptide fusions to >95% purity by subsequent anion exchange chromatography or to 80% purity by using a linear gradient of imidazole on the Ni- affinity column. Employing reverse-phase HPLC, we were able to robustly purify the cleaved peptide to homogeneity after either of these protocols. This finding significantly shortened our production time by eliminating the ion-exchange purification step.

An additional advantage of the method described is that it is unnecessary for the peptide and the protease to be tagged with the same affinity tag, as is the case in some commercial protein-fusion purification systems. The method described here is compatible with any fusion partner and any protease. Since both the fusion partner and the protease are likely to be much larger than peptide products, it will be possible to separate the product from the contaminants in the final reverse-phase HPLC purification step. This fact is of particular importance in production of peptide libraries where additional amino acids at either the N- or C-termini can have significant effects on the overall properties of the peptide. Since any protease can be used, the overhanging amino acids from the

protease cleavage site can be matched with the desired peptide sequence, or a protease that does not leave any overhang can be selected37. The facts that this method is not dependent on use of any particular affinity tags or on utilizing matching affinity tags also limit sub-cloning steps that are necessary to use the library production method.

This method of peptide production using genetically encoded peptides offers clear advantages in the length and fidelity of peptides that can be produced and the resultant purity of the final samples. In the field of chemical peptide synthesis, the concept of

“difficult” sequences (e.g. hydrophobic sequences) exists. Conversely, the genetically- encoded production method described here was insensitive to peptide sequence such that every sequence attempted worked the first time with no optimization of expression or purification procedure required. To make this method truly competitive with chemical synthesis, it is essential that libraries can be produced on the same general time scale as chemical synthesis. The scheduling strategy for the twenty-member library we present here lays the foundation for production of much larger libraries using the same processes.

To this end, we have time-optimized our expression, purification and characterization protocols. Using the protocols we report and have routinely executed, we have constructed a map for methodological synchronization (Fig. 2.8). By coordinating mutagenesis cycles, sequencing, transformation, expression, purification and characterization, this peptide library can be produced rapidly. We generated this map reflecting the methods that were actually used; however, we have also performed sequencing reactions after an eight-hour growth of cultures followed by mini-prepping of

DNA for sequencing. If this were applied to each peptide construct, it would save one

Figure 2.8 Methodological Synchronization. An experimental map showing how synchronization of QuikChange-based mutagenesis and protein preparatory procedures allows the production of 20 peptides in a five-week time frame. Mutagenesis daily cycle (top) outlines the number of days in which cloning steps such as QuikChange mutagenesis, transformation, DNA purification and sequencing proceeds. 1:3, 2:3, and 3a:3 represents mutagenesis steps to create first peptide construct B3. Protein daily cycle (bottom) outlines the number of days in which expression, purification, and characterization for each peptide proceeds. The resultant DNA constructs are represented in light gray and purified peptides are represented in dark gray. day from each segment of the mutagenesis portion of the plan. In addition, we have

developed methodology so that QuikChange-mutagenized plasmids can be directly transformed into an expression strain of bacteria (such as BL21(DE3)) rather than into a cloning strain, which can save one step in the protocol. With these developments, the entire library could be generated even more rapidly. As presented, this methodological synchronization scheme allows construction of twenty peptide-expressing plasmids and purification of the resultant peptides to homogeneity in a five-week period using one

chromatography system, one HPLC and one LC-MS.

In this work, we have used no more than two liters of E. coli culture per peptide and have accepted losses at the affinity purification step. Nevertheless, the protocols discussed here could facilely be scaled up to produce even greater yields of pure peptide.

For some of the peptide fusions we have constructed, the yield of the expressed fusion protein as a fraction of total E. coli proteins is high enough (45% of total E. coli protein) that we can nearly envision skipping purification of the fusion protein altogether. By co- expressing the protease and the fusion protein it may be possible to perform a one-step purification by HPLC to yield large quantities of pure peptide.

2.4. Materials and Methods

2.4.1. Cloning of Recombinant Peptide-Fusion Variants

In order to create the parent vector, pET32-Peptide, for the peptide-fusion DNA variants, the aPP gene was amplified by PCR from the pJC2035 vector (from Alana

Schepartz and Doug Daniels). Using the forward primer 5’GTACAACCATGGCTGGA-

AGTGCTGTTTCAGGGTCCGTCCCAGCCGACCTACCC3’ that included the human rhinovirus 3C cleavage sequence (bold) and NcoI endonuclease restriction site (italics) and the reverse primer 5’TCGAGCCTCGAGCTAGTAACGGTGACGGGTAACAACG-

TTCAGG3’ which encodes the XhoI restriction site (italics) and stop codon (bold), the desired gene was produced. This gene product was then ligated into the thioredoxin fusion tag, 6xHis purification tag containing vector, pET32b (Novagen) via restriction sites NcoI and XhoI. Insertion was confirmed by sequence analysis (Genewiz, Inc.).

The newly constructed vector, pET32-Peptide, was used for PCR-based site directed

mutagenesis via QuikChange (Stratagene) to create the variable peptide sequences. Initial base constructs, B3 and B3II were created using two sets of primers. Mutational sites include Q4, T6, L17, I18, Y21, D23, Q25, L24 and Y27. Secondary base constructs

B3DQ and B3NQ were produced from the B3 parent sequence by mutating sites D10,

D11, and D16 while B3IQ and 3INQ were produced from the B3II sequence using two sets of primers. Remaining peptide sequences were produced from single step mutagenesis at position D11 utilizing one set of primers. The nucleotide sequences were optimized for expression in E. coli and all constructs were verified via sequence analysis.

2.4.2. Expression and Purification of Recombinant Peptide-Fusion Variants

DNA sequences encoding desired peptide sequences were transformed into E. coli strain BL21(DE3) competent cells (Novagen) for expression. These cells were inoculated into 2xYT media containing 100 μg/mL ampicillin (Sigma). Cells were grown at 37°C until an OD600 of 0.6 was reached. Isopropyl β-D-1-thiogalactopyranoside (IPTG),

(Anatrace) was then added to a final concentration of 1mM. Cells were allowed to grow at 37°C for an additional 3 hrs and then harvested by centrifugation at 5000 rpm (Sorvall

SLC-4000 rotor) for 10 minutes.

Cells were resuspended in lysis buffer (50 mM NaH2PO4 pH 8.0, 300 mM NaCl,

2mM imidazole) and lysed using a microfluidizer system (Microfludics). The resulting solution was then centrifuged at 15,000 rpm (Sorvall SS-34 rotor) for one hour at 4°C to remove cellular debris. Lysates were passed over HiTrapTM Chelating HP columns (GE

Healthcare). aPP, B3, B3DQ, B3DE, B3DK, B3DR, B3NE, B3NQ, B3NK, and B3II bound proteins were washed and eluted using a step gradient that consisted of a wash step

using 50 mM NaH2PO4 pH 8.0, 300 mM NaCl, 10 mM imidazole and an elution step using 50 mM NaH2PO4 pH 8.0, 300 mM NaCl, 250 mM imidazole. Peptide fusions B3,

B3DK, B3DR, B3NQ, B3NE, B3NK, B3II eluted proteins were subjected to ion exchange chromatography to obtain additional purity. The previously eluted proteins were diluted 1:5 using Buffer A (20 mM Tris pH 8.0, 2mM dithiothreitol (DTT)) and bound to Macro-Prep HighQ cartridge (Biorad) at 5% Buffer B (Buffer B: 20 mM Tris pH 8.0, 2 mM DTT, 1M NaCl). Peptide-fusion proteins were eluted via a linear gradient that ranged from 50 mM to 350 mM NaCl over 240 minutes. Peptide fusions B3NR,

B3IQ, B3IK, B3IE, B3IR, B3IL, 3INQ, 3INE, 3INR and 3INK were purified by a Ni- affinity gradient going from 0 mM imidazole to 50 mM imidazole over 300 minutes. All peptide-fusions were analyzed by 16% SDS-PAGE gels that were stained with comassie brilliant blue (Sigma). Peptide fusions were stored at -20°C until required for cleavage and separation.

2.4.3. Expression and purification of Human Rhinovirus 3C Protease

The human rhinovirus 3C gene in the pGEX vector (GE Healthcare) was transformed into E. coli strain BL21(DE3) competent cells (Novagen) for expression.

These cells were inoculated into 2xYT media containing 100 μg/mL ampicillin (Sigma).

Cells were grown at 37°C until an OD600 of 0.6 was reached. IPTG was then added to a final concentration of 1mM. Cells were allowed to grow at 20°C for 18 hours and then harvested by centrifugation at 5000 rpm (Sorvall SLC-4000 rotor) for 10 minutes.

Cells were resuspended in binding buffer (20 mM NaH2PO4 pH 7.4, 150 mM NaCl, 2mM

DTT) and lysed using a microfluidizer system (Microfludics). The resulting solution was

then centrifuged at 15,000 rpm (Sorvall SS-34 rotor) for one hour at 4°C to remove cellular debris. Lysates were passed over GSTrapTM FF column (GE Healthcare) and washed with 50 mM Tris HCl, 5mM reduced glutathione, 2mM DTT to remove loosely binding contaminants. The majority of the protease was eluted using 50 mM Tris HCl,

15mM reduced glutathione, 5mM DTT followed by a second elution step of 50 mM Tris

HCl, 35mM reduced glutathione, 5mM DTT to release remaining protease and recharge the column.

The elution was diluted 1:5 using Buffer A (20 mM Tris pH 8.0, 2mM DTT) and bound to Biorad Macro-Prep HighQ cartridge at 2% Buffer B (Buffer B: 20 mM Tris pH

8.0, 2mM DTT, 1M NaCl). Protease was eluted via a linear gradient from 20 mM to 100 mM NaCl over 50 minutes.

2.4.4. Peptide-Fusion Cleavage by Human Rhinovirus 3C and Separation from Fusion Partners

Peptide-fusion and protease were incubated at 4°C for a 5-hour digestion period in a 1:25 protease to protein ratio. Reverse phase HPLC was carried out on the cleaved peptide-fusion samples to separate the peptide from the protein components. Cleaved peptide-fusion samples were filtered through 0.22 μm syringe filter and loaded onto a

Waters Sunfire Prep C18 column (10x50 mm, 5μm, and 300Å). Separation of the peptide from fusion protein and contaminants was carried out by reverse phase HPLC on a

Shimadzu HPLC system equipped with a SPD-20AV Prominence UV/Vis detector and

LC-20AT Prominence liquid chromatograph. The mobile phase was 0.1% trifluoroacetic acid in water (Buffer A), with an elutant of 0.1% trifluoroacetic acid in acetonitrile

(Buffer B). The column was developed with a three phase gradient consisting of a steep

change in organic (5-30% Buffer B over 2.5 minutes) for buffer component elution, a shallow step (30-45% Buffer B over 15 minutes) for peptide elution, followed by a steep gradient (45-100% Buffer B over 4 minutes) to regenerate the column and elute larger proteins such as thioredoxin and human rhinovirus 3C. Absorbance was monitored at

214nm and 280 nm.

2.4.5. Mass spectrometry

To determine peptide molecular weight and degree of purity, peptide samples were analyzed using electrospray mass spectrometry. Lyophilized peptide samples were resuspended in water to a peptide concentration of 50 μM. 5 μl of sample was analyzed using an Esquire-LC electrospray ion trap mass spectrometer (Bruker Daltonics, Inc) set up with an ESI source and positive ion polarity. This system was equipped with an

HP1100 HPLC system (Hewlett Packard). Scanning was carried out between 200 m/z and

2000 m/z, and the final spectra obtained were an average of 10 individual spectra.

Equipment used was located at the Mass Spectrometry Center of the University of

Massachusetts Amherst.

2.5. References

1. Winston, R. L. & Gottesfeld, J. M. Rapid identification of key amino-acid–DNA contacts through combinatorial peptide synthesis. Chemistry & Biology 7, 245- 251, (2000).

2. Rodriguez, M., Li, S. S. C., Harper, J. W. & Songyang, Z. An Oriented Peptide Array Library (OPAL) Strategy to Study Protein-Protein Interactions. Journal of Biological Chemistry 279, 8802, (2004).

3. Sweeney, M. C. et al. Decoding protein-protein interactions through combinatorial chemistry: sequence specificity of SHP-1, SHP-2, and SHIP SH2 domains. Biochemistry 44, 14932–14947, (2005).

4. Baltrusch, S., Lenzen, S., Okar, D. A., Lange, A. J. & Tiedge, M. Characterization of Glucokinase-binding Protein Epitopes by a Phage-displayed Peptide Library Identification of 6-Phosphofructo-2-kinase/fructose-2, 6-Bisphosphatase as a Novel Interaction Partner. Journal of Biological Chemistry 276, 43915-43923, (2001).

5. Newman, J. R. S. & Keating, A. E. Vol. 300 2097-2101 (American Association for the Advancement of Science, 2003).

6. Choe, Y. et al. Substrate Profiling of Cysteine Proteases Using a Combinatorial Peptide Library Identifies Functionally Unique Specificities. Journal of Biological Chemistry 281, 12824, (2006).

7. Gosalia, D. N., Salisbury, C. M., Ellman, J. A. & Diamond, S. L. High Throughput Substrate Specificity Profiling of Serine and Cysteine Proteases Using Solution-phase Fluorogenic Peptide Microarrays*. Molecular & Cellular Proteomics 4, 626-636, (2005).

8. Gosalia, D. N., Salisbury, C. M., Maly, D. J., Ellman, J. A. & Diamond, S. L. Profiling serine protease substrate specificity with solution phase fluorogenic peptide microarrays. PROTEOMICS 5, 1292-1298, (2005).

9. Harris, J. L. et al. Vol. 97 7754-7759 (National Acad Sciences, 2000).

10. Turk, B. E., Huang, L. L., Piro, E. T. & Cantley, L. C. Determination of protease cleavage site motifs using mixture-based oriented peptide libraries. Nature Biotechnology 19, 661-667, (2001).

11. Cuerrier, D., Moldoveanu, T. & Davies, P. L. Determination of Peptide Substrate Specificity for {micro}-Calpain by a Peptide Library-based Approach: The Importance of Primed Side Interactions. Journal of Biological Chemistry 280, 40632, (2005).

12. Garaud, M. & Pei, D. Substrate profiling of protein tyrosine phosphatase PTP1B by screening a combinatorial peptide library. J. Am. Chem. Soc 129, 5366-5367, (2007).

13. Huyer, G. et al. Affinity Selection from Peptide Libraries to Determine Substrate Specificity of Protein Tyrosine Phosphatases. Analytical Biochemistry 258, 19-30, (1998).

14. Vetter, S. W. & Zhang, Z. Y. Probing the Phosphopeptide Specificities of Protein Tyrosine Phospha-tases, SH2 and PTB Domains with Combinatorial Library Methods. Current Protein and Peptide Science 3, 365-397, (2002).

15. Hutti, J. E. et al. A rapid method for determining protein kinase phosphorylation specificity. Nature Methods 1, 27-29, (2004).

16. Mah, A. S. et al. Substrate specificity analysis of protein kinase complex Dbf2- Mob1 by peptide library and proteome array screening. BMC Biochemistry 6, 22, (2005).

17. Yaffe, M. B. Study of Substrate Specificity of MAPKs Using Oriented Peptide Libraries. Methods in Molecular Biology 250, 237-250, (2004).

18. Turk, B. E., Hutti, J. E. & Cantley, L. C. Determining protein kinase substrate specificity by parallel solution-phase assay of large numbers of peptide substrates. Nat Protoc 1, 375-379, (2006).

19. Bartsevich, V. V. & Juliano, R. L. Regulation of the MDR1 Gene by Transcriptional Repressors Selected Using Peptide Combinatorial Libraries. Molecular Pharmacology 58, 1-10, (2000).

20. El-Mousawi, M. et al. A Vascular Endothelial Growth Factor High Affinity Receptor 1-specific Peptide with Antiangiogenic Activity Identified Using a Phage Display Peptide Library*. Journal of Biological Chemistry 278, 46681- 46691, (2003).

21. Jahnke, A. et al. Leukemia targeting ligands isolated from phage display peptide libraries. Leukemia 21, 411, (2007).

22. Obata, T. et al. Peptide and Protein Library Screening Defines Optimal Substrate Motifs for AKT/PKB. Journal of Biological Chemistry 275, 36108-36115, (2000).

23. Benoiton, N. L. Chemistry Of Peptide Synthesis. (Taylor & Francis, 2006).

24. Scriba, G. K. E. in Encyclopedia of Analytical Chemistry (2006).

25. Tam, J. P. & Lu, Y. A. Coupling Difficulty Associated with Interchain Clustering and Phase Transition in Solid Phase Peptide Synthesis. Journal of the American Chemical Society 117, 12058-12063, (1995).

26. Kent, S. B. H. in Proceedings of the 9th Annual American Peptide Symposium. (ed VJ Hruby and KD Kopple CM Deber) 407-414.

27. Tam, J. P. in Proceedings of the 9th Annual American Peptide Symposium. (ed VJ Hruby and KD Kopple CM Deber) 423-425.

28. Weaver, R. F. Molecular Biology. (McGraw-Hill College, 2007).

29. Smith, D. B. & Johnson, K. S. Single-step purification of polypeptides expressed in Escherichia coli as fusions with glutathione S-transferase. Gene 67, 31-40, (1988).

30. Guan, C., Li, P., Riggs, P. D. & Inouye, H. Vectors that facilitate the expression and purification of foreign peptides in Escherichia coli by fusion to maltose- binding protein. Gene 67, 21-30, (1988).

31. LaVallie, E. R. et al. A Thioredoxin Gene Fusion Expression System That Circumvents Inclusion Body Formation in the E. coli Cytoplasm. Bio/Technology 11, 187-193, (1993).

32. Blundell, T. L., Pitts, J. E., Tickle, I. J., Wood, S. P. & Wu, C. W. X-Ray Analysis (1. 4-angstrom Resolution) of Avian Pancreatic Polypeptide: Small Globular Protein Hormone. Proceedings of the National Academy of Sciences 78, 4175-4179, (1981).

33. Tonan, K., Kawata, Y. & Hamaguchi, K. Conformations of isolated fragments of pancreatic polypeptide. Biochemistry 29, 4424-4429, (1990).

34. Zondlo, N. J. & Schepartz, A. Highly Specific DNA Recognition by a Designed Miniature Protein. Nature (London) 363, 38, (1993).

35. Chin, J. W., Grotzfeld, R. M., Fabian, M. A. & Schepartz, A. Methodology for optimizing functional miniature proteins based on avian pancreatic polypeptide using phage display. Bioorganic & Medicinal Chemistry Letters 11, 1501-1505, (2001).

36. Shimba, N., Nomura, A. M., Marnett, A. B. & Craik, C. S. Herpesvirus Protease Inhibition by Dimer Disruption. Journal of Virology 78, 6657-6665, (2004).

37. Feeney, B., Soderblom, E. J., Goshe, M. B. & Clark, A. C. Novel protein purification system utilizing an N-terminal fusion protein and a caspase-3 cleavable linker. Protein Expression and Purification 47, 311-318, (2006).

CHAPTER III

INHIBITION OF CASPASE-9 BY STABILIZED PEPTIDES TARGETING

THE DIMERIZATION INTERFACE

This chapter has been in collaboration with Sumana Ghosh. Sumana perfomed all of the solid phase peptide synthesis and structural characterization of the resultant peptides by circular dichroism spectroscopy. The author performed all other aspects of the work described here.

Abstract

Caspases comprise a family of dimeric cysteine proteases that control apoptotic programmed cell death and are therefore critical in both organismal development and disease. Specific inhibition of individual caspases has been repeatedly attempted, but has not yet been attained. Caspase-9 is an upstream or initiator caspase that is regulated differently from all other caspases, as interaction with natural inhibitor XIAP-BIR3 occurs at the dimer interface maintaining capsase-9 in an inactive monomeric state. One route to caspase-9-specific inhibition is to mimic this interaction, which has been localized to the α5 helix of XIAP-BIR3. We have developed three types of stabilized peptides derived from the α5 helix, using incorporation of amino-isobutyric acid, the avian pancreatic polypeptide-scaffold or aliphatic staples. The stabilized peptides are helical in solution and achieve up to 32 μM inhibition, indicating that this allosteric site at the caspase-9 dimerization interface is regulatable with low-molecular weight synthetic ligands and is thus a druggable site. The most potent peptides are the avian pancreatic polypeptide-scaffolded peptides. Given that all of the peptides attain helical structures but cannot recapitulate the high-affinity inhibition of the intact BIR3 domain, it has become clear that interactions of caspase-9 with the BIR3 exosite are essential for high-affinity binding. These results explain why the full XIAP-BIR3 domain is required for maximal

inhibition and suggest a path forward for achieving allosteric inhibition at the dimerization interface using peptides or small molecules.

3.1. Introduction

The apoptotic process of programmed cell death has been well studied for its involvement in development in all multi-cellular organisms as well as for its roles in conditions including cancer, heart attack, stroke, Alzheimer's and Huntington disease.

Irregularities in apoptosis may contribute up to 50% of all diseases in which there are no suitable therapies1, underscoring the need to both understand and control apoptosis.

One promising approach to controlling apoptosis is through caspase regulation. The caspases are cysteine aspartate proteases known to propagate apoptosis through a cascade of cleavage reactions ultimately leading to the demise of the cell. Apoptotic caspases are typically classified into two groups, the initiator caspases, caspase-8 and -9, and the executioners, caspase-3, -6 and -7. Caspase-8 is activated in response to upstream signals, which are transmitted to caspase-8 by the DISC complex2-4 while caspase-9 is activated by association with the apoptosome5-7. Once activated, caspase-8 and -9 further propagate the apoptotic cascade by cleaving and thus activating the executioner caspases, caspase-3,

-6, and -78-11. The executioner caspases are responsible for cleaving specific intracellular targets, ultimately resulting in cell death. Due to its central role in initiating and propagating apoptosis and it’s unique regulatory mechanism12, we have focused on controlling the apoptosis initiator, caspase-9.

Caspase-9 is synthesized as this three-domain polypeptide, which is able to form homodimers. Structurally, caspase-9 consists of three domains. A prodomain, categorized as a Caspase Activation and Recruitment Domain (CARD), a large subunit, which houses

the catalytic Cys-His dyad, and the small subunit which comprises the main portion of the dimer interface 13. The zymogen caspase-9 has very low activity as a monomer.

Activity increases upon dimerization13 and increases even more profoundly upon interaction with the apoptosome5-7. The apoptosome is a heptameric complex formed by

Apaf-1 and cytochrome c in an ATP dependent process10,14,15. The uncleaved zymogen form of caspase-9 is recruited to the apoptosome by binding of its CARD domain to the

Apaf-1 CARD domain. For many years the oligomeric state of caspase-9 bound to the apoptosome has been debated. Evidence is emerging from high-resolution cryo electron microscopy that caspase-9 monomers are activated while bound to the apoptosome16.

Caspase-9 does not require intersubunit cleavage for activation. However, when bound to the apoptosome, caspase-9 is processed at Asp315, resulting in a CARD-large subunit portion of the enzyme and a small subunit beginning with the N-terminal sequence of

ATPF. Proteolytic removal of the CARD domain from the large subunit, which results in decreased activity, has also been observed. Processed caspase-9 can be displaced from the apoptosome by additional molecules of procaspase-917, resulting in release of cleaved caspase-9, which can form active dimers. As a cleaved dimer, the L2 loop from one half of the dimer is able to interact with the L2` loop from the other half in the presence of substrate similar to other caspases13. In this state, the protein is catalytically competent to process substrate. Like other caspases, caspase-9 recognizes its substrates and cleaves after specific aspartic acid residues.

Once the caspase cascade is activated, apoptosis rapidly ensues. Treating diseases in which apoptosis is activated requires specific apoptotic caspase inhibitors. A naturally occurring family of caspase inhibitors, the inhibitor of apoptosis (IAP) proteins, show

promise as models for caspase-specific inhibition. Caspase-9 is inhibited by the x-linked inhibitor of apoptosis protein (XIAP) in a manner that is distinct from the inhibition of other apoptotic caspases, including caspase-3 and -7. XIAP comprises three baculovirus inhibitory repeats (BIR). The linker between BIR1 and BIR2 binds to the active sites of caspase-3 and -7 dimers, blocking access to substrate and thereby inhibiting these caspases. The third domain of XIAP, BIR3, docks at the dimer interface of caspase-9 holding capase-9 in an inactive monomeric state. The most critical interactions occur between the α5 helix in BIR3 and the 8 strand of caspase-9 (Fig 3.1 A). In addition, the

N-terminus of the caspase-9 small subunit binds to an exosite on the BIR3 domain.

Critical interactions within these two regions of the BIR3 domain have been highlighted as essential for BIR3’s interactions with caspase-9 (Fig 3.1 B) and its inhibitory properties12. Many of these interactions are hydrogen bonds and salt bridges between the

α5 helix of BIR3 and caspase-9, often BIR3 side chains interacting with caspase-9 backbone atoms. In addition, some hydrophobic interactions are critical. For example, mutations of both polar H343 and hydrophobic L344 in the α5 helix of BIR3 result in a complete loss of its inhibitory properties12,18. Substitution of these two residues, within similar but non-functional IAP’s, such as cIAP1 and ML-IAP, restored caspase-9 inhibition enforcing the idea of the α5 helix as a main component in the BIR3 inhibitory mechanism19,20. Another region of BIR, comprised of the amino acid sequence WYPG

(residues 323 to 326), also improved the inhibitory properties when substituted in other

IAP’s, suggesting that this region is also important for the interaction. Although these key regions are localized to the first three helices of BIR3, truncation of BIR3’s N- terminal residues 245-261 resulted in an extremely unstable protein with low potency for

Figure 3.1 Interactions required for inhibition of caspase-9 are clustered in the 5 helix. (A) Structure of a caspase-9 monomer (purple) bound to XIAP-BIR3 (blue) observed in 1NW9 (upper panel). A structural zinc (red sphere) is observed in XIAP-BIR3. The exosite is marked with a circle. The boxed region is highlighted in the lower panel. Hydrogen bonds (black dotted lines) between caspase-9 monomer and the 5 helix from XIAP-BIR3 provide recognition specificity to the complex. These interactions are recapitulated in the native peptides. (B) Specific interactions observed in the structure of caspase-9 bound to XIAP-BIR3 and those present in the three classes of designs are listed. caspase-9 inhibition21. Additionally, the C-terminal truncation, BIR3 residues 348-356 had no affect on protein stability. Potency for caspsae-9 was increased only in the presence of the N-terminal region indicating that stabilization of the C-terminal region of

BIR3 is required for its inhibitory properties against caspase-921. We hypothesize that this unique mechanism of BIR3 inhibition of caspase-9 may enable development of caspase-9 specific inhibitors if the α5 helix can be sufficiently stabilized.

To date, the vast majority of work towards caspase-specific inhibition has focused on creating small-molecule and peptide-based active-site inhibitors. Achieving specificity at the active site has been difficult because all caspases have a stringent requirement for binding aspartic acids in the S1 pocket. This means that both apoptotic and inflammatory caspases are subject to inhibition by very similar inhibitors. Most caspase inhibitors have consisted of a peptide or peptidomimetic with an acid functionality to bind in the S1 pocket of the active site and some kind of cysteine-reactive warhead. Utilizing the BIR3 mechanism of inhibition is promising in that it is unique to caspase-9 and should avoid inhibition of other caspases. In this work we have developed peptide-based inhibitors that mimic BIR3 inhibition of caspase-9.

Peptide mimics as surrogates for large protein-protein interactions have been extensively pursued. In one early example, an entire interface of the vascular endothelial growth factor was reduced to a 20-amino acid peptide which could then be optimized via phage display to achieve the needed binding affinity22. The atrial natriuretic peptide was similarly minimized to half the original size using rational design and phage display and still bound23. Some proteins, like the β-adrenergic receptor, can be controlled with small, unstructured peptides in which the principal requirement for binding is an available amine. On the other hand, successful mimics of the vast majority of protein-protein interactions attempted to date have required a structured peptide to mimic the native protein interactions. Thus a number of approaches have been developed to stabilize fragments of proteins in their native structure. For helices the prominent methods are to initiate or stabilize helix formation or use small scaffolding proteins. Some approaches have targeted naturally occurring capping motifs or side-chain cross-linking methods

such as disulfide bonds, lactam bridges and metal mediated bridges24-33. Main chain conformational restrictions using unnatural α-methylated amino acids, such as aminoisobutyric acid (Aib), have also been explored. The placement of Aib in peptides restricts the conformations of the peptide bonds, promoting proper torsion angles for α- helical formation34. More recently, all hydrocarbon cross-linking side chains, sometimes called peptide staples have been introduced to maintain helical conformations of small peptides35. These hydrocarbon staples have been shown to improve helicity as well as cell permeability of peptides derived from a BID BH3 helix36, the NOTCH transcription factor complex37, and a p53 helix38. In this ‘stapling’ method, unnatural α-methylated amino acids containing olefinic side-chains of the appropriate length, are placed at the i and i+4 or i and i+7 positions of the helix to be stabilized. These positions are selected because one turn of an α-helix is 3.6 residues in length thus the staples would span one or two turns of the helix. Cyclization of these amino acids is performed through an olefin metathesis reaction, facilitated by Grubbs catalyst, thus locking the amino acids into a staple formation reinforcing nucleation of the α-helix39. Others have enforced main-chain interactions through hydrogen bond surrogates which places a covalent link between the backbone of the i and i+4 positions utilizing a similar olefin metathesis reaction40. An orthogonal approach utilizing stabilized scaffolds called miniature proteins has also proven useful in helix stabilization41,42. The critical amino acids for the protein-protein interface are grafted onto a stable helical scaffold in the appropriate register to obtain the proper amino acid orientation for the interaction. For example, the scaffold of avian pancreatic polypeptide (aPP) has been used to design Bcl-2 based inhibitors43, p53-hDM2 interaction inhibitors44, herpes virus protease dimer disrupters45, as well as mimicking

protein-DNA interactions46-48. More recently, Peptide YY has also been used as a miniature protein scaffold42.

Upon analysis of the caspase-9/BIR3 complex and by understanding the successes of stabilized helices, our approach to caspase-9 inhibition is two-fold. First, we aim to investigate which regions of the α5 helix of BIR3 are essential to achieve BIR3-like inhibition. Secondly, we aim to utilize the α5 helix region of BIR3 in order to achieve caspase-9 inhibition via small α-helical peptides42.

3.2. Results

To achieve inhibition of caspase-9 utilizing the interactions observed between the

BIR3 α5 helix and the caspase-9 8 strand (Fig 3.1A), we predicted that some type of helix stabilization or helical scaffold would be required. We have designed and synthesized three different classes of α5-stabilized peptides and tested them for their ability to inhibit caspase-9. These three classes: native and Aib-stabilized, aPP- scaffolded, and aliphatic stapled peptides take three entirely different approaches to helix stabilization, but each of them recapitulates the most important interactions from the α5 helix (Fig 3.1 B). The stabilized peptides are markedly more helical and are better inhibitors than the analogous native, non-stabilized peptides.

3.2.1. Native and Aib-Stabilized Peptides

Our first approach was to isolate native sequences from the BIR3 α5 helix (Fig

3.1 A, 3.2). In some sequences, aminoisobutyric acid (Aib) was inserted at various positions to stabilize the helical conformation of the peptide. Aib is an α-branched amino acid analogous to the naturally-occurring amino acid alanine, but derivatized at the α- carbon with two equivalent methyl groups. The presence of two methyl groups on the α-

carbon locks adjacent peptide bonds into the α- helical confirmation which supports the desired dihedral angles, and thus has been widely used to stabilize helical Figure 3.2 Sequences of the α5 region of XIAP-BIR3 (BIR3) and peptides34,49-51. We peptides 1-9. * represents aminoisobutyric acid (Aib) designed peptides ranging from 11 to 35 amino acids in length. The longest,

Peptide 1, composed of BIR3 residues 315-350, was designed to contain the α3-α5 helices. The α3 helix forms part of the exosite; the 4 helix orients the WYPG region and the α5 helix contains the major elements for recognition of caspase-9. Peptides 2-4 are

21-25 amino acids long and contain all of the core interactions from α3, α4 and α5 helices. Peptide 9 is similar to Peptide 2 but is stabilized in four non-interacting positions by Aib. Peptide 5 is 11 residues long and derived directly from the α5 helix. In Peptide 6 we added helix capping residues at the N- and C-termini. Peptides 7 and 8 are stabilized at non-interacting position by Aib. The circular dichroism (CD) spectra suggested that the native peptides were less helical than the Aib-stabilized peptides (Fig 3.3), predicting that the Aib-containing peptides were closer to their target structures than the peptides constructed solely from native amino acids derived directly from BIR3.

Caspase-9 is most active prior to catalytic removal of the 16 kD caspase activation and recruitment domain (CARD). The CARD domain is responsible for associating caspase-9 with the apoptosome. However, in the absence of the apoptosome,

the location of the CARD domain remains uncharacterized.

Due to the size of the CARD and its ability to Figure 3.3 Circular dichroism spectra of the native Peptide 5 and the Aib-stabilized Peptide 8. These spectra demonstrate that the enhance caspase-9’s basal presence of Aib increases the helicity of the peptides. catalytic activity through an as yet unknown mechanism, we reasoned that the peptide inhibitors may show a preference for interaction with either full-length caspase-9 (C9FL: containing the CARD, large and small subunits) due to positive interactions with the

CARD or a preference for

ΔN caspase-9 (C9ΔN: containing only the large and small subunits) due to negative interactions with the CARD. Finding no difference in the ability of the inhibitors to influence full-length or ΔN caspase-9 would suggest that the peptides do not interact in Figure 3.4 Native and Aib-stabilized peptides show some any way with CARD. We inhibition of caspase-9 activity. Peptides (1-9) exhibit some inhibition of (A) full-length caspase-9 (C9 FL) or (B) the N- terminal CARD-domain deleted caspase-9 (C9 N) activity to tested both C9FL (Fig 3.4 cleave a natural caspase-9 substrate, the caspase-7 zymogen (C7 C186A) to the caspase-7 large (C7 Lg) and small (C7 Sm) A) and C9ΔN (Fig 3.4 B) subunits in an in vitro cleavage assay monitored by gel mobility.

in an in vitro gel-based caspase-9 assay, which monitors cleavage of caspase-7 (substrate)

(Fig 3.4, Table 3.1) and in a fluorescent peptide cleavage assay in which we observed no difference between the two enzymes (Table 3.1). Peptide concentrations were calculated by absorbance at 280 nm or by using densitometry measurements on the purified peptides analyzed by SDS-PAGE. Although it is difficult to quantify peptides in this manner, we have confirmed that the error in the determination of peptide concentrations is no more than 2-fold.

For peptides in this study an apparent IC50 (IC50 app) was fit from an inhibitor titration at a fixed substrate concentration. This was compared to the amount of inhibition observed in a gel-based assay where the peptide concentration was fixed at 66.7 μM, a concentration in excess of the best IC50 app observed (Table 3.1). The full BIR3 domain, with a reported Ki of 10-20 nM (18, 20, 52) is an effective inhibitor in both the fluorescence assay, which uses a small fluorogenic peptide substrate mimic and in the gel based assay, which uses a natural substrate. In both activity assay formats, we observed moderate inhibition by many of the peptides. Other investigators have traditionally relied on gel-based assays to assess caspase-9 inhibition and activation. This is likely due to the greater reproducibly and lowered requirements for peptide consumption in the gel-based assay, which we have also observed. The inhibition that we observe in the two assays formats, agrees qualitatively for the peptides we have tested. All of the native and Aib- stabilized peptides showed an IC50 app of >100 μM.

Peptide 2 routinely activated caspase-9 in samples of aged peptide (incubated for three weeks at 4°C). Initially we were intrigued by the ability of aged Peptide 2 to activate caspase-9. We confirmed that samples of aged Peptide 2 were of the same

Table 1: Overall Inhibition of Caspase-9 Full Length by Peptides a b Peptide % Inhibition IC50 app (66.7 μM peptide) (μM) 1 0 >100 2 0 >100 3 0 >100 4 1.1 ± 2.4 >100 5 0.45 ± 1.1 >100 6 0 >100 7 6.3 ± 11 >100 8 13 ± 16 >100 9 15 ± 14 >100 10 n.d. 64 ± 13 11 24 ± 13 32 ± 11 12 8 ± 11 >50 13 0 >100 14 13 ± 7.1 50 ± 3.8 15 0 >100 16 6.2 ± 2.6 >100 17 22 ± 23 50 ± 5.1 18 17 ± 9.6 >100 19 24 ± 8.4 80 ± 4.8 20 2.2 ± 1.2 >100 21 2.9 ± 3 >100 22 0 >100 23 0 >100 24 1.8 ± 2.6 >100 25 2.5 ± 3.5 >100 26 8.9 ± 12 >100 27 0 > 100 28 0 >100 a Proteolytic cleavage gel-based caspase-9 activity assay which monitors cleavage of natural substrate, caspase-7. Percent inhibition was calculated using Gene Tools (SynGene by producing a standard curve from densiometry values for processed and unprocessed substrate, (caspase-7 C186A). b Fluorescence-based caspase-9 activity assays which monitor cleavage of fluorogenic substrate (LEHD-AFC).

molecular weight as the freshly diluted Peptide 2 immediately following synthesis, confirming that there were no chemical modifications upon aging

(Fig 3.5 A). Freshly diluted Peptide 2 samples were incapable of Figure 3.5 Peptide 2 non-specifically activates caspase-9. (A) Peptide activation, whereas the 2 (MW 3118) is chemically stable even after a 3-week, 4˚C incubation. (B) Aged peptide 2 activates caspase-9 activity (C) 3-week incubated peptides were aged peptide non-specifically activates both caspase-9 and caspase-7 in a manner similar to the polymeric crowding agent PEG 8,000 (Fig 3.5 B). Although aged Peptide 2 samples were capable of activating caspase-9 four-fold over basal rates of hydrolysis of the fluorogenic LEHD-AFC peptide, they were similarly capable of activating other caspases, including caspase-7 (Fig 3.5 C). Caspase active sites are highly mobile and are thus responsive to molecular crowding agents, including polymers like polyethylene glycol (PEG)52. The type of activation from aged Peptide 2 is similar to that observed by PEG 8000 (Fig 3.5 C), suggesting that aggregation occurring over a three- week aging process at 4°C was responsible for this level of activation.

3.2.2. aPP-Scaffolded Peptides

Our second approach for developing stabilized α5 helices used the avian pancreatic polypeptide (aPP) scaffold (Fig 3.6). aPP is a 36-amino acid peptide derived from the pancreas of turkey. Its native role is in the feedback inhibition loop halting

Figure 3.6 BIR3 interactions grafted onto stabilized miniature protein aPP. (A) Modeled interactions of designed peptides based on an aPP scaffold (yellow) were designed to mimic the interactions between caspase-9 (purple) and the 5 helix. The polyproline helix labeled in the upper panel, has been removed from the lower panel for clarity. (B) Sequences of native BIR3, derivative Peptides 10-26 and native aPP, in which critical residues from the 5 region of XIAP-BIR3 have been inserted into the aPP scaffold. Highly varied positions are shaded. pancreatic secretion after a meal (reviewed in53). In the crystal structure, aPP is composed of a standard alpha helix flanked by a polyproline helix (Fig 3.6 A)54,55. The interaction between these two helices provides stability for the folded conformation. aPP is amenable to substitution on many faces41,48 and can withstand a C-terminal truncation55 without negatively effecting the structure or function. α5-mimicing peptides were designed by structurally aligning the α-helical region of aPP with the α5 helix in the

BIR3-Caspase-9 complex. aPP was aligned principally by superposition of the Cα positions and tested for maximum overlap in all possible helical registers. We ultimately selected one register (Fig 3.2, 3.6 A) because it maximized overlap of productive interactions between naturally-occurring aPP amino acids and caspase-9, such as the

WYPG amino acid region which was found to be critical in converting other IAP’s to inhibit caspase-919,20,45. In addition this orientation predicts no steric clashes between the

aPP N-terminus, the polyproline helix, and caspase-9. Ultimately we chose to truncate aPP to prevent steric clash between the aPP C-terminus and caspase-9. To make Peptide

10, we grafted the critical BIR3 residues 323-6, 336-7 and 340-4 onto aPP at positions

17-18 and 21-25 (Fig 3.6 B). We also grafted residues 323-326, which compose the sequence WYPG between the α3 and α4 helices of BIR3, at positions 6-9 of aPP. aPP positions 6-9 are in the polyproline helix and are designed to support the interactions of the WYPG region of BIR3 with caspase-9. In Peptides 11-14, we varied the identity of position 11 to enable novel interactions with the loop region in caspase-9 and changed all of the aspartates to glutamates. Caspases cleave exclusively following aspartate residues, so substitution of these residues was designed to limit their caspase substrate characteristic. In Peptides 15-18, the aspartates were substituted by asparagines to prevent caspase cleavage and position 11 was varied to introduce novel interactions. In

Peptides 19-26, we systematically returned positions that had previously been shown to be important for the fold and stability of aPP to the identity in the native aPP sequence.

To produce this series of caspase-9 inhibitors, Peptides 10-26, we developed a novel expression system that allowed robust production and purification of aPP-based peptides56. The mass spectra of peptides 10-26 indicated that each of these peptides are produced accurately in bacteria and that the peptides could be purified to a very homogenous state using our methodology56. The conformation of these truncated versions of aPP were predicted by Rosetta57,58 to fold into a helical structure (Fig 3.7). In all of our aPP scaffolded designs, additional interactions are created between Y7, D10 and D11 of aPP’s polyproline helix and the caspase-9 interface. Neither truncation of aPP nor the insertion of the α5-derived residues appears to have a substantial effect on the

Figure 3.7 aPP and peptide structures predicted computationally (black) by Rosetta. Predicted structures are superimposed on the known structure of aPP (gray). RMSD for backbone atoms suggests the majority of the designed aPP-based peptides should adopt a helical conformation in aqueous solution. structure of aPP-derived peptides as the CD spectra of the designed peptides show the same conformation as the aPP parent (Fig 3.8 A). All spectra indicate that these peptides are mainly composed of polyproline helix character with a negative peak at 198-206 nm59,60 and a predicted 5% of α-helical content61. Although these data suggest that the aPP peptides fold into a less helical conformation that the target helical conformation of the scaffold, the aPP-based peptides were still tested for their ability to inhibit full-length

Figure 3.8 Properties of aPP based peptides. (A) The CD spectra of the aPP and Peptides 11 and 25 indicate that the structure of the designed peptides is very similar to that of native aPP in solution. (B) aPP-based peptides show some inhibition of full-length caspase-9 (C9 FL) or the N-terminal CARD- domain deleted caspase-9 (C9 N) to cleave a natural caspase-9 substrate, the caspase-7 zymogen (C7 C186A) to the caspase-7 large (C7 Lg) and small (C7 Sm) subunits in an in vitro cleavage assay monitored by gel mobility.

and N caspase-9 in the gel based (Fig 3.8 B) and fluorescent peptide cleavage assays

(Table 3.1). Of this class, the best inhibitors were peptides 10, 11, 14, 17 and 19 with

IC50 app of 64, 32, 50, 50 and 80 M respectively. The majority of the aPP-based peptides had IC50 app greater than 100 M. Our four best peptides 10, 11, 14, and 17 each contain all of the interacting residues that make up the BIR3 interface while Peptide 19 incorporates those residues important for the stability of the aPP scaffold. Exclusion of any of the critical BIR3 residues appears to be unfavorable for the peptides inhibitory properties based on our findings considering Peptide 19 could potentially pose as an active site competitor with three aspartate groups at positions ten, eleven and sixteen.

Peptides 11, 14 and 17 are devoid of aspartate residues, which decreases the propensity of the peptides to interact with the caspase-9 substrate binding groove and thus increases the probability of interaction with the caspase-9 dimer interface. Interestingly, the only difference between peptides 10 and 11 is the substitution of three aspartates present in peptide 10 for three glutamates. Thus the improved potency of Peptide 11 over Peptide

10 could be related to differences in caspase-9-mediated cleavage. In addition, position eleven on the scaffold in Peptide 11, 14, and 17 contains a glutamate, arginine or lysine, respectively, at residue 11. These residues were designed to form an additional salt bridge with the caspase-9 interface (not present in the native BIR3 domain) thus enhancing their ability to inhibit caspase-9 activity. This appears to have been successful as these peptides have the best inhibition properties. Interestingly, the addition of a glutamine at position 11 did not show any favorable effects on the inhibitory properties of the peptides, suggesting that glutamate, arginine, and lysine can form specific interactions that are not possible for glutamine.

3.2.3. Aliphatic Stapled Peptides

The third approach to achieve caspase-9 allosteric inhibition at the dimer interface was to incorporate aliphatic staples to stabilize short helices derived directly from the α5 helix in BIR3 (Fig

Figure 3.9 Aliphatic stapled peptide design. (A) Modeled interactions of α5- 3.9A). Aliphatic derived peptides stabilized by aliphatic staples (green) with caspase-9 (purple). (B) Synthetic route for production of the pentyl alanine amino acid staples have been used to generate the aliphatic peptide staples used successfully to stabilize helices and increase cell permeability of the peptides in which they are incorporated36,62. To synthesize the stapled peptides, we adapted the synthetic route reported by Seebach et.al63-65 to synthesize three unnatural Fmoc- protected amino acids used in the stapling reactions (Fig 3.9B). In this synthetic approach we used a mild organic base, potassium trimethyl-silanolate that could perform both the

CBz deprotection and five-membered ring opening reactions in a single step to obtain the final α-α-disubstituted amino acids. This method is less hazardous than traditional two- step methods35,66, which require the use of sodium in liquid ammonia for hydrogenolysis of the five-membered ring, followed by a reaction with TFA to deprotect the Boc group

to obtain α-α-disubstituted amino acids. Our adapted method produced the non-native amino acids at yields comparable to the traditional published routes generally used to obtain the unnatural amino acids used in peptide stapling. We focused on two pairs of stapling peptides.

Peptide 27 is 11 amino acids long and composed of residues 336 to 345 from the α5 helix of BIR3. In peptide Figure 3.10 Properties of aliphatically stapled peptides. (A) Sequences of BIR3 and stapled Peptides 27 and 28. x, y and z represent the synthetic, non-native amino 27 amino acids Y338 and I342 from acids as listed. (B) Ring-closing metathesis reaction performed on the unstapled Peptide 27 to form the BIR3, which point away from the aliphatic stapled Peptide 27 with an 8-carbon macrocyclic linker. The structure of aliphatically caspase-9 dimer interface, were both stapled Peptide 28 contains an 11-carbon macrocyclic linker replaced by S-2-(4’pentyl)alanine (Fig 3.10A). Catalysis of the ring closing metathesis forms an 8–carbon alkyl macrocyclic cross-link (Fig 3.10B). Peptide 28 is composed of an N-terminal threonine for helix capping of BIR3 residues 336-348. In Peptide 28 BIR3 residues Y338 and T345 were replaced by R-2-(4’pentyl) alanine and S-2-

(4’octyl)alanine respectively. Threonine was added to the N-terminus of Peptide 28 because it has a higher helix capping propensity than the native residue, glycine

(reviewed in67). The C-terminus was likewise extended because of the improved helical propensity of the residues SLE (reviewed in67). Ring closing metathesis results in an 11-

carbon aliphatic linker (Fig 3.10B) which has been shown to be the optimal staple for i to i+7 stapling35. As predicted, both peptides appeared to be substantially more helical in aqueous solution following the ring closing metathesis reaction to generate the aliphatic staple than prior to closure of the staple (Fig 3.11A). Thus in both cases we can conclude that the peptides attain the designed helical structure. The stapled peptides were both tested against caspase-9 full-length or ΔN caspase-9 (Fig 3.11B), but did not show strong inhibition at concentrations below 100 μM in either assay format on either caspase-9 construct.

Figure 3.11 Analysis of aliphatic stapled peptides. (A) The CD spectra of Peptides 27 and 28 indicate a significant increase in the helicity following the formation of the aliphatic staple. (B) Stapled Peptides 27 and 28 show some inhibition of full-length caspase-9 (C9 FL) or the N-terminal CARD-domain deleted caspase-9 (C9 ΔN) to cleave a natural caspase-9 substrate, the caspase-7 zymogen (C7 C186A) to the caspase-7 large (C7 Lg) and small (C7 Sm) subunits in an in vitro cleavage assay monitored by gel mobility.

3.3. Discussion

All three classes of peptides are capable of attaining helical structures to present the appropriate amino acids to the caspase-9 dimer interface. The length and the type of scaffold did appear to have a tremendous influence on the potency of the peptides. The aPP-scaffolded peptides were more successful than the native, Aib-stabilized or stapled peptides. Our data suggest peptides composed of 15 amino acids or less in length such as

Peptides 5-8 of the native and Aib stabilized class or Peptides 27-28 of the stapled peptides may be too short to have the requisite interaction for high-affinity binding to caspase-9. Models of these small peptides bound to caspase-9 bury 900Å2 of surface area in contrast to the 1700Å2 for models of the aPP-scaffolded peptides and the 2200Å2 interface found in the native BIR3-caspase-9 complex. It is possible that other non- critical residues found in the large interface potentially provide a hydrophobic interaction, which may be lacking in the short peptide sequences, thus decreasing their ability to efficiently inhibit caspase-9.

Peptides ranging in length from 11 to 39 amino acids have achieved high affinity binding utilizing the various stabilizing methods we have employed here37,38,44-47,68-70. An efficient peptide inhibitor of the hdm2–p53 interaction was shown to improve potency based on additional helical constraints imposed by Aib where incorporation of four Aib residues into a 12 amino acid peptide was required68. Our peptides utilized two Aib residues for an 11 amino acid stretch, so perhaps a greater number of Aib residues could further improve binding properties. The aPP-scaffolded Kaposi’s sarcoma-associated herpesvirus protease (KSHV Pr) inhibiting peptide, which was 30 amino acids long, was also designed with a five amino acid C-terminal truncation and altered ten amino acids within the α-helical region of aPP. This peptide was successful at disruption of dimerization, however it required 200-fold molar excess of peptide in order to obtain

50% inhibition of protease activity45. Our best peptides were at least as effective. We observe 50% inhibition with an average of ~30-fold molar excess peptide. Additionally, a

13 amino acid stapled peptide co-activator mimic of the estrogen receptor was able to achieve high affinity binding. The authors noted however, that in these short peptides, the

hydrocarbon staple can alter the binding geometry via its ability to interact with hydrophobic interfaces thus perturbing the desired interactions required for inhibition70.

Thus, although our peptides were not as potent as the parent BIR3 domain, they are of similar efficacy as other stabilized peptides. Most importantly, these peptides demonstrate that the dimerization interface of caspase-9 can be targeted by small- molecular weight inhibitors, indicating that this is indeed a druggable site.

Based on spectroscopic data, it is clear that all three classes of peptides we designed and synthesized have been stabilized in a helical conformation by incorporation of Aib, aliphatic staples or by the aPP scaffold. Despite the measured helicity, we have not been able to recapitulate the 10-20 nM interaction of BIR3 with caspase-918,20,71.

Consideration of critical residues, additional interface interactions, scaffold requirements, and fold led to production of some peptide inhibitors of caspase-9 in the micromolar range. The most potent inhibitors were all members of the aPP scaffolded class of peptides. This class provided the secondary structural requirements needed to mimic the

α5 helix interactions, provided the additional interactions of the WYPG region of BIR3 as well as providing a larger surface area coverage, which the other peptide classes lack.

Although native and Aib-stabilized Peptides 1-9 and the stapled Peptides 27-28 contain all of the critical contacts for interacting with caspase-9 through the α5 helix and were helical in solution, they were still not highly potent inhibitors. This suggests that the α5 helix alone is not sufficient for full inhibition of the intact BIR3 domain even when it is presented in a stable, helical structure. This observation further suggests that the exosite of BIR3 (composed of residues 306-314) which binds the N-termini of the small subunit of caspase-9 provides additional, mainly hydrophobic interactions, required for high

affinity inhibition (Fig 3.12 A,B). Prior reports suggested that lack of stability explained the inability of truncations of the BIR3 domain to inhibit caspase-921. Together our data on stabilized α5 helices provide a model for why full length BIR3 is required for the interaction. Although the α5 helix has been shown to contain the most critical residues for recognizing caspase-9, the BIR3 exosite also appears to be essential. The BIR3 exosite is composed of residues 306-314. These residues do not fold into any regular element of secondary structure, but exist in an ordered loop. This loop is held in the appropriate conformation from behind by two other loops from the 270s and 290s region of BIR3

(Fig 3.12 B). Furthermore, this region is stabilized via the N-terminal loop of BIR3 indicating the lack of this region leaves the exosite residues unordered. The exosite must

Figure 3.12 Full-length BIR3 is required for high-affinity caspase-9 inhibition. (A) Sequence of BIR3 and regions of BIR3 contained in the caspase-9 inhibitory peptides. (B) Interactions of caspase-9 monomer (purple) small subunit N-terminus (L2’ loop) with BIR3 (blue) exosite residues, which are contained within the box.

interact with the very flexible N-terminus of the caspase-9 small subunit also called the

L2’ loop. Because the exosite region does not adopt any regular secondary structure it is difficult to envision how to recapitulate the high-affinity BIR3:caspase-9 interaction with any smaller, contiguous region of BIR3. Thus, we now understand why full-length BIR3 is required for full-potency inhibition of caspase-9 via the dimer interface. Furthermore, these studies provide a roadmap for design of future BIR3-like small-molecule or peptide inhibitors. High potency inhibitors that block caspase-9 inhibition by a BIR3-type mechanism must necessarily encompass all the critical interactions in the exosite as well those in the α5 helix.

3.4. Materials and Methods

3.4.1. Caspase-9 Expression and Purification

The caspase-9 full-length gene (human sequence) construct in pET23b (Addgene plasmid 1182972) was transformed into the BL21 (DE3) T7 Express strain of E. coli

(NEB). The cultures were grown in 2xYT media with ampicillin (100 mg/L, Sigma-

Aldrich) at 37°C until they reached an optical density of 1.2. The temperature was reduced to 15°C and cells were induced with 1 mM IPTG (Anatrace) to express soluble

6xHis-tagged protein. Cells were harvested after 3 hrs to obtain single site processing.

Cell pellets stored at -20°C were freeze-thawed and lysed in a microfluidizer

(Microfluidics, Inc.) in 50 mM sodium phosphate pH 8.0, 300 mM NaCl, 2 mM imidazole. Lysed cells were centrifuged at 17K rpm to remove cellular debris. The filtered supernatant was loaded onto a 5 ml HiTrap Ni-affinity column (GE Healthcare).

The column was washed with 50mM sodium phosphate pH 8.0, 300 mM NaCl, 2 mM imidazole until base lined. The protein was eluted using a 2-100mM imidazole gradient

over the course of 270 mL. The eluted fractions containing protein of the expected molecular weight and composition were diluted by 10-fold into 20 mM Tris pH 8.5, 10 mM DTT buffer to reduce the salt concentration. This protein sample was loaded onto a 5 ml Macro-Prep High Q column (Bio-Rad Laboratories, Inc.). The column was developed with a linear NaCl gradient and eluted in 20 mM Tris pH 8.5, 100 mM NaCl, and 10 mM

DTT buffer. The eluted protein was stored in -80°C in the above buffer conditions. The identity of the purified caspase-9 was analyzed by SDS-PAGE and ESI-MS to confirm mass and purity.

Caspase-9ΔCARD was expressed from a two plasmid expression system52,73,74.

Two separate constructs, one encoding the large subunit, residues 140-305 and the other encoding the small subunit, residues 331-416, both in the pRSET plasmid, were separately transformed into the BL21 (DE3) T7 Express strain of E. coli (NEB). The recombinant large and small subunits were individually expressed as inclusion bodies.

Cultures were grown in 2xYT media with ampicillin (100 mg/L, Sigma-Aldrich) at 37°C until they reached an optical density of 0.6. Protein expression was induced with 0.2mM

IPTG. Cells were harvested after 3 hrs at 37°C. Cell pellets stored at -20°C were freeze- thawed and lysed in a microfluidizer (Microfluidics, Inc.) in 10mM Tris pH 8.0 and 1mM

EDTA. Inclusion body pellets were washed twice in 100mM Tris pH 8.0, 1mM EDTA,

0.5M NaCl, 2% Triton, and 1M urea, twice in 100mM Tris pH 8.0, 1mM EDTA and finally resuspended in 6M guanidine hydrochloride. Caspase-9 large and small subunit proteins in guanidine hydrochloride were combined in a ratio of 1:2, large:small subunits, and rapidly diluted dropwise into refolding buffer composed of 100mM Tris pH 8.0, 10% sucrose,0.1% CHAPS, 0.15M NaCl, and 10mM DTT, allowed to stir for one hour at

room temperature and then dialyzed four times against 10mM Tris pH 8.5, 10mM DTT, and 0.1mM EDTA buffer at 4°C. The dialyzed protein was spun for 15 minutes at

10,000rpm to remove precipitate and then purified using a HiTrap Q HP ion exchange column (GE Healthcare) with a linear gradient from 0 to 250mM NaCl in 20 mM Tris buffer pH 8.5, with 10 mM DTT. Protein eluted in 20 mM Tris pH 8.5, 100 mM NaCl, and 10 mM DTT buffer was stored in -80°C. The identity of the purified caspase-

9ΔCARD was analyzed by SDS-PAGE and ESI-MS to confirm mass and purity.

3.4.2. Caspase-7 WT and Caspase-7 C186A Expression and Purification

The caspase-7 full-length human gene in the pET23b vector or the caspase-7

C186A variant (zymogen), made by Quikchange mutagenesis (Stratagene) of the human caspase-7 gene in pET23b (gift of Guy Salvesen72) were transformed into BL21 (DE3)

T7 Express strain of E. Coli. Induction of caspase-7 was at 18°C for 18 hrs. These proteins were purified as described previously for caspase-775. The eluted proteins were stored in -80°C in the buffer in which they were eluted. The identity of purified caspase-7 was assessed by SDS-PAGE and ESI-MS to confirm mass and purity.

3.4.3. Peptide Production

Peptides 1-9 were synthetically produced by New England Peptide (Gardner,

MA). The aPP scaffolded peptides 10-26 were designed and purified using methods previously described56. Unnatural amino acids for Peptides 27-28 were synthesized by the following reactions:

3.4.3.1. Synthesis of N-Fmoc-S-2-(2’-pentyl)alanine (5)

(2S,4S)-2-Phenyl-3-(carbobenzyloxy)-4-methyloxazolidin-5-one(2S,4S), (1)

To a stirred solution of Z-D-Alanine (0.4 g, 1.8 mmol) and benzaldehyde dimethyl acetal

(0.4 ml, 2.61 mmol) in Et2O (5 mL) was added 2 ml (15.7 mmol) of BF3• Et2O slowly by maintaining the reaction temperature at -78 °C. The mixture was allowed to reach -20°C and then stirring was continued at -20 °C for 4 days. At the end of this time period, the reaction mixture was slowly added to ice-cooled, saturated aqueous NaHCO3 (10 ml), and the mixture was stirred for additional 30 min at 0°C. After the aqueous work-up, the separated organic layer was washed thrice with saturated NaHCO3 and H2O and then dried over Na2SO4. The solvent was removed in vacuo. The residue was dissolved in 11 ml of Et2O/hexane (4/7) for recrystallization and afforded white crystals of (2S,4S)-1

1 (0.37 g, 70%). H NMR (CDCl3, 400 MHz): δ ppm 1.59 (d, J = 6.8 Hz, 3H), 4.49 (q, J =

6.8 Hz, 1H), 5.13-5.16 (m, 2H), 6.64 (s, 1H), 7.26-7.42 (m, 10H).

(2S,4R)-Benzyl-4-methyl-5-oxo-4-(pent-4-enyl)-2-phenyloxazolidine-3carboxylate, (2S,4R), (3)

(2S,4S)-1 (0.22 g, 0.71 mmol) was dissolved in anhydrous THF/HMPA (4:1, 2 ml) under an argon atmosphere. The resultant solution was cooled to -78o C. Into that 1M

LiHMDS solution (1.1 ml, 1.065 mmol) in THF was added slowly under nitrogen at -

78°C. After the addition of LiHMDS, the slightly yellow solution was stirred at this temperature for 1 h. Next 5-iodo-1-pentene (0.21 g, 1.06 mmol, synthesized from 5-

Bromo-1-pentene) was added dropwise into the reaction mixture under an argon atmosphere and the resultant mixture was slowly allowed to reach room temperature. The resultant reaction mixture was stirred at room temperature overnight. At end, saturated

NH4Cl solution was added, and the reaction mixture was extracted in ether. The organic layer was washed subsequently with saturated NaHCO3 and saturated aqueous NaCl solutions, respectively. The organic phase was dried over Na2SO4 and evaporated. The

product was purified by flash column chromatography and elution with 30% DCM in

1 hexane gives the gummy product (0.08 gm) with 30 % isolated yield. H NMR (CDCl3,

400 MHz): δ ppm 1.26-1.33 (m, 2H), 1.69 (s, 1H), 1.79 (s, 3H), 2.06-2.08 (m, 2H), 2.14-

2.196 and 2.49-2.56 (m, 1H), 4.91-5.035 (m, 4H), 5.56-5.77 (m, 1H), 6.4 (d, 1H), 6.88 (d,

J = 7.0 Hz, 1H), 7.20-7.416 (m, 10 H).

S-2-Amino-2-methylhept-6-enoic-acid, (4)

A solution of 3 (1.1 gm, 2.9 mmol) in tetrahydrofuran (52 ml) was treated with potassium trimethylsilanolate (90% pure; 1.1 gm, 8.6 mmol) and the mixture was heated at 75°C for 2 h 30 min by which time all starting material was consumed. The mixture was diluted with methanol and the solvents were removed under reduced pressure. The resultant mixture was diluted with dichloromethane and evaporated under vacuum. The crude solid was used directly for the next step without further purification.

Fmoc-S-2-(2’-pentyl)alanine, (5)

To a solution of 4 (0.1 gm, 0.64 mmol) in water (8 ml), DIEA (0.167 ml, 0.96 mmol) was added followed by the addition of Fmoc-OSu (0.237gm, 0.7 mmol) in acetonitrile (8 ml). The resultant mixture was stirred at room temperature for 5-6 h. The solvent was evaporated and the resultant solution was dissolved in water and subsequently acidified with 2N HCl solution. The aqueous layer was extracted with ethylacetate for 3 times and purified by silica column chromatography using hexane/DCM and later MeOH/DCM as eluent. The product elutes with 3% MeOH/DCM mixture. This gives rise to 0.12 gm of the product with 50% isolated yield. 1H NMR

(CDCl3, 400 MHz): δ ppm 1.24-1.43 (m, 2H), 1.63 (s, 3H), 1.87-1.89 (m, 1H), 2.06-2.16

(m, 3H), 4.22 (t, J = 6.48 Hz, 1H), 4.41(bs, 2H), 4.96-5.03 (m, 2H), 5.52 (bs, 1H), 5.76

(m, 1H), 7.315 (t, J1= J2= 7.4 Hz, 2H), 7.40 (t, J = 7.4 Hz, 2H), 7.6 (d, J = 7.4 Hz, 2H),

7.76 (d, J = 7.48 Hz, 2H), 10.66 (bs, 1H).

5-iodo-1-pentene, (2)

5-Bromo-1-pentene (0.8 ml, 6.71 mmol) was added to a solution of sodium iodide

(2.0 gm, 13.3 mmol) in acetone (22 ml). The reaction mixture was heated at 60 oC for 2 h. The mixture was cooled to room temperature and diluted with water (100 ml) and extracted with pentane 3-times. The pentane layers were combined, washed with brine, dried over sodium sulfate and concentrated to give 5-iodo-1-pentene. The product was

1 obtained as 90% isolated yield (1.2 gm). H NMR (CDCl3, 400 MHz): δ ppm 1.91 (p, J1

= 7.12 Hz, J2 = 7.0 Hz, 2H), 2.16 (q, J1 = 6.7 Hz, J2 = 6.82 Hz, 2H), 3.19 (t, J1 = 6.88 Hz,

J2 = 6.92 Hz, 2H), 5.00-5.10 (dd, J1 = 17.12 Hz, J2 = 10.12 Hz, 2H), 5.69-5.80 (m, 1H).

Similarly Fmoc-S-2-(2’-octyl alanine)alanine, was obtained starting from Z-D-alanine using 8-iodo-1octene. Fmoc-R-2-(2’-pentyl)alanine was obtained starting from Z-L- alanine using similar reaction conditions as described above.

3.4.3.2. Synthesis of N-Fmoc-S-2-(2’-octyl)alanine

(2S,4R)-Benzyl-4-methyl-5-oxo-4-(oct-4-enyl)-2-phenyloxazolidine-3carboxylate, (2S,4R)

1 H NMR (CDCl3, 400 MHz): δ ppm 1.05-1.31 (m, 8H), 1.69 (s, 1H), 1.8 (s, 3H), 2.03-2.2

(m, 3H), 4.94-5.35 (m, 4H), 5.78-5.82 (m, 1H), 6.41 (d, 1H), 6.89 (m, 1H), 7.21-7.42 (m,

10H).

N-Fmoc-S-2-(2’-octyl)alanine

1 H NMR (CDCl3, 400 MHz): δ ppm 1.09-1.37 (m, 8H), 1.66 (s, 3H), 1.88 (bs, 1H), 2.03-

2.18 (m, 3H), 4.25 (s, 1H), 4.43-4.72 (m, 2H), 4.96-5.05 (m, 2H), 5.71 (bs, 1H), 5.78-

5.88 (m, 1H), 7.34 (t, J1 = 7.24, J2 = 7.32 Hz, Hz, 2H), 7.42 (t, J1 = 7.24, J2 = 7.0, 2H),

7.63 (bs, 2H), 7.79 (d, J = 7.44 Hz, 2H), 11.6 (bs, 1H).

8-iodo-1-octene

1 H NMR (CDCl3, 400 MHz): δ ppm 1.25-1.43 (m, 6H), 1.82 (p, J1 = 7.12 Hz, J2 = 7.4

Hz, 2H), 2.04 (q, J1 = 6.76 Hz, J2 = 6.84 Hz, 2H), 3.18 (t, J1 = J2 = 7.04 Hz, 2H), 4.92-

5.02 (m, 2H), 5.74-5.81 (m, 1H).

Peptides 27-28 were synthesized on solid phase by sequentially adding appropriate amino acids along with the peptide coupling reagents (HATU, DIEA) onto the Fmoc-Rink Amide resin76,77. Each amino acid coupling was done for 1.5 h to 2 h.

Both peptides were obtained in 60-80% isolated yields after purification by RP-HPLC and characterized by ESI-MS. All solvents and reagents used were of highest purity available. Fmoc-Asp(tBu)-OH, 1 was purchased from Novabiochem and N,O-dimethyl hydroxylamine hydrochloride was bought from Acros. 1H-NMR spectra were recorded on Bruker 400 spectrometer. Chemical shifts () are reported in ppm downfield from the internal standard (TMS).

Peptide concentrations were tested by absorbance at 280 nm or densitometry measurements for the gel-based activity measurements. A consistent error of the peptide concentration was determined to be no more than two-fold more concentrated than the expected amount. The inhibitory constant, Ki, was adjusted accordingly. However, the ranking of our best peptides were not changed, therefore our overall conclusions remain the same.

3.4.4. Activity Assays

For measurements of caspase activity, 700 nM freshly purified protein was assayed over the course of 10 minutes in a caspase-9 activity assay buffer containing 100 mM MES pH 6.5, 10% PEG 8,000 and 10 mM DTT78. 300 μM fluorogenic substrate,

LEHD-AFC, (N-acetyl-Leu-Glu-His-Asp-AFC (7-amino-4-fluorocoumarin), Enzo

Lifesciences) Ex395/Em505, was added to initiate the reaction. Assays were performed in duplicate at 37°C in 100 μL volumes in 96-well microplate format using a Molecular

Devices Spectramax M5 spectrophotometer. Initial velocities versus inhibitor concentration were fit to a rectangular hyperbola using GraphPad Prism (Graphpad

Software) to calculate Ki app.

Proteolytic cleavage gel-based caspase-9 activity assays were performed for testing the ability of each peptide to inhibit caspase-9 against a natural substrate.

Cleavage of the full-length procaspase-7 variant C186A, which is catalytically inactive and incapable of self-cleavage, was used to report activity. 1 μM full-length procaspase-

7 C186A was incubated with 1 μM active caspase-9 in a minimal assay buffer containing

100 mM MES pH 6.5 and 10 mM DTT in a 37°C water bath for 1 hr. Samples were analyzed by 16% SDS-PAGE to confirm the exact lengths of the cleavage products.

Percent inhibition was calculated using Gene Tools (SynGene) by producing a standard curve from densiometry values for process and unprocessed substrate (caspase-7 C186A).

3.4.5. Mass Spectrometry

Peptide 2 in water was diluted in α-cyano-4-hydroxycinnamic acid matrix to a final concentration of 5 µM. 1µl was spotted onto a target for analysis by Omniflex

MALDI-TOF mass spectrometer (Bruker Daltonics, Inc, Billerica, MA) using linear

detection mode. Peptides 27 and 28 in water were diluted in 0.1% formic acid to a final concentration of 5 µM for direct injection onto Esquire-LC electrospray ion trap mass spectrometer (Bruker Daltonics, Inc.) set up with an ESI source and positive ion polarity.

The MS instrument system was equipped with an HP1100 HPLC system (Hewlett-

Packard). Scanning was carried out between 600-1400 m/z and the final spectra obtained were an average of 10 individual spectra. All mass spectral data were obtained at the

University of Massachusetts Mass Spectrometry Facility, which is supported, in part, by the National Science Foundation.

3.4.6. Secondary Structure Analysis by Circular Dichroism

The secondary structure of the peptide variants was monitored via CD spectra

(250-190 nm) measured on a J-720 circular dichroism spectrometer (Jasco) with a peltier controller. Peptides 2, 3, 5, 8, 27, and 28 in water and Peptides 11, 25 and aPP in 20mM

Tris pH 8.0 were diluted to 15 μM and analyzed at room temperature. Peptides 27 and 28 were monitored for secondary structure formation before and after the metathesis reaction.

3.4.7. Computational Structure Prediction

The structures of aPP variants were predicted computationally using Rosetta57,58.

Folding simulations were performed on the designed peptide sequences 10-26 using the scaffold aPP as a control. The structure of aPP was reasonably well simulated as assessed by backbone alignment and rotomer confirmation of the top 15 lowest scored predicted conformations. Structural visualization and model interrogation was performed in The

PyMOL Molecular Graphics System, Version 1.3, Schrödinger, LLC79.

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93 CHAPTER IV

MECHANISM OF ZINC-MEDIATED INHIBITION OF CASPASE-9

Abstract

Zinc-mediated inhibition is implicated in global caspase regulation, with relief of zinc-mediated inhibition central to both small-molecule and natively induced caspase activation. As an initiator, caspase-9 regulates the upstream stages of the apoptotic caspase cascade, making it a critical control point. Here we identify two distinct zinc- binding sites on caspase-9. The first site, composed of H237, C239, C287 includes the active site dyad and is primarily responsible for zinc-mediated inhibition. The second binding site at C272 plays is distal from the active site. Given the amino-acid conservation in both regions, these sites appear to be conserved across the caspase family.

4.1. Introduction

Apoptotic cell death is carried out by caspases, a family of cysteine proteases.

Caspases function in a cascade of cleavage events leading to orderly destruction of cells, culminating in cell death. Initiator caspase-8 and -9 activate the executioner caspases-3,

-6 and -7, which cleave select targets enabling cell death. Apoptosis is important for both organismal development and homeostasis and is implicated in diseases ranging from ischemic injury to cancer. Understanding the mechanisms of the native regulatory pathways of caspases provides new insights for controlling and harnessing apoptosis.

Exposure of the apoptotic machinery to metal ions has been reported to influence apoptosis. Zinc, a metal commonly used for both biological structure and function, has been highlighted as a specific regulator of apoptosis, where even the smallest fluctuations

in cellular zinc concentrations can tip a cell towards survival or apoptotic cell death1.

Zinc-based regulation of apoptosis had previously been thought to act upstream of the caspases2. However, in vitro and in vivo studies implicate caspases as the targets for zinc- mediated inhibition of apoptosis3,4.

The basis of the necessity for zinc regulation of caspases is not well understood, but may relate to protection of the essential active-site sulfhydryls during oxidative stress

(reviewed in5). Misregulation of zinc and changes in caspase activity have been linked to a number of diseases. Patients with asthma and chronic bronchitis show a correlation between zinc deficiency and increased levels of apoptosis in airway epithelial cells6,7, suggesting a protective role for zinc-mediated caspase inhibition. Heliobacter Pylori based infections have also been linked to zinc-mediated regulation of the caspases8,9, where release of zinc from the bacterium inhibits caspase activity to avoid cell death.

PAC-1, a serendipitous small-molecule caspase activator, relieves zinc-mediated caspase-

3 inhibition10 further suggesting that regulation of zinc-inhibition of caspases may be therapeutically exploited. In this work, we aim to understand the molecular mechanism of zinc-mediated caspase inhibition.

4.2. Results

4.2.1. Metal Affects on the Properties of Caspase-9

Caspase inhibition by zinc has been explored in the executioner caspases -3, -6, and -7, however, little is known about zinc’s function on initiator caspases, including caspase-9. Inhibition of the initiator caspases is particularly critical as the initiators

control the upstream steps of the proteolytic cascade. Previous studies have suggested that zinc disrupts caspase-9 activation and thus activity11,12; however a full analysis of zinc’s affect on caspase-9 or the mechanism of inhibition has not been investigated. We interrogated the effects of a panel of metal cations on the ability of caspase-9 to cleave its natural substrate, caspase-7. Both full-length caspase-9 (Fig 4.1 A) and caspase-9 lacking its pro domain (Caspase Activation and Recruitment Domain, CARD) (Fig 4.1 B) were inhibited by zinc but not by any other cations, including cobalt (Fig 4.1 C). Full inhibition of both caspase-9 variants by zinc indicates that the CARD domain is not involved in zinc’s mechanism of inhibition. Other caspases including caspase-3, -6, -7 and -8 have

Figure 4.1 Zinc exclusively inhibits caspase-9 activity. (A, B) Zinc is the predominant metal cation to inhibit full-length caspase-9 (C9 FL) (A) or the N-terminal CARD-domain deleted caspase-9 (C9 N) (B) when monitored by cleavage of a natural caspase-9 substrate, the caspase-7 zymogen (C7 C186A) to the caspase-7 large (C7 Lg) and small (C7 Sm) subunits in an in vitro cleavage assay monitored by gel mobility. (C) Caspase-9 full-length activity was not inhibited by cobalt or cadmium as judged by cleavage of fluorogenic peptide substrate LEHD-AFC. (D) Competition with metal chelator, EDTA, indicates zinc inhibition is reversible, as monitored by the cleavage of fluorogenic peptide substrate LEHD-AFC.

previously been shown to be inhibited in the presence of zinc3; however other metal cations, including copper also inhibit caspase-3 function4. In contrast, caspase-9 appears to be inhibited specifically by zinc but not by other metals. Zinc-mediated inhibition was fully reversible by the metal chelator EDTA (Fig 4.1 D) suggesting a regulatory role for zinc rather than a non-specific mechanism such as promotion of an irreversible protein aggregate. Inhibition by zinc lead to Michaelis-Menten like curves that fit best to a mixed model of inhibition (Fig 4.2). The observed error within the fit of the individual curves was not due to quenching of the fluorophore, AFC, by zinc (data not shown). This suggests zinc inhibition functions allosterically or binds near the active site thus influencing the binding of substrate. The inhibitory constant, Ki, of 1.5 ± 0.3 μM is similar that reported for zinc inhibition of caspase-3, -6, -7 and -8 (IC50 = 8.8, 0.3, 1.7,

Figure 4.2 Kinetics of full-length caspase-9 wild type in the presence of 0-50 μM ZnCl2. Measured data were fit to competitive, noncompetitive, and mixed models of inhibition. R2 values are presented for individual zinc concentrations and for the overall global fit of the curves fit by each model in which mixed model of inhibition represented the best fit model.

1.9 μM, respectively)3. Although physiological “free” zinc concentrations are reported to be in the femto- to pico-molar range13,14, the “available” zinc pool is believed to be much higher. As a whole, the eukaryotic cell contains approximately 200 μM zinc14 where small shifts in the glutathione concentration or slight oxidative stress causes release of zinc from the metallothioneins15 or vesicles. Thus inhibition constants for caspases in the low micromolar range are likely to be functionally relevant.

Zinc has been shown to influence oligomerization or structure for a variety of proteins including β-2-microglobulin16,17, α-synuclein18, and prion proteins19, so we likewise investigated the effect of zinc binding on the biophysical properties of caspase-

9. Unlike other caspases which are constitutive dimers, caspase-9 is predominantly a monomer in solution, which dimerizes upon binding of substrate20. Dimerization can be observed by native gel analysis (Fig 4.3 A) and size exclusion chromatography (Fig 4.3

B). The presence of zinc does not influence dimerization (Fig

4.3 A, B). Zinc also has no observed effect on the secondary structure of caspase-9 as measured by circular dichrosim spectroscopy (Fig 4.3

Figure 4.3 Zinc does not alter the biophysical properties of caspase- C), further suggesting that zinc- 9. (A, B) Native gel analysis (A) and size exclusion chromatography (B) both indicate zinc does not alter the mediated inhibition does not oligomeric state of caspase-9 from monomer to dimer like the substrate-mimic z-VAD-FMK. (C) CD spectra of caspase-9 full influence the overall structure of length in the presence and absence of ZnCl2 indicate that zinc does not alter the caspase-9 secondary structure.

caspase-9 monomers.

4.2.2. Determining the Location of Zinc Binding

Zinc binding to caspase-9 was assessed by inductively coupled plasma-optical emissions spectroscopy (ICP-OES). Each wild-type caspase-9 monomer was observed to bind two zinc molecules. Caspase-9 monomers contain thirteen cysteine and eight histidine residues (Fig 4.4 A, B), all potential ligands for zinc. Most of these residues are located in the large subunit, either clustered near the active site or at the bottom of the

210’s helix (Fig 4.4 C, D) suggesting the locations of two potential zinc-binding sites.

Figure 4.4 Zinc binding in caspase-9. (A) Caspase-9, monomer, highlights clusters of cys and his metal- binding residues. (B) Cys and his residues are mainly located in the large subunit of caspase-9. (C) Cys- His clusters are conserved throughout the caspase family. (D) Location of the conserved active-site and 210’s helix exosite-ligand clusters on caspase-9 (pdb ID 1JXQ).

The active site cluster comprises the conserved catalytic residues C287 and H237, in proximity to two cysteines that are unique to caspase-9, C172 and C239. Interestingly, the

210’s helix region contains a conserved cysteine-histidine-cysteine triad of unknown function, comprising residues 272, 224, 230 respectively. Furthermore, both the active site region and the triad are highly conserved in the apoptotic class of caspases (Fig 4.4

C).

We interrogated five positions within Table 4.1 Zinc bincing as monitored by ICP-OES. these two cys-his clusters in caspase-9 for involvement in zinc binding (Table 4.1).

Whereas two zinc molecules bind to one monomer of wild-type caspase-9, caspase-9 variants which substitute the active-site residues

C287A or H237A bind just one molecule of zinc. This suggests that the active-site region is one of the zinc binding sites. The two most likely proximal zinc-binding ligands were also substituted. C172A had little effect on zinc binding, but C239S lead to a more substantial loss of zinc binding. The

C287A/C239S variant also lost binding of one molecule of zinc, suggesting that the active site zinc binding cluster comprises residues C239, H237 and C287. Although the identity at residue C239 is not conserved across caspases, it is Glu in all the executioner caspases, suggesting that this residue might be a conserved zinc ligand position across the apoptotic caspase family.

Substitution of the residue C272A to alanine in the conserved 210’s-helix triad,

100

also resulted in loss of one zinc molecule (Table 4.1), suggesting that C272 is at the core of the second zinc-binding site. The loss of zinc binding at C272A was not due to unfolding of the protein as the substitution C272A has no effect on basal activity of the enzyme. The catalytic efficiency of C272A is nearly the same as wild-type caspase-9, 2.0

± 0.16 and 2.7 ± 0.25 respectively. Given the proximity, we suggest that the second binding site comprises residues H224, C230 and C272. Together these data suggest that there are two distinct zinc-binding sites in caspase-9 (Fig 4.4 D). The presence of two independent binding sites was confirmed by caspase-9 C287A/C272A variant in which zinc binding is ablated.

Binding of zinc to both the catalytic residues of caspase-9 is expected to block nucleophilic attack of the peptide bond by C287 rendering the enzyme inactive. The potential contribution of the second binding site to inhibition is less obvious. If the C272 site contributes directly to zinc-mediated caspase-9 inhibition, then the caspase-9 C272A variant C272A might be expected to show a competitive mechanism of inhibition.

Surprisingly, the C272A variant also shows a mixed model of inhibition (Fig 4.5), like wild-type caspase-9 and exhibited a comparable Ki value of 5.0 ± 2.8 μM. The Ki for zinc with the C272A variant is statically indistinguishable from that observed with wild type enzyme (Student’s t-test), suggesting this site does not contribute appreciably to the mechanism of caspase-9 inhibition by zinc.

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Figure 4.5 Kinetics of full-length caspase-9 C272A variant in the presence of 0-50 μM ZnCl2. Measured data were fit to competitive, noncompetitive, and mixed models of inhibition. R2 values are presented for individual zinc concentrations and for the overall global fit of the curves fit by each model in which mixed model of inhibition represented the best fit model. 4.3. Discussion

Although no regulatory function was observed in the case of the caspase-9 second zinc binding site, this 210’s helix region (the 90’s helix in caspase-6) has been implicated in the mechanism of caspase-6 activation21. A 21° pivot about the bottom of this helix has been observed. The open conformation is adopted when caspase-6 is in an inactive state whereas the closed conformation is characteristic of the active enzyme. In light of the fact

3,22 that zinc inhibition is most potent in caspase-6 (IC50 = 0.3 μM ) , and that this structural region is sensitive and critical for caspase-6 activity, it is tempting to hypothesize that zinc could control caspase-6 allosterically, via binding to this caspase conserved exosite triad.

Our findings indicate that the reversible binding of zinc to the caspase-9 active

102

site residues H237, C239, and

C287 is the predominant mechanism for caspase-9 inhibition. A plausible model with reasonable metal-to-ligand bond distances between active- site residues and zinc was obtained simply by changing the rotomeric conformations of

H237, C239, and C287 (Fig Figure 4.6 A model of caspase-9 active-site ligand interactions with a modeled zinc ion. Model was obtained by altering the H237, C239 and C287 rotomers. 4.6). Like the majority of zinc- binding sites in proteins, the binding site is likely to be four-coordinate, tetrahedral. The fourth ligand could be water, solvent, or potentially a nearby glutamate residue, E290

(Fig 4.6). E290 resides on a flexible loop just 3.8Å from the proposed position of the zinc ion. Optimal E290 ligand-binding distances may be achieved by loop movement or by a water-mediated interaction.

Taken together, these results suggest a model for how zinc binds to the active site and inhibits caspase-9 and caspases generally. The function of the secondary zinc-binding exosite remains an open question; however the conservation of this site suggests that perhaps zinc does play a physiological role at this site as well. The fact that zinc does not function in a strictly competitive manner with substrate at the active site, may suggest some flexibility in the zinc-binding ligands. Given that the zinc-mediated inhibition

103

constants for all the caspases are similar and that ligands are conserved in the active site region, suggests that the mechanism of active-site inhibition of the apoptotic caspases is conserved. Thus, utilization of zinc-based inhibition of caspases, like that used by PAC-

110, remains a promising avenue for controlling caspases and apoptosis.

4.4. Materials and Methods

4.4.1. Caspase-9 Expression and Purification

The caspase-9 full-length gene (human sequence) construct, encoding amino acids

1-416, in pET23b (Addgene plasmid 1182923) was transformed into the BL21 (DE3) T7

Express strain of E. coli (NEB). The cultures were grown in 2xYT media with ampicillin

(100 mg/L, Sigma-Aldrich) at 37°C until they reached an optical density at 600 nm of

1.2. The temperature was reduced to 15°C and cells were induced with 1 mM IPTG

(Anatrace) to express soluble 6xHis-tagged full-length protein. Cells were harvested after 3 hrs to obtain single site processing at Asp315. Cell pellets stored at -20°C were freeze-thawed and lysed in a microfluidizer (Microfluidics, Inc.) in 50 mM sodium phosphate pH 8.0, 300 mM NaCl, and 2 mM imidazole. Lysed cells were centrifuged at

17,000 rpm to remove cellular debris. The filtered supernatant was loaded onto a 5-ml

HiTrap Ni-affinity column (GE Healthcare). The column was washed with a buffer containing 50 mM sodium phosphate pH 8.0, 300 mM NaCl, 2 mM imidazole until 280 nm absorbance returned to base line. The protein was eluted using a linear imidazole gradient of 2 to100 mM over the course of 270 mL. The eluted fractions containing protein of the expected molecular weight and composition were diluted by 10-fold into a buffer composed of 20 mM Tris pH 8.5, 10 mM DTT to reduce the salt concentration.

104

This protein sample was loaded onto a 5-ml Macro-Prep High Q column (Bio-Rad

Laboratories, Inc.). The column was developed with a linear NaCl gradient and eluted in

20 mM Tris pH 8.5, 100 mM NaCl, and 10 mM DTT buffer. The eluted protein was stored in -80°C in the above buffer conditions. The identity of the purified caspase-9 was analyzed by SDS-PAGE and ESI-MS to confirm mass and purity. Caspase-9 variants in the full-length expression construct (C287A, H237A, C172A, C239S, C287A/C239S,

C272A, C272A/C287A) were purified by the same method as described here for the wild-type protein.

Caspase-9 ΔCARD was expressed from a two-plasmid expression system. Two separate constructs, one encoding the large subunit, residues 140-305, and the other encoding the small subunit, residues 331-416, each in the pRSET plasmid, were separately transformed into the BL21 (DE3) T7 Express strain of E. coli (NEB). The recombinant large and small subunits were individually expressed as inclusion bodies for subsequent reconstitution. Cultures were grown in 2xYT media with ampicillin (100 mg/L, Sigma-Aldrich) at 37°C until they reached an optical density at 600 nm of 0.6.

Protein expression was induced with 0.2 mM IPTG. Cells were harvested after 3 hrs at

37°C. Cell pellets stored at -20°C were freeze-thawed and lysed in a microfluidizer

(Microfluidics, Inc.) in 10 mM Tris pH 8.0 and 1 mM EDTA. Inclusion body pellets were washed twice in 100 mM Tris pH 8.0, 1 mM EDTA, 0.5 M NaCl, 2% Triton, and 1M urea, twice in 100 mM Tris pH 8.0, 1 mM EDTA and finally resuspended in 6 M guanidine hydrochloride. Caspase-9 large and small subunit proteins in guanidine hydrochloride were combined in a ratio of 1:2, large:small subunits, and rapidly diluted

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dropwise into refolding buffer composed of 100 mM Tris pH 8.0, 10% sucrose, 0.1%

CHAPS, 0.15 M NaCl, and 10 mM DTT, allowed to stir for one hour at room temperature and then dialyzed four times against 10 mM Tris pH 8.5, 10 mM DTT, and

0.1mM EDTA buffer at 4°C. The dialyzed protein was centrifuged for 15 minutes at

10,000 rpm to remove precipitate and then purified using a HiTrap Q HP ion exchange column (GE Healthcare) with a linear gradient from 0 to 250 mM NaCl in 20 mM Tris buffer pH 8.5, with 10 mM DTT. Protein eluted in 20 mM Tris pH 8.5, 100 mM NaCl, and 10 mM DTT buffer was stored in -80°C. The identity of the purified caspase-

9ΔCARD was analyzed by SDS-PAGE and ESI-MS to confirm mass and purity.

4.4.2. Prediction of Metal Ligands and Construction of Ligand Substitution Variants

Potential zinc-binding ligands were predicted via the computational server which predicts metal ion-binding sites, HotPatch version 4.0 , by a computational program specific for zinc binding sites, PREDZINC version 1.1 , and by visual inspection of zinc ligand clusters and distances based on the caspase-9ΔCARD crystal structure (pdb ID:

1JXQ) via the PyMOL Molecular Graphics System, Version 1.3 (Schrödinger, LLC).

Sites predicted as potential zinc ligands were C172, C239, C272, H237, and C287.

Individual active-site knockout variants, H237A and C287A, and cysteine knockout variants, C172A, C239S, and C272A, in the caspase-9 full-length gene were made by a single round of QuikChange site directed mutagenesis (Stratagene). Double knockout variants, C239S/C287A and C272A/C287A, were made by an additional round of

QuikChange site directed mutagenesis in the background of the caspase-9 full-length

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C287A variant. Formation of the proper variant was confirmed by sequence analysis

(Genewiz, Inc.).

4.4.3. Caspase-7 C186A Expression and Purification

The active site knockout variant, C186A, was made by QuikChange mutagenesis

(Stratagene) of the human caspase-7 gene in pET23b (gift of Guy Salvesen ) The construct was transformed into BL21 (DE3) T7 Express strain of E. coli and protein expression was induced with 1 mM IPTG at 18°C for 18 hrs. The protein was purified as described previously for caspase-7. The eluted protein was stored in -80°C in the buffer in which they eluted. The identity of purified caspase-7 C186A was assessed by SDS-

PAGE and ESI-MS to confirm mass and purity.

4.4.4. Activity Assays

For measurements of caspase activity, proteolytic cleavage gel-based caspase-9 activity assays were performed for testing the ability of a panel of metal cations to inhibit caspase-9 activity against a natural substrate. Cleavage of the full-length procaspase-7 variant C186A, which is catalytically inactive and incapable of self-cleavage, was used to report caspase-9 activity. 1μM caspase-9 full-length in a minimal assay buffer containing

100 mM MES pH 6.5 and 10 mM DTT was incubated in the presence of 1mM metal cation for 1hr at room temperature. 1 μM full-length procaspase-7 C186A was added to the reaction mix and incubated in a 37°C water bath for 1 hr. Samples were analyzed by

16% SDS-PAGE to confirm the exact lengths of the cleavage products.

To test the reversibility of zinc binding, 700 nM freshly purified caspase-9 full- length was incubated in the presence and absence of 5 equivalents excess ZnCl2 for 1hr at

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room temperature. Half of each protein sample was treated with 25-fold excess EDTA just prior to fluorescence activity assay measurements. Samples were assayed for activity over the course of 10 minutes in a caspase-9 activity assay buffer containing 100 mM

MES pH 6.5, 10% PEG 8,000 and 10 mM DTT. 300 μM fluorogenic substrate, LEHD-

AFC, (N-acetyl-Leu-Glu-His-Asp-7-amino-4-fluorocoumarin), (Enzo Lifesciences)

Ex395/Em505, was added to initiate the reaction. Assays were performed in duplicate at

37°C in 100 μL volumes in 96-well microplate format using a Molecular Devices

Spectramax M5 spectrophotometer.

The inhibitory constant for zinc, Ki, was determined by incubating 700 nM caspase-9 full-length protein, diluted in 100 mM MES pH 6.5, 10% PEG 8,000, and 5

mM DTT, in the presence of 0-50 μM ZnCl2 for 1.5 hours at room temperature.

Samples from each ZnCl2 concentration were subjected to a substrate titration, performed in the range of 0-300 μM fluorogenic substrate, LEHD-AFC (Ex365/Em495) which was added to initiate the reaction. Assays were performed in duplicates at 37°C in 100 μL volumes in 96-well microplate format using a Molecular Devices Spectramax M5 spectrophotometer. Initial velocities versus substrate concentration were globally fit to competitive, noncompetitive and mixed models of inhibition using GraphPad Prism

(Graphpad Software) to determine inhibitory constant, Ki. Determination for the type of inhibition was based on four criteria. R2 values for the individual curve fits, criterion for the α-value reported by the mixed model of inhibition, overall visual inspection of the curve fits with the data and knowledge of the system were all taken into consideration.

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The R2 values and visual inspection of the curve fits with the data ruled out competitive inhibition whereas these criteria were less clear between the noncompetitive and mixed models of inhibition. However, a slight improvement in the R2 values for the individual curve fits for the mixed model of inhibition was observed, suggesting that this is the best fit for the data. The α-value obtained from the mixed model of inhibition fits also reports on the inhibition mechanism. If the α-value is equal to one, the mixed model of inhibition is identical to the non-competitive model whereas if this value is very large, the model becomes identical to the competitive model of inhibition. The reported α-value for the fits of both full-length caspase-9 wild-type and variant C272A match neither of these criteria, therefore the mechanism of inhibition is appears to follow a mixed model.

Furthermore, based on our enzymatic system and the determined location of metal binding, a mixed model of inhibition appears to be the most likely.

4.4.5. Oligomeric-State Determination

Caspase-9 wild-type, full-length protein samples in 20 mM Tris pH 8.5, 110 mM

NaCl, and 5 mM DTT were incubated alone (monomer), with covalent inhibitor z-VAD-

FMK (carbobenzoxy-Val-Ala-Asp-fluoromethylkeytone) (dimer) or with 5 equivalents excess of ZnCl2 for 2 hours at room temperature. The oligomeric state of the caspase-9 samples were determined via gel filtration and native gel analysis. 100 μL of 0.75 mg/ml protein sample was loaded onto a Superdex 200 10/300 GL (GE Healthcare) gel-filtration column. Apo and z-VAD-FMK incubated protein samples were eluted with 20 mM Tris pH 8.0, 100 mM NaCl, and 2 mM DTT while ZnCl2-incubated protein was eluted in the same buffer and one containing 50 μM ZnCl2 to ensure zinc was not be lost during the gel

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filtration process. Eluted peaks were analyzed by SDS-PAGE to ensure protein identity.

Four different molecular weight standards from the gel-filtration calibration kit LMW

(GE Healthcare) were run in the same conditions and a standard plot was generated to determine whether the peaks were caspase-9 monomer or dimer. For native gel analysis, samples were mixed with glycerol loading dye and fractionated on a 10% Tris/Glycine pH 8.3 polyacrylamide gel. The oligomeric state of the ZnCl2 treated caspase-9 was identified by comparison to the Apo (monomer) and z-VAD-FMK (dimer) caspase-9 full- length samples.

4.4.6. Secondary Structure Analysis by Circular Dichroism

Caspase-9 full-length protein was buffer exchanged via dialysis against 100 mM sodium phosphate pH 7.0, 110 mM NaCl, and 2 mM DTT, and diluted to 7 μM. The sample was split in half and incubated in the presence or absence of five equivalents of

ZnCl2 for 1hr at room temperature. The circular dichroism spectra of caspase-9 full- length in the presence and absence of zinc was monitored from 250-190 nm, measured on a J-720 circular dichroism spectrometer (Jasco) with a peltier controller.

4.4.7. Zinc Binding Analysis by ICP-OES

Purified caspase-9 full-length, wild-type and variants (15 μM) in 20 mM Tris pH

8.5, 110 mM NaCl, and 5 mM DTT were incubated with 5 equivalents excess of ZnCl2 for 1 hour at room temperature. Unbound zinc was removed by incubating samples of 1 mL volume in the presence of approximately 125 mg of Chelex® 100 for 1 hour, mixing every 20 minutes. A control sample of 1 mL of 20 mM Tris pH 8.5, 110 mM NaCl, and 5 mM DTT buffer only was treated in the exact same manner to judge the completeness of

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unbound zinc chelation. Wild-type caspase-9 full-length samples either treated with zinc or untreated were also used as controls for metal content pre and post zinc incubation.

Zinc content of the sample supernatant was quantified using a PerkinElmer Optima 4300

DV inductively coupled plasma-optical emission spectrometer (ICP-OES) equipped with a 40 MHz free-running generator and a segmented-array charge-coupled device (SCD) detector and a sample introduction system consisted of a concentric nebulizer with a cyclonic spray chamber. The concentration of zinc in each sample was determined at

206.2 nm and converted from ppm to μM to obtain the zinc to monomer of caspase-9 full-length ratio. Post ICP-OES analysis, remaining samples were tested via absorbance at 280 nm determine protein concentration. A 16% SDS-PAGE gel visualized with the

Pierce Silver Stain Kit (Thermo Scientific) was used to determine the percentage of low concentration contaminants so the total caspase-9 concentration could be adjusted accordingly. This adjustment did not alter the overall trend observed.

4.4.8. Model of Zinc Binding to Caspase-9

A proposed model for zinc binding at the active site of caspase-9 ΔCARD was developed in PyMOL Molecular Graphics System, Version 1.3 (Schrödinger, LLC). The zinc ionic radius was contoured to the reported values for a tetrahedral geometry of ligand binding. The rotomeric conformations of the ligands which coordinate the zinc ion, H237, C239, and C287, as well as a putative ligand, E290, were sampled to obtain the best metal to ligand geometry and allowed bond distances for ligand character.

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4.5. Structure Determination Trials of Caspase-9 and Zinc

4.5.1. Crystallization of Caspase-9 in the Presence and Absence of Zinc

To further understand the molecular details of zinc-mediated inhibition at the active site as well as zinc binding at the secondary binding site we aimed to obtain the crystal structure of caspase-9 full-length, wild-type protein in the presence of zinc.

Previous caspase-9 structures have shown a dimeric caspase-9 enzyme without the N- terminal CARD domain (ΔCARD) in complex with covalent peptide inhibitor z-VAD-

FMK (PDB ID: 1JXQ)20, a variant which shifts the enzymes oligomeric state from monomer to dimer by re-engineering the dimerization interface to mimic caspase-3 (PDB

ID: 2AR9)24, and monomeric caspase-9 ΔCARD in complex with the protein inhibitor

XIAP-Bir3 (PDB ID: 1NW9)25. Crystals from dimeric caspase-9 enzyme lacking the N- terminal CARD domain (ΔCARD) in complex with a covalent peptide inhibitor grew in a

0.1 M MES pH 6.0 and 12% PEG 20,000 solution when incubated at 20 °C. The sitting- drop vapor diffusion method was utilized for crystal growth. On the other hand, the dimeric caspase-9 variant, (residues 140–416) made by altering the dimeric interface to mimic that of caspase-3 and knocking out the catalytic Cys (C287S), crystallized in grown in 5% (w/v) PEG 5,000 monomethylether, 0.1M MES pH 6.0 and 3% tacsimate when incubated at 17 °C utilizing the hanging drop vapor diffusion method. Crystals of the caspase-9/XIAP-Bir3 inhibitory complex were grown also grown using this method where drops were set up in 0.1 M Tris pH 8.0, 1.0 M potassium monohydrogen phosphate, and 0.2 M sodium chloride. These three conditions served as a starting point for initial screening for caspase-9 crystals in the presence of zinc.

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Purified wild type caspase-9 full-length protein (residues 1-416) in 20 mM Tris pH 8.5, 100 mM NaCl, and 5 mM DTT was concentrated to 11 mg/mL using an Amicon

Ultracell 3K concentrator (Millipore). The concentrated protein was then incubated with five equivalents excess of ZnCl2 for one hour at room temperature and filtered using a durapore-PVDF 0.22 μM filter (Millipore) to remove any dust, debris or protein precipitate prior to crystallization trials. Initial crystallization trials were set up using the hanging drop, vapor diffusion method in all known conditions previously described.

However upon drop set-up, immediate protein precipitate was observed and no crystal formation was seen over time. Therefore, naïve crystallization screens were tested with 8 to 20 mg/mL of the full-length form of caspase-9 wild-type protein pre-incubated with zinc. No crystal formation was observed in the naïve screens when using Crystallization

Screen I & II, Index Screen, and JCSG+ Suite from Hampton Research. Crystallization trials of the apo wild-type caspase-9 in the full-length form were then pursued using the known crystallization conditions using 11 mg/mL protein. Initial drops result in a white cloudy precipitate in the drop, however, crystal growth was observed after 24 hours in the condition 0.1 M MES pH 6.0 and 12% PEG 20,000 in a 1:1 protein-to-mother-liquor ratio incubated at 20 °C. The crystals obtained formed clusters of thin plates measuring

730 x 260 x 50 μM in size, which were unusable for data collection (Fig 4.7 A).

Further optimization was performed to obtain single crystals of caspase-9.

Optimizations included varying the pH between 5.2 to 6.0, the identity of the PEG precipitant from PEG 200 to 20,000 average molecular weight and the concentration of

PEG from 10 to 20%. Final conditions were 0.1 M MES pH 5.8 and 14% PEG 6,000,

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Figure 4.7 Crystals of wild type caspase-9 full-length (A) pre and (B) post optimization.

resulting in thin single crystals of approximately 100 x 40 x 10 μM in size and form in the shape of a half of a hexagon or a full hexagon in some instances (Fig 4.7 B).

Microseeding, macroseeding, and additive screen experiments in addition to further purifying the protein via a HiLoad 26/600 Superdex gel filtration column in 20 mM Tris pH 8.5, 100 mM NaCl, and 2 mM DTT were used to improve consistency of the crystal form within a drop and overall thickness and size of crystals. Unfortunately these approaches did not result in a significant improvement in crystal morphology or growth.

4.5.2. Data Collection on Crystals of Caspase-9

Crystals of full length caspase-9 wild-type apo protein in final mother liquor conditions of 0.1 M MES pH 6.0 and 15% PEG 6,000 were incubated in 20% glycerol, ethylene glycol or PEG 400 as a cryoprotectant for one hour and flash frozen in liquid nitrogen for storage. During crystal screening at a variety of different angles at the

Brookhaven National Laboratory synchrotron x-ray light source, the caspase-9 wild-type

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crystals diffracted to approximately 3.0 Å, with nice rounded spot shape and minimal mosaicity, but a relatively weak intensity (Fig 4.8). This indicates that optimal crystallization and freezing conditions were obtained. Indexing the diffraction images was performed with ease, and indicated that the crystals showed C2 symmetry, similar to that of the previously reported N-terminal CARD domain (ΔCARD) in complex with covalent peptide inhibitor z-VAD-FMK (PDB ID: 1JXQ)20.

Figure 4.8 Diffraction image of apo wild type caspase-9 crystals soaked in a 20% PEG 400 cryoprotectant for one hour.

4.5.3. Zinc Soaks of caspase-9 Crystals

Therefore, these conditions were used for further experimentation. In order to obtain a structure of caspase-9 in the presence of zinc, full-length caspase-9 wild-type crystals needed to be soaked in mother liquor containing the zinc ion. The 0.1 M MES pH

5.8 and 15% PEG 6,000 mother liquor was prepared to include 10 mM ZnCl2. Crystals

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were transferred from their growth drops to a 3 μl drop of zinc containing mother liquor.

Soaking of the crystals lasted for the duration of 15 minutes, 30 minutes, 1 hour, 2 hours,

3 hours, 5 hours, or 18 hours. Crystals appeared to remain intact during the entire soaking process as no microfractures were observed microscopically. The zinc-soaked crystals were then either preserved in cryoprotectant containing zinc to ensure the bound zinc ions are not lost during the cryoprotectant incubation time due to the on/off rate of zinc to the binding sites or backsoaked in cryoprotectant to ensure excess zinc binding to random free thiols did not occur. Cryoprotectant incubation times lasted anywhere from 15 minutes to overnight. Crystals were preserved by flash freezing in liquid nitrogen. Data collection was carried out at Brookhaven National Labs Synchrotron x-ray Light Source beamline X6A on the caspase-9 wild type crystals in the presence of zinc. These crystals did diffract, however, the diffraction was limited to approximately 7Å (Fig 4.9). In addition, within the diffraction image, a smeary/mosaic spot shape was observed as well as an anisotropic diffraction pattern. These characteristics provide clues as to the changes in conformation that may have occurred within the caspase-9 crystal upon binding zinc.

The observed smeary spot shape, suggests increased mosaicity in the crystal form, which could not be corrected by annealing. In addition the diffraction images themselves indicate that crystals are directionally dependent or anisotropic in nature.

Based on the quality of the diffraction images for the apo caspase-9 crystals, conditions and methods for soaking of the caspase-9 crystal in zinc soak should be reevaluated. Alternate metal salts such as zinc sulfate and zinc acetate which also inhibit enzymatic function could be explored. Furthermore, soaking methodology such as

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Figure 4.9 Diffraction image of wild type caspase-9 soaked in ZnCl2. Crystals were soaked for 2 hours and a 20% ethylene glycol for ten minutes. the zinc concentration and time could also improve diffraction of zinc bound caspase-9 crystal.

References:

1. Zalewski, P. D., Forbes, I. J. & Betts, W. H. Correlation of apoptosis with change in intracellular labile Zn(II) using zinquin [(2-methyl-8-p-toluenesulphonamido-6- quinolyloxy)acetic acid], a new specific fluorescent probe for Zn(II). Biochem J 296 ( Pt 2), 403-408, (1993).

2. Wolf, C. M., Morana, S. J. & Eastman, A. Zinc inhibits apoptosis upstream of ICE/CED-3 proteases rather than at the level of an endonuclease. Cell Death Differ 4, 125-129, (1997).

3. Stennicke, H. R. & Salvesen, G. S. Biochemical characteristics of caspases-3, -6, - 7, and -8. J Biol Chem 272, 25719-25723, (1997).

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4. Perry, D. K. et al. Zinc is a potent inhibitor of the apoptotic protease, caspase-3. A novel target for zinc in the inhibition of apoptosis. J Biol Chem 272, 18530- 18533, (1997).

5. Truong-Tran, A. Q., Carter, J., Ruffin, R. E. & Zalewski, P. D. The role of zinc in caspase activation and apoptotic cell death. Biometals 14, 315-330, (2001).

6. Carter, J. E. et al. Involvement of redox events in caspase activation in zinc- depleted airway epithelial cells. Biochem Biophys Res Commun 297, 1062-1070, (2002).

7. Truong-Tran, A. Q., Grosser, D., Ruffin, R. E., Murgia, C. & Zalewski, P. D. Apoptosis in the normal and inflamed airway epithelium: role of zinc in epithelial protection and procaspase-3 regulation. Biochem Pharmacol 66, 1459-1468, (2003).

8. Kohler, J. E. et al. Monochloramine-induced toxicity and dysregulation of intracellular Zn2+ in parietal cells of rabbit gastric glands. Am J Physiol Gastrointest Liver Physiol 299, G170-178, (2010).

9. Kohler, J. E. et al. Monochloramine impairs caspase-3 through thiol oxidation and Zn2+ release. J Surg Res 153, 121-127, (2009).

10. Peterson, Q. P. et al. PAC-1 activates procaspase-3 in vitro through relief of zinc- mediated inhibition. J Mol Biol 388, 144-158, (2009).

11. Chimienti, F., Seve, M., Richard, S., Mathieu, J. & Favier, A. Role of cellular zinc in programmed cell death: temporal relationship between zinc depletion, activation of caspases, and cleavage of Sp family transcription factors. Biochem Pharmacol 62, 51-62, (2001).

12. Schrantz, N. et al. Zinc-mediated regulation of caspases activity: dose-dependent inhibition or activation of caspase-3 in the human Burkitt lymphoma B cells (Ramos). Cell Death Differ 8, 152-161, (2001).

13. Bozym, R. A., Thompson, R. B., Stoddard, A. K. & Fierke, C. A. Measuring picomolar intracellular exchangeable zinc in PC-12 cells using a ratiometric fluorescence biosensor. ACS Chem Biol 1, 103-111, (2006).

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14. Krezel, A. & Maret, W. Zinc-buffering capacity of a eukaryotic cell at physiological pZn. J Biol Inorg Chem 11, 1049-1062, (2006).

15. Krezel, A., Hao, Q. & Maret, W. The zinc/thiolate redox biochemistry of metallothionein and the control of zinc ion fluctuations in cell signaling. Arch Biochem Biophys 463, 188-200, (2007).

16. Eakin, C. M., Knight, J. D., Morgan, C. J., Gelfand, M. A. & Miranker, A. D. Formation of a copper specific binding site in non-native states of beta-2- microglobulin. Biochemistry 41, 10646-10656, (2002).

17. Morgan, C. J., Gelfand, M., Atreya, C. & Miranker, A. D. Kidney dialysis- associated amyloidosis: a molecular role for copper in fiber formation. J Mol Biol 309, 339-345, (2001).

18. Goers, J., Uversky, V. N. & Fink, A. L. Polycation-induced oligomerization and accelerated fibrillation of human alpha-synuclein in vitro. Protein Sci 12, 702- 707, (2003).

19. Millhauser, G. L. Copper binding in the prion protein. Acc Chem Res 37, 79-85, (2004).

20. Renatus, M., Stennicke, H. R., Scott, F. L., Liddington, R. C. & Salvesen, G. S. Dimer formation drives the activation of the cell death protease caspase 9. Proc Natl Acad Sci U S A 98, 14250-14255, (2001).

21. Vaidya, S., Velazquez-Delgado, E. M., Abbruzzese, G. & Hardy, J. A. Substrate- induced conformational changes occur in all cleaved forms of caspase-6. J Mol Biol 406, 75-91, (2011).

22. Takahashi, A. et al. Cleavage of lamin A by Mch2 alpha but not CPP32: multiple interleukin 1 beta-converting enzyme-related proteases with distinct substrate recognition properties are active in apoptosis. Proc Natl Acad Sci U S A 93, 8395- 8400, (1996).

23. Stennicke, H. R. et al. Caspase-9 can be activated without proteolytic processing. J Biol Chem 274, 8359-8362, (1999).

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24. Chao, Y. et al. Engineering a dimeric caspase-9: a re-evaluation of the induced proximity model for caspase activation. PLoS Biol 3, e183, (2005).

25. Shiozaki, E. N. et al. Mechanism of XIAP-mediated inhibition of caspase-9. Mol Cell 11, 519-527, (2003).

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CHAPTER V

CASPASE-9 CARD:CORE DOMAIN INTERACTIONS REQUIRE A

PROPERLY-FORMED ACTIVE SITE

Abstract

The activation mechanism of caspase-9 is unique in that the addition of individual domains or when bound to the large protein complex, the apoptosome, has a great affect on enzymatic activity. The N-terminal prodomain, CARD, of caspase-9 is the smallest unit capable of increasing caspase-9 activity. We investigated CARD with respect to the activation, oligomerization and stability of caspase-9. Our data suggest an increase in dimerization due to the presence of CARD is not the cause of the increased activity observed. We determined that caspase-9 undergoes conformational acrobatics by adopting one conformation in the uncleaved, monomeric form and cleaved active dimeric state where the active site is in an ordered state, while adopting another in the cleaved monomeric form in the presence of a disordered active site.

5.1. Introduction

Caspase-9 is as a critical initiator of the intrinsic pathway of apoptosis or programmed cell death. It is responsible for activating downstream executioner caspases-

3, -6 and -7 which ultimately results in the breakdown of the cell. Defects in this pathway are a characteristic of diseases ranging from autoimmune disorders to cancer. In these diseases, activation of caspase-9 is particularly critical1-3, however, a full mechanistic understanding of caspase-9 activation is lacking.

In general, caspases, or cysteine aspartate proteases, are functional as homodimers of monomeric units that comprise an N-terminal prodomain and a catalytic large and

121 small subunit connected by an intersubunit linker. Upon dimerization and cleaveage at a specific aspartate residue within the intersubuint linker, the enzyme becomes active.

Unlike the executioner caspases, caspase-9 is active even in the absence of cleavage of the intersubunit linker4. Furthermore, the executioner caspases have short N-terminal prodomains whereas caspase-9’s catalytic core is preceded by a long regulatory domain called the Caspase Activation and Recruitment Domain (CARD). The caspase-9 CARD facilitates a protein-protein interaction with the CARD of the apoptotic protease activation factor 1(Apaf-1), which anchors caspase-9 to the activation platform known as the apoptosome5-7. This platform is created upon release of cytochrome c from the mitochondria following an internal cellular stress signal. Cytochrome c is then able to bind to Apaf-1. This binding event, initiates a conformational change in Apaf-1, resulting in the heptameric oligomerization of the protein, driven in an ATP-dependent manner.

This Apaf-1 platform is then responsible for the recruitment and activation of caspase-9.

The molecular details of caspase-9 activation by the apoptosome have not been elucidated.

The interaction of caspase-9 with the apoptosome increases caspase-9’s activity by approximately 2000-fold4. However, even in the absence of this platform, the presence of individual domains influence caspase-9 activity. Caspase-9 is 20% more active when caspase-9’s catalytic core remains covalently linked to the CARD domain8 (Fig 5.1). The authors hypothesized that increased tangling of CARD domains lead to an increase in the dimeric fraction of full-length caspase-9 over ΔCARD-caspase-9. Addition of the Apaf-1

CARD in isolation further enhances caspase-9 activity by five-fold, in vitro. The greatest increase in activity is when full-length caspase-9, including both the CARD and core

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Figure 5.1 Model depicting the increase in enzymatic activity of caspase-9. Activity is lowest for the monomeric unit of the catalytic core (C9ΔCARD) and increases to the highest when full-length caspase-9 is bound to the apoptosome. domains, bind to the apoptosome through Caspase-9 CARD: Apoptosome CARD interactions.

Early models of the apoptosome activation of caspase-9 argued that the increased activity is due to a change in the oligomeric state of the enzyme via increasing the local concentration of monomeric caspase-9. Recruitment of additional molecules were hypothesized to participate as partners in dimerization9 or facilitate dimerization amongst the apoptosome-bound monomers10. This model is supported by the evidence that enhanced activity is associated with dimerized capase-9 molecules11. Alternative, competeting views of the activation mechanism invoke induced conformational changes around the active site region of the enzyme12. This conformational change model is potentially supported by the new evidence emerging from high-resolution cryo electron microscopy that shows that caspase-9 is monomeric when bound to the apoptosome in the highly active state and that caspase-9 activation can be achieved without apoptosome formation13.

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Taking inspiration from the proposed mechanisms of capase-9 activation by the apoptosome we want to further investigate the affect of CARD with respect to the activation, oligomerization and stability of caspase-9. A goal in this work is to understand whether CARD influences the oligomeric state of caspase-9, specifically the ratio of monomer to dimer in vitro, thus causing the observed increase in activity. We probe interactions between the caspase-9 CARD and core domains. Finally we investigate whether the caspase-9 CARD induces or takes advantage of conformational changes in the loops that form the active site region as has been observed for caspase-6 and -714,15. In caspase-6 and -7 conformational changes in the active-site loops can be observed by both thermal denaturation and circular dichroism spectral measurements. Defining CARD’s role in the activation of caspase-9 will provide much needed insight for exploiting caspase-9 for therapeutic means.

5.2. Results

5.2.1. The Influence of CARD on the Oligomeric State and Stability of Caspase-9

Activity measurements of caspase-9’s catalytic core (caspase-9 ΔCARD), in the absence of the apoptosome platform, have been previously observed to have a lower catalytic efficiently when compared to the full-length version of the enzyme. We interrogated the affect the presence of the CARD has on these specific biophysical properties of the capase-9 enzyme. To ensure that the caspase-9 ΔCARD and full-length reagents used for this study were comparable to the studies that show CARD’s ability to enhance caspase-9 activity, measurements of the enzymes catalytic properties were performed (Table 5.1). As previously reported, the ΔCARD variant of caspase-9 has a decreased catalytic efficiency when compared to the full-length version. Therefore we

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Table 5.1 Measured kinetic parameters of full-length caspase-9 (C9FL) and caspase-9 in the absence of the N-terminal CARD domain (C9ΔCARD). reasoned that any differences observed between the two caspase-9 variants would be due to the presence or absence of the CARD domain.

One potential reason for the increased in activity of full-length caspase-9 would be due to an altered oligomeric state of the enzyme in the presence of CARD as predicted previously8. Full-length and ΔCARD versions of the caspase-9 enzyme were subjected to size exclusion chromatography to determine the oligomeric state of both enzymes (Fig

5.2). Both enzymes are predominantly monomeric in solution to such a great extent that no dimeric fraction could be observed. Both the full-length and ΔCARD versions of caspase-9 are both capable of completely converting to the dimeric state as observed when the enzyme is in the presence of an active-site ligand such as the covalent active- site inhibitor z-VAD-FMK. Thus the CARD does not have a great influence on the

Figure 5.2 Size exclusion chromatography of full-length caspase-9 (C9F) and caspase-9 ΔCARD (C9 ΔCARD) in the presence and absence of active site ligand z-VAD-FMK.

125 oligomeric state of the enzyme and cannot be the cause of the increased activity observed in the presence of CARD.

Another potential reason for the increased activity of caspase-9 full-length is an increased in stability in the presence of the CARD. Thermal denaturation studies were then pursued to determine if the presence of CARD changes the thermal stability of the enzyme and thus the reason for increased activity. Both the full-length and ΔCARD caspase-9 enzymes in their monomeric and dimeric forms were analyzed for changes in their thermal stability measurements. Full-length capase-9, which has been cleaved at the intersubunit linker between the large and small subunits, and includes the

CARD domain, shows a three-state unfolding curve (Fig 5.3 A). The first melting transition occurs at 49 ± 2 °C while the second occurs at 62 ± 2 °C.

To determine which component of the full-length caspase-9 enzyme was contributing to which melting transition, the catalytic core and CARD domains were expressed independently and interrogated in a similar fashion.

The catalytic core of the enzyme

(caspase-9ΔCARD), which is also Figure 5.3 Thermal denaturation analysis and circular dichroism spectrum of monomeric, cleaved, caspase- cleaved at the intersubunit linker, is 9 (A) full-length, (B) ΔCARD, and (C) CARD only.

126 responsible for the first melting transition at 49 ± 1 °C (Fig 5.3 B) while the second transition corresponds to CARD only with a melting temperature of 61 ± 2 °C (Fig 5.3

C). These results indicate that when caspase-9 is in its cleaved monomeric state, the presence of CARD does not affect the overall thermal stability of the caspase-9 catalytic core and the two portions of the enzyme unfold independently. Caspase-9 can be forced to a dimeric state by binding of active-site ligand. Thermal denaturation studies were performed on the cleaved full-length and ΔCARD versions of caspase-9 when the enzyme is in a dimeric state with active site ligand bound (Fig 5.4). Both versions of caspase-9 show an increase in thermal stability. Caspase-9ΔCARD (Fig 5.4 A) has a

14°C increase in thermal stability which is similar to the increase in stability observed

Figure 5.4 Thermal denaturation analysis and circular dichroism spectrum of dimeric, cleaved, caspase-9 (A) full-length, (B) ΔCARD.

when caspase-7 binds active site ligand15, while full-length caspase-9 (Fig 5.4 B) shows only a 6 °C increase in thermal stability, more similar to the 3 °C increase in stability observed for caspase-6 upon ligand bidning14. Strikingly, the three-state unfolding of the full-length caspase-9 is no longer observed in the presence of substrate. It appears that binding ligand to the active site of caspase-9, which induces dimerization and ordering of the active site loop bundle, transitions caspase-9 to a two-state unfolding mechanism.

127

Full-length caspase-9 is completely unfolded at 90 °C as observed in the circular dichroism spectrum. The single transition observed when cleaved full-length caspase-9 in the dimeric state indicates that binding substrate and dimerization either results in the complete unfolding of CARD or causes the catalytic core and CARD to unfold as one cooperative unit. An overlay of the circular dichorism spectra from the full-length cleaved caspase-9 in the monomeric and dimeric states (Fig 5.5) indicates that there is not a change in the secondary structure content.

Therefore, CARD does not appear to unfold in the presence of substrate so the CARD and the caspase-9 catalytic core must be unfolding as a single cooperative unit. These results suggest that a physical interaction between the Figure 5.5 Overlay of the circular dichroism spectrum for full-length caspase-9 (C9F) in CARD and core domains occurs, which the presence and absence of active site ligand z-VAD-FMK causes the two domains to unfold as a single unit.

It has been reported that uncleaved caspase-9 acts as an active zymogen 4 possibly due to its increased intersubunit linker length which enables the enzyme to support a properly formed active site. Therefore, this form of caspase-9 can be utilized to interrogate whether the interaction observed between CARD and the catalytic core of caspase-9 is due to changes in the active site conformation. Analysis of the full-length monomeric caspase-9 in the uncleaved state, where the complete polypeptide chain is still intact, resulted in the similar two-state unfolding mechanism as observed for the cleaved active dimeric state (Fig 5.6). Therefore, the monomeric, uncleaved caspase-9 enzyme

128 appears to support the interaction of the core and CARD domains because they unfold as a single unit.

To further test this mechanism, we cleaved the same zymogen construct with caspase-3 (Fig 5.7).

Cleavage of the intersubunit linker breaks the interaction of the CARD and catalytic core domains observed Figure 5.6 Thermal denaturation analysis and circular by the independent three-state dichroism spectrum of (A) monomeric, uncleaved caspase-9 and (B) monomeric, cleaved caspase-9. unfolding property as seen in the cleaved wild-type enzyme (Fig 5.6 B, 5.7). Together, these data imply that the CARD domain and the catalytic core of caspase-9 do not physically interact and unfold independently in monomeric caspase-9 with a disordered active site, whereas, these domains physically interact and unfold cooperatively, as a single unit, in the uncleaved monomeric and cleaved dimeric states where the active site Figure 5.7 SDS-PAGE analysis of full-length is in an ordered state. uncleaved caspase-9 (C9F-C287A) in the presence and absence of 3% active caspase-3 enzyme (C3). Full length caspase-9 remains uncleaved due to Ala substitution of the catalytic residue Cys287.

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5.2.2. Determining an Interaction Site Between Caspase-9 Catalytic Core and CARD Domains.

The cooperative unfolding observed between the CARD domain and the catalytic core of dimeric caspase-9 implies a physical interaction between the two domains. To confirm this interaction, the interaction between the isolated CARD and catalytic core domains in trans was interrogated (Fig 5.8). The catalytic core in its monomeric or dimeric forms was incubated with the CARD domain and analyzed by native gel analysis for an interaction between the two domains (Fig 5.8 A). An interaction between the two domains, would result in a band the molecular weight of the full-length enzyme by native gel analysis. The monomeric and dimeric forms of full-length caspase-9 served as reliable controls, but no interaction between the catalytic core and CARD domain was observed. Further confirmation of this result was observed via Ni-NTA pull-down analysis (Fig 5.8 B) where the bound 6x His-tagged CARD domain was not able to pull- down the untagged catalytic core. In both gel-filtration and native gel analysis, the CARD was not observed to have a strong interaction with the catalytic core of caspase-9

Figure 5.8 Analysis of a CARD: caspase-9 core domain interactions in trans. Interactions analyzed by by (A) native gel analysis, (B) Ni-NTA pull-down, and (C) activity assays using fluorogenic substrate LEHD-AFC. All experiments were performed in trans. Caspase-9 full-length (C9F); Caspase-9ΔCARD (C9Δ). These analyses all suggest that the interaction between these domain is too weak or too transient for detection.

130 suggesting that the interaction between the CARD and core domains is either too weak or too transient for detection by these methods or is exquisitely sensitive to buffer conditions.

It is possible that the conditions and experimental requirements for the native gel and pull-down analysis could disrupt the CARD and core interaction, so CARD’s ability to increase caspase-9 activity in trans was tested under conditions in which activity can be monitored (Fig 5.8 C). This equilibrium-based experiment should report on the strength of the interaction between the CARD and catalytic core domains and define the importance of the linker that attaches the CARD and core domains together. No change in caspase-9 ΔCARD activity was observed in the presence of excess CARD alone or in the presence of excess lysozyme as a crowding agent. This indicates that the tether between CARD and the catalytic core of caspase-9 is necessary for CARD’s impact on enzymatic activity.

Since the linker between the CARD and core domains is essential to increase the activity of caspase-9, we reasoned that perhaps there were either specific interactions with the tether and the adjacent domains or a length-dependence to the interaction. To test this, a five amino acid Ser-Gly extension was inserted within the linker between CARD Figure 5.9 Characteristics of the Ser-Gly linker extension caspase-9 variant. Kinetic parameters, and the large subunit of caspase-9’s thermal denaturation analysis, and circular dichroism spectra of the Ser-Gly linker extension catalytic core (Fig 5.9). This variant caspase-9 variant.

131 behaves like the native full-length-form of caspase-9 in both its catalytic parameters and thermal stability suggesting that longer potentially more flexible linker does not negatively impact the function of caspase-9. Further investigation into the linker length would be required to determine if a certain distance or flexibility can disrupt the activation property of CARD or if a tether alone is sufficient to obtain the desired effect no matter the length. Moreover, it is intriguing to consider, shortening the linker length between the CARD and core domains to determine if restricting the flexibility of CARD will reduce the activation effect of CARD. This will provide information on the shortest length required for the interaction to occur.

The previous studies suggest that the interaction between the CARD and core domains of caspase-9 are present, but are weak and potentially transient. Docking studies between reported crystal structures of the CARD and the dimeric form of the caspase-9 catalytic core were performed using the RosettaDock server16 to predict potential interactions between the CARD and core domain to guide mutational studies. Protein structure files for CARD (PDB ID: 3YGS17) and the catalytic core of caspase-9 (PDB ID:

1JXQ11) were submitted for docking studies. The top docking models were analyzed using two criteria i) avoiding interactions with residues involved in the caspase-9

CARD/Apaf-1 CARD complex and ii) not allowing negative interactions to occur in the presence of Apaf-1 CARD. Two models, Model 2 and Model 10, fit these criteria (Fig

5.10). In both, the CARD is positioned behind the substrate-binding groove where it could potentially affect active site loops L3 and L4 and there are no negative interactions observed. Charge swap variations were made in both Model 2, which places CARD in a helix-to-helix interaction with the catalytic core of caspase-9 (Fig 5.10 A), and Model 10,

132

Figure 5.10 Top RosettaDock models of the potential interaction site between CARD and the catalytic core of caspase-9. (A) Model 2 provided a helix-to-helix mode of binding while (B) Model 10 provides a loop-to-helix mode of binding.

which places CARD in a loop-to-helix interaction with the catalytic core of caspase-9

(Fig 5.10 B), in an attempt to disrupt the activating effect of CARD. Single variations of

Arg7 and Arg11 and in combination, both changed to glutamate, would test Model 2 while Asp23 and Arg51 charge swapped to glutamate would test Model 10. These variations were selected to be in the CARD domain only so the observed effect would report on CARD and not on change the catalytic activity, which is solely due to the catalytic portion of the enzyme. One variant, Glu365Arg, was selected in the catalytic core which had potential to disrupt either model of CARD binding to the catalytic core of caspase-9.

We expected that mutations that disrupted the interaction between the CARD and core domain would decrease the catalytic efficiency of the variant versions of caspase-9.

Upon analysis of the charge swap affects on the catalytic efficiency of caspase-9, a two- fold decrease in catalytic efficiency was observed for variant R7E (Table 5.2) which would support Model 2, the helix-to-helix binding mode of CARD to caspase-9. A similar affect was observed by the single site variant R11E which further supports a helix-to-helix binding model of CARD and the catalytic core of caspase-9. However the

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Table 5.2 Measured kinetic parameters of caspase-9 variants designed to disrupt the activation affect of the CARD domain. double site variant of R7E/R11E did not show an enhancement of this affect (Table 5.2).

This could indicate that R7 and R11E are included in the activating affect of CARD; however, the combination of both these variants is not strong enough to completely eliminate the activation property of CARD, as the catalytic efficiency of the ΔCARD version of the enzyme was not recapitulated. This suggests that this region of the CARD domain is responsible for its activating effects however additional interactions are required.

5.3. Discussion

Caspase activation mechanisms are relevant for treatment of diseases in which apoptosis has gone awry. Caspase-9 in particular has a unique activation mechanism including changes in its conformational and oligomeric state and its association with an activation platform called the apoptosome. Furthermore, individual domains which are linked to the enzyme such as CARD or which are associated with the caspase-9 activation mechanism, such as the Apaf-1 CARD also have the ability to increase enzymatic

134 activity. The property of changing enzymatic activity by presence of additional domains such as the one observed with CARD and caspase-9 is also relevant in other proteins such as PAS Kinase18, Dnmt1 DNA methyltransferase19, and ADAMTS-420. Therefore studying the individual activation effects of a particular domain provides further insights towards how caspase-9 becomes activated on the apoptosome.

Here we have investigated the effect of the caspase-9 CARD domain with respect to its ability to increase enzymatic activity. We have determined that the oligomeric state of both caspase-9 full-length and ΔCARD were comparable with respect to the presence of the monomeric and dimeric caspase-9 states, indicating the presence of CARD does not shift the oligomeric state equilibrium from monomer to dimer. Thus, the presence of

CARD is not the cause for the increased enzymatic activity observed. Furthermore, we observed that the CARD and catalytic core domains of caspase-9 unfold independently and do not physically interact in the monomeric state of the cleaved enzyme where the active site is not able to form a properly ordered substrate-binding groove. However, these domains unfold cooperatively, or as a single unit, in a dimeric state indicating a physical interaction when the cleaved enzyme dimerizes and has a properly ordered active site. Moreover, cooperative unfolding of CARD and the catalytic core of caspase-9 is also observed in the uncleaved form of full-length caspase-9 zymogen. Intriguingly this

CARD-core domain interaction is alleviated by cleavage of the intersubunit linker by caspase-3. Thus, CARD appears to be interacting with any version of caspase-9 presenting a properly formed substrate-binding groove. The substrate-binding groove is ordered in dimeric cleaved caspases (-1, -3, -6, -7 and -9) and can also be formed in uncleaved zymogen of caspase-9, due to linkage effects which allow the intersubunit

135 linker to buttress the L3 and L4 loops in an ordered conformation (Fig 5.11). Shortening of the intersubunit linker would show the length requirement necessary this stabilizing effect.

This alteration in conformational state of CARD observed between the cleaved inactive monomer and the uncleaved monomer or cleaved active dimer states is reminiscent of the conformational acrobatics observed in caspase-6. In this case, a helical conformation is observed in the mature enzyme which converts to a β-strand conformation in the active state. Furthermore, the uncleaved zymogen of caspase-6 adopts a similar β-strand as observed in the active state of the enzyme. Our data suggest caspase-9 follows a similar trend to caspase-6 by adopting one conformation in the uncleaved, monomeric form and cleaved active dimeric state while adopting another in the cleaved monomeric state (Fig. 5.11).Therefore, it appears that the most unique regions within the caspase class of enzymes, the pro domain and intersubunit linker may combine efforts to regulate the activity of caspase-9, providing evidence for the active site stabilizing model of apoptosome activation.

Figure 5.11 Model of caspase-9 activation states in the presence of CARD.

136

Further studies on the interconversion properties of the caspase-9 catalytic core and CARD domains, utilizing a preformed caspase-9 dimer21, could provide insight into the dependence of the oligomeric state on the cooperative unfolding property of the full- length enzyme. Single charge swap mutations on the surface of the protein distal from the active site were not strong enough or properly positioned to disrupt the activating affect of CARD. A more extensive alanine scanning mutagenesis study or charge repulsion analysis around the substrate-binding groove could further define the region of interaction between the CARD and core domains mediated by the active-site region of the enzyme.

5.4. Materials and Methods

5.4.1. Caspase-9 Expression and Purification

The caspase-9 full-length gene (human sequence) construct, encoding amino acids

1-416, in pET23b (Addgene plasmid 118294) was transformed into the BL21 (DE3) T7

Express strain of E. coli (NEB). The cultures were grown in 2xYT media supplemented with ampicillin (100 mg/L, Sigma-Aldrich) at 37°C until they reached an optical density at 600 nm of 1.2. The temperature was reduced to 15°C and cells were induced with 1 mM IPTG (Anatrace) to express soluble 6xHis-tagged full-length protein. Cells were harvested after 3 hrs to obtain single-site processing at Asp315. Cell pellets stored at -

20°C were freeze-thawed and lysed in a microfluidizer (Microfluidics, Inc.) in 50 mM sodium phosphate pH 8.0, 300 mM NaCl, and 2 mM imidazole. Lysed cells were centrifuged at 17,000 rpm to remove cellular debris. The filtered supernatant was loaded onto a 5-ml HiTrap Ni-affinity column (GE Healthcare). The column was washed with a buffer containing 50 mM sodium phosphate pH 8.0, 300 mM NaCl, and 2 mM imidazole

137 until 280 nm absorbance returned to base line. The protein was eluted using a linear imidazole gradient of 2 to100 mM over the course of 270 mL. The eluted fractions containing protein of the expected molecular weight and composition were diluted by 10- fold into a buffer composed of 20 mM Tris pH 8.5, 10 mM DTT to reduce the salt concentration. This protein sample was loaded onto a 5-ml Macro-Prep High Q column

(Bio-Rad Laboratories, Inc.). The column was developed with a linear NaCl gradient and eluted in 20 mM Tris pH 8.5, 100 mM NaCl, and 10 mM DTT buffer. The eluted protein was stored in -80°C in the above buffer conditions. Purified caspase-9 was analyzed by

SDS-PAGE and ESI-MS to confirm mass and purity. Caspase-9 variants, C287A, R7E,

R11E, D23E, R51E E365R, R7E/R11E and the Ser-Gly linker extension, were produced by the Quikchange mutagenesis method (Stratagene), in the full-length expression construct, and were purified by the same method as described here for the wild-type protein.

Caspase-9ΔCARD was expressed from a two-plasmid expression system. Two separate constructs, one encoding the large subunit, residues 140-305, and the other encoding the small subunit, residues 331-416, each in the pRSET plasmid, were separately transformed into the BL21 (DE3) T7 Express strain of E. coli (NEB). The recombinant large and small subunits were individually expressed as inclusion bodies for subsequent reconstitution. Cultures were grown in 2xYT media supplemented with ampicillin (100 mg/L, Sigma-Aldrich) at 37°C until they reached an optical density at

600 nm of 0.6. Protein expression was induced with 0.2 mM IPTG. Cells were harvested after 3 hrs at 37°C. Cell pellets stored at -20°C were freeze-thawed and lysed in a microfluidizer (Microfluidics, Inc.) in 10 mM Tris pH 8.0 and 1 mM EDTA. Inclusion

138 body pellets were washed twice in 100 mM Tris pH 8.0, 1 mM EDTA, 0.5 M NaCl, 2%

Triton, and 1M urea, twice in 100 mM Tris pH 8.0, 1 mM EDTA and finally resuspended in 6 M guanidine hydrochloride. Caspase-9 large and small subunit proteins in guanidine hydrochloride were combined in a ratio of 1:2, large:small subunits, and rapidly diluted dropwise into refolding buffer composed of 100 mM Tris pH 8.0, 10% sucrose, 0.1%

CHAPS, 0.15 M NaCl, and 10 mM DTT, allowed to stir for one hour at room temperature and then dialyzed four times against 10 mM Tris pH 8.5, 10 mM DTT, and

0.1mM EDTA buffer at 4°C. Typically 5 mL of of mixed caspase large and small subunits was diluted into 80 mL in refolding buffer and dialyzed against 5 L of dialysis buffer. The first and last dialysis steps were allowed to proceed for 4 hours at 4 °C while the second dialysis proceeded overnight at 4 °C. The dialyzed protein was centrifuged for

15 minutes at 10,000 rpm to remove precipitate and then purified using a HiTrap Q HP ion exchange column (GE Healthcare) with a linear gradient from 0 to 250 mM NaCl in

20 mM Tris buffer pH 8.5, with 10 mM DTT. Protein eluted in 20 mM Tris pH 8.5, 100 mM NaCl, and 10 mM DTT buffer was stored in -80°C. The identity of the purified caspase-9ΔCARD was analyzed by SDS-PAGE and ESI-MS to confirm mass and purity.

5.4.2. Oligomeric-State Determination

Caspase-9 wild-type, full-length and ΔCARD variant protein samples in 20 mM

Tris pH 8.5, 110 mM NaCl, and 5 mM DTT were incubated alone (monomer) or with covalent inhibitor z-VAD-FMK (carbobenzoxy-Val-Ala-Asp-fluoromethylketone, Enzo

Lifesciences) (dimer) for 2 hours at room temperature. The oligomeric state of the caspase-9 samples were determined via gel filtration. 100 μL of 0.5 mg/ml protein sample was loaded onto a Superdex 200 10/300 GL (GE Healthcare) gel-filtration column. Apo

139 and z-VAD-FMK-incubated protein samples were eluted with 20 mM Tris pH 8.0, 100 mM NaCl, and 2 mM DTT. Eluted peaks were analyzed by SDS-PAGE to identify the eluted protein. Four different molecular weight standards from the gel-filtration calibration kit LMW (GE Healthcare) were run in the same conditions and a standard plot was generated to determine whether the peaks were caspase-9 monomer or dimer.

5.4.3. CARD Expression and Purification

The CARD only construct (amino acids 1-138) in pET23b was made by

QuikChange mutagenesis (Stratagene) using the oligo-nucleotide primer 5`-CCCAGA-

CCAGTGGACATTGGTTCTGGAGGATTCGGTGATCACCACCACCACCACCAC

TAAGTCGGTGCTCTTGAGAGTTTGAGGGGAAATGCAGATTTGG-3`and its reverse compliment on the caspase-9 full-length gene (Addgene plasmid 11829). These oligo-nucleotide primers insert a 6xHis-tag and a stop codon after the last amino acid of the CARD domain (D138), leaving the remaining portion of the caspase-9 gene in the plasmid. The construct was transformed into BL21 (DE3) T7 Express strain of E. coli.

The cultures were grown in 2xYT media supplemented with ampicillin (100 mg/L,

Sigma-Aldrich) at 37°C until they reached an optical density at 600 nm of 0.6. The temperature was reduced to 15°C and cells were induced with 1 mM IPTG (Anatrace) to express soluble 6xHis-tagged full-length protein. Cells were harvested after 18 hrs. Cell pellets stored at -20°C were freeze-thawed and lysed in a microfluidizer (Microfluidics,

Inc.) in 50 mM sodium phosphate pH 8.0, 300 mM NaCl, and 2 mM imidazole. Lysed cells were centrifuged at 17,000 rpm to remove cellular debris. The filtered supernatant was loaded onto a 5-ml HiTrap Ni-affinity column (GE Healthcare). The column was washed with a buffer containing 50 mM sodium phosphate pH 8.0, 300 mM NaCl, 2 mM

140 imidazole until 280 nm absorbance returned to base line. The column was washed with

50 mM phosphate pH 8.0, 300 mM NaCl, 50 mM imidazole and the protein was eluted with 50 mM phosphate pH 8.0, 300 mM NaCl, 250 mM imidazole. The eluted fraction was diluted by 10-fold into a buffer containing 20 mM Tris pH 8.0 and 2 mM DTT to reduce the salt concentration. This protein sample was loaded onto a 5 ml Macro-Prep

High Q column (Bio-Rad Laboratories, Inc.). The column was developed with a linear

NaCl gradient. Protein eluted in 20 mM Tris pH 8.0, 2 mM DTT, and 130 mM NaCl.

Eluted protein was analyzed by SDS-PAGE to assess purity and stored in -80°C.

5.4.4. Thermal Stability and Secondary Structure Analysis by Circular Dichroism

Caspase-9 variants and CARD proteins were buffer exchanged via dialysis against

100 mM sodium phosphate pH 7.0, 110 mM NaCl, and 5 mM TCEP and diluted to 7 μM.

The samples were split in half and incubated in the presence or absence of four molar equivalents of active site ligand VAD-FMK for 3 hours at room temperature. To ensure complete binding of the active site ligand to the protein, remaining enzymatic activity was assayed using 300 μM substrate, LEHD-AFC (N-acetyl-Leu-Glu-His-Asp-7-amino-

4-fluorocoumarin), (Enzo Lifesciences). Once full inhibition was achieved, samples were buffer-exchanged six times with 100 mM phosphate buffer pH 7.0, 100 mM NaCl and 5 mM TCEP using an Amicon Ultracell 3K concentrator (Millipore) to remove unbound inhibitor. For cleavage of the unprocessed caspase-9 C287A variant, the 7μM protein sample was incubated with 3% active caspase-3 protein for two hours at room temperature. Full processing of caspase-9 C287A by caspase-3 was determined by SDS-

PAGE analysis. Thermal denaturation of caspase-9 variants and CARD was monitored by loss of CD signal at 222 nm over a range of 20–90 °C. The circular dichroism spectra

141 were monitored from 250-190 nm. Both were performed on a J-720 CD spectrometer

(Jasco) with a Peltier controller. Data were collected four separate times on different days from different batches of purified proteins. Curves were fit with Origin Software

(OriginLab) using sigmoid fit to determine the melting temperature.

5.4.5. Caspase-3 Expression and Purification

Caspase-3 full-length gene (human sequence) in pET23b (Addgene plasmid

1182122) was transformed into BL21 (DE3) T7 Express strain of E. coli and protein expression was induced with 1 mM IPTG at 30°C for 3 23. The protein was purified as described previously for caspase-3 24. The eluted protein was stored in -80°C in the buffer in which they eluted. The identity of purified caspase-3 was assessed by SDS-PAGE and

ESI-MS to confirm mass and purity.

5.4.6. Native Gel Analysis and Ni-NTA Pull Down Assay to Determine in trans Interactions

For native gel analysis to diagnose an interaction between caspase-9ΔCARD and caspase-9 CARD in trans, full-length caspase-9, caspase-9 ΔCARD and the CARD domain only were dialyzed twice against 100 mM phosphate pH 7.0 and 2 mM DTT for

90 minutes to rid of excess salt. Samples were incubated either alone or combined with

CARD to achieve a 1:1 ratio of caspase-9ΔCARD plus CARD. Each protein sample was diluted to a final concentration 10 μM in the dialysis buffer. To induce dimerization samples were incubated with 5-fold excess z-VAD-FMK. All samples were allowed to incubate at room temperature for one hour. All samples were mixed with glycerol loading dye and fractionated on a 10% Tris/Glycine pH 8.3 polyacrylamide gel. The oligomeric state of the mixed caspase-9ΔCARD and CARD samples were identified by comparison

142 to the Apo (monomer) and z-VAD-FMK (dimer) of both the caspase-9 full-length and caspase-9ΔCARD protein in addition to the CARD only sample.

For Ni-NTA pull-down analysis of caspase-9ΔCARD plus CARD in trans, samples were diluted to 10 μM in 100 mM phosphate pH 8.0, 5 mM TCEP and 100 mM

NaCl, to avoid nonspecific interactions of the caspase-9 enzyme with the Ni-NTA beads.

Samples were incubated either alone or combined with CARD to achieve a 1:1 ratio of caspase-9ΔCARD (no tag) plus CARD (6x His-tag). Each protein sample was diluted to a final concentration of 10 μM in the dialysis buffer. To induce dimerization samples were incubated with 5-fold excess z-VAD-FMK for 15 minutes. 200 μL of protein sample was added to a tube containing 50 μL of Ni-NTA beads (Qiagen) that were washed three times in water and twice in buffer. Ni-NTA plus caspase-9ΔCARD and CARD samples were incubated for 3 hours at 4°C while rocking. Samples were centrifuged for 5 minutes at 13,000 rpm to pellet the Ni-NTA beads. Supernatant was aspirated and the beads were washed five times with 100 mM phosphate pH 8.0, 5 mM TCEP and 100 mM NaCl to remove any unbound or weakly bound protein. Protein elution was then carried out by incubating the Ni-NTA beads with phosphate buffer containing 300 mM imidazole for 10 minutes at room temperature. Samples were centrifuged for 5 minutes at 13,000 rpm to pellet the Ni-NTA beads and the supernatant was used for SDS-PAGE analysis. The Ni-

NTA beads were also tested to ensure there were no remaining proteins bound to the resin.

5.4.7. Activity Assays

For kinetic measurements of caspase activity, 700 nM caspase-9 full-length protein was diluted in 100 mM MES pH 6.5, 10% PEG 8,000, and 5 mM DTT. Each

143 sample was subjected to a substrate titration, performed in the range of 0-300 μM fluorogenic substrate, LEHD-AFC, (Ex365/Em495) which was added to initiate the reaction. Assays were performed in duplicates at 37°C in 100 μL volumes in 96-well microplate format using a Molecular Devices Spectramax M5 spectrophotometer. Initial velocities versus substrate concentration were fit to a rectangular hyperbola using

GraphPad Prism (Graphpad Software) to determine kinetic parameters Km and kcat.

Enzyme concentrations were determined by active site titration with quantitative, inhibitor z-VAD-FMK. Active site titrations were incubated over a period of 3 h in 100 mM MES pH 6.5, 10% PEG 8,000, and 5 mM DTT. Optimal labeling was observed when protein was added to VAD-FMK solvated in dimethylsulfoxide in 96-well V- bottom plates, sealed with tape, and incubated at room temperature in a final volume of

200 μL. 90 μL aliquots were transferred to black-well plates in duplicate and assayed with 300 μM substrate. The protein concentration was determined to be the lowest concentration at which full inhibition was observed.

To test the ability of CARD to activate caspase-9ΔCARD in trans, 700 nM freshly purified caspase-9ΔCARD was incubated in the presence and absence of 2-, 10-, or 20-fold excess of CARD protein for 15 minutes. Half of each protein sample was treated with excess lysozyme as a crowding agent. Samples were assayed for activity over the course of 10 minutes in a caspase-9 activity assay buffer containing 100 mM

MES pH 6.5, 10% PEG 8,000 and 10 mM DTT. 300 μM of the fluorogenic substrate,

LEHD-AFC (Ex365/Em495), was added to initiate the reaction. Assays were performed in duplicate at 37°C in 100 μL volumes in 96-well microplate format using a Molecular

Devices Spectramax M5 spectrophotometer.

144

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14. Vaidya, S. & Hardy, J. A. Caspase-6 latent state stability relies on helical propensity. Biochemistry 50, 3282-3287, (2011).

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15. Witkowski, W. A. & Hardy, J. A. L2' loop is critical for caspase-7 active site formation. Protein Sci 18, 1459-1468, (2009).

16. Lyskov, S. & Gray, J. J. The RosettaDock server for local protein-protein docking. Nucleic Acids Res 36, W233-238, (2008).

17. Qin, H. et al. Structural basis of procaspase-9 recruitment by the apoptotic protease-activating factor 1. Nature 399, 549-557, (1999).

18. Rutter, J., Michnoff, C. H., Harper, S. M., Gardner, K. H. & McKnight, S. L. PAS kinase: an evolutionarily conserved PAS domain-regulated serine/threonine kinase. Proc Natl Acad Sci U S A 98, 8991-8996, (2001).

19. Fatemi, M., Hermann, A., Pradhan, S. & Jeltsch, A. The activity of the murine DNA methyltransferase Dnmt1 is controlled by interaction of the catalytic domain with the N-terminal part of the enzyme leading to an allosteric activation of the enzyme after binding to methylated DNA. J Mol Biol 309, 1189-1199, (2001).

20. Kashiwagi, M. et al. Altered proteolytic activities of ADAMTS-4 expressed by C- terminal processing. J Biol Chem 279, 10109-10119, (2004).

21. Chao, Y. et al. Engineering a dimeric caspase-9: a re-evaluation of the induced proximity model for caspase activation. PLoS Biol 3, e183, (2005).

22. Zhou, Q. et al. Target protease specificity of the viral serpin CrmA. Analysis of five caspases. J Biol Chem 272, 7797-7800, (1997).

23. Stennicke, H. R. & Salvesen, G. S. Caspases: preparation and characterization. Methods 17, 313-319, (1999).

24. Stennicke, H. R. & Salvesen, G. S. Biochemical characteristics of caspases-3, -6, - 7, and -8. J Biol Chem 272, 25719-25723, (1997).

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CHAPTER VI

A SURFACE-WIDE VIEW OF THE

NATIVE REGULATORY MECHANSIMS OF CASPASE-9

6.1. Regulation of Caspase-9 Occurs at a Variety of Locations on its Surface

Control of cellular processes plays an essential role in maintaining the homeostasis of a cell. This control is maintained by a complex network composed of specific regulatory mechanisms for each individual protein within the cell. These checkpoints monitor a variety of different protein characteristics ranging from their initial synthesis to their final activation states and even their degradation pathways. In order to fully understand how to externally regulate a particular protein, a fundamental understanding of the native regulatory pathways and their interactions within those native pathways is required.

Protein regulatory mechanisms can be categorized into a variety of different classes. Regulation can occur by altering the genetic expression levels and timing of activation of a particular protein or enzyme. This allows the cell to control not only when a protein will be produced but the quantity of protein production, resulting in an increased versatility during the lifespan of the cell or organism. Post protein production, sequestration or compartmentalization of the protein from its functional partner(s) is utilized to control a protein function. In the case of proteases, processing of the zymogen form of an enzyme is frequently required to initiate its activation pathway. Alternate means of regulation also include post-translational modifications or even allosteric regulation. Through the utilization of all or a variety of these regulatory checkpoints, a tight control of a protein’s function can be obtained. Ideally, each point within these

147 native regulatory pathways can be targeted to create tools to further study enzymatic function or for therapies in cases where there has been disruption of the regulation of a particular protein.

This dissertation focused on discovering new regulatory pathways for caspase-9 as well as exploiting a known regulatory pathway of the enzyme due to its pivotal role in apoptosis and disease. Our work aimed to mimic the native inhibitory complex of caspase-9 with XIAP-BIR3 by stabilized peptides to create a tool to control enzymatic activity. Instead, this work underscored the importance of bi-functional binding sites.

Critical interactions at the protein-protein interface as well as at a distal exosite region provide two distinct avenues for protein control. In the case of caspase-9 and XIAP-

BIR3, both sites are necessary for potent inhibition. In other protein:inhibitor complexes this bi-functionality determines specificity or can individually be used as independent regulatory sites. The viral inhibitor, human adenovirus type 5, of human granzyme B, a protein that is essential for targeting cell death of natural killer cells and cytotoxic CD8+

T-cells, utilizes exosite interactions as a specificity factor1. Common homologues of human granzyme B are capable of recognizing human adenovirus type 5 in the substrate binding pocket, however, the exosite was critical for specific inhibition of the human enzyme. A similar exosite requirement has also been observed with plasminogen activator inhibitor-1 which prevents conversion of plasminogen into plasmin through its interactions with both tissue-type plasminogen activator and urokinase-type plasminogen2. Furthermore, when designing blockades to prevent heparin cofactor II binding to α-thrombin, targeting of the exosite position was an essential requirement for specificity and affinity3,4. These examples not only suggest that the presence of an exosite

148 is a common feature within protein complexes; they also provide an alternate avenue for designing regulatory molecules.

In addition to underscoring the importance of targeting the multiple regulatory regions within a native protein complex, our work has also provided insights towards alternate regulatory sites within caspase-9. When determining the regulatory mechanism of zinc-mediated inhibition of caspase-9, not only did we discover that one zinc ion binds to the catalytic region of the protein, an alternate zinc binding site also exists. The main regulatory site, at the active site, sequesters the active site residues which would prevent catalysis of the peptide bond of a substrate. Besides controlling function, a possible role for active-site zinc binding is to protect the active-site ligands from oxidation via a similar mechanism to that observed for protein tyrosine phosphatase5. The other binding site identified for zinc is distal from the active site and lies within a conserved Cys-His region of the enzyme. Although the functional details of this site remain unidentified, this region has been previously shown to be conformationally sensitive in caspase-66.

Therefore, zinc binding to this site could potentially have a site-specific impact on caspase-6 activity via an allosteric mechanism. Furthermore, the distal zinc binding site could potentially be utilized as a stabilization factor for a particular conformation in caspase-6 or other caspases, similar to that observed in voltage-gated potassium channel7.

The non-native control of protein activity by a metal also occurs in a variety of enzymes which natively do not utilize zinc. Glycerol 3-phosphate dehydrogenase8, tyrosine phosphatase9, aldehyde dehydrogenase10, and the caspases11, are all capable of binding zinc which results in a loss of enzymatic function. This suggests that a wide variety of non-metalloproteins have evolved to be influenced by metals as an underlying mode of

149 regulation and that these sites also exist in locations distal from the active site of these enzymes.

Furthermore, our work in elucidating the mechanism by which CARD activates the enzymatic core of caspase-9 revealed yet another alternate regulatory site on the surface of caspase-9. The mechanism reveals that CARD not only mediates interactions with Apaf-1 of the apoptosome, it potentially facilitates proper organization of the active- site loop bundle and strengthens these interactions resulting in a fully active enzyme. An alternate role for CARD has also been observed where free caspase-9 CARD has the ability to cause activation of NF-κB12. Furthermore, identification of an interaction between the CARD domain of a caspase-9 and the substrate-binding groove provides a unique regulatory mechanism instilled on this cell-death protease that could be targeted for further study. Although this interaction is unique to caspase-9 due to the secondary structural elements involved, globally similar observations have been made with the prodomains of the executioner caspases -3, -6, and -7 and their individual catalytic cores.

The prodomain of caspase-3 has been shown to play a role in determining the proper active-site conformation13, in which the presence of the prodomain enhances the efficiency of forming the native active-site conformation as if it was an intermolecular chaperone. In caspase-6, a similar structural conformation is observed in the zymogen and active states of the enzyme6 where an alternate form is adopted for the cleaved mature form as seen in caspase-9. Unlike caspase-9, caspase-7 has an observed increase in catalytic activity upon removal of the prodomain14, suggesting that the prodomain inhibits the function of the enzyme and possibly the proper formation of the active site loop bundle. Nonetheless, in all cases, the prodomain affects the catalytic function of the

150 caspase class of enzymes. It appears that the prodomains play more active role in regulating caspase activity than previously thought and affect function through an allosteric binding event. Therefore, investing time into understanding this mode of regulation and the location of these particular interactions sites would provide valuable information not only to the field of caspase function but also to the field of allosteric regulation.

By gaining an understanding of the additional modes of regulation imposed on caspase-9 function, it is clear that this enzyme uses the majority of its surface for regulatory control

(Fig 6.1). This characteristic can also be found for a variety of other proteins15-17 which exemplifies the Figure 6.1 Structure of caspase-9 with mapped regulatory surfaces. dynamic nature of the caspase class of enzymes and opens up the possibility of the caspases involvement in alternate roles within the cell. Furthermore, the discovery of these new regulatory sites native to the enzyme and possibly specific to caspase-9, increases the potential for specific drug targeting of an individual caspases and even makes the search for additional sites within the caspase class of enzymes an alluring concept.

Determination of these novel sites would be critical to distinguish one caspase from the next in a regulatory or drug targeting standpoint. Although their general

151 structural properties may be similar, it is suggested by Ranganathan and coworkers, that a statistical coupling between amino acids exists within a protein’s architecture and this coupling has the potential to impose functional constraints on the enzyme18-22. These networks provide a means of communication from a surface site on the protein to its functional active site and provide the underlying scheme that drives the co-evolution of new “hot spots” for protein regulation23,24. Investigation into these networks allowed for design of non-native allosteric control of dihydrofolate reductase24 and the PDZ family of proteins23. Therefore it is plausible to think that all proteins have these sensitive regulatory sites over the entire surface and at some point these sites will be exploited by nature to increase the ultimate fitness of the cell.

A study into the co-evolution of these “hot spot” networks via and computational simulations of correlated motions within the family of caspases could help distinguish unique patterns within the caspase architecture. An alternate yet parallel investigation utilizing NMR dynamics analysis or even thermal fluctuation analysis of crystal structures could provide insight into the dynamic properties of these enzymes and provide hints into the location of these networks as observed in the case of PDZ domains 20,25-27.

Hints towards these mechanisms within the caspase class of enzymes may already exist in the literature. Because thermodynamic values relating to protein stability under various conditions has been linked to functional cooperativity (reviewed in28), the presence of these networks in caspases seems likely. These studies would provide the information necessary for unique regulation of the caspases via a diverse set of novel allosteric sites.

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A potential venue to observe such networks may already exist by understanding the phosphorylation mechanisms of the caspase class of enzymes. Phosphorylation of a protein has been shown to increases the possibilities for protein-protein interactions as well as conformational regulation in a phosphorylation pathway. Therefore, it is intriguing to speculate how the co-evolving networks might change as a result of the phosphorylation patterns imposed on the enzyme during cellular signaling. Within the caspase class of enzymes, the function of some phosphorylation sites have been characterized to disrupt active site formation such as phosphorylation of S257 in caspase-

6 (unpublished data). However, the molecular details of most phosphorylation sites are unknown and determining those mechanisms could potentially link or even expose alternate “hot spot” networks to diversify the caspase function at a selected point in cellular signaling.

In summary, the emergence of allosteric mechanisms that occurs during the evolutionary process has the ability to diversify members of a single protein family.

Therefore, these mechanisms provide a means of specifically controlling a single protein from that of an otherwise undistinguishable class of enzymes. For this reason it is important to further explore native regulatory mechanisms of not only the caspases but other proteins as well to aid in understanding the protein function as well as be a guide for future design.

6.2. References

1. Andrade, F., Casciola-Rosen, L. A. & Rosen, A. A novel domain in adenovirus L4-100K is required for stable binding and efficient inhibition of human granzyme B: possible interaction with a species-specific exosite. Mol Cell Biol 23, 6315-6326, (2003).

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2. Ibarra, C. A., Blouse, G. E., Christian, T. D. & Shore, J. D. The contribution of the exosite residues of plasminogen activator inhibitor-1 to proteinase inhibition. J Biol Chem 279, 3643-3650, (2004).

3. Rogers, S. J., Pratt, C. W., Whinna, H. C. & Church, F. C. Role of thrombin exosites in inhibition by heparin cofactor II. J Biol Chem 267, 3613-3617, (1992).

4. Becker, D. L., Fredenburgh, J. C., Stafford, A. R. & Weitz, J. I. Exosites 1 and 2 are essential for protection of fibrin-bound thrombin from heparin-catalyzed inhibition by antithrombin and heparin cofactor II. J Biol Chem 274, 6226-6233, (1999).

5. Haase, H. & Maret, W. Protein tyrosine phosphatases as targets of the combined insulinomimetic effects of zinc and oxidants. Biometals 18, 333-338, (2005).

6. Vaidya, S., Velazquez-Delgado, E. M., Abbruzzese, G. & Hardy, J. A. Substrate- induced conformational changes occur in all cleaved forms of caspase-6. J Mol Biol 406, 75-91, (2011).

7. Bixby, K. A. et al. Zn2+-binding and molecular determinants of tetramerization in voltage-gated K+ channels. Nat Struct Biol 6, 38-43, (1999).

8. Maret, W., Yetman, C. A. & Jiang, L. Enzyme regulation by reversible zinc inhibition: glycerol phosphate dehydrogenase as an example. Chem Biol Interact 130-132, 891-901, (2001).

9. Zander, N. F. et al. Purification and characterization of a human recombinant T- cell protein-tyrosine-phosphatase from a baculovirus expression system. Biochemistry 30, 6964-6970, (1991).

10. Maret, W., Jacob, C., Vallee, B. L. & Fischer, E. H. Inhibitory sites in enzymes: zinc removal and reactivation by thionein. Proc Natl Acad Sci U S A 96, 1936- 1940, (1999).

11. Stennicke, H. R. & Salvesen, G. S. Biochemical characteristics of caspases-3, -6, - 7, and -8. J Biol Chem 272, 25719-25723, (1997).

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12. Stephanou, A., Scarabelli, T. M., Knight, R. A. & Latchman, D. S. Antiapoptotic activity of the free caspase recruitment domain of procaspase-9: a novel endogenous rescue pathway in cell death. J Biol Chem 277, 13693-13699, (2002).

13. Feeney, B. & Clark, A. C. Reassembly of active caspase-3 is facilitated by the propeptide. J Biol Chem 280, 39772-39785, (2005).

14. Denault, J. B. & Salvesen, G. S. Human caspase-7 activity and regulation by its N-terminal peptide. J Biol Chem 278, 34042-34050, (2003).

15. Gopal, V. K., Francis, S. H. & Corbin, J. D. Allosteric sites of phosphodiesterase- 5 (PDE5). A potential role in negative feedback regulation of cGMP signaling in corpus cavernosum. Eur J Biochem 268, 3304-3312, (2001).

16. Birdsall, N. J., Lazareno, S., Popham, A. & Saldanha, J. Multiple allosteric sites on muscarinic receptors. Life Sci 68, 2517-2524, (2001).

17. Hardy, J. A., Lam, J., Nguyen, J. T., O'Brien, T. & Wells, J. A. Discovery of an allosteric site in the caspases. Proc Natl Acad Sci U S A 101, 12461-12466, (2004).

18. Ferguson, A. D. et al. Signal transduction pathway of TonB-dependent transporters. Proc Natl Acad Sci U S A 104, 513-518, (2007).

19. Hatley, M. E., Lockless, S. W., Gibson, S. K., Gilman, A. G. & Ranganathan, R. Allosteric determinants in guanine nucleotide-binding proteins. Proc Natl Acad Sci U S A 100, 14445-14450, (2003).

20. Lockless, S. W. & Ranganathan, R. Evolutionarily conserved pathways of energetic connectivity in protein families. Science 286, 295-299, (1999).

21. Shulman, A. I., Larson, C., Mangelsdorf, D. J. & Ranganathan, R. Structural determinants of allosteric ligand activation in RXR heterodimers. Cell 116, 417- 429, (2004).

22. Suel, G. M., Lockless, S. W., Wall, M. A. & Ranganathan, R. Evolutionarily conserved networks of residues mediate allosteric communication in proteins. Nat Struct Biol 10, 59-69, (2003).

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23. Reynolds, K. A., McLaughlin, R. N. & Ranganathan, R. Hot spots for allosteric regulation on protein surfaces. Cell 147, 1564-1575, (2011).

24. Lee, J. et al. Surface sites for engineering allosteric control in proteins. Science 322, 438-442, (2008).

25. Ota, N. & Agard, D. A. Intramolecular signaling pathways revealed by modeling anisotropic thermal diffusion. J Mol Biol 351, 345-354, (2005).

26. Fuentes, E. J., Der, C. J. & Lee, A. L. Ligand-dependent dynamics and intramolecular signaling in a PDZ domain. J Mol Biol 335, 1105-1115, (2004).

27. Kong, Y. & Karplus, M. Signaling pathways of PDZ2 domain: a molecular dynamics interaction correlation analysis. Proteins 74, 145-154, (2009).

28. Luque, I., Leavitt, S. A. & Freire, E. The linkage between protein folding and functional cooperativity: two sides of the same coin? Annu Rev Biophys Biomol Struct 31, 235-256, (2002).

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APPENDIX A

SMALL MOLECULE ACTIVATION OF CASPASE-9

A.1. Introduction

A major factor in the activation of caspase-9 is regulation of its oligomeric state where the enzyme transitions from an inactive monomer to an active dimer1. Disruption of this mechanism leads to a down regulation of apoptosis, which is a characteristic of a variety of many cancer cells2-6. For this reason, molecules to control caspase activation are highly desired. To date, the molecule PAC-1 has been developed to activate the caspases through relieving their zinc mediated inhibition7,8, however, a molecule which directly acts on the initiator of the intrinsic pathway of the apoptotic cascade (caspase-9) has not been developed. An external trigger of caspase-9 dimerization would be a useful tool for understanding the detailed mechanism of caspase-9 activation and provide a proof of principal for small molecule development of caspase-9 activators.

The small fluorescent molecule, 4`, 5`-bis(1, 3, 2-dithioarsolan-2-yl)fluorescein-

(1, 2-ethanedithiol), otherwise known as Fluorescein Arsenical Hairpin or FlAsH9 recognizes a genetically encoded peptide tag making it a useful tool for studying protein localization, folding, stability and conformational changes10-14 and thus a tool for controlling caspase-9 dimerization. This fluorescein based molecule contains As(III) substituents on the 4`- and 5`- positions (Fig A.1) Figure A.1 Molecular structure of FlAsH-EDT2. which form a covalent bond with a reduced sulfhydryl found on a cysteine residue. The As(III) substituents are protected by 1,2-dithiol such as

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ethandithiol (EDT) to prevent non-specific binding to cysteine pairs in the protein, in addition to limiting toxicity and background fluorescence. The FlAsH recognition sequence includes two cysteine pairs separated by a two amino acid spacer (C-C-X-X-C-

C) where X could be any amino acid, although tightest binding is observed using Pro-Gly in those positions15. FlAsH is non-fluorescent in its unbound state. Upon binding to the

FlAsH recognition motif the small molecule becomes fluorescent with an excitation maximum of 508 nm and emission of 528 nm.

We aimed to engineer a FlAsH binding caspase-9 variant in which the FlAsH specific binding motif was incorporated across the dimer interface, similar to the split motif designed as a structural sensor of CRABP I10. FlAsH binding at this engineered site would lock the caspase-9 in the dimeric active form of the enzyme. Furthermore, the

FlAsH fluorescence will be used to confirm binding and the global location of this interaction within the enzyme. Ultimately, this study would provide a proof of concept that a small molecule can facilitate activation of caspase-9 through inducing dimerization of the enzyme and thus activation.

A.2. Results

A.2.1. Design of a FlAsH-Activatable Caspase-9

Harnessing the natural activation mechanism of caspase-9 will allow development of tools to further study this form of caspase regulation and therapeutics for diseases such as testicular cancer3 and infection of the vaccinia virus 16 in which the caspase-9 activation pathway is targeted. Therefore, our goal is to lock caspase-9 into a dimeric state via the small fluorescent molecule FlAsH. This would shift the enzymes equilibrium

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from inactive monomer to active dimer which would result in an overall increase in enzymatic activity.

Suitable sites for positioning the FlAsH binding motif across the dimer interface were selected based on four criteria; (1) the site would be able to accommodate not only the aresnic binding end of the molecule but also the bulky fluorescent portion of the probe, (2) the location of binding must be able to facilitate proper FlAsH binding geometry, (3) the designs must consider positions that would not disrupt the active site loop conformations and (4) if available, place half of the FlAsH motif on a homogeneous caspase-9 monomer where upon dimerization, the full binding motif could be formed.

Based on these outlined criteria, inspection of the wild-type caspase-9 structure

(PDB ID: 1JXQ) resulted in two FlAsH-binding models (Fig A.2 A). The first model,

C9-X, would place FlAsH on the periphery of the protein (Fig A.2 B). This site would link the C-terminal end of the α5 helix in caspase-9 to its symmetry partner on the other half of the dimer. This model is called C9-helix or C9-X. Amino acid variations at

Figure A.2 Models of the caspase-9 FlAsH binding variants. (A) An overall view of the caspase-9 dimer (blue/cyan) representing FlAsH binding sites for C9-X and C9-β within the dimer interface. (B) C9-X model of FlAsH binding to the α5 helix of caspase-9 at positions Q381C and S382C across the dimer interface. (C) C9-β model of FlAsH binding to the β6 strands of caspase-9 at positions C403 and N405C across the dimer interface.

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Gln381 and Ser382 to cysteine would facilitate the formation of the FlAsH binding motif upon dimerization of caspase-9 and create a covalent cross-link between the small subunit of one monomer to the small-subunit of the opposite monomer. These surface exposed residues would allow free movement of the FlAsH molecule while the positioning of this site is distant from the active site loop bundle which allows the C9-X model of FlAsH binding to fit all the desired criteria.

The second designed FlAsH binding motif focuses on the allosteric cavity observed within the dimer interface of the caspases17 (Fig A.2 C). The base of this site is lined with two β6 strands which are critical for the dimerization of caspase-918. This strand contains a native cysteine residue, Cys403, which became a focus for the design due to its location in the allosteric pocket and its close proximity to its symmetry related partner about the two-fold axis. The nearby residue, Asn405, sits directly across the dimer interface from the native cysteine, Cys403, and is in proper positioning for FlAsH binding. A cross-link at this position would link the small subunit from one monomer to the small-subunit of the opposite monomer. Therefore, Asn405 would complete the

FlAsH binding motif with native Cys403 for the C9-β model of FlAsH activation. Further inspection also suggests this pocket would be able to accommodate the entire FlAsH molecule. However, the C9-β model could potentially disrupt the inactive down conformation of the active site loops. Although avoiding disruption of loop conformation was a criterion for the design, at this position, binding of FlAsH could potentially shift the active site loop equilibrium towards the up or active conformation. Therefore, this model would not affect the overall goal of designing an active caspase-9 enzyme. Both

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the C9-X and C9-β FlAsH binding motifs fit the design criteria for activation of caspase-

9 via FlAsH binding.

A.2.2. Production of Caspse-9 FlAsH Variants

The caspase-9 gene construct encoding amino acids for the small subunit only, residues 331-416, in the pRSET was used for converting the wild type gene sequence into a version of the caspase-9 ΔCARD FlAsH variants. The oligo-nucleotide pimer 5`-

GCTCACTCTGAAGACCTGTGTTGCCTCCTCTTAGGGTCGC-3`was used to convert caspase-9 wild type sequence into the C9small-X variant for FlAsH binding studies via converting Gln381 and Ser382 to cysteine. Due to an error within the original primer sequence, a second oligo-nucleotide primer, 5`-CCTGCAGTCCCTCCTGCTTAGGG-

TCGCTAATGC-3`, was designed for correction of a base pair deletion at Leu384 which can be found in position 54 of the Hardy Lab DNA Archive Box 1. To make C9small-β, the oligo-nucleotide primer 5`-CAGATGCCTGGTTGTTTTTGCTTCCTCCGGAAA-

AAACTTTTC-3` was used for converting N405 to cysteine, found in position 51 of the

Hardy Lab DNA Archive Box 1. The previously described primers were also used for conversion of the caspase-9 full-length gene (human sequence), encoding amino acids 1-

416, in the pET23b vector (Addgene plasmid 11829) into a full-length version of the caspase-9 FlAsH variants, C9F-X and C9F-β, located in position 33 and 35 of the Hardy

Lab DNA Archive Box 2 Respectively. Furthermore, a FlAsH binding control variant was produced in the caspase-9 full-length construct (C9F-FC). The oligo-nucleotide primer, 5`- CCGGAAAAAACTTTTCTTTAAAACATCATGCTGCCCGGGCTGCT-

GCCACCACCACCACCACCACTAATCTAGCG-3`, was designed to incorporate the

FlAsH binding motif on the C-terminal end of the protein sequence just prior to the 6xHis

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tag. All caspase-9 FlAsH variants were produced via the Quikchange mutagenesis method (Stratagene).

The C9small-X and -β FlAsH constructs were separately transformed into the BL21

(DE3) T7 Express strain of E. coli (NEB). The recombinant large and small subunits were individually expressed as inclusion bodies for subsequent reconstitution. Cultures were grown in 2xYT media with ampicillin (100 mg/L, Sigma-Aldrich) at 37°C until they reached an optical density at 600 nm of 0.6. Protein expression was induced with 0.2 mM IPTG. Cells were harvested after 3 hrs at 37°C. Cell pellets stored at -20°C for future use. The resultant inclusion bodies were refolded and purified ion exchange column purification step only which is outlined in Chapter III.

C9F-X and -β FlAsH variants were also transformed into the BL21 (DE3) T7

Express strain of E. coli (NEB). The cultures were grown in 2xYT media with ampicillin

(100 mg/L, Sigma-Aldrich) at 37°C until they reached an optical density at 600 nm of

0.8. The temperature was reduced to 20°C and cells were induced with 1 mM IPTG

(Anatrace) to express soluble 6xHis-tagged full-length protein. Cells were harvested after 18 hrs and stored at -20°C. The resultant protein was purified using the standard caspase-9 imidazole gradient purification followed by an ion-exchange column purification step outlined in Chapter IV. The purified C9F-X and -β FlAsH variants proteins are estimated to be approximately 95% pure with a two-site cleavage (at residues

D315 and E306 (Large) or D330 (small)) within its intersubunit linker was used for further analysis.

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A.2.3. Analysis of FlAsH Binding

Initial FlAsH binding studies were performed with the ΔCARD version of caspase-9. Caspase-9 ΔCARD -X, -β, and wild-type enzymes were concentrated to 10

μM in Amicon Ultracell 3K concentrator (Millipore). The protein samples were then diluted to 4.5 μM in a degassed buffer that contains 0.1 M MES pH 6.5, 10% sucrose, 0.1

% CHAPS and 10 mM DTT which was then incubated at 4 °C to reduce the cysteine thiols. After an incubation time of 15 hours, each caspase-9 variant was incubated with 0,

0.5 or 2-fold excess FlAsH at room temperature. Samples of caspase-9 proteins with

FlAsH were anaylzed at 0.5, 3, 7 and 24 hours of incubation time by SDS-PAGE for detection of the cross-linked facilitated by FlAsH between two caspase-9 small subunits

(20 kDa) followed by an in-gel fluorescence for detection of FlAsH binding (Fig A.3). A

Figure A.3 Analysis of FlasH binding to caspase-9 ΔCARD. Analysis was performed via SDS- PAGE and in gel fluorescence at 0.5 hr (A) or 24 hr (B). FlAsH proteins assessed for small subunit cross-linking and FlAsH binding are the caspase-9 wild type (WT), and the engineered caspase-9 variants C9-X (X) and C9-β (β).

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fluorescent signature was observed just below the 20 kDa molecular weight marker as expected of a cross-link between two small subunits within the dimer of caspase-9. Upon further inspection of the wild type control, it was discovered that FlAsH was non- specifically binding to the large subunit of the enzyme (18 kDa). This phenomenon has been reported prior for other cysteine rich proteins19. To alleviate fluorescence within the expected 20 kDa region, the caspase-9 -X and -β variants were produced in the full length version of the enzyme in which the large subunit is connected to the CARD domain resulting in a 36.5 kDa fragment while the small subunit still remains at approximate 10 kDa. By using this form of the enzyme, a clear image of FlAsH fluorescence can be observed around the 20 kDa marker. A full-length caspase-9 variant which incorporates the FlAsH binding motif -CCPGCC- on the C-terminus was also created for a FlAsH binding control, labeled as C9F-FC (position 38 in the Hardy Lab DNA Archive Box 2).

10 μM of the C9F-WT, C9F-X, C9F-β and C9F-FC FlAsH variants were diluted to 4.5

μM in a reducing buffer containing 0.1 M MES pH 6.5, 10 % sucrose, 50 mM TCEP, which was used as a non-thiol based reductant. This mixture was allowed to incubate for one hour at room temperature. Samples were then subjected to FlAsH binding by addition of 0.5 mM 1,2-ethandithiol and a 2-fold excess of the FlAsH fluorescent compound.

Samples were then anayzed by SDS-PAGE (Fig A.4 A) and in-gel fluorescence methods

(Fig A.4 B). The small subunit of C9F-FC variant appears as a fluorescent band indicating these conditions result in a functional assay for detecting FlAsH fluorescence.

Furthermore, fluorescent bands are also observed for a molecular weight band corresponding to large + CARD (36 kDa) and large + small (28 kDa) as expected from the non-specific binding of the FlAsH molecule observed in the caspase-9ΔCARD

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Figure A.4. Analysis of FlasH binding to full-length caspase-9. Analysis was performed via (A) SDS-PAGE and (B) in gel fluorescence. FlAsH proteins assessed for small subunit cross-linking and FlAsH binding are the full-length caspase-9 FlAsH binding control (FC), full-length caspase-9 wild type (WT), and the engineered caspase-9 variants C9F-X (X) and C9F-β (β). version of the enzyme. Although the non-specific FlAsH binding is observed for the large subunit, under these conditions, detectable fluorescence of a molecular weight band is visible corresponding to a ~20 kDa fragment. This particular fragment is identified to contain the small subunit as judged by western blot analysis with an anit-caspase-9 small subunit antibody (Fig A.5). However, this band is also observed in the SDS-PAGE analysis, in the absence of FlAsH, suggesting potential disulfide bond cross-linking at the designed sights. Reductant concentration, identity as well as time of incubation was tested however this band still remained intact. It is possible that this band is a contaminant protein or the

Figure A.5. Western blot analysis of FlasH binding to full- caspase-9 large subunit only cross length caspase-9. Analysis was performed using an antibody specific for the small subunit of caspase-9. reacts with the caspase-9 antibody FlAsH. proteins assessed for small subunit cross-linking and FlAsH binding are the full-length caspase-9 FlAsH binding control (FC), full-length caspase-9 wild type and is cysteine rich explaining the (WT), and the engineered caspase-9 variants C9F-X (X) and C9F-β (β).

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minimal FlAsH fluorescence detected. Further studies are needed to determine whether this 20 kDa band is a pre-formed small-small dimer, caspase-9 large subunit only, or a protein contaminate.

Considering there was not a robust appearance of FlasH induced caspase-9 dimerization or fluorescence, efforts turned towards identifying the location of the non- specific FlAsH binding for further characterization and potential future design. The full- length caspase-9 contains thirteen cysteines in its amino acid sequence. Therefore, the cysteine residues that are solvent exposed were targeted as potential FlAsH binding sites.

Selected positions, C162, C172, C229, and C239 were interrogated for their role in the non-specific FlAsH binding observed due to their proximity to other cysteine residues.

Each position was converted to either a serine or an alanine by Quikchange mutagenesis

(Stratagene) and tested for a reduction in FlAsH binding. Single caspase-9 variants,

C162A, C172A, C229A, C239S, and the double variant C229A/C230A did not show a significant decrease in FlAsH fluorescence. This suggests that an alternate positioning of the FlAsH binding site exists or there is a need for construction of a caspase-9 variant which incorporates multiple cysteine locations other than C229/C230 due to the fact that fluorescence by FlAsH has been reported to occur when bound to less than four cysteines19.

A.3. Discussion

The initiator caspase of the intrinsic pathway of apoptosis, caspase-9, has the potential to be a good target for regulation by a small molecule. Caspase-9 plays a pivotal role in the activation of the cleavage cascade of apoptosis and its activation mechanism is targeted in a variety of diseases anywhere from viral infections to cancer. Taking

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inspiration from natural activation pathways of caspase-9, such as dimer formation, we designed two variants of caspase-9, C9-X and C9-β that could potentially dimerize upon binding of a small fluorescent molecule known as FlAsH. This would provide a proof-of- concept that caspase activation can be controlled by a small molecule regulator.

Upon analysis, distinct induction of the caspase-9 dimer was not observed, in addition to the lack of a robust fluorescence signal expected from binding of the FlAsH molecule, which was observed for the control protein. Although precautions were taken when designing FlAsH binding sites across the dimer interface, C9-X and C9-β were unsuccessful. Unforeseen pitfalls such as improper geometric binding angles for FlAsH binding within the designed cysteine motif or limited access to the designed site could have prevented FlAsH from binding the engineered caspases. In addition, it is possible that FlAsH could only bind to a pre-formed dimer of caspase-9 which is limited in vitro1.

This hypothesis can be tested by addition of a covalent active site inhibitor which shifts the enzyme’s equilibrium from monomer to dimer. Further analysis of the geometric restrictions imposed by these sites as well as the discovery of alternative FlAsH binding sites or alternative molecules could result in a caspase-9 variant that can be activated through a small molecule trigger.

Although the desired outcome was not obtained, new information was discovered about caspase-9. A natural FlAsH binding site exists within the large subunit of caspase-

9. Once the location of the non-specific FlAsH site is determine, this site can be optimized to obtain a more robust FlAsH signal and potentially exploit the site for allosteric control as in the case of protein tyrosine phosphate 1B20.

A.4. References

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1. Renatus, M., Stennicke, H. R., Scott, F. L., Liddington, R. C. & Salvesen, G. S. Dimer formation drives the activation of the cell death protease caspase 9. Proc Natl Acad Sci U S A 98, 14250-14255, (2001).

2. Palmerini, F., Devilard, E., Jarry, A., Birg, F. & Xerri, L. Caspase 7 downregulation as an immunohistochemical marker of colonic carcinoma. Hum Pathol 32, 461-467, (2001).

3. Mueller, T. et al. Failure of activation of caspase-9 induces a higher threshold for apoptosis and cisplatin resistance in testicular cancer. Cancer Res 63, 513-521, (2003).

4. Mizutani, Y. et al. Downregulation of Smac/DIABLO expression in renal cell carcinoma and its prognostic significance. J Clin Oncol 23, 448-454, (2005).

5. Sekimura, A. et al. Expression of Smac/DIABLO is a novel prognostic marker in lung cancer. Oncol Rep 11, 797-802, (2004).

6. Kempkensteffen, C. et al. The equilibrium of XIAP and Smac/DIABLO expression is gradually deranged during the development and progression of testicular germ cell tumours. Int J Androl 30, 476-483, (2007).

7. Peterson, Q. P. et al. PAC-1 activates procaspase-3 in vitro through relief of zinc- mediated inhibition. J Mol Biol 388, 144-158, (2009).

8. Wolan, D. W., Zorn, J. A., Gray, D. C. & Wells, J. A. Small-molecule activators of a proenzyme. Science 326, 853-858, (2009).

9. Griffin, B. A., Adams, S. R. & Tsien, R. Y. Specific covalent labeling of recombinant protein molecules inside live cells. Science 281, 269-272, (1998).

10. Krishnan, B. & Gierasch, L. M. Cross-strand split tetra-Cys motifs as structure sensors in a beta-sheet protein. Chem Biol 15, 1104-1115, (2008).

11. Luedtke, N. W., Dexter, R. J., Fried, D. B. & Schepartz, A. Surveying polypeptide and protein domain conformation and association with FlAsH and ReAsH. Nat Chem Biol 3, 779-784, (2007).

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12. Ignatova, Z. & Gierasch, L. M. Monitoring protein stability and aggregation in vivo by real-time fluorescent labeling. Proc Natl Acad Sci U S A 101, 523-528, (2004).

13. Ignatova, Z. & Gierasch, L. M. A method for direct measurement of protein stability in vivo. Methods Mol Biol 490, 165-178, (2009).

14. Ignatova, Z. et al. From the test tube to the cell: exploring the folding and aggregation of a beta-clam protein. Biopolymers 88, 157-163, (2007).

15. Adams, S. R. et al. New biarsenical ligands and tetracysteine motifs for protein labeling in vitro and in vivo: synthesis and biological applications. J Am Chem Soc 124, 6063-6076, (2002).

16. Zhai, D. et al. Vaccinia virus protein F1L is a caspase-9 inhibitor. J Biol Chem 285, 5569-5580, (2010).

17. Hardy, J. A., Lam, J., Nguyen, J. T., O'Brien, T. & Wells, J. A. Discovery of an allosteric site in the caspases. Proc Natl Acad Sci U S A 101, 12461-12466, (2004).

18. Chao, Y. et al. Engineering a dimeric caspase-9: a re-evaluation of the induced proximity model for caspase activation. PLoS Biol 3, e183, (2005).

19. Stroffekova, K., Proenza, C. & Beam, K. G. The protein-labeling reagent FLASH-EDT2 binds not only to CCXXCC motifs but also non-specifically to endogenous cysteine-rich proteins. Pflugers Arch 442, 859-866, (2001).

20. Zhang, X. Y. & Bishop, A. C. Site-specific incorporation of allosteric-inhibition sites in a protein tyrosine phosphatase. J Am Chem Soc 129, 3812-3813, (2007).

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APPENDIX B

DESIGN OF AN ACTIVATABLE INITIATOR CASPASE

B.1. Introduction

Caspases, proteases that control apoptosis, have been a target in cancer research for many years. In particular, caspase-9, an initiator of the caspase cleavage cascade, has been shown to be down regulated in tumor cells1,2. Therefore, activating the caspase cleavage cascade, via the intrinsic pathway of apoptosis by utilizing caspase-9 within these tumors, would be a useful tool to study apoptosis in tumors and to halt tumor progression. Critical steps within the caspase-9 activation pathway, such as dimerization and active site loop conformation, can be used as inspiration for designing such a tool.

Caspase-9’s equilibrium in solution lies toward the monomeric state3. Therefore designing a way to shift the equilibrium of caspase-9 from monomer to dimer would aid in increasing the active population of caspase-9 molecules. Additionally, if creating a dimeric variant of caspase-9 could also be facilitated via control of the active site loop bundle, further enhancement of caspase-9 activity would be expected.

Nature provides naturally occurring amino acids, such as cysteines, that form covalent, redox-controllable cross-links, which became a useful protein-engineering tool for controlling a proteins conformation. The disulfide cross-link has been used for a variety of applications including stabilization of protein conformational states and analyzing functional mechanisms4-10 on anywhere from viral proteins11 to critical cellular transcription factors12 and even for release from a delivery system13-17. Wide applicability and controlability of this technique, based upon redox environment18,19, makes disulfide cross-linking an intriguing tool for activating caspase-9. Therefore we aim to rationally

170 design a caspase-9 variant to become activated by inducing dimerization and formation of the active site loop bundle via a disulfide cross-link.

B.2. Results

B.2.1. Rational Redox Controllable Caspase-9

Like other caspases, caspase-9 undergoes distinct structural conformation changes based upon the state of the enzyme. Caspase-9 exists as an inactive monomer within the cytosol and becomes active upon dimerization3. Crystallographic evidence shows an active site loop bundle that adopts a disordered conformation in the absence of substrate and changes conformation to a more ordered state upon binding substrate3. The active state of the enzyme is a dimer in which the active site loop, L2, from one half of the dimer interacts with the L2` loop from the other half. Our goal is to lock caspase-9 into the active state, overcoming the slow step of the caspase-9 activation process and thus preparing it to cleave substrate.

The natural residue, cysteine, was utilized to create a redox controllable link between both halves of the dimer. Wild-type caspase-9 protein contains thirteen native cysteine residues, where a natural redox regulation event or a native disulfide bond linkage has not been observed within the enzyme to date. These thirteen native cysteines within the caspase-9 amino acid sequence were initially investigated for potentially forming a natural disulfide link. The only native cysteine capable of this would be C403, which resides within the dimer interface. Based on conformation with the β-sheet secondary structure and distance from the corresponding C403 across the two fold axis of the dimer interface, disulfide cross-linking at this site would not be feasible. Therefore a computational tool to predict the availability of residues for disulfide engineering, known

171 as MODIP, was utilized to predict which amino acid positions would facilitate proper amino acid pairing for potential disulfide linkage. The best prediction across the dimer interface comprised the residues Lys399, found at the top of the L4 loop, and Pro339 of the L2` loop from the opposite monomer. However, cross-linking two active site loops which have different functions would most likely have a negative effect on activity of the enzyme. Therefore, the caspase-9 crystal structure, in the active state, was inspected for alternate designs. Inspired by MODIP’s prediction of locking active site loop, interaction between L2 and L2`of caspase-9 in the substrate bound state (PDB ID: 1NW9) was investigated. The L2 and L2` loops naturally interact upon binding of substrate therefore the addition of a cross-link at this position would enhance the probability of the conformational state. Upon inspection of the caspase-9 active conformation, Glu297 on the L2 loop and Ser334C on L2` appeared to be within optimal distance and conformation as they reside just past the intersection point of the L2 and L2` (Fig B.1).

Optimal distance for disulfides was defined as 4.4-6.8 Å from Cα-Cα20. A disulfide at this site would link the large and small subunits of the enzyme together. Therefore these two sites were chosen as the link site for the redox controllable caspase-9 variant.

B.2.2 Production of a Redox

Controllable Caspase-9 Figure B.1 Model of the designed caspase-9 disulfide The caspase-9 full-length cross linked variant.

172 gene (human sequence) construct, encoding amino acids 1-416, in pET23b (Addgene plasmid 11829 ) was used to design a genetic construct for expression and purification from Bl21(DE3) E.coli strain of bacteria of the disulfide activatiable caspase-9 variant.

The oligo-nucleotide pimer 5’ GCAGAAAGACCATGGGTTTTGCGTGGCCTCCA-

CTTCCCC 3’converted position 297 from a Glu to a Cys while and a second primer was designed, 5’ GCTGGACGCCATATCTTGTTTGCCC-ACACCC 3’, to convert position

344 from a Ser to a Cys utilizing the Quikchange mutagenesis method. First, C9F-E297 was changed to cysteine, found in position 37 of the Hardy Lab DNA Archive Box 2. The final construct of C9F-E297C/S334C, alternately named C9F-DS was made by performing Quikchange mutagenesis using the Ser334 oligo-primer in the background of the C9F-E298 construct stored in the Hardy Lab DNA Archive Box 2 position 38.

The C9F-DS construct was transformed into the BL21 (DE3) T7 Express strain of

E. coli (NEB) for expression. The cultures were grown in 2xYT media with ampicillin

(100 mg/L, Sigma-Aldrich) at 37°C until they reached an optical density at 600 nm of

0.8. The temperature was reduced to 20°C and cells were induced with 1 mM IPTG

(Anatrace) to express soluble 6xHis-tagged full-length protein. Cells were harvested after 18 hrs and stored at -20°C. The resultant protein was purified using the standard caspase-9 imidazole gradient purification followed by an ion exchange column purification step, in which a detailed protocol can be found in Appendix D: Protocol D1.

The purified C9F-DS protein, estimated to be approximately 95% pure with a two site cleavage within its intersubunit linker was used for further analysis.

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B.2.3 Analysis of Designed Caspase-9 Disulfide Variant

The caspase-9 disulfide variant was tested for its ability to cross-link caspase-9 into an active state. Activity and cross-linking of the large and small subunits were used as readouts for success of the design. Initial titrations with reductant, DTT, did not show a change in activity between the engineered mutant, C9F-DS, and wild type protein.

Therefore, the purified proteins in 20 mM Tris pH 8.5, 100 mM NaCl, and 5 mM DTT were subjected to a Nap-5 desalting column (GE Healthcare) for removal of the DTT reductant and to buffer exchange the proteins into an activity buffer of 10% sucrose and

0.1 M MES pH 6.5. Caspase-9 exhibits a high propensity of precipitation in these conditions; however, sufficient protein of both C9F-DS and WT was recovered from the soluble fraction for experimentation. Proteins were diluted to 5 μM, as judged by absorbance at 280 nm, in assay buffer containing either 10 mM DTT for reduction of any pre-formed disulfide cross-links, 500 μM oxidixed glutathione, or slowly bubbled with air as an oxidant to facilitate formation of the engineered disulfide bond (Fig B.2).

Figure B.2. Disulfide cross-linking of wild type caspase-9 full-length and disulfide variant. SDS-PAGE analysis of caspase-9 wild type and disulfide variant in the presence and absence DTT, oxidized glutathione (GSSG), and air. Expected disulfide cross link would be represented as the full length form of the enzyme.

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Samples of the reaction were pulled at time points 0, 0.5, 1, and 2 hours to monitor the potential cross-link. A cross-link between the large and small subunits, forming the expected intact monomer, was not observed pre or post treatment with oxidized glutathione. However, a minimal amount of cross-linked monomer was detected upon oxidation with air at the one and two hour time points. Reduction in protein concentration was also observed due to precipitation of protein during the experimental timeframe, indicating a need for further optimization of experimental conditions. Protein precipitate was also observed for the wild type version of the enzyme (data not shown), therefore alternate ways to obtain the cross-linked form of C9F-DS need further exploration.

The designed redox controllable variant, C9F-DS, was successful to some degree under air oxidizing conditions only. Due to protein precipitation issues, studies of enzymatic activity posed to be troublesome, therefore investigating whether this variant can induced dimerization of the small and large subunits via a cross-link between the L2 and L2` loops was perused. This would verify this site alone can enforce the dimer form of the enzyme, which is required for activity. Small maleimide-based cross-linkers,

BMOE (bis-maleimidoethane) and BMDB (1,4 bismaleimidyl-2,3-dihydroxybutane)

(Pierce), of arm spacer lengths of 8.0 Å and 10.2 Å respectively, were used as cross- linking agents (Fig B.3 A-B). 300 μM of BOME or BMDB was added to 10 μM C9F-DS and WT protein that was reduced for 30 minutes in 5mM TCEP and equilibrated in 0.1 M phosphate buffer pH 7.0, 0.15 M NaCl and 5 mM EDTA. Samples were incubated for two hours at room temperature and immediately analyzed via SDS-PAGE (Fig B.3 C-D).

For both BMOE and BMDB cross-linking compounds, the expected large plus small molecular weight band is present; however, it is only slightly more concentrated than

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Figure B.3. Chemical cross-linking of wild type caspase-9 full-length and disulfide variant. (A-B) Small spacer arm maliemide based cross linking compounds. (A) BMOE (B) BMDB. (C-D) SDS- PAGE analysis of caspase-9 wild type and disulfide variant in the presence and absence of BMOE (C) and BMDB (D). Expected disulfide cross link would be represented as the full length form of the enzyme. samples that do not include the maleimide compounds. This result indicates that the

E297C/S334C site is capable of associating the large and small subunits via sulfhydryl cross-linking compounds in addition to formation of this complex under different buffer conditions.

B.3. Discussion

The initiator of the caspase cleavage cascade for the intrinsic pathway of apoptosis, caspase-9, is a promising target for activating apoptosis in tumor cells. Taking inspiration from natural activation pathways of caspase-9, such as dimer formation and active site loop orientation, we designed a disulfide cross-linked version of the caspase-9 enzyme. A cross-link between the large and small subunits of caspase-9, due to the altered amino acids E297C and S334C within the intersubunit linker, was observed under air oxidation as well as the presence of sulfhydryl cross-linking agents. However, precipitation of both the control wild type protein, as well as the disulfide variant posed

176 as a large issue within these experiments potentially due to intramolecular disulfides being formed between the thirteen native cysteines within the enzyme. The incidence of cross-linking between the large and small subunits of caspase-9 was observed to a greater degree (Fig B.3.D) than in the control experiments, suggesting induced dimerization of the enzyme. However protein precipitation and the intrinsic requirement of reductant prevented activation analysis.

Verification of caspase-9 increased activation is critical for making this disulfide mediated caspase-9 variant a useful tool. Therefore, optimization of the assay conditions is required to enforce the disulfide cross-link, while still being able to assay for activity and avoid precipitation of the enzyme. Identification of the lowest amount of reductant necessary for caspase-9 activity and solubility would also be useful for solving these issues. Potentially a redox buffer that includes both reduced and oxidized glutathione is more closely related to that of the chemical cross-linking studies, would be more appropriate. In addition, removal of any surface exposed cysteines would aid in this endeavor.

B.4. References

1. Mueller, T. et al. Failure of activation of caspase-9 induces a higher threshold for apoptosis and cisplatin resistance in testicular cancer. Cancer Res 63, 513-521, (2003).

2. Palmerini, F., Devilard, E., Jarry, A., Birg, F. & Xerri, L. Caspase 7 downregulation as an immunohistochemical marker of colonic carcinoma. Hum Pathol 32, 461-467, (2001).

3. Renatus, M., Stennicke, H. R., Scott, F. L., Liddington, R. C. & Salvesen, G. S. Dimer formation drives the activation of the cell death protease caspase 9. Proc Natl Acad Sci U S A 98, 14250-14255, (2001).

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4. Jeong, M. Y., Kim, S., Yun, C. W., Choi, Y. J. & Cho, S. G. Engineering a de novo internal disulfide bridge to improve the thermal stability of xylanase from Bacillus stearothermophilus No. 236. J Biotechnol 127, 300-309, (2007).

5. Ma, K., Temiakov, D., Anikin, M. & McAllister, W. T. Probing conformational changes in T7 RNA polymerase during initiation and termination by using engineered disulfide linkages. Proc Natl Acad Sci U S A 102, 17612-17617, (2005).

6. Mitchinson, C. & Wells, J. A. Protein engineering of disulfide bonds in subtilisin BPN'. Biochemistry 28, 4807-4815, (1989).

7. Qureshi, S. H., Yang, L., Manithody, C., Iakhiaev, A. V. & Rezaie, A. R. Mutagenesis studies toward understanding allostery in thrombin. Biochemistry 48, 8261-8270, (2009).

8. Seeger, M. A. et al. Engineered disulfide bonds support the functional rotation mechanism of multidrug efflux pump AcrB. Nat Struct Mol Biol 15, 199-205, (2008).

9. Shandiz, A. T., Capraro, B. R. & Sosnick, T. R. Intramolecular cross-linking evaluated as a structural probe of the protein folding transition state. Biochemistry 46, 13711-13719, (2007).

10. Witkowski, W. A. & Hardy, J. A. A designed redox-controlled caspase. Protein Sci, (2011).

11. Lee, J. K., Prussia, A., Snyder, J. P. & Plemper, R. K. Reversible inhibition of the fusion activity of measles virus F protein by an engineered intersubunit disulfide bridge. J Virol 81, 8821-8826, (2007).

12. Zheng, M., Aslund, F. & Storz, G. Activation of the OxyR transcription factor by reversible disulfide bond formation. Science 279, 1718-1721, (1998).

13. Bauhuber, S., Hozsa, C., Breunig, M. & Gopferich, A. Delivery of nucleic acids via disulfide-based carrier systems. Adv Mater 21, 3286-3306, (2009).

14. Hong, R. et al. Glutathione-mediated delivery and release using monolayer protected nanoparticle carriers. J Am Chem Soc 128, 1078-1079, (2006).

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15. Navath, R. S. et al. Dendrimer-drug conjugates for tailored intracellular drug release based on glutathione levels. Bioconjug Chem 19, 2446-2455, (2008).

16. Ouyang, D., Shah, N., Zhang, H., Smith, S. C. & Parekh, H. S. Reducible disulfide-based non-viral gene delivery systems. Mini Rev Med Chem 9, 1242- 1250, (2009).

17. Singh, R. & Lillard, J. W., Jr. Nanoparticle-based targeted drug delivery. Exp Mol Pathol 86, 215-223, (2009).

18. Russo, A., DeGraff, W., Friedman, N. & Mitchell, J. B. Selective modulation of glutathione levels in human normal versus tumor cells and subsequent differential response to chemotherapy drugs. Cancer Res 46, 2845-2848, (1986).

19. Jones, D. P., Brown, L. A. & Sternberg, P. Variability in glutathione-dependent detoxication in vivo and its relevance to detoxication of chemical mixtures. Toxicology 105, 267-274, (1995).

20. Richardson, J. S. The anatomy and taxonomy of protein structure. Adv Protein Chem 34, 167-339, (1981).

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APPENDIX C

EXPRESSION AND PURIFICATION OF A YEAST METACASPASE YCA1

C.1. Introduction

Saccharomyces cerevisiea yeast has served as a model organism for a huge number of genetic and regulatory pathways due to its ease of genetic manipulation and strikingly conserved pathways in comparison to other eukaryotic organisms.

Furthermore, this model system has demonstrated apoptotic-like cell death with such markers as DNA fragmentation and chromatin condensation1. Since the discovery of apoptosis in yeast, a number of yeast orthologs to key players in the mammalian apoptotic cycle have been identified, indicating conserved cell death pathways for proteasomal and mitochondrial-type cell death2.

An apoptotic-like phenotype in yeast has even been observed upon a variety of triggers including exposure to reactive oxygen species 3 and aging4. Characteristic homologues for apoptotic regulators such as the Bcl-2 family of proteins have not yet been discovered in the yeast model system; however, a caspase-like protein has been identified in the S. cerevisiea genome. Yeast caspase-1 (YCA1 also known as

YOR197W) has been classified as an evolutionary ancestor to human caspases5, or a metacaspase, due to primary sequence identification of a catalytic cysteine-histidine diad as well as a predicted caspase like fold. YCA1 has been highlighted as the protein responsible for apoptotic like cell death in yeast6. For example, knockout of the YCA1

7 gene in S. cervisiea has been shown to abolish an H2O2 induced form of apoptosis . In addition to YCA1’s ability to control the apoptotic properties of yeast, this protease shows caspase like character in both its cleavage properties and substrate recognition

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sequences, VEID and IETD. YCA1 studies have mainly been performed in whole yeast cells to understand how overexpression of the enzyme affects cell survival rates as well as how different apoptosis inducing factors, such as H2O2, affect survival when in the presence and absence of the YCA1 gene4,6,8,9. Additional studies involved removal of the soluble protein fractions from the yeast cells for measurement of YCA1 expression levels under normal and stress conditions7 and utilizing cellular extracts to test the enzymatic activity of YCA110. These caspase-like characteristics therefore make this metacaspase an intriguing target for protein engineering and structural studies. We aimed to express and purify YCA1 to perform biochemical characterizations for further elucidation of the proteases similarities to its human homologues. This ultimately would allow solution of the crystal structure of YCA1 to observe its structural similarities and to design alternate ways to regulate the enzyme though protein engineering which can be easily tested in yeast as the model system.

C.2. Results

C.2.1. Gene Construction

The S. cervisiea YCA1 gene was used to design and prepare a genetic construct for expression and purification from Bl21(DE3) E.coli strain of bacteria. The gene was amplified from a yeast plasmid by PCR for insertion into the pET21(+)b expression vector. Oligo-nucleotide pimers, 5`-AGTCGTGCTAGCATGTATCCAGGTAGTGGA-

CGTTACACC-3` and 5`-AGTATACTCGAGCTACATAATAAATTGCAGATTTAC-

GTCAATAGGG-3`, encode the NheI and Xho1 endonuclease restriction sites for the 5` and 3` primers, respectively, were utilized for the amplification. The PCR product was digested to expose complementary DNA overhangs and gel purified, resulting in a 64

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ng/μl digested YCA1 DNA gene product. Ligation of the YCA1 gene to the digested the pET21(+)b expression vector was performed through incubating one molar ratio of vector with three molar ratios of YCA1 gene insert with T4 DNA ligase at 16°C overnight. The ligation reactions were transformed into the TAM1 cloning strain of E.coli cells and plated onto 100 mg/ml ampicillin plates. Transformants were tested for successful ligations through enzymatic digest with NheI and XhoI to reveal the insertion of the

YCA1 gene which was subsequently confirmed through DNA sequencing. The resulting plasmid encodes the YCA1 gene with a C-terminal 6xHis tag driven by the T7 promoter with apmicillin resistance (Fig C.1). This plasmid is housed in the Hardy Lab DNA archive box 1 position 9.

Figure C.1. Schematic representation of expression vector, pYCA1. The T7 promoter, YCA1 gene, 6xHis tag, and ampicillin resistance are shown.

C.2.2. Protein Sequence

10 20 30 40 50 60 MYPGSGRYTY NNAGGNNGYQ RPMAPPPNQQ YGQQYGQQYE QQYGQQYGQQ NDQQFSQQYA

70 80 90 100 110 120 PPPGPPPMAY NRPVYPPPQF QQEQAKAQLS NGYNNPNVNA SNMYGPPQNM SLPPPQTQTI

130 140 150 160 170 180 QGTDQPYQYS QCTGRRKALI IGINYIGSKN QLRGCINDAH NIFNFLTNGY GYSSDDIVIL

182

190 200 210 220 230 240 TDDQNDLVRV PTRANMIRAM QWLVKDAQPN DSLFLHYSGH GGQTEDLDGD EEDGMDDVIY

250 260 270 280 290 300 PVDFETQGPI IDDEMHDIMV KPLQQGVRLT ALFDSCHSGT VLDLPYTYST KGIIKEPNIW

310 320 330 340 350 360 KDVGQDGLQA AISYATGNRA ALIGSLGSIF KTVKGGMGNN VDRERVRQIK FSAADVVMLS

370 380 390 400 410 420 GSKDNQTSAD AVEDGQNTGA MSHAFIKVMT LQPQQSYLSL LQNMRKELAG KYSQKPQLSS

430 SHPIDVNLQF IM

C.2.3. YCA1 Expression

The YCA1 gene is expected to produce ~50 kDa 6xHis-tagged protein of 438 amino acids in length. YCA1 is reported to undergo autoprocessing upon induction of apoptosis or overexpression of the enzyme in yeast. Therefore, two cleavage products would be observed, a ~12 kDa portion of the enzyme which is noted as the small subunit and ~38 kD molecular weight piece of the remaining enzyme that would comprise the large and pro-domains.

In order to obtain a homogenous sample of the active form of the YCA1 protein, expression and purification of the YCA1 was performed. Cell growth of the BL21(DE3) strain of E.coli transformed with the YCA1-pET21(+)b plasmid in 2xYT media was performed. When OD600 reached 0.6, cells were induced for protein expression with

1mM IPTG at 14, 30, or 37 °C. A standard 6x-His tag purification protocol was utilized in order to obtain the desired protein from cellular lysates as outlined by the Current

Protocols in Protein Science11. Under these standard expression and purification conditions, protein induced at OD600= 0.6 with 1 mM IPTG 14°C for 18 hrs produced an elution fraction that contained minimal amount of protein of the expected molecular weight of full length YCA1 as judged by Western blot analysis with an antibody specific

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for the 6xHis-purification tag (Fig. C.2) However, in addition to low expression levels, the enzymatic autoprocessing was incomplete and breakdown products are observed.

Activity of protein identified in the elution fraction was assessed though cleavage of fluorogenic substrate VEID-AMC (Ex= 380nm; Em= 460), a previously reported cleavage sequence for YCA1 in yeast. Unfortunately, no activity was detected. Human homologue, caspase-6, was used as a control for activity against the fluorogenic

Figure C.2. Purification of YCA1. (A-B) SDS-PAGE of the loaded supernatant (L), flow through (FT), wash (W) and elution (E) fractions from a Ni-affinity purification of YCA1. Molecular weights determined from marker (M) and caspase-7 was used as a control (C7). (A) Coomassie stain. (B) Anti-his western blot. substrate. ARR-2NA, a previously reported fluorogenic substrate for YCA1 was also tested. Activity was observed within the supernatant of lysed cells, however, no detectable activity was observed for the purified protein. The lack of observed YCA1 activity is thought to be due to the presence of the inactive full length version of the protein or the multiple breakdown products of the processed enzyme.

In order to optimize expression conditions for overexpression of the YCA1 protein in E.coli with the proper molecular weight products, theYCA1-pET21(+)b

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expression vector was transformed into various forms of the BL21(DE3) E.coli cell line including the original BL21(DE3) strain as well as the PLysS, Codon +, and Rosetta strains. Induction of protein expression was performed at an OD600 = 0.4 and 0.6 in a 5 mL 2xYT media culture format by addition of IPTG in a concentration intervals of 0.05,

0.2, 0.5, 0.75, and 1 mM of protein growth or through utilizing auto-induction media.

Protein expression was allowed to be induced for a time course of 3, 6 or 18 hours at temperatures of 15, 28 or 37 °C. Cells cultures were stored in a 96-well format at -80°C for future analysis. Stored cells were collected by centrifugation and resuspended in phosphate buffer pH 8.0 containing lysozyme. The samples were subjected to five freeze- thaw cycles to lyse cells, spun at 3700 rpm to remove cellular debris. Supernatants diluted in 0.1M HEPES pH 7.5, 0.1% CHAPS, 10% sucrose, and 5mM β-Me. buffer were then analyzed for activity using the fluorogenic substrate VEID-AMC. Activity was only observed for control protein caspase-6, indicating that changing the bacterial strain induction conditions such as cell density, temperature, and [IPTG] did not improve

YCA1 expression levels or cleavage patterns to result in an active enzyme judged by protease activity.

C.3. Discussion

Yeast caspase 1, a homologue of human caspases, would be a good surrogate for studying regulation of YCA1 in the yeast apoptotic cascade. A genetic construct containing the YCA1 gene was created for expression of YCA1 in E.coli. Upon analysis of expression, induction, and purification conditions, minimal amounts of the inactive full length form of protein was obtained. In addition, a non-homogeneous mixture of cleavage products was observed, which did not possess any enzyme activity. Due to this lack of

185

activity, the observed cleaved forms of the enzyme are thought to be breakdown products which occurred during expression or purification by an endogenous E.coli protease. This would also reflect on the low yield that was obtained. Due to these issues, further progression of this project was halted.

Future studies could include purifications in which protease inhibitors such as

PMSF are utilized. These inhibitor cocktails, however, will also target the enzyme of interest, limiting analysis to only structural based methods in which a peptide is bound to the active site. Additionally, avenues to express YCA1 in its native system, yeast, could also be explored. Although over-expression of YCA1 in yeast has been shown to induce apoptotic cell death, potential purification from natural abundance or leaky expression would most likely lead to the proper cleavage of the enzyme resulting in the active enzyme.

C.4. References

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4. Herker, E. et al. Chronological aging leads to apoptosis in yeast. J Cell Biol 164, 501-507, (2004).

5. Uren, A. G. et al. Identification of paracaspases and metacaspases: two ancient families of caspase-like proteins, one of which plays a key role in MALT lymphoma. Mol Cell 6, 961-967, (2000).

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6. Madeo, F. et al. A caspase-related protease regulates apoptosis in yeast. Mol Cell 9, 911-917, (2002).

7. Khan, M. A., Chock, P. B. & Stadtman, E. R. Knockout of caspase-like gene, YCA1, abrogates apoptosis and elevates oxidized proteins in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 102, 17326-17331, (2005).

8. Bettiga, M., Calzari, L., Orlandi, I., Alberghina, L. & Vai, M. Involvement of the yeast metacaspase Yca1 in ubp10Delta-programmed cell death. FEMS Yeast Res 5, 141-147, (2004).

9. Reiter, J., Herker, E., Madeo, F. & Schmitt, M. J. Viral killer toxins induce caspase-mediated apoptosis in yeast. J Cell Biol 168, 353-358, (2005).

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