Regulation of Caspase-9 by Natural and Synthetic Inhibitors

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5-2012

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

Kristen L. Huber University of Massachusetts Amherst

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REGULATION OF CASPASE-9 BY NATURAL AND SYNTHETIC INHIBITORS

A Dissertation Presented

by

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


A Dissertation Presented

by

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

v

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

vi

ABSTRACT


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

vii

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.

viii

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

ix

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

x

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

xi

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


5.4.2. 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


5.4.6. 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

xii

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

xiii

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

xiv

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

xv

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

1

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

2

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

3

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

4

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

5

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.

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.

6

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

properties: Km in the range of 4 to 408 μM and a kcat of 0.2 to 9.1 s-1

37

. 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

7

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

thought to be involved in substrate differentiation38-40

. 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

.

8

Executioner caspases prefer to bind acidic amino acids in their S4 pocket41

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

mechanisms (Fig. 1.2 B) as that of other cysteine proteases (reviewed in43

) 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

9

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.

10

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

11

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

12

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

active as dimers. The dimerization constant, KD, for caspase-9 is in the μM range62,63

,

enforcing a fairly weak complex considering cellular concentrations of ~20 nM54

. In

comparison, KD of the executioner caspases is <50 nM64

. 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

13

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

14

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

distal from the active site of the enzyme74

with a Ki of 0.2-5 nM is present73,75

. In the case

of caspase-9, the BIR3 domain, residues 251-350, causes inhibition of the enzyme73,76

with a Ki of 10-20 nM76-78

(Fig 1.4). BIR3 binds to the dimerization interface of the

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

.

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.

15

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

16

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).

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.

17

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

an IC50 of 25 nM92

, 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

,

18

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.

19

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

20

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.

21

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30

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.

31

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 interactions

2-5,

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 targets

19-22. 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

32

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

33

Figure 2.1 aPP structure. (A) aPP backbone (lightest

gray) has a hydrophobic core created by three

prolines located on the poly-proline helix (medium

gray) and various residues on the α-helix (dark gray).

(B) aPP’s internal hydrogen bond network,

highlighted in dashed lines, is responsible for the

impressive structural stabilization of a peptide that is

just 36 amino acids long.

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

sequence34,35

or to a c-terminal

truncation33,36

.

A common difficulty with peptide

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.

34

Figure 2.2 Schematic representation of expression vector,

pET32-Peptide. The T7 promoter, a fusion partner

thioredoxin (TrxA), 6xHis tag, human rhinovirus 3C

cleavable linker, gene for peptide, and ampicillin resistance

are shown.

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

human rhinovirus 3C protease

cleavage site (LEVLFQ) and

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

35

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

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.

36

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.

37

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 2.4 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.

38

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 2.5 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.

39

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.

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.

40

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

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

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.

41

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

42

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

43

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

44

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

45

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

46

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

47

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

48

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

49

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).

50

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


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).

51

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).

52

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).

53

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).

54

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

55

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

56

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

57

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

58

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.

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.

59

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

60

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

61

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-

62

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

peptides34,49-51

. We

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,

Figure 3.2 Sequences of the α5 region of XIAP-BIR3 (BIR3) and

peptides 1-9. * represents aminoisobutyric acid (Aib)

63

the location of the CARD

domain remains

uncharacterized.

Due to the size of

the CARD and its ability to

enhance caspase-9’s basal

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

any way with CARD. We

tested both C9FL (Fig 3.4

A) and C9ΔN (Fig 3.4 B)

Figure 3.3 Circular dichroism spectra of the native Peptide 5 and

the Aib-stabilized Peptide 8. These spectra demonstrate that the

presence of Aib increases the helicity of the peptides.

Figure 3.4 Native and Aib-stabilized peptides show some

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

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.

64

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

65

Table 1: Overall Inhibition of Caspase-9 Full Length by Peptides

Peptide % Inhibitiona IC50 appb

(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).

66

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

activation, whereas the

incubated peptides were

(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 3.5 Peptide 2 non-specifically activates caspase-9. (A) Peptide

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

aged peptide non-specifically activates both caspase-9 and caspase-7

in a manner similar to the polymeric crowding agent PEG 8,000

67

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

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.

68

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

69

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.

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.

70

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.

71

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

3.9A). Aliphatic

staples have been

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

Figure 3.9 Aliphatic stapled peptide design. (A) Modeled interactions of α5-

derived peptides stabilized by aliphatic staples (green) with caspase-9

(purple). (B) Synthetic route for production of the pentyl alanine amino acid

used to generate the aliphatic peptide staples

72

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

27 amino acids Y338 and I342 from

BIR3, which point away from the

caspase-9 dimer interface, were both

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-

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

acids as listed. (B) Ring-closing metathesis reaction

performed on the unstapled Peptide 27 to form the

aliphatic stapled Peptide 27 with an 8-carbon

macrocyclic linker. The structure of aliphatically

stapled Peptide 28 contains an 11-carbon macrocyclic

linker

73

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.

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

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.

74

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 2200Å

2

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

75

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

76

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.

77

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.


3.4.1. 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

78

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

79

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)

80

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

(0.37 g, 70%). 1H 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

81

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

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

82

(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

obtained as 90% isolated yield (1.2 gm). 1H 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)

1H 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

1H 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-

83

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

1H 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.

84

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

85

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

.

86

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domain. Nature 408, 1004-1008, (2000).

72. Stennicke, H. R. & Salvesen, G. S. Caspases: preparation and characterization.

Methods 17, 313-319, (1999).

73. Rotonda, J. et al. The three-dimensional structure of apopain/CPP32, a key

mediator of apoptosis. Nat Struct Biol 3, 619-625, (1996).

74. Garcia-Calvo, M. et al. Inhibition of human caspases by peptide-based and

macromolecular inhibitors. J Biol Chem 273, 32608-32613, (1998).

75. Witkowski, W. A. & Hardy, J. A. L2' loop is critical for caspase-7 active site

formation. Protein Sci 18, 1459-1468, (2009).

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76. Barany, G., Kneib Cordonier, N. & Mullen, D. G. Solid phase peptide synthesis: a

silver anniversary report*. International Journal of Peptide and Protein Research

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fluorenylmethoxycarbonyl amino acids. International Journal of Peptide and

Protein Research 35, 161-214, (1990).

78. Stennicke, H. R. & Salvesen, G. S. Biochemical characteristics of caspases-3,-6,-

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79. Schrödinger, L. The PyMOL Molecular Graphics System, Version 1.3, (2010).

94

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

95

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

96

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.

97

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.

98

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

C), further suggesting that zinc-

mediated inhibition does not

influence the overall structure of

Figure 4.3 Zinc does not alter the biophysical properties of caspase-

9. (A, B) Native gel analysis (A) and size exclusion

chromatography (B) both indicate zinc does not alter the

oligomeric state of caspase-9 from monomer to dimer like the

substrate-mimic z-VAD-FMK. (C) CD spectra of caspase-9 full

length in the presence and absence of ZnCl2 indicate that zinc does not alter the caspase-9 secondary structure.

99

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).

100

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

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,

Table 4.1 Zinc bincing as monitored by

ICP-OES.

101

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.

102

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

that zinc inhibition is most potent in caspase-6 (IC50 = 0.3 μM )3,22

, 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

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.

103

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

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

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.

104

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.



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.

105

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


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

106

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

107

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.


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

108

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 spectro-

photometer. 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.

109

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

110

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

111

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.

112

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,

114

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

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

115

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

.

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

Figure 4.8 Diffraction image of apo wild type caspase-9 crystals soaked in a 20% PEG 400

cryoprotectant for one hour.

116

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

117

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).

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.

118

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).


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).


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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

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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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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

.

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.

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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

125

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

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).

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.

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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

cleaved at the intersubunit linker, is

Figure 5.3 Thermal denaturation analysis and circular

dichroism spectrum of monomeric, cleaved, caspase-

9 (A) full-length, (B) ΔCARD, and (C) CARD only.

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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

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.

Figure 5.4 Thermal denaturation analysis and circular dichroism spectrum of dimeric, cleaved,

caspase-9 (A) full-length, (B) ΔCARD.

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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

CARD and core domains occurs, which

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

Figure 5.5 Overlay of the circular dichroism

spectrum for full-length caspase-9 (C9F) in

the presence and absence of active site ligand

z-VAD-FMK

129

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

by the independent three-state

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

is in an ordered state.

Figure 5.6 Thermal denaturation analysis and circular

dichroism spectrum of (A) monomeric, uncleaved

caspase-9 and (B) monomeric, cleaved caspase-9.

Figure 5.7 SDS-PAGE analysis of full-length

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.

130

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.

131

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

and the large subunit of caspase-9’s

catalytic core (Fig 5.9). This variant

Figure 5.9 Characteristics of the Ser-Gly linker

extension caspase-9 variant. Kinetic parameters,

thermal denaturation analysis, and circular

dichroism spectra of the Ser-Gly linker extension

caspase-9 variant.

132

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,

133

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

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.

134

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

Table 5.2 Measured kinetic parameters of caspase-9 variants designed to disrupt the activation affect

of the CARD domain.

135

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

136

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.

137

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.



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

138

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


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

139

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

140

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

141

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

142

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.


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

143

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.


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

144

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.

145

5.5. References

1. Zhai, D. et al. Vaccinia virus protein F1L is a caspase-9 inhibitor. J Biol Chem

285, 5569-5580, (2010).

2. Oztas, P. et al. Caspase-9 expression is increased in endothelial cells of active

Behcet's disease patients. Int J Dermatol 46, 172-176, (2007).


lung cancer. Oncol Rep 11, 797-802, (2004).


J Biol Chem 274, 8359-8362, (1999).


Genes Dev 13, 3179-3184, (1999).

6. Zou, H., Li, Y., Liu, X. & Wang, X. An APAF-1.cytochrome c multimeric

complex is a functional apoptosome that activates procaspase-9. J Biol Chem 274,

11549-11556, (1999).


caspase-9 by dimerization. Mol Cell 22, 269-275, (2006).

8. Shiozaki, E. N., Chai, J. & Shi, Y. Oligomerization and activation of caspase-9,

induced by Apaf-1 CARD. Proc Natl Acad Sci U S A 99, 4197-4202, (2002).

9. Acehan, D. et al. Three-dimensional structure of the apoptosome: implications for

assembly, procaspase-9 binding, and activation. Mol Cell 9, 423-432, (2002).

10. Salvesen, G. S. & Dixit, V. M. Caspase activation: the induced-proximity model.

Proc Natl Acad Sci U S A 96, 10964-10967, (1999).



Natl Acad Sci U S A 98, 14250-14255, (2001).

12. Shi, Y. Caspase activation: revisiting the induced proximity model. Cell 117, 855-

858, (2004).

13. Manns, J. et al. Triggering of a novel intrinsic apoptosis pathway by the kinase

inhibitor staurosporine: activation of caspase-9 in the absence of Apaf-1. FASEB J

25, 3250-3261, (2011).

14. Vaidya, S. & Hardy, J. A. Caspase-6 latent state stability relies on helical

propensity. Biochemistry 50, 3282-3287, (2011).

146

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).



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).


7, and -8. J Biol Chem 272, 25719-25723, (1997).

147

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

148

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

149

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 dehydrogenase

10, and the caspases

11, 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

150

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

151

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

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

Figure 6.1 Structure of caspase-9 with mapped regulatory

surfaces.

152

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.

153

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).

154

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).


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).

156

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).

157

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)

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

Figure A.1 Molecular structure

of FlAsH-EDT2.

158

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

159

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.

160

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

161

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

162

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




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.

163

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).

164

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

165

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

caspase-9 large subunit only cross

reacts with the caspase-9 antibody

and is cysteine rich explaining the

Figure A.5. Western blot analysis of FlasH binding to full-

length caspase-9. Analysis was performed using an

antibody specific for the small subunit of caspase-9.

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).

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).

166

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

167

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

168



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170

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

171

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

172

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

The caspase-9 full-length

Figure B.1 Model of the designed caspase-9 disulfide

cross linked variant.

173

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




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.

174

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.

175

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

176

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

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.

177

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


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(2003).


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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).

180

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 aging

4. 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

gene in S. cervisiea has been shown to abolish an H2O2 induced form of apoptosis7. 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

181

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

182

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.

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

Figure C.1. Schematic representation of expression vector, pYCA1. The T7 promoter, YCA1 gene,

6xHis tag, and ampicillin resistance are shown.

183

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

184

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

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

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.

185

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

186

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

1. Madeo, F., Frohlich, E. & Frohlich, K. U. A yeast mutant showing diagnostic

markers of early and late apoptosis. J Cell Biol 139, 729-734, (1997).

2. Carmona-Gutierrez, D. et al. Apoptosis in yeast: triggers, pathways, subroutines.

Cell Death Differ 17, 763-773, (2010).

3. Madeo, F. et al. Oxygen stress: a regulator of apoptosis in yeast. J Cell Biol 145,

757-767, (1999).

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).

187

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).

10. Watanabe, N. & Lam, E. Two Arabidopsis metacaspases AtMCP1b and

AtMCP2b are arginine/lysine-specific cysteine proteases and activate apoptosis-

like cell death in yeast. J Biol Chem 280, 14691-14699, (2005).

11. Petty, K. J. Metal-Chelate Affinity Chromatography. Vol. 2 (John Wiley and

Sons, Inc., 2000).

188

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