PURIFICATION AND CHARACTERIZATION OF TWO MEMBERS OF THE

PROTEIN TYROSINE PHOSPHATASE FAMILY: DUAL SPECIFICITY

PHOSPHATASE PVP AND LOW MOLECULAR WEIGHT PHOSPHATASE WZB

By

Paula A. Livingston

A Thesis Submitted to the Faculty of

The Charles E. Schmidt College of Science

in Partial Fulfillment for the Degree of

Master of Science

Florida Atlantic University

Boca Raton, Florida

December 2009 i

ACKNOWLEDGMENTS

The author wishes to thank her thesis advisor, Dr. Stefan Vetter, and the members

of her committee, Dr. Estelle Leclerc and Dr. Predrag Cudic, for their advice and support

throughout her years at Florida Atlantic University. The author also wishes to thank her family for their unending support and encouragement without which this would not have been possible. The author would also like to thank her nephew and niece, Griffin and

Skye, for inspiring her throughout her entire graduate experience.

iii

ABSTRACT

Author: Paula A. Livingston

Title: Purification and Characterization of Two Members of the Protein Tyrosine Phosphatase Family: Dual Specificity Phosphatase PVP and Low Molecular Weight Phosphatase WZB

Institution: Florida Atlantic University

Thesis Advisor: Dr. Stefan W. Vetter

Degree: Master of Science

Year: 2009

Two protein tyrosine phosphatases, dual specificity phosphatase PVP and low molecular weight phosphatase WZB were purified and characterized. PVP was expressed as inclusion bodies and a suitable purification and refolding method was

devised. Enzyme kinetics revealed that p-nitrophenylphosphate and β-naphthyl phosphate were substrates with KM of 4.0mM and 8.1mM respectively. PVP showed no

reactivity towards phosphoserine. Kinetic characterization of WZB showed that only p- nitrophenylphosphate was a substrate with no affinity for β-naphthyl phosphate and phosphoserine. Optimal conditions for activity with PNPP were found at a pH of 5 with

-1 -1 -1 a KM of 1.1mM, kcat of 35.4s and kcat/KM of 32.2s mM . Inhibition studies showed that

phosphate, fluoride, and molybdate were competitive inhibitors with Ki of 3.2mM,

71.7mM, and 50.4μM respectively and hydrogen peroxide abolished activity. Active site

mutants of WZB Cys9Ser and Asp115Asn showed no activity.

iv

PURIFICATION AND CHARACTERIZATION OF TWO MEMBERS OF THE

PROTEIN TYROSINE PHOSPHATASE FAMILY: DUAL SPECIFICITY

PHOSPHATASE PVP AND LOW MOLECULAR WEIGHT PHOSPHATASE WZB

List of Tables ...... viii

List of Figures ...... ix

Chapter 1 – Introduction ...... 1

1.1 – The protein phosphatase superfamily ...... 1

PPP family ...... 2

PPM family ...... 2

PTP family ...... 3

Aspartate based family ...... 4

1.2 – Overview of protein tyrosine phosphatases ...... 4

1.3 – Classification of PTPs ...... 5

Phosphotyrosine specific PTPs ...... 6

Dual specificity phosphatases ...... 7

Cdc25 phosphatases ...... 9

Low molecular weight phosphatases ...... 10

1.4 – Biological functions of PTPs ...... 11

PTPs and cancer ...... 11

PTPs and Alzheimer’s...... 13

v PTPs and diabetes ...... 14

1.5 – Molecular structure of PTPs ...... 17

1.6 – Mechanism of catalysis of PTPs ...... 20

1.7 – PTPs in eukaryotes and prokaryotes ...... 23

1.8 – Poxvirus phosphatases ...... 26

1.9 – Low molecular weight phosphatase WZB in Escherichia coli ...... 27

Chapter 2 – Materials and Methods ...... 29

2.1 – Instrumentation and Buffers ...... 29

2.2 – Transformation of plasmid into expression hosts ...... 30

2.3 – Cell growth and expression of phosphatases ...... 32

2.4 – Isolation of inclusion bodies ...... 36

2.5 – Nickel affinity chromatography ...... 37

2.6 – Ion exchange chromatography ...... 38

2.7 – Gel filtration chromatography...... 38

2.8 – Refolding matrix design ...... 39

2.9 – Refolding protocol ...... 40

2.10 – Enzyme kinetics ...... 41

2.11 – Fitting of enzyme kinetic data...... 42

2.12 – Synthesis of non-detergent sulphobetaines (NDSBs) ...... 43

2.13 – Molecular cloning of WZB mutants ...... 45

Chapter 3 – Results: cell growth, expression, purification and kinetic characterization

of PVP ...... 46

3.1 – Growth and expression of PVP ...... 46

vi 3.2 – Purification of PVP ...... 48

3.3 – Refolding of His6-tagged PVP ...... 51

3.4 – Enzyme kinetics of PVP ...... 56

Chapter 4 – Results: cloning, cell growth, expression, purification and kinetic

characterization of WZB ...... 60

4.1 – Growth and expression of WZB ...... 60

4.2 – Purification of WZB ...... 61

4.3 – Enzyme kinetics of WZB ...... 63

4.4 – Inhibitory effects on WZB ...... 68

4.5 – Comparison of enzymatic activity between the two forms of WZB ...... 76

4.6 – Enzyme activity of WZB mutants...... 78

Chapter 5 – Summary ...... 80

References ...... 83

vii

LIST OF TABLES

Table 1 – Cdc25 overexpression found in different cancers...... 13

Table 2 – Prokaryotic PTPs ...... 24

Table 3 – DNA and amino acid sequence of both forms of PVP ...... 35

Table 4 – DNA and amino acid sequence of WZB ...... 36

Table 5 – Initial refolding buffer systems for His6-tagged PVP ...... 39

Table 6 – Additional refolding buffer systems for His6-tagged PVP ...... 40

Table 7 – Enzymatic results of PVP with various substrates ...... 56

Table 8 – Results of inhibition studies with WZB ...... 69

Table 9 – Comparison of WZB activity between two forms of WZB ...... 76

viii

LIST OF FIGURES

Figure 1 – Protein phosphorylation as regulated by kinases and phosphatases ...... 1

Figure 2 – Catalytic mechanism of PP1 (member of PPP family) ...... 3

Figure 3 – Classification of the PTP superfamily ...... 7

Figure 4 – Cdc25 phosphatases and cyclin-dependent kinases ...... 10

Figure 5 – Cdc25 phosphatases in cell cycle control ...... 12

Figure 6 – PTPs in insulin signaling ...... 16

Figure 7 – X-ray structure of PTP1B ...... 18

Figure 8 – Structure of the P-loop from bovine LMW-PTP ...... 19

Figure 9 – Amino acid sequence comparison between WPD loop in different PTPs ...... 20

Figure 10 – Catalytic mechanism of PTPs ...... 21

Figure 11 – Suggested transition state for phosphorylation/dephosphorylation in PTPs ..23

Figure 12 – Map of pET101 expression vector ...... 31

Figure 13 – pET vector backbone ...... 32

Figure 14 – Isolation of inclusion bodies scheme ...... 37

Figure 15 – Synthesis scheme for all NDSBs ...... 43

Figure 16 – Yield and structure of NDSBs after synthesis ...... 44

Figure 17 – SDS-PAGE after isolation of inclusion bodies ...... 49

Figure 18 – Chromatogram of PVP His6-tagged after nickel affinity column ...... 50

Figure 19 – Chromatogram of PVP non-tagged after ion exchange column ...... 50

ix Figure 20 – SDS-PAGE of purified PVP His6-tagged and non-tagged ...... 51

Figure 21 – Enzymatic results after first refolding screen ...... 53

Figure 22 – Enzymatic results after second refolding screen ...... 54

Figure 23 – Results from dilution of NDSB256 in refolding buffer...... 56

Figure 24 – Michaelis-Menten graph of PVP-PNPP kinetics ...... 57

Figure 25 – Michaelis-Menten graph of PVP-pNaph kinetics...... 58

Figure 26 – Structure of β-naphthyl phosphate and p-nitrophenyl phosphate ...... 59

Figure 27 – Chromatogram of eluted WZB after nickel affinity column and gel

filtration column ...... 62

Figure 28 – SDS-PAGE gel of eluted WZB from nickel affinity column and gel

filtration column ...... 63

Figure 29 – Buffer effect on WZB enzyme activity ...... 64

Figure 30 – Kinetic parameters of WZB with PNPP as substrate over pH range of 4.5

to 7.5 ...... 66

Figure 31 – Michaelis-Menten plot of WZB-PNPP kinetics ...... 67

Figure 32 – Lineweaver Burk and Dixon plots of the inhibitor acting on WZB ...... 72

Figure 33 – Tetrahedral structure of phosphate, vanadate, and molybdate ...... 73

Figure 34 – Effect of hydrogen peroxide on WZB activity ...... 75

Figure 35 – Oxidation of catalytic cysteine followed by disulfide bond formation with

neighboring cysteine ...... 76

Figure 36 – Michaelis-Menten plots for the two forms of WZB ...... 77

Figure 37 – Role of cysteine, arginine, and aspartic acid in catalytic mechanism ...... 78

Figure 38 – SDS-PAGE of Cys9Ser and Asp115Asn WZB mutants ...... 79

x

CHAPTER 1 – INTRODUCTION

1.1 – The protein phosphatase superfamily

Many cellular processes are regulated by reversible protein phosphorylation. Two types of enzymes are responsible for this: protein kinases which attach phosphate groups to amino acid side chains and protein phosphatases which dephosphorylate the amino acids. The addition or removal of this phosphate group can cause diverse changes in cellular life (figure 1). It has been Figure 1 – Protein phosphorylation as regulated by shown that metabolic processes, cellular signaling, gene kinases and phosphatases regulation and cell cycle control are regulated by protein phosphorylation (D. D. Barford

1998). The diversity and importance of protein phosphorylation can be seen in the large portion of intracellular proteins (30%) that are subject to reversible phosphorylation. In eukaryotes, the combined genes that encode for protein kinases (~2000) and phosphatases

(~1000) constitute 3-4% of their genomes (Barford D. D. 1998, Barford D. 1995). In

humans, there are 518 genes encoding for protein kinases (Manning 2002) and 148 genes

encoding protein phosphatases (Kerk 2008).

Proteins can be phosphorylated on nine amino acids – tyrosine, threonine, serine,

histidine, cysteine, arginine, lysine, aspartate and glutamate. Phosphorylation on 1 tyrosine, threonine and serine are the most common in eukaryotes, with phosphorylation

on histidine and aspartate common in bacterial two-component systems. The enzymes

that dephosphorylate these amino acids are separated into groups (explored later) based

on unique catalytic signature motifs and substrate specificity (Moorehead 2009).

Dephosphorylation at serine and threonine residues is accounted for by two groups of protein phosphatases: the classic phosphoserine/phosphothreonine phosphoprotein phosphatases (PPPs) and the metallo-dependent protein phosphatases

(PPMs). The two families are unrelated in sequence and are encoded by two separate gene families (Moorehead 2009, Burke 1998). Though the families are unrelated in primary structure, it has been found that members of both families have similar three- dimensional structures and similar catalytic mechanism while members of the other two protein phosphatase families (the protein tyrosine phosphatases and the aspartate based catalysis phosphatases) have differing three-dimensional structure and catalytic mechanism (Zhang W. 2004). The three-dimensional structure of the PPP and PPM families consist of a core of mixed β-sheets that form a β-sandwich structure. The main difference between the two families is the metal ion found in the binuclear metal ion center. Members of the PPM have Mg2+ or Mn+ ions while the PPP family has Fe2+ and

Zn2+/Mn2+ (Barford D. D. 1998, Ingebritsen 2004, Zhang W. 2004). The catalytic

mechanism for the two families of phosphatases occurs via a single step in which a water

molecule bound by the metal acts as a nucleophile that attacks the phosphate group of the

phosphoserine/phosphothreonine (refer to figure 2). The metal acts as a Lewis acid to

enhance nucleophilicity of the water, as well as, electrophilicity of the phosphorous atom

by coordination of the oxyanions of the phosphate group. Additionally, a His sidechain

2 of the catalytic site acts as a general acid to protonate the leaving oxygen (Zhang W.

2004, Mildvan 1997).

Figure 2 – Catalytic mechanism of PP1 (member of PPP family) (Ingebritsen 2004)

Protein tyrosine phosphatases (PTPs) include phosphatases that dephosphorylate phosphotyrosine substrates. This large superfamily of phosphatases has many members, which can be subdivided into further groups, with little sequence similarity but all maintain a common catalytic motif, C(X)5R. This similarity allows all members of this family to follow the same catalytic mechanism, and though they also, at times, dephosphorylate phosphoserine and phosphothreonine residues like members of the PPP and PPM families, the two catalytic mechanisms are dissimilar (Barford D. D. 1998,

Zhang Z. Y. 1998). Members of the PTP family use no metal cations in their catalysis and its catalytic mechanism involves a two step reaction involving the formation of a phosphoenzyme (a complex between the phosphate and the enzyme) intermediate that is

3 then hydrolyzed by water (Zhang Z. Y. 1998). Further insights into the catalytic mechanism of the PTP family will be explained later on.

The last group of the protein phosphatase family is the newly discovered phosphatases that use aspartate based catalysis. The catalytic motif of this family of phosphatases is led by an aspartic acid – DXDXT/V. The members of this family can be classified into two groups that belong to the haloacid dehalogenase (HAD) family of enzymes (Moorehead 2009, Kerk 2008). Another feature of phosphatases includes the addition of other domains to the catalytic module. This additional domain(s) increase the specificity of function of the phosphatase. One of the domains commonly found is the

BRCT domain which may function in directing the enzyme to the phosphorylated substrates (Moorehead 2009).

1.2 – Overview of protein tyrosine phosphatases (PTPs)

Cellular signaling is often controlled by signal transduction processes in which extracellular signals are converted into intracellular signals that are taken to the nucleus and an appropriate cellular response is generated. A common mode of signal transduction involves the phosphorylation/dephosphorylation of proteins at the cellular level. This addition/removal of phosphate has been known to create certain patterns that control protein-protein interactions as well as enzyme activity. A common and important signaling process involves the phosphorylation of tyrosine residues (Wang 2003).

Phosphorylation of tyrosine residues is controlled by two opposing forces: the protein tyrosine kinases (PTKs) and the protein tyrosine phosphatases (PTPs). Both classes of

4 enzymes are largely complex and diverse with PTPs rivaling the complexity of PTKs.

Unlike PTKs, which share sequence similarity with protein serine/threonine kinases,

PTPs share no sequence homology with the serine/threonine phosphatases (Chiarugi

2005). PTPs can have both positive and negative effects on signaling pathways and they play crucial roles in mammalian tissues and cells. Additionally, deregulation of PTP activity can lead to pathogenesis of human diseases (Burke 1998, Chiarugi 2005, Zhang

Z. 2002). This large superfamily of signaling enzymes has been linked to many cellular functions including the regulation of the cell cycle, mitogenesis, cell-cell interactions, metabolism, gene transcription, and immune response (Burke 1998). Since phosphorylation of tyrosine residues is governed by both PTPs and PTKs, characterization and understanding of PTPs as well as PTKs in healthy and disease cells must be achieved. As PTKs, PTPs are now molecular targets for understanding pathogenesis of certain human diseases as well as medicinal targets (Zhang Z. 2002).

1.3 – Classification of PTPs

Protein tyrosine phosphatases make up a large superfamily (figure 3) whose substrates range from protein (the most common substrate) to phosphoinositides and mRNAs (substrates specific for certain protein tyrosine phosphatases). A key feature of this superfamily is the PTP active site signature motif, C(X)5R. Each PTP has this conserved catalytic motif comprised of ~240 amino acid residues. This constitutes to 30-

40% identity between individual PTPs. Individual PTPs can be determined according to

5 their unique noncatalytic domains that are connected to either end of the catalytic

domain. These domains usually have some regulatory roles (Tonks 1996).

The first group of the PTP family is the classical phosphotyrosine (pTyr) specific

PTPs. This family has approximately 40 members all with the PTP catalytic domain within an ~250 amino acid residues. In relation to their subcellular location and similarities in the structural and functional domains in the regulatory segments, the pTyr

PTPs can be further divided into intracellular and receptor-like PTPs (figure 3) (Tonks

2003, Zhang Z. 2002). The intracellular PTPs have a single catalytic domain and several extensions at the amino and carboxyl terminus which may include the SH2 (Src homology 2) domain that could play roles in targeting and regulation through phosphate interactions with other proteins (Ninfa 1994). An example of an intracellular PTP is

PTP1B which contains a 35 residue carboxyl-terminal extension that is necessary to target PTP1B to the cytoplasmic face of the endoplasmic reticulum. This may be the cause for PTP1B to inactivate receptor PTKs and in the prevention of the premature activation of kinases before reaching their final destination in the cytoplasmic membrane

(Wang 2003). The receptor-like PTPs are characterized by generally having an extracellullar, ligand binding domain, a single transmembrane region, and one or two cytoplasmic PTP domains. The structural features of the extracellular domain further classifies this group into six subtypes. These include domains that resemble immunoglobins, fibronectin III repeats, carboxyl anhydrase, MAM (Meprin-Xenophus

A2-Mu) domains or cysteine rich regions. These structural differences led to the suggestion that receptor-like PTPs may be involved in cell-cell or cell-matrix adhesion

(Wang 2003). Additionally, the role of the tandem cytoplasmic PTP domains is not

6 completely known, however, it may be that there are distinct regulatory functions of the two domains. It is believed that most of the phosphatase activity lies in the proximal PTP domain and the second domain is believed to be inactive (Wang 2003).

Figure 3 - Classification of PTP superfamily (Z. Zhang 2003)

The second group of the PTP family is the dual specificity phosphatases (DSPs).

Similarly to the classical pTyr PTPs, the DSPs can also be further subdivided based on their substrate specificity. This large group of PTPs, with ~50 members, is characterized by their ability to dephosphorylate phosphoserine (pSer), phosphothreonine (pThr), and phosphotyrosine (pTyr) substrates. The DSPs share very little sequence similarity with the classical pTyr PTPs except for conservation in the catalytic domain. Additionally, the

7 catalytic core of the DSPs has a more open active site than the classical pTyr PTPs which

account for their ability to fit the different phosphorylated substrates (Tonks 2003,

Stauffacher 2000). The first group of the DSPs is the VH1-like based on VH1, a protein

from the open reading frame of the small pox virus, the founding member of the DSPs.

The group contains VHR, the first DSP identified in humans, and several MAP kinase

phosphatases (MKPs), which show high affinity towards MAP kinases. Members of this

subgroup contain a single catalytic domain and MKPs have a variable terminal domain

that accounts for their specificity towards specific MAP kinases (Wang 2003). The

second group of DSPs is known as the Cdc14-like phosphatases and include Cdc14A and

B, and KAP. Both Cdc14 and KAP are involved in cell cycle regulation with KAP

dephosphorylating of pTyr of cyclin dependent kinases which lead to inactivation of the

kinase activity. Cdc14 phosphatases play critical roles in events of the cell cycle,

especifically the transition from late anaphase to G1 (Wang 2003). The next group of the

DSPs, the phosphoinositide phosphatases, include two groups of phosphatases, PTEN

and Myotubularin, whose substrates are not proteins. The PTEN phosphatase is a tumor

suppressor that is mutated is many human tumor types. The specificity of PTEN is for

the 3 position of the inositol ring in phosphoinositide signaling molecules. There have

been several PTEN homologs found, one of which is a novel phosphoinositide 3- phosphatase, TPIP. TPIP occurs in two forms, TPIPα and TPIPβ. TPIPα has transmembrane domains and it is found in the endoplasmic reticulum while TPIPβ is cytosolic and does not appear to have any phosphatase activity. It is believed that TPIPα

plays a role in signaling on the endoplasmic reticulum and might be a functional

homologue of PTEN in tumor suppresion (Walker 2001). Myotubularin is produced

8 from a gene mutation in X-linked myotubular myopathy, a congenital muscle disorder

generalized by muscle weakness. This phosphatase dephosphorylates the 3 phosphate of

the lipid phosphate, phosphatidyl 3-phosphate (Laporte 1996). There are approximately

10 genes that encode for myotubularin like phosphatases in humans. Both PTEN and

Myotubularin can antagonize the phosphoinositide 3-kinase pathway, PTEN in cellular

growth and Myotubularin in vacuolar protein sorting (Wang 2003). The last group of

DSPs are grouped into the RNA triphosphatases. Similarly to the phosphoinositide phosphatases, the RNA triphosphatases have weak affinity for protein substrates but great activity towards mRNA 5’-triphosphates. These phosphatases are important in the first step in 5’cap formation in mRNA processing, by the cleavage of the γ-phosphate in pre-

mRNA that forms a diphosphate that may then be capped with GMP by RNA

guanyltransferase (Wang 2003). One of these phosphatases, CEL-1 RNA triphosphatase,

is interesting because it contains two domains, a guanyltransferase domain and a PTP

domain. The PTP domain shows no activity towards pTyr substrate but it maintains the

nucleophilic cysteine necessary for nucleophilic attack. It is hypothesized that although

CEL-1 RNA triphosphatase shows no activity towards pTyr substrate, the

dephosphorylation of the nascent pre-mRNA occurs by the same catalytic mechanism of

other PTPs as seen by mutation studies that abolish activity when the catalytic cysteine is

mutated (Takagi 1997).

The next group of the PTP family is the Cdc25 phosphatases. Cdc25

phosphatases play important roles in the cell division cycle and the transitions between

the various phases by activating specific cyclin-dependent kinases (CDKs) through

dephosphorylation (Ducruet 2005, Buotros 2006). Members of the Cdc25 phosphatases

9 are more structurally diverse than other members of the PTP family. They possess

distinct differences in the primary and tertiary structure but they remain a part of the PTP

family because of the conserved PTP signature motif (Wang 2003). There are three human Cdc25 phosphatases (Cdc25A, Cdc25B, and Cdc25C) that are responsible for the dephosphorylation of CDKs on pThr14 and/or pTyr15 residues and the three isoforms are responsible for regulating cell cycle control and progression (Rudolph 2007).

Figure 4 - Cdc25 phosphatases and cyclin-dependent kinases (Ducruet 2005)

The last group of the PTP family is the low molecular weight phosphatases

(LMW-PTPs). These cytoplasmic PTPs have a molecular weight of ~18,000 Da and they

share little sequence homology with other members of the PTP family except for the

conserved PTP catalytic domain. The catalytic signature motif of the LMW-PTPs is located near the C-terminus of the catalytic domain while in the other members of the

PTP family, the signature motif is most oftern found in the N-terminus part of the catalytic domain (Stauffacher 2000). These phosphatases have strong affinity towards pTyr substrates with little affinity towards pSer and pThr substrates. In addition, it has been recently determined that LMW-PTPs show high activity towards the 5’-phosphate 10 of the flavin mononucleotide, although its significance has not been uncovered (Van

Etten 2003).

1.4 - Biological functions of PTPs

As mentioned earlier, PTPs have positive and negative effects on signaling

pathways. PTPs can act as “on” and “off” switches for events leading to signal

transduction pathways. The results of PTPs rely on the effect that they exert on PTK

signaling. Since PTPs are the natural antagonists of PTKs, PTPs can act negatively on

pTyr signaling pathways (because the dephosphorylation of the tyrosine residue may lead

to incapacitation of the signal transduction pathway). Alternatively, PTPs may act

positively of PTK signaling by assisting signal transduction. This can occur through the

repriming of the system or by the activation of downstream PTKs through

dephosphorylation of certain tyrosine residues (Burke 1998). PTPs, through its many

members, can have regulatory effects on many human diseases ranging from cancer,

Alzheimer’s and autoimmune diseases (Zhang Z. Y. 1998).

Cancer is one of the most widespread human diseases with ~10 million new cases

each year that cause about 6% - 12% of all human deaths. Although all cancers have different causes, similarities between all cancers include changes within the cell cycle and other irregularities such as overexpression of proteins or mutations in proteins that

control the cell cycle. (Kristjansdottir 2004). The cell cycle is mediated by cyclin-

dependent kinases (CDKs) and Cdc25 phosphatases. There are three Cdc25 phosphatases

in humans – Cdc25A, Cdc25B, and Cdc25C. These Cdc25 phosphatases

11 dephosphorylate, and thus activate, specific CDKs. These CDKs are in turn maintained

in an inactive state by WEE1 and MYT1, which phosphorylate CDKs. The three human

isoforms have been linked in roles during G1-S and G2-M transitions during the cell cycle

(figure 5) as well as in mitosis, where Cdc25s show activity towards CDK1/cyclin B

(important for mitotic progression) (Ducruet 2005).

Figure 5 - Cdc25 phosphatases in cell cycle control. a - phosphorylation/dephosphorylation of

CDKs. b - Cdc25 phosphatases and their roles in the cell cycle (Buotros 2006)

Overexpression of Cdc25A and Cdc25B has been found in many types of cancer

with high levels resulting in poor prognosis (Ducruet 2005). Recent studies have found correlation between specific cancers and overexpression of either Cdc25A or Cdc25B

(table 1). Alternatively, although it plays important roles in G2-M transition and during

12 the S phase of the cell cycle, overexpression of Cdc25C has not been noted in any cancer

studied (Kristjansdottir 2004).

Cdc25s Type of Cancer Studied Overexpressed hepatocellular carcinoma A, B A (78%) prostate cancer B B (97%) esophageal squamous cell carcinoma A, B, C A, B (40%, 17%) breast cancer A A (47%) breast cancer A, B, C B (32%) colorectal carcinoma A, B, C A, B (53%, 67%) pancreatic ductal adenocarcinoma A, B, C B (72%) head and neck cancers A, B, C A, B (80%, 50%) gastric carcinomas A, B, C B (78%) nonsmall cell lung cancer A, B, C A, B (60%, 45%) ovarian cancer A, B, C A, B (30%, 30%) non-Hodgkins lymphoma A, B, C A, B (35%, 39%) Table 1- Cdc25 overexpression found in different human cancers (Kristjansdottir 2004)

Aside from cancer, Cdc25 overexpression also has profound effects in

neurodegenerative diseases such as Alzheimer’s. Cdc25A and Cdc25B have been found

expressed active in the brain of patients with Alzheimer’s (Ducruet 2005). Alzheimer’s is a neurodegenerative disease that affects millions of people worldwide. Alzheimer’s disease is a degenerative disease of neurons in the cortex of the brain that eventually leads to cell death in specific areas of the brain. It is often characterized with amyloid plaques and neurofibrillary tangles, malformed nerve tangles. It has been previously believed that adult neurons of the central nervous system remain in the quiescent G0

13 phase, however, recent evidence points to neurons being able to reenter the cell cycle

(Nagy 1998). The activation of the cell cycle thus leads to the neurodegenerative nature of the disease, as seen by studies showing an increase of Cdc25A activity in Alzheimer’s disease neurons. The increase of activity of Cdc25A in Alzheimer’s disease leads to support evidence that reactivation of the cell cycle leads to neurodegeneration seen in

Alzheimer’s disease (Ding 2000).

Other diseases in which protein tyrosine phosphatases play a role are autoimmune diseases. There are over 80 diseases that can be classified as autoimmune and these affect ~5% of the population. In the United States, autoimmune diseases are among the top 10 causes of death and they affect ~10 million people (Vang 2008). A type of autoimmune disease, type 1 diabetes (T1D) is characterized by an immune attack against insulin producing pancreatic β-cells. T-cells, whose development is controlled by T-cell receptor signaling pathways, play key roles in the autoimmune destruction of β-cells. A single nucleotide polymorphism (SNP) of LYP, a protein tyrosine phosphatase encoded by the PTN22 gene located on chromosome 1p13, correlates with T1D in several populations (Bottini 2006). This SNP changes a critical amino acid residue at position

620 from Arg to Trp in LYP. The R620 in LYP is important to bind the Src homology 3

(SH3) domain of the Csk tyrosine kinase, and is an important negative regulator of T-cell receptor signaling. The SNP to W620 fails to bind Csk, therefore altering T-cell receptor signaling. This SNP, unexpectedly, produces a “gain-of-function” mutant; it is a more active phosphatase that suppresses T-cell signaling better than normal LYP. It is therefore believed, that this mutant, predisposes autoimmunity by suppressing T-cell receptor signaling resulting in survival of T cells that would otherwise be deleted (Bottini 14 2006). Another way that PTPs can influence diabetes is its role in the insulin signaling

pathway. Phosphorylation of tyrosine is important in initiation and propagation of insulin

action mediated by the insulin receptor (IR), a transmembrane glycoprotein and member

of the protein tyrosine kinases (Zhang Z. Y. 1998). The IR contains two extracellular α subunits and two intracellular β subunits. Insulin binding causes autophosphorylation on several tyrosine residues on the β subunits. The phosphorylation of the tyrosine residues creates docking sites for molecules, insulin receptor substrates (IRSs), containing SH2 or phosphotyrosine binding (PTB) domains that convey the downstream insulin signal.

Therefore the dephosphorylation of the insulin receptor will lead to the termination of insulin signal (Zhang Z. Y. 1998, Cheng 2002). It is for this reason that PTPs are molecules associated with negative regulation of the insulin receptor.

There are several PTPs that have been connected to negative regulation of the

insulin receptor (figure 6). Four PTPs that have shown increasing activity in rat

adipocytes and in insulin resistant patients include PTPα, LAR, SHP-2 and PTP1B

(Cheng 2002). PTPα is a PTP that has been found expressed in brain and kidney tissues.

It has been found to cause dephosphorylation of the IR which results in inhibition of

insulin-mediated cell rounding. Additionally, PTPα, has been found to decrease Glut4,

an insulin-responsive glucose transporter (Stoker 2005). LAR, another PTP with roles in

negative regulation of the IR, is a member of the receptor PTPs subfamily. LAR is a

direct negative regulator of the IR, as seen by studies linking overexpression of LAR with

decreased insulin action, by dephosphorylation of insulin receptor substrates (IRS)

(Cheng 2002). SHP-2 is a PTP with two N-terminal SH2 domains, a C-terminal catalytic

15 domain, and a C-terminal segment containing two tyrosyl phosphorylation sites. Studies

have shown that SHP-2 binds both the IR and IRS-1.

Figure 6 - PTPs and insulin signaling; PTP enzymes are in purple with dephosphorylation shown by

red lines. Phosphorylation by kinases is seen by green lines; black lines indicate downstream

pathways stimulated by the IR; inhibition of PTPs by peroxide is seen in blue (Stoker 2005).

In addition to binding to the IR and IRS-1, SHP-2 also binds other growth factor

receptors. SHP-2 does not dephosphorylate growth factor receptors but actually seems to

enhance growth receptor signaling. Recently, it was proposed, that binding of IRS-1 to

SHP-2 increases phosphatase activity towards IRS-1, thus resulting in dephosphorylation

(Cheng 2002, Stoker 2005). PTP1B is another PTP implicated in negative regulation of the IR. The IR receptor has been demonstrated to be a substrate of PTP1B. PTP1B becomes phosphorylated at key tyrosine residues upon insulin treatment. The phosphorylation of these residues is important for binding to the IR, as seen by mutation

16 studies in which mutation of these residues results in less involvement of PTP1B with the

activated IR. PTP1B binds to a certain region of the IR, which contains three critical

tyrosine residues. IRS-1 is also a substrate of PTP1B. Therefore, PTP1B is a negative

regulator of insulin signalling by working not only on one but on two components of the

insulin pathway (Cheng 2002). There are additional candidates that act upon the IR. One

possible candidate is PTPσ, which shows structure similarity to LAR and it is found

overexpressed in insulin sensitive tissues. PTPε, which shows similiarities with PTPα,

also inhibits insulin mediated cells. Further in vitro studies have expanded the list of

PTPs with roles in the IR: TC-PTP, PTPδ, and Sap-1. These three PTPs were shown to bind to phosphorylated IR ‘baits’ in substrate trapping experiments. They were also shown to be able to dephosphorylate an IR target derived peptide sequence based on the key tyrosine residues targeted by PTPs (Cheng 2002).

1.5 - Molecular Structure of PTPs

One of the most significant relationships between all members of the PTP family is the conserved PTP signature motif, C(X)5R, contained within the ~250 amino acid

residues catalytic domain. The PTPs are α+β proteins with tertiary folds formed from

very twisted β-sheets flanked by α-helices on both sides (figure 7). The active site can be located in a crevice of the protein surface and below it is the phosphate binding loop (P- loop) formed by the PTP signature motif (Zhang Z. Y. 1998).

17

Figure 7 - X-ray structure of PTP1B; A is the PTP loop, B is the movable WPD loop and C is the

pTyr binding pocket (Burke 1998)

The PTP active site is located in a crevice, ~9 Å for tyrosine specific phosphatases and ~6Å for dual-specificity phosphatases, on the molecular surface flanked by several loops that are important for catalysis and substrate recognition (Burke

1998). An interesting feature of the PTP loop (figure 8) is a strand-loop helix motif that contains the PTP signature motif, C(X)5R. The active site contains the P loop at the base

and is surrounded by loops that provide an essential Arg residue, which binds oxyanions

supported by hydrogen bonds between the oxygen of the oxyanion and the guanidium

group of the Arg residue. The Cys residue within the P-loop is essential for the formation of the covalent phosphoenzyme intermediate. Studies between the complex of PTP and

18

Figure 8 - Structure of P-loop from bovine LMW-PTP (Tabernero 2008)

tungstate reveal that the Cys residue acts as a nucleophile in the catalytic mechanism.

Additonally, studies have also shown that the Asp residue acts as a general acid in

catalysis. Crystal structures of the complex between PTP and an oxyanion also reveal

that the Asp residue is across the Cys residue and points towards the oxyanion (Burke

1998, Zhang Z. Y. 1998). This essential Asp residue resides in a specific surface loop

known as the WPD loop. This particular loop is pretty diverse among members of the

PTP family except for the conserved Asp and Trp residues near the hinge position of the

loop (figure 9). This particular loop acts like a “flap” to cover the active site upon substrate binding. Following catalysis, the loop opens up once again to facilitate release of the product. The opening and closing of the WPD loop is believed to work for the entire PTP family, however, this has not been confirmed with the dual specificity phosphatases and the low molecular weight phosphatases since only the ligand-bound structures have been determined (Zhang Z. 2002).

19

Figure 9 - Amino acid sequence comparison between WPD loops in different PTPs (Zhang Z. Y.

1998)

As a whole, the core structural elements (the P-loop and the WPD loop) are common to all the classical PTPs, the dual specificity phosphatases, and the low molecular weight phosphatases. Superposition of the structural elements between members of the different subclasses show structural conservation and thus leads to the belief that the PTPs follow the same, if not similar, catalytic mechanism.

1.6 - Mechanism of catalysis for PTPs

The catalytic mechanism of PTPs has been determined through numerous studies on different members of the PTP family. It has been revealed, that the mechanism of catalysis of PTPs differs from other protein phosphatases such as members of the PPP family. The PPP family uses metal cations in a one step mechanism involving water in their catalytic mechanism, whereas, PTPs do not use metal cations and proceed through a

20 two-step mechanism involving the formation of a covalent cysteine phosphoenzyme

intermediate followed by hydrolysis (Mildvan 1997).

The common two-step catalytic mechanism is shown in figure 10. This

mechanism requires the side chain of a Cys residue to act as a nucleophile that attacks the

phosphoryl group of the substrate and thus forms the covalent cysteinyl phosphate

intermediate. The Arg residue, found in the PTP active site, creates hydrogen bonds

through its guanidinium group with the phosphoryl group of the substrate and aids in

substrate binding as well as transition state stability. An Asp residue, from the WPD

loop, functions as an acid by protonating the ester leaving group. It also helps in

improving the formation of the covalent phosphoenzyme intermediate. During the

second step of the catalytic mechanism, a nucleophilic water, with the help of the Asp

residue now acting as a general base, dephosphorylates the phosphoenzyme intermediate

and releases the free enzyme and the inorganic phosphate (Wang 2003).

Figure 10 - Catalytic mechanism of PTPs (Wang 2003).

There are residues within the PTP signature catalytic motif that are essential for

PTP catalysis. The Cys residue is important in forming the covalent cysteine phosphoenzyme intermediate. This has been supported by site directed mutagenesis

21 studies in which mutation of the Cys residue within the PTP signature motif leads to no

phosphatase activity (Zhang Z. Y. 1998). The Arg residue in the P-loop is important for substrate binding and transition state stabilization. As with the Cys residue, mutation of this residue leads to loss of phosphatase activity. Reasoning behind transition state stabilization of the Arg residue stems from the ability of the guanidinium group to complex the phosphoryl oxygens and to stabilize the trigonal bipyramidal transition state

(Burke 1998). Another important feature involved in the catalytic mechanism of PTPs is the utilization of general acid/base catalysis to increase catalytic turnover. In the PTP catalytic mechanism, the Asp residue of the WPD loop, acts as an acid during the first step and as a base during the second step. Mutation of this residue to an Asn results in lowering of catalytic activity by three orders of magnitude (Burke 1998).

Another important aspect of the PTP catalytic mechanism is the stabilization of the transition state. The active site Asp residue is important in stabilizing the transition state, specifically, the dissociative transition state for E-P complex formation (figure 11).

Here the Asp residue facilitates the leaving of the phenoxide. It has been found that

mutations of the Asp residue more greatly affect the formation of the intermediate in

comparison to the breakdown. Properties of transition states also explain the conserved

Ser/Thr residue seen in the PTP signature motif. Differences of amino acid residues at

this position result in great decrease in product decomposition. The hydroxyl group of

Ser/Thr is suggested to facilitate the breakdown of the phosphoenzyme intermediate, by

charge stabilization of the thiolate in Cys (figure 11) (Burke 1998).

22

Figure 11 - Suggested transition state for phosphorylation/dephosphorylation in PTPs (Zhang Z. Y.

1998)

1.7 - PTPs in eukaryotes and prokaryotes

Proteins that are phosphorylated undergo post translation modifications to achieve this covalent alteration. Protein phosphorylation was first seen, in the 1950s, in eukaryotes. About 20 years later, phosphorylation in prokaryotes was observed

(Cozzonne 1988). At the beginning, protein phosphorylation was thought to be unique to

‘higher’ eukaryotic organisms, since it is a key element in signal transduction and it was early associated with hormone action. Later on, in the 1980s, it become known that protein phosphorylation could also be detected in ‘lower’ eukaryotic organisms as wells

23 as prokaryotes. Even though phosphorylation was evident in prokaryotes (Kennelly

1999), the phosphorylation mechanism did not resemble the mechanism seen in

eukaryotes (Shi 1998). Three phosphorylating systems in prokaryotes emerged: the

“two-component system”, the phosphoenol pyruvate:carbohydrate phosphotransferase

(PTS) system, and a mechanism that closely resembles the “classical” ATP/GTP- dependent system seen in eukaryotes (Cozzone 2004).

Protein- Protein-tyrosine phosphatase Organism tyrosine kinase Name Family Gram-negative eubacteria Acinetobacter johnsonii Ptk Ptp Low MW Acinetobacter lwoffii Wzc Wzb Low MW Erwinia amylovora AmsA AmsI Low MW Escherichia coli K-12 WzcCa Wzb Low MW Escherichia coli K-12/K-30 Etk Etp Low MW Escherichia coli K-30 WzcCPS Wzb Low MW Gram-positive eubacteria Bacillus subtilis YwqD YwqE PHP - PtpA Low MW

Mycobacterium tuberculosis - MPtpA, MPtpB Low MW Staphylococcus aureus Cap1B, Cap1A Cap1C PHP Cap5B, Cap5A Cap5C PHP Cap8B, Cap8A Cap8C PHP - PtpA, PtpB Low MW Streptococcus agalactiae CpsD (+CpsC) CpsB PHP

Table 2- Prokaryotic PTPs (adapted from Cozzone 2004)

In bacteria, PTPs come from two of the main families: the classical pTyr phosphatases and DSPs as well as LMW-PTPs. Recently, in 2002, a new manganese- dependent PTP has been discovered (Morona 2002). This PTP belongs to a third class of

prokaryotic PTPs, the polymerase and histidinol phosphatase (PHP) family, found mainly

24 in gram-positive bacteria. As indicated in table 2, there are some PTPs that have an

associated PTK, as specific substrate for the phosphatase. An example of this is the

kinase and phosphatase seen in E.coli, WzcCA and Wzb respectively. Wzb is a LMW-

PTP that is able to dephosphorylate the autokinase WzcCA. Another example is the Etp phosphatase that can dephosphorylate the Etk kinase.. The kinase/phosphate pairing is most common between a LMW-PTP and its corresponding kinase generally in gram- negative bacteria. In gram-positive bacteria, pairings are common between PHPs and their kinase (Cozzone 2004).

Eukaryotic PTPs are more diverse and correlations have been made to trace some back to bacteria. The human genome encodes over 100 PTPs with 11 being inactive and

16 having substrates that are not proteins. Most, however, are believed to dephosphorylate phosphotyrosine and, in the case of DSPs, phosphoserine and phosphothreonine. Evolutionarily, it is now believed that dephosphorylation of phosphotyrosine coincides with the appearance of tyrosine kinases, since the expansion of the PTP family is related to the appearance of tyrosine kinases. The classical PTPs and the DSPs are believed to have evolved from a common ancestor, as seen by studies about their overall structure. The LMW-PTPs and Cdc25 phosphatases are thought to have evolved from bacteria, as seen through sequence similarities studies (Moorehead

2009).

Several genomic studies reveal that tyrosine phosphorylation was present in bacteria but to a limited degree. The emergence of tyrosine kinases created a plethora of

PTP evolution. PTPs evolved by addition of different domains and docking mechanisms.

25 This, it is believed, explains the nature of eukaryotic and prokaryotic PTP differences

observed (Moorehead 2009).

1.8 – Poxvirus phosphatases

Interferons (IFNs), type I and type II, are major factors in the defense against viruses. Binding of type I IFNs, IFN-α and IFN-β, to its receptor results in the triggering of tyrosine phosphorylation of the Janus kinases Tyk2 and Jak1 then followed by tyrosine phosphorylation of Stat1 and Stat2 (signal transducer and activator of transcription) proteins. The activated Stat1 and Stat2 heterodimerize with IFN-regulatory factor 9

(IRF-9) and translocate to the nucleus to activate IFN-stimulated genes (ISGs) through

IFN-stimulated responsive element (ISRE) (F. B. Wang 2009). On the other hand, binding of type II IFNs, IFN-γ, to its IFN-γ receptor activates Jak1 and Jak2 which results in the tyrosine phosphorylation of only Stat1. Consequently, Stat1 homodimerizes and migrates to the nucleus and activates ISGs through the γ-activated sequence (GAS) (F. B.

Wang 2009).

The important role of the Jak-Stat pathway in defense against viruses is evident in the evolution of viruses’ ability to inhibit the pathway at different points. For example, the Japanese encephalitis virus and the West Nile virus inhibit Tyk2 and Jak1 tyrosine phosphorylation downstream of the IFN receptors while human cytomegallvirus and adenovirus decrease Jak1 expression (F. B. Wang 2009). Additionally, mumps virus V protein disrupts the ability of Stat1 to associate with type I and type II IFN receptors while measles V virus protein associates with both Stat1 and Stat2 and blocks function

26 but not activation (Mann 2008). The members of the poxvirus family all encode unique

immune evasion molecules that challenge the host’s immune response, in particularly,

they antagonize IFN-induced antiviral ISGs. However, only two distinct mechanisms on

how poxviruses challenge the Jak-Stat pathway have been determined. The first mechanism in which poxviruses inhibit the pathway is by acting as IFN ligand scavengers and in so doing prevent initiation of IFN signal transcution The second mechanism involves a dual specificity phosphatase that dephosphorylates Stat1 following type II IFN stimulation or both Stat1 and Stat2 following type I IFN treatment (F. B.

Wang 2009).

One of these poxviruses, the vaccinia virus, encodes a dual specificity phosphatase, the vaccinia virus H1 gene product (VH1), in its genome. VH1 was the first discovered dual specificity phosphatase and it is found in a central part of the viral genome, open reading frame H1 (ORF H1), a region that is highly conserved among members of the poxvirus family (Mann 2008). VH1 has been found to block IFN-γ signaling by dephosphorylation of Stat1 (Najarro 2001, Mann 2008). Additionally, the vaccinia virus has also been shown to inhibit Stat1-independent signaling pathways.

Together, the mechanisms that the vaccinia virus employ to disrupt antiviral pathways promotes the successful reproduction during poxvirus replication (Mann 2008).

1.9 – Low Molecular Weight Phosphatase WZB in Escherichia coli

The presence of protein tyrosine kinases in bacteria, found much later than in eukaryotes, was first suggested in Escherichia coli from the finding of phosphotyrosine

27 residues. Later, phosphotyrosine residues were also noticed in proteins from other bacterial species. However, the presence of phosphotyrosine phosphatases has just begun to be reported in bacteria, though the role of reversible tyrosine phosphorylation in bacteria is still unclear (Vincent 1999).

Gram-positive and gram-negative bacteria utilize bacterial capsules (made up of capsular polyssacharides (CPS)) to evade the immune system by playing critical roles in the interactions between the bacteria and their environments (Reid 2005). These capsules act by increasing adhesion to host tissue and by building up resistance to phagocytosis.

Often the capsule resembles host cell components thus impairing the host’s immune response (Reid 2005). There have been several methods described for CPS biosynthesis and assembly. One of these methods is known as the Wzy-dependent pathway, which is found in many gram-positive and gram-negative bacteria (Hagelueken 2009). In the

Wzy-dependent pathway, intermediates carrying individual repeats of the CPS are exported across the cytoplasmic membrane where they form substrates for the polymerization step, which requires the oligosaccharide polymerase Wzy. Full polymerization requires other polysaccharide co-polymerases such as Wzc in E.coli

(Hagelueken 2009, Raetz 2002).

In E.coli K30 two proteins, Wzc and Wzb, are responsible for the biosynthesis of colanic acid, an exopolysaccharide. Wzb is a low molecular weight phosphatase which dephosphorylates the autokinase Wzc, with the phosphorylation state of Wzc dictating colanic acid production. When Wzc is phosphorylated no colanic acid is synthesized, but, if, Wzc is dephosphorylated by Wzb, colanic acid is produced (Lescop 2006).

28

CHAPTER 2 - MATERIALS AND METHODS

2.1 – Instrumentation and buffers

Sonication of protein cell slurry for protein purification was done using a Misonix

Sonicator 3000. Cell were sonicated on ice in 20sec intervals for 3 min. Typical power

output was 8-12 Watt.

All protein purifications were done on a BioLogic DuoFlow Chromatography

System from BioRad using BioLogic DuoFlow software version 5.0.

All absorbance readings for concentration determination and kinetic analysis were

performed on a Hewlett Packard Agilent 8453 UV-Vis spectrophotometer with a deuterium and tungsten lamp and a range from 190nm to 1100nm.

All buffers and media were made from ACS or biochemical grade chemicals.

Chemicals were purchased from standard suppliers including Sigma-Aldrich, Fisher,

VWR, Difco, BDH, Amresco and others.

LB broth was made from 10g/l yeast extract, 5g/l tryptone, 10g/l NaCl, 2.2g/l

KH2PO4, 9.8g/l K2HPO4, pH 7.5. LB-agar plates contained 15g/l of agar. Media were

heat sterilized in an autoclave at 120°C for 30 min. Antibiotica (ampicillin or

kanamycin) were added after sterilization at a final concentration of 100 μg/l.

Buffers used in nickel affinity chromatography were composed of 50mM KPi,

300mM NaCl, 20mM imidazole, pH 7.8 (Buffer A) and 50mM KPi, 300mM NaCl, 29 500mM imidazole, pH7.8 (Buffer B); gel filtration buffer was 25mM Tris, 50mM NaCl,

pH6.

WZB kinetic reaction buffer was composed of 50mM Tris, 50mM BisTris, 50mM

NaCl, pH5

2.2 – Transformation of plasmid into expression hosts

Plasmids encoding the phosphatases were transformed into expression hosts by either heat shock or electroporation.

In heat shock transformation, 2μL of plasmid DNA was incubated with 100μL of chemically competent expression cell slurry on ice for 30 minutes followed by incubation at 42°C (water bath) for exactly 30 seconds and then quickly chilled on ice for at least 2 minutes. The cells were recovered with 1 ml LB medium and incubated in a rotating shaker at 37°C for one hour. Cells were then plated onto LB agar plates containing an appropriate selection antibiotic.

Transformation by electroporation was done as follows – electroporation cuvettes were incubated on ice for 10 minutes. 2μL of plasmid DNA and 90μL of electro competent expression cell slurry were mixed in the cuvette and placed in electroporator. An electroporation voltage of 1,600V was used. The cells were recovered with 1mL of LB medium and incubated in shaker at 37°C for one hour. The transformation mixture was then plated onto appropriate antibiotic containing LB plates.

30 E.coli strains used for protein expression were either Origami2(DE3)

(Invitrogen), BL21Star(DE3) (Invitrogen) or, T7Express (NEB Biolabs).

Expression plasmids for WZB and PVP were provided by Dr. Vetter. The PVP gene was cloned into the pET101 vector (Invitrogen) using the topo-cloning methodology. A stop codon at the 5’ end of the PVP gene sequence resulted in expression of PVP without a C-terminal V5 epitope and 6xHis tag (PVP non-tagged). A clone without the additional stop codon resulted in expression of PVP with V6 epitope and

6xHis tag (PVP His6-tagged). A vector map, gene and protein sequences of both forms of

PVP are shown below.

Figure 12 - Map of pET101 expression vector (Invitrogen)

Two variants of His tagged WZB were expressed from pET28 (Novagen) and pET29 (Novagen) plasmids, respectively. The pET28 construct used the NcoI and EcoRI restriction sites. The EcoRI site was proceeded by a stop codon to produce WZB with a

31 N-terminal 6xHis tag (His-WZB). The pET29 construct used the NdeI and XhoI

restriction sites to produce WZB with a C-terminal 6xHis tag. Vector maps for pET28 and pET29 are shown below.

Figure 13 - pET vector backbone (Novagen)

2.3 – Cell growth and expression of phosphatases

For both forms of PVP and WZB, cells were grown first in 2mL volumes and the

grown cells were used to inoculate 400mL of LB-broth media and incubated at 37°C to

an optical density of 0.5-0.8. Upon this OD value, the PVP cells were induced with 1mM

IPTG overnight at room temperature. The WZB cells were transferred to a room

temperature shaker and allowed to cool down for 30 minutes. Cells were then induced by

addition of 1mM IPTG and allowed to express for 6-8 hours. For both PVP and WZB,

the cells were harvested and the cell pellet was resuspended in 50mM KPi, 300mM NaCl,

20mM imidazole, pH7.8 and stored at -80°C. Another medium was used in the cell growth of PVP - auto-inducing medium. This is a medium whose components promote 32 high-density cell growth and automatic induction of lac promoter protein expression.

Additionally, this method has been found to be suitable for expression in pET vector systems and it is, at times, preferable to induction by IPTG (Grabski, 2003). The cells were grown at 37°C with continuous shaking and harvested upon reaching an OD600 of approximately 20. Confirmation of protein expression was monitored by SDS-Page and solubility of expressed protein was further tested by enzymatic activity with PNPP.

Nucleotide PVP His6- ATGGATAAAAAATCTCTGTATAAATATCTGCTTCTG sequence tagged CGTAGCACCGGCGATATGCATAAAGCTAAATCTCC GACCATTATGACTCGCGTAACCAATAACGTTTATCT GGGTAACTATAAGAACGCGATGGATGCACCGTCCT CTGAAGTTAAATTTAAGTATGTACTGAACCTGACT ATGGATAAATATACTCTGCCGAACTCCAACATTAA CATTATTCACATTCCACTGGTAGATGATACTACCAC TGATATCAGCAAGTACTTTGATGATGTTACTGCTTT TCTGTCCAAATGCGATCAGCGTAACGAACCGGTTC TGGTACATTGTGCGGCTGGTGTGAACCGCAGCGGT GCAATGATTCTGGCTTATCTGATGTCTAAGAATAA AGAGTCTCTGCCGATGCTGTACTTCCTGTATGTGTA TCATAGCATGCGTGATCTGCGTGGCGCGTTCGTTGA AAACCCGTCCTTTAAACGCCAAATCATTGAAAAAT ACGTGATTGATAAAAATTCTGGAACCAAGGGCGAG CTCAATTCGAAGCTTGAAGGTAAGCCTATCCCTAA CCCTCTCCTCGGTCTCGATTCTACGCGTACCGGTCA TCATCACCATCACCATTG

PVP non- ATGGATAAAAAATCTCTGTATAAATATCTGCTTCTG tagged CGTAGCACCGGCGATATGCATAAAGCTAAATCTCC

33 GACCATTATGACTCGCGTAACCAATAACGTTTATCT GGGTAACTATAAGAACGCGATGGATGCACCGTCCT CTGAAGTTAAATTTAAGTATGTACTGAACCTGACT ATGGATAAATATACTCTGCCGAACTCCAACATTAA CATTATTCACATTCCACTGGTAGATGATACTACCAC TGATATCAGCAAGTACTTTGATGATGTTACTGCTTT TCTGTCCAAATGCGATCAGCGTAACGAACCGGTTC TGGTACATTGTGCGGCTGGTGTGAACCGCAGCGGT GCAATGATTCTGGCTTATCTGATGTCTAAGAATAA AGAGTCTCTGCCGATGCTGTACTTCCTGTATGTGTA TCATAGCATGCGTGATCTGCGTGGCGCGTTCGTTGA AAACCCGTCCTTTAAACGCCAAATCATTGAAAAAT ACGTGATTGATAAAAAT

Amino PVP His6- MDKKSLYKYLLLRSTGDMHKAKSPTIMTRVTNNVYL acid tagged GNYKNAMDAPSSEVKFKYVLNLTMDKYTLPNSNINII sequence HIPLVDDTTTDISKYFDDVTAFLSKCDQRNEPVLVHC AAGVNRSGAMILAYLMSKNKESLPMLYFLYVYHSM RDLRGAFVENPSFKRQIIEKYVIDKNSGTKGELNSKLE GKPIPNPLLGLDSTRTGHHHHHH

Amino Acids: 206

Molecular weight: 23,483 Da

Isoelectric point, pI: 9.5

PVP non- MDKKSLYKYLLLRSTGDMHKAKSPTIMTRVTNNVYL tagged GNYKNAMDAPSSEVKFKYVLNLTMDKYTLPNSNINII HIPLVDDTTTDISKYFDDVTAFLSKCDQRNEPVLVHC AAGVNRSGAMILAYLMSKNKESLPMLYFLYVYHSM RDLRGAFVENPSFKRQIIEKYVIDKN

34 Amino Acids: 171

Molecular weight: 19,698 Da

Isoelectric point, pI: 9.4

Table 3 - DNA and protein sequence of both forms of PVP

Nucleotide His6- ATGGGCAGCAGCCATCATCATCATCATCACAGCAG Sequence tagged CGGCCTGGTGCCGCGCGGCAGCCATATGTTTAACA WZB ACATCTTAGTTGTCTGTGTCGGCAATATTTGCCGTT CCCCGACGGCGGAACGCTTACTGCAACGTTATCAC CCGGAGCTGAAAGTGGAGTCCGCTGGACTCGGCGC GCTGGTCGGTAAGGGCGCTGATCCTACCGCTATCA GCGTCGCCGCAGAACATCAACTGTCTCTGGAAGGT CACTGTGCCCGTCAAATCAGCCGCCGTCTGTGTCG CAACTACGACCTGATTTTGACCATGGAAAAGCGCC ATATCGAACGCTTATGCGAGATGGCACCGGAGATG CGCGGCAAAGTGATGCTGTTTGGTCACTGGGATAA CGAATGTGAAATCCCCGATCCGTATCGCAAAAGCC GGGAAACGTTTGCAGCGGTGTACACATTACTTGAA CGGTCTGCCCGCCAGTGGGCGCAGGCATTGAACGA GCAGGTA WZB ATGTTTAACAACATCTTAGTTGTCTGTGTCGGCAAT His6- ATTTGCCGTTCCCCGACGGCGGAACGCTTACTGCA tagged ACGTTATCACCCGGAGCTGAAAGTGGAGTCCGCTG GACTCGGCGCGCTGGTCGGTAAGGGCGCTGATCCT ACCGCTATCAGCGTCGCCGCAGAACATCAACTGTC TCTGGAAGGTCACTGTGCCCGTCAAATCAGCCGCC GTCTGTGTCGCAACTACGACCTGATTTTGACCATGG AAAAGCGCCATATCGAACGCTTATGCGAGATGGCA CCGGAGATGCGCGGCAAAGTGATGCTGTTTGGTCA

35 CTGGGATAACGAATGTGAAATCCCCGATCCGTATC GCAAAAGCCGGGAAACGTTTGCAGCGGTGTACACA TTACTTGAACGGTCTGCCCGCCAGTGGGCGCAGGC ATTGAACGAGCAGGTATCACTCGAGCACCACCACC ACCACCAC Amino His6- MGSSHHHHHHSSGLVPRGSHMFNNILVVCVGNICRS Acid tagged PTAERLLQRYHPELKVESAGLGALVGKGADPTAISVA Sequence WZB AEHQLSLEGHCARQISRRLCRNYDLILTMEKRHIERL CEMAPEMRGKVMLFGHWDNECEIPDPYRKSRETFAA VYTLLERSARQWAQALNEQV Amino Acids: 167 Molecular Weight: 18,873 Da Isoelectric point: 8.0 7.85 WZB MFNNILVVCVGNICRSPTAERLLQRYHPELKVESAGL His6- GALVGKGADPTAISVAAEHQLSLEGHCARQISRRLCR tagged NYDLILTMEKRHIERLCEMAPEMRGKVMLFGHWDN ECEIPDPYRKSRETFAAVYTLLERSARQWAQALNEQ VSLEHHHHHH Amino Acids: 155 Molecular Weight: 17,791 Da Isoelectric point: 7.31 Table 4 - DNA and protein sequence of WZB protein; red portion signifies the histidine tag at either the N- or C-terminus

2.4 – Isolation of inclusion bodies

The frozen bacterial cell slurry was thawed and diluted with 1x TBS to reduce sample viscosity. Lysozyme was added to a final concentration of 1mg/ml and the solution was allowed to stir at room temperature for 1 hour. The stirred solution was then

36 thoroughly sonicated (at least 3 minutes in intervals of 20-30 seconds) followed by centrifugation (18,500 g for 30 minutes). The cell pellet was resuspended in a wash buffer (50mM Tris, 150mM NaCl, 2M Urea, 1% Triton X-100 (w/v), pH8) and centrifuged again (18,500 g for 30 minutes). The process was repeated four times with the final wash excluding Triton X-100. The pellet, now white, was stored at -80°C. The inclusion body isolation scheme can be seen below (figure 14)

Figure 14 - Isolation of inclusion bodies scheme

2.5 – Nickel affinity chromatography

All the phosphatases that contained the histidine tag functionality were purified

using a nickel affinity column. For PVP, the inclusion body pellet was solubilized in 8M

urea, 50mM KPi, 20mM imidazole, pH8 and purified on a nickel affinity column

37 (HisTrap high performance column, 1mL volume from Pharmacia/GF) and ran on a step

gradient at a flow rate of 1mL/min: 100% buffer A for 20mL, 100% buffer B for 10mL

and 100% buffer A for 10mL. Buffer A: 6M urea, 50mM KPi, 20mM imidazole, pH8;

Buffer B: buffer A + 0.5M imidazole. Alternatively, for larger amounts of protein

purified, an IMAC Sepharose – nickel loaded high performance 20mL column form

Pharmacia/GF was used as follows: 100% buffer A for 60mL; 100% buffer B for 20mL,

100% buffer A for 20mL where buffer A is 50mM KPi, 300mM NaCl, 20mM imidazole, pH7.8 and buffer B is buffer A plus 500mM imidazole.

2.6 – Ion exchange chromatography

The phosphatase that did not contain a histidine tag, PVP non-tagged, was

purified using an SP-ion exchange chromatography column (Q sepharose High

Performance 20mL column). The PVP inclusion was solubilized in 50mM Tris, 8M urea,

0.5% β-mercaptoethanol, pH8 and purified on a step gradient as follows: 100% buffer A

for 60mL, 50% buffer A, 50% buffer B for 40mL, 100% buffer B for 30mL then 100%

buffer A for 40mL with buffer A: 50mM Tris, 7.5M Urea, 0.1% β-mercaptoethanol,

pH7.2 and buffer B: buffer A plus 1M NaCl.

2.7 – Gel filtration chromatography

After initial purification using nickel affinity column, WZB was further purified

by gel filtration using a HiPrep Sephacryl S100HR 320mL column. The eluting buffer 38 was 25mM Tris, 50mM NaCl, pH6. The protein peak was collected and DTT (10 mM final concentration) and EDTA (1 mM final concentration) were added.

2.8 – Refolding matrix design

The purified denatured protein, PVP, was then subjected to several refolding buffers to ascertain the most suitable refolding conditions for the phosphatase. Different buffer systems at different pHs were tested. Additionally, ionic strength was varied as well as additives and reducing agents. The three buffers systems tested are shown below:

NaCl, 100mM Tris Glycerol β-mercaptoethanol, NDSB256, 100uM (+) pH8.2 10% 10mM 250mM 500uM (-) 1 + + 2 + + + 3 + + + 4 + + + 5 + + + + + 6 + - 7 + - + 8 + - + 9 + - + 10 + - + + + Table 5 – Initial refolding buffer systems tested for His6-tagged PVP

The same refolding screens were done with 100mM borate pH9.5 and 100mM potassium phosphate pH5.6 in place of Tris pH 8.2. A second set of refolding buffer systems was

39 created to compare two buffer pH values as well as two non-detergent sulphobetaines

(table 6).

NDSB201, NaCl, β- Tris pH8.2 (+) Glycerol 250mM (+) 100uM (+) mercaptoethanol, Borate pH9.5 (-) 10% NDSB256, 500uM (-) 10mM 250mM (-) 11 + + + 12 - + + 13 + + + + + 14 + + + + - 15 - + + + + 16 - + + + - 17 + - + + + 18 + - + + - 19 - - + + + 20 - - + + - Table 6 - Additional refolding buffer systems for His6-tagged PVP

2.9 – Refolding protocol

Purified His6-tagged PVP, in a volume of 50μL, was added to 1mL of refolding buffer, stirred, and allowed to sit overnight at 4°C. Enzyme activity was checked by addition of the substrate, p-nitrophenyl phosphate, incubated at room temperature for

10min and subsequently quenched with 1M NaOH with absorbance readings taken at

405nm. For non-tagged PVP, The refolding buffer consisted of 50mM Tris, 125mM

NaCl, 0.5% β-mercaptoethanol, 0.8M NDSB256, pH7. The pooled fractions from the ion

40 exchange column were added dropwise to the refolding buffer on ice. The reaction

mixture was allowed to stir overnight at 4°C.

2.10 – Enzyme kinetics

For refolded PVP, a continuous assay was used as follows: A reaction mixture

containing refolded PVP (in 50mM Tris, 125mM NaCl, 0.5% β-mercaptoethanol, 0.8M

NDSB256, pH7) and substrate, pNPP (prepared in refolding buffer) was prepared to a total volume of 110μL. The reaction was continuously monitored for release of p- nitrophenolate at 405nm using a UV-Vis spectrophotometer. Substrate concentration ranged from 1mM to 15mM.

For WZB, a quenched assay was used as follows: The total reaction volume was

1000μL with 50μL being the enzyme, WZB, a specific concentration of pNPP and the remaining volume was made up with 50mM Tris, 50mM Bis-Tris, 50mM NaCl, pH5.

The reaction mixture was mixed and allowed to proceed. At specific time intervals, every 10 seconds, 100μL of the reaction mixture was quenched with 5M NaOH.

Absorbance readings were measured at 405nm using a UV-Vis spectrophotometer. For kinetic reaction using β-naphthyl phosphate (pNaph) and phosphoserine (pSer) a colorimetric assay adapted from Katewa et al was used. This assay is dependent on the development of the molybdenum blue color after a reducing agent is introduced (a mixture of hydrazine sulfate and ascorbic acid) (Katewa, 2003). A reaction mixture was prepared with a specific substrate concentration equaling 1000μL minus the volume of

41 WZB to be added. WZB was added in volumes of 100μL and the reaction was allowed

to proceed. At specific time intervals, every 30 seconds to 1 minute, 100uL aliquots of

reaction mixtures was taken and added to the quench solution (100μL of 5% SDS). To

the quenched reaction mixture, the following was added: 250μL of DI water, 250μL of

1.5M (3N) H2SO4, and 100μL of 0.5% ammonium molybdate in 3N H2SO4. The

reagents were vortexed thoroughly before adding 200μL of the reducing reagent

(10mg/ml hydrazine sulfate and 10mg/ml ascorbic acid in 1N H2SO4) and the reaction

vessel was mixed. The reaction was allowed to sit for 60 minutes and absorbance at

820nm was recorded.

2.11 – Fitting of enzyme kinetic data

All graphs were created on KalediaGraph 4.01 (Synergy Software). For the kinetic reaction between pNPP and PVP and WZB, the molar extinction coefficient of pNPP was used - ε = 18,000 M-1cm-1. A plot of absorbance versus time was then converted

into a product concentration versus time graph using the molar absorptivity coefficient of

pNPP. This graph yielded the initial velocity at specific substrate concentration and a

Michaelis-Menten graph was then obtained and fitted using the Michaelis-Menten

equation - yielding Km, Vmax, and kcat values. For the kinetic reaction

between the phosphatases and pNaph and pSer, the experimental extinction coefficient,

from the molybdenum blue color formed, found from a standard curve was used (ε =

8,000 M-1cm-1) and the graphs were fitted as described above.

42 2.12 – Synthesis of non-detergent sulphobetaines (NDSBs)

All the NDSBs were synthesized by the reaction of 1,3-propane sultone and a tertiary amine in 1,2-dichloroethane according to reference (L. B.-B. Vuillard 1995). The reaction mixtures were stirred at the reflux for 3-6 hours, and then, continued at room temperature for two days (figure 15). The products (figure 16) were collected by filtration, washed with EtOAc, dried and characterized by HPLC, MALDI-TOF MS, and

NMR spectroscopy.

O O N+ S O N O 1,2-DCE S Reflux 3-6 hrs O 1,3-propane RT 48 hrs -O tertiary amine NDSB195 sultone

Figure 15 – General synthesis scheme for NDSBs

43 N+ N+ - SO3

NDSB 195 - 70% yield - SO3 NDSB 256 - 93% yield - SO3

- O3S N+

+ NDSB201 - 78% yield N

- SO3 NDSB257 - 88% yield

- O3S N N+

NDSB204 - 93% yield - O3S

+ - N N+ SO3

N+ NDSB360 - >99% yield

- SO3 O NDSB223 - 65% yield

N N+ N+

- NDSB246 - 93% yield SO3

NDSB223b - 65% yield

Figure 16 – Yield and structure of NDSBs after synthesis

44 2.13 – Molecular cloning of WZB mutants

Molecular cloning was performed by Dr. Vetter. Mutation of the two catalytic amino acids, cysteine 9 and aspartic acid 115, was conducted with the use of the quick change mutagenesis kit from Stratagene. The mutagenic primers were added to the denatured DNA template and using thermal cycling, the mutant strand was synthesized.

A restriction endonuclease, Dpn I, specific for methylated DNA was added to the thermal cycling products to denature the parental DNA. The mutated DNA was then transformed in XL1-Blue and the plasmid was then sequenced for verification of the appropriate mutation. After confirmation of the mutation, the plasmid was transformed into NEB

T7Express electro-competent cells for protein expression.

45

CHAPTER 3 – RESULTS: CELL GROWTH, EXPRESSION,

PURIFICATION AND KINETIC CHARACTERIZATION OF

DUAL SPECIFICITY PHOSPHATASE, PVP

3.1 – Growth and expression of PVP

The dual specificity phosphatase, PVP, was cloned in two forms, non-tagged PVP

and His6-tagged PVP, into a pET101 expression vector. The tagged formed of PVP contained a histidine tag at the C terminus and the non-tagged PVP form contained no histidine tag at either terminus. For protein expression, the PVP plasmids were transformed into BL21 Star (DE3), an E.coli expression host specifically designed for high level recombinant protein expression. In order to optimize the amount of soluble protein expressed, several conditions, such as type of media, temperature and expression time, were tested and altered.

Initially standard conditions for cell growth and protein expression were tried.

The transformed plasmids were grown in Luria-Bertani (LB) medium, a high nutrient cell growth medium, at 37°C until an optical density at 600nm (OD600) of ~0.8 was obtained.

Upon reaching the desired OD600, protein expression was initiated by addition of 1mM

isopropyl β-D-thiogalactopyranoside (IPTG), an analog of lactose that triggers

46 transcription of the T7/lac operon, and allowed to proceed overnight. The cells were then harvested and stored at -80°C. Protein expression was confirmed via sodium dodecyl

sulfate polyacrylamide gel electrophoresis (SDS-PAGE). The presence of soluble protein expressed was tested, via enzymatic activity with PNPP, and it was discovered that the

majority of expressed protein was present as inclusion bodies, protein aggregates. Due to

this, different factors influencing cell growth and protein expression were changed in

order to acquire the most soluble protein possible.

The first factor altered was the temperature after induction of protein expression.

The cells were grown regularly in LB medium at 37°C until an OD600 was reached. The

cells were induced with 1mM IPTG and the temperature was lowered to room

temperature, 25°C, and allowed to grow overnight. As with the previous attempt, tests showed that the protein was again expressed as inclusion bodies. It was therefore decided to reduce the expression time from overnight (approximately 12 hours) to 4-6 hours. Testing with PNPP revealed that, once again, the protein was expressed as inclusion bodies. In order to increase the amount of soluble protein being expressed, it was then decided to change the growth medium. The medium was changed to an auto- inducing medium. However, as with the previous attempts, the expressed protein was found to be mostly inclusion bodies. A final attempt was made using the auto-inducing medium. The temperature was lowered to room temperature (25°C) and allowed to proceed to an OD600 of approximately 20. Protein expression was confirmed by SDS-

PAGE and presence of soluble protein was again tested by enzymatic activity with PNPP.

As with all other attempts, it was determined that the majority of protein expressed was found as inclusion bodies. Both forms of PVP, His6-tagged PVP and non-tagged PVP,

47 were expressed as inclusion bodies in all conditions tested. At this time, it was decided to

continue to the purification steps with the protein expressed as inclusion bodies.

3.2 – Purification of PVP

After altering several factors during cell growth and protein expression, most of

the protein was expressed as inclusion bodies, usually resulting of high-level of

recombinant protein expression (Singh 2005). The inclusion bodies are protein aggregates of misfolded protein that have no protein activity. At the beginning, the use of the inclusion bodies was attempted to be bypassed because solubilization of inclusion

bodies requires use of strong denaturing agents, such as urea or guanidinium chloride, which leads to protein denaturation. Protein purification can then be performed with the solubilized inclusion bodies; however, the purified protein is in an unfolded, inactive state. Protein refolding, thus, leads to various complications since finding conditions for in vitro refolding becomes problematic.

The first step involved the degrading of the cell wall by the use of lysozyme followed by sonication. Initial purification of the inclusion bodies was then accomplished by the use of a wash buffer containing urea and triton X-100 in conjunction with centrifugation steps in between the washes. The scheme followed can be seen in the

Materials and Methods section. Purity of the inclusion bodies was visualized using SDS-

PAGE (figure 17). The isolation of inclusion bodies procedure was carried out for both forms of PVP, His6-tagged PVP and non-tagged PVP, in similar fashion.

48 Inclusion bodies

Figure 17 - SDS-Page gel after isolation of inclusion bodies

The purification for the two forms of PVP then took different turns because of the

histidine tag functionality. The His6-tagged PVP form was preferably purified using

nickel affinity chromatography (figure 18) while the non-tagged PVP was purified using

ion exchange chromatography (figure 19). His6-tagged PVP was chosen to be worked with first due to personal preferences. Initially two purification steps were thought to be appropriate to achieve the most pure protein possible. The solubilized inclusion bodies were loaded onto a histidine trap column (the nickel affinity chromatography column)

and the first purification step followed. Subsequently, the eluted protein was subjected to

49 Eluted PVP His6-tagged

Figure 18 – Chromatogram of PVP His6-tagged after nickel affinity column; top darker line corresponds to UV absorbance with the peak being the eluted protein and the dotted box showing the flow through after loading; bottom lighter line corresponds to conductivity

refolding to achieve functionally active protein which was then further purified via gel

filtration chromatography. Problems began to arise after the gel filtration

chromatography steps. The protein being eluted was found to be inactive through

enzymatic activity tests with PNPP. It was therefore decided to exclude the gel filtration

step and work with the eluted protein from the nickel affinity column to achieve active,

refolded protein. Purity of the eluted protein is seen with SDS-PAGE. A similar

approach was used for the purification of the non-tagged PVP but instead of using a

nickel affinity column, ion exchange chromatography was used.

Eluted PVP non-tagged

Figure 19 – Chromatogram of PVP non-tagged after ion exchange chromatography; for explanation of lines and dotted box refer to figure above

50 The gel filtration step was again eliminated and refolding conditions were tested after elution from the ion exchange column. As with His6-tagged PVP, a SDS-PAGE gel was ran to check purity of eluted protein (figure 20).

PVP His6-tagged

PVP non-tagged

Figure 20 – SDS-Page gel of purified PVP His6-tagged (left) and PVP non-tagged (right)

3.3 – Refolding of His6-tagged PVP

Since purification of the two forms of PVP was achieved through the

solubilization of the inclusion bodies containing unfolded protein, refolding of the

purified protein was essential in order to obtain functionally active protein. Finding

conditions in vitro for protein refolding is a challenging ordeal with many factors

influencing the outcome. The challenge thus becomes this “hit and miss” project in

which the addition or subtraction of specific additives severely influencing protein 51 refolding conditions are tested. A useful technique to screening different reagents in

refolding buffers is to add the reagents in a positive/negative fashion with

positive/negative meaning either presence or absence of the reagent or, alternatively, a

different concentration of the reagents (Willis 2005). The refolding screen tested in this

study, also included a relative new low molecular weight additive that is a member of a

class of solubilization agents known as non-detergent sulphobetaines (NDSBs) (L. B.-B.

Vuillard 1995). NDSBs have been shown to increase yield of active enzyme on several

proteins including lysozyme and they have been shown to effectively solubilize proteins

at non-denaturing conditions and are, thus, suggested to be ‘anti-aggregants’ in protein

renaturation (Goldberg 1995). Therefore, to enhance the possibility of determining

refolding conditions for the purified PVP protein, several factors including buffer

systems, ionic strength, reducing agents and other additives including the before

referenced NDSBs were tested in the refolding screens.

As with the protein purification step, the first form of PVP that was subjected to

the refolding screen was His6-tagged PVP. As seen in tables 5 and 6, in the materials

and methods section, two refolding screens were tested for the refolding of His6-tagged

PVP. Initially three buffer systems, each with distinct physiological pH believed to be

appropriate for activity of phosphatases, were tested. In conjunction with the buffer system, other factors were introduced to the refolding buffer to see its effect on protein

refolding. In total, thirty buffers were tested in the initial His6-tagged PVP refolding screen. Temperature became an important factor in the purification, refolding and storage of PVP since it was noted that when left at room temperature the protein

52 precipitated. The protein in the refolding buffers was tested for enzymatic activity with

PNPP and the absorbance at 405nm was recorded.

Figure 21 - Enzymatic results after first refolding screen (for buffer reference, see table 3 in

materials and methods section)

From the results shown in figure 21, it appeared that buffer systems three, five,

and ten had the greatest effect in protein refolding, with not much change after incubation

of the enzyme with PNPP after one hour of incubation. These three buffers were then put

through another buffer screen in which an additional NDSB was tested for efficacy in

protein refolding (see table 6). As with the previous results, it appeared that none of the

tested refolding buffers were suitable for the refolding of His6-tagged PVP (figure 22).

53

Figure 22 - Enzymatic activity after second refolding screen (for buffer reference, see table 4 in

materials and methods section)

At this point, it was decided to proceed to the refolding of non-tagged PVP. The refolding of non-tagged PVP proved to be a simpler process than the previously attempted His6-tagged PVP. From information learned from the previous attempts and intuitive knowledge, it was decided to begin with a simple buffer at neutral physiological pH (50mM Tris, pH7). A reducing agent was added to maintain the protein in the reduced state since cysteines in the primary structure have possibilities of forming disulfide bonds. This buffer was initially tested but proved to be ineffective. It was determined, from the previous studies, that ionic strength could play an important role in maintaining the protein in the folded, active state. Additionally, NDSBs were added to the refolding buffer as they are good protein anti-aggregants. After testing three salt concentrations, it was determined that a concentration of 125mM NaCl in the refolding 54 buffer was adequate for protein refolding; buffer without NaCl proved to be inactive and

buffer containing over 250mM NaCl resulted in protein precipitating. It was determined that the best refolding buffer for non-tagged PVP included 50mM Tris, 125mM NaCl,

0.5% β-mercaptoethanol, 0.8M NDSB256, pH7. The protein was added dropwise to the

refolding buffer and allowed to stir overnight at 4°C. The refolded protein was then

tested for enzymatic activity with PNPP and activity recorded with absorbance at 405nm.

The effect of NDSB256 on non-tagged PVP refolding was further explored. To see the extent in which NDSB256 assisted in the refolding of non-tagged PVP, dialysis of

the refolded protein against a refolding buffer without NDSB256 was done. After

dialysis overnight, the protein was tested for activity with PNPP. NDSB256 was also

diluted in the refolding buffer in increments of 0.05M to see the effects on enzyme

activity. It was noted that as NDSB256 concentration decreased, the enzymatic velocity

also decreased and finally ceased at a concentration of 0.16M (figure 23). Therefore,

NDSB256 was necessary to maintain the refolded state of non-tagged PVP.

55

Figure 23 - Results from dilution of NDSB256 in refolding buffer

3.4 – Enzyme kinetics of PVP

After finding suitable conditions for non-tagged PVP protein refolding, enzyme kinetics with various small molecule substrates were carried out (table 7). The first substrate examined was p-nitrophenyl phosphate (PNPP). Since product formed between non-tagged PVP and PNPP was visible, the reaction was continuously monitored at

405nm for 5 minutes using a UV-Vis spectrophotometer. The observed absorbancies

-1 Substrate KM (mM) Vmax (mM/min) kcat (min ) PNPP 4.01 0.31 76.62 pNaph 8.81 0.06 15.1 pSer ------Table 7 – Enzyme kinetics results of PVP with various substrates

56 could then be transformed into concentration of product formed by the use of Beer’s law

and the absorption coefficient of PNPP of 18,000 M-1cm-1 and the reaction velocity was

determined. Subsequent plotting of a Michaelis-Menten graph showed the Michaelis

constant (KM) to be 4.01mM, the maximum velocity (Vmax) to be 0.31 mM/min, and the

-1 turnover number (kcat) value to be 76.62 min (figure 24).

Figure 24 - Michaelis-Menten graph of PVP-PNPP kinetics

The kinetic reaction between non-tagged PVP and the other two substrates, β- naphthyl phosphate (pNaph) and phosphoserine (pSer), were harder to monitor because product formation was not detectable in the visible range. Due to this, quenched phosphatase kinetic assays had to be performed. After performing the phosphatase assay and consequent plotting of a Michaelis-Menten graph, the following kinetic parameters

-1 were found for pNaph: KM = 8.81mM, Vmax = 0.0604 mM/min, kcat = 15.1 min (figure

57 25). Studies with pSer revealed that it was not a substrate of PVP because no enzymatic

activity was seen for any substrate concentration tested.

Figure 25 - Michaelis-Menten graph of PVP-pNaph kinetics

The results obtained for the kinetic characterization of PVP with the three small molecule substrates correlate well with other DSPs. PNPP is known to be a substrate of

PTPs in general, therefore, its activity with PVP was expected. The activity yielded by

the reaction between PVP and PNPP was similar to other DSPs, though a little slower.

For instance, the human DSP VHR, was shown to have a KM of 7.9mM which is within

range of the PVP results – 4.0mM (Denu 1995). However, it was noticed that the

turnover number of PVP was 76.65min-1 (1.27s-1) which is slower that the turnover

number of VHR, 3.65s-1 (Denu 1995). Additionally, the reaction of PVP with pNaph

-1 -1 yielded a high KM, 8.01mM, and a very slow turnover number 15.1min (0.25s ). A

58 possible reason for the slow reaction could be the size of the substrate (figure 26). β-

naphthyl phosphate is a bulkier substrate than p-nitrophenyl phosphate and although the

active site pocket of DSPs is more open, and shallower active site, the bulk of β-naphthyl

phosphate could hinder the proper position of the phosphate in the active site for

catalysis.

O O O2N O P O OH P - O -O OH

β-naphthyl phosphate p-nitrophenyl phosphate

Figure 26 – Structure of β-naphthyl phosphate and p-nitrophenyl phosphate

The most surprising result was the inactivity of PVP with phosphoserine. Dual

specificity phosphatases are known for their ability to dephosphorylate phosphotyrosine

as well as phosphoserine and phosphothreonine substrates. Possibilities for no activity

seen with phosphoserine could be that the reaction was too slow to be detectable or that the enzyme used in the assay was not in the proper refolded state. This could also explain the slow reaction with the other two substrates.

59

CHAPTER 4 – RESULTS: CLONING, CELL GROWTH,

EXPRESSION, PURIFICATION AND KINETIC

CHARACTERIZATION OF LOW MOLECULAR WEIGHT

PHOSPHATASE, WZB

4.1 – Growth and expression of WZB

WZB, a low molecular weight phosphatase, was cloned in two forms; both forms

contained a histidine tag at either the N- or C- terminus. WZB was cloned into pET

expression vectors with the N-terminal histidine tag WZB (His6-tagged WZB) being cloned into pET28a vector and the C-terminal histidine tag WZB (WZB His6-tagged) cloned into the pET29b expression vector. The WZB plasmids were transformed into

Origami (DE3) cells ideal for recombinant protein expression. Conditions for expression of soluble protein were quickly found and all subsequent cell growth and protein expression was conducted using found method.

Standard conditions for cell growth were initially tried. Cells were inoculated in

LB medium and incubated at 37°C in shaker and upon reaching the appropriate OD, the protein expression was induced by addition of 1mM IPTG and expression was allowed to proceed for 4-8 hours. The solubility of the protein was tested by enzymatic activity tests

60 with PNPP and the level of protein expression was determined by SDS-Page. The above

conditions proved to give soluble protein after kinetic tests with PNPP. Further

experiments were made to see if the temperature after protein induction played a role in

the solubility of protein expressed. Therefore, after the cell OD600 reached 0.8, protein

expression was induced with 1mM IPTG and the temperature was lowered to room

temperature and again expression was allowed to proceed for 4-8 hours. Solubility of the

expressed protein was tested with PNPP and level of expression was shown by SDS-

PAGE. As with the first attempt, the protein expressed was shown to be soluble and the

level of expression was comparable between the two methods. It was therefore decided

to use induction at room temperature due to personal preferences, though both methods worked equally.

4.2 – Purification of WZB

The purification of both forms of WZB (figure 27) proved to be fairly straight

forward with the same method being able to be used for both. The purification protocol

began with cell lysis triggered by the addition of lysozyme followed by sonication. Due

to the functionality of the histidine tag, initial protein purification was achieved with a

histidine trap, nickel affinity column. The eluted protein was obtained in a partially

purified form, as seen by SDS-PAGE. Therefore the eluted protein was subsequently run

through a gel filtration column. Purity of the purified protein was seen through SDS-

PAGE (figure 28).

a) Nickel affinity chromatogram

61 Eluted WZB His6-Tagged

b) Gel filtration chromatogram

Eluted WZB His6-tagged

Figure 27 - Chromatogram of eluted WZB (a) His6-tagged after nickel affinity column and (b) gel filtration; in both spectra the top darker line corresponds to UV absorbance and bottom lighter line corresponds to conductivity

Throughout the purification steps, several obstacles had to be overcome. One of the most important factors for protein stability during protein purification was temperature. Since the protein was most stable in the cold cabinet, a temperature of 4°C, the entire purification chromatography steps were thus performed in the cold cabinet and cell lysis was done while the protein was in ice. Additionally, after purification, the eluted protein had to be maintained at 4°C to avoid protein precipitation. Maintaining the protein in a soluble, active state also proved to be challenging. It was noted that the protein was best stable under conditions with low salt concentration as well as acidic pH (below 5).

Addition of other additives, such as glycerol, caused the protein to precipitate and

62 become inactive. As noted before, the temperature had to also be closely monitored with

best results at 4°C and freezing of the protein resulting in protein precipitation. Due to

these factors, it was determined that purification of the protein had to be done fairly

quickly with all the purification steps done in one day and use of the protein for further

analysis quickly following on the next day.

WZB eluted form WZB eluted from gel nickel affinity column filtration column

Figure 28 - SDS-Page gel of eluted WZB from nickel affinity column and gel filtration column

4.3 – Enzyme kinetics of WZB

After obtaining the purified protein, enzyme kinetics between WZB and various small molecule substrates were performed. Initially, the enzyme kinetic reactions were carried out in a phosphate buffer with pNPP as the substrate. However, it was seen that the enzymatic activity was not desirable with KM of above 2.5mM, as it is known that the

approximate Km of WZB with pNPP is approximately 1mM (Vincent 1999), and these

were not the results seen with the phosphate buffer. Therefore, a comparison of the

effects of different buffers on enzyme activity was tested. Three buffers – citrate, tris and

63 phosphate – at pH6 were tested and the turnover number, kcat, were compared. It was determined (figure 29) that a Tris buffer was the most suitable buffer for enzyme activity.

Initially, a buffer of 50mM Tris pH6 was used but since Tris has low buffer capacity at low pH, it was decided to use a mixture of Tris and BisTris as the buffer system.

Therefore a buffer consisting of 50mM Tris, 50mM BisTris, 50mM NaCl, AcOH was used for further kinetic studies.

Figure 29 - Buffer effect on WZB enzyme activity with all buffers at pH6

Prior to commencing the kinetic reactions of the enzyme with the small molecule substrates, a study to determine the optimal pH for enzyme activity was first done. The kinetic parameters, kcat, KM, and kcat/KM, were determined for WZB with pNPP as the substrate over a pH range of 4.5 to 7.5 in increments of 0.5. Comparison of the three kinetic parameters revealed that pH 5 was the

64 a)

b)

65 c)

Figure 30 - Kinetic parameters of WZB with pNPP as substrate over pH range of 4.5 to 7.5: a) kcat

values, b) KM values, c) kcat/KM values

optimal pH for enzyme activity, as seen in figure 30, not a surprising result as it is known

that most low molecular weight phosphatases work best under acidic conditions. The pH

study also revealed that as the pH was increased, activity of the enzyme decreases

dramatically and almost lost all activity by pH 7.5. Comparison of the kcat values revealed that at pH 4.5 and 5 kcat values were comparable with the turnover number being

-1 approximately 38s and as the pH was increased the kcat values decreased with almost no

turnover at pH 6.5 and 7, and no activity at pH 7.5. A look at the KM values throughout

the entire pH range showed little fluctuation between the pH range 5.0 to 7.0 with KM

values ranging from 0.9mM to 1.45mM. In contrast the Km at pH 4.5 jumped to almost

3mM. Additionally, the efficiency of the enzyme was determined by the kcat/KM ratio.

-1 -1 The best enzyme efficiency was seen at a pH of 5 with the kcat/KM of 35mM s whereas

-1 -1 the kcat/KM for the other pH values were all under 15mM s with a steady decrease as the 66 pH was raised. Overall, taking into consideration the three kinetic parameters tested, it

was found that optimal enzyme activity could be achieved at a pH of 5 since it produced

a high turnover number, a low Michaelis constant and showed high efficacy with the

kcat/Km parameter. Therefore, the buffer used for further kinetic studies consisted of

50mM Tris, 50mM BisTris, 50mM NaCl, AcOH pH 5. The above results are in

correlation with other LMW-PTPs (Vasilios 2004).

Figure 31 - Michaelis-Menten plot of WZB with pNPP as substrate

Upon finding of the optimal conditions for WZB activity, kinetic studies with several small molecule substrates were conducted. The first substrate tested was pNPP.

The reaction between WZB and pNPP releases p-nitrophenol which concentration increase during the course of the reaction can be monitored at 405nm. The reaction was allowed to proceed for 80 seconds and quenched every 10 seconds with 5M NaOH. The amount of product released was determined from the absorbance readings at 405nm and the absorption coefficient of pNPP – 18,000M-1cm-1. A Michaelis-Menten graph was 67 plotted and the kinetic parameters were determined. For the substrate pNPP, the

Michaelis constant, KM, was found to be 1.08±0.12mM and the turnover number (kcat),

determined from the maximum velocity of the reaction and the enzyme concentration,

was found to be 37.59±8.34s-1 (figure 31). Another two small molecule substrates, β-

naphthyl phosphate and phosphoserine, were tested for affinity to WZB. Since the

release of inorganic phosphate catalyzed by these reactions is colorless, a new

colorimetric assay had to be designed to measure the amount of phosphate released. A

modified version of the ammonium molybdate assay was used (Katewa 2003). A

standard calibration curve with potassium phosphate was used to determine the

experimental absorbance coefficient for the assay, 8000 M-1cm-1. The two substrates

were tested in range from 1mM to 15mM and no reaction was observed. Therefore, it

was determined that neither β-naphthyl phosphate or phosphoserine are substrates for

WZB.

4.4 – Inhibitory effects on WZB

The inhibition of WZB was tested with several compounds (phosphate, fluoride, hydrogen peroxide, molybdate) under the buffer conditions previously determined –

50mM Tris, 50mM BisTris, 50mM NaCl, AcOH pH5; the inhibition constant and type of inhibition was determined for each of the inhibitors tested – phosphate, fluoride, hydrogen peroxide, and molybdate – all sodium salts except for hydrogen peroxide. A

Michaelis Menten curve was plotted for each of the reactions in the presence of inhibitor.

The Michaelis Menten curve showed that inhibition by all compounds except hydrogen

68 peroxide was competitive. It was found that the Michaelis constant, KM, increased as the amount of inhibitor was increased and that the maximum velocity, Vmax, of the curve,

remained relatively the same. To confirm this finding, the Michaelis Menten curves were

converted to Lineweaver Burk plots. For competitive inhibitors, all the reciprocal plots

from Michaelis Menten curve will intercept at the y-axis whereas non-competitive

inhibitors intercept the x-axis and mixed inhibition usually intercept at other parts of the

Lineweaver Burk plot. The Lineweaver Burk plots showed that phosphate, molybdate

and fluoride all intercepted at the y-axis, thus confirming that they were competitive

inhibitors of WZB, while hydrogen peroxide was found to have a suicide inhibition effect

on WZB. In order to determine the inhibition constant, Ki, for these inhibitors a Dixon plot was used to see the affinity of these inhibitors to WZB. Interestingly, it was found that the inhibitor with the lowest inhibition constant was molybdate with 50.4μM and the other two inhibitors, phosphate and fluoride, had much higher inhibition constants,

3.2mM and 71.7mM, respectively (table 8, figure 32).

Type of Inhibitor K Std Dev Inhibition i phosphate competitive 3.2mM ±0.6mM hydrogen suicide-like - - peroxide fluoride competitive 71.7mM ±27.0mM molybdate competitive 50.4μM ±7.8μM Table 8 - Results of inhibition studies with WZB

To better understand why these salts are competitive inhibitors of WZB and the reason behind their wide range inhibition activities, a closer look at their properties and structure was taken.

69 a)

70 b)

71 c)

Figure 32 – Lineweaver Burk and Dixon plots of the inhibitors acting on WZB – a) phosphate; b) fluoride; c) molybdate

72 All inhibitors tested were sodium salts except for hydrogen peroxide. The first

inhibitor to be tested was phosphate and it was determined to be a competitive inhibitor

with the second best inhibition constant of 3.155mM. Phosphate as an inhibitor would

actively compete for the active site with any substrate and, therefore, be a competitive

inhibitor. To understand the mechanism of why vanadate and molybdate are competitive

inhibitors of WZB as well, a quick look at their structure (figure 33) in comparison to phosphate is taken. Vanadate and molybdate, metal oxyanions, are transition state analogs of phosphate and adopt a similar shape to phosphate, a tetrahedral structure, as

seen below. These early transition state metals oxoanions have been shown to be potent

inhibitors of PTPs (Zhang M. Z. 1997, Zhang Z. 1990).

O O O

P V Mo HO O- HO O- HO O OH OH O-

Figure 33 - Tetrahedral structure of phosphate, vanadate and molybdate anions

It has long been thought, that these metal oxoanions, are potent inhibitors of PTPs

because they bind as transition state analogs, through covalent bonds, to the enzyme due to their ability to form a trigonal bipyramidal structure. They are also thought to be better inhibitors than phosphate because of their rate of exchange at the central atom, with oxygen exchange with the oxyanions being faster than with phosphate (M. Z. Zhang

1997).

Vanadate and molybdate have several effects on biological systems. Vanadate is an insulin mimetic and has been shown to inhibit many kinases and phosphatases while both vanadate and molybdate can act as stabilizers of glucocorticoid receptors (M. Z.

73 Zhang 1997). The mechanism by which these oxyanions bind to PTPs and, thus, inhibits

them has not been well investigated in the past, but recent studies have given more

information as to their mechanism of inhibition (Huyer 1997). Recently, solving of crystal structures of PTPs with the transition metal analogs, revealed that the oxyanions bind to the phosphatase at the catalytic cysteine residue of the PTP loop (Madhurantakam

2007, Huyer 1997). For instance, it was determined that when the Yersinia PTP is complexed with vanadate, the vanadate is in the active site within covalent linkage between the thiol of the catalytic cysteine residue, forming a thiol-vanadyl linkage similar to the thiol-phosphate linkage formed during catalysis (Huyer 1997). This is similar to studies with PTP1B and molybdate, which showed that molybdate adopted a trigonal bipyramidal structure at the active site and showed a linkage between molydeum at the thiol group of the catalytic cysteine residue. Additionally, the oxygen atoms of molybdate formed hydrogen bonds to the amino groups in the backbone of the PTP signature motif (Heo 2002).

Another inhibitor tested was fluoride and it was found to be a competitive inhibitor of WZB. However, it was shown to have a lower affinity to WZB with a ki of

70mM. This is not a surprising finding as it is known that fluoride is a weak inhibitor of

low molecular weight PTPs such as WZB (Z. Zhang 2002). This key feature, the weak

inhibition by fluoride, is one of the characteristics that differentiates LMW-PTPs from

the higher molecular weight PTPs (Z. Y. Zhang 1990). The last inhibitor tested on WZB

was hydrogen peroxide. Initially, hydrogen peroxide was treated as the previous

inhibitors, but it was seen that the longer the hydrogen peroxide was left in solution with

the purified enzyme, the reaction rate slowed down, thus not exhibiting steady state

74 kinetics behavior. Therefore, the hydrogen peroxide was allowed to incubate with WZB

for specific times and, at a hydrogen peroxide concentration of 0.333mM, under these

experimental conditions, it was found that after three minutes most of WZB activity

(figure 34) ceased leading to the assumption that hydrogen peroxide alters a part of the

catalytic site. Upon re-evaluation of the properties of hydrogen peroxide, it was

concluded that hydrogen peroxide, as an oxidizer, reversibly oxidizes the catalytic cysteine residue in the PTP catalytic motif which leads to inactivation of the enzyme.

Figure 34 – Effect of hydrogen peroxide on WZB activity

Recently, other studies have led to the finding that oxidation by hydrogen peroxide can lead to the formation of a disulfide bond between the catalytic cysteine and a neighboring cysteine (five amino acids away) (Chiarugi 2005). Formation of this disulfide bond

(figure 35) is reversible by addition of a reducing agent. A possible biological function for the formation of the disulfide bond is to protect the oxidized catalytic cysteine from further irreversible oxidation. The disulfide bond is reversible with addition of reducing 75 agent such as DTT, however, further oxidation of the cysteine to cysteine-sulfinic acid

(Cys-SO2H) or cysteine-sulfonic acid (Cys-SO3H) is irreversible (Cho 2004).

Figure 35 – Oxidation of catalytic cysteine (12) followed by disulfide bond formation with neighboring cysteine (17) (Cho 2004)

4.5 – Comparison of enzymatic activity between two forms of WZB

At the beginning of the project two forms of WZB were cloned - His6-tagged

WZB and WZB His6-tagged. All subsequent kinetic studies were performed with the

His6-tagged form. In the present study, the kinetic assays for the two forms were

compared to assess whether the position of the histidine tag influenced the enzyme’s

activity. As the results below show, there is no significant difference in enzymatic activity between the two forms of WZB, leading to the conclusion that the position of the histidine tag had no bearing on the enzyme’s activity (table 9, figure 36).

-1 -1 Form KM, mM kcat/KM, s mM His6-tagged WZB 1.09±0.11 32.84±8.83 WZB His6-tagged 0.70±0.11 26.16±1.10 Table 9 – Comparison of WZB activity between two forms of WZB

76

Figure 36 – Michaelis-Menten plots for the two forms of WZB

4.6 – Enzyme activity of WZB mutants

Using site-directed mutagenesis, two His6-tagged WZB mutants were created –

Cys9Ser and Asp115Asn. It is known that the catalytic mechanism of PTPs involves three important amino acid residues – a cysteine, an arginine, and an aspartic acid (figure 77 39). The catalytic cysteine is essential in forming the covalent phosphoenzyme

intermediate through the formation of the thiol-phosphate bond. The arginine is important in orienting the substrate towards the thiol group of the cysteine as well as forming key hydrogen bonds and stabilization of the transition state. The aspartic acid is also important by acting as a general acid in the first step of the catalytic mechanism and

as a general base in the second step. It is expected that mutation of any of the above

amino acids would inactivate the enzymatic activity or dramatically reduce it. First Step Second Step

Figure 37 – Role of cysteine, arginine, and aspartic acid in catalytic mechanism (Z. Y. Zhang 1998)

After purification (figure 40) and subsequent testing of the enzyme’s activity using pNPP as the substrate, it was observed that the activity of the Cys9Ser mutant was completely abolished, which is in support of other studies where the catalytic cysteine was mutated using site directed mutagenesis (Zhang Z. 2003, Zhang Z. Y. 1998). The other His6-tagged WZB mutant, Asp115Asn, yielded similar results. Upon testing the

enzymatic activity with pNPP, in standard conditions, no activity was observed. Other

studies in which the aspartic residue was mutated showed a significant decreased in the

phosphatase activity. To further support the results of the enzymatic assay with 78 Asp115Asn, a new enzymatic assay was performed over a long period of time (over one

hour) and using higher concentration of pNPP (15mM). However, no activity was

determined. These experiments clearly showed that mutation at the cysteine and aspartic

acid residue severely influences WZB acitivity to the point of completely abolishing it.

75kDa 75kDa 50kDa 50kDa 37kDa 37kDa 25kDa 20kDa 25kDa Asp115Asn mutant 20kDa Cys9Ser 15kDa mutant 10kDa 10kDa

Figure 38 – SDS-Page of Cys9Ser WZB mutant (left) and Asp115Asn WZB mutant (right)

79

CHAPTER 5 – SUMMARY

Protein phosphatases are a large superfamily of enzymes that catalyze the removal

of phosphate from specific amino acids residues. The superfamily can be further divided

into families according to their catalytic mechanism and substrate specificity. One of

these families is the protein tyrosine phosphatases (PTPs). PTPs are a large family of

enzymes that dephosphorylate primarily phosphotyrosine, and to some extent,

phosphoserine and phosphothreonine amino acid residues. This large superfamily can be further broken down into four subfamilies – the classical phosphotyrosine phosphatases, the dual specificity phosphatases, the Cdc25 phosphatases, and the low molecular weight phosphatases – with each of these subfamilies having the ability to be further subdivided.

Even though these subfamilies share very little similarities and appear rather diverse, they all possess the signature PTP catalytic motif, C(X)5R, which allows all of them to follow

the same catalytic mechanism. In the present study, two members of the PTP family

were investigated, the dual specificity phosphatase PVP and the low molecular weight

phosphatase WZB.

PVP from the smallpox virus, is a member of the dual specificity subfamily of

PTPs specifically the VH1-like DSPs. Interestingly, it has been shown that members of

orthopoxviruses, all contain this conserved dual specificity phosphatase encoded in their

genome. It has been suggested that this conserved PTP plays a role in the virus’

80 pathogenesis. In this study, two forms of the recombinant enzyme were cloned into the

pET101 vector, PVP His6-tagged and PVP non-tagged. After expression, under several

conditions tested, most protein was in the form of inclusion bodies. Therefore, prior to

purification, solubilization of the inclusion bodies in urea was necessary. Purification of

the His6-tagged form was done using a nickel affinity column. The purification of the

non-tagged form was done using an ion exchange column and refolding conditions were

found, a buffer consisting of 25mM Tris, 125mM NaCl, 0.5% β-mercaptoethanol, 0.8M

NDSB256, pH7. Enzyme kinetics of PVP with several small molecule substrates

revealed that pNPP and pNaph were substrates while pSer was not. Using pNPP, it was

-1 found that the KM = 4.01mM and the kcat = 76.62min . For pNaph, the KM = 8.81mM

-1 and the kcat = 15.1 min .

WZB is a low molecular weight phosphatase found in E.coli. Along with the

protein phosphatase kinase WZC, WZB aids in the transport and synthesis of colanic

acid. When WZC is phosphorylated no colanic acid is synthesized, but, if, WZC is

dephosphorylated by WZB, colanic acid is produced. Colanic acid is an

exopolyssacharide which are believed to mediate the interaction between the bacteria and

its surrounding environments. This interaction is believed to be an important factor in the

virulence of the bacteria and thus, may be part of pathogenicity of the bacteria. In the

present study, two forms of WZB, both with histidine tags at either the N- or C- terminus,

were cloned. Protein expression was straight forward and soluble protein was acquired.

Purification methods for both forms of WZB were the same starting with a nickel affinity

column followed by gel filtration. Subsequent kinetic studies were performed with His6-

tagged WZB. To determine the best buffer, several buffers systems were tested and the

81 enzymatic activity was compared. Determination of the most active pH was done using

the optimal buffer found – 50mM Tris, 50mM BisTris, 50mM NaCl, AcOH. Comparison of the KM, kcat, and kcat/KM of all the pH’s tested, led to the conclusion that at pH5 the

enzyme had its optimal activity. Kinetic studies with several small molecule substrates

revealed that pNPP was the only substrate, while pNaph and pSer not being substrates of

WZB. The kinetic parameters of WZB with pNPP as substrate were: KM = 1.08mM and

-1 kcat = 37.59s . After studying the enzyme kinetics of WZB with pNPP, several

compounds were tested to determine their inhibitory effects on WZB. Phosphate,

fluoride, and molybdate were found to all be competitive inhibitors with molybdate being

the best inhibitor with a Ki of 50.4μM. Phosphate and fluoride both had inhibition

constants in the millimolar range, 3.2mM and 71.7mM respectively. The last compound

tested was hydrogen peroxide and it was determined that incubation of the enzyme with

hydrogen peroxide led to the inactivation of the enzyme. This might be due to the

oxidative characteristic of hydrogen peroxide; it oxidizes the catalytic cysteine in the

active site and ceases the activity of WZB.

There were two forms of WZB cloned, which differed in the position of the

histidine tag. It was found that the position of the histidine tag did not affect the catalytic

activity of the enzyme. Two active site mutants of WZB were generated: Cys9Ser and

Asp115Asn. Both mutants activity showed no clearly detectable activity, supporting the assumption that the Cys9 and Asp115 act as a general acid/base and primary nucleophile in the catalytic cycle.

82

REFERENCES

Barford, D. "Protein phosphatases." Curr. Opin. Struct. Biol. 5 (1995): 728-734.

Barford, D.S., Das, A.K., and Egloff, M. "The structure and mechanism of protein

phosphatases: insights into catalysis and regulation." Annu. Rev. Biophys. Biomol.

Struct., 27 (1998): 133-164.

Bottini, N., Vang, T., Cucca, F., and Mustelin, T. "Role of PTPN22 in type 1 diabetes and

other autoimmune diseases." Semin. Immunol. 18 (2006): 207-213.

Buotros, R., Dozier, C. and Ducommun, B. "The when and wheres of CDC25

phosphatases." Curr. Opin. Cell Biol. 18 (2006): 185-191.

Burke, T.R. and Zhang, Z. Y. "Protein-tyrosine phosphatases: structure, mechanism, and

inhibitor discovery." Biopolymers (Peptide Science) 47 (1998) 225-241.

Cheng, A., Dube, N., Gu, F., and Tremblay, M.L. "Coordinated action of protein tyrosine

phosphatases in insulin signal transduction." Eur. J. Biochem. 269 (2002): 1050-

1059.

Chiarugi, P. "PTPs versus PTKs: the redox side of the coin." Free Rad. Research 39

(2005): 353-364.

83 Cho, S.H., Lee, C.H., Ahn, Y., Kim, H., Kim, H., Ahn, C.Y., Yang, K.S., and Lee S.R.

"Redox regulation of PTEN and protein tyrosine phosphatases in H2O2-mediated

cell signaling." FEBS Letters 560 (2004): 7-13.

Cozzone, A.J., Grangease, C., Doublet, P., and Duclos, B. "Protein phosphorylation on

tyrosine in bacteria." Arch. Microbiol. 81 (2004): 171-181.

Cozzonne, A.J. "Protein phosphorylation in prokaryotes." Annu. Rev. Microbiol. 51

(1988): 97-125.

Crans, D.C, Bunch, R.L., and Theisen, L.A. "Interaction of trace levels of vanadium (IV)

and vanadium (V) in biological systems." JACS 111 (1989): 7597-7607.

Denu, J.M., Zhou, G., Zhao, R., Yuvaniyama, J., Saper, M.A., and Dixon, J.E. "The

purification and characterization of a human dual-specific protein tyrosine

phosphatase." J. Biol. Chem. 270 (1995): 3796-3803.

Ding, X.L., Husseman, J., Tomashevski, A., Nochlin, D., Jin, L.W., and Vincent, I. "The

cell cycle Cdc25A tyrosine phosphatase is activated in degenerating postmitotic

neurons in Alzheimer's disease." Am. J. Patho. 157 (2000): 1983-1990.

Ducruet, A.P., Vogt, A., Wipf, P. and Lazo, J.S. "Dual specificity protein phosphatases:

therapeutic targets for cancer and Alzheimer's disese." Annu. Rev. Pharmacol.

Toxicol. 45 (2005): 725-750.

84 Goldberg, M.E., Expert-Bezancon, N., Vuillard, L., and Rabilloud, T. "Non-detergent

sulphobetaines: a new class of molecules that facilitate in vitro protein

renaturation." Folding & Design 1 (1995): 21-27.

Grabski, A., Spangler, N., Ginos, M., and Drott, D. "Overnight express instant TB

medium: autoinduction of protein expression is now even easier!" inNovations 21

(2003): 3-7.

Grangeasse, C., Obadia, B., Mijakovic, I., Deutscher, J., Cozzone, A.J., and Doublet, P.

"Autophosphorylation of the Escherichia coli protein kinase Wzc regulates

tyrosine phosphorylation of Ugd, a UDP-glucose dehydrogenase." J. Biol. Chem.

278 (2003): 39323-39329.

Hagelueken, G., Huang, H., Mainprize, I.L., Whitfield, C., and Naismith, J.H. "Crystal

structures of Wzb of Escherichia coli and CpsB of Streptococcus pneumoniae,

representatives of two families of tyrosine phosphatases that regulate capsule

assembly." J. Mol. Biol. 392 (2009): 678-688.

Hakes, D.J., Martell, K.J., Zhao, W.G., Massung R.F., Esposito, J.J., and Dixon, J.E. "A

protein phosphatase related to the vaccinia virus VH1 is encoded in genomes of

several orthopoxviruses and a baculovirus." Proc. Natl. Acad. Sci. USA 90 (1993):

4017-4021.

Heo, Y.S., Ryu, J.M., Park, S.M., Park, J.H., Lee, H.C., Hwang, K.Y., and Kim, J.

"Structural basis for inhibition of protein tyrosine phosphatases by Keggin

85 compounds phosphomolybdate and phosphotungstate." Exp. Mol. Med. 34 (2002):

211-223.

Huyer, G., Liu, S., Kelly, J., Moffat, J., Payette P., Kennedy, B., Tsaprailis, G., Gresser,

M.J., and Ramachandran, C. "Mechanishm of inhibition of protein-tyrosine

phosphatases by vanadate and pervanadate." J. Biol. Chem 272 (1997): 843-851.

Ingebritsen, T.S. "Serine/threonine phosphatases." Encyclopedia of Biol. Chem. 4 (2004):

27-32.

Katewa, S.D. and Ketyare, S.S. "A simplified method for inorganic phosphate

determination and its application for phosphate analysis in enzyme assays." Anal.

Biochem. 323 (2003): 180-187.

Kennelly, P.J., and Potts, M. "Life among the primitives: protein o-phosphatases in

prokaryotes." Front. Biosci. 4 (1999): d372-385.

Kerk, D., Templeton, G. and Moorehead, G.B.G. "Evolutionary radiation pattern of novel

protein phosphatases revealed by analysis of protein data from the completely

sequenced genomes of humans, green algae, and higher plants." Plant Phys. 146

(2008): 351-367.

Kristjansdottir, K. and Rudolph, J. "Cdc25 phosphatases and cancer." Chem. Biol. 11

(2004): 1043-1051.

Laporte, J., Hu, L.J., Kretz, C., Mandel, J.L., Kioschis, P., Coy, J.F., Klauck, S.M.,

Poutska, A., and Dahl, N. "A gene mutated in X-linked myotubular myopathy

86 defines a new putative tyrosine phosphatase family conserved in yeast." Nat.

Genet. 13 (1996): 175-182.

Lescop, E., Hu, Y., Xu, H., Hu, W., Chen, J., Xia, B., and Jin, C. "The solution structure

of Escherichia coli Wzb reveals a novel substrate recognition mechanism of

prokaryotic low molecular weight protein tyrosine phosphatases." J. Biol. Chem.

285 (2006): 19570-19577.

Li, Liwu and Dixon, J.E. "Form, function, and regulation of protein tyrosine

phosphatases and their involvement in human diseases." Sem. Immunol. 12 (200):

75-84.

Madhurantakam, C., Chavali, V.R.M., and Das, A.K. "Analyzing the catalytic

mechanism of MPtpA: a low molecular weight protein tyrosine phosphate from

Mycobacterium tuberculosis through site-directed mutagenesis." Proteins: Struc.,

Func., Bioninfor. 71 (2007): 706-714.

Mann, B.A., Huang, J.H., Li, P., Chang, H.C., Slee, R.B., O'Sullivan, A., Mathur, A.,

Yeh, N., Klemsz, M.J., Brutkiewicz, R.R., Blum, J.S., and Kaplan, M.H.

"Vaccinia virus blocks Stat1-dependent and Stat1-independent gene expression

induced by type I and type II interferons." J. Inter. & Cyt. Research 28 (2008):

367-380.

Manning, G., Whyte, D.B., Martinez, R., Hunter, T., and Sudarsanam, S. "The protein

kinase complement of the human genome." Science 298 (2992): 1912-1934.

87 McCain, D.F. and Zhang, Z.Y. "Assays for protein tyrosine phosphatases." In Methods in

Enzymology, 507-518. Academic Press, 2002.

Mildvan, A.S. "Mechanisms of signaling and related enzymes." Proteins: Struct., Func.,

and Gen. 29 (1997): 401-416.

Moorehead, G.B.G., De Wever, V., Templeton, G. and Kerk, G. "Evolution of protein

phosphatases in plants and animals." Biochem. J. 417 (2009): 401-409.

Morona, J.K, Morona, R., Miller, D.C., and Paton, J.C. "Streptococcus pneumoniae

capsule biosynthesis protein CpsB is a novel manganese-dependent

phosphotyrosine-protein phosphatase." J. Bacteriol. 184 (2002): 577-583.

Nagy, Z.S., Esiri, M.M., and Smith, A.D. "The cell division cycle and the

pathophysiology of Alzheimer's disease." Neuroscience 87 (1998): 731-739.

Najarro, P., Traktman, P., and Lewis, J.A. "Vaccinia virus blocks gamma interferon

signal transduction: viral VH1 phosphatase reverses Stat1 activation." J. Virol. 75

(2001): 3185-3196.

Ninfa, E.G. and Dixon, J.E. "Protein tyrosine phosphatases in disease processes." Trends

in Cell Biol. 4 (1994): 427-430.

Pulido, R. and Hooft van Huijsduijnen, R. "Protein tyrosine phosphatases: dual-

specificity phosphatases in health and diease." FEBS Journal 275 (2008): 848-

866.

88 Raetz, C.R.H., and Whitfield, C. "Lipopolysaccharide Endotoxins." Annu. Rev. Biochem.

71 (2002): 635-700.

Reid, A.N., and Whitfield, C. "Functional analysis of conserved gene products involved

in assembly of Escherichia coli capsules and exopolysaccharides: evidence for

molecular recognition between Wza and Wzc for colanic acid biosynthesis." J.

Bacteriol. 187 (2005): 5470-5481.

Rudolph, J. "Cdc25 phosphatases: structure, specificty, and mechanism." Biochemistry 46

(2007): 3595-3604.

Shi, L., Potts, M., and Kennelly, P.J. "The serine, threonine, and/or tyrosine-specific

protein kinases and protein phosphatases of prokaryotic organisms: a family

portrait." FEMS Microbiol. Rev. 22 (1998): 229-253.

Singh, S.M., and Panda, A.K. "Solubilization and refolding of bacterial inclusion body

proteins." J. Biosci. Bioeng. 99 (2005): 303-310.

Stauffacher, C. V. and Charbonneau, H. "Protein tyrosine phosphatases." In Principles of

Molecular Recognition, by P.M. and Means, A. R., eds Conn, 323-347. Totowa,

NJ: Humana Press Inc., 2000.

Stoker, A.W. "Protein tyrosine phosphatases and signaling." J. Endocrin. 185 (2005): 19-

33.

89 Tabernero, L., Ariscescu, A.R., Jones, E.Y., and Szedlacsek, S.E. "Protein tyrosine

phosphatases: structure-function relationships." FEBS Journal 275 (2008): 867-

882.

Takagi, T., Moore, C.R., Diehn, F., and Buratowski, S. "An RNA 5'-triphosphatase

related to the protein tyrosine phosphatases." Cell 89 (1997): 867-873.

Tonks, N.K. "Overview of protein tyrosine phosphatases." In Handbook of Cell

Signaling, Volume 1, 641-651. USA: Elsevier Science, 2003.

Tonks, N.K. "Protein tyrosine phosphatases." In Signal Transduction, 253-269. Chapman

& Hill, 1996.

Tonks, N.K. "Protein tyrosine phosphatases: from genes, to function, to disease." Nat.

Rev. 7 (2006): 833-846.

Tracey, A.s., and Gresser, M.J. "Vanadium (V) oxyanions: interaction of vanadate with

1,1,1-tris(hydroxymethyl)ethane and with the buffer

tris(hydroxymethyl)aminomethane." Inorg. Chem. 27 (1988): 1269-1275.

Tremblay, M.L. "The PTP family photo album." Cell 136 (2009): 213-214.

Van Etten, R.L. "Low molecular weight protein tyrosine phosphatases." In Handbook of

Cell Signaling, Volume 1, 733-740. USA: Elsevier Science, 2003.

Vang, T., Miletic, A.V., Arimura, Y., Tautz, L., Rickert, R.C., and Mustelin, T. "Protein

tyrosine phosphatases in autoimmunity." Annu. Rev. Immunol. 26 (2008): 29-55.

90 Vasilios, M, and Ross, J.D. "Isolation and partial characterization of acid phosphatase

isoenzymes from dormant oilseed of Corylus avellana L." Planta 219 (2004):

346-358.

Vincent, C., Doublet, P., Grangeasse, C., Vaganay, E., Cozzone, A.J., and Duclos, B.

"Cells of Escherichia coli contain a protein-tyrosine kinase, Wzc, and a

phosphotyrosine-protein phosphatase, Wzb." J. Bacteriol. 181 (1999): 3472-3477.

Vuillard, L., Braun-Breton, C., and Rabilloud, T. "Non-detergent sulphobetaines: a new

class of mild solubilization agents for protein purification." Biochem. J. 305

(1995): 337-343.

Vuillard, L., Rabilloud, T., and Goldberg, M.E. "Interactions of non-detergent

sulfobetaines with early folding intermediates facilitate in vitro protein

renaturation." Eur. J. Biochem. 256 (1998): 128-135.

Walker, S.M., Downer, R. and Leslie, N.R. "TPIP: a novel phosphoinositide 3-

phosphate." Biochem. J. 360 (2001): 277-283.

Wang, F., Barrett, J.W., Shao, Q., Gao, X., Dekaban, G.A., and McFadden, G. "Myxoma

virus selectively disrupts type I interferon signaling in primary human fibroblasts

by blocking the activation of the Janus kinase Tyk2." Virology 387 (2009): 136-

146.

Wang, W.Q., Sun, J.P., and Zhang, Z. Y. "An overview of the protein tyrosine

phosphatase superfamily." Curr. Top. Med. Chem. 3 (2003): 739-748.

91 Willis, M.S., Hogan, J.K., Prabhakar, P., Liu, X., Tsai, K., Wei, Y., and Fox, T.

"Investigation of protein refolding using a fractional factorial screen: a study of

reagent effects and interactions." Pro. Sci. 14 (2005): 1818-1826.

Zhang, M. Zhou, M., Van Etten, R.L., and Stauffacher, C.V. "Crystal structure of bovine

low molecular weight phosphotyrosyl phosphatase complexed with the transition

state analog vanadate." Biochemistry 36 (1997): 15-23.

Zhang, W. and Shi, L. "Evolution of PPM-family protein phosphatases in Streptomyces:

duplication of catalytic domain and lateral recuitment of additional sensory

domains." Microbiology 150 (2004): 4189-4197.

Zhang, Z. Y. "Protein-tyrosine phosphatases: biological function, structural

characteristics, and mechanism of catalysis." Cri. Rev. Biochem. Mol. Biol. 33

(1998): 1-52.

Zhang, Z.Y. and Van Etten, R.L. "Purification and characterization of a low-molecular-

weight acid phosphatase - a phosphotyrosyl protein phosphatase from bovine

heart." Arch. Biochem. Biophys. 282 (1990): 39-49.

Zhang, Z.Y. "Chemical and mechanistic approached to the study of protein tyrosine

phosphatases." Acc. Chem. Res. 36 (2003): 385-392.

Zhang, Z.Y. "Protein tyrosine phosphatases: structure and function, substrate specificity,

and inhibitor development." Annu. Rev. Pharmacol. Toxicol. 42 (2002): 209-234.

92