ABSTRACT Development of an Aptamer-Based Detection Assay

ABSTRACT

Development of an Aptamer-based Detection Assay Against Bacillus anthracis Lethal Factor and Escherichia coli

Mieke Lahousse, Ph.D.

Mentor: Sung-Kun Kim, Ph.D.

The impressive capacity of bacteria to adapt to their environment has led to the development of mechanisms that can convey resistance to currently FDA approved antibiotics. In an attempt to find a solution to the possibility that soon there will be no antibiotic treatment available for bacterial infections, new approaches are being explored.

One such approach relies on the effective detection and/or diagnosis of pathogens or unique bacterial proteins to aid in prevention or better treatment of infectious diseases.

Within this work we describe the development of ssDNA aptamers by SELEX that can bind to their target with high affinity and can be used to develop colorimetric assays that can positively detect the presence of the target. Our targets for aptamer development were whole E. coli cells, which are currently used in water quality assessment, and a unique protein toxin (lethal factor) produced by B. anthracis, the etiological agent of anthrax. As a result of the SELEX experiments we recovered one aptamer that binds to

LF protein with high affinity and we demonstrated that its use in the development of an aptamer-based colorimetric assay for LF was successful. We also demonstrate that the aptamer found for LF protein can act as an inhibitor of the catalytic activity of LF, suggests its potential use as a therapeutic agent. The SELEX experiment using whole E. coli cells generated a diverse number of aptamers with binding affinities in the low µM range. Additionally, studies of aptamer specificity showed that three aptamers had no binding affinity for other coliform strains, making them suitable aptamers for further characterization. This work also describes the characterization of a glutaredoxin protein expressed by Synechocystis sp. involved in arsenate reduction. Elucidation of the mechanism by which cyanobacteria can survive in highly arsenic contaminated environments could enlighten possible remediation approaches. In conclusion, we successfully developed ssDNA aptamers that can bind with high affinity and specificity to its targets with demonstrated potential as detecting agents and contributed to the enrichment of the knowledge available for glutaredoxin GrxA involved in arsenate oxidoreduction by

Synechocystis sp.

LIST OF FIGURES ...... vii

LIST OF TABLES ...... xi

LIST OF ABBREVIATIONS ...... xii

CHAPTER ONE ...... 1

Introduction...... 1

CHAPTER TWO ...... 6

Detection and Inhibition of Anthrax Lethal Factor by ssDNA Aptamers ...... 6

Introduction...... 6

Materials and Methods ...... 13

Experimental Results ...... 18

Discussion ...... 32

CHAPTER THREE ...... 36

Development of Double Strand and Single Strand DNA Aptamers Against Escherichia coli Bacterial Cells ...... 36

Introduction...... 36

Material and Methods ...... 40

Experimental Results ...... 45

Discussion ...... 577

CHAPTER FOUR ...... 61

Site Directed Mutagenesis Study on GRXA a Protein from Cyanobacteria Synechocystis sp pcc6803 to Elucidate its Mechanism of Action ...... 61

Introduction...... 61

Materials and Methods ...... 67

Experimental Results ...... 72

Discussion ...... 77

CHAPTER FIVE ...... 90

Conclusions ...... 90

APPENDIX A ...... 92

APPENDIX B ...... 99

APPENDIX C ...... 101

REFERENCES ...... 109

LIST OF FIGURES

Figure 1. Diagram of the assembly and mode of action of the three protein involved in toxicity by Bacillus anthracis ...... 8

Figure 2. Schematic representation of a complete round of selection ...... 11

Figure 3. 12% SDS-PAGE stained with coomasie blue showing purified LF protein ...... 19

Figure 4. 6% native polyacrylamide gel electrophoresis (PAGE) stained with EtBr showing the product from asymetric PCR ...... 20

Figure 5. 6% PAGE stained with SYBR gold showing the separation of complexed ssDNA with LF from unbound ssDNA (EMSA) ...... 21

Figure 6. PCR confirmation of DNA aptamer insertion in pGem-T Easy plasmid ...... 22

Figure 7. Determination of binding affinity by EMSA using ML12 aptamer ...... 23

Figure 8. Determination of binding affinity of ML12 using a Cy3 attached ML12 aptamer ...... 24

Figure 9. Binding specificty determination of ML12 against BSA using Cy3 attached ML12 aptamer ...... 26

Figure 10. Development of a colorimetric assay for LF protein using a biotinylated ML12 aptamer ...... 26

Figure 11. Predicted secondary structure for the 30-mer ML12 aptamer using Mfold...... 28

Figure 12. CD spectra of 30-mer ML12 aptamer ...... 29

Figure 13. 12% SDS-PAGE stained with coomasie blue showing GST-MEK1 protein purification ...... 30

Figure 14. Diagram representation of the catalytic activity of LF on GST-MEK1 and the mass of the products generated after catalytic activity...... 30

Figure 15. Inhibition study of ML12 on the catalytic activity of LF protein activity by 12% SDS-polyacrilamide gel electrophoresis ...... 31

vii

Figure 16. Figure 16. IC50 determination of the inhibitory activity of ML12 aptamer over LF catalytic activity...... 32

Figure 17. Specificity test using Cy3 labeled dsDNA aptamers developed for E. coli DH5α ...... 47

Figure 18. Generation of an 80 oligonucleotide long ssDNA library for cell-SELEX experiments ...... 49

Figure 19. PCR product from Cell-SELEX round one ...... 50

Figure 20. PCR product form Counter-SELEX round two...... 51

Figure 21. PCR product from Cell-SELEX round three ...... 52

Figure 22. PCR product form Counter-SELEX round four ...... 53

Figure 23. PCR product from Cell-SELEX round five...... 54

Figure 24. Graphical representation of the aligment of 15 aptamer sequences ...... 56

Figure 25. Catalytic reaction cycle of Trx-coupled arsenate reductase from S. aureus ...... 63

Figure 26. Grx-coupled arsenate reductase mechanism of action ...... 64

Figure 27. Schematic representation of the Grx system involved in reduction of dithiol groups...... 64

Figure 28. Dithiol reduction mechanism by Grx protein ...... 66

Figure 29. Proposed mechanism of action for Synechocystis sp ...... 67

Figure 30. Graphical representation of the gene sequence alingment for the two new mutant GrxA proteins ...... 73

Figure 31. Study of the Oxidation/reduction potential of GrxA protein………………..75

Figure 32. Determination of the redox midpoint potential of GrxA ...... 78

Figure 33. Comparison of the redox stage between the three GrxA variants studied with or without binding interaction with 2 kDa malPEG polymer ... 79

Figure 34. Determination of the redox midpoint potential for GrxA using a fluorescent probe ...... 80

viii

Figure 35. Oxidized GrxA wild type deconvoluted mass spectrometry analysis ...... 81

Figure 36. Reduced GrxA wild type deconvoluted mass spectrometry analysis ...... 82

Figure 37. Oxidized GrxA double mutant C18S/C36A deconvoluted mass spectrometry analysis ...... 83

Figure 38. Reduces GrxA double mutant C18S/C36A deconvoluted mass spectrometry analysis ...... 84

Figure 39. Oxidized GrxA triple mutant C18S/C36A/C70S deconvoluted mass spectrometry analysis ...... 85

Figure 40. Mass spectrometry analysis for Grx triple mutant C18S/C36A/C70S ...... 86

Figure A1. Aptamer generation for LF protein. Asymmetric PCR product using 60 bp long oligonuclotide library ...... 94

Figure A2. Aptamer generation for LF protein. Asymmetric PCR product using the DNA recovered from round one of selection ...... 95

Figure A3. Aptamer generation for LF protein. Asymmetric PCR product using the DNA recovered from round two of selection ...... 96

Figure A4. Aptamer generation for LF protein. Asymmetric PCR using as template the product from the fourth round of selection ...... 97

Figure A5. Aptamer generation for LF protein. Asymmetric PCR performed with the DNA generated from the fifth round of selection ...... 98

Figure B1. PCR product from asymmetric PCR using the recovered ssDNA from round one of cell SELEX...... 99

Figure B2. PCR product from asymmetric PCR using the recovered ssDNA from round two of counter cell SELEX ...... 100

Figure B3. PCR product from asymmetric PCR using the ssDNA recovered from round three of cell SELEX ...... 101

Figure B4. PCR product from asymmetric PCR using the ssDNA recovered from round four of cell SELEX ...... 102

Figure C1. LCMS GrxA oxidized (-100 mV) ...... 103

Figure C2. LCMS GrxA reduced (-330 mV) ...... 104

Figure C3. LCMS GrxA double mutant oxidized (-100 mV) ...... 105

Figure C4. LCMS GrxA double mutant reduced (-330 mV) ...... 106

Figure C5. LCMS GrxA triple mutant oxidized (-100 mV) ...... 107

Figure C6. LCMS GrxA triple mutant reduced (-330 mV) ...... 108

LIST OF TABLES

Table 1. Buffer conditions for each round of SELEX ...... 20

Table 2. Sequence of aptamers against LF and their Kd values ...... 22

Table 3. Binding affinity of a dsDNA aptamer developed against E. coli DH5α cells ...... 46

Table 4. Dissociation constant determination of a dsDNA aptamer to different bacterial cells...... 47

Table 5. Sequence of the 40 oligonicleotide long random region for each ssDNA aptamers developed against E. coli DH5α cells...... 57

Table 6. Middle point potential for GrxA wild type protein and two cystine mutants ...... 76

LIST OF ABBREVIATIONS

AMP˗˗˗Adenosil mono phosphate

B. anthracis˗˗˗Bacillus anthracis

BSA˗˗˗Bovine serum albumin

Bio-ML12˗˗˗Biotinylated Mieke Lahousse 12 aptamer

BoNT˗˗˗Botulinum neurotoxin

CD spectra˗˗˗Circular dichroism spectra

CMG2˗˗˗Capillary morphogenesis protein 2

Cy3˗˗˗Cyanine 3

Cys˗˗˗Cysteine

DNA˗˗˗Deoxy ribonucleic acid dsDNA˗˗˗Double strand DNA dNTPs˗˗˗Deoxynucleotide triphosphates

DTT˗˗˗dithiothreitol

E. coli ˗˗˗Escherichia coli

EcoRI˗˗˗ Escherichia coli restriction enzyme I

EF˗˗˗Edema factor

ELISA˗˗˗Enzyme-linked immunosorbent assay

EMSA˗˗˗Electrophoretic mobility shift assay

EPA˗˗˗Environmental Protection Agency

ERK˗˗˗Extracellular signal-regulated kinase

EtBr˗˗˗Ethidium bromide

xii

6-FAM˗˗˗6-Carboxifluorecein

FC˗˗˗Fecal coliform

FDA˗˗˗Food and Drug Administration

G-quadruplex˗˗˗Guanosine-quadruplex

GrxA˗˗˗Glutaredoxin A

GST-MEK1˗˗˗Glutathion S-transferase-MEK

GSH˗˗˗Glutathione

GSSG˗˗˗Glutathione disulfide

HEPES˗˗˗4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

6-His-Tag˗˗˗6-histidine-tagged

H2SO4˗˗˗Sulfuric acid

IC50˗˗˗Half maximal inhibitory concentration

IDT˗˗˗Integrated DNA Technologies

IEF˗˗˗Isoelectric focusing

IPTG˗˗˗isopropiyl-β-D-thiogalactopyranoside

JNK˗˗˗c-Jun N-terminal kinase

KCl˗˗˗Potassium chloride

Kd˗˗˗Dissociation constant kDa˗˗˗kilo dalton

LF˗˗˗lethal factor

LRP6˗˗˗LDL-receptor-related protein mM˗˗˗milli Molar

MAPK˗˗˗Mitogen-activated protein kinase

xiii

MAPKK˗˗˗Mitogen-activated protein kinase kinases malPEG˗˗˗ Maleimide functionalized polyethylene glycol mBBr˗˗˗Monobromobimane

MEK˗˗˗Serine-threonine protein kinases

MDR˗˗˗Multi-drug resistance

MgCl2˗˗˗Magnesium chloride

ML12˗˗˗Mieke Lahousse 12 aptamer

MRSA˗˗˗Mehticillin resistant Staphylococcus aureus

NaCl˗˗˗Sodium chloride

Na3PO4˗˗˗Sodium phosphate

NADPH˗˗˗Nicotinamide adenine dinucleotide phosphate

ND˗˗˗Not determined

N-terminal˗˗˗Amino terminal

NZY Broth˗˗˗NZ amine, yeast extract based medium with no casamino acids

O.D.600˗˗˗Optical density 600 nm wavelength

PA˗˗˗protective antigen

PAGE˗˗˗Polyacrylamide gel electrophoresis

PCR˗˗˗Polymerase chain reaction

RNA˗˗˗Ribonucleic acid

Rpm˗˗˗revolutions per minute

RT-PCR˗˗˗Reverse transcription polymerase chain reaction

SELEX˗˗˗Systematic evolution of ligands by exponential enrichment

Ser˗˗˗Serine

xiv

SDS-PAGE˗˗˗Sodium-dodecyl-sulfate polyacrylamide gel electrophoresis ssDNA˗˗˗Single strand DNA

SynArsC˗˗˗Synechocystis-Arsenate reductase C

SWNT-FET˗˗˗ Shell–single Wall Nanotubes-Field Effect Transistor

TBST˗˗˗Tris-buffered saline Tween-20

TA˗˗˗Tris-acetate buffer

TC˗˗˗Total coliform

TCA˗˗˗Trichloroacetic acid

TEM8˗˗˗Tumor endothelium marker 8

TMB˗˗˗3,3’,5,5’-tetramethyl benzidine

Tris-HCl˗˗˗Trisaminomethane hydrochloric acid

Trx˗˗˗Thioredoxin

UNICEF˗˗˗United Nations Children's Fund

VEGF˗˗˗Vascular endothelial growth factor

Zn+2˗˗˗Zinc

CHAPTER ONE

Introduction

Historically human life expectancy was very low, between 25 to 30 years, compared to modern life expectancy of 80 years or more in some countries. The increase in life expectancy is in part the result of the evolution of human civilization, which introduced improved living conditions such as clean water sources, proper sewage disposal, access to better nutrition, and tremendous advances in medicine (McMichael T., 2001). Before the

1930’s the main causes of death were attributed to infectious diseases and hunger. With the discovery and clinical use of antimicrobial agents in the late 1930’s, humans appeared to have won the battle against infectious diseases, giving room for new non-pathogenic diseases that are nowadays becoming the leading cause of death in some countries. It has been predicted that by the year 2020 the most prevalent diseases will be heart diseases, obesity, and cancer in developed countries (McMichael T., 2001).

The introduction of antibiotic treatments and vaccination to combat bacterial infections seemed at the time to have put an end to the threat posed by pathogens to human health and life expectancy. The first known vaccine was used in 1798 and it consisted of the inoculation of smallpox from infected people in to the skin of healthy people, which proved to provide protection against subsequent exposures to smallpox. These results fueled the interest of doctors and scientists to study the mechanism of protection and to develop more vaccines that could protect against prevalent infectious diseases. By the use of antigens, that are portions or complete inactivated viruses or bacteria that elicit an immune response,

1 vaccines have been developed that can induce an adaptive immune response against the live pathogens and convey protection against recurrent encounters with the same pathogen. As a result of those efforts we now have vaccines for diseases like smallpox, diphtheria, tetanus, yellow fever, pertussis, haemophilus influenza type b, poliomyelitis, measles, mumps, rubella, hepatitis B, meningitis, pneumococcal infections amongst other vaccines (Bazin H.,

2003). But development of new vaccines has been slow and very difficult, even when promising antigens have been found; the translation process from research to development is very complex. For a vaccine to be approved by the FDA requires it to be well characterized with mandatory safety studies, reproducible results in small and large scale production, and provide long term protection (Buckland B., 2005); requirements difficult to satisfy, which was the case for a vaccine developed for B. anthracis, the causal agent of anthrax (Auerbach

S. et al. 1955; Ivins G. et al. 1998). The vaccine provides short term protection, with reported adverse reactions by immunized humans at high risk of exposure at military or industrial settings. The vaccine is only approved by the FDA to be used in people at very high risk of exposure (U.S. Department of Health and human services).

Antibiotics, on the other hand, have the capacity to inhibit bacterial reproduction once an infection has been established in the body. They act by inhibiting essential functions in bacteria like DNA and RNA synthesis, protein and cell wall synthesis, and cell membrane integrity (McMichael T., 2001). The use of these broad spectrum antibiotics has accelerated the generation of resistance mechanisms by bacteria, due to the increase in selective pressure posed by such antibiotics (Alanis A., 2005). This outcome was predicted to happen by the time antibiotics were first introduced into clinical settings, and indeed, short after their introduction, the first strains of resistant bacteria to penicillin appeared (McMichael T. 2001).

Presently, gram-negative multi-drug resistant bacteria (MDR) and meticillin resistant

Staphylococcus aureus (MRSA) are becoming a serious worldwide health problem.

Development of new and novel antibiotics has been very slow and new alternatives to antibiotic treatment are becoming more popular targets for development of novel treatments.

Other approaches have made use of anti-toxins or inhibitors of essential enzymes, in combination with antibiotics, to enhance potency of treatment and reduce development of resistance (Moradpour Z., 2011).

In the case of anthrax disease, antibiotic treatment is readily effective to treat bacteremia, but the potent toxins released by bacteria, if not neutralized, leads to death of the patient. The particular mode of action of B. anthracis has overemphasized the need for effective toxin inhibitors as well as diagnostic tools that will aid in fast diagnosis and effective treatment. Infection with anthrax in animals is diagnosed on site using polychrome methylene blue-stain from blood of carcasses, it is used to visualize the encapsulated bacilli and the test has a sensitivity of 5x104 cell/ml or more for positive diagnosis. Other available tests are very expensive and not easily performed in the field. In humans, diagnosis of cutaneous anthrax is confirmed by cultures from fluids of untreated lesions, while diagnosis of gastrointestinal or pulmonary anthrax, the deadliest forms of infection, are difficult to diagnose if there is no clear history of exposure. Positive identification is typically accomplished after the patient has died. Current clinically used diagnostic tools are: bacterial culture from bodily fluids of untreated patients, which can take over 4 days for positive confirmation; when possible, immunohistochemical stains of bronchial biopsies from treated patients are good confirmatory tests; PCR with commercial kits is becoming increasingly accepted as a diagnostic tool, especially if it is complemented with cultured samples. Other

3 more sophisticated methods that are culture free have been devised, like antigen based diagnosis, but are only used in specialized research laboratories at a very high cost (World

Health Organization, 2003)

Before new effective vaccines or therapies are developed, the best approach to combat emerging infections is by prevention. Strategies have been implemented in which frequent hand washing, use of protective respiratory devices, and minimal contact with infected persons has limited the spread of infectious diseases (Mao L., 2012). Initiatives like

“start smart then focus”, undertaken in Europe to prevent negligent use of antibiotics, have been successfully undertaken. Strict regulation on detection, diagnosis and antibiotic prescription are important parameters for the reversion of bacterial resistance (Hogberg L.

D., 2010). In order to allow for these initiatives to be successful, more probes for pathogen detection and/or diagnosis are required.

Future patterns of disease will be defined by the age of the population, environmental burden of population growth, and demand of fresh water and food. To ensure fresh water supply that is safe for human consumption, constant monitoring of drinking water sources has become mandatory. Current water testing assays are slow, lack specificity, and some are very expensive to perform, requiring specialized training and instrumentation. In addition to microbial monitoring, other tests are also required to measure the concentration of harmful contaminants, like heavy metals and pharmaceutical, which have detrimental effects on human health. New probes that can specifically identify bacterial indicators, that are fast and easy to perform, and most importantly that are cost effective are in demand to ensure good water quality.

The scope of this work is to develop new detecting probes that can facilitate environmental monitoring of microorganism like E. coli and B. anthracis, to aid in prevention of disease and/or proper diagnosis to ensure effective treatment. Within this work, we report the development of a new detecting assay and possible therapeutic agent based on DNA fragments specifically design to target bacterial toxins, as is the case of LF from B. anthracis, or whole cell bacteria like E. coli, that represent potential threats to human health. We also present compiling results on the characterization of cyanobacterial mechanisms evolved to overcome heavy water contamination with arsenic, information that contributes to our understanding of the mechanism and open the possibility to generate remediation approaches to providing safe drinking water.

CHAPTER TWO

Detection and Inhibition of Anthrax Lethal Factor by ssDNA Aptamers

Introduction

Anthrax is caused by Bacillus anthracis, a Gram-positive, non-motile, spore forming bacterium commonly found in soil (Mock M. et al. 2001). Infection with the bacterium can cause three different clinical manifestations depending on the site of infection: cutaneous, gastrointestinal, or pulmonary anthrax. Pulmonary infection represents the most serious manifestation of the disease, usually having very poor prognosis and a death rate of 100% after manifestation of respiratory distress (Mock et al. 2001).

The high toxicity of B. anthracis infection is due to the production and secretion of three proteins encoded in a virulent plasmid named pXO1, the proteins are: protective antigen (PA), Edema factor (EF), and lethal factor (LF) (Koehler T.M., 2009; Young et al.

2007). These three proteins act as an AB type toxin complex where PA plays a role as the non-catalytic portion (B portion) and either EF or LF is the catalytic portion (A portion) (Cao

Sha et al. 2007; Scobie et al. 2005). PA is an 83 kDa protein that binds to cell-surface receptors like tumor endothelium marker 8 (TEM8) or capillary morphogenesis protein 2

(CMG2), receptors that are expressed by several cell types including cells from the immune system (Bradley K. A. et al. 2001; Loving C. et al. 2009). Upon receptor recognition, PA83 is cleaved to form a free 20 kDa fragment and a receptor bound PA63 fragment that assembles into a heptameric or octameric ring-shaped complex (Kintzer et al. 2009). After receptor recognition and oligomerization, the pre-pore interacts with EF or LF and the entire complex is internalized forming a phagosome (Loving C. et al. 2009). Subsequently the phagosome

6 becomes acidified, and EF and LF are translocated into the cytoplasm throughout the pore formed by PA63 oligomer. Once in the cytoplasm, EF, a calcium/calmodulin dependent adenylate cyclase, increases the intracellular levels of cyclic AMP causing cell swelling; while LF, a zinc-dependent metalloprotease that cleaves most of the mitogen-activated protein kinase kinases (MAPKKs, also known as MEKs), interrupts the transcription of several genes including genes involved in immune response (Young et al. 2007). Figure 1 shows a summary of the internalization process for the AB type anthrax toxins and the targets for each toxin within the cell (Bouzianas D., 2010).

From the toxins generated by B. anthracis, LF has shown to be the most potent, causing on its own the death of rodents that were intravenously exposed to the toxin; the loss of the gene encoding for LF would produce substantially less pathogenic bacteria forms

(Moayeri et al. 2003; Duesbery et al. 1999; Pezard et al. 1991). Characterization of the protein revealed that LF is a 90 kDa protein, consisting of four domains: domain I that binds to PA for consequent cell internalization; and domain II, III, and IV that together form the binding site for the amino-terminal portion of MAPKK and the catalytic site (Pannifer et al.

2001). LF cleaves all mammalian MAPKKs, except for MAPKK5 (Tonello et al. 2009), and thereby shuts down the extracellular signal-regulated kinase (ERK), the c-Jun N-terminal kinase (JNK), and p38 MAPK pathways. LF cleaves near the amino-terminal site from the

MAPKK which is involved in binding to MAPKs therefore reducing their binding affinity and phosphorylation of MAPKs (TURK B. 2007; Xu et al. 2008).

One well studied effect of LF catalytic activity is the suppression of pro-inflammatory gene expression in macrophages and induction of apoptosis in dendritic cells and

7 macrophages; furthermore, it also inhibits T-cell and B-cell activation as well as blocking neutrophil chemotaxis and superoxide production (TURK B. 2007; Fang H. et al. 2010).

Figure 1. Diagram of the assembly and mode of action of the three proteins involved in toxicity by Bacillus anthracis. The diagram represents the assembly of PA into a channel and internalization of LF and ED into the cell and their respective mode of action. The monomeric form of the PA protein binds to its receptor on the cell surface where it is cleaved to generate two fragments, the largest one remains bound to the receptor and the smaller fragment is released. The receptor bound fragment then forms a heptamer or octamer complex that can interact with LF and EF proteins for internalization. Once internalized the vesicle becomes acidified causing a change in conformation of the PA complex that allows the release of the two proteins, LF and EF, in the cytoplasm were they exert their catalytic activity (figure modified from Bouzianas D., 2010).

Baldary reports in his review that EF and LF have shown inhibitory activity on oxidative burst and phagocytosis by the immune response. Taken all together, the response of most cellular components of the innate immunity is neutralized and the adaptive response

8 is halted (Baldary et al. 2006). These reports provide strong evidence that LF plays a pivotal role in the biological action of anthrax.

Given the destructive potential of the toxins produced by B. anthracis, approaches to promptly and effectively diagnose anthrax disease have proven to be particularly challenging because the initial symptoms of the infection are very similar to flu symptoms, and characterization of the bacteria is rather complicated due to its similarity to other members of the Bacillus group (Edwards et al. 2006). Hence, prompt diagnosis is required to treat the infection effectively with antibiotic treatments; however, antibiotics are not able to neutralize the toxins secreted from the bacteria into the body. Efforts to develop antidotes for LF, the main virulence factor, have generated a monoclonal antibody that neutralize LF toxin and can be used to prevent or treat toxemia (Lim NK. Et al. 2005); peptide hydroxamate inhibitors that mimic the substrate binding site and protease activity inhibitors like sulfonamide-based inhibitor (Montecucco C. et al. 2004). The last type of inhibitor is a zinc binding molecule that interacts with the active site of LF. The disadvantage of zinc binding inhibitors and their use in humans is that they can cause adverse side effects during treatment like musculoskeletal syndrome, which causes joint stiffness, pain, inflammation and tendinitis, therefore limiting the use of such therapies (Xiong et al. 2006; Jiao G. 2010; Montecucco et al, 2004; Jacobsen J. 2010). Additionally, major problems of cell permeability, stability, and specificity of other potential inhibitors need to be resolved before finding good antidotes to anthrax toxins. Since LF is a secreted protein that acts as a toxin, it offers an attractive venue for a detection and therapeutic target. Additionally, it has been reported that although the PA concentration in rabbit sera was highly variable, the concentration of LF at 48 h after infection was generally higher than those of PA and EF (particularly the concentration ratio

9 of LF and EF was 5:1 and the concentrations measured were in the low µg/ml range) (Molin et al. 2008). This fact also supports the notion that LF could be an attractive target for detecting the presence of B. anthracis.

In order to detect the toxins, SELEX technology was explored. SELEX is a combinatorial chemistry procedure developed to select synthetic oligonucleotides capable of binding with high affinity to a target molecule. The oligonucleotides can either be single stranded DNA (ssDNA), RNA, or double stranded DNA (dsDNA). Single stranded oligonucleotides are preferentially used to generate aptamers because they can fold into complex secondary and tertiary structures like stems, loops, bulges, hairpins, tetra loops, pseudoknots, etc, and are more stable than RNA (Kim et al. 2008; review). The particular structure of ssDNA aptamers facilitates their interaction with the target through hydrogen bonding, van der Waals forces, and electrostatic interactions. The procedure was first reported by three independent research laboratories (Tuerk et al. 1990; Joyce G. F. 1989;

Ellington et al. 1990). The SELEX technique is an iterative procedure that consists of three steps: 1) Incubation of the oligonucleotide library with the target molecule; 2) Separation of the complex (target:oligonucleotide) from the unbound oligonucleotides; and 3)

Amplification of the bound sequences by PCR. Affinity of the aptamers developed will depend on the number of rounds performed, salt concentration in buffers, and the nature of the sample.

SELEX was developed more than 15 years ago and since its development, SELEX has been applied to several research fields generating aptamers for virtually any target which include: proteins, peptides, nucleic acids, polysaccharides, small organic molecules, virus particles, whole cells, and tissues (Kulbachinskiy et al. 2007; Shamah et al. 2008).

Figure 2. Schematic representation of a complete round of selection. A ssDNA library with a diverse composition of secondary structures adopted by ssDNA (red lines: linear DNA; blue and black: secondary structures of differing complexity) can be combined with the target protein during binding and the unbound ssDNA is then removed during the separation step. Bound ssDNA is then amplified by PCR and the product is used in subsequent rounds of selection. The final round of selection is amplified and sequenced to characterize the composition of the aptamers generated by SELEX.

In the diagnostic field, aptamers have been developed that can identify proteins like

MPT64 antibody for detection of Mycobacterium tuberculosis infection (Qin et al. 2009).

Aptamers that bind to B. anthracis spores have also been developed and proposed to be used as potential detecting agents (Bruno et al. 1999) and more recently aptamers that bind to anthrax PA have also been proposed as a detecting and diagnostic tool (Choi, Ji Sun 2011).

For research purposes, aptamers have been developed to study intracellular processes involved in cell signaling and protein localization (Mallikaratchy et al. 2006; Strehlitz et al.

2008). Whole cell SELEX has been extensively used in cancer research to develop tracers against specific cancer cell markers and to collect and detect multiple cancer cell lines by the combination of aptamers with emergent technologies like nanoparticles (Chen et al. 2009;

Smith et al. 2007).

Because of their successful performance, aptamers have been used to develop new drugs that can act as inhibitors. For example, aptamers developed against botulinum neurotoxin (BoNT), a toxin produced by Clostridium botulinum, prevents the docking of the toxin to gangliosides and the synaptic vesicle, therefore inhibiting their activity (Jeffrey et al.

2007). More relevant is the commercialization of a product developed using SELEX technology named Macugen®, which is produced and marketed by OSI

Pharmaceuticals/Pfizer Inc. (Chapman et al. 2006). The product uses an aptamer developed against a vascular endothelial growth factor (VEGF) and is currently used to treat age-related macular degeneration disease (Ng E. W., 2006).

Through the process of SELEX, tightly binding dsDNA, ssDNA, or RNAs can be found. SELEX technique has been employed to discover aptamers against B. anthracis spores (Bruno, J. G, 1999) and PA (Choi et al. 2011), and yet there are no reports of aptamers against LF. Here, we have developed ssDNA aptamers against LF using SELEX and determined their binding affinity and specificity. We also designed an aptamer-based ELISA assay to visually detect LF. In addition, binding sites of the aptamer and LF have been studied with the combination of experimental and computational analyses. Furthermore, the possibility of LF inhibition with the found aptamer was assessed.

Materials and Methods

Protein Expression and Purification

To obtain purified six-histidine tagged LF protein from transformed E. coli BL21

(DE3) strain, harboring the pET28a-LF plasmid, the transformed bacteria was grown in

Luria-Bertani (LB) medium containing 50 µg/ml of kanamycin and incubated at 37 °C and

200 rpm in a shaking incubator until the optical density of the culture measured at 600 nm reached 0.6. Once the desired O.D.600 was reached, expression of the recombinant protein was induced by the addition of isopropyl-β-D-thioglactopyranoside (IPTG) at a final concentration of 1 mM and continued to incubate the cells at 30°C for an additional 6 hours.

Cells were harvested and stored at -80°C. Protein purification was performed as described previously (Park S. et al. 2000) but with slight modifications. Briefly, cells were resuspended in resuspension buffer (30mM Tris-HCl pH=8.0, 500mM NaCl,) and lysed using a French press under 18,000 psi three times with intervals of 5 min. Cell lysate was centrifuged to remove cell debris by the use of a Beckman Coulter Avanti J-26 XP at 20,000 rpm (38762.3 g force) using a JA 25-50 rotor. Supernatant was filtered using a 0.45 µm filter

(EMD Millipore Corporation, Billerica, MA) prior to loading on to a Ni2+-affinity chromatography column equilibrated with resuspention buffer (GE Healthcare). After loading, the column was washed with 5 column volumes of a 3% mixture of elution buffer

(30mM Tris-HCl pH=8.0, 500mM NaCl, and 500mM imidazole) and resuspension buffer.

Elution of the protein was achieved by a gradient of elution buffer up to 50%. Purity of the protein extract was assessed by 12% sodium-dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) stained with Coomassie brilliant blue (Thermo Fisher

Scientific, Inc, Pittsburgh, PA). An additional step of purification was performed using size

13 exclusion chromatography on a GE AKTA purifier (high load 26/60 superdex 200 column) using a 30mM Tris buffer pH=8.0. The column was washed for 1 hour and 2 ml of protein sample were loaded onto the column. Protein retention time in the column was 3 hours.

Purity was confirmed by SDS-PAGE.

For the purification of Glutathione S-transferase (GST)-tagged mitogen-activated extracellular signal-regulated kinase kinase 1 (GST-MEK1), the gene was cloned and expressed using a pGEX-2T vector cloned into E. coli BL21 (DE3) cells. Cultured cells were grown for 6 hours at 30°C after IPTG induction and the pellet was saved at -80°C.

Cells were disrupted by French press and the supernatant from the centrifugation step was loaded into a GSH-affinity chromatography column. Protein was recovered from the column by using a gradient from 0-50% mixture of buffer B into buffer A (Buffer A: 30 mM Tris-

HCl Ph=8.0; 500 mM NaCl and Buffer B: 30 mM Tris-HCl pH=8.0; 500 mM NaCl; 5 mM reduced glutathione). After protein purification, purity of the recovered GST-MEK1 was evaluated by SDS-PAGE. An additional purification step was required for GST-MEK1 after affinity chromatography purification. The method chosen was size exclusion chromatography using a 30 mM Tris-HCL pH=8 buffer and purity of the recovered protein was confirmed by SDS-PAGE.

Generation of ssDNA Library and SELEX Process

To begin SELEX, a random 30 oligonucleotide long DNA fragment flanked by a forward primer region and a reverse primer region, 5'-GCGCGGATCCCGCGC-N(30)-

CGCGCGAAGCTTGCG-3’, was purchased from Integrated DNA Technologies (IDT) (IDT,

Coralville, IA). Additionally, two primers were purchased with the following sequence: forward primer: 5'-GCGCGGATCCCGCGC-3' and reverse primer: 5'-

CGCAAGCTTCGCGCG-3'. The template was amplified by asymmetric PCR using a thermocyler with a 10:1 forward:reverse primer ratio, and the ssDNA was then recovered from a 6% polyacrylamide native gel using a previously described crush-and-soak method and concentrated by ethanol precipitation (Kim et al. 2009).

To obtain aptamers against LF, purified protein resuspended in a binding buffer (BB)

(50 mM Tris-HCl pH=7.4, 5.0 mM KCl, 100 mM NaCl, 1.0 mM MgCl2) was incubated at

30°C with the randomized ssDNA library for 30 min in a final volume of 50 µL. The complex between protein and ssDNA was resolved by an electrophoretic mobility shift assay

(EMSA) using a 6% polyacrylamide native gel and visualized with either ethidium bromide or SYBR gold stain. Recovery of the complexed ssDNA from the gel was achieved by a crush-and-soak method followed by ethanol precipitation and regular PCR amplification.

The double stranded DNA (dsDNA) was then amplified by asymmetric PCR to produce the ssDNA for the next SELEX round. This process was repeated up to round 5, at which point the ssDNA was used to perform a counter SELEX step using bovine serum albumin (BSA,

Sera Care Life Sciences, Birch St Milford, MA). After one round of counter SELEX, the selection process continued for 3 more rounds with an increased NaCl and KCl concentration. The product of the final round of SELEX was amplified by PCR and the resulting dsDNA was purified by running a 6% polyacrylamide gel electrophoresis. The dsDNA band on the gel was recovered by crush-and-soak and then inserted into pGem-T

Easy vector according to the manufacturer’s instructions (Promega Corporation, Madison,

WI). E. coli BL21 (DE3) cells were transformed and grown on an agar plate containing 50

M kanamycin. The presence of the insert was confirmed by plasmid restriction enzyme digestion using EcoRI (BioLabs, Inc. Lawrenceville, GA). Plasmids from positive colonies

15 were sent to Functional Biosciences for sequencing using T7 forward and reverse primers

(Functional Biosciences, Inc. Madison, WI).

Determination of Dissociation Constant (Kd)

To estimate the Kd for the found aptamers to LF, we purchased the aptamers with a fluorescent label (cyanine, Cy3) at the 3’-end (IDT, Coralville, IA). A fixed amount of LF

(0.5 g) in 100 L of 20 mM HEPES (pH= 7.4) at room temperature was loaded onto a 96- well white opaque immunoplate (SPL, Kyounggi-do, South Korea) and was incubated for 1 h with shaking to immobilize LF on the plate. The wells containing LF were washed with 200

L of Tris-buffered saline Tween-20, called TBST (0.5 M Tris-HCL pH=8.4, 9% NaCl,

0.5% Tween-20) three times. Subsequently, 100 L of 2% BSA was added and incubated for 1 h. An equal volume of TBST was again used to wash the wells three times. Varied concentrations of ssDNA aptamers (0 – 50 nM) were added to the wells and incubated for 1 h with shaking. Excess ssDNA was eliminated by washing with 200 L of 20 mM HEPES

(pH=7.4) three times. The fluorescent intensities of the complex between ssDNA and LF were measured by Gemini EM fluorescence microplate reader (excitation: 545 nm; emission:

572 nm) (Molecular Devices, Sunnyvale, CA).

An additional method was used to observe the binding of the aptamer ML12 to the LF protein. A constant amount of 5.9 µM protein was incubated with increasing concentrations of ML12 aptamer (0-10 nM) in 1X BB. Samples were incubated for 20 min at 30°C and then electrophoresed in a 6% PAGE. The ssDNA was stained using SYBR gold according to manufacturer’s instructions and pictures were taken using an excitation wavelength of 495 nm and emission wavelength at 537 nm. Fluorescent intensities were calculated using the computational program ImageJ (freely available at http://rsbweb.nih.gov/ij/).

Aptamer-based ELISA method

LF protein (0.5 g/well) was immobilized onto a 96 polystyrene white opaque immnunoplate (SPL, Kyounggi-do, South Korea) as described in the preceding section.

Varied concentrations of the aptamer ML12 labeled with biotin at the 3’-end (purchased from

IDT) were added to the wells possessing immobilized LF and then incubated for 1 h with shaking. Excess ssDNA was removed by washing with 200 L TBST three times.

Subsequently, 100 L of streptavidin-attached horseradish peroxidase (purchased from BD

Biosciences, Franklin Lakes, NJ) was added to the wells with 1:1000 dilution with TBST.

Incubation was carried out for 1 h with shaking. Subsequent to washing the wells with 200

L of TBST three times, blue color was developed by adding 90 L of TMB substrate solution (3,3’,5,5’-tetramethyl benzidine; R&D Systems, Minneapolis, MN). After waiting

15 min the reaction was terminated by the addition of 90 L of 1M H2SO4 to yield a yellow color. The color intensities in each well were measured with the Biotrak multiwall plate reader (Amersham Biosciences, Piscataway, NJ) at 450 nm.

CD Spectra Analysis

To determine possible secondary structure characteristics, aptamers (5 M) were resupended in 20 mM Na3PO4 (pH 7.5) that contained 100 mM KCl or 100 nM NaCl.

Samples were heated at 90oC for 5 min, followed by gradual cooling to room temperature.

CD spectra were collected by a JASCO J-810 spectropolarimeter (JASCO, Inc., Easton, MD) from 320 – 220 nm, with a 1 cm path length cuvette.

In vitro Proteolytic Activity Assay of Lethal Factor Protein

GST-MEK1 was expressed in E. coli DH5α and purified as described previously

(Kim et al. 2003). LF (0.25 µg) was pre-incubated with or without ML12 aptamer at 37oC for 15 min. The reaction was initiated by adding substrate mixture (25 mM HEPES pH=7.4,

1 mM DTT, 10% glycerol, 50 µM CaCl2, 5 mM MgCl2, 0.7 g GST-MEK1), terminated by the addition of 5x SDS-polyacrylamide loading buffer after 1 h, and the reaction was completed by boiling. Samples were assessed on a 12% SDS-polyacrylamide gel and the cleavage of GST-MEK1 was visualized by coomassie blue staining (Kim et al. 2003). To quantify the intensity of the stained bands, ImageJ was used (freely available at http://rsbweb.nih.gov/ij/).

Experimental Results

Protein Purification and SELEX Process

Purification of LF protein, required for SELEX experiments, was achieved by a two- step purification using two different chromatographic techniques. Figure 3 shows the results of an SDS-PAGE with the protein product from the two purification steps. The middle column on the SDS-PAGE shows that two proteins were recovered from affinity chromatography; one with a molecular weight of 94 kDa that represents LF protein with the

6-His-tag tail, and the second one with a lower molecular weight of 40 kDa representing an unknown protein. The recovered protein fraction from affinity chromatography was further purified by gel filtration chromatography to separate the two different proteins. The right column shows only one band at 94 kDa that represents a pure fraction of LF protein (Figure

3).

Once we obtained a pure fraction of LF protein, we proceeded to perform the SELEX experiment. For the selection experiments we used a ssDNA library generated by asymmetric PCR. Figure 4 shows the result from the amplification of the library by asymmetric PCR. The ssDNA library was recovered from the gel by a crush and soak method as described in the material and methods section. Table 1 summarizes the buffer conditions in which the selection process was performed. Note that for the final two selection rounds, the salt concentration was increased to increase the stringency of the selection. Addition of counter-SELEX rounds within the SELEX procedure is commonly used to improve the specificity of the final product; Table 1 shows the conditions under which an intermediate round of counter-SELEX was performed.

Figure 3. 12% SDS-PAGE stained with coomassie brilliant blue showing purified LF protein. Line 1 contains a molecular weight ladder, Line 2 contains LF protein purified by affinity chromatography, and Line 3 contains LF protein purified by gel filtration chromatography.

Table 1. Buffer Conditions of Each Round of SELEX.

Round BSA counter Round Round 1st – 5th SELEX 6th 7th – 8th 30 mM Tris 30 mM Tris 30 mM Tris 30 mM Tris (pH 8.0) (pH 8.0) (pH 8.0) (pH 8.0) 60 mM NaCl 60 mM NaCl 60 mM NaCl 100 mM NaCl 3.0 mM KCl 3.0 mM KCl 3.0 mM KCl 5.0 mM KCl 0.6 mM MgCl2 0.6 mM MgCl2 0.6 mM MgCl2 0.6 mM MgCl2

Figure 4. 6% native polyacrylamide gel electrophoresis (PAGE) stained with EtBr showing the product from asymetric PCR. Line 1 shows the DNA ladder with increments of 20 bp, line 2 shows a sample of the template DNA used to generate the ssDNA library, and line 3 shows a sample of the asymetric PCR product with two bands of DNA: top one for dsDNA and botom one for ssDNA.

To separate the ssDNA:LF complex from the unbound ssDNA at the end of each round of selection, we performed an EMSA assay. Figure 5 shows the results of the sixth round of selection, where is clearly observed the separation between the ssDNA bound to LF and the unbound ssDNA. The band on EMSA containing the complex was recovered and

20 amplified to generate the new ssDNA used for a subsequent SELEX round. Amplification of the recovered DNA for further SELEX rounds presented some challenges as observed in the results provided in appendix A, Figures A1-A5, where the product of amplification generated more than two bands and as the rounds of selection progressed, it became more difficult to generate a clear ssDNA band. For this reason SELEX was terminated after the eight round.

ssDNA:LF

ssDNA

Figure 5. 6% PAGE stained with SYBR gold showing the separation of complexed ssDNA with LF from unbound ssDNA (EMSA). EMSA was loaded with the product from the sixth round of selection. The top band on the EMSA represents the complex of LF protein and bound ssDNA, the bottom band represents the unbound ssDNA.

After completing the final round of SELEX, the ssDNA product was recovered and amplified by PCR to generate the dsDNA required for cloning into a vector for sequencing.

Before sending the plasmids for sequencing we confirmed the presence of a 60 oligonucleotide long insert by restriction enzyme treatment. Figure 6 shows the results of a cloning experiment where the plasmid of several colonies were recovered and treated with restriction enzymes, the product from the reaction was then run on a 6% polyacrylamide gel to confirm the presence of the dsDNA insert. A total of 12 plasmids containing an insert

21 were sent for sequencing and only four out of the twelve plasmids had an insert that matched the 60 oligonucleotide sequence of the starting library. Table 2 shows the three different sequences obtained as a product from SELEX and the frequency they were found within the sequenced plasmids.

Figure 6. PCR confirmation of DNA aptamer insertion in pGEM-T Easy plasmid. Picture of a 6% PAGE stained with EtBr showing the product of restriction enzyme treated plasmid to confirm the presence of a 60 oligonucleotie long DNA insert generated by SELEX. Lines 1, 2, 4, and 5 show the product from restriction enzyme treatment; line 3 shows a DNA ladder with increments of 20 bp. Only the plasmids showing a DNA insert of 60 nucleotides were send for sequencing.

Table 2. Sequence of Aptamers Against LF and Their Kd Values

Aptamer ssDNA Sequence (30-bp) Frequency Kd value ML6 GGACCAGCCGCCGCGCCTTGACCGGGGGTA 1 ND* ML7 GGAGAGAGGGAGACGCGCAACCTCGACCCT 1 ND* ML12 CGAGGGAGACGCGAACCTTCTCGCCTTGGG 2 11.0nM *Not determined within the µM range. The underlined sequence indicates the portion of the oligonucleotide that may contribute to the formation of a hairpin secondary structure.

Binding Affinity and Specificity

To determine the binding affinity of the found aptamers, we initially used an EMSA assay to visually explore the binding affinity of ML12 to the target LF protein. Figure 7A shows the result of an EMSA experiment performed with increasing concentrations of the aptamer from where the change in intensity of the fluorescent signal obtained from the interaction of SYBR gold with the increasing concentrations of ssDNA allowed us to calculate the Kd value using a Langmuir isotherm fit. From the plot shown in Figure 7B the

Kd value was estimated to be 524 pM.

Figure 7. Determination of binding affinity by EMSA assay using ML12 aptamer. A: picture of a 6% PAGE stained with SYBR gold showing the interaction of LF and ML12; binding reaction was carried out using a constant concentration of LF [5.9 µM] with increasing concentrations of ML12 aptamer. B: determination of the Kd value by Langmuir isotherm fit of fluorescent intensity vs ML12 concentration.

Due to difficulties in the reproducibility of the data generated by EMSA we decided to use a fluorometric method in which the fluorophore Cy3 was attached to the 3’-end of each aptamer. Binding affinity tests for each aptamer showed that only ML12 aptamer had a binding affinity with a Kd value of 11.0 ± 2.7 nM while the other two aptamers shoed binding affinities out of the mM range and therefore were not tested any further. Figure 8 shows the results of the Kd determination for ML12 with the florescent Cy3 molecule and the data points fit into a single ligand binding hyperbolic curve.

1.0

0.8

0.6

0.4

0.2 Normalized Fluorescence Intensity Fluorescence Normalized 0.0 0 20 40 60 80 100 ML12 (nM)

Figure 8. Determination of binding affinity of Cy3-ML12 aptamer. Binding reactions were carried out with a constant concentration of LF protein in a 96-well black plate and varied concentrations of Cy3-attached ML12 (0 – 50 nM). The fluorescent intensity of the bound ML12 to LF was measured after a couple of washing steps. The Kd value was determined by nonlinear fit of the saturation binding curve.

We also tested Cy3-ML12 including the flanking sequences (i.e., a 60-mer) and observed no binding up to a concentration of 1 µM. To ensure that the binding observed by

Cy3-ML12 was not a product of experimental error, a scramble sequence with the same size of ML12 was used in the assay as a control (i.e., ssDNA 30-mer; 5’CGCACTGAAGTG

GATACCGCCTAAACGGA-3’). No binding was observed with the scramble sequence up to a concentration of 1 µM.

To test the specificity of the ML12 aptamer for its target LF the % binding of Cy3-

ML12 to LF and BSA was compared using a constant aptamer and protein concentration.

BSA was used to test specificity because is a protein found in high abundance in serum samples and is commonly used during assay development to block nonspecific binging sites, therefore it was fundamental to make sure that the ML12 aptamer had low to no binding affinity for BSA. Figure 9 shows the % binding affinities of ML12 for LF and BSA. It can be observed that there is a significant difference between the % binding of the ML12 aptamer to its target LF protein and the % banding of the ML12 aptamer to BSA indicating that ML12 aptamer has a higher binding affinity for LF when compared to BSA.

Aptamer-Based ELISA Development

In order to use ML12 aptamer as a detection tool, an aptamer-based ELISA using a

96-well clear plate was developed. Control experiments testing non-specific binding of the biotinylated ssDNA aptamer (Bio-ML12) to the plate showed no binding of this molecule to the plate, as was previously reported by other researchers (Choi et al. 2011). When binding of the bio-ML12 aptamer to LF protein was tested, we were able to observe an increased intensity in the color generated by the reaction of the HRP-conjugated streptavidin with TMB and H2SO4 that was dependent on the concentration of bio-ML12 used. This approach for an

ELISA-based detecting method is shown in Figure 10, from where a clear increase in the color intensity was generated as the aptamer concentration increased, with saturated

25 intensities at the highest concentrations of aptamer used. The quantification of color intensity revealed a Kd value of 530 ± 10.8 nM. Additionally, we tested the scramble sequence oligonucleotide used for the preceding fluorescent measurement method and found that no binding to LF was observed in this ELISA method.

Figure 9. Binding specificty of ML12 against BSA. Fluorescent intensity measurement of bound Cy3-ML12 to either LF or BSA. The concentration of the proteins (LF and BSA) kept constant at 0.5 M, and 10 nM of ML12 was used for initial incubation against the proteins.

Figure 10. Use of biotinylated ML12 aptamer to develop a colorimetric assay for LF protein. The first row contains all the components of the reaction without LF protein and the second row contains a fixed concentration of LF (0.5 g/well). Each row was treated with various concentrations of biotinylated ML12 (0-2.0 µM) and the colorimetric reaction was performed as described in the method section.

Characterization of the Aptamer ML12

To explore potential binding sites of the aptamer, MFold was used to predict possible secondary structures of the aptamers (Zuker M, 1989). Although our focus of the binding sites was on the complex between ML12 and LF, we examined all three aptamers to ensure that there was no structural relationship among them. As a result, MFold predicted all dissimilar secondary structures, inferring that no common binding sites would be present among the three aptamers. In the case of ML12, two different secondary structures were predicted by MFold with weak ΔG binding values of -2.66 and -1.70 kcal/mol, respectively.

The secondary structure with the lowest energy value is shown in Figure 11. In this secondary structure prediction, a 14-mer sequence forms a stem-loop that may be a good candidate for the binding site of ML12. When the isolated 14-mer (5’-

GCGAACCTTCTCGC-3’) sequence was subject to the DNA folding prediction, an identical secondary structure was formed with a ΔG value of -2.01 kcal/mol, which is slightly lower than the value obtained for the full sequence. In an effort to obtain the experimental data of interaction between the 14-mer and LF, we attempted to determine Kd value between them.

However, there was no significant fluorescent signals up to 1 µM by the 14-mer, suggesting that instead of the 14-mer, the 30-mer would be a better choice for detection. It should be noted here that we also predicted the secondary structure of the ML12 with the flanking sequences (i.e., 60-mer containing ML12 30-mer sequence and two flanking sequences) and found that the predicted secondary structures are different from those of the 30-mer ML12.

Taken together, these observations suggest that in this research, the prediction by MFold may not be essential for binding site evaluation.

Figure 11. Predicted secondary structure for the 30-mer ML12 aptamer using the Mfold program.

In view of the aptamer composition, differences in guanosine (G) content between the full sequence (30-mer) and the 14-mer can be observed. In the case of the 30-mer, the G content is 36.7%, whereas the 14-mer contains only 21.4%. One might argue that due to the high G content in the 30-mer, the 30-mer may form a G-quadruplex structure instead of a stem-and-loop structure. To assess the possibility of the presence of a G-quadruplex, we used circular dichroism (CD). Figure 12 shows the CD spectra of ML12 generated under three different salt conditions: 100 mM NaCl, 100 mM KCl, and a control using only the phosphate buffer. NaCl and KCl were assed because earlier literature reported that the presence of potassium or sodium ions in solution and appropriate heat-and-cooling procedures would facilitate the formation of correctly folded G-quadruplex structures

(Michalowski et al. 2008). We studied the CD spectra of the 30-mer ML12 in the presence of K+ or Na+ and found that no G-quadruplex spectra were observed. In fact, the CD spectra obtained with the 30-mer ML12 in the presence or absence of K+ or Na+ showed almost identical spectra with a positive maximum peak near 280 nm and a negative peak near 240 nm. These results indicate that a secondary structure with B-form double helix is formed, rather than a G-quadruplex (Shum et al. 2011). Combined, these observations suggest that there is only one possible secondary structure (14-mer) in the 30-mer ML12 and that the entire 30-mer dictates tight binding to LF.

Evaluation of an Aptamer-Based LF Inhibitor

Since the aptamer ML12 tightly binds to LF, there is a possibility that ML12 would interfere with the interaction between LF and its substrate MEK1 and act as an inhibitor of the reaction. To test this hypothesis a GST-MEK1 protein was expressed and purified

(Figure 13) in order to perform an enzymatic assay to test LF catalytic activity (Kim et al.

2003). The GST modification on the amino terminal portion of MEK1 increases the molecular mass of the cleaved portion enabling the successful observation of the catalytic activity of LF enzyme on GST-MEK1 by the generation of two new fragments of know molecular weight. The two new fragments can be identified on an SDS-PAGE, and the original GST-MEK1 band disappears due to the proteolytic activity of LF. Figure 14 shows the molecular mass of each protein and the mass of the two fragments generated after LF cleavage of the GST-MEK1 construct.

Figure 12. CD spectra of 30-mer ML12 aptamer. The red line represents the CD spectrum of ML12 aptamer measured in only 20 mM Na3PO4 (pH=7.5); the green line represents the CD spectrum of ML12 in 20 mM Na3PO4 (pH=7.5) and 100 mM NaCl; the blue line represents the CD spectrum of ML12 in 20 mM Na3PO4 (pH=7.5) and 100 mM KCl.

Figure 13. 12% SDS-PAGE stained with coomassie brilliant blue showing GST-MEK1 protein purification. Line 1 contains a protein ladder and line 2 column contains the product from two step protein purification for GST-MEK1 with an estimated molecular size of 71 kDa (First step: affinity chromatography and second step gel filtration chromatografy).

Figure 14. Diagram representation of the catalytic activity of LF on GST-MEK1 and the mass of the products generated after catalytic activity.

To ensure that the assay method is valid, two controls were carried out. The first control experiment was treated with no pre-incubation time for LF reaction to GST-MEK1, and the second control experiment was performed to observe complete conversion of the

GST-MEK protein by LF into the two smaller size proteins observed in the diagram. As shown in Figure 15 for the first control experiment (Lane 1), the GST-MEK1 is intact, and for the second control experiment (Lane 2), the GST-MEK1 band disappears as a result of LF enzyme activity. To assess the possibility of inhibition of LF by ML12, varied concentrations of ML12 ranging from 0 to 60 µM were used in the same gel as was used for the control experiments. From Figure 15 it can be observed that as the concentration of

ML12 increased, there was a decrease in the activity of LF that resulted in a more intense band for GST-MEK1 protein. By measuring the intensity of the GST-MEK1 band from the inhibition test we were able to construct a dose/response curve and estimate the inhibitor concentration required to obtain a 50% inhibition of the enzyme activity. The data shown in

Figure 16 shows a good fit to a single exponential decay curve with a calculated IC50 of 15

µM ± 1.5.

Figure 15. Inhibition study of ML12 on the catalytic activity of LF protein activity by 12% SDS-polyacrilamide gel electrophoresis. SDS-PAGE electrophoresis showing the catalytic Reaction of LF over GST-MEK1 enzyme with increasing concentrations of ML12 aptamer from 0-60µM.

Figure 16. IC50 determination of the inhibitory activity of ML12 aptamer over LF catalytic activity. Plot of the relative LF activity at increasing ML12 aptamer concentration.

Discussion

To date, most efforts to find an inhibitor of LF produced by B. anthracis focused on the use of hydroxamate functional group containing-compounds that effectively inhibit the

Zn+2 methaloproteinase activity of LF, or peptides (Kocer et al. 2005; Tonello et al. 2002;

Turk et al. 2004). The disadvantage of such an approach is that hydroxamate has a wide spectrum of targets, has the ability to reversibly bind a number of metal ions including zinc ions, and cause serious side effect such as musculoskeletal syndrome (Peterson J. T. 2006).

Thus, new type of inhibitors like aptamers can be developed against specific protein targets.

A wide collection of evidence has been reported and some of the most relevant aptamers created and used in clinical applications are for example: Pegaptanib which is an RNA aptamer approved by the FDA to treat macular degeneration; other aptamers are in phase one and two of clinical trials for the treatment of type two diabetes, heart disease, leukemia, and

32 age-related macular degeneration (Ni et al. 2011). Consequently, exploration of the possibility of developing ssDNA aptamers that could bind to LF and possibly inhibit its activity is a promising approach to develop new drugs for this protein. The SELEX experiment performed using LF protein as target generated three aptamers with differing sequences, out of which only one showed significant binding affinity in the low nanomolar range (Kd 11.0 ± 2.7 nM) measured by a fluorescent method and with a higher binding affinity for the target protein when compared to BSA. This result is most likely explained by the performance of a counter SELEX round using BSA protein as target, therefore making

BSA an ideal protein to be used in subsequent experimental design. From the study of the aptamer structure it was found that it has a unique secondary folding structure that forms a stem-loop of 14 nucleotides and linear sequences on each side. CD spectra analysis ruled out the possibility that the aptamer could form a very stable G-quadruplex structure known to be very important in aptamer binding and affinity. The CD spectra provided additional proof that the aptamer folded into a B-form double helix, supporting the prediction generated by

Mfold. Additional studies of the aptamer sequence were only the 14 nucleotides were tested for binding affinity revealed that we completely lost binding affinity for LF in the concentration range tested. This evidence shows that the entire 30 nucleotide long sequence is involved in binding of ML12 to LF protein. With a good understanding of the ML12 binding characteristics we developed a colorimetric assay based on the ELISA method in which the use of an antibody was replaced by the ML12 aptamer complex with a biotin molecule. By applying this approach we were able to visually detect the presence of a constant LF enzyme concentration by varying the aptamer concentration. The Kd value obtained for this approach showed that at an aptamer concentration of 530 nM we could

33 effectively identify the presence of LF with little to no background noise. It should be mentioned that in this aptamer-based ELISA method, the Kd value obtained was much higher than the value obtained by the Cy3 fluorescence detection method. This could be explained by the requirement of multiple steps of washing during the procedure for color development in the ELISA method as well as the reaction between biotin and streptavidin (Choi et al.

2011; Oh et al. 2011). Hence, future investigation is warranted to further refine the ELISA method in an effort to reduce the apparent Kd value of the aptamer.

We additionally explored the possibility that ML12 could act as an inhibitor of the catalytic activity of LF over its substrate MEK1. By utilizing a previously reported method to measure LF enzymatic activity we demonstrated that the enzyme purified not only retained its activity but also that ML12 was able to interfere with the catalytic activity of LF by the observation of a decrease in the product generated after catalytic activity. The results from this experiment clearly provide evidence that ML12 inhibits the LF cleavage reaction.

Furthermore, to determine the potency of inhibition the data of the band intensities was plotted and gave a good fit to a single decay exponential curve with an IC50 value of 15 µM ±

1.5. This observation suggests that ML12 might block the active site of LF to prevent protease activity. The mode of action of the inhibitor is not clear but we suspect that it could be related to a competitive type of inhibition based on the observation that the IC50 and the

Kd value of ML12 are not equal. To confirm this possibility, further investigation of the kinetics of inhibition must be performed.

We performed SELEX against LF from B. anthracis and have demonstrated that the ssDNA aptamer ML12 binds to LF with high affinity. Additionally, visualization of binding using an aptamer-based ELISA method provides more evidence that ML12 tightly binds to

LF. Efforts to specify the binding sites of ML12 led to the conclusion that the full length 30- mer ML12 sequence is required for effective binding to LF. Although we do not yet possess sufficient data to propose exact binding sites of LF, a model might well involve the binding of ML12 to an active site of LF based on the inhibition results. The discovery of the aptamer against LF possessing a dual function (detection and therapeutics) may make a contribution to developing a sensitive detecting device and drug to solve problems arising from B. anthracis infection.

CHAPTER THREE

Development of Double Stranded and Single Stranded DNA Aptamers Against Escherichia coli Bacterial Cells

Introduction

Waterborne diseases are an ongoing problem worldwide. Large efforts have been put into monitoring and improving the quality of drinking water to decrease the incidence of waterborne diseases, but at the present moment the assays available to monitor water quality are slow, lack specificity, and are often expensive. Efforts to ensure the quality of drinking water involve routine monitoring of coliform bacteria in water, which are used as indicator organisms to determine if the water is contaminated with fecal waste. Monitoring of indicator organisms is easier to perform and more cost effective than specific pathogen identification in water (Berger et al. 2009). A good indicator organism must possess the following characteristics:

 The organism is present in all types of drinking water.

 The organism is found in sewage and polluted water at much higher concentrations than fecal pathogens.

 The survival time of the organism in water must be at least that of waterborne pathogens.

 The organism is at least as resistant to disinfection treatments as the waterborne pathogens.

 The organism can be detected by simple, inexpensive, and fast laboratory tests with accurate results.

 The organism is stable and nonpathogenic.

 The organism is generally not present in pristine water uncontaminated by mammalian feces.

Total coliforms (TC) are currently used as indicator organisms; they include most species of the genera Enterobacter, Klebsiella, Citrobacter, and Escherichia. TC bacteria have the following characteristics: they are aerobic or facultative anaerobes, gram-negative, non-spore-forming, lactose fermenting, and β-D-galactosidase-positive (Rompré et al. 2002).

If more detailed information about the microorganisms present in water is required then fecal coliforms are assessed (FC). FC is a subset of the total coliforms that include E. coli and some strains of Klebsiella (Bergen et al. 2009).

Because of the specific characteristics of the indicator organisms, it has been established that any water treatment that reduces the presence of these organisms will also reduce the presence of pathogens. Some of the currently used water treatments are: coagulation, flocculation, and sedimentation that remove large particulate material and bacteria; filtration using fast or slow filters to remove suspended materials that have not settled; and finally disinfection by chlorination, ozone, and ultraviolet light to inactivate pathogenic microorganisms (Berger et al. 2009). The US Environmental Protection Agency

(EPA) has approved the following methods to test for the presence of coliforms in water: multiple tube fermentation that tests for the ability of bacteria to use lactose as a carbon source, and membrane filtering that traps bacteria on a filter that can grow colonies when incubated with the appropriate media (Rompré et al. 2002). The presence/absence test is a current microbiological test employed to assess microbial diversity in water. It involves the growth of bacteria on supplemented media, specific for coliforms, which results in the presence or absence of coliform bacteria.

Molecular methods like immunological recognition of enterobacterial common antigen, PCR, and in situ hybridization are sensitive methods developed to identify TC but

37 showed cross reactivity with other members of the coliform group (reviewed by Rompré et al. 2002). The molecular techniques described suffer of low specificity, low discriminative power between live and dead bacteria, poor quantification of total bacteria in water, and depend on cell culture to increase bacterial count prior to positive identification (Rompré et al. 2002). Consequently there is an increasing need for faster and more reliable bacterial screening tests.

SELEX has been used to develop DNA or RNA aptamers that can detect or inhibit specific targets. This technique has gained particular interest in environmental and medical field for the generation of probes that can recognize complex targets like whole cells. The method for aptamer development for whole cell targets is termed cell-SELEX, and is often performed in combination with subtractive-SELEX and/or counter-SELEX using other whole cell targets. Development of aptamers by SELEX that can recognize whole bacterial cells could be a plausible approach for new bacterial detecting probes.

Cell-SELEX has extensively been used in cancer research but for the purpose of this research, we will focus our attention on applications using pathogenic bacteria. Development of aptamers against pathogens can be performed using purified proteins, membrane extracts, or whole cells. The advantage of performing SELEX with purified membrane targets is that the final product is highly specific for its target, while the use of membrane extracts or whole cells can produce aptamers with lower affinity due to the complexity of the target used. In the case of purified membrane targets, ssDNA aptamers resistant to nuclease activity have been developed against lipopolysaccharides from E. coli O111:B4 and used in the development of aptamer-immuno conjugates (Bruno et al. 2008). Protein lysates have also been used to generate aptamers against membrane fragments from Francisella tularensis,

38 resulting in the isolation of 25 unique ssDNA aptamers that were used to conduct detecting assays for F. tularensis with no cross reactivity to unrelated Gram-negative members

(Vivekananda et al. 2006). Finally, outer membrane proteins (OMP) from Salmonella enterica serotype Typhimurium were used to perform counter-SELEX against OMP from E. coli. A single ssDNA aptamer with high affinity for S. typhimurium was recovered and conjugated to magnetic beads, allowing fast concentration of whole cells in complex samples prior to PCR or RT-PCR identification of S. typhimurium (Joshi et al. 2009). The main disadvantage of this approach is that there is a decrease in the affinity of the aptamers for its target in living cells; this is due to native conformational changes generated during isolation and purification (Kim L. et al. 2009).

To eliminate the problem of target conformational changes due to isolation and purification, life whole cells have been used as targets for the generation of aptamers with potential use as microbiological probes. Bruno and collaborators used anthrax spores as targets to develop high affinity ssDNA aptamers. They used the aptamers to develop an aptamer-colorimetric assay that was able to identify the spores, at albeit low specificity

(Bruno et al. 1999). The method published by Bruno et al. (1999) served as a platform to develop a ssDNA aptamer-quantum dot fluorescent probe that was able to detect Bacillus thuringiensis spores with no interference of autofluorescence from the spores, this method was developed by Ikanovic et al. (2007). Unfortunately due to the rather simple nature of the spore, the aptamer developed had low specificity for its target (Ikanovik et al. 2007). On the same note, E. coli DH5α cells were used to develop RNA aptamers that were functionalized for a shell–single wall nanotubes-field effect transistor (SWNT-FET) assay, where binding of

E. coli cells to the aptamer generated a measurable change in conductivity that enabled the

39 positive confirmation of cell binding. Binding specificity was tested against other common food pathogens and there was no significant decrease in conductivity showing that the probe can specifically target E. coli cells (So et al. 2008). RNA aptamers have also been developed using whole cell-SELEX to specifically identify the pathogenic E. coli O157:H7 serotype, with potential use as a diagnostic ligand for food borne illness (Lee et al. 2012).

Development of aptamers against such complex targets, like whole cells, capsules, and cell walls, has proven to be challenging due to the negatively charged nature of this cell components, as well as the high similarity in their composition within species; but the advantages of developing aptamer based probes for whole cells, as opposed to the use of antibodies, makes this approach worth considering. Some of the advantages are that aptamers have equal or higher affinity for their targets, can be synthesized in vitro, do not require living organism to be developed, can be selected against a large number of targets, are highly stable and reproducible, have low to no immunogenicity or toxicity, and can be reused because of their thermal stability (WANG et al. 2009).

This work describes the characterization of a previously developed dsDNA aptamer against E. coli DH5α cells in our lab. We also described the development of a new set of ssDNA aptamers against E. coli DH5α to improve the affinity and specificity of the aptamers developed for this target.

Material and Methods

Bacterial Strains

Escherichia coli DH5α, Klebsiella pneumoniae, Citrobacter fruendii, Enterobacter aerogenes, and Staphylococcus epidermis were obtained from Carolina Biological Supply

Company (Burlington, NC). All strains were grown under aerobic conditions at 37 °C in

Luria-Bertani (LB) liquid media.

Double Stranded DNA Library

The library used for dsDNA whole cell SELEX is a random 30 oligonucleotide long

DNA fragment flanked by a forward primer region and a reverse primer region 5'-

GCGCGGATCCCGCGC-N(30)-GCGCAAGCTTCGCGC-3’, which was purchased from

IDT (IDT, Coralville, IA). Additionally two primers were purchased with the following sequence: forward primer: 5'-GCGCGGATCCCGCGC-3' and reverse primer: 5'-

GCGCAAGCTTCGCGC-3'. Template DNA was amplified by PCR (Eppendorf AG,

Austin TX) based on the method outlined in Hamula et al. (Hamula et al. 2008). Following

PCR, the dsDNA was purified by 12% native polyacrylamide gel electrophoresis (PAGE) with EtBr in 1x Tris-acetate (TA) buffer. The dsDNA band was excised and recovered using a crush-and-soak method and concentrated by ethanol precipitation (Kim et al. 2009).

Single Stranded DNA Library

The library was developed by introducing a 40 oligonucleotide long random sequence flanked by 20 oligonucleotide long forward primer region and reverse primer region. The sequence of the forward biotinylated primer is: 5’-AGCAGCACA-GAGGTCAGATG-3’ and the sequence for the reverse primer is 5’- TTCACGGT-AGCACGCATAGG-3’. To obtain the ssDNA library the template DNA sequence was amplified by an asymmetric PCR reaction with a ratio of forward to reverse primer of 4:1 and 20 rounds of PCR reaction. To purify the forward ssDNA we used streptavidin magnetic beads and followed manufacturer’s

41 instructions (New England BioLabs, Ipswich, MA). Confirmation of pure ssDNA was performed by running a 6% PAGE gel.

SELEX Using ssDNA

The method used for aptamer selection against E. coli whole cells was based on the method published by Hamula et al. (Hamula et al. 2008). Briefly, in this method a 10 ml overnight culture of E. coli is subcultured in LB medium until the O.D.600 reaches a reading of 0.5, cells are then pellet by centrifugation at 4750 g in a Beckman Coulter Allefra X-115R centrifuge at 4°C and resuspended in 1x binding buffer (1x BB: 50 mM Tris-HCl pH=7.4, 5 mM KCl, 100 mM NaCl, 1 mM MgCl2). A total reaction volume of 500 µL was used for the selection process (an appropriate volume of cell culture, that would generate 108 colonies, was mixed with ssDNA up to a final concentration of 100 nM and a final concentration of

0.25 mg/ml BSA). The mixture was incubated for 45 min at room temperature after which unbound DNA is removed by pelleting the cells and collecting the supernatant, after this step two more washing steps were performed using 1x BB. Bound DNA is eluted by exposing the cells at 94°C for 10 min, cooled on ice for 10 min and cell debris pelleted by centrifugation. The supernatant containing the eluted DNA is ethanol precipitated to concentrate and desalt the DNA fraction that was then used to generate the new ssDNA to be used for the next round of selection. To increase aptamer specificity for its target, after the second round of selection the ssDNA product was used to perform a counter-SELEX round using E. aerogenes; the supernatant from this round was amplified following the previously outlined method for ssDNA and the product was used to perform a second counter-SELEX round with a mixture of gram-negative cells (K. pneumoniae, C. fruendii, and E. aerogenes).

Finally the product from this round was used to perform a final selection round against the

42 target E. coli cells. The eluted ssDNA was amplified by regular PCR and the product was used for cloning into a pGEM-T Easy vector (Promega Corporation, Madison, WI).

Cloning and Sequencing of ssDNA Aptamers

The final round of SELEX was amplified by PCR and the dsDNA from the reaction was purified by running a 6% PAGE. The dsDNA was recovered from the gel by a crush- and-soak method and then inserted in a pGEM-T Easy vector system following manufacturer’s instructions (Promega Corporation, Madison, WI). E. coli BL21(DE3) cells

(Invitrogen Life technologies, Grand Island, NY) were transformed and grown on an agar plate containing 10 mM kanamycin. Individual colonies were picked up and cultured to extract the plasmid with an SV miniprep kit, following manufacturer’s instructions (Promega

Corporation, Madison, WI) and the presence of the insert was confirmed by restriction enzyme digestion using EcoRI (BioLabs, Inc. Lawrenceville, GA). Plasmids from colonies containing an insert of the expected size were sent to Functional Biosciences for sequencing

(Functional Biosciences, Inc. Madison, WI). The secondary structure of the isolated ssDNA aptamers was predicted using Mfold.

Kd Value Measurement for dsDNA Using a Microplate Reader

To obtain a fluorescently labeled aptamer the forward primer was modified to add a fluorescent probe to its 5’ end while the reverse primer remained unchanged. The fluorescent probe selected for this method was 6-carboxyfluorescein (6-FAM). The PCR product was purified following a previously described method. To obtain the Kd value, a constant E. coli cell concentration (107) was incubated at room temperature for 45 min with increasing concentrations of fluorescently labeled dsDNA aptamer (1-1365 pM). After

43 incubation, cells were pelleted using a centrifuge at 5000g for 10 min and the supernatant was removed. Two washing step of 100 µL of 1x BB were performed after which the cells were resuspended in 100 µL of 1x BB and incubated at 94°C for 10 min. Dead cells were removed by centrifugation and the supernatant was collected to obtain fluorescent readings on a Thermo Flouroskan Ascent FL with an excitation wavelength of 485 nm and an emission wavelength of 535 nm. Data collected by Shipley S as part her thesis work

(Shipley S. 2010).

Kd Value Measurement and Specificity Test for dsDNA Aptamer Using Fluorescent Imaging

To test specificity the dsDNA aptamer generated for E. coli we used a different fluorescent molecule to decrease cell fluorescent interference. In this method we fluorescently labeled the forward strand of the aptamer with cyanine 3 (Cy3) on the 3’ end of the sequence. The aptamer was then conjugated with the reverse strand and heated up to

95°C, followed by a short period of incubation at the aptamer annealing temperature, after which the sample was slowly cooled down to obtain dsDNA. The cell lines used to test specificity were grown overnight in LB medium and then subcultured in 30 ml of LB medium up to an O.D.600 of 0.5. Each culture was centrifuged at 3000 rpm to obtain a cell pellet and remove the LB medium. Pellets were washed two times with 2 ml 1x BB and finally resuspended to a volume of 1 ml in 1x BB. Aliquots of 100 µl of the cell culture were transferred to an ependorff tube and cells were pellet to remove the supernatant. To perform the binding assay, cell pellets were exposed to increasing concentrations of dsDNA (0 nM to

2000nM) and the volume was filled up to 100 µL of 1x BB, followed by incubation in the dark for 30 min at room temperature. After incubation, unbound dsDNA was removed by centrifugation and the pellet was washed two times to remove remaining unbound dsDNA.

Then cell pellets were resuspended in 100 µL of 1x BB and incubated at 94°C for 10 min to recover the bound dsDNA from the cells by centrifugation at 14000 rpm. Supernatant was transferred to a 96 well plate and images were capture using a Typhoon FLA9000 (GE

Healthcare Life Sciences, Pittsburgh, PA). Fluorescent images were analyzed with ImageJ program (freely available at http://rsbweb.nih.gov/ij/), and the fluorescent intensity of each sample was plotted to obtain a non-linear regression to calculate the Kd for each bacterial strain used.

Kd Determination for ssDNA Aptamers Developed Against E. coli DH5α

To test the binding affinity of the ssDNA aptamers developed against whole cells, we used a low binding cellulose filter with a molecular weight cutoff of 30,000 Da to ensure that unbound ssDNA could pass through the filter and only bound ssDNA to the cells was retained. Two washing steps were performed to ensure no unbound DNA remained in the filter. The absorbance of ssDNA was measured at 260 nm and the difference between the absorbance of unbound and bound ssDNA was used to calculate the binding affinities for each aptamer.

Experimental Results

Selection of dsDNA Aptamers

For the selection of dsDNA aptamers that bind to E. coli DH5α cells, the dsDNA library was incubated with the bacterial cells and was heat eluted for further amplification of the dsDNA by PCR to enrich the pool of aptamers in each round of selection. To increase the specificity of the aptamers for its cellular target two rounds of counter-SELEX were performed, one of them against the gram-positive bacteria B. subtilis and the other round was

45 performed against the gram-negative bacteria E. aerogenes. After six rounds of cell-SELEX and two rounds of counter-SELEX the resulting dsDNA was cloned into a vector and 15 transformed colonies were sent for sequencing. Only four colonies showed a 30 bp long insert, three of them with the same sequence and one that only differed from the rest by 1 bp.

(Table 3). Data collected by Shipley S as part her thesis work (Shipley S. 2010).

Table 3. Binding Affinity of a dsDNA Aptamer Developed Against E. coli DH5α Cells

Aptamer dsDNA sequence Frequency Kd value

SS1 5’-AGCGAAGCACCTTTCACTGTACCTATTTCC-3’ 3 96.98 pM

SS2 5’-AGCGAAGCACCTTTCACTCTACCTATTTCC-3’ 1 NA

* Oligonucleotide sequence for the aptamers random region developed by Shipley S. 2010

Specificity and Dissociation Constant for dsDNA

Two rounds of counter-SELEX were performed to increase specificity of the aptamers for its target. The first round was performed using the non-related bacterial strain

B. subtilis, this strain vastly exist in water and is commonly used in water treatment to help breakdown complex molecules. The second round was performed using E. aerogenes, which is a member of TC commonly assessed in water quality screenings. To test the effectiveness of the counter-SELEX rounds, we performed four independent tests using the dsDNA aptamer modified with a fluorescent molecule and incubated the aptamer against B. subtilis,

C. freundii, S. epidermis, E. aerogenes, as well as E.coli to visually compare the binding affinities. Results of the specificity tests shown in Figure 17 revealed that the dsDNA aptamer can bind to all strains of bacteria studied, but binding affinities are different. The aptamer showed the highest affinity for E. aerogenes with a Kd value of 474.5 nM and the

46 lowest affinity for S. epidermis with a Kd value of 1.4 µM, while the previously reported Kd value for the target was 96.98 pM. Table 3 and Figure 14 show the Kd values for each strain and the fluorescent image from specificity tests using the Cy-3 dsDNA aptamer.

Table 4. Dissociation Constant Determination of a dsDNA Aptamer to Different Bacterial Cells.

Bacteria Dissociation constant E. coli 142.8 µM* C. freundii 744.3 nM E. aerogenes 474.5 nM B. subtilis 343.6 nM* S. epidermis 1.4 µM *No saturation point was reached for these aptamers in the range of concentrations studied.

Figure 17. Specificity test using Cy3 labeled dsDNA. A constant numbers of cells were exposed to increasing concentrations of a fluorescently labeled aptamer as explained in the methods section. Image was acquired using a Typhoon image system with an excitation wavelength 532 nm and emission wavelength of 575 nm.

Selection of ssDNA Aptamers

For the selection of ssDNA aptamers that can bind to E. coli DH5α cells, an 80 oligonuclotide long ssDNA library generated by asymmetric PCR was incubate with the target cells (Figure 18). After incubation, bound ssDNA to the E. coli cells was recovered and amplified by asymmetric PCR and the product was resolved on a 6% native polyacrylamide gel.

Figure 19 shows the result of the first cell-SELEX round perform using E. coli DH5α cells. There are five fractions recovered but only the ssDNA recovered from the elution fraction is of interest and was used to generate a new ssDNA pool to perform a subsequent counter-SELEX round. Figure 20 shows the product recovered from the second round of selection, where the fraction recovered from the incubation is further amplified to perform the next round of selection. Figure 21 shows the third round of selection using E. coli cells and the eluted DNA was further amplified to perform a counter-SELEX round using a combination of three gram-negative bacterial strains. Figure 22 shows the results of the counter-SELEX using a mixture of three gram-negative bacteria. Figure 23 shows the final round of selection in which E. coli cells were used and the eluted fraction was amplified to generate a dsDNA pool that was cloned into pGEM-T Easy vector. A total of 15 colonies were isolated and sent for sequencing.

Figure 18. Generation of an 80 oligonucleotide long ssDNA library for cell-SELEX experiments. An 80 oligonucleotide long ssDNA library was amplified by asymmetric PCR with a ratio of 4:1 forward:reverse primer. Separation of the ssDNA from the dsDNA was performed by the use of streptavidin magnetic beads and the elution product was electrophoresed in a 6% native gel. Line 1 shows the molecular size ladder and line 2 shows the ssDNA recovered from the streptavidin beads. Bands belonging to ssDNA were excised and the ssDNA recovered by crush-and-soak. Salts were removed by ethanol precipitation.

Figure 19. Cell-SELEX round one. The product from PCR amplification of each fraction collected during the selection protocol for round one was run on a 6% native gel. Line1 shows the PCR amplification of the DNA eluted after heat denaturation and supernatant recovery, line 2, 3, and 4 show the PCR amplified fractions recovered during each washing step, line 5 shows the product of PCR amplification of the supernatant after incubation and cell pelleting.

Figure 20. Counter-SELEX round two. PCR amplification of each fraction collected during selection using E. aerogenes. Line 1 shows the PCR product from amplification of the supernatant, line 2 and 3 show the PCR amplification of the two washing steps, line 4 shows the PCR product from a control with no 80 oligonucleotide ssDNA, and line 5 shows the PCR product from the eluted DNA that was bound to the cells. Samples were run on a 6% native gel and the band of interest recovered by a crush-and-soak method.

Figure 21. Cell-SELEX round three. PCR amplification of each fraction recovered from the SELEX procedure using E. coli DH5α. Line 1 contains the PCR product from the incubation fraction, line 2 and 3contain the PCR product from the two washing steps, and line 4 contains the PCR product from the DNA recovered from the cell pellet. Samples were run on a 6% native gel and the band of interest recovered by a crush-and-soak method.

Figure 22. Counter-SELEX round four. PCR amplification of each fraction collected from the counter-SELEX step using a pool of three gram-negative bacterial strains (C. freundii, E. aerogenes and K. pneumonia). Line 1 contains the PCR amplified supernatant collected from the incubation step, line 2 and 3 contain the PCR product from the two washing steps, and line 4 contains the PCR amplified eluted fraction from the cell pellet. Samples were run on a 6% native gel and the band of interest recovered by a crush-and-soak method.

Figure 23. Cell-SELEX round five. PCR amplification of each fraction collected from the SELEX step using E. coli DH5α. Line 1 contains the PCR amplified supernatant collected from the incubation step, line 2 and 3 contain the PCR product from the two washing steps, and line 4 contains the PCR amplified eluted fraction from the cell pellet. Samples were run on a 6% native gel and the band of interest recovered by a crush-and-soak method.

The sequencing results show great diversity in the final ssDNA product form the

SELEX experiment, possibly due to the low number of selection rounds (Figure 24).

Attempts were made to perform further rounds of selection but amplification and extraction of ssDNA for further rounds failed to produce a clear band of ssDNA that could be used for further selection. Appendix B (Figures B1-B5) shows the product form asymmetric PCR for each round. It can be observed from these results that as the selection of ssDNA for E. coli progresses the PCR reaction generated less ssDNA. The PCR product was also increasingly enriched with larger fragments of DNA that were preferentially amplified during PCR. The difficulty encounter to generate ssDNA during the final rounds of SELEX was the determining factor that forced us to stop the experiment after the fifth round. By comparing the aptamer sequences it becomes clear that there are no fragments of the sequence that are similar. Determination of the dissociation constant for each aptamer shows that the binding affinity of the aptamers for E. coli DH5α cells is roughly in the low µM range. For the aptamers EW1, EW14, and EW18, there was no significant binding to the target in the ssDNA concentration range used (0.05 µM to 0.5 µM) so they were not used for further characterization of their binding properties. To assess the specificity of the remaining aptamers the same method was employed, but the target cell for binding was exchanged for

C. freundii, E. aerogenes and S. epidermis. Results for specificity revealed that most of the aptamers had some interaction with one or more of the bacteria used to test specificity, and only aptamers EW3, EW15, and EW22, showed no binding to C. freundii, E. aerogenes and

S. epidermis in the concentration range of ssDNA used (0.05 µM to 0.5 µM).

Figure 24. Graphical representation of the alignment of the15 aptamers. 15 ssDNA aptamers developed for E. coli DH5α were isolated and sequenced from the last round of selection showing unique sequences (EW˗˗˗Ethan White).

Table 5. Sequence of The 40 Oligonicleotide Long Random Region for Each ssDNA Aptamers Developed Against E. coli DH5α Cells.

Aptamer ssDNA sequence (5’ to 3’) Kd value (µM)

EW1 TGCGTAGGGCGCCCGGTTCACCGTCGACCCCTTGAGCACC N/A

EW2 GCCAGGTCTGTGACGATAGCATCAGTTGTTCCTAGTGTAC 3.93

EW3 CCCCCCACAAGTCCCANCCCCCACCAACCCACTCTCTACC 2.58

EW4 CCGAACGCTTCCTCTGTCGCGCCCATCGTTCGCAACCGAC 5.51

EW5 GGCGACCAAACCAATCACGACGTCAATTGGCGATGCCTAC 14.3

EW13 CAGGCAGGGGCCCCGACAGGCGCTCTGTGGAATGTGACT 24.6

EW14 GCGATCGGCGGCTAAGCCTGGGCAATCATCCTGAGCCCCC N/A

EW15 TGCAGTACTTGTCCGATACGCGGGTATCTCGTCGATGCGG 2.27

EW16 CCGCTAGACCGCCCCCGCTCCACGGCGCCTTTGCAAACTC 1.75

EW18 GCGGCGACAGGGGGCGAGCGGTGACCAGCCATGGAGGTGT N/A

EW22 GACGGTACCTGCTCCAACCACACCTTGAGGGTGAGCCGTA 1.49

EW23 GGTCTACGATGATTACGCTAGGTTGGTGACCATCGCGGGG 1.57

EW24 GCAGAGACCAGGGCGCCGGAGACAGCTCTCGAGATGCGTT N/A

EW25 CCAGGGGGGTGACCCTCGTCGTGCCTCACGTATCTGGGGC 14.2

EW26 GGCCGCACACGTGATTGCCCACCCGTTCCCCTGCCTCGGC 9.96

Kd determination by low binding regenerated cellulose membrane separation

Discussion

In this study we used a cell-SELEX method combined with counter-SELEX to obtain dsDNA aptamers that bind to E. coli DH5α cells. The bacterium was selected as the target cell for the experiment because it is a derivative from E. coli K12, which is an extensively

57 studied organism with a complete genome sequence, safely used in laboratory experiments, and currently used in water quality screenings. Additionally the use of E. coli DH5α to successfully develop RNA aptamers against whole cells shows that this organism is a good target for cell-SELEX (So et al. 2008). As a result of the cell-SELEX on E. coli DH5α using a dsDNA library, only one dsDNA aptamer was isolated after eight rounds of selection.

Previous study of the binding affinity of the aptamer for its target initially performed by

Shipley S. using a fluorescently labeled aptamer-FAM revealed that the aptamer binds with a binding affinity of 96.98 pM (Shipley S. 2010). The recent study of the same aptamer using a different fluorescently label (Cy3) aptamer and florescent imaging system showed an estimated binding affinity of 143 µM. The difference in binding affinity found between the two techniques employed could be explained by the possibility that the FAM labeled aptamer had a background fluorescent reading coming from the bacterial cells that was not taken in consideration during previous experimental set up. During this research it was found that proteins and also whole cells can emit a signal at the excitation and emission wavelengths used to test FAM fluorophores (Exc. 494 nm and Em. 521 nM) thereby introducing an unexpected error in the readings.

Specificity tests performed for the dsDNA aptamer show that the aptamer has a low specificity for its selected target. The dsDNA aptamer binds to other bacterial strains with differing affinities. From the bacterial strains used, the results show that the aptamer binds more tightly to E. aerogenes with a Kd value of 474 nM and less strongly to S. epidermis with a Kd value of 1.4 µM. The lack of specificity shown by the aptamer can be explained by the simple nature of the dsDNA which folds in a specific helical structure that significantly decreases the possibility of the aptamer to interact with membrane molecules based on their

58 shape, either by fitting into a grove of a protein or by surrounding a particular segment of a target as the oligonucleotide folds into their three-dimensional structure (Hermann et al.

2000). Therefore most likely the dsDNA aptamer is interacting with the cell by electrostatic forces and not by shape compatibility. Another explanation can be found in the possibility that the aptamer is interacting with a commonly expressed lipopolysaccharide or membrane protein that is present in all five bacterial strains used and that the change in Kd could be a consequence of differential expression of the lipopolysaccharide by the different strains. To further investigate this possibility, a lipopolysaccharide characterization of the membranes of all five bacterial strains could be undertaken and then binding affinity evaluation of the aptamer against the purified lipopolysaccharides could be determined. The fact that the aptamers can bind to other bacteria belonging to the coliform group is a positive finding but it raises the question as to how specific it can be in a complex sample that has other bacteria not necessarily from the coliform group. Another concern with the aptamer is its sensitivity; the fluorescent imaging test shows that we are able to detect a positive signal at an aptamer concentration of 1500 nM for all five bacterial strains at a constant cell number of 105 cell/ml and the current tests requires a positive coliform identification of 1 cell in 100 ml of water to consider that the water is not safe for human consumption. This results indicate that in order for our dsDNA aptamer to be able to positively detect the bacterial cells, an enrichment step by filtration or culture will be required.

In the case of ssDNA aptamers selected for E. coli DH5α, we obtained a large diversity of aptamers from the final round of selection. This clearly indicates that further rounds of selection needed to be done. Before concluding the selection, attempts to amplify the ssDNA product from the fifth round were undertaken by performing changes in the

59 primer ratio and annealing temperatures for PCR, but despite the changes made we failed to produce a clear band of ssDNA that could be used for subsequent SELEX rounds. One possible explanation for this outcome is that by the fifth round, the ssDNA recovered was enriched with sequences that had a higher tendency to form dimmers and multimers, therefore generating a final PCR product with enlarged size, and amplification of these fragments was predominantly favored during the PCR reaction. The determination of the binding affinities for each ssDNA aptamer showed that they all had very similar binding affinities in the low µM range, showing that five rounds of selection where sufficient to generate aptamers with decent binding affinities but was not sufficient to enrich the library with a particular sequence which would present tighter binding properties. Studies of the binding specificity for each of the aptamers against other cell types indicated that only a few aptamers appeared to be specific to E. coli cells in the range of concentrations used for the experiment. Future investigation of binding affinities employing more sensitive methods like the use of fluorescently labeled aptamers can be performed with aptamers EW3, EW15, and

EW22.

CHAPTER FOUR

Site Directed Mutagenesis Study on GrxA a Protein from Cyanobacteria Synechocystis sp pcc6803 to Elucidate its Mechanism of Action

Introduction

Arsenic is the 20th most common element on earth and is rarely encountered as a free element. It is present in rocks and in soil generally as a metal or it can be found in soil,

3- water, and living organisms as oxyanions of arsenic like arsenate (V, AsO 4) or arsenite (III,

- AsO 2 or As(OH)3) (Messens et al. 2006). Arsenic is also released into the environment by human activity (coal burning, metal smelting, acidic mining, and the semiconductor industry) and by microbial activity in the form of arsenite (Mukhopadhyay et al. 2002). Arsenic species produce different levels of toxicity that varies in the following order: arsenite > arsenate > monomethilarsonate > dimethilarsinate. The high toxicity of arsenite is a consequence of its high affinity for sulfhydryl groups, lipoic acids, and cysteinyl residues that are commonly found in many enzymes. The formation of a bond between arsenite and sulfur results in the inhibition of enzyme activity affecting enzymes like glutathione oxidoreductases and thioredoxin oxidoreductases (Sharma et al. 2009). Depending on the concentration of toxins ingested by humans, two different toxic effects can be observed: acute effects that are generally caused by ingestion of contaminated food or water and requires immediate medical attention to avoid severe dehydration and shock, whereas the sub-acute toxic effects are caused by long term exposition and can cause a number of different serious clinical conditions (Jain C. K. 2000). Arsenite is the product of arsenate reduction by microorganisms that have evolved a mechanism to overcome the accidental

61 incorporation of arsenate by the phosphate transport systems that mistakenly incorporates arsenate into the organism due to its chemical similarity with phosphorous. The mechanism involves two steps that allow the reduction of arsenate to arsenite by an enzyme named arsenate reductase, and the product of the reaction is then transported out of the cell by an inducible transporter (Li et al. 2003). Arsenate reductases can be divided in three families depending on the redox proteins with which they are coupled with: the thioredoxin-coupled family (Trx) characterized by S. aureus ArsC enzyme from plasmid pI258; the glutaredoxin- couple family (Grx/glutathione (GSH)) characterized by E. coli ArsC from plasmid R773; and the thioredoxin/glutaredoxin hybrid family characterized by Synechocystis sp. ArsC from strain PCC 6803. For the purpose of this research, we will focus our attention on the mechanism of action of the last Grx family that structurally belongs to the thioredoxin dependent family but uses the Grx/GSH system as electron donors for reduction (Messens et al. 2006; Lopez-Maury et al. 2009). Based on the amino acid sequence of the SynArxC enzyme, we would expect this protein to undergo a mechanism of reaction similar to the one outlined in Figure 25 where the enzyme in the fully reduced state will have an anionic Cys residue that will perform a nucleophilic attack on the arsenate substrate forming a covalent intermediate that is then attacked by an intramolecular Cys residue to form an intramolecular disulfide intermediate and release of arsenite. Finally another intramolecular Cys residue attacks the disulfide bridge formed and restores the thiolate form of the initial ArsC enzyme and the intramolecular disulphide is later reduced by Trx. This mechanism depends on three

Cys residues all provided by ArsC (Messens et al. 2006).

The requirement of Grx/GSH by SynArxC can be explained by the study of the

Grx/GSH coupled mechanism outlined in Figure 26. In this mechanism the enzyme reacts

62 with arsenate to generate a covalent intermediate that then interacts with GSH to generate a tertiary glutathionylated arsenate intermediate that is consequently attacked by reduced Grx to produce dihydroxy arsenite and oxidized Grx. The reaction is then followed by the formation of a monohydroxilated arsenite intermediate that is then further hydroxylated and released with the regeneration of reduced ArsC. This mechanism requires three Cys residues and they are provided by ArsC, Grx, and GSH (Messens et al. 2006). Oxidized Grx is further reduced by GSH to form disulfide GSSG, which is subsequently reduced by glutathione reductase at the expense of NADPH, as it can be appreciated from the mechanism outlined in

Figure 27 (Herrero et al. 2007).

Figure 25. Catalytic cycle of Trx-coupled arsenate reductase from S. aureus. Cycle characterized by four steps that start with the formation of a complex between ArsC and arsenate, followed by the formation of a disulfide bridge that transfers two electrons to arsenate to form arsenite and release the arsenite leaving ArsC in the oxidized form that is latter reduced by Trx to reconstitute the initial Trx.

Figure 26. Grx-coupled arsenate reductase mechanism of action. Abbreviations: Grx, glutaredoxin; GSH, glutathione; ArsC, arsenate reductase C. (modified from Messens et al. 2006).

Figure 27. Schematic representation of the Grx system involved in reduction of dithiol groups.

Research into the mechanism of action of SynArsC has revealed so far that the ArsC requires Cys8, Cys80, and Cys82 for reduction of arsenate to arsenite resembling the mechanism of action of the Ars reductases belonging to the Trx-coupled mechanism.

Additionally, mutations on any of the previously mentioned Cys residues completely abolish

64 the enzyme activity (Kim et al. 2012; Li et al. 2003). Sequencing analysis of Synechocystis sp. genome identified three genes that code for three Grx proteins: GrxA, GrxB, and GrxC.

Comparison of the gene sequences with existing Grx proteins from other well studied organisms showed that GrxA and GrxB belong to the dithiol glutaredoxins group, with a characteristic active site containing two cysteine residues (CXXC motif) and are involved in the reduction of protein disulfides and reduction of glutathione (GSH)-protein mixed disulfides (deglutathionylation), whereas GrxC belongs to the monothiol glutaredoxin group with a single cysteine active site (CGFS motif) that can only catalyze deglutathionylation reactions (Gao et al. 2009). The mechanism of action for both Grx groups has a common first step as shown in Figure 28 that involves the nucleophilic attack of a glutathione-mixed disulfide by the amino-terminal cysteine of the active site, resulting in the release of the deglutathionylated substrate. In the monothiol mechanism, the glutathionylated Grx is then reduced by a second molecule of GSH forming a glutathione disulfide (GSSG) and reconstitutes the reduced form of Grx. In the dithiol mechanism the second step consists of a nucleophilic attack performed by a vicinal Cys on the active site of Grx, leading to the formation of oxidized Grx and GSH (Herrero et al. 2007). Recently it has been demonstrated that only GrxA and GrxB are involved in arsenate reduction by SynArsC and that there is no activity when GrxC was tested (Lopez-Maury et al. 2009). Additionally, studies on the activity of GrxA from E. coli coupled with SynArsC showed that only Cys11 from E. coli

GrxA was essential for ArsC reduction, indicating that GrxA reduces a mixed ArsC- glutathione disulfide and not an internal cysteine-cysteine disulfide on ArsC (Li et al. 2003).

Results that are further supported by studies performed using a GrxA from Synechocystis sp

65 conclusively showing that GrxA is involved in deglutathionylation reactions to restore the active form of ArsC (Kim et al. 2012).

The proposed mechanism of action based on experimental results for this type of arsenate reduction is outlined in Figure 29. In this mechanism, Cys 80 and 82 are initially in an oxidized state and steps 1 to 3 resemble the mechanism encountered for the GSH/Grx- coupled ArsC reduction producing a glutathionylated ArsC on Cys 8; in step 4 an internal disulfide exchange takes place shifting the mixed disulfide from the active site to a more solvent accessible site were the GrxC can deglutathionylate Cys82 followed by disulfide interchange in ArsC to restore the original oxidized form of the enzyme (Li et al. 2003).

Based on the information available from the mechanism of action for SynArsC reductase, it remains unclear if GrxA can participate in reduction of protein disulfide bonds, therefore we proposed to perform a mutagenic study on GrxA to test whether or not this enzyme is capable of performing disulfide reduction reactions and what would be the implications of this type of activity on the mechanism proposed for arsenate reduction by

Synechocystis sp.

Figure 28. Dithiol reduction mechanism by Grx protein. A) Two Cys-mechanism involving two Cys residues for the reduction of dithiols requiring two GSH molecules for reconstitution of the reduced Grx. B) One Cys-mechanism involving only one Cys residue for the reduction of dithiols requiring only one GSH molecule.

Figure 29. Proposed mechanism of action for Synechocystis sp. ArsC reductase. Abbreviations: Grx, glutaredoxin; GSH, glutathione; GSSG, glutathione disulfide; SynArsC, Synechocystis sp. arsenate reductase (Kim et al. 2012).

Materials and Methods

Material

All primers were purchased from Integrated DNA Technologies (IDT, Coralville,

IA). Quik-Change site directed mutagenesis kit was purchased from Agilent Technologies

(Agilent, Santa Clara, CA.) and an SV mini prep kit was purchased from Promega for plasmid extraction (Promega Corporation, Madison, WI.).

Site Directed Mutagenesis

Previously in the laboratory a set of GrxA mutants were developed using the GrxA gene (Gene ID 954155). The mutants generated carried an amino acid substitution of a Cys

67 residue for a Ser residue. The mutants used for site directed mutagenesis had the following mutations: mutant one had a mutation on Cys18 and is identified as C18S, and mutant two had a double mutation on Cys18 and Cys70 identified as C18S/C70S. To generate mutants caring an additional mutation at Cys36 we used the two previously developed mutants and designed two primers for the introduction of an alanine in position 36 to remove the cysteine residue.

The sequences of the primers designed for site directed mutagenesis are as follows:

Forward primer: 5’-GAGTTCCAAGAATATGCCATTGACGGCGACAAC-3’

Reverse primer: 5’-GTTGTCGCCGTCAATGGCATATTCTTGGAACTC-3’

Plasmid (pQE80) with the gene of interest was extracted from transformed BL21

(DE3) cells and used as templates for the PCR reaction with the primers designed for site directed mutagenesis. The PCR components were mixed following manufacturer’s instructions for the use of the Quik-change site directed mutagenesis kit. PCR reaction mixture was made by combining the following reagents: 1 µL dNTPs, 1 µL plasmid DNA

(about 200ng), 1 µL forward primer (2 µM final concentration), 1 µL reverse primer (2 µM final concentration), 5 µL 10X reaction buffer, 41 µL double distilled water, and 1 µL pfu turbo DNA polymerase (2.5 u/µL). PCR program was set as: step one denaturing at 95°C for

40 sec, step two denaturing at 95°C for 30 sec, step three annealing at 58°C for 1 min, step four elongation at 69°C for 5 min and 8 sec; step two throughout four was repeated 15 times with a final elongation time of 6 min. PCR product was then treated with 1 µL Dpn1 and incubated at 37°C for 3 hours. 2 µL of the Dpn1 treated PCR product was incubated with competent E. coli BL21 (DE3) cells for 30 min on ice, cells were then briefly heat shocked at

42°C for 30 sec and placed on ice for 20 min. 500 µl of NZY Broth was added to the

68 transformed cells and they were incubated at 37°C for 1 hour in a shaker set at 205 rpm. 100

µL of transformed cells were spread on an agar plate containing 50 µg ampicillin and grown overnight. The next day three single colonies were picked up and grown to extract the plasmid following manufacturer’s instructions for the SV miniprep kit (Promega

Corporation, Madison, WI). Three plasmid samples for each mutant were sent to Functional

Biosciences for sequencing.

Protein Expression and Purification

Protein was expressed in E. coli BL21 (DE3) cells. Briefly, transformed cells were grown overnight at 37°C in a 10 ml culture tube containing 50 µg/ml ampicillin in LB medium. The next day the culture was transferred to a 1 L, 50 µg/ml ampicillin LB medium and grown at 37°C until O.D.600 reached 0.6, at which point IPTG was added and the culture was further grown for 5 h at 30°C. Cells were collected using a Beckman Coulter bench top ultracentrifuge and the pellet was saved at -80°C until use. To extract the protein, cells were resuspended in 30 mM Tris buffer pH=8.0 and sonicated using 5 times 30 sec pulses followed by 30 sec of cooling down periods. Cell debris was pelleted by ultracentrifugation and supernatant was filtered using a 0.45 µm filter (Millipore Corporation, Billerica, MA).

Filtered sample was loaded onto a Ni+2 column pre-equilibrated with resuspension buffer.

Column was washed with 12 column volumes of 3% imidazole and the protein sample was eluted using a step gradient from 3%-12% imidazole, followed by two column volumes of

12% imidazole and a final step of elution from 12% - 30% imidazole. Fractions containing the protein were concentrated and further purified using a gel filtration system using a 30 mM Tris-HCl pH=8 buffer, the protein retention time on the column was of 3 hours.

Determination of the Oxidation/Reduction Potential of Mutant GrxA Proteins

To establish the oxidation/reduction potential of the new mutant GrxA proteins two different methods were used.

Method 1: Fluorescent Protein Titration

To test the protein oxidation/reduction potential in the region between -100 mV and -

240 mV, a 2 mM glutathione (GSH) and glutathione disulphide (GSSG) solution was freshly made in 100 mM HEPES buffer pH=7.0 before starting the titration experiment. The right proportion of oxidizing and reducing agents were mixed to generate the different voltage states. A constant protein volume of 10µl from the stock was added to the mixture and incubated for 3 h at room temperature. After incubation, 7 µl of a saturated solution of the fluorescent molecule monobromobimane (mBBr) was added and incubated at room temperature, and protected from light for 20 min. Protein was then precipitated by the addition of 500 µl 20% trichloroacetic acid (TCA) and incubated on ice for 30 min. After incubation the precipitated protein was pelleted by centrifugation at 14,000 rpm for 10 min.

Supernatant was removed and the pellet was washed with 500 µl of a 1% TCA solution. The pellet was then resuspended with 400 µl of 100 mM Tris-HCl pH=8.0 and 1% SDS. The volume of each sample was adjusted to 2.6 ml using 100 mM Tris-HCL pH=8.0 buffer and fluorescent readings were acquired using a 3 ml quartz cuvet and a spectrofluorophotometer

(RF-5301PC; Shimadzu, Houston TX) with an excitation wavelength of 380 nm and emission wavelength of 450 nm. Titration data was fitted to a single-component Nernst

Equation for a two-electron couple.

Method 2: Methoxy-PEG Maleimide (2,000 Da.) Modifying Thiol Group Titration

Reduced and oxidized GSSG and GSH were freshly prepared and mixed in the right proportion to obtain a solution with potentials ranging from -100 mV to -240 mV. A fixed amount of protein was mixed in the titrating solution and incubated for 3 h at room temperature. After incubation the mixture was treated with 500 µl of a 20% TCA solution and incubated on ice for 30 min. Precipitated protein was pelleted by centrifugation and the supernatant was decanted. The pellet was washed with 1% TCA solution followed by an additional washing step using cold acetone. Excess acetone was removed by vacuum and the pellet was resuspended with 40 µl of 2 mM malPEG (2 kDa) in 50 mM HEPES buffer pH=7.0 and 2.5% SDS. Solution was incubated at room temperature for 1.5 h. After incubation, samples are loweded on a 15% PAGE for electrophoresis. The gel was stained with coomassie brilliant blue and a picture of the gel was taken with a Canon camera.

Mass Spectrometry

Mass spectrometry studies were conducted to test the effect of the reducing and oxidizing agents, like GSH and DTT, on the wild-type and mutant Grx proteins. The mass analysis utilized an Accela liquid chromatography coupled with LTQ Orbitrap Discovery mass spectrometer (Thermo Electron, Bremen, Germany), while using positive electrospray ionization (+ESI). The wild-type and mutant Grxs (~100 µM) were injected (5 µl) into a 15 cm x 2.1 mm extended-C-18 column (Agilent Technologies, Palo Alto, CA) with a mobile phase gradient (20 – 97%) consisting of acetonitrile and 0.1% (v/v) formic acid in water.

The column temperature was set at 30°C with a 400 µL/min flow rate. Orbitrap mass analyzer was used to obtain a full-scan mass spectra (m/z range: 200 – 4000) of eluting compounds, and Xcalibur v.2.0.7 software was used to deconvolute the spectrum. The

71 following electrospray source conditions were also applied: sheath and auxiliary gas flow 60 and 10 arbitrary units (a.u.), respectively; heated capillary temperature 275°C; electrospray voltage 4.5 kV; capillary voltage 27 V; tube lens voltage 240 V.

Experimental Results

Site Directed Mutagenesis

To evaluate the participation of Cys residues in the formation of intra disulfide bonds to stabilize the enzyme after electron donation, two mutants were created. By the use of the forward primer and the reverse primer in the PCR reaction, using as template the pQE80 plasmid containing the GrxA-C18S mutant protein, we obtained a new plasmid containing a new GrxA mutant protein with two Cys mutations (C18S/C36A). To obtain the triple GrxA mutant the same primers were used but the template GrxA sequence used for PCR was different; the template contained a double Cys mutation (C18S/C70S) that after PCR generated the new triple Cys mutant (GrxA-C18S/C36A/C70S). Figure 30 A shows the sequence of the wild type GrxA protein with the translated amino acid sequence. Figure 30

B shows the graphical representation of the gene sequences belonging to the wild type GrxA gene, and the two new mutants developed by site directed mutagenesis. The shaded regions on the graphical representation show areas of identical sequences, while the clear regions represent areas where the sequence does not match. The red squares show the sites where the cysteine residue was changed for another amino acid. The first mutation is found in the amino acid sequence number 18 where the Cys is replaced by a serine. The second mutation is found in the amino acid sequence number 36 where the Cys is replaced by an alanine, and third mutation is found in the amino acid sequence number 70 where the Cys was replaced by

72 a serine. The predicted molecular weight for each protein was calculated to be 11,031 Da for the wild type; 10,983 Da for the double mutant; and 10.967 Da for the triple mutant.

1 ATG CGC GCG CTG GCG CTG CTG AAA CGC AAA GGC GTG GAA TTT CAG 45 1 M R A L A L L K R K G V E F Q 15

46 GAA TAT TGC ATT GAT GGC GAT AAC GAA GCG CGC GAA GCG ATG GCG 90 16 E Y C I D G D N E A R E A M A 30

91 GCG CGC GCG AAC GGC AAA CGC AGC CTG CCG CAG ATT TTT ATT GAT 135 31 A R A N G K R S L P Q I F I D 45

136 GAT CAG CAT ATT GGC GGC TGC GAT GAT ATT TAT GCG CTG GAT GGC 180 46 D Q H I G G C D D I Y A L D G 60

181 GCG GGC AAA CTG GAT CCG CTG CTG CAT AGC 210 61 A G K L D P L L H S

Figure 30 A and B representing the sequence for wild type GrxA and the newly generated mutants. A: wild type GrxA sequence with the corresponding amino acid sequence. B: graphical representation of the gene sequence alingment for the two new mutant GrxA proteins and the wild type GrxA protein. The shaded area shows identical sequence while the clear areas show the sites at which the cysteine residues were mutated, these areas are also highlighted by red squares.

Determination of Middle Point Potential for the Will Type GrxA and the Two GrxA Mutants

To elucidate the mechanism of the GrxA protein and find if this protein can participate in GSH disulfide oxidoreduction as well as protein disulfide reduction, we measured the redox potential of the double and triple mutant using two different methods, as outlined in the material an methods section, and compared the potential of the two mutants with the potential of the wild type protein. Results obtained for the oxidation/reduction titration of each protein using the malPEG method showed on the SDS-polyacrylamide gel that the wild type GrxA exposed to a potential gradient from -100 to -240 mV had a change in mass corresponding to more than one interaction with malPEG polymers (Figure 31 and

32), therefore generating more than one band on the gel as the potential changed. For the two Cys mutant the same range of potential showed that there are two distinctive bands on the gel corresponding to non-malPEG bound (lower band) and malPEG bound protein (upper band). Finally, the triple mutant showed two bands like in the case of the double mutant, but when comparing the distance traveled on the gel it can be noticed that the triple mutant appears to have a smaller change in molecular weight when compared to the double mutant

(Figure 31 and 32). Figure 33 shows the difference in mass increment due to 100% reduced protein with malPEG bound to the protein, the mass increment showed the following pattern: wild type was larger than double mutant and triple mutant, and the difference between the mutants was very small; result that correlates with the number of Cys residues available in each protein. Figure 33 also shows a second band at about 36 kDa that was treated with DTT to assess the possibility of the formation of a dimer Grx, but this was not the case, indicating that the band observed was a co-eluting protein other than Grx. Because of this observation we decided to test a more electronegative potential between -240 mV and -320 mV and

74 found that there was no binding of malPEG to any of the mutants while for the wild type we observed an increase in mass up to -240 mV, where 100% of the enzyme was in the reduced form.

Figure 31. Study of the oxidation/reduction potential of GrxA protein. A) Mutant GrxA in C18S/C36A; B) Mutant GrxA in C18S/C36A/C70S; C) GrxA wild type. All proteins were exposed to different potentials and then treated with 2 kDa malPEG to differentiate the oxidized and reduced form of the protein. Samples were run on a non-reducing SDS-PAGE at 100 mV for 1 h 45 min and stained with coomassie brilliant blue stain.

When analyzing the enzyme redox potential by the fluorescent method we found that all three enzymes had a change in fluorescence as the enzyme changed from the oxidized form to the reduced form and the data collected showed a good fit to a single-component

Nernst Equation for a two electron couple (Figure 34), a result that was not expected at least for the triple mutant because there is no Cys available to form a disulfide bridge. Most likely this result could be explained by the capacity of GrxA to form a disulfide bridge with GSH.

The calculated enzyme redox potential for the wild type and mutant enzymes using the florescent method is summarized in Table 6, from where we can observe that the predicted potential for the wild type (-174 mV) is lower than the calculated potential by Kim et al.

(2012) of -240 mV using the malPEG method. These results highlight the observation made by Kim et al. that the mBBr titration method (fluorescent method) is no very reliable when dealing with mixed glutathione disulfide species and therefore we cannot trust the data produced by this method to make predictions about enzyme redox potential.

Table 6. Middle Point Potential for GrxA Wild Type Protein and Two Cystine Mutants

GrxA protein Middle point potential (mV)* Wild type -175 C18S/C36A -156 C18S/C36A/C70S -148

*Values calculated using single-component Nernst Equation for a two-electron couple mechanism.

Mass Spectrometry Analysis

Mass spectrometry analysis of the proteins exposed to oxidizing or reducing environments shows only one significant peak under oxidizing conditions for the GrxA wild type, while under reducing conditions it shows two significant peaks (Figure 35 and 36, respectively). The average mass determination for GrxA in the oxidized form was 11,306.6

Da while the mass for the reduced forms was 10,953.6 Da and 11,209.6 Da. We observed there is a difference of 97 Da between the oxidized and reduced protein. On the other hand, there is a difference of 256 Da between the two peaks observed under the reducing condition that could be attributed to the formation of an unknown adduct. The calculated mass of the

76 protein based on the amino acid sequence including the six histidine residues linked to the protein is 11,031 Da and the difference between the calculated mass and the mass obtained by mass spectrometry experiments reveal that there is a 77.4 Da difference. Figure 37 and 38 show the mass spectra for the double mutant GrxA exposed to the same conditions as the wild type GrxA. In the case of wild type protein there are two peaks with a mass for the oxidized protein of 11,258.8 Da and 11,515.3 Da and a mass for reduced protein of 10,953.6

Da and 11,209.6 Da. The difference between reduced and oxidized forms for each peak is of

305.7 Da, indicating that the oxidized form of the protein has a glutathione molecule bound to Cys15. The difference between the two peaks observed for the oxidized or reduced form is of 256 Da. Finally the results for the triple mutant shown in Figure 39 and 40 are also evidence of the presence of a glutathione molecule bound to the oxidized form of GrxA triple mutant protein.

Discussion

Comparison of the amino acid sequence for GrxA from Synechocystis sp. with amino acid sequences from other glutaredoxins revealed that the sequence for GrxA contains the characteristic CXXC active site motive (Lilling et al. 2008), suggesting that this enzyme is involved in deglutathionylation reactions as well as disulfide reduction reactions.

Cumulative research on glutaredoxins has demonstrated that all Grx enzymes are involved in deglutathionylation of other proteins, but not all Grxs containing the CXXC motive in their active site are involved in protein disulfide oxidation (Lopez-Maury et al. 2009). Data collected for SynGrxA has demonstrated that the enzyme can act as a deglutathionylation reductase with the use of only one Cys in the active site (Li et al. 2003). Additionally a mutagenesis study on the cysteines present in SynGrxA showed that only Cys15 appeared to

Figure 32. Determination of the redox middle point potential of GrxA. Three individual non reducing SDS-polyacrylamide gels run with protein samples for wild, double, and triple mutant protein GrxA treated with 2 kDa malPEG under oxidizing and reducing conditions.

Figure 33. Comparison of the redox stage between the three GrxA proteins studied with or without binding interaction with 2 kDa malPEG polymer. From left to right: first column represents the molecular weight marker; second column is the as purified wild type GrxA protein; third column represents the wild type protein exposed to a reducing environment (DTT) in the absence of malPEG; fourth column represents the wild type protein exposed to a reducing environment and treated with malPEG; fifth column represent the as isolated double Cys GrxA mutant; sixth column represents the double mutant exposed to a reducing environment in the absence of malPEG; seventh column represents the double mutant exposed to a reducing environment and treated with malPEG; eighth column represent the as isolated triple Cys GrxA mutant; ninth column represent the triple mutant exposed to a reducing environment in the absence of malPEG; and the tenth column represent the triple mutant exposed to a reducing environment and treated with malPEG.

Figure 34. Determination of the redox midpoint potential for GrxA using a fluorescent probe. (A) Oxidation/reduction titration for the control wild type GrxA protein. (B) Oxidation/reduction titration for the double mutant GrxA C18S/C36A. (C) Oxidation/reduction titration for the triple mutant GrxA C18S/C36A/C70S. Plots obtained by single-component Nernst Equation for a two-electron couple fit to estimate the midpoint potential for mutant GrxA proteins.

Figure 35. Oxidized GrxA wild type deconvoluted mass spectrometry analysis. Sample treated with GSSG to obtain the oxidized form of the protein (-100 mV)

Figure 36. Reduced GrxA wild type deconvoluted mass spectrometry analysis. Sample treated with DTT to obtain the reduced form of the protein (-330 mV).

Figure 37. Oxidized GrxA double mutant C18S/C36A deconvoluted mass spectrometry analysis. GrxA treated with GSSG to obtain the oxidized form of the protein (-100 mV).

Figure 38. Reduces GrxA double mutant C18S/C36A deconvoluted mass spectrometry analysis. GrxA protein treated with DTT to obtain de reduced form of the protein (-330 mV).

Figure 39. Oxidized GrxA triple mutant C18S/C36A/C70S deconvoluted mass spectrometry analysis. GrxA treated with GSSG to obtain the oxidized form of the protein (-100 mV).

Figure 40. Mass spectrometry analysis for Grx triple mutant C18S/C36A/C70S. GrxA protein treated with DTT to obtain de reduced form of the protein (-330 mV).

86 be essential for enzyme activity, and mutations on any other cystine residues present in

SynGrxA did not alter the catalytic activity of the enzyme in the presence of glutathione, which also supports the hypothesis that SynGrxA acts as a deglutathionylation enzyme (Kim et al. 2012). However, data collected from enzyme redox potential determination showed that the wild type GrxA can form a mixture of proteins interacting with one or more thiol binding malPEG polymers. Mass spectrometry analysis of the different bands generated on non-reducing SDS-PAGE displayed the different molecules: one GrxA with three malPEG polymers in a reducing state, one GrxA with two malPEG polymers around the midpoint potential of the enzyme, and one GrxA with only one malPEG in a oxidized state that could have two other Cys residues forming a disulfide bond. To help elucidate these puzzling results, we proposed to generate two mutants and study their middle point potential by a fluorescent method and malPEG method. As a result, it becomes apparent that there is a discrepancy in the middle point potential determined by the two methods. The fluorescent results suggested that the potential of the wild type and mutant proteins was between -145 mV and -174 mV (Table 6), and the malPEG method showed that the wild type had a midpoint potential of around -230 mV with three different bands corresponding to a mixture of GrxA proteins that contain one, two, or three malPEG polymers. These results are consistent with previous reports by Kim et al. (2012), while for the double mutant there were two bands corresponding to no binding and two malPEG binding to the enzyme. The triple mutant showed two bands, one for no malPEG binding and the other with one malPEG binding. The differences can be attributed to the presence of more than one glutathionylated

GrxA. There is no dimmer formation in any of the GrxA proteins observed, as is confirmed by mass spectrometry and SDS-PAGE with DTT treatment.

The mass spectrometry data collected shows that in all three proteins tested there is a single glutathione molecule interacting with a Cys residue on the protein. Additionally, a mutant GrxA on three of the four Cys residues present in the protein (Cys18, Cys36, and

Cys70) was able to interact with glutathione, indicating that Cys15 is responsible for glutathione binding. This evidence suggests that GrxA can be involved in monothiol reactions of deglutathionylation as proposed by Li et al. (2003); but it does not rule out the possibility that the GrxA protein can form disulfide bonds with any of the Cys present in its structure as suggested by Kim et al. (2012). They have found that the active site of GrxA is present in the reduced dithiol form and not in the oxidized disulphide form. In addition they demonstrated that Cys15 is the only Cys essential for arsenate reductase enzyme activity; evidence that confirms the participation of GrxA in deglutathionylation reactions. The evidence collected does not demonstrate its participation in disulfide reduction, however results obtained from the determination of the middle point potential of GrxA by malPEG, along with mass spectrometry analysis, suggested that there is a possible disulfide bond forming within the enzyme (Kim et al. 2012). All experimental data collected from our research group indicate that GrxA is involved in deglutathionylation reactions, but it remains unclear from the data collected if GrxA can be involved in disulfide reduction reactions. Our data also supports the hypothesis proposed by Li et al. (2003), where they proposed that the

GrxA protein from Synechocystis sp. was involved in a deglutathionylation step during arsenate reduction (Li et al. 2003). To explore the possibility that GrxA can also be involved in disulfide reduction reactions, an additional experiment can be performed where the ribonucleotide reductase enzyme is used to measure enzyme activity. This enzyme requires a dithiol/disulfide reaction in order to transfer the electrons from the hydrogen donor to the

88 enzyme thiols. What we expect to find after performing this enzymatic test, as described by

Bushweller et al. (1992), is that the wild type GrxA capable of forming intra-disulfide bonds will show activity in the assay, but mutants on Cys15 and/or Cys18, Cys36, and Cys70 will have decreased or no activity. If the mutants have no activity, this will clearly indicate that

GrxA is also involved in disulfide reduction reactions, as suggested by the sequence of its active site. To further confirm this hypothesis we can perform an isoelectric focusing (IEF) electrophoresis using GrxA. Examination of the IEF from the alkylated wild type GrxA and its mutants with iodoacetamide, which removes the negative charge due to thiols, will reveal the total number of thiol groups present in each protein and by comparison we could establish if the wild GrxA can form intra-disulfide bonds with any of the other Cys residues

(Bushweller et al. 1992). Results obtained by this method would conclusively confirm the hypothesis and help elucidate the mode of action of GrxA from Synechocystis sp.

CHAPTER FIVE

Conclusions

The use of SELEX technique to develop ssDNA aptamer that can bind to B. anthracis

LF protein proved to be a successful approach to develop ssDNA aptamers. The aptamer

ML12 showed to have a tight binding affinity to its target of 11 ± 2.7 nM and low nonspecific interaction with BSA. Characterization of the binding properties of the aptamer revealed that the full length of the random region was necessary for tight binding. The biding characteristics of ML12 enabled the development of an aptamer-base ELISA assay that was used to visually assess the presence of LF protein. The binding affinity of the aptamer determined by aptamer-based ELISA was lower to the one observed by Cy3-ML12 fluorescent determination. This discrepancy between the two Kd values is most likely a consequence of multiple washing steps involved in the aptamer-based ELISA assay. We also found that the aptamer ML12 had inhibitory effects over the catalytic activity of LF protein with an IC50 value of 15 M ± 1.5. These results suggest that the binding site of the aptamer is located within the active site of LF, but further characterization of their interaction is required to conclusively establish the actual binding site of the aptamer. The evidence collected through this research supports the idea that the ML12 aptamer has the potential to be used in the development of a detecting and therapeutic agent against LF. Development of such probes will require aptamer modification to resist nuclease digestion and improve cell permeability; the study of macrophage IC50 determination to test inhibitory effects in living cells; characterization of clearance time in vivo, and determination of the binding site of the aptamer. The use of whole cell-SELEX to develop dsDNA and ssDNA aptamers that can

90 bind to E. coli cells generated aptamers with binding affinities in the µM range. The dsDNA aptamer isolated by Shipley S. in 2010 proved to have very low specificity. The aptamer could bind with higher affinity to other strains belonging to TC group, which reduces the applicability of this aptamer to develop a detecting assay. The lack of specificity of this aptamer is most likely explained by the unique double strand conformation adopted by this aptamer that dramatically reduces the capability of the aptamer to bind specific targets. The aptamers obtained from ssDNA SELEX after five rounds showed large diversity; from the 12 aptamers sequenced, none of them showed sequence similarities that could indicate enrichment of a particular sequence by SELEX. Characterization of the Kd value of all the aptamers showed they all had very similar binding affinities in the low µM range. From the

12 aptamers studied only three of them showed good specificity for its target when tested against other strains belonging to the TC group. The similarity in the binding affinities and the lack of specificity can be attributed to the low number of SELEX rounds performed. In order to develop a detecting assay for E. coli using the ssDNA aptamers isolated, further characterization of their binding affinity is required.

Studies of the redox potential of the Synechocystis sp wild type GrxA protein and derived mutants on the cys residues of the protein corroborate that the protein is involved in deglutationilation of other proteins, data obtained from mass spectrometry analysis.

Measurement of the redox potential of each mutant and wild type GrxA protein suggests that the mechanism employed by this protein is a monothiol redox mechanism. The experiments performed within this research failed to conclusively demonstrate if the GrxA is capable to be involved in dithiol mechanisms as suggested by the amino acid sequence of the active site; never the less our data aggress with the observation that glutaredoxins with active sites

91 containing two cys residues not always are involved in dithiol redox reactions. To conclusively demonstrate if the GrxA from Synechocystis sp. can be involved in dithiol redox reactions further experiments need to be performed as suggested in the discussion section of chapter

APPENDICES

APPENDIX A

ssDNA SELEX for LF Protein, Rounds1-5

Figure A1. Aptamer generation for LF protein using the ssDNA product from asymmetric PCR. Primers used for amplification were mixed in a 10:1 ratio of forward:reverse primer respectively and the product (60bp long library) was resolved on a 6% PAGE.

Figure A2. Aptamer generation for LF protein. Product from asymmetric PCR using the DNA recovered from round one of selection and resolved on a 6% PAGE. The PCR reaction generated three bands; one belonging to the ssDNA from as indicated by the arrow followed on top by the double strand DNA and a third band of nonspecific amplification.

Figure A3. Aptamer generation for LF protein. Product from asymmetric PCR using as template the DNA recovered from round two of selection and resolved on a 6% PAGE. Four bands are generated by the PCR reaction; on the gel the arrow points to the ssDNA band that was excised and recovered for further SELEX rounds and on top of this band we find the double strand DNA fragment. The two other fragments are nonspecific amplifications.

Figure A4. Aptamer generation for LF protein. Product from asymmetric PCR using as template the product from the fourth selection round and resolved on a 6% PAGE. Confirmation of the location of the ssDNA band was performed by running a sample of ssDNA from the first ssDNA generated for the SELEX procedure.

Figure A5. Aptamer generation for LF protein. Product from asymmetric PCR performed with the DNA generated from the fifth round of selection and resolved on a 6% PAGE. The ssDNA recovered from this round was used to perform the counter SELEX step.

APPENDIX B

ssDNA SELEX for Whole E. coli Cells, Rounds 1-5

Figure B1. PCR product from asymmetric PCR using the recovered ssDNA from round one of cell SELEX. Line 1 contains the eluted fraction from streptavidin bead incubation; line 2 contains the wash after NaOH treatment; line 3 contains the NaOH treatment of dsDNA; line 4 contains the supernatant from streptavidin incubation; line 5 contains the wash from previous PCR reaction as a marker. 6% PAGE stained with EtBr.

Figure B2. PCR product from asymmetric PCR using the recovered ssDNA from round two of counter cell SELEX. Line 1 contains the elute fraction from streptavidin beads; line 2 contains the wash after NaOH treatment from streptavidin beads; line 3 contains ssDNA from denaturation with NaOH; line 4 contains the supernatant from incubation with streptavidin beads. 6% PAGE stained with EtBr.

100

Figure B3. PCR product from round three of cell SELEX. Line 1 contains the wash step of a previous round to use as control; line 2 contains dsDNA recovered from round 2 of selection to use as template for asymmetric PCR; line 3 contains the supernatant after incubation with streptavidin beads; line 4contains DNA recovered from NaOH treatment; line 5 contains the wash after NaOH treatment; line 6 contains the eluted fraction from streptavidin beads.

101

Figure B4. PCR product from round four of cell SELEX. Line 1 contains template PCR product used as control for ssDNA; line 2 contains the supernatant from incubation with streptavidin beads; line 3 contains DNA recovered from NaOH treatment; line 4 contains the wash after NaOH treatment; line 5 contains the eluted fraction from streptavidin beads.

102

APPENDIX C

Mass Spectrometry Analysis of GrxA Protein

E:\GRX\10-24-12\GRX_A_-100_121024175043 10/24/2012 5:50:43 PM

RT: 0.00 - 10.10 SM: 5B 3.78 NL: 100 1.06E10 TIC MS 90 GRX_A_- 100_12102 80 4175043

40 RelativeAbundance 30

20 4.24 10 7.92 2.28 2.49 2.70 3.13 3.31 4.97 5.54 5.69 6.08 6.61 6.82 7.12 7.45 8.53 8.74 8.97 9.17 9.56 9.97 0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0 Time (min)

GRX_A_-100_121024175043 #128-157 RT: 3.64-3.92 AV: 30 NL: 4.31E7 T: FTMS + p ESI Full ms [200.00-2000.00] 808.6817 870.7335 z=14 100 z=13 964.6368 90 z=12

70 707.7228 1028.8655 60 z=16 z=11 1052.2393 50 z=11 40 1131.6513 z=10

RelativeAbundance 666.1512 30 z=17 1257.2783 z=9 20 1285.7356 1414.4378 629.1431 z=9 10 284.1322 427.1535 553.5301 z=8 1508.6035 1616.2169 1695.9449 1883.7154 1968.5832 z=18 z=? z=? z=? z=? z=7 z=? z=? z=? 0 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000 m/z

Figure C1. LCMS GrxA oxidized (-100 mV). Top chromatogram represents the total ion mode output (TIC). Bottom chromatogram represents the mass spectrum obtained for GrxA.

103

E:\GRX\10-24-12\GRX_A_-330 10/24/2012 7:13:10 PM

RT: 0.00 - 10.10 SM: 5B 3.81 NL: 100 9.30E9 TIC MS 90 GRX_A_- 330 80

40 RelativeAbundance 30

10 4.25 2.22 2.35 2.62 2.98 3.31 5.28 5.59 5.92 6.10 6.47 6.81 7.15 7.35 7.60 7.93 8.54 8.84 9.469.33 10.01 0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0 Time (min)

GRX_A_-330 #134-156 RT: 3.72-3.94 AV: 23 NL: 5.40E7 T: FTMS + p ESI Full ms [200.00-2000.00] 917.8702 847.3425 z=12 100 z=13

80 734.4982 1001.2213 z=15 z=11 70 1024.5042 60 z=11 50 1101.2425 688.6551 z=10 40 z=16

RelativeAbundance 1223.4913 30 z=9

20 1251.9481 648.2050 1376.3016 z=17 z=9 10 284.1319 356.0653 427.1204 565.5643 z=8 1467.9242 1572.9167 1713.5139 1883.6447 1997.5647 z=? z=? z=? z=? z=? z=7 z=? z=? z=? 0 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000 m/z

Figure C2. LCMS GrxA reduced (-330 mV). Top chromatogram represents the total ion mode output (TIC). Bottom chromatogram represents the mass spectrum obtained for GrxA.

104

E:\GRX\10-24-12\GRX_18_36_-100 10/24/2012 8:09:47 PM

RT: 0.00 - 10.09 SM: 5B 3.70 NL: 100 7.51E9 TIC MS 90 GRX_18_3 6_-100 80

40 RelativeAbundance 30

10 4.03 2.20 2.41 2.63 3.03 3.23 4.28 4.82 5.28 5.55 5.77 5.99 6.53 6.81 7.01 7.30 7.51 7.93 8.47 8.77 8.92 9.51 9.92 0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0 Time (min)

GRX_18_36_-100 #126-142 RT: 3.64-3.80 AV: 17 NL: 3.81E7 T: FTMS + p ESI Full ms [200.00-2000.00] 805.1848 z=14 867.0448 100 z=13 960.6403 90 z=12

80 704.7253 z=16 70 1024.5058 z=11 60 1047.8799 50 z=11 40 1126.8553 z=10 663.2714 1251.9497 RelativeAbundance 30 z=17 z=9 20 1280.6291 z=9 1408.3183 626.4232 1202.2271 10 284.1319 427.1265 546.0888 z=8 1519.1384 1609.2190 1702.6699 1883.6689 1982.1452 z=18 z=1 z=? z=? z=? z=? z=7 z=? z=? z=? 0 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000 m/z

Figure C3. LCMS GrxA double mutant oxidized (-100 mV). Top chromatogram represents the total ion mode output (TIC). Bottom chromatogram represents the mass spectrum obtained for GrxA mutant.

105

E:\GRX\10-24-12\GRX_18_36_-330 10/24/2012 8:52:31 PM

RT: 0.00 - 10.09 SM: 5B 3.71 NL: 100 7.37E9 TIC MS 90 GRX_18_3 6_-330 80

40 RelativeAbundance 30

10 4.18 2.22 2.38 2.62 3.20 3.31 4.79 5.28 5.56 5.77 6.10 6.566.81 7.087.22 7.47 7.93 8.56 8.69 8.95 9.40 9.55 9.83 0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0 Time (min)

GRX_18_36_-330 #126-148 RT: 3.64-3.86 AV: 23 NL: 3.24E7 T: FTMS + p ESI Full ms [200.00-2000.00] 843.6530 913.7900 z=13 z=12 100

90 731.3005 996.7703 80 z=15 z=11

70 1020.1441 60 z=11 1096.3462 50 685.6574 z=10 z=16 40 1217.9395

RelativeAbundance z=9 30 1246.5081 20 645.2660 z=9 1370.1803 z=17 z=8 10 284.1314 389.1771 472.7337 577.5540 1461.4433 1565.7792 1706.1121 1883.6190 1953.3000 z=? z=? z=? z=? z=? z=7 z=? z=? z=? 0 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000 m/z

Figure C4. LCMS GrxA double mutant reduced (-330 mV). Top chromatogram represents the total ion mode output (TIC). Bottom chromatogram represents the mass spectrum obtained for GrxA mutant.

106

E:\GRX\10-24-12\GRX_18_36_70_-100 10/24/2012 9:24:31 PM

RT: 0.00 - 10.09 SM: 5B 3.69 NL: 100 4.52E9 TIC MS 90 GRX_18_3 6_70_-100 80

40 RelativeAbundance 30

10 3.93 4.16 4.28 2.25 2.47 2.65 3.302.95 4.83 5.28 5.55 5.76 5.95 6.32 6.83 7.26 7.43 7.96 8.38 8.72 9.02 9.31 9.63 9.93 0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0 Time (min)

GRX_18_36_70_-100 #124-141 RT: 3.62-3.79 AV: 18 NL: 1.85E7 T: FTMS + p ESI Full ms [200.00-2000.00] 767.6461 z=15 100 885.5902 z=13 90 959.2223 z=12 80 719.6693 z=16 70

60 1046.4235 z=11 50

40 677.4537 1150.9652 z=17 z=10 1278.7383 RelativeAbundance 30 z=9

20 1406.3144 639.8176 1212.6047 10 284.1310 391.2846 543.7421 z=8 1517.2678 1607.2210 1781.1838 1883.6219 1981.0364 z=18 z=1 z=? z=? z=? z=? z=7 z=? z=? z=? 0 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000 m/z

Figure C5. LCMS GrxA triple mutant (-100 mV). Top chromatogram represents the total ion mode output (TIC). Bottom chromatogram represents the mass spectrum obtained for GrxA mutant.

107

E:\GRX\10-24-12\GRX_18_36_70_-330 10/24/2012 10:07:12 PM

RT: 0.00 - 10.10 SM: 5B 3.70 NL: 100 6.03E9 TIC MS 90 GRX_18_3 6_70_-330 80

40 RelativeAbundance 30

20 3.93 10 4.17 2.23 2.44 2.64 2.91 3.31 4.82 5.27 5.57 5.90 6.05 6.66 6.82 6.99 7.31 7.49 7.96 8.65 8.78 9.05 9.41 9.70 9.98 0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0 Time (min)

GRX_18_36_70_-330 #126-144 RT: 3.64-3.82 AV: 19 NL: 2.56E7 T: FTMS + p ESI Full ms [200.00-2000.00] 933.8000 862.1239 782.2508 z=12 100 z=13 z=14 90

80 747.3084 z=15 1018.5995 70 z=11

60 1042.4047 z=? 50 700.6026 z=16 1120.3583 40 z=10 1244.8429 RelativeAbundance 30 z=9

20 659.4496 z=17 1368.0567 10 284.1307 356.0998 470.9488 553.5266 z=8 1493.5939 1563.3493 1643.1909 1765.6787 1883.7838 1951.2024 z=? z=? z=? z=? z=? z=7 z=? z=? z=? z=? 0 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000 m/z

Figure C6. LCMS GrxA triple mutant (-330 mV). Top chromatogram represents the total ion mode output (TIC). Bottom chromatogram represents the mass spectrum obtained for GrxA.

108

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