Regulation of Virulence Gene Transcripts by the Francisella Orphan Response

Regulator PmrA: Role of Phosphorylation and Evidence of MglA/ SspA

Interaction

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of

Philosophy in the Graduate School of The Ohio State University

By

Brian Len Bell, B.S.

Integrated Biomedical Science

The Ohio State University

2009

Dissertation Committee:

John Gunn, Ph.D., Advisor

Chad Rappleye, Ph.D.

Larry Schlesinger, M.D.

Mark Wewers, M.D.

ABSTRACT

Francisella tularensis subspecies tularensis is the etiologic agent of tularemia and has been designated a category A biothreat agent by the CDC.

Tularemia is characterized by replication and dissemination within host phagocytes. Intramacrophage growth is dependent upon the regulation of

Francisella Pathogenicity Island (FPI) virulence genes, which is poorly understood. Two-component regulatory systems (TCS) are widely employed by

Gram negative bacteria to monitor and respond to environmental signals.

Virulent strains of F. tularensis are devoid of classical, tandemnly arranged TCS, but orphaned members, such as the response regulator PmrA, have been identified. In the F. novicida model system, previous work has shown that a pmrA mutant is deficient for intramacrophage growth and is avirulent in the mouse model. Here we determine that phosphorylation aids PmrA binding to regulated promoters pmrA and the FPI encoded pdpD, and KdpD is the histidine kinase primarily responsible for phosphorylation of PmrA at the aspartic acid at position 51 (D51). A strain expressing PmrA D51A retains some DNA binding but exhibits reduced expression of the PmrA-regulon, is deficient for intramacrophage replication and is attenuated in the mouse model. PmrA co- precipitates with the FPI transcription factors MglA and SspA, which bind RNA ii polymerase. Together this data suggests a model of Francisella gene regulation that includes a TCS consisting of KdpD and PmrA. Once phosphorylated, PmrA binds to regulated gene promoters recruiting free or RNA polymerase bound

MglA and SspA to initiate FPI gene transcription.

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DEDICATION

To Erin, whose belief and support have made this possible.

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ACKNOWLEDGEMENTS

I have been blessed with many excellent mentors: Janet Staab started my career as a bench scientist. Dave Brown taught me to see the big picture.

Richard Warren gave me an important nudge into graduate school. Mick Arthur and Tom Dreier showed me how to do the job well. John Gunn helped me become a real student of science and patiently corrected my scientific errors.

I have shared laboratories with many talented scientists. The long hours, piles of disappointing data, and the struggle to find meaning were all made bearable by experiencing them together. I hope you have all enjoyed working together as much as I have. This work has been positively affected by every member of the Gunn lab and the Center for Microbial Interface Biology.

I would like to thank Denise Monack of Stanford University for generously providing the F. novicida His-SspA strain and Daniel Wozniak for supplying purified CheA.

This work was sponsored by the NIH/NIAID Regional Center of Excellence for Bio-defense and Emerging Infectious Diseases Research (RCE) Program and by the CMIB fellowship through NIH Grant T32AI065411.

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VITA

May 12, 1971 ...... Born – Columbus, Ohio

1995 ...... B.S. Zoology, The Ohio State University

1995 – 1998...... Research Associate Department of Microbiology The Ohio State University Columbus, Ohio

1998 – 2000...... Project Scientist Life Sciences Test Facility Dugway Proving Ground Dugway, Utah

2000 – 2006...... Principal Research Scientist Biotechnology Battelle Memorial Institute Columbus, Ohio

2006-2008 ...... Graduate Research Associate Molecular Virology, Immunology and Medical Genetics The Ohio State University Medical Center Columbus Ohio

2008 – present ...... Graduate Research Fellow Center for Microbial Interface Biology The Ohio State University Medical Center Columbus, Ohio

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PUBLICATIONS

Identification of an orphan response regulator required for the virulence of Francisella spp. and transcription of pathogenicity island genes. Nrusingh P. Mohapatra, Shilpa Soni, Brian L. Bell, Richard Warren, Robert K. Ernst, Artur Muszynski, Russell W. Carlson, John S. Gunn. Infection and Immunity, July 2007;75(7):3305-14

Final Report for the Joint Vaccine Acquisition Program (JVAP) for the Francisella tularensis Vaccine Project. Brian L. Bell, Robert M. Miceli, Lisa M. Moore, Sheri L. Schanaman, Katie Liljestand, Jennifer Thermos, Jeff Varelman, Stephen Parker, John D. Wright, Richard Warren, and Bruce Harper. West Desert Test Center, Life Sciences Division, U.S. Army Dugway Proving Ground. October 1999.

Effects of Decontamination Solutions on Traditional and Molecular Detection Methodologies: TaqMan LightCycler PCR, ORIGEN Electrochemiluminescent, Hand-Held (HHA) and Colony-Forming Unit (cfu) Assays. Sheri L. Schanaman, Jeff Varelman, Brian L. Bell, Lloyd Larsen, Bruce G. Harper, David H. Brown, Michael Glass, Robert M. Miceli. West Desert Test Center, Life Sciences Division, U.S. Army Dugway Proving Ground. May 1999.

Biological Weapons Convention Treaty Reference Guide. Daniel D. Martin, Brian L. Bell, and Stephen J. Parker. West Desert Test Center; Life Sciences Division; U.S. Army Dugway Proving Ground. February 1999.

Biological Sampling Protocols for Use During On-Site Visits (1992 Biological Weapons Trilateral Agreement). Daniel D. Martin, Brian L. Bell, and Lester J. Richoux. West Desert Test Center; Life Sciences Division; U.S. Army Dugway Proving Ground. December 1998.

Molecular Probe for Typing Strains of Candida albicans. Pamela Postlethwait, Brian L. Bell, W. Todd Oberle, and Paula Sundstrom. Journal of Clinical Microbiology, Feb. 1996, p. 474-476.

FIELDS OF STUDY

Major Field: Integrated Biomedical Science

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TABLE OF CONTENTS

ABSTRACT ...... ii

DEDICATION ...... iv

ACKNOWLEDGEMENTS ...... v

VITA ...... vi

LIST OF TABLES ...... xi

LIST OF FIGURES ...... xii

INTRODUCTION ...... 1

1.1. Nomenclature ...... 2

1.2. Ecology ...... 4

1.3. Disease ...... 5

1.4. Diagnosis ...... 6

1.5. Treatment ...... 7

1.6. Animal models of tularemia ...... 8

1.7. Epidemiology ...... 9

1.8. Laboratory safety ...... 10

1.9. Biological Warfare ...... 11

1.10. Vaccines ...... 12

1.11. Pathogenesis ...... 14

1.12. Virulence Factors ...... 18

1.13. The Francisella Pathogenicity Island (FPI) ...... 19

1.14. Regulation of the FPI ...... 24 viii

1.15. Other Francisella Virulence Factors ...... 26

1.16. Two Component Regulatory Systems and Francisella ...... 27

PmrA Regulates Francisella Virulence Factors by an Independent Pathway from other Virulence Determinants ...... 32

2.1. Introduction ...... 32

2.2. Results ...... 35

2.3. Discussion ...... 38

2.4. Materials and Methods ...... 66

The Francisella Orphan Response Regulator PmrA Binds to Regulated Gene Promoters and Binding is Mediated by Phosphorylation...... 68

3.1. Introduction ...... 68

3.2. PmrA binds to the promoter regions of regulated genes and binding is increased upon treatment with a phosphate donor...... 70

3.3. MglA does not bind to the pmrA or pdpD promoters and does not interact with DNA bound PmrA...... 73

3.4. PmrA may bind to multiple sites in regulated gene promoters...... 73

3.5. Discussion ...... 75

3.5. Materials and Methods ...... 86

PmrA Aspartate 51 is the Site of Phosphorylation and is Essential for Francisella Intramacrophage Replication and Virulence...... 89

4.1. Introduction ...... 89

4.2. PmrA is phosphorylated at aspartate 51...... 91

4.3. PmrA D51 is important for gene regulation...... 93

4.4. PmrA D51 is required for intramacrophage growth...... 94

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4.5. PmrA D51 is required for mouse virulence...... 95

4.6. PmrA is a primary target of the histidine kinase KdpD...... 96

4.7. Discussion ...... 98

4.8. Materials and Methods ...... 108

PmrA Interacts with other Francisella Transcription Factors ...... 112

5.1. Introduction ...... 112

5.2. PmrA co-precipitates with MglA and SspA...... 114

5.3. MglA co-precipitates with PmrA from a F. tularensis Schu4 lysate...... 116

5.4. Discussion ...... 117

5.5. Materials and Methods ...... 122

DISCUSSION ...... 124

BIBLIOGRAPHY ...... 141

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LIST OF TABLES

Table 2.1: Differentially regulated genes when comparing F. novicida ∆pmrA to F. novicida wild-type bacteria by microarray...... 45

Table 2.2. Confirmation of differential expression by qRT-PCR...... 49

Table 2.4: Differentially regulated genes (1.5 fold or greater) when comparing F. novicida ∆kdpD to F. novicida wild-type bacteria by microarray...... 55

Table 2.5: Differentially regulated genes (1.5 fold or greater) when comparing F. novicida ∆qseC to F. novicida wild-type bacteria by microarray...... 62

Table 2.6: Differentially regulated genes (1.5 fold or greater) when comparing F. novicida ∆FTN_1355 to F. novicida wild-type bacteria by microarray...... 63

Table 6.1. Phenotypic characterization of various Francisella pmrA mutants. . 140

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LIST OF FIGURES

Figure 1.1. The Francisella Pathogenicity Island (FPI) ...... 31

Figure 2.1. RT-PCR analysis of pmrA and iglC in F. novicida mutant strains .... 42

Figure 2.2. Venn diagram of the F. novicida ∆pmrA, ∆mglA and ∆kdpD regulons as determined by microarray analysis ...... 43

Figure 2.3. Venn diagram of the F. novicida ∆FTN_1355, ∆qseC and ∆kdpD regulons as determined by microarray analysis...... 44

Figure 3.1. Primer extension results for pmrA (FTN_1465) ...... 77

Figure 3.2. Primer extension results for pdpD (FTN_1325) ...... 79

Figure 3.3. EMSA: PmrA Binding ...... 81

Figure 3.4. EMSA: PmrA Phosphorylation ...... 82

Figure 3.5. EMSA of His-PmrA D51A binding to pmrA ...... 83

Figure 3.6. His-PmrA D46A and His-PmrA D46G retain the ability to bind DNA promoters and that binding is increased with acetyl phosphate treatment ...... 84

Figure 3.7. His-PmrA binding to regulated gene promoters ...... 85

Figure 4.1. Phosphotransfer from CheA ...... 101

Figure 4. 2. Phosphotransfer from membrane fractions ...... 102

Figure 4.3. Phosphotransfer from membrane fractions ...... 103

Figure 4.4. RT-PCR analysis of PmrA-regulated genes in F. novicida ∆pmrA complemented with PmrA or PmrA D51A ...... 104

Figure 4.5. PmrA D51A is important for intramacrophage replication...... 105

Figure 4.6. PmrA D51A is important for mouse virulence ...... 106

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Figure 4.7. Phosphotransfer control reaction ...... 107

Figure 5.1. Co-precipitation. PmrA, MglA and SspA co-precipitate...... 120

Figure 5.2. Co-precipitation. MglA precipitates with PmrA from a F. tularensis Schu 4 soluble fraction...... 121

Figure 6.1. Model of PmrA Regulation of the FPI ...... 141

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CHAPTER 1

INTRODUCTION

Bacterium tularense was first isolated in Tulare County, California in 1911 from a squirrel that had succumbed to a plague like disease (Chapin, 1921). The first human cases were documented in Ohio in 1925 with an outbreak involving six patients that contracted ocular infections (Nigrovic and Wingerter, 2008). In

1928, Dr. Edward Francis published a report of over 600 human cases of tularemia (Francis, 1983). This landmark article described the forms of the disease and identified rabbits, rodents and blood feeding insects as transmission vectors. In honor of Dr. Francis’s contributions to the study of tularemia and to acknowledge that this bacteria deserved its own genus, the bacteria was renamed Francisella tularensis in 1947 (Olsufiev et al., 1959). Tularemia has been known as rabbit fever, lemming fever and deertick fever. At risk populations have included hunters, trappers, cooks, landscapers, farmers, veterinarians, meat handlers and laboratory workers (Nigrovic and Wingerter,

2008). Further study has characterized F. tularensis as a Gram negative facultative intracellular pathogen that is capable of manipulating the

1 host innate immune system to cause disease. Today tularemia is a relatively rare disease but the possibility of weaponization has renewed interest into the molecular mechanisms of its virulence (Ellis et al., 2002).

1.1. Nomenclature

Two species of Francisella are currently recognized, Francisella tularensis and Francisella philomiragia. Four subspecies of Francisella tularensis have been identified: tularensis, holarctica, mediasiatica, and novicida. Francisella tularensis subspecies tularensis (F. tularensis) has been designated as group type A which are the most virulent strains and have been recovered only in North

America. Type B strains, comprised of Francisella tularensis subspecies holarctica (F. holarctica), are less virulent and are found in North America and

Europe. Francisella tularensis subspecies novicida (F. novicida) is also markedly less virulent than F. tularensis and has been isolated primarily in North America.

F. novicida is closely related to the F. tularensis strains; sharing greater than 96 percent DNA homology, but does not cause disease in immunocompetent humans (McLendon et al., 2006). With the exception of a recent discovery of F. novicida in Australia, francisellae have not been found in the Southern hemisphere. However, little is known about the geographical distribution of F. philomiragia, F. tularensis subspecies mediasiatica or F. novicida, in part due to the fact that they rarely cause disease in man. Francisella mediasiatica is not a

2 clinically relavent strain but has been found in Central Asia (Keim et al., 2007).

F. philomiragia is not a pathogen in mammals but has been described as the causative agent of disease in fish (Ottem et al., 2009;Ottem et al., 2007).

Based on 16S ribosomal DNA (rDNA) sequencing, francisellae are most closely related to proteobacteria and do not share similarity with any other characterized genera (Forsman et al., 1994). No other genus of bacteria currently belong to the francisellae family, but some soil bacteria and fish pathogens have been suggested as members (Forsman et al., 1994;Ottem et al.,

2009). The slow evolving rRNA typing provides some relationship to other families of bacteria; however it is not sufficient to differentiate the evolutionary differences between the closely related subspecies of F. tularensis.

Whole genome sequencing identification of single nucleotide polymorphisms (SNPs) has generated data that differentiate members of the

Francisella family. F. philomiragia is distantly related to the human disease- causing members. F. novicida is distinct from the other subspecies of F. tularensis and is the ancestral progenitor from which the remaining three subspecies evolved. The more virulent F. tularensis strains are more diverse than the F. holarctica strains (Keim et al., 2007).

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1.2. Ecology

Francisella species have been isolated from a very broad host range. F. tularensis subspecies have been isolated from over 100 species of mammals; especially from rodents and lagomorphs. Interestingly, F. tularensis has a more terrestrial distribution; found in rabbits, squirrels and ticks, while F. holarctica is more aquatic being recovered from muskrats, beavers and mosquitoe larvae

(Morner, 1992). In addition to mosquitoes, ticks have been identified as vectors.

F. tularensis has been shown to be very hardy in aquatic environments with documented survival in water and mud for as long as four months (Feldman et al., 2001). One of the reasons for this may be the ability of F. tularensis to survive and replicate within amoeba (Lauriano et al., 2004;Abd et al.,

2003;Lauriano et al., 2004). Residence within amoeba may provide a protective environmental niche in which the bacteria can exist. A reservoir of F. tularensis has yet to be discovered. Risk of contracting tularemia from domesticated animals has been highlighted in the literature with transmission being documented from, cats and captive rodents (Avashia et al., 2004;Meinkoth et al.,

2004;Capellan and Fong, 1993;Cooper and Ewell, 1973;Tarnvik and Chu, 2007).

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1.3. Disease

The incubation period for F. tularensis is typically 3-10 days with acute, rapid onset of increasing fever with chills, body aches and headache (Tarnvik and Chu, 2007). Several forms of tularemia can occur and are related to the route and dose of infection. Subcutaneous inoculation by the bite of an infected arthropod or exposure of a break in the skin to infected meat or water results in the ulceroglandular tularemia. Ulceroglandular tularemia accounts for approximately 75% of the reported cases. This form is characterized by an ulcer which forms at the site of infection. Draining lymph nodes can also become infected leading to lymphadenopathy. Glandular tularemia results when lymphadenopathy is not accompanied by a subcutaneous ulcer. The consumption of improperly prepared contaminated meat and/or water can result in the oropharnygeal and/or gastrointestinal form of the disease. Oropharyngeal tularemia is quite rare and is characterized by pharyngitis and excessive regional neck lymphanapathy. Symptoms of gastrointestinal tularemia include diarrhea and vomiting. The first human cases described were of the ocularglandular form which resulted from infection via the eye conjunctiva. Infection via inhalation of aerosolized viable F. tularensis results in the pneumonic form of the disease, which is the most serious. It should be noted that pneumonia can be a complication of systemic spread of the bacteria resulting from any form of tularemia. Any form of the disease can progress to systematic tularemia but gastrointestinal and pneumonic forms have a higher incidence. Progression to 5 systemic disease is related to the infectious dose of viable bacteria delivered at infection. Systemic tularemia results from the infection spreading from the regional lymphatics to other organs including the liver and speen. These sites have a new population of host phagocytes that the bacteria can infect. The bacteria essentially travel throughout the host in macrophages protected from immune challenge to new sites of infection. This behavior has prompted some to describe F. tularensis as a stealth pathogen and refer to the macrophage as a

Trojan horse (Sjostedt, 2006). Systemic or typhoidal forms of the disease can be extremely serious with untreated mortality rates being reported as high as 60 percent. There are a variety of complications associated with tularemia including septicemia, meningitis, endocarditis, hepatitis and renal failure. Dissemination of the bacteria may occur to the point where the patient may experience disseminated intravascular coagulation or acute respiratory distress. Cause of death related to tularemia may be the result of septic shock (Tarnvik and Chu,

2007).

1.4. Diagnosis

Dr. Edward Francis, the namesake of F. tularensis wrote in 1925,

―Knowledge of at least the name of a disease must precede its diagnosis; a physician cannot be expected to diagnose a disease of which he has never heard.‖ Even today this is an appropriate statement. The most important step in

6 the diagnosis of tularemia is recognition that the symptoms could be caused by

F. tularensis exposure. In 1994, tularemia was removed from the list of reportable diseases but was reinstated in 2000 in part to raise awareness of the possibility of infection. Definitive diagnosis of tularemia is primarily made be identifying anti-F. tularensis antibodies in circulation or from culturing the organism from wounds, lymphnode exudates or blood. F. tularensis is rarely detected by Gram staining. The signs and symptoms of tularemia can mimic those of other diseases, complicating diagnosis. It may take up to two weeks for measurable antibodies to be present, delaying a positive diagnosis (Feldman,

2003). Other diagnostic tests using fluorescent antibody detection or polymerase chain reaction have been developed and are in use in public health laboratories throughout the United States.

1.5. Treatment

Treatment of F. tularensis is very effective and results in total recovery of more than 97% of cases. Streptomycin has been very effective in combating serious F. tularensis infections greatly reducing the mortality rate since its introduction in the 1940s. Due to complications with handling and treating with streptomycin, today most cases are treated with other aminoglycosides like gentamicin (Tarnvik and Chu, 2007). Serious systemic cases of tularemia can require extensive supportive care due to multi-organ failure. Systemic tularemia

7 is life threatening and survivors face a long and difficult recovery but may not ever completely regain full health (Sandrock, 2008;Sunderrajan et al.,

1985;Dennis et al., 2001).

1.6. Animal models of tularemia

Many models of tularemia infection have been investigated for their suitability to determine the effectiveness of treatment and/ or prophylaxis.

Humans, non-human primates, rodents and lagomorphs have all been used as model systems. Non-human primates respond similarly to vaccines as humans, but are more susceptible to Type B strains than Type A. That combined with high cost, will limit their use for investigation of tularemia. Rabbits are very susceptible to the Schu4 strain of F. tularensis but there is limited information about response to vaccination. Tools for evaluating rabbit immunological response are also lacking. Guinea pigs are highly susceptible to F. tularensis challenge but little information exists regarding vaccination. Rats respond to vaccination in a similar manner as humans but are more resistant to both Type A and B strains of F. tularensis. F. novicida causes a tularemia-like disease in mice. Accordingly, mice have been used extensively to determine the virulence of native and genetically manipulated strains of F. novicida as this strain is more amenable to genetic manipulation. Tools for evaluating immunological responses are widely available for mice and genetically altered mouse strains

8 also exist. In general, mice are more susceptible than humans to F. tularensis and mice response differently to vaccination than humans (Rick Lyons and Wu,

2007). For obvious reasons, the data collected for human infection with F. tularensis is as complete as it will get. To date, no ideal animal model has been identified that responds similarly to humans with respect to susceptibility to F. tularensis strains and outcome of vaccination.

1.7. Epidemiology

Cases of tularemia have dramatically declined since the 1940’s being attributed to the broad use of antibiotics and personal protective measures. The febrile nature of the symptoms of tularemia and their relatedness to other non-life threatening infections probably results in an underreporting of disease. Due to the decline in cases, tularemia was removed from the list of reportable diseases in 1994; however, it was reinstated in 2000 in response to fears of its use as a biological weapon (Feldman, 2003).

There have been a two tularemia outbreaks of note on the island of

Martha’s Vineyard. These outbreaks demonstrate that F. tularensis is endemic to this area. The first outbreak was in 1978 involving 7 cases of primary pneumonic tularemia. In 2000, there were 11 cases of tularemia, 5 of which were the pneumonic form. The pneumonic cases pointed to an aerosol source of infection and a higher risk of infection was established for anyone working with

9 lawnmowers and brush cutters. Further investigation by the CDC confirmed that it is possible to generate an aerosol of infectious F. tularensis by mowing over infected material (e.g. contaminated carcasses or feces). A small sampling of the local wildlife trapped two rodents were that were seropositive for antibodies against F. tularensis. These data established the possibility of natural occurring infections without coming in direct contact with infected flesh or being bitten by an infected arthropod (Feldman et al., 2001).

1.8. Laboratory safety

F. tularensis is one of the most infectious agents known with an infectious dose (ID) below 10 colony forming units (CFU) for humans via the subcutaneous or aerosol routes of infection. (Saslaw et al., 1961). Due to the extremely low infectious dose of F. tularensis and its hardiness in diverse environments, research is performed under the extremely controlled laboratory conditions of

Biosafetly level 3 (BSL3). Organisms that are assigned for use under BSL3 conditions typically have a risk of aerosol exposure and the resulting disease may have serious or life threatening consequences

(http://www.cdc.gov/od/ohs/biosfty/bmbl5/bmbl5toc.htm). Even under these controlled laboratory conditions, the rate of laboratory acquired tularemia infections has been documented to be high compared to other bacterial and viral infectious agents manipulated in the laboratory (Pike et al., 1965).

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1.9. Biological Warfare

F. tularensis was extensively researched and prepared as a biological weapon by at least two state sponsored programs, the United States and the former Soviet Union. The extremely low infectious dose and the relative hardiness of the bacteria made it a suitable candidate for weaponization. Testing and evaluation of F. tularensis preparations proved its effectiveness as a tactical weapon. In fact, by 1955 F. tularensis was being produced at the Pine Bluff

Arsenal in Pine Bluff, Arkansas. The virulent bacteria preparations were loaded into anti-personnel munitions on a rotating basis to keep it fresh and ready for deployment. The signing of the Unilateral Biological Weapons Convention by

President Nixon in 1972 signaled the end of the United States offensive weapons program and eventually resulted in the decommissing of the Pine Bluff Arsenal.

Today we know that while the former Soviet Union signed similar treaties with the

United States, their biological weapons programs, known as BioPreparat continued into the 1990’s. A former director of BioPreparat and defectee from the former Soviet Union claims that tularemia was employed as a biological weapon in World War II. The same scientist has warned that genetic modifications were made to F. tularensis to increase its lethality and/or decrease effectiveness of treatment or prophylaxis (1999).

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1.10. Vaccines

A live attenuated vaccine was developed and tested in the United State in the 1950’s. An F. tularensis Type B strain was passaged in mice resulting in a human attenuated strain that regained virulence in mice. This strain was designated the F. tularensis subspecies holarctica Live Vaccine Strain or Ft LVS.

Early work on this strain revealed the presence of two distinct agar colony morphologies (Eigelsbach et al., 1951;Eigelsbach et al., 1952). To this day, neither the nature nor the cause of the variation is known (Hartley et al., 2006a).

Oblique light can reveal the variants which have become known as blue and gray colonies. The blue colonies have a light blue hazy ring around them while the gray colonies appear dull and granular (Eigelsbach et al., 1951). Isolation of the blue and gray colonies led to the discovery that the blue colonies are immunogenic while the gray are not (Eigelsbach et al., 1952). The mechanism of this variation remains unclear and has been a substantial road block in the FDA approval process for this vaccine. The growth conditions for F. tularensis have been optimized to minimize the generation of the gray phenotype but all vaccine preparations contain a population of grays (Eigelsbach et al., 1951).

The efficacy of the Ft LVS vaccine has been extensively tested. The vaccine strain was derived from a strain of F. tularensis subspecies holarctica originally cultured at the Gamaleia Institute in the former Soviet Union. The strain was transferred to USAMRIID and virulence was decrease by lyophilization and five serial passages in mice (Eigelsbach and Downs, 1961). The LVS 12 vaccine has proven to be affective against subcutaneous infection of virulent F. tularensis but not as affective against aerosol challenge (Saslaw et al., 1961).

LVS was tested as a vaccine against challenge with virulent F. tularensis in human volunteers in 1956. Vaccincation with LVS proved to limit subcutaneous and pneumonic disease. Only two of the nineteen volunteers that were intradermally vaccinated with Ft LVS exhibited signs of disease after subcutaneous challenge with approximately 10 CFU of F. tularensis Schu4. This study also showed that disease was initiated by subcutaneous inoculation of 11 of 12 non-vaccinated volunteers with the same dose of Schu4 (Saslaw et al.,

1961). Intradermal vaccination with LVS resulted in protection of 8 or 10 volunteers to be protected from aerosol challenge with 10-51 CFU of Schu4.

Approximately 10 CFU of F. tularensis Schu4 was confirmed to cause disease in non-vaccinated individuals when delivered via aerosol (Saslaw et al., 1961). The efficacy of the LVS vaccine is not 100% but it does provide protection and has been administered to thousands of at risk laboratory and military personnel. The

1956 study also determined that a killed whole bacteria vaccine was less effective than the Ft LVS vaccine (Saslaw et al., 1961). In those cases in which vaccinated volunteers developed disease, the symptoms were less severe and recovery was more rapid. To date, a more effective vaccine has not been submitted for approval by the FDA and the use of the LVS vaccine is very limited.

FDA approval of the current LVS vaccine is unlikely due to the presence of non- immunogenic gray phenotypes and the fact that the mechanism of LVS

13 attenuation has not been discovered (Wayne Conlan and Oyston, 2007;Hartley et al., 2006a). The need for an effective, safe vaccine for the protection of laboratory and other at risk populations is very high.

Proteinaceous subunit vaccines have not been effective at providing protection against F. tularensis infection. The O-antigen of F. tularensis LPS provides very limited protection, and is not as effective as LVS at preventing disease (Thomas et al., 2007). Live attenuated vaccines have been extensively researched. There are several examples of attenuated strains of F. novicida providing homologous protection but unable to protect against fully virulent strains (Mohapatra et al., 2007b;Lauriano et al., 2004). Additionally, as genetic tools have improved attenuated F. tularensis strains have been used to vaccinate laboratory animals but have failed to provide substantial protection against F. tularensis Schu4 challenge.

1.11. Pathogenesis

A tularemia infection is characterized by replication of the bacteria within host phagocytes. All subspecies of F. tularensis have been shown to replicate within macrophages. F. tularensis replicates within human, mice, rats, guinea pigs and rabbit macrophages. Macrophages are thought to be the dominant reservoir of bacteria during a tularemia infection (Santic et al., 2006;Oyston,

2008). F. tularensis bacilli encounter macrophages at the source of infection and 14 are phagocytosed by a unique pseudoloop mechanism. A voluminous loop of the host cell membrane encircles the invading F. tularensis bacteria. Once the bacteria are enclosed in the spacious vacuole of host membrane the pseudoloop quickly collapses to just contain the invading bacteria (Clemens et al., 2005).

The combinations of host cell receptors that are involved in the process are not completely described. Studies have shown that complement receptor 3 (CR3) and the Fcγ receptor are involved in the uptake of F. novicida (Clay et al., 2008).

The presence of complement increases the phagocytosis of F. novicida but does not appear to be required for uptake. Virulent strains of F. tularensis are resistant to lysis by complement. Attenuated strains that do not elicit an immune response are much more susceptible to complement mediate lysis (Clay et al.,

2008). In the absence of serum opsonination, the mannose receptor becomes more involved in phagocytosis. Additionally, opsonization by surfactant protein A

(SP-A) has been shown to increase phagocytosis of F. novicida (Balagopal et al.,

2006).

The LPS of Francisella does not activate TLR4 like LPS from other Gram negative bacteria. Neither whole F. tularensis bacteria nor its purified LPS activate TLR2 or TLR4. This undetected entry of the bacteria is very important for the outcome of infection because F. tularensis species are unable to replicate within activated macrophages (Elkins et al., 2003). Once F. tularensis escapes to the cytosol it can be recognized by nod-like receptors (NOD) triggering activation of the inflammasome (Gavrilin et al., 2006;Parsa et al., 2006;Henry

15 and Monack, 2007). Involvement of the inflammasome complex results in the initiation of the apoptotic pathway. It is unclear whether apoptosis furthers infection by facilitating escape from the dying cell and allowing infection of neighboring phagocytes or if apoptosis helps control the infection.

Intracellular pathogens affect the formation of the phagosome in which they are taken up and reside (Duclos and Desjardins, 2000). The Francisella containing vacuole (FCV) is uniquely altered acquiring a different subset of phagosomal markers than phagosomes containing other intracellular pathogens.

The FCV acquires the early endosomal antigen 1 and late endosomal markers

(LAMP1 and LAMP2) indicating maturation to a late endosome. It has been demonstrated that treatment to prevent acidification of the vacuole blocks

Francisella replication suggesting that recruitment of an ATPase proton pump is required for intramacrophage growth (Fortier et al., 1995). Departing from the default endosomal-lysosomal maturation process, the FCV does not acquire cathepsin-D. Francisella escape from the FCV has been measured in a couple of ways; first the ability of an anti-Francisella antibody to stain bacteria within plasma membrane selectively permeabilized infected cells provides an indication of when the phagosomal membrane is compromised. This method estimates phagosomal escape to be within 1 hour of phagocytosis. Electron microscopy studies estimating the extent to which F. tularensis has escaped from the phagosomal compartment have determined that the bacteria escape approximately 3-4 hours post infection. Combined these data indicate

16 degradation of the phagosomal membrane followed by bacterial escape to the cytosol within a few hours of phagocytosis (Clemens et al., 2005;Chong et al.,

2008;Santic et al., 2005). Once in the cytosol, the bacteria are able to replicate rapidly and increase by 100-fold or more within 12-24 hours. Late in the infection of macrophages, F. tularensis is taken up within autophagosomal membranes.

This capture may be a last ditch effort by the host cell to control the infection before the bacteria has a chance to escape and infect other cells; however, containment in autophagosomes may provide the bacteria with access to the endocytic pathway and promote cellular exit by exocytosis (Checroun et al.,

2006). Infection with F. tularensis has been shown to affect host expression of genes responsible for mediating autophagy (Butchar et al., 2008). The F. tularensis gene ripA has also been identified that is involved in Ft LVS’s ability to interact with the autophagocytic pathway (Fuller et al., 2008). Strains missing ripA are deficient for replication within macrophages and attenuated in the mouse. The ability of F. tularensis to modify the innate host defense mechanisms, typically those of phagocytic cells, is critical for the bacteria’s ability to cause disease.

The role of polymorphonuclear leucocytes (neutrophils) in F. tularensis infection is understudied. The evidence that has been presented suggests that binding and subsequent phagocytosis of F. tularensis by neutrophils is dependent upon complement (McLendon et al., 2006). Opsonization of bacteria with immune serum results in more phagocytosis than by non-immune serum.

17

Attenuated strains of Ft LVS and F. novicida are more susceptible to neutrophil killing than F. tularensis (Lofgren et al., 1983). The role of neutrophils in controlling F. tularensis infection is unclear. Depletion of neutrophils with monoclonal antibodies has been shown to increase the lethality of F. tularensis

LVS infection in mice when infected subcutaneously or intraperitoneally (Sjostedt et al., 1994). However, neutropenia does not affect disease progression in aerosol infection (Conlan et al., 2002). Taken together, the data suggest that neutrophils may not be able to control pneumonic tularemia (McLendon et al.,

2006).

1.12. Virulence Factors

Genomic Islands (GIs) are regions of genetic material that have been acquired by lateral or horizontal gene transfer. GIs have G+C contents that are significantly different than the rest of the genome. This indicates that the region did not evolve with the rest of the genome. GIs are typically flanked by transposable elements (e.g. transposons, direct repeats, insertion elements) allowing the region of DNA to be mobile. Pathogenicity islands are genomic islands that contain genes responsible for conferring virulence. Numerous pathogenicity islands have been described in Gram positive and Gram negative bacteria (Gal-Mor and Finlay, 2006). PIs can contain a set of genes that confer a new phenotype to the recipients. Examples include PIs encoding exotoxins,

18 antimicrobial resistance and secretion systems. If these islands are acquired and then expressed in the proper context, the bacteria may be capable of surviving in a new ecological niche. The acquisition of PIs represents an evolutionary leap and can result in a phenotypic switch from non-virulent to virulent (Groisman and

Ochman, 1996).

1.13. The Francisella Pathogenicity Island (FPI)

The FPI was identified to have a G+C content of 26.6 percent, which is 6.6 percent less than the rest of the Francisella genome. This 30-kb region contains

4 large ORFs (2.5 to 3.9 kb) and 12 small ones (less than 1.5 kb) (Figure 1.1).

Large regions of the FPI have been identified to be required for virulence. The

FPI is flanked by the transposases tnpAB on one end and tnpA on the other serving as mobile genetic elements. There are also 16 base pair repeats at either end of the FPI. A putative operon exist consisting of pdpD, iglABCD

(intramacrophage growth locus) (Stephen A. Rodriguez, ). Genes of the igl locus are important for replication within macrophages and for virulence. PdpA, pdpB and pdpC are transcribed in the opposite direction as pdpD with several short

ORFs between that aren’t known to encode functional proteins (Nano et al.,

2004). The genes pdpABCD share homology with genes of Plasmodium. This indicates that part of the FPI may have been acquired from a eukaryotic pathogen in a horizontal gene transfer. Genes within the FPI are upregulated in

19 response to stress and within macrophages. Two identical copies of the FPI are present in the subspecies F. tularensis and holarctica. The second copy of the

FPI is missing in F. novicida. Differential gene usage and expression of FPI genes has been suggested as a possible explanation for the difference in virulence for such closely related strains.

Recent research has suggested that the FPI may encode a secretion system (Ludu et al., 2008b). Most intracellular pathogens employ secretion systems to modify the endocytic pathway of host immune cells (Santic et al.,

2006). Currently there are seven distinct secretion systems described (Types I-

VII). Each system is made up of several components which serve to translocate effectors across the bacterial periplasmic and outer membranes. The components of Type III and IV secretion systems form a needle like appendage that can penetrate host cell plasma membrane and deliver molecules directly from the cytosol of the bacteria into the cytoplasm of target cells. Secretion system types I, II and IV directly secrete effectors across the bacterial inner and outer membrane to the extracellular space without a periplasmic intermediate.

Types II and V use the Tat and Sec beta barrel proteins to move effectors across the inner membrane into the periplasmic space. Once effectors are in the periplasmic space, they are moved across the outer membrane by a second secretion protein. With the exception of the Type II Tat inner membrane translocation, the secretion of effectors is an active process requiring ATP as an energy source. Effector proteins of pathogenic bacteria include exotoxins like

20 hemolysins and mediators of actin polymerization. For example, Salmonella enterica secretes the effectors SopE,E2 and D into the host cytoplasm via a type

III secretion system encoded by a pathogenicity island (SPI-1) that initiates actin polymerization and mediates bacterial initiated phagocytosis. The process is essential for envasion of M cells in the intestinal epithelium and for Salmonella pathogenesis. IglA and IglB are homologous to protein in type six secretion systems (T6SSs) and pdpB encodes a protein homologous to IcmF a protein that is found in the Type IV secretion apparatus of Legionella (Ludu et al., 2008b).

Type VI secretion systems (T6SSs) were first described in Vibrio cholera and Pseudomonas aerugninosa and since have been identified in approximately a dozen Gram negative bacteria (Cascales, 2008). The type VI style of secretion systems were first identified by homologs to the type IV secretion system proteins IcmF and DotU in genomes that do not contain homologs to the other components of type IV secretion system. The remaining components are unique to type VI systems. DotU/ IcmF are required for L. pneumophila modification of the host endocytic pathway via secretion by type IV secretion machinery. In silico analysis predict T6SSs in about 100 bacterial genomes, but most described systems have been identified by screens looking for loss of virulence. Many

T6SSs are encoded within a pathogenicity island in which the G+C content is significantly but not dramatically different from the rest of the genome (Cascales,

2008). The FPI has a G+C content of 26.6 percent while the rest of the genome contains 33.2 percent. The core components which are present in T6SSs

21 include the DotU/IcmF, the ClpV ATPase and the putative secreted effectors Hcp

(lysin co-regulated protein) and VgrG (valine-glycine repeats) proteins.

Upregulation of most type VI secretion clusters is dependent upon contact with or growth inside host cells (Cascales, 2008). The FPI encodes homologs of T6SSs but is distinct from other well characterized systems (Filloux et al., 2008). The

FPI genes pdpB and pdpD encode IcmF-like proteins (Cascales, 2008;Filloux et al., 2008). Homologs of ClpV (FTN_1311/FTT_1701 and FTT_1346), Vgr

(FTN_1312/FTT_1702 and FTT_1347) and DotU(FTN_1315/FTT_1705 and

FTT_1350) are encoded downstream of pdpB. The T6SS component Hcp is represented by a gene downstream of pdpD right after the iglABCD locus,

FTN_1320/FTT_1710 and FTT_1355. It remains to be shown whether the FPI is a functional secretion system and what the secreted effectors are and what functional role they might play in intramacrophage survival.

Variability in pdpD, the leading gene of the putative pdpDiglABCD operon has been noted at the subspecies level. PdpD is present in the fully virulent F. tularensis Schu4 strain but is missing in the less virulent F. holarctica strains. In

F. novicida PdpD is modified by a hydrophilic 48 amino acid insertion in the middle of the protein (Ludu et al., 2008b). Different phenotypes have been reported for F. novicida pdpD mutants. One phenotype demonstrated that F. novicida null mutant is defective for intramacrophage growth and virulence in mice (Nano et al., 2004); however, this mutant had polar effects on downstream genes (iglABCD). An in frame deletion of pdpD that does not affect the

22 expression of downstream genes retained the ability to replicate in macrophages and yet was still attenuated when used to infect mice intradermally (Ludu et al.,

2008b). The construction of another pdpD mutant showed only a mild defect for growth in macrophages and maintained high virulence in mice (Stephen A.

Rodriguez, ). The data on the affect of pdpD deletion on the virulence of F. novicida is still inconclusive. PdpD localizes to the outer membrane and associates with IglA and IglB. Overexpression of pdpD increases the amount of lglA and IglB that are detectable from membrane fractions suggesting that PdpD,

IglA and IglB physically interact (Ludu et al., 2008b).

The gene encoding IglC was identified by transposon mutagenesis to be important for survival and replication within macrophages and for virulence (Gray et al., 2002;Weiss et al., 2007). Production of the IglC protein is increased upon infection of macrophages. An allelic replacement mutant of IglC proved that the virulence phenotype was not due to polar affects on downstream genes

(Golovliov et al., 2003). IglC is required for phagosomal escape and replication within the cytosol (Lindgren et al., 2004;Santic et al., 2005;Santic et al., 2007).

IglC has also been shown to be necessary for Francisella to survive in arthropod cells and amoebae linking the ability to survive in a mammalian host to survival in the environment (Santic et al., 2009;Read et al., 2008;Lauriano et al., 2004). The genes upstream of iglC, iglA and iglB have also been shown to be required for intramacrophage growth and virulence in the F. novicida model (de Bruin et al.,

2007).

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1.14. Regulation of the FPI

Two genes arranged in an operon were identified to be required for the replication of F. tularensis within macrophages, named mglA and mglB

(macrophage growth locus). MglA is upregulated upon infection of macrophages

(Baron and Nano, 1998;Lauriano et al., 2004;Santic et al., 2005b) and an mglA mutant is not capable of escaping the macrophage phagosome (Santic et al.,

2005b). F. novicida mglA null mutants are attenuated in the mouse model

(Santic et al., 2005b;Lauriano et al., 2004;Guina et al., 2007;Brotcke et al.,

2006;West et al., 2008). MglA has been characterized as a transcription factor that is responsible for regulating approximately 100 genes including the genes within the FPI (Brotcke et al., 2006;Lauriano et al., 2004;Guina et al., 2007).

Strains deficient in mglA have much lower concentrations of the FPI encoded protein IglC. An F. novicida ∆mglA mutant is attenuated when delivered via the aerosol route but does not provide protection against homolgous or heterologous challenge. The mglA mutant does replicate within the lungs and control of the mglA mutant infection involves MyD88 and IFN-γ (West et al., 2008). In addition to the FPI, other virulence factors such as the metalloprotease PepO are regulated by MglA (Guina et al., 2007)

PepO is a homologue for a proendothelin which is a precursor for a potent vasoconstrictor, endothelin. PepO secretion should result in increased vasoconstriction at the site of infection that may limit the spread of the bacteria

(Guina et al., 2007). There is a mutation in the secretion machinery responsible 24 for secretion of PepO in the more virulent F. tularensis subspecies that results in lower levels of PepO secretion and increased dissemination as compared to F. novicida that does not have this mutation (Oyston, 2008;Hager et al., 2006).

Therefore the strains most virulent in humans disseminate more rapidly from the site of infection and may explain in part the enhanced virulence of subspecies tularensis.

MglA is homologous to SspA from other Gram negative bacteria including

E. coli. Another SspA homologue (FTN_0549, FTT_0458) has been appropriately dubbed SspA and shown to be important for the regulation of

Francisella virulence genes. MglA and SspA both regulate similar sets of genes including the positive regulation of the FPI (Charity et al., 2007). MglA and SspA interact with one another and both can bind Francisella RNA polymerase

(RNAP). Francisella RNAP is unique in that it contains two distinct forms of the alpha subunit. Two separate genes encode these proteins and they are incorporated into the enzyme (Charity et al., 2007). Research suggests that

MglA and SspA cooperate to regulate genes important for Francisella virulence.

SspA is maximally expressed during stationary phase while MglA is maximally expressed during lag and exponential phages of growth (Brotcke et al., 2006).

MglA is upregulated during macrophage infection (Baron and Nano, 1998).

Another regulator of the FPI that is essential for intramacrophage replication and virulence in the mouse model has been named Francisella effector of virulence regulation or FevR. The expression of fevR itself is 25 regulated by MglA but both proteins are necessary for replication in macrophages and virulence indicating that they work in parallel to affect changes in FPI expression. FevR has poor homology to DNA binding proteins and appears to act upstream of MglA and SspA RNA polymerase binding (Brotcke and Monack, 2008).

1.15. Other Francisella Virulence Factors

Acid phosphatases have been shown to be important for survival within the phagosome of several intracellular pathogens. The four acid phosphatases encoded within the F. tularensis genome are also important for virulence. A F. novicida quadruple mutant of acpABC and hapA reduces acid phosphotase acitivity by 90 percent and renders the strain unable to replicate with macrophages and is attenuated in the mouse (Mohapatra et al., 2008;Mohapatra et al., 2007a).

Whole genome genetic screens have been used to identify Francisella virulence proteins. One of these studies has identified genes whose deletion results in strains with unique phenotypes. These mutants retain the ability to replicate with host derived phagocytes but are attenuated. An in vivo negative selection technique identified most of the known F. tularensis virulence factors

(Weiss et al., 2007). An F. novicida transposon library was constructed, pooled and used to infect mice. The mutants that did not survive in the mouse were 26 identified by microarray analysis. Forty-four previously unidentified genes of unknown function were identified that were involved in virulence. Of these, null mutants of FTT0398 and FTT01048 were shown to have a significant virulence defect and yet retained their ability to replicate within macrophages in vitro

(Weiss et al., 2007). These studies are the only evidence for attenuation of F. tularensis without disrupting the ability to replicate within macrophages. The function of these genes has not yet been determined.

1.16. Two Component Regulatory Systems and Francisella

Two Component Regulatory Systems (TCS) are widely used by Gram negative bacteria to react to environmental changes and respond by regulating gene expression. TCS are typically composed of a periplasmic membrane bound sensor kinase and a cytosolic response regulator. The kinase has a sensor or input domain that is exposed to the periplasmic space and a cytosolic localized domain called a transmitter domain. The input domain monitors the extracellular milieu and reacts to a signal (e.g. pH, cations or peptides) resulting in a conformational change in the transmitter domain (Mascher et al., 2006). The conformational change allows the transmitter domain to accept a phosphate from

ATP to a conserved histidine residue. A phosphotransfer event allows the phosphate to be transferred to a conserved aspartate residue in the receiver domain of the response regulator. Phosphorylation of the response regulator

27 results in a conformational change that allows the output domain to affect changes in gene regulation. Most response regulators can bind to regulated gene promoters upon phosphorylation initiating or upregulated transcription

(Beier and Gross, 2006). The genes encoding the components of TCS are traditionally arranged in an operon under control of a single promoter with the histidine kinase directly downstream of the response regulator. Several thousand TCS have been described in over 100 species. Some organisms, like

E. coli and Pseudomonas, employ dozens of TCS to respond to changes in their environment (Beier and Gross, 2006).

TCS have been extensively studied in Salmonella enterica serovar

Typhimurium (S. typhimurium) which has 38 such systems and is a model organism for studying TCS control of virulence traits (Ashby, 2004). The S. typhimurium PmrAB TCS is activated in response to the host macrophage environment by the PhoPQ TCS. Activation of PhoPQ results in the upregulation of the pmrCAB operon which encodes the PmrAB TCS. PmrAB is upregulated upon infection of host cells by the PhoPQ dependent mechanism or independently by PmrB detection of lowered pH and phosphorylation of PmrA

(Gunn et al., 1998). PmrA has been shown to regulate somewhere between 20 and 100 genes either directly or indirectly (Tamayo et al., 2005a;Tamayo et al.,

2002;Marchal et al., 2004). This regulon plays a role in host-pathogen response as well as cellular functions. The PmrAB TCS is activated in the presence of antimicrobial peptides (AMPs) and is responsible for mediating the decoration of

28

Salmonella LPS resulting in an overall change in cell wall charge and resistance to AMPs (Moskowitz et al., 2004;Navarre et al., 2005). The role that this modification plays in vivo is not completely understood. A pmrA null mutant has a virulence defect when infection occurs via the oral but not the intraperitoneal route of infection (Gunn et al., 2000). LPS modification plays a role in protecting invading bacteria from host innate immune factors providing a fitness advantage.

Orthologs of the PmrAB system have been discovered in several other

Gram negative bacteria and are involved in the regulation of virulence factors.

PmrA mediates LPS modifications in bacteria such as Yersina pseudotuberculosis, Pseudomonas aeruginosa and Escherishia coli (Flamez et al., 2007b;McPhee et al., 2006;Moskowitz et al., 2004;Raetz et al., 2007). PmrA regulates the icm/dot type IV secretion system in Legionella pneumophilia and

Coxiella burnetii. Regulation of the icm/dot type IV secretion system is required for virlence in both of these pathogens (Zusman et al., 2007;Al-Khodor et al.,

2009).

An ortholog to the Salmonella response regulator PmrA exists in all

Francisella subspecies; however, the histidine kinase, PmrB, is not present. This orphaned response regulator was investigated to determine its importance for virulence. To determine if PmrA is involved in regulation of virulence determinants, a PmrA null mutant was constructed in F. novicida. This mutant is more susceptible to antimicrobial peptides suggesting that PmrA may be involved in regulating LPS modification in Francisella. Microarray analysis indicated that 29

PmrA is involved in transcription of the FPI that is required for replication within host phagocytes. Indeed, F. novicida ∆pmrA is defective for intramacrophage replication. Additionally, the pmrA null mutant is attenuated in mouse model of infection (Mohapatra et al., 2007b). This report investigates the mechanism by which PmrA regulates transcription of the FPI, thereby mediating replication in host macrophages and virulence. We hypothesize that PmrA is a DNA binding protein that recognizes regulated virulence gene promoters and that the binding of PmrA is dependent upon phosphorylation by a histidine kinase.

30

31

Figure 1.1. The Francisella Pathogenicity Island (FPI). Schematic of the genomic arrangement of the FPI. Note the variations in pdpD and the surrounding region between Francisella subspecies. aa = amino acid. bp = base pair. Adapted from (Nano et al., 2004) and (Ludu et al., 2008b).

31

CHAPTER 2:

PmrA Regulates Francisella Virulence Factors Independent of MglA and

SspA

2.1. Introduction

Francisella tularensis subspecies tularensis (F. tularensis) is a Gram negative non-motile facultative intracellular pathogen and the causative agent of tularemia. There are four subspecies of Francisella tularensis (tularensis, holarctica, mediasiatica, and novicida). Type A strains (F. tularensis) recovered primarily from North America are the most virulent. Less virulent Type B (F. tularensis subspecies holarctica) strains are found in Europe. Francisella tularensis subspecies novicida (F. novicida) is closely related to Type A F. tularensis (>96% DNA homology) (Keim et al., 2007) and causes a tularemia-like disease in mice but does not cause disease in immunocompetent humans.

Infections with F. tularensis are characterized by invasion of and replication within host phagocytes. Upon entry into macrophages, Francisella modifies the

32 endocytic pathway, preventing phagolysosomal fusion. Once the phagosomal maturation is halted, the bacteria escape to the cytosol where they replicate

(Santic et al., 2006b). It is clear that in vivo expression of the Francisella

Pathogenicity Island (FPI) is critical for the ability of this pathogen to cause disease (Santic et al., 2006;Lai et al., 2004). Mutations in the FPI result in attenuation, inability to escape the phagosome, and deficient replication within macrophages (Nano et al., 2004). MglA, SspA, FevR and PmrA have been shown to be necessary for Francisella virulence and transcription of the FPI, and

MglA and SspA have been shown to bind to RNA polymerase (Charity et al.,

2007;Mohapatra et al., 2007b;Lauriano et al., 2004;Lauriano et al., 2004). How these proteins coordinate the regulation of the FPI is not well understood.

Two-component regulatory systems (TCS) play a critical role in the regulation of virulence determinants for many bacterial pathogens. TCS are composed of a sensor kinase and a response regulator. Typically, response regulators are phosphorylated at a conserved aspartate residue by the sensor kinase that has autophosphorylated at a conserved histidine residue.

Autophosphorylation occurs in response to an environmental signal that is detected by the membrane-bound sensor kinase. The phosphorylated response regulator then causes changes in transcription by binding to gene promoters.

Traditional TCS are arranged tandemly in the genome within an operon consisting of a single promoter followed by the response regulator and then the sensor kinase (Flamez et al., 2007a;Stock et al., 2000;Mitrophanov and

33

Groisman, 2008;Beier and Gross, 2006). There are no tandemly-arranged TCS in virulent Francisella, but orphaned members including PmrA are present

(Mohapatra et al., 2007b).

Our previous report showed that the absence of the orphaned response regulator pmrA rendered F. novicida defective for intramacrophage survival and attenuated virulence in mice. We hypothesize that PmrA is involved in the regulation of known virulence genes such as the FPI. A comparison of the expression profiles of F. novicida wild-type and various mutants was performed using a custom Affymetrix GeneChip. This chip contains probesets for each of the 1,804 open reading frames (ORFs) that have been identified in F. tularensis

Schu4 by whole genome sequencing. Each probeset contains 11 perfect match and 11 mis-match probes. Each perfect match probe is a unique sequence, and the set spans the length of the ORF. Mis-match probes are different from perfect match probes by a single nucleotide. The ratio and intensities of perfect match to mis-match probes are used to determine the expression level of the ORF the probeset represents.

34

2.2. Results

PmrA regulates the FPI. Microarray analysis was performed on RNA extracted from broth-grown bacteria harvested at an optical density at 600 nanometers of 0.5 corresponding to exponential growth phase. Reverse transcriptase was used to generate cDNA which was incompletely digested with

DNase I. The cDNA fragments were labeled and hybridized to a Genechip with probesets to each of the ORFs of F. tularensis Schu4. Genes were selected as being differentially regulated if they had a p-value less than 0.05 and were 1.5 fold or greater change in expression. This selection criteria eliminated genes, like house keeping genes, that were not expected to be differentially regulated.

For example, dnaK (FTT_1269, fold-change = -1.15) and tul4 (FTT_0093, fold- change = 1.26) were not identified as being differentially regulated in the F. novicida ∆pmrA strain. Comparison of the gene expression profiles of an F. novicida ∆pmrA mutant to that of wild-type bacteria identified 56 PmrA regulated genes. Twelve of the 17 genes encoded within the FPI were positively regulated by PmrA. Interestingly, pdpD, iglA and iglC were not identified as being differentially regulated, but iglB and iglD were significantly different than wild- type. This result is unusual because pdpD iglABCD is transcribed as an operon

(Stephen A. Rodriguez, ) and by definition the genes within an operon are transcribed together. Five of the 56 differentially regulated genes were those composing the pmrA operon (pmrA lepB rnc truB rnr). Not surprisingly, pmrA

35 had the largest difference in expression at 143-fold change compared to wild- type. This analysis also identified fevR (FTT_0383) as being differentially regulated in the pmrA null mutant, but neither mglA (FTT_1275, fold-change = -

1.01) or sspA (FTT_0458, fold-change = 1.03) were regulated by PmrA (Table

2.1). RT-PCR analysis of select genes generally agreed with the microarray data

(Table 2.2). Interestingly, pdpD was dramatically differentially expressed (950 fold) in the ∆pmrA mutant when analyzed using RT-PCR but not in the microarray analysis. This data indicates that PmrA either directly or indirectly regulates the expression of the FPI but not the expression of MglA or SspA which also regulate the FPI.

MglA regulates the FPI. The F. tularensis microarray was used to compare the expression profile of an mglA null mutant to wild-type bacteria. The analysis identified 84 genes that were differentially regulated by at least three- fold in the mglA mutant (Table 2.3). A cut off of three-fold or greater was arbitrarily assigned to keep the list of differentially regulated genes manageable.

More than 300 genes were identified that were differentially regulated by more than 1.5 fold than in wild-type bacteria. Another microarray analysis of a mglA mutant reported a very similar set of approximately 100 genes differentially regulated by greater than two-fold (Brotcke et al., 2006). Both microarray analyses of different MglA mutants identified all of the genes within the FPI as being differentially regulated, indicating that MglA positively regulates the FPI.

MglA does not regulate PmrA, SspA or FevR. The microarray used in our study

36 is an Affymetrix custom array while the previously published data employed a

DNA spotted array. This data confirms the previous published data and supports the use of both microarray technologies for expression analysis in Francisella.

The histidine kinases KdpD and FTN_1453 regulate the FPI and the

PmrA operon. Microarray analysis of a null mutant of the orphaned response regulator PmrA determined that it is required for transcription of the FPI. If PmrA is functioning as a response regulator, there should be a linked histidine kinase that phosphorylates PmrA. Transposon mutants of the three putative histidine kinase genes identified in the F. novicida genome (http://go.francisella.org/cgi- bin/frangb/genomelist.cgi), FTN_1453 (JSG2890), FTN_1617 (qseC, JSG2892), and FTN_1715 (kdpD, JSG2894)(Gallagher et al., 2007) were used to attempt to identify which kinase is linked to PmrA. Microarray analysis compared the gene expression profile of each of these kinase mutants to that of wild-type bacteria.

Of the three kinases, the gene expression profile of the kdpD transposon mutant is most similar to that of the pmrA mutant. Ninety-five genes were differentially regulated in the kdpD transposon mutant. As one would expect, kdpE, the response regulator linked to kdpD, was identified as being regulated by KdpD in

F. novicida. Three members of the pmrA operon and 12 genes within the FPI were positively regulated by KdpD (Table 2.4). Mutation of qseC had little effect on the gene expression profile with only three genes differentially regulated by more than 1.5-fold (Table 2.5). One of these genes was lepB, which is a part of the pmrA operon (Sammons-Jackson et al., 2008). Fifty-two genes were

37 differentially regulated in the FTN_1355 transposon mutant. Four of these genes are within the FPI and three are a part of the pmrA operon (Table 2.6).

Surprisingly, pmrA was not identified in microarray analysis of any of the kinase mutants even when other members of the pmrA operon were identified as being differentially regulated. qRT-PCR analysis of the expression of pmrA and iglC in the kdpD and qseC transposon mutants do not agree with the microarray analysis. Expression of pmrA and iglC are not significantly different from wild- type bacteria when analyzed using qRT-PCR (Figure 2.1). The expression analysis of the virulence factors in histidine kinase mutants is thus far inconclusive; however, microarray data indicates that KdpD and FTN_1355 affect transcription of the FPI and the pmrA operon and that the pmrA and kdpD regulons most closely resemble one another.

2.3. Discussion

Microarrays are commonly used to screen the bacterial transcriptome to reveal changes in gene expression that may explain phenotypic differences.

Using a microarray containing probes for all of the ORFs in F. tularensis Schu4, it was determined that PmrA is involved in the regulation of FPI. Our analysis of an mglA mutant showed that MglA is also involved in regulation of the FPI. This data is in agreement with a previous study that reported that MglA is involved in the regulation of about 100 genes (Brotcke et al., 2006). PmrA does not regulate 38

MglA nor does MglA regulate PmrA and SspA is not regulated by either PmrA or

MglA. The microarray data indicates FevR is regulated by PmrA confirming qRT-

PCR data that showed decreased expression of fevR in pmrA mutants (Denise

Monack, Stanford University, personal communication). The identified regulons of PmrA and MglA have 29 genes in common, of which 14 are encoded within the FPI. The function and relevance of the remaining 15 genes they share are unknown (Figure 2.2). PmrA, MglA, SspA, and FevR have been shown to regulate the FPI (Charity et al., 2007;Brotcke and Monack, 2008;Mohapatra et al., 2007b;Brotcke et al., 2006), indicating that regulation of the FPI is complicated, requiring at least four proteins for transcription.

The availability of transposon mutants in the three F. novicida putative histidine kinases allowed expression analysis comparing these mutants to wild- type bacteria. By microarray, KdpD is involved in regulating twelve of the fourteen genes within the FPI and the PmrA operon. FTN_1453 was shown to be involved in regulating four genes within the FPI, including iglC (FTT_1357), and the three genes in the pmrA operon. Comparing genome-wide expression of a qseC transposon mutant to wild-type showed only three differentially regulated genes. The gene expression profile of the kdpD transposon mutant most closely matches that of the pmrA null mutant. The regulons identified for FTN_1453 and kdpD are quite similar, sharing 31 differentially regulated genes (Figure 2.3).

Only four of these are FPI genes, and three are a part of the pmrA operon. The majority of the remaining differentially regulated genes shared between

39

FTN_1453 and kdpD are hypothetical. All three kinase mutant lists of differentially regulated genes have one gene in common, lepB (Figure 3). LepB is encoded by the ORF directly downstream of pmrA. Taken together, this data suggests that KpdD and the kinase encoded by FTN_1453 are involved in regulating PmrA and the FPI, but KdpD is more likely to be PmrA’s linked histidine kinase.

Microarray analysis is a powerful tool because it can screen the expression profile of a whole genome at one time using a single sample.

Microarray’s strength is also directly related to its weakness (Faucher et al.,

2006). The complicated statistical analysis required to make thousands of multiple gene comparisons can affect the results in unpredictable ways (Hinton et al., 2004). For this reason, a secondary analysis, like quantitative real time- polymerase chain reaction (qRT-PCR), is required to confirm the results . In the analysis presented here, unexplained anomalies were noted. The differentially regulated genes that were identified in this study for kdpD and FTN_1453 included lepB, rnc, truB, and rnr but not the lead gene of the operon pmrA

(Sammons-Jackson et al., 2008). Comparison of the pmrA mutant to wild-type bacteria identified iglB and iglD but not the other members of the pdpD operon

(Stephen A. Rodriguez, ). It is likely that the members of these operons that were not identified by microarray are in fact differentially regulated. These genes may not have been reported because their fold change compared to wild-type did not meet the criteria for biological significance. The modest fold change in gene

40 expression reported here may be the result of the conditions used to generate the RNA for analysis. Bacteria were grown in liquid broth and harvested at equivalent optical densities. The virulence factors of F. tularensis are upregulated in response to host cell interaction; however, these culture conditions do not mimic the intramacrophage environment, and therefore the virulence genes identified may not be activated. The fold changes reported for

MglA differentially regulated genes were much higher. This makes sense, taking into account that the MglA is expressed even in broth cultures as indicated by the fact that ∆mglA mutants have a growth defect (Lauriano et al., 2004;Brotcke et al., 2006;Santic et al., 2005). A better picture of the gene expression profile and identification of PmrA regulated genes can be determined by microarray analysis of bacteria that are harvested from an environment which mimics host cell interaction, activating virulence gene systems.

The microarray results demonstrate that PmrA and MglA regulate that FPI but do not regulate one another. PmrA, MglA and SspA all regulate the FPI but not one another indicating that these proteins cooperatively regulate the pathogenicity island. KdpD and the product of FTN_1453 regulate the FPI and

PmrA, suggesting they play a role in the regulation of PmrA. Further study will be needed to determine whether KdpD or FTN_1453 phosphorylate PmrA and if this phosphorylation leads to an increase in PmrA’s ability to bind the promoter region of regulated genes.

41

Figure 2.1. qRT-PCR analysis of pmrA and iglC in F. novicida mutant strains. F. novicida ∆pmrA expresses very little iglC and virtually no pmrA. The F. novicida ∆kdpD and ∆qseC mutants express equilavent amount of pmrA and iglC as wild type bacteria. Black bars = pmrA. Grey bars = iglC. RCN = relative copy number

42

Figure 2.2. Venn diagram of the F. novicida ∆pmrA, ∆mglA and ∆kdpD regulons as determined by microarray analysis. Numbers represent the number of differentially regulated genes that had at least 1.5 (∆pmrA and ∆kpdD) or 3.0 (∆mglA) fold change when compared to wild-type bacteria.

43

Figure 2.3. Venn diagram of the F. novicida ∆FTN_1355, ∆qseC and ∆kdpD regulons as determined by microarray analysis. Numbers represent the number of differentially regulated genes that had at least 1.5 fold change when compared to wild-type bacteria.

44

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

Two-component response 1 FTT1557c_at pmrA 1.62E-12 -143.854 regulator

2 FTT0029c_at hypothetical protein NA 0.000447658 -4.73025

3 FTT1555c_at Ribonuclease III rnc 1.02E-08 -4.72911

4 FTT1556c_at signal peptidase I lepB 6.15E-11 -4.11701

tRNA pseudouridine 5 FTT1554c_at truB 2.56E-08 -3.48811 synthetase B

6 FTT0783_at Arylsulfatase ars 0.00226224 -2.64372

7 FTT1351_s_at hypothetical protein NA 0.00021087 -2.42773

diaminopimelate 8 FTT0027c_at lysA 0.0225724 -2.39449 decarboxylase

9 FTT1346_s_at hypothetical protein NA 0.00267679 -2.32642

10 FTT1353_s_at hypothetical protein NA 0.00294948 -2.24355

11 FTT0786_at hypothetical protein NA 0.020893 -2.17426

12 FTT0784_at hypothetical protein NA 0.0209019 -2.0073

13 FTT0241c_at hypothetical protein NA 0.0263488 -2.00302

14 FTT1352_s_at hypothetical protein NA 0.00709283 -1.97347

cystathionine beta-synthase 15 FTT1287_at cbs 0.0033774 -1.94308 (cystein synthase)

16 FTT1344_s_at hypothetical protein pdpA 0.0025581 -1.86818

conserved hypothetical 17 FTT1180_at NA 0.00533776 -1.85594 protein, pseudogene

18 FTT1350_s_at hypothetical protein NA 0.011412 -1.84608

Continued

Table 2.1: Differentially regulated genes when comparing F. novicida ∆pmrA to F. novicida wild-type bacteria by microarray. 45

Table 2.1 continued.

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

19 FTT1349_s_at hypothetical protein NA 0.00825986 -1.80895

20 FTT0611c_at beta-lactamase NA 0.0139178 -1.78039

conserved hypothetical 21 FTT0852_at NA 0.0171499 -1.73733 protein, pseudogene

22 FTT1347_s_at hypothetical protein NA 0.0242315 -1.73713

23 FTT1345_s_at hypothetical protein pdpB 0.0179171 -1.65009

24 FTT1354_s_at hypothetical protein NA 0.0120281 -1.64562

conserved hypothetical 25 FTT1032_at membrane NA 0.0453979 -1.63502 protein,pseudogene

26 FTT0403_at peptide deformylase def1 0.00733194 -1.60275

27 FTT0809c_at Recombination protein recR recR 0.00804579 -1.59396

28 FTT0383_at hypothetical protein NA 0.00604246 -1.58403

conserved membrane 29 FTT0101_at NA 0.00542161 -1.58217 hypothetical protein

outer membrane 30 FTT0421_at NA 0.000474065 -1.57757 lipoprotein, pseudogene

31 FTT1542c_at outer membrane protein 26 omp26 0.0115477 -1.56399

intracellular growth locus, 32 FTT1356c_s_at iglD 0.0242764 -1.56284 subunit D

intracellular growth locus, 33 FTT1358c_s_at iglB 0.00230781 -1.54681 subunit B

34 FTT0813_at hypothetical protein NA 0.0409866 -1.54517

hypothetical membrane 35 FTT0297_at NA 0.045887 -1.53701 protein

36 FTT1553c_at Ribonuclease R rnr 0.00256858 -1.53522

Continued

46

Table 2.1 continued.

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆prmA vs. WT)

37 FTT0559c_at cytidylate kinase cmk 0.0137657 -1.53231

Rhodanese-like family 38 FTT1127_at NA 0.00494136 -1.5224 protein

39 FTT1541c_at hypothetical protein NA 0.014424 -1.51895

40 FTT0394_at hypothetical protein NA 0.0155197 -1.50306

Sodium hydrogen 41 FTT0669_at NA 0.0227026 1.5382 exchanger family protein

hypothetical membrane 42 FTT0263_at NA 0.00442426 1.66304 protein

conserved hypothetical 43 FTT0193c_at NA 0.000988585 1.72826 membrane protein

44 FTT0180_at Acetyltransferase NA 0.00408648 1.75961

45 FTT1140_at hypothetical protein NA 0.0176831 1.81463

46 FTT0465_at hypothetical protein NA 0.00747885 1.9122

47 FTT1145_at cation transport regulator chaB 0.0321222 1.91705

major facilitator 48 FTT0488c_at superfamily (MFS) transport NA 0.0100722 1.98102 protein

Short-chain dehydrogenase 49 FTT1144_at reductase (SDR) family NA 0.00171609 2.02711 protein, pseudogene

50 FTT1143_at hypothetical protein NA 0.000373195 2.25231

conserved hypothetical 51 FTT1141_at NA 0.000980579 2.26303 protein, pseudogene

Continued

47

Table 2.1 continued

Gene p-value Fold-Change Probeset ID Gene Title # Symbol (∆pmrA vs. WT) (∆prmA vs. WT)

glycosyl hydrolase, family 3, 52 FTT1565c_at NA 5.95E-05 2.90943 pseudogene

53 FTT0401_at hypothetical protein NA 0.000943353 2.99822

hypothetical membrane 54 FTT0272_at NA 4.34E-07 4.88705 protein

hypothetical membrane 55 FTT0172_at NA 1.15E-05 5.64777 protein, fragment

56 FTT1242_at hypothetical protein NA 6.00E-08 13.2638

48

Fold-change

Gene Symbol Description Microarray qRT-PCR FTT_0029 NA Hypothetical protein -4.73 4.89 ± 0.45 FTT_1346 NA Hypothetical protein -2.33 6.00 ± 0.823 FTT_1347 NA Hypothetical protein -1.74 4.3 ± 0.65 FTT_1354 NA Hypothetical protein -1.65 3.2 ± 0.38 FTT_1356 NA Intracellular growth locus, subunit D -1.56 7.5 ± 1.1 FTT_1357 iglC Intracellular growth locus, subunit C NDR 1.8 ± 0.25 FTT_1360 pdpD pdpD NDR 950 ± 8.5 FTT_1555 rnc RNase III -4.73 8.2 ± 1.5 FTT_1556 lepB Signal peptidase I -4.12 68 ± 3.45 FTT_1242 NA Hypothetical protein +13.26 +96 ± 5.5

Table 2.2. Confirmation of differential expression by qRT-PCR. A subset of the differentially regulated genes in F. novicida ∆pmrA compared to F. novicida wild- type bacteria identified by microarray analysis were analyzed by qRT-PCR (adapted from (Mohapatra et al., 2007b)). NDR = not differentially regulated.

49

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

1 FTT0430_at S-adenosylmethionine speH 0.0332215 -5.70584 decarboxylase ABC transporter, 2 FTT0176c_at membrane protein, NA 0.264813 -5.23356 pseudogene 3 FTT1786_at conserved hypothetical, NA 0.0287203 -4.77127 pseudogene 4 FTT0303c_at L-lactate dehydrogenase lldD 0.371975 -3.97801 5 FTT0362c_at hypothetical protein NA 0.0246786 -3.6107 6 FTT0776c_at Ribonuclease D Rnd 0.0690291 -3.54436 7 FTT1564_at hypothetical protein NA 0.0027869 -3.38819 8 FTT0678c_at hypothetical lipoprotein NA 0.0372677 -3.35085 9 FTT0736_at hypothetical protein NA 0.290057 -3.29814 Iron-containing alcohol 10 FTT0517_at dehydrogenase,pseudog NA 0.0850669 -3.2933 ene 11 FTT0948c_at Aldo keto reductase NA 0.00742846 -3.21573 12 FTT0175c_at ABC transporter, ATP- NA 0.266918 -3.15775 binding protein 13 FTT1078c_at hypothetical protein NA 0.316455 -3.108 phosphatidylserine 14 FTT0384c_at decarboxylase Psd 0.000168788 -3.04444 proenzyme 15 FTT0392c_at hypothetical protein NA 0.00550767 -3.01363 mandelate racemase 16 FTT0735_at muconate lactonizing NA 0.0147645 -3.00781 enzyme family protein, pseudogene major facilitator 17 FTT0488c_at superfamily (MFS) NA 0.0253555 3.03321 transport protein

Continued

Table 2.3: Differentially regulated genes (3-fold or greater) when comparing F. novicida ∆mglA to F. novicida wild-type bacteria by microarray.

50

Table 2.3 continued.

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

18 FTT0090c_at conserved hypothetical NA 0.00473526 3.13864 membrane protein 19 FTT0982_at hypothetical membrane NA 0.0622631 3.14701 protein 20 FTT1558c_at conserved hypothetical NA 0.022351 3.15101 protein, pseudogene 21 FTT0272_at hypothetical membrane NA 0.13513 3.15885 protein glycerophosphoryl diester 22 FTT0726c_at NA 0.000292768 3.19661 phosphodiesterase family protein Proton-dependent 23 FTT0953c_at oligopeptide transport NA 0.00191734 3.20475 (POT) family protein 24 FTT0172_at hypothetical membrane NA 0.215208 3.27251 protein, fragment 25 FTT1143_at hypothetical protein NA 0.00428303 3.32092 26 FTT1665_at aspartate pyrB 0.00105454 3.64631 carbamoyltransferase 27 FTT0663_at hypothetical protein NA 0.00848232 3.65593 28 FTT1141_at conserved hypothetical NA 0.0194262 3.67751 protein, pseudogene 30S ribosomal protein S6 29 FTT0178c_at modification protein- rimK 7.92E-05 3.76144 related protein outer membrane protein 30 FTT1542c_at omp26 0.000958812 3.7926 26 31 FTT0029c_at hypothetical protein NA 0.00680358 3.88534 32 FTT1431_at threonine efflux protein rhtC 0.00729041 3.93951 33 FTT1657c_at hypothetical protein NA 0.00144858 3.99459 major facilitator 34 FTT0070c_at superfamily (MFS) ampG 0.0191665 4.17126 tranport protein

Continued 51

Table 2.3 continued.

Gene p-value Fold-Change Probeset ID Gene Title # Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

35 FTT0421_at outer membrane NA 0.00254899 4.18824 lipoprotein, pseudogene 36 FTT1191_at Aminoacylase NA 0.000689432 4.29471 37 FTT1324_at conserved hypothetical NA 0.00215304 4.32103 membrane protein 38 FTT1140_at hypothetical protein NA 0.0109979 4.59371 39 FTT0297_at hypothetical membrane NA 0.00807957 4.66299 protein 40 FTT1070c_at conserved hypothetical NA 0.00392877 4.68357 protein, pseudogene 41 FTT1354_s_at hypothetical protein NA 0.00190979 4.79915 42 FTT1345_s_at hypothetical protein pdpB 0.00274583 5.14872 43 FTT0101_at conserved membrane NA 0.00352889 5.19373 hypothetical protein 44 FTT0611c_at beta-lactamase NA 0.00335719 5.25006 45 FTT1346_s_at hypothetical protein NA 0.000843353 5.40901 Proton-dependent 46 FTT0651_at oligopeptide transport NA 0.00413795 5.47405 (POT) family protein 47 FTT1541c_at hypothetical protein NA 0.0145995 5.58226 D-alanyl-D-alanine 48 FTT0724c_at carboxypeptidase dacB1 0.00229556 5.64981 (Penicillin binding protein) family protein, 49 FTT1347_s_at hypothetical protein NA 0.000283154 5.76946 50 FTT0296_at Pyrrolidone-carboxylate Pcp 5.47E-05 6.13172 peptidase 51 FTT1771_at hypothetical protein NA 0.00060796 6.19559 52 FTT0613c_at hypothetical protein NA 0.000540945 6.48354 53 FTT1682_at conserved hypothetical NA 4.48E-05 6.53867 protein, pseudogene 54 FTT1351_s_at hypothetical protein NA 0.00408871 6.93722 55 FTT0988_at hypothetical protein NA 0.0077561 7.01417

Continued 52

Table 2.3 continued.

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

56 FTT1350_s_at hypothetical protein NA 0.000875701 7.15075 57 FTT1360c_s_at hypothetical protein pdpD 0.00229986 7.18161 58 FTT1358c_s_at intracellular growth IglB 0.00127452 7.32454 locus, subunit B 59 FTT1353_s_at hypothetical protein NA 0.00133397 7.44621 60 FTT1349_s_at hypothetical protein NA 0.000179626 7.61572 61 FTT1091_at isochorismatase NA 0.0024697 7.76692 hydrolase family protein 62 FTT1352_s_at hypothetical protein NA 0.000299382 8.11156 63 FTT1357c_s_at intracellular growth IglC 0.00271922 8.7707 locus, subunit C glycosyl hydrolases 64 FTT1033_at family 31 yihQ 1.90E-05 9.49163 protein,pseudogene 65 FTT0254c_at hypothetical protein NA 0.000511943 10.5197 66 FTT0310_at amino acid permease NA 0.000188023 11.2928 67 FTT0998_at hypothetical lipoprotein NA 7.53E-05 11.7424 68 FTT1356c_s_at intracellular growth iglD 0.000188873 11.9729 locus, subunit D 69 FTT1565c_at glycosyl hydrolase, family NA 0.000928271 12.2433 3, pseudogene 70 FTT0989_at hypothetical protein NA 0.000679949 12.6513 71 FTT1090_at conserved hypothetical NA 0.00069957 12.7677 membrane protein 72 FTT1344_s_at hypothetical protein pdpA 0.000681278 12.9788 73 FTT1359c_s_at intracellular growth IglA 9.48E-05 13.259 locus, subunit A 74 FTT1089_at isochorismatase NA 0.000255061 14.531 hydrolase family protein 75 FTT0981_at hypothetical protein NA 0.0004472 14.5516 76 FTT0255c_at hypothetical protein NA 0.00018722 20.0031 77 FTT1275_at macrophage growth mglA 2.68E-05 20.8118 locus, subunit A 78 FTT0566_at hypothetical protein NA 0.0018847 20.9415

Continued 53

Table 2.3 continued.

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

metallopeptidase family 79 FTT1209c_at M13 protein, NA 0.000601081 30.9836 pseudogene 80 FTT0852_at conserved hypothetical NA 0.000114005 31.0054 protein, pseudogene conserved hypothetical 81 FTT1032_at membrane NA 0.000130647 31.4124 protein,pseudogene 82 FTT0241c_at hypothetical protein NA 0.00191123 33.1346 83 FTT1650c_at chorismate mutase NA 0.000328111 34.9855 84 FTT0980_at Aminotransferase, class II NA 4.85E-05 40.9387

54

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

1 FTT0029c_at hypothetical protein NA 0.016725 -3.46746

2 FTT0310_at amino acid permease NA 0.004722 -3.0328

VacJ lipoprotein, 3 FTT1182c_at vacJ 0.000285 -3.01386 pseudogene

4 FTT0241c_at hypothetical protein NA 0.004685 -2.99774

5 FTT1353_s_at hypothetical protein NA 0.003428 -2.63427

conserved membrane 6 FTT0028c_at NA 0.044625 -2.59244 protein

7 FTT1555c_at Ribonuclease III rnc 0.00141 -2.51195

oligopeptide transporter, subunit B, ABC transporter, 8 FTT0123_at oppB 0.017369 -2.42374 membrane protein, pseudoge

9 FTT1388_at hypothetical protein NA 0.010619 -2.35717

Uroporphyrinogen III 10 FTT1408c_at hemD 0.010688 -2.35424 synthase

11 FTT0786_at hypothetical protein NA 0.037571 -2.33469

12 FTT1350_s_at hypothetical protein NA 0.006668 -2.23956

13 FTT0566_at hypothetical protein NA 0.014159 -2.21933

transcriptional regulatory 14 FTT1735c_at kdpE 0.001137 -2.21344 protein, pseudogene

15 FTT1346_s_at hypothetical protein NA 0.019672 -2.1895

16 FTT0311c_at hypothetical protein NA 0.001575 -2.1836

Continued

Table 2.4: Differentially regulated genes (1.5-fold or greater) when comparing F. novicida ∆kdpD to F. novicida wild-type bacteria by microarray.

55

Table 2.4 continued.

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

3-methyl-2-oxobutanoate 17 FTT1389_at panB 0.020048 -2.1713 hydroxymethyltransferase

18 FTT1348_s_at hypothetical protein NA 0.022334 -2.1435

19 FTT1391_at Aspartate-1-decarboxylase panD 0.013727 -2.12482

20 FTT1774c_at Carboxypeptidase,fragment NA 0.016989 -2.11281

Proton-dependent 21 FTT0651_at oligopeptide transport NA 0.041452 -2.09722 (POT) family protein

22 FTT1349_s_at hypothetical protein NA 0.007003 -2.09461

D-methionine binding transport protein, ABC 23 FTT1125_at metIQ 0.002948 -2.08554 transporter, membrane and periplasmi

conserved hypothetical 24 FTT1004c_at NA 0.022515 -2.08538 membrane protein

conserved hypothetical 25 FTT0003c_at membrane NA 0.038316 -2.07781 protein,fragment

conserved hypothetical 26 FTT1779_at NA 0.031334 -2.04174 membrane, pseudogene

27 FTT1602_at hypothetical lipoprotein NA 0.037934 -2.03886

3-methyl-2-oxobutanoate 17 FTT1389_at panB 0.020048 -2.1713 hydroxymethyltransferase

18 FTT1348_s_at hypothetical protein NA 0.022334 -2.1435

19 FTT1391_at Aspartate-1-decarboxylase panD 0.013727 -2.12482

20 FTT1774c_at Carboxypeptidase,fragment NA 0.016989 -2.11281

Continued

56

Table 2.4 continued.

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

Proton-dependent 21 FTT0651_at oligopeptide transport NA 0.041452 -2.09722 (POT) family protein

22 FTT1349_s_at hypothetical protein NA 0.007003 -2.09461

D-methionine binding transport protein, ABC 23 FTT1125_at metIQ 0.002948 -2.08554 transporter, membrane and periplasmi

conserved hypothetical 24 FTT1004c_at NA 0.022515 -2.08538 membrane protein

conserved hypothetical 25 FTT0003c_at membrane NA 0.038316 -2.07781 protein,fragment

conserved hypothetical 26 FTT1779_at NA 0.031334 -2.04174 membrane, pseudogene

27 FTT1602_at hypothetical lipoprotein NA 0.037934 -2.03886

conserved hypothetical 28 FTT1032_at membrane NA 0.019951 -2.01995 protein,pseudogene

29 FTT1347_s_at hypothetical protein NA 0.020027 -2.00668

cystathionine beta-synthase 30 FTT1287_at cbs 0.01087 -2.00464 (cystein synthase)

31 FTT1439c_at Deoxyribonuclease NA 0.02479 -1.99743

32 FTT1351_s_at hypothetical protein NA 0.01379 -1.98889

33 FTT0222c_at hydrolase subunit ybgK 0.036961 -1.97206

34 FTT1556c_at signal peptidase I lepB 0.001644 -1.97134

Proton-dependent 35 FTT0686c_at oligopeptide transport NA 0.015102 -1.96094 (POT) family protein

Continued 57

Table 2.4 continued.

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

36 FTT1345_s_at hypothetical protein pdpB 0.009598 -1.95917

conserved hypothetical 37 FTT0224c_at NA 0.006484 -1.95269 protein, pseudogene

hypothetical membrane 38 FTT1740c_at NA 0.025631 -1.94944 protein, fragment

39 FTT0108c_at tRNA nucleotidyltransferase cca 0.028931 -1.92104

glutamine 40 FTT1166c_at amidotransferases class-II NA 0.021886 -1.91473 family protein

41 FTT0855c_at hypothetical protein NA 0.047035 -1.91427

conserved hypothetical 42 FTT1180_at NA 0.01667 -1.89756 protein, pseudogene

intracellular growth locus, 43 FTT1356c_s_at iglD 0.008948 -1.89329 subunit D

44 FTT0255c_at hypothetical protein NA 0.040115 -1.87751

chromosomal replication 45 FTT0001_at dnaA 0.006262 -1.87681 initiator protein dnaA

46 FTT0906c_at DNA topoisomerase I topA 0.005754 -1.86277

Type IV pili glycosylation 47 FTT0905_at NA 0.008942 -1.85094 protein

DNA polymerase III, delta 48 FTT0460_at holB 0.00813 -1.84603 prime subunit

49 FTT1344_s_at hypothetical protein pdpA 0.01441 -1.83551

conserved hypothetical 50 FTT1682_at NA 0.027906 -1.81953 protein, pseudogene

51 FTT1431_at threonine efflux protein rhtC 0.042251 -1.80178

Continued

58

Table 2.4 continued

Gene p-value Fold-Change Probeset ID Gene Title # Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

52 FTT1793c_at Aminopeptidase N pepN 0.008568 -1.79553

conserved hypothetical 53 FTT1090_at NA 0.044533 -1.79018 membrane protein

hypothetical membrane 54 FTT0297_at NA 0.029098 -1.77811 protein

Aldose 1-epimerase 55 FTT1146c_at galM 0.009444 -1.77799 (pseudogene)

DNA RNA endonuclease 56 FTT0610_at NA 0.018323 -1.77449 family protein

tRNA pseudouridine 57 FTT1554c_at truB 0.015671 -1.76581 synthetase B

3-demethylubiquinone-9 3- 58 FTT1590c_at ubiG 0.044294 -1.76227 methyltransferase

hypothetical membrane 59 FTT0119_at NA 0.009407 -1.76136 protein

conserved hypothetical 60 FTT0277c_at NA 0.040619 -1.75712 membrane protein

NADH dehydrogenase I, N 61 FTT0044_at nuoN 0.01244 -1.74914 subunit

62 FTT0981_at hypothetical protein NA 0.004239 -1.7471

conserved hypothetical 63 FTT0194c_at NA 0.026851 -1.74664 membrane protein

DNA polymerase III (CHI 64 FTT0298_at holC 0.027006 -1.73159 subunit) protein

65 FTT0254c_at hypothetical protein NA 0.03833 -1.71309

nucleoside permease NUP 66 FTT0115_at nupC1 0.038465 -1.71155 family protein

67 FTT1354_s_at hypothetical protein NA 0.026004 -1.7064

Continued 59

Table 2.4 continued

Gene p-value Fold-Change Probeset ID Gene Title # Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

68 FTT1541c_at hypothetical protein NA 0.010984 -1.7014

intracellular growth locus, 69 FTT1357c_s_at iglC 0.004831 -1.69959 subunit C

70 FTT0499_at hypothetical protein NA 0.038168 -1.69936

71 FTT1504_at hypothetical protein NA 0.039995 -1.69912

72 FTT0857c_at hypothetical protein NA 0.043191 -1.68975

ABC transporter, membrane 73 FTT1609_at NA 0.046064 -1.68333 protein

74 FTT0234c_at hypothetical protein NA 0.044094 -1.68044

75 FTT0663_at hypothetical protein NA 0.026499 -1.67925

2,3-bisphosphoglycerate- 76 FTT1329_at independent gpmI 0.025408 -1.65153 phosphoglycerate mutase

Biofunctional protein, glutaredoxin 3 protein 77 FTT0532c_at nrdB 0.011586 -1.64792 Ribonucleoside- diphosphate reducta

host factor I for 78 FTT0630_at bacteriophage Q beta hfq 0.038977 -1.64121 replication

79 FTT0838_at TolR protein tolR 0.022003 -1.62594

cytochrome d terminal 80 FTT0279c_at oxidase, polypeptide cydA 0.012302 -1.61652 subunit I

Pyrrolidone-carboxylate 81 FTT0296_at pcp 0.047212 -1.61475 peptidase

Ribonucleoside- 82 FTT0534c_at diphosphate reductase, nrdA 0.00135 -1.59787 alpha subunit

Continued 60

Table 2.4 continued

Gene p-value Fold-Change Probeset ID Gene Title # Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

83 FTT1467c_at L-aspartate oxidase nadB 0.035858 -1.59121 23S rRNA (Uracil-5-)- 84 FTT0705_at rumA 0.023173 -1.58592 methyltransferase

formyltetrahydrofolate 85 FTT0717_at purU 0.048209 -1.58422 deformylase, pseudogene

86 FTT1055c_at hypothetical protein NA 0.047419 -1.57944

87 FTT1371_at 50S ribosomal protein L32 rmpF 0.042044 -1.57506

88 FTT1059c_at Replicative DNA helicase dnaB 0.021513 -1.57031

89 FTT1573c_at outer membrane protein NA 0.034887 -1.56761

90 FTT0375_at S-transferase NA 0.042208 -1.56122

phosphate 91 FTT1754_at pta 0.047944 -1.52775 acetyltransferase

multidrug resistance 92 FTT1727c_at NA 0.014226 -1.52228 protein, membrane located

93 FTT0369c_at hypothetical protein NA 0.02457 -1.52005

Glutamate-1-semialdehyde- 94 FTT0927_at hemL 0.039741 -1.51134 2,1-aminomutase

95 FTT1508c_at GTP pyrophosphokinase relA 0.032781 -1.50135

61

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

1 FTT1556c_at signal peptidase I lepB 0.023744 -1.60681 conserved hypothetical 2 FTT0193c_at NA 0.029173 1.53207 membrane protein hypothetical membrane 3 FTT0272_at NA 0.029189 2.09263 protein

Table 2.5: Differentially regulated genes (1.5-fold or greater) when comparing F. novicida ∆qseC to F. novicida wild-type bacteria by microarray.

62

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (∆pmrA vs. WT) (∆pmrA vs. WT)

1 FTT1392_at NA 0.000178 -3.49278 transcriptional regulator 2 FTT0566_at NA 0.006825 -2.42471 hypothetical protein 3 hypothetical membrane FTT0474_at NA 0.041498 -2.33965 protein 4 FTT1555c_at rnc 0.00358 -2.29817 Ribonuclease III 5 FTT0241c_at NA 0.029546 -2.29139 hypothetical protein 6 FTT1589c_at NA 0.017243 -2.24881 hypothetical protein 7 hypothetical membrane FTT1178c_at NA 0.046172 -2.22989 protein 8 FTT0222c_at ybgK 0.020229 -2.13904 hydrolase subunit 9 FTT1353_s_at NA 0.020237 -2.12675 hypothetical protein 10 conserved hypothetical FTT1588c_at NA 0.043141 -2.12299 protein, fragment 11 FTT1341_at NA 0.032447 -2.1224 Membrane protein host factor I for 12 FTT0630_at bacteriophage Q beta hfq 0.003258 -2.06301 replication oligopeptide transporter, 13 subunit B, ABC transporter, FTT0123_at oppB 0.050862 -2.05257 membrane protein, pseudoge 14 Uroporphyrinogen III FTT1408c_at hemD 0.034383 -2.01646 synthase 15 FTT1602_at NA 0.045136 -1.98668 hypothetical lipoprotein 16 Peptidase M16 family FTT1322_at NA 0.022932 -1.95221 protein 17 FTT1556c_at lepB 0.00366 -1.86128 signal peptidase I

Continued

Table 2.6: Differentially regulated genes (1.5-fold or greater) when comparing F. novicida ∆FTN_1355 to F. novicida wild-type bacteria by microarray.

63

Table 2.6 continued.

Gene p-value Fold-Change # Probeset ID Gene Title Symbol (PMRA vs. WT) (PMRA vs. WT)

18 ATP-dependent DNA FTT1087c_at rep 0.018276 -1.85921 helicase 19 acetoacetate FTT0930c_at NA 0.030088 -1.84682 decarboxylase, fragment 20 3-demethylubiquinone-9 3- FTT1590c_at ubiG 0.030346 -1.84467 methyltransferase 21 conserved hypothetical FTT0224c_at NA 0.01259 -1.83901 protein, pseudogene 21 conserved hypothetical FTT0224c_at NA 0.01259 -1.83901 protein, pseudogene 22 FTT1439c_at NA 0.047811 -1.83327 Deoxyribonuclease 23 FTT1349_s_at NA 0.030277 -1.79547 hypothetical protein 24 FTT0483c_at NA 0.017848 -1.7869 hypothetical protein 25 FTT1350_s_at NA 0.046461 -1.78672 hypothetical protein 26 FTT0311c_at NA 0.01612 -1.78663 hypothetical protein 27 cystathionine beta-synthase FTT1287_at cbs 0.043546 -1.72224 (cystein synthase) 28 conserved hypothetical FTT1180_at NA 0.042301 -1.71322 protein, pseudogene 29 FTT0857c_at NA 0.040186 -1.70357 hypothetical protein 30 DNA polymerase III, delta FTT0460_at holB 0.021769 -1.69243 prime subunit 31 Type IV pili glycosylation FTT0905_at NA 0.02433 -1.69048 protein 32 hypothetical membrane FTT0297_at NA 0.048225 -1.67945 protein UDP-N-acetylmuramate:L- 33 alanyl-gamma-D-glutamyl- FTT0438_at mpl 0.046074 -1.66681 me so-diaminopimelate ligase 34 tRNA pseudouridine FTT1554c_at truB 0.029192 -1.6653 synthetase B 2,3-bisphosphoglycerate- 35 FTT1329_at independent gpmI 0.023904 -1.66078 phosphoglycerate mutase

Continued

64

Table 2.6 continued.

Gene p-value Fold-Change Probeset ID Gene Title # Symbol (PMRA vs. WT) (PMRA vs. WT)

36 nucleoside permease NUP FTT0115_at nupC1 0.050712 -1.65856 family protein cytochrome d terminal 37 FTT0279c_at oxidase, polypeptide cydA 0.008875 -1.65487 subunit I 38 FTT1476_at galK 0.023395 -1.6407 Galactokinase 39 FTT1793c_at pepN 0.026631 -1.629 Aminopeptidase N DNA polymerase IV, devoid 40 of proofreading,damage FTT0529c_at dinP 0.049245 -1.61038 inducible protein P, pseudogene 41 intracellular growth locus, FTT1357c_s_at iglC 0.015174 -1.57092 subunit C 42 Type IV pili fiber building FTT1314c_at NA 0.041434 -1.56182 block protein 43 FTT0573_at alr 0.05041 -1.52988 alanine racemase 44 Rhodanese-like family FTT1127_at NA 0.021893 -1.5094 protein 45 Phenylalanyl-tRNA FTT1003c_at pheS 0.004603 -1.50392 synthetase, alpha subunit 46 3-oxoacyl-[acyl carrier FTT1373_at fabH 0.040718 -1.50177 protein] synthase III 47 FTT1388_at NA 3.25E-06 5.51723 hypothetical protein 48 Pantoate-beta-alanine FTT1390_at panC 3.33E-08 12.5951 ligase 49 3-methyl-2-oxobutanoate FTT1389_at panB 2.36E-10 13.5862 hydroxymethyltransferase 50 FTT1391_at panD 6.71E-12 14.9131 Aspartate-1-decarboxylase

65

2.4. Materials and Methods

Microarray. Bacterial RNA (10 μg) was prepared according to protocols supplied in the Affymetrix GeneChip Expression Analysis Prokaryotic manual

(Santa Clara, CA). Briefly, TSB-cysteine broth was inoculated with a colony from a 48-h chocolate II agar plate and grown to an optical density at 600 nm of 0.5.

The cells were harvested, and RNA was isolated with the RNeasy kit (QIAGEN,

Valencia, CA). The assay utilizes reverse transcriptase and random hexamer primers to generate DNA complementary to the RNA. The cDNA is then fragmented by DNase I and labeled with terminal transferase and biotinylated

GeneChip DNA labeling reagent at the 3′ termini. The labeled samples were hybridized to a custom GeneChip containing probes to the 1,804 open reading frames of the F. tularensis Schu S4 genome (supplied by Battelle). Washing and staining of the chips with streptavidin-phycoerythrin were performed using the

Affymetrix Fluidics Station 450. Scanning of the chips was done using the

Affymetrix Genechip Scanner 3000.

Microarray data analysis. The Partek Discovery Suite was used to identify changes in gene expression. The CEL files for all samples were loaded into Partek where they were normalized utilizing RMA. Using a filtering criterion of a 1.5-fold or greater change in expression and a P value of <0.05 from analysis of variance, a list of differentially expressed genes was generated.

66

Quantitative real-time PCR (qRT-PCR). Expression analysis was performed as described previously (Mohapatra et al., 2007b). RNA was extracted from bacteria grown in TSB + 0.1% cysteine to mid-log phase (OD600,

0.4 to 0.5) F. novicida (JSG1819), F. novicida ∆pmrA (JSG2845), F. novicida

∆pmrA pKK214pgroEL(pmrA) (JSG2847), and F. novicida ∆pmrA pKK

214pgroEL(pmrA D51A) (JSG3033) bacteria using the RNeasy Kit (QIAGEN,

Valencia, CA). The RNA quality and quantity were determined with the Experion automated electrophoresis system (Bio-Rad, Hercules, CA) and NanoDrop spectrophotometry (NanoDrop products. Wilmington, DE). One microgram of total RNA was reverse transcribed to cDNA with Superscript II RNase Hˉ reverse transcriptase (Invitrogen, Carlsbad, CA) and normalized according to the concentration. Two nanograms of the converted cDNA were used for quantitative PCR with the SYBR green PCR master mixture in the Bio-Rad iCycler apparatus (Bio-Rad, Hercules, CA). Relative quantification was used to evaluate the expression of chosen genes. All primers were designed to give

200- to 220-nucleotide amplicons, which have a G+C range of 30 of 50% and a melting temperature of 58 to 60°C. Relative copy numbers and expression ratios of selected genes were normalized to the expression of two housekeeping genes

(the 16S rRNA gene and dnaK) and calculated as described by Gavrilin et al.

(Gavrilin et al., 2006).

67

CHAPTER 3

The Francisella Orphan Response Regulator PmrA Binds to Regulated Gene

Promoters and Binding is Mediated by Phosphorylation.

3.1. Introduction

Francisella tularensis subspecies tularensis (F. tularensis) is a Gram negative non-motile facultative intracellular pathogen and the causative agent of tularemia. F. tularensis has been extensively researched as a biological weapon and has been designated a category A biothreat agent by the Centers for

Disease Control (CDC). There is a low frequency of tularemia in the United

States, and in those occurring cases, misdiagnosis can lead to a poor prognosis

(Nigrovic and Wingerter, 2008). F. tularensis can be acquired from the bite of an infected arthropod, contact with an infected animal, or ingestion of contaminated food, water or air (Oyston et al., 2004). Different routes of entry can lead to several forms of the disease, with pneumonic tularemia being most serious

(Nigrovic and Wingerter, 2008).

68

The Francisella Pathogenicity Island (FPI) is critical for the ability of this pathogen to cause disease (Santic et al., 2006;Lai et al., 2004). Mutations in the

FPI result in attenuation, inability to escape the phagosome, and deficient replication within macrophages (Nano et al., 2004). MglA, SspA, FevR and

PmrA have been shown to be necessary for Francisella virulence and transcription of the FPI, and MglA and SspA have been shown to bind to RNA polymerase (Charity et al., 2007;Mohapatra et al., 2007b;Lauriano et al.,

2004;Lauriano et al., 2004). How these proteins coordinate regulation of the FPI is not understood.

Thousands of TCS have been described that are employed by Gram- negative bacteria to respond to changes in environmental conditions by modulating gene expression. The components of TCS include a membrane bound sensor kinase and a cytosolic response regulator. The periplasmic domain of the sensor kinase responds to a specific signal which causes a conformational change for the cytosolic portion of the protein. This conformational change results in autophosphorylation of the kinase at a conserved histidine residue. The phospohate residue is transferred to a conserved aspartate residue of the response regulator. Phosphorylation of the response regulator results in a conformational change that increases its affinity for regulated gene promoters. TCS components are traditional under the control of a single promoter (Gunn, 2008). Genome sequencing of virulent strains of

69

Francisella revealed the lack of tandemnly arranged TCS; however, individual or orphaned components were identified (Mohapatra et al., 2007b).

A F. novicida pmrA null mutant is defective for intramacrophage survival and is attenuated in mice. This is likely due to the fact that PmrA regulates its own transcription and that of the FPI (Mohapatra et al., 2007b). In the chapter, we demonstrate that PmrA binds to regulated gene promoters and that, though lacking a linked kinase, DNA binding is enhanced by phosphorylation. In addition, we investigate the site of PmrA phosphorylation, the role of this phosphorylation site in DNA binding, and the effect of this phosphorylation site on intramacrophage replication and virulence.

3.2. PmrA binds to the promoter regions of regulated genes and binding is increased upon treatment with a phosphate donor.

In order to perform DNA binding studies with PmrA, it was necessary to define the promoter regions of PmrA regulated genes. The transcriptional initiation sites for pmrA and pdpD were determined by primer extension.

Fluorescently labeled primers were designed to bind to the RNA 50-100 base pairs downstream of the start codon of the gene of interest. These primers were annealed to F. novicida wild-type RNA and used as a template for a reverse transcription reaction. The genes pmrA and pdpD were chosen for initial analysis

70 as pmrA is positively autoregulated and unaffected by MglA/ SspA, while pdpD transcription requires MglA, SspA and PmrA (Brotcke et al., 2006;Charity et al.,

2007;Mohapatra et al., 2007b). The start of transcription of pmrA was determined to be 40 base pairs upstream of the ATG and pdpD 32 base pairs upstream of its ATG (Figures 3.1 and 3.2). This confirmed the existence of promoters upstream for both of these genes, indicating they are the first genes of their respective operons (Sammons-Jackson et al., 2008;Ludu et al., 2008a).

The primer extension data was used to design PCR primers that would amplify a region of DNA flanking the start of transcription. The pmrA promoter region PCR product is approximately 350 base pairs extending 190 base pairs upstream of the start of transcription. The pdpD probe is 200 base pairs in length extending approximately 128 base pairs upstream of the start of transcription.

Radioactive PCR reactions generated labeled probes that were incubated with purified His-tagged PmrA in electropheretic mobility shift assays (EMSA). The results show that purified His-PmrA binds to the promoter regions of pmrA and pdpD in a dose dependent manner (Figure 3.3). Binding of His-PmrA to the promoter region was interrupted when unlabelled DNA was added as a specific competitor; however, similar amounts of a non-specific competitor (an internal region of the iglC open reading frame) did not affect binding (Figure 3.3).

Phosphorylation of response regulators typically results in an increase in

DNA binding, providing a mechanism for regulation by phosphotransfer. We treated PmrA with acetyl phosphate, which has been shown to phosphorylate 71 response regulators in vitro (McCleary and Stock, 1994). The phosphorylated protein was added to radiolabeled DNA in EMSA. Treatment of PmrA with acetyl phosphate increased its binding to the promoter regions of pmrA and pdpD

(Figure 3.4). Phosphorylation resulted in a 15-20 percent increase versus unphosphorylated PmrA.

Alignment of PmrA with other response regulators indicated that the site of phosphorylation should be at an aspartate residue at or about position 50.

Francisella PmrA has aspartate residues at positions 46 and 51. Mutations in

PmrA D46 and PmrA D51 were constructed and the mutant proteins were purified by His-tag affinity chromatography. The site directed mutant proteins

His-PmrA D51A, His-PmrA D46A, and His-PmrA D46G were evaluated for their ability to bind DNA promoters by EMSA. Purified His-PmrA D51A retains the ability to bind to both the pmrA and the pdpD promoters. However, surprisingly, binding appears to be even more robust for His-PmrA D51A as compared to the same amount of His-PmrA (Figure 3.4). Using lower concentrations of His-PmrA

D51A that did not result in a 100% shift of the pmrA promoter fragment, the addition of acetyl phosphate did not enhance promoter binding (Figure 3.5).

Purified His-PmrA D46A and His-PmrA D46G retained the ability to bind to the pmrA promoter and treatment with acetyl phosphate increase binding to the promoter region (Figure 3.6). Combined this data indicates that the site of PmrA phosphorylation is at D51.

72

3.3. Bt EMSA MglA does not bind to the pmrA or pdpD promoters and does not interact with DNA bound PmrA.

F. novicida mglA was cloned into a His-tag vector, purified and used in

EMSAs. Purified His-tagged MglA was unable to bind to these same promoter regions. As much as 4 µg of His-MglA was added to promoter fragments and no binding was observed (data not shown). When His-MglA and His-PmrA were added to the labeled promoter DNA, no supershift was observed, suggesting that these two proteins do not physically associate under these conditions. This assay, however, did not have other transcription factors or RNA polymerase that might be required for the interaction to occur. Another protein (e.g. SspA) may interact directly with PmrA forming a protein complex that is required for interaction with MglA. Attempts to purify SspA to test this hypothesis have been unsuccessful.

3.4. PmrA may bind to multiple sites in regulated gene promoters.

Typically, response regulators and DNA binding proteins adhere to a consensus binding site(s) that determine their affinity for regions of DNA.

Determination of a response regulator binding site allows in silico identification of regulated promoters. DNA footprinting is often used to identify the DNA recognition site of a DNA binding protein. In this procedure, DNA is bound to a 73 known protein binding partner, and then the complex is incompletely digested with DNase. The fragments are then analyzed to identify the DNA region that was protected from digestion by the bound protein. Attempts to determine the

PmrA DNA binding site have been unsuccessful. A specific region of protection has not been identified comparing His-PmrA bound DNA (pmrA or pdpD promoter regions) to this identical DNA incubated with a non-DNA binding protein

(BSA or His-MglA). Perhaps His-PmrA bound DNA is not protected from DNase digestion, or perhaps the conditions for DNA digestion have not been optimized, though many different variables have been tested.

In an attempt to narrow the location of the PmrA DNA binding site, a series of PCR primers were designed to generate fragments of the promoter probes that bound His-PmrA. Figure 3.7 graphically represents the lengths and locations of probes generated to further investigate the region to which PmrA binds (Figures 3.3-3.6). The only probe that demonstrated the same amount of binding to PmrA as the full length pmrA probe was composed of the nucleotides stretching from base 90 to 350 (Figure 3.7). From this and other probe fragments, it is clear that the first 90 bases of the full length probe are not necessary for His-PmrA binding. Attempts to further narrow the promoter region to which His-PmrA binds were not successful. Full binding was not achieved unless the probe contained two regions: the region stretching from approximately 90-140 base pairs and that from 225-350. This suggests that two regions of the promoter are required for His-PmrA to bind. The results from the

74 pdpD promoter fragments also support this conclusion. His-PmrA did not bind to smaller units of the pdpD promoter fragment; again suggesting that binding is dependent upon two separate regions of DNA.

3.5. Discussion

Primer extension was used to determine the start of transcription for two

PmrA regulated genes: pmrA and pdpD. The pdpD gene is regulated by PmrA,

MglA, and SspA; however, pmrA is positively auto-regulated by PmrA but not by

MglA or SspA. The start of transcription for pdpD was identified as the thiamine

32 bases upstream of its ATG. This identifies the pdpD promoter to be located in the 101 base pair intergenic region between pdpD and FTN1326. Due to the lack of significant intergenic spaces, it is likely that pdpD is the lead gene of an operon including iglABCD. The identical 101 bases are upstream of pdpD in the fully virulent F. tularensis Schu4 strain, indicating the start of transcription and the promoter are likely the same for both species. The start of transcription of pmrA is located 40 bases upstream of its ATG. There is a 441 nucleotide intergenic region upstream of pmrA to the end of FTN1466, the first 324 bases of which are identical in F. tularensis Schu4, suggesting that the pmrA promoter is conserved.

The pmrA operon contains five genes consisting of pmrA, lepB, rnc, truB, and rnr

(Sammons-Jackson et al., 2008).

75

His-PmrA binds to the promoter fragment of both pmrA and pdpD, and phosphorylating His-PmrA with acetyl phosphate increased binding to both fragments. The pmrA promoter region required approximately 700 ng of purified protein to observe a shift in labeled DNA, while the pdpD promoter region required a greater amount (~1400 ng) of protein to visualize binding. This suggests that PmrA has a greater affinity for its own promoter than it does for the promoter of pdpD. EMSA experiments performed with PmrA homologs have used similar amounts of protein to shift labeled promoter fragments (Tamayo et al., 2005b). From scanning the fragments and the likely binding locations on these fragments, we were unable to identify a consensus binding site. The identification and characterization of additional promoters will undoubtedly aid the defining of such a sequence. Further complicating the issue was our finding that smaller, overlapping fragments of the shifted promoters did not bind His-PmrA, suggesting the involvement of multiple, non-tandem binding sites. We also tested the ability of His-MglA to bind to these same promoter fragments, but no binding was observed with as much as 4 µg of purified protein. A supershift was not observed when both purified His-MglA and His-PmrA were used together in these assays. This suggested either that PmrA and MglA do not physically interact or that their physical interaction is dependent upon other molecules (e.g.

SspA, RNA polymerase or FevR). Based on other data gained here and discussed below, we favor the latter hypothesis.

76

Figure 3.1. Primer extension results for pmrA (FTN_1465). Total RNA from F. novicida wild-type bacteria was annealed with 6-FAM-labeled primer complementary to a region of pmrA and extended as described in Material and Methods. Dideoxy sequencing ladders were generated with the same primer and F. novicida wild-type DNA as a template (top panel). Reactions were appropriately diluted and run in a capillary electrophoresis sequencer to correctly estimate the size and position of the 6-FAM-labeled primer extension product (bottom panel). The signal from each chromatographic peak is reported as relative fluorescence units. The bases corresponding to the transcription start on the coding strand is shaded. The start of transcription of pmrA is 40 bases upstream of its ATG.

77

Figure 3.1

78

Figure 3.2. Primer extension results for pdpD (FTN_1325). Total RNA from F. novicida wild-type bacteria was annealed with 6-FAM-labeled primer complementary to a region of pdpD and extended as described in Material and Methods. Dideoxy sequencing ladders were generated with the same primer and F. novicida wild-type DNA as a template (top panel). Reactions were appropriately diluted and run in a capillary electrophoresis sequencer to correctly estimate the size and position of the 6-FAM-labeled primer extension product (bottom panel). The signal from each chromatographic peak is reported as relative fluorescence units. The bases corresponding to the transcription start on the coding strand is shaded. The start of transcription for pdpD is 32 bases upstream from the start of the gene.

79

Figure 3.2.

80

Free probe

Figure 3.3. EMSA: PmrA Binding. Mobility shift assays were used to determine if purified proteins bound to regulated gene promoters. Panel A: His- PmrA binds to the pmrA (12 ng lanes 1-6) and pdpD (12 ng lanes 7-12) in a dose dependent manner. Panel B: His-PmrA binding to the pmrA promoter is affected by specific but not non specific competitor DNA. Each lane contains 12 ng of labeled pmrA promoter. Panel C. His-PmrA binding to the pdpD promoter is affected by specific but not non-specific competitor DNA. Each lane contains 12 ng of labeled pdpD promoter. The blot shown is representative of at least three replicate experiments.

81

Figure 3.4. EMSA: PmrA Phosphorylation. Phosphorylating His-PmrA increases binding to regulated promoters. His-MglA does not bind to either the pdpD or the pmrA promoters. Adding His-PmrA and His-MglA to promoters does not result in a supershift. His-PmrA D51A retains its ability to bind regulated promoters. Lanes 1-9 = 12 ng of labeled pmrA promoter and 1 µg of each indicated protein. Lanes 10-18 = 12 ng of labeled pdpD promoter and 2 µg of each indicated protein. –P indicates the protein was treated with acetyl phosphate. The blot shown is representative of at least three replicate experiments.

82

Figure 3.5. EMSA of His-PmrA D51A binding to pmrA. Phosphorylating His- PmrA increases binding to the pmrA promoter. His-MglA does not bind the pmrA promoter. Adding His-PmrA and His-MglA to promoters does not result in a supershift. His-PmrA D51A retains its ability to bind regulated promoters but binding is not affected by phosphorylation. Lanes contain 12 ng of labeled pmrA promoter and 1 µg of each indicated protein. –P indicates the protein was treated with acetyl phosphate. The blot shown is representative of at least three replicate experiments.

83

Figure 3.6. His-PmrA D46A and His-PmrA D46G retain the ability to bind DNA promoters and that binding is increased with acetyl phosphate treatment. Phosphorylating His-PmrA increases binding to the pmrA promoter. His-MglA does not bind the pmrA promoter. Adding His-PmrA and His-MglA to promoters does not result in a supershift. His-PmrA D46A and His-PmrA D46G retain the ability to bind regulated promoters and the binding is increased by phosphorylation. Lanes contain 12 ng of labeled pmrA promoter and 1 µg of each indicated protein. –P indicates the protein was treated with acetyl phosphate. The blot shown is representative of at least three replicate experiments.

84

pmrA

SOT ATG

0 25 50 75 100 125 150 175 200 225 250 275 300 325 bps

pdpD

SOT ATG

0 25 50 75 100 125 150 175 200 bps

85

Figure 3.7. His-PmrA binding to regulated gene promoters. Graphical representation of His-PmrA binding to regulated gene promoters and pieces of the promoter. Top row for each promoter represents the promoter fragments (pmrA = 300 bps, pdpD = 200 bps) that was used to determine His-PmrA binding by EMSA. Remaining rows indicate smaller promoter fragments that were generated to narrow the region to which His-PmrA binds. Promoter fragments were labeled, incubated with His-PmrA and electrophoresed to observe a shift in migration attributed to protein binding. The top row represents the full length probe that has been used to characterize the interaction of PmrA with regulated promoters examined here. The color of each line represents the level of binding observed when each probe was incubated with His-PmrA in EMSAs. Green represents binding equal to that of the full length probe. Red indicates the PmrA did not bind to the DNA fragment. An orange colored line means that His-PmrA bound to the DNA fragment but not as well as to the full length probe. SOT = start of transcription. ATG = beginning of gene. 85

3.5. Materials and Methods

Protein Purification. The entire mglA gene was amplified by PCR from

F. novicida genomic DNA. The primers (JG1071, 5’

CGGGATCCGAGGATACAATCTTGCTTTTATACAC 3’ and JG1072, 5’

AACTGCAGTTAAGCTCCTTTTGCTTTGATAGT 3’) were engineered to include a BamHI restriction site on the forward primer and a PstI restriction site on the reverse. The PCR products were digested and cloned into the pQE30 His- tagged expression vector (Qiagen, Valencia, CA). His-tagged proteins were purified from E. coli lysates using IMAC native affinity and desalting columns with the Profinia protein purification system (Bio-Rad, Hercules, CA). The concentration of the purified protein was determined using the BCA Protein

Assay Kit (Peirce, Rockford, IL). The purified protein was also analyzed by SDS-

PAGE separation and staining with Gel-Code Blue (Pierce Biotechnology Inc.,

Rockford, IL).

Site Directed Mutagenesis. The site directed mutant PmrA D51A was generated by using a procedure with overlapping PCR primers to create a single base pair substitution that resulted in the desired amino acid conversion. The pKK214pgroEL plasmid carrying pmrA was purified from F. tularensis pmrA complemented strain (JSG2846). Purified plasmid was methylated with CpG methyltransferase (New England Biolabs, Ipswich, MA). The methylated plasmid

DNA was used as a template for PCR with primers JG1412 (5’

86

GTATGATATAGTCGTCTTAGCCATTGGTATGCCAATAAAAAC 3’) and JG1413

(5’ TAAGACGACTATATCATACAATCCAGATTCTATAAAAGTTTGCGC 3’). The resulting PCR products were transformed into F. novicida where the methylated template was degraded, leaving the PCR produced plasmid DNA with the mutated sequence. The constructed pmrA D51A gene was digested from the pKK214 plasmid and cloned into pQE30 His expression vector (Qiagen,

Valencia, CA) using the restriction sites BamHI and PstI. The pQE30 [pmrA

D51A] plasmid was maintained in E. coli JM109.

Primer Extension. These studies were performed essentially as described previously (Li et al., 1999). FAM labeled primers (JG1514, 5’ FAM –

GCCTTCACCCAAATGAAGATC 3’ for pmrA and JG1515, 5’ FAM –

TAGCCATGACATCCATCGTTT 3’ for pdpD) were designed to bind downstream of the ATG of the gene of interest. The primer was bound to a PCR product that contained the fluorescent probe binding site and the predicted location of the promoter. The fluorescently labeled fragment was then sequenced. One nanomole of the fluorescently labeled primer was also bound to 50 µg of RNA and used as a template to generate an ssDNA using SuperScript III reverse transcriptase (Invitrogen, Carlsbad, CA). Denatured single-stranded DNAs were analyzed in an ABI 3770 capillary electrophoresis sequencer. The length of the fluorescently labeled ssDNA fragment is equal to the distance from the primer to the start of transcription. The fluorescent signal from the sequence reaction and

87 the ssDNA reaction were aligned to determine the exact base at which transcription was initiated.

EMSA. Promoter regions for pmrA and pdpD were amplified and labeled by PCR from F. novicida wild-type genomic DNA. Standard PCR conditions were used and the reactions were spiked with [γ-32P] dATP. Primer pairs

JSG1508 (5’ AAAAGTTTGATGTAACTTTAGAAAACATTTTCA 3’) and JSG1463

(5’ ATAAAAGTTTGCGCTGCCTCACCA 3’) were used to amplify the pmrA promoter region. Primers JG1510 (5’ GCAACCGGAGCAAAAAGTAG 3’) and

JG1511 (5’ GAGGTCATCAGTATCATATAATAAATCGTT 3’) were used to amplify the pdpD promoter region. Non-radioactive control reactions were used to estimate the concentration and purity of the PCR products by NanoDrop spectrophotometry and gel electrophoresis followed by staining with ethidium bromide. Purified His-tagged proteins were added to 12 ng of labeled DNA and incubated for 30-40 minutes at room temperature in binding buffer (0.1 mM DTT,

2 mM MgCl2, 1% glycerol, 0.2 mM EDTA, 20 mM KCl and 2 mM Tris-HCl pH 7.5 and 1 µg poly[d(I-C)]). Agarose loading buffer was added to each sample, then electrophoresed on pre-run 5% acrylamide TBE gels at 20 milliamps. Gels were dried and the DNA was detected by autoradiography. Purified protein was phosphorylated by incubating with acetyl phosphate (40 mM acetyl phosphate,

50 mM Tris-HCl, 20 mM MgCl2 and 0.1 mM DTT) for 30-40 minutes at 37°C prior to adding to the DNA.

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CHAPTER 4

PmrA Aspartate 51 is the Site of Phosphorylation and is Essential for Francisella Intramacrophage Replication and Virulence.

4.1. Introduction

Intracellular pathogens replicate within host cells, and in so doing interrupt the normal maturation of the phagasome along the endocytic pathway. F. tularensis halts lysosomal fusion and then mediates degradation of the phagosomal membrane to escape and replicate within the cytosol. The

Francisella pathogenicity island (FPI) has been shown to be required for escape from the phagosome and intramacrophage replication (Nano et al.,

2004;Clemens et al., 2004). The functions of the proteins encoded with this genomic island are unknown, but homology suggests that they are a part of a type six secretion system (Ludu et al., 2008b). Transcription of the FPI is upregulated during infection of host cells (Brotcke et al., 2006;Santic et al.,

2005). Four FPI transcription factors have been identified: MglA, SspA, FevR, and PmrA.

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MglA and SspA are homologous and regulate similar sets of genes, including the FPI. Additionally, both of these proteins bind to RNA polymerase

(Charity et al., 2007). How MglA and SspA coordinate expression of virulence determinants is not understood. Recent work has established FevR as a regulator of the FPI and is itself regulated by MglA and SspA. Because the FevR regulon is different than the MglA/ SspA regulon, it is believed that these regulatory elements operate in parallel to affect transcription of the FPI. PmrA also regulates the FPI but not MglA or SspA, and MglA/ SspA do not affect expression of Pmra. PmrA is homologous to two component system response regulators.

Two component systems (TCS) are a means for Gram negative bacteria to monitor the extracellular milieu and respond by modulating gene expression.

TCS are composed of a membrane-bound histidine kinase and a response regulator that is free in the cytosol. The system is regulated by phosphorylation.

The periplasmic portion of the histidine kinase reacts to environmental signals by undergoing a conformation change that results in autophosphorylation at a conserved histidine residue on the cytosolic face of the protein. The phosphate group is available for transfer to a conserved aspartate residue in the receiver domain of the response regulator. Response regulators are typically DNA binding proteins that target DNA promoters. Phosphorylation of the response regulator results in increased affinity for regulated gene promoters.

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PmrA is homologous to response regulators described in Escherichia coli and Salmonella enteric. We have shown that PmrA is required for F. tularensis replication in host phagocytes and therefore virulence (Mohapatra et al., 2007b).

EMSA data demonstrates that PmrA is a DNA binding protein and that phosphorylation mediates binding to regulated gene promoters. Here we investigate the site of PmrA phosphorylation and its importance to F. tularensis’ regulation of virulence. Several thousand TCS systems have been described from over a hundred different species of bacteria. Some Gram negative bacterial pathogens have dozens of TCS that respond to a variety of environmental signals. Due to the conserved nature of TCS, these systems cross-talk, creating a complicated network of phosphorelays that interact to modulate gene expression to survive in varying environmental conditions. Cross-talk between

TCS is possible because histidine kinases can phosphorylate multiple response regulators.

4.2. PmrA is phosphorylated at aspartate 51.

By amino acid alignment with other characterized response regulators, it was determined that the site of PmrA phosphorylation was an aspartate at either position 46 or 51. To determine which aspartate is the site of phosphorylation, we again utilized the site-directed mutant proteins, His-PmrA D51A and His-

PmrA D46A. The enteric histidine kinase CheA has been used to phosphorylate 91 other response regulators in vitro (Whitchurch et al., 2002). The ability of CheA to phosphorylate His-PmrA,His-PmrA D51A, and His-PmrA D46A was compared by phospho-transfer experiments. CheA was autophosphorylated by incubation with gamma labeled 32P ATP. CheA was able to phosphorylate His-PmrA and

His-PmrA D46A but not His-PmrA D51A (Figure 4.1A). A duplicate SDS-PAGE gel stained for total protein showed equal amounts of His-PmrA and His-PmrA

D51A, ruling out the possibility that PmrA D51A was degraded during the reaction (Figure 4.1B). It is important to note that His-PmrA D51A runs slightly faster on SDS-PAGE than His-PmrA even though the gene sequence is identical to pmrA except for the codon 51. His-PmrA D46A was phosphorylated less than

His-PmrA; however, there was less His-PmrA D46A added to the reaction, as indicated by the total protein staining (Figure 4.1B). Subsequent total protein determination confirmed that the purified His-PmrA D46A preparation had degraded over time in storage thus explaining why less intact protein was present when equal concentrations (from BCA assays) were thought to be added. EMSA results showed that His-PmrA D46A responded to acetyl phosphate treatment while His-PmrA D51A did not (Figure 3.5 and 3.6).

Together, this data indicates that PmrA is phosphorylated at D51.

To determine if PmrA is phosphorylated by Francisella kinases, we prepared a membrane fraction of F. novicida wild-type bacteria. This membrane fraction was incubated with His-PmrA or His-PmrA D51A in the presence of gamma labeled 32P ATP. Phosphorylation was detected after separation by

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SDS-PAGE. His-PmrA was phosphorylated while the mutated D51A protein was not (Figure 4.2B, lanes 1 and 2). Duplicate reactions were run using unlabeled

ATP and probed for PmrA by Western blot to confirm that neither His-PmrA nor

His-PmrA D51A were degraded during the reaction (Figure 4.2C, lanes 3 and 4).

Finally, non-radioactive duplicate reactions were run and probed with an anti-F. novicida antibody to assess that equal amounts of membrane fractions were added to each reaction (Figure 4.2A). This data, combined with the results from the CheA phosphorylation experiment, demonstrates the site of PmrA phosphorylation is aspartate 51.

4.3. PmrA D51 is important for gene regulation.

If phosphorylation of PmrA increases binding to regulated promoters, it should have a positive effect on the expression of regulated genes. Similarly, if the site of phosphorylation (D51) is removed, then the expression of regulated gene promoters should decrease. To test if phosphorylation of PmrA is important for expression of regulated genes, we performed qRT-PCR for pmrA and iglC in

F. novicida ∆pmrA(JSG2845), F. novicida ∆pmrA complemented with pmrA carried on pKK214pgroEL (JSG2847), and F. novicida ∆pmrA complemented with pmrA D51A on pKK 214pgroEL (JSG3033). As expected, expression of pmrA and iglC was almost undetectable in the ∆pmrA (JSG2845), and transcript levels were much higher in the ∆pmrA complemented strain (JSG2847). The 93

∆pmrA strain complemented with PmrA D51A (JSG3033) had decreased expression of both pmrA and iglC compared to the strain complemented with native PmrA (JSG2847) with iglC being dramatically more reduced (Figure 4.4).

This data indicates that expression of PmrA-regulated genes is dependent upon the aspartate at position 51 and therefore, phosphorylation of PmrA is important for gene regulation.

4.4. PmrA D51 is required for intramacrophage growth.

Host macrophages have been widely described as a site of replication for francisellae (Ellis et al., 2002;Clemens et al., 2005;Fortier et al., 1994;Sjostedt,

2006). Previous data demonstrated that F. novicida ∆pmrA (JSG2845) is defective for replication within macrophages (Mohapatra et al., 2007b). To determine the importance of phosphorylation of PmrA on Francisella replication within macrophages, we infected PMA-induced THP-1 cells with F. novicda wild- type, F. novicida ∆pmrA (JSG2845), F. novicida ∆pmrA complemented with native PmrA (JSG2847), and F. novicida ∆pmrA complemented with PmrA D51A

(JSG3033). As expected, the wild-type and native complement strains were capable of replication within THP-1 macrophages, increasing by more than two logs at 24 hours post-infection. Conversely, the ∆pmrA strain and the strain complemented with the D51A mutant were defective for replication within macrophages. Some replication was observed for the D51A-complemented 94 strain, but at 24 hours post-infection the increase was less than one half log

(Figure 4.5). These data indicate that PmrA aspartate 51 and likely phosphorylation of PmrA are important for replication of F. novicida within macrophages.

4.5. PmrA D51 is required for mouse virulence.

Since PmrA D51 is required for intramacrophage survival, and replication within macrophages is closely tied to virulence, this residue may also be important for virulence. The lethality of F. novicda wild-type, F. novicida ∆pmrA

(JSG2845), F. novicida ∆pmrA complemented with native PmrA (JSG2847), and

F. novicida ∆pmrA complemented with PmrA D51A (JSG3033) was compared.

Groups of BALB/C mice were infected via the intranasal route. One thousand wild-type bacteria resulted in 100% lethality within four days while no mice died after receiving 1000 CFU of the pmrA mutant complemented with PmrA D51A.

Another group of mice was infected with 106 D51A complemented bacteria.

Eighty percent of these mice were killed within ten days of infection (Figure 4.6).

Our previous report showed that the F. novicida ∆pmrA strain was completely attenuated in mice at a dose of 108 bacteria delivered intranasally (Mohapatra et al., 2007b). Therefore, the PmrA D51A-complemented strain is attenuated, but not to the extent of a strain that is completely lacking PmrA. This data indicates

95 that D51 and likely phosphorylation of PmrA are important for F. novicida virulence.

4.6. PmrA is a primary target of the histidine kinase KdpD.

The aspartate at position 51 of PmrA is important for survival within macrophages and virulence of F. novicida. Though we have demonstrated that this amino acid was the site of phosphotransfer from CheA and F. novicida membrane extracts, the kinase responsible for phosphorylating PmrA at D51 has not been identified. Transposon mutants of the three putative histidine kinase genes identified (http://go.francisella.org/cgi-bin/frangb/genomelist.cgi) in the F. novicida genome: FTN1453 (JSG2890), FTN1617 (qseC, JSG2892), and

FTN1715 (kdpD, JSG2894)(Gallagher et al., 2007) were used to attempt to identify which kinase is responsible for phosphorylation of PmrA. Membrane fractions from these mutants were incubated with gamma labeled 32P-ATP and either His-PmrA or His-PmrA D51A. Phosphorylated proteins were separated on an SDS-PAGE gel and detected by autoradiography. Duplicate reactions were run without His-PmrA and His-PmrA D51A as a control to identify phosphorylated proteins unrelated to PmrA (Figure 4.7). His-PmrA was phosphorylated, to some degree, by each of the membrane fractions. According to densitometry readings, the FTN1453 mutant membrane fraction phosphorylated His-PmrA twice as much as F. novicida wild-type membranes, while the qseC mutant 96 phosphorylated His-PmrA the same as wild-type. The kdpD mutant membrane preparations phosphorylated His-PmrA only 20 percent of the level of wild-type membranes. This suggested that the kinase primarily responsible for phosphorylation of PmrA was the histidine kinase KdpD (Figure 4.2B, lanes 3-5).

Duplicate reactions were run with cold ATP and analyzed by Western blot with anti-PmrA anti-sera to confirm that neither His-PmrA nor His-PmrA D51A were degraded during the reaction and that equal amounts of target proteins were present (Figure 4.2C, lanes 5-7).

Further phosphorylation studies were conducted to determine the relative amount of PmrA phosphorylation in different Francisella strains. Although there are no typical tandemly arranged two component systems in virulent francisellae, there is one in F. novicida comprised of KdpD and KdpE. We hypothesized that in the absence of KdpE, phosphorylation of PmrA by KdpD would be increased.

Indeed, using a membrane fraction from a KpdE (FTN_1714) transposon mutant

(Gallagher et al., 2007), His-PmrA was phosphorylated 14 times more than by F. novicida wild-type (Figure 4.3). The fully virulent F. tularensis Schu4 strain does not have homologs to either kdpE or FTN_1453; therefore, phosphorylation of

PmrA should be higher in this strain. When His-PmrA is mixed with an F. tularensis Schu4 membrane preparation in the presence of radiolabeled ATP, it is phosphorylated 50 times more compared to that observed with F. novicida wild-type membranes (Figure 4.3).

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4.7. Discussion

By amino acid alignment with known response regulators from other Gram negative bacteria, the PmrA phosphorylation site was predicted to be D46 or

D51. His-PmrA D46A and His-PmrA D51A mutant proteins were constructed to test this prediction. We demonstrated that His-PmrA and His-PmrA D46A but not

His-PmrA D51A are phosphorylated by the enteric histidine kinase CheA and F. novicida wild-type membrane fractions. The His-PmrA D51A mutant still bound

DNA, demonstrating that the substitution did not result in a non-functional protein. The role of the putative histidine kinases in the Francisella genome in phosphorylating PmrA was also investigated. This data indicated that KdpD is the histidine kinase primarily responsible for phosphorylating PmrA; however, there appeared to be some cross talk or target promiscuity as His-PmrA was phosphorylated to a small degree in the absence of KdpD. Membrane fractions missing the putative sensor kinase FTN_1453 resulted in increased PmrA phosphorylation when compared to wild-type, suggesting that it regulated, or acted itself, as a phosphatase. Interestingly, F. tularensis Schu4 has no

FTN1453 homolog. This result supports the microarray studies that suggested that both KdpD and FTN_1453 are involved in the regulation of PmrA.

Phosphorylation of PmrA affects the expression of PmrA-regulated genes.

The genes iglC and pmrA are downregulated in a pmrA null strain complemented with pmrA D51A compared to a strain complemented with pmrA. This

98 demonstrates the importance of phosphorylation for expression of the PmrA regulon. Since KdpD is the kinase most affecting PmrA phosphorylation, it is reasonable to hypothesize that the PmrA regulon will be downregulated in a

KdpD mutant. In support of this, microarray analysis comparing a kdpD mutant to the wild-type strain demonstrated KdpD-dependent regulation of FPI genes and the pmrA operon (Table 2.4).

Growth of a pmrA null strain complemented with pmrA D51A within macrophages was diminished as compared to a strain complemented with native pmrA; however, some replication of the pmrA D51A-complemented strain was observed as compared to a pmrA null mutant. Similarly, the PmrA D51A- complemented strain was more virulent in the mouse model than strains devoid of PmrA. This is likely a result of PmrA D51A overexpression, such that increased copy number coupled with the residual PmrA D51A DNA binding resulted in modest expression of the PmrA regulon, resulting in some replication within macrophages and retention of virulence.

Data presented thus far have demonstrated that PmrA is a DNA binding protein whose binding to regulated gene promoters is enhanced upon phosphorylation. In this chapter, the results show that the site of PmrA phosphorylation is at aspartate 51. In addition to this, expression analysis combined with intramacrophage replication and mouse virulence assays determined that D51 is critical for the ability F. novicida to cause disease.

Phosphotransfer data identified KdpD as the histidine kinase most likely 99 responsible for phosphorylating PmrA. This establishes KdpD and PmrA as a functional TCS that is implicated in regulating transcription of the pdpD operon that encodes known virulence factors. In this system, KdpD would respond to an environmental signal by transferring a phosphate to PmrA which increases binding to the regulated gene promoters, specifically pmrA and pdpD. This would up-regulate expression of the genes within these operons including member of the putative T6SS contained in the FPI. Other virulence factors

(MglA, SspA) have been described that are required for transcription of the FPI.

MglA and SspA bind to one another and to RNA polymerase. This suggests that

PmrA binds to regulated promoters and interacts with MglA and/ or SspA which recruit RNA polymerase to the site to initiate transcription of the FPI. This model of regulation is dependent upon PmrA physically interacting with MglA and/or

SspA.

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Figure 4.1. Phosphotransfer from CheA. Panel A: PmrA and PmrA-D46A but not PmrA D51A are phosphorylated by the enteric histidine kinase CheA. Each lane consists of 0.1 ng phosphorylated CheA and 1 µg His-PmrA, His-PmrA D51A, or His-PmrA D46A. Panel B: Duplicate reactions were stained for total protein to demonstrate that neither His-PmrA nor His-PmrA D51A was degraded during the reaction. Less intact His-PmrA D46A purified protein was added to the reaction, resulting in a lower phosphorylation signal. The blot shown is representative of at least three replicate experiments.. 101

P-PmrA

Figure 4. 2. Phosphotransfer from membrane fractions. Panel A: Western blot analysis with an anti-F. novicida antibody on duplicate phospho-transfer reactions. Panel C: Western blot analysis for PmrA of duplicate phospho- transfer reactions. Panel B: Francisella membrane fractions phosphorylate His- PmrA but not His-PmrA D51A. The putative histidine kinase KdpD is primarily responsible for phosphorylating His-PmrA. Each lane consists of 5 µg of membrane fractions and 1 µg His-PmrA or His-PmrA D51A. P = phosphorylated. The blot shown is representative of at least three replicate experiments.

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Figure 4.3. Phosphotransfer from membrane fractions. His-PmrA is phosphorylated more by ∆kdpE mutant and F. tularensis Schu4 membrane fractions. In each lane: 5 µg of membrane fractions (top labels) and 1 µg His- PmrA. The blot shown is representative of at least three replicate experiments.

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Figure 4.4. qRT-PCR analysis of PmrA-regulated genes in F. novicida ∆pmrA complemented with PmrA or PmrA D51A. Complementation of F. novicida ∆pmrA with pmrA results in restored expression of pmrA and iglC while complementation with pmrA D51A results in decreased expression. Black bars = pmrA. Grey bars = iglC. Error bars represent the standard deviation of three replicate experiments.

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Figure 4.5. PmrA D51A is important for intramacrophage replication. PmrA D51 is required for replication within macrophages. Closed circles = F. novicida wild-type. Open circles = F. novicida ∆pmrA. Open squares = F. novicida ∆pmrA pKK214pgroEL [pmrA]. Closed squares = F. novicida ∆pmrA pKK214pgroEL [pmrA D51A]. Error bars represent the standand deviation of three replicate experiments.

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Figure 4.6. PmrA D51A is important for mouse virulence. PmrA D51 is required for full virulence of F. novicida in mice. Closed circles = 1 x 103 F. novicida ∆pmrA pKK214pgroEL [pmrA]. Closed squares with solid line = 1 x 103 F. novicida ∆pmrA pKK214pgroEL [pmrA D51A]. Closed squares with dashed line = 1 x 106 F. novicida ∆pmrA pKK214pgroEL [pmrA D51A]. This experiment was performed once with five mice per group.

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Figure 4.7. Phosphotransfer control reaction. F., novicida wild-type and histidine kinase transposon mutant membrane preparations were incubated with and without His-PmrA in the presence of [γ-32P] ATP. His-PmrA was identified as the phosphorylated protein present at approximately 25 Kd but absent in reactions that did not contain His-PmrA. The blot shown is representative of at least three replicate experiments.

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4.8. Materials and Methods

Protein Purification. Materials and methods are the same as that presented in Chapter 3.

Site Directed Mutagenesis. The site-directed mutant PmrA D46A was generated by using a procedure identical to that described in Chapter 3 using

PCR primers designed to create the D46 to A mutation: JG1818 (5’

TACAGAATACTACTTTAAATAGCAAGGGTT 3’) and JG1820 (5’

GCTTCTAATAAGCCTTCACCC 3’).

Phosphotransfer with CheA. The conditions used for autophosphorylation and phosphotransfer from CheA were previously described

(Whitchurch et al., 2002). Autophosphorylation of CheA was performed with 50

µCi of [γ-32P] ATP at room temperature in 10 µl of phosphorylation buffer (100 mM Tris-HCL [pH 7.4], 5 mM MgCl2, 50 mM KCl) for 1 hour. Phosphotransfer was performed with 1 µg of purified His-PmrA or His-PmrA D51A with 100 ng of autophosphorylated CheA in phosphorylation buffer at 37°C for 1 hour. The reaction was stopped with an equal volume of Laemmli sample buffer (Bio-Rad,

Hercules, CA) plus 0.5% β-mercaptoethanol (β-ME). Radiolabelled products were separated on a 12.5% SDS-PAGE gel and detected by autoradiography.

Duplicate reactions with non-radioactive ATP were electrophoresed through a

12.5% SDS-PAGE gel and proteins were detected by staining with GelCode Blue

(Peirce, Rockford, IL). 108

Phosphotransfer from bacteria membrane fractions.

Phosphostransfer from membrane fractions was performed as described by

Gunn et. al (Gunn et al., 1996). F. novicida wild-type (JSG1819), F.novicida

∆FTN1453 (JSG2890), F. novicida ∆FTN1617 (qseC, JSG2892), F. novicida

∆FTN1715 (kdpD, JSG2894), F. novicida ∆FTN1714 (kdpE, JSG2893), and F. tularensis Schu4 strains were grown in 100 ml of TSB + 0.1% cysteine to an

OD600 of 1.0. Bacteria were harvested by centrifugation at ~9,000 x g for 15 minutes at 4°C. Pellets were resuspended in 5 mL of Bugbuster protein extraction buffer (Novagen , Madison, WI) and sonicated for 2 minutes in 10 second bursts at an output of 50 using a Sonics VibraCell (Sonics and Materials,

Inc., Newtown, CT). Lysates were centrifuged at 100,000xg for 1 hour at 4°C.

The pellet was resuspended in 10% glycerol. Total protein was measured using the BCA Protein Assay Kit (Peirce, Rockford, IL) and the NanoDrop spectrophotometer. Membranes (5 µg), His-PmrA or His-PmrA D51A (1µg), and

[γ-32P] ATP (3 pmol) were incubated at 37°C for 1 hour. The reaction was stopped with an equal volume of Laemmli sample buffer (Bio-Rad, Hercules, CA) plus 0.5% β-ME. Radiolabelled proteins were separated on a 12.5% SDS-PAGE gel and detected by autoradiography or exposed to a phosphoscreen and imaged with a Typhoon Variable Mode Imager (GE Healthcare, Pittsburgh, PA).

Duplicate control reactions with an equal amount of non-radioactive ATP were analyzed by Western blotting. Growth and lysis of Type A Schu S4 strain was performed in CDC-approved BSL3/aBSL3 suites at The Ohio State University.

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Quantitative real-time PCR (qRT-PCR). Expression analysis was performed as described previously (Mohapatra et al., 2007b). RNA was extracted from mid-log phase (OD600, 0.4 to 0.5) F. novicida (JSG1819), F. novicida ∆pmrA (JSG2845), F. novicida ∆pmrA pKK214pgroEL(pmrA)

(JSG2847), and F. novicida ∆pmrA pKK 214pgroEL(pmrA D51A) (JSG3033) bacteria using the RNeasy Kit (QIAGEN, Valencia, CA). The RNA quality and quantity were determined with the Experion automated electrophoresis system

(Bio-Rad, Hercules, CA) and NanoDrop spectrophotometry (NanoDrop products.

Wilmington, DE). One microgram of total RNA was reverse transcribed to cDNA with Superscript II RNase Hˉ reverse transcriptase (Invitrogen, Carlsbad, CA) and normalized according to the concentration. Two nanograms of the converted cDNA were used for quantitative PCR with the SYBR green PCR master mixture in the Bio-Rad iCycler apparatus (Bio-Rad, Hercules, CA). Relative quantification was used to evaluate the expression of chosen genes. All primers were designed to give 200- to 220-nucleotide amplicons, which have a G+C range of 30 of 50% and a melting temperature of 58 to 60°C. Relative copy numbers and expression ratios of selected genes were normalized to the expression of two housekeeping genes (the 16S rRNA gene and dnaK) and calculated as described by Gavrilin et al. (Gavrilin et al., 2006).

Intramacrophage survival assay. Gentamicin protection assays were performed as previously described (Mohapatra et al., 2007b). F. novicida

(JSG1819), F. novicida ∆pmrA (JSG2845), F. novicida ∆pmrA

110 pKK214pgroEL(pmrA) (JSG2847), and F. novicida ∆pmrA pKK 214pgroEL(pmrA

D51A) (JSG3033) were used to infect phorbol myristate acetate (PMA)-induced

(10 ng/ml) THP-1 cells, a human macrophage-like cell line, at a multiplicity of infection of 50:1. Wells were seeded with 2 x 105 macrophages and 1.0 x 107 bacteria were added to each well. After 2 hours of incubation at 37°C and 5%

CO2, gentamicin (50 µg/ml) was added to the medium to eliminate extracellular organisms. After 30 minutes, wells were washed twice with PBS and incubated with their respective media supplemented with 10 µg/ml gentamicin. The macrophages were lysed with 0.1% sodium dodecyl sulfate (SDS) at 2 h, 12 h, and 24 h postinfection, and the lysates were serially diluted in PBS and plated on chocolate II agar plates (BD, Franklin Lakes, NJ) for determination of viable counts.

Mouse survival studies. Virulence experiments were performed as described in Mohapatra et. al.(Mohapatra et al., 2007b). Francisella strains were given intranasally to groups of five anesthetized female 4 to 6 week-old BALB/c mice (Harlan Sprague, Indianapolis, IN) at a dose of 1 x 103 to 1 x 106 CFU/20 µl

PBS. Actual bacterial counts delivered were determined by plate counts from the inoculum. Mice were monitored for 5 weeks postinfection.

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CHAPTER 5

PmrA Interacts with other Francisella Transcription Factors

5.1. Introduction

Tularemia is caused by F. tularensis, a Gram negative intracellular bacterial pathogen that invades and replicates within host phagocytes (Baron and Nano, 1998). This pathogen was extensively studied by multiple state sponsored biological weapons programs. In fact, in the United States, F. tularensis was produced at large scale and deployed in tactical weaponry ready for use (2007). For these reasons F. tularensis has been categorized as a

Category A biothreat agent by the CDC. There are several forms of the disease tularemia, that are determined by the route of entry of the bacteria and the infective dose (Nigrovic and Wingerter, 2008;Oyston et al., 2004). Replication with host phagocytes is a critical aspect of disease. Mutants that cannot replicate within macrophages are not capable of causing disease (Weiss et al.,

2007;Ludu et al., 2008b). Expression of the Francisella pathogenicity island

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(FPI) is crucial for bacterial escape from the phagosome, intramacrophage survival and virulence (Nano et al., 2004). Transcription of the FPI is dependent on PmrA, MglA, SspA and FevR, but how these proteins interact to mediate expression of these virulence factors is not understood (Mohapatra et al.,

2007b;Brotcke and Monack, 2008;Baron and Nano, 1998;Charity et al., 2007).

There are four subspecies of Francisella tularensis (tularensis, holarctica, mediasiatica, and novicida) and all are very closely related. Type A strains (F. tularensis) are extremely infectious (infectious dose less than 10 CFU), while F. novicida has caused disease only in immunocompromised individuals; however, these subspecies share greater than 99 percent DNA homology (Keim et al.,

2007). This suggests that gene regulation rather than gene content is responsible for strain differences in virulence.

F. novicida mglA null mutants are attenuated in the mouse model (Santic et al., 2005b;Lauriano et al., 2004;Guina et al., 2007;Brotcke et al., 2006;West et al., 2008). MglA is upregulated upon infection of macrophages (Baron and Nano,

1998;Lauriano et al., 2004;Santic et al., 2005b) and an mglA mutant escape poorly from the macrophage phagosome (Santic et al., 2005b). MglA is known as a master regulator of transcription in Francisella, as it is responsible for regulating approximately 100 genes including the genes within the FPI (Brotcke et al., 2006;Lauriano et al., 2004). MglA and its homolog SspA have been shown to interact with one another and to bind RNA polymerase. (Charity et al., 2007).

SspA homologs in E. coli bind to the alpha subunit(s) of RNA polymerase and

113 participate in gene regulation (Hansen et al., 2003). Unlike other Gram negative bacteria, F. tularensis subspecies have two distinct RNA polymerase alpha subunits. The existence of different alpha subunits may explain why F. tularensis species have two SspA homologs while other bacteria encode only one (Charity et al., 2007). SspA, like MglA, is also required for transcription of the FPI and is maximally expressed during stationary phase while MglA is maximally expressed during lag and exponential phages of growth in vitro (Brotcke et al., 2006;Baron and Nano, 1998). MglA and SspA cooperate to positively regulate virulence factors.

5.2. PmrA co-precipitates with MglA and SspA.

PmrA, MglA, SspA and FevR are required for Francisella virulence and regulate the FPI (Brotcke et al., 2006;Charity et al., 2007;Baron and Nano,

1998;Lauriano et al., 2004;Brotcke and Monack, 2008). FevR has poor homology to DNA binding proteins and appears to act upstream of MglA, SspA, and PmrA RNA polymerase binding (Brotcke and Monack, 2008). One hypothesis to explain the involvement of PmrA, MglA, and SspA is that PmrA binds to regulated promoters and interacts with MglA and/or SspA, which in turn recruit RNA polymerase to the site (or interacts with MglA and/or SspA bound to

RNA polymerase), thereby initiating transcription of regulated genes. This model predicts PmrA physically interacts with MglA and/or SspA. We showed earlier 114 that adding MglA and PmrA together did not result in a EMSA supershift, which would be an indication of an interaction between the two proteins (Figure 5.1).

This in vitro assay does not contain many of the components that would be present in vivo (e.g. SspA and RNA polymerase), so we attempted to co- precipitate PmrA with MglA and SspA from whole cell lysates. Soluble fractions were prepared from lysates of F. novicida ∆pmrA (JSG2845), F. novicida ∆mglA

(JSG2250), and F. novicida His-SspA (JSG2970). His-PmrA was added to the soluble fraction of the F. novicida ∆pmrA (Figure 5.1, lane 1). Once precipitated with His-affinity resin, PmrA and associated proteins were detected by Western blotting. PmrA was detected by an anti-PmrA anti-sera (Figure 5.1, panel A, lane

1) and by an anti-His antibody (Figure 5.1, panel C, lane 1). MglA was detected in the PmrA precipitated sample by anti-MglA anti-sera (Figure 5.1, panel B, lane

1), showing that MglA co-precipitates with PmrA. Similarly, His-MglA was added to the soluble fraction from F. novicida ∆mglA. Precipitation with His-affinity resin allowed detection of MglA using an anti-MglA anti-sera (Figure 5.1, panel B, lane

2) and the anti-His antibody (Figure 5.1, panel C, lane 2). PmrA precipitated with

MglA and was detected with anti-PmrA anti-sera (Figure 5.1, panel A, lane 2).

This also demonstrates that PmrA and MglA co-precipitate. Finally SspA was precipitated from the soluble fraction of F. novicida His-SspA and detected with the anti-His antibody (Figure 5.1, panel C, lane 3). PmrA was present in the

SspA precipitated sample (Figure 5.1, panel A, lane 3), as was MglA (Figure 5.1, panel B, lane 3). Using the identical protocol without adding His-tagged proteins

115 did not result in immunoprecipitation of PmrA, MglA or SspA. Although FevR is required for regulation of the FPI, it does not co-precipitate with MglA, SspA or

PmrA (data not shown and (Brotcke and Monack, 2008)). These data indicate that PmrA, MglA and SspA physically interact with one another.

5.3. MglA co-precipitates with PmrA from a F. tularensis Schu4 lysate.

The data presented shows that PmrA, MglA and SspA co-precipitate from

F. novicida whole cell lysate soluble fractions. F. novicida serves as a good model organism for tularemia since it is very closely related to fully virulent F. tularensis strains like Schu4 and causes a tularemia like disease in the murine model. To confirm that our model of FPI regulation also applies to virulent strains, co-precipitation was attempted from an F. tularensis Schu4 lysate. The

F. tularensis Schu4 strain was grown and harvested from broth and then lysed.

A soluble fraction was prepared from the lysates by centrifugation at 100,000 x g.

Purified His-tagged PmrA was added to the soluble fraction and then precipitated using His-bind resin. PmrA was detected in the imunnoprecipitate using an anti-

PmrA polyclonal anti-sera and an anti-His monoclonal antibody (Figure 5.2, lane

4, panels A and C). It was interesting to note that His-PmrA and native PmrA were both detected by the anti-PmrA antibody when PmrA was precipitated from the Schu4 soluble fraction (Figure 5.2, lane 4, panel A). MglA precipitated with

His-PmrA as it was detected using anti-MglA polyclonal anti-sera (Figure 5.2, 116 lane 4, panel B). This result shows that PmrA and MglA interact in F. tularensis

Schu4 as well as F. novicida and suggests that the PmrA, MglA complex is also present in the virulent strain.

5.4. Discussion

Co-immunoprecipitation experiments reported here showed that PmrA,

MglA, and SspA physically interact with one another. The data was obtained using the model organism F. novicida, but the amino acid sequence of MglA and

SspA are nearly identical between F. novicida and the virulent strain F. tularensis

Schu4. The PmrA protein sequence is identical between these subspecies. In addition to the F. novicida data presented, PmrA also co-precipitates with MglA from a F. tularensis Schu4 whole cell soluble fraction (Figure 5.2). The protein interactions, combined with the promoter binding behavior of PmrA, suggest a model of gene regulation. DNA bound PmrA interacts with MglA and/ or SspA to regulate expression of gene transcription. The data presented, however, do not indicate whether PmrA binds to MglA, SspA, or to both proteins. We predict RNA polymerase is a part of this complex; however, other binding partners/ complex components may also exist. The sequence of protein binding for initiating transcription is also unclear. PmrA does bind promoters in the absence of MglA and SspA, but the EMSA data suggest that MglA will not interact with unbound or

DNA bound PmrA in vitro without the presence of other factors. MglA may need 117 to interact with SspA before it can bind PmrA, or Francisella RNA polymerase may need to interact with MglA and/ or SspA before binding to PmrA. SspA has a role in regulating gene expression in other pathogens. Examples include

Neisseria gonorrhoeae (De Reuse and Taha, 1997) and Vibrio cholera (Merrell et al., 2002). In E. coli, SspA is a transcriptional activator of bacteriophage P1 late genes. In this case, the DNA binding protein Lpa acts as a coactivator of P1 expression. The sigma subunit of RNA polymerase interacts with the -10 and -35 elements where it initiates transcription. Lpa has been shown to regulate expression of E. coli promoters that do not contain a -35 transcriptional element.

Lpa serves as an intermediate between the DNA promoter and sigma subunit of

RNA polymerase (Hansen et al., 2003). This example is similar to the model proposed for PmrA interaction with regulated gene promoters. PmrA may also mediate the interaction of RNA polymerase with DNA promoters that may by missing transcriptional elements. Unlike E. coli, Francisella expresses two proteins like SspA (MglA and SspA) but this may be explained by the fact that F. tularensis species have two distinct RNA polymerase alpha subunits. Each SspA homolog may be a binding partner for a single type of alpha subunit. The data presented in this chapter confirms that PmrA, MglA and SspA physically interact with one another. This supports the hypothesis that PmrA is functioning as a response regulator by controlling the transcription of regulated genes. Once phosphorylated by the histidine kinase KdpD, PmrA binds to regulated gene promoters. This binding is likely independent of the other transcriptional

118 regulators (MglA, Sspa or FevR) because PmrA binding in EMSAs occurs without MglA or SspA. PmrA then interacts with MglA and/ or SspA which may or may not already be bound to RNA polymerase. Once bound this protein complex initiates expression of the regulated gene.

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Figure 5.1. Co-precipitation. PmrA, MglA and SspA co-precipitate. Western blot analysis of soluble lysates to which His-PmrA or His-MglA were added and precipitated with His-Bind resin. Panel A = anti-PmrA sera. Panel B = anti-MglA sera. Panel C = anti-His monoclonal antibody. Lane 1 = His-PmrA added to an F. novicida ∆pmrA soluble fraction and precipitated with His-bind resin. Lane 2 = His-MglA added to an F. novicida ∆mglA soluble fraction and precipitated with His-bind resin. Lane 3 = His-SspA from F. novicida His-sspA precipitated with His-bind resin. Lane 4 = His-bind resin was added to an F. novicida wild-type soluble fraction without the addition of a His-tagged protein to determine the level of non-specific binding to the resin. The blot shown is representative of at least three replicate experiments.

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Figure 5.2. Co-precipitation. PmrA, MglA and SspA co-precipitate and MglA precipitates with PmrA from a F. tularensis Schu 4 soluble fraction. Western blot analysis of soluble lysates to which His-PmrA or His-MglA were added and precipitated with His-Bind resin. Panel A = anti-PmrA sera. Panel B = anti-MglA sera. Panel C = anti-His monoclonal antibody. Lane 1 = His-PmrA added to an F. novicida ∆pmrA soluble fraction and precipitated with His-bind resin. Lane 2 = His-MglA added to an F. novicida ∆mglA soluble fraction and precipitated with His-bind resin. Lane 3 = His-SspA from F. novicida His-sspA precipitated with His-bind resin. Lane 4 = His-PmrA was added to an F. tularensis Schu4 soluble fraction and precipitated with His-bind resin. The blot shown is representative of at least three replicate experiments.

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5.5. Materials and Methods

Immunoprecipitation. Co-precipiation experiments were performed essentially as previously described (Brotcke and Monack, 2008). F. novicida

∆pmrA (JSG2845), F. novicida ∆mglA (JSG2250), F. novicida SspA-His

(JSB2970), and F. tularensis Schu4 were grown in TSB + 0.1% cysteine at 37°C to a OD600 of 1.0. The bacteria were harvested by centrifugation at ~9000 x g for

15 minutes at 4°C. Pellets were resuspended in 5 mL of Bugbuster protein extraction buffer (Novagen , Madison, WI) and sonicated for 2 minutes in 10 second bursts at an output of 50 using a Sonics VibraCell (Newtown, CT).

Lysates were centrifuged at 100,000 x g for 1 hour at 4°C. The supernatant was recovered for immunoprecipitation. Purified His-PmrA or His-MglA were added to 1 ml of the appropriate soluble fractions and incubated for 1 hour at 4°C. His- tagged and associated proteins were precipitated by incubating with 75 µl of His-

Bind Resin (Novagen, Madison, WI) overnight at 4°C with purification as per the manufacturer’s instructions. An equal volume of Laemmli sample buffer (Bio-

Rad, Hercules, CA) plus 0.5% β-ME was added to the samples and they were stored at -70°C. Proteins were detected by Western blotting. F. tularensis

Schu4 was grown and manipulated in a CDC approved BSL3 on the campus of

The Ohio State University.

Western Blotting. Proteins were separated on SDS-PAGE gels. The gels were transferred to nitrocellulose using the Trans-Blot SD semi-dry transfer cell (Bio-Rad, Hercules, CA) for 1 hour at 12 volts. Immuno-blots were blocked 122 with 1% casein (Novagen, Madison, WI). PmrA was detected using anti-PmrA rabbit antisera. MglA was detected using anti-MglA rabbit antisera. Antisera were produced by Alpha Diagnostic Intl. Inc. (San Antonio, TX). His tagged proteins were detected using a His-Tag monoclonal antibody (Novagen,

Madison, WI).

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CHAPTER 6

DISCUSSION

Francisella tularensis is one of the most infectious pathogens ever described. It can cause human disease by subcutaneous entry or pulmonary inoculation with less than 10 CFU (Saslaw et al., 1961). There is a lack of host innate immune response early in infection indicating a very efficient, active mechanism for eluding host defenses (Parsa et al., 2006). Upon entry into a host, the invading bacteria are phagocytosed by resident macrophages where they are capable of delaying maturation of the phagosome along the endocytic pathway and blocking fusion with the lysosome (Santic et al., 2006). F. tularensis escapes the phagosome to the cytosol where it can replicate rapidly

(Lauriano et al., 2004;Chong et al., 2008;Clemens et al., 2005). Once in the cytosol, F. tularensis is recognized by NOD-like receptors; this leads to activation of the apoptotic pathway in naïve macrophages (Parsa et al., 2006;Henry and

Monack, 2007;Parsa et al., 2006). Apoptosis is a non-inflammatory process which may allow the bacteria to escape and infect other host phagocytes without activating the innate immune system. F. tularensis

124 cannot replicate within activated macrophages, and once infected these cells are more likely to proceed to pyroptosis, an inflammatory process, which will result in an immune response. Recruitment of activated immune cells to the site(s) of infection is important for controlling infection (Bergsbaken et al., 2009).

The ability to replicate within macrophages is key to the bacteria’s ability to cause disease. The FPI is upregulated upon bacterial containment in the host phagosome (Lauriano et al., 2004). The proteins encoded by the FPI operon, pdpDiglABCD, have been extensively studied. The iglABCD locus has proven to be essential for escape from the phagosome and subsequent replication in the cytosol, and the IglC protein is produced in greater abundance upon infection of host macrophages (Santic et al., 2005b;Lai et al., 2004). PdpD has been shown to be located in the bacterial membrane, and its expression affects the localization of the remaining members of the operon (Ludu et al., 2008a). All of the data collected to date support the hypothesis that the FPI encodes a secretion system that is activated within host macrophages. How the system assembles and the identity of the secreted effectors have not been determined.

The F. tularensis subspecies are very closely related on the genomic level but have different virulence phenotypes (Keim et al., 2007). Significant years of research have been aimed at identifying the genetic differences and the related mechanisms that affect disease progression. F. tularensis and F. novicida replicate within murine phagocytes and are extremely lethal in mice when administered via the subcutaneous or aerosol routes of infection. These 125 subspecies also replicate within human phagocytes, but only F. tularensis is highly virulent in humans (Santic et al., 2005;Clemens and Horwitz, 2007). The mechanisms that cause these virulence phenotypes are not understood. It has been suggested that variation in, or the regulation of, the FPI may provide an explanation (Nano et al., 2004).

The FPI is present in virulent and attenuated strains of F. tularensis alike.

The island is well conserved among the subspecies; however, some differences in genes have been discovered. Notable variations in pdpD and its upstream regulatory region have been identified (Table 1.1). The FPI gene pdpD is not present in the F. holarctica subspecies (Sammons-Jackson et al., 2008).

Additionally, the F. novicida pdpD contains a 150-base pair insertion in the middle of the gene that is not present in Type A F. tularensis strains (Ludu et al.,

2008b). Despite these differences, each of these subspecies is able to replicate within host macrophages, but they differ in their ability to cause disease in man

(Feldman, 2003). Since the FPI is conserved and necessary for intramacrophage replication, it is reasonable to assume that regulation of these genes is an important factor in determining virulence. The F. tularensis subspecies (but not F. novicida) contain two copies of the FPI, but expression of two copies does not explain the differences in virulence because disruption of one copy of the FPI does not result in a virulence phenotype (Nano et al., 2004).

The regulatory network responsible for upregulating the FPI upon infection of host phagocytes may play an important role in determining virulence.

126

Our previous study investigated whether the Francisella pmrA homolog played a role in virulence (Mohapatra et al., 2007b). This study confirmed that an

F. novicida ∆pmrA is more susceptible to antimicrobial peptides, is defective for replication within macrophages, and is attenuated in the mouse. Because of the similarity to other TCS response regulators that affect transcription, an F. tularensis microarray was used to identify genes that PmrA is involved in regulating. These genes included ten of the fourteen of those encoded within the

FPI and the entire pmrA operon. Conspicuously missing from the list of differentially regulated genes was pdpD, even though the other members of the operon were present. Those genes within the pdpDiglABCD operon that were differentially regulated in the ∆pmrA mutant were iglB and iglD. Analysis of the same data using another microarray analysis software identified the entire pdpD operon to be differentially regulated. This may be due to the normalization procedures used to reduce the background associated with microarray comparisons. A hypothesis that may explain this penomenon is that there is low expression of these operons in wild-type broth-grown bacteria, and inducing environmental signals are not present. However, RT-PCR analysis of pmrA and pdpD in wild type and ∆pmrA strains disagrees with the microarray results and suggests that these genes are regulated by PmrA (Mohapatra et al., 2007b).

While the reason for these genes not being identified as being regulated by PmrA from every microarray analysis remains unclear, it is reasonable to assume that these genes and the entire FPI are differentially regulated by PmrA.

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This study provides evidence for a mechanism by which PmrA regulates virulence in Francisella. We have identified the first two component regulatory system (TCS) in Francisella. TCS regulatory systems include a histidine kinase that communicates environmental changes via phosphotransfer mediating the activity of the second component, a response regulator, that affect changes in gene expression by binding to targeted gene promoters. The regulation of TCS is dependent upon the phosphorylation of the response regulator at a conserved aspartate from a conserved histidine of the kinase. PmrA functions as a response regulator that binds to regulated gene promoters, and this binding is enhanced by phosphorylation at an aspartate residue. Microarray and RT-PCR data indicate that PmrA regulates the expression of pmrA and pdpD. We know from primer extension data that there are promoters upstream of the genes pmrA and pdpD. The EMSA experiments presented in Chapter 3 show that PmrA is a

DNA binding protein, binding upstream of pmrA and pdpD. Site directed mutagenesis and phosphotransfer experiments demonstrate that PmrA is phosphorylated at aspartate 51, and that phosphorylation affects DNA binding.

Regulation of PmrA by phosphorylation is important for the ability of F. tularensis to replicate in host cells and cause disease. Expression analysis demonstrated that transcription of iglC is dependent upon PmrA D51.

Complementation of the F. novicida ∆pmrA mutant recovers the wild-type phenotype, but complementation with the site directed mutant D51A protein results in a strain that is deficient for intramacrophage replication and virulence in

128 mice. The F. novicidia ∆pmrA pKK214pgroEL[pmrA D51A], although defective for intramacrophage replication and virulence, did display more replication and virulence than F. novicida ∆pmrA. This can be explained by overexpression of

PmrA D51A on the pkk214pgroEL plasmid. His-PmrA D51A is capable of binding to regulated gene promoters as determined by EMSAs, and this binding combined with the plasmid-mediated expression probably resulted in a low level expression of the pmrA and FPI operons. This expression explains low virulence and some intramacrophage replication observed for this strain.

KdpD is the histidine kinase primarily responsible for phosphorylating

PmrA. Phosphotransfer experiments show that PmrA phosphorylation from a F. novicida membrane fraction is dramatically reduced in the absence of KdpD.

Microarray analysis of F. novicida histidine kinase transposon mutants was inconclusive but suggests that KdpD and the protein encoded by FTN_1453 are involved in regulation of the pmrA operon. A KdpD transposon mutant is deficient for growth in macrophages, indicating a virulence phenotype similar to the pmrA mutant (data not shown). The KdpD/ KdpE TCS is widely distributed among Gram negative and positive bacteria. This regulatory system has been shown to be involved in regulation of potassium and controlling osmotic pressure.

Phosphorylated KdpE activates the kdpFABC that encodes a high affinity potassium transporter that is synthesized in potassium limiting environments

(Ballal et al., 2007). All of the kdp operons studied are repressed in the presence of high extracellular potassium (Ballal and Apte, 2005). In E. coli, the KdpD/

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KdpE system is activated in response to changes in medium osmolarity (Sugiura et al., 1992). The phagosome is derived from the plasma membrane and therefore would contain pumps that could transport potassium from the lumen of the phagosome to the cytoplasm (Amaral et al., 2007). Creation of a potassium limiting environment within the phagosome could be the signal to which KdpD responds. In Francisella, activation of KdpD would result in phosphorylation of

PmrA and then upregulation of the FPI.

Phosphotransfer experiments indicate that PmrA is phosphorylated to a greater extent in the absence of FTN_1453 and KdpE. Additionally, phosphotransfer experiments with PmrA indicate that phosphorylation of this FPI regulator is increased in the virulent F. tularensis Schu4 strain compared to the attenuated F. novicida. We attribute this to the natural absence of KdpE and the kinase encoded by FTN_1355 in the virulent Schu4 strain. KdpE is the linked response regulator to KdpD and would most likely be the primary target for

KdpD. The absence of KdpE would allow KdpD to be permiscuous and phosphorylate other targets (like PmrA) to a greater extent. Phosphotransfer experiments demonstrated that FTN_1355 was inhibitory to PmrA phosphorylation, and its absence should result in increased phosphorylation of

PmrA. Increased phosphorylation of PmrA should result in increased expression of the pmrA and pdpD operons. Increased expression of the pdpD should increase the production of virulence factors like IglC (encoded in the pdpD

130 operon). This suggests a connection between regulation of virulence determinants and virulence.

Several research groups besides ours have undertaken the creation of pmrA mutations in Francisella subspecies. The pmrA gene is 687 base pairs and is conserved at the nucleotide level in F. novicida, F. tularensis, and F. tularensis subspecies holarctica (F. holarctica). The data generated for this report used an

F. novicida pmrA deletion mutant that was created by homologous recombination that deleted minus 150 bases upstream of the ATG to base 595. The mutant is defective for intramacrophage survival and is attenuated in mice. The deletion of pmrA surprisingly does not dramatically affect the expression of the downstream genes in the pmrA operon (qRT-PCR, data not shown). Additionally, complementation of this mutant by expression of pmrA from the pKK214pgroEL plasmid restored the wild-type phenotype. Further, complementing the F. novicida pmrA mutant by expression the four genes downstream of pmrA (lepB, rnc, truB, rnr) on a plasmid does not rescue the pmrA phenotype (data not shown). A transposon mutant of lepB does have a virulence phenotype but not to a degree to account for the data observed with the pmrA mutant. It is surprising that although lepB is the only signal peptidase annotated in the whole genome sequence of F. novicida, its interruption is not a lethal mutation. It is reasonable to predict that mutation in the genes downstream of pmrA would have a general fitness phenotype. A growth defect has not been detected for the F. novicida ∆pmrA mutant, but those experiments have not been exhaustive. The

131 data is not conclusive as to whether this mutation is polar or if polar effects on downstream genes contribute to the phenotype.

A F. novicida ∆pmrA transposon mutant was also evaluated, and it was deficient for intramacrophage replication and attenuated in mice (Gallagher et al.,

2007;Mohapatra et al., 2007b). This transposon mutant does have polar effects on downstream genes. A third F. novicida ∆pmrA was constructed as a clean in- frame deletion of the gene. This mutant shares the deficiency in intramacrophage survival and mouse attenuation, but the phenotypes are not as strong as the other F. novicida ∆pmrA mutations. That is, the third ∆pmrA mutant was capable of some replication in macrophages and displayed more virulence than the other F. novicida mutations (Denise Monack, Stanford University, personal communication and data not shown) (Table 6.1).

Other pmrA mutations have been made in Francisella tularensis subspecies holarctica Live Vaccine Strain (F. tularensis LVS). Sammons-

Jackson et. al. reported on an F. tularensis LVS mutant that deleted nucleotides

26 through 645 of the pmrA gene. This mutant is also deficient for intramacrophage growth and attenuated in the mouse model (Sammons-Jackson et al., 2008). Another deletion of pmrA was constructed in F. tularensis LVS; however, this mutant does not have an appreciable phenotype in macrophage survival or the mouse model of virulence and was stated to only regulate two genes by microarray (Simon Dove, Harvard University, personal communication).

A third F. tularensis LVS mutant is also competent for replication within

132 macrophages and retains virulence in mice (Tom Zahrt, Medical College of

Wisconsin, personal communication). Our lab has made a F. tularensis LVS clean non-polar pmrA mutation that has an intermediate phenotype displaying some replication in macrophage cell lines and is more virulent in mice than the F. novicida ∆pmrA strain but not as much as the wild-type strain (Table 6.1). PmrA binds upstream of pdpD and presumably promotes transcription of virulence factors encoded within the FPI. The holarctica subspecies is unique in that pdpD and its upstream regulatory region are missing. Due to the missing PmrA binding site, it is unclear what role PmrA would play in regulating the FPI in F. tularensis

LVS and other strains of the holarctica subspecies. This may account for the lack of phenotype in F. tularensis LVS but does not explain why one of these mutants was attenuated in mice and incapable of intramacrophage replication

(Table 6.1).

Finally, our laboratory has constructed a F. tularensis Schu4 pmrA null mutant. This mutant is deficient for intramacrophage replication and attenuated in the mouse model of infection. However, this mutant displays increased intracellular replication and greater virulence than the F. novicida ∆pmrA mutant.

We have described this mutant, the F. novicida mutant constructed by Monack et. al. and the LVS pmrA mutants constructed by both Sammons-Jackson et. al. and by our laboratory to have an intermediate phenotypes. This designation indicates that they are less virulent than wild-type but more virulent that the original mutation constructed in F. novicida. The characteristics and phenotypes

133 of the pmrA mutants are summarized in Table 6.1. Six of the eight Francisella pmrA mutants are deficient in intramacrophage replication and are attenuated.

The varying phenotypes of these mutations are difficult to explain, but the variations in the regulatory region of the pdpD operon may play a role. Also, the expression of pmrA may very among the wild-type colonies from which the deletions were derived. Further investigation comparing each of the mutations to one another is required to understand how deletion of pmrA affects virulence in the three medically relevant subspecies of F. tularensis.

In recent years, a great deal of Francisella research has focused on developing a vaccine suitable for FDA approval. Many attenuated vaccine preparations have been reported in the literature, but none have shown greater efficacy against fully virulent F. tularensis than the live vaccine strain (LVS)

(Mohapatra et al., 2007b;Lauriano et al., 2004;Wayne Conlan and Oyston,

2007;Twine et al., 2005;Wu et al., 2005; Saslaw et al., 1961). The F. tularensis

LVS vaccine has not received approval due to the unknown mechanism of attenuation and the existence of non-immunogenic phase variants in vaccine preparations (Wayne Conlan and Oyston, 2007). Blue variants are immunogenic while the grey variants are non-immunogenic and the appearance of grey variants increases during infection. A possible response to the host environment is for Francisella to change to this grey or non-immunogenic phenotype in order to evade recognition by the host immune system. Research into the difference between blue and grey phase variants suggest that modification to LPS accounts

134 for the differences (Hartley et al., 2006b;Cowley et al., 1996). Because the mechanism of variation is unknown, the regulation of switching between the phenotypes has not been determined. F. tularensis phase variation has been noted in other subspecies besides holarctica. In fact, blue/ grey variants were first reported for the fully virulent F. tularensis Schu4 strain (Eigelsbach et al.,

1952). The existence of the grey variants may represent an in vivo non- immunogenic bacterial phase that provides avoidance of immune recognition and is therefore critical for virulence. Attenuation of Francisella for vaccine purposes has been attempted by deletion of transcription factors known to upregulate virulence factors during infection. While these mutants are protective against homologous challenge, they are not effective vaccines against fully virulent strains of Francisella (Lauriano et al., 2004;West et al., 2008;Mohapatra et al.,

2007b). It is possible that these transcription factors are directly or indirectly connected to the regulatory process that mediates blue/ grey phase variation.

These attenuated strains may be locked into the grey variant and are therefore unable to elicit an immune response that would be necessary for the development of protective immunity. Understanding the mechanisms of F. tularensis blue/ grey variation may be key in developing a targeted mutation for use as a tularemia vaccine.

Regulation of the FPI in F. holarctica strains is an interesting research problem. PmrA binds to the region just upstream of pdpD, and the data presented here indicates that it contributes to initiating transcription of the FPI.

135

The majority of the pdpD gene is missing in F. holarctica along with almost three kilobases upstream of this gene (Figure 1.1). This deletion removes the region to which PmrA binds. Despite this deletion, F. holarctica strains express the FPI and are able to replicate within host macrophages. Understanding the binding sequence of PmrA would allow in silico analysis to target promoters for further study aimed at determining how the FPI is regulated in F. holarctica. Perhaps this deletion interrupts the normal regulation of the FPI and accounts for as yet unexplained attenuation of the F. tularensis live LVS.

To date, five genes have been identified that are involved in regulating the

FPI: MglA, SspA, PmrA, FevR and Hfq. The products of these genes participate in a regulatory network that affects expression of the FPI. Phosphorylation of

PmrA results in upregulation of its own transcription, but may also increase transcription of FevR. MglA and SspA also regulate FevR (Brotcke and Monack,

2008). The function of FevR is unknown; it shares little similarity with non-

Francisella proteins. Slight similarity to the helix-turn-helix (DNA binding motif) of the MerR family of transcription factors suggest that FevR may bind to DNA

(Brotcke and Monack, 2008). The MerR family of transcription factors bind to

DNA independent of RNA polymerase. Therefore, FevR may bind to DNA independent of the PmrA/ MglA/ SspA complex described here. Hfq is a post transcriptional regulator that binds mRNA and has been shown to regulate the

FPI in F. tularensis LVS (Meibom et al., 2009). By binding FPI transcripts, Hfq may regulate translation and fine tune the response. The complexity of the

136 regulation of these genes has not been completely described; more research is required to understand the cooperative regulation of virulence in Francisella.

PmrA, MglA and SspA physically interact with one another as shown by co-precipiation data presented in this report. PmrA, MglA and SspA co- precipitated from F. novicida soluble fractions. PmrA, MglA, and SspA protein sequences are very similar between the F. tularensis subspecies. The amino acid sequence of PmrA is identical in the F. tularensis and F. novicida genomes.

The sequences of MglA and SspA are very similar between the two subspecies, and PmrA precipitates with MglA from a F. tularensis Schu4 whole-cell lysate soluble fraction. The data does not allow elucidation of the order of binding of

PmrA, MglA, SspA and RNA polyermase to one another. Previous data has established that MglA and SspA bind to one another and to RNA polymerase in the absence of PmrA. EMSAs results have shown that MglA does not bind PmrA bound to DNA. It is unclear whether PmrA binds to MglA and/or SspA and whether the interacting partner(s) must first bind RNA polymerase.

The data presented regarding the regulation of virulence in Francisella suggests a model for transcription of the FPI. The membrane-bound histidine kinase KdpD responds to an as yet unidentified environmental signal by phosphorylating PmrA. Phosphorylated PmrA binds to regulated promoters recruiting MglA and/ or SspA, which are bound to or will bind RNA polymerase.

In the model, this complex of proteins cooperates to initiate transcription of the

FPI (Figure 6.1). This is a model for initiating transcription of the pdpD operon; 137 however, it does not explain how PmrA regulates the pmrA operon.

Transcription of the FPI is dependent upon MglA, SspA, PmrA and FevR while expression of the pmrA operon relies on PmrA, but not the other regulators

(Brotcke and Monack, 2008;Charity et al., 2007;Brotcke et al., 2006;Mohapatra et al., 2007b). The F. tularensis RNA polymerase has two distinct alpha subunits.

This may allow for the assembly of four different RNA polymerase core enzymes.

Two of these core enzymes would have homodimer alpha subunits. The other two would be heterozygous for the alpha subunits but differ in their placement relative to the beta subunits (Charity et al., 2007). If each of these core enzymes is present, it may allow for another mechanism of regulation. MglA and SspA may only bind to a particular RNA polymerase alpha subunit and therefore would not be capable of binding to both of the core enzymes containing homodimer alpha units. The makeup of the core enzyme may determine which genes are transcribed by it. Additional research will need to be conducted to reveal the components necessary for transcription of the pmrA and pdpD operons.

The data clearly demonstrate that PmrA is involved in the regulation of virulence factors encoded within the FPI and supports a model of regulation in which PmrA functions as a response regulator in a TCS with the histidine kinase

KdpD. To date, PmrA is among four transcription factors that regulate expression of the FPI (Brotcke and Monack, 2008;Charity et al., 2007;Baron and

Nano, 1998;Mohapatra et al., 2007b), but more genes may be involved. The

138 function of each of the proteins and how they cooperate to control Francisella virulence to subvert host innate immune functions still remains to be determined.

139

Background Description* Intramacrophage Attenuated in Polar Source replication mouse

1 F. novicida Deletion replaces from -150 to 595 nt Deficient Yes No Mohapatra et. al.

2 F. novicida Transposon insertion Deficient Yes Yes (Gallagher et al., 2007)

3 F. novicida Complete in-frame deletion Deficient Yes No Monack, Stanford, personal intermediate** intermediate** communication

4 F. tularensis LVS Deletion replacing 26–645 Deficient Yes No (Sammons-Jackson et al., 2008)

5 F. tularensis LVS In-frame deletion Competent No ??? Dove, Harvard, personal communication

6 F. tularensis LVS In-frame deletion Competent No ??? Zahrt, Wisconsin, personal 139 communication

7 F. tularensis LVS Complete in-frame deletion Deficient Yes No Gunn, unpublished intermediate** intermediate**

8 F. tularensis Complete in-frame deletion Deficient Yes No Gunn, unpublished Schu4 intermediate** intermediate**

*pmrA is 687 nucleotides (nt) long. **Intermediate refers to a phenotype that is not a strong as the original F. novicida mutation.

Table 6.1. Phenotypic characterization of various Francisella pmrA mutants.

140

Figure 6.1. Model of PmrA Regulation of the FPI. The data presented in this report as well as that in the literature support this model in which: A. KdpD responds to an environmental signal and autophosphorylated, B. KdpD phosphorylates PmrA at aspartate 51, C. Phosphorylated PmrA (shown as a dimer) binds to regulated gene promoters, and D. DNA bound PmrA recruits MglA and/ or SspA, which bind RNA polymerase. Arrows indicate protein— protein interaction; however, the order of binding remains to be determined. This complex cooperates to initiate transcription of the pdpDiglABCD operon.

141

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