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Theses and Dissertations
2005
Identification of Francisella tularensis outer
membrane proteins
Amanda Melillo
Medical College of Ohio
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Identification of Francisella tularensis Outer Membrane Proteins
Amanda Melillo
Medical College of Ohio
2005
DEDICATION
This dissertation is dedicated to my mother, Adeline Melillo. She was an
amazing woman who always inspired me and believed in me. She always helped me to
believe in myself and let me know that I could accomplish anything I put my mind to.
She is a constant source of inspiration for which I will always be grateful. I love you and
miss you Mom.
ii
ACKNOWLEDGEMENTS
I would like to acknowledge my original advisor, Dr. Darren Sledjeski for all his
help and input in my project. I would also like to thank my new advisor Dr. Eric
Lafontaine for not only all of his help and insight, but also for allowing me to join his lab
once Dr. Sledjeski started his new job. Without their assistance and support, I would
never have been able to complete my master’s degree. The knowledge and expertise both
of you have shared with me will undoubtedly be useful throughout my lifetime. It has
been both a pleasure and privilege to work in both of these labs during my time at MCO.
I would like to acknowledge my committee members, Dr. Mark Wooten and Dr.
Robert Blumenthal, for taking time out of their busy schedules. Their suggestions and
discussions were a great help and extremely useful. I would also like to acknowledge Dr.
Venkatesha Basrur for his help with the Mass Spectrometry section of my project, and for
all of his input.
The members of Club Coli Conclave, Meenakshi Kaw, Serena Vanlerberg, Rachel
Balder, Brian Bullard, Christine Akimana, John Lazarus, Kylie Roach, Maria Kay,
Robert Lintner, and Pankaj Mishra, have not only been a great help during our weekly lab
meetings, but they have also become great friends. I would like to thank them all for
making work an enjoyable place to be.
I would like to thank both the Graduate School and the Medical Microbiology and
Immunology Department for allotting me the opportunity to obtain my master’s degree at
the Medical College of Ohio. I would also like to thank the secretaries of the Medical
Microbiology and Immunology Department, Joyce Rodebaugh, Sharon Ellard, and Susan
Payne, for all their help over the past 2 years.
iii
Without my friends at MCO I would have never been able to make it through the
last year. All of the good times and laughs kept me a float when times were tough. An
extra thanks to the members of the Westrink lab, where I could always escape to and get
a fresh cup of espresso. Thank you for your wonderful friendships and I will miss you
all.
Finally I would like to thank my family, especially my Dad and my brother Joe.
Thank you for always standing by me and believing in me.
iv
TABLE OF CONTENTS
Dedication…………………………………………………………………………………ii
Acknowledgements………………………………………………………………………iii
Table of Content…………………………………………………………………………..v
Introduction………………………………………………………………………………..1
Literature Review………………………………………………………………………….4
Material and Methods……………………………………………………………………27
Results……………………………………………………………………………………45
Discussion………………………………………………………………………………..75
Conclusions………………………………………………………………………………83
References………………………………………………………………………………..84
Abstract…………………………………………………………………………………..96
v
INTRODUCTION
Francisella tularensis (F. tularensis), a Gram-negative bacterium, is the
etiological agent of the zoonotic disease tularemia. Ulceroglandular tularemia is the most
common form of the disease and typically is contracted through arthropod vectors that
have fed on an infested animal. Treatment includes antibiotics such as streptomycin or
tetracycline (www.cdc.gov, 2005). Respiratory tularemia, although rare, is a deadly
disease. Due to its extreme infectious does (<10 organisms), the Center for Disease
Control and Prevention (CDC) considers F. tularensis a category A biological weapon.
Presently, there is no licensed vaccine available for use in the United States. If the
symptoms are recognized early, antibiotic therapy such as doxycycline and ciprofloxacin
can be used for treatment; however, without antibiotic therapy the mortality rate can be as
great as 30% (Oyston et al., 2004). For this reason, a new found interest has been
established in the field of F. tularensis research.
Francisella tularensis is part of the γ-proteobacteria family. Francisella
tularensis includes three main studied subspecies. These subspecies include F. tularensis
subsp. tularensis, which is the most virulent strain, and typically is the cause of
respiratory tularemia. Francisella tularensis subsp. holartica is less virulent than subsp.
tularensis in humans and typically causes the more common form of tularemia,
ulceroglandular. Subspecies holartica is the strain from which the live vaccine strain
(LVS) was derived, however, this vaccine is not used in the United States due to its
method of attenuation. Francisella tularensis subsp. novicida is rarely virulent in
1
humans, however it is fully virulent in rodents. Subspecies F. tularensis LVS and subsp.
novicida both are closely related to subsp. tularensis and make good models to study the
virulent form of tularemia without working under BioSafety Lab (BSL-3) level
conditions.
Currently, little is known regarding F. tularensis subspecies mechanisms of
virulence. It is known that F. tularensis has the ability to survive within a macrophage
(Oyston et al., 2004). A handful of virulence factors have been previously identified
including two genetic loci, intracellular growth locus, iglABCD, and macrophage growth
locus, mglAB. Genes that compose these loci are known to be upregulated during
macrophage growth (Lauriano et al., 2004). Both of these loci seem to be essential for F.
tularensis survival within a macrophage. Recently, the genomic sequence of the highly
virulent subspecies tularensis (Schu4 strain) was completed. This will help to further
understand the virulence mechanisms used by the bacterium.
Outer membrane proteins (OMPs) play a central role in the virulence of many
Gram-negative pathogenic bacteria. Since OMPs are expressed on the surface of the
bacterium, they often play an important role in bacterial-host interaction. Outer
membrane proteins can act as adhesins, which mediate adherence to host cells, such as
the Hag (Holm et al., 2003), McaP, and OMPCD proteins (Holm et al., 2004) that
mediate adherence of Morxella catarrhalis. Another important type of OMPs are porins,
that play an essential role by allowing the influx of essential nutrients into the cell and
keep out deadly toxins.
2
The relative paucity of information regarding the pathogenesis of F. tularensis
subsp. in part is due to the lack of genetic tools available. It is difficult to genetically
manipulate F. tularensis subsp. (Golovliov et al., 2003b; Lauriano et al., 2003), which in
turn makes it extremely challenging to discover what different genes are regulating.
Recently, more useful and successful methods of transformation and allelic exchange
have been described by Golovliov et al. (2003b) and Lauriano et al. (2003) in both F.
tularensis subsp. novicida and F. tularensis LVS.
In this study we identify and examine a number of F. tularensis LVS outer
membrane proteins using a membrane impermeable biotinylation agent to identify
surface proteins. We hypothesize that certain OMPs on the surface of F. tularensis
mediate adherence to lung epithelium.
3
LITERATURE REVIEW
Background on Francisella tularensis
The bacterium F. tularensis causes tularemia, a deadly zoonotic disease. This
disease is found throughout the northern hemisphere with the majority of United States
cases occurring in south-central and western states. In humans, tularemia occurs in
several different forms including ulceroglandular, oropharyngeal, glandular, typhoidal,
and respiratory (www.cdc.gov, 2005). Ulceroglandular tularemia is rarely fatal and is
contracted via infection by arthropod vectors, such as tick bites or deer flies. It is the
most common form found in the United States, with several hundred cases reported each
year. Most non-laboratory infections occur after handling infected animals. Three to 5 d
after exposure, lesions appear and lymph nodes become enlarged due to the
dissemination of F. tularensis (Evans et al., 1985). Ulceroglandular tularemia can be
treated effectively with antibiotics such as tetracyline, aminoglycosides and most
commonly streptomycin (www.cdc.gov, 2005).
Respiratory tularemia is the most deadly form of tularemia. In some cases it can
progress from the ulceroglandular form, but typically results from inhalation of the
bacteria (Evans et al., 1985). Respiratory tularemia is the most threatening form of the
disease in terms of intentional release, and is one of the main reasons why F. tularensis is
designated as a Category A agent of bioterrorism by the CDC. Without the use of
antibiotics, respiratory tularemia has a mortality rate of 5-30% (Larsson et al., 2005).
Antibiotic treatment for a respiratory outbreak of tularemia includes doxycycline and
ciprofloxacin, administered orally (Russell et al., 1998; www.cdc.gov, 2005).
4
Respiratory tularemia may lead to bronchial hemorrhaging and mediastinal
lymphadenopathy (Kawula et al., 2004). Although respiratory tularemia is less frequent,
it can occur naturally by the spread of dust on a farm with infected animals, or in rare
events, such as the following. The first case of respiratory tularemia in the United States
was reported in Martha’s Vineyard, Massachusetts, in 1978 (Feldman et al., 2001). In
2001, an infected rabbit carcass was run over by a lawnmower, which aerosolized F.
tularensis. Fifteen patients were diagnosed with tularemia, including 11 respiratory form
cases. One individual died and was found to be infected with F. tularensis Type A
(Feldman et al., 2001). Similar cases have been seen in Martha’s Vineyard within the last
few years, mostly among landscape workers.
F. tularensis as a Bioweapon
Francisella tularensis is considered a potential biological weapon because of its
extreme infectivity (infectious dose, ID50 in humans, of 10 – 50 organisms via
inhalation), ease of dissemination via aerosols, and substantial capacity to cause illness
and death. Francisella tularensis was investigated for use as a biological weapon in
Japan, the former Soviet Union and the United States. There were stockpiles of
weaponized F. tularensis made in the Soviet Union in the mid 1930s (Dennis et al.,
2001). By the 1960s there were reports that the Soviet Union had antibiotic resistant
strains as well as strains resistant to the immunological responses elicted by
immunization with F. tularensis LVS (Dennis et al., 2001). In the 1970s, all stockpiles in
the United States were declared to have been destroyed, followed by the reported
destruction of the Soviet Union’s stockpiles in the early 1990s.
5
It is important to study tularemia because of recent biowarfare threats and its
potential use by terrorists. It is not known if any countries possess weaponized F.
tularensis today, but due to its extreme infectivity, it remains a threat. The World Health
Organization (WHO) predicts that the “release of 50 kg of dried F. tularensis over a city
with a population of 5 million would result in 250,000 cases of the disease and 19,000
deaths” (Oyston et al., 2004). It was also predicted that illnesses from such an attack
would persist for many weeks and several disease relapses would occur.
F. tularensis
Francisella tularensis, a Gram-negative pleomorphic coccobacillus, is a
ubiquitous, nonmotile, aerobic bacterium that has the ability to survive in various
environments such as soil and animal carcasses. This organism ranges in length from 0.5
to 10 µm (Ketterer, 2003), and produces a polysaccharide capsule (Sorokin et al., 1996).
Natural reservoirs for F. tularensis include mammals, birds, rodents and amphibians.
Francisella tularensis has been isolated from approximately 250 different species (Hopla,
1974).
Francisella tularensis is a member of the γ-proteobacteria and can be divided into
four subspecies based upon 16S rRNA sequence analysis. Other human bacterial
pathogens most closely related to F. tularensis are Coxiella burnetii and Legionella
pneumophila (Titball et al., 2003) based on 16S rRNA sequence analysis. The four F.
tularensis subspecies possess varying virulence for humans, but are all highly virulent in
mice. Francisella tularensis subsp. tularensis (Type A, strain Schu4) is the category A
biological agent, and the most virulent of the subspecies for humans, with an estimated
6
infectious dose in humans of <10 CFU (Oyston et al., 2004). Following the arrival of
potential vaccine strains to the United States in 1956, research on these strains was
substantial enough, at that time, to begin human studies. During these studies it was
shown that nonvaccinated individuals could become ill with tularemia through the
inhalation of between 10-50 organisms (Ellis et al., 2002).
Francisella tularensis subsp. holarctica (Type B) is less virulent than subsp.
tularensis in humans and is distributed throughout Europe and North America, with an
estimated ID50 in humans <103 CFU (Ellis et al., 2002). Infection with subsp. holarctica
most commonly results in ulceroglandular tularemia and the F. tularensis LVS was
derived from a human isolate of this subspecies. A mixture of potential live vaccine
strains were transferred to the U.S. in 1956, and an isolate was derived, passaged through
mice, then tested for protective immunity against subsp. tularensis challenge (Sandstrom,
1994). Because of this procedure the genetic basis for the attenuation of virulence in F.
tularensis LVS is not known. Therefore, F. tularensis LVS is not in use for vaccination
in the United States due to the possibility of this strain reverting back to its virulent form;
however, it has been used by some countries, including the United States, to immunize
lab workers (Oyston et al., 2004; www.cdc.gov, 2005). F. tularensis LVS retains
virulence for mice. Francisella tularensis subsp. mediaasiatica is isolated only in limited
areas of the former Soviet Union and Central Asia, and is not well studied. Francisella
tularensis subsp. novicida is found throughout the Northern Hemisphere, and recently in
Australia, and is virulent for most rodents, but typically avirulent in humans (Oyston et
al., 2004).
7
Virulence Factors of F. tularensis
Very little is known about the molecular mechanism for pathogenesis of
tularemia. Survival within macrophages appears to be critical to the pathogenicity of all
F. tularensis subsp. A hallmark of all F. tularensis subsp. infection is its ability to
survive and replicate within alveolar macrophages (Lauriano et al., 2004). Francisella
tularensis subsp. are all highly virulent in mice and have the ability to infect and replicate
in murine macrophages as well. Infected murine macrophages (J774A.1) are unable to
clear the bacteria and eventually undergo apoptosis, leading to the release of the bacteria
and dissemination throughout the host (Lai et al., 2004). Two genetic loci (mglAB and
iglABCD) have been shown to be essential for the survival of subsp. novicida within
murine macrophages. The mglAB locus is one of the few known virulence factors of
subsp. novicida. Recent work has shown that the protein MglA is a critical determinant
of virulence in subsp. novicida (Lauriano et al., 2004). The MglA protein functions as a
transcriptional regulator of the intracellular growth locus (Igl) A, B, C, and D (Lauriano
et al., 2004) and the pathogenicity determinant protein locus (pdpABC) (Nano et al.,
2004), once the bacterium enters the macrophage (Figure 1) (Gray et al., 2002). High
sequence similarity is seen between MglA and the stringent starvation protein (SspAB) in
Escherichia coli (E. coli) (Gray et al., 2002). During transcription SspAB modulates the
function of RNA polymerase. When cells are faced with amino acid starvation, SspAB is
heavily
8
Figure 1. Genes Regulated by MglA
Diagram of the genes surrounding iglC, shown as arrows, oriented in the direction
of transcription. The region following iglD contains 11 additional ORFs. This entire
gene cluster is regulated by MglA (Lauriano et al., 2004).
9
synthesized which upregulates SspAB and helps keep the cell alive (Serizawa and
Fukuda, 1987).
Through the use of shuttle mutagenesis methodology, Gray et al. (2002) identified
four genes clustered into the same locus that appear to be essential for subsp. novicida
intracellular growth and labeled them intracellular growth loci (Igl) A, B, C, and D
(Figure 1) (Lauriano et al., 2004). A 23 kD protein is expressed by iglC when subsp.
novicida infects a murine macrophage (cell line J774.1). IglA and IglB show high
similarity to a group of conserved proteins in Rhizobium leguminosarum, with unknown
function (Lauriano et al., 2004). Protein family (Pfam) database searches also show that
IglA and IglB share conserved domains, DUF770 and DUF877, respectively, in bacteria
such as Pseudomonas aeruginosa, Yersinia pestis, and Vibrio cholerae. The function of
IglD is unknown and no significant sequence similarity was found through protein
database searches. When the phenotypes of isogenic mutants that lack expression of the
proteins were assayed, it was found that iglC mutants demonstrated poor intracellular
growth in murine macrophages (Lai et al., 2004). This could lead to the conclusion that
the entire operon is necessary for intracellular growth, but more research must be done to
confirm this. Expression of IglC in subsp. novicida is also increased during oxidative
stress, such as the presence of hydrogen peroxide (Gray et al., 2002; Lauriano et al.,
2004). An iglC isogenic mutant of subsp. nocivida showed decreased growth within
murine macrophages, and was readily cleared from macrophages. This suggests that IglC
might have the ability to interrupt key antimicrobial effector mechanisms (Lai et al.,
10
2004). F. tularensis LVS iglC mutants were found to have similar phenotypes in murine
macrophages (Kawula et al., 2004; Lai et al., 2004).
Francisella tularensis subsp. novicida has recently been shown to survive and
replicate within the amoebae Acanthamoebae castellanii (A. castellanii) (Lauriano et al.,
2004). Both mglA and iglC mutations demonstrate decreased growth and survival within
amoebae as well as macrophages (Lauriano et al., 2004). Legionella pneumophila, also
an intracellular pathogen that also infects human macrophages, has been shown to utilize
protozoa as a natural reservoir (Neumeister et al., 1997). It has been shown that
Legionella has similar growth and survival mechanisms within amobae and
magrophages. This suggests that subsp. novicida’s ability to survive within macrophages
may have similar mechanisms to its survival and growth in A. castellanii. Francisella
tularensis, like Legionella, may have evolved its ability to survive within macrophages
from its ability to survive in protests (Lauriano et al., 2004). Francisella tularensis LVS
also has been demonstrated to survive and grow within the amoebae A. castellanii (Abd
et al., 2003; Neumeister et al., 1997).
Factors Involved in the Survival and Escape from the Phagosome
Little is known regarding the intracellular survival of F. tularensis subsp. Three
typical pathways of intracellular survival are 1) extraphagosomal, in which bacteria
escape from the phagosome and reside in the cytoplasm, as seen in and Shigella subsp.
(Clerc et al., 1987); 2) phagolysosmal, in which the phagosome and lysosome fuse, as
seen in Coxiella burnetii (Burton et al., 1971); and 3) phagosomal, in which the bacterial
reside in the phagosome, but fusion to the lysosome is blocked, as seen in Mycobacterium
11
tuberculosis (Armstrong and Hart, 1971). Francisella tularensis research has not yet
confirmed any of these strategies with certainty. Anthony et al. (1991) demonstrated that
there is no fusion with the lysosome, while Golovliov et al. (2003a) and Clemens et al.
(2004) observed the presence of lysosome-associated membrane glycoproteins (LAMPs).
Golovliov et al. (2003a) showed that after initial infection of macrophages there was a
significant increase in the presence of LAMPs, and after 1 h their levels were decreased,
however, no fusion between lysosome and phagosome was seen. Golovliov et al (2003a)
and Clemens et al. (2004) both show the escape of F. tularensis (LVS and subsp.
tularensis) from the phagosome into the cytoplasm following the increase in LAMP
levels. The extraphagosomal pathway is the most likely strategy of F. tularensis, as
established by this research. However, there is contradictory data regarding the pH of the
endosomal environment.
When F. tularensis LVS is grown in acidic conditions, ammonia is released
(Chamberlain, 1965). This may help to buffer acidic environments such as those in
endosomes or macrophages, which typically have a pH ~5. Genes have been identified in
F. tularensis LVS, orthologs of which are typically responsible for production of
ammonia. These include, L-glutaminase, L-asparginase and citrulline ureidase. Each of
these enzymes has been used in earlier research to help characterize differences between
subsp. tularensis (highly virulent) from subsp. holartica (low virulence); for example
lower levels of glutaminase activity are associated with decreased virulence (Larsson et
al., 2005). However, according to Gray et al. (2002) and Fortier et al. (1995), F.
tularensis LVS requires acidification of the phagosome, in order to help with the
12
acquisition of iron. Although the phagosome containing F. tularensis has not been
shown to fuse with lysosomes, the environment is still considered to be acidic from this
data (Fortier et al., 1994; Rhinehart-Jones et al., 1994). Additional pH experiments were
performed by Clemens et al. (2004), which demonstrated the pH of the endosomal
compartment is not to be significantly acidic. More research must be done in this area
before any conclusions can be drawn.
Iron Uptake
It is postulated that the acidic phagosome allows intraphagosomal bacteria to strip
iron from transferrin (Fortier et al., 1995). The genome sequence of Schu4 (subsp.
tularensis) contains a ferric uptake regulator, fur, which is important for the uptake of
iron by other bacteria. However, F. tularensis LVS has been seen to escape the
phagosome (Golovliov et al., 2003a), therefore, one would expect to find genes for
complex-bound iron uptake in order to utilize the highly complex iron in the cytoplasm.
The TonB-dependent system is an example of a gene that functions to allow the
bacterium to utilize complex iron. The TonB-dependent system is seen in pathogenic
bacteria such as Pasteurella (Nemish et al., 2003), Neisseria (Nemish et al., 2003), and E.
coli (K12 and enteropathogenic)(Nikaido, 2003). No obvious orthologs were identified
in subsp. tularensis genome (Larsson et al., 2005).
Lipopolysaccharide (LPS)
A common virulence factor among pathogenic Gram-negative bacteria is the
lipopolysaccharide (LPS). Lipopolysaccharide is composed of an amphipathic glycolipid
with a hydrophobic lipid (known as Lipid A), that is covalently attached to hydrophilic
13
complex polysaccharide. The presence of LPS in bacteria, such as E. coli, typically
induces release of proinflammatory cytokines by activating Toll-like receptors (TLRs)
(Telepnev et al., 2003). The LPS of both F. tularensis subsp. tularensis and holartica
(including LVS) exhibits unusual biological properties such as low endotoxicity, and are
a poor inducers of proinflammatory cytokines (Sandstrom et al., 1992). The LPS of F.
tularensis subsp. is significantly defective in the induction of TNF-α, IL-1, and NO
production in macrophages, compared to the LPS of other Gram-negative bacteria
(Cowley et al., 1996). Francisella tularensis LPS has been observed to undergo phase
variation due to changes in both the O-antigen and Lipid A portions, with reduced
expression of LPS, in subsp. tularensis and holartica (Eigelsbach et al., 1951). Phase
variation typically occurs in the polysaccharide portion, leading to changes in surface
antigenicity and resistance to antibody mediated clearance. The phase variation of subsp.
holartica LPS possibly leads to the decreased toxicity of the LPS, however, the
significance of phase variation of LPS in relation to human disease is unknown (Oyston
et al., 2004).
F. tularensis subsp. tularensis Genome
The complete genome sequence of the Schu S4 strain (subsp. tularensis) has
recently been determined by Larsson et al. (2005). The circular chromosome includes
1,892,819 base pairs (bp), containing 1,804 recognized coding sequences of which 1,281
have orthologs to γ-proteobacterial genomes. Identified proteins were clustered into 61
groups each consisting of two or more members. A number of new proteins and potential
virulence factors were identified; including one cluster consisting of five novel proteins,
14
three of which contain putative signal peptides and coil-coil domains. Five families of
insertion sequence (IS) elements were identified in subsp. tularensis. Surprisingly, one
of the insertion sequences (ISFtu1), shares similarity to the IS630 Tc-1 mariner element,
which is more typically seen in eukaryotes. It is thought that ISFtu1 may have been
acquired from insects, which are a common host of F. tularensis subsp. One copy of
ISFtu1 is located in the O-antigen cluster. A similar IS630 element has been seen in
Shigella sonnei (S. sonnei) and plays a role in stable expression of the particular Oantigen, which is essential in pathogenesis of S. sonnei. Other IS elements families found
include IS5, IS4, IS982 and IShpaI. The latter possesses a terminal inverted repeat
sequence that has not been previously identified, and is therefore, a new type of IS
element (Larsson et al., 2005).
A number of potential virulence factors also were identified, including a type IV
pilus, as predicted in earlier work (Gil et al., 2004; Hubalek et al., 2004). Gil et al.
(2004) described a number of type IV pili that had been identified in F. tularensis LVS
via ultrastructure examination of broth grown cells. In other Gram-negative bacteria,
such as Pseudomonas aeruginosa, Neisseria gonorrheae, Ralstonia solancearum (Sundin
et al., 2002), type IV pili are a known virulence factor. These pili have numerous
functions including the ability to mediate adherence to host cells (Strom and Lory, 1993),
DNA uptake (Fussenegger et al., 1997), and biofilm formation (Merz et al., 2000). In F.
tularensis subsp. the purpose of the type IV pili is not yet determined, but they may have
similar functions.
15
A putative pathogenicity island also was identified. This area is the only area of
duplication, and some genes found within this area are known virulence factors such as
iglC and mglA (Larsson et al., 2005). Evidence of an F. tularensis pathogenicity island
include clusters of genes encoding virulence factors (Lauriano et al., 2004), high G + C
content (Nano et al., 2004), and transposable elements adjacent to the area (Nano et al.,
2004). Phospholipase C and a phospholipase D orthologs were identified in Schu4, and
may be important for degrading the phagosomal membrane and bacterial escape into the
cytosol, which has been observed in vitro (Golovlivov et al., 2003a). An ortholog of the
Mammalian Cell Entry (mce) gene (Kumar et al., 2003) was identified; mce is involved in
gaining entry and surviving in macrophages in order to escape the host immune defenses,
by Mycobacterium tuberculosis (Arruda et al., 1993). A gene denoted FTT1043 is
related to a gene found in Legionella pneumophila (L. pneumophila) that is a macrophage
infectivity protein, and plays a role in the virulence of L. pneumophila (Larsson et al.,
2005).
Polysaccharide Capsule
Computer analysis of the subsp. tularensis genome identified a cluster of genes
that could be involved in the synthesis of the poorly characterized F. tularensis
polysaccharide capsule. In F. tularensis subsp tularensis, capB and capC genes were
identified (Larsson et al., 2005). Although their function is unknown in F. tularensis at
this time, these genes are responsible for the production of capsule in Bacillus anthracis
and may play a similar role in F. tularensis. The F. tularensis capsule has been reported
to have similar composition to the LPS O-antigen. One report demonstrated that strains
16
lacking a capsule are more susceptible to serum killing, but appear to survive better
within neutrophils (Sorokin et al., 1996).
Several virulence genes that are typically found in pathogenic bacteria, such as
type III, IV or V protein export systems as well as a TonB-dependent system, or other
systems known to acquire complex iron (Larsson et al., 2005), were not identified in the
complete sequence of the Schu4 genome. This suggests that novel proteins or other
macromolecules might play a role in F. tularensis pathogenicity. Further studies must be
conducted to determine the methods of virulence, including identification of outer
membrane proteins (OMPs). Outer membrane proteins may potentially play an important
role in the pathogenesis of all F. tularensis subsp. by mediating adherence to the host
cell. Discovery of new virulence genes in F. tularensis subsp. will not only increase
knowledge regarding F. tularensis pathogenesis, but also enable rational searches for
similar virulence mechanisms in other bacteria.
Pathogenesis
Disease pathogenesis is important to understand when studying a disease and is
essential for rational vaccine design. Little is known regarding the pathogenesis of F.
tularensis or any of its subspecies. The current research and knowledge of pathogenesis
of tularemia is based on studies of F. tularensis LVS and subsp. novicida (99.6%
sequence identity to subsp. tularensis, Schu4). This is due to their relatively low
virulence in humans compared to Schu4 which must be handled in BSL-3 containment
laboratories (Eigelsbach et al., 1975).
17
Frequently, the first step of pathogenesis is adherence of the bacterium to the host
(Nikaido, 2003). Two main mechanisms that mediate bacterial adherence to host cells
involve the polysaccharide capsule and surface proteins called adhesins. Adhesins are
found on most pathogenic bacteria. For example H. influenzae, uses the Hia (Ecevit et
al., 2004) and Hap (Hendrixson and St. Geme, 1998) adhesins for binding to human cells
(Hendrixson and St. Geme, 1998). Until recently, no adhesins and few outer membrane
proteins had been identified on F. tularensis subsp. It is important to identify adhesins,
not only because of the information they can provide regarding pathogenesis of the
bacterium, but also as they are good vaccine candidates. This is due to the fact that they
are expressed on the surface of bacteria, making them a prime target of the immune
system.
A good example of an adhesin that was used as a vaccine candidate is the 155
kDa adhesin produced by non-typable Haemophilus influenzae (NTHI). This protein is
expressed by most NTHI isolates. In a mouse model of infection, antibodies against this
protein were cross-reactive to the 155 kDa protein of most NTHI strains. When the
antibody is preincubated with cells expressing the Hap cell binding domain (CBD)
adherence was decreased, indicating that the antibodies are blocking the adhesin. Further
tests showed that the antibodies against CBD blocked colonization in a mouse model of
infection (Liu et al., 2004). These data indicate that the Hap adhesin is an excellent
vaccine candidate. A similar approach could therefore be used for an F. tularensis
adhesin vaccine, though other surface proteins could also be used for this purpose.
18
The most likely use of F. tularensis as a biological weapon is through inhalation
of aerosolized bacteria, leading to respiratory tularemia. In this scenario, the likely
sequence of events would be initial infection at the lung interface by F. tularensis,
followed by phagocytosis by macrophages, and then dissemination via the migratory
macrophages through the lymphatic system (Dennis et al., 2001). One report has
demonstrated that F. tularensis LVS-infected macrophages undergo apoptosis (Lai et al.,
2001), but no secreted products or even secretion systems that could mediate this process
have been identified. Figure 2 illustrates our proposed steps leading to the pathogenesis
by F. tularensis. We hypothesize that F. tularensis produces adhesins that are crucial for
initial infection on the lung surface. Once bound to the host, the bacteria multiply and
attract macrophages, which phagocytose the bacteria. These macrophages can then
migrate via the lymphatic, where they undergo F. tularensis induced apoptosis, allowing
the bacteria to survive and disseminate within the host.
Outer Membrane Proteins
Surface-exposed proteins play vital roles in bacterial virulence and pathogenesis
(Navarre and Schneewind, 1999). As of spring 2005, no data on initial bacterial-host cell
interactions have been reported for F. tularensis. Also, little is known regarding the
surface antigens important in stimulating host immunity. Different methods have been
used to identify outer membrane proteins which are possible vaccine candidates in other
bacteria. These include the whole genome approach, in S. pneumoniae (Wizemann et al.,
2001), which uses sequence scanning and bioinformatics analysis, and DNA microarray,
in Neisseria meningitidis (Grifantini et al., 2002), by identifying genes that are regulated
19
Figure 2. Model for F. tularensis Pathogenesis
This model represents a possible mechanism for F. tularensis pathogenesis. Research
presented in this thesis focuses on identifying and characterizing outer membrane
proteins of F. tularensis that mediate adherence to human cells (first step in figure).
20
during adhesion and then using microarray data to find antigens with protective immunity
among the adhesion-induced surface proteins. Some of these OMPs were found to
mediate adherence to host cell and blocking this interaction could be beneficial in vaccine
discovery.
Outer membranes are a second lipid bilayer membrane that is typical of Gramnegative bacteria. The components of the outer membrane often play important roles in
the interaction of pathogenic bacteria with their host organisms; however, the major role
of this membrane appears to be as a permeability barrier that prevents the entry of many
toxic molecules, while allowing influx of nutrient molecules and efflux of waste products
(Nikaido, 2003). Outer membranes have necessary channels, called porins which are
composed of OMPs. Porins allow the passage of essential nutrients into the cell.
Examples of porins include, OmpF, OmpC and PhoE which are all part of the outer
membrane of E. coli (Nikaido, 2003). These porins are expressed under different growth
conditions; for example, PhoE is only expressed during phosphate starvation (Nikaido,
2003).
Two types of proteins are found within the outer membrane, lipoproteins and
integral proteins. Lipoproteins are attached to the outer membrane via the N-terminal
lipid tail. Integral proteins contain one or more membrane spanning regions. Most
proteins destined for the outer membrane are synthesized in the cytoplasm as precursors
with N-terminal signal sequences, which are essential for translocation across the inner
membrane. Proteins are typically translocated to the outer membrane through the Sec
system, which translocates unfolded proteins (Bos and Tommassen, 2004).
21
The OMPs constitute an interface between the host and bacterium, which makes
them essential to study. Some bacteria use OMPs to mediate adherence to the host cells,
as in the Hia and Hap adhesins of H. influenzae, and the Hag and OmpCD proteins of
Moraxella catarrhalis. The OMPs can also interact with mucosal surfaces or the immune
system. For example, Pandher et al. (1998) demonstrated that the OMP PlpE is an
important surface antigen in Pasteurella haemolytica, by suggesting that it is induces
protective immunity (Pandher et al., 1998). The OMPs also have been used as markers of
virulence between strains. For example, the majority of clinical cases of H. influenzae
Type b express an OMP, called 1H, while both 1H and non-1H strains can colonize hosts,
1H strains are more pathogenic (Barenkamp et al., 1981).
Gilmore et al. (2004) described several putative outer membrane proteins on
subsp. tularensis (Schu4). Following the identification of loci essential for intracellular
growth, mglAB and iglC, it is now important to identify surface-associated proteins on F.
tularensis subsp. because many virulence-associated proteins are exported to the cell
surface. Gilmore et al. (2004) used the transposon, TnphoA, which utilizes phoA, to
identify surface-associated proteins on subsp. tularensis. Alkaline phosphatase (AP) is
the product of phoA, and must be exported extracytoplasmically to exhibit enzymatic
activity. An AP gene without a signal sequence cannot be exported. However, if it is put
in frame with a target gene to create a hybrid protein, then if that protein contains a signal
sequence, PhoA activity will result. This method revealed 11 potential OMPs, including
two potential virulence factors. One identified gene, P52, has significant amino acid
sequence similarity to penicillin-binding proteins. Pathogenic streptococci are resistant
22
to β-lactams due to over use of antibiotics. By modification of penicillin binding
proteins, recombination of PBPs causes low affinity for β-lactams. Francisella.
tularensis is resistant to penicillin, therefore, P52 could be a potential virulence factor.
Another gene, P15, resembles VirK, which is a virulence factor in both Shigella and
Salmonella.
Genetic Manipulation of Francisella tularensis
The relative paucity of information regarding pathogenesis by F. tularensis subsp.
is due to, in part, the poorly developed genetic tools available. Most mutant strains have
been generated by random transposon mutagenesis. Although this method works, it has
been found that transposon insertions are unstable in subsp. novicida as well as F.
tularensis LVS, however, subsp. novicida has been shown to be easier to genetically
manipulate than F. tularensis LVS (Lauriano et al., 2003). Also subsp. novicida and F.
tularensis LVS are resistant to different antibiotics, so the same protocol cannot typically
be used for both. Although F. tularensis subsp. are very closely related genetically
(99%), their virulence towards humans differs greatly. Therefore, it is important to
improve genetic manipulation of all F. tularensis subsp. in order to have more efficient
methods to dissect mechanisms of pathogenesis in humans. Transformation and allelic
exchange have been recently described by Golovliov et al. (2003b) and Lauriano et al.
(2003), which are further discussed below.
Electroporation (Anthony et al., 1991; Baron et al., 1995), cryotransformation
(Pavlov et al., 1996), and conjugation (Pomerantsev et al., 1991) are all methods that
have been used to make isogenic mutants of F. tularensis subsp. Golovliov et al. (2003b)
23
describes a method to successfully recombine a mutant gene into F. tularensis LVS via
conjugation. This method makes use of a plasmid carrying sacB, which expresses a
secreted levansucrase (expression is toxic for Gram-negative bacteria in the presence of
sucrose). The gene of intrest, iglC, was mutated by deleting the majority of its sequence
via restriction enzyme digestions, followed by ligation. The cloned DNA was ligated
into pPV, which contains a chloramphenicol resistance gene and a sacB gene. The
plasmid was transformed into E. coli and via conjugation and a single crossover event,
the plasmid was transferred to F. tularensis LVS. Transformants which grow on
chloramphenicol were recovered and would either have undergone illegitimate
recombination or contain the truncated gene. To check which recombination event
occurred the transformants were plated with sucrose and only transformants which
contained truncated iglC would grow (Golovliov et al., 2003b).
Transposon mutagenesis using mTn10Km has been successfully used to isolate
macrophage growth mutants in subsp. novicida, but they were found to be unstable due to
further transposition of the mTn10Km (Lauriano et al., 2003). To overcome this problem
Lauriano et al. (2003) reports mutagenesis by allelic exchange with PCR products. One
of the few known virulence factors in F. tularensis subsp is mglA. The activity of mglA
is removed by deleting approximately 100 codons by PCR, followed by the insertion of
an erythromycin resistance cassette. In order to transform the knockout into subsp.
novicida, it was shown that increasing the flanking regions of the gene greatly increases
the transformation efficiency. The knockout PCR products were introduced by
cryotransformation (Lauriano et al., 2004). Deletions were also made by this same
24
method in iglC (upregulated during growth in macrophages), tul4 (T-cell reactive protein,
unknown function) and bla (beta-lactamase gene) in subsp. novicida. Following
transformation, the four gene knockouts were characterized in subsp. novicida. Tul4 may
be required for full resistance to macrophage killing mechanisms, however, strains with a
mutation in bla were still able to grow in the presence of ampicillin, therefore, there must
be other ampicillin resistance genes present in subsp. novicida genome. This method
could not be used in F. tularensis LVS because it is naturally resistant to erythromycin.
Due to the problems using the mTn10Km and Tn1721 transposons, Kawula et al.
(2004) describes the use of transposon Tn5 (kanamycin resistance gene) to disrupt
potential virulence genes in F. tularensis LVS. The Tn5 derivative was used in earlier
work by Goryshin et al. (2000), where they describe its use to create random mutations in
Salmonella, Proteus and E. coli. Kawula et al (2004) identified 21 potential open reading
frames by creating random insertions using electroporation in F. tularensis LVS. They
consistently achieved a low insertion frequency (2.6 x 10-8), but the mutations were
distributed evenly throughout the chromosome. To test whether Tn5 moved throughout
the chromosome due to instability, as Tn10 and Tn1721, they passed five strains for ten
days on kanamycin media and performed southern blots. Results showed the transposon
to be stable within the genome and not susceptible to chromosomally encoded
transposases (Kawula et al., 2004).
A plasmid shuttle vector that replicates in F. tularensis LVS, subsp. novicida and
E. coli has been developed. This vector was successfully used to express the green
florescent protein in F. tularensis LVS. A specific shuttle vector, pFNL10, was
25
developed from a native plasmid of subsp. novicida, and has the ability to replicate in
both F. tularensis LVS and subsp. novicida, and is stably inherited (Maier et al., 2004).
Due to pFNL10 inability to replicate in E. coli as well as the lack of selectable markers, it
is not highly used. Maier et al. (2004) describes the manipulation of this shuttle vector,
pFNL10, into a more stable and efficient vector, pFNLTP1. The use of pFNLTP1, allows
for autonomous replication in F. tularensis spp. as well as E. coli expression of antibiotic
resistance markers, transformation at increased efficiency, stability in vitro and in vivo
and no effect on the growth or virulence of the bacterium (Maier et al., 2004). This
potentially expands the tools of genetic analysis of F. tularensis.
We hypothesize F. tularensis produces adherence to target cells and does so via
surface adhesins. In order to identify adhesins several methods may be used. These
include, (1) transposon mutagenesis, in which random mutations are made and the cells
are tested for decreased adherence, (2) construction of a library in a non-adherent bacteria
such as E. coli followed by testing for enhanced adherence, and (3) biotinylation in which
OMPs are identified and compared to adhesins within protein databases.
In this study, we utilize a non-genetic method for identification of surface
expressed proteins. This method uses the membrane impermeable reagent,
sulfosuccinimidyl-6-(biotin-amido)hexanoate (sulfo-NHS-LC-Biotin, Pierce) to
specifically biotinylate the amine groups on surface exposed proteins. Following the
identification of these proteins, two specific putative OMPs were examined as possible
adhesins.
26
MATERIAL AND METHODS
Strains and Media
Francisella tularensis subspecies holartica (LVS) and subsp. novicida are both
used in this study. We obtained the F. tularensis subspecies holartica (LVS) from
Francis Nano at the University of Victoria in Canada, Department of Biochemistry and
Microbiology. We obtained the F. tularensis subspecies novicida from Bernard
Arulanandam, University of Texas at San Antonio, Department of Biology. Both
bacteria are grown on chocolate II agar medium supplemented with hemoglobin and
isovitaleX, from Fisher Scientific. To see isolated colonies on agar plates, F. tularensis
LVS requires ~36 h incubation with 7.5% CO2 at 37°C, while subsp. novicida needs ~24
h.
The E. coli cloning strain Epi300 (Epicentre) was used for all genetic
manipulations in this study. Table I summarizes the plasmids used.
The human cell lines A549 (type II human alveolar epithelium; ATCC CCL85)
and Chang (conjuctival epithelium; ATCC CCL20.2) were cultured in tissue culture (TC)
medium (HAM’s F12 medium (Cellgro) supplemented with 2 mM glutamine, 10% heatinactived fetal bovine serum, 0.15% sodium bicarbonate, 100 µg/ml
penicillin/streptomycin) at 37°C with 7.5% CO2. The human cell line NCIH292
(mucoepidermoid lung epithelium; ATCC CRL-1848) was cultured in the same medium
supplemented with 10 mM HEPES, pH 7.5 (GIBCO), 1 mM sodium pyruvate, and 4.5 g
of glucose/liter.
27
Table I. Summary of Plasmids.
Name
Insert
Insert
Size
Vector
Antibiotic
Resistance
E. coli
Cell line
Used for
pCC_FopA.5
FopA w/ 2kb
flanking regions
~6kb
pCC1.1
Chloramphenicol
Epi300
inserting Kan cassette
pCC_pomp1.5
pOMP1 w/ 2kb
flanking regions
~6kb
pCC1.1
Chloramphenicol
Epi300
inserting Kan cassette
pCR_fopA.5
FopA w/ 2kb
flanking regions
~6kb
pCR2.1
Ampicilin
DH5α
Adherence Assays
pCR_pomp1.5
pOMP1 w/ 2kb
flanking regions
~6kb
pCR2.1
Ampicilin
DH5α
Adherence Assays
fopA_ORF
FopA ORF
~1kb
pCC1.1
Chloramphenicol
Epi300
Adherence Assays
pomp1_ORF
pOMP1 ORF
~1kb
pCC1.1
Chloramphenicol
Epi300
Adherence Assays
pCC_pOMP1.5K
pOMP1 ORF
with kanamycinR
cassette
~2.3kb
pCC1.1
Chloramphenicol
Epi300
Electroporation into
F. Tularensis
pET_pOMP1
pOMP1 ORF
~1kb
pETcoco
Chloramphenicol
Epi300 and
tuner
Antibody
development
Adherence Assay
The cell lines A549, Chang or NCI were cultured in TC medium, and once
confluent the cells were detached from the bottom of plastic flasks by adding 2 ml
polyvinylpyrrolidone-EGTA-trypsin solution (PVP; 1% polyvinylpyrrolindone, 0.02%
EGTA, 0.02% trypsin-0.02% EDTA) for 5 min at 37°C. Eight ml of tissue culture
medium supplemented with antibiotics were added to the detached cells, and 2 ml were
then removed to seed a new flask (containing 10 ml of tissue culture medium) in order to
maintain the cell line for subsequent experiments. The remaining cells were counted
using a hemocytometer, diluted, and 0.5 ml of ~1 x 105 cells were used to seed
individual wells of a 24 well TC plate (Greiner Bio-one). Following overnight
incubation, cells were washed with HEPES-buffered saline solution (HBSS; 0.2 M
HEPES, 0.12 M NaCl, 1.0 mM Glucose, 0.01 M Na2HPO4, 0.01 g phenol red, pH 7.5),
and 0.5 ml tissue culture medium lacking antibiotics was added to the monolayers.
Bacterial cells were prepared as follows: F. tularensis LVS and subsp. novicida
were grown on a chocolate agar plate for 16-36 h, resuspended in 5 ml phosphate
buffered saline (PBS) to a cell density of 230 Klett units (~109 CFU/ml). Epi300 E. coli
containing the FopA_ORF and pOMP1_ORF were grown in 5 ml Luria-Bertani (LB)
broth supplemented with 15 µg/ml chloramphenicol overnight with shaking (~200 rpm),
and the next day subcultured in 20 ml of LB medium containing 15 µg/ml
chloramphenicol and 250 µl of copy control 1000x inducer (Epicentre) in order to
increase plasmid copy number per cell for increased expression of the protein. Cells
were incubated for an additional 2 h at 37°C with shaking (~300rpm), then centrifuged at
29
3000x g and resuspended in 1 ml of PBS. The recombinant bacteria were diluted to 230
Klett units in 5 ml of PBS. Epi300 cells containing the pCR_FopA.5 and pCR_pOMP1.5
were grown overnight on LB agar supplemented with 100 µg/ml ampicillin. Bacterial
cells were scraped from the plate and resuspended in 5 ml of PBS to a cell density of 230
Kletts.
Twenty-five µl (~107 CFU) of bacterial suspension, at 230 Klett, were added to
each well of epithelial cells and plates were centrifuged for 5 min at 165xg at room
temperature, in order to facilitate binding of bacteria to the epithelial cells. Infected
monolayers were incubated for 5 min, 15 min, or 3 h at 37°C with 7.5% CO2. Following
incubation, monolayers were washed with 0.5 ml PBS supplemented with 0.15% gelatin
(PBSG) five times to remove non-adherent bacteria. For visual assays, cells were fixed
with 1 ml of 100% methanol for 5 min at room temperature. Cells then were stained
with 0.5 ml Giemsa stain (diluted 1:20), incubated for 30 min at room temperature,
washed with 0.5 ml of water to remove excess stain, and then viewed under a
microscope. The prokaryote to eukaryote ratio was counted per field. Ten fields were
counted per well and the average of all fields was determined. All experiments were
performed with duplicate samples and repeated at least once.
For viable assays, various dilutions of bacterial inoculums were plated prior to
adherence assay. Following the assay, cells were washed as described above and
incubated with a solution containing the detergent saponin (10% 10x PBS, 0.084%
EDTA, 0.152% gelatin, 0.52% saponin) for 5-10 min at 37°C, and vigorously mixed by
pipetting. Various dilutions of recovered bacteria were plated. Following the growth of
30
the bacteria, percent adherence was determined by comparing the number of cells on the
plates from dilutions from the inoculums with the number of the cells on the plates from
dilutions after the co-incubation. These results are expressed as percent (± standard
error) of bacteria that adhered to the monolayers. All experiments were performed with
duplicate samples and repeated at least once.
Biotinylation
Biotinylation was performed with the membrane impermeable reagent sulfoNHS-LC-biotin (EZ-link, Pierce). Francisella tularensis LVS and subsp. novicida were
grown for 16-36 h on chocolate agar plates and resuspended in 5 ml PBS to a cell density
of 230 Klett (~108 CFU). Bacteria were washed by pelleting and resuspending in 1 ml of
PBS three times (12,700x g for 1 min). Following the last wash bacteria were
resuspended in 0.5 ml of PBS then treated with 250 µL of 10 mg/ml of sulfo-NHS-LCbiotin (dissolved in sterile H2O) to biotinylate the amine groups of surface exposed
proteins. Bacteria were incubated at room temperature for 30 min. Cells were
centrifuged at 7,500x g for 1 min and washed with 1 ml of salt solution (50 mM tris, 300
mM NaCl) (7,500x g, 1 min), followed by two washes with PBS (1 ml PBS, 7,500x g, 1
min). Following the second wash bacteria were resuspended in 50 µL of PBS to prevent
clumping when the cells are lysed with the addition of 500 µL B-PERII (bacterial protein
extraction reagent, Pierce). In order to more completely lyse bacteria, cells were frozen
(-80°C) and thawed (70°C) three to four times and centrifuged at 15,200x g for 1 min.
The supernatant, containing biotinylated proteins, was transferred to a new 1.5 ml tube
and 200 µl of UltraLink immobilized streptavidin beads were added. Tubes were
31
incubated for 30 min at room temperature, with gentle rocking, in order to bind the
biotinylated proteins to the beads for purification. Cells then were washed (1 mL, 320x
g, 1 min) with Tris-buffered saline (TBS) supplemented with Tween-20 (50 mM NaCl,
25 mM Tris, 200 µl/l Tween-20, pH 7.5) five times. Following the last wash, proteins
were recovered by resuspending beads, in 25 µl of 2x tricine SDS sample buffer
(Invitrogen)(25 µl buffer and 25 µl water). Samples were heated at 95°C for 10 min and
centrifuged at 320x g to separate supernatant from beads. Fifteen µl of supernatant were
electrophoresed on a 16% tricine gel and stained with coomassie blue or silver stain using
the Pierce SilverSNAP Stain Kit II, when more sensitive staining was required.
Plasmid Construction
pCC1 and pCR2.1 Plasmids
To clone genes of interest into E. coli, primers were designed using Vector NTI.
The first set was designed to amplify the open reading frame (ORF) of fopA (FopAORF1,
5’GCTTAGTATCCTAGTATCAT 3’, FopAORF2, 5’TTTAACATAGTTTGTTGTA 3’)
for the adherence assays. FopAORF1 hybridizes 104 bp upstream of the ATG start
codon, while FopAORF2 lies 250 bp downstream of the stop codon, resulting in an
amplification product of 1548 bp. The other set consists of the ORF plus ~2.5 kb of
flanking DNA on either side, for transformation into F. tularensis LVS or subsp.
novicida, (FopA1, 5’TCTTTGACACCTACTCGCCA3’, FopA2,
5’GCTGGGGATATTGGTGAGCT 3’). These primers produce a 5030 bp product.
FopA1 is located 1994 bp upstream of the start codon and FopA2 is 1881 bp downstream
of the stop codon (Figure 3a).
32
Figure 3. Plasmids Maps: fopA and pOMP1
A. FopA inserted into the pCR2.1 vector. Thin arrows indicate ORFs surrounding fopA
(thick arrow). The primers used for cloning are also shown and labeled appropriately
FopA1 and FopA2 are ~2 kb up or down stream, respectively, from the ORF. B.
pOMP1 inserted into the pCR2.1 vector. Thin arrows indicate ORFs surrounding
pOMP1. The primers used for cloning are also shown. The pOMP1F and pOMP1R
primers are located ~2 kb up or down stream, respectively, from the ORF. C.
pCC_pOMP1.5K. The kanamycin resistance cassette is located 529 bp downstream of
the ATG start codon. Dark arrows indicate the pOMP1 ORF primers used for
sequencing, and light arrows indicated primers used to clone the ~5 kb region from F.
tularensis LVS.
33
Figure 3
fopA1 (100.0%)
1
FopA
A.
AmpR
AmpR
FopA ORF2 (100.0
FopA
ORF
R
Kan
kanR
pcr2.1-FopA
8981 bp
FopA
FopA
FopA 2
fopA ORF1 (89.5%)
FopA ORF
1
FopA 2 (100.0%)
B.
pOMP1R
pomp1R (100.0%)
AmpR
AmpR
KanR
pcr2.1-pomp1
kanR
pOMP1
pomp1 ORF2 (100.0
8938 bp
pOMP1
pOMP1
pOMP1F
pomp1F (100.0%)
pomp1 ORF1 (100.0%)
pOMP1
34
C.
KanR
ORF
pCC_pOMP1.5K
CmR
35
Primers were also designed to amplify the ORF of pomp1 (1314 bp)
(pOMPORF1, 5’GGGGTTAATTTACAGCTGGC3’, pOMPORF2
5’CCTGCTTGGAGTCGAGAAAA3’). pOMPORF1 binds 125 bp upstream of the start
codon and pOMPORF2 is positioned 117 bp downstream of the ORF. The other primer
set consists of the ORF plus ~2.5 kb of flanking DNA, (pOMP1F,
5’ATTTCTCACCCAGATGCTGG 3’, pOMP1R, 5’CCACAGCTCCTTTGACATCC
3’). This primers result in an amplification product of 4937 bp, and pOMP1F is located
1584 bp upstream of pOMP1, while pOMP1R is positioned 2020 bp downstream of the
ORF (Figure 3b).
Genes were amplified by PCR from F. tularensis LVS chromosomal DNA. DNA
was isolated using the Invitrogen EZ DNA kit. A toothpick was used to scrape bacteria
from the plate and resuspended in 200 µl PBS. Three hundred and fifty µl of EZ DNA
kit Solution A was added and the tube was vortexed. The cells then were incubated at
65°C for 10 min, followed by addition of 150 µl of EZ DNA Solution B. Cells were
vortexed and 500 µl of chloroform was added. Cells were vortexed again and centrifuged
for 10 min at 15,200x g. The supernatant was transferred to a new 1.5 ml tube, 1 ml of
100% ethanol was added, and the tube was inverted several times. The solution was
incubated at -20°C for 20 min and centrifuged for 15 min at 15,200x g. The supernatant
was removed, 500 µl of 70% ethanol was added, and the solution was vortexed, then
centrifuged for 5 min at 15,200x g. Supernatant was removed and the pellet was air dried
for ~10 min and resuspended in 100 µl of sterile water. The DNA was electrophoresed
on a gel in order to estimate the concentration.
36
One µl of undiluted, as well as 4 µl of diluted (1:10, 1:20, 1:40) chromosomal
DNA were used as template in the PCR reaction. Platinum Pfx DNA polymerase
(Invitrogen) was used instead of Taq polymerase to increase the fidelity of the reaction (1
µl MgSO4 buffer, 10 µl 10x PFX buffer, 1 µl of 10 mM dNPTs, 100 ng forward
primer,100 ng reverse primer, 5 µl enhancer, 22 µl sterile water, 1 µl PFX, DNA). All
primers required the same annealing temperature of 50°C. The extension times differed
according to the amplification region (2 min for the ORF primers, 6 min for the 5 kb
primers). The TA Topo cloning kit (Invitrogen) was used to construct pCR_FopA.5 and
pCR_pOMP1.5. PCR products were ligated into the pCR2.1 vector, which contains an
ampicillin resistance cassette. The plasmids were then transformed into EPI300 E. coli.
Colony PCR using vector-based primers was performed to ensure insertion. For the
development of the plasmids pCC_FopA.5, pCC_pOMP1.5, FopA_ORF and
pOMP1_ORF (all in pCC1 vector), PCR-generated DNA was precipitated using PCR
precipitation solution (Epicentre) and electrophoresed on an agarose gel to both purify
and estimate concentration. End-repair and ligation of the PCR products into pCC1
vector were performed using the CopyControl PCR Cloning Kit (Epicentre technologies).
Between 100-200 ng of ligation was transformed into EPI300 E. coli.
All plasmids then were transformed into electrocompetent Epi300 E. coli.
Electroporations include ~200 ng of plasmid, 25 µL of E. coli, and 100 µL of 10%
glycerol. This mixture was placed in a 1 mm cuvette and electroporated (1.6 kV). Cells
were recovered by the addition of 950 µL of SOC and incubated at 37°C with shaking
(~200 rpm) for 1 h. Cells then were pelleted, resuspended in 200 µl medium, and plated
37
on LB agar supplemented with 15 µg/ml chloramphenicol. The colonies were screened
by colony PCR the following day using primer sequences found within the pCC1 vector
(pCCRP2 and pCCT7) or the pCR2.1 vector (M13F and M13R) to ensure insertion of the
gene. For plasmids that contain ~5 kb DNA inserts (pCR_FopA.5, pCR_pOMP1.5,
pCC_FopA.5, pCC_pOMP1.5) the product is expected to be ~6.5 – 7 kb when
amplifying with the vector-based primers. The amplicon for ORF plasmids
(pOMP1_ORF and FopA_ORF) is expected to be ~1.3 kb. In all cases if there is no
insertion present the amplification will result in a small fragment (~300 kb).
The orientation of the gene was also verified to allow the FopA and pOMP1 gene
product to be expressed in E. coli. This was performed via colony PCR using one vectorspecific primer (pCCRP2, which is located near a vector borne promoter), and one primer
that is located within the insert, at the C-terminal end of the gene. Correct insertion of
the gene will result in an amplification product, but if inserted in the incorrect orientation,
the insert will not be amplified.
pETcoco Plasmid
Primers were designed by Invitrogen custom primers (pOMPF24AscI,
5’TTGGCGCGCCACATGGAAATATATACCGTT 3’, and pOMPRpacI, 5’
CCTTAATTAAAAATATTTATCTCTAAGTTT 3’). AscI and PacI restriction enzyme
sites were included at the end of the primers, as underlined. Primers were designed inframe relative to the restriction sites in the vector, in order for the protein to be expressed.
The ORF of pOMP1 was amplified with PFX, as described above, and precipitated using
PCR precipitation solution (Epicentre). Ends of the amplicon were digested with AscI
38
and PacI and incubated overnight at 37°C in order to produce sticky ends for ligation.
The product was precipitated and resuspended in 10 µl sterile water. The pETcoco vector
(Novagen) was digested with AscI and PacI for ~3 h at 37°C. T4 ligase was used to
ligate the PCR product into the vector ( 4:1 ratio of PCR:vector, T4 ligase, T4 ligase
buffer) and left at room temperature overnight. The plasmid was transformed by
electroporation into Epi300 electrocompetent cells as described earlier. This plasmid will
be referred to as the pET_POMP1. pET_POMP1 encodes the pOMP1 ORF of F.
tularensis LVS, which consists of 439 amino acids. This plasmid also adds a His-tag as
well as a S-tag. Following transformation, the colonies were screened by colony PCR
with S-tag and T7 terminator primers, which are located within the vector, to ensure the
presence of the PCR fragment within the vector. In order to guarantee the amino acid
sequence was error free, pET_POMP1 was sent out for sequencing to the University of
Michigan DNA Sequencing Core Facility (http://seqcore.brcf.med.umich.edu), and the
amino acid sequence was aligned with the known F. tularensis LVS pOMP1 sequence.
Plasmid Preparations
Cells were grown overnight with shaking (~200 rpm) in 5 ml LB broth
supplemented with 15 µg/mL chloramphenicol. The next day the cells were subcultured
in 20 ml of medium supplemented with chloramphenicol and 250 µl of 1000x copy
control inducer (Epicentre) to increase copy number for higher plasmid recovery.
Bacteria were incubated for 5 h at 37°C with shaking (~300 rpm). Cells then were
pelleted at 3000x g for 10 min and resuspended in 500 µL P1 solution (Qiagen miniprep
kit). Five hundred µl was divided equally between two 1.5 ml tubes, 250 µL of P2
39
solution was added to each, and the tubes were inverted several times. Three hundred
and fifty µL of N3 solution was added to each tube and mixed by inversion several times.
Solution was centrifuged at 12,700x g for 10 min and the supernatant was transferred to a
Qiagen miniprep tube and centrifuged to bind the plasmid to the membrane. The plasmid
was washed 3x with 750 µl of PE wash buffer (12,700x g, 1 min) and then plasmid was
eluted into a clean 1.5 ml tube with 75 µl of elution buffer. The plasmid was run on an
agarose gel to determine concentration. Samples were run with supercoiled plasmid with
a known concentration for comparison.
Transposon Mutagenesis
In order to insert the Kan-2 transposon into the plasmid, pCC1-based plasmids
(0.2 µg) containing the ~5 kb pOMP1 inserts were incubated with 1µl EZ:Tn<Kan-2>
transposon, 1µl EZ:Tn transposase, and 1µl EZ:Tn 10x reaction buffer for 2 h at 37°C.
Following the incubation, 1 µl 10x stop solution was added and the tubes were incubated
at 70°C for 10 min to inactivate the enzyme. One µl of plasmid was transformed into 25
µl Epi300 chemically competent cells and were co-incubated on ice for no less than 30
min, heat shocked at 42°C for 30 s and put back on ice for 2 min. Nine hundred and fifty
µl of SOC media was added and cells were left to recover for 1 h at 37°C with shaking
(~200 rpm), then plated on LB agar containing both µg/ml kanamycin and 15 µg/ml
chloramphenicol. The next day, colonies were screened by colony PCR using ORF
primers to identify colonies that had the Kanamycin cassette (1.3 kb) inserted within the
pOMP1. After screening ~50 colonies, one colony, clone #33, had the cassette inserted
within pOMP1 (in order to knockout expression of the protein). Clone #33, will be
40
referred to as pCC_pOMP1.5K (Figure 3c). It was identified on an agarose gel by
increased size of the fragment amplified with ORF primers (without cassette size is ~1
kb, with cassette size is ~2.3 kb). The ORF of pCC_pOMP1.5K was sequenced to
determine the location of the kanamycin cassette.
Transformation
Francisella tularensis LVS cells were grown for 36 h on chocolate agar plates.
The bacteria were resuspended in 5 ml of PBSG and vortexed. Cells were washed once
with cold 1 mM HEPES buffer (4°C, 3000x g, 10 min) then washed once with 10%
glycerol. Cells were resuspended in 0.5 ml of 10% glycerol and transferred to a 1.5 ml
tube, then centrifuged for 2 min at 12,700x g. The pellet was resuspended in 100 µl of
cold 10% glycerol. Varying amounts of precipitated PCR products (~6 kb region and
ORF alone, from pCC_pOMP1.5K), as well as whole pCC_pOMP1.5K were used for
electroporation. The plasmid or PCR were mixed with 15 µl of electrocompetent cells
and 100ul of 10% glycerol and immediately electroporated at 1.6 kV in a 1 mm cuvette.
Todd Hewitt Broth (200 µl) was added to cells and they were spread onto chocolate
plates with no antibiotics overnight for recovery. The next day all cells were scraped and
resuspended in 1 ml of PBSG. One hundred µL of cells were spread on 5-10 chocolate
plates supplemented with 7.5 µg/ml kanamycin. Cells were incubated for >48 h, until
growth was seen.
Cryotransformation also was used in order to insert pCC_pOMP1.5K into F.
tularensis LVS. Francisella tularensis LVS was grown for ~36 h on chocolate agar.
Cells were removed from the plate and resuspended to a cell density of 230 Kletts in 5 ml
41
of 0.2 M KCl. Twenty five µl of bacteria were added to 25 µl of transformation buffer
(0.2 M MgSO4, 0.01 M Tris-actate, pH 7.5), followed by the addition of varying amounts
(100 - 700 µg) of plasmid 33, or precipitated PCR product (of ORF and 5 kb region).
The solution was incubated at room temperature for 5 min and then frozen in liquid
nitrogen for 5 min, followed by incubation at 37°C for 5 min. Two hundred µl of ToddHewitt broth was added and the 100 µl of cells were plated on non-selective chocolate
agar and incubated overnight at 37°C with 7.5% CO2. The next day cells were removed
for the plate, resuspended in 1 ml of PBSG and replated on chocolate agar supplemented
with 7.5 µg/ml of kanamycin.
Sequencing
An ABI310 sequencing machine (Perkin Elmer) was used to sequence all
plasmids. Half reactions were performed by using 4 µl terminator ready reaction mix
(ABI), 200-500 ng plasmid DNA, 3.2 pmol primer, and deionized water up to 10 µl. This
reaction was amplified for 25 cycles as follows: 96°C for 10 s, 50°C for 5 s, and 60°C
for 4 m. The reaction was transferred to a 0.65 ml sequencing tube containing 32 µl of
95% ethanol and 6 µl of sterile water. The tubes were left at room temperature for 10-15
min to allow precipitation, and then centrifuged at 15,200x g for 30 min. The supernatant
was removed and the pellet was heated to 90°C for 1 min in order to dry. The pellet was
resuspended in 18 µl of TSR (template suppression reagent, PE Applied Biosystems) and
briefly vortexed. Finally, the sample was heated to 95°C for 2 min to denature and
chilled on ice for 10 min. The sample was placed in the ABI 310 Sequencer and left until
complete.
42
Protein Purification
PET_POMP1 (pOMP1 pETCOCO plasmid) was transformed into chemically
competent Tuner cells (Novagen) for in order to express the pOMP1 protein. Cells were
grown overnight in LB containing 0.2% glucose (to inhibit arabinose and keep plasmid at
a copy number of one) and 15 µl g/ml chloramphenicol. Colony PCR using vector
primers was performed to ensure the presence of the plasmid. Tuner cells containing
PET_POMP1 were grown in 5 ml of LB supplemented with 15 µg/ml chloramphenicol
and 0.2% glucose. The next day cells were subcultured into 20 ml LB (supplemented
with 15 µg/mL chloramphenicol and 0.2% glucose) and were grown for 1 h with shaking
(250 rpm). The optical density of the cells was read, 1ml was removed, and 3x SDSPAGE sample buffer was added (uninduced sample). Twenty five µl of 1 M isopropylbeta-D-thiogalactopyranoside (IPTG) was added to the remainder of the culture to induce
expression of the gene and cells were incubated for an additional 4 h. The OD was read,
and 1 ml of cells was recovered and added to 3x SDS-PAGE sample buffer (induced
sample). The remainder of the culture was centrifuged for 15 min at 4000x g, and the
pellet was resuspended in BugBuster (Novagen) plus 1 µl of recombinate lysozyme
(Novagen), in order to break open the cells. The mixture was lightly vortexed and
incubated at room temperature with gently rocking for 30 min to free proteins from the
cell. To separate the soluble protein from the proteins in inclusion bodies the solution
was centrifuged at 21,000x g for 20 min at 4°C. To determine whether pOMP1 is soluble
or insoluble, the following samples were electrophoresed for 1 h at 100 V on a 10%
acrylamide SDS-PAGE gel: uninduced fraction, induced fraction, soluble/insoluble
43
fraction, soluble fraction. The gel was stained with coomassie blue and pOMP1 was
located in both the soluble/insoluble lane as well as the soluble lane, therefore, pOMP1 is
a soluble protein.
Using the Novagen His-Bind Resin Kit, the pOMP1 was further purified. The
His-Bind Resin slurry was first treated as follows, in order to prepare it for binding with
the His-tag located in PET_POMP1. One ml of His-Bind Resin was added to a 2 ml
centrifuge tube and centrifuged for 1 min at 1000x g. The resin was washed (1 ml of
solution, incubated for 5 min, centrifuged at 1000x g for 1 min) with 1 ml of sterile water
two times, followed by similar three washes with 1x charge buffer. Next, the resin was
washed twice with 1x binding buffer pH 7.9. Following the second wash, 1 ml of 1x
binding buffer was added to the soluble pOMP1 fraction and incubated at room
temperature for 60 min while rotating. The solution was centrifuged for 1 min at 1000x g
and washed (1.5 ml buffer, centrifuged at 1000 x g for 1 min) with 1x binding buffer
three times. The pellet was washed with 1x wash buffer + 10 mM imidazole (pH 7.9)
twice followed by two washes with 1x wash buffer + 20 mM imidazole (pH 7.9). The
protein was eluted by the addition of 1 ml of 1x elute buffer (pH 7.9) and was rotated for
30 min. The solution was centrifuged for 1 min at 1000 x g, and the supernatant stored at
-20°C (elute 1). The same elution process was followed again and the supernatant was
stored at -20°C (elute 2). Two 10% SDS PAGE gels were run with the soluble protein
(from outer membrane preparation) and elutes 1 and 2. One gel was stained with
coomassie blue, and a western blot was preformed on the other using a histidine-specific
antibody.
44
RESULTS
Adherence of F. tularensis Subspecies to Human Lung Cell Line
In our model of F. tularensis pathogenesis, we have proposed that adherence to
human lung epithelial cells is critical for virulence. To test this, we have examined the
adherence of F. tularensis LVS and subsp. novicida to a cell line derived from human
alveolar epithelial cells (Figure 4). As stated earlier, F. tularensis subsp. holartica, from
which the live vaccine strain (LVS) was derived, has an estimated LD50 of <103 CFU in
humans (Ellis et al., 2002) and can cause ulceroglandular tularemia, while subsp.
novicida rarely causes disease in humans. Both subspecies, holartica and novicida, are
fully virulent in mice. As illustrated in the visual assay, F. tularensis LVS adheres
significantly to the human alveolar lung cell line, A549 (Figure 4A). In contrast, subsp.
novicida showed very little adherence (Figure 4B). Bacterial cells were incubated with
epithelial cells for either 15 min or 3 h. Results were similar for both incubation times,
so 15 min incubations were performed for all subsequent experiments.
Adherence was quantitated by Giemsa staining the monolayers and directly
counting bound F. tularensis in random microscopic fields from each well. All
experiments were performed in triplicate, and the average bacterial counts (± standard
error) were calculated. F. tularensis LVS and subsp. novicida showed significantly
different adherence (Figure 5). Furthermore, F. tularensis LVS has slightly increased
adherence when grown for 36 h prior to inoculating than when grown just 16 h (Figure
5). At both growth times, F. tularensis LVS shows significantly greater adherence to
45
Figure 4. F. tularensis LVS and subsp. novicida Visual Adherence Assay
In each case bacteria were co-incubated for 15 min.
A. Adherence of F. tularensis LVS to human alveolar epithelium stained with Giemsa.
B. Adherence of subsp. novicida to human alveolar epithelium stained with Giemsa.
46
Figure 4. F. tularensis LVS and subsp. novicida Visual Adherence Assay
A. F. tularensis LVS
B. Subspecies novicida
47
Figure 5. Adherence of F. tularensis LVS and Subsp. novicida to a Human Lung Cell
Line
Results are expressed as bacteria per microscopic field, either of F. tularensis LVS or
subsp. novicida, bound to A594 cells after a 3 h incubation. F. tularensis LVS was
grown for either 36 h or 16 h prior to adherence assay. The two bars shadings indicate
results from two experiments, each with triplicate samples. Only one experiment was
performed for F. tularensis LVS grown for 16 h prior to adherence assay.
48
Figure 5. Adherence of F. tularensis LVS and Subsp. novicida to Human Lung Cells
ND
49
A549 cells than subsp. novicida. Therefore, F. tularensis LVS was grown for ~36 h
prior to the assay for all remaining experiments.
Since the visual adherence assays do not indicate viability of bound cells, we
tested the adherence of F. tularensis LVS and subsp. novicida by assaying the recovery
of viable cells. Prior to the adherence assay, both F. tularensis LVS and subsp. novicida
were resuspended in PBS to a cell density of ~107 CFU, serially diluted and plated on
chocolate agar in order to determine the cell count of the inoculums. Twenty five µl of
the bacteria (~107) were incubated with ~105 A549 cells for 15 min and following the
incubation the monolayers were washed as normal then treated with saponin in order to
disrupt the epithelial cells. Serial dilutions of recovered bacteria were plated on
chocolate agar and incubated. The number of colonies, for both the inoculums and the
recovered bacteria were counted, and the number of recovered cells was normalized.
These experiments were performed in duplicate and repeated on two separate occasions.
In both cases, F. tularensis LVS demonstrates significantly greater adherence than subsp.
novicida (Figure 6). These results are expressed as recovered bacteria that adhered to the
monolayers.
To determine whether F. tularensis could adhere to other human epithelial cell
lines we compared the ability of F. tularensis LVS and subsp. novicida to adhere to three
different cell lines, A549 (human lung epithelial cells), Chang (human conjunctiva
epithelial cells) and NCI-H292 (mucoepidermoid lung epithelium) cells. Adherence
assays were again repeated for 15 min or 3 h, and similar results were seen at the two
times. Adherence was significantly less for F. tularensis LVS incubated with Chang or
50
Figure 6. Adherence Assay Viable Cell Count
Quantitative adherence assays. Following co-incubation, the monolayers were treated
with saponin, and bacterial cells were recovered and plated at different dilutions on
chocolate agar. The recovered bacteria were counted. The black bar and the striped bar
represent two independent experiments.
51
Figure 6. Adherence Assay Viable Cell Count
52
NCI-H292 cell lines than A549 cells. Figure 7 shows the visual assay results, where 10
random fields were viewed per well. Similar trends were seen when viable assays were
performed (data not shown). No adherence of subsp. novicida was seen to any of the cell
lines. In summary, F. tularensis LVS demonstrates significantly greater adherence than
subsp. novicida to the cell line A549. To determine what is mediating this adherence, we
identified specific proteins that are expressed on the outer membrane of F. tularensis
LVS.
Identification of F. tularensis Surface Proteins
Most bacterial adhesins are membrane proteins containing extracellular domains
that specifically interact with host cell ligands (Oyston et al., 2004). To help identify
putative adhesins we used a membrane-impermeable biotinylation reagent (Sulfo-NHSLC-Biotin, Pierce, Ill) to specifically biotinylate the amine groups of surface exposed
proteins. The biotinylated proteins then were purified using avidin-conjugated sepharose
beads. The recovered proteins were resolved on SDS-PAGE tricine gels and visualized
with Coomassie blue. At least 12 distinct bands can be seen in the lanes using
biotinylated cells (Figure 8, Lane 1-3). In contrast, only one darkly staining and a few
faintly staining bands are visible in the non-biotinylated control lane (Figure 8, Lane 5).
In collaboration with our core proteomics facility, we have identified nine of the
biotinylated proteins from F. tularensis LVS using tandem mass spectroscopy and the
genome sequence of F. tularensis Schu 4. The identified proteins are summarized in
Table II.
53
Figure 7. Adherence of F. tularensis to Different Human Cell Lines
Visual assay of the adherence of F. tularensis LVS (striped) or subsp. novicida (black) to
three different human cell lines: NCI-H292, Chang, or A549, and incubated for 3 h.
Results are expressed as average bacteria per microscopic field, ± standard error. Ten
fields for each of the triplicate samples were counted.
54
Figure 7. Adherence of F. tularensis to Different Human Cell Lines
55
Figure 8. Purification of Surface Proteins from F. tularensis
Francisella tularensis LVS was surface labeled with the membrane impermeable
biotinylation reagent Sulfo-NHS-LC –Biotin. Proteins were extracted with B-PerII,
purified with avidin-conjugated sepharose beads and separated on a 10%-20% Tricine
SDS-PAGE gel that was stained with Coomassie blue. Over 12 distinct proteins bands
are visible from biotinylated F. tularensis (+Bio, the same sample was loaded in
triplicate). Very little protein was purified from non-biotinylated F. tularensis control (BIO). A similar pattern is observed with subsp. novicida.
56
Figure 8. Purification of Surface Proteins from F. tularensis
57
Table II. Summary of Identified Surface Proteins.
Protein
Function
Surface
Associated
DnaK
Protein
Chaperone
Possibly
Surface associated in Coxiella,
Mycoplasma and Haemophilus
Erricson et al., 1994
Erricsson, et al., 1997
Zuber, et al., 1995
GroE
Protein
Chaperone
Possibly
Surface associated adhesion in
Legionella, Clostridium and
Chlamydiae
Erricsson et al., 1997
Garduno et al., 1998
Hennequin et al., 2001
KatG
CatalasePeroxidase
Perimlasmic?
Secreted?
Secreted by Mycoplasma
Sonnenberg et al., 1997
pOMP1
Unknown
Probably
Signal sequence
LysM domain
This work
pOMP2
Unknown
Unknown
Downstream of pdpA and B
Gray et al., 2002
Kapartal et al., 2002
IglA
Unknown
Unknown
First gene in a five gene operon
Gray et al., 2002
FopA
Adhesin?
Porin?
Adhesin?
Porin?
P. aeruginosa, M. catarrhalis homologs
are adhesins
Leslie et al., 1993
Nano et al., 1988
Biotin
Binding
Protein
Biotin
Transport
No
Covalently linked to biotin
bacterioferritin
Iron
sequestration
possibly
Role is not clear
Notes
References
Andrews et al., 2003
Two of these proteins, FopA and pOMP1, were examined further. FopA is a
known outer membrane protein of F. tularensis that has been studied for use as a vaccine
candidate, although its function is unknown (Fulop et al., 1996; Leslie et al., 1993).
FopA can also be expressed (Figure 9) and exported to the outer membrane of E. coli
(Nano, 1988). These features make FopA a good candidate to study further as a possible
F. tularensis adhesin.
Putative outer membrane protein 1 (pOMP1) is a 49.3 kDa protein. Using SignalP
1.1, DAS and the conserved domain (CD) database, we have identified a potential signal
sequence, two transmembrane domains and a LysM domain (Figure 10). These
properties, in addition to its ability to be labeled by Sulfo-NHS-LC-Biotin support its
localization to the outer membrane. The LysM domain is thought to function as general
peptidoglycan binding module (Bateman and Bycroft, 2000).
In summary, nine putative F. tularensis LVS OMPs have been identified. One of
the important roles that OMPs play is adherence to the host. There are two main ways to
test for adherence, fist it is possible to disrupt the putative adhesin in vivo and look for
decreased adherence, second it is possible to clone the putative adhesion into a nonadherent bacterium, such as E. coli and look for enhanced adherence. Preliminary data
suggests that FopA does not adhere to epithelial cells when expressed in a non-adherent
bacterium, and therefore, we did not attempt to transform the fopA mutant in vivo. This
will be discussed further in the next section.
As stated, one way to test for adherence is to disrupt the putative adhesion in vivo.
In order to eliminate expression of the protein in vivo, pOMP1 must be inactivated. This
59
Figure 9. SDS-PAGE of Recombinant E. coli
Recombinant E. coli cells were lysed and electrophoresed on a 10% tris-glycine gel to
assess protein expression. There is no obvious expression of the pOMP1 gene, or lack of
expression of pCC_pOMP1.5K. Both FopA and McaP are being expressed, as indicated
by the white ovals. The arrow indicates where pOMP1 would be expected to run on the
gel.
60
Figure 9. SDS-PAGE of Recombinant E. coli
61
Figure 10. pOMP1
Putative outer membrane protein 1 (pOMP1) is a 49.3 kDa protein. Using SignalP 1.1,
DAS and the conserved domain (CD) database we have identified a potential signal
sequence (Met1-Ser23), two transmembrane domains (Val8-Val21 and Ser196 –Gly211)
and a LysM domain (Tyr27– Pro81).
62
Figure 10. pOMP1
tm Val8-Val211
Tyr27-Pro81
LysM
signal sequence
Met1-Ser23
d
tm Ser196-Gly2112
i
438aa, 50 KDa
63
was done by inserting a kanamycin resistant gene into the ORF of pOMP1, after
amplifying the ORF plus flanking regions (~2 kb) using the pOMP1F primer located
~1.6kb upstream of the start codon and pOMP1R located ~2.0 kb downstream (Figure
3b). This gene was ligated into a CopyControl vector, pCC1, which allows high
expression of the cloned protein when inducer is added. Colony PCR and DNA
sequencing were used to verify the insertion of the gene into the vector. By using the
EZ:Tn<Kan> transposon kit (Epicentre), a kanamycin resistance cassette was randomly
inserted into pCC_pOMP1.5. Colonies were screened to find a clone in which the
kanamycin cassette was inserted into the pOMP1 ORF. The pCC_pOMP1.5K plasmid
was identified by colony PCR with ORF specific primers. The primers used typically
amplify ~1.3 kb, but with the addition of a kanamycin cassette (1.3 kb), the amplicon is
~2.6 kb (Figure 11). This region was sequenced to determine where the kanamycin
cassette was located in the ORF (Figure 3c). The kanamycin cassette was inserted 529 bp
downstream of the ATG start codon.
In order to eliminate activity of the pOMP1 gene in situ, the insert from
pCC_pOMP1.5K must undergo a double crossover event with the wild-type gene of F.
tularensis LVS. In theory, this should be accomplished by transforming the plasmid into
F. tularensis LVS and relying on recA-mediated recombination. Both electroporation
and cryotransformation were used to attempt to make this knockout. The DNA from
pCC_pOMP1.5K was prepared utilizing different protocols. Using the pOMP1 ORF
primers as well as the pOMP1 forward and reverse primers, (which amplify the ORF plus
flanking regions) of pCC_pOMP1.5K, the insert was amplified. Following amplification,
64
Figure 11. pCC_pOMP1.5K
Using the EZ:Tn<Kan> transposon kit (Epicentre), a kanamycin cassette (1.3 kb) was
randomly inserted into pCC_pOMP1.5. Colony PCR was performed using ORF specific
primers to identify a clone in which the cassette was inserted into pOMP1. The arrow
indicates clone #33 (pCC_pOMP1.5K), in which the kanamycin cassette was inserted
(~2.6 kb).
65
Figure 11. pCC_pOMP1.5K
3 kb
1.6 kb
1 kb
66
the product was precipitated using DNA precipitation solution (Epicentre) and
resuspended in 10 µl of sterile water. Varying amounts of the resuspended product (l µl 10 µl), as well as dilutions of the product (1:5, 1:10), were used to transform the pOMP1
kanamycin DNA from E. coli into F. tularensis LVS. We also attempted to electroporate
the complete pCC_pOMP1.5K into F. tularensis LVS. However, we were unable to
make a pOMP1 mutant in F. tularensis LVS. Because we were not able to directly
eliminate activity of pOMP1 in F. tularensis LVS, an alternative approach for testing
adherence was examined.
Adherence Assays: Alternative Approach for Testing Adherence to Human
Epithelial Cells
Since we were unable to create a F. tularensis pOMP1 chromosomal mutant, we
used an alternative approach to testing its potential as an adhesin. FopA and pOMP1
were cloned into vectors (pCR2.1 and pCC1) and used to transform E. coli, a nonadherent bacterium, to test for increased adherence due to the expression of the pOMP1.
This approach has been used successfully for several other bacteria (Timpe et al., 2003).
FopA and pOMP1, each with ~2 kb flanking regions, were cloned into pCR2.1 and tested
for adherence to human epithelial lung cells (A549). The positive control used in this
assay is McaP, an adhesin of Moraxella catarrhalis (cloned into pCC1). Two negative
controls, pCC1 vector and pCR2.1 vector, were used in this experiment. Monolayers
were stained with Giemsa and cells were viewed under a microscope. The number of
prokaryotic cells bound per eukaryotic cell were calculated in 10 random fields per well.
This experiment was performed in duplicate. The E. coli expressing FopA (pCR_FopA1)
demonstrated no significant adherence to the human epithelial cells (A549). However, E.
67
coli transformed with pCR_pOMP1.5 demonstrated significant adherence to A549 cell
(Figure 12). Similar trends were seen in viable counts (data not shown). Although the
binding of E. coli containing pCR_pOMP1.5 to A549 cells is statistically significant
relative to what we observe for the vector alone, the absolute level of binding is relatively
low. Thus, the importance of the binding during infection remains unclear.
One possible reason for the lack of adherence could be due to the protein not
being sufficiently expressed or post-translationally modified in E. coli. To increase
expression the ORFs of FopA and pOMP1 were cloned into pCC1 (FopA_ORF and
pOMP1_ORF), to control copy number, and used to transform Epi300 E. coli. The E.
coli was induced prior to the adherence assay to highly express each protein. Visual
adherence assays were performed with A549 cells and the results are expressed as
prokaryotes per eukaryotes as described previously. Epi300 E. coli with FopA_ORF
demonstrated no significant difference in adherence from the negative control, pCC1
(data not shown). Epi300 cells with pOMP1_ORF again adhered significantly more than
the negative control (Figure 13a). Again, the physiological significance of this binding
remains unclear due to its relatively low level. Figure 13b illustrates the difference in
binding between 15 min and 3 h co-incubation with monolayers: there is no significant
difference.
To make certain that the E. coli was expressing the proteins, whole cell lysates of
the E. coli cells containing FopA_ORF or pOMP1_ORF were electrophoresed on a 7.5%
tris-glycine gel. Figure 9, lanes two and four, illustrate the expression of both FopA and
the positive control, McaP. However, there is no clear band indicating pOMP1
expression. The arrow indicates where pOMP1 would be expected. From these data, we
68
Figure 12. Adherence of Recombinant E. coli, Adherence was Measured Via Visual
Assessment
Recombinant bacteria were grown in LB broth and induced to express the cloned gene.
Unbound bacteria were washed off A549 cells after 15 min incubation, and the
monolayers were stained with Giemsa.
69
Figure 12. Adherence of Recombinant E. coli, Adherence was Measured Via Visual
Assessment
70
Figure 13. Adherence of E. coli Transformed With pOMP1
Recombinant bacteria were grown in LB broth and induced to express the cloned gene.
Unbound bacteria were washed off A549 cells after 5 min incubation, and the monolayers
were stained with Giemsa. A. Adherence of pOMP1_ORF compared to the negative
(pCC1 vector alone) and the positive control (McmA cloned and expressed in pCC1). B.
The experiment from 13A was repeated to compare the binding of the recombinant
bacteria after 15 min or 3 h. The white bars indicate a15 min incubation and the shaded
bars indicate a 3 h incubation.
71
Figure 13a. Adherence of E. coli Transformed with pOMP1
72
Figure 13b. Adherence of E. coli Transformed with pOMP1, 15 min and 3h
73
can conclude that FopA is most likely not an adhesin for F. tularensis LVS. However,
we cannot conclude that pOMP1 does not mediate adherence of F. tularensis LVS to
human lung cells due to the possible lack of expression as well as the fact that pOMP1
demonstrated experimental significance.
Although further characterization is needed to determine if pOMP1 mediates F.
tularensis adherence to human lung cells, it is clear that pOMP1 is a novel OMP
identified on the surface of F. tularensis. Currently, antibodies are being produced in
mice in order for additional examination of pOMP1 role as an outer membrane protein.
We intend to then assess whether anti-pOMP1 anitsera can block adherence by either F.
tularensis LVS or E. coli producing pOMP1.
74
DISCUSSION
Although research into F. tularensis pathogenesis has recently increased, the
virulence mechanisms used by the bacterium have yet to be fully uncovered. F.
tularensis differs from other Gram-negative bacteria in that its genome lacks key
virulence components, such as recognizable type-III, IV or V secretion systems or a
TonB-dependent system (for acquisition of complex iron), which are typically seen other
pathogens (Larsson et al., 2005). This deficiency of known virulence factors makes it
difficult to understand how F. tularensis not only infects a cell, but also how it persists
and escapes the macrophage. The first step in further understanding the pathogenesis of
F. tularensis is to comprehend how it initially binds to a host cell during infection. This
can be studied by examining what potential roles the outer membrane proteins of F.
tularensis play in mediating adherence to the host, as well as, discovering the general
function of these proteins.
In this study, we found that F. tularensis LVS adheres to human alveolar
epithelium significantly better than subsp. novicida (Figure 4). Discovering what is
mediating this adherence will help to improve our understanding of F. tularensis
pathogenesis. As seen in many other pathogenic bacteria, OMPs have the ability to play
a role in mediating adherence to the host. For example, H. influenzae has several known
adhesins such as Hia (Ecevit et al., 2004) and Hap (Hendrixson and St Geme, 1998),
while Moraxella catarrhalis utilizes OmpCD (Holm et al., 2004), Hag (Holm et al.,
2003) and McaP (Timpe et al., 2003) for binding to human epithelium. In order to
identify OMPs on the surface of F. tularensis LVS, biotinylation was performed with the
membrane impermeable reagent sulfo-NHS-LC-biotin. Twelve potential surface proteins
75
were identified (Table II). The potential roles in F. tularensis for several of these
proteins are summarized below.
DnaK and GroE (Hsp 70 and 60) are heat shock proteins that can function as
molecular chaperones in many bacteria, including E. coli (Ericsson et al., 1994, 1997;
Garduno et al., 1998). Interestingly, both proteins also have been identified as surface
proteins in other pathogens, such as Clostridium difficile (Hennequin et al., 2001) and
Legionella pneumophila (Garduno et al., 1998) . Clostridium difficile GroEL is surface
associated and may play a role mediating adherence, since antibodies against GroEL
partially block adherence (Hennequin et al., 2001). In L. pneumophila GroEL is also
surface associated and accumulates in endosomal spaces (Garduno et al., 1998). DnaK is
surface associated in Coxiella burnetti (Macellaro et al., 1998), is part of the P1 adhesin
complex of Mycoplasma pneumoniae (Layh-Schmitt et al., 2000) and is a putative
surface adhesin in Helicobacter pylor (Huesca et al., 1998). In F. tularensis the synthesis
of both GroEL and DnaK increases in response to hydrogen peroxide (Ericsson et al.,
1994). GroEL was also found to accumulate in F. tularensis-containing phagosomes
isolated from macrophages (Kovarova et al., 2002). Finally, Ericsson et al. showed that
T cells from individuals vaccinated with live F. tularensis LVS had an increased
proliferative response to DnaK and GroEL. This suggests the involvement of the heat
shock proteins in protective immunity to tularemia (Ericsson et al., 1994). These
observations, together with our data, suggests that DnaK and GroE are surface associated
in F. tularensis and could be involved in adherence to human cells (Zuber et al., 1995).
IglA is encoded by the first gene in a five-gene operon (Gray et al., 2002). The
third gene in the operon, iglC, codes for a 23 KDa protein that is necessary for growth
76
and survival within macrophages (Kovarova et al., 2002; Lauriano et al., 2003). The
expression of iglC is increased during growth in macrophages, and the protein
accumulates in macrophage vacuoles containing F. tularnesis (Kovarova et al., 2002;
Lauriano et al., 2003). The functions of IglC and the other members of the operon are not
known, however, isogenic mutants of iglC and iglA show decreased growth in murine
macrophages (Lauriano et al., 2004).
KatG contains a conserved catalase/peroxidase domain, a signal sequence and
three putative transmembrane domains. A homologous protein in Mycoplasma is a
secreted virulence factor (Sonnenberg and Belisle, 1997). One possibility is that F.
tularensis exports KatG to its surface to protect against oxidative damage in the
macrophage phagosome.
Putative outer membrane protein 2 (pOMP2) is a 17.5 kDa protein that does not
contain a conserved signal sequence or obvious transmembrane regions. Unpublished
data by Nano et al. (Genbank accession number AY293579) suggest that the genes
around pOMP2 are necessary for intramacrophage growth, virulence in mice, and are part
of a large pathogenicity island (Kapatral et al., 2002).
We chose to further examine two of these biotinylated proteins, FopA and
pOMP1. FopA is an outer membrane protein of F. tularensis that has been studied for
use as a vaccine candidate (Fulop et al., 1996; Leslie et al., 1993). The function of FopA
is not known but its closest orthologs include the P. aeruginosa adhesin OprF and the M.
catarrhalis adhesin OMPCD (Azghani et al., 2002). Both of these proteins mediate
adherence to human lung epithelial cells, however, our data suggest that FopA does not
mediate adherence to any of the human cell lines examined, including alveolar
77
epithelium, conjuctival epithelium and mucoepidermoid lung epithelium. More research
must be done to determine the function of FopA.
Putative outer membrane protein 1 (pOmp1) is a 49.3 kDa protein composed of
438 amino acids (Genebank accession number AY774763). Using SignalP 1.1, DAS and
the conserved domain (CD) database we have identified a potential signal sequence, two
transmembrane domains and a Lysin motif (LysM) domain (Figure 10). Signal
sequences are typically located at the N-terminal of a secreted protein. This sequence is
necessary for transport through the cell membrane. The potential pOMP1 signal
sequence is 23 amino acids in length and is located at the N-terminus from Met1 to
Ser23. According to Hidden Markov Model (HMM) analysis, the signal peptide
probability is nearly 90%. The most likely cleavage site is between Ser23 and Met24.
The two transmembrane domains that were proposed, using DAS, are located from Val8Val21 and Ser196-Gly211. The LysM domain is found in many cell wall degrading
proteins (Bateman and Bycroft, 2000), but is also present in a number of bacterial
virulence proteins, including Staphylococcal IgG binding protein, the E. coli (EPEC)
adhesin intimin, and the elastin-binding protein of Staphylococcus aureus (Bateman and
Bycroft, 2000). The LysM domain is thought to function as general peptidoglycan
binding module (Bateman and Bycroft, 2000). In F. tularensis LVS, the LysM domain
contains 54 amino acids (Tyr27-Pro81). All of these domains support our suggestion that
pOMP1 is a novel F. tularensis OMP. In a sequence comparison between F. tularensis
LVS pOMP1 and subsp. tularensis pOMP1 there is only a one amino acid difference,
therefore they may play a similar role in the different subsp.
78
Interestingly, we have clearly demonstrated that F. tularensis LVS has the ability
to adhere to human lung cells while subsp. novicida lacks this ability. Similar
biotinylation experiments were performed with subsp. novicida and a similar pattern of
protein expression was observed. When performing the biotinylation experiments we
were interested in seeing if any proteins were being expressed in F. tularensis LVS and
not in subsp. novicida. Due to time restraints and lack of any polymorphisms, we were
unable to look further into the similarities between the F. tularensis LVS and subsp.
novicida. However, it is possible that this conserved banding patterns between
subspecies shows that these proteins are important because they are conserved. As seen
in other bacteria the higher the conservation of genes the more important the genes are to
the bacterium. In E. coli the enterohaemorrhagic strain 0157:H7 and the non-pathogenic
strain K12 are highly conserved. This demonstrates that the majority of the genes are
present in both strains meaning they are important for the survival of the bacteria.
Although the biotinylated proteins are similar between F. tularensis LVS and
subsp. novicida, F. tularensis LVS still demonstrates increased adherence to human lung
cells. One possible reason that F. tularensis LVS demonstrates increased adherence,
when compared with subsp. novicida, could be that the levels of expression of certain
proteins differ between subspecies. It is possible that F. tularensis LVS has other genes
which are regulating the expression levels of proteins important to adherence to human
lung. For example, different strains of M. catarrhalis have the ability to express different
levels of the Hag protein (Holm et al., 2003).
Genetic manipulation of F. tularensis LVS has proven to be a difficult task.
Lauriano et al. (2003) demonstrated mutagenesis by allelic exchange using PCR product,
79
which include flanking regions surrounding the gene of interest, in order to increase the
transformation efficiency. In our model, we used a kanamycin cassette to disrupt the
gene pOMP1. However, we were unable to transform F. tularensis LVS to make an
isogenic mutant. There are several possible explanations for this difficulty. As seen
throughout the literature, all tested F. tularensis subspecies are difficult to transform,
however, F. tularensis LVS has been found particularly difficult (Kawula et al., 2004;
Lauriano et al., 2003). The reasons for this difficulty are currently unknown. Not only
has it been proven difficult to mutagenize F. tularensis, tranposon insertions are also
typically unstable. This difficulty could be due to a restriction modification system
present in F. tularensis subsp. holartica. Restriction modification systems contain
restriction endonucleases, and are found throughout bacteria. They function in protecting
the cell against foreign DNA, by cleavage. Native DNA within the cell is protected by
methylation. Helicobacter pylori is just one of the many examples of a bacterium with
numerous restriction modification systems present (Pingoud et al., 2005). It contains ~20
restriction modification systems which is four percent of the total genome (Lin et al.,
2001). This demonstrates how important these systems are to a cell. No research has
been published regarding any restriction modification systems with in F. tularensis. The
complexity of this system may be one possible explanation of why it is so difficult to
mutangenize F. tualarensis.
Another possibility for this difficulty could be due to the inactivation of the
homologous recombination enzymes of F. tularensis. The RecA protein is an important
catalytic and regulatory component in homologous recombination. RecA works with a
number of other proteins, such as the single-stranded DNA binding protein (SSB) and
80
RecBCD. If RecA or one of the other proteins essential for homologous recombination is
being inactivated or is not functioning properly the DNA that is transformed into the cell
would not recombine with the wild-type DNA. A homolog of RecA has not currently
been identified in F. tularensis LVS, however, a gene locus functionally analogous to the
E. coli RecA has been identified (Berg et al., 1992). Berg et al. (1992) cloned the subsp.
novicida RecA homolog into a RecA minus strain of E. coli and made a RecA mutant in
subsp. novicida. These experiments demonstrate that subsp. novicida RecA functionally
helped in both recombination and DNA repair in the RecA deficient strain of E. coli. It
also showed that subsp. novicida mutant was deficient in homologous recombination.
These experiments were performed only in subsp. novicida which has been shown to be
easier to genetically manipulate than F. tularensis LVS. This leaves open the possibility
that there is a deficiency in the F. tularensis LVS RecA, which leads to decreased
homologous recombination.
The final explanation for poor transformation efficiency could be that pOMP1 is
an essential protein for the cell and without it functioning properly F. tularensis LVS
cannot grow. Since pOMP1 is an OMP, it could be an essential porin or play a key role
in the structure of the outer membrane. An example of an essential outer membrane
protein is OMP85 (Bos and Tommassen, 2004). This protein is evolutionarily conserved
among all Gram-negative genomes that have been sequenced to date. It has been found
that it is essential for viability of the cell (Bos and Tommassen, 2004). To date, there has
not been an OMP85 ortholog identified in F. tularensis subspecies.
In order to characterize the properties of pOMP1, it would be best to have
antibodies against the protein. Antibodies to pOMP1 will conclusively determine its
81
expression level by F. tularensis LVS, as well as E. coli transformed with the pOMP1
plasmid. It will also determine whether this protein contains surface expressed epitopes.
We are currently developing antibodies against pOMP1 in mice. Once the antibodies are
recovered, we will use flow cytometry, ELISA assays, and western blots to confirm
pOMP1 contains surface expressed epitopes, and to further characterize pOMP1 function.
Due to the recent threat of terrorists, it is becoming more desired to discover a
vaccine for F. tularensis. A better understanding of the virulence mechanisms used by F.
tularensis should allow identification of potential vaccine candidates. To date, little is
known regarding the pathogenesis of F. tularensis. In this study, we demonstrate that F.
tularensis LVS has the ability to adhere to human lung cells (A549) and subsp. novicida
lacks this ability to bind. We also identified 12 potential OMPs, including one novel
protein, pOMP1. Cloning of pOMP1 showed that it may play a role in mediating
adherence of F. tularensis LVS. This information along with the current production of
antibodies for pOMP1, will help reveal more information about this novel OMP.
82
CONCLUSIONS
1.
F. tularensis LVS demonstrates significantly greater adherence to the human
alveolar cell line A549 than subsp. novicida.
2.
Using surface biotinylation nine F. tularensis LVS outer membrane proteins were
identified, including two novel protein, pOMP1 and pOMP2.
3.
The known F. tularensis outer membrane protein, FopA, is most likely does not
function as an adhesin for F. tularensis LVS.
4.
A novel outer membrane protein, pOMP1, was identified.
83
REFERENCES
Abd, H., Johansson, T., Golovliov, I., Sandstrom, G., and Forsman, M. (2003). Survival
and growth of Francisella tularensis in Acanthamoeba castellanii. Appl Environ
Microbiol 69, 600-606.
Anthony, L. S., Gu, M. Z., Cowley, S. C., Leung, W. W., and Nano, F. E. (1991).
Transformation and allelic replacement in Francisella spp. J Gen Microbiol 137,
2697-2703.
Armstrong, J. A., and Hart, P. D. (1971). Response of cultured macrophages to
Mycobacterium tuberculosis, with observations on fusion of lysosomes with
phagosomes. J Exp Med 134, 713-740.
Arruda, S., Bomfim, G., Knights, R., Huima-Byron, T., and Riley, L. W. (1993). Cloning
of an M. tuberculosis DNA fragment associated with entry and survival inside
cells. Science 261, 1454-1457.
Azghani, A. O., Idell, S., Bains, M., and Hancock, R. E. (2002). Pseudomonas
aeruginosa outer membrane protein F is an adhesin in bacterial binding to lung
epithelial cells in culture. Microb Pathog 33, 109-114.
Barenkamp, S. J., Granoff, D. M., and Munson, R. S., Jr. (1981). Outer-membrane
protein subtypes of Haemophilus influenzae type b and spread of disease in daycare centers. J Infect Dis 144, 210-217.
Baron, G. S., Myltseva, S. V., and Nano, F. E. (1995). Electroporation of Francisella
tularensis. Methods Mol Biol 47, 149-154.
84
Bateman, A., and Bycroft, M. (2000). The structure of a LysM domain from E. coli
membrane-bound lytic murein transglycosylase D (MltD). J Mol Biol 299, 11131119.
Berg, J. M., Mdluli, K. E., and Nano, F. E. (1992). Molecular cloning of the recA gene
and construction of a recA strain of Francisella novicida. Infect Immun 60, 690693.
Bos, M. P., and Tommassen, J. (2004). Biogenesis of the Gram-negative bacterial outer
membrane. Curr Opin Microbiol 7, 610-616.
Burton, P. R., Kordova, N., and Paretsky, D. (1971). Electron microscopic studies of the
rickettsia Coxiella burneti: entry, lysosomal response, and fate of rickettsial DNA
in L-cells. Can J Microbiol 17, 143-150.
Chamberlain, R. E. (1965). Evaluation of Live Tularemia Vaccine Prepared in a
Chemically Defined Medium. Appl Microbiol 13, 232-235.
Clerc, P. L., Ryter, A., Mounier, J., and Sansonetti, P. J. (1987). Plasmid-mediated early
killing of eucaryotic cells by Shigella flexneri as studied by infection of J774
macrophages. Infect Immun 55, 521-527.
Cowley, S. C., Myltseva, S. V., and Nano, F. E. (1996). Phase variation in Francisella
tularensis affecting intracellular growth, lipopolysaccharide antigenicity and nitric
oxide production. Mol Microbiol 20, 867-874.
Dennis, D. T., Inglesby, T. V., Henderson, D. A., Bartlett, J. G., Ascher, M. S., Eitzen,
E., Fine, A. D., Friedlander, A. M., Hauer, J., Layton, M., et al. (2001). Tularemia
as a biological weapon: medical and public health management. Jama 285, 27632773.
85
Ecevit, I. Z., McCrea, K. W., Pettigrew, M. M., Sen, A., Marrs, C. F., and Gilsdorf, J. R.
(2004). Prevalence of the hifBC, hmw1A, hmw2A, hmwC, and hia Genes in
Haemophilus influenzae Isolates. J Clin Microbiol 42, 3065-3072.
Eigelsbach, H. T., Braun, W., and Herring, R. D. (1951). Studies on the variation of
Bacterium tularense. J Bacteriol 61, 557-569.
Eigelsbach, H. T., Hunter, D. H., Janssen, W. A., Dangerfield, H. G., and Rabinowitz, S.
G. (1975). Murine model for study of cell-mediated immunity: protection against
death from fully virulent Francisella tularensis infection. Infect Immun 12, 9991005.
Ellis, J., Oyston, P. C., Green, M., and Titball, R. W. (2002). Tularemia. Clin Microbiol
Rev 15, 631-646.
Ericsson, M., Golovliov, I., Sandstrom, G., Tarnvik, A., and Sjostedt, A. (1997).
Characterization of the nucleotide sequence of the groE operon encoding heat
shock proteins chaperone-60 and -10 of Francisella tularensis and determination
of the T-cell response to the proteins in individuals vaccinated with F. tularensis.
Infect Immun 65, 1824-1829.
Ericsson, M., Tarnvik, A., Kuoppa, K., Sandstrom, G., and Sjostedt, A. (1994). Increased
synthesis of DnaK, GroEL, and GroES homologs by Francisella tularensis LVS
in response to heat and hydrogen peroxide. Infect Immun 62, 178-183.
Evans, M. E., Gregory, D. W., Schaffner, W., and McGee, Z. A. (1985). Tularemia: a 30year experience with 88 cases. Medicine (Baltimore) 64, 251-269.
Feldman, K. A., Enscore, R. E., Lathrop, S. L., Matyas, B. T., McGuill, M., Schriefer, M.
E., Stiles-Enos, D., Dennis, D. T., Petersen, L. R., and Hayes, E. B. (2001). An
86
outbreak of primary pneumonic tularemia on Martha's Vineyard. N Engl J Med
345, 1601-1606.
Fortier, A. H., Green, S. J., Polsinelli, T., Jones, T. R., Crawford, R. M., Leiby, D. A.,
Elkins, K. L., Meltzer, M. S., and Nacy, C. A. (1994). Life and death of an
intracellular pathogen: Francisella tularensis and the macrophage. Immunol Ser
60, 349-361.
Fortier, A. H., Leiby, D. A., Narayanan, R. B., Asafoadjei, E., Crawford, R. M., Nacy, C.
A., and Meltzer, M. S. (1995). Growth of Francisella tularensis LVS in
macrophages: the acidic intracellular compartment provides essential iron
required for growth. Infect Immun 63, 1478-1483.
Fulop, M., Manchee, R., and Titball, R. (1996). Role of two outer membrane antigens in
the induction of protective immunity against Francisella tularensis strains of
different virulence. FEMS Immunol Med Microbiol 13, 245-247.
Fussenegger, M., Rudel, T., Barten, R., Ryll, R., and Meyer, T. F. (1997). Transformation
competence and type-4 pilus biogenesis in Neisseria gonorrhoeae--a review.
Gene 192, 125-134.
Garduno, R. A., Faulkner, G., Trevors, M. A., Vats, N., and Hoffman, P. S. (1998).
Immunolocalization of Hsp60 in Legionella pneumophila. J Bacteriol 180, 505513.
Gil, H., Benach, J. L., and Thanassi, D. G. (2004). Presence of pili on the surface of
Francisella tularensis. Infect Immun 72, 3042-3047.
87
Golovliov, I., Baranov, V., Krocova, Z., Kovarova, H., and Sjostedt, A. (2003a). An
attenuated strain of the facultative intracellular bacterium Francisella tularensis
can escape the phagosome of monocytic cells. Infect Immun 71, 5940-5950.
Golovliov, I., Sjostedt, A., Mokrievich, A., and Pavlov, V. (2003b). A method for allelic
replacement in Francisella tularensis. FEMS Microbiol Lett 222, 273-280.
Gray, C. G., Cowley, S. C., Cheung, K. K., and Nano, F. E. (2002). The identification of
five genetic loci of Francisella novicida associated with intracellular growth.
FEMS Microbiol Lett 215, 53-56.
Grifantini, R., Bartolini, E., Muzzi, A., Draghi, M., Frigimelica, E., Berger, J., Ratti, G.,
Petracca, R., Galli, G., Agnusdei, M., et al. (2002). Previously unrecognized
vaccine candidates against group B meningococcus identified by DNA
microarrays. Nat Biotechnol 20, 914-921.
Hendrixson, D. R., and St Geme, J. W., 3rd (1998). The Haemophilus influenzae Hap
serine protease promotes adherence and microcolony formation, potentiated by a
soluble host protein. Mol Cell 2, 841-850.
Hennequin, C., Porcheray, F., Waligora-Dupriet, A., Collignon, A., Barc, M., Bourlioux,
P., and Karjalainen, T. (2001). GroEL (Hsp60) of Clostridium difficile is involved
in cell adherence. Microbiology 147, 87-96.
Holm, M. M., Vanlerberg, S. L., Foley, I. M., Sledjeski, D. D., and Lafontaine, E. R.
(2004). The Moraxella catarrhalis porin-like outer membrane protein CD is an
adhesin for human lung cells. Infect Immun 72, 1906-1913.
88
Holm, M. M., Vanlerberg, S. L., Sledjeski, D. D., and Lafontaine, E. R. (2003). The Hag
protein of Moraxella catarrhalis strain O35E is associated with adherence to
human lung and middle ear cells. Infect Immun 71, 4977-4984.
Hopla, C. E. (1974). The ecology of tularemia. Adv Vet Sci Comp Med 18, 25-53.
Hubalek, M., Hernychova, L., Brychta, M., Lenco, J., Zechovska, J., and Stulik, J.
(2004). Comparative proteome analysis of cellular proteins extracted from highly
virulent Francisella tularensis ssp. tularensis and less virulent F. tularensis ssp.
holarctica and F. tularensis ssp. mediaasiatica. Proteomics 4, 3048-3060.
Huesca, M., Goodwin, A., Bhagwansingh, A., Hoffman, P., and Lingwood, C. A. (1998).
Characterization of an acidic-pH-inducible stress protein (hsp70), a putative
sulfatide binding adhesin, from Helicobacter pylori. Infect Immun 66, 4061-4067.
Kapatral, V., Anderson, I., Ivanova, N., Reznik, G., Los, T., Lykidis, A., Bhattacharyya,
A., Bartman, A., Gardner, W., Grechkin, G., et al. (2002). Genome sequence and
analysis of the oral bacterium Fusobacterium nucleatum strain ATCC 25586. J
Bacteriol 184, 2005-2018.
Kawula, T. H., Hall, J. D., Fuller, J. R., and Craven, R. R. (2004). Use of transposontransposase complexes to create stable insertion mutant strains of Francisella
tularensis LVS. Appl Environ Microbiol 70, 6901-6904.
Ketterer, M. R., Shao, J.Q. & Apicella, M.A. (2003). Ultrastructural Analysis of
Francisella tularensis. Paper presented at: ASM General Meeting (Washington,
DC).
Kovarova, H., Halada, P., Man, P., Golovliov, I., Krocova, Z., Spacek, J., Porkertova, S.,
and Necasova, R. (2002). Proteome study of Francisella tularensis live vaccine
89
strain-containing phagosome in Bcg/Nramp1 congenic macrophages: resistant
allele contributes to permissive environment and susceptibility to infection.
Proteomics 2, 85-93.
Kumar, A., Bose, M., and Brahmachari, V. (2003). Analysis of expression profile of
mammalian cell entry (mce) operons of Mycobacterium tuberculosis. Infect
Immun 71, 6083-6087.
Lai, X. H., Golovliov, I., and Sjostedt, A. (2001). Francisella tularensis induces
cytopathogenicity and apoptosis in murine macrophages via a mechanism that
requires intracellular bacterial multiplication. Infect Immun 69, 4691-4694.
Lai, X. H., Golovliov, I., and Sjostedt, A. (2004). Expression of IglC is necessary for
intracellular growth and induction of apoptosis in murine macrophages by
Francisella tularensis. Microb Pathog 37, 225-230.
Larsson, P., Oyston, P. C., Chain, P., Chu, M. C., Duffield, M., Fuxelius, H. H., Garcia,
E., Halltorp, G., Johansson, D., Isherwood, K. E., et al. (2005). The complete
genome sequence of Francisella tularensis, the causative agent of tularemia. Nat
Genet 37, 153-159.
Lauriano, C. M., Barker, J. R., Nano, F. E., Arulanandam, B. P., and Klose, K. E. (2003).
Allelic exchange in Francisella tularensis using PCR products. FEMS Microbiol
Lett 229, 195-202.
Lauriano, C. M., Barker, J. R., Yoon, S. S., Nano, F. E., Arulanandam, B. P., Hassett, D.
J., and Klose, K. E. (2004). MglA regulates transcription of virulence factors
necessary for Francisella tularensis intraamoebae and intramacrophage survival.
Proc Natl Acad Sci U S A 101, 4246-4249.
90
Layh-Schmitt, G., Podtelejnikov, A., and Mann, M. (2000). Proteins complexed to the P1
adhesin of Mycoplasma pneumoniae. Microbiology 146 ( Pt 3), 741-747.
Leslie, D. L., Cox, J., Lee, M., and Titball, R. W. (1993). Analysis of a cloned
Francisella tularensis outer membrane protein gene and expression in attenuated
Salmonella typhimurium. FEMS Microbiol Lett 111, 331-335.
Lin, L. F., Posfai, J., Roberts, R. J., and Kong, H. (2001). Comparative genomics of the
restriction-modification systems in Helicobacter pylori. Proc Natl Acad Sci U S A
98, 2740-2745.
Liu, D. F., Mason, K. W., Mastri, M., Pazirandeh, M., Cutter, D., Fink, D. L., St Geme, J.
W., 3rd, Zhu, D., and Green, B. A. (2004). The C-terminal fragment of the
internal 110-kilodalton passenger domain of the Hap protein of nontypeable
Haemophilus influenzae is a potential vaccine candidate. Infect Immun 72, 69616968.
Macellaro, A., Tujulin, E., Hjalmarsson, K., and Norlander, L. (1998). Identification of a
71-kilodalton surface-associated Hsp70 homologue in Coxiella burnetii. Infect
Immun 66, 5882-5888.
Maier, T. M., Havig, A., Casey, M., Nano, F. E., Frank, D. W., and Zahrt, T. C. (2004).
Construction and characterization of a highly efficient Francisella shuttle
plasmid. Appl Environ Microbiol 70, 7511-7519.
Merz, A. J., So, M., and Sheetz, M. P. (2000). Pilus retraction powers bacterial twitching
motility. Nature 407, 98-102.
Nano, F. E. (1988). Identification of a heat-modifiable protein of Francisella tularensis
and molecular cloning of the encoding gene. Microb Pathog 5, 109-119.
91
Nano, F. E., Zhang, N., Cowley, S. C., Klose, K. E., Cheung, K. K., Roberts, M. J., Ludu,
J. S., Letendre, G. W., Meierovics, A. I., Stephens, G., and Elkins, K. L. (2004).
A Francisella tularensis pathogenicity island required for intramacrophage
growth. J Bacteriol 186, 6430-6436.
Navarre, W. W., and Schneewind, O. (1999). Surface proteins of gram-positive bacteria
and mechanisms of their targeting to the cell wall envelope. Microbiol Mol Biol
Rev 63, 174-229.
Nemish, U., Yu, R. H., Tari, L. W., Krewulak, K., and Schryvers, A. B. (2003). The
bacterial receptor protein, transferrin-binding protein B, does not independently
facilitate the release of metal ion from human transferrin. Biochem Cell Biol 81,
275-283.
Neumeister, B., Schoniger, S., Faigle, M., Eichner, M., and Dietz, K. (1997).
Multiplication of different Legionella species in Mono Mac 6 cells and in
Acanthamoeba castellanii. Appl Environ Microbiol 63, 1219-1224.
Nikaido, H. (2003). Molecular basis of bacterial outer membrane permeability revisited.
Microbiol Mol Biol Rev 67, 593-656.
Oyston, P. C., Sjostedt, A., and Titball, R. W. (2004). Tularaemia: bioterrorism defence
renews interest in Francisella tularensis. Nat Rev Microbiol 2, 967-978.
Pandher, K., Confer, A. W., and Murphy, G. L. (1998). Genetic and immunologic
analyses of PlpE, a lipoprotein important in complement-mediated killing of
Pasteurella haemolytica serotype 1. Infect Immun 66, 5613-5619.
92
Pavlov, V. M., Mokrievich, A. N., and Volkovoy, K. (1996). Cryptic plasmid pFNL10
from Francisella novicida-like F6168: the base of plasmid vectors for Francisella
tularensis. FEMS Immunol Med Microbiol 13, 253-256.
Pingoud, A., Fuxreiter, M., Pingoud, V., and Wende, W. (2005). Type II restriction
endonucleases: structure and mechanism. Cell Mol Life Sci 62, 685-707.
Pomerantsev, A. P., Domaradskii, I. V., and Shishkova, N. A. (1991). [Hybrid plasmid
pSKEFT5 assures the transfer of plasmids Sa from Escherichia coli cells into
cells of the tularemia microbe vaccine strain cells]. Mol Gen Mikrobiol Virusol,
13-16.
Rhinehart-Jones, T. R., Fortier, A. H., and Elkins, K. L. (1994). Transfer of immunity
against lethal murine Francisella infection by specific antibody depends on host
gamma interferon and T cells. Infect Immun 62, 3129-3137.
Russell, P., Eley, S. M., Fulop, M. J., Bell, D. L., and Titball, R. W. (1998). The efficacy
of ciprofloxacin and doxycycline against experimental tularaemia. J Antimicrob
Chemother 41, 461-465.
Sandstrom, G. (1994). The tularaemia vaccine. J Chem Technol Biotechnol 59, 315-320.
Sandstrom, G., Sjostedt, A., Johansson, T., Kuoppa, K., and Williams, J. C. (1992).
Immunogenicity and toxicity of lipopolysaccharide from Francisella tularensis
LVS. FEMS Microbiol Immunol 5, 201-210.
Serizawa, H., and Fukuda, R. (1987). Structure of the gene for the stringent starvation
protein of Escherichia coli. Nucleic Acids Res 15, 1153-1163.
Sonnenberg, M. G., and Belisle, J. T. (1997). Definition of Mycobacterium tuberculosis
culture filtrate proteins by two-dimensional polyacrylamide gel electrophoresis,
93
N-terminal amino acid sequencing, and electrospray mass spectrometry. Infect
Immun 65, 4515-4524.
Sorokin, V. M., Pavlovich, N. V., and Prozorova, L. A. (1996). Francisella tularensis
resistance to bactericidal action of normal human serum. FEMS Immunol Med
Microbiol 13, 249-252.
Strom, M. S., and Lory, S. (1993). Structure-function and biogenesis of the type IV pili.
Annu Rev Microbiol 47, 565-596.
Sundin, C., Wolfgang, M. C., Lory, S., Forsberg, A., and Frithz-Lindsten, E. (2002).
Type IV pili are not specifically required for contact dependent translocation of
exoenzymes by Pseudomonas aeruginosa. Microb Pathog 33, 265-277.
Telepnev, M., Golovliov, I., Grundstrom, T., Tarnvik, A., and Sjostedt, A. (2003).
Francisella tularensis inhibits Toll-like receptor-mediated activation of
intracellular signalling and secretion of TNF-alpha and IL-1 from murine
macrophages. Cell Microbiol 5, 41-51.
Timpe, J. M., Holm, M. M., Vanlerberg, S. L., Basrur, V., and Lafontaine, E. R. (2003).
Identification of a Moraxella catarrhalis outer membrane protein exhibiting both
adhesin and lipolytic activities. Infect Immun 71, 4341-4350.
Titball, R. W., Johansson, A., and Forsman, M. (2003). Will the enigma of Francisella
tularensis virulence soon be solved? Trends Microbiol 11, 118-123.
Wizemann, T. M., Heinrichs, J. H., Adamou, J. E., Erwin, A. L., Kunsch, C., Choi, G. H.,
Barash, S. C., Rosen, C. A., Masure, H. R., Tuomanen, E., et al. (2001). Use of a
whole genome approach to identify vaccine molecules affording protection
against Streptococcus pneumoniae infection. Infect Immun 69, 1593-1598.
94
www.cdc.gov (2005). Centers for Disease Control and Prevention (CDC).
Zuber, M., Hoover, T. A., Dertzbaugh, M. T., and Court, D. L. (1995). Analysis of the
DnaK molecular chaperone system of Francisella tularensis. Gene 164, 149-152.
95
ABSTRACT
Francisella tularensis is a Gram-negative bacterium that causes tularemia. It is a
potential biological weapon because of its extreme infectivity (<10 organisms), ease of
dissemination, and substantial capacity to cause illness and death. Little is known about
the pathogenesis of F. tularensis infections. We hypothesize that adherence of F.
tularensis to lung epithelium is an important first step in pathogenesis and that surface
proteins (adhesins) are critical for this adherence. We have observed enhanced adherence
of F. tularensis LVS (live vaccine strain) to the human epithelial cell line, A549, when
compared to the adherence of subsp. novicida or laboratory strains of E. coli. To identify
surface proteins, bacteria were treated with a membrane impermeable biotinylation agent.
We have identified one known F. tularensis surface protein (FopA) and seven putative
outer membrane proteins. This work will give us a greater understanding of the initial
steps in F. tularensis pathogenesis.
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