impaired virulence factor production in a dihydroorotate .../67531/metadc...pyocyanin is a redox...
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APPROVED: Gerard A. O’Donovan, Major Professor Robert C. Benjamin, Committee Member Rollie R. Schafer, Committee Member Debrah A. Beck, Committee Member John E. Knesek, Committee Member Art Goven, Chair of the Department of Biological
Sciences Sandra L. Terrell, Dean of the Robert B. Toulouse
School of Graduate Studies
IMPAIRED VIRULENCE FACTOR PRODUCTION IN A DIHYDROOROTATE
DEHYDROGENASE MUTANT (pyrD) OF Pseudomonas aeruginosa
Pooja Ralli, B.S., M.S.
Dissertation Prepared for the Degree of
DOCTOR OF PHILOSOPHY
UNIVERSITY OF NORTH TEXAS
December 2005
Ralli, Pooja. Impaired virulence factor production in a dihydroorotate dehydrogenase
mutant (pyrD) of Pseudomonas aeruginosa. Doctor of Philosophy (Molecular Biology),
December 2005, 112 pp., 6 tables, 34 figures, references, 163 titles.
Previous research in our laboratory showed that when knockout mutations were created
in the pyrB and pyrC genes of the pyrimidine pathway in Pseudomonas aeruginosa, not only
were the resultant mutants auxotrophic for pyrimidines but they were also impaired in virulence
factor production. Such a correlation had not been previously reported for P. aeruginosa, a
ubiquitous opportunistic pathogen in humans. In an earlier study it was reported that mutants
blocked in one of the first three enzymes of the pyrimidine pathway in the non-pathogenic strain
P. putida M produced no pyoverdin pigment while mutants blocked in the later steps produced
copious amounts of pigment, just like the wild type. This study probed for the same connection
between pyrimidine auxotrophy and pigment production applied in P. aeruginosa. To that end a
knockout mutation was created in pyrD, the fourth step in the pyrimidine pathway which
encodes dihydroorotate dehydrogenase. The resulting mutant required pyrimidines for growth
but produced wild type pigment levels. Since the pigment pyoverdin is a siderophore it may also
be considered a virulence factor, other virulence factors were quantified in the mutant. These
included casein protease, hemolysin, elastase, swimming, swarming and twitching motility, and
iron binding capacity. In all cases these virulence factors were significantly decreased in the
mutant. Even supplementing with uracil did not attain wild type levels. Starvation of the
pyrimidine mutant for uracil caused increased specific activity of the pyrimidine enzymes,
suggesting that regulation of the pyrimidine pathway occurred at the level of transcription. This
effect has also been reported for P. oleovorans. The present research consolidates the idea that
pyrimidine auxotrophs cause decreased pathogenicity in P. aeruginosa. Such a finding may open
the search for chemotherapy targets in cystic fibrosis and burn victims where P. aeruginosa is an
infecting agent.
ii
Copyright 2005
by
Pooja Ralli
iii
ACKNOWLEDGEMENTS
I would like to sincerely thank Dr. O’Donovan for his support, generosity and
tremendous mentorship. I am indebted to my parents for believing in me. I would like to thank
my brother and his family who supported me in every possible way, and my fiancé for his
endless support and patience. I am thankful to all my friends here and back home and to Dr.
Avinash Srivastava for his invaluable suggestions and guidance. Thank you all for helping me
achieve this goal and making it possible.
TABLE OF CONTENTS
ACKNOWLEDGEMENTS ...........................................................................................................iii
LIST OF TABLES .......................................................................................................................... v
LIST OF FIGURES........................................................................................................................vi
INTRODUCTION........................................................................................................................... 1
MATERIALS AND METHODS.................................................................................................. 17
RESULTS...................................................................................................................................... 46
DISCUSSION ............................................................................................................................... 80
REFERENCES.............................................................................................................................. 88
v
LIST OF TABLES
Table 1. Bacterial strains and plasmids......................................................................................... 18
Table 2. Primers and PCR conditins. ............................................................................................ 35
Table 3. Complementation of E. coli auxotroph MA1004 with P. aeruginosa pyrD. .................. 52
Table 4. Generation time for PAO1 and PAOPR04 in Pseudomonas minimal medium.............. 61
Table 5. Generation time for PAO1 and PAOPR04 in PTSB medium......................................... 64
Table 6. Comparison of various virulence factors of the pyrimidine mutant with wild type
P. aeruginosa. ................................................................................................................... 86
vi
LIST OF FIGURES
Fig. 1. Overview of the infection process of Pseudomonas aeruginosa......................................... 4
Fig. 2. The quorum-sensing cascade of Pseudomonas aeruginosa................................................. 7
Fig. 3. The pyrimidine biosynthetic pathway in Pseudomonas aeruginosa. ................................ 10
Fig. 4. The pyrimidine salvage pathway in Pseudomonas aeruginosa. ........................................ 12
Fig. 5. Bradford protein standard curve of lysozyme.................................................................... 28
Fig. 6. Carbamoyl aspartate standard curve. ................................................................................. 30
Fig. 7. Construction of the plasmid pPR2. .................................................................................... 36
Fig. 8. Sequence of P. aeruginosa pyrD. ...................................................................................... 37
Fig. 9. Construction of the plasmid pPR3. .................................................................................... 39
Fig. 10. Scheme for the gene replacement method. ...................................................................... 41
Fig. 11. Construction of the plasmid pPR4. .................................................................................. 45
Fig. 12. PCR amplified pyrD gene of Pseudomonas aeruginosa. ................................................ 48
Fig. 13. Plasmid pPR2 digested with restriction enzymes EcoRI and BamHI.............................. 49
Fig. 14. SalI digest on the plasmid pPR2...................................................................................... 50
Fig. 15. Southern blot analysis of PAO1 and PAOPR04.............................................................. 51
Fig. 16. Specific activity of the pyrimidine enzymes from PAO1................................................ 55
Fig. 17. Specific activity of the pyrimidine enzymes from PAO1 and PAOPR04 cells grown with
uracil................................................................................................................................. 56
Fig. 18. Specific activity of the pyrimidine enzymes from PAO1 and PAOPR04 cells grown with
orotate............................................................................................................................... 57
vii
Fig. 19. Comparative analysis of specific activity of pyrimidine enzymes for PAO1 grown in
Pseudomonas minimal medium with 0.2% glucose. ........................................................ 58
Fig. 20. Comparison of specific activity of pyrimidine enzymes for PAOPR04 cells grown with
uracil.................................................................................................................................. 59
Fig. 21. Comparison of specific activity of pyrimidine enzymes for PAOPR04 cells grown with
orotate................................................................................................................................ 60
Fig. 22. Growth curves of PAO1 and PAOPR04 cells in Pseudomonas minimal medium ......... 62
Fig. 23. Growth curves of PAO1 and PAOPR04 cells in PTSB medium..................................... 65
Fig. 24. Pyocyanin production of PAO1 and PAOPR04 cells ..................................................... 66
Fig. 25. Measurement of pyoverdin for PAO1 and PAOPR04 cells ............................................ 67
Fig. 26. Growth of PAO1 and PAOPR04 on CAS medium ......................................................... 69
Fig. 27. Casein protease assay....................................................................................................... 70
Fig. 28. The Elastase Congo Red assay. ...................................................................................... 72
Fig. 29. Hemolysis of PAO1 and PAOPR04 strains on blood agar plates.................................... 73
Fig. 30. Swimming motility test.................................................................................................... 75
Fig. 31. Swarming motility test ..................................................................................................... 76
Fig. 32. Twitching motility test ..................................................................................................... 77
Fig. 33. Production of Rhamnolipid.............................................................................................. 79
Fig. 34. The chromosomal map of P. aeruginosa ......................................................................... 81
1
CHAPTER I
INTRODUCTION
Bacteria that belong to the family Pseudomonadaceae are called pseudomonads. They are
aerobic Gram-negative rods, with one or more polar flagella and include a wide variety of
species that thrive in diverse environments such as plants (Rahme et al., 1995, 1997), nematodes
(D’Argenio et al., 2002), insects (Mahajan-Miklos et al., 1999) and mammals (Lomholt et al.,
2001).
Extensive research on Pseudomonas aeruginosa has been carried out since it was first
mentioned by J. Schroeter, a German scientist in 1872 (who called it Bacterium aeruginosum)
(Brichta, 2003). Pseudomonas aeruginosa is a strict aerobe like all pseudomonads, but can
survive under anaerobic conditions utilizing nitrate in the absence of oxygen as the final electron
acceptor (Carlson, 1982). It is capable of utilizing a wide variety of sources of carbon for
survival (O’Sullivan and O’Gara, 1992). It is a ubiquitous organism usually harmless to a healthy
human being, but readily infects immunocompromised patients such as burn victims with open
wounds. It exudes “blue pus” from the wounds of such victims, which contains the bluish green
pigment, pyocyanin, and a florescent yellow pigment, initially called fluorescine, now called
pyoverdin.
Pyocyanin is a redox active phenazine compound which targets bacterial and mammalian
cells by generating reactive oxygen intermediates. Pseudomonas aeruginosa is not harmed by
this compound because of limited redox cycling of pyocyanin and also the increased production
of catalase and superoxide dismutase that eliminate the oxygen radicals generated (Hassett et al.,
1992). On the other hand pyoverdin is a siderophore that is involved in the transport of Fe+3 ions
into cells (Neilands, 1981) and possesses antibacterial activity (Kloepper et al., 1980; Misaghi et
2
al., 1982). Translocation of iron across the bacterial membrane is carried out with the help of
proteins TonB, ExbB, and ExbD that couple the electrochemical gradient across the cytoplasmic
membrane (Stinzi et al., 2000). Both pyocyanin and pyoverdin pigments are known to have a
crucial role in the virulence of P. aeruginosa. Pyocyanin interferes with the cytosolic
concentration of Ca+2 in human airway epithelial cells (Denning et al., 1998). Pseudomonas
aeruginosa also possesses another pigment pyochelin, a siderophore, which together with
pyocyanin and a reducing agent (NADH) causes tissue injury by catalyzing the formation of
reactive hydroxyl and superoxide radicals (Britigan, 1993). Iron uptake in a cell is very well
regulated and secretion of siderophores occurs under limited iron supply (Schwyn and Neilands,
1986). Pyoverdin deficient mutants show no virulence in vivo (Meyer et al., 1995), indicating the
importance of pyoverdin in virulence.
Pseudomonas aeruginosa is an opportunistic pathogen that causes a wide variety of
infections such as urinary tract infections, respiratory system infections, dermatitis, soft tissue
infections, and joint and bone infections especially in burn victims, cancer patients and cystic
fibrosis patients. The fatality rate in immunocompromised patients prone to P. aeruginosa
infections is about 50%. The reason for this bacterium to have such a wide range of attacking
mode is due to the presence of several virulence factors. For an infection to occur the body’s
defenses have to be breached and in immunocompromised patients this is a fairly easy task. The
organism attaches to the epithelial cells with the help of flagella and type IV pili for adhesion
and colonization (O’Toole and Kolter, 1998; Stickler, 1999). Flagellar movement is initiated by a
signal relay system of chemotaxis (Craven and Montie, 1985; Taguch et al., 1997) that helps the
bacterium to swim in an aqueous environment. Once the bacterium attaches to the cells,
twitching motility that requires type IV pili, helps in coalescing of cells into microcolonies
3
(O’Toole and Kolter, 1998). The bacterium also propagates via swarming motility, requiring
type IV pili on semi-solid surfaces under conditions of limited nitrogen (Köhler et al., 2000).
After colonization, P. aeruginosa releases several toxins that play a key role in virulence,
but the extent of tissue damage depends on the contribution of each one of these products and the
type of infection (Nicas and Iglewski, 1985) (Fig. 1). Proteases and hemolysins help in the
invasion of the host by the bacterium. Elastase (LasB), alkaline protease (AprA) and
staphylolysin (LasA) are the exoproduct proteases of P. aeruginosa (Passador and Iglewski,
1995). Elastin gives resilience to the red blood cells and helps in the expansion and contraction
of the lungs. Staphylolysin (LasA) nicks elastin when it is further degraded by LasB, AprA, and
neutrophil elastase (Galloway, 1991). LasB interferes with host defense mechanisms through a
wide range of effects. It degrades fibrin and collagen (Heck et al., 1986), inactivates human
immunoglobulin G and A (Heck et al., 1990), airway lysozyme (Jacquot et al., 1985), and
complement components (Hong & Ghebrehiwet, 1992). LasB is also α-1-proteinase inhibitor
(Morihara et al., 1979) that protects the respiratory tract against proteases and bronchial mucus
proteinase inhibitor (Johnson et al., 1982). Rhamnolipid and phospholipase C are two
hemolysins that break down lipids and lecithin. Rhamnolipid is a rhamnose containing glycolipid
biosurfactant with detergent properties that break down the phospholipid surfactant of the lungs
making it more susceptible to phospholipase C (Liu, 1974). Rhamnolipids are also capable of
inhibiting mucociliary transport and ciliary function in human respiratory epithelium (Read et
al., 1992).
Exotoxin A and exoenzyme S are two exotoxins released by P. aeruginosa. Exotoxin A is
responsible for local tissue damage and bacterial invasion (Woods and Iglewski, 1983) caused by
inhibiting protein synthesis and cell death (Wick et al., 1990) due to ADP-ribosylation
4
Fig. 1. Overview of the infection process of Pseudomonas aeruginosa
• Blood vessel invasion
• Multiple organ failure
• Death
Attachment and
colonization:
Chemotaxis
Flagellum
Type-IV pili
Adhesions
Chronic infection:
LPS
Alginate
Proteases
Hemolysin
Elastase
Exotoxins
Rhamnolipid
Pyocyanin
• Tissue damage
• [Ca+2] interference
in human airway
epithelium
Acute infection:
Biofilm
Quorum-sensing
Elevated levels of
virulence factors-
Proteases
Exotoxins
Pyocyanin
Hydrogen cyanide
Immuno-
compromised
patient
5
and inactivation of elongation factor-2. Exoenzyme S brings about ribosylation of GTP-binding
protein such as Ras (Iglewski et al., 1978) and may be responsible for direct tissue damage in
lung infections (Nicas et al., 1985a) and bacterial dissemination (Nicas et al., 1985b).
The role of the pigments pyocyanin and pyoverdin as virulence factors of P. aeruginosa was
explained previously.
Quorum sensing is the term that describes a bacterial intracellular signaling mechanism
that arises with increasing bacterial numbers that enable bacteria to coordinate the expression of
certain genes (Fig. 2). In Gram-negative bacteria, autoinducers and acylated homoserine lactone
are the chemicals that act as signaling molecules which are able to diffuse through bacterial
membranes or are actively transported (Pearson et al., 1999). After the bacteria reach a particular
density (usually late log to stationary phase) signaling molecules are released which enter the
neighboring cells and initiate a cascade of genes.
Pseudomonas aeruginosa is by far the best characterized bacterium for its quorum-
sensing system among Gram-negative bacteria (Rumbaugh et al., 2000). Initially two quorum-
sensing systems were identified, which were called las and rhl (Pesci and Iglewski, 1999). The
las system consists of a transcriptional activator LasR and an autoinducer synthase LasI that
directs the synthesis of the autoinducer N-(3-oxododecanoyl)-L-homoserine lactone, designated
as 3-oxo-C12-HSL (Gambello and Iglewski, 1991; Passador et al., 1993; Pearson et al, 1994).
The las system also has a repressor of virulence gene expression called RsaL (De Kievet et al.,
1999). When the concentration of the autoinducer (AI) is low, the repressor binds to the inducer
(I), repressing lasI transcription, thus preventing early induction of the quorum-sensing system.
As the bacterial cell density increases, the concentration of autoinducers increase and the LasR-
AI-I complex out-competes RsaL and binds to the lasI promoter leading to the synthesis of LasI.
6
The transcription of lasR is controlled by a global regulator called the virulence factor regulator,
Vfr (Albus et al., 1997). A global regulator, GacA, has been reported as a highly conserved
regulator in Gram-negative bacteria that positively regulates LasR and RhlR expression, and also
controls the production of secondary metabolites and exoenzymes in Pseudomonas spp.
(Reimann et al., 1997). The rhl system consists of a transcriptional activator RhlR and an
autoinducer N-butyryl-L-homoserine lactone (C4-HSL) synthesized by the autoinducer synthase
RhlI (Ochsner et al., 1994; Ochsner and Reiser, 1995; Pearson et al., 1995). RhlR binds to its
autoinducer and activates the transcription of its own autoinducer RhlI (Whitehead et al., 2001).
The two quorum-sensing systems independently control the transcription of LasB that encodes
the lasB elastase gene, which plays a major role in virulence (Brint and Ohman, 1995; Gambello
and Iglewski, 1991; Latifi et al., 1995; Pearson et al., 1997). Both the las and rhl systems also
control the transcription of genes that encode the production of elastase, proteases, exotoxins,
hemolysin and various other virulence factors. Between the las and rhl systems, the las quorum-
sensing system has hierarchy and controls the rhl quorum-sensing system at transcriptional and
posttranscriptional levels (Latifi et al., 1996; Pesci et al., 1997).
A third signaling molecule was identified as 2-heptyl-3-hydroxy-4-quinolone called the
Pseudomonas quinolone signal (PQS) by Pesci et al. (1995). Pseudomonas quinolone signal was
found only in the presence of an active LasR. RhlR was important for PQS bioactivity (Pesci et
al., 1995) and PQS was further found to induce rhlI. Pseudomonas quinolone signal also
activates the transcription of lasB gene and promotes the production of pyocyanin (Calfee et al,
2001; Gallagher et al, 2002). Pseudomonas quinolone signal was found to be present at a
maximum concentration only in the late stationary phase, which indicates that an intracellular
signal is not involved in sensing cell density (McKnight et al., 2000).
7
Fig. 2. The quorum-sensing cascade of Pseudomonas aeruginosa. Red arrows indicate negative
regulation while blue arrows indicate transcription. Adapted from McKnight et al., 2000.
lasR rsaL lasI
Vfr
PQS
LasR LasI
LasR
rhlR rhlI
RhlR RhlI
RhlR
LasB
lasB
3-oxo-C12-HSL
C4-HSL
Elastase (LasA & B)
Exotoxin A
Alkaline protease Rhamnolipid
Elastase (LasA & B)
Exotoxin A
Alkaline protease Rhamnolipid
Pyocyanin
Hydrogen cyanide
Pyocyanin
RsaL GacA
GacA
8
Biofilm is defined as the aggregation of a group of bacteria surrounded by a
polysaccharide capsule. This is a major factor in the treatment of patients with cystic fibrosis
(CF). Cystic fibrosis is a genetic disorder that results from mutations in the cystic fibrosis
transmembrane conductance regulator (CFTR) gene (Doull, 2001; Boucher, 2004), producing
defective chloride ion transport across epithelial cell surfaces and a build-up of dehydrated
mucus in the lung (Boucher, 2004). Biofilms composed of P. aeruginosa arise after the
bacterium is inhaled and reaches the airway of the epithelium. The bacterium adheres to the
epithelium and starts growing on the surface. Other bacteria adhere to the surface as they pass
through the epithelial airway. This adhesion later becomes irreversible due to the
polysaccharides present in the pili and glycocalyx. Certain genes of the bacteria get activated
under changed environmental conditions such as phosphate starvation, nitrogen starvation,
increased NaCl concentration and dehydration. This causes the production of
exopolysaccharides, mainly alginate (Linker and Jones, 1966; Costerton et al., 1994). As the
infection progresses the bacteria undergo a change in their phenotype and become mucoid,
adapting the biofilm mode of growth (Lam et al., 1980; Deretic et al., 1995; Singh et al., 2000).
The biopolymeric matrix thus formed within the airway protects the bacteria from host defense
mechanisms and antimicrobial agents (Mah and O’Toole, 2001; Dunne, 2002; Parsek and Singh,
2003). Alginate production and biofilm together encompass the host defenses and result in a poor
prognosis for CF patients (Govan and Deretic, 1996; Pier, 1998).
The virulence of an organism would not be possible without the metabolism that makes
the final product RNA and DNA. Pyrimidine metabolism is universal in all organisms so far
explored (O’Donovan and Neuhard, 1970; Grogan and Gonsalus, 1993) and UMP is the
precursor for all the pyrimidine nucleotides. The pathway has been found to be the same in
9
bacteria (Neuhard and Kelln, 1996), fungi (Denis-Duphil, 1989), plants (Doremus, 1986) and
mammals (Jones, 1980, a & b), with similar enzymatic reactions but varied structure of the
enzymes.
The pyrimidine biosynthetic pathway has been most extensively studied in Escherichia
coli and Salmonella typhimurium and is the same as found in Pseudomonas aeruginosa (Fig. 3).
The pathway comprises nine enzymatic steps culminating in the production of UTP and CTP.
The pathway commences with the formation of carbamoyl phosphate from glutamine (or
ammonia), bicarbonate and ATP, catalyzed by the enzyme carbamoyl phosphate synthetase
(CPSase), encoded by carAB operon (Anderson and Meister, 1965). Aspartate transcarbamoylase
(ATCase) encoded by the genes pyrBC’ in P. aeruginosa catalyzes the condensation of
carbamoyl phosphate and aspartate to form carbamoyl aspartate and inorganic phosphate (Pi)
(Jones et al., 1955; Vickrey, 1993).
The next step involves cyclization of the pyrimidine ring catalyzed by dihydroorotase
(DHOase), encoded by pyrC yielding dihydroorotate (Washabaugh and Collins, 1986).
Dihydroorotate dehydrogenase (DHOdehase), encoded by the pyrD gene producing the only
membrane-bound enzyme in this pathway brings about oxidation of dihydroorotate to orotate
(Larsen and Jensen, 1985; Karibian and Couchoud, 1974). Transfer of ribose-5’-phosphate from
γ-D-5’-phosphoribosyl-1’-pyrophosphate (PRPP) to orotate results in the formation of the first
pyrimidine nucleotide. Orotidine-5’-monphosphate (OMP) (Liebermann and Kornberg, 1954).
This reaction is catalyzed by orotate phosphoribosyl transferase (OPRTase) encoded by the pyrE
gene. Uridine-5’-monophosphate (UMP) is formed by decarboxylation of OMP catalyzed by
OMP decarboxylase (OMPdecase), encoded by the pyrF gene (Scapin et al., 1993). UMP kinase
encoded by the pyrH gene catalyzes the phosphorylation of UMP to uridine diphosphate (UDP)
10
Fig. 3. The pyrimidine biosynthetic pathway in Pseudomonas aeruginosa. The genes that encode
the enzymes are italicized.
Nucleoside
diphosphate
kinase
ndk
DHOdehase
pyrD
NH4+
CPSase
pyrA ATCase
pyrB
OMPdecase
pyrF
CTP
synthetase
pyrG
OPRTase
pyrE
Orotate
2 ADP + Pi
+
Glutamate
2 ATP
+
Glutamine
H2O
NADH + H+
NAD
PRPP
PPi
CO2
Bicarbonate Carbamoyl
phosphate
Carbamoyl
aspartate
Aspartate Pi
Dihydroorotate
DHOase
pyrC C2
Orotidine
monophosphate
Uridine
monophosphate
Uridine
diphosphate
Uridine
triphosphate
Cytidine
triphosphate UMP kinase
pyrH
DNA
Synthesis
RNA
Synthesis
ATP ATP ADP ATP ADP + Pi ADP
11
(O’Donovan and Gerhart, 1972; Serino et al., 1997). Nucleoside diphosphate kinase, encoded by
the gene ndk catalyzes the phosphorylation of UDP to uridine triphosphate (UTP) (Ginther and
Ingraham, 1974). Cytidine triphosphate (CTP) is produced by CTP synthetase, encoded by pyrG.
Adenosine triphosphate (ATP), and glutamine are involved in this reaction (Long and Koshland,
1978; Anderson, 1983).
The pyrimidine pathway is missing in some obligate parasites (Jones, 1980) but the
salvage pathway is present in all organisms and may differ from species to species. The salvage
pathway has been studied in about 40 different wild type and mutant strains of bacteria (Beck,
1995). The purpose of salvage is to replenish the pyrimidine bases, nucleosides and nucleotides,
primarily mononucleotides from mRNA degradation. In wild type prototroph strains
of bacteria salvage maintains a balance between the pyrimidine biosynthetic pathway and RNA
synthesis, whereas in pyrimidine auxotrophs the salvage pathway fulfills the pyrimidine
requirement in its entirety (O’Donovan and Shanley, 1995). Pyrimidine mutants in P. aeruginosa
are able to use exogenously supplied uracil, uridine, cytosine or cytidine for growth (Fig. 4).
Uracil gets into the cell via uracil permease, encoded by the uraA gene. Uracil and PRPP
together produce UMP, releasing a pyrophosphate (PPi), the reaction is catalyzed by uracil
phosphoribosyl transferase (UPRTase). UMP then follows the de novo pathway, finally resulting
in the formation of CTP from UTP. Exogenous cytosine gets into the cells via cytosine
permease, encoded by the gene codB, and gets oxidatively deaminated to uracil by cytosine
deaminase, encoded by codA. The entry of exogenous cytidine and uridine is facilitated by
nucleoside permease, encoded by the nup gene. Cytidine and uridine are irreversibly hydrolyzed
to cytosine and uracil respectively by the enzyme nucleoside hydrolase, encoded by nuh. This
enzyme also handles the purine nucleosides, adenosine, guanosine and inosine. Cytosine is then
12
Fig. 4. The pyrimidine salvage pathway in Pseudomonas aeruginosa. The genes that encode the
enzymes are italicized.
CTP
CDP
CMP
CR
C
Cytosine deaminase
codA
Nucleoside hydrolase
nuh
5’ Nucleotidase
CMP Kinase
cmk
CMP glycosylase
H2O
Ribose-5-
phosphate
UTP
UDP
UMP
UR
U
Nucleoside hydrolase
nuh
5’ Nucleotidase
PRPP
PPi
UPRTase
upp
Biosynthetic
pathway
H2O
Ribose
mRNA
Uracil
Orotate
Cytosine
codB uraA dctA
INSIDE
OUTSIDE
Cytidine Uridine
nup nup
13
deaminated to uracil, and uracil is converted to UMP by UPRTase. mRNA degradation results
in mononucleotides CMP or UMP which are degraded by 5’nucleotidase to cytidine or uridine
respectively. CMP can also be directly broken down to cytosine, releasing one ribose-5-
phosphate catalyzed by CMP glycosylase (West et al., 1985). Alternatively, CMP may be
phosphorylated to CDP via CMP kinase, encoded by cmk. All pseudomonads lack cytidine
deaminase and uridine/cytidine kinase encoded by the genes cdd and udk, respectively.
Regulation of pyrimidine biosynthesis in prokaryotes is carried out at the transcriptional
or translational level. In E. coli the pyrimidine biosynthetic pathway is regulated at the enzyme
synthesis level by attenuation (Roof et al., 1983; Turnbough et al., 1983) and at the enzyme
activity level by allosteric inhibition (Gerhart and Pardee, 1962; 1964). At the enzyme synthesis
level the genes pyrBI and pyrE that encode ATCase and OPRTase respectively have upstream
leader sequences that allow attenuation control (Navre and Schachmann, 1983).
At the level of enzyme activity there are three loci for regulation of the pyrimidine
biosynthetic pathway in E. coli. CPSase encoded by carAB is inhibited by UMP and activated by
ornithine and inosine-5’-monophosphate (IMP). Aspartate transcarbamoylase (ATCase),
encoded by pyrBI, catalyzes the second step, but is the first committed step of the pathway. It is
inhibited by CTP and activated by ATP (Foltermann et al., 1981). CTP synthetase, encoded by
pyrG, catalyzes the conversion of UTP to CTP, and is inhibited by CTP and activated by UTP
(Long and Pardee, 1967; Long and Koshland, 1978). In Pseudomonas, the regulation is brought
about by purine and pyrimidine nucleotide effectors (Chu and West, 1990). CPSase enzyme
activity is inhibited by UMP and activated by ornithine and N-acetylornithine (Abdelal et al.,
1983). ATCase, encoded by pyrBC’ is inhibited by ATP, CTP, and UTP with ATP being the
most effective inhibitor (Isaac and Holloway, 1968; Condon et al., 1976; Schurr et al., 1995;
14
Vickrey, 1993). In P. aeruginosa, a regulatory protein PyrR was identified that regulates the
expression of downstream pyrimidine biosynthetic genes pyrD, pyrE, and pyrF and the
expression of the pyrR gene (Patel, 2000).
The focus of research in our laboratory is to explore the pyrimidine pathway, as it could
be a target for drug therapy. To this end, mutants are isolated in the biosynthetic pathway
creating pyrimidine auxotrophs. These mutants are then tested in order to understand the
regulation of the pathway and virulence studies are also carried out. Blocks were created at the
second and third steps of the pyrimidine biosynthetic pathway and specific activity and
virulence-based experiments were performed. Virulence studies carried out on the mutants
blocked in the second (ATCase) and third (DHOase) step of the de novo pathway showed
significant virulent factor decreases when compared with the wild type strain (Hammerstein,
2004; Brichta, 2003). The aim of this research was the construction of a mutant blocked at the
fourth step of the pyrimidine biosynthetic pathway in Pseudomonas aeruginosa and to study its
effects on virulence factors and the effects on the other enzymes of the pathway.
Dihydroorotate dehydrogenase enzymes catalyze the only redox-reaction in the de novo
pyrimidine pathway. Dihydroorotate dehydrogenases are studied because it has been shown that
in certain parasites which lack the salvage pathway, the inhibition of dihydroorotate
dehydrogenase shuts down the biosynthetic pathway. In this way, the dehydrogenase becomes a
selective target for antiproliferative, antiparasitic and immunosuppressive drugs (DeFrees et al.,
1988; Davis and Copeland, 1997; Rückemann et al., 1998; Davis et al., 1996; Williamson et al,
1995; Löffler et al., 1998).
Dihydroorotate dehydrogenases are divided into two main classes (Bjornberg et al.,
1997) – class I and II. Class I enzymes are cytosolic and are further subdivided into class A and
15
class B. Class II enzymes are membrane-associated. Lactococcus lactis contains two
DHOdehases designated DHODA and DHODB (Andersen et al., 1994). Subclass A with
DHODA is a dimer formed by two identical PyrD subunits (Nielsen et al., 1996) and has
significant amino acid identity with the yeast enzyme (Andersen et al., 1994). Fumarate is the
natural electron acceptor for DHODA. Subclass B including DHODB has amino acid identity
with the Bacillus subtilis DHOdehase (Nielsen et al., 1996). It is a heterotetramer consisting of
two PyrDB and two PyrK subunits (Rowland et al., 2000; Andersen et al., 1996; Nielsen et al.,
1996) The PyrDB subunit couples with a prosthetic FMN group and is homologous to the PyrDA
subunits of DHODA (Nielsen et al., 1996; Björnberg et al., 1999). The two PyrK subunits have
[2Fe-2S] cluster and FAD as the cofactor and are capable of using NAD+ as an electron acceptor
(Nielsen et al., 1996). Class II enzymes are monomeric and crystalline structures of DHOdehase
of Escherichia coli (DHODC) and humans (Liu et al., 2000; Noranger et al., 2002) have been
studied. The E. coli DHOD is a homodimer with a molecular mass of 37KDa and contains one
FMN unit (Karibian et al., 1974; Larsen et al., 1985). In the presence of oxygen, ubiquinone is
the electron acceptor (Kerr and Miler, 1968) and under anaerobic conditions, the electron
acceptor is menaquinone (Andrews et al., 1977). P. aeruginosa DHOdehase (PyrD) is composed
of 342 amino acids, a total of 1026 nucleotides (PA # 3050), excluding the stop codon. It belongs
to class II and hence membrane-bound. This is the only membrane-bound enzyme in the
pyrimidine pathway. Structurally it is 70% similar to the PyrD of E. coli.
It has been reported that a C4-dicarboxylate transport protein, DctA, encoded by the dctA
gene is responsible for the transport of succinate, malate, and fumarate as carbon and energy
sources; it also mediates the entry of orotate into the cell (Baker et al., 1996; Yurgel et al., 2000).
Baker et al., 1996 also reported that orotate could satisfy the pyrimidine requirement in E. coli
16
and Salmonella typhimurium only in the presence of glycerol as the carbon and energy source,
but not in the presence of glucose. This aspect was tested in the P. aeruginosa pyrD mutant
where the formation of orotate in the pyrimidine biosynthetic pathway is blocked. Here orotate
satisfies the pyrimidine requirement when cells are grown in glucose but not in succinate.
17
CHAPTER II
MATERIALS AND METHODS
Bacterial strains and plasmids:
The bacterial strains and plasmids used in this study are listed in the Table 1.
Pseudomonas minimal medium (PsMM) adapted from Ornston and Stanier, 1966 was used to
grow Pseudomonas strains with 0.2% glucose as the carbon and energy source and an exogenous
supplement of uracil or orotate at 40 µg/ml was provided whenever required.
Media preparation:
Pseudomonas minimal medium (PsMM) was used for the cultivation of Pseudomonas
strains and contains the following compounds (Ornston and Stanier, 1966). 12.5 mM Na2HPO4,
12.5 mM KH2PO4, 0.1% (NH4)2SO4, and 10ml concentrated base. The composition of
concentrated base includes 14.45 g MgSO4 (anhydrous), 3.335 g CaCl2.2H2O, 0.00925 g
(NH4)6Mo7O24.4H2O, 0.099 g FeSO4.7H2O, and 50ml Metals 44. First 10 g nitrilotriacetic acid
was dissolved in 600 ml ddH2O and neutralized with 7.3 g KOH. Then the compounds
mentioned above were added, pH adjusted to 6.8 and brought up to a final volume of 1000 ml
with ddH2O. Metals 44 contains 2.5 g EDTA, 10.95 g ZnSO4.7H2O, 5 g FeSO4.7H2O, 1.54 g
MnSO4.7H2O, 0.392 g CuSO4.5H2O, 0.248 g Co(NO3)2.6H2O, 0.177 g Na2B4O7.10H2O and
ddH2O to a final volume of 1000 ml. A few drops of concentrated H2SO4 was added to avoid salt
precipitation.
Escherichia coli minimal medium (EcMM) was used to grow E. coli strain MA1004 (E.
coli genetic stock center). Escherichia coli minimal medium (Gause, 1934; Jost et al., 1972) has
the following ingredients per 1000 ml of ddH2O- 10.5 g K2HPO4 (dibasic), 4.5 g KH2PO4
18
Table 1. Bacterial strains and plasmids used in this study.
Strain or plasmid Genotype Source or Reference
P. aeruginosa
PAO1 Prototroph ATCC
PAOPR04 PAO1 pyrD∆ This study
E. coli
MA1004 lacZ43(Fs) λ- pyrD68 relA1 thi-1 Beckwith et al, 1962
DH5α recA1 hsdR17 endA1 supE44 thi-1 relA1 gyrA96 ∆(argF- lacZYA) GIBCO-BRL, 1986
U169 Ф80lacZ∆M15
SM10 KmR thi thr leu tonA lacY supE recA::Rp-2 Tc::Mu Simon et al, 1983
Top10 F- mcrA ∆(mrr-hsdRMS-mcrBC) Ф80lacZ∆M15 ∆lacX74 recA1 Invitrogen
araD139 ∆(ara-leu) galU galK rspL (StrR) endA1 nupG
Plasmids
pGEX2T GST fusion vector tac promoter lacIq AmpR Amersham Biosciences
pEX18Gm GmR oriT sacB+ gene replacement vector with multiple cloning site Hoang et al, 1998
from pUC18
pVDZ’2 IncPI mob+ tra tac lacZ’ lacIq ApR Deretic et al, 1987
pRK2013 ColEI mob+ tra+ (RK2) KmR Figurski and Helinski, 1979
pPR2 pGEX2T with 1.57Kb insert of P. aeruginosa pyrD This study
pPR2∆D pGEX2T containing the 1.47Kb deleted pyrD gene This study
pPR3 pEX18Gm containing the 1.47Kb deleted pyrD gene This study
pPR4 pVDZ’2 containing 1.57Kb insert of P. aeruginosa pyrD This study
19
(monobasic), 1.0 g (NH4)2SO4, and 0.5 g trisodium citrate. Once these compounds were
autoclaved and cooled, the following compounds were sterilized separately and added to the
medium- 1 ml 1 M MgSO4, 5 ml 0.337% B1 thiamine, and 0.2% glucose as carbon and energy
source.
Luria-Bertani (LB) medium supplied by Difco® contained 10 g Bacto® tryptone, 5 g
Bacto® yeast extract, and 10 g NaCl. LB medium with isopropylthio-β-D-thiogalactoside
(IPTG), X-gal (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside) and the appropriate choice
of antibiotic(s) was used for cloning experiments.
Pseudomonas Isolation Agar (PIA) supplied by Difco® is based on King’s A medium
(King et al., 1954) with slight modification in containing 0.025 g of the antibiotic Igrasan®
(Ciba Geigy) and was used as a selection medium for mating experiments of Pseudomonas
strains.
King’s A medium (King et al., 1954) was used to enhance pyocyanin production in liquid
culture and contains 2% Bacto peptone (Difco), 0.14% MgCl2, 1% K2SO4, and 1% glycerol.
Quantitation of pyocyanin assay was done according to Essar et al, 1990. King’s B medium
(King et al., 1954) was used for quantitative measurements of the pigment pyoverdin in liquid
culture. The medium contains 2% proteose peptone no. 3, 0.15% K2HPO4 (anhydrous), 0.15%
MgSO4.7H2O, and 1% glycerol.
Peptone tryptic soy broth (PTSB) medium containing 5% peptone (Bacto, Difco certified)
and 0.25% Tryptic soy (without dextrose, Difco) was used according to Pearson et al. (1997) to
quantitate the exoproducts casein protease and elastase.
Blood agar medium supplied by BBL® that contains tryptic soy agar base with 5%
sheep’s blood was used to test for the pathogenecity of the wild type and mutant strains of P.
20
aeruginosa. Exogenous supplement of uracil or orotate at a concentration of 40 µg/ml was also
added to the blood agar plates. The plates grown overnight were used and a single colony was
transferred with a toothpick and incubated for 24 hours. The zone of hemolysis implies the
degree of pathogenicity of the strains.
Chrome azurol S (CAS) medium (Schwyn and Neilands, 1987) also called Blue Agar was
used to detect the excretion of siderophores based on their affinity for iron. The excretion of
siderophore (ligand) causes the release of iron from the iron-dye complex to form the ligand-iron
complex, thereby causing the release of the free dye seen in the form of orange halos. The CAS
agar plates contained the following: MM9 salts modified from M9 (Sambrook et al., 1989) by
reducing the amount of phosphate to 0.03% KH2PO4 and replacing 0.1M phosphate buffer with
Pipes (1,4-piperazinediethanesulfonic acid). The MM9 10X stock solution contains per liter
Na2HPO4 (60 g), KH2PO4 (3 g), NaCl (5 g) and NH4Cl (10 g) in ddH2O. CAS-HDTMA
(hexadecyltrimethylammonium bromide) iron-dye solution was made by dissolving CAS (605
mg) to 500 ml ddH2O and then adding 100 ml iron (III) solution (1 mM FeCl3.6H2O in 10 mM
HCl). HDTMA solution was prepared by stirring to dissolve 729 mg in 400 ml ddH2O. The
concentration of HDTMA was crucial because too low could cause precipitation and too high
would be toxic for the bacterium.
To make a CAS-HDTMA solution, 600 ml of CAS-iron was added to HDTMA slowly by
stirring. Deferrated casamino acids was made by dissolving 10 g of casamino acids in 100 ml of
ddH2O and extracting with equal volume of 3% (w/w) 8-hydroxyquinoline in chloroform to
remove contaminating iron. Traces of 8-hydroxyquinoline were removed by extracting with
equal volume of chloroform. All these solutions were sterilized and stored at room temperature.
To make 1 liter of CAS agar medium, the following were added to 750 ml of ddH2O and
21
autoclaved- Pipes (30.24 g), NaOH (6 g) to raise the pH of the solution to the pKa of Pipes to 6.3,
10x MM9 (100 ml) and agar (15 g). The pH of the solution is a crucial factor and had to be
checked prior to the addition of agar as higher pH values could cause the change of color from
blue to green. After autoclaving and cooling the medium to about 50oC 30 ml deferrated
casamino acids (10%) and 0.2% glucose (20% stock) was added. Exogenous supplement of
uracil or orotate was provided at a concentration of 40 µg/ml. The CAS-HDTMA iron-dye
complex, a 100 ml volume was added slowly along the walls of the container and mixed by
stirring gently to avoid foaming. The plates that were obtained were blue in color and ready to be
used to test the affinity of siderophore for iron.
Rhamnolipid medium (Köhler et al., 2000) was used to detect the release of
rhamnolipids, a biosurfactant required for swarming motility in P. aeruginosa strains. The
preparation of the medium was modified from the protocol described by Siegmund and Wagner
(1991). The medium contained M8 salts (modified from M9, Sambrook et al., 1989) which
contains Na2HPO4 (6 g), KH2PO4 (3 g), and NaCl (0.5 g), 0.2% glucose (instead of 2%
glycerol), 2 mM MgSO4, 10 ml of 10X trace elements, 0.0005% methylene blue, 0.02%
cetyltrimethylammonium bromide (CTAB), 0.05% glutamic acid as nitrogen source and agar at a
final concentration of 1.6% (wt / vol). 10X trace elements contains 0.232g H3Bo3, 0.174 g
ZnSO4.7H2O, 0.23 g FeNH4(SO4)2.12H2O, 0.096 g CoSO4.7H2O, 0.022 g (NH4)6Mo7O24.4H2O,
8.0 mg CuSO4.5H2O, and 8.0 mg MnSO4.4H2O. The salts, agar, trace elements, methylene blue
and CTAB were autoclaved together. When the temperature reached 50oC, 0.2% glucose, 2 mM
MgSO4, and 0.05% glutamic acid were added slowly and mixed by stirring gently. Exogenous
supplement of uracil or orotate was provided at a concentration of 40 µg/ml. The plates so
22
obtained were pastel blue in color and ready to test the production of rhamnolipid seen in the
form of pink halos.
Swarm agar (Rashid and Kornberg, 2000) used to detect swarming motility contained
0.5% (wt/vol) Difco bacto agar, 0.8% Difco® nutrient broth, and 0.5% glucose. Swim agar
plates (Rashid and Kornberg, 2000) used to detect swimming motility contained tryptone broth
(1% tryptone, 0.5% NaCl), and 0.3% (w/v) agarose. Twitching motility was tested with the
medium (Rashid and Kornberg, 2000) that contained LB broth and 1% (wt / vol) Difco®
granulated agar. The motility plates were supplemented with uracil or orotate at 40 µg/ml
concentration.
Quantitation of casein protease and elastase was performed in Peptone tryptic soy broth
(PTSB) (Diener et al., 1972), which contained 5% Difco® peptone and 0.25% tryptic soy.
The antibiotics used for the selective growth of E. coli were Ampicillin (Ap – 100
µg/ml), Gentamycin (Gm – 15 µg/ml), Kanamycin (Km – 25 µg/ml), Tetracycline (Tc -
15µg/ml) and for P. aeruginosa Gentamycin (Gm – 100 µg/ml). Difco® agar was added at a
final concentration of 2% for the solid medium whenever required. All the strains were grown at
37oC by shaking at 250 rpm for liquid cultures.
Pyocyanin Quantitation assay:
The method of Essar et aI, 1990 was followed with slight modifications. The principle of
this assay is absorbance of pyocyanin at 520 nm in an acidic solution (Kurachi, 1958;
MacDonald, 1967). The strains were grown in King’s A solid medium and then cultured into its
liquid broth. An overnight grown liquid culture was inoculated into a 50 ml King’s A liquid
medium in a 250 ml Erlenmeyer flask and incubated at 37oC with shaking for 24 hours until the
23
wild type and mutant strains had the same optical density at 600 nm. A 5 ml aliquot was
transferred to a 15 ml conical flask and centrifuged at 1300xg for 25 minutes at 4oC in a Sorvall
RT 600B tabletop centrifuge. An aliquot of 1 ml supernatant was used to measure the optical
density at 600 nm to eliminate the background. The supernatant was filtered through a 0.45 µm
Ambion filter disc and pyocyanin was extracted from the filtered supernatant with 3 ml of
chloroform after vortexing to mix and centrifuging at 4oC in the tabletop centrifuge for 10
minutes at 1300xg. The pyocyanin (bottom layer) was transferred to a new 15 ml conical tube
and 1 ml of 0.2N HCl added. The mixture was centrifuged as described above and the pink color
obtained from the extraction was transferred to a cuvette and the absorbance was measured at
A520. Microgram quantities of pyocyanin were calculated by multiplying the absorbance at
520nm by 17.072 (Kurachi, 1958).
Pyoverdin Quantitation assay:
Strains were grown in King’s B solid medium and then cultured into its liquid broth. An
overnight liquid culture was inoculated into a 50 ml King’s B liquid medium in a 250 ml
Erlenmeyer flask and incubated at 37oC with shaking for 24 hours until the wild type and mutant
strains had the same O.D. reading at 600 nm. A 1ml aliquot was removed and the O.D. at 600 nm
was measured. The sample was then transferred to a microcentrifuge tube and centrifuged at
10000xg for 3-5 minutes at 4oC. The supernatant was measured at 405 nm. Pyoverdin levels
were expressed as a ratio of A405/A600 (Stinzi et al., 2000).
Elastolysis assay:
Elastolytic activity in P. aeruginosa was determined using the Elastin Congo Red (ECR)
24
assay (Pearson et al., 1997) with slight modifications. Cells were cultivated in PTSB medium for
24 hours at 37oC with shaking, until the wild type and mutant strains reached the same density at
O.D600. The cells were centrifuged as described above (Pyocyanin assay) and the supernatant
obtained was filtered through a 0.45 µm pore filter disc and the samples were stored at -70oC
until used. Filtered supernatant of 50 µl was added to a 1.0 ml buffer containing 0.1 M Tris (pH
7.2) and 1 mM CaCl2 (Brichta, 2003). The tubes were incubated at 37oC with shaking for 18
hours; 0.1 ml of 0.12 M EDTA added and then placed on ice for 10 minutes. Insoluble ECR was
removed by centrifugation at 10000xg for 1 minute at 4oC and the A495 was measured.
Absorption of the pigments produced by P. aeruginosa was corrected by subtracting the A495 of
the filtered supernatant that was incubated without ECR plus the A495 of the buffer and ECR
incubated with no filtered supernatant.
Casein protease assay:
Cells were cultivated and supernatants were obtained as described above. The method of
Kessler et al., 1993 was used with slight modification. Filtered supernatant of 50 µl was added to
1 ml of 0.3% azocasein in 0.05 M Tris-HCl (pH 7.5), 0.5 mM CaCl2 and incubated at 37oC with
shaking for 15 minutes. 0.5 ml of 10% trichloroacetic acid was added to stop the reaction and the
supernatant obtained after centrifugation at 10000 rpm for 1 minute at 4oC was measured at A400.
Absorption due to pigments produced by P. aeruginosa was subtracted from the A400 of the
filtered supernatant that was incubated without azocasein plus the A400 of the buffer and
azocasein incubated with no filtered supernatant.
.
25
Motility tests:
These tests were carried out according to the method of Rashid and Kornberg (2000) with
slight modifications. A small portion of a single colony was patched onto swim, swarm and
twitching plates from their respective overnight master plates. Swim plates were sealed with
parafilm to prevent dehydration and incubated at 30oC for 14 hours and the diameter of the
colony was measured. Swarm plates were incubated at 37oC for 14 hours and the diameter of the
colony was measured. Twitching motility plates were inoculated by stabbing through the
medium to the bottom of the plate and incubated at 37oC for 24 hours. The diameter of the
colony was measured at the agar/Petri dish interface. The solid medium was peeled and the plate
was stained with crystal violet and rinsed with water to measure the diameter of twitching
motility.
Conditions of growth:
Cells were grown in PsMM with 0.05% Triton-X as a surfactant and 0.2% glucose as the
carbon and energy source. Exogenous uracil or orotate was added at a concentration of 40 µg/ml.
50 ml cultures were grown in a 250 ml pyrex flask by shaking at 200 rpm and the cell density
was measured at 600 nm using a Perkin Elmer® Lambda 3A UV/VIS Spectrophotometer.
Preparation of cell extract:
Cells were harvested when they reached the exponential phase (O.D600 - 0.6 to 0.8) and
samples were collected by centrifugation at 3000xg for 30 minutes at 10oC in a Sorvall RT
6000B tabletop centrifuge. The cell pellet was resuspended in 20 ml 40 mM potassium
phosphate buffer pH 7.0 and centrifuged at 3000xg for 20 minutes at 10oC. This wash step was
26
carried out twice and the cell pellet obtained after the second washing was resuspended in 1ml
ATCase breaking buffer (2mM β-mercaptoethanol. 20 µM ZnSO4, 50 mM Tris-HCl pH 8.0, 20%
glycerol) and sonicated at 30 bursts per minute for 3 minutes using a Branson Sonifier Cell
Disrupter 200. The lysed cells were then centrifuged in a 1.5 ml microcentrifuge tube at 7000xg
for 4 minutes in a Savant high-speed centrifuge. Three hundred µl of the supernatant presumably
containing the membrane fraction was removed and saved to perform the dihydroorotate
dehydrogenase (DHOdehase) enzyme assay. The remaining cell lysate was centrifuged at
10000xg for 15 minutes and the supernatant was used for enzyme assays of the de novo
pyrimidine enzymes. The supernatant was dialyzed for 18-24 hours in 150 volumes ATCase
breaking buffer (without glycerol) at 4oC. Assays were performed the next day.
For the pyrimidine limitation (starvation) experiments the wild type and mutant strains of
P. aeruginosa were grown with uracil or orotate at 40 µg/ml until the cells reached an
absorbance of 0.48 at 600 nm (Haugaard & West, 2002). The cells were harvested as described
above and the cell pellet was resuspended in PsMM lacking either uracil or orotate and keeping
the other conditions sterile. After 2 hours the cells were collected and washed as described
above. The ultrasonicated cells were dialyzed and assays were performed as described below.
Enzyme Assays:
All enzyme assays were performed at 30oC and the measurements were obtained using a
Perkin Elmer® Lambda 3A UV/VIS Spectrophotometer. Specific activities of the enzymes are
expressed in nmol/min/mg protein and determined under conditions in which product formation
(or substrate utilization) is proportional to the amount of extract and the time of incubation.
Protein concentration was determined using the Bradford method (Bradford, 1976) with
27
lysozyme as the standard (Fig. 5).
Aspartate transcarbamoylase (ATCase): The method of Gerhart and Pardee, 1962 with
modification was used. The reaction mixture contained the following compounds in a total
volume of 1 ml: 40 µl tribuffer (51mM diethanolamine, 51 mM N-ethylmorpholine, and 100 mM
MES) adjusted to pH 9.5, 20 mM L-aspartic acid (monopotassium salt) pH 9.5, 5 mM carbamoyl
phosphate (dilithium salt), 20 µl cell extract and sterile double distilled water. The control tube
that was used as blank contained all components except cell extract and another control tube that
was used to check for background color contained all the components except carbamoyl
phosphate (the substrate). The tubes containing buffer, L-aspartate, water and cell extract were
preincubated at 30oC for 2 minutes. The substrate was then added and mixed by vortexing to
start the reaction and the mixture was incubated at 30oC for 10 minutes. The reaction was
stopped by adding 1 ml stop color mix of Prescott and Jones (1969) that contained freshly made
cocktail of 2:1 parts of antipyrine and monoxime solutions. The tubes were incubated at 65oC
water bath for 1 hour, cooled in the dark briefly following which the conversion of carbamoyl
phosphate to carbamoyl aspartate was read at O.D466. Specific activity was calculated using a
carbamoyl aspartate (CAA) standard curve (Fig. 6).
Dihydroorotase (DHO): The method of Beckwith et al., 1962 was used. The reaction mixture
contained in 1 ml volume 1 mM EDTA, 100 mM Tris/HCl buffer pH 8.6, 2 mM L-dihydroorotic
acid (DHO) dissolved in 0.1 M phosphate buffer, 50 µl cell extract and sterile ddH2O. Control
tubes were prepared to obtain a blank reading (without cell extract) and the background (without
DHO). All ingredients except dihydroorotate (DHO) were preincubated at 30oC for 2 minutes.
28
Fig. 5. Bradford protein standard curve of lysozyme. The absorbance A595 of lysozyme at
various concentrations was plotted by taking an average of three independent samples. With
permission of Dr. Dayna Brichta (Brichta, 2003)
29
Following the addition of dihydroorotate the reaction mixture was incubated for 10 minutes at
30oC. Stop color mix was added as described for the ATCase assay and incubated at 65
oC for 1
hour. After the tubes were cooled in the dark for a brief period of time the reverse reaction of
CAA from DHO was measured at O.D600. Specific activity was calculated using a CAA standard
curve (Fig. 6).
Dihydroorotate dehydrogenase (DHOdehase): The cell extract from the slow spin was used
for this assay. The assay mixture of 1 ml was prepared in a quartz cuvette containing 100 mM
Tris/HCl buffer (pH 8.6), 6 mM MgCl2, 1 mM DHO, 20 µl cell extract and sterile ddH2O. All
components except DHO were added in the quartz cuvette, mixed and preincubated at 30oC for 2
minutes. The reaction was initiated by adding DHO for its conversion to orotate by oxidation and
the absorbance at 290 nm was measured as zero minutes (T0). Following the incubation at 30oC
for 10 minutes the absorbance at (T10) was measured. An increase in the absorbance of 1.93 is
equivalent to a change in substrate concentration of 1 mM (Beckwith et al., 1962; Schwartz and
Neuhard, 1975).
Orotate phosphoribosyl transferase (OPRTase): The assay mixture of 1 ml was prepared in a
quartz cuvette containing 100 mM Tris/HCl buffer pH 8.6, 6 mM MgCl2, 0.25 mM orotate, 0.6
mM 5-phosphoribosyl-1-pyrophsphate (PRPP), 20 µl cell extract and sterile double distilled
water. All the components except PRPP were added in the quartz cuvette, mixed and
preincubated at 30oC for 2 minutes. The reaction was initiated by adding PRPP for its conversion
to orotidine-5’-monophosphate (OMP) and the absorbance at 295 nm was measured as zero
minutes (T0). Following the incubation at 30oC for 10 minutes the absorbance at (T10) was
30
y = 0.005x
0
0.1
0.2
0.3
0.4
0.5
0.6
0 10 20 30 40 50 60 70 80 90 100
nmol CAA
A466 nm
Fig. 6. Carbamoyl aspartate (CAA) standard curve. Various concentrations of carbamoyl
aspartate (nmol) were measured at 466nm by taking an average of three independent samples.
With permission of Dr. Kamran Azad (Azad, 2005).
31
measured. A decrease in the absorbance of 3.67 is equivalent to a change in substrate
concentration of 1 mM (Beckwith et al., 1962; Schwartz and Neuhard, 1975).
Orotidine-5’-monophosphate decarboxylase (OMPdecase): The assay mixture of 1ml was
prepared in a quartz cuvette containing 100 mM Tris/HCl buffer pH 8.6, 6 mM MgCl2, 0.2 mM
OMP, 20 µl cell extract and sterile double distilled water. All components except OMP were
added in the quartz cuvette, mixed and preincubated at 30oC for 2 minutes. The reaction was
initiated by adding OMP for its conversion to uridine-5’-monophosphate (UMP) and the
absorbance at 290 nm (West, 1997) was measured as zero minutes (T0). Following the incubation
at 30oC for 10 minutes, the absorbance at (T10) was measured. A decrease in the absorbance of
1.38 is equivalent to a change in substrate concentration of 1 mM (Beckwith et al., 1962;
Schwartz and Neuhard, 1975).
Isolation of P. aeruginosa genomic DNA:
The basic protocol of isolation of genomic DNA (Ausubel et al., 1997) was used with
slight modifications. Wild type P. aeruginosa (PAO1) cells were inoculated from an overnight
culture grown in PsMM with 0.2% glucose into a 5 ml PsMM glucose (0.2%) broth. Cells are
collected by centrifugation of a 3 ml culture in a microcentrifuge tube at 4oC, 10000xg for 4
minutes. After the supernatant was removed, the cell pellet was resuspended in 250 µl TENS
(0.1 M NaCl, 10 mM Tris pH 8.0, and 10 mM EDTA) solution. Cells were lysed by the addition
of 25 µl of 10% SDS solution and incubated at 65oC for 10 minutes. 25 µl of proteinase K (5
mg/ml) was then added and the suspension incubated at 37oC for 30 minutes. Extraction of the
genomic DNA from the protein and cell debris was carried out using equal volume of phenol :
32
chloroform, and chloroform : isoamyl alcohol. The DNA was precipitated by adding 100 µl of
7.5 M ammonium acetate and 750 µl of precooled 95% ethanol. The suspension was mixed by
inversion and incubated at -80oC for 30 minutes; followed by centrifugation at 10000xg at 4
oC
for 5 minutes. The ethanol was carefully poured off and the cell pellet was air-dried overnight
and then resuspended in 250 µl of TE (1 M Tris pH 8.0 and 0.5 M EDTA pH 8.0) containing
RNaseA at a concentration of 10 µg/ml. DNA was brought into solution by gentle mixing,
vortexing being avoided to prevent shearing.
Rapid plasmid isolation:
The procedure was modified from Zhou et al. (1990). Plasmid was extracted from a 5 ml
overnight culture obtained from a single colony. 1.5 ml culture was centrifuged at 10000xg for
10 seconds, supernatant discarded and the pellet resuspended in 1.5 ml culture, and centrifuged.
The pellet obtained after centrifugation at 10000xg for 10 seconds was resuspended in about 50-
100 µl of supernatant. Freshly prepared TENS solution in a quantity of 300 µl was added and
vortexed for 2-5 seconds until the mixture became sticky. A volume of 150µl 3.0 M sodium
acetate (pH 5.2) was added to the mixture and vortexed for 2-5 seconds. The mixture was then
centrifuged at 10000xg for 2 minutes and supernatant containing the plasmid transferred to a new
microcentrifuge tube. The plasmid DNA was precipitated by adding 900 µl of 95% ethanol
(precooled). The solution was mixed by inverting the tube and incubated at -80oC for at least 30
minutes, after which the tube was centrifuged at 10000xg for 2 minutes at 4oC to pellet the
plasmid DNA. The supernatant was aspirated and the DNA pellet washed with 1 ml cold 70%
ethanol twice. This washed pellet was dried under vacuum and resuspended in 40 µl sterile
ddH2O containing 2 µl RNaseA (10 mg/ml).
33
Preparation of competent cells:
Competent cells were made by the chemical method according to Mandel and Higa
(1970) with slight modifications. 50 ml LB culture was inoculated with an overnight 5 ml culture
in 1:100 ratio and the cells were grown at 37oC with shaking to an O.D600 of about 0.4. The cells
were transferred to a 50 ml conical flask, incubated on ice for 30 minutes and harvested for 20
minutes at 1300xg at 4oC. The cell pellet obtained was washed with sterile 20 ml ice cold 0.1 M
CaCl2, incubated on ice for 20 minutes and then the cells were centrifuged for 20 minutes at
1300xg at 4oC. The cell pellet was resuspended with 812 µl of sterile ice cold 0.1 M CaCl2,
packed in ice and left at 4oC overnight. Next morning 188 µl of sterile 80% glycerol was added
to the cells and aliquots of 100 µl were distributed in 1.5 ml microcentrifuge tubes. The
competent cells were stored at -80oC until used. Typically the competent cells are viable and
good for 5-6 months.
Transformation:
Transformation in E. coli DH5α was carried out by thawing the competent cells on ice.
30-200 ng of plasmid DNA was added to the competent cells gently mixed and incubated on ice
for 20-30 minutes. The cells were then heat-shocked at 42oC for 90 seconds and then placed on
ice immediately for 2 minutes. Sterile LB broth in 1ml quantity was added to them and the cells
were harvested after incubating at 37oC for 1 hour. Cells were plated on LB plates with the
appropriate antibiotic and IPTG and X-gal for blue-white selection of recombinants.
Ligations:
Cohesive ligations were performed according to the procedure of Sambrook et al. (1989).
34
Vector and insert were added in 1:3 ratio and T4 ligase and the buffer (New England Biolabs)
were added in final volume of 10 µl reaction and incubated at 16oC for 16 hours.
Construction of plasmid pPR2 and pPR2∆D:
The 970 bp pyrD gene was amplified from the genomic DNA by performing a
polymerase chain reaction (PCR). Primers (Table 2) were designed approximately 300 bp
upstream and downstream of the gene and were flanked with restriction sites EcoRI on the 5’ end
and BamHI on the 3’ end for convenience in cloning. The PCR conditions that were optimized to
get the 1570 bp product size are listed below (Table 2). The amplified fragment was digested
with restriction enzymes EcoRI and BamHI and ligated into the multiple cloning site of the
vector pGEX2T (4.9 Kb) digested with the same restriction enzymes (Fig. 7). The ligated
mixture was transformed into E. coli DH5α. Transformed white colonies on LB Amp100 X-gal
plates were selected and grown in 5 ml LB broth for extraction of the plasmid. EcoRI digestion
of the plasmid resulted in a total fragment size of 6.47 Kb. Sequencing confirmed the insertion of
the pyrD fragment into the vector.
The restriction enzyme SalI cuts the pyrD gene at two sites (Fig. 8). A digest of the
plasmid pPR2 with SalI resulted in two linearized sizes of DNA, a 96 bp band and a 6.37 Kb
band on an agarose gel. The DNA was extracted from linearized 6.37 Kb band using the
SephaglasTM
BandPrep Kit (Amersham Pharmacia Biotech Inc.) and ligated together. The
ligation mixture was transformed into E. coli DH5α and screened for transformants (white
colonies) on LB plates containing Amp100 and X-gal. Transformed white colonies were picked
and grown at 37oC in 5 ml LB broth with Amp100. The plasmid was extracted and digested with
EcoRI resulting in a 6.37 Kb plasmid, pPR2∆D. Restriction digest with EcoRI and BamHI
35
Table 2. Forward and Reverse primers incorporated with restriction sites EcoRI and BamHI were
designed for the amplification of the pyrD gene from P. aeruginosa chromosomal gene.
Primer 1F: 5’ CCG GAA TTC GGT TCA TTC AAG CGG GGG 3’
Primer 1R: 5’CGC GGA TCC CAT AAC CGG TAA ATC TCG TG 3’
PCR Conditions:
95oC for 5 minutes - Denaturation
95oC for 1 minute - Denaturation
29times 56oC for 1 minute - Annealing
72oC for 1 minute 30 seconds - Polymerization
72oC for 5 minutes - Extension
4oC for ∞
36
Fig. 7. Construction of the plasmid pPR2. The 1.57 Kb insert containing the pyrD gene was
force-ligated into the plasmid pGEX2T that was 4.9 Kb in size using the restriction enzymes
EcoRI and BamHI. The total size of the new plasmid pPR2 was about 6.5 Kb.
37
Fig. 8. Sequence of P. aeruginosa pyrD (www.pseudomonas.com). The restriction site SalI is
highlighted in blue and the start and stop codons are in red.
ATGTACACCCTGGCCCGCCAGCTGCTGTTCAAACTGTCCCCGGAAACCTCCCACGA
ACTGTCCATCGACCTGATCGGGGCCGGCGGCCGCCTGGGCCTGAACCGCCTGCTGA
CGCCCAAGCCGGCCAGTCTGCCGGTCTCTGTGCTGGGCCTGGAGTTTCCCAACCCG
GTGGGGCTGGCCGCGGGCCTGGACAAGAACGGCGACGCCATCGACGGGTTCGGCC
AGCTCGGTTTCGGCTTCATCGAGATCGGCACCGTCACTCCGCGACCGCAGCCGGGC
AATCCCAGGCCGCGCCTGTTCCGCCTGCCGCAAGCCAACGCGATCATCAATCGCAT
GGGTTTCAACAACCACGGCGTCGACCACCTGCTGGCGCGGGTGCGTGCGGCGAAGT
ATCGCGGCGTGCTGGGGATCAACATCGGCAAGAACTTCGATACCCCGGTGGAGCGC
GCGGTCGACGACTACCTGATCTGCCTGGACAAGGTCTACGCCGACGCCAGCTACGT
CACCGTCAACGTCAGCTCGCCGAACACGCCCGGCCTGCGCAGCCTGCAGTTCGGCG
ATTCGCTCAAGCAACTGCTGGAGGCACTGCGCCAGCGTCAGGAAGCGCTGGCGCTG
CGGCATGGCCGGCGGGTACCGCTGGCGATCAAGATCGCCCCGGACATGACCGACG
AAGAGACCGCGCTGGTCGCCGCGGCGCTGGTCGAGGCGGGGATGGACGCGGTGAT
CGCGACCAATACCACTCTCGGCCGCGAAGGCGTGGAAGGCCTGCCGCATGGCGAC
GAGGCGGGTGGCCTGTCCGGCGCGCCGGTGCGGGAAAAGAGCACGCACACGGTGA
AGGTGCTGGCCGGCGAACTGGGCGGACGGCTGCCGATCATCGCCGCCGGCGGTATC
ACCGAGGGCGCGCACGCGGCGGAGAAGATCGCCGCCGGGGCGAGCCTGGTGCAGA
TCTATTCGGGTTTCATCTACAAGGGACCGGCGTTGATCCGCGAAGCGGTGGACGCC
ATCGCCGCATTGCCGCGGCGGAATTGA
38
resulted in the excision of the deleted pyrD band.
Construction of plasmid pPR3:
The deleted pyrD fragment was digested out of the pGEX2T vector with restriction
enzymes EcoRI and BamHI and ligated into the gene replacement vector pEX18Gm (5831 bp)
that was digested with the same restriction enzymes (Fig. 9). The ligation mixture was
transformed into E. coli DH5α and screened for white colonies on LB plates supplemented with
Gm15 and X-gal.
The recombinants (white colonies) obtained were selected and grown overnight at 37oC
in a 5 ml LB broth with Gm15 for plasmid isolation. Digestion of the plasmid with EcoRI resulted
in a 7.2 Kb band and double digestion with EcoRI and BamHI excised the deleted pyrD band.
The positive clones were patched on to LB plates containing Gm15 first and then on to LB plates
containing Suc10%. Colonies that were Gm15 resistant and Suc10% sensitive were used further for
genetic manipulation.
Gene replacement:
Escherichia coli SM10 cells were grown in 50 ml LB broth with Kn25 and competent
cells were made using the chemical method as described previously. The plasmid pPR3 that was
GmR and Suc
S was transformed into E. coli SM10 cells and screened for colonies that grew on
LB Gm15 Kn25 plates grown overnight at 37oC. P. aeruginosa wild type cells (recipient) were
grown in 5 ml LB broth at 42oC to alter its host restriction modification system. The SM10 cells
carrying the plasmid (donor) were grown in a 5 ml LB broth with Gm15 and Kn25 at 37oC. Both
the donor and recipient cells were grown to an O.D600 of 0.6-0.8 and mixed in a microcentrifuge
39
Fig. 9. Construction of the plasmid pPR3. The 1.47 Kb insert containing the deleted pyrD gene
was force-ligated into the vector pEX18Gm that was about 5.7 Kb using the restriction enzymes
EcoRI and BamHI. The new plasmid pPR3 was about 7.3 Kb in size.
40
tube in the ratio of 20:1 recipient:donor in a total volume of 200 µl. The cells were centrifuged at
10000xg for 1 minute and the pellet was resuspended in 30 µl saline by pipetting up and down
and spotted onto a LB plate.
After overnight incubation at 37oC the cells were scraped from the LB plate and serial
dilutions made with saline up to 10-6. From each dilution tube 0.1 ml were plated on PIA plates
supplemented with Gm100 and incubated overnight at 37oC. The colonies that resulted from
single crossovers were patched onto PIA Gm100 plates first and then onto PIA sucrose (Suc
10%), for the selection of GmR and Suc
S phenotype. These cells were grown in a 5 ml LB broth
overnight at 37oC without any antibiotic. Serial dilutions were made up to 10
-4 and 0.1 ml plated
onto PIA sucrose. Emerging colonies were patched onto PIA sucrose plates first and onto PIA
Gm100, for the selection of GmS and Suc
R phenotype (double crossover). These colonies were
patched onto PsMM with 0.2% glucose plates first and then onto PsMM with 0.2% glucose and
uracil at 40 µg/ml. Colonies that appeared only on the latter plates were the auxotrophic mutant
of P. aeruginosa with a defect in the gene responsible for the enzyme dihydroorotate
dehydrogenase (pyrD). The pyrD mutant obtained was designated PAOPR04. The mutant
obtained was confirmed by sequencing and also by enzyme assay for dihydroorotate
dehydrogenase. An overview of the gene replacement method is shown in Fig. 10.
Southern blot:
A Southern blot was performed according to Sambrook et al., 1989 with slight
modifications. Genomic DNA was extracted from the wild type and the mutant strain.
Restriction digest with SalI (40U) was performed on 10 µg of DNA and digested overnight at
37oC. Half of the digest was loaded on 1% agarose gel made with TAE and run at 100 volts for
41
Fig. 10. Scheme for the gene replacement method to generate double crossovers. The deleted
pyrD gene was introduced into PAO1 resulting in an auxotroph that was unable to grow on
Pseudomonas minimal medium with 0.2% glucose.
PAO1 (42º C) pPR3 in SM10 (37º C)
20:1
Grown at 37ºC for 6 hours
Dilutions and platings on PIA Gm100
Grown in LB at 37ºC overnight
Dilutions and platings on PIA 10% sucrose
Patch on PIA Gm100 and Suc10% (GmS, Suc
R)
Patched on PIA Suc10% and Gm100 (Suc
s, Gm
R)
Patch onto PsMM with 0.2% glucose (no uracil) and with uracil (40 µg/ml)
42
the first 30 minutes, 20 volts for the next 12 hours and 100 volts for the last 15 minutes. The gel
was then subjected to two 15 minute washes in 100 ml denaturing solution (0.5 M NaOH, 1.5 M
NaCl) with gentle rocking at room temperature, followed by three washes with 100 ml
neutralizing solution (0.5 M Tris pH 7.4, 1.5 M NaCl). A gel tray wrapped with Whatman®
paper was placed in a dish containing the transfer solution. The gel, denatured and neutralized,
was placed with the wells facing down on the gel tray. A positively charged nylon membrane
(Nalgene) was placed on the gel carefully to avoid the formation of air bubbles underneath and
cut on one corner for the right orientation. Three properly cut Whatman papers (approximately 3
mm in thickness) were placed on top of the membrane. A stack of paper towels cut with the
exact size of the membrane were placed on top of the Whatman paper and a heavy weight placed
on them. An overnight transfer was allowed replacing the wet paper towels with fresh dry ones.
The following day the apparatus was disassembled and nylon membrane placed face up on a
Whatman paper wetted with 20X SSC. This was placed in a UV crosslinker (UVP CL-1000) and
exposed to ultraviolet radiation for 40 seconds at 1500 x 100 µJ/cm2. The membrane was then
rinsed briefly in 20X SSC and dried on Whatman paper for 10 minutes.
The DIG (dioxigenin) high prime DNA labeling and detection starter kit II provided by
Roche® was used for preparation of the probe, hybridization and detection. The probe was
created, labeled with DIG high prime and quantified with DIG-labeled control DNA. Antibody
anti-dioxigenin-AP was used for immunodetection and the chemiluminescence was observed
using a chemiluminiscent substrate CSPD®, ready-to-use to identify and quantitate the probe.
The membrane was prehybridized with DIG Easy Hyb solution in the hybridization oven for 30
minutes at 42oC (hybridization temperature). Meanwhile the DIG-labeled probe/DIG Easy Hyb
43
mixture was prepared and added to the membrane and incubated at the optimum temperature
(50oC) for 18 hours with gentle agitation.
Stringency washes were carried out twice for 5 minutes at room temperature with 50 ml
2X SSC and 0.1% SDS under constant agitation. The next two washes were performed for 15
minutes at 68oC with 0.5X SSC and 0.1% SDS (prewarmed to wash temperature) under constant
agitation.
For immunologic detection the membrane was rinsed with washing buffer for 5 minutes,
followed by 30 minute incubation at room temperature with 100 ml blocking solution. Antibody
solution (20 ml) was added to the membrane and incubated for 30 minutes. This was followed by
two 15 minute washes with 100 ml washing buffer and finally equilibrated with 20 ml detection
buffer for 5 minutes. For detection by chemiluminescence 1 ml CSPD® ready-to-use was spread
on the membrane and incubated for 5 minutes at room temperature, followed by 10 minute
incubation at 37oC. The treated membrane was placed in a development folder, exposed to x-ray
film and developed using an automated developer machine.
Construction of pPR4:
The vector used in the construction of this plasmid pPR4 was pVDZ’2, which is a low
copy number broad host range vector (Deretic et al., 1987). The intact pyrD gene with
approximately 300 bp upstream and downstream, a total of 1570 bp was amplified using PCR,
and purified. The purified DNA was digested with restriction enzymes EcoRI and BamHI and
force-ligated into the 20500 bp pVDZ’2 vector that was also digested with EcoRI and BamHI.
The ligated mixture after 16 hours was transformed into chemically competent E. coli One Shot®
TOP10 (Invitrogen) cells and transformants were plated onto LB plates supplemented with
tetracycline 15 µg/ml and 0.2% X-gal. The recombinant clones which had white colonies were
44
tested for the presence of the insert, by plasmid prep and restriction digest. The plasmid that was
constructed was named pPR4 and had a total size of 22070 bp (Fig. 11).
Complementation of pyrD:
The E. coli strain MA1004 obtained from E. coli Genetic Stock Center was used for this
experiment. MA1004 is a pyrD mutant, incapable of growing on EcMM with glucose as the
carbon and energy source, unless supplemented with exogenous uracil. Transformation of the
plasmid pPR4 into chemically competent MA1004 cells was carried out and plated on LB
medium with tetracycline at 15 µg/ml. Transconjugants that were obtained were further screened
by patching first onto EcMM with 0.2% glucose and tetracycline (15 µg/ml) and then onto
EcMM with 0.2% glucose, tetracycline (15 µg/ml) and uracil (40 µg/ml). Plasmid was extracted
from the colonies obtained from the latter plate and digested with restriction enzymes EcoRI and
BamHI to confirm the presence of the pyrD gene.
45
pyrD disrupted MCS
disrupted MCS
tet A
tet R
lacZ' EcoRI
BamHI
pPR4
22070 bp
Fig. 11. Construction of the plasmid pPR4. The insert containing the 1.57 Kb intact pyrD gene
was force-ligated into the vector pVDZ’2 that was 20.5 Kb in size using the restriction enzymes
EcoRI and BamHI. The new plasmid pPR4 obtained was 22.07 Kb in size.
46
CHAPTER III
RESULTS
PCR amplification of the pyrD gene:
The 980 bp active pyrD gene and approximately 300 bp upstream and downstream, a
total of 1570 bp was amplified by a polymerase chain reaction. The primers and optimized PCR
conditions are shown in Table 2. The 1.57 Kb PCR product obtained is shown in Fig. 12.
Creation of the deletion mutant:
The 1570 bp fragment was force-ligated into the vector pGEX2T, resulting in the plasmid
pPR2, of total size of 6.57 Kb (Fig. 13). A SalI digest performed on this plasmid caused a
deletion of 101 bp as shown in the Fig. 14.
Characterization and verification of the pyrD mutant:
The pyrD mutant PAOPR04 was created by allelic exchange of the native pyrD with the
deleted version. The auxotroph was confirmed phenotypically by its inability to grow on PsMM
with glucose as the carbon and energy source unless provided with an exogenous supplement of
uracil or orotate at 40 µg/ml concentration. The biochemical assay that measures the specific
activity of the pyrimidine biosynthetic enzymes showed no activity for the dihydroorotate
dehydrogenase enzyme encoded by the pyrD gene. To verify the deleted pyrD gene, PAO1
genomic DNA and PAOPR04 genomic DNA was digested with SalI independently. The probe
created for the Southern blot was the 1570 bp containing the intact pyrD gene resulting in 656
bp, 819 bp and 101 bp bands for PAO1. PAOPR04 had 656 bp and 819 bp bands, but the 101 bp
band was missing, due to the deletion of the 101 bp caused by excision of SalI (Fig. 15).
47
Complementation of E. coli pyrD- with P. aeruginosa pyrD:
The 1.57 Kb fragment was ligated into a broad host range vector pVDZ’2 resulting in the
plasmid pPR4. The plasmid pPR4 was able to complement the E. coli pyrD auxotroph MA1004
after transformation. Growth was observed on EcMM with 0.2% glucose and 15 µg/ml
tetracycline. The controls for the complementation studies are listed in Table 3.
Effects of PAOPR04 on the pyrimidine biosynthetic pathway in comparison with PAO1:
The effect of the mutant strain on the pyrimidine biosynthetic pathway was tested by
measuring the specific activity of enzymes of the de novo pathway and compared with the
activity of pyrimidine enzymes of the wild type strain. Cells were grown in PsMM with 0.2%
glucose as the carbon and energy source, supplemented with either uracil or orotate at a
concentration of 40 µg/ml as described in Materials and Methods. PAO1 cells were also starved
for different compounds such as carbon source, uracil or orotate. PAOPR04 cells were starved
for uracil or orotate. The conditions for starvation are described in Materials and Methods.
Dayna Brichta observed repression (Brichta, 2003) in the pyrC, pyrC2 double mutant
(Brichta et al., 2004), which was different from the results of Isaac and Holloway (1968). The
pyrD mutant was grown with uracil or orotate independently to see if there was any repression of
the pyrimidine pathway.
48
Fig. 12. PCR amplified 1.57 Kb band containing the pyrD gene of Pseudomonas aeruginosa run
on a 1% agarose gel. The PCR primers and optimized PCR conditions are described in Table 2.
2.0 Kb
1.5 Kb
1.0 Kb
1.57 Kb
1 Kb Ladder
49
Fig. 13. Plasmid pPR2 digested with restriction enzymes EcoRI and BamHI. The 1.57 Kb insert
containing the pyrD gene was separated from the 4.9 Kb vector pGEX2T.
1Kb Ladder
1.57 Kb
4.9 Kb
1.5 Kb
5.0 Kb
50
Fig. 14. SalI digest on the plasmid pPR2. The linearized plasmid resulting from the digest was
ligated to form the deleted pyrD gene. The excised 101 bp band is shown on this 1% agarose gel.
0.101 Kb
0.5Kb
6.4 Kb
1 Kb Ladder
51
Fig. 15. Southern blot analysis of PAO1 and PAOPR04. The picture on the right shows SalI
digest on the genomic DNA of the two strains run on 1% agarose gel. The picture on the left
shows the expected bands obtained after developing the x-ray film.
PAOPR04 PAO1 2 Log Ladder PAOPR04 PAO1
1.0 Kb
0.1 Kb
0.82 Kb
0.65 Kb
0.101 Kb
52
Plasmid/Strain Medium Result
pVDZ’2
MA1004
MA1004
E. coli K12
pPR4 in MA1004
E. coli minimal medium with
Tc 15 µg/ml
E. coli minimal medium
E. coli minimal medium with
uracil 40 µg/ml
E. coli minimal medium
E. coli minimal medium with
Tc 15 µg/ml
No growth
No growth
Growth
Growth
Growth
Table 3. Complementation of E. coli auxotroph MA1004 with P. aeruginosa pyrD. The plasmid
and the strains used in this part of the study are listed. Transformants were selected on E. coli
minimal medium with appropriate requirements. The results obtained are listed.
53
Specific activity values are reported in nmol/min/mg protein. Repression was seen in
PAO1 cells when grown in PsMM with glucose and uracil as compared with that of PAO1 cells
grown in PsMM with glucose (Fig. 16). Repression was seen in the case of ATCase (0.5-fold),
OPRTase (0.6-fold), and OMPdecase (0.8-fold). Dihydroorotate dehydrogenase showed 8.2-fold
derepression and an insignificant change was observed for DHOdehase. The exogenous supply
of orotate to PAO1 cells brought about derepression of ATCase (1.03-fold), DHOase (5.3-fold),
DHOdehase (1.1-fold), OPRTase (1-fold) and OMPdecase (insignificant level) when compared
with PAO1 cells grown in the presence of glucose (Fig. 16).
In order to compare the specific activity data of pyrimidine enzymes of PAO1 with
PAOPR04, cells were grown in PsMM with glucose and uracil (Fig. 17). Repression of ATCase
(0.9-fold), and OPRTase (0.9-fold) was observed while DHOase showed 8.2-fold derepression
and OMPdecase showed 1.09-fold derepression. On the other hand all the enzymes showed
derepression when PAOPR04 cells were grown in orotate (Fig. 18); ATCase (1.3-fold), DHOase
(3-fold), OPRTase (1.2-fold), and OMPdecase (1.2-fold). As expected no activity was observed
for DHOdehase enzyme in the mutant strain even when the cell extract volume was doubled.
Derepression usually occurs when cells are grown under limited levels of pyrimidines as
reported in Salmonella typhimurium (Smith et al, 1980). This finding was tested in P. aeruginosa
by performing starvation experiments on the mutant strain. The conditions to test the effect of
pyrimidine limitation on the regulation of the pyrimidine pathway are described in Materials and
Methods
Comparing PAO1 specific activity data for starved versus non-starved conditions gave
interesting results. No activity was observed for DHOase when cells were limited for glucose as
the carbon and energy source. Derepression was observed for ATCase (1.06-fold), DHOdehase
54
(4-fold), and OPRTase (1.05-fold), whereas a 1.8-fold repression was observed for OMPdecase
(Fig. 19). The pyrD mutant, PAOPR04 could not be starved for glucose as the carbon and energy
source in PsMM since it is an auxotroph and requires exogenous pyrimidines for growth. Hence,
the cells were subjected to uracil or orotate limitation independently and compared with non-
starved conditions to check for derepression. When starved for uracil (Fig. 20), PAOPR04
showed derepression of ATCase (3-fold), OPRTase (2-fold) and OMPdecase (2.8-fold). A two-
fold repression was observed for DHOase. Growth under limited conditions of orotate (Fig. 21)
caused slight derepression of ATCase (1.2-fold) and OMPdecase (1.3-fold); and repression of
DHOase (2.9-fold) and OPRTase (insignificant level) in PAOPR04.
Growth analysis:
Growth curves were performed for the wild type and mutant strain in minimal and rich
media. The media were inoculated with fresh overnight broth cultures and their growth
monitored using a spectrophotometer as described in Materials and Methods. All the cultures
were started at the same optical density to maintain a consistent basal starting point.
Table 4 shows the generation times of PAO1 and PAOPR04 grown in PsMM with
glucose as the carbon and energy source, and with the exogenous supply of orotate or uracil
independently. The PAO1 cells grew faster when supplied with either uracil or orotate, whereas
PAOPR04 cells prefer uracil as a supplement to orotate. The logarithmic curves are shown in
Fig. 22. The striking difference amongst the growth patterns of the two strains was that of
PAOPR04 supplied with orotate.
55
0
50
100
150
200
250
300
350
400
450
ATCase DHOase DHOdehase OPRTase OMPdecase
nm
ol/m
in/m
g p
rote
in
PAO1
PAO1 + U
PAO1 + O
Fig. 16. Specific activity of the pyrimidine enzymes from wild type PAO1 expressed in
nmol/min/mg protein. PAO1 cells were grown in Pseudomonas minimal medium with 0.2%
glucose, and uracil or orotate at a concentration of 40 µg/ml. Cells were also grown with 0.2%
glucose only. Assays were performed individually three times and the error bars reported.
56
0
50
100
150
200
250
300
350
400
450
ATCase DHOase DHOdehase OPRTase OMPdecase
nm
ol/m
in/m
g p
rote
in
PAO1
PAO1 + U
PAOPR04 + U
Fig. 17. Specific activity of pyrimidine enzymes from wild type PAO1 and mutant PAOPR04
expressed in nmol/min/mg quantity of protein. PAO1 and PAOPR04 cells were grown in
Pseudomonas minimal medium with 0.2% glucose and uracil, supplied exogenously at a
concentration of 40 µg/ml. Assays were performed individually three times and the error bars
reported.
57
0
100
200
300
400
500
600
700
ATCase DHOase DHOdehase OPRTase OMPdecase
nm
ol/m
in/m
g p
rote
in
PAO1
PAO1 + O
PAOPR04 + O
Fig. 18. Specific activity of the pyrimidine enzymes expressed in nmol/min/mg quantity of
protein. PAO1 and PAOPR04 cells were grown in Pseudomonas minimal medium with 0.2%
glucose and orotate, supplied exogenously at a concentration of 40 µg/ml. Assays were
performed individually three times and error bars reported.
58
0
50
100
150
200
250
300
350
400
450
Not starved Starved
nm
ol/m
in/m
g p
rote
in
ATCase
DHOase
DHOdehase
OPRTase
OMPdecase
Fig. 19. Comparative analysis of specific activity of pyrimidine enzymes for PAO1 grown in
Pseudomonas minimal medium with 0.2% glucose. The condition where PAO1 cells were grown
without starvation is described in Materials and Methods. Cells were starved for glucose for 2
hours before harvesting, as described in Materials and Methods. All assays were performed
individually in triplicate and the error bars reported.
59
0
100
200
300
400
500
600
700
Not starved Starved
nm
ol/m
in/m
g p
rote
inATCase
DHOase
DHOdehase
OPRTase
OMPdecase
Fig. 20. Comparison of specific activity of pyrimidine enzymes for PAOPR04 cells grown in
Pseudomonas minimal medium with 0.2% glucose with uracil supplemented at a concentration
of 40 µg/ml. The condition where PAOPR04 cells were grown without starvation is described
under Materials and Methods. Cells were starved for uracil for 2 hours before harvesting as
described under Materials and Methods. All assays were performed individually in triplicate and
the error bars reported.
60
0
100
200
300
400
500
600
700
800
Not starved Starved
nm
ol/m
in/m
g p
rote
in
ATCase
DHOase
DHOdehase
OPRTase
OMPdecase
Fig. 21. Comparison of specific activity of pyrimidine enzymes for PAOPR04 cells grown in
Pseudomonas minimal medium with 0.2% glucose with orotate supplemented at a concentration
of 40 µg/ml. The condition where PAOPR04 cells were grown without starvation is described
under Materials and Methods. Cells were starved for orotate for 2 hours before harvesting as
described under Materials and Methods. All assays were performed individually in triplicate and
the error bars reported.
61
PAO1 PAO1
with
uracil
PAO1
with
Orotate
PAOPR04
with
uracil
PAOPR04
with
orotate
60 54 54 84 102
Table 4. Generation time in minutes for PAO1 and PAOPR04 grown in Pseudomonas minimal
medium with 0.2% glucose and uracil (40 µg/ml) or orotate (40 µg/ml). PAO1 cells were also
grown in Pseudomonas minimal medium with 0.2% glucose only.
62
0.01
0.1
1
10
00.5 1
1.5 2
2.5 3
3.5 4
4.5 5
5.5 6
6.5 7
7.5 8
8.5 9
9.5 10
10.5 11
11.5 12
12.5 13
13.5 14
14.5 15
Time in hours
log #
cells
at 600nm
PAO1
PAO1 with U
PAO1 with O
PAOPR04 with U
PAOPR04 with O
Fig. 22. Growth curves of PAO1 and PAOPR04 cells grown in Pseudomonas minimal medium
with 0.2% glucose as the carbon and energy source. Cells were grown in the presence or absence
of uracil or orotate supplied at a concentration of 40 µg/ml. An average of three individual
growth curves is reported.
63
As expected growth curve analysis observed in rich medium (PTSB) shows a different set
of results when compared to that in minimal medium (Table 5). The generation time of PAO1
cells grown in rich medium was almost half that of minimal medium. All the growth curves seem
very similar. The wild type and the mutant strain give the same generation time when grown in
the presence or absence of orotate. Exogenously supplied uracil caused the cells to grow for a
longer period of time (Fig. 23).
Quantitation of pigment production:
Pseudomonas aeruginosa produces two pigments: pyocyanin, a greenish yellow pigment
and pyoverdin, a blue green pigment. It has been shown that these pigments contribute to the
virulence of the organism (Reimmann et al., 1997). This aspect of virulence was tested for the
wild type and mutant strain. Two different media were used to enhance these pigments
independently. Pyocyanin was enhanced using King’s A medium and pyoverdin was enhanced
using King’s B medium. The composition of the two media and the extraction of these pigments
are described in detail under Materials and Methods. The production of pyocyanin in the mutant
was slightly less than that of the wild type when grown in King’s A medium with no supplement,
and it was 2.5-fold higher in the mutant when uracil was supplied. Supplementing with
exogenous orotate brought about a 1.4-fold increase in pyocyanin levels in the mutant (Fig. 24).
Pyoverdin levels were found to be the slightly higher in the mutant than the wild type when
grown in King’s B medium with no additional supplement, but a 2.1-fold increase was observed
in the mutant when supplied with uracil. Supplementing with exogenous orotate caused the
mutant to have an increased pyoverdin level by a factor of 1.7 (Fig. 25).
64
PAO1 PAO1
with
uracil
PAO1
with
orotate
PAOPR04 PAOPR04
with
Uracil
PAOPR04
with
orotate
36 60 36 36 48 36
Table 5. Generation time in minutes for PAO1 and PAOPR04 grown in PTSB medium in the
presence of uracil (40 µg/ml) or orotate (40 µg/ml). Cells were also grown in PTSB medium
without supplements.
65
0.01
0.1
1
10
00.5 1
1.5 2
2.5 3
3.5 4
4.5 5
5.5 6
6.5 7
7.5 8
8.5 9
9.5 10
10.5 11
Time in hours
log#cells
at 600nm
PAO1
PAO1 with U
PAO1 with O
PAOPR04
PAOPR04 with U
PAOPR04 with O
Fig. 23. Growth curves of PAO1 and PAOPR04 cells grown in PTSB medium in the presence of
uracil or orotate supplied at a concentration of 40 µg/ml. Cells were also grown in PTSB medium
without supplements. An average of three individual growth curves is reported.
66
0
0.5
1
1.5
2
2.5
3
3.5
4
4.5
PAO1 PAOPR04
ug/5
ml
No supplement
U40ug/ml
O40ug/ml
Fig. 24. Measurement of pyocyanin for PAO1 and PAOPR04 cells grown in King’s A medium
in the presence of uracil or orotate supplied at a concentration of 40 µg/ml. PAO1 and PAOPR04
cells were also grown in King’s A medium without supplements. Cells were grown for 24 hours
with shaking at 37oC till they reached a similar optical density at 600 nm. Pyocyanin was
extracted from 5 ml supernatant as described in Materials and Methods and µg quantities
reported. Samples were grown individually three times and the error bars reported.
67
0
0.05
0.1
0.15
0.2
0.25
0.3
PAO1 PAOPR04
A405/A
600
No supplement
U40ug/ml
O40ug/ml
Fig. 25. Measurement of pyoverdin for PAO1 and PAOPR04 cells grown in King’s B medium in
the presence of uracil or orotate supplied at a concentration of 40 µg/ml. PAO1 and PAOPR04
cells were also grown in King’s B medium without supplements. Cells were grown for 24 hours
with shaking at 37oC till they reached a similar optical density at 600 nm. Extraction of
pyoverdin obtained from a 1 ml supernatant is described under Materials and Methods.
Pyoverdin levels are reported as the absorbance at 405 nm over 600 nm. Samples were grown
individually three times and the error bars reported.
68
The production of pyoverdin was also tested using Chrome Azurol S (CAS) plates
containing an iron-dye complex (Fig. 26). Pyoverdin is a siderophore that binds to the iron and
releases the dye, resulting in an orange halo. PAOPR04 was unable to grow in CAS plates
without supplement because CAS plates act as minimal medium, but PAO1 showed growth.
However when supplied with uracil or orotate independently, PAOPR04 was able to grow and
release the dye, which is seen as an orange halo around the growth of the bacterium. When CAS
plates were incubated for 24 hours, the halo around the mutant strain PAOPR04 was 25% of that
of the wild type PAO1 level (Fig. 26 A). The plates were then incubated for another 24 hours for
a total of 48 hours to see if the mutant strain came up to the wild type level in producing the
orange halo. After 48 hours, PAOPR04 produced about half of that seen for PAO1 (Fig. 26 B).
The plates were then incubated at room temperature over a week, but the halo produced by the
mutant never approached the wild type level.
Casein protease and Elastase analysis:
Kessler et al. (1993) showed that some genes produce certain enzymes capable of
enhancing the virulence in P. aeruginosa. Casein protease and elastase are two such enzymes
that were tested in the wild type and the mutant strains. PTSB medium was used to grow the
cells and to further test for the enzymes casein protease and elastase. When no supplement was
provided in this rich medium, a decrease of about 40% in casein protease level was observed in
the mutant strain. Exogenously supplied uracil or orotate independently brought about 10%
decrease each in casein protease level in the mutant when compared with the wild type strain
(Fig. 27). The results obtained for elastase were very similar to that of casein protease. When not
supplemented with uracil or orotate there was a 40% decrease in the mutant strain, whereas
69
A.
B.
Fig. 26. Growth of PAO1 and pyrD- strains on CAS medium with 0.2% glucose as carbon and
energy source. Plate (a) was supplied only with 0.2% glucose, plate (b) was supplied with 0.2%
glucose and 40 µg/ml uracil, plate (c) was supplied with 0.2% glucose and 40 µg/ml orotate. A
minute portion of a single colony was touched with a sterile toothpick and transferred on the
medium.
Fig. 25 A. Result obtained after 24 hours incubation of plates at 37oC
Fig. 25 B. Result obtained after 48 hours incubation of plates at 37oC.
* pyrD- signifies PAOPR04 for convenience in labeling.
70
0
0.5
1
1.5
2
2.5
3
PAO1 PAOPR04
Abs 4
00nm
No supplement
U40ug/ml
O40ug/ml
Fig. 27. Quantitation of casein protease for PAO1 and PAOPR04 cells. Cells were cultivated in
PTSB medium in the presence of uracil or orotate at a concentration of 40 µg/ml for 24 hours at
37oC with shaking, until the strains reached the same optical density at 600 nm. The PAO1 and
PAOPR04 cells were also grown in PTSB medium without supplements. Measurement of casein
protease at A400 from the supernatant is described under Materials and Methods. Samples were
tested individually three times and error bars reported.
71
providing uracil or orotate independently brought only 11% decrease for each in the mutant
strain (Fig. 28).
Production of hemolysin:
Hemolysin is the enzyme that causes the breakdown of red blood cells, which is visible as
hemolysis on a blood agar plate (BAP). Hemolysis can be categorized as α (partial), β (complete)
or γ (none). Organisms capable of causing β hemolysis are mostly pathogenic. PAOPR04 strain
was compared with the wild type to see the extent of hemolysis being produced in the mutant.
The wild type showed clear β hemolysis on the plate with uracil after 24 hours, whereas the
mutant showed no sign of any kind of hemolysis (γ). After 48 hours the mutant still showed no
hemolysis on the plate with no supplement. About 50% β hemolysis was observed on the uracil
supplemented plate and about 25% β hemolysis was observed on the orotate supplemented plate
for the mutant. The wild type cells showed very clear pattern of β hemolysis on all the blood agar
plates (Fig. 29).
Motility studies:
The ability of P. aeruginosa to be opportunistic and cause infections by producing
virulence factors would not be possible without the adherence factor. Swimming, swarming, and
twitching motilities was tested on PAO1 and PAOPR04 strains to see if there was any significant
difference in the mutant when compared to the wild type. Swimming motility requires flagella,
while swarming and twitching motility require type IV pili.
Swimming motility was completely impaired in the mutant when tested after 14 hours,
and it did not improve even after 24 hours. The wild type strain exhibited free swimming ability
72
0
1
2
3
4
5
6
7
8
9
10
PAO1 PAOPR04
Abs 4
95nm
No supplement
U40ug/ml
O40ug/ml
Fig. 28. The Elastase Congo Red assay for the measurement of elastase produced by PAO1 and
PAOPR04 cells. Cells were cultivated in PTSB medium in the presence of uracil or orotate at a
concentration of 40 µg/ml for 24 hours at 37oC with shaking, until the strains reached the same
optical density at 600 nm. PAO1 and PAOPR04 cells were also grown in PTSB medium without
supplements. Measurement of the enzyme elastase at A495 from the supernatant is described
under Materials and Methods. Samples were tested individually three times and error bars
reported.
73
A.
B.
Fig. 29. Production of hemolysin by PAO1 and pyrD- strains on blood agar plates (Tryptic soy
agar medium supplied with 5% sheep’s blood). Plate (a) was not supplied with any exogenous
nutrient, plate (b) was supplied with 40 µg/ml uracil, plate (c) was supplied with 40 µg/ml
orotate. A minute portion of a single colony was touched with a sterile toothpick and transferred
on the medium.
Fig. 28 A. Result obtained after 24 hours incubation of plates at 37oC.
Fig. 28 B. Result obtained after 48 hours incubation of plates at 37oC.
* pyrD- signifies PAOPR04 for convenience in labeling.
74
after 14 hours (Fig. 30 A) and more so after 24 hours (Fig. 30 B). The swimming pattern seemed
restricted for wild type PAO1 on plates that were supplemented with uracil. Swimming plates
that contained no supplement or supplemented with orotate exhibited free flow swimming
movements.
Swarming motility results were different from swimming. Swarming was totally impaired
for PAOPR04 in plates that had no supplement or an orotate supplement after 14 and 24 hours.
The PAOPR04 strain showed the ability to swarm in the plate that was supplemented with uracil.
After 14 hours, the mutant was almost 30% less than the wild type (Fig. 31 A), and was up to the
wild type swarming capacity after 24 hours (Fig. 31 B).
Twitching motility was observed by peeling off the agar after the growth of bacteria and
staining the empty Petri dish to find the zone of attachment on the surface of the plate. Observing
the plates for bacterial growth suggests that the growth of the mutant is less on plates with no
supplement or when supplied with orotate, whereas growth is the same as the wild type on the
plate supplied with uracil (Fig. 32 A). The results obtained after peeling the agar and staining
them with crystal showed that the mutant strain was about 15% less than the wild type after 24
hours (Fig. 32 B).
Rhamnolipid is a biosurfactant which is involved in swarming motility (Kohler et al.,
2000). Production of rhamnolipid was seen in the form of a faint pink halo around the growth of
the bacterium. The rhamnolipid test was performed on a minimal plate and the mutant failed to
grow on a plate with no supplement. After 24 hours, the mutant was about 40% less than that of
the wild type on the plate with uracil, whereas with orotate the level of rhamnolipid
75
A.
B.
Fig. 30. Swimming motility of PAO1 and pyrD- strains on swim medium supplied with 0.2%
glucose as carbon and energy source. Plate (a) was supplied only with 0.2% glucose, plate (b)
was supplied with 0.2% glucose and 40 µg/ml uracil, plate (c) was supplied with 0.2% glucose
and 40 µg/ml orotate. A minute portion of a single colony was touched with a sterile toothpick
and transferred on the medium.
Fig. 29 A. Result obtained after 14 hours incubation of plates at 30oC.
Fig. 29 B. Result obtained after 24 hours incubation of plates at 30oC.
* pyrD- signifies PAOPR04 for convenience in labeling.
76
A.
B.
Fig. 31. Swarming motility of PAO1 and pyrD- strains on swarm medium supplied with 0.2%
glucose as carbon and energy source. Plate (a) was supplied only with 0.2% glucose, plate (b)
was supplied with 0.2% glucose and 40 µg/ml uracil, plate (c) was supplied with 0.2% glucose
and 40 µg/ml orotate. A minute portion of a single colony was touched with a sterile toothpick
and transferred on the medium.
Fig. 30 A. Result obtained after 14 hours incubation of plates at 37oC.
Fig. 30 B. Result obtained after 24 hours incubation of plates at 37oC.
* pyrD- signifies PAOPR04 for convenience in labeling.
77
A.
B.
Fig. 32. Twitching motility of PAO1 and pyrD- strains on twitch medium supplied with 0.2%
glucose as carbon and energy source. Plate (a) was supplied only with 0.2% glucose, plate (b)
was supplied with 0.2% glucose and 40 µg/ml uracil, plate (c) was supplied with 0.2% glucose
and 40 µg/ml orotate. A minute portion of a single colony was touched with a sterile toothpick
and transferred on the medium.
Fig. 31 A. Result obtained after 24 hours incubation of plates at 37oC.
Fig. 31 B. Result obtained after 24 hours incubation of plates at 37oC., peeled and stained with
crystal violet.
* pyrD- signifies PAOPR04 for convenience in labeling.
78
production was not visible (Fig. 33 A). However after 48 hours the halo produced by the
mutant on the uracil supplemented plate was about half that of the wild type. The orotate
supplemented plate showed about 75% less production in the mutant strain. (Fig. 33 B).
79
A.
B.
Fig. 33. Rhamnolipid production of PAO1 and pyrD- strains on the medium supplied with 0.2%
glucose as carbon and energy source. Plate (a) was supplied only with 0.2% glucose, plate (b)
was supplied with 0.2% glucose and 40 µg/ml uracil, plate (c) was supplied with 0.2% glucose
and 40 µg/ml orotate. A minute portion of a single colony was touched with a sterile toothpick
and transferred on the medium.
Fig. 32 A. Result obtained after 24 hours incubation of plates at 37oC.
Fig. 32 B. Result obtained after 48 hours incubation of plates at 37oC.
* pyrD- signifies PAOPR04 for convenience in labeling.
80
CHAPTER IV
DISCUSSION
This study of regulation of the pyrimidine pathway was initiated by studying mutants that
were blocked in the pyrimidine pathway. Isaac and Holloway in 1968 previously studied various
mutants of the pyrimidine pathway generated by chemical mutagenesis, but they never
succeeded in isolating a mutant blocked in pyrC encoding DHOase. Dayna Brichta in our lab
sought to rectify this problem and cloned the DHOase gene and created a double knockout
(DKO) after she discovered that P. aeruginosa has two active pyrC genes located on separate
regions of the chromosome (Brichta, 2003) each with an active DHOase as illustrated in Fig. 34.
Not only was the DKO a pyrimidine auxotroph, but it established a novel link between the
pyrimidine pathway and virulence. Brichta (2003) discovered a decrease in various exoproducts
that are responsible for the virulence of P. aeruginosa. She attributed this result to the build up of
carbamoyl aspartate (CAA) thereby increasing the negative charge within the cell and disturbing
the balance of ions. This hypothesis of Brichta was tested by creating a mutant that would cause
no buildup of CAA. A knockout of the second step of the pyrimidine pathway resulted in the
inactivation of the enzyme ATCase, which caused no buildup of CAA (Hammerstein, 2004).
Inactivation of the pyrB gene like the DKO also caused a decrease in the exoproducts found for
the DKO mutant. Hammerstein then proposed that the virulence factor production was affected
in the DKO due to a buildup of CAA and in pyrB- due to the buildup of carbamoyl phosphate
(CP). The increase in negative charge of CAA or CP was assumed to disrupt the energetics of the
cell, thereby decreasing the virulence. No such link between pyrimidine auxotrophy and
virulence factor production had ever been reported for P. aeruginosa.
81
Fig. 34. The chromosomal map of P. aeruginosa with the location of the pyrD gene that encodes
dihydroorotate dehydrogenase and the two active pyrC genes (C and C2) that encode
dihydroorotate are shown. This information was obtained from www.pseudomonas.com
Pseudomonas aeruginosa
PAO1
6264403 bp
pyrC2
pyrC
pyrD
82
Our next step was to determine if all pyrimidine mutants behave in a similar fashion in
showing decreased virulence. We therefore created a block in pyrD encoding DHOdehase to
cause a buildup of DHO and prevents the production of orotate (OA). The pyrD mutant,
PAOPR04 was constructed by cloning as described in Materials and Methods. The pyrD mutant
exhibits a pleiotropic phenotype, which was unable to grow on PsMM with just glucose as the
carbon and energy source, unless an exogenous pyrimidine supplement, such as uracil or orotate
was provided. The mutant showed no sign of growth defect in PsMM with glucose and uracil or
orotate.
The effect of the pyrD mutant on the pyrimidine pathway was tested by measuring the
specific activity of the pyrimidine enzymes. Isaac and Holloway (1968) reported that providing
uracil to P. aeruginosa did not cause repression and starving a mutant for uracil did not bring
about derepression. I observed repression in PAO1 for ATCase, OPRTase and OMPdecase and
derepression of DHOase when cells were grown in the presence of uracil. Repression of ATCase
and OPRTase and derepression of DHOase and OMPdecase were observed for uracil grown cells
of the mutant PAOPR04 in comparison to uracil grown cells of wild type PAO1. A similar
pattern of repression and derepression was previously reported in P. oleovorans ATCC 8062
(Lee and Chandler, 1941) when cultured with glucose as carbon and energy source and uracil
addition (Haugaard and West, 2001). When orotate rather than uracil was used as the
supplement, all the pyrimidine enzymes showed increased activity in PAO1 and PAOPR04 with
DHOase being the highest amongst them at 5-fold for PAO1 and 3-fold for PAOPR04. The C4-
dicarboxylate transport protein, DctA is responsible for the entry of succinate and also mediates
the entry of orotate into the cells (Baker et al., 1996; Yurgel et al., 2000). Pseudomonas
aeruginosa mutants blocked prior to pyrD are unable to grow with succinate as a carbon and
83
energy source and orotate as the pyrimidine source. This is so because succinate outcompetes
orotate for entry into the cell (Yurgel et al., 2000). Other mutant species of Pseudomonas are
capable of growth with succinate and orotate such as P. oleovorans (Haugaard and West, 2001),
P. reptilivora (West, 2003), P. alkanolytica (West, 2004), and P. resinovorans (West, 2005).
Uracil is converted to UMP via UPRTase in all pseudomonads (Neuhard and Kelln, 1996).
Likewise, in P. aeruginosa the only way uracil is converted to UMP is via the enzyme UPRTase.
UMP degradation product was shown to be responsible for the increased enzyme levels of
ATCase and DHOase in P. oleovorans (Haugaard and West, 2002). A similar situation could be
possible in P. aeruginosa where UMP degradation product(s) could be responsible for high
levels of enzyme activity for DHOase under conditions of excess uracil. Providing excess orotate
increased the specific activity of all the pyrimidine biosynthetic enzymes. A possible explanation
for this is that orotate is converted to OMP first and then to UMP, increasing the specific
activities of their respective enzymes. UMP degradation releases a product that acts as an inducer
thereby bringing an increase in the activities of ATCase, DHOase and DHOdehase (not present
in the pyrD mutant).
Pyrimidine starvation experiments are typically carried out to identify repression or
derepression in pyrimidine auxotrophs. To this end, the pyrD mutant was starved for uracil or
orotate. Starving a pyrimidine auxotroph for uracil has shown depression of the de novo
pyrimidine pathway enzymes (West, 1994; Haugaard and West, 2002). Derepression was
observed for ATCase (3-fold), OPRTase (2-fold) and OMPdecase (3-fold) after 2 hours of uracil
starvation. When starved for orotate 1.2-fold increase of ATCase and 1.3-fold increase of
OMPdecase was observed after 2 hours. Dihydroorotate dehydrogenase showed 2 and 3-fold
repression when starved for uracil or orotate respectively after 2 hours. A uracil auxotroph of P.
84
alkanolytica PT112 grown with glucose as a carbon and energy source and starved for uracil
showed a 5.4-fold repression after 1 hour and a 5.7-fold derepression after 2 hours of starvation
(West, 2004). It is possible that the pyrD mutant of P. aeruginosa may be derepressed when
starved for uracil or orotate independently for longer than 2 hours. Pyrimidine ribonucleotide
pools are known to drop when cells are starved for uracil (West et al., 1983). Significant change
in derepression of biosynthetic pyrimidine enzymes indicates reduced cellular pyrimidine
ribonucleotide levels indicating transcriptional regulation of pyrimidine enzymes by their
ribonucleotides (Haugaard and West, 2002). In this study pyrimidine starvation experiments
have demonstrated that the pyrimidine biosynthetic enzymes can be derepressed and repression
seen in the case of pyrimidine-supplemented compounds indicates that regulation is occurring at
the transcriptional level.
Carbon starvation has a major influence on amino acid synthesis and degradation in P.
aeruginosa (Kay and Gronlund, 1969). The decrease of certain amino acids to a constitutive
basal level of degradative enzymes was reported due to the depletion of the endogenous amino
acid pool thus limiting the exogenous supply of carbon. In E. coli, carbon starvation brought
about a decrease in amino acid transport (Britten and McClure, 1962).
The influence of carbon (glucose) starvation on pyrimidine enzymes was tested on PAO1
cells grown in PsMM in our lab. The most striking feature that was observed with carbon
starvation of PAO1 was finding zero activity for DHOase and a four-fold increase of
DHOdehase activity when compared with PAO1 cells grown under normal conditions with 0.2%
glucose. The reason for this is unknown, but perhaps the pyrC, pyrC2 genes are affected here and
may play a role in amino acid transport. Further studies need to be carried out by starving the
wild type cells for other carbon sources such as succinate or glycerol.
85
In a related study a block in the genes that encode CPSase, ATCase and DHOase showed
no synthesis of the pigment pyoverdin in Pseudomonas putida M, whereas mutations in the
genes that encode DHOdehase and OPRTase had no affect on the formation of pyoverdin
(Maksimova et al., 1993). The reasoning behind this result was that dihydroorotate (DHO) was
thought to be a precursor of pyoverdin (Maksimova et al., 1993) in Pseudomonas putida which
produces no pyocyanin. A block in the pyrB gene encoding ATCase and pyrC, pyrC2 encoding
DHOase in P. aeruginosa showed diminished levels of pyoverdin and pyocyanin. A block in the
pyrD gene encoding DHOdehase showed results similar to what was observed in P. putida. The
pyoverdin level was almost the same as wild type PAO1. The pyocyanin pigment level also was
up to the level of the wild type P. aeruginosa. Further studies to test the pyoverdin level in pyrE
and pyrF mutants of P. aeruginosa are underway. If these mutants show no impairment of
pyoverdin production, it may suggest that DHO is be a precursor for pyoverdin, even in P.
aeruginosa.
The pyrD mutant was tested for various secondary metabolites produced and compared
with wild type P. aeruginosa and also with the pyrB and pyrC, pyrC2 double mutant. When
compared to wild type, the levels produced were significantly lower in casein protease, elastase,
and hemolysin production in the pyrD mutant. Motility studies showed impaired movement in
swimming, swarming and twitching. Rhamnolipid production was reduced and so was the iron
gathering capability on CAS plates. Table 6 compares the effects of different mutations in the
pyrimidine genes studied so far in our laboratory.
The proton motive force drives various systems within a bacterial cell such as the twin
arginine translocase (TAT) apparatus (Chaddock et al., 1995) and the Mex transport system
(Paulsen et al., 1996). Proper functioning of such transport apparatus within the cell requires the
86
Table 6. Comparison of various virulence factors between the pyrimidine mutants blocked at the
second, third and fourth steps of the biosynthetic pathway with wild type P. aeruginosa. pyrE
and pyrF mutants are currently being studied.
Virulence PAO1 pyrD pyrC, pyrC2 pyrB
factors
Pyocyanin + + - -
Pyoverdin + + - -
Casein protease + - - -
Elastase + - - -
Hemolysin + - - -
Swimming + - - -
Swarming + - - -
Twitching + - - -
Rhamnolipid + - - -
87
ions to be in equilibrium. We hypothesize that a build up of negative ions within the cell in a
pyrC, pyrC2 mutant (CAA) and pyrB mutant (CP) may have been the reason behind decreased
virulence. DHO has one negative charge and a build up of DHO could also disturb the
equilibrium of ions, thereby causing an effect at the enzyme level of pyrimidines, secondary
metabolites and motility. Also, DHO can be converted to CAA via a reversible reaction (Fig. 3),
thereby increasing the negative charges within the cell, a situation similar to that of the DHOase
double mutant.
In conclusion, the pyrD mutant PAOPR04 showed no growth defect when grown in
PsMM with pyrimidine supplements (uracil or orotic acid). The growth rate of the mutant was
similar to that of the wild type in minimal and rich medium. The production of the pigments
pyocyanin and pyoverdin was not hampered and the levels were at wild type strain levels. The
exoproducts (casein protease, elastase, and hemolysin) were produced at a diminished level and
motility (swimming, swarming, and twitching) was impaired. The level of the biosurfactant
rhamnolipid was also diminished in the mutant, as was the iron binding capacity. Further
research needs to be carried out to test the antibiotic susceptibility of the pyrD mutant and a role
that the mutant might have to play in the elimination of the infection caused by P. aeruginosa in
immunocompromised patients.
This laboratory is currently creating mutants with defects in the pyrE and pyrF genes of
the pyrimidine pathway to understand their effects on the pathway enzymes and virulence. A
purine auxotroph and an amino acid auxotroph are being studied to determine if the results
obtained from the mutants of the pyrimidine pathway pertained solely to pyrimidines or to other
auxotrophs as well. The net result of this body of work corroborates the finding of Brichta
(Brichta, 2003) that couple pyrimidine auxotrophy with virulence production.
88
REFERENCES
Abdelal, A.T., Bussey, L., Vickers, L., 1983. Carbamoylphosphate synthetase from
Pseudomonas aeruginosa: subunit composition, kinetic analysis and regulation. Eur J Biochem.
129, 697-702.
Albus, A.M., Pesci, E.C., Runyen-Janecky, L.J., West, S.E., and Iglewski, B.H., 1997. Vfr
controls quorum-sensing in Pseudomonas aeruginosa. J Bacteriol. 179, 3928-35.
Anderson, P.M., 1983. CTP synthetase from Escherichia coli: an improved purification
procedure and characterization of hysteretic and enzyme concentration effects on kinetic
properties. Biochem. 22, 3285-92.
Andersen, P.S., Jansen, P.J., and Hammer, K., 1994. Two different dihydroorotate
dehydrogenases in Lactococcus lactis. J Bacteriol. 176, 3975-82.
Andersen, P.S., Martinussen, J., and Hammer, K., 1996. Sequence analysis and identification
of the pyrKDbF operon from Lactococcus lactis including a novel gene, pyrK, involved in
pyrimidine biosynthesis. J Bacteriol. 178, 5005-12.
Anderson, P.M., and Meister, A., 1965. Evidence for an activated form of carbon dioxide in
the reaction catalyzed by Escherichia coli carbamyl phosphate synthetase. Biochem. 4, 2803-9.
89
Andrews, S., Cox, G.B., and Gibson, F., 1977. The anaerobic oxidation of dihydroorotate by
Escherichia coli K-12. Biochim Biophys Acta. 462, 153-60.
Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Smith, J.A., and Struhl, K., 1997.
Short protocols in molecular biology. 3rd ed. John Wiley & Sons Inc.
Azad, K.N., 2005. Pyrimidine enzyme specific activity at four different phases of growth in
minimal and rich media, and concomitant evaluation of virulence factor production in
Pseudomonas aeruginosa. Ph.D. Dissertation. University of North Texas, Denton, Texas.
Baker, K.E., Dutillo, K.P., Neuhard, J., and Kelln, R.A., 1996. Utilization f orotate as a
pyrimidine source by Salmonella typhimurium and Escherichia coli requires the dicarboxylate
transport protein encoded by dctA. J Bacteriol. 178, 7099-105.
Beck, D.A., 1995. The pyrimidine salvage pathway in prokaryotes: Labyrinths of enzymatic
diversity. Ph.D. Dissertation. University of North Texas, Denton, Texas.
Beckwith, J.R., Pardee, A.B., Austrian, R., and Jacob, F., 1962. Coordination of the synthesis
of enzymes in the pyrimidine pathway of E. coli. J Mol Biol. 5, 618-34.
Björnberg, O., Grüner, A.C., Roepstorff, P., and Jensen, K.F., 1999. The activity of
Escherichia coli dihydroorotate dehydrogenase is dependent on a conserved loop identified by
sequence homology, mutagenesis, and limited proteolysis. Biochem. 38, 2899-908.
90
Björnberg, O., Rowland, P., Larsen, S., and Jensen, K.F., 1997. Active site of dihydroorotate
dehydrogenase A from Lactococcus lactis investigated by chemical modification and
mutagenesis. Biochem. 36, 16197-205.
Boucher, R.C., 2004. New concepts of the pathogenesis of cystic fibrosis lung disease. Eur
Respir J. 23, 146-58.
Bradford, M.M., 1976. A rapid and sensitive method for the quantitation of microgram
quantities of protein utilizing the principle of dye-binding. Anal Biochem. 72, 249-54.
Brichta, D.M., 2003. Construction of a Pseudomonas aeruginosa dihydroorotase mutant and the
discovery of a novel link between pyrimidine biosynthetic intermediates and the ability to
produce virulence factors. Ph.D. Dissertation. University of North Texas, Denton, Texas.
Brichta, D.M., Azad, K.., Ralli, P., and O’Donovan, G.A., 2004. Pseudomonas aeruginosa
dihydroorotases: a tale of three pyrCs. Arch Microbiol. 182, 7-17.
Brint, J.M., and Ohman, D.E., 1995. Synthesis of multiple exoproducts in Pseudomonas
aeruginosa is under the control of RhlR-RhlI, another set of regulators in strain PAO1 with
homology of the autoinducer-responsive LuxR-LuxI family. J Bacteriol. 177, 7155-63.
91
Britigan, B.E., 1993. Role of reactive oxygen species in Pseudomonas infection. In
Pseudomonas aeruginosa: The opportunist. Pathogenesis and disease. Fick, Jr. R.B. (ed). Boc
Raton; CRC Press, pp. 113-40.
Britten, R.J., and McClure, F.T., 1962. The amino acid pool in Escherichia coli. Bacteriol
Rev. 26, 293-335.
Calfee, M.W., Coleman, J.P., and Pesci, E.C., 2001. Interference of Pseudomonas quinolone
signal synthesis inhibits virulence factor expression by Pseudomonas aeruginosa. Proc Natl
Acad Sci U.S.A. 98, 11633-7.
Carlson, C.A., 1982. The physiological genetics of denitrifying bacteria. Antonie Van
Leeuwenhoek. 48, 555-67.
Chaddock, A.M., Mant, A., Karnauchov, I., Brink, S., Herrmann, R.G., Klösgen, R.B., and
Robinson, C., 1995. A new type of signal peptide: central role of a twin-arginine motif
in
transfer signals for the delta pH-dependent thylakoidal protein translocase. EMBO J. 14, 2715-
22.
Chu, C.P., and West, T.P., 1990. Pyrimidine biosynthetic pathway of Pseudomonas
flourescens. J Gen Microbiol. 136, 875-80.
92
Craven, R., and Montie, T.C., 1985. Regulation of Pseudomonas aeruginosa: involvement of
methylation. J Bacteriol. 154, 780-6.
Condon, S., Collins, J.K., and O’Donovan, G.A., 1976. Regulation of arginine and pyrimidine
biosynthesis in Pseudomonas putida. J Gen Microbiol. 92, 375-83.
Costerton, J.W., Lewandowski, Z., DeBeer, D., Caldwell, D., Korber, D., and James, G.,
1994. Biofilms, the customized microniche. J Bacteriol. 176, 2137-42.
D’Argenio, D.A., Calfee, M.W., Rainey, P.B., and Pesci, E.C., 2002. Autolysis and
autoaggregation in Pseudomonas aeruginosa colony morphology mutants. J Bacteriol. 184,
6481-9.
Davis, J.P., Cain, G.A., Pitts, W.J., Magolda, R.L., and Copeland, R.A., 1996. The
immunosuppressive metabolite of leflunomide is a potent inhibitor of human dihydroorotate
dehydrogenase. Biochem. 35, 1270-3.
Davis, J.P., and Copeland, R.A., 1997. Histidine to alanine mutants of human dihydroorotate
dehydrogenase. Identification of a brequinar-resistant mutant enzyme. Biochem Pharmacol. 54,
459-65.
93
DeFrees, S.A., Sawick, D.P., Cunningham, B., Heinstein, P.F., Morre, D.J., and Cassady,
J.M., 1988. Structure-activity relationships of pyrimidines as dihydroorotate dehydrogenase
inhibitors. Biochem Pharmacol. 37, 3807-16.
De Kievit, T., Seed, P.C., Nezezon, J., Passador, L., and Iglewski, B.H., 1999. RsaL, a novel
repressor of virulence gene expression in Pseudomonas aeruginosa. J Bacteriol. 181, 2175-84.
Denis-Duphil, M., 1989. Pyrimidine biosynthesis in Saccharomyces cerevisiae: the uraA cluster
gene, its multifunctional enzyme product, and other structural or regulatory genes involved in de
novo UMP synthesis. Biochem Cell Biol. 67, 612-31.
Denning, G.M., Railsback, M.A., Rasmussen, G.T., Cox, C.D., and Britigan, B.E., 1998.
Pseudomonas pyocyanine alters calcium signaling in human airway epithelial cells. Am J
Physiol Lung Cell Mol Physiol. 274, L893-900.
Deretic, V., Chandrasekharappa, S., Gill, J.F., Chatterjee, D.K., and Chakrabarty, A.M.,
1987. A set of cassettes and improved vectors for genetic and biochemical characterization of
Pseudomonas genes. Gene. 57, 61-72.
Deretic, V., Schurr, M.J., and Yu, H., 1995. Pseudomonas aeruginosa, mucoidy and the
chronic infection phenotype in cystic fibrosis. Trends Microbiol. 3, 351-6.
94
Diener, B., Carrick, L. Jr, and Berk, R.S., 1973. In vivo studies with collagenase from
Pseudomonas aeruginosa. Infect Immun. 7, 212-7.
Doremus, H.D., 1986. Organization of the pathway of de novo pyrimidine nucleotide
biosynthesis in pea (Pisum sativum L. cv Progress No. 9) leaves. Arch Biochim Biophys. 20,
112-9.
Doull, I.J., 2001. Recent advances in cystic fibrosis. Arch Dis Childhood. 85, 62-6.
Dunne, J.W.M., 2002. Bacterial adhesion: seen any good biofilms lately? Clin Microbiol Rev.
15, 155-66.
Essar, D.W., Eberly, L., Hadero, A., and Crawford, I.P., 1990. Identification and
characterization of genes for a second anthranilate synthase in Pseudomonas aeruginosa:
interchangeability of the two anthranilate synthases and evolutionary implications. J Bacteriol.
172, 884-90.
Figurski, D.H., and Helinski, D.R., 1979. Replication of an origin-containing derivative of
plasmid RK2 dependent on a plasmid function provided in trans. Proc Natl Acad Sci USA. 76,
1648-52.
95
Foltermann, K.F., Wild, J.R., Zink, D.L., and O’Donovan, G.A., 1981. Regulatory variance
of aspartate transcarbamoylase among strains of Yersenia enterocolitica-like organisms. Curr
Microbiol. 6, 43-7.
Gallagher, L.A., McKnight, S.L., Kuznetsova, M.S., Pesci, E.C., and Manoil, C., 2002.
Functions required for extracellular quinolone signaling by Pseudomonas aeruginosa. J
Bacteriol. 184, 6472-80.
Galloway, D.R., 1991. Pseudomonas aeruginosa elastase and elastolysis revisited: recent
developments. Mol Microbiol. 5, 2315-21.
Gambello, M.J., and Iglewski, B.H., 1991. Cloning and characterization of the Pseudomonas
aeruginosa lasR gene: a transcriptional activator of elastase expression. J Bacteriol. 173, 3000-9.
Gerhart, J.C., and Pardee, A.B., 1962. The enzymology of control by feedback inhibition. J
Biol Chem. 237, 891-6.
Gerhart, J.C., and Pardee, A.B., 1964. Aspartate transcarbamoylase, an enzyme designed for
feedback inhibition. Fed Proc. 23, 727-35.
Ginther, C.L., and Ingraham, J.L., 1974. Nucleotide diphosphokinase of Salmonella
typhimurium. J Biol Chem. 249, 3406-11.
96
Govan, J.R.W., and Deretic, V., 1996. Microbial pathogenesis in cystic fibrosis: mucoid
Pseudomonas aeruginosa and Burkholderia cepacia. Microbiol Rev. 60, 539-74.
Grogan, G.W., and Gonsalus, R.P., 1993. Sulfolobus acidocaldarius synthesizes UMP via a
standard de novo pathway: results of biochemical-genetic study. J Baceriol. 175, 1500-7.
Hammerstein, H.C., 2004. Isolation of a Pseudomonas aeruginosa aspartate transcarbamoylase
mutant and the investigation of its growth characteristics, pyrimidine biosynthetic enzyme
activities, and virulence factor production. M.S. Thesis. University of North Texas, Denton,
Texas.
Hassett, D.J., Charniga, L., Bean, K., Ohman, D.E., and Cohen, M.S., 1992. Response of
Pseudomonas aeruginosa to pyocyanin: mechanisms of resistance, antioxidant defenses, and
demonstration of a manganese-cofactored superoxide dismutase. Infect Immun. 60, 328-36.
Haugaard, L.E., and West, T.P., 2002. Pyrimidine biosynthesis in Pseudomonas oleovorans. J
Appl Microbiol. 92, 517-25.
Heck, L.W., Alarcon, P.G., Kulhavy, R.M., Morihara, K., Russell, M.W., and Mestecky,
J.F., 1990. Degradation of IgA proteins by Pseudomonas aeruginosa elastase. J Immunol. 6,
2253-7.
Heck, L.W., Morihara, K., McRae, W.B., and Miller, E.J., 1986 Specific cleavage of human
type III and IV collagens by Pseudomonas aeruginosa elastase. Infect Immun. 51, 115-8.
97
Hong, Y., and Ghebrehiwet, B., 1992. Effect of Pseudomonas aeruginosa elastase and alkaline
protease on serum complement and isolated components C1q and C3. Clin Immunol
Immunopathol. 62, 133-8.
Iglewski, B.H., Sadoff, J., Bjorn, M.J., and Maxwell, E.S., 1978. Pseudomonas aeruginosa
exoenzyme S: an adenosine diphosphate ribosyltransferase distinct from toxin A. Proc Natl Acad
Sci U.S.A. 75, 3211-5.
Isaac, J.H., and Holloway, B.W., 1968. Control of pyrimidine biosynthesis in Pseudomonas
aeruginosa. J Bacteriol. 96, 1732-41.
Jacquot, J., Tournier, J., and Puchelle, E., 1985. In vitro evidence that human airway
lysozyme is cleaved and inactivated by Pseudomonas aeruginosa elastase and not by human
leukocyte elastase. Infect Immun. 47, 555-60.
Johnson, D.A., Carter-Hamm, B., and Dralle, W.M., 1982. Inactivation of human bronchial
mucous proteinase inhibitor by Pseudomonas aeruginosa elastase. Am Rev Respir Dis. 126,
1070-3.
Jones, M.E., 1980. The genes for and regulation of enzyme activities of to multifunctional
proteins required for the de novo pathway for UMP biosynthesis in mammals. Mol Bio Biochem
Biophys. 32, 165-82.
Jones, M.E., 1980. Pyrimidine nucleotide biosynthesis in animals: genes, enzymes and
regulation of UMP biosynthesis. Ann Rev Biochem. 49, 253-79.
98
Jones, M., Spector, L., and Lipmann, F., 1955. Carbamoyl phosphate, the carbamoyl donor in
enzymatic citrulline synthesis. J Am Chem Soc. 77, 819-20.
Kay, W.W., and Gronlund, A.F., 1969. Influence of carbon or nitrogen starvation on amino
acid transport in Pseudomonas aeruginosa. J Bacteriol. 100, 276-82.
Karibian, D., and Couchoud, P., 1974. Dihydro-orotate oxidase of Escherichia coli K12:
purification, properties and relation to the cytoplasmic membrane. Biochem Biophys Acta. 364,
218-32.
Kerr, C.T., and Miller, R.W., 1968. Dihydrorotate-ubiquinone reductase complex of
Escherichia coli B. J Biol Chem. 243, 2963-8.
Kessler, E., Safrin, M., Olson, J.C., and Ohman, D.E., 1993. Secreted LasA of Pseudomonas
aeruginosa is a staphylolytic protease. J Biol Chem. 268, 7503-8.
King, E.O., Ward, M.K., and Raney, D.E., 1954. Two simple media for demonstration of
pyocyanin and flourescin. J Lab Clin Med. 44, 301-7.
Kloepper, J.W., Leong, J., Teintza, M., and Schroth, M.N., 1980. Enhanced plant- growth by
siderophores produced by plant growth-promoting Rhizobacteria. Nature. 286, 885-6.
99
Köhler, T., Curty, L.K., Barja, F., Van Delden, C., and Pechère, J.C., 2000. Swarming of
Pseudomonas aeruginosa is dependent on cell-to-cell signaling and requires flagella and pili. J
Bacteriol. 182, 5990-6.
Kurachi, M., 1958. Studies of the biosynthesis of pyocyanine (II). Bull Inst Chem Res., Kyoto
University. 36, 174-87.
Lam, J., Chan, R., Lam, K., and Costerton, J.R.W., 1980. Production of mucoid
microcolonies by Pseudomonas aeruginosa within infected lungs in cystic fibrosis. Infect
Immun. 28, 546-56.
Larsen, J.N., and Jensen, K.F., 1985. Nucleotide sequence of the pyrD gene of Escherichia
coli and characterization of the flavoprotein dihydroorotate dehydrogenase. Eur J Biochem. 151,
59-65.
Latifi, A., Foglino, M., Tanaka, K., Williams, P., and Lazdunski, A., 1996. A hierarchical
quorum-sensing cascade in Pseudomonas aeruginosa links the transcriptional activators LasR
and RhlR to expression of the stationary-phase sigma factor RpoS. Mol Microbiol. 21, 1137-46.
Latifi, A., Winson, M.K., Foglino, M., Bycroft, B.W., Stewart, G.S.A.B., Lazdunski, A., and
Williams, P., 1995. Multiple homologues of LuxR and LuxI control expression of virulence
determinants and secondary metabolites through quorum sensing in Pseudomonas aeruginosa
PAO1. Mol Microbiol. 17, 333-43.
Lee, M., and Chandler, A.C., 1941. A study of the nature, growth and control of bacteria in
cutting compounds. J Bacteriol. 41, 373-86.
100
Liebermann, I., Kornberg, A., 1954. Enzymatic synthesis and breakdown of a pyrimidine,
orotic acid. II. Dihydroorotic acid, ureidosuccinic acid and 5-carboxymethyl hydantoin. J Biol
Chem. 207, 911-24.
Linker, A., and Jones, R.S., 1966. A new polysaccharide resembling alginic acid isolated from
Pseudomonads. J Biol Chem 241, 3845-51.
Liu, P.V., 1974. Extracellular toxins of Pseudomonas aeruginosa. J Infect Dis. 130, 94-9.
Liu, S., Neidhardt, E.A., Grossman, T.H., Ocain, T., and Clardy, J., 2000. Structures of
human dihydroorotate dehydrogenase in complex with antiproliferative agents. Struture Fold
Des. 8, 25-33.
Löffler, M., Grein, K., Knecht, W., Klein, A., and Bergjohann, U., 1998. Dihydroorotate
dehydrogenase. Profile of a novel target for antiproliferative and immunosuppressive drugs. Adv
Exp Med Biol. 431, 507-13.
Lomholt, J.A., Poulsen, K. and Kilian, M., 2001. Epidemic population structure of
Pseudomonas aeruginosa: evidence for a clone that is pathogenic to the eye and that has a
distinct combination of virulence factors. Infect Immun. 69, 6284-95.
Long, C., and Koshland, D.E.Jr., 1978. Cytidine triphosphate synthetase. Materials and
Methods Enzymol. 51, 79-83.
101
Long, C.W., and Pardee, A.B., 1967. Cytidine triphosphate synthetase of Escherichia coli B. I.
Purification and kinetics. J Biol Chem. 242, 4715-21.
MacDonald, J.C., 1966. Production of pyocyanin by Pseudomonas aeruginosa ATCC 9027.
Can J Microbiol. 12, 771-4.
Mah, T.C., and O'Toole, G.A., 2001. Mechanisms of biofilm resistance to antimicrobial agents.
Trends Microbiol. 9, 34-9.
Mahajan-Miklos, S., Tan, M.W., Rahme, L.G., and Ausubel, F.M., 1999. Molecular
mechanisms of bacterial virulence elucidated using a Pseudomonas aeruginosa-Caenorhabditis
elegans pathogenesis model. Cell. 96, 47-56.
Maksimova, N.P., Blazhevich, O.V., and Fomichev, Yu.K., 1993. The role of pyrimidines in
the biosynthesis of fluorescent pigment pyoverdin Pm in Pseudomonas putida M bacteria.
Molekul Gen, Mikrobiol i Virusol. 5, 22-6.
McKnight, S.L., Iglewski, B.H., and Pesci, E.C., 2000. The Pseudomonas quinolone signal
regulates rhlI quorum sensing in Pseudomonas aeruginosa. J Bacteriol. 182, 2702-8.
Meyer, J.M., Neely, A., Stinzi, A., Georges, C., and Holder, I.A., 1995. Pyoverdin is essential
for virulence of Pseudomonas aeruginosa. Infect Immun. 64, 518-23.
102
Miller, J.H., 1972. Experiments in Molecular Genetics. Cold Spring Harbor, NY: Cold Spring
Harbor Laboratory Press.
Misaghi, I.J., Stowell, L.J., Grogan, R.G., and Spearman, L.C., 1982. Fuhgistatic activity of
water-soluble florescent pigments of fluorescent Pseudomonads. Phytopathol. 72, 33-6.
Morihara, K., Tsuzuki, H., and Oda, K., 1979. Protease and elastase of Pseudomonas
aeruginosa: inactivation of human a 1-proteinase inhibitor. Infect Immun. 24, 188-93.
Navre, M., and Schachmann, H.K., 1983. Synthesis of aspartate transcarbamoylase in
Escherichia coli: Transcriptional regulation of the pyrBI operon. Proc Natl Acad Sci U.S.A. 80,
1207-11.
Neilands, J.B., 1981. Iron absorption and transport in microorganisms. Annu Rev Nutr. 1, 21-46.
Nielsen, F.S., Andersen, P.S., and Jensen, K.F., 1996. The B form of dihydroorotate
dehydrogenase from Lactococcus lactis consists of two different subunits, encoded by the pyrDb
and pyrK genes, and contains FMN, FAD, and [FeS] redox centers. J Biol Chem. 271, 29359-65.
Neuhard, J., and Kelln, R., 1996. Biosynthesis and conversion of pyrimidines. In F.C
Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Linn, K. B. Low, B. Magasanik, W. S.
Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella:
Cell and Mol Biol., 2nd ed. Am Soc for Microbiol. pp. 580-99.
103
Nicas, T.I., Bradley, J., Lochner, J.E., and Iglewski, B.H., 1985. The role of exoenzyme S in
infections with Pseudomonas aeruginosa. J Infect Dis. 152, 716-21.
Nicas, T.I., Frank, D.W., Stenzel, P., Lile, J.D, and Iglewski, B.H., 1985. Role of exoenzyme
S in chronic Pseudomonas aeruginosa lung infections. Eur J Clin Microbiol. 4, 175-9.
Nicas, T.I., and Iglewski B.H., 1985. The contribution of exoproducts to virulence of
Pseudomonas aeruginosa. Can J Microbiol. 31, 87-92.
Nielsen, F.S., Andersen, P.S., and Jensen, K.F., 1996. The B form of dihydroorotate
dehydrogenase from Lactococcus lactis consists of two different subunits, encoded by the pyrDb
and pyrK genes, and contains FMN, FAD, and [FeS] redox centers. J Biol Chem. 271, 29359-65.
Nielsen, F.S., Rowland, P., Larsen, S., and Jensen, K.F., 1996. Purification and
characterization of dihydroorotate dehydrogenase A from Lactococcus lactis, crystallization and
preliminary X-ray diffraction studies of the enzyme. Protein Sci. 5, 852-856.
Noranger, S., Jensen, K.F., Björnberg, O., and Larsen, S., 2002. E. coli dihydroorotate
dehydrogenase reveals structural and functional distinctions between different classes of
dihydroorotate dehydrogenases. Structure. 10, 1211-23.
Ochsner, U.A., Koch, A.K., Fiechter, A. and Rierser, J., 1994. Isolation and characterization
of a regulatory gene affecting rhamnolipid biosurfactant synthesis in Pseudomonas aeruginosa. J
Bacteriol. 176, 2044-54.
Ochsner, U.A., and Reiser, J., 1995. Autoinducer-mediated regulation of rhamnolipid
biosurfactant synthesis in Pseudomonas aeruginosa. Proc Natl Acad Sci U.S.A. 92, 6424-8.
104
O’Donovan, G.A., and Gerhart, J.C., 1972. Isolation and partial characterization of regulatory
mutants of the pyrimidine pathway in Salmonella typhimurium. J Bacteriol. 109, 1085-96.
O’Donovan, G.A., and Neuhard, J., 1970. Pyrimidine metabolism in microorganisms.
Bacteriol Rev. 34, 278-343.
O’Donovan, G.A., and Shanley, M.S., 1995. Pyrimidine metabolism in Pseudomonas. Paths to
pyrimidines. 3, 49-59.
Ornston, L.N., and Stanier, R.Y., 1966. The conversion of catechol and protochatechaute to
beta-ketoadipate by Pseudomonas putida. J Biol Chem. 16, 3776-86.
O’Sullivan, D.J., and O’Gara, F., 1992. Traits of fluorescent Pseudomonas spp. Involved in
suppression of plant root pathogens. Microbiol Rev. 56, 662-676.
O’Toole, G.A., and Kolter, R., 1998. Flagellar and twitching motility are necessary for
Pseudomonas aeruginosa biofilm development. Mol Microbiol. 28, 449-61
Parsek, M., and Singh, P.K., 2003. Bacterial biofilms: an emerging link to disease
pathogenesis. Annu Rev Microbiol. 57, 677-701.
Passador, L., Cook, J.M., Gambello, M.J., Rust, L., and Iglewski, B.H., 1993. Expression of
105
Pseudomonas aeruginosa virulence genes requires cell-to-cell communication. Science. 260,
1127-30.
Passador, L., and Iglewski, B.H., 1995. Quorum sensing and virulence gene regulation in
Pseudomonas aeruginosa. In Roth JA, editor. Virulence mechanisms of bacterial pathogens. 2nd
ed. Washington: Am Soc for Microbiol. pp. 65-78.
Patel. M.V.P., 2000. The regulatory roles of PyrR and Crc in pyrimidine metabolism in
Pseudomonas aeruginosa. Ph.D. Dissertation. University of North Texas, Denton, Texas.
Paulsen, I.T., Brown, M.H., and Skurray, R.A., 1996. Proton-dependent multidrug efflux
systems. Microbiol Rev. 60, 575-608.
Pearson, J.P., Gray, K.M., Passador, L. Tucker, K.D., Eberhart, A., Iglewski, B.H., and
Greenberg, E.P., 1994. Structure of the autoinducer required for expression of Pseudomonas
aeruginosa virulence genes. Proc Natl Acad Sci U.S.A. 91, 197-201.
Pearson, J.P., Passador, L., Iglewski, B.H., and Greenberg, E.P., 1995. A second N-
acylhomoserine lactone signal produced by Pseudomonas aeruginosa. Proc Natl Acad Sc U.S.A.
92, 1490-4.
Pearson, J.P., Pesci, E.C., and Iglewski, B.H., 1997. Roles of Pseudomonas aeruginosa las and
rhl quorum-sensing systems in control of elastase and rhamnolipid biosynthesis genes. J
Bacteriol. 179, 5756-67.
106
Pearson, J.P., Van Delden, C., and Iglewski, B.H., 1999. Active efflux and diffusion are
involved in transport of P.aeruginosa cell-to-cell signals. J Bacteriol. 181, 1203-10.
Pesci, E.C., and Iglewski, B.H., 1999. Quorum sensing in Pseudomonas aeruginosa. In G.M.
Dunny and S.C. Winnans (ed), Cell-cell signaling in bacteria. Am Soc Microbiol. pp. 147-55.
Pesci, E. C., Milbank, J.B.J., Pearson, J.P., McKnight, S., Kende, A.S., Greenberg, E.P.,
and Iglewski, B.H., 1999. Quinolone signaling in the cell to cell communication system of
Pseudomonas aeruginosa. Proc Natl Acad Sci U.S.A. 96,11229-34.
Pesci, E. C., Pearson, J.P., Seed, P.C., and Iglewski, B.H., 1997. Regulation of las and rhl
quorum sensing in Pseudomonas aeruginosa. J Bacteriol. 179, 3127-32.
Pier, G.B., 1998. Pseudomonas aeruginosa: a key problem in cystic fibrosis. ASM News. 6,
339-47.
Prescott, L.M., and Jones, M.E., 1969. Modified Materials and Methods for the determination
of carbamyl aspartate. Anal Biochem. 32, 408-19.
Rahme, L.G., Stevens, E.J., Wolfort, S.F., Shao, J., Tompkins, R.G., and Ausubel, F.M.,
1995. Common virulence factors for bacterial pathogenecity in plants and animals. Science. 268,
1899-1902.
107
Rahme, L.G., Tan, M.W., Le, L., Wong, S.M., Tompkins, R.G., Calderwood, S.B., and
Ausubel, F.M., 1997. Use of model plant hosts to identify Pseudomonas aeruginosa virulence
factors. Proc Natl Acad Sci U.S.A. 94, 13245-50.
Rashid, M.H., and Kornberg, A., 2000. Inorganic polyphosphate is needed for swimming,
swarming, and twitching motilities of Pseudomonas aeruginosa. Proc. Natl. Acad. Sci U S A.
97, 4885-90.
Read, R.C., Roberts, P., and Munro, N., 1992. Effect of Pseudomonas aeruginosa
rhamnolipids on mucociliary transport and ciliary beating. J Appl Physiol.72, 2271-7.
Reimmann, C., Beyeler, M., Latifi, A., Winteler, H., Foglino, M., Lazdunski, A., and Haas,
D., 1997. The global activator GacA of Pseudomonas aeruginosa PAO positively controls the
production of the autoinducer N-butyryl-homoserine lactone and the formation of the virulence
factors pyocyanin, cyanide, and lipase. Mol Microbiol. 24, 309-19.
Rowland, P., Nørager, S., Jensen, K.F., and Larsen, S., 2000. Structure of dihydroorotate
dehydrogenase B: electron transfer between two flavin groups bridged by an iron-sulphur cluster.
Struct Fold Des. 8, 1227-38.
Roof, W.D., Foltermann, K.F., and Wild, J.R., 1982. The organization and regulation of the
pyrBI operon in Escherichia coli includes a rho-dependent attenuator sequence. Mol Gen Genet.
187, 391-400.
108
Rückemann, K., Fairbanks, L.D., Carrey, E.A., Hawrylowicz, C.M., Richards, D.F.,
Kirschbaum, B., and Simmonds, H.A., 1998. Leflunomide inhibits pyrimidine de novo
synthesis in mitogen-stimulated T-lymphocytes from healthy humans. J Biol Chem. 273, 21682-
91.
Rumbaugh, K.P., Griswold, J.A., and Hamood, A.N., 2000. The role of quorum sensing is the
in vivo virulence of Pseudomonas aeruginosa. (rev) Microbes Infect. 2, 1721-31.
Sambrook, J., Fritsch, E.F., and Maniatis, T., 1989. Molecular Cloning A Lab manual, 2nd ed.
Cold Spring Harbor Laboratory Press.
Scapin, G., Sacchettini, J.C., Dessen, A., Bhatia, M., and Grubmeyer, C., 1993. Primary
structure and crystallization of orotate phosphoribosyltransferase from Salmonella typhimurium.
J Mol Biol. 230, 1304-8.
Schurr, M.J., Vickrey, J.F., Kumar, A.P., Campbell, A.L., Cunin, R., Benjamin, R.C.,
Shanley, M.S., and O’Donovan, G.A., 1995. Aspartate transcarbamoylase genes in
Pseudomonas putida: Requirement for an inactive dihydroorotase for assembly into dodecameric
holoenzyme. J Bacteriol. 177, 1751-59.
Schwartz, M., and Neuhard, J., 1975. Control of expression of pyr genes in Salmonella
typhimurium: effects of variation in uridine and cytidine nucleotide pool. J Bacteriol. 121, 814-
22.
109
Schwyn B., and Neilands, J.B., 1987. Universal chemical assay for the detection and
determination of siderophores. Anal Biochem. 160, 47-56.
Serino, L., Reimmann, C., Visca, P., Beyeler, M., Della Chiesa, V., and Haas, D., 1997.
Biosynthesis of pyochelin and dihydroaeruginoic acid requires the iron-regulated pchDCBA
operon in Pseudomonas aeruginosa. J. Bacteriol. 179, 248-57.
Siegmund, L., and Wagner, F., 1991. New method for detecting rhamnolipids excreted by
Pseudomonas species during growth in mineral agar. Biotechnol Tech. 5, 265-8.
Singh, P.K., Schaefer, A.L., Parsek, M.R., Moninger, T.O., Welsh, M.J., and Greenberg,
E.P., 2000. Quorum-sensing signals indicate that cystic fibrosis lungs are infected with bacterial
biofilms. Nature. 407, 762-4.
Smith, J.M., Kelln, R.A., and O’Donovan, G.A., 1980. Repression and derepression of the
enzymes of the pyrimidine pathway in Salmonella typhimurium. J Gen Microbiol. 121, 27-38.
Stickler, D., 1999. Biofilms. Curr Opin. Microbiol. 2, 270-5.
Stinzi, A., Barnes, C., Xu, J., and Raymond, K.N., 2000. Microbial iron transport via a
siderophore shuttle: a membrane ion transport paradigm. Proc Natl Acad Sci U.S.A. 97, 10691-6.
110
Taguchi, K., Fukutomi, H., Kuroda, A., Kato, J., and Ohtake, H., 1997. Genetic
identification of chemotactic transducers for amino acids in Pseudomonas aeruginosa.
Microbiol. 143, 3223-9.
Turnbough, C.L., Hicks, K.L, and Donahue, J.P., 1983. Attenuator control in pyrBI operon
expression in Escherichia coli K12. Proc Natl Acad Sci U.S.A. 80, 368-72.
Vickrey, J. F., 1993. Isolation and Characterization of the Operon Containing Aspartate
Transcarbamoylase and Dihydroorotase from Pseudomonas aeruginosa. Ph.D. thesis, University
of North Texas, Denton, Texas.
Washabaugh, M.W., Collins, K.D., 1986. Dihydroorotase from Escherichia coli: Sulfhydryl
group-metal ion interactions. J Biol Chem. 261, 5920-9.
West, T.P., 1994. Control of the pyrimidine biosynthetic pathway in Pseudomonas
pseudoalcaligenes. Arch Microbiol. 162, 75-9.
West, T.P., 2003. Regulation of pyrimidine nucleotide formation in Pseudomonas reptilivora.
Lett Appl Microbiol. 38, 81-6.
West, T.P., 2004. Control of pyrimidine biosynthesis in “Pseudomonas alkanolytica” ATCC
21034. J Basic Microbiol. 44, 253-7.
111
West, T.P., 2005. Regulation of pyrimidine biosynthesis in Pseudomonas resinovorans. Lett
Appl Microbiol. 40, 473-8.
West, T.P., Herlick, S.A., and O’Donovan, G.A., 1983. Inverse relationship between
thymidylate synthetase and cytidine triphosphate activities during pyrimidine limitation in
Salmonella typhimurium. FEMS Microbiol Lett. 18, 275-8.
West, T.P., Traut, T.W., Shanley, M.S., and O’Donovan, G.A., 1985. A Salmonella
typhimurium strain defective in uracil catabolism and beta-alanine synthesis. J Gen Microbiol.
131, 1083-90.
Whitehead, N.A., Barnard, A.M., Slater, H., Simpson, N.J., and Salmon, G.P., 2001.
Quorum-sensing in gram-negative bacteria. FEMS Microbiol Rev. 25, 365-404.
Wick, M.J., Hamood, A.N., and Iglewski, B.H., 1990. Analysis of the structure-function
relationship of Pseudomonas aeruginosa exotoxin A [review]. Mol Microbiol. 4, 527-35.
Williamson, R.A., Yea, C.M., Robson, P.A., Curnock, A.P., Gadher, S., Hambleton, A.B.,
Woodward, K., Bruneau, J.M., Hambleton, P., and Moss, D., 1995. Dihydroorotate
dehydrogenase is a high affinity binding protein for A77 1726 and mediator of a range of
biological effects of the immunomodulatory compound. J Biol Chem. 270, 22467-72.
Woods, D.E., and Iglewski, B.H., 1983. Toxins of Pseudomonas aeruginosa: new perspectives
[review]. Rev Infect Dis. 4, S715-22.
112
Yurgel, S., Mortimer, M.W., Rogers, K.L., and Kahn, M.L., 2000. New substrates for the
dicarboxylate transport system in Sinorhizobium meliloti. J Bacteriol. 182, 4216-21.
Zhou, C., Yang, Y., and Jong, A.Y., 1990. Mini-prep in ten minutes. Biotech. 8, 172-3.