The role of catalase in Pseudomonas aeruginosa biofilm resistance to hydrogen peroxide
by James Garrett Elkins
A thesis submitted in partial fulfillment of the requirements for the degree of Master of Science in Land
Resources and Environmental Sciences
Montana State University
© Copyright by James Garrett Elkins (1999)
Abstract:
Microbial biofilm formation on man-made surfaces can profoundly impact human health and welfare.
Biofilms readily develop on medical implants such as catheters and artificial joints causing chronic
infections. Industrial processing systems are also plagued with the accumulation of biofilm in pipes,
heat exchangers, cooling towers, and other equipment resulting in the loss of system efficiency.
Biofilm bacteria are tremendously more difficult to kill with antimicrobial agents than freely suspended
organisms. The basis for biofilm resistance to antimicrobial agents has been investigated but, the
primary mechanisms responsible for this phenomena are still poorly understood. It has been
hypothesized that biofilm bacteria are able to rapidly adapt to an antimicrobial agent and neutralize it
with protective proteins/enzymes. This hypothesis was addressed in this study by investigating the
adaptive response of Pseudomonas aeruginosa biofilms to the oxidizing biocide, hydrogen peroxide
(H2O2). P. aeruginosa expresses two catalase enzymes in defense against H2O2 known as KatA and
KatB. These enzymes catalyticaly degrade H2O2 into oxygen and water. P. aeruginosa mutants that are
unable to synthesize either KatA or KatB were tested for their susceptibility to H2O2 when grown as
planktonic cells and biofilms. Biofilms lacking KatA activity were more susceptible to H2O2 than the
wild-type or KatB- strain but remained much more resistant to the biocide than planktonic cultures.
However, the absence of catalase activity in KatA- biofilms allowed efficient biofilm removal with
H2O2. Biofilms unable to synthesize the inducible KatB catalase were nearly equal to the wild-type
strain with respect to H2O2 resistance. Using spectrophotometric assays, activity gel staining
techniques, and reporter enzyme measurements, catalase expression was monitored in biofilms.
Specific catalase levels were essentially equal for biofilms and planktonic cells. Biofilms exposed to
relatively high concentrations of H2O2 induced the KatB catalase but induction patterns did not differ
from planktonic cells exposed to lower H2O2 concentrations. In conclusion, constitutive catalase
expression is necessary for optimal biofilm resistance to H2O2 but other mechanisms of resistance
must exist. Biofilms are capable of adapting to exogenous H2O2 by inducing KatB synthesis but this
enzyme is relatively insignificant to overall biofilm resistance to short-term H2O2 exposure. THE ROLE OF CATALASE IN Pseudomonas aeruginosa
BIOFILM RESISTANCE TO HYDROGEN PEROXIDE
by
James Garrett Elkins
A thesis submitted in partial fulfillment
of the requirements for the degree
of
Master of Science
in
Land Resources and Environmental Sciences
MONTANA STATE UNIVERSITY-BOZEMAN
Bozeman, Montana
May 1999
APPROVAL
of a thesis submitted by
James Garrett Elkins
This thesis has been read by each member of the thesis committee and has been
found to be satisfactory regarding content, English usage, format, citations, bibliographic
style, and consistency, and is ready for submission to the College of Graduate Studies.
Timothy R. McDermott
Approved for the Department of Land Resources and Environmental Sciences
Jeffery S. Jacobsen
A? 7
(Sjgnafiife)
Date
Approved for the College of Graduate Studies
Bruce R. McLeod
/r
(Signature)
Date
iii
STATEMENT OF PERMISSION TO USE
In presenting this thesis in partial fulfillment of the requirements for a master’s
degree at Montana State University-Bozeman, I agree that the Library shall make it
available to borrowers under rules of the Library.
If I have indicated my intention to copyright this thesis by including a copyright
notice page, copying is allowable only for scholarly purposes, consistent with “fair use”
as prescribed in the U.S. Copyright Law. Requests for permission for extended quotation
from or reproduction of this thesis in whole or in parts may be granted only by the
copyright holder.
Signature
Date
7
iv
ACKNOWLEDGMENTS
Special thanks must be given to several individuals who helped me complete this
thesis. I especially thank Dr. Timothy R. McDermott for his guidance and advice over
the last several years. Dr. Phil Stewart and I enjoyed many hours of insightful discussion
regarding this project and he provided outstanding direction. Dr. Thamir Al-Niemi often
provided help with difficulties encountered during my work and Dr. Michael L. Summers
proved to be a fountain of wisdom in and out of the laboratory. I would also like to thank
Dr. Dan Hassett for conveying his knowledge of oxidative stress physiology and for
providing the catalase mutants used in this study. Dr. Herbert Schweizer and Dr. Michael
Franklin deserve credit for furnishing many of the plasmids and strains needed for my
research. Many close friends and family members provided support during my graduate
work, especially my fiance Melissa, Sam and Louise Sampson, Bill and Jana McRae,
Nicole and McKenzie Elkins, Tracy Elkins, Tom and Lauren Mackay, and Scott Korell.
Most of all, I would like to thank my parents, Jim and Charlene, for their tremendous
amount of love and support throughout my educational endeavors.
TABLE OF CONTENTS
LIST OF TABLES............................................................................................................ vii
LIST OF FIGURES........................................................................................................... ix
ABSTRACT....................................................................................................................... x
1. INTRODUCTION........................................................................................................ I
References............................................................................................................ 6
2. LITERATURE REVIEW............................................................................................
Introduction..........................................................................................................
Mechanisms of Biofilm Resistance to Antimicrobial Agents.............................
Role of Extracellular Polysaccharide Matrix..........................................
Resistance Due to Low Physiological Activity......................................
Biofilm Physiological Adaptation to Antimicrobial Agents...................
Reactive Oxygen Intermediates..............................
Superoxide (O2")......................................................................................
Hydrogen Peroxide (H2O2).....................................................................
The Hydroxyl Radical (HO )...................................................................
Oxidizing Biocides............................................ :...............................................
Chlorine..................................................................................................
Ozone................................................................................
Hydrogen Peroxide.................................................................................
Mechanisms of Oxidative Stress Resistance.............................................
Superoxide Response and SoxRS Regulon.............................................
Regulation of Superoxide Dismutases....................................................
SOD regulation in E. coli............................................................
SOD regulation in P. aeruginosa................................................
Hydrogen Peroxide Response.................................................................
OxyR Regulon.......................................................................................
Regulation of Catalase in E. coli.............................................................
Regulation of katG in E. coli..............................................
Regulation of katE in E. coli............................ ,.........................
Regulation of Catalase in P. aeruginosa...........................................
Summary.................................................
8
8
10
10
13
15
17
17
18
19
20
21
22
23
24
25
26
27
28
29
29
31
32
34
36
38
vi
References..........................................
3. Pseudomonas aeruginosa BIOFILM RESISTANCE TO HYDROGEN
PEROXIDE: PROTECTIVE ROLE OF CATALASE............................................
Introduction........................................................................................................
Materials and Methods........................................................................................
Bacterial Strains and Plasmids..........................................................
Media and Growth Conditions................................................................
Planktonic Disinfection...........................................................................
Biofilm Disinfection...............................................................................
Preparation of Cell Free Extracts...:....................
Catalase Assays and Native-PAGE........................................................
DNA Manipulation..............................
Construction of pJGE02....................
Construction of pJGE03................................
Isolation of PAOI(katB\\lacZ)................
Reporter Gene Assays.........................................................,..................
Statistical Analysis........................................
Results.....................................................................................
Catalase Activity in katA and katB mutants............................................
H2O2 Sensitivity of Planktonic Cells......................................................
H2O2 Sensitivity of Biofilm Cells...........................................................
Biofilm Removal with Hydrogen Peroxide............................................
Catalase Activity of P. aeruginosa Biofilms..........................................
katB::IacZ Reporter Construction and Analysis.....................................
Discussion.........................................................
References........................................................ ................. ,.............:...............
40
49
49
52
52
52
54
55
56
57
57
57
58
58
59
59
60
60
62
62
63
67
70
72
77
4. CONCLUSIONS......................................................................................................... 81
APPENDIX
86
vii
LIST OF TABLES
Table
Page
3-1. List of Strains and Plasmids Used in this Study................................... .............. 53
3-2. Planktonic Specific Catalase Activity of PAOl wt,
KatA", and KatB" Before and After Treatment with H2O2................................. 61
A-1. Percent Survival of PAOl(wt) Planktonic Cells During Disinfection................. 86
A-2. Percent Survival of PA01(KatA") Planktonic Cells During Disinfection......... 86
A-3. Percent Survival of PAOI (KatB') Planktonic Cells During Disinfection.......... 86
A-4. Percent Survival of PAOl (wt) Biofilm Cells During Disinfection..................... 87
A-5. Percent Survival of PAOl(KatA') Biofilm Cells During Disinfection.............. 87
A-6. Percent Survival of PAOI (KatB") Biofilm Cells During Disinfection............... 87
A.-1. Percent Removal of PAOl(wt) Biofilm Cells During Disinfection.................... 88
A-8. Percent Removal of PAOl(KatA ) Biofilm Cells During Disinfection............... 88
A-9. Percent Removal of PAOl(KatB') Biofilm Cells During Disinfection............... 88
A-10. Specific Catalase Activity of Stationary Phase, Planktonic wt,
KatA", and KatB" Cells Before and After Treatment with 2mM
Pulses of H2O2 Every 10 min for I hr.................................................................. 89
A -II. Specific Catalase Activity (U/mg protein) of PAOl wt
Biofilm During Disinfection................................................................................ 89
A-12. Specific Catalase Activity (U/mg protein) of PAOl KatB"
Biofilm During Disinfection............................................................................
90
viii
LIST OF TABLES (Cont)
Table
A-13. Specific Catalase Activity (U/mg protein) of PAOl KatB"
Biofilm During Disinfection..................................................
Page
90
A-14. PAOI QcatB::lacT) Reporter Activity in Miller. Units
for Biofilms Exposed to 50 mM H2O2 for I hr.................................................... 90
A-15. PAOI QcatB::lacZ) Reporter Activity in Miller Units
for Biofilms Not Exposed to H2O2...................................................................... 91
A-16. PAOI QcatBwlacZ) Reporter Activity in Miller Units
for Planktonic Cells Exposed to 2mM Pulses of H2O2
Every 10 min. for I hr.......................................................................................... 91
A-17. PAOI QiatB::lacZ) Reporter Activity in Miller Units
for Planktonic Cells Exposed to 1-50 mM Dose of H2O2.................................... 91
A-18. PAOI QiatB::lacZ) Reporter Activity in Miller Units
for Planktonic Cells Not Exposed to H2O2.......................................................... 92
ix
LIST OF FIGURES
Figure
Page
2- 1. Induction of HPI During Exponential Growth By the OxyR System............... 33
2- 2. Model of Catalase Regulation in Stationary Phase E. coli............................... . 35
3- 1. Planktonic Cell Disinfection with 50 mM H2O2................................................ 64
3- 2. Biofilm Disinfection with 50 mM H2O2............................ ........'...................... 65
3-3. Biofilm Removal During Disinfection.................................................................. 66
3-4. Biofilm Specific Catalase Activity........................................................................ 68
3-5. Biofilm Catalase Expression During Disinfection................................................ 69
3-6. PAOl (katB::lacZ) Expression in Response to Hydrogen Peroxide
71
ABSTRACT
Microbial biofilm formation on man-made surfaces can profoundly impact human
health and welfare. Biofilms readily develop on medical implants such as catheters and
artificial joints causing chronic infections. Industrial processing systems are also plagued
with the accumulation of biofilm in pipes, heat exchangers, cooling towers, and other
equipment resulting in the loss of system efficiency. Biofilm bacteria are tremendously
more difficult to kill with antimicrobial agents than freely suspended organisms. The
basis for biofilm resistance to antimicrobial agents has been investigated but, the primary
mechanisms responsible for this phenomena are still poorly understood. It has been
hypothesized that biofilm bacteria are able to rapidly adapt to an antimicrobial agent and
neutralize it with protective proteins/enzymes. This hypothesis was addressed in this
study by investigating the adaptive response of Pseudomonas aeruginosa biofilms to the
oxidizing biocide, hydrogen peroxide (H2O2). P- aeruginosa expresses two catalase
enzymes in defense against H2O2 known as KatA and KatB. These enzymes catalyticaly
degrade H2O2 into oxygen and water. P. aeruginosa mutants that are unable to
synthesize either KatA or KatB were tested for their susceptibility to H2O2 when grown
as planktonic cells and biofilms. Biofilms lacking KatA activity were more susceptible to
H2O2 than the wild-type or KatB' strain but remained much more resistant to the biocide
than planktonic cultures. However, the absence of catalase activity in KatA" biofilms
allowed efficient biofilm removal with H2O2. Biofilms unable to synthesize the inducible
KatB catalase were nearly equal to the wild-type strain with respect to H2O2 resistance.
Using spectrophotometric assays, activity gel staining techniques, and reporter enzyme
measurements, catalase expression was monitored in biofilms. Specific catalase levels
were essentially equal for biofilms and planktonic cells. Biofilms exposed to relatively
high concentrations of H2O2 induced the KatB catalase but induction patterns did not
differ from planktonic cells exposed to lower H2O2 concentrations. In conclusion,
constitutive catalase expression is necessary for optimal biofilm resistance to H2O2 but
other mechanisms of resistance must exist.. Biofilms are capable of adapting to
exogenous H2O2 by inducing KatB synthesis but this enzyme is relatively insignificant to
overall biofilm resistance to short-term H2O2 exposure.
I
CHAPTER I
INTRODUCTION
Prokaryotic success in many different environments can likely be attributed to the
tendency of bacteria to form biofilms. Biofilms arise when planktonic cells adhere to a
surface and undergo phenotypic changes including the expression of genes involved in
exopolysaccharide (EPS) synthesis (Davies et al., 1993, Davies and Geesey, 1995).
Through binary fission, single cells become microcolonies and eventually, a highly
complex microbial community is established. Biofilms consist of microorganisms, a
polymer matrix surrounding each cell, a system of voids and flow channels allowing fluid •
convection throughout the biofilm, and other abiotic materials that become trapped by the
EPS matrix (Costerton et al., 1995). As a common soil/water microorganism and
opportunistic pathogen, Pseudomonas aeruginosa occupies many diverse niches in
nature. Since P. aeruginosa is often isolated from natural biofilms, this species serves as
an appropriate model organism for biofilm research.
Biofilm formation is problematic in many different medical and industrial
situations. For instance, the colonization of catheters and prosthetic devices by
microorganisms results in chronic systemic infections, especially in
immunocompromised patients. P. aeruginosa colonization and biofilm formation is often
the cause of morbidity and mortality in patients suffering from cystic fibrosis. Regrowth
of coliform biofilms in drinking water distribution systems can also impact human health.
In industrial systems, heat exchangers are less efficient when fouled with biofilms.
Biofilm accumulation in pipe systems results in pressure loss and poor flow
characteristics. The warm, oxygenated waters found in cooling towers are especially
conducive to biofilm formation, resulting in loss of cooling efficiency and structural
integrity. The corrosion of metal surfaces is also enhanced when colonized with bacteria
(Costertonetal., 1988).
Antimicrobial agents are widely used to control unwanted microbial growth in
medical and industrial situations. Antimicrobial agents include all classes of antibiotics
(i. e. (3-lactams) as well as low molecular weight, broad spectrum biocides such as
hypochlorous acid and hydrogen peroxide. The use of these compounds to kill or remove
biofilms often proves difficult since sessile bacteria are much more resistant to killing
than freely suspended organisms (Ashby et ah, 1994; Brown and Gilbert, 1993;
LeChevallier et ah, 1988). Currently, the basis for the reduced efficacy of antimicrobial
agents against biofilm organisms is poorly understood. Knowing the underlying
mechanisms responsible for biofilm resistance will allow for the development of
improved antimicrobial compounds as well as better application protocols for compounds
already in use.
Several biofilm structural and physiological characteristics have been assessed for
their role in biofilm resistance to antimicrobial agents. The EPS matrix may provide
protection by establishing a reaction/diffusional barrier that prevents penetration of the
antimicrobial compound into the biofilm (Nichols et al., 1989; Tresse and Junter, 1995),
Poor biocide penetration has been shown to enhance the survival of biofilm bacteria
3
although some investigators argue that slow penetration alone is insufficient to account
t
for the resistance biofilms display to many bactericidal compounds (Brown and Gilbert,
1994; Stewart, 1996). Low physiological activity has been demonstrated in P.
aeruginosa biofilms due to oxygen limitation (Xu et al., 1998). The mode of action for
many antimicrobial agents such as P-Iactams is growth dependent; therefore, growth
limiting conditions within a biofilm may account for reduced susceptibility.
Another mechanism that may contribute to the reduced efficacy of antimicrobial
agents against biofilm organisms is physiological adaptation to the compound itself.
Upon exposure to a particular agent, bacteria may possess the genetic and physiological
capability of synthesizing proteins/enzymes that are able neutralize or catalyticaly
degrade the compound in use. Physiological adaptation to antimicrobial agents has been
observed in biofilm disinfection experiments but the actual mechanisms responsible for
adaptation are not understood (Sanderson and Stewart, 1997).
Adaptive stress responses to oxidizing agents have been well characterized in
enteric bacteria. One of the best described adaptive responses to an oxidizing agent
includes the expression of catalase in defense against the oxidizing biocide, hydrogen
peroxide (H2O2). Catalase enzymatically degrades H2O2 into oxygen and water. E. coli
increases total cellular catalase activity when exposed to sub-lethal doses of H2O2,
enabling the organism to become resistant to much higher concentrations of the oxidant
(Christman et al., 1985). Catalase expression and its role in scavenging H2O2 has been
studied in organisms such as E. coli for nearly 30 years and a comprehensive review of
this work is provided in this thesis. Since past research has been conducted almost
exclusively with planktonic organisms, little is known about the importance of catalase
expression in biofilm resistance to H2O2.
Hassett and co-workers (Brown et al., 1995; Hassett et al, 1992) have studied
catalase expression in planktonic P. aeruginosa cells under both oxidative and nonoxidative conditions. Two genes, termed TcatA and katB, encoding two distinct catalases
have been identified in P. aeruginosa. The JeatA gene is constitutively expressed
throughout the aerobic growth cycle while IeatB transcription occurs only when cells are
under oxidative attack by H2O2. Since adaptation to H2O2 has been described for
planktonic cells of P. aeruginosa, relatively simple yet fundamentally important
questions could be addressed in this thesis regarding the importance of adaptive
responses in biofilm resistance to H2O2.
By assessing the susceptibility of mutant biofilms that are unable to synthesize either
KatA or KatB to H2O2, the following hypotheses were tested:
1. P. aeruginosa biofilms that cannot constitutively express catalase (KatA")
will be hypersusceptible to H2O2.
0 KatA mutants have essentially no catalase activity. What fraction of overall
biofilm resistance to" H2O2 can be attributed to constitutive catalase
expression?
2. Catalase (KatB) induction in response to H2O2 is necessary for biofilm resistance
to H2O2.
0 If P. aeruginosa biofilms are unable to initiate an adaptive response to
peroxide by inducing KatB, will they be more susceptible to H2O2 relative to
wild-type biofilms.
By studying the expression patterns of both enzymes under oxidative and non-oxidative
stress conditions, these hypotheses were tested:
3. Even in the absence OfH2Ch, catalase genes (JeatA, katB, or both) are upregulated
in biofilms resulting in a high specific catalase activity relative to planktonic
organisms.
o Perhaps biofilms are resistant to H2Oa because of differential gene
regulation that results in relatively high biofilm specific catalase
activity.
4. P. aeruginosa biofilms initiate an oxidative stress response upon exposure to
H2O2.
o Although this has been demonstrated in planktonic cells (Brown et ah, 1995),
catalase expression, specifically katB induction, may be differentially
regulated in biofilms.
This thesis begins with a comprehensive literature review (Chapter 2) covering
pertinent topics such as possible mechanisms of biofilm resistance to antimicrobial
agents, evidence for physiological adaptation to antimicrobials, the use of oxidizing
biocides to control problematic biofilms, oxygen radical chemistry and biology, and
oxidative stress genetics and physiology in enteric bacteria and P. aeruginosa. Chapter 3
features the main scientific study completed for this thesis in a professional publication
format. In addition to assessing the importance of adaptive responses in biofilm
resistance to oxidizing agents, this research represents a novel approach toward
understanding basic differences in oxidative stress physiology between planktonic
organisms and biofilms. Also resulting from this work, we have described an excellent
model system for examining resistance mechanisms in biofilms (Hassett et ah, 1999a) as
well as uncover other aspects of biofilm physiology that affect biofilm resistance to
oxidizing agents (Hassett et ah, 1999b). These contributions should provide further
insight regarding differences in biofilm and planktonic cell physiology and will hopefully
assist in the development of improved antimicrobial agents and biofilm control strategies.
6
References
Ashby, M. J., J. E. Neale, S. J. Knott, and I. A. Critchley. 1994. Effect of antibiotics
on non-growing planktonic cells and biofilms of Escherichia coli. J. Antimicroh
Chemother. 33:443-452.
Brown, M. R. W., and P. Gilbert. 1993. Sensitivity of biofilm to antimicrobial agents. J.
AppL Bacteriol. Symp. Suppl. 74:87S-97S.
Brown, S. M., M. L. Howell,, M. L. Vasil, A. J. Anderson, and B. J. Hassett. 1995.
Cloning and characterization of the katB gene of Pseudomonas aeruginosa encoding a
hydrogen peroxide-inducible catalase: purification of KatB5cellular localization, and
demonstration that it is essential for optimal resistance to hydrogen peroxide. J. Bacteriol.
177:6536-6544.
Christman, M. F., R. W. Morgan, F. S. Jacobson, and B. N. Ames. 1985. Positive
control of a regulon for defenses against oxidative stress and some heat-shock proteins in
Salmonella typhimurium. Cell. 41:753-762.
Costerton, J. W., Z..Lewandowski, B. E. Caldwell, D. R. Korber, and H. M. LappinScott. 1995. Microbial biofilms. Annu. Rev. Microbiol. 49:711-745.
Costerton, J. W., G. G. Geesey, and P. A. Jones. 1988. Bacterial biofilms in relation to
internal corrosion monitoring and biocide strategies. Materials Performance. 27:49-53.
Davies D. G., A. M. Chakrabarty, and G. G. Geesey. 1993. Exopolysaccharide
production in biofilms: substratum activation of alginate gene expression by
Pseudomonas aeruginosa. AppL Environ. Microbiol. 59:1181-1186.
Davies D. G., and G. G. Geesey. 1995. Regulation of the alginate biosynthesis gene
algC in Pseudomonas aeruginosa during biofilm development in continuous culture.
AppL Environ. Microbiol. 61:860-867.
Hassett, D. J., L. Charniga, K. Bean, D. E. Ohman, and M. S. Cohen. 1992. Response
of Pseudomonas aeruginosa to pyocyanin: mechanisms of resistance, antioxidant
defenses, and demonstration of a manganese-cofactored superoxide dismutase. Infect.
Immun. 60:328-336.
Hassett, D. J., J. G. Elkins, D.-J. Ma, and T. R. McDermott. 1999a. Pseudomonas
aeruginosa biofilm sensitivity to biocides: use of hydrogen peroxide as a model
antimicrobial agent for examining resistance mechanisms. Meth. EnzymoL In press.
7
Hassett, D. J., J.-F. Ma, J. G. Elkins, T. R. McDermott, U. A. Ochsner, S. E. H.
West, C.-T. Huang, J. Fredericks, S. Burnett, P. S. Stewart, G. McFeters, L.
Passador, and B. H. Iglewski. 1999b. Quorum sensing in Pseudomonas aeruginosa
controls catalase and superoxide dismutase genes and mediates biofilm susceptibility to
hydrogen peroxide. J. Bacteriol. Submitted.
Lechevallier M. W., C. D. Cawthon, and R. G. Lee. 1988. Inactivation of biofilm
bacteria. AppL Environ. Microbiol. 54:2492-2499.
Nichols, W. W., M. J. Evans, M. P. E. Slack, and H. L. Walmsley. 1989. The
penetration of antibiotics into aggregates of mucoid and non-mucoid Pseudomonas
aeruginosa. J. Gen. Microbiol. 135:1291-1303.
Sanderson, S. S., and P. S. Stewart. 1997. Evidence of bacterial adaptation to
monochloramine in Pseudomonas aeruginosa biofilms and evaluation of biocide action
model. Biotechnol. Bioeng. 56:201-209.
Stewart, P. S. 1996. Theoretical aspects of antibiotic diffusion into microbial biofilms.
Antimicrob. Agents. Chemother. 40:2517-2522.
Tresse, O., T. Jouenne, and G.-A. Junter. The role of oxygen limitation in the
resistance of agar-entrapped, sessile-like Escherichia coli to aminoglycoside and 13lactam antibiotics. J. Antimicrob. Chemother. 36:521-526.
Xu, K. D., P. S. Stewart, F. Xia, C.-T. Huang, and G. A. McFeters. 1998.
Physiological heterogeneity in Pseudomonas aeruginosa biofilm is determined by
oxygen availability. AppL Environ. Microbiol. 64:4035-4039.
8
CHAPTER 2
LITERATURE REVIEW
Introduction
One of the primary reasons behind the problematic nature of biofilms is their
ability to resist killing by antimicrobial agents. Biofilm resistance to numerous
antimicrobial compounds has been demonstrated repeatedly (Brown and Gilbert, 1993;
LeChevallier et ah, 1988; Sanderson and Stewart, 1997) and, to date, no antimicrobial
agent has been shown to eliminate biofilms as effectively as it can suspended organisms.
Biofilm resistance appears to be universal for all types of compounds including
antibiotics such as 3-lactams and aminoglycosides (Anwar et al., 1990) as well as low
molecular weight biocides like hypochlorous acid and monochloramine (Berman et al.,
1988; Sanderson and Stewart, 1997). Biofilm resistance to such a wide range of
antimicrobial agents is the basis for the problematic nature of biofilms in so many
different situations. This is especially true in the prevention of nosocomial infections
where conventional antibiotics are proving to be poorly effective against many types of
pathogenic organisms. Biofilm formation by inherently antibiotic resistant strains of
bacteria greatly increases the potential for disease causing organisms to evade
disinfection.
9
Control of biofilms in industrial settings generally relies on the use of broad
spectrum, low molecular weight biocides that can be purchased in bulk quantities. Of
the many types of biocides employed in industry, members of the oxidizing class of
biocides remain the most widely used (Kuo and Smith, 1996). Oxidizing biocides
include halogenated compounds such as hypochlorous acid (HOC1), monochloramine
(NH2CI), chlorine dioxide (CIO2), and hypobromous acid (HOBr), or oxygen derivatives
such as ozone (O3) and hydrogen peroxide (H2O2). Biofilms composed of single
organisms or consortia of bacterial species have demonstrated resistance to oxidizing
biocides. LeChevallier et al. (1988) challenged planktonic and biofilm cultures of
heterotrophic bacteria isolated from drinking water and Klebsiella pneumoniae isolates
with varying doses of hypochlorous acid and monochloramine. Planktonic cells required
only 0.08 mg L'1 of hypochlorous acid or 94 mg L"1 of monochloramine to achieve a 99%
reduction in viability within I minute of exposure. Biofilms grown on the surfaces of
granular activated carbon particles, metal coupons, and glass microscope slides were 150to 3000 fold more resistant to equal doses of hypochlorous acid and 2- to 100 fold more
resistant to equal doses of monochloramine. The authors speculated that the decreased
reactivity of the monochloramine allowed the agent to penetrate deeper into the biofilm
thus improving its efficacy relative to hypochlorous acid.
Currently, the mechanisms responsible for biofilm resistance to antimicrobial
agents are poorly understood. Several unifying characteristics of biofilms and their
structure have been implicated in resistance. These hallmarks include cell encapsulation
within an extra-cellular polysaccharide matrix; decreased physiological activity due to
limitation of electron donors/acceptors or other essential nutrients; and the physiological
10
production of cell constituents capable of neutralizing biocides. These processes and
their role in biofilm resistance to antimicrobial agents will now be reviewed.
Mechanisms of Biofilm Resistance to Antimicrobial Agents
Role of Extracellular Polysaccharide Matrix
One of the prominent features of any biofilm is the presence of an
exopolysaccharide (EPS) matrix surrounding each cell. Biofilm formation is initiated
when bacteria originating from the external milieu attach to a surface. The ability of a
single cell to remain attached to a surface and initiate biofilm formation has been shown
to occur through the expression of genes responsible for polysaccharide biosynthesis
(Davies et al., 1993; Davies and Geesey, 1995). Cell division and the accretion of
additional planktonic cells results in the development of a mature biofilm which includes
an EPS matrix. Biofilm polymer matrices are highly hydrated containing up to 95%
water and are responsible for the heterogeneous three-dimensional structure of biofilms
)
which include clusters of cells (mushrooms) and flow channels (Costerton et al., 1995).
The EPS matrix of surface attached bacteria has been implicated in biofilm
resistance to antimicrobial agents (Nichols et al., 1989). Primarily, the EPS matrix is
thought to either slow or prevent the transport of antimicrobial agents from the bulk
phase to the encapsulated biofilm cells. This could occur with the matrix serving as a
simple physical barrier to convective/diffusive mass transport. Also, the antimicrobial
agent could physically adsorb to the matrix thus immobilizing it. Alternatively, reactive
biocides such as hypochlorous acid may act upon the EPS matrix, being
stoichiometrically consumed. The role of the EPS matrix in protecting biofihn bacteria
11
from various antimicrobial agents has been determined experimentally. Using
microelectrodes, DeBeer et al. (1994) directly measured the penetration of chlorine
(added as hypochlorous acid) into P. aeruginosa and K. pneumoniae biofilms cultured on
stainless steel slides. Sharp concentration gradients were established throughout the
depth of the biofilm within a few minutes following the addition of chlorine. The ability
of chlorine to penetrate these biofilms varied widely with concentration and exposure
times, although this variation was thought to be mainly due to heterogeneity in the
biofilm structure. Nevertheless, the EPS matrix of these two-species biofilms did
establish a reaction-diffusion barrier, enhancing resistance to chlorine.
Artificial biofilms constructed of bacteria entrapped in agarose gel slabs (Chen
and Stewart, 1996) or alginate and agarose beads (Xu et al., 1996) have been used as
models to study chlorine transport into polymer matrices. Such artificial biofilms posses
homogeneous matrices and consistent geometries which minimizes the variablility
otherwise encountered with real biofilms. In these studies, chlorine reacted slowly with
agarose and penetrated ca. 500 pm thick slab comparatively rapidly (15 min.) when cells
of P. aeruginosa were absent. However, the entrapment of bacteria in the agarose matrix
greatly retarded chlorine transport with penetration equivalent to cell free slabs occurring
after I hour of exposure. Unlike agarose, alginate presents a significant reactiondiffusion barrier for chlorine. After 45 hours of exposure to 20 mg L'1 chlorine,
concentrations did not reach 50% of the bulk phase at the center of 3 mm agarose beads
containing 2% (w/w) alginate. Agarose/alginate beads containing P. aeruginosa at a
density 2.5 x IO9 cfu/cm3further retarded chlorine penetration. Entrapment of cells in
polymer beads resulted in remarkable resistance to chlorine with 6 hours of exposure
12
killing only the cells near the surface of the bead.
Others have argued that the biofilm EPS matrix provides only limited or no
antimicrobial resistance. Using povidone-iodine as a disinfectant against P. .aeruginosa.
Brown et al. (1995) showed that the iodine demands imparted by biofilm and planktonic
samples of equal biomass were similar. Consequently, iodine reacting with the matrix
could not explain the difference in resistance observed between planktonic and biofilm
cells. Similarly, Nichols et al. (1989) measured the penetration of tobramycin and
cefsulodin through mucoid and non-mucoid aggregates of P. aeruginosa and concluded
that the rates of penetration were not significantly different between the two cell types.
Gristina et al. (1989) tested the susceptibility of mucoid and non-mucoid strains of
Staphylococcus epidermis to a number of antibiotics and concluded that polymer
encapsulation provided no protection.
The specific role of the EPS matrix in the protection of natural biofilms from
antimicrobial agents remains unclear. In general, the establishment of a
diffusion/convection barrier by the EPS layer is insufficient to explain the recalcitrance of
biofilms to disinfection (Quesnel et al., 1993; Stewart et al., 1996). This is supported by
the fact that biofilm cells remain resistant to biocides even after system equilibrium is
reached and solute concentrations within the biofilm equal those in the bulk fluid. This
suggests that other mechanisms are involved in biofilm resistance to disinfection. It is
likely that the nature of the biocide, primarily its reactivity with EPS, that will be
important in determining the degree to which the polysaccharide matrix will have
possible protective affects. Specifically, the level of protection afforded by the matrix
probably depends on biofilm species composition, age, hydraulic flow conditions, and the
13
chemical nature of the antimicrobial agent in question.
Resistance Due to Low Physiological Activity
Gradients of metabolic activity have been demonstrated within biofilms (Huang et
ah, 1998; Huang et ah, 1995; Xu et ah, 1998). Due to reaction/diffusion limitations on
the penetration of electron donors/acceptors, organisms residing near the substratum or
deep within microcolonies of a biofilm are likely to be experiencing a nutrient limited
physiology. Microelectrodes have been employed to demonstrate that in aerobic,
heterotropbic biofilms, oxygen concentrations sharply decline with biofilm depth
(Costerton et ah, 1995). Xu et ah (1998) recently concluded that oxygen was the primary
determinant of physiological activity in P. aeruginosa biofilms. Using the ability to
induce alkaline phosphatase under phosphate starvation conditions as an indicator of
protein synthesis, it was found that only the oxygenated fraction of the biofilm (ca. upper
30 |_im) was capable of de novo protein synthesis. Kinniment and Wimpenny (1992) used
a different approach to demonstrate physiological heterogeneity in P. aeruginosa
biofilms. Frozen sections of biofilm were sliced horizontally and then assayed for
adenylated nucleotides. Based on ratios of ATP, ADP, and AMP, an adenylate energy
charge (ECa) index could be established to predict cellular energy status. Interestingly,
AMP was the dominant species of adenine nucleotide in either 55 or 105 hour old
biofilms except near the biofilm surface where the ATP/ADP ratio increased. These
results suggested that a large percentage of the biofilm was in a metabolically quiescent.
Antibiotic efficacy is often dependant on the target organisms being metabolically
active. This is especially true for antibiotics that target growth related phenomena such
14
as peptidoglycan, nucleic acid, and protein synthesis. Gristina et al. (1989) determined
nafcillin, vancomycin, gentamicin, and daptomycin efficacy against suspended and
adherent Staphylococcus epidermidis and concluded that the increased resistance S.
epidermidis biofilms displayed to all of the antibiotics was likely due to slow growth.
Other studies have shown, however, that slow microbial growth rates were not
responsible for increased resistance to several antibiotics applied to differing
experimental conditions. Slow growth rates may not lead to Staphylococcus aureus
biofilm resistance to high molecular weight antibiotics such as benzylpenicillin,
tetracycline, and vancomycin (Williams et ah, 1997). The efficacies of halogenated
biocides such as iodine (Brown and Gauthier, 1993) and fluoride (Embleton et ah, 1998)
against biofilms have also been shown to be independent of specific growth rate.
Other phenomena related to growth are often implicated in biofilm resistance to
antimicrobial agents. Profound changes occur in cells when nutrients such as carbon,
nitrogen, phosphate, and/or iron become limiting (Kjelleberg, 1993). Nutrient limitation
is also known to induce significant changes in cell permeability to many antimicrobial
agents (Brown et ah, 1990), therefore decreasing susceptibility to disinfection.
Phenotypic changes elicited by nutrient limitation vary widely and many authors
speculate that some aspect of a nutrient limited physiology is responsible for the
increased resistance of biofilms to antimicrobial agents. Increased EPS production at the
onset of stationary phase has often been attributed to antibiotic resistance (Evans et ah,
1994). Although, as in many cases noted above, the presence of the EPS envelope alone
cannot account for decreased susceptibility. Also, the production of an EPS matrix has
been shown to result from just the initial attachment of P. aeruginosa to a surface (Davies
et al., 1995) which implies that formation of a polymer matrix is independent of
starvation; however, extracellular polymer formation likely plays some role in the long
term survival of biofilm organisms (Costerton et al., 1981).
Biofilm Physiological Adaptation to Antimicrobial Agents
Bacteria residing in a biofilm may physiologically adapt to antimicrobial agents,
therefore increasing their resistance. Adaptation, in this sense, occurs when the biofilm
cell expresses a phenotype that renders the bacterium less susceptible to a particular
agent. Since microorganisms have always been confronted with naturally occurring
antimicrobial agents, resistance mechanisms have evolved to overcome them. Adaptation
strategies include the expression of genes/operons that provide nonspecific protection to
various agents while other mechanisms act by irreversible enzymatic degradation.
Examples include the induction of membrane bound permeases to pump antibiotics such
as tetracycline out of the cell; the generation of intracellular reducing agents like
glutathione which can neutralize oxidizing biocides (Chapman et al. 1993; Chesney et al.
1996); and the induction of enzymes that possess high substrate specificity against a
particular agent such as (3-lactamases for [3-lactam antibiotics or catalase for hydrogen
peroxide. The onset of physiological resistance to antimicrobials in biofilms can be
elicited by exposure to the agent itself, or perhaps by the induction/repression of genes
associated with biofilm formation which co-incidentally affect antimicrobial
susceptibility.
Physiological adaptation to antimicrobial agents has been observed in biofilms.
Giwercman et al. (1991) recorded increases in p-lactamase production in P. aeruginosa
16
biofilms exposed to imipenem and peperacillin; the ability of the biofilms to induce
(3-lactamase upon exposure to the antibiotics afforded the cells increased protection. This
report demonstrated that exposure to the antimicrobial agent was the cause of increased
P-Iactamase activity rather than biofilm formation itself. When testing a mathematical
model designed to predict the kinetics of P. aeruginosa biofilm disinfection with
monochloramine, Sanderson and Stewart (1997) reported that biofilm cells became
increasingly resistant to subsequent doses of monochloramine at increasing
concentrations. They concluded that the adaptation to the oxidizing agent,
monochloramine, was perhaps due to an oxidative stress response being elicited by the
biofilm organisms.
Oxidizing biocides serve as the primary tools used to combat unwanted biofilm
accumulation and biofouling in industrial settings. Oxidants in the form of reactive
oxygen intermediates (ROIs) (see below) are also employed to kill invading
microorganisms in vivo via phagocytosis and the respiratory burst (Babior, 1987). Over
the last several decades, an impressive scientific effort has been focused on
characterizing the genetic, physiological, and biochemical mechanisms responsible for
the bacterial evasion of oxidizing agents. However, a weakness in our knowledge
regarding how bacteria adapt to and evade external oxidants exists since, in the past, only
planktonic organisms cultured in highly artificial environments have been studied. The
information obtained in this manner has unquestionably been crucial to our current
understanding of microbial genetics and physiology; nevertheless, bacteria in situ prefer
to associate with surfaces. Oxidative stress responses evoked by biofilm organisms are
yet to be explored and understood. The remainder of this review will cover the known
17
chemical species that induce oxidative stress responses in microorganisms as well as the
genetics and physiology of oxidative stress as it is known to occur in Eschehcia coli and
Pseudomonas aeruginosa.
Reactive Oxygen Intermediates
In aerobic organisms, the dependency on molecular oxygen (O2) as a terminal
electron acceptor is not without consequence. Toxic, reactive oxygen intermediates
(ROIs) are readily formed during the reduction of O2 to H2O through aerobic metabolism.
This is due to the paramagnetic nature of O2 which possesses unpaired electrons in
separate orbitals of parallel spin. In order to avoid formation of ROIs, simultaneous
reduction of each unpaired electron must occur by another electron of complementary
spin. Univalent, single electron reductions frequently occur through aberrant electron
flow in membrane associated electron transport systems, thus forming both oxygen
radicals and nonradical, but unstable, compounds (Fridovich, 1978; Demple, 1991).
ROIs are known to cause damage to nucleic acids (Imlay and Linn 1988), proteins
(Tamarit et al., 1998), and lipids (Farr and Kogoma, 1991). As shown in Equation I,
three distinct oxygen species may be created as O2 is reduced to water:
O2
j^i
^
HO2
^ H2O2
^ HO' + H2O
p+
2H2O
(Equation I)
Superoxide (PT)
The superoxide anion radical is generated via a single electron reduction of
molecular oxygen and is the most readily formed of the ROIs (Fridovich, 1978). In
18
biological systems, superoxide production mainly occurs via redox cycling agents and, to
a lesser extent, non-cyclic autoxidation of biomolecules by O2. Cyclic production of
superoxide is mediated through a large class of cellular and environmental compounds
which undergo reduction via electron carriers, usually NADH or NADPH, followed with
oxidation by O2. Redox cycling agents include quinones and the herbicide paraquat.
Imlay and Fridovich (1991) used membrane preparations of superoxide dismutase (SOD)
deficient E. coli cells to measure the flux of superoxide being generated by aerobic
respiration. These authors reported that respiring membrane vesicles generated 3 O2"
molecules for every 10,000 electrons entering the electron transport system which
correlated to an intracellular superoxide production rate of 5 p,mol/s. SODs restricted the
calculated intracellular concentration of superoxide to ca. 2x10"111M, which is sufficient
to cause oxidative damage. Superoxide is capable of oxidizing thiols, ascorbate,
tocopherols, and catecholamines, and especially proteins containing (Fe-S)4 centers (Farr
and Kogoma, 1991; Fridovich, 1978). An important characteristic of superoxide is its
ability to reduce both free and complexed transition metals, especially ferric iron, as
shown by the following reaction:
O f + M(n+1)+ ---------- ► O2 + Mn+
(Equation 2)
Hydrogen Peroxide (H2O2)
Superoxide electrons may be redistributed in vivo to generate O2 and H2O2. This
dismutation is mediated either by redox-active metals, such as iron, or enzymatically by
superoxide dismutases (Fridovich, 1978). Since hydrogen peroxide can freely permeate
cellular membranes, Gonzalez-Flecha and Demple (1995) could determine the
intracellular steady-state concentration OfH2O2 in oxygen respiring, intact cells offs, coli
by measuring the concentrations of peroxide present in extracellular buffers. They
concluded that exponentially growing cells maintain an intracellular H2O2 concentration
of 0.15 pM even in the presence of catalase. Using various inhibitors of the electron
transport system, these authors also determined the primary sources of H2O2 generation
within the cell. They reported that less than 20% of the steady state intracellular H2O2
concentration originated from the cytosolic fraction of the cell. Most H2O2originated
from O2' which was produced via aberrant electron flow from NADH dehydrogenase and
ubiquinone. Hydrogen peroxide is relatively unreactive with biomolecules but may
oxidize thiols and reduced glutathione (Farr and Kogoma, 1991). The bactericidal effects
of hydrogen peroxide are likely not due to the peroxide itself but the tendency of this
molecule to react with reduced metals, mainly ferrous iron, to form hydroxyl radicals
(Imlay et al., 1988).
The Hydroxyl Radical (HO )
The oxidative effects of O2" and H2O2 are relatively mild compared to those
caused by the hydroxyl radical. This species is capable of reacting with any biological
molecule at rates limited only by diffusion. It is believed that the cytotoxic effects of
superoxide and hydrogen peroxide are imparted by the participation in the formation of
intracellular hydroxyl radicals when hydrogen peroxide undergoes univalent reduction by
superoxide (Equation 3; Ascenzi, 1996).
O2 - + H2O2
> OH + OH' + O2
(Equation 3)
20
Hydroxyl radicals are also generated in the presence of ferrous iron acts as a catalyst in
radical formation from H2O2 (Equation 4; Imlay et al, 1988):
Fe2+ + H2O2 + H+
►
Fe3+ + HO' + H2O
(Equation 4)
It is also important to revisit the chemistry represented in Eq. 2 where superoxide
can reduce the ferric iron produced in Eq. 4 back to the ferrous species, allowing it to
participate in another Fenton reaction. Attempting to elucidate the cytotoxic mechanism
of hydrogen peroxide, Imlay et al. (1988) were able to generate severe DNA strand
breaks in an in vitro system that contained ferrous iron and H2O2. Also, the cytotoxic
effects of hydrogen peroxide could be eliminated when iron chelators were included with
cultures of exponentially growing E. coli undergoing H2O2 disinfection. Interestingly,
low doses ( 1 - 3 mM) of hydrogen peroxide were more effective at generating hydroxyl
radicals while higher concentrations seemed to inhibit Fenton chemistry. These authors
concluded that the primary killing mechanism of hydrogen peroxide was the generation
of hydroxyl radicals through Fenton reactions at or near the surface of DNA molecules. A
follow-up report by the same group demonstrated that NADH and NADPH could also
reduce ferric iron bound to DNA (Imlay and Linn, 1988). The reduced nucleotides would
then serve as a source of reductant to fuel ongoing Fenton reactions at the DNA surface,
which would cause strand breaks and damage individual nucleotides.
Oxidizing Biocides
ROIs and other oxidizing agents, when added to the external milieu, can be
powerful antimicrobial agents. Generally referred to as oxidizing biocides, these
21
compounds are known to be very powerful killers of planktonic organisms, but are often
inadequate to eliminate biofilm problems such as biofouling of cooling towers and heat
exchangers. Many oxidizing biocides are believed to generate intracellular hydroxyl
radicals and the ROI chemistry described above is likely to be involved in the killing
mechanisms possessed by many different compounds. Commonly used biocides that are
either known or hypothesized to involve oxygen radical generation will be reviewed
below.
Chlorine
Chlorine is still the most widely used biocide employed in industry since it can be
purchased as an inexpensive bulk commodity and is relatively easy to apply (Ascenzi,
1996). Chlorine can be administered as a compressed gas, neutralized salt solution such
as sodium hypochlorite (bleach), dry calcium salt form, or as organo-chlorides (Ascenzi,
1996). The compressed gas form is commonly employed in municipal water treatments
as well as industrial water process systems. Chlorine gas forms hydrochloric and
hypochlorous acid in water. The hypochlorous acid can further dissociate into the
hypochlorite ion which is ca. 80 times less effective than the acid species. Hypochlorous
acid is an electrophilic, strong oxidizing compound that can disrupt cell walls and
membranes at very low concentrations (Ascenzi, 1996). Thiol and amino groups of
proteins and other cellular constituents are also attacked.
Candelas et al. (1994) demonstrated that hydroxyl radicals could be generated in
vitro from hypochlorous acid and reduced iron by the following Fenton-type reaction
(Equation 5):
22
Fe2+ + HOCl
---------- ►
Fe3+ + HO' + Cl"
(Equation 5)
Similar to Eq. 3, superoxide and hypochlorous acid also produced hydroxyl radicals in
vitro by this reaction:
O2 ' + HOCl
^
OH' + Cl" + O2
(Equation 6)
Dukan and Touati (1996) investigated the mechanisms that may be responsible
for hypochlorous acid killing of E. coli. It was found that mutants lacking various genes
responsible for hydrogen peroxide resistance (see below) were more susceptible to HOCl
suggesting that HOCl toxicity did involve ROIs. Also,/hr mutants, which accumulate
elevated levels of iron, were also more susceptible to hypochlorous acid disinfection
which implies the generation of hydroxyl radicals via a Fenton-like reaction. Another
report (Dukan et al., 1996) ruled out the possibility that intracellular superoxide radicals
were generated by hypochlorous acid since strains lacking protection against superoxide
(sodA and sodB) were no more susceptible to HOCl than wild-type controls.
Ozone
Stricter environmental regulations are requiring non-toxic discharges from
industrial sources. The use of chlorinated biocides as a means to control biofouling
results in the formation of disinfection by-products. Disinfection by-products are created
when organic matter reacts with chlorine to form chlorinated organics such as
trihalomethane. The use of ozone (O3) as a disinfectant is gaining in popularity since it is
a strong oxidant, and therefore bactericidal, but does not generate chlorinated disinfection
23
byproducts (Kuo and Smith, 1996). O3 has been shown to be moderately effective
against suspended organisms and biofilms at concentrations ranging from 0.2 to 3.5 ppm
(Videla et ah, 1995; Lund and Ormerod, 1995) O3 is a highly reactive form of oxygen
which decomposes in pure water to form hydroxyl and superoxide radicals (Farr and
Kogoma, 1991). O3 will also generate intracellular ozonides (cyclic peroxides) and
hydrogen peroxide. Since ozone primarily generates ROIs, its killing mechanism relies
on that of the ROIs described above.
Hydrogen Peroxide
The external application of H2O2 serves as an effective biocide since it is a
relatively stable ROL H2O2 is often used as a disinfectant for medical and dental
equipment (Ascenzi, 1996) and is also gaining in popularity as an industrial and
municipal biocontrol agent since its use avoids the production of disinfection by-products
(Kuo and Smith, 1996). Christensen et al. (1990) demonstrated that sub-lethal doses of
H2O2, in the presence of ferrous iron, could induce biofilm detachment. These authors
cultured mixed Pseudomonas spp. biofilms in micro-annular reactors and exposed them
to 1.5 mM H2O2 and either 0.03 to 0.1 mM Fe2+ or 100 mM ascorbate. Ascorbate, like
ferrous iron, may participate in the univalent, single electron reduction of H2O2 to form
hydroxyl radicals. H2O2 treatments alone had no affect on biofilm removal although
when either reduced iron or ascorbate was added, as much as 90% of the biofilm biomass
was removed. These authors speculated that the peroxide was reacting with the reducing
agents to form hydroxyl radicals which in turn, were degrading the EPS matrix
surrounding the cells, causing the biofilm to slough. Interestingly, it was noted that after
24
disinfection, biofilm regrowth rapidly occurred and that the resulting bacterial population
was much more resistant to subsequent doses of peroxide and iron/ascorbate. A similar
result was reported by Sanderson and Stewart (1997) where P. aeruginosa biofilms
displayed resistance to subsequent doses of monochloramine of increasing concentration.
Since microorganisms have been exposed to ROIs for much of their evolution, resistance
or adaptation to oxidizing agents has naturally been a part of that evolutionary process.
In fact, microorganisms possess elaborate genetic and physiological oxidative stress
protection mechanisms which will now be reviewed.
■Mechanisms of Oxidative Stress Resistance
Driven by the presence of internal and extracellular ROIs, multi-layered
protection systems have evolved in bacteria. In addition to neutralizing the constant flux
of superoxide and hydrogen peroxide produced during aerobic metabolism,
microorganisms may also physiologically respond to extracellular sources of potentially
lethal ROIs. The genetics and physiology of oxidative stress in enteric bacteria, primarily
E. coli and S. Iyphimurium, has been studied for nearly 30 years. As a result of this
research, distinct responses to specific ROIs have been elucidated and patterns of global
gene regulation have emerged. Still, the function and regulation of many genes known to
be ROI sensitive remain unknown.
Superoxide and hydrogen peroxide each elicit distinct oxidative stress responses
in E. coli and S. typhimurium. This is due to the presence of cognate regulatory elements
present in the cell that are able to sense these oxidants and then transduce a signal that
initiates a physiological response. The physiological response includes the de novo
synthesis of a large number of proteins/enzymes that have some protective function.
These adaptive responses are a key survival strategy when killing by ROIs is eminent.
Alsoi oxidative stress responses appear to be integrated with other general, but very
complex, cellular adaptations to nutritional downshift conditions. Several genes with an
oxidant protective product are known to be induced as the cell begins to experience
nutrient starvation (Kjelleberg, 1993). The major adaptive responses to superoxide and
hydrogen peroxide, as well as the protective role of nutrient limited or stationary phase
physiology will be reviewed below. An emphasis will be placed on the hydrogen
peroxide response since this oxidant is generated through the elimination of superoxide
from the cell by the action of superoxide dismutases; a flux of superoxide results in a
hydrogen peroxide response as well (Demple, 1991). Most information regarding
bacterial oxidative stress has been obtained using enteric organisms. However, more
recently, the oxidative stress physiology of P. aeruginosa has been partially
characterized. Similarities and differences between the enteric models and P. aeruginosa
with respect to ROI defense systems will be highlighted.
Superoxide Response and SoxRS Regulon
When pre-treated with low doses of superoxide generating (redox cycling) drugs
such as paraquat, E. coli becomes increasingly resistant to subsequent doses of
superoxide generators presented at higher concentrations (Hassan and Fridovich, 1977).
This adaptive resistance is made possible via the induction of ca. 40 proteins as
determined by 2-dimensional gel analysis (Greenberg and Demple, 1989). At least nine
of these proteins are members of the soxRS regulon of superoxide inducible (soi) genes
26
Greenberg et al., 1990; Tsaneva and Weiss, 1990). The soxRS oxidative stress regulon is
controlled by two divergently transcribed genes soxR, and soxS (Wu and Weiss, 1991).
The SoxR protein contains redox sensitive [2Fe-2S] centers which upon oxidation,
activate this DNA binding protein, allowing it to enhance transcription of its only target
gene, soxS (Ding et al., 1996; Gaudu et al., 1997; Nunoshiba et al., 1992). SoxS belongs
to the AraC family of transcriptional activator proteins and induces transcription of
genes/operons included in the soxRS regulon. Therefore, the SoxR protein serves as the
sensor of superoxide stress and the SoxS protein transduces the signal to superoxide
adaptive response genes. Several genes included in the soxRS regulon have been
identified in E. coli. These include manganese-cofactored superoxide dismutase (sodA),
endonuclease IV info), glucose-6-phosphate dehydrogenase (zwf), NAD(P)Hdehydrogenase {ndh), heat shock protein GroEL (groEL), and a repressor of OmpF
synthesis (micF) (Farr and Kogoma, 1991; Greenberg and Demple, 1989).
Regulation of Superoxide Dismutases
Although all of the above mentioned genes are important to a superoxide adaptive
response, superoxide dismutases (SODs) serve as a first line of defense against
superoxide since these enzymes can directly scavenge superoxide from the intracellular
environment as shown in Equation 7.
2 0 2‘ + 2H+
__ H2O2 + O2
(Equation 7)
E. coli possesses both a manganese-cofactored superoxide dismutase (MnSOD) (Keele et
al., 1970) encoded by the sodA gene and an iron-cofactored superoxide dismutase
(FeSOD) (Yost and Fridovich, 1973) encoded by sodB. Carlioz and Touati (1986) found
that the aerobic growth rates of E. coli single mutants lacking either SOD were equal to
wild-type cells although the aerobic growth of double mutants in rich medium was
severely retarded. These authors also found that mutants lacking total SOD activity were
unable to grow in minimal media due to the inactivation of a, (3-dihydroxyisovalerate
dehydratase, a superoxide sensitive enzyme essential for leucine and isoleucine
biosynthesis.
SOD regulation in E. coli. SOD regulation is complex, displaying multilayer
control of MnSOD and FeSOD. FeSOD is constitutively expressed in anaerobically and
aerobically grown cells (Hassan and Fridovich, 1977), while MnSOD is not expressed
anaerobically but is inducible upon exposure to oxygen or superoxide-generating redox
cycling agents (Hassan and Fridovich, 1979). In addition to soxRS control, Tardat and
Touati (1991) showed that MnSOD is also regulated by the Fur (ferric uptake regulatory)
protein as well as the Arc regulatory system (aerobic respiratory control). Hassan and
Sun (1992) also observed multi-layered regulation of MnSOD by monitoring sodAr.lacZ
expression in several mutant backgrounds. These authors reported that under anaerobic
conditions, sodA is tightly repressed by the Fur, Arc, and Fnr (fumarate nitrate
reduction/regulator of anaerobic respiration) regulatory proteins, all of which are
independent of SoxRS regulation. Transcription of the sodB gene, on the other hand, is
positively regulated by the Fur protein and is negatively regulated by iron starvation
(Neiderhofferetal., 1990).
28
SOD regulation in P. aeruginosa. Like E. coli, P. aeruginosa possesses both an
iron (Steinman, 1985) and manganese (Hassett et al, 1992) cofactored superoxide
dismutase. Both P. aeruginosa SODs show high homology (50-67% deduced amino acid
sequence similarity) to their E. coli counterparts (Hassett et al., 1993). Similar to SOD
expression patterns in E. coli, P. aeruginosa FeSOD is expressed throughout the aerobic
growth cycle while MnSOD is tightly repressed. Hassett et al. (1996) found that P.
aeruginosa strains that expressed mutant Fur proteins, also possessed increased levels of
MnS OD, suggesting that Fur has a repressive function in P. aeruginosa similar to that in
E. coli. Interestingly, the same fur mutants also produce increased alginate levels
(Hassett et al., 1997). The addition of iron chelators such as 2,2’-dipyridyl to cultures of
P. aeruginosa increases the expression of MnSOD which further supports the regulatory
role of the Fur protein in P. aeruginosa MnSOD expression. Hassett et al. (1997) latter
reported that sodA is included in a four gene operon that contained two Fur binding sites
immediately upstream of the transcriptional start. Sequence analysis of the region
upstream of the sodB open reading frame suggests that this gene is regulated by a
promoter of the a 70 type (Hassett et al., 1993). As of yet, a superoxide sensing,
transcriptional activation system similar to the soxRS system in E. coli has not been
characterized in P. aeruginosa.
Recent studies conducted by Hassett et al., (submitted) have shown that quorum
sensing (for review see Fuqua et al., 1996) affects MnSOD expression in P. aeruginosa.
These authors observed that P. aeruginosa mutants that are unable to synthesize the
autoinducer N-Q-xoxdedecanoyl)-L-homoserine lactone (a.k.a. Pseudomonas
autoindercer-1; PAI-1) were also unable to express MnSOD in stationary phase cells.
29
Growth of these mutants in the presence of externally added PAI-I restored MnSOD
activity to near wild-type levels. As expected, quorum sensing mutants were also more
susceptible than the wild type strain to the superoxide generating redox cycling
compound paraquat.
Hydrogen Peroxide Response
Enteric bacteria are able to sense and respond to potentially lethal concentrations
of peroxide by inducing several genes that have products of some protective function.
For example, S. typhimurium treated with low doses (60 pM) of hydrogen peroxide for a
period of one hour induces the synthesis of 30 proteins (Christman et al., 1985; Morgan
et al., 1986) Of these, 12 (early proteins) are maximally induced during the first 30
minutes of exposure while the remaining 18 (late proteins) continue to be synthesized at
an elevated rate for the remainder of the dosing period. The induction of these genes
renders S. typhimurium resistant to subsequent doses of hydrogen peroxide at
concentrations of up to I OmM. A similar adaptive response is elicited by E. coli as
determined by 2-dimensional protein gel analysis (Farr and Kogoma, 1991; VanBogelen,
1987). Similar results were obtained in a study using randomly inserted Xp/acMu53
promoter probes showing sensitivity to hydrogen peroxide (Schellhom and Stones, 1992).
Similar to the soxRS system, a peroxide sensing system has been well characterized in E.
coli and S. typhimurium and is described below.
OxyR Regulation
Similar to superoxide adaptive responses, many H2O2 inducible genes and the
regulatory circuitry that governs their expression remain to he identified. A major
30
regulator of peroxide stress in S. typhimurium and E. coli was identified when Christman
et al. (1985) subjected S. typhimurium to chemical mutagenesis and selected mutants that
demonstrated increased resistance to hydrogen peroxide without prior exposure to the
oxidant. Two-dimensional protein gel analysis revealed that nine of the twelve ‘early’
proteins were constitutively overexpressed in a mutant strain designated oxyRl. A Tn5
insertion mutant that expressed the same phenotype as oxyRl allowed identification of
the regulatory component controlling the overexpressed genes. Designated oxyR, the
protein encoded by this gene was determined to be a positive transcriptional activator of a
regulon which included hydrogen peroxide stress inducible genes as well as three heat
shock proteins. Both the S. typhimurium (Christman et al., 1989) and E. coli (Tao et al.,
1989) oxyR homologues were later identified as members of a large class of bacterial
regulatory proteins known as the LysR family of positive transcriptional activators.
OxyR homologues have also been identified in Mycobacterium spp. (Deretic et al., 1995)
and Xanthomonas campestris (Mongkolsuk et al., 1998).
The mechanism allowing transcriptional activation of hydrogen peroxide
inducible genes OxyR has been elucidated (Kullik et al., 1995). When a cysteine residue
at position 199 becomes oxidized by H2O2, OxyR is conformationally changed such that
it will now bind to its target promoter sequences. Toledano et al. (1994) employed a
series of synthetic binding sequences to determine that the oxidized OxyR protein
recognizes a binding motif comprised of four ATAGnt elements spaced at 10 bp intervals
along the DNA helix. Oxidized OxyR, once bound to its target sequence, allows contact
of the target nucleotide sequence with the a subunit of RNA polymerase, which then
initiates transcription of the affected gene (Tao et al., 1995). In contrast, reduced OxyR
31
on the other hand, binds to only two ATAGnt elements which enables it to act as a
repressor by blocking transcription of the target gene. Like other LysR transcriptional
regulators, OxyR autoregulates its own expression during normal growth by repressing
oxyR (Christman et ah, 198.9; Tao et ah, 1991).
OxyR regulates the synthesis of several genes that participate in the protection
against oxidative stress as well as the maintenance of homeostatic, intracellular H2O2
concentrations (Gonzalez-Flecha and Demple, 1997) Initially, JcatG (encoding HPI
catalase), ahpCF (encoding an alkyl hydroperoxide reductase), dps (encoding a
nonspecific DNA-binding protein), gorA (encoding glutathione reductase), and oxyS
(encoding a small untranslated, regulatory RNA) were identified as being under OxyR
control (Altuvia et al., 1994; Christman et ah, 1985; Storz et ah, 1990). Mukhopadhyay
and Schellhom (1997) tested previously constructed libraries of hydrogen peroxidesensitive (hps) XpZacMuS3 promoter probes (Schellhom and Stones, 1992) to screen for
genes controlled by OxyR. By transducing the library into OxyRir and AoxyR E. coli
backgrounds and comparing IacZ reporter activities, these authors reported that 8 of the
hps phenotypes were independent of OxyR, 10 alleles required OxyR for activation, and
6 fusions were always repressed in the presence of oxidized or reduced OxyR. Using
sequence information obtained from the reporter junctions, several previously unknown
genes activated by OxyR were identified, including rcsC, which encodes a sensorregulator protein of capsular polysaccharide synthesis genes.
Regulation of Catalase in E. coli
With a role similar to superoxide dismutase, catalase (a.k.a. hydrpperoxidase) is
32
an essential component of a hydrogen peroxide adaptive response (Schellhorn, 1995).
Catalases are heme containing enzymes that catalyze the disproportionation of hydrogen
peroxide into the products oxygen and water as shown by Equation 8.
2H2O2
— catalase
^
2H2O + O2
(Equation 8)
The importance of catalase in the protection of bacteria from H2O2 has been
repeatedly demonstrated with the use of catalase mutants (Ma and Eaton, 1992; Brown et
ah, 1995; Rocha et ah, 1996) and with regulatory mutants unable to synthesize wild type
levels of catalase (Christman et ah 1985; Seymour et ah 1996).
The best model for the regulation of catalases has been constructed from work
done on E. coli, which expresses two distinct inducible catalases termed HPI and HPII
(Claiborne et ah, 1979; Hassan and Fridovich, 1978). HPI, encoded by katG, is a
tetramer with subunits of 80 kDa in molecular mass. The HPII holoenzyme is a hexamer
with subunits of 93 kDa. HPII is also able to scavenge peroxide at a much high rate with
a Vmax of 8924 pmol min'1mg ' enzyme compared to 1496 pmol min ' mg"' enzyme for
HPh Unlike HPII, HPI possesses a peroxidase activity in addition to its catalase activity
that catalyses the reduction of hydrogen peroxide by oxidizing an intracellular reductant
such as NAD(P)H (Loewen et ah, 1985).
Regulation of katG in E. coli. During exponential growth, HPI composes only a
small fraction of the total catalase activity expressed by E. coli', however, this catalase is
the most important determinant for hydrogen peroxide resistance (Schellhorn, 1995).
HPI has been shown to have several protective functions including recovery of proton
33
Exponential Phase
Oxidative Stress
I
H 2O 2
OxyR
(reduced/inactive)
OxyR
(oxidized/active)
9 other proteins
katG
I
Figure 2-1. Induction of HPI during exponential growth by the OxyR system
(Adapted from Schellhorn, 1995).
34
motive force following oxidative attack (Farr et al., 1988). As mentioned above, HPI is a
member of the OxyR regulon and is therefore, highly inducible upon exposure to sublethal levels of hydrogen peroxide (Christman et al., 1985) (See Fig. 2-1). Transcription
of katG has been shown to increase 100-fold when oxidized OxyR protein is present
(Storz et al. 1990). This inductive capability makes HPI expression a marker enzyme for
the hydrogen peroxide adaptive response in E. coli. The rapid induction of HPI due to
exposure to hydrogen peroxide has been characterized with exponentially growing cells;
however, little is known about the role of OxyR activation in nutrient limited cultures.
In the absence of hydrogen peroxide, HPI is also constitutively expressed at low levels
during normal aerobic and anaerobic growth. HPI activities have also been shown to
dramatically increase when cultures of A. coli reach stationary phase. This induction in
HPI activity during nutrient limiting conditions is controlled by another layer of catalase
regulation in E. coli which is also the primary regulatory mechanism responsible for HPII
synthesis (Ivanova et al., 1994) (See Fig. 2-2).
Regulation of katE in E. coli. HPII, encoded by katE, constitutes the largest
fraction of catalase activity in aerobically grown E. coli and it is well known that catalase
levels increase 10-20 fold as cultures of E. coli enter stationary phase. HPII is not
inducible by hydrogen peroxide and transcription of this catalase is essentially the same
in a AoxyR mutant as it is in wild type cells suggesting that katE regulation is independent
of OxyR (Schellhom and Hassan, 1988). By screening E. coli TnfO insertion mutants
that displayed a constitutive catalase activity phenotype, Loewen and Triggs, (1984) were
able to identify a regulatory element that was responsible for increased stationary phase
35
Stationary Phase
Starvation
------ ► ribosomal stalling
ReIA
ATP + GTP
PpGpp + AMP, Pi
stringent induction of
biosynthetic operons
homoserine
-<
RspA
homoserine lactone
I
induction of rpoS
weak acids
30 other proteins
katG
i
i
HPII
HPI
Figure 2-2. Model of catalase regulation in stationary phase E. coli. (Adapted from
Schellhom, 1995)
36
catalase production. The regulatory element, termed KatF, was latter identified as
an alternative RNA polymerase sigma factor now commonly referred to as a 38 or RpoS
(encoded by the rpoS gene) (see Loewen and Hengge-Aronis, 1994 for review). As cells
enter stationary phase, RpoS increases the synthesis of more that 30 proteins in
E. coli including HPII, exonuclease III, glycogen synthetase, trehalose synthesis, acid
phosphatase, and bacterial virulence factors (Loewen and Hengge-Aronis, 1994).
The regulation of rpoS is complex and RpoS itself is subject to transcriptional,
translational, and post-translational regulation. RpoS expression has been shown to be
influenced by stringent response inducing metabolites such as guanosine tetraphosphate
(ppGpp) (Gentry et al., 1993) as well as cell density dependent, homoserine lactone
mediated signaling pathways (Huisman and Kolter, 1994).
Regulation of Catalase in P. aeruginosa
Compared to E. coli, relatively little is known about the hydrogen peroxide
adaptive response and regulation of catalase in P. aeruginosa. The work of Hassett and
co-workers (Brown et al., 1995; Hassett et al., 1992; Hassett et al., 1996) has uncovered
many similarities in the overall peroxide stress response and catalase expression patterns
in P. aeruginosa although evidence also suggests likely differences. In defense against
hydrogen peroxide, P. aeruginosa expresses two major catalases designated KatA and
KatB (Hassett et al., 1992). Under certain conditions, a third catalase, KatC, may also be
' expressed at low levels (Fredricks, unpublished observations). Similar to HPII in E. coli,
KatA constitutes the major fraction of total catalase activity in P. aeruginosa and is
expressed during the entire aerobic growth cycle. Also like HPII, KatA expression
37
increases markedly at the onset of stationary phase. P. aeruginosa possesses an rpoS
homologue which, as in E. coli, directs the synthesis of many stationary phase inducible
genes (Fujita et al., 1994; Tanaka and Takahashi, 1994) Although direct evidence has not
been provided thus far, KatA is likely under RpoS control much the same as HPII
synthesis is in E. coli (Hassett, 1998). Indirect evidence suggesting RpoS regulation of
KatA was recently discovered by Hassett et al. (submitted) by showing that P.
aeruginosa mutants unable to synthesize cell-to-cell signaling molecules, especially PAI1, express nearly 2-fold less catalase in stationary phase compared to wild-type
organisms. A katAv.lacZ transcriptional fusion was used to confirm that the decrease in
total catalase activity could be attributed to sub-optimal KatA expression. Since quorum
sensing is known to positively influence RpoS expression in P. aeruginosa (Latifi et al.,
1996), it is possible that the observed decrease in katA transcription was due to restricted
Rpo S-mediated stationary phase gene expression resulting from the lack of quorum
sensing ability in the mutants. Catalase activity in Rhizobium leguminosarum bv.
phaseoli, a Gram-negative bacterium related to P. aeruginosa, is also affected by cell
density-dependent gene regulation (Crockford et al., 1995). Iron also affects the
regulation of KatA in P. aeruginosa since Fur mutants of this organism express roughly
2-fold less catalase than Fur+cells and the addition of iron chelators such as 2,2’dipyridyl severely decrease total catalase activities (Hassett et al., 1996). It is not known,
however, whether iron limitation or defective Fur binding affects KatA at the
transcriptional or post-translational level.
P. aeruginosa KatB is typically not expressed during the aerobic growth cycle of
P. aeruginosa but is highly inducible upon exposure to sub-lethal doses of hydrogen
38
peroxide (Brown et al., 1995). Although the regulation of KatB is currently unknown, P.
aeruginosa must possess some system that is able to sense peroxide attack and transduce
a signal to oxidative stress sensitive genes like katB. Like the E. coli katG loci, katB
transcription increases in response to peroxide. Unlike HPI, however, KatB activity is
virtually silent in the absence of H2O2. Again, iron availability and the Fur system likely
play a role in KatB regulation since P. aeruginosa Fur mutants constitutively express
KatB in the absence of inducing agents (Hassett et ah, 1996). This suggests the regulation
of katB by a redox sensitive transcriptional repression system (i.e. Fur protein) rather that
a transcriptional activating element like OxyR.
Summary
Biofilms undoubtedly affect human health and prosperity by creating chronic
infections in medical situations and retarding industrial system performance. Much effort
has been focused on improving methods of controlling microbial biofilms either by
preventing their formation or by eliminating them with antimicrobial agents. Since most
existing antimicrobial control strategies have been developed using planktonic organisms,
new challenges exist in developing methods of killing highly resistant biofilms. In order
to develop effective biofilm management strategies, the underlying mechanisms
responsible for biofilm resistance to antimicrobial agents must be understood. Unifying
hallmarks of biofilm structure and physiology such as EPS encapsulation and retarded
growth rates have been implicated in biofilm resistance, but these attributes are
sometimes inadequate for explaining why biofilms are less susceptible to antimicrobial
agents.
39
Microorganisms are extremely ‘talented’ in their capacity to adapt to adverse
environmental conditions. Since many toxic compounds (or at least structural analogs)
used to control biofilms are found in nature, bacteria are likely to have evolved
mechanisms to effectively deal with them. Toxic oxygen derivatives such as hydrogen
peroxide have existed through much of biological evolution and therefore
microorganisms possess effective physiological weapons for their defense. Ironically,
toxic oxygen derivatives are often applied in large quantities to control biofilms. Much
work has focused on elucidating oxygen chemistry in living systems as well as the
microbial genetics and physiology that are associated with oxygen respiration and
oxidative stress. It is well known that microorganisms can sense and adapt to oxidizing
agents but the role of these adaptive responses in protecting biofilms from oxidizing
biocides is poorly understood. Determining the importance of oxidative stress
physiology in biofilm bacteria will broaden our understanding of biofilms as whole and
lead to more effective strategies for controlling biofilms.
40
References
Altuvia, S., M. Almiron, G. Huisman, R. Kolter, and G. Storz. 1994, The dps
promoter is activated by OxyR during growth and by IHF and crs in stationary phase.
Mol. Microbiol. 13:265-272.
Anwar, H., M. K. Dasgupta, and J. W. Costerton. 1990. Testing the susceptibility of
bacteria in biofilms to antibacterial agents. Antimicrob. Agents Chemother. 34:20432046.
Ascenzi, J. M. 1996. Handbook of Disinfectants and Antiseptics. Marcel Dekker, Inc.
New York, New York.
Babior, B. M. 1988. The respiratory burst oxidase. Basic Life Sci. 49:815-821.
Berman, D., E. W. Rice, and J. C. Hoff. 1988 . Inactivation of particle-associated
coliforms by chlorine and monochloramine. AppL Environ. Microbiol. 54:507-512.
Brown, M. L., H. C. Aldrich, and J. J. Gauthier. 1995. Relationship between
glycocalyx and povidone-iodine resistance in Pseudomonas aeruginosa (ATCC 27853)
biofilms. AppL Environ. Microbiol. 61:187-193.
Brown, M. R. W., P. J. Collier, and P. Gilbert. 1990. Influence of growth rate on
susceptibility to antimicrobial agents: modification of the cell envelope and batch and
continuous culture studies. Antimicrob. Agents Chemother. 34:1623-1628.
Brown, M. L., and J. J. Gauthier. 1993. Cell density and growth phase as factors in the
resistance of a biofilm of Pseudomonas aeruginosa (ATCC 27853) to iodine. AppL
Environ. Microbiol. 59:2320-2322.
Brown, M. R. W., and P. Gilbert. 1993. Sensitivity of biofilm to antimicrobial agents. J.
AppL BacterioL Symp. SuppL 74:87S-97S
Brown, S. M., M. L. Howell, M. L. Vasil, A. J. Anderson, and D. J. Hassett. 1995.
Cloning and characterization of the katB gene of Pseudomonas aeruginosa encoding a
hydrogen peroxide-inducible catalase: purification of KatB, cellular localization, and
demonstration that it is essential for optimal resistance to hydrogen peroxide. J. BacterioL
177:6536-6544.
Carlioz, A., and D. Touati. 1986. Isolation of superoxide dismutase mutants in
Escherichia coir, is superoxide dismutase necessary for aerobic life? EMB0. 5:623-630.
41
Candelas, L. P., M. R. L. Stratford, and P. Wardman. 1994. Formation of hydroxyl
radicals on reaction of hypochlorous acid with ferrocyanide, a model Iron(II) complex.
Free Radical Res. 20:241-249.
Chapman, J. S., M. A. Diehl, and R. C. Lyman. 1993. Biocide susceptibility and
intracellular glutathione in Escherichia coli. J. Ind. Microbiol. 12:403-407.
Chesney, J. A., J. W. Eaton, and J. R. Mahoney, Jr. 1996. Bacterial, glutathione: a
sacrificial defense against chlorine compounds. J. Bacteriol. 178:2131-2135.
Chen, X., and P. S. Stewart. 1996. Chlorine penetration into artificial biofilm is limited
by a reaction-diffusion interaction. Environ. Sci. Technoh 30:2078-2083.
Christensen, B. E., H. N. Tronnes, K. Vollan, O. Smidsrod, and R. Bakke. 1990.
Biofilm removal by low concentrations of hydrogen peroxide. Biofouling 2:165-175.
Christman, M. F., R. W. Morgan, F. S. Jacobson, and B. N. Ames. 1985. Positive
control of a regulon for defenses against oxidative stress and some heat-shock proteins in
Salmonella typhimurium. Cell. 41:753-762.
Christman, M. F., G. Storz, and B. N. Ames. 1989. OxyR, a positive regulator of
hydrogen peroxide-inducible genes in Escherichia coli and Salmonella typhimurium, is
homologous to a family of bacterial regulatory proteins. Proc. Natl. Acad. Sci- 86:34843488.
Claiborne, A., D. P. Malinowski, and I. Fridovich. 1979. Purification and
characterization of hydroperoxidase II of Escherichia coli B. J. Biol. Chem. 254:1166411668.
Costerton, J. W., R. T. Irvin, and K.-J. Cheng. 1981. The bacterial glycocalyx in
nature and disease. Annu. Rev. Microbiol. 35:299-324.
Costerton, J. W., Z. Lewandowski, D. E. Caldwell, D. R. Korber, and H. M. LappinScott. 1995. Microbial biofilms. Annu. Rev. Microbiol. 49:711-745.
Crockford, A. J., G. A. Davis, and H. D. Williams. 1995. Evidence for cell-densitydependent regulation of catalase activity in Rhizobium leguminosarum bv. phaseoli.
Microbiology. 141:843-851.
Davies D. G., A. M. Chakrabarty, and G. G. Geesey. 1993. Exopolysaccharide
production in biofilms: substratum activation of alginate gene expression by
Pseudomonas aeruginosa. Apph Environ. Microbiol. 59:1181-1186.
)
42
Davies D. G., and G. G. Geesey. 1995. Regulation of the alginate biosynthesis gene
algC in Pseudomonas aeruginosa during biofilm development in continuous culture.
Appl. Environ. Microbiol. 61:860-867.
de Beer, D., R. Srinivasan, and P. S. Stewart. 1994. Direct measurement of chlorine
penetration into biofilms during disinfection. Appl. Environ. Microbiol. 60:4339-4344.
Demple B. 1991. Regulation of bacterial oxidative stress genes. Annu. Rev. Genet.
25:315-337.
Deretic, V., W. Philipp, S. Dhandayuthapani, M. H. Mudd, R. Curcic, T. Garbe, B.
Heym, L. E. Via, and S. T. Cole. 1995. Mycobacterium tuberculosis is a natural mutant
with an inactivated oxidative-stress regulatory gene: implications for sensitivity to
isoniazid. Mol. Microbiol. 17:889-900.
Ding, H., E. Hidalgo, and B. Demple. 1996. The redox state of the [2Fe-2S] clusters in
the SoxR protein regulates its activity as a transcription factor. J. Biol. Chem. 271:3317333175.
Dukan, S., S. Dadon, D. R. Smulski, and S. Belkin. 1996. Hypochlorous acid activates
the heat shock and soxRS systems of Escherichia coli. Appl. Environ. Microbiol.
62:4003-4008.
Dukan, S., and D. Touati. 1996. Hypochlorous acid stress m. Escherichia coir.
resistance, DNA damage, and comparison with hydrogen peroxide stress. J. Bacteriol.
178:6145-6150.
Embleton, J. V., H. N. Newman, and M. Wilson. 1998. Influence of growth mode and
sucrose on susceptibility of streptococcus sanguis to amine fluorides and amine fluorideinorganic fluoride combinations. Appl. Environ. Microbiol. 64:3503-3506.
Evans, E., M. R. Brown, and P. Gilbert. 1994. Iron chelator, exopolysaccharide and
protease production in Staphylococcus epidermidis: a comparative study of the effects of
specific growth rate in biofilm and planktonic culture. Microbiology. 140:153-157.
Fujita, M., K. Tanaka, H. Takahashi, and A. Amemura. 1994. Transcription of the
principal sigma-factor genes, rpoD and rpoS, in Pseudomonas aeruginosa is controlled
according to the growth phase. Mol. Microbiol. 13:1071-1077.
F a rr S. B., and T. Kogoma. 1991. Oxidative stress responses in Escherichia coli and
Salmonella typhimurium. Microbiol. Rev. 55:561-585.
F arr S. B., D. Touati, and T. Kogoma. 1988. Effects of oxygen stress on membrane
functions in Escherichia coli: role of HPI catalase. J. Bacteriol. 170:1837-1842.
43
Fridovich, I. 1978. The biology of oxygen radicals. Science 201:875-880.
Fuqua, W. C., S. C. Winans, and E. P. Greenberg. 1994. Quorum sensing in bacteria:
the LuxR-LuxI family of cell density-responsive transcriptional regulators. J. Bacteriol.
176:269-275.
Gaudu, P., N. Noon, and B. Weiss. 1997. Regulation of the soxRS oxidative stress
regulon: reversible oxidation of the Re-S centers of SoxR in vivo.. J. Biol. Chem.
272:5082-5086.
Gentry, D. R., V. J. Hernandez, L. H. Nuygen, D. B. Jensen, and M. Cashel. 1993.
Synthesis of the stationary phase sigma factor a s is positively regulated by ppGpp. J.
Bacteriol. 175:7982-7989.
Giwercman, B., E. T. Jensen, N. Hoiby, A. Kharazmi, and J. W. Costerton. 1991.
Induction of the (3-lactamase production in Pseudomonas aeruginosa biofilm.
Antimicrob. Agents Chemother. 35:1008-1010.
Greenberg, J. T., and B. Demple. 1989. A global response induced in Escherichia coli
by redox-cycling agents overlaps with that induced by peroxide stress. J. Bacteriol.
171:3933-3939.
Greenberg J. T., P. A. Monach, J. H. Chou, P. D. Josephy, and B. Demple. 1990.
Positive control of a global antioxidant defense regulon activated by superoxide­
generating agents in Escherichia coli. Proc. Natl. Acad. Sci. 87:6181-6185.
Gristina, A. G., R. A. Jennings, P. T. Naylor, Q. N. Myrvik, and L. X. Webb. 1989.
comparative in vitro antibiotic resistance of surface-colonizing coagulase-negative
staphylococci. Antimicrob. Agents Chemother. 33:813-816.
Gonzalez-Flecha, B., and B. Demple. 1997. Homeostatic regulation of intracellular
hydrogen peroxide concentration in aerobically growing Escherichia coli. J. Bacteriol.
179:382-388.
Gonzalez-Flecha, B., and B. Demple. 1995. Metabolic sources of hydrogen peroxide in
aerobically growing Escherichia coli. J. Biol. Chem. 270:13681-13687.
Hassan, H. M., and I. Fridovich. 1977. Regulation of the synthesis of superoxide
dismutase in Escherichia coil. J. Biol. Chem. 252:7667-7672..
Hassan, H. M., and I. Fridovich. 1978. Regulation of the synthesis of catalase and
peroxidase in Escherichia coli. J. Biol. Chem. 253:6445-6450.
44
Hassan, H. M., and H. C. Sun. 1992. Regulatory roles of Fur, Fur, and Arc in
expression of manganese-containing superoxide dismutase in Escherichia coli. Proc.
Natl. Acad. Sci. 89:3217-3221.
Hassett, D. J. 1998. Personal communication.
Hassett, D. J., L. Charniga, K. Bean, B. E. Ohman, and M. S. Cohen. 1992. Response
of Pseudomonas aeruginosa to pyocyanin: mechanisms of resistance, antioxidant
defenses, and demonstration of a manganese-cofactored superoxide dismutase. Infect.
Immun. 60:328-336.
Hassett, D. J., M. L. Howell, U. A. Ochsner, M. L. Vasil, Z. Johnson, and G. E.
Dean. 1997. An operon containing/wmC and sodA encoding lumarase C and manganese
superoxide dismutase is controlled by the ferric uptake regulator in Pseudomonas
aeruginosa: fur mutants produce elevated alginate levels. J. Bacteriol. 179:1452-1459.
Hassett, D. J., J.-F. Ma, J. G. Elkins, T. R. McDermott, U. A. Ochsner, S. E. H.
West, C.-T. Huang, J. Fredericks, S. Burnett, P. S. Stewart, G. McFeters, L.
Passador, and B. H. Iglewski. 1999. Quorum sensing in Pseudomonas aeruginosa
controls catalase and superoxide dismutase genes and mediates biofilm susceptibility to
hydrogen peroxide. J. Bacteriol. Submitted.
Hassett, D. J., P. A. Sokol, M. L. Howell, J.-F. Ma, H. T. Schweizer, U. Ochsner, and
M. L. Vasil. 1996. Ferric uptake regulator (Fur) mutants of Pseudomonas aeruginosa
demonstrate defective siderophore-mediated iron uptake, altered aerobic growth, and
decreased superoxide dismutase and catalase activities. J. Bacteriol. 178:3996-4003.
Hassett, D. J., W. A. Woodruff, D. J. Wozniak, M. L. Vasil, M. S. Cohen, and D. E.
Ohman. 1993. Cloning and characterization of the Pseudomonas aeruginosa sodA and
sodB genes encoding manganese- and iron-cofactored superoxide dismutase:
demonstration of increased manganese superoxide dismutase activity in alginateproducing bacteria. J. Bacteriol. 175:7658-7665.
Huang, C.-T., K. D. Xu, G. A. McFeters, and P. A. Stewart. 1998. Spatial patterns of
alkaline phosphatase expression within bacterial colonies and biofilm in response to
phosphate starvation. AppL Environ. Microbiol. 64:1526-1531.
Huang, C.-T., F. P. Yu, G. A. McFeters, and P. S. Stewart. 1995. Nonuniform spatial
patterns of respiratory activity within biofilms during disinfection. AppL Environ.
Microbiol. 61:2252-2256.
Huisman, G. W., and R. Kolter. 1994. Sensing starvation: a homoserine lactonedependent signaling pathway m. Escherichia coli. Science. 265:537-539.
45
Imlay, J. A., S. M. Chin, and S. Linn. 1988. Toxic DNA damage by hydrogen peroxide
through the Fenton reaction in vivo and in vitro. Science. 240:640-642.
Imlay, J. A., and S. Linn. 1988. DNA damage and oxygen radical toxicity. Science.
240:1302-13089.
Imlay, J. A., and I. Fridovich. 1991. Assay of metabolic superoxide production in
Escherichia coli. J. Biol. Chem. 266:6957-6965.
Ivanova, A., C. Miller, G. Glinsky, and A. Eisenstark. 1994. Role of rpoS (katF) in
oxyR-independent regulation of hydroperoxidase I in Escherichia coli. Mol. Microbiol.
12:571-578.
Keele, B. B., Jr., J. M. McCord, and I. Fridovich. 1970. Superoxide dismutase from
Escherichia coli B. A new manganese containing enzyme. J. Biol. Chem. 245:61766181.
Kinniment, S. L., and J. W. T. Wimpenny. 1992. Measurements of the distribution of
adenylate concentrations and adenylate energy charge across Pseudomonas aeruginosa
biofilms. Appl. Environ. Microbiol. 58:1629-1635.
Kjelleberg, S. 1993. Starvation in Bacteria. Plenum Press. New York, New York.
Kullik, I., J. Stevens, M. B. Toledano, and G. Storz. 1995. Mutational analysis of the
redox-sensitive transcriptional regulator OxyR: regions important for DNA binding and
multimerization. J. Bacteriol. 177:1285-1291.
Kullik, I., M. B. Toledano, L. A. Tartaglia, and G. Storz. 1995. Mutational analysis of
the redox-sensitive transcriptional regulator OxyR: regions important for oxidation and
transcriptional activation. J. Bacteriol. 177:1275-1284.
Kuo, J.-F., S. O. Smith. 1996. Disinfection. Wat. Environ. Res. 68:503-510.
Latifi, A., M. Foglino, K. Tanaka, P. Williams, and A. LazdunskL 1996. A
hierarchical quorum-sensing cascade in Pseudomonas aeruginosa links the
transcriptional activators LasR and RhlR (VsmR) to expression of the stationary-phase
sigma factor RpoS. Mol. Microbiol. 21:1137-1146.
Lechevallier M. W., C. D. Cawthon, and R. G. Lee. 1988. Inactivation of biofilm
bacteria. Appl. Environ. Microbiol. 54:2492-2499.
Loewen, P. C., and R. Hengge-Aronis. 1994. The role of the sigma factor Gs (KatF) in
bacterial global regulation. Annu. Rev. Microbiol. 48:53-80.
46
Loewen, J. C., and B. L. Triggs. 1984. Genetic mapping of katF, a locus that the katE
affects the synthesis of a second catalase species in Escherichia coli. J. BacterioL
160:668-675.
Lund, V., and K. Ormerod. 1995. The influence of disinfection processes on biofilm
formation in water distribution systems. Wat. Res. 29:1013-1021.
Ma, M., and J. W. Eaton. 1992. Multicellular oxidant defense in unicellular organisms.
Proc. Natl. Acad. Sci. 89:7924-7928.
Mongkolsuk, S., R. Sukchawalit, S. Loprasert, W. Praituan, and A. Upaichit 1998.
Construction and physiological analysis of Xanthomonas mutant to examine the role of
the oxyR gene in oxidant-induced protection against peroxide killing. J. Bacteriol.
180:3988-3991.
Morgan, R. W., M. F. Christman, F. S. Jacobson, G. Storz, and B. N. Ames. 1986.
Hydrogen peroxide-inducible proteins in Salmonella typhimurium overlap with heat
shock and other stress proteins. Proc. Natl. Acad. Sci. 83:8059-8063. ■
Mukhopadhyay, S., and H. B. ScheIIhorn. 1997. Identification and characterization of
hydrogen peroxide-sensitive mutants of Escherichia coli: genes that require OxyR for
expression. J. Bacteriol. 179:330-338.
Niederhoffer, E. C., C. M. Naranjo, K. L. Bradley, and J. A. Fee. 1990. Control of
Escherichia coli superoxide dismutase (sodA and sodB) genes by the ferric uptake
regulation (fur) locus. J. Bacteriol. 172:1930-1938.
Nichols, W. W., M. J. Evans, M. P. E. Slack, and H. L. Walmsley. 1989. The
penetration of antibiotics into aggregates of mucoid and non-mucoid Pseudomonas
aeruginosa. J. Gen. Microbiol. 135:1291-1303.
Nunoshiba, T., E. Hidalgo, C. F. A. Cuevas, and B. Bemple. 1992. Two-stage control
of an oxidative stress regulon: the Escherichia coli SoxR protein triggers redox-inducible
expression of the soxS regulatory gene. J. Bacteriol. 174:6054-6060.
Rocha, E. R., T. Selby, J. P. Coleman, and C. J. Smith. 1996. Oxidative stress
response in an anaerobe, Bacteriodes fragilis: a role for catalase in protection against
hydrogen peroxide. 178:6895-6903.
Rosner, J. L. 1993. Susceptibilities of oxyR regulon mutants of Escherichia coli and
Salmonella typhimurium to isoniazid. Antimicrob. Agents. Chemother. 37:2251-2253.
Sanderson, S. S., and P. S. Stewart. 1997. Evidence of bacterial adaptation to
monochloramine in Pseudomonas aeruginosa biofilms and evaluation of biocide action
model. Biotechnol. Bioeng. 56:201-209.
47
Schellhorn, H. E. 1995. Regulation of hydroperoxidase (catalase) expression in
Escherichia coli. FEMS Microbiol. Lett. 131:113-119.
Schellhorn, H. E., and H. M. Hassan. 1988. Transcriptional regulation of katE in
Escherichia coli K-12. J. Bacteriol. 170:4286-4292.
Schellhorn, H. E., and V. L. Stones. 1992. Regulation of katF and katE in Escherichia
coli K-12 by weak acids. J. Bacteriol. 174:4769-4776.
Seymour, R. L., P. V. Mishra, M. A. Khan, and M. P. Spector. 1996. Essential roles
of core starvation-stress response loci in carbon-starvation-inducible cross-resistance and
hydrogen peroxide-inducible adaptive resistance to oxidative challenge in Salmonella
typhimurium. Mol. Microbiol. 20:497-505.
Stewart, P. S. 1996. Theoretical aspects of antibiotic diffusion into microbial biofilms.
Antimicrob. Agents Chemother. 40:2517-2522.
Storz, G., L. A. Tartaglia, and B. N. Ames. 1990. Transcriptional regulator of
Oxidative stress-inducible genes: direct activation by oxidation. Science. 248:189-194.
Steinman, H. M. 1985. Bacteriocuprein superoxide dismutases in Pseudomonads. J.
Bacteriol. 162:1255-1260.
Tamarit, J., E. Cabiscol, and J. Ros. 1998. Identification of the major oxidatively
damaged proteins in Escherichia coli cells exposed to oxidative stress. J. Biol. Chem.
273:3027-3032.
Tanaka, K., and H. Takahashi. 1994. Cloning, analysis, and expression of an rpoS
homologue gene from. Pseudomonas aeruginosa PAOI. Gene. 150:81-85.
Tao, K., K. Makino, S. Yonei, A. Nakata, and H. Shinahawa. 1989. Molecular cloning
and nucleotide sequencing of oxyR, the positive regulatory gene of a regulon for an
adaptive response to oxidative stress in Escherichia coli: homologies between OxyR
protein and a family of bacterial activator proteins. Mol. Gen. Genet. 218:371-376.
Tao, K., K. Makino, S. Yonei, A. Nakata, and H. Shinahawa. 1991. Purification and
characterization of the Escherichia coli OxyR protein, the positive regulator for a
hydrogen peroxide-inducible regulon. J. Biochem. 109:262-266.
Tao, K., C. Zon, N. Fujita, and A. Ishihama. 1995. Mapping of the OxyR protein
contact site in the c-terminal region of RNA polymerase a subunit. J. Bacteriol.
177:6740-6744.
48
Toledano, M. B., I. Kullik, F. Trinh, P. T. Baird, T. B. Schneider, and G. Storz.
1994. Redox-dependent shift of OxyR-DNA contacts along an extended DNA-binding
site: a mechanism for differential promoter selection. Cell. 78:897-909.
Tsaneva, L. R., and B. Weiss. 1990. soxR, a locus governing a superoxide response
regulon'm Escherichia coli K-12. J. Bacteriol. 170:4415-4419.
VanBogelen, R. A., P. M. Kelley, and F. C. Neidhardt. 1987. Differential induction of
heat shock, SOS, and oxidative stress regulons and accumulation of nucleotides in
Escherichia coli. J. Bacteriol. 169:26-32.
Videla H. A., M. R. Viera, P. S. Guiamet, and J. C. S. Alais. 1995. Using ozone to
control biofilms. Materials Performance. 124:40-44. ■
Williams, I., W. A. Venables, D. Lloyd, F. Paul, and I. Critchley. 1997. The effects of
adherence to silicone surfaces on antibiotic susceptibility in Staphylococcus aureus.
Microbiology. 143:2407-2413.
Wu, J., and B. Weiss. 1991. Two divergently transcribed genes, soxR and soxS, control a
superoxide response regulon of Escherichia coli. J. Bacteriol. 173:2864-2871.
Xu, X., P. S. Stewart, and X. Chen. 1996. Transport limitation of chlorine disinfection
of Pseudomonas aeruginosa entrapped in alginate beads. Biotechnol. Bioeng. 49:93-100.
Xu, K. D., P. S. Stewart, F. Xia, C.-T. Huang, and G. A. McFeters. 1998.
Physiological heterogeneity in Pseudomonas aeruginosa biofilm is determined by
oxygen availability. AppL Environ. Microbiol. 64:4035-4039.
Yost, F. J., Jr., and I. Fridovich. 1973. An iron-containing superoxide dismutase from
Escherichia coli. J. Biol. Chem. 248:4905-4908.
49
CHAPTER 3
Pseudomonas aeruginosa BIOFILM RESISTANCE TO
HYDROGEN PEROXIDE: PROTECTIVE ROLE OF CATALASE
Introduction
Bacterial biofilms profoundly affect industrial systems by promoting material
fouling and loss of ‘pump and pipe’ system efficiency. Biofilms also impact human
health by causing persistent infections stemming from bacterial accumulations on tooth
surfaces and medical implants. Biofilm formation, especially in Gram-negative bacteria,
is thought to be an effective survival strategy for bacteria against environmental
challenges such as desiccation (Costerton et ah, 1981) or nutrient limitation (Ostling et
ah, 1 993). The importance of biofilm formation to the survival of microbial populations
is best demonstrated by the tremendous resistance biofilms exhibit against antimicrobial
agents (Ashby et ah, 1994; Brown and Gilbert, 1993; LeChevallier et ah, 1988). Biofilm
resistance to antimicrobials is much greater than planktonic organisms, and it is this
innate recalcitrance to a broad spectrum of bactericidal compounds that makes biofilm
control difficult. An understanding of the primary mechanisms responsible for the
reduced susceptibility of biofilms to antimicrobial agents will aid in the successful
eradication of problem biofilms in the future.
Biofilm structural characteristics have been assessed for their contribution to
50
biofilm resistance to antimicrobial agents. The retarded penetration of antibiotics into
microbial aggregates due to exopolysaccharide (EPS) encapsulation has been implicated
in biofilm resistance (Nichols et ah, 1989; Tresse and Junter, 1995). Also, reaction, and
therefore neutralization, between strongly oxidizing biocides such as hypochlorous acid
and biofilm constituents have been shown to provide some protection against Tcilling
(Chen and Stewart, 1996). Undoubtedly, reaction and diffusional barriers to
antimicrobial penetration established by the complex EPS matrices surrounding biofilm
organisms provide some degree of protection. However, it has been suggested that
reaction-diffusional limitation alone cannot totally account for biofilm resistance to many
antimicrobial agents (Brown and Gilbert, 1993; Stewart, 1996).
It has recently been shown that physiological heterogeneity exists throughout the
depth of a biofilm (Huang et al., 1998), with physiological variation perhaps occurring as
a function of oxygen availability (Xu et al., 1998). Physiological heterogeneity is likely
to contribute to the reduced susceptibility of biofilms to antimicrobial agents, especially
when growth dependent antibiotics (i.e. p-lactams) are administered. Also, portions of a
biofilm experiencing nutrient limitation may be more resistant to antimicrobial agents
due to differential stationary-phase gene regulation which is well known to render
microorganisms more resistant to many adverse environmental conditions (Morita, 1993).
In addition to reaction-diffusion limitation and low physiological activity,
physiological adaptation to antimicrobial agents may also be important. Specific
physiological responses to antimicrobial agents have been well characterized in bacteria,
especially oxidative stress responses against bactericidal reactive oxygen intermediates
(ROIs) (See Demple, 1991; Farr and Kogoma, 1991 for reviews). ROIs include the
superoxide anion (0% ), hydrogen peroxide (H2O2), and the powerfully oxidizing
hydroxyl radical (OH ). ROIs primarily result from the partial, univalent reduction of
oxygen by electron transport components during aerobic metabolism (Gonzalez-Flecha
and Demple, 1995). Bacteria may also encounter extracellular fluxes of ROIs from
macrophage when attempting to colonize animal tissues or when ROIs are employed as
disinfectants. Although adaptive responses against oxidative stress have been extensively
studied with planktonic cells (Demple, 1991; Farr and Kogoma, 1991), comparatively
little is known about biofilm responses to biocide attack. Biofilms have been shown to
become increasingly resistant to repeated doses of antibiotics (Giwercman et al., 1991) or
non-specific oxidizing biocides such as monochloramine (Sanderson and Stewart, 1997),
but the basis for this acquired resistance is currently unknown.
In this study, a previously described model system for examining biofilm
resistance mechanisms (Hassett et al., 1999a) was employed to assess the significance of
catalase expression in the protection of Pseudomonas aeruginosa biofilms against the
oxidizing biocide H2O2. P. aeruginosa expresses several catalases, which catalyze the
disproportionation OfH2O2 into oxygen and water. The primary ‘housekeeping’ catalase,
KatA, is expressed constitutively throughout the growth cycle with increased expression
at the onset of stationary phase (Brown et al., 1995; Hassett et al., 1992). KatB
expression is repressed during aerobic growth but is highly inducible upon exposure to
H2O2 (Brown et al., 1995). Another catalase, KatC, has been observed in P. aeruginosa
at very low levels and the importance and regulation qf this enzyme remain currently
unknown (Hassett, personal communication). In this study, the importance of catalase
for the protection of P. aeruginosa biofilms was examined by comparing the wild type
strain against KatA' and KatB" mutants for their susceptibility to H2O2.
Spectrophotometric enzyme assays, activity stains in non-denaturing polyacrylamide
gels, and reporter gene experiments were employed to characterize the oxidative stress
response in biofilms and planktonic cells.
Materials and Methods
Bacterial Strains and Plasmids
The Escherichia coli and P. aeruginosa strains as well as the plasmids used in this
study are listed in Table 3-1.
Media and Growth Conditions
Planktonic cells were cultured in Pseudomonas Basal Mineral (PBM) medium
(Atlas, 1997), which contained I g/L glucose as carbon source. Iron was added as 10 pM
FeCl3, which was found in preliminary studies to yield maximum catalase activity in
PBM medium (results not shown). Strains were maintained on PBM medium solidified
with 1.5% Noble Agar (Difco) and containing 300 mg/L gentamicin (Sigma) when
appropriate. All planktonic cultures were incubated at 25° C in a rotary shaker water bath
at 300 rpm. Culture volumes did not exceed 10% of the flask volume to ensure
maximum aeration. Biofilms were cultured using a drip-flow reactor system previously
described by Huang et al. (1998) and included 316L stainless steel slides (1.3 x 7.6 cm)
as the substratum. Briefly, 10 mis of PBM medium were added to each chamber (four
chambers per reactor), followed by inoculation with I ml of stationary phase culture of
the test strain grown in the same medium. The reactor was then incubated horizontally at
53
Table 3-1. List of Strains and Plasmids Used in This Study.
Strain or Plasmid
E. coli
DHSoc
Source or Reference
F" ZacZDMlS recAl hsdRl 7 supE44
Liss, 1987
EilacZYA argF)
thi thr leu tonA IacY supE recA Muc1"
Simon et al., 1983
recA thi pro leu hsdRM'
Boyer and R.-Dussoix, 1969
SMlO
HBlOl
P. aeruginosa
PAOl
PAOl(AaM)
P A O l( M )
PAOI (katB\:lacZ)
PAOl(ZacZ)
Plasmids
pRK2013
pZ1918
pMini-CTX
pFLP2
pJE26
Relevant Genotype or Characteristic
Wild-type
PAOI, AaMxGmr
PAOI, AatSxGmr
PAOI, katB reporter strain
PAOI, IacZcontrol strain
This study
This study
This study
This study
This study
Kmr, orzT helper plasmid.
Figurski and Helinski, 1979
Apr, pUC19/18 with 3.2 kb IacZcassette
Schweizer, 1993
Tcr, attP site specific integration vector
H. P. Schweizer
Apr, Sucs, broad-host-range Flp expression
H. P. Schweizer
vector
Apr, Xbal-HindLll katB fragment in pQFSO
This study
forming a katBv.lacZ transcriptional fusion
pJGE02
Tcr, pMini-CTX containing IacZ cassette
from pZ1918 on BamlAl fragment.
pJGE03
Tcr, pJGE02 with Xhol-Hindlll fragment from
pJE26 containing katB promoter.
This study
This study
Abbreviations include: Apr, ampicillin resistance; Gmr, gentamicin resistance;
Kmr, kanamycin resistance; Sucs, sucrose sensitivity; Tcr, tetracycline
resistance.
54
room temperature for 24 hrs to allow bacterial attachment to the substratum. Following
the attachment period, the reactor was inclined 10° and a constant drip of half-strength
PBM (except 200 mg/L glucose) was allowed to flow over the slides at a rate of 50 mis
hr ^for 72 hrs. Biofilm thickness was determined using cryosectioning and microscopic
analysis as previously described (Huang et ah, 1998). For construction and isolation of
catalase IacZ reporter strains, Pseudomonas Isolation Agar (PIA) (Difco) was employed
as the sole medium for P. aeruginosa strain selection and maintenance and was
supplemented with carbenicillin (Cb; 300 mg/L), tetracycline (Tc; 100 mg/L), or sucrose
(Sc; 5%) when appropriate. Plates were incubated at 37° C for 24-48 hrs. E. coli strains
were cultured in LB (1% Triptone, 0.5% Yeast Extract, and 0.5% NaCl) broth or on LB
plates solidified with 1.5% Bacto agar (Difco). For maintenance and selection of
plasmids, liquid and solid LB media were supplemented with tetracycline (Tc; 15 mg/L),
ampicillin (Ap; 100 mg/L) and 5-bromo-4-chloro-3-indolyl-|3-D-galactopyranoside (XGal; 40 mg/L). Liquid cultures were incubated overnight at 37° C in a rotary shaker
water bath set at 300 rpm; plates were also incubated overnight at 37° C.
Planktonic Disinfection
P. aeruginosa wild-type (wt) strain PAOI, katA mutant, and katB mutant strains
were cultured as described above for broth cultures. Stationary phase Cells were
harvested and diluted to a cell density of approximately 3.0 x IO7 CFU/ihl in PBM
medium that did not contain glucose. To determine initial cell viability, an aliquot of
cells from each flask was serially diluted in phosphate buffered saline (PBS) (pH 7.2) that
contained 0.2% sodium thiosulfate (w/v) to neutralize H2O2, then plated on R2A agar
(Difco). Hydrogen peroxide (Sigma) was then added to a final concentration of 50 mM
and each flask was immediately returned to shaking at 25° C. At 20 minute intervals for
a period of I hour, culture aliquots were collected and serially diluted in the peroxide
neutralizing PBS buffer. Appropriate dilutions were plated on duplicate R2A agar plates
which were allowed to incubate overnight at 37° C. Following incubation, CPUs were
enumerated and disinfection was expressed as the percentage of surviving cell's relative to
the initial cell viability.
Biofilm Disinfection
P- aeruginosa wt, KatA", and KatB" biofilms were generated with a drip-flow
reactor under the conditions described above. Following the 72 hr growth period, a
biofilm sample was harvested as a zero H2O2 control and the culture medium was
exchanged for PBM medium that included 50 mM H2O2. The peroxide containing
medium was allowed to flow over the biofilms at a rate of 50 mls/hr for a period of I hr.
At 20 minute intervals, biofilm slides were harvested and scraped into 50 mis of PB S (pH
7.2) containing 0.2% sodium thiosulfate to neutralize H2O2. Biofilm biomass was
homogenized by using a PT 10/35 Brinkman homogenizer (Brinkman Instruments,
Westbury, NY) for 15 sec. at setting 4. Biofilm cell viability was determined before and
during disinfection by diluting biofilm homogenate and enumerating viable cells on R2A
agar as described for the planktonic disinfection experiments. In addition to the viable
cell count, the total number of cells harvested from each biofilm was determined using
nonspecific staining and epifluorescence microscopy. Cells from appropriate dilutions
were collected on black polycarbonate filters (Poretics, Livermore, CA) and stained with
4 ’-6 diamidino-2-phenylindole dihydrochloride (DAPI; Molecular Probes, Inc, Eugene,
Oregon). Stained cells were visualized using an Olympus BH-2 microscope (Lake
Success, NY) with ultra-violet, epiflourescent illumination. In order to eliminate
potential sampling bias that may arise from detachment of biofilm cells during
disinfection, biofilm disinfection is expressed as a ratio between the viable fraction of the
total number of cells present in each biofilm before and after disinfection. All
experiments were repeated at least three times.
Preparation of Cell Free Extracts
Planktonic cultures were allowed to achieve stationary phase and then subjected
to a 2 mM pulse OfH2O2 every 10 min. for a period of I hr. while biofilms were treated
with a constant stream of peroxide as described above. Planktonic cultures were
harvested before and after the dosing protocol while biofilm samples were collected at 20
minute intervals during the I hr. disinfection period. Batch cultures and biofilm
homogenates were neutralized with 0.2% sodium thiosulfate, and chloramphenicol (Cm;
300 mg/L) Was added to arrest protein synthesis. Cell free extracts and catalase enzyme
assays for planktonic and biofilm cells were prepared according to Hassett et al. (1999a).
Briefly, treated and control planktonic cultures and biofilm homogenates were
centrifuged at 8,000 x g for 8 minutes at 4° C, washed twice with 10 mis of ice-cold 50
mM potassium phosphate buffer (PPB; pH 7.0), then resuspended in 0.5 ml of PPB and
transferred to an Eppendorftube for sonication. Cell extracts were generated by 2-30 sec.
pulses with a Fisher Scientific Model #550 sonicator at power setting 1.8. Sonicate was
then centrifuged at 13,000 x g for 10 minutes at 4° C to remove unbroken cells and cell
debris, and the supernatant was then transferred to a fresh tube. Total protein
concentration was determined by the method of Bradford (1976) with bovine serum
albumin fraction V as standard.
Catalase Assays and Native-PAGE
Specific catalase activities for planktonic and biofilm cell free extracts were
determined as previously described (Beers and Sizer, 1952; Brown etal., 1995; Hassett et
al., 1999a). Catalase expression was also analyzed using non-denaturing polyacrylamide
gel electrophoresis according to Hassett et al. (1999a). Briefly, cell free extracts (15 pg
protein/well) were electrophoresed through vertical 5% continuous polyacrylamide gels
for approximately 10 hours at 10 mA constant current. Gels were stained for catalase
activity as previously described (Brown et al., 1995; Wayne and Diaz, 1986).
DNA Manipulation and Cloning
Protocols used to manipulate nucleic acids were obtained from Sambrook et al.
(1989). Enzymes were purchased from either Promega (Madison, WI) or New England
Biolabs (Beverly, MA). Oligonucleotides were designed with OLIGO software and
purchased from Integrated DNA Technologies Inc. (Coralville, IA).
Construction of pJGE02
A promoterless IacZ cassette was excised from pZ1918 (Schweizer, 1993) on a
3.4-kb BamRl fragment which was then ligated into the unique BamRl site located in the
multi-cloning region of the integration vector pMini-CTX (Schweizer, 1999). The
ligation products were transformed into E. coli DH5a and transformants yielding light
blue colonies, on LB plates containing 20 mg/L tetracycline and 40 mg/L X-Gal were
selected for further analysis. Restriction analysis was used to screen recombinant
plasmids for constructs containing the 3.4-kb IacZ cassette in the correct orientation.
Construction of pJGEOB
A 580 bp fragment containing the katB promoter was isolated from pJE26, a
previously constructed katB::lacZ transcriptional reporter vector (Hassett et al., 1999b),
on Z-XhoI-HindlIl fragment and sub-cloned into the corresponding sites of the pJGE02
polylinker. Restriction analysis of plasmids isolated from tetracycline resistant
transformants revealed several constructs containing the 580-bp XhoI-HindIII fragment.
Isolation of PAOI (katB:\lacZ)
A novel system developed by Schweizer (1999) was employed to introduce a site
directed, single-copy katBwlacZ transcriptional fusion into the P. aeruginosa PAOl
genome. Briefly, the pMini-CTX suicide vector (pJGE03 backbone) integrates into the
P- aeruginosa genome at the attB site via an integrase {inf) mediated recombination. Tcr
PAOl colonies are selected and a second plasmid, pFLP2, is introduced. The pFLP2
construct is a broad-host-range, Flp-expression vector that excises DNA sequences
included within Flp recombinase target (FRT) sites. The integrated pMini-CTX vector
contains two FRT sites that flank the Tcr, int, and onT markers. Therefore, introduction
of the pFLP2 construct eliminates these markers from the integrate. After the pFLP2 .
plasmid is cured from the reporter strain, only the inserted gene fusion is left remaining
which is also flanked by T4-derived transcriptional terminators. The pJGEOS construct
was introduced into PAOl by tri-parental conjugal mating using E. coli DH5a as the
donor and E. coli HB101(pRK2013) as the helper strain. Tcr colonies on PIA containing
Tc 100 mg/L were selected and E. coli SMlO (pFLP2) was used to introduce the excision
plasmid into the Tcr transconjugates. Several colonies showing Cbr and Tcs were
selected. To cure the pFLP2 plasmid from putative reporter strains after FRT excision,
cultures were grown overnight in the absence of antibiotics at 42° C and then plated on
PIA containing 5% sucrose. Plasmid preps were used to verify absence of the plasmid.
A control strain, PAOl(ZacZ), which contained a promoterless IacZ gene inserted at the
attB site, was also constructed using the same approach. PCR primers (designed from
template obtained from GenBank accession number D13407) were designed to verify
integration of pJGE02 and excision of the unwanted vector sequences. PCR products
were sequenced using an ABI Prism BigDye Kit (PE Applied Biosystems, Foster City,
.
CA) and an ABI model 310 Genetic Analyzer (PE Applied Biosystems, Foster City, CA).
Reporter Gene Assays
Planktonic cells and biofilms were cultured and treated with H2O2 as described
above except that in addition, stationary phase planktonic cells were also treated with a
single 50 mM dose of H2O2. Biofilms and planktonic cells were harvested as described
above but instead of preparing cell free extracts, biofilm homogenates and planktonic
cultures were washed twice in PBS (pH 7.2) and resuspended in Z-buffer (pH 7.2; Miller, •
1972). Resuspended cells were then assayed for (3-galactosidase activity by the method
described by Miller (1972).
Statistical Analysis
When comparing sample means, a Student’s Z-test was performed to generate
60
probability values (P). Statistical significance was declared with 95% confidence
(a=0.05).
Results
Catalase Activity in IcatA and katB mutants
The oxidative stress response in the catalase mutants was characterized relative to
the wild-type (wt) strain. Previous work had established that KatA is expressed
constitutively in P. aeruginosa and represents the primary defense against H2O2, whereas
KatB is the secondary resistance mechanism and is inducible upon exposure of the cells
to H2O2 (Brown et ah, 1995; Hassett et ah, 1992). Catalase specific activity levels in wt
stationary phase planktonic cultures were typically about 400 U/mg protein (Table 3-2).
After repeated doses of low levels of H2O2, total catalase levels increased by roughly
50%. No catalase activity was observed in untreated stationary phase cultures of the hatA
mutant, nor was it detectable in equivalent cultures exposed to H2O2. In the latter case,
the lack of the constitutively expressed KatA catalase resulted in almost complete killing
within minutes (see below) and thus the cells were not capable of inducing katB.
Inducibility of katB was verified, however, using very small doses of H2O2 (Table 3-2).
Catalase levels in uninduced katB cells were similar to wt cells, but would not increase
when the mutant was stressed with H2O2. The results of these studies established the
range of catalase activity to be expected with each strain and verified that the phenotype
matched the mutations introduced into P. aeruginosa.
Table 3-2. Planktonic Specific Catalase Activity of wt, KatA", and KatB' Before and
After Treatment with H2O2.
Strain
PAOl
Catalase Activity
Before Disinfection
(U/mg protein)
426 ± 6
Catalase Activity
After Disinfection1
(U/mg protein)
609 ± 14
PAOl(AnM)
ND2
ND
PAOl(WB)
430 ± 10
432 ± 9
1 Stationary phase cultures were treated with 2 mM doses of H2O2 every 10 min, for I
hr. while shaking at 25° C.
2 ND, not detectable.
62
H2O2 Sensitivity of Planktonic Cells
In order to assess the importance of catalase in the protection of planktonic cells
from hydrogen peroxide, each strain was subjected to a single dose of 50 mM H2O2.
Reductions in cell viability were determined at 20 min. intervals for a period of I hour
(see Materials and Methods). Shown in Fig. 3-1, all strains were variably sensitive to the
peroxide treatment. Even with a full complement of catalase activity, wt cells displayed a
logic 3.5 cfu/ml decrease in viability during the I hour sampling period. The KatB"
mutant appeared to be more sensitive to peroxide than the wt (mean logic reduction of
5.0 at I hr.), particularly at the longer H2O2 exposure periods, but these differences were
not statistically significant (P-0.119). Cells lacking KatA, and therefore having virtually
no catalase activity (Table 3-2), were hypersensitive to peroxide. In all experiments^
viable KatA" cells could not be detected at the 20 min. sampling time. Therefore, at least
a logic 6.3 reduction in viable cells occurred as soon as 20 min. after addition of
peroxide. The lack of KatA clearly had a significant effect on the ability of planktonic
cells to survive H2O2 treatment when compared to the wt (P < 0.01) or KatB' strains (P <
0 .01).
H2O2 Sensitivity of Biofilm Cells
Biofilms formed by the wt and catalase mutant strains were also tested for their
susceptibility to 50 mM H2O2. However, in these experiments peroxide was delivered as
a constant flow (50 mls/hr.) rather than a single pulse. The catalase mutations had no
apparent affect on the ability of these strains to form a biofilm. Biofilm thickness of .
these strains was determined to be 114 +/-13 pm and total cell counts estimated that
63
initial cell densities ranged from 2.1xl09 to 8.7xl09 total cells/cm2; these are typical
ranges for both parameters. To better represent actual biofilm cell survival, percent
survival is shown as the viable fraction of the total number of cells present on the
substratum at each sampling point (Fig. 3-2). When compared to the rates of killing
observed with planktonic organisms, all strains were relatively much more resistant to
peroxide. For the wt and KatB' strains, survival did not differ and was nearly constant at
about 80% and 77%, respectively. A different result was obtained with the KatA' mutant.
During the first 20 min. of H2O2 exposure, a logic 1.03 reduction in the viable fraction
occurred. At 40 min., the total number of cells showed consistently higher viability and
nearly equaled those of the wt and KatB" strains. Then at 60 min., viability decreased
again approximately 90%. Although not as dramatic as the affect seen with planktonic
cells, the lack of KatA significantly (P< 0.05) increased the susceptibility of biofilms to
the 50 mM H2O2 treatment.
Biofilm Removal with Hydrogen Peroxide.
From total cell counts, the relative amount of biofilm removal that occurred
during exposure to H2O2 could be determined and compared (Fig. 3-3). Even with
profuse effervescence due to high specific catalase activity (Fig. 3-4), wt and KatB'
biofilms remained largely intact with greater than 70% of the initial total biomass
remaining on the stainless steel surface at I hr. Unlike the wt and KatB", virtually no
fizzing was observed with the KatA" biofilms during treatment with H2O2. Nevertheless,
biofilm breakup occurred such that approximately 90% of the initial biomass was
removed, leaving a very thin, and apparently more resistant, layer of cells remaining on
64
Planktonic Cell Disinfection with 50 mM H2O2
1.0E+03 -
'o Surviv
1.0E+01 .
1.0E-01 .
1.0E-03 .
1.0E-05
Time (min.)
Figure 3-1. Planktonic disinfection of wt (■ ), KatA" (♦ ), and KatB" ( • ) strains
exposed to 50 mM H2O2 for I hr. Cells were grown to stationary phase in
PBM medium and then diluted to approx. 3.0 x IO7cfu/ml in PBM w/o
carbon source. Hydrogen peroxide was added as a single dose and samples
were collected and neutralized at 20 min. intervals for I hr. Data are
represented as the mean +/- S.E. for 3 independent experiments.
65
Biofilm Disinfection with 50 mM H2O2
Surviv
100.0
10. 0
.
Time (min.)
Figure 3-2. Biofilm disinfection of wt (■ ), KatA" (♦ ), and KatB" ( • ) strains exposed
to a constant flow of 50 mM H2O2 for I hr. Percent survival is expressed as
the viable fraction of the total number of cells present on the substratum
relative to f=0 min. or, [(V,/T,) / (V0ZT0)] x 100. Data are represented as the
mean +/- S.E. for 3 independent experiments.
66
Biofilm Removal During Disinfection
1 0 0 .,
80.
ro 60
o
E
<D
cr
40
20.
wild-type
KatA-
KatB-
Figure 3-3. Total cell removal from wt, KatA', and KatB'biofilms after 60 min. of
constant exposure to 50 mM H2O2. Each data point represents the
mean +/- S. E. for three independent experiments.
the slide. Differences in remaining katA mutant biofilm biomass and that measured with
the wt and katB strains were statistically significant (P=O-Ol).
Catalase Activity of P. aeruginosa Biofilms
In addition to assessing susceptibility to hydrogen peroxide, the specific catalase
activities of biofilms (Fig. 3-4) before and after exposure to H2O2 were determined for P.
aeruginosa PAOl wt and catalase mutant strains. Biofilms were collected at 20 min.
intervals during exposure to a constant flow of 50 mM H2O2. Initial specific catalase
activities of wt biofilms did not differ from stationary phase planktonic cultures (compare
to Table 3-2). Like planktonic cells, biofilm catalase expression is influenced by
peroxide. As soon as 20 min. after exposure, specific catalase activity had increased in
wt biofilms and continued to increase throughout the exposure period reaching
approximately 700 U/mg protein in I hr. The 47% increase over the initial activity was
again almost identical to planktonic cells. KatB" biofilms yielded similar initial specific
catalase activities but displayed no increase during exposure to peroxide indicating that
katB induction was primarily responsible for the significant (P=0.01) increase in catalase
activity observed for the wt biofilms. Total specific catalase activities for KatA" biofilms
could only be determined at 0 and 20 min. after exposure since most of the biomass
sloughed from the substratum after 30 min. No catalase activity was detected in cell free
extracts collected from KatA" biofilms at either time point (data not shown). As with
planktonic cells (Table 3-2), KatA" biofilm cells appeared to be rapidly overwhelmed by
the H2O2, and could not survive long enough to synthesize the KatB catalase. Attributing
the increase in biofilm catalase activity to katB induction is supported by the data shown
68
Biofilm Specific Catalase Activity
800 n
700 .
600 .
500 --
Time (min.)
Figure 3-4. Total specific catalase activity of wt ( a ) and katB- ( * ) biofilms exposed
to 50 mM H2O2 for I hr. Cell free extracts were prepared from biofilms
during the disinfection period and assayed for catalase activity as described
in materials and methods. Specific catalase activity is expressed as ^mol
H2O2degraded per min. per mg of total protein. Each data point represents
the mean +/- S. E. for three independent experiments. Duplicate assays were
performed on each cell free extract preparation.
Biofilm Catalase Expression During Disinfection
Figure 3-5. Biofilm catalase expression patterns during disinfection. Biofilms were
exposed to 50 mM H2O2 and samples were harvested at 20 min. intervals for
a period of I hour. Cell free extracts were loaded on a 5% non-denaturing
polyacrylamide gel allowing separation of the KatA and KatB enzymes.
Each lane contains 15 pg of total protein.
In Fig. 3-5. Examining catalase activity in nondenaturing polyacrylamide gels, though
lacking the precision of spectrophotometric assays, allows a sensitive assessment of
which isozymes are being expressed. Before exposure to H2O2, no KatB activity is
apparent in cell free extracts collected from wt biofilms. Following H2O2 exposure, KatB
is apparent at the first sampling point during the treatment period. Although the lack of
KatB did not have a significant affect on biofilm susceptibility to the dosing procedure
used in these experiments, the induction of KatB activity strongly suggests that the
biofilms cells are experiencing oxidative stress due to the constant exposure to hydrogen
peroxide.
katBwlacZ Reporter Construction and Analysis
To further investigate the physiological response of P. aeruginosa biofilms to
H2O2, a PAOKkatBwlacZ) reporter strain was constructed using a novel approach for
introducing single-copy gene fusions into the P. aeruginosa genome (Schweizer et al,
1999). This method is advantageous for biofilm studies since the chromosomal fusions
are stable in the absence of antibiotic selection and of particular importance, the
promoter/gene of interest is not insertionally inactivated. To verify correct insertion,
PCR primers specific for the flanking chromosomal DNA and integrated fusion were
used to amplify the junction sequences. Single amplification products of the predicted
size were obtained from genomic templates (data not shown). These PCR products were
also sequenced as an additional verification that the constructions were correct. Sequence
data verified that both the pJGE03 and pMini-CTX (control) vectors had integrated into
the chromosome at the attP site and that pFLP2 mediated excision had occurred
PAOI(katB/.lacZ) Expression in Response to Hydrogen Peroxide
Time (min.)
Time (min.)
Figure 3-6. PAOl (katB. dacZ) reporter activity of biofilm (A) and planktonic cultures
(B) exposed to hydrogen peroxide. Biofilms were treated with a continuous
dose of OmM ( • ) or 50 mM (O) H2O2 for a period of I hr. Planktonic,
stationary phase cultures were exposed to single 0 mM ( • ) and 50 mM (O)
doses or 2 mM pulses every 10 min. for I hr. (■ ). Data represent the mean
+/- S. E. for three independent experiments.
72
precisely.
The ^AO\{katBwlacZ) reporter strain provided a means of studying katB
expression in both planktonic and biofilm cells (Fig. 3-6A and 3-6B). The induction
profile was similar for both cell types, although the conditions required for induction
differed somewhat. In preliminary measurements, it was empirically determined that
culture cfu/ml optical density units for washed cell suspensions of both cell types were
equivalent (results not shown), and therefore allowed reporter enzyme units to be
conveniently calculated in terms of optical density. Similar to the results of experiments
depicted in Table 3-2 and Fig. 3-4, PAOI {katB::lacZ) biofilms exhibited a gradual, two­
fold induction of reporter gene activity over the one hour exposure period. Reporter
enzyme levels did not increase in ?AOl{katBwlacZ) biofilms that were not exposed to
H2O2. When planktonic cells were exposed to 2 mM doses OfH2O2 every 10 min. for I
hr., a response similar to that of biofilm cells (Fig. 3-6A) was observed. However, when
stationary phase planktonic cells of the reporter strain were exposed to a single 50 mM
dose OfH2O2, no katBwlacZ induction was observed (Fig. 3-6B). Initial levels of biofilm
and planktonic cells gave nearly equal levels of reporter enzyme activity, averaging
roughly 30 Miller units. The promoterless IacZ control construct, PAOl(ZacZ),
consistently yielded approximately 30 Miller units and was unaffected by single 50 mM
or multiple 2 mM H2O2 treatments.
Discussion
Due to its ubiquity in nature and tendency to form biofilms, P. aeruginosa has
been used as a model organism for studying biofilm behavior and for the development of
biofilm control strategies (Costerton et al., 1994). Though recent progress has been
significant, P. aeruginosa biofilm cell physiology is still only poorly understood.
Gradients in metabolic activity have been shown to exist in P. aeruginosa biofilms, and
some information regarding adaptive gene regulation has also been recently published
(Huang et al. 1998; Xu et al., 1998). These studies examined P. aeruginosa gene
regulation in response to environmental stress, showing for example that induction of
phoA in response to phosphate limitation was limited to the upper region (ca. 20%) of the
biofilm (Huang et al. 1998). It was proposed that oxygen limitation within the biofilm
imposes restrictions on cell activities that include de novo protein synthesis in response to
environmental stress (Xu et al., 1998). It is anticipated that spatially differentiated gene
regulatory responses within biofilms will prove to be an important issue in biofilm
control. In continuing attempts to expand our understanding of biofilm cell physiology,
we are also examining oxidative stress responses in P. aeruginosa biofilms. Significant
genetic and biochemical information regarding the oxidative stress response to H2O2 in
this organism is already available [Hassett and co-workers papers (1992, 1995, 1999a,
1999b)], This should facilitate efficient progress in determining the basis for the
significant resistance oiP. aeruginosa biofilms to oxidizing biocides.
In the data reported here, the extraordinary recalcitrance of P. aeruginosa
biofilms to H2O2 treatment is in complete agreement with previous studies using other
oxidizing biocides (LeChevallier et al., 1988; Sanderson and Stewart, 1997). Even after
one hour of continuous exposure to 50 mM H2O2, wt biofilm integrity remained largely
intact (Fig. 3-3) and nearly 80% of the cells survived (Fig. 3-2). We were interested in
determining whether such biofilms employed special mechanisms that are especially
effective in guarding the cell against oxidative damage by H2O2. An obvious physiologic
trait of importance to inactivating H2O2 would be the cellular level of catalase activity,
and therefore we focussed our initial efforts on this enzyme. While the catalase specific
activity of P. aeruginosa grown in a defined medium is lower than that measured with
cultures grown in complex media (Brown et al., 1995; Hassett et al., 1999b), regulation
of the isozymes appears to be the same. KatA is expressed throughout the growth cycle
with a marked increase at the onset of stationary phase (Brown et al., 1995). KatB is not
expressed during the aerobic growth cycle but is inducible upon exposure to peroxide,
making KatB expression a marker enzyme for oxidative stress in P. aeruginosa. With
planktonic cultures grown in minimal media, expression patterns of KatA and KatB were
unaltered (Table 3-2 and other data not shown). In the absence OfH2O2, biofilms formed
by the wt strain contained only KatA (Fig. 3-5), and at levels that were equivalent to
planktonic cultures (Table 3-2., Fig. 3-4). As the KatB catalase represents the primary
oxidative stress response catalase in P. aeruginosa, we also examined the expression of
this enzyme to determine whether significantly high levels of expression could explain
the elevated levels of resistance to H2O2. As was observed with planktonic cells, catalase
specific activity increased in biofilms treated with H2O2; however, the increase observed
in biofilm cells was of the same magnitude as observed in planktonic cultures (Table 3-2,
Fig. 3-4). Native gel analysis and reporter gene data were also in agreement and
suggested that the increased catalase activity was due to KatB (Fig. 3-5, Fig. 3-6).
As another approach to assessing the relative contribution of these enzymes to
biofilm resistance to H2O2, we created defined katA and JcatB mutations in the wt strain
PAOI. The loss of KatA resulted in planktonic cells being hypersensitive to H2O2 (Fig.
3-1), and biofilm resistance to H2O2 decreasing by approximately an order of magnitude
relative to the wt strain (Fig. 3-2). Further, biofilms formed by the KatA' mutant
essentially disintegrated when exposed to H2O2 (Fig. 3-3), suggesting that constitutive
catalase expression is necessary for maintaining biofilm structural integrity in the
presence O fH 2O2. In contrast, katB mutants were very similar to the wt strain with
regards to planktonic cell survival and biofilm resistance to H2O2 (Figs. 3-1 and 3-2).
Therefore, even though katB is readily induced in both cell types in response to H2O2
exposure, its contribution to the initial biofilm oxidative stress resistance phenotype
appears to be minimal. It is important to note however, that biofilms were able to induce
katB in the presence OfH2O2 concentrations that were lethal to planktonic cells (Figs 3-6a
and 3-6b). This suggests that at the community level, biofilms are better adapted than
planktonic cells in initiating a multi-cellular oxidant defense against H2O2.
In the current study, we have assessed the relative importance of the different
catalase enzymes to protection against H2O2. Based on the analysis of KatA and KatB
catalase expression and activity levels, we conclude that the remarkable resistance of the
wt biofilm to H2O2 cannot be attributed to abnormally high initial or induced levels of
catalase activity. Catalase expression is critical to survival, however, other mechanisms
of resistance appear to be involved. Indeed, it is important to note that under the
conditions of the experiments used in this study, even the katA mutant still survived at
relatively high levels. Other possible mechanisms contributing to the reduced
susceptibility of biofilm cells to oxidizing agents have been investigated. The failure of
reactive agents to penetrate the EPS matrix of a biofilm has been shown to contribute to
the reduction in susceptibility to oxidizing agents (Chen and Stewart, 1996); but,
penetration failure alone may only be partially responsible for resistance (Brown and
Gilbert, 1994; Stewart, 1996). Gradients in metabolic activity have been shown to exist
in biofilms (Huang et ah, 1998; Xu et ah, 1998); and, slow microbial growth rates are
also correlated with increased resistance to oxidants such as hydrogen peroxide (Ivanova
et ah, 1994; Loewen and Hengge-Aronis, 1994; Seymor et ah, 1996). It is likely that
several mechanisms including physiological adaptation contribute simultaneously to
biofilm resistance to antimicrobial agents. Further investigation of the unique aspects of
biofilm physiology will eventually reveal assailable approaches to eradicating
problematic biofilms in the future.
77
References
Ashby, M. J., J. E. Neale, S. J. Knott, and I. A. Critchley. 1994. Effect of antibiotics
on non-growing planktonic cells and biofilms of Escherichia coli. J. Antimicrob.
Chemother. 33:443-452.
Atlas, R. M. 1997 in “Handbook of Microbiological Media”, L. C. Parks, ed., CRC
Press, Ann Arbor, Mich.
Beers, R. F., Jr., and I. W. Sizer. 1952. A spectrophotometric method for measuring the
breakdown of hydrogen peroxide by catalase. J. Biol. Chem. 195:133-140.
Boyer, H. W., and D. Roulland-Dussoix. 1969. A complementation analysis of the
restriction and modification of DNA in Escherichia coli. J. Mol. Biol. 41:459.
Bradford, M. M. 1976. A rapid and sensitive method for the quantitation of microgram
quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem.
72:248-254.
Brown, M. R. W., and P. Gilbert. 1993. Sensitivity of biofilm to antimicrobial agents. J.
AppL Bacteriol. Symp. Suppl. 74:87S-97S.
Brown, S. M., M. L. Howell, M. L. Vasil, A. J. Anderson, and D. J. Hassett. 1995.
Cloning and characterization of the katB gene of Pseudomonas aeruginosa encoding a
hydrogen peroxide-inducible catalase: purification of KatB, cellular localization, and
demonstration that it is essential for optimal resistance to hydrogen peroxide. J. Bacteriol.
177:6536-6544.
Chen, X., and P. S. Stewart. 1996. Chlorine penetration into artificial biofilm is limited
by a reaction-diffusion interaction. Environ. Sci. Technol. 30:2078-2083.
Costerton, J. W., R. T. Irvin, and K.-J. Cheng. 1981. The bacterial glycocalyx in
nature and disease. Annu. Rev. Microbiol. 35:299-324.
Costerton, J. W., Z. Lewandowski, D. DeBeer, D. Caldwell, D. Korber, and G.
James. 1994. Biqfilms, the customized microniche. J. Bacteriol. 176:2137-2142.
DempIe B. 1991. Regulation of bacterial oxidative stress genes. Annu. Rev. Genet.
25:315-337.
Farr S. B., and T. Kogoma. 1991. Oxidative stress responses in Escherichia coli and
Salmonella typhimurium. Microbiol. Rev. 55:561-585.
78
Figurski, D. H., and D. R. Helinski. 1979. Replication of an origin-containing
derivative of plasmid RK2 dependent on a plasmid function provided in trans. Proc. Natl.
Acad. Sci. USA 76:1648-1652.
Giwercman, B., E. T. Jensen, N. Heiby, A. Kharazmi, and J. W. Costerton. 1991.
Induction of the p-lactamase production in Pseudomonas aeruginosa biofilm.
Antimicrob. Agents Chemother. 35:1008-1010.
Gonzalez-Flecha, B., and B. Demple. 1995. Metabolic sources of hydrogen peroxide in
aerobically growing Escherichia coli. I. Biol. Chem. 270:13681-13687.
Hassett, D. J., L. Charniga, K. Bean, D. E. Ohman, and M. S. Cohen. 1992. Response
of Pseudomonas aeruginosa to pyocyanin: mechanisms of resistance, antioxidant
defenses, and demonstration of a manganese-cofactored superoxide dismutase. Infect.
Immun. 60:328-336.
Hassett, D. J., J. G. Elkins, D.-J. Ma, and T. R. McDermott. 1999a. Pseudomonas
aeruginosa biofilm sensitivity to biocides: use of hydrogen peroxide as a model
antimicrobial agent for examining resistance mechanisms. Meth. Enzymol. In press.
Hassett, D. J., J.-F. Ma, J. G. Elkins, T. R. McDermott, U. A. Ochsner, S. E. H.
West, C.-T. Huang, J. Fredericks, S. Burnett, P. S. Stewart, G. McFeters, L.
Passador, and B. H. Iglewski. 1999b. Quorum sensing'm Pseudomonas aeruginosa
controls catalase and superoxide dismutase genes and mediates biofilm susceptibility to
hydrogen peroxide. J. Bacteriol. Submitted.
Huang, C.-T., K. D. Xu, G. A. McFeters, and P. A. Stewart. 1998. Spatial patterns of
alkaline phosphatase expression within bacterial colonies and biofilm in response to
phosphate starvation. AppL Environ. Microbiol. 64:1526-1531.
Ivanova, A., C. Miller, G. Glinsky, and A. Eisenstark. 1994. Role of rpoS (katF) in
oxyiZ-independent regulation of hydroperoxidase I 'm Escherichia coli. Mol. Microbiol.
12:571-578.
Lechevallier M. W., C. D. Cawthon, and R. G. Lee. 1988. Inactivation of biofilm
bacteria. AppL Environ. Microbiol. 54:2492-2499.
Liss, L. 1987. New M13 host: DH5aF’ competent cells Bethesda Res. Lab Focus 9:13.
Loewen, P. C , and R. Hengge-Aronis. 1994. The role of the sigma factor Gs (KatF) in
bacterial global regulation. Annu. Rev. Microbiol. 48:53-80.
Miller, J. H. 1972. Experiments in Molecular Genetics. Cold Spring Harbor Laboratory,
Cold Spring Harbor, New York.
79
Morita, R. Y. 1993. Bioavailability of energy and the starvation state, in “Starvation in
Bacteria”. Plenum Press, New York, New York.
Nichols, W. W., M. J. Evans, M. P. E. Slack, and H. L. Walmsley. 1989. The
penetration of antibiotics into aggregates of mucoid and non-mucoid Pseudomonas
aeruginosa. J. Gen. Microbiol. 135:1291-1303.
Ostling, J., L. Holmquist, K. Flardh, B. Svenblad, A. Jouper-Jaan, and S.
KjelIeberg. 1993. Starvation and Recovery of Vibrio, in “Starvation in Bacteria”.
Plenum Press, New York, New York.
Sambrook, J., E. F. Frisch, and T. A. Maniatis. 1989. Molecular Cloning: A
Laboratory Manual. 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.
Sanderson, S. S., and P. S. Stewart. 1997. Evidence of bacterial adaptation to
monochloramine in Pseudomonas aeruginosa biofilms and evaluation of biocide action
model. Biotechnol. Bioeng. 56:201-209
Schweizer, H. P. 1993. Two plasmids, X1918 and Z1918, for easy recovery of xylE and
IacZ reporter genes. Gene. 134:89-91.
Schweizer, H. P. 1999. Manuscript in preparation
Seymour, R. L., P. V. Mishra, M. A. Khan, and M. P. Spector. 1996. Essential roles
of core starvation-stress response loci in carbon-starvation-inducible cross-resistance and
hydrogen peroxide-inducible adaptive resistance to oxidative challenge in Salmonella
typhimurium. Mol. Microbiol. 20:497-505.
Simon, R., U. Priefer, and A. Puehller. 1983. A broad host range mobilization system
for in-vivo genetic engineering: transposon mutagenesis in gram negative bacteria.
Bio/Technology 1:784-791.
Stewart, P. S. 1996. Theoretical aspects of antibiotic diffusion into microbial biofilms.
Antimicrob. Agents. Chemother. 40:2517-2522.
Tresse, O., T. Jouenne, and G.-A. Junter. The role of oxygen limitation in the
resistance of agar-entrapped, sessile-like Escherichia coli to aminoglycoside and Plactam antibiotics. J. Antimicrob. Chemother. 36:521-526
Wayne, L. G., and G. A. Diaz. 1986. A double staining method for differentiating
between two classes of mycobacterial catalases in polyacrylamide gels. Anal. Biochem.
157:89-92.
80
Xu, K. D., P. S. Stewart, F. Xia, C.-T. Huang, and G. A. McFeters. 1998.
Physiological heterogeneity in Pseudomonas aeruginosa biofilm is determined by
oxygen availability. Appl. Environ. Microbiol. 64:4035-4039.
CHAPTER 4
CONCLUSIONS
The use of P, aeruginosa as a model biofilm organism and H0O0 as a model
biocide allowed several pertinent questions regarding biofilm physiological adaptation to
antimicrobial agents to be addressed. However, the work presented in this thesis
specifically examines the importance of catalase in protecting P. aeruginosa biofilms
from H2O2. It should be emphasized that studies using other biofilm organisms exposed
to different antimicrobial agents may exhibit results dissimilar to the ones obtained in this
investigation. Nevertheless, determining the importance of oxidative stress metabolism
in protecting biofilms from oxidizing agents represents a relatively new area of research
in microbiology since studies conducted in the past have focused only on the physiology
of planktonic cells. Since little is known about the importance of catalase in protecting
sessile populations of bacteria, relatively simple questions could be directly addressed
with the model system employed in this study.' Stated below are the hypotheses I directly
tested in this work followed by a brief discussion of the results that support or reject each
hypothesis.
Hypothesis I . P. aeruginosa biofilms that cannot constitutively express catalase (KatA )
will be hypersusceptible to H2O2.
o
The data displayed in Fig. 3-2 shows that the VAQKkatA) strain was more susceptible
to the H3O3 treatment than the wild-type in the biofilm disinfection experiments.
However, the survival of the KatA- mutant was much greater in biofilms relative to ;
the substantial and rapid drop in viability that occurred in the disinfection
experiments using planktonic cells. Therefore, constitutive catalase activity due to
KatA expression cannot account for all of the resistance PAOl biofilms display
against H2O2.
Hypothesis 2. Catalase (KatB) induction in response to H2O2 is necessary for biofilm
resistance to H2O2.
o
Catalase induction has little influence on overall biofilm resistance to H2O2. This is
shown in Fig. 3-2 where the survival of the PAOI (katB) strain was nearly identical to
that of the wild-type. Since biofilms were exposed to H2O2 for only I hour, the role
of catalase induction in protecting against prolonged or repeated exposures to
peroxide could not be assessed.
Hypothesis 3. Even in the absence of H2O2, catalase genes (katA, katB, or both) are
upregulated in biofilms resulting in a high specific catalase activity
relative to planktonic organisms.
o
A comparison of specific catalase activities between planktonic (Table 3-2) and
biofilm cells (Fig. 3-3) confirms that catalase levels are nearly equal for both cell
types. Catalase expression patterns in biofilms before and during exposure to
peroxide (Fig. 3-5) also demonstrate that there are no differences in catalase
expression between planktonic cells and biofilms (i.e. katB is not expressed prior to
H2O2 treatment).
Hypothesis 4. P. aeruginosa biofilms initiate an oxidative stress response upon exposure
o
Shown by the rapid increase in catalase activity upon exposure to H2O2 (Fig. 3-4). P.
aeruginosa biofilms do physiologically respond to the oxidant. Also, lack of
induction with the PAQl(WB) confirms that the increase in catalase activity is due
solely to KatB expression. This is also demonstrated by the appearance of the KatB
activity band after peroxide exposure in Native-PAGE gels (Fig. 3-5). Reporter gene
studies also confirmed katB induction (Fig. 3-6b) in biofilms and also showed that
planktonic cells could not adapt to 50 mM H2O2 by inducing katB (Fig. 3-6a) whereas
the biofilm ‘community’ could physiologically respond This data also suggests that the catalase activity produced by the KatA isozyme was insufficient to prevent
oxidative stress in biofilms since expression of KatB is an indicator of an oxidative
stress response.
By using the KatA- and KatB- mutants, the possibility of H2O2 penetration failure
was eliminated along with the biofilms ability to adapt to H2O2 by increasing catalase
activity. However, physiological activity of the cells was likely a variable that impacted
the overall susceptibility of the biofilms to H2O2 (See Chapter 2 for review). Since
biofilms were cultured for 72 hours, the cells near the substratum at the time of
disinfection were 72 hours old. During treatment with 50 mM H2O2, the KatA- biofilm
sloughed from the stainless steel surface leaving behind a deeper (closer to the
substratum), and therefore older, layer of apparently more resistant cells. The survival
84
rates of the KatA- mutant biofilms shown in Fig. 3-2 reflect a relatively high viable
fraction of the total number of cells in the biofilm but it must be emphasized that these
biofilms are also rapidly sloughing from the substratum while the wild-type and KatBmutant biofilms largely retained their integrity.
Planktonic cultures were tested for their sensitivity to H2O2 at 24 hours (stationary
phase). Future antimicrobial efficacy studies should consider overall cell age as an
important factor in resistance rather than just growth phase. Natural populations of
bacteria, whether they occur on the surface of an endo-prosthetic medical devise or in a
drinking water distribution system, are likely to consist of cells that vary widely with age
which is likely to contribute to antimicrobial evasion.
Stress responses to adverse environmental conditions such as nutrient starvation,
heat shock, osmotic shock, desiccation, and oxidative attack have been well characterized
genetically and physiologically in a number of different organisms. Bacteria
predominately exist as biofilms in their natural micro-environments; therefore, future
studies regarding stress responses should be conducted with biofilms rather than
planktonic cells. Conclusions based only on the results of the planktonic disinfection
experiments conducted for this study would erroneously predict that catalase is absolutely
essential in protecting P. aeruginosa from H2O2. However, data collected from biofilms
suggest that other resistance mechanisms are involved since biofilms retained relatively
high viability in the complete absence of catalase activity. I strongly feel that this work
represents ‘a step in the right direction’ regarding the way microbiological research
(specifically microbial physiology) should be conducted in the future. Adopting the
biofilm mode of growth as the standard method for cell cultivation in the laboratory
should yield a more accurate understanding of microbial physiology and the way
microorganisms interact with and survive in their natural environment.
86
APPENDIX
Table A-1. Percent Survival of PAOl (wt) Planktonic Cells During Disinfection.
Time (min.)
Exp. I
Exp. 2
Exp. 3
Mean
S. E.
O
1.0x1002
1.0x1002
1.0x1002
. 1.0x1002
0.0x1000
20
2.0x1001
2.0x1000
7.3x10-01
7.4xlOoo
6.1x1000
40
4.6x1000
7.7x10-03
6.1x10-04
1.5x1000
1.5x1000
60
8.5x10-02
4.2x10-03
3.7x10-05
3.0x10-02
2.8x10-02
Table A-2. Percent Survival OfPAOl(KatAi) Planktonic Cells During Disinfection.
Time (min.)
Exp. I
Exp. 2
Exp. 3
Mean
0
1.0x1002
1.0x1002
1.0x1002
1.0x1002
0.0x1000
20
7.7x10-05
3.6x10-05
3.0x10-05
4.8x10-05
1.5x10-05
40
7.7x10-05
3.6x10-05
3.0x10-05
4.8x10-05
1.5x10-05
60
7.7x10-05
3.6x10-05
3.0x10-05
4.8x10-05
1.5x10-05
S. E.
Table A-3. Percent Survival of PAOI (KatB-) Planktonic Cells During Disinfection.
Time (min.)
Exp. I
Exp. 2
Exp. 3
Mean
0
1.0x1002
1.0x1002
1.0x1002
1.0x1002
0.0x1000
20
9.2x10-01
2.3xl0-oo
3.8xlO-oo
2.3xlO-oo
8.3x10-01
40
5.4x10-04
1.7xlO-oi
2.9x10-03
5.6x10-02
5.4x10-02
60
5.0x10-05
3.0x10-03
1.7x10-04
1.1x10-03
9.6x10-04
S.E.
87
T ab le A -4 . P ercen t S urvival o f P A O I (w t) B io film C ells D uring D isin fectio n ..
Time (min.)
Exp. I
Exp. 2
Exp. 3
Mean
S.E.
100.0
100.0
100.0
100.0
0.0
20
56.0
69.1
78.5
67.9
6.5
40
64.4
94.7
50.9
70.0
13.0
60
98.8
92.8
48.1
79.9
16.0
O
Table A-5. Percent Survival of PAOI (KatA ) Biofilm Cells During Disinfection.
Time (min.)
Exp. I
Exp. 2
Exp. 3
0
100.0
100.0
100.0
100.0
0.0
20
15.3
8.3
15.9
13.2
2.4
40
91.7
24.5
23.5
46.3
22.6
60
1.6
25.2
1.3
9.3
7.9
Mean
S.E.
Table A-6. Percent Survival of PAOl(KatB ) Biofilm Cells During Disinfection.
Time (min.)
Exp. I
Exp . 2
Exp. 3
0
100.0
100.0
100.0
100.0
0.0
20
84.5
68.2
.80.7
77.8
4.9
40
80.8
50.5
38.1
56.5
12.7
60
69.1
60.1
100.4
76.5
12.2
Mean
S. E.
88
T ab le
A -I . P ercent R em oval o f P A O I (w t) B io film C ells D uring D isin fectio n .
Time (min.)
Exp. I
Exp. 2
O
0.0
0.0
20
18.1
40
60
Exp. 3
Mean
S.E.
0.0
0.0
0.0
48.9
4.5
23.8
6.5
78.1
19.2
88.5
61.9
13.0
32.1
4.9
41.0
26.0
16.0
Table A-8. Percent Removal of PAOI (KatA-) Biofilm Cells During Disinfection.
Time (min.)
Exp. I
Exp. 2
Exp. 3
Mean
S.E.
0
0.0
0.0
0.0
0.0
0.0
20
42.7
66.2
5.8
38.2
17.6
40
94.6
96.5
83.0
91.4
4.2
60
94.5
95.8
88.2
92.8
2.4
Table A-9. Percent Removal of PAOI (KatB-) Biofilm Cells During Disinfection.
Time (mm.)
Exp. I
Exp. 2
Exp. 3
0
0.0
0.0
20
5.7
40
60
Mean
S.E.
0.0
0.0
0.0
4.9
53.2
38.2
16.0
-0.1
20.2
57.9
91.4
17.0
20.7
47.5
52.9
92.8
10.0
I
89
Table A-10. Specific Catalase Activity of Stationary Phase, Planktonic PAOl wt, KatA-,
and KatB- Cells Before and After Treatment with 2mM Pulses of H O9
. Every 10 min. for I hr.
Specific Catalase Activity (U/mg protein)
Strain
wild-type
- H2O2
+ H2O2
Exp. I
Exp. 2
Exp. 3
Mean
S.E.
420
613
441
577
418
637
426
609
6.0
14.2
430
432
10.4
9.0
KatA mutant
- H2O2
+ H2O2
None Detected
None Detected
None Detected
None Detected
KatB mutant
- H2O2
+ H2O2
409
423
453
419
None Detected
None Detected
429
454 ■
Table A-I I . Specific Catalase Activity (U/mg protein) of PAOl wt Biofilm During
Disinfection.
Time (min.)
Exp. I
Exp. 2
Exp. 3
0
0.0
0.0
20
5.7
40
60
Mean
S.E.
0.0
0.0
0.0
4.9
53.2
38.2
16.0
-0.1
20.2
57.9
91.4
17.0
20.7
47.5
52.9
92.8
10.0
90
T ab le A -1 2 . S p e c ific C atalase A c tiv ity (U /m g protein) o f P A O l w t B io film D uring
D isin fectio n .
Time (min.)
Exp. I
Exp. 2
0
507
20
Exp. 3
Mean
S.E.
423
495
475
26.2
603
626
517
582
33.2
40
606
625
752
661
45.8
60
618
719
795
717
51.3
Table A-13. Specific Catalase Activity (U/mg protein) of PAOI KatB- Biofilm During
Disinfection.
Time (min.)
Exp. I
Exp. 2
Exp. 3
Mean
S.E.
0
383
478 ■
519
460
40.3
20
. 421
515
470
469
27.1
40
497
460
428
462
19.9
60
421
530
434
462
34.4
Table A-14. PAOI {katB::lacT) Reporter Activity in Miller Units for Biofilms Exposed
to 50 mM H2O2 for I hr.
Time (min.)
Exp. I
Exp. 2
0
■35
33
20
43
40
60
Exp. 3
Mean
S. E.
25
31
3.0
52
37
44
4.2
51
59
47
52
3.5
79
64
43
62
10.5
91
Table A-15. VAO\(katBwlacZ) Reporter Activity in Miller Units for Biofilms Not
Exposed to H2O2.
Time (min.)
Exp. I
Exp. 2
0
23
29
20
29
40
60
Exp. 3
Mean
S. E.
36
29
3.5
27
36
31
2.5
24
28
36
29
3.3
19
28
31
26
3.6
Table A-16. BAO\{katBv.lacZ) Reporter Activity in Miller Units for Planktonic Cells
Exposed to 2mM Pulses of H2O2 Every I Omin. for I hr.
Time (min.)
Exp. I
Exp. 2
0
29
29
20
69
40
60
Exp. 3
Mean
S.E.
34
31
1.7
68
54
64
4.8
82
61
62
68
6.8
87
71
67
75
6.1
Table A-17. PAOl(WRxZacZ) Reporter Activity in Miller Units for Planktonic Cells
Exposed to 1-50 mM Dose OfH2O2.
Time (min.)
Exp. I
Exp. 2
0
33
33
31
32
0.7
20
30
30
30
30
0.0
40
34
32
30 .
32
1.2
60
29
29
29 '
29
0.0
Exp. 3
. Mean
S.E.
.
92
Table A-18. VAO\(katBwlacZ) Reporter Activity in Miller Units for Planktonic Cells
Not Exposed to H2O2.
Time (min.)
Exp. I
Exp. 2
0
26
27
20'
29
■ 40
29
60
31
'
Exp. 3
Mean
S.E.
33
29
2.2
27
30
29
0.9
30
34
31
1.5
29
35
32
1.8