MICROSCOPIC INVESTIGATIONS OF THE ADHESION OF BACTERIA AND
ALGAE ON BIOMATERIAL SURFACES
A Thesis
Presented to
The Graduate Faculty of The University of Akron
In Partial Fulfillment
of the Requirements for the Degree
Master of Science
Pooja Pathak
May, 2007
MICROSCOPIC INVESTIGATIONS OF THE ADHESION OF BACTERIA AND
ALGAE ON BIOMATERIAL SURFACES
Pooja Pathak
Thesis
Approved:
Accepted:
Advisor
Dr. Rex D. Ramsier
Dean of the College
Dr. Ronald F. Levant
Committee Member
Dr. Martha M. Kory
Dean of the Graduate School
Dr. George R. Newkome
Department Chair
Dr. Robert R. Mallik
Date
ii
ABSTRACT
The aim of this investigation was to study the adhesion of bacteria and algae using
optical microscopy, and consists of two parts. The first part involves Pseudomonas
aeruginosa and Staphylococcus epidermidis bacterial adhesion on modified polyvinyl
chloride (PVC – a material with various biomedical applications) surfaces. In this
investigation, PVC substrates were modified by imprint lithography and bacterial
adhesion was performed to determine the influence of the imprinted surface structures on
bio-adhesion. The effects of incubation condition were also studied by performing the
bacterial exposures of the samples under both shaken and stationary conditions. Data
were generated by analyzing the optical microscopy images of the substrates after Gram
staining, using Sigma Scan Pro (Jandel Corporation) software. The results indicate that
bacterial adhesion depends on the surface properties, incubation condition and the type of
bacteria.
The second part of this investigation involves adhesion of cyanobacteria or bluegreen algae on oxidized Zr-705 alloys. Optical microscopy and Sigma Scan Pro software
were used to study the effects of substrate oxide thickness on the propensity for algae
adhesion and growth. The results show that algae adhesion depends on how much the Zr705 surfaces are oxidized. This is relevant to water purification and transport
technologies, since algae growth on piping can adversely affect water quality.
iii
ACKNOWLEDGEMENTS
I would like to express my sincere thanks to my advisor Dr. Rex D. Ramsier for
his support, patience and encouragement throughout my graduate studies. His valuable
suggestions and the time he has given to my thesis will always be appreciated. I’ll always
be grateful to him throughout my life.
I would also like to thank my instructors Dr. Jutta Luettmer- Strathmann and Dr.
Alper Buldum for their guidance and support throughout my course of studies.
I would like to thank Dr. Natalia Farkas for stamping the PVC samples, Ms. Sarah
Smith and Mr. Brad Buczynski for performing the bacterial adhesion, Ms. Deb LaForest
for algae adhesion on Zr-705, and Dr. Nenad Stojilovic for providing some reference
materials.
In addition, I would like to thank my committee members Dr. Martha M. Kory
and Dr. Robert R. Mallik.
iv
TABLE OF CONTENTS
Page
LIST OF FIGURES……………………………………………………………………vii
CHAPTER
I INTRODUCTION……………………………………………………………………1
II LITERATURE REVIEW I ………………………………………………………….4
2.1 Introduction..……………………………………………………………………….4
2.2 Dynamics of Biofilm Formation..………………………………………………….5
2.3 Properties of Biofilms.……………………………………………………………..7
2.3.1 Heterogeneity…………………………………………………………………..7
2.3.2 Resistance.……………………………………………………………………..8
2.4 Biofilms and Disease………………………………………………………………8
2.5 Infection Causing Organisms………………………………………………………9
2.6 Infections and Medical Devices…………………………………………………..11
2.7 Intervention Strategies or Therapy for Infection or Biofilm Control……………..11
III BACTERIAL ADHESION ON PVC SURFACES……………………………….14
3.1 Experimental Details………………………………………………………………14
3.2 PVC Sample Preparation…………………………………………………………..15
3.3 Bacterial Adhesion on PVC………………………………………………………..15
v
3.4 Use of Gram-Staining Techniques…………………………………………………17
3.5 Optical Microscopy.……………………………………………………………….18
3.6 Use of Sigma Scan Pro Software…………………………………………………..21
3.7 Analysis…………..………………………………………………………………..23
3.8 Results and Discussion.……………………………………………………………23
IV LITERATURE REVIEW II.……………………………………………………….32
4.1 Introduction.……………………………………………………………………….32
4.2 Cyanobacterial Toxins and Their Effects.…………………………………………32
4.3 Removal of Cyanobacteria.………………………………………………………..34
V ALGAE ADHESION ON Zr-705 ………………………………………………….36
5.1 Experimental Details.………………………………………………………………36
5.2 Metal Substrate Preparation.……………………………………………………….36
5.3 Algae Sample Preparation.…………………………………………………………37
5.4 Optical Microscopy.………………………………………………………………..37
5.5 Sigma Scan Pro Software.………………………………………………………….37
5.6 Analysis………….…………………………………………………………………39
5.7 Results and Discussion.…………………………………………………………….39
VI CONCLUSIONS……………………………………………………………………41
LITERATURE CITED…………………………………………………………………43
vi
LIST OF FIGURES
Figure
Page
1. Optical images of stamped samples at different magnifications..………………….16
2. Optical images of Pseudomonas aeruginosa after Gram-staining
on stamped and unstamped PVC samples. Images A and C are from
stamped samples and images B and D are from unstamped samples ……………..19
3. Optical images of Staphylococcus epidermidis after Gram-staining
on stamped and unstamped PVC samples. Images E and G are from
stamped samples and F and H are from unstamped samples……………………….20
4. Images of stamped samples after the application of Sigma Scan Pro
software. The red regions are the areas covered by bacteria……….……………….22
5. Percentage area covered by Pseudomonas aeruginosa on unstamped
shaken and stationary samples………………………………………………………24
6. Percentage area covered by Pseudomonas aeruginosa on stamped
shaken and stationary samples………………………………………………………24
7. Percentage area covered by Staphylococcus epidermidis on unstamped
shaken and stationary samples………………………………………………………25
8. Percentage area covered by Staphylococcus epidermidis on stamped
shaken and stationary samples………………………………………………………25
9. Percentage area covered on stamped shaken samples by
Pseudomonas aeruginosa and Staphylococcus epidermidis………………………..27
10. Percentage area covered on stamped stationary samples by
Pseudomonas aeruginosa and Staphylococcus epidermidis……………………….27
11. Percentage area covered by two kinds of bacteria on unstamped
shaken samples…………………………………………………………………….28
12. Percentage area covered by two kinds of bacteria on unstamped
stationary samples…………………………………………………………………28
vii
13. Percentage area of shaken stamped and unstamped samples
covered by Pseudomonas aeruginosa……………………………………………..30
14. Percentage area of stationary stamped and unstamped samples
covered by Pseudomonas aeruginosa……………………………………………..30
15. Percentage area of shaken stamped and unstamped
samples covered by Staphylococcus epidermidis…………………………………..31
16. Percentage area of shaken stamped and unstamped
samples covered by Staphylococcus epidermidis…………………………………31
17. Optical images of blue-green algae on Zr-705 surfaces…………………………..38
18. Percentage area covered by blue-green algae on Zr-705 samples
with different oxide layer thicknesses on their surfaces…………………………..40
viii
CHAPTER I
INTRODUCTION
Biofilms adhering to implants and medical devices present a considerable risk of
infection to patients. According to an estimate, biofilms account for more than half of the
microbial infections in the body [1]. Many of these infections are associated with the use
of biomedical devices such as prosthetic heart valves, contact lenses, cardiovascular
implants, orthopedic replacements, intravascular and urinary catheters, and dental unit
water lines [2]. Global utilization of bio-medical devices is valued at $86 billion per
annum with a growth rate of 7% each year [3], which indicates a move toward the use of
more and more bio-medical devices and thus an eventual rise in device-related infections.
In most cases, removal and replacement of the device is the only effective therapy for
these infections [4]. However, removal can cause complications and impact human
health, as well as increase medical care costs [5]. Therefore, it is important to develop
preventive measures to eliminate or reduce device-related infections.
One method to accomplish this is to modify the physicochemical interactions
between bacteria and the device (substrate) through surface modification [6]. Various
techniques have been used, such as oxygen plasma treatment [7], wet chemical methods
using organic reactions [8], Ni-P-PTFE (Nickel-phosphorus-polytetrafluoroethylene)
coatings [9], and diamond-like carbon (DLC) films [4]. Such techniques have proven to
1
be effective in reducing bacterial adhesion and biofilm formation. The technique used in
this study was nanoimprint lithography (NIL), a method which allows complete freedom
in designing the size, shape, and spacing of a pattern and is capable of producing sub 10
nm features over a large area with high throughput and low cost [10]. NIL was used in
this research to investigate its potential to alter the physicochemical properties of
polyvinyl chloride (PVC) surfaces.
PVC is extensively used in biomedical devices due to its wide range of
advantageous properties such as ease of processing, chemical resistance, and low cost
[4,8]. The modified PVC surfaces were studied with respect to adherence of two
clinically relevant bacterial strains of Pseudomonas aeruginosa and Staphylococcus
epidermidis under different incubation conditions. The results show a dependence of the
bacterial adhesion on the surface characteristics of the modified polymer. The samples
were analyzed with optical microscopy and Gram staining techniques and the results are
described and discussed in Chapter III.
Separately in this thesis there is also a study of another relevant issue concerned
with bio-adhesion i.e., contamination of drinking water by blue-green algae or
cyanobacteria. Cyanobacteria are a division of bacteria that are extremely toxic and are
capable of causing serious illness or even death if consumed [11]. The adherence of
cyanobacteria to drinking water lines is one source of such contamination [12].
Removing cyanobacteria from water by harsh physical or chemical treatment leads to
lysis of the cells and the release of cyanobacterial toxins, most of which are water soluble
[13]. Therefore, this problem is attracting the attention of environment agencies, water
authorities and human and animal health organizations [14].
2
Our proposal to deal with this problem is to replace the pipe material with alloys
that can resist or reduce algae adhesion. Usually drinking water pipes are made of PVC,
copper, stainless steel, polythene, or cement and generally drinking water is disinfected
by chlorine [15,16]. However, according to M. Lehtola et al., the pipeline material can
modify the effectiveness of disinfectants in drinking water distribution systems [15]. In
this research Zr-705 was used to test its propensity for algae adhesion as a potential
piping material. Zr-705 was chosen because previous work [17] on zirconium alloys
provided evidence for reduced bacterial adhesion and biofilm formation on their surfaces.
Zr-705 as a potential material for water pipes was investigated in this project by changing
the thickness of the oxide layer on its surface and investigating the effect of oxidation
with respect to algae adhesion using optical microscopy and Sigma Scan pro software
(Jandel Corporation).
3
CHAPTER ІІ
LITERATURE REVIEW Ι
2.1 Introduction
Bacteria in natural settings attach to surfaces, aggregate and form a complex
structure known as a biofilm [18]. The existence of bacteria in biofilm form profoundly
differs from bacteria in planktonic (free-floating) form, in that biofilms are inherently
resistant to anti-microbial agents [19]. This property is primarily attributed to the
production (secretion) of extra-cellular polymeric substances (EPS) during their
formation [20]. Because of the “resistance” property, biofilms cause considerable adverse
effects in the medical [19], environmental [1], and industrial [21] fields. Therefore,
researchers from various disciplines such as chemistry, engineering, mathematics, and
biology are investigating biofilms [22].
It was known in the early 1980’s that 99.9% of bacteria in aquatic ecosystems
lived on surfaces [23]. However, up to the late 1980’s it was thought that biofilms were
random accumulations of cells in matrix material [23]. Early in the 1990’s it became
clear that most biofilms were composed of structured micro-colonies and water channels
through which nutrients and oxygen circulated [21,23]. Further, micro-colonies were
composed of matrix material and cells, and this basic micro colony/water channel pattern
was observed in most natural biofilms like dental plaque [23].
4
The exact definition of biofilms has evolved over the past few decades and can be
defined as highly organized [21] and complex communities [23] of microorganisms that
are irreversibly attached to a surface or interface or to each other, secrete extra-cellular
polymeric substances (EPS) to form a matrix, and exhibit an altered growth rate and gene
transcription [2]. This definition considers two characteristics of biofilm organisms,
altered growth rate and gene transcription that planktonic organisms do not possess,
which are particularly useful in differentiating biofilms from “nonbiofilms”. The matrix,
formed by microorganisms, protects them from UV radiation, extreme pH values,
dehydration, antibiotics, and the immune system of the host organism. In this way, by
forming a biofilm, bacteria survive inside and outside the host organism [21].
Bacterial cells in a biofilm have close proximity and high density, with a potential
for inter-species communication, competition, and cooperation [22]. Specifically, the
cell-cell or interspecies communication system called Quorum sensing [22] is one of the
basic [2] and essential biofilm processes [23], which provides a continuous exchange of
information through specific chemical molecules [23]. It is an important feature of
microbial communities and allows bacteria to live as multicellular organisms [21]. For
instance, in some cases, it provides aid in different stages of biofilm formation (initial
attachment, maturation) [1,24].
2.2 Dynamics of Biofilm Formation
The first stage of biofilm formation is initial adhesion in which bacteria in
planktonic (floating) form reach a surface and under favorable conditions attach to a
surface to form clusters [21]. This is called the first maturation stage. This stage is
5
governed by two types of interactions, van der Waals and electrostatic. Van der Waals
interactions are usually attractive and electrostatic forces are usually repulsive (due to
both bacteria and surfaces being negatively charged in nature) [25]. Bacteria have a sense
of touch, which enables them to detect a surface [1]. Clusters of bacteria further combine
to form micro-colonies (which are the basic building blocks of biofilms [2]), and this is
called the second maturation stage. Further, these micro-colonies combine to form an
exopolymer matrix and secrete EPS to form a complex structure [21]. Finally, the last
stage called dissociation or detachment is when bacteria enclosed in the EPS matrix
detach from the matrix and return to planktonic form, depending on the availability of
nutrients, oxygen and iron [1,21].
Bacteria must be able to detach from mature biofilms in order to avoid starvation
due to the population density of attached bacteria and to colonize new surfaces [26]. In
order to detach, a polysaccharide lyase (an enzyme that degrades polyanionic substrates
[27]) helps bacteria to return to their planktonic state. Otherwise, the matrix can become a
graveyard for bacteria under low nutrient conditions, unfavorable environmental factors
and unwanted neighbors [28]. The biofilm formation process exhibits directionality,
meaning that there are more chances of bacteria to accumulate and form a biofilm than
moving back to planktonic states [22]. In this way, biofilm development is sustained and
the detached bacteria find new surfaces for colonization under favorable conditions.
6
2.3 Properties of Biofilms
Heterogeneity and resistance (towards antimicrobial agents) are two remarkable
properties of biofilms and are discussed below.
2.3.1 Heterogeneity
Previously, there was an assumption that biofilms are homogenous in composition
and properties such as porosity, pore size division, density and microbial populations
varied little with depth. Due to EPS there would be confined heterogeneity, but as a
whole the biofilms were assumed to be homogenous [29]. However, in recent years, it
has been shown that biofilms are structurally, chemically and physically heterogeneous,
thus making them complex [22].
The heterogeneity can be explained on the basis of gradients of nutrients, waste
products and signaling factors [19]. Specifically, in the case of structural heterogeneity,
gradients of nutrients and oxygen determine the difference in density of bacteria
throughout the biofilm [30]. As the cells at the edge of the biofilm have better access to
nutrients as compare to cells within the interior, this results in the formation of
metabolically active and inactive cells, respectively, eventually making biofilms
structurally heterogeneous [22]. In addition, heterogeneity can be enhanced by the
presence of multiple species. For example, the biofilm involved in endocarditis involves a
single species, but in tooth decay, multi-species biofilms are present [31].
7
2.3.2 Resistance
Resistance is the ability of microorganisms to grow in the presence of high levels
of anti-microbial agents like antibiotics, disinfectants, germicides, etc., and is attributed
to the biofilm structure and its organism’s physiology [2,32]. Since the structure of
biofilms consists of an EPS matrix, the anti-microbial species must diffuse through this
matrix in order to inactivate the enclosed cells. However, EPS creates a barrier for the
anti-microbial species and either influences their rate of movement to the biofilm interior
or their reaction with the enclosed cells [2]. In addition, the growth rates of biofilmassociated cells are slower than planktonic cells, and slow growing cells are not as
susceptible to many anti-microbial agents [1,2]. The fact that biofilm bacteria are
different from their planktonic counterparts is clear from the data reported by Lewis et.
al, which show that the concentration of anti-microbial agents needed to kill biofilm
bacteria is 10-1000 times that for planktonic bacteria [32].
2.4 Biofilms and Disease
The “Resistance” property of biofilms toward anti-microbial agents makes them
adaptable to various environments. As such, biofilms cause infectious diseases like cystic
fibrosis, peridontitis, etc., and infections due to the use of medical devices like prosthetic
heart valves, catheters, etc., [2]. In fact, more than half of the infectious diseases in the
body are due to biofilms [1]. However, the exact processes by which biofilm organisms
cause disease are not completely understood. The current understanding of the
mechanism describes the processes in four stages. These are:
8
(1) Detachment: Detachment of cells from biofilms, as a result of cell growth and
division within the biofilms, has the greatest impact on the outcome of chronic bacterial
infections [2]. For example, the shedding or detachment can likely increase the risk of
stroke in the case of heart valves and a very small number of bacteria can cause
bloodstream and urinary tract infections [2].
(2) Production of endotoxin by Gram-negative bacteria inside the biofilms
attached to indwelling medical devices. Endotoxin, a lipopolysaccharide (LPS), is a cell
wall constituent of Gram-negative bacteria [33]. When Gram-negative bacteria multiply
or die, LPS is released [34]. Endotoxin production may draw out an immune response in
the human host [2].
(3) Organisms detaching from medical device biofilms could exhibit resistance to
the host immune system and cause an infection.
(4) Conjugation: Conjugation is an exchange of genetic information between
different members of the same bacterial species for the continued survival of an organism
[35]. Through conjugation within biofilms, bacteria can exchange plasmids [2]. Plasmids
are substances that carry genes that encode proteins required for mating pair formation
[35]. From these plasmids, resistance factors may be carried forward, providing an
environment for future generations of resistant organisms [2].
2.5 Infection Causing Organisms
Organisms can be divided into two categories, pathogenic and non-pathogenic.
Pathogenic organisms are those associated with infection such as Staphylococcus
epidermidis [3] and Pseudomonas aeruginosa [36]. Non-pathogenic organisms are those,
9
which colonize on surfaces but are not associated with infection, for example,
Lactobacillus [3]. Common pathogens in medical settings are Staphylococcus
epidermidis and Pseudomonas aeruginosa, whose biofilms are well known for their
antibiotic resistance [37].
Out of these two, Staphylococcus epidermidis has become a major cause of
nosocomial (hospital) infections. In fact, according to the 1998 National Nosocomial
Surveillance System report, Staphylococcus epidermidis was the most important
pathogen involved in nosocomial blood stream infections, cardiovascular infections, and
infections of the eye, ear, nose and throat [38]. It is categorized under staphylococcus, a
Gram- positive bacterium having spherical shape that aggregates in clusters [39]. The
infections caused by S. epidermidis are usually chronic because of the lack of severely
tissue damaging toxins [38]. It is the normal inhabitant of skin and mucous membranes of
human body [38]. Under normal circumstances, it is a non-pathogen. But in the presence
of a mildly compromised individual or a medical device, it becomes the leading cause of
infection [4] because of its ability to adhere to surfaces, colonize and to form a multilayered biofilm [38].
Another common bacterium related to infection is Pseudomonas aeruginosa, a
pathogen and one of the top three causes of opportunistic human infections [36]. It has
the ability to form biofilms readily on most surfaces and plays a role in catheter and
contact lens related infections, etc. [1]. It is also one of the major causes of cystic fibrosis,
a chronic lung infection [1] and is associated with the high rate of patient mortality. The
clinical strains of P. aeruginosa are characterized by the formation of EPS, which
provides protection from anti-microbial agents [21]. In case of ventilator associative
10
pneumonia (VAP), colonization of the intubations by P. aeruginosa leads to a mortality
rate of over 60% despite antibiotic therapy [7].
2.6 Infections and Medical Devices
Polymers are among the various materials used for medical devices. They are
widely used because they are durable, shapeable and have low production cost [40].
Apart from that, they are highly versatile and have excellent physicochemical properties
[8]. Among all the polymers, polyvinyl chloride (PVC) is outstanding because it has
broad range of processing potential and shows high mechanical and chemical resistance
[8]. For more than 70 years, PVC has been one of the most important biopolymers [41].
The biomedical devices that are made of PVC are blood bags, ventilation tubes, blood
transfusion tubes, indwelling catheters and urinary catheters. In fact, more than 5% of the
PVC produced is utilized in the medical arena [8].
Regarding infection, most plastic and metal surfaces of medical devices will
develop bacterial biofilms when bacteria are present in the body fluids. Almost all body
fluids (such as blood, urine, saliva, etc.,) provide sufficient organic nutrients for bacterial
growth. In spite of shear forces, such as those generated on heart valves by blood flow
and on contact lenses by blinking, the development of biofilms occurs [42]. Therefore, it
is highly likely that infection will occur with the use of medical devices.
2.7 Intervention Strategies or Therapy for Infection or Biofilm Control
The adverse effects caused by common pathogens on medical devices can be
avoided or minimized by adopting a few strategies. First and foremost is to prevent initial
11
device contamination by using disinfectants “in time” before a biofilm develops. As the
initial step of biofilm formation (attachment) is a fast process and takes only a few hours,
disinfectants should be used quickly [2,43]. For example, in the case of S. epidermidis
infections during the insertion of catheters, specific hygienic procedures such as the use
of antibiotics during surgery and covering the implanted material with antibiotics should
be used [38].
A second strategy is to minimize or inhibit initial microbial cell attachment to the
device by incorporating biocides into the surface or by selecting surface materials that do
not promote attachment [2,43]. A third strategy is to penetrate the biofilm matrix and kill
the biofilm-associated cells using harsh disinfectants or biocides. As the chemical
composition of EPS significantly differs from biofilm to biofilm, non-specific measures
should be used [2,43]. Finally, if everything else fails and medical devices become
infected, the last option is to remove the device [2]. Out of all the strategies, surface
modification has been widely studied as a preventive measure, to minimize the
attachment and growth of microbes. This has been accomplished by modifying the
surfaces of existing bio-medical devices. Various techniques (such as plasma, corona,
laser treatment) have been used for surface modification [44].
Apart from the above-mentioned treatments, thermal energy and ultrasound can
also be used for inhibiting bacterial growth. According to a report by Mott et al.,
ultrasound treatment reduced biofilm growth and increased the efficiency of biocides
against biofilms [45]. Heat treatment at 65º C with sodium hypochlorite (an antimicrobial
agent), has been used to inactivate L. monocytogenes biofilms associated with stainless
steel surfaces [46]. Finally, ultraviolet (UV) radiation can modify polymer surfaces from
12
hydrophobic to hydrophilic. This modification enhances cell growth and therefore has
potential applications in tissue engineering [40,47].
The methods mentioned above may be preferred to others depending on the
medical device, implantation site, and restrictions imposed by fabrication methods [44].
However, biofilms are diverse due to factors such as the surface in question, availability
of oxygen and nutrients, type of microbial species, flow velocity of surrounding liquid,
etc. Therefore, information gathered from studies of one particular biofilm cannot
necessarily be transferred to another biofilm [43]. This makes the task of preventing or
reducing biofilm-related infection challenging. Considering this, another surface
modification technique, nano-imprint lithography (NIL) was used in this research.
Imprint lithography has applications in bio-medical [48] and semiconductor [10] devices.
In imprint lithography, a pattern is created by pressing a three dimensional stamp into a
resist [49] and a variety of shapes with varying physicochemical properties can be
constructed [50]. As compared to traditional photolithography, it is less expensive and
takes fewer steps to complete [50].
13
CHAPTER III
BACTERIAL ADHESION ON PVC SURFACES
3.1 Experimental Details
PVC is widely used as a material for bio-medical devices such as urinary
catheters, blood bags, and ventilation tubes [8]. One of the major disadvantages with
PVC is its susceptibility to infections related with these devices, which adversely affects
humans and health care costs [51]. Several approaches have been used to overcome this
issue. In the work reported here, surface modification by nano-imprint lithography was
performed on PVC to test its potential to minimize bacterial adhesion.
In total, eight samples of PVC and two types of bacteria were used. Four were
imprinted (stamped) samples and four non-imprinted (unstamped) samples. Out of these
eight, four samples (two stamped and two unstamped) were exposed to Pseudomonas
aeruginosa, and the remaining four to Staphylococcus epidermidis. Also, out of four
samples exposed to P. aeruginosa, two were kept stationary during exposure (one
stamped and one unstamped) and two were exposed under shaken condition (one stamped
and one unstamped). Similarly, four such samples were exposed to S. epidermidis
bacterium.
14
3.2 PVC Sample Preparation
Nano-imprint lithography was performed on four samples, out of eight, with a
silicon stamp as the master stamp. The dimensions of each sample were 1.5 X 2.5 cm.
The silicon stamp was prepared by standard photolithography and consisted of different
sized and shaped geometrical features.
The PVC sample and silicon stamp were clamped between two microscope
slides and placed into a commercial microwave oven for 4 seconds. During the
imprinting process, the PVC melts by selective heating and flows into the protrusions of
the silicon stamp. In this way, four samples of PVC were prepared by imprinting. Figure
1 presents optical images of stamped samples at different magnifications.
3.3 Bacterial Adhesion on PVC
Two types of bacteria, Pseudomonas aeruginosa and Staphylococcus epidermidis
were used in this investigation because of their clinical relevance. First of all, tryptic soy
broth (TSB) was prepared and sterilized in 500 ml flasks. Then, P. aeruginosa and S.
epidermidis were grown in 5 ml of TSB. All eight samples were sterilized in an
autoclave. Four samples were then aseptically added to four different flasks with P.
aeruginosa and four with S. epidermidis. Two flasks with P. aeruginosa and two with S.
epidermidis were left stationary in a 37° C incubator for three days. The remaining flasks
were placed in an incubated shaker at 37° C, with continuous agitation at 17 rpm for three
days.
15
⎯⎯− 10 µm
50X
⎯⎯⎯− 100µ m
-
100X
− 10 µm
10X
⎯ 100 µm
2X
Figure 1. Optical images of stamped samples at different magnifications.
16
After three days, the samples were aseptically removed from the flasks and rinsed
in 200 ml of sterile water for one minute. This was performed by holding a corner of the
sample with a pair forceps and dipping the sample in and out of the water. The purpose of
rinsing was to remove the unattached and loosely attached bacteria from the substrate
surface. Liquids commonly used for rinsing include sterile water, normal saline, and
phosphate buffered saline [52]. After rinsing, the samples were placed into sterile petri
dishes with sterile filter paper.
3.4 Use of Gram-Staining Techniques
Gram-staining was applied to all the samples after bacterial adhesion as an aid to
identifying bacteria microscopically. This method was originally developed by Hans
Christian Gram, a Danish clinician [53]. This method differentiates bacteria into two
fundamental groups, Gram-positive and Gram-negative.
Gram-positive bacteria are those that retain the initial crystal violet stain, whereas
Gram-negative bacteria are those that are decolorized and stain red with safranin [53].
Gram-positive bacteria have a thick cell wall made of peptidoglycan that resists decolorization [53]. Peptidoglycan can be regarded as a functional analogue of
lipopolysaccharide. Both molecules activate intrinsic immune defense mechanisms
[54,55]. Gram-negative bacteria have a thin peptidoglycan layer plus an overlying liquidprotein bi-layer known as the outer membrane, which can be disrupted by decolorization. Staining does not alter the shape and form of bacteria. Thus, this method
17
also permits a determination of the overall structure of the cells for example, cocci, rods,
spirals, filaments, cubic packets, etc. [53].
S. epidermidis is a Gram-positive bacterium [39] and P. aeruginosa is Gramnegative [7]. For a typical PVC sample with bacteria, the sample was held at a corner and
then flooded with crystal violet dye for 10 seconds and washed with running water. Then,
the sample was flooded with iodine for 10 seconds and again washed with water. After
about 30 seconds, the sample was rinsed with ethyl alcohol, a decolorizing fluid and then
with safranin (red dye) for 10 seconds followed by washing with water.
3.5 Optical Microscopy
After Gram-staining, the samples were observed and analyzed using an optical
microscope (Mitutoyo). Each sample was held at a corner and mounted onto a glass slide.
Images of each sample were obtained with different magnifying power i. e. 2X, 10X, 50X
and 100X. Images were captured with a digital camera (Duncan DT 4000 RGB CCD)
camera, which was interfaced to a computer. At different magnifying powers of the
objective lens, images of a 1 mm scale were also captured to calibrate the images. Figures
2 and 3 are representative of the images collected in this work.
18
⎯⎯− 10 µm
A
⎯⎯− 10 µm
50X
B
50X
⎯⎯− 10 µm
C
⎯⎯− 10 µm
50X
D
50X
Figure 2. Optical images of Pseudomonas aeruginosa after Gram-staining on stamped
and unstamped PVC samples. Images A and C are from stamped samples and images B
and D are from unstamped samples.
19
⎯⎯− 10 µm
E
50X
⎯⎯− 10 µm
F
50X
⎯⎯− 10 µm
G
50X
⎯⎯−
H
10 µm
50X
Figure 3. Optical images of Staphylococcus epidermidis after Gram-staining on stamped
and unstamped PVC samples. Images E and G are from stamped samples and F and H are
from unstamped samples.
20
3.6 Use of Sigma Scan Pro Software
Optical microscopy images were analyzed using Sigma Scan Pro software. This
software provides a complete image analysis package for studying the structure and size
of objects in images. Its powerful image analysis and data manipulation techniques
transform images into reliable statistics. In the case of biological sciences, this software
quickly counts and measure cells, organisms, or features [56]. In this research, this
software was employed to measure the area covered by bacteria on PVC samples from
optical microscopy images. For areal measurements, calibration or rescaling was
performed. Calibration can be performed in two modes, spatial or intensity calibration. In
this research, spatial calibration was applied. Spatial calibration calibrates distance and
area by converting raw pixel coordinates into specified measurement units. It can be
performed in three ways; one-point, two-point or three-point. In this research, two-point
rescaling was applied.
After selecting the calibration option and defining the number of pixels equal to 1
unit (1 unit = 730 pixels=0.1 mm), the areal measurement option was selected. This basic
setting was applied on all the images with one particular magnifying power of the
objective lens, i. e. 50X. For each sample, the portion selected for measurement was that
which contained the maximum number of bacteria. The total area covered by bacteria
was measured, as was the area of the entire image. Figure 4 demonstrates how the images
were analyzed.
21
⎯⎯− 10 µm
⎯⎯− 10 µm
Figure 4. Images of stamped samples after the application of Sigma Scan Pro software.
The red regions are the areas covered by bacteria.
22
3.7 Analysis
The percentage area covered by bacteria was calculated by taking the ratio of the
area covered by bacteria to the total area of the image. These data are presented in
histogram format with an experimental uncertainty of ± 5%.
3.8 Results and Discussion
Results were organized under three categories. (1) Incubation condition (shaken
and stationary), (2) Bacterial strains (S. epidermidis and P. aeruginosa), and (3) Surface
modification (stamped and unstamped). The samples obtained with S. epidermidis were
stamped shaken, stamped stationary, unstamped shaken and unstamped stationary.
Similar samples were prepared with P. aeruginosa bacterium.
Figure 5 and Figure 6 are representative of the first category, i. e. incubation
condition. Figure 5 represents unstamped samples with P. aeruginosa. Samples exposed
to bacteria under shaken conditions exhibit more adhesion than stationary samples.
Similar results were obtained with stamped samples also, as shown in Figure 6. For S.
epidermidis bacterium, Figure 7 represents unstamped samples. Samples under shaken
conditions had more adhesion than stationary ones, however stamped samples under
shaken conditions had less bacteria than stationary samples (Figure 8). These results
indicate that except for one system, i. e. stamped shaken (S. epidermidis), all the samples
under shaken conditions had more bacterial growth than stationary samples. These results
generally agree with previous results obtained from Zry-2 and Zr-705 surfaces [57].
23
20
18
16
Area(%)
14
12
10
8
6
4
2
0
Unstamped Shaken
Unstamped Stationary
Incubation Condition
Figure 5. Percentage area covered by Pseudomonas aeruginosa on unstamped shaken and
stationary samples.
35
30
Area(%)
25
20
15
10
5
0
Stamped Shaken
Stamped Stationary
Incubation Condition
Figure 6. Percentage area covered by Pseudomonas aeruginosa on stamped shaken and
stationary samples.
24
14
12
Area(%)
10
8
6
4
2
0
Unstamped Shaken
Unstamped Stationary
Incubation Condition
Figure 7. Percentage area covered by Staphylococcus epidermidis on unstamped shaken
and stationary samples.
14
12
Area(%)
10
8
6
4
2
0
Stamped Shaken
Stamped Stationary
Incubation Condition
Figure 8. Percentage area covered by Staphylococcus epidermidis on stamped shaken and
stationary samples.
25
For the next category, type of bacteria, Figures 9-12 represent the data. Only in
the stamped shaken case did P. aeruginosa cover more percentage area than S.
epidermidis (Figure 9). In all other samples, stamped stationary (Figure 10), unstamped
shaken (Figure 11) and unstamped stationary (Figure 12), S. epidermidis covered more
percentage area than P. aeruginosa.
These results indicate that overall S. epidermidis had more growth on PVC
samples than P. aeruginosa. One possible explanation can be based on their shapes. P.
aeruginosa is rod-shaped [58] and S. epidermidis is spherical [39]. A rod has a large
surface area per unit volume [59]. The rate of bacterial growth depends on the rate of
intake of nutrients, and this takes place through the cell surface. Large surface area will
lead to increases in bacterial growth [59]. On this basis, P. aeruginosa should have more
growth than S. epidermidis. However, in some conditions such as desiccation or osmotic
shock (dehydration and re-hydration within 1 second, [60]), the reverse can be true. In
hypertonic environments (where the concentration of solutes is larger in the environment
than in the cell, such as salt water [61]), organisms with large surface area will lose
water. Therefore in this case S. epidermidis has an advantage over P. aeruginosa.
26
35
30
Area(%)
25
20
15
10
5
0
P. aeruginosa
S. epidermidis
Bacteria
Figure 9. Percentage area covered on stamped shaken samples by Pseudomonas
aeruginosa and Staphylococcus epidermidis.
13
12.5
Area(%)
12
11.5
11
10.5
10
9.5
P. aeruginosa
S. epidermidis
Bacteria
Figure 10. Percentage area covered on stamped stationary samples by Pseudomonas
aeruginosa and Staphylococcus epidermidis.
27
Area(%)
20
18
16
14
12
10
8
6
4
2
0
P. aeruginosa
S. epidermidis
Bacteria
Area(%)
Figure 11. Percentage area covered by two kinds of bacteria on unstamped shaken
samples.
7.2
7
6.8
6.6
6.4
6.2
6
5.8
5.6
5.4
5.2
P. aeruginosa
S. epidermidis
Bacteria
Figure 12. Percentage area covered by two kinds of bacteria on unstamped stationary
samples.
28
Surface modification is the last category. Figure 13 represents samples shaken
with P. aeruginosa having stamped and unstamped surfaces. The stamped samples had
more bacteria than the unstamped ones. A similar result was obtained under stationary
conditions as well (Figure 14). In the case of S. epidermidis, under shaken conditions
stamped samples had less adhesion than unstamped ones (Figure 15), opposite to the
trend for stationary conditions (Figure 16).
These results show that stamping (imprinting) increased bacterial adhesion overall
on these PVC samples. A possible reason could be the roughening of the surface by
imprint lithography, which leads to increased bacterial growth. According to a report by
Harris et al, surface roughness enhances bacterial growth [62] and it has been shown that
surface modification can be used to guide cells to selected areas of medical devices [63].
Therefore, our results are consistent with literature by demonstrating that our imprinted
PVC surfaces do encourage, on average, more bacterial growth.
29
35
30
Area(%)
25
20
15
10
5
0
Stamped Shaken
Unstamped Shaken
Imprinting
Figure 13. Percentage area of shaken stamped and unstamped samples covered by
Pseudomonas aeruginosa.
14
12
Area(%)
10
8
6
4
2
0
Stamped Stationary
Unstamped Stationary
Imprinting
Figure 14. Percentage area of stationary stamped and unstamped samples covered by
Pseudomonas aeruginosa.
30
Area(%)
20
18
16
14
12
10
8
6
4
2
0
Stamped Shaken
Unstamped Shaken
Imprinting
Figure 15. Percentage area of shaken stamped and unstamped samples covered by
Staphylococcus epidermidis.
14
12
Area(%)
10
8
6
4
2
0
Stamped Stationary
Unstamped Stationary
Imprinting
Figure 16. Percentage area of shaken stamped and unstamped samples covered by
Staphylococcus epidermidis.
31
CHAPTER ΙV
LITERATURE REVIEW ΙΙ
4.1. Introduction
The remainder of this thesis deals with the topic of cyanobacteria and their effects
in drinking water distribution systems. Cyanobacteria or blue-green algae are among the
most ancient groups of organisms on earth [64]. This is clear from the calcareous
accumulations formed by cyanobacterial mats in the estuarine waters of western
Australia, which are more than one billion years old [64]. They evolved as the oxygenic
plant-type photosynthetic organisms [65], and were the first to produce molecular oxygen
as a byproduct of photosynthetic activity [66]. Over 2,000 species of cyanobacteria have
been identified in the form of single cells, colonies, and filaments [13]. They are formed
globally, in all types of aquatic environments ranging in size from garden ponds to
oceans [64]. Cyanobacteria can undergo rapid growth to form blooms in the presence of
chlorophyll and necessary photosynthesis pigments, and blooms are widely recognized as
a source of taste and odors in water supplies [11].
4.2 Cyanobacterial Toxins and Their Effects
Apart from bad taste and odor, cyanobacteria produce toxins called cyanotoxins
with which various health issues are associated [66]. There are more than 60 identified
32
toxins of cyanobacteria, which are classified as neurotoxins, hepatotoxins, cytotoxins,
skin irritants and gastrointestinal toxins [66]. They pose a potential health risk in
recreational waters and drinking water reservoirs. Adverse effects due to hepatotoxins
can be liver damage and tumor growth promotion. In the case of neurotoxins, acute
poisoning can result in death by paralysis and respiratory failure. Endotoxins can cause
eye irritation, skin rashes and respiratory allergies [11]. Cyanobacteria can affect the
human body through various exposure routes such as skin contact, inhalation, and
ingestion. Cases of dermal contact have been reported for at least 30 years in which
rashes, blisters, asthma, conjunctivitis, and ear and eye irritation and allergic reaction
occurred after contact with cyanobacteria in coastal waters [14].
The ingestion or oral route is the most investigated because of normal water
intake and accidental recreational intake [14]. According to the report of I. R. Falconer,
maximum outbreaks of poisoning by toxic blue-green algae occurred in chlorinated tap
water supplies [67]. An outbreak of hepatoenteritis due to ingestion of cyanobacteria
from a drinking water supply reservoir was observed in Australia in 1979. This severe
outbreak resulted in about 70% of the patients receiving intravenous treatment. The
symptoms of this outbreak were headache, abdominal pain, lethargy, diarrhea, acidosis,
and injury to the liver, kidneys, lungs, and intestines [14].
Microcystins, which is one of the most common and toxic [12] of the
cyanobacterial toxins, is present in surface waters used for drinking water production.
Due to its toxicity, the World Health Organization (WHO) has published a provisional
guideline for its toxicity in drinking water [68]. Guidelines for cyanobacterial toxins in
water exist in several countries worldwide [68].
33
4.3 Removal of Cyanobacteria
Because of the adverse effects of cyanobacteria, it is necessary to check all
potential sources of cyanobacterial contamination that may contact with drinking water
[13]. Removing cyanobacteria from water by harsh physical or chemical treatment leads
to lysis of the cells and release of cyanobacterial toxins, most of which are water soluble
[13]. Therefore remedial procedures must involve reducing or completely removing the
toxins from drinking water [13]. Drinking water is usually disinfected by chlorine,
chlorine dioxide or chloramines [15]. However, in case of household plumbing the
microbial concentration can increase because of higher temperatures, concentrations of
metals like copper and iron, and the decrease or absence of chlorine. This makes
household plumbing the most problematic part of the distribution system.
Cyanobacteria and micro-algae are major components of biofilms associated with natural
aquatic environments [69]. For disinfection, UV irradiation is another possibility.
However, this treatment is possible in low nutrient concentrations and in low water
temperature systems [70]. Under other conditions, this treatment is less effective than
chlorine dioxide because it can increase bacterial regeneration and proliferation [71].
Drinking water pipes are usually made of steel, PVC, copper, etc. [15,16]. Materials used
in pipes and coatings may promote microbial growth due to leaching of chemical
compounds, which microorganisms may use as nutrients [71]. On this basis the problem
of blue-green algae in drinking water may be solved by replacing the pipe material.
Keeping this point in mind, the present research investigated the potential of Zr-705 as a
pipeline material. Selection of this particular material is due to the oxide layer on its
surface, which makes it corrosion resistant and this property has been shown to give
34
satisfactory results in bacterial adhesion studies [57]. Zr-705 samples with different oxide
layer thicknesses on their surfaces were studied in this research with respect to inhibiting
algae growth.
.
35
CHAPTER V
ALGAE ADHESION ON Zr-705
5.1 Experimental Details
Zr-705 was the material used for this part of the investigation, with different
thicknesses of oxide layers on its surface. This work builds upon our previous efforts to
study bacterial adhesion on zirconium alloys [17,57].
5.2 Metal Substrate Preparation
All the samples for this section were prepared using Zircadyne-705 (Zr-705,
nominally 2.5 % Nb, balance Zr + Hf). Five samples were prepared with different oxide
layer thickness. One sample was un-oxidized, the other samples had oxide layer
thicknesses of 2.1 µm, 4 µm, 11.1 µm and 12 µm. Before oxidation in an oven, each
sample was polished on one side using diamond paste and an alumina suspension, and
then ultrasonically cleaned and degreased. For producing oxide layers the samples were
placed in a pre-heated furnace at annealing temperatures in the range of 500°-600° C for
the desired time. The coupons were allowed to cool in air at a relative humidity of
approximately 50%. Each sample was further autoclaved at 121° C for 15 minutes at 17
psi to ensure sterility [17].
36
5.3 Algae Sample Preparation
After substrate preparation, algae adhesion was performed on the samples. For
this, samples were placed inside a fish tank with neon tetra fish. First of all, algae started
to form in the fish tank as stray algae cells, which come off of the fish or their food. With
time, algae were observed depositing on the Zr-705 samples.
5.4 Optical Microscopy
Samples with algae were analyzed using the same optical microscope and digital
camera as described previously in section 3.5. For each sample, images at different
magnifying power (2X, 10X, 50X, 100X) were captured. For one particular
magnification (50X), five images of each sample were captured, one from each of the
four corners and one from the center of the sample. Figure 17 is representative of the
images taken with the optical microscope.
5.5 Sigma Scan Pro Software
Two-point rescaling and area measurement options were selected as in section
3.5. Total areas of the images were measured and the percentage area covered by algae
for each sample was calculated. For each sample, five images were captured therefore
five different values of area were obtained and their average was calculated.
37
⎯⎯− 10 µm
⎯⎯− 10 µm
50X
50X
⎯⎯− 10 µm
⎯⎯− 10 µm
50X
50X
Figure 17. Optical images of blue-green algae on Zr-705 surfaces.
38
5.6 Analysis
The values obtained from the previous step were plotted using Microsoft Excel.
Figure 18 shows the comparison of algae growth for different oxide layer thicknesses on
Zr-705 surfaces.
5.7 Results and Discussion
Zr-705 was considered as a material for this preliminary investigation because of
previous work in our lab involving zirconium and its alloys [17,57]. The histogram plot
(Figure 18) shows that as the thickness of the oxide layer increases, initially from
unoxidized to an oxide layer thickness of 2.1 µm, the growth of algae on the surface
decreases. This is consistent with what was found in ref. [17] concerning bacterial
growth. After that, as the oxide thickness increases the growth also increases, again
consistent with ref. [17]. There is a sudden transition of algae growth between oxide layer
thicknesses of 11.1 µm and 12 µm. This transition was not identified in our bacteria
studies previously [17,57], since we did not use materials with such thick oxides.
However, this may be associated with the structural and kinetic transitions known to
occur in oxide layers on Zr surfaces.
39
Area (%)
100
90
80
70
60
50
40
30
20
10
0
0
2.1
3.1
11.1
12
Oxide Layer Thickness (micrometer)
Figure 18. Percentage area covered by blue-green algae on Zr-705 samples with different
oxide layer thicknesses on their surfaces.
40
CHAPTER VI
CONCLUSIONS
Infections caused by biofilms adversely affect human health and the associated
health care costs. Several techniques have been used to address this issue. Among these
techniques surface modification has demonstrated great potential in reducing the
detrimental effects of biofilms. The applicability of this technique will likely be enhanced
by a better understanding of the physical environment surrounding the bacteria, types of
bacteria, and also the factors that determine the rate and extent of bacterial adhesion.
In this research the adhesion of clinical strains of bacteria Staphylococcus
epidermidis and Pseudomonas aeruginosa on PVC were studied. It was demonstrated
that samples under shaken conditions had generally more bacterial adhesion than
stationary ones. Also, it was shown that S. epidermidis samples covered more percentage
area than P. aeruginosa. In most of the samples, those that were stamped had more
adhesion than unstamped ones. Thus, it is possible that stamping can be used to guide
bacteria to selected areas of bio-medical devices.
In the second part of this investigation the adhesion of blue-green algae on Zr-705
with different oxide layer thicknesses was studied. It was found that with increasing
oxide layer thickness, adhesion also increases but after an oxide layer thickness of 11.1
µm adhesion abruptly decreases. The trends observed here for algae growth are consistent
41
with our previous work with bacteria, i. e. that a thin oxide is better than no oxide at all,
but a thicker oxide can then be worse (with respect to bio-film growth) [17].
It can be concluded that the diversity of biofilms makes the task of eradication of
infection due to bio-medical devices difficult. There are still enormous challenges in this
approach of surface modification and the related systems and applications. Nevertheless,
a multidisciplinary approach and continued investigation will play a valuable role in
furthering our understanding of bacterial adhesion and minimizing the related health
hazards.
42
LITERATURE CITED
[1] J. W. Costerton, P. S. Stewart, E. P. Greenberg, Science 284, 1318 (1999).
[2] R. M. Donlan, J. W. Costerton, Clinical Microbiology Reviews 15, 167 (2002).
[3] G. Reid, H. J. Busscher, S. Sharma, M. W. Mittelman, S.T McIntyre, Surface Science
Reports 21, 251 (1995).
[4] M. Katsikogianni, I. Spilliopoulou, D.P. Dowling, Y. F. Missirlis, Journal of Material
Science: Materials in Medicine 17, 679 (2006).
[5] J. Chandra, J. D. Patel, J. Li, G. Zhou, P. K. Mukherjee, T. S. McCormick, J. M.
Anderson, and M. A. Ghannoum, Applied and Environmental Microbiology 71, 8795
(2005).
[6] M. Harmansson, Colloids and Surfaces, B, Biointerfaces 14, 105 (1999).
[7] D. Balazs, D. Favez, Y. Chevolot, N. Xanthopoulos, C. Granges, B. -O. Aronsson, F.
Sidouni, P. Descouts, H. J Mathieu, European Cells and Materials 1, 18 (2001).
[8] M. Herrero, R. Navarro, H. Reinecke, C. Mijangos, Polymer Degradation and
Stability 91, 1915 (2006).
[9] Y. Liu, Q. Zhao, Biophysical Chemistry 117, 39 (2005).
[10] S.Y. Chou, P. R. Krauss, W. Zhang, L. Guo, L. Zhuang, Journal of Vacuum Science
& Technology. B, Microelectronics and Nanometer Structures Processing, Measurement
and Phenomena 15, 2897 (1997).
[11] D. Steffensen, M. Burch, B. Nicholson, M. Drikas, P. Baker, Environmental
Toxicology 14, 183 (1999).
[12] R. Sekar, K. V. K. Nair, V. N. R. Rao, V. P. Venugopalan, Freshwater Biology 47,
1893 (2002).
[13] S. Haider, V. Naithani, P. N. Viswanathan, P. Kakkar, Chemosphere 52, 1 (2003).
43
[14] G. A. Codd, S.G. Bell, K. Kaya, C. J. Ward, K. A. Beattie, J. S. Metcalf, European
Journal of Phycology 34, 405 (1999).
[15] M. J. Lehtola, I.T. Miettinen, T.Lampola, A. Hirvonen, T. Vartiainen, P. J.
Martikainen, Water Research 39, 1962 (2005).
[16] S. Skraber, J. Schijven, C. Gantzer, A. M. de Roda Husman, Biofilms 2, 105 (2005).
[17] B.W. Buczynski, M.M. Kory, R. P. Steiner, T. A. Kittinger, R. D. Ramsier, Colloids
and Surfaces B: Biointerfaces 30, 167 (2003).
[18] S. S. Branda, A. Vik, L. Friedman, R. Kolter, Trends in Microbiology 13, 20
(2005).
[19] T. C. Mah, G. A. O’Toole, Trends in Microbiology 9, 34 (2001).
[20] P. Gilbert, D. G. Allison, A. J. McBain, Journal of Applied Microbiology
Symposium Supplement 92, 98S (2002).
[21] Y. M. Romanova, T. A. Smirnova, A. L. Andreev, T. S. Il’na, l. V. Didenko, A.L.
Gintsburg, Microbiology 75, 481 (2006).
[22] M. R. Parsek, C. Fuqua, Journal of Bacteriology 186, 4427 (2004).
[23] W. J. Costerton, M. Wilson, Biofilms 1, 1 (2004).
[24] M. R. Parsek, E. P. Greenberg, Trends in Microbiology 13, 27 (2005).
[25] M. C. M. van Loosdrecht, J. Lyklema, W. Norde,A. J. B. Zehnder, Microbiological
Reviews 54, 75 (1990).
[26] D. G Allison, B. Ruiz, C. SanJose, A. Jaspe, P. Gilbert, FEMS Microbiology Letters
167, 179 (1998).
[27] I. W. Sutherland, FEMS Microbiology Reviews 16, 323 (1995).
[28] P. Watnick, R. Kolter, Journal of Bacteriology 182, 2675 (2000).
[29] P. L. Bishop, Water Science and Technology 36, 287 (1997).
[30] E. Drenkard, Microbes and Infection 5, 1213 (2003).
[31] K. K. Jefferson, FEMS Microbiology Letters 236, 163 (2004).
[32] K. Lewis, Antimicrobial Agents and Chemotherapy 45, 999 (2001).
44
[33] M. W. Potter, S. A. Shah, K. K. Elbirt, M. P. Callery, Journal of Surgical Research
97, 54 (2001).
[34] D. Heumann, T. Roger, Clinica Chimica Acta 323, 59 (2002).
[35] C. S. Mintz, Microbes and Infection 1, 1203 (1999).
[36] C. K Stover, X. Q. Pham, A. L. Erwin, S. D. Mizogucchi, P. Warrener, M. J. Hickey,
F. S. L. Brinkman, W. O. Hufnagle, D. J. Kowalik, M. Lagrou, R. L. garber, L. Goltry, E.
Tolentino, S. Westbrock-Wadman, Y. Yuan, L. L. Brody, S. N. Coulter, K. R. Folger, A.
Kas, K. Carbig, R. Lim, D. Spencer, G. K. S. Wong, Z. Wu, I. T. Paulsen, J. Reizer, M.
H. Saier, R. E. W Hancock, S. Lory, M. V. Olson, Nature 406, 959 (2000).
[37] P. S. Stewart, International Journal of Medical Microbiology 292, 107 (2002).
[38] C. Vuong, M. Otto, Microbes and Infection 4, 481 (2002).
[39] K. Jackson, B. R. Keyser, and D. J. Wozniak, Seminars in Respiratory and Critical
Care Medicine 24, 663 (2003).
[40] S. Choi, W. Choi, H. Jung, J. Park, B. Chung, Y. Yoo, S. Koh, Journal of Applied
Polymer Science 73, 41 (1999).
[41] D. Braun, Journal of Polymer Science: Part A: Polymer Chemistry 42, 578 (2004).
[42] J. W. Costerton, Z. Lewandowski, D. E. Caldwell, D. R. Korber, H. M. LappinScott,
Annual Review of Microbiology 49, 711(1995).
[43] B. Meyer, Biodeterioration and Biodegradation 51, 249 (2003).
[44] F. Abbasi, H. Mirzadeh, A. A. Katbab, Polymer International 50, 1279 (2001).
[45] I. E. C. Mott, D. J. Stickler, W. T. Coakley, T. R. Bott, Journal of Applied
Microbiology 84, 509 (1998).
[46] S. H. Lee, J. F. Frank, Proceedings of the XXIII International Dairy Congress,
Montreal, QC, Canada, 153 (1990).
[47] B. M. Callen, M. L. Ridge, S. Lahooti, A. W. Neumann, R. N. S. Sohdi, Journal of
Vacuum Science and Technology A 13, 2023 (1995).
[48] C. A. Mills, E. Martinez, A. Errachid, G. Gomila, A. Samso, A J. Samitier,
Contributions to Science 341, 47 (2005).
[49] K. Derbyshire, Semiconductor Manufacturing 5, 18 (2004).
45
[50] V. N. Tuskett, M. P. C. Watts, Trends in Biotechnology 24, 312 (2006).
[51] L. D. Landro, C. Capone, F. Inzoli, P. E. Malacari, Journal of Vinyl and Additive
Technology 11, 111 (2005).
[52] Y. H. An, R. J. Friedman, Journal of Microbiological Methods 30, 141 (1997).
[53] T. J. Beveridge, Biotechnic and Histochemistry 76, 111 (2001).
[54] I. A. Schrijver, M. V. Meurs, M. J. Melief, C. W. Ang, D. Buljevac, R. Ravid, M. P.
Hazenberg, J. D. Laman, Brain 124, 1544 (2001).
[55] J. A. Hoffman, F. C. Kafatos, C. A. Janeway, Jr., R. A. B. Ezekowitz, Science 284,
1313 (1999).
[56] E. Fox, C. G. Ulrich, Jandel Corporation, revision 2.0, SigmaScan and SigmaScan
Pro May (1995).
[57] "Influence of Exposure Conditions on Bacterial Adhesion to Zirconium Alloys",
E.A. Yamokoski, B.W. Buczynski, N. Stojilovic, J.W. Seabolt, L.M. Bloe, R. Foster, N.
Zito, M.M. Kory, R.P. Steiner and R.D. Ramsier, J. ASTM Intern. 2(7), July/August
(2005), paper ID JAI12812. Reprinted in Titanium, Niobium, Zirconium, and Tantalum
for Medical and Surgical Applications, L.D. Zardiackas, M.J. Kraay and H.L. Freese,
eds., pp. 225-238, (ASTM Int., West Conshohocken , PA , 2006).
[58] E. P. Greenberg, Nature 406, 947 (2000).
[59] M. Watve, Resonance 2, 79 (1997)
[60] Y. Mille, L. Beney, P. Gervais, Biochemica et Biophysica Acta 1567, 41 (2002).
[61] P. D’haeseleer, Nature Biotechnology 23, 941 (2005).
[62] L. G. Harris, S. Tosatti, M. Wieland, Biomaterials 25, 4135 (2004).
[63] H. G. Craighead, S. W. Turner, R. C. Davis, C. James, A. M. Perez, P. M. St. John,
M. S. Isaacson, L. Kam, W. Shain, J. N. Turner, G. Banker, Biomedical Microdevices 1,
49 (1998).
[64] B. Guven, A. Howard, Science of the Total Environment 368, 898 (2006).
[65] G. Regelsberger, C. Jakopitsch, L. Plasser, H. Schwaiger, P. G. Furtmuller, G. A.
Peschek, M. Zamocky, C. Obinger, Plant Physiology and Biochemistry 40, 479 (2002).
46
[66] T. Kutser, L. Metsamaa, N. Strombeck, E. Vahtmae, Estuarine, Coastal, Shelf
Science 67, 303 (2006).
[67] I. R. Falconer, Environmental Toxicology 14, 5 (1999).
[68] S. J. Hoeger, B. C. Hitzfeld, D. R. Dietrich, Toxicology and Applied Pharmacology
203, 231 (2005).
[69] M. E. Callow, Biofouling 7, 313 (1993).
[70] W. Uhl, G. Schaule, R. Gimbel, IWA Second World Water Congress, Berlin, 15
(2001).
[71] T. Schwartz, S. Hoffmann U. Obst, Water Research 32, 2787 (1998).
47