Treatment of Staphylococcus Epidermidis Biofilms

Home , Eperezolid, Pefloxacin, Staphylococcus caprae

PEPTIDE-BASED SYSTEMS FOR THE TARGETED DISRUPTION AND

TREATMENT OF STAPHYLOCOCCUS EPIDERMIDIS BIOFILMS

CHRISTOPHER MICHAEL HOFMANN

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Thesis Advisor: Dr. Roger E. Marchant

Department of Biomedical Engineering

CASE WESTERN RESERVE UNIVERSITY

May, 2012

CASE WESTERN RESERVE UNIVERSITY SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Christopher Michael Hofmann

Candidate for the Doctor of Philosophy degree *.

(signed) Roger E. Marchant (chair of the committee)

James M. Anderson

Anirban Sen Gupta

Brian Cobb

(date) March 29, 2012

*We also certify that written approval has been obtained for any proprietary material contained therein.

To my wife and parents, who taught me that anything is possible.

TABLE OF CONTENTS

TABLE OF CONTENTS 1

LIST OF TABLES 4

LIST OF FIGURES 5

ACKNOWLEDGEMENTS 7

ABSTRACT 8

CHAPTER 1: INTRODUCTION 10 1.1 Clinical Incidence of Infection 10 1.2 Pathogenicity of Implanted Medical Device Infections 11 1.3 Staphylococcus epidermidis Biofilms 13 1.3.1 The Bacteria 13 1.3.2 Primary Adhesion to Surfaces 18 1.3.2.1 Nonspecific Binding 18 1.3.2.2 Bacterial-Mediated Nonspecific Adhesion 23 1.3.2.3 Bacterial-Mediated Specific Binding 24 1.3.2.3.1 Role of Bacterial Polysaccharides as Ligands 25 1.3.2.3.2 Role of Bacterial Proteins as Ligands 26 1.3.3 Biofilm Formation 29 1.3.3.1 Polysaccharide Intercellular Adhesin (PIA) 29 1.3.3.2 Accumulation Associated Protein (AAP) 33 1.3.3.3 Other Matrix Components (eDNA, Bhp, EC TA) 35 1.3.3.4 Quorum Sensing System (Agr, LuxS) 37 1.4 The Host Response 40 1.5 References 45

CHAPTER 2: BACTERIAL RESISTANCE TO ANTIBIOTICS 71 2.1 Antibiotics and Resistance to Treatment 71 2.2 Overview of Antibiotics 72 2.3 Antibiotic Resistance 80 2.4 The Biofilm as a Source of Protection 87 2.5 References 98

CHAPTER 3: STRATEGIES FOR PREVENTING AND TREATING 109 DEVICE INFECTIONS 3.1 Modifications of the Biomaterial 109 3.1.1 Surface Modifications of Materials to Prevent Adhesion 109 3.1.2 Incorporation of Antimicrobial Agents 116 3.2 Specific Targeting Strategies 127 3.1.1 Bacteriophages 127 3.1.2 Antibodies and Opsonization 130 3.3 Specific Aims and Hypothesis 133 3.4 References 135

CHAPTER 4: FIBRINOGEN-BASED LIGAND FOR SPECIFIC 143 TARGETING AND DELIVERY TO SURFACE-ADHERENT S. EPIDERMIDIS 4.1 Introduction 143 4.2 Materials and Methods 146 4.3 Results 151 4.4 Discussion 158 4.5 Conclusions 163 4.6 Future Directions 164 4.7 Acknowledgements 167 4.8 References 168

CHAPTER 5: DISRUPTION OF S. EPIDERMIDIS BIOFILM 173 FORMATION USING A TARGETED CATIONIC PEPTIDE 5.1 Introduction 173 5.2 Materials and Methods 175 5.3 Results 178 5.4 Discussion 183 5.5 Conclusions 187 5.6 Acknowledgements 188 5.7 References 189

CHAPTER 6: TARGETED DELIVERY OF VANCOMYCIN TO 197 S. EPIDERMIDIS BIOFILMS USING A FIBRINOGEN- DERIVED PEPTIDE 6.1 Introduction 197 6.2 Materials and Methods 200 6.3 Results 207 6.4 Discussion 212 6.5 Conclusions 218 6.6 Acknowledgements 218 6.7 References 219

CHAPTER 7: CONCLUSIONS AND PERSPECTIVES 227 7.1 Summary of Completed Work 227 7.2 Targeted Vancomycin: Synthesis Considerations 229 7.3 Future Directions - Flow System 246 7.4 Acknowledgements 246 7.4 References 250

CHAPTER 8: BIBLIOGRAPHY 253

LIST OF TABLES

TABLE 2.1 Clinically Administered Antibiotics 73

TABLE 2.2 Modes of Antibiotic Resistance 82

TABLE 6.1 MIC and MBC Values for vancomycin, 208 6-20-PEG3400-VAN, and 6-20-PEG5000-Van

LIST OF FIGURES

FIGURE 1.1 Schematic of biofilm formation process on 12 cardiovascular biomaterials

FIGURE 1.2 Structure of Polysaccharide Intercellular Adhesin (PIA) 32

FIGURE 2.1 Binding interaction between vancomycin and 86 Peptidoglycan Acyl-D-Ala-D-Ala

FIGURE 4.1 6-20-NG synthesis scheme 147

FIGURE 4.2 Analysis of 6-20 purity by MALDI-TOF mass 152 spectrometry and RP-HPLC

FIGURE 4.3 Blocking non-specific 6-20-NG adhesion to substrate 153

FIGURE 4.4 Normalized 6-20-NG adhesion to PET and S. epidermidis 155

FIGURE 4.5 Scanning electron microscope images of peptide 156 blocking studies

FIGURE 4.6 Peptide blocking of 6-20-NG to S. epidermidis 157

FIGURE 5.1 Optical density growth curves of surface-adherent 179 S. epidermidis

FIGURE 5.2 Effects of peptide on the composition of surface-adherent 181 biofilms as determined by quantitative fluorescence microplate readings

FIGURE 5.3 Biofilm structure after 21 hours as observed by 182 SEM and PIA staining with wheat germ agglutinin

FIGURE 5.4 Biofilm composition after 21 hours as determined by the 184 XTT metabolic assay

FIGURE 6.1 Chemical structure of vancomycin 202

FIGURE 6.2 Two-step reaction scheme for synthesis of 6-20-PEGX-VAN 203

FIGURE 6.3 Retention of 6-20-PEG3400-VAN, 6-20-PEG5000-VAN, 210 and vancomycin by S. epidermidis

FIGURE 6.4 Retention of targeted antibiotics by 24 hour 216 S. epidermidis biofilms as determined by indirect ELISA

FIGURE 7.1 MALDI-TOF of vancomycin 231

FIGURE 7.2 MALDI-TOF of SM-PEG(12)-Vancomycin synthesized 232 in aqueous conditions (PBS, pH 7.4)

FIGURE 7.3 MALDI-TOF of SM-PEG(2)-Vancomycin synthesized 234 in organic conditions (DMF w/ TFA)

FIGURE 7.4 MALDI-TOF of SM-PEG(12)-Vancomycin synthesized 235 in organic conditions (DMF w/ TFA)

FIGURE 7.5 MALDI-TOF of 6-20-PEG(2)-Vancomycin synthesized 237 in organic conditions (DMF w/ TFA)

FIGURE 7.6 pKa values of vancomycin 239

FIGURE 7.7 Charge vs. pH plot for vancomycin and PEGx-Vancomycin 240

FIGURE 7.8 Proposed synthesis scheme for soluble 242 6-20-PEGx-Vancomycin

FIGURE 7.9 Proposed synthesis scheme for on-resin modification of 244 6-20 peptide and attachment of MAL-PEGx-NHS crosslinker

FIGURE 7.10 MALDI-TOF of MAL-PEG(2)-6-20 synthesized 245 according to scheme in Figure 7.8

ACKNOWLEDGEMENTS

The Marchant lab has been a wonderful place to learn, and without its many members, past and present, I could never have finished this project. I greatly appreciate the freedom Dr. Marchant allowed me in developing my project, as it allowed me to truly take ownership of the work and its outcome. Former graduate students Sharon Sagnella, Eric Anderson, Arya Kumar, Coby Larsen, and Jeff Beamish were great sources of knowledge and insight as I started out on this journey, while current graduate students Lynn Dudash, Lin Lin, Derek

Jones, and Jenny Bastijanic have been great sources of support as I worked to wrap things up. I would especially like to thank Kyle Bednar, who worked with me during his undergraduate time at Case. Without his contributions this work would have taken far longer, and I wish him the best of luck as he now pursues his own Ph.D. at the University of Cincinnati.

Most importantly, I want to thank my wife Chrissy for her constant support and encouragement over the years. Her enthusiasm and motivation served as a constant source of inspiration, giving me the strength to keep moving forward even when things weren’t looking so bright.

Such an accomplishment would not have been possible without the help and support of countless other people over the years. To those that I wasn’t able to thank by name, please know that I will be forever grateful for everything you have done on my behalf.

Peptide-Based Systems for the Targeted Disruption and Treatment of Staphylococcus epidermidis Biofilms

Abstract

CHRISTOPHER MICHAEL HOFMANN

Complications due to nosocomial infections of implanted medical devices pose a significant health risk to patients, with Staphylococcus epidermidis often implicated in the case of blood-contacting biomaterials. One method by which S. epidermidis initially adheres to biomaterials uses the plasma protein fibrinogen as an intermediary, where the S. epidermidis surface protein SdrG binds to a short amino acid sequence near the amino terminus of the B chain of fibrinogen. This study reports on the use of this fibrinogen-derived 6-20 peptide for the targeted disruption of S. epidermidis biofilm formation using a cationic peptide, as well as the specific delivery of vancomycin to S. epidermidis biofilms.

S. epidermidis virulence relies mainly upon its ability to form a biofilm, the main component of which is polysaccharide intercellular adhesin (PIA). The synthetic 6-20 peptide was utilized to deliver a cationic polylysine peptide (G3K6) to the bacterial surface and disrupt the charge-charge interactions needed for PIA retention and biofilm stability. The effects of the 6-20-G3K6 peptide on biofilm formation were assessed using optical density, fluorescently labeled wheat germ

agglutinin, nucleic acid stain (SYTO 9), and a metabolic assay (XTT, 2,3-bis(2- methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide inner salt).

Biofilms formed in the presence of 6-20-G3K6 peptide (100 M) resulted in a

37.9% reduction in PIA content and a 17.5% reduction of adherent bacteria relative to biofilms grown in the absence of peptide. These studies demonstrate the targeting ability of the 6-20 peptide towards biomaterial-adherent S. epidermidis, and highlight the potential for disrupting the early stages of biofilm formation.

Targeted 6-20-PEGx-VAN vancomycin derivatives were then synthesized using a flexible, variable length poly(ethylene glycol) linker between the peptide and antibiotic. Initial binding to surface adherent S. epidermidis was increased in a concentration-dependent manner relative to vancomycin for all equivalent concentrations ≥4 g/ml, with targeted vancomycin content up to 22.9 times that of vancomycin alone. Retention of the targeted antibiotics was measured after an additional 24 hour incubation period, revealing levels 1.3 times that of vancomycin. The results demonstrate the improved targeting and retention of vancomycin within a biofilm due to the incorporation of a specific targeting motif.

CHAPTER 1

INTRODUCTION

1.1 CLINICAL INCIDENCE OF INFECTION

There are close to 40 million surgical procedures involving the insertion of

artificial devices performed each year in the United States[1]. Of these implanted

devices, close to 1 million will become infected, accounting for nearly 45% of the total nosocomial infections reported annually[2, 3]. Approximately 835,700 cardiovascular devices are implanted each year, including mechanical heart valves, vascular grafts, pacemaker-defibrillators, and ventricular assist devices[1,

2], of which 31,680 devices are projected to become infected[2]. While the overall incidence of infection is relatively low at just under 4%, the associated morbidity/mortality rate can exceed >25%. In addition to the health risks, infected medical devices have a significant financial impact on patients. The average cost of medical and surgical treatments for infected cardiovascular devices ranges from $35,000 to $50,000, with an estimated annual cost of $1.2 billion[2]. The total annual cost of treatment for all nosocomial infections (both device and non-device related) is estimated to be $11 billion[3].

1.2 PATHOGENICITY OF IMPLANTED MEDICAL DEVICE INFECTIONS

Staphylococci, enterococci, enterobacteriaceae, and Candida spp. are common

pathogens associated with infections of indwelling medical devices[4], with the likelihood of infection, as well as the organism implicated in the infection, greatly

dependent upon the type and location of the implant itself[4]. In the case of

intravascular implants, coagulase-negative staphylococci (CoNS), particularly

Staphylococcus epidermidis, are the most common cause of infection[1, 5].

Staphylococcus epidermidis is a naturally occurring commensal organism found

on the surface of the human body, making up the majority of the bacterial

microflora[6]. The only infection known to be caused by the typically

noninvasive S. epidermidis in the immunocompetent adult is native valve

endocarditis; all other infections require the presence of a foreign body[7]. It is

likely that introduction of these microorganisms to an implanted device occurs at

the time of surgery, as it has been found that 5,000 to 50,000 skin particles are

transferred daily from physicians skin flora in intensive care units, and even

under aseptic conditions 90% of clean wounds contain pathogenic bacteria.[3]

S. epidermidis infections rely upon the ability of the bacteria to adhere to a

surface and successfully form a biofilm, as opposed to S. aureus infections which

are often associated with toxins and other virulence factors[5, 8]. A two-stage

•Mature biofilm •Leukocytes release oxidative/nitrogen and intermediates contentsgranular •Bacterial •Bacterial proliferation, aggregation, andslime formation iovascular biomaterials. Bacteria first adhere e and form an extracellular polysaccharide matrix ng which time leukocytes are able to phagocytose the Time •Specific bacterial to thrombus adhesion formations via protein intermediaries Biomaterial

•Platelet activation and adhesion •Leukocytes phagocytosis of bacteria S. epidermidis Leukocyte Activated Platelet Plasma Protein Plasma : Schematic of biofilm formation process on card •Plasma proteins proteins adsorb•Plasma onto biomaterial •Non-specific bacterial adhesion •Leukocyte chemotaxis and migration Figure 1.1 to surface-adsorbed proteins and/or platelets, duri bacteria. Following adhesion, the bacteria proliferat that protects them from the host defenses.

process can be used to describe the course of such foreign-body infections

(Figure 1.1), wherein initial adhesion of the bacteria to the material occurs first, followed by proliferation, matrix secretion, and cell-cell adhesion leading to the formation of a mature, multi-layered biofilm[5]. While planktonic S. epidermidis is known to be susceptible to a large number of antibiotics[9], the biofilm environment offers the encapsulated bacteria increased resistance, often times able to survive antibiotic concentrations several orders of magnitude higher than the minimum inhibitory concentration (MIC) measured in planktonic suspensions[10-12].

1.3 STAPHYLOCOCCUS EPIDERMIDIS BIOFILMS

1.3.1 The Bacteria

Staphylococcus epidermidis is a gram positive bacterium characterized by a low

G-C content and typically aggregating in grape-like clusters[8, 13]. As one of several gram positive, low G-C bacteria that share the cocci shape, further description is necessary in order to uniquely characterize S. epidermidis.

Staphylococcus test positive for the enzyme catalase, and as such can be easily distinguished from the similarly shaped and catalase-negative Streptococcus[13].

In order to distinguish Staphylococcus form Micrococcus, the oxidation- fermentation test is used. Micrococcus (an obligate aerobe) can only produce acid from glucose aerobically, while Staphylococcus (a falcultative aerobe), can produce acid from glucose both aerobically and anaerobically[13]. Finally, the

genus Staphylococcus is divided into two main groups based on expression of the

clotting enzyme coagulase. Staphylococcus epidermidis is non-pigmented and

coagulase-negative, whereas Staphylococcus aureus is yellow-pigmented and

coagulase-positive[8, 14].

As a gram positive bacterium, S. epidermidis is characterized by a thick cell

wall surrounding the cell’s cytoplasmic membrane. The cytoplasmic membrane

is an 8 nm thick phospholipid bilayer stabilized by hopanoids (similar to sterols

found in eukaryotic cells)[13]. The membrane acts as a diffusion barrier, and

along with active transport systems located within the membrane, allows the

bacteria to concentrate large amounts of dissolved solutes within the cell. The

resulting turgor pressure is substantial, equal to about 2 atmospheres in

Escherichia coli[13]. In order to counter this pressure, a thick peptidoglycan cell wall is constructed that provides the strength necessary to maintain membrane integrity. Inhibition of peptidoglycan synthesis or degradation of an existing cell wall will result in cell lysis[15].

Peptidoglycan is a linear polymer made up of two alternating sugar residues,

N-acetylglucosamine (GlcNAc) and N-acetylmuramic acid (MurNAc), linked

through β-1→4 bonds[15]. The D-Lactoyl group of each MurNAc residue is

substituted with a tetrapeptide sequence of L-Alanine−γ-D-Glutamic acid− L-

Lysine (or in some cases 2,6 Diaminopimelic Acid)−D-Alanine, although the

exact sequence is known to have some variations from strain to strain[15, 16].

The D-L-D-L-D sequence prevents the formation of α-helical secondary structure and gives the tetrapeptide more flexibility[17]. The peptidoglycan strands, with an average length of 18 disaccharide units in S. aureus, form sheets around the

cytoplasmic membrane which are subsequently crosslinked by an interpeptide

bridge[15]. The interpeptide bridge forms a crosslink between the amino group

of the L-lysine (or 2,6 diaminopimelic acid) and the carboxyl group of the

terminal D-alanine residue on a neighboring tetrapeptide[15]. While most gram negative bacteria form crosslinks using a direct amide bond between the two residues, gram positive bacteria utilize a peptide bridge consisting of one to

seven amino acids of varying composition[13, 15]. This peptide interbridge is the

greatest source of diversity in the bacterial peptidoglycan structure. S. aureus, for

example, typically utilizes a five-reside glycine linker[13]. S. epidermidis, on the

other hand, shows more variation. While 20% of S. epidermidis strain Texas 26

possessed penta-glycine bridges, 55% were found to have Gly-Gly-Ser-Gly-Gly,

15% were Ser-Gly-Ser-Gly-Gly, and 10% were Gly-Ser-Gly-Gly-Gly [18].

Meanwhile, S. epidermidis strain 66 contains an alanine residue in place of one of

the glycines of the penta-glycine bridge[17].

The peptidoglycan structure serves as an anchor for both polysaccharides and cell wall proteins. The principal polysaccharide attached to the bacterial surface of gram positive bacteria is teichoic acid (TA), which can either be anchored to the cell wall (cell wall teichoic acid, CW TA) or anchored to the cytoplasmic

membrane by way of a lipid-spanning anchor (lipoteichoic acid, LTA)[19].

Teichoic acid is generally described as a (1,3)-phosphodiester-linked polyglycerol

phosphate (S. epidermidis) or polyribitol phosphate (S. aureus) with an overall

negative charge[20, 21]. The 2-position of the glycerol residues in S. epidermidis

can be substituted with α-glucose (α-Glc), α-N-acetylglucosamine (α-GlcNAc), D- alanine, or α-6-alanyl glucose (α-Glc6Ala)[22]. D-alanine is of particular importance, as each D-ala substitution imparts a positive charge into the TA molecule, countering some (but not all) of the negative charge provided by the phosphate backbone. An S. aureus mutant lacking the ability to incorporate D-ala onto the TA molecule lost its ability to colonize glass and plastic surfaces, presumably due to an increased negative surface charge of the bacteria that resulted in unfavorable electrostatic interactions with the surfaces[23]. It is also speculated that the net negative charge of TA is utilized to anchor non-covalently bound extracellular molecules such as autolysin E (AtlE) and polysaccharide intercellular adhesin (PIA).

Bacterial surface proteins are also anchored to the cell wall, with eleven genes encoding putative surface-anchored proteins already identified[24] These cell- wall-anchored (CWA) proteins often times play key roles in bacterial adherence and evasion from the host immune system[25]. S. aureus Protein A is a prototypical surface protein, and most CWA proteins of gram positive bacteria can be described by the same general structure[26]. At the N-terminus, there is

an ~40 amino acid secretion signal responsible for mediating translocation across the cell’s cytoplasmic membrane[24]. Bordering the secretion signal there is an A- region containing the ligand-binding site, a B-repeat region containing tandemly repeated sequences, and a proline-rich wall-spanning region.[27] Finally, there is a sorting signal region required for properly locating the protein on the bacterial surface. This sorting signal consists of a wall-anchoring LPXTG motif, a hydrophobic transmembrane segment, and a positively charged cytoplasmic tail[26]. While the LPXTG motif is highly conserved amongst gram positive bacteria, the hydrophobic domain and the positively charged tail are variable in sequence and length[26].

Attachment of CWA proteins to the cell wall is mediated by transpeptidases known as sortases. The S. aureus transpeptidase Sortase A (SrtA) is considered the archetypical sortase for gram positive bacteria, although five distinct subfamilies of sortases have been identified, with most bacteria encoding two or more distinct sortases[28]. S. epidermidis RP62A encodes just one sortase, SrtA[24,

28]. After a protein is exported into the extracellular space, it is retained within the cell membrane at its C-terminal by the sorting signal’s positively charged tail.

Following retention, SrtA proteolytically cleaves the protein between the threonine and glycine of the LPXTG motif, and the carboxyl group of the threonine is amide-linked to a free amino group of the peptidoglycan crossbridge[25]. Since a mature cell wall is highly crosslinked and therefore

contains very few free crossbridge amines, it is likely that protein anchoring

takes place prior to completion of the peptidoglycan crosslinking process[25].

1.3.2 Primary Adhesion to Surfaces

Prior to colonization, bacteria present in the bulk fluid environment are

transported towards the surface by Brownian motion, sedimentation, and

convective mass transport[29]. For static systems, sedimentation is the most important factor, and Brownian motion and convection can often times be neglected. Meanwhile, for flow systems, convective mass transport and diffusion

become much more significant[30]. Once S. epidermidis is brought in to close

proximity with the indwelling biomaterial, both nonspecific interactions and

specific binding events work to anchor the bacteria to the surface.

1.3.2.1 Nonspecific Binding

Bacterial adhesion to abiotic surfaces was first proposed by Marshall et al. to involve an initial reversible sorption step, followed by a surface-dependent irreversible sorption process[31]. As such, it was suggested that the adhesion of a bacterium to a surface could be explained by the DLVO colloidal stability theory

(after Derjaguin, Landau, Verwey, and Overbeek)[31]. The DLVO theory has since become a commonly accepted approach to modeling bacterial adhesion[32,

33].

The classical DLVO theory describes the total interaction energy (WDLVO) between a sphere (bacteria) and planar surface (biomaterial surface) in aqueous media as a balance between the attractive van der Waals energy (WvdW) and repulsive electrostatic double layer energy (WEl)[32, 33]. The van der Waals and

electrostatic double layer energies are additive and can be represented as a

function of separation distance (d) as follows:

(1.1)

Adhesion between the bacteria and biomaterial is favored when the overall

energy WDLVO is negative, whereas repulsion will occur if WDLVO is positive.

Hence, the balance between the negative WvdW and positive WEl energies will

determine the adhesion between a bacterial cell and an abiotic surface.

Despite the widely accepted use of the DLVO theory to describe bacterial

adhesion, many studies have shown deviations due to non-DLVO energies[33].

In particular, the DLVO theory assumes both the bacterium and biomaterial to be

chemically inert. This assumption is invalid, as hydrogen and chemical bonding

play an important role in adhesion when polar solvents (such as water) are involved[32]. Hydrophobic interactions, which are polar (but not electrostatic) in

nature may be up to two orders of magnitude larger than either WvdW or WEl encountered in the classical DLVO approach[34]. In addition, it is estimated that

~90% of all noncovalent interactions in aqueous media involve hydrophobic

interactions, which rely on electron acceptor-electron donor interactions[35]. As

such, these hydrophobic interactions have been termed Lewis acid-base (AB)

interactions[34]. The Dupré thermodynamic model of adhesion takes in to

account the AB interactions, defining the bacterial adhesion energy as:

Δ Υ Υ Υ (1.2)

Where BS, BL, and SL are the interfacial tensions of the bacteria-substrate,

bacteria-liquid, and substrate-liquid interfaces, respectively[33]. Interfactial tensions in equation 1.2 can be calculated using Young’s equation by way of contact angles[36]. Similar to the DLVO model, bacterial adhesion is favored if the total energy ∆Gadh < 0.

As was the case with the DLVO model, a number of reports indicate that the

thermodynamic model is not accurate under various experimental

conditions[33]. As such, Van Oss suggested an extension of the classical DLVO

theory which incorporates the hydrophobic ∆Gadh surface energy component

developed under the thermodynamic theory[34]. This is represented by the

extended DLVO (xDLVO) equation:

(1.3)

Where Wadh is the adhesion energy due to the hydrophobic Lewis acid-base (AB)

interactions, defined in equation 1.2. In aqueous environments, these AB

interactions are never negligible[37].

In order to obtain an analytical solution for WvdW, WEl, and Wadh, approximations are made according to Derjaguin wherein interaction energies between a sphere and planar surface are calculated as the surface integral of plate-plate free energies along the separation distance, d [32, 38]. The resulting equations for van der Waals, electrostatic double layer, and lewis acid-base energies are:

∙ (1.4)

2 ln 1 (1.5)

2Δ (1.6)

Where A is the Hamaker constant, r is the sphere radius, εr is the relative

permittivity of water (=79), ε0 is the dielectric permittivity in a vacuum

(=8.854x10-12CV-1m-1), κ is the inverse Debye length (=3.28x109*I1/2m-1, where I is

the ionic strength in mol/L), ψb and ψs are the surface (zeta) potentials of the sphere and planar surface, λ is the decay length of acid-base interactions in water

(=0.6), d0 is the minimum separation distance (=0.157), and is the acid-base

component of free energy of interaction at contact (where d=d0)[32, 38, 39].

Adhesion between a bacterium and substrate occurs at the primary energy

minimum predicted by equation 1.3, typically at a separation distance < 1 nm. In order to reach this minimum, there is a repulsive energy barrier that must be overcome. Once this barrier is overcome and the minimum is reached, adhesion

is considered to be irreversible. If the energy barrier is not overcome, there is

usually a very shallow secondary energy minimum wherein the bacteria may

reversibly adhere to the surface. The characteristic separation distance for this

secondary minimum is ~8.5 nm[37].

Ionic concentration, I, plays a critical role in DLVO adhesion, as the inverse

Debye length utilized in equation 1.5 is dependent upon I. As a result, κ can take

on value from < 0.2 to > 20 nm when ionic concentrations are > 500 mM and < 1

mM, respectively[33]. At high ionic strengths, the electrostatic double layer is

very small, which in turn minimizes the contribution of WEl to the total energy

WDLVO. Consequently, the overall WDLVO may be attractive, despite both a

negatively charged bacterium and substrate. Another interesting consequence of

the dependence of κ on ionic strength is manifested at intermediate values of I, wherein the secondary energy minimum is created.[33]

A further extension of this theory can be used to predict the adhesion of S. epidermidis to various biomaterial surfaces[36]. With liquid surface tensions less than bacterial surface tensions, ∆Gadh decreases with decreasing biomaterial

surface tension. However, if the liquid surface tension is greater than the

bacterial surface tension, ∆Gadh increases with decreasing biomaterial surface

tension. Taken together, the thermodynamic model indicates that S. epidermidis

adhesion to biomaterial surfaces will increase as the material hydrophobicity

increases[36].

1.3.2.2 Bacterial Mediated Nonspecific Adhesion

Despite the ability of the xDLVO theory to reasonably model nonspecific

bacterial adhesion to abiotic surfaces, deviations from the theory are fairly

common as the xDLVO theory assumes the bacteria to be a smooth, uniform

sphere. In reality, S. epidermidis has a number of proteins and carbohydrates on

its surface that can influence both nonspecific and specific adhesion. Heilmann et

al utilized transposon mutants of S. epidermidis O-47 to identify a surface protein

responsible for adhesion to bare polystyrene[40, 41]. This protein shares a 61% similarity with the amino acid sequence of the major autolysin (Atl) from S.

aureus, and was therefore name Autolysin E (AtlE)[41]. Absence of AtlE from the

S. epidermidis surface resulted in the reduction of bacterial hydrophobicity, a decrease in adhesion to hydrophobic polystyrene surfaces, and a significant

increase in binding to hydrophilic glass surfaces[40]. Furthermore, the absence of

AtlE in an S. epidermidis O-47 mutant strain showed significantly less virulence in

a rat central venous catheter infection model when compared to the wild type

strain[42].

Extracellular DNA (eDNA) also plays an important role in nonspecific

adhesion to indwelling devices. Treatment of S. epidermidis with DNase I

prevented bacterial adhesion to polystyrene and glass surfaces, while treatment

with RNase or heat-activated DNase I had no effect on adhesion[43, 44]. PCR

analysis revealed that this eDNA was likely genomic DNA originating from

bacterial lysis, likely mediated by the autolytic activity of AtlE[43]. Inactivation

of the atlE gene reduced DNA release by more than 90%[43]. Similar results were found with S. aureus and P. aeruginosa, where DNase I and DNaseIL2 were able to suppress biofilm formation on plastic surfaces[45, 46]. Given the findings on the importance of eDNA in initial adhesion to plastic and glass surfaces, the possibility exists that AtlE may not play a direct role in initial adhesion to polystyrene as previously thought. Instead, the role of AtlE may be to mediate cell lysis in order to release genomic DNA for the purpose of adhesion[47].

1.3.2.3 Specific Binding

Plasma proteins are intrinsically surface active, and as such they rapidly

adhere to blood contacting biomaterials through electrostatic and hydrophobic

interactions[48]. Initial findings by Vroman demonstrated that fibrinogen adsorbed onto surfaces from blood plasma within seconds, only to be replaced at a subsequent time point by factor XII[49]. These findings became known as the

Vroman effect and indicated that protein adsorption was a dynamic process

involving time-, concentration-, and spatially-dependent absorption maxima[50].

Subsequent studies extended the principles of the Vroman effect to many other plasma proteins, including albumin, IgG and high molecular weight kininogen

(HMWK)[51]. Surface plasmon resonance (SPR) studies have shown that albumin absorbs quickly onto a polystyrene interface, soon to be displaced by

IgG and fibrinogen. Fibrinogen then displaces the IgG, with fibrinogen finally

removed by larger more surface active proteins[52]. As a general rule, smaller

proteins with high solution concentrations will adsorb first, after which they will

be displaced by larger, more surface active proteins[50, 52]. In addition, there is the possibility that an adsorbed protein will change its conformation so as to increase its surface activity to the point that it becomes irreversibly bound to the surface[50].

The consequences of the Vroman effect are quite significant in regards to infection of indwelling devices, as blood-contacting surfaces will display a multitude of potential binding sites for initial bacterial adhesion. To demonstrate this point, studies utilizing a canine arteriovenous shunt model have revealed that fibrinogen and fibronectin sufficiently coat polyethylene and polyvinyl chloride tubing after 5 minutes of exposure to canine blood to promote specific binding of S. aureus [53]. In fact, host proteins adsorb so rapidly to coat implanted devices that specific adhesion is thought to be the primary means by which S. epidermidis begins to colonize a surface[5, 20]

1.3.2.3.1 Role of Bacterial Polysaccharides as Ligands

Teichoic acid (TA), a component found on the surface of many gram positive

bacteria, is known to bind several host proteins. Purified TA was used to

significantly enhance adhesion to fibronectin coated surfaces in three S.

epidermidis strains (KH11, RP62A, and AB9)[54]. This adhesion was dose- dependent, and demonstrated that TA acts as a bridge between the staphylococcal cell surface and the surface-adherent fibronectin.[54] Meanwhile, the lipid component of lipotechoic acid (LTA) that is typically exposed to the extracellular environment in most gram-positive strains was shown to mediate adhesion to fibrin-platelet clots in vitro[55].

1.3.2.3.2 Role of Bacterial Proteins as Ligands

Staphylococcus epidermidis has the ability to adhere specifically to a variety of

host extracellular matrix proteins as well as plasma proteins that adsorb readily

onto implanted surfaces[56]. This adhesion is mediated by bacterial surface

proteins that are part of a family of adhesins termed microbial surface components recognizing adhesive matrix molecules (MSCRAMMs)[57, 58],

several of which have been identified in S. epidermidis[24].

S. epidermidis Extracellular matrix binding protein (Embp) was identified by

shotgun phage display cloning as the major protein responsible for binding to

fibronectin[59]. In the absence of the 1 MDa Embp protein, S. epidermidis does not adhere to fibronectin coated surfaces[60]. RT-PCR analysis revealed that Embp is expressed throughout all growth phases of S. epidermidis, and scanning force

microscopy studies revealed that Embp binds fibronectin at a single site on the

type III12-14 domains near the C-terminus[60, 61]. Heparin is known to bind

fibronectin at this same location, and in doing so can competitively inhibit the

Embp binding event[59, 61]. The binding interaction between Embp and

fibronectin is distinct from that of the S. aureus fibronectin binding proteins,

FnBPA and FnBPB, which bind to fibronectin type I domains at its N-

terminus[59, 60].

Two autlysins associated with the S. epidermidis surface, AtlE and Aae, are

known to bind to vitronectin[41, 62, 63]. AtlE has specific ligand-receptor

interactions with surface adherent vitronectin in addition to its ability to adhere

to bare polystyrene surfaces through hydrophobic interactions[41]. Aae was

demonstrated to bind to vitronectin with a Kd of 149 nM[63]. Aae also binds to

fibrinogen and fibronectin, but does not bind to polystyrene[63]. The specific

binding sites on vitronectin for AtlE and Aae are unknown.

The S. epidermidis lipase GehD was demonstrated to bind to type I, type II,

and type IV collagen[64]. Thus, in a manner similar to AtlE, GehD is a

bifunctional molecule, acting as both a lipase and an adhesive ligand.

Interestingly, the structure of GehD may be very similar to that of the

mammalian integrin α1 collagen-binding I-domain[64].

S. epidermidis is also known to specifically bind both soluble and surface-

adherent thrombospondin, although the exact mechanism is unknown. It is, however, believed that the bacterial component responsible for this interaction is proteinaceous in nature[65].

SdrG, also referred to as Fbe, is the most studied S. epidermidis MSCRAMM

and it binds fibrinogen[66-70] by way of a 331 amino acid fibrinogen binding

region (aa269-599) located in the SdrG A-Domain[24, 69, 71]. SdrG binds to the

-7 amino-terminus of the fibrinogen B chain [66, 72] with a KD of 0.9x10 M [66].

A synthetic peptide sequence representing the first 25 amino acid residues of the

B-chain of fibrinogen (1-25) has been used to mimic the SdrG binding region,

-7 and this peptide bound to fibrinogen with a KD of 1.4x10 M [66]. This indicates

that the significant amino acids necessary for the SdrG-Fibrinogen binding

interaction are present in the 1-25 peptide, and that this linear sequence is in a

nearly optimal conformational state[66]. The fibrinogen binding domain was

further narrowed down to the 6-20 peptide, consisting of the amino acid

sequence NEEGFFSARGHRPLD[66], with amino acids 10-15 contributing most

significantly to the SdrG-fibrinogen binding scheme. 6-9 and 19-20 at either

end of the peptide are not involved in significant physical interactions with the

bacterial surface proteins and are in fact extended into the solvent

environment[73]. Exchanging any single amino acid residue in this 10-15

binding region for an alanine residue (or serine in the case of Ala13) significantly

reduces the binding affinity of the peptide[73].

The SdrG-fibrinogen interaction has been shown to play a significant role in

the initial stages of bacterial adhesion. Recombinant SdrG is able to block the

adherence of S. epidermidis to fibrinogen in a concentration dependent

manner[67]. Adding SdrG under saturating conditions prevents S. epidermidis from binding to fibrinogen[72], and mutating the fibrinogen-binding region of the SdrG A-domain of S. epidermidis greatly impairs the ability to bind to immobilized fibrinogen.[73, 74]. Furthermore, both the gene encoding for SdrG

[24] and the SdrG protein itself [68] are present in 100% of clinical S. epidermidis isolates tested, and studies of infected and recovering patients reveal anti-SdrG antibodies in the serum, confirming that the bacteria express SdrG during the

infectious stages[68, 75]. An in-vivo intravascular catheter infection model

confirmed the importance of SdrG: an SdrG-negative S. epidermidis mutant had a

20% infection rate, compared to a 100% rate for the SdrG-positive strain[76].

1.3.3 Biofilm Formation

After initial adhesion to a substrate, S. epidermidis begins the process of

biofilm formation through proliferation, extracellular matrix production, and

intercellular adhesion. Both polysaccharides and proteins are involved, and

research is ongoing into the exact mechanisms of biofilm formation.

1.3.3.1 Polysaccharide Intercellular Adhesin (PIA)

A crucial step in the formation of a mature biofilm is the production of

exopolysaccharides[77], with various S. epidermidis exopolysaccharides described

in the literature. A capsular polysaccharide adhesin (PS/A) was first described in

1988 by Tojo et al[78], while Mack et al (1996) reported the structure of a -1-6- linked N-acetylglucosamine they termed polysaccharide intercellular adhesin

(PIA)[79]. McKenney et al subsequently identified the structure of PS/A as a -1-

6-linked N-succinylglucosamine (PNSG)[80]. Further studies by Maira-Litran et

al revealed that PS/A did not contain N-succinylglucosamine, but was instead -

1-6 N-acetylglucosamine (PNAG)[81]. A second variation of PNAG was reported

by Baldassarri et al as the S. epidermidis slime associated antigen (SAA)

containing 70% N-acetylglucosamine[82]. Despite the often times conflicting

reports on the structure of these various exopolysaccharides, it is now widely

accepted that PS/A, PIA, PNAG, and SAA are the same chemical entity, known

collectively as PIA[83-87].

The most accurate description of the structure of PIA (Figure 1.2) is provided

by Mack et al as a linear homopolymer of at least 130 -1,6-linked

N-acetylglucosamine (GlcNAc) residues with a molecular weight of

Mr=30,000[79]. However, the analytical technique used to make the

measurements appears to be highly influential on the results obtained, as other research groups have reported molecular weights of up to 460 kDa[85]. PIA consists of a major polysaccharide I (>80%) and a minor polysaccharide II

(<20%)[79]. Approximately 80-85% of the glucosamine residues in polysaccharide I are acetylated, while the remaining residues are deacetylated to give an overall positive charge to the molecule. The distribution of the

deacetylated residues was found to be more or less random. Meanwhile, polysaccharide II has significantly less deacetylated residues, instead containing

O-linked succinate groups that result in a slight anionic charge[79]. It was originally thought that polysaccharide II also contained O-linked phosphate groups, but it was later shown that the presence of phosphates was an experimental artifact due to the use of phosphate buffer[88].

PIA is a product of enzymes encoded by the intercellular adhesion (ica) icaADBC operon[89, 90]. IcaA contains the N-acetylglucosaminyltransferase activity needed for PIA synthesis, but activity levels are low when expressed on its own[89, 90]. When IcaA is co-expressed with the IcaD protein, enzymatic activity is increased 20-fold and oligomers can reach a maximum length of 20 residues. The exact function of IcaD is unknown[89]. IcaC is necessary for the assembly of full length polymer chains (~130 residues) and is likely responsible for the subsequent transport of the assembled PIA through the cytoplasmic membrane[83, 89]. IcaA, IcaD, and IcaC are transmembrane proteins and presumed to be co-expressed as an assembled complex in the membrane through ionic, rather than covalent, interactions [90, 91]. Typically, the PIA associates with the bacterial surface in the form of fibrous strands[92]. Expression of the icaADBC gene products is partly regulated by icaR, located upstream of icaADBC, whose gene product is a transcriptional repressor controlled by environmental conditions[93].

O O O O O HO HO O O HO NH HO O R O O NH3 O x R y NH

O O z

O R = H or

Figure 1.2: Structure of Polysaccharide Intercellular Adhesin (PIA). The x and y subunits are found primarily in Polysaccharide I, with approximately 15-20% of the residues taking on the deacetylated y structure. Polysaccharide II is composed mainly of the x and z subunits.

Homologous icaADBC operons are found in S. aureus[80, 94], S. caprae[95], S.

lugdunensis[96], and S. haemolyticus[97]. In addition, genes orthologous to

icaADBC have been found in various gram-negative organisms such as E. coli,

Yersina pestis, Aggregatibacter actinomycetemcomitans, Bordetella sp., and

Actinobacillus pleuropneumoniae[85, 98]. Due to the presence of icaADBC

homologues and orthologues in such a diverse group of bacteria, it follows that

PIA is a common feature of bacterial biofilm formation.

Multiple functions are attributed to PIA, including protection from host

polymorphonuclear leucocytes and antimicrobial peptides[91, 92], virulence in

several in vivo models[42, 99-101], hemagglutination[102], and intercellular

adhesion[79, 90, 103, 104]. However, with the creation of an icaB negative mutant,

the non-deacetylated PIA was unable to associate with the bacterial surface and

was instead released into the culture media[91]. The resulting inability to retain

PIA at the bacterial surface led to the elimination of biofilm formation on plastic

surfaces, reduced bacterial adhesion to human epithelial cells, and impaired

resistance to neutrophil phagocytosis and antimicrobial peptides in vitro, while

reducing the persistence of infection in an in vivo murine model[91].

1.3.3.2 Accumulation Associated Protein (AAP)

Recently, S. epidermidis biofilms have been observed that form in the absence

PIA. In a study of 66 coagulase-negative staphylococci (CoNS) strains, 15% were

found to form biofilms without PIA production[105], while a study of 44 CoNS

prosthetic joint infection isolates revealed that 54% were icaA-nagative[106].

Additional studies have confirmed the ability of PIA-negative clinical strains of

S. epidermidis to form biofilms utilizing proteinacious factors. Such biofilms were

readily disrupted by proteinase K, rather than the PIA-degrading

metaperiodate[107].

Hussain et al identified an extracellular protein, referred to as accumulation associated protein (AAP), which played an essential role in S. epidermidis biofilm

formation[108]. Of the 58 CoNS strains tested, 55% were AAP-positive and

formed larger biofilms than the AAP-negative strains[108]. AAP (sometimes

referred to as SesF) was later shown to mediate biofilm formation in a PIA-

independent manner, and expression of AAP could be exogenously

induced[109]. The AAP protein consists of an N-terminal Domain A, a repetitive

Domain B, and a C-terminal LPXTG anchoring domain common to S. epidermidis

surface anchored proteins[109]. The repetitive Domain B is also known as the G5

domain due to the presence of conserved glycine residues[110], with a purported

ability to bind N-acetylglucosamine in a Zn2+ dependent fashion[110, 111].

Structural analysis of the AAP Domain B predicts it to fold as a lectin, further

increasing the possibility that AAP may mediate intercellular adhesion by

associating with PIA[24]. The 280-kDa AAP forms tufts of hydrophobic fibrils on

the S. epidermidis surface, but is unable to mediate biofilm formation. However,

upon proteolytic cleavage of Domain A (either through staphylococcal or host

proteases), the mature 140-kDa surface protein, with exposed Domain B,

becomes biofilm-positive[112]. S. epidermidis RP62A expresses the 140-kDa

version of AAP. Monoclonal antibodies directed against the S. epidermidis RP62A

AAP were able to reduce biofilm formation by up to 87%[113] , while biofilm formation was significantly inhibited when grown in the presence of a Zn2+ chelator[111]. However, in contrast to the reports implicating AAP as the sole mediator of biofilm formation, reports have shown that biofilm formation is in fact dependent on PIA[114]. Preformed S. epidermidis RP62A biofilms were disrupted by dispersin B (an enzyme with PIA-hydrolyzing activity), while the proteolytic enzymes proteinase K and trypsin were unsuccessful at dispersing the RP62A biofilms[115]. It is now apparent that both PIA and AAP play important roles in S. epidermidis biofilm formation, with environmental conditions and strain variations determining the relative contribution of each. It is highly likely that the two systems are designed to be redundant and complimentary to each other[116].

1.3.3.3 Other Matrix Components (eDNA, Bhp, EC TA)

Extracellular DNA (eDNA) has been shown to be present in the matrix of S.

epidermidis and S. aureus biofilms[46]. However, the role of eDNA differs between

the two species. With S. epidermidis, eDNA plays a very minor role in the final

biofilm architecture, demonstrated by the inability of DNase I to disrupt or

disperse mature biofilms[43, 46]. The S. aureus biofilm structure, on the other

hand, is significantly disrupted by treatment with DNase I, indicating an

important intercellular adhesive role of eDNA[47].

Biofilm associated protein (Bap) was been identified in a bovine strain of S.

aureus, implicated in the process of biofilm formation[117]. Bap is a surface

anchored protein containing the LPXTG motif, and Bap expression leads to

strong biofilm formation[117]. While absent from all human strains of S. aureus, a

similar protein, termed Bap homologue protein (BhP) is present in S.

epidermidis[118]. It is likely that Bhp plays a role in intercellular adhesion, although there has been no study into the mechanism or significance.

Extracellular teichoic acid (EC TA) was discovered to be present in significant

quantities in the S. epidermidis matrix[119, 120]. The structure of EC TA is very similar to that of cell wall teichoic acid (CW TA), and the growth conditions

(aerobic vs. anaerobic, shaking vs. non-shaking, etc) dictated the relative amounts of EC TA and PIA found in the biofilm’s extracellular matrix[88]. As

PIA is cationic and hydrophobic, while EC TA is anionic and hydrophilic, it seems likely that S. epidermidis is able to adapt and regulate the charge and hydrophobicity of its biofilm in response to surface and environmental

challenges.

1.3.3.4 Quorum Sensing System (Agr, LuxS)

As biofilm production is considered to be the main virulent characteristic of

S. epidermidis, an effective means of controlling biofilm production is needed.

Quorum sensing is a mechanism by which a bacterial population can regulate

gene expression in response to cell density. In a general sense, bacterial cells

constantly release a low level of pheromones, also known as autoinducers. As the bacterial population increases, the local concentration of pheromones increases, and once above a certain threshold, a signal transduction pathway is activated

that leads to gene suppression and/or activation. Gram positive bacteria utilize

post-translationally modified peptides as their pheromones, and these peptides

are detected by a membrane bound receptor protein that works in tandem with a

cytoplasmic protein that binds to DNA[121]. This is in contrast to Gram negative bacteria, whose pheromone is typically an N-acyl-homoserine lactone (AHL) that diffuses directly into the bacteria and binds to a specific transcriptional activator protein[121, 122] Two quorum sensing systems are present in S. epidermidis, Agr and LuxS, with the ability to regulate initial adhesion, aggregation, and dispersal of the bacteria.

The accessory gene regulator (agr) was originally thought to be the only quorum sensing system present in Staphylococcus epidermidis[123]. The agr gene locus contains two divergent transcription units, driven by promoters P2 and

P3[124]. The P2 operon encodes four gene products, AgrA, AgrC, AgrD, and

AgrB, while the P3 operon encodes the actual effector of the agr system, a 0.5 kb

transcript known as RNA III[124]. AgrD is the prepheromone signal of the agr

quorum sensing system, processed at both the N- and C-terminals by AgrB to

form the mature autoinducing peptide (AIP).[122] The S. epidermidis AIP is an

octapeptdie (DSVCASYF) containing a unique 5 amino acid thiolactone ring

between a conserved central cysteine and the peptide’s C-terminal carboxyl

group[125]. This ring structure is absolutely essential for the full biological

activity of the AIP[126]. After export to the extracellular space (presumably by

AgrB), the AIP binds to and activates the membrane-bound AgrC protein (a

histidine kinase), which in turn activates the cytoplasmic AgrA[122]. The

activated AgrA is responsible for upregulating its own promoter (P2) as well as

the promoter of RNA III (P3)[122]. Thus, in response to increasing cell density,

the agr system results in an exponential increase in expression of both the agr

products and RNA III.

RNA III is the main effector molecule of the agr system, responsible for

encoding the S. epidermidis -toxin[125] as well as up- and down-regulation of

over 135 genes in S. aureus[127]. Through construction of an S. epidermidis agr

deletion mutant, it was revealed that the agr system down-regulates expression of atlE, even during early stages of growth where agr activity is low[121]. Loss of the agr system resulted in a higher concentration of AtlE on the bacterial surface, ultimately leading to significantly increased initial adhesion to polystyrene

surfaces[121]. Interestingly, staphylococcal pheromones demonstrate the

phenomenon of cross-inhibition, where AIPs from self induce the agr response,

while AIPs from non-self inhibit the agr response[128]. S. epidermidis pheromones

have a stronger inhibiting effect on S. aureus than vice versa, potentially

explaining the prevalence of S. epidermidis in implant infections[128]. As deletion

of the agr genes from S. epidermidis results in increased biofilm production,

attempts at inhibiting or removing the agr quorum sensing system as a means of

biofilm control is likely inappropriate. In fact, it has been demonstrated that

treating agr-competent cells with inhibiting AIPs mimics the effects of the agr deletion mutant[123], and an agr-deletion of a clinical strain of S. epidermidis showed increased persistence in an in vivo rabbit infection model[129]. The agr system does not appear to control polysaccharide intercellular adhesin

(PIA)[123].

In 2006, a second quorum sensing system was found to be functional in S. epidermidis, known as luxS[130]. While most details regarding the luxS system are still under investigation, the signaling molecule has been identified as AI-2[130].

While AI-2 has no effect on growth patterns of S. epidermidis, a luxS deletion mutant showed considerably enhanced biofilm formation due to increased PIA production[130]. Thus, in a similar fashion to the agr quorum sensing system which suppresses the atlE gene, luxS works to suppress the icaADBC operon. As

such, inhibiting or deleting luxS enhances PIA production and is likely an

unsuitable target for treatment of biofilms.

1.4 THE HOST RESPONSE

The innate immune system is the body’s primary line of defense against infection, with neutrophils and monocytes playing a key role. Immediately following implantation of a biomaterial, the resulting tissue damage, thrombus formation, and inflammation leads to the release of cytokines that trigger the migration of leukocytes[131, 132]. Following emigration to the site of inflammation, the neutrophils and monocytes are activated, with the initial cell population primarily neutrophils [133]. However, neutrophils are generally short-lived (24-48 hours) and disappear from the implant site rapidly.

Meanwhile, activated monocytes quickly differentiate into macrophages, with a lifetime of up to several months[133].

The primary role of neutrophils and macrophages in an infection scenario is phagocytosis of the invading organisms, as well as release of granular contents[133]. Granular leukocytes contain a variety of granules that serve as a reservoir for both oxygen-dependent and oxygen-independent antimicrobial components prior to their delivery into the phagosome or extracellular space[134]. Such diversity allows the leukocytes to target bacteria in different ways, depending on the conditions encountered[135].

The main oxygen-dependent bactericidal system within neutrophils and macrophages is based on the enzyme nicotinamide adenine dinucleotide

phosphate (NADPH) oxidase that catalyzes the generation of the superoxide

anion (O ) from oxygen[136]. NADPH assembles at the phagosomal membrane

and transfers electrons from the bacterial cytoplasm to oxygen molecules within

the phagosome, forming the reactive oxygen species[137]. O is a mild oxidizing agent that plays a direct role in phagosomal killing, as evidenced by the ability of superoxide dismutase (SOD) to inhibit killing[137]. SOD is known to convert the superoxide radical into hydrogen peroxide (H2O2), which accounts for the

majority of oxygen consumed by the neutrophil[137]. Myeloperoxidase (MPO) is

also released into the phagosome upon neutrophil activation, reacting with H2O2 and oxidizing chloride, resulting in the formation of hypochlorous acid (HOCl), an extremely potent nonradical oxidant[135]. It is believed that some

combination of O and HOCl is responsible for the oxygen-mediated killing of

bacteria in the neutrophil phagosomes.

A second oxygen-dependent bactericidal system found within macrophages

results in the production of nitric oxide (NO). Macrophage activation results in

the expression of inducible nitric oxide synthase (iNOS), which uses L-arginine and oxygen as substrates to synthesize NO[138]. Interestingly, under depleted

L-arginine conditions, iNOS produces a combination of NO and O , which then

interact to form peroxynitrite (ONOO), a potent oxidant that increases

antibacterial activity of macrophages[138]. iNOS is dependent upon calmodulin

binding and functions independently of Ca2+[138].

Both azurophil and specific granules of neutrophils contain a number of oxygen-independent components used to kill invading bacteria, with the specific organism engulfed by the neutrophil determining which granule contents are delivered into the phagosome[139]. For example, azurophil granules contain

-defensins, naturally occurring antimicrobial peptides[140] that kill bacteria through formation of multimeric transmembrane pores[141]. Specific granules contain lactoferrin, an iron-binding glycoprotein known to both impair bacterial growth through iron sequestration and induce cell lysis[142-144]. Both azurophil and specific granuless contain lysozyme, a cationic peptide that cleaves bacterial cell walls and leads to cell lysis[145]. Interestingly, granule contents such as defensins also act to recruit immature dendritic cells in addition to CD4+ and

CD8+ T cells[146]. Thus, the oxygen-independent granule contents serve as both antimicrobial agents and recruiters of the adaptive immune response.

Evidence has shown that implanted biomaterials impair the normal inflammatory response, providing an opportunity for invading organisms to colonize a surface. Initial adhesion of polymorphonuclear leukocytes (PMNs) to poly(etherurethane urea) (PEUU) was shown to be shear sensitive within the range of 0-17 dynes/cm2, with the leukocytes unable to adhere to the biomaterial

at shear stress values >7 dynes/cm2[147, 148]. In addition, PEUU-adherent

neutrophils experience decreased phagocytic capabilities and symptoms of

apoptosis after <60 minutes of adhesion at shear stress levels >6 dynes/cm2[149], while surface-adherent human monocytes exhibit signs of apoptosis in the presence of shear after just 4 hours[150]. Meanwhile, S. epidermidis adhesion is independent of shear, with adhesion occurring even at shear stress values >70 dynes/cm2[151]. Therefore, it is highly plausible that implanted cardiovascular

biomaterials, in the presence of physiological shear stresses, compromise the

ability of leukocytes to adhere to the surface and eliminate colonizing bacteria.

Biomaterial surface chemistry also plays an important role in the host

response. Leukocyte adhesion to biomaterials was found to be highly dependent upon surface chemistry, with monocytes adhering more readily than neutrophils[152]. In addition, neutrophil NO generation is compromised after exposure to biomaterials, with surface chemistry regulating the amount of NO produced[153]. Finally, neutrophil mobility on a biomaterial surface was found to be dependent upon both surface chemistry and adsorbed plasma proteins. In the absence of adsorbed proteins, mobility increased with increasing surface hydrophobicity, whereas the presence of adsorbed proteins resulted in mobility decreasing with increasing hydrophobicity[154].

S. epidermidis also has its own mechanism for inhibiting the host response. As part of the normal blood coagulation process, thrombin acts on the fibrinogen B chain, cleaving the Arg14-Gly15 peptide bond to release the 14 amino acid

fibrinopeptide B (FpB). This cleavage facilitates fibrin assembly and releases FpB to act as a chemotactic agent for blood leukocytes[155]. However, if S. epidermidis manages to bind to the intact fibrinogen B chain by way of its SdrG surface protein, it is able to prevent subsequent cleavage of FpB[66]. In fact, even in the absence of S. epidermidis, only 30% of FpB is released by thrombin from clots of whole blood, with the other 70% remaining intact to provide ample binding sites for subsequent S. epidermidis adhesion[156]. The SdrG-fibrinogen binding interaction therefore provides a significant route for initial bacterial adhesion, as well as a potential mechanism for evasion of the host immune system.

1.5 REFERENCES

1. Darouiche RabihÂ O, Device‐Associated Infections: A Macroproblem that

Starts with Microadherence. Clinical Infectious Diseases, 2001. 33(9): p. 1567‐

1572.

2. Darouiche RO, Treatment of Infections Associated with Surgical Implants. N

Engl J Med, 2004. 350(14): p. 1422‐1429.

3. Schierholz and J., Implant infections: a haven for opportunistic bacteria.

Journal of Hospital Infection, 2001. 49(2): p. 87‐93.

4. Schierholz JM and Beuth J, Implant infections: a haven for opportunistic

bacteria. Journal of Hospital Infection, 2001. 49(2): p. 87‐93.

5. Vuong C and Otto M, Staphylococcus epidermidis infections. Microbes Infect,

2002. 4(4): p. 481‐9.

6. OʹGara JP and Humphreys H, Staphylococcus epidermidis biofilms:

importance and implications. Journal of Medical Microbiology, 2001. 50(7): p.

582‐587.

7. von Eiff C, Peters G, and Heilmann C, Pathogenesis of Infections Due to

Coagulase‐Negative Staphylococci. The Lancet Infectious Diseases, 2002.

2(11): p. 677‐685.

8. Queck SY and Otto M, Staphylococcus epidermidis and other Coagulase‐

Negative Staphylococci, in Staphylococcus Molecular Genetics, J.A. Lindsay,

Editor. 2008, Caister Academic Press: Norfolk, UK.

9. Sabath LD, Garner C, Wilcox C, and Finland M, Susceptibility of

Staphylococcus aureus and Staphylococcus epidermidis to 65 antibiotics.

Antimicrob Agents Chemother, 1976. 9(6): p. 962‐9.

10. Cerca N, Martins S, Cerca F, Jefferson KK, Pier GB, Oliveira R, and

Azeredo J, Comparative assessment of antibiotic susceptibility of coagulase‐

negative staphylococci in biofilm versus planktonic culture as assessed by

bacterial enumeration or rapid XTT colorimetry. Journal of Antimicrobial

Chemotherapy, 2005. 56(2): p. 331‐336.

11. Ceri H, Olson ME, Stremick C, Read RR, Morck D, and Buret A, The

Calgary Biofilm Device: New Technology for Rapid Determination of Antibiotic

Susceptibilities of Bacterial Biofilms. J. Clin. Microbiol., 1999. 37(6): p. 1771‐

1776.

12. Olson M, Ceri H, Morck D, Buret A, and Read R, Biofilm bacteria: formation

and comparative susceptibility to antibiotics. Can. J. Vet. Res., 2002. 66(2): p.

86‐92.

13. Madigan MT and Martinko JM, Brock Biology of Microorganisms. 11 ed.

2006, Upper Saddle River, NJ: Pearson Prentice Hall.

14. Otto M, Staphylococcus epidermidis ‐ the ʹaccidentalʹ pathogen. Nat Rev Micro,

2009. 7(8): p. 555‐567.

15. Vollmer W, Blanot D, and De Pedro MA, Peptidoglycan structure and

architecture. FEMS Microbiology Reviews, 2008. 32(2): p. 149‐167.

16. van Heijenoort J, Formation of the glycan chains in the synthesis of bacterial

peptidoglycan. Glycobiology, 2001. 11(3): p. 25R‐36R.

17. Seltmann G and Holst O, The Bacterial Cell Wall. 2002, New York: Springer.

18. Tipper DJ and Berman MF, Structures of the cell wall peptidoglycans of

Staphylococcus epidermidis Texas 26 and Staphylococcus aureus Copenhagen. I.

Chain length and average sequence of cross‐bridge peptides. Biochemistry, 1969.

8(5): p. 2183‐2192.

19. Otto M, Virulence factors of the coagulase‐negative staphylococci. Front Biosci,

2004. 9: p. 841‐63.

20. Otto M, Staphylococcal biofilms. Curr Top Microbiol Immunol, 2008. 322: p.

207‐28.

21. Endl J, Seidl HP, Fiedler F, and Schleider KH, Chemical composition and

structure of cell wall teichoic acids of staphylococci. Archives of Microbiology,

1983. 135(3): p. 215‐223.

22. Sadovskaya I, Vinogradov E, Li J, and Jabbouri S, Structural elucidation of

the extracellular and cell‐wall teichoic acids of Staphylococcus epidermidis

RP62A, a reference biofilm‐positive strain. Carbohydrate Research, 2004.

339(8): p. 1467‐1473.

23. Gross M, Cramton SE, Gotz F, and Peschel A, Key role of teichoic acid net

charge in Staphylococcus aureus colonization of artificial surfaces. Infect

Immun, 2001. 69(5): p. 3423‐6.

24. Bowden MG, Chen W, Singvall J, Xu Y, Peacock SJ, Valtulina V, Speziale

P, and Hook M, Identification and preliminary characterization of cell‐wall‐

anchored proteins of Staphylococcus epidermidis. Microbiology‐Sgm, 2005. 151:

p. 1453‐1464.

25. Schneewind O, Fowler A, and Faull KF, Structure of the Cell Wall Anchor of

Surface Proteins in Staphylococcus aureus. Science, 1995. 268(5207): p. 103‐

106.

26. Schneewind O, Mihaylova‐Petkov D, and Model P, Cell wall sorting signals

in surface proteins of gram‐positive bacteria. Embo J, 1993. 12(12): p. 4803‐11.

27. Nagarajan R, Antibacterial activities and modes of action of vancomycin and

related glycopeptides. Antimicrob Agents Chemother, 1991. 35(4): p. 605‐9.

28. Comfort D and Clubb RT, A Comparative Genome Analysis Identifies Distinct

Sorting Pathways in Gram‐Positive Bacteria. Infect. Immun., 2004. 72(5): p.

2710‐2722.

29. Palmer J, Flint S, and Brooks J, Bacterial cell attachment, the beginning of a

biofilm. Journal of Industrial Microbiology & Biotechnology, 2007.

34(9): p. 577‐588.

30. Sjollem J, Busscher HJ, and Weerkamp AH, Experimental approaches for

studying adhesion of microorganisms to solid substrata: applications and mass

transport. Journal of Microbiological Methods, 1989. 9(2): p. 79‐90.

31. Marshall KC, Stout R, and Mitchell R, Mechanism of the Initial Events in the

Sorption of Marine Bacteria to Surfaces. J Gen Microbiol, 1971. 68(3): p. 337‐

348.

32. Bayoudh S, Othmane A, Mora L, and Ben Ouada H, Assessing bacterial

adhesion using DLVO and XDLVO theories and the jet impingement technique.

Colloids and Surfaces B: Biointerfaces, 2009. 73(1): p. 1‐9.

33. Hermansson M, The DLVO theory in microbial adhesion. Colloids and

Surfaces B: Biointerfaces, 1999. 14(1‐4): p. 105‐119.

34. Van Oss CJ, Chaudhury MK, and Good RJ, Interfacial Lifshitz‐van der Waals

and polar interactions in macroscopic systems. Chemical Reviews, 1988. 88(6):

p. 927‐941.

35. van Oss CJ, Long‐range and short‐range mechanisms of hydrophobic attraction

and hydrophilic repulsion in specific and aspecific interactions. Journal of

Molecular Recognition, 2003. 16(4): p. 177‐190.

36. Absolom DR, Lamberti FV, Policova Z, Zingg W, van Oss CJ, and

Neumann AW, Surface thermodynamics of bacterial adhesion. Appl. Environ.

Microbiol., 1983. 46(1): p. 90‐97.

37. Van Oss CJ, Interfacial Forces in Aqueous Media. 2 ed. 2006, Boca Raton, FL:

CRC Press.

38. Hoek EMV and Agarwal GK, Extended DLVO interactions between spherical

particles and rough surfaces. Journal of Colloid and Interface Science, 2006.

298(1): p. 50‐58.

39. Boks NP, Norde W, van der Mei HC, and Busscher HJ, Forces involved in

bacterial adhesion to hydrophilic and hydrophobic surfaces. Microbiology, 2008.

154(10): p. 3122‐3133.

40. Heilmann C, Gerke C, Perdreau‐Remington F, and Gotz F, Characterization

of Tn917 insertion mutants of Staphylococcus epidermidis affected in biofilm

formation. Infect. Immun., 1996. 64(1): p. 277‐282.

41. Heilmann C, Hussain M, Peters G, and Gotz F, Evidence for autolysin‐

mediated primary attachment of Staphylococcus epidermidis to a polystyrene

surface. Mol Microbiol, 1997. 24(5): p. 1013‐24.

42. Rupp ME, Fey PD, Heilmann C, and Gotz F, Characterization of the

importance of Staphylococcus epidermidis autolysin and polysaccharide

intercellular adhesin in the pathogenesis of intravascular catheter‐associated

infection in a rat model. Journal of Infectious Diseases, 2001. 183(7): p. 1038‐

1042.

43. Qin Z, Ou Y, Yang L, Zhu Y, Tolker‐Nielsen T, Molin S, and Qu D, Role of

autolysin‐mediated DNA release in biofilm formation of Staphylococcus

epidermidis. Microbiology, 2007. 153(7): p. 2083‐2092.

44. Das T, Sharma PK, Busscher HJ, van der Mei HC, and Krom BP, Role of

Extracellular DNA in Initial Bacterial Adhesion and Surface Aggregation. Appl.

Environ. Microbiol.: p. AEM.03119‐09.

45. Eckhart L, Fischer H, Barken KB, Tolker‐Nielsen T, and Tschachler E,

DNase1L2 suppresses biofilm formation by Pseudomonas aeruginosa and

Staphylococcus aureus. British Journal of Dermatology, 2007. 156(6): p. 1342‐

1345.

46. Izano EA, Amarante MA, Kher WB, and Kaplan JB, Differential Roles of

Poly‐N‐Acetylglucosamine Surface Polysaccharide and Extracellular DNA in

Staphylococcus aureus and Staphylococcus epidermidis Biofilms. Appl. Environ.

Microbiol., 2008. 74(2): p. 470‐476.

47. Rice KC, Mann EE, Endres JL, Weiss EC, Cassat JE, Smeltzer MS, and

Bayles KW, The cidA murein hydrolase regulator contributes to DNA release

and biofilm development in Staphylococcus aureus. Proceedings of the

National Academy of Sciences, 2007. 104(19): p. 8113‐8118.

48. Brash John L and Horbett Thomas A, Proteins at Interfaces, in Proteins at

Interfaces II. 1995, American Chemical Society. p. 1‐23.

49. Vroman L and Adams AL, Findings with the recording ellipsometer

suggesting rapid exchange of specific plasma proteins at liquid/solid interfaces.

Surface Science, 1969. 16: p. 438‐446.

50. Brash JL and Horbett TA, Proteins at Interfaces, in Proteins at Interfaces II.

1995, American Chemical Society. p. 1‐23.

51. Slack SM and Horbett TA, The Vroman Effect: A Critical Review, in Proteins

at Interfaces II: Fundamentals and Applications, T.A. Horbett and J.L. Brash,

Editors. 1995, American Chemical Society. p. 112‐128.

52. Green RJ, Davies MC, Roberts CJ, and Tendler SJB, Competitive protein

adsorption as observed by surface plasmon resonance. Biomaterials, 1999. 20(4):

p. 385‐391.

53. Francois P, Vaudaux P, and Lew PD, Role of Plasma and Extracellular Matrix

Proteins in the Physiopathology of Foreign Body Infections. Annals of Vascular

Surgery, 1998. 12(1): p. 34‐40.

54. Hussain M, Heilmann C, Peters G, and Herrmann M, Teichoic acid enhances

adhesion of Staphylococcus epidermidis to immobilized fibronectin. Microbial

Pathogenesis, 2001. 31(6): p. 261‐270.

55. Chugh TD, Burns GJ, Shuhaiber HJ, and Bahr GM, Adherence Of

Staphylococcus‐Epidermidis To Fibrin‐Platelet Clots Invitro Mediated By

Lipoteichoic Acid. Infection And Immunity, 1990. 58(2): p. 315‐319.

56. Bale MD, Wohlfahrt LA, Mosher DF, Tomasini B, and Sutton RC,

Identification of Vitronectin as a Major Plasma‐Protein Adsorbed on Polymer

Surfaces of Different Copolymer Composition. Blood, 1989. 74(8): p. 2698‐2706.

57. Patti JM and Hook M, Microbial Adhesins Recognizing Extracellular‐Matrix

Macromolecules. Current Opinion In Cell Biology, 1994. 6(5): p. 752‐758.

58. Patti JM, Allen BL, Mcgavin MJ, and Hook M, MSCRAMM‐Mediated

Adherence of Microorganisms to Host Tissues. Annual Review of

Microbiology, 1994. 48: p. 585‐617.

59. Williams RJ, Henderson B, Sharp LJ, and Nair SP, Identification of a

fibronectin‐binding protein from Staphylococcus epidermidis. Infection And

Immunity, 2002. 70(12): p. 6805‐6810.

60. Christner M, Franke GC, Schommer NN, Wendt U, Wegert K, Pehle P,

Kroll G, Schulze C, Buck F, Mack D, Aepfelbacher M, and Rohde H, The

giant extracellular matrix‐binding protein of Staphylococcus epidermidis

mediates biofilm accumulation and attachment to fibronectin. Molecular

Microbiology, 2010. 75(1): p. 187‐207.

61. Arciola CR, Bustanji Y, Conti M, Campoccia D, Baldassarri L, Samori B,

and Montanaro L, Staphylococcus epidermidis ‐ fibronectin binding and its

inhibition by heparin. Biomaterials, 2003. 24(18): p. 3013‐3019.

62. Li DQ, Lundberg F, and Ljungh A, Characterization of vitronectin‐binding

proteins of Staphylococcus epidermidis. Current Microbiology, 2001. 42(5): p.

361‐367.

63. Heilmann C, Identification and characterization of a novel autolysin (Aae) with

adhesive properties from Staphylococcus epidermidis. Microbiology, 2003.

149(10): p. 2769‐2778.

64. Bowden MG, Visai L, Longshaw CM, Holland KT, Speziale P, and Höök

M, Is the GehD Lipase from Staphylococcus epidermidis a Collagen Binding

Adhesin? Journal of Biological Chemistry, 2002. 277(45): p. 43017‐43023.

65. YANAGISAWA N, LI D‐Q, and LJUNGH A, The N‐terminal of

thrombospondin‐1 is essential for coagulase‐negative staphylococcal binding. J

Med Microbiol, 2001. 50(8): p. 712‐719.

66. Davis SL, Gurusiddappa S, McCrea KW, Perkins S, and Hook M, SdrG, a

fibrinogen‐binding bacterial adhesin of the microbial surface components

recognizing adhesive matrix molecules subfamily from Staphylococcus

epidermidis, targets the thrombin cleavage site in the B beta chain. Journal of

Biological Chemistry, 2001. 276(30): p. 27799‐27805.

67. Hartford O, OʹBrien L, Schofield K, Wells J, and Foster TJ, The Fbe (SdrG)

protein of Staphylococcus epidermidis HB promotes bacterial adherence to

fibrinogen. Microbiology‐Sgm, 2001. 147: p. 2545‐2552.

68. McCrea KW, Hartford O, Davis S, Eidhin DN, Lina G, Speziale P, Foster

TJ, and Hook M, The serine‐aspartate repeat (Sdr) protein family in

Staphylococcus epidermidis. Microbiology‐Uk, 2000. 146: p. 1535‐1546.

69. Nilsson M, Frykberg L, Flock JI, Pei L, Lindberg M, and Guss B, A

fibrinogen‐binding protein of Staphylococcus epidermidis. Infection and

Immunity, 1998. 66(6): p. 2666‐2673.

70. Pei L, Arvholm IL, Lonnies L, and Flock JI, GST‐Fbe can recognize beta‐

chains of fibrin(ogen) on explanted materials. Journal of Chromatography B‐

Analytical Technologies in the Biomedical and Life Sciences, 2003. 786(1‐

2): p. 319‐325.

71. Hall AE, Patel PR, Domanski PJ, Prater BD, Gorovits EL, Syribeys PJ,

Vernachio JH, Patti JM, and Hutchins JT, A panel of monoclonal antibodies

recognizing the Staphylococcus epidermidis fibrinogen‐binding MSCRAMM

SdrG. Hybridoma, 2007. 26(1): p. 28‐34.

72. Pei L, Palma M, Nilsson M, Guss B, and Flock JI, Functional studies of a

fibrinogen binding protein from Staphylococcus epidermidis. Infection and

Immunity, 1999. 67(9): p. 4525‐4530.

73. Ponnuraj K, Bowden MG, Davis S, Gurusiddappa S, Moore D, Choe D, Xu

Y, Hook M, and Narayana SVL, A ʺdock, lock, and latchʺ structural model for

a staphylococcal adhesin binding to fibrinogen. Cell, 2003. 115(2): p. 217‐228.

74. Pei L and Flock JI, Lack of fbe, the gene for a fibrinogen‐binding protein from

Staphylococcus epidermidis, reduces its adherence to fibrinogen coated surfaces.

Microbial Pathogenesis, 2001. 31(4): p. 185‐193.

75. Rennermalm A, Nilsson M, and Flock JI, Fibrinogen binding protein of

Staphylococcus epidermidis is a target for opsonic antibodies. Infection and

Immunity, 2004. 72(5): p. 3081‐3083.

76. Guo BN, Zhao X, Shi YG, Zhu DM, and Zhang YY, Pathogenic implication of

a fibrinogen‐binding protein of Staphylococcus epidermidis in a rat model of

intravascular‐catheter‐associated infection. Infection and Immunity, 2007.

75(6): p. 2991‐2995.

77. Watnick P and Kolter R, Biofilm, City of Microbes. J. Bacteriol., 2000. 182(10):

p. 2675‐2679.

78. Tojo M, Yamashita N, Goldmann DA, and Pier GB, Isolation and

Characterization of a Capsular Polysaccharide Adhesin from Staphylococcus

epidermidis. Journal of Infectious Diseases, 1988. 157(4): p. 713‐722.

79. Mack D, Fischer W, Krokotsch A, Leopold K, Hartmann R, Egge H, and

Laufs R, The intercellular adhesin involved in biofilm accumulation of

Staphylococcus epidermidis is a linear beta‐1,6‐linked glucosaminoglycan:

purification and structural analysis. J Bacteriol, 1996. 178(1): p. 175‐83.

80. McKenney D, Pouliot KL, Wang Y, Murthy V, Ulrich M, DÃ¶ring G, Lee

JC, Goldmann DA, and Pier GB, Broadly Protective Vaccine for

Staphylococcus aureus Based on an in Vivo‐Expressed Antigen. Science, 1999.

284(5419): p. 1523‐1527.

81. Maira‐Litran T, Kropec A, Abeygunawardana C, Joyce J, Mark Iii G,

Goldmann DA, and Pier GB, Immunochemical Properties of the Staphylococcal

Poly‐N‐Acetylglucosamine Surface Polysaccharide. Infect. Immun., 2002. 70(8):

p. 4433‐4440.

82. Baldassarri L, Donnelli G, Gelosia A, Voglino MC, Simpson AW, and

Christensen GD, Purification and characterization of the staphylococcal slime‐

associated antigen and its occurrence among Staphylococcus epidermis clinical

isolates. Infect. Immun., 1996. 64(8): p. 3410‐3415.

83. Gotz F, Staphylococcus and biofilms. Molecular Microbiology, 2002. 43: p.

1367‐1378.

84. Mack D, Mechanisms of biofilm formation in and : functional molecules,

regulatory circuits, and adaptive responses. International Journal of Medical

Microbiology, 2004. 294(2‐3): p. 203‐212.

85. Mack D, Davies A, Harris L, Knobloch J, and Rohde H, Staphylococcus

epidermidis Biofilms: Functional Molecules, Relation to Virulence, and Vaccine

Potential, in Glycoscience and Microbial Adhesion, T.K. Lindhorst and S.

Oscarson, Editors. 2009, Springer Berlin / Heidelberg. p. 157‐182.

86. Mack D, Davies AP, Harris LG, Rohde H, Horstkotte MA, and Knobloch

JKM, Microbial interactions in Staphylococcus epidermidis biofilms. Analytical

and Bioanalytical Chemistry, 2006. 387(2): p. 399‐408.

87. Maira‐Litran T, Kropec A, Goldmann D, and Pier GB, Biologic properties

and vaccine potential of the staphylococcal poly‐N‐acetyl glucosamine surface

polysaccharide. Vaccine, 2004. 22(7): p. 872‐879.

88. Sadovskaya I, Vinogradov E, Flahaut S, Kogan G, and Jabbouri S,

Extracellular Carbohydrate‐Containing Polymers of a Model Biofilm‐Producing

Strain, Staphylococcus epidermidis RP62A. Infection and Immunity, 2005.

73(5): p. 3007‐3017.

89. Gerke C, Kraft A, Süßmuth R, Schweitzer O, and Götz F, Characterization of

the N‐Acetylglucosaminyltransferase Activity Involved in the Biosynthesis of the

Staphylococcus epidermidisPolysaccharide Intercellular Adhesin. Journal of

Biological Chemistry, 1998. 273(29): p. 18586‐18593.

90. Heilmann C, Schweitzer O, Gerke C, Vanittanakom N, Mack D, and Gotz

F, Molecular basis of intercellular adhesion in the biofilm‐forming Staphylococcus

epidermidis. Mol Microbiol, 1996. 20(5): p. 1083‐91.

91. Vuong C, A Crucial Role for Exopolysaccharide Modification in Bacterial

Biofilm Formation, Immune Evasion, and Virulence. Journal of Biological

Chemistry, 2004. 279(52): p. 54881‐54886.

92. Vuong C, Voyich JM, Fischer ER, Braughton KR, Whitney AR, DeLeo FR,

and Otto M, Polysaccharide intercellular adhesin (PIA) protects Staphylococcus

epidermidis against major components of the human innate immune system. Cell

Microbiol, 2004. 6(3): p. 269‐75.

93. Conlon KM, Humphreys H, and OʹGara JP, icaR Encodes a Transcriptional

Repressor Involved in Environmental Regulation of ica Operon Expression and

Biofilm Formation in Staphylococcus epidermidis. Journal of Bacteriology,

2002. 184(16): p. 4400‐4408.

94. Cramton SE, Gerke C, Schnell NF, Nichols WW, and Gotz F, The

Intercellular Adhesion (ica) Locus Is Present in Staphylococcus aureus and Is

Required for Biofilm Formation. Infect. Immun., 1999. 67(10): p. 5427‐5433.

95. Allignet J, Aubert S, Dyke KGH, and El Solh N, Staphylococcus caprae

Strains Carry Determinants Known To Be Involved in Pathogenicity: a Gene

Encoding an Autolysin‐Binding Fibronectin and the ica Operon Involved in

Biofilm Formation. Infect. Immun., 2001. 69(2): p. 712‐718.

96. Frank KL and Patel R, Poly‐N‐Acetylglucosamine Is Not a Major Component

of the Extracellular Matrix in Biofilms Formed by icaADBC‐Positive

Staphylococcus lugdunensis Isolates. Infect. Immun., 2007. 75(10): p. 4728‐

4742.

97. Fredheim EGA, Klingenberg C, Rohde H, Frankenberger S, Gaustad P,

Flaegstad T, and Sollid JE, Biofilm Formation by Staphylococcus haemolyticus.

J. Clin. Microbiol., 2009. 47(4): p. 1172‐1180.

98. Rohde H, Frankenberger S, Zähringer U, and Mack D, Structure, function

and contribution of polysaccharide intercellular adhesin (PIA) to Staphylococcus

epidermidis biofilm formation and pathogenesis of biomaterial‐associated

infections. European Journal of Cell Biology, 2010. 89(1): p. 103‐111.

99. Li H, Xu L, Wang J, Wen Y, Vuong C, Otto M, and Gao Q, Conversion of

Staphylococcus epidermidis Strains from Commensal to Invasive by Expression of

the ica Locus Encoding Production of Biofilm Exopolysaccharide. Infection and

Immunity, 2005. 73(5): p. 3188‐3191.

100. Rupp ME, Ulphani JS, Fey PD, Bartscht K, and Mack D, Characterization of

the importance of polysaccharide intercellular adhesin/hemagglutinin of

Staphylococcus epidermidis in the pathogenesis of biomaterial‐based infection in a

mouse foreign body infection model. Infection and Immunity, 1999. 67(5): p.

2627‐2632.

101. Rupp ME, Ulphani JS, Fey PD, and Mack D, Characterization of

Staphylococcus epidermidis polysaccharide intercellular adhesin/hemagglutinin

in the pathogenesis of intravascular catheter‐associated infection in a rat model.

Infection and Immunity, 1999. 67(5): p. 2656‐2659.

102. Mack D, Riedewald J, Rohde H, Magnus T, Feucht HH, Elsner HA, Laufs

R, and Rupp ME, Essential functional role of the polysaccharide intercellular

adhesin of Staphylococcus epidermidis in hemagglutination. Infection and

Immunity, 1999. 67(2): p. 1004‐1008.

103. Mack D, Nedelmann M, Krokotsch A, Schwarzkopf A, Heesemann J, and

Laufs R, Characterization of transposon mutants of biofilm‐producing

Staphylococcus epidermidis impaired in the accumulative phase of biofilm

production: genetic identification of a hexosamine‐containing polysaccharide

intercellular adhesin. Infect. Immun., 1994. 62(8): p. 3244‐3253.

104. Mack D, Siemssen N, and Laufs R, Parallel induction by glucose of adherence

and a polysaccharide antigen specific for plastic‐adherent Staphylococcus

epidermidis: evidence for functional relation to intercellular adhesion. Infect

Immun, 1992. 60(5): p. 2048‐57.

105. Chokr A, Watier D, Eleaume H, Pangon B, Ghnassia J‐C, Mack D, and

Jabbouri S, Correlation between biofilm formation and production of

polysaccharide intercellular adhesin in clinical isolates of coagulase‐negative

staphylococci. International Journal of Medical Microbiology, 2006. 296(6):

p. 381‐388.

106. Frank KL, Hanssen AD, and Patel R, icaA Is Not a Useful Diagnostic Marker

for Prosthetic Joint Infection. Journal of Clinical Microbiology, 2004. 42(10):

p. 4846‐4849.

107. Kogan G, Sadovskaya I, Chaignon P, Chokr A, and Jabbouri S, Biofilms of

clinical strains of Staphylococcus that do not contain polysaccharide intercellular

adhesin. FEMS Microbiology Letters, 2006. 255(1): p. 11‐16.

108. Hussain M, Herrmann M, von Eiff C, Perdreau‐Remington F, and Peters

G, A 140‐kilodalton extracellular protein is essential for the accumulation of

Staphylococcus epidermidis strains on surfaces. Infect. Immun., 1997. 65(2): p.

519‐524.

109. Rohde H, Burdelski C, Bartscht K, Hussain M, Buck F, Horstkotte MA,

Knobloch JKM, Heilmann C, Herrmann M, and Mack D, Induction of

Staphylococcus epidermidis biofilm formation via proteolytic processing of the

accumulation‐associated protein by staphylococcal and host proteases. Molecular

Microbiology, 2005. 55(6): p. 1883‐1895.

110. Bateman A, The G5 domain: a potential N‐acetylglucosamine recognition

domain involved in biofilm formation. Bioinformatics, 2004. 21(8): p. 1301‐

1303.

111. Conrady DG, Brescia CC, Horii K, Weiss AA, Hassett DJ, and Herr AB, A

zinc‐dependent adhesion module is responsible for intercellular adhesion in

staphylococcal biofilms. Proceedings of the National Academy of Sciences,

2008. 105(49): p. 19456‐19461.

112. Banner MA, Cunniffe JG, Macintosh RL, Foster TJ, Rohde H, Mack D,

Hoyes E, Derrick J, Upton M, and Handley PS, Localized Tufts of Fibrils on

Staphylococcus epidermidis NCTC 11047 Are Comprised of the Accumulation‐

Associated Protein. Journal of Bacteriology, 2007. 189(7): p. 2793‐2804.

113. Sun D, Accavitti MA, and Bryers JD, Inhibition of Biofilm Formation by

Monoclonal Antibodies against Staphylococcus epidermidis RP62A

Accumulation‐Associated Protein. Clin. Diagn. Lab. Immunol., 2005. 12(1): p.

93‐100.

114. Conlon KM, Humphreys H, and OʹGara JP, Inactivations of rsbU and sarA

by IS256 Represent Novel Mechanisms of Biofilm Phenotypic Variation in

Staphylococcus epidermidis. J. Bacteriol., 2004. 186(18): p. 6208‐6219.

115. Chaignon P, Sadovskaya I, Ragunah C, Ramasubbu N, Kaplan J, and

Jabbouri S, Susceptibility of staphylococcal biofilms to enzymatic treatments

depends on their chemical composition. Applied Microbiology and

Biotechnology, 2007. 75(1): p. 125‐132.

116. Rohde H, Burandt E, Siemssen N, Frommelt L, Burdelski C, Wurster S,

Scherpe S, Davies A, Harris L, and Horstkotte M, Polysaccharide

intercellular adhesin or protein factors in biofilm accumulation of Staphylococcus

epidermidis and Staphylococcus aureus isolated from prosthetic hip and knee joint

infections☆. Biomaterials, 2007. 28(9): p. 1711‐1720.

117. Cucarella C, Solano C, Valle J, Amorena B, Lasa I, and Penades JR, Bap, a

Staphylococcus aureus Surface Protein Involved in Biofilm Formation. J.

Bacteriol., 2001. 183(9): p. 2888‐2896.

118. Tormo MA, Knecht E, Gotz F, Lasa I, and Penades JR, Bap‐dependent biofilm

formation by pathogenic species of Staphylococcus: evidence of horizontal gene

transfer? Microbiology, 2005. 151(7): p. 2465‐2475.

119. Hussain M, Hastings JGM, and White PJ, Isolation and Composition of the

Extracellular Slime Made by Coagulase‐Negative Staphylococci in a Chemically

Defined Medium. The Journal of Infectious Diseases, 1991. 163(3): p. 534‐

541.

120. Hussain M, Hastings JGM, and White PJ, Comparison of cell‐wall teichoic

acid with high‐molecular‐weight extracellular slime material from Staphylococcus

epidermidis. J Med Microbiol, 1992. 37(6): p. 368‐375.

121. Otto M, Quorum‐sensing control in Staphylococci ‐ a target for antimicrobial

drug therapy? Fems Microbiology Letters, 2004. 241(2): p. 135‐141.

122. Novick RP and Geisinger E, Quorum Sensing in Staphylococci. Annual

Review of Genetics, 2008. 42(1): p. 541‐564.

123. Vuong C, Gerke C, Somerville GA, Fischer ER, and Otto M, Quorum‐

sensing control of biofilm factors in Staphylococcus epidermidis. J Infect Dis,

2003. 188(5): p. 706‐18.

124. Novick R, Projan S, Kornblum J, Ross H, Ji G, Kreiswirth B, Vandenesch F,

Moghazeh S, and Novick R, The agr P2 operon: An autocatalytic sensory

transduction system in Staphylococcus aureus. Molecular and General

Genetics MGG, 1995. 248(4): p. 446‐458.

125. Otto M, Sussmuth R, Jung G, and Gotz F, Structure of the pheromone peptide

of the Staphylococcus epidermidis agr system. FEBS Lett, 1998. 424(1‐2): p. 89‐

94.

126. Mayville P, Ji G, Beavis R, Yang H, Goger M, Novick RP, and Muir TW,

Structure‐activity analysis of synthetic autoinducing thiolactone peptides from

Staphylococcus aureus responsible for virulence. Proceedings of the National

Academy of Sciences, 1999. 96(4): p. 1218‐1223.

127. Dunman PM, Murphy E, Haney S, Palacios D, Tucker‐Kellogg G, Wu S,

Brown EL, Zagursky RJ, Shlaes D, and Projan SJ, Transcription Profiling‐

Based Identification of Staphylococcus aureus Genes Regulated by the agr and/or

sarA Loci. J. Bacteriol., 2001. 183(24): p. 7341‐7353.

128. Otto M, Echner H, Voelter W, and Gotz F, Pheromone Cross‐Inhibition

between Staphylococcus aureus and Staphylococcus epidermidis. Infection and

Immunity, 2001. 69(3): p. 1957‐1960.

129. Vuong C, Kocianova S, Yao Y, Carmody AB, and Otto M, Increased

colonization of indwelling medical devices by quorum‐sensing mutants of

Staphylococcus epidermidis in vivo. J Infect Dis, 2004. 190(8): p. 1498‐505.

130. Xu L, Li H, Vuong C, Vadyvaloo V, Wang J, Yao Y, Otto M, and Gao Q,

Role of the luxS Quorum‐Sensing System in Biofilm Formation and Virulence of

Staphylococcus epidermidis. Infection and Immunity, 2005. 74(1): p. 488‐496.

131. Anderson JM, Chapter 4 Mechanisms of inflammation and infection with

implanted devices. Cardiovascular Pathology. 2(3, Supplement 1): p. 33‐41.

132. Ziats NP, Miller KM, and Anderson JM, In vitro and in vivo interactions of

cells with biomaterials. Biomaterials, 1988. 9(1): p. 5‐13.

133. Anderson JM, BIOLOGICAL RESPONSES TO MATERIALS. Annual

Review of Materials Research, 2001. 31(1): p. 81‐110.

134. Dale DC, Boxer L, and Liles WC, The phagocytes: neutrophils and monocytes.

Blood, 2008. 112(4): p. 935‐945.

135. Faurschou M and Borregaard N, Neutrophil granules and secretory vesicles in

inflammation. Microbes and Infection, 2003. 5(14): p. 1317‐1327.

136. Babior BM, Lambeth JD, and Nauseef W, The Neutrophil NADPH Oxidase.

Archives of Biochemistry and Biophysics, 2002. 397(2): p. 342‐344.

137. Hampton MB, Kettle AJ, and Winterbourn CC, Inside the Neutrophil

Phagosome: Oxidants, Myeloperoxidase, and Bacterial Killing. Blood, 1998.

92(9): p. 3007‐3017.

138. Xia Y and Zweier JL, Superoxide and peroxynitrite generation from inducible

nitric oxide synthaseâ€‰inâ€‰macrophages. Proceedings of the National

Academy of Sciences, 1997. 94(13): p. 6954‐6958.

139. Joiner KA, Ganz T, Albert J, and Rotrosen D, The opsonizing ligand on

Salmonella typhimurium influences incorporation of specific, but not azurophil,

granule constituents into neutrophil phagosomes. The Journal of Cell Biology,

1989. 109(6): p. 2771‐2782.

140. Ganz T, Selsted ME, Szklarek D, Harwig SS, Daher K, Bainton DF, and

Lehrer RI, Defensins. Natural peptide antibiotics of human neutrophils. The

Journal of Clinical Investigation, 1985. 76(4): p. 1427‐1435.

141. Wimley WC, Selsted ME, and White SH, Interactions between human

defensins and lipid bilayers: Evidence for formation of multimeric pores. Protein

Science, 1994. 3(9): p. 1362‐1373.

142. Chapple DS, Mason DJ, Joannou CL, Odell EW, Gant V, and Evans RW,

Structure‐function relationship of antibacterial synthetic peptides homologous to

a helical surface region on human lactoferrin against Escherichia coli serotype

O111. Infection and Immunity, 1998. 66(6): p. 2434‐2440.

143. Cramer E, Pryzwansky KB, and Villeval JL, Ultrastructural localization of

lactoferrin and myeloperoxidase in human neutrophils by immunogold. Blood,

1985. 65(2): p. 423‐432.

144. Oram JD and Reiter B, Inhibition of bacteria by lactoferrin and other iron‐

chelating agents. Biochimica et Biophysica Acta (BBA) ‐ General Subjects,

1968. 170(2): p. 351‐365.

145. Lollike K, Kjeldsen L, Sengelov H, and Borregaard N, Lysozyme in human

neutrophils and plasma. A parameter of myelopoietic activity. Leukemia, 1995.

9(1): p. 159‐164.

146. Yang D, Chen Q, Chertov O, and Oppenheim JJ, Human neutrophil

defensins selectively chemoattract naive T and immature dendritic cells. Journal

of Leukocyte Biology, 2000. 68(1): p. 9‐14.

147. Sapatnekar S, Kao WJ, and Anderson JM, Leukocyte—biomaterial

interactions in the presence of Staphylococcus epidermidis: Flow cytometric

evaluation of leukocyte activation (Student Research Award in the Hospital

Intern, Resident, or Clinical Fellow Category, 23rd Annual Meeting of the

Society for Biomaterials, New Orleans, LA, April 30–May 4, 1997). Journal of

Biomedical Materials Research, 1997. 35(4): p. 409‐420.

148. Shive MS, Hasan SM, and Anderson JM, Shear stress effects on bacterial

adhesion, leukocyte adhesion, and leukocyte oxidative capacity on a

polyetherurethane. Journal of Biomedical Materials Research, 1999. 46(4): p.

511‐519.

149. Shive MS, Salloum ML, and Anderson JM, Shear stress‐induced apoptosis of

adherent neutrophils: A mechanism for persistence of cardiovascular device

infections. Proceedings of the National Academy of Sciences, 2000. 97(12):

p. 6710‐6715.

150. Shive MS, Brodbeck WG, Colton E, and Anderson JM, Shear stress and

material surface effects on adherent human monocyte apoptosis. Journal of

Biomedical Materials Research, 2002. 60(1): p. 148‐158.

151. Wang I‐w, Anderson JM, and Marchant RE, Staphylococcus epidermidis

Adhesion to Hydrophobic Biomedical Polymer Is Mediated by Platelets. Journal

of Infectious Diseases, 1993. 167(2): p. 329‐336.

152. Patel JD, Ebert M, Stokes K, Ward R, and Anderson JM, Inhibition of

bacterial and leukocyte adhesion under shear stress conditions by material surface

chemistry. Journal of Biomaterials Science, Polymer Edition, 2003. 14: p.

279‐295.

153. Patel JD, Krupka T, and Anderson JM, iNOS‐mediated generation of reactive

oxygen and nitrogen species by biomaterial‐adherent neutrophils. Journal of

Biomedical Materials Research Part A, 2007. 80A(2): p. 381‐390.

154. Zhou Y, Doerschuk CM, Anderson JM, and Marchant RE, Biomaterial

surface‐dependent neutrophil mobility. Journal of Biomedical Materials

Research Part A, 2004. 69A(4): p. 611‐620.

155. Senior R, Skogen W, Griffin G, and Wilner G, Effects of Fibrinogen

Derivatives Upon the Inflammatory Response. J. Clin. Invest., 1986. 77: p.

1014‐1019.

156. Brummel K, Butenas S, and Mann K, An Integrated Study of Fibrinogen

During Blood Coagulation. J. Biol. Chem., 1999. 274(32): p. 22862‐22870.

CHAPTER 2

BACTERIAL RESISTANCE TO ANTIBIOTICS

2.1 ANTIBIOTICS AND RESISTANCE TO TREATMENT

Penicillin, despite being discovered in 1928, was not first clinically applied until the mid-1940’s[1, 2]. The following twenty years saw a surge in discovery of additional antibiotics, mainly through screening of soil bacteria for the production of antimicrobial compounds. However, with very few exceptions, all of the antibiotics approved for clinical use since the early 1960’s have been synthetic derivatives of preexisting antibiotic scaffolds[3]. In fact, four such scaffolds (cephalosporins, penicillins, quinolones, and macrolides) accounted for

73% of all new antibacterial compounds between 1981 and 2005[3]. One of the main obstacles to novel drug discovery in the antibacterial field is financial in nature. When a large pharmaceutical company considers developing an antibiotic drug, the product must compete with all other drugs offered by the company in terms of profitability and bottom line. When compared to other drugs, antibiotics are utilized by a narrow spectrum of patients for only a short period of time. In addition, antibiotic resistance can arise rapidly within a bacterial population, rendering the new drug unprofitable. Finally, regulatory requirements for new antibiotics don’t take in to account the fact that the superiority of the new drug might reside in its ability to kill drug resistant

organisms. As a result, clinical trials are often carried out on patients infected

with non-resistant bacteria, and the new drug is not seen as an improvement

over existing drugs[4]. Development of novel antibiotics is therefore seen as a

risky, often times unprofitable venture for a pharmaceutical company, making it much safer and cheaper to instead modify existing drugs.

2.2 OVERVIEW OF ANTIBIOTICS

Antibiotics exert their affect on bacterial targets by interfering with vital

metabolic and physiologic components of the cell, mainly translation, DNA

replication, metabolism, cell wall biosynthesis, and cell membrane integrity[2].

Depending on the bacterial species and mode of action, antibiotics can be either

bacteriostatic or bactericidal. Bacteriostatic antibiotics arrest cell growth, but

upon removal of the antibiotic, the cells will begin growing again. Bactericidal

antibiotics, on the other hand, are able to permanently damage and kill the cell.

Antibiotic classes that are currently utilized in clinical settings are listed in Table

2.1, organized by their intended target. The following section will provide a brief

overview of several relevant classes of antibiotics.

Table 2.1: Clinicaly Administered Antibiotics

Target Antibiotic Class Examples

DNA Replication Quinolones Ciprofloxacin

Metabolism Pyrimidines Trimethoprim

Sulfonamides Sulfamethoxazole

Peptidoglycan Biosynthesis -Lactams Penicillins (ampicillin) Cephalosporins (cephamycin) Penems (meropenem) Monobactams (aztreonam)

Glycopeptides Vancomycin Teicoplanin

Translation Aminoglycosides Gentamicin Streptomycin Spectinomycin

Tetracyclines Minocycline Tigecycline

Macrolides Erythromycin Azithromycin

Lincosamides Clindamycin

Streptogramins Synercid

Oxazolidinones Linezolid

Phenicols Chloramphenicol

Rifamycins Rifampin

Cell Membrane Lipopeptides Daptomycin

Cationic Peptides Colistin

Inhibition of DNA replication

Quinolones

Quinolones were first utilized in 1962 with the introduction of nalidixic acid[5]. Over the subsequent 50 years, second-, third-, and fourth-generation fluoroquinolones have been developed through structural modifications of the original quinolone scaffold. While the first generation of quinolones were only effective in treating gram-negative bacteria, successive generations of fluoroquinolones became better suited for treating both gram-negative and gram-positive bacteria[5, 6].

Quinolones have excellent oral absorption and bioavailability, exhibit concentration dependent bactericidal activity, with optimal serum concentrations of 30 times the MIC being reached after just 1 to 2 hours[7]. The mode of action for quinolones is to inhibit DNA synthesis through interactions with both DNA gyrase and topoisomerase IV[8]. Sensativities of DNA gyrase and topoisomerase

IV vary in a given bacterium, with DNA gyrase more sensitive in gram-negative bacteria and topoisomerase IV more sensitive in gram-positive bacteria[5, 8].

Ultimately, the formation of a quinolone-DNA-DNA gyrase or quinolone-DNA- topoisomerase IV complex results in cleavage of the DNA that rapidly kills the bacteria[5, 8]. As such, quinolones do not require the bacteria to be growing and/or dividing in order to exert their affect. However, the fluoroquinolones are rarely used as a first line of defense against Stapholoccus spp, as bacterial

resistance is quickly acquired, and resistance to a single quinolone could render

the species resistant to all quinolones[7].

Metabolic Inhibitors

Sulfonamides (Sulfa Drugs)

Sulfonamides target the enzyme dihydropteroate synthase (DHPS), which is

known to catalyzes the formation of dihydropteroic acid, the precursor to dihydrofolic acid[9]. In a sense, the Sulfa drugs starve the bacterial cells. Since

mammalian cells are able to utilize dietary folates and thus lack DHPS,

sulfonamides were the first class of antibiotics that could be used to selectively

target a broad spectrum of prokaryotic cells[9]. Despite the effectiveness of the

sulfonamide antibiotics, their use is very limited over the past few decades. Due to excessive use in its early years, sulfonamide-resistance is very common. In addition, sulfonamide allergies have become more widespread[9].

Inhibition of Peptidoglycan Synthesis

-lactams

-lactam antibiotics, including penicillin, ampicillin, and methicilllin, are arguably the most important class of antibiotics in the course of modern medicine. Despite finding widespread use in the 1940’s, it wasn’t until the mid

1960’s that the mechanistic action of penicillin was understood. Tipper and

Strominger first proposed in 1965 that penicillin interfered with the terminal

transpeptidation step in bacterial cell wall synthesis[10]. Specifically, it was

proposed that -lactam ring of penicillin is a structural analog for the acyl-D-

alanyl-D-alanine sequence found in the linear glycopeptides chains prior to

crosslinking. As such, penicillin is able to inhibit the transpeptidase responsible

for creating the crossbridges necessary for structural rigidity[10]. Subsequent

studies revealed that there were between three and eight distinct penicillin- binding proteins (PBPs) in a given bacterium[11]. Penicillin works by forming an ester bond with these PDBs, sequestering the enzyme and prevening crossbridge formation[12]. Substitution of the penicillin molecule’s -lactam side group

forms a new drug (ampicillin or methicillin, for example), imparting unique

properties upon the new drug, such as resistance to degradation by penicillinase,

tolerance of acidity, or the ability to penetrate the gram-negative cell

envelope[13]. Unfortunately, -lactam resistance is becoming widespread, as

indicated by the prevalence of methicillin resistant S. aureus and S. epidermidis

(MRSA and MRSE, respectively) in the clinical setting[12].

Glycopeptides

Glycopeptide antibiotics are large, rigid, complex molecules synthesized by

both the Streptomyces and Actinoplanes species. Vancomycin was the first such

antibiotic of this class, discovered in 1956 by researchers at Eli Lilly and

Company[14]. Due to toxic side effects, glycopeptide antibiotics did not initially

see general clinical use and were instead superseded by the -lactam antibiotics[15]. However, it has since been determined that the initial toxic side

effects were due to poor purification of the glycopeptides, and the glycopeptide antibiotics are now seeing increased clinical usage[16].

The general structure of the glycopeptide antibiotics consists of a central heptapeptide domain that has undergone extensive oxidative crosslinking, with five of the seven amino acid residues conserved among all glycopeptides[17, 18].

Glycopeptides typically differ at positions 1 and 3 of the heptapeptide chain. In addition, the substituents of the aromatic amino acids show variability, with various chlorine, hydroxyl, methyl, sugar, or aminosugar substitutions[18]. The basic crosslinked heptapeptide structure is the biologically active portion of the antibiotic, and is known as the aglycone. The sugars and aminosugars are important for stabilizing the molecule and imparting various pharmacokinetic properties[17, 18].

Glycopeptide antibiotics are too large to penetrate the bacterial cytoplasmic membrane, and as such act primarily against gram positive bacteria by preventing the incorporation of N-acetylglucosamine and N-acetylmuramic acid into the growing peptidoglycan matrix[19]. Vancomycin forms a series of five hydrogen bonds with the terminal acyl-D-Alanine-D-Alanine residues of the peptidoglycan intermediates, thereby interfering with the biosynthetic pathway used to build a stable cell wall[20]. In contrast to the -lactam antibiotics which sequester the transpeptidase enzymes necessary for forming the peptidoglycan crossbridges, glycopeptide antibiotics interact with the enzyme’s substrate and shield it from the transpeptidase’s activity[18, 21].

Inhibition of Translation

Aminoglycosides

Aminoglycosides are a large, highly-polar class of antibiotics that exert their

effect through inhibiting one or more biochemical steps involved in protein

translation on the prokaryotic ribosome[22]. The primary structure consists of

several aminated sugars joined to a dibasic cyclitol (such as 2-deoxystreptamine,

streptidine, or fortamine)[23]. After electron transport across the cytoplasmic

membrane, the aminoglycosides bind to the 30S subunit of the bacterial ribosome and impairs the proofreading process controlling translational accuracy[23].

Although very specific for prokaryotic ribosomes, care must be taken in selecting a proper dosage for these drugs, as high concentrations can begin to affect eukaryotic ribosomes through nonspecific binding[23]. Aminoglycosides are primarily reserved for severe gram-negative infections, but recently they have also been used in serious gram-positive infections involving Staphylococcus and

some streptococci. However, due to the poor nature of uptake across the

cytoplasmic membrane, as well as toxicity to the patient in high doses,

aminoglycosides are much less effective against gram-positive bacteria[24]. Due

to poor oral absorption, this class of antibiotics is administered by parenteral

injection[24].

Macrolides, Lincosamides, and Streptogramin B

Macrolides, lincosamides, and streptogramins (MLS), despite being quite

different structurally, all function in an identical manner[25]. The MLS antibiotics

have a spectrum targeting mainly gram-positive cocci (staphylococci and

streptococci in particular), as well as gram-positive bacilli[26]. Structurally,

macrolides contain 14-, 15-, or 16- membered lactone rings substituted with several neutral or amino sugars. Erythromycin, the most frequently used

macrolide, has a 14-membered ring[27]. Lincosamides do not contain a lactone

ring, and of the hundreds of synthetic derivates available, only clindamycin and

lincomycin A are suitable for clinical use[25, 26]. Meanwhile, streptogramins are

produced by the Streptomyces genus and are comprised of two subgroups, type

A and type B, produced in a 70:30 ratio. Type A streptogramins are cyclic,

polyunsaturated macrolactones, while Type B streptogramins are cyclic hepta-or

hexa-depsipeptides[28]. The MLS antibiotics block ribosomal protein synthesis

by inhibiting the peptidyltransferase reaction on the prokaryotic 50S ribosomal

subunit[25]. The antibiotics essentially block the exit path for growing peptides from the 50S ribosomal subunit, causing premature dissociation of peptidyl- tRNAs from the ribosome[27].

Oxazolidinones

Oxazolidinones are the only truly new class of antibiotics to be discovered and enter into clinical usage within the last 30 years[29-31]. Linezolid was the first oxazolidinone to be clinically available, launched in 2000 by Amersham

Pharmacia (Pfizer)[29]. Oxazolidinones are extremely effective at treating gram- positive organisms such as methicillin-resistant Staphylococcus aureus, penicillin- resistant Streptococcus pneumonia, and vancomycin-resistant Enterococcus

faecium[30]. Oxazolidinones target the bacterial ribosome and inhibit protein synthesis, although the exact mechanism remains controversial[29]. One plausible explanation is that oxazolidinones act by interfering with the formation of the tRNA-mRNA-70S (or 30S) complex necessary for initiation of translation[31].

Rifamycins

Rifamycins are an old class of antibiotics, first isolated in 1959 as a mixture of natural products[32]. Rifamycins are potent against gram-positive bacteria and particularly mycobacteria, and as such are the drug of choice for treating tuberculosis[32]. The drugs inhibit DNA-dependent RNA synthesis in prokaryotic cells by binding to RNA polymerase and obstructing the growing oligonucleotide chain after either the first or second chain elongation step[32].

Rifampin is the prototypical rifamycin, highly active against both coagulase- positive and coagulase-negative staphylococci (CoNS)[33]. As such, the

American Heart Association recommends administering rifampin orally as part of the standard treatment for S. epidermidis prosthetic valve endocarditis[33].

2.3 ANTIBIOTIC RESISTANCE

Not long after the discovery of penicillin, and several years before this first antibiotic even found widespread clinical use, bacterial resistance to antibiotics was encountered in the form of -lactamase. Since that point, is has become evident that bacteria have the ability to alter their susceptibility to antibiotics on

both the genotypic and phenotypic level. This section gives a brief summary of the most common mechanisms for antibiotic resistance, mainly efflux pumps, enzymatic degradation of antibiotics, modification of antibiotic targets, and reprogramming peptidoglycan synthesis. Antibiotic classes affected by these resistance mechanisms are summarized in Table 2.2.

Antibiotic Efflux Pumps

Efflux pumps are one of the most common means by which bacteria regulate intracellular antibiotic concentrations. In general, efflux pumps confer a moderate resistance to antibiotics (up to 64-fold increase in MIC)[34], particularly with macrolides, tetracyclines, and fluoroquinolones[35]. These antibiotic transporters, based upon structural similarities, can be divided into superfamilies. There are six such superfamiles currently recognized in prokaryotic cells: the Major Facilitator Superfamily (MFS), Resistance Nodulation

Division (RND), Drug Metabolite Transporter (DMT), Proton-dependent

Oligopeptide Transporter (POT), Multidrug/Oligosaccharidyl- lipid/Polysaccharide Flippase (MOP), and the ATP Binding Cassette (ABC) superfamilies[34]. As a testament to the ubiquitous nature of efflux pumps in nature, all six of these superfamilies are also present in eukaryotic cells[34].

Gram-positive bacteria possess a wide array of efflux pumps, typically consisting of a single protein component with 12 transmembrane domains.

Meanwhile, efflux pumps found in gram-negative bacteria often times contain

Table 2.2: Modes of Antibiotic Resistance

Mode of Resistance Antibiotic Classes Affected

Efflux Pumps -Lactams Aminoglycosides Tetracylcines Macrolides Lincosamides Streptogramins Oxazolidinones Phenicols Quinolones Pyrimidines Sulfonamides Rifamycins Cationic Peptides

Hydrolysis -Lactams Macrolides

Acetylation Aminoglycosides Streptogramins Phenicols Quinolones

Altered Target -Lactams Aminoglycosides Tetracyclines Macrolides Lincosamides Streptogramins Oxazolidinones Phenicols Quinolones Pyrimidines Sulfonamides Rifamycins Lipopeptides Cationic Peptides

Reprogramming Glycopeptides Peptidoglycan Biosynthesis

multiple protein components due to the presence of both an inner and outer

membrane[34, 36]. In general, efflux pumps have very poor selectivity in regards

to molecular targets. Most targets simply must be amphiphilic and have a

functional group susceptible to ionization[34, 35]. As a result, many pumps are

known to transport multiple antibiotics that are structurally unrelated.

Interestingly, due to their largely hydrophilic nature, efflux pumps for both

aminoglycoside and glycopeptides antibiotics are very rare[35]. In addition,

glycopeptides antibiotics (such as vancomycin) function in the extracellular space

and never cross the cell membrane. Therefore, efflux pumps would be ineffective

against this class of antibiotics.

Hydrolysis of -lactam Antibiotics

One of the most significant modes of antibiotic resistance involves enzymes

capable of hydrolyzing the -lactam family of antibiotics. The first pencillinase,

-lactamase, was discovered by Abraham and Chain in 1940, well before the

drug even saw widespread clinical use[37]. Since then, thousands of similar

enzymes have been discovered in a wide variety of bacterial species[38]. The - lactamase enzymes exert their effect by hydrolyzing the -lactam ring, rendering the antibiotic inactive[38]. Two such classes of enzymes are known to exist. The first class acts by forming a covalent antibiotic-enzyme intermediate followed by a rapid hydrolysis step to inactivate the -lactam antibiotic. The seccond class

utilizes a divalent Zn2+ ion to activate a water molecule that attacks the lactam

bond directly[38].

As enzymatic hydrolysis mechanisms become more widespread in the

bacterial population, -lactam drugs are continuously modified in an attempt to evade inactivation. However, bacteria respond with modifications and mutations of their own -lactamase enzymes, making it a constant struggle to create new, effective drugs[38, 39].

Chemical Modification of Target Molecules

In addition to removing and inactivating antibiotics through efflux pumps

and hydrolyzing enzymes, bacteria are also known to chemically modify their

cellular components in order to prevent binding between the antibiotic and its

target site. Common means of modification include acetylation, methylation,

phosphorylation of binding sites, as well as point mutations of the relevant

genes. While these modifications greatly reduce the function of the antibiotics,

they have little effect on cell function[39].

An excellent example of target modification explains the broad spectrum

antibiotic resistance of some S. aureus and E. coli strains towards Phenicols,

Lincosamides, Oxazolidinones, Pleuromutilins, and Streptogramin A antibiotic

classes (known as the PhLOPSA phenotype)[40]. Despite their differences in structure, all 5 of these classes exert their antibacterial affects through binding to

the peptidyl transferase center on the bacterial 23S rRNA. Bacterial members of

the PhLOPSA phenotype contain the cfr gene and express a Cfr rRNA methyltransferase responsible for methylating a nucleotide in the 23S rRNA peptidyl transferase center. As a result, the affected antibiotics have greatly reduced binding and MICs 4- to ≥4096-fold higher than the cfr negative strains[40].

Reprogramming Peptidoglycan Biosynthesis

When exposed to glycopeptide antibiotics, rather than chemically modifying the target of the antibiotic, the bacteria often times choose to bypass the antibiotic-sensitive system altogether. Instead, they create an entirely new pathway that is not sensitive to the antibiotic. As an example, the most common reason for vancomycin resistance in a microbial population is a mutation that results in the peptidoglycan precursors terminating with –D-Alanine-D-Lactate instead of the typical –D-Alanine-D-Alanine (figure 2.1). By replacing an amide bond with an ester bond, one of the hydrogen bonds with vancomycin is eliminated, electronic repulsion is introduced, and binding affinity is lowered by up to three orders of magnitude[39, 41]. This complicated reprogramming of cell wall synthesis involves activation of five new genes[39]. It is believed that the origin of these genes conferring vancomycin resistance arose through a transfer from the microorganisms that produce glycopeptides, as Marshall et al demonstrated that six glycopeptides producing strains (Streptomyces toyacaensis

A47934, Amycolatopsis orientalis C329.2, A. orientalis 18098, A. orientalis subspecies

Figure 2.1: Binding interaction between vancomycin and peptidoglycan acyl-D- Ala-D-Ala. A series of five hydrogen bonds are formed. Vancomycin resistant bacteria reprogram their peptidoglycan synthesis, replacing an amide bond with an ester bond, forming acyl-D-Ala-D-Lac. This eliminates one of the hydrogen bonds (shown in red) and introduces electrostatic repulsion. As a result, binding affinity is reduced by three orders of magnitude.

lurida, and A. coloradensis subspecies labela) contained all the genes necessary to

confer glycopeptide resistance[42].

2.4 THE BIOFILM AS A SOURCE OF PROTECTION

The success of S. epidermidis infections relies upon the ability of the bacteria to

form a biofilm after initial adhesion to a substrate[43, 44]. While planktonic S.

epidermidis is known to be susceptible to a large number of antibiotics[45], the

biofilm environment offers the encapsulated bacteria increased resistance, often

times able to survive antibiotic concentrations several orders of magnitude

higher than the minimum inhibitory concentration (MIC) and minimum

bactericidal concentration (MBC) measured in planktonic suspensions[46-51].

While specific genetic resistance mechanisms may develop within a subset of the

bacterial population due to exposure to antibiotics, the biofilm environment itself

can also contribute to increased resistance and tolerance in a non-genetic manner.

The most likely factors contributing to such resistance to antibiotics within a

biofilm will be reviewed here. Mainly, slow or incomplete antibiotic penetration

due to the polysaccharide matrix, an altered microenvironment leading to slower

bacterial growth, and an adaptive stress response leading to a small population

of bacteria that differentiate into persister cells.

1) The Polysaccharide Matrix

The mature biofilm matrix is comprised primarily of water [50, 51], and

theoretical calculations have indicated that, in the absence of significant

irreversible sorption or chemical inactivation, such a matrix would lack the ability to considerably retard the penetration of antibiotics into the biofilm[52].

These predictions are supported by Rani et al, wherein confocal microscopy was used to show that low molecular weight fluorescent tracers (approximately the same size as most antibiotics) were able to saturate the interior of S. epidermidis biofilm clusters several hundred micrometers in diameter within just a few minutes[53]. However, subsequent in vitro studies have yielded conflicting results regarding these claims.

Various studies have implicated the Staphylococcal polysaccharide matrix in the reduction of antibiotic efficacy. After isolating the polysaccharide component of the biofilm matrix, it was demonstrated that this polysaccharide extract interfered with the action of vancomycin, resulting in up to a four-fold increase in the minimum inhibitory concentration (MIC). Similar results were seen for teicoplanin, while there was no change in the MIC for clindamycin, cefazolin, or rifampin[54]. The authors of this study proposed that the polysaccharide either complexes with vancomycin and renders it inactive, or that the polysaccharide coats the walls of the bacteria and interferes with the action of vancomycin[54]. A subsequent study demonstrated that the matrix produced by clinical isolates of S. epidermidis were able to significantly decrease the effectiveness of vancomycin, teicoplanin, and pefloxacin (63%, 52%, and 30% reductions, respectively), while rifampin effectiveness was not affected (0.99% reduction). Meanwhile, 18 other antimicrobial agents tested showed a moderate decrease in effectiveness (1.4-16%

reduction)[55]. König and Blaser, using crude slime extract (CSE) from S. epidermidis biofilms, demonstrated that introducing CSE into a suspension of S. epidermidis increased the vancomycin MICs and MBCs by eight- and sixteen-fold, respectively. Similarly, teicoplanin MICs and MBCs increase four- to eight-fold, depending on the source of the CSE. Activities of fleroxacin, flucloxacillin, and rifampin were not significantly influenced by the CSE[56]. Interestingly, exposing solutions of vancomycin to high concentrations of CSE for 30 minutes had no effect on the free concentration of vancomycin. This indicates that a binding or adsorption process between vancomycin and the S. epidermidis polysaccharide matrix is highly unlikely. Instead, the authors hypothesized that glycopeptides activity is not compromised by limited diffusion or adsorption due to the biofilm matrix. Instead, it is likely that the polysaccharides interact directly with the bacterial cell in order to affect their influence[56]. Mathur et al described a linear increase in vancomycin and teicoplanin MICs with increasing concentrations of added polysaccharides to the culture media. MICs increased two- to eight-fold with the addition of 5 mg/ml slime extract, while MICs increased four- to sixteen-fold in the presence of 20 mg/ml slime extract.

Meanwhile, quinupristin/dalfopristin and linezolid showed just a slight increase in MICs (one- to two-fold), and ranbezolid and rifampicin MICs were completely unaffected by the addition of purified slime extract[57]. Most recently, Singh et al used S. epidermidis and S. aureus biofilms grown on porous membranes to demonstrate the inability of vancomycin and the -lactams oxacillin and

cefotaxime to fully penetrate a biofilm. At the same time, amikacin and ciprofloxacin were unaffected by the biofilms and fully penetrated through the underlying membrane[58].

There are also a number of reports demonstrating the successful penetration of the biofilm by a variety of antibiotics. S. epidermidis biofilms formed on dialysis membranes allowed both vancomycin and rifampin to penetrate the biofilm and cross the membrane. When bulk concentrations of vancomycin (40 g/ml) or rifampin (20 g/ml) were placed in contact with the biofilm, concentrations of 8

g/ml vancomycin or 3.1 g/ml rifampin were observed on the opposite side of the membrane after 72 hours[59]. Both of these values are well above the MICs and MBCs for S. epidermidis grown in suspension. However, they are well below the MICs and MBCs observed for surface-adherent biofilms, and as such viable bacteria were recovered from the membrane-bound biofilms. A separate study conducted using biofilms formed on stainless steel prostheses showed the biofilm’s ability to uptake significant quantities of vancomycin (1184 g/ml), well in excess of the MIC and MBC. However, despite the high concentration of vancomycin within the biofilms, viable cells were still recovered after treatment[60]. Interestingly, after dispersing the cells in a biofilm, the newly suspended bacteria are once again susceptible to the same MIC and MBC levels found in planktonic bacteria, indicating that antibiotic resistance within a biofilm is not obtained through irreversible genotypic changes[60]. One study conducted using confocal microscopy and a fluorescently tagged version of

vancomycin demonstrated that the glycopeptide antibiotic was able to fully

penetrate an S. aureus biofilm. However, it took >1 hour to completely penetrate the biofilm and bind to bacteria at the deepest levels of the biofilm, whereas

vancomycin was able to bind to planktonic bacteria within 5 minutes[61]. Most

importantly, polysaccharide intercellular adhesin (PIA) was shown to play no

role in the decreased rate of S. aureus biofilm penetration by vancomycin. In fact,

it was found that the amount of vancomycin binding to PIA positive cells was

greater than the amount of vancomycin binding to PIA negative cells[61]. In a

similar experiment, fluorescently labeled daptomycin was able to fully penetrate

S. epidermidis biofilms in just 1-2 minutes. The relative effective diffusivity of

daptomycin in a biofilm was estimated to be 28% of the diffusion coefficient in pure water. This diffusivity is in line with values observed for other antibiotics, and a consensus value of 25% has been recommended by Stewart for the effective diffusivity of organic solutes in S. epidermidis biofilms[62, 63].

Taken together, these studies indicate that the polysaccharide matrix present in S. epidermidis biofilms cannot be fully responsible for the ineffectiveness of antibiotics in eradicating such biofilms. In some cases, antibiotics have no problems penetrating the biofilm. In other cases, penetration is either retarded or eliminated. However, in most situations the polysaccharide matrix itself is found to be responsible for no more than a sixteen-fold reduction in antibiotic effectiveness…far less than the thousand-fold reduction observed in many experimental settings.

2) Microenvironment Effects: Slower Growth Rate

The biofilm itself is a heterogeneous environment, characterized by the

presence of oxygen, nutrient, and waste gradients. Such gradients undoubtedly

lead to changes in bacterial growth rates within the biofilm. Many antibiotics are

only functional against bacteria while they are actively growing and dividing. As

such, decreases in bacterial growth rates will lead to a reduced antibiotic efficacy

within the biofilm[64].

The primary suspect in reduced growth rates is oxygen starvation within the

biofilm. A simple yet clever experiment monitoring alkaline phosphatase (APase)

expression within P. aeruginosa biofilms was used to spatially monitor oxygen

content within the biofilm[65]. APase is only expressed under phosphate-poor,

oxygen-rich conditions. Therefore, by carrying out experiments in phosphate-

depleted media, the presence or absence of APase was used to characterize the

oxygen levels within the biofilm. Biofilms ranging from 117-151 m thick were

shown to have approximately 1/15th the APase activity of planktonic cultures,

with all of the APase activity localized to a 30 m thick band adjacent to the

media. Furthermore, measurements with an oxygen microelectrode within the biofilm confirmed that oxygen was completely absent at depths greater than 30

m[65].

Anderl et al, after confirming both ciprofloxacin and ampicillin are able to penetrate Klebsiella pneumonia biofilms, demonstrated that the bacteria could survive prolonged chemotherapy with these antimicrobial agents[66]. In order to

investigate the significance of nutrient limitations and growth rate on antibiotic

susceptibility, planktonic exponential-phase (E) and stationary-phase(S) K.

pneumonia was inoculated into either nutrient-rich (N+) or nutrient-poor (N-)

media. The E/N+ combination resulted in the bacteria most susceptible to

antibiotic killing, while the E/N- and S/N+ combinations resulted in reduced

bacterial killing. The S/N- combination resulted in bacteria least susceptible to

killing, with results closely approximating those observed in mature biofilms.

Furthermore, resuspending biofilm bacteria into nutrient-rich media restored

antibiotic susceptibility, while resuspending in nutrient-poor media allowed the

bacteria to retain their resistance to both ciprofloxacin and ampicillin[66].

Relating the nutrient limitations to bacterial growth, K. pneumonia specific

growth rates within biofilms were found to be only 0.032 h-1, compared to 0.59 h-1 found in planktonic bacteria. Such low growth rates are indicative of stationary phase bacteria, which was confirmed by the presence of the enzyme catalase[66].

The authors hypothesize that after entering stationary phase due to oxygen and glucose limitations deep within the biofilm, the bacteria are protected from antibiotics as long as they remain in nutrient-poor conditions.

Oxygen limitation and its affect on P. aeruginosa was studied on biofilms after either 4 hours (1.14 h-1 specific growth rate) or 48 hours (0.02 h-1 specific growth

rate) of growth. Biofilms were challenged with multiple antibiotics, and the

results were compared to oxygen concentration readings taken with

microelectrodes. 48 hour biofilms were killed poorly by ciprofloxacin,

tobramycin, ceftazidime, chloramphenicol, carbenicillin, and tetracycline, with ciprofloxacin showing the highest reduction in viable bacteria (1.13 log reduction)[67]. Oxygen concentration measurements with microelectrodes on the

48 hour biofilms (210 m thick) indicated that oxygen only penetrated to a depth of 50 m, while an inducible green fluorescent protein (GFP) was only active in a

32 m layer within the oxygen rich region[67, 68]. Thus, approximately 75% of the bacteria within a P. aeruginosa biofilm reside in an inactive, anaerobic state[67]. The younger 4 hour biofilms showed GFP activity in all regions, and these bacteria were susceptible to all antibiotics tested. However, by changing to an anaerobic growth environment, the 4 hour biofilms quickly stopped growing, showed reduced GFP expression, and demonstrated reduced antibiotic killing[67]. These findings demonstrate that oxygen availability plays a crucial role in determining antibiotic susceptibility of P. aeruginosa, confirming similar findings from Walters et al[69].

3) Adaptive Response & Persister Cells

A third, and potentially most important, mechanism for bacterial survival in the presence of extreme antibiotic concentrations is due to an adaptive response which creates a small amount of persister cells within the biofilm. Joseph Bigger was the first to discover persister cells while studying ability of the new drug penicillin to kill Staphylococcus pyogenes[70]. After treating exponentially growing, planktonic S. pyogenes with penicillin, a small volume of media was transferred onto agar or into a tube of fresh broth. In all cases, a small number of

cells, termed persisters, had survived the penicillin treatment and were able to reestablish the infection. Interstingly, the new population of S. pyogenes was identical to the original population, consisting of a penicillin-sensitive bulk and a new population of persister cells. Multiple studies have yielded similar results, confirming that persister cells are phenotypic variants, rather than genetic mutants[70-72]. Keren et al recently repeated a similar study, confirming that a small population of E. coli, P. aeruginosa, or S. aureus persisters was able to completely regenerate the original population[72]. Furthermore, analysis of persister presence in relation to antibiotic response and growth phase revealed that the persister cells do not represent a particular stage of the bacterial growth cell cycle, and they are not formed in response to antibiotic treatment. Persister cells are not resistant to antibiotics, but instead represent a specialized cell able to tolerate and survive high antibiotic concentrations[72]. It is also believed that persister cell formation is dependent upon the level of metabolic activity within the population, rather than a quorum-sensing phenomenon[73]. Persisters are essentially cells that have exchanged rapid propagation for a non-growing life that ensures survival of the population. After a biofilm is exposed to high concentrations of antibiotics that kill both planktonic and metabolically active cells, the persister cells can then reenter the cell cycle and repopulate the biofilm[73].

Additional studies further confirm that persister sells are tolerant, rather than resistant, to extreme antibiotic concentrations. Ofloxacin, fluoroquinolone

antibiotic, acts on bacterial DNA and does not require the bacterial cell to be actively growing and dividing. Ofloxacin resulted in significant killing of both log-phase planktonic cells and established biofilm cells. However, 0.001% of the planktonic cells and 0.1% of the biofilm cells were identified as persister cells and

were unaffected by the antibiotic[74]. Tobramycin is yet another antibiotic that

acts on non-growing cells by binding to bacterial ribosomes and interfering with

protein synthesis. Once again persister cells were found to be highly tolerant to

tobramycin in both stationary-phase planktonic cells and established biofilm

cells[74].

In summary, multiple explanations exist to explain biofilm resistance to

antibiotic concentrations that typically kill planktonic suspensions. The species

and strain of bacteria, the location of the infection, and the antibiotic being used

for treatment all play significant roles in determining which resistance

mechanism(s) come in to play. In some cases, the antibiotic has difficulty

diffusing through the biofilm matrix, In other cases, oxygen and nutrient

gradients within the biofilm result in a heterogeneous population of cells, which

leads to regions of slow growing, metabolically inactive bacteria that are

unaffected by antibiotics which only act on growing cells. Finally, and likely

most significantly, a small population of bacteria alter their phenotype to become

persister cells. These persister cells are highly resistant to most antibiotics, and

even after >99% of a biofilm is eradicated by antibiotic treatment the persister

cells are able to resume growing and replenish the site with a population of cells identical to the original infection.

2.5 REFERENCES

1. Davies J and Davies D, Origins and Evolution of Antibiotic Resistance.

Microbiol. Mol. Biol. Rev., 2010. 74(3): p. 417‐433.

2. Morar M and Wright GD, The Genomic Enzymology of Antibiotic Resistance.

Annual Review of Genetics, 2010. 44(1): p. 25‐51.

3. Fischbach MA and Walsh CT, Antibiotics for emerging pathogens. Science,

2009. 325(5944): p. 1089.

4. Nathan C and Goldberg FM, The profit problem in antibiotic R&D. Nat Rev

Drug Discov, 2005. 4(11): p. 887‐891.

5. Oliphant CM and Green GM, Quinolones: a comprehensive review. American

family physician, 2002. 65(3): p. 455.

6. Hooper DC, New Uses for New and Old Quinolones and the Challenge of

Resistance. Clinical Infectious Diseases, 2000. 30(2): p. 243‐254.

7. Turnidge J, Pharmacokinetics and pharmacodynamics of fluoroquinolones.

Drugs, 1999. 58(S2): p. 29‐36.

8. Hooper DC, Mode of Action of Fluoroquinolones. Drugs, 1999. 58(S6): p. 6‐10.

9. Sköld O, Sulfonamide resistance: mechanisms and trends. Drug Resistance

Updates, 2000. 3(3): p. 155‐160.

10. Tipper DJ and Strominger JL, Mechanism of action of penicillins: a proposal

based on their structural similarity to acyl‐D‐alanyl‐D‐alanine. Proceedings of

the National Academy of Sciences, 1965. 54(4): p. 1133‐1141.

11. Waxman DJ and Strominger JL, Penicillin‐Binding Proteins and the

Mechanism of Action of Beta‐Lactam Antibiotics1. Annual Review of

Biochemistry, 1983. 52(1): p. 825‐869.

12. Hujer AM, Kania M, Gerken T, Anderson VE, Buynak JD, Ge X, Caspers P,

Page MGP, Rice LB, and Bonomo RA, Structure‐Activity Relationships of

Different {beta}‐Lactam Antibiotics against a Soluble Form of Enterococcus

faecium PBP5, a Type II Bacterial Transpeptidase. Antimicrob. Agents

Chemother., 2005. 49(2): p. 612‐618.

13. Blumberg PM and Strominger JL, Interaction of penicillin with the bacterial

cell: penicillin‐binding proteins and penicillin‐sensitive enzymes. Microbiology

and Molecular Biology Reviews, 1974. 38(3): p. 291.

14. McCormick MH, Stark WM, Pittenger GE, Pittenger RC, and McGuire JM,

Vancomycin, a new antibiotic. I: Chemical and Biologic Properties. Antibiotics

Annual, 1956. 1955‐56: p. 606‐611.

15. Barna JCJ and Williams DH, The structure and mode of action of glycopeptide

antibiotics of the vancomycin group. Annual Reviews in Microbiology, 1984.

38(1): p. 339‐357.

16. Pace JL and Yang G, Glycopeptides: update on an old successful antibiotic class.

Biochemical pharmacology, 2006. 71(7): p. 968‐980.

17. Kahne D, Leimkuhler C, Lu W, and Walsh C, Glycopeptide and

lipoglycopeptide antibiotics. Chemical reviews, 2005. 105(2): p. 425‐448.

18. Reynolds PE, Structure, biochemistry and mechanism of action of glycopeptide

antibiotics. European Journal of Clinical Microbiology & Infectious

Diseases, 1989. 8(11): p. 943‐950.

19. Watanakunakorn C, The antibacterial action of vancomycin. Rev Infect Dis,

1981. 3 suppl: p. S210‐5.

20. Arimoto H, Nishimura K, Kiniumi T, Hayakawa I, and Uemura D, Multi‐

valent polymer of vancomycin ‐ enhanced antibacterial activity against VRE.

Chem. Commun., 1999(15): p. 1361‐1362.

21. Beauregard DA, Williams DH, Gwynn MN, and Knowles DJ, Dimerization

and membrane anchors in extracellular targeting of vancomycin group

antibiotics. Antimicrob. Agents Chemother., 1995. 39(3): p. 781‐785.

22. Davies J and Wright GD, Bacterial resistance to aminoglycoside antibiotics.

Trends in Microbiology, 1997. 5(6): p. 234‐240.

23. Mingeot‐Leclercq M‐P, Glupczynski Y, and Tulkens PM, Aminoglycosides:

Activity and Resistance. Antimicrob. Agents Chemother., 1999. 43(4): p. 727‐

737.

100

24. Hermann T, Aminoglycoside antibiotics: old drugs and new therapeutic

approaches. Cellular and Molecular Life Sciences, 2007. 64(14): p. 1841‐1852.

25. Spizek J and Rezanka T, Lincomycin, clindamycin and their applications.

Applied Microbiology and Biotechnology, 2004. 64(4): p. 455‐464.

26. Leclercq R, Mechanisms of Resistance to Macrolides and Lincosamides: Nature

of the Resistance Elements and Their Clinical Implications. Clinical Infectious

Diseases, 2002. 34(4): p. 482‐492.

27. Tenson T, Lovmar M, and Ehrenberg M, The Mechanism of Action of

Macrolides, Lincosamides and Streptogramin B Reveals the Nascent Peptide Exit

Path in the Ribosome. Journal of Molecular Biology, 2003. 330(5): p. 1005‐

1014.

28. Mukhtar TA and Wright GD, Streptogramins, Oxazolidinones, and Other

Inhibitors of Bacterial Protein Synthesis. Chemical Reviews, 2005. 105(2): p.

529‐542.

29. Wilson DN, Schluenzen F, Harms JM, Starosta AL, Connell SR, and Fucini

P, The oxazolidinone antibiotics perturb the ribosomal peptidyl‐transferase center

and effect tRNA positioning. Proceedings of the National Academy of

Sciences, 2008. 105(36): p. 13339‐13344.

30. Shinabarger DL, Marotti KR, Murray RW, Lin AH, Melchior EP, Swaney

SM, Dunyak DS, Demyan WF, and Buysse JM, Mechanism of action of

101

oxazolidinones: effects of linezolid and eperezolid on translation reactions.

Antimicrob. Agents Chemother., 1997. 41(10): p. 2132‐2136.

31. Swaney SM, Aoki H, Ganoza MC, and Shinabarger DL, The Oxazolidinone

Linezolid Inhibits Initiation of Protein Synthesis in Bacteria. Antimicrob.

Agents Chemother., 1998. 42(12): p. 3251‐3255.

32. Aristoff PA, Garcia GA, Kirchhoff PD, and Hollis Showalter HD,

Rifamycins ‐ Obstacles and opportunities. Tuberculosis, 2010. 90(2): p. 94‐118.

33. Chen LF and Kaye D, Current use for old antibacterial agents: polymyxins,

rifamycins, and aminoglycosides. Infectious disease clinics of North America,

2009. 23(4): p. 1053‐1075.

34. Van Bambeke F, Glupczynski Y, Plesiat P, Pechere JC, and Tulkens PM,

Antibiotic efflux pumps in prokaryotic cells: occurrence, impact on resistance and

strategies for the future of antimicrobial therapy. Journal of Antimicrobial

Chemotherapy, 2003. 51(5): p. 1055.

35. Van Bambeke F, Balzi E, and Tulkens PM, Antibiotic efflux pumps.

Biochemical pharmacology, 2000. 60: p. 457‐470.

36. Poole K, Efflux‐mediated antimicrobial resistance. Journal of Antimicrobial

Chemotherapy, 2005. 56(1): p. 20.

37. Abraham E and Chain E, An enzyme from bacteria able to destroy penicillin.

Nature, 1940. 146(3713): p. 837‐837.

102

38. De Pascale G and Wright GD, Antibiotic resistance by enzyme inactivation:

from mechanisms to solutions. ChemBioChem, 2010. 11(10): p. 1325‐1334.

39. Wright GD, Molecular mechanisms of antibiotic resistance. Chem. Commun.,

2011.

40. Long KS, Poehlsgaard J, Kehrenberg C, Schwarz S, and Vester B, The Cfr

rRNA methyltransferase confers resistance to phenicols, lincosamides,

oxazolidinones, pleuromutilins, and streptogramin A antibiotics. Antimicrobial

Agents and Chemotherapy, 2006. 50(7): p. 2500.

41. Williams DH, The glycopeptide story‐‐how to kill the deadly ʹsuperbugsʹ. Nat

Prod Rep, 1996. 13(6): p. 469‐77.

42. Marshall C, Lessard I, Park IS, and Wright G, Glycopeptide antibiotic

resistance genes in glycopeptide‐producing organisms. Antimicrobial Agents

and Chemotherapy, 1998. 42(9): p. 2215‐2220.

43. Queck SY and Otto M, Staphylococcus epidermidis and other Coagulase‐

Negative Staphylococci, in Staphylococcus Molecular Genetics, J.A. Lindsay,

Editor. 2008, Caister Academic Press: Norfolk, UK.

44. Vuong C and Otto M, Staphylococcus epidermidis infections. Microbes Infect,

2002. 4(4): p. 481‐9.

103

45. Sabath LD, Garner C, Wilcox C, and Finland M, Susceptibility of

Staphylococcus aureus and Staphylococcus epidermidis to 65 antibiotics.

Antimicrob Agents Chemother, 1976. 9(6): p. 962‐9.

46. Cerca N, Jefferson KK, Maira‐Litran T, Pier DB, Kelly‐Quintos C,

Goldmann DA, Azeredo J, and Pier GB, Molecular basis for preferential

protective efficacy of antibodies directed to the poorly acetylated form of

staphylococcal poly‐N‐acetyl‐beta‐(1‐6)‐glucosaminev. Infection and

Immunity, 2007. 75(7): p. 3406‐3413.

47. Ceri H, Olson ME, Stremick C, Read RR, Morck D, and Buret A, The

Calgary Biofilm Device: New Technology for Rapid Determination of Antibiotic

Susceptibilities of Bacterial Biofilms. J. Clin. Microbiol., 1999. 37(6): p. 1771‐

1776.

48. Olson M, Ceri H, Morck D, Buret A, and Read R, Biofilm bacteria: formation

and comparative susceptibility to antibiotics. Can. J. Vet. Res., 2002. 66(2): p.

86‐92.

49. Costerton JW, Bacterial Biofilms: A Common Cause of Persistent Infections.

Science, 1999. 284(5418): p. 1318‐1322.

50. Stewart P and Williamcosterton J, Antibiotic resistance of bacteria in biofilms.

The Lancet, 2001. 358(9276): p. 135‐138.

104

51. Stewart PS, Mechanisms of antibiotic resistance in bacterial biofilms.

International Journal of Medical Microbiology, 2002. 292(2): p. 107‐113.

52. Stewart PS, Theoretical aspects of antibiotic diffusion into microbial biofilms.

Antimicrob Agents Chemother, 1996. 40(11): p. 2517‐22.

53. Rani SA, Pitts B, and Stewart PS, Rapid Diffusion of Fluorescent Tracers into

Staphylococcus epidermidis Biofilms Visualized by Time Lapse Microscopy.

Antimicrob. Agents Chemother., 2005. 49(2): p. 728‐732.

54. Farber B, Kaplan M, and Clogston A, Staphylococcus epidermidis Extracted

Slime Inhibits the Antimicrobial Action of Glycopeptide Antibiotics. The Journal

of Infections Diseases, 1990. 161(1): p. 37‐40.

55. Souli M and Giamarellou H, Effects of Slime Produced by Clinical Isolates of

Coagulase‐Negative Staphylococci on Activities of Various Antimicrobial Agents.

Antimicrob. Agents Chemother., 1998. 42(4): p. 939‐941.

56. König C, Schwank S, and Blaser J, Factors compromising antibiotic activity

against biofilms of Staphylococcus epidermidis. European Journal of Clinical

Microbiology & Infectious Diseases, 2001. 20(1): p. 20‐26.

57. Mathur T, Singhal S, Khan S, Upadhyay D, Fatma T, and Rattan A, Adverse

effect of staphylococci slime on in vitro activity of glycopeptides. Jpn J Infect Dis,

2005. 58(6): p. 353‐357.

105

58. Singh R, Ray P, Das A, and Sharma M, Penetration of antibiotics through

Staphylococcus aureus and Staphylococcus epidermidis biofilms. Journal of

Antimicrobial Chemotherapy, 2010. 65(9): p. 1955‐1958.

59. Dunne WM, Mason E, and Kaplan S, Diffusion of Rifampin and Vancomycin

through a Staphylococcus epidermidis Biofilm. Antimicrobial Agents and

Chemotherapy, 1993. 37(12): p. 2522‐2526.

60. Darouiche RO, Dhir A, Miller AJ, Landon GC, Raad, II, and Musher DM,

Vancomycin penetration into biofilm covering infected prostheses and effect on

bacteria. J Infect Dis, 1994. 170(3): p. 720‐3.

61. Jefferson KK, Goldmann DA, and Pier GB, Use of Confocal Microscopy To

Analyze the Rate of Vancomycin Penetration through Staphylococcus aureus

Biofilms. Antimicrob. Agents Chemother., 2005. 49(6): p. 2467‐2473.

62. Stewart PS, Davison WM, and Steenbergen JN, Daptomycin Rapidly

Penetrates a Staphylococcus epidermidis Biofilm. Antimicrob. Agents

Chemother., 2009. 53(8): p. 3505‐3507.

63. Stewart PS, Diffusion in Biofilms. J. Bacteriol., 2003. 185(5): p. 1485‐1491.

64. Brown MRW, Allison DG, and Gilbert P, Resistance of bacterial biofilms to

antibiotics a growth‐rate related effect? Journal of Antimicrobial

Chemotherapy, 1988. 22(6): p. 777‐780.

106

65. Xu KD, Stewart PS, Xia F, Huang C‐T, and McFeters GA, Spatial

Physiological Heterogeneity in Pseudomonas aeruginosa Biofilm Is Determined

by Oxygen Availability. Appl. Environ. Microbiol., 1998. 64(10): p. 4035‐

4039.

66. Anderl JN, Zahller J, Roe F, and Stewart PS, Role of Nutrient Limitation and

Stationary‐Phase Existence in Klebsiella pneumoniae Biofilm Resistance to

Ampicillin and Ciprofloxacin. Antimicrob. Agents Chemother., 2003. 47(4):

p. 1251‐1256.

67. Borriello G, Werner E, Roe F, Kim AM, Ehrlich GD, and Stewart PS,

Oxygen Limitation Contributes to Antibiotic Tolerance of Pseudomonas

aeruginosa in Biofilms. Antimicrob. Agents Chemother., 2004. 48(7): p. 2659‐

2664.

68. Werner E, Roe F, Bugnicourt A, Franklin MJ, Heydorn A, Molin S, Pitts B,

and Stewart PS, Stratified Growth in Pseudomonas aeruginosa Biofilms. Appl.

Environ. Microbiol., 2004. 70(10): p. 6188‐6196.

69. Walters MC, III, Roe F, Bugnicourt A, Franklin MJ, and Stewart PS,

Contributions of Antibiotic Penetration, Oxygen Limitation, and Low Metabolic

Activity to Tolerance of Pseudomonas aeruginosa Biofilms to Ciprofloxacin and

Tobramycin. Antimicrob. Agents Chemother., 2003. 47(1): p. 317‐323.

107

70. Bigger J, Treatment of Staphylococcal Infections with Penicillin by Intermittent

Sterilisation. The Lancet, 1944. 244(6320): p. 497‐500.

71. Balaban NQ, Merrin J, Chait R, Kowalik L, and Leibler S, Bacterial

persistence as a phenotypic switch. Science, 2004. 305(5690): p. 1622.

72. Keren I, Kaldalu N, Spoering A, Wang Y, and Lewis K, Persister cells and

tolerance to antimicrobials. FEMS Microbiology Letters, 2004. 230(1): p. 13‐

18.

73. Lewis K, Persister cells and the riddle of biofilm survival. Biochemistry

(Moscow), 2005. 70(2): p. 267‐274.

74. Spoering AL and Lewis K, Biofilms and planktonic cells of Pseudomonas

aeruginosa have similar resistance to killing by antimicrobials. Journal of

bacteriology, 2001. 183(23): p. 6746.

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CHAPTER 3

STRATEGIES FOR PREVENTING AND TREATING DEVICE INFECTIONS

Traditional antibiotic treatment regimes have proven to have significant

limitations when it comes to treating infections associated with biofilms and

implanted medical devices. Bacteria have the ability to rapidly develop antibiotic

resistance through a variety of mechanisms. As a result, a great deal of recent

research has centered on the concept of non-traditional treatments, including

surface modifications to prevent adhesion, drug-eluting devices, surface-grafted

antibiotics, and immunotherapy. This chapter will provide an overview of the current strategies under active development for preventing and treating material-based infections, with relevant examples pertaining to S. epidermidis.

3.1 MODIFICATIONS OF THE BIOMATERIAL

3.1.1 SURFACE MODIFICATIONS OF MATERIALS TO PREVENT ADHESION

As a result of increased resistance to antibiotics, managing infections through the prevention of initial adhesion to biomaterials has taken on a prominent role.

It is well known that factors such as surface charge and hydrophobicity play a

key role in nonspecific adhesion to bare surfaces[1]. In addition, multiple

bacterial surface components interact with biomaterial substrates after a

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conditioning layer of host proteins is adsorbed. Therefore, modifications of

biomaterials in order to alter surface chemistry and protein adsorption behavior

have been widely studied as a means for preventing bacterial adhesion.

One of the simplest strategies for altering surface chemistry has been to

position poly(ethylene glycol) (PEG) chains on the material surface. An early

study utilized a surface physical interpenetrating network (SPIN) to modify

poly(ethylene terephthalate) (PET) surfaces, wherein the PET surface was

swollen and PEG (MW 18,500) diffused into the material. The material was then

deswelled in a rapid manner, trapping PEG on the surface of the PET substrates.

Results of this study indicated that initial adhesion of Staphylococcus epidermidis,

Staphylococcus aureus, and Pseudomonas aeruginosa to PET was reduced by 70-95%

due to the presence of PEG chains[2]. However, the authors observed that

fibrinogen absorbed readily to the PEG-coated surfaces, and when present,

greatly increased the amount of initial adhesion. Therefore, it was suggested that

such PEG coatings may be more appropriate when used for short term and/or non-blood contacting devices.

Another study devised a means for attaching PEG chains to stainless steel

substrates in order to examine changes in bacterial adhesion[3]. Poly(ethylene

imine) (PEI) was first adsorbed onto stainless steel surfaces, followed by grafting

of PEG (MW 5,000) to the positively charged amines. These surfaces were able to

prevent adsorption of the model protein -lactoglobulin, although other proteins

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were not tested. However, despite their ability to repel proteins, the surfaces had no effect on adherence of gram-positive (Listeria monocytogenes) or gram-negative

(Pseudomonas sp.) organisms[3]. The authors speculate that the growth media itself may provide enough protein adsorption to facilitate bacterial adhesion, or that the bacteria may possess the ability to physically remove the surface- adsorbed PEI-PEG coating. Thus, covalent linkage to the surface may be necessary in order to effectively prevent surface adhesion. In addition, this study only examined PEG with a molecular weight of 5,000 Da. Previous studies have yielded somewhat conflicting results, indicating that higher molecular weight

PEG is necessary to prevent bacterial adhesion[2]. Still other studies show that extremely short hexa(ethylene glycol) monolayers were sufficient to prevent S. epidermidis adhesion[4]. In the case of stainless steel, it is likely that short PEG chains may be insufficient at shielding the surface from colonizing bacteria.

Polymer brush coatings are yet another strategy employed for creating non- adhesive surfaces. In order to create such brushes, polymer chains are attached to the surface with a high enough density to force the chains to extend away from the surface, thereby shielding the surface from potential adherent cells or proteins. While PEG brushes are fairly well established in their ability to repel protein adsorption[5], they are only recently being characterized regarding their ability to prevent bacterial adhesion. Poly(ethylene glycol)-poly(propylene oxide)-poly(ethylene glycol) triblock copolymers (PEG-PPO-PEG), which are

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readily absorbed onto hydrophobic biomaterials, have been used to create PEG

brush coatings and studied for their ability to prevent surface adhesion of

various bacterial species. A dip-coating method has been used to create a PEG-

PPO-PEG brush coating on poly(styrene), and these surfaces demonstrated an

order of magnitude reduction in Staphylococcus epidermidis and Serratia marcescens

adhesion over 24 hours when compared to uncoated poly(styrene)[6]. Another

study covalently grafted PEG brushes to silanized glass surfaces by using

methacryl-terminated PEG (MW 9,800) and measured the surface adhesion

properties for 5 bacterial strains (S. epidermidis HBH 276, S. aureus ATCC 12600,

Streptococcus salivarius GB 24/9, E. coli O2K2, and P. aeruginosa AK1) and 2 yeast strains (C. albicans GB 1/2 and C. tropicalis GB 9/9)[7]. After 4 hours of exposure to the various organisms, initial adhesion was significantly lower for S. epidermidis, S. aureus, S. salivarius, and E. coli (95-98% reduction) when compared to bare glass substrates. Interestingly, P. aeruginosa, C. albicans, and C. tropicalis exhibited a smaller 70% reduction in adhesion, which the authors attributed to the increased hydrophobicity of these organisms, and in the case of the two yeast species, the increased size[7].

While most studies demonstrate a significant reduction in initial bacterial adhesion to polymer brush surfaces, few go on to examine the fate of the bacteria that actually adhere to the surface. After demonstrating that brushes formed on silicone by the adsorption of PEG-PPO-PEG tri-block copolymers are able to

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reduce initial adhesion of S. epidermidis and S. aureus by an order of magnitude,

Nejadnik et al showed that even the relatively few bacteria that did adhere were

sufficient to lead to the formation of a mature biofilm[8]. However, these mature

biofilms differed from those formed on bare silicone, in that they formed more

slowly and they were more easily detachable from the surface. 100% of S.

epidermidis and S. aureus biofilms grown on brushed surfaces were removed by 4

Pa of shear stress, whereas detachment from bare silicone surfaces was just

71±47% for S. epidermidis biofilms and 58±20% for S. aureus. In addition, a significantly larger portion of the bacteria present in the biofilms formed on brushed surfaces were viable for both S. epidermidis (99±0%) and S. aureus

(81±5%) when compared to those formed on bare silicone (20-25% viable)[8]. The

implications of altering the viability of adherent bacteria remain to be seen.

Recent studies have also investigated the use of atom transfer radical

polymerization (ATRP) to grow poly(acrylamide) PAAm polymer brush chains

directly on silicon wafers[9]. Growing brushes directly from the surface allows for much greater control of chain length and density, in contrast to most other methods wherein existing polymer chains are grafted or adsorbed onto a surface.

The PAAm brushes were studied for their effectiveness in preventing S. aureus,

S. salivarius, and C. albicans adhesion over a 4 hour time period in a parallel plate

flow chamber. All three organisms demonstrated slower initial deposition rates

on the PAAm brush surfaces when compared to bare silicon. In addition, after 4

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hours of flow, S. aureus showed a 92±4% reduction in adhesion, S. salivarius

showed a 70±9 reduction, and C. albicans showed a 83±7% reduction.

Furthermore, the authors concluded that S. aureus and S. salivarius adhesion

forces to the brushed surface were much lower, as 65-87% of the adherent

bacteria were detached when a large bubble was introduced into the flow system

(compared to 11-17% detachment from the bare silicon surface)[10].

Finally, the use of zwitterionic surfaces has grown increasingly popular due

to their simplicity and effectiveness. The rationale for the use of zwitterionic

surfaces is derived from the knowledge that the outer lipid membrane of

erythrocytes is non-thrombogenic, and that the electrically neutral

phosphorylcholine moiety was largely responsible for this

haemocompatibility[11]. Since this discovery, phosphorylcholine (PC) groups have been incorporated into a wide variety of systems in order to improve biocompatibility. Such procedures have been termed ‘biomembrane mimicry’ and have been used to prevent protein, cellular, and bacterial interactions with biomaterial surfaces[12]. 2-methacryloyloxyethyl phosophorylcholine (MPC) has been used to coat PET coverslips, significantly reducing the number of S. aureus,

Streptococcus mutans, P. aeruginosa, and C. albicans adhering to the surfaces after a

1 hour static incubation when compared to bare PET[13]. In order to study the effects of MPC surfaces on biofilm formation, Fujii et al [14] coated stainless steel plates with poly(MPC-co-n-butyl methacrylate) (PMB) and exposed the surfaces

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to S. aureus, S. epidermidis, and P. aeruginosa for 48 hours. Bacteria were then

stained with ethidium bromide for nucleic acid staining and with a FITC- conjugated concanavalin A solution for polysaccharide labeling. Confocal laser

scanning microscopy (CLSM) then revealed that the coated surfaces were unable

to support biofilm growth for any of the strains tested, whereas the bare stainless steel supported growth of mature biofilms. In addition, an ATP assay was used to assess total adherent bacteria with and without the addition of antiobitics. The coatings alone resulted in a 20-fold (S. aureus) to 50-fold (S. epidermidis) reduction in adherent bacteria, whereas the addition of cefazolin or gentamicin resulted in a 2000-fold (S. aureus) and 1200-fold (S. epidermidis) when compared to uncoated surfaces[14]. This indicates that phosphorylcholine coatings are not only able to prevent the formation of mature biofilms, but they can also render surface- adherent bacteria more susceptible to antibiotics. Most recently, alternative zwitterionic polymers such as poly(sulfobetaine methacrylate) (pSBMA) surface grafted onto gold substrates by ATRP have been shown to possess excellent resistance to protein and bacterial adhesion[15]. The pSBMA-brushed surfaces were exposed to S. epidermidis and P. aeruginosa in a parallel plate flow chamber for 3 hour, 24 hour, and 48 hour adhesion/growth studies. Both species showed significant decreases in adhesion to pSBMA at all time points, indicating that the highly hydrated surfaces are able to interfere with the interactions necessary for adhesion. Significant reductions in non-specific fibrinogen adhesion to the

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pSBMA surfaces were also demonstrated, offering another possible explanation

for the lack of bacterial adhesion[15].

3.1.2 INCORPORATION OF ANTIMICROBIAL AGENTS

While PEG and zwitterionic coatings of biomaterials have proven effective at

preventing adhesion and biofilm growth, these materials are still unable to

ensure a 100% elimination of adhesion. Therefore, it is necessary to explore the

use of a more active means of eliminating adherent bacteria: incorporating antimicrobial agents into biomaterials.

One strategy that has been widely studied for the prevention of bacterial

colonization is the release of antibiotics from coatings applied to the biomaterial

or from directly impregnating the material. Such site-specific delivery of

antibiotics offers several advantages over systemic delivery, including localized

higher concentrations that avoid the issues associated with systemic toxicity.

However, kinetic issues come in to play with this type of strategy, wherein the

rate of release must be carefully controlled in order to ensure therapeutic levels

of the antibiotic. Too high of an antibiotic concentration could result in toxicity to

the surrounding tissue, while too low of a concentration could result in the

development of antibiotic resistance. In addition, the antibiotic supply within the

device or coating will eventually be depleted, at which point the material will no

longer be protected from microorganisms[16]. The potential solution to these

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problems is to covalently graft antibiotic compounds directly to the biomaterial surface.

Silver-Releasing Coatings

Silver has been utilized for its antibacterial properties for thousands of years.

Although inert in its metallic state, silver becomes highly active when ionized in fluids and is able to bind to bacterial cell walls and membranes (interfering with

cell respiration). Silver ions are also capable of binding and denaturing bacterial

DNA and RNA (interfering with reproduction)[17]. In addition, it is relatively

uncommon for bacteria to develop any sort of resistance towards silver ions.

Directly coating devices with a layer of silver oxide or silver alloy has been

explored and such devices have found their way into clinical settings. However,

silver ions in direct coatings are easily bound by albumin or precipitated as AgCl when bound by chloride ions. Furthermore, most clinical studies suggesting

successful application of silver-coated devices consist of a small study

population, and one of the few large-scale studies has indicated that metallic

silver coatings fail to improve antibacterial activity[18, 19]. Recent advances in nanotechnology have enabled scientists to re-explore the use of silver as an

antibiotic. The high surface-to-volume ratio of nanoparticles yields completely new chemical and physical properties for the silver, with most current research focusing on either bulk impregnation of biomaterials with silver nanoparticles or exploring the use of silver-releasing polymeric coatings.

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Direct impregnation of bulk silicone with silver nanoparticles was recently

studied by Furno et al[20]. Organometallic precursors, dissolved in supercritical

carbon dioxide, were used to swell and subsequently permeate silicone discs at

4000 psi for 24 hours at 40°C. The discs were then exposed to hydrogen gas in order to decompose the organometallic precursors into uniformly dispersed silver nanoparticles 10-100 nm in diameter. It was found that the discs had a significant amount of antibacterial effect on both adherent and planktonic

bacteria, but these effects were likely due to a ‘burst’ release of nanoparticles that

were located close to the surface. This burst effect could be eliminated by simply

washing the discs prior to use. However, even after washing, there was a

residual, slow release of silver nanoparticles from deep within the silicone,

allowing the discs to retain their ability to kill adherent S. epidermidis[20].

The majority of research regarding silver nanoparticles as antibacterial agents involves silver-impregnated coatings, rather than bulk impregnation of the biomaterial itself. Silver-doped phenyltriethoxysilane (PhTEOS) sol-gel coatings of glass slides have been used to demonstrate inhibition of S. epidermidis biofilm growth and toxicity towards planktonic S. epidermidis[21]. The silver- impregnated sol-gel surfaces were shown to provide a large initial burst of silver ions, followed by a slow, sustained release over a 10 day period. Biofilms were unable to form on the silver-impregnated sol-gel, whereas the undoped sol-gel surfaces supported robust biofilm development. In addition, nearly 100% of

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planktonic S. epidermidis could be killed by exposure to the silver-impregnated

PhTEOS sol-gel coatings[21]. Other common methods for preparing silver nanoparticle coatings include layer-by-layer [22], ion reduction and chemical vapor deposition (CVD)[23], and hydrogel-based designs[24].

One particularly elegant design involves a polymer coating that is loaded with silver nanoparticles and subsequently surface-modified with PEG. Thus, this strategy incorporates both anti-adhesion and anti-bacterial characteristics[25]. Poly(ethylene imine) (PEI) was reacted with methacryloyl chloride to introduce polymerizeable double bonds, and this solution was thereafter polymerized with 2-hydroxyethyl acrylate (HEA) on trimethyoxymethacryloxypropyl silane modified glass slides to form surface- adherent films approximately 20 m thick. The polymer films were then loaded with silver nanoparticles by chemical reduction of silver ions within the polymer network. Finally, PEG chains were tethered to the free hydroxyl groups contained within the polymerized film. After incubation in the presence of S. aureus for 12 hours, significant numbers of adherent colonies were able to grow on PEI films lacking PEG and silver. Introduction of the PEG chains (but no silver) resulted in a 4- to 5-fold reduction in adherent colonies, demonstrating the anti-adhesive nature of the PEG. Meanwhile, Silver-impregnated, PEG- containing films showed 100% elimination of colony formation[25]. However, even in the presence of silver, longer-term incubations eventually resulted in

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bacterial growth. This indicates that these coatings are most likely bacteriostatic as opposed to bactericidal.

Despite their promising results in vitro, silver-containing biomaterials have potential drawbacks. At high concentrations such as those found adjacent to silver-releasing implants, silver nanoparticles have the potential to become toxic to host tissue. While most studies report that silver nanoparticles are nontoxic under specific conditions, very little in vivo data is available. One particular study reported toxicity towards rat liver cells, wherein mitochondrial cells became abnormally sized, displaying cellular shrinkage and irregular shape.

Another study demonstrated cytotoxic effects on keratinocytes and fibroblast cultures exposed to commercially available silver-containing wound dressings[16, 17]. It is clear that more research needs to be done in regards to the body’s tolerance towards silver nanoparticles.

Antibiotic-Releasing Coatings

Whereas systemic delivery of antibiotics is limited due to toxic effects at high concentrations to various host tissues and organs, localized delivery at the site of infection allows for much higher antibiotic doses to be used. Ideally, a device or device coating would be able to release a large, fast initial burst of antibiotics in the first 6 hours in order to prevent initial colonization of the implant. Thereafter, it is desirable for the device to maintain a slow release of antibiotics in order to

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maintain the coating’s prophylactic nature and protect the surface from future

infections[16].

As was the case with silver-releasing materials, silicone makes an ideal

candidate for direct incorporation of antibiotics. Silicone catheters were impregnated with a combination of chlorhexidine and triclosan and tested for their ability to prevent bacterial colonization in an in vitro urinary tract model[26]. The impregnated catheters successfully prevented colonization over a

20-30 day period by K. pneumonia, E. aerogenes, P. mirabilis, methicillin resistant S. aureus (MRSA), vancomycin-resistant E. faecium, E. faecalis, E. coli, and C. albicans.

Meanwhile, uncoated catheters were infected and colonized within 1-2 days.

Earlier studies by the same group had revealed that silicone catheters impregnated with chlorhexidine, silver sulfadiazine, and triclosan were able to prevent colonization over a 15-24 day period by S. epidermidis, S. aureus, E. coli, and C. albicans[27].

A much broader area of research revolves around the concept of biomaterial coatings capable of releasing entrapped antibiotics with a specific rate and duration. Dacron surfaces (PET) have been coated with polymerized cyclodextrins (CDs) and their toxicity towards human epithelial cells was assessed over the course of 6 days[28]. Results indicated that the CD coated surfaces were not toxic towards the cells, and similar results were obtained with the coated and uncoated surfaces. Structurally, cyclodextrins are cyclic

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oligosaccharides with a hydrophilic exterior and a hydrophobic internal cavity.

Thus, the CDs are able to capture various molecules in their core and slowly

release them over time. Once it was confirmed that the surfaces were not harmful

to human cells, the surfaces were loaded with vancomycin by submersing in a 5 g/L solution. The samples were then rinsed and placed in physiologic medium

for 80 days. Quantification of vancomycin in the media indicated a linear release

during the first 50 days at a rate of 0.13 to 0.3 mg/L/day, depending on the

amount of CDs contained on the surfaces (5%-wt and 10%-wt, respectively).

Uncoated surfaces released 100% of their vancomycin within just 1 hour[28].

Thus, extended release of vancomycin was achieved through the use of a cyclodextrin coating on PET substrates. However, the Dacron surfaces became more hydrophobic upon modification[29], which could potentially increase the propensity for bacteria to adhere to the surface. These surfaces have not yet been tested in vitro for microbial adhesion or growth.

Prabu et al [30] constructed biodegradable scaffolds from collagen and poly(caprolactone) (PCL), and subsequently incorporated either amikacin or

gentamycin. Scaffolds demonstrated an initial burst release of both drugs over

the first 4 hours, followed by a sustained release over 48 hours. However,

concentrations of gentamicin were generally much lower than those of amikacin, which the authors attributed to a strong interaction between gentamicin and

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PCL. The authors envision such a scaffold could be applied to wound dressing in order to provide sustained release of antibiotics.

Biomaterial-Grafted Antibiotics

The most likely problem to arise with drug-releasing biomaterials, regardless of whether the biomaterial is directly impregnated or modified with a drug- releasing surface coating, is that eventually the drug reservoir will be exhausted and the material will lose its antibacterial properties. In order to solve this problem, the obvious solution is to graft the antibacterial agents directly to the surface of the material. A wide variety of strategies exist attempting to produce a

‘permanent’ bactericidal surface.

Perhaps analogous to silver nanoparticles in the drug-releasing category, surface-bound quaternary ammonium compounds (QACs) are a widely studied solution that falls outside the realm of traditional antibiotics. QACs are permanently charged cationic molecules well known for their bactericidal properties against both gram-negative and gram-positive bacteria when utilized in surface-bound applications. While it is commonly believed that the mode of action for QACs is the insertion of their long tails into the bacterial membrane,

Kugler et al [31] have recently demonstrated that it may instead be a high density of positive charges interacting with the bacteria that causes cell death. For static

(ie non-dividing) bacterial populations, a charge density of 1014 N+ cm-2 was

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sufficient to kill both S. epidermidis and E. coli. Meanwhile, for rapidly dividing

13 + -2 populations, a charge density of 10 N cm was sufficient to kill S. epidermidis, and a charge density of 1012 N+ cm-2 was sufficient to kill E. coli. The authors propose that by adsorbing onto the highly charged surface, divalent counterions are removed from the bacteria, resulting in disruption of the bacterial envelope[31]. Thus, highly charged cationic surfaces may be a viable substitute for antibiotics, especially in regards to antibiotic-resistant strains.

As an example of QACs utilized in biomaterial applications, 3-

(trimethoxysilyl)-propyldimethyloctadecylammonium chloride (QAS) coatings were covalently attached to silicone discs and studied both in vitro and in vivo for

their antibacterial properties against S. epidermidis, S. aureus, E. coli, and P. aeruginosa[32]. After 1 hour in a parallel plate flow chamber, bacteria adhering to the uncoated silicone rubber were 82-97% viable, while bacteria adhering to the

QAS-coated silicone were 0-37% viable, depending on the organism. For in vivo studies with pre-seeded implants, the QAS-coated silicone developed an infection with just 1 out of 8 implants, whereas the uncoated silicone implants were infected 7 out of 8 times[32]. As is the case with silver nanoparticles, concerns do exist regarding QACs, since the mode of action is not selective towards prokaryotic organisms. It is very possible that neighboring tissue could be damaged by the same mechanisms involved in killing invading bacteria.

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Covalent attachment of more traditional antibiotics to biomaterial surfaces is

perhaps the most promising method for providing permanent protection against

bacterial colonization and biofilm formation on implanted biomaterials.

However, care must be taken to ensure that the function, mechanism of action,

and activity of the antibiotics is not compromised during the grafting process.

Jose et al [33] examined the possibility of covalently tethering vancomycin to a

titanium surface, with potential applications in orthopaedic and dental implants.

Titanium has a Ti-OH surface layer that can be readily silanized with 3-

aminopropyltriethoxysilane (APTS). After silanization, vancomycin was tethered

to the titanium surface utilizing solid phase peptide synthesis techniques,

attaching the vancomycin by way of its C-terminal carboxylic acid. Two flexible

8-amino-3,6,dioxaoctanoate-acetate (AEEA) linkers (~40 Å total) were included

between the antibiotic and titanium-APTS surface in order to increase mobility of the antibiotic, allowing it to enter the cell well and bind to its -D-Ala-D-Ala

target. Indirect immunofluorescence was used to confirm surface grafting, and

results indicated that the functionalized titanium was stable for at least 3 weeks

at room temperature in Dulbecco’s modified essential medium (DMEM)

containing 10% fetal bovine serum. Radiolabeled peptidoglycans containing L-

Lys-D-(14C)Ala-D--(14C)Ala were used to confirm that surface grafted

vancomycin retained its binding activity. Vancomycin-grafted titanium particles

were incubated with S. aureus for 2 hours, reducing colony formation by 88±16%

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when compared to bare titanium. In addition, most of the surface adherent

bacteria were dead on the grafted surfaces, while most bacteria were alive on the

bare titanium.

In yet another approach to coat a titanium surface with antibiotics, a

derivatized form of vancomycin was covalently tethered to a Ti-6Al-4V alloy[34].

First, vancomycin was reacted with N-hydroxysuccinimide-PEG(3400)-acrylate

to attach a polymerizable acrylate group to the vancomycin V3 postion by way of

a flexible PEG spacer. The MIC (11.5±0.8 g/ml) and MBC (15±1 g/ml) values of this newly synthesized VPA monomer were determined by the broth dilution method. Although less active than the native vancomycin molecule

(MIC=MBC=1.7±0.06 g/ml), it was speculated that this difference may not be of clinical importance, as both MIC and MBC are solution-based parameters that don’t translate directly to surface-bound scenarios. Oxidized Ti-6Al-4V was then silanized with methacryloxypropyltrimethoxysilane, providing surface-grafted polymerizable methacrylate groups. Finally, the VPA monomer solution was photopolymerized on the methacrylate-functionalized titanium surfaces, resulting in a covalently bound coating containing active vancomycin molecules.

The VPA-coated titanium discs were able to reduce the level of viable S.

epidermidis adhering to the surfaces by approximately 5-fold after just 4 hours

when compared to uncoated Ti-6Al-4V. Furthermore, after a 17 hour incubation period, the VPA-coated surfaces reduced adherent S. epidermidis to undetectable

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levels while the uncoated surfaces supported significant numbers of bacteria.

The same research group obtained similar results when vancomycin-PEG(X)- acrylate (VPA(X)) and vancomycin-acrylamide derivatives were synthesized and photografted to dithiocarbamate (DTC) functionalized substrates[35].

3.2 SPECIFIC TARGETING STRATEGIES

3.2.1 BACTERIOPHAGES

First discovered in the 1920’s, bacteriophages represent a class of naturally occurring, targeted antibacterial treatments. Lytic phages are viruses that infect and kill microorganisms, replicating within the targeted bacteria and leading to a very high concentration of phages at the site of the infection. Despite initial results indicating phages had little value in treating infections in vivo and the rapid rise in popularity and success of antibiotic treatments, further work has revealed that phage treatment of E. coli infections in vivo can be more effective than treatments with antibiotics[36]. Mice were treated with lethal intramuscular inoculations of E. coli (3x107 viable organisms) in their left gastrocnemius muscle, followed by injections in the right gastrocnemius muscle with either a single dose of 3x108 viable anti-K1 phage particles or doses of 25 mg/kg of antibiotics

(tetracycline, ampicillin, chloramphenicol, trimethoprim plus sulphafurazole, or streptomycin). Results indicated that introducing phage into the right leg led to a

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rapid elimination of viable E. coli in the left leg muscle. Phage was at least as effective at preventing death as multiple doses of streptomycin, and more effective than multiple doses of all other drugs tested. In addition, due to its self- perpetuating nature in the presence of bacteria, even a small intravascular dose was typically sufficient to prevent the mouse from dying[36].

In order to study the effectiveness of bacteriophages at preventing catheter infection, an in vitro study was carried out wherein hydrogel-coated Foley catheters were pretreated with phage prior to exposure to S. epidermidis[37].

Catheters were soaked for 1 hour in Mueller-Hinton broth (MHB) containing

1x1010 to 2.2x1010 viable phages/ml (phage 456), after which the S. epidermidis in

MHB (108-109 cfu/ml) was pumped through the catheter for 2 hours. Finally,

sterile MHB was pumped through the catheters for 22 hours before enumerating

the number of viable adherent organisms by the viable count method. Results

indicated that 24 hour biofilms grown on untreated catheters supported

7.01±0.47 log cfu/cm2 of catheter. However, pretreatment of the catheters

resulted in up to a 4.47 log reduction of viable adherent S. epidermidis[37]. SEM analysis confirmed that phage-treated catheters were unable to support biofilm growth.

Cerca et al [38] investigated the influence of S. epidermidis phenotype on the effects of the polyvalent antistaphylococcus bacteriophage K. Eleven different clinical strains of S. epidermidis were tested under three phenotypic conditions:

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biofilm cells, stationary phase planktonic cells, and log phase planktonic cells.

Only 1 of the 11 strains tested failed to be susceptible to phage K when challenged in a planktonic culture. Most strains were also susceptible to infection by phage K when grown as biofilms, with a significant reduction in biomass noticeable after 21 hours. Interestingly, log phase planktonic cells were more susceptible to killing than either stationary phase planktonic cells or biofilm cells, indicating that the growth state of the bacteria is important when it comes to treatments with bacteriophages.

While phages have been identified that can infect various organisms, dissolve the exopolysaccharide matrix of existing biofilms, and even reduce the need for clinical use of antibiotics (and subsequent antibiotic resistance)[39], there are several drawbacks to utilizing the phage approach to manage device related infections. First, phages are very specific when it comes to which organism(s) they will target and infect. A phage may target a certain strain, multiple related strains, or even a few species. Most of this specificity is due to the structural differences in bacteria, with gram-negative phages targeting lipopolysaccharides and outer membrane proteins, and gram-positive phages targeting peptidoglycan, teichoic acids, and lipoteichoic acids[40]. Second, since phages attack their target by binding to a receptor on the bacterial surface, there is the chance that the bacteria could alter the target receptor and attain phage resistance (much the same as they do with antibiotics)[39]. Finally, when it comes

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to in vivo clinical usage, phages are considered antigens and will be treated

accordingly by the host immune system. It has long been known that phage will

be cleared from the blood stream, and anti-phage antibodies will be present in

the host serum after treatment[41]. Further studies are necessary to ensure the safety and successful application of bacteriophages to treat infections of implanted biomaterials.

3.2.2 ANTIBODIES AND OPSONIZATION

One of the more intriguing concepts for the prevention and treatment of

biofilms is the use of antibodies as opsonizing agents. Rennermalm et al [42]

have demonstrated that S. epidermidis opsonized with anti-SdrG antibodies are

ingested and killed much more effectively by rat macrophages in vitro than nonopsonized bacteria. Rat macrophages were attached to the bottom of 24-well plates, and 107 S. epidermidis cells were opsonized and added for 2 hours. After

rinsing away extracellular bacteria, it was shown that macrophages ingested ~10

opsonized bacteria per macrophage, and just 0.01 nonopsonized bacteria per

macrophage. In addition, only 1% of ingested bacteria were able to survive inside

the macrophage for longer than 18 hours. Finally, opsonized and nonopsonized

S. epidermidis were given intravenously to mice, and bacterial numbers were

measured in the kidneys after 4 days[42]. Preopsonized bacteria resulted in

3.2x104 CFU in the kidneys, compared to 1.3x105 CFU with nonopsonized bacteria. These results indicate that antibodies directed against SdrG have great 130

potential for protecting against S. epidermidis infections, with potential

applications in vaccine development.

Polyclonal IgG antibodies specific to clumping factor A (ClfA) and SdrG have

been studied extensively for their ability to prevent and treat both S. epidermidis

and S. aureus infections[43]. The bacterial cells were first labeled fluorescently

with 5(6)-FAM, after which they were exposed to the polyclonal IgG mixture for

30 minutes. Phagocytosis by murine macrophages was then monitored by flow

cytometery, with an increase in fluorescence intensity of the macrophages

indicating increased phagocytosis. Opsonization with the ClfA/SdrG specific

antibodies significantly increased phagocytosis of both S. epidermidis and S.

aureus in a concentration dependant manner when compared to bacteria opsonized with nonspecific IgG. Next, the antibodies were tested for their ability

to protect against infection in animals with an immature immune system.

Neonatal rats were injected with antibodies, followed 20 hours later by injection

with S. epidermidis. Within 3 days, 100% of the animals receiving the nonspecific

IgG antibodies had died, whereas 72% of the animals receiving the ClfA/SdrG

antibodies survived a full 7 days as a result of the single antibody injection.

Finally, a rabbit cardiovascular catheter model was examined. Prophylactic doses

of antibodies were given 24 hours prior to injection with sub-lethal doses of

methicillin resistant S. epidermidis (MRSE) and S. aureus (MRSA). Animals

receiving the ClfA/SdrG specific antibodies developed infections just 27%

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(MRSE) and 7% (MRSA) of the time, compared to a 63% infection rate with

nonspecific IgG. Finally, the polyclonal antibodies were used to treat established

infections on the vascular catheters. Treatments with 300 mg/kg ClfA/SdrG

antibodies significantly reduced MRSE or MRSA levels in the blood, kidney, and

cardiac valves, as well as overall bacteremia when compared to control

treatments[43]. Recently, these antibodies (known as INH-A21) were studied in

phase II clinical trials for the prevention of nosocomial infections in low-birth-

weight infants. Initial results indicate that a dose of 750 mg/kg had the ability to

reduce sepsis and mortality cause by S. aureus[44]. However, the INH-A21 drug

(Veronate) failed to meet its target endpoints in phase III trials[45].

As demonstrated above, favorable in vitro results have led to the pursuit of

developing antibody therapies for human use. However, significant work still remains, as indicated by the failure and/or postponement of various clinical trials. The fact remains that bacteria targeted by antibodies may adapt and alter their targeted surface molecules so as to develop resistance. A potential strength of antibody-mediated treatments, however, is that new monoclonal or polyclonal

mixtures could be produced using the already established procedure in order to

deal with resistant strains.

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3.3 SPECIFIC AIMS AND HYPOTHESIS

This project focuses on the development of novel methods for the treatment of Staphylococcus epidermidis infections. S. epidermidis is known to adhere to intravascular biomaterials through a number of mechanisms, including a specific binding event between the S. epidermidis surface protein SdrG and the amino acid sequence NEEGFFSARGHRPLD (6-20) of the human plasma protein fibrinogen.

The hypothesis of this work is that we can engineer a robust biofilm targeting system based on the 6-20 peptide, and utilize this system for targeted biofilm disruption and drug delivery. Through the inhibition of biofilm formation, we hope to demonstrate new approaches for improving the effectiveness of antimicrobial treatment. Specifically, the aims are to:

Aim 1: Develop a fibrinogen-derived 6-20 peptide for specific targeting and delivery to surface-adherent S. epidermidis. The 6-20 peptide

(NEEGFFSARGHRPLD) will be studied for its suitability as a targeting motif and its ability to deliver small particles to the bacterial surface. Nanogold particles will be covalently coupled to the amino terminus of6-20 peptide, allowing for direct visualization of peptide binding events by scanning electron microscopy (SEM). A series of unlabeled peptides of varying binding affinity

(NEEGFFSARGHRPLD, NEEGFFSARAHRPLD, NEEGFFSAAGHRPLD) will be used in conjunction with the Nanogold labeled peptide to demonstrate and quantify the specificity of the 6-20 peptide towards S. epidermidis.

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Aim 2: Engineer a cationic 6-20-G3K6 peptide capable of disrupting biofilm formation in vitro. A positively charged peptide (G3K6) and will be coupled to the 6-20 targeting peptide. This 6-20-G3K6 combination peptide will bind to

S. epidermidis through its 6-20 region, tethering the positively charged lysine residues to the bacterial surface. The 6-20-G3K6 will work to reduce adhesion of the positively charged PIA to the negatively charged bacteria, which in turn will inhibit biofilm formation on relevant cardiovascular biomaterials. Biofilm formation after exposure to the peptide will be monitored, with adherent bacteria and PIA content within the biofilms quantified as a function of both time and peptide concentration.

Aim 3: Engineer a vancomycin delivery system with improved antibacterial properties. Vancomycin will be systematically modified in order to specifically target the antibiotic to S. epidermidis biofilms. The 6-20 peptide will be incorporated into the antibiotic so as to engineer a new targeted antibiotic.

Various crosslinking chemistries will be studied in order to determine the effects of linker length and composition on antibacterial effectiveness. PEG linkers of various lengths and orientations will be studied, as well as zero-length crosslinkers. The system will be optimized to determine the optimal separation distance and spatial orientation between peptide and antibiotic resulting in the greatest improvement in antibacterial activity.

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3.4 REFERENCES

1. MacKintosh EE, Patel JD, Marchant RE, and Anderson JM, Effects of

biomaterial surface chemistry on the adhesion and biofilm formation of

Staphylococcus epidermidis in vitro. J Biomed Mater Res A, 2006. 78(4): p.

836‐42.

2. Desai NP, Hossainy SFA, and Hubbell JA, Surface‐immobilized polyethylene

oxide for bacterial repellence. Biomaterials, 1992. 13(7): p. 417‐420.

3. Wei J, Ravn DB, Gram L, and Kingshott P, Stainless steel modified with

poly(ethylene glycol) can prevent protein adsorption but not bacterial adhesion.

Colloids and Surfaces B: Biointerfaces, 2003. 32(4): p. 275‐291.

4. K Ista L, Fan H, Baca O, and P LÃ³pez G, Attachment of bacteria to model

solid surfaces: oligo (ethylene glycol) surfaces inhibit bacterial attachment. FEMS

microbiology letters, 1996. 142(1): p. 59‐63.

5. Leckband D, Sheth S, and Halperin A, Grafted poly(ethylene oxide) brushes as

nonfouling surface coatings. Journal of Biomaterials Science, Polymer

Edition, 1999. 10(10): p. 1125‐1147.

6. Marsh LH, Coke M, Dettmar PW, Ewen RJ, Havler M, Nevell TG, Smart

JD, Smith JR, Timmins B, Tsibouklis J, and Alexander C, Adsorbed

poly(ethyleneoxide)–poly(propyleneoxide) copolymers on synthetic surfaces:

Spectroscopy and microscopy of polymer structures and effects on adhesion of

135

skin‐borne bacteria. Journal of Biomedical Materials Research, 2002. 61(4): p.

641‐652.

7. Roosjen A, Kaper HJ, van der Mei HC, Norde W, and Busscher HJ,

Inhibition of adhesion of yeasts and bacteria by poly(ethylene oxide)‐brushes on

glass in a parallel plate flow chamber. Microbiology, 2003. 149(11): p. 3239‐

3246.

8. Nejadnik MR, van der Mei HC, Norde W, and Busscher HJ, Bacterial

adhesion and growth on a polymer brush‐coating. Biomaterials, 2008. 29(30): p.

4117‐4121.

9. Cringus‐Fundeanu I, Luijten J, van der Mei HC, Busscher HJ, and

Schouten AJ, Synthesis and Characterization of Surface‐Grafted Polyacrylamide

Brushes and Their Inhibition of Microbial Adhesion. Langmuir, 2007. 23(9): p.

5120‐5126.

10. Costerton J, Lewandowski Z, Caldwell D, Korber D, and Lappin‐Scott H,

Microbial biofilms. Annu Rev Microbiol, 1995. 49: p. 711‐45.

11. Hayward JA and Chapman D, Biomembrane surfaces as models for polymer

design: the potential for haemocompatibility. Biomaterials, 1984. 5(3): p. 135‐

142.

12. Lewis AL, Phosphorylcholine‐based polymers and their use in the prevention of

biofouling. Colloids and Surfaces B: Biointerfaces, 2000. 18(3‐4): p. 261‐275.

136

13. Hirota K, Murakami K, Nemoto K, and Miyake Y, Coating of a surface with

2‐methacryloyloxyethyl phosphorylcholine (MPC) co‐polymer significantly

reduces retention of human pathogenic microorganisms. FEMS Microbiology

Letters, 2005. 248(1): p. 37‐45.

14. Fujii K, Matsumoto HN, Koyama Y, Iwasaki Y, Ishihara K, and Takakuda

K, Prevention of biofilm formation with a coating of 2‐methacryloyloxyethyl

phosphorylcholine polymer. Journal of Veterinary Medical Science, 2008.

70(2): p. 167‐173.

15. Cheng G, Zhang Z, Chen S, Bryers JD, and Jiang S, Inhibition of bacterial

adhesion and biofilm formation on zwitterionic surfaces. Biomaterials, 2007.

28(29): p. 4192‐4199.

16. Vasilev K, Cook J, and Griesser HJ, Antibacterial surfaces for biomedical

devices. Expert Review of Medical Devices, 2009. 6(5): p. 553‐567.

17. Rai M, Yadav A, and Gade A, Silver nanoparticles as a new generation of

antimicrobials. Biotechnology advances, 2009. 27(1): p. 76‐83.

18. Riley DK, Classen DC, Stevens LE, and Burke JP, A large randomized clinical

trial of a silver‐impregnated urinary catheter: Lack of efficacy and staphylococcal

superinfection. The American Journal of Medicine, 1995. 98(4): p. 349‐356.

19. Schierholz and J., Implant infections: a haven for opportunistic bacteria.

Journal of Hospital Infection, 2001. 49(2): p. 87‐93.

137

20. Furno F, Morley KS, Wong B, Sharp BL, Arnold PL, Howdle SM, Bayston

R, Brown PD, Winship PD, and Reid HJ, Silver nanoparticles and polymeric

medical devices: a new approach to prevention of infection? Journal of

Antimicrobial Chemotherapy, 2004. 54(6): p. 1019‐1024.

21. Stobie N, Duffy B, McCormack DE, Colreavy J, Hidalgo M, McHale P, and

Hinder SJ, Prevention of Staphylococcus epidermidis biofilm formation using a

low‐temperature processed silver‐doped phenyltriethoxysilane solâ€“gel coating.

Biomaterials, 2008. 29(8): p. 963‐969.

22. Lee H, Lee Y, Statz AR, Rho J, Park TG, and Messersmith PB, Substrate‐

Independent Layer‐by‐Layer Assembly by Using Mussel‐Adhesive‐Inspired

Polymers. Advanced Materials, 2008. 20(9): p. 1619‐1623.

23. Cao H and Liu X, Silver nanoparticles‐modified films versus biomedical device‐

associated infections. Wiley Interdisciplinary Reviews: Nanomedicine and

Nanobiotechnology, 2010. 2(6): p. 670‐684.

24. Vimala K, Samba Sivudu K, Murali Mohan Y, Sreedhar B, and Mohana

Raju K, Controlled silver nanoparticles synthesis in semi‐hydrogel networks of

poly(acrylamide) and carbohydrates: A rational methodology for antibacterial

application. Carbohydrate Polymers, 2009. 75(3): p. 463‐471.

138

25. Ho CH, Tobis J, Sprich C, Thomann R, and Tiller JC, Nanoseparated

polymeric networks with multiple antimicrobial properties. Advanced

materials, 2004. 16(12): p. 957‐961.

26. Gaonkar TruptiÂ AP, Caraos LBS, and Modak SP, Efficacy of a Silicone

Urinary Catheter Impregnated with Chlorhexidine and Triclosan Against

Colonization With Proteus mirabilis and Other UropathogensÂ â€¢â€‚.

Infection Control and Hospital Epidemiology, 2007. 28(5): p. 596‐598.

27. Gaonkar TAP, Sampath LABA, and Modak SMP, Evaluation of the

Antimicrobial Efficacy of Urinary Catheters Impregnated with Antiseptics in an

In Vitro Urinary Tract ModelÂ â€¢â€‚. Infection Control and Hospital

Epidemiology, 2003. 24(7): p. 506‐513.

28. Blanchemain N, Haulon S, Martel B, Traisnel M, Morcellet M, and

Hildebrand HF, Vascular PET prostheses surface modification with cyclodextrin

coating: development of a new drug delivery system. European journal of

vascular and endovascular surgery, 2005. 29(6): p. 628‐632.

29. Blanchemain N, Haulon S, Boschin F, Marcon‐Bachari E, Traisnel M,

Morcellet M, Hildebrand HF, and Martel B, Vascular prostheses with

controlled release of antibiotics: Part 1: Surface modification with cyclodextrins of

PET prostheses. Biomolecular Engineering, 2007. 24(1): p. 149‐153.

139

30. Prabu P, Dharmaraj N, Aryal S, Lee BM, Ramesh V, and Kim HY,

Preparation and drug release activity of scaffolds containing collagen and

poly(caprolactone). Journal of Biomedical Materials Research Part A, 2006.

79A(1): p. 153‐158.

31. Kugler R, Bouloussa O, and Rondelez F, Evidence of a charge‐density

threshold for optimum efficiency of biocidal cationic surfaces. Microbiology,

2005. 151(5): p. 1341.

32. Gottenbos B, van der Mei HC, Klatter F, Nieuwenhuis P, and Busscher HJ,

In vitro and in vivo antimicrobial activity of covalently coupled quaternary

ammonium silane coatings on silicone rubber. Biomaterials, 2002. 23(6): p.

1417‐1423.

33. Jose B, Antoci Jr V, Zeiger AR, Wickstrom E, and Hickok NJ, Vancomycin

Covalently Bonded to Titanium Beads Kills Staphylococcus aureus. Chemistry

& Biology, 2005. 12(9): p. 1041‐1048.

34. Lawson MKC, Bowman CN, and Anseth KS, Vancomycin derivative

photopolymerized to titanium kills S. epidermidis. Clinical orthopaedics and

related research, 2007. 461: p. 96.

35. Lawson MC, Shoemaker R, Hoth KB, Bowman CN, and Anseth KS,

Polymerizable vancomycin derivatives for bactericidal biomaterial surface

140

modification: structure‐function evaluation. Biomacromolecules, 2009. 10(8):

p. 2221‐34.

36. Smith HW and Huggins MB, Successful treatment of experimental Escherichia

coli infections in mice using phage: its general superiority over antibiotics.

Journal of General Microbiology, 1982. 128(2): p. 307.

37. Curtin JJ and Donlan RM, Using Bacteriophages To Reduce Formation of

Catheter‐Associated Biofilms by Staphylococcus epidermidis. Antimicrobial

Agents and Chemotherapy, 2006. 50(4): p. 1268‐1275.

38. Cerca N, Oliveira R, and Azeredo J, Susceptibility of Staphylococcus

epidermidis planktonic cells and biofilms to the lytic action of staphylococcus

bacteriophage K. Letters in Applied Microbiology, 2007. 45(3): p. 313‐317.

39. Donlan RM, Preventing biofilms of clinically relevant organisms using

bacteriophage. Trends in microbiology, 2009. 17(2): p. 66‐72.

40. Kutter E, Raya R, and Carlson K, Molecular mechanisms of phage infection, in

Bacteriophages: Biology and Applications, E. Kutter and A. Sulakvelidze,

Editors. 2005. p. 165‐222.

41. Jerne NK and Avegno P, The Development of the Phage‐Inactivating

Properties of Serum During the Course of Specific Immunization of an Animal:

Reversible and Irreversible Inactivation. The Journal of Immunology, 1956.

76(3): p. 200‐208.

141

42. Rennermalm A, Nilsson M, and Flock JI, Fibrinogen binding protein of

Staphylococcus epidermidis is a target for opsonic antibodies. Infection and

Immunity, 2004. 72(5): p. 3081‐3083.

43. Vernachio JH, Bayer AS, Ames B, Bryant D, Prater BD, Syribeys PJ,

Gorovits EL, and Patti JM, Human immunoglobulin G recognizing fibrinogen‐

binding surface proteins is protective against both Staphylococcus aureus and

Staphylococcus epidermidis infections in vivo. Antimicrobial Agents and

Chemotherapy, 2006. 50(2): p. 511‐518.

44. Bloom B, Schelonka R, Kueser T, Walker W, Jung E, Kaufman D, Kesler K,

Roberson D, Patti J, and Hetherington S, Multicenter study to assess safety

and efficacy of INH‐A21, a donor‐selected human staphylococcal

immunoglobulin, for prevention of nosocomial infections in very low birth weight

infants. The Pediatric infectious disease journal, 2005. 24(10): p. 858.

45. Ohlsen K and Lorenz U, Immunotherapeutic strategies to combat

staphylococcal infections. International Journal of Medical Microbiology,

2010. 300(6): p. 402‐410.

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CHAPTER 4

FIBRINOGEN-BASED LIGAND FOR SPECIFIC TARGETING AND DELIVERY TO SURFACE-ADHERENT STAPHYLOCOCCUS EPIDERMIDIS

4.1 INTRODUCTION

Staphylococcus epidermidis is a coagulase negative, gram positive bacteria that has been implicated in the nosocomial infections of many blood-contacting biomaterials[1]. A two-stage process can be used to describe the course of such foreign-body infections, wherein initial adhesion of the bacteria to the material occurs first, followed by proliferation, matrix secretion, and cell-cell adhesion leading to the formation of a mature, multi-layered biofilm[1]. The requisite initial adhesion can occur either nonspecifically to the bare surface through hydrophobic and charge interactions, or specifically through protein mediated ligand-receptor interactions. Plasma proteins coat blood contacting surfaces almost immediately upon implantation, and consequently specific protein- mediated binding is likely to have a much more significant contribution towards initial bacterial adhesion[1].

S. epidermidis has the ability to adhere specifically to a variety of host plasma proteins that adsorb readily onto implanted surfaces[2]. Teichoic acid, a component found on the surface of many gram positive bacteria, has been shown to bind to both fibronectin[3] and fibrin[4], while surface-associated proteins are

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known to bind to fibrinogen[5-7], fibronectin[8], and vitronectin[9, 10]. These

bacterial proteins are part of a family of adhesins termed microbial surface

components recognizing adhesive matrix molecules (MSCRAMMs)[11, 12],

several of which have been identified in S. epidermidis[13]. AtlE, in addition to

aiding in nonspecific adhesion to bare surfaces, is able to bind specifically to

vitronectin[9], while Embp is known to provide an additional route for binding fibronectin[8]. SdrG, also referred to as Fbe, is the most studied S. epidermidis

MSCRAMM and it binds fibrinogen[5, 14-16] through a 331 amino acid

fibrinogen binding region (aa269-599) located in the SdrG A-Domain[5, 13, 17].

SdrG binds to the amino-terminus of the fibrinogen B chain[7, 14] with a KD of 0.9x10-7 M[14]. A synthetic peptide sequence representing the first 25 amino acid residues of the Bchain of fibrinogen (1-25) was used by Davis et al to

mimic the SdrG binding region, and this peptide bound to fibrinogen with a KD of 1.4x10-7 M [14]. This indicates that the significant amino acids necessary for

the SdrG-Fibrinogen binding interaction are present in the 1-25 peptide, and

that this linear sequence is in a nearly optimal conformational state[14]. The

fibrinogen binding domain was further narrowed down to the 6-20 peptide,

consisting of the amino acid sequence NEEGFFSARGHRPLD[14], with amino

acids 10-15 contributing most significantly to the SdrG-fibrinogen binding

scheme. 6-9 and 19-20 at either end of the peptide are not involved in

significant physical interactions with the bacterial surface proteins and are in fact

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extended into the solvent environment[18]. Exchanging any single amino acid residue in this 10-15 binding region for an alanine residue (or serine in the case of Ala13) significantly reduces the binding affinity of the peptide[18].

The SdrG-fibrinogen interaction has been shown to play a significant role in the initial stages of bacterial adhesion. Recombinant SdrG is able to block the adherence of S. epidermidis to fibrinogen in a concentration dependent manner[15]. Adding SdrG under saturating conditions prevents S. epidermidis from binding to fibrinogen[7], and mutating the fibrinogen-binding region of the

SdrG A-domain of S. epidermidis greatly impairs the ability to bind to immobilized fibrinogen.[18, 19]. Furthermore, both the gene encoding for

SdrG[13] and the SdrG protein itself[16] are present in 100% of clinical S. epidermidis isolates tested, and studies of infected and recovering patients reveal anti-SdrG antibodies in the serum, confirming that the bacteria express SdrG during the infectious stages[16, 20]. An in-vivo intravascular catheter infection model confirmed the importance of SdrG: an SdrG-negative S. epidermidis mutant had a 20% infection rate, compared to a 100% rate for the SdrG-positive strain[21].

Given the importance and prevalence of the SdrG-fibrinogen binding scheme in the initial stages of S. epidermidis infections, simulating and characterizing this binding behavior in vitro could provide a mechanism for specifically targeting or preventing S. epidermidis infections in vivo. The experiments presented here were

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designed to examine the ability of the synthetic 6-20 peptide to specifically target S. epidermidis in vitro.

4.2 MATERIALS AND METHODS

Peptide Synthesis

The peptides NEEGFFSARGHRPLD (6-20), NEEGFFSAAGHRPLD (6-20-

R14A), and NEEGFFSARAHRPLD (6-20-G15A), were synthesized by standard solid phase peptide synthesis on a 433a ABI peptide synthesizer, using 9- fluoronylmethoxycarbonyl (fmoc) amino acids on a Knorr resin[22]. The peptides were cleaved and deprotected using trifluoroacetic acid (TFA), with water, ethandithiol (EDT), triisopropylsilane (TIS), and phenol as scavengers

(82.5:5:2.5:1 TFA:Water:EDT:TIS; 100 mg phenol; 100 mg resin). Reverse phase high performance liquid chromatography (RP-HPLC) was used to purify the cleaved peptides, using water (with 0.1% TFA) and acetonitrile (with 0.082%

TFA) as the aqueous and organic components of the mobile phase. MALDI-TOF mass spectrometry was used to verify that the peptides were of the correct molecular weight, and absence of additional peaks indicated a pure product. The dried, purified peptides were stored at -20ºC until needed.

Peptide Labeling and Conjugate Purification

The 6-20 peptide was labeled for imaging (scheme shown in figure 4.1) using

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Figure 4.1: 6-20-NG synthesis scheme. A 20-fold excess of 6-20 peptide was added to Sulfo-NHS-Nanogold in 0.02 M HEPES buffer (pH 7.5) for 1 hour at room temperature. Unbound peptide was removed by centrifuge filtration. The central 10-18 residues (FFSARGHRP) are necessary for binding to SdrG.

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sulfo-N-hydroxy succinimide Nanogold (Nanoprobes, Inc; Yaphank, NY).

Nanogold is a discrete, uncharged gold label 1.4nm in diameter with a single sulfo-N-hydroxy succinimide moiety which covalently binds to the primary

amine at the N-terminus of the 6-20 peptide. A 6 nmol quantity of Nanogold

was dissolved in 0.2 mL of Millipore water. A 28-fold excess of the 6-20 peptide

was dissolved in 1 mL of 0.02 M HEPES buffer (4-(2-hydroxyethyl)-1-

piperazineethanesulfonic acid, pH 7.5 using dilute NaOH). The 2 solutions were

then mixed and incubated at room temperature for 1 hour while gently shaking.

The excess of peptide ensured that all of the Nanogold covalently bound to a peptide.

Centrifuge filtration was used to separate the unbound peptide from the peptide-Nanogold conjugate (6-20-NG) based on molecular weight. The

molecular weights for the unbound peptide and the Nanogold labeled peptide

were approximately 1.73 kDa and 16.73 kDa, respectively. The labeled peptide

was separated from the unlabeled peptide using a Millipore Centricon YM-10

centrifuge filter with a molecular weight cutoff of 10 kDa. The 1.2 mL

peptide/Nanogold mixture was pipetted into the filter device and the filter was

spun at 4000x g. After 2 hours, the filtrate was removed and discarded, while the

filter was inverted and spun for 3 minutes at 1000x g to recover the labeled

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peptide into a new vial. This procedure was repeated twice to enhance the

purification.

Seeding of Sample Surfaces

S. epidermidis strain RP62A was used for all procedures. An inoculating loop

was used to retrieve a small sample of S. epidermidis from a refrigerated culture

plate not more than two weeks old. The bacteria were incubated in TSB for 18-24

hours at 37ºC while shaking at 120 rpm, then pelleted and washed with fresh PBS

before being resuspended in PBS to a final concentration of 1x108 cfu/ml, determined by optical density readings (550 nm).

Bacteria were grown on poly(ethylene terephthalate) (PET) surfaces (15mm diameter). Prior to seeding, surfaces were cleaned with ethanol, placed into sterile 24-well plates, and secured in place using sterilized silicone rings.

Bacteria solution (200 L, 1x108 cfu/ml) was then added to each well and the

plates were covered and incubated at 37ºC for 2 hours.

Targeting/Specificity Studies

Plates were removed from the incubator after 2 hours and the sample surfaces

were gently rinsed with fresh PBS to remove non-adherent bacteria. To test the

specificity of the 6-20 probe towards S. epidermidis, Nanogold labeled peptide

(6-20-NG) was added directly to one set of substrates, while another set was

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blocked with a solution of 1% BSA in PBS prior to adding the 6-20-NG peptide

to reduce nonspecific peptide adhesion.

To further examine specificity, three additional samples sets were prepared.

Unlabeled peptide (6-20, 6-20-G15A, or 6-20-R14A) was added to the bacteria in 1% BSA-PBS for 1 hour at room temperature, followed by a gentle rinse with fresh 1% BSA-PBS and the subsequent addition of the 6-20-NG peptide for 1 hour. After binding, all wells were rinsed with 1% BSA-PBS.

The 1.4 nm Nanogold particles must be enhanced in order to make them clearly visible in the scanning electron microscope (SEM). GoldEnhance

(Nanoprobes, Inc; Yaphank, NY) was used to deposit additional gold onto the

Nanogold particles already bound to the bacterial surface. The wells were rinsed three times with fresh Millipore water, after which 20 l of the freshly prepared

GoldEnhance solution was added to 200 l water in each well for 15 minutes, resulting in Nanogold particles with diameters of 20-50 nm. To halt the development process, the wells were rinsed three times with Millipore water.

Finally, samples were covered with a 2.5% buffered glutaraldehyde solution and bacteria were fixed at 4ºC overnight.

Preparing Samples for SEM

Substrates were dehydrated for imaging using a water/ethanol gradient of

50%, 70%, 95%, and 100% ethanol. Samples were then covered 2x15 min with

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hexamethyldisilazane (HMDS) and allowed to air dry while covered in a fume

hood. When dry, the substrates were mounted onto SEM stubs and sputter

coated with a 50 Å layer of palladium. The samples were imaged using a Hitachi

S-4500 Scanning Electron Microscope

Statistics

Statistical analysis was done using Minitab 15, with data represented as mean plus standard error. Comparisons were made using an un-paired student’s t-test with p < 0.05 considered significant.

4.3 RESULTS

Purity of the 6-20, 6-20-R14A, and 6-20-G15A peptides was assessed using

MALDI-TOF mass spectrometry. Single peaks at the expected molecular weights

(1730, 1645, and 1744 m/z respectively) were indicative of a pure product.

Complementary analysis by RP-HPLC showed a single peptide elution peak

when monitored at 220 nm. Results of the 6-20 peptide analysis are shown in

figure 4.2 (6-20-R14A and 6-20-G15A results are similar, but not shown).

In order to test the specificity of the 6-20 peptide, a Bovine Serum Albumin

(BSA) blocking experiment was carried out. Figure 4.3 is representative of the

images obtained, wherin the amount of background nonspecific binding is

significantly decreased when BSA is used to passivate the exposed substrate

(Figure 4.3A vs. Figure 4.3B). More importantly, the addition of BSA did not

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MALDI-TOF

RP-HPLC

3.0

2.5 nm) 2.0 (220 1.5

1.0

Absorbance 0.5

0.0 10 15 20 25 30 35 Retention Time (minutes)

Figure 4.2: Analysis of 6-20 purity by MALDI-TOF mass spectrometry (top) and RP-HPLC (bottom). MALDI-TOF results indicate a single peptide product with the expected molecular weight of 1730.767. RP-HPLC using a C-18 analytical scale column (25 minute linear gradient, 2-30% acetonitrile in water) confirmed the purity of the 6-20 peptide.

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Figure 4.3: Blocking non-specific 6-20-NG adhesion to substrate. S. epidermidis was grown on PET, followed by the addition of 6-20-NG peptide with (B,D) or without (A,C) a 1% BSA blocking step. BSA greatly reduced nonspecific binding, but did not reduce specific binding. White arrows identify several Nanogold labels.

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prevent the Nanogold labels from binding to the bacteria (Figure 4.3C vs. Figure

4.3D)

ImageJ software[23] was used to quantify peptide binding. The surface area

covered by bacteria (Ab) and the exposed PET substrate (As) was measured, and

the total number of Nanogold labels adhering to bacteria (Nb) and substrate (Ns)

were counted. Normalized nonspecific binding (NB) and normalized specific

binding (SB) were defined as:

N N NB  s SB  b A A s b

Figure 4.4 depicts the binding data obtained from these calculations. In the

absence BSA, there is no significant difference between nonspecific and specific

binding of the 6-20-NG peptide to the bacteria. However, by using a 1% BSA

blocking solution prior to adding the 6-20-NG peptide, nonspecific binding

decreases significantly (p< 0.022) when compared to nonspecific binding in the

absence of BSA. This results in a significant difference between specific and

nonspecific binding when BSA is used to block the exposed PET substrate (p<

0.011). The addition of BSA does not result in any significant change in specific

binding of 6-20-NG to the bacteria (p< 0.765).

To further examine the targeting ability of the 6-20 peptide, blocking studies

were carried out, with representative SEM images shown in Figure 4.5. The 6-

20 blocking sample (A) shows minimal Nanogold labels bound to the surface of the bacteria, while the samples with no blocking (B), 6-20-R14A blocking (C),

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Figure 4.4: Normalized 6-20-NG adhesion to PET (white) and S. epidermidis (shaded). Using BSA to passivate the exposed substrate (right) significantly decreases nonspecific binding (p< 0.022) while having no effect on specific binding (p< 0.765). Data represents mean plus standard error.

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Figure 4.5: Scanning electron microscope images of peptide blocking studies. Bacteria were exposed to A) 6-20 peptide, B) no peptide (control), C) 6-20-R14A peptide, or D) 6-20-G15A peptide prior to addition of the Nanogold-labeled 6-20-NG peptide. Nanogold labels (identified by white arrows) are abundant on the control, 6-20-R14A, and 6-20-G15A blocking samples (B,C,D) while there are minimal labels bound to the bacteria in the 6-20 blocking sample (A).

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Figure 4.6: Peptide blocking study. There was no significant difference in specific binding between the positive control and the 6-20-R14A or 6-20-G15A negative control blocking peptides (p< 0.744 and p< 0.592, respectively). Using the 6-20 peptide to block specific binding, however, resulted in a significant decrease relative to the positive control (p< 0.021). Data represents mean plus standard error.

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and 6-20-G15A blocking (D) all show Nanogold labels specifically binding to the bacteria.

Figure 4.6 depicts the results obtained from the peptide blocking experiments,

as determined by analysis with ImageJ. The control data (no peptide) represents

the labeled 6-20-NG peptide binding directly to the bacteria without an initial blocking step. When sample surfaces are incubated with either the 6-20-R14A or 6-20-G15A peptide prior to adding the labeled peptide, there are no significant differences in nonspecific binding when compared to the control (p<

0.744 and p< 0.592, respectively). However, when samples are incubated with the unlabeled 6-20 peptide prior to adding the labeled peptide, there is a significant decrease in specific binding relative to the control (p< 0.021).

4.4 DISCUSSION

Staphylococcus epidermidis is part of the normal human flora and considered to

be noninvasive and nonpathogenic under most circumstances. However,

implantation of a medical device provides a foreign surface that can be colonized

by bacteria that may be unintentionally introduced during the procedure.

Infections by S. epidermidis, unlike those by S. aureus, are rarely associated with

toxins and virulence factors. Instead, the determining factors for the success of

these infections are the initial adhesion of the bacteria to the surface, followed by

the ability to evade the host’s immune system through biofilm formation[24].

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This study strives to identify a binding ligand capable of specifically targeting

S. epidermidis. Since primary adhesion is such an important factor in the success of S. epidermidis infections, the MSCRAMM family of proteins, several of which are present on S. epidermidis, presents a suitable model for the development of such a targeting ligand. MSCRAMMs are responsible for binding a variety of host proteins such as fibronectin, vitronectin, and fibrinogen, all of which readily adsorb onto blood-contacting biomaterial surfaces. While S. epidermidis is capable of binding fibronectin through multiple surface molecules, heparin is known to inhibit the S. epidermidis-fibronectin interaction, rendering this binding model unattractive for the purposes of targeted binding[25]. The S. epidermidis- vitronectin binding interaction, while potentially more robust, has not been well characterized and the peptide sequence involved in the binding has not yet been identified. The S. epidermidis-fibrinogen interaction, on the other hand, has been examined in great detail, with studies revealing that the bacterial surface protein

SdrG binds to the fibrinogen 6-20 peptide sequence.

Fibrinogen represents a major route for potential bacterial colonization. In addition to its high plasma concentration, fibrinogen is a major component of thrombus formations. As part of the normal blood coagulation process, thrombin acts on the fibrinogen B chain, cleaving the Arg14-Gly15 peptide bond to release the 14 amino acid fibrinopeptide B (FpB). This cleavage facilitates fibrin assembly, eliminates the S. epidermidis SdrG binding site, and releases FpB to act

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as a chemotactic agent for blood leukocytes[26]. However, if SdrG manages to

bind to the intact fibrinogen B chain, it is able to prevent subsequent cleavage of

FpB[14]. In fact, even in the absence of S. epidermidis, only 30% of FpB is released

by thrombin from clots of whole blood, with the other 70% remaining intact to

provide ample binding sites for subsequent S. epidermidis adhesion[27]. The

SdrG-fibrinogen binding interaction therefore provides a significant route for

initial bacterial adhesion, as well as a potential mechanism for evasion of the host

immune system. Taking advantage of this known binding interaction will allow

for the development of a robust targeting ligand.

Four peptides were prepared for this study: the NEEGFFSARGHRPLD 6-20

peptide, a negative control NEEGFFSAAAHRPLD peptide (6-20-R14A), a

negative control NEEGFFSARAHRPLD peptide (6-20-G15A), and the 6-20 peptide with a single 1.4nm Nanogold label covalently attached to the amino terminus (6-20-NG). Due to the nature of the binding scheme, the 6-20 peptide sequence has four amino acids extending into solution at the amino terminus that are not involved in binding to SdrG[18]. Thus, attaching the Nanogold label to the amino terminus reduces the likelihood that the gold particle will interfere with the binding process. Furthermore, Nanogold labeling and scanning electron microscopy were chosen for this study because they provide the ability to visualize individual peptide labels adhering to the surface of the bacteria.

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In order to verify that the 6-20-NG peptide specifically binds to S. epidermidis, bovine serum albumin (BSA) was used to examine the tendency of the peptide to bind nonspecifically to the exposed substrate. In the absence of

BSA blocking (Figure 1A), the 6-20-NG peptide binds to both the bacteria and the exposed substrate, with no significant difference between the two (Figure 2, left). However, when surfaces are coated with BSA prior to adding 6-20-NG

(Figure 1B), the amount of peptide adhering to the substrate is significantly reduced (p < 0.022) while specific binding to the bacteria remained nearly constant (Figure 2, right). This demonstrates that the 6-20 interaction with biomaterial-adherent S. epidermidis is specific, as BSA blocking does not decrease the ability of the 6-20-NG probe to bind to its receptor on the bacterial surface

(Figure 1C,D).

In order to further characterize the peptide’s targeting abilities, the effects of the 6-20, 6-20-R14A, and 6-20-G15A peptides were studied in regards to their effectiveness at preventing 6-20-NG from binding to S. epidermidis. When the bacteria were exposed to the 6-20 peptide prior to adding 6-20-NG (Figure 3A), there were very few visible Nanogold-labeled peptides bound to the surface of the bacteria, indicating that the unlabeled peptide has occupied all of the available binding sites. When the labeled peptide was subsequently introduced, there were no SdrG proteins available for binding, and as a result the labeled peptide was washed away during the rinsing step.

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The 6-20-R14A and 6-20-G15A peptides were used as negative controls in

these studies. These two peptides have a greatly reduced binding affinity for

SdrG[18], so it is expected that they will have a reduced ability to block 6-20-NG

binding. Results in Figure 3C and 3D show that there are many Nanogold labels

adhering to the surface of the bacteria, therefore verifying the inability of the 6-

20-R14A and 6-20-G15A peptides to bind and block the SdrG proteins on the

bacterial surface. These results are similar to those shown in Figure 3B, where no blocking peptide was used.

Quantitatively (Figure 4), when using the 6-20 blocking peptide, specific

binding is significantly reduced (p< 0.021) relative to the control. However, the

negative control peptides, 6-20-R14A and 6-20-G15A, are unable to

significantly reduce this specific binding. Taken together, these results indicate

that the 6-20 peptide is able to bind specifically to the S. epidermidis SdrG

protein in vitro and prevent any subsequent binding to the receptor. Alteration

of the 6-20 peptide by even a single amino acid significantly impairs this

blocking ability.

The Fibrinogen-SdrG interaction is an important component of bacterial

adhesion to blood-contacting biomaterials, and the specificity of the 6-20

peptide demonstrated in vitro indicates the potential for using this peptide to

interfere with initial bacterial adhesion. The 6-20 peptide was able to

specifically deliver Nanogold particles to biomaterial-adherent bacteria, pointing

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towards a possible application as an in vivo targeting motif, wherein the

Nanogold label could be replaced by either an antibacterial drug (or drug carrier) for localized treatment or an imaging agent for diagnostic purposes. Subsequent chapters will examine different nanogold substitutions and their effect on biofilm formation. If successfully developed, a similar technique could be applied to target and treat infections of other bacterial species.

4.5 CONCLUSIONS

The results from this study show that the fibrinogen-based 6-20 peptide

sequence NEEGFFSARGHRPLD specifically targets biomaterial-adherent

Staphylococcus epidermidis RP62A under in vitro conditions, as demonstrated by

both BSA and peptide blocking studies. In these experiments, when binding sites

are selectively blocked, the labeled peptide is unable to bind to the bacteria, and

when BSA is used to coat the exposed substrate, nonspecific binding is greatly reduced while specific binding is unaffected. In these studies, nanogold particles

are used primarily as a tool for observing targeted binding. However, in

delivering the nanogold to the bacterial surface, the 6-20 peptide is identified as

a viable targeting motif for delivering antibacterial agents to S. epidermidis

infections.

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4.6 FUTURE STUDIES

A key area for future study will involve a more thorough investigation regarding the specificity of the 6-20 peptide. The studies presented in this chapter confirm that the 6-20 peptide is able to target S. epidermidis strain

RP62A, as confirmed by blocking studies using bovine serum albumin and non- binding substitution peptides. However, these controls alone do not fully characterize the binding specificity of the 6-20 peptide; two additional studies involving different strains of bacteria are necessary to fully confirm specificity.

First, binding studies using the 6-20-NG peptide should be carried out using an alternative species such as Staphylococcus aureus, which is very closely related to S. epidermidis. S. aureus displays two fibrinogen-binding proteins, clumping factor A and B (ClfA and ClfB), that are structurally similar to the S. epidermidis

SdrG protein. As a result, comparing binding behavior of the 6-20 peptide towards S. epidermidis and S. aureus would demonstrate that the peptide is in fact specifically targeting S. epidermidis, rather than generically targeting multiple bacteria. Second, targeting studies should be carried out with a strain of S. epidermidis lacking the SdrG protein. Since the gene encoding for SdrG[13] and the SdrG protein itself[16] have been shown to be present in 100% of clinical S. epidermidis isolates tested, an SdrG-negative S. epidermidis mutant will need to be developed and studied. Such a mutant would lack the 6-20 binding site (either through deletion or mutation), and as such should negate the ability of the

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peptide to specifically target S. epidermidis. The results of a study involving SdrG-

negative S. epidermidis would confirm the role of the SdrG protein in the 6-20

delivery scheme. When combined with the aforementioned study utilizing S.

aureus, in addition to the results presented in this chapter, the data would support the claim that the 6-20 peptide specifically targets the SdrG protein of S.

epidermidis.

Another area for investigation involves competition between the 6-20

peptide and the native fibrinogen molecule. Published studies have

demonstrated that recombinant SdrG is able to block the adherence of S.

epidermidis to fibrinogen in a concentration dependent manner[15], and when

SdrG is added under saturating conditions, S. epidermidis is unable to bind to fibrinogen[7]. Thus, it is clear that fibrinogen and the 6-20 peptide are competing for the same binding site on S. epidermidis. While the experiments presented in this chapter used the unlabeled 6-20 peptide to block subsequent binding of the nanogold labeled 6-20-NG peptide, a useful study that would extend the scope of this project would be to utilize the native fibrinogen protein as the blocking molecule instead of the 6-20 peptide. Such studies would bring to light potential limitations of the 6-20 peptide, given that any future in vivo

use of the peptide would involve a fibrinogen-rich environment that may be able

to compromise targeting binding and delivery. Additional targeting studies

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involving 6-20-NG in the presence of physiological concentrations of fibrinogen would elucidate the necessary conditions for targeted delivery.

Finally, targeting of S. epidermidis should be examined at different stages of biofilm formation. Studies in this chapter examined targeted delivery to surface- adherent bacteria prior to biofilm formation. As SdrG is an adhesion molecule important in the early stages of biofilm formation, it is likely that SdrG expression will decrease as the biofilm matures. Therefore, delivery of 6-20-NG to S. epidermidis biofilms at various developmental time points (2, 4, 6, 8, 12, 18,

24, 48 hrs) will provide data detailing the efficiency of targeting biofilms at various stages of growth. The need for this data will become even more evident after examining results of experiments presented in subsequent chapters, wherein the targeted cationic peptide and targeted vancomycin molecule performs significantly better during the early stages of biofilm development.

Knowledge of targeting ability in relation to biofilm maturity will play a crucial role in determining the focus of future research.

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4.7 ACKNOWLEDGEMENTS

The project described here was funded by Award

Number 5R01EB000279 from the National Institute of Biomedical Imaging and

Bioengineering, and work was carried out using the facilities of the Center for

Cardiovascular Biomaterials. I would also like to thank Eric Anderson, a former graduate student in this lab, for his support and guidance in teaching me the art of solid phase peptide synthesis.

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4.8 REFERENCES

1. Vuong C and Otto M, Staphylococcus epidermidis infections. Microbes And

Infection, 2002. 4(4): p. 481‐489.

2. Bale MD, Wohlfahrt LA, Mosher DF, Tomasini B, and Sutton RC,

Identification of Vitronectin as a Major Plasma‐Protein Adsorbed on Polymer

Surfaces of Different Copolymer Composition. Blood, 1989. 74(8): p. 2698‐2706.

3. Hussain M, Heilmann C, Peters G, and Herrmann M, Teichoic acid enhances

adhesion of Staphylococcus epidermidis to immobilized fibronectin. Microbial

Pathogenesis, 2001. 31(6): p. 261‐270.

4. Chugh TD, Burns GJ, Shuhaiber HJ, and Bahr GM, Adherence Of

Staphylococcus‐Epidermidis To Fibrin‐Platelet Clots Invitro Mediated By

Lipoteichoic Acid. Infection And Immunity, 1990. 58(2): p. 315‐319.

5. Nilsson M, Frykberg L, Flock JI, Pei L, Lindberg M, and Guss B, A

fibrinogen‐binding protein of Staphylococcus epidermidis. Infection and

Immunity, 1998. 66(6): p. 2666‐2673.

6. Pei L, Arvholm IL, Lonnies L, and Flock JI, GST‐Fbe can recognize beta‐

chains of fibrin(ogen) on explanted materials. Journal of Chromatography B‐

Analytical Technologies in the Biomedical and Life Sciences, 2003. 786(1‐

2): p. 319‐325.

168

7. Pei L, Palma M, Nilsson M, Guss B, and Flock JI, Functional studies of a

fibrinogen binding protein from Staphylococcus epidermidis. Infection and

Immunity, 1999. 67(9): p. 4525‐4530.

8. Williams RJ, Henderson B, Sharp LJ, and Nair SP, Identification of a

fibronectin‐binding protein from Staphylococcus epidermidis. Infection And

Immunity, 2002. 70(12): p. 6805‐6810.

9. Heilmann C, Hussain M, Peters G, and Gotz F, Evidence for autolysin‐

mediated primary attachment of Staphylococcus epidermidis to a polystyrene

surface. Molecular Microbiology, 1997. 24(5): p. 1013‐1024.

10. Li DQ, Lundberg F, and Ljungh A, Characterization of vitronectin‐binding

proteins of Staphylococcus epidermidis. Current Microbiology, 2001. 42(5): p.

361‐367.

11. Patti JM and Hook M, Microbial Adhesins Recognizing Extracellular‐Matrix

Macromolecules. Current Opinion In Cell Biology, 1994. 6(5): p. 752‐758.

12. Patti JM, Allen BL, Mcgavin MJ, and Hook M, MSCRAMM‐Mediated

Adherence of Microorganisms to Host Tissues. Annual Review of

Microbiology, 1994. 48: p. 585‐617.

13. Bowden MG, Chen W, Singvall J, Xu Y, Peacock SJ, Valtulina V, Speziale

P, and Hook M, Identification and preliminary characterization of cell‐wall‐

169

anchored proteins of Staphylococcus epidermidis. Microbiology‐Sgm, 2005. 151:

p. 1453‐1464.

14. Davis SL, Gurusiddappa S, McCrea KW, Perkins S, and Hook M, SdrG, a

fibrinogen‐binding bacterial adhesin of the microbial surface components

recognizing adhesive matrix molecules subfamily from Staphylococcus

epidermidis, targets the thrombin cleavage site in the B beta chain. Journal of

Biological Chemistry, 2001. 276(30): p. 27799‐27805.

15. Hartford O, OʹBrien L, Schofield K, Wells J, and Foster TJ, The Fbe (SdrG)

protein of Staphylococcus epidermidis HB promotes bacterial adherence to

fibrinogen. Microbiology‐Sgm, 2001. 147: p. 2545‐2552.

16. McCrea KW, Hartford O, Davis S, Eidhin DN, Lina G, Speziale P, Foster

TJ, and Hook M, The serine‐aspartate repeat (Sdr) protein family in

Staphylococcus epidermidis. Microbiology‐Uk, 2000. 146: p. 1535‐1546.

17. Hall AE, Patel PR, Domanski PJ, Prater BD, Gorovits EL, Syribeys PJ,

Vernachio JH, Patti JM, and Hutchins JT, A panel of monoclonal antibodies

recognizing the Staphylococcus epidermidis fibrinogen‐binding MSCRAMM

SdrG. Hybridoma, 2007. 26(1): p. 28‐34.

18. Ponnuraj K, Bowden MG, Davis S, Gurusiddappa S, Moore D, Choe D, Xu

Y, Hook M, and Narayana SVL, A ʺdock, lock, and latchʺ structural model for

a staphylococcal adhesin binding to fibrinogen. Cell, 2003. 115(2): p. 217‐228.

170

19. Pei L and Flock JI, Lack of fbe, the gene for a fibrinogen‐binding protein from

Staphylococcus epidermidis, reduces its adherence to fibrinogen coated surfaces.

Microbial Pathogenesis, 2001. 31(4): p. 185‐193.

20. Rennermalm A, Nilsson M, and Flock JI, Fibrinogen binding protein of

Staphylococcus epidermidis is a target for opsonic antibodies. Infection and

Immunity, 2004. 72(5): p. 3081‐3083.

21. Guo BN, Zhao X, Shi YG, Zhu DM, and Zhang YY, Pathogenic implication of

a fibrinogen‐binding protein of Staphylococcus epidermidis in a rat model of

intravascular‐catheter‐associated infection. Infection and Immunity, 2007.

75(6): p. 2991‐2995.

22. Fields G and Noble R, Solid phase peptide synthesis utilizing 9‐

fluorenylmethoxycarbonyl amino acids. Int J Pept Protein Res, 1990. 35(3): p.

161‐214.

23. Abramoff M, Magelhaes P, and Ram S, Image Processing with ImageJ.

Biophotonics International, 2004. 11(7): p. 36‐42.

24. Queck SY and Otto M, Staphylococcus epidermidis and other Coagulase‐

Negative Staphylococci, in Staphylococcus Molecular Genetics, J.A. Lindsay,

Editor. 2008, Caister Academic Press: Norfolk, UK.

171

25. Arciola CR, Bustanji Y, Conti M, Campoccia D, Baldassarri L, Samori B,

and Montanaro L, Staphylococcus epidermidis ‐ fibronectin binding and its

inhibition by heparin. Biomaterials, 2003. 24(18): p. 3013‐3019.

26. Senior R, Skogen W, Griffin G, and Wilner G, Effects of Fibrinogen

Derivatives Upon the Inflammatory Response. J. Clin. Invest., 1986. 77: p.

1014‐1019.

27. Brummel K, Butenas S, and Mann K, An Integrated Study of Fibrinogen

During Blood Coagulation. J. Biol. Chem., 1999. 274(32): p. 22862‐22870.

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CHAPTER 5

DISRUPTION OF STAPHYLOCOCCUS EPIDERMIDIS BIOFILM FORMATION USING A TARGETED CATIONIC PEPTIDE

Based on: Hofmann CM, Bednar KJ, Anderson JM, Marchant RE. Disruption of Staphylococcus epidermidis biofilm formation using a targeted cationic peptide. J Biomed Mater Res Part A, 2012; 100A:1061-1067.

5.1 INTRODUCTION

Staphylococcus epidermidis has become one of the most common infecting

organisms of implanted medical devices[1-3]. In its planktonic state,

S. epidermidis is known to be susceptible to a large number of antibiotics[4].

However, the S. epidermidis biofilm environment offers the encapsulated bacteria

increased resistance to antibiotics. With this protection, the bacteria can survive antibiotic concentrations several orders of magnitude higher than the minimum

inhibitory concentration (MIC) measured in planktonic suspensions[5-7]. Due to the reduced effectiveness of traditional antimicrobial treatments towards biofilms, removal of infected devices is often required[8].

Biofilm formation begins with the adhesion of the bacteria to the implanted

device[9]. One common mechanism of adhesion involves specific binding

between S. epidermidis and host plasma proteins that adsorb onto intravascular devices immediately following implantation[10-19]. This is mediated by

microbial surface components recognizing adhesive matrix molecules

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(MSCRAMMs)[20, 21]. SdrG is a well studied MSCRAMM responsible for S.

epidermidis binding to amino acids 6-20 of the fibrinogen B chain[15, 22]. The

fibrinogen-derived β6-20 peptide binds to S. epidermidis with an affinity similar to that of the native fibrinogen molecule[22].

After adhering to the substrate, S. epidermidis begins the process of biofilm formation through proliferation, polysaccharide production, and intercellular adhesion. A crucial step in the formation of a mature biofilm is the production of exopolysaccharides[23].

Polysaccharide intercellular adhesin (PIA), a -1,6-linked N-acetylglucosamine

(GlcNAc), is the main exopolysaccharide associated with S. epidermidis biofilms[24].

Multiple functions are attributed to PIA, including protection from host polymorphonuclear leucocytes and antimicrobial peptides[25, 26], virulence in several in

vivo models[27-30], hemagglutination[31], and intercellular adhesion[24, 32-34].

PIA is a cationic exopolysaccharide, with approximately 15-20% of the glucosamine

residues deacetylated by IcaB, a product of the icaADBC gene locus[24, 25]. PIA is

believed to associate with the bacterial surface and form fibrous strands through

electrostatic interactions [25, 26, 32]. However, with the creation of an icaB negative mutant, the non-deacetylated PIA was unable to associate with the bacterial surface and was instead released into the culture media[25]. The resulting inability to retain PIA at the bacterial surface led to the elimination of biofilm formation on plastic surfaces, reduced bacterial adhesion to human epithelial cells, and impaired resistance to neutrophil phagocytosis and antimicrobial peptides in vitro, while reducing the persistence of infection in an in vivo murine model[25].

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Most current methods of treating and preventing biofilms utilize bactericidal or

bacteriostatic approaches in an attempt to reduce the risk of an infection, as attempts to

eradicate established S. epidermidis infections are seen as difficult and potentially

counterproductive[1]. Antibiotic-impregnated, antibiotic-coated, and silver-coated

intravascular catheters have been tested with measured success at preventing

infections[35, 36], while hydrogel coatings have been used on silicone catheters to absorb

bacteriophages and significantly reduce the extent of S. epidermidis biofilm

development[37]. In an attempt to treat established biofilms, silver ions have been shown

to have a destabilizing effect on biofilm structure through interactions with

exopolysaccharides, although these ions remain largely ineffective as bactericidal agents

towards sessile bacteria[38].

Here, we utilize the 6-20 targeting peptide as a means to tether a positively charged

polylysine sequence to the bacterial surface. The aim of this work is to disrupt the

electrostatic interaction between S. epidermidis and PIA, inhibiting the ability of the PIA

to associate with the bacteria and resulting in disruption of the biofilm formation process.

5.2 MATERIALS & METHODS:

Peptide Synthesis

The peptides NEEGFFSARGHRPLD (6-20, MW 1730), GGGKKKKKK (G3K6,

MW 958), and NEEGFFSARGHRPLDGGGKKKKKK (6-20-G3K6, MW 2672) were

synthesized by solid phase peptide synthetic methods on a 433a ABI peptide synthesizer

(Applied Biosystems), using 9-fluoronylmethoxycarbonyl (fmoc) amino acids on a Knorr

resin[39]. The peptides were cleaved and deprotected using trifluoroacetic acid (TFA)

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and precipitated into cold ether. The solvents were then evaporated and the peptides were

dried by lyophilization. Reverse phase high performance liquid chromatography (HPLC)

was used to purify the cleaved peptides, using water (0.1 % TFA) and acetonitrile

(0.082% TFA) as the aqueous and organic components of the mobile phase.

MALDI-TOF mass spectrometry was used to confirm the purified peptide products were

of the correct molecular weight.

Seeding of Sample Surfaces

S. epidermidis strain RP62A was used for all procedures as a representative biofilm-

positive strain. An inoculating loop was used to retrieve a small sample of S. epidermidis

from a refrigerated culture plate not more than 2 weeks old. The bacteria were incubated

in tryptic soy broth (TSB) for 18 hours at 37°C while shaking at 120 rpm, then pelleted

and washed with fresh PBS before being resuspended in TSB to a final concentration of

0.5x108 cfu/ml, as determined by optical density (OD). For OD, microplate fluorescence,

and colorimetric analysis, bacteria were seeded on 96-well plates. For SEM and

epifluorescence microscopy analysis, bacteria were seeded on 24-well plates containing

poly(ethylene terephthalate) (PET) surfaces (15mm diameter). Prior to seeding, the PET

surfaces were cleaned with ethanol and secured in place using sterilized silicone tubing.

Bacteria were seeded onto the sample surfaces using the 0.5 x108 cfu/ml bacterial

suspension (150L for 96-well plates, 200 l for 24-well plates). The plates were incubated at 37°C for 2 hrs, after which they were gently rinsed with sterile PBS. 6-20,

G3K6, or 6-20-G3K6 peptide was then added to each well (175 L, 0-100 M in TSB), followed by an additional 21 hour incubation at 37°C.

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Optical Density Analysis of Bacterial Growth

For growth analysis, 96-well plates were incubated in an optical plate reader

(Molecular Devices) at 37°C and OD readings were taken at 15 minute intervals (550 nm) over the course of 21 hrs.

Fluorescence Analysis of Biofilm Composition

Fluorescent labeling was used to determine PIA and adherent bacteria content in treated biofilms. Samples were rinsed with PBS, followed by a blocking step with 0.5%

BSA (in PBS) solution. PIA was then labeled using wheat germ agglutinin (100 μl, 20

μg/ml in 0.85% NaCl, Invitrogen) conjugated to either Alexa Fluor 488 (WGA-488) or

Alexa Fluor 647 (WGA-647) for 15 minutes at room temperature. Samples labeled with

WGA-647 were subsequently labeled with SYTO 9 (100 L, 6M in water, Invitrogen) for 5 minutes. Epifluorescent microscopy (Nikon Diophot 200, 40x objective) was used to image samples grown on the PET substrates and labeled with WGA-488. Quantitative fluorescence intensity readings were taken of samples grown in 96-well plates and labeled with WGA-647 (excitation/emission: 650/665 nm) and SYTO-9

(excitation/emission: 485/530 nm).

XTT Assay for Bacterial Viability and Biofilm Composition

An XTT assay was used to assess bacterial viability and biofilm composition. For planktonic analysis, 30 μl culture media was removed from each well and transferred to the corresponding well of a sterile 96-well plate, to which 170 μl of a freshly prepared

XTT/PMS solution was added to yield a final concentration of 0.125 mM XTT (2,3-

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Bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide inner salt, Sigma

Aldrich) and 0.01 mM PMS (Phenazine methosulfate, Sigma Aldrich). For sessile analysis, wells were rinsed 3 times with PBS, followed by the addition of 200 μl of a freshly prepared XTT/PMS solution (0.125 mM XTT, 0.01 mM PMS). Both plates were then incubated at 37° for 3 hours, after which optical density readings were taken at 490 nm.

Scanning Electron Microscopy

Samples for scanning electron microscopy (SEM) analysis were dehydrated using a water/ethanol gradient of 50%, 70%, 95%, and 100% ethanol. Samples were then covered

2x15min with hexamethyldisilazane (HMDS) and allowed to air dry. The samples were mounted onto SEM stubs and sputter coated with a 50 Ǻ layer of palladium and imaged using a Hitachi S-4500 SEM.

Statistics

Comparisons were made using a one-way ANOVA with Dunnett’s post-hoc test

(α < 0.01 was considered to be significant). Statistical analysis was carried out using

Minitab 15, with data represented as mean ± standard deviation for a minimum of n=3.

5.3 RESULTS:

The effects of the various peptides on the growth of S. epidermidis biofilms were determined by optical density (Figure 5.1). Bacteria were allowed to grow for 21 hours in

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Figure 5.1: Optical density growth curves of surface-adherent S. epidermidis incubated in the presence of 50 μM β6-20, G3K6, or β6-20-G3K6 peptide (left) or various concentrations (0-100 μM) of the β6-20-G3K6 peptide (right). Data represents mean (n=3) for all curves.

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the presence of 50 μM β6-20, G3K6, and β6-20-G3K6 peptides (left) and 0-100 μM β6-20-

G3K6 peptide (right), with data normalized to the t=0 readings in each well. In all cases, the lag time prior to growth, the rate and duration of exponential growth, and the maximal bacterial concentration remain unchanged by both the type and concentration of peptide in the growth medium. These results indicate that peptide concentrations up to

100 μM were neither toxic nor inhibitory to the overall growth of the bacteria.

Wheat germ agglutinin (WGA) specifically labels polysaccharide intercellular adhesin (PIA), while SYTO 9 is a nucleic acid stain capable of penetrating the bacterial cell wall in order to assess cell numbers. As shown in figure 5.2, neither the β6-20 nor the

G3K6 peptides have an effect on the amount of PIA or bacteria present in surface-

adherent biofilms over the 0-100 μM concentration range tested. However, the β6-20-

G3K6 peptide significantly decreases PIA content and adherent bacteria relative to the 0

μM control when peptide concentration is greater than 50 μM. Specifically, 100 μM β6-

20-G3K6 was able to reduce PIA content of a biofilm by 37.9%, with a corresponding

17.5% reduction in surface-adherent bacteria.

SEM was used to visualize biofilm structures (Figure 5.3a,b), while epifluorescence

microscopy was used to visualize PIA content in biofilms (Figure 5.3c,d). A mature,

mutli-layered biofilm is visible with a significant PIA presence following a 21 hr

incubation period in the absence of peptide (Figure 5.3a,c). However, biofilms are

immature and large portions of uncolonized substrate remain visible when S. epidermidis

is grown in the presence of β6-20-G3K6 peptide (50 μM, Figure 5.3b,d). Biofilms grown

in the presence of β6-20 or G3K6 (50 μM) were indistinguishable from the biofilms

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Figure 5.2: Effects of peptide on the composition of surface-adherent biofilms as determined by quantitative fluorescence microplate readings. WGA-647 was used to monitor PIA content (left), while SYTO 9 was used to monitor the amount of surface- adherent bacteria (right). Data represents mean (n=12) ± standard deviation. Significance determined by one-way ANOVA (<0.01) with Dunnett’s post-hoc test.

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Figure 5.3: Biofilm structure after 21 hours as observed by SEM (a,b) and PIA staining with wheat germ agglutinin conjugated to Alexa Fluor 488 (c,d). Differences in biofilm formation were observed with the addition of 50 μM β6-20-G3K6 peptide (b,d) when compared to biofilms grown in the absence of peptide (a,c),

182

grown without peptide (data not shown). The SEM and epifluorescence images further

confirm the quantitative fluorescence readings obtained using WGA and SYTO 9,

indicating a reduction in PIA content and surface-adherent bacteria due to the β6-20-

G3K6 peptide.

The XTT assay was used to analyze the bacteria in each well in terms of sessile vs.

planktonic populations. When exposed to the assay reagent, metabolically active cells

reduce the tetrazolium salt to a water soluble formazan dye that absorbs strongly at 490

nm, with the amount of dye produced linearly related to the number of cells present.

Figure 5.4 illustrates the composition of each well (n=3) after exposure to 0-100 μM

peptide. While the β6-20 and G3K6 peptides have minimal effect on the sessile vs.

planktonic composition of the biofilms, there is a discernable shift of bacteria from the sessile to planktonic state in response to increasing concentrations of the β6-20-G3K6 peptide.

5.4 DISCUSSION:

Staphylococcus epidermidis biofilms contain significant amounts of polysaccharide intercellular adhesin (PIA), and it is believed that the cationic nature of this exopolysaccharide allows it to associate with the negatively charged bacterial surface during the biofilm accumulation process. Biofilm formation is reduced or eliminated and the bacteria’s resistance to phagocytosis and antimicrobial peptides is greatly reduced after eliminating the positive charges on the PIA [25]. This ability of S. epidermidis to form a biofilm is thought to be the most critical factor in determining its virulence[18].

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Figure 5.4: Biofilm composition after 21 hours as determined by the XTT metabolic assay. Surface-adherent bacteria were grown in the presence of β6-20, G3K6, or β6-20- G3K6 peptide (10-100 μM) and changes in the amount of surface-adherent sessile bacteria and free floating planktonic bacteria were monitored. Significance was determined by comparing each data point to the untreated control using one-way ANOVA with Dunnet’s post-hoc test (<0.01, n=3 for all data).

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The goal of the present study is to utilize a targeted cationic peptide in order to

disrupt the natural electrostatic interaction between S. epidermidis and polysaccharide

intercellular adhesin in an attempt to reduce biofilm formation. The work presented here

centers on the use of the fibrinogen-based 6-20 peptide, which is known to specifically

bind to the S. epidermidis surface. By incorporating a cationic sequence at the C-terminus

of the peptide, we have engineered a means of specifically delivering a positive charge to

the bacterial surface, with the potential to interfere with the process of biofilm formation.

Three peptides were prepared: the cationic targeting peptide 6-20-G3K6, as well as

the two individual components of the combination peptide, 6-20 and G3K6, which were

used as controls. Surface-adherent bacteria were exposed to these peptides and the effects

of the peptides on biofilm formation were assessed. As seen in Figure 5.1 (left), biofilms

grown in the presence of 50 μM peptide had similar growth curves to that of the 0 μM

control, indicating that the peptides are not toxic to the bacteria. In addition, biofilm

growth in the presence of 0-100 μM β6-20-G3K6 peptide (right) further indicated that this

particular peptide had minimal effects over a wider range of concentrations. Previous

studies have demonstrated that S. epidermidis is highly sensitive to the bactericidal

effects of both ε- and α-poly-L-lysine[40, 41], with a minimum of 10 lysine repeat units

necessary to show antimicrobial behavior[41]. The α-poly-L-lysine sequence used in

this study was 6 residues in length and did not alter bacterial growth curves, indicating

that this particular cationic sequence is not cytotoxic to S. epidermidis.

Wheat germ agglutinin conjugated to Alexa Fluor 647 (WGA-647) and SYTO 9 were used to assess the PIA content and the amount of surface-adherent bacteria present in biofilms (Figure 5.2). Neither the β6-20 nor the G3K6 peptide had any significant impact

185

on PIA or bacterial content of the biofilms over the range of peptide concentrations tested

(0-100 μM) when compared to biofilm controls. However, significant reductions were seen in both PIA content (35.9±6.2%, 37.9±6.0%) and adherent bacterial content

(17.3±5.8%, 17.5±5.2%) when grown in the presence of β6-20-G3K6 peptide

concentrations of 50 μM and 100 μM, respectively. These reductions indicate that the

targeting component plays a significant role in the effectiveness of the cationic peptide in

disrupting biofilm formation, since the β6-20 targeting peptide and the G3K6 cationic peptide did not elicit such reductions on their own. SEM and epifluorescent microscopy further support these findings, visually demonstrating that bacteria grown in the presence of 50 μM β6-20-G3K6 peptide failed to form mature biofilms.

While optical density data indicates that the β6-20-G3K6 peptide does not inhibit

bacterial growth, microplate fluorescence data indicates a significant reduction in biofilm

formation due to the peptide. In order to examine this apparent discrepancy, a partitioning

experiment was carried out using an XTT metabolic assay. After a 21 hour incubation in

the presence of 0-100 M, the culture media was carefully separated from the adherent

biofilm, and the XTT assay was used to elucidate the sessile vs. planktonic nature of each

biofilm. Increasing peptide concentrations did not alter the makeup of the biofilm grown

in the presence of the β6-20 and G3K6 peptides, with changes in the amount of sessile and planktonic bacteria statistically insignificant across the 0-100 μM peptide concentration

range. However, increasing concentration of the β6-20-G3K6 peptide showed a significant increase in planktonic bacteria (+28.2%), and a corresponding decrease in sessile bacteria

(-18.7%). This 18.7% reduction in sessile bacteria observed through the XTT assay agrees with the 17.5% reduction observed using the SYTO 9 assay. Thus, the β6-20-G3K6

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peptide significantly influenced whether or not the bacteria remained surface-adherent,

while having no statistical effect on the total amount of bacteria present in each well as

observed by OD growth analysis.

The technique presented here utilizes the β6-20-G3K6 peptide to target an existing

bacterial surface protein and manipulate the naturally occurring electrostatic interactions

between S. epidermidis and PIA. Most currently practiced preventative antibacterial

measures utilize antibiotics and other chemotherapeutic means to kill the bacteria, leading

to the distinct possibility of an adaptive response by the bacteria and the formation of a

resistant strain. The β6-20-G3K6 peptide, on the other hand, does not directly attack any

of the chemical or biological systems within the bacteria. Instead, the peptide acts in the

extracellular space to disrupt the biofilm formation process and lends itself to the

possibility of a synergistic relationship with the natural host response. Due to this passive

mode of action, the β6-20-G3K6 peptide will not suffer from long-term resistance issues more common with antibacterial treatments currently employed.

5.5 CONCLUSION:

The results from this study show that the β6-20-G3K6 peptide is able to disrupt the

process of biofilm formation. The G3K6 sequence on its own was shown to possess no native inhibition or cytotoxicity. However, the targeting ability provided by the 6-20

sequence imparted significant biofilm disrupting potential. As the β6-20-G3K6 concentration was increased, bacteria in a growing biofilm were shifted from a sessile to planktonic state. Fluorescent staining as well as SEM further confirmed these findings, demonstrating a significant reduction of PIA content and adherent bacterial within the

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biofilms. The data presented supports the hypothesis that targeted delivery of a positive charge to the surface of Staphylococcus epidermidis can interfere with PIA retention, which in turn hinders bacterial adhesion within the biofilm.

5.6 ACKNOWLEDGEMENTS:

The project described here was funded by Award

Number 5R01EB000279 from the National Institute of Biomedical Imaging and

Bioengineering, and work was carried out using the facilities of the Center for

Cardiovascular Biomaterials. I am extremely grateful for the assistance of Kyle

Bednar in completing much of this work. I would also like to thank Faina

Kligman for her assistance with MALDI-TOF analysis.

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5.7 REFERENCES

1. Otto M, Staphylococcus epidermidis ‐ the ʹaccidentalʹ pathogen. Nat Rev Micro,

2009. 7(8): p. 555‐567.

2. Rogers KL, Fey PD, and Rupp ME, Coagulase‐Negative Staphylococcal

Infections. Infectious Disease Clinics of North America, 2009. 23(1): p. 73‐

98.

3. Uçkay I, Pittet D, Vaudaux P, Sax H, Lew D, and Waldvogel F, Foreign

body infections due toStaphylococcus epidermidis. Annals of Medicine, 2009.

41(2): p. 109‐119.

4. Sabath LD, Garner C, Wilcox C, and Finland M, Susceptibility of

Staphylococcus aureus and Staphylococcus epidermidis to 65 antibiotics.

Antimicrob Agents Chemother, 1976. 9(6): p. 962‐9.

5. Cerca N, Jefferson KK, Maira‐Litran T, Pier DB, Kelly‐Quintos C,

Goldmann DA, Azeredo J, and Pier GB, Molecular basis for preferential

protective efficacy of antibodies directed to the poorly acetylated form of

staphylococcal poly‐N‐acetyl‐beta‐(1‐6)‐glucosaminev. Infection and

Immunity, 2007. 75(7): p. 3406‐3413.

6. Ceri H, Olson ME, Stremick C, Read RR, Morck D, and Buret A, The

Calgary Biofilm Device: New Technology for Rapid Determination of Antibiotic

189

Susceptibilities of Bacterial Biofilms. J. Clin. Microbiol., 1999. 37(6): p. 1771‐

1776.

7. Olson M, Ceri H, Morck D, Buret A, and Read R, Biofilm bacteria: formation

and comparative susceptibility to antibiotics. Can. J. Vet. Res., 2002. 66(2): p.

86‐92.

8. Darouiche RO, Treatment of Infections Associated with Surgical Implants. N

Engl J Med, 2004. 350(14): p. 1422‐1429.

9. OʹToole G, Kaplan HB, and Kolter R, Biofilm Formation as Microbial

Development. Annual Review of Microbiology, 2000. 54(1): p. 49‐79.

10. Bale MD, Wohlfahrt LA, Mosher DF, Tomasini B, and Sutton RC,

Identification of Vitronectin as a Major Plasma‐Protein Adsorbed on Polymer

Surfaces of Different Copolymer Composition. Blood, 1989. 74(8): p. 2698‐2706.

11. Chugh TD, Burns GJ, Shuhaiber HJ, and Bahr GM, Adherence Of

Staphylococcus‐Epidermidis To Fibrin‐Platelet Clots Invitro Mediated By

Lipoteichoic Acid. Infection And Immunity, 1990. 58(2): p. 315‐319.

12. Heilmann C, Hussain M, Peters G, and Gotz F, Evidence for autolysin‐

mediated primary attachment of Staphylococcus epidermidis to a polystyrene

surface. Mol Microbiol, 1997. 24(5): p. 1013‐24.

190

13. Hussain M, Heilmann C, Peters G, and Herrmann M, Teichoic acid enhances

adhesion of Staphylococcus epidermidis to immobilized fibronectin. Microbial

Pathogenesis, 2001. 31(6): p. 261‐270.

14. Li DQ, Lundberg F, and Ljungh A, Characterization of vitronectin‐binding

proteins of Staphylococcus epidermidis. Current Microbiology, 2001. 42(5): p.

361‐367.

15. Nilsson M, Frykberg L, Flock JI, Pei L, Lindberg M, and Guss B, A

fibrinogen‐binding protein of Staphylococcus epidermidis. Infection and

Immunity, 1998. 66(6): p. 2666‐2673.

16. Pei L, Arvholm IL, Lonnies L, and Flock JI, GST‐Fbe can recognize beta‐

chains of fibrin(ogen) on explanted materials. Journal of Chromatography B‐

Analytical Technologies in the Biomedical and Life Sciences, 2003. 786(1‐

2): p. 319‐325.

17. Pei L, Palma M, Nilsson M, Guss B, and Flock JI, Functional studies of a

fibrinogen binding protein from Staphylococcus epidermidis. Infection and

Immunity, 1999. 67(9): p. 4525‐4530.

18. Vuong C and Otto M, Staphylococcus epidermidis infections. Microbes Infect,

2002. 4(4): p. 481‐9.

191

19. Williams RJ, Henderson B, Sharp LJ, and Nair SP, Identification of a

fibronectin‐binding protein from Staphylococcus epidermidis. Infection And

Immunity, 2002. 70(12): p. 6805‐6810.

20. Patti JM, Allen BL, Mcgavin MJ, and Hook M, MSCRAMM‐Mediated

Adherence of Microorganisms to Host Tissues. Annual Review of

Microbiology, 1994. 48: p. 585‐617.

21. Patti JM and Hook M, Microbial Adhesins Recognizing Extracellular‐Matrix

Macromolecules. Current Opinion In Cell Biology, 1994. 6(5): p. 752‐758.

22. Davis SL, Gurusiddappa S, McCrea KW, Perkins S, and Hook M, SdrG, a

fibrinogen‐binding bacterial adhesin of the microbial surface components

recognizing adhesive matrix molecules subfamily from Staphylococcus

epidermidis, targets the thrombin cleavage site in the B beta chain. Journal of

Biological Chemistry, 2001. 276(30): p. 27799‐27805.

23. Watnick P and Kolter R, Biofilm, City of Microbes. J. Bacteriol., 2000. 182(10):

p. 2675‐2679.

24. Mack D, Fischer W, Krokotsch A, Leopold K, Hartmann R, Egge H, and

Laufs R, The intercellular adhesin involved in biofilm accumulation of

Staphylococcus epidermidis is a linear beta‐1,6‐linked glucosaminoglycan:

purification and structural analysis. J Bacteriol, 1996. 178(1): p. 175‐83.

192

25. Vuong C, A Crucial Role for Exopolysaccharide Modification in Bacterial

Biofilm Formation, Immune Evasion, and Virulence. Journal of Biological

Chemistry, 2004. 279(52): p. 54881‐54886.

26. Vuong C, Voyich JM, Fischer ER, Braughton KR, Whitney AR, DeLeo FR,

and Otto M, Polysaccharide intercellular adhesin (PIA) protects Staphylococcus

epidermidis against major components of the human innate immune system. Cell

Microbiol, 2004. 6(3): p. 269‐75.

27. Li H, Xu L, Wang J, Wen Y, Vuong C, Otto M, and Gao Q, Conversion of

Staphylococcus epidermidis Strains from Commensal to Invasive by Expression of

the ica Locus Encoding Production of Biofilm Exopolysaccharide. Infection and

Immunity, 2005. 73(5): p. 3188‐3191.

28. Rupp ME, Fey PD, Heilmann C, and Gotz F, Characterization of the

importance of Staphylococcus epidermidis autolysin and polysaccharide

intercellular adhesin in the pathogenesis of intravascular catheter‐associated

infection in a rat model. Journal of Infectious Diseases, 2001. 183(7): p. 1038‐

1042.

29. Rupp ME, Ulphani JS, Fey PD, Bartscht K, and Mack D, Characterization of

the importance of polysaccharide intercellular adhesin/hemagglutinin of

Staphylococcus epidermidis in the pathogenesis of biomaterial‐based infection in a

193

mouse foreign body infection model. Infection and Immunity, 1999. 67(5): p.

2627‐2632.

30. Rupp ME, Ulphani JS, Fey PD, and Mack D, Characterization of

Staphylococcus epidermidis polysaccharide intercellular adhesin/hemagglutinin

in the pathogenesis of intravascular catheter‐associated infection in a rat model.

Infection and Immunity, 1999. 67(5): p. 2656‐2659.

31. Mack D, Riedewald J, Rohde H, Magnus T, Feucht HH, Elsner HA, Laufs

R, and Rupp ME, Essential functional role of the polysaccharide intercellular

adhesin of Staphylococcus epidermidis in hemagglutination. Infection and

Immunity, 1999. 67(2): p. 1004‐1008.

32. Heilmann C, Schweitzer O, Gerke C, Vanittanakom N, Mack D, and Gotz

F, Molecular basis of intercellular adhesion in the biofilm‐forming Staphylococcus

epidermidis. Mol Microbiol, 1996. 20(5): p. 1083‐91.

33. Mack D, Nedelmann M, Krokotsch A, Schwarzkopf A, Heesemann J, and

Laufs R, Characterization of transposon mutants of biofilm‐producing

Staphylococcus epidermidis impaired in the accumulative phase of biofilm

production: genetic identification of a hexosamine‐containing polysaccharide

intercellular adhesin. Infect. Immun., 1994. 62(8): p. 3244‐3253.

34. Mack D, Siemssen N, and Laufs R, Parallel induction by glucose of adherence

and a polysaccharide antigen specific for plastic‐adherent Staphylococcus

194

epidermidis: evidence for functional relation to intercellular adhesion. Infect

Immun, 1992. 60(5): p. 2048‐57.

35. Trautner BW and Darouiche RO, Catheter‐Associated Infections: Pathogenesis

Affects Prevention. Arch Intern Med, 2004. 164(8): p. 842‐850.

36. Riddles PW, Blakeley RL, Zerner B, and C.H.W. Hirs SNT, [8] Reassessment

of Ellmanʹs reagent, in Methods in Enzymology. 1983, Academic Press. p. 49‐

60.

37. Curtin JJ and Donlan RM, Using Bacteriophages To Reduce Formation of

Catheter‐Associated Biofilms by Staphylococcus epidermidis. Antimicrobial

Agents and Chemotherapy, 2006. 50(4): p. 1268‐1275.

38. Chaw KC, Manimaran M, and Tay FEH, Role of Silver Ions in Destabilization

of Intermolecular Adhesion Forces Measured by Atomic Force Microscopy in

Staphylococcus epidermidis Biofilms. Antimicrob. Agents Chemother., 2005.

49(12): p. 4853‐4859.

39. Fields G and Noble R, Solid phase peptide synthesis utilizing 9‐

fluorenylmethoxycarbonyl amino acids. Int J Pept Protein Res, 1990. 35(3): p.

161‐214.

40. Delihas N, Riley LW, Loo W, Berkowitz J, and Poltoratskaia N, High‐

Sensitivity of Mycobacterium Species to the Bactericidal Activity by Polylysine.

Fems Microbiology Letters, 1995. 132(3): p. 233‐237.

195

41. Shima S, Matsuoka H, Iwamoto T, and Sakai H, Antimicrobial action of

epsilon‐poly‐L‐lysine. J Antibiot (Tokyo), 1984. 37(11): p. 1449‐55.

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CHAPTER 6

TARGETED DELIVERY OF VANCOMYCIN TO STAPHYLOCOCCUS EPIDERMIDIS BIOFILMS USING A FIBRINOGEN-DERIVED PEPTIDE

Based on: Hofmann CM, Anderson JM, Marchant RE. Targeted delivery of vancomycin to Staphylococcus epidermidis biofilms using a fibrinogen-derived peptide. J. Biomed Mater Res Part A, 2012; in press.

6.1 INTRODUCTION

There are approximately 40 million surgical procedures involving artificial

devices performed each year in the United States[1]. Complications arising from

nosocomial infections pose a significant health risk to patients with synthetic

implants. Staphylococci, enterococci, enterobacteriaceae, and Candida spp. are

the common pathogens associated with infections of indwelling medical

devices[2], with the likelihood of infection, as well as the organism implicated in the infection, greatly dependent upon the type and location of the implant[2]. In the case of intravascular implants, coagulase-negative staphylococci (CoNS), particularly Staphylococcus epidermidis, are the most common cause of infection[1,

3].

The initial stage of S. epidermidis infection of intravascular devices involves bacterial adhesion through interactions with host plasma proteins that adsorb onto the biomaterial surface immediately following implantation[3-12]. Specific binding between S. epidermidis and host proteins is commonly mediated by

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microbial surface components recognizing adhesive matrix molecules

(MSCRAMMs)[13, 14], with SdrG responsible for S. epidermidis binding to amino

-7 acids 6-20 of the fibrinogen B chain[9, 11, 15] with a KD of 0.9x10 M[15]. A

synthetic peptide sequence representing the first 25 amino acid residues of the B chain of fibrinogen (1-25) was used by Davis et al[15] to mimic the SdrG

-7 binding region, and this peptide bound to SdrG with a KD of 1.4x10 M. Thus,

the significant amino acids necessary for the SdrG-fibrinogen binding interaction

are present in the 1-25 peptide, and this linear sequence is in a nearly optimal

conformational state. The fibrinogen binding domain was further localized to

the 6-20 peptide, consisting of the amino acid sequence

NEEGFFSARGHRPLD[15], with amino acids 10-15 contributing most

significantly to the SdrG-fibrinogen binding[16]. The affinity of the 6-20 peptide

towards SdrG can be significantly reduced by creating substitution peptides,

such as 6-20-R14A and 6-20-G15A, in which individual amino acid residues

within the 10-15 binding region are replaced with an alanine residue[16].

Targeted therapy of bacterial infections has been studied in various forms.

Lytic bacteriophages represent a class of naturally occurring targeted

antibacterial treatments, with phages identified that can infect various

organisms, dissolve the exopolysaccharide matrix of existing biofilms, and in

some cases even reduce the need for clinical use of antibiotics[17]. Another

promising concept is the use of antibodies as opsonizing agents for colonizing

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bacteria. Opsonized S. epidermidis experience increased phagocytosis, resulting in a significantly lower incidence of infections in vivo[18-21]. However, targeted delivery of existing antibiotics remains a relatively unexplored area.

Vancomycin is a glycopeptide antibiotic that acts against gram positive bacteria by preventing the incorporation of N-acetylglucosamine and N- acetylmuramic acid into the growing peptidoglycan matrix[22]. Vancomycin forms a series of five hydrogen bonds with the terminal -D-Alanine-D-Alanine residues of the peptidoglycan intermediates, thereby interfering with the biosynthetic pathway used to build a stable cell wall[23]. The most common reason for Vancomycin resistance in a microbial population is a mutation that results in the peptidoglycan precursors terminating with -D-Alanine-D-Lactate, which eliminates one of the hydrogen bonds with vancomycin and lowers binding affinity by a factor on the order of 1000[24]. In order to overcome this resistance, glycopeptide antibiotics form non-covalent dimers that are thought to improve antibiotic effectiveness by preferentially locating the antibiotic at the site of action. After the first vancomycin molecule binds its target, the second binding event can be considered an intramolecular event that is more difficult to antagonize than a solution-phase bimolecular binding event[25]. Thus, facilitating the binding interaction between vancomycin and its ligand can lead to improved antibiotic treatment by recovering binding efficiency lost due to the

-D-Ala-D-Lac mutation.

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Given the importance and prevalence of the SdrG-fibrinogen binding scheme in the initial stages of S. epidermidis infections, this binding behavior should be explored as a mechanism for specifically delivering therapeutic agents to S. epidermidis infections. The experiments presented here were designed to mimic the benefits obtained by dimerization of glycopeptide antibiotics, while adding a specific targeting and delivery aspect to the system through use of the synthetic

6-20 peptide. By covalently linking vancomycin to the 6-20 peptide, we have developed a system to specifically deliver vancomycin to the S. epidermidis biofilm.

6.2 MATERIALS & METHODS:

Peptide Synthesis

(Ac)-CNEEGFFSARGHRPLD (Cys-β6-20) was synthesized by solid phase peptide synthesis on a 433a ABI peptide synthesizer (Applied Biosystems), using

9-fluoronylmethoxycarbonyl (fmoc) amino acids on a Knorr resin[26]. The Cys-

6-20 amino terminus was acetylated using acetic acid in a final automated coupling step on the synthesizer. The peptide was cleaved and deprotected using trifluoroacetic acid (TFA). Reverse phase high performance liquid chromatography (RP-HPLC) was used to purify the cleaved peptide, using water

(w/ 0.1% TFA) and acetonitrile (w/ 0.082% TFA) as the aqueous and organic

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components of the mobile phase. MALDI-TOF mass spectrometry was used to

confirm the purified peptide product was of the correct molecular weight.

Peptide-Vancomycin Conjugation

The 6-20-PEGx-Vancomycin (6-20-PEGx-VAN) conjugate was synthesized

using a Maleimide-PEGx-Succinimide heterofunctional crosslinker (x=3400 or

x=5000 MW). Vancomycin contains both a primary amine and a secondary

methylamine suitable for conjugation reactions, commonly described as the V3 and X1 positions, respectively (figure 6.1). In order to ensure that the PEG chain

attaches to vancomycin at the desired V3 position as opposed to the X1 position, a

two-step reaction was used (figure 6.2). The first step of the conjugation was

carried out as described by Greenwald et al[27]. Briefly, MAL-PEG3400-SCM or

MAL-PEG5000-SCM (Creative PEGWorks, Winston-Salem, NC) was added to a

two-fold excess of vancomycin hydrochloride (Sigma) in anhydrous

dimethylformamide (DMF) supplemented with a twenty-fold molar excess of triethylamine (TEA). The reaction was carried out under nitrogen, while stirring at room temperature for 12 hours, after which the mixture was added drop-wise

to cold diethyl ether to precipitate the product. The product was pelleted by

centrifugation, and rinsed an additional two times with fresh diethyl ether.

Excess vancomycin was removed from the crude MAL-PEGx-VAN product by

dialysis (3500 MWCO) against water adjusted to pH 5.0 with 1N HCl (in order to

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Figure 6.1: Chemical structure of vancomycin. Dashed circles indicate the potential locations for amine-targeted modifications. Reaction conditions can be tailored to favor the primary V3 amine over the secondary X1 methylamine.

202

-Vancomycin. Step 1 Combines x

ng formation of an amide bond at the 6-20-PEG 

-Vancomycin with Cys- 6-20 in PBS to create a x

argeting peptide and the PEG linker. amine. Step 2 combines MAL-PEG 3 -SCM with vancomycin in DMF/TEA, favori Two-step reaction scheme for synthesis of x vancomycin V Figure 6.2: MAL-PEG stable thioether bond between the t

203

prevent maleimide hydrolysis). The intermediate product was dried and residual vancomycin was quantified by RP-HPLC (C18 column). Residual, unconjugated

1 MAL-PEGx-SCM was estimated using H NMR.

To prepare the final β6-20-PEGx-VAN product, a 1.5 molar excess of purified

Cys-β6-20 peptide was added to MAL-PEGx-VAN in phosphate buffered saline

(pH 7.4, 1 mM EDTA) and stirred for 2 hours at room temperature. Ellman’s

assay was used to assess the extent of thiol disappearance in the reaction

mixture, corresponding to formation of the desired thioether bond. Dialysis

against water (3500 MWCO) was then used to remove excess peptide. Residual

peptide and vancomycin were quantified by RP-HPLC (C18 column).

Peptide:PEG:Vancomycin ratios were estimated using 1H NMR.

Determination of Minimum Inhibitory Concentration (MIC) and Minimum Bactericidal

Concentration (MBC)

The broth dilution method was used to determine the MIC and MBC of the newly synthesized targeted antibiotics. Tryptic soy broth (TSB, 1 ml) containing

5 S. epidermidis (1x10 cfu/ml) and vancomycin, 6-20-PEG3400-VAN, or 6-20-

PEG3400-VAN (0-128 g/ml equivalent concentrations) was prepared. The

samples (n=3) were then incubated for 24 hours at37°C with gentle shaking, after which the turbidity of the vials was observed. The MIC was taken as the lowest antibiotic concentration at which the TSB was no longer clear. In order to

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determine the MBC, 10 l of the media from each tube was plated on a trypticase

soy agar plate and incubated for an additional 24 hours at 37°C. The MBC was

taken as the lowest antibiotic concentration at which no colonies were observed.

Seeding of Sample Surfaces

S. epidermidis strain RP62A was used for all procedures as a representative

biofilm-positive strain. An inoculating loop was used to retrieve a small sample

of S. epidermidis from a refrigerated culture plate not more than two weeks old.

The bacteria were incubated in tryptic soy broth (TSB) for 18 hours at 37ºC while

shaking at 120 rpm, then pelleted and washed with fresh PBS before being

resuspended in TSB to a final concentration of 0.5x108 cfu/ml, determined by

optical density readings taken at 550nm. Bacteria solution was then added to 96-

well plates (150 L) and incubated at 37ºC for 2 or 24 hours.

Vancomycin Targeting

In order to examine targeted delivery of vancomycin, 96-well plates were seeded and incubated for 2 or 24 hours, after which the sample surfaces were gently rinsed with PBS to remove non-adherent bacteria. 6-20-PEGx-VAN or free vancomycin (150 μl, 0-86.2 μM in TSB) was added to the wells and incubated at 37°C for 1 or 24 hours. All wells were then rinsed twice with fresh PBS, followed by an additional incubation period in TSB at 37°C (0 or 24 hours). The

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vancomycin-targeted samples were then analyzed for vancomycin retention,

biofilm composition, and bacterial viability.

Vancomycin Retention Assay

Vancomycin-targeted samples were rinsed with 0.5% BSA, followed by the

addition of an anti-vancomycin primary antibody (100 μl in 3% BSA, 5 g/ml,

rabbit IgG, Abcam) for 60 minutes at room temperature. All samples were then

rinsed 3 times with fresh 0.5% BSA, followed by the addition of a secondary

antibody conjugated to horseradish peroxidase (HRP, 100 μl, 1:1000 dilution,

goat IgG anti-rabbit IgG, Molecular Probes) for an additional 60 minutes at room

temperature, and then a final rinse with PBS. ABTS (2,2’-Azino-bis(3-

ethylbenzothiazoline-6-sulfonic acid diammonium salt) was then added as an

HRP substrate (200 l, 0.1 mg/ml ABTS, 1 l/ml hydrogen peroxide, 50 mM

phosphate-citrate buffer, pH 5.0). After 20 minutes, 100 l was transferred from

each well into a new 96-well plate, and the wells were examined quantitatively for vancomycin content using a microplate reader (412 nm).

Statistics

Statistical analysis was done using Minitab 16. Comparisons between individual samples were made using an un-paired student’s t-test, while

206

comparisons within groups were made using one-way ANOVA and Dunnett’s

post-hoc test. α < 0.05 was considered significant.

6.3 RESULTS:

Synthesis of the 6-20-PEG3400-VAN and 6-20-PEG5000-VAN products was

1 monitored for purity by RP-HPLC and H NMR in D2O (data not shown). RP-

HPLC showed that the intermediate MAL-PEG3400-VAN and MAL-PEG5000-VAN

products contained 1.27% (w/w) and 0.71% (w/w) residual vancomycin,

respectively. In addition, 1H NMR characterization of the maleimide hydrogens

(=6.8 ppm, 2H) and vancomycin methyl hydrogens (=0.9 ppm, 6H) indicated

that the intermediate products were approximately 70 mol% MAL-PEGx-VAN and 30 mol% unreacted MAL-PEGx-COOH. After the Cys-6-20 + MAL-PEGx-

VAN conjugation step, RP-HPLC of the final 6-20-PEG3400-VAN and 6-20-

PEG5000-VAN products indicated no residual peptide, and showed 0.13% (w/w)

and 0.08% (w/w) residual vancomycin, respectively. The amounts of residual vancomycin and unreacted PEG were taken in to account when weighing out the final products for retention studies, and as such, the term equivalent vancomycin concentration is used when referring to these products.

Minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) were measured for the new targeted vancomycin derivatives (table 6.1). Vancomycin alone showed an MIC of 2 g/ml and an

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MIC (g/ml) MBC (g/ml) Vancomycin 2 8

‐PEG3400‐VAN 16 32

‐PEG5000‐VAN 32 64

Table 6.1: Minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) values for vancomycin, 6-20-PEG3400-VAN, or 6-20-

PEG5000-VAN.

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MBC of 8 g/ml. However, increases in MIC and MBC were seen for both 6-20-

PEG3400-VAN and 6-20-PEG5000-VAN. The 6-20-PEG3400-VAN product showed

MIC and MBC values of 16 g/ml and 32 g/ml, respectively. The 6-20-

PEG5000-VAN product showed MIC and MBC values double those seen with the

6-20-PEG3400-VAN product, at 32 g/ml and 64 g/ml, respectively.

The 6-20-PEGx-VAN products were tested for their ability to target S.

epidermidis and improve vancomycin retention. Retention was first measured on

S. epidermidis in the early stages of biofilm formation by allowing the bacteria to

adhere to 96-well plates for 2 hours prior to exposure to antibiotics (1 hour

exposure, 0 g/ml to 64 g/ml vancomycin equivalents). All data (n=3) was

normalized to the 0 g/ml control within each plate. Vancomycin showed no

sigificant retention (p=0.48) at any concentration tested when compared to the 0

g/ml control. Both 6-20-PEG3400-VAN and 6-20-PEG5000-VAN showed

significantly increased retention (p<0.001) relative to vancomycin for all

equivalent concentrations ≥4 g/ml (Figure 6.3a), with retention increasing in a concentration-dependent manner. Maximum retention was observed at 64

g/ml, with a 22.9-fold increase for 6-20-PEG3400-VAN and a 21.1-fold increase

for 6-20-PEG5000-VAN. For equivalent concentrations <4 g/ml, the targeted

antibiotics showed no difference in retention when compared to vancomycin.

Following the initial 1 hour exposure, all samples were incubated for an

additional 24 hours before re-measuring the amount of antibiotics retained in the

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(a) (b) 25.0 2.0 Vancomycin VancomycinVancomycin

b66‐‐2020‐PEG3.4k‐PEG3400‐VAN‐Vancomycin b6‐620‐20‐PEG3.4k‐PEG3400‐VAN‐Vancomycin 20.0 b66‐‐2020‐PEG5k‐PEG5000‐VAN‐Vancomycin 1.5 b6‐620‐20‐PEG5k‐PEG‐VAN5000‐Vancomycin

15.0 1.0 Vancomycin Vancomycin

10.0

0.5 Relative Relative 5.0

0 1 2 4 8 16 32 64 0 1 2 4 8 16 32 64 Antibiotic Concentration (g/ml) Antibiotic Concentration (g/ml) (c) 2.0 VancomycinVancomycin b66‐20‐20‐PEG3.4k‐PEG3400‐VAN‐Vancomycin 1.5 b66‐20‐20‐PEG5k‐PEG‐5000VAN‐Vancomycin

1.0 Vancomycin

0.5 Relative

0 1 2 4 8 16 32 64 Antibiotic Concentration (g/ml)

Figure 6.3: Retention of 6-20-PEG3400-VAN, 6-20-PEG5000-VAN, and vancomycin

by S. epidermidis after (a) 2 hour bacterial adhesion, 1 hour antibiotic exposure, (b) 24

hour biofilm growth, 1 hour antibiotic exposure, and (c) 24 hour biofilm growth, 24

hour antibiotic exposure. All data (n=3) are normalized to the 0 g/ml concentration

210

biofilms. Samples that had been treated with 32 g/ml and 64 g/ml

vancomycin equivalents continued to show significant improvements, with

retention of 1.3- and 1.8-fold for 6-20-PEG3400-VAN and 1.3- and 1.6-fold for 6-

20-PEG5000-VAN (p<0.001, data not shown). For equivalent vancomycin

concentrations ≤16 g/ml, there were no differences in retained vancomycin

between any of the samples.

Retention was also measured with more mature biofilms grown for 24 hours

prior to exposure to antibiotics (1 hour exposure, 0 g/ml to 64 g/ml

vancomycin equivalents). Vancomycin showed no sigificant retention (p=0.16) in

the biofilm at any concentration tested when compared to the 0 g/ml control.

6-20-PEG3400-VAN showed improved retention (p<0.001) relative to

vancomycin for equivalent concentrations ≥16 g/ml, while 6-20-PEG5000-VAN showed improved retention (p<0.001) for equivalent concentrations ≥32 g/ml

(Figure 6.3b). Retention ranged from 1.2-1.4-fold increases for both 6-20-

PEG3400-VAN and 6-20-PEG5000-VAN. Following the initial 1 hour exposure, all wells were incubated for an additional 24 hours before re-measuring the amount of antibiotics retained in the biofilms. There were no differences in retained vancomycin between any of the samples at this time point (p=0.79, data not shown).

Finally, retention was measured using 24 hour biofilms with a longer period of exposure to antibiotics (24 hour exposure, 0 g/ml to 64 g/ml vancomycin

211

equivalents). Vancomycin showed no sigificant retention in the biofilm at any

concentration tested when compared to the 0 g/ml control, with the exception of 4 g/ml (p=0.01). Both 6-20-PEG3400-VAN and 6-20-PEG5000-VAN showed

significant increases in retention (p<0.001) relative to vancomycin for equivalent

concentrations ≥32 g/ml (Figure 6.3c). Retention ranged from 1.3-1.6-fold

increases for 6-20-PEG3400-VAN and 1.2-1.4-fold increases for 6-20-PEG5000-

VAN. Following the initial 24 hour exposure, all wells were incubated for an

additional 24 hours before re-measuring the amount of antibiotics retained in the

biofilms (data not shown). 6-20-PEG3400-VAN showed 1.3- and 1.4-fold

increases in retention (p=0.01) for concentrations of 32 g/ml and 64 g/ml,

while 6-20-PEG5000-VAN showed a 1.3-fold increase in retention at 32 g/ml

(p=0.01).

6.4 DISCUSSION:

Staphylococcus epidermidis is part of the normal human flora and considered to

be noninvasive and nonpathogenic under most circumstances. However,

implantation of a medical device provides a foreign surface that can be colonized

by bacteria that may be unintentionally introduced during the procedure. The

SdrG-fibrinogen interaction has been shown to play a significant role in the

initial stages of bacterial adhesion: recombinant SdrG is able to block the

adherence of S. epidermidis to fibrinogen in a concentration dependent

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manner[28], adding SdrG under saturating conditions prevents S. epidermidis

from binding to fibrinogen[11], and mutating the fibrinogen-binding region of

the SdrG A-domain of S. epidermidis greatly impairs the ability to bind to

immobilized fibrinogen.[16, 29]. Furthermore, both the gene encoding for

SdrG[30] and the SdrG protein itself[31] are present in 100% of the various

clinical S. epidermidis isolates tested, and studies of infected and recovering

patients reveal anti-SdrG antibodies in the serum, confirming that the bacteria express SdrG during the infectious stages[20, 31]. An in-vivo intravascular

catheter infection model confirmed the importance of SdrG: an SdrG-negative S.

epidermidis mutant had a 20% infection rate, compared to a 100% rate for the

SdrG-positive strain[32].

The success of S. epidermidis infections depends upon the ability of the bacteria to form a biofilm following initial adhesion to a material[3, 33]. The

biofilm environment offers the encapsulated bacteria increased resistance to

antibiotics, often times able to survive antibiotic concentrations several orders of

magnitude higher than the minimum inhibitory concentration (MIC) and

minimum bactericidal concentration (MBC) measured in planktonic

suspensions[34-36]. However, after dispersing the cells in a biofilm, the newly

suspended bacteria are once again susceptible to the same MIC and MBC levels

found in planktonic bacteria[37]. Furthermore, S. epidermidis biofilms formed on

dialysis membranes allow vancomycin to penetrate the biofilm and cross the

213

membrane[38], while biofilms formed on stainless steel prostheses have been

shown to uptake significant quantities of vancomycin[37]. It is likely that

antibiotic resistance within a biofilm is not obtained through irreversible

genotypic changes, and that biofilm penetration is not a limiting factor in

antibiotic effectiveness. As such, it is possible that increasing the binding

capabilities of vancomycin through use of a targeting ligand could prove

beneficial in treating infections.

Covalently tethering vancomycin to a synthetic targeting ligand yields

significant increases in antibiotic retention by the S. epidermidis biofilm.

Interestingly, the amount of 6-20-PEG3400-VAN product retained by the S. epidermidis samples was slightly greater than the amount of 6-20-PEG5000-VAN

retained under most conditions tested. One likely reason for this difference is the

potential for the longer PEG chain to obstruct the binding interactions between

peptide and bacteria. In addition, it is also possible that the higher molecular

weight of the 6-20-PEG5000-VAN product decreases the speed at which the

molecule can penetrate the growing biofilm. These findings correlate well with

the MIC and MBC values observed for the new targeted vancomycin derivatives.

While both 6-20-PEG3400-VAN and 6-20-PEG5000-VAN were shown to be active

against S. epidermidis, the MIC and MBC values for 6-20-PEG5000-VAN product

were twice as high as those observed for 6-20-PEG3400-VAN.

214

Within the typical clinical range for vancomycin concentrations (figure 6.4),

initial retention of the targeted antibiotics after 1 hour or 24 hour exposure was

20-30% greater than retention of vancomycin alone. The amount of vancomycin

retained by the mature biofilms after an additional 24 hour incubation period

depended upon the initial duration of vancomycin exposure prior to the rinsing

step. The biofilms that were treated for just 1 hour ended up with targeted

vancomycin levels that were not significantly different from those of vancomycin alone. A potential explanation for this inability to retain the antibiotics is due to the larger size of the targeted vancomycin: 1 hour may not be enough time for such a large molecule to fully penetrate a mature biofilm, and as such, there was no real advantage over vancomycin. However, when the mature biofilms were exposed to the targeted vancomycin for a full 24 hours prior to the final incubation step, there were negligible changes in retained targeted vancomycin when comparing initial and final values, with final retention 27-34% higher than seen with vancomycin alone.

When surface adherent bacteria are exposed to the targeted vancomycin prior to biofilm formation, the targeting peptide significantly improves the vancomycin retention, with 22.1-22.9-fold increases observed after just a 1 hour exposure. However, as the biofilm develops, it is likely that the size difference between vancomycin and the two targeted species becomes more of a factor,

215

2 6‐20‐PEG3400‐Vancomycin 2 6‐20‐PEG5000‐Vancomycin 0 hr Retention 0 hr Retention 24 hr retention 24 hr Retention vancomycin **vancomycin * * Retention Retention

* *

1 1

Antibiotic Antibiotic

Relative Relative 0 0 1 hour 24 hour 1 hour 24 hour Antibiotic Exposure Time Antibiotic Exposure Time

Figure 6.4: Retention of targeted antibiotics by 24 hour S. epidermidis biofilms as determined by indirect ELISA. S. epidermidis was exposed to vancomycin, 6-20-

PEG3400-VAN, or 6-20-PEG5000-VAN (32 g/ml vancomycin equivalents) for 1 hour or 24 hours. Vancomycin levels were measured 0 and 24 hours post- treatment. All data (n=3) are reported as increases in retention relative to untargeted vancomycin, with (*) indicating a statistically significant increase

(<0.05).

216

resulting in the observed decrease in retention of the targeted molecules relative to the untargeted vancomycin. This becomes most evident when the biofilm is allowed to mature prior to being exposed to the targeted vancomycin molecules.

Extending the duration of exposure to the targeted vancomycin allows more time for the larger molecule to penetrate the biofilm, which in turn results in a higher relative retention.

It is expected that increased retention within the biofilm will provide benefits analogous to those seen with vancomycin dimerization. Dimerization of glycopeptide antibiotics such as Vancomycin occurs naturally, improving effectiveness of the antibiotic by preferentially locating the antibiotic at the site of action where subsequent binding interactions are thought to be more of an intramolecular event[25]. Dimerization has been shown to improve antibiotic effectiveness in a number of ways. A series of 40 covalently linked vancomycin dimers were synthesized, many of which displayed improved effectiveness against both vancomycin-susceptible organisms and vancomycin-resistant

Enterococci (VRE)[39]. Meanwhile, a multi-valent polymer of vancomycin demonstrated enhanced antibacterial activity against VRE that was 8- to 60-fold more effective than traditional vancomycin. While vancomycin binds to -D-Ala-

-6 D-Ala with a KD of ~1x10 M, a trivalent system of vancomycin binds a trivalent

-17 -D-Ala-D-Ala ligand with a KD of ~4x10 , which is 25 times tighter than biotin- avidin[23]. Finally, vancomycin covalently tethered to a titanium surface

217

maintained its ability to bind -D-Ala-D-Ala as well as kill S. aureus, and indicated

that a high surface density of antibiotic is equivalent to an enormous solution

concentration of free vancomycin[40]. Similar results can be seen with the

antibiotic teicoplanin, which does not form dimers but instead anchors to the

bacterial membrane by way of its fatty acid chain. Teicoplanin, once anchored to

a surface, functions intramolecularly and displays improved effectiveness when

compared to binding events in the absence of a membrane[25, 41].

6.5 CONCLUSION:

The Fibrinogen-SdrG interaction is an important component of bacterial

adhesion to blood-contacting biomaterials. The covalently tethered antibiotic can

be used for localized treatment with a higher retention rate within the biofilm.

Significant improvements in initial binding of vancomycin to S. epidermidis were obtained utilizing this targeted vancomycin scheme, and 27-34% improvements in retention rates were achieved. If successfully developed, a similar technique could be applied to target and treat infections of other bacterial species.

6.6 ACKNOWLEDGEMENTS:

The project described here was funded by Award

Number 5R01EB000279 from the National Institute of Biomedical Imaging and

Bioengineering, and work was carried out using the facilities of the Center for

Cardiovascular Biomaterials.

218

6.7 REFERENCES

1. Darouiche RabihÂ O, Device‐Associated Infections: A Macroproblem that

Starts with Microadherence. Clinical Infectious Diseases, 2001. 33(9): p. 1567‐

1572.

2. Schierholz JM and Beuth J, Implant infections: a haven for opportunistic

bacteria. Journal of Hospital Infection, 2001. 49(2): p. 87‐93.

3. Vuong C and Otto M, Staphylococcus epidermidis infections. Microbes Infect,

2002. 4(4): p. 481‐9.

4. Bale MD, Wohlfahrt LA, Mosher DF, Tomasini B, and Sutton RC,

Identification of Vitronectin as a Major Plasma‐Protein Adsorbed on Polymer

Surfaces of Different Copolymer Composition. Blood, 1989. 74(8): p. 2698‐2706.

5. Chugh TD, Burns GJ, Shuhaiber HJ, and Bahr GM, Adherence Of

Staphylococcus‐Epidermidis To Fibrin‐Platelet Clots Invitro Mediated By

Lipoteichoic Acid. Infection And Immunity, 1990. 58(2): p. 315‐319.

6. Heilmann C, Hussain M, Peters G, and Gotz F, Evidence for autolysin‐

mediated primary attachment of Staphylococcus epidermidis to a polystyrene

surface. Molecular Microbiology, 1997. 24(5): p. 1013‐1024.

219

7. Hussain M, Heilmann C, Peters G, and Herrmann M, Teichoic acid enhances

adhesion of Staphylococcus epidermidis to immobilized fibronectin. Microbial

Pathogenesis, 2001. 31(6): p. 261‐270.

8. Li DQ, Lundberg F, and Ljungh A, Characterization of vitronectin‐binding

proteins of Staphylococcus epidermidis. Current Microbiology, 2001. 42(5): p.

361‐367.

9. Nilsson M, Frykberg L, Flock JI, Pei L, Lindberg M, and Guss B, A

fibrinogen‐binding protein of Staphylococcus epidermidis. Infection and

Immunity, 1998. 66(6): p. 2666‐2673.

10. Pei L, Arvholm IL, Lonnies L, and Flock JI, GST‐Fbe can recognize beta‐

chains of fibrin(ogen) on explanted materials. Journal of Chromatography B‐

Analytical Technologies in the Biomedical and Life Sciences, 2003. 786(1‐

2): p. 319‐325.

11. Pei L, Palma M, Nilsson M, Guss B, and Flock JI, Functional studies of a

fibrinogen binding protein from Staphylococcus epidermidis. Infection and

Immunity, 1999. 67(9): p. 4525‐4530.

12. Williams RJ, Henderson B, Sharp LJ, and Nair SP, Identification of a

fibronectin‐binding protein from Staphylococcus epidermidis. Infection And

Immunity, 2002. 70(12): p. 6805‐6810.

220

13. Patti JM, Allen BL, Mcgavin MJ, and Hook M, MSCRAMM‐Mediated

Adherence of Microorganisms to Host Tissues. Annual Review of

Microbiology, 1994. 48: p. 585‐617.

14. Patti JM and Hook M, Microbial Adhesins Recognizing Extracellular‐Matrix

Macromolecules. Current Opinion In Cell Biology, 1994. 6(5): p. 752‐758.

15. Davis SL, Gurusiddappa S, McCrea KW, Perkins S, and Hook M, SdrG, a

fibrinogen‐binding bacterial adhesin of the microbial surface components

recognizing adhesive matrix molecules subfamily from Staphylococcus

epidermidis, targets the thrombin cleavage site in the B beta chain. Journal of

Biological Chemistry, 2001. 276(30): p. 27799‐27805.

16. Ponnuraj K, Bowden MG, Davis S, Gurusiddappa S, Moore D, Choe D, Xu

Y, Hook M, and Narayana SVL, A ʺdock, lock, and latchʺ structural model for

a staphylococcal adhesin binding to fibrinogen. Cell, 2003. 115(2): p. 217‐228.

17. Donlan RM, Preventing biofilms of clinically relevant organisms using

bacteriophage. Trends in microbiology, 2009. 17(2): p. 66‐72.

18. Bloom B, Schelonka R, Kueser T, Walker W, Jung E, Kaufman D, Kesler K,

Roberson D, Patti J, and Hetherington S, Multicenter study to assess safety

and efficacy of INH‐A21, a donor‐selected human staphylococcal

immunoglobulin, for prevention of nosocomial infections in very low birth weight

infants. The Pediatric infectious disease journal, 2005. 24(10): p. 858.

221

19. Ohlsen K and Lorenz U, Immunotherapeutic strategies to combat

staphylococcal infections. International Journal of Medical Microbiology,

2010. 300(6): p. 402‐410.

20. Rennermalm A, Nilsson M, and Flock JI, Fibrinogen binding protein of

Staphylococcus epidermidis is a target for opsonic antibodies. Infection and

Immunity, 2004. 72(5): p. 3081‐3083.

21. Vernachio JH, Bayer AS, Ames B, Bryant D, Prater BD, Syribeys PJ,

Gorovits EL, and Patti JM, Human immunoglobulin G recognizing fibrinogen‐

binding surface proteins is protective against both Staphylococcus aureus and

Staphylococcus epidermidis infections in vivo. Antimicrobial Agents and

Chemotherapy, 2006. 50(2): p. 511‐518.

22. Watanakunakorn C, The antibacterial action of vancomycin. Rev Infect Dis,

1981. 3 suppl: p. S210‐5.

23. Arimoto H, Nishimura K, Kiniumi T, Hayakawa I, and Uemura D, Multi‐

valent polymer of vancomycin ‐ enhanced antibacterial activity against VRE.

Chem. Commun., 1999(15): p. 1361‐1362.

24. Williams DH, The glycopeptide story‐‐how to kill the deadly ʹsuperbugsʹ. Nat

Prod Rep, 1996. 13(6): p. 469‐77.

222

25. Beauregard DA, Williams DH, Gwynn MN, and Knowles DJ, Dimerization

and membrane anchors in extracellular targeting of vancomycin group

antibiotics. Antimicrob. Agents Chemother., 1995. 39(3): p. 781‐785.

26. Fields G and Noble R, Solid phase peptide synthesis utilizing 9‐

fluorenylmethoxycarbonyl amino acids. Int J Pept Protein Res, 1990. 35(3): p.

161‐214.

27. Greenwald RB, Zhao H, Xia J, and Martinez A, Poly (ethylene glycol)

transport forms of vancomycin: a long‐lived continuous release delivery system.

Journal of medicinal chemistry, 2003. 46(23): p. 5021‐5030.

28. Hartford O, OʹBrien L, Schofield K, Wells J, and Foster TJ, The Fbe (SdrG)

protein of Staphylococcus epidermidis HB promotes bacterial adherence to

fibrinogen. Microbiology‐Sgm, 2001. 147: p. 2545‐2552.

29. Pei L and Flock JI, Lack of fbe, the gene for a fibrinogen‐binding protein from

Staphylococcus epidermidis, reduces its adherence to fibrinogen coated surfaces.

Microbial Pathogenesis, 2001. 31(4): p. 185‐193.

30. Bowden MG, Chen W, Singvall J, Xu Y, Peacock SJ, Valtulina V, Speziale

P, and Hook M, Identification and preliminary characterization of cell‐wall‐

anchored proteins of Staphylococcus epidermidis. Microbiology‐Sgm, 2005. 151:

p. 1453‐1464.

223

31. McCrea KW, Hartford O, Davis S, Eidhin DN, Lina G, Speziale P, Foster

TJ, and Hook M, The serine‐aspartate repeat (Sdr) protein family in

Staphylococcus epidermidis. Microbiology‐Uk, 2000. 146: p. 1535‐1546.

32. Guo BN, Zhao X, Shi YG, Zhu DM, and Zhang YY, Pathogenic implication of

a fibrinogen‐binding protein of Staphylococcus epidermidis in a rat model of

intravascular‐catheter‐associated infection. Infection and Immunity, 2007.

75(6): p. 2991‐2995.

33. Queck SY and Otto M, Staphylococcus epidermidis and other Coagulase‐

Negative Staphylococci, in Staphylococcus Molecular Genetics, J.A. Lindsay,

Editor. 2008, Caister Academic Press: Norfolk, UK.

34. Cerca N, Jefferson KK, Maira‐Litran T, Pier DB, Kelly‐Quintos C,

Goldmann DA, Azeredo J, and Pier GB, Molecular basis for preferential

protective efficacy of antibodies directed to the poorly acetylated form of

staphylococcal poly‐N‐acetyl‐beta‐(1‐6)‐glucosaminev. Infection and

Immunity, 2007. 75(7): p. 3406‐3413.

35. Ceri H, Olson ME, Stremick C, Read RR, Morck D, and Buret A, The

Calgary Biofilm Device: New Technology for Rapid Determination of Antibiotic

Susceptibilities of Bacterial Biofilms. J. Clin. Microbiol., 1999. 37(6): p. 1771‐

1776.

224

36. Olson M, Ceri H, Morck D, Buret A, and Read R, Biofilm bacteria: formation

and comparative susceptibility to antibiotics. Can. J. Vet. Res., 2002. 66(2): p.

86‐92.

37. Darouiche RO, Dhir A, Miller AJ, Landon GC, Raad, II, and Musher DM,

Vancomycin penetration into biofilm covering infected prostheses and effect on

bacteria. J Infect Dis, 1994. 170(3): p. 720‐3.

38. Dunne WM, Mason E, and Kaplan S, Diffusion of Rifampin and Vancomycin

through a Staphylococcus epidermidis Biofilm. Antimicrobial Agents and

Chemotherapy, 1993. 37(12): p. 2522‐2526.

39. Griffin JH, Linsell MS, Nodwell MB, Chen Q, Pace JL, Quast KL, Krause

KM, Farrington L, Wu TX, Higgins DL, Jenkins TE, Christensen BG, and

Judice JK, Multivalent Drug Design. Synthesis and In Vitro Analysis of an

Array of Vancomycin Dimers. Journal of the American Chemical Society,

2003. 125(21): p. 6517‐6531.

40. Jose B, Antoci Jr V, Zeiger AR, Wickstrom E, and Hickok NJ, Vancomycin

Covalently Bonded to Titanium Beads Kills Staphylococcus aureus. Chemistry

& Biology, 2005. 12(9): p. 1041‐1048.

41. Westwell MS, Bardsley B, Dancer RJ, Try AC, and Williams DH,

Cooperativity in ligand binding expressed at a model cell membrane by the

225

vancomycin group antibiotics. Chemical Communications, 1996(5): p. 589‐

590.

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CHAPTER 7

CONCLUSIONS AND PERSPECTIVES

5.1 SUMMARY OF COMPLETED WORK:

This project investigated the hypothesis that a biofilm targeting system

utilizing the 6-20 targeting peptide can be developed for disruption and treatment of Staphylococcus epidermidis biofilms. S. epidermidis adheres to

intravascular biomaterials through a number of mechanisms, including a specific

binding event between the S. epidermidis surface protein SdrG and the amino acid

sequence NEEGFFSARGHRPLD (6-20) of the human plasma protein fibrinogen.

In Chapter 4, the 6-20 peptide sequence NEEGFFSARGHRPLD was used to

target biomaterial-adherent Staphylococcus epidermidis under in vitro conditions.

Specificity was demonstrated by both BSA and peptide blocking studies. By

selectively blocking SdrG binding sites with unlabeled peptide, the labeled

peptide was unable to bind to the bacteria. In addition, by coating the samples

with BSA, nonspecific binding to the underlying PET substrate was greatly

reduced and specific binding to S. epidermidis was unaffected. In this study,

nanogold particles are used primarily as a tool for observing targeted binding.

However, in delivering the nanogold to the bacterial surface, the 6-20 peptide

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was identified as a viable targeting motif for delivering antibacterial agents to S. epidermidis infections. Subsequent chapters investigated this possibility.

In Chapter 5, the β6-20 peptide was modified to incorporate a positively charged poly(lysine) sequence. This resulting β6-20-G3K6 peptide was able to disrupt the process of biofilm formation. The G3K6 sequence on its own was shown to possess no native inhibition or cytotoxicity. However, the targeting ability provided by the 6-20 sequence imparted significant biofilm disrupting potential. As the β6-20-G3K6 concentration was increased, bacteria in a growing biofilm were shifted from a sessile to planktonic state.

The data presented supports the hypothesis that targeted delivery of a positive charge to the surface of Staphylococcus epidermidis can interfere with PIA retention, which in turn hinders bacterial adhesion within the biofilm.

In Chapter 6, vancomycin was modified so as to incorporate the β6-20 targeting peptide. A flexible PEG linker was used between the peptide and antibiotic, allowing vancomycin to be tethered to S. epidermidis. Results indicated that the targeted vancomycin can be used for localized treatment with a higher retention rate within the biofilm. Significant improvements in initial binding of vancomycin to

S. epidermidis were obtained utilizing this targeted vancomycin scheme, and 27-

34% improvements in retention rates were achieved. If successfully developed, a similar technique could be applied to target and treat infections of other bacterial species.

Targeted therapy of bacterial infections has been studied in various forms.

Lytic bacteriophages represent a class of naturally occurring targeted

228

antibacterial treatments[1], while the use of antibodies as opsonizing agents

results in a significantly lower incidence of infection in vivo[2-5]. This project

strived to develop novel means of antibacterial treatment through the use of a

fibrinogen-derived targeting peptide. The remainder of this chapter will outline

important findings uncovered in the process of synthesizing the targeted

vancomycin derivatives, in addition to highlighting future directions for such

work.

7.2 TARGETED VANCOMYCIN: SYNTHESIS CONSIDERATIONS

Targeted delivery of existing antibiotics remains a relatively unexplored area.

With a wide selection of current antibiotics being slowly rendered ineffective due

to acquired resistance, the work presented in Chapter 6 strived to lay the

groundwork for overcoming the problem of resistance. Several issues and

complications regarding the synthesis of such targeted vancomycin derivatives will be presented below. While specific to the work carried out in this project, the general conclusions should be taken in to consideration for development of any future targeted antibiotics.

The first main variable examined in this work was crosslink position.

Vancomycin contains both a primary amine and a secondary methylamine suitable for conjugation reactions, commonly described as the V3 and X1 positions, respectively. In theory the primary V3 amine should be more reactive

229

than the secondary X1 amine, lending itself to a simple succinimide (NHS)

coupling step in water or dimethyl sulfoxide (DMSO). Such assumptions were

made by Lawson et al[6, 7] in the synthesis of polymerizeable vancomycin derivatives using acrylate-PEG-NHS in DMSO. However, experimental data collected over the course of this project indicates that NHS reactions with

vancomycin carried out in aqueous environments yield a heterogeneous mixture

of V3- and X1-linked products that was not easily purified.

Figure 7.1 depicts the MALDI-TOF spectrum of vancomycin. In addition to

the main product peak (1448 m/z), two fragment products are typically observed

due to the vancosamine (M-117 m/z) and glucosamine (M-305 m/z) sugar

residues being sequentially cleaved by the instrument’s laser. After coupling SM-

PEG(12)-NHS to vancomycin using aqueous conditions (PBS, pH 7.4), it was

expected that the PEG would couple to the primary V3 amine. However, MALDI-

TOF results indicated that the product was heterogeneous, with coupling at both

the V3 and X1 positions (figure 7.2). In addition to unreacted SM-PEG(12)-NHS

and unreacted vancomycin, there were three additional product peaks. The main

product peak (2298 m/z) could correspond to attachment at either the V3 or X1 position. However, the two additional peaks at 2081 m/z (M-117 m/z) and 1893 m/z (M-305 m/z) indicate that a significant portion of the PEG crosslinker had attached at the X1 secondary amine, rather than the primary amine. While the

uncreacted crosslinker and vancomycin could be removed by dialysis and HPLC,

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Vancomycin

Vancosamine cleaved

Vancosamine + glucose cleaved

Figure 7.1: MALDI-TOF of vancomycin, showing the typical vancomycin

‘fingerprint’ of three peaks with m/z values of M (ie product), M-117, and M-305

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= Product + Na

= Product + H

Unreacted Vancomycin

Figure 7.2: MALDI‐TOF of SM‐PEG(12)‐Vancomycin synthesized in aqueous

conditions (PBS, pH 7.4). The presence of the M‐117 (2054 m/z) and M‐305 (1895 m/z) peaks indicate undesired coupling to the X1 amine.

232

it would not be possible to easily separate the two final products with any

significant yield. At this point, it became obvious that aqueous conditions would

not yield consistent, reproducible results, and other synthesis conditions were

investigated.

Greenwald et al [8] reported that DMSO was an inferior solvent for

Vancomycin-NHS reactions. Instead, they found that dimethyl formamide

(DMF) in combination with an excess of triethylamine (TEA) could be used to

generate high yields of a homogeneous V3-linked product. Greenwald also

investigated the use of other bases in place of TEA, such as N,N-

Diisopropylethylamine (DIEA) or 4-Dimethylaminopyridine (DMAP), but they

were found to yield di-substituted vancomycin derivatives. As a result, the reaction conditions employed for this work were modeled after Greenwald’s studies so as to favor a singly-substituted, homogeneous, high-yield product.

Using Greenwald’s organic synthesis conditions[8], vancomycin was coupled to the heterofunctional crosslinker SM-PEG(n)-NHS, with n representing a discrete number of ethylene glycol repeat units. MALDI-TOF was used to determine the PEG binding site for the SM-PEG(2)-Vancomycin and SM-

PEG(12)-Vancomycin products (Figure 7.3 and 7.4, respectively). The SM-

PEG(2)-Vancomycin product peak was observed at 1759 m/z, while the SM-

PEG(12)-Vancomycin product was observed at 2198 m/z. In addition, large peaks were observed at 454 m/z and 894 m/z, representing SM-PEG(2)-

233

= Product + H

Figure 7.3: MALDI-TOF of SM-PEG(2)-Vancomycin product synthesized in DMF with TFA. Fragment pattern indicates coupling at correct V3 position.

234

1143.281 894.471 1305.338

1115.280 959.463 2198.821 = Product + H 1448.437 1535.811 2220.827 = Product + Na 2236.779 = Product + K

Figure 7.4: MALDI-TOF of SM-PEG(12)-Vancomycin product synthesized in

DMF with TFA. Fragment pattern indicates coupling at correct V3 position.

235

vancosamine and SM-PEG(12)-vancosamine fragments (respectively) created by

the instrument’s laser. Notably absent are the fragment peaks at (M-117) m/z

and (M-305) m/z that were observed after synthesis in aqueous conditions.

Together, these results indicate that under organic conditions, the SM-PEG(n)-

NHS crosslinker attaches at the V3 position, creating a homogeneous product with no evidence of X1 attachment.

After confirming the successful synthesis of homogeneous SM-PEG(n)-

Vancomycin intermediate products under organic conditions, the 6-20 targeting

peptide was attached to the PEG maleimide group under aqueous conditions

(PBS, pH 7.4, 1 mM EDTA). MALDI-TOF was used to examine the final 6-20-

PEG(2)-Vancomycin product and confirm a successful conjugation (Figure 7.5).

The main product peak was clearly evident at 3633 m/z, while three main fragment peaks were observed at 1875, 2329, and 2491 m/z. Together, these peaks indicated the successful synthesis of the desired targeted vancomycin product.

The second main variable examined in this project was the length of the PEG linker. While Greenwald et al[8] report that small, low molecular weight groups yield insoluble final products, Lawson et al report no such difficulties[6, 7]. Once again, experimental data collected in this project agree with Greendwald’s findings, rather than Lawson’s. As described above, initial attempts were made to synthesize 6‐20‐PEG(n)‐Vancomycin products utilizing discrete, low

236

= Product

Figure 7.5: MALDI-TOF of 6-20-PEG(2)-Vancomycin product synthesized in

DMF with TFA. Fragment pattern indicates coupling at correct V3 position.

237

molecular weight PEG crosslinkers with 2 and 12 ethylene glycol repeat units.

After attaching the SM‐PEG(n)‐NHS crosslinkers to the vancomycin molecule at

the desired V3 position in organic conditions, the final synthesis step required

reaction in aqueous conditions. As seen in Figure 7.5, this reaction was successful

for the 6-20-PEG(2)-Vancomycin product. However, after the synthesis was

complete, it was discovered that the final product had very poor solubility in all

solvents tested (aqueous, organic, polar, non-polar, protic, aprotic). Furthermore,

the SM-PEG(12)-Vancomycin intermediate product was insoluble in aqueous

conditions, and as a result it was not possible to attach the targeting peptide in

order to finish the synthesis. Finally, higher molecular weight (MW 2000, 3400,

and 5000) polydisperse PEG crosslinkers were studied. As with the discrete

PEG(2) and PEG(12) linkers, the MW 2000 PEG linker resulted in an insoluble

product. However, with the MW 3400 and MW 5000 linkers, the solubility

problem was overcome due to the length of the hydrophilic PEG chain.

Under most pH conditions, vancomycin has a net positive charge of +1.

However, examination of the synthesis conditions revealed that coupling the

PEG chain to either the V3 or X1 amine on vancomycin results in the loss of a single positive charge from the molecule (Figure 7.6). The resulting PEG-

Vancomycin molecule no longer carries a net positive charge, but is instead

neutrally charged between pH 3 and pH 8 (Figure 7.7). It is suspected that this

238

pKa 7.75

pKa 2.18 pKa 8.89

pKa 12 pKa 10.4 pKa 9.59

Figure 7.6: Vancomycin typically carries two positive charges and one negative

charge at physiological pH. Covalent attachment of a heterofunctional

crosslinker to either of the amine groups results in the loss of a positive charge.

239

+3 VancomycinVancomycin +2 PEGxPEG‐Vancomycin‐Vancomycin

0 ‐1 2468101214

Charge ‐2

‐3

‐4 ‐5 pH

Figure 7.7: Vancomycin typically carries a net positive charge below a pH of ~8.

Attaching the PEG crosslinker to the V3 or X1 amine results in a neutrally charged molecule between pH 3 and pH 8.

240

loss of charge is the main reason vancomycin becomes insoluble after attaching

low molecular weight PEG linkers to either of the positively charged amine

groups. This suspicion was confirmed by studying the solubility of SM-PEG(2)-

Vancomycin and SM-PEG(12)-Vancomycin in aqueous environments ranging

from pH 1.0 to pH 12.0 (water, adjusted with 1N HCl in pH 1.0 increments). It

was observed that both products were readily soluble at pH values ≤3.0 and

≥10.0.

Due to the limitations imposed on crosslink length due to solubility issues,

this project was unable to study the effects of crosslink length on vancomycin

targeting in as much detail as was desired. It is expected that linker length will

play a crucial role on the effectiveness of the final products, as indicated by

results with 6‐20‐PEG3400‐VAN and 6‐20‐PEG5000‐VAN. The shorter MW 3400

linker demonstrated improved targeting ability at lower concentrations, likely due to decreased steric interference compared to the longer MW 5000 linker.

Therefore, it is of great interest going forward to devise a synthesis scheme that would enable us to create and study targeted vancomycin molecules with

increasingly shorter crosslinks. One potential scheme that was preliminarily

investigated over the course of this project is shown in Figure 7.8. It is believed

that the main reason for product insolubility when utilizing short PEG linkers is

due to the loss of charge on the vancomycin molecule. Therefore, the new

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3 -Vancomycin. The use of of -Vancomycin. The use x 6-20-PEG  the positive charge at the vancomycin V

Figure 7.8: Proposed synthesis scheme for soluble Traut’s reagent (2-iminothiolane) will maintain position.

242

scheme was devised in order to maintain this positive charge by utilizing 2‐

iminothiolane (Traut’s reagent) to introduce a maleimide‐reactive thiol group at

the vancomycin V3 position.

Meanwhile, the MAL‐PEGx‐NHS crosslinker would be attached by way of its

succinimide group to a selectively deprotected lysine residue at the amino‐

terminus of the resin‐bound 6‐20 peptide (Figure 7.9). By incorporating an

MTT‐protected lysine residue into the Peptide secquence, it is possible to

deprotect the lysine residue with 1.8% trifluoroacetic acid (TFA) in dichloromethane (DCM). As a result, the PEG chain can be coupled to the

peptide while it is still attached to the Knorr resin. Subsequent cleavage and

deprotection with 88% TFA then allows the 6‐20‐PEGx‐MAL product to be recovered in high yield. A preliminary synthesis was carried out using this on‐ resin modification technique, and MALDI‐TOF was used to examine the success of the 6‐20‐PEG(2)‐MAL conjugation (Figure 7.10). The main product peak is

identified at 2226 m/z. However, additional peaks between 2403 and 2713 m/z

indicate that the final 88% TFA cleavage/deprotection step was incomplete.

Further studies involving this synthesis step would strive to optimize the cleavage step, which is perhaps hindered by the presence of the PEG linker.

243

Selective Deprotection of Lys(Mtt): • 1.8% TFA in DCM • 9 x 3 minutes (room temperature) • 10 ml solvent per 1 g resin

Conjugation: • DMF • DIEA (10 equiv) • 4 hours (room temperature) • Argon Atmosphere

Cleavage (Reagent B): • TFA/Phenol/Water/TIPS (88/5/5/2) • 2 hours (room temp) • Argon Atmosphere

Figure 7.9: Synthesis scheme for on-resin modification of 6-20 peptide and attachment of MAL-PEGx-NHS crosslinker.

244

M+H Incomplete Deprotection

Mass (m/z)

Figure 7.10: MALDI-TOF of MAL-PEG(2)-6-20 product, synthesized according to the scheme presented in Figure 7.8. The main product peak is present at 2226 m/z, along with several peaks resulting from incomplete deprotection.

245

7.3 FUTURE DIRECTIONS - FLOW SYSTEM:

The studies presented in this project have served to demonstrate the

feasibility of specifically targeting antibacterial moieties to surface-adherent S.

epidermidis. A targeted cationic peptide was able to disrupt biofilm formation,

while a targeted vancomycin derivative was able increase vancomycin retention

at the biofilm. These results lay the groundwork for future studies striving to

characterize the targeted binding and retention behavior in more complex

systems, such as flow environments and in vivo models. While not examined directly in these studies, it is envisioned that the true benefit of the targeted products will be observed under flow conditions, where the improved binding ability will impart significant advantages.

Implementation of a flow system will allow for the study of peptide performance in the presence of human plasma proteins. A key unanswered question pertains to thrombin’s ability to bind the 6-20 peptide. While it has been demonstrated that the 6-20 peptide contains the Arg14-Gly15 thrombin cleavage site[9], it is unknown if thrombin can bind and/or cleave the peptide in solution. Studies have indicated that thrombin is unable to successfully cleave fibrinopeptide B from the fibrinogen B chain in the absence of the A and 

chains due to long-range interactions that require the intact fibrinogen

molecule[10, 11], despite the thrombin cleavage site being readily accessible on

the fibrinogen surface[11]. Kaczmarek and McDonagh[12] later showed that

246

thrombin is able to bind the fibrinogen chain (fibrinopeptide B removed).

Together, these findings indicate that while thrombin may be able bind to the B chain, it is unable to cleave fibrinopeptide B unless the fibrinogen molecule remains intact. Through the use of a flow system, targeted delivery to the 6-20 peptide to S. epidermidis biofilms could be studied in the presence of thrombin, potentially confirming that thrombin will be unable to bind or cleave the 6-20 peptide.

An additional question exists regarding the 6-20 peptide’s ability to compete with soluble fibrinogen in vivo. Since both the peptide and the protein are known to bind to the S. epidermidis SdrG protein, it is essential to confirm that soluble fibrinogen will not saturate the SdrG binding sites and render the peptide unable to bind. Preliminary studies addressing this issue were discussed in Chapter 4.6, using the 6-20-NG peptide to study targeting ability in the presence of fibrinogen. As S. epidermidis utilizes the fibrinogen binding pathway during biomaterial colonization, it is assumed that soluble fibrinogen is not able to block all available SdrG proteins. A flow system would allow us to test this hypothesis, wherein an S. epidermidis biofilm would be grown in the presence of circulating fibrinogen, and the 6-20 peptide could then be tested for its ability to target the bacteria.

In regards to the targeted vancomycin, it is crucial that the newly synthesized vancomycin derivatives possess the ability to kill the biofilms they are designed

247

to treat. Minimum inhibitory concentrations (MIC) and minimum bactericidal

concentrations (MBC) were determined for the 6‐20‐PEG3400‐VAN and 6‐20‐

PEG5000‐VAN products in Chapter 6. While these values were higher than those observed for vancomycin alone, it is important to note that these are solution‐

based parameters with no direct translation to biofilm environments. The

targeted products still had the ability to inhibit and kill S. epidermidis, and it is

very likely that the improved binding and retention properties would overcome

the slight decreases in antibacterial activity resulting from the synthesis process.

A study designed around a flow system would provide critical data regarding

the feasibility of using such targeted antibiotics in vivo. Data presented in

Chapter 6 confirmed that the native vancomycin molecule is not retained within

the biofilm, even after a 24 hour exposure period. However, the targeted

vancomycin molecules demonstrated a significant increase in retention. A flow

system would allow for wash‐out and dosing experiments, examining the ability

of targeted vancomycin to be delivered in small doses over extended periods of time. It is believed that such an experiment would yield a targeted, locally

elevated concentration of vancomycin within the biofilm. Studies would be

conducted in order to quantify the amount of vancomycin retained as well as its

location within the biofilm by way of confocal microscopy, in addition to biofilm

viability.

248

7.4 ACKNOWLEDGEMENTS:

The work described here was funded by Award Number 5R01EB000279 from the National Institute of Biomedical Imaging and Bioengineering, and work was carried out using the facilities of the Center for Cardiovascular Biomaterials. I would like to thank Faina Kligman for her assistance with MALDI-TOF analysis.

249

7.5 REFERENCES

1. Donlan, R.M., Preventing biofilms of clinically relevant organisms using

bacteriophage. Trends in microbiology, 2009. 17(2): p. 66-72.

2. Bloom, B., R. Schelonka, T. Kueser, W. Walker, E. Jung, D. Kaufman, K.

Kesler, D. Roberson, J. Patti, and S. Hetherington, Multicenter study to

assess safety and efficacy of INH-A21, a donor-selected human staphylococcal

immunoglobulin, for prevention of nosocomial infections in very low birth weight

infants. The Pediatric infectious disease journal, 2005. 24(10): p. 858.

3. Ohlsen, K. and U. Lorenz, Immunotherapeutic strategies to combat

staphylococcal infections. International Journal of Medical Microbiology,

2010. 300(6): p. 402-410.

4. Rennermalm, A., M. Nilsson, and J.I. Flock, Fibrinogen binding protein of

Staphylococcus epidermidis is a target for opsonic antibodies. Infection and

Immunity, 2004. 72(5): p. 3081-3083.

5. Vernachio, J.H., A.S. Bayer, B. Ames, D. Bryant, B.D. Prater, P.J. Syribeys,

E.L. Gorovits, and J.M. Patti, Human immunoglobulin G recognizing

fibrinogen-binding surface proteins is protective against both Staphylococcus

aureus and Staphylococcus epidermidis infections in vivo. Antimicrobial

Agents and Chemotherapy, 2006. 50(2): p. 511-518.

6. Lawson, M.C., R. Shoemaker, K.B. Hoth, C.N. Bowman, and K.S. Anseth,

Polymerizable vancomycin derivatives for bactericidal biomaterial surface

250

modification: structure-function evaluation. Biomacromolecules, 2009. 10(8):

p. 2221-34.

7. Lawson, M.K.C., C.N. Bowman, and K.S. Anseth, Vancomycin derivative

photopolymerized to titanium kills S. epidermidis. Clinical orthopaedics and

related research, 2007. 461: p. 96.

8. Greenwald, R.B., H. Zhao, J. Xia, and A. Martinez, Poly (ethylene glycol)

transport forms of vancomycin: a long-lived continuous release delivery system.

Journal of medicinal chemistry, 2003. 46(23): p. 5021-5030.

9. Davis, S.L., S. Gurusiddappa, K.W. McCrea, S. Perkins, and M. Hook,

SdrG, a fibrinogen-binding bacterial adhesin of the microbial surface components

recognizing adhesive matrix molecules subfamily from Staphylococcus

epidermidis, targets the thrombin cleavage site in the B beta chain. Journal of

Biological Chemistry, 2001. 276(30): p. 27799-27805.

10. Hageman, T.C. and H.A. Scheraga, Mechanism of action of thrombin on

fibrinogen: Reaction of the N-terminal CNBr fragment from the BÎ² chain of

bovine fibrinogen with bovine thrombin. Archives of Biochemistry and

Biophysics, 1977. 179(2): p. 506-517.

11. Nagy, J.A., Y.C. Meinwald, and H.A. Scheraga, Immunochemical

determination of conformational equilibria for fragments of the B.beta. chain of

fibrinogen. Biochemistry, 1985. 24(4): p. 882-887.

251

12. Kaczmarek, E. and J. McDonagh, Thrombin binding to the A alpha-, B beta-,

and gamma-chains of fibrinogen and to their remnants contained in fragment E.

Journal of Biological Chemistry, 1988. 263(27): p. 13896-13900.

252

CHAPTER 8

BIBLIOGRAPHY

Abraham, E and Chain, E, An enzyme from bacteria able to destroy penicillin. Nature,

1940. 146(3713): p. 837‐837.

Abramoff, M, Magelhaes, P, and Ram, S, Image Processing with ImageJ.

Biophotonics International, 2004. 11(7): p. 36‐42.

Absolom, DR, Lamberti, FV, Policova, Z, Zingg, W, van Oss, CJ, and Neumann,

AW, Surface thermodynamics of bacterial adhesion. Appl. Environ. Microbiol.,

1983. 46(1): p. 90‐97.

Allignet, J, Aubert, S, Dyke, KGH, and El Solh, N, Staphylococcus caprae Strains

Carry Determinants Known To Be Involved in Pathogenicity: a Gene Encoding an

Autolysin‐Binding Fibronectin and the ica Operon Involved in Biofilm Formation.

Infect. Immun., 2001. 69(2): p. 712‐718.

Anderl, JN, Zahller, J, Roe, F, and Stewart, PS, Role of Nutrient Limitation and

Stationary‐Phase Existence in Klebsiella pneumoniae Biofilm Resistance to

Ampicillin and Ciprofloxacin. Antimicrob. Agents Chemother., 2003. 47(4): p.

1251‐1256.

253

Anderson, JM, Chapter 4 Mechanisms of inflammation and infection with implanted

devices. Cardiovascular Pathology. 2(3, Supplement 1): p. 33‐41.

Anderson, JM, BIOLOGICAL RESPONSES TO MATERIALS. Annual Review of

Materials Research, 2001. 31(1): p. 81‐110.

Arciola, CR, Bustanji, Y, Conti, M, Campoccia, D, Baldassarri, L, Samori, B, and

Montanaro, L, Staphylococcus epidermidis ‐ fibronectin binding and its inhibition

by heparin. Biomaterials, 2003. 24(18): p. 3013‐3019.

Arimoto, H, Nishimura, K, Kiniumi, T, Hayakawa, I, and Uemura, D, Multi‐

valent polymer of vancomycin ‐ enhanced antibacterial activity against VRE. Chem.

Commun., 1999(15): p. 1361‐1362.

Aristoff, PA, Garcia, GA, Kirchhoff, PD, and Hollis Showalter, HD, Rifamycins ‐

Obstacles and opportunities. Tuberculosis, 2010. 90(2): p. 94‐118.

Babior, BM, Lambeth, JD, and Nauseef, W, The Neutrophil NADPH Oxidase.

Archives of Biochemistry and Biophysics, 2002. 397(2): p. 342‐344.

Balaban, NQ, Merrin, J, Chait, R, Kowalik, L, and Leibler, S, Bacterial persistence as

a phenotypic switch. Science, 2004. 305(5690): p. 1622.

Baldassarri, L, Donnelli, G, Gelosia, A, Voglino, MC, Simpson, AW, and

Christensen, GD, Purification and characterization of the staphylococcal slime‐

associated antigen and its occurrence among Staphylococcus epidermis clinical

isolates. Infect. Immun., 1996. 64(8): p. 3410‐3415.

254

Bale, MD, Wohlfahrt, LA, Mosher, DF, Tomasini, B, and Sutton, RC, Identification

of Vitronectin as a Major Plasma‐Protein Adsorbed on Polymer Surfaces of Different

Copolymer Composition. Blood, 1989. 74(8): p. 2698‐2706.

Banner, MA, Cunniffe, JG, Macintosh, RL, Foster, TJ, Rohde, H, Mack, D, Hoyes,

E, Derrick, J, Upton, M, and Handley, PS, Localized Tufts of Fibrils on

Staphylococcus epidermidis NCTC 11047 Are Comprised of the Accumulation‐

Associated Protein. Journal of Bacteriology, 2007. 189(7): p. 2793‐2804.

Barna, JCJ and Williams, DH, The structure and mode of action of glycopeptide

antibiotics of the vancomycin group. Annual Reviews in Microbiology, 1984.

38(1): p. 339‐357.

Bateman, A, The G5 domain: a potential N‐acetylglucosamine recognition domain

involved in biofilm formation. Bioinformatics, 2004. 21(8): p. 1301‐1303.

Bayoudh, S, Othmane, A, Mora, L, and Ben Ouada, H, Assessing bacterial adhesion

using DLVO and XDLVO theories and the jet impingement technique. Colloids and

Surfaces B: Biointerfaces, 2009. 73(1): p. 1‐9.

Beauregard, DA, Williams, DH, Gwynn, MN, and Knowles, DJ, Dimerization and

membrane anchors in extracellular targeting of vancomycin group antibiotics.

Antimicrob. Agents Chemother., 1995. 39(3): p. 781‐785.

Bigger, J, Treatment of Staphylococcal Infections with Penicillin by Intermittent

Sterilisation. The Lancet, 1944. 244(6320): p. 497‐500.

255

Blanchemain, N, Haulon, S, Boschin, F, Marcon‐Bachari, E, Traisnel, M,

Morcellet, M, Hildebrand, HF, and Martel, B, Vascular prostheses with controlled

release of antibiotics: Part 1: Surface modification with cyclodextrins of PET

prostheses. Biomolecular Engineering, 2007. 24(1): p. 149‐153.

Blanchemain, N, Haulon, S, Martel, B, Traisnel, M, Morcellet, M, and

Hildebrand, HF, Vascular PET prostheses surface modification with cyclodextrin

coating: development of a new drug delivery system. European journal of vascular

and endovascular surgery, 2005. 29(6): p. 628‐632.

Bloom, B, Schelonka, R, Kueser, T, Walker, W, Jung, E, Kaufman, D, Kesler, K,

Roberson, D, Patti, J, and Hetherington, S, Multicenter study to assess safety and

efficacy of INH‐A21, a donor‐selected human staphylococcal immunoglobulin, for

prevention of nosocomial infections in very low birth weight infants. The Pediatric

infectious disease journal, 2005. 24(10): p. 858.

Blumberg, PM and Strominger, JL, Interaction of penicillin with the bacterial cell:

penicillin‐binding proteins and penicillin‐sensitive enzymes. Microbiology and

Molecular Biology Reviews, 1974. 38(3): p. 291.

Boks, NP, Norde, W, van der Mei, HC, and Busscher, HJ, Forces involved in

bacterial adhesion to hydrophilic and hydrophobic surfaces. Microbiology, 2008.

154(10): p. 3122‐3133.

256

Borriello, G, Werner, E, Roe, F, Kim, AM, Ehrlich, GD, and Stewart, PS, Oxygen

Limitation Contributes to Antibiotic Tolerance of Pseudomonas aeruginosa in

Biofilms. Antimicrob. Agents Chemother., 2004. 48(7): p. 2659‐2664.

Bowden, MG, Chen, W, Singvall, J, Xu, Y, Peacock, SJ, Valtulina, V, Speziale, P,

and Hook, M, Identification and preliminary characterization of cell‐wall‐anchored

proteins of Staphylococcus epidermidis. Microbiology‐Sgm, 2005. 151: p. 1453‐

1464.

Bowden, MG, Visai, L, Longshaw, CM, Holland, KT, Speziale, P, and Höök, M, Is

the GehD Lipase from Staphylococcus epidermidis a Collagen Binding Adhesin?

Journal of Biological Chemistry, 2002. 277(45): p. 43017‐43023.

Brash, JL and Horbett, TA, Proteins at Interfaces, in Proteins at Interfaces II. 1995,

American Chemical Society. p. 1‐23.

Brash John, L and Horbett Thomas, A, Proteins at Interfaces, in Proteins at Interfaces

II. 1995, American Chemical Society. p. 1‐23.

Brown, MRW, Allison, DG, and Gilbert, P, Resistance of bacterial biofilms to

antibiotics a growth‐rate related effect? Journal of Antimicrobial Chemotherapy,

1988. 22(6): p. 777‐780.

Brummel, K, Butenas, S, and Mann, K, An Integrated Study of Fibrinogen During

Blood Coagulation. J. Biol. Chem., 1999. 274(32): p. 22862‐22870.

257

Cao, H and Liu, X, Silver nanoparticles‐modified films versus biomedical device‐

associated infections. Wiley Interdisciplinary Reviews: Nanomedicine and

Nanobiotechnology, 2010. 2(6): p. 670‐684.

Cerca, N, Jefferson, KK, Maira‐Litran, T, Pier, DB, Kelly‐Quintos, C, Goldmann,

DA, Azeredo, J, and Pier, GB, Molecular basis for preferential protective efficacy of

antibodies directed to the poorly acetylated form of staphylococcal poly‐N‐acetyl‐beta‐

(1‐6)‐glucosaminev. Infection and Immunity, 2007. 75(7): p. 3406‐3413.

Cerca, N, Martins, S, Cerca, F, Jefferson, KK, Pier, GB, Oliveira, R, and Azeredo,

J, Comparative assessment of antibiotic susceptibility of coagulase‐negative

staphylococci in biofilm versus planktonic culture as assessed by bacterial

enumeration or rapid XTT colorimetry. Journal of Antimicrobial Chemotherapy,

2005. 56(2): p. 331‐336.

Cerca, N, Oliveira, R, and Azeredo, J, Susceptibility of Staphylococcus epidermidis

planktonic cells and biofilms to the lytic action of staphylococcus bacteriophage K.

Letters in Applied Microbiology, 2007. 45(3): p. 313‐317.

Ceri, H, Olson, ME, Stremick, C, Read, RR, Morck, D, and Buret, A, The Calgary

Biofilm Device: New Technology for Rapid Determination of Antibiotic

Susceptibilities of Bacterial Biofilms. J. Clin. Microbiol., 1999. 37(6): p. 1771‐1776.

Chaignon, P, Sadovskaya, I, Ragunah, C, Ramasubbu, N, Kaplan, J, and Jabbouri,

S, Susceptibility of staphylococcal biofilms to enzymatic treatments depends on their

258

chemical composition. Applied Microbiology and Biotechnology, 2007. 75(1): p.

125‐132.

Chapple, DS, Mason, DJ, Joannou, CL, Odell, EW, Gant, V, and Evans, RW,

Structure‐function relationship of antibacterial synthetic peptides homologous to a

helical surface region on human lactoferrin against Escherichia coli serotype O111.

Infection and Immunity, 1998. 66(6): p. 2434‐2440.

Chaw, KC, Manimaran, M, and Tay, FEH, Role of Silver Ions in Destabilization of

Intermolecular Adhesion Forces Measured by Atomic Force Microscopy in

Staphylococcus epidermidis Biofilms. Antimicrob. Agents Chemother., 2005.

49(12): p. 4853‐4859.

Chen, LF and Kaye, D, Current use for old antibacterial agents: polymyxins,

rifamycins, and aminoglycosides. Infectious disease clinics of North America,

2009. 23(4): p. 1053‐1075.

Cheng, G, Zhang, Z, Chen, S, Bryers, JD, and Jiang, S, Inhibition of bacterial

adhesion and biofilm formation on zwitterionic surfaces. Biomaterials, 2007. 28(29):

p. 4192‐4199.

Chokr, A, Watier, D, Eleaume, H, Pangon, B, Ghnassia, J‐C, Mack, D, and

Jabbouri, S, Correlation between biofilm formation and production of polysaccharide

intercellular adhesin in clinical isolates of coagulase‐negative staphylococci.

International Journal of Medical Microbiology, 2006. 296(6): p. 381‐388.

259

Christner, M, Franke, GC, Schommer, NN, Wendt, U, Wegert, K, Pehle, P, Kroll,

G, Schulze, C, Buck, F, Mack, D, Aepfelbacher, M, and Rohde, H, The giant

extracellular matrix‐binding protein of Staphylococcus epidermidis mediates biofilm

accumulation and attachment to fibronectin. Molecular Microbiology, 2010. 75(1):

p. 187‐207.

Chugh, TD, Burns, GJ, Shuhaiber, HJ, and Bahr, GM, Adherence Of Staphylococcus‐

Epidermidis To Fibrin‐Platelet Clots Invitro Mediated By Lipoteichoic Acid.

Infection And Immunity, 1990. 58(2): p. 315‐319.

Comfort, D and Clubb, RT, A Comparative Genome Analysis Identifies Distinct

Sorting Pathways in Gram‐Positive Bacteria. Infect. Immun., 2004. 72(5): p. 2710‐

2722.

Conlon, KM, Humphreys, H, and OʹGara, JP, icaR Encodes a Transcriptional

Repressor Involved in Environmental Regulation of ica Operon Expression and

Biofilm Formation in Staphylococcus epidermidis. Journal of Bacteriology, 2002.

184(16): p. 4400‐4408.

Conlon, KM, Humphreys, H, and OʹGara, JP, Inactivations of rsbU and sarA by

IS256 Represent Novel Mechanisms of Biofilm Phenotypic Variation in

Staphylococcus epidermidis. J. Bacteriol., 2004. 186(18): p. 6208‐6219.

Conrady, DG, Brescia, CC, Horii, K, Weiss, AA, Hassett, DJ, and Herr, AB, A

zinc‐dependent adhesion module is responsible for intercellular adhesion in

260

staphylococcal biofilms. Proceedings of the National Academy of Sciences, 2008.

105(49): p. 19456‐19461.

Costerton, J, Lewandowski, Z, Caldwell, D, Korber, D, and Lappin‐Scott, H,

Microbial biofilms. Annu Rev Microbiol, 1995. 49: p. 711‐45.

Costerton, JW, Bacterial Biofilms: A Common Cause of Persistent Infections. Science,

1999. 284(5418): p. 1318‐1322.

Cramer, E, Pryzwansky, KB, and Villeval, JL, Ultrastructural localization of

lactoferrin and myeloperoxidase in human neutrophils by immunogold. Blood, 1985.

65(2): p. 423‐432.

Cramton, SE, Gerke, C, Schnell, NF, Nichols, WW, and Gotz, F, The Intercellular

Adhesion (ica) Locus Is Present in Staphylococcus aureus and Is Required for Biofilm

Formation. Infect. Immun., 1999. 67(10): p. 5427‐5433.

Cringus‐Fundeanu, I, Luijten, J, van der Mei, HC, Busscher, HJ, and Schouten,

AJ, Synthesis and Characterization of Surface‐Grafted Polyacrylamide Brushes and

Their Inhibition of Microbial Adhesion. Langmuir, 2007. 23(9): p. 5120‐5126.

Cucarella, C, Solano, C, Valle, J, Amorena, B, Lasa, I, and Penades, JR, Bap, a

Staphylococcus aureus Surface Protein Involved in Biofilm Formation. J. Bacteriol.,

2001. 183(9): p. 2888‐2896.

261

Curtin, JJ and Donlan, RM, Using Bacteriophages To Reduce Formation of Catheter‐

Associated Biofilms by Staphylococcus epidermidis. Antimicrobial Agents and

Chemotherapy, 2006. 50(4): p. 1268‐1275.

Dale, DC, Boxer, L, and Liles, WC, The phagocytes: neutrophils and monocytes.

Blood, 2008. 112(4): p. 935‐945.

Darouiche, RabihÂ O, Device‐Associated Infections: A Macroproblem that Starts with

Microadherence. Clinical Infectious Diseases, 2001. 33(9): p. 1567‐1572.

Darouiche, RO, Treatment of Infections Associated with Surgical Implants. N Engl J

Med, 2004. 350(14): p. 1422‐1429.

Darouiche, RO, Dhir, A, Miller, AJ, Landon, GC, Raad, II, and Musher, DM,

Vancomycin penetration into biofilm covering infected prostheses and effect on

bacteria. J Infect Dis, 1994. 170(3): p. 720‐3.

Das, T, Sharma, PK, Busscher, HJ, van der Mei, HC, and Krom, BP, Role of

Extracellular DNA in Initial Bacterial Adhesion and Surface Aggregation. Appl.

Environ. Microbiol.: p. AEM.03119‐09.

Davies, J and Davies, D, Origins and Evolution of Antibiotic Resistance. Microbiol.

Mol. Biol. Rev., 2010. 74(3): p. 417‐433.

Davies, J and Wright, GD, Bacterial resistance to aminoglycoside antibiotics. Trends

in Microbiology, 1997. 5(6): p. 234‐240.

262

Davis, SL, Gurusiddappa, S, McCrea, KW, Perkins, S, and Hook, M, SdrG, a

fibrinogen‐binding bacterial adhesin of the microbial surface components recognizing

adhesive matrix molecules subfamily from Staphylococcus epidermidis, targets the

thrombin cleavage site in the B beta chain. Journal of Biological Chemistry, 2001.

276(30): p. 27799‐27805.

De Pascale, G and Wright, GD, Antibiotic resistance by enzyme inactivation: from

mechanisms to solutions. ChemBioChem, 2010. 11(10): p. 1325‐1334.

Delihas, N, Riley, LW, Loo, W, Berkowitz, J, and Poltoratskaia, N, High‐Sensitivity

of Mycobacterium Species to the Bactericidal Activity by Polylysine. Fems

Microbiology Letters, 1995. 132(3): p. 233‐237.

Desai, NP, Hossainy, SFA, and Hubbell, JA, Surface‐immobilized polyethylene oxide

for bacterial repellence. Biomaterials, 1992. 13(7): p. 417‐420.

Donlan, RM, Preventing biofilms of clinically relevant organisms using bacteriophage.

Trends in microbiology, 2009. 17(2): p. 66‐72.

Dunman, PM, Murphy, E, Haney, S, Palacios, D, Tucker‐Kellogg, G, Wu, S,

Brown, EL, Zagursky, RJ, Shlaes, D, and Projan, SJ, Transcription Profiling‐

Based Identification of Staphylococcus aureus Genes Regulated by the agr and/or

sarA Loci. J. Bacteriol., 2001. 183(24): p. 7341‐7353.

263

Dunne, WM, Mason, E, and Kaplan, S, Diffusion of Rifampin and Vancomycin

through a Staphylococcus epidermidis Biofilm. Antimicrobial Agents and

Chemotherapy, 1993. 37(12): p. 2522‐2526.

Eckhart, L, Fischer, H, Barken, KB, Tolker‐Nielsen, T, and Tschachler, E,

DNase1L2 suppresses biofilm formation by Pseudomonas aeruginosa and

Staphylococcus aureus. British Journal of Dermatology, 2007. 156(6): p. 1342‐

1345.

Endl, J, Seidl, HP, Fiedler, F, and Schleider, KH, Chemical composition and structure

of cell wall teichoic acids of staphylococci. Archives of Microbiology, 1983. 135(3):

p. 215‐223.

Farber, B, Kaplan, M, and Clogston, A, Staphylococcus epidermidis Extracted Slime

Inhibits the Antimicrobial Action of Glycopeptide Antibiotics. The Journal of

Infections Diseases, 1990. 161(1): p. 37‐40.

Faurschou, M and Borregaard, N, Neutrophil granules and secretory vesicles in

inflammation. Microbes and Infection, 2003. 5(14): p. 1317‐1327.

Fields, G and Noble, R, Solid phase peptide synthesis utilizing 9‐

fluorenylmethoxycarbonyl amino acids. Int J Pept Protein Res, 1990. 35(3): p. 161‐

214.

Fischbach, MA and Walsh, CT, Antibiotics for emerging pathogens. Science, 2009.

325(5944): p. 1089.

264

Francois, P, Vaudaux, P, and Lew, PD, Role of Plasma and Extracellular Matrix

Proteins in the Physiopathology of Foreign Body Infections. Annals of Vascular

Surgery, 1998. 12(1): p. 34‐40.

Frank, KL, Hanssen, AD, and Patel, R, icaA Is Not a Useful Diagnostic Marker for

Prosthetic Joint Infection. Journal of Clinical Microbiology, 2004. 42(10): p. 4846‐

4849.

Frank, KL and Patel, R, Poly‐N‐Acetylglucosamine Is Not a Major Component of the

Extracellular Matrix in Biofilms Formed by icaADBC‐Positive Staphylococcus

lugdunensis Isolates. Infect. Immun., 2007. 75(10): p. 4728‐4742.

Fredheim, EGA, Klingenberg, C, Rohde, H, Frankenberger, S, Gaustad, P,

Flaegstad, T, and Sollid, JE, Biofilm Formation by Staphylococcus haemolyticus. J.

Clin. Microbiol., 2009. 47(4): p. 1172‐1180.

Fujii, K, Matsumoto, HN, Koyama, Y, Iwasaki, Y, Ishihara, K, and Takakuda, K,

Prevention of biofilm formation with a coating of 2‐methacryloyloxyethyl

phosphorylcholine polymer. Journal of Veterinary Medical Science, 2008. 70(2): p.

167‐173.

Furno, F, Morley, KS, Wong, B, Sharp, BL, Arnold, PL, Howdle, SM, Bayston, R,

Brown, PD, Winship, PD, and Reid, HJ, Silver nanoparticles and polymeric

medical devices: a new approach to prevention of infection? Journal of

Antimicrobial Chemotherapy, 2004. 54(6): p. 1019‐1024.

265

Ganz, T, Selsted, ME, Szklarek, D, Harwig, SS, Daher, K, Bainton, DF, and

Lehrer, RI, Defensins. Natural peptide antibiotics of human neutrophils. The

Journal of Clinical Investigation, 1985. 76(4): p. 1427‐1435.

Gaonkar, TruptiÂ AP, Caraos, LBS, and Modak, SP, Efficacy of a Silicone Urinary

Catheter Impregnated with Chlorhexidine and Triclosan Against Colonization With

Proteus mirabilis and Other UropathogensÂ â€¢â€‚. Infection Control and

Hospital Epidemiology, 2007. 28(5): p. 596‐598.

Gaonkar, TAP, Sampath, LABA, and Modak, SMP, Evaluation of the Antimicrobial

Efficacy of Urinary Catheters Impregnated with Antiseptics in an In Vitro Urinary

Tract ModelÂ â€¢â€‚. Infection Control and Hospital Epidemiology, 2003.

24(7): p. 506‐513.

Gerke, C, Kraft, A, Süßmuth, R, Schweitzer, O, and Götz, F, Characterization of the

N‐Acetylglucosaminyltransferase Activity Involved in the Biosynthesis of the

Staphylococcus epidermidisPolysaccharide Intercellular Adhesin. Journal of

Biological Chemistry, 1998. 273(29): p. 18586‐18593.

Gottenbos, B, van der Mei, HC, Klatter, F, Nieuwenhuis, P, and Busscher, HJ, In

vitro and in vivo antimicrobial activity of covalently coupled quaternary ammonium

silane coatings on silicone rubber. Biomaterials, 2002. 23(6): p. 1417‐1423.

Gotz, F, Staphylococcus and biofilms. Molecular Microbiology, 2002. 43: p. 1367‐

1378.

266

Green, RJ, Davies, MC, Roberts, CJ, and Tendler, SJB, Competitive protein

adsorption as observed by surface plasmon resonance. Biomaterials, 1999. 20(4): p.

385‐391.

Greenwald, RB, Zhao, H, Xia, J, and Martinez, A, Poly (ethylene glycol) transport

forms of vancomycin: a long‐lived continuous release delivery system. Journal of

medicinal chemistry, 2003. 46(23): p. 5021‐5030.

Griffin, JH, Linsell, MS, Nodwell, MB, Chen, Q, Pace, JL, Quast, KL, Krause, KM,

Farrington, L, Wu, TX, Higgins, DL, Jenkins, TE, Christensen, BG, and Judice,

JK, Multivalent Drug Design. Synthesis and In Vitro Analysis of an Array of

Vancomycin Dimers. Journal of the American Chemical Society, 2003. 125(21):

p. 6517‐6531.

Gross, M, Cramton, SE, Gotz, F, and Peschel, A, Key role of teichoic acid net charge

in Staphylococcus aureus colonization of artificial surfaces. Infect Immun, 2001.

69(5): p. 3423‐6.

Guo, BN, Zhao, X, Shi, YG, Zhu, DM, and Zhang, YY, Pathogenic implication of a

fibrinogen‐binding protein of Staphylococcus epidermidis in a rat model of

intravascular‐catheter‐associated infection. Infection and Immunity, 2007. 75(6):

p. 2991‐2995.

Hageman, TC and Scheraga, HA, Mechanism of action of thrombin on fibrinogen:

Reaction of the N‐terminal CNBr fragment from the BÎ² chain of bovine fibrinogen

267

with bovine thrombin. Archives of Biochemistry and Biophysics, 1977. 179(2): p.

506‐517.

Hall, AE, Patel, PR, Domanski, PJ, Prater, BD, Gorovits, EL, Syribeys, PJ,

Vernachio, JH, Patti, JM, and Hutchins, JT, A panel of monoclonal antibodies

recognizing the Staphylococcus epidermidis fibrinogen‐binding MSCRAMM SdrG.

Hybridoma, 2007. 26(1): p. 28‐34.

Hampton, MB, Kettle, AJ, and Winterbourn, CC, Inside the Neutrophil Phagosome:

Oxidants, Myeloperoxidase, and Bacterial Killing. Blood, 1998. 92(9): p. 3007‐3017.

Hartford, O, OʹBrien, L, Schofield, K, Wells, J, and Foster, TJ, The Fbe (SdrG)

protein of Staphylococcus epidermidis HB promotes bacterial adherence to fibrinogen.

Microbiology‐Sgm, 2001. 147: p. 2545‐2552.

Hayward, JA and Chapman, D, Biomembrane surfaces as models for polymer design:

the potential for haemocompatibility. Biomaterials, 1984. 5(3): p. 135‐142.

Heilmann, C, Identification and characterization of a novel autolysin (Aae) with

adhesive properties from Staphylococcus epidermidis. Microbiology, 2003. 149(10):

p. 2769‐2778.

Heilmann, C, Gerke, C, Perdreau‐Remington, F, and Gotz, F, Characterization of

Tn917 insertion mutants of Staphylococcus epidermidis affected in biofilm formation.

Infect. Immun., 1996. 64(1): p. 277‐282.

268

Heilmann, C, Hussain, M, Peters, G, and Gotz, F, Evidence for autolysin‐mediated

primary attachment of Staphylococcus epidermidis to a polystyrene surface.

Molecular Microbiology, 1997. 24(5): p. 1013‐1024.

Heilmann, C, Hussain, M, Peters, G, and Gotz, F, Evidence for autolysin‐mediated

primary attachment of Staphylococcus epidermidis to a polystyrene surface. Mol

Microbiol, 1997. 24(5): p. 1013‐24.

Heilmann, C, Schweitzer, O, Gerke, C, Vanittanakom, N, Mack, D, and Gotz, F,

Molecular basis of intercellular adhesion in the biofilm‐forming Staphylococcus

epidermidis. Mol Microbiol, 1996. 20(5): p. 1083‐91.

Hermann, T, Aminoglycoside antibiotics: old drugs and new therapeutic approaches.

Cellular and Molecular Life Sciences, 2007. 64(14): p. 1841‐1852.

Hermansson, M, The DLVO theory in microbial adhesion. Colloids and Surfaces B:

Biointerfaces, 1999. 14(1‐4): p. 105‐119.

Hirota, K, Murakami, K, Nemoto, K, and Miyake, Y, Coating of a surface with 2‐

methacryloyloxyethyl phosphorylcholine (MPC) co‐polymer significantly reduces

retention of human pathogenic microorganisms. FEMS Microbiology Letters, 2005.

248(1): p. 37‐45.

Ho, CH, Tobis, J, Sprich, C, Thomann, R, and Tiller, JC, Nanoseparated polymeric

networks with multiple antimicrobial properties. Advanced materials, 2004.

16(12): p. 957‐961.

269

Hoek, EMV and Agarwal, GK, Extended DLVO interactions between spherical

particles and rough surfaces. Journal of Colloid and Interface Science, 2006.

298(1): p. 50‐58.

Hooper, DC, Mode of Action of Fluoroquinolones. Drugs, 1999. 58(S6): p. 6‐10.

Hooper, DC, New Uses for New and Old Quinolones and the Challenge of Resistance.

Clinical Infectious Diseases, 2000. 30(2): p. 243‐254.

Hujer, AM, Kania, M, Gerken, T, Anderson, VE, Buynak, JD, Ge, X, Caspers, P,

Page, MGP, Rice, LB, and Bonomo, RA, Structure‐Activity Relationships of

Different {beta}‐Lactam Antibiotics against a Soluble Form of Enterococcus faecium

PBP5, a Type II Bacterial Transpeptidase. Antimicrob. Agents Chemother., 2005.

49(2): p. 612‐618.

Hussain, M, Hastings, JGM, and White, PJ, Isolation and Composition of the

Extracellular Slime Made by Coagulase‐Negative Staphylococci in a Chemically

Defined Medium. The Journal of Infectious Diseases, 1991. 163(3): p. 534‐541.

Hussain, M, Hastings, JGM, and White, PJ, Comparison of cell‐wall teichoic acid with

high‐molecular‐weight extracellular slime material from Staphylococcus epidermidis.

J Med Microbiol, 1992. 37(6): p. 368‐375.

Hussain, M, Heilmann, C, Peters, G, and Herrmann, M, Teichoic acid enhances

adhesion of Staphylococcus epidermidis to immobilized fibronectin. Microbial

Pathogenesis, 2001. 31(6): p. 261‐270.

270

Hussain, M, Herrmann, M, von Eiff, C, Perdreau‐Remington, F, and Peters, G, A

140‐kilodalton extracellular protein is essential for the accumulation of

Staphylococcus epidermidis strains on surfaces. Infect. Immun., 1997. 65(2): p. 519‐

524.

Izano, EA, Amarante, MA, Kher, WB, and Kaplan, JB, Differential Roles of Poly‐N‐

Acetylglucosamine Surface Polysaccharide and Extracellular DNA in Staphylococcus

aureus and Staphylococcus epidermidis Biofilms. Appl. Environ. Microbiol., 2008.

74(2): p. 470‐476.

Jefferson, KK, Goldmann, DA, and Pier, GB, Use of Confocal Microscopy To Analyze

the Rate of Vancomycin Penetration through Staphylococcus aureus Biofilms.

Antimicrob. Agents Chemother., 2005. 49(6): p. 2467‐2473.

Jerne, NK and Avegno, P, The Development of the Phage‐Inactivating Properties of

Serum During the Course of Specific Immunization of an Animal: Reversible and

Irreversible Inactivation. The Journal of Immunology, 1956. 76(3): p. 200‐208.

Joiner, KA, Ganz, T, Albert, J, and Rotrosen, D, The opsonizing ligand on Salmonella

typhimurium influences incorporation of specific, but not azurophil, granule

constituents into neutrophil phagosomes. The Journal of Cell Biology, 1989.

109(6): p. 2771‐2782.

271

Jose, B, Antoci Jr, V, Zeiger, AR, Wickstrom, E, and Hickok, NJ, Vancomycin

Covalently Bonded to Titanium Beads Kills Staphylococcus aureus. Chemistry &

Biology, 2005. 12(9): p. 1041‐1048.

K Ista, L, Fan, H, Baca, O, and P LÃ³pez, G, Attachment of bacteria to model solid

surfaces: oligo (ethylene glycol) surfaces inhibit bacterial attachment. FEMS

microbiology letters, 1996. 142(1): p. 59‐63.

Kaczmarek, E and McDonagh, J, Thrombin binding to the A alpha‐, B beta‐, and

gamma‐chains of fibrinogen and to their remnants contained in fragment E. Journal

of Biological Chemistry, 1988. 263(27): p. 13896‐13900.

Kahne, D, Leimkuhler, C, Lu, W, and Walsh, C, Glycopeptide and lipoglycopeptide

antibiotics. Chemical reviews, 2005. 105(2): p. 425‐448.

Keren, I, Kaldalu, N, Spoering, A, Wang, Y, and Lewis, K, Persister cells and

tolerance to antimicrobials. FEMS Microbiology Letters, 2004. 230(1): p. 13‐18.

Kogan, G, Sadovskaya, I, Chaignon, P, Chokr, A, and Jabbouri, S, Biofilms of

clinical strains of Staphylococcus that do not contain polysaccharide intercellular

adhesin. FEMS Microbiology Letters, 2006. 255(1): p. 11‐16.

König, C, Schwank, S, and Blaser, J, Factors compromising antibiotic activity against

biofilms of Staphylococcus epidermidis. European Journal of Clinical

Microbiology & Infectious Diseases, 2001. 20(1): p. 20‐26.

272

Kugler, R, Bouloussa, O, and Rondelez, F, Evidence of a charge‐density threshold for

optimum efficiency of biocidal cationic surfaces. Microbiology, 2005. 151(5): p.

1341.

Kutter, E, Raya, R, and Carlson, K, Molecular mechanisms of phage infection, in

Bacteriophages: Biology and Applications, Kutter, E and Sulakvelidze, A, Editors.

2005. p. 165‐222.

Lawson, MC, Shoemaker, R, Hoth, KB, Bowman, CN, and Anseth, KS,

Polymerizable vancomycin derivatives for bactericidal biomaterial surface

modification: structure‐function evaluation. Biomacromolecules, 2009. 10(8): p.

2221‐34.

Lawson, MKC, Bowman, CN, and Anseth, KS, Vancomycin derivative

photopolymerized to titanium kills S. epidermidis. Clinical orthopaedics and

related research, 2007. 461: p. 96.

Leckband, D, Sheth, S, and Halperin, A, Grafted poly(ethylene oxide) brushes as

nonfouling surface coatings. Journal of Biomaterials Science, Polymer Edition,

1999. 10(10): p. 1125‐1147.

Leclercq, R, Mechanisms of Resistance to Macrolides and Lincosamides: Nature of the

Resistance Elements and Their Clinical Implications. Clinical Infectious Diseases,

2002. 34(4): p. 482‐492.

273

Lee, H, Lee, Y, Statz, AR, Rho, J, Park, TG, and Messersmith, PB, Substrate‐

Independent Layer‐by‐Layer Assembly by Using Mussel‐Adhesive‐Inspired

Polymers. Advanced Materials, 2008. 20(9): p. 1619‐1623.

Lewis, AL, Phosphorylcholine‐based polymers and their use in the prevention of

biofouling. Colloids and Surfaces B: Biointerfaces, 2000. 18(3‐4): p. 261‐275.

Lewis, K, Persister cells and the riddle of biofilm survival. Biochemistry (Moscow),

2005. 70(2): p. 267‐274.

Li, DQ, Lundberg, F, and Ljungh, A, Characterization of vitronectin‐binding proteins

of Staphylococcus epidermidis. Current Microbiology, 2001. 42(5): p. 361‐367.

Li, H, Xu, L, Wang, J, Wen, Y, Vuong, C, Otto, M, and Gao, Q, Conversion of

Staphylococcus epidermidis Strains from Commensal to Invasive by Expression of the

ica Locus Encoding Production of Biofilm Exopolysaccharide. Infection and

Immunity, 2005. 73(5): p. 3188‐3191.

Lollike, K, Kjeldsen, L, Sengelov, H, and Borregaard, N, Lysozyme in human

neutrophils and plasma. A parameter of myelopoietic activity. Leukemia, 1995. 9(1):

p. 159‐164.

Long, KS, Poehlsgaard, J, Kehrenberg, C, Schwarz, S, and Vester, B, The Cfr rRNA

methyltransferase confers resistance to phenicols, lincosamides, oxazolidinones,

pleuromutilins, and streptogramin A antibiotics. Antimicrobial Agents and

Chemotherapy, 2006. 50(7): p. 2500.

274

Mack, D, Mechanisms of biofilm formation in and : functional molecules, regulatory

circuits, and adaptive responses. International Journal of Medical Microbiology,

2004. 294(2‐3): p. 203‐212.

Mack, D, Davies, A, Harris, L, Knobloch, J, and Rohde, H, Staphylococcus

epidermidis Biofilms: Functional Molecules, Relation to Virulence, and Vaccine

Potential, in Glycoscience and Microbial Adhesion, Lindhorst, TK and Oscarson,

S, Editors. 2009, Springer Berlin / Heidelberg. p. 157‐182.

Mack, D, Davies, AP, Harris, LG, Rohde, H, Horstkotte, MA, and Knobloch,

JKM, Microbial interactions in Staphylococcus epidermidis biofilms. Analytical and

Bioanalytical Chemistry, 2006. 387(2): p. 399‐408.

Mack, D, Fischer, W, Krokotsch, A, Leopold, K, Hartmann, R, Egge, H, and

Laufs, R, The intercellular adhesin involved in biofilm accumulation of

Staphylococcus epidermidis is a linear beta‐1,6‐linked glucosaminoglycan:

purification and structural analysis. J Bacteriol, 1996. 178(1): p. 175‐83.

Mack, D, Nedelmann, M, Krokotsch, A, Schwarzkopf, A, Heesemann, J, and

Laufs, R, Characterization of transposon mutants of biofilm‐producing

Staphylococcus epidermidis impaired in the accumulative phase of biofilm production:

genetic identification of a hexosamine‐containing polysaccharide intercellular

adhesin. Infect. Immun., 1994. 62(8): p. 3244‐3253.

275

Mack, D, Riedewald, J, Rohde, H, Magnus, T, Feucht, HH, Elsner, HA, Laufs, R,

and Rupp, ME, Essential functional role of the polysaccharide intercellular adhesin

of Staphylococcus epidermidis in hemagglutination. Infection and Immunity, 1999.

67(2): p. 1004‐1008.

Mack, D, Siemssen, N, and Laufs, R, Parallel induction by glucose of adherence and a

polysaccharide antigen specific for plastic‐adherent Staphylococcus epidermidis:

evidence for functional relation to intercellular adhesion. Infect Immun, 1992. 60(5):

p. 2048‐57.

MacKintosh, EE, Patel, JD, Marchant, RE, and Anderson, JM, Effects of biomaterial

surface chemistry on the adhesion and biofilm formation of Staphylococcus

epidermidis in vitro. J Biomed Mater Res A, 2006. 78(4): p. 836‐42.

Madigan, MT and Martinko, JM, Brock Biology of Microorganisms. 11 ed. 2006,

Upper Saddle River, NJ: Pearson Prentice Hall.

Maira‐Litran, T, Kropec, A, Abeygunawardana, C, Joyce, J, Mark Iii, G,

Goldmann, DA, and Pier, GB, Immunochemical Properties of the Staphylococcal

Poly‐N‐Acetylglucosamine Surface Polysaccharide. Infect. Immun., 2002. 70(8): p.

4433‐4440.

Maira‐Litran, T, Kropec, A, Goldmann, D, and Pier, GB, Biologic properties and

vaccine potential of the staphylococcal poly‐N‐acetyl glucosamine surface

polysaccharide. Vaccine, 2004. 22(7): p. 872‐879.

276

Marsh, LH, Coke, M, Dettmar, PW, Ewen, RJ, Havler, M, Nevell, TG, Smart, JD,

Smith, JR, Timmins, B, Tsibouklis, J, and Alexander, C, Adsorbed

poly(ethyleneoxide)–poly(propyleneoxide) copolymers on synthetic surfaces:

Spectroscopy and microscopy of polymer structures and effects on adhesion of skin‐

borne bacteria. Journal of Biomedical Materials Research, 2002. 61(4): p. 641‐

652.

Marshall, C, Lessard, I, Park, IS, and Wright, G, Glycopeptide antibiotic resistance

genes in glycopeptide‐producing organisms. Antimicrobial Agents and

Chemotherapy, 1998. 42(9): p. 2215‐2220.

Marshall, KC, Stout, R, and Mitchell, R, Mechanism of the Initial Events in the

Sorption of Marine Bacteria to Surfaces. J Gen Microbiol, 1971. 68(3): p. 337‐348.

Mathur, T, Singhal, S, Khan, S, Upadhyay, D, Fatma, T, and Rattan, A, Adverse

effect of staphylococci slime on in vitro activity of glycopeptides. Jpn J Infect Dis,

2005. 58(6): p. 353‐357.

Mayville, P, Ji, G, Beavis, R, Yang, H, Goger, M, Novick, RP, and Muir, TW,

Structure‐activity analysis of synthetic autoinducing thiolactone peptides from

Staphylococcus aureus responsible for virulence. Proceedings of the National

Academy of Sciences, 1999. 96(4): p. 1218‐1223.

277

McCormick, MH, Stark, WM, Pittenger, GE, Pittenger, RC, and McGuire, JM,

Vancomycin, a new antibiotic. I: Chemical and Biologic Properties. Antibiotics

Annual, 1956. 1955‐56: p. 606‐611.

McCrea, KW, Hartford, O, Davis, S, Eidhin, DN, Lina, G, Speziale, P, Foster, TJ,

and Hook, M, The serine‐aspartate repeat (Sdr) protein family in Staphylococcus

epidermidis. Microbiology‐Uk, 2000. 146: p. 1535‐1546.

McKenney, D, Pouliot, KL, Wang, Y, Murthy, V, Ulrich, M, DÃ¶ring, G, Lee, JC,

Goldmann, DA, and Pier, GB, Broadly Protective Vaccine for Staphylococcus

aureus Based on an in Vivo‐Expressed Antigen. Science, 1999. 284(5419): p. 1523‐

1527.

Mingeot‐Leclercq, M‐P, Glupczynski, Y, and Tulkens, PM, Aminoglycosides:

Activity and Resistance. Antimicrob. Agents Chemother., 1999. 43(4): p. 727‐

737.

Morar, M and Wright, GD, The Genomic Enzymology of Antibiotic Resistance.

Annual Review of Genetics, 2010. 44(1): p. 25‐51.

Mukhtar, TA and Wright, GD, Streptogramins, Oxazolidinones, and Other Inhibitors

of Bacterial Protein Synthesis. Chemical Reviews, 2005. 105(2): p. 529‐542.

Nagarajan, R, Antibacterial activities and modes of action of vancomycin and related

glycopeptides. Antimicrob Agents Chemother, 1991. 35(4): p. 605‐9.

278

Nagy, JA, Meinwald, YC, and Scheraga, HA, Immunochemical determination of

conformational equilibria for fragments of the B.beta. chain of fibrinogen.

Biochemistry, 1985. 24(4): p. 882‐887.

Nathan, C and Goldberg, FM, The profit problem in antibiotic R&D. Nat Rev Drug

Discov, 2005. 4(11): p. 887‐891.

Nejadnik, MR, van der Mei, HC, Norde, W, and Busscher, HJ, Bacterial adhesion

and growth on a polymer brush‐coating. Biomaterials, 2008. 29(30): p. 4117‐4121.

Nilsson, M, Frykberg, L, Flock, JI, Pei, L, Lindberg, M, and Guss, B, A fibrinogen‐

binding protein of Staphylococcus epidermidis. Infection and Immunity, 1998.

66(6): p. 2666‐2673.

Novick, R, Projan, S, Kornblum, J, Ross, H, Ji, G, Kreiswirth, B, Vandenesch, F,

Moghazeh, S, and Novick, R, The agr P2 operon: An autocatalytic sensory

transduction system in Staphylococcus aureus. Molecular and General Genetics

MGG, 1995. 248(4): p. 446‐458.

Novick, RP and Geisinger, E, Quorum Sensing in Staphylococci. Annual Review of

Genetics, 2008. 42(1): p. 541‐564.

OʹGara, JP and Humphreys, H, Staphylococcus epidermidis biofilms: importance and

implications. Journal of Medical Microbiology, 2001. 50(7): p. 582‐587.

OʹToole, G, Kaplan, HB, and Kolter, R, Biofilm Formation as Microbial Development.

Annual Review of Microbiology, 2000. 54(1): p. 49‐79.

279

Ohlsen, K and Lorenz, U, Immunotherapeutic strategies to combat staphylococcal

infections. International Journal of Medical Microbiology, 2010. 300(6): p. 402‐

410.

Oliphant, CM and Green, GM, Quinolones: a comprehensive review. American

family physician, 2002. 65(3): p. 455.

Olson, M, Ceri, H, Morck, D, Buret, A, and Read, R, Biofilm bacteria: formation and

comparative susceptibility to antibiotics. Can. J. Vet. Res., 2002. 66(2): p. 86‐92.

Oram, JD and Reiter, B, Inhibition of bacteria by lactoferrin and other iron‐chelating

agents. Biochimica et Biophysica Acta (BBA) ‐ General Subjects, 1968. 170(2):

p. 351‐365.

Otto, M, Quorum‐sensing control in Staphylococci ‐ a target for antimicrobial drug

therapy? Fems Microbiology Letters, 2004. 241(2): p. 135‐141.

Otto, M, Virulence factors of the coagulase‐negative staphylococci. Front Biosci, 2004.

9: p. 841‐63.

Otto, M, Staphylococcal biofilms. Curr Top Microbiol Immunol, 2008. 322: p. 207‐

28.

Otto, M, Staphylococcus epidermidis ‐ the ʹaccidentalʹ pathogen. Nat Rev Micro, 2009.

7(8): p. 555‐567.

280

Otto, M, Echner, H, Voelter, W, and Gotz, F, Pheromone Cross‐Inhibition between

Staphylococcus aureus and Staphylococcus epidermidis. Infection and Immunity,

2001. 69(3): p. 1957‐1960.

Otto, M, Sussmuth, R, Jung, G, and Gotz, F, Structure of the pheromone peptide of

the Staphylococcus epidermidis agr system. FEBS Lett, 1998. 424(1‐2): p. 89‐94.

Pace, JL and Yang, G, Glycopeptides: update on an old successful antibiotic class.

Biochemical pharmacology, 2006. 71(7): p. 968‐980.

Palmer, J, Flint, S, and Brooks, J, Bacterial cell attachment, the beginning of a biofilm.

Journal of Industrial Microbiology & Biotechnology, 2007. 34(9): p. 577‐

588.

Patel, JD, Ebert, M, Stokes, K, Ward, R, and Anderson, JM, Inhibition of bacterial

and leukocyte adhesion under shear stress conditions by material surface chemistry.

Journal of Biomaterials Science, Polymer Edition, 2003. 14: p. 279‐295.

Patel, JD, Krupka, T, and Anderson, JM, iNOS‐mediated generation of reactive

oxygen and nitrogen species by biomaterial‐adherent neutrophils. Journal of

Biomedical Materials Research Part A, 2007. 80A(2): p. 381‐390.

Patti, JM, Allen, BL, Mcgavin, MJ, and Hook, M, MSCRAMM‐Mediated Adherence

of Microorganisms to Host Tissues. Annual Review of Microbiology, 1994. 48: p.

585‐617.

281

Patti, JM and Hook, M, Microbial Adhesins Recognizing Extracellular‐Matrix

Macromolecules. Current Opinion In Cell Biology, 1994. 6(5): p. 752‐758.

Pei, L, Arvholm, IL, Lonnies, L, and Flock, JI, GST‐Fbe can recognize beta‐chains of

fibrin(ogen) on explanted materials. Journal of Chromatography B‐Analytical

Technologies in the Biomedical and Life Sciences, 2003. 786(1‐2): p. 319‐325.

Pei, L and Flock, JI, Lack of fbe, the gene for a fibrinogen‐binding protein from

Staphylococcus epidermidis, reduces its adherence to fibrinogen coated surfaces.

Microbial Pathogenesis, 2001. 31(4): p. 185‐193.

Pei, L, Palma, M, Nilsson, M, Guss, B, and Flock, JI, Functional studies of a

fibrinogen binding protein from Staphylococcus epidermidis. Infection and

Immunity, 1999. 67(9): p. 4525‐4530.

Ponnuraj, K, Bowden, MG, Davis, S, Gurusiddappa, S, Moore, D, Choe, D, Xu, Y,

Hook, M, and Narayana, SVL, A ʺdock, lock, and latchʺ structural model for a

staphylococcal adhesin binding to fibrinogen. Cell, 2003. 115(2): p. 217‐228.

Poole, K, Efflux‐mediated antimicrobial resistance. Journal of Antimicrobial

Chemotherapy, 2005. 56(1): p. 20.

Prabu, P, Dharmaraj, N, Aryal, S, Lee, BM, Ramesh, V, and Kim, HY, Preparation

and drug release activity of scaffolds containing collagen and poly(caprolactone).

Journal of Biomedical Materials Research Part A, 2006. 79A(1): p. 153‐158.

282

Qin, Z, Ou, Y, Yang, L, Zhu, Y, Tolker‐Nielsen, T, Molin, S, and Qu, D, Role of

autolysin‐mediated DNA release in biofilm formation of Staphylococcus epidermidis.

Microbiology, 2007. 153(7): p. 2083‐2092.

Queck, SY and Otto, M, Staphylococcus epidermidis and other Coagulase‐Negative

Staphylococci, in Staphylococcus Molecular Genetics, Lindsay, JA, Editor. 2008,

Caister Academic Press: Norfolk, UK.

Rai, M, Yadav, A, and Gade, A, Silver nanoparticles as a new generation of

antimicrobials. Biotechnology advances, 2009. 27(1): p. 76‐83.

Rani, SA, Pitts, B, and Stewart, PS, Rapid Diffusion of Fluorescent Tracers into

Staphylococcus epidermidis Biofilms Visualized by Time Lapse Microscopy.

Antimicrob. Agents Chemother., 2005. 49(2): p. 728‐732.

Rennermalm, A, Nilsson, M, and Flock, JI, Fibrinogen binding protein of

Staphylococcus epidermidis is a target for opsonic antibodies. Infection and

Immunity, 2004. 72(5): p. 3081‐3083.

Reynolds, PE, Structure, biochemistry and mechanism of action of glycopeptide

antibiotics. European Journal of Clinical Microbiology & Infectious Diseases,

1989. 8(11): p. 943‐950.

Rice, KC, Mann, EE, Endres, JL, Weiss, EC, Cassat, JE, Smeltzer, MS, and Bayles,

KW, The cidA murein hydrolase regulator contributes to DNA release and biofilm

283

development in Staphylococcus aureus. Proceedings of the National Academy of

Sciences, 2007. 104(19): p. 8113‐8118.

Riddles, PW, Blakeley, RL, Zerner, B, and C.H.W. Hirs, SNT, [8] Reassessment of

Ellmanʹs reagent, in Methods in Enzymology. 1983, Academic Press. p. 49‐60.

Riley, DK, Classen, DC, Stevens, LE, and Burke, JP, A large randomized clinical trial

of a silver‐impregnated urinary catheter: Lack of efficacy and staphylococcal

superinfection. The American Journal of Medicine, 1995. 98(4): p. 349‐356.

Rogers, KL, Fey, PD, and Rupp, ME, Coagulase‐Negative Staphylococcal Infections.

Infectious Disease Clinics of North America, 2009. 23(1): p. 73‐98.

Rohde, H, Burandt, E, Siemssen, N, Frommelt, L, Burdelski, C, Wurster, S,

Scherpe, S, Davies, A, Harris, L, and Horstkotte, M, Polysaccharide intercellular

adhesin or protein factors in biofilm accumulation of Staphylococcus epidermidis and

Staphylococcus aureus isolated from prosthetic hip and knee joint infections☆.

Biomaterials, 2007. 28(9): p. 1711‐1720.

Rohde, H, Burdelski, C, Bartscht, K, Hussain, M, Buck, F, Horstkotte, MA,

Knobloch, JKM, Heilmann, C, Herrmann, M, and Mack, D, Induction of

Staphylococcus epidermidis biofilm formation via proteolytic processing of the

accumulation‐associated protein by staphylococcal and host proteases. Molecular

Microbiology, 2005. 55(6): p. 1883‐1895.

284

Rohde, H, Frankenberger, S, Zähringer, U, and Mack, D, Structure, function and

contribution of polysaccharide intercellular adhesin (PIA) to Staphylococcus

epidermidis biofilm formation and pathogenesis of biomaterial‐associated infections.

European Journal of Cell Biology, 2010. 89(1): p. 103‐111.

Roosjen, A, Kaper, HJ, van der Mei, HC, Norde, W, and Busscher, HJ, Inhibition of

adhesion of yeasts and bacteria by poly(ethylene oxide)‐brushes on glass in a parallel

plate flow chamber. Microbiology, 2003. 149(11): p. 3239‐3246.

Rupp, ME, Fey, PD, Heilmann, C, and Gotz, F, Characterization of the importance of

Staphylococcus epidermidis autolysin and polysaccharide intercellular adhesin in the

pathogenesis of intravascular catheter‐associated infection in a rat model. Journal of

Infectious Diseases, 2001. 183(7): p. 1038‐1042.

Rupp, ME, Ulphani, JS, Fey, PD, Bartscht, K, and Mack, D, Characterization of the

importance of polysaccharide intercellular adhesin/hemagglutinin of Staphylococcus

epidermidis in the pathogenesis of biomaterial‐based infection in a mouse foreign body

infection model. Infection and Immunity, 1999. 67(5): p. 2627‐2632.

Rupp, ME, Ulphani, JS, Fey, PD, and Mack, D, Characterization of Staphylococcus

epidermidis polysaccharide intercellular adhesin/hemagglutinin in the pathogenesis of

intravascular catheter‐associated infection in a rat model. Infection and Immunity,

1999. 67(5): p. 2656‐2659.

285

Sabath, LD, Garner, C, Wilcox, C, and Finland, M, Susceptibility of Staphylococcus

aureus and Staphylococcus epidermidis to 65 antibiotics. Antimicrob Agents

Chemother, 1976. 9(6): p. 962‐9.

Sadovskaya, I, Vinogradov, E, Flahaut, S, Kogan, G, and Jabbouri, S, Extracellular

Carbohydrate‐Containing Polymers of a Model Biofilm‐Producing Strain,

Staphylococcus epidermidis RP62A. Infection and Immunity, 2005. 73(5): p.

3007‐3017.

Sadovskaya, I, Vinogradov, E, Li, J, and Jabbouri, S, Structural elucidation of the

extracellular and cell‐wall teichoic acids of Staphylococcus epidermidis RP62A, a

reference biofilm‐positive strain. Carbohydrate Research, 2004. 339(8): p. 1467‐

1473.

Sapatnekar, S, Kao, WJ, and Anderson, JM, Leukocyte—biomaterial interactions in

the presence of Staphylococcus epidermidis: Flow cytometric evaluation of leukocyte

activation (Student Research Award in the Hospital Intern, Resident, or Clinical

Fellow Category, 23rd Annual Meeting of the Society for Biomaterials, New Orleans,

LA, April 30–May 4, 1997). Journal of Biomedical Materials Research, 1997.

35(4): p. 409‐420.

Schierholz and J., Implant infections: a haven for opportunistic bacteria. Journal of

Hospital Infection, 2001. 49(2): p. 87‐93.

286

Schierholz, JM and Beuth, J, Implant infections: a haven for opportunistic bacteria.

Journal of Hospital Infection, 2001. 49(2): p. 87‐93.

Schneewind, O, Fowler, A, and Faull, KF, Structure of the Cell Wall Anchor of

Surface Proteins in Staphylococcus aureus. Science, 1995. 268(5207): p. 103‐106.

Schneewind, O, Mihaylova‐Petkov, D, and Model, P, Cell wall sorting signals in

surface proteins of gram‐positive bacteria. Embo J, 1993. 12(12): p. 4803‐11.

Seltmann, G and Holst, O, The Bacterial Cell Wall. 2002, New York: Springer.

Senior, R, Skogen, W, Griffin, G, and Wilner, G, Effects of Fibrinogen Derivatives

Upon the Inflammatory Response. J. Clin. Invest., 1986. 77: p. 1014‐1019.

Shima, S, Matsuoka, H, Iwamoto, T, and Sakai, H, Antimicrobial action of epsilon‐

poly‐L‐lysine. J Antibiot (Tokyo), 1984. 37(11): p. 1449‐55.

Shinabarger, DL, Marotti, KR, Murray, RW, Lin, AH, Melchior, EP, Swaney, SM,

Dunyak, DS, Demyan, WF, and Buysse, JM, Mechanism of action of

oxazolidinones: effects of linezolid and eperezolid on translation reactions.

Antimicrob. Agents Chemother., 1997. 41(10): p. 2132‐2136.

Shive, MS, Brodbeck, WG, Colton, E, and Anderson, JM, Shear stress and material

surface effects on adherent human monocyte apoptosis. Journal of Biomedical

Materials Research, 2002. 60(1): p. 148‐158.

287

Shive, MS, Hasan, SM, and Anderson, JM, Shear stress effects on bacterial adhesion,

leukocyte adhesion, and leukocyte oxidative capacity on a polyetherurethane. Journal

of Biomedical Materials Research, 1999. 46(4): p. 511‐519.

Shive, MS, Salloum, ML, and Anderson, JM, Shear stress‐induced apoptosis of

adherent neutrophils: A mechanism for persistence of cardiovascular device

infections. Proceedings of the National Academy of Sciences, 2000. 97(12): p.

6710‐6715.

Singh, R, Ray, P, Das, A, and Sharma, M, Penetration of antibiotics through

Staphylococcus aureus and Staphylococcus epidermidis biofilms. Journal of

Antimicrobial Chemotherapy, 2010. 65(9): p. 1955‐1958.

Sjollem, J, Busscher, HJ, and Weerkamp, AH, Experimental approaches for studying

adhesion of microorganisms to solid substrata: applications and mass transport.

Journal of Microbiological Methods, 1989. 9(2): p. 79‐90.

Sköld, O, Sulfonamide resistance: mechanisms and trends. Drug Resistance Updates,

2000. 3(3): p. 155‐160.

Slack, SM and Horbett, TA, The Vroman Effect: A Critical Review, in Proteins at

Interfaces II: Fundamentals and Applications, Horbett, TA and Brash, JL, Editors.

1995, American Chemical Society. p. 112‐128.

288

Smith, HW and Huggins, MB, Successful treatment of experimental Escherichia coli

infections in mice using phage: its general superiority over antibiotics. Journal of

General Microbiology, 1982. 128(2): p. 307.

Souli, M and Giamarellou, H, Effects of Slime Produced by Clinical Isolates of

Coagulase‐Negative Staphylococci on Activities of Various Antimicrobial Agents.

Antimicrob. Agents Chemother., 1998. 42(4): p. 939‐941.

Spizek, J and Rezanka, T, Lincomycin, clindamycin and their applications. Applied

Microbiology and Biotechnology, 2004. 64(4): p. 455‐464.

Spoering, AL and Lewis, K, Biofilms and planktonic cells of Pseudomonas aeruginosa

have similar resistance to killing by antimicrobials. Journal of bacteriology, 2001.

183(23): p. 6746.

Stewart, P and Williamcosterton, J, Antibiotic resistance of bacteria in biofilms. The

Lancet, 2001. 358(9276): p. 135‐138.

Stewart, PS, Theoretical aspects of antibiotic diffusion into microbial biofilms.

Antimicrob Agents Chemother, 1996. 40(11): p. 2517‐22.

Stewart, PS, Mechanisms of antibiotic resistance in bacterial biofilms. International

Journal of Medical Microbiology, 2002. 292(2): p. 107‐113.

Stewart, PS, Diffusion in Biofilms. J. Bacteriol., 2003. 185(5): p. 1485‐1491.

289

Stewart, PS, Davison, WM, and Steenbergen, JN, Daptomycin Rapidly Penetrates a

Staphylococcus epidermidis Biofilm. Antimicrob. Agents Chemother., 2009. 53(8):

p. 3505‐3507.

Stobie, N, Duffy, B, McCormack, DE, Colreavy, J, Hidalgo, M, McHale, P, and

Hinder, SJ, Prevention of Staphylococcus epidermidis biofilm formation using a low‐

temperature processed silver‐doped phenyltriethoxysilane solâ€“gel coating.

Biomaterials, 2008. 29(8): p. 963‐969.

Sun, D, Accavitti, MA, and Bryers, JD, Inhibition of Biofilm Formation by Monoclonal

Antibodies against Staphylococcus epidermidis RP62A Accumulation‐Associated

Protein. Clin. Diagn. Lab. Immunol., 2005. 12(1): p. 93‐100.

Swaney, SM, Aoki, H, Ganoza, MC, and Shinabarger, DL, The Oxazolidinone

Linezolid Inhibits Initiation of Protein Synthesis in Bacteria. Antimicrob. Agents

Chemother., 1998. 42(12): p. 3251‐3255.

Tenson, T, Lovmar, M, and Ehrenberg, M, The Mechanism of Action of Macrolides,

Lincosamides and Streptogramin B Reveals the Nascent Peptide Exit Path in the

Ribosome. Journal of Molecular Biology, 2003. 330(5): p. 1005‐1014.

Tipper, DJ and Berman, MF, Structures of the cell wall peptidoglycans of

Staphylococcus epidermidis Texas 26 and Staphylococcus aureus Copenhagen. I.

Chain length and average sequence of cross‐bridge peptides. Biochemistry, 1969.

8(5): p. 2183‐2192.

290

Tipper, DJ and Strominger, JL, Mechanism of action of penicillins: a proposal based on

their structural similarity to acyl‐D‐alanyl‐D‐alanine. Proceedings of the National

Academy of Sciences, 1965. 54(4): p. 1133‐1141.

Tojo, M, Yamashita, N, Goldmann, DA, and Pier, GB, Isolation and

Characterization of a Capsular Polysaccharide Adhesin from Staphylococcus

epidermidis. Journal of Infectious Diseases, 1988. 157(4): p. 713‐722.

Tormo, MA, Knecht, E, Gotz, F, Lasa, I, and Penades, JR, Bap‐dependent biofilm

formation by pathogenic species of Staphylococcus: evidence of horizontal gene

transfer? Microbiology, 2005. 151(7): p. 2465‐2475.

Trautner, BW and Darouiche, RO, Catheter‐Associated Infections: Pathogenesis

Affects Prevention. Arch Intern Med, 2004. 164(8): p. 842‐850.

Turnidge, J, Pharmacokinetics and pharmacodynamics of fluoroquinolones. Drugs,

1999. 58(S2): p. 29‐36.

Uçkay, I, Pittet, D, Vaudaux, P, Sax, H, Lew, D, and Waldvogel, F, Foreign body

infections due toStaphylococcus epidermidis. Annals of Medicine, 2009. 41(2): p.

109‐119.

Van Bambeke, F, Balzi, E, and Tulkens, PM, Antibiotic efflux pumps. Biochemical

pharmacology, 2000. 60: p. 457‐470.

Van Bambeke, F, Glupczynski, Y, Plesiat, P, Pechere, JC, and Tulkens, PM,

Antibiotic efflux pumps in prokaryotic cells: occurrence, impact on resistance and

291

strategies for the future of antimicrobial therapy. Journal of Antimicrobial

Chemotherapy, 2003. 51(5): p. 1055.

van Heijenoort, J, Formation of the glycan chains in the synthesis of bacterial

peptidoglycan. Glycobiology, 2001. 11(3): p. 25R‐36R. van Oss, CJ, Long‐range and short‐range mechanisms of hydrophobic attraction and

hydrophilic repulsion in specific and aspecific interactions. Journal of Molecular

Recognition, 2003. 16(4): p. 177‐190.

Van Oss, CJ, Interfacial Forces in Aqueous Media. 2 ed. 2006, Boca Raton, FL: CRC

Press.

Van Oss, CJ, Chaudhury, MK, and Good, RJ, Interfacial Lifshitz‐van der Waals and

polar interactions in macroscopic systems. Chemical Reviews, 1988. 88(6): p. 927‐

941.

Vasilev, K, Cook, J, and Griesser, HJ, Antibacterial surfaces for biomedical devices.

Expert Review of Medical Devices, 2009. 6(5): p. 553‐567.

Vernachio, JH, Bayer, AS, Ames, B, Bryant, D, Prater, BD, Syribeys, PJ, Gorovits,

EL, and Patti, JM, Human immunoglobulin G recognizing fibrinogen‐binding

surface proteins is protective against both Staphylococcus aureus and Staphylococcus

epidermidis infections in vivo. Antimicrobial Agents and Chemotherapy, 2006.

50(2): p. 511‐518.

292

Vimala, K, Samba Sivudu, K, Murali Mohan, Y, Sreedhar, B, and Mohana Raju,

K, Controlled silver nanoparticles synthesis in semi‐hydrogel networks of

poly(acrylamide) and carbohydrates: A rational methodology for antibacterial

application. Carbohydrate Polymers, 2009. 75(3): p. 463‐471.

Vollmer, W, Blanot, D, and De Pedro, MA, Peptidoglycan structure and architecture.

FEMS Microbiology Reviews, 2008. 32(2): p. 149‐167. von Eiff, C, Peters, G, and Heilmann, C, Pathogenesis of Infections Due to Coagulase‐

Negative Staphylococci. The Lancet Infectious Diseases, 2002. 2(11): p. 677‐685.

Vroman, L and Adams, AL, Findings with the recording ellipsometer suggesting rapid

exchange of specific plasma proteins at liquid/solid interfaces. Surface Science, 1969.

16: p. 438‐446.

Vuong, C, A Crucial Role for Exopolysaccharide Modification in Bacterial Biofilm

Formation, Immune Evasion, and Virulence. Journal of Biological Chemistry,

2004. 279(52): p. 54881‐54886.

Vuong, C, Gerke, C, Somerville, GA, Fischer, ER, and Otto, M, Quorum‐sensing

control of biofilm factors in Staphylococcus epidermidis. J Infect Dis, 2003. 188(5):

p. 706‐18.

Vuong, C, Kocianova, S, Yao, Y, Carmody, AB, and Otto, M, Increased colonization

of indwelling medical devices by quorum‐sensing mutants of Staphylococcus

epidermidis in vivo. J Infect Dis, 2004. 190(8): p. 1498‐505.

293

Vuong, C and Otto, M, Staphylococcus epidermidis infections. Microbes Infect, 2002.

4(4): p. 481‐9.

Vuong, C, Voyich, JM, Fischer, ER, Braughton, KR, Whitney, AR, DeLeo, FR, and

Otto, M, Polysaccharide intercellular adhesin (PIA) protects Staphylococcus

epidermidis against major components of the human innate immune system. Cell

Microbiol, 2004. 6(3): p. 269‐75.

Walters, MC, III, Roe, F, Bugnicourt, A, Franklin, MJ, and Stewart, PS,

Contributions of Antibiotic Penetration, Oxygen Limitation, and Low Metabolic

Activity to Tolerance of Pseudomonas aeruginosa Biofilms to Ciprofloxacin and

Tobramycin. Antimicrob. Agents Chemother., 2003. 47(1): p. 317‐323.

Wang, I‐w, Anderson, JM, and Marchant, RE, Staphylococcus epidermidis Adhesion

to Hydrophobic Biomedical Polymer Is Mediated by Platelets. Journal of Infectious

Diseases, 1993. 167(2): p. 329‐336.

Watanakunakorn, C, The antibacterial action of vancomycin. Rev Infect Dis, 1981. 3

suppl: p. S210‐5.

Watnick, P and Kolter, R, Biofilm, City of Microbes. J. Bacteriol., 2000. 182(10): p.

2675‐2679.

Waxman, DJ and Strominger, JL, Penicillin‐Binding Proteins and the Mechanism of

Action of Beta‐Lactam Antibiotics1. Annual Review of Biochemistry, 1983. 52(1):

p. 825‐869.

294

Wei, J, Ravn, DB, Gram, L, and Kingshott, P, Stainless steel modified with

poly(ethylene glycol) can prevent protein adsorption but not bacterial adhesion.

Colloids and Surfaces B: Biointerfaces, 2003. 32(4): p. 275‐291.

Werner, E, Roe, F, Bugnicourt, A, Franklin, MJ, Heydorn, A, Molin, S, Pitts, B,

and Stewart, PS, Stratified Growth in Pseudomonas aeruginosa Biofilms. Appl.

Environ. Microbiol., 2004. 70(10): p. 6188‐6196.

Westwell, MS, Bardsley, B, Dancer, RJ, Try, AC, and Williams, DH, Cooperativity

in ligand binding expressed at a model cell membrane by the vancomycin group

antibiotics. Chemical Communications, 1996(5): p. 589‐590.

Williams, DH, The glycopeptide story‐‐how to kill the deadly ʹsuperbugsʹ. Nat Prod

Rep, 1996. 13(6): p. 469‐77.

Williams, RJ, Henderson, B, Sharp, LJ, and Nair, SP, Identification of a fibronectin‐

binding protein from Staphylococcus epidermidis. Infection And Immunity, 2002.

70(12): p. 6805‐6810.

Wilson, DN, Schluenzen, F, Harms, JM, Starosta, AL, Connell, SR, and Fucini, P,

The oxazolidinone antibiotics perturb the ribosomal peptidyl‐transferase center and

effect tRNA positioning. Proceedings of the National Academy of Sciences,

2008. 105(36): p. 13339‐13344.

295

Wimley, WC, Selsted, ME, and White, SH, Interactions between human defensins

and lipid bilayers: Evidence for formation of multimeric pores. Protein Science,

1994. 3(9): p. 1362‐1373.

Wright, GD, Molecular mechanisms of antibiotic resistance. Chem. Commun., 2011.

Xia, Y and Zweier, JL, Superoxide and peroxynitrite generation from inducible nitric

oxide synthaseâ€‰inâ€‰macrophages. Proceedings of the National Academy of

Sciences, 1997. 94(13): p. 6954‐6958.

Xu, KD, Stewart, PS, Xia, F, Huang, C‐T, and McFeters, GA, Spatial Physiological

Heterogeneity in Pseudomonas aeruginosa Biofilm Is Determined by Oxygen

Availability. Appl. Environ. Microbiol., 1998. 64(10): p. 4035‐4039.

Xu, L, Li, H, Vuong, C, Vadyvaloo, V, Wang, J, Yao, Y, Otto, M, and Gao, Q, Role

of the luxS Quorum‐Sensing System in Biofilm Formation and Virulence of

Staphylococcus epidermidis. Infection and Immunity, 2005. 74(1): p. 488‐496.

YANAGISAWA, N, LI, D‐Q, and LJUNGH, A, The N‐terminal of thrombospondin‐1

is essential for coagulase‐negative staphylococcal binding. J Med Microbiol, 2001.

50(8): p. 712‐719.

Yang, D, Chen, Q, Chertov, O, and Oppenheim, JJ, Human neutrophil defensins

selectively chemoattract naive T and immature dendritic cells. Journal of Leukocyte

Biology, 2000. 68(1): p. 9‐14.

296

Zhou, Y, Doerschuk, CM, Anderson, JM, and Marchant, RE, Biomaterial surface‐

dependent neutrophil mobility. Journal of Biomedical Materials Research Part A,

2004. 69A(4): p. 611‐620.

Ziats, NP, Miller, KM, and Anderson, JM, In vitro and in vivo interactions of cells

with biomaterials. Biomaterials, 1988. 9(1): p. 5‐13.

297