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Innovative coatings for anti-bacterial surfacesSwartjes, Jan

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Innovative coatings for anti-

bacterial surfaces

Johannes Julius Thomas Marinus Swartjes

The printing of this thesis was financially supported by Ducom Instruments.

Innovative coatings for anti-bacterial surfaces

By Johannes Julius Thomas Marinus Swartjes

University Medical Center Groningen, University of Groningen

Groningen, The Netherlands

Copyright © 2014 by Johannes Julius Thomas Marinus Swartjes

Printed by Off-page Amsterdam, The Netherlands

ISBN (printed version): 978-90-367-7562-5

ISBN (electronic version): 978-90-367-7561-8

Innovative coatings for anti-bacterial surfaces

Proefschrift

ter verkrijging van de graad van doctor aan de

Rijksuniversiteit Groningen

op gezag van de

rector magnificus prof. dr. E. Sterken

en volgens besluit van het College voor Promoties.

De openbare verdediging zal plaatsvinden op

maandag 26 januari 2015 om 16:15 uur

door

Johannes Julius Thomas Marinus Swartjes

geboren op 8 oktober 1986

te Hoogezand-Sappemeer

Promotores

Prof. dr. ir. H.J. Busscher

Prof. dr. H.C. van der Mei

Copromotor

Dr. P.K. Sharma

Beoordelingscommissie

Prof. dr. A. Herrmann

Prof. dr. P. Buma

Prof. dr. L. W. van Rhijn

To my dear parents

Paranimfen

Deepak H. Veeregowda

Edward T.J. Rochford

CONTENTS

Chapter 1 General introduction: current developments in antimicrobial surface coatings

9

Aim of Thesis 31

Chapter 2 Length-scale mediated differential adhesion of mammalian cells and microbes

41

Chapter 3 A functional DNase I coating to prevent adhesion of bacteria and the formation of biofilm

59

Chapter 4 A PLGA-coating releasing inulin protected DNase I to prevent adhesion of bacteria and the formation of biofilm

75

Chapter 5 Normally oriented adhesion versus friction forces in bacterial adhesion to polymer-brush functionalized surfaces under fluid flow

93

Chapter 6 Antigen I/II mediated binding of S. mutans to salivary films provides an anchoring mechanism to withstand shear

109

Chapter 7 General discussion

121

Summary 127

Samenvatting 133

CHAPTER 1

GENERAL INTRODUCTION AND

AIM OF THE THESIS

CURRENT DEVELOPMENTS IN ANTIMICROBIAL SURFACE COATINGS FOR

BIOMEDICAL APPLICATIONS

Jan J.T.M. Swartjes, Prashant K. Sharma, Theo G. van Kooten, Henny C. van der Mei,

Morteza Mahmoudi, Henk J. Busscher and Edward T.J. Rochford

Curr. Med. Chem. 2014, DOI:10.2174/0929867321666140916121355

CHAPTER 1

10

ABSTRACT

Bacterial adhesion and subsequent biofilm formation on material surfaces represent a serious problem in

society from both an economical and health perspective. Surface coating approaches to prevent bacterial

adhesion and biofilm formation are of increased importance due to the increasing prevalence of antibiotic

resistant bacterial strains. Effective antimicrobial surface coatings can be based on an anti-adhesive

principle that prevents bacteria to adhere, or on bactericidal strategies, killing organisms either before or

after contact is made with the surface. Many strategies, however, implement a multi-functional approach

that incorporates both of these mechanisms. Notwithstanding the ubiquitous nature of the problem of

microbial colonization of material surfaces, this review focuses on the recent developments in

antimicrobial surface coatings with respect to biomaterial implants and devices. In this biomedical arena,

to rank the different coating strategies in order of increasing efficacy is impossible, since this depends on

the clinical application aimed for and whether expectations are short- or long term.

CURRENT DEVELOPMENTS IN ANTIMICROBIAL SURFACE COATINGS

11

INTRODUCTION

Bacterial adhesion and subsequent biofilm formation on material surfaces represent a serious problem in

society from both an economical and health perspective [1–3]. Biofilms formed in industrial settings like

pipelines, water treatment plants, heat exchangers and on ship hulls, contribute to decreased efficiency

accompanied with huge increases in operating costs. Furthermore, microbial adhesion and growth on food

processing equipment, but even more so on medical devices and implants, can cause serious

complications to human health. Whereas mechanical removal of biofilms in industrial settings is expensive,

but usually effective, in medical applications removal represents a last resort solution. Biofilm detachment

and mechanical removal from biomaterial-associated infections means extensive debridement and high-

risk revision surgery accompanied by increased risks of further infectious complications. Treatment and

prevention methods include the use of antibiotics, but the low sensitivity of bacteria to antibiotics induced

by the biofilm mode of growth, together with the increasing number of multi-resistant strains, makes their

use currently less effective than it has ever been [4–6].

As an alternative to the use of antibiotics to prevent bacteria from causing infection or to treat established

biofilms, the development of new materials or surface coatings that prevent viable bacteria from adhering

has been the center of attention in many studies [7–9]. Since the first step of bacteria in developing into

a highly resistant biofilm, is to adhere firmly onto a surface, interfering with this step can reduce infection

risks. This is achieved not only by preventing biofilm formation, but also by maintaining bacteria in their

planktonic, non-adhering state, which means these pathogens are more sensitive to antibiotics and

clearance by the immune system. For decades, the method of choice to create these so-called anti-

adhesive coatings has been the modification of surfaces with polymer brushes [10, 11]. When sufficiently

long polymer chains are grafted to a surface at a high enough surface density, a steric barrier is created

which can prevent adhesion of proteins and bacteria. Polyethylene glycol (PEG) was one of the first

polymers used to this end and demonstrated log reductions in adhesion of both proteins and bacterial

cells. This led to a period in which several variants of PEG-based brush coatings were designed and

evaluated [11–13]. However, the use of polymer brushes never achieves complete adhesion prevention

and the few bacteria that do manage to adhere still demonstrate the capacity of growing into a biofilm

[11].

To date, the original thought that rather simple polymer brushes would be sufficient for preventing

implants and medical devices to become colonized by bacteria has been surpassed by the realization that

multiple functionalities, including tissue integrative ones, need to be combined in one surface coating in

order to effectively prevent implant and device colonization [14]. Additionally, the diverse range of

implants applied in the clinical setting requires the design of any future antimicrobial coating to be

carefully matched to the intended application. For example, the requirements of a coating for a short-

term catheter differ dramatically from those of a permanent hip implant. Consequently, during the design

process a number of variables must be considered, the first of which is the duration of the coating efficacy.

CHAPTER 1

12

Microorganisms can be encountered pre-operatively from wound contamination, peri-operatively from

the operating room or contaminated equipment and post-operatively over the lifetime of the implant via

hematogenous seeding [15]. For a short term implant these three contamination mechanisms can be

treated similarly; however, for long term implants a compromise needs to be made for the duration of

protection afforded by the coating: simply over the early high risk period or also the long-term low risk

period. A second consideration is whether the mechanism applied should release antimicrobials or present

the active component bound to the surface. The release of antimicrobial compounds is beneficial as it not

only kills microorganisms associated with the implant surface directly, but also any susceptible pathogens

in the surrounding area. However there is a caveat, the release profiles of such coatings are often difficult

to effectively control and often inappropriate concentrations of antimicrobials are released. For many of

these coatings an initial massive burst of the active component is delivered followed by a longer period of

diminishing release. It is during this latter phase that bacteria may be exposed to sub-inhibitory

concentrations of antimicrobials which is conducive to the development of resistance and therefore may

render the coating ineffective [16].

In this review, we aim to provide an overview of the current developments in antimicrobial surface

coatings for use in the biomedical field, over the past few years. An overview will be given of the main

types of antimicrobial strategies for surface coating: use of antimicrobial peptides (AMPs), antibiotics,

enzymes, nanoparticles (NPs), quaternary ammonium compounds (QACs), anti-adhesive polymers, super

hydrophobic coatings and chitosan based strategies. Figure 1 shows four example strategies for creating

antimicrobial surface coatings, e.g. surface immobilization of antimicrobials, surface coatings designed to

release antimicrobial into the surrounding, hydrogel or other matrix structures containing bound

antimicrobials and antimicrobials tethered to a surface through spacer-molecules.

FIGURE 1 Schematic presentation of four example strategies for antimicrobial surface coating of materials. Combinations of strategies to achieve optimum results are often applied.

Surface immobilized antimicrobials

Hydrogel contained antimicrobials Surface tethered antimicrobials

Antimicrobial releasing surface coating


13

OVERVIEW OF ANTIMICROBIAL SURFACE COATINGS

Antibiotics

The most commonly used antimicrobials are antibiotics, of which penicillin is perhaps the most well-known

and one of the earliest to be applied in a medical setting. In the decades after its discovery, manufacturing

methods were simplified and new formulations were discovered, making the use of antibiotics widespread

[6]. The dark side of the wide availability and use of antibiotics turned out to be the rise of bacterial strains

that had developed resistance against one or more antibiotic agents. Methicillin resistant Staphylococcus

aureus for example is one of the most notorious among these strains and although the name suggests it

to be only resistant to methicillin, in reality the resistance profile is often not just restricted to methicillin

[17]. Despite the rise of resistant strains, antibiotics are still widely used and subject of new research to

develop antibacterial coatings for a number of reasons. Firstly, because of the relative ease of translating

techniques involving currently used antibiotics to new generations of antibiotics. Secondly, because

developments in surface coating technology permit controlled localized release which decreases the risk

of bacterial resistance development compared to systemic administration [18]. However, sub-inhibitory

concentrations of antibiotics remain to form a high risk factor in antibiotic releasing materials and coatings

[16].

Clinically, the application of antibiotic releasing hydroxyapatite (HAP) is common practice in orthopedic

surgery; bone implants are often coated with antibiotic releasing HAP to prevent infection while at the

same time promoting bone ingrowth [19, 20]. Belcarz et al. modified HAP by addition of β-1,3-glucan,

creating an elastic composite coating that was able to bind antibiotics by ionic interactions and released

the majority of the drug during the first 48 h, with a very short period of drug release at sub-inhibitory

concentrations [21]. Avoiding the problem of release of sub-inhibitory concentrations was approached in

a different way by Noble et al., who created a poly(2-hydroxyethyl methacrylate) (pHEMA) polymeric

monolith and added a self-assembled multilayer (SAM) coating of long methylene chains [22]. Addition of

ciprofloxacin resulted in an antibiotic releasing coating that could be switched “on” and “off” by using

ultrasound. After application of ultrasound the methylene chains re-organized to a relatively impermeable

self-assembled coating stopping the release of antibiotic. Although a small amount of background release

was observed, the applied system is a promising way of delivering antibiotics on-demand.

In addition, release of antibiotics by coating degradation is possible by using degradable polymers such as

poly(D,L-lactide), poly(ε-caprolactone) or poly(trimethylene carbonate) [23–25]. Combining different

degradable polymers into a multilayer system offers the opportunity to include multiple antibiotics that

allow modulation of the release profile per antibiotic [24] and additionally degradable surfaces may be

inherently resistant to infection [26]. An alternative method to obtain multilayer systems has been

described by Shukla and co-workers who applied tetra-layers of (poly-2-dextran

sulfate/vancomycin/dextran sulfate) by spray coating [27]. To this end, a vacuum was applied to the back

of a porous gelatin surface and each individual layer was sprayed on, followed by a rinsing step. The tetra-

CHAPTER 1

14

layer system on gelatin sponges showed more linear release kinetics compared to flat substrates,

expanding the time of release by 100 h. Additionally, hydrolytically degradable polyelectrolyte multilayers

manufactured by Wong et al. consisted of a non-degradable bactericidal base bilayer of N,N-

dodecyl,methyl-poly(ethyleneimine) (DMLPEI) and poly(acrylic acid) (PAA) on plasma-etched silicon

topped with the degradable gentamicin sulfate (GS) containing top layer. This top layer consisted of

(PAA/GS/PAA) tetra-layers in which hydrolytically degradable poly(β-amino-ester) was included [28].

These films showed high burst release of gentamicin in the first hours, while the bactericidal base-coating

prevented bacterial colonization of substrates by S. aureus for up to two weeks.

In contrast to release coatings, surface binding of antibiotic agents creates a high local concentration,

minimizing the risk of bacterial exposure to sub-inhibitory concentrations and thereby reduces the risk of

resistance. Current immobilization studies focus mainly on binding of vancomycin, which is considered to

be a last resort in treatment of infections caused by multi-resistant bacterial strains [29]. Since the working

mechanism of vancomycin requires penetration of the cell wall, surface tethering is generally performed

by including spacers that allow for a certain degree of freedom to penetrate the cell wall. Jose et al. used

a double aminoethoxyethoxyacetate linker combined with a 3-aminopropyltriethoxysilane modified

titanium surface, to provide a vancomycin surface distance of about 4 nm [30]. Surface coating of titanium

particles confirmed that the vancomycin-surface distance was sufficient to retain antimicrobial activity,

reducing S. aureus colony-forming units by 88% over two hours, while repeated exposure to bacterial

suspensions did not alter the antimicrobial activity. Swanson et al. passivated titanium surfaces to increase

the amount of hydroxide groups, which were then changed to amine functional groups through a 3-

aminopropyl-triethoxysilane reaction. The amine functional groups were converted to aldehyde groups

via a glutaraldehyde reaction and were bonded to the amine functional group of chitosan. This layer was

used to subsequently promote the binding of a chitosan-vancomycin mixture, creating a surface coating

capable of giving a zone of inhibition for S. aureus similar to the use of standard freely soluble vancomycin

[31].

With increased antibiotic resistance, focus on alternative antimicrobial therapeutics is gaining, but

controlled antibiotic therapy by means of surface coating remains an enticing topic. Not only to maintain

the current last-resort antibiotics such as vancomycin, but also to be able to responsibly use new, future

formulations of antibiotics in order to avoid development of resistant strains shortly after they are

introduced.

Antimicrobial peptides

The concept of using peptides against microbial attack, is not a recent development and in fact has been

employed by nature, as shown by the antimicrobial peptides that are part of the innate immune system

[32, 33]. The current augmented attention in science towards the use of AMPs in antimicrobial applications

is largely due to their broad antimicrobial spectrum which includes both Gram-positive and Gram-negative


15

bacteria and even viruses [34, 35] with relatively little induction of resistance among its target organisms

[36, 37]. Additional to the broad range of susceptible microorganisms, AMPs are effective against strains

that have developed a high degree of antibiotic resistance; for example, methicillin resistant S. aureus [38].

AMPs generally have an overall positive charge and contain a large portion of hydrophobic residues. Their

antibacterial activity comes from association with the negatively charged bacterial cell wall, after which

hydrophobic interactions of accumulated AMPs disrupt the cell wall [35]. Together these characteristics

mean that AMPs are well suited for incorporation in surface coatings and therefore this area of research

has obtained the attention of many investigations.

Built-up from a variety of amino-acids, AMPs are suitable for surface attachment by various coupling

mechanisms [39]. The primary amine-groups associated with most amino acids can be used to directly

couple AMPs to activated surfaces containing aldehyde, carboxyl or NHS modifications [34, 40, 41]. N-

terminal coupling of AMPs to gold has been achieved by modification of the surface using 11-

mercaptoundecanoic acid (MUA), which is subsequently activated using EDC and NHS. After this, the

amine group of the AMP can react with the activated carboxylic terminal of the established MUA

monolayer (see Figure 2) [42]. The stability of this approach is emphasized by Humblot et al. who

demonstrated a 40% reduction in bacterial adhesion after six months of storage at 4°C, including exposure

of the coated surfaces to four bacterial adhesion assays during this six months period [43]. Coupling of the

AMP gramicidin A onto gold has also been successfully achieved by modifying gold surfaces using

cystamine, which was then allowed to react with the aldehyde functional group at the NH2 terminus of

gramicidin A, formed by natural formylation [44].

Direct and rather uncomplicated coupling of AMPs is preferable and possible on e.g. gold surfaces as well

as titanium [45], but coating of implants for orthopedic applications might require additional surface

modification to make the implant more suitable to fulfill its function inside the body. Calcium phosphate

(CaP) has been known to enhance bone growth on orthopedic implants and a system in which an AMP

(Tet213) was added by absorbing it into micro-porous CaP coated titanium showed high antimicrobial

activity against Pseudomonas aeruginosa [46]. In another study, the antimicrobial peptide HHC-36 was

incorporated into a multilayer system of CaP on TiO2 nanotubes [47]. The titanium nanotubes were loaded

with AMPs using vacuum-assisted physical adsorption, while the CaP was loaded by applying an AMP

solution in ethanol and letting it dry in air. For a better controlled release profile a phospholipid layer was

added on top of the CaP, to create a bio-inspired cell membrane. The modified surfaces showed sufficient

release of AMPs to kill S. aureus and P. aeruginosa, while osteoblast-like cells were able to attach to the

implants and no cytotoxicity against these cells was observed after five days. The difference of these AMP

loaded surfaces compared to directly coupled AMPs by the aforementioned surface chemistry is that

rather than killing bacteria upon contact, the AMPs are released and bacteria in the vicinity of the surface

are killed before they can adhere.

CHAPTER 1

16

FIGURE 2 Scheme showing magainin I immobilization on gold by 11-mercaptoundecanoic acid and 6-mercaptohexanol modification (1:3 ratio) of the surface, followed by esterification using NHS/EDC and ultimately coupling of magainin I. Adapted from [43] and reprinted with permission.

An additional method to load AMPs onto the surface of materials from which they then are released and

kill bacteria close-by, is to apply hydrogels with incorporated AMPs. This mechanism has been applied by

immersion of a dry poly(2-hydroxyethyl methacrylate) or poly(methacrylic acid) (PMAA) hydrogel in

solutions containing the desired AMP [48, 49]. Further to hydrogel coatings that release their AMP load,

AMPs can also be attached to the surface of, or within, the hydrogel, employing contact killing combined

with the anti-adhesiveness that some hydrogels exert [50]. PEG based hydrogels containing AMPs have

been prepared by mixing photo-polymerizable epsilon-poly-L-lysine-graft-methacrylamide with

poly(ethylene glycol) diacrylate and dimethyl-acrylamide followed by UV treatment [51]. These hydrogels

were attached to fluoroalkyl fumarate copolymer disks by plasma-UV induced surface grafting

polymerization; after argon plasma treatment of the surface the hydrogel precursor solution was cross-

linked by UV exposure. These hydrogel modified surfaces subsequently demonstrated 1 to 6-log

reductions in adhering microorganisms for six different strains (Escherichia coli, P. aeruginosa, Serratia

marcescens, S. aureus, Candida albicans, Fusarium solani) demonstrating the potential of these coatings.

Alternatively, surface tethering of AMPs using larger polymer chains, offer the non-adhesive advantage of

brush-like structures, while at the same time allowing freedom of movement for AMPs to optimize their

efficacy. The increased efficacy offered by more mobile adhesion of AMPs has been demonstrated by

comparing cathelin LL37 directly coupled to epoxy-silanized titanium surfaces with attachment including

a PEG spacer (using α-amino-ω-carboxy-PEG), and is supported by the observation that immobilization of

Au Au Au


17

AMPs reduces their activity compared to free soluble peptides [52, 53]. However, the efficacy of AMPs

depends on the appropriate chain length and the AMP used. For example, some AMPs require the

penetration of bacterial cell walls to function, if a short chain length prevents this the AMP is rendered

ineffective.

Shalev et al. used a bio-inspired approach by depositing a polydopamine layer on several kinds of surfaces

and subsequently coupling an ultra-short lipopeptide to the formed polydopamine layer, which resulted

in a non-leaching coating of covalently coupled AMPs with high killing efficiency against E. coli and S. aureus

[54]. In a study using a similar coating approach, a catechol derivative was used to attach a double amine-

functionalized PEG linker to titanium surfaces after which Magainin I, a well-known AMP, was attached.

By combining anti-adhesive with antimicrobial properties in this way, reductions in bacterial adhesion of

more than 90% were achieved [55].

AMPs offer high antimicrobial efficiency and the wide variety of possibilities to incorporate them into

surface coatings demonstrates the relative ease by which they can be chemically modified. However, for

future applications and surface coating development based on the antimicrobial properties of peptides, it

is important to consider the mode of action of the desired coating. Releasing coatings can deplete rapidly

if AMPs are released too quickly, while for surface tethered AMPs the efficacy can largely depend on the

chain length of the spacer molecule [56]. Although most AMPs are considered biocompatible, and indeed

do not show any direct toxicity to eukaryotic cells, worries are expressed because of their resemblance to

some eukaryotic signaling peptides [35] and possible hemolytic effects [57]. This alternative form of

toxicity by mimicking host peptides could potentially induce unwanted cell responses and requires

additional attention, before the use of AMPs can be considered completely safe.

Antibacterial enzymes

The use of enzymes is common in detergents, industrial processes and the food industry. Considered as

non-toxic bioactive non-fouling compounds, enzymes are being recognized as a valuable source for

production of antimicrobial surface coatings [58]. The biocompatibility of enzymes is evident due to the

natural source of these agents and presence in the human body. Enzymes serve as catalysts for chemical

reactions, increasing the rate and efficiency at which they take place by lowering the activation energy of

the reaction. Regarding the adhesion of bacteria, enzymes can either interfere with the adhesion

mechanism used by bacteria to adhere to a surface, or they can kill bacteria. Killing is achieved by catalyzing

hydrolysis of parts of the peptidoglycan cell wall, leading to lysis of the cell. Whilst interference with the

adhesion mechanism can be achieved by enzymatic degradation or rearrangement of molecules, or

molecular-assemblies, essential for adhesion, e.g. extracellular DNA (eDNA), adhesive proteins or

carbohydrates.

CHAPTER 1

18

Retaining enzymatic activity is a pre-requisite for any effective enzyme surface coatings; however, this can

be difficult to achieve. Although most enzymes demonstrate optimal efficacy at physiological conditions,

stability beyond these conditions can be limited. Additionally, the conformational structure of an enzyme

is of key importance for their activity to ensure optimal accessibility to the active site. Providing flexibility

of the enzyme is one way to keep its activity after surface immobilization, as Muszanska et al.

demonstrated by using poly-ethylene oxide (PEO) to attach lysozyme to silicone rubber [59]. To this end,

Pluronic F-127 (PEO99-PPO65-PEO99) was modified to change the PEO hydroxyl end-groups into aldehyde

functionalities, which reacted with the amine groups of lysozyme from chicken egg white. The hydrophobic

polypropyleneoxide (PPO) backbone of the Pluronic induced the formation of micelles which were

adsorbed to hydrophobic silicone rubber surfaces, creating a polymer brush with lysozyme functionalities.

They showed that lysozyme functionalization of 1% of the Pluronic preserved the anti-adhesive properties

of the brush against Bacillus subtilis whilst the lysozyme remained active, based on the increased fraction

of dead bacteria. Yuan et al. used PEG and lysozyme in a “grafting from” approach, by dopamine mediated

coating of a terminal alkyl halide initiator on stainless steel surfaces, followed by surface-initiated atom

transfer radical polymerization (ATRP) of PEG-monomethacrylate, after which lysozyme was coupled to

the chain end of PEG branches using 1,1`-carbonyldiimidazole as a bio-functional linker [60]. Because of

the broad-spectrum of lysozyme as an antimicrobial, it has been extensively used in many more types of

surface coatings, including layer-by-layer assembly based on electrostatic interactions [61–63],

immobilization using Fischer carbine complex [64] and in mesoporous release systems [65].

As an important component of extracellular polymeric substances, eDNA was shown to be vital for

bacterial adhesion as well as biofilm formation in several bacterial strains [66]. Swartjes et al.

demonstrated that enzymatic cleavage of eDNA by a functional DNase I surface coating was effective in

disrupting the extracellular polymeric substances of bacteria, and yielded a reduction in adhering bacteria

of 99% for P. aeruginosa and 95% for S. aureus, while 14 h biofilms formed by these strains were reduced

to thicknesses of 0.2 and 3 µm, respectively [67]. By applying polydopamine as an intermediate coupling

layer on polymethylmethacrylate, DNase I was bound by Michael addition reactions, yielding a DNase I

coating (see Figure 3) that retained its ability to degrade DNA for at least 14 h without leakage of active

enzyme.


19

FIGURE 3 Formation of polydopamine films on polymethylmethacrylate (PMMA) and attachment of DNase I by Michael addition reaction. Reprinted with permission from [67].

A glycoside hydrolase called dispersin B (DspB), which cleaves poly-N-acetylglucosamine polysaccharides,

is another example of an enzyme known to disturb biofilm formation by specifically attacking an

extracellular polymeric substances component necessary for biofilm formation [68]. Pavlukhina et al.

showed that a DspB loaded coating was able to reduce Staphylococcus epidermidis surface coverage by

98% [69]. PMAA surface hydrogels on silicon wafers were prepared by depositing bilayers of

PMAA/poly(allylamine hydrochloride) (PAH) which were cross-linked using glutaraldehyde. Incorporation

of DspB was performed overnight by submersion in a 0.5 mg mL-1 solution and the resulting coating

showed complete retention of DspB at a wide pH range. In another study, DspB was “grafted onto”

surfaces by using a poly(dimethylaminoethyl methacrylate) with quinone functionalities as a glue layer on

top of which five bilayers of PAH together with oxidized dopamine moieties (Pox(mDOPA)) were cross-

linked [70]. By applying a top layer of Pox(mDOPA), the surface was then rendered active towards grafting

of DspB. Coating of DspB by this method decreased the number of viable S. epidermidis bacteria in 24 h

old biofilms by 97%.

TRIS pH 8.5

PMMA

Polydopamine film on PMMA

DNase I coated PMMA

Dopamine

CHAPTER 1

20

Several enzymes that are able to interfere with bacterial adhesion have been coated and whether they

attack and kill bacteria or whether they target essential parts of the adhesion mechanism, their enzymatic

activity can reduce adhesion and proliferation. Essential to the efficacy of surface coating enzymes is to

retain their full, or at least most of their function. The examples highlighted here show that several

approaches are possible and enzymes can be used in combination with other anti-adhesive coatings, like

polymer brushes, to increase the overall effect.

Nanoparticles

When the size of certain materials reaches the nano-scale, the chemical, electrical, mechanical and optical

properties can change completely compared to the bulk material. Nanoparticles (NPs) have been known

to possess antibacterial properties for quite some time and besides for their effects in solution, NPs have

been applied in surface coatings and release systems. Whereas most antibacterial agents, such as

antibiotics, are developed for a specific target within bacteria, e.g. the cell wall or vital components in the

cytoplasm, NPs were generally designed for other applications and more or less serendipitously found to

have properties which make them suitable against bacterial adhesion and growth. Even though in many

cases the exact mechanisms of NP toxicity against bacteria are not fully understood, it is clear that in some

cases NPs are able to attach to the bacterial cell wall by electrostatic interactions and disrupt the cell

membrane [55, 56]. Another general mechanism of bacterial toxicity by NPs is through the generation of

reactive oxygen species which induces oxidative stress by free radical formation [57]. A more extensive

overview of the killing mechanism by different kinds of NPs for several strains of bacteria is presented by

Mahmoudi et al. [71].

The bactericidal effect of silver is known for many years and silver-ions and silver-based materials have

been used as disinfectants and as an antimicrobial in paints [72]. When considering antibacterial NPs, silver

is still most abundantly represented, although other metals are increasingly being studied. Direct

immobilization of silver NPs (Ag-NPs) to glass can be achieved by modification of glass surfaces with

aminopropyl-triethoxysilane and placing it in a colloidal suspension of Ag-NPs afterwards. The survival of

S. epidermdis on these Ag-NP modified glass surfaces after 24 h incubation at 37°C was 105 times lower

than that on control glass surfaces. Chen et al. have successfully incorporated Ag-NP into layered double

hydroxides (LDHs) on titanium plates [73]. The nanoporous Mg-Al LDHs resulting from hydrothermal

attachment to Ti were immersed in AgNO3 solution at 100°C, resulting in the formation of Ag-NPs on the

LDH covered surface. Transmission electron microscopy images revealed that the Ag-NPs were well

dispersed on the surface and that the majority of the particles was in the range of 5–20 nm. Experiments

on their antibacterial properties showed 99% reduction in the number of adhering organisms after 3 h of

exposure to bacterial suspensions of E. coli, P. aeruginosa, S. aureus and B. subtilis, even after 4 runs with

the same coating, showing that the coating was stable and retained its antibacterial activity. The high

temperature and aggressive way of coating makes it suitable only for metals and other hard materials.

Coating of polymeric materials can require a more delicate approach and is more easily achieved using


21

polymer based coatings. A N-vinylpyrrollidinone and n-butyl methacrylate based hydrophilic surface

coating has been described by Stevens et al., who embedded both Ag-NPs and heparin in the coating,

designed for application on central venous catheters [74]. The embedded Ag-NPs were found to have a

bactericidal effect against several S. aureus strains, even enhanced by the presence of heparin which at

the same time improved the non-thrombogenic behavior of the coating. Another way of coating medically

relevant polymer materials is given by Huda et al. where cooperative electrostatic adsorption was used. In

this system, NPs stabilized with SAMs of ω-functionalized alkane thiols, were given opposite charges using

N,N,N-trimethyl(11-mercaptoundecyl)-ammonium chloride (positive charges) and mercaptoundecanoic

acid (negative charges) and deposited on polypropylene and polyvinylchloride. The adsorbed NP coatings

showed antibacterial effects due to the release of Ag+ ions and were stable for several months [75]. A

release-system based coating has been described by Liu et al. who used poly(lactic-co-glycolic) acid (PLGA)

as a degradable reservoir for Ag-NPs [76]. Stainless steel was dip-coated by immersing it three times in

17.5% (w/v) PLGA in chloroform containing spherical Ag-NPs of 20-40 nm diameter for 30 s and incubating

for 12 h at 37°C. A 2% silver containing PLGA coating not only inhibited growth of S. aureus and P.

aeruginosa in vitro, but using a rat femoral canal model they observed no sign of bacterial survival around

the coated implant after 8 weeks. In addition, at the same time the coated implants significantly improved

the generation of bone.

Next to silver, other metal NPs used for antibacterial surface coatings include Cu or CuO. Akhavan et al.

used a sol–gel procedure to synthesize silica thin films containing copper-based NPs on soda lime glass

substrates [77]. Depending on the temperature of the subsequent heat treatment step, the films

contained either mainly CuO (reduction at 300°C) or mainly Cu (reduction at 600°C) NPs. Bactericidal

effects of the coating were tested against E. coli and it showed that use of the CuO-NPs decreased bacterial

killing in absolute numbers, however, when the antibacterial activity was normalized by its Cu/Si ratio,

heat treatment actually improved the antibacterial activity. They concluded that Cu-NPs were a stronger

antibacterial material compared to CuO-NPs, due to increased photo-inactivation of bacteria. Cu-NPs

coated cellulose films have also been demonstrated to have a bactericidal effect [78]. By dissolving cotton

linter in aqueous cuprammonium and casting it on glass substrates followed by exposure to 10 wt% NaOH

aqueous solution for 10 min, Cu/cellulose coatings were produced. Subsequent placement of the film in

0.3 M NaBH4 aqueous solution at 5°C for 5 h resulted in Cu/cellulose nanocomposite films, containing Cu-

NPs with a mean size of 47.5 nm, which completely killed S. aureus and E. coli bacteria in suspension within

1 h.

Rai et al. have used the antibiotic cefaclor as both a reducing agent for tetrachloroauric(III) acid (HAuCl4)

as well as a capping agent for the resulting gold NPs (22-52 nm diameter) [79]. Coating on glass surfaces

was achieved using PEI, which resulted in extremely stable coatings even at highly acidic (pH 3) and alkaline

(pH 10) conditions. Coatings were effective in completely eradicating S. aureus and E. coli from suspension

within 6 h and binding of cefaclor to gold NPs lowered the minimum inhibition concentration (MIC) from

50 mg mL-1 to 10 mg mL-1, showing the beneficial effect of antibiotic coating to gold NP.

CHAPTER 1

22

Besides metals, silica NPs have been applied as antibacterial surface coatings as well. However, since silica

NPs do not display any known form of antibacterial activity, they require addition of components offering

them such properties. One strategy is to coat silica NPs with a quaternary ammonium cationic surfactant,

as was achieved by Botequim et al., who used didodecyldimethylammonium bromide (DDAB) [80]. In their

work, silica NPs of either 8 or 80 nm in size were coated with DDAB and subsequently coated to glass

coverslips using dopamine hydrochloride as a coupling agent. Coated substrates showed antibacterial

activity by completely preventing the adhesion of living cells of C. albicans, E. coli, and S. aureus, after 6 h

incubation with 1 × 105, 1 × 103 and 1 × 106 cells mL-1, respectively.

This large collection of studies demonstrates the extent of the field and the wide range of methods

available to apply these ultra-small particles to fight bacterial adhesion and biofilm formation. However,

with new methods also come new restrictions. A point of concern for the use of nanoparticles in

antibacterial surface coatings is the quick assembly of a layer of adsorbed proteins on the nanoparticle

surface, called the protein corona, when exposed to bodily fluids [81-83]. The protein corona can partly

obstruct functional molecules on the nanoparticle surface and reduce the overall desired effect, requiring

a higher dose to achieve the same net effect as would be achieved in the absence of these surface

associated proteins.

Whether applied on their own, or in combination with other antimicrobial compounds, some NPs display

excellent antibacterial properties. Initially, worries were expressed towards the toxicity against human

cells and tissue, as for example seen in amino-modified polystyrene nanoparticles [84], but most studies

incorporate these concerns into their experimental set-up and seldom find any negative effects on

proliferation or adhesion of mammalian cells. Nevertheless, it is important to continue taking this aspect

in consideration.

Quaternary ammonium compounds

The general chemical structure of quaternary ammonium compounds (QACs) is represented by R1R2R3R4N+

X− (Figure 4), in which R depicts a hydrogen atom, an alkyl group or an alkyl group with other functional

groups, and X represents an anion [85]. The efficacy of QACs towards killing of bacteria has turned out to

be greatly dependent on whether the positive charge density in a coating exceeds the required threshold

of 1015 N+ cm−2 [86, 87] and on the length of the alkyl chain. Generally, when the alkyl chain length falls

below 4, or above 18, the antimicrobial effects are almost completely diminished [85, 88]. The chain length

dependence of the efficacy towards the antibacterial properties is related to the mechanism by which

QACs inhibit or kill bacterial cells in solution, but it is uncertain whether this mechanism also prevails for

QACs immobilized on a surface. Generally it is assumed that the positively charged quaternary nitrogen of

a QAC molecule is strongly attracted to the negative cell wall of bacteria interacting with negatively

charged phospholipid head groups. Once the QAC molecule becomes associated with the cell wall, the

hydrophobic alkyl tail of the QAC becomes incorporated into the hydrophobic bacterial cell membrane.


23

When the concentration of QACs in the cell membrane becomes high enough, this causes disruption of

the cell membrane with subsequent leakage of the of the bacterial cytosol, resulting in lysis of the cell [85].

FIGURE 4 The general structure of a quaternary ammonium ion. R can represent a hydrogen atom or an alkyl group that can be substituted with other functional groups and X represents the anion.

Since the antimicrobial activity of QACs is mainly expressed by incorporation into the bacterial cell

membrane, QAC surface coatings require a certain degree of freedom for the molecule, similar to

antibiotics and enzymes as previously discussed. Polymer mediated surface tethering is one way to achieve

sufficient flexibility for QACs to retain their antimicrobial properties. Hyperbranched polyurea coatings

have shown to be an effective way of tethering PEI to silicon substrates, after which amino groups of the

PEI coating could be converted into hydrophobic, poly-cationic species by a consecutive two-step

alkylation process [89]. Fabricated coatings were more hydrophobic than the underlying silicon and

showed to have a charge density of 1015 N+ cm−2, above the required threshold positive charge density,

and killed adhering S. epidermidis up to challenge numbers of 1600 CFU cm-2. Importantly, whereas the

majority of papers describing contact-killing of adhering bacteria neglect to demonstrate absence of

leachables that may contribute to bacterial killing, killing by the above described hyperbranched coating

was confirmed to be in the absence of leachables. Due to the hyperbranched nature of the coating, QACs

do not only have more spatial flexibility, but also allow for multiple contact points to develop between an

adhering bacterium and the coating Accordingly, Asri et al. strengthened the current perception that

electrostatic attractions, strong enough to extract anionic lipids from the bacterial cell membrane with

subsequent leakage of the bacterial cytosol, play a major role in the working mechanism for immobilized

QACs in a coating [89,90]. Moreover, this also explains why bacterial strains, not susceptible to QACs in

solution, are contact-killed by immobilized QACs as it provides for an entirely different working mechanism

than operative in solution [91].

Wong et al. described a method of coating antibacterial thin films assembled from layer by layer

application of polycationic N-alkylated PEIs and polyanions on silicon substrates [92]. Layer by layer films

were built up from alternating polycationic PEIs with polyanions, using three different PEI-based

polycations and varying the number of bi-layers in the films. Focusing on their result with linear DMLPEI

as the polycation component and PAA as the polyanion, they found that the bactericidal activity was

CHAPTER 1

24

dependent on the number of bilayers and influenced by the pH of the PAA solution at the time of layer

formation. At low pH the PAA remained relatively uncharged, which resulted in a bilayer with fewer

interaction points between DMLPEI and PAA, thereby creating a bilayer with more loops and a rougher

surface, displaying a higher number of cations available for interaction with bacteria (see Figure 5). These

systems proved to be more bactericidal, only requiring 1.5 bilayers for complete killing of airborne S.

aureus, compared to 14.5 bilayers being required for the same effect when PAA with a pH of 5 was used

to create bilayers. Additionally, the authors observed that deposition of only DMLPEI on negatively charged

Si wafers lacked any antibacterial activity, confirming the thought that tight binding of the positive surface

charges is detrimental for the antibacterial activity of these QACs. Similar results were found in sort-like

multilayer system using different QACs [93].

FIGURE 5 Schematic representation of the different conformations of polymer chains resulting from PAA at different pH values. At pH 3.0, most of the PAA chains (blue) are uncharged, which results in a conformation of the DMLPEI cation (red) with most of its positive charges available, leading to a high bactericidal effect. As the pH increases the PAA chains become more negatively charged, crosslinking with more of the positive charges of DMLPEI leaving less cations available for interaction with the bacterial cell membrane and hence less bactericidal activity. Adapted from [92] and reprinted with permission.

pH 3.0

pH 7.0

pH 5.0


25

An alternative method to ensure availability of the positive charges of QACs to keep their antimicrobial

activity is by covalent immobilization on glass using a short linker molecule. Recently, Iarikov et al.

functionalized glass surfaces with epoxide groups using 3-glycidoxypropyltrimethoxysilane (GOPTS) [94].

Modified glass surfaces were then exposed to poly-allylamine (PA) so that the PA could bind via reaction

of a part of its amine groups. Covalently bound PA showed to have more extended chains compared to

electrostatically adsorbed PA, while chain length also increased with increased GOPTS reaction time and

lower molecular weight. Accordingly, glass surfaces modified with PA resulting in the most extended chains

showed the highest overall killing of S. epidermidis, S.aureus and P. aeruginosa, showing reductions in

adhered bacteria of 97%, 97% and 88%, respectively.

Siedenbiedel et al. developed an antimicrobial coating of a quaternized amphiphilic star block copolymer

with a semi-permanent character [95]. By creating a hydrophilic antimicrobially active outershell, together

with a hydrophobic core, micellar structures assembled in water and lead to differently structured

antimicrobial coatings being developed on the surfaces after drying. The star-shaped structures made

from a polystyrene core and poly(4-vinyl-N-methylpyridinium) outer shell were, when the polymers were

applied in the right proportions, capable of being coated on a surface from water and showed

antimicrobial activity against S. aureus. The antimicrobial activity was maintained even after rinsing, while

deliberately streaming water over the surface for longer periods of time, removed the coating so that the

unmodified surface was recovered, showing the non-permanent nature of their coating.

Although quaternary compounds were first used in solutions as disinfectants, due to their high stability

they are currently mainly studied for use in permanent surface coatings. The high stability contact-killing

mechanism causes bacteria to disintegrate leaving the coating intact and capable of protecting the surface

against more bacteria. The absence of decreased efficacy by shielding of the positive charge of QACs due

to adhered bacterial debris or protein adsorption is shown in in vivo studies, in which QAC coatings

remained effective in preventing infection for multiple days and even induced bone healing [96, 97]. The

advantages of permanent surface binding of QACs make that there are only few studies on the report of

QAC releasing coatings, especially since such a strategy bears the risk of QAC-induced hemolysis [98].

Polymers in passive coatings

PEG is often considered the gold standard for polymer brush surface modification, designed to resist

fouling of the surface by many different substances [99]. By forming an osmotically driven, steric barrier

to which bacteria cannot adhere, PEG modification is an example of a polymer which passively protects

the surface from bacterial adhesion [100]. The low adhesion forces between bacteria and polymer brushes

is believed to cause them to keep their planktonic phenotype, missing the stimulation to develop into a

biofilm [101]. The passive mechanism by which PEG prevents adhesion of bacteria is typical for how

polymer coatings protect surfaces. On their own, most polymers do not possess any antibacterial activity

and hence their only way to stop bacterial colonization is by passive prevention of adhesion. Polymers can,

CHAPTER 1

26

however, be very effective in preventing bacteria to adhere, and this is why there have been many

examples in which polymer surface coatings have been combined with the use of antimicrobials,

combining the non-adhesiveness of the polymer brush with the killing efficacy of an antimicrobial to

improve the overall result. The possible antimicrobials that can be combined with polymer surface coatings

have been mentioned before and include, but are not limited to, QACs, peptides and antibiotics. Surface

coatings consisting of these combinations have been discussed separately in the above sections and

therefore this section will mainly focus on polymer surface coatings that passively prevent bacterial

adhesion.

To keep bacteria from adhering to a polymer coated surface, the attached polymer layer has to be in a

well hydrated state, which is generally achieved by covalent immobilization or physisorption of hydrophilic

polymer chains to the surface. This strategy has been used for many years and the latest developments

are driven towards the novel application of known polymers rather than the design and use of new

chemicals. The application of bio-inspired attachment of polymer chains is recent, and dopamine

molecules which are found to be important in the strong adhesion of marine-mussels, or derivatives of

dopamine, are often applied to this end. Polydopamine, formed by coating a surface with dopamine, can

be directly functionalized with polymer brushes, or can be used to attach an ATRP initiator for a grafting-

from approach [102, 103]. Amine terminated PEO was grafted to a thin layer of polydopamine by Pop-

Georgievski et al. by dip-coating of dopamine coated samples into PEO solution [103]. The resulting brush

coatings were shown to be stable for multiple days, based on their ability to resist protein adsorption.

ATRP formation of brushes showed anti-adhesive properties, but inclusion of a quaternized group was

necessary for the desired antibacterial properties. The requirement for the inclusion of these antimicrobial

groups, which as mentioned previously is often performed, depicts the consensus that for most

applications anti-adhesiveness is not sufficient to prevent infection. This is supported by the fact that when

polymer brushes are subjected to bacteria in growth media, even without firm attachment of initially

adhering bacteria, a biofilm may still form [11].

The hydrated state of polymer brushes is vital to the anti-adhesive capacity of these coatings. To further

hydrate a coating, crosslinks can be formed between the polymer chains on a surface to create a hydrogel

which can hold more water than a brush-structure without collapsing and thus can increase the anti-

adhesive properties of a surface. Wang et al. cross-linked PEG using an electron beam and created micro-

patterned surfaces of PEG hydrogels separated at different distances [50]. When the micron-sized

hydrogel spots where separated at distances of 0.5 μm, bacteria could not adhere between the structures

thus preventing bacterial adhesion. By slightly increasing the space between hydrogels anti-adhesive

properties were still observed, while at the same time tissue cells where able to adhere to the surface due

to their larger size with respect to bacteria. Alternatively, end-functionalization of PEG with dopamine

molecules leads to crosslinking between PEG molecules and results in hydrogel formation. This strategy

was demonstrated to decrease bacterial adhesion by 80%; however, this was not as effective as most other

PEG modifications [104]. Zhao et al. crosslinked poly(N-hydroxyethylacrylamide) and loaded the resulting


27

hydrogel with salicylate. Using this method the authors created hydrogels exhibiting both anti-adhesive

properties as a result of hydrogel formation as well as antibacterial properties, attributed to the release

of salicylate [105]. The combination of anti-adhesive and antibacterial properties resulted in the ability to

withstand bacterial adhesion of S. epidermidis and E. coli for over 24 h.

Polymer attachment to surfaces is effective in reducing bacterial adhesion, but due to the passive nature

cannot prevent biofilm formation over longer periods of time. However, in many temporary applications

the non-adhesive nature of polymer surface modifications may be adequate to prolong the lifespan of a

device or implant, e.g. of urinary or intra-vascular catheters. The weak adhesion forces of bacteria on

polymer brushes and the occurrence of fluid induced shear forces in these situations provide physical

removal of bacteria not found under static fluid conditions.

Super-hydrophobicity

Hydrophobic interactions play a role in the adhesion of bacteria to surfaces, by favoring the attraction of

two hydrophobic components to remove interfacial water and lower the free energy of a particular system

[106, 107]. However, extremely hydrophobic surfaces have been shown to possess extraordinary anti-

adhesive properties [108].

The Aizenberg group recently published a paper in which they added lubricating fluids, consisting of

perfluorinated liquids, to porous polytetrafluorethylene (PTFE) to fabricate liquid-infused surfaces [109].

These surfaces were shown to be extremely resistant to bacterial adhesion and biofilm formation. Biofilm

attachment of P. aeruginosa after 7 days was effectively zero, showing excellent anti-adhesive properties

and stability of the coating. In another study from the same group, it was shown that a nanostructured

surface based on an epoxy-resin could be infused with the same perfluorinated liquids to obtain a similar

anti-adhesiveness and even demonstrating self-repairing behavior after physical damage [110]. Li and co-

workers have also used a liquid infusion technique to create slippery, bacterial adhesion resistant surfaces

[111]. By preparing a porous polymer surface of a mixture of butyl methacrylate and

ethylenedimethacrylate on glass and the subsequent addition of perfluoropolyether, slippery surfaces

were made that resisted biofilm formation of most P. aeruginosa strains included in their study. One of

the used multi-resistant strains however, was still able to form a biofilm, suggesting that the results were

strain dependent.

Privett et al. prepared super-hydrophobic surfaces by depositing fluorinated silica colloids onto glass slides

[112]. Briefly, (heptadecafluoro-1,1,2,2-tetrahydrodecyl)trimethoxysilane and tetraethylorthosilicate

were sonicated and added to a solution of ethanol and ammonium-hydroxide to form silica colloids, which

were spread-cast onto ozone/ultraviolet (UV)-treated glass slides. Static water contact angles on the

coated surfaces exceeded 150 degrees and showed an over 1.75 log reduction in adhesion of S. aureus

and P. aeruginosa. Water contact angles did not change after immersing the coating in water for over 15

CHAPTER 1

28

days, showing good stability of the coating, although no bacterial adhesion experiments were performed

after storage.

Whereas the previously mentioned studies all require fluorinated substances to achieve super-

hydrophobicity, Hu et al. described an electro-spraying method to apply a super-hydrophobic

biodegradable coating, without the use of such liquids [113]. In their work they present how co-electro-

spraying of poly(L-lactide) and modified silica NPs onto titanium plates resulted in a coating with a water

contact angle of 157°. Bacterial adhesion was reduced by 75% compared to poly(L-lactide) films, however,

no numbers for bare titanium plates were reported. Although, like other super-hydrophobic coatings, the

adhesion of mammalian cells was reduced as well, the biodegradable nature or the coating could still allow

for tissue integration of an implant after degradation of the coating, thereby showing a novel feature to

make super-hydrophobic surfaces more applicable to medical implants.

The main disadvantage of super-hydrophobic coatings is that they not only restrict the adhesion of

bacteria but of mammalian cells as well, which means they cannot be used in applications requiring tissue

ingrowth, although the previously mentioned study by Hu et al. showed that tissue integrating variants

could be made as well, by making the coating biodegradable [113]. However, even without allowing

attachment of mammalian cells, there are many possible applications for which super-hydrophobic

surfaces are suitable and could reduce the infection rate. Especially in the presence of flowing liquids, e.g.

in catheter, the non-adhesiveness would promote clearance of unwanted contaminants.

Chitosan

The use of naturally derived components is an important current theme in surface coating of materials

[114–116]. Examples of such materials include, hyaluronic acid, alginate, collagen, chitosan and dextran.

Chitosan however, is the only one among these materials possessing an inherent antibacterial activity,

albeit that this antibacterial activity depends on the degree of chitosan acetylation [117, 118]. Yang et al.

used a biomimetic anchor and chitosan functionalized polymer brushes to prevent bacterial adhesion on

stainless steel surfaces [117]. Barnacle cement, harvested from live barnacles, was used to attach an ATRP

initiator for formation of surface initiated PHEMA polymer brushes. Subsequently, the hydroxyl groups of

the PHEMA brushes were converted into carboxyl groups that were allowed to react with the amine groups

of chitosan, to achieve chitosan functionalized polymer brush surfaces. The viability of E. coli that managed

to adhere on the chitosan modified polymer brush coated stainless steel was 80% lower compared to bare

stainless steel, due to the bactericidal effect of chitosan. Surface composition of the coated surfaces after

30 days showed a less than 10% loss of the barnacle cement, indicating good stability, however, the

authors did not test for bacterial adhesion after this time period. Combining the anti-adhesive nature of

polymers with the antibacterial properties of chitosan was achieved by Wang et al. by using a multilayer

system in which the base layer consisted of a heparin/chitosan film held together by electrostatic

interactions [119]. On top of the heparin/chitosan base layer a (polyvinylpyrrolidone/poly(acrylic acid))


29

(PVP/PAA) layer was added by alternate deposition of PVP and PAA, after which the top layer was then

cross-linked using heat treatment. The final coating was initially anti-adhesive, but demonstrated contact

killing of S. aureus after the anti-adhesive top-layer had degraded after 24 h in phosphate buffered saline,

exposing the bactericidal heparin/chitosan base layer. Another study demonstrated that a

heparin/chitosan multilayer had antibacterial functionality and at the same time served as an osteo-

inductive coating, offering a good prospective to improve the outcome of bone allograft procedures [120].

Chitosan could also be incorporated into hydroxyapatite, resulting in good antibacterial properties of the

coating against S. aureus, while at the same time the porous character of the hydroxyapatite enhanced

osteoblast cell response, as long as chitosan concentrations remained below cytotoxic values [121].

Although chitosan already possesses antibacterial properties by itself, many studies have been performed

on how to improve the bactericidal effect by the use of additional antibacterial compounds. Since QACs

have strong bactericidal capabilities, chitosan has been quaternized in several studies to increase the

antibacterial properties [122–124]. Lee and co-workers studied quaternary ammonium modified chitosan

brush layers [125]. Modification of chitosan by performing a Michael reaction with an acryl reagent in

water, proved to be effective in introducing quaternary ammonium groups. To evaluate the effect of QAC

modification, 25% and 50% QAC substitution was tested while chitosan only brushes were used as control.

To immobilize the resulting chitosan and chitosan-QAC complex (CH-Q), silicon oxide surfaces were treated

with GOPTS and an aqueous solution of CH (or CH-Q) was added and allowed to evaporate slowly. After

evaporation, the film was left at 60°C for 12 h and the resulting films expressed a pH dependent swelling

behavior which allowed fine tuning of the film thickness. Brush layers decreased in thickness with

increasing quaternization, whereas the antimicrobial activity increased. The 50% CH-Q coating showed

antibacterial activity against S. aureus, decreasing the amount of CFUs after 6 h exposure by 97%

compared to uncoated control surfaces. Later studies confirmed these results and showed that CH-Q

coatings effectively prevented bacterial adhesion in flow conditions as well [126]. Ding et al. showed that

addition of alkynyl groups to chitosan led to increased antibacterial activity of hydrogel coatings against S.

aureus and E. coli [127].

Even though chitosan possesses a limited bactericidal effect, its biocompatibility, along with the ease by

which it can be modified, makes it a popular building block for many antibacterial surface coatings. Little

is known about the possible development of bacterial resistance against chitosan and it remains to be seen

whether or not this occurs upon its increasing use.

CONCLUSIONS

Antimicrobial coatings are of ubiquitous importance, but requirements set to such coatings are most

stringent in biomedical applications, constituting the focus of this review. Prevention of bacterial adhesion

and killing of bacteria, either by coatings that release antibacterial substances or through surface-

associated mechanisms, are the most prevalent approaches. A trend towards developing multi-functional

CHAPTER 1

30

coatings is becoming more apparent. Approaches based on the release of substances bear the risk of a

depleted coating when needed most. Surface-associated mechanisms may suffer from attenuated efficacy

due to coverage by proteins adsorbing from body fluids, but hitherto QACs coatings have been

demonstrated to remain antimicrobially active in animal studies. From a general perspective it is

impossible to tell which coating strategy will yield the best options, since this all depends on the clinical

application aimed for and whether expectations are short- or long term. However, taking into

consideration that the era of antibiotics to control infectious biofilms will eventually come to an end, it

becomes evident that the future for biofilm control on biomaterial implants and devices is with surface-

associated modification of surfaces that are non-antibiotic related.


31

AIM OF THE THESIS

The general aim of this thesis is twofold:

The first aim of this thesis is to develop antibacterial coatings preventing bacterial adhesion and biofilm

formation by making it difficult for bacteria to adhere, while at the same time allowing for rapid tissue

integration, as this constitutes the best protection against bacterial contamination. To this end, we

developed micro-patterned surfaces of PEG-hydrogels, with anti-adhesive properties towards bacteria,

while offering mammalian cells, which are larger in size, enough possibilities to firmly adhere. Additionally,

using a completely different strategy, we studied the possibility of incorporating DNase I into surface

coatings to attack extracellular DNA, as an important component of bacterial extracellular polymeric

substances in biofilms.

The second aim is to increase our knowledge of bacterial adhesion mechanisms based on lateral force

microscopy, rather than using more common normal force microscopy. Lateral forces arise when adhering

bacteria are forced to move over a surface, and can have different origins depending on the type of

substratum surface involved. In this thesis we studied lateral adhesion forces on a synthetic polymer-brush

coating, which resists bacterial adhesion, and a salivary coating, able to interact with adhering bacteria

through specific receptor-ligand bonds.

CHAPTER 1

32

LIST OF ABBREVIATIONS

AMP = Antimicrobial peptide

ATRP = Atom transfer radical polymerization

CaP = Calcium phosphate

CH-Q = Chitosan-QAC complex

DDAB = Didodecyldimethylammonium bromide

DMLPEI= Dodecyl,methyl-poly(ethyleneimine)

DOPA = Dopamine

DspB = Dispersin B

GOPTS = 3-Glycidoxypropyltrimethoxysilane

GS = Gentamicin sulfate

HAP = Hydroxyapatite

LDH = Layer double hydroxide

MUA = Mercaptoundecenoic acid

NP = Nanoparticle

PAA = Poly(acrylic acid)

PAH = Poly(allylamine hydrochloride)

PEG = Poly(ethylene glycol)

PEI = Poly(ethyleneimine)

PEO = poly(ethylene oxide)

pHEMA = Poly(2-hydroxyethyl methacrylate)

PLGA = Poly(lactic-co-glycolic) acid

PMAA = Poly(methacrylic acid)

PMMA = Polymethymethacrylate

PPO = Poly(propylene oxide)

PVP = Polyvinylpyrrolidone

QAC = Quaternary ammonium compound

SAM = Self-assembled monolayer


33

CONFLICT OF INTEREST

This study was funded by the UMCG, Groningen, The Netherlands. H.J. Busscher is also director of a

consulting company, SASA BV (GN Schutterlaan 4, 9797 PC Thesinge, The Netherlands). The authors

declare no potential conflicts of interest with respect to authorship and/or publication of this article.

Opinions and assertions contained herein are those of the authors and are not construed as necessarily

representing views of the funding organization or their respective employers.

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CHAPTER 2

LENGTH-SCALE MEDIATED DIFFERENTIAL ADHESION

OF MAMMALIAN CELLS AND MICROBES

Yi Wang, Guruprakash Subbiahdoss, Jan J.T.M. Swartjes, Henny C. van der Mei,

Henk J. Busscher, and Matthew Libera,

Adv. Funct. Mater. 2011,.21;3916–23

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42

ABSTRACT

Surfaces of implantable biomedical devices are increasingly engineered to promote their interactions with

tissue. However, surfaces that stimulate desirable mammalian cell adhesion, spreading, and proliferation

also enable microbial colonization. The biomaterials-associated infection that can result is now a critical

clinical problem. We have identified an important mechanism to create a surface that can simultaneously

promote healing while reducing the probability of infection. We created surfaces with submicron-sized,

non-adhesive microgels patterned on an otherwise cell-adhesive surface. Quantitative force

measurements between a staphylococcus and a patterned surface show that the adhesion strength

decreases significantly at inter-gel spacings comparable to bacterial dimensions. Time-resolved flow-

chamber measurements show that the microbial deposition rate dramatically decreases at these same

spacings. Importantly, the adhesion and spreading of osteoblast-like cells is preserved despite the sub-

cellular non-adhesive surface features. Since such length-scale-mediated differential interactions do not

rely on antibiotics, this mechanism can be particularly significant in mitigating biomaterials-associated

infection by antibiotic-resistant bacteria such as MRSA.


43

INTRODUCTION

Restoration of human function using implantable biomedical devices and prostheses is indispensable to

modern medicine, and the surfaces of modern biomaterials are now highly engineered to regulate their

interactions with physiological systems. However, many of the same surface properties that influence

mammalian cell interactions also enable bacterial adhesion (Figure 1A). The subsequent biomaterials-

associated-infection that can occur is now recognized as a major clinical problem. When bacteria win the

race with mammalian cells to colonize an implant surface [1-4], they can develop into biofilms where they

are both extremely resistant to antibiotics and able to evade the host immune system [5-9]. While

antibiotics can mitigate the short-term symptoms of systemic infection, they are usually unable to resolve

the localized biomaterials-associated infection. In such cases, the implant is removed, the infection is

resolved over periods of week to months, and a revision surgery, with a higher probability of re-infection,

is performed in a site with less native tissue [10-12]. Biomaterials-associated infection is a concern with all

biomedical devices that contact tissue and is usually assumed as a given in cases involving, for example,

long-term percutaneous structures or serious trauma involving large and contaminated wounds. The

general problem is being increasingly exacerbated by the growing preponderance of antibiotic-resistant

bacteria and the concomitant decline of new antibiotic development [13, 14].

Surface modification to mitigate bacterial colonization has largely followed one of two routes. The first

incorporates antimicrobials by various drug-delivery mechanisms [15-17]. Such an approach, however,

must determine a priori the appropriate antimicrobial and often delivers it when not needed, thus

promoting antibiotic resistance. Consequently, there is a growing focus on alternatives involving, for

example, metal ions, cationic peptides, and quorum-sensing targets. A second route concentrates on the

modification of the surface itself, much of which has focused on antifouling coatings. Among these,

surfaces that incorporate poly(ethylene glycol) (PEG) have been extensively studied because of their ability

to resist both protein adsorption and cell adhesion [18, 19]. These have served as a model for the

development of other highly hydrophilic non-adhesive and multi-functional surfaces [20-23]. Such surfaces

protect against microbial colonization, but they also resist mammalian cell adhesion (Figure 1B). Thus,

while they can mitigate biomaterials-associated infection, they simultaneously compromise healing.

A fundamental problem in biomaterials science centers is how to create surfaces that differentially interact

with different cell types. In the case of biomaterials-associated infection, this problem sharpens to the

question of how to create a surface that differentially promotes interactions with desirable mammalian

cells while simultaneously inhibiting microbial colonization [24, 25]. Significantly, there are important

physico-chemical differences between mammalian cells and bacteria around which differentially

interactive surfaces can be designed. For example, mammalian cells, such as osteoblasts, are typically 10-

100 µm in diameter, have flexible cell membranes that can conform to a substratum, and adhere to

surfaces through multiple, integrin-mediated submicron-sized focal contacts [26]. In contrast, microbes

have well-defined shapes and are a few micrometers or less in size. In particular, the staphylococci most

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44

often implicated in biomaterials-associated infection – Staphylococcus epidermidis and Staphylococcus

aureus - are spherically shaped with diameters of about 1 µm and have relatively rigid cell walls. Combining

these differences with emerging concepts of surface patterning and compartmentalization [27, 28], we

hypothesize that a surface whose cell adhesiveness is laterally modulated at microscopic length scales will

enable mammalian cell adhesion while reducing microbial adhesion (Figure 1C). Using a combination of

electron-beam surface patterning, quantitative staphylococcal-surface adhesive force measurement, and

in situ characterization of microbe/osteoblast surface colonization, we show this hypothesis to be true:

laterally modulated adhesiveness significantly reduces microbial colonization, in the absence of antibiotics,

when the spacing between non-adhesive features is comparable to microbial dimensions (1-2 µm), while

osteoblast-like cells can nevertheless adhere to and spread over these surfaces. Such a differentially

adhesive surface is one that can promote healing while simultaneously reducing the risk of infection.

FIGURE 1 (A) Both mammalian cells and microbes can adhere to a fully cell-adhesive biomaterial surface; (B) Neither mammalian cells nor microbes can adhere to a fully non-adhesive biomaterial coating; (C) A surface patterned with submicron-sized, non-adhesive features on an otherwise cell-adhesive surface can enable mammalian-cell adhesion but reduce microbial adhesion due to their smaller size.


45

RESULTS AND DISCUSSION

Surface-patterned PEG gels

To test the concept of length-scale-mediated differential adhesion, we created submicron-sized PEG

hydrogels patterned in square arrays on an otherwise cell-adhesive substratum. We assessed the response

of various microbes and an osteoblast-like cell line to these as a function of inter-gel spacing, δ. Focused

electron beams (e-beams) can locally crosslink PEG films to form surface-bound microgels [29-31]. We

have shown that such gels resist nonspecific protein adhesion [30, 31] as well as the adhesion of S.

epidermidis [32], astrocytes, and neurons [33]. We have also previously shown that the exposed surface

between microgels remains adhesive to protein adsorption [31, 33]. Because of the flexibility afforded by

e-beam patterning, the spatial distribution of these discrete microgels can be easily varied. Figure 2A

shows atomic force microscopy (AFM) images of surface-patterned PEG microgels in the dry and hydrated

states for three intergel spacings (δ = 0.5, 1.5, and 3.0 µm). When hydrated, each microgel is about 400

nm in diameter and about 120 nm in height. They swell from the dry state by a factor of about 2, primarily

in the direction perpendicular to the substratum. Importantly, the irradiation conditions used to create

each microgel are identical, so their size and swelling behaviour are very reproducible.

FIGURE 2 Surface-patterned PEG microgels modulate the adhesion force experienced by an individual S. aureus bacterium. AFM images (upper) of patterned surfaces in the dry and hydrated states for three inter-gel spacings (δ). These images were collected in non-contact mode using a Si3N4 tip. The surfaces were hydrated in PBS buffer. The lower plots show examples of the spatially resolved adhesion force between a S. aureus bacterial probe and the three patterned surfaces. The adhesion force is minimal for δ = 0.5 µm and develops a substantial spatial dependence as δ is increased to dimensions comparable to or larger than the size of an individual bacterium.

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46

By immobilizing bacteria on an AFM cantilever in place of a typical Si or Si3N4 AFM tip, an atomic force

microscope was used to measure the adhesion force that an S. aureus bacterium feels as it interacts with

a silanized glass surface modulated by PEG microgels. The probe was made by adhering S. aureus to a poly-

L-lysine-treated AFM cantilever. In contrast to a typical AFM tip, this bacterial tip is substantially larger and

provides a biochemical interaction of bacterial origin. Bacterial probes were rastered across patterned

surfaces, and the adhesion force was determined at each point in the raster. Figure 2 (lower plots)

illustrates the adhesion force as a function of lateral distance for surfaces with δ = 0.5, 1.5, and 3.0 µm.

The force is negligible between the bacterium and the PEG microgels, consistent with the non-adhesive

characteristics of highly hydrated PEG. The magnitude of the adhesive force increases between the gels.

The maximum force is presented in Table 1 as a function of inter-gel spacing. It is lowest for δ = 0.5 µm

where the individual gels almost overlap. In this case, the bacterium is shielded from the glass surface by

the intervening PEG. As δ increases, the maximum adhesion force increases. For δ between 1.0 and 1.5

µm, the adhesion force increases substantially. It further increases as δ increases to 3.0 µm. This finding

confirms that: (i) an individual bacterium experiences an adhesive interaction with the patterned

substrates despite the non-adhesive features; and (ii) the magnitude of the adhesive interaction increases

as the spacing between the non-adhesive features increases.

TABLE 1 Maximum adhesion force of S. aureus on patterned glass surfaces with different inter-gel spacing (δ). Each data point represents the average ± the standard deviation from at least nine values of maximum and minimum adhesive force

Microbial adhesion

We used a parallel-plate flow chamber to measure the initial deposition rate of various microbes. Glass

slides were prepared with square arrays of patterned gels, each array having a fixed δ. Triplicate arrays

with δ ranging from 0.5 µm to 8 µm were patterned on the same substratum. Thus, the interactions of a

given microbe with a range of patterns could be evaluated in a single flow-chamber experiment. Inocula

with fixed microbial concentrations were passed over the surface for 3 h, and we measured the number

of adhering bacteria on each patterned array as a function of time. Nutrient-free media were used to

isolate microbial adhesion from growth.

Inter-gel spacing (µm) 0.5 1.0 1.5 2.0 2.5 3.0

Adhesion Force (nN) -0.04±.04 -.3±0.0 -2.2± 0.2 -2.8± 0.3 -.5±0.8 -3.6±0.3


47

FIGURE 3 The microbial deposition rate decreases as δ approaches microbial dimensions. (A) Representative optical micrographs of S. aureus adhered to various surfaces after 180 min deposition time from an inoculum of 3x106 bacteria/mL at a shear rate of 11/s. (B) The number of adhering S. aureus increases linearly with time for a given δ. The solid line is a linear least squares fit to one data set. (C) A Weibull model (lines) of normalized microbial deposition rate shows that surfaces with cell adhesiveness laterally modulated over microscale distances resist microbial adhesion. Each data point represents the average, and the error bars represent the standard deviation, of slopes of linear least squares fits to data from three independent bacterial culture experiments.

Typical image data are presented in Figure 3A for S. aureus adhesion after 180 min. Note that the bacteria

appear primarily as individuals rather than as multi-bacteria clusters. Figure 3B plots the number of

adherent bacteria per unit area over 180 min. For all inter-gel spacings, the number of adhering S. aureus

increases linearly with time, indicating that the surface fraction covered by bacteria remains small during

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48

the experiment. Interactions between incoming and adhering bacteria can thus be neglected. We

determined the initial deposition rate from the slope of straight lines fit to such data (Figure 3B). The initial

deposition rate on the unpatterned glass surface, , was always the highest. The deposition rate on

a surface with a particular inter-gel spacing, , decreased as δ decreased. For a given microbial species,

we defined a normalized initial deposition rate as:

(1)

Normalized rates are presented in Fig. 3C. We used a cumulative Weibull function to model these rates as

a function of inter-gel spacing according to:

(2)

The solid lines (Figure 3C) represent Weibull fits, and Table II reports values of α and β for the three data

sets. The Weibull function has historically been used to model statistical phenomena associated with

random events, most notably for instances of failure in engineered systems, and it provides two

parameters, α and β, which give further insight into the microbial adhesion process. The scale parameter,

α, represents the δ value at which the initial deposition rate is 63.2% of that for unpatterned glass. The

different α values indicate that S. aureus and C. albicans are affected more strongly by the gels than S.

epidermidis. The shape factor, β, exceeds 1 for all three microbes, consistent with our finding that the

deposition rates increase with increasing δ. The fact that β is close to 2 is consistent with the idea that the

adhesion rate is related to the available area of adhesive surface, since from eq. [2] it follows that

jo,δ ≈(δ/α)β jo,glass. Deviations from β =2 again indicate species-specific differences in the nature of the

microbe-surface adhesive interaction.

TABLE 2 The initial deposition rates on unpatterned glass of the different microbial strains, together with Weibull scale (α) and shape (β) factors describing initial deposition rates on patterned surfaces as a function of inter-gel spacing.

jo, glass

jo,

glassoj

jF

,

,)(

exp1)(F

Microbial strain jo, glass (cm-2 sec-1) α (µm) β

S. epidermidis 15 ± 3 1.2 ± 0.2 1.3 ± 0.06

S. aureus 46 ± 20 2.6 ± 0.2 1.7 ± 0.02

C. albicans 76 ± 35 2.4 ± 0.2 2.2 ± 0.1


49

Mammalian cell adhesion and spreading

We studied surface interactions with an osteoblast-like cell line (U-2 OS) in the same flow-chamber system

in which the microbial studies were performed. U-2 OS is an immortalized human cell line derived from

osteosarcoma cells [34] and was chosen from a broad selection of human osteoblastic cell lines used

previously [4, 35]. Though there are well-known differences between cancerous and primary osteoblastic

cells, Clover et al. [36] have demonstrated that osteosarcoma cell lines exhibit meaningful osteoblastic

phenotypes. For our experiments, these mammalian cells were suspended in culture medium to enable

both adhesion and spreading. After a deposition period of 1.5 h to allow for static U-2 OS binding, cell-

free medium was passed through the chamber. Our analysis concentrated on 1.5 h and 48 h to assess the

initial adhesion and cell spreading, respectively. Figure 4 shows typical image data demonstrating the very

different U-2 OS behaviour when interacting with surfaces patterned with microgels at different inter-gel

spacings. The cells clearly adhere to the surrounding unpatterned surface. They are completely absent

from surface with δ = 0.5 µm. SEM imaging (Figure 5A) confirms this behavior and further illustrates that

U-2 OS cells spread nicely on adjacent unpatterned surface.

FIGURE 4 Osteoblast-like U-2 OS cells are able to adhere and spread on microgel-patterned surfaces with an inter-gel spacing of 1.0 µm or more. Top: phase-contrast microscope images collected in situ during cell culture within a flow chamber. Bottom: fluorescent microscope images showing the nuclei (blue) and cytoskeletal actin (red).

A significant number of cells can be seen on the patterned δ = 1 µm surface (Figure 4) for both 1.5 and 48

h. SEM imaging (Figure 5B) shows that the cells are able to grow and spread over the non-adhesive

microgels.

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50

FIGURE 5 Scanning electron micrographs showing that osteoblast-like U-2 OS cells are able

to (A) adhere and spread on unpatterned glass but not on microgel-patterned surfaces

with an inter-gel spacing of δ= 0.5 µm located in the right bottom corner of this

micrograph; and (B) adhere and spread on microgel-patterned surface with an inter-gel

spacing of δ= 1.0 µm despite the presence of non-adhesive PEG microgels at sub-cellular

length scales. Note that lamellapodia extend over and between non-adhesive microgels to

reach the adhesive glass surface.

Time-resolved measurements further capture the dynamics of mammalian-cell interactions with the

various surfaces. Fig. 6A describes the number of adhering U-2 OS cells as a function of δ after 1.5 and 48

h. At 1.5 h, we calculated the number of cells and total coverage of cells from the phase contrast images.

Whereas at 48 h, the number of cells and surface coverage of adhering U-2 OS cells were obtained from

fluorescent stained CLSM images. The number of cells decreases with time for δ = 0.5 µm, indicating that

the cells have a high motility on this surface and migrate to the adjacent unpatterned surface. In contrast,

the number of cells on the δ = 1.0 µm surface remains high after 48 h. The effect is even more pronounced

for surfaces with δ = 1.5 µm and above, indicating that both the fraction of adhesive area and its spatial

distribution are sufficient to promote stable cell adhesion. Note that the number of cells per unit area on

unpatterned and patterned surfaces with δ ≥ 1 µm do not change significantly over 48 h indicating that

only cell spreading and no proliferation takes place during this time period. Data on the area fraction of

the patterned square arrays covered after 1.5 h and 48 h (Figure 6B) provide insight into the spreading

behaviour. The fraction for δ = 0.5 µm decreases with time, because the cells migrate away from this

surface. For larger δ, however, the fraction covered increases significantly with time despite the fact that

the number of cells does not change (Figure 6B). This finding indicates that the cells spread on these

surfaces, and, for δ = 1.5 µm or more, do so in a manner similar to that on unpatterned surface.

Our findings that osteoblast-like cells are able to adhere to a surface with modulated adhesiveness are

consistent with a number of studies showing similar results involving different mammalian cells and

different patterning methods. For example, Chen et al. [37] used soft lithography to pattern circular

fibronectin patches 5 µm in diameter and separated from each other by PEGylated thiols at inter-patch

distances of 10 µm. Fibroblasts could adhere to these modulated surfaces and cellular vinculin was co-

localized with the patterned fibronectin, indicating that focal adhesions could form on the adhesive

patches and the cells could bridge the intervening non-adhesive regions. More recently, Spatz et al. [38,

39] have studied the dynamics of integrin-mediated mammalian-cell binding and focal-contact formation

on surfaces with nanoscale modulations of adhesiveness provided by RGD-functionalized gold

nanoparticles in clusters of controllable size and spacing on an otherwise non-adhesive surface. Despite

the fact that these experimental configurations are the inverse of ours i.e. adhesive patches on a largely

non-adhesive background rather than non-adhesive patches on an adhesive background, this and other


51

work [40-43] again clearly demonstrates that mammalian cells are able to efficiently bind to 2-D surfaces

with sub-cellular modulations of adhesiveness.

FIGURE 6 Osteoblast-like U-2 OS cells adhere to and spread on surfaces with δ ≥ 1 µm: (A) the number of adhering U-2 OS cells 1.5 h (white) and 48 h (grey) after seeding; and (B) the percent surface coverage by adhering U-2 OS cells immediately after seeding at 1.5 h (white) and after 48 h (grey) of growth on gel-patterned glass. *Indicates a significant difference (p<0.05) from 1.5 h.

Length-scale mediated differential adhesion

We have studied both mammalian cell and microbe response to surfaces intermediate between the two

extremes of fully adhesive and fully repulsive. Our experimental platform provides for a controllable area

fraction and spatial distribution of non-adhesive microgels on an otherwise cell-adhesive surface. We

attribute the differential response exhibited by these mammalian cells and microbes to differences in their

size, their adhesion mechanism(s), and the relative mechanical rigidity/fluidity of bacterial cell walls and

mammalian cell membranes.

Flow-chamber measurements indicate that microbial adhesion is very strongly reduced when the spacing

between microgels is approximately 1.5 µm or less. AFM force measurements show that this spacing is a

critical one, below which the adhesion force between the surface and an individual bacterium substantially

decreases. Staphylococci are approximately 1 µm in diameter and spherical, and they have highly

crosslinked and relatively rigid cell walls. The fact that staphylococcal adhesion decreases substantially

when they sense non-adhesive features at spacings comparable to their own dimension is consistent with

the idea that reconfiguring their shape to better access exposed patches of cell-adhesive surface is

energetically unfavorable. Interestingly, Candida albicans exhibits similar adhesive behavior despite the

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52

fact that it is physically larger than S. epidermidis and S. aureus and could be perhaps expected to show a

decreased adhesion rate at correspondingly larger microgel spacings. We attribute this behavior to the

fact that the surface of C. albicans is decorated with cell surface structures [44] that afford a degree of

conformational flexibility and an adhesion mechanism unavailable to staphylococci.

Significantly, we find that osteoblast-like cells can adhere to and spread on surfaces with microgels

patterned at inter-gel spacings of 1 µm or more, including the range where staphylococcal adhesion is

substantially reduced. In contrast to the highly crosslinked cell walls of Gram-positive bacteria, the

membranes of mammalian cells are highly fluid. Furthermore, such cells adhere to surfaces via clusters of

transmembrane integrins that ultimately develop into focal contacts and focal adhesions whose

dimensions can be in the range of tens to hundreds of nanometers. Consistent with published reports

establishing mammalian cell adhesion and spreading on surfaces with modulated adhesiveness, we

speculate that the osteoblast-like cells studied are able to adhere to the adhesive patches in between non-

adhesive microgels by sub-micron focal contacts/adhesions. The energetic price to extend cell membranes

over the nonadhesive microgels is apparently insufficient to prevent adhesion except in the case of

microgels spaced at 0.5 µm intervals, which corresponds to a pseudo-continuous PEG film.

The finding of length-scale-mediated differential cell adhesion is important, because it is a non-specific

mechanism by which a surface can differentially interact with different types of cells. By reducing the

probability of microbial adhesion, such a surface can both minimize biofilm formation and force bacteria

at a wound site to reside planktonically where they can be effectively cleared by the innate immune

system. In the particular case of osteoblast-like mammalian cells and staphylococci, this finding has

important technological implications for a broad range of orthopedic implant materials, which must

promote tissue integration necessary for healing while simultaneously reducing the bacterial colonization

that leads to infection.

MATERIALS AND METHODS

PEG hydrogel surface patterning

Patterned glass slides were prepared using established procedures [31, 45]. Slides were sonicated in

ethanol for 5 min and dried with N2 gas. Piranha etch (3:1 98% sulphuric acid and 30% H2O2) was used for

a second cleaning. After a water rinse and N2 dry, the slides were exposed to an O2 plasma (300 mTorr,

1.75 W) for 10 min and silanized with 2% [v/v] vinyl-methoxy siloxane (Gelest) in ethanol for 10 min, rinsed

with ethanol and baked at 110°C (2 h). A solution of 2 wt% PEG (6 kDa; Fluka) in tetrahydrofuran was used

to make thin films by spin casting on these slides. The film thickness after solvent evaporation was ~60

nm.


53

E-beam patterning used an FEI Helios SEM controlled by a Nanometer Pattern Generation System (Nabity).

A typical point dose (dose per single microgel) was 10 fC (2 keV) to locally crosslink the PEG. The

crosslinking of PEG under such e-beam irradiation has been attributed to a free-radical polymerization

mechanism after ionization of C-H bonds [27, 45]. After exposure, the slides were washed in de-ionized

water (30 min) to remove unexposed PEG. The resulting surface consisted of surface-bound microgels

separated from each other by silanized glass. Patterns were made as 200 μm × 200 μm microgel arrays.

Within a given array, δ was fixed. Three identical copies with a particular δ were created on each

substratum. Arrays sampling inter-gel spacings from 0.5 - 8 µm were patterned on the same substratum.

Each array was separated from an adjacent array by 100 µm. After patterning, substrata were stored under

vacuum (50 mTorr). Prior to adhesion experiments, the substrates were placed in phosphate buffered

saline (PBS) for at least 30 min.

Experiments with bacteria and yeast

Bacterial inoculum

Two different staphylococcal strains were used: S. aureus ATCC 12600 and S. epidermidis ATCC 35983. For

each flow-chamber experiment a colony from an agar plate was inoculated into 10 mL of tryptone soya

broth (TSB), cultured for 24 h at 37 oC, and used to inoculate a second culture grown for 17 h in 200 mL

TSB. Bacteria were harvested by centrifugation (6500 g) for 5 min at 10 oC and washed twice in sterile

phosphate-buffered saline (PBS; 10 mM potassium phosphate and 0.15 M NaCl, pH 7). Bacterial

aggregates were broken by mild sonication on ice for 3 × 10 s at 30 W (Wibra Cell model 375, Sonics and

Materials Inc., Danbury, Connecticut, USA) and then resuspended to a concentration of 3 × 106

bacteria/mL in sterile PBS.

Yeast cell inoculum

Candida albicans GB1/2 yeast cells were used. A colony from a brain heart infusion (BHI) agar plate was

inoculated in 10 mL BHI, cultured for 24 h at 37 oC, and used to inoculate a second culture grown for 17 h

in 200 mL BHI. Yeast cells were harvested by centrifugation (5000 g) for 5 min at 10 oC and washed twice

in sterile PBS. The harvested cells were resuspended in 10 mL sterile PBS and diluted to 3 x 106 cells/mL in

PBS.

Time-resolved microbial adhesion

Microbial adhesion was studied by incorporating patterned slides into a parallel-plate flow chamber [4].

Microbial deposition was monitored by digital phase-contrast microscopy (Olympus BH-2; 40x). After

removing air bubbles by flowing PBS, bacteria or yeast cells in PBS were perfused through the chamber

(11 s-1 shear rate) for 3 h at room temperature. Images were taken from each patterned array, and from

the unpatterned silanized glass (control), at 1 min intervals. From these, the number of adhering bacteria

or yeast on each array was determined. Since yeast tended to sediment, the yeast inoculum reservoir was

gently stirred.

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54

Mammalian cell experiments

Mammalian cell culturing and harvesting

U-2 OS osteosarcoma cells were cultured in low-glucose Dulbecco’s Modified Eagles Medium (DMEM)

supplemented with 10% fetal calf serum (FBS) and 0.2 mM of ascorbic acid-2-phosphate. U-2 OS cells were

maintained in T75 culture flasks at 37 oC in humidified 5% CO2 and harvested at 90% confluency using

trypsine/ethylenediamine–tetraacetic acid. The harvested cells were diluted to 6 x 105 cells/mL.

Time-resolved mammalian cell adhesion

U-2 OS adhesion and spreading on patterned substrata were studied by in situ imaging in the parallel-plate

flow chamber and by ex situ immunofluorescence/SEM imaging. The flow chamber was kept at 37 °C. Once

fully filled and bubble free, a U-2 OS cell suspension in modified medium (DMEM + 10% FBS and 2% TSB

[4]) was introduced. After filling the chamber, flow was stopped (1.5 h) to allow for U-2 OS adhesion.

Modified culture medium supplemented with 2% HEPES buffer was then flowed at 0.14 s-1 shear rate for

48 h. Phase-contrast images were taken from each patterned array and from the silanized-glass control at

1.5 h to determine the number of adhering cells per unit area and total surface coverage of spread U-2 OS

cells. Statistical analysis of variance used Tukey’s HSD post hoc test and a p-value of <0.05 was considered

significant.

After 48 h the substratum was removed. Substrata were fixed in 30 mL of 3.7% formaldehyde in

cytoskeleton stabilization buffer (CS; 0.1 M Pipes, 1 mM EGTA, 4% (w/v) PEG 8000, pH 6.9). After 5 min,

this fixation step was repeated. Slides were then incubated in 0.5% Triton X-100 (3 min), rinsed with PBS,

and stained for 30 min in 5 mL PBS containing 49 µL DAPI and 2 µg/mL of TRITC–Phalloidin. After 4x wash

in PBS, the slides were examined by fluorescence microscopy (Leica DM 4000B). The number of adhering

cells per unit area and the total surface coverage of adhered cells on the patterned surface were

determined using Scion image software. To further visualize cell morphology after 48 h, cells were fixed

with 2% glutaraldehyde in 0.1 M cacodylate buffer, post fixed with OsO4 (1% in 0.1 M cacodylate buffer)

for 1 h and dehydrated using an ethanol series. Cells were incubated in tetramethylsilane for 15 min, air

dried and sputter coated with Au/Pd. Secondary electron imaging was done using a JEOL JSM 6301F SEM

at 2 kV.

AFM force measurement

Bacterial probe preparation

S. aureus ATCC 12600 was immobilized on tipless V-shaped cantilevers (VEECO, DNP-0) via electrostatic

interaction with poly-L-lysine (PLL). A drop of PLL solution was placed on a glass slide and the tip of the

cantilever was dipped in the droplet for 1 min. After air drying, the cantilever was dipped in bacterial

suspension (1 min). Bacterial probes were freshly prepared for each experiment. The bacterial probes

usually yielded single-cell contact with a substratum. Images with double contour lines indicated double

cell contacts, and a probe exhibiting such behavior was discarded. Each patterned surface was

interrogated by at least nine different S. aureus bacterial AFM probes. Three probes were made from each

of three different cultures of S. aureus.


55

Atomic force microscopy

Force measurements were done at room temperature in PBS (pH 7) using a Bioscope Catalyst AFM with

PeakForceTM Quantitative NanoMechanical Mapping software (Bruker). Each curve was analyzed to

produce the maximum adhesion force as the control feedback signal and the mechanical properties of the

sample (Adhesion, Modulus, Deformation, and Dissipation). By collecting data from different specimen

locations, comparative values were derived by subtracting the most adhesive (glass) and most repulsive

(gel). Calibration of bacterial probes was done using by thermal tuning to yield spring constants of 0.044

± 0.008 Nm-1.

ACKNOWLEDGEMENTS

We thank Joop de Vries (W.J. Kolff Institute, University Medical Center Groningen, The Netherlands) for

assisting with the AFM force measurements. Y. Wang and M. Libera thank the U.S. Army Research Office

for support (grant W911NF-07-0543). E-beam patterning was done at the Center for Functional

Nanomaterials, Brookhaven Nat. Lab. (U.S. DOE, Basic Energy Sciences, Contract #DE-AC02-98CH10886).

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[11] Engelsman, A. F.; Saldarriaga-Fernandez, I. C.; Nejadnik, M. R.; Van Dam, G. M.; Francis, K. P.; Ploeg, R. J.; et al. The risk of biomaterial-associated infection after revision surgery due to an experimental primary implant infection. Biofouling, 2010, 26, 761-767.

[12] Tetsworth, K.; Dennis, J. Contemporary management of infected total knee replacement. Curr. Orthopaed. Pract., 2008, 19, 175-180.

[13] Douthwaite, S. Designer drugs for discerning bugs. Proc. Nat. Acad. Sci. U S A, 2010, 107, 17065-17066.

[14] Projan, S. J. Why is big Pharma getting out of antibacterial drug discovery? Curr. Opin. Microbiol., 2003, 6, 427-430.

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[15] Richards, R. G.; Harris, L. G.; Schneider, E.; Haas, N. Antiseptics and antibiotics on implants. Injury, 2006, 37, S113-S116.

[16] Tiller, J. C.; Lioa, C-J.; Lewis, K.; Klibanov, A. M. Designing surfaces that kill bacteria on contact. Proc. Nat. Acad. Sci. U S A, 2001, 98, 5981-5985.

[17] Vasilev, K.; Cook, J.; Griesser, H. J. Antibacterial surfaces for biomedical devices. Expert Rev. of Med. Devic., 2009, 6, 553-567.

[18] Israelachvili J. The different faces of poly(ethylene glycol). Proc. Nat. Acad. Sci. U S A, 1997, 94, 8378-8379.

[19] Prime, K. L.; Whitesides, G. M. Adsorption of proteins onto surfaces containing end-attached oligo(ethylene oxide): A model system using self-assembled monolayers. J. Am. Chem. Soc., 1993, 115, 10714-10721.

[20] Hucknall, A.; Rangarajan, S.; Chilkoti, A. In pursuit of zero: Polymer brushes that resist the adsorption of proteins. Adv. Mater., 2009, 21, 2441-2446.

[21] Lichter, J. A.; Van Vlietpa, K. J.; Rubner, M. F. Design of antibacterial surfaces and interfaces: Polyelectrolyte multilayers as a multifunctional platform. Macromolecules, 2009, 42, 8573-8586.

[22] Magin, C. M.; Cooper, S. P.; Brennan, A. B. Non-toxic antifouling strategies. Mater. Today, 2010, 13, 36-44.

[23] Wagner, V. E.; Koberstein, J. T.; Bryers, J. D. Protein and bacterial fouling characteristics of peptide and antibody decorated surfaces of PEG-poly(acrylic acid) co-polymers. Biomaterials, 2004, 25, 2247-2263.

[24] Hu, X.; Neoh, K. G.; Shi, Z.; Kang, E. T.; Poh, C.; Wang, W. An in vitro assessment of titanium functionalized with polysaccharides conjugated with vascular endothelial growth factor for enhanced osseointegration and inhibition of bacterial adhesion. Biomaterials, 2010, 31, 8854-8863.

[25] Vasilev, K.; Sah, V.; Anselme, K.; Ndi, C.; Mateescu, M.; Dollmann, B.; et al. Tunable antibacterial coatings that support mammalian cell growth. Nano Lett., 2010, 10, 202-207.

[26] Dubash, A. D.; Menold, M. M.; Samson, T.; Boulter, E.; Garcia-Mata, R.; Doughman, R.; et al. Focal Adhesions: New Angles on Old Structures. Intern. Rev. Cell. Mol. Biol., 2009, 277, 1-65.

[27] Christman, K. L.; Enrique-Rios, V. D.; Maynard, H. D. Nanopatterning proteins and peptides. Soft Matter, 2006, 2, 928-939.

[28] Mitragotri, S.; Lahann, J. Physical approaches to biomaterial design. Nat. Mater., 2009, 8, 15-23.

[29] Christman, K. L.; Schopf, E.; Broyer, R. M.; Li, R. C.; Chen, Y.; Maynard, H. D. Positioning multiple proteins at the nanoscale with electron beam cross-linked functional polymers. J. Am. Chem. Soc., 2009, 131, 521-527.

[30] Hong, Y.; Krsko, P.; Libera, M. Protein surface patterning using nanoscale poly(ethylene glycol) nanohydrogels. Langmuir, 2004, 20, 11123-11126.

[31] Krsko, P.; Mansfield, M.; Sukhishvili, S.; Clancy, R.; Libera, M. Electron-beam patterned poly(ethylene glycol) microhydrogels. Langmuir, 2003, 19, 5618-5625.

[32] Krsko, P.; Kaplan, J.; Libera, M. Spatially controlled bacterial adhesion using surface-patterned poly(ethylene glycol) hydrogels. Acta Biomater., 2009, 5, 589-596.

[33] Krsko, P.; McCann, T.; Thach, T-T.; Laabs, T.; Geller, H.; Libera, M. Length-scale mediated adhesion and directed growth of neural cells by surface-patterned poly(ethylene glycol) hydrogels. Biomaterials, 2009, 30, 721-729.

[34] Pautke, C.; Schieker, M.; Tischer, T.; Kolk, A.; Neth, P.; Mutschler, W.; et al. Characterization of osteosarcoma cell lines MG-63, Saos-2 and U-2 OS in comparison to human osteoblasts. Anticancer Res., 2004, 24, 3743-3748.

[35] De Ruijter, J. E.; Ter Brugge, P. J.; Dieudonne, S. C.; Van Vliet, S. J.; Torensma, R.; Jansen, J. A. Analysis of integrin expression in U2OS cells cultured on various calcium phosphate ceramic substrates. Tissue Eng., 2001, 7, 279-289.

[36] Clover, J.; Gowen, M. Are MG-63 and HOS TE85 human osteosarcoma cell lines representative models of the osteoblastic phenotype? Bone, 1994, 15, 585-591.

[37] Chen, C. S.; Mrksich, M.; Huang, S.; Whitesides, G. M.; Ingber, D. E. Geometric control of cell life and death. Science, 1997, 27, 1425-1428.

[38] Arnold, M.; Schwieder, M.; Blommel, J.; Cavalcanti-Adam, E. A.; Lopez-Garcia, M.; Kessler, H.; et al. Cell interactions with hierarchically structured nano-patterned adhesive surfaces. Soft Matter, 2009, 5, 72-77.


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[39] Cavalcanti-Adam, E. A.; Volberg, T.; Micoulet, A.; Kessler, .; Geiger, B.; Spatz, JP. Cell spreading and focal adhesion dynamics are regulated by spacing of integrin ligands. Biophys. J., 2007, 92, 2964-2974.

[40] George, P. A.; Doran, M. R.; Croll, T. I.; Munro, T. P.; Cooper-White, J. J. Nanoscale presentation of cell adhesive molecules via block copolymer self-assembly. Biomaterials, 2009, 30, 4732-4737.

[41] Jaehrling, S.; Thelen, K.; Wolfram, T.; Pollerberg, G. E.; Nanopatterns bio-functionalized with cell adhesion molecule dm-grasp offered as cell substrate: Spacing determines attachment and differentiation of neurons. Nano Lett., 2009, 9, 4115-4121.

[42] Lehnert, D.; Wehrle-Haller, B.; David, C.; Weiland, U.; Ballestrem, C.; Imhof, B. A.; et al.

Cell behaviour on micropatterned substrata: Limits of extracellular matrix geometry for spreading and adhesion. J. Cell Sci., 2004, 117, 41-52.

[43] Malmstrom, J.; Christensen, B.; Jakobsen, H. P.; Lovmand, J.; Foldbjerg, R.; Sorensen, E. S.; et al. Large area protein patterning reveals nanoscale control of focal adhesion development. Nano Lett., 2010, 10, 686-694.

[44] Chaffin, W. L,; Candida albicans cell wall proteins. Microbiol. Mol. Biol. Rev. 2008, 72, 495-544.

[45] Krsko, P.; Saaem, I.; Clancy, R.; Geller, H.; Soteropoulos, P.; Libera, M. E-beam patterned hydrogels to control nanoscale surface bioactivity In: Nanofabrication: Technologies, Devices, and Applications II. (Lai WY, Ocola LE, Pau S, eds) Bellingham: SPIE, 2005, p. 60020.

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CHAPTER 3

A FUNCTIONAL DNASE I COATING TO PREVENT ADHESION OF

BACTERIA AND THE FORMATION OF BIOFILM

Jan J.T.M. Swartjes, Theerthankar Das, Shahriar Sharifi, Guruprakash Subbiahdoss,

Prashant K. Sharma, Bastiaan P. Krom, Henk J. Busscher, and Henny C. van der Mei

Adv. Funct. Mater., 2013, 23, 2843-2849

CHAPTER 3

60

ABSTRACT

Biofilms are detrimental in many industrial and biomedical applications and prevention of biofilm

formation has been a prime challenge for decades. Biofilms consist of communities of adhering bacteria,

supported and protected by extracellular-polymeric-substances (EPS), the so-called “house of biofilm

organisms”. EPS consists of water, proteins, polysaccharides and extracellular DNA (eDNA). eDNA, being

the longest molecule in EPS, connects the different EPS components and therewith holds an adhering

biofilm together. eDNA is associated with bacterial cell surfaces by specific and non-specific mechanisms,

mediating binding of other biopolymers in EPS. eDNA therewith assists in facilitating adhesion, aggregation

and maintenance of biofilm structure. Here, we describe a new method to prevent biofilm formation on

surfaces by applying a DNase I enzyme coating to polymethylmethacrylate, using dopamine as an

intermediate. The intermediate coupling layer and final DNase I coating were characterized by water-

contact-angle measurements and X-ray photoelectron-spectroscopy.

A FUNCTIONAL DNASE I COATING TO PREVENT ADHESION OF BACTERIA

61

INTRODUCTION

Bacteria exist in nature on nearly all surfaces and have a strong potential to form biofilms that has been

beyond prevention for centuries [1,2]. Bacterial biofilms are defined as groups or clusters of bacteria,

embedded in a self-produced matrix of extracellular polymeric substances (EPS) forming a three-

dimensional structure [3–5]. Despite extensive studies over many decades, the mechanisms involved in

biofilm formation are still not fully understood and as a consequence, prevention of biofilm formation

remains a prime challenge in many industrial and biomedical applications. In industrial applications,

biofilms inflict major damage when formed on processing equipment or in pipes used to transport

resources. In the biomedical field, infections that arise after implantation of a biomaterial implant (e.g.

prosthetic hips and knees) or device (pace makers) are known to be very persistent and difficult to treat

due to the formation of biofilms by adhering bacteria and in worst cases cause death of the patient.

Moreover dental caries, the number one infectious disease in the world, is due to biofilms [6].

Biofilm formation is a cyclic and step-by-step process, starting when bacteria adhere to a substratum

surface, which is first and foremost a reversible process [1]. Initial adhesion however, is rapidly followed

by a transition to irreversible adhesion through removal of interfacial water and adjustment of bacterial

cell surface adhesion sites, generally occurring within several minutes. Next, aggregation, co-adhesion and

growth lead to microcolony formation while continued production of EPS ultimately results in the

construction of “the house” of biofilm organisms,[7] providing mechanical stability to the biofilm [6,8]. At

later stages after biofilm maturation, biofilms may release single bacteria for colonization of new surfaces

[9].

EPS can be associated with the bacterial cell surface or excreted in the environment and is involved in all

stages of biofilm formation including initial adhesion of planktonic bacteria [10,11]. EPS is primarily

composed of proteins, polysaccharides and extracellular DNA (eDNA) and is often referred to as the glue

or cement that holds a biofilm together in its 3-dimensional housing [7]. eDNA, being the longest molecule

in EPS and shown to be associated with proteins and polysaccharides,[12,13] could well be the most

important component of EPS keeping all its constituents together. Goodman and co-workers recently

showed that Escherichia coli strain U91 produces two types of proteins (HU and IHF, both belonging to the

DNABII family) that are responsible for arrangement of eDNA and also help eDNA in maintaining the

integrity of the EPS in a biofilm [14]. eDNA is generally considered to originate either from the

chromosomes of lysed cells in a biofilm or to be actively released via vesicles by metabolically active

bacteria.[15–17] These bacterial membrane-derived vesicles have long been known to contain DNA,[18]

but only recently their presence in bacterial biofilms has been shown [19].

The essential role of eDNA as a component of EPS was first reported by Whitchurch et al. for Pseudomonas

aeruginosa biofilms [20]. They showed that presence of DNase I, an enzyme which non-specifically cleaves

DNA by breaking phosphodiester bonds of the phosphate backbone through hydrolysis, in growth medium

CHAPTER 3

62

could prevent biofilm formation and also rinsing a biofilm with DNase I could effectively disintegrate 60 h

old biofilms.[20] Similar effects of DNase I treatment on whole biofilms have been later reported for Gram-

positive and Gram-negative bacterial species [21], including Enterococcus faecalis and Staphylococcus

aureus [22,23].

The important role of eDNA in biofilms (see overview in Table 1) can be attributed to its supporting

function in adhesion and aggregation of bacteria, both crucial in the primary events leading to biofilm

formation. Several studies reported that eDNA is involved in initial adhesion of bacteria to substratum

surfaces. For instance, in S. aureus eDNA release due to cell lysis during initial stages of biofilm formation

is essential for bacterial adhesion to the surface, and removal of eDNA or blocking of eDNA production at

any time during biofilm formation results in a decrease in bacterial adhesion [24]. Similarly, initial adhesion

of Staphylococcus epidermidis, S. aureus and Streptococcus mutans to a glass surface was inhibited by

removal of eDNA.[25] Besides adhesion, removal of eDNA also reduced aggregation in S. mutans, S.

epidermidis, P. aeruginosa [26] and E. coli.[27] Similar as observed for adhesion, aggregation of DNase I

treated bacteria could not be fully restored by the addition of purified DNA [12,28].

TABLE 1 Different roles proposed in the literature for extracellular DNA in biofilm formation.

Bacterial species Role of eDNA in biofilm formation References

Listeria monocytogenes,

Bacillus cereus,

S. epidermidis,

S. aureus,

S. mutans

Increased initial adhesion [19,29,30]

S. mutans,

S. epidermidis,

P. aeruginosa,

E. coli,

S. aureus

Increased aggregation [3,14,25,27]

P. aeruginosa,

E. faecalis,

S. aureus,

Enhanced mechanical stability of biofilm [2,21,31]

Caulobacter crescentus Inhibition of hold-fast adhesion [32]


63

eDNA can interact non-specifically with bacterial cell surfaces, but different bacterial species also possess

DNA binding proteins that specifically interact with eDNA in addition to their non-specific interaction [13].

However, in many bacterial species, non-specific mechanisms are involved in eDNA-mediated adhesion

and aggregation. Liu et al. indicated that aggregation was independent of the source of the DNA and

therefore concluded that DNA enhances aggregation of E. coli through a non-specific mechanism [27]. In

support of this, it was shown that DNA-mediated aggregation is due to a combination of attractive Lifshitz-

Van der Waals forces and attractive acid-base interactions [26].

Since eDNA is essential in bacterial adhesion and biofilm formation, attacking this pivotal component of

EPS as the cement that holds a biofilm together by DNase I strands, has been considered as a possible

approach to prevent biofilm formation. Hitherto, the effectiveness of DNase I in inhibiting biofilm

formation has been shown either by pre-treating bacteria prior to their adhesion or post-treating biofilms,

neither of which would be a strategy to prevent biofilm formation in nature. Pre-treatment of bacteria

requires continued exposure of new generations to DNase I, while post-treatment involves penetration of

the DNase I into a biofilm, which can be a cumbersome process depending on the strains and

environmental conditions for growth involved. Rather, a better and completely new strategy would be to

have DNase I present at the time and place that bacteria first come in contact with a substratum surface.

In this study, we created a functional DNase I coating on a surface, while maintaining its biological activity.

A DNase I coating on substratum surfaces was achieved through a dopamine coupling agent and we

established this DNase I to be capable of inhibiting bacterial adhesion and biofilm formation by disrupting

eDNA in EPS. The DNase I coating strongly reduced bacterial adhesion and prevented biofilm formation up

to 14 h, without affecting mammalian cell adhesion and proliferation.

RESULTS

Polymethylmethacrylate (PMMA), a commonly used biomaterial, was coated with dopamine as an

intermediate coupling layer (see Figure 1), reducing the water contact angle on PMMA from 65 ± 6 degrees

to 48 ± 4 degrees. Determination of the elemental surface composition of the coatings by X-ray electron

spectroscopy (XPS) showed that the % nitrogen upon coating the PMMA surface with dopamine increased

to 7.7%. Concurrently, the (C-N) component at 286.1 eV of the C1s peak increased (see Table 2), attesting

an increase in amine content as related to the presence of a dopamine layer. Next, dopamine coated

PMMA surfaces were immersed in a DNase I solution for 6 - 8 h at room temperature. This caused a further

decrease in water contact angle to 25 ± 10 degrees and a reduction of the % nitrogen to 4.1%, while the

(O=C-N) component at 288.0 eV of the C1s peak increased, due to an increase in the number of peptide

bonds in the DNase I overcoat. Note that full chemical analysis of the coating is impossible, since all

elements other than nitrogen appear in PMMA, as well as in the coating components.

CHAPTER 3

64

TABLE 2 Decomposition of the C1s photo-electron peaks (binding energies given in eV) obtained in the process of coating PMMA with a DNase I layer, into four components, due to different chemical functionalities.

Sample C284.8

(C-C)

C286.1

(C-N)

C288.0

(O=C-O)

C289.2

(O=C-O)

PMMA 67.9% 18.4% 13.8% 0.0%

Dopamine coating 59.8% 33.5% 4.5% 2.2%

DNase I overcoat 49.0% 36.7% 10.6% 3.7%

FIGURE 1 Adhesion of dopamine to PMMA by dip coating and subsequent DNase I coupling to the polydopamine film.

The activity of freshly prepared and heat inactivated DNase I coatings was established by studying the

hydrolysis of plasmid DNA in solution droplets placed on the coatings. Full hydrolysis of plasmid DNA on

freshly prepared DNase I coated PMMA occurred within 30 min (see Figure 2A), while in contrast no

hydrolysis of plasmid DNA occurred within 60 min on coatings of heat inactivated DNase I. Note that

immobilized DNase I and soluble DNase I showed complete hydrolysis of plasmid DNA, indicating that the

covalent immobilization did not affect the activity of DNase I (Figure 2B).

TRIS pH 8.5

PMMA

Polydopamine film on PMMA

DNase I coated PMMA

Dopamine


65

In order to evaluate the stability of the DNase I coating, phosphate buffered saline (PBS) droplets were left

on the coating for 1, 4, 8, or 24 h (Figure 2B), after which plasmid DNA was placed on the coating on the

same spot as the PBS droplet. Hydrolysis of plasmid DNA on the surface was observed up to 8 h, while loss

of activity of the DNAse I coating was indicated between 8 and 24 h (Figure 2B). The PBS droplet removed

was mixed with DNA in order to detect DNase I possibly released from the coating, no activity in the PBS

droplet was observed (see also Figure 2B).

FIGURE 2 (A) Agarose gel, showing the degradation of plasmid DNA in a droplet on PMMA coated with DNase I with and without heat inactivation. The plasmid DNA is hydrolyzed within 30 min when placed on PMMA coated with active DNAse I. (B) Agarose gel, showing the result of the DNase I release from the coating and the activity of remaining DNase I coating after exposure to PBS for up to one day. Plasmid DNA is shown as a negative control (-), whereas plasmid DNA after mixing with DNase I in PBS (+, left lane) and plasmid DNA on freshly coated DNase I (+, right lane) are shown as positive controls. Time-indicated gels (1 h to 24 h) show hydrolysis of plasmid DNA on the DNase I coating after exposure to PBS (left lanes) and hydrolysis of plasmid DNA with PBS droplets containing the released DNase I from the coating (right lanes).

The initial adhesion of bacteria to the DNase I coating on PMMA was strongly decreased compared to

substrata without an active coating (see Figure 3A). After 60 min, the number of bacteria that adhered to

DNase I coated PMMA was reduced by 99% for P. aeruginosa PAO1 and 95% for S. aureus ATCC 12600

compared to PMMA, dopamine coated PMMA and heat inactivated DNase I coated PMMA. Confocal laser

scanning microscopic (CLSM) images show biofilms of P. aeruginosa PAO1 on all control substrata that

fully cover the surface, whereas on DNase I coated PMMA only few adhering bacteria were observed, most

of which were dead (Figure 3B). Similar observations were made for S. aureus ATCC 12600 biofilms.

Quantitative analysis of CLSM images of P. aeruginosa PAO1 and S. aureus ATCC 12600 biofilms after 14 h

on control substrata showed average biofilm thicknesses of approximately 10 and 18 µm, respectively,

whereas on PMMA with DNase I coating average biofilm thickness was reduced to 0.2 and 3 µm,

respectively (Figure 3C).

Heat inactivated DNase I coated

PMMA

DNase I coated PMMA

Droplet containing

plasmid DNA

0 15 30 60 min0 15 30 60 min

24h8h4h1h- + +

A

B

CHAPTER 3

66

FIGURE 3 (A) Initial adhesion of P. aeruginosa PAO1 and S. aureus ATCC 12600 after 60 min on various substrata. Error bars represent the standard deviations over three experiments with separately grown bacteria. (B) CLSM images of 14 h old P. aeruginosa PAO1 biofilms on different coatings, scale bar represents 100 µm. Green and red dots represent live and dead bacteria, respectively. (C) Average thickness of 14 h old P. aeruginosa PAO1 (green) and S. aureus ATCC 12600 (grey) biofilms. Error bars represent the standard deviations over three experiments with separately grown bacteria and three spots taken per sample. *indicates significant decrease (p<0.05) in initial bacterial adhesion (A) and biofilm thickness (C) on DNase I coated on PMMA substratum in comparison to all other substrata.

For biomedical application of the coating, it is necessary to demonstrate the absence of adverse effects

on the interaction with mammalian cells. Therefore the effect of the DNase I coating on adhesion and

proliferation of human osteosarcoma U-2 OS cells was studied. Coating of PMMA with dopamine, or DNase

I with and without heat inactivation showed no effect on the adhesion and proliferation of U-2 OS cells

(see Figure 4). Cells were able to adhere to all substrata and proliferate to form a confluent layer within

24 h. Furthermore, no significant differences were observed between the various substrata with respect

to the total number of adhering U-2 OS cells and the area covered by cells after 24 h of growth (Figure 4).

Bare PMMA Dopamine coated PMMA

Heat inactivated DNase I coated PMMA DNase I coated PMMA

0

1

2

3

4

5

6

7

8

9

10

Bare Dopamine coated Heat inactivatedDNase I coated

DNase I coated

Tota

l nu

mb

er

of

adh

eri

ng

bac

teri

a af

ter

60

min

(×

10

6/

cm2)

P. aeruginosa PAO1

S. aureus ATCC 12600

0

5

10

15

20

25

30

Bare Dopaminecoated

Heat inactivatedDNase I coated

DNase I coated

Ave

rage

bio

film

th

ickn

ess

afte

r 1

4 h

(μ

m)

PMMA

P. aeruginosa PAO1S.aureus ATCC 12600

A

C

B


67

FIGURE 4 (A) CLSM images of U-2 OS cell adhesion and proliferation on different substrata. Scale bar represents 100 µm. (B) The total number of U-2 OS cells adhering after 24 h incubation on various substrata. Error bars represent the standard deviations over three experiments with separately grown U-2 OS cells. No significant difference in cell adhesion and proliferation was observed between DNase I coated on the PMMA substratum in comparison to other substrata.

DISCUSSION

DNA is associated with the bacterial cell surface by both specific and non-specific interactions. Present at

the bacterial cell surface, eDNA acts as a bridge between the bacterial cell wall and EPS by binding

biopolymers in EPS, most likely polysaccharides and proteins, through attractive, short-range acid-base

interactions. Therewith eDNA is not only responsible for the presence of structurally intact EPS on bacterial

cell surfaces, but is also a crucial component in constructing the house of biofilm that holds the organisms

together and on a surface during biofilm formation (Figure 5A). Degradation of eDNA by our DNase I

coating disrupts the EPS glue of adhering bacteria, interfering with the initial step of biofilm formation and

preventing the construction of the house of biofilm organisms (Figure 5B). Degradation of eDNA upon

contact with the DNase I coating prevents bacteria from adhering. In case where a bacterium does manage

to adhere to the coated surface, degradation of the eDNA leads to disintegration of the EPS formed during

growth of the adhering bacteria, leaving it unable to develop into a structurally stable house [24].

Additionally, the presence of DNase I at the bacterial-substratum interface interferes with the aggregation

of bacteria and cell-cell interactions at the substratum surface, since these processes are both mediated

by eDNA [25,26].

Dopamine coated PMMA

Heat inactivatedDNase I coatedPMMA

DNase I coated PMMA

Bare PMMA

0

1

2

3

4

5

6

7

Bare Dopamine coated Heat inactivatedDNase I coated

DNase I coated

Nu

mb

er o

f ad

her

ing

U2

OS

cells

aft

er 2

4 h

(

×1

04

/ cm

2 )

PMMA

A B

CHAPTER 3

68

FIGURE 5 (A) eDNA acting as a bridge between a bacterial cell surface and various biopolymers in EPS, like proteins and polysaccharides, therewith playing an important role in bacterial adhesion. (B) Disruption of EPS by DNase I coating attacking the eDNA component of the EPS, preventing bacterial adhesion to the substratum surface.

In this study it was demonstrated that a DNase I coating is effective against biofilm formation for up to a

minimum of 14 h, while hydrolysis of DNA by the coating did not occur anymore after 24 h. No release of

active DNase I into PBS was observed within 24 h, suggesting that either the coated enzyme became

inactive after 8 h or release from the coating caused DNase I to become inactive after that period. Since it

has been shown that DNase I is amongst the most stable enzymes known,[33] we believe it is likely that

desorption of DNase I will occur.

The loss of activity of our DNase I coating within 24 h does not necessarily limit its applicability. Application

in industrial processes, running 24 h per day for months, will not be possible when activity is lost within 24

h, but in the biomedical field many prophylactic measures with respect to the development of biomaterial-

associated infection, i.e. infections associated with the surgical implantation of biomaterial implants and

devices (prosthetic hips and knee, pace makers, etc.) are only active for 24 h. Currently, bacteria that are

inevitably introduced during implantation of biomaterial implants or devices are eliminated through

systemic antibiotic administration and local-antibiotic delivery materials. Local antibiotic delivery

materials, such as gentamicin-loaded bone cements in orthopedics for the fixation of hip and knee

prostheses, are also known to be active for 24 h utmost,[34] similar to the period during which our current

DNase I coating is antimicrobially active. Therefore, in an era where the efficacy of many antibiotics is

Bacterium

Substratum

Bacterium

Substratum

proteins

polysaccharides

eDNA

DNase I

A

B


69

fading with respect to many pathogens involved in biomaterial-associated infection, a DNase I coating is a

badly needed addendum to the antimicrobial armamentarium in modern medicine. Important for the

further downstream translation of the DNase I coating toward clinical application is that DNase I is

naturally produced in the human body by the pancreas, kidneys, liver and subsequently released into body

fluids [35]. Moreover, aerosolized DNase I is often used via an inhaler for reducing the viscosity of the

sputum of cystic fibrosis patients through hydrolysis of DNA [36,37].

In conclusion, our biocompatible DNase I coating on PMMA resulted in unprecedented reductions of initial

bacterial adhesion for 60 min and further biofilm inhibition for up to 14 h by a Pseudomonas and

Staphylococcus strain. Although optimization of the coating might extend this time period, prevention of

biofilm formation over a time span of 14 h is sufficient to prevent biomaterial-associated infection due to

contamination of a biomaterial implant or device during surgery.


DNase I coating on PMMA

PMMA was coated with DNase I (Fermentas life sciences, Roosendaal, The Netherlands) using dopamine

as an intermediate coupling layer. DNase I is a non-toxic enzyme and no report has been published showing

bactericidal effects of DNase I, since it only degrades eDNA on the outside of a bacterium [38]. Dopamine

was chosen as a coupling agent, because it has been shown not to interfere with mammalian cell adhesion

and proliferation.[39,40] Adhesion of dopamine to organic surfaces involves oxidation of catechol to

quinone, which then reacts with the surface, and other catechols or quinones to form a polymer film [41–

43]. Although the resulting polymer film is chemically heterogeneous and its composition is not precisely

known, quinone functional groups are believed to be present at the interface and capable of covalently

binding nucleophiles by Michael addition reactions [41,44].

PMMA (Vink, Didam, The Netherlands) substrata (1.5 × 1.5 cm) were cleaned by sonication for 3 min in

2% RBS35 (Omnilabo International BV, The Netherlands) followed by rinsing with tap water, methanol, tap

water and finally with sterile ultrapure water. The PMMA substrata were then immersed in 8 ml of

dopamine (Sigma-Aldrich, St. Louis, USA) solution (6 mg ml-1 dopamine-HCl in 10 mM Tris-HCl (Merck

KGaA, Darmstadt, Germany) at pH 8.5) for 48 h at room temperature under static conditions. After 48 h,

the PMMA substrata were dried in an oven at 40°C for 6 h, followed by rinsing with sterile ultrapure water,

drying again for 2 h at 40°C and stored at 4°C. To coat with active DNase I or heat inactivated (at 65°C for

10 min) DNase I, dopamine coated PMMA surfaces were immersed in 5 ml (20 units ml-1) of a DNase I

solution in PBS (150 mM NaCl, 10 mM potassium phosphate, pH 6.8) in the presence of 10 mM MgCl2 for

6 - 8 h at room temperature under static conditions. A schematic description of the coating procedure is

CHAPTER 3

70

shown in Figure 1. After coating, the samples were rinsed with sterile ultrapure water, stored at -20C and

used within 14 h.

Water contact angles were measured on bare PMMA, dopamine coated PMMA, and on heat inactivated

and active DNase I coatings using the sessile drop method and a home-made contour monitor. The

elemental composition of PMMA, the dopamine coupling layer and the entire DNase I coating were

determined using XPS (S-probe, Surface Science Instruments, Mountain View, CA, USA). First, samples

were placed in the pre-vacuum chamber of the XPS, and then subjected to a vacuum of 10-7 Pa. X-rays (10

kV, 22 mA), at a spot size of 250 1000 m, were produced using an aluminum anode. Scans of the overall

spectrum in the binding energy range of 1-1100 eV were made at low resolution (pass energy 150 eV). The

area under each peak was used to yield elemental surface concentrations for C, O, N, and other minor

elements occurring after correction with sensitivity factors provided by the manufacturer. The C1s peaks

were decomposed in four components, i.e. for carbon involved in alkyl bonds (C-(C,H); 284.8 eV), amine

bonds (C-N); 286.1 eV), amide groups (O=C-N; 288.0 eV) and carbon arising from carboxylic acid (O=C-OH;

289.2).

Analysis of DNase I activity and stability

The activity of DNase I coatings was determined by placing a 40 µl droplet of ultrapure water containing

90 ng µl-1 plasmid DNA on DNase I coated PMMA samples for 60 min at 37C under static conditions. For

control, PMMA samples were employed that were coated with DNase I with or without heat inactivation.

After every 15 min time intervals up till 60 min, 5 µl aliquots of the DNA solution were removed and

analyzed for hydrolysis following electrophoresis on a 1.5% agarose gel. 5 µl samples were loaded using 1

µl 6x loading buffer (Roti® load DNA, Carl Roth GmbH, Karlsruhe, Germany), after which the gel was left

running for approximately 1.5 h at a constant potential of 85 V. Images were taken directly after running

the gels, under UV light using a digital camera.

To assess the stability of the coatings, release of DNase I from PMMA samples coated with active DNAse I

into the surrounding environment was measured by placing a droplet of 100 µl PBS on five coupons at

37C for 1, 4, 8 and 24 h respectively. At the end of this time period, 25 µl was taken from the droplet,

mixed with 25 µl of plasmid DNA (10 ng µl-1) and incubated for 1 h at 37C, followed by agarose gel

electrophoresis (see above). The activity of DNAse I remaining in the coating was measured by removing

the droplets of PBS completely and placing a new 50 µl droplet containing plasmid DNA (10 ng µl-1) on the

same five spots of the coating. After incubating at 37C for 1 h, hydrolysis of plasmid DNA in this 50 µl

droplet was determined using agarose gel electrophoresis.

Bacterial species and culture conditions

P. aeruginosa PAO1 and S. aureus ATCC 12600 were stored in 7% DMSO at -80C. Strains were inoculated

onto blood agar plates and incubated overnight under aerobic conditions at 37C. Single colonies from the

agar plates were used to inoculate 10 ml pre-cultures in tryptone soya broth (TSB, OXOID, Basingstoke,


71

UK) and cultured for 24 h. This culture was in turn used to inoculate a 200 ml main culture in TSB, which

was grown for 16 h prior to harvesting. Bacteria were harvested by centrifugation at 5000 × g for 5 min at

10C, and washed twice with PBS. S. aureus ATCC 12600 aggregates were disrupted by sonication on ice

for 3 x 10 s at 30 W (Vibra Cell model 375, Sonics and Materials Inc., Danbury, Connecticut, USA). The

bacterial densities of the suspensions were determined using a Bürker-Türk counting chamber and

bacteria were diluted in 50 ml PBS to a final density of 3 × 108 bacteria per ml for adhesion and biofilm

experiments.

Initial bacterial adhesion and biofilm growth

Both initial adhesion of bacteria and biofilm growth were studied on bare PMMA, dopamine coated

PMMA and PMMA coated with active and heat inactivated DNAse I. For initial adhesion, 5 ml of a bacterial

suspension was added to each substratum, inserted in 6 wells plates and bacteria were allowed to adhere

at 37C at 60 rpm. Bacterial suspensions were removed after 60 min and the substrata were gently washed

with PBS. The total number of adhering bacteria were then counted after staining with Live/Dead stain

(BacLightTM, Invitrogen, Breda, The Netherlands) for 15 min in the dark using fluorescence microscopy

(Leica DM4000B, Leica Microsystems GmbH, Heidelberg, Germany).

Biofilm growth was initiated after 60 min of initial bacterial adhesion. The bacterial suspension in the wells

was replaced by 5 ml of TSB and wells were incubated for 14 h at 37C at 60 rpm. After 14 h, growth

medium was decanted and the biofilms were gently rinsed with PBS in order to remove any planktonic

bacteria followed by staining with Live/Dead stain for 15 min in the dark. Subsequently, optical cross-

sections of the biofilms were made using CLSM (Leica DMRXE TCS-SP2; Leica Microsystems Heidelberg

GmbH, Heidelberg, Germany). Optical cross-sections were further examined for the average biofilm

thickness employing COMSTAT, a Matlab (The Mathworks, Inc.) based analysis program.

Human osteosarcoma U-2 OS cell culturing and harvesting

In order to determine possible adverse effects of the DNase I coating on mammalian cells, U-2 OS

osteosarcoma cells were cultured in Dulbecco’s modified Eagle’s low glucose medium, supplemented with

10% fetal calf serum, 0.2 mM ascorbic acid-2-phosphate containing antibiotics (penicillin/streptomycin).

U-2 OS cells were maintained in a T-75 cell culture flask at 37°C in a humidified, 5% CO2 atmosphere and

harvested at 90% confluency using trypsin/ethylenediaminetetraacetic acid. The harvested cells were

counted using a Bürker-Türk counting chamber and subsequently, diluted with growth medium to a

density of 6 × 105 cells per ml. Cell adhesion was initiated by addition of 5 ml of cell suspension to each

substratum, inserted in a 6-well plate and cells were allowed to grow for 24 h at 37°C in a humidified, 5%

CO2 atmosphere. To assess the morphology of U-2 OS cells adhering on different substrata after 24 h,

samples were prepared for immuno-cytochemical staining. For fixation, 2 ml of 3.7% formaldehyde in

cytoskeleton stabilization buffer (0.1 M Pipes, 1 mM ethylene glycol tetra-acetic acid, 4% (w/v)

polyethylene glycol 8000, pH 6.9) was added to the substrata. Subsequently, cells were incubated in 0.5%

Triton X-100 for 3 min, rinsed with PBS and stained for 30 min with 1 ml PBS containing 10 µl DAPI and 2

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72

µg ml-1 of TRITC-Phalloidin. The substrata were washed 4 times in PBS and examined with CLSM and the

number of adhering cells per unit area was determined.

Statistical analysis

The effects of functional DNase I coated PMMA on initial bacterial adhesion and on biofilm thickness were

analyzed using a two-tailed Student’s t-test. Differences were considered significant if p < 0.05.

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CHAPTER 4

A PROTECTIVE, BIODEGRADABLE PLGA-COATING RELEASING INULIN-

PACKAGED DNASE I TO PREVENT BACTERIAL

ADHESION AND BIOFILM FORMATION

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76

ABSTRACT

Enzyme coating of biomaterial surfaces has proven to be a promising in vitro concept to prevent

biomaterial associated infections. However, coating stability and robustness can be a critical factor,

depending on the site of the implant or device in the human body, especially enzymes can easily become

inactive due to changes in their tertiary structure. By using poly-(lactic-co-glycolic)-acid (PLGA) as a

biodegradable protective overcoat on titanium with inulin-packaged DNase I, bacterial adhesion of two

Staphylococcus aureus strains was drastically reduced and the biovolume of a 20 h biofilm was significantly

lowered with respect to bare titanium, a common orthopedic biomaterial. More importantly, the PLGA

coating combined with inulin-packaging of DNase I withstood different storage and handling conditions,

without negatively affecting antibacterial efficacy. With antibiotic treatment of infections becoming less

effective due to increasing antibiotic resistance worldwide, a protective, biodegradable coating of PLGA

with inulin-packaged DNase I could offer a potential new strategy to prevent the infection of medical

devices and implants, possibly even without the need for post-operative antibiotic treatment.

A PLGA-COATING RELEASING INULIN-PACKAGED DNASE I

77

INTRODUCTION

Driven by a high level of protection and increased survival rate, bacteria seek shelter on material surfaces

in their biofilm mode of growth rather than by staying planktonic in suspension [1,2]. In these sophisticated

biofilm communities, bacteria produce extracellular polymeric substances (EPS) that protect biofilm

inhabitants against predators and penetrating antimicrobials [3,4]. Combined with a low metabolic state,

bacteria in a biofilm mode of growth can easily become resistant to treatment. Accordingly, biofilm

formation represents a major problem in many industrial and biomedical applications [5–7]. When biofilms

form on biomaterial implants and devices, persistent infections develop and frequently removal of the

colonized implant or device is the only successful treatment option [4].

EPS consists mainly of proteins, polysaccharides and extracellular DNA (eDNA). The exact roles of the

different individual components of EPS are not exactly known, but collectively they provide mechanical

stability to a biofilm [8,9]. Genetically, eDNA resembles chromosomal DNA and is therefore suspected to

originate from cell lysis, although also active secretion of DNA containing vesicles has been observed

[10,11]. The presence of eDNA favors acid-base mediated crosslinking between the various components

of the EPS with a distinct role of Ca2+ ions [12]. The resulting network can trap bacteria to bring them close

enough to each other to allow cell-cell communication and facilitate metabolic interactions [1,13,14].

eDNA is not only a key factor in providing stability to mature biofilms, but also plays an important role in

the initial adhesion and aggregation of bacteria [15–18]. The recognition of the role of eDNA in maintaining

the mechanical stability of biofilms, opens a new pathway to the disruption of biofilms, i.e. their EPS matrix

using enzyme treatments. DNase I is an enzyme which catalyzes hydrolysis of the phosphodiester bonds

in the phosphate backbone of DNA and was involved in the studies leading to the initial findings on the

role of eDNA in bacterial adhesion, biofilm formation and stability [19,20]. Treatment of Pseudomonas

aeruginosa biofilms with DNase I dispersed biofilms that were up to 60 h old, whilst on 84 h old biofilms

only minor effects were seen [19]. Addition of DNase I to bacterial suspensions prior to adhesion drastically

reduced adhesion of Staphylococcus epidermidis [21] and Listeria monocytogenes [22], while initial

adhesion of an environmental Pseudomonas isolate in the presence of DNase I was decreased by more

than 95% after 30 min of adhesion [17]. Recently, we attached DNase I to polymethylmethacrylate

(PMMA) using dopamine as a coupling agent and showed that surface attached DNase I can decrease

initial bacterial adhesion and lower biofilm formation during 14 h [9]. However, the use of a biomolecule

to attach DNase I to surfaces leaves the enzyme unprotected and prone to detachment, while its surface

exposure can also result in loss of activity. Poly-(lactic-co-glycolic)-acid (PLGA) has been used as a

protective overcoat on titanium surfaces spray-coated with gentamicin [23] and as a bio-degradable,

gentamicin-loaded coating on tibia nails [24] to prevent infection. The degradation time of PLGA can be

adjusted to the application aimed for by varying the coating thickness, the ratio of poly-lactic to glycolic

acid and the molecular weight. Formation of a PLGA coating however, requires the polymer to be dissolved

in a volatile solvent that can be applied to the desired material, yielding a hard, protective layer of PLGA

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78

after evaporation of the solvent [25]. Accordingly, PLGA could be an ideal candidate to protect DNase I on

implant materials in order to prevent bacterial adhesion and biofilm formation, with the possibility to fine-

tune its release.

However, the use of volatile solvents poses a major disadvantage of using PLGA for the protection of DNase

I. Even though DNase I is one of the most stable enzymes known, when brought in solution its stability at

4°C is limited to a few days and mixing DNase I with PLGA dissolved in a volatile solvent will yield a severe

reduction of activity [26]. The stability of DNase I can be increased by spray-drying DNase I with inulin [27],

yielding inulin-packaged DNase I that retains more than 80% of enzyme activity after weeks of storage,

even at 85°C.

In this study, we combine the clinically applied concept of using a protective, biodegradable PLGA [25]

coating with inulin-packaging [27] to yield a stable DNase I releasing coating on titanium that prevents

bacterial adhesion and biofilm formation. Titanium is a common biomaterial used in various biomaterial

implants prone to bacterial colonization, such as hip and knee arthroplasties, dental implants and bone

fixator pins. By using a degradable polymer coating, DNase I is released over an extended time period,

providing breakdown of bacterial eDNA at the bacteria-biomaterial interface. In addition, inulin protects

DNase I during coating preparation, while protection by inulin also increases the stability of DNase I during

storage.


Particle preparation

Inulin-DNase I mixed (weight ratio 4 : 1) and single powders of inulin and DNase I were produced by

dissolving DNase I (from bovine pancreas, purity ≥ 86%, 400 Kunitz units/mg, Sigma Aldrich, St. Louis, MO,

USA) and inulin (DP23, Sensus, Roosendaal, The Netherlands) in water up to a total concentration of 5

mg/ml. The solutions were spray-dried using a B-90 spray-dryer (Büchi Labortechnik AG, Flawil,

Switzerland) in combination with a B-296 dehumidifier and a two-fluid nozzle. The inlet air temperature

was set at 80°C, the aspirator at 150 l/min, liquid feed flow at 1 ml/min and atomizing air flow at 50 mm.

Particle size was determined by laser diffraction (HELOS, Sympatec, Clausthal-Zellerfeld, Germany).

Surface coating

Titanium substrata (1.50 × 1.50 x 0.10 cm, Goodfellow Cambridge Ltd., Cambridge, United Kingdom) were

cleaned by sonication for 3 min in 2% RBS35 (Omnilabo International BV, The Netherlands) followed by

rinsing with water, methanol and water again. PLGA (PURASORB PDLG 5002, Corbion, Diemen, The

Netherlands) was dissolved (10% w/v) in acetonitrile (Merck KGaA, Darmstadt, Germany) while stirring.

Particle formulations consisting of either only inulin, only DNase I, or inulin-packaged DNase I particles

were added (1% w/v) to the PLGA solution in acetonitrile and stirred for an additional 3 h. 100 µl of the

suspension was applied to titanium substrata and left to dry overnight at room temperature, resulting in


79

a total of 4.4 mg PLGA/cm2, containing 0.44 mg of particles/cm2 (in the case of inulin-packaged DNase I

particles, the ratio of inulin over DNase I was 4 to 1). Coated substrata were stored at room temperature

and used within five days. The thickness of the PLGA layer was determined by making a cut through the

coating, exposing the bare titanium, and measuring the depth using white light interferometry (Proscan

2000, Scantron Industrial Products Ltd, Taunton, England).

Release kinetics

After surface coating, titanium substrata were placed in 2 ml PBS (150 mM NaCl, 10 mM potassium

phosphate, pH 7) and incubated at 37°C and 60 rpm, in order to determine the release kinetics of DNase I

and inulin from the protective PLGA coating. Individual samples were removed from PBS at different time

points (1, 4, 8, 24, 48, 72 and 96 h) after which the concentration DNase I and inulin was determined in

PBS using photospectrometry.

The amount of inulin released was determined using an anthrone assay; a commonly used assay to

determine carbohydrate concentrations [28,29]. Anthrone reagent (Sigma Aldrich) was freshly prepared

for each measurement at a concentration of 1 mg/ml in sulfuric acid (H2SO4, 95-97%, Merck KGaA,

Darmstadt, Germany). First, a calibration curve of the absorbance of inulin solutions with known

concentrations was constructed, using 0, 20, 40, 60, 80 and 100 µg/ml inulin in PBS. To measure the

absorbance, 1 ml of each solution was mixed with 2 ml anthrone reagent and vortexed. After letting the

mixture cool down for 30 min, it was transferred to a plastic cuvette and the absorbance at 630 nm was

measured (Smart Spec™ 3000, Bio-Rad, Hercules, Ca, USA). When necessary, samples were diluted 10

times in PBS and measured again. The calibration curve was used to determine the concentration of inulin

released by the different samples.

The amount of DNase I released from the samples was measured by mixing 100 µl PBS containing DNase

I released from samples with 100 µl Bradford reagent (Sigma Aldrich) and measuring the absorbance using

a NanoDrop® spectrophotometer (ND-1000, NanoDrop Technologies, Inc, Wilmington, DE) at 595 nm. The

absorbance was then compared to a calibration curve, prepared similarly as for inulin, to calculate the

amount of DNase I released.

Bacterial culture and harvesting

Green fluorescent protein (GFP) expressing Staphylococcus aureus ATCC 12600 and S. aureus Newman

D2C (also known as S. aureus ATCC 25904) were obtained by introduction of pMV158GFP [30] into

competent bacteria by electroporation, as described by Li et al. [31]. Transformants were selected on

tryptone soya broth (TSB, OXOID, Basingstoke, UK) agar (bactoagar, BD Le Pont de Claix, France) agar plates

containing 10 μg/ml of tetracycline (hydrochloride, purity ≥ 95%, Sigma). S. aureus ATCC 12600GFP and S.

aureus NewmanGFP were stored in 7% DMSO at -80 °C. Green fluorescent protein (GFP) expressing bacteria

were used, as they allow easy quantification of adhesion numbers and biofilm formation without

additional staining.

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S. aureus ATCC 12600 is a common reference strain in biofilm research, while S. aureus Newman D2C is a

highly virulent clinical isolate, known to cause highly persistent infections [32]. Strains were inoculated on

TSB agar plates containing 10 µg/ml tetracycline and incubated overnight under aerobic conditions at

37°C. Single colonies were used to inoculate 10 ml pre-cultures in TSB containing 10 µg/ml tetracycline

and cultured for 24 h. Pre-cultures were used to inoculate 200 ml main cultures in TSB in absence of

antibiotics, which were grown for 16 h. Cells were harvested by centrifugation at 5000 x g for 5 min at

10°C and washed twice with PBS, before being sonicated at 30 W (Vibra Cell model 375, Sonics and

Materials Inc., Danbury, CT, USA) for 3 x 10 s on ice with 30 s breaks in between in order to break up

bacterial aggregates. Bacterial densities were determined using a Bürker-Türk counting chamber and

suspensions were diluted to 3 x 108 bacteria per ml in PBS for all experiments.


For initial adhesion, 5 ml of bacterial suspension supplemented with 1 mM CaCl2 and 10 mM MgCl2 was

added to the various substrata in 6-well plates and incubated at 37°C and 60 rpm. After 1 h, samples were

rinsed in PBS and the total number of adhering bacteria was counted using fluorescence microscopy (Leica

DM4000B, Leica Microsystems GmbH, Heidelberg, Germany).

Biofilm growth was initiated after 1 h of initial bacterial adhesion. Samples were removed from their

suspensions and placed into new 6-well plates containing 5 ml TSB, supplemented with 1 mM CaCl2 and

10 mM MgCl2 and incubated for 20 h at 37°C and 60 rpm. After incubation, biofilm-covered samples were

carefully transferred to 6-well plates containing PBS and examined by confocal laser scanning microscopy

(CLSM) (Leica DMRXE TCS-SP2; Leica Microsystems Heidelberg GmbH, Heidelberg, Germany). Optical

cross-sections of the biofilms were taken at three different positions and their biovolumes calculated

employing COMSTAT [33,34], a Matlab (The MathWorks, Natick, MA, USA ) based analysis program.

Biofilms grown on bare titanium and protective PLGA coatings containing inulin-DNase I particles were

also evaluated using optical coherence tomography (OCT) (Telesto-II 1300, Thorlabs, Newton, NJ, USA) in

addition to CLSM, in order to rule out possible effects of DNase I on fluorescence.

Stability of the PLGA-inulin/DNase I coating

To determine the stability of the PLGA coatings containing DNase I and inulin-packaged DNase I, several

conditions were applied that could affect the efficacy of the coating during storage or handling. In a first

condition, coated substrata were placed in 6-well plates containing 5 ml PBS and incubated for 2 h at 37°C.

In a second condition, the robustness of the coating and its resistance against physical handling was

determined by pressing a finger on coated substrata with a force of 20 N for 10 s, resulting in a pressure

of approximately 3.2 x 104 Pa, after which DNase I activity was determined. Finally, coated samples were

stored at ambient conditions for 4 weeks prior to assessing DNase I activity. Efficacies of the coatings were

evaluated by assessing bacterial adhesion and biofilm formation, as described above.


81

Cell adhesion assay using U-2 OS

The presence of possible adverse effects of the coatings on mammalian cells was determined using U-2

OS osteosarcoma cells. Cells were cultured in Dulbecco’s modified Eagle’s low glucose medium

supplemented with 10% fetal bovine calf serum and 0.2 mM ascorbic acid-2-phosphate. Cells were

maintained in T-75 cell culture flasks at 37°C in a humidified, 5% CO2 atmosphere until 95% confluency.

Cells were harvested using trypsin/ethylenediaminetetraacetic acid and counted using a Bürker-Türk

counting chamber and diluted to 5.5 x 105 cells per ml in growth medium. Cell adhesion assays were

performed in 6-well plates containing each of the substrata and 5 ml of the cell suspension. After 24 h,

substrata were prepared for immuno-cytochemical staining by fixation in 3.7% paraformaldehyde in

cytoskeleton stabilization buffer (0.1 M Pipes, 1 mM ethylene glycol tetra acetic acid, 4% (w/v)

polyethylene glycol 8000, pH 6.9) and subsequent treatment with 0.5% Triton-X100. After rinsing with PBS,

substrata were incubated in 1 ml PBS containing 2 µg/ml TRITC-Phalloidin and 4 µg/ml DAPI for 30 min in

the dark. Before being examined using fluorescence microscopy, substrata were washed 4 times with PBS

to remove any excess stain, and the number of cells per cm2 on each sample was determined by counting

the number of blue-stained nuclei.

Statistics

Bacterial adhesion, biofilm formation and mammalian cell adhesion data followed a normal distribution

(Shapiro-Wilk test, p < 0.05). Differences were analyzed using a one-way ANOVA and considered significant

if p < 0.05.

RESULTS

Surface coating thickness and release kinetics

Spray-drying of inulin, DNase I or inulin-packaged DNase I, resulted in particles with a mean diameter of 1

μm. Resulting particles were added to a solution of PLGA in acetonitrile and used to coat titanium. The

thickness of the applied PLGA layer on titanium substrata was approximately 6 μm in absence and 15 μm

in presence of inulin-packaged DNase I particles, while appearing more rough in the presence of particles,

as determined using white light interferometry (Fig.1).

CHAPTER 4

82

FIGURE 1 Height map of a protective PLGA coating on titanium in absence and presence of inulin-packaged DNase I particles obtained interferometrically. The blue region represents a cut made throughout the PLGA layer, revealing the bare titanium surface. (A) PLGA coating only. (B) Coating of PLGA with inulin-packaged DNase I. Total area is 5 x 5 mm and colors are artificially generated to yield a height map.

PLGA-protected coatings containing inulin-packaged DNase I particles showed a burst release of DNase I

and inulin within the first hour of placement in PBS, after which release continued at a slower rate (Fig. 2).

While the burst release amounted approximately 25% of the total content of the coating, after 96 h more

than half of the total amounts incorporated had been released (total inulin and DNase I contents were 800

and 200 µg, respectively). Note that the ratio of inulin over DNase I released was 4 : 1 on average, in line

with the ratio of inulin over DNase I incorporated during particle preparation (see Materials and Methods

section).

FIGURE 2 (A) Calibration curves of DNase I (595 nm) and inulin (630 nm) solutions as a function of their concentration

1 2 3 4 5µm

mm 1 2 3 4 5µm

mm0

1

2

3

4

5

A B

0

1

2

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5

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25.0

30.0

0

50

100

150

200

1 4 8 24 48 72 96

DN

ase

I re

leas

ed

(µ

g)

Time (h)

DNase I

0

200

400

600

800

Inu

lin r

ele

ase

d (

µg)

Inulin

Calibration curves Cumulative release

A B

0.0

0.5

1.0

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2.0

2.5

0

0.01

0.02

0.03

0.04

0.05

0.06

0 50 100 150 200

Inu

lin a

bso

rban

ce

DN

ase

I ab

sorb

ance

Concentration (µg/ml)

DNase I

Inulin


83

in solution obtained photospectrometrically. (B) Cumulative amounts of DNase I and inulin released from PLGA-protected coatings with inulin-packaged DNase I immersed in PBS for different periods of time obtained photospectrometrically, using the calibration curves in Fig. 2A. Note that the amount of released DNase I is shown on the left axis, while the released inulin is depicted on the right axis. Error bars indicate standard deviations over three experiments with separately prepared coatings.


Fluorescent microscope images show that bacterial adhesion was greatly reduced by the release of DNase

I from PLGA coatings for both S. aureus 12600GFP and S. aureus Newman D2CGFP (Fig. 3A). The number of

initially adhering staphylococci on titanium substrata was not affected by applying a coating consisting of

only PLGA, or PLGA containing inulin particles (see Fig. 3B). Addition of particles containing DNase I to the

PLGA coating significantly reduced staphylococcal adhesion, regardless of whether particles consisted of

only DNase I, or inulin-packaged DNase I. Interestingly, S. aureus Newman D2CGFP adhered in higher

numbers than S. aureus ATCC 12600GFP to bare titanium and coatings consisting of PLGA or PLGA

containing inulin particles, but on PLGA-protected coatings containing DNase I and inulin-packaged DNase

I lower numbers of S. aureus Newman D2CGFP were found, indicating that this strain is more sensitive to

DNase I treatment than S. aureus ATCC 12600GFP.

FIGURE 3 (A). Fluorescence microscopy images of staphylococcal adhesion after 1 h in PBS to titanium surfaces and various coatings. Scale bar denotes 75 µm. (B) Number of adhering S. aureus ATCC 12600GFP and S. aureus Newman D2CGFP after 1 h adhesion in PBS to titanium surfaces and various coatings. Error bars represent the standard deviations

0

2

4

6

8

10

Nu

mb

er

of

adh

eri

ng

bac

teri

a (1

06

cm-2

)

S. aureusATCC 12600GFP

Titanium PLGA PLGA-inulin PLGA-DNase I

PLGA-inulin-packaged DNase I

S. aureusNewman D2CGFP

Titanium PLGA PLGA-inulin

PLGA-DNase I


B

A

S. aureus ATCC 12600GFP

S. aureus Newman D2CGFP

* **

*

CHAPTER 4

84

over three experiments with separately grown bacteria. *Indicates a significant difference (p < 0.05) in the numbers of adhering bacteria, within the same strain, compared to titanium and coatings not containing DNase I.

Both CLSM (Fig. 4A) and non-fluorescence-based OCT (Fig. 5) images of staphylococcal biofilms grown for

20 h show abundant biofilm formation on bare titanium, while biofilm formation is nearly absent on PLGA

coatings containing inulin-packaged DNase I particles. In line with effects on initial adhesion, coating of

titanium with either only PLGA, or PLGA containing inulin particles in absence of DNase I did not

significantly affect the biovolumes for both S. aureus strains (Fig. 4B). Importantly, S. aureus ATCC 12600GFP

biovolume was hardly reduced on PLGA coatings containing DNase I over the growth period of 20 h, but

S. aureus Newman D2CGFP showed a significant decrease in biovolume compared to titanium and titanium

with either only PLGA, or PLGA containing inulin particles. Inulin-packaging of DNase I yielded much

stronger reductions in biovolume for both strains, indicating that DNase I activity is better preserved by

inulin-packaging than by solely protecting it in PLGA. Note that the amount of DNase I being five-fold lower

in inulin-packaged DNase I coatings, compared to DNase I only coatings.

FIGURE 4 (A). CLSM overlay images of 20 h old biofilms of S. aureus ATCC 12600GFP and S. aureus Newman D2CGFP on titanium surfaces and various coatings. Scale bar denotes 75 µm.(B) Average biovolumes of 20 h old biofilms of S. aureus ATCC 12600GFP and S. aureus Newman D2CGFP grown in TSB on titanium surfaces and various coatings. Error bars

A

B

S. aureusATCC 12600GFP

Titanium PLGA PLGA-inulin PLGA-DNase IPLGA-inulin-packaged DNase I

S. aureusNewman D2CGFP

0

20

40

60

80

100

Bio

volu

me

(µm

3µ

m-2

)


PLGA-DNase I


S. aureus ATCC 12600GFP

S. aureus Newman D2CGFP

*

*

*


85

represent standard deviations over three experiments with separately grown bacteria and different batches of coated samples. *Indicates a significant difference (p < 0.05) in biovolume, within the same strain, compared to titanium and coatings not containing DNase I.

FIGURE 5 3-D (top) and side (bottom) views of staphylococcal biofilms obtained using non-fluorescence-based, optical coherence tomography. (A) S. aureus Newman D2CGFP biofilms grown on bare titanium. (B) S. aureus Newman D2CGFP biofilms grown on titanium coated with PLGA containing inulin-packaged DNase I particles.

Stability of the PLGA-inulin-packaged DNase I coating

To assess the stability of the protective PLGA coating with unpackaged and inulin-packaged DNase I,

staphylococcal adhesion and biofilm formation were also determined after different conditions of storage

and handling (see Fig. 6) and compared with adhesion and biofilm formation on bare titanium and a

previously described [9] dopamine-coupling of DNase I on PMMA.

High numbers of adhering staphylococci and large biovolumes were seen on bare titanium, regardless of

storage and handling conditions for both staphylococcal strains. PLGA yielded effective protection when

judged on the reductions achieved in initial adhesion numbers and when evaluated based on 20 h biofilm

formation; no significant differences were seen after storage or handling. Benefits of inulin-packaging of

DNase I become most evident with respect to 20 h biofilm formation. The necessity of protecting the

DNase I with PLGA and inulin follows directly from the comparison with the efficacy of dopamine-coupled

DNase I on PMMA, that is clearly far less than of our new protective, biodegradable PLGA-coating releasing

inulin-packaged DNase I particles.

A B

(mm)(mm)

0.2

(mm) 0.0

0.2

(mm) 0.0

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86

FIGURE 6 Effects of different storage and handling conditions on the efficacy of different coatings on titanium, including immediate use, 2 h exposure to PBS, exposure to a pressure of 3.2 x 104 Pa for 10 s and 4 weeks of storage in ambient air. (A, B) Numbers of adhering staphylococci after 1 h adhesion in PBS for S. aureus ATCC 12600GFP (A) and S. aureus Newman D2CGFP (B). (C, D) Biovolumes of staphylococcal biofilms grown for 20 h, for S. aureus ATCC 12600GFP (C) and S. aureus Newman D2CGFP (D). Error bars represent standard deviations over three experiments with separately grown bacteria and different batches of coated samples. *Indicates a significant difference (p < 0.05) from titanium within the same storage and handling conditions. # indicates a significant difference (p < 0.05) between PLGA-DNase I and PLGA-inulin-packaged DNase I within the same storage and handling conditions.

Cell adhesion assay using U-2 OS

For use in biomedical applications, it is important to exclude harmful effects of the coating towards

mammalian cells. The coating was therefore assessed for its influence on the adhesion and proliferation

of human osteosarcoma U-2 OS cells. None of the coatings showed any negative effect on the adhesion

and proliferation of U-2 OS cells after 24 h of incubation. Cells adhered to all substrata and formed a

confluent layer within 24 h. No significant differences in the number of adhering cells were observed

between any of the coatings (Fig. 7).

*0

1

2

3

4

5

6

7

8

0

1

2

3

4

5

6

7

8

Nu

mb

er

of

adh

eri

ng

bac

teri

a(x

10

6cm

-2)

*#

Titanium PLGA-DNase I

PLGA-inulin-packagedDNase I

No storage or handling

2 h storage in PBS

Pressure

Storage in ambient air

S. aureus ATCC 12600GFP S. aureus Newman D2CGFP

A B

C D

No storage or handling

2 h storage in PBS

Pressure

Storage in ambient air

0

20

40

60

80

100

*

Dopamine-DNase I

0

20

40

60

80

100

Bio

volu

me

(µm

3µ

m-2

)

#

* *

*

*

*

**

*

* * * * * * * *

*

*

*

*

*

*

* **

* * *

* *

*

* * * *

* *#

#

Titanium PLGA-DNase I

PLGA-inulin-packagedDNase I

Dopamine-DNase I


87

FIGURE 7 (A) Fluorescence microscopy images of adhering U- 2 OS after 24 h of growth on titanium surfaces and various coatings showing complete coverage. Scale bar denotes 75 µm. (B) Number of adhering U-2 OS cells after 24 h of growth on titanium surfaces and various coatings. Error bars represent standard deviations over three experiments with separately grown U- 2 OS cells and different batches of coated samples.

DISCUSSION

Previously, we reported on a biomaterials coating of DNase I coupled through dopamine PMMA, which

resulted in the prevention of initial bacterial adhesion, as well as reduced biofilm formation over a

timescale of 14 h [9]. Loss of enzyme activity occurred between 8 en 24 h. Combined, this indicates that

the stability of this DNase I coating varied between 14 to 20 h. Whereas it is clear that this timescale will

not be able to prevent late biomaterial-associated infection arising from the spreading of

hematogeneously introduced bacteria to the infection site, it is a matter of debate whether a timescale of

14-20 h is sufficiently long to prevent infection arising from peri-operatively introduced bacteria as is

currently done with a dose of post-operatively administered antibiotics. Moreover, dopamine-coupled

DNase I may not be able to withstand physical handling and exposure to fluids as common during insertion

of a biomaterial implant or device. The conditions under which an orthopedic implant is inserted will

inevitably lead to mechanical stress and exposure to fluids, such as for example PBS or blood. In the current

study, we have developed a PLGA coating on titanium in which DNase I is embedded. DNase I is released

upon degradation of the PLGA and disrupts the EPS matrix surrounding bacteria at the biomaterial-bacteria

interface through hydrolysis of eDNA, similar as our previous dopamine-coupled DNase I on PMMA. The

0

5

10

15

20

25

30

35

40

Nu

mb

er o

f ad

her

ing

U-2

OS

cells

(x

104

cm-2

)


PLGA-DNase I


Titanium PLGA PLGA-inulin PLGA-DNase I


A

B

CHAPTER 4

88

hardened PLGA provides protection against storage and handling conditions. Importantly, even in the

situation where an implant needs to be hammered into a tight bone junction and the integrity of the

coating will be inevitably compromised, PLGA containing DNase I will remain to be accumulated at the

implant-bone junction where protection is needed. Our coating allows for unhindered tissue integration

(Fig. 7), which is a prerequisite for a successful outcome of biomaterial associated surgery in many

applications and at the same time offers the best long-term protection against bacterial colonization [35].

To protect DNase I from the harsh volatile solvents used to dissolve and coat PLGA, we packaged DNase I

in inulin. Inulin is a sugar glass which helps to prevents damage to the tertiary structure of enzymes and

has been used for stabilizing DNase I in applications for patients suffering from cystic fibrosis [27].

Stabilization of DNase I using inulin increases the activity of coated DNase I by protecting it during the

coating process, as well as it helps to increase the shelf life. The beneficial effect is indicated by the

enhanced activity of inulin-packaged DNase I in this study. Even though coatings containing inulin-

packaged DNase I harbored only one fifth of the amount of DNase I compared to coatings with DNase I

only particles, their effectiveness against bacterial adhesion and biofilm formation was at least equal and

in some cases even increased (Fig. 6). The choice of PLGA as a protective material above other possible

degradable polymers has the advantage of already being approved by the U.S. Food and Drug

Administration (FDA) for biomedical applications, including implantation [36]. Besides being approved for

biomedical applications, the degradation rate of PLGA can be easily tuned by varying the ratio of lactic to

glycolic acid, or the molecular mass, in order to comply with the desired application requirements [37].

The rise of bacterial strains resistant to antibiotics, together with the limited development of new

antibiotics, calls for antibiotic independent mechanisms to prevent biomaterial associated infections

[38,39]. The use of DNase I seems extremely suitable in this matter, as it leaves bacteria unable to adhere

and form biofilms, causing them to stay planktonic, meaning they more easily eradicated by the host

immune system. Additionally, the authors have no knowledge of any literature reporting bacterial

resistance against enzymatic treatment. Compared to regular antibiotic treatment of biofilms on non-

adhesive coatings and similar systems releasing antibiotics rather than enzymes, the reduction in biofilm

formation by our coating is comparable [40,41]. This indicates that the results obtained using DNase I

might not require the use of additional antibiotic treatment. Moreover, the presented method of

protecting and packaging can be applied using other enzymes that are known to possess the ability to

prevent bacterial adhesion and biofilm formation, such as lysozyme and dispersin B [42,43]. Not using

antibiotics after biomaterial implant surgery, which is the premise of this coating, may be considered too

much of a risk by treating physicians and therewith impede downward clinical translation. In this respect

it is important to note that the coating can be used together with common post-operative antibiotic

treatment and it may even be expected that the antibiotic efficacy will be increased as enzymatic

disruption of the EPS matrix will prevent biofilm formation. Alternatively, antibiotics may be directly

included in the biodegradable PLGA coating [36,44].


89

CONCLUSIONS

In this study, we studied the effect of a protective, biodegradable PLGA coating containing inulin-packaged

DNase I on bacterial adhesion and biofilm formation. We demonstrated that coating of titanium drastically

decreases initial bacterial adhesion and biofilm formation in vitro, without affecting the ability of tissue

cells to adhere. By packaging DNase I in inulin, DNase I underwent less damage during the coating process

and activity was not lost during exposure to several conditions aimed to mimic handling of an implant

before surgery.

Infections associated with biomaterial implant and devices are accompanied by a large use of antibiotics,

mostly in vain, and therewith contribute to the development of antibiotic resistant strains and species.

Underlined by a recently published first report of the World Health Organization on antibiotic resistance

[45], the rise of antibiotic resistant bacterial strains and the limited number of newly discovered antibiotics

indicate the importance of finding alternatives to battle biomaterial associated infections. Our

biodegradable PLGA coating containing inulin-packaged DNase I provides such an alternative, that is not

based on antibiotics but rather on the disruption of the integrity of the EPS matrix in which biofilm

organisms find shelter.

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CHAPTER 5

NORMALLY-ORIENTED ADHESION VERSUS FRICTION FORCES IN

BACTERIAL ADHESION TO POLYMER-BRUSH FUNCTIONALIZED

SURFACES UNDER FLUID FLOW

Jan J.T.M. Swartjes, Deepak H. Veeregowda, Henny C. van der Mei,

Henk J. Busscher and Prashant K. Sharma,

Adv. Funct. Mater. 2014, 24, 4435–4441

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94

ABSTRACT

Bacterial adhesion is problematic in many diverse applications. Coatings of hydrophilic polymer chains in

a brush configuration reduce bacterial adhesion by orders of magnitude, but not to zero. Here we advance

our understanding of the mechanism by which polymer-brush functionalized surfaces reduce bacterial

adhesion from a flowing carrier fluid by relating bacterial adhesion with normally-oriented adhesion and

friction forces on polymer (PEG)-brush coatings of different softness. Softer brush coatings deform more

than rigid ones, which yields extensive bond-maturation and strong, normally-oriented adhesion forces,

accompanied by irreversible adhesion of bacteria. On rigid brushes, normally-oriented adhesion forces

remain small, allowing desorption and accordingly lower numbers of adhering bacteria result. Friction

forces, generated by fluid flow and normally-oriented adhesion forces, are required to oppose fluid shear

forces and cause immobile adhesion. Summarizing, inclusion of friction forces and substratum softness,

provides a more complete mechanism of bacterial adhesion from flowing carrier fluids than available

hitherto.

NORMALLY-ORIENTED ADHESION VERSUS FRICTION FORCES

95

INTRODUCTION

Bacteria have a strong preference to adhere to and colonize surfaces by forming biofilm communities

rather than remaining in a planktonic state [1,2]. Bacteria can adhere to almost any surface, which

represents a major problem in both industrial and biomedical applications [3,4] In biomedical applications,

biomaterial-associated infections caused by biofilms are persistent and hard to eradicate due to their high

resistance to antimicrobials [5,6]. In industrial settings, like pipelines, water treatment plants, heat

exchangers, food processing equipment and on ship hulls, biofilms are associated with public health risks,

lower heat transfer efficiencies and high fuel costs, respectively.

Several strategies to prevent bacterial adhesion and biofilm formation have been described, either by

simply changing the charge or hydrophobicity of the substratum surface or by more complex chemical

modifications of the interface between bacteria and substratum surfaces [7-10]. Surface modification by

means of attaching hydrophilic polymer chains in a brush configuration has been highly successful in

decreasing adhesion of different bacterial strains and species by forming an osmotically driven, steric

barrier to which bacteria have difficulties adhering [11,12]. Polyethylene glycol (PEG) is one of the most

commonly applied polymers for creating polymer-brush coatings and these brush coatings have been

shown to reduce protein adsorption and decrease bacterial adhesion by at least one order of magnitude.

Although no other non-adhesive coating is able to reduce bacterial adhesion by an order of magnitude,

this is still by far insufficient to prevent biofilm formation and biofilms eventually form also on polymer-

brush coatings [13,14]. The mechanisms through which polymer-brushes reduce bacterial adhesion have

been related to brush density and structure, resulting in strongly reduced adhesion forces between a

substratum surface and a bacterium [15-18]. Attachment of polymer-brushes to a surface can be achieved

by various means, like for instance adsorption mediated by hydrophobic interaction of tri-block-

copolymers, or covalent grafting [19]. So far, key factor in creating a polymer-brush, effective in preventing

protein adsorption and bacterial adhesion seems to be the structure of the brush [15]. However, further

understanding of the influence of polymer-brush properties on bacterial adhesion is required in order to

develop polymer-brush coatings with amplified reductions in bacterial adhesion.

In many industrial and bio-medical applications, bacteria approach a surface from a flowing carrier fluid

attracted by non-specific, DLVO-type forces, acting in a direction normal to the substratum surface (Figure

1). Upon close approach of a bacterium to a surface, the bond between a substratum and an adhering

bacterium may mature over time and the normally-oriented adhesion force Fadh becomes stronger

through re-arrangements within the interface [20]. The influence of fluid flow on bacterial mass transport

and adhesion to surfaces has been amply studied, but only tells part of the story [21]. The existence of a

normally-oriented adhesion force in combination with convective fluid flow parallel to a surface will lead

to friction forces, Ff, (Figure 1) between a bacterium and a surface that act perpendicularly to the normally-

oriented adhesion forces, i.e. laterally to the substratum surface. The influence of normally-oriented

adhesion versus friction forces has never been considered in bacterial adhesion in general and the

CHAPTER 5

96

reduction of bacterial adhesion caused by polymer-brush coatings in particular. Normally-oriented

adhesion forces between bacteria and surfaces can be measured with atomic force microscopy (AFM) by

equipping the cantilever with a bacterial probe [22]. Similarly, using the AFM in its lateral mode, friction

forces can be measured, but to our knowledge this has never been done with a bacterial probe on a

material surface [23].

FIGURE 1 Forces experienced by a bacterium approaching a surface from a flowing carrier fluid. Fadh represents the normally-oriented adhesion force between the substratum surface and a bacterium, whereas Ff represents the friction force.

The aim of this paper is to advance our understanding of the mechanisms by which polymer-brush

functionalized surfaces reduce bacterial adhesion, or stated otherwise, the mechanisms by which bacteria

still manage to adhere to a brush coating despite the existing, osmotically driven, steric barrier. To this

end, we determine both the normally-oriented adhesion and friction forces between bacteria and two

distinct, self-assembling-monolayers of PEG, applied at different concentrations. Normally-oriented

adhesion and friction forces will be determined using AFM in its normal and lateral mode for a

Staphylococcus epidermidis strain, while the softness of the polymer-brush coatings will be characterized

using a quartz crystal microbalance with dissipation monitoring (QCM-D). All data will be related with

staphylococcal adhesion to the polymer-brush coatings under convective-diffusional mass transport

conditions.

RESULTS

Softness of polymer-brush coatings

The degree to which steric hindrance can contribute to the reduction of bacterial adhesion by polymer-

brush coatings will depend in part on the softness of the brush coating. Softness can be inferred from the

FadhFf

Convective flow


97

ratio of the change in dissipation (D) and frequency (f) of a gold-coated crystal sensor in QCM-D upon

attachment of a polymer-brush to the crystal surface [24]. The softness of a polymer-brush, as expressed

by the ratio (-ΔD/Δf) varies differently with PEG-concentration for maleimide-PEG (MPEG) and PEG-thiol

(TPEG; see Figure 2). Softness decreases with increasing concentrations of MPEG during brush formation

and increases with increasing concentrations of TPEG, until a similar softness is observed for MPEG and

TPEG at 80 mM. We speculate that the large differences in softness with varying concentrations of MPEG

are caused by the bulky nature of its maleimide coupling group and the accompanying slower reaction

rate as compared to TPEG (see Figures 2a and 2b), which couples directly at a much faster rate, leading to

much smaller differences in softness over the range of applied concentrations.

FIGURE 2 (a) Example of the changes in frequency (Δf3) as measured by the QCM-D for 0.2 mM TPEG and 0.2 mM MPEG. Arrows indicate start of rinsing, after frequency reached stable values. (b) Example of the changes in dissipation (-ΔD3) as measured by the QCM-D for 0.2 mM TPEG and 0.2 mM MPEG. Arrows indicate start of rinsing, after dissipation reached stable values. (c) Softness, expressed as the ratio (-ΔD/Δf) obtained using QCM-D, of two different polymer-brush coatings adsorbed onto gold-coated crystal sensors from solutions with different concentrations of PEG. Smaller values of (-ΔD/Δf) point to a more rigid brush structure. Error bars represent standard deviations over three experiments. *indicates a significant difference (p < 0.05) between MPEG and TPEG polymer-brushes from solutions with the same concentration.

Normally-oriented adhesion forces between bacteria and different polymer-brush coatings

Normally-oriented adhesion forces between staphylococci and the different polymer-brush coatings as

applied in QCM-D, were measured by approaching the coating with a staphylococcal probe until a force of

0

0.4

0.8

1.2

1.6

2

2.4

0.2 2 20 80

-ΔD

/Δf

(10

-7)

PEG concentration (mM)

TPEG

MPEG

0.0

1.5

3.0

4.5

6.0

7.5

0 2000 4000 6000 8000

Δf 3

(Hz)

Time (s)

0

15

30

45

60

75

0 2000 4000 6000 8000

-ΔD

3(1

0-7

)

Time (s)

a

c

b

*

**

CHAPTER 5

98

5 nN was reached and subsequently retracting the probe. The maximal force recorded during retract was

taken as the normal adhesion force. In order to account for bond-maturation, both the initial adhesion

force and the adhesion force after 30 s of contact were measured (see Figure 3a). Initial normal adhesion

forces were very low (0.2 to 0.5 nN) and similar across both types of PEG coatings and all concentrations

applied (see Figure 3b). However, adhesion forces after 30 s of bond-maturation decreased with increasing

concentrations of MPEG and increased with increasing concentrations of TPEG until an adhesion force of

1.5 nN was reached at 80 mM. The differences in softness between TPEG and MPEG brushes were only

large enough to cause significant differences in adhesion forces after 30 s bond-maturation at the two

lowest concentrations applied (0.2 and 2 mM).

Figure 3 (a) Example of a force-distance curve between S. epidermidis ATCC 35983 and a MPEG coating on a gold-coated wafer surface applied from a 0.2 mM solution of MPEG, taken immediately upon contact and after 30 s of bond-maturation. Arrows indicate the distances at which forces reached their maximal values upon retract. These maximal forces were taken as the normally-oriented adhesion forces presented in Figure 3b. (b) Normally-oriented adhesion forces (Fadh) between S. epidermidis ATCC 35983 and TPEG and MPEG polymer-brush coatings prepared from PEG-solutions of different concentrations. Forces were recorded immediately upon contact between the staphylococcal probe and the coating or after 30 s of bond-maturation. Error bars represent standard errors of the mean over at least twelve force-distance curves, comprising three separately prepared coatings and bacterial probes. *indicates a significant difference (p < 0.05) between 0 s and 30 s adhesion forces on polymer-brush surfaces from solutions of the same concentration MPEG or TPEG. #indicates a significant difference (p < 0.05) between adhesion forces with the same bond-maturation time on polymer-brush surfaces of TPEG and MPEG from solutions of the same concentration.

Friction forces between bacteria and polymer-brush coatings

Immediately after taking force-distance curves to confirm successful attachment of bacteria to the AFM

cantilever, friction forces were measured using the AFM at different applied normal forces. Friction forces

were obtained upon both increasing and decreasing the normal force from 0 to 10 nN and vice versa.

Coefficients of friction (COF) between the staphylococci and the polymer-brush coatings were calculated

from the ratio between the measured friction force and the normal force applied. In general, friction forces

were similar upon increasing or decreasing the applied normal force (Figure 4a). Interestingly, COFs

decreased as a function of the normal force applied in a non-linear fashion, as shown in Figure 4a. Since

on hard surfaces, the COF is independent of the normal force applied, this attests to the soft nature of the

-5-4-3-2-1012345

0 100 200 300 400 500 600

Forc

e (

nN

)

Distance (nm)

Initial adhesion force

30 s bond-maturation

Approach

A B

0

0.5

1

1.5

2

2.5

3

3.5

4

4.5

0.2 2 20 80

Ad

he

sio

n f

orc

e (

nN

)


TPEG 0s TPEG 30s

MPEG 0s MPEG 30s

*

#*

#*

*#

*

**

*

#


99

polymer-brush coatings. In order to deal with this effect, COFs were averaged over the values obtained for

increasing and decreasing normal forces and plotted as a function of the normal forces applied on a double

log-scale (Figure 4b).

Friction forces at a 1 nN applied normal force, i.e. within the range of normally-oriented adhesion forces

between the staphylococci and the polymer-brush coatings, are presented for TPEG and MPEG polymer-

brush coatings prepared from PEG-solutions of different concentrations in Figure 4c, showing a decrease

in friction force with increasing MPEG concentrations in solution and little or no effect of the concentration

of TPEG in solution as used for preparation of a coating.

FIGURE 4 (a) Example of the coefficient of friction (COF) as a function of normal force applied upon increasing and decreasing the normal forces for a MPEG polymer-brush coating applied from a solution of 0.2 mM MPEG. COF were calculated from the ratio between the measured friction force and the normal force applied. (b) Double log-scale representation of the average COF upon increasing and decreasing the normal force (see Figure 4a), including results of a linear regression analysis (R2 = 0.70 for the example given). (c) Friction forces at a 1 nN normal force derived from linear regression analyses as shown in Figure 4b for TPEG and MPEG polymer-brush coatings prepared from PEG-solutions of different concentrations. Error bars represent standard errors of the mean over nine measurements, comprising three separately prepared coatings and bacterial probes.

Bacterial adhesion numbers and polymer-brush characteristics

The numbers of staphylococci adhering to the different polymer-brush coatings were determined under

convective-diffusional mass transport from a flowing carrier fluid (wall shear rate 10 s-1). Adhesion

a

-1.5

-1

-0.5

0

0.5

1

1.5

-0.4 -0.2 0 0.2 0.4 0.6 0.8 1 1.2

Log

(CO

F)

Log (normal force (nN))

b

c

0

1

2

3

4

5

0 2 4 6 8 10

Co

effi

cie

nt

of

fric

tio

n

Normal force (nN)

Increasing normal force

Decreasing normal force

0

1

2

3

4

5

0.2 2 20 80

Fric

tio

n fo

rce

at

1 n

Nn

orm

al f

orc

e (

nN

)


TPEG

MPEG

CHAPTER 5

100

experiments were continued until stable adhesion numbers over time were obtained, which usually

occurred within 2 h. The numbers of staphylococci adhering after 2 h to the different polymer-brush

coatings are summarized in Figure 5 as a function of the friction force (see Figure 5a), the normally-

oriented adhesion force (see Figure 5b) and the softness of the polymer-brush coating (see Figure 5c).

Staphylococcal adhesion numbers increased with all three brush characteristics, with the strongest

correlations existing for friction force and softness.

FIGURE 5 (a) Friction forces between staphylococci and polymer-brush coatings at an applied normal force of 1 nN, as a function of bacterial adhesion numbers after 2 h of exposure of the coatings to a flowing staphylococcal suspension. (b) Normally-oriented adhesion forces after 30 s bond-maturation as a function of bacterial adhesion numbers after 2 h of exposure of the coatings to a flowing staphylococcal suspension. (c) Softness of the polymer-brush, expressed as the ratio (-ΔD/Δf) in QCM-D, as a function of bacterial adhesion numbers after 2 h of exposure of the coatings to a flowing staphylococcal suspension. R2 values assuming a linear correlation are given in each graph, while the dotted lines represent the 95% confidence intervals.

DISCUSSION

In this paper, we determined the normally-oriented adhesion and friction forces between bacteria and

two distinct, self-assembling-monolayers of PEG, and related these forces and the softness of the polymer-

brush coatings with the number of staphylococci adhering under convective-diffusional mass transport

R² = 0.92

0.4

0.8

1.2

1.6

2.0

0 2 4 6 8 10

-ΔD

/Δf

( x1

0-7

)

Bacterial adhesion (x 106 cm-2)

TPEG

MPEG

a

b

c

R² = 0.90

0

1

2

3

4

5

0 2 4 6 8 10

Fric

tio

n fo

rce

(n

N)

R² = 0.78

0

1

2

3

4

0 2 4 6 8 10

30

s ad

he

sio

n fo

rce

(n

N) R² = 0.92

0.4

0.8

1.2

1.6

2.0

0 2 4 6 8 10

-ΔD

/Δf

( x1

0-7

)


TPEG

MPEG

a

b

c

R² = 0.90

0

1

2

3

4

5

0 2 4 6 8 10

Fric

tio

n fo

rce

(n

N)

R² = 0.78

0

1

2

3

4

0 2 4 6 8 10

30

s ad

he

sio

n fo

rce

(n

N)

R² = 0.92

0.4

0.8

1.2

1.6

2.0

0 2 4 6 8 10

-ΔD

/Δf

( x1

0-7

)


TPEG

MPEG

a

b

c

R² = 0.90

0

1

2

3

4

5

0 2 4 6 8 10

Fric

tio

n fo

rce

(n

N)

R² = 0.78

0

1

2

3

4

0 2 4 6 8 10

30

s ad

he

sio

n fo

rce

(n

N)

R² = 0.92

0.4

0.8

1.2

1.6

2.0

0 2 4 6 8 10

-ΔD

/Δf

( x1

0-7

)


TPEG

MPEG

a

b

c

R² = 0.90

0

1

2

3

4

5

0 2 4 6 8 10

Fric

tio

n fo

rce

(n

N)

R² = 0.78

0

1

2

3

4

0 2 4 6 8 10

30

s a

dh

esi

on

forc

e (

nN

)

R² = 0.92

0.4

0.8

1.2

1.6

2.0

0 2 4 6 8 10

-ΔD

/Δf

( x1

0-7

)


TPEG

MPEG

a

b

c

R² = 0.90

0

1

2

3

4

5

0 2 4 6 8 10

Fric

tio

n fo

rce

(nN

)

R² = 0.78

0

1

2

3

4

0 2 4 6 8 10

30

s ad

hes

ion

forc

e (n

N)


101

conditions. Hitherto, studies on mechanisms of microbial adhesion have focused on normally-oriented

adhesion forces. However, normally-oriented forces can only explain why a bacterium adheres and keeps

a certain distance from a substratum surface, but cannot explain why it becomes immobilized under fluid

flow, which requires forces operating laterally to a substratum surface [25]. When laterally moving

bacteria encounter a surface, friction forces will arise, as demonstrated for the first time in this paper.

Such friction forces can considerably slow-down bacteria in a flowing carrier fluid, to facilitate their

immobile adhesion despite low, normally-oriented adhesion forces [26].

Bacteria suspended in a flowing carrier fluid adapt the velocity of the fluid, which depends on their height

above the substratum surface. According to Poiseuille’s law for a PPFC

𝑣(𝑦) =3

4×

Qy

b2w× (2-

y

b) (1)

in which Q is the volumetric flow rate, y the perpendicular distance from the bottom plate, b the half depth

of the chamber and w its width. Close to the substratum surface, within the diffusion-boundary layer, the

velocity of bacteria parallel to the surface follows from Poiseuille’s law, while both its own weight and

diffusion drive the suspended bacteria closer to the surface to decrease their velocity and bring them

under the influence of the normally-oriented adhesion forces arising from the substratum [27]. Image-

sequence analysis of bacteria approaching a polymer-brush coating from a flowing carrier fluid clearly

shows that once bacteria appear within the focus depth of the microscope system (up to 3 µm above the

substratum surface), they have a velocity of around 20 x 10-6 – 40 x 10-6 m s-1 (Figure 6). According to

Equation 1, this indeed indicates that they are located in fluid flow lines approximately 3 µm above the

surface, far outside the reach of the normally-oriented adhesion forces arising from the substratum, but

within the diffusion-boundary layer [27]. As a consequence, a diffusion-driven approach to the surface

occurs resulting in contact when the velocity has decreased to around 6 x 10-6 m s-1 at 0.5 µm (i.e. half the

staphylococcal diameter) above the surface (see also Figure 6), a phase that has been described as “mobile

adhesion” [25]. Mobile adhesion subsequently transforms into “immobile adhesion” as friction forces

develop laterally to the substratum surface that oppose the shear force acting on the decelerating

bacterium. The shear force acting on a fully immobilized bacterium follows from

𝐹 = 𝜂 × 𝜎 × 𝐴𝑏𝑎𝑐 (2)

in which 𝜂 is the viscosity of the medium, σ is the shear rate and Abac is the cross-sectional area of a

staphylococcus. Under a fluid shear rate of 10 s-1, Equation 2 allows to calculate a shear force acting on

immobilized staphylococci of around 9 x 10-6 nN, which is orders of magnitude smaller than the friction

forces measured in this study. Considering that shear forces arising from lateral convection are much

CHAPTER 5

102

smaller than the measured friction forces, we conclude that the friction forces are large enough to oppose

the fluid shear forces and contribute to immobilization of a bacterium on the surface, needed for

subsequent bond-maturation.

FIGURE 6 Examples of the velocities of two staphylococci in a flowing carrier fluid derived from image-sequence analysis as a function of the time prior to immobilization on a substratum surface modified with a 0.2 mM MPEG polymer-brush.

Once the bacterial velocity has decreased by friction arising from contact with the surface and adhesion

forces have been established, the normally-oriented adhesion forces increase over time and already within

30 s, a considerable maturation of the bond has occurred. The origin of this type of bond-maturation is

physico-chemical in nature, as the time-scale over which it occurs in a nutrient-free suspension will not

allow for metabolic processes such as EPS-production, to have a major impact on the adhesion force [20].

Moreover, similar bond-maturation of normally-oriented adhesion forces has been observed for inert

polystyrene particles. Physico-chemical bond-maturation has been suggested to be due to removal of

interfacial water, re-arrangement of bacterial cell surface structures and conformational changes of cell

surface macromolecules [28].] This type of bond-maturation has been associated with the residence-time

dependent desorption of particles and bacteria from surfaces, based on the fact that particulate

desorption disappears within 2 min, i.e. the same time-scale as over which AFM-measured adhesion forces

mature to a stable value [29].

As a new aspect of this study, we demonstrate that bond-maturation is much more extensive on softer

polymer-brushes than on more rigid ones (see Figure 3B) with important consequences for bacterial

adhesion. Effective polymer-brush coatings must possess a certain degree of rigidity in order to prevent

their deformation under the influence of the normally-oriented adhesion forces between the substratum

surface and a bacterium. Therewith adhesion forces remain relatively small, allowing bacterial desorption

and keeping the numbers of adhering bacteria low (Figure 7) [29]. Soft polymer-brushes on the other hand,

deform upon contact with adhering bacteria, causing more extensive bond-maturation and stronger

normally-oriented adhesion forces than encountered on a more rigid brush. Consequently, bacterial

desorption will be low, and higher numbers of adhering bacteria will develop over time (see also Figure 7).

0

10

20

30

40

50

0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0

Bac

teri

al v

elo

city

(1

0-6m

s-1

)

Time prior to immobilization (s)

0

10

20

30

40

50

0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0

Bac

teri

al v

elo

city

(1

0-6

m s

-1)

Time prior to immobilization (s)


103

In conclusion, we show that friction forces are involved in the adhesion and immobilization of bacteria to

polymer-brush coated surfaces from a flowing carrier fluid. Friction forces are governed by an interplay of

the normally-oriented adhesion forces and the properties of the surface and are required to cause

immobilization of adhering bacteria once they have approached the surface and established contact. On

more rigid polymer-brush coatings, bacteria desorb more readily, while softer polymer-brush coatings

deform upon adhesion causing a strong increase in the normally-oriented adhesion force that impedes

desorption. Summarizing, inclusion of both friction forces and substratum softness provides a more

complete mechanism of bacterial adhesion to polymer-brush coated surfaces than can be obtained

considering only normally-oriented adhesion forces and will help to unveil additional options in the search

for materials properties and coatings that resist bacterial adhesion. Important in this respect, is that bond-

maturation after initial adhesion is avoided, which can be established with more rigid brushes.

FIGURE 7 Schematic presentation of bacteria approaching from a flowing carrier fluid to a substratum surface and their immobilized adhesion on a soft polymer-brush (top panel) and approach to more rigid one (bottom panel), from which a bacterium readily desorbs due to the absence of deformation and extensive bond-maturation.


O-[2-(3-mercaptopropionylamino)ethyl]-O’-methylpolyethyleneglycol 5,000 (TPEG), methoxy-

polyethylene glycol maleimide 5,000 (MPEG), DL-dithiothreitol and octanedithiol were purchased from

Sigma-Aldrich (St. Louis, USA). Ethanol (99.9%), ammonia (28-30%) and hydrogen peroxide (30%) were of

analytical grade and purchased from Merck, Darmstadt, Germany.

Polymer-brush functionalization of surfaces

Gold-coated silica wafers (Litcon, Gothenburg, Sweden) and gold-plated AT-cut quartz crystals (Q-sense,

Gothenburg, Sweden) with a sensitivity constant of 17.7 ng cm-2 and a 5 MHz resonance frequency were

CHAPTER 5

104

cleaned by UV/ozone treatment followed by immersion in a 5:1:1 solution of ultrapure water (18.2 MΩ

cm-1, Sartorius, Göttingen, Germany), H2O2 and NH3 for 10 min at 80 ºC. Mono-functional PEG-thiol was

attached to cleaned substrata by immersing cleaned wafers in a 1 ml solution of PEG-thiol in ultrapure

water for 30 min, or by flow of PEG-thiol solution over gold-plated quartz crystals in the QCM-D. Mono-

layers of octanedithiol, required for the attachment of maleimide conjugated PEG, were formed by

immersion in 20 mM octanedithiol in ethanol and incubation in a dark, oxygen free environment for 20 h.

Next, substrata were washed repeatedly with ethanol, sonicated for 2 min and washed again with ethanol

before being transferred to a solution of 20 mM DL-dithiothreitol in TRIS-HCl 50 mM/EDTA 5 mM pH 8.0

for 2 h to re-activate the dithiol monolayer. The entire cycle of exposure to octanedithiol and DL-

dithiothreitol was repeated once to obtain a homogenous monolayer [30]. Substrata were then

functionalized by exposure to MPEG solution (sodium hydrogen phosphate 100 mM, EDTA 5 mM at pH

6.5).

Softness of Polymer-brush Coating: QCM-D (model Q-sense E4, Q-sense Gothenburg, Sweden) was used

to measure the softness of the polymer-brush coatings. Polymer-brush coatings were applied by flowing

a PEG solution through the QCM-D chamber at a flow rate of 2 ml h-1. The QCM-D chamber is 14 mm in

diameter and has a volume of approximately 40 μl. Flow with PEG solution was continued until stable

values of frequency (f) and dissipation (D) where observed, which generally lasted 30 min when flowing

TPEG and 120 min when MPEG was used. As soon as stable values were obtained, the crystals were washed

by flow of the appropriate buffer, in order to remove unbound PEG molecules. ΔD and Δf values obtained

after washing were used to evaluate the softness of the brush layer by calculating the -ΔD/Δf ratio, higher

ratios representing softer brushes (higher dissipation shifts results from a softer brush dissipating more

energy upon vibration of the crystal in an aqueous environment than in case of a more rigid brush) [24].

Normally-oriented adhesion and friction forces between staphylococci and polymer-brush coatings

Staphylococci were grown overnight on blood agar plates at 37 ºC for 24 h from frozen stock. Plates were

kept at 4 ºC and stored no longer than two weeks. Pre-cultures were prepared by inoculating single

colonies from the blood agar plate into 10 ml tryptone soya broth (TSB, OXOID, Basingstoke, England) and

incubated at 37 ºC for 24 h. This pre-culture was used to inoculate a main-culture of 200 ml TSB, incubated

for 17 h at 37 ºC. Bacteria were harvested by centrifugation for 5 min at 3000 g and washed twice using

phosphate buffered saline (PBS, 150 mM NaCl, 10 mM potassium phosphate, pH 6.8). Bacterial suspension

was sonicated for 3 x 10 s (Vibra Cell model 375; Sonics and Materials, USA), with 30 s breaks in between,

on an ice/water bath to break up bacterial aggregates.

For AFM, S. epidermidis ATCC 35983 was immobilized on tipless cantilevers (MikroMasch, Sofia, Bulgaria)

via electrostatic interaction with poly-L-lysine (PLL; Sigma-Aldrich, USA) using a micromanipulator

(manufactured by Narishige Groups, Tokyo, Japan). To this end, a droplet of PLL solution was placed on a

glass slide and the far end part of the cantilever was dipped in the droplet for 1 min. After air drying (2

min), the cantilever was dipped in a droplet of a S. epidermidis ATCC 35983 suspension for 1 min to allow


105

bacterial attachment to the cantilever. Bacterial probes were used for AFM immediately after preparation.

Coefficients of friction and normally-oriented adhesion forces towards bacteria were measured with an

AFM (Nanoscope IV DimensionTM 3100) equipped with a Dimension Hybrid XYZ SPM scanner head (Veeco,

NY, USA). Rectangular tipless cantilevers (length (l), width (w), and thickness (t) of 300, 35, and 1 μm,

respectively) with an average stiffness of 0.05 Nm−1 were calibrated for their exact torsional and normal

stiffness using AFM Tune IT v2.5 software [31-33]. The normal stiffness (Kn) was in the range of 0.02 to

0.08 Nm−1, while the torsional stiffness (Kt) was in the range of 2 to 6 × 10−9 Nm rad−1. The deflection

sensitivity (α) of the cantilever was recorded on bare glass to calculate the applied normal force (Fn) using

Fn = ∆Vn x α x Kn (3)

where ΔVn is the voltage output from the AFM photodiode due to normal deflection. The torsional stiffness

and estimated geometrical parameters of the bacterial probe were used to calculate the friction force (Ff)

according to

𝐹f = ∆VL x Kt

2δ (d+t2) (4)

where t is the thickness of the cantilever, δ is the torsional detector sensitivity of the AFM and ΔVL

corresponds to the voltage output from the AFM photodiode due to lateral deflection of the bacterial

probe [32,34].

Normally-oriented adhesion force measurements

In order to verify prior to and in between measurements that a bacterial probe enabled a single contact

with the surface, a scanned image in AFM contact mode with a loading force of 1 to 2 nN was made at the

onset of each experiment and examined for double contour lines. Double contour lines indicate that the

AFM image is not obtained from contact of a single bacterium with the surface but that multiple bacteria

on the probe are in simultaneous contact with the substratum. Such probes were discarded both in the

measurement of normally-oriented adhesion forces as well as in friction force measurements.

Force-distance curves were measured immediately upon contact and after 30 s of bond-maturation at a

scan rate of 1 Hz and a trigger threshold of 5 nN. The adhesion force was calculated according to

F = Kn × D (5)

where D is the deflection of the cantilever. Force–distance curves were measured on three different spots,

before and after each friction measurement to verify that the bacteria were still attached to the probe,

i.e. if a hard contact was observed upon approach, the probe and the measurement were discarded.

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106

Friction force measurements

Lateral deflection was observed at a scanning angle of 90 degrees over a distance of 5 μm and a scanning

frequency of 1 Hz. The scanning angle, distance, and frequency were kept constant throughout all friction

force measurements. The bacterial probe was incrementally loaded and unloaded in steps of 0.5 nN up to

a maximal normal force of 10 nN. Negative friction values, resulting from operating the AFM near its lower

limits, were taken as zero [34]. COF was measured on three different spots on each substratum surface

and separate bacterial probes were used for measurements on at least three separately prepared

substratum surfaces. The coefficient of friction (COF) was calculated for each applied normal force rather

than from the slope in the friction versus load curves, because on soft polymer brushes the COF depended

in a non-linear way on the applied normal force, making it impossible to use the slope for obtaining the

COF.

Staphylococcal adhesion from a flowing carrier fluid

Adhesion of S. epidermidis ATCC 35983 to polymer-brush coatings on the gold-coated wafers was studied

using a parallel plate flow chamber, using a staphylococcal suspension with 3x108 bacteria ml-1 [35]. Prior

to each experiment, the flow system was perfused with PBS to remove all air bubbles, after which the

staphylococcal suspension was flown through the chamber at a constant shear rate (10 s-1) for 2 h.

Bacterial deposition was monitored real-time using a CCD-MRXi camera (Hight Technology, Eindhoven,

The Netherlands) mounted on top of a metallurgical microscope (Olympus BH-2) with a x40 long working

distance objective (depth of view 3 µm) and images were taken 2 h after flow, from which the number of

adhering bacteria was determined.


QCM-D data followed a normal distribution (Shapiro-Wilk test, p < 0.05) and differences between

polymer-brush coatings from solutions of TPEG and MPEG of the same concentration were analyzed using

Student’s t-test. Adhesion forces were not normally distributed (Shapiro-Wilk, p < 0.05), and the non-

parametric Kolmogorov–Smirnov test was used to compare adhesion forces. Differences were considered

significant if p < 0.05.

ACKNOWLEDGEMENTS

This study was entirely funded by the University Medical Center Groningen, Groningen, The Netherlands.

D.H. Veeregowda is also manager of Ducom Instruments Europe (Center for Innovation, L.J. Zielstraweg 2,

9713 GX, Groningen, The Netherlands), while H.J. Busscher is also director of a consulting company, SASA

BV (GN Schutterlaan 4, 9797 PC Thesinge, The Netherlands). The authors declare no potential conflicts of

interest with respect to authorship and/or publication of this article. Opinions and assertions contained

herein are those of the authors and are not construed as necessarily representing views of the funding

organization or their respective employers.


107

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CHAPTER 6

NORMAL AND LATERAL ADHESION FORCES MEDIATED BY ANTIGEN I/II

BINDING OF S. MUTANS TO SALIVARY FILMS

Jan J.T.M. Swartjes, Yun Chen, Joop de Vries, Prashant K. Sharma,

Henny C. van der Mei, Henk J. Busscher, and Deepak H. Veeregowda

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ABSTRACT

Streptococcus mutans is a cariogenic oral bacterial strain, adhering to salivary-conditioning-films (SCFs) on

teeth through both specific and non-specific interactions. Specific adhesion of S. mutans is mediated by a

surface-localized adhesin, called antigen I/II, interacting with salivary-agglutinins, composed of

glycoprotein complexes in the SCF. Here we use atomic-force-microscopy to compare the normal and

lateral adhesion forces between SCFs and two isogenic S. mutans strains, with (LT11) and without

(IB03987) antigen I/II.. First, normally-oriented adhesion forces between the streptococci and the different

substrata were measured, after which lateral forces were measured at an applied normal force equal to

the adhesion force arising from the substratum. The differences between normal and lateral adhesion

force values are due to direction dependent differences in the adhesion mechanism of bacteria. The

combination of normal and lateral force microscopy as applied here to reveal differences between specific

and non-specific adhesion mechanisms of bacteria to surfaces is new, and applicable in all industrial,

environmental and biomedical applications where bacteria adhere under fluid flow through a combination

of non-specific and specific adhesion mechanisms involving localized, high affinity sites.

NORMAL AND LATERAL ADHESION FORCES OF S. MUTANS

111

INTRODUCTION

Streptococcus mutans is a highly cariogenic oral bacterial strain, known to adhere to salivary conditioning

films (SCF) on teeth through both specific and non-specific adhesion mechanisms. Specific adhesion of S.

mutans is mediated by a cell surface-localized adhesin, called antigen I/II [1,2]. Antigen I/II is a stalk-like

structure with a length of 60 nm that extends from the bacterial cell surface of S. mutans and possesses a

hydrophobic end. Salivary agglutinin, a glycoprotein complex secreted by the salivary glands, forms the

human receptor for antigen I/II in SCFs [2]. Non-specific adhesion of S. mutans occurs through attractive,

long-range Lifshitz-Van der Waals forces as arising from all molecules constituting an organism and the

substratum, be they located at the surface or inside, while at short range non-specific adhesion is through

electrostatic and acid-base interactions [3]. As an important difference between specific and non-specific

adhesion forces, specific adhesion originates from localized high affinity ligand-receptor groups on the

interacting surfaces [4].

The forces by which S. mutans adheres to salivary conditioning films have been measured using bacterial

probe atomic force microscopy (AFM) through contacting the SCF-coated surface with a bacterial probe

and subsequent retraction to reveal the normal adhesion force [5,6]. Although bacterial probe AFM, or

also called single-cell force spectroscopy [7], is more complex and less reproducible than more traditional

ways of using AFM, it represents the only way to make a force analysis of the complex interaction of

bacterial cell surface structures with a substratum surface. Using bacterial probe microscopy, initial

colonizers of dental hard surfaces presented stronger adhesion forces to dental restorative materials

covered with a SCF (4.7 to 0.6 nN) than cariogenic strains, including S. mutans (1.8 and 0.5 nN) [5].

Adhesion forces of a S. mutans strain to SCF-coated enamel surfaces amounted 0.8 nN [6]. Non-specific

adhesion forces between oral bacteria and bovine serum albumin (BSA) coatings strengthened twofold

faster than their specific interactions with salivary conditioning films, likely because specific interactions

require a closer approach of the interacting surfaces with the removal of interfacial water and a more

extensive rearrangement of surface structures [8]. After bond-strengthening, bacterial adhesion forces

with a SCF-coated surface remained stronger (1.2 to 0.4 nN) than those with BSA coatings (0.3 to 0.2 nN).

Mutually perpendicular forces are known to act independently, which raises the question how organisms

in the oral cavity deal with the presence of shear forces, arising from fluid flow, movement of the tongue

and mastication. Withstanding shear forces requires laterally-acting forces, which need not necessarily be

the same as the normally-oriented adhesion forces. Lateral adhesion forces can be measured using the

AFM in its lateral mode [9], taking the force required for lateral movement after contact of the probe with

a surface as the lateral adhesion force. In addition, a delay in lateral movement after contact can be applied

to determine the lateral adhesion force after bond-strengthening.

Lateral force microscopy is relatively new and seldom applied using a bacterial probe. Here we compare

normally-oriented and lateral forces by which S. mutans LT11 (with antigen I/II) and S. mutans IB03987

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112

(without antigen I/II) adhere to glass surfaces and SCF-coated glass to yield a better understanding of the

mechanisms of S. mutans adhesion to salivary films.


Bacterial cultures

S. mutans was stored in 7% DMSO at -80°C, plated onto brain heart infusion (BHI, OXOID, Basingstoke, UK)

containing 1.5% agar and incubated overnight at 37°C in 5% CO2. A single colony was used to inoculate 0.5

mL BHI pre-culture without (LT11) or with (IB03987) 500 μg/mL kanamycin (OXOID) and incubated for 24

h. This pre-culture was used to inoculate a 10 mL main culture in BHI, which was grown for 16 h. Bacteria

were harvested by centrifugation at 5000 g for 5 min at 10°C and washed twice with 10 mM potassium

phosphate, pH 7. Bacterial suspension was sonicated for 3 x 10 s (Vibra Cell model 375; Sonics and

Materials, USA), with 30 s breaks in between, on an ice/water bath to break up bacterial aggregates.

AFM bacterial probe preparation

Bacterial probes were prepared by immobilizing a bacterium onto a tipless cantilever (MikroMasch, Sofia,

Bulgaria). The cantilever was mounted to the end of a micromanipulator and, under microscopic

observation, the tip of the cantilever was dipped into a droplet of 0.01% α-poly-L-lysine with a molecular

weight of 70,000 to 150,000 (Sigma-Aldrich) for 1 min to create a positively charged layer. After 2 min of

air drying, the tip of the cantilever was carefully dipped into a streptococcal suspension droplet for 1 min

to allow bacterial attachment through electrostatic attraction and dried in air for 2 min. Successful

attachment of streptococci on the cantilever follows directly from a comparison of the adhesion force-

distance curves of a streptococcal probe and the one of a poly-L-lysine-coated cantilever in absence of

streptococci. In order to verify that a bacterial probe had a single contact with the substratum surface, a

scanned image in the AFM contact mode with a loading force of 1 to 2 nN was made at the onset of each

experiment and examined for double contour lines. Double contour lines indicate that the AFM image was

not from the contact of a single bacterium with the surface but that multiple streptococci on the probe

were in simultaneous contact with the substratum. Any probe exhibiting double contour lines was

discarded. Bacterial probes were used for AFM immediately after preparation.

Normally- and laterally-oriented adhesion forces for both streptococcal strains were measured with an

AFM (Nanoscope IV DimensionTM 3100) equipped with a Dimension Hybrid XYZ SPM scanner head (Veeco,

NY, USA). Rectangular tipless cantilevers (length (l), width (w), and thickness (t) of 300, 35, and 1 μm,

respectively) were calibrated for their exact torsional and normal stiffness using AFM Tune IT v2.5 software

[10–12]. The normal stiffness (Kn) was in the range of 0.03 to 0.12 Nm−1, while the torsional stiffness (Kt)

was in the range of 2 to 6 × 10−9 Nm rad−1. The deflection sensitivity (α) of the cantilever was recorded on

bare glass to calculate the applied normal force (Fn) using


113

Fn = ∆Vn x 𝛼 x Kn (1)

where ΔVn is the voltage output from the AFM photodiode due to normal deflection. The torsional stiffness

and estimated geometrical parameters of the bacterial probe were used to calculate the lateral force (F)

according to

F = ∆VL x Kt

2𝛿 (d + t2)

(2)

where t is the thickness of the cantilever, δ is the torsional detector sensitivity of the AFM, d is the diameter

of bacterial probe (1 µm, for a single streptococcus) and ΔVL corresponds to the voltage output from the

AFM photodiode due to lateral deflection of the bacterial probe [11,13].

Normally-oriented adhesion forces

Force-distance curves were measured immediately upon contact and after 30 s of bond-strengthening at

a scan rate of 1 Hz and a trigger threshold of 1 nN. The adhesion force was calculated according to

F = Kn × D (3)

where D is the deflection of the cantilever.

Laterally-oriented forces

Lateral deflection was observed at a scanning angle of 90 degrees over a distance of 5 μm, at a scanning

frequency of 1 Hz and at an applied normal force equal to the normal adhesion force measured. Lateral

forces follow from the force required to move a bacterium along the surface and were measured on two

different spots on each substratum surface with three separately prepared bacterial probes and at least

three separately prepared substratum surfaces. Lateral forces have been measured immediately after

contact and after 30 s of bond-strengthening and represent averages of lateral force measurement taken

over a scanning length of 80 µm (i.e. 16 scan lines of 5 µm each) in order to obtain reproducible values.


Retraction and lateral force data did not follow a normal distribution (Shapiro-Wilk test, p < 0.05) and

hence differences were analyzed using Mann-Whitney U test. Differences were considered significant if p

< 0.05.

RESULTS

Normally-oriented adhesion forces follow from retract force-distance curves measured by AFM (see Fig. 1

A for schematics of the method), of which examples are presented in Fig. 1 B. The rupture path in the

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114

force-distance curves of S. mutans LT11 (possessing antigen I/II) started at smaller distances than of S.

mutans IB03987 (lacking antigen I/II), while also its adhesion forces on glass were higher than of S. mutans

IB03987 (Fig. 1 C). This was regardless of whether the streptococci were allowed to contact the substratum

surface for a longer period of time (30 s bond-strengthening) or whether the bacteria were retracted

immediately after contact (0 s bond-strengthening). Normally-oriented adhesion forces were extremely

small on SCF-coated glass surfaces immediately upon contact and were below detection for S. mutans

IB03987, while for S. mutans LT11 the normal adhesion force amounted 0.2 ± 0.1 nN (Fig. 1 D). After 30 s

of bond-strengthening, normally-oriented adhesion forces increased slightly for S. mutans IB03987 to 0.3

± 0.2 nN. The normal adhesion force of S. mutans LT11 however, increased to 1.3 ± 0.5 nN after bond-

strengthening, significantly exceeding the one of S. mutans IB03987.

FIGURE 1 Normally-oriented adhesion forces of S. mutans strains with and without antigen I/II. (A) Schematic presentation of normal adhesion force measurements by AFM. (B) Example of retract force-distance curves for S. mutans LT11 and S. mutans IB03987 on glass, from which adhesion forces are calculated. (C) Average adhesion forces to glass for S. mutans LT11 and S. mutans IB03987. Error bars represent the standard deviations over 15 force-distance curves, measured with six different probes, prepared from three separately grown bacterial cultures. (D) As in (C), now for a SCF-coated glass surface. * Indicates significant differences (p < 0.05) in adhesion forces between

0

1

2

3

4

5

6

7

LT11 IB03987

Forc

e (

nN

)

0

1

2

3

4

5

6

7

LT11 IB03987

Forc

e (

nN

)

-3

-2

-1

0

1

2

3

4

5

6

0 50 100 150 200 250 300

Ret

ract

ion

forc

e (

nN

)

Tip separation (μm)

S. mutans LT11S. mutans IB 03987

0 s bond-strengthening

30 s bond-strengthening

Glass SCF-coated glass

A

D

* **#

C

laserphotodetector

cantilever

B


115

S. mutans LT11 and IB03987. # Indicates significant differences (p < 0.05) in adhesion forces between 0 and 30 s bond-strengthening.

Lateral force microscopy (see Fig. 2 A for schematics of lateral force AFM) generates the lateral force at

an applied normal force. The laterally-oriented adhesion forces follow from the force required to move S.

mutans along the surface, at an applied normal force equal to the normally-oriented adhesion forces

measured (see Figs. 1 C and D). Lateral adhesion forces for S. mutans LT11 and IB03987 on glass after 30

s of contact were 5.2 ± 1.7 and 0.8 ± 0.3 nN (Fig. 2 B) respectively, while on SCF-coated glass they amounted

4.2 ± 0.8 and 1.0 ± 0.6 nN (Fig. 2 C).

FIGURE 2 Normally- and laterally-oriented adhesion forces of S. mutans strains after 30 s bond-strengthening. Lateral adhesion forces were measured at an applied normal force equal to the normally-oriented adhesion forces measured (see Fig. 1). (A) Schematic presentation of lateral force measurements by AFM. Upon lateral movement of the AFM cantilever in contact with the surface, lateral forces cause torsional deformation of the cantilever leading to changes in the lasers deflection detected by the photo detector from which the lateral force is calculated. Note that for clarity the twist of the cantilever has been exaggerated. (B) Normal and lateral adhesion forces for S. mutans LT11 and S. mutans IB03987 on glass. (C) As in (B), now for a SCF-coated glass surface.* indicates significant differences (p < 0.05) in adhesion forces between S. mutans IB03987 and S. mutans LT11. # indicates significant differences (p < 0.05) between normal and lateral adhesion forces.

0

1

2

3

4

5

6

7

LT11 IB03987

Forc

e (

nN

)

0

1

2

3

4

5

6

7

LT11 IB03987

Forc

e (

nN

)

Fadh normal

Fadh lateral

Glass SCF-coated glass

**

*#

*#B C

A laserphotodetector

cantilever

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116

DISCUSSION

Two isogenic strains of streptococci displayed completely different adhesion patterns both when adhesion

forces were measured in an orientation normal to the surface as well as in a direction lateral to the surface.

S. mutans LT11, possessing an antigen I/II based interaction mechanisms with salivary agglutinins in a SCF,

adhered more strongly than S. mutans IB03987 (see Fig. 2). Its lateral adhesion force was approximately

threefold higher than its normal one, both on bare glass and on a SCF. This study was carried out on glass

as a substratum surface rather than on enamel, because enamel surfaces are extremely difficult to polish

to the smoothness required for lateral force measurements while furthermore differences in surface

physico-chemical properties are minor, especially after coating with a SCF [14,15].

Lateral force microscopy as applied here is a derivative of colloidal force microscopy commonly used to

measure friction coefficients of homogeneous surfaces in absence of localized binding phenomena

[16,17], that has to our knowledge not been employed before to this end. For friction measurements, the

colloidal probe is incrementally loaded and unloaded up to a specified maximal normal force, while

measuring the lateral force which is then interpreted as a friction force. By taking the ratio of the friction

force to the applied normal force the friction coefficient can be obtained. We measured lateral forces of

two isogenic strains of streptococci in order to measure the lateral force that caused immobilization of the

organisms under shear, as occurring in the oral cavity. Importantly, this can only be done in combination

with normal adhesion force measurements as the lateral force needs to be determined at an applied

normal force equal to the adhesion forces arising from the substratum material. Since normally-oriented

adhesion forces strengthen with increasing contact times between a bacterium and a substratum surface,

the same may be true for lateral forces. We chose to use a 30 s bond-strengthening time for lateral

adhesion force measurements, as this usually yields stable normal adhesion force values [8]. Moreover,

considering the time scale of bacterial adhesion processes, this seems reasonable [18]. Lateral force

microscopy as applied here, bears not only relevance for bacterial adhesion under the dynamic conditions

of the human oral cavity, but is applicable in all industrial, environmental and biomedical applications

where bacteria adhere through a specific adhesion mechanisms involving localized, high affinity sites.

Strengthening of the bond between adhering bacteria and substratum surfaces is a common phenomenon

of physico-chemical nature and similar bond-strengthening has also been observed for inert polystyrene

particles [11]. Biological processes contributing to strengthening of bacterial binding to substratum

surfaces through excretion of extracellular polymeric substances for instance, occur over a much longer

time scale compared to the physico-chemical processes observed here [12] and generally require a more

nutrient-rich environment [13]. Bond-strengthening is not only observed in the case of non-specific

interactions between adhering bacteria and surfaces, but even more so in the case of specific ligand-

receptor interactions [7,14]. Non-specific bond-strengthening is caused by physico-chemical re-

arrangement of the interface, whereas in case of specific bond-strengthening the development of ligand-

receptor binding through conformational changes of cell surface polymers and optimal positioning,


117

contributes to strengthening of the bond [7]. In the current study, two isogenic S. mutans strains have

been used, that differ in their ability to form specific ligand-receptor bonds between cell surface associated

antigen I/II and salivary agglutinins in the SCF. However, even though one of the strains possessed a

specific antigen I/II based binding mechanism that created stronger adhesion forces than the strain lacking

antigen I/II, normal adhesion forces to glass and saliva-coated glass remained low also after strengthening

and amounted between 1 and 2 nN. Similarly small normal adhesion forces have been measured for a

staphylococcal strain adhering to polymer-brush coated surfaces [15], generally recognized as the most

non-adhesive surfaces known to date [16]. Yet, despite their small normal adhesion forces with SCFs, S.

mutans is able to maintain itself as a member of the oral microbiome on teeth and withstand naturally

occurring oral shear forces [17, 18]. On polymer brush coatings, lacking high affinity binding sites, lateral

adhesion forces have been interpreted as friction forces and these friction forces have been described to

cause immobilization of adhering bacteria under shear due to convective fluid flow [19]. SCFs possess

localized high affinity binding sites to antigen I/II and therefore the lateral forces measured for S. mutans

LT11 must be interpreted differently to represent localized binding.

Lateral adhesion forces were approximately three-fold stronger than normal adhesion forces for S. mutans

LT11, while for S. mutans IB03987 no significant differences were found. Directional dependence of

resistance to applied lateral force has been previously studied for proteins. Resistance to an applied force

for the proteins ubiquitin and E2lip3 depended on the direction in which forces were applied to the protein

[20–22]. In case of E2lip3, the directional dependence was attributed to the high content in β-sheets,

known to express a strong dependence on the direction in which it is being pulled [23] due to the way the

bonds are broken; upon a laterally-applied force, multiple bonds are simultaneously involved in resisting

the applied force while upon application of a normal force multiple bonds disrupt consecutively. Also, the

crucial role played by cellular integrin α4β1 and vascular cell adhesion molecule-1 (VCAM-1) in both rolling

and firm adhesion of leukocytes onto the vascular endothelium in the bloodstream, was determined by

their kinetics of dissociation, as governed by the rate at which a laterally-applied force was imposed [24].

A similar mechanism can be envisaged for S. mutans possessing antigen I/II, showing smaller normal

adhesion forces than lateral ones. Analysis of the structure of antigen I/II has revealed a stalk-like protein

of approximately 60 nm long, containing two independent adherence sites at opposite ends [1,2]. One of

the interaction sites is located within the A3VP1 region at the distal end, while the second one is positioned

within the C12 region near the anchor point (see Fig. 3 A). The existence of two independent interaction

domains points towards a heterogeneous adhesion mechanism [2], which suggests that directional

dependence not only originates from bonds within the interacting sites between antigen I/II and salivary

agglutinin, but also from consecutive or combined breaking of the two interaction sites, depending on

whether the applied force is normally- or laterally-oriented (see figure 3 B, C), respectively. This creates

an elongated rupture path [25], very similar to the bonding-debonding mechanism in Velcro (26).

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FIGURE 3 Schematic presentation of the mechanisms proposed for the differences observed between normal and lateral adhesion forces of S. mutans with salivary conditioning films, based on structural information on antigen I/II provided in [2]. (A) Model of the tertiary structure for antigen I/II, including the two adherence sites located at the A3VP1 region and the C12 region. (B) Subsequent disruption of a ligand-receptor complex due to an applied normal force. A normal force slightly larger than an individual ligand-receptor bond suffices to break a single bond. Disruption of the entire ligand-receptor complex requires consecutive disruption a two single bonds. (C) Effect of an applied lateral force on ligand-receptor bonds. In order to achieve detachment, the lateral force has to exceed the bond-strength of the entire ligand-receptor complex.

Using S. mutans LT11 and an isogenic mutant strain lacking antigen I/II, we identified antigen I/II as the

main contributor to the directional dependence of the adhesion forces of S. mutans to SCF-coated

surfaces. Strong lateral adhesion forces between antigen I/II and salivary agglutinins in SCF provide an

anchoring mechanism for S. mutans LT11 adhering to tooth surfaces in the oral cavity to maintain position

despite the presence of high naturally occurring shear forces. Lateral adhesion force based anchoring

mechanisms may also be operative for other bacterial strains adhering under shear conditions.

ACKNOWLEDGEMENTS

This study was entirely funded by the University Medical Center Groningen, Groningen, The Netherlands.

D.H. Veeregowda is also manager at Ducom Instruments Europe (Center for Innovation, L.J. Zielstraweg 2,

9713 GX, Groningen, The Netherlands), while H.J. Busscher is also director of a consulting company, SASA

BV (GN Schutterlaan 4, 9797 PC Thesinge, The Netherlands). The authors declare no potential conflicts of

V-region

P1A3

C1

C2

C3LPxTG anchor

Cell wall

A B C

Flateral

Salivary conditioning filmSAG SAG

Fn


Fn


Fn


Tim

e

Flateral



119

interest with respect to authorship and/or publication of this article. Opinions and assertions contained

herein are those of the authors and are not construed as necessarily representing views of the funding

organization or their respective employers.

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[2] Larson, M. R.; Rajashankar, K. R.; Crowley, P. J.; Kelly, C.; Mitchell, T. J.; Brady, L. J.; Deivanayagam, C. Crystal structure of the c-terminal region of Streptococcus mutans antigen I/II and characterization of salivary agglutinin adherence domains. J. Biol. Chem. 2011, 286, 21657–21666.

[3] Busscher, H. J.; Weerkamp, A. H. Specific and non-specific interactions in bacterial adhesion to solid substrata. FEMS Microbiol. Lett. 1987, 46, 165–173.

[4] Kisiela, D. I.; Rodriguez, V. B.; Tchesnokova, V.; Avagyan, H.; Aprikian, P.; Liu, Y.; Wu, X.-R.; Thomas, W. E.; Sokurenko, E. V. Conformational inactivation induces immunogenicity of the receptor-binding pocket of a bacterial adhesin. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 19089–19094.

[5] Mei, L.; Busscher, H. J.; Van der Mei, H. C.; Ren, Y. Influence of surface roughness on streptococcal adhesion forces to composite resins. Dent. Mater. 2011, 27, 770–778.

[6] Wessel, S. W.; Chen, Y.; Maitra, A; Van den Heuvel, E. R.; Slomp, A. M.; Busscher, H. J.; Van der Mei, H. C. Adhesion forces and composition of planktonic and adhering oral microbiomes. J. Dent. Res. 2014, 93, 84–88.

[7] Beaussart, A.; El-Kirat-Chatel, S.; Sullan, R. M. A.; Alsteens, D.; Herman, P.; Derclaye, S.; Dufrêne, Y. F. Quantifying the forces guiding microbial cell adhesion using single-cell force spectroscopy. Nat. Protoc. 2014, 9, 1049–1055.

[8] Van der Mei, H. C.; Rustema-Abbing, M.; de Vries, J.; Busscher, H. J. Bond strengthening in oral

bacterial adhesion to salivary conditioning films. Appl. Environ. Microbiol. 2008, 74, 5511–5515.

[9] Filleter, T.; Bennewitz, R. Structural and frictional properties of graphene films on sic(0001) studied by atomic force microscopy. Phys. Rev. B 2010, 81, 155412.

[10] Green, C. P.; Lioe, H.; Cleveland, J. P.; Proksch, R.; Mulvaney, P.; Sader, J. E. Normal and torsional spring constants of atomic force microscope cantilevers. Rev. Sci. Instrum. 2004, 75, 1988.

[11] Pettersson, T.; Nordgren, N.; Rutland, M. W.; Feiler, A. Comparison of different methods to calibrate torsional spring constant and photodetector for atomic force microscopy friction measurements in air and liquid. Rev. Sci. Instrum. 2007, 78, 093702.

[12] Sader, J. E.; Chon, J. W. M.; Mulvaney, P. Calibration of rectangular atomic force microscope cantilevers. Rev. Sci. Instrum. 1999, 70, 3967.

[13] Pettersson, T.; Naderi, A.; Makuska, R.; Claesson, P. M. Lubrication properties of bottle-brush polyelectrolytes: an AFM study on the effect of side chain and charge density. Langmuir 2008, 24, 3336–3347.

[14] Hannig, M.; Herzog, S.; Willigeroth, S. F.; Zimehl, R. Atomic force microscopy study of salivary pellicles formed on enamel and glass in vivo. Colloid Polym. Sci. 2001, 279, 479–483.

[15] Hannig, M. Transmission electron microscopic study of in vivo pellicle formation on dental restorative materials. Eur. J. Oral Sci. 1997, 105, 422–433.

[16] Landherr, L. J. T.; Cohen, C.; Agarwal, P.; Archer, L. A. Interfacial friction and adhesion of polymer brushes. Langmuir 2011, 27, 9387–9395.

[17] Raftari, M.; Zhang, Z.; Carter, S. R.; Leggett, G. J.; Geoghegan, M. Frictional properties of a polycationic brush. Soft Matter 2014, 10, 2759–2766.

[18] Busscher, H. J.; Norde, W.; Sharma, P. K.; Van der Mei, H. C. Interfacial re-arrangement in initial

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microbial adhesion to surfaces. Curr. Opin. Colloid Interface Sci. 2010, 15, 510–517.

[19] Swartjes, J.J.T.M., D.H. Veeregowda, H.C. van der Mei, H.J. Busscher, and P.K. Sharma. 2014. Normally oriented adhesion versus friction forces in bacterial adhesion to polymer-brush functionalized surfaces under fluid flow. Adv. Funct. Mater. 2014, 24, 4435-4441.

[20] Carrion-Vazquez, M.; Li, H.; Lu, H.; Marszalek, P. E.; Oberhauser, A. F.; Fernandez, J. M. The mechanical stability of ubiquitin is linkage dependent. Nat. Struct. Biol. 2003, 10, 738–743.

[21] Brockwell, D. J.; Paci, E.; Zinober, R. C.; Beddard, G. S.; Olmsted, P. D.; Smith, D. A.; Perham, R. N.; Radford, S. E. Pulling geometry defines the mechanical resistance of a beta-sheet protein. Nat. Struct. Biol. 2003, 10, 731–737.

[22] West, D. K.; Brockwell, D. J.; Olmsted, P. D.; Radford, S. E.; Paci, E. Mechanical resistance of

proteins explained using simple molecular models. Biophys. J. 2006, 90, 287–297.

[23] Rohs, R.; Etchebest, C.; Lavery, R. Unraveling proteins: a molecular mechanics study. Biophys. J. 1999, 76, 2760–2768.

[24] Zhang, X.; Craig, S. E.; Kirby, H.; Humphries, M. J.; Moy, V. T. Molecular basis for the dynamic strength of the integrin alpha4beta1/VCAM-1 interaction. Biophys. J. 2004, 87, 3470–3478.

[25] Beaussart, A.; Herman, P.; El-Kirat-Chatel, S.; Lipke, P. N.; Kucharíková, S.; Van Dijck, P.; Dufrêne, Y. F. Single-cell force spectroscopy of the medically important Staphylococcus epidermidis-Candida albicans interaction. Nanoscale 2013, 5, 10894–10900.

[26] Matouschek, A.; Bustamante, C. Finding a protein’s achilles heel. Nat. Struct. Biol. 2003, 10,

674–676.

CHAPTER 7

GENERAL DISCUSSION

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Bacterial adhesion and biofilm formation represent major problems in biomedical implants and devices,

with severe consequences to patients. Moreover, it yields a large economic impact on the health-care

system in general. The mechanisms of bacterial adhesion have been studied for decades, and results of

this research have led to the development of a variety of different strategies to prevent bacterial adhesion

and biofilm formation.

In this thesis we have investigated a number of different innovative coatings, including hydrogel-patterned

surfaces (Chapter 2), enzyme-based coatings (Chapter 3 and 4) and polymer brush coatings (Chapter 5). In

addition, the salivary coating to which lateral adhesion forces of adhering streptococci were studied

(chapter 6) represents the inevitable biological coating that forms on biomaterial surfaces once an implant

has been placed in the human body. These conditioning films have not been the true topic of this thesis,

but in most studies proteins present in the growth media used will form conditioning films [1]. In some

applications, these conditioning films can interfere with the working mechanisms of applied coatings [2,3],

but we anticipate that this effect will be minimal for the coatings presented in this thesis. Polymer brushes

are known to resist not only the adhesion of bacteria, but also of small molecules, such as proteins. The

non-adhesive hydrogels of our patterned coating prevent the adsorption of proteins that form

conditioning films, while the presence of a conditioning film on the adhesive patches does not change the

working mechanism of the coatings, as shown in later studies by allowing fibronectin to adsorb prior to

bacterial adhesion [4]. In addition, enzyme release from poly-(lactic-co-glycolic)-acid (PLGA) coatings will

not be hampered by conditioning films and the prevention of bacterial adhesion by this coating relies on

the removal of eDNA from the bacterial cell-wall, not on the absence of adhesion possibilities on the

surface. Moreover, this coating showed reductions in biofilm formation over 20 h in the presence of

bacterial growth medium, despite the conditioning film formed.

Mechanistically, the coatings studied are poor representatives of the multi-functional coatings that are

currently advocated for use on biomaterials implants and devices [5] (see Figure 1). However, the level of

multi-functionality of the currently presented coatings can be easily increased; the hydrogels used to

create patterned surfaces can be loaded with an antimicrobial [6], while the PLGA-enzyme coating can be

loaded with antimicrobial substances as well [7]. Besides antimicrobials, another possibility is to include

growth factors that facilitate the migration of appropriate host cells towards the implant and stimulate

tissue integration even further [8,9].

GENERAL DISCUSSION

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FIGURE 1 Overview of the surface properties of different antimicrobial functionalities that can be added to surface coatings and their possible clinical application. Reproduced with permission from [5].

In Chapters 5 and 6, the role of lateral forces in bacterial adhesion was determined using AFM in its lateral

mode. Even though bacteria most often adhere to surfaces from a flowing carrier fluid, the role of lateral

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forces is seldom studied, and in the few cases lateral forces are determined, it is by indirect measurements,

mostly related to the fluid shear present in a system. We revealed a strong correlation between the friction

forces experienced when a bacterium comes in contact with a polymer brush and the number of adhering

bacteria. Additionally, in the case of specific interactions, a strong direction dependence of the force can

occur, causing bacteria to be able to withstand lateral forces of much higher strength, compared to normal

forces. Although two completely different mechanisms, prevailing on very different surfaces, these

findings strongly suggest that lateral forces are involved in how and when bacteria adhere to surfaces, but

also in how they secure their bound state. Recently, liquid infused coatings have been developed to show

extremely low friction forces and thereby form non-fouling surfaces [10]. This is a novel example of how

decreasing lateral forces can play a role in reducing bacterial adhesion, and is one of the results of the

increased interest for the tribology of bacterial adhesion. It is therefore important to continue studying

the role of lateral forces and use the resulting knowledge to specifically engineer surfaces that disturb

bacterial adhesion mechanisms.

One of the drawbacks of many surface coatings that remains ill elicited in the current literature is the

suitability to be applied onto real implants. Whereas the inside of catheters is a quite protected

environment, implants such as hip and knee prostheses are subjected to large forces during surgical

implantation. Many types of coatings are claimed to have sufficient robustness, but often this is stated on

the basis of liquid exposure tests, not on the basis of any mechanical challenges. Chapter 4, describing the

robustness of our biodegradable PLGA coating, is a clear exception to this and it showed the PLGA coating

was not affected by different challenges. As an interesting aspect of these coatings, when they are

“hammered” into the bone as in orthopedic applications, even smearing out of the protective PLGA

coating would not necessarily disrupt its working mechanism as the active ingredient would still be

delivered to the relevant site. We did not perform similar tests on the hydrogel patterned surfaces, but

their application may be limited to applications where surgical placement is done less forcefully than in

orthopedics.

An important feature of surface coatings is the feasibility to be clinically applied. As shown in Figure 1,

there are many different clinical applications which all pose their own specific requirements to surface

coatings. In this perspective, the patterned hydrogel coating is most suitable for soft implants requiring

tissue integration and lacking high force implantation. In its current form, the robustness is not sufficient

to withstand the forceful placement as done in orthopedic surgery. Another drawback of the patterned

surface coating is the timely and costly nature of electron-beam lithography, making it not suitable for

large surface area modification, although these limitations can be overcome by using different techniques

that can yield the same result, but are better suited for larger-scale modifications, e.g. using photo-masks,

or other light mediated polymerization techniques [11,12]. In this respect, the clinical implementation of

the enzyme-based PLGA coating requires less additional work. By dip-coating, even large implants can be

readily coated in a relatively cost-effective and fast manner. Although animal studies are necessary to

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support the current in-vitro findings, the coating is robust enough to be studied in orthopedic implant

models.

REFERENCES

[1] Chen, M.Y.; Chen, M.J.; Lee, P.F.; Cheng, L.H.; Huang, L.J.; Lai, C.H.; Huang, K.H. Towards real-time observation of conditioning film and early biofilm formation under laminar flow conditions using a quartz crystal microbalance. Biochem. Eng. J. 2010, 53, 121–130.

[2] Furno, F.; Morley, K. S.; Wong, B.; Sharp, B. L.; Arnold, P. L.; Howdle, S. M.; Bayston, R.; Brown, P. D.; Winship, P. D.; Reid, H. J. Silver nanoparticles and polymeric medical devices: a new approach to prevention of infection? J. Antimicrob. Chemother. 2004, 54, 1019–1024.

[3] Everaert, E. P.; van de Belt-Gritter, B.; Van der Mei, H. C.; Busscher, H. J.; Verkerke, G. J.; Dijk, F.; Mahieu, H. F.; Reitsma, a. In vitro and in vivo microbial adhesion and growth on argon plasma-treated silicone rubber voice prostheses. J. Mater. Sci. Mater. Med. 1998, 9, 147–157.

[4] Wang, Y.; Subbiahdoss, G.; de Vries, J.; Libera, M.; Van der Mei, H. C.; Busscher, H. J. Effect of adsorbed fibronectin on the differential adhesion of osteoblast-like cells and Staphylococcus aureus with and without fibronectin-binding proteins. Biofouling 2012, 28, 1011–1021.

[5] Busscher, H. J.; Van der Mei, H. C.; Subbiahdoss, G.; Jutte, P. C.; van den Dungen, J. J. A. M.; Zaat, S. A. J.; Schultz, M. J.; Grainger, D. W. Biomaterial-associated infection: locating the finish line in the race for the surface. Sci. Transl. Med. 2012, 4, 153rv10.

[6] Singh, B.; Sharma, S.; Dhiman, A. Design of antibiotic containing hydrogel wound dressings: biomedical properties and histological study of wound healing. Int. J. Pharm. 2013, 457, 82–91.

[7] Yeh, M.; Chen, K.; Chang, N.; Chen, H. Prolonged antibiotic release by plga encapsulation on titanium alloy. 2011, 33, 17–22.

[8] Schmidmaier, G.; Wildemann, B.; Stemberger, A.; Haas, N. P.; Raschke, M. Biodegradable poly(d,l-lactide) coating of implants for continuous release of growth factors. J. Biomed. Mater. Res. 2001, 58, 449–455.

[9] Wildemann, B.; Sander, A.; Schwabe, P.; Lucke, M.; Stöckle, U.; Raschke, M.; Haas, N. P.; Schmidmaier, G. Short term in vivo biocompatibility testing of biodegradable poly(d,l-lactide)--growth factor coating for orthopaedic implants. Biomaterials 2005, 26, 4035–4040.

[10] Epstein, A. K.; Wong, T.S.; Belisle, R. a; Boggs, E. M.; Aizenberg, J. Liquid-infused structured surfaces with exceptional anti-biofouling performance. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 13182–13187.

[11] Poelma, J. E.; Fors, B. P.; Meyers, G. F.; Kramer, J. W.; Hawker, C. J. Fabrication of complex three-dimensional polymer brush nanostructures through light-mediated living radical polymerization. Angew. Chemie 2013, 125, 6982–6986.

[12] Rodriguez-Emmenegger, C.; Preuss, C. M.; Yameen, B.; Pop-Georgievski, O.; Bachmann, M.; Mueller, J. O.; Bruns, M.; Goldmann, A. S.; Bastmeyer, M.; Barner-Kowollik, C. Controlled cell adhesion on poly(dopamine) interfaces photopatterned with non-fouling brushes. Adv. Mater. 2013, 25, 6123–6127.

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SUMMARY

SUMMARY

128

SUMMARY

Bacterial adhesion is the main cause of implant failure, despite the existence of numerous preventive

strategies and use of antibiotics. Procedures regarding the sterile environment in the operating theatre

have been optimized to a near maximum and due to the fading discovery of new and better antibiotics

combined with the rise of antibiotic resistant bacterial strains, there are decreasing options for treatment

[1-4] . Surface coating of materials has been applied as a biomaterial surface modification to prevent the

adhesion of bacteria, the first step in biofilm formation (see fig. 1). In Chapter 1, we provide an overview

of the current developments in antimicrobial surface coatings. The method of choice has long been to

achieve anti-adhesive surfaces, for which polymer-brush coatings have been used extensively. By forming

a highly hydrated layer of extended polymer chains, polymer-brushes represent a steric barrier that

prevents bacteria from adhering to the surface. However, even the best performing polymer-brush surface

is not able to completely prevent all bacteria from adhering, and bacteria that do manage to adhere can

grow into a full biofilm when given the opportunity [5]. Inclusion of some sort of anti-bacterial substance

is therefore necessary to achieve enhanced results regarding bacterial adhesion and biofilm formation. In

addition, in applications where tissue incorporation is required, surface coatings need to allow tissue cells

to adhere and integrate the implant into the body, which in turn offers the best protection against further

bacterial contamination. In this thesis we describe a number of anti-bacterial surface coatings, while also

trying to enhance the current state of knowledge about the mechanisms by which bacteria adhere, since

this can offer valuable information for the design of new strategies to prevent bacterial adhesion and

biofilm formation.

Micro-patterned surfaces

Polyethylene glycol (PEG)-based surface coatings have long dominated the field of anti-adhesive surface

modifications [7]. As an excellent repellant to both proteins and bacterial cells, it has been a promising

candidate to serve as a coating for biomedical implants and devices to withstand bacterial adhesion and

biofilm formation. However, as a repulsive surface coating, PEG polymer-brushes also hinder tissue cells

to accommodate the surface and thereby prevent tissue integration in applications where it might be

desirable [8]. In Chapter 2, we used micro-patterned surfaces of PEG-hydrogels to create surfaces with a

repulsive nature towards bacteria, by taking advantage of its anti-adhesive character, while at the same

time by introducing unmodified patches, exposing cell-adhesive surfaces, we create anchor points for

mammalian cells to bypass the anti-adhesiveness of the coating. Whereas many previous studies have

used cell recognition peptide-sequences, i.e. RGD, to stimulate cell adhesion on PEG modified surfaces

[9,10], we showed that combining PEG hydrogel patches with the appropriate amount of bare surface can

effectively prevent bacterial adhesion, while mammalian cells are still able to adhere and proliferate. An

inter-gel spacing of 1 µm showed to be the optimum spacing which could sufficiently reduce bacterial

adhesion and allow osteoblasts to adhere to and spread on the surface. This inter-gel spacing, of the same

order of magnitude as bacteria, prevents bacteria from adhering while osteoblasts use their focal contacts

and focal adhesions, which can extend hundreds of nanometers, to reach the adhesive spaces in between

the hydrogels.

SUMMARY

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FIGURE 1 Schematic representation of the different steps in biofilm formation consisting of initial attachment, early growth phase, biofilm maturation, and dispersal of the biofilm [6]. Copyright by the Montana State University Center for Biofilm Engineering.

DNase I containing surface modification

eDNA has been pointed out as an important component of extrapolymeric substances (EPS) and playing

an important role in adhesion of bacteria and the subsequent formation and maintenance of biofilm.

Besides DNase I, several other enzymes already have been studied as possible surface coatings to prevent

bacterial adhesion, including lysozyme and dispersin B [11,12]. In Chapter 3, we have shown a proof of

principle of a functional DNase I coating, effective in preventing bacterial adhesion and biofilm formation,

by using dopamine as an intermediate for surface attachment of DNase I. By presenting DNase I at the

material-bacteria interface, bacterial adhesion could be prevented and biofilm formation after 14 h was

drastically reduced as well. Dopamine is generally considered as a chemically stable attachment method,

with studies reporting only a slight loss of surface attached compounds after exposure to phosphate

buffered saline (PBS) for time periods between 7 and 30 days [13,14]. However, as an implant is inserted

into the body, it is exposed to other harsh conditions that can be detrimental to any ‘soft’ surface coating.

The handling of the implant by the surgeon is one of these, but nevertheless mechanical testing of surface

coatings is seldom reported. The presence of DNase I and maintaining its activity is another challenge in

the surface coating process. Therefore, in Chapter 4, we used poly-(lactic-co-glycolic)-acid (PLGA) as a

‘hard’ polymer surface coating that could withstand mechanical stress, and incorporated DNase I in a sugar

SUMMARY

130

glass, inulin, to protect the enzyme and retain its activity during the coating process. As a degradable

polymer approved by the U.S. Food and Drug Administration, PLGA released the inulin-packaged DNase I

for an extended time period and coating of a titanium surface reduced the number of adhering bacteria

up to 99% and lowered the biomass of biofilms formed in 20 h significantly. More importantly, the PLGA

coating proved to be able to withstand a number of storage and handling conditions that mimicked the

environment to which an implant could be exposed in the operating theatre.

Lateral forces in bacterial adhesion

As the first step of biofilm formation, initial adhesion of bacteria is a main focus in the prevention of

biomaterial associated infections. However, despite the increasing number of potential strategies

developed to fight bacterial adhesion, the exact mechanisms remain poorly understood [15,16]. Major

advances have been made by the use of cell probe microscopy in which bacterial adhesion forces of single

bacteria to surfaces have been studied, and even the forces between specific cell-wall appendages and

surfaces can be measured this way [17–19]. However, all these studies have in common that the direction

of the forces between the bacteria and the surface are measured perpendicularly to the surface. In most

real-life situations, and in many studies on bacterial adhesion, bacteria approach a surface from a flowing

carrier liquid [20–22], which means the forces arising between these bacteria and the surface are actually

laterally oriented. In this perspective, we studied the lateral forces arising between Staphylococcus

epidermidis ATCC 35983 and several polymer brush coatings when moving a bacterium along the surface

using lateral force microscopy (Chapter 5). Using two types of PEG molecules in different concentrations,

we created brushes with different softness and could conclude that not only the friction forces between

polymer-brush modified surfaces and bacteria are highly correlated to the number of adhering bacteria,

but also the softness of a polymer-brush plays an important role in the success of resisting bacterial

adhesion. On soft polymer-brushes bond-maturation is much more extensive compared to rigid brushes,

from which bacteria easily desorb.

In addition to the lateral forces between bacteria and polymer-brushes, in Chapter 6 we studied the

influence of specific interactions between bacteria and surfaces on the lateral forces that arise when

bacteria move along the surface. To this extend we used Streptococcus mutans with antigen I/II on its

outer surface together with an isogenic mutant strain, not possessing this particular adhesin. The structure

of antigen I/II has been revealed into detail in the last decade and it has been demonstrated that both its

distal part as well as the cell-wall anchoring domain are able to interact with salivary agglutinins (SAGs)

present in salivary conditioning films (SCFs) [23,24]. Comparison of both the normally-oriented and the

lateral adhesion forces between these two strains and glass surfaces coated with SCFs suggests that the

specific interactions between antigen I/II and SAG are directional depended and are better capable to

resist lateral forces than forces oriented normally to the surface. This suggest that S. mutans has adapted

to the highly dynamic environment of the oral cavity, where mastication, tongue movement and presence

of fluids all create a high amount of shear forces. The nature of these high affinity binding site between

SUMMARY

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bacterial ligands and salivary receptors suggests that similar mechanisms could also hold true for other

bacterial strains capable of forming bonds using specific interactions.

CONCLUSIONS

Antibiotic-independent strategies to prevent infection of medical devices and implants are a crucial factor

for the success of future medicine. Problematic in the design of surface modifications for these

applications is the wide variety of existing devices and implants, of which each comes with its own

requirements regarding stability, tissue integration and duration of use. Our results suggest that surface

patterning of anti-adhesive patches can aid in the tissue integration component of implants, while still

offering a high degree of resistance against bacterial adhesion. Another approach to prevent bacterial

adhesion is by applying natural components, like enzymes, that have specific activities to disturb bacteria

in their adhesion and ability to form biofilms. Using biodegradable polymers makes it possible to create

robust surface coatings that release these substances and keep the implant free of bacteria until tissue

integration is achieved. However, as discussed in the final Chapter 7, these surface modification strategies

will not be universally suitable for all applications, and therefore it is important to keep studying the

mechanisms by which bacteria colonize surfaces, since this remains poorly understood. We made a start

in unravelling the nature of lateral, or friction, forces between bacteria and surfaces whereas before only

the normal adhesion forces have been directly measured. A better understanding of the role of lateral

forces can help in future engineering of new surface modifications to resist bacterial adhesion.

REFERENCES

[1] Lee, I.; Agarwal, R. K.; Lee, B. Y.; Fishman, N. O.; Umscheid, C. A. Systematic review and cost analysis comparing use of chlorhexidine with use of iodine for preoperative skin antisepsis to prevent surgical site infection. Infect. Control Hosp. Epidemiol. 2010, 31, 1219–1229.

[2] Palmer, A. C.; Kishony, R. Understanding, predicting and manipulating the genotypic evolution of antibiotic resistance. Nat. Rev. Genet. 2013, 14, 243–248.

[3] Chow, T. T.; Yang, X. Y. Ventilation performance in operating theatres against airborne infection: review of research activities and practical guidance. J. Hosp. Infect. 2004, 56, 85–92.

[4] Kåhrström, C. T. Entering a post-antibiotic era? Nat. Rev. Microbiol. 2013, 11, 146–146.

[5] Nejadnik, M. R.; Van der Mei, H. C.; Norde, W.; Busscher, H. J. Bacterial adhesion and growth on a polymer brush-coating. Biomaterials 2008, 29, 4117–4121.

[6] Costerton, B. Microbial ecology comes of age and joins the general ecology community. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 16983–16984.

[7] Konradi, R.; Acikgoz, C.; Textor, M. Polyoxazolines for nonfouling surface coatings: a direct comparison to the gold standard PEG. Macromol. Rapid Commun. 2012, 33, 1663–1676.

[8] Zhang, M.; Desai, T.; Ferrari, M. Proteins and cells on PEG immobilized silicon surfaces. Biomaterials 1998, 19, 953–960.

[9] Muszanska, A. K.; Rochford, E. T. J.; Gruszka, A.; Bastian, A. A.; Busscher, H. J.; Norde, W.; van der Mei, H. C.; Herrmann, A. Antiadhesive polymer brush coating functionalized with antimicrobial and rgd peptides to reduce biofilm formation and enhance tissue integration. Biomacromolecules 2014.

[10] Burdick, J. A.; Anseth, K. S. Photoencapsulation of osteoblasts in injectable rgd-modified peg

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hydrogels for bone tissue engineering. Biomaterials 2002, 23, 4315–4323.


[12] Pavlukhina, S. V; Kaplan, J. B.; Xu, L.; Chang, W.; Yu, X.; Madhyastha, S.; Yakandawala, N.; Mentbayeva, A.; Khan, B.; Sukhishvili, S. A. Noneluting enzymatic antibiofilm coatings. ACS Appl. Mater. Interfaces 2012, 4, 4708–4716.

[13] Yang, W. J.; Cai, T.; Neoh, K.G.; Kang, E.T.; Dickinson, G. H.; Teo, S. L.-M.; Rittschof, D. Biomimetic anchors for antifouling and antibacterial polymer brushes on stainless steel. Langmuir 2011, 27, 7065–7076.

[14] Pop-Georgievski, O.; Popelka, Š.; Houska, M.; Chvostová, D.; Proks, V.; Rypáček, F. Poly(ethylene oxide) layers grafted to dopamine-melanin anchoring layer: stability and resistance to protein adsorption. Biomacromolecules 2011, 12, 3232–3242.

[15] Hori, K.; Matsumoto, S. bacterial adhesion: from mechanism to control. Biochem. Eng. J. 2010, 48, 424–434.

[16] Katsikogianni, M.; Missirlis, Y. F. Concise review of mechanisms of bacterial adhesion to biomaterials and of techniques used in estimating bacteria-material interactions. Eur. Cell. Mater. 2004, 8, 37–57.

[17] Beaussart, A.; El-Kirat-Chatel, S.; Herman, P.; Alsteens, D.; Mahillon, J.; Hols, P.; Dufrêne, Y. F. Single-cell force spectroscopy of probiotic bacteria. Biophys. J. 2013, 104, 1886–1892.

[18] Harimawan, A.; Rajasekar, A.; Ting, Y.-P. Bacteria attachment to surfaces: AFM force spectroscopy

and physicochemical analyses. J. Colloid Interface Sci. 2011, 364, 213–218.

[19] Razatos, a; Ong, Y. L.; Sharma, M. M.; Georgiou, G. Molecular determinants of bacterial adhesion monitored by atomic force microscopy. Proc. Natl. Acad. Sci. U. S. A. 1998, 95, 11059–11064.

[20] Busscher, H. J.; Van der Mei, H. C. Microbial adhesion in flow displacement Systems. Clin. Microbiol. Rev. 2006, 19, 127–141.

[21] Drescher, K.; Shen, Y.; Bassler, B. L.; Stone, H. a. biofilm streamers cause catastrophic disruption of flow with consequences for environmental and medical systems. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 4345–4350.

[22] Rusconi, R.; Guasto, J. S.; Stocker, R. bacterial transport suppressed by fluid shear. Nat. Phys. 2014, 10, 212–217.

[23] Larson, M. R.; Rajashankar, K. R.; Crowley, P. J.; Kelly, C.; Mitchell, T. J.; Brady, L. J.; Deivanayagam, C. Crystal structure of the c-terminal region of Streptococcus mutans antigen I/II and characterization of salivary agglutinin adherence domains. J. Biol. Chem. 2011, 286, 21657–21666.

[24] Larson, M. R.; Rajashankar, K. R.; Patel, M. H.; Robinette, R. A.; Crowley, P. J.; Michalek, S.; Brady, L. J.; Deivanayagam, C. elongated fibrillar structure of a streptococcal adhesin assembled by the high-affinity association of alpha- and PPII-helices. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 5983–5988.

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SAMENVATTING

De belangrijkste oorzaak voor het falen van een biomedisch implantaat, ondanks de vele preventie

methoden en het gebruik van antibiotica, is de hechting van bacteriën. De procedures voor het creëren

van een steriele omgeving in de operatiekamer zijn bijna maximaal geoptimaliseerd en door de afname

van de ontdekking van nieuwe en betere antibiotica, samen met de toenemende antibiotica resistentie

van vele bacteriën, neemt het aantal mogelijkheden om infecties te bestrijden af [1-4]. Het coaten van het

oppervlak van een materiaal, met als doel het hechten van bacteriën te voorkomen, is daarom een veel

onderzochte methode ter preventie van de hechting van bacteriën, de eerste stap in biofilm vorming. In

Hoofdstuk 1 wordt een overzicht gepresenteerd van recente ontwikkelingen op het gebied van

antimicrobiële oppervlakte coatings. De meest toegepaste strategie tot dusver is het verminderen van

hechting, waarvoor veelal polymere-borstel coatings gebruikt worden. Door de formatie van een sterk

gehydrateerde laag van uitgestrekte polymere ketens, vormen polymere-borstel coatings een sterische

barrière die hechting van bacteriën op het oppervlak voorkomt. Echter, zelfs de beste polymere-borstel

coating is niet in staat om de hechting van bacteriën in zijn geheel te voorkomen, en daarnaast groeien de

enkele bacteriën die wel hechten uit tot een biofilm zodra ze de kans krijgen [5]. Het toevoegen van een

antibacteriële substantie is daarom vereist om betere resultaten te verkrijgen wat betreft bacteriële

hechting en biofilm vorming. Daarnaast zijn er ook applicaties waarbij weefsel integratie een voordeel is,

en de gebruikte coatings mogen in dat geval de hechting van weefselcellen en de integratie van het

materiaal in het lichaam, wat overigens de beste bescherming tegen bacteriële contaminatie biedt, niet

verstoren. In dit proefschrift worden een aantal antibacteriële oppervlakte coatings beschreven, terwijl

ook geprobeerd wordt de huidige kennis van mechanismen die bacteriën gebruiken om aan oppervlakken

te hechten te vergroten, omdat dit waardevolle informatie oplevert die kan helpen bij het ontwikkelen van

nieuwe strategieën om bacteriële hechting en biofilm vorming te voorkomen.

Coatings bestaande uit micro-patronen

Oppervlakte coatings gebaseerd op polyethyleen glycol (PEG) zijn lange tijd dominant geweest op het

gebied van oppervlakte modificaties die hechting tegen moeten gaan [7]. Omdat de hechting van zowel

eiwitten als bacteriën drastisch verminderde, werd het gezien als een veelbelovende coating voor

biomedische implantaten en apparaten om de hechting van bacteriën en biofilm vorming tegen te gaan.

Echter, als afstotende coating hinderen PEG polymere-borstel coatings ook de hechting van weefselcellen

en sluiten de mogelijkheid tot weefsel integratie op deze manier uit, ook in toepassingen waar die

integratie wenselijk, of zelf vereist is [8]. In Hoofdstuk 2 worden oppervlakken gebruikt met een micro-

patroon van PEG-hydrogelen om oppervlakken te creëren waarop bacteriën, door het afstotende karakter,

niet hechten, terwijl tegelijkertijd bepaalde delen van het met silaan gemodificeerd glas oppervlak

onveranderd worden gelaten, waardoor het voor weefselcellen mogelijk is om te hechten. Op deze manier

worden de niet-hechtende eigenschappen omzeild, voor wat betreft weefselcellen, terwijl het voor

bacteriën nagenoeg onmogelijk blijft om aan het oppervlak te hechten. Hoewel vele onderzoeken speciale

peptide-sequenties zoals RGD gebruiken om hechting van cellen mogelijk te maken op PEG

gemodificeerde oppervlakken [9,10], laten wij zien dat de combinatie van PEG hydrogelen en de juiste

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hoeveelheid ongemodificeerd glas oppervlak effectief kan zijn in het tegengaan van bacteriële hechting,

terwijl weefselcellen nog steeds in staat zijn om te hechten en te delen. Een ruimte van 1 µm tussen de

PEG hydrogelen bleek de meest optimale afstand te zijn om de hechting van bacteriën significant te

verlagen en osteoblasten de kans te geven om te hechten en te spreiden op het oppervlak. Deze ruimte

tussen de hydrogelen, die in dezelfde orde van grootte valt als een bacterie, voorkomt de hechting van

bacteriën, terwijl osteoblasten hun specifieke hechtingsmoleculen gebruiken die zich tot wel 100 µm

kunnen uitstrekken en op die manier de ongemodificeerde plekken tussen de hydrogelen kunnen

bereiken.

FIGUUR 1 Schematische weergave van de verschillende stappen van biofilm vorming, bestaande uit; initiële hechting, de vroege groeifase, biofilm maturatie, en tenslotte dispersie van de biofilm [6]. Auteursrecht in handen van Montana State University Center for Biofilm Engineering.

Oppervlakte modificaties met DNase I

eDNA is een zeer belangrijke component van de extracellulaire polymere substantie van bacteriën

gebleken en speelt een belangrijke rol in de hechting van bacteriën en de daaropvolgende vorming, en het

onderhoud, van biofilms. Behalve DNase I zijn er meerdere enzymen bestudeerd voor mogelijk gebruik in

oppervlakte coatings om bacteriële hechting tegen te gaan, waaronder lysozyme en dispersin B [11,12]. In

Hoofdstuk 3 wordt een “proof of principle” van een functionele DNase I coating getoond, die effectief is in

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het voorkomen van bacteriële hechting en biofilm formatie, door het gebruik van dopamine als een

intermediair voor de hechting van DNase I. Door DNase I te gebruiken op het grensvlak tussen materiaal

en bacterie werd de hechting van bacteriën voorkomen en biofilm vorming gedurende 14 uur drastisch

verminderd. Dopamine wordt beschouwd als een chemisch stabiele bindingsmethode, wat ondersteund

wordt door onderzoeken die slechts een klein verlies van aangebracht materiaal laten zien na blootstelling

aan een fysiologische zoutoplossing gedurende tijdsintervallen tussen 7 en 30 dagen [13,14]. Echter,

wanneer een materiaal in het lichaam geïmplanteerd wordt, wordt het blootgesteld aan extreme condities

die destructief zijn voor elke ‘zachte’ coating. Het vasthouden en plaatsen van het implantaat door de

chirurg is hier één van, maar het mechanisch testen van coatings wordt zelden genoemd in studies. De

aanwezigheid van DNase I en het behoud van de enzymatische activiteit is een andere uitdaging in het

coating proces. Vandaar dat in Hoofdstuk 4 poly-(lactic-co-glycolic)-acid (PLGA) gebruikt is als een hard

polymeer dat mechanische stress kan weerstaan, en DNase I verpakt in een suikerglas, inuline, om het

enzym te beschermen tijdens het coating proces. PLGA is een degradeerbaar polymeer dat is goedgekeurd

door de U.S. Food and Drug Administration. Het PLGA degradeerde langzaam en de toegevoegde DNase I

werd afgegeven over een langere tijd, waarbij de hechting van bacteriën op gecoat titanium met 99%

afnam en de biomassa van 20 uur oude biofilms significant lager was dan zonder DNase I coating.

Belangrijker, de PLGA coating bleek bestand tegen een aantal condities die simuleerden wat een

implantaat ondergaat in de operatie kamer.

Laterale krachten in de hechting van bacteriën

Initiële hechting van bacteriën is, als zijnde de eerste stap in het vormen van biofilms, het belangrijkste

focus punt in de preventie van biomateriaal geassocieerde infecties. Echter, terwijl het aantal potentiele

methoden dat wordt ontwikkeld om bacteriële hechting te voorkomen toeneemt, blijven de exacte

mechanismen waarmee bacteriën hechten slecht begrepen [15,16]. Grote stappen zijn gemaakt door het

gebruik van “bacterial probe” microscopie waarbij de hechtingskrachten tussen individuele bacteriën en

oppervlakken bestudeerd zijn en zelfs de krachten tussen specifieke structuren op de celwand en

oppervlakken gemeten kunnen worden [17-19]. Echter, in al deze onderzoeken worden de krachten

tussen bacteriën en het oppervlak loodrecht ten opzichte van het oppervlak gemeten. In de natuurlijke

situatie, en in veel onderzoeken naar bacteriële hechting, naderen bacteriën een oppervlak vanuit een

vloeistof die ze meevoert [20-22], wat inhoudt dat de krachten die ontstaan tussen deze bacteriën en het

oppervlak in werkelijkheid parallel aan het oppervlak lopen. Daarom zijn laterale krachten gemeten tussen

Staphylococcus epidermidis ATCC 25984 en verschillende polymere-borstel coatings. Hierbij werd een

bacterie langs het oppervlak bewogen en de kracht gemeten met behulp van laterale kracht microscopie

(Hoofdstuk 5). Door twee type PEG-moleculen in verschillende concentraties te gebruiken hebben we

coatings gemaakt die verschilden in zachtheid en kwamen tot de conclusie dat niet alleen de

wrijvingskrachten tussen polymere-borstel coatings en bacteriën sterk gerelateerd zijn aan het aantal

bacteriën dat hecht, maar ook de zachtheid van de polymere-borstel coatings speelt een belangrijke rol in

hoe succesvol de coating is in het weerstaan van bacteriële hechting. In het geval van zachte coatings

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treedt veel meer maturatie van de binding tussen bacterie en oppervlak op vergeleken met harde coatings,

waarvan bacteriën veel makkelijker loslaten na initieel contact.

Naast laterale krachten tussen bacteriën en polymere borstel coatings wordt in Hoofdstuk 6 de invloed

van specifieke interacties tussen bacteriën en oppervlakken op de laterale krachten bestudeerd. Hiervoor

is een Streptococcus mutans stam gebruikt met antigeen I/II op het oppervlak, samen met een isogene

mutant welke deze hechtings structuren niet heeft. De structuur van antigeen I/II is in de laatste tien jaar

tot in detail bepaald en het is bekend dat zowel het distale deel als het domein dat in de celwand vastzit,

in staat is om interacties aan te gaan met speciale receptoren in speeksel films [23,24]. Het vergelijken van

de hechtingskrachten loodrecht en lateraal tussen deze twee stammen en glas gecoat met een speeksel

film, suggereert dat de specifieke interacties tussen antigeen I/II en de speciale receptoren

richtingsafhankelijk zijn en meer weerstand bieden tegen laterale krachten dan tegen krachten loodrecht

op het oppervlak. Dit zou betekenen dat S. mutans zich heeft aangepast aan de zeer dynamische omgeving

van de mondholte, waarin kauwen, bewegingen van de tong en de aanwezigheid van allerlei vloeistoffen

zorgen voor grote laterale krachten. Het algemene karakter van de specifieke binding tussen bacteriën en

de receptoren in speeksel houdt in dat een vergelijkbaar mechanisme ook zou kunnen optreden in andere

bacteriële stammen die in staat zijn te hechten met behulp van specifieke interacties.

CONCLUSIES

Antibiotica onafhankelijke strategieën om infecties gerelateerd aan medische apparaten en implantaten

te voorkomen zijn een cruciale factor voor het succes van de toekomstige geneeskunde. Problematisch in

het ontwerp van oppervlakte modificaties voor deze toepassingen is de grote variatie in bestaande

apparaten en implantaten, ieder met eigen eisen wat betreft stabiliteit, weefsel integratie en

gebruiksduur. Onze resultaten laten zien dat oppervlakte modificaties bestaande uit patronen die hechting

kunnen voorkomen en delen die hechting toestaan, kunnen bijdragen aan de weefsel integratie van

implantaten, terwijl ze nog steeds zeer goed bestand zijn tegen de hechting van bacteriën. Een andere

aanpak om hechting van bacteriën te voorkomen is door het gebruik van natuurlijke componenten, zoals

enzymen, die een specifieke activiteit hebben om de hechting en biofilm vorming van bacteriën te kunnen

verstoren. Door het gebruik van biodegradeerbare polymeren is het mogelijk om een robuuste

oppervlakte coating te maken die deze substanties langzaam loslaat en het implantaat bacterie vrij houdt

totdat weefsel integratie heeft plaatsgevonden. Echter, zoals in Hoofdstuk 7 beschreven wordt, zullen deze

oppervlakte modificaties veelal niet universeel toepasbaar zijn, en daarom is het belangrijk om door te

gaan met het bestuderen van de mechanismen die bacteriën gebruiken om oppervlakken te koloniseren,

iets waar nog lang niet alles over bekend is. We hebben een start gemaakt met het ontrafelen van de

oorzaak van laterale, of wrijvings, -krachten tussen bacteriën en oppervlakken, waar voorheen alleen de

hechtingskrachten loodrecht op het oppervlak direct gemeten werden. Een beter begrip van de rol die

laterale krachten spelen kan helpen bij het toekomstig ontwerp van nieuwe oppervlakte modificaties om

de hechting van bacteriën te voorkomen.

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REFERENCES

[1] Lee, I.; Agarwal, R. K.; Lee, B. Y.; Fishman, N. O.; Umscheid, C. A. Systematic review and cost analysis comparing use of chlorhexidine with use of iodine for preoperative skin antisepsis to prevent surgical site infection. Infect. Control Hosp. Epidemiol. 2010, 31, 1219–1229.

[2] Palmer, A. C.; Kishony, R. Understanding, predicting and manipulating the genotypic evolution of antibiotic resistance. Nat. Rev. Genet. 2013, 14, 243–248.

[3] Chow, T. T.; Yang, X. Y. Ventilation performance in operating theatres against airborne infection: review of research activities and practical guidance. J. Hosp. Infect. 2004, 56, 85–92.

[4] Kåhrström, C. T. Entering a post-antibiotic era? Nat. Rev. Microbiol. 2013, 11, 146–146.

[5] Nejadnik, M. R.; Van der Mei, H. C.; Norde, W.; Busscher, H. J. Bacterial adhesion and growth on a polymer brush-coating. Biomaterials 2008, 29, 4117–4121.

[6] Costerton, B. Microbial ecology comes of age and joins the general ecology community. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 16983–16984.

[7] Konradi, R.; Acikgoz, C.; Textor, M. Polyoxazolines for nonfouling surface coatings: a direct comparison to the gold standard PEG. Macromol. Rapid Commun. 2012, 33, 1663–1676.

[8] Zhang, M.; Desai, T.; Ferrari, M. Proteins and cells on PEG immobilized silicon surfaces. Biomaterials 1998, 19, 953–960.

[9] Muszanska, A. K.; Rochford, E. T. J.; Gruszka, A.; Bastian, A. A.; Busscher, H. J.; Norde, W.; Van der Mei, H. C.; Herrmann, A. Antiadhesive polymer brush coating functionalized with antimicrobial and rgd peptides to reduce biofilm formation and enhance tissue integration. Biomacromolecules 2014, 15, 2019–2026.

[10] Burdick, J. A.; Anseth, K. S. Photoencapsulation of osteoblasts in injectable RGD-modified PEG hydrogels for bone tissue engineering. Biomaterials 2002, 23, 4315–4323.


[12] Pavlukhina, S. V; Kaplan, J. B.; Xu, L.; Chang, W.; Yu, X.; Madhyastha, S.; Yakandawala, N.;

Mentbayeva, A.; Khan, B.; Sukhishvili, S. A. Noneluting enzymatic antibiofilm coatings. ACS Appl. Mater. Interfaces 2012, 4, 4708–4716.

[13] Yang, W. J.; Cai, T.; Neoh, K.G.; Kang, E.T.; Dickinson, G. H.; Teo, S. L.-M.; Rittschof, D. Biomimetic anchors for antifouling and antibacterial polymer brushes on stainless steel. Langmuir 2011, 27, 7065–7076.

[14] Pop-Georgievski, O.; Popelka, Š.; Houska, M.; Chvostová, D.; Proks, V.; Rypáček, F. Poly(ethylene oxide) layers grafted to dopamine-melanin anchoring layer: stability and resistance to protein adsorption. Biomacromolecules 2011, 12, 3232–3242.

[15] Hori, K.; Matsumoto, S. Bacterial adhesion: from mechanism to control. Biochem. Eng. J. 2010, 48, 424–434.

[16] Katsikogianni, M.; Missirlis, Y. F. Concise review of mechanisms of bacterial adhesion to biomaterials and of techniques used in estimating bacteria-material interactions. Eur. Cell Mater. 2004, 8, 37–57.

[17] Beaussart, A.; El-Kirat-Chatel, S.; Herman, P.; Alsteens, D.; Mahillon, J.; Hols, P.; Dufrêne, Y. F. Single-cell force spectroscopy of probiotic bacteria. Biophys. J. 2013, 104, 1886–1892.

[18] Harimawan, A.; Rajasekar, A.; Ting, Y.-P. Bacteria attachment to surfaces: AFM force spectroscopy and physicochemical analyses. J. Colloid Interface Sci. 2011, 364, 213–218.

[19] Razatos, A.; Ong, Y. L.; Sharma, M. M.; Georgiou, G. Molecular determinants of bacterial adhesion monitored by atomic force microscopy. Proc. Natl. Acad. Sci. U. S. A. 1998, 95, 11059–11064.

[20] Busscher, H. J.; Van der Mei, H. C. Microbial adhesion in flow displacement Systems. Clin. Microbiol. Rev. 2006, 19, 127–141.

[21] Drescher, K.; Shen, Y.; Bassler, B. L.; Stone, H. A. Biofilm streamers cause catastrophic disruption of flow with consequences for environmental and medical systems. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 4345–4350.

[22] Rusconi, R.; Guasto, J. S.; Stocker, R. Bacterial transport suppressed by fluid shear. Nat. Phys. 2014, 10, 212–217.

[23] Larson, M. R.; Rajashankar, K. R.; Crowley, P. J.; Kelly, C.; Mitchell, T. J.; Brady, L. J.; Deivanayagam, C. Crystal structure of the c-terminal region of Streptococcus mutans antigen

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I/II and characterization of salivary agglutinin adherence domains. J. Biol. Chem. 2011, 286, 21657–21666.

[24] Larson, M. R.; Rajashankar, K. R.; Patel, M. H.; Robinette, R. A.; Crowley, P. J.; Michalek, S.;

Brady, L. J.; Deivanayagam, C. Elongated fibrillar structure of a streptococcal adhesin assembled by the high-affinity association of alpha- and PPII-helices. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 5983–5988.

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ACKNOWLEDGEMENTS

141

Henk, Henny, since you operate as a team, I won’t even try to thank you separately. Thank you for giving

me to chance to continue the research I did in my Master as a PhD student. The monthly progress meetings

have been extremely valuable and significant in shaping my thesis. Your feedback and ideas about science

have been a great source of help. I have great admiration for the dedication both of you have towards

science.

Prashant, as my daily supervisor you have helped me to become an independent scientist. The fact that

our meetings became scarcer as the end of my PhD was approaching, for me was a sign of more

independence. Thank you for all your help at every stage of my PhD, the guidance and advice in the

beginning, and the freedom you have given me near the end.

To the reading committee, Prof. dr. A. Herrmann, Prof. dr. P. Buma and Prof. dr. L.W. van Rhijn, thank you

for taking the time to read and evaluate my thesis.

Ina, Willy, thank you for all your help with the administrative affairs. Joop, thank you for all the help doing

XPS, and even more so AFM. Without you I would not have been able finish all my experiments. Hans, your

skills in Matlab and programming have helped me tremendously when I was doing flow experiments and

analyzing biofilms, thank you for this help. Betsy, Theo, Babs, Roel, Jelmer, Gesinda, Rene, Jelly, Marianne,

Roel, Willem, thank you all for actively or passively helping in all the lab work.

Guru, as my master thesis supervisor, you introduced me to the lab and guided my initial start as a scientist,

I am very grateful for the time you took for me during that period and everything you learned me has

greatly helped me during my PhD. Eva, working with you during my master resulted in my first publication.

Thank you for all the scientific help, but also for showing me around Hoboken. Das, you introduced me to

the world of DNase I; a significant part of my thesis. I am grateful for all your time and help in this area,

but even more so for your friendship. Ed, this is where I would like to say thank you for helping me write

the review that serves as the introduction for my thesis, I could not have finished it in time without you.

Adam, as the experienced PhD student you were when we shared the office, I have learned a lot from you

about science, regarding doing experiments, but also about writing papers. Besides the science, even more

valuable for me was the trip to Sweden you organized with some of the people from the lab, including me.

I was honored to meet your family and stay at your house. Thank you for showing me this part of Sweden

and it’s traditions.

Brandon, it was great having someone in the office that is so passionate about science. I greatly enjoyed

the trip, you, my sister and me made to Orlando, for which you were kind enough to invite us. Exploring

the United States guided by an American was an unforgettable experience and besides your kindness, your

family was so generous to let us stay at their house. Thank you for this, and also for the honor of asking

me as a paranymph for your defense.

ACKNOWLEDGEMENTS

142

Sara, Rebecca, it has been great to share the office with the two of you. Thank you for making it a pleasure

to be at work and even more for all the good times outside of the department.

Deepak, Agnieszka, Ed, Philipp, Stefan, Otto, Katya thank you all for the daily fun in, and even more so,

outside of the department. Lunch, coffee breaks, and labwork would not have been as much fun without

all of you. Not to mention the drinks, dinners, parties, etc. Thank you all!

Deepak, Ed, thank you for your willingness to be my paranymphs and help me organize this important day

for me.

To everyone else in the department I have not mentioned, I would like to say thank you for your help,

discussions, talks, company and presence; Ferdi, Brian, Anna, Song, Mark, Jesse, Steven, Hilde, Joana, Yun,

Helen, Bu, Rene, Niar.

Barbara, your endless love and support means the world to me and I cannot express how grateful I am to

you.

Oma, Grada en Gerhard, bedankt voor jullie oneindige interesse in mijn werk en publicaties, ook al waren

die laatste meestal moeilijk te begrijpen.

To the most important people in my live, my parents and sister; your endless and unconditional support

has made it possible for me to achieve all of this. Your presence and interest in my work have been my

greatest source of motivation. Thank you for always being there for me.

ACKNOWLEDGEMENTS

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University of Groningen Innovative coatings for anti-bacterial … · 2016-03-09 · developments in surface coating technology permit controlled localized release which decreases

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