University of Groningen Adaptive antimicrobial nanocarriers for the control of infectious biofilms Liu, Yong

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(1)University of Groningen. Adaptive antimicrobial nanocarriers for the control of infectious biofilms Liu, Yong. IMPORTANT NOTE: You are advised to consult the publisher's version (publisher's PDF) if you wish to cite from it. Please check the document version below.. Document Version Publisher's PDF, also known as Version of record. Publication date: 2019 Link to publication in University of Groningen/UMCG research database. Citation for published version (APA): Liu, Y. (2019). Adaptive antimicrobial nanocarriers for the control of infectious biofilms. University of Groningen.. Copyright Other than for strictly personal use, it is not permitted to download or to forward/distribute the text or part of it without the consent of the author(s) and/or copyright holder(s), unless the work is under an open content license (like Creative Commons). Take-down policy If you believe that this document breaches copyright please contact us providing details, and we will remove access to the work immediately and investigate your claim.. Downloaded from the University of Groningen/UMCG research database (Pure): http://www.rug.nl/research/portal. For technical reasons the number of authors shown on this cover page is limited to 10 maximum.. Download date: 27-06-2021.

(2) Chapter 4. CHAPTER 4 Nanocarriers with Conjugated Antimicrobials to Eradicate Pathogenic Biofilms Evaluated in Murine In Vivo and Human Ex Vivo Infection Models. 4. Liu, Y. Ren, Y. Li, Y. Su, L. Zhang, Y. Huang, F. Liu, J. Liu, J. van Kooten, T. G. An, Y. Shi, L. van der Mei, H. C. Busscher, H. J. Acta. Biomateria., 2018, 79, 331–343. Reproduced with permission from Elsevier.. 71.

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(4) Chapter 4. ABSTRACT Conventional antimicrobials are becoming increasingly ineffective for treating bacterial infection due to the emergence of multi-drug resistant (MDR) pathogens. In addition, the biofilm-mode-of-growth of infecting bacteria impedes antimicrobial penetration in biofilms. Here, we report on poly(ethylene) glycol-poly(β-amino esters) (PEG-PAE) micelles with conjugated antimicrobials, that can uniquely penetrate biofilms, target themselves to bacterial cell surfaces once inside the low-pH environment of a biofilm and release conjugated antimicrobials through degradation of their ester-linkage with PAE by bacterial lipases. In vitro, PEGPAE micelles with conjugated Triclosan (PEG-PAE-Triclosan) yielded no inadvertent leakage of their antimicrobial cargo and better killing of MDR Staphylococcus aureus, Escherichia coli and oral streptococcal biofilms than Triclosan in solution. In mice, PEG-PAE-Triclosan micelles with conjugated Triclosan yielded better eradication efficacy towards a MDR S. aureus-infection compared with Triclosan in solution and Triclosan-loaded micelles at equal Triclosan-equivalent concentrations. Ex vivo exposure of multi-species oral biofilms collected from orthodontic patients to PEG-PAE-Triclosan micelles, demonstrated effective bacterial killing at 30-40 fold lower Triclosan-equivalent concentrations than achieved by Triclosan in solution. Importantly, Streptococcus mutans, the main causative organism of dental caries, was preferentially killed by PEG-PAE-Triclosan micelles. Thus PEG-PAE-Triclosan micelles present a promising addendum to the decreasing armamentarium available to combat infection in diverse sites of the body. Statement of Significance pH-adaptive polymeric micelles with conjugated antimicrobials can efficiently eradicate infectious biofilms from diverse body sites in mice and men. An antimicrobial was conjugated through an ester-linkage to a poly(ethylene glycol) (PEG)/poly(β-amino ester) block copolymer to create micellar nanocarriers. Stable micelle structures were formed by the hydrophobic poly(β-amino ester) inner core and a hydrophilic PEG outer shell. Thus formed PEG-PAE-Triclosan micelles do not lose their antimicrobial cargo underway to an infection site through the blood circulation, but penetrate and accumulate in biofilms to release antimicrobials once inside a biofilm through degradation of its ester-linkage by bacterial lipases, to kill biofilm-embedded bacteria at lower antimicrobial concentrations than when applied in solution. PEG-PAE-Triclosan micelles effectively eradicate biofilms of multi-drug-resistant pathogens and oral bacteria, most notably highly cariogenic Streptococcus mutans, in mice and men respectively, and possess excellent clinical translation possibilities.. KEYWORDS: biofilm, multi-drug resistance, enzymatic pH-adaptive, polymeric micelle, infection model. 73. 4.

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(6) Chapter 4. INTRODUCTION Most bacterial infections are due to pathogens in their biofilm-mode of growth, in which bacteria grow embedded in their self-produced matrix of extracellular polymeric substances.1,2 On top of the antimicrobial resistance of biofilms due to low penetrability of many antimicrobials into biofilms, the armamentarium of conventional antimicrobials presently available to control bacterial infections is rapidly shrinking due to the emergence of intrinsic, antimicrobial resistance amongst pathogens.3–5 Nanotechnology has contributed in recent years to combat bacterial biofilms, although many of its potential assets have not yet been translated to clinical application.6–8 Micellar nanocarriers have been developed that can be loaded with antimicrobials9 and possessing unique trades, such as pH adaptivity to self-target to bacteria in the acidic environment of a biofilm,10,11 enhanced penetration and accumulation in a biofilm,12 antimicrobialrelease in response to bacterial enzymes breaking down the micellar carriers,13,14 or inducing direct cell lysis owing to their strong positive surface charge.15,16 Antimicrobials are often encapsulated in nanocarriers through relatively weak hydrophobic interactions, causing drug leakage during storage17 or circulation in blood underway to their target that can cause severe side effects.18 As an alternative to weak hydrophobic binding inside micelles, antimicrobials can also be conjugated rather than loaded in micellar nanocarriers through a biodegradable linkage, as also done with various chemotherapeutics.19,20 Micellar nanocarriers with antimicrobials conjugated through a linkage that would degrade under the influence of bacterial enzymes, will not inadvertently lose their cargo underway to an infection site in the human body. However, nanoengineered micelles with antimicrobials conjugated through a biodegradable linkage have not yet been made nor evaluated for their use in infection control. Poly(ethylene)glycol-poly(β-amino esters) (PEG-PAE) micelles are biologically invisible due to the PEG moieties and negative charge at physiological pH values, making them ideal for transportation of antimicrobials through the blood stream and penetration into a biofilm. Moreover, once inside the low pH environment of a biofilm, PAE becomes exposed and positively-charged to self-target the micelles to negatively-charged bacterial cell surfaces.21–24 Because of these properties of PEG-PAE micellar nanocarriers, we chose PEG-PAE micelles to build an antimicrobial in through a biodegradable ester-linkage, with the aim of making a PEG-PAE micellar nanocarrier exhibiting negligible inadvertent antimicrobial release. Aiming for applicability towards infection control in widely diverse sites of the human body, PEG-PAETriclosan micelles with conjugated Triclosan as an antimicrobial, will first be evaluated for their ability to penetrate and kill multi-drug resistant Staphylococcus aureus, Escherichia coli or oral streptococcal biofilms in vitro (see Table S1). Staphylococci and E. coli occur in a plethora of infections across the human body25,26 and accordingly in vivo efficacy of PEG-PAE-Triclosan micelles in eradicating a sub-cutaneous, multi-drug resistant S. aureus infection in mice was evaluated. PEG-PAE-Triclosan micelles were also evaluated with respect to their ability to eradicate human ex vivo, multi-species oral biofilm harvested from orthodontic patients. Increasing occurrence of orthodontic biofilms in a growing patient population presents an enormous burden to the healthcare system,27,28 as they can cause carious lesions around brackets on tooth surfaces that persist after treatment29,30 and therewith interfere with the functional and esthetic repair achieved. Triclosan was chosen as an antimicrobial because it is not only a common oral antimicrobial in toothpastes and mouthwashes,31 but it is also applied in antimicrobial sutures32 and other clinical infection control measures.33 However, the methodology designed allows to synthesize micelles with other conjugated antimicrobials through conjugation with PAE, provided they carry functional groups, that can be modified with acryloyl groups for further polymerization, such as rifampicin and quinolones.. 75. 4.

(7) Chapter 4. EXPERIMENTAL SECTION. 4. Conjugation of Triclosan to Poly(Ethylene)glycol-Poly(β-Amino Ester). Amphiphilic Triclosan-conjugated PAE was synthesized through a one-pot Michael addition (Figure 1A). Key-monomer is a Triclosan containing diacrylate that will react with acrylate-terminated poly(ethylene glycol) (PEG) and 4,4’-trimethylene dipiperidine (TDP) through Michael addition to form antimicrobially conjugated poly(ethylene)glycol-poly(β-amino esters) (Figure 1B) in which PEG serves to stabilize the micelles and enhance their biocompatibility. The key-monomer was synthesized from commercially available 2,2-bis(hydroxymethyl) propionic acid through two sequential esterification steps. Step 1 in the synthesis of the key-monomer was the synthesis of 3-(acryloyloxy)-2-((acryloyloxy)methyl)-2-methylpropanoic acid. To this end, 2,2-bis(hydroxymethyl) propionic acid (10.1 g, 75 mmol, 1.0 equivalent) was dissolved in anhydrous dichloromethane in a flame-dried round-bottomed flask, while cooling on ice. Subsequently, triethylamine (Et3N, 15.2 g, 150 mmol, 2.0 equivalent) was added dropwise followed by acryloyl chloride (13.64 g, 150 mmol, 2.0 equivalent) within 1.5 h. The resulting solution was stirred for 0.5 h in an ice bath and filtered. The filtrate was concentrated by removal of the solvent under vacuum and the residue was dissolved in 50 mL Na2CO3 aqueous solution (10% w/v). Hydrochloric acid (HCl, 6 M) was then dropped in under vigorous stirring, until the pH reached 2.0. Finally, dichloromethane (3 × 50 mL) was added. The dichloromethane solution was then concentrated by removal of the solvent under vacuum. The crude product was recrystallized with ethyl acetate/hexane mixed solvent to yield 2,2-bis(acryloxymethyl)propionic acid, white crystals (16.8 g, yield: 92%). The purity of the product was confirmed by 1H and 13C NMR spectroscopy (Figure S1). As the second step in the preparation of the key-monomer, Triclosan (2.89 g, 10 mmol, 1.0 equivalent), 2,2-bis(acryloxymethyl)propionic acid (3.15 g, 13 mmol, 1.3 equivalent), 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC•HCl) (2.49 g, 13 mmol), and 4-(N,N'-dimethylamino)pyridine (DMAP) (1.59 g, 13 mmol, 1.3 equivalent) were dissolved in methylene chloride (200 mL) and the resulting solution stirred for 12 h at room temperature under nitrogen atmosphere. The resulting solution was diluted by adding 200 mL methylene chloride, and washed with saturated Na2CO3 (1 × 100 mL) and a diluted HCl solution (1 × 100 mL), respectively. After drying over MgSO4, filtration and concentration under vacuum, the crude product was purified by silica gel column chromatography using petroleum ether/ethyl acetate (5:1) as an eluent to yield the key-monomer as a colorless oil (4.6 g, yield: 89%). The composition of the key-monomer was confirmed by 1H and 13C NMR spectroscopy (Figure S2). In order to conjugate Triclosan to PEG-PAE, first, PEG5k-OH was turned into PEG monoacrylate by reacting with acryloyl chloride at low temperature in the presence of trimethylamine, as an acid-binding agent.23 Subsequently, we investigated the polymerization of PEG, TDP and the key-monomer with different Triclosan molar feed ratios (Fig. 1C). PEG monoacrylate (1.0 equivalent) was added to the key-monomer (5.0 equivalent or 10.0 equivalent), 4,4′-trimethylene dipiperidine (TDP, 5.05 or 10.1 equivalent) in chloroform and stirred at 55°C under nitrogen for 3 days. The resulting mixture was precipitated into ice-cold (-20°C) dimethyl ether to yield PEG-PAE with either five (denoted PEG-PAE-T5) or eight (PEG-PAE-T8) blocks of conjugated Triclosan (for 1H NMR spectrum, see Figure S3), corresponding with a drug loading content of 20 wt% and 27 wt%, respectively (Figure 1C). Matrix assisted laser desorption/ionization time of flight (MALDI-ToF) mass spectrometry (Figure S4) confirmed the mass-to-charge ratios (m/Z) were around 8603 and 10893 for PEG-PAE-T5 and PEG-PAE-T8, respectively. PEG-PAE-T5 and PEG-PAE-T5 polymers showed low critical micelle concentrations of 1.02 μg mL –1 and 1.07 μg mL –1 in 10 mM potassium phosphate, respectively (Figure S5). PEG-PAE with different numbers (x) of conjugated Triclosan blocks are indicated as PEG-PAE-Tx in the forthcoming parts of this paper. Preparation of PEG-PAE-Tx Micelles. For the preparation of polymeric micelle suspensions with conjugated Triclosan, 5 mg mL –1 stock solutions of PEG-PAE-Tx in dimethyl sulphoxide (DMSO) were 76.

(8) Chapter 4. 4. Figure 1. Schematics of the synthesis of poly(ethylene)glycol-poly(β-amino ester) with a conjugated antimicrobial. (A) Synthesis of a Triclosan containing diacrylate (key-monomer). (B) Self-assembly of pH-adaptive (panel 1) PEG-PAE-Tx micelles with Triclosan-conjugated and lipase-triggered (panel 2) Triclosan release and bacterial killing. (C) PEG-PAE-Tx synthesized in this study. M1, M2, M3 refer to PEG-A, TDP, and key-monomer, respectively, while output ratio was determined by 1H NMR spectroscopy. Tx indicates the amount of Triclosan conjugated relative to PEG-A. Drug loading content (DLC) in weight percentages was calculated as 100% × (weight of loaded drug)/(weight of polymer). 77.

(9) Chapter 4. prepared. For PEG-PAE-Tx micelle preparation, 2 mL PEG-PAE-Tx stock solution was added dropwise into 7 mL ultrapure water at a rate of 1 droplet (20 μL) per 20 s under magnetic stirring for 4 h to form a micelle suspension. The micelle suspension was dialyzed in a dialysis bag with molecular weight cut-off of 6–8 kDa against ultrapure water for 48 h to remove DMSO. The dialyzed micelle suspension was diluted to a final concentration of 0.5 mg mL−1 and stored in a refrigerator at 4°C. Nile red-loaded micelles were essentially prepared as described above, the only difference being that 2 mL PEG-PAE-Tx solutions in DMSO was first mixed with a Nile red solution in DMSO (0.5 mg mL –1, 80 μL) prior to micelle preparation. In the resulting PEG-PAE-Tx/Nile red solution, the mass ratio of Nile red to PEGPAE-Tx was kept at 0.4 wt%. Then, the mixed PEG-PAE/Nile red solution was added to water, followed by dialysis to remove DMSO and subsequently used to form Nile red-loaded micelles.. 4. Coating of Micelles with Salivary Proteins. For selected experiments relevant to application of the micelles in the oral cavity, micelles were exposed to salivary proteins under shaking (60 rpm) for 24 h in 5 mL reconstituted human whole saliva to a final micelle concentration in saliva of 0.1 mg mL –1. Reconstituted human whole saliva was prepared from a stock of human whole saliva from at least 20 healthy volunteers of both genders, collected into ice-cooled beakers after stimulation by chewing Parafilm®,34 and subsequently pooled, centrifuged, dialyzed and lyophilized for storage. Prior to lyophilization, phenylmethyl sulfonyl fluoride was added to a final concentration of 1 mM as a protease inhibitor in order to reduce protein breakdown. Freeze-dried saliva (1.5 mg mL –1) was dissolved in a saliva buffer (1 mM K2HPO4, 1 mM KH2PO4, 50 mM KCl and 1 mM CaCl2, pH 7.0) and will be referred to in this paper as “saliva”. All volunteers gave their informed consent to saliva donation, in agreement with the guidelines set out by the Medical Ethical Committee at University Medical Center Groningen, Groningen, The Netherlands (letter 06-02-2009). Micelle Characterizations. Zeta potentials of the micelles were measured at 25°C using a Zetasizer Nano-ZS (Malvern Instruments, Worcestershire, UK) with a DTS1060-type cuvette in 10 mM potassium phosphate or saliva buffer over the pH range from 5.0 to 7.4. In addition, hydrodynamic diameters of the micelles were measured using the same instrument. UV-VIS absorption spectra were measured using an UV− VIS spectrophotometer (Shimadzu, Tokyo, Japan) for Triclosan detection. All experiments were done at a micelle concentration of 0.1 mg mL –1. Triclosan-Release by PEG-PAE-Tx Micelles. To determine the release of Triclosan from Triclosanconjugated PEG-PAE-Tx micelles, 2 mL of freshly prepared micelle suspensions (1.0 mg mL−1) was transferred into a dialysis bag (molecular weight cut off: 12−14 kDa) and subsequently immersed in 20 mL of a 10 mM phosphate buffer (pH 7.4) at 37°C. Aliquots (1 mL) of the dialysis solution were collected every 30 min up to 72 h, and the absorbance of the solutions at 281 nm was recorded on a UV−VIS spectrophotometer. The volume of the stock dialysis solution was kept constant by adding 1 mL of fresh buffer after each aliquot was taken. A key assumption in the current work is that bacterial enzymes, such as lipase, can hydrolyze the ester-linkage between Triclosan and PAE chains to release the conjugated antimicrobial. To demonstrate this mechanism, lipase-triggered release of Triclosan from PEG-PAE-T5 and PEG-PAE-T8 micelles (1 mg mL−1) was studied. Lipase (Sigma, from Pseudomonas cepacia, ≥30 U mg−1) was added to micelle suspension to a final enzyme concentration of 0.5 mg mL−1 and Triclosan release was measured as described above. Bacterial Culturing and Harvesting. All strains employed (see Table S1) were grown from cryopreservative beads (Protect Technical Surface Consultants Ltd., UK) onto agar plates, with or without antibiotic supplementation (see also Table S1) at 37°C in ambient air, except for streptococci which were grown under 5% CO2. For experiments, one colony was transferred to inoculate 10 mL of liquid growth medium (see Table S1) for 24 h. This pre-culture was subsequently diluted 1:20 in 200 mL liquid medium 78.

(10) Chapter 4. and grown statically for 16 h at 37°C. Staphylococci and E. coli cultures were harvested by centrifugation for 5 min at 5000 g, washed twice in phosphate buffered saline (PBS, 5 mM K2HPO4, 5 mM KH2PO4 and 150 mM NaCl, pH 7.0), sonicated for 3 × 10 s (Vibra cell model 375, Sonics and Material Inc., Danbury, CT) while cooling in an ice/water bath to break possible aggregates and finally suspended in 10 mL PBS to a concentration of 3 × 108 bacteria mL –1, as determined in a Bürker-Türk counting chamber. Streptococci were washed and suspended as described above, but in saliva buffer. Minimal inhibitory concentrations (MIC) of the staphylococci and E. coli used in this study against a selection of common antibiotics are given in Table S2, while MIC values of all strains used against Triclosan are presented in Table S3. Interaction of Nile Red-Loaded Micelles with Staphylococcal and Streptococcal Biofilms. In order to study the interaction of Nile red-loaded Triclosan-conjugated PEG-PAE-Tx micelles with staphylococcal or streptococcal biofilms, 500 μL bacterial suspension (108 bacteria mL –1) was pipetted into a 6-well-plate to allow adhesion for 1 h at 37°C. Next the suspension was discarded and the wells were washed once with 500 μL PBS or saliva buffer to remove planktonic bacteria and 2 mL appropriate liquid medium (see Table S1) was added to each well and incubated for biofilm growth during a total 48 h at 37°C. After 24 h, the medium was replaced with fresh one and the biofilm was left to grow for another 24 h. Finally, wells with biofilm attached were washed three times with PBS or saliva buffer and the biofilms were exposed to the appropriate buffer with pH ranges from 5.0 to 7.4 containing Nile red-loaded PEG-PAE-Tx micelles (0.1 mg mL –1). The penetration of the micelles into fluorescent staphylococcal biofilms was subsequently studied with CLSM (Leica TCS SP2 Leica, Wetzlar, Germany) with a HCX APO L40×/0.80 W U-V-1 objective. An argon ion laser at 488 nm and a green HeNe laser were used to excite the GFP and Nile red in the biofilms, respectively and fluorescence was collected at 500 – 535 nm (GFP) and 583 – 688 nm (Nile red). All data were acquired and analyzed using Leica software, version 2.0 and ImageJ software. To study penetration in streptococcal biofilms, biofilms were stained using SYTO 9 (Thermo Fisher Scientific, Waltham, Massachusetts, USA) for 15 min in the dark and subsequently studied with CLSM under the same excitation conditions as described for staphylococci, but collecting fluorescence at 500 – 540 nm (SYTO 9) and 583 – 688 nm (Nile red). Bacterial Killing in Biofilms In Vitro. In order to determine the killing efficacy of Triclosan in solution or conjugated in to PEG-PAE-Tx micelles, biofilms were grown by adding 100 μL of bacterial suspension (108 bacteria mL –1) either in PBS (S. aureus or E. coli) or in saliva buffer (streptococci) to 96-wells plates at 37°C for 1 h to allow bacteria to adhere. Next, bacterial suspensions were removed and the wells were washed with 100 μL of the appropriate buffer. Subsequently, 200 μL of the appropriate medium (see Table S1) was added and bacteria were allowed to grow for 48 h at 37ºC. Next, 100 μL solutions of Triclosan dissolved or conjugated in to PEG-PAE-Tx micelles either in 10 mM potassium phosphate or saliva buffer (pH 7.4) were added to the biofilms under shaking at 60 rpm. Exposure to either form of Triclosan for S. aureus and E. coli biofilms (Triclosan-equivalent concentration: 0.625, 20, 80 μg mL –1), and 2 min for streptococcal biofilms (Triclosanequivalent concentration: 100 and 200 μg mL –1), followed by retention, i.e. exposure of the biofilm to 300 μL saliva buffer for different periods of time at 37°C in a CO2 incubator in order to determine whether or not Triclosan was substantively present in the streptococcal biofilms, as required for oral applications. For evaluation of bacterial killing in biofilms of S. aureus Xen36 and E. coli Xen14, bioluminescence images were taken (IVIS® Lumina II, Imaging System, Perkin Elmer) every hour up to 5 h, starting 2 h after exposure to the different forms of Triclosan (image acquisition factors: 20 s exposure time, medium binning, 1 F/Stop, open emission filter). Images were automatically corrected for background noise. Regions of interest (ROIs) were manually defined for each well and average radiances over the ROIs were converted to photon fluxes (p/ s) using Living Image software (Perkin Elmer). For evaluation of bacterial killing in streptococcal biofilms, biofilms were stained with 200 μL solution of Live/Dead BacLightTM Bacterial Viability Kit (Thermo Fisher 79. 4.

(11) Chapter 4. Scientific, Waltham, Massachusetts, USA) for 15 min in the dark, and images recorded with CLSM (see above). An argon ion laser at 488 nm and a green HeNe laser were used to excite the SYTO 9 (live bacteria) and propidium iodide (dead bacteria) in the samples, respectively and fluorescence was collected at 500 – 535 nm (SYTO 9) and 580 – 630 nm (propidium iodide). All data were acquired and analyzed using Leica software, version 2.0 and ImageJ software.. 4. In Vivo Eradication of A Sub-Cutaneous, Multi-Drug Resistant S. aureus Infection in Mice. BALB/c nude mice (15 g to 16 g each) were obtained from Vital River Laboratory Animal Technology Co. (Beijing, China). All animals were housed in the on-site animal facility of Nankai University and experimental procedures were approved by the Institutional Animal Care and User Committee of Nankai University, Tianjin, China (Certificate number: SYXK(TIANJIN)2014-00003). Two sub-cutaneous infection sites were created in each mouse by injecting a dose (50 μL) of 2 × 109 bioluminescent S. aureus Xen36 in the left and right flanks of the mice to initiate infection. Bioluminescent intensity and area were recorded using a biooptical imaging system (IVIS®, 45 s exposure time, medium binning, 1 F/Stop, Open Emission Filter). Images were analyzed using “Image J” (NIH Research Services Branch, USA) software. Since bioluminescence reached a maximum at day 2 after injection, infected animals were randomly assigned after two days into four groups of six animals each, receiving every two days, an intravenous injection of (i) 200 μL PBS (untreated control), (ii) 200 μL Triclosan in PBS (0.8 mg mL –1), (iii) 200 μL Triclosan-loaded PEG-PAE micelles (4 mg mL –1 micelle concentration, drug loading content: 20%, see 23) or (iv) 200 μL Triclosan-conjugated PEGPAE-T5 micelles (4 mg mL –1 micelle concentration, drug loading content: 20%, see Fig. 1C). Accordingly, all treatments, except the PBS control were done at a Triclosan-equivalent dose of 160 μg. Treatment was initiated 2 days post-infection and continued for 5 consecutive days. Bioluminescent imaging was performed daily. For histopathology, infected tissue was aseptically dissected and fixed overnight at 4°C in 10% formalin. Tissue samples were dehydrated using a graded ethanol series from 70–100%, cleared with xylene to remove the dehydrate, and infiltrated with paraffin. Processed tissue was embedded in paraffin, cut in 5 μm sections, and placed on microscope slides and stained with hematoxylin/eosin and Gram-stain. Tissue samples were examined on a CX41 microscope (Olympus, Tokyo, Japan). Efficacy of PEG-PAE-T5 Micelles towards Human Ex Vivo Orthodontic Biofilms. Human ex vivo oral biofilm samples with an approximate equal volume of 0.5 mm3 were collected from 12 healthy, orthodontic adolescent patients during a routine visit to the orthodontic clinic, and without having obtained specific oral health care instructions. Biofilm was collected using a periodontal probe from the area surrounding the brackets on the buccal tooth surfaces in both arches, excluding molars. Three biofilm samples were obtained from each patient that were randomly assigned to three treatments (exposure to saliva buffer, Triclosan in solution or conjugated in to micelles). The study was performed according to the guidelines of the Medical Ethics Committee of the University Medical Center Groningen, Groningen, The Netherlands (letter 17-07-2017), including the signed informed consent by the patients and according to the tenets of the Declaration of Helsinki. Each biofilm sample was immersed in 300 μL saliva buffer in 1.5 mL sterilized Eppendorf tubes for 2 min exposure with Triclosan dissolved in saliva buffer or conjugated PEGPAE-T5 micelles suspended in saliva buffer. All experiments were done at 200 μg mL –1 Triclosan-equivalent concentrations, taking saliva buffer as a control. After 2 min exposure, substantive action of the Triclosan as required for effective intra-oral killing,35 was examined by placing the biofilm samples in 0.5 mL saliva buffer for 3 h at 37°C. After the entire treatment procedure, biofilm was dispersed by sonicating for 3 × 10 s (Vibra cell model 375, Sonics and Material Inc., Danbury, CT), while cooling in an ice/water bath after which total bacterial numbers (live and dead) in ex vivo orthodontic biofilm samples microscopically were examined in a Bürker80.

(12) Chapter 4. 4. Figure 2. Characterization of poly(ethylene)glycol-poly(β-amino ester) micelles with conjugated Triclosan carried out at 25°C. (A) UV-VIS absorption as a function of wavelength of PEG-PAETx micelles with conjugated Triclosan and unloaded PEG-PAE micelles. The 281 nm absorption peak represents conjugated Triclosan. (B) Cumulative release of Triclosan, ester-linked in PEGPAE-Tx micelles as a function of time during suspension of micelles in absence or presence of lipase, measured using UV-VIS absorption spectroscopy. Data are expressed as mean ± SD over 3 experiments with separately prepared micelles, relative to the initial amount of Triclosan conjugated. (C) Transmission electron micrographs of PEG-PAE-Tx micelles with conjugated Triclosan. Scale bars indicate 200 nm. (D) Example of the diameter distribution of PEG-PAETx micelles in a single experiment measured using dynamic light scattering at different pHs. (E) Diameter as a function of pH for PEG-PAE-Tx micelles measured using dynamic light scattering. Data are expressed as mean ± SD over 3 experiments with separately prepared micelles. (F) Same as (E), now for micelles in saliva buffer. (G) Zeta potentials of PEG-PAE-Tx micelles (micelle concentration of 0.1 mg mL –1) as a function of pH. Data are expressed as mean ± SD over 3 experiments with separately prepared micelles. (H) Same as (G), now for micelles in saliva buffer.. 81.

(13) Chapter 4. Türk counting chamber. Part of the dispersed biofilm samples were also live/dead stained (see above) to determine the total number of red and green fluorescent pixels using CLSM (see above), allowing to calculate the percentage viability of the biofilm as the ratio between the number of green fluorescent pixels over the sum of the numbers of green and red ones. Finally, the number of total bacterial CFUs or S. mutans CFUs in the biofilms were determined on blood agar plates or S. mutans selective agar, respectively. To this end, a 10 μL aliquot of the bacterial dispersion was serially diluted and 100 μL of each dilution plated on blood agar (Tryptic Soy Agar with 5% Sheep Blood) plates or S. mutans selective growing TSY20B agar plates, respectively. All plates were incubated at 37°C in an anaerobic incubator for 4 days, followed by 24 h growth at ambient temperature after which the number of total bacterial or S. mutans CFUs were counted. Statistical Methods. All data are expressed as means ± standard deviations (SD). Differences between groups were examined for statistical significance with two-tailed Student’s t test, accepting significance at p < 0.05.. RESULTS. 4. Characterization of PEG-PAE-Triclosan Micelles. UV-VIS spectra of PEG-PAE-Tx micelles with conjugated Triclosan suspended in potassium phosphate buffer showed an absorption peak at 281 nm due to Triclosan that was slightly higher for PEG-PAE-T8 than for PEG-PAE-T5, confirming that the amount of Triclosan conjugated in PEG-PAE-T8 is higher than in PEG-T5 (Figure 2A). Cumulative release of Triclosan from both PEG-PAE-T5 as well as from PEG-PAE-T8 micelles could not be demonstrated up to 72 h in absence of lipases, indicating strong conjugation and absence of inadvertent release (Figure 2B). In the presence of lipase however, more than 50% and 63% of the Triclosan-conjugated PEG-PAE-T5 and PEGPAE-T8 micelles, respectively was released within 72 h. TEM micrographs showed a uniform, spherical morphology of the micelles (Figure 2C) with an average diameter of around 100 nm, regardless of the number of Triclosan blocks conjugated. PEG-PAE-T8 micelles appeared more compact and had darker micelle cores in TEM micrographs than PEG-PAE-T5 micelles, because of their higher amount of hydrophobic Triclosan conjugated. Micelle diameters determined by DLS had a narrow distribution (Figure 2D) and were virtually independent of the ionic composition of the suspension fluid (compare Figures 2E and 2F). Exposure of PEG-PAE micelles to a 10% murine plasma solution or reconstituted saliva only slightly increased the micellar diameter to 148 and 143 nm at pH 5.0 respectively,36 but not at higher pH values (Figure S6A). The ionic composition of the suspension fluid affected the pH dependence of the micellar zeta potentials in different ways, when in 10 mM potassium phosphate buffer or in saliva buffer (compare Figures 2G and 2H), due to double-layer condensation in the presence of large amounts of calcium and phosphate ions in the saliva buffer. Zeta potentials of the negatively charged PEG-PAE-Tx micelles became positive up to 10 mV in a potassium phosphate buffer, when the surrounding pH dropped below pH 7 (Figure 2G). In a saliva buffer (Figure 2H), micelles also became positively charged at acidic pH values, but over the entire pH trajectory zeta potentials hovered around zero between –2.2 and +1.4 mV due to double-layer condensation. Exposure of PEG-PAE to a plasma protein solution36 or artificial saliva (Figure S6B) did not significantly affect the zeta potentials of the micelles in the respective buffers. Penetration, Accumulation and Retention of PEG-PAE-Tx Micelles in Biofilms. In order to gain insight in the interaction between PEG-PAE-Tx micelles and bacteria during penetration, accumulation and retention in biofilms, Nile red-loaded PEG-PAE-Tx micelles were prepared. Nile red-loaded micelles possessed similar pH dependent hydrodynamic diameters (compare Figures 2E and S7A) and zeta potentials (compare Figures 2G and S7B) in phosphate buffer as in absence of Nile red-loading. CLSM showed that after 2 h exposure of green-fluorescent S. aureus WHGFP biofilms to PEG-PAE-Tx micelles in potassium phosphate buffer, micelles penetrated and accumulated in biofilms only at pH 5.0, but not at pH 7.0 (Figures 3A and 3B). At higher pH values, both micelles (Figure S7B) and staphylococci are negatively charged, causing 82.

(14) Chapter 4. 4 Figure 3. pH-dependent penetration, accumulation and retention in staphylococcal biofilms of fluorescent, Nile red-loaded, PEG-PAE-Tx micelles (micelle concentration 0.1 mg mL –1). (A) CLSM cross-sectional images of an S. aureus WHGFP biofilm after 2 h exposure to a suspension of PEG-PAE-T5 micelles in phosphate buffer. Bar marker equals 50 μm. (B) Percentage micelle accumulation in an S. aureus WHGFP biofilm after 2 h exposure to a suspensions of PEG-PAE-Tx micelles in phosphate buffer. Percentage accumulation is expressed with respect to the number of red pixels observed in a biofilm after 2 h exposure to Nile red-loaded micelles at pH 5.0. Error bars represent SD values over three experiments with differently grown biofilms and separately prepared micelles. (C) CLSM images of an S. mitis ATCC9811 biofilm after 2 h exposure to a suspension of PEG-PAE-T5 micelles in saliva buffer. Bar marker equals 50 μm. (D) Percentage Nile-red fluorescence intensity as a function of depth in the biofilm and micelle presence in S. mitis ATCC9811 biofilms as a function of exposure time to a suspension of PEG-PAE-T5 micelles in saliva buffer, pH 7.0. Percentage micelle presence (right panel; red line) is expressed with respect to the number of red pixels observed in a biofilm after 2 h exposure to micelles. Note that in separate experiments after 2 min exposure to a micelle suspension, a retention phase was initiated by replacing the suspension with saliva buffer without micelles (right panel; green line), as relevant for oral application requiring short exposure times and substantive presence of an antimicrobial for effective oral biofilm eradication.37 (E) Same as (C), now for an S. mutans ATCC700610 biofilm prior to (panel 1) and after exposure to saliva (panel 2). (F) Percentage micelle presence in S. mutans ATCC700610 biofilms as a function of time during exposure to a suspension of PEG-PAE-T5 micelles in saliva buffer, pH 7.0 (red lines) and retention in buffer without micelles (green lines). Percentage micelle presence is expressed with respect to the number of red pixels observed in a biofilm after 2 h exposure to micelles. 83.

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(17). Figure 4. Bacterial killing efficacies by Triclosan in solution and conjugated PEG-PAE-Tx micelles of multi-drug-resistant S. aureus Xen36, E. coli Xen14 and various oral streptococcal biofilms in vitro. (A) Schematics of the evaluation of the killing efficacy by Triclosan of S. aureus and E. coli biofilms and examples of bioluminescence images of S. aureus biofilms prior to and after exposure in pseudo-colors. (B) Bioluminescence intensities expressed relative to those of S. aureus and E. coli biofilms exposed to phosphate buffer after different exposure times to Triclosan at different Triclosan-equivalent concentrations. All data are expressed as means ± SD over triplicate experiments with separately prepared micelles and different staphylococcal and E. coli cultures. Asterisks indicate statistical significance at p < 0.05 (*), p < 0.01 (**) and p < 0.001 (***, Students’ t-test) between Triclosan in solution and micelles with Triclosan conjugated.. 84.

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(20). (C) Schematics of the evaluation of the killing efficacy by Triclosan of oral streptococcal biofilms and examples of fluorescent images of LIVE/DEAD stained S. mutans ATCC700610 biofilms prior to exposure to a micelle suspension and after 3 h of subsequent retention following 2 min exposure. (D) Percentage bacterial viability in S. salivarius HB, S. mitis ATCC9811 and S. mutans ATCC700610 biofilms after 1, 3 and 5 h of retention following 2 min exposure to Triclosan at different Triclosan-equivalent concentrations. Viability was expressed as the ratio between green fluorescent pixels relative to green and red fluorescent pixels. All data are expressed as means ± SD over triplicate experiments with separately prepared micelles and different streptococcal cultures. Asterisks indicate statistical significance at p < 0.0001 (****, Students’ t-test) between Triclosan in solution and Triclosan conjugated to micelles.. 85.

(21) Chapter 4. 4. Figure 5. Eradication of a sub-cutaneous, multi-drug resistant S. aureus Xen36 infection in mice using PEG-PAE-T5 micelles. (A) Schematics of the murine infection model, indicating the staphylococcal injection sites. All groups of mice were treated with the same Triclosanequivalent concentration of 800 μg mL−1 or PBS (0.2 mL, repeated every 48 h, starting at day 2 post infection) (B) Percentage survival of mice as a function of post-treatment time for differently treated groups of mice. Each group comprised six mice at the day of staphylococcal injection. (C) Body weight of mice as a function of post-treatment time. Error bars represent SD values over the surviving mice in each group. (D) Time series of bioluminescence images in the same mouse taken at different days post-treatment for differently treated mice. Bioluminescence intensity is given on a pseudo-color scale. (E) Percentage bioluminescence intensity arising from the infection site as a function of post-treatment time after initiating the different treatments. Bioluminescence intensity at day 0 was set at 100%. Error bars represent SD values over the surviving mice in each group. (F) Percentage area of the infected site as a function of posttreatment time. Infection site area at day 0 was set at 100%. Error bars represent SD values 86.

(22) Chapter 4. electrostatic repulsion and impeding their accumulation in the biofilm. At pH 5.0, staphylococci and micelles carry opposite charges and thus attract each other. Interestingly, in the absence of a sizeable negative or positive zeta potential of the micelles in saliva buffer, penetration and accumulation in streptococcal biofilms from a saliva buffer was near complete in 2 h (Figures 3C and 3E). This suggests that positively-charged micelles attracted from a potassium phosphate buffer into negatively-charged staphylococcal biofilms (Figure 3A, left panel) may block biofilm entry and limit further micelle entry and diffusion. Oppositely, uncharged micelles like in saliva buffer may freely enter and diffuse to deeper streptococcal biofilm layers (Figures 3C, 3D and 3E). PEG-PAE micelles have the ability to remain in the blood stream for several days before being secreted through the urinary pathway,36 but for oral applications time is a critical factor. Most oral therapeutics are present in the oral cavity no longer than 2 min, and their substantive presence requires adsorption to abundantly available mucosal surfaces38 or absorption in oral biofilm,4 followed by subsequent slow release. Therefore micellar penetration and accumulation was studied for two streptococcal biofilms as a function of exposure time. Penetration and accumulation reached a level of 20% to 50% of their maximum as achieved after 2 h, within 2 min exposure for S. mitis (Figure 3D) and S. mutans (Figure 3F, panel 1), respectively. Prior adsorption of salivary proteins to the micelles, as abundantly present in the oral cavity, did not affect penetration and accumulation (compare Figure 3F, panels 1 and 2). Wash-out of the micelles from streptococcal biofilms after 2 min exposure to micellar suspensions in saliva buffer, hardly occurred demonstrating their substantive presence in the streptococcal biofilms (see also Figures 3D and 3F). Eradication of Bacteria in Their Biofilm-Mode of Growth In Vitro. Eradication of multi-drug resistant S. aureus and E. coli was evaluated as described schematically in Figure 4A. However, first it was established that unloaded micelles did not kill any of the bacterial strains used for micellar concentrations up to 4 mg mL -1, indicating that bacterial killing observed is solely due to Triclosan release from the micelles and not by the polymeric micelles themselves. In general, Triclosan-conjugated PEG-PAE-Tx micelles were more efficacious against both strains than Triclosan in solution, especially at elevated Triclosan-equivalent concentrations (Figure 4B). PEG-PAE-T8 micelles had similar killing efficacies as PEG-PAE-T5 micelles at the same Triclosan concentration, and in our later experiments, we have chosen PEG-PAE-T5 to access their efficacies. Note that at the lowest Triclosan concentration, which is sub-MIC (see Table S3), and for the shortest exposure time, an increase in S. aureus bioluminescence was observed, similar to previous observations on bioluminescent staphylococci exposed to sub-MIC concentrations of antibiotics. This phenomenon was attributed to increased metabolic activity of the organisms under chemical stress.39 Evaluation of the eradication of oral streptococcal biofilms was done according to a slightly different protocol (see Figure 4C), to better match the short exposure times in oral cavity and to include the possibility to determine the importance of micelle retention in oral biofilms. All streptococcal biofilms prior to exposure to Triclosan in solution possessed 95% to 100% viability. Using Triclosan-equivalent concentrations 3040 times lower than the concentration of Triclosan in commercial toothpastes or mouthwashes (0.3% w/ w which equals 3 mg mL -1), it was observed that neither Triclosan in solution nor conjugated PEG-PAE-T5 micelles killed S. salivarius (Figure 4D), a member of the healthy oral microbiome,40 which is in line with its relatively high MIC against Triclosan compared with other oral bacterial strains (Table S3). The viability of S. mitis, another member of the healthy oral microbiome, and cariogenic S. mutans biofilms was hardly affected by Triclosan in solution (see also Figure 4D), in line with the resistance of oral biofilms to antimicrobial over the surviving mice in each group. (G) Micrographs of Gram-stained murine skin tissues from mice at day 5 post different treatments. Yellow arrows indicate biofilm imbedded in subcutaneous tissues. Scale bar equals 100 μm. 87. 4.

(23) Chapter 4. penetration and killing.41 However, PEG-PAE-T5 micelles with conjugated Triclosan clearly broke the biofilm barrier of these two streptococcal strains, as evidenced by an increasing loss in viability with increasing retention times. The largest reduction in biofilm viability down to 4% was observed for cariogenic S. mutans upon 2 min exposure to and subsequent 5 h retention of PEG-PAE-T5 micelles, in comparison with the significantly higher viability at 13% of S. mitis biofilms achieved under similar conditions. Importantly, also in vitro cytocompatibility studies human umbilical vein endothelial cells, human skin fibroblasts or human epithelial colorectal adenocarcinoma cells (Figure S8) co-cultured with PEG-PAE-Tx micelles was not negatively affected by micellar presence, as judged from the metabolic activity of the cells. In Vivo Eradication of a Sub-Cutaneous, Multi-Drug Resistant S. aureus Infection. Eradication of a multi-drug resistant S. aureus infection (see Table S2 for MICs) was studied in a sub-cutaneous murine infection model using bio-optical imaging,36 applying a dose of 2 × 109 bioluminescent S. aureus Xen36 in the left and right flanks of mice to initiate infection at two sites (Figure 5A). Bioluminescence of the infected sites reached a maximum 2 days after injecting the staphylococci as established before,36 at which time treatment was initiated (denoted as day 0 post-treatment).. 4. Two mice died on day 2 and 4, respectively in the group treated with PBS, while one mouse died on day 3 in the group treated with Triclosan in solution. All mice in the micelle treated groups survived (Figure 5B). Body weight of the mice showed little variation post-treatment (Figure 5C). Time series of bioluminescence images for one and the same mouse after initiating each of the different treatments are shown in Figure 5D, while quantitative analysis of the bioluminescence arising from the infected sites and the area of the infected site are presented in Figures 5E and 5F. PBS and Triclosan in solution yielded significantly less eradication of infection in terms of bioluminescence intensity and bioluminescent area than micelles loaded with Triclosan or with Triclosan conjugated. However, Triclosan conjugated to micelles was more effective than Triclosanloaded micelles, especially with regard to the decreasing area of the infected sites (Figure 5F). Histological images (Figures 5G, panel a and 5G, panel b) confirmed the presence of large numbers of staphylococci in tissue of mice sacrificed at day 5 after treatment with PBS or Triclosan in solution. Moreover, hematoxylin/ eosin staining clearly showed infection present in subdermal layers, but not going into the muscle layer and also infiltration of immune cells in more superficial layers of the skin tissue (Figures S9, panel a and S9, panel b). Small numbers of bacteria were seen in tissue of mice treated with Triclosan-loaded micelles (see also Figure 5G, panel c), including tissue damage and signs of immune response (Figure S9, panel c). In the tissue of mice treated with Triclosan-conjugated PEG-PAE-T5 micelles (Figure 5G, panel d), no bacteria nor infiltration of immune cells were observed in the tissue samples (Figure S9, panel d). Ex Vivo Killing of Bacteria of Oral Biofilm Collected from Orthodontic Patients. Bacterial killing in human ex vivo, multi-species oral biofilms from 12 orthodontic adolescent patients by Triclosan in solution and PEG-PAE-T5 micelles was evaluated as schematically presented in Figure 6A, taking saliva buffer as a control. Equal amounts of oral biofilm were collected from the area around orthodontic brackets in 12 patients and exposed for 2 min to Triclosan in 0.3 mL solution or suspension at a Triclosan-equivalent concentration of 200 μg mL –1, after which biofilm was placed in 0.3 mL saliva buffer for substantive action. Killing efficacy was evaluated using fluorescence microscopy after LIVE-DEAD staining of dispersed biofilms (Figure S10), while dispersed bacteria were also evaluated in a counting chamber for total counts and plated on selective agar for S. mutans enumeration. All biofilm samples collected contained a highly comparable total number of bacteria, demonstrating that nearly equal amounts of biofilm were collected clinically (Figure 6B). Furthermore, collection of oral biofilm in equal amounts and of similar viability and composition can be inferred from biofilm viability, total and S. mutans CFU counts after biofilm exposure to saliva buffer as a control (Figures 6B-6E). Neither the viability of the biofilm samples evaluated by LIVE-DEAD staining (Figure 6C), nor the total or S. mutans CFUs counts in biofilm samples (Figures 6D and 6E, respectively) 88.

(24) Chapter 4. Figure 6. Ex vivo killing of bacteria of oral biofilm collected from orthodontic patients. (A) Schematics of the evaluation of the killing efficacy in human ex vivo orthodontic biofilms of Triclosan in solution and PEG-PAE-T5 micelles. Biofilms were exposed to saliva buffer as a control. (B) Total bacterial number (live and dead) in ex vivo orthodontic biofilm samples after exposure to saliva buffer, Triclosan in solution or conjugated micelles. Error bars represent SD values over the 12 orthodontic biofilm samples in each group. (C) Percentage bacterial viability in orthodontic biofilm samples treated ex vivo. Bacterial viability was quantified from fluorescence micrographs of LIVE-DEAD stained dispersed biofilms (see Figure S10) as the ratio of red fluorescent pixels of the total number of red and green fluorescent pixels. Error bars represent SD values over the 12 orthodontic biofilm samples in each group. (D) Number of colonies formed on blood agar plates in ex vivo orthodontic biofilms after exposure to saliva buffer, Triclosan in solution or conjugated micelles. (E) Number of S. mutans on S. mutans selective agar in ex vivo orthodontic biofilms after exposure to saliva buffer, Triclosan in solution or conjugated micelles. Error bars represent SD values over the 12 orthodontic biofilm samples in each group. Asterisks indicate statistical significance at p < 0.05 (*), p < 0.01 (**, Students’ t-test) between saliva buffer, Triclosan in solution and micelles with Triclosan conjugated. were reduced by exposure to saliva buffer or to Triclosan in solution. However, Triclosan-conjugated PEGPAE-T5 micelles reduced biofilm viability to less than 50% (see also Figure 6C) and caused two-log units reduction in S. mutans CFU counts (Figure 6E). Total CFU counts remained the same, albeit showing a wider variations amongst biofilm collected from different patients (see also Figure 6D). This demonstrates that S. mutans is preferentially killed by Triclosan-conjugated PEG-PAE-T5, which is highly advantageous because selective killing of the major pathogens, amongst which cariogenic S. mutans, leaves a great number of species in oral biofilm (“the healthy microbiome”) alive to provide protection against environmental insults and maintain oral health.. DISCUSSION In this paper we describe the conjugation of an antimicrobial, Triclosan, with PAE in order to form a micelle system with an antimicrobial conjugated to eradicate biofilm-associated infections. This approach is opposite to loading of an antimicrobial into micelles, that often suffers from inadvertent leakage of the antimicrobial. 89. 4.

(25) Chapter 4. Conjugation to PAE through a biodegradable linkage with PAE ensures degradation of the linkage once inside a biofilm by bacterial lipases to cause far better localized accumulation and higher local concentrations of the antimicrobial than can be achieved by systemic administration. Triclosan was chosen as an antimicrobial as it is applied both in oral health care products as well as in many clinical infection control measures,42 but several other antimicrobials are candidate for this approach, provided they carry amino or hydroxyl groups that can be modified with acryloyl groups, like rifampicin and fluoroquinolones. The conjugation of Triclosan in micelles through a biodegradable linkage not only yields the advantage of zero leakage (Figure 2B), but due to the excellent penetration properties of PEG-PAE-T micelles into biofilms and the highly localized release of Triclosan inside the biofilm, bacterial killing and biofilm eradication occurs at significantly lower Triclosan concentrations than observed for Triclosan in solution (Figure 4). Considering the regularly voiced concerns stimulated by the use of Triclosan in non-health care related applications such as toys, furniture, and paints, its high efficacy at low concentrations when conjugated in to micelles, may provide new avenues to address environmental concerns.43 Note that also Triclosan-loaded micelles composed of alendronate conjugated to different Pluronic copolymers and possessing tooth-binding abilities, as opposed to our pH responsive, targetable Triclosan-conjugated micelles, demonstrated killing efficacy against single-species S. mutans biofilms in vitro. However, this was not due to lipase-induced Triclosan release as in our micelles, making selective killing of S. mutans in a multi-species biofilm unlikely and killing efficacy was not evaluated against multispecies oral biofilms generated in humans.44. 4. Important in the increased efficacy of Triclosan when conjugated in to PEG-PAE micelles is the complete, stealthy penetration of such micelles into biofilms, combined with the ability to accumulate in biofilms through electrostatic attraction at acidic pH to negatively charged bacterial cell surfaces.45 Therefore, we were initially disappointed that PEG-PAE micelles were virtually uncharged in high ionic strength saliva buffer (Figure 2H), but to our surprise this did not hamper penetration, accumulation nor long-time retention of the micelles in oral biofilms (Figure 3). Thus in absence of strong electrostatic attraction, Lifshitz-Van der Waals forces must be sufficient to enable micellar accumulation and retention, while at the same time possibly allowing micelles to move more freely to deeper biofilm layers. PEG-PAE-T5 micelles were demonstrated to be effective against a sub-cutaneous, multi-drug resistant infection in mice (Figure 5). Histological evaluation showed no signs of negative tissue reaction around the infection site five days after micelles injection and tissue looked completely healthy. In addition, PEGPAE-Tx micelles showed excellent cytocompatibility in the in vitro studies using different mammalian cell lines. Accordingly, PEG-PAE-T micelles may be considered cytocompatible and potentially safe for clinical treatment of various type of bacterial infections. Application towards oral biofilms yielded similarly positive results with respect to biofilm eradication (Figure 6). Opposite to internal biofilms, oral biofilms are composed of hundreds to thousands of different bacterial strains and species, most of which are considered to belong to the oral microbiome at health and function to protect the host against invading pathogens. In case of a disbalance in the oral microbiome, oral pathogens like S. mutans can become prevalent to cause dental caries. Therefore it is of high interest for the control of oral biofilms and the prevention of caries that Triclosan-conjugated PEG-PAE-T micelles selectively kills S. mutans as compared with Triclosan in solution and Triclosan-loaded micelles46 at a lower Triclosan-equivalent concentration (Figure 6). This is probably due to the fact that many cariogenic S. mutans isolates exhibit esterase activity,47 through which they may degrade the ester-linkage applied to conjugate Triclosan with PAE. S. sobrinus, another cariogenic member of the oral microbiome, is also suspected to possess a high esterase activity46 and may thus also be preferentially killed by our PEG-PAE-T micelles, but this was not evaluated in the present study. Considering the high carcinogenicity of S. mutans,48 PEG-PAE-T micelles might contribute in an oral health care product to the restoration of a healthy oral microbiome with less unnecessary killing of 90.

(26) Chapter 4. the non-pathogenic bacterial species.49 The prospects for further clinical evaluation of PEG-PAE-T micelles are promising, especially for difficult-totreat biofilm-associated infections such as the diabetic foot,50 but moreover for many other internal infections by multi-drug resistant strains that cannot be eradicated by conventional administration of antibiotics. This makes PEG-PAE with conjugated antimicrobials, such as, but not limited to Triclosan, a direly needed addition to the shrinking armamentarium of antibiotics clinically effective. From a regulatory perspective, it is important, that these micelles are not only based on approved and clinically used antimicrobials, but also PEG is FDA approved for drug conjugation, modification and delivery.51,52 So far, PAE has been shown to be fully biodegradable and biocompatible, in different types of studies regarding gene delivery53,54 and bacterial encapsulation.55 Since cytocompatibility of the micelles in mice as well as in vitro was good (Figure 5G), with antimicrobial efficacy ranging from ex vivo oral biofilms (Figure 6) to sub-cutaneous, MDR S. aureus infections in mice (Figure 5). Apart from the difference in bacterial composition of the biofilms studied in vitro, the staphylococcal biofilms studied in mice and the multi-species oral biofilms studied ex vivo, the degree of maturation of the biofilms also differs greatly. How long a biofilm needs to grow before it may be called “mature”, and what mature actually means, can be debated. However, it is evident 24 h of culturing forms a less mature biofilm than the oral biofilms formed in orthodontic patients, not seldom representing biofilms that have been matured for several weeks, depending on patient’s oral hygiene that is hampered by the orthodontic appliances. Without making specific claims with respect of the suitability of the micelles evaluated here, their efficacy on biofilms with different degrees of maturation, suggest potential prophylactic (usually very thin, unmatured biofilms) as well as therapeutic use (often full-grown, mature biofilms with clinical signs of infection).. CONCLUSION In conclusion, we have developed pH-adaptive, PEG-PAE-Tx micellar nanocarriers with a conjugated antimicrobial that is uniquely released by degradation of its ester-linkage with PAE through bacterial lipases. Therewith no antimicrobial cargo is released underway to an infection site through the blood stream. These micelles fully penetrated infectious biofilms owing to their stealth properties and accumulated in biofilms due to their pH-adaptive properties. Triclosan release was triggered by bacterial lipases once micelles had accumulated in a biofilm, and released their conjugated antimicrobial to kill biofilm bacteria. In vitro, the micelles with conjugated Triclosan were found effective in eradicating multi-drug resistant S. aureus, E. coli and streptococcal biofilms at lower Triclosan concentrations than required with dissolved Triclosan, while having no negative effects on the growth of mammalian cells. Micelles with ester-linked Triclosan eradicated subcutaneous multi-drug resistant S. aureus infections in mice and oral bacteria in human, ex vivo biofilms, preferentially killing S. mutans as a highly cariogenic oral pathogen. Therewith, prospects for further downward clinical translation of PEG-PAE-Tx micelles are promising.. REFERENCES (1) 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 (153), 153rv110. (2) Flemming , H.-C.; Wingender, J.; Szewzyk, U.; Steinberg, P.; Rice, S. A.; Kjelleberg, S. Biofilms: An Emergent Form of Bacterial Life. Nat. Rev. Microbiol. 2016, 14 (9), 563–575. (3) Min, J.; Choi, K. Y.; Dreaden, E. C.; Padera, R. F.; Braatz, R. D.; Spector, M.; Hammond, P. T. Designer Dual. Therapy Nanolayered Implant Coatings Eradicate Biofilms and Accelerate Bone Tissue Repair. ACS Nano 2016, 10 (4), 4441–4450. (4) Koo, H.; Allan, R. N.; Howlin, R. P.; Stoodley, P.; HallStoodley, L. Targeting Microbial Biofilms: Current and Prospective Therapeutic Strategies. Nat. Rev. Microbiol. 2017, 15 (12), 740–755. (5) Levy, S. B.; Bonnie, M. Antibacterial Resistance Worldwide: Causes, Challenges and Responses. Nat. Med. 2004, 10 (12S), S122–S129. (6) Yu, S.; Li, G.; Liu, R.; Ma, D.; Xue, W. Dendritic 91. 4.

(27) Chapter 4. 4. Fe3O4@Poly(Dopamine)@PAMAM Nanocomposite as Controllable NO-Releasing Material: A Synergistic Photothermal and NO Antibacterial Study. Adv. Funct. Mater. 2018, 28 (20), 1707440. (7) Yang, Y.; Ma, L.; Cheng, C.; Deng, Y.; Huang, J.; Fan, X.; Nie, C.; Zhao, W.; Zhao, C. Nonchemotherapic and Robust Dual-Responsive Nanoagents with On-Demand Bacterial Trapping, Ablation, and Release for Efficient Wound Disinfection. Adv. Funct. Mater. 2018, 28 (21), 1705708. (8) Natan, M.; Banin, E. From Nano to Micro: Using Nanotechnology to Combat Microorganisms and Their Multidrug Resistance. FEMS Microbiol. Rev. 2017, 41 (3), 302–322. (9) Ng, V. W. L.; Chan, J. M. W.; Sardon, H.; Ono, R. J.; García, J. M.; Yang, Y. Y.; Hedrick, J. L. Antimicrobial Hydrogels: A New Weapon in the Arsenal against Multidrug-Resistant Infections. Adv. Drug Deliv. Rev. 2014, 78, 46–62. (10) Hu, D.; Li, H.; Wang, B.; Ye, Z.; Lei, W.; Jia, F.; Jin, Q.; Ren, K.-F.; Ji, J. Surface-Adaptive Gold Nanoparticles with Effective Adherence and Enhanced Photothermal Ablation of Methicillin-Resistant Staphylococcus aureus Biofilm. ACS Nano 2017, 11 (9), 9330–9339. (11) Radovic-Moreno, A. F.; Lu, T. K.; Puscasu, V. A.; Yoon, C. J.; Langer, R.; Farokhzad, O. C. Surface ChargeSwitching Polymeric Nanoparticles for Bacterial Cell WallTargeted Delivery of Antibiotics. ACS Nano 2012, 6 (5), 4279–4287. (12) Landis, R. F.; Gupta, A.; Lee, Y.-W.; Wang, L.-S.; Golba, B.; Couillaud, B.; Ridolfo, R.; Das, R.; Rotello, V. M. Cross-Linked Polymer-Stabilized Nanocomposites for the Treatment of Bacterial Biofilms. ACS Nano 2017, 11 (1), 946–952. (13) Li, Y.; Liu, G.; Wang, X.; Hu, J.; Liu, S. EnzymeResponsive Polymeric Vesicles for Bacterial-Strain-Selective Delivery of Antimicrobial Agents. Angew. Chemie - Int. Ed. 2016, 55 (5), 1760–1764. (14) Xiong, M.-H.; Li, Y.-J.; Bao, Y.; Yang, X.-Z.; Hu, B.; Wang, J. Bacteria-Responsive Multifunctional Nanogel for Targeted Antibiotic Delivery. Adv. Mater. 2012, 24 (46), 6175–6180. (15) Zhu, Y.; Xu, C.; Zhang, N.; Ding, X.; Yu, B.; Xu, F.-J. Polycationic Synergistic Antibacterial Agents with Multiple Functional Components for Efficient Anti-Infective Therapy. Adv. Funct. Mater. 2018, 28 (14), 1706709. (16) Lam, S. J.; O’Brien-Simpson, N. M.; Pantarat, N.; Sulistio, A.; Wong, E. H. H.; Chen, Y.-Y.; Lenzo, J. C.; Holden, J. A.; Blencowe, A.; Reynolds, E. C.; et al. Combating Multidrug-Resistant Gram-Negative Bacteria with Structurally Nanoengineered Antimicrobial Peptide Polymers. Nat. Microbiol. 2016, 1, 16162. 92. (17) Elsabahy, M.; Wooley, K. L. Design of Polymeric Nanoparticles for Biomedical Delivery Applications. Chem. Soc. Rev. 2012, 41 (7), 2545–2561. (18) Pelgrift, R. Y.; Friedman, A. J. Nanotechnology as a Therapeutic Tool to Combat Microbial Resistance. Adv. Drug Deliv. Rev. 2013, 65 (13–14), 1803–1815. (19) Liu, J.; Liu, W.; Weitzhandler, I.; Bhattacharyya, J.; Li, X.; Wang, J.; Qi, Y.; Bhattacharjee, S.; Chilkoti, A. RingOpening Polymerization of Prodrugs: A Versatile Approach to Prepare Well-Defined Drug-Loaded Nanoparticles. Angew. Chemie - Int. Ed. 2015, 54 (3), 1002–1006. (20) Callmann, C. E.; Barback, C. V.; Thompson, M. P.; Hall, D. J.; Mattrey, R. F.; Gianneschi, N. C. Therapeutic Enzyme-Responsive Nanoparticles for Targeted Delivery and Accumulation in Tumors. Adv. Mater. 2015, 27 (31), 4611–4615. (21) Gao, H.; Cheng, T.; Liu, J.; Liu, J.; Yang, C.; Chu, L.; Zhang, Y.; Ma, R.; Shi, L. Self-Regulated Multifunctional Collaboration of Targeted Nanocarriers for Enhanced Tumor Therapy. Biomacromolecules 2014, 15 (10), 3634– 3642. ( 2 2 ) Z ha ng , Z . ; Ma , R . ; Sh i , L . Co o p e rat i v e Macromolecular Self-Assembly toward Polymeric Assemblies with Multiple and Bioactive Functions. Acc. Chem. Res. 2014, 47 (4), 1426–1437. (23) Liu, Y.; Busscher, H. J.; Zhao, B.; Li, Y.; Zhang, Z.; Van Der Mei, H. C.; Ren, Y.; Shi, L. Surface-Adaptive, Antimicrobial ly Loaded , Micel lar Nanocarr iers with Enhanced Penetration and Killing Efficiency in Staphylococcal Biofilms. ACS Nano 2016, 10 (4), 4779– 4789. (24) Gao, G. H.; Lee, J. W.; Nguyen, M. K.; Im, G. H.; Yang, J.; Heo, H.; Jeon, P.; Park, T. G.; Lee, J. H.; Lee, D. S. pH-Responsive Polymeric Micelle Based on PEG-Poly(βAmino Ester)/(Amido Amine) as Intelligent Vehicle for Magnetic Resonance Imaging in Detection of Cerebral Ischemic Area. J. Control. Release 2011, 155 (1), 11–17. (25) Kaper, J. B.; Nataro, J. P.; Mobley, H. L. T. Pathogenic Escherichia Coli. Nat. Rev. Microbiol. 2004, 2 (2), 123–140. (26) Lowy, F. D. Medical Progress: Staphylococcus aureus Infections. N. Engl. J. Med. 1998, 339 (8), 520–532. (27) Ren, Y.; Jongsma, M. A.; Mei, L.; van der Mei, H. C.; Busscher, H. J. Orthodontic Treatment with Fixed Appliances and Biofilm Formation--a Potential Public Health Threat? Clin. Oral Investig. 2014, 18 (7), 1711– 1718. (28) Pihlstrom, B. L.; Michalowicz, B. S.; Johnson, N. W. Periodontal Diseases. Lancet 2005, 366 (9499), 1809– 1820. (29) Otten, M. P. T.; Busscher, H. J.; Abbas, F.; van der Mei, H. C.; van Hoogmoed, C. G. Plaque-Left-behind after Brushing: Intra-Oral Reservoir for Antibacterial Toothpaste.

(28) Chapter 4. Ingredients. Clin. Oral Investig. 2012, 16 (5), 1435–1442. (30) Paula, A. J.; Koo, H. Nanosized Building Blocks for Customizing Novel Antibiofilm Approaches. J. Dent. Res. 2016, 96 (2), 128–136. (31) Moharamzadeh, K. Biocompatibility of Oral Care Products; 2016. (32) Leaper, D. J.; Edmiston, C. E.; Holy, C. E. MetaAnalysis of the Potential Economic Impact Following Introduction of Absorbable Antimicrobial Sutures. Br. J. Surg. 2017, 104 (2), e134–e144. (33) Renko, M.; Paalanne, N.; Tapiainen, T.; Hinkkainen, M.; Pokka, T.; Kinnula, S.; Sinikumpu, J.-J.; Uhari, M.; Serlo, W. Triclosan-Containing Sutures versus Ordinary Sutures for Reducing Surgical Site Infections in Children: A Double-Blind, Randomised Controlled Trial. Lancet Infect. Dis. 2017, 17 (1), 50–57. (34) Navazesh, M.; Christensen, C. M. A Comparison of W hole Mouth Resting and Stimulated Salivar y Measurement Procedures. J. Dent. Res. 1982, 61 (10), 1158–1162. (35) Patel, M. P.; Cruchley, A. T.; Coleman, D. C.; Swai, H.; Braden, M.; Williams, D. M. A Polymeric System for the Intra-Oral Delivery of an Anti-Fungal Agent. Biomaterials 2001, 22 (17), 2319–2324. (36) Liu, Y.; van der Mei, H. C.; Zhao, B.; Zhai, Y.; Cheng, T.; Li, Y.; Zhang, Z.; Busscher, H. J.; Ren, Y.; Shi, L. Eradication of Multidrug-Resistant Staphylococcal Infections by Light-Activatable Micellar Nanocarriers in a Murine Model. Adv. Funct. Mater. 2017, 27 (44), 1701974. (37) Horev, B.; Klein, M. I.; Hwang, G.; Li, Y.; Kim, D.; Koo, H.; Benoit, D. S. W. pH-Activated Nanoparticles for Controlled Topical Delivery of Farnesol to Disrupt Oral Biofilm Virulence. ACS Nano 2015, 9 (3), 2390–2404. (38) Otten, M. P. T.; Busscher, H. J.; Van Der Mei, H. C.; Abbas, F.; Van Hoogmoed, C. G. Retention of Antimicrobial Activity in Plaque and Saliva Following Mouthrinse Use in Vivo. Caries Res. 2010, 44 (5), 459–464. (39) Daghighi, S.; Sjollema, J.; Dijkstra, R. J. B.; Jaspers, V.; Zaat, S. A. J.; van der Mei, H. C.; Busscher, H. J. Real-Time Quantification of Matrix Metalloproteinase and Integrin Αvβ3 Expression during Biomaterial-Associated Infection in a Murine Model. Eur. Cells Mater. 2014, 27, 26–38. (40) Wescombe, P. A.; Heng, N. C. K .; Burton, J. P.; Chilcott, C. N.; Tagg, J. R. Streptococcal Bacteriocins and the Case for Streptococcus salivarius as Model Oral Probiotics. Future Microbiol. 2009, 4 (7), 819–835. (41) Gordon, J. I.; Klaenhammer, T. R. A Rendezvous with Our Microbes. Proc. Natl. Acad. Sci. U. S. A. 2011, 108 (SUPPL. 1), 4513–4515. (42) Allegranzi, B.; Zayed, B.; Bischoff, P.; Kubilay, N. Z.; de Jonge, S.; de Vries, F.; Gomes, S. M.; Gans, S.; Wallert, E. D.; Wu, X.; et al. New WHO Recommendations on. Intraoperative and Postoperative Measures for Surgical Site Infection Prevention: An Evidence-Based Global Perspective. Lancet Infect. Dis. 2016, 16 (12), e288–e303. (43) Bhargava, H. N.; Leonard, P. A . Triclosan: Applications and Safety. Am. J. Infect. Control 1996, 24 (3), 209–218. (44) Chen, F.; Rice, K. C.; Liu, X.-M.; Reinhardt, R. A.; Bayles, K. W.; Wang, D. Triclosan-Loaded Tooth-Binding Micelles for Prevention and Treatment of Dental Biofilm. Pharm. Res. 2010, 27 (11), 2356–2364. (45) Wang, L.-S.; Gupta, A.; Rotello, V. M. Nanomaterials for the Treatment of Bacterial Biofilms. ACS Infect. Dis. 2016, 2 (1), 3–4. (46) Bourbia, M.; Ma, D.; Cvitkovitch, D. G.; Santerre, J. P.; Finer, Y. Cariogenic Bacteria Degrade Dental Resin Composites and Adhesives. J. Dent. Res. 2013, 92 (11), 989–994. (47) Hansel, C.; Leyhausen, G.; Mai, U. E. H.; Geurtsen, W. Effects of Various Resin Composite (Co)Monomers and Extracts on Two Caries-Associated Micro-Organisms in Vitro. J. Dent. Res. 1998, 77 (1), 60–67. (48) Warinner, C.; Rodrigues, J. F. M.; Vyas, R.; Trachsel, C.; Shved, N.; Grossmann, J.; Radini, A.; Hancock, Y.; Tito, R. Y.; Fiddyment, S.; et al. Pathogens and Host Immunity in the Ancient Human Oral Cavity. Nat. Genet. 2014, 46 (4), 336–344. (49) Duran-Pinedo, A . E.; Frias-Lopez, J. Beyond Microbial Community Composition: Functional Activities of the Oral Microbiome in Health and Disease. Microbes Infect. 2015, 17 (7), 505–516. (50) Neut, D.; Tijdens-Creusen, E. J.; Bulstra, S. K.; van der Mei, H. C.; Busscher, H. J. Biofilms in Chronic Diabetic Foot Ulcers--a Study of 2 Cases. Acta Orthop. 2011, 82 (3), 383–385. (51) Greenwald, R . B.; Choe, Y. H.; McGuire, J.; Conover, C. D. Effective Drug Delivery by PEGylated Drug Conjugates. Adv. Drug Deliv. Rev. 2003, 55 (2), 217–250. (52) Veronese, F. M.; Pasut, G. PEGylation, Successful Approach to Drug Delivery. Drug Discov. Today 2005, 10 (21), 1451–1458. (53) Zhou, D.; Cutlar, L.; Gao, Y.; Wang, W.; O’KeeffeAhern, J.; McMahon, S.; Duarte, B.; Larcher, F.; Rodriguez, B. J.; Greiser, U. The Transition from Linear to Highly Branched Poly(β-Amino Ester)s: Branching Matters for Gene Delivery. Sci. Adv. 2016, 2 (6), e1600102. (54) Shah, N. J.; Hyder, M. N.; Moskowitz, J. S.; Quadir, M. A.; Morton, S. W.; Seeherman, H. J.; Padera, R. F.; Spector, M.; Hammond, P. T. Surface-Mediated Bone Tissue Morphogenesis from Tunable Nanolayered Implant Coatings. Sci. Transl. Med. 2013, 5 (191), 191ra183. (55) Li, Y.; Beitelshees, M.; Fang, L.; Hill, A.; Ahmadi, M. K.; Chen, M.; Davidson, B. A.; Knight, P.; Smith, R. 93. 4.

(29) Chapter 4. J.; Andreadis, S. T.; et al. In Situ Pneumococcal Vaccine Production and Delivery through a Hybrid BiologicalBiomaterial Vector. Sci. Adv. 2016, 2 (7), e1600264.. 4. 94.

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(34) . . . . . . . . . . . . . . . . . . . . . . . . . . . Chapter 4. SUPPORTING INFORMATION. . . Figure S1. NMR spectra of 2,2-bis(acryloxymethyl)propionic acid in d-CHCl3 at 0 °C. (A) 1H NMR and (B) 13C NMR.. . 4. . . Figure S2. NMR spectra of the key-monomer in d-CHCl3 at 0°C. (A) 1H NMR, (B) 13C NMR.. 95.

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(38) . . . . . . Chapter 4. . . . Figure S3. 1H NMR spectra of polymeric PEG-PAE-Tx in d-CHCl3 at 0°C. (A) PEG5k-b-P(AEco-T)5 and (B) PEG5k-b-P(AE-co-T)8 . “T” denotes Triclosan..

(39) Chapter 4. Figure S4. Matrix assisted laser desorption/ionization time of flight (MALDI-ToF) mass spectrometry of PEG-PAE-Tx measured on an AutoflexIII LRF200-CID mass spectrometer (Bruker, Massachusetts, United States). (A) PEG5k-b-P(AE-co-T)5 and (B) PEG5k-b-P(AE-co-T)8 . “T” denotes Triclosan.. Figure S5. Scattered light intensity (kcps) as a function of PEG-PAE-Tx concentration (μg mL –1) in 10 mM potassium phosphate. The critical micelle concentration (CMC) of PEG-PAETx in 10 mM potassium phosphate (pH 7.4) was determined by dynamic light scattering. All samples were measured at 25°C using a Zetasizer Nano-ZS with a DTS1060-type cuvette.. 97. 4.

(40) Chapter 4. Figure S6. Diameters and zeta potentials of PEG-PAE-T5 micelles exposed to salivary proteins (1.5 mg mL –1 in saliva buffer) for 2 h. (A) Diameter as a function of pH for PEG-PAE-T5 micelles measured using dynamic light scattering in saliva buffer at 25°C. Data are expressed as mean ± SD over 3 experiments with separately prepared micelles. (B) Zeta potentials of PEGPAE-T5 micelles (micelle concentration of 0.1 mg mL –1) as a function of pH in saliva buffer. Data are expressed as mean ± SD over 3 experiments with separately prepared micelles.. 4. Figure S7. Diameters and zeta potentials of Nile red-loaded PEG-PAE-Tx micelles. (A) Diameter as a function of pH for Nile red-loaded PEG-PAE-Tx micelles measured using dynamic light scattering in 10 mM potassium phosphate buffer at 25°C. (B) Same as (A), now for the zeta potentials (micelle concentration of 0.1 mg mL –1).. 98.

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(42). Figure S8. Cytocompatibility of PEG-PAE-Tx micelles with Triclosan conjugated towards mammalian cells. Relative metabolic activity, expressed as percentage MTT conversion of human umbilical vein endothelial cells (HUVEC), human skin fibroblasts (HSF) and human epithelial colorectal adenocarcinoma cells (Caco-2) after 24 h growth in the presence of PEG-PAE-Tx at different micelle concentrations. MTT conversion in absence of micelles was set at 100%.. 4. Figure S9. Examples of hematoxylin/eosin stained histological micrographs of murine skin tissues from mice at day 5 post different treatments. Red arrows indicate the infiltration with immune cells, while white arrows indicate the infected area within the tissue. Scale bar equals 200 μm.. 99.

(43) Chapter 4. 4. Figure S10. Examples of CLSM micrographs of orthodontic biofilms after ex vivo treatment. Treatment involved 2 min exposure to saliva buffer or Triclosan at equal Triclosan-equivalent concentrations (200 μg mL –1) followed by 3 h retention in saliva buffer at 37°C. Images represent dispersed biofilms after LIVE-DEAD staining. Dead bacteria appear red-fluorescent, live bacteria are green-fluorescent. Scale bar equals 20 μm. Table S1. Bacterial strains and species together with their agar types and liquid growth media as used in this study.* Strains Staphylococcus aureus Xen36 WHGFP Escherichia coli Xen14 Streptococcus mutans ATCC700610 HG985 Streptococcus sobrinus ATCC33478 HG1025 Streptococcus oralis J22 Streptococcus sanguinis ATCC10556 Streptococcus mitis. 100. Agar Type. Growth medium. TSB agar with 200 μg mL –1 kanamycin TSB agar with 10 μg mL –1 tetracycline. TSB TSB with 10 μg mL –1 tetracycline. LB agar. LB. Blood agar (5% sheep blood) Blood agar (5% sheep blood). THB THB. Blood agar (5% sheep blood) Blood agar (5% sheep blood). THB THB. Blood agar (5% sheep blood). THB. Blood agar (5% sheep blood). THB.

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