An N-halamine-based rechargeable antimicrobial and biofilm controlling polyurethane

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An N-halamine-based rechargeable antimicrobial and biofilmcontrolling polyurethane

Xinbo Sun a, Zhengbing Cao a, Nuala Porteous b, Yuyu Sun a,⇑a Biomedical Engineering Program, The University of South Dakota, 4800 North Career Avenue, Sioux Falls, SD 57107, USAb Department of Comprehensive Dentistry, University of Texas Health Science Center at San Antonio, San Antonio, TX 78229-4404, USA

a r t i c l e i n f o

Article history:Received 15 September 2011Received in revised form 27 November 2011Accepted 20 December 2011Available online 27 December 2011

Keywords:AntimicrobialBiofilm controllingN-HalamineChlorinationRechargeable

a b s t r a c t

An N-halamine precursor, 5,5-dimethylhydantoin (DMH), was covalently linked to the surface of polyure-thane (PU) with 1,6-hexamethylene diisocyanate (HDI) as the coupling agent. The reaction pathwayswere investigated using propyl isocyanate (PI) as a model compound. The results suggested that theimide and amide groups of DMH have very similar reactivities toward the isocyanate groups on PU sur-faces activated with HDI. After bleach treatment the covalently bound DMH moieties were transformedinto N-halamines. The new N-halamine-based PU provided potent antimicrobial effects against Staphylo-coccus aureus (Gram-positive bacterium), Escherichia coli (Gram-negative bacterium), methicillin-resis-tant Staphylococcus aureus (MRSA, drug-resistant Gram-positive bacterium), vancomycin-resistantEnterococcus faecium (VRE, drug-resistant Gram-positive bacterium), and Candida albicans (fungus), andsuccessfully prevented bacterial and fungal biofilm formation. The antimicrobial and biofilm controllingeffects were stable for longer than 6 months under normal storage in open air. Furthermore, if the func-tions were lost due to prolonged use they could be recharged by another chlorination treatment. Therecharging could be repeated as needed to achieve long-term protection against microbial contaminationand biofilm formation.

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1. Introduction

Thanks to its wide availability, low cost, ease of fabrication, andexcellent physical and biological properties polyurethane (PU) hasbecome one of the most versatile polymers in medical, dental,industrial, institutional, and environmental applications [1–3].Unfortunately, like most conventional polymers, the PU surface issusceptible to contamination by microorganisms, which can actas sources of cross-contamination and cross-infection [4–6]. More-over, adherent microbial species can secrete ‘‘slimes’’ (extracellularpolymeric substances or EPS) that attach them firmly to the mate-rial surfaces, resulting in irreversible adhesion and biofilm forma-tion [7–9]. Once formed, biofilms are very difficult to destroy.Protected by the EPS, microbes living in a biofilm are up to1000 times more resistant to disinfection, causing serious prob-lems, including device-related infections, healthcare-acquiredinfections, waterborne/foodborne illnesses, corrosion, blockage offilters, etc.

Consequently, there has been a growing interest in the develop-ment of antimicrobial polymers, including antimicrobial PU, tosolve these problems. The majority of reported antimicrobial PU

was prepared by either physically mixing antimicrobial agents(e.g. antibiotics, metal ions, quaternary ammonium salts, iodine,etc.) into or covalently binding the antimicrobial agents onto thetarget polymers [11–14]. Nonetheless, since microbial contamina-tion and biofilm formation mainly occur on polymer surfaces, itcould be advantageous to utilize surface modification to impartthe desired antimicrobial effects without causing bulk structural/property changes in the polymers [10,15]. The research interest ofthis laboratory focuses on N-halamine-based antimicrobial and bio-film controlling polymers. N-Halamines are compounds containingone or more nitrogen–halogen covalent bonds. N-Halamines haveantimicrobial efficacies similar to that of hypochlorite bleach, oneof the most widely used disinfectants, but they are more stable, lesscorrosive, and have much less tendency to generate halogenatedhydrocarbons. Therefore, N-halamines have found wide applica-tions as food and water disinfectants [16,17]. Moreover, once N-hal-amine structures are covalently linked to the backbones of ordinarypolymers the antimicrobial effects can be retained and the resultingpolymers are transformed into antimicrobial polymers [18–23].

Building on these results, in the current study we report a strat-egy to covalently bind an N-halamine precursor, 5,5-dimethylhy-dantoin (DMH), onto the PU surface using hexamethylenediisocyanate (HDI) as a coupling agent. The bound DMH moietieson the PU surface are transformed into N-halamines through a sim-ple bleach treatment, leading to potent antimicrobial and biofilm

1742-7061/$ - see front matter � 2011 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.doi:10.1016/j.actbio.2011.12.027

⇑ Corresponding author. Tel.: +1 605 367 7776.E-mail address: [email protected] (Y. Sun).

Acta Biomaterialia 8 (2012) 1498–1506

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controlling functions against Staphylococcus aureus (Gram-positivebacterium), Escherichia coli (Gram-negative bacterium), methicil-lin-resistant Staphylococcus aureus (MRSA, drug-resistant Gram-positive bacterium), vancomycin-resistant Enterococcus faecium(VRE, drug-resistant Gram-positive bacterium), and Candidaalbicans (fungus), representative microorganisms that are responsi-ble for various infections [24–27]. The antimicrobial and biofilmcontrolling effects of the new polymers are both durable andrechargeable, pointing to even greater potential of the new systemsfor a broad range of related applications.

2. Experimental procedures

2.1. Materials

5,5-Dimethylhydantoin (DMH), 1,6-hexamethylene diisocya-nate (HDI), and propyl isocyanate (PI) were purchased from Sig-ma–Aldrich and used as received. Polyurethane (PU) (Estane�

5707), a polyester-based thermoplastic PU, was kindly suppliedby Lubrizol Advanced Materials Inc. (Cleveland, OH). A householdbleach (Clorox� regular bleach) containing 6.0 wt.% sodium hypo-chlorite was used throughout this study. Other chemicals were ofanalytical grade and were used without further purification.

The microorganisms S. aureus (ATCC 6538), E. coli (ATCC 15597),MRSA (ATCC BAA-811), VRE (ATCC 700221), and C. albicans (ATCC10231) were obtained from the American Type Culture Collection(ATCC).

2.2. Instruments

Attenuated total reflectance (ATR) spectra of the samples wererecorded by a Thermo Nicolet 6700 infrared spectroscope (Woburn,MA). 1H NMR studies were carried out using a Varian Unity-300spectrometer (Palo Alto, CA) at ambient temperature in DMSO-d6.A FEI Quanta 450 scanning electron microscope was used to studythe surface morphology of the samples. UV/vis spectra wererecorded on a Beckman DU� 520 UV/vis spectrophotometer.

2.3. Surface modification of PU

PU pellets were extracted with methanol and distilled water for24 h to remove low molecular weight components and other po-tential contaminants. After extraction the pellets were dried in avacuum drier at 25 �C for 48 h. PU films were obtained by solutioncasting. To prepare PU films 2 g PU was dissolved in 20 ml of tetra-hydrofuran (THF) at room temperature. After filtration the PU solu-tion was poured into a glass dish (100 � 15 mm) and allowed todry in a fume hood overnight at room temperature. The resultingfilm was uniform and optically transparent with a thickness ofaround 0.1 mm and was cut into square shaped test films.

DMH was covalently linked to the PU surface through a two-step procedure, as illustrated in Scheme 1. In the first step 1,6-hexamethylene diisocyanate (HDI) was dissolved in anhydroustoluene at a volume ratio of 1:10. PU films were immersed in150 ml of HDI–toluene in a 250 ml three-necked round flask withthe presence of 0.5 vol.% dibutyltin dilaurate (DBTDL) as catalyst.The mixture was stirred under an N2 atmosphere at 60 �C for 2 h.The isocyanate-containing PU (PU–HDI) films were taken out andwashed repeatedly with anhydrous toluene to remove unreactedHDI. In the second step DMH was immobilized onto the PU–HDIsurface. PU–HDI films were dispersed in 100 ml of anhydroustoluene containing 10 g DMH, and the mixtures were stirred underan N2 atmosphere at 70 �C for 3 h. All films after surface modifica-tions (PU–HDI–DMH) were extracted with anhydrous toluene andethanol three times, and dried under vacuum.

2.4. Synthesis of the model compound

To study the reaction mechanism between PU–HDI and DMH, PIwas used as a model compound of PU–HDI. In the reaction equalmolar concentrationsof DMH and PI were dissolved in anhydroustoluene containing 0.5 vol.% DBTDL. The reaction continued in a250 ml three-necked round flask under an N2 atmosphere at70 �C for 3 h. After reaction the solvent was removed under re-duced pressure, and the residuals were dried in a vacuum oven.The final product was recrystallized twice from deionized water,and dried over CaCl2 in a vacuum oven.

2.5. Determination of isocyanate content

The –NCO content on the surface of PU–HDI was determined byreacting the isocyanate groups on PU–HDI with an excess of dibu-tylamine (Bu2NH) in anhydrous toluene, followed by back-titrationof the remaining Bu2NH with hydrochloric acid in anhydrous eth-anol using bromphenol blue as an indicator, as reported previously[28,29]. The mechanism is shown in Eqs. (1) and (2):

RNCOþ Bu2NH! RNHCONBu2 ð1Þ

Bu2NHþHCl! Bu2NHþ2 Cl� ð2Þ

The unmodified original PU was titrated under the same condi-tions to serve as a control. Each test was repeated three times.

2.6. Chlorination of PU–HDI–DMH

To transform the grafted DMH moieties into N-halamines, PU–HDI–DMH films were immersed in 10% bleach solution at roomtemperature for 45 min with constant stirring. Afterwards thesamples were washed copiously with deionized water (the wash-ing water was tested with potassium iodide/starch to ensure thatthe free chlorine was removed), air dried and stored in a desiccatorfor 48 h to reach constant weight. The content of active chlorine onthe surface of the chlorinated PU–HDI–DMH film was determinedby iodometric titration following our published procedures[18–20,22,23].

2.7. Antibacterial and antifungal functions

In the antibacterial study S. aureus (ATCC 6538) and E. coli (ATCC15597) were used as examples of non-resistant Gram-positive andGram-negative bacteria, respectively. MRSA (ATCC BAA-811) andVRE (ATCC 700221) were selected to represent drug-resistantstrains because these species have caused serious healthcare-asso-ciated infections (HAI) and community-acquired infections [30]. C.albicans (ATCC 10231) was employed to challenge the antifungalactivities of the samples.

To prepare the bacterial and fungal suspensions an inoculatingloop (10 ll) of S. aureus, E. coli, MRSA, and VRE 700221 were grownin corresponding 5 ml of broth solutions (see Table 1) at 37 �C for24 h, and C. albicans was grown in 5 ml of YM broth at 26 �C for36 h. Cells were harvested by centrifugation, washed twice withsterile phosphate-buffered saline (PBS), and then resuspended insterile PBS.

In the current study waterborne microbial challenge conditionswere used to evaluate the antimicrobial effects of the film samples.All microbial tests were performed in a Biosafety Level-2 hood toensure safety, and the films were sterilized for 15 min on each sideunder UV light before microbial studies. In each test 10 ll of anaqueous suspension containing 108–109 CFU ml�1 of each bacteria(S. aureus, E. coli, MRSA, and VRE) or the fungus (C. albicans) was

X. Sun et al. / Acta Biomaterialia 8 (2012) 1498–1506 1499


placed on the surface of a chlorinated PU–HDI–DMH film(3 � 3 cm). Another identical film was used to cover it, so the bac-terial/fungal aqueous suspension was trapped inside the films, likethe ‘‘filling’’ in a sandwich to improve contact between the bacteriaand the film surfaces. After a certain period of contact time theentire ‘‘sandwich’’ was transferred to 10 ml of 0.03 wt.% sodiumthiosulfate aqueous solution to quench the active chlorine and stopthe microbial tests [20]. The mixture was vortexed for 1 min andthen sonicated for 5 min. An aliquot of the solution was seriallydiluted, and 100 ll of each dilution was plated onto the

corresponding agar plates. The same procedure was also appliedto the original PU film to serve as a control. Bacterial colonies werecounted after incubation at 37 �C for 24 h, and fungal colonies werecounted after incubation at 26 �C for 36 h. Each test was repeatedthree times.

2.8. Kirby–Bauer test

In this test the surfaces of a tryptic soy agar plate and Luria–Bertant (LB) agar plate were overlaid with 1 ml of 108–109 CFU ml�1

Scheme 1. Covalently binding DMH onto PU surfaces.

Table 1Microorganisms tested in this study and the media used for their growth.

Bacteria Drug-resistant bacteria Fungi

S. aureusa E. colib MRSRa VREa C. albicans 10231

Broth Tryptic soy broth LB broth Tryptic soy broth Tryptic soy broth YM brothAgar Tryptic soy agar LB agar Tryptic soy agar Tryptic soy agar YPD agar

a Gram-positive bacteria.b Gram-negative bacterium.

1500 X. Sun et al. / Acta Biomaterialia 8 (2012) 1498–1506


of S. aureus and E. coli, respectively. The plates were then allowed tostand at 37 �C for 2 h. Each chlorinated PU–HDI–DMH film(2 � 2 cm) was placed on the surface of each of the bacteria-containing agar plates. The film was gently pressed with sterile for-ceps to ensure full contact between the film and the agar. Afterincubation at 37 �C for 24 h the inhibition zone around the films(if any) was measured. Afterwards the films were sterilely removedfrom the agar plates, and washed gently with non-flowing sterilePBS (3 � 10 ml) to remove loosely attached bacteria and/or possibleagar components. The resulting films were vortexed for 1 min andsonicated for 5 min in 10 ml of PBS to detach adherent bacteria.The sonicated solution was serially diluted, and 100 ll of each dilu-tion was placed on the corresponding agar plates (see Table 1).Recoverable microbial colonies were counted after incubation at37 �C for 24 h. The same procedure was also applied to the originalPU film to serve as a control.

2.9. Biofilm controlling function

The ability of the chlorinated PU–HDI–DMH films to preventbiofilm formation was evaluated using scanning electron micros-copy (SEM). In this study S. aureus, E. coli and C. albicans weregrown and harvested as described above. A chlorinated PU–HDI–DMH film (2 � 2 cm) was immersed in 10 ml of sterile PBS contain-ing 108–109 CFU ml�1 of S. aureus, E. coli or C. albicans. The mixturewas gently shaken at 37 �C for 30 min to allow microbial adhesion.The films were taken out of the bacterial or fungal solution, gentlywashed three times with 10 ml of sterile PBS to remove loosely at-tached cells. The films were then immersed in broth solutions(tryptic soy broth for S. aureus, LB broth for E. coli, and YM brothfor C. albicans), and incubated at 37 �C for 72 h to facilitate biofilmformation. The films were taken out, gently washed three timeswith 10 ml of sterile PBS, and fixed with 3% glutaraldehyde in0.1 M sodium cacodylate buffer (SCB) at 4 �C for 24 h. After beinggently washed with SCB the samples were dehydrated throughan alcohol gradient, dried in a critical point drier, mounted on sam-ple holders, sputter coated with gold–palladium, and observed in aFEI Quanta 450 scanning electron microscope to check for the pres-ence of biofilms. The same procedure was also applied to theunchlorinated PU–HDI–DMH films and the original PU film to serveas controls.

2.10. Stability and rechargeability of the N-halamines

To evaluate the stability of the covalently bound chorines in theN-halamines under wet conditions a series of chlorinated PU–HDI–DMH films (3 � 3 cm) were immersed in 10 ml of PBS with constantshaking (50 r.p.m.) at room temperature. After a certain period oftime 1 ml of solution was taken out of the immersing PBS and testedwith a Beckman DU� 520 UV/vis spectrophotometer in the range190–400 nm to determine whether DMH or Cl–DMH containingcompounds were released from the PU film into solution (the char-acteristic absorption peak of pure DMH is at 218 nm, that ofCl–DMH at 348 nm). The chlorine contents and the antibacterialand antifungal functions of the film samples were tested periodi-cally over a 1 month period.

The stability of the N-halamine-based PU was also investigatedunder dry storage conditions. A series of chlorinated PU–HDI–DMHfilms with known chlorine contents were stored under normallaboratory conditions (25 �C, 40–60% relative humidity) in openair. The chlorine contents and the antibacterial and antifungalfunctions were tested periodically over a 6 month storage period.

To test rechargeability the N-halamine-based PU films werefirst treated with 0.1 M sodium thiosulfate aqueous solution atroom temperature for 24 h to quench the bound chlorine, and thenrecharged using a 10% Clorox� regular bleach aqueous solution for

45 min. The chlorinated PU films were washed with distilled water,air dried, and then quenched and recharged again. After differentcycles of this ‘‘quenching–recharging’’ treatment the chlorine con-tents and antibacterial and antifungal functions of the resultingfilms were retested to fully evaluate rechargeability. Each testwas repeated three times.

2.11. Statistical analysis

Data management and analysis was conducted using SPSS v.11.0 (SPSS Inc., Chicago, IL). One-way analysis of variance was usedto assess the statistical significance at a 95% confidence level.

3. Results and discussion

3.1. Surface modification of PU

DMH was covalently linked to the surface of PU using a two-stepchemical reaction using HDI as the coupling agent, as shown inScheme 1. The reaction was confirmed by attenuated total reflec-tion (ATR)–Fourier transform infrared spectroscopy studies (seeFig. 1). In the spectrum of PU the bands around 3420–3200 cm�1

were attributable to N–H stretching vibrations, the peaks at3000–2800 cm�1 were assigned to CH2 and CH3 stretching, the peakat 1725 cm�1 was caused by C@O (amide I) stretching, and the sig-nal at 1530 cm�1 was attributable to N–H (amide II) deformation, ingood agreement with the literature data [31]. Each HDI has two iso-cyanate (–NCO) groups, both of which could react with the ure-thane groups of PU, causing crosslinking of the polymer (seebelow). On the other hand, some HDI molecules could contributeonly one –NCO group to react with PU, leaving a free –NCO groupon the PU surface for further functionalization with DMH.

The presence of free –NCO groups on PU–HDI was confirmed byboth ATR studies and Bu2NH/HCl titration. In the ATR spectrum ofPU–HDI (Fig. 1), in addition to the characteristic bands of PU, a newband at 2260 cm�1 could be clearly detected, which was attribut-able to the stretching oscillation of free –NCO groups. In Bu2NH/HCl titration, although the original PU did not have any detectable–NCO groups, after reaction with HDI under our reported condi-tions the resulting PU–HDI surface had (1.81 ± 0.15) � 1016 groupscm�2 free –NCO (see Table 2).

Fig. 1. FTIR spectra of PU, PU–HDI, and PU–HDI–DMH.



The free –NCO groups on PU–HDI enabled the polymer to cova-lently bind DMH on the film surface. As shown in Fig. 1, after reac-tion with DMH the 2260 cm�1 isocyanate signal disappeared. Onthe other hand, a new band at 1760 cm�1 could be observed inthe ATR spectrum of PU–HDI–DMH as a weak shoulder, whichmust be caused by the carbonyl groups of the covalently boundDMH [32].

Each DMH molecule has two potential sites, the amide –NH andimide –NH, for reaction with the free isocyanate groups on PU–HDI. It was of great interest to determine which site(s) reactedwith –NCO. Normally such information could be readily obtainedthrough 1H NMR studies. However, under our reaction conditions(see Experimental procedures) the resulting PU–HDI and PU–HDI–DMH were only partially soluble in DMSO, DMF, THF, andother good solvents of PU. These results suggest that part of thePU polymer chains were crosslinked by HDI, making 1H NMR stud-ies difficult to perform on these polymers. On the other hand, sincethe reaction of PU–HDI with DMH occurred as one free –NCO onPU–HDI choosing a reaction site with DMH (amide –NH or imide–NH), we believe that the reaction pathway could be reasonably

deduced using a mono isocyanate, such as PI, as a model of PU–HDI to react with DMH.

Fig. 2 presents the 1H NMR spectra of DMH and the product ofDMH reaction with PI (DMH–PI). In the spectrum of DMH the sig-nals for the imide protons and amide protons appeared at 10.56and 7.92 ppm, respectively. The signal at 1.23 ppm correspondsto the protons of methyl groups. Upon reaction with PI the signalat 10.56 ppm in the spectrum of the resulting DMH–PI would de-crease significantly if the binding site was primarily the imide –NH group of DMH. On the other hand, if the reaction mainly oc-curred at the amide –NH group of DMH the signal of the productaround 7.92 ppm would become much weaker [32,33]. However,in the 1H NMR spectrum of DMH–PI both signals, of the imideand amide protons, were clearly detectable, with very similar pro-ton integration values (1.82 vs. 1.87). These results might suggestthat due to the high reactivity of isocyanate groups the imide –NH and amide –NH groups of DMH had very similar probabilitiesof reacting with PU–HDI. Thus the resulting PU–HDI–DMH couldhave both amide and imide –NH groups for chlorination to produceN-halamines.

After treatment with 10% bleach solution the amide and theimide groups on PU–HDI–DMH were readily transformed to N-hal-amine structures. Iodometric titration revealed that the chlorinatedPU–HDI–DMH had a total chlorine atom area density of (1.76 ±0.11) � 1016 atoms cm�2, which was relatively unchanged afterstorage for up to 6 months in the open air. As a comparison, afterthe same chlorine bleach treatment the original PU film had(1.18 ± 0.13) � 1014 atoms cm�2 chlorine. After only 1 week of

Table 2Determination of NCO content and active chlorine content.

Sample NCO (groups cm�2) Active chlorine (atoms cm�2)

Original PU 0 (1.18 ± 0.13) � 1014

PU–HDI (1.81 ± 0.15) � 1016 N/APU–HDI–DMH 0 (1.76 ± 0.11) � 1016

Fig. 2. 1H NMR spectra of DMH and DMH–PI.



storage in the open air almost no chlorine could be detected by titra-tion. These findings strongly suggest the presence of stable N-hal-amines on the surface of chlorinated PU–HDI–DMH films.

3.2. Antibacterial and antifungal activity

In this study the antibacterial and antifungal efficacies of thechlorinated PU–HDI–DMH films were evaluated in waterbornecontact mode [18,20,34]. The antimicrobial results are shown inTable 3 using the untreated original PU film as a negative control.The chlorinated PU–HDI–DMH films provided a 4-log reduction(the highest antimicrobial potency that could be observed underthe current test conditions) in S. aureus (Gram-positive bacterium)in 60 min, and a 4-log reduction in E. coli (Gram-negative bacte-rium) in 30 min, respectively. On the other hand, the freshly pre-pared chlorinated original PU films only provided a 90% reduction

of the same bacteria after as long as 8 h contact (data not shownin Table 3), suggesting that the antimicrobial effects of the chlori-nated PU–HDI–DMH mainly originated from the N-halamines onthe covalently bound DMH moieties.

Drug-resistant bacteria, including MRSA and VRE, are majorconcerns in healthcare settings and a wide range of related com-munity facilities, causing serious healthcare-related infectionsand community-acquired infections. It was encouraging to findthat the chlorinated PU–HDI–DMH films provided potent antimi-crobial effects against MRSR and VRE, achieving a 4-log reductionin both species within 30 min (see Table 3). This phenomenoncould be explained by the antimicrobial mechanisms. MRSA isresistant to certain antibiotics via the mecA gene, and VRE acquiresresistance to vancomycin through the introduction of new DNA toprevent vancomycin-binding [6]. On the other hand, N-halaminesprovide antimicrobial effects by the oxidative reaction of Cl+ fromN-halamines with microbial cells, which was apparently not af-fected by the drug-resistant characteristics of MRSR or VRE.[35].

Candida spp. are emerging as important nosocomial pathogens,and implanted devices with detectable biofilms are frequentlyassociated with these species [36]. The antifungal action of thechlorinated PU–HDI–DMH samples was evaluated againstC. albicans, which showed that the new N-halamine-based PUsamples provided a 4-log reduction in the yeast within 60 min,pointing to even greater potential of the new samples.

3.3. Kirby–Bauer test

The antibacterial activity of the chlorinated PU–HDI–DMHsamples was further assessed using the Kirby–Bauer test. As shownin Fig. 3, the original PU film did not show any inhibition zones. In

Table 3Log reduction in microbial cells in the antimicrobial tests.a

Contact time (min) S. aureus E. coli MRSA VRE C. albicans

5 1-log 2-log 1-log 1-log 1-log10 2-log 3-log 3-log 3-log 2-log30 3-log 4-logb 4-log 4-log 3-log60 4-log 4-log

a S. aureus, E. coli, MRSA, VRE, and C. albicans concentrations were 108–109 CFU ml�1, and 10 ll of such microbial suspensions were used in each test (seetext for details). The chlorinated PU–HDI–DMH surface contained (1.76 ± 0.11)� 1016 atoms cm�2 active chlorine.

b The highest antimicrobial potency that could be observed under the currenttesting conditions.

Fig. 3. Inhibition zone of (a) the original PU film against S. aureus, (b) the chlorinated PU–HDI–DMH film against S. aureus, (c) the original PU film against E. coli, and (d) thechlorinated PU–HDI–DMH film against E. coli.



the case of the chlorinated PU–HDI–DMH samples containing(1.76 ± 0.11) � 1016 atoms cm�2 active chlorine a clear inhibitionzone could be detected, with a zone size of around 1.8 mm against

S. aureus and about 0.9 mm against E. coli. These findings suggestthat during the antimicrobial tests active chlorines diffused awayfrom the chlorinated PU–HDI–DMH to kill the bacteria. To confirmthis, chlorinated PU–HDI–DMH films were immersed in deionizedwater under constant shaking at room temperature and a UV/visspectrophotometer was used to test the immersion solutions.Within the test period of 72 h the soaking solution was very clear,with no suspension/precipitation being observed. In the wave-length range 190–400 nm no UV absorption could be detected,suggesting that no detectable DMH or Cl–DMH containing com-pounds were released into the water system. On the other hand,iodometric titration found that the solution contained about0.2 ppm active chlorine, indicating that the inhibition zones weremost likely caused by Cl+ originating from the disassociation of

Table 4Inhibition zones of the films and levels of recoverable bacteria.

Inhibition zone (mm) Bacteria recovered (CFU cm�2)

S. aureus E. coli S. aureus E. coli

Controla 0 0 (5.4 ± 0.11) � 106 (4.9 ± 0.18) � 106

Sampleb 1.8 ± 0.1 0.9 ± 0.1 (4.9 ± 0.36) � 102 (3.1 ± 0.21) � 102

a The original PU.b Chlorinated PU–HDI–DMH with (1.76 ± 0.11) � 1016 atoms cm�2 active

chlorine.

Fig. 4. Biofilm controlling function of (a) the original PU film against S. aureus, (b) the chlorinated PU–HDI–DMH film against S. aureus, (c) the original PU film against E. coli,(d) the chlorinated PU–HDI–DMH film against E. coli, (e) the original PU film against C. albicans, and (f) the chlorinated PU–HDI–DMH film against C. albicans.



N-halamines. Such a low level of chlorine release pointed to goodstability and safety of the new samples (as a comparison, the EPAallows drinking water to contain up to 4 ppm active chlorine asdisinfection residues).

To provide more information about the antimicrobial effects,immediately after the Kirby–Bauer test the films were washedand sonicated to remove surface adherent bacteria. As shown inTable 4, as high as (5.4 ± 0.11) � 106 CFU cm�2 S. aureus or(4.9 ± 0.18) � 106 CFU cm�2 E. coli could be recovered from the ori-ginal PU film (n = 3). From the chlorinated PU–HDI–DMH films,however, the recoverable levels of S. aureus and E. coli were onlyin the range 102 CFU cm�2, further confirming the antimicrobialeffects of the new samples.

3.4. Biofilm controlling effects

The potent antimicrobial activity of the new chlorinated PU–HDI–DMH samples pointed to effective biofilm controlling func-tions. To evaluate this effect, Fig. 4 presents the SEM results ofthe samples in biofilm controlling studies. After 72 h growth inthe corresponding broth solutions a large quantity of layered S.aureus (Fig. 4a), E. coli (Fig. 4c) and C. albicans (Fig. 4e) attachedand aggregated on the original PU surfaces, indicating the forma-tion and development of biofilms. Similar results were observedon the unchlorinated PU–HDI–DMH films (images not shown).On the surfaces of the chlorinated PU–HDI–DMH films, however,no adherent microbial cells could be observed, suggesting powerfulbiofilm controlling functions of the new N-halamine-based PUagainst Gram-positive bacteria, Gram-negative bacteria, and fungi.The biofilm controlling function of the chlorinated PU–HDI–DMHfilms could be attributed to the antimicrobial activities of the N-halamines, i.e. when microbial cells came into contact with thefilms most of them were inactivated during and/or after adher-ence/colonization. The inactivated cells could be easily removedduring the SEM preparation process (e.g. rinsing, fixation, and alco-hol dehydration), resulting in cleaner surfaces [20].

3.5. Stability and rechargeability of the N-halamines

In the soaking studies the soaking solution was very clear andno suspension/precipitation could be observed during the test per-iod of 1 month with constant stirring. In the wavelength range190–400 nm no UV absorption could be found, suggesting thatno detectable DMH or Cl–DMH containing compounds were re-leased from the chlorinated PU–HDI–DMH films. Moreover, after1 month in solution the antimicrobial efficacies of the samplesagainst the bacterial and fungal species were unchanged.

In the dry storage studies, under normal laboratory conditions,after 6 months storage in the open air 91% the original active chlo-rines were retained on the chlorinated PU–HDI–DMH films (asshown in Table 5). The antimicrobial activity against the bacterialand fungal species were unchanged at the end of the 6 month stor-age tests. Furthermore, after 10 cycles of ‘‘quenching–recharging’’treatment the chlorine contents and antimicrobial activities of thenew N-halamine-based PU samples were essentially unchanged,indicating that the antimicrobial function was fully rechargeable.

4. Conclusions

DMH, an N-halamine precursor, was covalently bound to PUsurfaces using HDI as the coupling agent. The reactions were con-firmed by ATR, 1H NMR, –NCO content determination, and iodo-metric titration. Model compound studies suggested that theimide and amide groups of DMH have very similar reactivity to-wards –NCO groups on the PU–HDI surfaces. Upon bleach treat-ment the DMH moieties on the PU–HDI–DMH surfaces weretransformed into N-halamines, providing powerful, durable, andrechargeable antimicrobial activities against Gram-positive bacte-ria, Gram-negative bacteria, drug-resistant bacteria, and fungi.The Kirby–Bauer test showed that the antimicrobial function wasat least partly provided by the positively charged chlorines gener-ated from the disassociation of the newly formed N-halaminestructures. SEM studies demonstrated that the N-halamine-basedPU could effectively prevent the formation of bacterial and fungalbiofilms. These encouraging results point to the great potential ofthe new N-halamine-based PU for a wide range of long-term anti-microbial and biofilm controlling applications.

Acknowledgement

This study was sponsored by the NIH, NIDCR (Grant No.R01DE018707).

Appendix A. Figures with essential colour discrimination

Certain figures in this article, particularly Figure 3, are difficultto interpret in black and white. The full colour images can be foundin the on-line version, at doi:10.1016/j.actbio.2011.12.027.

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Table 5Durability and rechargeability of the N-halamines in the chlorinated PU–HDI–DMH.

Sample Chlorine atom area density(atoms cm�2)

Chlorineleft (%)

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recharging’’(1.74 ± 0.21) � 1016 98.6



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An N-halamine-based rechargeable antimicrobial and biofilm controlling polyurethane

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