Health and Safety via Surface Modification of ... - ACS Publications

Chapter 18

Downloaded by VIRGINIA TECH on July 17, 2013 | http://pubs.acs.org Publication Date (Web): July 8, 2013 | doi: 10.1021/bk-2013-1135.ch018

Health and Safety via Surface Modification of Polyurethanes Kenneth J. Wynne,*,1 Pinar Kurt,1 Kennard Brunson,1 Asima Chakravorty,1 Murari Gupta,1 Wei Zhang,1 Lynn Wood,2,3 and Dennis E. Ohman2,3 1Department

of Chemical and Life Science Engineering, Virginia Commonwealth University, 601 West Main Street, Richmond, Virginia 23284 2Department of Microbiology and Immunology, VCU Medical Center, Virginia Commonwealth University Richmond, 5-040 Sanger Hall, 1101 E. Marshall Street, Richmond, Virginia 23298 3McGuire Veterans Affairs Medical Center, 1201 Broad Rock Blvd., Richmond, Virginia 23249 *E-mail: [email protected].

Alkylammonium containing polymers are well known to effect contact kill of pathogenic bacteria. For coatings, alkylammonium function is needed only at the surface and not in the bulk. This paper describes a blend strategy that is “processing friendly” and aims at development of modifiers for conventional coating or molding operations. Specifically, the temporal instability of a polyurethane modifier with a copolyoxetane soft block having trifluoroethoxymethyl (3F) and C12 quaternary (dimethyldodecyl ammonium) side chains was overcome by utilizing a –CF2CF2H (4F) fluorous side chain and increasing the C12 mole fraction (U-4F-C12(0.34)). Characterization of a 2 wt% base polyurethane modified with 2 wt% U-4F-C12(0.34) validated a soft block surface modification model. Biocidal effectiveness (100% kill of spray on challenges of Gram+/- bacteria) was retained after storage for at least a month under ambient conditions of temperature © 2013 American Chemical Society In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.

and humidity. This stability was correlated with zeta potential measurements of accessible near surface charge. These results suggest zeta potential measurements hold promise as a physical method for assessing effectiveness of polycation contact kill and polycation stability in contact with aqueous systems.


1. Introduction Infection is a serious medical complication associated with health care environments. The 5-10% incidence of infections for hospital patients is well documented (1, 2) Sources of bacteria include health care equipment such as cell phones, pagers, computer keyboards, doorknobs, and electrocardiograph leads (3). Modes of transmission of antibiotic resistant pathogens occur in obvious ways such as touches, coughs, and sneezes. A study of Pennsylvania hospitals found that catheter associated urinary tract infections, designated CAUTIs, constituted over 60% of nosocomial infections (4). Women are particularly susceptible to UTIs (5). The majority of UTIs are caused by resistant strains of Escherichia coli (6) but infections are also caused by other Gram +/- bacterial strains (7). Biocidal polymers offer promise in helping curb the spread of infections by providing coatings for biomedical devices or molded articles. Such polymers may be classified as biocide release or non-release contact kill. Biocide release systems are the most advanced in that commercial products are available. A recent review covers recent advances in contact kill systems (8). A brief description of biocide release coatings is provided below followed by the development of surface modifiers that introduce contact kill into conventional polymers. A. Biocide Release The most commonly encountered strategy is biocide release. This approach is implemented by mixing polymeric materials with an FDA approved antibiotic or biocide (9–11). The active agent leaches from the tubing, coating, or other article and effects bacterial kill. A test of efficacy is carried out by placing a disk or similar object in contact with bacteria seeded on agar. A characteristic “zone of inhibition” forms a halo of killed bacteria around the article due to leached biocide. Although few laboratories report results, this is an important test to distinguish between biocide leaching and contact kill. Polymer materials employing release of silver (Ag+) are increasingly common (12, 13). However, some bacterial strains are inherently resistant to silver (14) and some develop resistance (15, 16). More research is needed to understand whether this situation is as serious as the buildup of antibiotic resistance and to understand effects on eukaryotic cells (17). Bacterial strains develop resistance against antibiotics since the mechanism of action is based on disruption of metabolic pathways via genetic alteration in cells. Some bacteria strains such as methicillin-resistant Staphylococcus aureus (MRSA) are of current concern because of increasing resistance to antibiotics (18, 19). 304 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.

B. Contact Kill

As the designation indicates, contact kill occurs when bacteria impinge on a solid surface. The two main classes of materials that effect contact kill do so either by a covalently bound surface oxidant that is renewed periodically or by membrane disruption, typically brought about by surface accessible positive charge.


1. Oxidative Kill

This area was pioneered by Worley and Sun, who showed that introduction of hydantoin generates oxidative biocidal surfaces after conversion of amide (-CON-H) to chloramide (-CON-Cl) (20–24). Oxidative kill is unparalleled for rapid kinetics, typically effecting 100% kill in a few minutes. Chloramine and related functionality must be renewed periodically by treatment with bleach, which is convenient for applications such as water disinfection and treatment of fabrics used in nosocomial environments (23, 25). Worley has continued to develop various strategies to effect oxidative kill and applications of this technology (21, 26–30). Inspired by this research, we developed hydantoin based surface modified polyurethanes (31). These findings on oxidative kill will not be reviewed in this article, which focuses on surface concentration of quaternary (quat) polycations.

2. Polycations at Surfaces

Contact kill occurs when a bacterium impinges on a surface with tethered polycations. As noted above, we focus on solid surfaces with quaternary nitrogen functionality. The mechanism by which surface quat functionality interacts with pathogenic organisms for contact kill has been discussed but the details are not clear (32–35). For coatings and thin films, a mechanistic analogy is drawn below to cell wall disruption proposed for solution borne polycations that have been studied extensively by authors including Tew (36–38), Kuroda (39), and Tiller (40). In a 1986 solution study, Ikeda showed that a 1 h exposure of Bacilus subtilis cells and protoplasts to polycations resulted in release of K+ (41). This release likely plays an important role in cell death as K+ concentration in cells is much higher than the surrounding medium. That is, thermodynamically driven K+ diffusion is one factor thought responsible for rapid cell lysis from osmotic shock (Eq 1). Solution borne polycations with multiple sites for conjugation through the formation of polysalts account for rapid chemisorption, release of K+, and the formation of insoluble bioconjugates (41). Along with release of K+ (or other inorganic cations) is likely that disruption of cell proton motive force (Δp) occurs (Δp results from the difference in pH between intracellular pH (pHi) and extracellular pH (pHo)) (42, 43). 305 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


Figure 1. A model for contact kill by disruption of bacterial membrane (negative, yellow) and K+ / H+ diffusion leading to cell death; coating (green, positive) has near surface accessible quaternary polycations, represented by red bars.

A combination of (1) negatively charged phospholipid exchange (8) with negatively charged counter ions associated with quat cations (Eq 1) and (2) outflow of K+ driven by osmotic pressure analogous to findings for water soluble polycations (41) and (3) disruption of cell proton motive force (Δp) seems likely for contact kill by surface concentrated polycations (Figure 1). A combination of these thermodynamically driven processes is consistent with 100% kill for high concentrations of sprayed on pathogen challenges in 30 min (44) as diffusion of species such as K+ and H+ would be rapid. While evidence for proposed mechanisms of kill is difficult to acquire (8), Santore has confirmed that bacterial chemisorption to solid surfaces bearing a positive charge is strongly favored (45).

2. Quats at Surfaces Figure 2 provides a schematic that represents homogeneous, covalently bound polycations (Figure 2A) and surface modified coatings (Figure 2B). Either can give surface accessible quaternary function. 306 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


Figure 2. Schematic for A, polymers containing quaternary function and B, surface modified coatings.

A. Homo-Quats A good deal of research has described the homogeneous incorporation of alkylammonium function in coatings (46–49), and thin films (50, 51). An example of homogeneous incorporation of quaternary function is provided by Cooper’s report of chain extender modification (33). Depending on the percent alkylpyridinium content, water uptake was 22 to 49 wt %, which negatively affected mechanical properties. For optimum compositions, contact kill was up to 95% against a sprayed challenge of Staphylococcus aureus. Hard block functionality trapped in the bulk may have resulted in an insufficient surface accessible quaternary charge for higher efficiency. With a similar functional group, quaternized polystyrene-b-poly(4-vinylpyridine) copolymers containing fluorous moieties showed good antimicrobial activity against Staphylococcus aureus. Bactericidal effectiveness was correlated with the molecular composition and organization in the top 2-3 nm (52). Quaternized polyethylene imine has been extensively studied by Klibanov (53). Thin coatings have been found to generate highly effective C12-quaternary antimicrobial surfaces. Thus, in the absence of inter- or intramolecular associative forces that sequester charge, surface accessible C12-quat function is robust and strongly antimicrobial (54).

B. Surface Modification Coatings with quaternary function concentrated at the surface are depicted in Figure 2B. Surface modified coatings represented by Figure 2B have been obtained by several methods including plasma treatment and subsequent functionalization (55, 56) and photografting (57) and grafting to or from surfaces (32, 51, 57). Due to entropic considerations, chain ends tend to be excluded from the bulk (58). End group surface concentration was used by Tiller in the photopolymerization of acrylate monomers and crosslinkers along with polyoxazoline macromonomers having biocidal quarternary ammonium end groups (59). 307 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.

In the research described below, we focus on a straightforward blend method for surface functionalization of polyurethanes. Because this approach involves an additive for conventional polyurethanes, this blend surface modification may be translated to a large scale process.


1. Generation 1 Surface Modifiers (SMs) The attractive features of “processing friendly” surface modifiers motivated our research in polyurethanes (31, 60, 61). Prior work showed that polyurethane soft blocks are surface-concentrated (62–64). Thus, we have explored functional P[AB]-copolyoxetane diols 1 in a new approach to surface modification. Here, A and B designate repeat units or side chains. A model for this concept applied to quaternary surface function is shown in Figure 3. The surface concentrated functional soft block of the SM is shown in red along with side chains A and B. Economy is thus ensured by using the P[AB]-soft block polyurethane as a minor constituent in the blend.

Figure 3. Schematic of P[AB]-polyurethane surface modification. Here, A is a fluorous group and B is a C12 quat. The model shown in Figure 3 has been used to guide the development of quaternary alkylammonium antimicrobial SMs (31, 44, 65). Herein, we focus on SMs with fluorous side chains A having Rf = -CF3 (3F) and -CF2CF2H (4F), 308 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


respectively and B = C12 quat. Both of these modifiers effect contact kill due to quat functionality (44, 65) but the morphological and functional stability of the modified surfaces are dramatically different. The synthesis, initial surface characterization and biotesting for the 3F modifier have been published (44, 65). Details for the 4F modifier, which was prepared in a similar fashion, will be published separately. The bulk polyurethane is H12MDI-BD(50)-PTMO-1000, where the hard block is 4,4′-(methylene bis(p-cyclohexyl isocyanate) (H12MDI) and butanediol (BD) and poly(tetramethylene oxide) PTMO-1000 is the soft block. This base polyurethane with a Shore D hardness of 47 ±2 was selected for its similarity to commercially available aliphatic polyurethanes that are used in many applications such as biomedical materials and sports equipment. A notable difference for the base polyurethane described herein is a purification scheme that ensures no water contamination during a sensitive dynamic contact angle experiment (66). The focus of initial discussion is a “first generation” SM H12MDI-BD(50)3F-C12(0.11), Figure 3, (1-q) = 0.11. This SM is designated U-3F-C12(0.11). Mn for the soft block is 5.1 kDa. Coatings were generated by dipping or spreading tetrahydrofuran (THF) or THF-isopropanol (THF-IPA) solutions of 2 wt% U-3FC12(0.11) and base polyurethane. The compositional rationale and other details may be found elsewhere (44). 2 wt% U-3F-C12(0.11) coatings were tested against aerosol challenges (106 CFU/mL) of Gram negative (Pseudomonas aeruginosa, Escherichia coli) and Gram positive (Staphylococcus aureus) bacteria. After 30 min residence, bacteria were stripped, serially diluted and plated for colony counts. 2 wt% U-3F-C12(0.11) coatings effected 100% kill, a 3.6-4.4 log reduction of Gram +/bacteria. A zone of inhibition test showed no biocide release for SM modified compositions. This encouraging result was attributed to good block/soft block phase separation, a cation/co-repeat group ratio mimicking natural biocidal proteins (67), and a semifluorinated “chaperone” or low surface energy moiety that aids in alkylammonium surface-concentration.

2. Generation 1 Not Stable!

A denouement began with biocidal testing on the same coatings as a function of time to assess durability of contact kill. Figure 4 shows aerosol test results for 2 wt% U-3F-C12(0.11) coatings within a few days and after two weeks at ambient temperature. To our surprise, the modified coating had lost biocidal effectiveness. Several sets of tests showed that contact biocidal kill for 2 wt% PU-8-5100 ranged from 0-50%. To gain insight into loss of contact biocidal effectiveness, TM-AFM imaging was performed as a function of time. Figure 5 shows the TM-AFM 3D height image for a 2 wt% U-3F-C12(0.11) coating after 2 d and 2 wk at ambient temperature. One to two days after the coating was generated, the image in Figure 5A was almost featureless, as reported earlier (44). 309 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


Figure 4. Agar plates after a spray challenge of P. aeruginosa: Left, base polyurethane control; Right, 2 wt% U-3FC12(0.11).

The 3D height image shown in Figure 5B shows a radical change after two weeks aging at ambient temperature. The vertical (z scale) is increased to 1000 nm compared to the image in Figure 5A to capture the formation of multiple features up to ~10 μm wide and 100 – 200 nm high. Longer aging in air results in further complexities that we assign to surface phase separation of the U-3F-C12(0.11) modifier.

Figure 5. TM-AFM 3D height image for a 2 wt% U-3F-C12(0.11) coating after 2 d (A) and 2 wk (B) at ambient temperature.

Based on biotesting and AFM imaging, we now realize that the solution coating process results in a metastable state. Quaternary charge is accessible and contact antimicrobial effectiveness is high. With time, phase separation occurs, partly driven by the immiscibility of the 3F side chains. The phase separated domains must sequester quaternary charge leading to ineffective contact antimicrobial kill. 310 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


3. Streaming Potential Measurements for Quantifying near Surface Quaternary Charge Solution test methods provide quantitative estimates of antimicrobial effectiveness including minimum inhibitory concentrations (MICs), minimum biocidal concentrations (MBCs) and kill kinetics (68). However, methods for estimating biocidal effectiveness for surface bound polycation coatings are widely varying and do not often permit unambiguous comparisons. Regardless of the procedure, testing coatings for antimicrobial effectiveness is tedious and carries an element of personal risk, as Gram negative (Pseudomonas aeruginosa, Escherichia coli) and Gram positive (Staphylococcus aureus) are opportunistic pathogenic bacteria. A rapid, quantitative measurement of surface accessible quaternary ammonium charge was therefore sought. Measurement of quaternary charge density has often employed fluorescein dye binding and release of bound dye by an ion exchange surfactant such as dodecyl trimethyl ammonium chloride (34, 69–72). This method failed for polyurethanes modified with quaternary SMs because of nonspecific dye adsorption and slow desorption (73). Streaming potential measurements have long been known for quantifying surface charge, particularly on colloidal particles (74). Assessing surface accessible charge for thin films formed by alternating polyelectrolyte deposition was elegantly demonstrated by Adamczyk (75). Alternating positive and negative potentials attested to the charge of the last polyelectrolyte deposited. Streaming potentials have also been employed for studying antimicrobial function of polyelectrolyte multilayers comprising poly(allyl amine) hydrochloride and poly(sodium 4-styrene sulfonate (76).

Figure 6. Streaming potentials for a 2 wt% U-3F-C12(0.11) coating as a function of capillary pumping cycle (~20 s per cycle).

In an initial examination of streaming potential measurements on modified polyurethanes the microfluidic method developed by Alvarez (77) was adapted (73). The inside of a 100 μm capillary was coated with the desired polyurethane and streaming potentials were obtained using a 1 mM KBr solution (Figure 6) (73). The good news was an initial streaming potential above 100 mV for a 2 wt% U-3FC12(0.11) coating. The high near surface accessible charge agrees with excellent 311 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


near term contact antimicrobial effectiveness (Figure 4, 1-2 d). The bad news was a rapid decrease in streaming potentials over the course of only four pumping cycles (73). Streaming potentials scale to charge density so that the results for the 2 wt% U-3F-C12(0.11) coating indicate a precipitous decrease (~50%) in accessible near surface quaternary charge in < 2 min. Compared to the relatively slow (2 wk) deactivation of quaternary charge density in air (biotesting, AFM), streaming potential measurements show loss of surface accessible positive charge for 2 wt% U-3F-C12(0.11) takes only a few minutes. Apparently, water plasticization of the near surface SM domain results in an acceleration of phase separation that sequesters quat function.

Figure 7. Image of a 2 wt% U-4F-C12(0.34) coated glass slide.

4. Generation 2 Modifier The unexpected temporal instability of 2 wt% U-3F-C12(0.11) has led to the development of alternative SMs that will achieve long term contact antimicrobial effectiveness. One of these is based on the replacement of –CF3 by –CF2CF2H (Figure 3). H-bonding was proposed by Ellison to explain decreased surface tension upon replacement of terminal -CF3 by –CF2H (78). Thus replacement of 3F with amphiphilic 4F seemed attractive based on the notion that CF2H hydrogen bonding would provide an enthalpic contribution enhancing the free energy of mixing. The polyurethane modifier U-4F-C12(0.34) was prepared by a method paralleling that described previously for the 3F analog (Figure 3, Rf = -CF2CF2H, q = 0.34) (44). The weight fraction of C12 was increased to 0.34 compared to 0.11 for generation 1 modifier based on preliminary experiments that showed enhanced modified surface stability. By 1H-NMR end group analysis, Mn for 4F-C12(0.34) diol was 6.0 kDa. Copolyoxetane diol and polyurethane solution characterization (1H-NMR, GPC) will be provided in a separate publication. Unlike the 3F polyurethane SM, U-4F-C12(0.34) was insoluble in THF. DMAC (dimethyl acetamide) was used as an alternative solvent for deposition of 2 wt% U-4F-C12(0.34). An overnight thermal treatment in vacuum (120 °C) removed solvent. Optically transparent coatings resulted (Figure 7); coating thickness was typically 160 μm. 312 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


Based on the hypothesis that stable, accessible near surface charge would be a predictor of long term antimicrobial effectiveness, zeta potential measurements were a priority. As noted above, we developed a capillary method for this purpose (73). However, coating the inside of a 100 μm capillary is tedious and subsequent characterization of the coating surface problematical. In a new approach, zeta potential measurements were carried out on drip coated microscope slides using a SurPASS electrokinetic analyzer and clamping cell (Anton PAAR). For these initial measurements, pH was constant at ~7. For the SurPASS, each run consists of a forward and backward pumping cycle taking ~15 min. Three runs (~ 45 min) provided a more demanding test of stability in dilute polyelectrolyte (10-3 M KBr) compared to the capillary method (~ 2 min) used previously (73).

Table 1. Zeta potentials for 2 wt% U-4F-C12(0.34) relative to base polyurethane Zeta potential (mV) Run

1 wk

4 wks

1

42.1

39.8

2

41.0

37.6

3

38.8

35.8

Reported here are initial zeta potential measurements for 2 wt% U-4F-C12(0.34) relative to base polyurethane HMDI-BD(50)-PTMO(1000). Zeta potentials for polymer coatings are known to be affected by pH (79). Initial measurements reported here were carried out at pH 7. Compared to base polyurethane (-37± 4 mV), zeta potentials for 2 wt% U-4FC12(0.34) were positive. Table 1 lists zeta potentials relative to base polyurethane at < 1 wk and > 4 wk. Zeta potentials for 2 wt% U-4F-C12(0.34) were diminished 5-10% after one month storage in air at ambient temperature. Each set of zeta potential tests amounted to ~2 h immersion in 1 mM NaBr. Interestingly, the diminution observed over the course of the three runs for the first test at < 1 wk (42.1 → 38.8 mV) was restored to some degree for the first run after 4 wk in air (39.8 mV). However, the final value after four weeks “aging” in air (35.8 mV). was10% less than the corresponding value after 1 wk. The stability of zeta potentials for 2 wt% U-4F-C12(0.34) supports the hypothesis that -CF2H hydrogen bonding provides enthalpic stabilization of the modified surface. However, a detailed study is warranted to confirm these initial results. Spray challenges of Pseudomonas aeruginosa, Escherichia coli, and Staphylococcus aureus were carried out as described previously (44). The residence time for each bacterial challenge (106 cfu/ml) was 1 h. Table 2 shows 313 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.

percent kill for the pathogen challenges after one hour residence time. Within experimental error, complete kill was observed within 1 wk and after 4 wks at ambient temperature. These results are important in validating the model shown in Figure 3 for surface concentration of the U-4F-C12(0.34) modifier. Importantly, the biocidal effectiveness after storage of at least a month under ambient conditions of temperature and humidity also provide support for the use of zeta potentials as a predictor for polycation contact kill.


Table 2. Percent kill of spray challenges for U-4F-C12-34. Residence time for each challenge (106 cfu/ml) was 1 h Challenge

< 1 wk

> 4 wk

Pseudomonas aeruginosa

100

95

Escherichia coli

100

100

Staphylococcus aureus

99

90

3. Conclusion Fluorous side chain A in polyurethanes with P[AB] copolyoxetane soft blocks act as chaperones for surface concentration of functional side chain B. However, unanticipated new surface morphologies and new surface phenomena can arise from A-side chain / B-side chain interactions invalidating the simple model shown in Figure 3. In the present work, the temporal instability of the U-3F-C12(0.11) modifier was avoided by utilizing a –CF2CF2H (4F) fluorous side chain and increasing the C12 mole fraction. The results for U-4F-C12(0.34) are important in validating the model shown in Figure 3. Biocidal effectiveness was demonstrated after storage for at least a month under ambient conditions of temperature and humidity. This stability was correlated with zeta potential measurements of accessible near surface charge. These results suggest zeta potential measurements hold promise as a physical method for assessing effectiveness of polycation contact kill and polycation stability in contact with aqueous systems. In summary, polyurethanes with P[AB] copolyoxetane soft blocks are effective and potentially processing-compatible surface modifiers for conventional polyurethanes. This surface modification approach is “green” in the sense that only a minimal amount of surface modifier (2 wt% or less) is leveraged so as to effect a radical change in surface function. In addition, the 4F chaperone is “green” in avoiding the use of perfluorinated eight carbon moieties that are likely PFOA precursors.

314 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.

Acknowledgments K.J.W. thanks the National Science Foundation (DMR- grants DMR0207560, DMR-0802452, and DMR-1206259) and the VCU School of Engineering Foundation for support of this research. This work was also supported by Public Health Service Grant AI-19146 from the National Institute of Allergy and Infectious Disease (D.E.O.), and in part by Veterans Administration Medical Research Grant I01BX000477 (D.E.O.). K.J.W. also thanks Ray Ottenbrite for being a great colleague and friend for over 30 years.


References 1. 2. 3.

4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19.

20. 21. 22.

Friedman, N. D.; Korman, T. M.; Fairley, C. K.; Franklin, J. C.; Spelman, D. W. J. Infect. 2002, 45, 47. Hacek, D. M.; Suriano, T.; Noskin, G. A.; Krusynski, J.; Reisberg, B.; Peterson, L. R. Am. J. Clin. Pathol. 1999, 111, 647. Borer, A.; Gilad, J.; Smolyakov, R.; Eskira, S.; Peled, N.; Porat, N.; Hyam, E.; Trefler, R.; Riesenberg, K.; Schlaeffer, F. Emerging Infect. Dis. 2005, 11, 1160. Maki, D. G.; Tambyah, P. A. Emerging Infect. Dis. 2001, 7, 342. Nicolle, L. E. Women’s Health 2005, 1, 39. Shigemura, K.; Tanaka, K.; Adachi, M.; Yamashita, M.; Arakawa, S.; Fujisawa, M. J. Infect. Chemother. 2011, 17, 646. Shigemura, K.; Arakawa, S.; Tanaka, K.; Fujisawa, M. J. Infect. Chemother. 2009, 15, 18. Bieser, A. M.; Tiller, J. C. Macromol. Biosci. 2011, 11, 526. Medlin, J. Environ. Health Perspect. 1997, 105, 290. Hazzizalaskar, J.; Helary, G.; Sauvet, G. J. Appl. Polym. Sci. 1995, 58, 77. Nurdin, N.; Helary, G.; Sauvet, G. J. Appl. Polym. Sci. 1993, 50, 663. Muller, G.; Winkler, Y.; Kramer, A. J. Hosp. Infect. 2003, 53, 211. Holder, I. A.; Durkee, P.; Supp, A. P.; Boyce, S. T. Burns 2003, 29, 445. Johnson, J. R.; Delavari, P.; Azar, M. Antimicrob. Agents Chemother. 1999, 43, 2990. Silver, S. FEMS Microbiol. Rev. 2003, 27, 341. Silver, S.; Phung, L. T.; Silver, G. J. Ind. Microbiol. Biotechnol. 2006, 33, 627. Marambio-Jones, C.; Hoek, E. M. V. J. Nanopart. Res. 2010, 12, 1531. Diekema, D. J.; Pfaller, M. A.; Schmitz, F. J.; Smayevsky, J.; Bell, J.; Jones, R. N.; Beach, M. Clin. Infect. Dis. 2001, 32, S114. Borer, A.; Gilad, J.; Yagupsky, P.; Peled, N.; Porat, N.; Trefler, R.; ShprecherLevy, H.; Riesenberg, K.; Shipman, M.; Schlaeffer, F. Emerging Infect. Dis. 2002, 8, 966. Worley, S. D.; Sun, G. Trends Polym. Sci. 1996, 4, 364. Eknoian, M. W.; Worley, S. D. J. Bioact. Compat. Polym. 1998, 13, 303. Eknoian, M. W.; Worley, S. D.; Bickert, J.; Williams, J. F. Polymer 1999, 40, 1367. 315

In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


23. Sun, G.; Worley, S. D. In Modified Fibers with Medical and Specialty Applications; Edwards, J. V., Buschle-Diller, G., Eds.; Springer: Dordrecht, The Netherlands, 2006; p 81. 24. Kenawy, E. R.; Worley, S. D.; Broughton, R. Biomacromolecules 2007, 8, 1359. 25. Sun, G.; Worley, S. D. J. Chem. Educ. 2005, 82, 60. 26. Sun, G.; Allen, L. C.; Luckie, E. P.; Wheatley, W. B.; Worley, S. D. Ind. Eng. Chem. Res. 1995, 34, 4106. 27. Kocer, H. B.; Worley, S. D.; Broughton, R. M.; Huang, T. S. React. Funct. Polym. 2011, 71, 561. 28. Cerkez, I.; Kocer, H. B.; Worley, S. D.; Broughton, R. M.; Huang, T. S. React. Funct. Polym., 72, 673. 29. Cerkez, I.; Kocer, H. B.; Worley, S. D.; Broughton, R. M.; Huang, T. S. J. Appl. Polym. Sci., 124, 4230. 30. Cerkez, I.; Kocer, H. B.; Worley, S. D.; Broughton, R. M.; Huang, T. S. Cellulose, 19, 959. 31. Makal, U.; Wood, L.; Ohman, D. E.; Wynne, K. J. Biomaterials 2006, 27, 1316. 32. Lee, S. B.; Koepsel, R. R.; Morley, S. W.; Matyjaszewski, K.; Sun, Y. J.; Russell, A. J. Biomacromolecules 2004, 5, 877. 33. Grapski, J. A.; Cooper, S. L. Biomaterials 2001, 22, 2239. 34. Tiller, J. C.; Liao, C. J.; Lewis, K.; Klibanov, A. M. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 5981. 35. Lin, J.; Qiu, S. Y.; Lewis, K.; Klibanov, A. M. Biotechnol. Prog. 2002, 18, 1082. 36. Tew, G. N.; Scott, R. W.; Klein, M. L.; De Grado, W. F. Acc. Chem. Res. 2009, 43, 30. 37. Som, A.; Navasa, N.; Percher, A.; Scott, R. W.; Tew, G. N.; Anguita, J. Clin. Vaccine Immunol. 2012, 19, 1784. 38. Tew, G. N.; Scott, R. W.; Klein, M. L.; Degrado, W. F. Acc. Chem. Res. 2010, 43, 30. 39. Sovadinova, I.; Palermo, E. F.; Urban, M.; Mpiga, P.; Caputo, G. A.; Kuroda, K. Polymers 2011, 3, 1512. 40. Waschinski, C. J.; Barnert, S.; Theobald, A.; Schubert, R.; Kleinschmidt, F.; Hoffmann, A.; Saalwachter, K.; Tiller, J. C. Biomacromolecules 2008, 9, 1764. 41. Ikeda, T.; Hirayama, H.; Yamaguchi, H.; Tazuke, S.; Watanabe, M. Antimicrob. Agents Chemother. 1986, 30, 132. 42. Kashket, E. R. Annu. Rev. Microbiol. 1985, 39, 219. 43. Kakinuma, Y. Microbiol. Mol. Biol. Rev. 1998, 62, 1021. 44. Kurt, P.; Wood, L.; Ohman, D. E.; Wynne, K. J. Langmuir 2007, 23, 4719. 45. Gon, S.; Kumar, K. N.; Nusslein, K.; Santore, M. M. Macromolecules 2012, 45, 8373. 46. Majumdar, P.; Lee, E.; Patel, N.; Ward, K.; Stafslien, S. J.; Daniels, J.; Chisholm, B. J.; Boudjouk, P.; Callow, M. E.; Callow, J. A.; Thompson, S. E. M. Biofouling 2008, 24, 185. 316 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


47. Pant, R. R.; Buckley, J. L.; Fulmer, P. A.; Wynne, J. H.; McCluskey, D. M.; Phillips, J. P. J. Appl. Polym. Sci. 2008, 110, 3080. 48. Majumdar, P.; He, J.; Lee, E.; Kallam, A.; Gubbins, N.; Stafslien, S. J.; Daniels, J.; Chisholm, B. J. J. Coat. Technol. Res. 2010, 7, 455. 49. Ye, S. J.; Majumdar, P.; Chisholm, B.; Stafslien, S.; Chen, Z. Langmuir 2010, 26, 16455. 50. Milovic, N. M.; Wang, J.; Lewis, K.; Klibanov, A. M. Biotechnol. Bioeng. 2005, 90, 715. 51. Huang, J. Y.; Koepsel, R. R.; Murata, H.; Wu, W.; Lee, S. B.; Kowalewski, T.; Russell, A. J.; Matyjaszewski, K. Langmuir 2008, 24, 6785. 52. Krishnan, S.; Ward, R. J.; Hexemer, A.; Sohn, K. E.; Lee, K. L.; Angert, E. R.; Fischer, D. A.; Kramer, E. J.; Ober, C. K. Langmuir 2006, 22, 11255. 53. Schaer, T. P.; Stewart, S.; Hsu, B. B.; Klibanov, A. M. Biomaterials 2012, 33, 1245. 54. Park, D.; Wang, J.; Klibanov, A. M. Biotechnol. Prog. 2006, 22, 584. 55. Gottenbos, B.; van der Mei, H. C.; Klatter, F.; Nieuwenhuis, P.; Busscher, H. J. Biomaterials 2002, 23, 1417. 56. Vasilev, K.; Griesser, S. S.; Griesser, H. J. Plasma Processes Polym. 2011, 8, 1010. 57. Huang, J.; Murata, H.; Koepsel, R. R.; Russell, A. J.; Matyjaszewski, K. Biomacromolecules 2007, 8, 1396. 58. Jalbert, C.; Koberstein, J. T.; Hariharan, A.; Kumar, S. K. Macromolecules 1997, 30, 4481. 59. Waschinski, C. J.; Tiller, J. C. Biomacromolecules 2005, 6, 235. 60. Makal, U.; Wynne, K. J. Langmuir 2005, 21, 3742. 61. Wynne, K. J.; Makal, U.; Kurt, P.; Gamble, L. Langmuir 2007, 23, 10573. 62. Andrade, J. D.; Tingey, K. G. Langmuir 1991, 7, 2471. 63. Garrett, J. T.; Lin, J. S.; Runt, J. Macromolecules 2002, 35, 161. 64. Ratner, B. D.; Yoon, S. C.; Kaul, A.; Rahman, R. In Polyurethanes in biomedical engineering II; Planck, H., Syre, I., Dauner, M., Egbers, G., Eds.; Elsevier: New York, 1986; Vol. 3, p 213. 65. Kurt, P.; Gamble, L. J.; Wynne, K. J. Langmuir 2008, 24, 5816. 66. Uilk, J. M.; Mera, A. E.; Fox, R. B.; Wynne, K. J. Macromolecules 2003, 36, 3689. 67. Tew, G. N.; Liu, D. H.; Chen, B.; Doerksen, R. J.; Kaplan, J.; Carroll, P. J.; Klein, M. L.; DeGrado, W. F. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 5110. 68. Chakrabarty, S.; King, A.; Kurt, P.; Zhang, W.; Ohman, D. E.; Wood, L. F.; Lovelace, C.; Rao, R.; Wynne, K. J. Biomacromolecules 2011, 12, 757. 69. Goddard, J. M.; Hotchkiss, J. H. Prog. Polym. Sci. 2007, 32, 698. 70. Kugler, R.; Bouloussa, O.; Rondelez, F. Microbiology 2005, 151, 1341. 71. Ledbetter, J. W., Jr.; Bowen, J. R. Anal. Chem. 1969, 41, 1345. 72. Murata, H.; Koepsel, R. R.; Matyjaszewski, K.; Russell, A. J. Biomaterials 2007, 28, 4870. 73. Gupta, M. L.; Brunson, K.; Chakravorty, A.; Kurt, P.; Alvarez, J. C.; LunaVera, F.; Wynne, K. J. Langmuir 2010, 26, 9032. 74. Stanley, J. S. J. Phys. Chem. 1954, 58, 533. 317 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.


75. Adamczyk, Z.; Zembala, M.; Warszynski, P.; Jachimska, B. Langmuir 2004, 20, 10517. 76. Lichter, J. A.; Rubner, M. F. Langmuir 2009, 25, 7686. 77. Pu, Q. S.; Elazazy, M. S.; Alvarez, J. C. Anal. Chem. 2008, 80, 6532. 78. Ellison, A. H.; Fox, H. W.; Zisman, W. A. J. Phys. Chem. 1953, 57, 622. 79. Kirby, B. J.; Ernest, F.; Hasselbrink, J. Electrophoresis 2004, 25, 203.

318 In Tailored Polymer Architectures for Pharmaceutical and Biomedical Applications; Scholz, C., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2013.

Health and Safety via Surface Modification of ... - ACS Publications

Recommend Documents