Poly(oxazoline)s with Telechelic Antimicrobial Functions

Sitaraman Krishnan, Rebekah J. Ward, Alexander Hexemer, Karen E. Sohn, Kristen L. Lee, Esther ... Jordan , Marina Sokolski-Papkov , Alexander V. Kaban...
0 downloads 0 Views 211KB Size
Download PDF
Biomacromolecules 2005, 6, 235-243

235

Poly(oxazoline)s with Telechelic Antimicrobial Functions Christian J. Waschinski and Joerg C. Tiller* Freiburg Materials Research Center and Institute for Macromolecular Chemistry, Department of Chemistry, Albert-Ludwigs-Universita¨t Freiburg, Stefan-Meier-Str. 21, 79104 Freiburg, Germany Received August 3, 2004; Revised Manuscript Received September 29, 2004

Poly(2-alkyl-1,3-oxazoline)s (alkyl ) methyl, ethyl) with terminal quarternary ammonium groups were synthesized. It could be shown by NMR and ESI-MS that the termination of the living polymerization with N,N-dimethylalkyl(butyl to hexadecyl)amines was quantitative. The novel functions were investigated regarding their antimicrobial potential toward the bacterium Staphylococcus aureus revealing that only quarternary ammonium functions with 12 and more carbons are antibacterial. Using a novel bifunctional initiator, 3-[(tert-butoxycarbonyl)amino]benzyl-p-toluenesulfonate, poly(oxazoline) with a primary amino group at the starting end and an antimicrobial function at the terminal could be synthesized, as confirmed by NMR and ESI-MS measurements. Comparing the bioactivity of polymers with different functions at the starting end and terminated with dimethyldodecylamine revealed that the starting group has a great effect on the antibacterial properties of the distant terminal. The minimal inhibitory concentrations varied from 0.1 mM for polymer derivatives with a BOC-NH-phenyl starting group to 4 mM for poly(oxazoline)s with a free primary amine at the starting end. Introduction Spreading microbial infections and increasing numbers of resistant microbial strains are great problems of the modern society and medicine. It constantly demands new antimicrobial agents, e.g., antibiotics, and materials that do not allow microbes to survive on their surfaces. Polymeric biocides may provide solutions to both of these problems, because they have a great antimicrobial potential in dissolved form and are capable of rendering materials surfaces upon grafting in a way that the surfaces kill microbes on contact.1,2 Antimicrobial polymers have been known since 1979 and are usually based on a nonbioactive polymer backbone that is modified with antimicrobial functions at the side chains.3 Naturally, the antimicrobial activity of a single biocidal group did not increase upon attaching it to a polymer side group; that is, the antimicrobial activity of polymers is often lower or equal than the sum of the attached bioactive functions in free state. Nevertheless, the polymer considered as one molecule exhibits sometimes a higher antimicrobial activity than the respective low molecular weight compound. Although the mechanism of antibiotics is well understood in many cases, the antimicrobial action of polymers is mostly not known. Yet, the few existing studies reveal the potential of antimicrobial polymers. For instance, although quarternary ammonium polymers such as poly(hexylene-N,N-dimethylimine)s are more active against bacteria than the respective monomers,4 quarternerized chitosan derivatives show lower antimicrobial activity compared to the respective monomer units.5 In the case of poly(4-vinyl-N-alkylpyridinium) salts, * To whom correspondence should be addressed. Tel.: +49 761 203 4735. Fax: +49 761 203 4709. E-mail: [email protected].

the polymers are not only more active than their monomers, but also obviate the resistance mechanism of Gram-positive bacteria against quarternary ammonium compounds.6 Even less is known about surface grafted polymers. Yet, in the light of recent health concerns, it is of great importance to understand the mechanism of such biocidal polymers and surfaces to be capable of creating new weapons against infectious microbes. All known antimicrobial polymers can be understood as polymerized biocides. Therefore the role of the polymer backbone is hard to distinguish from the multiplied activity of the antimicrobial functions attached to it. To explore the influence of the polymer backbone to the function of antimicrobial structures tethered to the polymer, it is from our perspective necessary to attach one bioactive function to the end of a polymer well defined in length and nature. Many antibiotics and disinfectants have their target structure in the cytoplasm or the cytoplasm membrane of bacteria. This means that these biocides must diffuse through the microbial cell wall, which is a network consisting of polysaccharides cross-linked with short peptides (Figure 1). Therefore, an active biocide-terminated polymer must be water soluble and should not have interactions with the microbial cell wall.3b These two requirements are met best by the polymers poly(ethylene glycol) (PEG) and various poly(2-alkyl-1,3-oxazoline)s (POX). Both types of polymers repel bacteria indicating that they do not bind to the surface of such microorganisms.7,8 Another advantage of those polymers is their preparation via a living polymerization mechanism, resulting in macromolecules of almost uniform chain lengths.9,10 The microbe-repelling properties and uniform length make PEG and POX ideal candidates to explore the mechanism of grafted antibacterial functions.

10.1021/bm049553i CCC: $30.25 © 2005 American Chemical Society Published on Web 11/13/2004

236

Biomacromolecules, Vol. 6, No. 1, 2005

Figure 1. Principle of the interaction between an antimicrobial function at the terminal of a polymer and a Gram-positive bacterium.

PEG, prepared by means of anionic polymerization, is one of the best investigated biomedical polymers. It has been proven to preserve the activity of drugs, e.g., cytokines, after being linked with the polymer.11 Antimicrobial functions attached to PEG were rarely explored.12 The other suited polymers are poly(oxazoline)s which have been prepared by cationic polymerization leading to narrowly distributed macromolecules.13 Several compounds, such as double bonds, epoxy-functions, siloxanes, hydroxyl groups, and amines, were introduced as functional end-groups.14 The obtained polymers were used as polymeric surfactants, as cross-linker in amphiphilic networks, and as antifouling coatings.15 Surprisingly, poly(oxazoline)s were not yet investigated as covalent drug-polymer-conjugates. Here, we report on novel POX derivatives carrying an antimicrobial end-group, which can penetrate bacterial cell walls. Some POX-derivatives were further modified with an NH2 function at the starting end of the polymer to give narrowly distributed bitelechelic polymers. Experimental Part Materials. The monomers, 2-methyl-1,3-oxazoline (MOX, Fluka) and 2-ethyl-1,3-oxazoline (EOX, Merck) were freshly distilled from KOH prior to use. Pyridine and chloroform (all Roth) were freshly distilled prior to use. PEG monomethyl ether mesylate (average molecular weight ) 5000 g/mol) was obtained from Fluka. All other compounds were of analytical grade or purer and used without further purification. All distillations and reactions were carried out under an atmosphere of argon. Instrumentation. 1H and 13C NMR spectra were recorded using a Bruker ARX 300 spectrometer operating at 300 MHz for 1H and 75.4 MHz for 13C. Gel permeation chromatography (GPC) was carried out on a Knauer Mikrogelset C11 equipped with an LS- and RI-detector and using DMF as eluent and poly(styrene) and poly(ethyloxazoline) standards (all standards from Aldrich) at 25 °C. UV-vis analyses were performed using a Perkin-Elmer Lambda 11 UV-vis spectrometer. MALDI-TOF-MS-spectra were collected by using a Bruker Reflex II mass spectrometer following a procedure given by Jordan et al.14c ESI-MS-spectra were recorded using a Finnigan Mat TSQ 700 mass spectrometer in positive

Waschinski and Tiller

mode. The polymer samples were dissolved in aqueous solutions containing 50 vol.-% MeOH and 6 vol.-% acetic acid to obtain solutions with c ) 10 pmol/µL and then injected into the mass spectrometer with 10 µL/min. Fluorescence spectra were recorded using a Perkin-Elmer LS 50B luminescence spectrometer. Syntheses. Initiator Synthesis. 3-[(tert-Butoxycarbonyl)amino]benzyl alcohol (0a). Modifying a general procedure of Moroder et al.,16 di-tert-butyl-dicarbonate (7.09 g; 32.5 mmol) was added dropwise to a solution of 4.00 g (32.5 mmol) of 3-aminobenzyl alcohol in 15 mL of CHCl3, at room temperature. After stirring the mixture for another 2 h, the solution was concentrated in vacuo followed by washing the crude product with water to give 5.92 g (82%) of the slightly yellow product. 1 H NMR (CDCl3): δ (ppm) ) 7.40 (m, 1H, NCCHCHarCHCCH2OH), 7.30-7.20 (m, 2H, NCCHarCHCHarCCH2OH), 7.10-7.02 (m, 1H, NCCHarCCH2OH), 6.69 (br s, 1H, NH), 4.63 (s, 2H, CH2OH), 2.40 (br s, 1H, OH), 1.54 (s, 9H, C(CH3)3). 13 C NMR (CDCl3): δ (ppm) ) 152.9 (N-COO), 141.9 (Car-CH2-OH), 138.4 (Car-NH), 128.9 (NCCHCarCHCCH2OH), 121.4 (NCCHCCarHCCH2OH), 117.6 (NCCarHCCH2OH), 117.1 (NCCarHCCHCCH2OH), 80.4 (-C(CH3)3), 64.6 (C6H4CH2OH), 28.2 (C(CH3)3). C12H17NO3 (223.27): Calcd (%): C, 65.55; H, 7.69; N, 6.27. Found (%): C, 65.34; H, 7.51; N, 6.47. 3-[(tert-Butoxycarbonyl)amino]benzyl-p-toluenesulfonate (0b). According to a varied literature method,17 ptoluenesulfonyl chloride (1.13 g; 5.9 mmol) was slowly added to a mixture containing 1.32 g (5.9 mmol) of 3-[(tertbutoxycarbonyl)amino]benzyl alcohol 0a, 0.02 g (0.24 mmol) of tetrabutylammonium hydrogen sulfate, 10 mL of chloroform, and 5 mL of 30% aqueous NaOH solution under stirring at -5 °C. After maintaining the mixture at 0 °C for 2 h, the organic layer was washed three times with water and dried over anhydrous Na2SO4. Chromatography on silica gel using chloroform as eluent afforded 1.00 g (90%) of the product. 1 H NMR (CDCl3): δ (ppm) ) 7.78 (d, 2H, 3JH,H ) 8.60 Hz, CHarCSO3-), 7.39-7.18 (m, 5H, NC(CHar)3CCH2OSO2 and CHarCCH3), 6.92-6.85 (m, 1H, NCCHarCCH2OSO2), 6.52 (br s, 1H, NH), 4.99 (s, 2H, CH2OSO2), 2.43 (s, 3H, CH3C6H4), 1.49 (s, 9H, C(CH3)3). 13 C NMR (CDCl3): δ (ppm) ) 152.6 (N-COO), 144.8 (Car-SO3), 138.7 (Car-CH2-OSO2), 134.3 (Car-NH), 133.3 (CarCH3), 129.8 (CarCCH3), 129.3 (NCCHCarCHCCH2OSO2), 128.0 (CarCSO3) 122.8 (NCCHCCarHCCH2OSO2), 118.9 (NCCarHCCH2OSO2), 118.3 (NCCarHCCHCCH2OSO2), 80.7 (C(CH3)3), 71.6 (C6H4CH2OSO2), 28.3 (-C(CH3)3), 21.6 (CH3-C6H4-SO3). C19H23NO5S (377.46): Calcd (%): C, 60.45; H, 6.15; N, 3.71; S, 8.49. Found (%): C, 60.59; H, 6.05; N, 3.42; S, 8.86. Oxazoline Polymerization (General Procedure). Three grams of MOX and EOX, respectively, were mixed with 15 mL of chloroform in an argon atmosphere and cooled to 0 °C. The initiator (methyltrifluoromethanesulfonate, trifluoromethanesulfonic acid, 0b, amount calculated according to

Biomacromolecules, Vol. 6, No. 1, 2005 237

POZs with Telechelic Antimicrobial Functions Table 1. MS Data of Selected Telechelic Poly(oxazoline)sj polymera CH3-PMOX-OH 1 CH3-PEOX-Py 2 CH3-PEOX-Py 2 H-PMOX-DDA 8 CH3-PMOX-DDA 7a BAB-PMOX-DDA 12a AB-PMOX-DDA 13

DPNMR

DPMS

23 22 22 25 22 25 24

23g

MMUb 85.11 99.13 99.13 85.11 85.11 85.11 85.11

19g 20h 27h 20h 19h 25h

Mstartc 15.04 15.04 15.04 1.01 15.04 206.27 106.14

Mtermd 17.01 79.10 79.10 213.41 213.41 213.41 213.41

zex Mcharge 6.9 6.9 1.0i 1.0 1.0 1.0 1.0

ze 1 1 2 2 2 2 2

mtheo/z

mexp/z

1989.58 1984.56 1038.39 1256.14 965.28 1018.33 1223.65

1990.23g 1903.03g 1038.81h 1256.42h 965.51h 1018.23h 1223.21h

mexp,minorf 1920.06/1945.14 1765.62 1682.24/1725.21 1746.40 1652.21/1660.35/1669.73 1722.63/1745.87

a It was presumed in all cases that the main MS population belongs to the target molecule. b Mass of a monomer unit. c Mass of group at starting end. Mass of group at terminating end. e Number of positive charges per polymer. f Examples for mass peaks of side populations deriving from side reactions or fragmentation. g calculated frommost intensive peak in MALDI-TOF. h Calculated from the most intensive peak inESI-MS. i One positive charge deriving from quarternary end group, the second one from protonation. j Additional Analytical Data of the Derivatives Are Given in Tables 2 and 3.

d

Table 2. Analytical Data of Telechelic Synthesized Poly(oxazoline)s polymer

DPcalca

DPNMRb

Mn,NMRc

Mw,GPC

Mn,GPC

CH3-PMOX-OH 1 CH3-PEOX-Py 2 CH3-PMOX-Py 3 CH3-PMOX-DBA 4 CH3-PMOX-DHA 5 CH3-PMOX-DOA 6 CH3-PMOX-DDA 7a CH3-PMOX-DHDA 8

20 20 20 20 20 20 20 20

23 22 26 24 26 22 22 23

1990 2120 2750 2310 2510 2190 2250 2390

2130 2420 3120 3320 2720 3580 2720 5820

2030 2330 2910 3130 2530 3360 2570 5260

MICe

PDGPC

CMC [*10-4mol/L]

[mg/mL]

[*10-4mol/L]

1.05 1.12 1.07 1.06 1.07 1.07 1.05 1.11

n.d.d n.d. n.d. 1.5 1.6 0.8 0.6 0.3

>10 >10 >10 >10 >10 >10 0.4 0.3

1.9 1.1

a Calculated from [M ]/[I], reaction time was 2 d for DP20, 4 d for DP50, and 6 d for DP100. b Calculated from the signal of CH at the starting end (2.91 0 3 ppm) and the backbone signal at 3.58-3.20 ppm. c Calculated presuming that the polymers are of the proposed structure. d Not detectable, i.e., the polymer did not show surfactant-like behavior. e All experiments have been performed in triplicate. The standard deviation was between 15 and 21%.

Table 3. Analytical Data of Telechelic Synthesized Poly(oxazoline)s polymer

DPcalca

DPNMRb

Mn,NMRc

Mw,GPC

Mn,GPC

CH3-PMOX-DDA7b CH3-PMOX-DDA7c CH3-PEOX-DDA 9a CH3-PEOX-DDA 9b CH3-PEOX-DDA 9c CH3-PEG-DDA 10

50 100 20 50 100 125d

53 96 20 44 96 116

4890 8550 2360 4740 9890 5330

5540 9550 3420 5820 10940 5550

5130 8740 3200 5260 10340 5210

MICe

PDGPC

CMC [*10-4mol/L]

[mg/mL]

[*10-4mol/L]

1.08 1.09 1.07 1.11 1.06 1.07

1.8 2.5 1.0 1.9 2.4 1.0

1.0 2.0 0.2 0.5 1.0 4.7

2.0 2.3 0.8 1.0 1.0 9.4

a Calculated from [M ]/[I], reaction time was 2 d for DP20, 4 d for DP50, and 6 d for DP100. b Calculated from the signal of CH at the starting end (2.91 0 3 ppm) and the backbone signal at 3.58-3.20 ppm. c Calculated presuming that the polymers are of the proposed structure. d Calculated from the molecular e weight given by the provider. All experiments have been performed in triplicate. The standard deviation was between 15 and 21%.

[I] ) [M0]/DPcalc.) was added to this mixture and the latter was left at 0 °C for 1 h. The solution was then quickly heated to reflux and stirred for several days (reaction times see Tables 2 and 3) followed by the addition of an 10-fold to 20-fold excess of terminating agent and then kept refluxing for 24 h. After a 3-fold precipitation in diethyl ether, dialysis in water (benzoylated cellulose membrane by Aldrich with MWCO of 1200 g/mol), and drying in vacuo, the polymers could be obtained in yields between 79 and 95%. 1H and 13 C NMR spectra of the polymer backbone showed the same signals described in the literature.14 Additional signals deriving from the new initiator and the new terminating agents are listed in the following. Signals DeriVing from Initiation with 0b. 1H NMR (CDCl3): δ (ppm): 7.61 (d, 2H, 3JH,H ) 8.17 Hz, CHarCSO3-), 7.42-7.12 (m, 3H, NC(CHar)3CCH2-POX), 7.08 (d, 2H, 3 JH,H ) 8.16 Hz, CHarCCH3), 6.74-6.69 (m, 1H, NCCHarCCH2-POX), 4.50-4.41 (m, 2H, C6H4CH2-POX), 2.27 (s, 3H, p-CH3C6H4SO3-), 1.42 (s, 9H, (CH3)3C). 13 C NMR (CDCl3): δ (ppm): 151.8 (N-COO), 146.6 (CarSO3-), 139.6 (Car-CH2-POX), 136.9 (Car-NH), 129.5 (Car-

CH3), 128.6 (CarHCCH3), 125.5 (CarHCSO3-), 80.2 ((CH3)3C), 28.2 ((CH3)3C), 21.6 (p-CH3C6H4SO3-). The signals deriving from dimethylalkylamine-termination are similar for all used amines and given on the example of DDA-termination: 1 H NMR (CDCl3): δ (ppm): 3.64-3.55 (m, 2H, POXCH2CH2N+), 3.16-3.06 (m, 10H, CH2(CH3)2N+CH2), 1.681.61 (m, 2H, POX-N+CH2CH2), 1.30-1.01 (m, 18H, N+(CH2)2(CH2)9CH3), 0.79 (t, 3H, 3JH,H ) 6.02, N+(CH2)11CH3). 13 C NMR (CDCl3): δ (ppm): 68.1 (-CH2(CH3)2N+CH2), 50.8 ((CH3)2N+), 13.9 (N+(CH2)11CH3). Deprotection of BOC-Protected Aminogroups. One gram of a polymer sample was dissolved in a mixture containing equivalent volumes of CHCl3 and trifluoroacetic acid (3+3 mL), a mixture commonly used for this reaction.18 After stirring the solution at room temperature for 1 h, the polymer was precipitated three times in diethyl ether and dried in vacuo affording the deprotected sample. The analytical data are given in results and discussion as well as in Tables 1 and 3.

238

Biomacromolecules, Vol. 6, No. 1, 2005

PEG-Derivatization. Commercially available poly(ethylene glycol) monomethyl ether mesylate (1.0 g; 0.2 mmol) was dissolved in 5 mL of CHCl3. DDA (0.85 g, 1.10 mL; 4.0 mmol) was added to this solution and the reaction mixture was heated under reflux at 70 °C for 24 h. After a 3-fold precipitation in diethyl ether, dialysis in water and drying in vacuo, 1.01 g (95%) of the derivative could be obtained as a colorless solid. 1 H NMR (CDCl3): δ (ppm): 3.94-3.89 (m, 2H, O-CH2CH2N+), 3.71-3.45 (m, 4nH, CH2-O-CH2), 3.35-3.23 (m, 10H, CH2(CH3)2N+CH2), 1.68-1.64 (m, 2H, PEG-N+CH2CH2),1.30-1.1 (m, 18H, N+(CH2)2(CH2)9CH3), 0.81 (t, 3H, 3JH,H ) 6.88, N+(CH2)11CH3). 13 C NMR (CDCl3): δ (ppm): 70.7-69.2 (CH2-O-CH2) 65.1 (-CH2(CH3)2N+CH2), 58.8 (CH3-O), 51.5 ((CH3)2N+), 39.4 (CH3SO3-), 31.7 (N+(CH2)9CH2), 29.4-28.9 (N+(CH2)2(CH2)7), 26.1 (N+CH2CH2), 22.5 (N+(CH2)10CH2), 14.0 (N+(CH2)11CH3). Analytical Procedures. Antimicrobial Susceptibility Determination. The Gram-positive S. aureus cells (ATCC, strain 25923) were cultivated by adding 100 µL of a stock suspension of the bacterial cells in PBS (1011 cells per mL) to 50 mL of a standard growth medium from Merck (composition in g/L: Peptone 15.0; yeast extract 3.0; NaCl 6.0; D-glucose 1.0, pH 7.5) and incubating it under shaking at 37 °C for 6 h. The polymers were dissolved in 10 mL of the growth medium in a concentration range of 0.1-10 mg per mL and inoculated with 100 µL of a the S. aureus suspension in the growth medium (107 cells per mL). The same procedure was carried out without polymer as control. After 6 h of shaking at 37 °C, the absorbance at 600 nm was measured, determining the concentration of the microbial cells, which is directly proportional to the optical density.19 The minimal inhibitory concentration (MIC) is the concentration of the biocide, where E600 (control)/E600 (sample) > 100, i.e., when 99% of the bacteria are inhibited in growth. Titration of NH2 Groups.20 A mixture containing 10 mg of polymeric material and 2 mL of aqueous NaHCO3 solution (2 g/L) was incubated at 37 °C for 10 min. Then, 25 µL of 1 M aqueous solution of 2,4,6-trinitrobenzene sulfonic acid were quickly added, and the mixture was incubated at 37 °C under stirring for another 4 h. The solution was diluted with 3 mL of 25 wt.-% HCl and the UV-vis spectrum was measured in a dilution of 1:40 with water. The maximum of absorbance of the formed compound appears at 346 nm; absorbance coefficient  (346 nm) ) 14500 M-1*cm-1. Thus, with the aid of the law of Lambert-Beer: E ) cd, whereas c is the concentration of NH2 groups with one amino function per polymer chain, the DP was calculated. The thickness of the cuvette was d ) 1 cm. Critical Micelle Concentration (CMC).21 Sample solutions were prepared in a concentration range from 10-2 to 10-7 mol/L by dissolving the carefully dried polymer in a 0.2 µM solution of the fluorescence dye 6-(p-toluidino)-2-naphthalenesulfonic acid in bidistilled water followed by fluorescence analysis of these polymer solutions at an excitation of 306 nm and an emission of 415 nm. The abrupt increase in emission intensity indicates the formation of micelles.

Waschinski and Tiller

Results and Discussion Synthesis, Characterization, and Microbial Testing of PMOX with Terminal Quarternary Ammonium Functions. It was established that living cationic polymerization of 2-alkyl-1,3-oxazolines started with bifunctional initiators and terminated with various nucleophils leads to well-defined macromolecules functionalized at both terminal ends.14b,22 Applying this general concept, we chose to introduce various functions to the starting end and antimicrobial functions to the terminal of poly(oxazoline)s to explore if the functions are still bioactive and how the polymers nature as well as the starting group influence this activity. Quarternary ammonium functions were used as covalently attached biocidal groups. Such compounds act antimicrobial if they contain an alkyl chain of adequate length interacting with the bacterial membrane by binding to it. Then the membrane’s equilibrium is disturbed due to the positive charge of the bioactive function eventually leading to cell death. As shown in cases of, e.g., pyridine14b and N,N-dimethylaminopropylmethacrylamide,23 the termination of the polymerization with tertiary amines results in quarternary ammonium end groups. For initial experiments, 2-methyl-1,3-oxazoline (MOX) has been cationically polymerized using the common methyltrifluoromethanesulfonate as initiator, adjusting the monomer/ initiator ratio to 20 and terminating the polymer with various tertiary amines. All synthesized polymers showed degrees of polymerization (DP) in good agreement with the monomerinitiator-ratio [M0]/[I] and low polydispersities (PD) between 1.05 and 1.12 as displayed in Table 2. The molecular weights Mn of the GPC measurements could be confirmed by the 1H NMR end group signal. Although this is usually accepted as proof for complete termination, there are still doubts, since nonterminated ends are hardly seen in the NMR or the GPC. For testing the antimicrobial activity, it was crucial that the polymers were indeed completely terminated with quarternary ammonium groups. MALDI-TOF is a promising method to find structure and molecular weight of polymeric compounds with molecular weights of up to 10.000 g/mol. As demonstrated by Jordan et al., poly(oxazoline)s with different end-groups can be well characterized by this method.14c However, applying the same protocol for the polymers shown in Table 2 resulted not in the expected discrete molecular masses of the macromolecules. The MALDI-TOF spectrum of CH3-PEOX-Py 2 is shown as an example in Figure 2a. The measurements roughly confirmed the molecular weight and DP of the polymer as measured by GPC. The distances of the peaks show exactly the mass of the monomer unit (see Table 1). Unfortunately, closer inspection of the peaks revealed that they do not present the exact molecular weights of the target molecule. One possibility is that pyridine is cleaved by dealkylation, which is typical for N-alkylpyridinium compounds in MS spectra. However, the exact masses of the macromolecules could not be found and with that the structure could not be confirmed by MALDI-TOF. Alternatively, electrospray ionization mass spectroscopy (ESI-MS) a method commonly used for the analysis of proteins and peptides, was recently described as milder method to confirm the structure of macromonomers.24 Applying ESI-MS to CH3-

POZs with Telechelic Antimicrobial Functions

Figure 2. (a) MALDI TOF mass spectrum, (b) ESI-MS of CH3PEOX-Py 2, and (c) ESI-MS of CH3-PMOX-DDA 7a.

PEOX-Py 2 resulted in a spectrum shown in Figure 2b. The major signals are direct repeating units of once, twice, and thrice positively charged poly(oxazoline) molecules. All of them contain a terminating methyl group originating from the polymerization initiator and one pyridinium-group as result of the termination (see also Table 1). Thus, the structure of a POX derivative could be proven by ESI-MS for the first time. The minor signals in Figure 2b could, according to their molar mass, e.g., m/z 1765.6, originate from a PEOXmolecule with a hydrogen at the starting end and pyridinium at the terminal (Table 1). Such molecules can be created by initiating the polymerization with an acid, a possible impurity in the methyltriflate-starter. Other possibilities are sidereactions such as chain transfer25 or fragmentation of the

Biomacromolecules, Vol. 6, No. 1, 2005 239

molecules during ionization. However, no nonterminated polymers could be detected indicating that the termination with pyridine was complete. The termination of the polymerization of MOX with N,Ndimethylalkylamines also led to completely terminated polymers. On the example of N,N-dimethyldodecylamine (DDA) the kinetics of the termination was followed at 70 °C at 20-fold excess using the 1H NMR-signal of the methyl group of the dodecyl-residue at 0.79 ppm to calculate the conversion rate. According to these measurements more than 95% of the living polymer chain ends were terminated with the amine after only 4 h, and termination was completed after 6 h. As seen in Figure 2c the ESI-MS of CH3-PMOXDDA 7a showed that the product after termination consists exclusively of polymer PMOX molecules terminated with DDA. The minor peaks could be interpreted as H-PMOXDDA molecules. The lack of PMOX without DDA end-group indicates that the termination reaction was quantitative. With this, the NMR-measurements could be confirmed. Termination with butyl-, hexyl-, octyl-, and hexadecyl-N,N-dimethylamine, respectively, led to similar results. The antimicrobial potential of the polymers was tested by determining the minimal inhibitory concentration (MIC) of the polymer against the ubiquitous Gram-positive bacterium Staphylococcus aureus. MIC is the standard quantification for antimicrobial substances and can be defined as the minimal required concentration to inhibit the growth of 99% of the bacterial cells in a suspension. In our experiments, we defined samples with MICs higher than 10 mg/mL as antimicrobially ineffective. The results of the microbial susceptibility test are given in Table 2. CH3-PMOX-OH 1 was used as control and showed no antimicrobial activity proving that the poly(methyloxazoline) itself is not bioactive. Also CH3-PMOXPy 3 did not exhibited a bacteriostatic effect indicating that the mere presence of a quarternary ammonium group at the polymer terminal is not sufficient to attack bacteria. Also aliphatic quarternary ammonium functions with alkyl chain lengths of 4, 6, and 8 carbons afforded no antimicrobial activity when attached to CH3-PMOX (derivatives 4-6, Table 2). Derivatives with alkyl chains of 12 (7a) and 16 carbons (8) were able to inhibit the growth of S. aureus cells effectively at concentrations of 0.19 and 0.11 mM, respectively. This is qualitatively coherent with the antimicrobial activity of the low molecular weight alkyl-trimethylammonium compounds,26 indicating that the antimicrobial effect originates from the telechelic function. It has been established that antimicrobial quarternary ammonium compounds are binding to the membrane of Gram-positive bacteria and must therefore penetrate the bacterial cell wall. Since the here described POX derivatives only show bacteriostatic activity if equipped with ammonium compounds that are active as low molecular weight pendants, we hypothesis that the polymers follow the same mechanism, i.e., they penetrate the bacterial cell wall. The MIC-values of the CH3-PMOX-DDA 7a and CH3PMOX-DHDA 8 are showing in microbiological way that the tertiary amine truly terminated the polymer, because only the quarternary ammonium compound could be antimicro-

240

Biomacromolecules, Vol. 6, No. 1, 2005

bial. For example, the MIC value of the polymer-bond DDA function is about 10 times higher than the one of the respective quarternary low-molecular-weight compound, dodecyltrimethylammonium chloride (DTAC), but more than 50 times lower than that of the terminating agent DDA. Molecules that bind to a bacterial cell membrane should either show specific binding toward specific structures, such as proteins and lipids, or should be incorporated into the selforganized supramolecular structure of the lipid membrane. In the latter case, the antibacterial molecules must behave like surfactants, a behavior already described for lipid-like polymers including poly(oxazoline)s,27 which are characterized by the critical micelle concentration (CMC). Lower CMCs should then result into lower MICs, i.e., higher antimicrobial activity. In fact, the CMCs of the CH3PMOX-dimethylalkylamines (4-8) determined fluorometrically with 6-(p-toluidino)-2-naphthalenesulfonic acid depicted in Table 2 show this trend. However, the DDA-function attached to PMOX showed a 10 times lower antimicrobial activity then the comparable low molecular weight compound DTAC. This is not surprising since the antimicrobial function is attached to an about 10 times greater and inactive molecule. However, it is also possible that the loss in activity is due to either a lower binding affinity to the bacterial cell membrane or to limited diffusion through the bacterial cell wall (see Figure 1). A variation of length and nature of the attached polymer should give more insights in this problem. Influence of Polymer Chain Length and Nature on the Terminal Antimicrobial DDA Function. Having established that DDA and DHDA are bacteriostatic when coupled to CH3-PMOX, the effect of the polymer chain length and the nature of the polymer backbone were explored. To this end, longer CH3-PMOX-DDA derivatives with 53 (7b) and 96 (7c) repeating units were synthesized. To investigate the influence of the nature of the polymer backbone, CH3PEOX-DDA 9a-c and CH3-PEG-DDA 10 polymers were prepared. The syntheses of the PEOX derivatives were carried out in the same way as for PMOX and lead to similar, completely terminated polymers with narrow molecular weight distributions as shown in Table 3. The PEG derivative 10 was prepared by the reaction of the commercially available PEG monomethyl ether mesylate with DDA in 20fold excess. According to NMR measurements the conversion of the PEG terminal was quantitative. As seen in Table 3 and Figure 3, the antimicrobial activity with regard to the concentration of the terminal function does not depend on the polymer chain length in the range of 22 to 96 repeating units. This is an unexpected result, because the increasing polymer chain length was suspected to hinder the biofunctional group from reaching its target of the microbial cell membrane. However, this result supports that the polymer does not influence the diffusion of the antimicrobial function through the bacterial cell wall indicating that the activity of the polymer is only dependent on the binding affinity to the microbial cell membrane. The increasing CMC values for greater molecules further indicate that the binding the microbial cell membrane is rather specific and not driven by supramolecular structuring.

Waschinski and Tiller

Figure 3. MIC values of antimicrobially terminated polymers. All experiments have been performed in triplicate. The error bars indicate the standard deviation.

Investigating the antimicrobial potential of a different polymer backbone, PEOX (9a-c) resulted in a chain length independent antimicrobial activity, which is twice as great as the activity of the respective PMOX derivatives 7a-c (see Figure 3). The CH3-PEG-DDA 10 exhibits a low antimicrobial effect being 5 times lower than that of the respective PMOX derivative 7c and even 9 times lower than that of the respective PEOX compound 9c. This is surprising, since PEG is the only polymer successfully used to stabilize bioactive molecules upon covalently binding them to the polymers terminal.11 In general, PMOX and PEOX seem to be well suited polymers for carrying antimicrobial functions through the bacterial cell walls. The POX derived antimicrobials are even more bioactive than the respective PEG derivative. Influence of the Starting End on the Terminal Antimicrobial DDA Function. As shown above, the nature of the polymer backbone attached to a quarternary ammonium function significantly influences the bioactivity of this compound. Another possibly influencing function is the group at the starting end of the polymer, which was a methyl group in all previous experiments. To investigate the impact of this function, a hydrogen and an NH2 group, respectively, were introduced to the starting end of the PMOX. In the case of the hydrogen at the starting end, the polymerization was simply initiated with trifluoromethanesulfonic acid. The monomer/initiator ratio was adjusted to 20, and the resulting polymer after termination with the DDA showed a molecular weight of ca. 2500 and had a PD of 1.31. According to 1H NMR data and ESI-MS results, the polymer chains carry a hydrogen at the starting end and were quantitatively terminated with DDA (Tables 1 and 4). The NH2 group was introduced by a novel bifunctional starter via a BOC (tert-butoxycarbonyl) strategy (see Scheme 1). MOX was polymerized with the initiator 0b and terminated with DDA to give the polymers BAB-PMOXDDA 12a-c (see Scheme 2). Although the DPs of the polymer are somehow higher than expected from the monomer/initiator ratio, they still show excellent narrow polydispersities in a range of 1.03 and 1.04 (see Table 4); that is, the novel initiator 0b afforded a living cationic polymerization of oxazolines. According to 1H NMR and ESI-MS data, the major compound was the target molecule (Figure 4). Two of the minor peak generations seen in Figure 4b and calculated in Table 1 could be interpreted

Biomacromolecules, Vol. 6, No. 1, 2005 241

POZs with Telechelic Antimicrobial Functions Table 4. Analytical Data for Bitelechelic Poly(oxazoline)s

MIC

polymer

DPcalca

DPNMRb

Mn,NMRc

Mw,GPC

Mn,GPC

PDGPC

Mtitrad

CMC [*10-4mol/L]

[mg/mL]

[*10-4mol/L]

H-PMOX-DDA 11 BAB-PMOX-DDA 12ae BAB-PMOX-DDA 12b BAB-PMOX-DDA 12c AB-PMOX-DDA 13f BAB-PEOX-DDA 14 BAB-PMOX-Py 15a BAB-PMOX-Py 15b BAB-PMOX-Py 15c

20 20 50 100 20 50 20 50 100

25 25 65 132 24 57 22 61 128

2490 2720 6120 11820 2530 6240 2330 5650 11350

3430 2950 6340 10230 2420 5730 2980 6980 13180

2620 2860 6150 9840 2310 5240 2830 6740 12680

1.31 1.03 1.03 1.04 1.05 1.09 1.05 1.04 1.04

3130 5470 11100 3130 6540 2710 5500 11330

0.3 0.2 0.4 0.5 0.2 1.8 n.d.g n.d. n.d.

2.5 0.2 0.4 1.0 10.0 0.5 >10 >10 >10

10.0 0.8 0.7 0.8 39.5 0.8

a Calculated from [M ]/[I] ratio, reaction times similar to derivatives in Table 2. b Calculated in case of 11-14 from CH -DDA signal at 0.79 ppm and 0 3 in case of 15 from pyridinium signal at 7.41 ppm in relation to backbone signal at 3.58-3.20 ppm. c Calculated presuming that the polymers are of the presumped structure. d Determined by photometrical titration. e BAB ) BOC-amino benzyl. f AB ) amino benzyl. g Not detectable, i.e., the polymers do not behave surfactant-like. h All experiments have been performed in triplicate. The standard deviation was between 15 and 21%.

Scheme 1. Initiator with BOC-Protected NH2 Function and a Sulfonic Acid Ester Function

as fragments of the polymer and the third minor generation corresponds to a molar mass of PMOX carrying the BOCprotected amino compound at the starting and a hydroxyl group at the terminating end. This could be due to an incomplete termination, followed by hydrolysis during the isolation procedure or during the sample preparation for the MS measurements. However, according to 1H NMR data and ESI-MS, more than 90% of the PMOX carry a DDA function at the terminating end (see Figure 4a). The protective BOC group could be cleaved easily by the standard procedure for BOC deprotection18 that includes treatment with a mixture of chloroform/trifluoro acetic acid (1:1, v:v) at room temperature. The deprotection was quantitatively proven by the complete disappearance of the BOC signal at 1.4 ppm in the 1H NMR spectrum. Further, no BOC carrying polymers could be found in the ESI-MS of 13 anymore (see Table 1). All main as well as the side populations in this spectrum present deprotected molecules of the respective populations in the ESI-MS of BABPMOX-DDA 12a. The lack of additional populations indicate that the polymer backbone was not affected by the deprotection procedure. To determine if the BOC deprotection resulted in active NH2 functions, the latter groups were photometrically titrated with 2,4,6-trinitrobenzene sulfonic acid, a method well-known from protein chemistry but rarely used in polymer science.20 Even though old-fashioned, this method is still about 1000-times more sensitive than for instance NMR-based determination of functional end-groups. Calculating Mn of the polymers from the amount of photometrically titrated NH2 groups resulted in molecular weights similar to those obtained by NMR and GPC (see Table 4). These results indicate that all formerly BOC-protected NH2

Figure 4. (a) 1H NMR and (b) ESI-MS of BAB-PMOX-DDA 12a.

are fully intact after deprotection and that the functional groups at the initiating polymer end could be introduced quantitatively. The polymerization of EOX as monomer and 0b as initiator lead to similar results as shown in Table 4 (BABPEOX-DDA 14). To have a control with a nonbioactive endgroup, MOX was polymerized with pyridine as the terminating reagent resulting in the expected bitelechelic polymers (BAB-PMOX-Py 15a-c, Table 4). The biological activity of the POXs with antimicrobial DDA end group and a hydrogen, a BOC protected aromatic NH2 group, and a free aromatic NH2 function were investigated by determining their MIC values against S. aureus as described above. The antimicrobial activity of the polymers seen in Table 4 are strongly dependent on the nature of the starting function. H-PMOX-DDA 11 is about

242

Biomacromolecules, Vol. 6, No. 1, 2005

Waschinski and Tiller

Scheme 2. Synthesis of PMOX with Telechelic NH2-Group and Antimicrobial Function

5 times less antimicrobially active than the respective methyl started polymer 7a. On the other hand BOC-protected functions at the starting end (BAB-PMOX-DDA 12a-c) seemingly increased the activity of the DDA group by more than a factor of 2 compared to 7a-c. In case of PEOX as polymer backbone, the BOC-protected amino group did not significantly increase the bacteriostatic activity as seen upon comparison of the MIC values of the derivatives BABPEOX-DDA 14 and 9b. In contrast, PMOX-DDA with free aromatic terminal function (AB-PMOX-DDA 13) almost completely diminished the antimicrobial effect of the terminal DDA group. All pyridine terminated polymers did not exhibit bactericidal properties, revealing that the BAB function at the starting end is not bioactive by itself. Apparently, the nonactive starting group of POX influences the DDA function’s activity in a synergistic way. The latter result is surprising, because it seems quite unusual that a nonbioactive function at the starting end of a polymer with more than 100 repeating units has the potential to influence the bioactivity of the group at the other end so greatly. This might open a new way to control the activity of polymer attached antimicrobial functions. Comparing the CMC values of the investigated polymers did not reveal a relation between supramolecular structuring and antimicrobial activity (Tables 2 and 3). Apparently, the bioactivity of the DDA terminated polymers is due to more specific interactions between polymer and bacterial cell membrane. Conclusions Narrowly distributed poly(oxazoline)s with hydrogen, methyl, BOC-protected aromatic amine, and free aromatic NH2 groups at the starting end and quarternary alkylamines (dimethylalkyl (butyl to hexadecyl)ammonium triflates and tosylates) at the terminating end with DPs in a range of 20 to 132 were synthesized. The structures of the polymers were proven by NMR and ESI-MS. It could be demonstrated that the POX attached quarternary dimethyldodecyl and dimethylhexadecylammonium groups were antimicrobial toward the Gram-positive bacterium S. aureus independent of the length of the polymer chain. This

means the polymer can apparently penetrate microbial cell walls. The nature of the polymeric backbone influenced the bioactivity of the terminating antimicrobial functions. Comparing PEOX, PMOX, and PEG, the first seems to be the best suited polymer for retaining the antimicrobial function. The functions at the starting end had an even greater effect to the bioactivity of the polymers. The MIC values of these compounds varied in a range of almost 2 orders of magnitude with derivatives carrying BOC-NH-groups showing the highest and the polymers containing free NH2 functions exhibiting the lowest activity. This dramatic controllability of a biological function by a nonactive group up to 300 atoms away from it is new in polymer science and might open an interesting field in drug design via designed polymeric functionalization. Investigating and optimizing the synergistic effect of these starting groups is part of current investigations. Further the activity of other bioactive groups attached to the end of polymers will be explored. Particularly, the polymers carrying aromatic NH2 functions will be coupled to surfaces via this functional group to hopefully reveal the true mechanism of surface-grafted antimicrobial polymers as contact-active biocidal materials. Acknowledgment. The authors thank the Deutsche Forschungsgemeinschaft for financing this work in the EmmyNoether-Programm and the Fonds der Chemischen Industrie for financial support. We thank Vera Herdes for preparation of some polymers, Bettina Knapp for measuring the ESIMS, Dr. R. Hanselmann for the MALDI TOF spectra, and Dr. Pelz for providing the S. aureus cells. References and Notes (1) Tiller, J. C.; Liao, C. J.; Lewis, K.; Klibanov, A. M. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 5981. (2) Lee, S. B.; Koepsel, R. R.; Morley, S. W.; Matyjaszewski, K.; Sun, Y. J.; Russell, A. J. Biomacromolecules 2004, 5, 877. (3) (a) Vogl, O.; Tirrel, D. J. Macromol. Sci. Chem. 1979, A13, 415. (b) For review, see: Tashiro, T. Macromol. Mater. Eng. 2001, 286, 63. (4) Ikeda, T.; Yamaguchi, H.; Tazuke, S. J. Biocat. Compat. Polym. 1990, 5, 31. (5) Kim, C. H.; Choi, J. W.; Chun, H. G.; Choi, K. S. Polym. Bull. 1997, 38, 387. (6) Lin, J.; Tiller, J. C.; Lee, S. B.; Lewis, K.; Klibanov, A. M. Biotechnol. Lett. 2002, 24, 801.

POZs with Telechelic Antimicrobial Functions (7) Desai, N. P.; Hossainy, S. F.; Hubbell, J. A. Biomaterials 1992, 13, 417. (8) Harder, P.; Grunze, M.; Dahint, R.; Whitesides, G. M.; Laibinis, P. E. J. Phys. Chem. B 1998, 102, 426. (9) Stone, F. W.; Stratta, J. J. Encycl. Polym. Sci. Technol. 1967, 6, 103. (10) (a) Seeliger, W.; Aufderhaar, E.; Diepers, W.; Feinauer, R.; Nehring, R.; Thier, W.; Hellmann, E. Angew. Chem. 1966, 20, 913. (b) Saegusa, T.; Ikeda, H.; Fuji, H. Macromolecules 1972, 5, 359. (c) Saegusa, T.; Ikeda, H.; Fujii, H. Polym. J. 1973, 4, 87. (11) (a) Francis, G. E. Inn. Pharm. Technol. 1998, 1, 41. (b) Greenwald, R. B. J. Controlled Release 2001, 74, 159. (12) Kimura, D.-H.; Kim, J.; Sugiyama, Y. Imanishi. Langmuir 1999, 15, 4461. (13) Kobayashi, S. Prog. Polym. Sci. 1990, 15, 751. (14) (a) Kobayashi, S.; Masuda, E.; Shoda, S.-I. Macromolecules 1989, 22, 2878. (b) Einzmann, M.; Binder, W. H. J. Polym. Sci: Part A: Polym. Chem. 2001, 39, 2821. (c) Jordan, R.; Martin, K.; Ra¨der, H. J.; Unger, K. K. Macromolecules 2001, 34, 8858. (d) Gross, A.; Maier, G.; Nuyken, O. Macromol. Chem. Phys. 1996, 197, 2811. (15) (a) Kobayashi, S.; Iijima, S.; Igarashi, T.; Saegusa, T. Macromolecules 1987, 20, 1729. (b) Christova, D.; Velichkova, R.; Goethals, E. J.; Du Prez, F. E. Polymer 2002, 43, 4585. (c) Chapman, R. G.; Ostuni, E.; Takayama, S.; Holmlin, R. E.; Yan, L.; Whitesides, G. M. J. Am. Chem. Soc. 2000, 122, 8303.

Biomacromolecules, Vol. 6, No. 1, 2005 243 (16) Moroder, L.; Hallet, A.; Wu¨nsch, E.; Keller, O.; Wersin, G. Z. Physiol. Chem. 1976, 357, 1651. (17) Malanga, C.; Mannucci, S.; Lardicci, L. J. Chem. Res. (M) 2000, 6, 0701. (18) Lundt, B. F. Int. J. Peptide Protein Res. 1978, 12, 258. (19) Hogt, A. H.; Dankert, J.; Feijen, J. J. Biomed. Mater. Res. 1986, 20, 533. (20) Tiller, J. C.; Bonner, G.; Pan, L.-C.; Klibanov, A. M. Biotechnol. Bioeng. 2001, 73, 246. (21) Persigehl, P.; Jordan, R.; Nuyken, O. Macromolecules 2000, 33, 6977. (22) Shimano, Y.; Sato, K.; Kobayashi, S. J. Polym. Sci: Part A: Polym. Chem. 1995, 33, 2715. (23) Schulz, R. C.; Schwarzenbach, E. Macromol. Symp. 1988, 13, 495. (24) (a) Krishnan, R.; Srinivasan, K. S. V. Macromolecules 2004, 37, 3614. (b) Jiang X. L.; Schoenmakers P. J.; van Dongen J. L. J.; Lou X. W.; Lima V.; Brokken-Zijp J. Anal. Chem. 2003, 75, 5517. (25) Litt, M.; Levy, A.; Herz, J. J. Macromol. Sci. Chem. 1975, A9, 703. (26) Ceruti, M.; Balliano, G.; Viola, F.; Cattel, L.; Gerst, N.; Schubert, F. Eur. J. Med. Chem. 1987, 22, 199. (27) Naumann, C. A.; Brooks, C. F.; Fuller, G. G.; Lehmann, T.; Ruehe, J.; Knoll, W.; Nuyken, O.; Frank, C. W. Langmuir 2001, 17, 2801. (28) Green, T. W.; Wuts, P. G. ProtectiVe Groups in Organic Synthesis, 2nd ed.; John Wiley & Sons: New York 1991; p 315.

BM049553I