Insights in the Antibacterial Action of Poly(methyloxazoline)swith a Biocidal End Group and Varying Satellite Groups

Christian J. Waschinski,† Sabine Barnert,‡ Alice Theobald,‡ Rolf Schubert,‡

Felix Kleinschmidt,§ Anke Hoffmann,§ Kay Saalwachter,| and Joerg C. Tiller*,⊥

Freiburg Material Research Center, University of Freiburg, Stefan-Meier-Str. 21,D-79104 Freiburg, Germany, Institute for Pharmaceutical Sciences, University of Freiburg,

Hermann-Herder-Str. 9, D-79104 Freiburg, Germany, Institute for Macromolecular Chemistry, University ofFreiburg, Stefan-Meier-Str. 31, D-79104 Freiburg, Germany, Department of Physics, University of Halle,Friedemann-Bach-Platz 6, D-06108 Halle, Germany, and Department of Bio- and Chemical Engineering,

TU Dortmund, Emil-Figge-Str. 66, D-44227 Dortmund, Germany

Received December 15, 2007; Revised Manuscript Received April 14, 2008

The antimicrobial activity of poly(2-methyl-1,3-oxazoline)s (PMOX) with the antimicrobial N,N-dimethyldode-cylammonium (DDA) end group is greatly dependent on the nature of the group at the distal end of the polymer,the satellite group. Three comparable PMOX with a DDA end group and different satellite groups (methyl, decyl,hexadecyl) were investigated with respect to the reasons for the huge differences in their biocidal behavior. Staticlight scattering (SLS) and pulsed field gradient diffusion NMR measurements revealed that the samples showcomparable aggregation conduct, thus, not being responsible for the varying biological activity. Experimentsusing different liposomal systems as models for bacterial cell membranes have been performed. It was found thatdifferential interactions between the respective polymers and the phospholipid membranes constitute the reasonfor the varying effectiveness observed in antimicrobial susceptibility determinations.

Introduction

Bacterial infections are one of the major scourges for humanmankind. Particularly, the increasing resistance against antibiot-ics demands the development of conceptually new agents againstbacteria, which turns out to be an everlasting campaign.1,2 Upto date numerous ways of killing bacteria or inhibiting theirproliferation are known.

Typical bacterial cell specific targets addressed by antibioticsare for example the cell wall (i.e., vancomycin), the cellmembrane (i.e., polymyxin), the DNA (i.e., tetracycline), andseveral proteins, including gyrase (i.e., ciprofloxacin) andtranspeptidase (i.e., penicillin).3–6 Various less specific antibac-terial agents are not completely understood. This is particularlytrue for antibacterial peptides and polymers. The first class ofmolecules is based on naturally occurring peptides, such asmagainin.7 Recently, a number of highly potent antimicrobialpeptides have been discovered.8 During the last few years,several active ingredients based on macromolecules have gainedincreasing relevance, whereas the effect is accessible in eitherpolymerizing biocidal monomers or in modifying polymersantibacterially.9,10 Because the antimicrobial activity of qua-ternary ammonium compounds has been known for decades,11–13

many of these systems gain their antimicrobial activity in beingcovalently equipped with positively charged functions, especiallyammonium groups.10,14–19 The often observed advantages ofpolymers are their nontoxicity, possible contact activity, lowerbacterial resistance, and enhanced activity compared to their

monomers.20–22 Furthermore, macromolecular systems can beequipped with different biologically active groups to studysynergistic effects and promising combinations.

However, understanding the mechanisms of these relativelynew classes of biocides is the basis to improve their perfor-mance, hopefully enabling the development of novel antibiotics.One major advantage of peptides and polymers over establishedantibiotics is their ability to fold in numerous different confor-mations, depending on their environment. One example is theconformation switching of magainin from a purely hydrophiliccoil in aqueous solution into an amphiphilic molecule on aphospholipid membrane, that is, the molecule reveals its activeform only in direct proximity to the target structure.23 This so-called amphipathic character is typical for many peptides andcan be predicted for most antimicrobial polymers.

In previous work, we have synthesized poly(methyloxazo-line)s (PMOX) with the antimicrobial N,N-dimethyldodecylam-monium (DDA) group at one polymer terminal and variousinherently nonbiocidal functions at the diametrically opposedend, the so-called satellite groups.24–26 It was found that thebactericidal activity of these macromolecules is independent onthe degree of polymerization (DP) in the range from 2000 g/molto 12000 g/mol, but is tremendously affected by the satellitegroups, whereby these groups have to be in the same polymerchain as the DDA function to achieve a change in antibacterialaction. Mixing polymers with different telechelic groups didnot show this effect. According to the obtained data, wesuggested a model that interprets these polymers as amphipathicmacromolecules that change their conformation on a phospho-lipid membrane.

In this study, we present investigations on selected PMOX-DDA polymers with different satellite groups regarding theiraggregation behavior in solution and their interactions withliposomes as model for the bacterial phospholipid membrane.

* To whom correspondence should be addressed. Tel.: +49 231 2552479. Fax: +49 231 755 2480. E-mail: tiller@bci.uni-dortmund.de.

† Freiburg Material Research Center, University of Freiburg.‡ Institute for Pharmaceutical Sciences, University of Freiburg.§ Institute for Macromolecular Chemistry, University of Freiburg.| University of Halle.⊥ TU Dortmund.

Biomacromolecules 2008, 9, 1764–17711764

10.1021/bm7013944 CCC: $40.75 2008 American Chemical SocietyPublished on Web 06/24/2008

Experimental Section

Materials. 2-Methyl-1,3-oxazoline (MOX, Fluka) was freshlydistilled from KOH prior to use. Chloroform (Roth) was freshly distilledprior use. All other compounds were of analytical grade or purer andused without further purification. All distillations and polymerizationswere carried out under an atmosphere of argon.

Methods. 1H and 13C NMR spectra were recorded using a BrukerARX 300 spectrometer operating at 300 MHz for 1H and 75.4 MHzfor 13C in CDCl3.

1H pulsed gradient diffusion NMR experiments were performed ona Bruker Advance 500 solid-state NMR spectrometer operating at500.12 MHz, equipped with a Bruker diff30 imaging/diffusion systemholding 5 mm sample tubes (∼5 µs 90° pulses), using the stimulated-echo technique.27 Gradient pulses with trapezoidal shape and lengthδ ) 1.5 ms were used. Experiments were performed at 24 °C at aconcentration of 30 mg/mL of the respective polymer sample in D2O.

Fourier transformed infrared (FT-IR) spectra were collected inattenuated total reflection (ATR) technique using a Bruker IFS 88 FT-IR spectrometer equipped with a Golden Gate unit.

Gel permeation chromatography (GPC) was carried out on a KnauerMikrogelset C11 equipped with a LS and RI detector and using DMFas eluent and poly(styrene) and poly(ethyl-oxazoline) standards (allstandards from Aldrich) at 25 °C.

Fluorescence spectra were recorded using a Perkin-Elmer LS 50Bluminescence spectrometer.

Static light scattering (SLS) was performed using a light scatteringapparatus by SOFICA with vertically polarized laser light at 632.8 nmwith a power output of 10 mW. The respective aqueous solutions withconcentrations ranging from 2 mg/mL to 10 mg/mL were filteredthrough 0.45 µm Minisart SRP 4 filters and then measured within anangular range from 50 to 130° (5° steps) at 20 ( 0.1 °C.

Transmission electron microscopy (TEM) was performed using anULTRACUT UCT microscope (120 keV) equipped with a cryochamberEM FCS (both by Leica).

Polymerization Procedure. Under an atmosphere of argon, 0.5-3mmol of initiator (decylbromide, hexadecylbromide, methyltrifluo-rmethanesulfonate) were reacted with 30-50 mmol of methyl-oxazolinein 15 mL of chloroform at 0 °C for 1 h. Afterward, the mixture wasallowed to react at 70 °C for two days. Polymerizations were terminatedby adding an at least 10-fold excess of N,N-dimethyldodecylamine(polymer 1-3) and a 1:1 mixture of Na2CO3/H2O (polymer 0),respectively, related to the amount of initiator and then kept at 70 °Cfor 24 h.

The macromolecules were precipitated thrice in diethyl ether,dialyzed in water (benzoylated cellulose membrane by Aldrich with amolecular weight cutoff of 1200 g/mol), and finally dried in vacuo.The polymers could be obtained in yields between 85-95%, FT-IRand 1H and 13C NMR measurements confirmed the structures.

Critical Micelle Concentration (CMC).28 Sample solutions wereprepared in a concentration range from 10-2 to 10-7 mol/L bydissolving the carefully dried polymer in a 0.2 µM solution of thefluorescence dye 6-(p-toluidino)-2-naphtalenesulfonic acid in bidistilledwater followed by fluorescence analysis of these polymer solutions atan excitation of 306 nm and an emission of 415 nm. The abrupt increasein emission intensity indicates the formation of micelles.

Antimicrobial Susceptibility Determination. The respective bacterialcells (Staphylococcus aureus ATCC 25923; Staphylococcus epidermis;Escherichia coli ATCC 25922; Pseudomonas aeruginosa ATCC 27853)were cultivated by adding 100 µL of a stock suspension of the bacterialcells in PBS (1011 cells per mL) to 50 mL of a standard growth mediumfrom Merck (composition in g/L: Peptone 15.0; yeast extract 3.0; NaCl6.0; D-glucose 1.0, pH 7.5; in the case of P. aeruginosa, the pH wasadjusted to 6.8) and incubating it under shaking at 37 °C for 16 h.Then, the respective polymer was dissolved in 10 mL of the growthmedium and a dilution series was prepared bisecting the concentrationof each solution. Each solution (5 mL) was inoculated with 50 µL ofa bacterial suspension (108 cells per mL) in the growth medium. After

6 h of shaking at 37 °C, the absorbance at 600 nm was measured. Thisway, the concentration of the microbial cells was determined, which isdirectly proportional to the optical density (A600 ) 1, which is equalto 109 cells per mL). The minimal inhibitory concentration (MIC) isthe concentration of the biocide, where A600 (control)/A600 (sample)< 100, that is, when 99% of the bacteria are inhibited in growth.24,25

To explore the temperature dependence on the antimicrobial activity,the MIC of the polymers was determined at room temperature as well,with the difference that the incubation time had be extended to up to10 days. The measured MICs were the same as that determined at 37°C.

Liposomes and Liposomal Systems. The Hepes buffer for prepara-tion and investigation of all of the liposomal systems consisted of asolution of N-(2-hydroxyethyl)piperazine-N′-(2-ethanesulfonic acid) (10mM) and NaCl (150 mM) in deionized water, whereby the pH wasadjusted with 1 N NaOH to give a value of 7.4. The size and sizedistribution (polydisperity index) of all liposomes were determined byphoton correlation spectroscopy (PCS) using a Zetamaster S (Malverninstruments).

Kinetics. The syntheses of liposomes for the membrane permeabi-lization kinetics are exemplified by means of preparation of eggphosphatidylcholin (EPC)/egg phosphatidylglycerol (EPG)/cholesterol(CHOL) systems. A prepared lipid film of EPC, EPG, and CHOL in amolar ratio of 4:3:3 was resuspended at a lipid concentration of 50mmol/L in Hepes buffer containing the fluorescence marker trinatrium-8-hydroxypyrene-1,3,6-trisulfonate (HPTS, 30.5 mM) and the quencherp-xylen-bis(N-pyridinium bromide) (DPX, 27.8 mM) for 1 h, followedby subjecting to three freeze-thaw cycles. The dispersion was extruded17 times through a 200 nm and 21 times through an 80 nmpoly(carbonate) membrane. Nonencapsulated HPTS and DPX were thenfiltered off in chromatographing the liposomal solution using a CL-4Bcolumn by Pharmazia and Hepes buffer as eluent. The liposomes weresome 190 nm in diameter with a polydispersity index of 0.12. Thefluorescence measurements were performed at λexc ) 403 nm and λem

) 510 nm at 25 °C, whereby solutions of the respective polymers inHepes buffer were added to agitated liposomal dilutions (lipidconcentration during measurement ) 25 µmol/L), and fluorescencespectra of the mixture were recorded over a period of 20 min.

RelatiVe Surface Polarity/MSP. (Exemplified for EPC-EPG-CHOLliposomes). The fluorescence marker 6-lauroyl-2-dimethylaminonaph-talene (Laurdan; 2.5 mM) dissolved in chloroform was added dropwiseto a solution of EPC/EPG/CHOL (molar ratio ) 4:3:3) in EtOH. Afterremoving the solvents the resulting film was dried under high vacuumfollowed by resuspending in Hepes buffer to give a mixture with a 20mM concentration of liposomes containing 0.2 mol % of Laurdan. Themixture was extruded 21 times through a 200 nm, 21 times through an80 nm poly(carbonate) membrane, and then further diluted with Hepesbuffer to give a dispersion with a lipid concentration of 0.46 mmol/L(diameter of liposomes ∼ 120 nm; polydispersity index ) 0.04).

The respective polymer solution was added to agitated liposomaldilutions (concentration of liposomes during measurement ) 230 µmol/L). The time-dependent intensity of fluorescence (λexc ) 355 nm) wascontinuously determined at the maximum (Imax, 434 nm) and theshoulder (IS, 487 nm) of the Laurdan spectrum at 25 °C over a periodof 60 min. The membrane state parameter (MSP) is defined as thequotient of IS/Imax and was automatically calculated for every readingpoint. MSP attributes a numerical value to a specific membrane stateat a defined temperature and pH value and does not change in the courseof time for the pure liposomes containing Laurdan giving MSPbase.Addition of a membrane active substance results in a change of thecontinuous fluorescence spectrum and thus different MSP data. Thesevalues are divided by MSPbase, resulting in MSPnorm, and MSPnorm ofthe pure liposomes always equals 1, representing a convenient guidancelevel.

For TEM investigations, the respective polymers were added to 10mM solutions of blank EPC-EPG-CHOL liposomes (molar ratio ) 4:3:3), and the resulting mixtures containing 2.88 mM of the macromol-

Antibacterial Action of Poly(methyloxazoline)s Biomacromolecules, Vol. 9, No. 7, 2008 1765

ecules were superimposed on a copper grid, freeze-dried in liquid ethane(Cryogen, 90 K), and investigated with TEM.

Results and Discussion

Three DDA-terminated poly(2-methyl-1,3-oxazoline)s (PMOX),with different satellite groups, were chosen for investigationregarding their antimicrobial mechanism. The structures of thepolymers are shown in Scheme 1. The basic analytical data ofthe polymers are given in the columns 3-5 of Table 1.

The MIC values of these polymers were determined with thebacterial stains S. aureus, S. epidermidis, E. coli, and P.aeruginosa (shown in columns 6-9 of Table 1). N,N-Trimeth-yldodecylammonium chloride [MIC (S. aureus) ) 20 µmol/L,MIC (S. epidermidis) ) 10 µmol/L, MIC (E.coli) ) 110 µmol/L] was used as a low molecular weight reference structurallysimilar to the DDA group.

In the cases of the first three bacterial strains, the polymerwith the methyl satellite group (1) showed a moderate antibacte-rial activity, while the decyl group (2) increased the activity ofthe polymer above that of the DDA group. The hexadecyl (3)function nearly completely diminished the activity of themolecule. In the case of P. aeruginosa, only polymer 2 wasinhibiting the growth of the cells. PMOX without DDA andsatellite group (0) did not show any antimicrobial activity.

Obviously, the antimicrobial properties of the polymers aredependent on the nature of the bacterial cell. However, thedifferences in the activity between the polymers against eachbacterium seem qualitatively consistent and are most likelycontrolled by the satellite group.

One explanation for the strikingly different biological activityof the PMOX-DDA derivatives could be that the satellite groupsinfluence the formation of large polymer aggregates that cannotenter the bacterial cell and are, therefore, not antimicrobial. Ifthis scenario was true, the MIC value would be falsified. In thecase that the remaining concentration of nonaggregated polymermolecules is below their true MIC, they would not beantimicrobial, as it could be the case for polymer 3.

The investigated DDA-terminated polymers are amphiphilicmolecules and should therefore be able to form micelle-likestructures in aqueous solution, as found for blockcopolymers29,30

and low molecular weight quarternary ammonium surfactantsas well.31 This was supported by the successful determinationof the critical micelle concentration (CMC), which indicatedthat the polymers with the DDA terminal truly form aggregates

at a certain concentration. The ability to form such supramo-lecular structures can have two consequences: On the one hand,the affinity of the polymers to hydrophobic surfaces such asphospholipid membranes should increase with decreasing CMCbecause of the increasing tendency to avoid water molecules.On the other hand, the formed micelles can hide the hydrophobicantimicrobial DDA group inside. Further, such large aggregatesmight not be able to penetrate the bacterial cell wall. Both effectswould result in a total loss of antimicrobial activity of theaggregates. Additionally, the maximal possible concentrationof single molecules in solution might not be high enough tokill bacterial cells. The CMC values of the investigatedmacromolecules decrease expectedly with increasing hydro-phobic character of the end groups in the order 1 > 2 > 3 (cf.Table 2). However, the antimicrobial activity increases comingfrom 1 to 2, but drops to very low activity in the case of polymer3. To investigate the aggregation behavior of the DDA-terminated PMOX, static light scattering and diffusion NMRexperiments were performed at first.

Correlation of Biological Activity with Polymer Aggrega-tion in Solution. Static light scattering was performed inaqueous solution at concentrations ranging from 2 mg/mL to10 mg/mL. The results are listed in Table 2.

In all LS measurements, comparatively large aggregates withmean radii of gyration (rg) between 280-317 nm were found.The radii rg of 1 and 2 of some 320 nm are virtually the same.The nonbiocidal polymer 3 shows somewhat smaller aggregateswith an rg of 280 nm. Given the great differences of the MICsof the polymers, a direct relation between the biological activityand the large aggregates seems irrelevant.

To analyze single molecules and smaller aggregates, furtherinvestigations were performed using diffusion NMR. Thismethod allows the determination of the hydrodynamic radiusof polymers with relatively low molecular weight and theiraggregates by measuring their diffusion coefficients. Further,coexisting species can be detected and quantified.27,32–35 Themeasurements were carried out with polymer solutions in D2Oat concentrations of 30 mg/mL (ranges between 4.5 and 6.8 ×102 µmol/L), which is above CMC for all polymers (see Table2). The decay of the stimulated echo intensity as a function ofgradient strength, g, gradient pulse duration, δ, and diffusiontime, ∆, is given by

II0) exp{-γH

2 δ2g2Dself(∆- δ ⁄ 3)} (1)

Where γH is the magnetogyric ratio and Dself is the self-diffusion coefficient, which is related to the Stokes hydrody-namic radius via the viscosity η:

Dself ) kT ⁄ (6πη0rS) (2)

Eq 1 constitutes a Gaussian-type decay as function of thevariable gradient strength, g, which is exemplified in its linearform in Figure 1. In the case of samples 1-3, a bicomponentdecay (three fitting parameters: two Dself and a relative weight)provided the best fit to the data. The resulting Stokes radii (rS)are displayed in Table 2.

The percentages given in brackets indicate the relative masscontent of the detected species. Because the large aggregatesare not visible in NMR measurements, their content wasdetermined by comparing the total area of the 1H NMR spectrumwith that of the spectrum of polymer 0, which does not formmicelles. It was found that the large aggregates measured inSLS experiments account for a small moiety (some 5%) in thesamples 1-3. In the remaining fraction of polymers 1-3 and

Scheme 1. Structures of the Investigated Polymers

1766 Biomacromolecules, Vol. 9, No. 7, 2008 Waschinski et al.

the complete sample 0, respectively, one species is existentwhose size of rS ) 2.6-2.9 nm is comparable to the size of therespective isolated macromolecules calculated according to theestablished theories of random/unpertubated polymer coils insolution. This species is, as already suggested, the only onefound in the case of sample 0, as was expected, because thispolymer does not possess structural elements stimulatingaggregation. In the case of the amphiphilic polymers 1 and 2,a second species was discovered with a particle size of rS )9.7 nm. Strikingly, these aggregates represent, with 5 and 6%,respectively, only minor fractions as well. Most of the polymerchains are still existent as isolated molecules. In the case ofpolymer 3, there are also two species present. The major fractionof 70% consists of larger aggregates (rS ) 18.0 nm) comparedto 1 and 2. However, the mixture still contains about 25% ofsingle molecules, with an rS of 2.9 nm. This represents aconcentration of about 110 µmol of single molecules per liter.Taking into account that the CMC of this molecule is 3 µmol/L, polymer 3 obviously does not act like a typical surfactant

because increasing the concentration in regions above CMCshould not increase the number of single molecules in solution.36

Thus, small aggregates should not be visible in the diffusionNMR measurements of polymer 3. The same is true forpolymers 1 and 2.

Summarizing the previous results leads to the conclusion thataggregation processes of the investigated polymers are not thedecisive factor for the observed differences in the biologicalactivity. The polymers 1 and 2 are almost identical in referenceto their respective particle size and distribution. In both cases,most of the polymers in solution exist as free molecules andshould be able to attack the bacterial cells. In spite of this thereis a great difference in their bactericidal impact. Even polymer3 contains a relatively large fraction of 25% of nonaggregatedmolecules in solution. Thus, its poor antimicrobial activitycannot be sufficiently explained by aggregation. The antimi-crobial activity of 3 could only be corrected by a factor of 4,which does not explain the huge difference in MIC betweenpolymers 2 and 3. The only remaining cause of the differentantimicrobial activity of the macromolecules might be theirinteractions with the bacterial cell membrane.

Correlation of Biological Activity with Different Interac-tions between Polymers and Phospholipid Membranes. Toexplore the direct interaction of the bioactive polymers withphospholipid membranes, liposomes were used as models forthe bacterial membrane. Because liposomes are artificial vesicleswith a shell of one or more lipid membranes (mostly bilayer)and an aqueous interior, the opportunity is provided to synthesizeentities with well-defined compositions. One common methodto measure the interactions between membrane-active com-pounds and its targets is measuring the release rate of anliposome entrapped marker in the presence of the compound.First, we chose a liposome model that resembles the membraneof E. coli.37 These liposomes for the membrane permeabilizationkinetics determinations have been prepared with the molarcomposition egg phosphatidylethanolamine (EPE)/egg phos-phatidylglycerol (EPG)/cardiolipin (CL) ) 80:15:5. Theirdiameter was some 160 nm.

The fluorescence marker, trinatrium-8-hydroxypyrene-1,3,6-trisulfonate (HPTS), and the fluorescence quencher, p-xylen-bis(N-pyridinium bromide) (DPX), were liposomally encapsu-lated, and the resulting liposomes were suspended in a buffersolution of N-(2-hydroxyethyl)piperazine-N′-(2-ethanesulfonicacid) and NaCl. As long as the liposomes are intact, nofluorescence is detectable. Addition of a membrane-activecompound results in damage of the membrane, the encapsulatedagents are released and diluted, leading to a strong increase influorescence.38 In our experiments, the liposomes containingbuffer solution were propounded, followed by the addition ofbuffer mixtures, including the respective polymers 0-3. Addingthe nonactive polymer 0 in a concentration as high as 2.88 mMto these negatively charged liposomes led to no release of dye,and thus, no increase in fluorescence was detectable (curve notshown). Thus, polymer 0possesses no membranous active

Table 1. Basic Analytical Data and MIC Values of the Investigated Polymers

MICa

polymer DPNMR Mn,NMR g/mol Mn,GPC g/mol PDGPC

S. aureusµmol/L

S. epidermisµmol/L

E. coliµmol/L

P. aeruginosaµmol/L

CH3-PMOX53-DDA 1 53 4890 5130 1.08 210 105 510 <10200C10H21-PMOX47-DDA 2 47 4430 5700 1.18 6 1.25 56 1120C16H33-PMOX71-DDA 3 71 6560 5120 1.15 760 1420 4700 >9400CH3-PMOX64-OH 0 64 5600 5640 1.03 >8900 >8900 >8900 >8900

a All experiments have been performed in triplicate, giving the MIC value at the same dilution factor in all cases.

Table 2. Size of the Polymer Aggregates Determined by the SLSand NMR Investigations

polymerCMCa

[µmol/L] rg [nm] cb [µmol/L] rS [nm]

1 180 317 (5%)c 615 2.9(90%) 9.7(5%)2 25 318 (5%) 675 2.8(89%) 9.7(6%)3 3 285 (5%) 456 2.9(25%) 18.0(70%)0 n.d.d n.d. 534 2.6(100%)

a CMC values were determined in triplicate and the standard deviationof these experiments were less than 5% in all cases. b Concentration forNMR measurements. c Relative mass content of the detected species inpercent. d n.d. ) not detectable.

Figure 1. Sample diffusion NMR results using a diffusion time ∆ )24.114 ms and a gradient pulse duration δ ) 1.4 ms for sample 0([), ∆ ) 24.712 ms and δ ) 1.7 ms for sample 1 (•), ∆ ) 24.113 msand δ ) 1.4 ms for sample 2 (gray square), ∆ ) 24.713 ms and δ )1.7 ms for sample 3 (2), and g was varied between 0 and 250 Gauss/cm.

Antibacterial Action of Poly(methyloxazoline)s Biomacromolecules, Vol. 9, No. 7, 2008 1767

functions and, therefore, features no interactions with lipo-somes. Figure 2a shows the measured fluorescence intensityof the liposome mixtures with time after addition of thepolymers 1-3, with resulting concentrations of 2.88 mM(polymer/lipid ratio ) 1:120), which is above the MICs ofthe polymers 1 and 2 against E. coli cells and below that ofpolymer 3.

Addition of the polymers 1 and 2, respectively, resulted inan outburst of the fluorescence dye indicating a rapid perforationof the membrane. The release stopped abruptly after about 20 sat some 80% (1) and 60% (2), respectively. Apparently, theholes formed by both polymers were healed quickly. Therespective experiment with polymer 3 resulted in a very slowrelease of the dye, which could be expected, because thecompound is not antimicrobial against E. coli at this concentra-tion. Unfortunately, for technical reasons, the concentration of3 could not be further increased in the experiments. Neverthe-less, compound 3 is able to perforate the phospholipid membraneto some extent, even below its MIC value of 4.7 mM (releaseof 1.5% of the dye in 100 s).

To examine the relation between polymer concentration andrelease rate, the experiment was repeated at different concentra-tions. As seen in Figure 2b, the release rate drops significantly

when adding 1 at a concentration of 510 mM, which is the MICvalue of the polymer against E. coli (see Table 1). However,the release at MIC is still 50% within 7 min, which could indeedbe the reason for the antimicrobial activity of the polymer.Lowering the concentration of polymer 1 to 0.288 and to 0.029mM, respectively, results in a nearly complete drop in membraneperforating activity. A similar scenario was found in the caseof polymer 2, shown in Figure 2c. In contrast to the findings of1, the perforation ability of 2 is still very high at its MIC valueagainst E. coli (0.056 mM, 75% release in 20 s). In this case,the release rate lowers significantly starting a concentration of0.014, which is below the MIC against E. coli. The membraneperforating activity is nearly gone at 0.003 mM.

Altogether, the membrane perforation experiments of E. coliresembling membranes reveal that the polymers show significantperforation activity at their respective MIC value against thebacterium and strongly drop this activity below MIC. Thus, theantimicrobial activity of the polymers seems to originate frompolymer membrane interactions. Further, the fluorescence dyeleaching experiment seems to be a good method to evaluatethe antimicrobial activity of membrane-active antimicrobialcompounds.

To generalize the concept of membrane/polymer interactionsand to get closer insights into the still unresolved reason forthe different antibacterial behavior of the polymers, neutral andnegatively charged liposomes of a generic composition wereused for the membrane permeabilization kinetics determinations.These liposomes were prepared with the molar composition eggphosphatidylcholin (EPC)/egg phosphatidylglycerol (EPG)/cholesterol (CHOL) ) 4:3:3 and egg phosphatidylcholin (EPC)/cholesterol (CHOL) ) 7:3, respectively.

The results of the release of the fluorescence dye in thepresence of 2.880 mM of each respective polymer are shownin Figure 3a,b. As found in the case of the E. coli membraneresembling liposomes, polymers 1 and 2 afford a high releaserate, while polymer 3 is not active at the investigated concentra-tion. Most surprisingly, the neutral liposomes (EPC-CHOL,Figure 3b) are effected in the same way as the negativelycharged ones (EPC-EPG-CHOL, Figure 3a). Thus, the mem-brane perforation activity of the polymers seems to not bedependent on the charge of the surface, but is most likely dueto hydrophobic interactions between polymer and membraneonly.

To get a closer insight into these hydrophobic interactions,the experiments were repeated at different polymer concentra-tions on the neutral liposome system. As seen in Figure 3c,d,the release rate of the fluorescent dye decreases with lowerconcentration of both polymers. The perforation tendency ofpolymer 1 for the neutral liposomes is similar to that of the E.coli membrane resembling liposomes shown in Figure 2b.Polymer 2 shows a higher perforation activity than polymer 1.In contrast to the latter, 2 is less active against the neutralliposomes compared to the negatively charged liposomes shownin Figure 2c. The general differences in perforation activity ofthe polymers are qualitatively similar for liposomes of verydifferent compositions.

Because the striking differences of the polymers in antibacte-rial activity are obviously caused by interactions with thephospholipid membrane, investigations concerning the relativesurface polarity were performed to see if the activity isinfluenced by the affinity of the polymers to the surface ofphospholipid membranes.

To this end, liposomes were prepared with the fluorescenceprobe 6-lauroyl-2-dimethylaminonaphtalene (Laurdan) incor-

Figure 2. Membrane permeabilization kinetics. EPE-EPG-CL lipo-somes (a) with polymers 1-3 at 2.88 mmol/L), (b) with polymer 1 atdifferent concentrations (given as numbers in mmol/L), and (c) withpolymer 2 at different concentrations (given as numbers in mmol/L).The experiments were reproduced at least once. The deviation wasless than 1% in slope and less than 5% in final release. The arrowsmark the time of polymer addition.

1768 Biomacromolecules, Vol. 9, No. 7, 2008 Waschinski et al.

porated into the membrane via noncovalent interactions.39–41

The fluorescence spectrum of liposomes containing Laurdanshows a maximum at 434 nm (Imax) and a shoulder at 487 nm(IS). The quotient IS/Imax of these two values defines themembrane state parameter (MSP), which attributes a numericalvalue that senses the polarity of the environment, that is, ahydrophobic group that attaches to the membranes surface willchange the MSP significantly to lower values.42

The MSP determination was performed with the EPC-EPG-CHOL liposomes. We chose to use polymers 1 and 2 at aconcentration of 210 µM, where the first compound has a lowperforation activity with respect to the used liposomes, and thesecond compound affords a very high leaching rate. Polymer 3was used at a concentration of 760 µM, where it does not showany perforation activity. Figure 4 depicts the MSPnorm versustime plots of the labeled liposome suspension after the additionof the polymers 1-3.

Surprisingly, the MSP values changed to above 1, whichindicates that the surface becomes more hydrophilic. Commonly,

hydrophobic compounds such as the DDA group of the usedpolymers afford MSP values lower than 1.43 This unusualbehavior can be explained by the fact that the hydrophobic endgroups of the polymer represent only a small volume fractionof the molecule. If attached to a surface, most of the surfacearea will be covered with the hydrophilic PMOX chains. Thiswill result in higher surface polarity with increasing numbersof surface attached molecules.

Applying 1 to the liposomes did change the MSP value to1.07, indicating that a very small number of molecules areattached to the liposome surface. Polymer 2, on the other hand,changes the MSP to 1.25 at the same concentration. Obviously,the difference in the antimicrobial behavior and membraneperforation activity of these two polymers can be explained bytheir different affinity to the surface of phospholipid membranes.Polymer 3 displays a different scenario. Although the moleculeclearly adsorbs to the surface of the liposomes (MSP 1.69),which is expected because of the more hydrophobic nature ofits end groups, it shows no perforation activity at the usedconcentration. Apparently, the molecule adsorbs to the surfacebut cannot penetrate the membrane. One explanation of thiseffect might be that the C16 satellite group is too long to beincorporated into the membrane, as is the case for respectivephospholipids.44

To visualize the impact of the bactericidal polymers 1 and2 on liposomes, cryo transmission electron microscopy (TEM)images of the liposomes were recorded after their contactwith the antimicrobial polymers. Due to the previous resultsof the experiments with the liposomes, the concentrationsof the polymers were chosen to be 2.1 mM in the case of 1and 0.06 mM in the case of 2, because the liposomes showeda high leaching rate at these concentrations. The respectivepolymers were added to solutions of the negatively chargedEPC-EPG-CHOL liposomes (lipid concentration 10 mmol/

Figure 3. Membrane permeabilization kinetics. (a) EPC-EPG-CHOL liposomes with polymers 1-3 at 2.880 mmol/L), (b) EPC-CHOL liposomes withpolymers 1-3 at 2.880 mmol/L, and EPC-CHOL liposomes with (c) polymer 1 and (d) polymer 2 at different concentrations (-, 2.880 mmol/L; - -,0.288 mmol/L; •, 0.029 mmol/L). The experiments were reproduced at least once. The deviation was less than 1% in slope and less than 5% in finalrelease. The arrows mark the time of polymer addition.

Figure 4. Course of MSPnorm in EPC-EPG-CHOL liposomes at aconcentration of 210 µmol/L for polymers 1 and 2 and at 760 µmol/Lfor polymer 3. The numbers given in the graphic are that of therespective polymer. The experiment was reproduced once. Thedeviation of the MSPs was less than 3%.

Antibacterial Action of Poly(methyloxazoline)s Biomacromolecules, Vol. 9, No. 7, 2008 1769

L), and the resulting mixtures were freeze-dried and inves-tigated with cryo TEM. Unmodified liposomes (not shown)appear as unilamellar spheres with 100-150 nm in diameter,and the entities are evenly spread. As expected, addition ofthe inactive reference 0 had no impact. Treating the liposomeswith polymer 1 at a concentration as high as 2100 µmol/Ldoes surprisingly show no effect as demonstrated in Figure5a. The entities are still uniform without any changes in theirsize, so attachment to and integration into the liposomalmembrane sufficient for lethal action on respective bacteriaand a steady release of dye in the kinetics experiments,respectively, cannot be visualized in this case. The TEMimages of the blank liposomes and the liposomes in thepresence of polymer 0 all look similar to the image shownin Figure 5a. Addition of 2 to liposomes with a resultingpolymer concentration of 60 µM, which is more than 30 timeslower than in the case of 1, has a considerable impact, asdisplayed in Figure 5b. The TEM images of the liposomereveal the formation of aggregates, so polymer 2 may act asa seed for aggregation due to its high membrane affinity andactivity. This could also be the reason for the change of theoverall size of the liposomes and their appearance. Besidesthe still present spheres with a diameter of 100-150 nm,some larger structures of up to 200 nm in diameter haveemerged. Moreover, many liposomes exhibit one or evenmore smaller spheres (d ) 50-100 nm) encapsulated in theirinterior, and a few species with multilamellar membranesare identifiable. So the interactions of 2 seem to compel acomplete reorganization process of the liposomes that alsoprovokes the strong outburst of fluorescence dye in the above-described kinetics determinations.

Conclusions

Three comparable PMOX polymers with a DDA end groupand different satellite groups (methyl (1), decyl (2), hexadecyl(3)) were investigated with respect to the reasons for the hugedifferences in their biocidal behavior. It was found that allpolymers form large aggregates of some 600 nm in diameter,smaller clusters of 19 nm in the cases of polymer 1 and 2,and 38 nm in the case of 3 in aqueous solution. However,even far above the respective CMC of the polymers, a largeamount of single molecules was found in the solution, beingsome 90% in the cases of 1 and 2 and 25% in the case of 3.Thus, aggregation of the molecules cannot likely be thereason for the different antimicrobial behavior. Studies on

the polymer induced release of a fluorescence dye entrappedin liposomes, which match the composition of E. colimembrane, revealed that the antimicrobial action of thepolymers mainly originates from the perforation of thebacterial phospholipid membrane. Further release studies withneutral and negatively charged liposomes showed that themembrane disruption mechanism of the polymers does notdepend on the charge of the membrane. The reason for thedifferent antimicrobial potential of the polymers 1 and 2 ismost likely their different affinity to phospholipid membranes,as demonstrated on surface polarity measurements on lipo-somes. The reason for the extremely low activity of polymer3 could not be explained with these measurements, becausethe compound had a very high membrane affinity, but didnot show any perforation activity at the investigated con-centration. We propose that this polymer adheres to themembrane surface but cannot be incorporated properly, thatis, the long hexadecyl satellite group prevents the penetrationof the antimicrobial DDA function. Thus, the satellite effectof the distal nonbioactive groups is truly controlling theantimicrobial activity of the polymers. The results providenew and interesting opportunities in the area of designingbiologically active ingredients based on polymer systems.

Acknowledgment. The authors thank the DeutscheForschungsgemeinschaft for financing this work in theEmmy-Noether-Programm, the Fonds der Chemischen In-dustrie for financial support, and Dr. Klaus Pelz for providingthe bacterial cell lines.

References and Notes

(1) Tiller, J. C.; Lee, S. B.; Lewis, K.; Klibanov, A. M. Biotechnol. Bioeng.2002, 79 (4), 465–471.

(2) Tiller, J. C.; Liao, C. J.; Lewis, K.; Klibanov, A. M. Proc. Natl. Acad.Sci. U.S.A. 2001, 98 (11), 5981–5.

(3) Chen, C. C. H.; Feingold, D. S. Biochemistry 1973, 12 (11), 2105–2111.

(4) Perkins, H. R.; Nieto, M. Pure Appl. Chem. 1973, 35 (4), 371–381.(5) Wolfson, J. S.; Hooper, D. C. Antimicrob. Agents Chemother. 1985,

28 (4), 581–6.(6) Chopra, I.; Roberts, M. Microbiol. Mol. Biol. ReV. 2001, 65 (2), 232–

260.(7) Zasloff, M. Proc. Natl. Acad. Sci. U.S.A. 1987, 84 (15), 5449–5453.(8) Sato, H.; Feix, J. B. Biochim. Biophys. Acta 2006, 1758 (9), 1245–

1256.(9) Vogl, O.; Tirrell, D. J. Macromol. Sci., Pure Appl. Chem. 1979, A13

(3), 415–439.(10) Tashiro, T. Macromol. Mater. Eng. 2001, 286 (2), 63–87.

Figure 5. TEM pictures of EPC-EPG-CHOL liposomes after (a) addition of 1 at 2.1 mM and (b) after addition of 2 at 0.06 mM.

1770 Biomacromolecules, Vol. 9, No. 7, 2008 Waschinski et al.

(11) Shelton, R. S.; Campen, M. G. V.; Tilford, C. H.; Lang, H. C.;Nisonger, L.; Bandelin, F. J.; Rubenkoenig, H. L. J. Am. Chem. Soc.1946, 68, 753–5.

(12) Shelton, R. S.; Campen, M. G. V.; Tilford, C. H.; Lang, H. C.;Nisonger, L.; Bandelin, F. J.; Rubenkoenig, H. L. J. Am. Chem. Soc.1946, 68, 757–9.

(13) Harris, G. H.; Shelton, R. S.; Van Campen, M. G.; Andrews, E. R.;Schumann, E. L. J. Am. Chem. Soc. 1951, 73, 3959–3963.

(14) Hazziza-Laskar, J.; Nurdin, N.; Helary, G.; Sauvet, G. J. Appl. Polym.Sci. 1993, 50 (4), 651–662.

(15) Nurdin, N.; Helary, G.; Sauvet, G. J. Appl. Polym. Sci. 1993, 50 (4),663–670.

(16) Hazziza-Laskar, J.; Helary, G.; Sauvet, G. J. Appl. Polym. Sci. 1995,58 (1), 77–84.

(17) Dizman, B.; Elasri, M. O.; Mathias, L. J. J. Polym. Sci., Part A: Polym.Chem. 2006, 44, 5965–5973.

(18) Fuchs, A. D.; Tiller, J. C. Angew. Chem., Int. Ed. 2006, 118, 6911–4.

(19) Kenawy, E.-R.; Abdel-Hay, F. I.; Abou El-Magd, A.; Mahmoud, Y.React. Funct. Polym. 2006, 66 (4), 419–429.

(20) Lin, J.; Tiller, J. C.; Lee, S. B.; Lewis, K.; Klibanov, A. M. Biotechnol.Lett. 2002, 24 (10), 801–5.

(21) Ikeda, T.; Yamaguchi, H.; Tazuke, S. Antimicrob. Agents Chemother.1984, 26 (2), 139–144.

(22) Ikeda, T.; Hirayama, H.; Yamaguchi, H.; Tazuke, S.; Watanabe, M.Antimicrob. Agents Chemother. 1986, 30 (1), 132–6.

(23) Oren, Z.; Shai, Y. Biopolymers 1998, 47 (6), 451–463.(24) Waschinski, C. J.; Herdes, V.; Schueler, F.; Tiller, J. C. Macromol.

Biosci. 2005, 5, 149–156.(25) Waschinski, C. J.; Tiller, J. C. Biomacromolecules 2005, 6 (1), 235–

243.

(26) Waschinski, C. J.; Zimmermann, J.; Salz, U.; Hutzler, R.; Sadowski,G.; Tiller, J. C. AdV. Mater. 2008, 20, 104–107.

(27) Tanner, J. E. J. Chem. Phys. 1970, 52, 2523–6.(28) Persigehl, P.; Jordan, R.; Nuyken, O. Macromolecules 2000, 33 (19),

6977–6981.(29) Antonietti, M.; Forster, S. AdV. Mater. 2003, 15, 1323–1333.(30) Bonne, T. B.; Ludtke, K.; Jordan, R.; Papadakis, C. M. Macromol.

Chem. Phys. 2007, 208 (13), 1402–1408.(31) Roy, S.; Dasgupta, A.; Das, P. K. Langmuir 2007, 23 (23), 11769–

11776.(32) Stejskal, E. O. J. Chem. Phys. 1965, 43 (10), 3597–3603.(33) Stejskal, E. O.; Tanner, J. E. J. Chem. Phys. 1965, 42 (1), 288–292.(34) Stilbs, P. Prog. Nucl. Magn. Reson. Spectrosc. 1987, 19 (1), 1–45.(35) Soderman, O.; Stilbs, P. Prog. Nucl. Magn. Reson. Spectrosc. 1994,

26 (5), 445–482.(36) Yeung, A. S.; Frank, C. W. Polymer 1990, 31 (11), 2089–2100.(37) Morein, S.; Andersson, A.-S.; Rilfors, L.; Lindblom, G. J. Biol. Chem.

1996, 271 (12), 6801–6809.(38) Smolarsky, M.; Teitelbaum, D.; Sela, M.; Gitler, C. J. Immunol.

Methods 1977, 15 (3), 255–265.(39) Chong, P. L.; Wong, P. T. Biochim. Biophys. Acta 1993, 1149 (2),

260–266.(40) Krasnowska, E. K.; Gratton, E.; Parasassi, T. Biophys. J. 1998, 74

(4), 1984–1993.(41) Parasassi, T.; Krasnowska, E. K.; Bagatolli, L.; Gratton, E. J. Fluoresc.

1998, 8 (4), 365–373.(42) Viard, M.; Gallay, J.; Vincent, M.; Meyer, O.; Robert, B.; Paternostre,

M. Biophys. J. 1997, 73, 2221–2234.(43) Schubert, R.; Schmidt, K. H. Biochemistry 1988, 27, 8787–8794.(44) Iden, D. L.; Allen, T. M. Biochim. Biophys. Acta 2001, 1513, 207–216.

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