Biomacromolecules 2004, 5, 1452-1456
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Toward Non-Toxic Antifouling: Synthesis of Hydroxy-, Cinnamic Acid-, Sulfate-, and Zosteric Acid-Labeled Poly[3-hydroxyalkanoates] Roland Hany,*,† Christine Bo¨hlen,† Thomas Geiger,† Manfred Schmid,‡ and Manfred Zinn‡ Swiss Federal Laboratories for Materials Testing and Research (EMPA), CH-8600 Du¨bendorf, Switzerland, and Swiss Federal Laboratories for Materials Testing and Research (EMPA), CH-9014 St. Gallen, Switzerland Received January 14, 2004; Revised Manuscript Received March 26, 2004
The side-chain double bonds of bacterial poly[3-hydroxyalkanoate-co-3-hydroxyalkenoate] (PHAE, 1) were transformed into thioether bonds (derivative 2) via the radical addition reaction of 11-mercapto-1-undecanol. The terminal hydroxy functionalities of derivative 2 were subsequently esterified with cinnamic acid (derivative 3), sulfatized with ClSO3H (derivative 4), or coupled with tert-butyldimethylsilyl-protected coumaric acid, to give, after deprotection with tetrabutylammonium fluoride (derivative 5) followed by sulfatization, p-(sulfooxy) cinnamic acid- (zosteric acid) labeled PHAE (derivative 6). The reactions proceeded with good yields and little side reactions, which was confirmed with 1H NMR and GPC experiments. These functionalized polyesters are currently investigated as environmentally friendly coatings to protect surfaces from biofouling. Introduction The undesired deposition of cells on a surface and the subsequent formation of a cell layer (biofilm) are called biofouling.1 Conventional strategies to remove a biofilm or protect a surface use toxic compounds that are gradually released over time and that are aimed at killing the bacteria. For example, the most effective solution to marine biofouling is the self-polishing copolymer organotin coating; unfortunately, it is also the most toxic one. Likewise, attempts to use conventional antibiotics to retard the formation of bacterial films on implanted medical devices suffer from the ubiquitous problem of emerging resistance from the most problematic bacterial strains. Several modifications of polymer surfaces were demonstrated to prevent bacterial adherence. Co-polyamines2 or polymers with ionic side chains such as poly(methyl methacrylate)-based terpolymers bearing sulfonate and carboxylate groups3 or poly(vinyl-N-hexylpyridinium bromide)4 are recent examples where the antimicrobial efficacy has been attributed only to the final polymer and not to leaching low molecular weight monomers.5 Other nonrelease strategies to construct nonbiofouling surfaces have consisted of changing the hydrophobicity/hydrophilicity balance using thermoresponsive polymer coatings6 or stable coatings with surfacereactive copolymers containing poly(ethylene glycol).7,8 Here we report on the synthesis of p-(sulfooxy) cinnamic acid- (zosteric acid) labeled poly[3-hydroxyalkanoate] (PHA), designed for the environmentally safe protection of surfaces * To whom correspondence should be addressed. Phone: +4118234084. Fax: +4118234038. E-mail:
[email protected]. † EMPA, Du ¨ bendorf. ‡ EMPA, St. Gallen.
from biofouling. Zosteric acid is a natural product made by the seagrass Zostera marina L. and has been found to prevent biofouling at nontoxic concentrations.9 The antifouling property of zosteric acid results from binding to attachment sites on cell surfaces, preventing cell adhesion.1 This nontoxic, anti-adhesion mode of action should side-step the risk of bacteria developing resistance. PHAs are high molecular weight, biocompatible, and biodegradable polyesters produced by many bacteria as intracellular carbon and energy sources.10 Recently, poly[3-hydroxybutyrate] (PHB) has been used as the matrix for pesticides,11 and the microbial induced surface erosion and surface renewal of PHB and poly[3-hydroxybutyrate-co-3-hydroxyvalerate] coatings have been proposed to be useful for foul-release at low ship speeds.12 We used the terminal side-chain double bonds of poly[3-hydroxyalkanoate-co-3-hydroxyalkenoate] (PHAE) for covalent attachment of zosteric acid. Polymer-analogous reactions on PHAEs have been studied in detail, and crosslinking,13-15 epoxidation,16 and conversion of double bonds to diol17 and carboxylic groups18,19 have been reported so far. As an additional method, we transformed the PHAE sidechain alkenes into hydroxy groups via the free-radical coupling reaction of a substituted thiol,20 followed by esterification. In addition, the cinnamic acid- and sulfatelabeled PHAEs were synthesized, since it was suggested that the sulfate group only was required for the antifouling effect of zosteric acid.9 These structural analogues will be included in tests on the antifouling properties of functionalized PHA surface coatings. Experimental Section Biosynthesis of Poly[3-hydroxyalkanoate-co-3-hydroxyalkenoate], PHAE. PHAE (1) was produced in a chemostat
10.1021/bm049962e CCC: $27.50 © 2004 American Chemical Society Published on Web 05/01/2004
Toward Non-Toxic Antifouling
culture of Pseudomonas putida Gpo1 (ATTC 29347) at a dilution rate of 0.1 h-1 under multiple (C,N) nutrient limited growth conditions.21 Cells were fed with octanoic acid (75 mol % or 90 mol %, respectively), 10-undecenoic acid (25 mol %, 10 mol %), and a mineral medium. The medium was designed in such a way that only nitrogen and carbon limited growth, whereas all other nutrients were in excess. Synthesis of PHAE (1) Derivatives 2-4 (1 with 25 mol % Double Bonds). P-(sulfooxy) cinnamic acid (zosteric acid) was purchased from R. C. Zimmerman,22 and all other reagents were used as purchased from Fluka or Aldrich. For the synthesis of 2, 10 g of the starting polyester 1 (17.6 mmol PHAE double bonds), 14.4 g of 11-mercapto-1-undecanol (70.5 mmol), and 0.58 g of AIBN (3.5 mmol, 2,2′azoisobutyronitrile) were dissolved in 100 mL toluene under argon. The solution was heated to 75 °C for 20 h, cooled to room temperature, and dropped into 2 L of ice-cold methanol. The raw product was dissolved twice in a minimal amount of CH2Cl2 and precipitated in a 10-fold excess of methanol for further purification, and then 2 was dried under high vacuum. 12.1 g of 2 was obtained (89% yield; all yields were calculated assuming complete conversion of precursor functional groups). For the synthesis of 3, a solution of 1.07 g of DCC (5.2 mmol, dicyclohexylcarbodiimide) and 0.14 g of 4-pyrrolidinopyridine (0.94 mmol) in 10 mL of dry CH2Cl2 was added under argon to a 0 °C cold solution of 3.6 g of 2 (4.7 mmol -OH groups) and 0.77 g of cinnamic acid (5.2 mmol) in 60 mL of dry CH2Cl2. The mixture was stirred for 2 h at 0 °C and 20 h at room temperature, filtered, precipitated twice in methanol, and dried. 3.7 g of 3 was obtained (88% yield). For the synthesis of 4, 1.1 mL of chlorosulfonic acid (ClSO3H, 16.5 mmol) was added dropwise to a solution of 5 g of 2 and 2.6 g of pyridine (32.9 mmol) in 25 mL of CH2Cl2 under argon at room temperature. After 2-3 h, 1 mL of H2O and MeOH was added, the solution basified (pH ) 8, 2 M NaOH), and then MeOH was removed under vacuum followed by freeze-drying. The raw product was dissolved in MeOH, filtered, dried, and further purified by repeated washing with acetone and H2O. The yield of 4 was 4.3 g (76%). Synthesis of Zosteric Acid-Labeled PHAE (1 with 10 mol % Double Bonds). The phenolic -OH group of 4-coumaric acid was protected with tert-butyldimethylchlorosilane (TBDMSCl) as described recently.23 The raw product was purified by recrystallization from hexane (-20 °C, 40% yield). 4-O-tert-Butyldimethylsilylcoumaric acid was esterified with 2 as described above (2 f 3, 78% yield). CH2Cl2 was continuously added during the reaction to decrease the viscosity of the reaction mixture and to keep the polymer in solution. The TBDMS groups were then cleaved in the presence of tetrabutylammonium fluoride (TBAF) to yield 5. A solution of 6.5 g of polymer in 150 mL of THF was cooled to -10 °C, and 3.7 mL of a solution of TBAF (3.7 mmol) in THF was added. After 3 min, the reaction was quenched by the addition of 500 mL of 2 M NH4+Cl- solution. The reaction mixture was extracted with AcOEt, and the organic layer washed with brine and dried over Na2SO4. 5 was precipitated in ice-cold methanol. 5.2 g (85% yield) of 5 was obtained. Zosteric acid-labeled PHAE
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was synthesized as described above (2 f 4); however, the washing with acetone was omitted. From 4.8 g of 5, 4.7 g of product 6 was obtained (93% yield). Polymer Characterization. Proton NMR experiments in solution were performed on a Bruker ASX-400 spectrometer. The measurements were carried out at 300 K with samples of typically 10-20 mg of polymer dissolved in 0.7 mL solvent. Chemical shifts are given in ppm relative to the remaining signals of chloroform at 7.26 ppm, methanol at 3.31 ppm, or acetone at 2.05 ppm as internal references. Molecular weights were determined by gel permeation chromatography (GPC, Viscotek, Houston, TX) equipped with a RI detector. The system was calibrated by using 10 polystyrene standards with known Mw (2 × 103 to 2.13 × 106 g mol-1) and low polydispersity (Mw/Mn e 1.09). 40 mg of every sample was dissolved in 10 mL of THF overnight. Aliquots of 100 µL of the polymer solution were chromatographed at 35 °C with pure THF as solvent phase through 2 GPC-columns (Mixed-Bed, Viscotek, Houston, TX) at a flow rate of 1 mL min-1. Results and Discussion PHAE (1) production was carried out using Pseudomonas putida GPo1 (ATCC 29347) in a chemostat culture as described earlier.21 By taking advantage of the adjustable effect of feed mixture on polymer composition, the steadystate conditions in a continuous culture are ideally suited to produce PHAs with tailored copolymer composition. The carbon feeds consisted of mixtures of 75 or 90 mol % octanoate and 25 or 10 mol % undecenoate, which indeed resulted in PHAE with 25 or 10 mol % monomer units containing unsaturated side chains. Because of the conversion of the carbon source by fatty acid degradation (β-oxidation) to monomer units which had two carbons less, 3-hydroxyhexanoate, 3-hydroxynonenoate, and 3-hydroxyheptenoate units were also incorporated into the resulting polymers.20 The unsaturated side-chain double bonds of PHAE were transformed into hydroxy functionalities via the free-radical addition of 11-mercapto-1-undecanol. The reaction was carried out in toluene with AIBN as the radical initiator. PHAE derivative 2 was isolated from the reaction mixture and purified by precipitation into cooled methanol. The hydroxy functions of 2 were then esterified with cinnamic acid (2 f 3) or sulfatized using ClSO3H (2 f 4). The synthesis of PHAE derivatives 2, 3, and 4 is summarized in Figure 1. Reaction conversions were monitored with solution 1H NMR spectroscopy. Figure 2 shows parts of the corresponding spectra with the crucial resonances assigned. In each spectrum, the intensity of the methine backbone protons was set to 100 (mol %). The alkene resonances of 1 with 25 mol % double bonds are observed at 4.95 and 5.77 ppm in Figure 2a. These resonances are completely absent in the spectrum of derivative 2 (Figure 2b), suggesting quantitative conversion of the alkene functions. In addition, the resonance of the methylene protons adjacent to the hydroxy groups appears at 3.63 ppm, and the intensity value of 50.8 for these two equivalent protons suggests the complete conversion of the alkene groups to thioethers in derivative 2. This observation
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Hany et al. Table 1. Solubility of PHAs 1-6 with 10 mol % (e.g., 1(10)) or 25 mol % Functional Side Chainsa solvent 1(10, 25) 2(10, 25) 3(10,25) 4(25) 5(10, 25) 6(10) 6(25) CHCl3 THF acetone MeOH DMSO
+ + + -
+ + + -
+ + + -
+ +
+ + + -
( ( ( +
+ + +
a + denotes soluble (at least 10 mg/mL), ( denotes cloudy solution, denotes insoluble. The chemical structures of 1-6 are shown in Figures 1 and 3.
Figure 1. Synthesis of PHAE derivatives 2-4.
Figure 3. Synthetic route to zosteric acid-labeled PHAE derivative 6.
Figure 2. Parts of 1H NMR spectra of (a) PHAE, 1, (b) derivative 2, (c) derivative 3, and (d), derivative 4. The numbers 1-4 above refer to the chemical structures shown in Figure 1, whereas the numbers under certain resonances are relative intensity values. Residual solvent signals are marked with * at 7.26 and 4.77 ppm.
agrees with the smooth free-radical addition of 11-mercaptoundecanoic acid to the side-chain double bonds of 1 we carried out recently.20 The NMR spectra as displayed in parts c and d of Figure 2 prove that also the esterification reaction with cinnamic acid (95% conversion of hydroxy groups) and the sulfatization reaction were nearly quantitative and free from significant side reactions. The chemical shift of -CH2OSO3-Na+ of 4 agreed with commercially available dodecyl sulfate at 3.99 ppm in the solvent methanol. While the starting polymer 1 and derivatives 2 and 3 were rather sticky, derivative 4 was a fibrous and colorless material. Also the solubility changed by introducing the ionic sulfate group (Table 1). 1, 2, and 3 were soluble in CHCl3, THF, or
acetone, but not soluble in MeOH or DMSO. The opposite is correct for 4. None of the materials were soluble in water. The direct coupling of derivative 2 with zosteric acid, either present as di-sodium salt or protonated at pH ) 2, was not successful. We therefore esterified 2 with TBDMSprotected coumaric acid, followed by deprotection with TBAF and sulfatization with ClSO3H (Figure 3). For derivative 2 containing 25 mol % hydroxy groups (2(25)), the polymer precipitated during the esterification reaction in CH2Cl2. The solvent was removed under vacuum and the residue purified by repeated washing with methanol. The deprotection with TBAF was then carried out in the solvent N-methyl-2-pyrrolidone. After quenching the reaction with NH4+Cl- solution, precipitated derivative 5(25) was dissolved in CH2Cl2, filtered, and purified by washing with methanol and water. The deprotection reaction conditions turned out to be crucial. When the reaction mixture was stirred for 1 h at room temperature as described,23 derivative 5(25) was obtained as a yellow oil, and the NMR spectrum pointed to significant side reactions and polymer backbone degradation. The reaction time was therefore gradually reduced to 3 min and the temperature to -10 °C. The NMR spectrum of 5(25) then revealed that 90% of the aliphatic hydroxy groups were esterified and that the silyl ether deprotection reaction was still quantitative. Zosteric acidlabeled PHAE, 6(25), was obtained from 5(25) as described above (Experimental Section, 70% sulfatization of phenolic -OH groups, 41% yield). Similar to derivative 4(25), 6(25) was neither soluble in CHCl3 nor THF. To follow the course of molecular weights and to study the delicate TBDMS deprotection step, we synthesized derivatives 2(10), 3(10), 5(10), and 6(10) starting from PHAE
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Figure 4. Parts of 1H NMR spectra of (a) PHAE derivative 2, (b) derivative 5, and (c) derivative 6. 2, 5, and 6 with 10 mol % functionalized side chains. The numbers above refer to the chemical structures shown in Figure 3, whereas the numbers under certain resonances are relative intensity values. Table 2. Molecular Weights [g/mol] of PHAE (1) and Derivatives 2, 3, 5, and 6 Shown in the Reaction Sequences of Figures 1 and 3a
a
PHA
Mn
Mw/Mn
1(10) 2(10) 3(10) 5(10) 6(10)
113 000 125 200 187 700 96 800 38 400
2.02 2.22 2.40 2.73 1.52
1, 2, 5, and 6 with 10 mol % functional side chains (e.g., 1(10)).
with 10 mol % side-chain double bonds. By decreasing the amount of ionic groups in 6(10), we expected to increase the solubility in THF, which is usually used for GPC measurements. Figure 4 shows parts of the NMR spectra for 2(10), 5(10), and 6(10) in the solvent acetone. The degree of conversion of the aliphatic hydroxy groups of derivative 2(10) into the ester functionalities in 5(10) was 78%, and 90% of the phenolic -OH groups were sulfatized (5(10) f 6(10)). The appearing signal at 3.9 ppm in the NMR spectrum of 6(10) (Figure 4c) suggests that parts of the remaining aliphatic -OH groups in 5(10) were sulfatized as well. The NMR spectrum of 5(10) was immediately restored by adding a drop of D2SO4 to an acetone solution of 6(10) in the NMR tube. This proved the successful sulfatization reaction and the rapid acid hydrolysis of the sulfate bond.9 As compared to 6(25), the hydrophilicity decreased with decreasing content of sulfate groups, and 6(10) was slightly soluble in THF (Table 1), but still had to be filtered for the GPC measurements. Molecular weights are shown in Table 2. The molecular weight remained essentially constant upon addition of 11-mercapto-1-undecanol to the PHAE double bonds and increased when the hydroxy groups of 2 were transformed into the ester (3) derivative. This course of molecular weights is consistent with the chemistry and confirms the smooth esterification reaction. In contrast to
Figure 5. ESEM micrographs of coatings after 6 h of exposure to activated sludge. (a) unprotected PHAE coating, (b) PHAE coating with dispersed zosteric acid, (c) PHAE coating with encapsulated zosteric acid, (d) coating of PHAE derivative 4 shown in Figure 1. Biological organisms are observable as small particles and thin threads covering the surfaces in Figure 5a-c, no bacterial settlement is observed on the surface coating shown in Figure 5d.
TBDMS-protected 5 containing 25 mol % side-chain functional groups, TBDMS-5(10) was clearly soluble in THF and Mn ) 141 800 g/mol (Mw/Mn ) 2.95) was measured. Deprotection with TBAF for 3 min at -10 °C yielded 5(10) with Mn ) 96 800 g/mol and Mw/Mn ) 2.73. The optimization of the deprotection conditions for efficient use in polymer-analogous reactions was important, and we observed for the reaction conditions 10 min at 0 °C, already significant polymer backbone degradation (Mn ) 17 000 g/mol). This is in keeping with reported problems found for the removal of phenolic silanes using TBAF.24,25 The molecular weight decreased when 5(10) was sulfatized with ClSO3H. However, as was mentioned above, 6(10) was only partly soluble in THF and had to be filtered for the GPC measurement. Therefore, it might be that the insoluble, higher-molecular fraction of 6(10) was removed by the filtration process and only the low-molecular fraction was actually detected. It should also be considered that the hydrodynamic volume of 6(10) in THF probably changed significantly due to the incorporation of ionic side chains. The covalent attachment of antifouling compounds to PHAE described in this work typifies the design of a passive antifouling coating. Such surface coatings could find applications in systems where liquids are moved fast and protection over a long period is desired. Active (leaching) PHAE surface coatings, where zosteric acid was mixed in, were prepared as well. The flux of zosteric acid from a leaching surface followed the Fickian law and was therefore only efficient over a short time. The interactive coating finally consisted of zosteric acid-loaded polystyrene microcapsules embedded in a PHAE coating. Since PHAE is biodegradable, the surface erodes through microbial action until new globules are uncovered and zosteric acid is secreted. The interactive surface represents an adaptive
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method that enables controlled release of agent in direct response to the presence of fouling microorganisms. To examine the mode of action, biofilm formation experiments were carried out. Therefore, PHAE coatings were exposed to activated sludge and the initial steps of biofilm formation (cell adhesion and attachment) were analyzed with environmental scanning microscopy. As an illustrative example, electron micrographs of various coatings after 6 h are shown in Figure 5. Clusters of microorganisms are observable as small particles covering the coating surfaces. The extent of cell adhesion on the surfaces followed the expected trend and the bacterial settlement was highest on the unprotected PHAE surface (Figure 5a), whereas no cell attachment was observed on the passive coating (Figure 5d). Detailed experimental results on the efficiency of these model surface coatings to control biofilm formation will be reported separately.26 Acknowledgment. We thank E. Pletscher for his help in the fermentation process and A. Hinz for the GPC measurement. References and Notes (1) Zinn, M.; Zimmermann, R. C.; White, D. C. In Biofilms: Recent AdVances in Their Study and Control; Evans, L. V., Ed.; Harwood Academic Publishers: Chichester, U.K., 2000; pp 361-380. (2) Tho¨lmann, D.; Kossmann, B.; Sosna, F. Europ. Coatings J. 2003, 1-2, 16. (3) Berlot, S.; Aissaoui, Z.; Pavon-Djavid, G.; Belleney, J.; Jozefowicz, M.; He´lary, G.; Migonney, V. Biomacromolecules 2002, 3, 63. (4) Tiller, J. C.; Lee, S. B.; Lewis, K.; Klibanov, A. M. Biotechnol. Bioeng. 2002, 79 (4), 465. (5) Lin, J.; Qiu, S.; Lewis, K.; Klibanov, A. M. Biotechnol. Bioeng. 2003, 83 (2), 168.
Hany et al. (6) Cunliffe, D.; Smart, C. A.; Tsibouklis, J.; Young, S.; Alexander, C.; Vulfson, E. N. Biotechnol. Lett. 2000, 22, 141. (7) Bearinger, J. P.; Terrettaz, S.; Michel, R.; Tirelli, N.; Vogel, H.; Textor, M.; Hubbell, J. A. Nat. Mater. 2003, 2, 259. (8) Jon, S.; Seong, J.; Khademhosseini, A.; Tran, T.-N. T.; Laibinis P. E.; Langer, R. Langmuir 2003, 19 (24), 9989. (9) Todd, J. S.; Zimmerman, R. C.; Crews, Ph.; Alberte, R. S. Phytochemistry 1993, 34, 401. (10) Steinbu¨chel, A.; Valentin, H. E. FEMS Microbiol. Lett. 1995, 128, 219. (11) Savenkova, L.; Gercberga, Z.; Muter, O.; Nikolaeva, V.; Dzene, A.; Tupureina, V. Process Biochem. 2002, 37, 719. (12) Yu, J. Biofouling 2003, 19 (Suppl.), 83. (13) Dufresne, A.; Reche, L.; Marchessault, R. H.; Lacroix, M. Int. J. Biol. Macromol. 2001, 29, 73. (14) de Koning, G. J. M.; van Bilsen, H. M. M.; Lemstra, P. J.; Hazenberg, W.; Witholt, B.; Preusting, H.; van der Galien, J. G.; Schirmer, A.; Jendrossek, D. Polymer 1994, 35, 2090. (15) Hazer, B.; Demirel, S. I.; Borcakli, M.; Eroglu, M. S.; Cakmak, M.; Erman, B. Polym. Bull. (Berlin) 2001, 46, 389. (16) Bear, M.-M.; Leboucher-Durand, M.-A.; Langlois, V.; Lenz, R. W.; Goodwin, S.; Guerin, P. React. Functi. Polym. 1997, 34, 65. (17) Lee, M. Y.; Park, W. H.; Lenz, R. W. Polymer 2000, 41, 1703. (18) Kurth, N.; Renard, E.; Brachet, F.; Robic, D.; Guerin, P.; Bourbouze, R. Polymer 2002, 43, 1095. (19) Stigers, D. J.; Tew, G. N. Biomacromolecules 2003, 4, 193. (20) Hany, R.; Bo¨hlen, Ch.; Geiger, T.; Hartmann, R.; Kawada, J.; Schmid, M.; Zinn, M.; Marchessault, R. H. Macromolecules 2004, 37, 385. (21) Durner, R.; Zinn, M.; Witholt, B.; Egli, T. Biotechnol. Bioeng. 2001, 72, 278. (22) Zimmerman, R. C., Moss Landing Marine Laboratories, 8272 Moss Landing Rd, Moss Landing, CA 95039. (23) Matsuno, M.; Nagatsu, A.; Ogihara, Y.; Mizukami, H. Chem. Pharm. Bull. 2001, 49, 1644. (24) Greene, T. W.; Wuts, P. G. M. ProtectiVe Groups in Organic Synthesis; John Wiley & Sons: New York, 1999; Chapter 3. (25) Wilson, N. S.; Keay, B. A. Tetrahedron Lett. 1997, 38, 187. (26) Geiger, T.; et al., Polym. Bull. (Berlin) manuscript in preparation. Zinn, M.; et al. Biofouling manuscript in preparation.
BM049962E
