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Biocidal Activity of a Light-Absorbing Fluorescent Conjugated Polyelectrolyte† Liangde Lu, Frauke H. Rininsland, Shannon K. Wittenburg, Komandoor E. Achyuthan, Duncan W. McBranch, and David G. Whitten* QTL Biosystems, 2778 Agua Fria, Santa Fe, New Mexico 87507-5491 Received December 7, 2004. In Final Form: January 31, 2005 Herein we describe studies that indicate a cationic conjugated polyelectrolyte shows biocidal activity against Gram-negative bacteria (Escherichia coli, E. coli, BL21, with plasmids for Azurin and ampicillin resistance) and Gram-positive bacterial spores (Bacillus anthracis, Sterne, B. anthracis, Sterne). These studies were carried out with aqueous suspensions of the conjugated polyelectrolyte, with the polyelectrolyte in supported formats and with samples in which the conjugated polyelectrolyte was coated on the bacteria. The results are interesting in that the biocidal activity is light-induced and appears effective due to the ability of the conjugated polyelectrolyte to form a surface coating on both types of bacteria. The effects observed here should be general and suggest that a range of conjugated polyelectrolytes in different formulations may provide a useful new class of biocides for both dark and light-activated applications.
Introduction There has been much recent interest from several different sectors in interfacial coatings (solid-liquid and solid-vapor) that exhibit efficient biocidal activity against bacteria, bacterial spores, and other agents. Among the systems that have been proposed and/or developed are metal ion containing formulations,1-6 coated and uncoated semiconductor particles,3,7 and polymer blends or surfactants containing pendant reactive organic functionalities (quaternary ammonium groups, hydantoins, tetramisole derivatives, or alkyl pyridinium structures) that may or may not require additional reagents for activation of biocidal function.8-19 * To whom all correspondence should be addressed. † Part of the Bob Rowell Festschrift special issue. (1) Collins, T. J.; Banerjee, D.; Khetan, S. K.; Yano, T. Book of Abstracts; 226th National Meeting of the American Chemical Society, New York, 2003; IEC-158. (2) Ignatova, M.; Labaye, D.; Lenoir, S.; Strivay, D.; Jerome, R.; Jerome, C. Langmuir 2003, 19, 8971-8979. (3) Goebbert, C.; Schichtel, M.; Nonninger, R. Farbe + Lack 2002, 108, 20-25. (4) Stoimenov, P. K.; Klinger, R. L.; Marchin, G. L.; Klabunde, K. J. Langmuir 2002, 18, 6679-6686. (5) Brunt, K. D.; Thompson, S. M. Adv. Coat. Technol. 1998, 35/ 31-35/18. (6) Koper, O. B.; Klabunde, J. S.; Marchin, G. L.; Klabunde, K. J.; Stoimenov, P.; Bohra, L. Curr. Microbiol. 2002, 44, 49-55. (7) Brunt, K. D.; Wood, P. N. Surf. Coat. Int. 1997, 80, 473-475. (8) Worley, S. D.; Eknoian, M.; Bickert, J.; Williams, J. F. Book of Abstracts; 216th National Meeting of the American Chemical Society, Boston, 1998; POLY-410. (9) Hazziza-Laskar, J.; Helary, G.; Sauvet, G. J. Appl. Polym. Sci. 1995, 58, 77-84. (10) Hamouda, T.; Baker, J. R. J. Appl. Microbiol. 2000, 89, 397403. (11) Henning, J.; Muller, F.; Peggau, J. SOFW J. 2001, 127, 30-35. (12) Maillard, J.-Y. Soc. Appl. Microbiol. Symp. Ser. 2002, 31, 16S21S. (13) Cen, L.; Neoh, K. G.; Kang, E. T. Langmuir 2003, 19, 1029510303. (14) Mandeville, R.; Kournikakis, B.; Brousseau, P.; Richard, L. PCT Int. Appl., WO 2003037331, 2003. (15) Werle, P.; Trageser, M.; Stober, R. DE 904002404, 1991. (16) Sauvet, G.; Fortuniak, W.; Kazmierski, K.; Chojnowski, J. J. Polym. Sci., Part A: Polym. Chem. 2003, 41, 2939-2948. (17) Chen, C. Z.; Beck-Tan, N. C.; Dhurjati, P.; Dyk, T. K. V.; LaRossa, R. A.; Cooper, S. L. Biomacromolecules 2000, 1, 473-480. (18) Shirai, A.; Maeda, T.; Masayo, K.; Kawano, G.; Kourai, H. Biocontrol Sci. 2000, 5, 97-102. (19) Wang, H.-H.; Lin, M. S. J. Polym. Res. 1998, 5, 177-186.
Conjugated polyelectrolytes (CPs) exhibit limited water solubility and spontaneously coat close to monolayer coverage when exposed to solid surfaces having surface charge opposite to the CP.20-23 The properties of specific CPs may be tuned so that the coating process is irreversible such that the coatings are robust and stable in the presence and absence of interfacial water.23 In particular, assemblies containing CP have been shown to be the basis of practical biosensors since the anchored CPs may exhibit the important combination of properties of efficient light harvesting, excitonic delocalization, and excited-state superquenching that can be coupled with biodetection by the use of synthetic quencher conjugates.20,22-26 In the present report, we show that a cationic CP shows biocidal activity against (Gram-negative) bacteria (Escherichia coli, E. coli, BL21) and bacterial (Grampositive) spores (Bacillus anthracis, Sterne, B. anthracis, Sterne). These studies were carried out with aqueous suspensions of the polymer, with the polymer in supported formats, and with samples in which the CP was coated on the bacteria. The results are interesting in that CP’s biocidal activity is white light induced (little or no biocidal activity was observed under yellow light) and appears effective due to the ability of the CP to form a surface coating on both types of bacteria. The effects observed here should be general and suggest that a range of CPs in different formulations may provide a useful new class of biocides. Materials and Methods Fluorescent Conjugated Polyelectrolyte, Coated Polyelectrolyte, and Chemicals. The polymer used in these (20) Chen, L.; McBranch, D. W.; Wang, H.-L.; Helgeson, R.; Wudl, F.; Whitten, D. G. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 12287-12292. (21) Jones, R. M.; Bergstedt, T. S.; McBranch, D.; Whitten, D. J. Am. Chem. Soc. 2001, 123, 6726-6727. (22) Jones, R. M.; Lu, L.; Helgeson, R.; Bergstedt, T. S.; McBranch, D. W.; Whitten, D. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 1476914772. (23) Lu, L.; Jones, R. M.; McBranch, D.; Whitten, D. Langmuir 2002, 18, 7706-7713. (24) Jones, R. M.; Bergstedt, T. S.; Buscher, T. C.; McBranch, D.; Whitten, D. Langmuir 2001, 17, 2568-2571. (25) Kushon, S. A.; D.Ley, K.; Bradford, K.; Jones, R. M.; McBranch, D.; Whitten, D. Langmuir 2002, 18, 7245-7249. (26) Kushon, S. A.; Jordan, J. P.; Seifert, J. L.; Nielsen, H.; Nielsen, P. E.; Armitage, B. A. J. Am. Chem. Soc. 2001, 123, 10805-10813.
10.1021/la046987q CCC: $30.25 © 2005 American Chemical Society Published on Web 03/11/2005
Light-Induced Biocidal Activity
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Figure 1. Molecular structures of the biocidal reagents, the fluorescent-conjugated polyelectrolyte polyphenylene ethynylene, PPE (1, CP), Methylene Blue (MB), Rose Bengal lactone (RBL), cetylpyridinium chloride (CPC), and dodecyltrimethylammonium bromide (DTAB). investigations, 1 (structure of 1 and other compounds shown in Figure 1), has been used in biosensing experiments.25,26 It is water soluble, yet forms insoluble coatings on oppositely charged particles such as carboxyl-functionalized polystyrene microspheres. Matrix-assisted laser desorption/ionization (MALDITOF) investigations indicate that the polymer may have ∼144 polymer repeat units (PRUs). The PRU length was estimated from MALDI-TOF analyses of several different batches and lots of the fluorescent conjugated polyelectrolyte preparations, and thus the above figure for PRUs represents the average length among various syntheses. Even with end-capping, it is difficult to control the polymerization process to exact lengths within a synthesis and especially so across synthetic lots of the polymer. Rose Bengal lactone (RBL), Methylene Blue (MB), dodecyltrimethylammonium bromide (DTAB), and cetylpyridinium chloride (CPC) were purchased from Sigma-Aldrich (St. Louis, MO). Microspheres were coated with 1 as described previously.20-23 Bacterial Growth Experiments. Initial experiments involved incubating B. anthracis spores and E. coli bacteria with the polymer and comparing the survival rate with bacteria not exposed to 1. Both types of bacterial cells could be stained using MB (vide infra).29,30 Bacterial cell counts were made using a Marienfeld haemocytometer (with Neubauer rulings) at 400× total magnification using the 40× objective of a phase contrast microscope (VWR Scientific, West Chester, PA) under bottom illumination. E. coli was grown in Luria-Bertani (LB) medium in the presence of 50 µg/mL ampicillin. E. coli cells were grown at either 37 or 25 °C according to the conditions described previously.27 E. coli cells were incubated for 2 h with different concentrations of the fluorescent conjugated polyelectrolyte or other biocidal agents as a suspension under yellow light. Then, (27) Hartley, H. A.; Baeumner, A. J. Anal. Bioanal. Chem. 2003, 376, 319-327. (28) Dang, J. L.; Heroux, K.; Kerarney, J.; Arasteh, A.; Gostomski, M.; A.Emanuel, P. Appl. Environ. Microbiol. 2001, 67, 3665-3670. (29) Venkateswaran, K.; Murakoshi, A.; Satake, M. Appl. Environ. Microbiol. 1996, 62, 2236-2243. (30) Brauwer, E. D.; Jacobs, J.; Nieman, F.; Bruggeman, C.; Drent, M. J. Clin. Microbiol. 1999, 37, 427-429.
the bacterial cells were centrifuged (4000g for 20 min). The supernatant was removed, and the bacterial pellet was resuspended in 1.0 mL of LB+amp media and exposed to laboratory white light for an additional 2 h. Then, 0.2 mL of the bacterial suspension was added to the wells of a microplate and absorbance from light scattering was measured. E. coli growth and multiplication rates were monitored by measuring light scattering at λ560 nm using a Molecular Devices (Sunnyvale, CA) microplate absorption spectrometer. Control experiments were carried out simultaneously in the absence of the fluorescent conjugated polyelectrolyte. Additional details are described in the legends to the appropriate figures. B. anthracis spores were germinated at 37 °C on 5% sheepblood-agar (SBA) plates (BD Biosciences, Cockeysville, MD) as described previously.28 B. anthracis spores were incubated for 2 h with different concentrations of the fluorescent conjugated polyelectrolyte or other biocidal agents, as a suspension under yellow light. The spores were then centrifuged (13000g for 35 min). The supernatant was removed, and the spore pellet was resuspended in 1.0 mL of deionized water before being further diluted in deionized water in order to plate the spores to check for viability. A 0.1 mL portion of the diluted spore suspension was asceptically spread on 5% SBA plates. The plates were incubated at 37 °C overnight. The following day, colony forming units were counted on the plate to determine viability. Control experiments were carried out simultaneously in the absence of the fluorescent conjugated polyelectrolyte. Additional details are described in the legends to the appropriate figures and tables.
Results and Discussion Results of the initial experiments are summarized in Table 1. These experiments were carried out with initial polymer concentrations of 2 × 10-6 to 1 × 10-5 M. For both bacteria it was found that incubation with polymer resulted in ∼40% reduction of bacterial survival. Both bacteria were treated with microsphere-supported suspensions of 1; in these cases there was also found a ∼40% reduction of bacterial survival following incubation over
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Table 1. Biocidal Activity of Various Formulations toward B. anthracis Sporea
Table 2. Concentration Effects on the Biocidal Activity of PPE toward B. Anthracis Sporea
treatment
spore colonies
% killing
concn of PPE-NR3+
spore colonies
% killing
control PPE-NR3+ (1) control Bead 1266 Bead 1268 Bead 1255 Bead-NR3+ Bead-CO2DTAB
130 ( 10 93 ( 4 122 ( 11 71 ( 3 73 ( 9 70 ( 3 122 ( 8 84 ( 4
0 30 0 42 40 43 0 40
0 (control) 1.1 × 10-3 M 2.8 × 10-4 M 1.1 × 10-4 M 2.8 × 10-5 M 1.1 × 10-5 M 2.8 × 10-6 M 1.1 × 10-6 M 2.8 × 10-7 M
72 ( 8 75 ( 3 55 ( 2 59 ( 3 64 ( 4 48 ( 3 52 ( 2 45 ( 1 46 ( 4
0 0 23 18 11 33 28 38 37
a Polymer (1) concentration is 10-5 M. Control is spores alone in deionized water. The concentration of DTAB is 2 × 10-5 M, 1266 is a “control” polystyrene-Neutravidin microsphere (0.6 µm), 1268 is a polystyrene-Neutravidin microsphere (0.6 µm) with 1 at a level of 1.1 × 106 PRUs/microsphere, 1255 is a polystyreneNeutravidin microsphere (0.6 µm) with 1 at a level of 7.8 × 106 PRUs/microsphere, Bead-NR3+ is a 0.2 µm bead with quaternary ammonium groups while Bead-COO- is a carboxylate functionalized microsphere. The bead concentration in each case is 500 microspheres per spore.
1.5 h. Anionic (carboxyl functionalized) microspheres by themselves had no effects on B. anthracis (Sterne) spore survival. Similar experiments with ammonium derivatized microspheres resulted in a reduction of survival corresponding to the microsphere-supported 1. Experiments with the quaternary ammonium surfactant DTAB (2 × 10-5 M) showed a ∼40% reduction in bacterial survival following 1.5 h of incubation under fluorescent laboratory lighting conditions; for the surfactant it was found that reduction of bacterial survival increased with a decrease in DTAB concentration over the range 3 × 10-5 to 1 × 10-3 M.
a Spores were treated with the indicated concentrations of PPE and then exposed to white light illumination. The spore suspension was then plated on to sheep-blood-agar (SBA) plates the survival estimated by colony counting.28
Association of 1 with Spores and Bacteria. Studies by fluorescence and phase contrast microscopy indicated that 1 is taken up by both bacteria (Figure 2) and that the polymer coated on the either the spores or bacteria is strongly fluorescent. Since 1 absorbs broadly through the visible region, it seemed possible that samples of bacteria incubated in room light could be undergoing both dark and photoinitiated interactions with the polymer. Preliminary attempts to separate the two effects indicated that there was somewhat lower reduction of B. anthracis survival when bacterial spores and 1 were incubated under yellow light (not absorbed to an appreciable extent by 1). Very interestingly, it was found (Table 2) that incubation of B. anthracis spores with 1 (1 × 10-5 to 1 × 104 M) under fluorescent lighting for 2 h showed an inverse dependence of reduction of bacterial survival with polymer concentra-
Figure 2. Phase-contrast and fluorescence microscope images of PPE-treated E.coli (upper panel) and PPE-treated B. anthracis Sterne spores (lower panel). Exposure of E. coli or B. anthracis to PPE resulted in the CP coating and/or penetrating (phagocytosis?) the cell membrane or cell wall of the bacteria or spore. Such cells glowed yellow-green, typical of the emission spectra of PPE. Phase-contrast images were taken at 100 ms and fluorescent images at 5.86 s exposures. The scale bar (bottom left of each photomicrograph) equals 5 µm.
Light-Induced Biocidal Activity
Figure 3. Schematic representation of the inner filter effect of PPE. For additional details please refer to the text.
tion. Thus at moderate to high polymer concentrations, there is almost no reduction of bacterial survival. The near complete “protection” of the spores afforded by high polymer concentrations suggested that quite possibly the reduction of bacterial survival was due entirely to a photoinitiated process and that large excesses of polymer in solution beyond that taken up by the bacteria might be affording protection of the polymer-coated bacteria by an inner filter effect (Figure 3). To test this possibility we performed a series of experiments to determine the level of adsorptive coating of 1 on B. anthracis spores and to isolate the behavior of the 1-coated spores. By use of different concentrations of 1, a “subtractive” assay of polymer uptake by spores was obtained by measuring the optical density before and following exposure to the spores and removal of the spores by centrifugation. The average uptake of PRUs/spore was found to be (2-3) × 107. It is reported that the size of a single B. anthracis spore is approximately 0.95 µm × 3.5 µm.31,32 It is also known that the E. coli bacterium is nominally 0.5 µm × 2 µm in size.33,34 The areas of B. anthracis spore and of E. coli were calculated by the equation: area ) 2πr2 + 2πrh (where π is, nominally, 22/7; r is the radius; h is the height). The area of B. anthracis spore was calculated to be 11.9 µm2 and the area of E. coli was computed to be 3.5 µm2. These dimensions then equal to 11.9 × 108 and 3.5 × 108 Å2, respectively. The surface area of 1 is estimated to be ∼120 Å2 per PRU. Given these values, the experimentally determined PRUs/spore for B. anthracis was ∼2 × 107 or about 2-fold compared to monolayer coverage. Thus the spores take up about two times more than “monolayer coverage” of the polymer. The excess could be due to spore penetration by 1. In a parallel experiment, spores incubated with a solution of 1 were collected by centrifugation, resuspended in aqueous medium, and exposed to white light for various periods. We found that, more-or-less independent of exposure time, the level of bacterial survival (as measured by spore growth in sheep (31) Weis, C. P.; Intrepido, A. J.; Miller, A. K.; Cowin, P. G.; Durno, M. A.; Gebhardt, J. S.; Bull, R. J. Am. Med. Assoc. 2002, 288, 28532858. (32) Dull, P. M.; Wilson, K. E.; Kournikakis, B.; Whitney, E. A. S.; Boulet, C. A.; Ho, J. Y. W.; Ogston, J.; Spence, M. R.; Mckenzie, M. M.; Phelan, M. A.; Popovic, A.; Ashford, D. Emerging Infect. Dis. 2002, 8, 1044-1047. (33) Miao, J.; Hodgson, K. O.; Ishikawa, T.; Larabell, C. A.; LeGros, M. A.; Nishino, Y. Proc. Natl. Acad. Sci. U.S.A. 2003, 100. (34) Maki, N.; Gestwicki, J. E.; Lake, E. M.; Kiessling, L. L.; Adler, J. J. Bacteriol. 2000, 182.
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Figure 4. Biocidal activity of PPE toward E. coli: E. coli (8 × 105 cells) were grown in Luria-Bertani broth containing ampicillin (LB + amp) at 37 °C in the presence (closed circles) or absence (open circles) of 2 × 10-6 M PPE. Growth was monitored by measuring the absorbance at 560 nm over 16 h at half-hour intervals. The absorbance was corrected by incorporating various controls including the absorbance from E. coli growth media alone. The absorbance of E. coli grown in the presence of 2 × 10-6 M PPE was indistinguishable from the absorbance of the media alone over the entire growth kinetics.
blood agar growth medium) was reduced to
