JAC Advance Access originally published online on February 8, 2006
Journal of Antimicrobial Chemotherapy 2006 57(4):691-698; doi:10.1093/jac/dkl012
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Comparative study of the antimicrobial activity of bis(N
-caproyl-L-arginine)-1,3-propanediamine dihydrochloride and chlorhexidine dihydrochloride against Staphylococcus aureus and Escherichia coli
1 Institute for Chemistry and Environmental Research, CSIC, c/Jordi Girona 18-26, 08034 Barcelona, Spain; 2 Serveis Científico-Tècnics, Universitat de Barcelona, c/Josep Samitier 1-5, 08028 Barcelona, Spain; 3 Laboratori de Microbiologia, Facultat de Farmàcia, Universitat de Barcelona, Av. Joan XXIII s/n, 08028 Barcelona, Spain
* Corresponding author. Tel: +34-934024496; Fax: +34-934024498; E-mail: amanresa{at}ub.edu
Received 6 June 2005; returned 20 September 2005; revised 20 October 2005; accepted 5 January 2006
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Abstract |
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Objectives: The aim of this study is to gain insight into the mechanism of the antimicrobial action of a novel arginine-based surfactant, bis(N
-caproyl-L-arginine)-1,3-propanediamine dihydrochloride [C3(CA)2]. Methods: To this end, we compared its effects against Staphylococcus aureus and Escherichia coli with those caused by the commercial and widely known antiseptic, chlorhexidine dihydrochloride (CHX).
Results: Both disrupted the cell membrane of the target bacteria to cause potassium leakage and morphological damage. The effect of C3(CA)2 on E. coli was concentration dependent, causing loss of membrane potential and membrane integrity leading to cell death, whereas CHX did not have these effects on E. coli. The effect of C3(CA)2 on S. aureus was the formation of mesomes and cytoplasmic clear zones, but the loss of membrane potential and membrane integrity was slightly lower than that with CHX.
Conclusions: We propose that C3(CA)2 acts preferentially against Gram-negative bacteria through strong initial binding to the surface lipopolysaccharides and subsequently partitioning into the cell membrane to cause membrane damage, followed by cell death.
Keywords: antimicrobial activity , flow cytometry , viability reduction , transmission electron microscopy , potassium leakage , cationic surfactants
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Introduction |
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Arginine-based cationic surfactants are amphiphilic compounds that possess excellent self-assembling properties, a low toxicity profile, high biodegradability and a broad antimicrobial activity, which make them candidates of choice as preservatives and antiseptics in pharmaceutical, food and dermatological formulations.1–5 Among the arginine-based surfactants synthesized in our laboratory, bis(N
-caproyl-L-arginine)-1,3-propanediamine dihydrochloride [C3(CA)2] is a novel gemini (double-chain/double-polar head) compound which shows antimicrobial activity against a wide range of Gram-positive and Gram-negative bacteria.1,6 Bacterial cells offer three regions for biocide interaction: the cell wall, the cytoplasmic membrane and the cytoplasm. In general, microbicides lack target specificity, though in the case of biguanide compounds such as chlorhexidine dihydrochloride (CHX), and cationic surfactants, their main target is the bacterial membrane.7–10 Both biguanides and cationic surfactants have a similar mechanism of action.11 As shown in Figure 1, both compounds are dicationic but with different structures: C3(CA)2 has two arginine residues connected by three methylene groups, and two alkyl chains of ten carbon atoms that form the hydrophobic moieties. CHX, on the other hand, possess two biguanides as the polar heads connected by a spacer chain of six methylenes, and two chloro-phenyl groups as the hydrophobic core.
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These cationic compounds initiate their interaction with the cytoplasmic membrane by binding to the phospholipid surface through electrostatic forces. They are then absorbed in the hydrophobic core of the membrane perturbing the packing of the lipids, leading to dissolution of the proton motive force and leakage of essential molecules.8,10 The specificity of these compounds towards prokaryotes, rather than eukaryotes, is thought to be due to the higher amount of anionic phospholipids in prokaryotic membranes.12–14
The synthesis of new biocompatible (i.e. low toxicity profile and high biodegradability) powerful antimicrobial agents is of paramount importance to prevent infectious diseases.10,15 The design of new molecules can be aided by an understanding of their mechanism of action, and we have recently reported the interaction of arginine-based surfactants with biomembrane models (i.e. liposomes and phospholipid monolayers).6 The purpose of this work is to elucidate the effects of the gemini surfactant C3(CA)2 on cell viability and on cell envelope integrity in Staphylococcus aureus and Escherichia coli. The well known biguanide CHX was included in these studies for comparison.
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Materials and methods |
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Antibacterial products and chemicals
C3(CA)2 was synthesized in our laboratory as previously described.3,16 CHX was purchased from Sigma (St Louis, MO, USA). Molecular dyes Syto-13, propidium iodide (PI) and bis-1,3-dibutylbarbutiric acid (bis-oxonol) were supplied by Molecular Probes Europe BV (Leiden, The Netherlands). The solvents and reagents used were of analytical grade. Ultra pure water, produced by a Nanopure purification system coupled to a Milli-Q water purification system, resistivity = 18.2 M·
·cm, was used for the aqueous solutions.
Microorganisms
Strains were obtained from the American Type Culture Collection (ATCC; Manassas, VA, USA): S. aureus ATCC 9144 and E. coli ATCC 10536. They were sub-cultured weekly on trypticase soy agar (TSA; Pronadisa, Barcelona, Spain). Strains were maintained frozen in cryovials (AES Laboratoire, Combourg, France) at –80°C.
MICs
The MICs of C3(CA)2 and CHX were determined by using a broth micro-dilution assay.17 Briefly, serial dilutions of the antimicrobials were made in Mueller–Hinton broth (Oxoid, USA). A 96-well polypropylene microtitre plate (Costar, Corning Incorporated, Corning, NY, USA) was used. Each well was inoculated with 100 µL of the test organism in Mueller–Hinton broth to a final concentration of 
105 cfu/mL. The MIC was taken as the lowest antimicrobial concentration at which growth was inhibited after 24 h of incubation at 37°C.
Exposure of microorganisms to biocides
Suspensions of the microorganisms were obtained from an overnight culture of each strain on tryptone soy broth (TSB) (Oxoid, USA) at 30°C. Cultures were then centrifuged at 8000 g for 15 min, washed twice in sterile Ringer's solution (Scharlau, Barcelona, Spain) and resuspended in Ringer's solution to obtain a cell suspension of about 107–108 cfu/mL. Three millilitres of the respective cell suspensions was used to inoculate flasks containing 27 mL of Ringer's solution to obtain a cell density of about 106–107 cfu/mL.
Different aliquots (7.5–360 µL, depending on the final 1/2 or 3/2 MIC) of the stock solution of C3(CA)2 and CHX (see Table 1) were added to flasks with the respective bacterial suspensions.
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After a contact time of 30 min, the biocide effect was immediately neutralized by dilution with sterile Ringer's solution, then the bacterial suspension was centrifuged for 20 min at 8000 g and the pellet was resuspended with Ringer's solution to obtain the convenient cell suspension concentration (107–108 cfu/mL). The inoculated flasks were kept at room temperature.
For flow cytometry (FC) experiments, 15 mL samples were diluted, centrifuged at 8000 g for 30 min and the bacterial pellets were resuspended in 1 mL of Ringer's solution.
For transmission electronic microscopy (TEM) observations, the contact time was 30 min at biocide concentration 3/2 MIC. Then, samples of 5 mL were taken, diluted with Ringer's solution and centrifuged at 4500 g for 30 min. The bacterial pellets were then rinsed with 0.1 M phosphate buffer (pH 7.4).
In all cases, control experiments lacking microbicide were conducted in parallel.
Staining procedure
Staining protocols for FC experiments were as follows: 1 µL of a 500 µM stock solution of Syto-13 in dimethyl sulphoxide and 10 µL of a 1 mg/mL stock solution of PI in distilled water were added to 500 µL of the bacterial suspension in filtered Ringer's solution. The S. aureus and E. coli suspensions were incubated with the dyes for 3 and 20 min, respectively. Stains were performed at room temperature before the FC analysis. Experiments were conducted in triplicate.
In the case of the membrane potential dye, 2 µL of a 250 µM stock solution of bis-oxonol in ethanol was added to 500 µL of the bacterial suspension to a final concentration of 1 µM and incubated for 2 min. Cells killed by heat exposure (30 min at 70°C) were used as controls for bis-oxonol staining. Experiments were conducted in duplicate.
Flow cytometry (FC)
A Coulter Epics Elite flow cytometer (Coulter Corporation, Florida, USA) equipped with a 15 mW air-cooled 488 nm argon-ion laser (for Syto-13, PI and bis-oxonol excitation) was used. The green emission from Syto-13 and bis-oxonol was collected through a 525 nm band-pass filter. The red emission from PI was collected with a 675 nm band-pass filter. Although the maximum emission is at 620 nm, a 675 nm band-pass filter was used to minimize interference with the strong fluorescence of Syto-13.
Bacteria were counted using a Cytek Flow Module (Cytek Development, CA, USA) adapted to the flow cytometer. Forward, side-scatter and fluorescence signals were collected in logarithmic scale. A significant percentage of the bacterial population that can be detected by its Syto-13 fluorescence appears in the first channel of the scatter. Consequently, fluorescence is used to discriminate bacteria rather than scatter, thus obtaining a better resolution and decreasing the background. Data were analysed with Elitesoft version 4.1 (Coulter Corporation) and WinMDI version 2.8 software.
Particle size analysis
Cell suspensions were analysed with a Multisizer II (Coulter) using a 30 µm aperture. Cell suspensions were diluted (1/1000) in 0.9% NaCl previously passed through a 0.2 µm filter. Data were analysed by AccuComp software version 1.15 (Coulter).
Bacterial count
Viable counts (cfu/mL) were obtained on trypticase soy agar (TSA). After an appropriate dilution in Ringer's solution, the sample was inoculated on plates and incubated at 30°C for 24–48 h. Rapid separation of bacteria from the antimicrobial was achieved by centrifugation at 4500 g in a bench top centrifuge for 15 min and subsequent dilution in Ringer's solution prior to plating. Cell counting was performed in triplicate.
Transmission electron microscopy (TEM)
After treatment of cell suspensions with the biocides at 3/2 MIC for 30 min for each microorganism [3 and 12 mg/L of C3(CA)2 with S. aureus and E. coli, respectively, and 0.75 and 3 mg/L of CHX with S. aureus and E. coli, respectively] the bacterial pellets were rinsed with 0.1 M phosphate buffer (pH 7.4), washed three times and fixed with 2.5% buffered glutaraldehyde for 1 h at 4°C. The cells were then post-fixed in 1% buffered osmium tetroxide for 1 h, stained with 1% uranyl acetate, dehydrated in a graded series of ethanol, and embedded in LR (London Resin Co. Ltd, London, UK) white resin. Ultra-thin sections were prepared and stained with 1% uranyl acetate and sodium citrate. Microscopy was performed with a Philips EM 30 (Eindhoven, Holland) microscope with an acceleration of 60 kV.
Potassium leakage
Potassium leakage was measured as described previously.4 Cells were grown on TSA at 37°C for 12 h, then harvested in 10 mL of Ringer's solution, washed three times with Ringer's solution and finally centrifuged at 5000 g for 30 min at 15°C and resuspended in 25 mL of 1 mM glycil-glycine buffer solution pH 6.8 to obtain a cell density of 2 x 108 cfu/mL and 3 x 108 cfu/mL for S. aureus and E. coli, respectively. The microorganisms were treated with the antimicrobials at 3/2 MIC for 30 min. Then, 5 mL of cell suspension was removed, diluted and centrifuged at 4500 g for 15 min. Positive controls were lysed cell suspension obtained by thermal shock (70°C for 30 min). Experiments were conducted in triplicate.
The potassium concentration in the supernatant was measured using an atomic absorption Philips PU9200X spectrophotometer (Philips Cambridge, UK). Absorbance values were converted into potassium ion concentration (ppm) by reference to a curve using potassium ion solution of 0, 0.1, 0.2, 0.5 and 1 ppm.
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Results |
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As shown in Table 1, the MICs of C3(CA)2 and CHX for S. aureus were 2 and 0.5 mg/L, respectively, and those for E. coli were 8 and 2 mg/L, respectively. To study the effect of the antimicrobial agents on the bacterial population, two concentrations were selected on an MIC basis: the first was 50% greater than the corresponding MIC (3/2 MIC) and the second was 50% lower than the MIC (1/2 MIC). The concentrations of the surfactant, C3(CA)2, were always lower than the corresponding critical micellar concentration (i.e. 4 x 10–5 M).18
Flow cytometry (FC) and viability reduction
To assess the effect of C3(CA)2 and CHX on bacterial populations, dual staining of cells was performed with Syto-13 (a nucleic acid stain that penetrates all types of cellular membranes) and PI, a nucleic acid stain not taken up by intact cells.19 Hence, three types of stained cells could be observed after the treatment: cells stained with Syto-13 (intact cells), double stained cells (partially damaged cells) and cells stained with PI (severely damaged cells).
To validate the FC results, and to examine the putative relation between cell damage and cell viability, viable cell counts were determined after exposure to the microbicides. The results with S. aureus are shown in Figure 2(a–c) and Table 2. As seen in Figure 2(a), the control population was mainly stained with Syto-13 (99 ± 1%). After 30 min of contact with 3 mg/L (3/2 MIC) C3(CA)2, 95 ± 1% of the cells were stained with Syto-13, 2 ± 1% were double stained and 3 ± 1% were stained with PI, indicating slight damage caused by C3(CA)2. Varying the C3(CA)2 concentration did not alter the proportion of damaged cells (Table 2, entries 2 and 3). Likewise, cell counts on plates did not change substantially with biocide concentration; viable cells dropped from 79 ± 10% at 1 mg/L (1/2 MIC) to 69 ± 6% at 3 mg/L (3/2 MIC). When S. aureus was treated with 0.75 mg/L CHX (3/2 MIC) (Figure 2c), 11 ± 4% partially damaged (double stained), and 15 ± 1% severely damaged cells (PI stained cells) were seen. Decreasing the biocide concentration from 3/2 to 1/2 MIC did not affect the proportion of partially damaged cells, since double stained cells only decreased from 11 ± 4 to 5 ± 1% (Table 2, entries 4 and 5). These results agree with the cell counts, where varying biocide concentration caused only a slight effect on cell viability (Table 2, entries 4 and 5).
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A different pattern was observed in E. coli populations when they were treated with C3(CA)2 and CHX (Figure 2d–f and Table 3). The control population showed 91 ± 4% of intact cells (stained by Syto-13). After treatment with 12 mg/L C3(CA)2 (3/2 MIC), a dramatic decrease in intact cells was detected, only 5 ± 1% of the population retained Syto-13 whereas 77 ± 10% were severely damaged (stained by PI), and the remaining 18 ± 3% were partially damaged (double stained, Figure 2e). At 4 mg/L C3(CA)2 (1/2 MIC), 42% (PI and PI + Syto-13) of the population showed signs of damage and the reduction in viability was 77 ± 1% (Table 3, entry 2). Three sub-populations could clearly be observed (Figure 2f) when E. coli cultures were treated with CHX at 3/2 MIC: 63 ± 5% remained intact (Syto-13); 21 ± 2% showed partial damage (double stained); and 16 ± 3% of cells were severely damaged (stained by PI). As presented in Table 3 (entries 4 and 5), variation of CHX concentration did not appear to have any effect as observed by FC; the population stained by Syto-13 decreased from 67 ± 5 to 63 ± 5%. However, the viability reduction changed significantly with CHX concentration, from 44 ± 1 to 61 ± 1 (i.e. 17 ± 2% of viability reduction).
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Membrane potential
Membrane depolarization was measured by determining the relative fluorescence intensity of bis-oxonol labelling using FC. As shown in Figure 3, the negative controls (i.e. cell suspensions in the absence of microbicides) showed the minimum relative fluorescence intensity (Figure 3a and e). We considered the range M1 [0-350 fluorescence units (FU)] as undamaged cells showing no significant depolarization of the cytoplasmic membrane. As expected, the positive control (i.e. bacteria that underwent a thermal shock at 70°C for 30 min) showed the maximum relative fluorescence intensity (Figure 3d and h). We considered the range M2 (between 350 and 1024 FU) as damaged and dead cells. The effect of C3(CA)2 on S. aureus showed a bimodal profile (Figure 3b) with two distinct populations of cells: almost 50% of the population emitted fluorescence in the region of non-damaged cells; while the rest emitted in the region of the dead cells. A different pattern was observed on exposure of E. coli to C3(CA)2; 84% of cells were damaged or dead (increased fluorescence); and 16% remained intact. When cells were exposed to CHX, a bimodal pattern was observed (i.e. 49% intact and 51% damaged cells) with E. coli (Figure 3g), whereas, for the most part, S. aureus was undamaged (Figure 3c).
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Potassium ion leakage
Potassium ion (K+) leakage reflects an increase in the cytoplasmic membrane. Here, intracellular potassium leakage resulting from exposure to C3(CA)2 and CHX of S. aureus and E. coli at the corresponding 3/2 MIC was measured. The cell density was 2 x 108 and 3 x 108 cfu/mL for S. aureus and E. coli, respectively. The negative blank was the K+ leakage measured from the bacterial suspensions in the absence of the antimicrobials (363 ± 75 x 10–3 ppm K+ from S. aureus and 769 ± 42 x 10–3 ppm K+ from E. coli). The positive blank was the K+ released from the bacterial suspensions heated at 70°C for 30 min (509 ± 29 x 10–3 ppm K+ from S. aureus and 918 ± 33 x 10–3 ppm K+ from E. coli). Thus, the percentage of K+ leakage was the ratio of net to total amount of K+ released.
Experiments with the Gram-positive organism showed that C3(CA)2 caused a potassium loss of 401 ± 32 x 10–3 ppm K+ (26%) and CHX 375 ± 63 x 10–3 ppm K+ (8%). The results with the Gram-negative organism showed that both antimicrobials released higher amounts of potassium ion than the Gram-positive organism: the surfactant, C3(CA)2, released 951 ± 52 x 10–3 ppm K+ (>100%) and CHX released, 889 ± 42 x 10–3 ppm K+ (81%).
TEM
Mesosome-like structures (i.e. spiral-bodies of the cytoplasmic membrane forming within the cytoplasm) and some cytoplasmic clear zones could be seen in S. aureus treated with C3(CA)2 (Figure 4a). In the case of E. coli more severe perturbations could be seen; numerous vesicles protruded from around the cell, severe damage to the membranes, appearance of cytoplasmic white spots, and abnormal septation (Figure 4b). The morphological disturbance caused by CHX on S. aureus was mesosome formation, clear zones and membrane-like convoluted structures in the cytoplasm (Figure 4c). In E. coli, cytoplasmic damage was clearly seen with the appearance of cytoplasmic white spots, as well as clear zones, and perturbations of the cytoplasmic membrane (Figure 4d).
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Discussion |
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C3(CA)2, and the comparator molecule CHX, change the polarity of the cell membrane bilayer of the bacteria and cause leakage of K+ ions from the cytoplasm. Both phenomena are involved in the cell death mechanism. But, these two compounds have different bacterial selectivity suggesting a different mode of action; C3(CA)2 has a concentration dependent microbicidal effect on E. coli, but has little effect on S. aureus; whereas CHX has little effect on E. coli, and does cause some damage (though not loss of viability) to S. aureus.
Potassium leakage is a first indication of membrane damage in microorganisms.20–22 Ultrastructural changes are also observed early on in the progress of membrane damage, with mesosome-like structures often appearing before any other effects on cell viability are noted.23 Both C3(CA)2 and CHX induced the first stages of cytoplasmic membrane damage in E. coli and in S. aureus, as shown by loss of intracellular potassium ions from washed cell suspensions, and the presence of intracellular mesosomes. This may be due to both the physicochemical properties of the microbicides and the composition of the cell membrane of Gram-positive and Gram-negative bacteria.7 Both, C3(CA)2 and CHX have two negative charges but the former is an amphiphilic molecule with surfactant properties whereas the later has a weak surface activity.6,24,25
The fluorochromes PI and Syto-13 were chosen because of their potential ability to distinguish between intermediate states of dead and live cells; PI confers fluorescence on cells which have lost their membrane integrity, whereas Syto-13 confers fluorescence on intact cells. The fluorochrome bis-oxonol is an anionic lipophilic dye that does not accumulate to any great extent in cells with a negative transmembrane potential, and fluorescence increases as membrane potential decreases.26 These labels, in combination with ultrastructural studies, and cell viability assays demonstrated marked differences between the microbicides in their action on the bacterial cells: when E. coli was exposed to C3(CA)2, 84% showed loss of membrane potential, 27–77% of cells showed membrane damage and 77–95% loss of cell viability, depending on the concentration of C3(CA)2 used. It is remarkable that the bis-oxonol fluorescence measured from the C3(CA)2-treated E. coli was close to that of the positive control. This indicates a complete membrane depolarization that caused irreversible damage leading to loss of viability. Electron micrographs agree with these results. Intracytoplasmic black spots were observed, similar to those reported in E. coli heat-treated cells.27 The most striking ultrastructural effect observed in E. coli was the formation of vesicles blebbing out from the outer membrane in the presence of C3(CA)2, similar to those reported for benzalkonium chloride-treated Pseudomonas aeruginosa.28 CHX had little effect on E. coli membrane integrity, shown by PI and Syto-13 labelling, though some intracytoplasmic changes were observed by electron microscopy. A moderate effect on E. coli membrane potential (51%) and cell viability (61% reduction) were also observed.
S. aureus membranes seem to be little affected by exposure of the bacteria to C3(CA)2, with just 5–7% loss of integrity shown by PI staining; a low emission of bis-oxonol fluorescence, indicating a moderate effect on the membrane potential and a low impact on the viability of the cells. Exposure to CHX caused some loss of membrane integrity (the proportion of PI stained cells increased), and a moderate reduction in membrane potential, but no reduction in viability. These data were supported by ultrastructural studies. These data are consistent with the idea that C3(CA)2 and CHX are probably working through similar overall mechanisms, but with different bacterial specificities due to their charge properties and the nature of the bacterial targets: Gram-negative cells offer a lipopolysaccharide (LPS) barrier that Gram-positive cells do not posses.7 The anionic character of the LPS layer14 could enhance the binding of the cationic surfactant C3(CA)2, which would not be so strong with CHX due to its weak surface activity and surfactant properties. This, together with the ability of C3(CA)2 to partition between the aqueous solution and the biological membrane, would make it an excellent molecule for targeting Gram-negative bacteria.
A lack of correlation between the membrane damaging action and cell viability seen for S. aureus suggests that membrane perturbation may be an important step in cell death, but is not necessarily a lethal event. Previous studies with S. aureus and E. coli have also indicated that membrane perturbation caused by antimicrobials and recovery of cells on agar are not always directly related.4,29 This may be due to technical factors such as the conditions for recovery of cells;8,30,31 or that membrane perturbation does not necessarily lead to dissolution of the proton motive force and cell death.8,30,31
It is notable that two different populations (50%) were registered using the bis-oxonol dye. This points to the presence of sub-populations with different degrees of damage, suggesting reversible damage or it could even represent viable and non-viable organisms.32
Conclusions
C3(CA)2 and CHX induce similar types of damage to S. aureus and E. coli, though they differ in the extent of membrane depolarization and reduction in cell viability. These differences suggest that both compounds damage bacteria in a different way, and that the surfactant properties of C3(CA)2 and its ability to partition into the biological membrane from aqueous solution might be important characteristics in its microbicidal action.
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No declarations were made by the authors of this paper.
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Acknowledgements |
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This work was supported by the MCYT project PPQ2000-1687-CO2-01 and CTQ-2004-7771-CO2-01/PPQ. J. A. C. acknowledges the CSIC I3P postgraduate scholarship programme for financial support.
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References |
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