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Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Immobilization of photosensitizers in polymers opens prospects for their continuous and reusable application. Methylene blue (MB) and Rose Bengal were immobilized in polystyrene by mixing solutions of the photosensitizers in chloroform with a polymer solution, followed by air evaporation of the solvent. This procedure yielded 15–140 μm polymer films with a porous surface structure. The method chosen for immobilization ensured 99% enclosure of the photosensitizer in the polymer. The antimicrobial activity of the immobilized photosensitizers was tested against Gram-positive and Gram-negative bacteria. It was found that both immobilized photosensitizers exhibited high antimicrobial properties, and caused by a 1.5–3 log10 reduction in the bacterial concentrations to their total eradication. The bactericidal effect of the immobilized photosensitizers depended on the cell concentration and on the illumination conditions. Scanning electron microscopy was used to prove that immobilized photosensitizers excited by white light caused irreversible damage to microbial cells. Photosensitizers immobilized on a solid phase can be applied for continuous disinfection of wastewater bacteria.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Photodynamic antimicrobial chemotreatment (PACT) is based on illumination caused excitation of a photosensitizer (PS) when PS molecules undergo a transition from a low energy ground state to a higher energy state. The excited PS can follow two pathways, named Type I and Type II reactions. In the Type I mechanism, PS molecules react with bio-organic molecules producing active free radicals and radical ions. The Type II reaction is accompanied by an energy transfer to molecular oxygen dissolved in an aqueous phase resulting in production of reactive oxygen species. The active products of both reactions cause direct and indirect damage to cellular membrane phospholipids and proteins, leading to membrane leakage and cytolysis inducing bacterial cell death [1-3]. The photosensitizers (PSs) are inactive and harmless in the dark [1-3]. Several Gram-positive and Gram-negative microorganisms were found to be sensitive to various PSs under illumination in vitro [2-17]. A large number of compounds have been shown to exhibit photodynamic properties, including porphyrin compounds, phenothiazine dyes, xanthene dyes, macrocyclic molecules such as phthalocyanines, and a group of natural PSs, including psoralens and perylenequinonoids [3].

The idea of the inclusion of a PS into a solid phase was raised in the early 1970s [18, 19]. Application of immobilized PSs exhibits several obvious advantages over their use in solution: immobilized PSs can be used in solvents in which free PSs are insoluble, they can be easily removed at the end of the treatment, can be reused, can be introduced into continuous processes [20], and are more resistant to bleaching by light and oxygen than free PSs [21]. There are a number of methods for solid phase immobilization of PSs—adsorption onto a solid support, formation of ionic bonds between the PS and ion-exchange resins, formation of a covalent bond between the PS and a polymer support and incorporation of the PS into a polymeric film [22]. Most of the studies in the field concentrate on a search for effective methods for covalent attachment of the PS molecules to solid supports. Covalent attachment of a large number of PSs, such as Rose Bengal (RB), eosin, fluorescein, chlorophyllin, hematoporphyrin and Zn(II) phthalocyanine tetrasulfonic acid to various supports, including silica gel, poly(styrene-co-vinylbenzyl chloride), poly[(N-isopropylacrylamide)-co-(vinylbenzyl chloride)], poly[(sodium p-styrenesulfonate)-co-(4-vinylbenzyl chloride)] and chitosan has been reported [20-25]. PSs that were covalently attached to polymers demonstrated high (up to 0.91) quantum yields of singlet oxygen formation [23]. The ability of the PS molecules, immobilized on a solid support, to maintain their photodynamic properties for a long period was shown in the works of Faust, Jiménez-Hernández and Bonnett [25-27].

In this work, we used a simple and cheap method of PS inclusion into a polymeric film by mixing chloroform solutions of PSs—Methylene blue (MB) and RB, with polystyrene solutions in the same solvent with subsequent air evaporation of the latter. This approach enabled the preparation of PS-polystyrene films or coatings with a predetermined thickness. The chosen PSs are soluble in water as well as in organic solvents. The photodynamic properties of the PSs in their free forms have been examined previously in several works [1, 3, 13, 14, 28, 29] and by us [15]. The aim of this work was to study bacterial photoinactivation by PSs that were incorporated into polymer films.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Bacterial growth

Cultures of Staphylococcus aureus (ATCC 25923) and Escherichia coli (ATCC 10798) were grown on brain-heart agar (BHA; Acumedia, USA) for 24 h, then were transferred into brain-heart broth (BH; Acumedia) and were grown at 37°C and at a 170 rpm speed of shaking up to 3 × 108 cells mL−1 concentration. Cells were harvested by centrifugation, washed twice with 0.05 m with phosphate-buffered saline (PBS), pH 6.5, diluted with PBS to a final concentration of 108 cells mL−1 and then were serially diluted in several (two to five) 10-fold dilutions.

Immobilization of photosensitizers in a polymer

A quantity of 25 g of polystyrene was dissolved in 100 mL of chloroform and mixed with a solution of 25–250 mg of a PS (MB or RB) in 20 mL of chloroform. The solutions were dispersed into disposable Petri plates, having a 55 mm diameter in portions of 4 mL, and the solvent was air-evaporated in a hood. After drying, the polymer films were washed with distilled water to get rid of free PSs. The washings were collected and a non included PS portion was determined by measuring absorption with the help of UV–Visible Spectrophotometer Cary 50 Bio (Varian, Australia) at the appropriate for each PS wavelength (Table 1). The degree of PS inclusion into the polymer was determined as the difference between the amount of PS taken for immobilization and the amount that leaked from the polymer during the washing procedure, relative to the amount taken. To test possible leakage of PS from the polymer surface, the Petri plates with PS-polystyrene coatings were filled with 4 mL of sterile PBS and incubated for 3 h at room temperature under the white lamp illumination and in the dark. PBS solutions were tested using visible absorption spectrophotometry and for antimicrobial activity.

Table 1. Photosensitizers used in this studyThumbnail image of

For experiments in scanning electron microscopy (SEM) imaging the above described chloroform solutions of the PSs and polystyrene and of polystyrene alone were dispensed on glass plates. After air evaporation of the solvent, the obtained polymer films were separated from the glass support and washed by distilled water.

Test of antibacterial activity of the polymer immobilized photosensitizers

Volumes of 4 mL portions of bacterial suspension at concentrations of 103 or 104 cells mL−1 in sterile PBS were dispensed into Petri plates having PS-polymer inner coating. The plates were illuminated for 0.5–3 h with a white luminescent lamp emitting in the range of 400–700 nm and having 1–3 mW cm−2 light intensity (1.8–5.4 J cm−2). The emission spectrum of the lamp was registered using a UV/Vis Spectrometer Ocean Optics USB 2000 (USA) and the light intensity was measured with a LX-102 Light-meter (Lutron, Taiwan). Quantities of 100 μL bacteria samples at various decimal dilutions were evenly spread over BHA plates with a Drigalsky spreader. The plates were incubated at 37°C overnight, and colony forming units (CFU) were counted. Control experiments were carried out with bacteria cultures on plates with or without polystyrene coating under illumination, and on plates with PS-polymer coating in the dark.

Microscopy characterization of photosensitizer-polystyrene surfaces and visualization of photosensitizer-polystyrene antibacterial effect

The thickness of the PS-polystyrene films cross-sections was measured using Axiolab Zeiss optical microscope. Surfaces of PS-polystyrene films were examined using scanning electron microscope SEM JSM-6510LV.

For SEM imaging of bacteria, the PS-polystyrene films underwent the following procedure: incubation in the suspension of bacteria (S. aureus and E. coli) in liquid Nutrient Broth growth medium (NB; Acumedia) for 24 h (initial concentration of bacteria was 106 CFU mL−1), easy rinse with 0.01 m PBS buffer, pH 7.5, initial fixation with 2% glutaraldehyde in 0.01 m PBS, pH 7.2, dehydration with varying concentrations of aqueous ethanol solutions (50–100%) and with varying concentrations of aqueous freon solutions (70–100%). Then, the films were coated with gold layer. Micrographs were obtained using a SEM microscope JEOL JSM 840.

Statistical methods

The results obtained from at least three independent experiments fulfilled with duplicates were statistically analyzed by ANOVA single factor or by ANOVA two-factor analyses. The difference between the results was considered significant if the P-value was less than 0.05.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Immobilization of photosensitizers in polystyrene

The applied method of immobilization afforded to achieve thin polymer films with inclusions of the PS. Polystyrene demonstrated a high ability to mix with the examined PSs in various ratios, and the applied method was supposed to provide a homogeneous distribution of the PS in the polystyrene. The PS-polystyrene films were washed by distilled water under monitoring of absorption spectra of the washings (Fig. 1). The first portions of washings contained high concentration of free MB (Fig. 1a) and tangible concentration of RB (Fig. 1b), but the last washings were actually devoid of free PS. It was shown that percent of PS inclusion into the polystyrene was more than 99% for each PS. The inclusion percent was calculated taking into consideration the amounts of the washed-out PS, measured at appropriate for each PS wavelength (Table 1). The final concentration of both PSs in polystyrene was more than 9.99 mg per g of the polymer.

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Figure 1. Visible absorption spectra of the first and the last portions of aqueous washings from plates with immobilized MB (a) and RB (b) and of samples after the 3 h incubation of PBS on the plates with PS-polystyrene coating under illumination and in the dark.

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The PS-polystyrene films exhibited high adhesion to the material of disposable Petri dishes, and were used for microbiological experiments without separation from the support. For optical and SEM imaging, PS-polystyrene films were obtained by spreading PS-polystyrene solutions on glass supports, which were separated after drying. The films were shown to have a porous structure with pores ranging from 1 to 3 μm (Fig. 2a). The cross-section view of the polymer films shows that the thickness of the films ranged from 15 to 140 μm (Fig. 2b–d).

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Figure 2. Microphotographs of the RB-polystyrene film (10 mg (RB) per g polystyrene) at ×1000 magnification (a) made by a SEM JSM-6510LV microscope; film cross-sections of microphotographs obtained with the help of a light Axiolab Zeiss microscope: (b) and (c) MB-polystyrene; (d) RB-polystyrene.

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To find out if PS leaked from the polymer, the Petri plates with PS-polystyrene coatings were filled with sterile PBS and part of the plates was incubated under illumination and another part—in the dark. After the incubation, visible spectra of the buffer solutions were registered. As can be seen from Fig. 1, in all the cases PS leaked from the polymer at a low degree and what is more, the leakage under illumination was higher than that in the dark, resulting in 0.48 μm MB and 0.26 μm RB by the end of the incubation. These solutions of leaked PSs were used as controls in the further PACT experiments.

The bactericidal activity of the polymer immobilized photosensitizers

The bactericidal activity of the immobilized PSs was tested against Gram-positive S. aureus and Gram-negative E. coli bacteria in the PACT experiments. For this purpose, bacterial suspensions in various concentrations were added to Petri plates with inner coatings of immobilized PSs, and illuminated with white light. The emission spectrum of the lamp (Fig. 3) overlapped the absorption regions of the used PS (Table 1) and at the same time, it contained no UV bands. Petri plates coated by polystyrene only, without any PS, served as controls. After testing several PS to polystyrene ratios for immobilization of PS, it was found that antimicrobial activity of the films, containing less than 10 mg of PS per gram of polystyrene, was low. Loading of MB in a higher concentration was problematic because of difficulties to get a homogeneous solution of MB and polystyrene in chloroform. For this reason in the PACT experiments, the loading of 10 mg (PS) per g (polystyrene) was used.

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Figure 3. Emission spectrum of the white luminescent lamp.

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As can be seen from Fig. 4, the immobilized PSs caused a decrease in the bacterial concentration. The concentrations of S. aureus and of E. coli dropped by 3 log10 and by 2.5 log10, respectively, in the presence of the immobilized RB (Fig. 4a, b). Taking into consideration bacterial growth in the control series, it can be concluded that actually, bacterial concentrations decreased by more than 4 log10 in both cases (P-values for comparison with control series were 0.0029 for S. aureus and 0.0038 for E. coli). Immobilized MB demonstrated lower than RB efficiency—S. aureus and E. coli concentrations were reduced by only 1.5 log10 and 1 log10, respectively (Fig. 4c, d), but when compared to the control samples at the end of experiments, the drop in the cell concentrations can be evaluated as ca 3 log10 for both bacteria (P-value in the case of S. aureus was 0.0027 and in the case of E. coli—0.013). In all cases, the initial bacterial concentration was 104 cells mL−1. No statistically significant difference was found between the control groups of bacteria incubated in the light and in the dark as well as between illuminated and darkened bacterial controls in the presence of polystyrene without PS inclusion (P-values of 0.5–0.9).

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Figure 4. PACT effect of RB (a and b) and MB (c and d) immobilized in polystyrene on the viability of S. aureus (a and c) and E. coli (b and d). Bacterial cells at the concentration of 104 cells mL−1 were incubated in the Petri plates coated by RB-polystyrene (RB/Ps), MB-polystyrene (MB/Ps) or polystyrene without PSs (Ps) for 0–3 h under illumination with white light having a 2.1 mW cm−2 fluence rate. Control groups of bacteria were incubated in uncoated Petri plates under illumination.

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To find out if the observed antibacterial activity was caused only by PSs immobilized or also by PSs leaked into the solutions during incubation of bacteria, the PBS solutions obtained after the incubation in the PS-polymer plates were tested for antibacterial activity in the conditions of the PACT experiment with the PS-polystyrene films. It can be seen from the Fig. 5a, b that MB, leaked in the dark or in the light, did not affect on viability of both tested bacteria—the 0.1–0.5 log10 differences between the control experiments and incubations under illumination of bacteria with leaked MB were not significant (P-values lied between 0.207 and 0.681). In the case of RB (Fig. 5c, d), there was no significant difference between the control experiments and RB leaked in the dark (for E. coli P-value was 0.69 and for S. aureus—0.28), but in the case of RB leaked under illumination, there were 0.9 and 2 log10 drops in bacterial concentration of E. coli (P-value = 0.032) and S. aureus (P-value = 0.038) correspondingly.

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Figure 5. PACT activity of PSs leaked from the PS-polymers upon 3 h incubation with PBS under illumination (“light washings”) or in the dark (“dark washings”). PACT experiments were carried out under additional 3-h illumination with white light having a 2.1 mW cm−2 fluence rate. Viability of S. aureus (a and c) and of E. coli (b and d) was tested in the presence of the leaked MB (a and c) or leaked RB (b and d). Control groups of bacteria were incubated in uncoated Petri plates under illumination.

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The activity of the immobilized PSs depended on the illumination intensity and on the initial bacterial concentration. At low bacterial concentrations (103 cells mL−1), both S. aureus (Fig. 6a) and E. coli (Fig. 6b) did not survive in the plates with RB-polystyrene coatings under a 30 min illumination at a light fluence rate of 2.1 mW cm−2. However, the same fluence rate applied for the same time was not enough for total eradication of bacteria at initial concentration of 104 cells mL−1, and ca 2% of S. aureus and ca. 5% of E. coli cells survived (Fig. 6c, d). At lower light intensities, the bacteria were either partially inactivated (1.1–1.5 log10 decrease in S. aureus concentration, Fig. 6a, c) or not affected at all (E. coli, Fig. 6b, d). Control experiments showed that illumination of non-treated cells in the presence of polystyrene alone as well as incubation of cells on plates coated with PS-polystyrene plates in the dark did not cause any decrease in the number of viable cells of both bacterial types.

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Figure 6. Photodynamic effect of RB immobilized in polystyrene on the viability of S. aureus (a and c) and E. coli (b and d) under 30 min illumination with white light at various fluence rates. Bacterial cells were incubated in the presence of RB immobilized on polystyrene (RB/Ps) or polymer without photosensitizer (Ps). Control groups of bacteria were incubated in uncoated Petri plates under illumination. Initial concentrations of bacteria were 103 cells mL−1 (a and b) or 104 cells mL−1 (c and d). After the treatment bacterial samples were serially diluted in 10-fold dilutions and evenly spread over BHA plates with a Drigalsky spreader. Plates were incubated at 37°C overnight and CFU were counted.

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SEM imaging of bactericidal effect of the photosensitizer-polystyrene surfaces

To visualize antibacterial properties of the immobilized PSs, suspensions of S. aureus and E. coli were incubated for a day under 1.8 mW cm−2 illumination on the PS-polystyrene films and on the polystyrene films alone (as a control), after which the films were examined under a SEM. The obtained micrographs (Figs. 7-9) show a drastic difference between the state of bacteria in the presence and in the absence of PS in the polymer. In control experiments on polystyrene films not containing PS, both S. aureus and E. coli were distributed over the entire polymer surface and were attached to it (Fig. 7a, c).

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Figure 7. SEM microphotographs of the control polystyrene films after incubation with suspensions of S. aureus (a and b) and E. coli (c and d) containing 106 cells mL−1 under 24 h illumination with light at a 1.8 mW cm−2 fluence rate at room temperature obtained with the help of a JEOL JSM 840 microscope at (a) ×1,527, (b) ×21 204, (c) ×2,400 and (d) ×30 000 magnification. Framed cells are in a process of biofilm formation.

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Figure 8. SEM microphotographs of the MB-polystyrene films after incubation with suspensions of S. aureus (a) and E. coli (b) containing 106 cells mL−1 under 24 h illumination with light at a 1.8 mW cm−2 fluence rate at room temperature obtained with the help of a JEOL JSM 840 microscope at (a) ×20 000 and (b) ×10 000 magnifications.

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Figure 9. SEM microphotographs of the RB-polystyrene films after incubation with suspensions of E. coli containing 106 cells mL−1 under 24 h illumination with white light at a 1.8 mW cm−2 fluence rate at room temperature obtained with the help of a JEOL JSM 840 microscope at ×10 000 magnification.

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Bacterial cells incubated in the presence of polystyrene films appeared to have normal shapes; they multiplied and propagated (Fig. 7b, d). Moreover, as seen in Fig. 7b, d, some of the cells started biofilm formation for stronger attachment to the polystyrene surface (see framed regions). In contradistinction, S. aureus cells (Fig. 8a) and E. coli cells (Fig. 8b) were very sparse on the MB-polystyrene films, and those which could be detected were in various stages of destruction. Actually, only bacterial remains are seen in Fig. 8. S. aureus cells were not found at all in the RB-polystyrene films, and the sporadic E. coli cells that were observed were all dead (Fig. 9).

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Hetero-phase bactericidal agents seem to be very attractive due to their wide range of potential medical and environmental applications. Immobilized PSs in a solid phase are a promising solution for development of long-term disinfection systems suitable for a continuous modus operandi. The photodynamic effectiveness of immobilized PSs was shown earlier by several groups. PSs immobilized on silicon polymer complexes of Ru(II) with poly-azo-heterocyclic compounds were shown to generate reactive oxygen species (ROS) under sunlight illumination [26]. Zn(II) phthalocyanine tetrasulfonic acid, covalently connected to a membrane polymer, was shown to have a high photodynamic efficacy in wastewater disinfection [25]. Several PSs incorporated under pressure and heat into poly(vinylidene fluoride) demonstrated high antimicrobial properties against Gram-positive and Gram-negative bacteria, decreasing their concentration by 4 log10 [30]. However, in some cases the rate of ROS formation by immobilized PSs can be significantly lower than with the free PSs [27].

The PSs studied in this work dissolve well in water, which allowed using them in a free form in aqueous solutions on the one hand, and demonstrate good solubility in organic solvents on the other hand, thus enabling dissolving them along with various polymers. This PS-polymer solution can be applied on chosen surfaces, and after solvent evaporation, the surfaces become coated with the active polymer. In this study, we chose polystyrene as the support polymer, which was able to mix with the tested PSs in a wide range of ratios due to the presence of aromatic rings in both PSs (Table 1) as well as in the polystyrene structure. Another advantage of the applied method was a high efficiency of immobilization (more than 99%), suggesting that only a negligible part of the PS may be released from the polymer support into the treated aqueous phase. The polymer was washed after the immobilization to remove traces of free PS, to ensure an antimicrobial effect of the immobilized PS without hindrance of free PS. However, in future practical applications of this system, it will be possible to skip this step because of low release of the free PS. The observed leakage of PS from the polymer can be presumably assigned to a minor destruction of the polystyrene under illumination. The data obtained in our study indicate that immobilized PSs caused a significant decrease in the concentration of the Gram-positive S. aureus and Gram-negative E.coli bacteria, up to their total eradication. The results of the experiments with the MB leaked from the polymer, show that the antibacterial activity of the MB-polystyrene was due only to the immobilized MB. In the case of RB-polystyrene, the total PACT effect was composed of the effect of the immobilized RB and of the effect of the RB leaked to the solution. It is important to notice that the observed antibacterial activity of the RB-polystyrene could not be solely explained by the presence of the leaked RB. We propose an explanation for the process of bacterial eradication by PSs immobilized in a solid phase. As shown in Fig. 2, round pores were formed on the surface of the polystyrene, which were big enough for enabling bacteria cells to enter into them, so that the effective surface area was enlarged compared to even and smooth polymer surfaces. Furthermore, according to the results obtained after a continual incubation of both S. aureus and E. coli on polystyrene films that did not contain a PS, the cells readily attached to the polymer surface, probably owing to its high hydrophobic properties. These factors led to increased contact between the polymer and the bacterial cells and to a possibility of high cell sorption by the polymer. Since a photodynamic effect can occur only in close proximity between the PS and the cells, the high polymer contact area was an important factor for exhibiting bactericidal activity by the examined system. Polystyrene by itself was not toxic to the cells at all, and both types of cells not only grew and multiplied on the polymer surface, but even started to form biofilms. In contradistinction, polymer films with the incorporated PSs did not contain cell aggregates and moreover, the single bacterial cells which were detected on the polystyrene surface had actually already been destroyed. These facts indicate that the hydrophobic attachment of the cells to the polymer, amplified by the porous polystyrene structure, facilitated the photodynamic activity of the immobilized PSs. PS molecules excited by light transferred energy to dissolved oxygen in the nearby polymer layer, promoting ROS formation, which in turn eradicated the bacterial cells attached to the polymer surface. Interestingly, the immobilized PS exhibited equal killing effects on E. coli and S. aureus. Apparently, for exhibiting the PACT effect, PS must not necessarily bind to the cell envelopes or enter into the cells. The obtained data open wide prospects for further applications of immobilized PS, for instance for inner coating of baths for aseptic holding of medical devices and for wastewater disinfection in a continuous regime. The latter application is now being studied by our group. In addition, using for immobilization of PSs polymer supports labile under illumination, systems of controlled release of PSs can be constructed.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

This research was supported in part by the Research Authority of the Ariel University Center of Samaria, by the Samaria and Jordan Rift Valley Regional R&D Center, by the Cherna Moskowitz Foundation, California, USA and by the Rappaport Foundation for Medical Microbiology, Bar Ilan University, Israel (to Y.N.). We acknowledge Mrs. Natalya Litvak (Ariel University Center of Samaria) and Dr. Yaakov Langzam, (Head of the E-SEM Unit of Bar-Ilan University) for their assistance in the SEM imaging of the samples).

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References