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 ~  Abstract
 ~ Introduction
 ~  Materials and Me...
 ~ Results
 ~ Discussion
 ~ Conclusion
 ~ Acknowledgement
 ~  References
 ~  Article Figures
 ~  Article Tables

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  Table of Contents  
ORIGINAL ARTICLE
Year : 2015  |  Volume : 33  |  Issue : 1  |  Page : 101-109
 

Anti-biofilm efficacy of silver nanoparticles against MRSA and MRSE isolated from wounds in a tertiary care hospital


1 Nanotechnology and Antimicrobial Drug Resistance Research Lab, Department of Microbiology , Jawaharlal Nehru Medical College and Hospital, Aligarh Muslim University, Aligarh, India
2 Department of Anatomy , Jawaharlal Nehru Medical College and Hospital, Aligarh Muslim University, Aligarh, India
3 Institute of Microbial Technology (IMTECH) , Sector 39 A, Chandigarh, India
4 Department of Medical Laboratories College of Applied Medical Science Buraydah Colleges, Buraydah 51452, Saudi Arabia

Date of Submission11-Jul-2013
Date of Acceptance29-Jan-2014
Date of Web Publication5-Jan-2015

Correspondence Address:
M A Ansari
Nanotechnology and Antimicrobial Drug Resistance Research Lab, Department of Microbiology , Jawaharlal Nehru Medical College and Hospital, Aligarh Muslim University, Aligarh
India
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Source of Support: The authors would like to acknowledge SAIFDST, Department of Anatomy, All India Institute of Medical Sciences (AIIMS), New Delhi, India, for HR-TEM and Advanced Instrumentation Research facility (AIRF), Jawaharlal Nehru University, New Delhi, India for SEM and Confocal laser scanning electron microscopy observation of AgNPs nanoparticle and bacterial interaction, Conflict of Interest: None


DOI: 10.4103/0255-0857.148402

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 ~ Abstract 

Purpose: Different approaches have been used for preventing biofilm-related infections in health care settings. Many of these methods have their own de-merits, which include chemical-based complications; emergent antibiotic resistant strains, etc. The formation of biofilm is the hallmark characteristic of Staphylococcus aureus and S. epidermidis infection, which consists of multiple layers of bacteria encased within an exopolysachharide glycocalyx. Nanotechnology may provide the answer to penetrate such biofilms and reduce biofilm formation. Therefore, the aim of present study was to demonstrate the biofilm formation by methicillin resistance S. aureus (MRSA) and methicillin resistance S. epidermidis (MRSE) isolated from wounds by direct visualisation applying tissue culture plate, tube and Congo Red Agar methods. Materials and Methods: The anti-biofilm activity of AgNPs was investigated by Congo Red, scanning electron microscopy (SEM) and confocal laser scanning microscopy (CLSM) techniques. Results: The minimum inhibitory concentration (MIC) was found to be in the range of 11.25-45 μg/ml. The AgNPs coated surfaces effectively restricted biofilm formation of the tested bacteria. Double fluorescent staining (propidium iodide staining to detect bacterial cells and fluorescein isothiocyanate concanavalin A (Con A-FITC) staining to detect the exopolysachharides matrix) technique using CLSM provides the visual evidence that AgNPs arrested the bacterial growth and prevent the glycocalyx formation. In our study, we could demonstrate the complete anti-biofilm activity AgNPs at a concentration as low as 50 μg/ml. Conclusions: Our findings suggested that AgNPs can be exploited towards the development of potential anti-bacterial coatings for various biomedical and environmental applications. In the near future, the AgNPs may play major role in the coating of medical devices and treatment of infections caused due to highly antibiotic resistant biofilm.


Keywords: Anti-biofilm, AgNPs, confocal laser scanning microscopy, exopolysachharide, scanning electron microscopy


How to cite this article:
Ansari M A, Khan H M, Khan A A, Cameotra S S, Alzohairy M A. Anti-biofilm efficacy of silver nanoparticles against MRSA and MRSE isolated from wounds in a tertiary care hospital. Indian J Med Microbiol 2015;33:101-9

How to cite this URL:
Ansari M A, Khan H M, Khan A A, Cameotra S S, Alzohairy M A. Anti-biofilm efficacy of silver nanoparticles against MRSA and MRSE isolated from wounds in a tertiary care hospital. Indian J Med Microbiol [serial online] 2015 [cited 2019 Nov 22];33:101-9. Available from: http://www.ijmm.org/text.asp?2015/33/1/101/148402



 ~ Introduction Top


Staphylococci are most often associated with chronic infections of implanted medical devices. [1] The use of indwelling medical devices is important in the treatment of critically and chronically ill patients, however, bacterial colonisation of implanted foreign material can cause major medical and economic sequel. The increased use of indwelling medical devices has had considerable impact on the role of staphylococci in clinical medicine. The predominant species isolated in these infections are Staphylococcus epidermidis and S. aureus, their major pathogenic factor being ability to form biofilm on polymeric surfaces. [2] Biofilm producing staphylococci frequently colonise catheters and medical devices and may cause foreign body-related infections. They easily get attached to polymer surfaces. [3] Bacterial biofilms are a predominant challenge to wound healing. [4]

The first recorded observation concerning biofilm was probably given by Henrici in 1933, who observed that water bacteria are not free floating but grow upon submerged surfaces. [5] Biofilm consists of multilayered cell clusters embedded in a matrix of extracellular polysaccharide (slime), which facilitates the adherence of these microorganisms to biomedical surfaces and protect them from host immune system and anti-microbial therapy. [6] Biofilm formation is regulated by expression of polysaccharide intracellular adhesin (PIA), which mediates cell to cell adhesion and is the gene product of icaADBC.[7] Various reports attest to the presence of icaADBC gene in S. aureus and S. epidermidis isolated from infections associated with indwelling medical devices. [8]

It is now well documented that biofilms are notoriously difficult to eradicate and are often resistant to systemic antibiotic therapy and removal of infected device becomes necessary. [3],[9] According to National Institute of Health, more than 60% of all infections are caused by biofilm. [10] Biofilm organisms have an inherent resistance to antibiotics, disinfectants and germicides. Unlike planktonic populations, bacterial cells embedded in biofilms exhibit intrinsic resistance to antibiotics due to several specific defense mechanisms conferred by the biofilm environment, including the inactivation of anti-microbial agents by exopolysachharide (EPS), over expression of stress-responsive genes, oxygen gradients within the biofilm matrix and differentiation of a subpopulation of biofilm cells into resistant dormant cells. [11],[12] The intrinsic resistance of bacterial cells within biofilms to conventional anti-microbials has motivated new approaches for the treatment of biofilm-associated infections, including the use of silver preparations. Several silver-containing dressings are recommended for long-term de-contamination and wound healing based on silver's broad-spectrum, high-level anti-microbial activity. [13] The difficulty in eradicating a chronic infection associated with biofilm formation lies in the fact that biofilm bacteria are able to resist higher antibiotic concentration than bacteria in suspension. [14] Nanotechnology may provide the answer to penetrate such biofilms and reduce biofilm formation. Silver nanotechnology chemistry can prevent the formation of life-threatening biofilms on medical devices. Silver is one of the oldest known anti-microbials. It has recently been demonstrated that AgNPs hydrogel hybrid with different sizes of AgNPs can be effectively employed as anti-bacterial agents. [15] Saxena et al. studied that propylene-based sutures immobilised AgNPs show anti-bacterial activity against S. aureus and  Escherichia More Details coli. [16] Due to the strong anti-bacterial properties and low toxicity towards mammalian cells, AgNPs have been applied in a wide range of areas including wound dressing, coatings on medical devices to reduce nosocomial infection rates, [17] protective clothing, anti-bacterial surfaces, water treatment, food preservation and cosmetics as biocidal and disinfecting agents. [18] Although the literature reports that some studies are related to the anti-bacterial activity of AgNPs, to the authors' knowledge, there are very few studies concerning the effect of these particles against adhered cells and biofilms of methicillin resistance S. aureus (MRSA) and methicillin resistance S. epidermidis (MRSE). Thus, the aim of the present study was to evaluate the anti-biofilm potential of AgNPs against biofilms of MRSA and MRSE by Congo Red Agar (CRA), scanning electron microscopy (SEM) and confocal laser scanning microscopy (CLSM).


 ~ Materials and Methods Top


Characterisation of silver nanoparticles

A stock solution of commercially available water soluble AgNPs (5-10 nm) were procured from Nanoparticle Biochem, Inc. (Columbia, USA). The subsequent dilutions were made in autoclaved Milli Q water. The morphological features and particle size of the procured NPs were characterised by high-resolution transmission electron microscopy (HR-TEM, Technai, FEI, Electron Optics, USA) [Figure 1].
Figure 1: HR-TEM image of AgNPs dispersion

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Bacterial strains, media and materials

A total of 62 non-repetitive clinical isolates of Staphylococcus spp were recovered over a period of 6 months (September 2012 to February 2013) from skin lesion such as pus, wounds, burn, etc., were subjected to the study.

In vitro antibiotic susceptibility test

Individual isolates were tested, based on the recommendations of the Clinical and Laboratory Standards Institute (CLSI), [19] by the Kirby-Bauer disc diffusion method for susceptibility to the following antibiotics: Amikacin (AK, 30μg), Gentamycin (G, 10μg), Oxacillin (Ox, 1 μg), Ceftriaxone (Ci, 30 μg), Cefotaxime (Ce, 30 μg), Vancomycin (Va, 30 μg), Chloramphenicol (C, 30 μg), Tobramycin (Tb, 10 μg), Novobiocin (No, 30 μg), Levofloxacin (Le, 5 μg), Clindamycin (Cd, 1 μg), Erythromycin (E, 1 μg). Antibiotic discs used were procured by Hi-Media (Mumbai, India).

Screening for methicillin resistance

Resistance to oxacillin was determined in all S. aureus isolates using an oxacillin broth screening and disc diffusion test recommended by the British Society for Antimicrobial Chemotherapy (BSAC), [20] respectively. For the oxacillin broth screening test, isolated colonies from an 18 to 24 h sheep blood agar plate were used to prepare a direct inoculum equivalent to a 0.5 McFarland and suspended in 2 ml nutrient broth, supplemented with 2% NaCl. Breakpoints published by the CLSI were used: oxacillin susceptible (S) <2 mg/L and resistant (R) ≥4 mg/L, on Mueller-Hinton agar (MHA, Hi-Media, Mumbai, India) plate with 2%NaCl incubated at 35°C for overnight.

For the disc diffusion test, a sterile swab was dipped in the same S. aureus suspension as used for the oxacillin broth screening test. The swab was plated on MHA plate in order to get confluent growth (approx. 0.5 McFarland). Oxacillin discs (1 μg) were applied on to the surface of inoculated agar. Plates were incubated overnight at 30°C. An isolate was classified as resistant to oxacillin when the inhibition zone was ≤14 mm diameter. [20]

Detection of biofilm formation

Tissue culture plate method

The tissue culture plate (TCP) assay, described by Christensen et al., is most widely used and was considered as standard test for detection of biofilm formation. [21] A total of 10 ml of Trypticase soy broth (TSB) with 1% glucose was inoculated with a loopful of test organism from overnight culture on nutrient agar. The broth was incubated at 37°C for 24 h. The culture was further diluted 1:100 with fresh medium. 96 wells flat bottom TCPs were filled with 0.2 ml of diluted cultures individually. Only sterile broth was served as blank. Similarly control organisms were also diluted and incubated. The culture plates were incubated at 37°C for 24 h. After incubation gentle tapping of the plates was done. The wells were washed with 0.2 ml of phosphate buffer saline (pH 7.2) four times to remove free floating bacteria. Biofilms, which remained adherent to the walls and the bottoms of the wells, were fixed with 2% sodium acetate and stained with 0.1% crystal violet. Excess stain was washed with de-ionised water and plates were dried properly. Optical densities (OD) of stained adherent biofilm were obtained with a micro ELISA autoreader at wave length 570 nm. Experiment was performed in triplicate and repeated thrice. Average of OD values of sterile medium were calculated and subtracted from all test values. [25]

Tube method

A qualitative assessment of biofilm formation was determined as previously described. [22] A total of 10 ml TSB with 1% glucose was inoculated with a loopful of microorganism from overnight culture plates and incubated for 24 h at 37°C. The tubes were decanted and washed with phosphate buffered saline (PBS) 0.1% (pH 7.3) and dried. Dried tubes were stained with crystal violet (0.1%). Excess stain was removed and tubes were washed with de-ionised water. Tubes were than dried in inverted position and observed for biofilm formation. Biofilm formation was considered as positive, when a visible film lined the wall and bottom of the tube. Ring formation at the liquid interface was not indicative of biofilm formation.

Congo red agar method

Freeman et al. had described an alternative CRA method for screening biofilm formation by Staphylococcus isolates. [23] Plates were inoculated and incubated aerobically for 24-48 h at 37°C. Positive result was indicated by black colonies with a dry crystalline consistency. Weak slime producers usually remained pink, though occasional darkening at the centres of colonies was observed. A darkening of the colonies with the absence of a dry crystalline colonial morphology indicated an indeterminate result. The experiment was performed in triplicate and repeated three times.

Scanning electron microscopy

Biofilms were assessed as previously described with or without AgNPs. Briefly, the cells were washed with PBS, fixed with 2.5% glutaraldehyde, then fixed samples were subsequently washed again with PBS and dehydrated gently by washing in a series of ethanol alcohol (30%, 50%, 70%, 80%, 95% and 100%) for 10 min at room temperature, and critical point drying was performed. Afterwards, cells were then oriented, mounted on the aluminium stubs and coated with gold before imaging. The topographic features of the biofilms were visualised with a SEM (Carl Zeiss EVO 40, Germany) with accelerating voltage of 20 kV.

Confocal laser scanning microscopy

Biofilms for confocal analysis were grown on glass coverslips as previously described. [24] Briefly, 12-well microtitre plate seeded with glass coverslips were incubated for 24 h at 37°C in 5 ml of brain heart infusion (BHI) with 5% sucrose. The wells were inoculated with 100 μl of mid-exponential grown culture of S. aureus After 24 h, the cover slips were removed and gently washed three times with sterile PBS and were first stained with 15 μM Propidium iodide for 5 min at room temperature to detect bacterial cells in red. After being washed in PBS, the sections were incubated with 50 μg/ml of concanavalin A-fluorescein isothiocyanate (Con A-FITC) (C7642; Sigma-Aldrich Inc, St Louis, MO, USA) for 5 min at room temperature to stain the glycocalyx matrix green. The Propidium iodide was excited at 520 nm, the emission was monitored at 620 nm and Con A-FITC was excited and monitored at 495 and 525 nm, respectively. Intact biofilms were examined non-destructively using a Fluoview FV1000 Espectral Olympus CSLM (Olympus Latin America, Miami, FL, USA) equipped with a UPlanSApo 100X/1.40 oil UIS2 Olympus oil immersion lens. Optical sections of 0.87 μm were collected from the complete thickness of the biofilms. Then, for each sample, images from three randomly selected positions were obtained and analysed using an Olympus Fluoview FV 1000.


 ~ Results Top


Prevalence of bacterial isolates

Sixty-two Staphylococcus species were isolated from the patients associated generally with the diseases of purulent dermatitis, burns, wounds and other localised skin infections. Out of 62 isolates screened, 25 (43.9%) isolates were MRSA, 27 (47.5%) were methicillin sensitive S. aureus (MSSA), 6 (5.3%) were methicillin resistance Coagulase negative Staphylococcus (MR-CONS) and 4 (3.5%) methicillin sensitive Coagulase negative Staphylococcus (MS-CONS).

Antibiotic susceptibility testing

Results obtained from investigation of resistance in Staphylococcus spp isolates, which were susceptible or resistant to individual antibiotics, are displayed in [Table 1].
Table 1: Antimicrobial susceptibility pattern of Staphylococcus species (N=62)

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Evaluation of anti-bacterial activity of AgNPs

The MIC and MBC of AgNPs dispersion tested against MRSA, MSSA and MRSE were observed in the range of 11.25-45 μg/ml [Table 2]a and b]. The result clearly shows that AgNPs exhibit excellent bacteriostatic and bactericidal effect regardless of their drug-resistant mechanisms.
Table 2: MIC and MBC values of AgNPs tested against clinical isolates of Staphylococcus species

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Screening of biofilm formation On CRA

[Table 3]a shows that out of 52 isolates of S. aureus and 10 of S. epidermidis tested for biofilm formation by CRA method, 41 (78.85%) isolates of S. aureus and 8 (80%) isolates of S. epidermidis produced black colonies. However, only 25 (48.08%) isolates of S. aureus and 5 (50%) isolates of S. epidermidis colonies were black in colour with dry crystalline consistency, which is indicative of biofilm formation. A total of 16 (30.77%) S. aureus and 3 (30%) S. epidermidis isolates were black in colour but were not dry and crystalline and were indeterminate for biofilm formation. These isolates were also taken as negative for biofilm formation. A total of 11 (21.15%) S. aureus and 2 (20%) S. epidermidis isolates produced pink colonies, which were taken as negative for biofilm formation.
Table 3: Screening of biofilm formation of Staphylococcus species by (a) Congo Red Agar, (b) Tissue Culture Plate and (c) Tube methods

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Screening of biofilm formation by TCP method

In the quantitative assay for the biofilm production, the isolates were classified as highly biofilm producing (strongly adherent), moderate adherent isolates and non-biofilm producers (weak/non-adherent). Quantitative microtitre assay for biofilm formation was strongly positive in six (11.54%) isolates of S. aureus and three (30%) S. epidermidis, while the remaining isolates were either moderate adherent 30 (57.69%) S. aureus and 4 (40%) S. epidermidis or weak/non-biofilm producers 16 (30.77%) S. aureus and 3 (30%) S. epidermidis [Table 3]b]. Isolates screened for biofilm formation by microtitre plate assay method are shown in [Figure 2].
Figure 2: Screening of biofilm producers by TCP method: High, moderate and non-slime producers differentiated with crystal violet staining were shown on 96 well tissue culture plates

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Screening of biofilm formation by tube method

Qualitative tube method of biofilm screening of bacterial isolates showed that 33 (63.46%) isolates of S. aureus and 7 (70%) isolates of S. epidermidis were positive for biofilm production [Table 3]c]. Most of the isolates showed thick blue ring at the liquid-air interface.

Characterisation of anti-biofilm activity of AgNPs on CRA Plates

Biofilm formation is detected in many organisms synthesising exopolysachharides. The biofilm is made up of microorganisms adhering to the surface coated with slime - the exopolysachharide matrix which protects the microbes from the unfavourable environmental factors. Biofilm formation by S. aureus isolates were tested by growing the organism in brain-heart infusion agar supplemented with Congo Red (BHIC) with and without silver nanoparticles. When the colonies were grown without AgNPs in the medium, the organisms appeared as dry crystalline black colonies, indicating the production of exopolysachharides, which is the prerequisite for the formation of biofilm [Figure 3]. Whereas when the organisms were grown on BHIC with AgNPs, the organisms did not survive. During the treatment with reduced concentrations of AgNPs (10 μg/ml), the organisms continued to grow, but AgNPs treatment has inhibited the synthesis of glycocalyx matrix, indicated by the absence of dry crystalline black colonies [Figure 3]. It was found that at higher concentration of AgNPs inhibited bacterial growth by more than 98%. When the glycocalyx matrix synthesis is arrested, the organism cannot form biofilm. It was also observed that 20 μg/ml of AgNPs significantly arrested biofilm formation without affecting viability, whereas 50 μg/ml completely blocked the biofilm formation and inhibited the growth of the organism itself.
Figure 3: Ability of the organisms was checked for biofilm formation in BHI agar supplemented with Congo Red. The appearance of black crystalline colonies (a and b) indicate the exopolysachharide production by MRSA (a) and MRSE (b), whereas the addition of 50 μg/ml AgNPs blocked the exopolysachharide synthesis by MRSA (c) and MRSE (d) and also inhibited the growth of the organism itself

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Characterisation of anti-biofilm activity of AgNPs by scanning electron microscopy

The biofilm grown on glass slide for 24 h was observed using SEM. The biofilm formed by the native strain have aggregated and clumped bacterial cells [[Figure 4]a]. The S. aureus cultures grown without AgNPs exhibit the expected normal cellular morphology with smooth cell surfaces [[Figure 4]a]. Under the same growth conditions but in the presence of AgNPs (20 μg/ml) S. aureus cell shows changes in morphology and it was also examined that AgNPs inhibit bacterial colonisation on the surfaces [[Figure 4]b]. The apparent biofilm formed by S. aureus has very few cells individually scattered along the surface. The cells were arranged in short chains with absence of exopolysachharides matrix. The biofilm formed by the S. aureus was very much patchy [[Figure 4]a]. More specifically, an obvious increase in the roughness of the cell surface suggested that it has been damaged by the nanoparticles. Microscope evaluation of the surfaces clearly shows that the AgNPs treated glass surfaces do not allow bacterial colonisation and biofilm formation compared with the untreated controls. Untreated glass surfaces supported a massive biofilm formation by S. aureus [[Figure 4]a], while AgNPs treated glass surfaces shows dramatically restricted bacterial colonisation and biofilm formation [[Figure 4]b]. These results suggest that AgNPs are effective in restraining bacterial colonisation of the surface.
Figure 4: SEM images of S. aureus biofilms developed on the glass slide surface at 24 h of incubation. Untreated (a) and treated (b) with AgNPs (20 μg/ml)

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Characterisation of anti-biofilm activity of AgNPs by Confocal laser scanning microscopy

CLSM analysis of biofilms formation by MRSA [[Figure 5]a] and MRSE [[Figure 5]b] was performed to examine the effects of coating of AgNPs on biofilm architecture using AgNPs-coated glass cover slips in a 12-well microtitre plate. To visualise bacterial cells and the surrounding glycocalyx matrix (which is indicative of bacterial biofilm formation), double staining was performed using propidium iodide and Con A-FITC. Bacterial cells stained red and were easily identified by their size and morphologic features, when using propidium iodide. Whereas Con A-FITC binds to mannose residues resulting in green staining and indicating the presence of a bacterial glycocalyx. Although, we observed the marked co-localisation of green Con A-FITC staining with clusters of bacterial cells, the staining of the matrix was not homogeneously distributed. The presence of dark areas within the biofilm can be explained by the existing water channels, the heterogeneous production of the matrix and the types of exopolysachharides within the biofilm, as well as the absence of Con A-FITC binding to the matrix. When CLSM images with red and green fluorescent intensities were superimposed, yellow colour (green + red) revealed that the extracellular-Con A-FITC-reactive polysaccharide (green) was produced in the intra-cellular spaces (red), indicating thereby that extracellular polysaccharides were produced as a capsular component in biofilm. We observed that interconnected bacteria were encased in a scaffolding network composed of extracellular matrix, suggesting a 3-dimensional architecture of biofilm formations. The micrographs suggest that the biofilm formed on uncoated AgNPs surfaces covered a larger surface area and had a definite architecture ([Figure 5]a and b, first panel). However, the biofilms formed on coverslips coated with AgNPs showed no or few spread cells with no distinct pattern of arrangement. S. aureus cells exhibited shape distortions, indentations and slight elongation ([Figure 5]a and b second panel), after exposure to AgNPs. The cells grown in the presence of 25 μg/ml AgNPs show a complete absence of clumped cells and were individually scattered over the surface rather than in any arrangement. Uncoated cover slips surfaces supported a massive biofilm formation of all bacterial cells, whereas AgNPs-coated surfaces dramatically restricted bacterial colonisation.
Figure 5: CLSM micrographs of MRSA (a) and MRSE (b) biofilm. Panel 1 from left to right represents CLSM images of native biofilm (without AgNPs). Whereas panel 2 from left to right represent CLSM images of MRSA (a) and MRSE (b) biofilm treated with AgNPs and most of the cells exhibited shape distortions and slight elongation and almost no exopolysachharides (green fluorescent) was observed. The number of live bacterial cells was reduced significantly, and the 3-dimensional structure was also disrupted. Magnification ×100

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 ~ Discussion Top


In recent years, there has been considerable interest in the problems posed by the biofilm mode of bacterial and fungal growth. According to public announcement from national institute of health, more than 60% of all microbial infection is caused by biofilms. [25] Infections resulting from microbial biofilm formation remain a serious threat to patients worldwide. Particularly problematic are wound infections, [26] with chronic wounds such as foot, leg and pressure ulcers being particularly susceptible to biofilm infections. [27] In order to kill or remove biofilms, anti-microbials must penetrate the polysaccharide matrix to gain access to the microbial cells. Nanotechnology may provide the answer to penetrate such biofilms and reduce biofilm formation by the use of 'nano functionalisation' surface techniques to prevent the biofilm formation. Biofilm formation by clinical isolates of S. aureus was tested by CRA, tube and TCP methods. However, the anti-biofilm efficacy of AgNPs was investigated by growing the organism on CRA supplemented with and without AgNPs. When the colonies were grown without AgNPs, the organisms appeared as dry crystalline black colonies, indicating the production of exopolysachharides, which is the prerequisite for the formation of biofilm [Figure 3]. Whereas when the organisms were grown with AgNPs, the organisms did not survive. During the treatment with reduced concentrations of AgNPs (10 μg/ml), the organisms continued to grow, but AgNPs treatment has inhibited the synthesis of glycocalyx matrix, indicated by the absence of dry crystalline black colonies. However, at higher concentration of AgNPs (50 μg/ml) almost no growth was observed [Figure 3]. Thus, when the glycocalyx synthesis is arrested, the organism cannot form biofilm. Similar results were also reported by Kalishwaralal et al. against P. aeruginosa and S. epidermidis biofilms and found that 100 nM of AgNPs resulted in a 95-98% reduction in biofilm. [28] It was also cited in literature that if the surface of medical devices has an AgNPs coating, then it was helpful in preventing bacterial adhesion and subsequent biofilm formation on the medical devices. [29]

Based on these results we continued to address the mechanism of action of AgNPs on biofilm forming bacterial species by applying CLSM and SEM. SEM was used to examine cell morphologies following exposure to the nanoparticles. S. aureus cultures grown without nanoparticles exhibit the expected normal cellular morphology with smooth cell surfaces [[Figure 4]a]. Under the same growth conditions but in the presence of suspended AgNPs (30 μg/ml), S. aureus bacteria presented a change in morphology [[Figure 4]b]. More specifically, an obvious increase in the roughness of the cell surface suggested that it has been damaged by the nanoparticles. EPS within S. aureus biofilms was not detected by SEM. SEM observations clearly indicate that AgNPs reduced the surface coverage by S. aureus [[Figure 4]b] biofilms. Our results are in agreement with previously reported anti-biofilm activity of nanocrystalline silver by Kostenko et al. [30] They observed that nanocrystalline silver dramatically decreased viable cell numbers within the tested biofilms. SEM results shows that nanocrystalline silver reduced the surface coverage by P. aeruginosa biofilms. Prolonged treatment with nanocrystalline silver provided a further reduction in the surface coverage by MRSA biofilms. The surface coverage by E. coli biofilms was reduced by the tested dressings by approximately 20% after the first day. [30]

In order to achieve a fundamental understanding of the formation and presence of bacterial biofilms, the analysis should include detection of the bacteria and the matrix. The most common methods of assessing biofilm heterogeneity are direct microscopic imaging of local biofilm morphology or microscopic measurement of the local biofilm thickness. For many applications, time lapse microscopy using CLSM is an ideal tool for monitoring at a spatial resolution of the order of micrometers, and allows the non-destructive study of biofilms through an examination of all the layers at different depths, thus making it is possible to reconstruct a 3-dimensional structure. [31] The detection of the matrix can be achieved using a double-staining technique in combination with CLSM, which allows the simultaneous imaging of bacterial cells as well as glycocalyx within biofilms. [32]

Therefore, to visualise the bacterial cells and the surrounding glycocalyx matrix (which is indicative of bacterial biofilm formation), we used this powerful technique (CLSM) to examine the effect of nanoparticles on glycocalyx matrix/exopolysachharides synthesis; double staining was performed using propidium iodide and Con A-FITC. In case of untreated S. aureus ([Figure 5]a and b, panel 1), PI stain bacterial nucleic acids and fluorescent red, while, green fluorescent (Con A-FITC) around bacteria indicates the presence of exopolysachharides. Whereas AgNPs treated samples of S. aureus ([Figure 5]a and b, panel 2) shows that most of the cells were dead and no glycocalyx matrix (green fluorescent) was observed. The number of live bacterial cells was reduced significantly, and the 3-dimensional structure was also disrupted. We investigated that this inhibitory effect of AgNPs on the existing biofilm may due to be presence of water channels though out the biofilm. Since in all biofilms, water channels (pores) are present for nutrient transportation, AgNPs may directly diffuse through the glycocalyx matrix layer through the pores and may impart anti-microbial function. Masurkar et al. investigated the anti-biofilm activity of AgNPs (51 nm) synthesised from B. megaterium and reported that AgNPs showed enhanced quorum quenching activity against S. aureus biofilm and prevention of biofilm formation, which can be seen under inverted microscope. They concluded that AgNPs might be involved in neutralising these adhesive substances, thus preventing biofilm formation. [33]

However, the exact mechanism of action of AgNPs in biofilm-related studies is yet to be demonstrated. Kostenko et al. reported that Acticoat nanocrystalline silver has the highest anti-biofilm efficacy compared with Aquacel silver and Silverlon and the silver concentration alone cannot account for the anti-biofilm efficacy of the silver dressings. The type of silver species present also plays a role. [30] The reduction of the silver particle to the nanoscale level increases the relative surface area, which provides higher Ag + release rates than for elemental silver particles. [34] Moreover, nanoparticles have a higher capacity to attach to and penetrate bacterial membranes and accumulate inside cells, providing a continuous release of silver ions inside the cell. [35],[36],[37]


 ~ Conclusion Top


When MRSA and MRSE assume the biofilm phenotype, these infections are often extremely difficult to treat. The infection may fail to respond to antibiotic therapy or it may initially respond only to relapse weeks or months later. In such cases, invasive treatments, such as surgical removal and replacement of the infected tissue or device, may be required. So for proper treatment of S. aureus infection screening for biofilm production is necessary. The presence of biofilms on medical devices or surfaces can only be observed using a limited number of techniques. The reason why the demonstration of bacterial biofilms is challenging is because it is difficult to stain both the bacteria and glycocalyx. Furthermore, light and electron microscopy techniques require a dehydration process that reduces the total volume of the matrix and alters its architecture. CLSM is an ideal tool for monitoring at a spatial resolution of the order of micrometers, and allows the non-destructive study of biofilms through an examination of all the layers at different depths, thus making it possible to reconstruct a 3-dimensional structure. The detection of the glycocalyx matrix can be achieved using a double-staining technique in combination with CLSM, which allows the simultaneous imaging of bacterial cells as well as glycocalyx within biofilms. In the near future, the AgNPs may play major role in the coating of medical devices and treatment of infections caused due to highly antibiotic-resistant biofilm.


 ~ Acknowledgement Top


The authors would like to thank the Indian Council of Medical Research (ICMR) New Delhi-India, grant number 35/15/BMS-11 for their partial support and funding of this project.

 
 ~ References Top

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    Figures

  [Figure 1], [Figure 2], [Figure 3], [Figure 4], [Figure 5]
 
 
    Tables

  [Table 1], [Table 2], [Table 3]



 

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2004 - Indian Journal of Medical Microbiology
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