Characterization of an exopolysaccharide from probiont Enterobacter faecalis MSI12 and its effect on the disruption of Candida albicans biofilm

Abstract

Biofilm-forming pathogens are a potential threat to indwelling medical devices and infectious diseases. Management of device-associated Candida infections remains challenging with the existing drug discovery platforms. The available antifungal drugs are effective for the control of free-living pathogens but not effective on biofilm-forming pathogens. Thus an antifungal drug synergized with an antibiofilm agent would be an effective strategy to treat Candida biofilms. Mature C. albicans biofilms are anchored by a complex architecture in terms of distribution of fungal cells stabilized by exocellular polymeric substances. The findings of the present study provide a new insight on the possible development of enterococci probiotics and/or its exopolysaccharide (EPS) as synergistic with existing antifungal drugs to treat biofilm infections. The probiont Enterococcus faecalis MSI12 was picked from 142 seawater isolates screened for EPS production using a congo red plate assay. The probiotic characteristics of the isolate MSI12 were evaluated based on the temperature, pH, acid tolerance, autoaggregation, hydrophobicity and antioxidant activity. The biofilm disruption ability of the lyophilized EPS was determined in a microtitre plate assay using fluconazole as reference drug. Scanning electron microscope and confocal laser scanning microscope images were used for analysis of antibiofilm activity. The cell viability of E. faecalis MSI12 was very high at higher temperature, acidic pH, bile salt and salt concentration when compared to the reference strain Lactobacillus plantarum. Therefore the strain MSI12 might survive in the niche like human gut without prebiotics. The EPS from Enterococcus sp. MSI12 showed significant reduction of the treated Candida biofilm. The antibiofilm potential of EPS was much stronger than the standard antifungal drug fluconazole. This study revealed that biofilm disruption/control using a probiont EPS could deliver a synergistic approach as the probiotic strain can colonize in the host to prevent the formation of Candida biofilms.

---

1. Introduction

Biofilms are formed by a biomass of bacterial cells housed in exo-polymeric substances on biotic and abiotic surfaces. Biofilm-forming pathogens are a major threat to human health and a challenge to existing drug discovery platforms. It has been reported that up to 80% of clinical infections are caused by biofilms and the formation of biofilms on indwelling medical devices are responsible for a large proportion of nosocomial infections.¹ The pathogenic species of the genus Candida have gained recognition as nosocomial agents in recent years which may be attributed to specific risk factors associated with modern therapeutics like immunosuppressive agents, cytotoxic drugs, and broad-spectrum antibiotics that suppress the normal bacterial microbiota. Candidiasis is normally associated with indwelling medical devices such as dental implants,² catheters, heart valves, vascular bypass grafts, ocular lenses, artificial joints, central nervous system shunts etc. which can act as abiotic substrates for biofilm growth.³ Even with current antifungal therapy, mortality of patients with invasive candidiasis can be as high as 40%.^3,4 A biofilm is a shield of progressive pathogenic Candida cells, which facilitate escape from the host immune system and dramatically reduce susceptibility to clinically used antifungal drugs like fluconazole, nystatin, amphotericin B, and chlorhexidine.³ The available antifungal drugs are effective on the control of Candida pathogens but not effective on their biofilms as the drugs cannot cross the biofilm boundary to target the cells. The exo-polymer matrix in the biofilm can restrict access of the antimicrobial agents or the microbial cells present deep in the colony escape the drug or changes in the membrane sterol composition during biofilm development makes the microbial cells resistant to antibiotics.⁵ Thus an antifungal agent synergized with antibiofilm agents would be an effective strategy to treat Candida biofilms.

The genus Enterococcus belongs to lactic acid bacteria (LAB), a prominent group of probiotics in human and animal healthcare. This genus is typically characterized as non-spore forming Gram-positive bacteria which appear as cocci, diplococci or short chains, and produce lactic acid as the major end product of glucose fermentation.⁴ This genus is a proven group of probiotics and third-largest genus of LAB after the genera Lactobacillus and Streptococcus.⁵ Among the LAB, the enterococci can be designated as robust probiotics due to their high salt, temperature and pH tolerance. A well-characterized and commercial probiotic in the genus enterococci is E. faecium SF68 which is used in the treatment of diarrhoea.⁷ The literature shows that enterococci probiotics are effective in the prevention of antibiotic-associated diarrhoea.⁶ But their potential for the disruption/prevention of pathogenic biofilms has not been reported. The enterococci genus occurs in diverse habitats including soil, surface waters, ocean water, sewage, on plants and in the gastrointestinal tract.⁶ In this study, the isolation was performed from seawater to isolate an enterococci genus with high salt, pH and temperature tolerance. Marine ecosystems represent a large and as yet largely under explored reservoir of biodiversity with respect to industrially useful enzymes, bioactive molecules and biosurfactants. These ecosystems can range from coastal environments to deep-sea hydrothermal vents with high hydrostatic pressure and temperatures as high as approximately 400 °C.⁸

Exopolysaccharides (EPSs) are carbohydrate polymers that make up a substantial component of the extracellular polymers surrounding most microbial cells in the marine environment.⁹ The EPSs support the microbial communities to survive extremes of temperature, salinity, and nutrient limitation. EPSs can be used as bioflocculants for the synthesis of silver nanoparticles, bioabsorbents, encapsulating materials, heavy metal removing agents, drug delivery agents, natural immunomodulators, antioxidant and antibiofilm agents.^10–12 It has been reported that mature C. albicans biofilms have a highly complex architecture in terms of distribution of fungal cells in extracellular material.³ The common antifungal drugs are not effective to treat established (biofilm) Candida infections. We hypothesized that probiotics with antibiofilm potential would be a synergistic approach to treat established pathogens. The findings of the present study provide a new insight on the possible development of enterococci probiotics and their EPSs as synergistic to the antifungal drugs being used for antiinfective therapy. In this study, we report EPS isolation from a probiotic strain Enterococcus faecalis MSI12 isolated from the marine environment and its potential as an antibiofilm agent and/or disruptor of pre-established biofilms of the most prominent human pathogen C. albicans. This is a first report of probiotic EPS produced by marine E. faecalis disrupting the pathogenic biofilm of C. albicans.

2. Materials and methods

2.1 Isolation and identification of EPS producing marine bacterium

Seawater samples were collected aseptically at 150 m depth from the coastal region of Pondicherry, India (11° 52′ 56 N′′; 11° 59′ 53′′ E). Bacterial isolation was performed on Zobell marine agar (Himedia). Based on the stability in sub-culturing steps, 142 isolates were screened for EPS production. The congo red plate assay was performed for screening of EPS producing isolates. The stain congo red binds to the extracellular matrix such as polysaccharides and cellulose to give a dark-red stained background around the colony.¹³ Among the 11 positive isolates, the maximum EPS producing isolate MSI12 was selected for further characterization. The selected isolate MSI12 was identified by 16S rRNA gene sequence analysis. Briefly, genomic DNA from strain MSI12 was extracted with suitable modification using the previously described method.¹⁴ The 16S rRNA gene was amplified using universal primers 8F (5′-AGA GTT TGA TCC TGG CTC AG-3′) and 1492R (5′-GGT TAC CTT GTT ACG ACT T-3′) and the gene product was cloned using the TOPO TA cloning kit according to the manufacturer’s instructions. The 16S rRNA sequence of the isolate was then analyzed using the megaBLAST algorithm of GenBank (http://www.ncbi.nlm.nih.gov/genbank). Sequences of the isolate MSI12 were confirmed with seqmatch program of RDPII (http://rdp.cme.msu.edu/seqmatch/seqmatch_intro.jsp). Multiple alignments of these sequences were performed using ClustalW 1.83 version of EBI (http://www.ebi.ac.uk/cgi-bin/clustalw/) with 0.5 transition weight. Phylogenetic trees were constructed using MEGA 6.0 version (http://www.megasoftware.net) with maximum parsimony algorithm. The maximum parsimony bootstrapping was performed with 1000 replicates with pseudorandom number generators. The nucleotide composition of each aligned sequence was predicted by BioEdit software. The nucleotide sequences were deposited in the genebank with an accession number KR559728.

2.2 Production and quantification of EPS

The bacterium MSI12 was inoculated in 250 ml of Zobell marine broth and incubated at 30 °C for 72 h at 200 rpm on a shaking incubator (Orbitek). After incubation the culture was centrifuged at 15 000g for 20 min at 4 °C. The proteins and nucleic acids were precipitated out with 20% trichloroacetic acid at 4 °C for 30 min. Two volumes of ice cold acetone were added to the supernatant and left for precipitation of the EPS at 4 °C for 24 h. The precipitated EPS was separated using a filter membrane 0.45 μm (Millipore) and dialyzed against deionized water for 24 h. The separated EPS was lyophilized (Yamato) and stored at 30 °C. The biochemical composition of the EPS was determined for carbohydrates,¹⁵ uronic acids¹⁶ and protein.¹⁷

2.3 Anti-Candida activity

The anti-Candida effect of the EPS on C. albicans was investigated in a plate assay. The C. albicans strain was collected from microbial type culture collection (MTCC) with accession number 227. The lyophilised EPS from MSI12 was aliquoted in sterilized distilled water to obtain different concentrations of EPS (5 μg to 20 μg ml⁻¹). The inoculums were prepared from a 48 h culture of C. albicans containing 1 × 10⁶ cells and were uniformly spread over Sabouraud dextrose agar (SDA) plates using a sterile cotton swab (HiMedia). Absorption of excess moisture was allowed to occur for 10 min. Wells with a diameter of 4 or 7 mm were then punched in the swabbed plates and loaded with different concentrations of the EPS aliquot. The drug fluconazole was also used at a concentration of 10–150 mg ml⁻¹ to determine the MIC. The plates were incubated at 37 °C and were observed for zone of inhibition after 48 h.

2.4 Evaluation of the strain MSI12 for probiotic characteristics

2.4.1 Stress tolerance assay. Heat stress tolerance of the isolate MSI12 was determined as per Stack et al.¹⁸ with necessary modifications. The isolate Enterococcus sp. MSI12 was allowed to grow overnight in a 250 ml conical flask at 30 °C with shaking at 200 rpm. Then 1 ml of culture was centrifuged at 5000 × g for 10 min, the supernatant was removed, and the pellet was resuspended in 1 ml of preheated (58 °C) nutrient broth and incubated at 58 °C with agitation for different time intervals (10 min to 1 h). The incubated broth at various time intervals was serially diluted and plated on nutrient agar plates and incubated at 37 °C for 48 h. For the acid stress assay 1 ml of overnight culture of MSI12 was centrifuged at 5000 × g for 10 min and the pellet obtained was resuspended in 2 ml of nutrient broth of pH 2.0 preadjusted with 1 M HCl. The OD_{600 nm} was recorded at 30 min intervals for 5 h. For the salt stress assay the cell pellet was resuspended in nutrient broth containing 5 M NaCl and incubated with agitation at 37 °C. Bile tolerance was determined using the cell pellet resuspended in 2 ml nutrient broth with 1% of bovine bile (Sigma-Aldrich). The OD_{600 nm} was recorded at 30 min intervals for 5 h. For all the above assays, L. plantarum MTCC 2621 was used as control and maintained under similar assay conditions except for the medium (MRS broth). All the assays were performed in triplicates and each value presented is the mean value ± standard deviation.

2.4.2 Survival of MSI12 in a simulated gastric environment. Simulated gastric juice was prepared as per Corcoran et al.¹⁹ The ability of survival of MSI12 in gastric juice was compared with the control L. plantarum. Overnight grown cultures of MSI12 (NB) and L. plantarum (MRS) were centrifuged at 10 000 × g at 4 °C for 10 min. The pellet was washed in an equal volume of cold 0.25 × Ringer’s solution. Pellets were then resuspended in an equal volume of simulated gastric juice and incubated at 37 °C in a shaker incubator (Orbitek) at 150 rpm for 90 min. Samples were taken at frequent time intervals 0, 10, 30, 60, and 90 min and plated on respective media and incubated at 37 °C for 72 h.

2.4.3 Auto aggregation assay. The auto aggregation assay was performed with minor modification of the method described by Del Re et al.²⁰ Briefly, 48 h culture of E. faecalis was centrifuged at 13 000 × g for 5 min at 4 °C. The pellet thus obtained was washed and resuspended in sterile phosphate-buffered saline (PBS, pH 7.2) to obtain an OD₆₀₀ of 0.5. The absorbance was measured at OD₆₀₀ after 1 h of incubation. The percentage of auto aggregation was expressed as: 1 − (A_t/A₀) × 100, where A_t represents the absorbance at 1 h and A₀ was the absorbance at 0 h.

2.5 Hydrophobicity and antioxidant assay

Bacterial adhesion to hydrocarbon was performed as per the procedure described by Rosenberg²¹ and Kiran et al.²² Microbial hydrophobicity was measured in the exponential growth phase of strain MSI12. Phosphate urea magnesium sulfate buffer (PUM) (g l⁻¹): 19.7 K₂HPO₄, 7.26 KH₂PO₄, 1.8 urea and 0.2 MgSO₄·7H₂O was used for the hydrophobicity test. After centrifugation, cells were washed twice with PUM buffer, resuspended in the PUM buffer to get an optical density of ca. 1.0 (A₀). The optical density was measured at 600 nm on a UV-vis Spectrophotometer (PG Instruments, UK). Then, 400 μl of hydrocarbons such as xylene or toluene were added to 2 ml of microbial suspension and vortexed for 2 min. After 10 min, the optical density of the aqueous phase was measured (A₁). The degree of hydrophobicity is calculated as [1 − (A₁/A₀)] × 100%]. The antioxidant activity of EPS was measured by DPPH radical scavenging activity.²³ The percentage of DPPH scavenging ability was calculated as 1 − (A_sample − A_blank)A_control] × 100%. All these assay conditions were performed in triplicate.

2.6 Characterization of EPS

The functional groups of the EPS were determined using Fourier transformed infrared (FTIR) spectroscopy analysis. The dialysed EPS was thoroughly mixed with KBr and dried. The dried sample was subjected to IR spectral analysis using a Fourier Transform Infrared Spectrophotometer (Thermo Nicolet Model: 6700) with spectral range of 4000–400 cm⁻¹. The lyophilized EPS was subjected to gas chromatography (GC) (JEOL GCMATE II GC-MS system) analysis to detect the alditol-acetate derivative as described in Kavita et al.²⁴ The peaks of the GC were subjected to mass-spectral analysis and the spectra were analyzed by using the NIST database (version 2.0). The ¹H NMR spectra was acquired by dissolving the EPS in deuterated water at a concentration of 20 mg ml⁻¹ and analyzed on a Bruker Model: Avance-II at 22 °C.

2.7 Thermal gravimetric (TG) and differential scanning calorimetric (DSC) analysis

Thermal stability and weight loss of EPS was performed and determined using TA instruments, Q600 SDT and DSC. Approximately 7–8 mg of sample was placed in standard 70 μl aluminium pans. The analysis was carried out over the temperature range from 0 °C to 500 °C at a rise in temperature of 10 °C min⁻¹. The flow rate of the gas was 50 ml min⁻¹. The weight loss is recorded as a function of temperature.²⁵

2.8 Antibiofilm activity of EPS

The biofilm disruption ability of lyophilized EPS was determined in a microtitre plate assay. The 48 h old C. albicans culture was inoculated in 96 well polystyrene microtiter plates using a sterilized 96-pin replicator. The well plate was prepared with aliquoted EPS of varying concentration (50–250 μg ml⁻¹) in Sabourauds dextrose (SD) broth. The biofilm formed by C. albicans in the absence of EPS was used as the control. The plate was incubated for 48 h, and then the medium was aspirated. The wells were gently rinsed with phosphate buffer (50 mM, pH 7.0), air-dried, and the biofilms were quantified using the crystal violet assay.^22,25 Fluconazole (50–250 mg ml⁻¹) was used as standard antifungal drug. The data related to these experiments are depicted as average values of triplicate observations and error bars indicate standard deviation. The results are expressed in terms of percent biofilm formed in the presence of EPS compared to untreated wells (indicating 100% biofilm coverage).

2.9 Disruption of pre-formed C. albicans biofilm using the EPS

The C. albicans biofilm was developed in a cover glass immersed in a 50 ml Erlenmeyer flask containing SD broth and incubated at 28 °C for 48 h. To study the biofilm disruption, the mature preformed biofilm was treated with varying concentrations of EPS (50–250 μg ml⁻¹) and fluconazole ranging from 50–250 mg ml⁻¹ and incubated for 24 h. The biofilm developed by the C. albicans in the absence of EPS was used as the control. The exposed cover slip was washed, dried and stained with a 0.4% crystal violet solution (w/v) and observed under a phase-contrast microscope (Optica) at ×40 magnification.

3. Confocal laser scanning microscopy (CLSM) and SEM analysis

To prepare the culture for confocal imaging, the cells were grown in SD broth at 37 °C for 48 h. This fresh culture was used to form biofilms on sterilized glass slides and the pre-formed biofilm was treated for 24 h with EPS (50–250 μg ml⁻¹) and fluconazole of varying concentrations ranging from 50–250 mg ml⁻¹. Untreated biofilms were used as controls and the biofilm coverage thus formed on glass slides was stained with 0.1% acridine orange and subjected to visualization in a CLSM (LSM 710, Carl Zeiss). The 488 nm Ar laser and 500–640 nm band pass emission filter were used to excite and detect the stained cells. Multiple (20) images were scanned and processed using Zen 2009 image software. The effective disrupted concentration of EPS was selected for SEM and confocal biofilm analysis with BacLight Live/Dead stain (Molecular Probes, Eugene). The cell viability was observed and the images were recorded using CLSM (Carl Zeiss).

4. Results

4.1 Identification of probiont MSI12

Isolation of bacteria from seawater was performed on Zobell marine agar and 142 stable isolates were picked for screening. Of the 11 positive isolates (S1), the strain designated as MSI12 produced maximum EPS and was selected for further evaluation. The production of EPS by MSI12 was 580 μg ml⁻¹ after 72 h of incubation at 30 °C. The isolate was characterized as Gram positive, facultative anaerobe, non-motile, catalase negative and oxidase positive. The megaBLAST analysis of the 16S rRNA gene sequence of MSI12 showed a similarity of 98% with strains of Enterococcus faecalis (Fig. 1). Based on the biochemical characteristics and phylogenetic analysis, the isolate MSI12 was identified as Enterococcus faecalis. The RDPII analysis confirmed the isolate had closest match with E. faecalis.

Fig. 1 Phylogenetic tree of Enterococcus faecalis MSI12. Maximum parsimony phylogenetic tree based on 16S rRNA gene sequence of Enterococcus faecalis MSI12 showing representatives of other related taxa. The analysis was performed with bootstrap values of 1000 replications greater than 50% are shown at branching points.

4.2 Probiotic characteristics E. faecalis MSI12

The strain MSI12 was evaluated for probiotic potential along with L. plantarum as reference strains (control). After 10 min exposure at 58 °C, a significant increase in viability was observed for MSI12 over the control strains. The increase in viability of MSI12 was observed at various exposure periods of 20, 30, 40, and 50 min respectively. At 40 min, a 50-fold difference in survival of MSI12 was observed when compared to the control. The control strain was reduced to 4.4 × 10³ CFU ml⁻¹ from an initial biomass of 8.5 × 10⁸ CFU ml⁻¹ whereas the strain MSI12 was reduced to 4 × 10⁷ CFU ml⁻¹ from 2 × 10⁹ CFU ml⁻¹. Even after 60 min exposure to 58 °C the cell viability of MSI12 was significantly higher than the controls.

The isolate MSI12 showed increased acid tolerance over the control strain L. plantarum and survived at pH values as low as 2.0 for 5 h. The results showed the survival rate of E. faecalis was 60% in 1% bile acids when compared to the control L. plantarum (40%) after 5 h. The strain MSI12 showed high tolerance to salt concentration up to 5 M NaCl. Auto aggregation was investigated on the basis of sedimentation characteristics after 1 h. The strain MSI12 showed strong aggregation (81.2%) and was strongly hydrophobic with 61.47% adhesion to xylene, and 60% to toluene (Table 1). The DPPH scavenging activity of EPS from E. faecalis was 39.24 ± 4.25% at a concentration of 200 μg ml⁻¹ which was higher than the standard ascorbic acid (37.54 ± 4.55%) at a concentration of 250 μg ml⁻¹ (Fig. 2a). We also observed that the scavenging activity increases with an increase in EPS concentration. The survival of probiotic strain MSI12 in simulated gastric juice was compared with the control L. plantarum from 0 to 90 min. Decrease in growth was observed in both the strains at 90 min of incubation. Comparatively, the MSI12 has better survival in gastric juice with more than 70% at 90 min of incubation suggesting MSI12 can be a potential probiotic strain (Fig. 2b). The control has a lesser survival of 40% in gastric juice.

Table 1 Results showing the mean value of triplicate experiments on acid, bile, autoaggregation after 5 h of incubation and hydrophobicity assays. The results for MSI12 were compared with the standard probiotic L. plantarum, and all the results indicated the isolate MSI12 has increased resistance than control

Strain	A 600 nm hydrophobicity (%)
	NB (pH 2.0, 5 h)	NB + bile (5 h)	NB + salt	Auto-aggregation (%)	Xylene	Toluene
Lactobacillus plantarum	6.69 ± 0.21	5.74 ± 0.33	6.36 ± 0.31	55.11 ± 0.32	38.22 ± 0.14	20.92 ± 0.43
MSI12	7.42 ± 0.10	6.48 ± 0.23	7.42 ± 0.12	81.20 ± 0.19	61.47 ± 0.09	60.01 ± 0.29

---

Fig. 2 (a) Antioxidant activity of EPS. Ascorbic acid was used as standard in the DPPH scavenging activity of EPS from E. faecalis MSI12. The activity (39.24 ± 4.25%) of EPS (200 μg ml⁻¹) was equal to the standard (37.54 ± 4.55% at a concentration of 250 μg ml⁻¹). (b) Survival rate of MSI12 in the simulated gastric juice. The survival rate of MSI12 in the simulated gastric juice was compared with the control L. plantarum. The analysis showed the survival rate of the MSI12 was better than the control L. plantarum.

4.3 Chemical characterization of EPS

Chemical analysis showed that the purified EPS composed of 76.6% (w/w) total sugar and 12.8% (w/w) total protein and 5% uronic acids indicating that MSI12 derived EPS was mainly composed of polysaccharide. The FTIR spectrum of MSI12-EPS showed characteristic functional groups as shown in Fig. 3. The strong absorption band at 3449 cm⁻¹ corresponds to the presence of hydrogen bonded –OH functional group. The peaks at 2926 and 2962 cm⁻¹ correspond to the different types of C–H stretching frequencies. The weak absorption at 1708 cm⁻¹ showed the presence of carbohydrate containing carbonyl group. The medium absorption at 1657 cm⁻¹ showed the stretching frequency of carbohydrates such as glucose and mannose. The C–C stretching frequency appeared around 1445 cm⁻¹ and C–O stretching frequencies appeared in the range of 1000–1300 cm⁻¹.

Fig. 3 FT-IR spectra of EPS from E. faecalis MSI12. Absorption band at 3448.72 cm⁻¹ corresponds to the presence of hydrogen bonded –OH functional group, peaks at 2926 and 2962 cm⁻¹ correspond to C–H stretching frequencies, carbohydrate containing carbonyl group is shown in absorption at 1708 cm⁻¹.

The well resolved ¹H signal appearing in the range of δ 4.4–5.4 ppm, reveals the presence of carbohydrate anomeric protons. The tertiary carbon hydrogen (R–CH–OR′) was observed as a multiplet at δ 4.3–3.6 ppm. The sharp single peak at δ 3.5 ppm reveals the presence of a methoxy (–OCH₃) functional group. The acid functional group –OH proton and carbon attached to the –OH proton appeared at δ 3.2 ppm and δ 2.8 ppm respectively. The sharp singlet peak at δ 2.1 ppm corresponded to the presence of a carbonyl group attached to CH₃. The alkyl –CH₂– protons appeared in between δ 1.8 and 1.3 ppm as a multiplet. A singlet peak was observed at δ 1.1 ppm, this reveals the presence of a –CH₃ group attached to quaternary carbon (Fig. 4a). GC-MS analysis of the EPS showed the presence of sugars like D-mannose, D-galactose, glucose, D-arabinose with retention times of 5.32, 5.93, 6.61, and 10.67 min respectively. The molar percentage of glucose (47.4%) was highest followed by galactose, mannose and arabinose (Fig. 4b).

Fig. 4 (a) ¹H NMR spectra of MSI12-EPS. The well resolved ¹H signal appearing in the range of δ 4.4–5.4 ppm, reveals the presence of carbohydrate anomeric protons. The tertiary carbon hydrogen (R–CH–OR′) was observed as a multiplet at δ 4.3–3.6 ppm. The sharp singlet peak at δ 3.5 ppm reveals the presence of a methoxy (–OCH₃) functional group. The acid functional group –OH proton and carbon attached to the –OH proton appeared at δ 3.2 ppm and δ 2.8 ppm respectively. The sharp singlet peak at δ 2.1 ppm showed the presence of a carbonyl group attached to CH₃. The alkyl –CH₂– protons appeared in between δ1.8 and 1.3 ppm as a multiplet. A singlet peak was observed at δ 1.1 ppm, this reveals the presence of a –CH₃ group attached to a quaternary carbon. (b) GC-MS spectra of the EPS. The spectra show the presence of sugars like D-mannose, D-galactose, glucose, D-arabinose with retention times of 5.32, 5.93, 6.61, and 10.67 min respectively.

4.4 Thermal stability of EPS

Degradation of EPS took place in two steps (S2). In the first step, a weight loss of (19.24%) was observed at 110 °C due to the loss of moisture molecules in the polymer. In the second step, depolymerisation occurred up to 240 °C and a weight loss of about 27.48% was observed. The thermal transition of the EPS was studied by differential scanning calorimetric analysis. The thermogram showed an exothermic peak of EPS with a crystallization temperature (T_c) of 83.06 °C accompanied by 196.4 J g⁻¹ latent energy. The melting transitions started at 253.27 °C (ESI†).

4.5 Antifungal and antibiofilm activity of EPS

The EPS of E. faecalis MSI12 extracted after 72 h of incubation showed a significant inhibition pattern on C. albicans MTCC226 when compared to the control. The crude and the purified EPS showed concentration dependent activity (Fig. 5). The C. albicans was more susceptible to the crude EPS at a MIC of 50 μg ml⁻¹ and purified EPS at a MIC of 10 μg ml⁻¹. No growth inhibition of C. albicans was observed at a micromolar concentration of fluconazole (150 μg ml⁻¹). But on raising the fluconazole concentration from micromolar to millimolar, growth inhibition was noticed at 100 mg ml⁻¹. The effect of EPS and fluconazole on biofilm formation of C. albicans was evaluated using microtitre plate assay and phase contrast microscopy observations. The control biofilms showed 100% biofilm coverage. In the presence of EPS (50 μg) a significant decrease (78%) in biofilm formation by C. albicans was observed when compared to fluconazole (50 mg) where the biofilm coverage was shown to decrease to 43%. A 98% reduction of biofilm was achieved with EPS at a concentration of 250 μg whereas the reference drug fluconazole (250 mg) was effective on an 83% reduction of biofilm coverage (Fig. 6). Phase contrast images (Fig. 7) showed 200 and 250 μg of EPS were effective in the disruption of C. albicans biofilm when compared to fluconazole. Comparatively the EPS was found to be effective in reducing the survival/biofilm of C. albicans and it also proves that the drug fluconazole was not effective in inhibiting the biofilm of C. albicans.

Fig. 5 Antifungal activity of MSI12-EPS on C. albicans. The well diffusion assay was performed using crude and purified EPS. A is 5 μg ml⁻¹ of crude EPS, B is 25 μg ml⁻¹ of crude EPS, C is 50 μg ml⁻¹ of crude EPS, D is 5 μg ml⁻¹ of purified EPS and E is 10 μg ml⁻¹ of purified EPS.

Fig. 6 Antibiofilm potential of MSI12-EPS. Antibiofilm activity of MSI12–EPS on C. albicans biofilm was assayed in a 96-microtitre plate with varying concentrations of EPS (μg ml⁻¹) and flucanazole (mg ml⁻¹). Each concentration was tested in triplicate. The reduction of biofilm was significantly high (98%) with 250 μg of EPS whereas the reference drug fluconazole (250 mg) showed reduction of biofilm to 83%.

Fig. 7 Phase contrast micrographs showing biofilm disruption potential of MSI12-EPS on C. albicans. The C. albicans biofilm was developed on a cover glass and then it was treated with varying concentrations of EPS 50–250 μg and fluconazole ranging from 50–250 mg. The treated cover glass was stained with crystal violet and observed under a phase-contrast microscope (Optica) at ×40 magnification. A – control biofilm, B1 – 50 mg fluconazole, B2 – 50 μg MSI12-EPS, C1 – 100 mg fluconazole, C2 – 100 μg MSI12-EPS, D1 – 150 mg fluconazole, D2 – 150 μg MSI12-EPS, E1 – 200 mg fluconazole, E2 – 200 μg MSI12-EPS and F1 – 250 mg fluconazole, F2 – 250 μg MSI12-EPS.

4.6 CLSM observations

Biofilms of C. albicans formed on glass slides were treated with EPS of 50 μg to 250 μg and fluconazole of varying concentrations ranging from 50 to 250 mg. The image analysis (Fig. 8) lucidly visualized that the EPS was potentially more effective on the disruption of C. albicans biofilm than the reference drug. The effect of various dilutions of EPS on the biofilm disruption potential showed that it is concentration dependent. Data revealed that at a lower concentration of EPS (50 μg) biofilm disruption was not significant when compared to 100 μg and above concentration. SEM images (Fig. 9) showed the representative image of the control biofilm and Fig. 8b depicts the biofilm disruption potential of EPS (150 μg) on C. albicans. The Bac/light staining method showed live and dead cells of C. albicans treated with EPS. The CLSM images showed the cells were stained in red colour indicating dead cells of C. albicans treated with 150 μg EPS whereas control live cells were stained green (Fig. 10). The live–dead differential staining procedure proves the potential effect of MSI12-EPS on the disruption/control of biofilm forming Candida infections.

Fig. 8 Confocal laser scanning micrographs showing biofilm disruption potential of MSI12-EPS on C. albicans. The pre-formed biofilm was treated for 24 h with EPS and fluconazole of varying concentrations ranging from 50–250 μg. Untreated biofilms were used as controls and the biofilm coverage thus formed on glass slides were stained with 0.1% acridine orange and subjected to visualization in a CLSM (LSM 710, Carl Zeiss). A – control biofilm, B1 – 50 mg fluconazole, B2 – 50 μg MSI12-EPS, C1 – 100 mg fluconazole, C2 – 100 μg MSI12-EPS, D1 – 150 mg fluconazole, D2 – 150 μg MSI12-EPS, E1 – 200 mg fluconazole, E2 – 200 μg MSI12-EPS and F1 – 250 mg fluconazole, F2 – 250 μg MSI12-EPS.

Fig. 9 Scanning electron microscope images showing antibiofilm potential of MSI12-EPS on C. albicans. (A) Control biofilm and (B) disrupted by EPS.

Fig. 10 CLSM analysis with BacLight Live/Dead stain. A. Control biofilms of C. albicans. B. EPS treated biofilms stained using Bac/light kit. Based on this analysis the red colour staining shows the EPS effectively kills the C. albicans cells.

5. Discussion

The use of probiotics in human health care is now emerging as a proven microbial therapeutic and/or as functional foods. The health benefits of probiotics are well-established including potential antagonists of infectious diarrhea,²⁶ immune stimulation,²⁷ treatment of inflammatory bowel diseases, reduction in allergic responses^28,29 and lowering serum cholesterol.¹⁸ The literature shows that probiotics are an effective method of treating diarrheal infection in particular and all types of bacterial infection in general. The existing probiotic-based anti-infective treatments are reactive and their potential for the treatment of established pathogens has been less explored. This investigation should impart a new dimension on the treatment of established pathogens like biofilms. Based on the present findings the potential implications of probiotics could be envisaged as preventive/control method to treat biofilm formation in implant medical devices and/or alternatively as synergistic to the existing antibiotics to disrupt Candida biofilms. The increasing number of indwelling medical implants are facing a potential threat due to device-associated biofilm infections caused by Candida spp.³⁰ Management of device-associated Candida infections remains challenging. In this context the present findings are promising to develop an effective probiotic strategy to maintain the implants free from Candida biofilms.

In principle, a probiotic strain can colonize, aggregate and adhere to the host. The stability and longevity of probiotic establishment depends on their tolerance potential in the host. In this study, we established that the probiotic strain E. faecalis MSI12 could grow in and tolerate the niche conditions like those found in the human gut. Most of the prominent probiotic LAB strains are heat tolerant, but their viability at 60 °C and above was a constraint to their utilization as technologically feasible. The isolate E. feacalis MSI12 survived and showed high tolerance to acid even after 5 h of incubation at pH 2.0. An approximate concentration of 0.3–0.5% bile level is found in the human gastrointestinal tract. The strain E. feacalis MSI12 showed 60% survival on treatment with 1% bovine bile. The EPS was used as a shield and adhesive by the producing microorganisms to protect the cells against heat and other stress. In this study, the EPS produced from E. faecalis MSI12 exhibited high antioxidant activity, increased auto-aggregation (81.2%) and hydrophobicity which evidences that the isolate MSI12 is a potential probiotic strain. The isolate MSI12 exhibited a strong auto-aggregation of 81.2% in 1 h and the percentage of aggregation was higher when compared to Lactobacillus 63.1% after 5 h as reported by Anwar et al.³¹ Del Re et al.²⁰ reported auto-aggregation as an important factor in the adhesion of bacterial cells to the intestinal epithelium and its maintenance in the gastrointestinal tract. To improve their heat tolerance, the LAB was treated or formulated with prebiotics. For example gum acacia treatment offered protection to Lactobacillus paracasei NFBC 338 during heat, bile, and H₂O₂ stress.³⁰ Dixit et al.³² examined 3 strains of L. acidophilus and found that the survival of strain NCIM 2285 was reduced by 50% and the strain NCIM 2903 by 21% after 4 h of incubation at pH 2.5. A previous report evidences the use of E. faecium and E. faecalis strains as probiotics and are ingested in high numbers to treat diarrhoea, antibiotic-associated diarrhoea or irritable bowel syndrome, to lower cholesterol levels or to improve host immunity.³³ The MSI12 was established as potential probiotic strain compared to L. plantarum. The cell viability was very high at higher temperature, acidic pH, bile salt, survival in simulated gastric juice and salt concentration and therefore the strain MSI12 can survive the gut like niche without prebiotics. The tolerance at higher temperature, acidic pH, bile salt, gastric juice and salt concentration are the prerequisite characteristics of a probiotic strain. The overall tolerance potential envisages that the strain MSI12 can colonize on any surfaces like gastrointestinal tract, vaginal, dental and mucosal surfaces.

A possible mechanism of destabilizing the EPS of biofilms includes breaking the bonds that connect polysaccharide residues in the EPS. This could be an effective means of dispersing biofilms and making the subjacent bacteria more susceptible to treatment by antibiotics.³⁴ Enterococci are known for secretion of bacteriocins³⁵ and therefore the EPS of E. faecalis MSI12 was expected to be synergized with antimicrobial secretions. This study revealed that biofilm disruption/control using a probiont could deliver a synergistic approach as the probiotic strain can colonize in the host to prevent the formation of Candida biofilms. The EPS from Enterococcus sp. MSI12 showed significant reduction of treated Candida biofilm. The antibiofilm potential of EPS was very stronger than the standard antifungal drug fluconazole. The strain C. albicans MTCC227 used in this study is resistant to fluconazole. Previously it was reported that even 3000 μg ml⁻¹ of fluconazole was not effective in inhibiting the growth of C. albicans MTCC 227.³⁶ The EPS produced by the probiotic E. faecalis MSI12 showed 100% disruption of C. albicans biofilm when compared with the drug flucanazole (80%). The literature shows that fluconazole is still used as drug in patients with candidemia or suspected invasive candidiasis.³⁷ Fluconazole has excellent in vitro activity against C. albicans. Fluconazole can also be effective against non-albicans Candida species, including Candida parapsilosis, Candida tropicalis and Candida glabrata, although higher doses may be required.^38,39 Usage of fluconazole has side effects and may cause gastrointestinal tract disorders like nausea, abdominal discomfort, vomiting, diarrhea. The exopolysaccharides released by the biofilm of E. faecalis act as an effective antifungal as well as biofilm inhibitory agent. This study is the first report on E. faecalis probiotics from a marine environment as effective antifungal/antibiofilm agent.

The production of EPS is beneficial to human health as well as EPS has been exploited in the food industry as viscosifiers, stabilizers, emulsifiers, or gelling agents.¹⁸ Based on the available reports probiotic mediated disruption of Candida biofilms is new and this report illustrates that MSI12-EPS is an effective antibiofilm agent to contain Candida infections. Thus using a probiotic strain which secretes EPS to prevent/control Candida biofilm is a novel, effective and safe method of developing an antibiofilm therapy. This study demonstrates a new safe way of Candida biofilm control. The EPS from Oceanobacillus iheyensis was effective on the disruption of biofilm formed by S. aureus.⁴⁰ Biosurfactant and rhamnolipids were shown to be effective for the disruption of pathogenic biofilms of Candida.^22,41,42 Carbohydrates consumed in the diet are the primary and preferred nutrient sources for Candida albicans^43–45 Based on the present findings E. faecalis is a probiont which can establish an EPS assisted biofilm which in turn is a potential antagonist to the Candida cells as well as an inhibitor to the formation of biofilm and/or disruption of established biofilm during Candida infections, thereby innovative approaches to contain Candida infection were demonstrated.

Acknowledgements

This work was funded by Department of Biotechnology, Ministry of Science and Technology. JS and GSK are the Investigators of DBT scheme on Metagenomic exploration PHB. We acknowledge language editing support by Dr Valanarasu. The CLSM facility provided Dr N. Thajuddin, Department of Microbiology, Bharathidasan University is thankfully acknowledged.

References

1. C. Wang, M. Li, D. Dong, J. Wang, J. Ren, M. Otto and Q. Gao, Microbes Infect., 2007, 9, 1376–1383.
2. N. Mishra, S. Ali and P. K. Shukla, Braz. J. Infect. Dis., 2014, 18, 287–293.
3. J. Chandra, P. K. Mukherjee, S. D. Leidich, F. F. Faddoul, L. L. Hoyer, L. J. Douglas and M. A. Ghannoum, J. Dent. Res., 2001, 80, 903–908.
4. R. Cebrián, A. Baños, E. Valdivi, M. Pérez Pulido, M. Bueno and M. Maqueda, Food Microbiol., 2012, 1–9.
5. A. A. Fattani and L. J. Douglas, J. Med. Microbiol., 2006, 55, 999–1008.
6. C. Franz, M. Huch, A. Hikmate, W. H. Holzapfel and A. Gálvez, Int. J. Food Microbiol., 2011, 151, 125–140.
7. G. Bellomo, A. Mangiagle, L. Nicastro and G. A. Curr, Curr. Ther. Res., 1980, 28, 927–936.
8. S. Kiran, A. N. Lipton, S. Priyadharshini, K. Anitha, L. E. Cruz, M. V. Arasu, K. C. Choi, J. Selvin and N. A. Al Dhabi, Microb. Cell Fact., 2014, 13, 114.
9. A. Mancuso Nichols, J. Guezennec and J. P. Bowman, Mar. Biotechnol., 2005, 7, 253–271.
10. B. Ismailand and K. M. Nampoothiri, Arch. Microbiol., 2010, 192, 1049–1057.
11. P. Kanmani, K. Suganya, N. Yuvaraj, V. Pattukumar, K. A. Paari and V. Arul, Appl. Biochem. Biotechnol., 2013, 169, 1001–1015.
12. G. Sathyanarayanan, G. S. Kiran and S. Joseph, Colloids Surf., B, 2013, 102, 13–20.
13. U. Romling, W. D. Sierralta, K. Eriksson and S. Normark, Mol. Microbiol., 1998, 28, 249–264.
14. J. Enticknap, M. KellyShanks, O. Peraud and R. T. Hill, Appl. Environ. Microbiol., 2006, 72, 4105–4119.
15. M. Dubois, K. A. Gilles, J. K. Hamilton, P. A. Rebers and F. Smith, Anal. Chem., 1956, 28, 350.
16. A. Knutson and A. Jeanes, Anal. Biochem., 1968, 24, 482–490.
17. H. Lowry, N. J. Rosebrough, A. L. Farr and R. J. Randall, J. Biol. Chem., 1951, 193, 265–275.
18. M. Stack, N. Kearney, C. Stanton, G. F. Fitzgerald and R. P. Ross, Appl. Environ. Microbiol., 2010, 76, 500–507.
19. M. Corcoran, C. Stanton, G. F. Fitzgerald and R. P. Ross, Microbiology, 2007, 153, 291–299.
20. B. Del Re, B. Sgorbati, M. Miglioli and D. Palenzona, Lett. Appl. Microbiol., 2000, 31, 438–442.
21. Rosenberg, FEMS Microbiol. Lett., 2006, 22, 289–295.
22. S. Kiran, B. Sabarathnam and J. Selvin, FEMS Immunol. Med. Microbiol., 2010, 59, 432–438.
23. H. Kao and B. H. Chen, J. Agric. Food Chem., 2006, 54, 7544–7555.
24. K. Kavita, A. Mishra and B. Jha, Carbohydr. Polym., 2013, 94, 882–888.
25. S. Kiran, A. N. Lipton, J. Kennedy, A. Dobson and J. Selvin, Bioengineered, 2014, 5, 305–318.
26. T. Arvola, K. Laiho, S. Torkkeli, H. Mykkanen, S. Salminen, L. Maunula and E. Isolauri, Pediatrics, 1999, 104(5), e64.
27. E. Isolauri, Curr. Allergy Asthma Rep., 2004, 4, 270–275.
28. M. Kalliomaki, P. Kirjavainen, E. Eerola, P. Kero, S. Salminen and E. Isolauri, J. Allergy Clin. Immunol., 2001, 107, 129–134.
29. K. Kukkonen, E. Savilahti, T. Haahtela, K. Juntunen-Backman, R. Korpela, T. Poussa, T. Tuure and M. Kuitenen, J. Allergy Clin. Immunol., 2007, 119, 192–198.
30. M. Kojic and R. O. Darouiche, Clin. Microbiol. Rev., 2004, 17, 255–267.
31. A. Anwar, A. A. Thikra and A. M. Saeed, Br. Microbiol. Res. J., 2014, 4(4), 381–391.
32. G. Dixit, D. Samarth, V. Tale and R. B. Eurasia, J. Biosci., 2013, 7, 1–9.
33. C. Franz, M. Huch, A. Hikmate, W. H. Holzapfel and A. Gálvez, Int. J. Food Microbiol., 2011, 151, 125–140.
34. M. Bales, E. M. Renke, S. L. May, Y. Shen and D. C. Nelson, PLoS One, 2013, 8, e67950.
35. E. Balla and L. M. Dicks, Int. J. Food Microbiol., 2005, 99, 33–45.
36. N. Devkatte, G. B. Zore and S. M. Karuppayil, FEMS Yeast Res., 2005, 5, 867–873.
37. E. Ha, K. R. Peck, E. J. Joo, S. W. Kim, S. I. Jung, H. H. Chang, K. H. Park and S. H. Han, Antimicrob. Agents Chemother., 2012, 56, 3950–3956.
38. P. Arévalo, A. Arias, A. Andreu, C. Rodríguez and A. Sierra, J. Chemother., 1994, 6, 226–229.
39. W. van’t Wout, Eur. J. Clin. Microbiol. Infect. Dis., 1996, 15, 238–242.
40. K. Kavita, V. K. Singh, A. Mishra and B. Jha, Carbohydr. Polym., 2014, 101, 29–35.
41. L. Fracchia, M. Cavallo, G. Allegrone and M. G. Martinotti, Current research technology and education topics in applied microbiology and microbial technology, 2010, pp. 827–837.
42. N. Singh, S. C. Pemmaraju, P. A. Pruthi, S. S. Cameotra and V. Pruthi, Appl. Biochem. Biotechnol., 2013, 169, 2374–2391.
43. C. J. Seneviratne, T. Zhang, H. H. Fang, L. J. Jin and L. P. Samaranayake, Mycopathologia, 2009, 167, 325–331.
44. P. Samaranayake, J. Oral Pathol., 1986, 15, 61–65.
45. Y. Jin, L. P. Samaranayake, Y. Samaranayake and H. K. Yip, Arch. Oral Biol., 2004, 49, 789–798.

---

Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c5ra10302a

---