Probiotics & Antimicro. Prot. (2015) 7:181–192 DOI 10.1007/s12602-015-9195-5
Screening, Characterization and In Vitro Evaluation of Probiotic Properties Among Lactic Acid Bacteria Through Comparative Analysis Sundru Manjulata Devi1 • Ann Catherine Archer1 • Prakash M. Halami1
Published online: 7 June 2015 Ó Springer Science+Business Media New York 2015
Abstract The present work aimed to identify probiotic bacteria from healthy human infant faecal and dairy samples. Subsequently, an assay was developed to evaluate the probiotic properties using comparative genetic approach for marker genes involved in adhesion to the intestinal epithelial layer. Several in vitro properties including tolerance to biological barriers (such as acid and bile), antimicrobial spectrum, resistance to simulated digestive fluids and cellular hydrophobicity were assessed. The potential probiotic cultures were rapidly characterized by morphological, physiological and molecular-based methods [such as RFLP, ITS, RAPD and (GTG)5]. Further analysis by 16S rDNA sequencing revealed that the selected isolates belong to Lactobacillus, Pediococcus and Enterococcus species. Two cultures of non-lactic, nonpathogenic Staphylococcus spp. were also isolated. The native isolates were able to survive under acidic, bile and simulated intestinal conditions. In addition, these cultures inhibited the growth of tested bacterial pathogens. Further, no correlation was observed between hydrophobicity and adhesion ability. Sequencing of probiotic marker genes such as bile salt hydrolase (bsh), fibronectin-binding protein (fbp) and mucin-binding protein (mub) for selected isolates revealed nucleotide variation. The probiotic binding domains were detected by several bioinformatic tools.
Electronic supplementary material The online version of this article (doi:10.1007/s12602-015-9195-5) contains supplementary material, which is available to authorized users. & Prakash M. Halami
[email protected] 1
Microbiology and Fermentation Technology Department, CSIR- Central Food Technological Research Institute, Mysore 570020, India
The approach used in the study enabled the identification of potential probiotic domains responsible for adhesion of bacteria to intestinal epithelial layer, which may further assist in screening of novel probiotic bacteria. The rapid detection of binding domains will help in revealing the beneficial properties of the probiotic cultures. Further, studies will be performed to develop a novel probiotic product which will contribute in food and feed industry. Keywords Probiotics Lactic acid bacteria Binding proteins Bile salt hydrolase Phylogeny Genetic comparison
Introduction Probiotics are live microorganisms, when administered in adequate amounts confer several health benefits on the host which include reduction in lactose intolerance, prevention of colon cancer, inflammatory bowel disease, antimutagenic, anticarcinogenic, reduction in allergies, cholesterol and blood pressure [1]. Among the studied probiotic microbiota, lactic acid bacteria (LAB) are the most predominant, widely distributed and common inhabitants of epithelial surface of humans [2]. Several genera of LAB are widely used in fermented foods that are of traditional and industrial importance. In the development of functional foods, LAB have gained more attention due to their application as probiotics, nutraceuticals, etc. [3]. The selection of a probiotic culture plays a crucial role, as these bacteria must survive and colonize the gastro intestinal tract (GIT) to confer the functional properties and health benefits [4]. The taxonomical identification of LAB is very important because of their advantages towards the improvement of
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health and nutritional status of humans [5]. Traditionally, most of the LAB were characterized based upon their morphological, physiological and carbohydrate fermentation patterns [6]. However, results obtained through the conventional methods may be inappropriate due to their low reproducibility and discriminatory power [7]. For more appropriate and sub-species level identification, several molecular typing techniques are employed for easy interpretation, discrimination, reproducibility and fast generation of data [8]. In the present study, we have utilized the techniques such as random amplified polymorphic DNA (RAPD), restriction fragment length polymorphism (RFLP), 16S-23S rDNA intergenic transcribed spacer (ITS) regions, repetitive PCR with (GTG)5 followed by sequencing of 16S rDNA gene for the strain level identification of the selected probiotic LAB. Most of the LAB used as probiotic bacteria are found to have an additional advantage in resetting a disturbed microbiota of human GIT to its normal beneficial composition [9]. In addition, probiotic bacteria can colonize the intestinal mucus layer, where they are found to affect the immune system, inhibit colonization of enteric pathogens and play a major role in cell signalling [9]. The ability to tolerate acid and bile conditions, adherence to intestinal surfaces, antimicrobial activity against pathogens and technological properties are considered as main criteria in the selection of probiotic bacteria [10]. Most of the Lactobacillus sp. are sub-dominant microbiota of intestinal ecosystem and are found to provoke different responses on host epithelium depending on the interaction with a strain [11]. Based on the environmental changes, several of the lactobacilli species are found to change drastically in their cell surface architecture, which impart to their strainspecific characteristics [12]. Cell adhesion to intestinal epithelial layer is another important criterion for a probiotic microorganism, and several in vitro and in vivo studies are performed [13]. However, several difficulties have been experienced for studying the adhesion properties under in vivo models [14]. To overcome these difficulties, many reports suggest utilizing the physical and chemical interaction assays (such as to hydrocarbons) that would reveal the tentative hydrophobic cell adhesion properties [15]. However, there is a need to validate the binding capabilities of lactobacilli or other probiotic LAB through PCR-based methods or by phylogenetic profiling and/or in silico comparative analysis. A study on functional characterization of a potential probiotic marker gene, especially mucin-binding protein (mub) and fibronectin-binding protein (fbp), in different species of LAB will help in developing a potent culture, which can be used in food fermentation to improve the human health [16]. Hence, there is a need to understand the
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genetic systems responsible for intestinal adhesion of probiotic bacteria by rapid detection methods. Sequencing of the probiotic marker genes such as bile salt hydrolase (bsh), mub, fbp and comparative genetic analysis will provide a platform for several unresolved functional analysis of genes or proteins. Moreover, very limited reports are available on such study. Further, our analysis helps in rapid detection which is non-laborious and cost-effective. Hence, in the present study we have evaluated the cell adhesion properties by chemical interaction to hydrocarbons as well as genetic level detection of binding proteins such as mub and fbp among different species of LAB. This method may provide an evidence to the adherence of probiotic bacteria to intestinal cells. Our aim to investigate the probiotic marker genes among selected LAB will provide a platform for the selection of a novel strain for food industry in general.
Materials and Methods Chemicals for Biochemical and Molecular Biology Sodium Chloride (NaCl), potassium chloride (KCl), sodium bicarbonate (NaHCO3), sodium hydroxide (NaOH) and Tris–HCl were procured from ICN Biomedical, USA. Glycerol, gelatin, calcium chloride (CaCl2), pepsin and pancreatin were obtained from SRL, Mumbai, India. Lysozyme, proteinase K, restriction enzymes (HaeIII, AluI and HindIII), dNTP mix, 10 Kb DNA molecular ladder, 3 Kb DNA molecular marker, were purchased from Merck, Bangalore, India. PCR product purification kit, gel extraction of PCR product kit, Taq DNA polymerase, primers, agarose, ethidium bromide (EtBr) and taurodeoxycholic acid were obtained from Sigma-Aldrich, USA. Bacteriological media such as MRS (de Man, Ragosa and Sharpe) broth, BHI (Brian heart infusion) broth, nutrient broth, agar-agar, Mueller– Hinton agar, Todd-Hewitt agar, Gram staining kit, potassium tellurite, antibiotic octadisc (Combi I, G-VIII plus, Combi 69, G-XI plus, Combi 77), discs for carbohydrate fermentation test (adonitol, arabinose, cellobiose, dextrose, dulcitol, galactose, fructose, inositol, inulin, lactose, maltose, mannitol, mannose, melibiose, raffinose, rhamnose, salicin, sorbitol, sucrose, trehalose, xylose), xylene, n-hexadecane, pepsin, pancreatin, Ox bile and bovine bile salt were procured from HiMedia, Mumbai, India. Collection of Samples and Storage Healthy human infant faecal samples were obtained from JSS Medical Hospital, Mysore, Karnataka, India. The samples were collected in sterile containers and stored on ice until conveyance to the laboratory. One gram of sample
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was weighed and used for further studies. Different faecal samples were screened for isolation of probiotic LAB. Simultaneously, several dairy samples such as milk, curd, paneer and buttermilk from a local market of Mysore, India, were procured. Isolation of Bacteria and Culture Maintenance One gram of solid (human faeces and paneer) homogenized sample and 1 ml of liquid (dairy) sample were mixed with 9 ml of sterile saline (0.85 % NaCl). Subsequently, the samples were serially diluted, and appropriate dilutions were pour plated on MRS agar with pH 3 and 0.3 % bile. Later the plates were incubated at 37 °C for 24 h in an anaerobic CO2 incubator (Thermo Scientific, USA). Single isolated colonies were picked up and purified by repeated streaking. The selected colonies were maintained in MRS soft agar stabs (0.8 % agar) at 4 °C. Bacterial Cultures and Growth Conditions Bacteriocin-producing LAB including Pediococcus acidilactici NCIM 5424 and Enterococcus faecium NCIM 5423 were obtained from CSIR-CFTRI culture collection centre, Mysore, were grown in MRS broth at 37 °C under static condition and were used as controls [17]. Food-borne pathogenic bacteria such as Listeria monocytogenes ScottA (obtained from Dr. AK Bhunia, Purdue University, USA), Staphylococcus aureus FRI1722, S. aureus 17A, Escherichia coli MTCC118 (from Dr. E. Notermans, National Institute of Public health, Netherlands) were used as indicator strains and grown in BHI broth at 37 °C under shaking condition (200 rpm) for 4–6 h. The above-mentioned cultures were maintained at -40 °C in MRS and BHI media with 40 % glycerol (v/v). Before being used, the cultures were propagated twice in their respective broths. Acid, Bile Tolerance Assay and Bile Salt Hydrolase (BSH) Activity The resistance to acid (pH 2 and 3) and bile salts (0.1, 0.3 and 0.5 % w/v of bovine salt) for the selected isolates was performed as described earlier [22]. After inoculation, the cells were withdrawn at 0, 1 and 3 h, and the bacterial count was enumerated as described above. The selected isolates were screened for BSH activity as described by Sieladie et al. [20] and observed for white precipitate or zone of halo precipitation around the colonies. Preliminary Characterization of Probiotic Bacteria The physiological and biochemical characterization of LAB isolates was carried out according to Bergey’s
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Manual of Systematic Bacteriology [18]. The carbohydrate fermentation pattern was assessed by using HiMedia sugar discs with 20 mg/ml concentration (cellobiose, lactose, dextrose, adonitol, inulin, trehalose, melibiose, galactose, inositol, fructose, salicin, sorbitol, sucrose, xylose, rhamnose, mannitol, arabinose, mannose, maltose, raffinose, dulcitol). The safety evaluation of LAB was performed by microbiological detection of haemolytic, gelatinase and lecithinase activity on nutrient agar plates supplemented with 7 % defibrinated sheep blood, Todd-Hewitt agar containing 3 % of gelatin and nutrient agar with 5 % egg yolk emulsion as well as with 1 % potassium tellurite, respectively, as described earlier [19–21]. Staphylococcus aureus 17A was used as a positive control for safety evaluation assays. The isolates which were non-haemolytic, non-gelatinase and non-lecithinase were considered for further studies. Antibiotic susceptibility test was performed by using antibiotic octadiscs on Mueller–Hinton agar as described earlier [20]. The Combi I, G-VIII plus, Combi 69, G-XI plus and Combi 77 antibiotic octadiscs were used in the present study. Antimicrobial Activity of Probiotic Bacteria The neutralized cell-free supernatants of the selected isolates were tested against the indicator organisms (L. monocytogenes ScottA, S. aureus FRI1722 and E. coli MTCC 118) as described by Devi and Halami [17]. Clear inhibitory zones [1 mm or more were considered as positive inhibition. Resistance to Simulated Gastric and Intestinal Fluids The resistance of bacterial cells to the simulated gastric fluid (NaCl—125 mM, KCl—7 mM, NaHCO3—45 mM and pepsin 0.3 % w/v at pH 2.5) and intestinal fluid (pancreatin—0.1 % w/v, bovine bile salt—0.15 % w/v at pH 8.0) was performed as described previously by Grimoud et al. [23]. Bacterial Adhesion to Hydrocarbons (Cellular Hydrophobicity) and Auto-Aggregation Assay The bacterial adhesion to hydrocarbons such as xylene and n-hexadecane and cell aggregation assay was determined as described by Kaushik et al. [24]. Briefly, the cell pellet was collected from freshly grown bacterial cultures in MRS broth and the pellet was washed twice with phosphate urea magnesium (PUM) buffer. The cell pellet was resuspended in PUM buffer, and the absorbance was adjusted to 0.7 OD at 600 nm. The probiotic bacterial cell suspension (3 ml) and hydrocarbons (xylene or n-hexadecane) (1 ml)
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were mixed by vortexing and incubated at 37 °C for 10 min. Once again the mixture was vortexed and incubated for 1 h at 37 °C for phase separation. The absorbance of aqueous phase was measured at 600 nm. The bacterial adhesion percentage was calculated as described by Kaushik et al. [24]. Similarly, auto-aggregation assay was performed by collecting the cell pellet as described above and the cell pellet was washed twice in phosphate buffered saline (PBS) buffer. The cell suspension was resuspended in PBS to obtain an absorbance of 0.5 at 600 nm. The cell suspension was dissolved in equal volume of broth and allowed to stand at 37 °C for 2 h. The upper suspension (1 ml) was taken, and the absorbance was recorded by using broth as control. The percentage of auto-aggregation was calculated as described earlier [24].
primers were designed by comparing the sequences retrieved from genomic and protein database (NCBI) using BLASTn, BLASTp and BLASTx algorithms [28]. The primers were designed by considering the conserved consensus sequences by using Primer3 Input (version 0.4.0) (http://frodo.wi.mit.edu/primer3). The conserved protein domains were detected using Pfam database (http://pfam. wustl.edu) with hidden Markov model used as a default parameter. The neighbour-joining phylogenetic dendrogram was constructed by using the analysed nucleotide sequences with MEGA 5.1 version [29] from 1000 bootstrap values by Kimura 2—parameter model. Simultaneously, RAPD and (GTG)5 scores were analysed, and the neighbour-joining phylogenetic tree was constructed by using 1000 bootstrap values with p-distance model.
Molecular Characterization of Probiotic Bacteria
Statistical Analysis
DNA was isolated from the selected LAB according to the procedure described by Mora et al. [25]. Molecular characterization of the probiotic LAB was performed by using typing methods such as restriction fragment length polymorphism (RFLP) of 16S rDNA, random amplified polymorphic DNA (RAPD) PCR with M13 primer, 16S-23S rDNA intergenic transcribed spacer (ITS) regions and repPCR with (GTG)5 primer as described previously [6, 17, 26]. All primers were procured from Sigma-Aldrich (Bengaluru, India), and supplementary Table 1 enlists the primers used in the present study. The taxonomical identification of the selected cultures was performed by sequencing of the purified 16S rDNA gene product as described earlier [17].
Statistical analysis for acid and bile tolerance, hydrophobicity and auto-aggregation was performed by one-way analysis of variance (ANOVA) and correlational analysis by using Microsoft Excel [30]. All the data are represented as mean ± standard deviation (SD). Significance for all the experiments was set at P \ 0.05.
Confirmation of Probiotic-Associated Genes The probiotic marker genes such as bile salt hydrolase (bsh), fibronectin-binding protein (fbp) and mucin-binding protein (mub) were targeted to determine their probiotic binding abilities. Each 25 ll of PCR mixture contained 1X concentration of PCR buffer, 3 mM of MgCl2, 200 lM of dNTP mix, 0.5 pm of each primer and 0.2 U of Taq DNA polymerase. The PCR products were separated on an agarose gel prestained with ethidium bromide. When amplicon was obtained for different species, the corresponding PCR product was sequenced at the Vimta Labs sequencing facility (Hyderabad) for further confirmation. Computational Analysis Multiple sequence alignment was performed by using ClustalW program [27] as well as GeneDoc software (http://www.psc.edu/biomed/genedoc). To detect the presence of probiotic marker genes (bsh, fbp and mub), the
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Nucleotide Sequence Accession Numbers The partial nucleotide sequences of the 16S rDNA amplicons have been deposited at the GenBank database under the accession numbers KM052201 to KM052209. The GenBank accession numbers for the sequences of other probiotic marker genes such as bsh and fbp are KM052210 to KM052215, respectively. The accession number obtained for mub gene Lactobacillus mucosae CM21 is KP238206. The partial sequence of mub gene of Lactobacillus plantarum LP has been submitted to DDBJ database under the accession number LC014928.
Results Screening of Native Probiotic Bacteria Among the obtained 71 isolates, 44 isolates survived at tested pH and bile concentrations. In addition to the different sources used, the isolates from human source showed maximum tolerance to acid and bile stress. Among the 44 acid and bile tolerant cultures, only 11 isolates showed negative reaction for haemolytic and gelatinase activity when compared to S. aureus 17A (a positive control). These 11 isolates were further tested for lecithinase activity where all the cultures were found to be nonpathogenic except the isolate F10a and 288. Further, the
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highest resistance to acid and bile was observed for the isolates P1, CM21, F14a and F2a. Table 1 highlights acid and bile tolerance results for the selected isolates. On studying their antibiotic sensitivity to 25 different antibiotics, the isolates F4C and E2 were found to be resistant to only two antibiotics, i.e. trimethoprim and sulphamethaxole. This suggests that the selected isolates (CM21, F4b2, F2a, E2, P1, F14a, FIX and F10a) are nonpathogenic in nature and sensitive to all the antibiotics tested. Subsequently, these isolates were further subjected to antimicrobial activity against pathogens such as L. monocytogenes ScottA, S. aureus FRI1722 and E. coli MTCC 118. The tested isolates (CM21, F2a, E2, P1, FIX, F10a) were able to inhibit L. monocytogenes ScottA and S. aureus FRI1722, whereas only three cultures, namely LP, F4b2 and F14a, were also able to inhibit only the growth of E. coli MTCC 118. Most of the isolates showed zone of inhibition [10 mm, suggesting their antagonistic activity against the food-borne pathogens. Further BSH assay revealed positive activity in all selected isolates (Table 1), except the culture F10a. Molecular Characterization of Selected Probiotic Bacteria To differentiate the selected probiotic LAB among each other, several molecular typing methods such as RFLP, ITS, RAPD and (GTG)5 PCR were used. The results obtained for fingerprints of RFLP and ITS for selected LAB were not clearly associated with species and strain level identification (Supplementary Fig. 1a and 1b). However, through RAPD analysis the isolates were differentiated at their species level. The isolates F14a and FIX that showed similar banding pattern with RFLP and ITS showed variation with RAPD PCR and suggested their species level variation. The isolates E2 and F2a showed similar banding profile, indicating their species level resemblance. Similar kind of observation was observed for the isolates F4C, F4b2 and P1 (Supplementary Fig. 1c). Strain level variation among the isolates F4C and F4b2 as well as for E2 and F2a was achieved with (GTG)5 (Supplementary Fig. 1d). However, discrimination and variation in the banding pattern among different species of same genera were observed. The phylogenetic dendrogram by binary scores of RAPD and (GTG)5 separately showed a clade switching of few strains at their genus-specific clusters (Fig. 1a, b). However, bifurcating neighbour-joining tree obtained for concatenated scores of RAPD and (GTG)5 revealed broad genus-specific clusters. The tree based on the concatenated scores of RAPD and (GTG)5 gave four major clusters with group I, II, III and IV represented as Lactobacillus,
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Pediococcus, Enterococcus and Staphylococcus group, respectively (Fig. 1c). The dendrogram obtained for 16S rDNA revealed their nominal species-specific clusters (Supplementary Fig. 2). The BLAST analysis of the selected probiotic LAB showed [98–99 % homology to their respective species. The selected isolates were concluded as follows—Staphylococcus epidermidis (F2a and E2), Pediococcus acidilactici (P1), Lactobacillus fermentum (F14a), Lactobacillus salivarius (FIX), Lactobacillus mucosae (CM21), Lactobacillus plantarum (LP), Enterococcus durans (F10a) and Pediococcus pentosaceus (F4b2 and F4C). Further, the carbohydrate fermentation studies of these isolates were found to be divergent at their species level and correlated towards their partial 16S rDNA sequence data (data not shown). In Vitro Adherence Capabilities of Probiotic LAB The cell surface hydrophobicity was determined for all the selected non-pathogenic isolates by using two hydrocarbons, namely n-hexadecane and xylene (Table 1). The values obtained for both the hydrocarbons were almost similar, with a maximum value above 50 % for the isolate P. acidilactici P1 and least value for S. epidermidis F2a with 30 %. Simultaneously, cellular auto-aggregation was evaluated, and highest value was observed for L. mucosae CM21 (61.7 %), followed by P. pentosaceus F4b2, L. fermentum F14a and P. acidilactici P1 with 51.8, 49.6 and 46.7 %, respectively. Lowest cellular auto-aggregation percentage was observed for S. epidermidis F2a (35 %), which also showed a similar trend for the tested hydrocarbons. Survival Under Simulated Gastric and Intestinal Fluids The selected cultures were further tested to observe their survival rates under simulated gastric (pH 2.5 with pepsin for 3 h) and intestinal fluids (0.1 % w/v bovine salt with pancreatin for 24 h) and the selected isolates were able to survive the adverse conditions (Table 2). Maximum percentage of survivability ([80 %) under gastric conditions was observed for L. salivarius FIX, L. mucosae CM21, P. acidilactici P1, P. pentosaceus F4b2 and L. fermentum F14a. The isolates P. acidilactici P1 and P. pentosaceus F4C showed [90 % survivability under simulated intestinal fluid conditions. Similarly, the isolates L. salivarius FIX, L. mucosae CM21 and L. fermentum F14a showed more than 80 % tolerance to bile–pancreatin conditions. The values obtained for the simulated gastric and intestinal fluids were found to be statistically significant (P [ 0.05) when compared to control.
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Cow milk
Human infant faecal
Human infant faecal
Human infant faecal
Paneer
Human infant faecal
Human infant faecal
Cow milk
Human infant faecal
Human infant faecal
Butter milk
L. plantarum LP
P. pentosaceus F4b2
L. fermentum F14a
S. epidermidis E2
P. acidilactici P1
P. pentosaceus F4C
L. salivarius FIX
L. mucosae CM21
S. epidermidis F2a
E. durans F10a
Enterococcus sp. 288
84.19 ± 0.006a 86.72 ± 0.006b 90.40 ± 0.007d 88.11 ± 0.005c 88.95 ± 0.003c 90.88 ± 0.002d 90.62 ± 0.008d 86.46 ± 0.007b 93.83 ± 0.006e
87.90 ± 0.007c 85.89 ± 0.005b 97.80 ± 0.004f 82.19 ± 0.006a 94.08 ± 0.003d 93.00 ± 0.007d 99.12 ± 0.005g 99.54 ± 0.006g 95.14 ± 0.007e
87.55 ± 0.005
b
c
87.25 ± 0.006
83.16 ± 0.004a
86.88 ± 0.007b
91.56 ± 0.004f
92.32 ± 0.006g
76.32 ± 0.005b
90.51 ± 0.004f
91.20 ± 0.005f
79.54 ± 0.007c
87.36 ± 0.008e
70.54 ± 0.003a
84.56 ± 0.006d
88.54 ± 0.005
e
85.36 ± 0.004d
?
-
?
?
?
?
?
?
?
?
?
BSH assay
ND
ND
32.6 ± 0.03a
47.5 ± 0.06d
46.3 ± 0.05c
34.0 ± 0.05b
54.0 ± 0.03e
33.0 ± 0.04a
48.6 ± 0.06d
47.3 ± 0.02
d
45.3 ± 0.06c
n-hexadecane
ND
ND
30.0 ± 0.02a
48.0 ± 0.03d
48.0 ± 0.04d
32.0 ± 0.04b
52.0 ± 0.04e
30.0 ± 0.05a
46.8 ± 0.05c
45.0 ± 0.03
c
45.8 ± 0.03c
Xylene
Hydrophobicity (%)
ND
ND
35.0 ± 0.06b
61.7 ± 0.04g
45.0 ± 0.06d
50.7 ± 0.05e
46.7 ± 0.03d
40.0 ± 0.06c
49.6 ± 0.04e
51.8 ± 0.03f
32.0 ± 0.04a
Autoaggregation (%)
ND
ND
22
21
16
20
20
21
23
20
–
Listeria monocyto genes ScottA
ND
ND
16
14
22
20
14
16
18
15
10
S. aureus FRI1722
ND
ND
–
–
–
–
–
–
13
11
10
E. coli MTCC118
Antimicrobial activity (zone of inhibition in mm)
The values represented are the mean ± SD (n = 3) of survival percentage of the bacterial cultures used, and the alphabetic superscripts (a, b, c, d, e, f, g) represented in the same columns followed by different letters were significantly different P\0.05
92.37 ± 0.002d
96.56 ± 0.005f
80.78 ± 0.005b
94.32 ± 0.006e
93.41 ± 0.005e
80.51 ± 0.004b
89.61 ± 0.006b
75.32 ± 0.008a
87.36 ± 0.007c
91.26 ± 0.006
d
88.32 ± 0.008c
1h
3h
1h
3h
Bile tolerance (0.3 %)
Acid tolerance test (pH 3)
? Positive activity, - Negative activity, ND not determined
Source
Isolates
Table 1 Probiotic attributes for selected lactic acid bacteria
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Fig. 1 Comparison of phylogenetic dendrogram constructed by using the scores of (GTG)5 (a), RAPD (b) and concatenated scores of (GTG)5 and RAPD (c)
Confirmation of Probiotic-Associated Genes For the detection of probiotic marker genes, the isolates L. fermentum F14a, L. plantarum LP, L. mucosae CM21, L. salivarius FIX, P. acidilactici P1 and P. pentosaceus F4b2 were selected. It was observed that the Pediococcus cultures (i.e. P1 and F4b2) showed an amplicon size of 155 bp with bshAF/R primer pair and that a product size of 384 bp was obtained with bshBF/R primers for L. plantarum LP and L. mucosae CM21, whereas bshCF/R primer amplified for L. salivarius FIX and L. fermentum F14a with a product size of 205 bp for bsh gene PCR. Simultaneously, Fbp protein was targeted to detect the presence of their adhesion ability by PCR. It was observed that L. plantarum LP, L. fermentum F14a, L. salivarius FIX and L. mucosae CM21 showed an amplicon size around 835 bp with Fbp1F/1R. Similarly, a product size around 1100 bp was obtained with Fbp2F/2R primer pair for Pediococcus cultures, i.e. F4b2 and P1. DNA sequencing of these products revealed their homology towards the fbp corresponding to their species-specific genes. Subsequently, the presence of another putative binding protein gene, i.e. mucin-binding protein, was investigated with different set of primers. A product size of 150 bp was obtained for all the selected isolates with PedLac_mubF1/ R1 primer pair, except for L. mucosae CM21. However, mub gene was targeted for L. mucosae CM21 with another set of primer (L.muc_mubF1/R1 primer pair) which gave an amplicon size of 550 bp. To confirm the presence of selected mub gene, the purified PCR product of around 150 and 550 bp from L. plantarum LP and L. mucosae CM21, respectively, was sequenced. The sequence data of these fragment from L. plantarum LP and L. mucosae CM21 revealed the presence of mub gene.
Comparative Genetic Analysis to Observe Diversity Among the Probiotic Marker Genes The complete genome sequences that were available publicly at online databases were retrieved for selected species of LAB. The in silico comparative genetic analysis for probiotic marker genes among different cultures of L. fermentum, L. plantarum, L. mucosae, L. salivarius, P. pentosaceus and P. acidilactici showed diversity among the binding domains of Mub and Fbp proteins (Fig. 2). The mucin-binding protein domain (MuBP) and fibronectinbinding protein domain (FBP) are defined according to the Pfam database. The analysis of fbp gene revealed that in most of the isolates only one binding domain was present. However, a lot of diversity was observed in case of mub gene. Highest number of MuBP domain within a mub gene was noticed for L. mucosae LM1 with 10 binding sites, and least was detected for Pediococcus sp.
Discussion Several experiments with different parameters were defined in the present study to isolate bacteria associated with probiotic properties from human infant faecal and dairy source. It is important for a probiotic strain to survive under gastric juice for 1–4 h and resist the stomach pH above 2 and also resist the simulated digestive juices [20]. Our results are in accordance with the above-mentioned report and found to survive under low pH and bile conditions. On further analysis for safety aspects, the tested cultures were found to be non-haemolytic and devoid of gelatinase, lecithinase activity and antibiotic resistance. It is essential for a probiotic strain to be sensitive to most of
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123
pH 7
pH 8
76.4 ± 0.4
24
76.7 ± 0.5
80.8 ± 0.1 83.1 – 0.2
85.4 ± 0.3 81.3 – 0.2
3
6
81.2 ± 0.2
76.0 ± 0.2
65.9 ± 0.2
24 83.4 ± 0.2
80.5 – 0.3
74.4 – 0.1
6
0
80.9 ± 0.1 76.8 ± 0.2
85.4 ± 0.1 82.1 ± 0.2
0 3
70.1 ± 0.2 81.4 – 0.2
84.9 ± 0.2 84.5 – 0.2
11/2
3
75.6 ± 0.2
81.3 – 0.2
85.9 ± 0.3
82.9 – 0.2
0
3
76.2 ± 0.2 72.6 ± 0.2
85.9 ± 0.1 84.2 ± 0.2
L. salivarius FIX
0
S. epidermidis F2a
11/2
Time intervals (h)
58.6 ± 0.1
75.5 – 0.2
78.1 ± 0.1
75.4 ± 0.2
55.4 ± 0.1
70.6 – 0.2
82.4 ± 0.2 79.3 ± 0.1
75.1 – 0.1
72.2 ± 0.2
72.5 ± 0.2
77.4 – 0.3
80.4 ± 0.1
81.8 ± 0.2
L. fermentum F14a
73.9 ± 0.2
85.5 – 0.2
75.0 ± 0.4
79.9 ± 0.2
81.4 ± 0.1
84.3 – 0.2
88.6 ± 0.2 89.8 ± 0.3
78.1 – 0.2
74.2 ± 0.2
80.4 ± 0.2
81.2 – 0.2
88.4 ± 0.1
90.9 ± 0.2
L. mucosae CM21
Bold values are statistically significant (P [ 0.05)
The values presented are the mean ±SD (n = 3) of survival percentage of the bacterial cultures used
Intestinal fluid
pH 2
Gastric fluid
pH 7
Parameters used
Simulated fluids
Table 2 Percentage tolerance of probiotic LAB to the simulated gastric and intestinal juices
67.7 ± 0.3
78.2 – 0.2
62.8 ± 0.2
44.4 ± 0.1
45.1 ± 0.2
63.0 – 0.2
60.2 ± 0.2 75.2 ± 0.1
74.9 – 0.1
70.8 ± 0.2
56.4 ± 0.2
83.0 – 0.1
68.7 ± 0.2
69.3 ± 0.1
P. pentosaceus F4b2
73.3 ± 0.2
83.5 – 0.2
71.8 ± 0.3
69.5 ± 0.2
93.4 ± 0.2
97.1 – 0.1
82.4 ± 0.2 86.2 ± 0.2
69.8 – 0.1
72.6 ± 0.2
88.6 ± 0.2
71.0 – 0.2
68.1 ± 0.1
76.3 ± 0.4
P. acidilactici P1
72.1 ± 0.2
74.3 – 0.1
74.2 ± 0.2
74.3 ± 0.1
90.7 ± 0.2
95.7 – 0.2
83.9 ± 0.2 94.1 ± 0.1
73.9 – 0.2
74.8 ± 0.1
75.6 ± 0.2
73.3 – 0.1
73.8 ± 0.2
76.8 ± 0.2
P. pentosaceus F4C
70.3 ± 0.1
72.8 – 0.2
71.3 ± 0.1
71.9 ± 0.1
97.2 ± 0.2
98.8 – 0.2
83.0 ± 0.2 99.3 ± 0.1
76.5 – 0.2
77.9 ± 0.1
80.3 ± 0.2
78.9 – 0.1
80.2 ± 0.2
81.2 ± 0.2
S. epidermidis E2
64.3 ± 0.1
75.4 – 0.2
85.4 ± 0.1
89.9 ± 0.2
66.5 ± 0.3
68.9 – 0.4
77.2 ± 0.3 75.1 ± 0.2
74.5 – 0.3
75.1 ± 0.1
70.4 ± 0.2
72.8 – 0.2
77.3 ± 0.1
78.6 ± 0.2
L. plantarum LP
188 Probiotics & Antimicro. Prot. (2015) 7:181–192
Probiotics & Antimicro. Prot. (2015) 7:181–192
189
Fig. 2 Comparative analysis among the functional domains of fibronectin- and mucin-binding proteins between different species of probiotic LAB
the antibiotics, as they may otherwise serve as agents in transfer of antibiotic resistance genes through conjugation [20, 31]. Many reports have been published in support for the production of bacteriocins among different species of LAB and are found to be effective against several foodborne pathogens present in the GIT of humans [4]. Moreover, this property serves as a good technological asset in storage of several food products and simultaneously lowers the risk factor to consumers [4]. It is important for probiotic bacteria to tolerate the bile salts in the upper small intestine and should have the ability to produce bile salt hydrolase (BSH) enzyme. The presence of BSH activity for selected isolates reveals hydrolysis of toxic conjugated bile salts to free bile acid and amino acids, thereby detoxifying the bile salt and simultaneously involving in the reduction in serum cholesterol level [20, 22]. Similar observation was reported for different probiotic Lactobacillus and Pediococcus species [19, 23, 24]. The administration of such bacteria with several probiotic properties may exert several beneficial properties to the human health.
Molecular characterization was used to differentiate the probiotic-associated bacteria. In the present study, the selected isolates were found to be divergent as analysed by molecular typing tools and phylogenetically they have clustered according to their species clades. Several probiotic Lactobacillus species have been differentiated successfully by RFLP and 16S-23S spacer region PCR methods [26]. Similarly, Devi and Halami [17] have characterized different bacteriocin-producing LAB by RFLP and RAPD typing tools. (GTG)5 PCR was reported to have the discriminatory power and reproducibility at species and sub-species level among LAB [32]. Based on the results obtained, (GTG)5 can be considered as a better method for characterization of probiotic cultures at their strain level. The phylogenetic tree deduced from individual scores of RAPD and (GTG)5 showed clade switching of the strains, and clear species-specific clusters were not obtained. Similar kind of observation was reported by Svec et al. [33] where a clade switching was observed for strain Enterococcus durans LMG 10746 from its species-specific
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190
cluster. However, the dendrogram obtained from concatenated scores of RAPD and (GTG)5 clearly resulted in genus-specific clusters and supports the phylogenetic correlation between different species of LAB. Hence, this method helped us to characterize the probiotic-associated LAB and non-LAB at their genus level. The present method employed in characterization of LAB was found more discriminating and reliable. The taxonomical identification of the selected isolates was further confirmed by sequencing of their 16S rDNA gene and simultaneously compared with their biochemical and physiological properties. The 16S rDNA gene analysis and the biochemical characterization were in complete agreement [14, 18]. The auto-aggregation and cell surface hydrophobicity properties were studied to assess the adherence ability of probiotic bacteria to intestinal epithelial mucosal cells and are found to be strongly co-related among several Lactobacillus and Bifidobacteria cultures [32]. Bacterial cells with high hydrophobicity were found to have either strong interaction towards mucosal cells or have a strong affinity of adherence to epithelial and/or mucous layers [34, 35]. Kos et al. [36] has reported that the cellular hydrophobicity may affect auto-aggregation as well as adhesion of bacteria to different cell surfaces. In contrast, Tuo et al. [34] has observed a correlation between hydrophobicity and adhesion ability among different Lactobacillus sp. However, in the present study the isolates were found to be statistically significant, and no much variation was observed between the different hydrocarbons used (P [ 0.05). No significant correlation (P [ 0.05) was found between the values of hydrophobicity and auto-aggregation, indicating their adherence ability to gut at various degrees. Though S. epidermidis E2 and F2a were able to withstand low pH and bile conditions, they were found to possess very less percentage of cellular hydrophobicity and auto-aggregation properties; hence, these cultures were not considered for further studies. Genetic screening was used to characterize the probiotic potential among the studied isolates to determine the presence of probiotic marker genes. Hence, in the present study the bsh gene (required to detoxify the bile salts), fbp and mub (involved in the binding mechanisms) were targeted. Our results are in accordance with the data stated by Turpin et al. [37] except for the isolate L. mucosae CM21 which was not reported earlier. However, sequencing of the bsh gene from L. mucosae CM21 showed maximum homology to L. fermentum, which suggests possible transfer of bsh gene. Mc Auliffe et al. [38] observed transfer of bsh gene from L. acidophilus NCFM to L. johnsonii NCC533 by insertional elements. Hence, this may be the reason for acquisition of bsh gene from L. fermentum to L. mucosae present in close vicinity of an ecological niche, where further investigation are essential. Pridmore et al. [39] and Kaushik et al. [24] reported that
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Probiotics & Antimicro. Prot. (2015) 7:181–192
the presence of functional bsh gene acts as a potential marker extensively used in the detection of GIT colonizing bacteria. Similar kind of observation was reported earlier for L. fermentum BCS 87 by Macias-Rodriguez et al. [40]. Then, in silico analysis of these sequenced products revealed the presence of these genes in several of the complete genome sequences of probiotic strains [41]. The putative binding proteins such as Mub and Fbp has been investigated only in L. plantarum Lp9 strain and was found to possess these genes at genetic level [24]. Ross et al. [42] first time reported a homologue of Mub protein in L. mucosae and their binding ability under in vitro conditions. The MUB protein contains several repeated functional domains, where they play a role in establishing host–microbial interactions with high affinity towards adherence and found to promote the evolution of GIT-associated organisms [13, 43]. Hence, the detection of binding-associated genes as well as binding domains strongly suggests the putative functional unit that plays a major role in adherence of probiotic bacteria to GIT of humans. The sequence alignments and comparative genetic tools are highly desirable in identifying the orthologous genes in a species, candidate genes associated with bacteriocin production, probiotic marker genes, immunomodulatory loci, adaptability, pathogenicity, etc. [44]. Many extracellular surface layer (S-layer) proteins are reported among LAB that are known to possess certain domains, which help the bacteria to get anchored to the host GI-tract mucus layer [45]. As complete genome sequence data for P. pentosaceus, P. acidilactici and L. mucosae are not available for many cultures, the prediction of protein domains in different isolates will be informative and helps in assessing biological function of such genes. A study on the adherence mechanisms of bacteria to the GIT of humans involves several expensive, exhaustive, laborious processes such as extracellular matrices, immunological detection, in vitro cell line studies, quantitative measurements and microscopic enumeration. Hence, genetic-based approaches are currently being used to reveal the bacterial S-layer proteinbinding mechanism to the host intestinal cells [12, 46]. In conclusion, the native isolates L. fermentum F14a, L. plantarum LP, L. mucosae CM21, L. salivarius FIX, P. pentosaceus F4b2 and P. acidilactici P1 were found to possess desirable in vitro probiotic properties. In our investigation, we have been able to detect probiotic marker genes by PCR and found that their presence correlated with in vitro probiotic properties. These methods are rapid, nonlaborious, easy to perform and cost-effective, aiding in identifying the adhesion domains of mucin- and fibronectin-binding proteins. Moreover, to exert the health benefits on the host, studying the colonization and adherence properties of probiotic bacteria is crucial. In this context, the results obtained indicated the variability
Probiotics & Antimicro. Prot. (2015) 7:181–192
among native isolates for adherence gene that was also supported with bioinformatic analysis and suggest playing a functional role in attaching. Hence, the cultures that are obtained in this investigation could be potential candidates for probiotic application. Presently, we are involved in in vivo evaluation and efficacy studies for the selected probiotic cultures. Further complete genome sequence and analysis of certain cultures will confirm their safety, probiotic properties, adhesion diversity and many more.
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12.
13. 14.
15. Acknowledgments We extend our gratitude to Prof. Ram Rajasekharan, Director, CSIR-CFTRI, Mysore, for encouragement and support. This work was carried out in the ICMR, New Delhi, funded project on probiotics (No. 5/9/1029/2011-RHN). Conflict of interest
The authors declare no conflicts of interest.
16.
17.
18.
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