Escherichia coli is considered as a member of the dominant flora inhabiting the human colonic region. Although most of the members of this species are harmless to the intestinal lumen, some acquired virulence factors and can cause a wide range of human diseases (Nataro et al., 1998). The pathogenic E. coli is causative of three clinical syndromes: urinary tract infections, enteric/diarrheal diseases and meningitis (Kaper et al., 2004). The key mechanisms by which E. coli cause enteric diseases include attachment and colonization of the intestinal mucosa, manipulation of the host cell cytoskeleton or evading host immune defenses, and production of toxins (Torres, 2009). Six categories of pathogenic E. coli are well-studied and comprise enteropathogenic E. coli, enterotoxigenic E. coli, enteroaggregative E. coli, enterohaemorrhagic E. coli, diffusely adherent E. coli and enteroinvasive E. coli (Croxen and Brett Finlay, 2012, Kaper et al., 2004). Diarrheal diseases caused by E. coli worldwide were estimated to be nearly 300–800 million clinical cases and 300,000–500,000 deaths every year (Torres, 2009); this highlights the significance of pathogenic E. coli in global health burden imposed by diarrheal diseases. Current interventions to inactivate/eliminate pathogenic E. coli involve the use of antibiotics. However, many pathogenic strains that are able to cause illness have become resistant to antibiotics (Collignon, 2009, Tadesse et al., 2012). The rise of antibiotic resistance has motivated research to find out antimicrobial alternatives of which probiotics have gained a growing interest.
The use of Lactobacillus spp. and Bifidobacterium spp. as probiotics to combat microbial infections and boosting human health inspired many studies. Probiotics have been associated with the treatment of gastroenteritis (Chai et al., 2013), antibiotic-associated diarrhoea (Friedman, 2012), necrotizing enterocolitis (Alfaleh et al., 2011), pouchitis (Wall et al., 2011), inflammatory bowel diseases (Schultz, 2008), allergic disorders and others (Minocha, 2009). The antimicrobial activity of a range of probiotics against pathogens including E. coli has been reported (Tejero-Sariñena et al., 2012). In addition, down-regulation of virulence genes expression in E. coli O157: H7 using bioactive molecules secreted by probiotics has been described (Medellin-Pena et al., 2007). Moreover, probiotics were capable of reduction of E. coli O157: H7 and E. coli O127: H6 adhesion to epithelial cells monolayers (Erdem et al., 2007). The ability of pathogenic E. coli to form biofilms that contribute to their pathogenicity was documented (Beloin et al., 2008, Martinez-Medina et al., 2009). The antibiofilm activity of probiotics against pathogenic E. coli is poorly studied. Here, we aimed to better utilize probiotics to combat multidrug-resistant E. coli and reduce their ability to form biofilms.
Bacterial strains and growth conditions
The bacterial strains used in this study are summarized in Table 1. Six strains of probiotics belonging to the genera Lactobacillus and Bifidobacterium were grown in Man-Rogosa-Sharpe (MRS, Oxoid, Hampshire, England) agar medium for 24 h at 37 °C with 5% CO2. A single colony from each strain was transferred into MRS broth under the same incubation conditions for 24 h for the preparation of the cell-free spent medium (CFSM). For pathogenic E. coli isolation and cultivation, clinical specimens from the intensive care unit (ICU) of Benha University Teaching hospital and sewage water samples from Benha city were collected. All samples were collected aseptically and transferred immediately to culture on MacConkey agar and Eosin methylene blue agar (Oxoid) followed by incubation aerobically at 37 °C for 24 h. The isolated colonies were further identified using Vitek 2 system (Biomerieux, USA). For the preparation of milk fermented by probiotics, a reconstituted skim milk powder (Nestle, Cairo, Egypt) was heated at 95 °C for 30 min and then cooled (4 °C) overnight. A 24 h fresh culture of the six probiotic strains was inoculated individually in the milk and then incubated under anaerobic conditions at 37 °C for 24 h.
Antibiotic susceptibility testing
Susceptibility testing was performed using the disc diffusion (modified Kirby Bauer) method (Biemer, 1973) for the following antibiotics (Oxoid, UK); Ampicillin (AM 10 µg), Cefotaxime (CTX 30 µg), Amikacin (AK 30 µg), Cefoxitin (FOX 30 µg), Amoxicillin + Clavulanic Acid (AMC 20 + 10 µg), Ceftriaxone (CRO 30 µg), Ciprofloxacin (CIP 5 µg), Clarithromycin (CL 15 µg), Ceftazidime (CAZ 30 µg). The results were inferred according to Clinical and Laboratory Standards Institute (CLSI) guidelines (CLSI 2013).
Preparation of the cell-free spent medium (CFSM) of probiotics
The preparation of CFSM of each probiotic strain was performed as described previously (Bayoumi and Griffiths, 2012). Briefly; overnight cultures of the six probiotic strains grown in MRS broth at 37 °C were diluted 1:100 with fresh medium and allowed to grow under same conditions to an optical density at 600 nm of 1.6 (∼1 × 108 cells/ml), the cells were then removed using centrifugation at 6000g at 4 °C for 10 min. The supernatant was filter-sterilized with 0.2 µm–pore-size filter, and referred to as cell-free spent medium (CFSM). In case of the milk fermented by probiotics, CFSM was obtained by centrifugation of the fermented milk at 10,000g at 15 °C for 15 min and then the supernatant was filter sterilized as mentioned above. The CFSM of all probiotic strains was stored at −20 °C until use for further assays.
Antibacterial activity of the probiotic CFSM
The E. coli test isolates were activated on trypticase soy agar (Oxoid) for 24 h at 37 °C. Suspension of each test bacteria of 109 CFU/ml was prepared by growing each bacterium in trypticase soy broth (Oxoid) for 24 h at 37 °C. The agar diffusion method was used to determine the inhibition zone of the test bacteria (Cruz et al., 2001). First, a bottom layer (10 ml) of TSA was prepared in the petri-dish. Secondly, a top layer of molten and cooled TSA (5 ml) mixed with each test bacteria suspension (109 CFU/ml) was poured on the bottom layer. Five 6 mm in diameter wells were prepared in each plate, and 100 µl of the probiotic CFSM was introduced in each well. The test bacteria were incubated for 24 h at 37 °C and the inhibition zones were measured in millimetres.
Initial bacterial inoculum for the biofilm experiment was prepared as mentioned previously (Arora and Kaur, 1999). A single colony of E. coli isolates was transferred into 5 ml of nutrient broth (Oxoid, UK) and incubated for 16 h at 37 °C to obtain the cells at exponential phase.
Evaluation of antibiofilm potential
The anti-biofilm formation activity of CFSM prepared from probiotics was assessed as previously described (Jadhav et al., 2013). Two groups labeled test agents (probiotic CFSM) along with their experimental control (broth medium) were prepared in the microtitre plate. Supplementation with 10% (vol/vol) of all probiotic CFSM was used as suggested previously (Medellin-Pena et al., 2007, Medellin-Peña and Griffiths, 2009). In each group, 40 µl of the test antibiofilm CFSM prepared was added in triplicate to the corresponding wells of sterile 96-well microtitre plate (Sigma Aldrich, USA) except for the negative controls. Then, 160 µl of E. coli cultures were added into wells in all the groups (the same volume of broth medium was added instead in case of experimental control) in a final volume of 200 µl per well. The microtitre plates were sealed and incubated for 16 h at 37 °C. The biofilm biomass compared to negative control was determined using crystal violet (CV) assay. The results were expressed as the percentage of inhibition.
After the treatment periods in preceding experiments, quantification of the biomass was carried out by CV assay (Djordjevic et al., 2002). The amount of biofilm formed on wells surface is reflected by the amount of stain absorbed. After incubation, the culture medium was discarded from each well and then all wells were washed three times with sterile distilled water to remove any loosely adhered cells. The microtitre plate was allowed to be air-dried, then further dried in the oven at 60 °C for 35 min and finally stained with 150 µl of 0.1% crystal violet. After incubating the microtitre at room temperature for 20 min, the wells were washed two times with sterile distilled water to remove the excessive unabsorbed stain. For estimating biofilm density, the absorbance was determined at 595 nm using an automated microplate reader (Sun Rise –TECAN. Inc.® USA). The mean absorbance (OD595 nm) of test organisms was determined and the percentage inhibition was calculated using the following formula (Eq. (1)):(1)Percentage inhibition=100-OD595nmtest for positive control well/OD595nmnegative control well×100
Coli isolation and antibiotic susceptibility profiling
In this work, six multidrug resistant E. coli were isolated. They were designated as E. coli IC1 to E. coli IC4, E. coli WW1 to E. coli WW2 as they were isolated from the intensive care unit and sewage water, respectively. All isolates were resistant to at least five antibiotics (Ceftazidime, Ampicillin, Clarithromycin, Amoxicillin + Clavulanic Acid and Ceftriaxone). Only one isolate showed no resistance to Cefoxitin. In addition, three isolates were intermediate in sensitivity to amikacin while two isolates exhibited intermediate sensitivity to cefoxitin and ciprofloxacin. These results demonstrated that the multidrug resistance pattern was exhibited by all isolates of which E. coli WW2 and E. coli IC2 were the most resistant to all antibiotics and were subjected to antibiofilm experiments. The antibiotic resistance patterns were summarized in Table 2.
Antibacterial activity of cell-free preparations of probiotics
The multidrug resistant E. coli isolates showed sensitivity to CFSM of all probiotics (Table 3). All the isolates were inhibited to a similar extent by the six probiotics (inhibition zones ∼ 13–14 mm) while highest antibacterial activity was observed in case of B. longum, L. acidophilus and B. bifidum against E. coli IC3, E. coli IC1 and E. coli WW2 (inhibition zones of 17.10 mm, 17.10 mm and 23.10 mm, respectively), respectively.
Antibiofilm activity of probiotics
The effect of CFSM of MRS fermented by probiotics on the initial attachment of E. coli IC2 and WW1 towards biofilm formation was observed and summarized in Table 4. In general, the two studied E. coli isolates showed a satisfactory ability to form biofilms while E. coli WW1 is a strong biofilm former as compared to IC2 strain. Interestingly, all probiotics resulted in inhibition of biofilm formation to a similar extent. B. longum caused the highest inhibition (57.94%) in case of E. coli IC2 while L. plantarum was responsible for 64.57% reduction of E. coli WW1 biofilms. E. coli WW1 biofilms were negatively influenced by CFSM of probiotics as compared to IC2 isolate. On the other hand, CFSM of skim milk fermented by probiotics exhibited a slight inhibitory activity against IC2 isolate using L. helveticus and L. rhamnosus (inhibition percentage of 31.52 and 17. 68, respectively). Unfortunately, the rest of probiotic strains had no inhibitory effect on IC2 biofilms (Table 5). In contrast, WW1 isolate was greatly affected by CFSM of fermented milk by all probiotics. B. longum and L. helveticus showed the strongest antibiofilm activity against the WW1 isolate (70.81 and 69.49 reduction percentages, respectively. These results demonstrated the promising inhibitory potential of CFSM of standard MRS broth medium and skim milk fermented by probiotics against biofilms formed by two multidrug resistant E. coli isolates.
The emergence of multidrug-resistant E. coli has been increasingly reported (Paterson and Bonomo, 2005). Nearly, 63% of total E. coli isolates in many countries were able to produce extended-spectrum beta-lactamase and the majority belonged to E. coli isolated from the ICU patients (Magiorakos et al., 2012, Nakai et al., 2016). In this work, all E. coli isolates were resistant to at least six antibiotics of different classes. Multidrug resistance of clinical pathogens imposes a rising threat to the human health by increasing the disease burden and spread. In previous studies, the resistance of E. coli to multiple drugs such as co-trimoxazole, penicillin and nitrofurantoin was at high frequencies (Mubita et al., 2008). High resistance rates of 60.6% of E. coli strains to cefazolin, tetracycline, ampicillin and trimethoprim/sulfamethoxazole was reported by (Li et al., 2017). The continuous emergence of multidrug resistance and even resistance to antibiotics of the last resort led to developing alternative intervention strategies to combat bacteria pathogenesis. Probiotics have received a growing interest in the prevention and treatment of infectious and other human diseases such as gastrointestinal, urogenital, respiratory and even periodontal diseases (Vuotto et al., 2014). In this work, cell-free preparations of different probiotics belong to Lactobacillus and Bifidobacterium species were able to reduce the growth of drug-resistant E. coli when investigated using agar well diffusion method. In other words, the inhibitory activity of selenium-enriched probiotics against pathogenic E. coli under in vitro and in vivo conditions was well-documented (Yang et al., 2009). In addition, probiotics isolated from yoghurts exhibited antibacterial effects against some common pathogens including E. coli (Kaboosi, 2011). Moreover, fifteen strains of probiotics belonging to many genera among which Lactobacillus and Bifidobacterium had antibacterial properties against gram negative and gram positive bacteria (Tejero-Sariñena et al., 2013). In the same cell, free supernatants of probiotics were used and the evidence of antibacterial properties was due to produced organic acids lowering the pH. In addition, bioactive compounds released by probiotics such as bacteriocins and hydrogen peroxide were responsible for their antimicrobial properties (Drider et al., 2016, Noordiana et al., 2013, Yang et al., 2014). These results support the potential use of probiotics/their bioactive compounds as antimicrobials against multidrug-resistant pathogens such as E. coli.
Biofilm formation is a phenomenon of microorganisms which results in a persistent microbial mass resistant to antimicrobial agents and related to about 80% of bacterial infections to humans. Infections due to biofilms formed by Staphylococcus sp. and enterobacteria such as E. coli are difficult to diagnose and can contribute to high healthcare costs and morbidity rates (Römling and Balsalobre, 2012). The antibiofilm properties of probiotics against biofilm-forming enteropathogens have been investigated, although the results obtained so far are few and conflicting. In this study, CFSM of probiotics grown in MRS broth or skim milk reduced the biofilm formation of two multidrug resistant E. coli. In previous studies single strains of L. acidophilus, L. plantarum, B. longum and B. lactis were effective is displacing Salmonella Typhimurium and E. coli from Caco-2 cell layer (Candela et al., 2008). In addition, exopolysaccharides produced by L. acidophilus decreases enterohaemorrhagic E. coli biofilms on polyvinyl chloride and polystyrene surfaces by affecting curli production genes (Kim et al., 2009). In vitro investigations focusing on bacteriocins production, adhesion, growth inhibition and co-aggregation of probiotics support their potential role in modulating microbial biofilms (Vuotto et al., 2014).
In conclusion, the antibacterial effects of cell-free preparations obtained from probiotics against multidrug-resistant E. coli support their effective use as antimicrobial alternatives and widen their applications in medicine and food bio-preservation as well as the possibility to eradicate biofilms formed by pathogenic E. coli.