Abstract
Background:
Escherichia coli is recognized as a common cause of infection. Long-lasting presence of bacteria on biotic and abiotic surfaces and failure of bacterial eradication are predicted by biofilm production. Some extracellular frills of E. coli can be implicated in productive events, leading to biofilm formation by surface colonization.Objectives:
In the present study, correlation of csgA (encoding curli fimbriae) and fimA (encoding a large subunit of type I fimberiae) gene expression with biofilm formation of extraintestinal bovine pathogenic E. coli strains was evaluated in different enrichment media.Methods:
The microtiter plate-based crystal violet method was applied to examine the biofilm production of 30 E. coli strains in Luria-Bertani (LB) and Brain-Heart Infusion (BHI) broth with 1% sucrose. PCR assay was performed to determine the presence of the studied genes.Results:
According to the results, 100% of isolates contained csgA genes, and 96.7% contained fimA genes. Using the BHI medium with 1% sucrose, 53.3% and 16.6% of strains were average and strong biofilm producers, respectively. On the other hand, by using the LB medium, 66.6% of isolates were poor biofilm producers, whereas none were strong biofilm producers.Conclusions:
The BHI medium containing 1% sucrose was better detected in biofilm production, compared to the LB medium. Since the studied genes were present in non-biofilm producing isolates, the correlation of these genes with biofilm-producing ability is questioned.Keywords
1. Background
Escherichia coli (E. coli) are facultative anaerobic bacteria, which are Gram-negative, rod-shaped, and motile by peritrichous flagella. Common classification of E. coli species is based on the mechanisms of pathogenicity, virulence factors, O and H antigenic serotyping, and clinical syndromes (1). E. coli may be expressed in different virulence factors related to different pathotypes, isolated from bovine infections, such as diarrhea and septicemia in calves and urinary infection, metritis, and mastitis in cows.
Microbial cell colonization of biofilms occurs on surfaces, and gentle rinsing cannot remove them. The similar structure of biofilms to the polysaccharide matrix material can support the survival and growth of bacteria in sessile environments (2). Pathogenic bacterial biofilms on indwelling devices or tissues cause infections and increase antimicrobial resistance and host immune responses (3). Production of these bacteria on medically relevant surfaces may be difficult to eradicate, resulting in the stability of infection (4).
Previous studies show that subinhibitory concentrations of antibiotics can trigger the formation of biofilms (5, 6). Pathogenic bacteria are embedded in a self-produced extracellular protein matrix and some micromolecules, including exopolysaccharide (EPS) and DNA (7, 8). Various definitions of biofilm indicate three major constituents, including the surface, microbes, and slime EPS; biofilm production can be terminated by removing any of these constituents (9).
Several superficial proteins in different organisms contribute to early biofilm formation and attachment to eukaryotic cell hosts. These surface determinants contribute to biofilm formation (e.g., flagella, autotransporter proteins, fimbriae, curli, EPS, and F-type conjugative pilus) (10). Synthesis of bacterial surface appendages (including flagella) in early stages of biofilm formation allows motility and reversible attachment, as major determinants of biofilm structure. In the second stage, adhesive organelles, such as type I fimbriae, which are encoded by fim genes, and curli fimbriae, encoded by csg operon, contribute to biofilm formation for irreversible attachment. At this stage, flagella synthesis is repressed (11, 12).
Curli are wiry, long, and thin protein fibers on bacterial surfaces (13). They not only improve the binding potential to surfaces, such as polystyrene, glass cover slips, and stainless steel in some Enterohemorrhagic Escherichia coli (EHEC) strains, however, they also have a greater capacity to bind intestinal cells in comparison with non-curli-producing strains (14). In some previous studies, association of biofilm formation by extraintestinal E. coli and different adhesins such as curli has been reported (15). Various bacterial species are known to form biofilms, which bacterial biofilm formation is clearly preferred in the majority of nutrient-sufficient environments.
2. Objectives
This study examined the biofilm-producing ability of pathogenic strains of E. coli via microtiter-plate crystal violet method in two different media and investigated its correlation with csgA and fimA gene expression.
3. Methods
3.1. Pathogenic E. coli Isolation
E. coli was isolated from the different infections of cattle (metritis, mastitis, and urinary infection) in the Khuzestan Province. After culturing the samples on blood agar, they were incubated at 37°C for one day. Suspicious colonies were again streaked on blood agar and examined by oxidase test, catalase test, Gram staining, and biochemical tests (i.e., MacConkey, triple sugar iron agar, urease, sulfur-indole motility, phenylalanine deaminase, simmon citrate, lysine iron agar, and methyl red Voges-Proskauer tests) (16). Generally, E. coli is a normal flora of some organs in animals and humans. Therefore, identification of E. coli as the main pathogen in clinical samples is important.
3.2. Biofilm Assay
The modified technique described by Stepanovic et al. was used for the assessment of biofilm-producing ability on polystyrene microtiter plates (17). The positive control was E. coli ATCC 25922. After separately growing the isolates on Brain-Heart Infusion (BHI) broth with 1% sucrose (Merck, Germany) and Luria-Bertani (LB) broth medium, they were incubated for one day at 37°C. Then, 100 microliter (μL) of overnight cultures, were added to 1 milliliter (mL) of fresh BHI broth (1) containing 1% sucrose, as well as LB broth. Following that, 200 μL of bacterial suspension (0.5 MacFarland) was incubated in triplicate on sterile, flat-bottomed, polystyrene microtiter plates (96 wells; Maxwell, China). Sterile, supplemented BHI and LB broth were used as negative controls in triplicate.
After 24 hours of incubation at 37°C, microtiter plates were aspirated and washed by 300 μL/well of sterile normal saline three times. After drying, for fixation of probably forming biofilms, 200 μL of methanol was used for 15 minutes. Afterwards, methanol was removed and the plates were dried at ambient temperature. Biofilm staining was performed using 200 μL of crystal violet 2% (Hucker’s solution). After five minutes, washing with distilled water and drying at ambient temperature were done. An enzyme-linked immunosorbent assay (ELISA) reader (Biotek SX2, USA) was used to measure the absorbance at 600 nanometer (nm) after adding ethanol-acetone (discoloring solution; 200 μL) for 15 minutes.
For each strain, the arithmetic mean of optical density (OD) in three wells was compared with the mean absorbance of negative controls (ODnc). Biofilm formation was classified as follows: strong (4.ODnc < ODs); moderate (2.ODnc < ODs < 4.ODnc); poor (ODnc < ODs < 2.ODnc); and no production (ODs < ODnc) (17, 18).
3.3. Polymerase Chain Reaction (PCR) Assay
According to a study by Silva et al. PCR assay was performed to identify curli (csgA) and Type I fimbria (fimA) genes in the isolated E. coli. From each strain, DNA was first extracted through boiling the bacterial suspension in tris-EDTA (TE) buffer with 2-mercaptoethanol (2%). Following centrifugation, the supernatant of suspension was used as the DNA source. Silva et al. designed specific primers for fimA and csgA genes. The specific primers for csgA gene included 5’-ATCTGACCCAACGTGGCTTCG-3’ and 5’-GATGAGCGGTCGCGTTGTTACC-3’, which detected the 178-bp segment. The specific primers for fimA gene, which amplified the 119-bp segment, were 5’-CTCTGGCAATCGTTGTTCTGTCG-3’ and 5’-GCAAGCGGCGTTAACAACTTCC-3’.
The PCR assay was performed in a total volume of 25 μL, including bacterial DNA (5 μL), forward and reverse primers (1 μL; 10 pmol/L), 2X PCR Master Mix (12.5 μL; Ampliqon), and nuclease-free water (5.5 μL). The assay included a four-minute cycle at 94°C; followed by 30 cycles for 30 seconds at 94°C, for 30 seconds at 60°C, and for 30 seconds at 72°C, as well as a final four-minute extension at 72°C in a thermal cycler (Eppendorf, Germany). E. coli ATCC 25922 and nuclease-free water were used as positive and negative controls, respectively. Via electrophoresis on 1% agarose gel (Max Pure, Spain), the PCR products were visualized. Then, a UV transilluminator (UVtech, Germany) was used for staining (safe stain 1 × SinaClon) (19).
4. Results
After collection of 30 E. coli isolates via cultivation of 64 clinical samples (mostly metritis samples; 83.3%), their biofilm-producing ability was evaluated using modified microtiter plates, presented by Stepanovic et al. Most isolates (66.6%) were poor biofilm producers in the LB medium, whereas in the BHI medium containing 1% sucrose, most isolates (53.3%) were moderate biofilm producers. In addition, the number of non-biofilm producing isolates was lower in the BHI medium (16.6%), compared to the LB medium (26.6%).
No isolate could strongly produce biofilm in the LB medium, while 16.6% of the isolates showed strong potentials. In fact, the BHI medium containing sucrose 1% had a greater biofilm-formation potential in comparison with the LB medium. Different levels of biofilm production by the studied media are demonstrated in the columns of Figure 1. According to the PCR assay, as demonstrated in Figures 2 and 3, all isolates (100%) were carriers of csgA gene, while only one isolate contained no fimA gene (3.3%) (Figures 2 and 3). Comparative diagram of the prevalence of studied genes is demonstrated in Diagram 1. In this research, all isolates with different biofilm-producing abilities contained csgA and fimA genes; therefore, the presence of csgA and fimA genes had no correlation with the biofilm-producing ability.
Comparison of biofilm inducing ability of LB medium (Blue) and BHI medium (Red) based on biofilm producing ability scale (Negative, Weak, Moderate and Strong) and number of studied strains.
CsgA gene in E.coli isolates; lane 1: 100 bp ladder; lane 2: positive control (178 bp); lane 3: negative control; lanes 4, 5, 6, 7: positive isolates.
FimA gene in E.coli isolates; lane 1: 100 bp ladder; lane 2: positive control (119 bp); lane 3: negative control; lanes 4, 5, 6, 7: positive isolates.
5. Discussion
Biofilm-producing bacteria cause various infections in humans and animals. Resistance of biofilm-producing bacteria to antibiotics and disinfectants is 500 - 5000 times higher than that of the planktonic type (20). Bacteria can be protected by the expression of specific resistance genes, besides the production of a large quantity of EPS during slow biofilm production (8). Dispersal of the planktonic type is required in new locations for biofilm production and colonization (21). The detected extraintestinal E. coli genotype may indicate the binding ability of E. coli strains to eukaryotic cells.
The improved binding ability of E. coli isolates to eukaryotic cells was indicated by the genotype of these extraintestinal isolates. Various surface organelles and extracellular molecules contribute to biofilm development of E. coli, including curli fimbriae, Type I pili, and flagella (22, 23). In several studies, correlation of different virulence factors in some bacteria with biofilm-producing ability was evaluated (24-27). Biofilm-producing ability has been examined by different quantification methods, including the commonly used microtiter plate systems (17, 28, 29). Biofilm formation by microtiter plate systems has been used for many different organisms and strains (18, 28, 30). Several studies have assessed the effects of enrichment medium type on biofilm assays. However, regulation of biofilm synthesis is a very complex process and there is scarce information in different species.
In this study, biofilm formation of 30 bovine extraintestinal E. coli isolates was assessed using the microtiter-plate crystal violet method in two culture media (LB and BHI + %1 sucrose), and correlation of attachment factors (Type 1 fimbria and curli fimbria) with biofilm-producing ability was examined in the isolates. By using two different culture media (LB and BHI + %1sucrose) in the microtiter plate system, 73.4% and 83.4% of the isolates were producers of biofilm in LB and BHI + %1 sucrose media, respectively.
The level of biofilm production was higher in the BHI medium containing sucrose (16.6%, strong production; 53.3%, moderate production) in comparison with the LB medium (0%, strong production; 66.6%, poor production). Some studies have examined the impact of enrichment medium type in biofilm assays. In this regard, Stepanovic et al. and Samet et al. found BHI medium to be superior to others (17, 31). In the study by Samet et al. BHI medium supplemented with 1% sucrose was used in biofilm production. In addition, no association was observed in uropathogenic E. coli strains between csgA and fimA genes.
Moori Bakhtiari and Javadmakoui in their studies detected no correlation between the biofilm-producing ability and presence of fimA and csgA genes in human uropathogenic E. coli strains. They recommended the BHI medium containing 1% sucrose for the study of biofilm-producing ability of these strains (32). In a study by Rijavec et al. biofilm production by pathogenic E. coli had no correlation with the presence of papC, usp, and sfa/foc virulence genes. However, Naves et al. showed that strong biofilm-producing strains of E. coli had a higher frequency of papG, sfa/foc, papC, hlyA, focG, and cnf1 genes (33, 34).
5.1. Conclusions
Since biofilm production is affected by different environmental factors, similar to the species and type of bacteria, extensive research is necessary on these isolates. In addition, considering the lack of association between the biofilm-producing ability and studied genes, evaluation of other involved genes is recommended.
Acknowledgements
References
-
1.
Montville TJ, Matthews KR. Food microbiol: an introduction. Washington (DC): ASM Press; 2005. 380 p.
-
2.
Hall-Stoodley L, Costerton J, Stoodley P. Bacterial biofilms: from the natural environment to infectious diseases. Natur Rev Microbiol. 2004;2(2):95-108. https://doi.org/10.1038/nrmicro821.
-
3.
Stewart PS, Franklin MJ. Physiological heterogeneity in biofilms. Natur Rev Microbiol. 2008;6(3):199-210. https://doi.org/10.1038/nrmicro1838.
-
4.
Da Re S, Le Quere B, Ghigo JM, Beloin C. Tight modulation of Escherichia coli bacterial biofilm formation through controlled expression of adhesion factors. Appl Environ Microbiol. 2007;73(10):3391-403. https://doi.org/10.1128/aem.02625-06.
-
5.
Chen C, Liao X, Jiang H, Zhu H, Yue L, Li S, et al. Characteristics of Escherichia coli biofilm production, genetic typing, drug resistance pattern and gene expression under aminoglycoside pressures. Environ Toxicol Pharmacol. 2010;30(1):5-10. https://doi.org/10.1016/j.etap.2010.03.004.
-
6.
Costa JCM, Espeschit Ide F, Pieri FA, Benjamin Ldos A, Moreira MA. Increased production of biofilms by Escherichia coli in the presence of enrofloxacin. Veterinar Microbiol. 2012;160(3-4):488-90. https://doi.org/10.1016/j.vetmic.2012.05.036.
-
7.
Gowrishankar S, Duncun Mosioma N, Karutha Pandian S. Coral-associated bacteria as a promising antibiofilm agent against methicillin-resistant and -susceptible staphylococcus aureus biofilms. Evidenc Based Complement Alternativ Med. 2012;2012:1-16. https://doi.org/10.1155/2012/862374.
-
8.
El-Feky MA, El-Rehewy MS, Hassan MA, Abolella HA, Abd El-Baky RM, Gad GF. Effect of ciprofloxacin and N-acetylcysteine on bacterial adherence and biofilm formation on ureteral stent surfaces. Pol J Microbiol. 2009;58(3):261-7. [PubMed ID: 19899620].
-
9.
Murugan K, Usha M, Malathi P, Al-Sohaibani AS, Chandrasekaran M. Biofilm forming multi drug resistant Staphylococcus spp. among patients with conjunctivitis. Pol J Microbiol. 2010;59(4):233-9. [PubMed ID: 21466040].
-
10.
Soto SM, Smithson A, Martinez JA, Horcajada JP, Mensa J, Vila J. Biofilm formation in uropathogenic Escherichia coli strains: relationship with prostatitis, urovirulence factors and antimicrobial resistance. J Urol. 2007;177(1):365-8. https://doi.org/10.1016/j.juro.2006.08.081.
-
11.
Pruss BM, Besemann C, Denton A, Wolfe AJ. A complex transcription network controls the early stages of biofilm development by Escherichia coli. J Bacteriol. 2006;188(11):3731-9. https://doi.org/10.1128/jb.01780-05.
-
12.
Wood TK, González Barrios AF, Herzberg M, Lee J. Motility influences biofilm architecture in Escherichia coli. Appl Microbiol Biotechnol. 2006;72(2):361-7. https://doi.org/10.1007/s00253-005-0263-8.
-
13.
Pawar DM, Rossman ML, Chen J. Role of curli fimbriae in mediating the cells of enterohaemorrhagic Escherichia coli to attach to abiotic surfaces. J Appl Microbiol. 2005;99(2):418-25. https://doi.org/10.1111/j.1365-2672.2005.02499.x.
-
14.
Boyer RR, Sumner SS, Williams RC, Pierson MD, Popham DL, Kniel KE. Influence of curli expression by Escherichia coli 0157:H7 on the cell's overall hydrophobicity, charge, and ability to attach to lettuce. J Food Prot. 2007;70(6):1339-45. [PubMed ID: 17612061].
-
15.
Uhlich GA, Cooke PH, Solomon EB. Analyses of the red-dry-rough phenotype of an Escherichia coli O157:H7 strain and its role in biofilm formation and resistance to antibacterial agents. Appl Environ Microbiol. 2006;72(4):2564-72. https://doi.org/10.1128/aem.72.4.2564-2572.2006.
-
16.
Quinn PJ, Carete RM, Markey B, Carter GR. Clinical veterinary microbiology: enterobacteriaceae. 2th ed. London UK: Mosby; 2004.
-
17.
Stepanovic S, Cirkovic I, Ranin L, Svabic-Vlahovic M. Biofilm formation by Salmonella spp. and Listeria monocytogenes on plastic surface. Letter Appl Microbiol. 2004;38(5):428-32. https://doi.org/10.1111/j.1472-765X.2004.01513.x.
-
18.
Stepanovic S, Vukovic D, Dakic I, Savic B, Svabic-Vlahovic M. A modified microtiter-plate test for quantification of staphylococcal biofilm formation. J Microbiol Method. 2000;40(2):175-9. [PubMed ID: 10699673].
-
19.
Silva VO, Espeschit IF, Moreira MAS. Clonal relationship of Escherichia coli biofilm producer isolates obtained from mastitic milk. Canadian J Microbiol. 2013;59(5):291-3. https://doi.org/10.1139/cjm-2013-0053.
-
20.
El-Shekh NA, Ayoub AMA, El-Hendawy HH, Abada EA, Khalifa SYE. In vitro activity of some antimicrobial agents against intact and disrupted biofilms of Staphylococci in the indwelling vascular catheter patients. World Appl Sci J. 2010;10(1):108-20.
-
21.
Costerton JW, Stewart PS, Greenberg EP. Bacterial biofilms: a common cause of persistent infections. Science. 1999;284(5418):1318-22. [PubMed ID: 10334980].
-
22.
Pratt LA, Kolter R. Genetic analyses of bacterial biofilm formation. Curr Opin Microbiol. 1999;2(6):598-603. https://doi.org/10.1016/s1369-5274(99)00028-4.
-
23.
Vidal O, Longin R, Prigent-Combaret C, Dorel C, Hooreman M, Lejeune P. Isolation of an Escherichia coli K-12 mutant strain able to form biofilms on inert surfaces: involvement of a new ompR allele that increases curli expression. J Bacteriol. 1998;180(9):2442-9. [PubMed ID: 9573197]. [PubMed Central ID: PMC107187].
-
24.
Ghasemian A, Najar Peerayeh S, Bakhshi B, Mirzaee M. High Frequency of icaAD, clumping factors A/B, fib and eno Genes in Staphylococcus aureus Species Isolated From Wounds in Tehran, Iran during 2012-2013. Arch Clin Infect Dis. 2015;10(4). e23033. https://doi.org/10.5812/archcid.23033.
-
25.
Reisner A, Krogfelt KA, Klein BM, Zechner EL, Molin S. In vitro biofilm formation of commensal and pathogenic Escherichia coli strains: Impact of environmental and genetic factors. J Bacteriol. 2006;188(10):3572-81. https://doi.org/10.1128/jb.188.10.3572-3581.2006.
-
26.
White-Ziegler CA, Um S, Perez NM, Berns AL, Malhowski AJ, Young S. Low temperature (23 C) increases expression of biofilm-, cold-shock- and RpoS-dependent genes in Escherichia coli K-12. Microbiol. 2008;154(1):148-66. https://doi.org/10.1099/mic.0.2007/012021-0.
-
27.
Skurnik M, Da Re S, Valle J, Charbonnel N, Beloin C, Latour-Lambert P, et al. Identification of commensal Escherichia coli genes involved in biofilm resistance to pathogen colonization. PLoS ONE. 2013;8(5). e61628. https://doi.org/10.1371/journal.pone.0061628.
-
28.
StepanoviĆ S, VukoviĆ D, Hola V, Bonaventura GD, DjukiĆ S, ĆIrkoviĆ I, et al. Quantification of biofilm in microtiter plates: overview of testing conditions and practical recommendations for assessment of biofilm production by staphylococci. Apmis. 2007;115(8):891-900. https://doi.org/10.1111/j.1600-0463.2007.apm_630.x.
-
29.
Christensen GD, Simpson WA, Younger JJ, Baddour LM, Barrett FF, Melton DM, et al. Adherence of coagulase-negative staphylococci to plastic tissue culture plates: a quantitative model for the adherence of staphylococci to medical devices. J Clin Microbiol. 1985;22(6):996-1006. [PubMed ID: 3905855]. [PubMed Central ID: PMC271866].
-
30.
Deighton MA, Balkau B. Adherence measured by microtiter assay as a virulence marker for Staphylococcus epidermidis infections. J Clin Microbiol. 1990;28(11):2442-7. [PubMed ID: 2254419]. [PubMed Central ID: PMC268203].
-
31.
Samet M, Ghaemi E, Jahanpur S, Jamalli A. Evaluation of biofilm-forming capabilities of urinary Escherichia coli isolates in microtiter plate using two different culture media. Int J Mol Clin Microbiol. 2013;1:244-7.
-
32.
Moori Bakhtiari N, Javadmakoei S. Survey on biofilm production and presence of attachment factors in human uropathogenic strains of Escherichia coli. Jundishapur J Microbiol. 2017;10(6). e13108. https://doi.org/10.5812/jjm-13108.
-
33.
Rijavec M, Muller-Premru M, Zakotnik B, Zgur-Bertok D. Virulence factors and biofilm production among Escherichia coli strains causing bacteraemia of urinary tract origin. J Med Microbiol. 2008;57(11):1329-34. https://doi.org/10.1099/jmm.0.2008/002543-0.
-
34.
Naves P, del Prado G, Huelves L, Gracia M, Ruiz V, Blanco J, et al. Correlation between virulence factors and in vitro biofilm formation by Escherichia coli strains. Microbial Pathogenesis. 2008;45(2):86-91. https://doi.org/10.1016/j.micpath.2008.03.003.