ISSN 0026-2617, Microbiology, 2016, Vol. 85, No. 4, pp. 509–513. © Pleiades Publishing, Ltd., 2016. Original Russian Text © V.K. Plakunov, S.V. Mart’yanov, N.A. Teteneva, M.V. Zhurina, 2016, published in Mikrobiologiya, 2016, Vol. 85, No. 4, pp. 484–489.
SHORT COMMUNICATIONS
A Universal Method for Quantitative Characterization of Growth and Metabolic Activity of Microbial Biofilms in Static Models V. K. Plakunov1, S. V. Mart’yanov, N. A. Teteneva, and M. V. Zhurina Winogradsky Institute of Microbiology, Research Center of Biotechnology, Russian Academy of Sciences, Moscow, Russia 1e-mail:
[email protected] Received February 18, 2016
DOI: 10.1134/S0026261716040147
According to the results of worldwide research in medical microbiology, many microbial infections are caused by microorganisms organized as biofilms. Laboratory simulation of these communities in vitro is therefore necessary for investigation of the patterns of their formation and for testing the effect of antimicrobial preparations. For these studies, the correlation between in vitro and in vivo activity of biocidal agents is of primary importance. Standard criteria of antimicrobial activity used for planktonic cultures, such as the minimal inhibiting concentration (MIC), at which growth of planktonic cultures is suppressed completely (growth is not registered by optical techniques), or minimal bactericidal concentration (MBC), which causes the death of 99.9% of microorganisms (determined as the number of surviving cells), are not applicable to biofilm microbial communities. The properties of biofilm microbial populations, which contain high numbers of persister cells insensitive to antimicrobial agents (Verstraeten et al., 2016), and of viable nonculturable cells (Lee et al., 2007; Li et al., 2014), prevent the application of these techniques. The new pharmacodynamic parameters proposed in order to overcome these difficulties include biofilm prevention/inhibiting concentration (MBPC or MBIC) (Sabaeifard et al., 2014), minimal biofilm bactericidal concentration (MBBC) (Macià et al., 2014), and minimal biofilm eradication concentration (MBEC) (Takei et al., 2013). Since they cannot be determined using the standard techniques for biofilm testing, new approaches are required, including the application of metabolic indicators (Peeters et al., 2008). Existing methods for the reconstruction and modeling of microbial biofilms belong to several major types (McBain, 2009; Coenye and Nelis, 2010; Lebeaux et al., 2013): (1) “closed” or static models based on the application of microtiter plates (usually with 96 wells) and colony biofilms (Vandecandelaere et al., 2016); (2) “open” or dynamic systems with constant flow of fresh medium (Palmer, 1999; McBain, 2009);
and (3) microcosms, i.e., multispecies biofilms formed on a surface similar to the one occurring in the environment and simulating the in situ situation (Rudney et al., 2012). Approaches based on microfluidic techniques and combining the flow method with the possibility of continuous microscopic monitoring of the process of biofilm formation became popular recently (Coenye and Nelis, 2010; Kim et al., 2012; Hassanpourfard et al., 2014). Due to their relation to the topic of this article, we will discuss the most widespread models of the first type in more detail. The main shortcoming of microtiter plates (which are made of polystyrene, polypropylene, of polycarbonate) is disordered growth of microbial biofilms. They may form at the bottom of the well, on its wall, or on the surface of the medium. In the latter case, separation of the biofilm and the planktonic culture is almost impossible. Limited volume of the wells and poor mixing of the medium result in oxygen and nutrient limitation, i.e., in starvation stress, which in combination with an antimicrobial agent may have a synergistic or antagonistic effect on microbial growth. Moreover, conditions in which pathogenic microorganisms exist within a host are different from starvation conditions. Biofilm growth in the wells is usually determined by staining the cells attached to the walls and bottom of the well with bacteriological stains, usually crystal violet (CV). Since CV stains both the living and dead cells, as well as the extracellular polymer matrix (EPM) in which the cells are embedded, determination of the number of metabolically active bacteria is possible only after additional staining by the dyes which can be metabolized: resazurin, fluorescein diacetate, or tetrazolium salts (Peeters et al., 2008). Numerous recent modifications of this method ameliorate some of its shortcomings to a certain degree (Chavant et al., 2007; Lyamin et al., 2012). Quantitative determination of live cells in biofilms is a serious challenge. The standard staining technique (live/dead) is practically inapplicable to biofilms due
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to interaction of the dyes with the extracellular polymer matrix (EPM) containing high levels of DNA, which produces a screening effect (Netuschil et al., 2014). Since quantitative separation of the cells and EPM, even with harsh homogenization techniques affecting microbial viability, is almost impossible for many biofilms, especially for mature ones, enumeration of colony-forming units (CFU) results in determination of the minimal probable number of culturable cells. Moreover, the presence of viable nonculturable cells mentioned above also prevents accurate determination of the number of live bacteria in biofilms. We developed a method of simultaneous analysis of the biofilm and planktonic cultures, which is an alternative to microtiter plates and makes it possible to characterize biofilm growth in general, EPM synthesis, metabolic activity, and the minimal probable number of viable microbial cells in the biofilm. Two variants of static models for biofilm cultivation were used: on Teflon cubes in test tubes and on glass fiber filters in petri dishes with the growth medium. Experimental subjects were typical gram-positive (Staphylococcus aureus, S. epidermidis) and gram-negative (Pseudomonas aeruginosa, P. chlororaphis, Escherichia coli, and Chromobacterium violaceum) bacteria. The methods described below were shown to be applicable to all these subjects, including oil-oxidizing bacteria isolated from oil-contaminated soil (Rhodococcus equi), and from stratal waters of oil fields (Kocuria rhizophila, Dietzia natronolimnaea). Application of Teflon Cubes Staining with CV (which stains both live and dead cells, as well as the EPM), 1,9-dimethylmethylene blue (DMMB) (a more specific dye for the EPM acidic polysaccharides), and metabolized stain 3-(4,5-dimethyl-2thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) were used for analysis of planktonic cultures and biofilms. The latter dye is commonly used for detection of viable, metabolically active cells, acting as an electron acceptor in the mitochondrial respiration chain; in the process it is converted to water-insoluble blue formazan (Berridge and Tan, 1993). It is presently used in microtiter plate methods for detection of metabolically active cells in bacterial biofilms (Grare et al., 2008; Wang et al., 2010). Teflon (polytetrafluoroethylene, PTFE) has been often used (mainly as plates) as a surface for biofilm formation (Kondoh and Hashiba, 1998; Planchon et al., 2006; Fuchslocher Hellemann et al., 2013). We used the standard 4-mm PTFE cubes (Ftoroplastovye tekhnologii, Russia). The sort of PTFE is unimportant for the purpose. Prior to the experiment, the cubes were treated with the standard bichromate mixture (5% potassium bichromate in concentrated sulfuric acid) for 24 h at room temperature or for 1 h in a boil-
ing water bath, and washed with distilled water to achieve the neutral reaction of rinsing water. The cubes may be used repeatedly. Washed cubes (3 g, 21– 22 pieces) were transferred into the standard 20-mL bacteriological test tubes with 3 mL of the nutrient medium (usually LB medium). The surface area of the cubes (~20 cm2) in a test tube corresponded approximately to the area of eight wells of a standard 96-well plate, providing for stability and reliability of the experimental results. Sterilization was carried out at 1 atm. After addition of the tested compounds (e.g., antimicrobial agents) and inoculation (usually with 5– 50 μL of a 24-h culture), the tubes were incubated on a shaker at 150 rpm at a required temperature value. Biofilms were formed on PTFE cubes, while planktonic cultures grew in the liquid medium in the same test tube. The tubes were then cooled to room temperature, the planktonic culture was decanted, and the cubes were washed with 1% NaCl. Growth of the planktonic culture was determined using relative optical density (absorption + light scattering) at 540 nm. Biofilms were fixed with 96% ethanol for 15 min, washed with distilled water, and stained. For staining with CV and DMMB, the standard procedure (Peeters et al., 2008) was used with some modifications. MTT staining was carried out (without fixing) using our modification of the published procedure (Wang et al., 2010). This method was used successfully for investigation of the effect of a number of antibacterial and antibiofilm agents on formation of microbial biofilms (Gannesen et al., 2015; Mart’yanov et al., 2015). CV staining. CV solution (0.1%) was prepared by dissolving 500 mg CV (Sigma) in 15 mL of 96% ethanol and diluting the mixture with distilled water to 500 mL. This solution may be stored at room temperature in the dark for several months. CV solution (3 mL) was added to the test tubes with the cubes washed of the planktonic culture. After 30-min incubation, the solution was removed and the cubes were washed with distilled water to remove the excess dye. CV was extracted with 3 mL of 96% ethanol (the results did not change significantly at extraction times from 60 to 120 min). The experimental and control variants were treated similarly. Optical density of the extract (OD590) was measured against the relevant uninoculated control (a test tube with the cubes incubated together with the experimental and control tubes). DMMB staining. DMMB (1,9-dimethyl-methylene blue zinc chloride double salt, Sigma) was dissolved in 96% ethanol (80 mg/25 mL) and diluted with 100 mL 1 M guanidine chloride supplemented with sodium formate (1 g) and 99% formic acid (1 mL). The mixture was slightly acidic (pH 3). This solution may be stored at room temperature in the dark for several months. DMMB solution (3 mL) was added to the test tubes with the cubes washed of the planktonic culture. After incubation for 60–120 min, the solution MICROBIOLOGY
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was removed and the cubes were washed with distilled water to remove the excess dye. The dye was extracted with 3 mL of the solution containing 1.64 g sodium acetate, 100 g guanidine chloride, and 40 mL isopropanol in 400 mL distilled water (pH 6.8). Optical density was measured against the relevant uninoculated control within the range from 620 to 670 nm (similar to CV staining). MTT staining. MTT dissolved in sterile distilled water may be used for analysis of planktonic cultures. For investigation of washed biofilms, MTT should be dissolved in the relevant sterile medium, since the amount of intracellular endogenous substrates may be insufficient for quantitative MTT reduction. Planktonic cultures were stained by adding 1 mL of 0.2% MTT solution to 1 mL of the culture (OD 1.0– 1.5), which was then incubated for 60–120 min at 30°C, depending on the rate of reduction. The mixture was then cooled to room temperature and the cells were precipitated by centrifugation for 15 min at 7000 rpm. The supernatant was discarded, and 3 mL of chemically pure dimethyl sulfoxide (DMSO) was added to the cell pellet. Extraction was carried out to almost complete decoloration of the cells. The mixture was then centrifuged, and OD of the supernatant was measured within the range of 500–600 nm. Direct proportionality between optical densities of the cell suspension and of the extract (as well as the CFU values) was observed in the control cultures. For biofilm staining, the cubes washed of planktonic culture were incubated with 3 mL of 0.1% MTT solution in the medium for 60–120 min at 30°C to the maximal staining. In experiments with biocidal agents, the biocide concentration in the mixture should be the same as in the experimental test tubes. Excess dye was removed, and DMSO (3 mL) was added to the cubes. Optical density was measured against the relevant uninoculated control within the range from 500 to 600 nm (similar to CV and DMMB staining). Application of Glass Fiber Filters Determination of metabolically active microbial cells. The publications on the application of glass fiber filters for investigation of biofilm formation are few (Trémoulet et al., 2002; Guillier et al., 2008), primarily due to the fact that the dyes used for biofilm staining bind to glass fiber, resulting in too high background coloration of the extracts. MTT was used to stain the biofilms formed on glass fiber filters, since formazan forming in the reaction is localized inside the cells, thus providing for no background staining of the filter. This method makes it possible to determine the number of viable, metabolically active cells in biofilms and is therefore suitable for quantitative characterizaMICROBIOLOGY
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tion of the pharmacodynamic parameters mentioned above (MBBC and MBEC). Glass fiber paper (GF/F, Whatman) was used for detection of metabolically active cells in developing biofilms. It was cut into 2 × 2-cm squares, sterilized at 1 atm, and placed on the surface of solid nutrient medium supplemented with the tested compounds. A relevant amount of the solvent for the tested compounds was added to the control plates. The studied microorganism was applied to the filters as 50 μL of a 24-h planktonic culture. In the case of motile bacteria, the inoculum was immobilized in the medium with 0.3% agar. The density of planktonic cultures was found to be important. Since the rates of MTT reduction by different microorganisms vary considerably, an express method for determination of the optimal density was developed. Serial dilutions of the culture in the medium were applied (50 μL) on a strip of regular filter paper. Immediately after absorption of the suspension, 50 μL of 0.1% MTT solution in the medium was then added. The control spot (without suspension) contained only the MTT solution. After incubation for 30 min at 30°C, the color was assessed visually and the dilution resulting in weak, albeit visible, coloration was chosen. The control spot should remain uncolored. Glass fiber filters were incubated at 30°С. Incubation time depended on the goal of the experiment. After incubation, the filters were transferred into weighing bottles with 0.1% MTT solution in the medium and incubated at 30°С to complete development of the stain (usually 15 to 30 min). Stained filters were washed with distilled water and dried on strips of filter paper. Quantitative measurement of the staining was achieved by densitometry (e.g., on a Sorbfil densitometer, Tekhnokom, Russia) or by photometry after formazan extraction with DMSO, as was described above. The results obtained by both techniques were similar. Glass fiber filters may be used in test tube instead of PTFE cubes. In this case, a filter (2 × 2 cm) was placed into a test tube with 5 mL of the medium. All procedures were similar to the experiments with PTFE cubes, but only MTT staining was used. CFU determination. As was mentioned above, close association between microbial cells and EPM is the main factor hindering CFU determination in biofilms. Approaches used to separate the cells and EPM are either ineffective (e.g., DNase treatment) or result in partial disintegration of the cells (sonication). Highspeed centrifugation in a density gradient and mechanical homogenization techniques proved to be milder and more efficient (Toyofuku et al., 2012). The glass fibers of the filters may act as an abrasive material, and intense shaking may result in good homogenization (Wang et al., 2010). For this purpose, a bio-
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film-covered filter was placed into a test tube with 5 mL of sterile medium, homogenized with a sterile glass rod, and shaken on a Vortex-ZX3 mixer at the maximal rate. After precipitation of the fragments of the filter, tenfold dilutions in sterile medium were prepared from the supernatant and then plated (50 μL) on petri dishes with agar medium. The material was spread with a sterile spatula, and the colonies were counted after incubation for 48 h at 30°C. While the results of CFU counts were shown to correlate well with those of MTT staining, due to the factors described above it is certainly a probabilistic estimate of the minimal possible number of living cells in a biofilm. The work was supported by the Russian Science Foundation, project no. 16-14-00028. REFERENCES Berridge, M.V. and Tan, A.S., Characterization of the cellular reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide (MTT): subcellular localization, substrate dependence, and involvement of mitochondrial electron transport in MTT reduction, Arch. Biochem. Biophys., 1993, vol. 303, pp. 474–482. Chavant, P., Gaillard-Martinie, B., Talon, R., Hébraud, M., and Bernardi, T., A new device for rapid evaluation of biofilm formation potential by bacteria, J. Microbiol. Methods, 2007, vol. 68, pp. 605–612. Christensen, G.D., Simpson, W.A., Younger, J.J., Baddour, L.M., Barrett, F.F., Melton, D.M., and Beachey, E.H., 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, vol. 22, pp. 996–1006. Coenye, T. and Nelis, H.J., In vitro and in vivo model systems to study microbial biofilm formation, J. Microbiol. Methods, 2010, vol. 83, pp. 89–105. Fuchslocher Hellemann, C., Grade, S., Heuer, W., Dittmer, M.P., Stiesch, M., Schwestka-Polly, R., and Demling, A.P., Three-dimensional analysis of initial biofilm formation on polytetrafluoroethylene in the oral cavity, J. Orofac. Orthop., 2013, vol. 74, pp. 458–467. Gannesen, A.V., Zhurina, M.V., Veselova, M.A., Khmel’, I.A., and Plakunov, V.K., Regulation of biofilm formation by Pseudomonas chlororaphis in an in vitro system, Microbiology (Moscow), 2015, vol. 84, no. 3, pp. 319– 327. Grare, M., Fontanay, S., Cornil, C., Finance, C, and Duval, R.E., Tetrazolium salts for MIC determination in microplates: Why? Which salt to select? How?, J. Microbiol. Methods, 2008, vol. 75, pp. 156–159. Guillier, L., Stahl, V., Hezard, B., Notz, E., and Briandet, R., Modelling the competitive growth between Listeria monocytogenes and biofilm microflora of smear cheese wooden shelves, Int. J. Food Microbiol., 2008, vol. 128, pp. 51–57.
Hassanpourfard, M., Sun, X., Valiei, A., Mukherjee, P., Thundat, T., Liu, Y., and Kumar, A., Protocol for biofilm streamer formation in a microfluidic device with micro-pillars, J. Vis. Exp., 2014, vol. 90. doi 10.3791/51732 Kim, J., Park, Y.-D., and Chung, S., Microfluidic approaches to bacterial biofilm formation, Molecules, 2012, vol. 17, pp. 9818–9834. Kondoh, K. and Hashiba, M., Inhibitory effect of macrolide antibiotics on biofilm formation by Pseudomonas aeruginosa, Nihon Jibiinkoka Gakkai Kaiho, 1998, vol. 101, pp. 25–36. Lebeaux, D., Chauhan, A., Renducles, O., and Beloin, C,. From in vitro to in vivo models of bacterial biofilm-related infections, Pathogens, 2013, vol. 2, pp. 288–356. Lee, D.G., Park, S.J., and Kim, S.J., Influence of pipe materials and VBNC cells on culturable bacteria in a chlorinated drinking water model system, J. Microbiol. Biotechnol., 2007, vol. 17, pp. 1558–1562. Li, L., Mendis, N., Trigui, H., Oliver, J.D., and Faucher, S.P., The importance of the viable but non-culturable state in human bacterial pathogens, Front. Microbiol., 2014, vol. 5, p. 258. doi 10.3389/fmicb.2014.00258 Lyamin. A.V., Botkin, E.A., and Zhestkov, A.V., Methods of biofilm detection in medicine, Klin. Mikrobiol. Antimikrob. Khimioter., 2012, vol. 14, no. 1, pp. 17–22. Macià, M.D., Rojo-Molinero, E., and Oliver, A., Antimicrobial susceptibility testing in biofilm growing bacteria, Clin. Microbiol. Infect., 2014, vol. 20, pp. 981–990. Mart’yanov, S.V., Zhurina, M.V., El’Registan, G.I., and Plakunov, V.K., Activation of formation of bacterial biofilms by azithromycin and prevention of this effect, Microbiology (Moscow), 2014, vol. 83, no. 6, pp. 723–731. McBain, A.J., Chapter 4: In vitro biofilm models: an overview, Adv. Appl. Microbiol., 2009, vol. 69, pp. 99–132. Netuschil, L., Auschill, T.M, Sculean, A., and Arweiler, N.B., Confusion over live/dead stainings for the detection of vital microorganisms in oral biofilms–which stain is suitable?, BMC Oral Health, 2014, vol. 14, p. 2. doi 10.1186/1472-6831-14-2 Palmer, R.J., Jr., Microscopy flowcells: perfusion chambers for real-time study of biofilms, Methods Enzymol., 1999, vol. 310, pp. 160–166. Peeters, E., Nelis, H.J., and Coenye, T., Comparison of multiple methods for quantification of microbial biofilms grown in microtiter plates, J. Microbiol. Methods, 2008, vol. 72, pp. 157–165. Planchon, S., Gaillard-Martinie, B., Dordet-Frisoni, E., Bellon-Fontaine, M.N., Leroy, S., Labadie, J., Hébraud, M., and Talon, R., Formation of biofilm by Staphylococcus xylosus, Int. J. Food. Microbiol., 2006, vol. 109, pp. 88–96. Rudney, J.D., Chen, R., Lenton, P., Li, J., Li, Y., Jones, R.S., Reilly, C., Fok, A.S., and Aparicio, C.A., A reproducible oral microcosm biofilm model for testing dental materials, J. Appl. Microbiol., 2012, vol. 113, pp. 1540– 1553. MICROBIOLOGY
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Vandecandelaere, I., Van Acker, H., and Coenye, T.A., Microplate-based system as in vitro model of biofilm growth and quantification, Methods Mol. Biol., 2016, vol. 1333, pp. 53–66. Verstraeten, N., Knapen, W., Fauvart, M., and Michiels, J., A historical perspective on bacterial persistence, Methods Mol. Biol., 2016, vol. 1333, pp. 3–13. Wang, H., Cheng, H., Wang, F., Wei, D., and Wang, X., An improved 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) reduction assay for evaluating the viability of Escherichia coli cells, J. Microbiol. Methods, 2010, vol. 82, pp. 330–333.
Translated by P. Sigalevich
