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A colorimetric microtiter plate method for assessment of phage effect on Pseudomonas aeruginosa biofilm
Journal of Microbiological Methods 74 (2008) 114–118
Contents lists available at ScienceDirect
Journal of Microbiological Methods j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / j m i c m e t h
Note
A colorimetric microtiter plate method for assessment of phage effect on Pseudomonas aeruginosa biofilm Petar Knezevic ⁎, Olga Petrovic Department of Biology and Ecology, Faculty of Sciences, University of Novi Sad, Trg Dositeja Obradovica 2, 21 000 Novi Sad, Vojvodina, Serbia
A R T I C L E
I N F O
Article history: Received 30 November 2007 Received in revised form 29 February 2008 Accepted 5 March 2008 Available online 14 March 2008
A B S T R A C T Bacteriophages have a potential in biofilm control. The aim of the study was to develop a method for selection of the most effective Pseudomonas aeruginosa phages for inhibition of biofilm formation and its eradication. The microtiter plate method is based on crystal violet staining and measuring of optical density. © 2008 Elsevier B.V. All rights reserved.
Keywords: Biofilm Crystal violet Microtiter plate Phage Pseudomonas aeruginosa Pseudomonas aeruginosa TTC
Attached, densely packed bacterial cells embedded in extracellular polysaccharide matrix on biotic or abiotic surfaces form a structure known as biofilm, showing many specific morphological and physiological characteristics in comparison to planktonic cells (O'Toole et al., 2000). One of the most important specificities of the attached cells is up to 1000 times higher resistance to antimicrobial agents, primarily as the result of slow growth and presence of impermeable exopolysaccharides on bacterial surface (Gilbert et al., 1997). P. aeruginosa is a species able to form biofilms on different abiotic surfaces, including artificial implants, contact lenses, urinary catheters and endotracheal tubes (reviewed in Davey and O'Toole, 2000). On the other hand, as an opportunistic pathogen, P. aeruginosa can cause various infections, which are difficult to treat with conventional antibiotics. Some of the infections, such as periodonitis, prostatitis and infections of patients with cystic fibrosis are considered to be in connection with bacterial attached mode of growth on the biotic surfaces (Hanlon, 2007; Lyczak et al., 2002). The biofilm forming P. aeruginosa produces a great amount of alginate — a linear polymanuronic–polyguluronic acid heteropolysaccharide (Linker and Johns, 1966) which binds bacteria together, plays a role in forming micro-
⁎ Corresponding author. Tel.: +381 214852681; fax: +381 21450620. E-mail addresses: [email protected] (P. Knezevic), [email protected] (O. Petrovic). 0167-7012/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.mimet.2008.03.005
colonies (Lam et al., 1980), and provides protection against unfavorable environmental factors, such as antimicrobials and humoral and cellular host defenses (Allison and Matthews, 1992; Abdi-Ali et al., 2006). Considering these facts, it seems useful to examine alternative solutions for degrading EPSs and controlling P. aeruginosa biofilms. In the last few years, the emergence of antibiotic resistant bacteria has increased interest in phage therapy. Phages have been examined to a lesser extent as potential agents for biofilm control (Doolittle et al., 1995; Hughes et al., 1998a; Hughes et al., 1998b; Hanlon et al., 2001; Sutherland et al., 2004; Hanlon, 2007). The main advantage of phage application is their ability to degrade exopolysacharides by enzymes attached to their baseplate — polysaccharide depolymerases and lyases, important for penetration into a host cell (Sutherland et al., 2004; Hughes et al., 1998b). The starting point for studying phages and/or their enzymes in biofilm control is their isolation and determination of potency to eliminate or inhibit biofilm formation. At the moment, there is only one method that allows comparison of different phages effect on bacterial growth in microplate format (McLaughlin, 2007), several microtiter plate methods for biofilm quantification (Christensen et al., 1985; O'Toole and Kolter, 1998; Stepanovic et al., 2000) and no microtiter plate method providing comparison of phages' effect on biofilm. The primary aim of the study is to establish a microtiter plate method for evaluation of phage effect on P. aeruginosa biofilm formation and removal, based on a combination of the above methods. The second aim is to apply the tests for assessing of isolated P. aeruginosa phages efficacy on biofilm removal, as well as to modify McLaughlin method (2007) for pseudomonas phage testing and
P. Knezevic, O. Petrovic / Journal of Microbiological Methods 74 (2008) 114–118
compare the results with the results of biofilm inhibition and eradication tests. Two different strains of P. aeruginosa were used in the study. The first strain, designated as PA-4u, was isolated from environmental biofilm using a selective medium for P. aeruginosa (Cetrimide agar, Serva Co., Feinbiochemica, Heidelberg, Germany). Confirmation of the species was based on morphological, tinctorial and biochemical characteristics (Gram staining, oxidase test, gelatin liquefaction, production of soluble pigment and nitrate utilization). The second strain was the reference strain P. aeruginosa ATCC 9027. Both strains were able to form a thick, compact biofilm when cultivated in Luria Bertain (LB) broth with glucose addition (Tryptone 10 g/l; yeast extract 5 g/l; NaCl 10 g/l; glucose 5 g/l; pH 7.5). Four lytic bacteriophages were isolated from different samples of municipal wastewater and designated as π-1, π-2, π-3 and π-4. Briefly, the samples were filtered through membrane filters (0.45 μm, Sartorius Co., Goettingen, Germany) and filtrate was added in double strengthen LB broth (v/v 1:1) inoculated with 100 μl of host strain PA4u. After enrichment (24 h at 35 °C and 120 rpm), the phages were isolated by the overlay agar method, picking a plaque by a sterile loop (Carlson, 2005). Each phage was reisolated three times and phage stocks were prepared by plate lysis and elution using SM buffer (50 mM Tris HCl [pH 7.5]; 0.1 M NaCl; 8 mM MgSO4; 0.01% w/v gelatin). The phage suspension was purified and concentrated by centrifugation (11,000 × g for 10 min at 4 °C), filtration (Minisart 0.2 μm, Sartorius Biotech BmbH, Goettingen, Germany), precipitation in NaCl and PEG6000 (Merck, Hohenbrunn, Germany) and ultracentrifugation in discontinuous glycerol gradient (110,000 × g for 1 h at 4 °C) (Sambrook and Russell, 2001). Plaque forming units per milliliter (PFU/ml) in the stocks were preliminary determined by the spot method, followed by overlay agar method for more precise count assessment (Carlson, 2005). The phage stocks were stored at 4 ± 1 °C and all dilutions of the original stocks were made in SM buffer with gelatin. Prior to the experiments, P. aeruginosa PA-4u and ATCC 9027 lawn were prepared by the overlay agar method, and 10 μl of each phage suspension (1 × 108 PFU/ml) was placed on the surface of semisolid medium in order to determine lytic spectra of the phages. Bacterial growth inhibition assay was carried out using a modified method developed by McLaughlin (2007). The modification included the following: triplicate application of each phage and bacterial count combination at the same microtiter plate, the reduction of incubation period after 2,3,5-triphenyltetrazolium chloride (TTC) addition, the use of double strengthen LB broth, and the reading absorbance at different wavelength. Briefly, bacterial cultures in LB broth were incubated until reached OD650 0.4 (approximately 2 × 109 CFU/ml) and used to inoculate double strengthen LB broth supplemented with 1% of glucose. Sterile 96-well microtiter plates with flat bottom (Spektar Cacak, Serbia) were filled with 100 μl of inoculated medium in such a way to get 1 × 102 CFU/well in the columns 1–3, 1 × 104 in columns 4–6, 1 × 106 in columns 7–9 and 1 × 108 CFU/well in the last three columns (10–12), with the exception of the wells H1, H4, H7 and H10 (Fig. 1A). These four wells did not contain bacteria and were filled with 100 μl of sterile double strengthen LB broth with glucose and 100 μl of sterile SM buffer. The other wells of the row H were filled additionally with 100 μl of sterile SM buffer (controls with appropriate bacterial CFU/ well without phage). Appropriate phage dilutions in SM buffer (100 μl) were added into the wells of rows from A to G in such a way to get 1 × 101–1 × 107 PFU/well, respectively. The microtiter plates were incubated in a thermostat with a water container (to elevate moisture level) at 35 °C overnight with shaking (120 rpm). After incubation, 50 μl of 0.1% filter-sterilized TTC (Sigma Chemical Co., St. Louis, MO, USA) was added into each well (final concentration 200 μg/ml, i.e. 50 μg/well) and incubated for additional 3 h. The absorbance was read at 540 nm using a microtiter plate reader (Multiskan EX, ThermoLabsystem, Vantaa, Finland).
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Fig. 1. Colorimetric microplate test of phage π-4 effect on P. aeruginosa PA-4u growth (A) and biofilm formation (B). Row H is a control (H1, H4, H7 and H10 are controls without bacteria and phages; the others are controls without phages — H2 and H3 contains 1 × 102 CFU/well of bacteria; H5 and H6 1 × 104 CFU/well; H8 and H9 1 × 106 CFU/ well and H11 and H12 1 × 108 CFU/well). Rows from A to G contain from 1 × 101 to 1 × 107 PFU/well of phage π-4, respectively. Columns 1–3 contain 1 × 102 CFU/well of PA-4u; columns 4–6 1 × 104 CFU/well; 7–9 1 × 106 CFU/well and 10–12 1 × 108 CFU/well.
For the study of bacteriophage effect on biofilm formation, the plates were filled and incubated in the same way as in the previous experiment (Fig. 1B). After overnight incubation, liquid content with planktonic cells was removed from the wells and each well was washed twice with 250 μl of PBS and left to dry. Attached bacterial cells were fixed with 250 μl of absolute methanol for 15 min. The fixative was removed and the plates were air-dried. Into each well, 200 μl of 0.4% crystal violet was added and after 15 min stain was removed. The plates were washed by stream of tap water in order to remove excessive amount of the stain and left to dry. Into each well of dried plates, 250 μl of 33% acetic acid was added and left for 20 min to allow stain to dissolve. The absorbance was measured at 595 nm using the microtiter plate reader. In order to determine phage effect on already formed biofilm, i.e. on its removal, the microtiter plates were filled with 100 μl of inoculated double strengthen medium as in the previous experiment, but instead of phages suspension in SM buffer, 100 μl of sterile SM buffer was added into each well. Consequently, the volume in each well, composition of the medium and bacterial CFU/well were the same as in the previous two experiments. The plates were incubated overnight to allow biofilm formation. When biofilm was formed after 24 hours, planktonic bacteria were removed and plates were washed once with PBS. The plates were left to dry for 10 min and then the wells were filled with 100 μl of appropriate phage dilution (as in previous cases) and with 100 μl of sterile double strengthen LB broth with glucose (to maintain the same experimental conditions). After incubation (for 24 h at 35 °C and 120 rpm), the plates were washed and stained as described above.
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Fig. 2. Bacteriophages on PA-4u (A) and P. aeruginosa ATCC 9027 lawns (B).
Each plate regarding one combination of phages and bacteria for growth inhibition, biofilm inhibition and biofilm removal assay was done in three independent experiments.
The bacterial count and/or biofilm reduction after the treatment with phages was calculated for each combination of phage and bacteria from the absorbance data. The average absorbance of wells filled with sterile medium and SM buffer was calculated and subtracted from the average absorbances of both controls containing only bacteria and the treated wells. The percentage of survived bacteria, for plates with TTC addition, was calculated by dividing the average absorbance of identically treated wells and the average absorbance of controls containing only bacteria multiplied by 100. The percentage was subtracted from 100% in order to get the percentage of growth inhibition. The calculations of biofilm formation inhibition and biofilm removal were carried out in the similar way for the plates stained with crystal violet. Reproducibility of the tests was estimated according to the coefficient of variation (S.D. / mean ⁎ 100). All isolated phages formed clear zones on PA-4u lawn when spot method was applied, but it was impossible to discriminate their lytic efficacy (Fig. 2A). Only phage π-2 was able to multiply on ATCC 9027 strain, and formed turbid zone (Fig. 2B).
Fig. 3. Effect of phage π-1 (a), π-2 (b), π-3 (c) and π-4 (d) on growth (A) and biofilm formation (B) of P. aeruginosa PA-4u: (■) represent 108 CFU/well of bacteria; (▲) 1 × 106 CFU/well; (♦) 1 × 104 CFU/well and (●) 1 × 102 CFU/well.
P. Knezevic, O. Petrovic / Journal of Microbiological Methods 74 (2008) 114–118
Fig. 4. Effect of phage π-4 on already formed P. aeruginosa PA-4u biofilm. (■) represent 1 × 108 CF/well of bacteria; (▲) 1 × 106 CFU/well; (♦) 1 × 104 CFU/well and (●) 1 ×102 CFU/ml.
The results of modified McLaughlin (2007) method regarding combination of phages with PA-4u are presented in Fig. 3(A). The modified method that uses less phage and bacterial combinations is more precise, because each combination of bacteria and phages can be tested at the same plate in triplicate. The absorbance was read after 3 h from the TTC addition, and as the plates were opened for reading in non-aseptic condition, any further change in color was not taken in consideration. McLaughlin (2007) reads absorbance at 590 nm, but our experience, as well as the experience of other authors (Gabrielson et al., 2002; Rahman et al., 2004) shows that 540 nm is optimal for reading after TTC addition, as it gives a higher resolution (absorbance values are not shown). The phages inhibition of bacterial growth is greater for isolate PA-4u than for ATCC 9027, as PA-4u is the original phage host. Phages π-2 and π-4 are very efficient and reduce bacterial biomass measured as A540 for 50% even at very low multiplicity of infection (MOI 10− 5 with bacterial CFU/ml 106 and 10− 6 with 108 CFU/ ml, respectively), while phages π-1 and π-3 barely achieve the same level of inhibition at much higher MOI (104 with CFU/ml 102). The results indicate a good potential of phages π-2 and π-4 application in growth control of P. aeruginosa populations. The results of phage effect on reference strain are not shown, as only phage π-2 had a negligible effect on growth inhibition. The phage effect on biofilm formation is presented in Fig. 3(B). All phages inhibited biofilm formation by more than 50% when PA-4u count was 1 × 102, 1 × 104 and 1 × 106 CFU/ml, while different effect was observed when CFU was 1 × 108. Similar to the previous experiment, in comparison to phages π-1 and π-3, phages π-2 and π-4 showed higher effect on PA-4u and inhibited biofilm formation by more than 50% with MOI 10− 5 and 10− 7, respectively. The different MOI needed for bacterial growth and biofilm formation inhibition is probably the result of phage enzymatic activity. For many phages, EPS acts as a secondary receptor, while primary receptors are located on outer membrane (Hughes et al., 1998a). Even if a phage cannot attach to primary receptors and infect cells, it can degrade EPS and inhibit biofilm formation. Moreover, as receptors on bacterial cells in planktonic and attached state are quantitatively and qualitatively different (Sutherland et al., 2004), it is obvious that phages can have higher or lower affinity to attached bacteria. Although phages showed considerable effect on growth inhibition and biofilm formation, their effect on already formed biofilm was very low. As this test is more discriminative than the previous two tests, it seems to be more appropriate for selection of phages for biofilm control — a significant effect on formed P. aeruginosa PA-4u biofilm was shown only by phage π-4 (Fig. 4). The phage almost completely eliminated biofilm at MOI 103 but only when bacterial count was 102 and 104, and reduced it for more than 50% with the MOI 10− 1 when bacterial count was 1 × 104 CFU/ml. Similarly, Hanlon et al. (2001) demonstrated a phage ability to reduce cell density of P. aeruginosa biofilm by 2 logs in intact biofilm and 3 logs in resuspended cells with a MOI 103. Our data confirm a good potential of phage application in
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P. aeruginosa biofilm control. However, the phages showed much greater effect on the host strain PA-4u than on the reference strain ATCC 9027 (the results are not shown), indicating that the phages are not universally applicable for the control of P. aeruginosa growth and biofilm. It is not surprising, as phages are usually highly specific for their receptors and slightly different receptors structure on bacterial surface may result in resistance to phage infection (Welkos et al., 1974). The coefficient of variation was different for the three tests: 18.3% for growth inhibition test, 16.7% for biofilm inhibition and 23.2% for biofilm removal assay. It should be emphasized that in a number of wells illogical coloration was detected in all three tests, the fact also observed for the growth inhibition test (McLaughlin, 2007). McLaughlin (2007) explains the phenomenon by an emergence of mutant bacteria resistant to phages. Mutants usually spontaneously appear during overnight incubation of bacteria and phages (Fridholm and Everitt, 2005), and this could be a reasonable explanation. However, it is important to point out that bacteria other than P. aeruginosa can reduce TTC, and that other bacteria are also able to form biofilms. On the other hand, phages are extremely selective and infect a single species or only few strains. The facts indicate that it is very important to maintain aseptic conditions throughout the experiments in order to reduce plates' contamination and non-specific reactions. The false results of the phages effect may have a great impact on standard deviation and hence on the results of experiments. If highly aseptic conditions are maintained, greater standard deviation may indicate higher bacterial potency to develop resistance to a phage, and the data should be taken in consideration during phage selection. The introduced tests for phage inhibition of biofilm formation and removal of already formed biofilm are easy, rapid, economical and satisfactory reproducible. The tests are appropriate to identify MOI needed for inhibition of biofilm formation or biofilm eradication, even in the absence of a microtiter plate reader by qualitative detection of color presence (“violet” or “colorless”). The method meets the criteria set for test development and could be useful for comparison of phage efficacy and selection of the most efficient phages in studies regarding their potential application for biofilm control. References Abdi-Ali, A., Mohammadi-Mehr, M., Agha Alaei, Y., 2006. Bactericidal activity of various antibiotics against biofilm-producing Pseudomonas aeruginosa. Int. J. Antimic. Agents 27, 196–200. Allison, D.G., Matthews, M.J., 1992. Effect of polysaccharide interactions on antibiotic susceptibility of Pseudomonas aeruginosa. J. Appl. Bacteriol. 73 (6), 484–488. Carlson, K., 2005. Working with bacteriophages: common techniques and methodological approaches. In: Kutter, E., Sulakvelidze, A. (Eds.), Bacteriophages: Biology and Applications. CRC Press, Boca Raton, Fla. Christensen, G.D., Simpson, W.A., Younger, J.J., Baddour, L.M., Barret, F.F., Melton, D.M., Beachey, E.H., 1985. Adherence of coagulase-negative staphylococci to plastic tissue culture plates: a quantitative model for the adherence of staphylococci to medical Devices. J. Clin. Microbiol. 22 (6), 996–1006. Davey, M.E., O'Toole, G.A., 2000. Microbial biofilms: from ecology to molecular genetics. Microbiol. Mol. Biol. Rev. 64 (2), 847–867. Doolittle, M.M., Cooney, J.J., Caldwell, D.E., 1995. Lytic infections of Escherichia coli biofilms by bacteriophage T4. Can. J. Microbiol. 41, 12–18. Fridholm, H., Everitt, E., 2005. Rapid reproducible infectivity end-point titration of virulent phage in a microplate system. J. Virol. Methods 128, 67–71. Gabrielson, J., Hart, M., Jarelov, A., Kuhn, I., McKenzie, D., Mollby, R., 2002. Evaluation of redox indicators and the use of digital scanners and spectrophotometer for quantification of microbial growth in microplates. J. Microbiol. Methods 50, 63–73. Gilbert, P., Das, J., Folez, I., 1997. Biofilm susceptibility to antimicrobials. Adv. Dent. Res. 11, 160–167. Hanlon, G.W., 2007. Bacteriophages: an appraisal of their role in the treatment of bacterial infections. Int. J. Antimic. Agents 30, 118–128. Hanlon, G.W., Denyer, S.P., Olliff, C.J., Ibrahim, L.J., 2001. Reduction in exopolysaccharide viscosity as an aid to bacteriophage penetration through Pseudomonas aeruginosa biofilms. Appl. Environ. Microbiol. 67 (6), 2746–2753. Hughes, K.A., Sutherland, I.W., Jones, M.V., 1998a. Biofilm susceptibility to bacteriophage attack: the role of phage-borne polysaccharide depolymerase. Microbiology 144, 3039–3047. Hughes, K.A., Sutherland, I.W., Clark, J., Jones, M.V., 1998b. Bacteriophage and associated polysaccharide depolymerases — novel tools for study of bacterial biofilms. J. Appl. Microbiol. 85, 583–598.
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Lam, J., Chan, R., Lam, K., Costerton, J.W., 1980. Production of mucoid microcolonies by Pseudomonas aeruginosa within infected lungs in cystic fibrosis. Infect. Immun. 28, 546–556. Linker, A., Johns, R.S., 1966. A new polysaccharide resembling alginic acid isolated from pseudomonads. J. Biol. Chem. 241, 3845–3851. Lyczak, J.B., Cannon, C.L., Pier, G.B., 2002. Lung infections associated with cystic fibrosis. Clin. Microbiol. Rev. 15, 194–222. McLaughlin, M.R., 2007. Simple colorimetric microplate test of phage lysis in Salmonella enterica. J. Microbiol. Methods 69 (2), 394–398. O'Toole, G.A., Kolter, R., 1998. Initiation of biofilm formation in Pseudomonas fluorescens WS365 proceeds via multiple, convergent signaling pathways: a genetic analysis. Mol. Microbiol. 28, 449–461. O'Toole, G., Kaplan, H.B., Kolter, R., 2000. Biofilm formation as microbial development. Annu. Rev. Microbiol. 54, 49–79.
Rahman, M., Kuhn, I., Rahman, M., Olsson-Liljeqist, B., Mollby, R., 2004. Evaluation of a scanner-assisted colorimetric MIC method for susceptibility testing of Gramnegative fermentative bacteria. Appl. Environ. Microbiol. 70 (4), 2398–2403. Sambrook, J., Russell, D.W., 2001. Molecular cloning: a laboratory manual, Third ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Stepanovic, S., Vukovic, D., Dakic, I., Savic, B., Svabic-Vlahovic, M., 2000. A modified microtiter-plate test for quantification of staphylococcal biofilm formation. J. Microbiol. Methods 40 (2), 175–179. Sutherland, I.W., Hughes, K.A., Skillman, L.C., Tait, K., 2004. The interaction of phage and biofilm. FEMS Microbiol. Lett. 232, 1–6. Welkos, S., Schreiber, M., Baer, H., 1974. Identification of Salmonella with the O-1 bacteriophage. Appl. Microbiol. 28 (4), 618–622.
Contents lists available at ScienceDirect
Journal of Microbiological Methods j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / j m i c m e t h
Note
A colorimetric microtiter plate method for assessment of phage effect on Pseudomonas aeruginosa biofilm Petar Knezevic ⁎, Olga Petrovic Department of Biology and Ecology, Faculty of Sciences, University of Novi Sad, Trg Dositeja Obradovica 2, 21 000 Novi Sad, Vojvodina, Serbia
A R T I C L E
I N F O
Article history: Received 30 November 2007 Received in revised form 29 February 2008 Accepted 5 March 2008 Available online 14 March 2008
A B S T R A C T Bacteriophages have a potential in biofilm control. The aim of the study was to develop a method for selection of the most effective Pseudomonas aeruginosa phages for inhibition of biofilm formation and its eradication. The microtiter plate method is based on crystal violet staining and measuring of optical density. © 2008 Elsevier B.V. All rights reserved.
Keywords: Biofilm Crystal violet Microtiter plate Phage Pseudomonas aeruginosa Pseudomonas aeruginosa TTC
Attached, densely packed bacterial cells embedded in extracellular polysaccharide matrix on biotic or abiotic surfaces form a structure known as biofilm, showing many specific morphological and physiological characteristics in comparison to planktonic cells (O'Toole et al., 2000). One of the most important specificities of the attached cells is up to 1000 times higher resistance to antimicrobial agents, primarily as the result of slow growth and presence of impermeable exopolysaccharides on bacterial surface (Gilbert et al., 1997). P. aeruginosa is a species able to form biofilms on different abiotic surfaces, including artificial implants, contact lenses, urinary catheters and endotracheal tubes (reviewed in Davey and O'Toole, 2000). On the other hand, as an opportunistic pathogen, P. aeruginosa can cause various infections, which are difficult to treat with conventional antibiotics. Some of the infections, such as periodonitis, prostatitis and infections of patients with cystic fibrosis are considered to be in connection with bacterial attached mode of growth on the biotic surfaces (Hanlon, 2007; Lyczak et al., 2002). The biofilm forming P. aeruginosa produces a great amount of alginate — a linear polymanuronic–polyguluronic acid heteropolysaccharide (Linker and Johns, 1966) which binds bacteria together, plays a role in forming micro-
⁎ Corresponding author. Tel.: +381 214852681; fax: +381 21450620. E-mail addresses: [email protected] (P. Knezevic), [email protected] (O. Petrovic). 0167-7012/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.mimet.2008.03.005
colonies (Lam et al., 1980), and provides protection against unfavorable environmental factors, such as antimicrobials and humoral and cellular host defenses (Allison and Matthews, 1992; Abdi-Ali et al., 2006). Considering these facts, it seems useful to examine alternative solutions for degrading EPSs and controlling P. aeruginosa biofilms. In the last few years, the emergence of antibiotic resistant bacteria has increased interest in phage therapy. Phages have been examined to a lesser extent as potential agents for biofilm control (Doolittle et al., 1995; Hughes et al., 1998a; Hughes et al., 1998b; Hanlon et al., 2001; Sutherland et al., 2004; Hanlon, 2007). The main advantage of phage application is their ability to degrade exopolysacharides by enzymes attached to their baseplate — polysaccharide depolymerases and lyases, important for penetration into a host cell (Sutherland et al., 2004; Hughes et al., 1998b). The starting point for studying phages and/or their enzymes in biofilm control is their isolation and determination of potency to eliminate or inhibit biofilm formation. At the moment, there is only one method that allows comparison of different phages effect on bacterial growth in microplate format (McLaughlin, 2007), several microtiter plate methods for biofilm quantification (Christensen et al., 1985; O'Toole and Kolter, 1998; Stepanovic et al., 2000) and no microtiter plate method providing comparison of phages' effect on biofilm. The primary aim of the study is to establish a microtiter plate method for evaluation of phage effect on P. aeruginosa biofilm formation and removal, based on a combination of the above methods. The second aim is to apply the tests for assessing of isolated P. aeruginosa phages efficacy on biofilm removal, as well as to modify McLaughlin method (2007) for pseudomonas phage testing and
P. Knezevic, O. Petrovic / Journal of Microbiological Methods 74 (2008) 114–118
compare the results with the results of biofilm inhibition and eradication tests. Two different strains of P. aeruginosa were used in the study. The first strain, designated as PA-4u, was isolated from environmental biofilm using a selective medium for P. aeruginosa (Cetrimide agar, Serva Co., Feinbiochemica, Heidelberg, Germany). Confirmation of the species was based on morphological, tinctorial and biochemical characteristics (Gram staining, oxidase test, gelatin liquefaction, production of soluble pigment and nitrate utilization). The second strain was the reference strain P. aeruginosa ATCC 9027. Both strains were able to form a thick, compact biofilm when cultivated in Luria Bertain (LB) broth with glucose addition (Tryptone 10 g/l; yeast extract 5 g/l; NaCl 10 g/l; glucose 5 g/l; pH 7.5). Four lytic bacteriophages were isolated from different samples of municipal wastewater and designated as π-1, π-2, π-3 and π-4. Briefly, the samples were filtered through membrane filters (0.45 μm, Sartorius Co., Goettingen, Germany) and filtrate was added in double strengthen LB broth (v/v 1:1) inoculated with 100 μl of host strain PA4u. After enrichment (24 h at 35 °C and 120 rpm), the phages were isolated by the overlay agar method, picking a plaque by a sterile loop (Carlson, 2005). Each phage was reisolated three times and phage stocks were prepared by plate lysis and elution using SM buffer (50 mM Tris HCl [pH 7.5]; 0.1 M NaCl; 8 mM MgSO4; 0.01% w/v gelatin). The phage suspension was purified and concentrated by centrifugation (11,000 × g for 10 min at 4 °C), filtration (Minisart 0.2 μm, Sartorius Biotech BmbH, Goettingen, Germany), precipitation in NaCl and PEG6000 (Merck, Hohenbrunn, Germany) and ultracentrifugation in discontinuous glycerol gradient (110,000 × g for 1 h at 4 °C) (Sambrook and Russell, 2001). Plaque forming units per milliliter (PFU/ml) in the stocks were preliminary determined by the spot method, followed by overlay agar method for more precise count assessment (Carlson, 2005). The phage stocks were stored at 4 ± 1 °C and all dilutions of the original stocks were made in SM buffer with gelatin. Prior to the experiments, P. aeruginosa PA-4u and ATCC 9027 lawn were prepared by the overlay agar method, and 10 μl of each phage suspension (1 × 108 PFU/ml) was placed on the surface of semisolid medium in order to determine lytic spectra of the phages. Bacterial growth inhibition assay was carried out using a modified method developed by McLaughlin (2007). The modification included the following: triplicate application of each phage and bacterial count combination at the same microtiter plate, the reduction of incubation period after 2,3,5-triphenyltetrazolium chloride (TTC) addition, the use of double strengthen LB broth, and the reading absorbance at different wavelength. Briefly, bacterial cultures in LB broth were incubated until reached OD650 0.4 (approximately 2 × 109 CFU/ml) and used to inoculate double strengthen LB broth supplemented with 1% of glucose. Sterile 96-well microtiter plates with flat bottom (Spektar Cacak, Serbia) were filled with 100 μl of inoculated medium in such a way to get 1 × 102 CFU/well in the columns 1–3, 1 × 104 in columns 4–6, 1 × 106 in columns 7–9 and 1 × 108 CFU/well in the last three columns (10–12), with the exception of the wells H1, H4, H7 and H10 (Fig. 1A). These four wells did not contain bacteria and were filled with 100 μl of sterile double strengthen LB broth with glucose and 100 μl of sterile SM buffer. The other wells of the row H were filled additionally with 100 μl of sterile SM buffer (controls with appropriate bacterial CFU/ well without phage). Appropriate phage dilutions in SM buffer (100 μl) were added into the wells of rows from A to G in such a way to get 1 × 101–1 × 107 PFU/well, respectively. The microtiter plates were incubated in a thermostat with a water container (to elevate moisture level) at 35 °C overnight with shaking (120 rpm). After incubation, 50 μl of 0.1% filter-sterilized TTC (Sigma Chemical Co., St. Louis, MO, USA) was added into each well (final concentration 200 μg/ml, i.e. 50 μg/well) and incubated for additional 3 h. The absorbance was read at 540 nm using a microtiter plate reader (Multiskan EX, ThermoLabsystem, Vantaa, Finland).
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Fig. 1. Colorimetric microplate test of phage π-4 effect on P. aeruginosa PA-4u growth (A) and biofilm formation (B). Row H is a control (H1, H4, H7 and H10 are controls without bacteria and phages; the others are controls without phages — H2 and H3 contains 1 × 102 CFU/well of bacteria; H5 and H6 1 × 104 CFU/well; H8 and H9 1 × 106 CFU/ well and H11 and H12 1 × 108 CFU/well). Rows from A to G contain from 1 × 101 to 1 × 107 PFU/well of phage π-4, respectively. Columns 1–3 contain 1 × 102 CFU/well of PA-4u; columns 4–6 1 × 104 CFU/well; 7–9 1 × 106 CFU/well and 10–12 1 × 108 CFU/well.
For the study of bacteriophage effect on biofilm formation, the plates were filled and incubated in the same way as in the previous experiment (Fig. 1B). After overnight incubation, liquid content with planktonic cells was removed from the wells and each well was washed twice with 250 μl of PBS and left to dry. Attached bacterial cells were fixed with 250 μl of absolute methanol for 15 min. The fixative was removed and the plates were air-dried. Into each well, 200 μl of 0.4% crystal violet was added and after 15 min stain was removed. The plates were washed by stream of tap water in order to remove excessive amount of the stain and left to dry. Into each well of dried plates, 250 μl of 33% acetic acid was added and left for 20 min to allow stain to dissolve. The absorbance was measured at 595 nm using the microtiter plate reader. In order to determine phage effect on already formed biofilm, i.e. on its removal, the microtiter plates were filled with 100 μl of inoculated double strengthen medium as in the previous experiment, but instead of phages suspension in SM buffer, 100 μl of sterile SM buffer was added into each well. Consequently, the volume in each well, composition of the medium and bacterial CFU/well were the same as in the previous two experiments. The plates were incubated overnight to allow biofilm formation. When biofilm was formed after 24 hours, planktonic bacteria were removed and plates were washed once with PBS. The plates were left to dry for 10 min and then the wells were filled with 100 μl of appropriate phage dilution (as in previous cases) and with 100 μl of sterile double strengthen LB broth with glucose (to maintain the same experimental conditions). After incubation (for 24 h at 35 °C and 120 rpm), the plates were washed and stained as described above.
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Fig. 2. Bacteriophages on PA-4u (A) and P. aeruginosa ATCC 9027 lawns (B).
Each plate regarding one combination of phages and bacteria for growth inhibition, biofilm inhibition and biofilm removal assay was done in three independent experiments.
The bacterial count and/or biofilm reduction after the treatment with phages was calculated for each combination of phage and bacteria from the absorbance data. The average absorbance of wells filled with sterile medium and SM buffer was calculated and subtracted from the average absorbances of both controls containing only bacteria and the treated wells. The percentage of survived bacteria, for plates with TTC addition, was calculated by dividing the average absorbance of identically treated wells and the average absorbance of controls containing only bacteria multiplied by 100. The percentage was subtracted from 100% in order to get the percentage of growth inhibition. The calculations of biofilm formation inhibition and biofilm removal were carried out in the similar way for the plates stained with crystal violet. Reproducibility of the tests was estimated according to the coefficient of variation (S.D. / mean ⁎ 100). All isolated phages formed clear zones on PA-4u lawn when spot method was applied, but it was impossible to discriminate their lytic efficacy (Fig. 2A). Only phage π-2 was able to multiply on ATCC 9027 strain, and formed turbid zone (Fig. 2B).
Fig. 3. Effect of phage π-1 (a), π-2 (b), π-3 (c) and π-4 (d) on growth (A) and biofilm formation (B) of P. aeruginosa PA-4u: (■) represent 108 CFU/well of bacteria; (▲) 1 × 106 CFU/well; (♦) 1 × 104 CFU/well and (●) 1 × 102 CFU/well.
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Fig. 4. Effect of phage π-4 on already formed P. aeruginosa PA-4u biofilm. (■) represent 1 × 108 CF/well of bacteria; (▲) 1 × 106 CFU/well; (♦) 1 × 104 CFU/well and (●) 1 ×102 CFU/ml.
The results of modified McLaughlin (2007) method regarding combination of phages with PA-4u are presented in Fig. 3(A). The modified method that uses less phage and bacterial combinations is more precise, because each combination of bacteria and phages can be tested at the same plate in triplicate. The absorbance was read after 3 h from the TTC addition, and as the plates were opened for reading in non-aseptic condition, any further change in color was not taken in consideration. McLaughlin (2007) reads absorbance at 590 nm, but our experience, as well as the experience of other authors (Gabrielson et al., 2002; Rahman et al., 2004) shows that 540 nm is optimal for reading after TTC addition, as it gives a higher resolution (absorbance values are not shown). The phages inhibition of bacterial growth is greater for isolate PA-4u than for ATCC 9027, as PA-4u is the original phage host. Phages π-2 and π-4 are very efficient and reduce bacterial biomass measured as A540 for 50% even at very low multiplicity of infection (MOI 10− 5 with bacterial CFU/ml 106 and 10− 6 with 108 CFU/ ml, respectively), while phages π-1 and π-3 barely achieve the same level of inhibition at much higher MOI (104 with CFU/ml 102). The results indicate a good potential of phages π-2 and π-4 application in growth control of P. aeruginosa populations. The results of phage effect on reference strain are not shown, as only phage π-2 had a negligible effect on growth inhibition. The phage effect on biofilm formation is presented in Fig. 3(B). All phages inhibited biofilm formation by more than 50% when PA-4u count was 1 × 102, 1 × 104 and 1 × 106 CFU/ml, while different effect was observed when CFU was 1 × 108. Similar to the previous experiment, in comparison to phages π-1 and π-3, phages π-2 and π-4 showed higher effect on PA-4u and inhibited biofilm formation by more than 50% with MOI 10− 5 and 10− 7, respectively. The different MOI needed for bacterial growth and biofilm formation inhibition is probably the result of phage enzymatic activity. For many phages, EPS acts as a secondary receptor, while primary receptors are located on outer membrane (Hughes et al., 1998a). Even if a phage cannot attach to primary receptors and infect cells, it can degrade EPS and inhibit biofilm formation. Moreover, as receptors on bacterial cells in planktonic and attached state are quantitatively and qualitatively different (Sutherland et al., 2004), it is obvious that phages can have higher or lower affinity to attached bacteria. Although phages showed considerable effect on growth inhibition and biofilm formation, their effect on already formed biofilm was very low. As this test is more discriminative than the previous two tests, it seems to be more appropriate for selection of phages for biofilm control — a significant effect on formed P. aeruginosa PA-4u biofilm was shown only by phage π-4 (Fig. 4). The phage almost completely eliminated biofilm at MOI 103 but only when bacterial count was 102 and 104, and reduced it for more than 50% with the MOI 10− 1 when bacterial count was 1 × 104 CFU/ml. Similarly, Hanlon et al. (2001) demonstrated a phage ability to reduce cell density of P. aeruginosa biofilm by 2 logs in intact biofilm and 3 logs in resuspended cells with a MOI 103. Our data confirm a good potential of phage application in
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P. aeruginosa biofilm control. However, the phages showed much greater effect on the host strain PA-4u than on the reference strain ATCC 9027 (the results are not shown), indicating that the phages are not universally applicable for the control of P. aeruginosa growth and biofilm. It is not surprising, as phages are usually highly specific for their receptors and slightly different receptors structure on bacterial surface may result in resistance to phage infection (Welkos et al., 1974). The coefficient of variation was different for the three tests: 18.3% for growth inhibition test, 16.7% for biofilm inhibition and 23.2% for biofilm removal assay. It should be emphasized that in a number of wells illogical coloration was detected in all three tests, the fact also observed for the growth inhibition test (McLaughlin, 2007). McLaughlin (2007) explains the phenomenon by an emergence of mutant bacteria resistant to phages. Mutants usually spontaneously appear during overnight incubation of bacteria and phages (Fridholm and Everitt, 2005), and this could be a reasonable explanation. However, it is important to point out that bacteria other than P. aeruginosa can reduce TTC, and that other bacteria are also able to form biofilms. On the other hand, phages are extremely selective and infect a single species or only few strains. The facts indicate that it is very important to maintain aseptic conditions throughout the experiments in order to reduce plates' contamination and non-specific reactions. The false results of the phages effect may have a great impact on standard deviation and hence on the results of experiments. If highly aseptic conditions are maintained, greater standard deviation may indicate higher bacterial potency to develop resistance to a phage, and the data should be taken in consideration during phage selection. The introduced tests for phage inhibition of biofilm formation and removal of already formed biofilm are easy, rapid, economical and satisfactory reproducible. The tests are appropriate to identify MOI needed for inhibition of biofilm formation or biofilm eradication, even in the absence of a microtiter plate reader by qualitative detection of color presence (“violet” or “colorless”). The method meets the criteria set for test development and could be useful for comparison of phage efficacy and selection of the most efficient phages in studies regarding their potential application for biofilm control. References Abdi-Ali, A., Mohammadi-Mehr, M., Agha Alaei, Y., 2006. Bactericidal activity of various antibiotics against biofilm-producing Pseudomonas aeruginosa. Int. J. Antimic. Agents 27, 196–200. Allison, D.G., Matthews, M.J., 1992. Effect of polysaccharide interactions on antibiotic susceptibility of Pseudomonas aeruginosa. J. Appl. Bacteriol. 73 (6), 484–488. Carlson, K., 2005. Working with bacteriophages: common techniques and methodological approaches. In: Kutter, E., Sulakvelidze, A. (Eds.), Bacteriophages: Biology and Applications. CRC Press, Boca Raton, Fla. Christensen, G.D., Simpson, W.A., Younger, J.J., Baddour, L.M., Barret, F.F., Melton, D.M., Beachey, E.H., 1985. Adherence of coagulase-negative staphylococci to plastic tissue culture plates: a quantitative model for the adherence of staphylococci to medical Devices. J. Clin. Microbiol. 22 (6), 996–1006. Davey, M.E., O'Toole, G.A., 2000. Microbial biofilms: from ecology to molecular genetics. Microbiol. Mol. Biol. Rev. 64 (2), 847–867. Doolittle, M.M., Cooney, J.J., Caldwell, D.E., 1995. Lytic infections of Escherichia coli biofilms by bacteriophage T4. Can. J. Microbiol. 41, 12–18. Fridholm, H., Everitt, E., 2005. Rapid reproducible infectivity end-point titration of virulent phage in a microplate system. J. Virol. Methods 128, 67–71. Gabrielson, J., Hart, M., Jarelov, A., Kuhn, I., McKenzie, D., Mollby, R., 2002. Evaluation of redox indicators and the use of digital scanners and spectrophotometer for quantification of microbial growth in microplates. J. Microbiol. Methods 50, 63–73. Gilbert, P., Das, J., Folez, I., 1997. Biofilm susceptibility to antimicrobials. Adv. Dent. Res. 11, 160–167. Hanlon, G.W., 2007. Bacteriophages: an appraisal of their role in the treatment of bacterial infections. Int. J. Antimic. Agents 30, 118–128. Hanlon, G.W., Denyer, S.P., Olliff, C.J., Ibrahim, L.J., 2001. Reduction in exopolysaccharide viscosity as an aid to bacteriophage penetration through Pseudomonas aeruginosa biofilms. Appl. Environ. Microbiol. 67 (6), 2746–2753. Hughes, K.A., Sutherland, I.W., Jones, M.V., 1998a. Biofilm susceptibility to bacteriophage attack: the role of phage-borne polysaccharide depolymerase. Microbiology 144, 3039–3047. Hughes, K.A., Sutherland, I.W., Clark, J., Jones, M.V., 1998b. Bacteriophage and associated polysaccharide depolymerases — novel tools for study of bacterial biofilms. J. Appl. Microbiol. 85, 583–598.
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Lam, J., Chan, R., Lam, K., Costerton, J.W., 1980. Production of mucoid microcolonies by Pseudomonas aeruginosa within infected lungs in cystic fibrosis. Infect. Immun. 28, 546–556. Linker, A., Johns, R.S., 1966. A new polysaccharide resembling alginic acid isolated from pseudomonads. J. Biol. Chem. 241, 3845–3851. Lyczak, J.B., Cannon, C.L., Pier, G.B., 2002. Lung infections associated with cystic fibrosis. Clin. Microbiol. Rev. 15, 194–222. McLaughlin, M.R., 2007. Simple colorimetric microplate test of phage lysis in Salmonella enterica. J. Microbiol. Methods 69 (2), 394–398. O'Toole, G.A., Kolter, R., 1998. Initiation of biofilm formation in Pseudomonas fluorescens WS365 proceeds via multiple, convergent signaling pathways: a genetic analysis. Mol. Microbiol. 28, 449–461. O'Toole, G., Kaplan, H.B., Kolter, R., 2000. Biofilm formation as microbial development. Annu. Rev. Microbiol. 54, 49–79.
Rahman, M., Kuhn, I., Rahman, M., Olsson-Liljeqist, B., Mollby, R., 2004. Evaluation of a scanner-assisted colorimetric MIC method for susceptibility testing of Gramnegative fermentative bacteria. Appl. Environ. Microbiol. 70 (4), 2398–2403. Sambrook, J., Russell, D.W., 2001. Molecular cloning: a laboratory manual, Third ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Stepanovic, S., Vukovic, D., Dakic, I., Savic, B., Svabic-Vlahovic, M., 2000. A modified microtiter-plate test for quantification of staphylococcal biofilm formation. J. Microbiol. Methods 40 (2), 175–179. Sutherland, I.W., Hughes, K.A., Skillman, L.C., Tait, K., 2004. The interaction of phage and biofilm. FEMS Microbiol. Lett. 232, 1–6. Welkos, S., Schreiber, M., Baer, H., 1974. Identification of Salmonella with the O-1 bacteriophage. Appl. Microbiol. 28 (4), 618–622.