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FEMS MicrobiologyLetters 9 (1980) 29-33 © CopyrightFederation of European MicrobiologicalSocieties Published by Elsevier/North-HollandBiomedicalPress 29 A D H E R E N C E O F B A C T E R I A TO H Y D R O C A R B O N S : A SIMPLE M E T H O D F O R MEASURING CELL-SURFACE HYDROPHOBICITY M. ROSENBERG, D. GUTNICK and E. ROSENBERG Department of Microbiology, George S. WiseFaculty of Life Sciences, Tel Aviv University, Ramat Aviv, Israel Received 20 May 1980 Accepted 20 June 1980 1. Introduction The hydrophobic nature of the outermost surface of various microbial cells has been implicated in such biological phenomena as interactions between bacteria and phagocytes [1], attachment of bacteria to host tissue [2,3], adherence of bacteria to nonwettable solid surfaces [4,5], partitioning of bacteria at liquid : liquid [6] and liquid : air [7,8] interfaces, and the ability of microbial cells to grow on hydrocarbons through direct contact with the immiscible substrate [9-12]. A number of methods for studying hydrophobic interactions of cells have been reported in the literature. These include binding of hydrocarbon and fatty acids to cells and cell components [12,13], measurement of the force required to remove hydrocarbonbound cells [9], partitioning of bacteria in aqueous polymer two-phase systems [ 14,15], hydrophobic interaction chromatography [3,15] and contact angle measurements of dried cell layers [1,16]. No single method adequately describes cell-surface hydrophobicity since (1) the experimental conditions employed influence the observed hydrophobic interactions to some degree and (2) various non-hydrophobic effects often interfere. Interest in the mechanism which enables direct contact between hydrocarbon-degrading cells and their water-insoluble alkane substrates has led us to develop a rapid quantitative assay for the hydrophobic interaction of ceils with liquid hydrocarbons. method is based on the degree of adherence of cells to various liquid hydrocarbons following a brief period of mixing. The present report describes this technique and its application in measuring the surface hydrophobicity of various bacterial cells. 2. Materials and Methods 2.1. Bacteria The following test bacteria were used: Acinetobacter calcoaceticus strain RAG-1 (ATCC 31012) was isolated previously in this laboratory [ 17,18] ; A. calcoaceticus strain BD 413 tryp E 27 was kindly provided by Dr. E. Juni. Escherichia cell B ilvA thy, E. coli K-12 CSH 57, E. coil J-5, Bacfflus subtilis 168 and Enterobacter aerogenes CDC 659]66 were kindly provided by Dr. E.Z. Ron. Micrococcus lysodeikticus ATCC 4698 was kindly pr6vided by Dr. I. Friedberg. Locally isolated strains of Staphylococcus aureus, Staphylococcus albus and Serratia marcescens were kindly provided by Ruth Zack. Pseudomonas aeruginosa PAS 279 was kindly provided by Dr. J. Shapiro. The test bacteria were grown at 30°C with shaking in nutrient broth. Unless otherwise stated, bacteria were harvested at earlY logarithmic growth phase, washed twice and resuspended in PUM buffer, pH 7.1 : 22.2 g K2I-IPO4 • 3H20, 7.26 g KH2PO4, 1.8 g urea, 0.2 g MgSO4 • 7H20 and distilled water to i 000 ml. 2.2. Assay procedure To round-bottom test tubes (10 mm diameter), containing 1.2 ml of washed cells suspended in PUM buffer, were added various volumes of test hydrocar- 30 bon (n-hexadecane, n-octane or p-xylene). Following 10 min preincubation at 30°C, the mixtures were agitated uniformly on a Thermolyne Maxi Mix (Sybron) for 120 s. After allowing 15 min for the hydrocarbon phase to rise completely, the aqueous phase was carefully removed with a Pasteur pipette and transferred to a 1 ml cuvette. Light absorbance was determined at 400 nm, using a Gilford Model 240 spectrophotometer. 3. Results The basic experiment used to measure bacterial hydrophobicity is shown qualitatively in Fig. 1. Hexadecane was layered onto a turbid aqueous suspension of Acinetobacter RAG-1 (Fig. 1, left tube) and then mixed for 120 s; on standing a clear bottom layer and a "creamy" upper layer formed (Fig. 1, middle tube). Microscopic examination of the upper layer revealed an oil-in-water emulsion consisting of hexadecane droplets covered with patches of bacteria (Fig. 2). Decrease in absorbance of the lower aqueous phase was used as a measure of cell surface hydrophobicity. Fig. 1. Adherence of Acinetobacter RAG-1 to hydrocarbon. Hexadecane was added to an aqueous suspension of Acinetobacter RAG-1 (left). After mixing for 120 s and allowing to stand, adherent cells rose with the hydrocarbon, forming a "creamy" upper layer and a clear aqueous phase (middle). Addition of isopropanol to a final concentration of 5% (v/v) results in breakage of the emulsion and release of cells back into the aqueous phase (right). The cell-stabilized emulsions did not break even after several days; however, addition of 5% (v/v) isopropranol brought about an immediate coalescence of droplets and release of cells into the aqueous phase (Fig. 1, right tube). Results illustrating the adherence of various bacteria to the test hydrocarbons are presented in Figs. 3 - 6 . Fig. 3 compares the affinity ofE. coli with those of two Acinetobacter strains. E. coli B showed no significant affinity towards hexadecane or octane and little or no affinity towards xylene. Similar results (not presented) were obtained with M. lysodeikticus, E. aerogenes, B. subtilis and P. aeruginosa. While Acinetobacter RAG-1 showed high affinity towards all three hydrocarbons, Acinetobacter BD exhibited higher affinity for octane and xylene than for hexadecane. More than 97% of the RAG-1 and over 99% of the BD strain cells were removed from the aqueous phase by 0.2 ml octane. The affinities of two species of Staphylococci towards the test hydrocarbons are presented in Fig. 4. Although S. albus did not adhere to hexadecane or xylene, S. aureus exhibited a high affinity towards all the test hydrocarbons. Although S. marcescens grown to early exponential phase showed relatively low affinity towards hexadecane and xylene and none towards octane, pink-pigmented early stationary phase cells adhered strongly to all three hydrocarbons (Fig. 5). Loss of oligosaccharide components on the outer surface ofE. coli rough mutant J-5 [19] was accompanied by a significant increase in its affinity towards Fig. 2. Phase micrograph showing RAG-1 cells adhering to hexadecane droplet following mixing. 825X. 100 ;; 2 75 r;: : 50 : 4 $? 25 0 0.1 HEXADECANE 0.2 <ml) 0.1 0.2 OCTANE 0.1 CmI) XYLENE 0.2 (ml) Fig. 3.0, E. coli B (1.562); a, Acinetobacter RAG-l (1.441); l,Acinezobwter BD (0.632). I- I loo I 1 I loo 0 ” 0 fj 2 75 75 50 50 25 25 1;: p a 4 $ I O 0.2 0.1 HEXADECANE <ml> 0.1 OCTANE 0.2 Cml) I I 0.1 XYLENE I 0.2 (ml> Fig. 4. 0, Staphylococcusalbus (0.716); 0, Staphylococcus aureus (1.175). - 0.1 HEXADECANE 0.2 (ml) 0.1 0.2 OCTANE Fig. 5. 0, Serratzizmareescens, early logarithmic phase (1.120); l, Cml) Senatia marcescens, 0.2 01 XYLENE (ml) early stationary phase (1.132). 25 32 I 1 I 100 1 u -O=C I I .3 O I r I O - 75 1OO 75 Z O < -- rn n,- O if) 50 50 <[ ~ 25 - I O I I HEXADECANE I I I 0.2 O. 1 (ml) I O.1 OCTANE I 0.2 (ml) I I I O.1 XYLENE 25 I 0.2 (,hi) Fig. 6. o, E. coli K-12 (1.558); o, E. coli J-5 (1.408). Figs. 3-6. Aff'mityof bacteria towards hydrocarbon as a function of hydrocarbon volume. Aqueous bacterial suspensions were mixed with varying volumes of hexadeeane, octane and xylene as described in Materials and Methods. Results are expressed as percentage of the initial absorbanee (A 400) of the aqueous suspension as a function of hydrocarbon volume. Initial absorbance is shown in parentheses. each of the three test hydrocarbons, as compared with E. coli K-12 (Fig. 6). 4. Discussion A simple quantitative method has been described for studying the outer cell surface of bacteria, based on the affinity of these cells for liquid hydrocarbons. Large differences in the aftrmities of various bacteria for hydrocarbons have been demonstrated using this technique. Previous observations have suggested that the ability to adhere to bulk hydrocarbon is a characteristic of hydrocarbon-degrading microorganisms [ 11 ]. We have shown that S. aureus and early stationary phase S. marcescens cells adhere to a variety of fiquid hydrocarbons despite their inability to degrade them. Thus, the ability of bacterial cells to adhere to hydrocarbon is not restricted to hydrocarbon-degrading bacteria. Moreover, both Acinetobacter strains adhered strongly to octane and xylene, hydrocarbons which they are incapable of metabolizing. This indicates that the adherence of these hydrocarbondegrading microorganisms is not limited to metabolizable hydrocarbons, and suggests that direct contact between Acinetobacter cells and bulk hydrocarbon is due to general hydrophobic interactions rather than specific recognition of the substrate. P. aeruginosa PAS 279 did not adhere significantly to the test hydrocarbons under the conditions studied, despite its ability to grow on hexadecane. This result suggests that affmity for hydrocarbon may vary among hydrocarbon-degrading bacteria. Further studies on the relationship between adherence to hydrocarbon and growth of hydrocarbon-degrading bacteria are underway. The large increase in hydrophobicity observed in S. marcescens with increasing age of the ceils agrees with previous reports [8,13]. The small, but significant increase in the affmity of the rough mutant E. coli J-5 as compared with that ofE. coli K-12 indicates the increased hydrophobicity of the rough mutant, presumably due to the increased exposure of inner core regions of the LPS layer. Similar results have been obtained with rough mutants of Salmonella typhimurium [1,14]. The method described here may prove useful in enabling the separation of certain cell mixtures, and in separating hydrophobic cell components. In addition, this technique may provide a means of enriching for and selecting mutants which are altered in their 33 surface hydrophobicity. A search for such mutants is currently in progress, with a view towards investigating the relationship between the metabolism o f hydrophobic substrates and modifications in cell surface hydrophobicity. Acknowledgements We thank Z.L. Zosim, L. Goldstein and E.A. Bayer for stimulating discussions and constructive criticism. References [ 1] Curmingham, R.K., S6derstriSm, T.O., Gillman, C.F. and Van Oss, C.J. (1975) Immunol. Commun. 4,429-442. [2] Peters, L., Andiiker, L., Edebo, L., Stendahl, O. and Tagesson, C. (1977) Acta Pathol. Mierobiol. Scand. Sect. B 85,308-316. [3] Smyth, C.J., Jonsson, P., Olsson, E., S~derlind, O., Rosengren, J., Hjert6n, S. and Wadstr6m, T. (1978) Infect. Immun. 22, 462-472. [4] Dexter, S.C., Sullivan, J.D., Jr., Williams, J., III and Watson, S.W. (1975) J. Gen. Microbiol. 30, 298-308. [5] Fletcher, M. and Loeb, G.I. (1979) Appl. Environ. Microbiol. 37, 67-72. [6] Marshall, K.C. and Cruickshank, R.H. (1973) Arch. Mikrobiol. 91, 29-40. [7] Stanley, S.O. and Rose, A.H. (1967) J. Gen. Microbiol. 48,9-23. [8] Blanchard, D.C. and Syzdek, L.D. (1978) Limnol. Oceanogr. 23,389-400. •[9] Miura, Y., Okazaki, M., Hamada, S.-I., Murakawa, S.-I and Yugen, R. (1977) Biotech. Bioeng. 19, 701-714. [10] McLee, A.G. and Davies, S.L. (1972) Can. J. Microbiol. 18,315-319. [ 11 ] Kennedy, R.S., Finnerty, W.R., Sudarsanan, K. and Young, R.A. (1975) Arch. Microbiol. 102, 75-83. [12] K~ippeli,O. and Fiechter, A. (1976) Biotech. Bioeng. 18,967-974. [13] KjeUeberg, S., Lagercrantz, C. and Larsson, Th. (1980) FEMS Microbiol. Lett. 7, 41-44. [14] Magnusson, K.-E. and Johansson, G. (1977) FEMS Microbiol. Lett. 2, 225-228. [ 15 ] Magnusson, K.-E., Stendahl, O., Stjernstr6m, I. and Edebo, L. (1978) Acta Pathol. Microbiol. Scand. Sect. B 86,113-120. [17 ] Reisfeld, A., Rosenberg, E. and Gutnick, D. (1972) Appl. Microbiol. 24,363-368. [18] Horowitz, A., Gutnick, D. and Rosenberg, E. (1975) Appl. Microbiol. 30, 10-19. [19] Braude, A.I. and Douglas, H. (1972) J. Immunol. 108, 505-512.