Ecole Nationale Supérieure des Industries Agricoles et Alimentaires, Laboratoire de Microbiologie Industrielle, 91744 Massy Cedex, France1
Institut Français du Pétrole, Division Chimie et Physico-chimie Appliquées, 92852 Rueil-Malmaison Cedex, France2
Author for correspondence: Murielle Bouchez-Naïtali. Tel: +33 1 69 93 51 40. Fax: +33 1 69 93 50 84. e-mail: naitali{at}ensia.inra.fr
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ABSTRACT |
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Keywords: alkane degradation, electrolytic respirometry, flocculating strains, Rhodoccocus sp., uptake kinetics
Abbreviations: HMN, 2,2,4,4,6,8-heptamethylnonane
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INTRODUCTION |
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Whatever the mechanisms involved, a point which is still only partially understood at the present time is the kinetics of long-chain alkane biodegradation. Studies focused on the kinetics of growth of yeasts on n-alkanes were conducted in the seventies and models were proposed to account for the complex kinetic patterns observed (Blanch & Einsele, 1973 ; Erickson et al., 1969
, 1970
; Erickson & Humphrey, 1969
; Gutierrez & Erickson, 1977
; Moo-Young & Shimizu, 1971
; Verkooyen & Rietma, 1980a
, b
; Yoshida & Yamane, 1974
). However, with other alkane-degrading micro-organisms, the interest was essentially oriented towards the production of biosurfactants and kinetic studies are scarce. In a recent work, we analysed the relative distribution of the modes of substrate uptake in a large series of bacterial strains degrading hexadecane (C16) (Bouchez-Naïtali et al., 1999
). In the present work, we studied in detail the kinetics of degradation of C16 by four Rhodococcus strains that had been identified as using direct interfacial uptake on the basis of the absence of biosurfactant production. Cell flocculation was found to constitute an important characteristic of the cultures of these strains on C16. The kinetic patterns of degradation were investigated in relation to the various phenomena involved in C16 uptake.
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METHODS |
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Strains.
Four Rhodococcus equi strains were employed: NapRu1, HdGe1, PyrGe1 and Fo2. They were isolated from soils as previously described (Bouchez-Naïtali et al., 1999 ) and were able to use C16 as sole source of carbon and energy.
Cultures in flasks.
Cultures were carried out in 250 ml flasks containing 100 ml medium and incubated on a rotary shaker (160 r.p.m.) at 30 °C. They were used as inocula for fermentation work and kinetic studies in the conditions described below. Flask cultures were also used for hydrophobicity determinations. In this case, the medium was MSM3 and the carbon source was C16 (alone or in the hydrophobic solvents studied).
Growth on hexadecane in laboratory fermenters.
Batch cultures were performed in 2 litre fermenters (Inceltech), in 1·47 litre MSM4 with 30 ml C16 as sole source of carbon and energy. The inoculum was grown in the same medium containing succinate as carbon source. To remove the residual carbon source from the inoculum, 30 ml of culture was centrifuged (10000 g, 10 min) and the cells were resuspended in 30 ml fresh MSM4 before inoculation of the fermenters. The fermentation conditions were: temperature, 30 °C; stirring speed, 500 r.p.m.; aeration, 1 vol. vol.-1 min-1. The pH was recorded but not regulated. Experiments were run in duplicate.
Kinetic studies by respirometry.
Time courses of oxygen consumed during growth on C16 were recorded using a sensitive respirometer (D12-S Sapromat, Voith) consisting of 12 conical flasks (500 ml) placed in a water bath maintained at 30 °C (Bouchez et al., 1997a ). Determination of electrolytically produced oxygen supplied to the cultures at constant pressure allowed quasi-continuous recording of oxygen consumption. The stirring speed being uniform (300 r.p.m.), two types of stirrers were employed in order to modify agitation efficiency: triangular-section bars (14x55 mm) in most cases, and smaller, round-section bars (6x20 mm) when specified. Culture flasks containing 237·5 ml MSM3 were inoculated with 12·5 ml of a preculture on MSM3 supplemented with succinate prepared as above. C16 was added alone or as a solution in a hydrophobic solvent, either 2,2,4,4,6,8,8-heptamethylnonane (HMN) or silicone oil 47V20 (Prolabo). Both solvents were non-toxic and non-water-soluble, and were not degraded by the strains. As indicated in the results, experiments were run several times but individual recordings are presented in the figures. Oxygen consumption blanks (inoculated flasks containing hydrophobic solvent without C16 and uninoculated flasks containing C16 and hydrophobic solvent) were run simultaneously. No significant oxygen consumption was detected in these blanks.
Parameters describing the kinetics obtained in different conditions were compared using analysis of variance. When considered as significantly different (P<0·05) or very significantly different (P<0·01), Students t-tests were performed.
Analyses.
Analyses were performed as described by Bouchez-Naïtali et al. (1999 ). Interfacial tension against C16 and surface tension were determined at 30 °C on filtered supernatant fluids of cultures by, respectively, the De Nouy ring and Wilhemy plate methods, using a K-12 tensiometer (Krüss). Each result was the mean of ten determinations and standard deviation was less than 0·4 mN m-1. Cell hydrophobicity was measured by the bacterial adherence to hydrocarbon test according to a method adapted from Rosenberg et al. (1980)
. Hydrophobicity was measured with respect to the hydrophobic phase used in the cultures: C16 when employed alone, or HMN or silicone oil. For each sample, three independent determinations were made and the standard deviation was within 5%. Residual nitrate concentration was determined in culture supernatant fluids using a commercial kit (Boehringer Mannheim). Glycosides were evaluated on supernatant fluids of cultures by the colorimetric method of Dubois et al. (1956)
using glucose as standard. Each result was the mean of three determinations and the standard deviation was within 5%.
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RESULTS |
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DISCUSSION |
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A different situation, however, prevailed in other cases. Flocculation of the cultures was always observed. Here, the model for interfacial uptake can account for the existence of a first phase of exponential growth. Higher solvent volumes did not increase the values of the specific growth rates, but this can be explained by the strong adsorption of the bacteria to their substrate, a well-recognized property of many alkane-degrading micro-organisms (Rosenberg, 1992 ). The interfacial uptake model, however, did not account for the fact that the exponential phase was very short and that the degradation rate during the subsequent phase of linear growth was not dependent on the solvent volume. The biological flocs formed were likely to be responsible for these characteristics. Indeed, a similar situation was discussed by Blanch & Einsele (1973)
in the case of growth of the yeast Candida tropicalis on C16. In flocs, contact with hydrocarbon or dissolved oxygen is rate-limiting for cells inside the aggregates. Interfacial substrate uptake in this case is no longer controlled by the parameters that modify the aqueous and solvent interface (such as the volume of the solvent phase) but by those that modify floc formation, since floc size determines cell access to the substrate. In such a case, stirring is a key parameter because it controls floc size, higher stirring rates promoting smaller flocs and thus higher transfer rates. This point is in agreement with the slower degradation kinetics observed in conditions of lower stirring efficiency. Cell aggregation can explain the onset of the linear phase. The tendency of the cells to agglomerate in flocs increases with increasing biomass. The transition occurs when agglomeration predominates over the turbulence of the medium, which breaks the floc particles. Growth in cell aggregates accounts for the long linear phase of degradation observed with these cultures. Differences in degradation rates during this phase appear to be essentially related, for a given rate of stirring, to differences in cell aggregation properties among strains. The floc formation observed here in almost all cases with Rhodococcus strains leads us to suggest that the analysis given by Blanch & Einsele (1973)
for C. tropicalis may be commonly applicable to micro-organisms growing on long-chain alkanes by strictly interfacial uptake. Actually, the propensity to aggregate appears quite significant among alkane-degrading micro-organisms and is obviously related to the high cell hydrophobicity of the strains concerned. Non-flocculent PyrGe1 cultures corresponded to the lowest values of cell hydrophobicity. In fact, because of their hydrophobicity, all our strains except HdGe1 exhibited some cell agglomeration in cultures on soluble substrates such as succinate, glucose or glycerol, and the growth kinetics of strain NapRu1 on glucose did not show a regular exponential growth pattern. However, growth of these strains on hexadecane clearly promoted cell aggregation.
Thus, the studies confirmed the diagnosis of an interfacial uptake mechanism for the four strains studied, but showed that aggregation of these hydrophobic cells was a major parameter that determined the degradation kinetics. For one strain, PyrGe1, interfacial uptake was either controlled or not by floc formation depending the organic phase used. The results help to clarify the hitherto insufficiently understood characteristics of interfacial uptake of alkanes. The phenomenon of flocculation has significant ecological implications. It is also important for biotechnological applications. The case of micro-organisms producing biosurfactants is quite different, not only because biosurfactants promote the dispersion of the hydrophobic substrate as emulsions or as micelles, but also because they affect cell hydrophobicity and bacterial adherence to hydrocarbons, as shown by Zhang & Miller (1994) for Pseudomonas aeruginosa.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Bouchez, M. (1995). La biodégradation des hydrocarbures aromatiques polycycliques: métabolisme de substrats non conventionnels. Doctorate Thesis, Ecole Nationale des Industries Agricoles et Alimentaires, Massy, France.
Bouchez, M., Blanchet, D., Besnainou, B., Leveau, J. & Vandecasteele, J.-P. (1997a). Kinetic studies of biodegradation of insoluble compounds by continuous determination of oxygen consumption. J Appl Microbiol 82, 310-316.[Medline]
Bouchez, M., Blanchet, D. & Vandecasteele, J.-P. (1997b). An interfacial uptake mechanism for the degradation of pyrene by a Rhodococcus strain. Microbiology 143, 1087-1093.
Bouchez-Naïtali, M., Rakatozafy, H., Marchal, R., Leveau, J. Y. & Vandecasteele, J.-P. (1999). Diversity of bacterial strains degrading hexadecane in relation to the mode of substrate uptake. J Appl Microbiol 86, 421-428.[Medline]
Boulton, C. A. & Ratledge, C. (1984). The physiology of hydrocarbon-utilizing microorganisms. In Enzyme and Fermentation Biotechnology , pp. 11-77. Edited by A. Wieseman. New York:Wiley.
Desai, J. D. & Banat, I. M. (1997). Microbial production of surfactants and their commercial potential. Microbiol Mol Biol Rev 61, 47-64.[Abstract]
Dubois, M., Gilles, K. A., Hamilton, J. K., Rebers, P. A. & Smith, T. (1956). Colorimetric method for determination of sugars and related substances. Anal Chem 28, 350-356.
Dunn, I. J. (1968). An interfacial kinetics model for hydrocarbon oxidation. Biotechnol Bioeng 10, 891-894.
Erickson, L. E. & Humphrey, A. E. (1969). Growth models of cultures with two liquid phases. II. Pure substrate in dispersed phase. Biotechnol Bioeng 11, 467-487.
Erickson, L. E., Humphrey, A. E. & Prokop, A. (1969). Growth models of cultures with two liquid phases. I. Substrate dissolved in dispersed phase. Biotechnol Bioeng 11, 440-466.
Erickson, L. E., Fan, L. T., Shah, P. S. & Chen, M. S. K. (1970). Growth models of cultures with two liquid phases. IV. Cell adsoption, drop size, and batch growth. Biotechnol Bioeng 12, 713-746.[Medline]
Gutierrez, J. R. & Erickson, L. E. (1977). Hydrocarbon uptake in hydrocarbon fermentations. Biotechnol Bioeng 19, 1331-1339.[Medline]
Haferburg, D., Hommel, R., Claus, R. & Kleber, H. P. (1986). Extracellular microbial lipids as biosurfactants. Adv Biochem Eng Biotechnol 33, 53-93.
Hommel, R. K. (1994). Formation and function of biosurfactants for degradation of water-insoluble substrates. In Biochemistry of Microbial Biodegradation , pp. 63-87. Edited by C. Ratledge. Dordrecht:Kluwer.
Moo-Young, M. & Shimizu, T. (1971). Hydrocarbon fermentations using Candida lipolytica. II. A model for cell growth kinetics. Biotechnol Bioeng 13, 761-778.[Medline]
Rosenberg, E. (1992). The hydrocarbon-oxidizing bacteria. In The Procaryotes , pp. 446-459. Edited by A. Balows, H. G. Trüper, M. Dworkin, W. Harder & K.-H. Schleifer. New York:Springer.
Rosenberg, M., Gutnick, D. & Rosenberg, E. (1980). Adherence of bacteria to hydrocarbons: a simple method for measuring cell surface hydrophobicity. FEMS Microbiol Lett 9, 29-33.
Singer, M. E. & Finnerty, W. R. (1984). Microbial metabolism of straight-chain and branched alkanes. In Microbial Metabolism of Straight-Chain and Branched Alkanes , pp. 1-60. Edited by R. M. Atlas. New York:Macmillan.
Verkooyen, A. H. M. & Rietma, K. (1980a). Growth of yeast on n-alkanes. I. Stochastic model. Biotechnol Bioeng 22, 571-595.
Verkooyen, A. H. M. & Rietma, K. (1980b). Growth of yeast on n-alkanes. III. Batch experiments. Biotechnol Bioeng 22, 615-637.
Westgate, S., Bell, G. & Halling, P. J. (1995). Kinetics of uptake of organic liquid substrates by microbial cells: a method to distinguish interfacial contact and mass-transfer mechanisms. Biotechnol Lett 17, 1013-1018.
Yoshida, F. & Yamane, T. (1974). Continuous hydrocarbon fermentation with colloidal emulsion feed. A kinetic model for two-liquid phase culture. Biotechnol Bioeng 16, 635-657.
Zhang, Y. & Miller, R. M. (1994). Effect of a Pseudomonas rhamnolipid biosurfactant on cell hydrophobicity and biodegradation of octadecane. Appl Environ Microbiol 60, 2101-2106.[Abstract]
Received 24 November 2000;
revised 12 April 2001;
accepted 1 June 2001.