Department of Biology, The University, D-78457 Konstanz, Germany
Correspondence
Alasdair Cook
alasdair.cook{at}uni-konstanz.de
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ABSTRACT |
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INTRODUCTION |
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Analysis of the sequence of the genome of Rhodopseudomonas palustris CGA009 revealed candidate genes to encode TauR and TauXY but not Xsc, and the organism was found to utilize taurine as a sole source of combined nitrogen under oxic and anoxic conditions (Denger et al., 2004b). R. palustris generated sulfoacetate quantitatively, and a sulfoacetaldehyde dehydrogenase was observed. It was concluded that taurine could be a major source of environmental sulfoacetate and that an exporter of sulfoacetate must be present in R. palustris (Denger et al., 2004b
).
We thus proceeded to test whether taurine was commonly used by bacteria as a sole source of combined nitrogen concomitant with release of sulfoacetate. The results showed that release of an organosulfonate was common under these conditions, but that sulfoacetate was never observed and that sulfoacetaldehyde was often the product which was released.
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METHODS |
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Organisms, growth media, growth conditions, cell disruption and enrichment cultures.
R. palustris CGA009 was grown in 1 l cultures in 5 l Erlenmeyer flasks which were shaken at 30 °C in the dark. The growth medium was a 50 mM potassium phosphate buffer, pH 7·2, which contained 0·25 mM MgSO4, 10 mM DL-malate, 2 mM taurine and trace elements (Thurnheer et al., 1986), as well as a seven-vitamin mixture (Pfennig, 1978
), which was added after autoclaving. Cultures were inoculated (1 %) from a growing culture in the same medium and incubated until the turbidity indicated a protein concentration of 10 mg protein l1 (78 days), when taurine was added to a final concentration of 2 mM. The culture was harvested 24 h later (30 000 g, 15 min, 4 °C), washed in 50 mM potassium phosphate buffer, pH 7·2, containing 5 mM MgCl2 and used immediately or stored frozen (20 °C). Cells at about 250 mg wet weight (ml wash buffer)1 were disrupted by eight passages through a chilled French pressure cell at 140 MPa. Debris was removed by centrifugation (20 000 g, 3 min, 4 °C). The specific activity of the sulfoacetaldehyde dehydrogenase in the resulting crude extract was about 1·3 mkat (kg protein)1.
The growth medium described above was used with minor modifications for enrichment cultures (5 ml in 50 ml tubes) with 2 mM taurine or 2 mM 2-aminoethanephosphonate (ciliatine) as sole source of nitrogen. The carbon source in different sets of enrichments was (i) 5 mM glucose plus 10 mM glycerol plus 10 mM DL-malate, (ii) 5 mM glucose plus 10 mM glycerol plus 10 mM succinate, (iii) 10 mM succinate, (iv) 5 mM adipate, (v) 5 mM benzoate, (vi) 20 mM acetate or (vii) 10 mM DL-lactate. Each set of enrichments was accompanied by controls containing either no added combined nitrogen or 2 mM NH4Cl instead of taurine or 2-aminoethanephosphonate. The inocula were derived from the aeration tank of the wastewater treatment plant in Konstanz, from forest soil or from agricultural soil; in each case, the material was shaken with several batches of 50 mM potassium phosphate buffer, pH 7·2, and decanted or centrifuged to remove any soluble nitrogenous material and inorganic solids. Inoculation left the cultures visibly turbid. Incubation for 13 days showed negligible growth in the negative controls and similar growth in both experiments and positive controls. After three passages in fresh medium, all cultures were plated on complex medium (LuriaBertani medium; Gerhardt et al., 1994) and common colony morphologies were picked to fresh selective medium. A culture was considered pure when three cycles of homogeneous plates and growth in selective medium were attained. Organisms isolated to utilize 2-aminoethanephosphonate were transferred to medium suitable to detect the possible release of phosphate from the source of nitrogen. The two alterations were to buffer with 50 mM MOPS/NaOH, pH 7·2 (instead of phosphate), and to add 0·5 mM potassium phosphate as a source of phosphorus. Outgrown cultures were analysed for the presence and concentrations of 2-aminoethanephosphonate and inorganic phosphate.
Growth of Acinetobacter calcoaceticus SW1 was followed in 50 ml cultures in 300 ml Erlenmeyer flasks shaken in a water bath at 30 °C. The growth medium was as described for R. palustris (above), except that the carbon source was 5 mM adipate and no vitamins were needed. Samples (2 ml) were taken at intervals to measure turbidity and to determine the concentrations of protein, taurine and other organosulfonates and the ammonium and sulfate ions. When larger amounts of cells were required for enzyme assays, 1 l cultures in 5 l Erlenmeyer flasks were used. Rupture of harvested cells of A. calcoaceticus was essentially as described above, but only three passages through the French pressure cell were required.
Rhodococcus opacus ISO-5 was grown with taurine as the sole source of carbon and energy for growth, as described elsewhere (Denger et al., 2004a). Crude extract was generated (Denger et al., 2004a
) to serve as positive controls for some enzyme assays.
Enzyme assays.
Taurine : pyruvate aminotransferase was assayed discontinuously in 1 ml reaction mixtures maintained at 37 °C in a water bath. The reaction mixture contained 50 µmol Tris/HCl, pH 9·0, 5 µmol MgCl2, 5 µmol taurine, 10 µmol pyruvate, 100 nmol pyridoxal 5'-phosphate and 0·3 mg protein, with which the reaction was started. Samples were taken at intervals and taurine and alanine were determined. Crude extract from Rhodococcus opacus ISO-5 was used as a positive control. The reaction mixture was adapted to assay any 2-oxoglutarate-dependent transamination (EC 2.6.1.55) by replacing pyruvate with 2-oxoglutarate. Alanine dehydrogenase was assayed by a standard method (Laue & Cook, 2000). Taurine dehydrogenase was assayed with either dichlorophenol indophenol (DCPIP) or beef-heart cytochrome c as the electron acceptor (Brüggemann et al., 2004
). Sulfoacetaldehyde dehydrogenase was assayed photometrically as the reduction of NAD+ at 365 nm. The reaction mixture (1 ml) at about 22 °C contained 50 µmol Tris/HCl, pH 9·0, 5 µmol MgCl2, 4 µmol NAD+, 400 nmol sulfoacetaldehyde (as the bisulfite addition complex) and 0·10·3 mg protein, with which the reaction was started.
Analytical methods.
Growth was followed as turbidity (an OD580 of 1·0 was taken as equivalent to 250 mg protein l1) and quantified as protein in a Lowry-type reaction (Cook & Hütter, 1981). Reversed-phase chromatography was used to quantify taurine, 2-aminoethanephosphonate, alanine or glutamate after derivatization with 2,4-dinitrofluorobenzene (Laue et al., 1997
) or free sulfoacetaldehyde (presumably in equilibrium with the added bisulfite addition complex) after derivatization with 2-(diphenylacetyl)indane-1,3-dione-1-hydrazone (Cunningham et al., 1998
). Sulfate was determined turbidimetrically as a suspension of BaSO4 (Sörbo, 1987
). Ammonium ion was assayed colorimetrically by the Berthelot reaction (Gesellschaft Deutscher Chemiker, 1996
). Sulfoacetate or inorganic phosphate was quantified by ion chromatography with suppression (Denger et al., 2004b
). The identity of sulfoacetaldehyde was confirmed by matrix-assisted, laser-desorption ionization time-of-flight mass spectrometry (MALDI-TOF-MS) in the negative ion mode with a matrix of 4-hydroxy-
-cyanocinnamic acid; these determinations were done under contract by K. Hollemeyer (Technical Biochemistry, Saarland University, Saarbrücken, Germany). Values of apparent Km (Kmapp) were derived by hyperbolic curve-fitting as cited elsewhere (Ruff et al., 2003
). SDS-PAGE and staining were done by standard methods (Laemmli, 1970
). Standard methods were used to establish Gram reaction, catalase and cytochrome c oxidase activity (Gerhardt et al., 1994
). A 1·4 kbp fragment of the 16S rRNA gene of several organisms under study was amplified by PCR, sequenced and analysed essentially as described elsewhere (Brüggemann et al., 2004
); primers 16S-27f and 16S-1492r (Weisburg et al., 1991
) were used.
Determination of sulfoacetaldehyde with sulfoacetaldehyde dehydrogenase from R. palustris.
This enzyme was determined to have Kmapp values of 26 µM for sulfoacetaldehyde and 92 µM for NAD+, as well as 12 mM for NADP+. NAD+ was obviously the physiological electron acceptor, and the high affinity for sulfoacetaldehyde made the catalyst a candidate for a specific (see Results) and sensitive enzymic determination of sulfoacetaldehyde (Bergmeyer, 1983). The enzyme could be enriched to about 28 mkat (kg protein)1 by two separations at different pH values on an anion-exchange column, but this preparation still contained about 10 proteins in similar amounts (SDS-PAGE). We found that the soluble fraction was adequate to measure sulfoacetaldehyde in cell-free culture medium of A. calcoaceticus, provided the medium did not contain malate, the carbon source for growth of R. palustris. It was also apparent that both the free sulfoacetaldehyde and the bisulfite addition complex were substrates for the enzyme. The reaction mixture (in 1·0 ml) contained 50 µmol Tris/HCl, pH 9·0, 5 µmol MgCl2, 4 µmol NAD+ and
60 nmol sulfoacetaldehyde, and the value of A365 was confirmed to be stable. The enzyme preparation (0·3 mg protein in 10 µl) was then added and the increase in the value of A365 was followed. The reaction was complete in about 5 min. It was possible to do the reaction with about 1 mM sulfoacetaldehyde in the test and a long reaction time. Under these conditions, 1 mM sulfoacetate was detected by ion chromatography after the enzyme reaction had ceased. We thus had confirmation that the required reaction was being catalysed.
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RESULTS |
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The organosulfonate generated by the other 12 isolates was at first unknown (Table 1); it was later shown to be isethionate (2-hydroxyethanesulfonate) (K. Styp von Rekowski, unpublished data). Three groups of organisms seemed to generate the compound. The largest group (eight organisms) was represented by strain TauN1, whose 16S rRNA gene was sequenced. This showed 99·8 % identity of position with the corresponding gene in the type strain of Klebsiella oxytoca, and its taxonomic and physiological properties supported this conclusion (Table 1
). K. oxytoca TauN1 was deposited with the DSMZ as DSM 16963. A similar train of argument identified two strains as Pseudomonas putida, represented by strain TauN2 (Table 1
). Two organisms were not identified.
A series of enrichment cultures yielded 33 isolates, all bacteria, able to utilize 2-aminoethanephosphonate as the sole source of combined nitrogen for growth. Each isolate utilized 2-aminoethanephosphonate quantitatively, but, in all cases, the phosphonate moiety was recovered quantitatively after growth as phosphate. There was no need to postulate an organophosphonate as an excreted product of metabolism of 2-aminoethanephosphonate.
Confirmation of the identity of sulfoacetaldehyde, its stability and reproducible quantification
We presumed that sulfoacetaldehyde would be labile, especially with the electron-withdrawing sulfonate moiety on the carbon atom adjacent to the aldehyde group. However, boiling the putative sulfoacetaldehyde in spent growth medium of A. calcoaceticus did not generate detectable sulfoacetate or alter the concentration of sulfoacetaldehyde, and the addition of several microlitres of fuming nitric acid also failed to generate sulfoacetate. The putative sulfoacetaldehyde was stable in growth medium.
Analysis of portions of growth medium by MALDI-TOF-MS in the negative-ion mode showed that neither negative controls nor taurine ([M1]=124) contained sulfoacetaldehyde, whereas spent taurine medium contained no detectable taurine but a product ([M1]=123) which confirmed the presence of sulfoacetaldehyde (M=124). The identification, however, offered no quantification of the product.
We generated a third, enzymic, determination of the product using sulfoacetaldehyde dehydrogenase (EC 1.2.1.-) from R. palustris (Fig. 2). The quantification was thus provided by the molar absorption coefficient of NADH, which has been in use for over 60 years, and we confirmed that the purity of our salt-contaminated synthetic material is about 53 % (see Denger et al., 2001
). When tested at higher substrate concentrations, the enzymic formation of the expected concentration of sulfoacetate could be determined by ion chromatography. We found no compound which interfered with the reaction (acetaldehyde, phosphonoacetaldehyde, glyoxylate, glycolaldehyde, propionaldehyde and succinate semialdehyde were tested). Whereas the standard curve of the hydrazone derivative suggested that 2 mol sulfoacetaldehyde (mol taurine)1 were formed (see above), the enzymic determination indicated unit stoichiometry (see below). We presume that our attempts to convert the bisulfite addition complex of sulfoacetaldehyde to the free aldehyde were inadequate (as observed elsewhere; Gritzer et al., 2003
), because Cunningham et al. (1998)
specify that the method only derivatizes the free aldehyde.
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DISCUSSION |
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The release of sulfoacetate by R. palustris was rationalized as both the NADH-yielding oxidation of the aldehyde to the acid and the irreversible conversion of the reactive aldehyde to the non-toxic acid (Denger et al., 2004b). The observation that sulfoacetaldehyde is neither labile nor especially reactive weakens the latter argument. The fact that many organisms need not carry out the oxidation at all, or even carry out a reduction to isethionate (K. Styp von Rekowski, unpublished data), also weakens the former argument. We suspect that the release of sulfoacetaldehyde into the environment is widespread, because bacteria able to dissimilate the compound have been observed (Lie et al., 1996
).
The pathway by which strain SW1 obtains the ammonium ion is short, with only three steps, two of which are transporters (Fig. 1). The specific activity of the metabolic enzyme, also presumably membrane-bound (Brüggemann et al., 2004
), was only 10 % of that required for growth, but the natural electron acceptor was unavailable. Both ATP-binding-cassette transporters [TC 3.A.1.17.1] and tripartite, ATP-independent transporters [TC 2.A.56.4.1] are presumed to be involved with the uptake of taurine in different organisms (Brüggemann et al., 2004
), but the nature of the sulfoacetaldehyde exporter is unknown, as is that for sulfoacetate in R. palustris (Denger et al., 2004b
). The putative sulfate exporter in e.g. Paracoccus pantotrophus is apparently novel (Rein et al., 2005
), so we hope to identify the genes involved in transport and oxidation of taurine and export of sulfoacetaldehyde, as well as the regulation, by characterizing taurine dehydrogenase, the corresponding gene(s) and its surroundings and developing testable, sequence-derived predictions.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Received 17 December 2004;
revised 20 January 2005;
accepted 21 January 2005.
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