1 Wageningen Centre for Food Sciences, PO Box 557, 6700AN Wageningen, The Netherlands
2 Wageningen University, Laboratory of Microbiology, Dreijenlaan 2, 6703HA Wageningen, The Netherlands
3 Wageningen University, Food and Bioprocess Engineering Group, PO Box 8129, 6700EV Wageningen, The Netherlands
Correspondence
George J. G. Ruijter
g.j.g.ruijter{at}lumc.nl
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ABSTRACT |
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Present address: Metabolic Diseases Laboratory, Centre for Human and Clinical Genetics, Leiden University Medical Centre, PO Box 9600, 2300RC Leiden, The Netherlands.
Present address: FGT Consultancy, PO Box 396, 6700AJ Wageningen, The Netherlands.
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INTRODUCTION |
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At low aw, micro-organisms try to prevent water loss from the cells by accumulating compatible solutes, such as ions, amino acids or polyols. Fungi generally accumulate polyols such as glycerol, erythritol, arabitol or mannitol. Very high levels, up to several molar, can be accumulated in the cells, but polyols are also secreted to the environment (Witteveen & Visser, 1995; Gutierrez-Rojas et al., 1995
). For SmF, polyol accumulation has been described for a number of species, such as glycerol production by Aspergillus niger (Witteveen & Visser, 1995
) and Aspergillus wentii (El-Kady et al., 1994
), glycerol and erythritol production by Aspergillus nidulans (Beever & Laracy, 1986
; Redkar et al., 1995
), and production of a mixture of polyols by Aspergillus repens (Kelavkar & Chhatpar, 1993
) and Aspergillus flavus (Mellon et al., 2002
). Much less is known about polyol production at low aw in SFF. A. niger cultured at low aw on Amberlite as an inert support produced glycerol and erythritol (Gutierrez-Rojas et al., 1995
), whereas Aspergillus ochraceus accumulated a mixture of polyols on agar media (Ramos et al., 1999
).
The metabolic pathways for polyol biosynthesis in fungi have only been elucidated in a few cases. In general, biosynthesis of a polyol from the corresponding phosphorylated sugar requires two steps, reduction and dephosphorylation, which may take place in either order. Saccharomyces cerevisiae produces glycerol from dihydroxyacetone phosphate via glycerol 3-phosphate using NAD+-dependent glycerol-3-phosphate dehydrogenase (Gpd1p) and glycerol-3-phosphate phosphatase (Gpp2p) (Albertyn et al., 1994; Norbeck et al., 1996
). Recently, it has been shown that A. nidulans utilizes a different route for glycerol biosynthesis, involving NADP+-dependent glycerol dehydrogenase and not glycerol-3-phosphate dehydrogenase (de Vries et al., 2003
; Fillinger et al., 2001
). In A. niger, mannitol biosynthesis from fructose 6-phosphate is accomplished by mannitol-1-phosphate dehydrogenase and mannitol-1-phosphate phosphatase (Ruijter et al., 2003
).
In this study, we have investigated the composition of the polyol pools in A. oryzae during SSF and the metabolic pathways involved in the biosynthesis of these polyols.
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METHODS |
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A single batch of wheat grains (Blok, Woerden, The Netherlands) was used in all experiments. Two types of solid media were used, wheat dough and whole grains. Wheat dough was prepared essentially as described by Nagel et al. (2001) with either 0·7 or 1 kg water (kg dry mass)1. The initial aw of the doughs was 0·981±0·002 and 0·991±0·003 for 0·7 and 1 kg water (kg dry mass)1, respectively. Discs of the wheat dough (Nagel et al., 2001
) were put into Petri dishes (60 mm) and inoculated by spreading 2·5x103 spores over the surface. Twenty Petri dishes were incubated without lids in a closed container, which was placed in a temperature-controlled cabinet at 35 °C. The container was flushed with 350 ml sterile air min1. The inlet air was humidified with water in the case of wheat dough with initial aw 0·991 or with a 0·6 M NaCl solution for dough with initial aw 0·981. Mycelium was cultured for 48 h.
Whole grains were soaked in excess water for 4 h at 50 °C. Following sieving, the grains were sterilized for 1·5 h at 121 °C. Wheat grains were put into a Petri dish (90 mm) in a packed layer and immersed in a sterile solution containing 1·5 % (w/v) agar such that the grains were just exposed to the surface of the agar. In order to decrease aw, 1·1 M NaCl was added to the agar solution. Plates were inoculated by spreading 104 spores over the surface. Five plates were incubated in a closed container as described above. Air was humidified with either water or 1·1 M NaCl. Mycelium was cultured for 48 h.
aw of solid substrates was measured using an Aqualab 3TE at 35 °C. aw and osmotic pressure are related by Vm=RTln(aw), where
is the osmotic pressure (Pa), Vm is the molar volume of water (mol m3), R=8·314 is the gas constant (J mol1 K1), T is the temperature (K) and aw is the water activity. Under the conditions used, aw 0·98 corresponds to osmotic pressure 2·9 MPa and aw 0·96 to 5·8 MPa.
Extraction and analysis of polyols.
Mycelium growing on solid substrates was rapidly frozen by pouring liquid nitrogen over the mycelium. Frozen mycelium was scraped from the substrate and lyophilized. Polyol extraction and determination were performed as described previously (Witteveen et al., 1994). In short, the method for polyol analysis was HPLC using a Dionex MA1 column combined with amperometric detection. This method will detect most polyols and sugars.
Preparation of cell extracts, enzyme assays and partial purification of polyol dehydrogenases.
Frozen mycelium (approx. 0·25 g), obtained as described above, was ground using a micro-dismembrator (B. Braun Biotech) and suspended in 0·5 ml of extraction buffer (50 mM potassium phosphate pH 7·0, 5 mM MgCl2, 5 mM 2-mercaptoethanol, 0·5 mM EDTA). Following centrifugation for 5 min at 15 000 g, the resulting supernatants were desalted by passage through a 2·5 ml Sephadex G25 column pre-equilibrated with extraction buffer.
Enzyme activities were determined at 30 °C. Polyol dehydrogenase activities were determined in 100 mM glycine pH 9·6 containing 0·5 mM NAD(P)+ and 100 mM glycerol, meso-erythritol, D-arabitol or D-mannitol. Reductase activities for aldoses, ketoses or their phosphorylated derivatives were assayed in 50 mM triethanolamine pH 7·4 containing 4 mM MgCl2, 0·2 mM NAD(P)H and one of the following substrates: 50 mM dihydroxyacetone, 50 mM D-erythrose, 50 mM D-ribulose, 50 mM D-xylulose, 50 mM D-ribose, 2 mM dihydroxyacetone phosphate, 1 mM D-ribulose 5-phosphate, 1 mM D-xylulose 5-phosphate or 1 mm D-ribose 5-phosphate. Mannitol-1-phosphate dehydrogenase activity was determined in 50 mM Tris pH 7·0 containing 0·2 mM NADH and 2·5 mM fructose 6-phosphate. Protein concentrations in cell extracts were determined with the Bicinchoninic acid protein kit (Sigma) according to the supplier's instructions using BSA as a standard.
Product analysis after incubation of cell extracts with various sugars was performed as follows. Desalted cell extract was incubated with 50 mM sugar in 50 mM triethanolamine pH 7·6, 2·5 mM MgCl2, 0·2 mM NADH. After different incubation times, 220 µl samples were taken and the reaction was stopped by addition of 20 µl of 9·1 M HClO4. Following 30 min incubation on ice, the samples were neutralized by addition of 40 µl of 5 M KOH and 20 µl of 2 M KHCO3. Precipitates were removed by centrifugation and reaction products were analysed in the supernatant by HPLC as described previously (Witteveen et al., 1994).
Fractionation of polyol dehydrogenases was performed by anion-exchange chromatography. A cell extract was prepared as described above using either 20 mM Tris pH 7·5 or 20 mM Bistris pH 6·5 as a buffer, in both cases containing 1 mM MgCl2, 5 mM 2-mercaptoethanol and 0·5 mM EDTA. Four millilitres of desalted cell extract was applied to a ResourceQ column (Amersham Biosciences) and, following rinsing with extraction buffer, protein bound to the column was eluted with a linear gradient of 00·5 M NaCl in extraction buffer. Polyol dehydrogenase activities were measured in the fractions obtained.
Northern analysis.
Powdered mycelium was prepared as described above. Total RNA was extracted using TRIzol (Life Technologies) according to the supplier's instructions. RNA (5 µg) was incubated with 3·3 µl of 6 M glyoxal, 10 µl DMSO and 2 µl of 0·1 M sodium phosphate buffer pH 7 in a total volume of 20 µl for 1 h at 50 °C to denature the RNA. Following electrophoresis in a 1·5 % agarose gel containing 10 mM sodium phosphate pH 7, RNA was transferred to nylon membranes (Hybond-N; Amersham Pharmacia Biotech) by capillary blotting in 10xSSC. Hybridization was performed at 42 °C in a solution containing 50 % (v/v) formamide, 0·9 M NaCl, 90 mM sodium citrate tribasic dihydrate (citric acid), 0·2 % (w/v) ficoll, 0·2 % (w/v) polyvinylpyrrolidone, 0·2 % (w/v) BSA, 0·1 % (w/v) SDS, 10 % (w/v) dextran sulphate and 100 µg single-stranded herring sperm DNA ml1. The following probes were used: a gldB cDNA from A. nidulans (de Vries et al., 2003) and a 0·7 kbp EcoRI fragment of the 18S rRNA subunit (Melchers et al., 1994
) as a loading control. Blots were washed with 0·6 M NaCl, 60 mM citric acid, 0·5 % (w/v) SDS at 56 °C (gldB) or with 30 mM NaCl, 3 mM citric acid, 0·5 % (w/v) SDS at 65 °C (18S).
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RESULTS |
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Upon a decrease in aw, NADP+-dependent glycerol dehydrogenase activity increased (Table 2). In contrast, NAD+-dependent glycerol-3-phosphate dehydrogenase activity was lower in mycelium cultured on wheat grains at low aw, whereas it did not vary with changes in aw in the case of mycelium cultured on wheat dough. Likewise, NADP+-dependent erythritol dehydrogenase (erythrose reductase) activity was increased in mycelium cultured at low aw (Table 2
). NAD+-dependent D-arabitol dehydrogenase activity did not vary in wheat-grain-cultured mycelium, but decreased with a reduction in aw in mycelium cultured on wheat dough. L-arabitol dehydrogenase activity was not detected and it was therefore assumed that the arabitol found in the cells was D-arabitol. The precursor of the D-arabitol produced by D-arabitol dehydrogenase could be either D-ribulose or D-xylulose. Incubation of cell extract with D-ribulose in the presence of NADH resulted in production of ribitol, whereas incubation with D-xylulose produced arabitol and xylitol.
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Partial purification of polyol dehydrogenases from a cell extract was performed by anion-exchange chromatography to find out which activities could be separated and hence were attributable to distinct enzymes. Four enzymes bearing highest activities with mannitol, glycerol, erythritol and D-arabitol, respectively, were well separated (Fig. 1). The major part of the mannitol dehydrogenase activity (70 %) did not bind to the anion-exchange column and eluted in the first two fractions, whereas glycerol dehydrogenase bound weakly. The glycerol dehydrogenase co-eluted with erythritol dehydrogenase activity, but a distinct erythritol dehydrogenase was eluted upon increasing the ionic strength of the buffer. Approximately 30 % of the erythritol dehydrogenase activity was catalysed by the glycerol dehydrogenase enzyme. Finally, D-arabitol dehydrogenase activity eluted after the erythritol dehydrogenase. The erythritol and arabitol dehydrogenase activities were not completely separated, but the first fractions containing erythritol dehydrogenase activity (fractions 1012) did not contain arabitol dehydrogenase activity while the opposite was observed for the last arabitol dehydrogenase fractions (1820). Since the erythritol dehydrogenase and the arabitol dehydrogenase have different cofactor specificities (NADP+ and NAD+, respectively) it is not likely that these activities are contained in one enzyme.
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DISCUSSION |
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A. oryzae mycelium that accumulated various polyols at low aw contained at least four distinct polyol dehydrogenases: a glycerol dehydrogenase that also had moderate activity toward erythritol, a distinct erythritol dehydrogenase, a D-arabitol dehydrogenase and a mannitol dehydrogenase. The level of NADP+-dependent glycerol dehydrogenase was higher at low aw and this increase in activity correlated very well to glycerol accumulation, suggesting that this enzyme is part of the biosynthetic pathway for glycerol in A. oryzae. The function of NADP+-dependent glycerol dehydrogenase in glycerol biosynthesis at low aw is well documented in A. nidulans (de Vries et al., 2003). The glycerol biosynthesis pathway in A. nidulans and probably also in A. oryzae comprises dephosphorylation of dihydroxyacetone phosphate followed by reduction of dihydroxyacetone to glycerol by glycerol dehydrogenase. Induction of glycerol dehydrogenase at low aw presumably is (part of) the response of A. oryzae to osmotic stress in order to be able to produce a larger quantity of polyols to maintain turgor of the cells. However, only a twofold increase in glycerol dehydrogenase activity was observed upon decreasing aw, while the glycerol concentration in the cells increased 10- to 15-fold. These observations indicate that glycerol biosynthesis is partially controlled by the quantity of glycerol dehydrogenase, but also by other mechanism(s). Possible other control mechanisms are an increase in dihydroxyacetone-phosphate phosphatase activity, regulation of glycerol dehydrogenase activity or better retention of glycerol in the cells. The latter mechanism is well documented in S. cerevisiae, where Fps1p controls glycerol accumulation and release (Tamás et al., 1999
).
Similar to glycerol, accumulation of erythritol correlated to NADP+-dependent erythritol dehydrogenase (erythrose reductase) activity. These data suggest that (i) A. oryzae developed similar pathways for biosynthesis of glycerol and erythritol, i.e. dephosphorylation of an intermediary metabolite followed by reduction of the resulting aldose/ketose to a polyol, and (ii) that erythritol dehydrogenase is induced at low aw. A different pathway for erythritol biosynthesis was reported for the lactic acid bacterium Leuconostoc oenos (Veiga-da-Cunha et al., 1993). In L. oenos, erythrose 4-phosphate is converted via erythritol 4-phosphate to erythritol, which is catalysed by erythrose-4-phosphate reductase and erythritol-4-phosphate phosphatase (Veiga-da-Cunha et al., 1993
). We did not detect any erythrose-4-phosphate reductase activity in A. oryzae, whereas high erythrose reductase activity was found in erythritol-accumulating mycelium. Erythrose reductases have been reported for Aureobasidium species (Tokuoka et al., 1992
), but these authors have not reported whether these enzymes are differentially expressed at low aw.
D-Arabitol production during osmostress has been described for a few fungal species, including Zygosaccharomyces rouxii (Blakeley & Spencer, 1962), Magnaporthe grisea (Dixon et al., 1999
), Aspergillus flavus (Mellon et al., 2002
), Epicoccum nigrum (Pascual et al., 2003
) and Cladosporium fulvum (Clark et al., 2003
). Z. rouxii and the marine fungus Dendryphiella salina convert D-xylulose 5-phosphate to D-xylulose which is subsequently reduced to D-arabitol by an nad+-dependent D-arabitol dehydrogenase (Blakeley & Spencer, 1962
; Low & Jennings, 1975
). In Candida albicans, it was first thought that D-arabitol was formed by dephosphorylation of D-ribulose 5-phosphate and subsequent reduction of D-ribulose by an nad+-dependent D-arabitol dehydrogenase (Wong et al., 1993
), but this model was later proven incorrect. A C. albicans ard mutant, lacking NAD+-dependent D-arabitol dehydrogenase, failed to grow on minimal D-arabitol and D-arabinose medium, but its D-arabitol production was not affected (Wong et al., 1995
). These results showed that NAD+-dependent D-arabitol dehydrogenase is involved in D-arabitol utilization, but that it is not required for D-arabitol biosynthesis. Our data are consistent with A. oryzae converting D-xylulose into D-arabitol using a pathway as described for Z. rouxii and D. salina.
In summary, we have shown in this study accumulation of glycerol, erythritol and arabitol by A. oryzae at low aw on solid-state substrate. Accumulation of a mixture of polyols might be typical for SSF due to the specific growth conditions present during growth on a solid substrate. NADP+-dependent glycerol and erythritol dehydrogenases were induced at low aw suggesting that these enzymes are involved in biosynthesis of glycerol and erythritol, respectively, but this needs to be proven by construction of mutants lacking these enzymes.
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ACKNOWLEDGEMENTS |
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Received 20 August 2003;
revised 18 November 2003;
accepted 23 December 2003.