Centre for Advanced Lipid Research, Department of Biological Sciences, University of Hull, Hull HU6 7RX, UK1
Department of Microbiology, Universitii Kebangsaan, Malaysia, 43600 Bangi, Salangor Darul Ehsan, Malaysia2
Author for correspondence: James P. Wynn. Tel: +44 1482 465507. Fax: +44 1482 465458. e-mail: j.p.wynn{at}biosci.hull.ac.uk
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ABSTRACT |
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Keywords: AMP deaminase, NAD+:isocitrate dehydrogenase, lipid accumulation
Abbreviations: dO2, dissolved oxygen concentration; NAD:ICDH, NAD+:isocitrate dehydrogenase; PFK, phosphofructokinase
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INTRODUCTION |
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In attempts to optimize these processes and to realize new ones, a sound understanding of the regulation of cell lipid accumulation in the production organism is required. Although the biochemical basis of microbial oleaginicity has been elucidated this work has been carried out almost exclusively in yeasts (Botham & Ratledge, 1979 ; Boulton & Ratledge, 1983
; Evans & Ratledge, 1983
, 1985a
, b
). A similar system has been tacitly assumed to apply to other groups of eukaryotic micro-organisms.
All the enzymes thought to be crucial for the accumulation of substantial amounts of storage lipid are present in both oleaginous yeasts and filamentous fungi. However, it has become apparent in the course of our more recent work that the regulation of lipogenesis may differ between filamentous fungi and yeasts. In the present study Mucor circinelloides was used as the principal model for oleaginous filamentous fungi. However, because of its commercial importance Mort. alpina was also included in order to verify the key results. This report indicates a number of significant differences in the biochemistry of lipid accumulation between oleaginous yeasts and oleaginous filamentous fungi. A revised and more concerted mechanism for the initiation of storage lipid synthesis when filamentous fungi experience N-limiting conditions is put forward. These findings have implications not only for the production of single-cell oils but also for other commercial fermentations using filamentous fungi (e.g. those producing citric acid and gibberellic acid) in which lipid synthesis represents an undesirable by-product and therefore an unnecessary waste of substrate.
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METHODS |
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Cultivation of fungi.
Mucor circinelloides CBS 108.16 and Mortierella alpina Peyron CBS 696.70 were initially cultivated in 1 litre magnetically stirred bottles containing 800 ml Kendrick medium (N-limiting) (Kendrick & Ratledge, 1992 ). These cultures were incubated for 16 h at 30 °C then used at 10% (v/v) to inoculate 4 litre (working volume) fermenters containing modified Kendrick medium containing ammonium tartrate at 2 g l-1 for both fungi, with glucose at 50 g l-1 for Mc. circinelloides and 30 g l-1 for Mort. alpina. Fermenters were incubated at 30 °C, stirred at 500 r.p.m. with aeration at 0·5 vol. vol.-1 min-1 and pH maintained at 5·5 by automatic addition of KOH and HCl.
Production of cell extracts.
Biomass was harvested by filtration (under reduced pressure) through a Whatman no. 1 filter and washed with cold distilled water. Cell extracts for the determination of enzyme activities were prepared by suspending mycelia in an extraction buffer (Wynn et al., 1998 ) and disrupted either by passage twice through a French pressure cell at 35 MPa or by a single pass through a One Shot cell disrupter (Constant Systems) at 640 MPa. Disrupted cell suspensions were centrifuged at 10000 g for 10 min and the supernatant retained for enzyme analysis. Protein concentrations were determined using the method of Bradford (1976)
with BSA as a standard.
Preparation of spheroplasts and isolation of mitochondria.
Mc. circinelloides was grown in 1 litre stirred bottles, as described above. Harvested biomass was washed twice with spheroplasting buffer (50 mM Tris/HCl, pH 6·5, containing 1·2 M sorbitol) then suspended in the same buffer containing chitinase/chitosanase preparation (Vanheeswijck, 1984 ) at 2 mg ml-1 and incubated for 3 h at 30 °C. The treated biomass was collected by centrifugation (10000 g for 10 min at 4 °C) and the spheroplasts formed were disrupted by resuspension in 50 mM Tris/HCl (pH 7·5) and incubated at 20 °C for 1 h. The disrupted cells were fractionated by centrifugation at 5000 g for 10 min and then the supernatant centrifuged at 16000 g for 20 min. The cell preparations were maintained at 4 °C throughout. The supernatant and the pellet formed at 16000 g were retained as the cytosolic and mitochondrial fractions respectively.
Estimation of enzyme activities.
NAD+:isocitrate dehydrogenase (NAD:ICDH) was assayed using the method of Kornberg (1955) . AMP deaminase activity was assayed using a reaction volume containing 100 mM KH2PO4/KOH (pH 7·1), 2 mM MgCl2, 0·5 mg BSA ml-1, 2 mM NaH2PO4, 2 mM ATP (pH 7·1), 5 mM AMP. The assay volume was incubated for 10 to 30 min at 30 °C and the
liberated was quantified using the indophenol method (Chaney & Marbach, 1962
). Adenylate kinase was determined using a method based on that of Sottocasa et al. (1967)
. The reaction volume contained 67 mM Tris/HCl (pH 7·5), 5 mM ADP, 0·5 units hexokinase/glucose-6-phosphate dehydrogenase mix (from bakers yeast), 5 mM MgSO4, 0·2 mM NADP+ and 10 mM glucose. The increase in A340 was measured. Phosphofructokinase (PFK) was assayed as described by Sols & Salas (1966)
, citrate synthase as described by Parvin (1969)
, succinate dehydrogenase as described by Schwitzguebel et al. (1981)
, pyruvate kinase as described by Worthington Enzymes (1979)
and pyruvate carboxylase as described by Seubert & Weicker (1969)
. The assay reactions for citrate synthase, succinate dehydrogenase, pyruvate kinase and pyruvate carboxylase were supplemented with 0·1% (w/v) Triton X-100.
Determination of nucleotide concentrations.
The concentrations of adenine nucleotides (ATP, ADP and AMP) were determined using the luciferin/luciferase system to quantify ATP (Speilmann et al., 1982 ). Samples (20 ml) from the fermenter were rapidly quenched (<0·5 s) by collection in sterile Universal tubes containing 4 ml conc. H2SO4. Quenched samples were diluted 1:10 with 5 mM glycylglycine/KOH buffer (pH 7·8) and, if necessary, the pH adjusted to 7·8. Cell debris was removed by centrifugation at 10000 g for 10 min and then the supernatant was diluted 1:10 with glycylglycine buffer. The diluted supernatant was either analysed immediately or stored at -20 °C for a maximum of 2 weeks prior to analysis. ATP was quantified by adding 50 µl of the sample prepared above to 50 µl 100 mM glycylglycine buffer (pH 7·8) containing 5 mM MgSO4, 0·5 mM phosphoenolpyruvate and 100 µl ATP assay mix (Sigma). ADP was quantified by conversion to ATP in the same reaction mixture as described above but with the addition of 1 unit pyruvate kinase (type 1: crude preparation from rabbit muscle) and incubated at 37 °C for 10 min prior to the addition of the ATP assay mix. ADP was calculated by subtraction of the amount of ATP detected previously. AMP was likewise quantified by its conversion to ATP and then subtraction of the amount of ATP+ADP detected in the sample. AMP was converted to ATP by addition of 1 unit myokinase (from porcine muscle) and 1 unit pyruvate kinase followed by a 10 min incubation at 37 °C prior to the addition of the ATP assay mix.
Determination of culture dry weight.
A 20 ml sample of the culture was harvested on to a washed, dried and pre-weighed filter (Whatman no. 1). The filtrate was retained and analysed for culture glucose and ammonium concentrations (see below). Harvested biomass was washed with distilled water and then dried at 110 °C to constant weight. The weight of the biomass was determined gravimetrically.
Analysis of the culture supernatant.
The glucose concentration in the culture medium was determined using a GOD-Perid test kit (Boehringer Mannheim) according to the manufacturers instructions. The ammonium concentration in the culture filtrate was determined using the indophenol test (Chaney & Marbach, 1962 ).
Analysis of intracellular metabolites.
Samples were collected into tubes containing 4 ml 5·8 M HClO4. Samples were centrifuged at 10000 g for 10 min and the supernatant adjusted to pH 7·4 with NaOH. The isocitrate and citrate concentrations in the samples were determined as described by Siebert (1974) and Dagley (1974)
respectively. Samples of culture medium were also analysed for isocitrate and citrate to allow corrections to be made for extracellular metabolites.
Determination of culture dO2 and CO2 evolution.
The dissolved oxygen concentration (dO2) of the culture medium in the fermenters was continuously recorded using galvanic O2 electrodes (Mettler-Toledo, Woburn, MA, USA) as a percentage of the O2 concentration in the equilibrated culture medium immediately prior to inoculation. The CO2 evolution by the fungal cultures was determined by measuring the CO2 in the fermenter exhaust using a CO2 analyser (Analytical Development Co., Hoddesdon, UK).
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RESULTS |
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When the activity of AMP deaminase was determined, in triplicate batch cultures of Mc. circinelloides, an increase in activity was reproducibly observed at the point when the cultures became N-limited and was coincident with the onset of lipid accumulation (see Fig. 3a). The increase in activity of AMP deaminase was, however, transitory and activity returned to basal levels within 5 h of N-exhaustion (see Fig. 3a
). Similar data were obtained with Mort. alpina (Fig. 3b
).
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The stimulation of AMP deaminase and adenylate kinase activities, at the time of N-exhaustion, led to an almost immediate decrease in the concentrations of ATP, ADP and AMP. The concentrations of all three adenine nucleotides decreased by 50% during the 23 h period immediately after N-exhaustion (see Fig. 4). ADP was the most abundant nucleotide [concentration varying from 3·7 to 1·1 nmol (mg dry wt)-1], followed by ATP [between 1·6 and 0·5 nmol (mg cell dry wt)-1]. The AMP concentration was consistently the lowest [0·90·4 nmol (mg dry wt)-1]. Although the cellular total nucleotide concentration decreased by 50% at the point when the cells became growth-limited by the depletion of the N-source, the cellular energy charge remained relatively constant at between 0·5 and 0·6 as there were concomitant changes in all three adenine nucleotides.
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DISCUSSION |
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To correlate this switch in metabolic flux to the overall down-regulation of metabolic activity is a large and complex task. However, with our focus being on the mechanism of lipid accumulation we have aimed to elucidate the biochemical events underlying this rapid cellular response to N-limitation and hence the transition from the trophophase to the idiophase when lipid and other secondary products are accumulated. Only by a sound understanding of this biochemistry can rational attempts at increasing lipid/secondary product accumulation using molecular techniques be made. Previous attempts to increase the production of secondary products by gene cloning without a sound biochemical background have highlighted the hit and miss (mainly miss) nature of this approach (Rangasamy & Ratledge, 2000 ; Ratledge, 2000
; Roehr et al., 1996
; Ruijter et al., 2000
).
The enzymes thought to be important in the initiation of lipid accumulation (ATP:citrate lyase, AMP deaminase and NAD+:isocitrate lyase) in yeasts (Evans & Ratledge, 1985a , b
) have all been reported in filamentous fungi (Botham & Ratledge, 1979
; Wynn et al., 1998
) and their presence in Mc. circinelloides has been confirmed in this and previous studies (Wynn et al., 1997
).
NAD+:ICDH in oleaginous fungi
The characteristics of the NAD:ICDHs of the two fungi examined were similar (though the enzyme from Mucor had a higher affinity for isocitrate than that from Mortierella) but were clearly distinct from those described for the NAD:ICDH from oleaginous yeasts (Botham & Ratledge, 1979 ; Evans et al., 1983
; Evans & Ratledge, 1985b
). Both fungi are oleaginous with Mc. circinelloides capable of accumulating 25% (w/w dry wt) lipid and Mort. alpina capable of accumulating 40% lipid (Wynn et al., 1999
) however, the NAD:ICDH activities in these fungi were not absolutely dependent on AMP and, therefore, more closely resembled the enzyme from non-oleaginous micro-organisms (Evans et al., 1983
; Atkinson et al., 1965
). It is important to note, however, that NAD:ICDH was only active in the absence of AMP at non-physiological concentrations of isocitrate; i.e. 3 mM and 5 mM for Mc. circinelloides and Mort. alpina respectively. Although we could not detect isocitrate within the cells it is likely that the concentration of isocitrate will be approximately 5% of the citrate concentration (Siebert, 1974
), which was measured maximally at 1520 nmol (mg dry wt)-1. This would give a concentration of isocitrate of less than 1 nmol (mg dry wt)-1, i.e. less than 0·5 mM assuming 24 µl H2O (mg dry wt)-1 (Knowles, 1977
). At this concentration of isocitrate, NAD:ICDH would be down-regulated by the decrease in AMP concentration observed in the cells as a result of the increase in AMP deaminase activity triggered by N-exhaustion from the culture medium: see below.
AMP deaminase activity
Activation of AMP deaminase plays a key role in decreasing the concentration of AMP in oleaginous yeasts entering N-limited growth (Evans & Ratledge, 1985a ). A similar situation occurred in the filamentous fungi studied, with a peak of AMP deaminase activity coincident with N-exhaustion causing a decrease in the intracellular AMP concentration (see below). The decrease in AMP concentration was coincident with the decrease in CO2 evolution (see Fig. 1
), which was indicative of a major decrease in the activity of the citric acid cycle. That the increase in activity of AMP deaminase was directly responsible for the decline in activity of NAD:ICDH, thereby affecting the overall activity of the citric acid cycle, is entirely consistent with these observations. The situation is more complex than in oleaginous yeasts in that AMP deaminase subsequently returned to its original activity, but this did not lead to any concomitant increase in AMP concentration or in the rate of CO2 evolution. Therefore, the effect of the increase in AMP deaminase, though transitory, nevertheless may have initiated a cascade of connected biochemical events that rapidly led to the initiation of lipid accumulation.
Changes in adenylate pool size and cellular energy charge
The increased activity of AMP deaminase led to a rapid decrease in the cellular AMP, from 0·8 nmol (mg dry wt)-1 (0·20·4 mM) prior to N-exhaustion to 0·5 nmol (mg dry wt)-1 (0·130·25 mM) after N-exhaustion. AMP remained at this concentration even after AMP deaminase returned to its initial activity (Fig. 4). This approximately 50% decrease in AMP concentration, although not as dramatic as the 90% decrease reported for some yeasts (Boulton & Ratledge, 1983
; Mitsushima et al., 1978
), would nevertheless still lead to a significant down-regulation of NAD:ICDH activity, and thereby rapidly diminish the carbon flux through the citric acid cycle. Although the AMP concentration measured represented the total intracellular, rather than intra-mitochondrial, concentration this is thought unimportant as the mitochondrial membrane in eukaryotic micro-organisms is permeable to AMP (Matsushima et al., 1978; Bartels & Jensen, 1979
). Therefore, the intracellular and intra-mitochondrial concentrations of AMP are likely to be equivalent.
Although a decrease in the AMP concentration associated with the depletion of the N-source in Mc. circinelloides resembles the situation reported for oleaginous yeasts (Boulton & Ratledge, 1983 ; Mitsushima et al., 1978
), changes in the adenylate pool and the cellular energy charge differed from those previously reported. Whereas in yeasts the decrease in AMP was accompanied by an increase in the intracellular ATP/AMP ratio (Solodovnikova et al., 1998
) and, therefore, energy charge, in Mc. circinelloides this did not happen. During the transition from N-replete to N-limited growth, the cellular concentrations of all three adenine nucleotides decreased by approximately 50% and the energy charge remained constant. This is similar to the situation reported in bacteria and fish, where during metabolic stress the cellular energy charge was maintained within closely defined limits at the expense of the total adenylate pool size, by the combined action of AMP deaminase and adenylate kinase (Atkinson, 1977
; Woo & Chiu, 1997
).
Regulatory properties of PFK
A preliminary study of the PFK of Mc. circinelloides demonstrated regulatory properties similar to those reported for the enzyme from other fungi (Roehr et al., 1996 ). Activity was stimulated by the presence of
and inhibited by citrate, with these effectors acting antagonistically. Therefore under N-limiting conditions a decrease in the intracellular
and an increase in citrate concentration would down-regulate the carbon flux through the glycolytic pathway, via inhibition of PFK activity. It is possible that the build-up of citrate due to the down-regulation of the citric acid cycle (see above) plays a role in regulating the rate of glycolysis (due to the inhibition of PFK) and therefore its own synthesis.
Subcellular localization of pyruvate carboxylase
A cytosolic pyruvate carboxylase activity in Mc. circinelloides is in accord with the observations in other fungi (Osmani & Scrutton, 1985 ; Klitsch et al., 1991
). In this regard filamentous fungi are distinct from yeasts, in which both mitochondrial and cytosolic pyruvate carboxylases have been reported (Evans et al., 1983
; Rohde et al., 1991
; Van Urk et al., 1989
), and animals, where pyruvate carboxylase is mitochondrial (Böttger et al., 1969
; Taylor et al., 1978
). A cytosolic pyruvate carboxylase would allow the cytosolic generation of NADPH for lipid biosynthesis via the co-operation of pyruvate carboxylase, malate dehydrogenase and malic enzyme to produce a transhydrogenase cycle generating NADPH at the expense of NADH and ATP.
Conclusions
The biochemical mechanism responsible for the initiation of lipid accumulation in oleaginous fungi differs in several fundamental ways from that observed in oleaginous yeasts. As a result, the hypothesis explaining the onset of lipid accumulation in oleaginous yeasts requires modification before it can be applied to filamentous fungi. Although many of the enzymes involved in both systems are the same, the mechanism that operates in filamentous fungi is a somewhat more complicated and concerted process than that reported for yeasts.
The following hypothesis is put forward to explain the switching from active growth to the stationary phase in filamentous fungi.
1. As the N-source in the medium reaches a concentration below the Km value of the uptake mechanisms the intracellular concentration decreases. Because
is an activator of PFK the decrease in its concentration will lead to a decrease in the carbon flux through the glycolytic pathway (Fig. 6
).
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3. As a result of the activation of AMP deaminase, the AMP concentration will decrease and further limit the activity of the citric acid cycle (via NAD:ICDH activity), further restricting ATP generation; this will lead to a feed-forward process so that the energy charge can only be maintained by decreasing the total adenylate pool to a minimum level (approx. 50% of its trophophase level). Flux through the citric acid cycle will be further limited to a basal level, thereby restricting the supply of intermediates for anabolic processes, which will be further down-regulated by the decreased cellular ATP concentration, bringing them into line with the growth-limited environment.
4. When NAD:ICDH activity falls, citrate accumulates, which could further limit PFK activity. Citrate is translocated into the cytosol for cleavage by ATP:citrate lyase to provide cytosolic acetyl-CoA for lipogenesis.
5. The cytosolic pyruvate carboxylase together with ATP:citrate lyase and malate dehydrogenase allow a transhydrogenase cycle to generate cytosolic NADPH independently from the citrate/malate cycle (Evans et al., 1983 ), thereby providing sufficient reducing power for lipogenesis.
That Mc. circinelloides and Mort. alpina have such an involved mechanism for the sensing of oncoming N-limited growth should not be a surprise. These fungi are commonly isolated from soil, which is a N-limited habitat, so that intricate mechanisms to deal with N-depletion from the environment would be an expected environmental adaptation.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Atkinson, D. E., Hathaway, J. A. & Smith, E. C. (1965). Kinetics of regulatory enzymes: kinetic order of the yeast diphosphopyridine nucleotide isocitrate dehydrogenase reaction and a model for the reaction. J Biol Chem 240, 2682-2690.
Bartels, P. D. & Jensen, P. J. (1979). Role of AMP in regulation of the citric acid cycle in mitochondria from bakers yeast. Biochim Biophys Acta 582, 246-259.[Medline]
Botham, P. A. & Ratledge, C. (1979). A biochemical explanation for lipid accumulation in Candida 107 and other oleaginous micro-organisms. J Gen Microbiol 114, 361-375.[Medline]
Böttger, I., Wieland, O., Brdiczka, D. & Pette, D. (1969). Intracellular localisation of pyruvate carboxylase and phosphoenolpyruvate carboxylase in rat liver. Eur J Biochem 8, 113-119.[Medline]
Boulton, C. A. & Ratledge, C. (1983). Use of transition studies in continuous cultures of Lipomyces starkeyi, an oleaginous yeast, to investigate the physiology of lipid accumulation. J Gen Microbiol 129, 2871-2876.
Bradford, M. M. (1976). A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of proteindye binding. Anal Biochem 72, 248-254.[Medline]
Chaney, A. L. & Marbach, E. P. (1962). Modified reagents for determination of urea and ammonium. Clin Chem 8, 130-132.
Dagley, S. (1974). Citrate UV spectrophotometric determination. Methods Enzymatic Anal 3, 1562-1565.
Evans, C. T. & Ratledge, C. (1983). Biochemical activities during lipid accumulation in Candida curvata. Lipids 18, 630-635.[Medline]
Evans, C. T. & Ratledge, C. (1985a). The role of NAD+:isocitrate dehydrogenase in lipid accumulation by the oleaginous yeast Rhodosporidium toruloides CBS 14. Can J Microbiol 31, 845-850.
Evans, C. T. & Ratledge, C. (1985b). Possible regulatory roles of ATP:citrate lyase, malic enzyme, and AMP deaminase in lipid accumulation by Rhodosporidium toruloides CBS 14. Can J Microbiol 31, 1000-1005.
Evans, C. T., Scragg, A. H. & Ratledge, C. (1983). A comparative study of citrate efflux from mitochondria of oleaginous and non-oleaginous yeasts. Eur J Biochem 130, 195-204.[Medline]
Kendrick, A. & Ratledge, C. (1992). Desaturation of polyunsaturated fatty acids in Mucor circinelloides and the involvement of a novel membrane-bound malic enzyme. Eur J Microbiol 209, 667-673.
Klitsch, W. M., Kubicek, C. P. & Scrutton, M. C. (1991). Intracellular location of enzymes involved in citrate production by Aspergillus niger. Can J Microbiol 37, 823-827.[Medline]
Knowles, C. J. (1977). Microbial metabolic regulation by adenine pools. Symp Soc Gen Microbiol 27, 241-283.
Kornberg, A. (1955). Isocitric dehydrogenase of yeast (TPN). Methods Enzymol 1, 705-709.
Kyle, D. J. (1997). Production and use of a single cell oil highly enriched in arachidonic acid. Lipid Technol 9, 116-121.
Mitsushima, K., Shinmyo, A. & Enatsu, T. (1978). Control of citrate and 2-oxoglutarate formation in Candida lipolytica mitochondria by adenine nucleotides. Biochim Biophys Acta 538, 481-492.[Medline]
Osmani, S. A. & Scrutton, M. C. (1985). The subcellular localisation and regulatory properties of pyruvate carboxylase from Rhizopus arrhizus. Eur J Biochem 147, 119-128.[Abstract]
Parvin, R. (1969). Citrate synthase from yeast. Methods Enzymol 13, 16-19.
Rangasamy, D. & Ratledge, C. (2000). Genetic enhancement of fatty acid synthesis by targeting rat liver ATP:citrate lyase into plastids of tobacco. Plant Physiol 122, 1231-1238.
Ratledge, C. (1997). Microbial lipids. In Biotechnology , pp. 133-197. Edited by H. J. Rehm & G. Reed. Weinheim:VCH.
Ratledge, C. (2000). Look before you clone A comment on Properties of Aspergillus niger citrate synthase and effects of citA overexpression on citric acid production by G. H. G. Ruijter, H. Panneman, D.-B. Xu & J. Visser FEMS Microbiol Lett 184 (2000), 3540. FEMS Microbiol Lett 189, 317-318.[Medline]
Roehr, M., Kubicek, C. P. & Komínek, J. (1996). Citric acid. In Biotechnology,vol. 6, Products of Primary Metabolism , pp. 311-145. Edited by H. J. Rehm, G. Reed, A. Pühler & P. Stadler. Weinheim:VCH.
Rohde, M., Lim, F. & Wallace, J. C. (1991). Electron microscopic localisation of pyruvate carboxylase in rat liver and Saccharomyces cerevisiae by immunogold procedures. Arch Biochem Biophys 290, 197-201.[Medline]
Ruijter, G. H. G., Panneman, H., Xu, D.-B. & Visser, J. (2000). Properties of Aspergillus niger citrate synthase and effects of citA overexpression on citric acid production. FEMS Microbiol Lett 184, 35-40.[Medline]
Schwitzguebel, J. P., Moller, I. M. & Palmer, J. M. (1981). Changes in density of mitochondria and glyoxysomes from Neurospora crassa: a re-evaluation utilizing silica sol gradient centrifugation. J Gen Microbiol 126, 289-295.
Seubert, W. & Weicker, H. (1969). Pyruvate carboxylase from Pseudomonas. Methods Enzymol 13, 258-260.
Siebert, G. (1974). Isocitrate UV spectrophotometric determination. Methods Enzymatic Anal 3, 1570-1576.
Solodovnikova, N. Y., Sharyshev, A. A., Medentsev, A. G., Voloshin, A. N., Morgunov, I. G. & Finogenova, T. V. (1998). Dynamics of the adenine nucleotide pool in Yarrowia lipolytica cells overproducing organic acids. Microbiology 67, 28-32.
Sols, A. & Salas, M. (1966). Phosphofructokinase III. Yeast. Methods Enzymol 9, 436-442.
Sottocasa, G. L., Kuylenstierna, B., Ernster, L. & Bergstrand, A. (1967). Separation and some enzymatic properties of the inner and outer membranes of rat liver mitochondria. Methods Enzymol 10, 448-463.
Speilmann, H., Jacob-Müller, U. & Schulz, P. (1982). Simple assay of 0·11·0 pmol of ATP, ADP and AMP in single somatic cells using purified luciferin luciferase. Anal Biochem 113, 172-178.
Taylor, D. J., Crabtree, B. & Smith, G. H. (1978). The intracellular location of pyruvate carboxylase, citrate synthase and 3-hydroxyacyl-CoA dehydrogenase in lactating rat mammary gland. Biochem J 171, 273-275.[Medline]
Van Urk, H., Schipper, D., Breedveld, G. J., Mak, P. R., Scheffers, W. A. & van Dijken, J. P. (1989). Localisation and kinetics of pyruvate-metabolizing enzymes in relation to aerobic alcoholic fermentation in Saccharomyces cerevisiae CBS 8066 and Candida utilis CBS 621. Biochim Biophys Acta 992, 78-86.[Medline]
Vanheeswijck, R. (1984). The formation of protoplasts from Mucor species. Carlsberg Res Commun 49, 597-609.
Woo, N. Y. S. & Chiu, S. F. (1997). Metabolic and osmoregulatory responses of the sea bass Lates calcarifer to nitrate exposure. Environ Toxicol Water Quality 12, 257-264.
Worthington Enzymes (1979). Pyruvate kinase. In Enzymes and Related Biochemicals, pp 179180. Bedford, MA: Millipore Corporation.
Wynn, J. P. (1998). Microorganisms as sources of nutritionally important polyunsaturated fatty acids. Proceedings of the 12th Forum on Applied Biotechnology, part 1, pp. 12151222. ISSN 0368-9697.
Wynn, J. P., Kendrick, A. & Ratledge, C. (1997). Sesamol as an inhibitor of growth and lipid metabolism in Mucor circinelloides via its action on malic enzyme. Lipids 32, 605-610.[Medline]
Wynn, J. P., Hamid, A. A., Midgley, M. & Ratledge, C. (1998). Widespread occurrence of ATP:citrate lyase and carnitine acetyltransferase in filamentous fungi. World J Microbiol Biotechnol 14, 145-147.
Wynn, J. P., Hamid, A. A. & Ratledge, C. (1999). The role of malic enzyme in the regulation of lipid accumulation in filamentous fungi. Microbiology 145, 1911-1917.[Abstract]
Received 16 March 2001;
revised 25 June 2001;
accepted 4 July 2001.