On the mechanism of action of the antifungal agent propionate

Propionyl-CoA inhibits glucose metabolism in Aspergillus nidulans

Authors


W. Buckel, Laboratorium für Mikrobiologie, Fachbereich Biologie, Philipps-Universität Marburg, D-35032 Marburg, Germany. Fax: +49 6421 2828979, Tel.: +49 6421 2821527,
E-mail: Buckel@staff.Uni-Marburg.de

Abstract

Propionate is used to protect bread and animal feed from moulds. The mode of action of this short-chain fatty acid was studied using Aspergillus nidulans as a model organism. The filamentous fungus is able to grow slowly on propionate, which is oxidized to acetyl-CoA via propionyl-CoA, methylcitrate and pyruvate. Propionate inhibits growth of A. nidulans on glucose but not on acetate; the latter was shown to inhibit propionate oxidation. When grown on glucose a methylcitrate synthase deletion mutant is much more sensitive towards the presence of propionate in the medium as compared to the wild-type and accumulates 10-fold higher levels of propionyl-CoA, which inhibits CoA-dependent enzymes such as pyruvate dehydrogenase, succinyl-CoA synthetase and ATP citrate lyase. The most important inhibition is that of pyruvate dehydrogenase, as this affects glucose and propionate metabolism directly. In contrast, the blocked succinyl-CoA synthetase can be circumvented by a succinyl-CoA:acetate/propionate CoA-transferase, whereas ATP citrate lyase is required only for biosynthetic purposes. In addition, data are presented that correlate inhibition of fungal polyketide synthesis by propionyl-CoA with the accumulation of this CoA-derivative. A possible toxicity of propionyl-CoA for humans in diseases such as propionic acidaemia and methylmalonic aciduria is also discussed.

Abbreviations
ABTS

2,2′-azinobis(3-ethylbenzo-6-thiazolinesulfonic acid

ACS

acetyl-CoA synthetase

DTNB

5,5′-dithiobis(2-nitrobenzoic acid)

GOD

glucose oxidase

LDH

lactate dehydrogenase

MDH

malate dehydrogenase

POD

peroxidase

Sodium propionate is widely used as a preservative due to its ability to inhibit fungal growth. Furthermore, this short-chain fatty acid (pion = fat) prevents the biosynthesis of polyketides such as ochratoxin A by Aspergillus sulphureus and Penicillium viridicatum[1]. On the other hand, many fungi are able to grow on propionate, although much more slowly than on glucose or acetate. Recently we have shown that in Aspergillus nidulans propionate is oxidized to pyruvate via the methylcitrate cycle [2,3]. Propionyl-CoA is formed from propionate, CoASH and ATP catalysed by acetyl-CoA synthetase, FacA [4,5], and by an additional acyl-CoA synthetase. The condensation of propionyl-CoA with oxaloacetate inside the mitochondria yields (2S,3S)-methylcitrate [2]. Isomerization of this tricarboxylic acid, most likely via cis-2-methylaconitate [6], yields (2R,3S)-2-methylisocitrate, which is cleaved to succinate and pyruvate [3]. Studies with 13C-labelled propionate indicated that in Escherichia coli the 2-oxo acid is further oxidized to acetyl-CoA, which is either funnelled into the citrate cycle or used for biosyntheses [7].

A clue to the mechanism of propionate toxicity was the construction of an A. nidulans methylcitrate synthase deletion strain (ΔmcsA), which was unable to grow on propionate as sole carbon and energy source. Unexpectedly, growth of ΔmcsA on glucose was more inhibited by propionate than that of a wild-type strain [2]. This result indicated that (2S,3S)-methylcitrate or (2R,3S)-2-methylisocitrate are unlikely to be responsible for this inhibitory effect. At high levels of propionyl-CoA yeast citrate synthase catalyses the slow formation of three of the four stereoisomers of methylcitrate [8], but their concentrations (< 10 µm) are very low and it remains controversial whether they may be able to act as significant inhibitors. Therefore, whether the finding that methylcitrate might be the causative agent of propionate toxicity in Salmonella enterica[9] is also true for eukaryotic cells, is questionable. Nevertheless, the identification of these isomers by GLC/MS is used for diagnosis of disorders in human propionate metabolism such as propionic acidaemia and methylmalonic aciduria [10,11].

The idea that propionyl-CoA itself could be the inhibitory agent is supported by previous work on bacterial and mammalian metabolism. The inhibition of growth of the bacterium Rodopseudomonas sphaeroides by propionate was most likely caused by propionyl-CoA, which acted as an inhibitor of pyruvate dehydrogenase, competitive with CoASH, Ki = 0.84 mm. The addition of sodium bicarbonate increased the growth rate again, probably because it stimulated the degradation of propionyl-CoA via methylmalonyl-CoA [12]. It was also shown that accumulation of propionyl-CoA in rat liver hepatocytes led to a decrease in the activity of pyruvate dehydrogenase [13].

In this investigation we examined carbon balances under different growth conditions. We found that growth of A. nidulans on glucose + propionate, especially of the ΔmcsA strain, led to the excretion of pyruvate and to high intracellular concentrations of propionyl-CoA, which inhibited pyruvate dehydrogenase, succinyl-CoA synthetase (GDP forming) and ATP-citrate lyase. We conclude that these observations can explain the toxicity of propionate towards cells growing on glucose as sole carbon and energy source. Furthermore, we were able to show a correlation between inhibition of polyketide formation and intracellular propionyl-CoA content.

Experimental procedures

Materials

Chemicals were from Sigma-Aldrich. Enzymes used for determination of acetate, glucose and pyruvate were from Roche. Columns and chromatographic media were, if not otherwise indicated, from Amersham Pharmacia Biotech.

A. nidulans strains, growth conditions and carbon balances

The A. nidulans strains used in this study are listed in Table 1. Supplemented minimal and complete media were prepared as described previously [14]. For the determination of specific enzyme activities on different carbon sources, growth times were strain and medium specific. Approximately 108 spores were used for inoculation of 100 mL medium and incubation was carried out in 250-mL flasks at 37 °C and 240 r.p.m. on a rotary shaker. On media containing 50 mm glucose as sole carbon source and 50 mm glucose + 100 mm acetate, all strains were incubated for 20 h; on 50 mm glucose + 100 mm acetate + 100 mm propionate, all strains were incubated for 23 h; on 50 mm glucose + 100 mm propionate the strains were incubated for 44 h, except strain SMB/acuA, which showed much less inhibition in the presence of propionate and was grown on this medium for 22 h. The presence of residual glucose in the medium (> 20 mm) was determined enzymatically. On 100 mm acetate and 100 mm acetate + 100 mm propionate all strains, with the exception of strain SMB/acuA, were grown for 36 and 41 h, respectively. To determine enzyme activities during growth on 100 mm propionate, we added 10 mm glucose to the medium to support initial growth. After total consumption of glucose cells were grown further for at least 12 h. Therefore, the wild-type strain was grown for 42 h, whereas the methylcitrate synthase deletion strain and the facB multi-copy strain were incubated for 94 h. Strain SMB/acuA was always grown in the presence of glucose, because the strain did not grow on acetate and growth on acetate/propionate was very poor. Therefore, we used the following composition of media and growth times: 10 mm glucose + 100 mm acetate harvest after 27 h; 10 mm glucose + 100 mm propionate harvest after 29 h; 10 mm glucose + 100 mm acetate + 100 mm propionate harvest after 29 h. Determination of the residual glucose concentration confirmed that the strains were incubated for at least 12 h after total consumption of glucose. In addition, we proved that acetate was still present under all conditions where it was used as a carbon source. Growth at all conditions and with all strains was replicated twice in order to confirm the results.

Table 1. A. nidulans strains used in this study. Strain RYQ11 was used throughout all experiments. Strain SΔmcsA1 was used in a previous work was taken as a control to confirm the results of spore colour formation, enzyme activities and carbon consumption.
StrainGenotypeSource
  1. a Two different methylcitrate synthase mutants ( Δ mcsA). FGSC, Fungal genetics stock center (http://www.fgsc.net).

SMB/acuAfacA303, yA2; veA1[2]
MH2671pabaA1; prn-309, cnxJ1[46]
Fab4-J3MH2671 cotransformed with pFAB4 and pAN222 (approx. 4–8 copies facB)[46]
A637yA2, pabaA1, pdhA1FGSC, Kansas City, KS, USA
A634yA2, pabaA1; pdhB4FGSC, Kansas City, KS, USA
A627yA2, pabaA1; pdhC1FGSC, Kansas City, KS, USA
A26biA1; veA1FGSC, Kansas City, KS, USA
SMI45yA2, pabaA1; wA3; veA1M. Krüger, Marburg, Germany
SRF200pyrG89; ΔargB::trpCΔB; pyroA4; veA1[47]
RYQ11aΔmcsA::argB, biA1; veA1N. Keller, UW-Madison, USA
mcsA1aΔmcsA::argB, pyrG89; ΔargB::trpCΔB; pyroA4; veA1[2]
SMB/B1pyrG89; ΔargB::trpCΔB; pyroA4; veA1 (alcA::mcsA, argB)[2]

For the determination of CO2 production, A. nidulans was grown at 37 °C in a 1-L gas wash bottle containing 600 mL medium (Schott, Mainz, Germany). The medium was stirred at 350 r.p.m and bubbled with CO2-free air. The CO2 was removed by washing the air with 2 m NaOH followed by sterile water to avoid the transfer of NaOH to the growth medium. The CO2 produced was trapped in a fourth wash bottle containing 400 mL 0.2 m Ba(OH)2. The insoluble BaCO3 that formed was dried at 60 °C for 20 h and weighed. Residual glucose and acetate contents in the growth medium were determined by enzymatic methods (see below). The mycelium was pressed to remove any liquid, frozen with liquid nitrogen, lyophilized, weighed, and ground to a fine powder. The CHN content of the mycelium was determined by elemental analysis (Zentrale Routineanalytik, Philipps-Universität Marburg, Lahnberge, Germany). Results from three independent samples were (%): N, 6.4 ± 0.1; C, 47.2 ± 0.3; H, 8.2 ± 0.1. Thus 1 g dried mycelium consists of 472 mg carbon equivalent to 39.3 mmol.

Sample preparation of intracellular acyl-CoA from lyophilized mycelium

The dried mycelium was ground to a fine powder in a mortar and suspended in 10 mL 2% HClO4 and 1 mL 0.1% trifluoroacetic acid. The suspension was sonicated three times for 4 min each at 70% full power and 60% pulses (Branson 250 sonifier; Branson, Dietzenbach, Germany) and neutralized to pH 4–5 by drop-wise addition of 2 m K2CO3. After incubation on ice for 15 min most of the perchloric acid was precipitated as insoluble KClO4. The solution was centrifuged at 120 000 g for 25 min and the supernatant was collected. For concentration and partial purification of the CoA-thioesters, the supernatant was applied on a C18-cartridge (Chromafix C18 ec®, 510 mg; Macherey-Nagel, Düren, Germany), previously rinsed with methanol and washed with 0.1% trifluoroacetic acid. The supernatant was slowly applied to the column and washed with 10 mL 0.1% trifluoroacetic acid. Elution was carried out with 1.5 mL 50% acetonitrile/0.1% trifluoroacetic acid and samples were collected in 2-mL micro centrifuge cups. The acetonitrile was evaporated in a Speed Vac Concentrator (Bachofer GmbH, Reutlingen, Germany) without heating and the residual volume of 200–500 µL was measured with an accuracy of ± 2 µL using a micropipette. An aliquot of the samples was used for the enzymatic determination of acetyl-CoA and propionyl-CoA concentrations.

Determination of the intracellular volume

Wet weight was determined after pressing the mycelium between several sheets of absorbent paper until no further liquid could be removed. Mycelium was dried for at least 20 h at 60 °C and weighed again; thereby 3.51 g wet cells yielded 1.0 g dry cells, the mean value of 20 independent samples.

Partial purification of ATP-citrate lyase and succinyl-CoA synthetase from A. nidulans

A. nidulans strain SMB/acuA [2] was grown for 20 h on glucose minimal medium. Mycelium was harvested over a Miracloth filter membrane (Calbiochem). The mycelium was dry-pressed for removal of residual medium and suspended in 50 mm Tris/HCl pH 8.0 containing 2 mm dithiothreitol (buffer A). The mycelium was homogenized by an Ultra Turrax (T25 basic, IKA Labortechnik, Staufen, Germany). Cells were broken by ultrasonication three times for 4 min at 80% full power and 60% pulses (Branson 250 sonifier). The extract was centrifuged at 96 000 g and the supernatant was applied to a Q-Sepharose column (Pharmacia Biotech, bed volume 25 mL), previously equilibrated with buffer A. The enzyme was eluted in buffer A with a 0–1 m NaCl gradient. Enzyme-containing fractions were checked for activity, collected and concentrated in an Amicon chamber over a PM 30 membrane (Millipore, Eschborn, Germany). Purity was sufficient for inhibition studies. Succinyl-CoA synthetase was partially purified as described above, except that buffer A did not contain dithiothreitol. No further column purification was necessary for the described activity measurements.

Enzymatic determination of glucose, acetate and pyruvate in the growth medium

Glucose concentrations were determined by the combined action of glucose oxidase (GOD, from A. niger), peroxidase (POD, from horseradish) and 2,2′-azinobis(3-ethylbenzo-6-thiazolinesulfonic acid). The test was a modification of a described procedure [15]. The composition of the test reagent was: 130 mm sodium phosphate, pH 7.0; 400 U POD (2 mg; 200 U·mg−1), 800 U GOD (4 mg; 200 U·mg−1) and 25 mg 2,2′-azinobis(3-ethylbenzo-6-thiazolinesulfonic acid), final volume 50 mL. Each assay, which contained 900 µL reagent and 100 µL sample, was incubated for 15 min at 37 °C and measured at 436 nm in a spectrophotometer. The assay was linear in a range of 0–30 µm glucose. A standard was run for every freshly prepared reagent.

Pyruvate concentrations were determined by the use of lactate dehydrogenase (LDH) from rabbit muscle. The oxidation of NADH was followed at 340 nm until no further change in absorbance was visible; ε340 =6.3 mm−1·cm−1[16]. The assay contained, in a final volume of 1 mL, 50 mm potassium phosphate pH 7.0, 0.2 mm NADH, 0.5 U LDH and 50–100 µL different dilutions of the medium.

Acetate concentrations were determined with citrate synthase and malate dehydrogenase [17]. Acetate was activated by an acetyl-CoA synthetase (ACS) from Saccharomyces cerevisiae (Roche) and the resulting acetyl-CoA was condensed with oxaloacetate by the use of citrate synthase from pig heart. Oxaloacetate was continuously provided from malate by use of NAD+ and malate dehydrogenase (MDH) from pig heart. A typical assay in a final volume of 1 mL contained (mm) 50 potassium phosphate, pH 7.0; 10 l-malate, 0.2 CoASH, 2 NAD+, 2 ATP, 4 MgCl2, 0.5 dithiothreitol, 0.5 U MDH, 0.5 U citrate synthase, 0.1 U ACS and 50–100 µL diluted medium. All components were added with the exception of MDH and citrate synthase and the resulting absorbance at 340 nm was measured (A1). MDH was added and the absorbance after reaching the equilibrium was taken as A2. Citrate synthase was added and the reaction was monitored until no further change in absorbance was visible (A3). Concentrations were calculated by the formula below [ɛ, absorbance (extinction) coefficient; d, length of light path of the cuvette], which considers the decrease of the concentration of oxaloacetate in equilibrium with l-malate during the formation of NADH (the concentrations of malate and NAD+ remain almost constant):

image

Determination of intracellular propionyl-CoA and acetyl-CoA

Concentrations of acyl-CoA were determined by the use of citrate synthase from pig heart and purified methylcitrate synthase from the overproducing A. nidulans strain SMB/B1 [2] by two independent methods. One method was performed as described above for the determination of the concentration of acetate from the growth medium. A 1-mL assay contained 50 mm potassium phosphate, pH 7.0; 10 mm l-malate, 2 mm NAD, 0.5 U MDH, 0.5 U citrate synthase, 0.5 U methylcitrate synthase and 50–100 µL sample. The concentration of acetyl-CoA was determined first by the use of citrate synthase. The reaction was followed at 340 nm until no further change in absorbance was detected. Methylcitrate synthase was added and the second change in absorbance was monitored.

The second method was based on the formation of a nitrothiophenolate (2-mercapto-5-nitrobenzoate dianion) during the reaction of 5,5′-dithiobis-(2-nitrobenzoate) (DTNB) with CoASH, which was released during the condensation of oxaloacetate with acetyl-CoA or propionyl-CoA. The assay contained, in a final volume of 1 mL, 50 mm Tris/HCl, pH 8.0; 1 mm oxaloacetate, 1 mm DTNB, 0.5 U citrate synthase, 0.5 U methylcitrate synthase and 20–100 µL sample. Change in absorbance was monitored at 412 nm; ε = 14.2 mm−1·cm−1[18,19]. Acetyl-CoA concentrations were determined first. When no further change in absorbance was visible, methylcitrate synthase was added.

Enzyme assays

ATP citrate lyase.  The assay [20] contained (mm) 50 Tris/HCl, pH 8.0; 0.2 NADH, 5 ATP, 0.34 CoASH, 20 citrate, 2 dithiothreitol, 2 MgCl2, 0.5 U MDH from pig heart, enzyme sample and water to a final volume of 1 mL. The reaction was started by addition of enzyme sample and decrease in absorbance at 340 nm was monitored. One unit of enzyme activity was defined as the amount of enzyme reducing 1 µmol NAD·min−1 under the assay conditions.

Succinyl-CoA synthetase  was measured by a modified method for the determination of citrate synthase activity [21]. A typical assay contained 50 mm Tris/HCl, pH 7.5; 0.14 mm succinyl-CoA, 1 mm DTNB, 0.5 mm GDP, 2 mm MgCl2, 5 mm potassium phosphate, enzyme sample and water to a final volume of 1 mL. One unit of enzyme activity was defined as the amount of enzyme producing 1 µmol CoASH·min−1 under the assay conditions.

Isocitrate lyase.  The assay [22] contained (mm) 50 potassium phosphate, pH 7.0; 1 threo-isocitrate, 10 phenylhydrazine HCl, 2 dithiothreitol, 2 MgCl2, 10–100 µL enzyme sample and water to a final volume of 1 mL. The formation of glyoxylate phenylhydrazone was followed at 324 nm; ε = 16.8 mm−1·cm−1. One unit of enzyme activity was defined as the amount of enzyme producing 1 µmol glyoxylate phenylhydrazone·min−1 under the assay conditions.

2-Methylisocitrate lyase.  The assay was based on the reduction of pyruvate with NADH catalysed by LDH, whereby the decrease in absorbance at 340 nm was recorded [3]. The composition of the reaction was (mm) 0.20 threo-2-methylisocitrate, 2 MgCl2, 2 dithiothreitol, 0.2 NADH, 1.5 U LDH, 50 potassium phosphate, pH 7.0; enzyme sample and water to a final volume of 1 mL. One unit of enzyme activity was defined as the amount of enzyme producing 1 µmol NADH·min−1 under the assay conditions.

Citrate synthase and methylcitrate synthase.  Citrate synthase and methylcitrate synthase activity was determined as described previously [2]. The reaction mixture contained (in mm), in a final volume of 1 mL, 50 Tris/HCl, pH 8.0; 1.0 5,5′-dithiobis-(2-nitrobenzoic acid), cell-free extract and 0.2 acetyl-CoA or propionyl-CoA, respectively. The assay was started by the addition on 1 mm oxaloacetate (final concentration) and monitored at 412 nm. One unit of enzyme activity was defined as the amount of enzyme producing 1 µmol CoASH·min−1 under the assay conditions.

Pyruvate dehydrogenase.  Pyruvate dehydrogenase (PDH) activity was measured according to a procedure described previously [23] with some modifications. The assay contained (in mm), in a final volume of 1 mL, 50 Tris/HCl, pH 8.0; 2 pyruvate, 0.8 thiamine pyrophosphate, 2.5 cysteine/HCl, 2 NAD, 2 MgCl2, cell-free extract and water to a final volume of 990 µL. The reaction was started by the addition of 0.02–0.17 mm CoASH and reduction of NAD+ to NADH was followed at 340 nm. One unit of enzyme activity was defined as the amount of enzyme producing 1 µmol NADH·min−1 under the assay conditions. The activity of 2-oxoglutarate dehydrogenase was determined by the analogous procedure, in which pyruvate was replaced by 2-oxoglutarate [24].

Acetyl-CoA synthetase.  Acetyl-CoA synthetase activity was determined in a coupled assay by the use of MDH and citrate synthase. In this method the acetyl-CoA produced reacts via citrate synthase with oxaloacetate, which is provided by MDH from malate. The assay contained (in mm), in a final volume of 1 mL, 50 potassium phosphate buffer, pH 7.0; 10 sodium acetate, 2 NAD, 20 d,l-malate, 0.4 CoASH, 2 dithiothreitol, 4 MgCl2, 6 U MDH (pig heart, Roche), 2 U citrate synthase (pig heart, Roche), cell-free extract and water to a final volume of 980 µL. The reaction was started by the addition of 20 µL of a 100 mm ATP solution (final concentration 2 mm) and the reduction of NAD was monitored at 340 nm. The extincition coefficient was set as 0.5 × 6.3 mm−1·cm−1, which compensates for the initial decrease of the oxaloacetate concentration in the equilibrium due to the accumulation of NADH [25]. Lineweaver–Burk diagrams were obtained by use of the worksheet of the program exel 98 (Microsoft Inc.).

Propionyl-CoA synthetase.  Propionyl-CoA synthetase activity was determined by the same method as described for the determination of acetyl-CoA synthetase activity, except sodium acetate was replaced by sodium propionate and citrate synthase by methylcitrate synthase (0.8 U) from A. nidulans[2].

CoA-Transferase.  CoA-Transferase activity was determined by using succinyl-CoA or propionyl-CoA as the CoA-donor and acetate or propionate as the acceptor. When acetate was the acceptor the assay was monitored by the use of citrate synthase, which released CoASH upon the condensation of newly generated acetyl-CoA with oxaloacetate as described for the determination of citrate synthase activity. When propionate was used as the acceptor, purified methylcitrate synthase was used to measure the CoASH release upon the condensation of propionyl-CoA with oxaloacetate as described for the determination of methylcitrate synthase activity. A typical assay contained (in mm), in a final volume of 1 mL, 50 Mops, pH 7.5; 0.4 CoA-donor (succinyl-CoA or propionyl-CoA, respectively), 2 U citrate synthase or 0.8 U methylcitrate synthase, respectively, 1 oxaloacetate and 10 CoA-acceptor (acetate or propionate, respectively) and cell-free extract.

Oxidative branch of the pentose phosphate pathway.  This was determined by the use of glucose-6-phosphate as the substrate and NADP as the hydrogen acceptor. Due to the use of cell-free extracts, not only the activity of glucose-6-phosphate dehydrogenase but also the activity of the 6-phosphogluconate dehydrogenase was measured. The described method was slightly modified [26]. A typical assay in a final volume of 1 mL contained (in mm) 50 Mops, pH 7.5; 1 glucose-6-phosphate, 1 NADP, 5 EDTA and cell-free extract. The reaction was monitored at 340 nm and specific activity was defined as the reduction of 2 µmol NADP·min−1·mg protein−1.

Determination of maintenance

In order to calculate the amount of glucose used for maintenance, the wild-type strain A26 was used. Four 100-mL aliquots of glucose minimal media in 250-mL flasks were inoculated with 4 × 108 spores and incubated for 13 h at 37 °C and 240 r.p.m. Two of the samples were harvested and dried at 70 °C to measure biomass formation as a control. The other two samples were washed with sterile 0.6 m KCl and transferred to fresh glucose minimal medium containing cycloheximide (200 µg·mL−1), which inhibits eucaryotic protein biosynthesis. The cultures were incubated for further 9 h at 37 °C and 240 r.p.m. The mycelium was dried and the biomass was compared to that of control samples. Glucose concentrations before and after the incubation with cycloheximide were measured as described above.

Results

Carbon balances on different growth media

Initial experiments showed that growth on glucose + propionate resulted in significant excretions of pyruvate into the medium (Table 2). In order to exclude substantial excretions of other carbon compounds, we measured the total carbon balances of wild-type and methylcitrate synthase deletion strain (ΔmcsA). Therefore, the consumption of substrates, formation of CO2, as well as excretion of pyruvate and the final pH were determined in media in which cells had been grown on different carbon sources. The measured carbon balances add up to almost 100% (Table 3) indicating that there was no substantial excretion of compounds other than CO2 and pyruvate or a significant consumption of propionate. The increase in the final pH (Table 2) correlated with the consumption of the carboxylates, by which protons are removed from the medium, whereas by oxidation of glucose no change in pH was observed. When grown only on glucose there was no significant difference between the wild-type and the ΔmcsA strain. In the presence of only acetate there was no difference between the strains; the approximate growth rate was only 50% of that with glucose and the increase in pH from 6.4 to 8.2 correlated with the high consumption of acetate. Growth on propionate alone was not included in this study, as the growth rate of the wild-type was extremely low and the morphology of the mycelium was quite different. Furthermore, on propionate the ΔmcsA strain did not grow at all.

Table 2. Carbon consumption and pyruvate excretion of wild-type and ΔmcsA strain under different growth conditions. The wild-type strain was SMI45 and the initial pH was 6.3–6.5. Consumption and excretion are data are given in mmol substrate·g dried mycelium−1. In all experiments the concentration of glucose was 50 mm and that of sodium propionate100 mm. The concentration of sodium acetate was 50 mm except when used in combination with propionate in which case 100 mm was used. Mycelia were harvested in the linear growth phase. ΔmcsA, methylcitrate synthase deletion mutant (RYQ11 and SΔmcsA1). For experiments marked by an asterisk see also Table 3.
StrainC-Source (final pH at harvest of mycelium) Glucose consumption (mmol/g) Acetate consumption (mmol/g) Pyruvate excretion (mmol/g)Growth time (h)
Wild-type *Glucose (6.6)10.60.14020
ΔmcsA*Glucose (6.7)9.90.05420
Wild-typeGlucose/acetate (7.3)10.01.00.07022
ΔmcsAGlucose/acetate (7.9)8.09.00.08722
Wild-type *Glucose/propionate (6.8)14.61.2744
ΔmcsA*Glucose/propionate (6.3)16.22.2172
Wild-typeGlucose/propionate/acetate (7.5)7.0240.3730
ΔmcsAGlucose/propionate/acetate (7.4)11.0190.5030
Wild-typeAcetate/propionate (8.0)490.14447
ΔmcsAAcetate/propionate (8.5)620.03547
Wild-typeAcetate (8.2)54< 0.0140
ΔmcsAAcetate (8.2)55< 0.0140
Table 3. Carbon balances of wild-type and ΔmcsA strain. Balances are calculated for 1 g of dried mycelium. The concentrations of the substrates are indicated in Table 2 (marked by asterisks). The wild-type strain was SMI45 and ΔmcsA strains were RYQ11 and SΔmcsA1.
Strain/C-sourceGlucose consumed (mmol C)Pyruvate (mmol C)CO2 recovered (mmol C)Biomass (mmol C)Total amount recovered [mmol C (%)]
Wild-type/glucose64 ± 4021 ± 43960 ± 4 (94)
ΔmcsA/glucose60 ± 4020 ± 43959 ± 4 (98)
Wild-type/glucose + propionate88 ± 4340 ± 23982 ± 2 (93)
ΔmcsA/glucose + propionate97 ± 4649 ± 43994 ± 4 (97)

Addition of acetate to a medium containing glucose did not change the growth rate significantly, but the lack of methylcitrate synthase in the mutant strain induced acetate consumption (Table 2). This observation is similar to strain Fab4-J3, which carries multiple copies of the transcriptional activator FacB of the acetate utilization genes. FacB is induced by acetate and acetylcarnitine [27]. Growth experiments with strain Fab4-J3 revealed that in the presence of both glucose and acetate, the latter substrate is mainly used. Thus cells grown on 50 mm glucose + 100 mm acetate consumed only 2.7 mmol glucose but 44.2 mmol acetate·g dried cells−1. That means that the higher basal level of the transcriptional activator FacB in a strain, which carries multiple integrations of the facB-gene in the genome, leads to preferred use of acetate as carbon source.

From our results we can conclude that propionate or an intermediary metabolite, most likely propionyl-CoA, is able to induce genes from propionate as well as from acetate metabolism (Table 4, see Icl, Micl and McsA). Therefore, in the ΔmcsA strain, accumulation of propionyl-CoA, derived from amino acid degradation, can cause the higher consumption of acetate as compared to the wild-type.

Table 4. Specific enzyme activities from cell-free extracts of different strains and growth conditions. Data are given in mU·mg protein−1. Acs, acetyl-CoA synthetase; Pcs, propionyl-CoA synthetase; Icl, isocitrate lyase; Micl, 2-methylisocitrate lyase; McsA, methylcitrate synthase. C-sources: G, glucose; A, acetate; P, propionate. Numbers denote the concentrations of C-sources (mm); G50/A100/P100 = 50 mm glucose + 100 mm acetate + 100 mm propionate.
EnzymeC-Source in mediumWild-type (A26)Fab4-J3ΔmcsASMB/acuA
AcsG5019 ± 319 ± 216 ± 30.5 ± 0.2
AcsG50/A10047 ± 4119 ± 1026 ± 41 ± 0.2
AcsG50/P10022 ± 254 ± 224 ± 22.3 ± 0.2
AcsG50/A100/P10059 ± 1137 ± 1027 ± 32 ± 0.3
AcsA100153 ± 5205 ± 10124 ± 217 ± 1
AcsG10/P100133 ± 4128 ± 10150 ± 1022 ± 2
AcsA100/P100135 ± 10289 ± 15167 ± 1018 ± 2
PcsG5010 ± 19 ± 18 ± 11 ± 0.5
PcsG50/A10016 ± 250 ± 513 ± 22.3 ± 0.2
PcsG50/P10010 ± 121 ± 110 ± 16 ± 0.4
PcsG50/A100/P10026 ± 238 ± 413 ± 13.5 ± 0.5
PcsA10058 ± 167 ± 342 ± 229 ± 2
PcsG10/P10077 ± 263 ± 376 ± 631 ± 1
PcsA100/P10059 ± 190 ± 274 ± 130 ± 1
IclG500.2 ± 0.10.1 ± 0.10.2 ± 0.10.6 ± 0.2
IclG50/A10023 ± 1108 ± 414 ± 27 ± 1
IclG50/P10035 ± 262 ± 941 ± 124 ± 2
IclG50/A100/P10085 ± 3170 ± 534 ± 326 ± 1
IclA10086 ± 5225 ± 563 ± 371 ± 4
IclG10/P100130 ± 5107 ± 7294 ± 1067 ± 5
IclA100/P100161 ± 1287 ± 15180 ± 1082 ± 5
MiclG507 ± 16 ± 26 ± 24 ± 1
MiclG50/A10010 ± 112 ± 19 ± 211 ± 1
MiclG50/P10030 ± 231 ± 262 ± 444 ± 1
MiclG50/A100/P10026 ± 127 ± 129 ± 233 ± 1
MiclA10026 ± 120 ± 129 ± 224 ± 1
MiclG10/P10074 ± 528 ± 2132 ± 164 ± 1
MiclA100/P10035 ± 136 ± 246 ± 163 ± 3
McsAG501 ± 02 ± 101 ± 0
McsAG50/A1005 ± 114 ± 207 ± 1
McsAG50/P10055 ± 252 ± 2057 ± 4
McsAG50/A100/P10037 ± 138 ± 1041 ± 1
McsAA10038 ± 220 ± 1042 ± 3
McsAG10/P100147 ± 672 ± 30153 ± 5
McsAA100/P10035 ± 183 ± 10133 ± 6

A dramatic effect on the growth rate was observed when propionate was added to the glucose medium; the growth time doubled with the wild-type and increased 3.6× with the ΔmcsA mutant. In both strains propionate caused an increase in glucose consumption and a huge enhancement of pyruvate excretion. The carbon balance excluded a significant excretion of other substances such as alanine [28], which may have escaped our analytical tools. Furthermore we found that the observed additional amount of consumed glucose was almost completely oxidized to CO2 (Table 3). Probably the increase in CO2 production caused by propionate (doubled with the wild-type and tripled with the mutant) was due to energy production required for maintenance (see below) during the extended growth times.

Upon addition of acetate to the media containing glucose and propionate, the growth rate of both strains increased and the effect of propionate became less apparent. Finally, in media containing acetate and propionate but no glucose, there was only a small delay (30%) in growth of the mutant as compared to the wild-type [2]. The higher acetate consumption of the mutant strain was probably due to higher maintenance requirement (see below) or to the action of a CoA-transferase, which is induced by propionate and seems to transfer the CoA-moiety from succinyl-CoA preferentially to acetate (see below and Table 5).

Table 5. CoA-transferase activity from wild-type and ΔmcsA grown on different carbon sources. Data are given in mU·mg protein−1; 1 U is defined as the release of 1 µmol CoASH·min−1 under the assay conditions. The wild-type strain was A26 and ΔmcsA was RYQ11. Succinyl-CoA > acetate, succinyl-CoA:acetate CoA-transferase; Succinyl-CoA > propionate, succinyl-CoA:propionate CoA-transferase; Propionyl-CoA > acetate, propionyl-CoA:acetate CoA-transferase.
CoA-donor > acceptorMedium used for growth
GlucoseGlucose/ acetateGlucose/ propionateGlucose/acetate/ propionateAcetatePropionate
Succinyl-CoA > acetate (WT)5.83878498665
Succinyl-CoA > acetate (ΔmcsA)12.74111563109143
Succinyl-CoA > propionate (WT)2.412.424153225
Succinyl-CoA > propionate (ΔmcsA)5.114.746162854
Propionyl-CoA > acetate (WT)< 0.54.49.65.69.65.3
Propionyl-CoA> acetate (ΔmcsA)0.95.811.77.18.713.3

The observed excretion of pyruvate prompted us to check strains, in each of which another of the three genes encoding pyruvate dehydrogenase [29] was mutated (A637, pdhA1-mutant = lipoate acetyltransferase; A634, pdhB4 = β-subunit of pyruvate decarboxylase; A627, pdhC1 = α-subunit of pyruvate decarboxylase). All three strains were unable to grow on glucose or propionate, but grew well on acetate. Growth of strain A627 on 50 mm acetate yielded 239 mg dried mycelium after 23 h (59 mmol acetate·g mycelium−1). Interestingly, growth of this mutant was enhanced rather than inhibited by the addition of 50 mm glucose, which led to the production of 313 mg mycelium in 23 h, whereby 26 mmol acetate and 4 mmol glucose were consumed and 0.9 mmol pyruvate were excreted. This can be explained by the fact that production of cell mass from glucose requires less ATP than from acetate, because the energy consuming gluconeogenesis via the glyoxylate cycle is not necessary. On the other hand consumption of acetate together with glucose was not expected, since CreA regulation should prohibit such a cometabolism. In the presence of glucose the wide-domain regulatory protein CreA forms a complex with target DNA binding sites and leads to a reduced transcription of genes coding for degradation of alternative carbon sources [30]. However, we cannot exclude the spontaneous formation of creA mutants, which derive from our cultivation conditions. This event would lead to a relieved carbon catabolite repression as also shown for other glyoxylate cycle mutants [5].

Determination of maintenance

Maintenance is the energy that is used for survival of cells without any biomass formation. Determination of maintenance was based on the inhibition of protein biosynthesis by the action of cycloheximide. Cycloheximide binds to the 80S-subunit of eukaryotic ribosomes and prevents the initiation and elongation reaction of protein biosynthesis. The mycelium of pregrown cultures was washed and transferred to fresh medium containing cycloheximide (200 µg·mL−1), which was sufficient to prevent biomass formation. Cultures were incubated for 8 h and dry mass as well as glucose consumption was determined. In this experiment significant glucose consumption was observed (8.75 ± 0.1 mmol·h−1·g dried cells−1). We conclude that indeed the prolonged growth time of both the wild-type and ΔmcsA strains on glucose/propionate medium led to the increased consumption of glucose as determined.

Intracellular acetyl-CoA and propionyl-CoA contents

To investigate whether propionyl-CoA accumulates in the methylcitrate synthase deletion strain during growth on different carbon sources, mycelium was harvested, directly frozen in liquid nitrogen and lyophilized. After opening the cells by sonication in the presence of perchloric acid, CoA-thioesters were partially purified and determined enzymatically as described in Experimental procedures. The suitability of this method was checked by mixing 16.5 nmol acetyl-CoA and 16.1 nmol of propionyl-CoA and performing the identical procedure as for the partial purification of the acyl-CoA ester from lyophilized mycelium, including addition of perchloric acid, neutralization, centrifugation, C18-cartridge and concentration. The recovery was 15.1 nmol (91.5%) acetyl-CoA and 14.4 nmol (89.5%) propionyl-CoA which showed that the method gave reliable results. Therefore we can conclude that the ratio between acetyl-CoA and propionyl-CoA remained constant during the procedure and the total yield was about 90% assuming that all cells were opened by the procedure described above.

After 20 h of growth on glucose as the sole carbon source, neither the wild-type nor the methylcitrate synthase deletion strain showed significant accumulation of propionyl-CoA (Fig. 1). Addition of propionate to the glucose medium led to an increase of the propionyl-CoA level in the wild-type strain. The methylcitrate synthase deletion strain showed an up to tenfold higher accumulation of propionyl-CoA under these conditions, as the thioester cannot be oxidized further. Addition of acetate to the glucose/propionate medium reduced the propionyl-CoA level of the cells, whereas an increase was observed again after growth on acetate + propionate without glucose. Despite this high level of propionyl-CoA, which was most probably due to an unspecific action of acetyl-CoA synthetase (described below), only a slight growth inhibition was visible [2] and Table 2. Remarkably, under the different growth conditions the intracellular acetyl-CoA concentrations were kept constant in a relatively narrow range (20–60 nmol·g−1 dried cells), even in the mutant strain.

Figure 1.

Intracellular contents of acetyl-CoA and propionyl-CoA from A. nidulans wild-type and ΔmcsA strain grown under different conditions. Carbon and energy sources were: 50 mm glucose; 50 mm glucose and 100 mm sodium propionate; 50 mm glucose, 100 mm sodium acetate, and 100 mm sodium propionate; 100 mm sodium acetate and 100 mm sodium propionate. The CoA-thioesters were released from the cells and determined as described in Experimental procedures.

Determination of the intracellular volume

In order to obtain the intracellular concentration of accumulated acyl-CoA esters, it was necessary to know the internal volume in relation to the mass of dried mycelium. The easiest way to calculate this volume was to measure the water content from the difference between the mass of wet and dry A. nidulans cells. Thus the internal volume was determined to be 2.51 ± 0.13 ml·g dry cells−1, which is in good agreement with that of Neurospora crassa (2.54 mL·g dried cells−1) [31]. Investigations on the intracellular concentrations of different metabolites of A. niger considered only the free intracellular water not bound to proteins, rather than the total water content, which was also similar to that of N. crassa. This content of free water was determined as 1.20 mL·g dried mycelium−1 by the use of xylitol and showed that ≈ 50% of the intracellular water is not available as a solvent for metabolites [32]. We therefore used this latter value for the calculation of the internal propionyl-CoA concentration of the methylcitrate synthase mutant and the wild-type after growth on 50 mm glucose + 100 mm propionate. Thus the ΔmcsA strain accumulated 0.21 mm propionyl-CoA, whereas in the wild-type strain only 0.03 mm propionyl-CoA could be found. Nevertheless, concentrations given here are just a simple mathematical calculation. Due to the very high concentration of macromolecules within the cell, accompanied by high viscosity, local concentrations may differ from that shown here. In addition, propionyl-CoA is supposed to be generated in the cytoplasm. For transport to the mitochondria a conversion into a carnitine-ester and a back-conversion to the CoA-ester inside the mitochondria has to be involved, which is most likely performed by cytoplasmic and mitochondrial acyl-carnitine transferases (AcuJ [33] and FacC [27]). The transporter involved in that process is most likely AcuH [34]. Mutants of the corresponding genes were unable to grow on propionate as sole carbon and energy source (data not shown). This transport mechanism implies a higher concentration of propionyl-CoA within the mitochondria. However, the fact that propionyl-CoA cannot be converted in a methylcitrate synthase deletion strain would lead to the formation of an equilibrium between propionyl-CoA and propionyl-carnitine in mitochondria and cytoplasm. Since the equilibrium constant between these two propionate esters is close to 1.0, we assume for our calculations that the concentration of propionyl-CoA is similar in all compartments.

Formation of acetyl-CoA and propionyl-CoA

For the determination of the substrate specificity of acetyl-CoA synthetase and a putative propionyl-CoA synthetase we used the acetate-grown strain Fab4-J3 and glucose/propionate grown SMB/acuA cells (10 mm glucose/100 mm propionate; 29 h). The high expression of the acetate utilization genes in the Fab4-J3 strain seemed to be suitable to measure mainly the acetate and propionate activating activity of acetyl-CoA synthetase. In comparison SMB/acuA carries a defective acetyl-CoA synthetase gene, which means that the activating activity must derive from alternative acyl-CoA synthetases, most likely a propionyl-CoA synthetase.

The kinetic constants were determined with an extract from acetate grown Fab4-J3 cells with acetate as substrate: Vmax = 205 mU·mg−1 protein and Km = 44 µm (Vmax/Km = 4700 U·g−1·mm−1); with propionate as substrate the values were: Vmax = 67 mU·mg−1 and Km = 640 µm (Vmax/Km = 100 U·g−1·mm−1); hence the enzyme is 47 times more specific for acetate than for propionate. In comparison, an extract from propionate grown SMB/acuA cells gave following values with acetate as substrate: Vmax = 22 mU·mg protein−1 and Km = 880 µm (Vmax/Km = 25 U·g−1·mm−1) and with propionate as substrate: Vmax = 31 mU mg protein−1 and Km = 90 µm (Vmax/Km = 344 U·g−1·mm−1); specificity ratio of acetate: propionate = 0.073. These data indicate that A. nidulans possesses both a highly active specific acetyl-CoA synthetase, and at least one additional synthetase which prefers propionate 14 times over acetate as substrate. The existence of two functional acetyl-CoA synthetases, ACS1 and ACS2, displaying different kinetics towards propionate, has also been shown in Sc. cerevisiae[35]. Furthermore, some bacteria such as E. coli and Salmonella typhimurium carry a specific propionyl-CoA synthetase, which is distinct from the acetyl-CoA synthetase [36]. A candidate for such a propionyl-CoA synthetase from A. nidulans is the hypothetical protein AN5833.2 (Accession No. EAA58342) from the conceptual translation of the A. nidulans genome (http://www.broad.mit.edu/annotation/fungi/aspergillus/geneindex.html). The protein possesses a conserved AMP-binding domain, which is also present in acetyl-CoA synthetases and shows 63% similarity (43% identity) to propionyl-CoA synthetases from bacterial sources such as Brucella melitensis (Accession No. AAL51488) or Vibrio parahaemolyticus (Accession No. BAC59907).

To determine the extent of acetate activation in comparison to propionate activation in the presence of both substrates we used the wild-type strain A26 grown on a medium containing 100 mm acetate + 100 mm propionate (Table 6). The cell-free extract was used to determine the inhibition of acetyl-CoA synthetase activity by propionate. The acetyl-CoA formed was measured in a coupled assay with citrate synthase, which displays no significant activity with propionyl-CoA. Therefore we exclusively monitored the activity for activation of acetate. In the presence of 0.5 mm acetate and 10 mm propionate (ratio 1 : 20) we observed still 50% acetyl-CoA synthetase activity. Therefore we conclude that in a wild-type background the activation of acetate is much favoured over the activation of propionate or, vice versa, acetate inhibits the formation of propionyl-CoA. This observation readily explains the decreased propionyl-CoA levels found in cells grown on glucose/acetate/propionate as compared to glucose/propionate.

Table 6. Acetyl-CoA synthetase activity from wild-type strain A26 grown on 100 mm acetate + 100 mm propionate. 100% acetyl-CoA synthetase activity refers to (135 ± 10) mU·mg protein−1.
SubstratesActivity (%)
Acetate (mm)Propionate (mm)Ratio Acetate : propionate
100100
60512 : 191
10201 : 286
10401 : 475
5401 : 861
1101 : 1066
0.5101 : 2050
0.1101 : 10025

Inhibition of CoASH-dependent enzymes of glucose metabolism

The high levels of propionyl-CoA in the mutant strain raised the question of whether the thioester might inhibit CoA-dependent enzymes in glucose metabolism. Initial experiments showed that pyruvate dehydrogenase, ATP citrate lyase and succinyl-CoA synthetase were inhibited by propionyl-CoA, but that 2-oxoglutarate dehydrogenase and also the acetyl-CoA dependent citrate synthase exhibited no effect with propionyl-CoA.

Pyruvate dehydrogenase.  In order to investigate the inhibitory effect of propionyl-CoA on the in vitro activity of the pyruvate dehydrogenase complex, cell-free extracts of glucose-grown wild-type cells (strain A26) were used. Activity was monitored by the reduction of NAD+ in the presence of pyruvate and CoASH. At low concentrations of CoASH (0.021 mm) and relatively high propionyl-CoA concentrations (0.32 mm) the formation of NADH from the complex was inhibited by 88%. At equimolar concentrations of both (0.17 mm CoASH and 0.16 mm propionyl-CoA), the inhibitory effect of propionyl-CoA was still around 50%. The Km for CoASH (7.2 µm) increased in the presence of 0.1 mm propionyl-CoA 3.6-fold (25 µm), whereas Vmax was reduced only by 30%, which demonstrated a mainly competitive inhibition with an apparent Ki of ≈ 50 µm. Addition of high concentrations of propionate (20 mm) did not produce any significant inhibition. Therefore we can conclude that the excretion of pyruvate during growth on glucose/propionate medium is caused by a direct inhibition of the pyruvate dehydrogenase complex by propionyl-CoA. Furthermore, the elevated pyruvate excretion of the methylcitrate synthase mutant is in agreement with the higher intracellular propionyl-CoA concentrations.

ATP citrate lyase and succinyl-CoA synthetase.  In order to measure the activities of ATP citrate lyase and succinyl-CoA synthetase more precisely, we partially purified both enzymes by chromatography over a Q-Sepharose column. Inhibition of ATP citrate lyase by acetyl-CoA, propionyl-CoA and butyryl-CoA was measured by addition of different concentrations of single acyl-CoA to the in vitro assay in the presence of 0.34 mm CoASH. Activity without addition of acyl-CoA (10 mU·mL−1) was set to 100%(Fig. 2A). Propionyl-CoA showed the strongest inhibitory effect, followed by acetyl-CoA and butyryl-CoA.

Figure 2.

Inhibition of ATP citrate lyase(A) and succinyl-CoA synthetase(B) from A. nidulans by different CoA-thioesters. Both enzymes were partially purified by chromatography over Q-Sepharose. Activity without addition of CoA-thioesters (≈ 10 mU·mL−1) was set as 100%.

Succinyl-CoA synthetase.  Succinyl-CoA synthetase (10 mU·mL−1) was assayed with succinyl-CoA, inorganic phosphate and GDP by trapping the liberated CoASH with 5,5′-dithiobis-2-nitrobenzoate (Fig. 2B). At concentrations of 0.4 mm acetyl-CoA or 0.4 mm propionyl-CoA the succinyl-CoA synthetase was inhibited by 70%. A combination of 0.2 mm acetyl-CoA and 0.4 mm propionyl-CoA, however, caused a 95% inhibition, whereas in the presence of 0.6 mm acetyl-CoA the inhibition was only 80%. Therefore, accumulation of propionyl-CoA in the mutant strain (≈ 0.2 mm) might lead to a partial block of the citric acid cycle at the level of succinyl-CoA synthetase.

CoA-transferase activity

As mentioned above, succinyl-CoA synthetase is almost completely blocked by the combined action of propionyl-CoA and acetyl-CoA. In the presence of both thioesters one might expect an accumulation of succinyl-CoA in the cell and a deadlock of further reactions of the citric acid cycle. The carbon balances revealed, however, that glucose is almost completely decomposed to CO2 and, furthermore, the oxidation of acetate is not inhibited by propionate. Therefore, we searched for an alternative reaction converting succinyl-CoA into succinate. For this purpose we determined the ability of cell-free extracts to transfer the CoA-moiety from succinyl-CoA to acetate or propionate as well as the ability to decompose propionyl-CoA by the transfer of the CoA-moiety to acetate by the action of a CoA-transferase. The wild-type and the methylcitrate synthase deletion strain were grown on different carbon sources and the presence of such a CoA-transferase was tested using succinyl-CoA + acetate, succinyl-CoA + propionate and propionyl-CoA + acetate as substrates (Table 4). In both strains highest CoA-transferase activity was determined by use of succinyl-CoA as the CoA-donor and acetate as the acceptor, followed by the transfer from succinyl-CoA to propionate (≈ 35% of the former activity) and the transfer from propionyl-CoA to acetate (≈ 11%). The enzyme was most active in strains grown in the presence of propionate and always higher in the ΔmcsA strain as compared to the wild-type. These CoA-transferase levels resemble the expression pattern of the gene encoding 2-methylisocitrate lyase, a specific enzyme of the methylcitric acid cycle (compare Table 4 to Table 3). Therefore, we conclude that an efficient transfer of the CoA-moiety from succinyl-CoA to acetate in the presence of both acetate and propionate is possible. In addition this might explain the low accumulation of propionyl-CoA during growth on glucose/acetate/propionate medium especially of the ΔmcsA strain, which is consistent with the higher growth rate and the elevated acetate consumption of both strains (Table 1). In the absence of acetate (glucose/propionate medium) the CoA-moiety, however, can only be transferred to propionate, which would on the one hand enable a completion of the citric acid cycle, but on the other hand produce even more propionyl-CoA, which accumulates especially in the ΔmcsA strain. CoA-transferases are already known from pro- and eukaryotic sources. However, the transfer of the CoA-moiety from succinyl-CoA to acetate or propionate has not been shown before in any organism at a reasonable rate. Further investigations on a purified enzyme will need to prove the substrate specificity and intracellular localization of the enzyme to manifest these observations.

The oxidative branch of the pentose phosphate pathway

We mentioned above that cells grown on glucose/propionate medium released approximately twice as much CO2 for the formation of 1 g dried mycelium; we attributed this mainly to the reduced growth rate and the consequent high consumption via maintenance (8 mmol·g−1·h−1). Another explanation of this apparent ‘uncoupling’ of glucose oxidation from growth could be the pentose phosphate cycle, in which no ATP is conserved. This pathway is involved in the metabolism of glucose and is essential for the generation of NADPH and ribose, which are necessary for biosynthetic processes such as fatty acid and nucleotide synthesis. If only NADPH is required, glucose can be completely oxidized via this pathway to CO2 without ATP formation. It was demonstrated that glucose-6-phosphate dehydrogenase, the first enzyme of this pathway, is essential for the viability of fungal cells, most likely due to its important biosynthetic role [37,38]. As shown in Table 7, A. nidulans contains relatively high amounts of glucose-6-phosphate dehydrogenase and gluconate-6-phosphate dehydrogenase, which were measured together in the same assay. The data indicate that the presence of propionate in the medium reduces the activity by ≈ 50% in the wild-type as well as in the ΔmcsA strain. Therefore it appears unlikely that an enhanced oxidation of glucose via the pentose phosphate cycle is responsible for the observed uncoupling of glucose oxidation and growth inhibition caused in the presence of propionate.

Table 7. Determination of the oxidative steps of the pentose phosphate pathway. Wild-type and ΔmcsA were grown on different carbon sources and the combined activity of glucose-6-phosphate dehydrogenase and gluconate-6-phosphate dehydrogenase was determined. One unit (U) is defined as the reduction of 1 µmol of NADP+ per min. The wild-type strain was A26 and ΔmcsA was RYQ11.
Growth conditionWild-type (U·mg protein−1)ΔmcsA (U·mg protein−1)
Glucose1.351.36
Glucose/acetate1.121.05
Glucose/propionate0.870.73
Glucose/acetate/propionate0.851.18

Correlation of spore colour formation to propionyl-CoA levels and enzymatic activities

The spore colour of conidia from A. nidulans derives from the polyketide naphtopyrone [39]. We have assumed a negative effect of propionyl-CoA on spore colour formation in an earlier study, without the knowledge about the accumulation of propionyl-CoA [2]. Recently, by screening for A. nidulans mutants with a defect in the synthesis of the polyketide sterigmatocystin (ST) a methylcitrate synthase deletion strain was identified. Further analysis of this mutant showed that it was not only disturbed in ST production but also in the formation of ascoquinone A, a polyketide, which is responsible for the red pigment of sexual spores (ascospores). Both polyketides are formed under conditions when carbon sources become limited (≥ 70 h of growth). Therefore, an accumulation of propionyl-CoA was predicted, which derives from the degradation of amino acids such as isoleucin, valine and methionine during starvation [40]. In this study we tried to correlate the inhibition of spore colour formation directly to the level of propionyl-CoA under different growth conditions.

In A. nidulans spore colour formation is prevented especially in a methylcitrate synthase deletion strain by the addition of propionate (Fig. 3, lines III, IV, V and VI). This effect is not observed upon the addition of acetate to the growth medium (Fig. 3, line II) and implies that the presence of propionyl-CoA or methylmalonyl-CoA inhibits polyketide synthases, for which fungi apparently only use acetyl-CoA and malonyl-CoA as substrates. As shown in lines III–VI of Fig. 3, the addition of increasing amounts of propionate also affects the wild-type and the facB multi-copy strain but not strain SMB/acuA, which carries a defective acetyl-CoA synthetase (n.b. acuA = facA). The order of the inhibitory effect on spore colour formation was: methylcitrate synthase deletion strain, followed by the facB multi-copy strain and the wild-type. This observation is in agreement with the activities for propionate activation in comparison to methylcitrate synthase activity (Table 3: compare Pcs and McsA on media G50/P100 and G10/P100). Strain SMB/acuA shows lowest propionyl-CoA synthetase activity but significant methylcitrate synthase activity. The facB multi-copy strain shows elevated propionyl-CoA synthetase activity without increasing methylcitrate synthase activity and therefore reacts more sensitively than the wild-type. However, it is noteworthy that strain Fab4-J3 in comparison to the wild-type shows similar activities of Acs, Pcs and Icl on propionate medium (G10/P100 of Table 3) but reduced levels of propionate specific enzyme activities such as methylcitrate synthase and methylisocitrate lyase (a canditate gene is AN8755.2 from the conceptual translation of the A. nidulans genome, which shows 46% identity to the methylisocitrate lyase from Sc. cerevisiae). This implies that the activating effect on glyoxylate cycle enzymes mediated by propionate is FacB independent and furthermore, higher basal levels of FacB seem to have a negative effect on methylcitrate cycle enzymes.

Figure 3.

Spore colour formation of different A. nidulans strains. Growth conditions are given on the right (G, glucose; P, propionate, A, acetate; e.g. G50/P10 = the medium contained 50 mm glucose + 10 mm propionate). Strains are A26, wild-type; Fab4-J3, facB multi copy strain; ΔmcsA, methylcitrate synthase deletion strain; SMB/acuA, facA303 mutation in the acetyl-CoA synthetase.

The inability of the methylcitrate synthase mutant to remove propionyl-CoA via the methylcitrate pathway leads to loss of spore colour formation even at low propionate concentrations. As shown in lines VII and IX of Fig. 3, the addition of acetate to glucose/propionate medium releases suppression of spore colour formation especially in the methylcitrate synthase mutant and the wild-type. The facB multi-copy strain Fab4-J3, however, is inhibited even more. This strain shows strongly increased acetyl-CoA and propionyl-CoA synthetase activity on media containing both acetate and propionate (Table 3), which leads to the accumulation of propionyl-CoA. On a medium containing 50 mm acetate and 10 mm propionate, the levels were 18 nmol acetyl-CoA and 40 nmol propionyl-CoA·g dry weight−1 (ratio 1 : 2.2); when the medium contained 100 mm acetate and 100 mm propionate, the levels rose to 20 nmol acetyl-CoA and 66 nmol propionyl-CoA ·g dry weight−1 (ratio 1 : 3.3). Furthermore lines VII and VIII show that this strain behaved very similarly on media without glucose, which is in agreement with the observation that the strain hardly uses glucose and acetate in parallel (see section entitled Carbon balances on different growth media). Nevertheless, the spore colour of strain Fab4-J3 in lanes IX and X is hard to visualize, because the number of spores at these growth conditions is greatly reduced. This is also true for the ΔmcsA-strain on G50/P100, which implies that at high propionyl-CoA concentrations not only spore colour formation but also conidiation is affected.

Utilization of acetate by the acetyl-CoA synthetase mutant is strictly dependent on the activity of the predicted propionyl-CoA synthetase. Strain SMB/acuA shows better growth on media containing only 10 mm propionate and 50 mm acetate instead of equimolar concentrations of these. This can be explained by the necessity of the presence of 10 mm propionate to induce propionyl-CoA synthetase activity (Table 3), which is then able to activate acetate. At equimolar concentrations of acetate and propionate, activation of propionate by the propionyl-CoA synthetase is much more likely than that of acetate (see section entitled Formation of acetyl-CoA and propionyl-CoA; Vmax/Km (acetate) = 25 U·g−1·mm−1; Vmax/Km (propionate) = 344 U·g−1·mm−1). From these results we conclude that the acetyl-CoA/propionyl-CoA ratio and also the ability to activate propionate to propionyl-CoA has to be well balanced with the methylcitrate synthase activity for successful spore colour formation and growth.

The data in Table 3 further imply that propionyl-CoA might be a direct inducer of methylcitrate cycle genes. On media G50/P100 and G10/P100, which lead to a strong accumulation of propionyl-CoA in the ΔmcsA-strain, the activity of methylisocitrate lyase is twice as high than that of the wild-type. The addition of acetate to these media not only lowered the propionyl-CoA level, but also that of methylisocitrate lyase activity. Therefore, a putative transcriptional activator of the methylcitrate cycle genes seems to be activated by propionyl-CoA (or propionyl-carnitine) rather than by methylcitrate as suggested for the procaryotic regulator of the propionate utilization genes from S. typhimurium[41].

Discussion

Growth of A. nidulans on glucose medium is inhibited by propionate in a concentration-dependent manner. In a strain carrying a defective methylcitrate synthase gene, this effect is even much more pronounced. When acetate was the main carbon source, addition of propionate had no growth inhibitory effect on the wild-type and little effect on the methylcitrate synthase deletion strain. One might assume that the inhibition observed on glucose is caused by a reduced glucose uptake, due to the presence of carboxylic acids. We were able to show that acetate and propionate did not inhibit uptake of glucose by measuring the total carbon consumption and carbon balances from different carbon sources. Measurements clearly indicated that in the presence of glucose and propionate, despite the reduced growth rate, an elevated level of glucose was required for the formation of 1 g dried mycelium. Furthermore, on glucose/acetate/propionate, which should inhibit glucose uptake even more, the growth rate was increased and was actually higher than that observed with acetate as sole carbon source [2].

On the other hand, we were able to correlate the growth inhibitory effect of propionate on glucose medium with the intracellular concentration of propionyl-CoA. Since this CoA-derivative also accumulated on acetate/propionate medium without showing significant growth retardation, we concluded that propionyl-CoA inhibits enzymes mainly involved in glucose rather than in acetate metabolism. We found that activities of CoA-dependent enzymes such as ATP citrate lyase, succinyl-CoA synthetase and the pyruvate dehydrogenase complex were strongly inhibited in the presence of propionyl-CoA.

ATP citrate lyase from A. nidulans provides cytosolic acetyl-CoA required for the biosynthesis of fatty acids and polyketides. The enzyme level was shown to be regulated by the carbon source present in the media: high levels on glucose and low levels on acetate. Unfortunately, the effect of propionate on enzyme levels was not investigated and remains unclear [42]. Therefore, further studies will also have to focus on the activity pattern of this enzyme on propionate containing media. We cannot evaluate the direct effect of a partial inhibition of ATP citrate lyase by propionyl-CoA on the metabolism, because this enzyme is not involved in glucose degradation. It is also not clear whether inhibition of ATP citrate lyase indirectly diminishes polyketide synthesis or whether a direct interaction of propionyl-CoA with polyketide synthetase is responsible for this effect.

Succinyl-CoA synthetase is directly involved in the degradation of glucose, acetate and propionate via the Krebs cycle. Therefore an inhibition of this enzyme would block the oxidation of all three substrates, which was not observed with acetate. An elegant way to bypass the inhibition of this synthetase is the transfer of the CoA-moiety from succinyl-CoA to either acetate or propionate. We were able to show the existence of such a CoA-transferase, which indeed seems to be induced by propionate but prefers acetate to propionate as CoA acceptor (Table 4). Hence, the CoA-transferase explains the higher growth rate, which was always observed when acetate was added to a medium containing propionate. In the absence of acetate, however, the transferase enhances the formation of propionyl-CoA, which traps the system into a loop.

A very important inhibition is attributed to the pyruvate dehydrogenase complex. The low Ki of ≈ 50 µm propionyl-CoA (compare to 840 µm for pyruvate dehydrogenase of R. sphaeroides) not only clarified the growth inhibition of both organisms but also the observed excretion of pyruvate, which was dependent on the intracellular propionyl-CoA content. The excretion of pyruvate clearly demonstrates that the target of propionyl-CoA is pyruvate dehydrogenase rather than Krebs cycle enzymes. Since pyruvate dehydrogenase catalyses an irreversible reaction, the inhibition of any enzyme of the cycle cannot lead to an accumulation of pyruvate. The inhibition of pyruvate dehydrogenase also explains the low growth rate on propionate. We showed that in addition to a functional methylcitrate cycle pyruvate dehydrogenase is required for the pathway of propionate oxidation. Therefore activation of propionate and the subsequent oxidation of propionyl-CoA to acetyl-CoA has to be well balanced and does not allow high turnovers.

Despite the different metabolism of propionyl-CoA in fungi and humans (methylcitrate cycle vs. methylmalonyl-CoA pathway) we conclude from our results that accumulation of propionyl-CoA might show severe effects not only on fungal but also on human cells, which carry defective genes of the methylmalonyl-CoA pathway. Mutated genes encoding propionyl-CoA carboxylase and methylmalonyl-CoA mutase cause the diseases propionic acidemia and methylmalonic aciduria, respectively. Both are generally diagnosed by the determination of methylcitrate in the urine generated from accumulated propionyl-CoA, especially in liver hepatocytes [43,44]. Hence, phenotypes of the diseases (dehydration, lethargy, nausea and vomiting as well as a risk for neurologic sequelae) might be caused not only by metabolites derived from propionyl-CoA as are propionate, β-hydroxypropionate, β-hydroxybutyrate, methylmalonyl-CoA and methylcitrate, but also directly by propionyl-CoA inhibiting pyruvate dehydrogenase as described in this study.

Besides the impairment caused by propionyl-CoA we cannot exclude a depletion of free CoASH, which would also lead to a strong disturbance of the metabolism and a reduction of pyruvate oxidation. However, the fact that the ΔmcsA strain also accumulates significant amounts of propionyl-CoA on acetate/propionate medium without showing a significant reduction in biomass formation compared to acetate as sole carbon source [2] seems to exclude this effect.

In order to get further insights into the mechanism of growth inhibition mediated by propionate, future work will focus on the phenotypic characterization of other mutants carrying defective genes of the methylcitrate cycle. Analysis of the fatty acid composition from the ΔmcsA strain grown on different carbon sources might also give an insight into substrate specificity of acetyl-CoA carboxylase and fatty acid synthases, depending on the existence of branched and odd chain fatty acids. Furthermore, we are trying to identify and purify the transcriptional activator of the propionate utilization genes and analyse its DNA recognition sequence. Knowledge of this sequence will facilitate the screening of other promoters for putative regulation by propionate, which might be helpful in the understanding of metabolic networks.

In summary the data presented here demonstrate how metabolites are shuttled between different pathways in fungal cells. However, exact flow rates cannot be determined by these methods. Flux measurements by 13C-NMR-spectroscopy could be helpful but are certainly difficult to interpret due the simultaneous use of mixtures of two or three substrates. Analysis of different mutants will give supporting evidence, but definite conclusions cannot be drawn, because ‘every change of enzyme activity in a metabolic network is able to disturb the metabolism’[45].

Acknowledgements

This work was supported by grants of the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie. We thank Jennifer Beier (Universität Hannover, Germany) for her technical assistance during activity determination, Richard B. Todd (The University of Melbourne, Australia) for providing strain Fab4-J3 and Professor Nancy Keller (University of Wisconsin-Madison, USA) for providing strain RYQ11.

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