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J. Biol. Chem., Vol. 280, Issue 6, 4469-4475, February 11, 2005

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Kinetic Properties and Metabolic Contributions of Yeast Mitochondrial and Cytosolic NADP⁺-specific Isocitrate Dehydrogenases^*

Veronica Contreras-Shannon, An-Ping Lin, Mark T. McCammon, and Lee McAlister-Henn

From the Department of Biochemistry, University of Texas Health Science Center, San Antonio, Texas 78229-3900

Received for publication, September 3, 2004 , and in revised form, December 1, 2004.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
To compare kinetic properties of homologous isozymes of NADP⁺-specific isocitrate dehydrogenase, histidine-tagged forms of yeast mitochondrial (IDP1) and cytosolic (IDP2) enzymes were expressed and purified. The isozymes were found to share similar apparent affinities for cofactors. However, with respect to isocitrate, IDP1 had an apparent K_m value 7-fold lower than that of IDP2, whereas, with respect to -ketoglutarate, IDP2 had an apparent K_m value 10-fold lower than that of IDP1. Similar K_m values for substrates and cofactors in decarboxylation and carboxylation reactions were obtained for IDP2, suggesting a capacity for bidirectional catalysis in vivo. Concentrations of isocitrate and -ketoglutarate measured in extracts from the parental strain were found to be similar with growth on different carbon sources. For mutant strains lacking IDP1, IDP2, and/or the mitochondrial NAD⁺-specific isocitrate dehydrogenase (IDH), metabolite measurements indicated that major cellular flux is through the IDH-catalyzed reaction in glucose-grown cells and through the IDP2-catalyzed reaction in cells grown with a nonfermentable carbon source (glycerol and lactate). A substantial cellular pool of -ketoglutarate is attributed to IDH function during glucose growth, and to both IDP1 and IDH function during growth on glycerol/lactate. Complementation experiments using a strain lacking IDH demonstrated that overexpression of IDP1 partially compensated for the glutamate auxotrophy associated with loss of IDH. Collectively, these results suggest an ancillary role for IDP1 in cellular glutamate synthesis and a role for IDP2 in equilibrating and maintaining cellular levels of isocitrate and -ketoglutarate.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The existence of multiple and differentially compartmentalized isozymes of NADP⁺-specific isocitrate dehydrogenase (IDP)¹ appears to be a common attribute of eukaroytic organisms. Whereas Saccharomyces cerevisiae has three isozymes encoded by different genes (1–4), the cytosolic and peroxisomal isozymes of mammalian cells are encoded by a single gene distinct from that encoding the mitochondrial isozyme (5–7). In Arabidopsis, genes for cytoplasmic and peroxisomal isozymes are distinct, but a third gene apparently encodes both mitochondrial and chloroplast isozymes (8). This duplication of genes and/or differential localization of the same enzyme suggests that the reaction catalyzed by IDP isozymes is important for optimal metabolic function of various cellular compartments. The current study is directed toward refining our understanding of these metabolic functions.

The yeast mitochondrial IDP1, cytosolic IDP2, and peroxisomal IDP3 isozymes are homodimers, and they share pairwise primary sequence identities of >70% (1–4). They differ with respect to organellar targeting sequences; IDP1 has a 16-residue amino-terminal sequence that is removed upon mitochondrial import (1), and IDP3 has a carboxyl-terminal Cys-Lys-Leu tripeptide essential for peroxisomal localization (3, 4). IDP2 also has a significantly lower isoelectric point than the two organellar isozymes (Table I). In addition, the IDP enzymes differ with respect to expression. IDP1 is essentially constitutively expressed, whereas IDP2 is expressed during growth on nonfermentable carbon sources, e.g. glycerol, ethanol, and lactate (5, 9). IDP2 is not expressed in cells growing logarithmically with glucose as the carbon source, but expression increases dramatically during the diauxic shift after glucose is exhausted and ethanol serves as the carbon source (10). IDP3 is expressed only with fatty acid carbon sources, reflecting the specific function of this enzyme in providing NADPH for the reduction of fatty acids with even-numbered double bonds during peroxisomal -oxidation (3, 4).

The IDP isozymes are quite distinct in terms of sequence, structure, and regulation from the mitochondrial NAD⁺-specific tricarboxylic acid cycle isocitrate dehydrogenase (IDH). Yeast IDH is an allosterically regulated octamer composed of four IDH1 and four IDH2 subunits (17, 18). Although residues in catalytic sites are primarily contributed by IDH2 and those in regulatory sites are primarily contributed by IDH1 (19–22), loss of either subunit eliminates cellular IDH activity and produces similar growth phenotypes. These include an inability to grow with acetate as a carbon source (23, 24), a phenotype shared with other yeast tricarboxylic acid cycle mutants (25–27). Growth in glucose medium in the absence of glutamate is also reduced with loss of IDH, but growth is eliminated only with the additional loss of IDP1 (28, 29), suggesting that both enzymes contribute to production of -ketoglutarate under conditions when IDP2 is not expressed. When IDP2 is expressed, i.e. with nonfermentable carbon sources, loss of all three enzymes (IDP1, IDP2, and IDH) is required to produce glutamate auxotrophy.

The goal of this report is to define further the physiological roles of the IDP1 and IDP2 isozymes. Detailed kinetic analyses of the purified enzymes have revealed some differences in kinetic parameters that may contribute to differential function in vivo. We have also directly examined contributions of the IDP1, IDP2, and IDH enzymes to cellular pools of isocitrate and -ketoglutarate, and we have determined the extent that IDP1 and IDP2 isozymes can augment cellular functions normally attributed to IDH.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Yeast Strains and Growth Conditions—Parental yeast strain S173-6B (MATa leu2-3,112 his3-1 ura3-57 trp1-289; see Ref. 30) was used for isozyme expression and metabolite analyses. Growth phenotype analyses were conducted with this parental strain and with MMY011 (MAT ade2-1 can1-100 his3-11,15 leu2-3,112 trp1-1 Ole⁺; see Ref. 31). Isocitrate dehydrogenase gene disruption mutants constructed by using these strains were described previously (27, 28). Strains were cultivated on rich YP medium (1% yeast extract, 2% Bacto-peptone) or on minimal YNB medium (0.17% yeast nitrogen base, 0.5% ammonium sulfate) containing nutrients to supplement auxotrophic requirements of the strains. Nitrogen-free medium was YNB lacking ammonium sulfate. Carbon sources were 2% glucose, 2% glycerol plus 2% lactate, or 2% ethanol. Growth rates in liquid cultures were monitored as absorbance at 600_nm (A₆₀₀). For plate spot assays of growth phenotypes, cells were pelleted from cultures and resuspended in H₂O to 2.0 A₆₀₀. The cells were serially diluted in 10-fold steps, and 10 µl of each dilution was spotted onto agar plates.

Expression and Purification of Histidine-tagged Forms of IDP1 and IDP2—To add histidine codons onto the 3' end of the coding region of IDP1, plasmid pBS-IDP1 (1) was used as a template for mutagenesis using the unique site transformer mutagenesis method from Stratagene. One oligonucleotide (5'-GTTTTTCTTTAGTTCAGCTATGAGGTGGTAGTGGTAGTGATTATTAGCTTAAATGCATCGG) was used to introduce five adjacent histidine codons (underlined) prior to the stop codon in the IDP1 coding sequence and to remove a PvuI site in this region for screening mutant clones; a second oligonucleotide (5'-CTCACTAAGGGAACAAGAGCTCCATGCCTGCAGGTCG) was used to delete a vector HindIII site for selection of mutagenized plasmids. The altered IDP1 gene was transferred on a 3.2-kbp SacI DNA fragment to plasmid pRS424 (32), which contains a 2-µm origin for multicopy replication and a yeast TRP1 gene for selection of transformants. The resulting plasmid, pRS424-IDP1^His, was subjected to DNA sequence analysis to confirm that the only change to the IDP1 coding sequence was introduction of the 3' histidine codons.

Plasmid pRS424-IDP2 (33) was used as the template to add histidine codons onto IDP2. Codons for six adjacent histidine residues were added at the 3' end of the IDP2 coding region using PCR with a forward oligonucleotide primer (5'-TGCTAGATCGATTAATGACAAAGATTAAGGTAGCTAACC) containing a ClaI restriction site (italics) and a reverse oligonucleotide primer (5'-CTCAAGCTCCGTCGACGTAACGTGGTGGTGGTGGTGGTGATTGACGTCAGGCTA) containing the histidine codons (underlined) and a PstI restriction site (italics). The resulting ClaI/PstI PCR product was used to replace the authentic IDP2 coding region in pRS424-IDP2, generating pRS424-IDP2^His. The IDP2^His coding sequence was verified by DNA sequence analysis.

The pRS424-IDP1^His and pRS424-IDP2^His plasmids were transformed into an S173-6B mutant strain (idp1idp2) containing disruptions in both IDP1 and IDP2 loci (5). For purification of histidine-tagged enzymes, transformants were cultivated in YP medium with glycerol/lactate as the carbon source. Cells were harvested at culture densities of A₆₀₀ = 5.0–7.0. Harvested cells were lysed with glass beads in 2.5 ml/g cells of lysis buffer (50 mM potassium phosphate (pH 7.75), 5 mM MgCl₂, 2 mM potassium citrate, and 5% glycerol) containing 10 mM phenylmethylsulfonyl fluoride (PMSF) and 1% (v/v) protease inhibitor mixture (Sigma). The extracts were diluted with an equal volume of a buffer containing 100 mM sodium phosphate (pH 8.0), 600 mM NaCl, 40 mM imidazole, 2 mM potassium citrate, and 15% glycerol, combined with 1.0 ml of a 50% slurry of Ni²⁺-nitrilotriacetic acid superflow resin (Qiagen), and rocked at 4 °C for 60 min. The mixture was loaded onto a 10 x 1-cm column, and the flow-through fraction was collected. The column was washed four times with 5.0 ml of buffer A (50 mM sodium phosphate (pH 8.0), 300 mM NaCl, 5 mM MgCl₂, 2 mM potassium citrate, and 1 mM PMSF) initially containing 20 mM imidazole and 15% glycerol. The concentration of imidazole was increased to 30 mM in the second wash, and the concentration of glycerol was gradually increased to 25% in the final wash. Bound protein was eluted with 0.75–1.5 ml of buffer A containing 200 mM imidazole and 25% glycerol. Purified enzymes were stored in the same buffer at –20 °C. The high concentrations of glycerol were determined to be necessary for stability during purification and storage. Protein concentrations were determined using calculated extinction coefficients (34).

Kinetic Analyses—pH optima for IDP enzymes were examined with isocitrate saturation curves using 50 mM potassium phosphate or Tris-HCl buffers at pH values ranging from 6.0 to 9.5. Assays (1.0 ml) contained 5 mM MgCl₂, 0.5 mM NADP⁺, and DL-isocitrate concentrations ranging from 0 to 5.0 mM. For kinetic assays of the oxidative decarboxylation reaction, IDP activity was assayed in 1.0-ml reactions containing 50 mM potassium phosphate (pH 7.75), 5 mM MgCl₂, and 10% glycerol. Kinetic parameters with respect to isocitrate were determined using 0.5 mM NADP⁺ and DL-isocitrate concentrations ranging from 1.0 µM to 10.0 mM. Kinetic parameters with respect to NADP⁺ were obtained using 5.0 mM isocitrate and NADP⁺ concentrations ranging from 1.0 µM to 0.25 mM. For inhibition studies, -ketoglutarate at concentrations ranging from 0.5 to 8.0 mM or glutamate at concentrations ranging from 1.0 to 17.5 mM was added with isocitrate as the substrate. These concentrations were chosen to represent low to high ranges of typical physiological conditions based on published values (35, 36).

For assays of the reductive carboxylation reaction, IDP activity was measured in 1.0 ml of 50 mM potassium phosphate (pH 6.5), 5 mM MgCl₂, 40 mM NaHCO₃ (54.5 mM CO₂), and 10% glycerol. Kinetic parameters with respect to -ketoglutarate were determined by using 0.25 mM NADPH and -ketoglutarate concentrations ranging from 1.0 µM to 10 mM. Kinetic parameters with respect to NADPH were obtained using 2.5 mM -ketoglutarate and NADPH concentrations ranging from 1.0 µM to 0.25 mM.

For all assays, rates of NADPH or NADP⁺ production were measured spectrophotometrically at A₃₄₀. Units are expressed as µmol of NADP(H) formed per min/mg of protein. Kinetic data were analyzed using Excel (Microsoft) and Sigma Plot (SPSS Inc.). Dixon plots (1/v versus [i] at various substrate concentrations) were used to analyze inhibition and to determine K_i values.

Analysis of Enzymes in Cellular Extracts—Whole cell protein extracts were prepared by glass bead lysis of cells harvested from cultures at A₆₀₀ = 2.0 in lysis buffer containing 2 mM PMSF. IDP activity in lysates was assayed in 1.0-ml reactions containing 50 mM potassium phosphate (pH 7.75), 5 mM MgCl₂, and 5.0 mM DL-isocitrate. The reaction was initiated by adding NADP⁺ to a final concentration of 0.5 mM. Lysate protein concentrations were determined by using the Bradford dye-binding assay (37) with bovine serum albumin as the standard.

Metabolite Extraction and Analysis—Rapid centrifugation (2–3 min at 4000 x g) was used to harvest cells grown to A₆₀₀ = 2.0 in 500 ml of minimal medium with glucose or glycerol/lactate as the carbon source. Cell pellets were resuspended in 1.5 ml of water and boiled for 10 min. Standard solutions containing -ketoglutarate and isocitrate were similarly boiled to ascertain the stability of both metabolites. After cooling on ice, a 75-µl sample of each cellular extract was collected for protein determination by using a modified Bradford assay (38). The remaining volume was cleared by using microcentrifugation for 15 min at 4 °C, and the supernatant was removed for metabolite analyses. The concentration of -ketoglutarate was determined using enzyme assays with beef liver glutamate dehydrogenase (Sigma) (39). The concentration of isocitrate in protein-free supernatants was determined by using enzyme assays with porcine heart NADP⁺-specific isocitrate dehydrogenase (Sigma). Similar concentrations were measured using yeast IDP1 purified as described above.

Electrophoresis and Immunoblot Analysis—Denaturing gel electrophoresis was conducted as described by Laemmli and Favre (40) using SDS-10% polyacrylamide gels. Immunoblot analyses of the IDP isozymes were conducted using an antiserum (diluted 1:500) prepared against purified yeast IDP1 (9). IDH antiserum was used as described previously (18). Immunoreactive polypeptides were detected using the enhanced chemiluminescent method (Amersham Biosciences) and autoradiography. Densitometric analyses were performed using Image-Quant (Amersham Biosciences).

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Purification of IDP Isozymes—To conduct kinetic comparisons of yeast mitochondrial IDP1 and cytosolic IDP2 isozymes, we purified carboxyl-terminal histidine-tagged forms of both enzymes. The enzymes were expressed using genes with authentic promoters on multicopy vectors in a yeast strain (idp1idp2) containing disruptions of both chromosomal genes. Transformants were cultivated in medium with glycerol and lactate as carbon sources, a condition that produces optimal expression of both isozymes (5, 9).

Conditions for purification of histidine-tagged IDP1 and IDP2 using Ni²⁺-NTA chromatography were empirically optimized to stabilize the enzymes during purification and to reduce levels of contaminating proteins. Recoveries, based on specific activities in crude lysates and in final elution fractions, were 60% for IDP1 and 40% for IDP2, with yields of 0.3 mg of IDP1/liter of culture and 1.8 mg of IDP2/liter of culture. Although the histidine-tagged forms of the purified enzymes have similar molecular weights, IDP1 and IDP2 polypeptides are resolved by denaturing gel electrophoresis (Fig. 1A). This is also the case for authentic forms of the polypeptides in extracts from cells grown with glycerol/lactate as the carbon source (Fig. 1B, immunoblot). Both isozymes are detected in an extract from the parental strain (lane 1), whereas only IDP2 is detected in an extract from an idp1 strain (lane 2), and only IDP1 is detected in an extract from an idp2 strain (lane 3).

View larger version (46K): [in this window] [in a new window]   FIG. 1. Electrophoretic comparisons of IDP1 and IDP2. A, Sypro Ruby staining was used following electrophoretic resolution of 0.12 µg each of histidine-tagged IDP1 (lane 1) and IDP2 (lane 2). Sizes of protein standards (lane 3) are indicated. B, immunoblot analysis using an antiserum that recognizes both yeast IDP isozymes (1) was conducted with 25-µg samples of cellular lysates from the parental strain (lane 1), an idp1 strain (lane 2), and an idp2 strain grown on YP medium with glycerol/lactate as the carbon source.

Kinetic properties were also determined for the reverse reductive carboxylation reaction (Table II). For these assays, it was necessary to use 10-fold higher concentrations of enzymes to obtain measurable activities. Apparent V_max values for IDP1 and IDP2 were found to be similar and much lower (10–25-fold) than values determined for the forward reaction. For IDP2, apparent K_m values for -ketoglutarate and for NADPH were quite similar to the respective K_m values for isocitrate and for NADP⁺. For IDP1, the apparent K_m values for cofactor, NADPH or NADP⁺, were similar. However, K_m values of IDP1 for substrate were strikingly different; the apparent K_m value for -ketoglutarate was >60-fold the value for isocitrate. This suggests that IDP1 would be less likely to catalyze the reverse reaction. However, the direction of the reaction catalyzed by IDP2 in vivo may be determined by relative levels of isocitrate and -ketoglutarate.

The potential for inhibition of the oxidative decarboxylation reaction by -ketoglutarate was also investigated. For both IDP1 and IDP2, -ketoglutarate acted as a competitive inhibitor (data not shown). Also, cooperativity of IDP2 with respect to isocitrate was not observed in the presence of -ketoglutarate. Similar apparent K_i values for -ketoglutarate, 3.6 mM for IDP1 and 2.0 mM for IDP2, were obtained. As described below, we estimate a cellular concentration of 1.0 mM for -ketoglutarate under certain growth conditions, suggesting the possibility for inhibitory effects in vivo. In other experiments, we found no evidence for inhibition of the oxidative decarboxylation reaction for either isozyme by glutamate.

Contributions to Cellular Pools of Isocitrate and -Ketoglutarate—The different kinetic properties of IDP1 and IDP2 suggest they may have different physiological functions. To examine aspects of this possibility, we compared levels of isocitrate and -ketoglutarate in cellular extracts from yeast mutants lacking each or both of these enzymes. In addition, we included extracts from yeast mutants with a disruption in the IDH2 gene, which eliminates cellular IDH activity (23), to compare metabolite contributions by the mitochondrial NAD⁺-specific enzyme. Extracts for measurements of metabolites were prepared from parental and mutant strains grown in minimal medium with glucose as the carbon source, a condition associated with expression of IDP1 and of IDH at glucose-repressed levels but not of IDP2, or with glycerol/lactate as the carbon source, a condition associated with expression of all three enzymes (9).

For the parental strain, as shown in Table III, cellular levels of isocitrate slightly exceeded those of -ketoglutarate, 1.5-fold in glucose and 2.6-fold in glycerol/lactate medium. Levels of these metabolites were remarkably similar with the different carbon sources, suggesting that increased flux through oxidative pathways with the nonfermentable carbon source (41) does not produce an increase in steady state levels of tricarboxylic acid cycle intermediates relative to levels in glucose-grown cells.

View this table: [in this window] [in a new window]   TABLE III Cellular concentrations of isocitrate and -ketoglutarate Concentrations of isocitrate and -ketoglutarate were determined enzymatically as described under "Experimental Procedures" in protein-free extracts prepared from parental and mutant yeast strains grown in minimal medium with either glucose or glycerol plus lactate as the carbon source.

With glycerol/lactate as the carbon source (Table III), loss of IDP2 alone produced a 22-fold increase in isocitrate levels and a slight increase in -ketoglutarate levels, whereas the major effect of loss of IDP1 or of IDH was a 10-fold decrease in -ketoglutarate levels. This suggests a major flux of isocitrate through the IDP2-catalyzed reaction with growth on this nonfermentable carbon source but that flux through the IDP1 and IDH reactions is important for maintenance of a cellular pool of -ketoglutarate. Data for mutants containing two gene disruptions and thus only one functional isocitrate dehydrogenase support this conclusion: function of IDH alone or IDP1 alone (in the idp1idp2 and idp2idh2 mutants, respectively) was adequate to maintain a substantial pool of -ketoglutarate, whereas function of IDP2 alone (in the idp1idh2 mutant) was not.

These data suggest that excess isocitrate and -ketoglutarate, produced as tricarboxylic acid cycle intermediates during growth on a nonfermentable carbon source, may be transferred to the cytosol for metabolism involving IDP2. They also provide some evidence for reversible function of IDP2 in vivo. The relative levels of isocitrate:-ketoglutarate (600:1) in the idp1idh2 mutant expressing only IDP2 on glycerol/lactate medium were similar to those (700:1) for the same mutant grown on glucose medium when IDP2 is not expressed, suggesting that IDP2 may normally function to equilibrate levels of these metabolites.

Correlates with Expression and Compensation by IDP Isozymes for Phenotypes Associated with Loss of IDH—Our previous analyses of growth phenotypes of yeast isocitrate dehydrogenase mutants (28, 29) have shown that, despite differences in cellular levels of -ketoglutarate (Table III), expression of any one of the IDP1, IDP2, or IDH isozymes is sufficient to permit parental rates of growth on nonfermentable carbon sources in the absence of glutamate. However, in the absence of glutamate with glucose as the carbon source, the growth rate of a strain lacking IDH is retarded 2-fold, but growth is eliminated only for a strain lacking both IDH and IDP1. This suggests that IDH is the predominant enzyme but that IDP1 can contribute to -ketoglutarate/glutamate synthesis under fermentative conditions despite the absence of dramatic effects on metabolite levels in a idp1 strain under such conditions (Table III).

To further examine the role of IDP1 as a source of -ketoglutarate, we transformed a strain (idh1idh2) lacking IDH with multicopy plasmids (pRS424-IDP1 and pRS424-IDP2) carrying the IDP1 or IDP2 genes. As shown in Fig. 2A, the transformant harboring pRS424-IDP1 grew better on minimal glucose plates lacking glutamate than the transformants harboring pRS424-IDP2 or the empty pRS424 vector. However, this growth was significantly less than that for a transformant expressing IDH subunits using a single-copy plasmid (YCp-IDH1/IDH2). All transformants grew equally well in the presence of glutamate. These results suggest that IDP1 can contribute to -ketoglutarate pools for glutamate synthesis. However, IDH is the major source, and even elevated levels of IDP1 (10-fold based on enzyme assays and immunoblots of cellular lysates, data not shown) cannot completely compensate for loss of IDH under fermentative conditions.

View larger version (40K): [in this window] [in a new window]   FIG. 2. Compensation by overexpression of IDP isozymes for growth phenotypes of a strain lacking IDH. An idh1idh2 strain derived from strain MMY011 (27) was transformed with the empty multicopy pRS424 plasmid, with pRS424-IDP1, pRS424-IDP2, or with YCp-IDH1/IDH1, a single copy plasmid carrying genes for both IDH subunits (29). The transformants were precultured overnight in YNB glucose medium supplemented with 1% casamino acids, and dilutions were plated on YNB glucose plates lacking or containing 0.1% glutamate. Similar phenotypes were observed with transformants of a mutant strain derived from strain S173-6B (data not shown).

View larger version (52K): [in this window] [in a new window]   FIG. 3. Immunoblot analyses of isocitrate dehydrogenase isozyme expression. A, carbon source and glutamate effects. Extracts were prepared from the parental strain grown on minimal medium with glucose or ethanol as the carbon source and in the presence or absence of 0.1% glutamate as indicated. 5-µg samples were analyzed with antisera for the IDP isozymes (1) or for the two subunits of IDH (18). B, ammonium effects. Extracts were prepared from the parental (wild type (wt)), idp1, and idp2 strains grown on minimal glucose medium containing or lacking ammonium sulfate (+/– nitrogen) as indicated. For immunoblot analysis, 25-µg samples were analyzed with IDP and IDH antisera. Values (averages of five independent determinations) for IDP-specific activity (µmol of NADPH/min/mg of protein) measured in these samples are shown. Differences are consistent with relative levels of IDP1 determined using densitometry. C, ammonium effects on IDP1 levels. Extracts were prepared from the parental strain grown on minimal glucose medium lacking ammonium sulfate (0 min) and at the indicated times following a shift of the cells to minimal glucose medium containing 0.5% ammonium sulfate. 15-µg samples were loaded for immunoblot analysis using the IDP antiserum.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
In the current report, we have investigated potential functions of the yeast mitochondrial IDP1 and cytosolic IDP2 isozymes by comparing their kinetic properties and by assessing some of the physiological consequences of either loss or overexpression of these enzymes. Although loss of either or both IDP enzymes produces no dramatic growth phenotypes (2, 9), our data suggest that IDP1 performs an ancillary function with IDH to maintain a cellular pool of -ketoglutarate for glutamate synthesis, and that the major flux of isocitrate to -ketoglutarate may be catalyzed by IDP2 in cells grown on nonfermentable carbon sources

IDH appears to be the major source of -ketoglutarate for glutamate synthesis in cells grown with glucose as the carbon source (Fig. 4A). Loss of IDH under this condition results in both a reduction in -ketoglutarate levels and an increase in isocitrate levels (Table III), which correlates with a reduction in growth rates in the absence of glutamate (29). Loss of IDP1, the only other isocitrate dehydrogenase expressed during growth on glucose, has a lesser effect on metabolite levels, but IDP1 can contribute to glutamate synthesis because concomitant loss of IDH and IDP1 is required to eliminate growth in the absence of glutamate (28, 29). Also, overexpression of IDP1 partially compensates for the slow glutamate growth phenotype of a strain lacking IDH (Fig. 2A). In parental glucose-grown cells, IDH expression is elevated by the absence of glutamate (Fig. 3A) (9), whereas IDP1 expression is increased by the absence of nitrogen (Fig. 3B), suggesting that IDP1 may be particularly important when nitrogen is limiting.

View larger version (21K): [in this window] [in a new window]   FIG. 4. Flux through isocitrate dehydrogenase reactions in yeast cells grown on glucose (A) or on a nonfermentable carbon source (B). Bold arrows indicate major flux as determined by metabolite measurements (Table III). Dotted arrows in B indicate expected flux in cells grown with a fatty acid carbon source.

By using conversion factors reported by others (5 µl of intracellular volume/mg of protein; see Refs. 43 and 44) and our value for levels of isocitrate and -ketoglutarate in glycerol/lactate grown parental cells, the respective cellular concentrations of these metabolites can be estimated to be 2.6 and 1.0 mM. Thus, the cellular concentration of isocitrate appears to exceed the apparent K_m values for the IDP enzymes (Table II) and for IDH (45). However, the cellular concentration of ketoglutarate is similar to the apparent K_m value for IDP1 and exceeds by 5-fold that for IDP2. The cellular level of -ketoglutarate is also similar (2–3-fold lower) to the apparent K_i values determined for inhibition of the oxidative decarboxylation reaction catalyzed by both IDP1 and IDP2. Thus, catalysis by these enzymes may be responsive to levels of -ketoglutarate in vivo, and the relative compartmental concentration of this metabolite, whereas unknown, could be physiologically relevant.

In the models illustrated in Fig. 4, the mitochondrial conversion of isocitrate to -ketoglutarate is shown to be irreversible, as suggested by the kinetic parameters obtained for IDP1 and those described previously for IDH (46). This is also supported by several lines of evidence suggesting that a substantial pool of -ketoglutarate is maintained primarily by IDH during growth on glucose (Fig. 4A) and by both mitochondrial enzymes during growth on a nonfermentable carbon source (Fig. 4B). A capacity for bidirectional catalysis is proposed for IDP2 (Fig. 4B) based on the similarity in apparent K_m values for isocitrate or -ketoglutarate and for NADP⁺ or NADPH. Finally, because the only known function for peroxisomal IDP3 is provision of NADPH for -oxidation, it is assumed that isocitrate would be diverted into peroxisomes (dotted lines in Fig. 4B), only when cells are grown with fatty acid carbon sources, and that catalysis by IDP3 is unidirectional. This model suggests that bidirectional catalysis by IDP2 may function to support metabolite shuttle cycles among the different compartments. With respect to a possible mitochondrial shuttle cycle, inner membrane transporters for tricarboxylic acids and for -ketoglutarate have been described for yeast (47, 48). The necessity for transport of these metabolites is implied by cytosolic localization of glutamate dehydrogenase and glutamine synthetase (49). With respect to a possible peroxisomal shuttle, it is known that peroxisomal membranes are impermeable to small molecules including NAD(H) (50), but metabolite carriers (e.g. for isocitrate or -ketoglutarate) analogous to those in mitochondria have not yet been described (13). Isocitrate is also a substrate for the glyoxylate cycle, which is partially compartmentalized in peroxisomes in yeast (31).

Whereas mammalian cells contain a mitochondrial transhydrogenase capable of interconversion of NADPH and NADH (51), the absence of a transhydrogenase in yeast mitochondria may increase reliance on shuttle cycles involving the isocitrate dehydrogenases. The current studies support a role for IDP2 in such shuttles. As described previously (11), IDP2 also functions with glucose-6-phosphate dehydrogenase to provide NADPH for thiol-dependent antioxidant enzymes. In a related study (33), we found that IDP2, when localized in mitochondria in lieu of IDP1, can support glutamate synthesis, but that IDP1, when localized in the cytosol in lieu of IDP2, does not fully restore antioxidant functions. Thus, some of the physical and kinetic differences between these isozymes may have evolved for specific compartmental functions and interactions.

	FOOTNOTES

 
^* This work was supported by National Institutes of Health Grants AG17477 and GM51265. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Present address: Dept. of Microbiology and Immunology, South Texas Centers for Biology in Medicine, University of Texas Health Science Center, San Antonio, TX 78245.

To whom correspondence should be addressed. Tel.: 210-567-3782; Fax: 210-567-6595; E-mail: henn{at}uthscsa.edu.

¹ The abbreviations used are: IDP, NADP⁺-specific isocitrate dehydrogenase; IDP1, mitochondrial NADP⁺-specific isocitrate dehydrogenase; IDP2, cytosolic NADP⁺-specific isocitrate dehydrogenase; IDP3, peroxisomal NADP⁺-specific isocitrate dehydrogenase; IDH, mitochondrial NAD⁺-specific isocitrate dehydrogenase; PMSF, phenylmethylsulfonyl fluoride.

² The K_m values for isocitrate of IDP1 and IDP2 cannot be directly compared due to the cooperativity displayed by IDP2.

	ACKNOWLEDGMENTS

 
We thank Dr. Karyl I. Minard for construction of the plasmid for expression of histidine-tagged IDP1 and for critical reading of the manuscript.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 

1. Haselbeck, R. J., and McAlister-Henn, L. (1991) J. Biol. Chem. 266, 2339–2345[Abstract/Free Full Text]
2. Loftus, T. M., Hall, L. V., Anderson, S. L., and McAlister-Henn, L. (1994) Biochemistry 33, 9661–9667[CrossRef][Medline] [Order article via Infotrieve]
3. Henke, B., Girzalsky, V., Berteaux-Lecelier, V., and Erdmann, R. (1998) J. Biol. Chem. 273, 3702–3711[Abstract/Free Full Text]
4. Van Roermund, C. W. T., Hettema, E. H., Kal, A. J., van den Berg, M., Tabak, H. F., and Wanders, R. J. A. (1998) EMBO J. 17, 677–687[CrossRef][Medline] [Order article via Infotrieve]
5. Minard, K. I., Jennings, G. T., Loftus, T. M., Xuan, D., and McAlister-Henn, L. (1998) J. Biol. Chem. 273, 31486–31493[Abstract/Free Full Text]
6. Geisbrecht, B. V., and Gould, S. J. (1999) J. Biol. Chem. 274, 30527–30533[Abstract/Free Full Text]
7. Haselbeck, R. J., Colman, R. F., and McAlister-Henn, L. (1992) Biochemistry 31, 6219–6223[CrossRef][Medline] [Order article via Infotrieve]
8. Hodges, M., Flesch, V., Gálvez, S., and Bismuth, E. (2003) Plant Physiol. Biochem. 41, 577–585[CrossRef]
9. Haselbeck, R. J., and McAlister-Henn, L. (1993) J. Biol. Chem. 263, 12116–12122
10. DeRisi, J. L., Iyer, V. R., and Brown, P. O. (1997) Science 273, 680–686
11. Minard, K. I., and McAlister-Henn, L. (1999) J. Biol. Chem. 274, 3402–3406[Abstract/Free Full Text]
12. Kunau, W.-H., Dommes, V., and Schulz, H. (1995) Prog. Lipid Res. 34, 267–342[CrossRef][Medline] [Order article via Infotrieve]
13. Hiltunen, J. K., Mursula, A. M., Rottensteiner, H., Wierenga, R. K., Kastaniotis, A. J., and Gurvitz, A. (2003) FEMS Microbiol. Rev. 27, 35–64[CrossRef][Medline] [Order article via Infotrieve]
14. Jo, S. H., Son, M. K., Koh, H. J., Lee, S. M., Song, I. H., Kim, Y. O., Lee, Y. S., Jeong, K. S. Kim, W. B., Park, J. W., Song, B. J., Huh, T. L., and Huhe, T. L. (2001) J. Biol. Chem. 276, 16168–16176[Abstract/Free Full Text]
15. Lee, S. M., Koh, H. J., Park, D. C., Song, B. J., Huh, T. L., and Park, J. W. (2002) Free Radic. Biol. Med. 32, 1185–1196[CrossRef][Medline] [Order article via Infotrieve]
16. Koh, H. J., Lee, S. M., Sohn, B. G., Lee, S. H., Ryoo, Z. Y., Chang, K. T., Park, J. W., Park, D. C., Song, B. J., Veech, R. L., Song, H., and Huh, T. L. (2004) J. Biol. Chem. 279, 39968–39974[Abstract/Free Full Text]
17. Barnes, L. D., Kuehn, G. D., and Atkinson, D. E. (1971) Biochemistry 10, 3939–3944[CrossRef][Medline] [Order article via Infotrieve]
18. Keys, D. A., and McAlister-Henn, L. (1990) J. Bacteriol. 172, 4280–4287[Abstract/Free Full Text]
19. Cupp, J. R., and McAlister-Henn, L. (1993) Biochemistry 32, 9323–9328[CrossRef][Medline] [Order article via Infotrieve]
20. Zhao, W.-N., and McAlister-Henn, L. (1997) J. Biol. Chem. 272, 21811–21817[Abstract/Free Full Text]
21. Lin, A.-P., and McAlister-Henn, L. (2002) J. Biol. Chem. 277, 22475–22483[Abstract/Free Full Text]
22. Lin, A.-P., and McAlister-Henn, L. (2003) J. Biol. Chem. 278, 12864–12872[Abstract/Free Full Text]
23. Cupp, J. R., and McAlister-Henn, L. (1991) J. Biol. Chem. 266, 22199–22205[Abstract/Free Full Text]
24. Cupp, J. R., and McAlister-Henn, L. (1992) J. Biol. Chem. 267, 16417–16423[Abstract/Free Full Text]
25. Kim, K., Rosenkrantz, M. S., and Guarente, L. (1986) Mol. Cell. Biol. 6, 1936–1942[Abstract/Free Full Text]
26. McAlister-Henn, L., and Thompson, L. M. (1987) J. Bacteriol. 169, 5157–5166[Abstract/Free Full Text]
27. Przybyla-Zawislak, B., Gadde, D. M., Ducharme, K., and McCammon, M. T. (1999) Genetics 152, 153–166[Abstract/Free Full Text]
28. Zhao, W.-N., and McAlister-Henn, L. (1996) Biochemistry 35, 7873–7878[CrossRef][Medline] [Order article via Infotrieve]
29. McCammon, M. T., and McAlister-Henn, L. (2003) Arch. Biochem. Biophys. 419, 222–233[CrossRef][Medline] [Order article via Infotrieve]
30. Botstein, D., Falco, S. C., Stewart, S. E., Brennan, M., Scherer, S., Stinchcomb, D. T., Struhl, K., and Davis, R. W. (1979) Gene (Amst.) 9, 17–24
31. McCammon, M. T., Veenhuis, M., Trapp, S. B., and Goodman, J. M. (1990) J. Bacteriol. 172, 5816–5827[Abstract/Free Full Text]
32. Sikorski, R. S., and Hieter, P. (1989) Genetics 122, 19–27[Abstract/Free Full Text]
33. Contreras-Shannon, V., and McAlister-Henn, L. (2004) Arch. Biochem. Biophys. 423, 235–246[Medline] [Order article via Infotrieve]
34. Pace, C. N., Vajdos, F., Fee, L., Grimsley, G., and Gray, T. (1995) Protein Sci. 4, 2411–2423[Medline] [Order article via Infotrieve]
35. Jones, E. W., and Fink, G. R. (1982) in The Molecular Biology of the Yeast Saccharomyces, Metabolism and Gene Expression (Strathern, J. N., Jones, E. W., and Broach, J. R., eds) pp. 181–299, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
36. DeLuna, A., Avendaño, A., Riego, L., and González, A. (2001) J. Biol. Chem. 276, 43775–43783[Abstract/Free Full Text]
37. Bradford, M. (1976) Anal. Biochem. 72, 248–254[CrossRef][Medline] [Order article via Infotrieve]
38. Gotham, S. M., Fryer, P. J., and Paterson, W. R. (1988) Anal. Biochem. 173, 353–358[CrossRef][Medline] [Order article via Infotrieve]
39. Bergmeyer, H., and Bernt, E. (1983) in Methods of Enzymatic Analysis (Bergmeyer, H., ed) Vol. I, pp. 1577–1580, Academic Press, New York
40. Laemmli, U. K., and Favre, M. (1973) J. Mol. Biol. 80, 575–599[CrossRef][Medline] [Order article via Infotrieve]
41. Sols, A., Gancedo, C., and De la Fuente, G. (1971) in The Yeasts (Rose, A. H., and Harrison, J. S., eds) Vol. 2, pp. 271–308, Academic Press, London
42. Liu, Z., Sekito, T., Epstein, C. B., and Butow, R. A. (2001) EMBO J. 20, 7209–7219[CrossRef][Medline] [Order article via Infotrieve]
43. De Koning, W., and van Dam, K. (1992) Anal. Biochem. 204, 118–123[CrossRef][Medline] [Order article via Infotrieve]
44. Uemura, H., and Fraenkel, D. G. (1999) J. Bacteriol. 181, 4719–4723[Abstract/Free Full Text]
45. Lin, A.-P., McCammon, M., and McAlister-Henn, L. (2001) Biochemistry 40, 14291–14301[CrossRef][Medline] [Order article via Infotrieve]
46. Barnes, L. D., McGuire, J. J., and Atkinson, D. E. (1972) Biochemistry 11, 4322–4328[CrossRef][Medline] [Order article via Infotrieve]
47. Palmieri, L., Runswick, M. J., Fiermonte, G., Walker, J. E., and Palmieri, F. (2000) J. Bioenerg. Biomembr. 32, 67–77[CrossRef][Medline] [Order article via Infotrieve]
48. Palmieri, L., Agrimi, G., Runswick, M. J., Fearnley, I. M., Palmieri, F., and Walker, J. E. (2001) J. Biol. Chem. 276, 1916–1922[Abstract/Free Full Text]
49. Ter Schure, E. G., van Riel, N. A. W., and Verrips, C. T. (2000) FEMS Microbiol. Rev. 24, 67–83[Medline] [Order article via Infotrieve]
50. Van Roermund, C. W., Elgersma, Y., Singh, N., Wanders, R. J., and Tabak, H. R. (1995) EMBO J. 14, 3480–3486[Medline] [Order article via Infotrieve]
51. Rydstrom, J., Hock, J. B., and Ernster, L. (1976) in The Enzymes (Boyer, P. D., ed) Vol. 13, pp. 1–88, Academic Press, New York[Medline] [Order article via Infotrieve]

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