Molecular Cloning of the Yeast Mitochondrial Aconitase ... - Europe PMC

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Vol. 10, No. 7

MOLECULAR AND CELLULAR BIOLOGY, JUlY 1990, p. 3551-3561 0270-7306/90/073551-11$02.00/0 Copyright C) 1990, American Society for Microbiology

Molecular Cloning of the Yeast Mitochondrial Aconitase Gene (ACOI) and Evidence of a Synergistic Regulation of Expression by Glucose plus Glutamate SERGE P. GANGLOFF, DIDIER MARGUET, AND GUY J.-M. LAUQUIN*

Institut de Biochimie Cellulaire et Neurochimie, Centre National de la Recherche Scientifique, 1, rue Camille Saint Saens, 33077 Bordeaux Cedex, France Received 27 November 1989/Accepted 3 April 1990

We have isolated genomic clones complementing the aconitase-deficient strain (glul-1) of Saccharomyces cerevisiae. Identification of the aconitase gene was established by enzymatic assays and molecular analyses. The corresponding mRNA has been characterized, and its direction of transcription has been determined. The complete nucleotide sequence revealed strong amino acid homologies with the sequences of some peptides isolated from the mammalian protein. Disruption of the gene by deletion-insertion led to glutamate auxotrophy. Expression of the aconitase gene was sensitive to glucose repression and was synergistically down regulated by glucose and glutamate.

Aconitase (citrate [isocitrate] hydro-lyase; EC 4.2.1.3), an enzyme of the Krebs cycle (18) located mainly in the mitochondrial matrix, catalyzes the reversible isomerization of the tricarboxylic acids (TCA) citrate and isocitrate. cisAconitate is formed as an intermediary product which normally does not dissociate from the enzyme during the course of the reaction (32, 45). Like the other TCA cycle enzymes, aconitase from Saccharomyces cerevisiae is encoded in the nucleus and transported into the mitochondria (39). This enzyme is composed of a single polypeptide chain of Mr about 80,000 (32, 42, 45). Aconitase has been purified from different organisms, and homologies in amino acid content of mammalian, Candida lipolytica, and S. cerevisiae species suggest a common evolutionary origin (42). Of considerable interest is the characterization of the enzyme as an Fe-S protein (36). As the enzymatic reaction involves only dehydration and hydration steps, the presence of an Fe-S cluster, normally implicated in electron transfer, was unexpected. This cluster appears to be a 4Fe-4S center capable of reversible rearrangement to yield a 3Fe-4S center; the labile iron seems to be involved in the binding of the enzyme substrate (2, 6, 15). Aconitase provides the best-characterized example of a nonheme iron enzyme that does not function as an agent of electron transport. In yeast cells, aconitase also takes part in the functioning of the glyoxylate cycle, and extramitochondrial aconitase activity has been found (5). This dual cell localization has also been described for citrate synthase activity, for which two functional nuclear genes have been characterized (17, 29). In S. cerevisiae, the first enzymes of the TCA cycle, i.e., citrate synthase and aconitase, are required both for synthesis of glutamate and for efficient utilization of nonfermentable energy sources (e.g., glycerol, lactate, or acetate) (17, 24). An aconitase-deficient strain (glul-J) has been isolated (24); its main phenotypic features are glutamate auxotrophy, citrate accumulation when glucose is present as a carbon source, and inability to grow on acetate, lactate, ethanol, and glycerol. Studies (28, 38, 47) of the effects of glucose repression on the activities of the TCA cycle enzymes have shown that aconitase activity is subject to catabolite regula*

tion. Recent work with Bacillus subtilis has demonstrated that the levels of aconitase are further reduced when a source of glutamate is supplied with a rapidly metabolized carbon source (34). By contrast, glutamate alone, or in combination with a poor carbon source (e.g., citrate), does not lead to reduction of the level of this enzyme (12). We report here the cloning of the S. cerevisiae gene coding for the mitochondrial aconitase by complementation of the glul-J mutation. We characterize its transcript and show that gene disruption leads to glutamate auxotrophy and the petite phenotype. Preliminary studies of ACOI gene expression are also presented, and subcellular localization of the gene product is discussed. MATERIALS AND METHODS Growth and transformation of S. cerevisiae. All experiments described were performed with S. cerevisiae GRF18 (MATa leu2-3,112 his3-11,15 canl-100), DBY747 (MATa leu2-3112 his3-AJ trpl-289 ura3-52) or its derivatives, and the aconitase-deficient strain GL153 (MATat leu2-3,112 his3Al ura3-52 glul-1), obtained by sporulation of the cross between MO-48-C (glul-1) and DBY747. SG3-2, SG4-6, SG6-1, and SG7-1 are the glul-J strain GL153 transformed with the complementing plasmids of the same name. YPD and SD media were prepared as described previously (41). YPR and SR were the same as YPD and SD except that 2% glucose was replaced by 2% raffinose; similarly, SDG and SRG were the same as SD and SR but with the addition of 1% glutamate. All synthetic media were supplemented with the required auxotrophic markers at a final concentration of 20 mg/liter for histidine and uracil, 30 mg/liter for leucine and tryptophan, and 100 mg/liter for glutamate. Yeast transformation was carried out either by the spheroplast method described by Beggs (1) or by the lithium method reported by Ito et al. (14). Bacterial strains and cloning vectors. Subcloning was performed with the Bluescript vector (Stratagene) and the shuttle vectors YEp24 (3) and pFL44 (a gift from F. Lacroute). The pFL44 vector was derived by insertion into the pUC19 Alul sites 629 and 747, respectively, of the URA3 HindIII fragment with BglII linkers and of the 2,um replication origin MstI fragment with ClaI linkers. The host Esch-

Corresponding author. 3551

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erichia coli strain was TG-1 [A(lac-pro) supE thi hsdD5 (F' traD6 proA+B+ laclIq lacZ AM15)]. Competent cells were prepared according to Maniatis et al. (20). Transformants were grown on LB plates supplemented with 100 pug of ampicillin per ml. Construction of plasmid pSE-3 AACO and gene replacement. The 0.65-kilobase-pair (kbp) SalI-EcoRI fragment from pAT153 was replaced with the 3.3-kbp SalI-EcoRI aconitase-bearing fragment from pSG7-1. The inner 0.35-kbp BglII fragment was then removed, and the 1.2-kbp URA3 marker was inserted. The 4.2-kbp newly generated SalIlEcoRI fragment was then introduced into pFL44, in which the BglII sites resulting from the URA3 deletion had been filled by reaction with Klenow enzyme and religated to form plasmid pSE-3 AACO. Gene disruption was performed according to the one-step method described by Rothstein (35). DNA sequence analysis. Sequencing of both strands of the EcoRI-SalI aconitase-bearing fragment subcloned in both orientations in the Bluescript vector was carried out as described by Sanger et al. (37). Sequences of the ACOJ gene were serially deleted from upstream or downstream first with exonuclease III and then with exonuclease VII (49). The ends were made blunt with T4 DNA polymerase, and the vectors were religated. The single-stranded DNAs, overlapping on at least 100 bp, were isolated by published methods (22), and primer extension was performed with the M13 universal 17-mer primer. Approximately 450 bp was read from each deleted fragment. Enzyme assays. Aconitase activity was assayed by an adaptation of the method described by Fansler and Lowenstein (7). Crude extracts were prepared from 100-ml cultures harvested in the early log phase. Cells were suspended in potassium phosphate buffer (20 mM, pH 7.4) supplemented with 1 mM phenylmethylsulfonyl fluoride and broken with glass beads by vortexing. After 5 min of centrifugation at 3,000 x g, the assays were performed on the supernatant fractions. Specific activities are given as nanomoles of cis-aconitate transformed per minute per milligram of protein. Protein concentration was determined by the biuret method (10). This regimen produced activity measurements that were reproducible within 15%. Assays were performed in duplicate, and results were averaged. Southern blots, Northern (RNA) blots, probes, and in vitro transcription experiments. S. cerevisiae genomic DNA was prepared by the method of Sherman et al. (41), digested with appropriate restriction enzymes, subjected to electrophoresis through 1% agarose gels, and blotted onto nitrocellulose as described by Southern (44). Yeast total RNA was extracted from early-log-phase cells (41), subjected to electrophoresis through formaldehyde-agarose gels, and blotted onto nitrocellulose (44). Blots were hybridized with purified nick-translated (30) or oligonucleotide-labeled (8) probes. In vitro transcription reactions were carried out as directed by the supplier (Stratagene).

RESULTS Isolation of the gene for mitochondrial aconitase. We separately transformed the aconitase-deficient strain GL153 with the three different pools of DNA from the YEp24 genomic libraries constructed by Carlson and Botstein (3). The uracil-independent transformants were plated onto glutamate-lacking medium, and 11 independent clones were selected. The DNA from the transformed colonies was prepared, and four different plasmid species were isolated (pSG3-2, -4-6, -6-1, and -7-1; Fig. 1A) which were character-

MOL. CELL. BIOL.

ized by a common restriction pattern within a 7.5-kbp region. To confirm that the complementation of the aconitase mutation (i.e., ability to grow without an exogenous supply of glutamate) was directly related to the presence of these plasmids, we reintroduced them into the glul-J mutant strain. The wild-type phenotype (i.e., glutamate independence) was restored in all cases and was accompanied by loss of the petite phenotype. We also assayed the enzymatic activities of crude extracts of the different transformants, together with activities of the parental strain controls, which had been grown on SD medium supplemented with 0.01% glutamate for strains SG3-2, SG4-6, SG6-1, SG7-1, and DBY747, enzyme activities were, respectively, 160, 160, 180, 160, and 70 nmol of cis-aconitate transformed per min per mg of protein; no activity was detectable for strain GL153. The transformed cells yielded aconitase activities up to 2.5-fold that obtained with wild-type strain DBY747, suggesting the presence of the aconitase structural gene on a multicopy plasmid. The mitochondrial localization of this activity has been investigated by cell fractionation experiments. In the mitochondrial fraction, we recovered 70% of the total activity and obtained with the transformed mutant at three- to fourfold-higher specific activity than with the wild-type strain. In addition, total DNA from wild-type strain DBY747 was digested with 10 different restriction enzymes, blotted onto nitrocellulose, and probed with the 2.6-kbp PvuII-EcoRI fragment (Fig. 1B). The results indicate that there are no BamHI, ClaI, EcoRI, or PvuII restriction sites within this region and that KpnI, PstI, and XbaI are present once and BglII and EcoRV are present twice in this fragment. The deduced genomic pattern is identical to the one determined by restriction analysis of the cloned DNAs and indicates that no recombination had occurred during the cloning steps. Messenger identification and subcloning experiments. To identify more precisely the region responsible for the recovery of aconitase activity, we restricted the yeast insert DNA from plasmid pSG3-2 with BamHI and KpnI into three fragments (1.5, 3.5, and 5.0 kbp) (Fig. 2A). After nick translation, these fragments were used as probes in Northern hybridization experiments with wild-type RNAs. Probe I (1.5 kbp) showed no autoradiographic signal, whereas probe II (3.5 kbp) identified a 2.6-kb messenger corresponding to the predicted size of the transcript for the 80-kilodalton aconitase protein (42). Probe III (5.0 kbp) detected the 2.6-kb messenger plus an unidentified 1.6-kb transcript (Fig. 2A). On the basis of these results, three different DNA fragments from plasmid pSG7-1, PstI-PstI (5.0 kbp), SaIlBamHI (5.6 kbp), and SalI-EcoRI (3.3 kbp), were subcloned into the corresponding polylinker restriction sites of the shuttle vector pFL44 to form plasmids pPP-1, pSB-2, and pSE-3, respectively. After transformation of the glul-1 mutant strain, the colonies were tested for glutamate prototrophy (Fig. 1B). The minimal region necessary for complementation of the aconitase glul-J mutation was located within the 3.3-kbp SalI-EcoRI fragment of plasmid pSG7-1. Since the Sal site of this subclone was derived from the YEp24 vector, the minimal yeast DNA necessary for complementing this aconitase mutation lies within the 3.0-kbp region proximal to the SalI restriction site of plasmid pSG7-1. This is in agreement with the 2.6-kb size of the mRNA detected by Northern blot hybridization experiments. Moreover, the PstI-PstI construction indicates that the region from the PstI site to the Sall site is necessary for the recovery of aconitase activity.

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Determination of transcription direction of the ACOI mesTo determine the orientation of the putative aconitase transcript, in vitro transcription experiments were performed. The 3.3-kbp SalI-EcoRI fragment from pSG7-1 was subcloned into the corresponding sites of the Bluescript+KS polylinker, whereas the 2.6-kbp PvuII-EcoRI was inserted into the Smal-EcoRI sites, thus yielding the reverse orientation. Transcription was performed from the T7 promoter, and the labeled RNA probes were hybridized with wild-type RNAs (Fig. 2B). The 2.6-kb autoradiographic signal was obtained with the latter construction only, indicating that transcription proceeds from the EcoRI to the PvuII site (Fig. 2B). Similar experiments carried out with single-stranded senger.

probes (13) derived from the 5.0-kbp BamHI-KpnI fragment cloned in M13mpl8 and -mpl9 vectors suggest that the 1.6-kb unidentified transcript is transcribed in the same direction as the ACOJ gene (data not shown). Disruption of the ACOI gene and genetic linkage analysis. If the ACOJ gene is essential for cell viability in the absence of exogenous glutamate, its disruption in haploid cells should lead to lethality in growth medium lacking glutamate. We inserted the 1.2-kbp Bglll URA3 selectable marker from the pFL44 vector, in which the inner PstI site had been removed by mutagenesis (F. Lacroute, personal communication), into the 0.35-kbp BglII-deleted aconitase gene borne on pSE-3

plasmid to form pSE-3 AACO (see Materials and Methods). The resulting 3.4-kbp PvuII-EcoRI linear fragment in which URA3 is inserted in the opposite transcriptional direction was used to transform strain DBY747 to uracil prototrophy (Fig. 3a). Transformed cells were then tested on synthetic medium lacking glutamate. All of the uracil prototrophs failed to grow when plated on glutamate-lacking medium, and no aconitase activity was detectable in crude extracts made from cultures of these colonies, indicating that the construction had inactivated the ACOI gene. In addition, all of the transformants were found unable to grow on glycerol medium and exhibited a petite phenotype when tested with the triphenyltetrazolium chloride technique (43). To confirm that the transformants carried the disrupted acol:: URA3 allele, we performed Southern analyses of genomic DNA digested with EcoRI and KpnI enzymes from untransformed and transformed cells (Fig. 3b). The 1.2-kbp intragenic EcoRV fragment spanning the BglII deletion was used as a probe. As expected, since there is one KpnI restriction site and no EcoRI site within the probe, the deletion-insertion event increased the size of the EcoRI and KpnI fragments spanning the BglII sites by about 0.8 kbp, leaving unchanged the size of the second KpnI fragment detected by the probe. To exclude the possibility that ACOJ is an extragenic suppressor of the glul-l aconitase mutation, we crossed the

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glul-J mutant strain GL153 (MATot leu2-3,112 his3-AJ ura352 glul-J) with the newly generated strain AACO1 (MATa ura3-52 leu2-3,112 his3-AJ trpl-289 acol::URA3). The resulting diploid cells (Ura+ Trp+) did not grow on synthetic medium lacking glutamate. Because of poor germination capacity of the haploid spores, random spore analysis was carried out. We analyzed 75 Trp+ colonies obtained after ether treatment of the sporulated diploids and found that all of them were Glu-, 38 were Ura+, and 37 were Ura-. We also verified that the analyzed clones were haploid by crossing them with tester strains. Thus, we can infer from that result that spores probably segregated 0+ :4- for glutamate auxotrophy and 2+:2- for uracil auxotrophy, indicating that the cloned ACOJ DNA is genetically linked to the GLUI DNA. To confirm this linkage, we integrated the ACOJ DNA at the glul-J locus of strain GL153 by transformation with the ClaI-linearized plasmid pBS-ESU (data not shown) and observed that a single copy of this DNA is capable of reversing the mutant phenotype. Plasmid pBSESU is a Bluescript vector in which the URA3 BglII fragment from pFL44 and the EcoRI-SalI ACOJ-bearing fragment have been inserted into the BamHI and EcoRI-SalI

polylinker sites, respectively. We crossed the resultant integrated GL153 strain (MATot leu2-3,112 his3-AJ ura3-52 gluJ-J: :pBS-ESU) with the glul-J mutant strain GL152 (MATa leu2-3,112 his3-AJ ura3-52 glul-J trpl-289) and selected the diploid cells on the basis of size. Random spore analysis was performed on 100 haploid spores, and 52 Glu+ and 53 Trp- spores were scored; among the 52 Glu+ spores, 27 were Trp+. This result is indicative of a 2+:2- segregation pattern for glutamate and tryptophan auxotrophies, confirming that we did not clone an extragenic suppressor of the glul-J mutation. These results indicate that the cloned ACOJ DNA is the GLUJ DNA. Sequence analysis of the ACOJ DNA. The EcoRI-SalI aconitase-bearing fragment on the Bluescript vector was entirely sequenced in both orientations as described in Materials and Methods; one single large open reading frame (ORF) was found on this DNA fragment. The nucleotide and derived polypeptide sequences are shown in Fig. 4. The ORF extends for 2,337 bp and can be translated into 779 amino acid residues (Mr 85,685). The first 24 residues of the protein encoded by the ORF include an abundance of hydrophobic residues, five basic and five hydroxylated (Ser

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FIG. 3. Disruption of wild-type aconitase gene by deletion-insertion. (a) Restriction maps of the aconitase gene in the disrupted and wild-type strains. The middle drawing represents the plasmid bearing the deleted-inserted aconitase gene whose PvuII-EcoRI linear fragment was used to transform the wild-type strain (see text). Open bars represent the aconitase gene; the hatched box represents the deleted portion; the direction of transcription of the inserted URA3 gene is shown by a black arrow within the box. The probe is the EcoRV fragment represented above the map of the wild-type strain. Enzyme restriction sites are as for Fig. 1. (b) Southern hybridization of chromosomal DNA from deletion mutant and wild-type strains. Mutant (lanes 1) and wild-type (lanes 2) DNAs were restricted by EcoRI (E) or KpnI (K). The probe is the labeled EcoRV fragment (see above). The indicated scale was generated by restricting lambda DNA with Hindlll and probing with 32P-labeled lambda DNA (lane m).

and Thr) amino acids, and a lack of acidic residues, a common feature of mitochondrial presequences (46). Therefore, this amino-terminal region is likely responsible for the mitochondrial targeting of the ACOJ product. The codon usage in ACOJ was compiled, and the codon bias index according to Sharp and Li (40) was calculated to be 0.43, which indicates a moderately expressed gene. We analyzed the 5' noncoding region and found a putative TATA box (TATATAAT; positions -141 to -134; underlined in Fig. 4). Interestingly, we also found at positions -291 to -284 a sequence corresponding to the inverted upstream activation sequence UAS2 UP1 (9), which has been shown to be a HAP2-HAP3-responsive site of the CYCI gene (25). Comparison of the deduced amino acid sequence with cysteinyl-tryptic peptides from beef heart aconitase. Former studies focused on isolation of the reactive cysteine of beef heart aconitase (26) have led to the determination of the amino acid sequence of seven cysteine-containing oligopeptides; no similarities between these sequences and that reported for the phenacyl bromide-reactive cysteine-containing peptide isolated from pig heart aconitase (11) have been reported. We compared our deduced amino acid sequence and those determined earlier and also found no apparent similarity with the pig heart aconitase peptide sequence, whereas all seven oligopeptides from beef heart aconitase were found to be present in the mitochondrial aconitase of S. cerevisiae, with similarities ranging from about 50 to nearly 100% (summarized in Fig. 5). These amino acid sequence comparisons together with the results presented above provide concrete evidence that the cloned ACOJ DNA corresponds to the mitochondrial aconitase structural gene. Regulation of aconitase expression. Recent studies on B. subtilis (34) have raised the problem of the regulation of aconitase expression. In this organism, aconitase is down regulated at the transcriptional level by glucose alone (catabolite regulation) or synergistically by glucose and glutamate.

To investigate whether the same situation occurs in S. cerevisiae, cultures were grown on different media. Cultures grown to early log phase were divided into two parts, one to be assayed for aconitase activity and the other to be assayed for extraction of total RNA, which was hybridized with the 1.25-kbp EcoRV intragenic probe. Preliminary experiments performed on cells grown on raffinose, a fermentable carbon source that does not cause catabolite repression to nearly the same extent as glucose (50), displayed degrees of aconitase activity and transcript similar to those obtained with a nonfermentable carbon source (e.g., glycerol), indicating that raffinose does not result in the repression of aconitase (data not shown). On the contrary, the results (Table 1) indicated that aconitase activity was down regulated about threefold by the presence of glucose as the sole carbon source in either rich or synthetic medium, as compared with the activity obtained with raffinose as the sole carbon source (catabolite regulation). Most of the mitochondrial enzymes are subject to a similar regulation (28, 38, 47). When glutamate was added to the culture (SDG and SRG media), the activity was further reduced twofold in the presence of glucose but not significantly with raffinose. These data suggest that glutamate severely decreases aconitase activity only when sufficient amounts of glucose are present in the medium. To define more accurately the level at which this regulation occurs, we analyzed the relative amounts of aconitase RNA present in the cells grown on the various media. Both catabolite regulation and synergistic repression by glucose and glutamate occurred principally at the mRNA accumulation level (Fig. 6). Indeed, the relative abundance of the transcripts as characterized by Northern blot experiments was closely related to the enzyme activities found for the same cultures. DISCUSSION

In this report, we describe the isolation and identification of a nuclear gene encoding the S. cerevisiae mitochondrial aconitase. Several key enzymes of intermediary metabolism

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432 144

GAA GTT TAT GAT TTC TTG GCC TCT GCC ACT GCG AAA TAT AAC ATG GGT TTC TGG Glu Val Tyr Asp Phe Leu Ala Ser Ala Thr Ala Lys Tyr Asn Met Gly Phe Trp

486 162

AAG CCA GGT TCC GGT ATC ATT QC CAA ATT GTT CTG GAA AAC TAC GCT TTC CCA Lys Pro Gly Ser Gly Ile Ile His Gln Ile Val Leu Glu Asn Tyr Ala Phe Pro

540

GGT GCT TTG ATC ATT GGT ACT GAC TCC OAT AOA CCA AAT GCT GGT GGT TTA GGT Gly Ala Leu Ile Ile Gly Thr Asp Ser His Thr Pro Asn Ala Gly Gly Leu Gly

594 198

CAA TTG GCT ATT GOT GTT GGT GGT GCT GAT GOC GTT GAT GTT ATG GCA GGT CGT Gln Leu Ala Ile Gly Val Gly Gly Ala Asp Ala Val Asp Val Met Ala Gly Arg

648 216

CCA TGG GAA TTG AAG GCT CCA AAG ATC TTA GGT GTT AAG TTG ACT GOT AAG ATG Pro Trp Glu Leu Lys Ala Pro Lys Ile Leu Gly Val Lys Leu Thr Gly Lys Met

702 234

AAC GGT TGG ACT TCT CCA AAG GAT ATT ATT TTG AAA TTG GCT GGT ATC AOA ACT Asn Gly Trp Thr Ser Pro Lys Asp Ile Ile Leu Lys Leu Ala Gly Ile Thr Thr

756 252

GTC AAA GGT GGT ACT GGT AAA ATT GTT GAA TAT TTT GGT GAT GOT GTT GAC ACT Val Lys Gly Gly Thr Gly Lys Ile Val Glu Tyr Phe Gly Asp Gly Val Asp Thr

810 270

TTC TOC GCT ACT GOT ATG GGT ACC ATT TGT AAT ATG GGT GOT GAA ATC GOT GCT Phe Ser Ala Thr Gly Met Gly Thr Ile Cys Asn Met Gly Ala Glu Ile Gly Ala

288

ACC AOA TCT GTT TTC CCA TTC AAC AAA TCT ATG ATT GAA TAT TTG GAA GCA ACT Ser Val Phe Pro Phe Asn Lys Ser Met Ile Glu Tyr Leu Glu Ala Thr

918 306

GGT CGT GGT AAG ATC GCT GAC ITT GCT AAA TTA TAC OC AAG GAT CTA TTA TCT Gly Arg Gly Lys Asp Phe Ala Lys Leu Tyr His Lys Asp Leu Leu Ser

972 324

GCT GAT AAG GAT GCT GAA TAC GAT GAG GTC GTC GAA ATT GAC TTG AAC ACT CTG Ala Asp Lys Asp Ala Glu Tyr Asp Glu Val Val Glu Ile Asp Leu Asn Thr Leu

1026 342

[EIThr

IlelAla

180

864

FIG. 4. Nucleotide and deduced amino acid sequences of the ACOI gene with its 5'- and 3'-flanking regions (GenBank accession number M33131). The noncoding strand and translation of the largest ORF are presented. The nucleotide sequence is numbered from nucleotide 1 of the presumed initiation codon. A potentially active TATA box and the inverted upstream activation sequence UAS2 UP1 are underlined (-145 to -134 and -291 to -284, respectively). The boxed amino acid sequences represent the oligopeptides corresponding to those sequenced by Plank and Howard (26). Oligopeptide numbering given in the text is depicted in Fig. 5. 3556

GAA CCA TAC ATC AT GGG CCA m ACC CCC GAT TTG GCT ACT CCA GTT TCT AAG Glu Pro Tyr Ile Asn Gly Pro Phe Thr Pro Asp Leu Ala Thr Pro Val Ser

Lysi

1080 360

ATG AAG GAA GTT GCT GTT GCT AAT AAC TGG CCA TTG GAT GTC AGA GTC GGT TTG 1134 Met Lys Glu Val Ala Val|Ala Asn Asn Trp Pro Leu Asp Val Arg|Val Gly Leu | 378 ATC GGT TCT TGT ACC AAT TCC TCT TAT GAA GAT ATG TCT CGT TCA GCA TCC ATT Ile Gly Ser Cys Thr Asn Ser Ser Tyr Glu Asp Met Ser Arg Ser Ala Ser Ile

1188 396

GTC AAG GAC GCT GCT GCT CAT GGT TTG AAA TCC AAG ACC ATr TTC ACT GTT ACT Val Lys Asp Ala Ala Ala His Gly Leu Lys Ser Lys Thr Ile Phe Thr Val Thr

1242 414

CCA GGT TCT GAA CAA ATC AGA GCC ACT ATT GAA CGT GAT GGC CAA TTA GAA ACC Pro Gly Ser Glu Gln Ile Arg Ala Thr Ile Glu Arg Asp Gly Gln Leu Glu Thr

1296 432

TTC AAA GAA TTT GGT GGT ATC GTT TTG GCA AAC GCC TGT GGC CCA TGT ATT GGT Phe Lys Glu Phe Gly Gly Ile Val Leu Ala Asn Ala Cys Gly Pro Cys Ile Gly

1350 450

CAA TGG GAT CGT AGA GAT ATC AAG AAA GGT GAC AAG AAT ACT ATT GTT TCC TCT Gln Trp Asp Arg Arg Asp Ile Lys Lys Gly Asp Lys Asn Thr Ile Val Ser Ser

1404 468

TAC AAC AGA AAT TTC ACT TCT AGA AAT GAT GGT AAC CCA CAA ACT CAT GCT TmT Tyr Asn Arg Asn Phe Thr Ser Arg Asn Asp Gly Asn Pro Gln Thr His Ala Phe

1458 486

GTT GCA TCT CCA GAA TTA GTA ACT GCG TTC GCC ATT GCG GGT GAT TTG AGA TTC Val Ala Ser Pro Glu Leu Val Thr Ala Phe Ala Ile Ala Gly Asp Leu Arg Phe

1512 504

AAC CCT CTA ACA GAC AAA TTA AAG GAC AAG GAT GGT AAT GAG TTC ATG TTG AAA Asn Pro Leu Thr Asp Lys Leu Lys Asp Lys Asp Gly Asn Glu Phe Met Leu Lys

1566 522

CCA CCA CAT GGT CGA TGG TmT GCC TCG AAA GAG GTT ATG ATG CTG GTG AGA ACA Pro Pro His Gly Arg Trp Phe Ala Ser Lys Glu Val Met Met Leu Val Arg Thr

1620 540

CTT ACC AAG CTC CAC CTG CAG ACC GTA GCC ACC GTT GAA GTT AAA GTT TCT CCA Leu Thr Lys Leu His Leu Gln Thr Val Ala Thr Val Glu Val Lys Val Ser Pro

1674 558

ACT TCA GAC CGT CTA CAA CTG TTG AAA CCA TTC AAA CCT TGG GAT GGT AAG GAT Thr Ser Asp Arg Leu Gln Leu Leu Lys Pro Phe Lys Pro Trp Asp Gly Lys Asp

1728

GCT AAA GAC ATG CCA ATC TTG ATT AAG GCC GTC GGT AAG ACA ACT ACT GAT CAT Ala Lys Asp Met Pro Ile Leu Ile Lys Ala Val Gly Lys Thr Thr Thr Asp His

1782 594

ATT TCT ATG GCT GGT CCA TGG TTG AAA TAC AGA GGT CAT TTA GAA AAC ATT TCT Ile Ser Met Ala Gly Pro Trp Leu Lys|Tyr Arg Gly His Leu Glu Asn Ile Ser

1836 612

AAT AAC TAT ATG ATT GGT GCT ATT AAT GCT GAA AAC AAG AAG GCT AAC TGT GTT Asn Asn Tyr Met Ile Gly Ala Ile Mn Ala Glu Asn Lys Lys Ala Asn Cys Val

1890 630

AAA AAT GTA TAT ACT GGT GAA TAC AAA GGT GTT CCA GAC ACT GCT AGA GAT TAC Lys Asn Val Tyr Thr Gly Glu Tyr Lys Gly Val Pro Asp Thr Ala Arg Asp Tyr

1944 648

AGA GAC CAA GGT ATC AAG TGG GIT GTT ATT GGT GAT GAA AAC TTT GGT GAA GGT Arg Asp Gln Gly Ile Lys Trp Val Val Ile Gly Asp Glu Asn Phe Gly Glu Gly

1998 666

TCC TCT CGT GAA CAC GCT GCT TTG GAA CCA AGA TTC TTG GGC GGT TTC GCT ATC Ser Ser Arg Glu His Ala Ala Leu Glu Pro Arg Phe Leu Gly Gly Phe Ala Ile

2052 684

ATC ACA AAG TCT TTC GCT CGT ATC CAT GAA ACT AAC TTG AAA AAA CAA GGT CTA Ile Thr Lys Ser Phe Ala Arg Ile His Glu Thr AMn Leu Lys Lys Gln Gly Leu

2106 702

TTG CCA TTG AAC TTC AAG AAC CCA GCT GAC TAT GAC AAG ATC AAC CCT GAT GAC Leu Pro Leu Asn Phe Lys AMn Pro Ala Asp Tyr Asp Lys Ile Asn Pro Asp Asp

2160 720

576

AGA ATC GAT ATT CTG GGT CTA GCT GAA TTG GCT CCA GGT AAG CCT GTA ACA ATG 2214 Arg Ile Asp Ile Leu Gly Leu Ala Giu Leu Ala Pro Gly Lys Pro Val Thr Met 738 AGA GTT CAT CCA AAG AAT GGT AAG CCA TGG GAT GCT GTG TTG ACC CAT ACT TTC Arg Val His Pro Lys Asn Gly Lys Pro Trp Asp Ala Val Leu Thr His Thr Phe

2268 756

AAC GAT GAG CAA ATT GAA TGG TTC AAA TAT GGT TCT GCC TTA AAT AAA ATT AAG Phe Lys Tyr Gly Ser Ala Leu Asn Lys Ile Lys

2322 774

Asn Asp Glu Gln Ile Glu T

GCC GAT GAG AAG AAA taatgaaaacattgttataatcttttaaaggttattatttattttgtcttc 2388 Ala Asp Glu Lys Lys 779

tgtacacgtacccttgtttatcttttctgccttaaatttaatgacgttcggctggagaagtcaagactatga 2460

aatatatctogtaatttatgatc

2483

3557

3558

GANGLOFF ET AL.

MOL. CELL. BIOL.

114 VAKPVTVHCDHLIQAQVGGEK **

*

*

******

**

****

peptide 5

VAVPSTIHCDHLIEAQLGGEK

255 GGTGKIVEYFGDGVDTFSATGMGTICNMGAEIGAT ****

****

*

*

***

***

************

GGTGAIVEYHGPGVDSISCTGMATICNMGAEIGAT

peptide 9

312 ADFAKL..YHKDLLSADKDAEYDEVVEIDLNTLEPYINGPFTPDLATPVSKMKEVAV **

*

**

*

*

*

**

**

*

*

*

**********

**

**

ADIANLADEFKDXLVPDSGCHYDQLIEINLSELYPHINGpFTpDLAAHpv. .AEVGS

peptide 10

376 VGLIGSCTNSSYEDMSR ***************

*

VGLIGSCTNSSYEDMGR

peptide 3

435 EFGGIVLANACGPCIGQWDR **************

***

DVGGIVLANACGPCIGEWDR

peptide 7

589 TTTDHISMAGPWLK ******

******

CTTDHISAAGPWLK

PePtide 4

729 ELAPGKPVTMRVHPKNGKPWDAVLTHTFNDEQIEW *****

*

**

*

**

*

***

DFAPGKPLTCIIKHPNGTQETILLNHTPNETXIEW

peptide 8 FIG. 5. Comparison of the ACOJ deduced amino acid sequence with sequences of cysteinyl-tryptic peptides from beef heart aconitase. The different amino acid sequences from S. cerevisiae are presented in the standard one-letter code and are aligned with the corresponding oligopeptides from beef heart aconitase by the Smith and Waterman program (43a). Breaks in the sequences are shown as periods and have been introduced to maximize identities. Homologies between the sequences are indicated by stars. The number above each pair of compared sequences indicates the position of the first amino acid in the S. cerevisiae peptide sequence. are known to have both cytosolic and mitochondrial isoforms. This is the case for citrate synthase (17, 29, 33), malate dehydrogenase (21), fumarase (48), and aconitase (5), which are present principally in the mitochondria and to a lesser extent in the cytoplasm, where they are implicated in the glyoxylate cycle. The ACOJ gene has been isolated from a genomic library in both orientations on YEp24 plasmids. The presence of the gene is indicated by the following results: (i) all of the cloned genomic DNAs share a common restriction pattern within 7.5 kbp and reverse the glutamate auxotrophy mutant phenotype; (ii) aconitase enzyme activities assayed on crude extracts of transformed cells were four times higher than those obtained on wild-type cells; (iii) and the nucleotide sequence revealed amino acid homologies of up to 100% with the seven cysteinyl-tryptic peptides from beef heart aconitase characterized by Plank and Howard (26). As previously noted for the beef enzyme (26), there is no clustering of cysteines. There are only 7 cysteine residues in the yeast enzyme but 11 in the beef heart aconitase. More specifically, according to the nomenclature of Howard and

Plank (26) for the beef enzyme, the cysteines equivalent to residues 199, 250, 305, 383, 565, and 714, found in peptides 1, 9, 10, 2, 4, and 6/8, respectively, do not exist within the yeast sequence. Conversely, cysteines 94 and 629 of the yeast aconitase are not present in the beef enzyme. Recent results by Robbins and Stout (31) indicated that the 4Fe cluster of beef heart aconitase has three thiols ligands, namely, cysteines 358, 421, and 424. Thus, possibly the equivalent cysteines in the yeast enzyme, i.e., 382, 445, and 448, respectively, are also the putative thiols ligands for the 4Fe-4S cluster. In contrast, Plank et al. (27) have recently suggested the possibility of a four-thiol ligand system; they propose that the cysteine 383 in peptide 2 could be the fourth ligand. As there is no equivalent cysteine in the yeast aconitase, the yeast 4Fe-4S cluster cannot involve a fourth thiol ligand, and the same might be true for the beef aconitase. These authors have also suggested that the linear 3Fe-4S cluster of purple aconitase uses both cysteines in peptide 7 and both cysteines in peptide 9 as ligands. The yeast enzyme cannot provide two cysteines within the equivalent peptide 9, and thus it would be interesting to

YEAST ACONITASE GENE

VOL. 10, 1990

TABLE 1. Aconitase activity determined on crude extracts of different yeast strains grown on various culture media

cn0

3559

e

0

Enzyme activity' (nmol of cis-aconitate

Medium

SD SDG SR SRG YPD YPR

transformed/min per mg of protein) DBY747

GRF18

68 25 180 180 140 290

55 20 160 160 140 270

' Average values determined on at least three independent experiments. Activity in strain GL153 and the AACO mutant strain was not detectable.

know whether yeast aconitase also exists as a stable purple aconitase. Our comparisons are similar to those drawn by Plank and Howard (26) regarding the similarities to other Fe-S proteins (thioredoxins or ferredoxins). Contrary to their findings, however, there is no cysteine residue in our peptide sequence corresponding to their oligopeptide 4, which they had implicated as containing the single cysteine that is found at or near the active site and which can be modified by a variety of sulfhydryl reagents. The absence of this cysteine residue in S. cerevisiae would seem to support the conclusions of Kennedy et al. (16), who suggested that this cysteine residue may interfere in substrate binding to the active site rather than being directly involved in the catalytic process. The ACOJ gene encodes the mitochondrial form in cells grown on a nonfermentable carbon source, and its disruption leads to both glutamate auxotrophy and petite phenotype. Therefore, if a second gene encoding cytosolic aconitase activity exists, it cannot compensate for the mitochondrial isoform defect. Furthermore, no aconitase activity is detectable in crude extracts of disrupted cells, and Southern analysis did not reveal any homologous sequences at high stringency of hybridization within the yeast genome; thus, any second gene would have diverged quite early from its mitochondrial counterpart. Taken together, these observations suggest the presence of a single-copy gene encoding both cytoplasmic and mitochondrial aconitase. Such a mechanism has been reported recently for the TCA enzyme fumarase (48). Differential sorting of products encoded by a single gene have already been described for invertase (3), some tRNA synthetases (4, 23), and more recently fumarase (48). All of these enzymes are synthesized as signal sequence-bearing precursors whether they are targeted to the mitochondria or secreted into the periplasmic space. It has been shown that this regulation is related to the presence of two transcription start sites and two initiation codons, one upstream and one downstream of the signal sequence. In the case of the mitochondrial aconitase of S. cerevisiae, no second translation start is found in the vicinity of the first ATG; the closest one would be Met-101, and therefore another kind of mechanism accounting for the dual cell localization must be considered. The observation that the strain carrying the disrupted acol allele displays no aconitase activity might be explained by the existence of a single-copy gene whose product is not localized solely within the mitochondria because of a low efficiency of the targeting signal. Thus, the aconitase enzyme fraction that does not enter the mitochondria could operate in the glyoxylate cycle. Another explanation for the results presented above may be the presence of

ACO _ ACT _

-

W

FIG. 6. Regulation of expression of the aconitase gene detected by Northern blot hybridization. Wild-type strain DBY747 was grown on SD (lane SD), SR (lane SR), SDG (lane SDG), and SRG (lane SRG). RNAs isolated from these cultures (8 ,ug) were subjected to electrophoresis, blotted onto nitrocellulose, and hybridized with a 32P-labeled EcoRV aconitase fragment. Actin DNA was used to probe the various amounts of RNA present in each lane. ACO, Aconitase transcript; ACT, actin messenger.

a related gene, the product of which is at a concentration too low to be detected by our biochemical assay. This product would not contain the necessary information to be targeted to the mitochondria and thus would be implicated only in the glyoxylate cycle. Further work is needed to understand the mechanism of aconitase partition between intra- and extramitochondrial compartments. Amino acid sequence analysis of the N-terminal part of both forms of aconitase would be of considerable help in this direction. In this study, we also explored the regulation of aconitase expression. This enzyme is required for growth on nonfermentable carbon sources and for biosynthesis of glutamate. It has been reported for the bacterium B. subtilis that levels of aconitase are regulated by the combination of a rapidly metabolized carbon source (e.g., glucose) and a source of glutamate (34); a similar control pathway operates for the citrate synthase of S. cerevisiae (17). We have shown by studying the mRNA steady-state levels and enzymatic activities of cells grown in various conditions that the pattern of regulation of aconitase in S. cerevisiae is superimposable on that observed for B. subtilis aconitase and S. cerevisiae citrate synthase; moreover, it would appear in these three cases that primary control is exercised at the level of transcription. Furthermore, recent results have shown that expression of the aconitase gene is under the control of both HAP2 and HAP3 genes (unpublished data). More accurate information about specific regulation will be provided by the study of mutant strains affected in the expression of both aconitase and citrate synthase. ACKNOWLEDGMENTS S. P. Gangloff is "allocataire" of the French Ministere de la Recherche et de l'Enseignement Superieur. This work was supported in part by research grants from the University of Bordeaux II, from la Fondation pour la Recherche Medicale, and from the Centre National de la Recherche Scientifique.

3560

GANGLOFF ET AL.

ADDENDUM IN PROOF

In our discussion on subcellular localization of aconitase we have overlooked previous work on shared gene products concerning the TRMI and MOD5 genes (J.-M. Li, A. K. Hopper, and N. C. Martin, J. Cell Biol. 109:1411-1419, 1989; D. Najarian, M. E. Dihanich, N. C. Martin, and A. K. Hopper, Mol. Cell. Biol. 7:185-191, 1987). A porcine aconitase cDNA was recently cloned and sequenced (L. Zheng, P. C. Andrews, M. A. Hermodson, J. E. Dixon, and H. Zalkin, J. Biol. Chem. 265:2814-2821, 1990). Comparison of the deduced amino acid sequence with that of yeast indicated 70% similarity between both sequences. LITERATURE CITED 1. Beggs, J. D. 1978. Transformation of yeast by a replicating hybrid plasmid. Nature (London) 275:104-109. 2. Beinert, H. 1986. Iron-sulfur clusters: agents of electron transfer and storage, and direct participants in enzymic reactions. Biochem. Soc. Trans. 14:527-533. 3. Carlson, M., and D. Botstein. 1982. Two differentially regulated mRNAs with different 5' ends encode secreted and intracellular forms of yeast invertase. Cell 28:145-154. 4. Chatton, B., P. Walter, J. P. Ebel, F. Lacroute, and F. Fasiolo. 1988. The yeast VAS1 gene encodes both mitochondrial and cytoplasmic valyl-tRNA synthetases. J. Biol. Chem. 263:52-57. 5. Duntze, W., D. Neumann, J. M. Gancedo, W. Atzpodien, and H. Holzer. 1969. Studies on the regulation and localization of the glyoxylate cycle enzymes in Saccharomyces cerevisiae. Eur. J. Biochem. 10:83-89. 6. Emptage, M. H., T. A. Kent, M. C. Kennedy, H. Beinert, and E. Munck. 1983. Mossbauer and EPR studies of activated aconitase: development of a localized valence state at a subsite of the (4Fe-4S) cluster on binding of citrate. Proc. Natl. Acad. Sci. USA 80:4674-4678. 7. Fansler, B., and J. M. Lowenstein. 1969. Aconitase from pig heart. Methods Enzymol. 13:26-31. 8. Feinberg, A. P., and B. Vogelstein. 1984. A technique for radiolabeling DNA restriction endonuclease fragments to high specific activity. Anal. Biochem. 137:266-267. 9. Forsburg, S. L., and L. Guarente. 1988. Mutational analysis of upstream activation sequence 2 of the CYCI gene of Saccharomyces cerevisiae: a HAP2-HAP3-responsive site. Mol. Cell. Biol. 8:647-654. 10. Gornall, A. G., C. J. Bardawill, and M. M. David. 1949. Determination of serum proteins by means of the biuret reaction. J. Biol. Chem. 177:751-766. 11. Hahm, K.-S., 0. Gawron, and D. Piszkiewicz. 1981. Amino acid sequence of a peptide containing an essential cysteine residue of pig heart aconitase. Biochim. Biophys. Acta 667:457-461. 12. Hanson, R. S., and D. P. Cox. 1967. Effect of different nutritional conditions on the synthesis of tricarboxylic acid cycle enzymes. J. Bacteriol. 93:1777-1787. 13. Hu, N.-T., and J. Messing. 1982. The making of strand-specific M13 probes. Gene 17:271-277. 14. Ito, H., Y. Fukuda, K. Murata, and A. Kimura. 1983. Transformation of intact yeast cells treated with alkali cations. J. Bacteriol. 153:163-168. 15. Kennedy, C., R. Rauner, and 0. Gawron. 1972. On pig heart aconitase. Biochem. Biophys. Res. Commun. 47:740-745. 16. Kennedy, M., G. Spoto, M. Emptage, and H. Beinert. 1988. The active site sulfhydryl of aconitase is not required for catalytic activity. J. Biol. Chem. 263:8190-8193. 17. Kim, K. S., M. S. Rosenkrantz, and L. Guarente. 1986. Saccharomyces cerevisiae contains two functional citrate synthase genes. Mol. Cell. Biol. 6:1936-1942. 18. Krebs, H. A., and J. M. Lowenstein. 1960. The tricarboxylic acid cycle, p. 129-303. In D. M. Greenberg (ed.), Metabolic pathways. Academic Press, Inc., New York. 19. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature (London) 227:680-685.

MOL. CELL. BIOL.

20. Maniatis, T., E. F. Fritsch, and J. Sambrook. 1982. Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 21. McAlister-Henn, L., and L. M. Thompson. 1987. Isolation and expression of the gene encoding yeast mitochondrial malate dehydrogenase. J. Bacteriol. 169:5157-5166. 22. Messing, J. 1983. New M13 vectors for cloning. Methods Enzymol. 101:20-78. 23. Natsoulis, G., F. Hilger, and G. R. Fink. 1986. The HTSl gene encodes both the cytoplasmic and mitochondrial histidinetRNA synthetases of S. cerevisiae. Cell 46:235-243. 24. Ogur, M., L. Coker, and S. Ogur. 1964. Glutamate auxotrophs in Saccharomyces. I. The biochemical lesion in the glt-l mutants. Biochem. Biophys. Res. Commun. 14:193-197. 25. Olesen, J., S. Hahn, and L. Guarente. 1987. Yeast HAP2 and HAP3 activators both bind to the CYCI upstream activation site, UAS2, in an interdependent manner. Cell 51:953-961. 26. Plank, D. W., and J. B. Howard. 1988. Identification of the reactive sulfhydryl and sequences of cysteinyl-tryptic peptides from beef heart aconitase. J. Biol. Chem. 263:8184-8189. 27. Plank, D. W., M. C. Kennedy, H. Beinert, and J. B. Howard. 1989. Cysteine labeling studies of beef heart aconitase containing a 4 Fe, a cubane, or a linear 3 Fe cluster. J. Biol. Chem. 264:20385-20393. 28. Polakis, E. S., and W. Bartley. 1965. Changes in the enzyme activities of Saccharomyces cerevisiae during aerobic growth on different carbon sources. Biochem. J. 97:284-297. 29. Rickey, T. M., and A. S. Lewin. 1986. Extramitochondrial citrate synthase activity in baker's yeast. Mol. Cell. Biol. 6:488-493. 30. Rigby, P. W. J., M. Dieckmann, C. Rgodes, and P. Berg. 1977. Labeling deoxyribonucleic acid to high specific activity in vitro by nick translation with DNA polymerase I. J. Mol. Biol. 113:237-251. 31. Robbins, A. H., and C. D. Stout. 1989. Structure of activated aconitase: formation of the [4Fe-4S] cluster in the crystal. Proc. Natl. Acad. Sci. USA 86:3639-3643. 32. Rose, I. A., and E. L. O'Connel. 1967. Mechanism of aconitase action. J. Biol. Chem. 242:1870-1879. 33. Rosenkrantz, M. S., T. Alam, K.-S. Kim, B. J. Clark, P. A. Srere, and L. P. Guarente. 1986. Mitochondrial and nonmitochondrial citrate synthase in Saccharomyces cerevisiae are encoded by distinct homologous genes. Mol. Cell. Biol. 6: 4509-4515. 34. Rosenkrantz, M. S., D. W. Dingman, and A. L. Sonenshein. 1985. Bacillus subtilis citB gene is regulated synergistically by glucose and glutamine. J. Bacteriol. 164:155-164. 35. Rothstein, R. 1983. One-step gene disruption in yeasts. Methods Enzymol. 101:202-211. 36. Ruzicka, F. J., and H. Beinert. 1978. The soluble "high potential" type iron-sulfur protein from mitochondria is aconitase. J. Biol. Chem. 253:2514-2517. 37. Sanger, F., S. Nicklen, and A. R. Coulson. 1977. DNA sequencing with chain-terminating inhibitors. Proc. Natl. Acad. Sci. USA 74:5463-5467. 38. Satrustegui, J., and A. Machado. 1977. The synthesis of yeast matrix mitochondrial enzymes is regulated by different levels of mitochondrial function. Arch. Biochem. Biophys. 184:355-363. 39. Schatz, G., and T. L. Mason. 1974. The biosynthesis of mitochondrial proteins. Annu. Rev. Biochem. 43:51-87. 40. Sharp, P. M., and W.-H. Li. 1987. The codon adaptation index-a measure of directional synonymous codon usage bias, and its potential applications. Nucleic Acids Res. 15:1281-1295. 41. Sherman, F., G. R. Fink, and J. B. Hicks. 1986. Methods in yeast genetics. Cold Spring Harbor Laboratory, Cold Spring

Harbor, N.Y. 42. Sholze, H. 1983. Studies on aconitase species from Saccharomyces cerevisiae, porcine and bovine heart, obtained by a modified isolation method. Biochim. Biophys. Acta 746:133137. 43. Slonimski, P. P., G. Perrodin, and J. H. Croft. 1968. Ethidium bromide induced mutation of yeast mitochondria: complete

VOL. 10, 1990 transformation of cells into respiratory deficient non chromosomal "petites." Biochem. Biophys. Res. Commun. 30:232239. 43a.Smith, T. F., and M. S. Waterman. 1981. Comparative biosequence metrics. J. Mol. Evol. 18:36-46. 44. Southern, E. M. 1975. Detection of specific sequences among DNA fragments separated by gel electrophoresis. J. Mol. Biol. 98:503-517.

45. Villafranca, J. J., and A. S. Mildvan. 1971. The mechanism of aconitase action. J. Biol. Chem. 246:772-779. 46. von Heijne, G. 1986. Mitochondrial targeting sequences may form amphiphilic helixes. EMBO J. 5:1335-1342. 47. Wales, D. S., T. G. Cartledge, and D. Lloyd. 1980. Effects of

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glucose repression and anaerobiosis on the activities and subcellular distribution of tricarboxylic acid cycle and associated enzymes in Saccharomyces carlsbergensis. J. Gen. Microbiol. 116:93-98. 48. Wu, M., and A. Tzagoloff. 1987. Mitochondrial and cytoplasmic fumarases in Saccharomyces cerevisiae are encoded by a single nuclear gene FUMI. J. Biol. Chem. 262:12275-12282. 49. Yanisch-Perron, C., J. Vieira, and J. Messing. 1985. Improved M13 phage cloning vectors and host strains: nucleotide sequences of the M13 mpl8 and pUC19 vectors. Gene 33:103-119. 50. Zitomer, R. S., D. L. Montgomery, D. L. Nichols, and B. D. Hall. 1979. Transcriptional regulation of the yeast cytochrome c gene. Proc. Natl. Acad. Sci. USA 76:3627-3631.