Aspartate Transcarbamoylase Genes of Pseudomonas putida

JOURNAL OF BACTERIOLOGY, Apr. 1995, p. 1751–1759 0021-9193/95/$04.0010 Copyright q 1995, American Society for Microbiology

Vol. 177, No. 7

Aspartate Transcarbamoylase Genes of Pseudomonas putida: Requirement for an Inactive Dihydroorotase for Assembly into the Dodecameric Holoenzyme MICHAEL J. SCHURR,1† JOHN F. VICKREY,1‡ ALAN P. KUMAR,1 ALAN L. CAMPBELL,1 RAYMOND CUNIN,2 ROBERT C. BENJAMIN,1 MARK S. SHANLEY,1 AND GERARD A. O’DONOVAN1* Department of Biological Sciences, University of North Texas, Denton, Texas 76203,1 and Department of Microbiology, Free University of Belgium, Brussels, Belgium2 Received 25 July 1994/Accepted 19 December 1994

The nucleotide sequences of the genes encoding the enzyme aspartate transcarbamoylase (ATCase) from Pseudomonas putida have been determined. Our results confirm that the P. putida ATCase is a dodecameric protein composed of two types of polypeptide chains translated coordinately from overlapping genes. The P. putida ATCase does not possess dissociable regulatory and catalytic functions but instead apparently contains the regulatory nucleotide binding site within a unique N-terminal extension of the pyrB-encoded subunit. The first gene, pyrB, is 1,005 bp long and encodes the 334-amino-acid, 36.4-kDa catalytic subunit of the enzyme. The second gene is 1,275 bp long and encodes a 424-residue polypeptide which bears significant homology to dihydroorotase (DHOase) from other organisms. Despite the homology of the overlapping gene to known DHOases, this 44.2-kDa polypeptide is not considered to be the functional product of the pyrC gene in P. putida, as DHOase activity is distinct from the ATCase complex. Moreover, the 44.2-kDa polypeptide lacks specific histidyl residues thought to be critical for DHOase enzymatic function. The pyrC-like gene (henceforth designated pyrC*) does not complement Escherichia coli pyrC auxotrophs, while the cloned pyrB gene does complement pyrB auxotrophs. The proposed function for the vestigial DHOase is to maintain ATCase activity by conserving the dodecameric assembly of the native enzyme. This unique assembly of six active pyrB polypeptides coupled with six inactive pyrC* polypeptides has not been seen previously for ATCase but is reminiscent of the fused trifunctional CAD enzyme of eukaryotes. polypeptide chains of 34 kDa each, and do not have associated with them a regulatory subunit (no pyrI-encoded counterpart) or the attendant heterotropic allosteric inhibition and activation. In B. subtilis (25, 43) and Bacillus caldolyticus (17), the pyrB gene (encoding ATCase) overlaps with the pyrC gene (encoding dihydroorotase [DHOase]). But the archetype of the smallest class of ATCases, the class C enzyme from B. subtilis, is distinct and does not interact with other enzymes of the pyrimidine pathway (43). The class A ATCases are large-molecular-mass enzymes (470 to 600 kDa) that were thought to be dimeric until Bergh and Evans (3) showed that the Pseudomonas fluorescens enzyme exists as a trimer with a 34-kDa catalytic chain in a dodecameric association with a 45-kDa chain of unknown origin and function. Similar results from studies with purified protein have been obtained by Shepherdson and McPhail (38) for the ATCase from Pseudomonas syringae. The 45-kDa chain was apparently not a homolog of the class B 17-kDa (pyrIencoded) regulatory polypeptide, since the binding site for nucleotide effectors was located on the 34-kDa chain. In those bacteria whose ATCase has been characterized, the enzyme exists separately from DHOase and carbamoylphosphate synthetase (CPSase). To date, the nucleotide sequences encoding 12 different ATCases have been determined and the primary protein structures have been deduced from the DNA sequences of their ATCase genes (Fig. 1). In eukaryotes, ATCase activity is found in a protein that also possesses activities for CPSase and sometimes DHOase as well. These polyproteins presumably arose as a result of gene fusion (22). In lower eukaryotes (Saccharomyces cerevisiae and Neurospora crassa), the enzymes for the first two steps of the pathway are encoded in a multifunctional

Aspartate transcarbamoylase (ATCase; EC 2.1.3.2) catalyzes the first committed step in the de novo synthesis of pyrimidine nucleotides. Properties of bacterial ATCases have been used by Bethell and Jones (5) to categorize ATCase enzymes into three different classes that follow phylogenetic lines. The ATCase from Escherichia coli is the archetypal class B enzyme, and it is a dodecamer of 310 kDa (47). Six identical catalytic polypeptides, organized as two enzymatically functional catalytic trimers (c3) and six identical regulatory polypeptides organized as three regulatory dimers (r2) define the holoenzyme. The 2c3:3r2 dodecameric structure is conserved among all enteric ATCases (50). The pyrBI operon of E. coli encodes the catalytic, pyrB, and the regulatory, pyrI, proteins of the native enzyme (32, 35). As expected, similar structural features are observed for those other enteric ATCases so far examined, specifically, Salmonella typhimurium (27), Serratia marcescens (2), and Proteus vulgaris (48). The 100-kDa-molecular-mass Bacillus subtilis enzyme is a typical class C enzyme, versus the larger 310-kDa enteric ATCase class B enzymes (5, 43). These class C enzymes correspond to the free catalytic trimers of the enteric enzyme (encoded by pyrB), are composed of three identical

* Corresponding author. Mailing address: Department of Biological Sciences, P.O. Box 5218, University of North Texas, Denton, TX 76203. Phone: (817) 565-3590. Fax: (817) 565-3821. Electronic mail address: [email protected]. † Present address: Department of Microbiology, University of Texas Health Science Center at San Antonio, San Antonio, TX 78284. ‡ Present address: Biochimie des Signaux de Regulation Moleculaire et Cellulaire, Universite´ Pierre et Marie Curie, 75006 Paris, France. 1751

1752

SCHURR ET AL.

J. BACTERIOL.

FIG. 1. Structural organization of glutamine amidotransferase (GATase), CPSase, ATCase, and DHOase genes in the pyrimidine pathway. The arrangements are shown for the genes encoding the first three enzymes of pyrimidine biosynthesis from E. coli (19, 32, 35), S. marcescens (2), S. typhimurium (27), P. vulgaris (48), B. subtilis (25, 34), B. caldolyticus (17), D. melanogaster (11, 15), M. auratus (39, 41), S. cerevisiae (42), D. discoideum (14), Pseudomonas aeruginosa (46), and P. putida (37; this paper). Genes that encode products that are known to have nontranslated bases between them in a polycistronic mRNA and that form a complex or copurify are depicted connected by a single black line. Genes for polyproteins that possess fused activities are drawn as continuous bars, for example, the CPSase, ATCase, and DHOase (CAD) complex from hamster cells. In those organisms whose genes are contiguous, the bars are drawn connected and on separate lines. Genes that display translational coupling are depicted by overlapping bars on separate lines. By indicating the genes on separate lines, no inference is intended with respect to the reading frame. Genes are not drawn to scale.

200-kDa polyprotein encoded by the ura2 gene in S. cerevisiae (12). This multifunctional CPSase-ATCase complex from S. cerevisiae cells contains, in addition, a defective DHOase-like domain that is not catalytically active (Fig. 1) (42). In higher eukaryotes, including Dictyostelium discoideum (14), Drosophila melanogaster (11, 15) and Mesocricetus auratus (39, 41), an additional enzyme activity, namely, DHOase, is found, such that the first three enzyme activities of the pyrimidine biosynthetic pathway are found in a single trifunctional protein of a molecular mass of 220 kDa (22). Like the pyrB and pyrC genes in Bacillus species, those of Pseudomonas putida overlap, but unlike the ATCase of Bacillus species, that of P. putida is dependent upon expression of the overlapping pyrC gene. In this communication, we use the DNA sequences of the ATCase genes to identify two polypeptides: a smaller 36.4-kDa catalytic polypeptide and a larger 44.2-kDa polypeptide. We provide evidence that the native enzyme is a dodecamer of a molecular mass of 482 kDa, that the active site and the nucleotide effector binding site are indeed located on the smaller polypeptide as previously reported (3), that the larger polypeptide is encoded by a cryptic pyrC gene that does not encode DHOase activity, and that DHOase is encoded by a gene residing elsewhere on the chromosome. Expression, transcriptional regulation, and promoter structure will be addressed in a future publication (7). MATERIALS AND METHODS Bacterial strains and plasmids. The bacterial strains and plasmids used in this study are listed in Table 1. Cosmid pMO020619 contains 25 kb of P. putida PPN1 chromosomal DNA. This was kindly provided by Bruce Holloway of Monash University, Clayton, Australia. Plasmid pMJS27 contains an 8.4-kb EcoRI fragment of pMO020619, while plasmid pMJS29 contains a 3.45-kb PstI fragment from pMJS27 with the P. putida pyrBC9 genes in the orientation identical to that of the lac promoter in the pUC series plasmids.

Media and growth conditions. E. coli and P. putida strains were grown in Luria-Bertani (LB) (4) broth at 37 and 308C, respectively, with aeration. Antibiotics were added to the medium, depending on which plasmids the cells contained, as follows: cells containing pUC-derived plasmids received 100 mg of ampicillin per ml, cells containing pMO020619 received 12.5 mg of tetracycline per ml, and cells containing pGP1-2 received 60 mg of kanamycin per ml. The Pseudomonas basal medium was that described by Ornston and Stanier (31) supplemented with Hutner’s Metals 44 (9), with succinate at a concentration of 10 mM as carbon source. The E. coli basal medium was as described by Miller (28) supplemented with thiamine (1.0 mg/ml) and uracil (50 mg/ml). The carbon source was glucose at a concentration of 0.2%. Genetic manipulations. Transformation of DNA into E. coli DH5a and TB2 strains was carried out by the method of Huff et al. (20). Restriction digestions were performed according to the specifications of the manufacturer. Ligation mixes typically contained 1 Weiss unit of T4 DNA ligase, DNA at 1 mg/ml and 13 ligase buffer in a total volume of 20 ml. These mixtures were incubated at room temperature overnight. Recombinant derivatives of pUC19 were screened on LB plates containing ampicillin (100 mg/ml) and 25 mg of X-Gal (5-bromo4-chloro-3-indolyl-b-D-galactopyranoside). Rapid plasmid preparations were done according to the protocol of Zhou et al. (52), and bulk plasmid preparations were done by a previously described method (26). Southern hybridizations. In order to locate P. putida pyrB, a purified 480-bp DdeI fragment from plasmid pEK2 (Table 1), which corresponded to the active site of the E. coli ATCase from the pyrB gene, was used as a probe for Southern analysis. EcoRI and PstI restriction digests of cosmid pMO020619 DNA were transferred to GeneScreen membranes per the manufacturer’s instructions. The 480-bp DdeI probe was end labeled with [g-32P]ATP by T4 polynucleotide kinase (26). The labeled probe was recovered free from unincorporated label by Sephadex G-50 chromatography and ethanol precipitation and was allowed to hybridize to the membrane for 12 h at 558C. The membrane was washed and exposed to Kodak XAR5 film for 3 days with a DuPont Lightning Plus intensifying screen. Nondenaturing activity polyacrylamide gels. Bacteria were grown at 378C for E. coli and 308C for P. putida overnight to an A590 of 1.0 in 5 ml of LB by the method of Kedzie (23). The bacterial cells were centrifuged, resuspended in 300 ml of 40 mM potassium phosphate buffer (pH 7.0)–1 mM dithiothreitol–0.2 mM zinc acetate, and disrupted by sonication for 30 s with a Branson model 200 sonifier. The cell extract was centrifuged for 10 min at 10,000 3 g. Aliquots (20 ml) were loaded onto a 4 to 20% nondenaturing polyacrylamide gradient gel, and the reaction mixtures were run at 90 V for 10 h. The gel was placed in 250 ml of ice-cold 50 mM histidine (free base), pH 7.0, for 5 min to which 5 ml of 1.0 M aspartate and 10 ml of 0.1 M carbamoylphosphate were added and allowed to

ASPARTATE TRANSCARBAMOYLASE GENES OF P. PUTIDA

VOL. 177, 1995

1753

TABLE 1. Strains and plasmids used in this study Strain or plasmid

Strains P. putida PPN1 PPN1137 P. fluorescens CW1012 CW1013 E. coli DH5a DH5aF9 TB2 HB101 X7014a Plasmids pUC19 pEK2 pLA2917 pMO020619 pMJS27 pMJS28 pMJS29 pT7-6 pT7-5 pGP1-2 pBJRC18

Genotype or description

Source

Wild type pyrB

B. W. Holloway B. W. Holloway

pyrB2 pyrC

T. P. West T. P. West

F2 f80dlacZDM15 D(lacZYA-argF)U169 deoR recA1 endA1 hsdR17 (rK2 mK1) supE44 l2 thi1-1 gyrA96 relA1 F9 f80dlacZDM15 D(lacZYA-argF)U169 deoR recA1 endA1 hsdR17 (rK2 mK1) supE44 l2 thi1-1 gyrA96 relA1 DpyrBI argF F2 supE44 hsdS 20(rB2 m9B2) recA13 ara-14 proA2 lacYI galK2 rpsL20 xyl5 mtl-1 thi leu l2 pyrC46 purB51 lacZ43 thi-1 malA1 mtl-2 xyl-7 rpsL125 (CGSC 5358)

GIBCO-BRL

R. L. Switzer

Apr lacZ DM15 E. coli pyrB in pUC7 Tcr cosmid pLA2917 with 25 kb of P. putida chromosome P. putida pyrBC9; 8.4-kb EcoRI insert in pUC19 P. putida pyrBC9; 2.8-kb insert in pUC19 with orientation identical to that of lac promoter P. putida pyrBC9; 3.45-kb partial PstI insert in pUC19 with orientation identical to that of lac promoter Apr, T7 promoter Apr, T7 promoter Kmr, T7 RNA polymerase S. typhimurium pyrC in pBR322

J. Messing E. R. Kantrowitz B. W. Holloway B. W. Holloway This study This study This study S. Tabor S. Tabor S. Tabor R. A. Kelln

react for 20 min. The gel was rinsed three times with ice-cold distilled water to remove reactants. The enzymatically released Pi trapped in the gel was precipitated by a reaction with 3 mM lead nitrate in ice-cold 50 mM histidine, pH 7.0, for 10 min. The soluble lead ions were removed with three changes of ice-cold water. The gel was then stained with 300 ml of 5% ammonium sulfide, which converted the white lead phosphate precipitate into dark lead sulfide bands (6, 23). Selective labeling of plasmid-encoded proteins. Plasmids pT7-5 and pT7-6 containing T7 promoters were used in conjunction with plasmid pGP1-2 encoding T7 RNA polymerase to express the cloned P. putida pyrBC9 genes. PstI fragments of 3.45 kb for the P. putida pyrBC9 genes were inserted into pT7-5 and pT7-6. These plasmids containing the insert in both orientations were transformed into E. coli HB101 containing plasmid pGP1-2 with the T7 RNA polymerase under the control of a temperature-inducible cI857 repressor and a lpL promoter. The method of Tabor was used to visualize the 35S-labeled pyrB and pyrC9 polypeptides (44, 45). ATCase enzyme assay. ATCase activity was assayed by measuring the amount of carbamoyl-L-aspartate produced in 30 min at 308C by the method of Gerhart and Pardee (16) with modifications and by the color development procedure of Prescott and Jones (33). Purification of the P. putida ATCase. The P. putida ATCase was purified from E. coli TB2 (Table 1) which contained the plasmid pMJS-29 carrying the cloned P. putida ATCase genes. The enzyme was purified from 5 liters of culture by the method of Adair and Jones (1) as modified by Bergh and Evans (3), with the following exceptions. The size exclusion chromatographic step employed a calibrated column (50 by 3 cm) of Sephacryl S-300. Instead of purification over hydroxyapatite, 100-ml aliquots of concentrated, partially purified protein from the size exclusion column were purified by linear gradient elution through a Dionex Propac ion exchange column (45 by 250 mm) run on a Waters highperformance liquid chromatography (HPLC) system. The system consisted of two model 510 pumps, a model 680 gradient controller, a U6K injector, a model 740 Data Module and a SpectroMonitor model 5000 photodiode array detector. The gradient was established by running from 0% buffer A (100 mM Tris-HCl [pH 8.0]) to 100% buffer B (100 mM Tris-HCl [pH 8.0], 1.0 M NaCl) over a period of 60 min at a flow rate of 1 ml/min. Fractions (0.5 ml each) were collected, assayed for the presence of ATCase activity, and pooled. Multiple runs and preparations were pooled to obtain quantities of the enzyme sufficient for subsequent molecular weight and kinetic analyses. Sequencing protocols. The chain-terminating sequencing method of Sanger et al. was utilized (36). A Sequenase system from United States Biochemical was used. Double-stranded sequencing was done with oligonucleotides synthesized by Biosynthesis (Lewisville, Tex.) as primers. Both strands of the 3.45-kb PstI fragment from pMJS29 containing P. putida pyrBC9 were sequenced. The ddITP

GIBCO-BRL W. D. Roof

and 7-deaza-GTP reactions of the Sequenase system were used to resolve compressions due to the high G1C content of the P. putida DNA. Nucleotide sequence accession numbers. The GenBank accession numbers for P. putida pyrB and pyrC9 are M97253 and M97254, respectively.

RESULTS Subcloning of P. putida pyrBC*. Cosmid pMO020619, containing 25 kb of P. putida chromosomal DNA in the parent plasmid pLA2917, complemented the pyrB auxotroph, P. putida PPN1137, upon conjugation from E. coli S17-1. This confirmed the presence of a functional pyrB gene in the cosmid. After digestion, transfer, and hybridization to an activesite probe derived from E. coli pyrB, two homologous fragments were identified. A 2.8-kb PstI fragment and an 8.4-kb EcoRI fragment hybridized to a 32P-labeled pyrB-specific probe (Fig. 2). Each fragment was ligated independently into pUC19, and the resulting plasmids were used to transform appropriate E. coli recipient strains. Only plasmid pMJS27, carrying the inserted 8.4-kb EcoRI hybridizing fragment, was able to complement the E. coli pyrB mutant TB2 strain (Table 1). The plasmid containing the hybridizing 2.8-kb PstI fragment was designated pMJS28 and did not complement pyrB mutant strains, nor did it produce measurable ATCase activity. Partial PstI digestion of pMJS27 resulted in a complementing clone (pMJS29) which contained a 3.4-kb PstI fragment. Plasmid pMJS29 harbors the 2.8-kb PstI fragment of pMJS28 plus an additional 600-bp PstI fragment. This 600-bp fragment included the promoter region and the first 28 nucleotides of the pyrB gene. Thus, the P. putida pyrB gene from the parent cosmid pMO020619, was subcloned as a 2.8-kb hybridizing PstI fragment (pMJS28) and as complementing 8.4-kb EcoRI (pMJS27)- and 3.4-kb PstI (pMJS29)-derived fragments. The P. putida pyrBC9 genes on pMJS29 complemented the pyrB E. coli strain (TB2) and not the E. coli pyrC strain

1754

SCHURR ET AL.

FIG. 2. Hybridization of E. coli pyrB DNA to P. putida ATCase clones. A 480-bp DdeI fragment of E. coli pyrB from plasmid pEK2 was labeled and used as a probe. Lane 1 contains PstI-cut cosmid pMO020619 carrying the pyrBC genes from P. putida. Lane 2 contains EcoRI-digested pMO020619. The size markers on the left correspond to HindIII-cut lambda DNA bands. The band denoted by I is the 8.4-kb EcoRI fragment shown in the diagram that was used to construct plasmid pMJS27. The band denoted by II is the 2.8-kb PstI fragment shown in the diagram that was used to construct plasmid pMJS28 and which does not carry the entire ATCase gene. The top band in lane 2 corresponds to an incompletely digested plasmid.

(X7014a). Moreover, the pyrC gene from S. typhimurium carried on plasmid pBJRC18 (Table 1) did complement the E. coli pyrC mutation, thereby restoring the strain to pyrimidine prototrophy. Since the cloned P. putida pyrB gene was capable of complementing the E. coli pyrB mutation, there were no intrinsic barriers to expression of the P. putida genes in the heterologous host. Nondenaturing ATCase activity polyacrylamide gels. Each of the plasmids, pMJS27, pMJS28, and pMJS29, and parent cosmid pMO020619 were tested for pyrB-encoded ATCase activity after separation on nondenaturing gradient gels specifically stained for enzyme activity (6, 23). These gradient gels afforded the separation by apparent molecular weight and the specific staining of ATCase activity in the clones. Cosmid pMO020619 produced a polypeptide with an approximate molecular mass of 480 kDa when expressed in the E. coli pyrB mutant strain, TB2 (Fig. 3, lane 1). Likewise, plasmids pMJS27 and pMJS29 produced a large-molecular-mass 480-kDa ATCase activity when expressed in E. coli (Fig. 3, lanes 2 and 4). The native E. coli enzyme was apparent at a relative position in the gel corresponding to its expected molecular mass of 310 kDa. P. putida pyrB subclone pMJS28 did not produce observable ATCase activity distinct from the E. coli enzyme activity of the parent strain (Fig. 3, lane 3).

FIG. 3. Nondenaturing ATCase activity polyacrylamide gradient gel. Lane 1 contains cell extract from E. coli TB2 with cosmid pMO020619 (cosmid pLA2917 with 25 kb of P. putida chromosomal DNA encoding ATCase). Lane 2 contains cell extract from E. coli DH5a with pMJS28 (pUC19 with the 8.4-kb EcoRI fragment encoding ATCase). Lane 3 contains cell extract from E. coli DH5a with pMJS28 (pUC19 with a 2.8-kb PstI fragment that lacks the 59 end of the ATCase genes). Lane 4 contains cell extract from E. coli DH5a containing pMJS29 (pUC19 with a 3.4-kb incompletely digested PstI fragment encoding the entire ATCase).

J. BACTERIOL.

FIG. 4. SDS-polyacrylamide gel electrophoresis of P. putida ATCase. Lane 1 contains denatured purified P. putida ATCase. The molecular weight (MW) standards were bovine plasma albumin (66 kDa), egg albumin (ovalbumin; 45 kDa), carbonic anhydrase (29 kDa), and lysozyme (14 kDa). After HPLC fractionation over an ion-exchange column, two polypeptides with apparent molecular masses of 34 and 46 kDa (arrows) were seen after silver protein staining.

Purification of P. putida ATCase. The enzyme was purified from 5-liter cultures of the recombinant E. coli strain carrying the cloned P. putida ATCase. The enzyme preparation obtained after elution from the HPLC stained as a single band on native 7% polyacrylamide gels. The same preparation produced two separate bands when separated on sodium dodecyl sulfate (SDS)-polyacrylamide gels (Fig. 4). The two bands had apparent molecular masses of approximately 34 and 46 kDa when compared to the standards. The purified enzyme preparation was again passed over the calibrated Sephacryl S-300 column and was eluted with an apparent molecular mass of 480 kDa. The purified ATCase from P. putida again produced a single band on nondenaturing gradient polyacrylamide gels and migrated to a position corresponding to a molecular mass of 480 kDa, as determined by both activity and protein staining. These values were similar to those for P. fluorescens ATCase (3) and suggested a possible dodecameric structure assembled from six 34-kDa polypeptides and six 46-kDa polypeptides. These approximate values for the molecular masses are refined with the values deduced from the translated DNA sequences below. DHOase activity was monitored throughout the purification. Fractions containing ATCase were distinct from DHOase-containing fractions. The ATCase activity in partially purified preparations eluted at a molecular mass of 480 kDa after sieving over Sephacryl S-300, while DHOase activity was found in two peaks at lower molecular masses. No DHOase activity was eluted from the HPLC ion-exchange column that was associated with fractions containing ATCase activity. DNA sequencing of P. putida pyrBC* genes. The nucleotide sequence of the 3.4-kb PstI fragment was determined by using the Sanger dideoxy method (36) and has been deposited in GenBank (accession numbers M97253 and M97254 for pyrB and pyrC9, respectively). Both strands were sequenced completely. This region contains two open reading frames (ORFs), one corresponding to the pyrB gene, as determined by its homology to the E. coli gene, and one corresponding to a pyrClike gene, herein designated pyrC9. The proximal ORF is the P. putida pyrB gene. It is 1,005 bp long, including its stop codon TGA, and exhibits a predicted translation of 334 amino acids and a deduced monomeric mo-

VOL. 177, 1995


FIG. 5. Identification of P. putida ATCase polypeptides. The 8.4-kb EcoRI fragment from plasmid pMJS27 containing the pyrBC9 genes of P. putida was used in a T7 expression assay. Molecular masses (in kilodaltons) for the standards (Std.) are indicated on the left. Lane 1 contains [35S]methionine-labeled cell extracts for the P. putida ATCase genes expressed in their proper transcriptional orientation. Lane 2 contains labeled extracts from cells carrying plasmids expressing the genes from the opposite orientation. Bands with molecular masses of 45 and 37 kDa were observed (lane 1) after electrophoresis (arrowheads) and correspond to the pyrC9 and pyrB polypeptides, respectively. The 29-kDa band corresponds to the b-lactamase of the vector. The 45-kDa DHOase-like polypeptide contains only 3 methionine residues, while the 37-kDa ATCase catalytic polypeptide contains 12 methionine residues; thus, the greater signal corresponds to the smaller subunit.

lecular mass of 36.4 kDa. The distal ORF is 1,275 bp long, including its TGA stop codon, and produces a polypeptide of 424 amino acids with a deduced monomeric molecular mass of 44.2 kDa. This second ORF shows about 30% sequence homology to known pyrC genes. The coding region of the pyrC9 gene overlaps the 39 end of the pyrB gene by 4 bp. The pyrC9 start codon is GTG. The GTG start of pyrC9 thus includes the 39 G of the terminal glutamine codon of pyrB, as well as the 59 TG of the TGA stop codon of pyrB. This GTG start codon was chosen because of its position relative to the consensus ShineDalgarno sequence, because the next methionine codon in pyrC9 is 200 codons away, because the amino terminus of the peptide encoded is homologous to the P. fluorescens polypeptide (3), and because this GTG start yields a polypeptide of the predicted molecular mass. T7 translational analysis of pyrBC* genes of P. putida. The T7 expression system of Tabor (44) was used to label the P. putida polypeptides produced from the two ORFs in plasmid pMJS29. The 3.4-kb PstI fragment gave rise to two translational products visible as bands corresponding to molecular masses of 37 and 45 kDa. No protein bands were seen when attempts were made to produce expression when the same fragment was inserted in the opposite orientation (Fig. 5). Since the 3.4-kb fragment contains two ORFs that yield the predicted translational products of 36.4 and 44.2 kDa driven by an inducible T7 promoter system and since this same fragment expresses a functional 480-kDa enzyme, we conclude that ATCase from P. putida is formed from the two polypeptide chains encoded by these genes (Fig. 5). Alignments of the deduced P. putida ATCase sequence with known ATCases. The deduced amino acid translations of the pyrB and pyrC9 genes of P. putida show significant similarity to the known sequences of these genes in other organisms (Fig. 6 and 7). The predicted ATCase and DHOase-like amino acid sequences were aligned and compared to other known published sequences using the PILEUP program of the University of Wisconsin Genetics Computer Group package (13).

1755

ATCase from B. subtilis exhibited the greatest amino acid sequence homology with the translational product of the P. putida pyrB gene (Fig. 6). Amino acid identity was observed at 46% of the positions. Only 4 residues (12 of 16 residues were identical) differed between the active site of E. coli and the corresponding position of the P. putida predicted enzyme sequence reported here. The exceptions are substitutions of methionine for leucine at position Met-265 of the E. coli ATCase, of glutamate for alanine at Ala-275, of serine for threonine at Thr-276, and of alanine for phenylalanine at Phe-295 of the E. coli ATCase (Fig. 6). Alignment of the deduced amino acid sequence of the pyrC* ORF with those of known DHOases. When a global homology search was performed by using the deduced amino acid sequence of P. putida pyrC9 with GenBank and National Biomedical Research Foundation databases, seven known DHOase sequences showed statistically significant similarity to the predicted amino acid sequence described in this paper (Fig. 7). The greatest sequence similarity was again observed with the corresponding enzyme from B. subtilis; 31% identity was seen. The deduced P. putida DHOase-like sequence was compared to those of the seven other known DHOases. The P. putida pyrC9 ORF is bigger than those of the other known enzymes, so that allowance must be made for substitutions and deletions in the alignment. Three regions of similarity, namely, middle, carboxy-terminal, and amino-terminal portions, have been proposed as the components of the active site in E. coli DHOase. The middle region of E. coli DHOase contains conserved Met-43, Pro-44, and Asn-45, while P. putida contains Pro-93, Pro-94, and Gln-95. Amino acids flanking this middle region are also conserved in the P. putida polypeptide. The carboxy-terminal region of E. coli DHOase contains conserved residues Asp-251, Ser-252, Ala-253, Pro-254, His-255, and Lys-260. The P. putida polypeptide contains His-306, His-307, Gln-308, Pro-309, His-310, and Lys-315 in this region. Within the amino-terminal region of the proposed active site are two histidine residues that are conserved in all known DHOases (40). The P. putida polypeptide lacks these conservative histidines and has glycine and serine at these positions. Other DHOases have five conserved histidines, which are believed to lie in the active-site regions. Only one of these active-site histidines (His-310) is found in the P. putida polypeptide (Fig. 7). The non-active site histidines are replaced by glutamine residues (Gln-232 and Gln-261). The lack of histidines in the P. putida pyrC9 polypeptide suggests only that it is not functional as a DHOase, since the precise threedimensional structure and catalytic mechanism have not been elucidated. In accordance with this finding was the inability of this pyrC9 gene to complement pyrC auxotrophs of E. coli. DHOase activity in crude cell extracts and partially purified ATCase preparations did not coelute or copurify, and the purified P. putida ATCase had no associated DHOase activity. Thus, it appears that the pyrC9-encoded polypeptide is not the functional DHOase of P. putida. DISCUSSION Pseudomonas ATCases or class A enzymes have been the subject of numerous studies (10, 21, 38, 49). ATCase has been purified and characterized to various degrees from a number of Pseudomonas species (1, 3, 5, 38, 46). Here, we report the first complete primary sequence of a Pseudomonas ATCase. Sequence analysis of the cloned P. putida ATCase genes revealed that the enzyme was expressed from two ORFs which overlap by 4 bp. The first ORF is pyrB, which encodes a 36.4kDa polypeptide. The deduced amino acid sequence for this gene shows 46% sequence homology to that of the B. subtilis ATCase and 40% homology to those of the E. coli and hamster

1756

SCHURR ET AL.

J. BACTERIOL.

FIG. 6. Amino acid comparisons of known ATCases with the ATCase from P. putida. The amino acid sequences from P. putida (Pputi [this paper]), B. subtilis (Bsubt [25, 34]), S. typhimurium (Styph [27]), E. coli (Ecoli [19, 32, 35]), S. marcescens (Smarc [2]) P. vulgaris (Pvulg [48]), D. discoideum (Ddisc [14]), M. auratus (Maura [39, 41]), S. cerevisiae (Scere [29]), and D. melanogaster (Dmela [11, 15]) are aligned. Residues shown to be involved in the binding of substrates at the active site of E. coli ATCase are indicated (F). Identical or conservatively changed residues in the aligned sequences are indicated by shaded boxes. The sequence numbers are indicated on the right.

enzymes. The second ORF is a pyrC-like gene which encodes a 44.2-kDa polypeptide with remarkable sequence homology to DHOases from B. subtilis (31%), hamster (30%), and E. coli (18%). Purified ATCase from P. putida exhibited an apparent molecular mass of 480 kDa in gel filtration experiments. The properties of the enzyme from cloned genes were identical to those of the wild-type ATCase. Velocity-substrate plots of the

wild type as well as of recombinant strains expressing the cloned genes of the native enzyme from P. putida showed Michaelis-Menten (hyperbolic) kinetics with ATP inhibition. The enzyme from P. putida contained two polypeptides that likewise assembled into a 480-kDa protein composed of 37and 45-kDa polypeptide units, as determined by gel electrophoresis after in vitro translation (Fig. 5) and as determined by SDS gel electrophoresis of the purified ATCase (Fig. 4). Thus,

VOL. 177, 1995


1757

FIG. 7. Amino acid comparisons of the known DHOases with the pyrC9 ORF of P. putida. The amino acid sequences from S. typhimurium (Styph [30]), E. coli (Ecoli [51]), S. cerevisiae (Scere [18, 42]), D. melanogaster (Dmela [11, 15]), M. auratus (Maura [39, 40]), D. discoideum (Ddisc [14]), P. putida (Pputi [37; this paper]) and B. subtilis (Bsubt [34]) are aligned. The residues conserved in all of the DHOases (F) and the five histidine residues that are also conserved and are thought to be involved in the binding of substrate at the active site (■) are indicated. Identical or conservatively changed residues in the aligned sequences are indicated by shaded boxes. The sequence numbers are indicated on the right.

the Pseudomonas ATCases are dodecamers of six 36.4-kDa pyrB polypeptides and six 44.2-kDa pyrC9 polypeptides, as concluded from the deduced amino acid sequences. Moreover, Bergh and Evans (3) have purified the related P. fluorescens ATCase and have sequenced the amino-terminal end of its 34-

and 45-kDa polypeptide subunits. In each of these chains, 14 of 19 amino acids are identical to the corresponding amino acids in our P. putida ATCase. However, these two enzymes show no sequence homology of their amino termini with those of any other known ATCase (Fig. 5). Computer comparisons of the

1758

SCHURR ET AL.

amino acid sequence of our P. putida ATCase with other known complete ATCase sequences revealed an N-terminal 11-amino-acid extension unique to the Pseudomonas ATCase (3, 39, 46). Of significance, this extension of the Pseudomonas pyrB polypeptides is homologous with that region of the E. coli and other enteric pyrI polypeptides which binds the effectors (2, 3, 27, 32, 37, 46). It has recently been shown (3) that labeled ATP, the negative effector, does indeed bind to the 34-kDa polypeptide and not to the 45-kDa polypeptide in P. fluorescens. We tested for the presence of such a functional ATP binding site on the amino-terminal extension of the P. putida enzyme by deleting the DNA encoding the 34 terminal amino acids. The ATCase expressed and assembled from this truncated pyrB gene was active but was no longer inhibited by ATP (24). The finding of a consensus ATP/GTP nucleotide binding site motif within this region, as revealed by the Genetics Computer Group Motifs program (13), further suggests that this N-terminal region provides the nucleotide effector binding site. With regard to the activity of the pyrC9 polypeptide, the P. putida pyrB PPN1137 strain lacks ATCase activity yet contains an active DHOase. A similar result was obtained for pyrB mutants of P. fluorescens (8). Taken together, these results suggest that DHOase activity in the cell can exist independently of ATCase activity. In accordance with the data above, there are pyrC mutants in P. fluorescens that have normal ATCase activity (8). Such pyrC mutations either must not affect ATCase assembly and activity or must reside in a separate pyrC unlinked to the pyrBC9 described here. Recent experiments by Vickrey (46) with purified P. aeruginosa ATCase, as well as the results of Bergh and Evans (3) with purified P. fluorescens ATCase, suggest that the 44.2-kDa pyrC9 polypeptide is not an active DHOase but is required for assembly of ATCase into an active form. In keeping with this idea, we have searched the P. putida genome with pyrC probes from B. subtilis (34) and S. typhimurium (30) and have found cross-hybridizing bands on Southern transfers distinct from and in addition to the known pyrC9 restriction fragments (7). Comparison of the predicted amino acid sequence of the pyrC9 polypeptide with those of known DHOases shows a number of substitutions at critical amino acid positions corresponding to the active site; specifically, only one of five critical histidines is present (Fig. 6). The seven other known DHOases have five conserved histidines. The P. putida pyrC9 polypeptide linked to pyrB has only one of these. Thus, the DHOase activity in P. putida is not coupled to the pyrBC9 genes and the functional pyrC gene resides elsewhere on the chromosome. On the basis of the optimized protein alignments produced by the Genetics Computer Group PILEUP program (13), P. putida ATCase, like other ATCase catalytic chains, possesses highly conserved amino acid homologies at regions responsible for catalysis and substrate binding. Despite the lack of the pyrI-encoded regulatory polypeptide, overall homology with the E. coli enzyme was approximately 40%, with sequence homology rising to 60% for these regions involved with catalysis and substrate binding. The P. putida ATCase has an amino-terminal carbamoylphosphate-binding domain and a carboxy-terminal aspartate-binding domain, like the E. coli ATCase. The Pseudomonas ATCase has a functional nucleotide effector binding site whose location has been shifted to the amino terminus of the 36.4-kDa catalytic polypeptide. In the absence of any regulatory dimers, ATCase in P. putida retains a similar dodecameric arrangement involving its 36.4kDa ATCase polypeptide and its 44.2-kDa pyrC9 analog. The only definitive function for this 44.2-kDa analog must be to maintain the quaternary structure, as without this 44.2-kDa

J. BACTERIOL.

polypeptide the ATCase does not have DHOase activity. The lack of DHOase activity for the pyrC9 polypeptide was demonstrated by the properties of the purified enzyme: the inability of pyrC9 clones to complement pyrC mutants, the presence of DHOase activity in fractions distinct from ATCase fractions during the purification, and the identification of the somatic pyrC gene from P. putida elsewhere on the chromosome. A role for this DHOase-like polypeptide other than to maintain the dodecameric structure of the active holoenzyme has not been identified. The importance of the pyrC9 polypeptide in the assembly is proposed on the basis of the fact that activity is not maintained upon dissociation; the presence of free catalytically active trimers has not been demonstrated (3). In our work, no active trimer could be assembled and detected from subclones of the enzyme that expressed only the pyrB catalytic subunit polypeptide. These subclones did not demonstrate any assembly of catalytic chains into large-molecular-weight catalytically active forms. The sequence overlap of the native genes in intact clones indicates that translational coupling might occur such that stoichiometric amounts of the two polypeptide chains would be produced for assembly into the active dodecamer. To our knowledge, an assembly such as this with six active pyrB ATCase catalytic chains coupled with six pyrC9 polypeptides is found nowhere else in prokaryotic systems. Curiously, in eukaryotes, the first three enzymes involved in pyrimidine biosynthesis (CPSase, ATCase, and DHOase) reside in the single trifunctional polypeptide, CAD. It is possible therefore that the 2A3:3D2 dodecamer described here may be a vestige of the eukaryotic CAD complex or even its progenitor. Regardless, in order to accommodate essential ATCase and DHOase functions, P. putida of necessity had to duplicate its pyrC gene, with one duplicate to provide catalytic activity and the other [now pyrC9] to maintain the dodecameric configuration of ATCase, without which it is not active. ACKNOWLEDGMENTS This work was supported in part by Faculty Research Awards (University of North Texas) to G.A.O. and M.S.S. and by an NSF Fellowship to J.F.V. We thank Dayna Brichta and Deb Beck for help with the preparation of the manuscript. John Houghton provided a critical review. T. P. West and B. W. Holloway freely and graciously provided strains. Jeffrey Davidson contributed unpublished DNA sequence data and critical advice about our work. REFERENCES 1. Adair, L. B., and M. E. Jones. 1972. Purification and characteristics of aspartate transcarbamylase from Pseudomonas fluorescens. J. Biol. Chem. 247:2308–2315. 2. Beck, D., K. M. Kedzie, and J. R. Wild. 1989. Comparison of aspartate transcarbamoylases from Serratia marcescens and Escherichia coli. J. Biol. Chem. 264:16629–16637. 3. Bergh, S. T., and D. R. Evans. 1993. Subunit structure of class A aspartate transcarbamoylase from Pseudomonas fluorescens. Proc. Natl. Acad. Sci. USA 90:9819–9822. 4. Bertani, G. 1951. Studies on lysogenesis. I. The mode of phage liberation by lysogenic Escherichia coli. J. Bacteriol. 62:293–300. 5. Bethell, M. R., and M. E. Jones. 1969. Molecular size and feedback-regulation characteristics of bacterial aspartate transcarbamylases. Arch. Biochem. Biophys. 134:352–365. 6. Bothwell, M. 1975. Ph.D. thesis. University of California, Berkeley. 7. Brichta, D. M., A. P. Kumar, and G. A. O’Donovan. Unpublished data. 8. Chu, C.-P., and T. P. West. 1990. Pyrimidine biosynthetic pathway of Pseudomonas fluorescens. J. Gen. Microbiol. 136:875–880. 9. Cohen-Bazire, G., W. R. Sistrom, and R. Y. Stanier. 1957. Kinetic studies of pigment synthesis by non-sulphur purple bacteria. J. Cell. Comp. Physiol. 49:25–68. 10. Condon, S., J. K. Collins, and G. A. O’Donovan. 1976. Regulation of arginine and pyrimidine biosynthesis in Pseudomonas putida. J. Gen. Microbiol. 92: 375–383. 11. Davidson, J. N., and C. B. Kem. 1994. Revision in sequence of CAD aspar-

VOL. 177, 1995


tate transcarbamoylase domain of Drosophila. J. Mol. Biol. 243:364–366. 12. Denis-Duphil, M. 1989. Pyrimidine biosynthesis in Saccharomyces cerevisiae: the ura2 cluster gene, its multifunctional enzyme product, and other structural or regulatory genes involved in de novo UMP synthesis. Biochem. Cell Biol. 67:612–631. 13. Devereux, J., P. Haeberli, and O. Smithies. 1984. A comprehensive set of sequence analysis programs for the VAX. Nucleic Acids Res. 12:387–395. 14. Faure, M., J. H. Camonis, and M. Jacquet. 1989. Molecular characterization of a Dictyostelium discoideum gene encoding a multifunctional enzyme of the pyrimidine pathway. Eur. J. Biochem. 179:345–358. 15. Freund, J. N., and B. P. Jarry. 1987. The rudimentary gene of Drosophila melanogaster encodes four enzymic functions. J. Mol. Biol. 193:25–36. 16. Gerhart, J. C., and A. B. Pardee. 1962. The enzymology of control by feedback inhibition. J. Biol. Chem. 237:891–896. 17. Ghim, S.-Y., and J. Neuhard. 1994. The pyrimidine biosynthesis operon of the thermophile Bacillus caldolyticus includes genes for uracil phosphoribosyltransferase and uracil permease. J. Bacteriol. 176:3698–3707. 18. Guyonvarch, A., M. Nguyen-Juilleret, J. C. Hubert, and F. Lacroute. 1988. Structure of the Saccharomyces cerevisiae URA4 gene encoding dihydroorotase. Mol. Gen. Genet. 212:134–141. 19. Hoover, T. A., W. D. Roof, K. F. Foltermann, G. A. O’Donovan, D. A. Bencini, and J. R. Wild. 1983. Nucleotide sequence of the structural gene (pyrB) that encodes the catalytic polypeptide of aspartate transcarbamoylase of Escherichia coli. Proc. Natl. Acad. Sci. USA 80:2462–2466. 20. Huff, J. P., B. J. Grant, C. A. Penning, and K. F. Sullivan. 1990. Optimization of transformation of Escherichia coli with plasmid DNA. BioTechniques 9:570–577. 21. Isaac, J. H., and B. W. Holloway. 1968. Control of pyrimidine biosynthesis in Pseudomonas aeruginosa. J. Bacteriol. 96:1732–1741. 22. Jones, M. E. 1980. Pyrimidine nucleotide biosynthesis in animals: genes, enzymes and regulation of UMP biosynthesis. Annu. Rev. Biochem. 49:253– 279. 23. Kedzie, K. M. 1987. Ph.D. thesis. Texas A & M University, College Station. 24. Kumar, A. P., D. M. Brichta, M. J. Schurr, and G. A. O’Donovan. 1994. Deletion of a 34 amino acid N-terminal region of the aspartate transcarbamoylase in Pseudomonas putida abolishes regulatory nucleotide effector responses, abstr. K-36, p. 282. In Abstracts of the 94th General Meeting of the American Society for Microbiology. American Society for Microbiology, Washington, D.C. 25. Lerner, C. G., and R. L. Switzer. 1986. Cloning and structure of the Bacillus subtilis aspartate transcarbamoylase gene (pyrB). J. Biol. Chem. 261:11156– 11165. 26. Maniatis, T., E. F. Fritsch, and J. Sambrook. 1982. Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 27. Michaels, G., R. A. Kelln, and F. E. Nargang. 1987. Cloning, nucleotide sequence and expression of the pyrBI operon of Salmonella typhimurium LT2. Eur. J. Biochem. 166:55–61. 28. Miller, J. H. 1972. Experiments in molecular genetics, p. 432. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 29. Nagy, M., M. Le Gouart, S. Potier, J.-L. Souciet, and G. Herve´. 1989. The primary structure of the aspartate transcarbamoylase region of the URA2 gene product in Saccharomyces cerevisiae. Features involved in activities and in nuclear localization. J. Biol. Chem. 264:8366–8374. 30. Neuhard, J., R. A. Kelln, and E. Stauning. 1986. Cloning and structural characterization of the Salmonella typhimurium pyrC gene encoding dihydroorotase. Eur. J. Biochem. 157:335–342. 31. Ornston, L. N., and R. Y. Stanier. 1966. The conversion of catechol and protocatechuate to b-ketoadipate by Pseudomonas putida. J. Biol. Chem. 16:3776–3786.

1759

32. Pauza, C. D., M. J. Karels, M. Navre, and H. K. Schachman. 1982. Genes encoding Escherichia coli aspartate transcarbamoylase: the pyrB-pyrI operon. Proc. Natl. Acad. Sci. USA 79:4020–4024. 33. Prescott, L. M., and M. E. Jones. 1969. Modified methods for the determination of carbamyl aspartate. Anal. Biochem. 32:408–419. 34. Quinn, C. L., B. T. Stephenson, and R. L. Switzer. 1991. Functional organization and nucleotide sequence of the Bacillus subtilis pyrimidine biosynthetic operon. J. Biol. Chem. 266:9113–9127. 35. Roof, W. D., K. F. Foltermann, and J. R. Wild. 1982. The organization and regulation of the pyrBI operon in E. coli includes a rho-independent attenuator sequence. Mol. Gen. Genet. 187:391–400. 36. Sanger, F., S. Nicklen, and A. R. Coulson. 1977. DNA sequencing with chain-terminating inhibitors. Proc. Natl. Acad. Sci. USA 74:5463–5467. 37. Schurr, M. J. 1992. Ph.D. thesis. University of North Texas, Denton. 38. Shepherdson, M., and D. McPhail. 1993. Purification of aspartate transcarbamoylase from Pseudomonas syringae. FEMS Microbiol. Lett. 114:201–206. 39. Shigesada, K., G. R. Stark, J. A. Maley, L. A. Niswander, and J. N. Davidson. 1985. Construction of a cDNA to the hamster CAD gene and its application toward defining the domain for aspartate transcarbamoylase. Mol. Cell. Biol. 5:1735–1742. 40. Simmer, J. P., R. E. Kelly, A. G. Rinker, Jr., B. H. Zimmermann, J. L. Scully, H. Kim, and D. R. Evans. 1990. Mammalian dihydroorotase nucleotide sequence, peptide sequences, and evolution of the dihydroorotase domain of the multifunctional protein CAD. Proc. Natl. Acad. Sci. USA 87:174–178. 41. Simmer, J. P., R. E. Kelly, J. L. Scully, D. R. Grayson, A. G. J. Rinker, S. T. Bergh, and D. R. Evans. 1989. Mammalian aspartate transcarbamoylase (ATCase): sequence of the ATCase domain and interdomain linker in the CAD multifunctional polypeptide and properties of the isolated domain. Proc. Natl. Acad. Sci. USA 86:4382–4386. 42. Souciet, J. L., M. Nagy, M. Le Gouar, F. Lacroute, and S. Potier. 1989. Organization of the yeast URA2 gene: identification of a defective dihydroorotase-like domain in the multifunctional carbamoylphosphate synthetaseaspartate transcarbamoylase complex. Gene 79:59–70. 43. Switzer, R. L., and C. L. Quinn. 1993. De novo pyrimidine nucleotide synthesis, p. 343–358. In A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis and other gram-positive bacteria. American Society for Microbiology, Washington, D.C. 44. Tabor, S. 1990. Expression using the T7 RNA polymerase promoter system, p. 16.2.1–16.2.8. In F. M. Ausubel, R. Brent, R. E. Kingston, D. D. Moore, J. D. Seidman, J. A. Smith, and K. Struhl (ed.), Current protocols in molecular biology. Wiley Interscience, New York. 45. Tabor, S., and C. C. Richardson. 1985. A bacteriophage T7 RNA polymerase/promoter system for controlled exclusive expression of specific genes. Proc. Natl. Acad. Sci. USA 82:1074–1078. 46. Vickrey, J. F. 1993. Ph.D. thesis. University of North Texas, Denton. 47. Weber, K. 1968. New structural model of E. coli aspartate transcarbamylase and the amino-acid sequence of the regulatory polypeptide chain. Nature (London) 218:1116–1119. 48. Wild, J. R. 1994. Personal communication. 49. Wild, J. R., and M. E. Wales. 1990. Molecular evolution and genetic engineering of protein domains involving aspartate transcarbamoylase. Annu. Rev. Microbiol. 44:193–218. 50. Wild, J. R., K. F. Foltermann, and G. A. O’Donovan. 1980. Regulatory divergence of aspartate transcarbamoylases within the Enterobacteriaceae. Arch. Biochem. Biophys. 201:506–517. 51. Wilson, H. R., P. T. Chan, and C. L. J. Turnbough. 1987. Nucleotide sequence and expression of the pyrC gene of Escherichia coli K-12. J. Bacteriol. 169:3051–3058. 52. Zhou, C., Y. Yang, and A. Y. Jong. 1990. Mini-prep in ten minutes. BioTechniques 8:172–173.