ABSTRACT
Protocatechuate 3,4-dioxygenase is a member of a family of bacterial enzymes that cleave the aromatic rings of their substrates between two adjacent hydroxyl groups, a key reaction in microbial metabolism of varied environmental chemicals. In an appropriate genetic background, it is possible to select for Acinetobacterstrains containing spontaneous mutations blocking expression ofpcaH or -G, genes encoding the α and β subunits of protocatechuate 3,4-dioxygenase. The crystal structure of the Acinetobacter oxygenase has been determined, and this knowledge affords us the opportunity to understand how mutations alter function in the enzyme. An earlier investigation had shown that a large fraction of spontaneous mutations inactivatingAcinetobacter protocatechuate oxygenase are either insertions or large deletions. Therefore, the prior procedure of mutant selection was modified to isolate Acinetobacter strains in which mutations within pcaH or -G cause a heat-sensitive phenotype. These mutations affected residues distributed throughout the linear amino acid sequences of PcaH and PcaG and impaired the dioxygenase to various degrees. Four of 16 mutants had insertions or deletions in the enzyme ranging in size from 1 to 10 amino acid residues, highlighting areas of the protein where large structural changes can be tolerated. To further understand how protein structure influences function, we isolated strains in which the phenotypes of three different deletion mutations in pcaH or -G were suppressed either by a spontaneous mutation or by a PCR-generated random mutation introduced into theAcinetobacter chromosome by natural transformation. The latter procedure was also used to identify a single amino acid substitution in PcaG that conferred activity towards catechol sufficient for growth with benzoate in a strain in which catechol 1,2-dioxygenase was inactivated.
Oxygenases are enzymes that split a molecule of oxygen and introduce one or both of the oxygen atoms into their substrates (18). The enzymes have long been the topic of study because of their ability both to modify a wide variety of stable substrates and to control highly reactive and potentially dangerous oxygen species. These studies have demonstrated the important roles of oxygenases in diverse organisms. In humans, for instance, the aromatic compound aspirin inhibits an oxygenase involved in the synthesis of steroids during inflammation (30). In plants, an enzyme that appears to regulate programmed cell death has significant sequence identity to bacterial oxygenases involved in aromatic catabolism (15). The enzyme protocatechuate 3,4-dioxygenase is a member of a family of bacterial oxygenases (18, 29) that use iron as a cofactor to cleave the aromatic rings of their substrates between two adjacent hydroxyl groups, a key step in mineralization of plant products (5). The crystal structure of protocatechuate 3,4-dioxygenase from a strain ofPseudomonas putida (originally classified asPseudomonas aeruginosa) has been determined (36, 37), and more recently, the structure of the enzyme fromAcinetobacter sp. strain ADP1 was solved (56, 57).
The product of protocatechuate oxygenase is carboxymuconate, andAcinetobacter strains blocked in its metabolism fail to grow with any substrate when they are exposed to either protocatechuate or its metabolic precursors. Thus, it is possible to select for strains in which spontaneous secondary mutations block earlier steps in either transport (6) or catabolism of aromatic compounds (Fig.1). Strains derived from such selections contained spontaneous mutations affecting residues important for structure or function in various oxygenases:p-hydroxybenzoate hydroxylase, encoded by pobA(8, 21), protocatechuate 3,4-dioxygenase, encoded bypcaH and pcaG (13), and vanillate demethylase, encoded by vanA and vanB(52). For genetic analysis of two regulatory proteins, PobR (governing expression of pobA) and PcaU (governing expression of pcaHG), PCR-generated mutations were introduced into the Acinetobacter chromosome by natural transformation, producing both loss-of-function and gain-of-function mutations (26, 27).
Positive selection of strains blocked in protocatechuate catabolism. Strain ADP500 with the engineered ΔpcaBDK1mutation cannot grow with succinate in the presence of protocatechuate because of the toxic accumulation of β-carboxy-cis,cis-muconate. ADP500 derivatives with spontaneous mutations in pcaH and -G, genes encoding the two subunits of protocatechuate 3,4-dioxygenase, can therefore be selected based on their resistance to protocatechuate. Analogous to the conversion of protocatechuate to carboxymuconate by protocatechuate 3,4-dioxygenase (PcaHG), catechol 1,2-dioxygenase (CatA) converts catechol to muconate in the benzoate branch of the β-ketoadipate pathway.
The present investigation is a genetic analysis ofAcinetobacter protocatechuate 3,4-dioxygenase complementing the recently determined crystal structure of the enzyme (56, 57). A refinement in the positive-selection protocol enabled rapid identification of strains with a heat-sensitive mutation inpcaH or -G, and subsequently, strains with spontaneous or PCR-generated second-site suppressors of some of these mutations were selected. Transformation-facilitated PCR mutagenesis was also used to select one strain in which a single amino acid substitution in PcaG allowed protocatechuate 3,4-dioxygenase to functionally replace catechol 1,2-dioxygenase (Fig. 1).
MATERIALS AND METHODS
Strains and culture conditions. Acinetobacter sp. strain ADP1, originally designated Acinetobacter calcoaceticus BD413 (24), was routinely grown with 10 mM succinate in a mineral medium (13). Unless otherwise indicated, cells were grown at 37°C and the mineral medium was supplemented with 5 mM p-hydroxybenzoate, 5 mM quinate, 3 mM protocatechuate, or 2.5 mM benzoate. Because of the instability of protocatechuate, we used only fresh plates with this carbon source, made with stock solutions (pH 7.0) that had been stored frozen until use. Escherichia coli strains (50), namely, DH5α (purchased as competent cells from Gibco BRL) and TG2 (gift of M. Biggin and T. Williams), were grown with Luria-Bertani medium. Ampicillin was added at 100 μg/ml to select for E. colicells with plasmids derived from pUC18 or pUC19 (59), and tetracycline was added at 12.5 μg/ml to select for E. colior Acinetobacter cells with plasmids derived from pRK415 (25).
DNA manipulations.Crude cell lysates ofAcinetobacter strains for use in transformation reactions were prepared by incubation of pelleted cells (from a 5-ml culture) in 0.5 ml of saline-citrate buffer with 0.05% sodium dodecyl sulfate at 60°C for 1 h (24). Plasmids were isolated from 5-mlE. coli cultures with a Wizard miniprep kit (Promega) and from 5-ml Acinetobacter cultures by the alkaline lysis miniprep procedure (50) with two extraction steps, the first with phenol-chloroform and the second with chloroform. To isolate template DNA for PCR, pelleted Acinetobacter cells from a 5-ml culture were washed and treated with InstaGene Matrix as recommended by the supplier (Bio-Rad); 5 μl of the resulting solution was then used in 50-μl PCR mixtures including 0.5 U of Taqpolymerase (Boehringer Mannheim), 10 pmol of each primer, and 10 nmol of each deoxynucleoside triphosphate. Standard PCR conditions were used: 30 cycles of denaturing at 94°C for 45 s, annealing at 56°C for 45 s, and elongation at 72°C for 1 min 30 s. PCR primers were synthesized by the Keck Biotechnology Resource Lab (Yale University).
Selection and characterization of strains with a spontaneous heat-sensitive mutation in pcaH or -G.Strain ADP500 (13, 20) contains the engineered ΔcatD101::Kmr and ΔpcaBDK1 mutations. Single colonies of succinate-grown ADP500 were transferred to patches on freshly prepared plates with 10 mM succinate and 3 mM protocatechuate. Cells with spontaneous secondary mutations blocking protocatechuate catabolism do not accumulate toxic levels of β-carboxy cis,cis-muconate and are able to grow (Fig. 1). To prevent analysis of siblings, only one mutant derivative was picked per single colony of ADP500.
After incubation for 2 days at 37°C, colonies of ADP500 derivatives with secondary mutations were transferred to each of three plates: one plate contained the original selective medium and two plates contained 10 mM succinate and 5 mM p-hydroxybenzoate. The latter two plates were incubated for 2 days, one at 37°C and one at 22°C. Strains with a conditional resistance to p-hydroxybenzoate (with significantly better growth at 37°C than at 22°C) were picked from the plate with protocatechuate and purified by two further rounds of growth on plates in the presence of protocatechuate. The ΔpcaBDK1 deletion was restored to wild type by transformation with plasmid pZR3 (13, 20) under nonselective conditions, followed by selection for growth with benzoate. Expression of PcaD in recombinants is sufficient for growth with benzoate and complements the engineered ΔcatD101::Kmr mutation inactivating the isofunctional enzyme in the benzoate branch of the β-ketoadipate pathway (13, 20).
Each of the resulting strains (ADP1102 to -1117) contained a spontaneous mutation blocking protocatechuate catabolism. Single colonies of succinate-grown mutant cells were then tested for growth, at 37 and 22°C, on plates with p-hydroxybenzoate as the sole carbon and energy source. The heat-sensitive mutation in eitherpcaH or -G was localized by marker rescue as described previously (13) except that quinate was used as the selective growth substrate (Fig. 1). DNA transformations for marker rescue or linkage experiments were performed by growing recipientAcinetobacter strains with 10 mM succinate overnight in 5-ml cultures. After addition of 10 μl of 1 M succinate, these cultures were then incubated (to induce competence) at 37°C for 30 min in a gyratory shaker before being spread onto selective plates onto which donor DNA was then spotted.
Recovery of Acinetobacter chromosomal DNA by gap repair.DNA fragments containing mutant pcaH and -G genes were recovered from the chromosome by gap repair with plasmid pZR1007 as previously described (2, 16). To reduce the chance that the recovered DNA would reintegrate into the chromosome, the duration of steps involving Acinetobactercells was minimized. The 2.4-kb HindIII restriction fragment containing pcaK′CHGquiB′ (13, 19) from each pZR1007 derivative was purified by preparative gel electrophoresis and ligated with HindIII-digested pUC18, and the resulting plasmids were amplified in E. coli DH5α. Cells with plasmids in which the insert was oriented so as to be expressed from the vector promoter formed distinctively small colonies on Luria-Bertani medium plates with ampicillin. Minipreps of such cells, however, gave low plasmid yields, so the plasmids were transferred toE. coli TG2 cells (in which lacIqdecreases the potentially toxic uninduced expression of the cloned genes).
Selection and characterization of a strain in which protocatechuate 3,4-dioxygenase functionally replaced catechol 1,2-dioxygenase.To select strains in which a gain-of-function mutation in protocatechuate 3,4-dioxygenase conferred activity towards catechol, pcaH and -G were first inserted into the gene for catechol 1,2-dioxygenase (catA) so that their expression would be regulated by benzoate (4). Plasmid pIB1344 (35), which contains catA, was cut withSalI and recircularized, generating pZR7559 with a uniqueSphI site within catA. A DNA fragment containingpcaC′HG (in which the Ω element from pHP45Ω [44] had been inserted into the BamHI site in pcaH) was amplified with Pfu polymerase (Stratagene) with plasmid pZR200Ω (gift of M. Stein) as the template and primers HG1 and HG6 (see below). The PCR DNA andSphI-digested pZR7559 were blunted with the DNA polymerase Klenow fragment and ligated together, and transformants of E. coli DH5α that were resistant to streptomycin, spectinomycin, and ampicillin and contained plasmids in which pcaHG and the disrupted catA were in the same orientation were selected.
The insert from one such plasmid was introduced into the chromosome ofAcinetobacter strain ISA25 (ΔcatBCIJF) by selecting for transformants in which inactivation of catAprevented the toxic accumulation of muconate generated during growth with succinate in the presence of benzoate (12, 58). PCR amplification across catA in one SmrSpcr Amps transformant with primers catA1 and catA2 (see below) and an extension time of 1 min 30 s generated a fragment of 3.4 kb, consistent with the absence of the 2-kb Ω element within pcaH. This PCR DNA was used, as before, to generate an ISA25 transformant strain resistant to benzoate, and thecatBCIJF deletion in one such strain was corrected by transformation with pPAN4 (35, 53) and selection for growth with muconate, generating ADP7559. Slow growth with muconate of ADP7559 suggested the existence of a spontaneous mutation affecting muconate transport (58), as has been observed for ISA25 derivatives in two separate studies (3, 58).
PCR mutagenesis of ADP7559 generated ADP7615. The pcaG7615gain-of-function mutation in the latter strain was recovered from the chromosome by PCR amplification with the primers catAK (5′-GGGGGTACCCTGATTCTACATGGCACG-3′) and catAR (5′-GGGGGAATTCATCGGTAATAATACTACGGCG-3′) whose 5′ ends include KpnI and EcoRI restriction sites (underlined), respectively. After digestion with KpnI andEcoRI, this PCR fragment was ligated with KpnI- and EcoRI-digested pUC19, generating pZR7615 in whichpcaH and -G are expressed from the plasmid promoter. pZR7615 was transferred from E. coli DH5α cells to TG2 cells, and the sequences of the cloned genes containingpcaG7615 were reconfirmed.
Transformation-facilitated PCR mutagenesis.For mutagenesis (26, 27), standard PCR was performed as described above except that the number of cycles was increased to 35. To generate DNA with suppressors of a heat-sensitive pcaH or -Gdeletion, primers HG1 (5′-CTACATTGTTCACTTTATGCAGGC-3′) and HG6 (5′-GATATACGGCCCGTTCCATAGTC-3′) were used, amplifying a 1.5-kb PCR fragment. To mutagenize the transposed pcaH and -G genes in ADP7559, primers catA1 (5′-GGTATAGAAACGACTATCG-3′) and catA2 (5′-CAAGTGTATGTCGTAACGC-3′) were used, amplifying a 3.4-kb fragment. PCR-amplified fragments, generated with template DNA from the planned recipient strain, were used directly in transformation reaction mixtures as follows. Two hundred microliters of a 5-ml culture of the recipient strain, grown overnight with 10 mM succinate, was transferred to a fresh 5-ml culture, and after growth in a gyratory shaker for 2 h (to induce competence), 500 μl of the culture was transferred to Falcon 15-ml polypropylene tubes together with 20 μl of the PCR mixture. After overnight shaking incubation, transformation reaction mixtures were spread onto selective plates.
DNA sequence analysis.To isolate template DNA for sequencing, PCR mixtures prepared under standard conditions were purified with 8 μl of GeneClean Glassmilk according to the recommendations of the supplier (Bio 101, Inc.) and resuspended in 25 μl of water, and 8 μl was used for ABI PRISM Dye Terminator Cycle Sequencing with AmpliTaq DNA polymerase FS (Perkin-Elmer). Cycle sequence reaction mixtures were processed as previously described (26).
RESULTS
Characterization of strains with a spontaneous heat-sensitive mutation in pcaH or -G.By positive selection for derivatives of Acinetobacter strain ADP500 that have a spontaneous mutation blocking the β-ketoadipate pathway, 94 strains (13) that had a mutation affecting protocatechuate 3,4-dioxygenase, the enzyme which cleaves the protocatechuate aromatic ring generating carboxymuconate (Fig. 1), were characterized. Only 11 of the 94 strains, however, had a missense mutation in pcaHor -G, a gene encoding the β or α subunit of the dioxygenase, respectively. Nearly one-quarter of the mutants had a deletion extending into DNA outside of these two genes (13). Six of the missense mutations caused a heat-sensitive phenotype, blocking growth with protocatechuate at 37°C but not at 22°C (13). It therefore seemed feasible to use this phenotype as a way to identify, early in the selection protocol, those ADP500 derivatives most likely to have a spontaneous missense mutation inpcaH or -G. This identification process would allow efforts to be focused on strains which could contribute to understanding how structure influences function in the dioxygenase enzyme and eliminate time spent identifying mutants with large deletions, frameshifts, or nonsense mutations.
To identify heat-sensitive pcaH or -G mutants in this study, ADP500 derivatives still containing the ΔpcaBDK1 deletion and able to grow in the presence of protocatechuate at 37°C were tested for resistance top-hydroxybenzoate during growth with succinate at both 37 and 22°C (Fig. 1). Screening was done withp-hydroxybenzoate because cells were found to grow poorly upon successive transfers onto plates containing protocatechuate, a compound that has inherent toxicity (41). Consistently, approximately 5% of the spontaneous mutants analyzed were resistant top-hydroxybenzoate at 37°C but not at 22°C. After correction of the ΔpcaBDK1 deletion in 17 such strains, the expected heat-sensitive phenotype was revealed: growth withp-hydroxybenzoate was completely blocked at 37°C but allowed at 22°C. The spontaneous mutation in each of these 17 strains, with one exception, was mapped by marker rescue topcaH and pcaG and sequenced. The one unique strain had a point mutation in the gene for PcaU, the transcriptional activator governing pca operon expression, and its characterization is part of a separate study (51).
The pcaH and -G mutations and their associated amino acid changes in the heat-sensitive mutants isolated in this study or previously (13) are listed in Table1. Figure 2shows the positions of the mutations in the primary sequence ofAcinetobacter PcaH and -G. The pcaG13 andpcaH1112 point mutations were each found in two independently isolated strains, and the pcaG13 andpcaG1114 mutations changed the same nucleotide but caused different amino acid substitutions (Table 1). Most of the remaining mutations affected residues distributed throughout the linear amino acid sequence of PcaH and -G (Fig. 2). Although all the mutants grew with p-hydroxybenzoate at 22°C but not at 37°C, protocatechuate 3,4-dioxygenase appeared to be impaired to various degrees: the most severely affected mutants (ADP6116 and ADP1116) grew with p-hydroxybenzoate extremely slowly even at 22°C.
Heat-sensitive mutations in pcaHand -G
Spontaneous and PCR-generated mutations in protocatechuate 3,4-dioxygenase from Acinetobacter sp. strain ADP1. Protocatechuate 3,4-dioxygenase is an oligomer (29) of heterodimers composed of PcaG (the α subunit) and PcaH (the β subunit). The two subunits have significant amino acid identity, and it has been suggested that they are derived from duplication and divergence of the gene for the homodimer subunit of the ancestral enzyme (36, 37). Acinetobacterprotocatechuate 3,4-dioxygenase subunits (GenBank accession no. L05770 ) are presented so that homologous amino acid sequences occupy corresponding positions in PcaG (top) and PcaH (bottom). In order to achieve this alignment, gaps indicated by dashes were introduced into the sequences. With the exception of pcaG7615, G425A, nucleotide sequence changes causing the amino acid alterations shown here are presented in Tables 1 and 2. Alleles with designations smaller than 1000 were identified in a previous investigation (13). To facilitate comparison with structural data from previous studies (37, 56), the same numbering system has been used for the primary structure with PcaG amino acids numbered 1 to 200 and PcaH amino acids numbered 300 to 540 (the five Acinetobacter PcaG residues that align with a gap between residue 88 and 89 in P. putida PcaG are designated 88a to -e). Amino acid substitutions are indicated by arrows pointing to the mutant amino acid. Duplicated residues are underlined, and the mutant repetitions are shown above the primary sequence. Shaded boxes superimposed on the primary sequence indicate residues missing in deletion mutations. Amino acids in shaded boxes separated by arrows from the primary sequence indicate suppressor mutations, and the borders of the boxes (solid, dotted, or not bordered) match those of the shaded boxes indicating the mutations that are suppressed. The double-dotted border indicates a mutation that confers the ability to grow at 37°C to a strain containing both ΔpcaG1102 and pcaG7554. The R133H substitution caused by pcaG7615 and conferring activity towards catechol is indicated by a box enclosing the structure of the compound. The glycyl residue at position 60 in PcaG appears to be a natural variant and is found in all the mutants described in this study (including ADP7615), whereas the wild-type strain ADP1 has serine (indicated in parentheses in the figure) at the corresponding position.
The majority of strains had a missense mutation, but unexpectedly, protocatechuate 3,4-dioxygenase was more severely altered in four newly isolated strains. Deletions in pcaH1116 andpcaG1102 caused deletions of 1 and 10 amino acid residues, respectively, and mutations in pcaH1111 andpcaG1103 caused insertions of 1 and 2 amino acids, respectively (Table 1; Fig. 2). Both deletions may have arisen by recombination between directly repeated DNA sequences, resulting in loss of one of the repeats, TAT in the ΔpcaH1116 strain and GATAC in the ΔpcaG1102 strain. The 3-bppcaH1116 deletion appears to lie within a local region of DNA instability, given its proximity to the previously described overlapping null mutations pcaH21 and pcaH13, duplications of 3 and 7 bp, respectively, (13). The cause at the DNA level of the duplication of TTC in pcaH1111 and AAATGC in pcaG1103 is likely to have been mediated by slippage at the tandem DNA repeats immediately flanking each of these mutations, GGTGGT and GAAGCAGAAGCA, respectively. As expected, phenotypic revertants were readily obtained from strains carrying either of these duplications.
Suppression by intragenic and extragenic point mutations.Strains in which a pcaH or -G deletion did not cause a null phenotype were particularly attractive candidates to use to identify second-site suppressor mutations since the encoded enzyme was still partly functional and a background of direct reversion would not be a concern. Suppression was first tried with a strain from a previous study, ADP6338, containing the ΔpcaH7 leaky 4-amino-acid deletion which causes slow growth withp-hydroxybenzoate at both 37 and 22°C (13) (Fig. 2). Liquid cultures of ADP6338 grown with 10 mM succinate and 5 mM p-hydroxybenzoate were serially transferred, and after the third transfer, one culture failed to develop the purple color indicative of protocatechuate accumulation due to deletion ofpcaH7. Partial suppression of the ΔpcaH7 leaky phenotype in a purified isolate from this culture was confirmed by comparing the growth on a plate with p-hydroxybenzoate of the derived strain, designated ADP7552, with that of ADP6338.
In addition, occasionally a liquid culture shaking at 37°C would not grow when it was provided with 5 mM p-hydroxybenzoate as the carbon source and inoculated with a single colony of succinate-grown ADP6338. In one case, the culture suddenly became turbid after 18 days of incubation. A purified isolate, designated ADP7553, grew slightly faster than the parental strain (but not as well as ADP7552) on a plate with p-hydroxybenzoate at 37°C. The cause of the varied behavior of ADP6338 is unknown but may reflect different physiological states conferring upon cells a range of sensitivities to the sudden intracellular accumulation of protocatechuate generated during growth with p-hydroxybenzoate.
Evidence that a mutation genetically linked to the pcaH7deletion was the cause of suppression in both ADP6338 derivatives came from results of a DNA transformation experiment using a recipient strain with the 440-bp pcaCH1 deletion (13) encompassing the pcaH7 deletion locus: recombinants that grew at 37°C with p-hydroxybenzoate like ADP7552 or ADP7553 were readily obtained with donor DNA from the corresponding strain either in crude cell lysate or as a 2.4-kbHindIII restriction fragment containingpcaCHG recovered from the chromosome by gap repair (2, 16). Confirming this linkage, sequencing of pcaH and -G in each strain revealed that protocatechuate 3,4-dioxygenase contained, in addition to the deletion of PcaH residues 319 to 322 due to the pcaH7 deletion, a G13S change in PcaG in ADP7552 and a P317L change in PcaH in ADP7553 (Fig. 2; Table2).
Suppressor mutations in pcaHand -G
During this investigation but as part of a separate study, PCR mutagenesis was combined with natural transformation to obtain loss-of-function as well as gain-of-function mutations in the PobR regulatory protein (26, 27). To show in that study that the technique was broadly applicable for genetic analysis of chromosomal genes in Acinetobacter, we identified PCR-generated mutations that suppressed the heat-sensitive pcaG1102deletion, causing loss of PcaG amino acid residues 76 to 85 (26) (Fig. 2). Each suppressor mutation caused an amino acid substitution near the deletion in the PcaG linear amino acid sequence and restored the ability to grow with p-hydroxybenzoate at 30°C but not at 37°C (26) (Fig. 2; Table 2). However, during a later experiment using cells with the pcaG7554suppressor, a single colony that could grow withp-hydroxybenzoate at 37°C appeared, suggesting that further spontaneous suppression of the heat-sensitive phenotype had been selected. Subsequent sequencing of pcaH and -G in this strain revealed an additional point mutation causing a D74N substitution, again near the original mutation in the PcaG primary sequence (Fig. 2; Table 2).
Of the heat-sensitive pcaH or -G mutations in Table 1, deletion of pcaH1116 caused one of the strongest phenotypes: strains with this mutation grew withp-hydroxybenzoate extremely slowly even at 22°C. Nevertheless, PCR mutagenesis also successfully suppressed deletion ofpcaH1116, generating a strain in which an L465V substitution in PcaH in combination with the deletion of Y437 caused by thepcaH1116 deletion restored growth withp-hydroxybenzoate even at 37°C (Fig. 2; Table 2). PCR amplification across both mutations in the suppressed strain created DNA that gave rise to over 1,000 times the number of transformants in the initial selection, consistent with the sequenced second-site mutation being the cause of suppression.
An alternate form of suppression.PCR-generated suppressor deletions of pcaG1102 were found in colonies that appeared on a plate with p-hydroxybenzoate after overnight incubation at 30°C. At this time there was no growth on control plates of ADP1102 cells that had not been transformed with PCR DNA. However, after several days’ further incubation, the control plates usually contained several colonies with exceptional properties. These strains maintained the ability to grow at 30°C withp-hydroxybenzoate even after the selective pressure was relieved during successive rounds of growth with succinate, suggesting that the strains had acquired a heritable suppressor mutation. Yet, DNA sequencing of pcaH and -G in two such strains revealed only the original ΔpcaG1102 mutation and DNA in crude cell lysates of these strains could not transform the new phenotype into the parental strain.
It has been noted that during suppression analysis of bacteria, often the most readily isolated cells are those in which amplification on the chromosome of the partly dysfunctional gene generates sufficient mutant protein activity to allow growth under the selective conditions (49). Such a phenomenon may account for the properties of the strains in which the pcaG1102 deletion appeared to have been spontaneously suppressed. DNA amplification has also been shown to be an adaptive response of wild-type bacterial cells (33, 46-49). In Acinetobacter, the ability to coamplify (49) all the genes necessary for catabolism of protocatechuate, quinate, and p-hydroxybenzoate to citric acid cycle intermediates is a selective benefit that may have acted continuously over evolutionary time to favor the current supraoperonic clustering (2, 14, 22) of these genes.
A single amino acid substitution in PcaG (R133H) confers catechol 1,2-dioxygenase activity.Isolation of strains with a PCR-generated suppressor mutation as described above suggested that the technique might be a powerful method of isolating gain-of-functionpcaH or -G mutations. In the benzoate branch of the β-ketoadipate pathway, catechol 1,2-dioxygenase occupies a position equivalent to that of protocatechuate 3,4-dioxygenase, and the respective substrates of these two enzymes differ by only a carboxyl group (Fig. 1). Comparison of the amino acid sequences of the two oxygenases indicates common ancestry (19, 34). An intriguing question, therefore, is to what extent the two enzymes have specialized to perform their respective tasks since their divergence from a common ancestor (55).
To address this problem, we constructed strain ADP7559, in which a copy of pcaH and -G was inserted near the middle ofcatA (34), the gene encoding the subunits of the catechol 1,2-dioxygenase homodimer (42). This insertion prevented growth with benzoate but presumably allowed benzoate to induce expression of the transposed genes (4). PCR amplification across pcaH and -G in this strain generated DNA which gave rise to transformants of ADP7559 that could grow with benzoate, although the brown staining produced during growth of the transformants indicated some accumulation of catechol. No background of spontaneous mutants was detected. Sequencing of the transposed pcaH and -G in 13 transformants, each independently generated by different PCRs, revealed in every case the same G425A point mutation (pcaG7615) causing an R133H substitution in PcaG (Fig. 2). The pcaH and -Ggenes from within catA in one of the transformants (ADP7615) were cloned by PCR into plasmid pZR7615, and after confirmation of the DNA sequence, transformation with this DNA was shown to confer the ability to grow with benzoate upon the parental strain.
DISCUSSION
Genetic analysis of protocatechuate 3,4-dioxygenase structure and function.In a previous investigation using a parental strain with the pcaBDK1 deletion, derivatives in which a conditional mutation in pcaH or -G provided resistance to the toxic intracellular accumulation of carboxymuconate produced from protocatechuate in the medium were isolated (13) (Fig. 1). These mutations were identified after pcaBDK DNA was restored to wild type. In contrast, this study describes a protocol for the identification of strains with a conditional pcaH or -G mutation before any genetic manipulation, thereby facilitating the generation of a large collection of such strains useful for an analysis of the effects on protocatechuate 3,4-dioxygenase function of relatively subtle alterations in the structure of the protein.
The pcaH and -G mutations isolated in this study and previously (13) have a range of heat sensitivity phenotypes. Unexpectedly, one-quarter of the mutants produced in this study had small deletions or insertions in pcaH or -G. The largest deletion (ΔpcaG1102) removed 10 amino acids from a surface loop in the α subunit that contacts the β subunit of the enzyme (37, 57). The sequences of this loop are highly varied among the organisms for which the sequences are known, with the Acinetobacter strain ADP1 andRhodococcus opacus 1CP enzymes having five additional residues and the Burkholderia cepacia and Pseudomonas marginata enzymes having two additional residues relative to the number of residues in the corresponding loop in the P. putida enzyme (10, 11, 19, 43, 60). This finding illustrates how the positive-selection protocol used here reveals not only general structural determinants of protein stability (1, 31, 32, 40) but also regions of the protein where large alterations can be tolerated.
Thermodynamic experiments have shown that the free energy of folding of proteins is relatively small, approximately 5 to 15 kcal/mol (45). This small value favoring protein folding is due to a delicate balance between two large numbers, the free energy of the noncovalent interactions in the folded state and that in the unfolded state. Interactions between hydrophobic residues are the principal thermodynamic force in the stabilization of the folded state, while solvation energies are the principal destabilizing force. Computational analyses of models of protein folding (7) have shown that mutations can lead to large changes in the denaturation temperature of a protein with little change to the overall structure of the protein. Thus, the protein can be active at reduced temperatures but inactive at higher temperatures.
In a method similar to the plating selection used to obtain mutants ofpcaH and -G, bacteriophage T4 lysozyme mutants in which the enzyme was phenotypically inactive at 42°C were obtained (17, 23). When purified, these enzymes were found to have reductions in thermal stability that could be rationalized upon examination of the structure. The recently determined crystal structure of protocatechuate 3,4-dioxygenase from Acinetobacter sp. strain ADP1 (57) similarly facilitates analysis of the mutations in this study for their possible effects. Figure3 shows the positions of all temperature-sensitive, deletion, and suppressor mutations superimposed on a drawing of the Cα trace of the Acinetobacter enzyme. Table 3 describes the structural consequences of the mutations. Typically, the mutations create van der Waals clashes, cavities in hydrophobic regions, or isolated buried charges destabilizing the native structure of an individual subunit, of an intersubunit interface, or of the interface between molecules in the dodecameric aggregate. For three of the mutations, residues in the active site are involved so that the reduction in activity may not arise from a lack of stability. T12I abolishes hydrogen bonds which stabilize the main chain around Pro15. Since the side chain of Pro15 forms part of the narrow waist of the substrate binding site, changes in its position may allow protocatechuate to bind to the iron in less productive orientations. W400S eliminates a residue that stacks against His462, an iron ligand. The substitution also removes the Nɛ that may form a hydrogen bond with molecular oxygen during the reaction cycle. P458L may move the end of Arg457, which has been proposed to stabilize the development of a negative charge on C-4 of protocatechuate, allowing an electrophilic attack by molecular oxygen (38).
Stereoview Cα trace of protocatechuate 3,4-dioxygenase from Acinetobacter sp. strain ADP1 showing the positions of spontaneous and PCR-generated mutations. Each protomer of protocatechuate 3,4-dioxygenase is composed of an α subunit (dashed line), a β subunit (solid line) and one nonheme iron (black sphere). Each mutation is labeled with a sphere and a letter which corresponds to a letter in Table 3. Point mutations and insertions are in white spheres. These mutations are scattered throughout the structure but usually lead to disruption of interfaces between secondary or tertiary structural elements. The deletion mutations X, Y, and Z are depicted in green, blue, and red spheres, respectively. Rescue mutations are shown in lighter shades of the corresponding color.
Mutations of Acinetobacter protocatechuate 3,4-dioxygenase and structural consequences
The three pcaH or -G deletion mutations for which second-site suppressors were identified in this study (Fig. 2 and 3; Table 2) appear to affect protocatechuate 3,4-dioxygenase in distinct ways: the pcaH7 deletion causes the loss of 4 amino acid residues in a loop containing Tyr324, which forms a hydrogen bond with the substrate in the native enzyme (38, 57); thepcaH1116 deletion causes the loss of one residue at the amino terminus of a β strand in the flattened β barrel of the β subunit; and deletion of pcaG1102 causes the loss of 10 residues in a loop that forms a portion of the interface between the α and β subunits (37, 57). The fact that spontaneous or PCR-generated suppressor mutations were readily identified for each of these deletions, with the pcaH7 deletion being suppressed by either an intragenic or an extragenic mutation (Fig. 2; Table 2), demonstrates the usefulness of combining positive selection and natural transformation for genetic analysis of protocatechuate 3,4-dioxygenase in Acinetobacter strain ADP1. Furthermore, it was discovered, unfortunately in a separate investigation after this study, that the combination of a mutation inactivating the regulatory genepcaU with the leaky ΔpcaH7 structural gene mutation completely blocked growth with p-hydroxybenzoate or protocatechuate although either mutation alone did not (27). This double mutant was used to identify PCR-generated gain-of-function mutations in pobR which could confer PcaU activity (27). However, the same strategy should also greatly facilitate the identification of PCR-generated mutations inpcaH or -G that suppress the pcaH7deletion or other leaky mutations that subtly alter the enzyme, since strains with such suppressors would appear against a background of no growth. Further biochemical and structural analysis of these suppressor mutations might shed light on the long slow process by which proteins evolve over time by accumulating beneficial mutations in combinations of two or more.
It has long been noted that the two parallel branches of the β-ketoadipate pathway (Fig. 1), the p-hydroxybenzoate branch encoded by genes in the pca-qui-pob supraoperonic cluster (14, 22) and the benzoate branch encoded by theben-cat supraoperonic cluster (14, 22), represent an attractive system for studying how the demands for a set of chemical reactions in a single cell line were met twice over evolutionary time to create each of the two branches of the pathway (55). At the two extremes of a continuum, the coenzyme A transferases encoded bypcaIJ and catIJ are over 99% identical (28) whereas the lactonizing enzymes encoded bypcaB and catB, although they act on closely related substrates, come from distinct protein families and do not have significant sequence identity (28). Between these extremes, the two dioxygenase subunits encoded by pcaH and -G have 26 to 36% amino acid identity over 127 to 213 aligned residues with the homodimer subunit encoded by catA, suggesting that the two enzymes have a common ancestor. Nevertheless, it was unexpected that it would take one and only one amino acid substitution in protocatechuate 3,4-dioxygenase to confer activity towards catechol sufficient to allow growth with benzoate in a strain in which catA was inactivated.
The ease with which directed evolution may generate a protocatechuate 3,4-dioxygenase capable of functionally replacing catechol 1,2-dioxygenase is intriguing given that there appears to be one trait which consistently distinguishes the two bacterial dioxygenases: the oligomeric complexity of the enzymes. Protocatechuate 3,4-dioxygenase forms oligomers composed of different numbers of αβ-subunit heterodimers in different bacteria (29): 12 inAcinetobacter (56) and P. putida(37) and 6 in Brevibacterium fuscum(9). Catechol 1,2-dioxygenases are not known to oligomerize beyond homo- and heterodimers (18). Oligomerization has been suggested to enhance thermal stability (37), perhaps indicating a trade-off between stability and catalytic efficiency (54). The active site in Acinetobacterprotocatechuate 3,4-dioxygenase is at the αβ interface near a neighboring protomer related by the local threefold-symmetry axis (57). Near the corresponding symmetry axis in the P. putida enzyme there is a second site where the substrate and inhibitors can bind (38, 39), being connected to the active site by a solvent-filled channel. These observations fueled speculation that the oligomeric state of these intradiol dioxygenases plays an important role in function. The finding of the R133H gain-of-function mutation shows that the oligomeric state may play only a small role in substrate specificity and that this can be overcome by a single mutation.
The opening to the active site of Acinetobacterprotocatechuate 3,4-dioxygenase is at the bottom of a 15-Å-deep cavity surrounded by a preponderance of basic residues. Calculation of the electrostatic potential around P. putida protocatechuate 3,4-dioxygenase shows that a large sphere of positive electrostatic potential originates around the iron and extends all the way to the outer edge of the active-site cavity (39). It has been proposed that the enzyme uses this excess positive potential to attract negatively charged substrates like protocatechuate into the active site. Arg133 is one of the positively charged residues in the midsection of the active-site cavity. It makes an intrasubunit salt link to Asp65 and an interprotomer salt link to Glu162. Arg133 and other positively charged residues near the entrance to the active-site cavity are not conserved in the catechol 1,2-dioxygenases. Modeling of the R133H mutation indicates that a histidine at this position would not be able to form a direct interaction with Glu162 of the neighboring protomer. Since both residues are exposed, the free-energy penalty of disrupting this interaction should be on the order of 1 kcal/mol or less. Choosing a common rotamer of the histidine would place the ring of the side chain within hydrogen-bonding distance of Tyr324 and Thr326. In the structure of the oxygenase with bound protocatechuate (38, 57), Tyr324 interacts with the carboxyl of the substrate. If the histidine rotated into this area it might interact directly with catechol, facilitating more productive modes of binding during catalysis with catechol.
Yet to be determined is the extent to which the R133H mutation deprives protocatechuate 3,4-dioxygenase of its capacity to support growth with protocatechuate in the wild-type genetic and physiological context ofpcaH and -G. In short, did the gain-of-function mutation make the enzyme a generalist, capable of acting effectively upon both catechol and protocatechuate, or a shifted specialist, with effective activity changed from protocatechuate to catechol?
ACKNOWLEDGMENTS
This research was supported by grants DAAG55-98-1-0232 from the Army Research Office and MCB-9603980 from the National Science Foundation to L.N.O. and by grants from the National Institutes of Health (GM-46436) and from the Minnesota Supercomputer Institute to D.H.O. M.W.V. acknowledges an NIH predoctoral training grant (GM-08277).
FOOTNOTES
- Received 14 May 1999.
- Accepted 21 July 1999.
- Copyright © 1999 American Society for Microbiology
