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J. Biol. Chem., Vol. 282, Issue 7, 4681-4692, February 16, 2007

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Characterization of the 2-Hydroxy-acid Dehydrogenase McyI, Encoded within the Microcystin Biosynthesis Gene Cluster of Microcystis aeruginosa PCC7806^*

From the School of Biotechnology and Biomolecular Sciences, University of New South Wales, Sydney, New South Wales 2052, Australia

Received for publication, July 24, 2006 , and in revised form, October 30, 2006.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The cyanobacterium Microcystis aeruginosa is widely known for its production of the potent hepatotoxin microcystin. This cyclic heptapeptide is synthesized non-ribosomally by the thio-template function of a large modular enzyme complex encoded within the 55-kb microcystin synthetase gene (mcy) cluster. The mcy gene cluster also encodes several stand-alone enzymes, putatively involved in the tailoring and export of microcystin. This study describes the characterization of the 2-hydroxy-acid dehydrogenase McyI, putatively involved in the production of D-methyl aspartate at position 3 within the microcystin cyclic structure. A combination of bioinformatics, molecular, and biochemical techniques was used to elucidate the structure, function, regulation, and evolution of this unique enzyme. The recombinant McyI enzyme was overexpressed in Escherichia coli and enzymatically characterized. The hypothesized native activity of McyI, the interconversion of 3-methyl malate to 3-methyl oxalacetate, was demonstrated using an in vitro spectrophotometric assay. The enzyme was also able to reduce -ketoglutarate to 2-hydroxyglutarate and to catalyze the interconversion of malate and oxalacetate. Although NADP(H) was the preferred cofactor of the McyI-catalyzed reactions, NAD(H) could also be utilized, although rates of catalysis were significantly lower. The combined results of this study suggest that hepatotoxic cyanobacteria such as M. aeruginosa PCC7806 are capable of producing methyl aspartate via a novel glutamate mutase-independent pathway, in which McyI plays a pivotal role.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The hepatotoxic microcystins compose the largest and most structurally diverse group of cyanobacterial toxins. Over 65 isoforms of microcystin varying by degree of methylation, hydroxylation, epimerization, peptide sequence, and toxicity have been identified (1). Underlying the extraordinary heterogeneity present among the microcystins is their common cyclic structure and possession of several rare nonproteinogenic amino acid moieties. Collectively, the microcystins may be described as monocyclic heptapeptides containing both D- and L-amino acids plus N-methyldehydroalanine and a unique -amino acid side group, 3-amino-9-methoxy-2,6,8-trimethyl-10-phenyldeca-4,6-dienoic acid (Fig. 1) (2). Although various amino acid substitutions can occur within the microcystin cyclic structure, the most common toxin isoform, microcystin LR, contains lysine and arginine at positions 2 and 4, respectively (Fig. 1).

The microcystin biosynthesis (mcy) gene cluster was the first complex metabolite gene cluster to be fully sequenced from a cyanobacterium. In Microcystis aeruginosa PCC7806, the mcy gene cluster spans 55 kb and comprises 10 genes arranged in two divergently transcribed operons, mcyA–C and mcyD–J. The larger of the two operons, mcyD–J, encodes a modular polyketide synthase (McyD), two hybrid enzymes comprising non-ribosomal peptide synthetase (McyE) and polyketide synthase (McyG) modules, and enzymes putatively involved in the tailoring (McyI, McyJ, and McyF) and transport (McyH) of the toxin. The smaller operon, mcyA–C, encodes three non-ribosomal peptide synthetases (McyA–C) (3).

Although most open reading frames within the mcy gene cluster have been assigned a function based on homologous entries in the data bases, no obvious function could be assigned to the 1014-bp gene mcyI. Preliminary sequence analysis of the inferred primary peptide sequence of mcyI by Tillett et al. (3) revealed a 41% identity to the catalytic region of the serA-encoded D-3-phosphoglycerate dehydrogenase (PGDH²; EC 1.1.1.95 [EC] ) from Methanobacterium thermoautotrophicum (4). Although this archaeal PGDH homolog has not yet been characterized, extensive research has been carried out on PGDH homologs from other organisms, including mammals, plants, and bacteria.

PGDH belongs to the 2-hydroxy acid family of dehydrogenases and catalyzes the first committed step in the phosphorylated serine biosynthesis pathway: the oxidation of 3-phosphoglycerate (3-PGA) to 3-phosphohydroxypyruvate, with the concomitant reduction of NAD to NADH. Interestingly, PGDH is also capable of catalyzing the interconversion of -ketoglutarate (-KG) and 2-hydroxyglutarate (2-HGA) (5). The D-2-hydroxy-acid dehydrogenase family also includes D-lactate dehydrogenase (LDH; EC 1.1.1.28 [EC] ) and malate dehydrogenase (MDH; EC 1.1.1.37 [EC] ), which catalyze the conversion of lactate to pyruvate and malate (Mal) to oxalacetate (OAA), respectively. The family members share an overall sequence identity of 22% and a similarity of 50% (6). These enzymes also possess a conserved nucleotide-binding motif that preferentially binds to either NAD(H) or NADP(H).

View larger version (23K): [in this window] [in a new window]   FIGURE 1. Structure of the cyanobacterial hepatotoxins microcystin and nodularin. The general numbering of residues is indicated. The two variable amino acids in microcystin are indicated by X and Y. The most common isoform is microcystin LR, where X is L-Leu and Y is L-Arg. MeDha, N-methyldehydroalanine; Adda, 3-amino-9-methoxy-2,6,8-trimethyl-10-phenyldeca-4,6-dienoic acid; MeDhb, methyldehydrobutyrine.

As McyI shares sequence homology with PGDH, the first committed step in the phosphorylated serine biosynthesis pathway, it is conceivable that this enzyme may also play a role in serine metabolism. However, as no other serine biosynthesis enzymes (e.g. phosphoserine transaminase and phosphoserine phosphatase) are encoded within the mcy gene cluster, it is unlikely that McyI is involved in the production of this nonpolar amino acid.

An alternative hypothesis is that McyI is involved in the production of the methyl aspartate unit (MeAsp³) of the microcystin cyclic structure. A feeding study with [1,2-¹³C]acetate suggested that the MeAsp residues in microcystin and nodularin are synthesized via a condensation reaction between acetyl-CoA and pyruvic acid. 2-Hydroxy-2-methylsuccinic acid is then converted to 2-hydroxy-3-methylsuccinic acid, oxidized to 2-oxo-3-methylsuccinic acid, and finally transaminated to MeAsp (9). As McyI is a 2-hydroxy-acid dehydrogenase homolog, we predict that this enzyme catalyzes the interconversion of 2-hydroxy-3-methylsuccinic acid (3-methyl malate) to 2-oxo-3-methylsuccinic acid (3-methyl oxalacetate), the penultimate step in methyl aspartate biosynthesis (Fig. 2).

This study describes the bioinformatics, genetic, and enzymatic characterization of McyI. The results of these experiments are discussed with respect to the role of this unique methyl-malate dehydrogenase in microcystin biosynthesis.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Cyanobacterial Strains and Cultures—The microcystin-producing strain M. aeruginosa PCC7806 was kindly provided by E. Dittmann (Institute for Biology, Humboldt University, Berlin). Other cyanobacteria were obtained from the Institute for Biology, the Microbiology Division of the Biocenter at the University of Helsinki (Helsinki, Finland), and the cyanobacterial culture collection (University of New South Wales).

Cyanobacteria were grown as batch cultures in BG11 medium (Fluka) under 16 µmol photons m^-2 s^-1 white light. Light intensities were measured using a LI-COR LI-250 light meter (LI-COR Biosciences). The absorbance (A₆₅₀) of the cultures was measured using an Ultrospec II spectrophotometer (Biochrom Ltd.).

Cyanobacterial DNA Extractions—DNA was isolated from cyanobacteria according to the xanthogenate-SDS method of Tillett and Neilan (10).

Screening Various Strains of Cyanobacteria for mcyI Orthologs—The degenerate oligonucleotide primers mcyIdegenF (5'-TGTGCGTTATCCTAMTAA-3') and mcyIdegenR (5'-GGCTTCTCDCCCTGAAGC-3') were designed to amplify, via PCR, an 790-bp sequence within the mcyI locus of M. aeruginosa PCC7806 and Anabaena sp. 90. These primers were used to screen chromosomal DNA samples from several toxic and nontoxic strains of cyanobacteria for mcyI orthologs (see Table 1).

View larger version (19K): [in this window] [in a new window]   FIGURE 2. Putative reaction pathway to MeAsp in microcystin and nodularin biosynthesis. Adapted from Moore (Ref. 37). Step 3 is putatively catalyzed by McyI (or NdaH in the case of nodularin production).

Phylogenetic Analysis—A PSI-BLAST search with the McyI sequence returned 26 sequences with BLAST scores >200 (see Table 2). These sequences were subsequently used to generate a multiple sequence alignment and corresponding phylogenetic tree (ClustalX). Four reference sequences from characterized enzymes (PGDH, LDH, MDH, and formate dehydrogenase) were also included, as was Escherichia coli NAD-independent D-lactate dehydrogenase. The latter was designated as an artificial outgroup because it is a membrane-bound FAD flavoenzyme and does not belong to the D-isomer-specific 2-hydroxy-acid dehydrogenase family (11). Phylogenetic trees were generated using the neighbor-joining method with gaps removed. Trees were displayed graphically using NJplot and AppleWorks Version 6 (Apple Computer, Inc.).

Purification of Histidine-tagged McyI—Cell pellets were thawed on ice, resuspended in 2% of the original culture volume of cold phosphate buffer containing 1 mM phenylmethylsulfonyl fluoride, passed several times through an 18-gauge needle, and then briefly sonicated (Branson Sonifier model 250; amplitude of 25, 50% duty cycle, pulsed). The lysate was cleared by centrifugation at 20,000 x g for 30 min at 4 °C and precipitated on ice using 40% ammonium sulfate. The final pellet was resuspended in 0.4% of the original culture volume of cold phosphate buffer containing 100 mM imidazole, filtered through a 22-µm membrane, and applied to a 5-ml HiTrap column (Amersham Biosciences) charged with Ni²⁺. The column was washed with 10 column volumes of phosphate buffer containing 100 mM imidazole and eluted with 2 column volumes of phosphate buffer containing 300 mM imidazole. The wash and eluate were collected in 1-ml volumes and analyzed by SDS-PAGE (12% polyacrylamide gels) and Western blotting using a nickel-nitrilotriacetic acid-alkaline phosphatase conjugate (Qiagen Inc.) with 5-bromo-4-chloro-3-indolyl phosphate/nitro blue tetrazolium staining (Sigma). Purified protein samples intended for enzyme assays were desalted by size exclusion filtration using Amicon Ultra 10-kDa cutoff columns and 50 mM HEPES exchange buffer (pH 7).

Determining the Subunit Organization of Native McyI—The molecular masses of the individual subunits of McyI were estimated by size exclusion chromatography using an ÄKTA Basic 900 series fast protein liquid chromatography apparatus fitted with a Frac-920 fraction collector, a UV detector, and a Superdex 200 10/300 GL column (Amersham Biosciences). The column was equilibrated with wash buffer (50 mM sodium phosphate (pH 7) and 150 mM NaCl) and calibrated with 1 mg each of seven molecular mass standards (Sigma) (see Fig. 6). One milligram of purified McyI was subsequently loaded. All samples were run at a flow rate of 0.5 ml/min. Eluted proteins were detected spectrophotometrically (220–280 nm) and collected in 250-µl fractions. Standard curves (log of subunit molecular mass versus volume eluted) were based on the molecular masses of the protein standards. The eluted McyI fractions were diluted to 5 µg/ml and checked for activity by performing an OAA reductase assay (see below).

Chemical Synthesis of 2-Hydroxy-3-methylsuccinic Acid— 2-Hydroxy-3-methylsuccinic acid (3-methyl malate (3-MeMal)) was synthesized by reduction of diethyl oxalpropionate (10 g, 0.05 mol; Aldrich) in 95% ethanol (100 ml) by the addition of an 8-fold excess of sodium borohydride (3.8 g, 0.1 mol). After the mixture was stirred at room temperature for 24 h, 4 M HCl (20 ml) was slowly added, and the most of the ethanol evaporated under reduced pressure. Saturated sodium chloride solution (100 ml) was added, and the mixture was extracted five times with ethyl acetate (100 ml). The combined extracts were filtered, dried (MgSO₄), and evaporated, yielding a viscous oil (4.5 g, 45%). The ¹H NMR spectrum (CDCl₃) showed signals for the threo-isomer ( 1.07 (d, J = 7.2 Hz, 3-Me) and 2.9 (m, H-3)) and the erythro-isomer ( 3.0 (m, H-3)). The ratio of the threo-to erythro-isomers was 1:6. The other signals of the two isomers (erythro and threo H-2 and erythro 3-Me signals) are underneath the CH₂ resonances ( 4.1–4.3) and the CH₃ resonances ( 1.1–1.2) of the ethyl ester groups (data not shown).

Hydrolysis of the crude esters (2.0 g) was carried out by refluxing with 4 M HCl (50 ml) for 4 h. The cooled mixture was diluted with water (100 ml) and extracted with ethyl acetate (2 x 50 ml) to remove any unhydrolyzed esters and some unsaturated acids formed. The aqueous phase was evaporated to dryness, giving an oil (1.2 g). This was left to stand at -20 °C for 72 h after approximately half had crystallized. The oily crystals were washed briefly with cold chloroform, in which hydroxymethylsuccinic acids are insoluble, leaving a colorless crystalline solid (0.54 g). The 600-MHz ¹H NMR (²H₂O) spectrum now showed only the clearly resolved resonances for the two isomers (threo/erythro ratio of 1:6): threo-isomer, 4.54 (d, J = 4.05 Hz, H-2), 2.91 (m, H-3), and 1.0 (d, J = 7.14 Hz, 3-Me); and erythro-isomer, 4.28 (d, J = 4.17 Hz, H-2), 2.97 (m, H-3), and 1.05 (d, J = 7.20 Hz, 3-Me). The latter assignments are identical to those reported for the erythro-isomer (12). The resonances are also consistent with earlier NMR data recorded at lower fields and thus lacking some of the dispersion reported here for both the free acids and some derivatives (13–15). No attempts were made to separate these two isomers, and they were used as inhibitors in the ratio indicated above.

Oxidoreductase Assays—The dehydrogenase and reductase activities of recombinant McyI were analyzed in triplicate in 1-ml cuvettes. The production or disappearance of NAD(P)/H was monitored spectrophotometrically at 340 nm using a Cary 100 UV spectrophotometer (Varian, Inc.). Reactions were initiated by the addition of substrate or enzyme.

The dehydrogenase activities of McyI were measured by monitoring the conversion of NADP to NADPH. Each reaction contained 1 mM NADP, 0.1–10 mM substrate (3-MeMal, D-Mal, L-Mal, D-2-HGA, D-3-PGA), and 50 µg of purified McyI in 1 ml of assay buffer. The production of NADPH was monitored spectrophotometrically at 340 nm for 0.5–1 min at 37 °C.

The reductase activities of McyI were measured by monitoring the conversion of NADPH to NADP. Each reaction contained 0.25 mM NADPH, 0.1–5 mM substrate (-KG or OAA), and 2–5 µg of purified McyI in 1 ml of assay buffer. The disappearance of NADPH was monitored spectrophotometrically at 340 nm for 0.5–1 min at 37 °C.

End Product Analysis—The end products of the Mal/OAA oxidoreductase reactions were analyzed by NMR spectroscopy. NMR spectroscopy was carried out on a Bruker Advance DMX-600 spectrometer using a triple broadband inverse probe. ¹H NMR spectra were acquired at 600.13 MHz with a 90 ° pulse of 9 µs, a spectral width of 8992 Hz, 33,000 data points and a 2-s delay between pulses. Samples were run in 0.5 ml of 95% potassium phosphate (pH 7) and 5% ²H₂O. The standard Bruker water saturation pulse program was used. The free induction decay was zero-filled to 64,000 data points and processed with a line-broadening factor of 1 Hz before Fourier transformation.

Determining the Optimal pH of the McyI OAA Reductase Assay—The pH optimum for the McyI OAA reductase assay was determined by monitoring the disappearance of NADPH at 340 nm over a pH range of 5.5–9 at 25 °C. Each 1-ml reaction contained 0.25 mM NADPH, 0.8 mM OAA, and 5 µg of purified McyI in 1 ml of assay buffer (1.0 mM dithiothreitol, 0.1 mM phenylmethylsulfonyl fluoride, and 100 mM buffer (MES at pH 5.5–6.5, HEPES at pH 7–8, or Tris-Cl at pH 8.5–9)).

Determining the Optimal Temperature and Thermostability of the McyI OAA Reductase Assay—The temperature optimum for the McyI OAA reductase assay was determined by monitoring the disappearance of NADPH at 340 nm over a temperature range of 25–60 °C. Each 1-ml reaction contained 0.25 mM NADPH, 0.8 mM OAA, and 5 µg of purified McyI in 1 ml of assay buffer (pH 7). The stability of McyI at different temperatures was determined by incubating the enzyme for 3 min at 25–60 °C and then chilling it immediately on ice prior to performing the OAA reductase assay at 37 °C as described above.

McyI Specificity—Various structurally related compounds (lactate, pyruvate, D-Ser, L-Ser, L-Phe, oxalic acid, cinnamate, phenyl acetate, phenyl lactate, phenylalanine, and diethyl oxalpropionate; 0.1–10 mM) were tested as substrates of McyI in the forward (dehydrogenase) or reverse (reductase) direction at 37 °C for 1 min at pH 7 as described above. To test whether McyI is able to use non-phosphorylated cofactors, the 3-MeMal, Mal, OAA, and -KG oxidoreductase assays were repeated using NAD/H in place of NADP/H.

McyI Kinetic Analysis—Kinetic analysis of the -KG and OAA reductase activities of McyI was carried out in 1-ml reactions each containing 0.25 mM NADPH, 5 µg of purified enzyme, and various amounts of substrate in reaction buffer (pH 7). Initial velocities were measured by monitoring the disappearance of NADPH at 340 nm for 0.5 min at 37 °C.

Kinetic analysis of the NADPH oxidase activity of McyI was performed in 1-ml reactions each containing 5 µg of purified enzyme, 0.8 mM OAA, and various amounts of NADPH in reaction buffer (pH 7). Initial velocities were measured as described above.

Inhibition Assays—The inhibitory effects of various substances on the OAA and -KG reductase activities of McyI were tested. Assays were performed in 1 ml of reaction buffer (pH 7) containing 0–100 mM putative inhibitor (L-Ser, 3-MeMal, L-Mal, D-Mal, L-Asp, D-Asp, D-2-HGA, or D-3-PGA), 0.25 mM NADPH, 5 µg of enzyme, and 0.2 mM OAA or 1 mM -KG. The disappearance of NADPH was monitored spectrophotometrically at 340 nm for 0.5–1 min at 37 °C.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Distribution of mcyI Orthologs in Various Species of Cyanobacteria—To assess the distribution of mcyI orthologs among toxic and nontoxic cyanobacteria, DNAs from 24 different strains were screened by PCR with degenerate oligonucleotide primers. The results are presented in Table 1. All microcystin-producing strains, except for Planktothrix agardhii CYA126, tested positive for mcyI. However, mcyI orthologs were not detected in the nodularin-producing strains. Interestingly, nearly half of all the nontoxic cyanobacteria screened also tested positive for mcyI.

Sequence Analysis of mcyI—The 1014-bp mcyI open reading frame was predicted to encode a 36.71-kDa peptide with a pI of 5.62. Although dense alignment surface (DAS) method analysis indicated two possible membrane-spanning domains within McyI (between residues 123 and 125 and residues 163 and 164), these were in the low confidence range. The Kyte-Doolittle plot (ProtScale) suggested that the McyI peptide is largely hydrophilic, although several small hydrophobic regions were identified (data not shown).

PSORTb gave a cytoplasmic localization score of 9.26, indicating that McyI is a cytoplasmic protein. The mcyI sequence contains 32 rare codons, including several repeated and/or consecutive rare codons. Subsequent heterologous expression studies were therefore carried out in the Rosetta(DE3)pLysS expression strain containing the pRARE plasmid encoding these rare E. coli tRNA codons.

Comparison of the inferred primary peptide sequence of mcyI with other sequences in the NCBI Database revealed significant similarity (up to 72% identity and 84% similarity using PSI-BLAST) to putative and experimentally characterized enzymes, mainly PGDHs, of bacterial (mostly cyanobacterial), archaeal, and eukaryotic origins. The top five PSI-BLAST results were sequences from N. spumigena (71% identity and 84% similarity), Anabaena sp. (72% identity and 84% similarity), Methanopyrus kandleri (43% identity and 64% similarity), Rubrobacter xylanophilus (42% identity and 60% similarity), and Methanothermobacter thermautotrophicus (41% identity and 61% similarity) (see Table 2 for accession numbers). A genomic BLAST search was also performed to identify previously characterized E. coli homologs. The returned sequences included 2-keto-D-gluconate reductase, PGDH, LDH, and erythronate-4-phosphate dehydrogenase, although scores were generally quite low (32% identity and 50% similarity). BL2SEQ analysis was also performed for McyI and E. coli PGDH (NCBI accession number P08328) and returned scores of 31% identity and 48% similarity. The individual domains of McyI (putative nucleotide and substrate domains) were also analyzed by blastp; however, the results generally reflected those of the full-length blastp search, i.e. no new sequences were returned.

A ScanProsite pattern search also identified D-isomer-specific 2-hydroxy-acid dehydrogenase NAD-binding signatures (LIVMA)(AG)(IVT)(LIVMFY)(AG)XG(NHKRQGSAC)(LIV) GX_13,14(LIVMFT)X₂(FYWCTH)(DNSTK) at residues 158–186 of the McyI peptide (Fig. 3). Enzymes possessing these signatures are structurally related and have similar enzyme activities (16–18).

The sequence alignment of McyI with other cyanobacterial homologs and E. coli PGDH revealed that McyI from M. aeruginosa and Anabaena sp. and NdaH (N. spumigena) lack 72 amino acids that compose the corresponding C-terminal regulatory (ACT) domain, making them, on average, 18% shorter than E. coli PGDH. Several other sequence variations between McyI and PGDH were also apparent (Fig. 3).

To gain an overview as to the relative phylogenetic position of McyI within the 2-hydroxy-acid dehydrogenase family, a phylogenetic analysis was performed. Sequence lengths used in the analysis ranged from 306 to 624 amino acids (Table 2). The phylogenetic tree (Fig. 4) partitioned into three main subgroups. Subgroup A, which contains McyI, is composed exclusively of sequences from the hepatotoxin biosynthesis gene clusters of cyanobacteria. Subgroup B is composed of putative dehydrogenases from Arabidopsis thaliana, several species of cyanobacteria, and the thermophilic actinobacterium R. xylanophilus. Subgroup C consists mostly of archaeal proteins, with the exception of two eubacterial sequences, Thermoanaerobacter tengcongensis (order Clostridiales) and Oceanobacillus iheyensis (order Bacillales).

Approximately one-third of all sequences in the phylogenetic tree possess amino acid C-terminal regulatory domains. Although these regulatory domains are present in each organismic lineage, they are most prevalent in the plant and archaeal subgroups. Clustering of ACT domain-possessing proteins within each organismic lineage is apparent.

The tree also demonstrates that subgroups B and C share a common ancestor that had an earlier divergence leading to the subgroup A lineage of cyanobacterial proteins. The distribution of the reference sequences within the tree supports the prediction that the subgroup A/B/C lineage evolved from a common PGDH ancestor rather than the ancestral MDH, LDH, or formate dehydrogenase.

Overexpression and Purification of Histidine-tagged McyI— McyI was overexpressed in E. coli with an N-terminal hexahistidine fusion tag. The recombinant peptide was highly expressed under all culture/induction conditions tested. Western blot analysis confirmed that McyI was expressed without truncation at the expected molecular mass of 40 kDa. Maximal yields of soluble protein (up to 10% total protein) were obtained after overnight induction at 25 °C. However, a 2.5-h induction at 30 °C also provided sufficient yields (5–10% total protein) for enzyme assays.

The recombinant McyI protein precipitated in the presence of 40% ammonium sulfate and eluted from the Ni²⁺-charged affinity column at an imidazole concentration of 300 mM. The recombinant protein eluate was of >95% purity and was eluted at concentrations of 1–2.5 mg/ml (Fig. 5).

Subunit Composition of McyI—The purified McyI protein eluted from the size exclusion column at 27.8 min, corresponding to a molecular mass of 85 kDa (dimeric form) (Fig. 6). The collected elution fractions (25–30 min) displayed OAA reductase activity of 12.5–14 µmol/min/mg, which was comparable with that of the unfractionated enzyme (data not shown).

Biochemical and Kinetic Properties of McyI—McyI was stable for >2 weeks at 4 °C without significant loss of activity and for at least 6 months at -20 °C in 15% glycerol. Plots of McyI OAA reductase activity after 1-min incubations at various temperatures were bell-shaped with an optimum of 45 °C at pH 7. However, McyI was relatively unstable at higher temperatures (90% drop in activity following preincubation at 50 °C). The pH optimum for the McyI OAA reductase activity was 7l however, the enzyme maintained at least 80% of its activity between pH 6.5 and 7.5 at 25 °C (Fig. 7).

McyI displayed weak 3-MeMal, D-Mal, and L-Mal oxidase activities (0.2–0.7 µmol/min/mg of protein under the described conditions). However, oxidation of D-2-HGA or D-3-PGA could not be detected. Because of strong product inhibition and the high concentration of enzyme required for each assay, we were unable to obtain kinetic constants for the oxidase reactions.

The reduction of OAA was the major in vitro activity observed for McyI, with V^app_max = 15.2 µmol/min/mg of protein at 37 °C (Fig. 8). However, high concentrations of substrate (>0.8 mM) inhibited the reaction. The enzyme was also able to reduce -KG, although the rates of catalysis were much lower (V^app_max 3.6 µmol/min/mg of protein). The OAA and -KG reductase reactions both obeyed regular Michaelis-Menten kinetics (Table 3).

End Product Analysis—NMR spectroscopic analysis confirmed that malate was the product of the OAA reductase reaction (supplemental Fig. S1).

Inhibition Assays—The OAA reductase activity of McyI was inhibited by D-Mal (I₅₀ = 2.2 mM) and 3-MeMal (I₅₀ = 0.7 mM) in a negatively cooperative manner. Interestingly, L-Mal did not inhibit the reaction, nor did L-Ser, D-Asp, L-Asp, D-2-HGA, or D-3-PGA. The -KG reductase activity of McyI was inhibited by 3-MeMal (I₅₀ = 2.3 mM) and D-2-HGA (I₅₀ = 20.5 mM) (Fig. 9 and Table 3).

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
To assess the role of mcyI in microcystin production, a variety of hepatotoxic and nontoxic strains of cyanobacteria were screened for mcyI orthologs. With the exception of P. agardhii CYA126, the degenerate primers successfully amplified an 790-bp PCR fragment from all microcystin-producing strains tested (Table 1). Interestingly, nearly half of all the nontoxic cyanobacterial strains examined tested positive for mcyI. These results reflect those of previous studies that identified mcy genes in several nontoxic Microcystis strains (3, 19, 20). It is likely that these strains have reverted to a nontoxic phenotype because of mutations within their mcy biosynthesis genes or within mcy regulatory regions. Previous studies have demonstrated that the mcy gene cluster does not well tolerate mutations with respect to toxin biosynthesis; in fact, the mutation of several individual mcy genes (mcyA, mcyB, mcyD, mcyE, mcyF, and mcyH) has led to the production of mutant strains with nontoxic phenotypes (3, 19, 21).

View larger version (58K): [in this window] [in a new window]   FIGURE 3. Alignment of McyI (M. aeruginosa, NCBI accession number AAF00955; and Anabaena sp., AA062580), NdaH (N. spumigena, AA064409), and PGDH (E. coli, P08328). Structurally important amino acid residues and the NAD-binding signature are indicated (see "Results" for details). Consensus residues are highlighted. Colored regions define the three domains of PGDH: the nucleotide-binding domain (residues 108–294; blue), the substrate-binding domain (residues 7–107 and 295–336; red), and the regulatory domain (residues 338–409; yellow).

View larger version (31K): [in this window] [in a new window]   FIGURE 4. Phylogenetic tree incorporating McyI and similar dehydrogenases. The phylogenetic tree encompasses PGDH homologs from cyanobacteria (boldface) and other organisms. An E. coli D-lactate dehydrogenase (DLD) constitutes the artificial outgroup. Asterisks indicate sequences with ACT (regulatory) domains. For a detailed list of the sequences used, see Table 2. Bootstrap values derived from 1000 replicates are given. The scale at the top of the tree corresponds to 5% divergence between sequences.

View larger version (53K): [in this window] [in a new window]   FIGURE 5. Purification of McyI. Hexahistidine-tagged McyI was eluted from a Ni2+-charged HiTrap column. Lane 1, broad-range molecular mass standards (shown in kilodaltons); lanes 2–10, McyI elution fractions 2–10.

Homology-based searches indicated that McyI is a cytosolic enzyme belonging to the 2-hydroxy-acid dehydrogenase family. Although its closest characterized relative, A. thaliana PGDH, is involved in Ser metabolism, sequence data suggest an alternative function for McyI. An immediately obvious difference between the peptide sequence of McyI and those of bacterial PGDHs is its lower subunit molecular mass (37 versus 44 kDa). This is largely due to the fact that McyI lacks an ACT domain (residues 338–409 in E. coli PGDH) (24). These regulatory domains have been linked to a wide range of metabolic enzymes that are controlled by amino acids in a concentration-dependent fashion (25).

An alternative regulatory mechanism such as transcriptional control may negate the need for an ACT domain in McyI and other cyanobacterial enzymes (e.g. those present in the phylogenetic tree in Fig. 4). The fact that McyI has two promoters, i.e. its own individual promoter as well as the central mcyD–J promoter, strongly supports this hypothesis (26, 27).

A close comparison of the primary peptide sequence of McyI with those of other 2-hydroxy-acid dehydrogenases in the data bases revealed further interesting characteristics of the protein. A large proportion of amino acids in McyI are highly conserved among most of the other dehydrogenases examined, including the NAD(P)-binding signature encompassing residues 158–186 of the McyI sequence. This nucleotide-binding domain is conserved among all 2-hydroxy-acid dehydrogenases, including PGDH, MDH, LDH, and glycerate dehydrogenase. The identification of a nucleotide-binding domain within McyI suggests that it is a NAD(P)-dependent enzyme with an oxidoreductase function similar to that of PGDH.

View larger version (20K): [in this window] [in a new window]   FIGURE 6. Determination of McyI subunit organization. A, elution standard curve for protein molecular mass standards; B, chromatogram of purified recombinant McyI. The peak at 27.8 min corresponds to the McyI dimer (85 kDa, including the His tag). All samples were run on an ÄKTA Basic 900 series fast protein liquid chromatography apparatus fitted with a Superdex 200 10/300 GL column.

View larger version (11K): [in this window] [in a new window]   FIGURE 7. Biochemical properties of the McyI OAA reductase activity. A and B, OAA reductase activity of McyI at different pH values and different temperatures, respectively; C, McyI thermal stability. See "Results" for details regarding reaction conditions. The data are presented as the means of two to three separate experiments.

View larger version (20K): [in this window] [in a new window]   FIGURE 8. Kinetic plots for the McyI OAA reductase activity. A, V versus [S] plot; B, Lineweaver-Burk plot. [S] equals millimolar substrate (OAA). The data presented are from a single experiment using 5 µg of purified McyI and 0.25 mM NADPH at pH 7 (see "Results" for details). These results were reproducible (at least four times on separate days) within an error margin of 10%.

View larger version (13K): [in this window] [in a new window]   FIGURE 9. Inhibition of McyI reductase activities. A, inhibition of OAA reductase activity by D-Mal and 3-MeMal; B, inhibition of -KG reductase activity by 3-MeMal. The x axes correspond to inhibitor concentration, and the y axes correspond to percentage enzyme activity remaining. One-hundred percent activity corresponds to activity in the absence of inhibitor. The data are presented as the means of two to three separate experiments.

His²⁹² is important for catalyzing the interconversion of 3-phosphohydroxypyruvate and 3-PGA in PGDH. In addition, Glu²⁶⁹ acts in tandem with His²⁹² to form a proton shuttle, as seen in many dehydrogenases (28). The conservation of these residues within McyI again supports an oxidoreductase function for this enzyme.

In PGDH, Gly²⁹⁴ and Gly²⁹⁵ form the link between the substrate-binding and nucleotide-binding domains, which form the active-site cleft of the enzyme. Mutations of these residues affect the K_cat of the enzyme without appreciably affecting sensitivity to serine (31). Although Gly²⁹⁵ is conserved in McyI and NdaH, Gly²⁹⁴ is replaced by Ala. Mutational studies have shown that G294A substitutions do not affect the catalytic capacity of PGDH (31), and therefore, this conservative substitution should not influence the K_cat of McyI or NdaH. Indeed, homologous enzymes can tolerate a number of different amino acids at this position, as evidenced by the alignment (Fig. 3) in which several sequences have either Ser or Glu in the first position and Lys, Arg, or Asn in the second position.

In an attempt to understand the evolution of McyI in the context of other 2-hydroxy-acid dehydrogenases such as PGDH, MDH, and LDH, a phylogenetic tree was constructed encompassing sequences from plants, Archaea, and eubacteria, including several species of cyanobacteria (Fig. 4). Interestingly, the cyanobacterial sequences cluster polyphyletically. Although the subgroup B cyanobacterial proteins appear to share a common ancestor with plant PGDHs, the hepatotoxin-associated proteins form a phylogenetically distinct subgroup (subgroup A). McyI and NdaH must have therefore diverged early in the evolution of PGDHs, prior to the divergence of the archaeal, plant, and subgroup B cyanobacterial protein lineages.

In a similar phylogenetic study in which PGDHs from a wide range of organisms were investigated, Ali et al. (32) reported that sequences from the order Bacillales also display a polyphyletic distribution. They concluded that many lateral gene transfer events, together with drastic insertion/deletion events, occurred during the evolution of PGDH, making the history of this superfamily very complex. The distribution of ACT domains among 2-hydroxy-acid dehydrogenases contributes to this complexity. In the phylogenetic tree presented in Fig. 4, it appears that the ancestral PGDH possessed an ACT domain, yet this domain was lost in some (but not all) subsequent generations. It is interesting to note that even proteins from closely related organisms (e.g. Synechocystis sp. PCC6803 and Synechocystis sp. WH8102) may differ when it comes to possession of an ACT domain. These results suggest that alternative regulatory mechanisms such as transcriptional control must be in place for many of these enzymes.

Although none of the reference sequences (PGDH, LDH, MDH, and formate dehydrogenase) partitioned within subgroups A–C, the fact that PGDH from A. thaliana fell within subgroup B suggests that the subgroup B cyanobacterial sequences, and probably the archaeal sequences, also function in serine metabolism. Conversely, the distantly related subgroup A cyanobacterial proteins (McyI and NdaH) are likely to have evolved functions specific to hepatotoxin biosynthesis.

To investigate the remarkable prospect that McyI may be a PGDH, the recombinant enzyme was assayed for 3-PGA dehydrogenase activity in vitro. However, despite extensive experimental variation, PGDH activity was not detected. The lack of PGDH activity also reflects the phylogenetic data presented in this study suggesting that McyI is evolutionarily distant to PGDH and belongs to a separate clade of enzymes, uniquely present in the hepatotoxic cyanobacteria. The comparative sequence data also support this notion and predict distinct functions for McyI and PGDH.

Although McyI is therefore clearly not involved in Ser metabolism, it is probable that M. aeruginosa possesses a true PGDH elsewhere in its genome. Genome sequencing projects and/or genetic screening studies should shed light on this topic in the near future.

Interestingly, McyI was able to utilize -KG/OAA and Mal as substrates, but not lactate or pyruvate. As with most 2-hydroxy-acid dehydrogenases, the major in vitro activity observed for McyI was in the reductase direction, i.e. the reduction of OAA to Mal. Oxalacetate showed a 30-fold decrease in K^app_m over -KG and a 100-fold increase in V^app_max/K^app_m. The higher catalytic efficiency of OAA is consistent with the hypothesis that the native substrate of McyI (3-methyl oxalacetate (3-MeOAA)) is a structural analog of OAA.

Both phosphorylated and dephosphorylated dinucleotide cofactors were utilized by McyI, yet NADPH was greatly preferred over NADH. It would be interesting to determine whether the putative cyanobacterial PGDHs used in this phylogenetic study are also NAD-specific. If so, it is likely that the nucleotide-binding domain of McyI evolved from a NAD-specific form to its current NADP-specific form.

As the reduction of OAA was the major in vitro activity observed for McyI, we predicted that the true physiological substrate of McyI must be a Mal/OAA analog. This led to the hypothesis that McyI catalyzes the conversion of 3-MeMal to 3-MeOAA, with subsequent transamination to MeAsp. To test this hypothesis, a racemic diastereomeric mixture of 3-MeMal was synthesized and tested as a substrate in an in vitro dehydrogenase assay. Unfortunately, the attempted synthesis of the unstable 3-MeOAA to test the reverse reaction (3-MeOAA to 3-MeMal) was not successful. McyI was able to oxidize 3-MeMal at a comparable rate to D- and L-Mal; however, there appeared to be strong end product inhibition, which significantly reduced the linear range of the assay (<30 s).

To understand the inhibitory effects of various substrates on the McyI reductase reactions, several inhibition assays were performed. Unlike PGDH, McyI was not allosterically regulated by Ser or by any other compound tested. This result was not surprising because McyI lacks an ACT regulatory domain. 3-MeMal and D-Mal inhibited McyI OAA reductase activity in a negatively cooperative fashion. 3-MeMal was the strongest inhibitor, with I₅₀ 0.7 mM. Interestingly, L-Mal had no inhibitory effects, suggesting that, like other 2-hydroxy-acid dehydrogenases, McyI preferentially binds substrates with D-conformations. It is therefore likely that the I₅₀ for 3-MeMal is overestimated because the 3-MeMal compound was a racemic diastereomeric mixture. These results support the hypothesis that McyI is a 3-MeMal/3-MeOAA oxidoreductase.

The nonproteinogenic amino acid 3-MeAsp is a common intermediate of the mesaconate pathway for (S)-glutamate fermentation in Clostridium spp. (33) and in members of the family Enterobacteriaceae (34). This unusual amino acid residue also occurs in the lipopeptide antibiotic friulimicin (35) and in the cyanobacterial hepatotoxins microcystin and nodularin (3, 7). In the mesaconate and friulimicin pathways, the route to MeAsp occurs via rearrangement of Glu to 3-MeAsp involving glutamate mutase. However, genome sequencing efforts have failed to identify glutamate mutase homologs in cyanobacteria. We propose that the cyanobacteria utilize an alternative glutamate mutase-independent biosynthetic pathway to 3-MeAsp, in which McyI plays a pivotal role.

Labeled precursor feeding experiments by Moore et al. (9) suggested that the MeAsp residues in microcystin and nodularin are produced via the rearrangement of citramalic acid, yet the authors were unable to identify the enzymes involved in this pathway. In this study, we have provided evidence that the 2-hydroxy-acid dehydrogenase McyI catalyzes the interconversion of 3-MeMal to 3-MeOAA. The final step in the MeAsp pathway, the conversion of 3-MeOAA to MeAsp, is likely to be catalyzed by a promiscuous Asp aminotransferase, as these enzymes occur in virtually all organisms, including cyanobacteria (36).

To our knowledge, McyI is the only reported example of a 3-MeMal/3-MeOAA oxidoreductase. On the basis of the data presented in this study, it appears that McyI and its homolog NdaH diverged from a PGDH ancestor to become NADP-dependent enzymes with specific roles in hepatotoxin production. Future mutagenesis experiments involving mcyI and ndaH will undoubtedly offer further insight into the role of these unique cyanobacterial enzymes.

	FOOTNOTES

 
^* This work was supported by the Australian Research Council and the Cooperative Research Centre for Water Quality and Treatment. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. S1.

¹ To whom correspondence should be addressed. Tel.: 612-9385-3235; Fax: 612-9385-1591; E-mail: b.neilan{at}unsw.edu.au.

² The abbreviations used are: PGDH, D-3-phosphoglycerate dehydrogenase; 3-PGA, 3-phosphoglycerate; -KG, -ketoglutarate; 2-HGA, 2-hydroxyglutarate; LDH, D-lactate dehydrogenase; MDH, malate dehydrogenase; Mal, malate; OAA, oxalacetate; MeAsp, methyl aspartate; 3-MeMal, 3-methyl malate; MES, 4-morpholineethanesulfonic acid; 3-MeOAA, 3-methyl oxalacetate; ACT, aspartokinase chorismate mutase and TyrA (prephenate dehydrogenase).

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