SHMT (serine hydroxymethyltransferase; EC 2.1.2.1) catalyses reversible hydroxymethyl group transfer from serine to H4PteGlun (tetrahydrofolate), yielding glycine and 5,10-methylenetetrahydrofolate. In plastids, SHMTs are thought to catalytically direct the hydroxymethyl moiety of serine into the metabolic network of H4PteGlun-bound one-carbon units. Genes encoding putative plastid SHMTs were found in the genomes of various plant species. SHMT activity was detected in chloroplasts in pea (Pisum sativum) and barley (Hordeum vulgare), suggesting that plastid SHMTs exist in all flowering plants. The Arabidopsis thaliana genome encodes one putative plastid SHMT (AtSHMT3). Its cDNA was cloned by reverse transcription–PCR and the encoded recombinant protein was produced in Escherichia coli. Evidence that AtSHMT3 is targeted to plastids was found by confocal microscopy of A. thaliana protoplasts transformed with proteins fused to enhanced green fluorescent protein. Characterization of recombinant AtSHMT3 revealed that substrate affinity for and the catalytic efficiency of H4PteGlu1-8 increase with n, and that H4PteGlu1-8 inhibit AtSHMT3. 5-Methyltetrahydrofolate and 5-formyltetrahydrofolate with one and five glutamate residues inhibited AtSHMT3-catalysed hydroxymethyl group transfer from serine to H4PteGlu6, with the pentaglutamylated inhibitors being more effective. Calculations revealed inhibition with 5-methyltetrahydrofolate or 5-formyltetrahydrofolate resulting in little reduction in AtSHMT3 activity under folate concentrations estimated for plastids.

INTRODUCTION

SHMT (serine hydroxymethyltransferase) catalyses reversible hydroxymethyl group transfer from serine to H4PteGlun (tetrahydrofolate), yielding glycine and 5,10-CH2-H4PteGlun (5,10-methylenetetrahydrofolate) [1,2]. Multiple SHMTs are encoded in plant genomes; bioinformatic prediction suggests the presence of seven enzymes in Arabidopsis thaliana [3,4] and five in rice (Oryza sativa) [5]. Previous studies have reported SHMT activity in mitochondria [1,68], plastids [1,8], the nucleus [8] and the cytosol [1,7,8] in plants. The gene for a mitochondrial SHMT from pea has been cloned, and the encoded enzyme has been detected in mitochondria using antibodies raised against the recombinant protein [6]. Also, the gene for a mitochondrial SHMT in which mutation causes a photorespiratory phenotype in A. thaliana has been identified [9]. Formation of a complex with ferredoxin-dependent glutamate synthase modulates activity of the enzyme encoded by this gene in vivo [10]. Little is known, however, about non-mitochondrial SHMTs in plants.

Plastid SHMTs were the focus of the present study. These enzymes provide one-carbon (C1) units for the biosynthesis of 5,10-CH2-H4PteGlun, which can then be oxidized to 5,10CH+-H4PteGlun (5,10-methenyltetrahydrofolate) and 10-HCO-H4PteGlun (10-formyltetrahydrofolate) by other catalysts [2]. 5,10-CH2-H4PteGlun, 5,10-CH+-H4PteGlun and 10-HCO-H4PteGlun are required in plastids for the biosynthesis of N-formylmethionine [2], thymidine nucleotide [2,11] and the inosine monophosphate precursors formylglycinamide ribonucleotide and 5-formaminoimidazole-4-carboxamide ribonucleotide [2,12]. Also, 5,10-CH+-H4PteGlun serves as a light-harvesting cofactor in a plastid-localized cryptochrome [13]. SHMT activity in plastids is thus potentially vital for photoreception and for the biosynthesis of purines, pyrimidines and N-formylmethionine.

Cellular folates, including those that serve as substrates and inhibitors of SHMTs, exist as conjugates with various numbers of glutamate residues, which are linked to a chain through the γ-carboxy group. Folates having up to 11 glutamate residues have been found in a diverse range of organisms, including plants [14,15]. Understanding how the degree of folate polyglutamylation of the substrate and inhibitor affects the catalytic properties of SHMTs is key to understanding how these enzymes function within the context of plant C1 metabolism.

Although SHMTs have been isolated and, in some cases, partially characterized from several plant species [3,16], the kinetic properties of those enzymes against folylpolyglutamate substrate species have yet to be investigated. Studies on inhibition of individual plant SHMTs by 5-HCO-H4PteGlun and 5-CH3-H4PteGlun (5-methyltetrahydrofolate), which are known to bind or inhibit SHMTs in other organisms [1723], are also lacking.

In the present paper, we report biochemical evidence for SHMT activity in plastids in both A. thaliana and barley (Hordeum vulgare), supporting the bioinformatic prediction suggesting that this enzyme resides in both monocotyledon and dicotyledon plastids. Also, we describe cDNA cloning, recombinant expression and biochemical characterization of a putative plastid SHMT from A. thaliana (AtSHMT3), and show evidence that AtSHMT3 resides in plastids from fluorescence microscopy of EGFP (enhanced green fluorescent protein)-fused protein constructs. The significance of these new findings for the overall understanding of C1 metabolism is discussed.

EXPERIMENTAL

Reagents and plant growth conditions

(6R,S)-H4PteGlu1, (6R,S)-5,10-CH2-H4PteGlu1, (6R,S)-5-HCO-H4PteGlu1&5, (6R,S)-5-CH3-H4PteGlu1-5 and PteGlu1-8 were purchased from Schircks Laboratories. Benzonase nuclease and recombinant enterokinase were from Novagen. Oligonucleotides were from MWG. Percoll was from GE Healthcare. Serine was from Sigma–Aldrich. KBH4 was from Acros Organics.

A. thaliana, ecotype Columbia, was grown in potting soil under 12 h light (100–120 μE·m−2·s−1)/12 h dark for 3 weeks at 22 °C. Pea, cv. Progress 9, and barley, cv. Bob, were respectively grown in coarse vermiculite and potting soil under 12 h light (75 μE·m−2·s−1)/12 h dark for 7–10 days at 22 °C during the light period and at 18 °C during the dark period.

Phylogenetic analysis

Putative SHMT protein sequences were obtained from GenBank®, http://www.phytozome.net and http://www.brachypodium.org databases using the BLASTP and TBLASTN prediction programs with AtSHMT3 as a query sequence. The phylogenetic analysis included SHMT protein sequences from nine representative plant species: A. thaliana, Populus trichocarpa, Vitis vinifera, O. sativa, Zea mays, Brachypodium distachyon, Chlamydomonas reinhardtii, Ostreococcus lucimarinus and Physcomitrella patens. These species were selected because their genomes have been fully sequenced. The accession numbers for the sequences used in the phylogenetic analysis are listed in Supplementary Table S1 at http://www.BiochemJ.org/bj/430/bj4300097add.htm. The phylogenetic tree was made with MEGA 4.0 [24]. The neighbour-joining method included Poisson correction, complete deletion and (1000) bootstrap replication.

cDNA cloning, constructs, sequence analysis and expression in Escherichia coli

The AtSHMT3 cDNA (At4g32520) was cloned by reverse transcription–PCR. Total A. thaliana RNA was isolated and reversetranscribed as described previously [25]. The AtSHMT3 open reading frame was amplified using Taq2000 DNA polymerase (Stratagene) and the primer pair 5′-GACGACGACAAGATGCAAGCTTGTTGTGG-3′ (AtSHMT3 Forward) and 5′-GAGGAGAAGCCCGGTTTAAACGCCAGGAATGGGAA-3′ (AtSHMT3 Reverse). Note that the vector-specific sequences (underlined), needed for cloning into the pET Ek/LIC expression vectors (Novagen), flank the gene-specific sequences of the primers. The amplified open reading frame was purified with a Wizard PCR Prep mini-column (Promega), and then cloned into the pGEM-T Easy vector (Promega) to generate the pGEM-AtSHMT3 construct.

The AtSHMT3 open reading frame, excluding the region coding for the putative organellar targeting peptide, was amplified from pGEM-AtSHMT3 using Pfu DNA polymerase (Stratagene) and the primer pair 5′-GACGACGACAAGATGAGAGCATCTTCAGT-3′ (AtSHMT3n Forward) and AtSHMT3 Reverse. The resulting PCR fragment was purified with a Wizard PCR column, treated with T4 DNA polymerase (Promega) and then inserted into the pET-44 Ek/LIC vector. All procedures were carried out in accordance with the manufacturer's protocols. All constructs were verified by DNA sequencing.

The expression vectors were introduced into the Rosetta strain of E. coli (Novagen) to produce the recombinant protein. Bacteria carrying the expression vector were cultured at 37 °C in LB (Luria–Bertani) medium containing 100 μg/ml kanamycin and 34 μg/ml chloramphenicol until the A600 reached 0.6–1. After addition of isopropyl β-D-thiogalactopyranoside (200 μM final concentration), the culture was incubated further at 15 °C overnight.

Purification of the recombinant protein

Induced E. coli cells expressing recombinant AtSHMT3 were harvested by centrifugation at 7500 g for 10 min at 4 °C. Bacterial cells were resuspended in 1 ml of buffer A {50 mM Tris/HCl, pH 8.5, 1 mM THP [tris-(3-hydroxypropyl)phosphine], 0.01 mM PLP (pyridoxal 5′-phosphate) and 10% glycerol} plus 25 units/ml Cenzonase® nuclease, and were mechanically broken with 0.1 mm zirconia/silica beads (BioSpec Products). Soluble protein extracts were cleared by centrifugation at 20800 g for 15 min at 4 °C, followed by filtering supernatants through a 0.45-μm pore-size PVDF membrane. The recombinant protein was purified by anion-exchange chromatography using an ÄKTA FPLC system equipped with a Mono Q 5/50 GL column (GE Healthcare); bound proteins were eluted using a linear gradient of buffer A to buffer A plus 1 M NaCl over 20 column volumes. Fractions containing the recombinant protein were desalted with PD-10 desalting columns (GE Healthcare) equilibrated with buffer A, and then digested with recombinant enterokinase. The digested recombinant enzyme was purified further on a Mono Q 5/50 GL column as described above; untagged AtSHMT3 was eluted in the flow-through. Fractions containing the recombinant enzyme were frozen in liquid N2 and stored at −80 °C until use. Freezing in liquid N2 had no effect on the activity of AtSHMT3. The recombinant enzyme for activity assays was desalted with 50 mM Ches [2-(N-cyclohexylamino)ethanesulfonic acid]/Hepes/citric acid buffer (pH 8.5) [26], containing 1 mM THP, 0.25 mM PLP and 10% glycerol.

Extraction of proteins from leaves and chloroplasts of pea and barley

To isolate soluble leaf proteins, pea or barley leaves were pulverized in liquid N2, and the material was suspended in 1.25 ml of buffer B (200 mM potassium phosphate, pH 7.5, 10 mM 2-mercaptoethanol, 1 mM EDTA, 1 mM PMSF and 10% glycerol) per 0.5 g of leaf material. Leaf protein extracts were mixed briefly by stirring, mixtures were cleared by centrifugation at 20800 g for 30 min at 4 °C, and then the supernatant was desalted with a PD-10 column equilibrated with buffer B without PMSF.

Chloroplasts of pea or barley were isolated by centrifugation on Percoll density gradients using published procedures [2729]. Isolated chloroplasts were resuspended in buffer B and broken by four cycles of freezing and thawing. Chloroplast protein extracts were cleared by centrifugation at 20800 g for 20 min at 4 °C, followed by desalting supernatants with Zeba Desalt spin columns (Pierce Biotechnology) equilibrated with buffer B without PMSF. The plastid marker glyceraldehyde-3-phosphate dehydrogenase and the mitochondrial marker fumarase were assayed for activity as described previously [30]. MTHFR (5,10-methylenetetrahydrofolate reductase) was assayed to evaluate contamination of isolated chloroplasts with the cytosol [31].

MTHFR activity was assayed as described previously [31], but using unlabelled assay components and HPLC with fluorescence detection. Assay mixtures (20 μl final volume) were incubated at 30 °C for 20 min. Assay blanks had the enzyme added after the incubation. The reactions were stopped by the addition of 1 M 2-mercaptoethanol (10 μl), followed by boiling reaction mixtures for 4 min; denatured proteins were pelleted by centrifugation at 20800 g for 30 min at 4 °C. Reaction products were separated isocratically on a Waters XTerra C18 column (4.6 mm×100 mm, 5 μm particle diameter), and monitored with 280 nm excitation and 359 nm emission wavelengths. The mobile phase contained 27 mM phosphoric acid and 7% methanol. The length of separation was 6 min. Authentic 5-CH3-H4PteGlu1 was used as a standard for quantification.

Leaf or chloroplast protein extracts were frozen in liquid N2 and stored at −80 °C until use. Protein concentrations were determined using the Bradford assay [32] with BSA as the standard.

Transient expression of EGFP-fused proteins in A. thaliana protoplasts

The full-length AtSHMT3-coding sequence was amplified with primers 5′-AAAAAGCAGGCTATGCAAGCTTGTTGTGGTGG-3′ (AtSHMT3 GFPF) and 5′-AGAAAGCTGGGTGAACGCCAGGAATGGGAAA-3′ (AtSHMT3 GFPR), and then reamplified with the attB adapter primer pair (Invitrogen). Resulting PCR fragments were purified with Wizard PCR columns, and then inserted into the pDNOR 221 vector (Invitrogen) using BP recombination. The cloned sequences were then fused to the N-terminus of the EGFP sequence segment in the p2GWF7 destination vector using LR recombination.

Leaf protoplasts were isolated from 4-week-old A. thaliana plants, and transformed with the EGFP-containing vectors following the procedure described in [33]. Fluorescence was monitored using an LSM 510 confocal laser-scanning microscope (Carl Zeiss MicroImaging). EGFP fluorescence was monitored with 488 nm excitation and 505–530 nm emission wavelengths, and chlorophyll fluorescence was monitored with 488 nm excitation and >650 nm emission.

Synthesis of H4PteGlu2-8

PteGlu2-8 were reduced to H4PteGlu2-8 with KBH4 [34]. The resulting H4PteGlu2-8 were purified on a Mono Q 5/50 GL column; bound folates were eluted with a linear gradient of 0.13–2.0 M sodium acetate (pH 6.9) plus 0.2 M 2-mercaptoethanol over 20 column volumes as described previously [35]. For use in enzyme activity assays, H4PteGlu2-8 were purified further by solid-phase extraction on a Chromabond C18 Hydra column (Macherey-Nagel). The column was washed with pure methanol and water before loading the samples; H4PteGlu2-8 were eluted with 10 mM THP. Fractions containing H4PteGlu2-8 were pooled, frozen in liquid N2 and then stored at −80 °C until use. H4PteGlu2-8 concentrations were determined spectrophotometrically at A298 [36].

SHMT activity assays

SHMT activity was measured using an HPLC-based fluorimetric assay as described previously [37]. Ches/Hepes/citric acid buffer (pH 8.5) replaced Tris/HCl in the assay mixture, as preliminary results indicated artefacts when assaying in Tris/HCl. Measurements were made using an Alliance 2695 separations module equipped with a 2675 fluorescence detector (Waters). Substrates were saturating, and product formation was proportional to time and enzyme concentration. Less than 10% of the substrates was typically consumed. Product formation was determined after subtraction of a blank, wherein serine was added after incubation.

Reaction products were separated by reversed-phase chromatography on a Waters XTerra C18 column (4.6 mm×100 mm, 5 μm particle size), and were measured by fluorescence detection with 280 nm excitation and 359 nm emission wavelengths. The mobile phase contained 27 mM phosphoric acid and 7, 9 or 10% methanol when substrates were H4PteGlu1-4, H4PteGlu5 or H4PteGlu6-8 respectively and the inhibitors were omitted; the length of the isocratic separation varied (6–15 min). Reaction products were separated isocratically when measuring inhibition with 5-CH3-H4PteGlu1&5. The mobile phase contained 27 mM phosphoric acid and 10% methanol or 4% acetonitrile for the assays with added 5-CH3-H4PteGlu1 or 5-CH3-H4PteGlu5 respectively. The length of separation was 9 min for 5-CH3-H4PteGlu1 and 20 min for 5-CH3-H4PteGlu5. Reaction products were separated under gradient conditions when measuring inhibition with 5-HCO-H4PteGlu1&5. The mobile phase contained 27 mM phosphoric acid and varied methanol content. For the assays with added 5-HCO-H4PteGlu1, methanol was held at 11% for 10 min, increased to 20% over 2 min and then decreased to 11% for 2 min. For the assays with added 5-HCO-H4PteGlu5, methanol was held at 9% for 13 min, increased to 20% over 2 min and then decreased to 9% for 2 min.

The 5-CH3-H4PteGlu1 standard was produced by reducing authentic 5,10-CH2-H4PteGlu1 using NaBH4 under conditions identical with those used for the assay products [37]. 5,10-CH2-H4PteGlu2-8 are commercially unavailable. Thus these assay products were quantified by first comparing the peak areas associated with the reduced products with that of the 5-CH3-H4PteGlu1 standard, and then multiplying those values by the ratio of the respective peak areas for 5-CH3-H4PteGlu2-5 and 5-CH3-H4PteGlu1. 5-CH3-H4PteGlu6-8 are commercially unavailable. Thus the 5-CH3-H4PteGlu6-8/5-CH3-H4PteGlu1 ratios were assumed to be 1 as deduced from the experimental data available for 5-CH3-H4PteGlu4&5. The concentrations of H4PteGlun, 5-CH3-H4PteGlun and 5,10-CH2-H4PteGlu1 were determined spectrophotometrically using published molar absorption coefficients [36].

Initial reaction rates were measured at pH 8.5 with various H4PteGlun concentrations and a constant serine concentration. To measure inhibition with 5-CH3-H4PteGlu1&5 or 5-HCO-H4PteGlu1&5, H4PteGlu6 was used as the substrate. Serine (5 mM) and H4PteGlu6 (2 and 4 μM) concentrations were held constant while the inhibitor concentrations were varied.

Apparent values for the kinetic parameters (Km, Ki, Vmax) were found by fitting measured initial reaction rates against substrate or substrate and inhibitor concentrations to suitable enzyme inhibition models in the Enzyme Kinetics Module 1.2 for SigmaPlot 9.0.1. The coefficient of determination, R2, helped single out the inhibition model (and derived parameters) that best fit one set of measurements. The standard error for kcat/Km and Ki/Km was calculated by error propagation [38].

In silico expression analysis

Gene expression was analysed in silico using the Meta Analyzer tool of the GENEVESTIGATOR software package (http://www.genevestigator.ethz.ch), and the publicly available Affymetrix microarray data for A. thaliana [39].

RESULTS

Bioinformatic sequence analysis

Protein sequences of representative dicotyledons (A. thaliana, P. trichocarpa and V. vinifera) and monocotyledons (O. sativa, Z. mays and B. distachyon), as well as a moss (P. patens) and two green algae (C. reinhardtii and O. lucimarinus) were retrieved from GenBank®, http://www.phytozome.net and http://www.brachypodium.org databases for use in the phylogenetic analysis (Figure 1). The complete genomes of the plant species used in this analysis have been published.

Phylogenetic tree of SHMTs from representative plant species

Figure 1
Phylogenetic tree of SHMTs from representative plant species

The sequence alignment for the phylogenetic tree included SHMT protein sequences of nine plant species: A. thaliana (At), P. trichocarpa (Pt), V. vinifera (Vv), O. sativa (Os), B. distachyon (Bd), Z. mays (Zm), P. patens (Pp), C. reinhardtii (Cr) and O. lucimarinus (Ol). Key to subcellular localization predicted within groups: Ia1, plastids; Ia2, cytosol; Ia3, cytosol; Ib, nucleus; IIa, plastids; IIb, mitochondria. The phylogenetic tree was made with MEGA 4.0. The neighbour-joining method included Poisson correction, complete deletion and (1000) bootstrap replication.

Figure 1
Phylogenetic tree of SHMTs from representative plant species

The sequence alignment for the phylogenetic tree included SHMT protein sequences of nine plant species: A. thaliana (At), P. trichocarpa (Pt), V. vinifera (Vv), O. sativa (Os), B. distachyon (Bd), Z. mays (Zm), P. patens (Pp), C. reinhardtii (Cr) and O. lucimarinus (Ol). Key to subcellular localization predicted within groups: Ia1, plastids; Ia2, cytosol; Ia3, cytosol; Ib, nucleus; IIa, plastids; IIb, mitochondria. The phylogenetic tree was made with MEGA 4.0. The neighbour-joining method included Poisson correction, complete deletion and (1000) bootstrap replication.

Overall, the phylogenetic analysis showed that SHMTs from the same plant species but putatively targeted to different subcellular compartments are less similar to each other than they are to SHMTs from other plant species putatively targeted to the same organelle. The SHMT protein sequences clustered into two main groups on the phylogenetic tree (Figure 1). The first group (group I) divided into two subgroups (Ia and Ib). Subgroup Ia comprised monocotyledon SHMTs with N-terminal extensions predicted as plastid targeting peptides (subgroup Ia2), as well as monocotyledon SHMTs (subgroup Ia1) and those of the remaining plant species (subgroup Ia3) with no N-terminal extensions and so predicted to localize in the cytosol.

The ZmSHMT4 protein sequence in subgroup Ia2 carries a putative plastid-targeting peptide at the N-terminus. The OsSHMT3 protein sequence in this subgroup is encoded by the Os12g0409000 (GenBank®) and Os12g22030 (http://www.phytozome.net) predicted genes. Their DNA sequences are identical except for the N-terminal extension encoding a putative plastid-targeting peptide, which is absent from the Os12g0409000 sequence, but present in the Os12g22030 sequence. The O. sativa EST (expressed sequence tag) database contains three ESTs (CT853747, CK063929 and CI307114) with the N-terminus present in the Os12g22030 sequence, and no ESTs with the N-terminus present in the Os12g0409000 sequence. Thus we think that the Os12g0409000 and Os12g22030 sequences represent the same gene, that the Os12g22030 sequence encodes the full-length plastid SHMT, and that, during annotation, the Os12g0409000 sequence was probably incorrectly spliced at the N-terminus. It is also possible that alternative splicing can produce mRNA and protein products encoded by the Os12g0409000 gene; we found, however, no support for this possibility upon analysing EST sequences. The BdSHMT3 protein sequence in subgroup Ia2 is encoded by the Bradi4g08100 predicted gene in the http://www.brachypodium.org database. The Bradi4g08100 sequence lacks the N-terminal extension predicted to encode the plastid-targeting peptide; we found, however, that the B. distachyon genome encodes such an extension upstream of the Bradi4g08100 sequence (see Supplementary Figure S1 at http://www.BiochemJ.org/bj/430/bj4300097add.htm). Thus we think that, during annotation, the Bradi4g08100 sequence was probably incorrectly spliced at the N-terminus. An alignment showing predicted protein sequences of putative plastid SHMTs from selected monocotyledons is shown in Supplementary Figure S1.

Subgroup Ib comprised SHMTs with N-terminal extensions generally not recognized by prediction programs as organellar-targeting peptides (results not shown). Members of subgroup Ib were also predicted to carry nuclear-targeting signals at the C-terminus; we therefore assigned putative nuclear localization to the proteins in subgroup Ib. Proteins from dicotyledons, monocotyledons and the moss were found in subgroups Ia and Ib. Only one protein for each species of algae was found in group I: OlSHMT3 from O. lucimarinus clustered with subgroup Ia; CrSHMT3 from C. reinhardtii shared roughly equal sequence similarity with both subgroups. The sequence distribution within group I suggests that plants evolved separate SHMTs for the nucleus and the cytosol after the evolution of algae, and before the evolution of mosses, and that plastid SHMTs in monocotyledons evolved within this clade from the cytosolic isoforms.

The second group (group II) comprised SHMTs with N-terminal extensions generally recognized by prediction programs as organellar-targeting peptides (results not shown). This group divided into two subgroups, with SHMTs predicted to localize either in plastids (IIa) or in mitochondria (IIb); this suggests that organellar SHMT isoforms evolved before the speciation of vascular plants. We found no SHMTs from monocotyledons that clustered with putative plastid SHMTs from the other plant species.

SHMT activity in chloroplasts

The above-described bioinformatic analysis predicting the existence of SHMTs in monocotyledon plastids contradicted a previous work reporting no SHMT activity in wheat chloroplasts [7]. Thus we examined SHMT activity in plastids of pea, a representative dicotyledon, and barley, a representative monocotyledon, with leaf extracts serving as positive assay controls. Chloroplasts of both species were found to have nearly equal levels of SHMT activity (Figure 2); this contradicts previous biochemical evidence [7] and supports the bioinformatic prediction for the occurrence of this activity in monocotyledon plastids.

SHMT, MTHFR, fumarase and glyceraldehyde-3-phosphate dehydrogenase activities in chloroplast and leaf extracts of pea and barley

Figure 2
SHMT, MTHFR, fumarase and glyceraldehyde-3-phosphate dehydrogenase activities in chloroplast and leaf extracts of pea and barley

Enzyme activity values from one of three independent determinations for representative enzyme activities. Assay conditions were as described in the Experimental section. The marker enzyme GAPDH was assayed to confirm identity of the isolated chloroplasts. The marker enzymes fumarase and MTHFR were assayed to evaluate contamination of the isolated chloroplasts with mitochondria and the cytosol respectively. Results are means+S.E.M. of triplicate determinations. GAPDH, glyceraldehyde-3-phosphate dehydrogenase; PC, pea chloroplasts; PL, pea leaves; BC, barley chloroplasts; BL, barley leaves.

Figure 2
SHMT, MTHFR, fumarase and glyceraldehyde-3-phosphate dehydrogenase activities in chloroplast and leaf extracts of pea and barley

Enzyme activity values from one of three independent determinations for representative enzyme activities. Assay conditions were as described in the Experimental section. The marker enzyme GAPDH was assayed to confirm identity of the isolated chloroplasts. The marker enzymes fumarase and MTHFR were assayed to evaluate contamination of the isolated chloroplasts with mitochondria and the cytosol respectively. Results are means+S.E.M. of triplicate determinations. GAPDH, glyceraldehyde-3-phosphate dehydrogenase; PC, pea chloroplasts; PL, pea leaves; BC, barley chloroplasts; BL, barley leaves.

Purification and biochemical characterization of AtSHMT3

Purification of recombinant AtSHMT3 is shown in Figure 3. Untagged recombinant enzymes were used in all subsequent work. AtSHMT3 was assayed for activity in the presence of various H4PteGlu1-8 concentrations and a constant serine concentration. Kinetic parameters derived from non-linear curve fitting to a model of uncompetitive substrate inhibition are presented in Table 1. Primary plots of steady-state kinetic data used to calculate the kinetic parameters are shown in Supplementary Figure S2 (at http://www.BiochemJ.org/bj/430/bj4300097add.htm). These data showed that the Km value decreases and the catalytic efficiency (kcat/Km) increases with n for H4PteGlu1-4, and that those two values remain nearly unchanged for H4PteGlu4-8. Increased polyglutamylation resulted in smaller relative decreases in the Ki values than in the Km values; the Ki/Km value increases approx. 18-fold for H4PteGlu2&3 and 54-fold for H4PteGlu4-8, compared with that for H4PteGlu1 (Table 1). A previous study in pea found increased affinity for folylpolyglutamate substrates by a mitochondrial SHMT for H4PteGlu1-3 [35]; however, the enzyme's Km values for H4PteGlu>2 were not determined because of the lack of sensitivity of the radioassay used. Thus a comparison of the kinetic properties of SHMTs from different subcellular compartments in plant cells awaits completion of their biochemical characterization. An SHMT activity assay we developed recently [37] may ease detailed biochemical characterization of other plant SHMTs because this assay is at least three orders of magnitude more sensitive than the standard radioassay [40] under comparable experimental conditions.

Purification of recombinant AtSHMT3 from E. coli cells

Figure 3
Purification of recombinant AtSHMT3 from E. coli cells

The recombinant enzyme was purified as described in the Experimental section. Samples were separated by SDS/PAGE on a 10% NuPAGE BisTris gel and stained with Coomassie Blue. Lane 1, 10 μg of crude E. coli extract expressing AtSHMT3; lane 2, 10 μg of partially purified tagged AtSHMT3 after Mono Q ion-exchange chromatography; lane 3, 2.5 μg of purified AtSHMT3 after a second passage on a Mono Q column; lane 4, 5 μg of molecular mass standard (sizes are indicated in kDa).

Figure 3
Purification of recombinant AtSHMT3 from E. coli cells

The recombinant enzyme was purified as described in the Experimental section. Samples were separated by SDS/PAGE on a 10% NuPAGE BisTris gel and stained with Coomassie Blue. Lane 1, 10 μg of crude E. coli extract expressing AtSHMT3; lane 2, 10 μg of partially purified tagged AtSHMT3 after Mono Q ion-exchange chromatography; lane 3, 2.5 μg of purified AtSHMT3 after a second passage on a Mono Q column; lane 4, 5 μg of molecular mass standard (sizes are indicated in kDa).

Table 1
Kinetic parameters of AtSHMT3 for H4PteGlu1-8

Kinetic parameters are best fits to a model of uncompetitive substrate inhibition (SigmaPlot 9.0.1). Assay conditions were as described in the Experimental section. Results are means±S.E.M. of three to six triplicate independent determinations.

Substrate Km (μM) Ki (μM) kcat (s−1kcat/Km (s−1·μM−1Ki/Km 
H4PteGlu1 217.73±3.35 85.2±2.3 15.8±0.7 0.07±0.00 0.39±0.01 
H4PteGlu2 13.30±0.78 93.9±4.0 7.6±0.7 0.57±0.06 7.06±0.51 
H4PteGlu3 3.06±0.03 22.0±0.3 8.7±0.2 2.84±0.07 7.19±0.12 
H4PteGlu4 0.83±0.03 15.7±1.6 4.3±0.2 5.18±0.31 18.92±2.05 
H4PteGlu5 0.64±0.07 13.3±2.2 3.5±0.4 5.47±0.87 20.78±4.12 
H4PteGlu6 0.69±0.05 15.0±1.4 3.5±0.4 5.07±0.69 21.74±2.57 
H4PteGlu7 0.67±0.11 13.9±3.4 3.8±0.2 5.67±0.98 20.75±6.11 
H4PteGlu8 0.55±0.04 11.8±2.2 3.3±0.6 6.00±1.17 21.45±4.29 
Substrate Km (μM) Ki (μM) kcat (s−1kcat/Km (s−1·μM−1Ki/Km 
H4PteGlu1 217.73±3.35 85.2±2.3 15.8±0.7 0.07±0.00 0.39±0.01 
H4PteGlu2 13.30±0.78 93.9±4.0 7.6±0.7 0.57±0.06 7.06±0.51 
H4PteGlu3 3.06±0.03 22.0±0.3 8.7±0.2 2.84±0.07 7.19±0.12 
H4PteGlu4 0.83±0.03 15.7±1.6 4.3±0.2 5.18±0.31 18.92±2.05 
H4PteGlu5 0.64±0.07 13.3±2.2 3.5±0.4 5.47±0.87 20.78±4.12 
H4PteGlu6 0.69±0.05 15.0±1.4 3.5±0.4 5.07±0.69 21.74±2.57 
H4PteGlu7 0.67±0.11 13.9±3.4 3.8±0.2 5.67±0.98 20.75±6.11 
H4PteGlu8 0.55±0.04 11.8±2.2 3.3±0.6 6.00±1.17 21.45±4.29 

Limited sensitivity of the standard radioassay is the most likely cause for the lack of published kinetic parameters for H4PteGlu>1 for SHMTs from other organisms. The exception is a study reporting kinetic parameters for H4PteGlu>1 for an SHMT from pig liver; the study reports a decrease in the Km value for H4PteGlun when n=1–3, 14-fold when n=1 or 2 and 2.3-fold when n=2 or 3 [41]. To increase understanding of how folylpolyglutamylation affects catalysis, earlier studies in mammalian SHMTs addressed the issue by determining the dissociation constants (Kd values) for binding of H4PteGlun to SHMT. Those studies found that the Kd value for H4PteGlun decreases when n=1–6 for an SHMT from rabbit liver mitochondria [42], and decreases when n=1–3 and remains constant when n=3–6 for an SHMT from the rabbit liver cytosol [43]. In an SHMT purified from pig liver, the Kd value decreases when n=1–3 and remains constant when n=3–7 [41]. Thus studies in two mammals suggest that increased H4PteGlun polyglutamylation affects affinity for these substrates in an enzyme-specific manner. Future work will establish whether this holds true for plant enzymes.

AtSHMT3 activity was inhibited by 5-CH3-H4PteGlu1&5 and 5-HCO-H4PteGlu1&5 (Table 2 and Supplementary Figure S3 at http://www.BiochemJ.org/bj/430/bj4300097add.htm). This is consistent with previous studies showing that 5-CH3-H4PteGlun species inhibit SHMTs from rabbit [17,18], and bind SHMTs from pig and E. coli [19,20]; and that 5-HCO-H4PteGlun species inhibit SHMTs from rabbit and pea mitochondria [17,18,30], and bind SHMTs from human and zebrafish [2123]. Furthermore, the pentaglutamate folates were more effective inhibitors than the monoglutamate folates (Table 2). This is consistent with previous studies showing that enzyme affinity for folylpolyglutamate inhibitors increases with the number of glutamate residues in mammalian SHMTs [18,19,21,44].

Table 2
Inhibition of AtSHMT3 with 5-CH3-H4PteGlu1&5 and 5-HCO-H4PteGlu1&5

Kinetic parameters are best fits to a model of mixed (partial) inhibition for 5-HCO-H4PteGlu1/5 and 5-CH3-H4PteGlu1, and to a model of mixed tight inhibition for 5-CH3-H4PteGlu5 (SigmaPlot 9.0.1). Inhibition by 5-CH3-H4PteGlu5 was biphasic; Ki and Vmax values for 5-CH3-H4PteGlu5 were determined from two phases of inhibition. Assay conditions were as described in the Experimental section. Results are means±S.E.M. of three to four triplicate independent determinations.

Inhibitor Ki (μM) 
5-CH3-H4PteGlu1 49.69±16.55 
5-CH3-H4PteGlu5 1.94±0.31/0.59±0.20 
5-HCO-H4PteGlu1 50.22±25.54 
5-HCO-H4PteGlu5 4.96±0.63 
Inhibitor Ki (μM) 
5-CH3-H4PteGlu1 49.69±16.55 
5-CH3-H4PteGlu5 1.94±0.31/0.59±0.20 
5-HCO-H4PteGlu1 50.22±25.54 
5-HCO-H4PteGlu5 4.96±0.63 

Transient expression of EGFP-fused proteins in A. thaliana protoplasts

Subcellular localization of AtSHMT3 was examined by fusing the full-length protein sequence to EGFP. Green fluorescence produced by expression of this fusion protein in A. thaliana protoplasts appeared in the form of granules that overlapped with the red autofluorescence of chloroplasts (Figure 4). The granular appearance of EGFP-fusion proteins in chloroplasts has been observed previously [45]. The cause for such an appearance is currently unclear. Green fluorescence was also visible throughout the cytoplasm in control protoplasts expressing EGFP alone (Figure 4). These findings thus establish that AtSHMT3 has a functional N-terminal peptide for its targeting to plastids. Consistently, AtSHMT3 has also been shown to localize in plastids by a proteomic approach [46].

Localization of AtSHMT3 in A. thaliana chloroplasts

Figure 4
Localization of AtSHMT3 in A. thaliana chloroplasts

C-terminal EGFP fusion to the full-length AtSHMT3 sequence (FL) was transiently expressed in A. thaliana protoplasts. EGFP expressed from the pUC18-GFP5T-sp plasmid was used as a control for targeting to the cytosol. Subcellular localization was analysed by fluorescence microscopy. (A and B) EGFP fluorescence; (C and D) chlorophyll autofluorescence; (E and F) merged images.

Figure 4
Localization of AtSHMT3 in A. thaliana chloroplasts

C-terminal EGFP fusion to the full-length AtSHMT3 sequence (FL) was transiently expressed in A. thaliana protoplasts. EGFP expressed from the pUC18-GFP5T-sp plasmid was used as a control for targeting to the cytosol. Subcellular localization was analysed by fluorescence microscopy. (A and B) EGFP fluorescence; (C and D) chlorophyll autofluorescence; (E and F) merged images.

In silico expression analysis

Organ- and development-specific expression of AtSHMT3 was studied in silico using publicly available microarray data and the GENEVESTIGATOR software package. AtSHMT3 was expressed in all plant organs and at all developmental stages examined; the expression level in germinated seeds was around twice as high as those in all other organs (Figure 5). These findings suggest that AtSHMT3 is a housekeeping enzyme needed by all plant organs throughout development.

In silico analysis of AtSHMT3 expression in A. thaliana

Figure 5
In silico analysis of AtSHMT3 expression in A. thaliana

Public Affymetrix expression analysis microarray data were analysed using the GENEVESTIGATOR reference expression database and meta-analysis system. GENEVESTIGATOR obtains the signal intensity values from raw array data by normalization using the Affymetrix MAS5 algorithm. Results are means±S.E.M.

Figure 5
In silico analysis of AtSHMT3 expression in A. thaliana

Public Affymetrix expression analysis microarray data were analysed using the GENEVESTIGATOR reference expression database and meta-analysis system. GENEVESTIGATOR obtains the signal intensity values from raw array data by normalization using the Affymetrix MAS5 algorithm. Results are means±S.E.M.

DISCUSSION

In plastids, SHMT catalytically directs the C1 moieties originating from serine into the metabolic network of H4PteGlunbound C1 units. The H4PteGlun-bound C1 moieties are then interconverted via reactions catalysed by a bifunctional 5,10methylenetetrahydrofolate dehydrogenase/5,10-methenyltetrahydrofolate cyclohydrolase [2] to provide C1 moieties for the biosynthesis of purines, thymidylate and N-formylmethionine [2,11,12]. Thus SHMT activity could potentially regulate important metabolic processes in plastids by regulating the influx of C1 units into the C1–H4PteGlun network. Before further studies can be attempted to find out the role of this enzyme in regulating metabolic fluxes, its presence in plastids needs to be confirmed and its catalytic properties need to be determined.

The first evidence of SHMT activity in dicotyledon plastids came from a study showing formation of [14C]serine in pea chloroplasts fed with [14C]formate [47], which suggested that formation of [14C]serine probably occurs via four reactions catalysed in sequence by 10-formyltetrahydrofolate synthetase, 5,10-methylenetetrahydrofolate dehydrogenase/5,10-methenyltetrahydrofolate cyclohydrolase and SHMT. An SHMT was subsequently isolated from spinach chloroplasts, but the corresponding gene remains to be cloned [1]. However, SHMT activity was not detected in wheat chloroplasts [7], raising the question of whether SHMTs exist in plastids of all flowering plants.

We found bioinformatic indications for the presence of genes encoding plastid SHMTs in monocotyledon genomes, and we detected SHMT activity in Percoll-isolated chloroplasts from pea (a representative dicotyledon) and barley (a representative monocotyledon) (Figure 2). These results provide evidence that monocotyledon plastids do contain SHMTs. Thus our results support the premise that plastids of monocotyledons and dicotyledons can draw C1 moieties from the endogenous serine pool.

Plastid folates are predominantly polyglutamylated, the most abundant forms being tetra-, penta- and hexa-glutamates [48,49]. We found decreases in the Km value and increases in the kcat/Km value of AtSHMT3 for H4PteGlu1-4, and no significant changes in those two values for H4PteGlu4-8. Thus if one assumes that, in plastids, the distribution of H4PteGlu1-8 species is nearly equal to that of the entire plastid folate pool reported previously [48,49], AtSHMT3 is then equally efficient at utilizing the three most abundant H4PteGlun species present in plastids (H4PteGlu4-6).

In A. thaliana chloroplasts, H4PteGlun was found to be below the detection limit of the assay used, and the total folate content was estimated to be approx. 40 pmol/mg of protein [50]. The protein concentration in chloroplast lumen and stroma in spinach was estimated to be >20 mg/ml [51]. Assuming that the protein concentrations in chloroplasts in A. thaliana and spinach have approximately equal values, we estimated that the total folate concentration in chloroplasts in A. thaliana is approx. 0.8 μM. Assuming further that H4PteGlun is <10% of the total folate pool in A. thaliana chloroplasts, we can then estimate that the H4PteGlun concentration in these organelles is <0.08 μM. A published value for folate concentration in pea chloroplasts is 1.7 μM [8]. On the basis of this value and considering that H4PteGlun is approx. 10% of the total folate pool [48], we estimated that the H4PteGlun concentration in pea chloroplasts is approx. 0.2 μM, which is close to that estimated for A. thaliana chloroplasts. Thus H4PteGlun concentration values in chloroplasts of both species are well below the Ki values we have determined for H4PteGlun (Table 1). Therefore inhibition of AtSHMT3 activity by H4PteGlun species probably lacks physiological significance.

Effects of polyglutamylation on the Km values of plant SHMTs have been studied only in an enzyme isolated from pea mitochondria [35]. The Km values for this enzyme are reported as 37.5 μM for H4PteGlu1, 13.5 μM for H4PteGlu2 and ≤3.7 μM for H4PteGlu3-6. The Km values for H4PteGlu≥3 could not be determined because of the limitations in assay sensitivity. Likewise, a decrease in Km values as polyglutamylation increases (56 μM for H4PteGlu1, 3.9 μM for H4PteGlu2 and 1.7 μM for H4PteGlu3) was found in an SHMT isolated from pig liver [41]. We are unaware of previous work reporting Km values of SHMTs for H4PteGlu>3, although Km values for H4PteGlu1 are known for several SHMTs [1,20,21,5256]. Neither are we aware of previous work reporting Ki values of SHMTs for H4PteGlun, although inhibition of SHMT activity by H4PteGlu1 has been described for enzymes from Crithidia fasciculata [55], Trypanosoma cruzi [56] and Leishmania donovani [57]. Thus the results in Table 1 represent, to our knowledge, the first set of kinetic parameters for H4PteGlu1-8 species of an SHMT from any organism.

5-CH3-H4PteGlun and 5-HCO-H4PteGlun are known to bind or inhibit SHMTs from several organisms [1723,30,52], and to be present in plastids [48,50]. 5-CH3-H4PteGlun is needed in plastids for methionine synthesis [58]. Current evidence supports the import of 5-CH3-H4PteGlu1 into plastids from the cytosol [2], where this folate is formed in a reaction catalysed by MTHFR [31], and supports its polyglutamylation in plastids, which is catalysed by a resident folylpolyglutamate synthetase [59]. 5-HCO-H4PteGlun is not known to act as a C1 donor, but only as an inhibitor of various enzymes. This folate can be formed from 5,10-CH+-H4PteGlun by catalysis in a side reaction of SHMT in the presence of glycine [21,60] or by chemical hydrolysis at acidic pH [61]. The stromal pH of illuminated chloroplasts is mildly basic [62], which is unfavourable for chemical formation of 5-HCO-H4PteGlun from 5,10-CH+-H4PteGlun. Thus the plastid 5-HCO-H4PteGlun pool is probably formed by catalysis. Applying the reasoning described above to estimate the H4PteGlun concentration, we arrived at ~0.22 μM 5-CH3-H4PteGlun and ~0.14 μM 5-HCO-H4PteGlun concentrations in chloroplasts in A. thaliana [50].

5-CH3-H4PteGlun and 5-HCO-H4PteGlun exist predominantly as tetra-, penta- and hexa-glutamates [48]. Accordingly, we consider that measuring inhibition of AtSHMT3 activity with H4PteGlu6 as a substrate, and 5-CH3-H4PteGlu5 or 5-HCO-H4PteGlu5 as an inhibitor provides physiologically relevant data as it utilizes substrate and inhibitor species that are abundant in plastids in vivo. Also, the difference between the Ki values obtained using the pentaglutamate inhibitors and those obtained using the monoglutamate inhibitors (Table 2), which are not abundant in plant cells, underscores the importance of assaying SHMTs using the polyglutamylated folate species.

By applying the model equations for mixed tight (5-CH3-H4PteGlun) or mixed (partial) (5-HCO-H4PteGlun) inhibition, we calculated that AtSHMT3 would be only approx. 10 and 3% inhibited respectively under the folate concentrations estimated for plastids. This calculation utilized the folate concentrations estimated above and the Ki values that we determined for 5-CH3-H4PteGlu5 (1.94 μM) and 5-HCO-H4PteGlu5 (4.96 μM), and considered that the Ki values for tetra-, penta- and hexa-glutamylated inhibitors are similar. Thus our data suggest that 5-CH3-H4PteGlun and 5-HCO-H4PteGlun do not significantly inhibit SHMT activity in plastids under physiological conditions, so they are probably not important as in vivo regulators of C1 metabolism in these organelles. Consistent with this finding, 5-formyltetrahydrofolate cycloligase, which eliminates 5-HCO-H4PteGlun from the folate pool by converting it into 5,10-CH+-H4PteGlun, is absent from plastids [30].

Abbreviations

     
  • 5-CH3-H4PteGlun

    5-methyltetrahydrofolate

  •  
  • 5,10-CH+-H4PteGlun

    5,10-methenyltetrahydrofolate

  •  
  • 5,10-CH2-H4PteGlun

    5,10-methylenetetrahydrofolate

  •  
  • 10-HCO-H4PteGlun

    10-formyltetrahydrofolate

  •  
  • Ches

    2-(N-cyclohexylamino)ethanesulfonic acid

  •  
  • EGFP

    enhanced green fluorescent protein

  •  
  • EST

    expressed sequence tag

  •  
  • MTHFR

    5,10-methylenetetrahydrofolate reductase

  •  
  • PLP

    pyridoxal 5′-phosphate

  •  
  • SHMT

    serine hydroxymethyltransferase

  •  
  • THP

    tris-(3-hydroxypropyl)phosphine

AUTHOR CONTRIBUTION

Yi Zhang participated in the purification and characterization of recombinant AtSHMT3, and in assaying enzyme activities in isolated plant organelles. He also conducted the transient expression of EGFP-fused proteins in A. thaliana protoplasts. Kehan Sun participated in assaying methylenetetrahydrofolate reductase activity. Francisco Sandoval isolated plant organelles. Katherine Santiago participated in purifying recombinant AtSHMT3. Sanja Roje cloned the AtSHMT3 cDNA. Yi Zhang and Sanja Roje analysed the data. Yi Zhang, Francisco Sandoval and Sanja Roje wrote the paper.

We thank Dr Matthew Willmann, University of Pennsylvania, Philadelphia, PA, U.S.A., for advice with the transformation of A. thaliana protoplasts. We also thank Dr Mechthild Tegeder, School of Biological Sciences, Washington State University, for providing the pUC18-GFP5T-sp plasmid; Craig Whitney and Julianna Gothard-Szamosfalvi for growing pea, barley and A. thaliana plants; and Dr Michael Knoblauch, Dr Christine Davitt and Dr Valerie Lynch-Holm from the Franceschi Microscopy and Imaging Center at Washington State University for help with the confocal laser-scanning microscope.

FUNDING

This work was supported by the National Science Foundation [grant number MCB-0429968 (to S.R.)] and by the Murdoch Foundation.

References

References
1
Besson
V.
Neuburger
M.
Rébeillé
F.
Douce
R.
Evidence for three serine hydroxymethyltransferases in green leaf cells: purification and characterization of the mitochondrial and chloroplastic isoforms
Plant Physiol. Biochem.
1995
, vol. 
33
 (pg. 
665
-
673
)
2
Hanson
A. D.
Gage
D. A.
Shachar-Hill
Y.
Plant one-carbon metabolism and its engineering
Trends Plant Sci.
2000
, vol. 
5
 (pg. 
206
-
213
)
3
Hanson
A. D.
Roje
S.
One-carbon metabolism in higher plants
Annu. Rev. Plant Physiol. Plant Mol. Biol.
2001
, vol. 
52
 (pg. 
119
-
137
)
4
McClung
C. R.
Hsu
M.
Painter
J. E.
Gagne
J. M.
Karlsberg
S. D.
Salomé
P. A.
Integrated temporal regulation of the photorespiratory pathway: circadian regulation of two Arabidopsis genes encoding serine hydroxymethyltransferase
Plant Physiol.
2000
, vol. 
123
 (pg. 
381
-
392
)
5
Ohyanagi
H.
Tanaka
T.
Sakai
H.
Shigemoto
Y.
Yamaguchi
K.
Habara
T.
Fujii
Y.
Antonio
B. A.
Nagamura
Y.
Imanishi
T.
, et al. 
The Rice Annotation Project Database (RAP-DB): hub for Oryza sativa ssp. japonica genome information
Nucleic Acids Res.
2006
, vol. 
34
 (pg. 
D741
-
D744
)
6
Turner
S. R.
Ireland
R.
Morgan
C.
Rawsthorne
S.
Identification and localization of multiple forms of serine hydroxymethyltransferase in pea (Pisum sativum) and characterization of a cDNA encoding a mitochondrial isoform
J. Biol. Chem.
1992
, vol. 
267
 (pg. 
13528
-
13534
)
7
Gardeström
P.
Edwards
G.
Henricson
D.
Ericson
I.
The localization of serine hydroxymethyltransferase in leaves of C3 and C4 species
Physiol. Plant.
1985
, vol. 
64
 (pg. 
29
-
33
)
8
Neuburger
M.
Rébeillé
F.
Jourdain
A.
Nakamura
S.
Douce
R.
Mitochondria are a major site for folate and thymidylate synthesis in plants
J. Biol. Chem.
1996
, vol. 
271
 (pg. 
9466
-
9472
)
9
Voll
L.
Jamai
A.
Renné
P.
Voll
H.
McClung
R.
Weber
A.
The photorespiratory Arabidopsis shm1 mutant is deficient in SHM1
Plant Physiol.
2006
, vol. 
140
 (pg. 
59
-
66
)
10
Jamai
A.
Salomé
P. A.
Schilling
S. H.
Weber
A. P.
McClung
C. R.
Arabidopsis photorespiratory serine hydroxymethyltransferase activity requires the mitochondrial accumulation of ferredoxin-dependent glutamate synthase
Plant Cell
2009
, vol. 
21
 (pg. 
595
-
606
)
11
Luo
M.
Orsi
R.
Patrucco
E.
Pancaldi
S.
Cella
R.
Multiple transcription start sites of the carrot dihydrofolate reductase-thymidylate synthase gene, and sub-cellular localization of the bifunctional protein
Plant Mol. Biol.
1997
, vol. 
33
 (pg. 
709
-
722
)
12
Zrenner
R.
Stitt
M.
Sonnewald
U.
Boldt
R.
Pyrimidine and purine biosynthesis and degradation in plants
Annu. Rev. Plant Biol.
2006
, vol. 
57
 (pg. 
805
-
836
)
13
Huang
Y.
Baxter
R.
Smith
B. S.
Partch
C. L.
Colbert
C. L.
Deisenhofer
J.
Crystal structure of cryptochrome 3 from Arabidopsis thaliana and its implications for photolyase activity
Proc. Natl. Acad. Sci. U.S.A.
2006
, vol. 
103
 (pg. 
17701
-
17706
)
14
Scott
J.
Rébeillé
F.
Fletcher
J.
Folic acid and folates: the feasibility for nutritional enhancement in plant foods
J. Sci. Food Agric.
2000
, vol. 
80
 (pg. 
795
-
824
)
15
Qi
H.
Atkinson
I.
Xiao
S.
Choi
Y. J.
Tobimatsu
T.
Shane
B.
Folylpoly-γ-glutamate synthetase: generation of isozymes and the role in one carbon metabolism and antifolate cytotoxicity
Adv. Enzyme Regul.
1999
, vol. 
39
 (pg. 
263
-
273
)
16
Cossins
E. A.
Chen
L.
Folates and one-carbon metabolism in plants and fungi
Phytochemistry
1997
, vol. 
45
 (pg. 
437
-
452
)
17
Schirch
L.
Ropp
M.
Serine transhydroxymethylase: affinity of tetrahydrofolate compounds for the enzyme and enzyme–glycine complex
Biochemistry
1967
, vol. 
6
 (pg. 
253
-
257
)
18
Stover
P.
Schirch
V.
5-Formyltetrahydrofolate polyglutamates are slow tight binding inhibitors of serine hydroxymethyltransferase
J. Biol. Chem.
1991
, vol. 
266
 (pg. 
1543
-
1550
)
19
Matthews
R. G.
Ross
J.
Baugh
C. M.
Cook
J. D.
Davis
L.
The role of folylpolyglutamates in the regulation of folate metabolism
Adv. Exp. Med. Biol.
1983
, vol. 
163
 (pg. 
35
-
44
)
20
Schirch
V.
Hopkins
S.
Villar
E.
Angelaccio
S.
Serine hydroxymethyltransferase from Escherichia coli: purification and properties
J. Bacteriol.
1985
, vol. 
163
 (pg. 
1
-
7
)
21
Fu
T. F.
Hunt
S.
Schirch
V.
Safo
M. K.
Chen
B. H.
Properties of human and rabbit cytosolic serine hydroxymethyltransferase are changed by single nucleotide polymorphic mutations
Arch. Biochem. Biophys.
2005
, vol. 
442
 (pg. 
92
-
101
)
22
Chang
W. N.
Tsai
J. N.
Chen
B. H.
Fu
T. F.
Cloning, expression, purification, and characterization of zebrafish cytosolic serine hydroxymethyltransferase
Protein Expression Purif.
2006
, vol. 
46
 (pg. 
212
-
220
)
23
Chang
W. N.
Tsai
J. N.
Chen
B. H.
Huang
H. S.
Fu
T. F.
Serine hydroxymethyltransferase isoforms are differentially inhibited by leucovorin: characterization and comparison of recombinant zebrafish serine hydroxymethyltransferases
Drug Metab. Dispos.
2007
, vol. 
35
 (pg. 
2127
-
2137
)
24
Tamura
K.
Dudley
J.
Nei
M.
Kumar
S.
MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0
Mol. Biol. Evol.
2007
, vol. 
24
 (pg. 
1596
-
1599
)
25
Sandoval
F. J.
Zhang
Y.
Roje
S.
Flavin nucleotide metabolism in plants: monofunctional enzymes synthesize FAD in plastids
J. Biol. Chem.
2008
, vol. 
283
 (pg. 
30890
-
30900
)
26
Newman
J.
Novel buffer systems for macromolecular crystallization
Acta Crystallogr. Sect. D Biol. Crystallogr.
2004
, vol. 
60
 (pg. 
610
-
612
)
27
Cline
K.
Import of proteins into chloroplasts. Membrane integration of a thylakoid precursor protein reconstituted in chloroplast lysates
J. Biol. Chem.
1986
, vol. 
261
 (pg. 
14804
-
14810
)
28
Brock
I. W.
Hazell
L.
Michl
D.
Nielsen
V. S.
Møller
B. L.
Herrmann
R. G.
Klösgen
R. B.
Robinson
C.
Precursors of one integral and five lumenal thylakoid proteins are imported by isolated pea and barley thylakoids: optimisation of in vitro assays
Plant Mol. Biol.
1993
, vol. 
23
 (pg. 
717
-
725
)
29
Schulz
A.
Knoetzel
J.
Scheller
H. V.
Mant
A.
Uptake of a fluorescent dye as a swift and simple indicator of organelle intactness: import-competent chloroplasts from soil-grown Arabidopsis
J. Histochem. Cytochem.
2004
, vol. 
52
 (pg. 
701
-
704
)
30
Roje
S.
Janave
M. T.
Ziemak
M. J.
Hanson
A. D.
Cloning and characterization of mitochondrial 5-formyltetrahydrofolate cycloligase from higher plants
J. Biol. Chem.
2002
, vol. 
277
 (pg. 
42748
-
42754
)
31
Roje
S.
Wang
H.
McNeil
S. D.
Raymond
R. K.
Appling
D. R.
Shachar-Hill
Y.
Bohnert
H. J.
Hanson
A. D.
Isolation, characterization, and functional expression of cDNAs encoding NADH-dependent methylenetetrahydrofolate reductase from higher plants
J. Biol. Chem.
1999
, vol. 
274
 (pg. 
36089
-
36096
)
32
Bradford
M. M.
A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding
Anal. Biochem.
1976
, vol. 
72
 (pg. 
248
-
254
)
33
Sheen
J.
Signal transduction in maize and Arabidopsis mesophyll protoplasts
Plant Physiol.
2001
, vol. 
127
 (pg. 
1466
-
1475
)
34
Scrimgeour
K. G.
Vitols
K. S.
The reduction of folate by borohydride
Biochemistry
1966
, vol. 
5
 (pg. 
1438
-
1443
)
35
Besson
V.
Rébeillé
F.
Neuburger
M.
Douce
R.
Cossins
E. A.
Effects of tetrahydrofolate polyglutamates on the kinetic parameters of serine hydroxymethyltransferase and glycine decarboxylase from pea leaf mitochondria
Biochem. J.
1993
, vol. 
292
 (pg. 
425
-
430
)
36
Temple
C.
Montgomery
J. A.
Blakley
R. L.
Benkovic
S. J.
Chemical and physical properties of folic acid and reduced derivatives
Folates and Pterins
1984
New York
John Wiley & Sons
(pg. 
79
-
80
)
37
Zhang
Y.
Sun
K.
Roje
S.
An HPLC-based fluorometric assay for serine hydroxymethyltransferase
Anal. Biochem.
2008
, vol. 
375
 (pg. 
367
-
369
)
38
Meyer
S. L.
Propagation of errors and least squares
Data Analysis for Scientists and Engineers
1975
New York
John Wiley & Sons
(pg. 
39
-
48
)
39
Zimmermann
P.
Hirsch-Hoffmann
M.
Hennig
L.
Gruissem
W.
GENEVESTIGATOR: Arabidopsis microarray database and analysis toolbox
Plant Physiol.
2004
, vol. 
136
 (pg. 
2621
-
2632
)
40
Taylor
R. T.
Weissbach
H.
Radioactive assay for serine transhydroxymethylase
Anal. Biochem.
1965
, vol. 
13
 (pg. 
80
-
84
)
41
Matthews
R. G.
Ross
J.
Baugh
C. M.
Cook
J. D.
Davis
L.
Interactions of pig liver serine hydroxymethyltransferase with methyltetrahydropteroylpolyglutamate inhibitors and with tetrahydropteroylpolyglutamate substrates
Biochemistry
1982
, vol. 
21
 (pg. 
1230
-
1238
)
42
Strong
W. B.
Cook
R.
Schirch
V.
Interaction of tetrahydropteroylpolyglutamates with two enzymes from mitochondria
Biochemistry
1989
, vol. 
28
 (pg. 
106
-
114
)
43
Strong
W. B.
Schirch
V.
In vitro conversion of formate to serine: effect of tetrahydropteroylpolyglutamates and serine hydroxymethyltransferase on the rate of 10-formyltetrahydrofolate synthetase
Biochemistry
1989
, vol. 
28
 (pg. 
9430
-
9439
)
44
Huang
T.
Wang
C.
Maras
B.
Barra
D.
Schirch
V.
Thermodynamic analysis of the binding of the polyglutamate chain of 5-formyltetrahydropteroylpolyglutamates to serine hydroxymethyltransferase
Biochemistry
1998
, vol. 
37
 (pg. 
13536
-
13542
)
45
Narayana Murthy
U. M.
Ollagnier-de-Choudens
S.
Sanakis
Y.
Abdel-Ghany
S. E.
Rousset
C.
Ye
H.
Fontecave
M.
Pilon-Smits
E. A.
Pilon
M.
Characterization of Arabidopsis thaliana SufE2 and SufE3: functions in chloroplast iron-sulfur cluster assembly and NAD synthesis
J. Biol. Chem.
2007
, vol. 
282
 (pg. 
18254
-
18264
)
46
Zybailov
B.
Rutschow
H.
Friso
G.
Rudella
A.
Emanuelsson
O.
Sun
Q.
van Wijk
K. J.
Sorting signals, N-terminal modifications and abundance of the chloroplast proteome
PLoS ONE
2008
, vol. 
3
 pg. 
e1994
 
47
Shingles
R.
Woodrow
L.
Grodzinski
B.
Effects of glycolate pathway intermediates on glycine decarboxylation and serine synthesis in pea (Pisum sativum L.)
Plant Physiol.
1984
, vol. 
74
 (pg. 
705
-
710
)
48
Orsomando
G.
Díaz de la Garza
R. D.
Green
B. J.
Peng
M.
Rea
P. A.
Ryan
T. J.
Gregory
J. F.
3rd
Hanson
A. D.
Plant γ-glutamyl hydrolases and folate polyglutamates: characterization, compartmentation, and co-occurrence in vacuoles
J. Biol. Chem.
2005
, vol. 
280
 (pg. 
28877
-
28884
)
49
Imeson
H.
Zheng
L.-l.
Cossins
E.
Folylpolyglutamate derivatives of Pisum sativum L. Determination of polyglutamate chain lengths by high performance liquid chromatography following conversion to p-aminobenzoylpolyglutamates
Plant Cell Physiol.
1990
, vol. 
31
 (pg. 
223
-
231
)
50
Klaus
S. M.
Kunji
E. R.
Bozzo
G. G.
Noiriel
A.
Díaz de la Garza
R. D.
Basset
G. J.
Ravanel
S.
Rébeillé
F.
Gregory
J. F.
3rd
Hanson
A. D.
Higher plant plastids and cyanobacteria have folate carriers related to those of trypanosomatids
J. Biol. Chem.
2005
, vol. 
280
 (pg. 
38457
-
38463
)
51
Kieselbach
T.
Hagman
A.
Andersson
B.
Schröder
W. P.
The thylakoid lumen of chloroplasts: isolation and characterization
J. Biol. Chem.
1998
, vol. 
273
 (pg. 
6710
-
6716
)
52
Mitchell
M. K.
Reynolds
P. H. S.
Blevins
D. G.
Serine hydroxymethyltransferase from soybean root nodules: purification and kinetic properties
Plant Physiol.
1986
, vol. 
81
 (pg. 
553
-
557
)
53
Strong
W. B.
Tendler
S. J.
Seither
R. L.
Goldman
I. D.
Schirch
V.
Purification and properties of serine hydroxymethyltransferase and C1-tetrahydrofolate synthase from L1210 cells
J. Biol. Chem.
1990
, vol. 
265
 (pg. 
12149
-
12155
)
54
Kruschwitz
H.
Ren
S. L.
Disalvo
M.
Schirch
V.
Expression, purification, and characterization of human cytosolic serine hydroxymethyltransferase
Protein Expression Purif.
1995
, vol. 
6
 (pg. 
411
-
416
)
55
Capelluto
D. G.
Purification and partial characterization of three isoforms of serine hydroxymethyltransferase from Crithidia fasciculata
Mol. Biochem. Parasitol.
1999
, vol. 
98
 (pg. 
187
-
201
)
56
Capelluto
D. G.
Hellman
U.
Cazzulo
J. J.
Cannata
J. J.
Purification and some properties of serine hydroxymethyltransferase from Trypanosoma cruzi
Eur. J. Biochem.
2000
, vol. 
267
 (pg. 
712
-
719
)
57
Vatsyayan
R.
Roy
U.
Molecular cloning and biochemical characterization of Leishmania donovani serine hydroxymethyltransferase
Protein Expression Purif.
2007
, vol. 
52
 (pg. 
433
-
440
)
58
Ravanel
S.
Block
M. A.
Rippert
P.
Jabrin
S.
Curien
G.
Rébeillé
F.
Douce
R.
Methionine metabolism in plants: chloroplasts are autonomous for de novo methionine synthesis and can import S-adenosylmethionine from the cytosol
J. Biol. Chem.
2004
, vol. 
279
 (pg. 
22548
-
22557
)
59
Ravanel
S.
Cherest
H.
Jabrin
S.
Grunwald
D.
Surdin-Kerjan
Y.
Douce
R.
Rébeillé
F.
Tetrahydrofolate biosynthesis in plants: molecular and functional characterization of dihydrofolate synthetase and three isoforms of folylpolyglutamate synthetase in Arabidopsis thaliana
Proc. Natl. Acad. Sci. U.S.A.
2001
, vol. 
98
 (pg. 
15360
-
15365
)
60
Stover
P.
Schirch
V.
Enzymatic mechanism for the hydrolysis of 5,10-methenyltetrahydropteroylglutamate to 5-formyltetrahydropteroylglutamate by serine hydroxymethyltransferase
Biochemistry
1992
, vol. 
31
 (pg. 
2155
-
2164
)
61
Baggott
J. E.
Hydrolysis of 5,10-methenyltetrahydrofolate to 5-formyltetrahydrofolate at pH 2.5 to 4.5
Biochemistry
2000
, vol. 
39
 (pg. 
14647
-
14653
)
62
Heldt
W. H.
Werdan
K.
Milovancev
M.
Geller
G.
Alkalization of the chloroplast stroma caused by light-dependent proton flux into the thylakoid space
Biochim. Biophys. Acta
1973
, vol. 
314
 (pg. 
224
-
241
)

Supplementary data