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J. Biol. Chem., Vol. 276, Issue 47, 43580-43588, November 23, 2001

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Pathway for the Synthesis of Mannosylglycerate in the Hyperthermophilic Archaeon Pyrococcus horikoshii

BIOCHEMICAL AND GENETIC CHARACTERIZATION OF KEY ENZYMES^*

Nuno Empadinhas^§¶, Joey D. Marugg^§, Nuno Borges, Helena Santos^**, and Milton S. da Costa^§

From the  Instituto de Tecnologia Química e Biológica, Universidade Nova de Lisboa, Rua da Quinta Grande 6, Apartado 127, 2780-156 Oeiras, Portugal, the ^§ Departamento de Bioquímica, Universidade de Coimbra, 3000 Coimbra, Portugal, and  Nestlé Research Center, CH-1000 Lausanne 26, Switzerland

Received for publication, August 21, 2001, and in revised form, September 18, 2001

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The biosynthetic pathway for the synthesis of the compatible solute -mannosylglycerate in the hyperthermophilic archaeon Pyrococcus horikoshii is proposed based on the activities of purified recombinant mannosyl-3-phosphoglycerate (MPG) synthase and mannosyl-3-phosphoglycerate phosphatase. The former activity was purified from cell extracts, and the N-terminal sequence was used to identify the encoding gene in the completely sequenced P. horikoshii genome. This gene, designated PH0927, and a gene immediately downstream (PH0926) were cloned and overexpressed in Escherichia coli. The recombinant product of gene PH0927 catalyzed the synthesis of -mannosyl-3-phosphoglycerate (MPG) from GDP-mannose and D-3-phosphoglycerate retaining the configuration about the anomeric carbon, whereas the recombinant gene product of PH0926 catalyzed the dephosphorylation of mannosyl-3-phosphoglycerate to yield the compatible solute -mannosylglycerate. The MPG synthase and the MPG phosphatase were specific for these substrates. Two genes immediately downstream from mpgs and mpgp were identified as a putative bifunctional phosphomannose isomerase/mannose-1-phosphate-guanylyltransferase (PH0925) and as a putative phosphomannose mutase (PH0923). Genes PH0927, PH0926, PH0925, and PH0923 were contained in an operon-like structure, leading to the hypothesis that these genes were under the control of an unknown osmosensing mechanism that would lead to -mannosylglycerate synthesis. Recombinant MPG synthase had a molecular mass of 45,208 Da, a temperature for optimal activity between 90 and 100 °C, and a pH optimum between 6.4 and 7.4; the recombinant MPG phosphatase had a molecular mass of 27,958 Da and optimum activity between 95 and 100 °C and between pH 5.2 and 6.4. This is the first report of the characterization of MPG synthase and MPG phosphatase and the elucidation of a pathway for the synthesis of mannosylglycerate in an archaeon.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The vast majority of microorganisms capable of osmotic adjustment to environmental alterations in salt levels accumulate small molecular weight organic solutes, termed compatible solutes or osmolytes, to maintain a positive turgor pressure and to protect enzymes from desiccation (1, 2). The accumulation of compatible solutes can be accomplished by specific uptake or by de novo synthesis of osmolytes. The uptake of organic solutes such as glycine betaine and trehalose, among other solutes, from the environment is preferred because it is energetically favorable (3). Many organisms, however, synthesize their own compatible solutes because specific solutes in the environment may not be freely available or do not fulfill the prerequisites of an osmolyte.

Ectoine, hydroxyectoine, glycine betaine, trehalose, and glutamate are probably among the most common compatible solutes of bacteria and archaea (4). However, slightly halophilic thermophilic and hyperthermophilic bacteria and archaea generally accumulate compatible solutes that are rare or unknown in mesophilic organisms. Moreover, these osmolytes usually have a negative charge that is neutralized by the accumulation of potassium (5-7). The archetypal compatible solutes of thermophiles and hyperthermophiles such as di-myo-inositol phosphate (DIP),¹ dimannosyl-di-myo-inositol phosphate, diglycerol phosphate, mannosylglyceramide, and mannosylglycerate (MG) have not been found in mesophilic bacteria and archaea. Mannosylglycerate appears to be a very common compatible solute in thermophilic and hyperthermophilic organisms, namely in Pyrococcus furiosus, the slightly halophilic Thermococcus spp., Aeropyrum pernix, Thermus thermophilus, Rhodothermus marinus, and Petrotoga miotherma (6, 8-11). Furthermore, some of the compatible solutes of thermophiles and hyperthermophiles, of which MG deserves special mention, have been shown to protect enzymes in vitro against thermal denaturation and could also have an important role in thermoprotection of cell components in vivo (12, 13).

Basic knowledge of the biosynthesis of specific compatible solutes is needed to understand the mechanisms underlying the events leading to salt and thermal tolerance, from water stress sensing to maintenance of the appropriate intracellular levels of compatible solutes (14). The pathways for the synthesis of osmolytes in thermophilic and hyperthermophilic organisms have only recently begun to be examined. Two pathways for the biosynthesis of MG exist in the thermophilic bacterium R. marinus. One pathway involves the single step conversion of GDP-mannose and D-glycerate to MG by mannosylglycerate synthase. An alternative pathway was also detected in R. marinus that leads to the synthesis of the phosphorylated intermediate, mannosyl-3-phosphoglycerate which, in turn, is converted to MG by a phosphatase (15). We deemed it important to investigate whether a similar strategy would hold in hyperthermophilic archaea, and the slightly halophilic archaeon Pyrococcus horikoshii (16) was selected for this purpose.

We identify the genes involved in the synthesis of MG in P. horikoshii and elucidate a pathway for the biosynthesis of this compatible solute, which involves a mannosyl-3-phosphoglycerate synthase (MPG synthase) and a specific mannosyl-3-phosphoglycerate phosphatase (MPG phosphatase). In addition, the genes for the synthesis of MG in P. horikoshii were cloned and overexpressed in Escherichia coli, and the recombinant enzymes were characterized in detail.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Strains, Plasmids, and Culture Conditions

The type strain of P. horikoshii (JCM 9974) was obtained from the Japanese Collection of Microorganisms (JCM), Saitama, Japan. The organism was cultivated as follows: the medium contained per liter, 5.0 g of peptone, 1.0 g of yeast extract, 25.0 g of NaCl, 1.0 mg of FeSO₄·7H₂O, 40.0 mg of KH₂PO₄, 19.6 ml of magnesium salts solution (180.0 g of MgSO₄·7H₂O and 140.0 g of MgCl₂·6H₂O/liter), 1.0 ml of solution A (4.0 g of trisodium citrate, 9.0 g of MnSO₄·4H₂O, 2.5 g of ZnSO₄·7H₂O, 2.5 g of NiCl₂·6H₂O, 0.3 g of AlK(SO₄)₂·12H₂O, 0.3 g of CoCl₂·6H₂O, 0.15 g of CuSO₄·5H₂O/liter), 2.0 ml of solution B (56.0 g of CaCl₂·2H₂O, 25.0 g of NaBr, 16.0 g of KCl, 10.0 g of KI, and 4.0 g of SrCl₂·6H₂O/liter), 2.0 ml of solution C (50.0 g of K₂HPO₄, 7.5 g of H₃BO₃, 3.3 g of Na₂WO₄·2H₂O, 0.15 g of Na₂MoO₄·2H₂O, and 0.005 g of Na₂SeO₃/liter), and 20.0 ml of solution D (0.1 g of citric acid, 0.75 g of nitrilotriacetic acid, 0.06 g of CoCl₂·6H₂O, 14.5 g of KCl, and 32.0 g of CaCl₂·2H₂O/liter). The final pH was adjusted to 7.0. The medium was gassed with N₂ and sterilized by autoclaving. Sterile sulfur (3.0 g/l) was added to the medium after cooling to 90 °C. Cultures were grown in a 5-liter fermentor at 98 °C with continuous gassing of N₂ and stirred at 80 rpm.

To examine the effect of osmotic stress on the synthesis of intracellular solutes by P. horikoshii, 15-45 g of NaCl/liter was added to the medium. Biomass production for enzyme purification was carried out in medium containing 4.5% NaCl (w/v). Cell growth was monitored by measuring the turbidity at 600 nm.

E. coli XL1-Blue was used as host for expression vectors pTRC99A, pKK223-3 (Amersham Pharmacia Biotech), and for a plasmid isolated from E. coli strain BL21-CodonPlus (Stratagene) that carries extra tRNA genes for codons commonly found in archaea but rarely used by E. coli. This organism was grown in YT medium, at pH 7.0 and 37 °C, containing 10.0 g of tryptone, 5.0 g of yeast extract, and 5.0 g of NaCl/liter. Ampicillin was added at a final concentration of 100 µg/ml for selection of plasmids pKK223-3 and pTRC99A. Chloramphenicol was added to a final concentration of 20 µg/ml for selection of plasmid carrying the tRNA genes. IPTG was obtained from Roche Molecular Biochemicals and added at a final concentration of 1 mM.

Extraction and Determination of Intracellular Solutes

Cells of P. horikoshii were harvested during mid-exponential phase of growth by centrifugation (7000 × g, 10 min, 4 °C) and washed once with a NaCl solution identical in concentration to that of the medium where the cells were grown. Cell pellets were extracted twice with boiling 80% ethanol as described previously (6). For solute quantification by ¹H NMR, the final pH of the samples was adjusted to ~8 by the addition of NaO²H. The protein content of the cells was determined by the Bradford assay (17) after lysis with 1 N NaOH (100 °C, 10 min) and neutralization with 1 N HCl.

Preparation of Cell-free Extracts

Cells were harvested by centrifugation (7000 × g, 10 min, 4 °C) during the late exponential phase of growth. The cell pellet was resuspended in Tris-HCl (20 mM (pH 7.6)) containing MgCl₂ (5 mM), DNase I (10 µg), and protease inhibitors, phenylmethylsulfonyl fluoride (80 µg), leupeptin (20 µg), and antipain (20 µg) per ml of the suspension. Cells were disrupted in a French press, followed by centrifugation (130,000 × g, 1 h, 4 °C). The supernatant was dialyzed against 20 mM Tris-HCl (pH 7.6) to remove endogenous mannosylglycerate (MG) and other low molecular weight compounds prior to measuring enzyme activities and purification procedures.

Enzyme Assays

To determine the combined activity of mannosyl-3-phosphoglycerate synthase (MPG synthase)/mannosyl-3-phosphoglycerate phosphatase (MPG phosphatase) in the cell extracts, the reaction mixture contained 2.5 mM GDP-mannose (Sigma) and 2.5 mM D-3-phosphoglycerate (sodium salt, Sigma) in 20 mM Tris-HCl (pH 7.6) with 10 mM MgCl₂. The reaction mixtures were incubated at 90 °C for 30 min, and the MG produced was quantified by ¹H NMR after freeze-drying and dissolving in ²H₂O. Formate was used as an internal concentration standard. This protocol was also used to examine the kinetic parameters, temperature, and pH optima, the effect of NaCl, KCl, and divalent cations, and thermal stability of the recombinant MPG synthase, but in this case MPG was the final product that was quantified by ¹H NMR.

To detect the presence of MPG synthase, during the purification of the native enzyme, the same procedure was followed, but after incubation at 90 °C, unless otherwise stated, for 30 min, the reaction mixture was cooled to 37 °C, and 2 units of alkaline phosphatase (Sigma) was added. The mixture was incubated for an additional 30 min at 37 °C, and the formation of MG was visualized by TLC.

The activity of the purified recombinant MPG phosphatase was measured with 2 mM MPG in 20 mM Tris-HCl (pH 7.6) containing 10 mM MgCl₂ at 98 °C by monitoring the release of inorganic phosphate using the spectrophotometric method described by Ames (18). All enzyme parameters of MPG phosphatase were examined using this reaction mixture, unless otherwise stated.

MPG used for these assays was obtained from a reaction catalyzed by pure recombinant MPG synthase with 12.5 mM GDP-mannose, 12.5 mM D-3-phosphoglycerate as substrates, in 20 mM Tris-HCl (pH 7.6) and 10 mM MgCl₂ at 98 °C for 15-20 min. Quantification of MPG was carried out by incubating an aliquot of the reaction mixture with MPG phosphatase for 15-20 min to ensure complete dephosphorylation of MPG, in 20 mM Tris-HCl (pH 7.6) with 10 mM MgCl₂. Inorganic phosphate released was quantified by the Ames method. The concentration of MPG in the reaction mixture was calculated from the concentration of inorganic phosphate produced. All reactions were stopped by freezing in liquid nitrogen.

Purification of Mannosyl-3-phosphoglycerate

MPG was partially purified from the reaction mixtures described above. Samples were loaded onto a QAE-Sephadex A-25 column previously equilibrated with 5.0 mM sodium bicarbonate (pH 8.0), and the elution was performed with 1 bed volume of the same buffer, followed by a linear gradient of 5.0 mM to 1 M NaHCO₃. The eluted fractions were analyzed for carbohydrate by the method of Dubois et al. (19). MPG was eluted at 0.5 M NaHCO₃. For a second chromatographic step, a column of activated Dowex 50W-X8 resin was used, and the elution was carried out with distilled water. Subsequently, the fractions were pooled and degassed under vacuum, and the pH was raised to 3.5 with 1.0 M KOH. Samples were lyophilized and dissolved in ²H₂O prior to NMR analysis.

NMR Spectroscopy

The identification of the phosphorylated intermediate, -mannosyl-3-phosphoglycerate, was accomplished using ¹³C, ³¹P, and ¹H NMR spectroscopy. Spectra were recorded on Bruker AMX300 or DRX500 spectrometers (9, 11). For quantification of substrates and products of enzyme reaction mixtures, ¹H NMR spectra were acquired on a Bruker AMX300 spectrometer with a broadband inverse probe head with presaturation of the water. Spectra were acquired with a repetition delay of 45 s, and formate was used as an internal concentration standard. Proton, carbon, and phosphorus chemical shifts are relative to 3-(trimethylsilyl)propanesulfonic acid, methanol (at 49.3 ppm), or 85% H₃PO₄, respectively.

Analysis of MG Formation by Thin Layer Chromatography

TLC was performed on Silica Gel 60 plates (Merck) with a solvent system composed of chloroform, methanol, acetic acid, and water (30:50:8:4, v/v). MG was visualized by spraying with -naphtholsulfuric acid solution followed by charring at 120 °C (20). Authentic standards of MG, mannose, guanosine, and GDP-mannose were used for comparative purposes.

Purification of Native MPG Synthase

Native enzyme was purified by fast protein liquid chromatography (Amersham Pharmacia Biotech) at room temperature from P. horikoshii cell extracts.

Ion-exchange Chromatography-- The cell-free extract was applied to a column (XK50/30, bed volume 250 ml) packed with DEAE-Sepharose fast flow equilibrated with Tris-HCl (20 mM (pH 7.6)). All the purification steps were carried out at pH 7.6. Elution was carried out with a two-step linear NaCl gradient (0.0-0.6 and 0.6-1.0 M) in the same buffer. MPG synthase activity was found in the fraction eluting between 0.4 and 0.5 M NaCl. Active fractions were pooled, concentrated, and equilibrated with 20 mM Tris-HCl. The sample was applied to a Q-Sepharose fast flow column equilibrated with the same buffer. Elution was carried out with a five-step discontinuous NaCl gradient (0.2, 0.4, 0.6, 0.8, and 1.0 M). The fractions eluting at 0.4 and 0.6 M NaCl contained MPG synthase activity. Fractions with MPG synthase activity were pooled, concentrated, and equilibrated with 20 mM Tris-HCl. This sample was applied to a 6-ml Resource Q column. Elution was carried out with a linear NaCl gradient (0.0-1.0 M). Fractions eluted between 0.29 and 0.34 M contained MPG synthase activity.

Gel Filtration Chromatography-- The active fractions were pooled and concentrated by ultrafiltration (30-kDa cutoff) and were applied to a Superdex 200 column equilibrated with 0.2 M NaCl in 50 mM Tris-HCl. The active MPG synthase fractions were concentrated by ultrafiltration (10-kDa cutoff) and applied to a Superose column equilibrated with 0.2 M NaCl in 50 mM Tris-HCl. The purity of the active fraction was assessed by SDS-PAGE (21). The sample was blotted on polyvinylidene difluoride membranes followed by N-terminal amino acid sequencing at Microchemical Facility, Emory University School of Medicine, GA.

DNA Techniques, Analysis, Cloning, and Functional Overexpression of mpgs (PH0927) and mpgp (PH0926) in E. coli

Most DNA manipulations followed standard molecular techniques and procedures (22). P. horikoshii chromosomal DNA was isolated according to Marmur (23). Based on the N-terminal amino acid sequence of the purified MPG synthase, a corresponding open reading frame (ORF), designated PH0927, was identified from the P. horikoshii OT3 complete genome sequence (24). ORF sequences surrounding PH0927 were also screened for homologies with known phosphatase genes using the (T)FASTA (25) and (T)BLAST (26) algorithms. PCR amplifications were carried out in a PerkinElmer Life Sciences GeneAmp PCR System 2400 in reaction mixtures (50 µl) containing 100 ng of P. horikoshii DNA, 100 ng of each primer, 10 mM Tris-HCl (pH 9.0), 1.5 mM MgCl₂, 50 mM KCl, 1 unit of Pwo DNA polymerase, and 0.2 mM of each deoxynucleoside triphosphate (Amersham Pharmacia Biotech). The mixture was preincubated for 5 min at 95 °C and then subjected to 30 cycles of denaturation at 95 °C for 1 min. Annealing was performed at 60 °C for 1 min, and primer extension was at 72 °C for 2 min. The extension reaction in the last cycle was prolonged for 5 min. Amplification products were purified from agarose gels (Bio-Rad).

Based on the complete gene sequence, mpgs was amplified by the forward primer (5'-GCGCCATGGTTCTAGAAGCTCC-3') and adding a recognition sequence for NcoI (underlined) including the ATG start codon. The reverse primer (5'-GCGCTGCAGTCATAGCTCAAACCTCAG-3') was constructed by adding an additional PstI recognition sequence (underlined) directly behind the TGA stop codon. Gene mpgp was amplified with the forward primer (5'-GCGGAATTCATGATTAGGTTAATATTC-3') constructed with an additional EcoRI recognition sequence (underlined) immediately upstream of the ATG start codon, and the reverse primer (5'-GCGCTGCAGTCATTTGATCACCTCC-3') with an additional PstI recognition sequence (underlined) directly behind the TGA stop codon. The PCR products were purified after digestion with NcoI and PstI for mpgs and with EcoRI and PstI for mpgp and ligated into corresponding sites of expression vectors pTRC99A and pKK223-3 to obtain plasmids pMPGS and pMPGP, respectively. Each construction was transformed into E. coli XL1-Blue cells previously transformed with the plasmid carrying tRNA genes for rare codons. Host cells containing pMPGS or pMPGP were grown to mid-exponential growth phase (A₆₀₀ = 0.6), induced with IPTG, and grown further for 6-8 h. Cells were harvested and treated as described above for the preparation of cell-free extracts.

Purification of Recombinant MPG Synthase

E. coli cell extracts containing MPG synthase were incubated for 20 min at 80 °C to denature the majority of the host proteins and centrifuged (25000 × g, 15 min, 4 °C). MPG synthase activity assay was performed as described above, and the enzyme was purified.

Ion-exchange Chromatography-- The supernatant was applied to a DEAE-Sepharose fast flow column (XK50/30), equilibrated with 20 mM Tris-HCl (pH 7.6). Elution was carried out with a two-step linear NaCl gradient (0.0-0.6 and 0.6-1.0 M) in the same buffer. MPG synthase activity was found in the fraction eluting between 0.25 and 0.5 M NaCl. Active fractions were pooled, concentrated, and equilibrated to 20 mM Tris-HCl. The sample was applied to a 6-ml Resource Q column and eluted with a linear NaCl gradient (0-1 M). Fractions with activity eluted between 0.2 and 0.6 M. Purity of the samples was determined by SDS-PAGE. Three different pools with different degrees of purity resulted from this purification step.

Gel Filtration Chromatography (Superose 12)-- The purest pool was concentrated by ultrafiltration (10-kDa cutoff). Fractions were applied to a gel Superose 12 column equilibrated with 0.35 M NaCl in 50 mM Tris-HCl (pH 7.6) and eluted with the same buffer. Active fractions were subjected to dialysis against 20 mM Tris-HCl (pH 8.0).

Anion-exchange Chromatography (Mono Q)-- The sample was loaded onto a Mono Q column that was eluted by a linear gradient of NaCl (0-1 M). The fraction with MPG synthase eluted between 0.55 and 0.6 M NaCl.

Purification of Recombinant MPG Phosphatase

Extracts for the purification of MPG phosphatase, as well as the assay for enzyme activity were as described above.

Ion-exchange Chromatography-- The MPG phosphatase-containing supernatant was applied to a DEAE-Sepharose fast flow column as described for the recombinant MPG synthase. Elution was carried out with a two-step linear NaCl gradient (0.0-0.5 and 0.5-1.0 M). MPG phosphatase eluted between 0.2 and 0.35 M NaCl. Active fractions were concentrated, dialyzed against 20 mM Tris-HCl (pH 7.9), and loaded onto a Mono Q column that was eluted by a linear gradient of NaCl (0.0-1.0 M). The fraction with MPG phosphatase eluted between 0.25 and 0.3 M NaCl.

Characterization of Recombinant MPG Synthase and MPG Phosphatase

All biochemical and kinetic parameters for these enzymes were determined using the assay conditions described above. The temperature profiles for activity of MPG synthase and MPG phosphatase were determined between 30 and 108 °C. The effect of pH on MPG synthase activity was determined at 98 °C in 50 mM BisTris/propane buffer (pH 6.5-9.5) and 50 mM CAPSO (pH 7.0-10.0). The effect of pH on MPG phosphatase activity was determined at 98 °C in 50 mM acetate buffer (pH 3.4-5.4), 50 mM BisTris/propane buffer (pH 6.5-9.5), and CAPSO (pH 7.0-10.0). All pH values were measured at room temperature (25 °C); pH values at 98 °C were calculated using the conversion factor pK_a/T °C = 0.015 for BisTris/propane and 0.018 for CAPSO. Enzyme thermostabilities were determined at 98 °C by incubating enzyme solutions (0.5 mg/ml) in 20 mM Tris-HCl (pH 7.6). At appropriate times, samples were withdrawn and immediately examined for residual activities at 98 °C.

Kinetic parameters for MPG synthase were determined in reaction mixtures containing GDP-mannose (0.1-5.0 mM) plus D-3-phosphoglycerate (5 mM) or GDP-mannose (5 mM) plus D-3-phosphoglycerate (0.1 to 5.0 mM). Reaction mixtures for the determination of the kinetic parameters of MPG phosphatase contained MPG (0.1-2.0 mM). Samples of MPG synthase and MPG phosphatase reactions were pre-heated for 3 and 2 min, respectively, and all reactions were initiated by the addition of the enzyme preparation. Kinetic parameters for all substrates were determined at 98 °C. All experiments were performed in duplicate. Values for V_max and K_m were determined from Hanes plots.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Effect of NaCl Concentration of the Medium on Growth and Solute Accumulation by P. horikoshii-- This organism had a behavior illustrative of slightly halophilic organisms, requiring 1.5-5.0% NaCl in the culture medium for growth, with an optimum for growth of about 2.5% NaCl (Fig. 1). Cells grown in medium with 1.5% NaCl were enlarged, compared with cells grown at higher NaCl concentrations, and the cell yield was significantly lower than after growth under the other conditions. At the lowest salinity examined for compatible solute accumulation (2.5% NaCl), the total pool of solutes was low. At this salinity MG was the major compatible solute (0.11 µmol/mg protein) compared with trace levels of DIP (0.04 µmol/mg protein). An increase of the salt concentration of the medium to 3.5% NaCl caused an increase in MG concentration to 0.24 µmol/mg protein and of DIP to 0.21 µmol/mg protein, whereas trehalose remained vestigial. At the highest salinity examined (4.5% NaCl), there was a large increase in MG levels to 0.84 µmol/mg protein, without a concomitant alteration in the levels of DIP. An increase in the trehalose concentration was also observed in this medium.

View larger version (24K): [in this window] [in a new window]   Fig. 1.   Effect of the NaCl concentration of medium on the growth rate () and on the accumulation of compatible solutes (, mannosylglycerate; , di-myo-inositol phosphate; , trehalose) during growth of P. horikoshii at the optimum growth temperature, 98 °C.

Synthesis of MG in Cell Extracts-- From an array of experiments using GDP-mannose, UDP-mannose, and ADP-mannose as possible sugar donors and D-3-phosphoglycerate and D-glycerate as the sugar acceptors, we only detected the formation of MG in cell extracts from GDP-mannose and D-3-phosphoglycerate. The specific activity for MG production in P. horikoshii cell extracts was 6.1 nmol/min·mg protein. MPG synthase/MPG phosphatase activities could not be measured independently in cell extracts nor was a phosphorylated intermediate detected by TLC, but the hypothesis of a two step pathway was confirmed during the purification of MPG synthase, because the phosphatase activity was separated after the Resource Q step. The analysis of the reaction mixture by TLC after this step showed a compound that did not co-migrate with standard MG. Incubation of this reaction mixture with alkaline phosphatase led to the formation of a compound that co-migrated with MG.

Purification of Native MPG synthase and Identification of mpgs and mpgp Genes-- MPG synthase was purified in five chromatographic steps, the final preparation containing three bands on SDS-PAGE (results not shown). The N-terminal sequence analyses of the three bands led to the identification of three separate ORFs in the P. horikoshii genome, one of which (PH0927) coded for a 45-kDa protein enclosing a conserved domain for family 2 glycosyltransferases in its sequence (NCBI, PSI-BLAST) (Fig. 2), the size of which was in agreement with one of the bands in the denaturing gel. Therefore, this protein was considered the most likely candidate for MPG synthase. Moreover, the analysis of the sequences surrounding ORF PH0927 revealed two ORFs (PH0925 and PH0923) putatively identified as mannose-1-phosphate-guanylyltransferase/phosphomannose isomerase (M1P-GT/PMI, EC 2.7.7.22/EC 5.3.1.8) and phosphomannomutase (EC 5.4.2.8), respectively. The presence of genes putatively related to the synthesis of mannose-containing compounds near PH0927 strengthened our hypothesis that this was the MPG synthase gene. Additionally, these findings also indicated PH0926 as the best candidate for a MPG phosphatase, which was further supported by the presence in its sequence of a conserved domain common to trehalose-6-phosphate phosphatases (Fig. 2).

View larger version (89K): [in this window] [in a new window]   Fig. 2.   Multiple sequence alignment of PH0927 (MPG synthase, MPGS) and PH0926 (MPG phosphatase, MPGP) homologues. The abbreviations are as follows: Pho, P. horikoshii; Pab, P. abyssi (NCBI accession numbers E75104 and F75104); Pfu, P. furiosus (Utah Genome Center PF610337 and PF609612) Ape, A. pernix (NCBI accession numbers H72683 and F72683). Conserved domains for glucosyltransferase 2 (MPGS) and for trehalose-P phosphatase (MPGP) are highlighted by boxes and consensus identical (black) or similar (gray) amino acids are shaded. Conserved DXD motif common to several glycosyltransferase families is indicated by arrows ().

Cloning, Functional Overexpression of mpgs and mpgp in E. coli, and Purification of Recombinant Enzymes-- PCR amplification of mpgs and mpgp from genomic DNA of P. horikoshii yielded products with the expected gene sizes. For overexpression in E. coli, the PCR-amplified mpgs and mpgp were separately cloned under the control of the strong inducible trc promoter (pTRC99A) and tac promoter (pKK223-3), respectively. The sequence of the insert for mpgp (PH0926) was identical to that of the native ORF. However, the cloning of mpgs (PH0927) required the introduction of a NcoI site in the coding region, resulting in a substitution of leucine at position 2 to valine in the recombinant gene product.

Activity assays carried out in E. coli cell extracts revealed MPG production by MPG synthase clones and MPG dephosphorylation by MPG phosphatase clones but not by the negative control E. coli XL1-Blue (pTRC99A or pKK223-3) cell extracts. SDS-PAGE analysis of cell extracts from E. coli XL1-Blue (containing pMPGS and pMPGP clones) grown with IPTG induction showed extra bands of 45 and 28 kDa, respectively, that were not observed in cell extracts from E. coli XL1-Blue with empty vectors. The specific activities of MPG synthase and of MPG phosphatase in crude extracts of E. coli XL1-Blue were 356 and 162 nmol/min·mg protein at 98 °C, respectively. Heat treatment of cell extracts at 80 °C for 20 min resulted in extensive purification of the 45 kDa (MPG synthase) and the 28 kDa (MPG phosphatase) proteins. The specific activity of MPG synthase in heat-treated cell extracts of E. coli XL1-Blue (pMPGS) was 10.1 µmol/min·mg protein at 98 °C. Specific activity of MPG phosphatase in heat-treated cell extracts of E. coli XL1-Blue (pMPGP) was 3.3 µmol/min·mg protein at 98 °C. The purity of recombinant MPG synthase and MPG phosphatase was judged by SDS-PAGE (Fig. 3).

View larger version (27K): [in this window] [in a new window]   Fig. 3.   SDS-PAGE of purified recombinant MPG synthase and MPG phosphatase. Lane 1, molecular mass markers; lane 2, recombinant MPG synthase; Lane 3, recombinant MPG phosphatase.

Catalytic Properties of MPG Synthase-- Nine sugar nucleotides, namely ADP-mannose, GDP-mannose, UDP-mannose, ADP-glucose, GDP-glucose, UDP-glucose, TDP-glucose, UDP-galactose, and ADP-ribose, were used as possible sugar donors, and six 3-carbon compounds (glycerol, D-3-phosphoglycerate, D-2-phosphoglycerate, L-glycerol-3-phosphate, 2,3-diphospho-D-glycerate, and phosphoenolpyruvate) were used as sugar acceptors. Of these, only GDP-mannose and D-3-phosphoglycerate formed MPG (results not shown). The unequivocal identification of the reaction product as -mannosyl-3-phosphoglycerate (MPG) was achieved by standard one- and two-dimensional NMR spectroscopy (Table I). In particular, the -configuration was established from the measurement of the coupling constant between the anomeric carbon and the directly bound proton (J = 171.8 Hz).

MPG synthase exhibited Michaelis-Menten kinetics and the K_m values for the substrates are shown in Table II. The activity of MPG synthase in the absence of Mg²⁺ was 46% that in the presence of this divalent cation, but NaCl and KCl, in the concentration range of 50 to 300 mM, inhibited enzyme activity (Table II). At 40 °C the activity of the enzyme was undetectable, and maximal activity of the enzyme was reached between 90 and 100 °C (Fig. 4). At 108 °C MPG synthase still had 25% of maximal activity. Within the pH range examined (5.4-9.0), the activity of the enzyme at 98 °C was maximal between pH 6.4 and pH 7.4 (Fig. 5). At 98 °C, the optimal temperature for growth of P. horikoshii, the half-life for MPG synthase activity was 16 min (Fig. 6).

View larger version (13K): [in this window] [in a new window]   Fig. 4.   Temperature dependence of the recombinant MPG synthase () and the recombinant MPG phosphatase () of P. horikoshii. The enzyme activities were determined between 30 and 108 °C under the conditions described under "Experimental Procedures."

View larger version (12K): [in this window] [in a new window]   Fig. 5.   pH dependence of the recombinant MPG synthase (solid symbols) and the recombinant MPG phosphatase (open symbols) of P. horikoshii. The enzyme activities were determined between pH 3.5 and 9.0 in acetate buffer (), BisTris/propane ( and ) and CAPSO ( and ).

View larger version (10K): [in this window] [in a new window]   Fig. 6.   Thermostability of recombinant MPG synthase () and MPG phosphatase (). Enzymes were incubated at 98 °C. Samples were withdrawn and examined for activity at the same temperature. The half-life for thermal inactivation at 98 °C of MPG synthase and MPG phosphatase was 16 and 15.6 min, respectively.

Catalytic Properties of MPG Phosphatase-- Several sugar phosphates, MPG, mannose 1-phosphate, mannose 6-phosphate, glucose 1-phosphate, glucose 1,6-bisphosphate, trehalose 6-phosphate, fructose 1-phosphate, fructose 6-phosphate, and ribose 5-phosphate, as well as GDP and GMP, were examined as possible substrates for MPG phosphatase. However, only MPG was dephosphorylated by the enzyme. MPG phosphatase exhibited Michaelis-Menten kinetics (Table II). NaCl and KCl between 50 and 300 mM had no effect on the enzyme activity, but Mg²⁺ was required for maximal activity. Below 40 °C the activity of the enzyme was undetectable, and maximal activity was reached between 95 and 100 °C (Fig. 4). Furthermore, 20% of maximal activity was retained at 108 °C. The optimum pH range for activity of the enzyme was 5.2-6.4, with 15% of the maximal activity observed even at pH 3.7 (Fig. 5). The enzyme activity had a half-life of 15.6 min at 98 °C (Fig. 6). The reaction catalyzed by MPG phosphatase led to the complete conversion of the substrate.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Mannosylglycerate serves as a compatible solute in several slightly halophilic or halotolerant bacteria and archaea that live at high temperatures, accumulating in direct response to an increase in the salinity of the medium (5, 6, 9-11). P. horikoshii, like the other slightly halophilic species of the order Thermococcales examined, also produces progressively higher amounts of MG in response to salt stress.

Here we propose that the pathway for the synthesis of MG in P. horikoshii proceeds via a two-step pathway where GDP-mannose and D-3-phosphoglycerate are converted to the phosphorylated intermediate, mannosyl-3-phosphoglycerate, which in turn is dephosphorylated to -mannosylglycerate (Fig. 7). Thus, the synthesis of MG can be traced from the key glycolytic metabolite glucose 6-phosphate as follows: glucose 6-phosphate is converted to fructose 6-phosphate by the action of a novel type of phosphoglucose isomerase (EC 5.3.1.9), recently identified in P. furiosus (27), which exhibits extensive sequence homology to P. horikoshii ORF PH1956 (83% sequence identity). Fructose 6-phosphate can then be isomerized to mannose 6-phosphate by phosphomannose isomerase (PH0925), putatively identified as a bifunctional PMI/M1P-GT (28, 29). Mannose 6-phosphate is converted to mannose 1-phosphate by phosphomannose mutase (PH0923). The putative bifunctional enzyme encoded by PH0925 could convert mannose 1-phosphate to GDP-mannose in the next step, using GDP as substrate. The assertion that PH0925 encodes bifunctional PMI/M1P-GT is strongly supported by the presence of two conserved domains that are commonly found in type II PMI enzymes that possess both catalytic activities (29). The subsequent step involves the formation of MPG from GDP-mannose and 3-phosphoglycerate by MPG synthase (PH0927). MPG is finally converted to MG by MPG phosphatase (PH0926) (Fig. 7).

View larger version (19K): [in this window] [in a new window]   Fig. 7.   Proposed pathway for the synthesis of mannosylglycerate in P. horikoshii. PGI, phosphoglucose isomerase (PH1956); PMI/M1P-GT (PH0925, NCBI accession number BAA30021); PMM, phosphomannose mutase (PH0923, NCBI accession number BAA30019); MPGS, manno- syl-3-phosphoglycerate synthase (PH0927, NCBI accession number BAA30023); MP GP,mannosyl-3-phosphoglycerate phosphatase (PH0926, NCBI accession number BAA30022).

The proposed pathway in P. horikoshii is also supported by the genetic organization of PH0927, PH0926, PH0925, and PH0923 (Fig. 8), because these four genes are organized in an operon-like structure. Moreover, the occurrence of a consensus archaeal AT-rich promoter sequence (TTTATATA) directly upstream of PH0927 indicates the formation of a polycystronic mRNA transcript (30, 31). It should be pointed out that other candidate mannose-1-phosphate guanylyltransferase (e.g. PH1697 and PH1022) and phosphomannose mutase (e.g. PH1210) genes are found in P. horikoshii that may be involved in the synthesis of mannose derivatives needed for other biosynthetic purposes, such as mannose-containing polysaccharides. Therefore, we hypothesize that genes PH0923 and PH0925 encoding phosphomannose mutase and bifunctional phosphomannose isomerase/mannose-1-phosphate guanylyltransferase, may be committed to MG synthesis, being expressed upon osmotic sensing.

View larger version (26K): [in this window] [in a new window]   Fig. 8.   Genomic organization and flanking regions of the MG operon in four archaeal species, P. horikoshii, P. abyssi, P. furiosus, and A. pernix. Arrows represent identified or putative genes (shaded) and unknown open reading frames (open) and their direction. The percentage of amino acid (aa) identity between homologous sequences is indicated inside the arrows. The enlargement shows the promoter region upstream mpgs gene with the consensus TATA box, possible ribosome-binding site, and ATG start codon in boxes.

Unlike R. marinus, where two pathways for the synthesis of MG have been identified (15), only the two-step pathway, proceeding through a phosphorylated intermediate, has been detected in the archaeon P. horikoshii. An enzyme activity for the direct conversion of GDP-mannose and D-glycerate to MG by cell extracts was not detected in this organism. A search of the P. horikoshii genome revealed an ORF (PH1879) that had 29% sequence similarity with the R. marinus MG synthase. However, the recombinant gene product had no measurable activity for the synthesis of MG.² The pathway for the synthesis of MG in P. horikoshii appears to be similar to those described for the synthesis of osmolytes like trehalose (32), glucosylglycerol (33), and galactosylglycerol (34), insofar as all proceed via two-step pathways involving a phosphorylated intermediate. The existence of a single pathway for the synthesis of MG in P. horikoshii instead of the branched pathway of R. marinus probably imposes a lower flexibility on the regulation of MG synthesis in response to osmotic and/or thermal stress. However, the significance of the presence of two pathways in R. marinus, as well as the accumulation of the compatible solute mannosylglyceramide, which is unique to this organism, remains elusive.

A search of data bases revealed that genes encoding MPG synthase (PH0927) and MPG phosphatase (PH0926) have homologues, with sequence identities of at least 87 and 69%, respectively, in Pyrococcus abyssi and P. furiosus genomes (National Centre for Sequencing, France (www.genoscope.cns.fr) and Utah Genome Center website (www.genome.utah.edu)). Moreover, the corresponding genes in these Pyrococcus spp. are also organized in operon-like structures with four genes that include putative phosphomannose mutase and bifunctional phosphomannose isomerase/mannose-1-phosphate guanylyltransferase. Lower sequence homology (37 and 27% identity) to the MPG synthase and MPG phosphatase genes of Pyrococcus spp. were also detected in the crenarchaeote A. pernix genome (37), but putative phosphomannose mutase and bifunctional phosphomannose isomerase/mannose-1-phosphate guanylyltransferase were not found immediately downstream from MPG synthase/MPG phosphatase genes (Fig. 8).

The MPG synthase sequence contains one highly conserved motif, DXD, found in several families of glycosyltransferases that use nucleoside diphosphate sugars as substrates (38). Glycosyltransferases are classified as "inverting" or "retaining" enzymes according to whether their catalytic mechanisms result in inversion or retention of the anomeric configuration of substrates (Carbohydrate-Active enzymes (afmb.cnrs-mrs.fr/~pedro/CAZY) and see Refs. 39, 41, and 42). A PSI-BLAST search with the MPG synthase sequence revealed a conserved domain specific of family 2 glycosyltransferases (Fig. 3), which are "inverting" enzymes. However, our data show that MPG synthase retains the -configuration of the substrate GDP-mannose and therefore cannot be classified as a family 2 glycosyltransferase. The same reasoning is applicable to the MG synthase characterized earlier in R. marinus (15). MPG synthase from P. horikoshii does not share significant amino acid sequence homology with other known sugar transferases nor with MG synthase from R. marinus (15). In view of the apparent peculiarity of the amino acid sequences of these two GDP-mannose-dependent enzymes, and the retaining character of their catalytic mechanism, the elucidation of their three-dimensional structures would be most important to reconcile these findings.

The sequence of P. horikoshii MPG phosphatase contained a conserved domain found in trehalose-6-phosphate phosphatases (Fig. 2); however, trehalose 6-phosphate was not a substrate for MPG phosphatase. Interestingly, MPG phosphatase has no significant sequence homology with other known phosphatases, and this divergence may explain the very high specificity of this enzyme for MPG. The glucosylglycerol-phosphate phosphatase from Synechocystis spp. is another example of a phosphatase that possesses a consensus motif common to acid phosphatases (43), very weak sequence similarity with other sugar phosphatases, and a very high specificity for its substrate (44).

MPG synthase and MPG phosphatase had maximum activity at very high temperatures but no detectable activity at low temperatures, as might be expected from results with other intracellular enzymes from hyperthermophilic organisms (35, 45). More interesting is the low thermostability of both enzymes at 98 °C, the optimum temperature for growth of the organism. The weak intrinsic thermostability of MPG synthase and MPG phosphatase, from an organism that can grow at temperatures close to the boiling point of water, implies the existence of extrinsic stabilizing factors and/or high turnover rates to ensure the functionality of these enzymes in vivo. It should be noted that other intracellular enzymes from hyperthermophiles also display various degrees of thermostability some of which are also surprisingly low (36).

In addition to mannosylglycerate P. horikoshii accumulated di-myo-inositol phosphate and small amounts of trehalose. It is interesting to note that genes for the synthesis of trehalose could not be identified in the genome of this organism, nor could we find the corresponding activities in cell extracts. Therefore, the accumulation of this disaccharide is exclusively due to uptake from the yeast extract in the growth medium, through a high affinity ABC maltose/trehalose transporter recently identified in Thermococcus litoralis (40).

The pathway for the synthesis of mannosylglycerate in the hyperthermophilic archaeon P. horikoshii proposed by us, as well as the characterization of the enzymes, genes, and operon-like structure, represents an essential step toward the elucidation of osmo/thermoregulation through the accumulation of compatible solutes in thermophilic and hyperthermophilic organisms.

	ACKNOWLEDGEMENT

We thank Prof. Pedro M. Coutinho, IST, Lisboa, for enlightening discussions on glycosyltransferase classification.

	FOOTNOTES

^* This work was supported in part by the European Commission 5th Framework Program, Project QLK3-CT-2000-00640, and FCT/FEDER Projects PRAXIS/P/BIO/12082/1998 and POCTI/35715/BIO/2000.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

^¶ Recipient of Ph.D. Grant BD/21665/99 from PRAXIS XXI.

Recipient of Ph.D. Grant BD/19868/99 from PRAXIS XXI.

^** To whom correspondence should be addressed: Instituto de Tecnologia Química e Biológica, Universidade Nova de Lisboa, Rua da Quinta Grande 6, Apartado 127, 2780-156 Oeiras, Portugal. Tel: 351-214469800; Fax: 351-214428766; E-mail: santos@itqb.unl.pt.

Published, JBC Papers in Press, September 18, 2001, DOI 10.1074/jbc.M108054200

² N. Empadinhas, J. D. Marugg, H. Santos, and M. S. da Costa, unpublished results.

	ABBREVIATIONS

The abbreviations used are: DIP, di-myo-inositol phosphate; MG, -mannosylglycerate; IPTG, isopropyl--D-thiogalactopyranoside; PAGE, polyacrylamide gel electrophoresis; PCR, polymerase chain reaction; BisTris, 2-[bis(hydroxyethyl)amino]-2-(hydroxymethyl)-propane-1,3-diol; MPG, -mannosyl-3-phosphoglycerate; MPG synthase, mannosyl-3-phosphoglycerate synthase; MPG phosphatase, mannosyl-3-phosphoglycerate phosphatase; ORF, open reading frame; CAPSO, 3-[cyclohexylamino]-2-hydroxy-1-propanesulfonic acid; M1P-GT/PMI, mannose-1-phosphate-guanylyltransferase/phosphomannose isomerase.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

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