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Biochem J. Apr 1, 2005; 387(Pt 1): 271–280.
Published online Mar 22, 2005. Prepublished online Oct 28, 2004. doi:  10.1042/BJ20041053
PMCID: PMC1134955

Identification and characterization of Sulfolobus solfataricus D-gluconate dehydratase: a key enzyme in the non-phosphorylated Entner–Doudoroff pathway

Seonghun Kim* and Sun Bok Lee*‡,1


The extremely thermoacidophilic archaeon Sulfolobus solfataricus utilizes D-glucose as a sole carbon and energy source through the non-phosphorylated Entner–Doudoroff pathway. It has been suggested that this micro-organism metabolizes D-gluconate, the oxidized form of D-glucose, to pyruvate and D-glyceraldehyde by using two unique enzymes, D-gluconate dehydratase and 2-keto-3-deoxy-D-gluconate aldolase. In the present study, we report the purification and characterization of D-gluconate dehydratase from S. solfataricus, which catalyses the conversion of D-gluconate into 2-keto-3-deoxy-D-gluconate. D-Gluconate dehydratase was purified 400-fold from extracts of S. solfataricus by ammonium sulphate fractionation and chromatography on DEAE-Sepharose, Q-Sepharose, phenyl-Sepharose and Mono Q. The native protein showed a molecular mass of 350 kDa by gel filtration, whereas SDS/PAGE analysis provided a molecular mass of 44 kDa, indicating that D-gluconate dehydratase is an octameric protein. The enzyme showed maximal activity at temperatures between 80 and 90 °C and pH values between 6.5 and 7.5, and a half-life of 40 min at 100 °C. Bivalent metal ions such as Co2+, Mg2+, Mn2+ and Ni2+ activated, whereas EDTA inhibited the enzyme. A metal analysis of the purified protein revealed the presence of one Co2+ ion per enzyme monomer. Of the 22 aldonic acids tested, only D-gluconate served as a substrate, with Km=0.45 mM and Vmax=0.15 unit/mg of enzyme. From N-terminal sequences of the purified enzyme, it was found that the gene product of SSO3198 in the S. solfataricus genome database corresponded to D-gluconate dehydratase (gnaD). We also found that the D-gluconate dehydratase of S. solfataricus is a phosphoprotein and that its catalytic activity is regulated by a phosphorylation–dephosphorylation mechanism. This is the first report on biochemical and genetic characterization of D-gluconate dehydratase involved in the non-phosphorylated Entner–Doudoroff pathway.

Keywords: gluconate dehydratase, metalloprotein, non-phosphorylated Entner–Doudoroff pathway, phosphoprotein, Sulfolobus solfataricus, thermoacidophilic archaeon
Abbreviations: BAP, bacterial alkaline phosphatase; ED, Entner–Doudoroff; nED, non-phosphorylated ED; pnED, partially nED; GalD, galactonate dehydratase; KDG, 2-keto-3-deoxy-D-gluconate; KDPG, 2-keto-3-deoxy-6-phospho-D-gluconate; ORF, open reading frame; PAP, potato acid phosphatase; TBA, thiobarbituric acid


Hyperthermophilic archaea, which grow optimally above 80 °C, are found in volcanically and geothermally heated hydrothermal environments, such as solfataric fields, hot springs and submarine hot vents [1]. In these natural biotopes, the Sulfolobales convert elemental sulphur into hydrogen sulphide using organic compounds or hydrogen as an electron donor [2]. Of the various members of the Sulfolobales, the most intensive physiological and genomic studies have been performed on Sulfolobus solfataricus [35]. S. solfataricus, which grows optimally at 80–85 °C and pH 2–4, can use glucose as a sole carbon and energy source [3]. Although S. solfataricus is a sulphur-oxidizing micro-organism, this archaeon can grow chemoheterotrophically to high cell densities using various kinds of carbohydrates [68].

The glucose metabolism of S. solfataricus was first analysed by De Rosa et al. [9] using [14C]glucose-label experiments. They found that S. solfataricus converts D-glucose into pyruvate through a modified ED (Entner–Doudoroff) pathway, which produces non-phosphorylated intermediates such as D-gluconate, KDG (2-keto-3-deoxy-D-gluconate) and D-glyceraldehyde. In the nED (non-phosphorylated ED) pathway in S. solfataricus, D-glucose is oxidized to D-gluconate by NAD(P)+-dependent D-glucose dehydrogenase, and dehydrated by D-gluconate dehydratase (EC to yield KDG, which is then cleaved by KDG aldolase (EC to pyruvate and D-glyceraldehyde. A modified ED pathway involving non-phosphorylated intermediates was also discovered in the thermoacidophilic archaeon Thermoplasma acidophilum [10]. In this micro-organism, D-glyceraldehyde is formed through a non-phosphorylated route and converted by D-glyceraldehyde dehydrogenase into D-glycerate, which is then phosphorylated to form 2-phosphoglycerate. This intermediate is then converted to generate one molecule of pyruvate by enolase and pyruvate kinase. The nED pathway has been found only in thermoacidophilic archaea such as S. solfataricus and T. acidophilum.

Another modified ED pathway that involves D-gluconate and KDG as intermediates is known for some micro-organisms. This metabolic pathway, which is named here as the pnED (partially nED) pathway, was first discovered by Szymona and Doudoroff [11] in the bacterium Rhodobacter sphaeroides, and was later found in other bacteria and halophilic archaea [12]. In the pnED pathway, D-glucose is converted into D-gluconate and KDG as in the nED pathway, but the KDG produced by D-gluconate dehydratase is then phosphorylated by KDG kinase to KDPG (2-keto-3-deoxy-6-phospho-D-gluconate). KDPG is then cleaved by KDPG aldolase to pyruvate and D-glyceraldehyde 3-phosphate, as in the unmodified ED pathway. This latter intermediate is oxidized to pyruvate by D-glyceraldehyde-3-phosphate dehydrogenase, D-phosphoglycerate mutase, enolase and pyruvate kinase.

D-Gluconate dehydratase is a key enzyme involved in the modified ED pathways (nED and pnED). These dehydratases have been purified and characterized only from bacteria (Achromobacter sp. and Clostridium pasteurianum) that metabolize D-gluconate through the pnED pathway [1316]. A comparison of the biochemical properties of two bacterial dehydratases shows that they are quite different despite performing the same catalytic reaction. Moreover, the biochemical properties and detailed mechanisms of the D-gluconate dehydratases involved in the nED pathway of thermoacidophilic archaea (S. solfataricus and T. acidophilum) are not known. Furthermore, no D-gluconate dehydratase has been sequenced to date. Accordingly, the gene encoding D-gluconate dehydratase has not yet been annotated in the S. solfataricus and T. acidophilum genome database [4,17].

In the present study, we described the detailed biochemical properties of D-gluconate dehydratase isolated from S. solfataricus. We determined the N-terminal sequences of the purified protein and identified the gene encoding this protein in the S. solfataricus P2 genome sequence.


Strains and culture conditions

S. solfataricus (DSM1617) was obtained from the Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (Braunschweig, Germany). The organism was cultivated in GYM medium [7], which contained (per litre): 3.0 g of glucose, 3.0 g of yeast extract, 1.3 g of (NH4)2SO4, 0.28 g of KH2PO4, 0.25 g of MgSO4·7H2O, 0.07 g of CaCl2·H2O, 1 ml of trace metal solution (20 mg of FeCl3·H2O, 4.5 mg of Na2B4O7·H2O, 1.8 mg of MnCl2·H2O, 0.05 mg of ZnSO4·H2O, 0.05 mg of CuCl2·H2O, 0.04 mg of VOSO4·H2O, 0.03 mg of Na2MoO4·H2O and 0.01 mg of CoSO4·H2O). The final pH was adjusted to pH 3.0 using 0.5 M H2SO4. Cultures were grown aerobically in a 2.5 litre fermenter (KLF 2000; Bioengineering AG, Wald, Switzerland) at 78 °C with stirring at 400 rev./min. Growth was monitored spectrophotometrically at 540 nm. D-Gluconate dehydratase induction experiments were performed in the culture medium mentioned above, omitting D-glucose and yeast extract, supplemented with each carbon source (0.2%), namely D-arabinose, D-xylose, D-fructose, D-galactose, D-glucose, D-mannose, D-fucose, D-lactose, D-maltose, D-gluconate and D-galactonate.

Purification and N-terminal sequencing of D-gluconate dehydratase

Cells of S. solfataricus (frozen wet cell weight 35 g) were harvested by centrifugation (5000 g, 30 min, 4 °C) and washed twice with 50 mM Tris/HCl (pH 7.2). Cell pellets were resuspended in 50 mM Tris/HCl (pH 7.2) and then disrupted by sonication. Crude extracts were heated at 90 °C for 20 min and heat-denatured proteins and cell debris were removed by centrifugation (50000 g, 1 h, 4 °C). Solid (NH4)2SO4 was then added to the supernatant up to 40% saturation to recover the fraction containing D-gluconate dehydratase activity. After centrifugation (50000 g, 1 h, 4 °C), the supernatant was dialysed in 50 mM Tris/HCl (pH 7.2). The dialysate was loaded on to a DEAE-Sepharose column (2.5 cm×16 cm) previously equilibrated with 50 mM Tris/HCl (pH 7.2) and eluted with three bed volumes of the buffer, followed by a linear gradient of 0.0–1.0 M NaCl. Fractions (5 ml each) were collected at a flow rate of 1 ml/min. Those containing D-gluconate dehydratase activity were pooled, concentrated using a Vivaspin™ concentrator membrane (Vivascience, Lincoln, U.K.) and loaded on to a phenyl-Sepharose column (1.0 cm×10 cm) equilibrated with 50 mM Tris/HCl buffer (pH 7.2) containing 1.0 M NaCl. After washing with the buffer, the enzyme was eluted using a decreasing salt gradient (1.0–0.0 M NaCl) at 0.5 ml/min. Active fractions were pooled, concentrated by ultrafiltration and loaded on to a Mono Q HR 5/5 column (0.5 cm×5 cm) equilibrated with 50 mM Tris/HCl (pH 7.2). The enzyme was then eluted using a linear gradient of 0.0–1.0 M NaCl. Active fractions, collected at a flow rate of 0.5 ml/min, were pooled, concentrated by ultrafiltration and desalted using a HiTrap™ desalting column (Amersham Biosciences, Uppsala, Sweden). For the N-terminal sequence analysis, purified D-gluconate dehydratase was transferred on to a PVDF membrane. Protein sequencing was performed by the Korea Basic Science Institute (Daejeon, South Korea). Protein concentrations were determined using the Bradford method. SDS/PAGE and Western blotting were performed using standard procedures [18].

Assay of D-gluconate dehydratase

D-Gluconate dehydratase activity was determined using semicarbazide- or TBA (thiobarbituric acid)-based methods [19,20]. The semicarbazide method was performed as follows [19]: the reaction mixture of total volume of 400 μl was incubated at 78 °C in 50 mM Tris/HCl buffer (pH 7.0), with 10 mM D-gluconate and enzyme solution. After 30 min, the enzyme reaction was stopped by adding 100 μl of 2.0 M HCl. To this solution, 300 μl of semicarbizide solution [1.0% (w/v) semicarbazide hydrochloride and 1.5% (w/v) sodium acetate dissolved in distilled water] was then added and the mixture was incubated at 30 °C for 15 min. Finally, the reaction mixture was diluted with 500 μl of distilled water and its absorbance A250 was measured (molar absorption coefficient ε=571 M−1·cm−1). This method could detect quantitatively almost all 2-keto-aldonic acids and showed linearity in an extended range of concentration against them. TBA assay was performed as follows [20]: the reaction mixture of 50 μl was oxidized using 125 μl of 25 mM periodic acid in 0.125 M H2SO4 at room temperature (20 °C) for 20 min. To terminate the oxidation, 250 μl of 2% (w/v) sodium arsenite dissolved in 0.5 M HCl was added. Finally, 1 ml of 0.3% TBA was added and the reaction mixture was heated at 100 °C for 10 min. The red chromophore produced was monitored at 549 nm after adding an equal volume of DMSO (molar absorption coefficient ε=6.78×104 M−1·cm−1). This assay method is very sensitive for detecting and measuring sugar acids ranging from C-4 to C-7 containing a 2-keto-3-deoxy group. D-Gluconate dehydratase activity (1 unit) was defined as the amount of enzyme producing 1 μmol of KDG per min from D-gluconate under standard assay conditions (78 °C and pH 7.0). All enzyme activities were determined in triplicate.

Substrate specificities and kinetic parameters

The substrate specificity of D-gluconate dehydratase for sugar acids was determined using a semicarbazide method by determining the levels of 2-keto-3-deoxy analogues produced from aldonic acids. The sugar acids used in this study were as follows: D-gluconate, D-galactonate, D-galactoheptonate, D,L-arabonate, D-glucuronate, D,L-gulonate, D,L-tartarate, D-glucarate, D,L-isovalerate, L-threonate, D-ribonate, D-galactarate, D-xylonate, D-galacturonate, D-glucitol, D-mannonate and D,L-glycerate. The kinetic parameters of D-gluconate dehydratase were determined using D-gluconate (0.1–40 mM) as substrate. All experiments were performed in triplicate, and the apparent Vmax and Km values (means±S.E.M.) were calculated by fitting the initial-rate data to the Michaelis–Menten equation with a non-linear regression analysis program (Sigma Plot, version 7.0).

Effect of bivalent ions and thiol reagents

The effects of bivalent ions (Co2+, Ni2+, Mg2+, Mn2+, Cd2+, Ba2+, Fe2+, Ca2+, Zn2+, Cu2+ and Hg2+) and EDTA on D-gluconate dehydratase were determined by measuring its enzymic activity after preincubation with 50 mM Tris/HCl (pH 7.2) at 30 °C for 1 h in the presence of bivalent ions. The effects of thiol compounds (glutathione, dithiothreitol and 2-mercaptoethanol) and thiol-modification reagents (iodoacetamide, p-chloromercuribenzoic acid and p-hydroxymecuribenzoic acid) on enzyme activity were determined by the TBA method using the same procedure with thiol reagents instead of bivalent ions.

Metal content measurements

To determine whether metals are associated with the purified protein, the contents of Co2+, Ni2+, Mg2+, Mn2+, Fe2+, Ca2+ and Zn2+ in the purified enzyme preparations were measured after dialysing the samples in metal-free buffers by using an inductively coupled plasma MS with atomic emission detection (Thermo Jarrell-Ash IRIS-AP, Franklin, MA, U.S.A.).

Optimum temperature, thermostability and optimum pH

The biochemical and kinetic parameters of the enzyme were determined using the above-mentioned substrate specificity assay. The temperature profile of the enzyme activity was determined between 40 and 100 °C. Enzyme thermostability was determined at 80, 90 and 100 °C by incubating the enzyme at 50 μg/ml in 50 mM Tris/HCl (pH 7.2). At appropriate times, samples were immediately cooled on ice, and the residual enzyme activity was determined by the TBA method. The effect of pH on D-gluconate dehydratase activity was determined at 78 °C in 50 mM citric acid/NaOH buffer (pH 2.7–6.0), 50 mM Tris/HCl buffer (pH 6.0–8.6) or 50 mM glycine/NaOH buffer (pH 8.5–10.5) by using the TBA method.

Molecular mass determination

Purified D-gluconate dehydratase (100 μg) was chromatographed at a flow rate of 0.5 ml/min on a Sephacryl S-200 column (1.0 cm×89 cm) using a gel-filtration calibration kit (Amersham Biosciences). The buffer used for column equilibrium and elution was 50 mM Tris/HCl (pH 7.2) containing 150 mM NaCl. The molecular-mass standards used were thyroglobulin (669 kDa), ferritin (440 kDa), aldolase (158 kDa), BSA (67 kDa) and ovalbumin (43 kDa). The A280 of the eluate was monitored and D-gluconate dehydratase activity was measured by the TBA method described above. The molecular mass of native D-gluconate dehydratase was calculated by interpolation on a plot of the log of molecular mass against the Kav values. [Kav is a function of the elution volume of a molecule and defined as (VeVo)/(VtVo), where Ve is the elution volume, Vo the void volume and Vt the total column volume.]

Identification of phosphorylated D-gluconate dehydratase

For radioactive labelling of S. solfataricus proteins, cells were cultivated in GM medium [8] containing 1 mCi (37 MBq) of [32P]Pi at 78 °C for 4 days, and then harvested and frozen. Crude extract was prepared from the labelled cells and then loaded on to a phenyl-Sepharose column (1.0 cm×10 cm) equilibrated with 50 mM Tris/HCl buffer (pH 8.0) containing 1 M ammonium sulphate. After washing the column with the same buffer, proteins were eluted with a linear gradient of 50 mM Tris/HCl (pH 8.0). Fractions containing D-gluconate dehydratase activity were collected and then dialysed in 50 mM Tris/HCl (pH 8.0). The dialysate was then loaded on to a Q-Sepharose column (1.0 cm×20 cm) equilibrated with 50 mM Tris/HCl (pH 8.0) containing 1 M NaCl. The fractions containing enzyme activity were collected and then concentrated by ultrafiltration. To identify the D-gluconate dehydratase phosphorylation, the partially purified 32P-labelled proteins were dephosphorylated with either PAP (potato acid phosphatase) or BAP (bacterial alkaline phosphatase). Dephosphorylation by PAP was performed at 37 °C for 3 h in reaction mixtures containing 20 mM Mes/NaOH (pH 6.5) and 100 mM NaCl. Dephosphorylation by BAP was performed similarly, except that the reaction mixtures contained 50 mM Tris/HCl (pH 8.5), 1 mM MgCl2 and 0.1 mM ZnCl2 instead of Mes/NaOH and NaCl. The protein samples treated with phosphatase were loaded on to SDS/12% polyacrylamide gel and the gel was then run and stained with Coomassie Brilliant Blue. After the stained gel had dried, autoradiography was performed by exposing the dried gel to an X-ray film overnight.


Effect of carbon sources on D-gluconate dehydratase activity

Although S. solfataricus is lithoautotrophic, this micro-organism grows well under chemoheterotrophic conditions using a sugar, such as glucose, as a sole carbon and energy source [3]. Since exogenous carbon compounds generally regulate specific metabolic enzymes in vivo, we examined D-gluconate dehydratase activity after cultivating S. solfataricus in a minimal medium supplemented with various carbon sources. D-Gluconate dehydratase activity was determined in cells harvested at late stationary phase. Enzyme activities in crude extracts were determined by using the TBA method. The specific activities of D-gluconate dehydratase were found to be carbon-source-dependent (Table 1). However, D-gluconate did not induce the production of D-gluconate dehydratase. When D-gluconate was used as a sole carbon source, D-gluconate dehydratase activities were similar to those observed for D-arabinose or D-glucose. For D-lactose, enzyme activities were found to be lower than those observed for the other carbon sources. On the other hand, when D-xylose or D-galactonate was used as a sole carbon source, no appreciable cell growth was observed up to 10 days of the cultivation.

Table 1
The effect of sole carbon sources on the production of D-gluconate dehydratase in S. solfataricus

Purification of D-gluconate dehydratase

The procedures used to purify D-gluconate dehydratase are summarized in Table 2. The heat treatment and 40% ammonium sulphate fractionation yielded a 2-fold purification and a 50% recovery of enzyme activity. After pretreatment, D-gluconate dehydratase was purified by 4-step column chromatography using DEAE-Sepharose, Q-Sepharose, phenyl-Sepharose and Mono Q columns to give a 400-fold purification and a yield of 10.9%.

Table 2
Purification of D-gluconate dehydratase

The molecular mass of the native enzyme was estimated to be 350 kDa using a calibrated Sephacryl S-200 column, and the molecular mass of the denatured enzyme by SDS/PAGE was approx. 44 kDa (results not shown). These results indicate that the S. solfataricus D-gluconate dehydratase in its native conformation is a homo-octamer.

Catalytic properties of D-gluconate dehydratase

A total of 22 aldonic acids were examined as possible substrates for D-gluconate dehydratase (Table 3). To determine the substrate specificities of D-gluconate dehydratase, 10 mM aldonic acid was incubated with 40 μg of the purified protein/ml. When product formation was measured by the semicarbazide method, a 100% D-gluconate conversion was observed after 30 min incubation under the standard reaction conditions (78 °C and pH 7.0). The results obtained for the various aldonic acids are summarized in Table 3. The catalytic activity of D-gluconate dehydratase was found to be specific for D-gluconate. Although D-galactonate and D-galactoheptonate were also substrates, their activities were <3% of the activity of D-gluconate. On the other hand, <1% activity was detected for the following substrates: D-glucuronate, L-gulonate, D-tartarate, D-glucarate, D,L-isovalerate, L-threonate, D-ribonate, L-tartarate, D-gulonate and D-galactarate; and no catalytic activity was observed for L-arabonate, D-xylonate, D-glacturonate, D-glucitol, D-mannonate and D,L-glycerate (Table 3). Therefore it appears that the enzyme has a preference for D-gluconate among the aldonic acid substrates tested in the present study.

Table 3
Substrate specificity of D-gluconate dehydratase

The dependence of enzyme reaction rate on D-gluconate concentration followed Michaelis–Menten kinetics. The Km and Vmax values obtained when D-gluconate was used as a substrate were 0.45±0.03 mM and 0.15±0.01 unit/mg. Turnover number (kcat) was calculated to be 7.6×102 min−1, which yielded a kcat/Km value of 1.7×103 mM−1·min−1.

The activity of D-gluconate dehydratase in the presence of bivalent cations such as Co2+, Ni2+ or Mg2+ increased (Figure 1). However, Zn2+, Cu2+, Hg2+ and EDTA decreased enzyme activity. In the case of Co2+, Ni2+ or Mg2+, the enzyme activities did not vary significantly over the concentration range 0.01–10 mM. To determine whether metals are associated with D-gluconate dehydratase, the purified enzyme was subjected to metal analysis using inductively coupled plasma MS. Measurements of metal ion content in the purified enzyme revealed an average content of 1.0±0.2 cobalt atom per enzyme monomer (means±S.E.M.). The contents of Ni2+, Mg2+, Mn2+, Fe2+, Ca2+ and Zn2+ were found to be below the detection limits.

Figure 1
Effect of bivalent cations and EDTA on the activity of D-gluconate dehydratase

Table 4 shows the effect of thiol compounds on the activity of D-gluconate dehydratase. Unlike the D-gluconate dehydratase purified from C. pasteurianum [15], these compounds inhibited the enzyme activity on D-gluconate. In the presence of 1 mM glutathione, 2-dithiothreitol and 2-mercaptoethanol, D-gluconate dehydratase activity decreased to 59, 76 and 83% respectively. On the other hand, thiol modification reagents such as iodoacetamide or p-chloromercuribenzoic acid had no detectable inhibitory effect on D-gluconate dehydratase (Table 4), which differs from the D-gluconate dehydratase purified from Achromobacter sp. [13].

Table 4
Effect of thiol reagents on D-gluconate dehydratase activity

Purified enzyme displayed optimal activity between 80 and 90 °C (Figure 2A), and its activity was almost undetectable below 60 °C. The thermostability of purified D-gluconate dehydratase was measured at 80, 90 and 100 °C (Figure 2B). At 80 °C, the optimal temperature for S. solfataricus P2 growth, D-gluconate dehydratase was stable for over 2 h, but at 90 °C its activity was decreased to <50% in 2 h and its half-life diminished to 120 min. At 100 °C, the enzyme had a half-life of less than 40 min. With respect to pH optimization over the range pH 2.7–10.5, purified enzyme showed optimal activity between pH 6.5 and 7.5 (Figure 3). The pKa1 (app) and pKa2 (app) values were found to be 6.3 and 7.3 respectively. The relative activities of the enzyme were slightly different, depending on the buffers used in the experiment.

Figure 2
Effect of temperature on the activity (A) and thermostability (B) of D-gluconate dehydratase from S. solfataricus
Figure 3
Effect of pH on the activity of D-gluconate dehydratase from S. solfataricus

Identification of the gene encoding D-gluconate dehydratase

To determine the N-terminal amino acid sequence of D-gluconate dehydratase, the purified enzyme was subjected to SDS/PAGE, blotted on to a PVDF membrane and excised. The N-terminal sequence of D-gluconate dehydratase purified from S. solfataricus was determined to be MRIREIEPIV. This amino acid sequence of D-gluconate dehydratase was in exact agreement with SSO3198; annotated as a muconate cycloisomerase in the S. solfataricus P2 genome database (GenBank® accession no. NC_002754). The molecular mass of the purified D-gluconate dehydratase in denaturing gel was 44 kDa, which corresponded to the molecular mass deduced from the nucleotide sequence (44.729 kDa). Consequently, we confirmed that the protein purified from S. solfataricus is D-gluconate dehydratase with an ORF (open reading frame) corresponding to the SSO3198, which we named gnaD, encoding gluconate dehydratase.

Identification of phosphorylated D-gluconate dehydratase

An analysis of the amino acid sequence of D-gluconate dehydratase indicated the presence of several putative phosphorylation sites on serine, threonine and tyrosine residues. The numbers of potential phosphorylation sites obtained from NetPhos 2.0 [21] were six, four and five for serine, threonine and tyrosine residues respectively. To ascertain whether D-gluconate dehydratase is a phosphoprotein, it was purified from a S. solfataricus culture containing 32P-labelled Pi. The purified active enzyme was incubated in reaction mixtures containing either PAP or BAP, to dephosphorylate the proteins. D-Gluconate dehydratase activities were then assessed using the TBA method. Interestingly, D-gluconate dehydratase activity was lost after incubation with phosphatase, whereas no such decrease in activity was observed when the enzymes were incubated in the same reaction mixtures without phosphatase (Figure 4). This abrogation of enzyme activity occurred only in the presence of PAP or BAP. To confirm that D-gluconate dehydratase was dephosphorylated by phosphatase, reaction mixtures were analysed electrophoretically and autoradiographically. Samples not treated with phosphatase showed radioactive signals at 44 kDa, whereas samples treated with either PAP or BAP displayed no or a negligible signal (see Figure 4). These results strongly indicate that the D-gluconate dehydratase of S. solfataricus is a phosphoprotein. Moreover, the loss of activity by dephosphorylation indicates that the phosphoamino acid residues are located in the catalytic domain of the enzyme and that phosphorylation and dephosphorylation control the activity of D-gluconate dehydratase.

Figure 4
Phosphorylation of S. solfataricus D-gluconate dehydratase in 32P-labelled cells


In the present study, we investigated the D-gluconate dehydratase in the archaeon S. solfataricus and identified the gene (SSO3198) encoding this enzyme in the S. solfataricus P2 genomic database. D-Gluconate dehydratase (EC activities have been reported in three domains, archaea (Sulfolobus sp., Thermoplasma sp. and Halobacterium sp.), bacteria (Achromobacter sp. and Clostridium sp.) and eukarya (Aspergillus sp.) [916,22]. Although their activities have been known for several decades, the nucleotide and amino acid sequences of D-gluconate dehydratase have not been reported, which is why the gene encoding D-gluconate dehydratase could not be discovered in genomic data published to date.

For example, the gene corresponding to D-gluconate dehydratase cannot be found in the genome sequences of S. solfataricus P2 [4] and T. acidophilum [17], although other genes encoding enzymes of the glycolysis pathway are annotated in the genomic database: D-glucose dehydrogenase (SSO3204, SSO3042, SSO3003 and Ta0897), KDG aldolase (SSO3197 and Ta0619), D-glycerate kinase (SSO0666 and Ta0453), enolase (SSO0913 and Ta0882), pyruvate kinase (SSO0981 and Ta0896) and D-glyceraldehyde-3-phosphate dehydrogenase (SSO3194 and Ta0809). Recently, Verhees et al. [23] have predicted a potential gluconate dehydratase in the genome of S. solfataricus.

Previous reports have shown that D-gluconate dehydratases from Alcaligenes (Achromobacter) and Clostridia are induced when D-gluconate is used as a carbon source [13,24]. This implies that bacterial D-gluconate dehydratases are inducible enzymes required for the metabolism of D-gluconate [12]. However, for S. solfataricus the activities of D-gluconate dehydratase were almost independent of the carbon sources (except D-lactose, see below). When one among D-arabinose, D-galactose, D-glucose and D-gluconate was added to the culture medium as a sole carbon source, D-gluconate dehydratase activities were relatively unchanged, indicating that S. solfataricus D-gluconate dehydratase might be a constitutive enzyme (see Table 1). This seems to be reasonable because the D-gluconate dehydratases of thermoacidophilic archaea (S. solfataricus and T. acidophilum) involve the main glycolysis pathway. On the other hand, bacterial D-gluconate dehydratases are required only in the presence of D-gluconate.

When D-lactose was used as a carbon source, the enzyme activity was considerably lower than that for other carbon sources (Table 1). The reproducibility of the result was confirmed by repeating the same experiment four times. In every case, D-gluconate dehydratase activity in cells grown on D-lactose was always less than one half of that in cells grown on D-glucose or D-gluconate. This indicates that the enzyme activities of S. solfataricus D-gluconate dehydratase are somehow influenced by the carbon sources with unknown mechanisms.

Using the N-terminal sequence of the D-gluconate dehydratase purified from S. solfataricus, we identified the gene (SSO3198; gnaD) in the S. solfaricus P2 genome database. In addition, based on the amino acid sequence of S. solfataricus D-gluconate dehydratase, PSI-BLAST searches were performed against the NCBI database. This search showed that the amino acid sequence of gnaD is highly homologous with proteins in other thermoacidiphilic archaea. Calculated sequence identities were 79 and 61% for the two dgoA proteins of Sulfolobus tokodaii (dgoA1, GenBank® accession no. BAB67476; dgoA2, GenBank® accession no. BAB67683), 62% for the muconate cycloisomerase of S. solfataricus (MuC, GenBank® accession no. AAK42783), 42% for the hypothetical proteins of Ferroplasma acidarmanus (hypoP1, GenBank® accession no. ZP_00001372), 42% for the GalD (galactonate dehydratase) of T. acidophilum (GenBank® accession no. CAC11889) and 41% for GalD1 (GenBank® accession no. BAB59311) of Thermoplasma volcanium (Figure 5). It has been shown that Sulfolobus and Thermoplasma cells convert D-gluconate into KDG [9,10]. However, it has not yet been determined whether there is an enzyme which catalyses the same reaction in Ferroplasma. The homologous protein in Ferroplasma (hypoP1) may have the activities of D-gluconate dehydratase and a similar role in the carbohydrate metabolism. It seems probable that the functions of enzymes in the nED pathway of thermoacidophilic archaea might be similar to each other [25].

Figure 5
Amino acid sequence alignment of the S. solfataricus D-gluconate dehydratase orthologues

The PSI-BLAST searches revealed that GnaD protein contains enolase-like domains at both the N- and C-termini. We found that the secondary structures of the N- and C-terminal regions of GnaD are predicted to have a α/β barrel domain typical of those found in the enolase superfamily (results not shown). Sequence alignments also showed conserved residues commonly found in the active sites of enolase superfamily enzymes, such as mandelate racemase, muconate lactonizing enzyme and GalD. In particular, enolase superfamily proteins have typical conserved metal-ion binding sites inside a barrel [2628]. On the basis of sequence alignments and structure superpositions of enolase superfamily proteins, it seems probable that the marked residues in Figure 5 (Lys-207, Lys-229 and Glu-264) are putative metal-binding sites and are responsible for co-ordinating with the bivalent metal ions required for D-gluconate dehydratase activity. The putative amino acid sequences of the gnaD gene in S. solfataricus contain one highly conserved motif, KXK (Lys-159 and Lys-161), which appears to be diagnostic for the presence of functional groups in the enolase superfamily. These properties are in line with the biochemical characteristics of S. solfataricus D-gluconate dehydratase. De Rosa et al. [9] also observed that in vivo D-gluconate dehydratase activity increased in the presence of Mg2+. The KXK sequence motif is involved in the abstraction of the α-proton from carboxylic acid substrates to initiate dehydration from substrate such as D-gluconate at the active site [27,28]. As is shown in Figure 1, Co2+, Ni2+ or Mg2+ bivalent metal ions activate D-gluconate dehydratase, whereas Zn2+, Cu2+, Hg2+ and EDTA decreased enzyme activity.

SSO3198 is located upstream of SSO3197, which encodes KDG aldolase, in the S. solfataricus P2 genome. Previously, no recognizable promoter sequence could be found upstream of the start codon of KDG aldolase [29], and no suitable transcriptional stop signal was found downstream of the KDG aldolase gene. It is noteworthy that the upstream region of SSO3198 encoding D-gluconate dehydratase contains a putative consensus archaeal AT-rich promoter sequence. The consensus promoter sequences TTTATA (box A) and TTGC (box B) [30,31], which are typical of promoter regions in most archaeal genes, appropriately spaced by 20 nt, are located at −69 to −64 and −43 to −40 ahead of the ORF respectively (Figure 6). The BRE (transcription factor B recognition element), which matches with a typical Crenarchaeal consensus sequence (A/G)N(A/T)AA(A/T) [32], is located immediately upstream of the TATA box. The 3′-downstream region of SSO3194, after the translational termination site, also contains the putative transcriptional termination signal [33], TTTTTTAATT. These indicate that D-gluconate dehydratase (SSO3198), KDG aldolase (SSO3197) and other downstream enzymes (SSO3195 and SSO3194) are organized in an operon-like structure. However, we could not identify the ribosome-binding site (consensus sequence of Crenarchaea: GGTGA [30]) in the upstream of gnaD operon. A comparison of the genome sequences of S. solfataricus and S. tokodaii revealed that putative ORFs involved in the glycolysis pathway are well ordered. In S. tokodaii, the putative KDG aldolase (ST2479) and other downstream enzymes (ST2478 and ST2477) are located in operon-type gene order as in S. solfataricus, except for the putative D-gluconate dehydratase gene (ST2366). It remains to be seen whether the transcription of genes in the nED-pathway operon is regulated by a single promoter to produce a polycistronic mRNA transcript.

Figure 6
Genomic organization and flanking regions of the putative glycolysis gene operon in thermoacidophilic archaea, S. solfataricus and S. tokodaii

The properties of D-gluconate dehydratase isolated from S. solfataricus (SSO_GnaD) differ from those of Alcaligenes faecalis and Clostridiun pasteurianum despite their similar reaction mechanisms (Table 5): the maximum activity of SSO_GnaD occurs between 80 and 90 °C, and is undetectable below 60 °C, and the optimum pH of SSO_GnaD is lower than those of C. pasteurianum and A. faecalis. Moreover, the Km value of SSO_GnaD for D-gluconate is lower than that of two eubacteria, and the substrate specificity of SSO_GnaD is relatively narrow compared with those of C. pasteurianum and A. faecalis. The effect of thiol reagents on the enzyme activity of S. solfataricus D-gluconate dehydratase is also different from those of A. faecalis and C. pasteurianum (see Table 4). The enzyme from A. faecalis is deactivated by p-chloromercuribenzoic acid, which specifically modifies cysteine residues of proteins. However, the enzyme from S. solfataricus is not affected by p-chloromercuribenzoic acid. In addition, the activity of S. solfataricus enzyme is decreased by the presence of glutathione, dithiothreitol and mercaptoethanol, whereas thiol compounds cause an increase (C. pasteurianum) or no appreciable change (A. faecalis) in the activities of bacterial gluconate dehydratases [1315].

Table 5
Biochemical properties of D-gluconate dehydratases

The common properties of the D-gluconate dehydratases of archaea and bacteria are their requirement for bivalent cations. Co2+ was most effective for activity of D-gluconate dehydratases from S. solfataricus, whereas Mg2+ or Mn2+ had the same effect on D-gluconate dehydratases obtained from S. solfataricus and A. faecalis. On the other hand, the D-gluconate dehydratase from C. pasteurianum, which is strongly inhibited by oxygen, was maximally activated by ferrous ions. All D-gluconate dehydratase activities were inhibited by the chelating agent EDTA. The relation between D-gluconate dehydratases and metal ions reveal that bivalent metal ions are involved in D-gluconate dehydration. From metal analysis of the purified protein, we have found that the D-gluconate dehydratase from S. solfataricus contains one Co2+ ion per enzyme monomer.

In the present study, we also found that the catalytic activity of S. solfataricus D-gluconate dehydratase is regulated by a phosphorylation–dephosphorylation mechanism. The D-gluconate dehydratase purified from the 32P-labelled cell lysate showed radioactive signals on film. Also, in vitro treatment of the enzyme with phosphatase decreased the activity, which implies that the phosphorylation of D-gluconate dehydratase is essential for catalytic activity. Protein phosphorylation in archaea was first reported from the extreme halophilic archaeon Halobacterium halobium [34]. Since then, several phosphoproteins whose activity or function is regulated by phosphorylation–dephosphorylation have been identified. These include protein kinases from several archaea [35], methyltransferase activation protein from Methanosarcina barkeri [36], Cdc6 protein from Methanobacterium thermoautotrophicum [37] and initiation factor 2α (aIF2α) from Pyrococcus horikoshii [38]. In S. solfataricus, phosphoglycerate mutase, another enzyme involved in the glycolysis pathway, was recently identified as a phosphoprotein [39]. Interestingly, this protein has the ability to autophosphorylate its serine catalytic site by using ATP as a phosphate donor.

Recently, eight protein kinases were identified by homology searching in the S. solfataricus P2 genome, and four of these were experimentally confirmed as protein serine/threonine kinases [35,40,41]. These protein kinases may be involved in the phosphorylation of D-gluconate dehydratase in S. solfataricus. A computer analysis of the amino acid sequence of D-gluconate dehydratase revealed that it contains 15 putative sites that can be phosphorylated by protein kinases. However, it remains to be seen which protein kinases are involved in the phosphorylation of D-gluconate dehydratase and which amino acid residues are phosphorylated.

Although the regulation of the protein function by phosphorylation is well established in bacteria and eukarya [35], the corresponding situation in archaea is largely unknown. It has been shown that several proteins associated with the glycolysis pathway in eukarya are phosphoproteins. These include glucose-6-phosphate dehydrogenase [42], phosphoglycerate mutase [43], phosphofructokinase [43], enolase [44] and lactate dehydrogenase [44]. Since the proteins associated with the glycolysis pathways in eukarya, bacteria and archaea share the module regulated by protein phosphorylation, the archaea might require a fine regulation system to control enzyme activity and metabolic flux. Thus they may have developed regulatory devices for protein phosphorylation as a strategy for optimizing their metabolic system. Considering that glycolysis is a core pathway, phosphorylation-modulated glycolysis pathway regulation might be useful as an evolutionary tracer.

In summary, the D-gluconate dehydratase from S. solfataricus was found to show biochemical properties distinct from those of bacteria. Unlike bacterial D-gluconate dehydratases, the D-gluconate dehydratase of S. solfataricus is an octameric phosphoprotein, which appears to be synthesized constitutively. From the N-terminal sequence of D-gluconate dehydratase, it has been found that the gene corresponding to D-gluconate dehydratase (gnaD) is SSO3198 in the S. solfataricus P2 genome database. This represents a first step towards understanding the D-gluconate dehydratases involved in the glycolysis of extreme thermoacidophilic archaea through non-phosphorylated intermediates. Further studies on the structure and the regulation of D-gluconate dehydratase are needed for a complete understanding of the mechanistics and the in vivo role of this enzyme in thermoacidophilic archaea.


This work was supported by a grant from POSCO through the POSTECH Biotechnology Center and by a grant from the 21C Frontier Microbial Genomics and Applications Center Program.


1. Stetter K. O. Extremophiles and their adaptation to hot environments. FEBS Lett. 1999;452:22–25. [PubMed]
2. Segerer A. H., Stetter K. O. The order Sulfolobales. In: Balows A., Trüper H. G., Dvorkin M., Harder W., Schleifer K.-H., editors. The Prokaryotes. 2nd edn. New York: Springer; 1992. pp. 684–701.
3. Grogan D. W. Phenotypic characterization of the archaebacterial genus Sulfolobus: comparison of five wild-type strains. J. Bacteriol. 1989;171:6710–6719. [PMC free article] [PubMed]
4. She Q., Singh R. K., Confalonieri F., Zivanovic Y., Allard G., Awayez M. J., Chan-Weiher C. C., Clausen I. G., Curtis B. A., De Moors A., et al. The complete genome of the crenarchaeon Sulfolobus solfataricus P2. Proc. Natl. Acad. Sci. U.S.A. 2001;98:7835–7840. [PMC free article] [PubMed]
5. Ciaramella M., Pisani F. M., Rossi M. Molecular biology of extremophiles: recent progress on the hyperthermophilic archaeon Sulfolobus. Antonie Van Leeuwenhoek. 2002;81:85–97. [PubMed]
6. Park C. B., Lee S. B. Constant-volume fed-batch operation for high density cultivation of hyperthermophilic aerobes. Biotechnol. Tech. 1997;11:277–281.
7. Park C. B., Lee S. B. Inhibitory effect of mineral ion accumulation on high density growth of the hyperthermophilic archaeon Sulfolobus solfataricus. J. Biosci. Bioeng. 1999;87:315–319. [PubMed]
8. Park C. B., Lee S. B., Ryu D. D. Y. L-pyroglutamate spontaneously formed from L-glutamate inhibits growth of the hyperthermophilic archaeon Sulfolobus solfataricus. Appl. Environ. Microbiol. 2001;67:3650–3654. [PMC free article] [PubMed]
9. De Rosa M., Gambacorta A., Nicolaus B., Giardina P., Poerio E., Buonocore V. Glucose metabolism in the extreme thermoacidophilic archaebacterium Sulfolobus solfataricus. Biochem. J. 1984;224:407–414. [PMC free article] [PubMed]
10. Budgen N., Danson M. J. Metabolism of glucose via a modified Entner-Doudoroff pathway in the thermoacidophilic archaebacterium Thermoplasma acidophilum. FEBS Lett. 1986;196:207–210.
11. Szymona M., Doudoroff M. Carbohydrate metabolism in Rhodopseudomonas sphreoides. J. Gen. Microbiol. 1958;22:167–183. [PubMed]
12. Conway T. The Entner-Doudoroff pathway: history, physiology and molecular biology. FEMS Microbiol. Rev. 1992;103:1–27. [PubMed]
13. Kersters K., Khan-Matsubara J., Nelen L., De Ley J. Purification and properties of D-gluconate dehydratase from Achromobacter. Antonie Van Leeuwenhoek. 1971;37:233–246. [PubMed]
14. Kersters K., De Ley J. D-gluconate dehydratase from Alcaligenes. Methods Enzymol. 1975;42:301–304. [PubMed]
15. Bender R., Gottschalk G. Purification and properties of D-gluconate dehydratase from Clostridium pasteurianum. Eur. J. Biochem. 1973;40:309–321. [PubMed]
16. Gottschalk G., Bender R. D-gluconate dehydratase from Clostridium pasteurianum. Methods Enzymol. 1982;90:283–287. [PubMed]
17. Ruepp A., Graml W., Santos-Martinez M. L., Koretke K. K., Volker C., Mewes H. W., Frishman D., Stocker S., Lupas A. N., Baumeister W. The genome sequence of the thermoacidophilic scavenger Thermoplasma acidophilum. Nature (London) 2000;407:508–513. [PubMed]
18. Sambrook J., Russell D. W. 3rd edn. Plainview, NY: Cold Spring Harbor Laboratory Press; 2001. Molecular Cloning: A Laboratory Manual.
19. MacGee J., Doudoroff M. A new phosphorylated intermediate in glucose oxidation. J. Biol. Chem. 1954;210:617–626. [PubMed]
20. Skoza L., Mohos S. Stable thiobarbituric acid chromophore with dimethyl sulphoxide. Application to sialic acid assay in analytical de-O-acetylation. Biochem. J. 1976;159:457–462. [PMC free article] [PubMed]
21. Blom N., Gammeltoft S., Brunak S. Sequence and structure-based prediction of eukaryotic protein phosphorylation sites. J. Mol. Biol. 1999;294:1351–1362. [PubMed]
22. Elzainy T. A., Hassan M. M., Allam A. M. New pathway for nonphosphorylated degradation of gluconate by Aspergillus niger. J. Bacteriol. 1973;114:457–459. [PMC free article] [PubMed]
23. Verhees C. H., Kengen S. W., Tuininga J. E., Schut G. J., Adams M. W., De Vos W. M., Van Der Oost J. The unique features of glycolytic pathways in Archaea. Biochem. J. 2003;375:231–246. [PMC free article] [PubMed]
24. Bender R., Andreesen J. R., Gottschalk G. 2-Keto-3-deoxygluconate, an intermediate in the fermentation of gluconate by clostridia. J. Bacteriol. 1973;107:570–573. [PMC free article] [PubMed]
25. Danson M. J. Central metabolism of the archaea. In: Kates M., Kushner D. J., Matheson A. T., editors. The Biochemistry of Archaea (archaebacteria) Amsterdam: Elsevier; 1993. pp. 1–21.
26. Babbitt P. C., Gerlt J. A. New functions from old scaffolds: how nature reengineers enzymes for new functions. Adv. Protein Chem. 2001;55:1–28. [PubMed]
27. Babbitt P. C., Mrachko G. T., Hansson M. S., Huisman G. W., Kolter R., Ringe D., Petsko G. A., Kenyon G. L., Gerlt J. A. A functionally diverse enzyme superfamily that abstracts the α protons of carboxylic acids. Science. 1995;267:1159–1161. [PubMed]
28. Babbitt P. C., Hasson M. S., Wedekind J. E., Palmer D. R. J., Barrett W. C., Reed G. H., Rayment I., Ringe D., Kenyon G. L., Gerlt J. A. The enolase superfamily: a general strategy for enzyme-catalyzed abstraction of the alpha-protons of carboxylic acids. Biochemistry. 1996;35:16489–16501. [PubMed]
29. Buchanan C. L., Connaris H., Danson M. J., Reeve C. D., Hough D. W. An extremely thermostable aldolase from Sulfolobus solfataricus with specificity for non-phosphorylated substrates. Biochem. J. 1999;343:563–570. [PMC free article] [PubMed]
30. Tolstrup N., Sensen C. W., Garrett R. A., Clausen I. G. Two different and highly organized mechanisms of translation initiation in the archaeon Sulfolobus solfataricus. Extremophiles. 2000;4:175–179. [PubMed]
31. Reiter W.-D., Hüdepohl U., Zillig W. Mutational analysis of an archaebacterial promoter: essential role of a TATA box for transcription efficiency and start-site selection in vitro. Proc. Natl. Acad. Sci. U.S.A. 1990;87:9509–9513. [PMC free article] [PubMed]
32. Bell S. D., Jackson S. P. Mechanism and regulation of transcription in archaea. Curr. Opin. Microbiol. 2001;4:208–213. [PubMed]
33. Reiter W.-D., Palm P., Zillig W. Transcription termination in the archaebacterium Sulfolobus: signal structures and linkage to transcription initiation. Nucleic Acids Res. 1988;16:2445–2459. [PMC free article] [PubMed]
34. Spudich J. L., Stoeckenius W. Light-regulated retinal-dependent reversible phosphorylation of Halobacterium proteins. J. Biol. Chem. 1980;255:5501–5503. [PubMed]
35. Kennelly P. J. Archaeal protein kinases and protein phosphatases: insights from genomics and biochemistry. Biochem. J. 2003;370:373–389. [PMC free article] [PubMed]
36. Daas P. J., Wassenaar R. W., Willemsen P., Theunissen R. J., Keltjens J. T., van der Drift C., Vogels G. D. Purification and properties of an enzyme involved in the ATP-dependent activation of the methanol: 2-mercaptoethanesulfonic acid methyl-transferase reaction in Methanosarcina barkeri. J. Biol. Chem. 1996;271:22339–22345. [PubMed]
37. Grabowski B., Kelman Z. Autophosphorylation of archaeal Cdc6 homologues is regulated by DNA. J. Bacteriol. 2001;183:5459–5464. [PMC free article] [PubMed]
38. Tahara M., Ohsawa A., Saito S., Kimura M. In vitro phosphorylation of initiation factor 2 alpha (aIF2 alpha) from hyperthermophilic archaeon Pyrococcus horikoshii OT3. J. Biochem. (Tokyo) 2004;135:479–485. [PubMed]
39. Potters M. B., Solow B. T., Bischoff K. M., Graham D. E., Lower B. H., Helm R., Kennelly P. J. Phosphoprotein with phosphoglycerate mutase activity from the archaeon Sulfolobus solfataricus. J. Bacteriol. 2003;185:2112–2121. [PMC free article] [PubMed]
40. Lower B. H., Kennelly P. J. Open reading frame sso2387 from the archaeon Sulfolobus solfataricus encodes a polypeptide with protein-serine kinase activity. J. Bacteriol. 2003;185:3436–3445. [PMC free article] [PubMed]
41. Lower B. H., Potters M. B., Kennelly P. J. A phosphoprotein from the archaeon Sulfolobus solfataricus with protein-serine/threonine kinase activity. J. Bacteriol. 2004;186:463–472. [PMC free article] [PubMed]
42. Napier M. A., Lipari M. T., Courter R. G., Cheng C. H. Epidermal growth factor receptor tyrosine kinase phosphorylation of glucose-6-phosphate dehydrogenase in vitro. Arch. Biochem. Biophys. 1987;259:296–304. [PubMed]
43. Sale E. M., White M. F., Kahn C. R. Phosphorylation of glycolytic and gluconeogenic enzymes by the insulin receptor kinase. J. Cell. Biochem. 1987;33:15–26. [PubMed]
44. Cooper J. A., Esch F. S., Taylor S. S., Hunter T. Phosphorylation sites in enolase and lactate dehydrogenase utilized by tyrosine protein kinases in vivo and in vitro. J. Biol. Chem. 1984;259:7835–7841. [PubMed]

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