The First Crystal Structure of Archaeal Aldolase

UNIQUE TETRAMERIC STRUCTURE of 2-DEOXY-D-RIBOSE-5-PHOSPHATE ALDOLASE FROM THE HYPERTHERMOPHILIC ARCHAEA Aeropyrum pernix*

Haruhiko SakurabaDagger §, Hideaki Tsuge§||, Ikuko ShimoyaDagger , Ryushi KawakamiDagger , Shuichiro GodaDagger , Yutaka Kawarabayasi**, Nobuhiko Katunuma, Hideo AgoDagger Dagger , Masashi MiyanoDagger Dagger , and Toshihisa OhshimaDagger §§

From the Dagger  Department of Biological Science and Technology, Faculty of Engineering, University of Tokushima, Tokushima 770-8506, Japan, the  Institute for Health Sciences, Tokushima Bunri University, Yamashiro-cho, Tokushima 770-8514, Japan, the ** National Institute of Advanced Industrial Science and Technology, Tsukuba, Ibaraki 305-8566, Japan, and the Dagger Dagger  Structural Biophysics Laboratory, RIKEN Harima Institute, Hyogo 679-5148, Japan

Received for publication, December 6, 2002, and in revised form, January 14, 2003

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES

A gene encoding a 2-deoxy-D-ribose-5-phosphate aldolase (DERA) homolog was identified in the hyperthermophilic Archaea Aeropyrum pernix. The gene was overexpressed in Escherichia coli, and the produced enzyme was purified and characterized. The enzyme is an extremely thermostable DERA; its activity was not lost after incubation at 100 °C for 10 min. The enzyme has a molecular mass of ~93 kDa and consists of four subunits with an identical molecular mass of 24 kDa. This is the first report of the presence of tetrameric DERA. The three-dimensional structure of the enzyme was determined by x-ray analysis. The subunit folds into an alpha /beta -barrel. The asymmetric unit consists of two homologous subunits, and a crystallographic 2-fold axis generates the functional tetramer. The main chain coordinate of the monomer of the A. pernix enzyme is quite similar to that of the E. coli enzyme. There was no significant difference in hydrophobic interactions and the number of ion pairs between the monomeric structures of the two enzymes. However, a significant difference in the quaternary structure was observed. The area of the subunit-subunit interface in the dimer of the A. pernix enzyme is much larger compared with the E. coli enzyme. In addition, the A. pernix enzyme is 10 amino acids longer than the E. coli enzyme in the N-terminal region and has an additional N-terminal helix. The N-terminal helix produces a unique dimer-dimer interface. This promotes the formation of a functional tetramer of the A. pernix enzyme and strengthens the hydrophobic intersubunit interactions. These structural features are considered to be responsible for the extremely high stability of the A. pernix enzyme. This is the first description of the structure of hyperthermophilic DERA and of aldolase from the Archaea domain.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES

Aldolases catalyze carbon-carbon bond formation and cleavage and are attractive as synthetic catalysts due to their ability to produce stereospecific carbohydrates (1). They are divided into two major classes based on the mode of stabilization of reaction intermediates. Class I and II aldolases employ a Schiff base mechanism (2, 3) and a divalent metal ion for intermediate stabilization (4), respectively. 2-Deoxy-D-ribose-5-phosphate aldolase (DERA1; EC 4.1.2.4) belongs to the class I aldolases and catalyzes a reversible aldol reaction between acetaldehyde and D-glyceraldehyde 3-phosphate to generate 2-deoxy-D-ribose 5-phosphate. DERA is unique in catalyzing the aldol reaction between two aldehydes as both the aldol donor and acceptor components. Its broad substrate specificity is an attractive characteristic for the production of a variety of stereospecific materials (5). The enzyme has high potential utility as an environmentally benign alternative to chiral transition metal catalysis of the asymmetric aldol reaction (6). However, the practical application of DERA is still not successful because of its low stability.

Since DERA was first described by Racker (7), who reported the presence of DERA in mammalian tissue as well as in Escherichia coli and Corynebacterium diphtheriae, a number of the enzymes from eucaryotes and bacteria have been studied and characterized (8, 9). Despite its wide distribution, the physiological role of the enzyme still remains obscure: nothing is known to date about its role in eucaryotes, whereas in bacterial cells, it has been proposed to function in the catabolism of deoxyribonucleosides (10, 11). In Bacillus cereus, the enzyme has been reported to play a key role in the utilization of the pentose moiety of exogenous nucleosides (12, 13). In Salmonella typhimurium and E. coli, the gene encoding DERA belongs to the deo regulon that contains four genes encoding enzymes involved in nucleoside catabolism (11, 14). The presence of DERA has not been described so far in Archaea, the third domain of life.

Recently, much attention has been paid to the isolation and characterization of enzymes from hyperthermophilic Archaea because the organisms have high potentiality as a new source of much more stable enzymes than the counterparts of mesophiles. A gene (open reading frame identification number APE2437) encoding a DERA homolog has been identified in the aerobic hyperthermophilic Archaea A. pernix via genome sequencing (15). In preliminary studies, we performed the cloning and expression of APE2437 in E. coli. However, no functional products could be obtained. We purified the native enzyme from A. pernix; analyzed the N-terminal amino acid sequence; and identified that TTG, which is present at 126 bp upstream from the 5' terminus of the predicted open reading frame, is the proper initial codon. In this study, we succeeded in the expression of the gene and purification and characterization of A. pernix DERA. In addition, we determined the crystal structure of the enzyme at 2.0-Å resolution and revealed that it has a unique tetrameric structure that might contribute to the high stability. We present here the first report of the characterization and crystal structure of hyperthermophilic DERA. These are essential steps in the effort to comprehend its function and stabilizing mechanisms and also to achieve the practical application of DERA.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES

Chemicals-- The pET-11a vector was obtained from Novagen (Madison, WI). The E. coli strain BL21-CodonPlusTM-RIL was purchased from Stratagene (La Jolla, CA). 2-Deoxy-D-ribose 5-phosphate, triose-phosphate isomerase, and glycerol-3-phosphate dehydrogenase were obtained from Sigma (Osaka, Japan). Restriction enzymes were purchased from New England Biolabs Inc. (Beverly, MA). KOD DNA polymerase was obtained from Toyobo (Osaka). All other chemicals were reagent-grade.

Organism and Growth Conditions-- The hyperthermophilic Archaea A. pernix K1 (JCM 9820) was obtained from the Japanese Collection of Microorganisms (Wako, Saitama, Japan). The microorganism was cultured in the modified medium of Sako et al. (16), consisting of natural seawater containing 5 g/liter trypticase peptone, 3 g/liter yeast extract, and 0.76 g/liter Na2S2O3 (pH 7.0 adjusted with 0.5 N NaOH). Cells were grown by shaking (100 rpm) in an air bath rotary shaker at 90 °C in 700 ml of medium in a 2-liter flask. After 24 h of cultivation, sterilized water (~100 ml) was replenished to avoid evaporation of the medium, and cultivation was continued for an additional 18 h. The cells were collected by centrifugation (7000 × g, 15 min) and washed twice with 3% NaCl solution. The washed cells were suspended in 10 mM Tris-HCl buffer (pH 8.0).

Purification of DERA from A. pernix K1-- Cells (~30 g (wet weight) from a 7-liter culture) were disrupted by ultrasonication. Cell debris was removed by centrifugation (15,000 × g, 20 min, 4 °C), and nucleic acids were removed by the addition of streptomycin sulfate to a final concentration of 1%, followed by centrifugation (15,000 × g, 20 min, 4 °C). The resultant supernatant was used as the crude extract. DERA was partially purified from the crude extract. The entire operation was done at room temperature (~25 °C). The crude extract (120 ml) was placed on a DEAE-Toyopearl column (5 × 15 cm; TOSOH, Tokyo, Japan) equilibrated with 10 mM Tris-HCl buffer (pH 8.0). After the column was washed with the same buffer, the enzyme was eluted with a 1000-ml linear gradient of 0-0.5 M NaCl in the same buffer. The active fractions were pooled, and the enzyme was dialyzed against 10 mM Tris-HCl buffer (pH 8.0). Solid (NH4)2SO4 was added to the enzyme solution up to 20% saturation. The enzyme solution was loaded onto a butyl-Toyopearl column (4 × 10 cm; TOSOH) that was previously equilibrated with the buffer supplemented with 20% (NH4)2SO4. After the column was washed with the same buffer (~3 column bed volumes), the enzyme was eluted with a 1000-ml linear gradient of 20 to 0% (NH4)2SO4 in the same buffer. The active fractions were collected and dialyzed against 10 mM Tris-HCl buffer (pH 8.0).

PAGE-- SDS-PAGE (12% acrylamide slab gel, 1 mm thick) was performed following the procedure of Laemmli (17). The protein band was stained with Coomassie Brilliant Blue R-250.

N-terminal Amino Acid Sequencing-- Approximately 3 µg of protein was subjected to SDS-PAGE, followed by electroblotting onto a polyvinylidene difluoride membrane. The membrane was then stained with Ponceau S and destained. A protein band was excised and subjected to automated Edman degradation using a Shimadzu Model PPSQ-10 protein sequencer.

Overexpression and Purification of Recombinant Protein-- The following set of oligonucleotide primers was used to amplify the DERA gene fragment by PCR: 5'-TATATCATATGCCGTCGGCCAGGGATATAC-3', containing a unique NdeI restriction site overlapping the 5'-initiation codon (TTG has been changed to ATG), and 5'-TAATGGATCCTTAGACTAGGGATTTGAAGC-3', containing a unique BamHI restriction site proximal to the 3'-end of the termination codon. The chromosomal A. pernix DNA was isolated as described (18) and used as the template. The amplified 0.7-kb fragment was digested with NdeI and BamHI and ligated with the expression vector pET-11a linearized with NdeI and BamHI to generate pEDERA. E. coli strain BL21-CodonPlusTM-RIL was transformed with pEDERA. The transformants were cultivated at 37 °C in 200 ml of medium containing 2.4 g of Tryptone, 4.8 g of yeast extract, 1 ml of glycerol, 2.5 g of K2HPO4, 0.76 g of KH2PO4, and 10 mg of ampicillin until the absorbance at 600 nm reached 0.6. Induction was carried out by the addition of 0.1 mM isopropyl-beta -D-thiogalactopyranoside to the medium, and cultivation was continued for 12 h at 18 °C. Cells (~50 g (wet weight) from a 2-liter culture) were harvested by centrifugation, suspended in 10 mM Tris-HCl buffer (pH 8.0), and disrupted by ultrasonication. The crude extract was heated at 80 °C for 10 min, and the denatured protein was then removed by centrifugation (15,000 × g, 20 min). Solid (NH4)2SO4 was added to the enzyme solution up to 20% saturation. The enzyme solution was loaded onto a butyl-Toyopearl column and eluted using the same procedure as described above. The active fractions were collected and dialyzed against 10 mM Tris-HCl buffer (pH 8.0), loaded onto a DEAE-Toyopearl column, and eluted as described above. The active fractions were pooled, dialyzed against 10 mM Tris-HCl buffer (pH 8.0), and used as the purified enzyme preparation.

Determination of Enzyme Activity-- Assay for DERA activity was conducted at 50 °C essentially as described by Wong et al. (19). The activity was determined by measuring the oxidation of NADH in a coupled assay with triose-phosphate isomerase and glycerol-3-phosphate dehydrogenase. The assay mixture contained 100 mM imidazole HCl buffer (pH 6.5), 0.1 mM NADH, 0.2 mM 2-deoxy-D-ribose 5-phosphate, 11 units of triose-phosphate isomerase (rabbit muscle), 4 units of glycerol-3-phosphate dehydrogenase (rabbit muscle), and the enzyme preparation. The absorbance of NADH was followed at 340 nm (epsilon  = 6.22 mM-1 cm-1). One enzyme unit is defined as the amount catalyzing the cleavage of 1 µmol of 2-deoxy-D-ribose 5-phosphate/min. Protein was determined by the method of Bradford (20) using the standard assay kit from Bio-Rad with bovine serum albumin as the standard.

Molecular Mass Determination-- The molecular mass of the purified enzyme was determined by analytical gel filtration on a Superdex 200 column (2.6 × 62 cm; Amersham Biosciences) pre-equilibrated with 50 mM Tris-HCl buffer (pH 8.0) containing 0.2 M NaCl. Gel filtration calibration kits (Amersham Biosciences) were used for molecular mass standards. The subunit molecular mass of the purified enzyme was determined by SDS-PAGE using eight marker proteins (New England Biolabs Inc.).

Stability, pH Optima, and Kinetic Parameters-- To determine thermostability, the enzyme solutions (0.5 mg/ml) in 10 mM Tris-HCl buffer (pH 8.0) were incubated at different temperatures, and the residual activity was determined by the standard assay method. To determine pH stability, the enzyme (0.5 mg/ml) in buffers of various pH values was incubated at 50 °C for 60 min, and the remaining activities were then assayed. The buffers (500 mM) used were acetate buffer, imidazole HCl buffer, Tris-HCl buffer, glycine/NaOH buffer, and KCl/NaOH buffer at pH ranges of 3.5-6.0, 6.0-7.5, 7.5-8.5, 8.5-11.0, and 11.0-12.0, respectively. The effects of organic solvents on enzyme stability were examined by measuring the activity remaining after incubation with the reagents. An aliquot of the incubation mixture was withdrawn, and the remaining activity of the enzyme was assayed. The water-miscible organic solvents used were methanol, ethanol, Me2SO, and N,N-dimethylformamide. The optimal pH of the enzyme was determined by running the standard assay at 50 °C using citrate buffer (0.1 M), imidazole HCl buffer (0.1 M), Tris-HCl buffer (0.1 M), and glycine/NaOH buffer (0.1 M) at pH ranges of 5.0-6.5, 6.0-7.5, 7.5-8.5, and 8.5-10.0, respectively. The Michaelis constants were determined from Lineweaver-Burk plots (21) of data obtained from the initial rate of 2-deoxy-D-ribose 5-phosphate cleavage at 50 °C.

Crystallization and Data Collection-- Crystals were obtained using the hanging-drop vapor diffusion method, in which 2 µl of 20 mg/ml protein solution was mixed with an equal volume of mother liquor, consisting of 0.7-1.2 M NaH2PO4, 250 mM K2HPO4, and 100 mM Na2HPO4/citrate buffer (pH 4.2). Crystals were grown at 20 °C for 3 days. The crystal belongs to the orthorhombic space group P21212 with the following unit cell parameters: a = 75.3, b = 83.2, and c = 87.2 Å. Heavy atom derivatives were prepared by soaking the crystals in a reservoir solution containing 1 mM thimerosal (10 h) or 1 mM K2Pt(SCN)6 (3 h). Data were collected using an ADSC Quantum4R CCD detector system (Area Detector Systems) on the BL-6A beamline at the Photon Factory in Tsukuba, Japan (see Table I). All diffraction measurements were carried out on crystals cryoprotected with ethylene glycol and cooled to 100 K in a stream of nitrogen gas. The native and derivative data were processed and integrated by DPS/mosflm (22) and scaled by scala (23).

Phasing and Refinement-- Native, mercury, and platinum data sets (2.0-Å resolution) were used for phase calculation (Table I) by the MIRAS (multiple isomorphous replacement with an anomalous scattering) method using SOLVE (24). The MIRAS map at 2.0 Å was subjected to maximum-likelihood density modification, followed by autotracing using RESOLVE (25). An initial model was built using Xtalview (26). The model was adjusted in Xtalview using both |Fo- |Fc| and 2|Fo- |Fc| maps. Several cycles of rigid-body refinement, positional refinement, and simulated annealing were performed at 2.0-Å resolution with CNS (27). The final Rcryst and Rfree were calculated to be 22.4 and 26.3%, respectively (28). Fig. 1 shows the final 2|Fo- |Fc| map. Model geometry was analyzed with PROCHECK (29), and 94.0% of the non-glycine residues were in the most favorable region of the Ramachandran plot and 6.0% in the additionally allowed region. The final coordinates have been deposited in the Protein Data Bank (code 1N7K).


                              
View this table:
[in this window]
[in a new window]
 
Table I
Statistics on data collection, phase determination, and refinement
The crystal belongs to the space group is P21212, with a = 75.2, b = 84.2, c = 87.2 Å and alpha  = beta  = gamma  = 90 °. Data were collected at the Photon Factory on the BL-6A beamline using lambda  = 1.00 Å. Rsymn = Sigma hSigma i|Ih,i - < Ih> |/Sigma hSigma iIh,i Hg, 1 mM thimerosal (10 h), Pt, 1 mM K2Pt(SCN)6 (3 h); FOM, figure of merit; r.m.s.d., root mean square deviation.


View larger version (108K):
[in this window]
[in a new window]
 
Fig. 1.   The 2.0-Å resolution final 2|Fo- |Fc| map superimposed on the refined 2.0-Å resolution coordinates of A. pernix DERA. The map was contoured at 1sigma .


    RESULTS AND DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES

Identification of the Gene Encoding DERA-- The complete sequence of the genome of A. pernix has been reported by Kawarabayasi et al. (15). APE2437 (582 bp, positions 1,543,210-1,544,351 on the entire genome) has been annotated as the gene encoding a putative DERA (a protein of 193 amino acids with a molecular mass of 19,942 Da). The estimated molecular mass is obviously smaller than that of the E. coli DERA (28 kDa). We performed the cloning and expression of APE2437 in E. coli. However, no functional products could be obtained (data not shown). The activity of DERA in the cell extract of A. pernix was observed to be ~0.0026 units/mg. The enzyme was purified 46-fold, and the specific activity was 0.12 units/mg. The final preparation still contained several contaminating proteins on the basis of SDS-PAGE analysis. The N-terminal amino acid sequence of the protein that appeared as a major band was analyzed, and the sequence of the protein with a molecular mass of ~24 kDa was determined to be PSARDILQQG. On the basis of the sequence, the upstream region of the 5' terminus of APE2437 was analyzed. As a result, TTG, which was present at 126 bp upstream from the 5' terminus of the predicted open reading frame, was identified to be the proper initial codon for the DERA gene (Fig. 2). Kawarabayasi et al. (15) assigned GTG as the start codon of this gene because sense codons starting with ATG or GTG were used as the criteria for assignment of the potential coding region in the genomic sequence. However, the N-terminal analysis of the native enzyme shows that the sense codon of the enzyme gene starts with a minor TTG codon, which is sometimes used for the start codon in cyanobacteria (30). The newly identified gene (708 bp) was estimated to code a protein of 235 amino acids with a molecular mass of 24,529 Da. The predicted amino acid sequence showed 28% identity to that reported for E. coli DERA (Fig. 3) (9).


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 2.   Nucleotide sequence of the upstream region of the 5' terminus of APE2437 and the deduced amino acid sequence. The underlined sequence was determined from the N-terminal region of DERA purified from A. pernix. The arrow shows the predicted N-terminal amino acid sequence based on the genome analysis.


View larger version (44K):
[in this window]
[in a new window]
 
Fig. 3.   Structure-based amino acid sequence alignment of A. pernix and E. coli DERAs. Sequences were aligned using ClustalX (42). alpha -Helices (alpha 1-alpha 10; green) and beta -sheets (beta 1-beta 8; yellow) are shown. Asterisks represent conserved residues in the enzymes. The residues involved in 2-deoxy-D-ribose 5-phosphate binding in E. coli DERA are boxed.

Expression of the Gene and Purification of the Recombinant Enzyme-- E. coli strain BL21-CodonPlusTM-RIL transformed with the expression vector pEDERA exhibited high activity for DERA, which was not lost upon incubation at 100 °C for 10 min. The enzyme was purified to homogeneity by heat treatment and two column chromatographies from the extract of E. coli cells. An efficient purification of the enzyme was achieved; ~180 mg of the purified enzyme was obtained from 1 liter of E. coli culture. The specific activity of the purified enzyme was estimated to be 4.5 units/mg at 50 °C for the 2-deoxy-D-ribose 5-phosphate cleavage reaction.

Characteristics of A. pernix DERA-- The biochemical characteristics of the purified enzyme were determined. SDS-PAGE of the purified enzyme gave only one band; the subunit molecular mass was determined to be ~24 kDa and was consistent with the molecular mass calculated from the amino acid sequence (24,529 Da). The N-terminal amino acid sequence was determined to be PSARDILQQGLD, which is identical to that determined with the enzyme purified from A. pernix cells. This suggests that the first methionine is processed in E. coli cells as well as in A. pernix cells. The native molecular mass of the enzyme determined by gel filtration is ~93 kDa (Fig. 4). This indicates that the enzyme consists of four subunits with identical molecular mass. E. coli DERA has a dimer structure (Protein Data Bank code 1JCL) composed of two identical subunits, which is most common for DERA. The A. pernix enzyme is the first example of tetrameric DERA.


View larger version (11K):
[in this window]
[in a new window]
 
Fig. 4.   Molecular mass of A. pernix DERA. The molecular mass of the purified enzyme was determined by analytical gel filtration on Superdex 200 as described under "Experimental Procedures."

The optimal pH for the 2-deoxy-D-ribose 5-phosphate cleavage reaction was ~6.5. Typical Michaelis-Menten kinetics were observed for the reaction at 50 °C. The apparent Km value for 2-deoxy-D-ribose 5-phosphate was calculated to be 0.057 mM. The enzyme retained full activity upon heating at 100 °C for 10 min and at 80 °C for 60 min (Fig. 5, A and B). The thermostability of DERA has so far been reported only for the E. coli enzyme. The enzyme is rapidly inactivated above 70 °C (7). Thus, the A. pernix enzyme is probably the most thermostable DERA among the enzymes from other organisms described to date. The stability of the enzyme at various pH values is shown in Fig. 5C. The enzyme was extremely stable over a wide pH range; upon heating at 50 °C for 60 min, the enzyme did not lose activity at pH 4.5-11.0. The enzyme was also highly resistant to organic solvents such as ethanol, methanol, N,N-dimethylformamide, and Me2SO at 50 °C. Loss of activity was not observed in the presence of these reagents even at a concentration as high as 40% (Fig. 5D). These results suggest that A. pernix DERA might be preferred as a synthetic catalyst in practical application.


View larger version (29K):
[in this window]
[in a new window]
 
Fig. 5.   Effects of temperature, pH, and organic solvents on A. pernix DERA stability. A, after treatment at various temperatures for 10 min, the remaining activity was assayed at 50 °C. B, the enzyme was incubated at 80 °C, and the activity of the sample was assayed at 50 °C at appropriate intervals. C, the enzyme in buffer of various pH values was incubated at 50 °C for 60 min, and the remaining activity was then assayed at 50 °C. D, the enzyme was incubated with various concentrations of water-miscible organic solvents at 50 °C for 30 min. After incubation, the activity of the aliquot was assayed at 50 °C. The organic solvents used were ethanol (open circle ), methanol (triangle ), N,N-dimethylformamide (), and Me2SO ().

Architecture of the Subunit-- The structure of A. pernix DERA was determined by the MIRAS method and refined at 2.0-Å resolution to an R-factor (Rfree) of 22.4% (26.3%). The subunit comprises 234 residues. The asymmetric unit consists of two homologous subunits. The present model contains the complete ordered residues 2-235 of each subunit and 199 water molecules. The subunit folds into an (alpha /beta )8-barrel carrying two additional helical segments. The secondary structure was mapped onto the amino acid sequence (Fig. 3), and the three-dimensional arrangement is shown in Fig. 6. The elements of secondary structure that create the barrel are beta 1-alpha 3-beta 2-alpha 4-beta 3-alpha 5-beta 4-alpha 6-beta 5-alpha 7-beta 6-alpha 8-beta 7-alpha 9-beta 8-alpha 10. The alpha 2-helix caps the N-terminal section of the barrel and is closely associated with the alpha 10-helix, the C-terminal helix of the barrel. The alpha 1-helix forms an "arm" protruding away from the barrel (Fig. 6), which provides an important component for subunit-subunit interactions.


View larger version (56K):
[in this window]
[in a new window]
 
Fig. 6.   Crystal structure of A. pernix DERA. The rainbow drawing shows the N terminus in blue and the C terminus in red. The alpha -helices and beta -sheets are numbered from the N terminus. The figure was prepared using MOLSCRIPT (43) and Raster 3D (44).

Monomeric Structural Comparison of A. pernix and E. coli DERAs-- The structure of E. coli DERA was recently reported (33). The structure of A. pernix DERA was compared with that of E. coli DERA. The main chain coordinate of the monomer of the A. pernix enzyme is quite similar to that of the E. coli enzyme (root mean square deviation of 1.11 Å for the C-alpha atoms of 153 residues) (Fig. 7A). A noteworthy difference between the two enzyme monomers is that the alpha 1 arm is not present in the E. coli enzyme. The structure of E. coli DERA has been determined in complex with 2-deoxy-D-ribose 5-phosphate (33). We predicted the substrate-binding site by the superposition of A. pernix DERA and the E. coli enzyme (Fig. 7B). Most of the residues involved in substrate binding in E. coli DERA (Asp16, Thr18, Asp102, Lys137, Lys167, Thr170, Gly171, Lys201, Gly204, Gly205, Gly236, Ser238, and Ser239) are conserved in the A. pernix enzyme, except Cys47, Lys172, Val206, and Arg234 are replaced with Val, Val, Ile, and Ile, respectively (residue numbers of E. coli DERA are given). In A. pernix DERA, Lys167 appears to play a critical role in substrate binding because the same residue in the E. coli enzyme forms a Schiff base with the substrate (33).


View larger version (55K):
[in this window]
[in a new window]
 
Fig. 7.   A, stereographic drawing of the A. pernix DERA monomer structure (green) superimposed on the E. coli DERA monomer structure (yellow). The 2-deoxy-D-ribose 5-phosphate molecules are represented by a ball-and-stick model (red). The figure was prepared using MOLSCRIPT (43) and Raster 3D (44). B, stereographic close-up of the active site of A. pernix DERA. The 2-deoxy-D-ribose 5-phosphate of E. coli DERA was built in for a better understanding (red). Highly conserved residues in A. pernix and E. coli DERAs are shown in green and are labeled. The figure was prepared using Rasmol (32).

Oligomeric Structural Comparison of A. pernix and E. coli DERAs-- The quaternary structure differs between the tetrameric A. pernix and dimeric E. coli enzymes. The functional tetramer (subunits A, B, C, and D) of the A. pernix enzyme is a ringed doughnut-like shape with approximate dimensions of 82 × 70 × 50 Å (the dimensions of the hole are 37 × 22 Å) (Figs. 8A and 9A). The A-B and A-D association represents two distinct subunit-subunit interfaces. The two subunits of the A. pernix DERA asymmetric unit (A-B) create an arrangement totally distinct from that observed for the functional dimer of E. coli DERA (Fig. 8C). The A-B interface is formed by the antiparallel alignment of the alpha 5-helices together with interactions with residues in the N-terminal region of the alpha 4-helix (Fig. 8B). The A-D interface is formed by the antiparallel alignment of the alpha 1-helices together with interactions with the alpha 9- and alpha 10-helices (Fig. 8B). Neither the A-B nor A-D form is similar to the E. coli dimer. A. pernix DERA is 10 amino acids longer than the E. coli enzyme in the N-terminal region, which results in the presence of an additional N-terminal alpha 1-helix (Fig. 3). This N-terminal helix might be essential for the formation of the tetramer of A. pernix DERA.


View larger version (38K):
[in this window]
[in a new window]
 
Fig. 8.   A, the functional A. pernix DERA tetramer, with subunits A, B, C, and D shown in different colors. B, the A-B and A-D interfaces. The alpha 4- and alpha 5-helices in the A-B interface and the alpha 1-, alpha 9-, and alpha 10-helices in the A-D interface, which are important for oligomerization, are labeled. C, an arrangement of the two subunits of the A. pernix DERA asymmetric unit (A-B; green) and that observed for the functional dimer of E. coli DERA (yellow). The A subunit of A. pernix DERA is superimposed on the one subunit of the E. coli DERA dimer. The figure was prepared using MOLSCRIPT (43) and Raster 3D (44).

Structural Features for Hyperthermostability-- Recent structural studies on hyperthermophilic proteins reveal an increase in the number of ion pairs and the formation of ion pair networks compared with mesophilic counterparts (34-37). 168 and 177 ion pairs were identified in the intrasubunits of A. pernix and E. coli DERAs, respectively, using a cutoff distance between oppositely charged residues of 3.0 Å. This indicates that there is no significant difference in the number of total ion pairs and ion pair networks in the intrasubunits between the two enzymes. In general, the decrease in the solvent-accessible surface area and the increase in the fraction of buried hydrophobic atoms have been discussed as the stabilizing principles for thermostable protein (38). As shown in Table II, the A. pernix DERA monomer (10,726 Å2) showed similar total solvent-accessible surface area compared with the E. coli enzyme monomer (10,751 Å2). No significant difference in the number of hydrophobic residues (134 and 138 residues in A. pernix and E. coli DERAs, respectively) was observed between the two enzymes. On the other hand, the area (4735 Å2) of the interface between subunits A, B, C, and D is extremely larger than that (576.5 Å2) of the interface of the E. coli enzyme dimer (Table II). The surface of the inner section between A-D and B-C (A-B interface) and that between A-B and D-C (A-D interface) are shown in Fig. 9 (B and C), respectively. The green area (hydrophobic area) in the inner section is remarkable compared with that in the outer section (Fig. 9D). This means that these tetrameric interactions are mainly hydrophobic (green). The hydrophobic and ionic residues involved in the A-B and A-D interactions are listed in Table III. The hydrophobic interaction is especially striking on the alpha 1-, alpha 9-, and alpha 10-helices in the A-D interface, and these hydrophobic residues are Pro2 (alpha 1), Ile7 (alpha 1), Leu8 (alpha 1), Ile206 (alpha 9), Leu210 (alpha 9), Leu234 (alpha 10), and Val235 (alpha 10). These results suggest that the increase in intersubunit hydrophobic interactions as a result of the formation of a tetramer plays important roles in the extremely high stability of the A. pernix enzyme. However, further study is required to prove this hypothesis.


                              
View this table:
[in this window]
[in a new window]
 
Table II
Comparison of solvent-accessible surface areas


View larger version (48K):
[in this window]
[in a new window]
 
Fig. 9.   A, the solvent-accessible surface of the functional A. pernix DERA tetramer, with subunits A, B, C, and D labeled. The active sites are indicated by arrows. B, the surface of the inner section between A-D and B-C (A-B interface). C, the solvent-accessible surface of the inner section between A-B and D-C (A-D interface). D, the solvent-accessible surface of the outer section of the A-B dimer. In B-D, the hydrophobic, acidic, and basic areas are shown in green, red, and blue, respectively. The figure was prepared using GRASP (31).


                              
View this table:
[in this window]
[in a new window]
 
Table III
Hydrophobic and ionic residues involved in the interactions between the A and B subunits and between the A and D subunits

A few enzymes from hyperthermophiles have been reported to assume a higher oligomerization state than that of their mesophilic counterparts. These include Thermococcus kodakaraensis KOD1 ribulose-bisphosphate carboxylase/oxygenase (39) and Thermotoga maritima dihydrofolate reductase (40) and phosphoribosylanthranilate isomerase (41). The higher oligomerization states of these proteins have been suggested to contribute to extreme thermostability. However, the details of the relationship between the oligomeric structure of these enzymes and thermostability are not still clear. In this study, the structure of hyperthermophilic DERA was first determined. The structure of aldolase from hyperthermophiles has not been reported so far. This study suggests that the thermostability of the aldolase can be enhanced by the formation of a unique quaternary structure unlike the mesophilic counterpart. In addition, thermostable DERA is expected to have high potentiality as a catalyst for the synthesis of some 2-deoxy-D-ribose 5-phosphate derivatives, and the information on the three-dimensional structure in the active site may be useful for the development of this application.

    ACKNOWLEDGEMENTS

Data collection was performed at the Photon Factory. We thank Drs. M. Suzuki, N. Igarashi, and N. Sakabe (Photon Factory) for assistance with data collection.

    FOOTNOTES

* This work was supported in part by the "National Project on Protein Structural and Functional Analysis" promoted by the Ministry of Education, Science, Sports, Culture, and Technology of Japan. Data collection performed at the Photon Factory was supported by the Tsukuba Advanced Research Alliance.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.

The atomic coordinates and the structure factors (code 1N7K) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).

§ Both authors contributed equally to this work.

|| Guest researcher in the Tsukuba Advanced Research Alliance.

§§ To whom correspondence should be addressed. Tel.: 81-88-656-7518; Fax: 81-88-656-9071; E-mail: ohshima@bio.tokushima-u.ac.jp.

Published, JBC Papers in Press, January 15, 2003, DOI 10.1074/jbc.M212449200

    ABBREVIATIONS

The abbreviation used is: DERA, 2-deoxy-D-ribose-5- phosphate aldolase.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES

1. Wong, C. H., Halcomb, R. L., Ichikawa, Y., and Kajimoto, T. (1995) Angew. Chem. Int. Ed. Engl. 34, 412-432
2. Gefflaut, T., Blonski, C., Perie, J., and Willson, M. (1995) Prog. Biophys. Mol. Biol. 63, 301-340[CrossRef][Medline] [Order article via Infotrieve]
3. Morse, D. E., Tsolas, O., and Lai, C. Y. (1972) in The Enzymes (Boyer, P. D., ed), 3rd Ed., Vol. 7 , pp. 213-253, Academic Press, Inc., New York
4. Morse, D. E., and Horecker, B. L. (1968) Adv. Enzymol. 31, 125-181[Medline] [Order article via Infotrieve]
5. Barbas, C. F., III, Wang, Y. F., and Wong, C. H. (1990) J. Am. Chem. Soc. 112, 2013-2014
6. Machajewski, T. D., and Wong, C. H. (2000) Angew. Chem. Int. Ed. Engl. 39, 1352-1375[CrossRef][Medline] [Order article via Infotrieve]
7. Racker, E. (1951) J. Biol. Chem. 196, 347-365
8. Feingold, D. S., and Hoffee, P. A. (1972) in The Enzymes (Boyer, P. D., ed), 3rd Ed., Vol. 7 , pp. 330-321, Academic Press, Inc., New York
9. Valentin-Hansen, P., Boetius, F., Hammer-Jespersen, K., and Svendsen, I. (1982) Eur. J. Biochem. 125, 561-566[Abstract]
10. Munch-Petersen, A. (1970) Eur. J. Biochem. 15, 191-202[Medline] [Order article via Infotrieve]
11. Blank, J., and Hoffee, P. A. (1972) Mol. Gen. Genet. 116, 291-298[Medline] [Order article via Infotrieve]
12. Tozzi, M. G., Sgarrella, F., Barsacchi, D., and Ipata, P. L. (1984) Biochem. Int. 9, 319-325[Medline] [Order article via Infotrieve]
13. Sgarrella, F., Del Corso, A., Tozzi, M. G., and Camici, M. (1992) Biochim. Biophys. Acta 1118, 130-133[Medline] [Order article via Infotrieve]
14. Valentin-Hansen, P., Hammer-Jespersen, K., and Buxton, R. S. (1979) J. Mol. Biol. 133, 1-17[Medline] [Order article via Infotrieve]
15. Kawarabayasi, Y., Hino, Y., Horikawa, H., Yamazaki, S., Haikawa, Y., Jin-no, K., Takahashi, M., Sekine, M., Baba, S., Ankai, A., Kosugi, H., Hosoyama, A., Fukui, S., Nagai, Y., Nishijima, K., Nakazawa, H., Takamiya, M., Masuda, S., Funahashi, T., Tanaka, T., Kudoh, Y., Yamazaki, J., Kushida, N., Oguchi, A., Aoki, K., Kubota, K., Nakamura, Y., Nomura, N., Sako, Y., and Kikuchi, H. (1999) DNA Res. 6, 83-101[Medline] [Order article via Infotrieve], 145-152
16. Sako, Y., Nomura, N., Uchida, A., Ishida, Y., Morii, H., Koga, Y., Hoaki, T., and Maruyama, T. (1996) Int. J. Syst. Bacteriol. 46, 1070-1077[Abstract]
17. Laemmli, U. K. (1970) Nature 227, 680-685[Medline] [Order article via Infotrieve]
18. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual , 2nd Ed. , pp. 9.14-9.23, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
19. Wong, C. H., Garcia-Junceda, E., Chen, L., Blanco, O., Gijsen, H. J. M., and Steensma, D. H. (1995) J. Am. Chem. Soc. 117, 3333-3339
20. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve]
21. Cleland, W. W. (1971) in The Enzymes (Boyer, P. D., ed), 3rd Ed., Vol. 2 , pp. 1-65, Academic Press, Inc., New York
22. Rossmann, M. G., and van Beek, C. G. (1999) Acta Crystallogr. Sect. D Biol. Crystallogr. 55, 1631-1640[CrossRef][Medline] [Order article via Infotrieve]
23. Collaborative Computational Project Number 4. (1994) Acta Crystallogr. Sect. D Biol. Crystallogr. 50, 760-763[CrossRef][Medline] [Order article via Infotrieve]
24. Terwilliger, T. C., and Berendzen, J. (1999) Acta Crystallogr. Sect. D Biol. Crystallogr. 55, 849-861[CrossRef][Medline] [Order article via Infotrieve]
25. Terwilliger, T. C. (2000) Acta Crystallogr. Sect. D Biol. Crystallogr. 55, 1863-1871[CrossRef]
26. McRee, D. E. (1992) J. Mol. Graphics 10, 44-46[CrossRef]
27. Brunger, A. T., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P., Grosse-Kunstleve, R. W., Jiang, J. S., Kuszewski, J., Nilges, M., Pannu, N. S., Read, R. J., Rice, L. M., Simonson, T., and Warren, G. L. (1998) Acta Crystallogr. Sect. D. Biol. Crystallogr. 54, 905-921[CrossRef][Medline] [Order article via Infotrieve]
28. Brunger, A. T. (1992) Nature 335, 472-474[CrossRef]
29. Laskowski, R. A., MacArthur, M. W., Moss, D. S., and Thornton, J. M. (1993) J. Appl. Crystallogr. 26, 283-291[CrossRef]
30. Sazuka, T., and Ohara, O. (1996) DNA Res. 3, 225-232[Medline] [Order article via Infotrieve]
31. Nicholls, A., Sharp, K. A., and Honig, B. (1991) Proteins 11, 281-296[Medline] [Order article via Infotrieve]
32. Bernstein, H. J. (2000) Trends Biochem. Sci. 25, 453-455[CrossRef][Medline] [Order article via Infotrieve]
33. Heine, A., DeSantis, G., Luz, J. G., Mitchell, M., Wong, C. H., and Wilson, I. A. (2001) Science 294, 369-374[Abstract/Free Full Text]
34. Rice, D. W., Yip, K. S., Stillman, T. J., Britton, K. L., Fuentes, A., Connerton, I., Pasquo, A., Scandura, R., and Engel, P. C. (1996) FEMS Microbiol. Rev. 18, 105-117[CrossRef][Medline] [Order article via Infotrieve]
35. Knapp, S., de Vos, W. M., Rice, D., and Ladenstein, R. (1997) J. Mol. Biol. 267, 916-932[CrossRef][Medline] [Order article via Infotrieve]
36. Russell, R. J., Ferguson, J. M., Hough, D. W., Danson, M. J., and Taylor, G. L. (1997) Biochemistry 36, 9983-9994[CrossRef][Medline] [Order article via Infotrieve]
37. Hashimoto, H., Inoue, T., Nishioka, M., Fujiwara, S., Takagi, M., Imanaka, T., and Kai, Y. (1999) J. Mol. Biol. 292, 707-716[CrossRef][Medline] [Order article via Infotrieve]
38. Chan, M. K., Mukund, S., Kletzin, A., Adams, M. W., and Rees, D. C. (1995) Science 267, 1463-1469[Medline] [Order article via Infotrieve]
39. Maeda, N., Kanai, T., Atomi, H., and Imanaka, T. (2002) J. Biol. Chem. 277, 31656-31662[Abstract/Free Full Text]
40. Dams, T., Auerbach, G., Bader, G., Jacob, U., Ploom, T., Huber, R., and Jaenicke, R. (2000) J. Mol. Biol. 297, 659-672[CrossRef][Medline] [Order article via Infotrieve]
41. Hennig, M., Sterner, R., Kirschner, K., and Jansonius, J. N. (1997) Biochemistry 36, 6009-6016[CrossRef][Medline] [Order article via Infotrieve]
42. Jeanmougin, F., Thompson, J. D., Gouy, M., Higgins, D. G., and Gibson, T. J. (1998) Trends Biochem. Sci. 23, 403-405[CrossRef][Medline] [Order article via Infotrieve]
43. Kraulis, P. (1991) J. Appl. Crystallogr. 24, 946-950[CrossRef]
44. Merritt, E. A., and Murphy, M. E. P. (1994) Acta Crystallogr. Sect. D Biol. Crystallogr. 50, 869-873[CrossRef][Medline] [Order article via Infotrieve]


Copyright © 2003 by The American Society for Biochemistry and Molecular Biology, Inc.