High-Resolution Structures of RmlC from Streptococcus ... - Cell Press

Structure, Vol. 11, 715–723, June, 2003, 2003 Elsevier Science Ltd. All rights reserved.

DOI 10.1016/S0969-2126(03)00098-4

High-Resolution Structures of RmlC from Streptococcus suis in Complex with Substrate Analogs Locate the Active Site of This Class of Enzyme Changjiang Dong,1 Louise L. Major,1 Andrew Allen,2,3 Wulf Blankenfeldt,1,4 Duncan Maskell,2 and James H. Naismith1,* 1 Centre for Biomolecular Science The University St. Andrews KY16 9ST United Kingdom 2 Centre for Veterinary Science University of Cambridge Cambridge CB3 0ES United Kingdom

Summary Nature achieves the epimerization of carbohydrates by a variety of chemical routes. One common route is that performed by the class of enzyme defined by dTDP-6-deoxy-D-xylo-4-hexulose 3,5-epimerase (RmlC) from the rhamnose pathway. Earlier studies failed to identify the key residues in catalysis. We report the 1.3 A˚ structure of RmlC from Streptococcus suis type 2 and its complexes with dTDP-D-glucose and dTDPD-xylose. The streptococcal RmlC enzymes belong to a separate subgroup, sharing only 25% identity with RmlC from other bacteria, yet the S. suis enzyme has similar kinetic properties and structure to other RmlC enzymes. Structure, sequence alignment, and mutational analysis have now allowed reliable identification of the catalytic residues and their roles. Introduction The molecular diversity of carbohydrates and their role in molecular recognition has for some time been an important area of biology. For instance, there are 16 diastereoisomers of D-hexulose. This range is possible because of the very dense functionalization of the molecule. Of the six carbons in hexulose, four are asymmetric centers and a fifth is created on cyclization. Further chemical modification of the hydroxyl groups leads to even greater diversity. Rather than synthesize each carbohydrate scaffold from scratch, biology has evolved enzymes to manipulate a relatively small number of carbohydrate skeletons to harness the diversity that the carbohydrate molecule allows. Bacteria utilize a number of sugars that are not found in humans, and their synthesis provides potential targets for novel anti-infectives. One such sugar is L-rhamnose, a 6-deoxysugar found in many complex carbohydrates, especially as a component of cell walls and envelopes of many pathogenic bacteria such as Mycobacterium tuberculosis (McNeil et al., 1990), Salmonella enterica *Correspondence: [email protected] 3 Present address: Arrow Therapeutics Ltd., Britannia House, 7 Trinity Street, Borough London SE1 1DA, United Kingdom. 4 Present address: Max Planck Institute of Molecular Physiology, Otto-Hahn-Str. 11, 44227 Dortmund, Germany.

serovar Typhimurium (Graninger et al., 1999), and Streptococcus suis serotype 2 (Elliott and Tai, 1978). The last organism is a particular problem in pigs, where untreated the infection can kill 20% of a herd (Staats et al., 1997). Rhamnose has been found as part of the capsule that is key to the organism’s virulence (Smith et al., 1999). L-rhamnose is synthesized from ␣-D-glucose-1-phosphate. When compared to ␣-D-glucose, L-rhamnose lacks a hydroxyl at C6 and the chirality is inverted at C3 and C5. The chemical transformations necessary for these changes are carried out by four enzymes (Melo and Glaser, 1968; Giraud and Naismith, 2000): ␣-D-glucose-1-phosphate thymidylyl transferase (Blankenfeldt et al., 2000; RmlA, EC: 2.7.7.24), dTDPD-glucose-4,6-dehydratase (Allard et al., 2001a, 2001b; RmlB, EC: 4.2.1.46), dTDP-6-deoxy-D-xylo-4-hexulose 3,5-epimerase (Giraud et al., 2000; Christendat et al., 2000; RmlC, EC: 5.1.3.13), and dTDP-6-deoxy-L-lyxo4-hexulose-4-reductase (Blankenfeldt et al., 2002; RmlD, EC: 1.1.1.133). The structures and mechanisms of all four enzymes have been studied in our laboratory. This report focuses on the mechanism of RmlC (Figure 1). Nature employs a variety of chemical reactions to epimerize the stereogenic carbon centers in the carbohydrate skeleton (Allard et al., 2001c). A common method is to activate a proton for removal by placing a keto group ␣ to the C-H bond. The result is to lower substantially the pKa of the proton. This is manifested in the well-known keto enol tautomerization (of such keto groups). Even an activated proton does not yet fall within the range of bases available in biology. However, many enzymes, not just those in carbohydrate epimerization, can abstract the proton ␣ to a keto function. A key feature in these reactions is that the enzyme stabilizes the negative charge of the enolate intermediate. RmlC, the third enzyme in the rhamnose pathway, was first identified over 30 years ago (Kornfeld and Glaser, 1961). Some controversy existed as to whether the enzyme performed the double epimerization itself or as a complex with the fourth enzyme, RmlD. This has now been settled by two papers (Graninger et al., 1999; Stern et al., 1999) that establish that RmlC is responsible for both epimerization at C3 and C5 (Figure 1). These studies also show that the equilibrium for the reaction lies heavily to the substrate with less than 10% conversion (Graninger et al., 1999; Stern et al., 1999). In vivo, the product dTDP-6-deoxy-L-lyxo-4-hexulose is consumed in the final thermodynamically favorable step, reduction to dTDP-L-rhamnose by RmlD (Graninger et al., 1999; Melo and Glaser, 1968). The substrate for RmlC, dTDP6-deoxy-D-xylo-4-hexulose, has a 4-keto group, which lowers the pKa of the protons attached to C3 and C5. In broad outline the mechanism seems clear: deprotonation from one face at the C3 position, followed by reprotonation at the opposite face at the C3 position, deprotonation from one face at C5, and reprotonation at the opposite face at C5. This constitutes four distinct chemical steps, all of which require careful steric control. This type of sequential chemistry defines a class of carbohy-

Structure 716

Figure 1. The Sequential Steps in the Conversion of dTDP-6-Deoxy-D-Xylo-4-Hexulose to dTDP-6-Deoxy-L-Lyxo-4-Hexulose by RmlC, as Conventionally Written Our data do not define the order of epimerization, and therefore the choice of C5 first is arbitrary. dTMP represents deoxythymidine monophosphate. The substrate mimics used in this study, dTDP-D-glucose and dTDP-D-xylose, are shown.

drate epimerases of which RmlC may be considered the paradigm. Examples of this enzyme activity are wide spread in bacterial biology. RmlC homologs are involved in the synthesis of carbohydrates such as gulose (Zhang et al., 1997) and antibiotics such as vancomycin (Chen et al., 2000) and tylosin (He et al., 2000). The structures of RmlC from Salmonella enterica serovar Typhimurium (Giraud et al., 2000) and Methanobacteria thermoautotrophicum (Christendat et al., 2000) have been reported recently. These studies showed that the protein existed as a functional dimer and was primarily formed of ␤ sheets. In addition, using dTDP (or an analog), they identified the nucleotide region of the binding site. Using this and inferences from sequence alignments, the location of the enzyme active site was predicted. To date, no substrate or substrate analog complex structures have been presented. The sequences from streptococci were not available when the two groups reported the RmlC structures. As a result, both studies highlighted a pair of apparently absolutely conserved His-Asp diads (Giraud et al., 2000; Christendat et al., 2000). However, in the streptococcal enzymes, only one diad is apparently conserved (EMBL AJ509828). An alignment of 25 RmlC sequences shows (data not shown) that the streptococcal homologs are outliers of the RmlC superfamily, having 25% identity rather than a more typical 40%–70%. To gain a better insight into the mechanism of C3 and C5 epimerization, we have confirmed that the S. suis enzyme is a competent RmlC, and determined the structure of this enzyme to very high resolution and in complex with two substrate analogs. Our results locate the active site experimentally, and unambiguously identify the key residues involved in catalysis. The results suggest that considerable rearrangement of substrate and/or protein side chains occurs during turnover.

Results and Discussion S. suis RmlC Is Kinetically Competent as a 3,5-Epimerase A simple kinetic analysis of the S. suis enzyme gives an apparent kcat of 10.5 ⫾ 0.3 s⫺1 and an apparent Km of 0.029 ⫾ 0.003 mM. The values are apparent, as the assay system involves coupling three enzymes together. Using identical conditions and employing the S. enterica serovar Typhimurium enzyme gives an apparent kcat of 19.2 ⫾ 0.5 s⫺1 and an apparent Km of 0.081 ⫾ 0.008 mM. Although not identical, these values are close enough to confirm that the S. suis protein is catalytically competent to perform the 3,5-epimerase reaction. Thus, the residues that perform the chemical steps in the catalysis in S. suis are likely to be conserved in sequence and space. The mutants H76N, K82A, Y133F, and D180A all showed decrease in activity. In particular, we were unable to detect activity above background from H76N, K82A, and Y140F (Table 1).

Table 1. Apparent Kinetic Constants for RmlC Mutants in an RmlB, RmlC, and RmlD Coupled Assay

S. enterica serovar Typhimurium RmlC S. suis RmlC S. suis RmlC H76N S. suis RmlC K82A S. suis RmlC Y140F S. suis RmlC D180A a

Km (mM)

Kcat (s⫺1)

0.081 ⫾ 0.008

19.2 ⫾ 0.5

0.029 ⫾ 0.003 n.d.a n.d.a n.d.a 0.071 ⫾ 0.008

10.5 ⫾ 0.3 n.d.a n.d.a n.d.a 0.148 ⫾ 0.007

Kinetic constants were not determined for enzymes exhibiting less than one-thousandth of the wild-type enzyme activity. This was close to background (no RmlC) activity in the assay.

Cocomplexes of S. suis RmlC 717

Figure 2. The Location of the Active Site A ribbon diagram of the RmlC homodimer from S. suis (monomer A is in green and monomer B is in orange); molecules of dTDP-D-glucose are shown in space-filling representation. Oxygen atoms are shown in red, nitrogen in blue, carbon in black, and phosphorus in purple.

Overall Structure RmlC from S. suis is a homodimer (Figure 2). Each monomer consists of 197 amino acids (Met1–Leu197), with a molecular mass of 22,433 Da. The residues Met1 and Thr2 were not located in electron density maps. As expected, the topology of the enzyme differs little from that reported for other RmlC proteins. It is mainly ␤ sheet with a jelly roll-like structure, consisting of 11 ␤ strands and 7 ␣ helixes. It can be divided into three parts, namely a near N-terminal portion, a core, and a C-terminal portion, which together form a sandwich-like shape. The near N-terminal part consists of a ␤ sheet (antiparallel ␤3 and ␤4) and some helical structure. The core consists of nine ␤ strands in two ␤ sheets that form the active site cleft. One ␤ sheet consists of six antiparallel strands (␤1, ␤2, ␤5, ␤6, ␤9, and ␤11), while the other consists of three antiparallel strands (␤7, ␤8, and ␤10). The active site is located in the core. The C terminus is mainly helical. Strand exchange from the monomers stabilizes the dimer in the same manner reported previously for RmlC from S. enterica and M. thermoautotrophicum (Giraud et al., 2000; Christendat et al., 2000). The exchanged strand also helps to form the active site. The dimer buries in total about 2700 A˚2, or 13% of available surface area. In comparing monomers within a crystal, residues 3–140 are relatively insensitive to crystal packing. In the three structures reported here, the root-meansquare deviation (rmsd) from perfect noncrystallographic symmetry (ncs) for this region varies between

0.2 A˚ and 0.28 A˚ for 138 C␣ atoms. Including the more flexible C-terminal region increases the rmsd from perfect ncs to above 0.7 A˚ in some cases. The structures show that the second of the highly conserved His-Asp diads (H120 D84, S. enterica numbering) is definitely absent. Instead, the N127 and G94 predicted on sequence alignment are found at these positions. The other His-Asp diad (H76 and D180) as well as the TyrAsn couple (Y140 and N63) and the hydrophilic residues (S65 and S67) are conserved. Comparing the structure of the S. suis enzyme with the S. enterica Typhimurium using the Brute Force method in LSQMAN (Kleywegt, unpublished program) gives an rmsd of over 1.4 A˚ for 140 C␣ atoms in the two apo structures. This degree of deviation probably confounded our molecular replacement approaches. Changes in Structure on Ligand Binding The dTDP-D-xylose complex has four monomers in the asymmetric unit. Although all four have essentially the same structure, monomer B is less well ordered than the others and precise distances between atoms of the carbohydrate and protein differ for this monomer. The other three monomers are essentially the same and we concentrate on one (monomer A in the PDB file) in the following analysis. There are two monomers in the asymmetric unit of the dTDP-D-glucose complex structure and in the native structure, and as there is no significant difference between them, we use monomer A in the

Structure 718

discussion. A comparison of the core structure region shows the two complexes have an rmsd of 0.38 A˚ (c.f. 0.27 A˚ for ncs). However, in comparison to the apo structure, this figure rises to about 1.0 A˚. This is a substantial change in the structure as a result of ligand binding. The conformation of the loop G30–G34 is significantly perturbed and helps form the substrate binding site of the neighboring monomer in the dimer. The change in structure permits R33 to move to form a salt bridge with the ␣-phosphate of the ligand in the other monomer. The conformation of R73 adjusts to make a salt bridge with the ␤-phosphate. The structure at W146–Y154 changes to form the pocket for the thymine ring, as previously noted (Giraud et al., 2000; Christendat et al., 2000). W146 moves in to stack against the thymine ring. This in effect closes the active site around the substrate. The Nucleotide Binding Pocket Is Unchanged from Previous Description We had used dTDP-phenol (Giraud et al., 2000; Christendat et al., 2000), and Christendat et al. (2000) had used dTDP to locate the nucleotide binding pocket of RmlC. The complexes from this study show that the thymine ring and phosphate groups are bound as reported previously in subunit A of the S. enterica serovar Typhimurium enzyme and the M. thermoautotrophicum enzyme. The ligand adopts a U shape in the protein rather than the extended conformation one would expect in solution. The Carbohydrate Rings Locate the Active Site The sugar ring of dTDP-D-glucose is located in the entrance to the ␤ sandwich (Figure 2). The carbohydrate ring is completely enveloped, with over 90% of the surface area of the sugar being withdrawn from solvent upon complexation. The O2 atom is bound to two water molecules that are part of an extensive hydrogen bond network. The O3 atom is hydrogen bonded to the NZ atom of the absolutely conserved K82 with a distance of 2.7 A˚. E78 makes a bidentate hydrogen bond to O2 (2.6 A˚) and O3 (3.5 A˚) of the sugar (Figure 2). This type of interaction is commonly observed in protein-carbohydrate complexes and correlates with affinity (Vyas, 1991). A water molecule is weakly bound to the O3 atom (3.4 A˚) and sits above the C3 position. The water is hydrogen bonded to W80 and Y140. The O4 atom, where the enolate is expected to form, is bound to ND2 of the nonconserved N127 (2.7 A˚) and a water molecule. dTDPD-glucose contains an additional hydroxyl group relative to the true substrate (Figure 1). This hydroxyl is at a 2.8 A˚ distance from the OG of the absolutely conserved S65 and 2.8 A˚ from the O5 oxygen of the sugar. The ND1 atom of H76 is 3.8 A˚ from the sugar C3 atom and 5.0 A˚ from the sugar C5 atom, the atoms from which the protons would be extracted. The hydroxyl group of Y140 is 3.7 A˚ from C3 and 3.5 A˚ from C5. Y140 also makes van der Waals contacts (⬍4.0 A˚) with the hydrophobic patch of the sugar (C4, C5, and C6; Vyas, 1991). Two aromatic residues, Y138 and F129, make van der Waals contacts with C6 and O6 of the substrate. Although not absolutely conserved in RmlC proteins,

these residues are very commonly both found as phenylalanine (⬎95% for F129 and ⬎50% for Y138). The substitutions that are found are conservative for bulky hydrophobic or aromatic residues. The dTDP-D-xylose complex is subtly different from the dTDP-D-glucose complex (Figure 3). Although the nucleotide positions are identical and the carbohydrate rings are coplanar, the pyranose ring is rotated by about 20⬚ around the glycoside linkage between the structures (Figure 3). O2 remains bonded to the same two water molecules. O3 is now 2.9 A˚ from the NZ atom of K82. E78 continues to make a bidentate hydrogen bond to O2 (3.5 A˚) and O3 (3.0 A˚). A water molecule sits in the same position above C3 and remains hydrogen bonded to O3, W80, and Y140. O4 is bonded to NZ of K82 (3.0 A˚) and to ND2 of N127 (2.9 A˚). There is less van der Waals contact than for the larger dTDP-D-glucose molecule, as might be expected for a smaller substrate. ND1 of H76 is now 4.1 A˚ from C3 and 5.4 A˚ from C5. The orientation of the His side chain relative to the pyranose ring in the A subunit is correct for proton abstraction; however, the angle of the H76 ring does vary in the other subunits. The OH of Y140 is 3.8 A˚ from C3 and 4.0 A˚ from C5. The methyl group that would be attached to C5 in the true substrate would point toward the same hydrophobic pocket that C6 in dTDP-D-glucose binds. The conformation of H76 differs between the complexes and is consistent with ncs differences in the dTDP-D-xylose structure, suggesting that this residue is flexible. From both structures, it appears that E78 plays an important role in recognition in both sugars, making the classical bidentate hydrogen bond (Vyas, 1991). However, this is puzzling because neither this residue nor this region is particularly well conserved in structure. The residue is in a stretch where there is a deletion of 4 residues relative to the majority of RmlC sequences. The result is that the loop that connects the absolutely conserved His and Lys residues (76 and 82 in S. suis; 63 and 73 in S. enterica) is pulled in to the active site in S. suis but bulges out and away in S. enterica. A number of nonconservative substitutions in this area would indicate that this bidentate interaction is unique to the S. suis structure. Below the carbohydrate ring, from where the proton(s) must be extracted, there is no other conserved residue other than H76, nor do any potential proton acceptors from nonconserved residues approach within 6 A˚ of either the C3 or C5 positions from below the face.

Mechanistic Implications The results of the structural analysis help greatly to determine the initial steps and to clarify the basic outline of the mechanism of the 3,5-epimerization. The first step of the reaction is the abstraction of the proton from either C3 or C5. Our results shed no light on which step occurs first. H76 is the only logical choice for this active site base. This is, of course, chemically reasonable, and is consistent with why both H76 and D180 are conserved in all RmlCs. The aspartic acid serves to increase the basicity of H76. The D180A mutant is catalytically competent but shows decreased activity consistent with this role. The second His-Asp diad is not found in any of the

Cocomplexes of S. suis RmlC 719

Figure 3. The dTDP-D-Glucose and dTDP-D-Xylose Complexes of S. suis RmlC (A) Superposition of dTDP-D-xylose and dTDP-D-glucose with the key residues mentioned in the text drawn. The color scheme of the dTDPD-glucose complex is as Figure 2A, except that carbon atoms are in green for the dTDP-D-xylose complex and orange for the dTDPD-glucose complex. The water molecule is in an identical position in the two structures. (B) The Fo ⫺ Fc map for the dTDP-D-glucose (left) and dTDP-D-xylose (right) complexes. In initial maps, the density for dTDP was clear but the sugars were less clear. Refinement continued with dTDP until density for each of the sugars was clear. dTDP was then removed from the model and further refinement was carried out. The resulting phases were used to calculate the above Fo ⫺ Fc maps. The maps are shown in green and are contoured at 2.5␴.

streptococcal enzymes and therefore seems unlikely to have a direct role in catalysis. The relatively conservative substitution of this second His residue to Asn in streptococcal sequences suggests that the actual role of this residue is limited to substrate recognition. After the first proton is abstracted from C3 or C5, an enolate anion will be formed. The pKa of these protons is likely to be substantially lowered by the presence of K82, which is positioned to make a salt bridge with O4 in the dTDP-D-xylose complex (3.0 A˚), thus stabilizing the negative charge. However, in the dTDP-D-glucose complex, the sugar is not orientated in such a way that this is possible (distance 5.0 A˚), possibly reflecting that dTDP-glucose is an imperfect mimic because it is a larger molecule than the true substrate.

The dTDP-D-glucose and dTDP-D-xylose complex structures show that the sugar ring can move within the active site but that the orientation of the plane of the carbohydrate ring relative to the protein remains fixed by the axial configuration of the glycosidic bond. However, these complexes also show that changing the orientation of the plane of the ring by 180⬚ would require substantial changes in protein structure, if severe steric clashes were not to occur during rotation. In addition, buried water molecules would have to be expelled and reenter the active site during such a rotation. A rotation of the carbohydrate ring underlies the chemistry of UDPgalactose epimerase (Thoden et al., 1996), but does not appear to have a role here. The UDP-galactose epimerase is a member of the SDR superfamily (Rossmann

Structure 720

Figure 4. The Principal Mechanistic Steps of RmlC The order of the epimerization is not defined by our data; the choice for this figure is arbitrary. The roles of the key residues are shown.

et al., 1975) and is unrelated in sequence or structure to RmlC. A number of experiments have shown that epimerization proceeds by proton exchange at C3 and C5. An acid must donate a different proton to the carbohydrate ring from the top face to invert the chirality. Examination of the top face reveals only the conserved tyrosine residue (Y140) in a position to donate a proton. Tyr has a pKa of around 10 and would not normally be considered a good acid. Contrary to the situation in the short-chain dehydrogenase superfamily (Rossmann et al., 1975), there is no electrostatic field to lower its pKa. There is a conserved asparagine residue (N63) that hydrogen bonds to the OH atom of Y140, which may marginally stabilize the resulting tyrosinate. The role of tyrosine as the acid seems inescapable as it is correctly positioned relative to C5 of the sugar, is absolutely conserved in all RmlC sequences (Figure 4), and its mutation compromises the enzyme. Whereas the residues involved in one of the epimerizations at one position seem clear, the mechanism of the second epimerization is less obvious. H76 must again act as the base and our structures have shown that its conformation is flexible. However, the residue must lose a proton before it can abstract a second one from the substrate. In both dTDP-D-xylose and dTDP-glucose complexes, a water molecule is hydrogen bonded to H76 and forms a wider network. The structural data do not reveal any alternative pathway of deprotonation of H76. K82 would have the same role for both epimerizations, as both generate an enolate intermediate (Figure 4).

Whether Y140 functions as an acid for the two epimerizations (in which case it would need to be reprotonated) is less clear. The structure shows a conserved water molecule above the C3 position. It may be that water functions as an acid for one epimerization. The water is found in a network aiding proton donation. If the water is not a proton donor, it may play a role in resetting the reprotonating Y140 after the first epimerization. It is clear from the discussion of the structures that the complexes presented here are not precise mimics of the situation during catalysis. The distances between key atoms are too long for proton transfer. Such a limitation is to be expected because neither sugar has the required keto function. In addition, one is larger by a hydroxyl group and the other is smaller by a methyl group than the true substrate. One possibility is that the sugar moves in the active site toward the base then acid in sequence as its conformation changes during the reaction. Alternatively, the protein active site clamps down more on the substrate during turnover, and that only by using a better substrate mimic will this be observed. Further studies will be required to address this detail. In summary, the structures and functional data presented here advance our understanding of this ubiquitous reaction. The enzyme uses a single base, a conserved His-Asp diad, to carry out proton abstractions from both positions, which are epimerized. The resulting enolate intermediate is formed twice during turnover and is stabilized by an absolutely conserved lysine residue. Epimerization requires that a proton is added to the

Cocomplexes of S. suis RmlC 721

sugar from the other face. Our work strongly indicates that for at least one of the epimerizations, a conserved tyrosine residue acts as the acid. The structures show that water could act as the second proton source. Biological Implications Bacteria present a continuing threat to human and animal health. Although antibiotics are controlling the situation, there is a growing realization that new antibiotics need to be developed. Bacteria evade the host immune system by a variety of means including use of a carbohydrate-rich capsule. Loss of the capsule can make the organism avirulent, as it often becomes susceptible to host defenses such as complement and phagocytosis. L-rhamnose is found in the capsules of streptococci as well as other bacteria, but is not found in humans. Its synthesis is thus an appealing target for intervention. L-rhamnose is synthesized by bacteria from D-glucose by inverting the chirality at C3 and C5 and dehydrating at C6. The inversion of chirality is carried out by a single enzyme, namely dTDP-6-deoxy-D-xylo-4-hexulose 3,5epimerase (RmlC). This enzyme defines a class of such epimerases that are found in many different carbohydrate biosynthetic pathways. Although this class of enzyme has been known for over 30 years, the catalytic site and hence the key residues were unknown. We have determined the structure of RmlC from Streptococcus suis, a pig pathogen, in complex with substrate analogs. Sequence and structural alignments coupled to site-directed mutagenesis has defined the active site, the catalytic residues, and their roles for this enzyme class. There are obvious applications of this knowledge in the design of specific inhibitors of this enzyme. There are also potential applications in altering the enzyme behavior for use in the synthesis of bioactive carbohydrates. Experimental Procedures Protein Expression, Purification, and Crystallization E. coli BL21 (DE3) cells were transformed with a pET-21 (⫹) plasmid containing the rmlc gene from S. suis type 2. The construct does not alter the sequence or add additional residues compared to the published gene sequence. Cells were grown in Terrific Broth culture medium containing 200 ␮g/ml of ampicillin at 37⬚C until A600 nm reached between 0.6 and 0.8. Overexpression was induced with 1 mM IPTG at 25⬚C for 6 hr. The induced cells were harvested at 5000 rpm for 15 min and resuspended in lysis buffer (100 mM NaCl, 2 mM DTT, 20 ␮M lysozyme, 20 mM Tris-HCl [pH 8.0]) at room temperature for 30 min. Five millimolar PMSF and 20 ␮g/ml DNase I were added, and the cells were disrupted by sonication for six cycles of 30 s, interrupted by 1 min periods on ice. After addition of 2 mM EDTA, the mixtures were centrifuged for 30 min at 20,000 rpm at 4⬚C. The supernatant was brought to 20% (NH4)SO4 saturation and kept at 4⬚C for 2 hr. The sample was centrifuged at 10,000 rpm for 30 min at 4⬚C and the supernatant was dialyzed against 50 mM NaCl, 2 mM DDT, 20 mM Tris-HCl (pH 8.5). The dialyzed sample was concentrated using an Amicon concentrator with a 10 KDa cutoff membrane. The sample was filtered by 0.22 or 0.45 ␮m syringe filters. The protein was applied to a POROS HQ/M 16 ⫻ 100 mm column and eluted by an increasing concentration gradient of NaCl. Fractions containing S. suis RmlC were pooled, brought to 45% saturated (NH4)SO4, and applied to a POROS HP2/M 10 ⫻ 100 mm column. Protein was eluted by a decreasing gradient of (NH4)SO4 concentration. Protein purity was confirmed by silver-stained SDS gels, integrity by MALDI-TOF mass spectrometry (whole protein and tryptic fragmentation), and its activity by coupled assay. Protein with

selenomethionine was expressed using the methionine biosynthesis pathway inhibition protocol (Doublie, 1997). This protein purified in the normal manner and substitution of sulfur for selenium was confirmed by mass spectroscopy. Crystals of native and selenomethionine protein were obtained from a sitting drop (4 ␮l protein at 7 mgml⫺1 and 4 ␮l reservoir) vapor diffusion against a 100 ␮l reservoir solution containing 0.2 M MgCl2, 0.1 M Tris-HCl (pH 8.5), 25% PEG 2000. Large plate crystals grew to full size (1.1 ⫻ 1.0 ⫻ 0.3 mm) in about 8 days at 20⬚C. These crystals were highly mosaic but addition of 4 mM NiCl2 during crystallization improved the quality and increased the size of the crystals. Crystals in complex with dTDP-D-glucose and dTDP-D-xylose were obtained by the same sitting drop procedure as native crystals with protein at 7 mgml⫺1. The reservoir for the dTDP-D-glucose complex was 0.1 M Tris-HCl (pH 8.4), 4 mM NiCl2, 40% PEG 2000 with the protein preincubated with 20 mM dTDP-D-glucose. The reservoir for the dTDP-D-xylose complex was 0.1 M Tris-HCl (pH 8.2), 4 mM NiCl2, 35% PEG 2000 with the protein preincubated with 10 mM dTDP-D-xylose. Mutagenesis Assay Site-specific mutations H76N, K82A, Y140F, and D180A were made using the plasmid encoding RmlC and Pfu DNA polymerase according to the Stratagene QuickChange mutagenesis protocol (Stratagene) with the following oligonucleotides: 3⬘-GTTCTTCGTGGTCTTGCCGCAGAGCCTTGGGAT-5⬘ and 3⬘-ATC CCAAGGCTCTGCGGCAAGACCACGAAGAAC-5⬘ for H76N; 3⬘-GCAG AGCCTTGGGATGCATACATTTCAGTAGCA-5⬘ and 3⬘-TGCTACTGA AATGTATGCATCCCAAGGCTCTGC-5⬘ for K82A; 3⬘-TTCGTGGCTTA CAGCTTCTTGGTTAACGACTAC-5⬘ and 3⬘-GTAGTCGTTAACCAAGAA GCTGTAAGCCACGAA-5⬘ for Y140F; 3⬘-GAAGTTTCAGAAGCAGCT GAAAACCACCCATTC-5⬘ and 3⬘-GAATGGGTGGTTTTCAGCTGCTT CTGAAACTTC-5⬘ for D180A. Mutated bases are underlined. The mutants were expressed and purified as the native protein. The mutants were confirmed by gene sequencing. The assay we used was based on the coupled assay of RmlC and RmlD developed by Graninger et al. (1999). The Graninger assay used the RmlC substrate dTDP-6-deoxy-D-xylo-4-hexulose. We were unable to synthesize sufficient quantity of pure compound to accurately quantitate it for the assay. Instead, we used the RmlB substrate dTDP-glucose and included RmlB in excess in the assay, thus coupling the activities of RmlB, RmlC, and RmlD. This strategy has previously been used to determine RmlD kinetic parameters (Blankenfeldt et al., 2002). Recombinant S. enterica serovar Typhimurium RmlB and RmlD proteins were used for all assays. These enzymes have native sequence, were overexpressed from pET vectors, and purified from E. coli BL21 (DE3) cells. In our assay we used 20 mM Tris (pH 7.5), 9 mM MgCl2, 0.3 mM NADPH (approximately 10,000-fold molar excess and 50-fold over the reported Km for NADPH with RmlD; Graninger et al., 1999), 1.3 ␮M RmlB (approximately 48-fold molar excess), 0.98 ␮M RmlD (approximately 30-fold molar excess), and activity was determined at 21⬚C. RmlC activity was determined by measuring the RmlD oxidation of NADPH at A340 nm. The rates measured were independent of the amount of RmlB or RmlD in the assay (both present in excess) and were dependent upon the amount of RmlC present in the assay. For the S. suis enzymes, dTDP-glucose concentrations ranged from 0.0025 to 0.34 mM, and for the S. enterica serovar Typhimurium, RmlC dTDP-glucose concentrations of 0.0125–1.7 mM were used. Our values differ from the values reported previously for the S. enterica serovar Typhimurium enzymes, but are within a factor of 10. In our analysis, it is the comparison between native and mutant rather than the absolute values that is important. Data Collection, Structure Determination, and Refinement Crystals were mounted in a loop and then placed in a stream of N2 at 120K for all data collection. The reservoir solution on its own was sufficient to prevent ice ring formation or crystal damage during freezing. A native data set, recorded on a single native crystal, was collected in two passes with a wavelength of 0.933 A˚ using QUANTUM CCD detector at ID14-1 ESRF. For the high-resolution

Structure 722

Table 2. Crystallographic Data Data Collection

SeMet

Native

dTDP-Xylose

dTDP-Glucose

Wavelength (A˚) Beamline Resolution (highest shell, A˚) Space group Cell constants (A˚; ⬚)

0.978765 BM14, ESRF 43.59–2.18 (2.31–2.18) P21 a ⫽ 46.0, b ⫽ 82.1, c ⫽ 52.4; ␣ ⫽ 90, ␤ ⫽ 108.49, ␥ ⫽ 90 2.1 329,957 19,088 6.6 (3.1) 7.0 (6.5) 98.7 (98.7) 97.1 (82.8) 5.9 (9.0) 32.0 10.4/6.1

0.934 ID 14.1, ESRF 28.30–1.30 (1.33–1.30) P21 a ⫽ 45.7, b ⫽ 81.6, c ⫽ 52.1; ␣ ⫽ 90, ␤ ⫽ 108.8, ␥ ⫽ 90 2.1 1,012,876 85,539 5.9 (2.8) 11.7 (3.4) 95.2 (84.0)

1.488 SRS 14.1, Daresbury 50.79–1.80 (1.81–1.80) P21 a ⫽ 50.9, b ⫽ 140.9, c ⫽ 53.7; ␣ ⫽ 90, ␤ ⫽ 92.7, ␥ ⫽ 90 2.1 659,635 63,995 5.3 (2.1) 3.8 (8.1) 91.8 (43.3)

0.933 ID14.2, ESRF 44.28–1.60 (1.69–1.60) P21 a ⫽ 50.6, b ⫽ 47.9, c ⫽ 72.4; ␣ ⫽ 90, ␤ ⫽ 99.0, ␥ ⫽ 90 2.0 383,364 43,352 3.5 (2.7) 5.2 (1.8) 95.7 (79.8)

3.9 (24.1) 14.11

11.7 (8.5) 21.39

8.3 (29.2) 24.3

Resolution (highest shell; A˚)

49.39–1.30 (1.33–1.30)

70.71–1.60 (1.64–1.60)

R (%) Rfree (%) Rmsd bonds (A˚)/angles (⬚) B factor deviation Bonds/angles (A˚2) Main chain Side chains Residues in Ramachandran core (%) Protein atoms Water atoms Ligand atoms Ion atoms (Ni) Average B factor (A˚2) PDB ID code

14.3 (19.5) 17.4 (22.0) 0.013/1.48

70.71–1.80 (1.847–1.80) 16.4 (14.8) 21.6 (26.0) 0.014/1.58

17.3 (31.2) 21.7 (30.2) 0.019/1.82

1.3/1.9 2.7/4.0 93.7 3214 633 0 0 9.3 1NXM

0.8/1.3 1.8/2.9 91.8 6304 874 136 2 17.9 1NZC

1.6/2.0 2.7/3.7 92.5 3151 508 72 0 18.9 1NYW

VM Total measurements Unique reflections Average redundancy I/␴ Completeness (%) Anomalous completeness Rmerge Wilson B factor (A˚2) f⬘/f″ Refinement

pass, 1.3 A˚ data were collected as 200 3 s 1⬚ oscillations. The lower resolution pass, 1.6 A˚ data, was collected as 200 1 s 1⬚ oscillations. The data were indexed and integrated using MOSFLM (Leslie, 1992). The crystal belongs to space group P21 with cell dimensions a ⫽ 45.7 A˚, b ⫽ 81.6 A˚, c ⫽ 52.1 A˚, ␣ ⫽ ␥ ⫽ 90⬚, ␤ ⫽ 108.8⬚. Data were merged using SCALA (CCP4, 1994). Despite intensive efforts, we were unable to obtain a molecular replacement solution based on models from existing RmlC structures. Data to 2.4 A˚ were collected on a selenomethionine crystal at a wavelength of 0.9787 A˚, chosen to maximize the anomalous signal, at 100K on BM14 at the ESRF. These data were indexed and integrated using DENZO and SCALEPACK (Otwinowski and Minor, 1997). Data on the dTDP-D-glucose complex crystals, cell a ⫽ 50.6 A˚, b ⫽ 47.9 A˚, c ⫽ 72.4 A˚, ␣ ⫽ ␥ ⫽ 90⬚, ␤ ⫽ 99.0⬚, were collected on ID14-2 at the ESRF to 1.6 A˚. Data on the dTDP-D-xylose complex crystals, cell a ⫽ 50.9 A˚, b ⫽ 140.9 A˚, c ⫽ 53.7 A˚, ␣ ⫽ ␥ ⫽ 90⬚, ␤ ⫽ 92.7⬚, were collected at the SRS Daresbury source on ID14-1 to 1.8 A˚. Data from both complexes were indexed and integrated with MOSFLM (Leslie, 1992) and merged in SCALA (CCP4, 1994). Statistics for all data sets are given in Table 1. Phases were determined by straightforward application of the SAD option in SOLVE and RESOLVE (Terwilliger and Berendzen, 1999). An initial model was built manually in O (Jones et al., 1991) by adjusting a polyalanine trace of S. enterica RmlC. This structure was refined against the SAD data using REFMAC5 (Murshudov et al., 1997). At that time, the dTDP-D-glucose data set was the highest resolution data at hand. Phases for these data were determined with AMoRe (Navaza, 1994) using the initial model from the free enzyme. These phases were improved by noncrystallographic averaging and extended to 1.6 A˚ using DMMULTI (CCP4, 1994). ARP/ wARP (Perrakis et al., 1999) tracing with these phases produced an essentially complete model. After a small amount of manual

adjustment, including removal of water molecules from the nucleotide binding and active sites, the model was refined using REFMAC5 (Murshudov et al., 1997). Refinement proceeded smoothly. Clear electron density for dTDP-D-glucose was visible in the 2Fo ⫺ Fc and Fo ⫺ Fc electron density maps (Figure 3). This was included in the model and automated inclusion of waters carried out using ARP/ wARP. The refined protein structure (without waters and ligands) from this data set was then used to phase the dTDP-D-xylose complex data. Refinement of both these complexes was carried out with REFMAC5 and ARP/wARP. Clear density for dTDP-D-xylose was seen before its inclusion in the model (Figure 3). Full statistics are given in Table 2 for the three structures discussed. Acknowledgments This work is supported by funding from the Wellcome Trust to J.H.N. C.D. holds an Overseas Research Studentship Award and a University Scholarship. J.H.N. is a BBSRC Career Development Fellow and A.A. was a Wellcome Trust Research Career Development Fellow. We thank David Rice for constructive critique of the manuscript. Received: December 20, 2002 Revised: February 24, 2003 Accepted: March 14, 2003 Published: June 3, 2003 References Allard, S.T., Giraud, M.F., Whitfield, C., Graninger, M., Messner, P., and Naismith, J.H. (2001a). The crystal structure of dTDP-D-glucose 4,6-dehydratase (RmlB) from Salmonella enterica serovar Typhimu-

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