Structure of Leishmania mexicana Phosphomannomutase Highlights Similarities with Human Isoforms

J. Mol. Biol. (2006) 363, 215–227

doi:10.1016/j.jmb.2006.08.023

Structure of Leishmania mexicana Phosphomannomutase Highlights Similarities with Human Isoforms Lukasz Kedzierski 1 †, Robyn L. Malby 2 †, Brian J. Smith 2 † Matthew A. Perugini 3 , Anthony N. Hodder 1 , Thomas Ilg 4 Peter M. Colman 2 and Emanuela Handman 1 ⁎ 1

Infection and Immunity Division, The Walter and Eliza Hall Institute of Medical Research, Melbourne, Australia 2

Structural Biology Division, The Walter and Eliza Hall Institute of Medical Research, Melbourne, Australia 3

Department of Biochemistry and Molecular Biology, The University of Melbourne, Australia 4 Intervet Innovation GmBH, Schwabenheim, Germany

Phosphomannomutase (PMM) catalyses the conversion of mannose-6phosphate to mannose-1-phosphate, an essential step in mannose activation and the biosynthesis of glycoconjugates in all eukaryotes. Deletion of PMM from Leishmania mexicana results in loss of virulence, suggesting that PMM is a promising drug target for the development of anti-leishmanial inhibitors. We report the crystallization and structure determination to 2.1 Å of L. mexicana PMM alone and in complex with glucose-1,6bisphosphate to 2.9 Å. PMM is a member of the haloacid dehalogenase (HAD) family, but has a novel dimeric structure and a distinct cap domain of unique topology. Although the structure is novel within the HAD family, the leishmanial enzyme shows a high degree of similarity with its human isoforms. We have generated L. major PMM knockouts, which are avirulent. We expressed the human pmm2 gene in the Leishmania PMM knockout, but despite the similarity between Leishmania and human PMM, expression of the human gene did not restore virulence. Similarities in the structure of the parasite enzyme and its human isoforms suggest that the development of parasite-selective inhibitors will not be an easy task. © 2006 Elsevier Ltd. All rights reserved.

*Corresponding author

Keywords: Leishmania; phosphomannomutase; crystal structure; virulence; mannose

Introduction Leishmania are protozoan parasites that shuttle between a sandfly insect vector, where they inhabit the gut as flagellated promastigotes, and mammalian hosts, where they exist as obligatory intracellular amastigotes in macrophages.1 Leishmania cause a spectrum of diseases ranging in severity from selfhealing skin lesions to fatal visceral disease, collectively known as leishmaniases. There are two million

† L.K., R.L.M. and B.J.S. contributed equally to this work. Abbreviations used: PMM, phosphomannomutase; HAD, haloacid dehalogenase; GDP-MP, GDP-mannose pyrophosphorylase; GDP-Man, GDP-mannose; M6P, mannose-6-phosphate; M1P, mannose-1-phosphate; G1,6P, glucose-1,6-bisphosphate. E-mail address of the corresponding author: [email protected]

new cases of leishmaniasis each year, and the burden of disease expressed in disability-adjusted life years (DALYs) is estimated to be over 2.3 million.2 The WHO has classified leishmaniasis as a category 1 disease, i.e. emerging and uncontrolled.3 In the absence of vaccines, chemotherapy is the main means of controlling the disease. Treatment relies on pentavalent antimony, but its use is becoming limited due to the emergence of drug resistance and loss of efficacy.4 Unfortunately, only a few drugs are available as a second-line therapy in case of antimonial failure, but cost and toxicity hamper their use.5 Phosphomannomutase (PMM, EC 5.4.2.8) belongs to a group of phosphotransferases with a conserved phosphorylated motif DxDx(T/V).6 The enzyme catalyzes the conversion of mannose-6-phosphate (M6P) to mannose-1-phosphate (M1P) (Figure 1) and mannose-1,6-bisphosphate (M1,6P) is required as a cofactor for enzymatic activity. PMM needs to be phosphorylated in order to be active.7 In humans, two isoforms have been described, PMM1 and PMM2.8

0022-2836/$ - see front matter © 2006 Elsevier Ltd. All rights reserved.

Crystal Structure of L. mexicana PMM

216

Figure 1. Schematic diagram of the reaction catalyzed by phosphomannomutase.

PMM2 is the dominant enzyme with widespread tissue distribution.9 In yeast, the pmm gene is essential for viability.10 In bacteria, PMM deficient mutants display attenuated virulence11 prompting the search for PMM/PGM inhibitors as potential drugs. Leishmania synthesize a range of mannose-rich glycoconjugates, which are considered virulence factors. A prerequisite for glycoconjugates biosynthesis in Leishmania, as in all eukaryotes, is the conversion of monosaccharides to activated sugar nucleotides. The activation of mannose comprises several enzymatic steps performed by phosphomannose isomerase (PMI), PMM, GDP-mannose pyrophosphorylase (GDP-MP) and dolicholphosphatemannose synthase (DPMS).12 The consecutive action of PMM and GDP-MP transforms M6P to GDPmannose (GDP-Man), which acts as the mannose donor. Deletion of the gene encoding PMM renders L. mexicana avirulent, but still viable in culture13 making PMM an attractive drug target. We report the crystal structure of L. mexicana PMM determined to 2.1 Å resolution. We compare the leishmanial PMM structure to the human PMM1 (hPMM1)14 and hPMM2 (PDB entry 2AMY) structures and provide insights into the mechanism of action of PMM that may lead to the design of structure-based inhibitors. We also describe the generation of PMM knockout parasites (ΔPMM) in Leishmania major, the cause of cutaneous leishmaniasis in the Old World, and ΔPMM parasites expressing human PMM2, which offer an excellent tool for testing the efficacy and specificity of potential inhibitors.

not shown). The protein was subjected to size exclusion chromatography and eluted as a single peak (data not shown) confirming the purity and integrity of the fusion protein observed on SDS– PAGE. Recombinant PMM is enzymatically active The enzymatic activity of recombinant PMM was determined in a colorimetric assay coupled to GDPMP and inorganic pyrophosphatase.15 The activity of recombinant PMM (mean A650 nm = 1.48) was comparable to that observed for GDP-MP (positive control, A650 nm = 1.52) (Figure 2) for an equivalent

Results Expression and purification of recombinant L. mexicana PMM The L. mexicana gene encoding PMM was cloned into the expression vector pPROEX HTb, and was expressed in Escherichia coli as an N-terminal hexaHis tagged protein. The recombinant protein was purified using TALON affinity resin resulting in high yields of a single protein band of 31.1 kDa (data

Figure 2. Enzyme activity assay for recombinant PMM. The activity is expressed as a change in A650 nm for 0.1 μg of enzyme over 30 min. Negative control, no PMM/GDP-MP added to reaction; positive control, GDP-MP activity assay; GDP-MP (M6P), GDP-MP alone (no PMM) with mannose-6-phosphate as substrate; PMM (M1P)/(M6P), PMM alone (no GDP-MP) with either mannose-1-phosphate or mannose-6-phosphate as substrate; no G1,6P, cofactor glucose-1,6-bisphosphate not included in the reaction; PMM + GDP-MP (M6P); assay with both GDP-MP and PMM present.

Crystal Structure of L. mexicana PMM

amount of protein (0.1 μg). In order to test for false positive results we checked if the GDP-MP and PMM could utilize M6P and M1P, respectively, as substrate in a single enzyme reaction. Both assays yielded negative results, A650 nm = 0.28 and 0.26, respectively, similar to the negative control A650 nm = 0.25. The activity of PMM could not be detected when GDP-MP was excluded from the reaction and M6P was used as a substrate (A650 nm = 0.35). These observations indicated that the increase in inorganic phosphate observed in the test reaction consisting of M6P as substrate and both PMM and GDP-MP, was due to the two enzymes acting consecutively. The assay also demonstrated that glucose-1,6-bisphosphate (G1,6P) is essential for PMM activity, since the omission of this cofactor from the reaction resulted in lack of enzymatic activity (A650 nm = 0.23). Recombinant PMM is a stable dimer in solution To evaluate the quaternary structure of PMM in aqueous solution, sedimentation velocity studies were performed in the analytical ultracentrifuge. Results indicated that PMM is primarily a single species with the apparent molar mass estimated to be 54.4 kDa (data not shown). This is in agreement with the theoretical mass of a PMM dimer and indicates that PMM exists as a stable homodimer in aqueous solution. Similar data were obtained when the PMM recombinant product was characterized under both non-reducing and reducing conditions (data not shown). This demonstrated that the PMM dimer associates via non-covalent interactions rather than inter-molecular disulphide bonds. PMM has a two domain structure The analysis of the L. mexicana PMM structure revealed that PMM is a member of the haloacid dehalogenase (HAD) family.16 The protein folds as two domains (Figure 3(a)), a classical HAD (core) domain comprising residues 4–84 and 187–245, and an inserted (cap) domain comprising residues 85– 186. The crystallographic asymmetric unit contains three polypeptides, one in a crystallographic dimer, referred to here as the A-dimer, and two others in a non-crystallographic dimer, the BC-dimer. The relative positions of the core and cap domains differ in the two dimers, the A-dimer being open, and the BC-dimer being closed. The orientations of the core domains differ by a 23° rotation (about the hinge region connecting domains) in the open and closed forms (Figure 3(a)). The cap domain is located after the second of three sequence motifs that have been identified in HAD proteins, defining L. mexicana PMM as a type II HAD protein.17 The core domain structure contains all the expected elements of the sequence motifs referred to above (Figure 3(b)). In the first, there are D10 and D12 of which D10 is expected to form the phosphoprotein intermediate. In the second, S46 is expected to form hydrogen bonds to the phosphoryl group,

217 and in the third, K188 and residues in and around the loop 206–215 should coordinate a magnesium ion and assist in binding to the phosphoryl group. In the L. mexicana PMM structure, a magnesium ion is observed coordinated to residues D10 and D207, with D215 (salt linked to K188) close by. The heart of the cap domain is a four-stranded antiparallel βsheet elaborated on one face with three α-helices. The solvent-exposed face of the sheet is oriented towards the active site of the core domain where the conserved sequence motifs are clustered. Predominantly hydrophilic residues project from this surface of the cap domain towards the active site of the core domain (Figure 3(b)), and are likely to be involved in substrate recognition and selectivity. The A-dimer and BC-dimer interfaces are identical to each other, and are formed from amino acid residues exclusively within the cap domain. The effect of dimer formation is to extend the solventexposed face of the β-sheet thus placing the two active sites adjacent to each other (Figure 4(a)). Residues D10 across the dimer interface are 38 Å apart, but residues E120 on the first strand of the cap domain are within 8 Å from each other (Figure 4(b)). A chloride ion, located on the molecular dyad axis, is coordinated by four peptide NH groups, those of F118 and V119 from dyad symmetry related molecules. The shape complementarity (Sc) equals 0.68 over a buried surface area of 700 Å2 on each monomer,18 which is typical of a protein–protein interface. The core domains of the BC-dimer in which the polypeptides are in the closed form, come close to each other in the vicinity of residue E57 (Figure 4(b)). The closest structural homologues of the Leishmania PMM in the Protein Data Bank are human PMM114 (PDB entry 2FUC) and PMM2 (currently unpublished; PDB entry 2AMY). A search using DALI19 for homologues of the cap domain identified human PMM2 (Z-score 19.9, rmsd of 0.7 Å over 101 Cα atoms) followed by a cluster of structures with similar Z-scores, the best fit being trehalose 6phosphate phosphatase related protein (PDB entry 1U02, Z-score 5.8, rmsd of 3.0 Å over 72 atoms), and including sucrose phosphatase (PDB entry 1U2S, Z-score 4.9, rmsd of 2.6 Å over 64 atoms). Structural alignment (LSQKAB)20 of the cap domain dimer with its counterpart in human PMM2 yielded an rmsd of 0.9 Å for 204 paired Cα atoms, confirming that the dimer interface is conserved between human PMM2 and L. mexicana PMM. A similar analysis of the entire polypeptides indicates that the relative conformation of core and cap domains in both human PMM1 and PMM2 crystals is very similar to that seen in the A-polypeptide of L. mexicana PMM, i.e. the open form. The cap domain in leishmanial and both human PMMs possesses a novel topology. The closest structural homologues, 1UO2 and 1U2S, have a similar topology, but with only two α-helices on the face of the β-sheet. Crystals that were nearly isomorphous with native crystals grew in the presence of 5 mM G1,6P, and electron density maps showed the ligand

218

Crystal Structure of L. mexicana PMM

Figure 3. PMM structure. (a) Comparison of the open (A polypeptide, magenta) and closed (B polypeptide, yellow) forms in the asymmetric unit of the crystal. (b) View into the active site. Residues that belong to sequence motifs I (D10, D12), II (S46) and III (K188, D207, D215) are coloured red, yellow and blue, respectively, with the magnesium ion in green. Residues from the cap domain, E120, R122, M125, N127, R133, R140, S172, G174, S178, D180 and F182, are presented in magenta.

Crystal Structure of L. mexicana PMM

219

Figure 4 (legend on next page)

220

Crystal Structure of L. mexicana PMM

bound only to the BC (closed) polypeptides. One phosphate group was associated with HAD domain residues including D10, K188, and N214, and the other with cap domain residues including R133, R140 and S178. Coordination of the glucose moiety is mostly via residues from the cap domain, including R122. At 2.9 Å resolution we cannot unambiguously determine the orientation of the sugar, and hence cannot assign which phosphate is associated with which domain. We favour an orientation with the 6-phosphate interacting with the HAD domain only because that places the hydrophobic face of the sugar against glycyl residues 174 and 175 (Figure 4(c)). A similar arrangement is observed in the high-resolution Xray structure of the complex of β-phosphoglucomutase with substrate, where the hydrophobic face of the carbohydrate abuts the highly conserved glycyl residue 46.21 The location of the magnesium ion is unaltered as a result of the binding of G1,6P. The closest approach of the G1,6P to its counterpart around the molecular dyad is 21 Å. Targeted gene replacement and complementation of PMM in L. major The original studies13 focused on the PMM from L. mexicana, the cause of cutaneous leishmaniasis in South America. Since the parasite biology, the disease pattern and its immunological correlates produced by L. major infection are different22 we set out to generate L. major PMM deletion mutants. After two rounds of targeted gene replacement several clones were isolated that lacked both alleles of the PMM open reading frame. The absence of the pmm gene and loss of its product was confirmed by PCR, Southern blot analysis, and Western blotting of cell lysates (Figure 5(a)–(c)). The pmm gene was not detectable by PCR in ΔPMM parasites, whereas the control gdp-mp gene was amplified from all genomic DNA samples. As a further control we performed PCR with ble specific primers, which detected the drug cassette in heterozygous and homozygous ΔPMM lines, but not in the wild-type (WT) parasites. The Southern blot analysis with a pmm and a ble probes (data not shown) confirmed the deletion of the pmm gene. Single bands of 2225 bp in NdeI and 4396 bp in EcoRI/EcoRV genomic digests were detected in WT and heterozygous parasites, but were absent in ΔPMM genomic DNA. Any consideration of PMM as a drug target for leishmaniasis has to take into account the similarity between the parasite and the human enzymes. Therefore, we have generated ΔPMM parasites

Figure 5. Analysis of L. major ΔPMM knockout parasites. (a) PCR analysis of wild-type (WT), heterozygote (PMM+/−) and knockout parasites (ΔPMM) with primers specific for the pmm, gdp-mp and BLE genes. (b) Southern blot analysis of genomic DNA from WT, PMM+/− and ΔPMM parasites probed with a DIGlabelled pmm probe. DNA size standards (in kb) are shown on the left of the panels. (c) Western blot analysis of WT and ΔPMM promastigote lysates; the membrane was probed with anti-PMM antibodies followed by antiGDP-MP antibodies without stripping; molecular mass standards (in kDa) are shown on the left of the panel.

expressing the human PMM2 to determine whether it can complement the activity of the parasite enzyme. The availability of this line should also allow us to test the selectivity of putative inhibitors in

Figure 4. L. mexicana dimer structure and complex with cofactor. (a) The dimer interface is contributed exclusively by the cap domains shown here, and results in an extension of the open face of the β-sheet and juxta-positioning of the two active sites. A chloride ion (yellow sphere) lies on the 2-fold axis. (b) Ribbon drawing of the BC (closed) dimer illustrating the close approach of core domain residues at residue E57, near the dyad axis. The dyad axis is vertical in the plane of the Figure, the cap domains below and core domains above. (c) Stereo image of the electron density omit map obtained from the refined complex with ligand removed. Coordinating groups from the core domain are the magnesium ion (green), D10, K188 and N214, and from the cap domain R122, S178, R133 and R140. The orientation of the cofactor is uncertain; the orientation shown places the carbohydrate hydrophobic face near glycine residues G174 and G175.

Crystal Structure of L. mexicana PMM

221

vivo. The ΔPMM mutants were complemented with hPMM2 expressed episomally on pX vector transfected into the knockout cell line. The complemented cell line was tested for the presence of hPMM2 by immunoblotting (Figure 6(a) and (b)).

Figure 7. Growth kinetics and virulence of ΔPMM and hPMM2 add-backs. (a) Growth curves of L. major WT, ΔPMM and ΔPMM + hPMM2 were compared under the same culture conditions. (b) In vitro infection of bone marrow-derived macrophages with L. major WT, ΔPMM and ΔPMM + hPMM2 parasites. The percent infected macrophages at 3 h and 48 h post-infection was calculated by counting duplicates of three independent samples. Error bars indicate SEM.

Figure 6. Human PMM2 expression in ΔPMM parasites. Promastigote lysates of WT, ΔPMM and human PMM2 addbacks (ΔPMM + hPMM2) and the human HeLa cell line lysate were immunoblotted with anti-hPMM2 antibodies; recombinant hPMM2 was included as a control (a); the same membrane was probed with antiPMM antibodies (b). Complementation of ΔPMM with hPMM2 was assessed by immunoblotting with antibodies to leishmanial glycoconjugates: (c) rabbit anti-PPG antibodies, (d) monoclonal 11E5 anti-PSA-2, (e) monoclonal WIC108.3 anti-LPG Gal(β1-4)Man backbone repeats, (f) monoclonal WIC79.3 specific for the galactose side-chains of L. major LPG. Molecular mass standards (in kDa) are shown on the left of each panel.

We examined the ability of the hPMM2 to restore the expression of some Leishmania glycoconjugates (Figure 6(c) and (d)). As expected, in the ΔPMM mutants there was a general down-regulation of biosynthesis of glycoconjugates such as LPG, PPG and PSA-2. The expression of the human enzyme in the mutants restored expression of these glycoconjugates to levels comparable with the wild-type as indicated by immunoblotting experiments. The growth kinetics of the ΔPMM and the hPMM2 transfected lines were followed over nine days. Under the normal culture conditions (26 °C, pH 7.5) ΔPMM parasites showed slower growth compared to WT parasites. The parasites complemented with either the L. major PMM (data not shown) or human PMM2 also showed slower growth in vitro (Figure 7(a)).

222 We then tested the L. major ΔPMM and hPMM2 add-back for their ability to infect macrophages and mice to determine whether the loss of PMM led to loss of virulence as described for the L. mexicana PMM.13 As expected the ΔPMM were avirulent, but interestingly, the hPMM2 transfected parasites were also avirulent as shown by the lack of lesion development and the inability to isolate parasites from infected mice (data not shown). The L. major ΔPMM parasites were also not able to survive in bone marrow derived macrophages (Figure 7(b)) or peritoneal macrophages (data not shown). Initially, mutant parasites were taken up at the same rate as WT parasites, but after 48 h no parasites could be detected. Although better than the PMM mutants, the ability to survive in macrophages was reduced in the hPMM2-complemented parasites compared to WT (Figure 7(b)).

Discussion Leishmania synthesize a range of mannose-rich glycoconjugates, and there is strong evidence that they are required for parasite virulence and survival.23 Activated mannose is essential for the biosynthesis of these glycoconjugates, and its biosynthesis requires action of four enzymes of which PMM plays a central role. Due to the importance of mannose-containing glycoconjugates to the parasite survival in the mammalian host, enzymes involved in the mannose activation pathway constitute attractive targets for anti-leishmanial drug development. In this study, the L. mexicana PMM was cloned and expressed as a recombinant protein, and we characterized it as a potential target for anti-Leishmania drug development. We have concentrated our initial efforts on the L. mexicana enzyme. However, the disease pattern and immune responses of mice infected with L. mexicana and L. major is different22 and their virulence factors differ as well.24 Therefore, we have generated PMM deletion mutants in L. major. L. mexicana and L. major PMM share 93.4% and 92.3% identity at protein and DNA level, respectively. The differences are located on the surface of the molecule and are unlikely to affect enzyme activity. We have shown that the recombinant PMM has enzymatic activity in a sensitive and simple colorimetric assay. The method is easy to perform and can be used in automated high-throughput screening analyses for the search of inhibitors of the enzyme. Our assay also confirmed that the presence of a bisphosphorylated sugar cofactor is essential for the enzymatic activity of PMM, and that the addition of glucose mono-phosphates neither inhibits the reaction nor can they serve as phosphate donors for PMM (data not shown). In preparation for the development of drugs targeting PMM, we sought to obtain the crystal structure of the enzyme. The structural analysis showed that L. mexicana PMM is a member of the haloacid dehalogenase (HAD) family of proteins.16

Crystal Structure of L. mexicana PMM

Members of the HAD family share a classical HAD domain and a cap domain with a variable function and fold. The leishmanial protein also folds as two domains, a classical core domain and a cap domain. The relative positions of the core and cap domains differ in the two dimers, the A-dimer being open and the BC-dimer being closed. The cap domain is located after the second of three sequence motifs that have been identified in HAD proteins, characterizing L. mexicana PMM as a type II HAD protein.17 Although the HAD family members do not usually share sequence similarities, they do share three conserved motifs. These include motif I, DX(D/T/Y)X(T/V)(L/V/I) that is transiently phosphorylated in HAD phosphatases;6 motif II, consisting of serine or threonine residues in a hydrophobic context,25 and motif III, that includes a conserved lysine residue that stabilizes the phosphorylated state of an intermediate in an active site. 25,26 All the motifs are present in L. mexicana PMM and are arranged adjacent to each other in the folded molecule and flank the active site. The structure of sucrose phosphatase from a cyanobacterium,27 the tenth hit in the DALI search for cap domain homologues described above, has been determined in complex both with glucose and with sucrose-6-phosphate. The phosphate moiety in the sucrose-6-phosphate complex is coordinated by core domain residues G42 (G45 in L. mexicana PMM), K163 (K188), N189 (N214), T41 (G44) and D11 (D12) illustrating the conservation of structures within the HAD core domains. In contrast, the cap domains have widely divergent sequences, reflective of different substrate specificities.17 The dimeric structure of L. mexicana PMM and of hPMM1 and hPMM2 appears novel among HAD family proteins, consistent with the novelty of the cap domain sequence and topology. It is not known whether catalysis requires the dimer, but the adjacent location of the active sites of the two dyadrelated subunits is suggestive of some communication between them. We have performed a MALDI mass spectrometry analysis of the leishmanial recombinant enzyme incubated for various times with a bis-phosphorylated cofactor in an attempt to detect the phosphorylated state of the enzyme. No shifts of peak were detected, indicating that it is likely that the enzyme is only transiently phosphorylated during the catalytic reaction (data not shown). The structure of PMM is consistent with a mechanism of catalysis that has been proposed for other phosphoryl transfer enzymes that involve a phosphorylated enzyme intermediate.28 The formation of this intermediate is facilitated by a 1,6-bisphosphocarbohydrate initiator. Thus, the first step in the reaction is the activation of the enzyme by transfer of a phosphate from the initiator to the enzyme aspartyl nucleophile, producing an aspartylphosphate, D10 in L. mexicana PMM. Following activation, substrate M6P binds, and a covalent bond is formed between the substrate C1 hydroxyl oxygen atom and the aspartylphosphate phosphorus atom. Formation of

Crystal Structure of L. mexicana PMM

the covalent bond with phosphorus produces a bisphospho-protein intermediate. This intermediate has been identified recently in the X-ray crystal structure of the functionally related enzyme βPGM.21 Subsequently, the bisphospho-carbohydrate is released from the enzyme, reorientates itself and transfers the 6-phosphate to the aspartyl nucleophile, replenishing the activated form of the enzyme, and forming the final product, M1P. Release of the phosphate from the bisphospho-carbohydrate to the aspartyl is facilitated by another acid/base group, D12, in the active site. Reorientation of the free bisphospho-carbohydrate is likely to require release from, and rebinding to, the enzyme active site.29 Unlike other phosphomutases, PMM from L. mexicana and both human forms are dimers. The dimeric nature of the enzyme, structurally conserved across different species, suggests functional importance. The mannose-1,6-bisphosphate intermediate when released from one monomer can either rebind to the same monomer or migrate and bind to the alternate monomer (but only if the alternate monomer is in the unphosphorylated state). The transfer of the bisphospho intermediate, synthesized in situ, from one active site on the dimer to the other ensures an efficient turnover, and also removes any requirement for reorientation of the substrate within the active site, a mechanism proposed for the structurally unrelated bacterial PMM/PGM.30 The crystal structure of the related sucrose-phosphatase reveals it is not dimeric, 27 but the proposed catalytic mechanism is a single-step process involving the substrate, and does not rely on multiple binding events of any intermediates. Whether a dimer is necessary for PMM activity in L. mexicana or not, the close proximity of the active sites suggests possible strategies for drug discovery based on linked groups able to simultaneously engage the two sites, in a manner akin to the recently developed influenza virus neuraminidase inhibitor dimers.31 In functionally related enzymes the second of the aspartate residues in the conserved DxDx(T/V) motif I, D12 in L. mexicana PMM, is responsible for release of the phosphate from the bisphosphocarbohydrate intermediate to the nucleophilic aspartyl group. In the X-ray crystal structures of other HAD family enzymes this acid binds the phosphate.32–34 However, in the structures of PMM, both in the presence and absence of ligand, this acid is sequestered away from the substrate phosphate through hydrogen-bonding interactions with the side-chain of a conserved glutamine (Q54) and the backbone amide nitrogen atom of R19, also conserved amongst PMM. Rotation about the Cα–Cβ bond by ∼180°, however, brings the side-chain of the acid into position to bind the phosphate. Similarities in the structure of the parasite and human PMM suggest that the development of parasite-selective inhibitors will not be trivial. Amino acid sequences adjacent to and embracing the three motifs in the HAD domain are similar in L. mexicana PMM and in both human isozymes PMM1 and PMM2, offering little scope for targeting. The

223 amino acid sequence identity ranges from 46% to 54.4% between leishmanial PMM and PMM1 and PMM2, respectively. Comparison of the structures of hPMM1, hPMM2, and PMM from L. mexicana also reveals a high level of similarity, especially at the domain interfaces where the substrate binds. In the L. mexicana PMM core domain, D207, which coordinates the magnesium ion and lies within 4 Å of the catalytic nucleophile D10, is replaced by an asparagine in hPMM1. Although in the structure of hPMM2 this region is poorly resolved, the sequence alignment with L. mexicana PMM indicates that the homologue of D207 is also an aspartate in hPMM2. Thus, there is little to differentiate between parasite and human enzymes in this region. Within the cap domain, all the residues on the open face of the β-sheet that face towards the active site on the HAD domain are conserved between parasite and human PMM1 and PMM2. The most significant structural variation between PMMs is found in the residues immediately following the third beta strand (G174, G175 in L. mexicana PMM), residues that abut S187. In hPMM1 the hydroxyl side-chain of this serine (S188) forms a hydrogen bond with a backbone carbonyl of this loop region. The structure of this loop differs slightly in the ligand-free and glucose1,6-bisphosphate-bound forms of leishmanial PMM, suggestive of an inherent flexibility in this region. The similarities between Leishmania and human PMMs, although somewhat discouraging, should not curb the search for selective inhibitors against the parasite enzyme or the mannose activation pathway in general. There are precedents where inhibitors can distinguish between virtually identical structures. The COX2 inhibitors are very selective in their mode of action despite the fact that the catalytic sites of COX1 and COX2 differ only by a single amino acid.35 Selective inhibitors recognizing a single amino acid change in the secondary binding site of protein tyrosine phosphatase 1B and T-cell protein tyrosine phosphatase have also been recently identified.36,37 The kinetic factors in the turnover rates of human and parasite enzymes may also play a significant role in drug selectivity. The human enzyme may be regenerated faster than its leishmanial counterpart minimising negative effects of anti-leishmanial therapy. Mutations in human PMM2 are associated with congenital disorder of glycosylation type Ia (CDGIa),38 a disorder that is lethal in 20% of subjects in the first year of life. Although the mutation data suggest a critical role for PMM activity during development, it is not known whether temporary inhibition of the enzyme, for example during treatment for leishmaniasis, would have adverse effects. It is worth pointing out that the majority of CDGIa patients display residual PMM activity of up to 10% of normal levels, but the heterozygous parents with approximately 50% of normal activity are healthy.39 Given these facts, it is likely that inhibition of PMM by an anti-leishmanial drug in individuals with normal PMM activity may be well tolerated. We have generated ΔPMM parasites expressing the human PMM2 that will allow testing the

Crystal Structure of L. mexicana PMM

224 selectivity of potential inhibitors. The hPMM2 complemented parasites synthesised glycoconjugates, but showed reduced virulence compared to the wild-type. In culture, these parasites also had slower growth kinetics, similar to the ΔPMM parasites. The slower growth rate was puzzling in view of the fact that the main glycoconjugate synthesis appeared to be restored, thus indicating that the effect of gene deletion had been complemented. These observations suggest that despite performing the same role and having a very similar structure, the human enzyme may be different from the leishmanial enzyme. Alternatively, PMM may perform as yet unknown functions in Leishmania, which cannot be complemented by the human enzyme. Despite the similarities between the human and parasite enzymes, the availability of the enzyme structure, the enzyme activity assay suitable for high throughput screening and specific parasite knockout and add-back lines, should facilitate the drug discovery process.

Materials and Methods Parasite and bacterial culture L. major MHOM/IL/80/Friedlin were maintained at 26 °C in M199 medium supplemented with 10% (v/v) heat inactivated fetal bovine serum (Trace Biosciences). DNA

manipulations were performed in E. coli strain DH5α and protein expression in the BL21 (DE3)pLysS. All bacterial strains were cultured in LB medium or in superbroth for protein expression with the addition of ampicillin (100 μg/ml). Subcloning of the pmm gene The L. mexicana pmm gene (AJ308232) was amplified from a genomic template by PCR using primers p240 (CCGGAATTCATGGGCTCCAAGGCTATTCTTC) and p241 (AAAACTGCAGTTACCGCGAATCCTCGAGAAG) under the conditions described.15 The restriction sites were engineered for cloning with a hexa-His tag into the expression vector pPROEX HTb (Invitrogen). Protein purification and crystallization Purification of the fusion protein was performed by affinity chromatography using TALON Metal Affinity Resin (BD Biosciences). The purity and integrity of the protein was assessed on SDS–PAGE gels stained with Coomassie blue. Recombinant PMM was dialysed against 10 mM Tris–HCl (pH 7.5), 1 mM DTT and concentrated using Vivaspin 2 concentrators (Vivascience AG). The PMM solution at 10 mg/ml was used to set up sitting drop vapour diffusion trays. Diffracting crystals were obtained by microseeding and grown in 14% (w/v) PEG3350, 10% PEG400, 0.1 M citrate buffer (pH5.5). Following crystal formation, 0.2 ml of 50% PEG3350 was added to the reservoir and crystals were equilibrated for five to seven days. Crystals were cryo-protected with the

Table 1. Crystallographic data collection and refinement

A. Data collection Space group Unit cell dimensions (Å) Resolution limit (Å) Observations Unique reflections Completeness (%) /c Rmerged B. Refinement Rworke Rfreef Wilson B factor (Å2) No. protein atoms (average B factor; Å2) No. solvent atoms (average B factor; Å2) No. ligand atoms (average B factor; Å2) r.m.s. deviation from ideal geometry Bond lengths (Å) Bond angles (°) C. Ramachandran plot Most favoured (%) Generously allowed (%)

Native

SeMeta peak (0.9794 Å)

P3121 a = b = 92.4 c = 173.3 2.1 243,915 50,650 99.7 (100.0) 12.1 (3.0) 0.046 (0.518)

P3121 a = b = 92.5 c = 174.4 2.5 623,148 30,444 100.0 (100.0) 41.5 (7.2) 0.080 (0.344)

SeMet edge (0.9796 Å)

2.5 306,856 30,690 99.7 (99.0) 36.1 (2.5) 0.077 (0.492)

SeMet remote (0.9840 Å)

G1,6Pb

2.5 283,288 30,308 98.2 (82.9) 31.5 (1.2) 0.075 (0.615)

P3121 a = b = 92.3 c = 172.8 2.9 64,339 17,084 90.6 (84.5) 16.1 (4.4) 0.065 (0.311)

0.190 (0.243) 0.230 (0.286) 45.5 5854(49.5) 350 (50.6) – 0.015

0.192 (0.270) 0.268 (0.391) 80.0 5841(62.9) 25 (60.6) 40 (58.6) 0.011

1.5

1.4

90.7 0.3

87.4 0.3

Values in parentheses refer to data in the highest resolution shell. a SeMet, selenomethionine. b G1,6P, glucose-1,6-bisphosphate. c / average signal to noise ratio for merged reflection intensities. d Rmerge = ∑ |Ii(h) –| / ∑ Ii(h), where Ii(h) and are the ith and mean measurements of reflection h. e R = ∑ ||Fo – k |Fc|| / ∑ |Fo|, where Fo and Fc are observed and calculated structure factor amplitudes of reflection h. f Rfree, test reflection (5% of data), selected randomly for cross-validation during crystallographic refinement.

Crystal Structure of L. mexicana PMM equilibrated reservoir and flash-cooled in liquid nitrogen for data collection. To facilitate the structure solution, a selenomethionine (SeMet)-substituted PMM was produced in E. coli by growing bacteria in minimal, auto-inducing medium PASM-5052 in the presence of SeMet (Sigma Chemical Co) and 17 amino acids (Sigma Chemical Co) as reported.40 The SeMet recombinant protein was purified according to the standard protocol. Incorporation of SeMet was confirmed by MALDI mass spectrometry that showed heterogeneity in the peak, presumably due to variable SeMet incorporation (data not shown). Native PMM crystals were used to seed into SeMet PMM drops containing 10 mg/ml protein. Initial SeMet crystals were subsequently used to obtain more SeMet crystals. Structure determination and refinement X-ray diffraction data (Table 1) were collected at the APS on beamlines 22-ID for the SeMet protein (SER-CAT) and 14-ID-B for the native protein (BioCARS), as well as at the SLS on a beamline XØ6SA, and processed using HKL200041 and the CCP4 program suite.20 Six Se sites were found by MAD phasing42 using HKL2MAP,43 but the resulting phases did not yield an interpretable electron density map. Heavy-atom refinement and phasing using SHARP44 and density modification with SOLOMON45 resulted in a traceable electron density map (ARP46 ) with evidence of three molecules in the asymmetric unit.47–49 Model building in O50 and refinement using REFMAC51 applying a TLS refinement protocol led to the structure described in Table 1. No residues are in disallowed regions of the Ramachandran analysis, and only two residues (R115 in A and C chains) are located in generously allowed regions as defined by PROCHECK.52 The refined native structure was used as the basis for initiating refinement of the G1,6P complex. The orientation reported here is the more stereochemically acceptable solution for the pyranose ring and positions the 6-P adjacent to the active site serine residue 46. PMM activity assay Enzymatic activity of recombinant PMM was determined using a modified colorimetric assay coupled with GDP-MP and inorganic pyrophosphatase.15 Through the activity of PMM, M6P is converted to M1P, which then acts as a substrate for GDP-MP. In addition, 10 μM G1,6P was added, which is required as a cofactor for PMM activity.7 The assay was carried out at 30 °C for 30 min in 100 μl reactions. Generation of gene knockout and add-back parasites The gene replacement strategy was performed as described.13 The 5′ and 3′ untranslated regions (UTR) of PMM were amplified by PCR from L. major Friedlin genomic DNA using primers p270 (AATGCGGCCGCAAAGACCAAATGGCTGGAGTG) and p271 (AGTACTAGTTTTGCTTTGTTGTGCTTTCG) for the 5′-UTR and p273 (ATCGATATCCAGATCCGTTAAAGAGCTGCG) and p274 (AGTACTAGTGGATCCATCCCTATCATCACCTGCG) for the 3′-UTR. Open reading frames (ORF) encoding phleomycin binding protein (BLE)24 and puromycin N-acetyltransferase (PAC)53 were used. Both UTRs and drug resistance genes were cloned into a pBSK plasmid and gene replacement cassettes were excised by

225 NotI/EcoRV digest and transfected into L. major Friedlin promastigotes as described.54 For the episomal gene addback, the human pmm2 ORF was amplified from clone 3611382, NIH Mammalian Gene Collection (Invitrogen), using primers p289 (CGCGGATCCATGGACGCGCCTGGCCCAGCG) and p291 (AAGGAAAAAAGCGGCCGCTCAGGAGAACAGCAGTTCAC) and cloned into the pX vector.55 The construct was transfected into L. major Friedlin ΔPMM promastigotes and transfectants were selected by growth in medium containing 100 μg/ml of G418 (Invitrogen). Southern blotting Purified L. major genomic DNA (2 μg)56 was subjected to overnight digestion with either NdeI or EcoRI/EcoRV restriction endonucleases. Digested DNA was transferred to nylon membranes (Amersham) and probed with DIGlabelled pmm and ble probes according to the manufacturer's instructions (Roche). In vitro infections of macrophages In vitro infection of bone marrow derived macrophages was performed as described.57 Results were plotted as a percentage of cells infected at different times after addition of parasites. Protein Data Bank accession codes The coordinates of the L. mexicana PMM crystal structure and the complex with cofactor have been deposited with RCSB, accession codes 2I54 and 2I55, respectively.

Acknowledgements We thank Michael Gorman, Trevor Huyton and Peter Czabotar, and staff at APS, Chicago, USA and SLS, Villigen, Switzerland for assistance with synchrotron data collection. This work was funded by the National Health and Medical Research Council, NHMRC/ARC Network for Parasitology and the Australian Synchrotron Research Program.

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Edited by M. Guss (Received 26 June 2006; received in revised form 4 August 2006; accepted 10 August 2006) Available online 12 August 2006