Kinetic Characterization and Phosphoregulation of the Francisella tularensis 1-Deoxy-D-Xylulose 5-Phosphate Reductoisomerase (MEP Synthase)

Deliberate and natural outbreaks of infectious disease underscore the necessity of effective vaccines and antimicrobial/antiviral therapeutics. The prevalence of antibiotic resistant strains and the ease by which antibiotic resistant bacteria can be intentionally engineered further highlights the need for continued development of novel antibiotics against new bacterial targets. Isoprenes are a class of molecules fundamentally involved in a variety of crucial biological functions. Mammalian cells utilize the mevalonic acid pathway for isoprene biosynthesis, whereas many bacteria utilize the methylerythritol phosphate (MEP) pathway, making the latter an attractive target for antibiotic development. In this report we describe the cloning and characterization of Francisella tularensis MEP synthase, a MEP pathway enzyme and potential target for antibiotic development. In vitro growth-inhibition assays using fosmidomycin, an inhibitor of MEP synthase, illustrates the effectiveness of MEP pathway inhibition with F. tularensis. To facilitate drug development, F. tularensis MEP synthase was cloned, expressed, purified, and characterized. Enzyme assays produced apparent kinetic constants (KMDXP = 104 µM, KMNADPH = 13 µM, kcatDXP = 2 s−1, kcatNADPH = 1.3 s−1), an IC50 for fosmidomycin of 247 nM, and a Ki for fosmidomycin of 99 nM. The enzyme exhibits a preference for Mg+2 as a divalent cation. Titanium dioxide chromatography-tandem mass spectrometry identified Ser177 as a site of phosphorylation. S177D and S177E site-directed mutants are inactive, suggesting a mechanism for post-translational control of metabolic flux through the F. tularensis MEP pathway. Overall, our study suggests that MEP synthase is an excellent target for the development of novel antibiotics against F. tularensis.


Introduction
The US Centers for Disease Control and Prevention (CDC) classify biothreat agents based upon their ease of dissemination, associated morbidity/mortality rates, projected social impact, and emergency response procedures. Category A agents (i.e. those of highest concern) include Bacillus anthracis (the causative agent of anthrax), Yersinia pestis (plague), and Francisella tularensis (tularemia), while category B agents (exhibiting lower morbidity/ mortality rates) include Brucella species (brucellosis), Burkholderia mallei (glanders), and Burkholderia pseudomallei (melioidosis). The 1984 Rajneeshee Salmonella attack, 2001 anthrax letter attacks, 2003 SARS outbreak, and 2009 H1N1 swine flu pandemic illustrate our vulnerability to both deliberate and natural outbreaks of infectious disease and underscore the necessity of effective vaccines and antimicrobial/antiviral therapeutics. The prevalence of antibiotic resistant strains and the ease by which antibiotic resistance can be engineered into bacteria further highlights the need for continued development of novel antibiotics against new bacterial targets.
Isoprenoids are a class of molecules fundamentally involved in a variety of crucial biological functions including electron transport (quinones), cell wall biosynthesis (dolichols), signal transduction (prenylated proteins), and the regulation of membrane fluidity (hopanoids and cholesterol). Despite their structural and functional diversity, all isoprenoids are derived from two building blocks, isopentenyl diphosphate (IPP) and dimethylallyl diphosphate (DMAPP), which originate from either the mevalonic acid (MVA) or methylerythritol phosphate (MEP) pathway (Fig. 1). Because mammalian cells exclusively utilize the MVA pathway, enzymes within the MEP pathway make an attractive target for the development of novel antimicrobials (reviewed in [1][2][3]).
Genome sequences reveal that Francisella, Brucella, Bacillus, Burkholderia, and Yersinia each harbor MEP pathway genes, but little else is known about isoprene biosynthesis in these biothreat agents. In this report we describe the characterization of F. tularensis MEP synthase, a MEP pathway enzyme and potential target for drug development. The F. tularensis MEP synthase gene was cloned, expressed in Escherichia coli, and the recombinant protein was purified and enzymatically characterized. Posttranslational modification (phosphorylation) was revealed by protein mass spectrometry and correlated with the loss of enzyme activity, suggesting a possible regulatory mechanism. The stability of F. tularensis MEP synthase in high-throughput type assays amenable to drug development was also evaluated. Overall, our results suggest that MEP synthase is an excellent target for the development of novel antibiotics against F. tularensis.

Construction of the F. tularensis MEP synthase expression plasmid
The MEP synthase coding region (ispC) was identified in the F. tularensis subsp. holarctica LVS genome (accession number NC_007880) via a BLAST search using the E. coli K12 homologous sequence as the query. Polymerase chain reaction (PCR) primer pairs, designed to flank ispC (FtIspC-f; 59-CACCATGTTTAAAAAAACTAAGATTAC -39 and FtIspC-r; 59-CCCCAAAACAGAATCCACATATTC -39), were purchased from Sigma-Genosys (The Woodlands, TX) and used to amplify the gene from F. tularensis subsp. holarctica LVS genomic DNA. FtIspC-f contains four additional 59 residues (CACC) to facilitate the unidirectional insertion of the PCR product into plasmid pET101/D-TOPO (Invitrogen). FtIspC-r is designed to eliminate the stop codon in the PCR product to permit the expression of a C-terminal His-tagged MEP synthase protein. PCR was performed with Platinum Pfx polymerase (Invitrogen) and the following parameters: 2 minutes at 94uC followed by 22 cycles of 15 sec at 94uC, 30 sec at 54uC, 1.5 min at 68uC, and a final elongation of 10 min at 68uC. The PCR product was purified using the Qiaquick PCR Purification Kit (Qiagen, Valencia, CA) and cloned into pET101/D-TOPO to create pFtIspC. Restriction mapping and DNA sequencing were used to confirm the fidelity of the PCR and the correct construction of the plasmid. pFtIspC was transformed into chemically competent E. coli BL21 CodonPlus (DE3)-RIL cells to express the protein.

Expression and purification of F. tularensis MEP synthase
A one liter shake flask was used for protein expression (37uC, 250 rpm). The flask was inoculated with a 10 mL overnight culture of E. coli BL21 CodonPlus (DE3)-RIL containing pFtIspC and upon reaching an OD 600 of 1.1, protein expression was induced by the addition of 0.5 mM isopropyl b-D-thiogalactopyranoside (IPTG). After incubation for an additional 18 hours, cells were harvested by centrifugation and stored at 280uC. To purify the His-tagged protein, the cell pellet was thawed then cells were lysed using Lysis Buffer A (100 mM Tris pH 8, .032% lysozyme; 3 mL per mg cell pellet), followed by Lysis Buffer B (0.1 M CaCl 2 , 0.1 M MgCl 2 , 0.1 M NaCl, .020% DNase; 0.3 mL per mg cell pellet). Clarified cell lysate was obtained by centrifugation (48,000 x g, 20 min) then passed through a TALON immobilized metal affinity chromatography column (Clontech Laboratories, Mountain View, CA). The column was washed with 15 column volumes of 1X equilibration buffer (50 mM HEPES pH 7.5, 300 mM NaCl), 10 column volumes of 1X wash buffer (50 mM HEPES pH 7.5, 300 mM NaCl, 10 mM imidazole), 10 column volumes of 2X wash buffer (100 mM HEPES pH 7.5, 600 mM NaCl, 20 mM imidazole), and the His-tagged protein was then eluted with 5 column volumes of 1X elution buffer (150 mM imidazole pH 7.0, 300 mM NaCl). Buffer was exchanged by addition of 0.1 M Tris pH 7.5, 1 mM NaCl, 5 mM DTT while concentrating the protein by ultrafiltration. Protein concentration was determined using the Advanced Protein Assay Reagent (Cytoskeleton, Denver, CO) with c-globulins (Sigma-Aldrich) as the standard. The protein was visualized via Coomassie stained SDS-PAGE and a Western blot with an anti-His antibody (Qiagen). The yield of purified MEP synthase averaged 5-10 mg per 1 L LB shake flask.

Size-Exclusion Chromatography
The molecular mass of MEP synthase and the mutant derivatives were estimated by loading 1 mg of protein onto a Sephacryl 200HR (Sigma Aldrich, St. Louis, MO) size-exclusion chromatography column equilibrated with 0.1 M Tris pH 7.5, 1 mM NaCl, 5 mM DTT (flow rate of 2 mL/min) and calibrated with a gel filtration standard kit purchased from Bio-Rad (Hercules, CA). Blue dextran was used to determine the void volume of the column.

Mutagenesis
The Ser 177 to Asp 177 mutant of MEP synthase was created via PCR based site directed mutagenesis using primers FT-IspC-SD177-FP (59 TTA ACA GCA gaT GGA GGT CCT TTT AG 39; lowercase residues indicate site of mutation) and FT-IspC-SD177-RP (59 AGG ACC TCC Atc TGC TGT TAA AAT TAT C 39), whereas the Ser 177 to Glu 177 mutant was created using primer pairs FT-IspC-SE177-FP (59 TTA ACA GCA gaa GGA GGT CCT TTT AGA G 39) and FT-IspC-SE177-RP (59 AGG ACC TCC ttc TGC TGT TAA AAT TAT C 39). Each primer pair is oriented outward from a central overlapping region. In the first step of the mutagenesis, four unidirectional PCRs were performed with each of the four individual PCR primers (FT-IspC-SD177-FP, FT-IspC-SD177-RP, FT-IspC-SE177-FP, or FT-IspC-SE177-RP) using the following parameters: 5 minutes at 95uC followed by 30 cycles of 30 sec at 95uC, 30 sec at 60uC, 40 sec at 72uC, and a final elongation of 7 min at 72uC. In the second step, 5 mL of each PCR product was appropriately combined with the product obtained from the companion primer (i.e. the product using FT-IspC-SD177-FP was combined with the FT-IspC-SD177-RP product, whereas the products obtained from the PCRs with FT-IspC-SE177-FP and FT-IspC-SE177-RP were combined) and each mixture served as template for a subsequent PCR using T7 forward and reverse primers. PCR conditions were as follows: 5 minutes at 95uC followed by 5 cycles of 30 sec at 95uC, 30 sec at 60uC, 40 sec at 72uC, then 25 cycles of 30 sec at 95uC, 30 sec at 52uC, 40 sec at 72uC and a final elongation of 10 min at 72uC. In the third step, the PCR products from step 2 were amplified using primers FtIspC-f and FtIspC-r (described above), and the resulting products were cloned into pET101/D-TOPO, transformed into XL1Blue cells, and sequence confirmed by restriction digestion and nucleotide sequencing. Each expression vector was introduced into chemically competent E. coli BL21 CodonPlus (DE3)-RIL cells to express the C-terminal His-tagged protein. The S177D and S177E mutants were purified as described for the wildtype His-tagged MEP synthase.

Fluorescence spectroscopy
Fluorescence spectra of MEP synthase and the mutant derivatives were measured using a Fluoromax-3 fluorometer (Horiba Jobin Yvon) at an excitation wavelength of 290 nm using cuvettes with an optical path length of 1 cm. The emission spectra of protein samples with a concentration of 5 mM in 0.1 M Tris pH 7.5, 1 mM NaCl, 5 mM DTT were measured from 310 to 400 nm (excitation and emission slit width = 5 nm). The temperature was maintained at 30uC. All fluorescence spectra were corrected for background with pure buffer.

Enzyme Assays
MEP synthase activity was evaluated at 22uC by spectrophotometrically monitoring the enzyme catalyzed oxidation of NADPH using an assay derived from that described by Takahashi et al [4]. All assays were performed in triplicate. To determine the apparent K M for 1-deoxy-D-xylulose 5-phosphate (DXP), assay mixtures (200 mL) contained 100 mM Tris pH 7.8, 25 mM MgCl 2 , 0.15 mM NADPH, 7 mM MEP synthase, and a variable concentration of DXP (Echelon Biosciences, Salt Lake City, UT). To determine cation specificity, assays were performed with 25 mM MgCl 2 , CaCl 2 , CoCl 2 , CuCl 2 , MnCl 2 , ZnCl 2 , or NiCl 2 . To determine the apparent K M for NADPH, assays were performed with fixed DXP concentration (0.4 mM) and variable NADPH concentration. Nonlinear regression fitting to the Michaelis-Menten equation was used to determine the kinetic constants. Half-maximal inhibition (IC 50 ) by fosmidomycin was determined using a plot of enzyme fractional activity as a function of inhibitor concentration. A plot of K M app as a function of inhibitor concentration was used to determine the K i for fosmidomycin (negative value of the x intercept). Because fosmidomycin is a slow, tight binding inhibitor [5], the enzyme was preincubated with fosmidomycin for 10 minutes prior to addition of substrate. Highthroughput assays were performed using 96-well plates with assay volumes adjusted to 100 mL. The Z-factor was calculated as described by Zhang et al [6] with fosmidomycin as the inhibitor control.

Mass spectrometry method for phosphopeptide identification
To obtain MEP synthase for phosphopeptide analysis, protein expression and purification was essentially as described above, with the exception of using 0.01 mM IPTG for induction. Purified MEP synthase was reduced with 10 mM dithiothreitol (DTT), alkylated by iodoacetamide (50 mM), and then digested by trypsin (Promega) in buffer containing ammonium bicarbonate (50 mM, pH 9) and urea (2 M). The digestion mixture was then desalted by a SepPak column (Waters, Milford, MA). Phosphopeptides were enriched from the tryptic peptides by a TiO 2 column as described by Thingholm et al [7] with modification. In brief, 30 cm fused silica capillary tubing (360 mm OD, 200 mm ID, Polymicro Technologies, Phoenix, AZ) was attached to the frit end of Inline MicroFilter Assembly (Upchurch Scientific), and TiO 2 loose media (GL Sciences, Inc) was slurry-packed into the tubing using a Pressure Cell (Brechbühler Inc.) to form a 200 mm 62 cm TiO 2 column. The SepPak-cleaned sample was mixed with equal volume of Loading Buffer (200 mg/mL 2,5-dihydroxybenzoic acid (DHB), 5% trifluoroacetic acid (TFA), 80% acetonitrile), and loaded into TiO 2 column by Pressure Cell with flow rate of 3 mL per minute. The column was washed with 200 mL Wash Buffer 1 (40 mg/ml DHB, 2% TFA, 80% acetonitrile) and 26200 mL Wash Buffer 2 (2% TFA, 50% acetonitrile) to remove nonphosphopeptides. Phosphopeptides were then eluted off the column by Elution Buffer (5% ammonia solution). Ammonia in the eluate was evaporated by SpeedVac (,3 min), acidified by adding glacial acetic acid to a final concentration of 2%, and desalted using a ZipTip (Millipore). The purified phosphopeptides were analyzed by reversed-phase liquid chromatography nanospray tandem mass spectrometry (LC-MS/MS) using an LTQ-Orbitrap mass spectrometer (ThermoFisher) using previously described methods [8]. Tandem mass spectra were searched using the program SEQUEST (Bioworks software, Thermo) with full tryptic cleavage constraints, static cysteine alkylation by iodoacetamide, and variable phosphorylation of Ser/Thr/Tyr. Phosphopeptide identification was determined using database match scoring criteria filters followed by manual evaluation of the raw data, as described [8].

Molecular Modeling
F. tularensis subsp. holarctica LVS MEP synthase (protein accession number CAJ78974) was homology-modeled using SWISS-MODEL [9] (http://swissmodel.expasy.org/) in automated modeling mode. The template used for modeling was identified via the template identification tool (with default parameters), which performs a primary sequence comparison of the query sequence with those in a structural database. The resolved structure that exhibited the greatest sequence homology with F. tularensis MEP synthase was chosen as the template for modeling (E. coli MEP synthase; PDB ID# 1T1R, BLAST e value = 3610 294 , 48% identity and 66% homology with the F. tularensis sequence). The quality of the F. tularensis MEP synthase model was evaluated using ProQRes [10], which uses atom-atom contacts, residue-residue contacts, solvent accessibility, and secondary structure information to assign an accuracy score from 0 (unreliable) to 1 (reliable). Swiss-PdbViewer 4.0 (http://spdbv. vital-it.ch/) was used to visualize and annotate the model.

Results and Discussion
The F. tularensis MEP pathway as an antimicrobial target Since MEP pathway orthologs are not found in the human genome, the pathway makes an attractive target for the development of novel antibiotics. To assess if MEP pathway inhibition would restrict the in vitro proliferation of F. tularensis subsp. novicida, we monitored bacterial growth in media supplemented with fosmidomycin [11], a strong and specific inhibitor of MEP synthase (E. coli genetically engineered to use mevalonate for IPP biosynthesis (Fig. 1) is unaffected by fosmidomycin when the culture medium is supplemented with mevalonate, but growth is inhibited by fosmidomycin when mevalonate is excluded [12], illustrating the specificity of fosmidomycin for the MEP pathway. Dose-dependent inhibition of purified E. coli MEP synthase [11] and a resolved crystal structure of E. coli MEP synthase bound to the inhibitor [13] further illustrate this specificity). As illustrated in the dose-response plot shown in Fig. 2, fosmidomycin inhibits F. tularensis growth, with half-maximal inhibition at 12 mM (2.2 mg/ mL). A transposon mutant library of F. tularensis [14] further confirms the essentiality of the pathway, as MEP pathway knockouts are lethal, further validating the pathway as an attractive target in F. tularensis. To facilitate drug development, we next set out to clone and evaluate MEP synthase.

Cloning, expression, and purification of F. tularensis MEP synthase
The F. tularensis subsp. holarctica LVS ispC gene, identified via a BLAST search of the genome using the E. coli K12 homologous sequence, is 1158 bp in length and encodes a polypeptide of 385 amino acids with a calculated molecular mass of 42.7 kDa. The subsp. holarctica MEP synthase amino acid sequence shares 99.7, 99.0, and 99.7% identity with the MEP synthase sequence from subsp. tularensis, subsp. novicida, and subsp. mediasiatica, respectively. PCR primer pairs, designed to flank the subsp. holarctica ispC, were used to amplify the gene. The PCR product was cloned into an expression plasmid engineered to express a C-terminal His-tagged protein in E. coli. Purified protein was visualized by SDS-PAGE and Western blot hybridization using an anti-His antibody (Fig. 3).   Size-exclusion chromatography using a calibrated column revealed that F. tularensis MEP synthase exists in solution as a dimer of ,94 kDa (Fig. S2), similar to MEP synthase from Mycobacterium tuberculosis [15] and Synechocystis sp. PCC6803 [16].

Kinetic characterization of F. tularensis MEP synthase
The kinetic activity of purified MEP synthase was spectrophotometrically evaluated by monitoring the substrate dependent enzyme catalyzed oxidation of NADPH (Fig. 1C). Nonlinear regression fitting of enzyme velocity versus substrate concentration was used to determine the apparent kinetic constants (Fig. 4 and Table 1). The K M app for 1-deoxy-D-xylulose 5-phosphate (DXP) was obtained using assays performed with a saturating concentration of NADPH (150 mM), whereas the K M app for NADPH was determined using assays with saturating levels of DXP (400 mM). The K M app, DXP and K M app, NADPH for recombinant F. tularensis MEP synthase are consistent with values reported for the enzyme from E. coli, M. tuberculosis, and Synechocystis sp. PCC6803 (Table 1). The apparent specificity constant of F. tularensis MEP synthase is also comparable to the Mycobacterium and Synechocystis enzymes, although it is 20 fold less than that reported for the E. coli enzyme (due to the difference in k cat DXP ). Assays performed with various divalent cations revealed that recombinant F. tularensis MEP synthase prefers MgCl 2 , although 60% of the enzyme activity is retained with MnCl 2 (Fig. 5).
Having established assay conditions for F. tularensis MEP synthase, we set out to evaluate the protein sensitivity to fosmidomycin. Half maximal activity (IC 50 ) was observed at 247 nM, similar to that reported for the Mycobacterium homolog ( Table 1). The K i for fosmidomycin (99 nM), obtained from a plot of K M app, DXP as a function of inhibitor concentration (Fig. 6), is greater than that reported for E. coli and Synechocystis sp. PCC6803 (Table 1). This may reflect structural differences in the MEP synthase homologs, although this remains to be determined.

F. tularensis MEP synthase in a high-throughput assay
One method of identifying lead molecules in the drug development process involves the screening of large molecular libraries for inhibitors of an assay. A crucial issue for reliable highthroughput screening is the quality and robustness of the assay, often described in terms of the Z-factor [6]. An assay with a Zfactor score between 0.5 and 1.0 is considered excellent for highthroughput screening. To determine the Z-factor for the assay using F. tularensis MEP synthase, we adjusted the assay volume to accommodate a 96-well plate, fixed the substrate concentration at the K M (104 mM), used a saturating concentration of NADPH (150 mM), and evaluated three separate lots of purified MEP synthase in a series of assays performed over three consecutive days. Fosmidomycin was used as a positive control for inhibition. The Z-factor with F. tularensis MEP synthase was found to be 0.8, indicative of an assay (and enzyme) well suited for use in a highthroughput screen.  Phosphorylation of F. tularensis MEP synthase The first committed and principle regulatory step in the MVA pathway is catalyzed by HMG-CoA reductase (reviewed in [17]). Multifaceted control mechanisms regulate HMG-CoA reductase activity, including the modulation of enzyme concentration, the modulation of membrane composition/fluidity, and the regulation of enzyme activity via reversible phosphorylation (specifically, enzyme inhibition by phosphorylation of a serine residue in the catalytic domain [18]). In comparison to the MVA pathway, much less is known about the regulatory mechanisms that control metabolic flux through the MEP pathway. Engineered alterations of MEP pathway gene expression suggest that several MEP pathway enzymes may share control over metabolic flux (reviewed in [19]). The role of posttranslational modification, such as reversible phosphorylation, remains unknown. Thus, we sought to evaluate if a phosphorylation site on F. tularensis MEP synthase could be identified.
Recombinant F. tularensis MEP synthase was purified from E. coli (induced with 10 mM IPTG), subjected to trypsinization, and phosphopeptides were isolated and identified via titanium dioxide chromatography-tandem mass spectrometry. Phosphoserine 177 (equivalent to Ser186 in the E. coli enzyme) was identified (Fig. 7). E. coli Ser186 is located in the substrate binding site, directly interacts with the substrate, and participates in additional interactions that contribute towards protein conformational changes that occur upon substrate binding [13]. Serine 177 presumably has a similar role in the F. tularensis homolog. Phosphorylation of Ser177 is likely to disrupt these interactions, influencing substrate binding and protein conformation, ultimately affecting enzyme activity. To test this assumption, two mutants of F. tularensis MEP synthase were created, S177D and S177E, wherein Ser177 was changed to an aspartate or glutamate, respectively, which serve to mimic a phosphoserine. Each mutant was expressed in E. coli and purified to near homogeneity via a C-terminal His-tag. Relative to wildtype MEP Figure 7. Predicted tertiary structure of F. tularensis MEP synthase, homology-modeled using SWISS-MODEL. A crystal structure of F. tularensis MEP synthase has not been reported. To permit the visualization of phosphoserine177 within the context of the tertiary structure, the F. tularensis MEP synthase was modeled based upon the resolved structure of the E. coli homolog [48] (48% identity, 66% homology). A cartoon representation of the model is shown, with alpha helices colored red, beta sheets colored yellow, and coiled regions colored gray. The beta sheets comprise the dimer interface in the E. coli structure. The quality of the model was evaluated with ProQRes ( Fig. S3) which provides scores ranging from 0 (unreliable) to 1 (reliable). Regions of the model scoring ,0.5 are colored light blue. Primary sequence alignment and the structure of E. coli MEP synthase were used to identify residues comprising the substrate binding site (colored dark blue with backbone and sidechain residues shown). Serine177 (colored green with backbone and sidechain residues shown) is equivalent to E. coli Ser186, which contributes to the substrate binding site and has been shown to participate in conformational changes that occur upon substrate binding [13]. Two tryptophan residues present in the F. tularensis MEP synthase model are colored pink. doi:10.1371/journal.pone.0008288.g007 synthase, a blue shift in the intrinsic fluorescence maximum of S177D suggests changes in the protein globular fold are brought about by the introduction of the Asp residue (Fig. 8A). The slight red shift in fluorescence emission spectrum of S177E is also indicative of a conformational change, although the change in S177E suggests exposure of a tryptophan residue to a hydrophilic environment whereas the blue shift observed with S177D is indicative of a conformational change sequestering a tryptophan in a hydrophobic environment. Like the wildtype enzyme, both mutants exist as dimers in solution, as determined by size-exclusion chromatography (Fig. S2). Enzyme assays performed using the purified mutants reveal that each is inactive (Fig. 8B). Collectively, these results suggest that phosphorylation of Ser177 leads to a conformational change in F. tularensis MEP synthase which inhibits the enzyme and may serve as a control mechanism to regulate metabolic flux through the MEP pathway.

Conclusions
In conclusion, we have shown that F. tularensis MEP synthase is a valid target for the development of novel therapeutics. Inhibition of MEP synthase is sufficient to inhibit the growth of F. tularensis in vitro. Purified MEP synthase is kinetically active and readily lends itself to use in high-throughput screens. Furthermore, our investigation is the first to show that metabolic flux through the F. tularensis MEP pathway may be regulated by phosphorylation of MEP synthase, similar to the regulatory control observed in the MVA pathway. Figure S1 Growth curves of F. tularensis subsp. novicida in minimal media supplemented with the indicated concentration of fosmidomycin (FoS). All conditions were evaluated in duplicate.

Supporting Information
Found at: doi:10.1371/journal.pone.0008288.s001 (3.20 MB TIF) Figure S2 Molecular weight determination by size-exclusion chromatography. Protein standards (&) were used to calibrate the column. Linear regression fitting (R 2 is indicated) generated the standard curve, which was used to determine the molecular weight of IspC, S177D, and S177E. Found at: doi:10.1371/journal.pone.0008288.s002 (1.44 MB TIF) Figure S3 ProQRes evaluation of the F. tularensis MEP synthase structural model generated by SWISS-MODEL. ProQRes uses atom-atom contacts, residue-residue contacts, solvent accessibility, and secondary structure information to score the model over a sliding window of 9 residues [10]. Scores range from 0 (unreliable) to 1 (reliable). Found at: doi:10.1371/journal.pone.0008288.s003 (3.95 MB TIF) Figure 8. Regulation of F. tularensis MEP synthase. A) Intrinsic fluorescence spectra of MEP synthase and its mutants. Wildtype and mutant (S177D and S177E) proteins were adjusted to 5 mM in 0.1 M Tris pH 7.5, 1 mM NaCl, 5 mM DTT and analyzed using an excitation wavelength of 290 nm. The emission spectra was measured from 310 to 400 nm. The Em l max of wildtype MEP synthase was detected at 335 nm, of S177E was detected at 337 nm, and of S177D was detected at 326 nm. The blue shift observed with S177D is indicative of a conformational change sequestering tryptophan residues into a hydrophobic environment. The slight red shift observed with S177E is indicative of a conformational change exposing tryptophan residues to a hydrophilic environment. The increased quantum yield observed with both S177D and S177E is also indicative of a structural change in MEP synthase. B) The relative catalytic activity of wildtype MEP synthase and the S177D and S177E mutants. Assays were performed with 300 mM DXP and 150 mM NADPH. doi:10.1371/journal.pone.0008288.g008