Mechanism of Substrate Recognition and Catalysis of the

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Mechanism of Substrate Recognition and Catalysis of the Haloalkanoic Acid Dehalogenase Family Member, #-Phosphoglucomutase Chunchun Zhang, Karen N Allen, and Debra Dunaway-Mariano Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.8b00396 • Publication Date (Web): 28 Jun 2018 Downloaded from http://pubs.acs.org on June 29, 2018

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Biochemistry

Mechanism of Substrate Recognition and Catalysis of the Haloalkanoic Acid Dehalogenase Family Member, α-Phosphoglucomutase# Chunchun Zhang1, Karen N. Allen2 and Debra Dunaway-Mariano3* 1Testing

& Analytical Center, Sichuan University, Chengdu, Sichuan 610064, P. R. China

2Department 3Department

of Chemistry, Boston University, Boston MA 02215-2521 of Chemistry and Chemical Biology, University of New Mexico, Albuquerque, New

Mexico 87131.

#This work was supported by NIH grant GM61099

*Corresponding author: Debra Dunaway-Mariano at the Department of Chemistry and Chemical Biology, University of New Mexico, Albuquerque New Mexico 87131. Email: [email protected]

Key words: phosphomannomutase, glucose phosphate, mannose phosphate, substrate recognition, domain association

Abbreviations used are: HAD (haloacid dehalogenase), PGM (phosphoglucomutase), PMM (phosphomannomutase), G1P (D-glucose 1-phosphate), M1P (D-mannose 1-phosphate), G6P (Dglucose 6-phosphate), M6P (mannose 6-phosphate), G1,6bisP (D-glucose 1,6-bisphosphate), M1,6bisP (D-mannose 1,6-bisphosphate), Pi (inorganic phosphate), HEPES (4-(2-hydroxyethyl)-1piperazineethanesulfonic acid), G6PD (glucose 6-phosphate dehydrogenase), NAD (nicotinamide dinucleotide), NADP (nicotinamide dinucleotide phosphate) Ll (Lactococcus lactis), Lm (Leishmania mexicana), Hs (Homo sapien), NMR (nuclear magnetic resonance), HPLC (high performance liquid chromatography) and SDS-PAGE (sodium dodecyl sulfate polyacrylamide gel electrophoresis).

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ABSTRACT α-Phosphoglucomutase (αPGM), in its phosphorylated state, catalyzes the interconversion of α-Dglucose 1-phosphate and α-D-glucose 6-phosphate. The αPGM of Lactococcus lactis is a type C2B member of the haloalkanoic acid dehalogenase (HAD) enzyme family, and is comprised of a Rossmann-fold catalytic domain and inserted α/β-fold cap domain. The active site is formed at the domain-domain interface. Herein, we report the results from a kinetic-based study of L. lactis αPGM catalysis which demonstrate enzyme activation by auto-catalyzed phosphorylation of Asp8 with αG1P, the intermediacy of αG1,6bisP in the phospho Ll-αPGM-catalyzed conversion of αG1P to G6P, and the reorientation of the αG1,6bisP intermediate via dissociation to solvent and rebinding. In order to provide insight into the structural determinants of L. lactis αPGM substrate recognition and catalysis, metal cofactor and substrate specificities were determined as were the contributions made by active-site residues towards catalytic efficiency. Lastly, the structure and catalytic mechanism of L. lactis αPGM are compared with those of HAD family phosphomutases L. lactis βphosphoglucomutase and eukayotic α-phosphomannomutase to provide insight into the evolution of phosphohexomutases from HAD family phosphatases.


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Biochemistry

INTRODUCTION α-Phosphoglucomutase (αPGM), in its phosphosphorylated state, catalyzes the interconversion of α-D-glucose 1-phosphate (αG1P) and D-glucose 6-phosphate (G6P) (Scheme 1). In plants and animals, αPGM functions in energy metabolism and in the synthesis and utilization of starch and glycogen.1,

2

In bacteria, αPGM is also required for the synthesis of glucose-containing

polysaccharides used in the formation of the cellular envelope and biofilm. 3-6

Scheme 1. The chemical reaction catalyzed by αPGM and the related pathways. Whereas the eukaryotic phosphoglucomutase is specific for αG1P/G6P, the mutase of bacteria can also catalyze the interconversion of α-D-mannose 1-phosphate (αM1P) and Dmannose 6-phosphate (M6P) and, therefore, is named αPGM/αPMM.7, 8 In eukaryotes, a second mutase, α-phosphomannomutase (αPMM), is employed in the synthesis of the mannose-containing glycan units of glycoproteins and glycolipids.9-11 Eukaryotic αPGM12 belongs to the α-Dphosphohexomutase enzyme family13,14, as does the αPGM/αPMM of Pseudomonas aeruginosa (PaαPGM/αPMM)15. These enzymes are comprised of four structural domains, circularly arranged with the active-site at the center (see Figure 1A). The eukaryote αPMM, on the other hand, is a member of the type C2B clade of the haloalkanoic acid dehalogenase (HAD) enzyme family.16, 17 X-ray crystallographic structural analyses have shown that αPMM is comprised of a conserved


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Rossmann-fold catalytic domain and an inserted α/β-fold cap domain.18,

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19

The domains are

connected by two solvated linkers and are therefore mobile. The structure of the αPMM from Leishmania mexicana (Lm-αPMM)19 is depicted in the domain-domain associated and disassociated conformations displayed in Figure 1B to illustrate the location of the active site at the domaindomain interface and the conformational cycling that takes place in a single catalytic turnover. Whereas catalysis takes place from the domain-domain associated state, substrate binding and product release requires opening of the active-site to solvent via domain-domain disassociation.

Figure 1. X-ray crystal structures of representative hexose phosphate mutases. (A) Phosphohexomutase family αPGM/αPMM from Pseudomonas aeruginosa (PDB accession code 1P5G) where the four domains are colored separately, the phosphorylated Ser108 and Zn2+ are shown in red stick and black sphere, respectively, and the G6P ligand is shown in cyan stick (red oxygen, cyan carbon and orange phosphorus). (B) Overlay of the HAD family type C2B Lm-αPMM structures (PDB accession codes 2I55, chains a and b), in which the catalytic domain is colored teal


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Biochemistry

and cap domain of the active-site open conformer is colored gray whereas that of the active-site closed conformer is colored purple. The Asp nucleophile D10 is shown in red stick, the αG1,6bisP ligand in stick with red oxygen, yellow carbon and orange phosphorus, and the Mg2+ cofactor in charcoal sphere. (C) Overlay of the HAD family type C1 Ll-βPGM structures (PDB accession codes 1LVH and 1O08) where the cap domain of the open conformer is colored gray and that of the closed conformer is colored blue. The Asp8 nucleophile, αG1,6bisP ligand and Mg2+ cofactor are represented the same as in (B). The bacterium Lactococcus lactis, known for its commercial use in the production of fermented milk products20, is unique in that it encodes two different phosphoglucomutases21, 22, both of which belong to the HAD enzyme family23, 24. One these enzymes, β-phosphoglucomutase (βPGM), catalyzes the conversion of the β-anomer of D-glucose 1-phosphate (βG1P) to G6P.21 The βG1P is derived from phosphorolytic cleavage of the disaccharides maltose and trehalose.25 In an earlier effort, we showed that the L. lactis βPGM (Ll-βPGM) is a type C1 HAD family member.23 The type C1 cap domain differs from the type C2B cap domain in its fold (α-helix bundle vs α/β) and in the site of insertion into the catalytic domain.17 The structure of the Ll-βPGM is depicted in the domain-domain associated vs dissociated conformations provided in Figure 1C. The other L. lactis phosphoglucomutase, Ll-αPGM, supports D-glucose-α-1-uridyldiphosphate synthesis and hence, cell wall biogenesis.22 Unlike Ll-βPGM, but the same as αPMM, Ll-αPGM is a type C2B HAD family member. The recently reported X-ray crystal structure of Ll-αPGM in complex with crystallization reagents glycerol and sulfate24 reveals only the domain-domain associated conformation (Figure 2). However, the cycling of Ll-αPGM between this conformation and the domain-domain


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disassociated conformation can be inferred from the reported X-ray crystallographic structures of its structural homolog Lm-αPMM represented in Figure 1B. 19

Figure 2. Illustration of the locations of the proximal and distal substrate binding sites in LlαPGM and the roles of the catalytic residues. (A) The structure of the Ll-αPGMglycerolsulfate complex (PDB accession code 4BND) in the domain-domain associated conformation. The cap domain is colored gray and catalytic domain teal, the glycerol and sulfate ligands are shown in stick (coloring: red oxygen, yellow carbon, orange sulfur) as are the residues of the distal site located in the cap domain (coloring: gray carbon, red oxygen, blue nitrogen) and the catalytic residues of motifs 1-4 (coloring: nucleophile Asp8 red, acid/base Asp10 magenta, the electrophiles Lys 189 blue and Ser41 green, and the Mg2+ ligand Asp212 silver). (B) Schematic depiction of the Ll-αPGM residues

that

form

the

four-motif

catalytic

platform


conserved

among

HAD

family 6

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Biochemistry

phosphotransferases using the color scheme of Figure 2A. Phosphorylated (cyan) D8 (red), the Mg2+ (black) cofactor and the αG1P substrate (cyan) are included. The hydrogen bonds are shown using narrow dashed lines and the coordination bonds between side chains and Mg2+ are depicted using wide dashed lines. Based on the mechanisms of action of eukaryotic αPGM26, bacterial αPGM/αPMM27 and LlβPGM28, we presumed that the chemical sequence for catalysis by Ll-αPGM would be initiated by phosphoryl-group transfer from the phosphorylated active-site nucleophile to the substrate to form αG1,6bisP as the reaction intermediate. The active-site nucleophile of the enzymes of the phosphohexomutase family is a serine residue14 (Figure 1A) whereas the nucleophile of the enzymes of the HAD family is an aspartate residue29,30 (Figure 2). In order to form product, and to regenerate the phosphoenzyme, the αG1,6bisP intermediate must first become reoriented in the active site, which allows the phosphoryl group originating from the substrate to be transferred to the active site nucleophile. As depicted by cartoon in Figure 3, the substrate phosphoryl group is bound by the cap domain at a location termed the “distal site” whereas phosphoryl-group transfer from the aspartylphosphate residue occurs at the “proximal site” of the catalytic domain. The locations of the two sites in HAD family mutases Lm-αPMM and Ll-βPGM are revealed by the X-ray crystallographic structures of the respective αG1,6bisP complexes shown in Figures 1B and 1C.23, 24

The corresponding locations of these two sites in Ll-αPGM are identified in Figure 2A.


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Figure 3. A depiction of the Ll-α αPGM catalytic cycle. In the catalytically active state “EP”, the proximal site (yellow shading) Asp8 nucleophile is phosphorylated. The  phosphoryl group is bound at the distal site located on the cap-domain (cyan shading). Phosphoryl-group transfer occurs in the active-site closed conformer and results in the formation of the αG1,6bisP intermediate. The intermediate must change orientation prior to phosphoryl-group transfer to Asp8 and product formation. Shown by the work reported herein, reorientation in Ll-αPGM occurs by dissociation of the intermediate from the active-site into solvent, followed by rebinding.

The residues of the catalytic domain responsible for mediating phosphoryl-group transfer are highly conserved among HAD family phosphotransferases whereas the residues of the cap domain responsible for substrate recognition are variable. The catalytic domain houses four


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Biochemistry

structural motifs, which form the catalytic platform consisting of an Asp nucleophile and Asp general acid/base in motif 1, a Thr/Ser electrophile in motif 2, a Lys in motif 3, and an Asp ligand to the Mg2+ cofactor in motif 4.30 These residues are identified in the Ll-αPGM structure shown in Figure 2A and their assumed roles in catalysis are depicted in Figure 2B. Residues of the cap domain that form the distal substrate binding site are also highlighted in Figure 2A. By binding to both domains, the substrate stabilizes the enzyme in the catalytically active domain-domain associated conformation. The overall goal of our research program is to gain insight into the process by which the HAD family mutases have evolved from ancestral phosphatases to catalyze intramolecular phosphorylgroup transfer. Herein, the results from a kinetic-based analysis of Ll-αPGM catalysis, which examined substrate recognition, metal-ion activation, enzyme activation by phosphorylation, and the mechanism of αG1,6bisP reorientation are reported. In addition, we compare and contrast the structural determinants and mechanisms of substrate recognition and catalysis operative in the type C2B HAD family Ll-αPGM, with those of the type C1A HAD family Ll-βPGM, the type C2B HAD family αPMM and the α-D-phosphohexomutase family αPGM/αPMM.

MATERIALS and METHODS General. αM1P, αG1P, αG1,6bisP, [14C]-αG1P (49.3 mCi/mmol), [14C]-G6P; (231 mCi/mmol), maltose, phosphoenol pyruvate (PEP), adenosine 5’-triphosphate (ATP), adenosine 5’-diphosphate (ADP), β-nicotinamide (NAD), β-adenine dinucleotide phosphate (NADP), buffers, metal ions, column chromatography materials and growth media were purchased from Sigma-Aldrich. Fructose 6-phosphate kinase type VII from Bacillus stearothermophilus, pyruvate kinase type III from rabbit muscle, L-lactate dehydrogenase type XI from rabbit muscle, glucose 6-phosphate


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dehydrogenase type IX from Baker's yeast (G6PD), phosphoglucose isomerase from E. coli and phosphomannose isomerase from E. coli were also purchased from Sigma-Aldrich. Plasmid DNA was prepared using the QIAprep Spin Miniprep Kit from QIAGEN, while sequencing and mutagenesis primers were purchased from Life Technology. Enzymes for DNA manipulation were purchased from Promega or New England Biolabs and used with the supplied buffers. E. coli BL21 (DE3) competent cells and the pET 3a expression vector were purchased from Novagen. Ni-NTA columns were obtained from QIAGEN and the PiBlueTM phosphate assay kit was purchased from BioAssay Systems. Recombinant L. lactis βPGM23 and maltose phosphorylase21 were prepared according to published procedures. 31P-NMR spectra were recorded at ambient temperature using a Bruker Avance 500 spectrometer. Protein MALDI mass spectra were measured at the University of New Mexico Mass Spectrometry Facility. Syntheses of the Hexose 1,6-bisphosphates. The protocols used for the synthesis of β-Dglucose 1,6-bisphosphate (βG1,6bisP), α-D-mannose 1,6-bisphosphate (αM1,6bisP) and [14C]αG1,6bisP are reported in Supporting Information. tion of the Recombinant lαPGM. Nde I/BamH I restriction sites were introduced at the 5’ and 3’ termini of the L. lactis αPGM gene (UniProtKB accession code Q9CHZ8) by

PCR

using

genomic

DNA

as

template

and

3’)GTTATAATGTAAGCGTTCATATGAAAAAAATAGGAGA

the and

oligonucleotides (5’

(5’

to to

3’)GCTTCGGTTGGAACGCGGATCCCATCAACAACGTGTG as the N-terminus (+) and C-terminus (-) primers. The resulting PCR product was digested with Nde I and BamH I, and after purification, the desired DNA fragment was ligated to a Nde I/BamH I-linearized pET3a vector. The resulting clone (pET3a-αpgm) was used to transform competent E. coli. BL21(DE3) cells. The cells were grown at 37 °C overnight on a LB-Agar plate containing 50 µM ampicillin. The plasmid DNA of one well-


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Biochemistry

isolated colony was extracted, purified and then shown to contain the desired gene by DNA sequencing (Center for Genetics in Medicine, University of New Mexico School of Medicine). The confirmed clone was then used as a source for expression of recombinant Ll-αPGM. Accordingly, transformed cells were grown at 25 oC in LB media for 6-7 h with mixing at 250 rpm and then used to inoculate 2 L × 3 LB medium containing 50 mg/L ampicillin. The cell culture was incubated at 25 oC

to an OD600nm of ~0.7 before the addition isopropyl β-D-1-thiogalactopyranoside (0.4 mM).

Following continued growth at 25 oC to an OD600nm ~1.5, the cells were harvested by centrifugation. Twenty grams of wet cells were obtained from a 6 L culture. The cell pellet was suspended in buffer A (50 mM HEPES containing 5 mM MgCl2 and 1 mM dithiothreitol at pH 7.5 and 4 oC) and then passed through a French Pressure cell press at 1500 psi. The supernatant, obtained by centrifugation of the cell-free mixture, was loaded onto a DEAE-Sepharose column (4.5 x 60 cm) equilibrated with buffer A, and then eluted using a 2 L linear gradient of KCl (0-0.6 M) in buffer A at 4 oC. Fractions containing Ll-αPGM, as judged by SDS-PAGE analysis, eluted at ~0.3 M KCl. These were combined, concentrated with an Amicon protein concentrator, and then loaded onto a phenyl Sepharose column (2.5 x 20 cm) equilibrated with 15% ammonium sulfate in buffer A. The column was eluted with buffer A with 5% ammonium sulfate at 4 oC. The homogeneity of the Ll-αPGMcontaining fractions was examined by SDS-PAGE chromatography. Fractions judged to be >95% pure were combined and then concentrated to 50 mg/mL. Yield: 10 mg protein per 1 g of cell paste. tion of the Recombinant L. lactis femD. The protocol used for gene cloning, recombinant protein preparation and αG1,6bisP synthase activity testing is reported in Supporting Information. Determination of the Equilibrium Constant of the αG1P Reaction. The reaction solution initially containing 0.1 µM Ll-αPGM, 20 µM [14C]-αG1P, 0.2 µM αG1,6bisP, 10 mM MgCl2 and 50 mM


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HEPES (pH 7.0, 25°C) was monitored until equilibrium was reached. Aliquots were removed at varying conversion times, combined with 1 M NaOH and the resulting mixtures were passed through a 5-kD filter to remove enzyme. The filtered samples were then subjected to chromatography by using an HPLC equipped with a CarboPAc PA1 Dionex (4 mm × 250 mm) column. The column was eluted at flow rate of 1 mL/min, first with 2 mL of solvent A (54 mM NaOH and 100 mM sodium acetate), then with a linear gradient (15 mL) of solvent A to 53.7% solvent B (75 mM NaOH and 500 mM sodium acetate), and finally with solvent B. The observed retention times are 5 min for αG1P, 15 min for G6P and 30 min for αG1,6bisP. The eluent fractions (1 mL) were collected and their respective 14C-content was determined by liquid scintillation counting. The Keq value was calculated from the ratio of [14C]-G6P to [14C]-αG1P. Determination of αPGM Steady-State Kinetic Constants. All kinetic assays were carried out at 25 °C in 50 mM HEPES (pH 7.0) containing 5 mM MgCl2, unless stated otherwise. The formation of G6P was monitored spectrophotometrically by measuring the increase in solution absorbance at 340 nm (Δε = 6.2 mM-1cm-1) resulting from the G6PD-catalyzed reduction of NADP. Ll-αPGM-catalyzed conversion of αM1P to M6P was tested by including phosphomannose isomerase (5 unit/mL) and phosphoglucose isomerase (5 unit/mL) in the assay solution. A typical α-G1P to D-G6P conversion reaction was carried out by adding a specified volume of Ll-αPGM stock solution to a 1 mL assay solution containing αG1P, αG1,6bisP, 5 mM MgCl2, 0.2 mM NADP, 5 units/mL G6PD and 50 mM HEPES (pH 7.0, 25 oC). The initial velocities measured as a function of αG1P concentration were fitted to eq 1, whereas those measured at varied αG1P and activator (αG1,6bisP or αM1,6bisP) concentration (0.5 to 5Km) were fitted to eq 2 where [A] is the αG1P concentration, [B] is the αG1,6bisP (or αM1,6bisP) concentration, V0 is the initial velocity, Vm is the


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Biochemistry

maximum velocity, KA is the Michaelis constant for αG1P, and KB is the Michaelis constant for αG1,6bisP (or αM1,6bisP). kcat was calculated from the ratio of Vmax to the enzyme concentration V0 = Vm [A]/(KA + [A])

(1)

V0 = Vm[A][B]/KB[A] + KA[B]+ [A][B]

(2)

pH Rate Profile Determination. Ll-αPGM steady-state kcat and kcat/Km values were measured as a function of reaction solution pH to define the apparent pKa values of ionizing residues. Data were fitted to eq 3, where Y is kcat or kcat/KM, C is a pH independent value of kcat or kcat/KM, [H+] is the proton concentration, Ka and Kb are the respective apparent ionization constants of essential acid and base residues. Log Y = log [C/(1 + [H+]/Ka +Kb/[H+])]

(3)

Assay solutions contained a specific amount of Ll-αPGM, αG1P at varying concentration (0.5 to 5 Km), 100 µM αG1,6bisP, 10 mM MgCl2, 200 µM NADP and 5 unit/mL G6PD in buffer at 25 oC. The reaction solutions were buffered at pH 4.5-5.5 with 50 mM acetic acid plus 50 mM 2-(Nmorpholino)ethanesulfonic acid (MES), pH 5.5-8.0 with 50 mM HEPES and 50 mM MES, pH 8.0-9.0 with 50 mM HEPES and 50 mM [tris(hydroxymethyl)methylamino]propanesulfonic acid (TAPS), and pH 9.0-9.5 with 50 mM TAPS and 50 mM N-cyclohexyl-3-aminopropanesulfonic acid (CAPSO). Control experiments were carried-out by pre-incubating the enzyme in the respective buffers and then assaying its activity at pH 7.0. Metal-Ion Activation and Inhibition of Ll-α αPGM. Activation: Metal-free Ll-αPGM was prepared by dialysis at 4 oC against four changes of 50 mM HEPES (pH 7.0) containing 20 mM EDTA and 1 mM DTT and then against 50 mM HEPES (pH 7.0) containing 1 mM DTT. This procedure removed ~90% of the Ll-αPGM activity, the remaining 10% activity is attributed to the presence of metal ions in the assay solution introduced as contaminants in the reagents. Full catalytic activity


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was restored to the metal-free Ll-αPGM by inclusion of Mg2+ in the assay solution. The initial velocity of G6P formation was measured as a function of MgCl2, NiCl2, CoCl2, MnCl2, ZnCl2 or LiCl concentration using freshly prepared, metal-free Ll-αPGM, in 1 mL assay solutions containing 50 µM αG1P, 100 µM αG1,6-bisP, 200 µM NADP and 5 unit/mL G6PD in 50 mM HEPES (pH 7.0, 25 oC). The initial velocity data were analyzed by fitting to eq 1, in which [A] is the metal ion concentration, KA is the metal ion activation constant, Vo is the initial velocity and Vm is the maximal velocity. In the case of Zn2+, which showed activation at low concentration and inhibition at high concentration, the initial velocity data measured at low ZnCl2 concentration (2-20 µM) were fitted to eq 1 the data measured at high ZnCl2 concentration (50-600 µM) were analyzed by using eq 4 (vide infra). Ca2+ Inhibition: Reaction solutions contained 0.004 µM (metal-free) Ll-αPGM, 10-200 µM MgCl2, 100 µM αG1P, 100 µM αG1,6bisP, 200 µM NADP, 5 unit/mL G6PD, and 0, 10, 20, 40, or 80 µM CaCl2 in 50 mM HEPES (pH 7.0, 25°C). The initial velocity data were fitted to eq 4 where [S] is the αG1P concentration, [I] is the metal cofactor concentration, V0 is the initial velocity, Vmax is the maximum velocity, Km is the Michaelis constant and Ki is the inhibition constant. Vo = Vm [S]/Km(1+ [I]/Ki) + [S]

(4)

Kinetic Analysis of Tungstate Inhibition. The 1 mL assay solutions contained 0.008 µM LlαPGM, αG1P at varying concentration (20-100 µM), 100 µM αG1,6-bisP, 5 mM MgCl2, 200 µM NADP, 5 unit/mL G6PD, and 0, 2, 5 or 10 mM tungstate in 50 mM HEPES (pH 7.0, 25 °C). The initial velocity data were fitted to eq 5 where [S] is the αG1P concentration, [I] is the tungstate concentration, V0 is the initial velocity, Vmax is the maximum velocity, Km is the Michaelis constant and Ki is the inhibition constant. V0 = Vmax[S]/Km + [S](1 + [I]/Ki))


(5)

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Biochemistry

metric Assay for α-G1,6bisP α and G6P Hydrolysis Reactions. αG1,6bisP hydrolysis. Reaction solutions initially contained 2 µM Ll-αPGM, 50 µM αG1,6bisP 5 mM MgCl2, in 50 mM HEPES (pH 7.0 and 25 °C) with and without inclusion of 5 unit/mL G6PD and 200 µM NADP. After varying incubation periods, 150 µL aliquots were removed and mixed with 300 µL of PiBlue phosphate assay reagent. The resulting mixture was incubated for 30 min at room temperature before measuring the measure absorbance at 620 nm (ε = 28.4 mM-1cm-1). G6P hydrolysis. Reaction solutions initially contained 1 mM Ll-αPGM or Ll-βPGM, 2.5 mM G6P and 5 mM MgCl2 in 50 mM HEPES (pH 7.0, 25 oC). Following specific incubation periods, 10 µL aliquots were removed and mixed with 40 µL of 0.2 M HCl, and then assayed for G6P content using 500 µL of 200 mM NADP and 5 unit/mL G6PD in 50 mM HEPES (pH 7.0 and 25 oC). 31P-NMR

Analysis of Ll-α αPGM-Catalyzed Reactions. Reaction solutions initially contained

1 mM Ll-αPGM and 5 mM αG1P, αM1P, αG1,6bisP or αM1,6bisP in 50 mM HEPES (pH 7.0 and 25 °C) containing 5 mM MgCl2.

31P-NMR

spectra were measured at 202 MHz after specific incubation

periods, using phosphoric acid as the chemical-shift standard. Transient Kinetic Analysis of Ll-α αPGM-catalyzed Single-turnover Reactions. The time courses for the Ll-αPGM-catalyzed single-turnover reactions were measured using a KinTek rapidmixing chemical-quench apparatus in conjunction with radiolabelled reactant ([14C]-αG1P or [14C]αG1,6bisP), HPLC-based separation and scintillation counter-based radiolabel measurement. The single-turnover reaction of [14C]-αG1,6bisP was carried out by mixing a 13 µL solution containing 100 µM Ll-αPGM, 2 mM MgCl2 and 50 mM HEPES (pH 7.0) with a 14 µL solution containing 25 µM [14C]-αG1,6bisP, 2 mM MgCl2 and 50 mM HEPES (pH 7.0). The single-turnover reaction of [14C]αG1P was carried out by mixing a 13 µL solution containing 40 µM Ll-αPGM, 2 mM MgCl2 and 50


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Page 16 of 46

mM HEPES (pH 7.0) with a 14 µL solution containing 10 µM [14C]-αG1P and 10 µM αG1,6bisP, 2 mM MgCl2 and 50 mM HEPES (pH 7.0). The reactions were quenched after specified periods of time with 193 µL 1 M NaOH (if the reaction time was shorter than 75 ms, then additional 1 M NaOH was added to adjust the total volume of the reagent to 193 µL). The resulting mixtures were passed through a 5 kDa filter to remove the enzyme and chromatographed on a HPLC equipped with a CarboPAc PA1 Dionex (4 mm × 250 mm) column. The elution program and product analysis used are the same as those described in the section for determination of the solution equilibrium constant (vide supra). The observed rate constants for the single-turnover reactions were obtained by curve simulation using KINSIM32 and by fitting to eq 6 or eq 7 where k is the first-order rate constant, [P]t and [S]t are the product and substrate concentrations at time “ t ”, [S]max is the initial concentration of substrate and [P]max is the product concentration at equilibrium.. [S]t = [S]max–([P]max(1- e-kt)) [P]t = [P]max(1 - e-kt)

(6) (7)

RESULTS and DISCUSSION

Biochemical and Biophysical Properties of Ll-α αPGM. Homogeneous, recombinant LlαPGM was prepared in a yield of 10 mg per g of cell paste. The SDS-PAGE gel of the purified enzyme is shown in Figure S1A. The enzyme was stable to storage in buffered stock solution for 12 h at room temperature and for 1 month at -80 oC. Mass spectral analysis defined the subunit molecular mass to be 28,277 Da, in close agreement with the theoretical mass of 28,276 Da. Because there exists a convenient coupled assay for G6P (G6PD-catalyzed reduction of NADP), and G6P formation from αG1P is thermodynamically favored, Ll-αPGM catalysis was


16

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Biochemistry

studied in the αG1P to G6P direction. The steady-state kinetic constants for Ll-αPGM-catalyzed conversion of αG1P to G6P in solution with 5 mM MgCl2, 50 µM αG1,6bisP and 50 mM HEPES (pH 7.0, 25 oC), are kcat = 44 ± 1 s-1, Km = 39 ± 3 µM and kcat/Km = 1.1 x 106 M-1s-1. The equilibrium constant for Ll-αPGM-catalyzed conversion of αG1P to G6P, in 50 mM HEPES (pH 7.0, 25 oC) containing 0.2 µM αG1,6bisP and 10 mM MgCl2, was determined to be 13 (the time course is shown in Figure S2). We note here that although it is the G6P α-anomer that is formed on the enzyme, once released into solution, it will undergo rapid conversion to the β-anomer (half-life ~1.5 s) to reach an equilibrium ratio of α:β = 4:6.33 Herein, no distinction is made between the G6P anomers and, thus, the term “G6P” represents one or both anomers. Hydrogen Ion and Metal Ion Dependency of Ll-aPGM Catalytic Efficiency. pH Rate Profiles: The variation in Ll-αPGM turnover rate with reaction solution pH, measured at saturating substrate concentration, is defined by the kcat profile. The kcat/Km profile defines the variation in catalytic efficiency measured at sub-saturating substrate. The observed bell-shaped curves of the log(kcat) and log(kcat/Km) vs pH plots (Figure 4) define an optimal pH range for catalysis centered at pH 7, which was used as the reaction solution pH for the experiments reported herein. The apparent pKa values obtained by data fitting are pKa = 5.6 ± 0.6 and pKb = 8.6 ± 0.1 in the case of the kcat profile and pKa = 5.0 ± 0.6 and pKb = 7.4 ± 0.1 in the case of the kcat/Km profile. No attempt was made to assign these values to specific residues, owing to the multiplicity of reaction steps and ionizing groups associated with the catalytic cycle.


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Page 18 of 46

) values measured as a function Figure 4. Log plots of the steady-state kcat () and kcat/Km ( of reaction solution pH for Ll-α αPGM-catalyzed conversion αG1P to G6P. Reaction mixtures contained varying concentrations of αG1P (0.5-10Km), 100 μM αG1,6-bisP, 0.2 mM NADP, 5 unit/mL G6PD, 10 mM MgCl2 and 100 mM buffer at 25 oC.

Metal Ion Specificity: HAD family phosphomutases and phosphatases require a divalent metal ion to serve as cofactor in catalysis, a role filled in vivo by Mg2+. X-ray crystallographic structures of HAD family phosphotransferases have shown that the cofactor binding sites are largely conserved.34 Although the structure of Ll-αPGM24 was determined in the absence of bound Mg2+, the residues that form the binding site can be inferred from the superimposed structure of the homolog Homo sapien-αPMM isozyme 1 (Hs-αPMM1)18 shown in Figure 5. In Hs-αPMM1, a six-coordinate, octahedral Mg2+ geometry is formed through coordination bonds to the carboxylate group of the Asp nucleophile D19, the main chain carbonyl oxygen atom of the downstream Asp acid/base D21,


18

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Biochemistry

the carbonyl oxygen of the N218 side chain and three water molecules. E219, N225 and D226 form hydrogen bonds with two of the water ligands, leaving the third water ligand free to exchange with the transferring phosphoryl group. The superimposed structures shown in Figure 5 indicate that Hs-αPMM1 D19 and D21 are conserved in Ll-αPGM as D8 and D10 and that N225 and D226 are conserved as N219 and D220. On the other hand, N218 and E219 in Hs-αPMM1 are replaced by D212 and M213 in Ll-αPGM. Although two of the Mg2+-binding site residues are not conserved, the net charges of the respective binding sites are the same.

Figure 5. Stereoview of the superposition of the Mg2+ binding site residues in Hs-α αPMM1 with the corresponding residues in Ll-α αPGM. The Mg2+ cofactor is shown in charcoal sphere, the water molecules in red sphere, and the bonding enzyme residues in stick with carbon atoms colored gray (Hs-αPMM1) or teal (Ll-αPGM), oxygen atoms red, nitrogen atoms blue and the sulfur atom yellow. The Mg2+ coordination bonds are represented by charcoal dashed lines and the hydrogen bonds between water ligands and active-site residues are represented by yellow dashed lines.


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The binding affinity of Mg2+ towards Ll-αPGM was examined by measuring the initial velocity of the conversion of αG1P to G6P as a function of Mg2+ concentration. αG1,6bisP was included in the assay to activate the metal-free enzyme by D8 phosphorylation. The plot of initial velocity vs [Mg2+] is shown in Figure S3A and the steady-state kinetic constants derived from the initial velocity data are reported in Table 1. The Mg2+ Kactivation = 29 µM.

Table 1. Steady-state kinetic constants kcat and Kactivation for Ll-α αPGM metal ion activation and/or Kinhibition for Ll-α αPGM metal ion inhibition. Reaction solutions contained metal-free LlαPGM, metal ion activator/inhibitor at varying concentration, 50 μM αG1P, 100 μM αG1,6bisP, 200 µM NADP, 5 unit/mL G6P dehydrogenase and 50 mM HEPES (pH 7.0; 25oC). Reaction solutions used in the determination of the Ca2+ inhibition also contained 20-200 µM MgCl2. See the experimental section for details.

Metal ion kcat (s-1) Mg2+ 23.7 ± 0.4 Ni2+ 19.7 ± 0.5 2+ Co 24.9 ± 0.3 2+ Zn 11.7 ± 0.1 Mn2+ < 10-5 2+ Ca < 10-5 Li+ < 10-5 aND is an abbreviation for “not determined”.

Kactivation (µ µM) 29 ± 0.4 13.3 ± 0.4 5.1 ± 0.3 1.8 ± 0.1 -----------

Ki (µ µM) ------------------~200 NDa 9.8 ± 0.4

Metal-ion binding sites that are formed by clustered carboxylate residues are known to discriminate between monovalent and divalent metal ions, yet they are less discriminatory between Mg2+ and other divalent metal ions.35, 36 Neves et al22 reported that the inclusion of 5 mM Li+, Zn2+,


20

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Biochemistry

Ca2+, Ni2+ or Mn2+ in assay solutions containing 50 µM Mg2+ did not increase Ll-αPGM activity. We too discovered that Li+ does not activate Ll-αPGM by showing that the addition of LiCl2 to the assay solution does not restore activity to the Mg2+-free enzyme. Of the divalent metal ions tested using Mg2+-free Ll-αPGM, Co2+, Ni2+, and Zn2+ showed activation but Mn2+ and Ca2+ did not (see Table 1). The kcat values determined for Co2+ (25 s-1), Ni2+ (20 s-1), Zn2+ (12 s-1) and Mg2+ (24 s-1)-activated LlαPGM are similar. The Kactivation values measured for Co2+ (5 µM), Ni2+ (13 µM) and Zn2+ (2 µM) reflect slightly higher binding affinities for these divalent metal ions than is observed for Mg2+ (Kactivation = 29 µM). However, the comparatively high concentration of Mg2+ in the cell (~1 mM)37, 38 supports Mg2+ in the role of physiological cofactor. An interesting observation made during the measurement Zn2+ activation, is that at higher concentrations, Zn2+ inhibits Ll-αPGM catalysis (Figure S3B). However, the affinity for the secondary binding site is too low (Ki ~200 µM) relative to the concentration of uncomplexed Zn2+ in the cell (picomolar)39 for the inhibition to take place under physiological conditions. Ca2+ displayed linear competitive inhibition vs Mg2+ with a Ki = 9 µM (Figure S3C, Table 1) by competing with the Mg2+ for binding at the cofactor site. Unlike Mg2+, Ca2+ binds to clustered carboxylate residues by forming seven-coordinate complexes in which bidentate coordination of a carboxylate ligand occurs.35 Indeed, this coordination geometry has been previously observed in the Ca2+ complex of the HAD family phosphoserine phosphatase wherein the bidentate coordination to the Asp nucleophile blocks its function.40 However, owing to the predominance of cellular Mg2+, inhibition by Ca2+ does not occur in vivo. 40 The Ll-α αPGM Catalytic Cycle. The Ll-αPGM catalytic cycle displayed in Figure 3 begins with the binding of αG1P to the active site of the phosphoenzyme, and is then followed by phosphorylgroup transfer to the substrate C(6)OH to generate αG1,6bisP as the reaction intermediate. In order


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Page 22 of 46

to form product, αG1,6bisP must first reorient in the active site to place the C(1)-phosphoryl group at the proximal site of the catalytic domain and the C(6)-phosphoryl group at the distal site of the cap domain (Figure 3). Phosphoryl-group transfer from the reoriented αG1,6bisP to the Asp nucleophile D8 forms the product G6P while regenerating the phosphoenzyme. The work described in this section was performed to probe the steps of this cycle. Steady-state Kinetic Mechanism. We began our investigation of the catalytic cycle by determining the steady-state kinetic mechanism of Ll-αPGM catalysis. This was accomplished by measuring the variation in the initial velocity of G6P formation as a function of αG1,6bisP and αG1P concentrations. The data of the parallel double reciprocal plot shown in Figure 6A, support a Bi Bi ping-pong mechanism in which αG1,6bisP binds to the dephosphoenzyme and αG1P binds to the phosphoenzyme. The kinetic constants were defined as: αG1,6bisP Km = 12 ± 0.7 µM, αG1P Km = 45 ± 3 µM and kcat = 133 ± 3 s-1. The Michaelis constants are indicators of the concentrations of αG1,6bisP and αG1P required for half maximal reaction velocity. Substrate inhibition was not observed, indicating that αG1P binding to the dephosphoenzyme is not significant in the presence of αG1,6bisP. The tighter binding of αG1,6bisP indicates that both phosphoryl groups make a significant contribution to the binding energy. Consistent with this conjecture, we observed that the phosphate analog, tungstate, is an uncompetitive inhibitor vs αG1P with a Ki = 4.3 ± 0.2 mM (see Figure 6B), indicative of the formation of the dead-end complex EαG1Ptungstate in which both the proximal and distal phosphoryl-group binding sites are occupied.


22

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Biochemistry

A

B

Figure 6. Plots of steady-state kinetic data measured for Ll-α αPGM activation and inhibition. (A) Double reciprocal plot of the steady-state initial velocity of (0.002 µM) Ll-αPGM-catalyzed formation of G6P from αG1P at varying concentration (20-200 µM), and in the presence of αG1,6bisP at varying concentration (()20, ()50, and () 100 µM). (B) Double reciprocal plot of the steady-state initial velocity of (0.008 µM) Ll-αPGM-catalyzed formation of G6P from αG1P at varying concentration (20-100 µM), and in the presence of tungstate at varying concentration (() 0, () 2, () 5 and () 10 mM). The reaction solutions also contained 100 µM αG1,6bisP, 5 mM MgCl2, 0.2 mM NADP, 5 unit/mL G6PD and 50 mM HEPES at pH 7.0 and 25 oC.

Cellular Ll-αPGM Activation: To enter the catalytic cycle, Ll-αPGM must be phosphorylated. αG1,6bisP is typically included in the in vitro activity assays to serve as the phosphoryl-group donor. However, in order for αG1,6bisP to function in the role of Ll-αPGM activator in L. lactis, it must be biosynthesized in situ. In higher organisms, αG1,6bisP is generated from the reaction of 1,3-diphosphoglycerate with αG1P or G6P, catalyzed by αG1,6bisP synthase.41 A search of the L.


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Page 24 of 46

lactis genome for the encoding gene was carried out by using the human αG1,6bisP synthase sequence as query. The homolog femD, which shares 36% sequence identity with the synthase, was identified as a candidate of interest. To pursue this lead, we cloned and expressed the L. lactis femD gene in E. coli and isolated the recombinant protein. The SDS-PAGE gel of the purified protein (Figure S1B) indicates a subunit mass of 50 kDa, consistent with the theoretical mass of 50635 Da. femD was tested as a catalyst of the phosphorylation of G6P and αG1P in the presence of 1,3diphosphoglycerate and Mg2+ at pH 7.0. However, the formation of αG1,6bisP was not detected (kcat < 1 x 10-4 s-1). Even though the possibility of an undiscovered biosynthetic pathway to αG1,6bisP remained, a more probable alternative was considered, namely that Ll-αPGM is activated via autocatalytic phosphoryl-group transfer from its substrate. Accordingly,

31P-NMR

techniques were used to test Ll-αPGM activation by αG1P. The

reaction solution initially contained 5 mM αG1P, 1 mM Ll-αPGM and 5 mM MgCl2 in 50 mM HEPES (pH 7.0, 25 oC). The

31P-NMR

spectrum measured immediately after mixing contained only one

signal (4.5 ppm), which was that derived from G6P (Figure S4A). The efficiency of G6P formation was then determined, by using steady-state kinetic methods, to define kcat = 23.1 ± 0.6 s-1, αG1P Km = 600 ± 50 µM and kcat/Km = 3.8 x 104 M-1s-1. For comparison, the kinetic constants determined for the conversion of αG1P to G6P in the presence of 50 µM αG1,6bisP are kcat = 44 ± 1 s-1, αG1P Km = 39 ± 3 µM and kcat/Km = 1.1 x 106 M-1s-1. Although less efficient than αG1,6bisP, αG1P can serve as a phosphoryl-group donor to Ll-αPGM. We posit that once the phosphoenzyme is formed, it binds a second αG1P and phosphorylates it to form αG1,6bisP, and through this process Ll-αPGM enters the catalytic cycle.


24

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Biochemistry

Reaction Intermediate: To demonstrate the intermediacy of αG1,6bisP, the single-turnover reaction of 20 µM Ll-αPGM with 10 µM αG1,6bisP and 5 µM [14C]-αG1P was carried-out. We anticipated that the unlabeled αG1,6bisP would be converted to phosphoenzyme and (unlabeled) G6P plus αG1P. Although ~10 µM dephosphoenzyme remains in solution, it is not expected to effectively compete with the phosphoenzyme for reaction with [14C]-αG1P. The time courses for the consumption of [14C]-αG1P and the formation of [14C]-G6P are shown in Figure 7A, along with the time course for the formation and consumption of [14C]-αG1,6bisP. [14C]-αG1,6bisP forms during the first 20 ms of the reaction, in parallel with formation of [14C]-G6P. Whereas the [14C]-G6P continues to accumulate over the next 300 ms, the amount of [14C]-αG1,6bisP peaks at ~10%, and then slowly decreases as a αG1,6bisP: αG1P: G6P equilibrium distribution of 1: 8: 41 is reached. The rate of formation of EαG1,6bisP occurs at 63 s-1, matching the rate of 60 s-1 at which it transitions to the EPG6P complex. Having demonstrated that the EαG1,6bisP complex is a kinetically competent intermediate in the Ll-αPGM catalytic cycle, we next examined the product distribution resulting from the reaction of αG1,6bisP with the dephosphoenzyme. Accordingly, the time courses for formation of [14C]-αG1P and [14C]-G6P were determined for a single-turnover reaction of 12.5 µM [14C]αG1,6bisP and 50 µM Ll-αPGM (Figure 7B). [14C]-αG1,6bisP was converted to [14C]-αG1P and [14C]G6P at a rate of 115 s-1. The reaction rate measured at 100 µM Ll-αPGM is the same as that measured at 50 µM Ll-αPGM (the time courses are reported in Fig S5) and therefore, it is assumed that the phosphoryl-group transfer step, rather than the αG1,6bisP-binding step, is rate limiting. EPαG1P and EPG6P form within the first 20 ms at the respective rates of 100 s-1 and 40 s-1. Over


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the ensuing 200 ms the EPαG1P converts to EαG1,6bisP and hence to EPαG6P, to reach an equilibrium ratio of αG1,6bisP: αG1P: G6P equal to 1:20:90.

Figure 7. Pre-steady state time courses measured for Ll-α αPGM catalysis which illustrate the intermediacy of αG1,6bisP and its efficiency as a phosphoryl group donor to Asp10. (A) Time course for the Ll-αPGM-catalyzed (20 µM) single-turnover reaction 5 µM [14C]-αG1P in the presence of 10 µM αG1,6bisP in 50 mM HEPES containing 5 mM MgCl2 at pH 7.0 and 25 oC. Key: [14C]-αG1P (), [14C]-αG1,6bisP (●) and [14C]-G6P (◆).(B) Time course for the Ll-αPGM-catalyzed (50 µM) single turnover reaction of [14C]-αG1,6bisP (12.5 µM) in 50 mM HEPES containing 5 mM MgCl2 at pH 7.0 and 25 oC. Key: [14C]-αG1,6bisP (●), [14C]-G6P (◆), and [14C]-αG1P ().

Reorientation of the αG1,6bisP Intermediate: Next, we carried out kinetic experiments to determine the mechanism by which the αG1,6bisP intermediate undergoes reorientation in the active site that is required for product formation. The two models considered are illustrated in


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Biochemistry

Figure 3. In one model, the αG1,6bisP changes orientation by “flipping” while remaining associated with the enzyme. In the other model, the αG1,6bisP diffuses from the open active site into solvent, and then rebinds in the opposite orientation. In order to distinguish between these two models, we used radiolabeled substrate, [14C]-αG1P, to test for exchange of the radiolabeled intermediate [14C]αG1,6bisP formed on the enzyme, with unlabeled αG1,6bisP present in the solvent. In the first experiment, a single-turnover reaction of excess phosphoenzyme with 5 µM [14C]αG1P was carried out in the presence of 5 µM unlabeled 5 αG1,6bisP in order to trap the [14C]αG1,6bisP released from enzyme via dilution. The time courses measured for the reaction of 20 µM Ll-αPGM, 25 µM αG1,6bisP, and 5 µM [14C]-αG1P are shown in Figure 8A. Data analysis showed that roughly half of the radiolabel is diverted from the [14C]-G6P fraction to the αG1,6bisP fraction, consistent with the mixing of the ~5 µM [14C]-αG1,6bisP formed in the active site with the ~5 µM unlabeled αG1,6bisP in the solvent. The second experiment was carried out under multipleturnover reaction conditions adapted from those used to test solvent exchange of the αG1,6bisP intermediate formed in the catalytic cycling of Pa-αPGM/αPMM.27 For this purpose, 5 µM Ll-αPGM was used to catalyze the equilibration of the radiolabel contained in 50 µM [14C]-αG1P with 500 µM αG1,6bisP and 500 µM G6P. The maximum amount of enzyme-bound [14C]-αG1,6bisP that can be formed is this reaction is limited to 5 µM. The large excess of unlabeled αG1,6bisP and G6P in solution results in the dilution of the [14C]-αG1,6bisP and [14C]-αG6P released from the enzyme. We note that the rate of equilibration, reflected by the time courses shown in Figure 8B, is slower than that predicted based on the initial velocity and transient kinetic determinations reported above, because αG1,6bisP and G6P sequester Ll-αPGM as the respective EαG1,6bisP and EPG6P complexes, which in turn inhibits the formation of the EPαG1P complex. Nevertheless, the release


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of [14C]-αG1,6bisP from the enzyme to solvent is apparent. Specifically, the radiolabel is almost equally partitioned between solvent αG1,6bisP and G6P. This result is in stark contrast to that observed for Pa-αPGM/αPMM in which αG1,6bisP reorientation takes place on the enzyme within the interdomain space (see Figure 1A).27, 42

Figure 8. Time courses for Ll-α αPGM-catalysis which illustrate the equilibration of the αG1,6bisP intermediate with solvent. (A) Time course for the single-turnover reaction of 20 µM Ll-αPGM, 25 µM αG1,6bisP, and 5 µM [14C]-αG1P in 50 mM HEPES (pH 7.0, 25 oC) containing 5 mM MgCl2. Key: ()[14C]-αG1P, (●)[14C]-αG1,6bisP and (◆)[14C]-G6P. The time courses were fitted with a first order rate equation to define the relative rates of formation kobs = 71 ± 2 s-1 for [14C]αG1,6bis6P and kobs = 8.5 ± 0.3 s-1 for [14C]-αG6P (B) Time course for the multiple-turnover reaction of 5 µM Ll-αPGM, 500 µM G6P, 500 µM αG1,6bisP, and 50 µM [14C]-αG1P in 50 mM HEPES (pH 7.0, 25 oC) containing 5 mM MgCl2. Key: ()[14C]-αG1P, (∆)[14C]-αG1,6bisP and ()[14C]-G6P.


28

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In conclusion, our findings show that the reorientation of the αG1,6bisP intermediate that must occur during the catalytic cycling of Ll-αPGM, proceeds via the αG1,6bisP dissociationrebinding pathway represented in Figure 3. The reorientation does not occur within the active site nor does it occur within the space encapsulated by the subunits of the biological homodimer as was first proposed by Nogly et al24. Substrate

Specificity.

Mutase

vs

Phosphatase

Activity:

The

HAD

family

phosphoglucomutases and phosphatases share a common catalytic scaffold and chemical mechanism. However, the mutases have evolved to suppress the hydrolysis of the phosphoenzyme intermediate, thus allowing phosphoryl transfer to the substrate to take place rather than phosphoryl-group transfer to solvent water. For the purpose of determining the extent to which phospho-Ll-αPGM suppresses phosphoryl-group transfer to water, Ll-αPGM (2 µM) was reacted with αG1,6bisP (50 µM) in the presence of G6PD and NADP to remove G6P formed as product. The rate of formation of inorganic phosphate (Pi) was monitored to define a turnover rate of 8.0 x 10-3 s1

(Figure 9). This rate is 8,000-fold slower than the rate of phosphoenzyme-catalyzed phosphoryl-

group transfer to αG1P (63 s-1).

50 40 [Pi] (µM)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biochemistry

30 20 10 0

0

20 40 60 80 100 120 140 160 Time (min)


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Figure 9. Determination of the rate of Ll-α αPGM-catalyzed phosphate ester hydrolysis in αG1,6bisP. The time course for orthophosphate (Pi) formation during the incubation of 2 µM LlαPGM with 50 µM αG1,6bisP in 50 mM HEPES buffer (pH 7.0, 25°C) containing 5 mM MgCl2, 0.2 mM NADP and 5 unit/mL G6PD (●) is compared with that measured in the absence of NADP and G6PD (□), and with that measured in the absence of Ll-αPGM (◇). Next, we determined the rate of Ll-αPGM-catalyzed hydrolysis of G6P. Accordingly, the time course for G6P hydrolysis was measured for the reaction of 1 mM Ll-αPGM with 2.5 mM G6P in 50 mM HEPES (pH 7.0, 25 oC) containing 5 mM MgCl2 (see Figure S6). The same reaction was carriedout using Ll-βPGM in place of the Ll-αPGM, so that the phosphatase activities of the two mutases could be compared (Figure S6). The Ll-αPGM turnover rate was calculated to be 8.3 x 10-5 s-1, which is 20-fold slower than the Ll-βPGM turnover rate of 1.7 x 10-3 s-1. In addition, the G6P phosphatase activity of Ll-αPGM is 100-fold lower than its αG1P phosphatase activity (8.0 x 10-3 s1).

Stereospecificity: The stereospecificity of Ll-αPGM catalysis was examined by comparing the relative substrate activities of αG1P, its anomer βG1P, and its C(2) epimer αM1P. Reaction solutions initially containing 5 µM αPGM, 100 µM αM1P or βG1P, 100 µM αG1,6bisP and 5 mM MgCl2 in 50 mM HEPES (pH 7.0, 25°C) were monitored for 1 h. During this period, neither βG1P or αM1P was converted to product. Therefore, the turnover rate of these stereoisomers must be

Mechanism of Substrate Recognition and Catalysis of the

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