Abstract
Pyridoxal 5’-phosphate (PLP)-dependent enzymes utilize a vitamin B6-derived cofactor to perform a myriad of chemical transformations on amino acids and other small molecules. Some PLP-dependent enzymes, such as serine hydroxymethyltransferase (SHMT), are promising drug targets for the design of small-molecule antimicrobials and anticancer therapeutics, while others have been used to synthesize pharmaceutical building blocks. Understanding PLP-dependent catalysis and the reaction specificity is crucial to advance structure-assisted drug design and enzyme engineering. Here we report the direct determination of the protonation states in the active site of Thermus thermophilus SHMT (TthSHMT) in the internal aldimine state using room-temperature joint X-ray/neutron crystallography. Conserved active site architecture of the model enzyme TthSHMT and of human mitochondrial SHMT (hSHMT2) were compared by obtaining a room-temperature X-ray structure of hSHMT2, suggesting identical protonation states in the human enzyme. The amino acid substrate serine pathway through the TthSHMT active site cavity was tracked, revealing the peripheral and cationic binding sites that correspond to the pre-Michaelis and pseudo-Michaelis complexes, respectively. At the peripheral binding site, the substrate is bound in the zwitterionic form. By analyzing the observed protonation states, Glu53, but not His residues, is proposed as the general base catalyst, orchestrating the retro-aldol transformation of L-serine into glycine.
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Introduction
Found in all living organisms, pyridoxal 5’-phosphate (PLP)-dependent enzymes utilize a phosphorylated, biologically active form of vitamin B6 (Fig. 1a) to catalyze a myriad of chemical reactions including transamination, β- and γ-elimination, α-decarboxylation, retro-aldol cleavages, and glycogen phosphorylation1,2. PLP-dependent enzymes have been organized into seven fold types based on their evolutionary lineages, however, within these grou**s, there are multiple catalytic activities3,4,5. The largest group of PLP-dependent enzymes is the aminotransferase superfamily (Fold I). Hence, PLP-dependent enzymes, owing to their versatility, are physiologically significant for the metabolism, interconversion, and synthesis of various amino acids. Unsurprisingly, many PLP-dependent enzymes are recognized as drug targets for the development of small-molecule therapeutics and have also been exploited for the synthesis of building blocks for pharmaceutical applications6,7.
a Chemical structures of vitamin B6, PLP, and the internal aldimine. Atom labels are given on the internal aldimine structure. b THF-dependent conversion of L-Ser to Gly catalyzed by SHMT. c Overview of the overall fold of SHMT proteins from human mitochondria (hSHMT2) and Thermus thermophilus (TthSHMT). Individual protomers are depicted in different color schemes and PLP cofactors are shown with CPK representation. hSHMT2 exists as a homotetramer made up of obligate dimers, whereas TthSHMT is a homodimer. d TthSHMT and hSHMT2 sequence alignment. Active sites are conserved between the two enzymes, highlighted in red. The sequence identity is 41%.
PLP-dependent catalysis is enabled by the stereoelectronic control of the labile covalent linkage, called the Schiff base, formed between the PLP cofactor and the side-chain amino group of a catalytic lysine (Lys) residue to create the internal aldimine functional state (Fig. 1a), and by the ability of the enzyme active sites to stabilize charged intermediates through the electron withdrawing properties of the pyridine ring8,9,10. These principles, however, do not fully explain the catalytic diversity of the PLP-dependent enzymes, but rather it appears that the manifold of PLP-dependent activities is governed by the surrounding protein environment and the electronic modulation of the cofactor through selective protonation. Consequently, PLP-dependent catalysis cannot be completely understood without knowing the protonation states of the cofactor and the surrounding residues that are dictated by the locations of hydrogen (H) atoms, which determine the electrostatic environment within the active site.
Serine hydroxymethyltransferase (SHMT), a ubiquitous PLP-dependent enzyme in the aminotransferase superfamily, catalyzes the reversible conversion of L-serine (L-Ser) to glycine (Gly), transferring a one-carbon unit to tetrahydrofolate (THF) to yield 5,10-methylenetetrahydrofolate (5,10-MTHF) (Fig. 1b)5,11. SHMT exhibits some catalytic promiscuity, similar to many PLP-dependent enzymes, and can catalyze THF-independent reactions such as the reversible cleavage of β-hydroxy amino acids12, decarboxylation of aminomalonates13, and the racemization and transamination of D- and L-alanine14. As an enzyme in one-carbon metabolism, SHMT is essential to the synthesis of thymine nucleotides, purines, methionine, and other essential biomolecules15, Columns for protein purification were purchased from Cytiva (Piscataway, New Jersey, USA). His-tagged Tobacco Etch Virus (TEV) protease was produced in-house. Crystallization reagents and supplies were purchased from Hampton Research (Aliso Viejo, California, USA). Crystallographic supplies for crystal mounting and X-ray and neutron diffraction data collection at room temperature were purchased from MiTeGen (Ithaca, New York, USA) and Vitrocom (Mountain Lakes, New Jersey, USA). Deuterated L-Ser and hydrogenous D-Ser were purchased from Millipore Sigma (St. Louis, Montana, USA). The glyA gene (Fig. S9) encoding SHMT enzyme from the bacterium Thermus thermophilus was codon optimized, synthesized, and cloned into kanamycin-resistance plasmid, pJ411 (ATUM, Newark, CA), in addition to a DNA sequence encoding for an N-terminal polyhistidine-(His6)-tag with a 34 amino acid long linker. A TEV protease cleavage tag, ENLYFQS, was introduced at the TthSHMT N-terminus sequence so that after cleavage the enzyme sequence started from Ser3 (Fig. S9). This was done because residues Met1 and Val2 are not visible in the electron density maps in the previously deposited TthSHMT structure (PDB ID 2DKJ). The plasmid was transformed into BL21(DE3) competent E.coli cells for expression. Transformed cells were grown in Luria-Bertani (LB) media supplemented with 50 μg/mL kanamycin antibiotic at 37 °C to an optical density of 0.8–1.0 and induced overnight with 1 mM isopropyl ß-D-1-thiogalactopyranoside (IPTG) at 22 °C (approximately 16–18 h). Induced cells were harvested by centrifugation at 5660 rpm at 4 °C, producing typical yields of 10 g of cells per 1 L of cell culture. A buffer containing 50 mM sodium phosphate pH 7.5, 500 mM NaCl, and 10 mM imidazole was used to resuspend the cell pellet, utilizing 5 mL of lysis buffer per gram of wet cell paste. The cells were stirred on ice for 30 minutes prior to mechanical sonication. The cell lysate was clarified by centrifugation at 17,000 rpm (~30,000 g) for 30 min. The supernatant was loaded onto a 5 mL HisTrap FF nickel column equilibrated with 20 mM HEPES pH 7.5, 100 mM NaCl, and 10 mM imidazole and washed with 5 column volumes (CV) of 20 mM HEPES pH 7.5, 100 mM NaCl, and 20 mM imidazole. The pure, tagged protein was eluted with 20 mM HEPES pH 7.5, 100 mM NaCl, and 500 mM imidazole in a linear gradient at relatively low imidazole concentrations (~50–60 mM). SHMT-containing fractions were pooled, and TEV protease was added to cleave the poly-histidine tag (1 mg TEV protease/100 mg of tagged protein). After room temperature overnight dialysis against 20 mM HEPES pH 7.5, 100 mM NaCl, and 1 mM EDTA, the TEV protease-treated fractions were loaded onto the 5 mL HisTrap FF nickel column and eluted in the flow-through. Pure TthSHMT, verified by SDS-PAGE, was then dialyzed overnight against 40 mM NaOAc pH 5.4 and 1 mM PLP at 4 °C, concentrated to 19 mg/mL, and stored at −30 °C in the presence of 20% (v/v) glycerol. The SHMT2 gene encoding the SHMT enzyme from human mitochondrion (residues 37-504, Uniprot ID P34897, Fig. S10) with mitochondrial leader sequence deleted was codon optimized, synthesized, and cloned into kanamycin-resistance plasmid, pJ411 (ATUM, Newark, CA). The DNA sequence encoding for a 35 amino acid long linking sequence containing an N-terminal His6-tag and the TEV protease cleavage tag so that after cleavage the enzyme sequence starts with Gly37 (Fig. S10). The plasmid was transformed into BL21(DE3) competent E. coli cells for expression and grown in LB media supplemented with 50 μg/mL kanamycin antibiotic at 37 °C. The culture was grown to an optical density of 0.8–1.0 and induced overnight with 1 mM IPTG at 22 °C (approximately 16–18 h). Induced cells were harvested by centrifugation at 5660 rpm at 4 °C. Using 5 mL of lysis buffer per gram of wet cell paste, the cell pellet was resuspended in a buffer containing 20 mM HEPES pH 7.5, 500 mM NaCl, and 10 mM imidazole. Lysozyme was added at 0.1 mg/mL as the cells were stirred on ice for 30 min and then subsequently mechanically sonicated. The lysates were clarified by centrifugation at 30,000 g for 30 min and then loaded onto a 5 mL HisTrap FF nickel column equilibrated with 20 mM HEPES pH 7.5, 500 mM NaCl, and 10 mM imidazole. The column was washed with 20 mM HEPES pH 7.5, 500 mM NaCl, and 20 mM imidazole, and tagged hSHMT2 was eluted with a linear gradient of 20 mM HEPES pH 7.5, 500 mM NaCl, and 500 mM imidazole. TEV protease was added to the pooled hSHMT2-containing fractions in order to cleave the poly-histidine tag (1 mg TEV protease/100 mg of tagged protein). The sample was dialyzed overnight at room-temperature against 20 mM HEPES pH 7.5, 250 mM NaCl, and 1 mM EDTA. The TEV protease-treated fractions were loaded onto a 5 mL HisTrap FF nickel column and eluted in the flow-through. Pure hSHMT2, verified by SDS-PAGE, was then dialyzed overnight against 20 mM HEPES pH 7.5, 300 mM NaCl, and 1 mM PLP at 4 °C and concentrated to 18 mg/mL for crystallization setups. hSHMT2 was crystallized in 50 mM Tricine pH 8.2–8.4 and 11-13% PEG 3350 in sitting drop vapor diffusion experiments. Aliquots of pure TthSHMT were thawed and dialyzed against 40 mM NaOAc pH 5.4 and 1 mM PLP to remove the glycerol. TthSHMT (19 mg/mL) was crystallized in 40 mM NaOAc pH 5.5, 1 M (NH4)2SO4, and 0.5 M Li2SO4 in sitting drop vapor diffusion experiments, producing showers of crystals and some crystal aggregates. Several crystal aggregates were crushed in their crystallization drops to create microcrystals for microseeding experiments. Large, single crystals for neutron diffraction were grown using a streak seeding method in 9-well glass plates and sandwich box setups. Specifically, after large crystallization drops were set up, a seeding tool from Hampton Research was dipped into the crushed crystals and then dipped quickly into the new crystal drops to transfer microcrystals. In this approach, only a few, but larger, crystals per drop of TthSHMT grow. A crystal of ~2 mm3 in volume suitable for neutron diffraction was mounted in a 2 mm-inner diameter quartz capillary with a liquid plug of 40 mM NaOAc pH 5.5, 1.0 M (NH4)2SO4, and 0.5 M Li2SO4 prepared in 100% D2O to perform H/D-vapor exchange. The same crystal was used for both neutron crystallographic experiments, in the absence and presence of substrate L-Ser-d7. To prepare the crystal for soaking with L-Ser, the quartz capillary was unsealed, and the H/D-exchange liquid plug was removed. The capillary was then filled with 500 mM deuterated L-Ser, 40 mM NaOAc pH 5.5, 1.0 M (NH4)2SO4, and 0.5 Li2SO4 left to soak overnight. The following day, the soaking solution was removed and replaced with a liquid plug of 40 mM NaOAc pH 5.5, 1.0 M (NH4)2SO4, and 0.5 Li2SO4 in 100% D2O to perform H/D-vapor exchange. TthSHMT crystals for D-Ser soaking experiments were first transferred to a fresh drop containing 0.1 M NaOAc pH 5.5 and 15% PEG 4000, to remove excess sulfate, then moved to a drop containing 0.5 M D-Ser in 0.1 M NaOAc pH 5.5 and 15% PEG 4000. hSHMT2 at 18 mg/mL was crystallized in 50 mM Tricine pH 8.2–8.4 and 11–13% PEG 3350 in sitting drop vapor diffusion experiments producing hexagonal, rod-shaped crystals. Neutron diffraction was tested at room temperature and a preliminary dataset was obtained on the LADI-DALI beamline88 at the Institut Laue-Langevin (ILL) in Grenoble. A complete room-temperature neutron diffraction dataset for the TthSHMT internal aldimine was collected on the IMAGINE89,90,91,92 single-crystal diffractometer at the HFIR at ORNL using a neutron wavelength range of 2.8–4.5 Å. Each neutron image was composed of a 20 h exposure of the crystal held in a stationary position. The crystal was rotated along the vertical axis (Δφ = 8°) before collecting each successive image. The crystal orientation was changed three times by tilting the capillary with respect to the incident neutron beam to improve data completeness. In total, 44 neutron diffraction images were collected. Neutron diffraction data processing was performed with a version of LAUEGEN93,94 modified to account for the geometry of the cylindrical image plate detector. The wavelength-normalization curve was determined using the intensities of symmetry-equivalent reflections at different wavelengths in LSCALE95. No explicit absorption corrections were applied. The data were scaled and merged in SCALA (Weiss 2001). Room-temperature neutron diffraction data for the TthSHMT-L-Ser pre-Michaelis complex were collected on the instrument MaNDi96,97 at the Spallation Neutron Source (SNS) at ORNL. The crystal was held stationary for 20 h exposures and rotated 10° around the φ-axis before collecting the next image. All neutrons between 2 and 4.16 Å were used to collect the frames, with a total of 25 images collected. Neutron diffraction data collection on MaNDi was processed and integrated with 3D time-of-flight profile fitting in Mantid98,99. Wavelength normalization of the data was performed with LAUENORM94,100 and the data were scaled and merged in SCALA101. Neutron data collection statistics for both datasets are shown in Table S1. Room temperature X-ray diffraction data collection for TthSHMT crystals was carried out on a Rigaku HighFlux HomeLab instrument equipped with a MicroMax-007 HF X-ray generator, Osmic VariMax optics, and a DECTRIS Eiger R 4 M detector at Oak Ridge National Laboratory. The data were indexed and integrated using the CrysAlis Pro software (Rigaku, The Woodlands, TX), then reduced and scaled with the AIMLESS program in the CCP4 software suite102,103. The TthSHMT room-temperature X-ray structures were solved by molecular replacement in PHASER104 using phases from PDB code 2DKJ. Room temperature X-ray diffraction data collection for hSHMT2 crystals was performed from a single crystal on the ID19 beamline at SBC-CAT using a Pilatus3 × 6 M detector at the Advanced Photon Source (APS). X-ray diffraction data were integrated and scaled using the HKL3000 software suite105. To minimize radiation damage to the hSHMT2 crystal, the X-ray beam intensity was attenuated 40 times and the data were collected with 0.2 sec/frame rate. The radiation damage to the crystal was estimated by the HKL3000 software to be less than 5%. The hSHMT2 structure was solved by molecular replacement using PHASER104. The cryo-temperature X-ray structure of hSHMT2 (PDB ID 4PVF)18 was used as a starting model. All the structures were subsequently refined against the room temperature data with Phenix.refine from the PHENIX suite106,107 and COOT108,109,110. Geometry validation was aided by Molprobity111. All ligand restraints were generated with eLBOW112 using geometry optimized by quantum mechanical calculations in Gaussian 16113 at B3LYP/6-31 g(d,p) level of theory. Final data collection and refinement statistics can be found in Table S2. Joint XN-refinement of the TthSHMT internal aldimine structure and TthSHMT pre-Michaelis complex were performed using the nCNS114,115 patch of the Crystallography & NMR Systems (CNS)115,116 software suite for macromolecular structure determination. The refinement procedure began with a single rigid body refinement followed by a series of atomic position, atomic displacement parameters, and D atom occupancy refinements. The structures were visualized in the graphics program COOT108,109,110, in between rounds of refinements to inspect side chain modeling and correctly rotate side chain hydroxyl, thiol, and ammonium groups, as well as rotate water molecules to make appropriate H-bonding networks based on both 2FO−FC and FO−FC nuclear scattering length density maps. All water molecules in the model are assumed to be and refined as D2O as a consequence of the H/D-vapor exchange. Because hydrogenous protein was used in this experiment, the protein was modeled with H atoms at non-exchangeable positions, that is in C-H bonds. All labile, thus exchangeable, H positions in the structure were initially modeled as D until their occupancies were refined. An individual occupancy of −0.56 is associated with the presence of pure H at that position, whereas occupancy of 1.00 is indicative of pure D, because the neutron scattering length of H is −0.56 times that of D. Before depositing the neutron structures to the PDB, coordinates of each D atom were split into two records corresponding to an H and a D partially occupying the same site, both with positive partial occupancies that add up to unity. The percent D at a specific site is calculated according to the following formula: % D = (occupancy(D) + 0.56)/1.56. Neutron refinement statistics can be found in Table S1. The 2D relaxed potential energy profiles of the C4′-NSB bond rotation around the PLP pyridine ring were calculated with a simplified model of the internal aldimine (Figure S7). The model was truncated at Cβ of the lysine-portion of the internal aldimine and the phosphate group was removed, making C5′ a methyl group. The scans were completed at B3PW91/Def2-TZVPP level of theory117,118 and reproduced the geometry observed in the experimental methods. Starting from the torsion angle observed in the TthSHMT joint XN substrate-free internal aldimine structure, the full rotation of C3-C4- C4′-NSB was scanned. The scan was also performed on the model with a protonated NSB. All calculations were performed with Gaussian16113. Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.Materials and methods
General information
TthSHMT expression and purification
hSHMT2 expression and purification
Crystallization and H/D-exchange
Neutron diffraction data collection
Room-temperature X-ray diffraction data collection and structure refinement
Joint XN refinement
Quantum chemical calculations
Reporting summary
Data availability
The structures and corresponding structure factors have been deposited into the protein data bank with the PDB accession codes 8SUJ for TthSHMT (Supplementary Data 1), 8SSJ for hSHMT2 (Supplementary Data 2) 8SUI for TthSHMT/L-Ser (Supplementary Data 3), and 8SSY for TthSHMT/D-Ser (Supplementary Data 4). Supporting information is available online.
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Acknowledgements
This research at ORNL’s High Flux Isotope Reactor (IMAGINE beamline) and at ORNL’s Spallation Neutron Source (MaNDi beamline) was sponsored by the Scientific User Facilities Division, Office of Basic Energy Sciences, U.S. Department of Energy. The Office of Biological and Environmental Research supported research at ORNL’s Center for Structural Molecular Biology (CSMB), a DOE Office of Science User Facility. ORNL is managed by UT-Battelle LLC for DOE’s Office of Science, the single largest supporter of basic research in the physical sciences in the United States. X-ray crystallographic data were in part collected at Argonne National Laboratory using Structural Biology Center (SBC) beamline ID19 at the Advanced Photon Source. Use of the Advanced Photon Source, an Office of Science User Facility operated for the U.S. Department of Energy (DOE) Office of Science by Argonne National Laboratory, was supported by the U.S. DOE under Contract No. DE-AC02-06CH11357. The authors thank the Institut Laue Langevin (beamline LADI-DALI) for awarded additional neutron beamtime. This research was supported by a grant from NIH-GMS (R01GM137008) to R.S.P. and A.K.
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R.S.P. and A.K. designed the study. V.N.D., C.C., M.H., A.C., O.G., K.L.W. and R.S.P. expressed and purified the proteins. V.N.D., C.C., O.G., and R.S.P. crystallized the proteins. V.N.D., M.H. and A.K. collected X-ray diffraction data. V.N.D. and A.K. reduced X-ray data and refined the structures. V.N.D., M.P.B., and A.K. collected and reduced neutron diffraction data. V.N.D. and A.K. refined joint XN structures. V.N.D. carried out quantum chemical calculations. V.N.D., R.S.P., and A.K. wrote the paper with help from all co-authors.
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Drago, V.N., Campos, C., Hooper, M. et al. Revealing protonation states and tracking substrate in serine hydroxymethyltransferase with room-temperature X-ray and neutron crystallography. Commun Chem 6, 162 (2023). https://doi.org/10.1038/s42004-023-00964-9
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DOI: https://doi.org/10.1038/s42004-023-00964-9
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