|USFDC Home | USF Electronic Theses and Dissertations||| RSS|
This item is only available as the following downloads:
Investigation into the Rate-Determining Step of Mammalian Heme Biosynthesis: Molecular Recognition and Catalysis in 5-Aminolevulinate Synthase by Thomas Lendrihas A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy Department of Molecular Medicine College of Medicine University of South Florida Major Professor: Gloria C. Ferreira, Ph.D. Samuel I. Beale, Ph.D R. Kennedy Keller, Ph.D. Randy W. Larsen, Ph.D. Gene C. Ness, Ph.D. Larry P. Solomonson, Ph.D. Date of Approval: June 30, 2009 Keywords: X-linked sideroblastic anemia, -oxoamine synthase, transient kinetics, pyridoxal 5'-phosphate, porphyria, photodynamic therapy Copyright 2009, Thomas Lendrihas
Acknowledgements I wish to ex press my gratitude to the members of my committee, Dr. R. Kennedy Keller, Dr. Randy W. Larsen, Dr. Gene C. Ness, and Dr. Larry P. Solomonson for their consistent guidance, understanding and suppor t throughout the course of my graduate work. Most of all, to Dr. Gloria C. Ferreir a, I am deeply apprecia tive for allowing me the privilege of working with her side-by-side. Her remarkable guidance as both a scientific mentor and cherished friend will never be forgotten. I am grateful to all the professors and colleagues in the Department of Mol ecular Medicine, for th eir intellectual and personal contributions. To Dr. Gregory A. Hunter and Dr. Tracy D. Turbeville, I am indebted for both their scientific and em otional counsel. I wish to express my appreciation to Ms. Kathy Zahn and Ms. Maxine Roth at the Office of Research and Graduate Affairs for their continuous admini strative assistance. Additionally, I would like to specifically thank Ms. Helen Chen -Duncan for her unwavering support and caring as both a colleague and treasured friend. I am forever grateful to my friends: Zena Y. Davis, Julia B. Huddleston, John K. Know les, Mitchell M. McNelly, Laura Jackson Roberts, Louis J. Smith and Thomas F. Za rella, for their enduring encouragement and love. Finally, I wish to acknowledge my fa mily, without whom, this journey would not have been possible.
i Table of Contents List of Tables iii List of Figures iv List of Abbreviations vi List of Schemes ix Abstract x Chapter One 1 INTRODUCTION: The central function of heme: bioge nesis, chemistry and health 1 Enzymes in the heme biosynthesis pathway 2 Aminolevulinate synthase 2 Porphobilinogen synthase 11 Porphobilinogen deaminase 13 Uroporphyrinogen III synthase 15 Uroporphyrinogen decarboxylase 19 Coproporphyrinogen oxidase 21 Protoporphyrinogen oxidase 24 Ferrochelatase 26 Enzymes in the heme degradation pathway 31 Heme oxygenase 31 Biliverdin reductase 34 Content of the dissertation 36 References 36 Chapter Two 51 SERINE-254 ENHANCES AN INDUCED FIT MECHANISM IN MURINE
ii 5-AMINOLEVULINATE SYNTHASE 51 Abstract 51 Introduction 53 Materials 59 Methods 59 Results 65 Discussion 74 Acknowledgements 79 References 79 Chapter Three 82 ARG-85 AND THR-430 IN MURINE 5-AMINOLEVULINATE SYNTHASE COORDINATE ACYL-COA-BINDING AND CONTRIBUTE TO SUBSTRATE SPECIFICITY 82 Abstract 82 Introduction 84 Materials 88 Methods 88 Results 93 Discussion 106 Acknowledgements 114 References 114 Chapter Four 117 HYPERACTIVE ENZYME VARIANTS ENGINEERED BY SYNTHETICALLY SHUFFLING A LOOP MOTIF IN MURINE 5-AMINOLEVULINATE SYNTHASE 117 Abstract 117 Introduction 119 Materials 123 Methods 123 Results 132 Discussion 150 Acknowledgements 156 References 156 Chapter Five 159 SUMMARY AND CONCLUSION 159 References 167 About the Author End Page
iii List of Tables Table 2.1. Summary of steadystate kinetic parameters. 66 Table 2.2. Gibbs free energy associated with the S254 variant-catalyzed reactions. 78 Table 3.1. Comparison of steady-state ki netic constants for wild-type ALAS, R85K, R85L, and R85L/T430V with CoA derivatives as substrates. 95 Table 3.2. Rates of quinonoid intermedia te formation and decay under singleturnover conditions. 103 Table 4.1. Designed mutations for incorporation at indicated positions within the ALAS ac tive site loop. 124 Table 4.2. Amino acids substitutions in active site lid variants. 139 Table 4.3. Kinetic parameters for the reactions of hyperactive ALAS enzymes. 141 Table 4.4. Thermodynamic activation para meters of wild-type ALAS and the SS2 variant. 149
iv List of Figures Figure 1.1. Enzymes and intermediates of the heme biosynthetic pathway. 4 Figure 1.2. The X-ray crystal stru cture of porphobilinogen deaminase from Homo sapiens 14 Figure 1.3. The three-dimensional structure of human uroporphyrinogen III synthase. 16 Figure 1.4. The X-ray crystal stru ctures of coproporphyrinogen III oxidase. 22 Figure 1.5. The three-dimensional st ructure of ferrochelatase from Homo sapiens 28 Figure 1.6. Enzymes in the heme degradation pathway. 32 Figure 2.1. Structural models for mu rine erythroid ALAS based on the R. capsulatus crystal structures. 57 Figure 2.2. Multiple sequence alignm ent of phylogenetically diverse members of the -oxoamine synthase family in the region of murine eALAS serine-254. 58 Figure 2.3. Circular dichroism and fluorescence emission spectra of ALAS and the S254 variants. 67 Figure 2.4. Reaction of th e S254 variants (60 M) with increasing concentrations of glycine. 69 Figure 2.5. Reaction of wild-type AL AS and the S254 variants (5 M) with ALA. 70 Figure 2.6. Reaction of wild-type ALASand S254 variant-glycine complexes with succinyl-CoA under single turnover conditions. 72
v Figure 2.7. Kinetic mechanisms of the S254 variant enzymes. 78 Figure 3.1 The acyl-CoA-binding cleft in R. capsulatus ALAS. 87 Figure 3.2. Comparison of normalized specificity constants for murine eALAS variants with differe nt CoA substrates. 98 Figure 3.3. Visible circular dichroism spectra of wild-type ALAS and the R85 and R85/T430 variants. 99 Figure 3.4. Reaction of wild-type ALAS, R85K, R85L and R85L/T430V (5 M) with ALA. 101 Figure 3.5. Reaction of wild-type AL ASand R85K-glycine complexes with different CoA deriva tives under single turnover conditions. 104 Figure 4.1. The position of the active site loop in the R. capsulatus ALAS crystal structure. 122 Figure 4.2. The generation and sc reening of the library. 126 Figure 4.3. Differential fl uorescence of ALAS varian t isolates streaked on expression agar. 128 Figure 4.4. Multiple alignment of the amino acid sequences of the ALAS loop region. 134 Figure 4.5. The single turnover reac tions of isolated hyperactive ALAS variants. 141 Figure 4.6. The SS2 variantcatalyzed reaction. 147 Figure 4.7. The thermal dependence of th e SS2-variant catalyzed reaction. 148 Figure 4.8. The simulated kinetic mech anism of the SS2 variant-catalyzed reaction. 149
vi List of Abbreviations A-site Acetyl-site AAT Aspartate aminotransferase AIP Acute intermittent porphyria ALA 5-Aminolevulinate ALAD 5-Aminolevulinate dehydratase ALAS 5-Aminolevulinate synthase ALAS1 5-Aminolevuinate synt hase (Non-specific isoform) ALAS2 5-Aminolevulinate synthase (Erythroid-specific isoform) AON 8-Amino-7-oxononanoate AONS 8-Amino-7-oxononanoate synthase Bach-1 Basic leucine transcription factor 1 BVR Biliverdin reductase CD Circular dichroism CO Carbon monoxide CO2 Carbon dioxide CoA Coenzyme-A CEP Congenital erythropoietic porphyria CPK Corey, Pauling and Koulton
vii CPO Coproporphyrinogen oxidase DEAE Diethylaminoethyl EPP Erythropoietic protoporphyria FAD Flavin adenine dinucleotide FC Ferrochelatase GATA1 Globin transcription factor 1 HCP Hereditary coproporphyria HEP Hepatoerythropoietic porphyria HEPES (N-[2-Hydroxyethyl] piperazi ne-N-[2-ethane sulfonic acid]) HMB Hydroxymethylbilane HO Heme oxygenase HRM Heme regulatory motif INH 4-Bromo-3-(5'-carboxy-4'-chlor o-2'-fluoro-phenyl)-1-methyl-5trifluoromethyl-pyrazol IRE Iron response element IRP IRE-binding protein KBL 2-Amino-3-ketobutyrate-CoA ligase meALAS Murine erythroid ALAS mno montalcino (zebrafish variant displa ying defective PPO activity) MOPS 4-Morpholinepropanesulfonic acid NAD+ -Nicotinamide adenine dinucleotide NADPH -Nicotinamide adenine dinucleotide phosphate O2 Diatomic oxygen
viii PBG Porphobilinogen PBGD Porphobilinogen deaminase PBGS Porphobilinogen synthase PCT Porphyria cutanea tarda P-site Propionyl-site PDB Protein data bank PLP Pyridoxal 5-phosphate PPO Protoporphyrinogen oxidase RMSD Root mean square deviation SAM S -adenosyl-L-methionine SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis SPT Serine palmitoyl transferase SS2 Synthetically shuffled variant #2 UROD Uroporphyrinogen decarboxylase UROS Uroporphyrinogen synthase VP Variegate porphyria XLSA X-linked sideroblastic anemia
ix List of Schemes Scheme 1.1. The chemical mechanism of ALAS. 8 Scheme 2.1. The role Ser-254 plays in the chemical mechanism of ALAS. 54 Scheme 3.1. The absorbance maxima of chemical species in the ALAScatalyzed reaction. 98
x Investigation into the Rate-Determining Step of Mammalian Heme Biosynthesis: Molecular Recognition and Catalysis in 5-Aminolevulinate Synthase Thomas Lendrihas Abstract The biosynthesis of tetrapyr olles in eukaryotes and the -subclass of purple photosynthetic bacteria is controlled by th e pyridoxal 5-phosphate (PLP)-dependent enzyme, 5-aminolevulinate synthase (ALAS). Aminolevulinate, the universal building block of these macromolecules, is produced together with Coenzyme A (CoA) and carbon dioxide from the condensation of glycine and succinyl-CoA. The threedimensional structures of Rhodobacter capsulatus ALAS reveal a conserved active site serine that moves to within hydrogen bonding distance of th e phenolic oxygen of the PLP cofactor in the closed, substrate-bound en zyme conformation, and simultaneously to within 3-4 angstroms of the thioester sulfur atom of bound succinyl-CoA. To elucidate the role(s) this residue play(s) in enzyme activity, the equivalent serine in murine erythroid ALAS was mutated to threonine or alanine. The S254A variant was active, but both the SCoA mK and kcat values were increased, by 25and 2-fold, respectively, suggesting the increase in turnover is i ndependent of succinyl-CoA-bindi ng. In contrast, substitution of S254 with threonine results in a decreased kcat, however the Km for succinyl-CoA is
xi unaltered. Removal of the side chain hydr oxyl group in the S 254A variant notably changes the spectroscopic proper ties of the PLP cofactor and the architecture of the PLPbinding site as inferred from circular dichro ism spectra. Experiments examining the rates associated with intrinsic protein fluoresce nce quenching of the variant enzymes in response to ALA binding show that S254 affect s product dissociation. Together, the data led us to suggest that succ inyl-CoA binding in concert wi th the hydrogen bonding state of S254 governs enzyme conformational equilibria. As a member of the -oxoamine synthase family, ALAS shares a high degree of structural similarity and reaction chemistr y with the other enzymes in the group. Crystallographic studies of the R. capsulatus ALAS structure show that the alkanoate component of succinyl-CoA is bound by a co nserved arginine a nd a threonine. To examine acyl-CoA-binding and substrate disc rimination in murine erythroid ALAS, the corresponding residues (R85 a nd T430) were mutated and a series of CoA substrate analogs were tested. The catalytic efficiency of the R85L variant with octanoyl-CoA was 66-fold higher than that calculated for the wild-type enzyme, suggest ing this residue is strategic in substrate binding. Hydrophobic subs titutions of the residues that coordinate acyl-CoA-binding produce ligandinduced changes in the CD spectra, indicating that these amino acids affect substrate-mediated changes to the microenvironment of the chromophore. Pre-steady-state kinetic analys es of the R85K variant-catalyzed reaction show that both the rates associated with pr oduct-binding and the para meters that define quinonoid intermediate lifetime are dependent on the chemical composition of the acylCoA tail. Each of the results in this study emphasizes the importance of the relationship between the bifurcate interac tion of the alkanoic acid component of succinyl-CoA and the
xii side chains of R85 and T430. From the X-ray crystal structures of Escherichia coli 8-amino-7-oxonoanoate synthase and R. capsulatus ALAS, it was inferred that a loop covering the active site moved 3-6 between the holoenzymic and acyl-CoA-bound conformations. To elucidate the role that the active site lid play s in enzyme function, we shuffled the portion of the murine erythroid ALAS cDNA co rresponding to the lid sequence (Y422-R439), and isolated functional variants based on ge netic complementation in an ALA-deficient strain. Variants with potentially greater enzymatic activity than the wild-type enzyme were screened for increased porphyrin overproduction. Turnover number and the catalytic efficiency of selected functional variants with both substrates were increased for each of the enzyme variants tested, suggest ing that increased activity is linked to alterations of the loop motif. The results of transient kinetics experiments for three isolated variants when compared to wild-t ype ALAS showed notable differences in the pre-steady-state rates that defi ne the kinetic mechanism, indicating that the rate of ALA release is not rate-limiting for these enzy mes. The thermodynamic parameters for a selected variant-catalyzed r eaction indicated a re duction in the amount of energy required for catalysis. This finding is consistent with the proposal that, in contrast to the wild-type ALAS reaction, a protein conformational chan ge associated with ALA release no longer limits turnover for this variant enzyme.
1 Chapter One Introduction The central function of heme: biogenesis, chemistry and health Living organisms utilize tetrapyrroles in many important cellular processes. Heme, a ferrous metallated tetrapyrolle, serves a crucial role as a prosthetic group in cytochromes and globins, proteins responsible for respiration (1). As a cofactor, heme is a component of reactions catalyzed by various enzymes including: catalases, peroxidases and the cytochrome P-450s (2, 3). Heme also modulates both steps of the central dogma of molecular biology. Transcription is affected via a signal transduction cascade, whereby heme activates transcripti onal repressors such as Bach-1 (4, 5). Message translation is highly regulated through the phosphorylation of eucaryotic translation initiation factor 2A (eIF2 ) by a number of heme-dependent kinases (6-9). Hierarchically, among eucaryotes and the -subclass of purple photosynthetic bacteria, the heme biosyntheti c pathway consists of eight enzymes (Figure 1.1). The biosynthesis begins with the condensation of glycine and succinyl-CoA and concludes with the chelation of ferr ous iron by protoporphyrin IX (10). Among cells with organelles, all eight enzymes are nuclear encoded; however, the enzymes are distributed in both the cytosol and mitochondria (11). Porphyrias, congenital disorders characterized by the accumulation of porphyrins and porphyrin precursors, occur when gene-derived
2 defects are present among the pathway enzymes (12). Aberrant iron metabolism and associated disorders stem from additional enzymatic deficiencies (13). The catabolism of heme is a tightly regul ated process involv ing heme oxygenase and biliverdin reductase, which together degrade heme to carbon monoxide, iron and bilirubin (14). Heme, present in hemoproteins, is de graded to bile pigments by the action of heme oxygenase (15, 16). Typical inducers of heme oxygenase include: heme, endotoxin, heavy metals and hypoxia (17-19). The role of these molecules and circumstances in the generation of reactive oxygen species has been documented, suggesting that the action of the rate-limitin g step of heme degradation is potently cytoprotective (20). Several disease states are asso ciated with defects in heme breakdown, including atherosclerosis and cancer, as well as a number of inflammatory, autoimmune, and degenerative diseases (21, 22). The enzymes of the heme biosynthetic and degradation pathways have been crystallized. These three-dimensional struct ures provide the framework for identifying structural components of both the molecular ba sis of heme-related disease and catalysis. This introduction describes each enzyme of these pathways in terms of structure and function with the congenital disorder at each step addressed. The focus of this dissertation is on aminolevulinate synthase, th e enzyme on which my theses are based. Enzymes in the heme biosynthesis pathway Aminolevulinate synthase 5-Aminolevulinate synthase (EC 126.96.36.199) (A LAS), is the first, and key regulatory enzyme of heme biosynthesis in non-plant eucaryotes and the -subclass of purple photosynthetic bacteria (Figure 1.1) (23). ALAS catalyzes the Claisen-like condensation
3 of glycine and succinyl-Coenzyme-A to yi eld coenzyme-A (CoA), carbon dioxide (CO2) and 5-aminolevulinate (ALA) (23). For the remaining monera, and in all plants, ALA is synthesized from an alternat ive pathway involving the five carbon skeleton of glutamate (24, 25). Interestingly, the phot osynthetic phytoflagellate Euglena gracilis synthesizes ALA via both pathways, with non-p lastid heme synthesized fr om glycine or the Shemin pathway, and chloroplast heme co ming from glutamate-derived ALA (26). The ALAS-catalyzed reaction takes pl ace in the mitochondria of non-plant eucaryotes (11). This reaction is tightly regulated, and the rate of reaction determines the anabolic production of downstrea m metabolites in the pathway (27, 28). Mammalian genomes contain two genes which code for tw o isoforms of ALAS. The gene encoding the non-specific, or housekeeping form of ALAS (ALAS1), which is constitutively expressed in all tissues, has been localized to chromosome band 3p21 (29). The erythroid specific form of the enzyme (ALAS2) is encoded by a gene located on the Xchromosome at band Xp11.21 (30). The nucleotide sequences of the two genes are notably different; however, the two protei n isoforms share significant similarity (29, 31). Mitochondrial import of ALAS is determined by an N-terminal transit sequence, which is cleaved prior to enzyme maturation (32). ALAS2 from Rattus rattus shows a ~9 kDa difference in monomeric molecular wei ght after the presequence is clipped (33). Heme and iron regulate both gene expres sion and sub-cellular localization of ALAS. Mammalian ALAS proteins contai n heme regulatory motifs (HRM) which consist of a conserved dipeptide, Cys-Pro (32). By way of HRMs, de pleted intracellular heme pools inhibit the mitochondr ial translocation of ALAS1 (34). Plentiful heme on the other hand in differentiating er ythrocytes does not contribute to
4 Figure 1.1 Enzymes and intermediates of the heme biosynthetic pathway. Mitochondrion C y tosol
5 mitochondrial import of the ALAS2 enzyme (34). Transcription of the ALAS1 gene is upregulated by peroxisome prolif erator-activated coactivator 1 (PGC-1 ) (35), through promoter-mediated interactions with both nuclear regulatory factor 1 (NRF-1) and the fox head family member FOX01 (36). Transcription of the AL AS2 gene is regulated by erythroid-specific factors incl uding GATA-binding pr otein 1 (globin transcription factor 1), a protein which is chiefl y responsible for the activati on of globin production in red cells (37). The resultant ALAS2 transcript cont ains a 5' iron regulatory element (IRE) which binds with the IRE-bi nding protein (IRP) in iron -poor conditions, rendering translation impossible (38). Under iron-rich conditions, the IRP-1 contains an Fe-S cluster (38). The incorporation of this prosthe tic group within the protein restricts formation of the IRE-IRP-1 complex, perm itting message translation by the ribosome (38). Ultimately, it is the bioavailability of iron that is the chief modulator of ALAS2 production in vivo (39). The reaction catalyzed by ALAS is markedly similar to those of 2-amino-3ketobutyrate-CoA ligase (KBL ), 7-amino-8-oxononanoate synt hase (AONS), and serine palmitoyl transferase (SPT) (40-43). Based on structure and f unction, ALAS is classified as a fold-type I pyridoxal-5phosphate (PLP)-dependent enzyme and as a member of the -oxoamine synthase subfamily; AONS, KBL a nd SPT represent the closest structural relatives, with the enzymes of the subfamily sharing a C RMSD of 1.5 (44). KBL catalyzes the degradation of threonine (45), AONS, the committed step in biotin biosynthesis (46) and SPT, the first step of sphingolipid biosynthesis (47). In all cases the reduced coenzyme is liber ated and the aminoketone prod uct of the enzyme-catalyzed reaction is further metabolized. Enzymes in the -oxoamine synthase family function as
6 homodimers, with each monomer containing a PLP cofactor at the subunit interface (44). In ALAS, there is one active site per subun it, comprised of residues from adjacent monomers at the dimeric boundary (48). The three-dimensional structure of ALAS from Rhodobacter capsulatus has been solved (49). The bacterial protein exists as a ho modimer, where each monomer consists of three domains. The N-terminal domain, disc rete from the remainder of the enzyme, is defined by an alpha-helix and an anti-parallel beta-sheet. The catalytic domain contains a core parallel beta-sheet flanked by alpha he lices. The C-terminal domain independently interacts with the N-terminal domain thr ough three alpha-helices and with the central core domain of the enzyme via a three-stranded, anti-paralle l beta-sheet. The orientation of the PLP cofactor can be c onsidered to occur through inter actions with three specific protein moieties. First, th e phosphate group is bound tightly via 6 hydrogen bonds (where three are intrasubunit, and three intersubunit). Sec ond, pi-stacking interactions between the conjugated systems of the cofactor and a conserved active site histidine also stabilize the position of PLP. Third, hydr ogen bonding between a conserved aspartate residue and the pyridinium nitrogen enhance the electron withdrawi ng properties of the cofactor. This PLP microenvironment a nd adjacent C-terminal domain delimit the substrate-binding channel, connecting the solv ent exposed surface of the enzyme with the hydrophobic core of the enzyme, where th e acyl-CoA-binding cleft is located (49). Studies focused on the role conserved ami no acids play in the reaction catalyzed by murine ALAS revealed several notable find ings. First, the cat alytic lysine (K313) (ALAS2 numbering), essential for enzyme activ ity and involved in forming a Schiff-base linkage with the PLP cofactor, has been elucidated (50). An aspartate residue (D279)
7 involved in enhancing the electron withdrawin g capacity of the PLP cofactor was also found (51). Recognition of the carboxyl group of th e glycine substrate and binding of the PLP cofactor were found to be dramatically affected when mutations were made to arginine residues, R149 and R439 (52-54). More recently, an active site histidine (H282), was reported to modulate the orientat ion and electronics of the PLP cofactor (55). The ALAS chemical mechanism (Schem e 1.1) is complex and involves: binding of glycine (I); transaldimination with the ac tive site lysine (K313) to yield an external aldimine (II); abstraction of the pro-R proton of glycine (III); condensation with succinylCoA (IV) and CoA release to generate an -amino-ketoadipate intermediate (V); decarboxylation resulting in an enol-quinonoid rapid equilibrium (VI); protonation of the enol to give an aldimine-bound molecule of ALA (VII); and ultimately release of the product (ALA) (VIII) (28). Transient kinetic analyses have indicated that the ratedetermining step of the ALAS-catalyzed reac tion is product release or a conformational change leading to product release (27, 28). The latter of the two possibilities is supported by the observation that the ALAS-catalyzed reac tion rates, when measured with a variety of acyl-CoA derivatives, are comparable (56). The proposed model of ALAS catalysis, base d on kinetic data obtained in both the steadyand pre-steady-states involves transition between open and closed conformations of the enzyme (28). The binding of the second substrate, succinyl-CoA, to ALAS increased the ALAS-catalyzed reaction rate over 250,000 times (57). This finding led to the proposal that part of the intrinsic bindin g energy of succinyl-Co A is utilized to
8 R=OPO3 2Scheme 1.1. The chemical mechanism of ALAS. The individual steps are: binding of glycine (I); transaldimination with the activ e site lysine (K313, mu rine erythroid ALAS numbering) to yield an external al dimine (II); abstraction of the pro-R proton of glycine (III); condensation with su ccinyl-CoA (IV) and CoA re lease to generate an -aminoketoadipate intermediate (V); decarboxyla tion resulting in an enol-quinonoid rapid equilibrium (VI); protonation of the enol to give an aldimine-bound molecule of ALA (VII); and ultimately release of the product (ALA) (VIII).
9 favor the conversion of the population of eq uilibrium conformers to a population of closed conformational species. This inter-conversion between the two conformational states is associated with progression of the reaction and ultimately to restoration of the open conformation, which is concomitant with ALA release. Mutations found in the ALAS2 gene can lead to sideroblastic anemia (58). Sideroblastic anemias are a group of disorders where the circulating erythrocytes appear hypochromic and the marrow is encumbered by ringed sideroblasts (59). The nuclei of these sideroblastic cells are surr ounded by iron-laden mitochondria (60), and thus the designation of ringed sideroblas ts. Diminished ALAS2 activity in red blood cells is the main reason why retained iron is a hallm ark of a defect in heme biosynthesis (61). The most common form of inherited sideroblastic anemia is X-linked sideroblastic anemia (XLSA), a sex-linked, congenital disorder (61). Hemizygous males present the most severe symp toms including: fatigue, diso rientation and both hepatoand splenomegaly (62, 63). The toxicity of excess iron ha s been well-documented and leads to heart disease, liver and kidney failure (62, 63). Specifically, potent oxidation of the cellular milieu by iron-burdened transferritin leads to decreased cell life via generation of reactive oxygen species (60, 64, 65). The majority of ALAS2 mutations l eading to pathological conditions (e.g., XLSA) are missense and are manifested in regions of the protein responsible for interactions between the enzyme and its cofactor (49, 66). In fact, these variants which turn over ALA with less efficiency are, to some extent, responsive to pyridoxine administration (61). Other cases, in which the stability or processing of the enzyme is perturbed, are refractory to this line of therapy (67). Advances regarding protein-protein
10 interactions between ALAS a nd the succinyl-CoA synthetase reveal that abrogation of this association may be res ponsible for the path ological presentation of the defect (49, 68). Until recently, all of the enzymes of the heme pathway in mammals, except ALAS, were recognized to have a porphyr in-associated disorder when defective (69). Now, ALAS2 gene deletions have been iden tified in eight families with the following genetic manifestations c.1706-1709 delAGT G (p.E569GfsX24) or c.1699-1700 delAT (p.M567GfsX24) (70). The corresponding gene product is a truncated form of the erythroid-specific ALAS enzyme (ALAS2), which may be responsible for increased circulating concentrations of protoporphyr in IX. Consequently, these variants demonstrate the first instance of an er ythroid ALAS-related porphyria, X-linked dominant protoporphyria. With this findi ng, several features of the biochemical mechanism of this mode of disease require fu rther research. First, since the experiments were performed on lysates of bacterial cells harboring the expression plasmid for the truncated proteins and not with deletion-vari ants purified to hom ogeneity, the opportunity for other proteins and metabolites affecting th e reaction cannot be rule d out. Next, while the investigators measured ALAS activity in addition to the concentration of reaction products and downstream heme pathway intermediates (71), they did not identify whether the stability of the enzyme or its message was unchanged. Certainl y, elimination of a protein sequence of this magnitude could poten tially affect protein degradation rates as well as message turnover. Thorough biochemical experiments are required to elucidate the complete nature of this intriguing finding.
11 Porphobilinogen synthase The means by which ALA exits the mitochondr ion of eukaryotic cells remains to be elucidated. Once in the cytosol, 2 molecules of ALA are asymmetrically condensed by the metalloenzyme porphobilinogen s ynthase (EC 188.8.131.52; PBGS or ALA dehydratase (ALAD)) (Figure 1.1) (72). The formation of porphobilinogen (PBG), a monopyrrole, is the first common step of tetrapyrrole biosynt hesis among all living organisms. PBGS isolated from different or ganisms, from bacteria to humans, share a high degree of sequence identity (73). These enzymes are large, exhibiting homooctameric quaternary structure, and mo lecular masses in excess of 280 kDa (74). The crystal structure for human PBGS has been determined (75). Each of the four compact homodimers embrace one another using an Nterminal arm, resulting in tetragonal trapezohedral (422) symmetry. With respect to oligomerization, PBGS is an example of a prototypical morpheein ensemble (76). Morpheeins are the building blocks of a group of polypeptides in which a monomer of an enzyme can exist in multiple conformations. Each monomeric conformation affects quaternar y structure differentl y, and the result is an oligomer of distinctive functionality (77). Catalysis by PBGS begins with the fo rmation of independent Schiff base bonds between 2 conserved active site lysi ne residues and the substrates (78, 79). The destination of the ALA molecule within the mo nopyrrole dictates the nomenclature of the active site. The A-site refers to one half of the active site that binds ALA destined for inclusion as the acety l component of PBG, while the pr opionyl-coordinating half of PBG derives from P-site ALA. P-site ALA binds before A-site ALA. Homo-bond formation (C-C) occurs when A-site ALA (C3 position) li nks with ALA in the P-site (C4) position
12via an aldol addition (78). Hetero-bond formation (C-N) follows whereby the P-site substrate amino group attacks the A-site Schiff base. The resultant Schiff-base exchange and liberation of the A-site catalytic lysine permit the energetically favorable steps of aromatization and product release (78). The role of metal ions in the reaction catalyzed by PBGS has been an item of contention. Th ree-dimensional struct ures obtained with a series of substrate and produc t analogs have been complete d, and the role of an activesite zinc has been partially addressed (78, 80). A PBGS structure from Pseudomonas aeruginosa shows that a magnesium ion can be replaced by a zinc ion through the introduction of cysteine resi dues to the metal binding site (78). This suggests that the direct involvement of magnesium ions in the mechanism of magnesium binding to PBGSs, is relatively plastic. Nevertheless, the functionality of metal ions in the mechanism of PBGS remains to be fully elucidated. Mutations that occur in the PBGS or ALAD gene result in a rare recessive autosomal disorder called ALAD porphyria (81). Less than ten cases have been reported that are consequence of a defective gene product (82). In addition to the inherited nature of the disease, heavy divalent metal ions can also illicit symptoms associated with PBGS deficiency (83). Over 80% of the lead found in the human body is bound to ALAD (84). It has been proposed that ALAD porphyria is a disease where pathology stems from defects in conformer equilibrium (85). Gel filtration data obtained using variants encoded by aberrant genes show that oligom erization of mutated enzymes occurs in a manner that favors the less active hexameric st ate. These hexamers, resulting from eight porphyria-associated variants, may be th e first example of a morpheein-based conformational disease. Diminished enzyme activity leads to the accumulation of ALA.
13 Symptoms associated with the poor por phobilinogen production include: intermittent acute neurovisceral attacks and a propensity toward poisoning by lead (86). Inhibition of this enzyme by succinylacetone, a metabolite f ound in among individuals with hereditary tyrosinemia type I, also causes pathological co nditions similar to thos e of lead poisoning and congenital ALAD (80, 87). Porphobilinogen deaminase A cytosolic polymerization reaction where four molecules of PBG are linked is catalyzed by porphobilinogen deaminase (EC 184.108.40.206; PBGD) or hydroxymethylbilane synthase) (Figures 1.1 and 1.2). The physiolo gically relevant re action product is the linear tetrapyrrole hydroxymethylbilane (HMB ) a.k.a. pre-uroporphyrinogen (74). HMB is exceedingly unstable and can undergo s pontaneous cyclization to form the nonphysiological isomer uroporphyrinogen I (88). PBGD sequences are highly conserved throughout evolution and among di verse phyla; to date, all isolated enzymes contain a unique cofactor, dipyrromethane (89). PBGD functions as a monomer and the human crystal structure was recently solved (90). Human PBGD consists of 344 residues with a molecular weight of ~37 kDa. The three-dime nsional structural anal ysis shows that the monomeric protein is organized into three equal-sized domai ns. Domain I houses most of the catalytic and substratebinding residues, while domain II is responsible for cofactor binding. The dipyrromethane cofactor is covalently linked to a loop of residues comprising domain III and is perched at the opening of the active site cavity, now known to be delimited by cleft formed by domains I and II (90). A novel mechanism defines the generation of the unique dipyrromethane cofactor (91). During turnover, the enzyme first binds HMB, then deaminates and polymerizes 2 molecules of PBG to form a
14 Figure 1.2. The X-ray crystal structure of porphobilinogen deaminase from Homo sapiens (PDB code: 3ecr) The monomeric protein is organized into three equal-sized domains, which are comprised of both -helices (green) and -sheets (rust) (A). Perched at the top of the enzyme is the unique dipyrro methane cofactor (depic ted as sticks in CPK color format) (B). A B
15 hexapyrrole. Subsequently, PBGD cleaves the di stal tetrapyrrole and releases HMB. A thioether linkage to an active site cysteine re tains the dipyrrole cofactor for the lifetime of the enzyme (91). Acute intermittent porphyria (AIP) is due to an autosomal dominant pattern of inherited mutations of the PBGD gene le ading to diminished enzyme activity (12). Hundreds of PBGD mutations have been id entified, with acute a ttacks of porphyria affecting 1:100000 individuals, with presen tation more common in women than men (92). However, only recently was a poly-dele tion mutant identified in exon 15 of the PBGD gene (93). The deletion occurs in a conserve d region of the pr otein where other disease causing mutations have been discovered (94). This particular defect clearly has a more substantial negative effect on catalysis, as the ALA concentration identified in the urine of the patient was 100 times that of normal (92). Symptoms of AIP include abdominal pain and other neurovisceral and ci rculatory disturbances ultimately resulting in tachycardia (95). A majority of the mutations id entified in the human PBGD gene perturb carboxylate binding betw een conserved arginines and the cofactor or substrate (96). However, recently, some of the mutati ons documented in patients suffering from AIP were found to be located distal from the active site (94). Uroporphyrinogen III synthase HMB serves as the substrate for the f ourth enzyme of the heme biosynthetic pathway, uroporphyrinogen III synthase (E C 220.127.116.11) (UROS) (Figure 1.3). UROS catalyzes closure of the tetrapyrrole m acrocycle by inverting the D-ring of HMB (74). All identified UROS enzymes function as a monomer with a molecular weight of ~30 kDa (97-100). UROS proteins from all kingdoms of life are similar with respect to
16 Figure 1.3. The three-dimensional structure of human uroporphyrinogen III synthase. (PDB code: 1jr2) The functional monomer c ontains two unambiguous domains connected by a short -ladder (yellow). Each dom ain is characterized by a sheet core (magenta) surrounded by -helices (teal).
17 molecular mass; however si gnificant sequence devia tions have been observed (101). Specifically, the sequence similarity between mammalian and bacterial UROS is less than 22%. Notably, recent three-dimensional studies on UROS from the gram-negative eubacterium Thermus thermophilus identified significant conformational information (102). In these experiments eight crysta llographically unique UROS structures (consisting of apoenzymic, ligand-bound and pr oduct-bound forms) were overlaid. From these maps, significant enzyme flexibility was observed, including a snapshot of the product-bound enzyme in the closed conformation (102). The X-ray crystal structure of human UROS revealed that the enzyme contains two una mbiguous domains connected by a short -ladder (103). Each domain is characterized by a -sheet core surrounded by -helices. The substrate bindi ng cleft is located at the dom ain interface and delimited by a series of evolutionarily conserved residues. The UROS-catalyzed reaction proceeds by way of a spirocyclic pyrrolenine intermediate (104). This intermediate occurs after a rearrangement of the A-ring of HMB, which results in the concomitant lo ss of the C20 hydroxyl group and the formation of a carbocation at C20. C16 of the D-ring is then susceptibl e to electroph ilic attack and the spirocyclic pyrrolenine intermediate is generated. Subseque ntly, azafluvene is formed permitting cyclization and D-ri ng inversion to yield the product uroporphyrinogen III. From the structural data, it was inferred that the interactions of the A and B ring carboxylate groups with both stru ctural domains of UROS are the chief modulators of the closed enzyme conformation (104). The C and D rings demonstrate increased flexibility, a characteristic consis tent with the sterically-mediated acts of catalytic cyclization and D ri ng inversion. Biological rele vance of all porphyrins is
18 demarcated by asymmetry about the D-ring of tetrapyrolles. Without enzymatic conversion, HMB spontaneously cyclizes to a toxic dead e nd product, which in patients with UROS deficiency results in accumulation of uroporphyrinogen I (105). Mutations to the UROS gene manifest as a rare form of porphyria called congenital erythropoietic porphyria (CEP) (105). The defective enzyme is inherited as an autosomal recessive trait. In CEP or Gnther disease, HMB is non-enzymatically converted to uroporphyrinogen I and is subseque ntly catalyzed by the fifth enzyme of the heme biosynthetic pathway to coproporphyrin I (105). Recently, a thorough study was conducted where 25 missense mutations were cl oned into expression vectors, and the respective proteins were purified to homogeneity and characterized (106). Kinetics experiments indicated that most mutated en zymes had significantly decreased activity, while others maintained reaction rates compar able to those of the wild-type enzyme. This suggested that mechanisms besides tu rnover may be responsible for the pathology observed in CEP. Located in -helix 3, perched above the ac tive-site, a conserved active site cysteine was the focus of experiments related to enzyme structure. Significantly, unfolding experiments performed on variants of this cysteine residue may be crucial for proper folding and turnover of uroporphyrinogen III (106). Coproporphyrin I is the causative agent of erythrodontia or red staining of the teeth (105). Patients are either homozygous for a single polymorphism or are compound heterozygotes for a variant form of the enzyme (105). In one particular case, CEP was diagnosed in a patient with a mu tation-free form of the enzyme (107). The causative agent for this pathology was faulty transcrip tion, a defect which was linked to a mutation
19 in the erythroid-specific GATA1 transcriptio n factor. These part icular data provide evidence that the functional responsiveness of erythroid specific promoters is different. Uroporphyrinogen decarboxylase The acetate side chains of uroporphyri nogen III are decarboxylated in four successive steps by uroporphyrinogen d ecarboxylase (EC 18.104.22.168; UROD), leaving the four methyl groups character istic of the product copr oporphyrinogen III (Figure 1.1) (74, 108). UROD exists as a homodimer, with a monomeric molecular mass of ~40 kDa (109, 110). Unlike most decarboxylases, UROD activity is independent of a prosthetic group or cofactor (111-113). Sequence similarity among isol ated UROD proteins is low; however common structural features have b een identified between the human and plant enzymes (109, 114). Three-dimensional studies have shown that the monomer contains a single domain characterized by a distorted ( / ) barrel, with the activ e site housed at the end of a deep cleft, delimited by the C-term inal loops of the barrel. Evolutionarily conserved residues line the cleft, and seve ral invariant basic re sidues are crucial for binding the propionate gr oups of the substrate (109, 110). A dynamic active site loop located at the head of the ac tive site undergoes conformationa l changes to allow substrate entry and reorganization of the catalytic cleft (109, 115). After the asymmetric D-ring of uroporphyrinogen III is decarboxylated, sequential removal of the acetate gro ups proceeds in a clockwise manner (114, 116, 117). To accomplish this, sequential decarboxylation requires the 180o reorientation of the intermediate, a process whereby the substr ate is flipped around its C10-C20 axis. Controversy exists as to the mechanism by wh ich the remaining steps take place. Several
20 theories state that the dimeric structure of UROD implies a dimer-dependent catalysis (114, 116). Since each subsequent decarboxylation only requires a 90o rotation, it has been postulated that the two monomers coll aborate during the decar boxylation of a single uroporphyrinogen substrate. It has been propos ed that the dimeric organization of UROD protects the reactants from solvent exposure, allowing the reaction intermediates to be passed and chemically modi fied between each monomer (114, 116). Further, analysis of UROD compared with another cofactorle ss decarboxylase, orotidine 5'-monophosphate decarboxylase (ODCase), indicates that a protonat ed basic residue assists the transition of the polar carboxylate groups from water to the comparatively less polar hydrophobic core by stabilizing the postscission carbanion (118). Alternatively, based on the structure for the human UROD bound to coproporphyrinogen, the UROD-catalyzed reaction may take place at a unique site on the enzyme surface (117). Mutations in the human gene are responsible for the familial form of porphyria cutanea tarda (PCT) (119). The disease has been classified into three sub-types. Type I PCT has decreased hepatic UROD activity, but normal erythrocyte UROD activity (119). Type II PCT has decreased UROD activity both in red cells and hepatocytes (119). Type III PCT is similar to type II, but erythrocyte UROD activity is normal (119). PCT is an autosomal dominant trait; however symptoms are rarely present in heterozygotes. Recently, three children from the same family presented with symptoms associated with early onset PCT (120). Analysis of the UROD gene fo r all three probands indicated two novel missense mutations and one previously identified polymorphism, giving these patients an unique and previously unide ntified compound heterozygote genotype (120). Dermatitic photosensitivity is the hallmark symptom of PCT (119); a clinical
21 manifestation which shows a marked increas e in intensity among individuals with comorbidities such as: hepatitis, HIV infection, or proclivity to imbibe (86). Instances where severe symptoms occur at early onset are indicative of an accessory condition named hepatoerythropoietic porphyria (HEP ). HEP is frequently diagnosed in homozygotes or among individuals with compound heterozygosity (121, 122). Coproporphyrinogen oxidase The antepenultimate enzyme of the heme biosynthetic pathway catalyzes the sequential oxidative decarboxylation of ri ngs A and B to form protoporphyrinogen IX (Figure 1.1). Coproporphyrinogen oxidase (EC 22.214.171.124; CPO) in humans is oxygendependent, found in the intermembranous space of mitochondria and produces coproporphyrinogen III, carbon di oxide and hydrogen peroxide (123, 124). The enzyme is targeted to the organelle by way of an unusually long leader sequence of 110 amino acids (125, 126). Oxygen-dependent CPOs are found in all eucaryotes and a select group of aerobic procaryotes (127). Sequence similarity among the oxygen-dependent enzymes is high, and to date a requirement for pros thetic groups or cof actors has not been identified (127). The human enzyme exists as a ho modimer, with a monomeric mass of ~37 kDa (Figure 1.4A) (128). CPO contains an elaborate subunit interface w ith multiple conserved residues, suggesting a role of dimeric assembly in stabilizing the catalytically competent conformers of the enzyme. Each monomer is composed of a central antiparallel -sheet flanked by -helices. The active site, de limited on both sides by the sheet and helices, elegantly houses the subs trate while minimizing contacts with the solvent. At the head of the active site, an -helix acts as a lid to modulate the solvationstate of the active-site cleft (128).
22 Figure 1.4. The X-ray crystal structures of coproporphyrinogen III oxidase. Each monomer of the dimeric human enzyme (PBD code: 2aex) is composed of a central antiparallel -sheet (red) flanked by -helices (green) (A). The E. coli enzyme (PDB code: 1olt) functions as a dimer (monomer shown) (B) and contains a catalytically essential Sadenosyl-L-methionine cofactor and a [4S-4S] cluster (C). B A C
23 Oxygen-independent CPOs are remarkab ly different from CPOs with oxygen requirements. CPO from Escherichia coli is a complex monomer of ~53 kDa (Figure 1.4B) (129). The enzyme, a member of the Radical SAM family of proteins, utilizes a particularly labile [4Fe-4S] cluster to facilitate electron transfer to S-adenosyl-Lmethionine (SAM) (Figure 1.4C). As an oxi dizing agent, SAM accepts one electron from the substrate in one catalytic turnover; and an as yet unidentified substrate accepts another electron. The identity of this acceptor molecule is of particular interest because anaerobes do not utilize oxygen, the physiologi cal electron depository for the product of the aerobic reaction. With respect to th e mechanism of catalysis, the anaerobic conversion of coproporphyrinogen III to pr otoporphyrinogen IX is only partially understood (130, 131), although two of the steps are well documented. These steps are the generation of the 5'-deoxyadenosyl radica l from the reductive cleavage of SAM and the radical-mediated proton abstraction from the B-carbon of the propionate side chain. A recent study, centered upon the conserved his tidines of human CPO, suggests that catalysis can occur despite alterations to these evolutionarily selected residues (132). Further work by the same group using substr ate analogs where the C and D rings were modified to replace alkyl gr oups with the ring 13and 17 -propionate moieties led the investigators to postulate that the propiona te side chains of rings C and D play a significant role in both substrate binding and turnover (133). Be that as it may, the order of ring decarboxylation and how the processive reorganization of the substrate takes place (i.e., the manner in which the substrate ro tates) remains to be elucidated. Mutation in the human CPO gene are asso ciated with hered itary coproporphyria (HCP), an acute condition of the liver (134). While the disease is inherited in an
24 autosomal dominant manner, variable penetrance of the trait is observed (12). Most of the disease-causing mutations have been mappe d to portions of the enzyme proposed to be responsible for maintaining enzyme stability (135). Work done by Stephenson et al. involving three invariant amino acids found in human CPO led to the identification of their roles in both the deprotonation of the s ubstrate and bifurcate interactions between the carboxylate tail of the subs trate and two arginine residues (136). However, only one arginine residue (Arg401) has been repor ted to be mutated in porphyric patients, suggesting that the pathological basis of HCP may be more complex. The most prominent biochemical feature of HCP is a marked increase in excreted coproporphyrinogen III; concentrations are typically 10200 times higher compared with controls (137). A severe variant form of HCP is known as harderoporphyria. This disorder is characterized by earlier onset of neurovisceral attacks compared to HCP, and massive excretion of a tricarboxylated porphyrin (harderoporphyrin) in the feces (138). Protoporphyrinogen oxidase Protoporphyrinogen IX is aromatized to protoporphyrin IX by the penultimate enzyme of the heme biosynthetic path way, protoporphyrinogen oxidase (EC 126.96.36.199; PPO) (Figure 1.1) (139). This step of the pathway c onstitutes a branch point whereby protoporphyrin IX is supplied to produce ch lorophyll or heme. PPOs from diverse phyla require FAD as a cofactor (140). This cofactor is coordinated by the macromolecule via a highly conserved N-terminal di nucleotide binding motif (GXGXXG) (141). Human PPO exists as a homodimer, with a single cofactor per dimer (142). Conversely, PPO of the facultative anaerobe Bacillus subtilis is a monomer of ~52 kDa (143). Structural information has been deduced from the crystal structure of PPO from Nicotiniana
25tabacum (144). This enzyme exists as a loosely associated dimer with three defined monomeric domains. The domains are res ponsible for FAD-, protoporphyrinogen IX-, and membrane-binding. The active site of PPO is located between the FADand the substrate-binding domains. Mutations associ ated with the human condition, variegate porphyria (VP) have been mapped using the plant model with notable success (145). A binding mechanism for the PPO-cataly zed reaction has been proposed based on experimental results obtained with the I NH (4-bromo-3-(5'-carboxy-4'-chloro-2'-fluorophenyl)-1-methyl-5-trifluoromethyl-pyr azol) and acifluorfen inhibitors (144, 146). The pyrazole ring of INH serves as an A ring mode l, which is coordinated in the active site by pi-stacking interactions w ith a conserved phenylalanin e residue. Ring B of the macrocycle substrate is stabilized by hydr ophobic interac tions provided by the side chains of two highly conserved leucine residu es. Oxidation of the C20 methylene bridge between rings A and D occurs by way of the FAD cofactor. Next, imine-enamine tautomerizations initiate a ll hydride transfers from C20. The three remaining oxidation reactions involving FAD generate three hydrogen peroxide mo lecules. Curiously, the candidates for the catalytic base involved in this reaction are not evolutionarily conserved (144). As such, a potential base which coul d execute proton abstraction from the substrate is the FADH--derived peroxide anion. PPO deficiency causes VP (147). Missense, nonsense, an d splice site mutations have been identified as the root of VP in most cases (145). The disorder, inherited as an autosomal dominant trait, is highly preval ent in South Africans of European descent (148). A founder mutation attribut ed to a missense mutation at position 59 (R59W) has been traced to an immigrant from the 17th century and results in a PPO with reduced
26 catalytic activity (148). The genetic drift of the abnorma l gene has resulted in very high prevalence; as many as twenty thousa nd South Africans may carry this gene (149). VP presents as acute neurovisceral attacks and/or dermatitic photosensitivity. It is the variability observed in pathological presentatio n of the disorder that is the basis of the nomenclature of the disease (86). Sudden death associated with the acute visceral attacks of VP, highlight the importance of identif ying silent carriers of mutated genes (150). Efforts to examine the mechanism of presenta tion and drift of the disorder are underway by a team at Harvard University (151). Their work involving a genetic screen of hematopoeitic chordate mutants, identified a zebrafish (Danio rerio) variant that showed defective PPO activity (151). This montalcino (mno) variant presented with hypochromic anemia and porphyria, which was partially ameliorated when hu man PPO mRNA was microinjected into mno embryos. Rescue of the mno phenotype by overexpression of human PPO suggests functiona l conservation of the enzy me across chordates. Consequently, mno appears an excellent model for inve stigation of PPO and a valuable tool for identification of therap eutic agents of VP. Increased plasma porphyrin in VP is detected by monitoring fluorescence emi ssion at 626628 nm, upon excitation at 405 nm (152). Other porphyrias including EPP and PC T are marked by fluorescence emission peaks 636 nm and 618622 nm, upon exci tation at 405 nm, respectively (153). Incidentally, patients with a rare homozygous form of the disorder have a notable increase in red cell Zn-protoporphyrin (154). Ferrochelatase Ferrochelatase (EC 188.8.131.52) (FC) is the last enzyme in the heme biosynthetic pathway, and it catalyzes the in sertion of ferrous iron into protoporphyrin IX to yield
27 protoheme (Figure 1.1) (155, 156). In vivo, the enzyme associates with the inner membrane of the mitochondrial matrix (155). The penultimate enzyme of the pathway, PPO, has been proposed to in teract with FC directly, by transferring protoporphyrinogen IX to the FC active site (155). However, experiments with isolated mitochondria have shown complexation between the two enzymes is not required for heme biosynthesis (157). Nevertheless, modeling based on the thre e-dimensional structur e of PPO with FC suggests early work on the topic is likely correct (144). Three X-ray crystal structures for FC have been solved (158-160). FC from humans, the yeast Schizocassharomyces pombe (161) and the Gram-negative oligotroph Caulobacter crescentus (161) contain a [2Fe2S] cluster. Curiously B. subtilis FC is devoid of this prosth etic group, and exists as a monomer (160). The function of these iron-s ulfur centers is largely unknown. Human FC is dimeric, and each monomer is defined by two domains (Figure 1.5) (158). The domains, structurally unrecognizable from each other, are composed of a four-stranded parallel -sheet delimited by an -helix in a motif (a Rossmann-type fold). A gene duplication event has been proposed based on the topological similarities shared between the two domains. A porphyrin binding model has been proposed based on the three-dimensional structure of N-methyl-protoporphyrin bound-ferrochelatase and and metallation kinetics using this inhibitor (160, 162). The substrate-binding cleft is deep within the macromolecule and is inter domain in nature. Metal binding and catalysis likely take place within this region and involve several evolutionarily conserved residues. Sub-cellular localization studi es with mammalian FC show that the active site is positioned near the membrane-associating side of the enzyme (163). This interaction includes formation of an active site access tu nnel, which permits substrate association
28 Figure 1.5. The three-dimensional st ructure of ferrochelatase from Homo sapiens (PBD code: 1hrk). The 2Fe-2S cluster is coordinate d by four cysteine residues. This prosthetic group is located at the s ubunit interface of the functional dimer.
29 and product release; a scenario mediated by the conserved hydr ophobic residues which partially define the site of catalysis. The FC-catalyzed reaction is characterized by two key steps: the binding of the substrate (protoporphyrin IX) and its s ubsequent metallation with ferrous iron (164). Distortion of the tetrapyrrole macrocycle is proposed to occur af ter its binding to the active site cleft of FC (165). This step has been identi fied as a defining feature of ferrochelatase-catalyzed metallation. In fa ct, resonance Raman spectroscopic studies showed that in the absence of metal, murine ferrochelatase is able to induce saddling of the porphyrin substrate (166). Additionally, quantum mechanical calculations of porphyrin binding to B. subtilis ferrochelatase permitted Sigfridsson and Ryde (2003) to describe a tilted pyrrole ring, according to a tetrapyrrole conformation which requires less energy for the insertion of metal (167). In short, most investigators believe that the microenvironment of the active site controls the planarity of the por phyrin, which in turn affects metallation efficiency (165). Chelatases catalyze the inse rtion of a metallic cation into a porphyrin to generate a variety of metallated tetrapyrroles (168). Concerning the role of ferrochelatase in the generation of metallated tetrapyr roles, several theories exis t. Proponents of one theory suggest that a channel is involved in shuttling metal ions from solution to the active site (169, 170). Several conserved residues in bacter ial ferrochelatase appear important in facilitating this process. An active site histidine residue (H183) is required for metal chelation (171), and together with a pair of nearby amino acids (glutamate and serine), define an internal metal-binding site (162, 169). It has been suggested that this inner coordination site is linked to a second solvent-exposed metal binding site (169, 172).
30 This proposed channel is defined by c onserved acidic amino acids comprising a -helix. While the function of this helix remains unc lear, hypotheses regard ing the regulatory nature of this motif have been suggested (169, 172). The mechanism by which metallation takes places requires further rese arch, however a notable theory regarding iron delivery to FC exists. Frataxin, as a monomer is a small protein, localized to the mitochondrion and plays a role in mitochondrial iron detoxification (173). As an oligomer, frataxin can be as large as a trimer to greater than a 48-mer (174). Based on the structures solved for frataxin ol igomer with iron bound, as well as in vitro assays proving protein-protein interactions between this protei n and FC, it is proposed that frataxin delivers ferrous iron to FC (165, 175). Certainly, the cha nneling of metal atoms by way of a chaperone-ferrochelatase comple x has the advantages of minimizing Fenton chemistry between the substrate and the aq ueous environment. Understanding the relationship between iron-delive ry and heme synthesis repres ents an intriguing future direction of ferrochelat ase-related research. Mutations in the FC gene result in a congenital disorder called, erythropoietic protoporphyria (EPP) (176). EPP is transmitted as an auto somal dominant trait; however evidence suggests that a second defective copy is necessary for pathological presentation (177, 178). Accumulation of protoporphyrin is lo calized to reticulocytes, and is a significant component of bile (12). Patients with protoporphyrin-rich bile often show cholestasis, cirrhosis and rema rkably fluorescent gall stones (12). Of all the porphyrias, EPP results in the greatest dermatitic photosen sitivity. Most EPP patients function with FCs that turnover substrate at a rate less than 20% of th at of the wild-type protein (12). While this diminished activity likely stem s from mutations in the protein; a common
31 wild-type FC allelic variant has been id entified and shows a marked reduction in expression (179). This decreased protein availability may be a reason why the disorder is not observed as exclusively dominant in natu re. The overproduction of protoporphyrin is also linked to acute liver damage, high lighting the necrotic effects of excess protoporphyrin, a circumstance noted in the treatment of several cancers (177, 180). Enzymes in the heme degradation pathway Heme oxygenase Heme oxygenase (EC 184.108.40.206; HO) (Figure 1.6) has 3 isoforms (14). The first, HO-1, is a highly inducible 32-kD a protein, that catalyzes the first and rate-limiting step in the degradation of heme from red blo od cells, yielding equimolar quantities of biliverdin IXa, carbon monoxide (CO), and iron (Figure 1.2) (181). Biliverdin (through the action of biliverdin reductase) is converted to bilirubin, and iron is sequestered into ferritin (181). Interestingly, HO-1 utilizes heme as both a prosthetic group and a substrate (182). The second isoform of hemoxygenase, HO-2, a constitutively synthesized 36-kDa protein, is generally unr esponsive to any of the inducers of HO-1 (181). The third isoform, HO-3, also catalyzes heme degradation, but much less so when compared with HO-2 (183). Although heme is the typical HO-1 inducer, others include endotoxin, heavy metals, oxidants, and hypoxia (182). A common feature of several of these inducers is their ability to generate reactive oxygen species, suggesting that HO-1 provides potent cytoprotective effects (21). These products ha ve physiological and pathological functions which include protect ion from oxidative stress, a circumstance linked to: atherosclerosis and cancer, as well as a number of inflammatory, autoimmune,
32 Figure 1.6. The enzymes and intermedia tes of the heme catabolic pathway.
33 and degenerative diseases (20, 184, 185). Further, the heme catabolic pathway is of major importance to the degr adation of the globins and other hemoproteins, many of which are affective stressors of the aforementioned disordered states. Mammalian HO-1 proteins share a high de gree of sequence similarity. Among eucaryotes, a conserved C-te rminal hydrophobic tail of ~20 re sidues appears to function in anchoring HO-1 to the microsomal memb rane. A number of conserved histidine residues are most likely to be important in heme binding (186). Human HO-1 is notably similar to bacterial HO with respect to sequenc e. Several bacterial HOs, including that from Neisseriae meningitides, have been crystallized and require an NADH reductase for enzymatic activity (187). These enzymes function as wa ter-soluble monomers of ~25 kDa. In procaryotes, HOs function to releas e iron to the environment, a process which increases microbial survival and pathogenesis, and mitigates heme toxicity (187). The Xray crystal structure of human HO-1 reveals many structural similarities to the bacterial proteins (188). Human HO exhibits a mostly helical content. Heme is found between two buried -helices (189). An evolutionarily conserved hi stidine in the proximal helix is the axial ligand for the substrate (189). On the opposite side, an -helix stretches over the active site and the heme molecule, by way of a glycine-rich loop, and terminates in a distal polar pocket. The -meso edge of heme is pointed toward the protein interior and is swaddled by a series of conserve d hydrophobic residues. Conformational heterogeneity is observed in the HO-1-heme co mplex and is likely due to flexibility of the distal pocket, a component of the enzyme which is proposed to contribute to the opening and closing of the active site (189).
34 The HO-1 reaction mechanism involves three oxygenation steps (16, 190). In the first step, the -meso heme position is oxi dized by diatomic oxygen (O2) in the presence of NADPH to yield -hydroxyheme. Subsequently, the -hydroxyheme intermediate is further oxidized by O2 to release CO and verdoheme. Finally, verdoheme is oxidized by O2 in the presence of NADPH to produce biliverdin and Fe3+. Among cyanobacteria, algae and plants, HO is ubiquitously expressed and plays a key role in the synthesis of photon-accepting chromophores for use in photosynthesis or light-sensing (191). Specifically both cyanobacteria and algae use the HO catabolic product biliverdin as a precursor for synthesis of phycobilin, th e main photoreceptor for photosynthesis. Transcriptional regulation of the HO-1 gene involves activ ators such as Nrf2 and repressors such as the heme-binding protein Bach-1 (192). Both the activating and repressing factors require heter odimerization with the small Maf proteins including MafK, MafF or MafG, which bind to the Maf recognition elements (MAREs) in HO-1 gene enhancers; a circumstance which a llows modulation of HO-1 gene expression (192). To date, no congenital disorder has been identified associated with mutations in the HO gene. Biliverdin reductase The conversion of biliverdin to bilirubin is controlled by biliverdin reductase (EC 220.127.116.11; BVR) (Figure 1.6 (193, 194), an enzyme that reduces the C10 bridge of biliverdin. BVR is evolutionarily cons erved among all metazoa; however a homolog exists in red algae (195, 196). Protein sequence comparison among diverse phyla shows a high degree of conservation. Among mamma lian species, BVR is greater than 80% identical, a degree of similarity bolster ed by a series of sequence features (197). BVRs
35 contain a leucine zipper motif (bzip), an adenine dinucleotide-binding motif, a serine/threonine kinase domain, two Sr c homology (SH2)-binding domains and a Zn/metal-binding motif (198-200). The reductase activity of BVR requires NADH as a substrate at acidic pH; however, NADP H is utilized in the basic range (201). While the structural basis for the unique cofactor/pH-depend ence activity profile is unclear, site-directed mutagenesis and X-ray crystallography have provided insight into which residues are responsible for much of the reductase activity (202). An N-terminal domain, complete with a Rossman fold, was identified from the three-dimensional structures of BVR with a nd without the cofactor bound (203). In the rat crystal, extensive interactions between the enzyme termini occur by way of a -sheet (203). Several point mutations to residues involved in defining the conserved binding domains of the enzyme (adenine dinucleotide, S/ T kinase domain, oxidoreducatse domain) abolish reductase activity (204). Most of these mutations have nearly the same negative impact on activity with both NADPH and NADH. In contrast, the loss of S44, which results in a 400% increase in only the NAD H-dependant activity, is due to reduced hindrance to NADH binding and NAD release (199). With respect to NADH-derived enzyme activity, the BVR-catalyzed reaction is accelerated in human renal carcinoma (205). The significance and cause of this increase in activity is ambiguous; however, attenuation of this effect woul d be a valid therapeutic target.
36 Content of the dissertation This dissertation focuses on three aspects of the ALAS-catalyzed reaction. First, the interconversion of ALAS between two forms, namely open and closed, is addressed with respect to hydrogen bonding in teractions between the enzyme and the cofactor. Second, substrate specificity rela ted to succinyl-CoA is examined through molecular interactions between two conser ved residues (Arg85 a nd Thr430 in murine ALAS) and the chemical nature of the acylCoA-derived tail. Third, the rate-determining step of the enzyme-catalyzed reaction is addresse d in the context of the conformational mobility of an active site loop. The conclusi ons set forth in each chapter are related to the advancement of knowledge regarding not only the ALAS-cat alyzed reaction, but reactions catalyzed by members of the -oxoamine synthase subfamily and PLPdependent enzymes as a group. References (1) Padmanaban, G., Venkateswar, V., and Rangarajan, P. N. (1989) Haem as a multifunctional regulator. Trends Biochem Sci 14, 492-6. (2) Guengerich, F. P., and MacDonald, T. L. (1990) Mechanisms of cytochrome P450 catalysis. Faseb J 4, 2453-9. (3) Chance, B. (1972) The nature of elec tron transfer and energy coupling reactions. FEBS Lett 23, 3-20. (4) Ryter, S. W., and Choi, A. M. (2005) Heme oxygenase-1: redox regulation of a stress protein in lung and cell culture models. Antioxid Redox Signal 7, 80-91. (5) Shan, Y., Lambrecht, R. W., Ghaziani, T., Donohue, S. E., and Bonkovsky, H. L. (2004) Role of Bach-1 in regulation of heme oxygenase-1 in human liver cells: insights from studies with small interfering RNAS. J Biol Chem 279, 51769-74. (6) Wek, R. C., Jiang, H. Y., and Anthony, T. G. (2006) Coping with stress: eIF2 kinases and translational control. Biochem Soc Trans 34, 7-11. (7) van den Beucken, T., Koritzinsky, M., a nd Wouters, B. G. (2006) Translational control of gene expression during hypoxia. Cancer Biol Ther 5, 749-55. (8) Berlanga, J. J., Herrero, S., and de Har o, C. (1998) Characterization of the heminsensitive eukaryotic initiati on factor 2alpha kinase from mouse nonerythroid cells. J Biol Chem 273, 32340-6. (9) Igarashi, J., Murase, M., Iizuka, A., Pi chierri, F., Martinkova, M., and Shimizu, T. (2008) Elucidation of the heme binding site of heme-regulated eukaryotic
37 initiation factor 2alpha kinase and the role of the regulatory motif in heme sensing by spectroscopic and catalytic st udies of mutant proteins. J Biol Chem 283, 18782-91. (10) Ferreira, G. C. (1999) 5-aminole vulinate synthase and mammalian heme biosynthesis, in Iron Metabolism. (11) Dailey, T. A., Woodruff, J. H., a nd Dailey, H. A. (2005) Examination of mitochondrial protein targeting of haem synthetic enzymes: in vivo identification of three functional haem -responsive motifs in 5-am inolaevulinate synthase. Biochem J 386, 381-6. (12) Badminton, M. N., and Elder, G. H. (2005) Molecular mechanisms of dominant expression in porphyria. J Inherit Metab Dis 28, 277-86. (13) Batts, K. P. (2007) Iron ove rload syndromes and the liver. Mod Pathol 20 Suppl 1, S31-9. (14) Tenhunen, R., Marver, H. S., and Schmi d, R. (1968) The enzymatic conversion of heme to bilirubin by microsomal heme oxygenase. Proc Natl Acad Sci U S A 61, 748-55. (15) May, B. K., Dogra, S. C., Sadlon, T. J., Bhasker, C. R., Cox, T. C., and Bottomley, S. S. (1995) Molecular regulat ion of heme biosynthesis in higher vertebrates. Progress in Nucleic Acid Re search & Molecular Biology 51, 1-51. (16) Noguchi, M., Yoshida, T., and Kikuchi, G. (1982) Identification of the product of heme degradation catalyzed by the heme oxyge nase system as bi liverdin IX alpha by reversed-phase high-perfor mance liquid chromatography. J Biochem 91, 147983. (17) Tuzuner, E., Liu, L., Shimada, M., Yilmaz, E., Glanemann, M., Settmacher, U., Langrehr, J. M., Jonas, S., Neuhaus, P., and Nussler, A. K. (2004) Heme oxygenase-1 protects human hepatocytes in vitro against warm and cold hypoxia. J Hepatol 41, 764-72. (18) Otterbein, L., Sylvester, S. L., a nd Choi, A. M. (1995) Hemoglobin provides protection against lethal endotoxemia in rats: the role of heme oxygenase-1. Am J Respir Cell Mol Biol 13, 595-601. (19) Tomaro, M. L., Frydman, J., and Frydman, R. B. (1991) Heme oxygenase induction by CoCl2, Co-protoporphyrin IX, phenylhydrazine, and diamide: evidence for oxidative stress involvement. Archives of Bioche mistry & Biophysics 286, 610-7. (20) Otterbein, L. E., and Choi, A. M. (2000) Heme oxygenase: colors of defense against cellular stress. Am J Physiol Lung Cell Mol Physiol 279, L1029-37. (21) Abraham, N. G., and Kappas, A. (2005) Heme oxygenase and the cardiovascularrenal system. Free Radic Biol Med 39, 1-25. (22) Willis, D., Moore, A. R., Freder ick, R., and Willoughby, D. A. (1996) Heme oxygenase: a novel target for the modulat ion of the inflammatory response. Nature Medicine 2, 87-90. (23) Akhtar, M., Abboud, M. M., Barnard, G., Jordan, P., and Zaman, Z. (1976) Mechanism and stereochemistry of enzy mic reactions involved in porphyrin biosynthesis. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 273, 117-136.
38 (24) Hansson, M., Rutberg, L., Schroder, I ., and Hederstedt, L. (1991) The Bacillus subtilis hemAXCDBL gene cluster, whic h encodes enzymes of the biosynthetic pathway from glutamate to uroporphyrinogen III. J Bacteriol 173, 2590-9. (25) Li, J. M., Brathwaite, O., Cosloy, S. D., and Russell, C. S. (1989) 5Aminolevulinic acid synthe sis in Escherichia coli. Journal of Bacteriology 171, 2547-52. (26) Weinstein, J. D., and Beale, S. I. (1983) Separate physiological roles and subcellular compartments for two tetr apyrrole biosynthetic pathways in Euglena gracilis. J Biol Chem 258, 6799-807. (27) Hunter, G. A., and Ferreira, G. C. (1999) Pre-steady-state reaction of 5aminolevulinate synthase. Evidence fo r a rate-determining product release. Journal of Biological Chemistry 274, 12222-8. (28) Hunter, G. A., Zhang, J., and Ferreira, G. C. (2007) Transient kinetic studies support refinements to the chemical and kinetic mechanisms of aminolevulinate synthase. J. Biol. Chem. 282, 23025-23035. (29) Bishop, D. F., Henderson, A. S., and Astrin, K. H. (1990) Human deltaaminolevulinate synthase: assignment of the housekeeping gene to 3p21 and the erythroid-specific gene to the X chromosome. Genomics 7, 207-14. (30) Cox, T. C., Bawden, M. J., Martin, A ., and May, B. K. (1991) Human erythroid 5aminolevulinate synthase: promoter an alysis and identification of an ironresponsive element in the mRNA. Embo J 10, 1891-902. (31) Cotter, P. D., Willard, H. F., Gorski, J. L., and Bishop, D. F. (1992) Assignment of human erythroid delta-am inolevulinate synthase (ALAS2) to a distal subregion of band Xp11.21 by PCR analysis of soma tic cell hybrids containing X; autosome translocations. Genomics 13, 211-2. (32) Lathrop, J. T., and Timko, M. P. (1993) Regulation by heme of mitochondrial protein transport through a conserved amino acid motif. Science 259, 522-5. (33) Goodfellow, B. J., Dias, J. S., Fe rreira, G. C., Henklein, P., Wray, V., and Macedo, A. L. (2001) The solution structur e and heme binding of the presequence of murine 5-aminolevulinate synthase. FEBS Lett 505, 325-31. (34) Munakata, H., Sun, J. Y., Yoshida, K., Nakatani, T., Honda, E., Hayakawa, S., Furuyama, K., and Hayashi, N. (2004) Ro le of the heme regulatory motif in the heme-mediated inhibition of mitochondria l import of 5-aminolevulinate synthase. J Biochem 136, 233-8. (35) Handschin, C., Lin, J., Rhee, J., Peyer, A. K., Chin, S., Wu, P. H., Meyer, U. A., and Spiegelman, B. M. (2005) Nutritional regulation of hepatic heme biosynthesis and porphyria through PGC-1alpha. Cell 122, 505-15. (36) Virbasius, J. V., and Scarpulla, R. C. (1994) Activation of the human mitochondrial transcription factor A ge ne by nuclear respiratory factors: a potential regulatory link between nuclear and mitochondrial gene expression in organelle biogenesis. Proc Natl Acad Sci U S A 91, 1309-13. (37) Srivastava, G., Borthwick, I. A., Magui re, D. J., Elferink, C. J., Bawden, M. J., Mercer, J. F., and May, B. K. (1988) Re gulation of 5-aminolevulinate synthase mRNA in different rat tissues. J Biol Chem 263, 5202-9. (38) Rouault, T. A. (2006) The role of iron regulatory proteins in mammalian iron homeostasis and disease. Nat Chem Biol 2, 406-14.
39 (39) Furuyama, K., Kaneko, K., and Vargas P. D. (2007) Heme as a magnificent molecule with multiple missions: heme determines its own fate and governs cellular homeostasis. Tohoku J Exp Med 213, 1-16. (40) Webster, S. P., Alexeev, D., Campop iano, D. J., Watt, R. M., Alexeeva, M., Sawyer, L., and Baxter, R. L. (2000) Mechanism of 8-amino-7-oxononanoate synthase: spectroscopic, kinetic, and crystallogr aphic studies. Biochemistry 39, 516-28. (41) Schmidt, A., Sivaraman, J., Li, Y., Laro cque, R., Barbosa, J. A., Smith, C., Matte, A., Schrag, J. D., and Cygler, M. (2001) Three-dimensional structure of 2-amino3-ketobutyrate CoA ligase from Escherichia coli complexed with a PLP-substrate intermediate: inferred reaction mechanism. Biochemistry 40, 5151-5160. (42) Ikushiro, H., Hayashi, H., and Kaga miyama, H. (2004) Reactions of serine palmitoyltransferase with serine and mo lecular mechanisms of the actions of serine derivatives as inhibitors. Biochemistry 43, 1082-1092. (43) Alexeev, D., Alexeeva, M., Baxter, R. L., Campopiano, D. J., Webster, S. P., and Sawyer, L. (1998) The crystal structur e of 8-amino-7-oxononanoate synthase: a bacterial PLP-dependent, acyl-CoA-condensing enzyme. Journal of Molecular Biology 284, 401-19. (44) Eliot, A. C., and Kirsch, J. F. (2004) Pyridoxal phosphate enzymes: mechanistic, structural, and evolutionary considerations. Annu. Rev. Biochem. 73, 383-415. (45) Bell, S. C., and Turner, J. M. (1976) Bacterial catabolism of threonine. Threonine degradation initiated by L-th reonine-NAD+ oxidoreductase. Biochem. J. 156, 449-458. (46) Eisenberg, M. (1987) Biosynthesis of bi otin and lipoic acid, Vol. 1, p. 544550. American Society of Mi crobiology, Washington D.C. (47) Hanada, K. (2003) Serine palmitoyltr ansferase, a key enzyme of sphingolipid metabolism. Biochim. Biophys. Acta 1632, 16-30. (48) Tan, D., and Ferreira, G. C. (1996) Ac tive site of 5-aminolevulinate synthase resides at the subunit interface. Evidence from in vivo heterodimer formation [published erratum appears in Biochemistry 1997 Apr 15;36(15):4712]. Biochemistry 35, 8934-41. (49) Astner, I., Schulze, J. O., van den Heuvel, J., Jahn, D., Schubert, W. D., and Heinz, D. W. (2005) Crystal structure of 5-aminolevulinate synthase, the first enzyme of heme biosynthesis, and its link to XLSA in humans. EMBO J. 24, 3166-3177. (50) Ferreira, G. C., Vajapey, U., Hafez, O ., Hunter, G. A., and Barber, M. J. (1995) Aminolevulinate synthase: lysine 313 is not essential for binding the pyridoxal phosphate cofactor but is essential for catalysis. Protein Science 4, 1001-6. (51) Gong, J., Hunter, G. A., and Ferre ira, G. C. (1998) Aspartate-279 in aminolevulinate synthase affects enzyme catalysis through en hancing the function of the pyridoxal 5'-phosphate cofactor. Biochemistry 37, 3509-17. (52) Tan, D., Harrison, T., Hunter, G. A., and Ferreira, G. C. (1998) Role of arginine 439 in substrate binding of 5aminolevulinate synthase. Biochemistry 37, 147884.
40 (53) Gong, J., Kay, C. J., Barber, M. J., a nd Ferreira, G. C. (1996) Mutations at a glycine loop in aminolevulinate syntha se affect pyridoxal phosphate cofactor binding and catalysis. Biochemistry 35, 14109-14117. (54) Gong, J., and Ferreira, G. C. (1995) Aminolevulinate synthase: functionally important residues at a glycine loop, a putative pyridoxal phosphate cofactorbinding site. Biochemistry 34, 1678-1685. (55) Turbeville, T. D., Zhang, J., Hunter, G. A., and Ferreira, G. C. (2007) Histidine 282 in 5-aminolevulinate synthase a ffects substrate binding and catalysis. Biochemistry 46, 5972-5981. (56) Shoolingin-Jordan, P. M., LeLean, J. E., and Lloyd, A. J. (1997) Continuous coupled assay for 5-aminolevulinate synthase. Methods Enzymol. 281, 309-316. (57) Zhang, J., and Ferreira, G. C. (2002) Transient state kineti c investigation of 5aminolevulinate synthase reaction mechanism. J. Biol. Chem. 277, 44660-44669. (58) Bottomley, S. S. (1999) Sideroblastic anemia, in Wintrobe's Clinical Hematology (Lee, G. R., Ed.) pp 1022-1045, Lippincott Williams & Wilkins, Baltimore. (59) Bottomley, S. S. (2004) Sideroblastic anemias in Wintrobe's Clinical Hematology (Greer, J. F., J. Lukens, J.N. Rodgers, G.M. Paraskevas, R. Glader, B., Ed.) pp 1012-1033, Lippincott, Williams, & Wilkins, Philadelphia. (60) Kohgo, Y., Ikuta, K., Ohtake, T., To rimoto, Y., and Kato, J. (2008) Body iron metabolism and pathophysiology of iron overload. Int J Hematol 88, 7-15. (61) Bottomley, S. S. (2006) C ongenital sideroblastic anemias. Curr. Hematol. Rep. 5, 41-49. (62) Fleming, M. D. (2002) The genetics of inherited sideroblastic anemias. Semin Hematol 39, 270-81. (63) MacKenzie, E. L., Iwasaki, K., and Ts uji, Y. (2008) Intr acellular iron transport and storage: from molecular mechanisms to health implications. Antioxid Redox Signal 10, 997-1030. (64) Lu, Z., Nie, G., Li, Y., Soe-Lin, S., Tao, Y., Cao, Y., Zhang, Z., Liu, N., Ponka, P., and Zhao, B. (2009) Overexpression of Mitochondrial Ferritin Sensitizes Cells to Oxidative Stress Via an Iron-Mediated Mechanism. Antioxid Redox Signal. (65) Ozment, C. P., and Turi, J. L. (2008) Iron overload following red blood cell transfusion and its impact on disease severity. Biochim Biophys Acta. (66) Cotter, P. D., May, A., Li, L., Al-Saba h, A. I., Fitzsimons, E. J., Cazzola, M., and Bishop, D. F. (1999) Four new mutati ons in the erythroid-specific 5aminolevulinate synthase (ALAS2) gene causing X-linked sideroblastic anemia: increased pyridoxine responsiveness after removal of iron overload by phlebotomy and coinheritance of hereditary hemochromatosis. Blood 93, 175769. (67) Furuyama, K., Fujita, H., Nagai, T ., Yomogida, K., Munakata, H., Kondo, M., Kimura, A., Kuramoto, A., Hayashi, N ., and Yamamoto, M. (1997) Pyridoxine refractory X-linked sideroblastic anem ia caused by a point mutation in the erythroid 5-aminolevul inate synthase gene. Blood 90, 822-30. (68) Furuyama, K., and Sassa, S. (2000) Inte raction between succinyl CoA synthetase and the heme-biosynthetic enzyme ALAS-E is disrupted in sideroblastic anemia. J. Clin. Invest. 105, 757-764.
41 (69) Heinemann, I. U., Jahn, M., and Jah n, D. (2008) The biochemistry of heme biosynthesis. Archives of Biochemistry & Biophysics 474, 238-51. (70) Whatley, S. D., Ducamp, S., Gouya, L., Grandchamp, B., Beaumont, C., Badminton, M. N., Elder, G. H., Holme, S. A., Anstey, A. V., Parker, M., Corrigall, A. V., Meissner, P. N., Hift R. J., Marsden, J. T., Ma, Y., MieliVergani, G., Deybach, J. C., and Puy, H. (2008) C-terminal deletions in the ALAS2 gene lead to gain of f unction and cause X-linked dominant protoporphyria without anemia or iron overload. Am J Hum Genet 83, 408-14. (71) Cotter, P. D., Rucknagel, D. L., a nd Bishop, D. F. (1994) X-linked sideroblastic anemia: identification of the mutation in the erythroid-specific deltaaminolevulinate synthase gene (ALAS2) in the original family described by Cooley. Blood 84, 3915-24. (72) Jaffe, E. K. (1995) Porphobilinogen synthase, the first source of heme's asymmetry. Journal of Bioenergetics & Biomembranes 27, 169-79. (73) Frankenberg, N., Moser, J., and Jahn, D. (2003) Bacterial heme biosynthesis and its biotechnological application. Appl Microbiol Biotechnol 63, 115-27. (74) Jordan, P. M. (1994) High lights in haem biosynthesis. Curr Opin Struct Biol 4, 902-11. (75) Breinig, S., Kervinen, J., Stith, L., Wasson, A. S., Fairman, R., Wlodawer, A., Zdanov, A., and Jaffe, E. K. (2003) Cont rol of tetrapyrrole biosynthesis by alternate quaternary forms of porphobilinogen synthase. Nat Struct Biol 10, 75763. (76) Tang, L., Breinig, S., Stith, L., Mischel, A., Tannir, J., Kokona, B., Fairman, R., and Jaffe, E. K. (2006) Single amino aci d mutations alter th e distribution of human porphobilinogen synthase quaternary structure isoforms (morpheeins). J Biol Chem 281, 6682-90. (77) Jaffe, E. K. (2005) Morpheeins--a ne w structural paradi gm for allosteric regulation. Trends Biochem Sci 30, 490-7. (78) Frere, F., Nentwich, M., Gacond, S., He inz, D. W., Neier, R., and FrankenbergDinkel, N. (2006) Probing the active site of Pseudomonas aeruginosa porphobilinogen synthase using newly developed inhibitors. Biochemistry 45, 8243-53. (79) Shemin, D. (1976) Proceedings: Stru cture, function and mechanism of deltaaminolevulinic acid dehydratase. J Biochem 79, 37P-38P. (80) Erskine, P. T., Newbold, R., Brindle y, A. A., Wood, S. P., Shoolingin-Jordan, P. M., Warren, M. J., and Cooper, J. B. (2001) The x-ray structure of yeast 5aminolaevulinic acid dehydratase complexe d with substrate and three inhibitors. Journal of Molecular Biology 312, 133-41. (81) Warren, M. J., Cooper, J. B., Wood, S. P., and Shoolingin-Jordan, P. M. (1998) Lead poisoning, haem synthesis and 5-aminolaevulinic acid dehydratase. Trends in Biochemical Sciences 23, 217-21. (82) Akagi, R., Shimizu, R., Furuyama, K ., Doss, M. O., and Sassa, S. (2000) Novel molecular defects of the delta-aminolevulin ate dehydratase gene in a patient with inherited acute hepatic porphyria. Hepatology 31, 704-8. (83) Fujita, H., Nishitani, C., and Ogawa, K. (2002) Lead, chemical porphyria, and heme as a biological mediator. Tohoku J Exp Med 196, 53-64.
42 (84) Bergdahl, I. A., Grubb, A., Schutz, A., Desnick, R. J., Wetmur, J. G., Sassa, S., and Skerfving, S. (1997) Lead binding to delta-aminolevulinic acid dehydratase (ALAD) in human erythrocytes. Pharmacol Toxicol 81, 153-8. (85) Jaffe, E. K., and Stith, L. (2007) AL AD porphyria is a conformational disease. Am J Hum Genet 80, 329-37. (86) Harper, P., and Wahlin, S. (2007) Treat ment options in acute porphyria, porphyria cutanea tarda, and erythropoietic protoporphyria. Curr Treat Options Gastroenterol 10, 444-55. (87) Mitchell, G., Larochelle, J., Lambert, M., Michaud, J., Grenier, A., Ogier, H., Gauthier, M., Lacroix, J., Vanasse, M., and Larbrisseau, A. (1990) Neurologic crises in hereditary tyrosinemia. New England Journal of Medicine 322, 432-7. (88) Jordan, P. M., Thomas, S. D., and Warren, M. J. (1988) Purification, crystallization and properties of porphob ilinogen deaminase from a recombinant strain of Escherichia coli K12. Biochem J 254, 427-35. (89) Jordan, P. M., Warren, M. J., Williams, H. J., Stolowich, N. J., Roessner, C. A., Grant, S. K., and Scott, A. I. (1988) Id entification of a cysteine residue as the binding site for the dipyrromethane cofactor at the active site of Escherichia coli porphobilinogen deaminase. FEBS Letters 235, 189-93. (90) Gill, R., Kolstoe, S. E., Mohammed, F., Al, D. B. A., Cooper, J. B., Wood, S. P., and Shoolingin-Jordan, P. M. (2009) The structure of human porphobilinogen deaminase at 2.8 A: the molecular ba sis of acute intermittent porphyria. Biochem J. (91) Warren, M. J., and Jordan, P. M. (1988) Investigation into the nature of substrate binding to the dipyrromethane cofactor of Escherichia coli porphobilinogen deaminase. Biochemistry 27, 9020-30. (92) Gregor, A., Schneider-Yin, X., Szle ndak, U., Wettstein, A., Lipniacka, A., Rufenacht, U.B. and Minder, E.I. (2002) Molecular study of the hydroxymethylbilane synthase gene (HMBS) among polish patients with acute intermittent porphyria. Hum. Mutat. 19, 310-320. (93) Yrjonen, A., Pischik, E., Mehtala, S., and Kauppi nen, R. (2008) A novel 19-bp deletion of exon 15 in the HMBS gene causing acute interm ittent porphyria associating with rhabdomyolys is during an acute attack. Clin Genet 74, 396-8. (94) Song, G., Li, Y., Cheng, C., Zhao, Y., Gao, A., Zhang, R., Joachimiak, A., Shaw, N., and Liu, Z. J. (2009) Structural insight into acute intermittent porphyria. Faseb J 23, 396-404. (95) Solis, C., Martinez-Bermejo, A., Naidic h, T. P., Kaufmann, W. E., Astrin, K. H., Bishop, D. F., and Desnick, R. J. (2004) Acute intermittent porphyria: studies of the severe homozygous dominant disease provides insights into the neurologic attacks in acute porphyrias. Arch Neurol 61, 1764-70. (96) Brownlie, P. D., Lambert, R., Louie, G. V., Jordan, P. M., Blundell, T. L., Warren, M. J., Cooper, J. B., and Wood, S. P. (1994) The three-dimensional structures of mutants of porphobilinogen deaminase: toward an understanding of the structural basis of acute intermittent porphyria. Protein Sci 3, 1644-50. (97) Tsai, S. F., Bishop, D. F., and De snick, R. J. (1988) Human uroporphyrinogen III synthase: molecular cloning, nucleotide se quence, and expression of a full-length cDNA. Proc Natl Acad Sci U S A 85, 7049-53.
43 (98) Kohashi, M., Clement, R. P., Tse, J., and Piper, W. N. (1984) Rat hepatic uroporphyrinogen III co-synthase. Purifica tion and evidence for a bound folate coenzyme participating in the bios ynthesis of uroporphyrinogen III. Biochem J 220, 755-65. (99) Hart, G. J., and Battersby, A. R. (1985) Purification and properties of uroporphyrinogen III synthase (co-synt hetase) from Euglena gracilis. Biochem J 232, 151-60. (100) Alwan, A. F., Mgbeje, B. I., and Jord an, P. M. (1989) Purification and properties of uroporphyrinogen III synthase (c o-synthase) from an overproducing recombinant strain of Escherichia coli K-12. Biochem J 264, 397-402. (101) O'Brian, M. R., and Thony-Meyer, L. (2002) Biochemistry, regulation and genomics of haem biosynthesis in prokaryotes. Adv Microb Physiol 46, 257-318. (102) Schubert, H. L., Phillips, J. D., Heroux, A., and Hill, C. P. (2008) Structure and mechanistic implications of a uroporphyrinogen III synthase-product complex. Biochemistry 47, 8648-55. (103) Mathews, A. M., Schubert, H.L., Whit by, F.G., Alexander, K.J., Schadick, K., Bergonia, H.A., Philips, J.D. and Hill, C.P. (2001) Crystal structure of human uroporphyrinogen III synthase. The EMBO Journal 20, 5832-5839. (104) Silva, P. J., and Ramos, M. J. ( 2008) Comparative densit y functional study of models for the reaction mechanis m of uroporphyrinogen III synthase. J Phys Chem B 112, 3144-8. (105) Bishop, D. F., Johansson, A., Phelps, R., Shady, A. A., Ramirez, M. C., Yasuda, M., Caro, A., and Desnick, R. J. (2006) Uroporphyrinogen III synthase knock-in mice have the human congenital erythropoi etic porphyria phenotype, including the characteristic light-indu ced cutaneous lesions. Am J Hum Genet 78, 645-58. (106) Fortian, A., Castano, D., Ortega, G., Lain, A., Pons, M., and Millet, O. (2009) Uroporphyrinogen III synthase mutations re lated to congenital erythropoietic porphyria identify a key helix for protein stability. Biochemistry 48, 454-61. (107) Phillips, J. D., Steensma, D. P., Pulsi pher, M. A., Spangrude, G. J., and Kushner, J. P. (2007) Congenital er ythropoietic porphyria due to a mutation in GATA1: the first trans-acting mutation causative for a human porphyria. Blood 109, 2618-21. (108) Mauzerall, D. a. G., S. (1958) Po rphyrin biosynthesis in erythrocytes. Uroporphyrinogen and its decarboxylase. J. Biol. Chem. 232, 1141-1162. (109) Martins, B. M., Grimm, B., Moc k, H. P., Richter, G., Huber, R., and Messerschmidt, A. (2001) Tobacco uroporphyrinogen-III decarboxylase: characterization, crystallization and preliminary X-ray analysis. Acta Crystallogr D Biol Crystallogr 57, 1709-11. (110) Fan, J., Liu, Q., Hao, Q., Teng, M., a nd Niu, L. (2007) Crystal structure of uroporphyrinogen decarboxylase from Bacillus subtilis. J Bacteriol 189, 3573-80. (111) Silva, P. J., and Ramos, M. J. (2005) Density-functional stud y of mechanisms for the cofactor-free decarboxylation performed by uroporphyrinogen III decarboxylase. J Phys Chem B 109, 18195-200. (112) Kawanishi, S., Seki, Y., and Sano, S. (1983) Uroporphyrinogen decarboxylase. Purification, properties, and inhibition by polychlorinated biphenyl isomers. J Biol Chem 258, 4285-92.
44 (113) Felix, F., and Brouillet, N. (1990) Pu rification and properties of uroporphyrinogen decarboxylase from Saccharomyces cerevisiae. Yeast uroporphyrinogen decarboxylase. Eur J Biochem 188, 393-403. (114) Whitby, F. G., Phillips, J. D., Kushner, J. P., and Hill, C. P. (1998) Crystal structure of human ur oporphyrinogen decarboxylase. Embo J 17, 2463-71. (115) Martins, B. M., Grimm, B., Mock, H. P., Huber, R., and Messerschmidt, A. (2001) Crystal structure and substrat e binding modeling of the uroporphyrinogenIII decarboxylase from Nicotiana tabacum. Implications for the catalytic mechanism. J Biol Chem 276, 44108-16. (116) Luo, J., and Lim, C. K. (1993) Or der of uroporphyrinogen III decarboxylation on incubation of porphobilinogen and uroporphyrinogen III with erythrocyte uroporphyrinogen decarboxylase. Biochem J 289 ( Pt 2), 529-32. (117) Phillips, J. D., Whitby, F. G., Kushner, J. P., and Hill, C. P. (2003) Structural basis for tetrapyrrole coordina tion by uroporphyrinogen decarboxylase. Embo J 22, 6225-33. (118) Lewis, C. A., Jr., and Wolfenden, R. (2008) Uroporphyrinogen decarboxylation as a benchmark for the catalytic proficiency of enzymes. Proc Natl Acad Sci U S A 105, 17328-33. (119) Elder, G. H. (1998) Porphyria cutanea tarda. Semin Liver Dis 18, 67-75. (120) Remenyik, E., Lecha, M., Badenas, C., Koszo, F., Vass, V., Herrero, C., Varga, V., Emri, G., Balogh, A., and Horkay, I. (2008) Childhood-onset mild cutaneous porphyria with compound heterozygotic mutations in the uroporphyrinogen decarboxylase gene. Clin Exp Dermatol 33, 602-5. (121) Elder, G. H., and Roberts, A. G. (1995) Uroporphyrinogen decarboxylase. Journal of Bioenergetics & Biomembranes 27, 207-14. (122) Phillips, J. D., Whitby, F. G., Stadtmueller, B. M., Edwards, C. Q., Hill, C. P., and Kushner, J. P. (2007) Two novel uroporphyrinogen decarboxylase (URO-D) mutations causing hepatoeryt hropoietic porphyria (HEP). Transl Res 149, 85-91. (123) Elder, G. H., and Evans, J. O. (1978) Evidence that th e coproporphyrinogen oxidase activity of rat liver is s ituated in the intermembrane space of mitochondria. Biochem J 172, 345-7. (124) Grandchamp, B., Phung, N., and Nordmann, Y. (1978) The mitochondrial localization of coproporphyrinogen III oxidase. Biochem J 176, 97-102. (125) Delfau-Larue, M. H., Martas ek, P., and Grandchamp, B. (1994) Coproporphyrinogen oxidase: gene organi zation and description of a mutation leading to exon 6 skipping. Hum Mol Genet 3, 1325-30. (126) Susa, S., Daimon, M., Ono, H., Li, S., Yoshida, T., and Kato, T. (2002) Heme inhibits the mitochondrial impor t of coproporphyrinogen oxidase. Blood 100, 4678-9. (127) Panek, H., and O'Brian, M. R. (2002) A whole genome view of prokaryotic haem biosynthesis. Microbiology 148, 2273-82. (128) Lee, D. S., Flachsova, E., Bodnarova, M., Demeler, B., Martasek, P., and Raman, C. S. (2005) Structural basis of hereditary coproporphyria. Proc Natl Acad Sci U S A 102, 14232-7.
45 (129) Layer, G., Moser, J., Heinz, D. W ., Jahn, D., and Schubert, W. D. (2003) Crystal structure of coproporphyrinogen III oxidase reveals cofactor geometry of Radical SAM enzymes. Embo J 22, 6214-24. (130) Layer, G., Verfurth, K., Mahlitz, E., and Jahn, D. (2002) Oxygen-independent coproporphyrinogen-III oxi dase HemN from Escherichia coli. J Biol Chem 277, 34136-42. (131) Sofia, H. J., Chen, G., Hetzler, B. G ., Reyes-Spindola, J. F., and Miller, N. E. (2001) Radical SAM, a novel protein supe rfamily linking unresolved steps in familiar biosynthetic pathways with radical mechanisms: functional characterization using new analysis a nd information visualization methods. Nucleic Acids Res 29, 1097-106. (132) Gitter, S. J., Cooper, C. L., Friesen, J. A., and Jones, M. A. (2007) Investigation of the catalytic and structural role s of conserved histidines of human coproporphyrinogen oxidase using site-directed mutagenesis. Med Sci Monit 13, BR1-10. (133) Morgenthaler, J. B., Barto, R. L., Lash, T. D., and Jones, M. A. (2008) Use of diand tripropionate substrate analogs to pr obe the active site of human recombinant coproporphyrinogen oxidase. Med Sci Monit 14, BR1-7. (134) Berger, H., and Goldberg, A. (1955) Hereditary coproporphyria. Br Med J 2, 858. (135) Martasek, P., Camadro, J. M., Raman, C. S., Lecomte, M. C., Le Caer, J. P., Demeler, B., Grandchamp, B., and La bbe, P. (1997) Human coproporphyrinogen oxidase. Biochemical characterizatio n of recombinant normal and R231W mutated enzymes expressed in E. coli as soluble, catalytically active homodimers. Cell Mol Biol (Noisy-le-grand) 43, 47-58. (136) Stephenson, J. R., Stacey, J. A., Morgenth aler, J. B., Friesen, J. A., Lash, T. D., and Jones, M. A. (2007) Role of aspart ate 400, arginine 262, and arginine 401 in the catalytic mechanism of human coproporphyrinogen oxidase. Protein Sci 16, 401-10. (137) Martasek, P. (1998) Hereditary coproporphyria. Semin Liver Dis 18, 25-32. (138) Nordmann, Y., Grandchamp, B., de Verneuil, H., Phung, L., Cartigny, B., and Fontaine, G. (1983) Harderoporphyria: a variant hereditary coproporphyria. J Clin Invest 72, 1139-49. (139) Brenner, D. A., and Bloomer, J. R. (1980) The enzymatic defect in variegate prophyria. Studies with human cultured skin fibroblasts. N Engl J Med 302, 7659. (140) Dailey, T. A., and Dailey, H. A. ( 1998) Identification of an FAD superfamily containing protoporphyrinogen oxidases, monoamine oxidases, and phytoene desaturase. Expression and characteri zation of phytoene desaturase of Myxococcus xanthus. J Biol Chem 273, 13658-62. (141) Bellamacina, C. R. (1996) The ni cotinamide dinucleotide binding motif: a comparison of nucleotide binding proteins. Faseb J 10, 1257-69. (142) Dailey, T. A., and Dailey, H. A. (1996) Human protoporphyrinogen oxidase: expression, purification, and charac terization of the cloned enzyme. Protein Sci 5, 98-105.
46 (143) Hansson, M., and Hederstedt, L. (1994) Bacillus subtilis HemY is a peripheral membrane protein essential for prot oheme IX synthesis which can oxidize coproporphyrinogen III a nd protoporphyrinogen IX. J Bacteriol 176, 5962-70. (144) Koch, M., Breithaupt, C., Kiefersa uer, R., Freigang, J., Huber, R., and Messerschmidt, A. (2004) Crystal struct ure of protoporphyrinogen IX oxidase: a key enzyme in haem and chlorophyll biosynthesis. Embo J 23, 1720-8. (145) Heinemann, I. U., Diekmann, N., Mas oumi, A., Koch, M., Messerschmidt, A., Jahn, M., and Jahn, D. (2007) Functional definition of the tobacco protoporphyrinogen IX oxidase substrate-binding site. Biochem J 402, 575-80. (146) Corradi, H. R., Corrigall, A. V., Bo ix, E., Mohan, C. G., Sturrock, E. D., Meissner, P. N., and Acharya, K. R. (2006) Crystal structure of protoporphyrinogen oxidase from Myxococcus xanthus and its complex with the inhibitor acifluorfen. J Biol Chem 281, 38625-33. (147) Baxter, D. L., Permowicz, S. E., and Fleischmajer, R. (1967) Variegate porphyria (mixed porphyria). Arch Dermatol 96, 98-100. (148) Groenewald, J. Z., Liebenberg, J., Gr oenewald, I. M., and Warnich, L. (1998) Linkage disequilibrium analysis in a recently founded population: evaluation of the variegate porphyria founder in South African Afrikaners. Am J Hum Genet 62, 1254-8. (149) Meissner, P. N., Dailey, T. A., Hift, R. J., Ziman, M., Corrigall, A. V., Roberts, A. G., Meissner, D. M., Kirsch, R. E., and Dailey, H. A. (1996) A R59W mutation in human protoporphyrinogen oxi dase results in decreased enzyme activity and is prevalent in South Africans with variegate porphyria. Nat Genet 13, 95-7. (150) Rossetti, M. V., Granata, B. X., Giudi ce, J., Parera, V. E., and Batlle, A. (2008) Genetic and biochemical studies in Argent inean patients with variegate porphyria. BMC Med Genet 9, 54. (151) Dooley, K. A., Fraenkel, P. G., La nger, N. B., Schmid, B., Davidson, A. J., Weber, G., Chiang, K., Foott, H., Dwyer, C., Wingert, R. A., Zhou, Y., Paw, B. H., and Zon, L. I. (2008) montalcino, A zeb rafish model for variegate porphyria. Exp Hematol 36, 1132-42. (152) Da Silva, V., Simonin, S., Deybac h, J. C., Puy, H., and Nordmann, Y. (1995) Variegate porphyria: diagnostic value of fluorometric scanning of plasma porphyrins. Clin Chim Acta 238, 163-8. (153) Enriquez de Salamanca, R., Sepulveda, P., Moran, M. J., Santos, J. L., Fontanellas, A., and Hernandez, A. ( 1993) Clinical utility of fluorometric scanning of plasma porphyrins for the diagnosis and typing of porphyrias. Clin Exp Dermatol 18, 128-30. (154) Norris, P. G., Elder, G. H., and Hawk, J. L. (1990) Homozygous variegate porphyria: a case report. Br J Dermatol 122, 253-7. (155) Ferreira, G. C., Franco, R., Lloyd, S. G ., Moura, I., Moura, J. J., and Huynh, B. H. (1995) Structure and function of ferrochelatase. Journal of Bioenergetics & Biomembranes 27, 221-9. (156) Porra, R. J., and Jones, O. T. ( 1963) Studies on ferrochelatase. 2. An in vestigation of the role offerrochelatas e in the biosynthesis of various haem prosthetic groups. Biochem J 87, 186-92.
47 (157) Proulx, K. L., Woodard, S. I., and Dailey, H. A. (1993) In situ conversion of coproporphyrinogen to heme by murine mito chondria: terminal st eps of the heme biosynthetic pathway. Protein Sci 2, 1092-8. (158) Wu, C. K., Dailey, H. A., Rose, J. P., Burden, A., Sellers, V. M., and Wang, B. C. (2001) The 2.0 A structure of human ferrochel atase, the terminal enzyme of heme biosynthesis. Nat Struct Biol 8, 156-60. (159) Schubert, H. L., Raux, E., Brindley, A. A., Leech, H. K., Wilson, K. S., Hill, C. P., and Warren, M. J. (2 002) The structure of Saccharomyces cerevisiae Met8p, a bifunctional dehydrogenase and ferrochelatase. Embo J 21, 2068-75. (160) Al-Karadaghi, S., Hansson, M., Ni konov, S., Jonsson, B., and Hederstedt, L. (1997) Crystal structure of ferrochelat ase: the terminal enzyme in heme biosynthesis. Structure 5, 1501-10. (161) Dailey, T. A. a. D., H.A. (2002) Identi fication of [2Fe-2S] clusters in microbial ferrochelatases. J. Bacteriology 184, 2460-2464. (162) Shipovskov, S., Karlberg, T., Fodje, M., Hansson, M. D., Ferreira, G. C., Hansson, M., Reimann, C. T., and Al-Kar adaghi, S. (2005) Metallation of the transition-state inhibitor N-methyl mes oporphyrin by ferrochelatase: implications for the catalytic r eaction mechanism. J Mol Biol 352, 1081-90. (163) Harbin, B. M., and Dailey, H. A. ( 1985) Orientation of fe rrochelatase in bovine liver mitochondria. Biochemistry 24, 366-70. (164) Ferreira, G. C. (1999) Ferrochelatase. International Journal of Biochemistry & Cell Biology 31, 995-1000. (165) Al-Karadaghi, S., Franco, R., Hansson, M ., Shelnutt, J. A., Isaya, G., and Ferreira, G. C. (2006) Chelatases : distort to select? Trends Biochem Sci 31, 135-42. (166) Franco, R., Ma, J. G., Lu, Y., Ferreira, G. C., and Shelnutt, J. A. (2000) Porphyrin interactions with wild-type a nd mutant mouse ferrochelatase. Biochemistry 39, 2517-29. (167) Sigfridsson, E., and Ryde, U. (2003) The importance of porphyrin distortions for the ferrochelatase reaction. J Biol Inorg Chem 8, 273-82. (168) Kappas, A., and Drummond, G. S. ( 1986) Control of heme metabolism with synthetic metalloporphyrins. J Clin Invest 77, 335-9. (169) Lecerof, D., Fodje, M. N., Alvarez Leon, R., Olsson, U., Hansson, A., Sigfridsson, E., Ryde, U., Hansson, M ., and Al-Karadaghi, S. (2003) Metal binding to Bacillus subtilis ferrochelatas e and interaction between metal sites. J Biol Inorg Chem 8, 452-8. (170) Sellers, V. M., Wu, C. K., Dailey, T. A., and Dailey, H. A. (2001) Human ferrochelatase: characterization of subs trate-iron binding and proton-abstracting residues. Biochemistry 40, 9821-7. (171) Kohno, H., Okuda, M., Furukawa, T., Tokuna ga, R., and Taketani, S. (1994) Sitedirected mutagenesis of hu man ferrochelatase: identific ation of histidine-263 as a binding site for metal ions. Biochim Biophys Acta 1209, 95-100. (172) Karlberg, T., Lecerof, D., Gora, M., Silvegren, G., Labbe-Bois, R., Hansson, M., and Al-Karadaghi, S. (2002) Metal binding to Saccharomyces cerevisiae ferrochelatase. Biochemistry 41, 13499-506. (173) Babcock, M., de Silva, D., Oaks R., Davis-Kaplan, S., Jiralerspong, S., Montermini, L., Pandolfo, M., and Kaplan, J. (1997) Regulation of mitochondrial
48 iron accumulation by Yfh1p, a put ative homolog of frataxin. Science 276, 170912. (174) Adamec, J., Rusnak, F., Owen, W. G., Naylor, S., Benson, L. M., Gacy, A. M., and Isaya, G. (2000) Iron-dependent self -assembly of recombinant yeast frataxin: implications for Friedreich ataxia. Am J Hum Genet 67, 549-62. (175) Bencze, K. Z., Yoon, T., Millan-Pachec o, C., Bradley, P. B., Pastor, N., Cowan, J. A., and Stemmler, T. L. (2007) Human frataxin: iron and ferrochelatase binding surface. Chem Commun (Camb), 1798-800. (176) Magnus, I. A., Jarrett, A., Pra nkerd, T. A., and Rimington, C. (1961) Erythropoietic protoporphyria A new porphyria syndrome w ith solar urticaria due to protoporphyrinaemia. Lancet 2, 448-51. (177) Todd, D. J. (1994) Er ythropoietic protoporphyria. Br J Dermatol 131, 751-66. (178) Li, C., Di Pierro, E., Brancaleoni, V ., Cappellini, M. D., and Steensma, D. P. (2009) A novel large deletion and thr ee polymorphisms in the FECH gene associated with erythr opoietic protoporphyria. Clin Chem Lab Med 47, 44-6. (179) Gouya, L., Puy, H., Lamoril, J., Da Silva, V., Grandchamp, B., Nordmann, Y., and Deybach, J. C. (1999) Inheritance in erythropoietic protoporphyria: a common wild-type ferrochelatase allelic variant with low ex pression accounts for clinical manifestation. Blood 93, 2105-10. (180) Krammer, B., and Plaetzer, K. (2008) ALA and its clinical impact, from bench to bedside. Photochem Photobiol Sci 7, 283-9. (181) Ortiz de Montellano, P. R. ( 2000) The mechanism of hemeoxygenase. Curr. Opin. Chem. Biol. 4, 221-226. (182) Ponka, P. (1999) Cell biology of heme. Am J Med Sci 318, 241-56. (183) Hayashi, S., Omata, Y., Sakamoto, H ., Higashimoto, Y., Hara, T., Sagara, Y., and Noguchi, M. (2004) Characterization of rat heme oxygenase-3 gene. Implication of processed pseudogenes derived from heme oxygenase-2 gene. Gene 336, 24150. (184) Doi, K., Akaike, T., Fujii, S., Tana ka, S., Ikebe, N., Beppu, T., Shibahara, S., Ogawa, M., and Maeda, H. (1999) Induc tion of haem oxygenase-1 nitric oxide and ischaemia in experimental solid tum ours and implications for tumour growth. Br J Cancer 80, 1945-54. (185) Keyse, S. M., and Tyrrell, R. M. (1 989) Heme oxygenase is the major 32-kDa stress protein induced in human skin fibroblasts by UVA radiation, hydrogen peroxide, and sodium arsenite. Proc Natl Acad Sci U S A 86, 99-103. (186) Bianchetti, C. M., Yi, L., Ragsdale S. W., and Phillips, G. N., Jr. (2007) Comparison of apoand heme-bound crys tal structures of a truncated human heme oxygenase-2. J Biol Chem 282, 37624-31. (187) Schuller, D. J., Zhu, W., Stojiljkovic, I., Wilks, A., and Poulos, T. L. (2001) Crystal structure of heme oxygenase from the gram-negative pathogen Neisseria meningitidis and a comparison with mammalian heme oxygenase-1. Biochemistry 40, 11552-8. (188) Schuller, D. J., Wilks, A., Ortiz de Montellano, P. R., and Poulos, T. L. (1999) Crystal structure of human heme oxygenase-1. Nat Struct Biol 6, 860-7.
49 (189) Schuller, D. J., Wilks, A., Ortiz de Montellano, P. R., and Poulos, T. L. (1999) Crystal structure of human heme oxygenase-1. [see comments]. Nature Structural Biology 6, 860-7. (190) Garcia-Serres, R., Davydov, R. M., Mats ui, T., Ikeda-Saito, M., Hoffman, B. M., and Huynh, B. H. (2007) Distinct reacti on pathways followed upon reduction of oxy-heme oxygenase and oxy-myoglobi n as characterized by Mossbauer spectroscopy. J Am Chem Soc 129, 1402-12. (191) Rhie, G., and Beale, S. I. (1994) Regulation of heme oxygenase activity in Cyanidium caldarium by light, glucose, and phycobilin precursors. J Biol Chem 269, 9620-6. (192) Reichard, J. F., Motz, G. T., and P uga, A. (2007) Heme oxygenase-1 induction by NRF2 requires inactivation of the transcriptional repressor BACH1. Nucleic Acids Res 35, 7074-86. (193) Maines, M. D. (1997) The heme oxy genase system: a regulator of second messenger gases. Annu Rev Pharmacol Toxicol 37, 517-54. (194) Tenhunen, R., Ross, M. E., Marver, H. S., and Schmid, R. (1970) Reduced nicotinamide-adenine dinucleotide phos phate dependent biliverdin reductase: partial purification and characterization. Biochemistry 9, 298-303. (195) Schluchter, W. M., a nd Glazer, A. N. (1997) Characterization of cyanobacterial biliverdin reductase. Conversion of biliverdi n to bilirubin is important for normal phycobiliprotein biosynthesis. Journal of Biological Chemistry 272, 13562-9. (196) Beale, S. I., and Cornejo, J. (1984) Enzymatic heme oxygenase activity in soluble extracts of the unicellular red alga, Cyanidium caldarium. Archives of Biochemistry & Biophysics 235, 371-84. (197) Fakhrai, H., and Maines, M. D. (1992) Expression and characterization of a cDNA for rat kidney biliverdin reducta se. Evidence suggest ing the liver and kidney enzymes are the same transcript product. J Biol Chem 267, 4023-9. (198) Lerner-Marmarosh, N., Miralem, T., Gibbs, P. E., and Maines, M. D. (2007) Regulation of TNF-alpha-activated PK C-zeta signaling by the human biliverdin reductase: identification of activating a nd inhibitory domains of the reductase. Faseb J 21, 3949-62. (199) Lerner-Marmarosh, N., Shen, J., Torno, M. D., Kravets, A., Hu, Z., and Maines, M. D. (2005) Human biliverdin reductase : a member of the insulin receptor substrate family with serine/th reonine/tyrosine kinase activity. Proc Natl Acad Sci U S A 102, 7109-14. (200) Ahmad, Z., Salim, M., and Maines, M. D. (2002) Human biliverdin reductase is a leucine zipper-like DNA-binding protein and functions in transcriptional activation of heme oxygenase-1 by oxidative stress. J Biol Chem 277, 9226-32. (201) Kutty, R. K., and Maines, M. D. ( 1981) Purification and characterization of biliverdin reductase from rat liver. Journal of Biological Chemistry 256, 3956-62. (202) McCoubrey, W. K., Jr., and Maines, M. D. (1994) Site-directed mutagenesis of cysteine residues in biliverdin reductase. Roles in substrate and cofactor binding. Eur J Biochem 222, 597-603. (203) Kikuchi, A., Park, S. Y., Miyatake, H ., Sun, D., Sato, M., Yoshida, T., and Shiro, Y. (2001) Crystal structure of rat biliverdin reductase. Nat Struct Biol 8, 221-5.
50 (204) Hunter, T., and Cooper, J. A. (1985) Protein-tyrosine kinases. Annual Review of Biochemistry 54, 897-930. (205) Maines, M. D., Mayer, R. D., Ertur k, E., Huang, T. J., and Disantagnese, A. (1999) The oxidoreductase, biliverdin re ductase, is induced in human renal carcinoma--pH and cofactor-spe cific increase in activity. J Urol 162, 1467-72.
51 Chapter Two Serine-254 enhances an induced fit m echanism in murine 5-aminolevulinate synthase Abstract 5-Aminolevulinate synthase (EC 18.104.22.168) (ALAS), is a homodimeric pyridoxal 5'-phosphate (PLP)-dependent enzyme and cat alyzes the initial step of the heme biosynthetic pathway in animals, fungi, and some bacteria. This reaction involves the condensation of glycine and succinyl-Coenz yme A to produce 5-aminolevulinate (ALA), Coenzyme A (CoA) and carbon dioxide. The X-ray crystal structures of Rhodobacter capsulatus ALAS reveal a conserved active site serine that moves to within hydrogen bonding distance of the phenolic oxygen of the PLP cofactor in the closed, substratebound enzyme conformation, and simultaneously to within 3-4 angstroms of the thioester sulfur atom of bound succinyl-CoA. To evalua te the potential roles of this residue in enzymatic activity, the equivalent serine in murine erythroid ALAS was mutated to alanine or threonine. The S254A variant is active, but both the SCoA mK and kcat values are increased, by 25and 2-fold, respectively, suggesting unusual functi onal complexity. In contrast, the S254T mutation resu lts in a significant decrease in kcat without alteringSCoA mK. Circular dichroism spectroscopy reve als that removal of the side chain hydroxyl group in the S254A vari ant dramatically alters the PLP microenvironment as
52 well as the responsiveness of this microe nvironment to succinyl-CoA binding. Protein fluorescence stopped-flow experiments confirm th at the mutations differentially alter the rates of conformational responsiveness to AL A binding. Taken together the data support the postulate that this serine residue is important for formation of a competent catalytic complex by coupling succinyl-CoA binding to enzyme conformational equilibria. Similar functions of this residue may be postulated for the other -oxoamine synthases.
53 Introduction 5-Aminolevulinate synthase (EC 22.214.171.124; ALAS) is a homodimeric PLPdependent enzyme that catalyzes the firs t and key regulatory enzyme of the heme biosynthetic pathway in nonplant eucaryotes and the -subclass of purple bacteria, involving the condensation of glycine and succinyl-CoA to produce CoA, carbon dioxide, and ALA (1). Animal genomes encode two highly conserved but differentially expressed ALAS genes, a housekeeping and an erythroid-specific (eALAS) gene (2). In humans, mutations in the eALAS gene can result in X-linked sideroblastic anemia, (3) a hypochromic and microcytic anemia char acterized by iron accumulation within erythroblast mitochondria (4). Approximately one-third of XLSA patients are pyridoxine responsive. In these patients mutations in ALAS are commonly observed in the PLPbinding site (5, 6). The ALAS chemical mechanism (Schem e 2.1) is complex and involves a high degree of stereoelectronic control, with indivi dual steps including: binding of glycine (I); transaldimination with the active site lysine (K313, murine eALAS numbering) to yield an external aldimine (II); abstraction of the pro-R proton of glycine (III); condensation with succinyl-CoA (IV) and CoA release to generate an -amino-ketoadipate intermediate (V); decarboxylat ion resulting in an enol-quinonoid rapid equilibrium (VI); protonation of the enol to give an aldimine -bound molecule of ALA (V II); and ultimately release of the product (ALA) (VIII) (7). This mechanistic complexity is manifested structurally as an enzyme with an unusua lly high degree of sequence conservation, as exemplified by the observation that the catalytic core of human eALAS and R. capsulatus ALAS are 49% identical and 70% similar (8).
54 Scheme 2.1. The role Ser-254 plays in the chemical mechanism of ALAS.
55 PLP-dependent enzymes are classified based on structural and mechanistic similarities (9). ALAS is evolutionarily related to transaminases and is grouped within class II of the fold type I PLP-dependent en zyme superfamily, for which the prototypical enzyme is generally considered to be aspartate aminotransferase (10-12). ALAS is most closely related to the three other members of the -oxoamine synthase subfamily, each of which catalyze reactions betw een small amino acids and Co A esters to generate 1,3aminoketones, while also sharin g high structural similarity (13, 14). Studies have demonstrated that aspartate aminotransferase exists in two predominant conformational forms, open and closed, and reactions catalyzed by PLP-dependent enzymes have been postulated to occur in a closed conf ormation, consistent with the induced fit hypothesis, where electrostatic and hydrophobic interactions between the substrates, cofactor, and amino acids comprising the activ e site provide the energetic impetus to stabilize this catalytically optimal conformation (15, 16). Prior to solution of an ALAS crystal stru cture, kinetic data led investigators to propose that ALAS transitions between open and closed conformations during the catalytic cycle (7, 17). Steady-state kinetic experiment s demonstrate that the kinetic mechanism is ordered, with glycine binding befo re succinyl-CoA, yet in transient kinetic studies binding of succ inyl-CoA accelerates the apparent rate at which glycine binds to ALAS by over 250,000-fold (18). This enhancement might occu r by utilization of part of the intrinsic binding energy for succinyl-CoA to shift the enzyme conformer equilibrium towards a closed conformation wherein tran saldimination of glycine with the PLP cofactor is rapid (17, 18). Return to the open conformation is considered to be the key step limiting ALA release and the overall catalytic rate.
56 The crystal structures of Rhodobacter capsulatus ALAS in holoenzymic and substrate-bound forms adopt open and clos ed conformations, respectively, further supporting the hypothesis that enzyme dynamics play a crucial role during the ALAS catalytic cycle (8). While the structure in genera l collapses slightly around the bound substrates, a more conformationally mobile loop of amino acids located between two sheets at the subunit interface cl oses directly over the channe l leading approximately 20 down into the deeply recesse d active site (Figure 2.1) (8). A conserved threonine at the apex of the mobile loop forms a strong hydr ogen bond (~2.5) with th e carboxylate tail of succinyl-CoA in the substrate-bound struct ure and appears to simultaneously provide molecular recognition for succinyl-CoA while he lping to lock this s ubstrate into optimal position for catalysis. Closer comparison of the active site structures reveals that, coincident with these changes, the side chain of S189 migrates from non-covalently associating with the peptide macroskeleton to within hydrogen bonding distance of the PLP phenolic oxygen, as well as the sulfur atom of succinyl-CoA (8, 19, 20). These interactions suggest that th is serine may be an important determinant in conformer equilibrium and catalysis by providing orienta tional binding energy be tween the cofactor and substrate, while stabilizing a clos ed Michaelis complex conformation. The conservation of this residue in ALAS and the other -oxoamine synthases suggests an important functionality that may be genera l to these enzymes (Figure 2.2). Here we present experiments aimed at probing the role of this serine in catalysis by murine eALAS. We have generated and purified th e positionally equivalent S254A and S254T variants and investigated the effects of th ese mutations on the kine tic and spectroscopic properties of the enzyme. The results s upport the postulate that S254 is a key
57 A B Cmultifunctional residue that couples succiny l-CoA binding to enzyme conformational equilibria and catalysis. Figure 2.1. Structural models for murine erythroid ALAS based on the R. capsulatus crystal structures. (A) Michaelis complex mo deled by alignment of open holoenzyme and closed glycine and succiny l-CoA bound monomeric st ructures. Serine254 is hidden by the succinyl-CoA ester in this view from the persp ective of the adjacent subunit, which has been removed. The active site loop is shown in yellow cartoon for the open and closed conformations, while all othe r structural features are for the closed conformation. (B) Serine-254 in the open conformation. (C ) Serine-254 in the closed conformation with succ inyl-CoA bound.
58 Figure 2.2. Multiple sequence alignment of phylogenetically diverse members of the -oxoamine synthase family in the reg ion of murine eALAS serine-254. The amino acid sequences were retrieved from public databases (NCBI) and aligned using CLUSTAL W (21). The conserved serine re sidue is high-lighted in cyan. The amino acid numbering in red refers to that of murine eryt hroid ALAS (mALAS2). Represented proteins are: M. mus. AL2 Mus musculus erythroid ALAS (156255176); H. sap. AL2 Homo sapiens erythroid ALAS (28586); H. sap. AL1 Homo sapiens housekeeping ALAS (40316939); S. cer. ALA Saccharomyces cerevisiae ALAS (151942209); R. cap. ALA Rhodobacter capsulatus ALAS (974202); A. nig. AON Aspergillus niger AONS (61696868); A. tha. AON Arabidopsis thaliana AONS (42573269); M. mar. AON, Methanococcus maripaludis AONS (1599054); E. col. AON Escherichia coli AONS (85674759); H. sap. KBL Homo sapiens KBL (3342906); C. kor. KBL Candidatus korarchaeum cryptofilum (17017433); E. col. KBL Escherichia coli KBL (169753078); H. sap. SPT Homo sapiens SPT (4758668); A. tha. SPT Arabidposis thaliana SPT (17221603); S. cer. SPT Saccharomyces cerivisiae SPT (706828), E. col. SPT Escherichia coli SPT (170517920). 254 M. mus. AL2 NDPGHLKKLLEKSDPK---------TPKIVAFETVH S MDGAICPLEELCD H. sap. AL2 NDPDHLKKLLEKSNPK---------IPKIVAFETVH S MDGAICPLEELCD H. sap. AL1 NDVSHLRELLQRSDPS---------VPKIVAFETVH S MDGAVCPLEELCD S. cer. ALA NDLNELEQLLQSYPKS---------VPKLIAFESVY S MAGSVADIEKICD R. cap. ALA NDVAHLRELIAADDPA---------APKLIAFESVY S MDGDFGPIKEICD A. nig. AON SCPRSLEDVLRREVEGDE-MVRNGKKNVFLVIESIY S MDGDIAPIREFVE A. tha. AON CDMYHLNSLLSNCKMKR----------KVVVTDSLF S MDGDFAPMEELSQ M. mar. AON NNTVDLIEIL-EKN-KN-------YENKFIVTDAVF S MDGDIAPVGELKK E. col. AON NDVTHLARLLASPCPGQ----------QMVVTEGVF S MDGDSAPLAEIQQ H. sap. KBL LDMADLEAKLQEAQKH---------RLRLVATDGAF S MDGDIAPLQEICC C. kor. KBL CDLADLEDKL-RQVHKK-------YNKILIITDGVF S MDGDIAPLDGITK E. col. KBL NDMQELEARLKEAREAG-------ARHVLIATDGVF S MDGVIANLKGVCD H. sap. SPT NNMQSLEKLLKDAIVYGQPRTRRPWKKILILVEGIY S MEGSIVRLPEVIA S. cer. SPT GDMVGLEKLIREQIVLGQPKTNRPWKKILICAEGLF S MEGTLCNLPKLVE A. tha. SPT NTPGHLEKVLKEQIAEGQPRTHRPWKKIIVVVEGIY S MEGEICHLPEIVS E. col. SPT NDAKDLERRMVRLGER--------AKEAIIIVEGIY S MLGDVAPLAEIVD : .: : * :
59 Materials Reagents. The following were purchased from Sigma-Aldrich Chemical Company (St. Louis, MO): ampicill in, DEAE-Sephacel, Ultrogel AcA-44, mercaptoethanol, PLP, bovine serum al bumin, succinyl-CoA, ALA-hydrochloride, ketoglutaric acid, -ketoglutarate dehydrogenase, HEPE S-free acid, MOPS, tricine, thiamine pyrophosphate, NAD+, and the bicinchoninic acid pr otein determination kit. Glucose, glycerol, glycine, disodium et hylenediamine tetraacetic acid dihydrate, ammonium sulfate, magnesium chloride he xahydrate, and pota ssium hydroxide were acquired from Fisher Scientific (Pittsburgh, PA ). Sodium dodecyl sulfate polyacrylamide gel electrophoresis reagents were acquired from Bio-Rad. Xba I, Bam HI restriction enzymes, Vent DNA Polymerase, and T4 DNA ligase were from New England Biolabs (Ipswich, MA). Oligonucleotides were s ynthesized by Integrated DNA Technologies (Coralville, IA). Methods Mutagenesis. The pGF23 expression plasmid encode d the full-length sequence for the murine, mature eALAS (22). Site-directed mutagenesi s for the S254A and S254T mouse ALAS variants was performed on th e whole plasmid pGF23 using a previously described method (23). The mutagenic oligonucleotides for S254A and S254T were: 5GAG ACT GTT CAT GCC ATG GAT GGT GCC-3 an d 5-GAG ACT GTT CAT ACC ATG GAT GGT GCC-3, respectively, with the introduced codon substitutions underlined. The PCR-generated DNAs were sequenced between the Blp I and Bam HI restriction enzyme sites to confirm the pr esence of the intende d mutation. The products
60 were then digested with Blp I and Bam HI and subcloned into pGF23 that was digested similarly. Protein purification, SDS-PAGE, protein determination and ste ady-state analysis. Recombinant murine eALAS and the S 254 variants were purified from DH5 Escherichia coli bacterial cells containing the ove rexpressed protein as previously described (22). Purity was determined by SDS-PAGE (24) and protein concentration determined by the bicinchoninic acid method us ing bovine serum albumin as the standard (25). All protein concentrations are reported on the basis of a subunit molecular weight of 56 kDa. Enzymatic activity was determin ed by a continuous spectrophotometric assay at 30oC (26). Structural analyses. The protein data base files 2BWN, 2BWO, and 2BWP, corresponding to the holoenzyme, succinyl-CoA-bound, and glycine-bound R. capsulatus ALAS crystal structures, were used as templates to model the PLP-binding core of the murine eALAS (8). Hydrogen bond determinations were accomplished using Deepview/Swiss-PdbViewer software (27, 28). Spectroscopic measurements. All spectroscopic measurements were performed with dialyzed enzyme in 20 mM HEPES, pH 7.5 with 10% (v/v) glycerol to remove free PLP. Circular dichroism (CD) spectra we re collected using an AVIV CD spectrometer calibrated for both wavelength maxima and signa l intensity with an aqueous solution of D-10 camphorsulfonic acid (29). CD spectra recorded fr om 190-240 nm were analyzed by the ridge regression method using a modi fied version of the computer program CONTIN developed by Provencher and Glckner (30). Protein concentrations were 10 M and 100 M for the near and far CD spectra, resp ectively. The final concentration of
61 succinyl-CoA was 100 M, giving a 1:1 molar ratio of enzyme to ligand for collection of the latter spectra. At least three CD spectra were collecte d per experiment and averaged, using a 0.1 cm path length cuve tte with a total volume of 300 l. Blank CD spectra contained all components of the solution ex cept enzyme. CD sp ectra containing the enzyme samples were collected immediately af ter adding the enzyme. The spectra of the samples containing enzyme were analyzed afte r substracting the blank spectra. Co-factor fluorescence spectra were collected on a Shimadzu RF-5301 PC spectrofluorophotometer using protein concentrations of 2 M. Spectra were measured at pH 7.5, 50 mM HEPES, and 20% (v/v) glycerol. Excitation was at 331 nm and the excitation and emission slit widths were each set to 10 nm. Emission was measured over the wavelength range 350 to 600 nm. Buffer blanks were subtracted from the spectra. Stopped-flow spectroscopy. All of the experiment s were carried out at 30C in 100 mM HEPES, pH 7.5 and 10% (v/v) glycerol. Th e concentration of reactants loaded into the syringes were always 2-fold greater than that present in the cell compartment after mixing. Because of the difference in Km for succinyl-CoA between the two variant enzymes, two different succinyl-CoA conc entrations were used to ensure the identification of a single enzyme-catalyzed ev ent. For the S254A-catalyzed reaction, the final concentra tions were: 120 M S254A, 130 mM glycine, and 30 M succinyl-CoA. The final concentrations of the reactants for S254T were: 50 M S254T, 130 mM glycine, and 10 M succinyl-CoA. Rapid scanning stoppe d-flow kinetic measurements were conducted using an OLIS model RS M-1000 stopped-flow spectrophotometer. The dead time of this instrument is approximately 2 ms, and the observation chamber optical path length is 4.0 mm. Scans covering the wavelength region 270-550 nm were acquired
62 at a rate of either 62 or 31 scans per second in order to condense the resulting data files to a tractable size for data fitting analysis. An external water bath was utilized to maintain constant temperature of the sy ringes and observation chamber. Specifically, spectral data covering the 270 550 nm range were analyz ed by global fitting to a triple exponential using the SIFIT program supplied with the stopped-flow instrument (OLIS, Inc.).(31). The quality of fits were j udged by visual analysis of the calculated residuals in conjunction with the Durbin-Watson statistic (32). Single turnover data were interpreted using a three kinetic step mech anism as described by Equation 2.1. (Equation 2.1) D C B A k k k 3 obs 2 obs 1 obs The observed rate constants were determ ined from at least three replicate experiments, and the reported values repr esent the average and standard error of measurement for each experime ntal condition. The forward and reverse rate constants depicted in the kinetic mechan ism (Figure 2.7) were obtained by modeling single wavelength kinetic traces at 510 nm with KinTekSim (Austin, TX) kinetic simulation software (33). The eight interior rate constants were allowed to float, while the KD values were held constant as determined separately. Transient kinetics of th e reaction of glycine w ith the variant enzymes. The reactions of the murine eALAS variants wi th glycine were perf ormed using the same instrument as was described for the single turnover reactions with the enzyme-glycine complex and succinyl-CoA. The fi nal enzyme concentration was 60 M. The glycine concentration was always at least 10-fold great er than the enzyme concentration to ensure that pseudo-first order kineti cs were observed. The treatment of the data was performed
63 using the fitting software supplied with the instrument. The time courses at 420 nm were fitted to Equation 2.2 (Equation 2. 2) 3 1) (n t k i tc e a Ai where At is the absorbance at time t a is the amplitude of each phase, k is the observed rate constant for each phase, and c is the final absorbance. The quality associated with the fit was determined by the calculated residuals and from the Durbin-Watson ratio (32) The observed rate constants were plotted against increasing c oncentrations of glycine and the resulting data were fitted to a two step reaction process represented by Equation 2.3. Data fitting to Equation 2.4 used the nonlinea r regression analysis program SigmaPlot10 (Systat, San Jose, CA) (Equation 2.3) C B A k k 2 obs 1 obs (Equation 2.4) 1 1] [ ] [ k S K S k kD obs where S is the concentration of substrate, k1 and k-1 are the forward and reverse rate constants, KD is the dissociation constant and kobs, the observed rate constant. Intrinsic protein fluorescence quenching. The pre-steady stat e kinetics of the product binding reaction of ALAS and the tw o serine variants were examined by measuring changes in the intrinsic protei n fluorescence intensity. An OLIS RSM-1000F rapid mixing spectrofluorimeter, equipped w ith a high-intensity xenon arc lamp, was used to monitor the reaction. The enzyme and ligand in 20 mM HEPES (pH 7.5) and 10% glycerol were maintained at 30oC in separate syringes prior to their mixing in the reaction chamber. The concentrations of enzyme a nd ligand in the reaction chamber were 1/2 of
64 those in the syringes. The intrinsic pr otein fluorescence, as measured with 5 M enzyme, was evaluated in the presence of increasing co ncentrations of the product, ALA. The excitation wavelength and the slit width were 280 and 5 mm, respectively. The emitted light was filtered using a cutoff filter (W G 320; 80% transmittance at 320 nm, (Edmund Optics, Barrington, NJ)). Typically, 500 time points were collected for varying lengths of time, and three or more experiments were averaged. Each averaged data set was then fitted to Equation 2.5, using the Global fitting software provided with the instrument. (Equation 2.5) 0 1) (A e A t Ft k obsobs where Fobs(t) is the observed fluorescence chan ge (in arbitrary units) at time t, kobs is the observed first-order rate constant, A1 is the pre-exponen tial factor and A0 is the offset. The observed rate constants were then plotted ag ainst ligand concentrati on and the data were fitted to Equation 2.6 by nonlinear regression. The rates of dissociation (koff) and association (kon) as well as the liga nd binding constants (KD) were calculated from the asymptotic maximal observed rate, the ordina te intercept, and the ligand concentration (x) in Equation 2.6. (Equation 2.6) x K x k k x fD off on ) (
65Results Kinetic characterizati on of the S254 variants. The steady-state kinetic parameters of the S254 variants were determined and th e results are summarized in Table 2.1. The mutation of serine-254 to alanine resulted in a kcat 2-fold higher than th at of the wild-type ALAS value. The Km for succinyl-CoA was increased 25-fold relative to ALAS, while the Km for glycine was not significantly affected. The ov erall catalytic efficiency for succinyl-CoA decreased 36-fold, while the value for glycine remained unchanged as compared to ALAS values. The replacemen t of serine-254 with threonine caused a 2fold decrease in the wild-type kcat value. The Michaelis c onstants for both substrates were indistinguishable from those of the wild-type enzyme. Spectroscopy. The orientation and average positioning of the PLP cofactor relative to the conserved serine and the activ e site were perturbed by the replacement of the serine with either an alanine or a threon ine. Gross structural changes evidenced by changes in the alpha helix and beta sheet cont ent of proteins can be identified from CD spectroscopic changes in the far-UV (ultraviolet) (34). The analysis of the UV CD spectra (Figure 2.3A) by CONTIN-CD indicates any changes in the secondary structure between wild-type ALAS and the two ALAS va riants are negligible. Locally chiral substructures that comprise the PLP mi croenvironment modulate the visible CD characteristics of the chromophore. Spectra for the wild-type and variant enzymes, as holoand succinyl-CoA-bound enzymes, were co llected (Figure 2.3). The spectra for wild-type ALAS and S254T holoenzymes disp layed positive dichroic bands at 330 and 420 nm. However, in relation to the wild -type ALAS, the S254A variant showed a
66 decrease in amplitude of the 330 nm band, while the amplitude in the 420 region increased. Table 2.1: Summary of steady-state kinetic parameters Enzyme Gly mK (mM) SCoA mK (M) catk (s-1) catk/Gly mK (mMs-1) catk/SCoA mK (Ms-1) Wild-type 25 4.2 1.3 0.9 0.14 0.02 0.01 0.11 S254A 18 1.7 32 7.7 0.27 0.01 0.02 9.0 x 10-3 S254T 27 2.9 1.2 0.3 0.050 0.004 2.0 x 10-3 0.05
67 Figure 2.3. Circular dichroism and fluorescence em ission spectra of ALAS and the S254 variants. Spectra of wild-type ALAS (), S254A () and S254T (---) (A) and (B) Holoenzymes; (C) in the the presence of 100 M succinyl-CoA; (D) upon excitation of the cofactor at 331 nm.
68 (Figure 2.3B). Wild-type ALAS and vari ant enzymes responded differently to the presence of 100 M ligand (Figure 2.3C). Compar ison of the CD spectra for the holoenzymes with succinyl-CoA showed that spectra of wild-type and S254T changed in the presence of substrate, while that of S254A maintained dichroic characteristics that were indistinguishable from those observed und er ligand-free conditions. Specifically, in wild-type and S254T, the amplitude of both dichroic bands decreased and the 330 nm peak red shifted to ~350 nm. The cofactor fluorescence spectra obtained at pH 7.5 for the S254T variant enzyme with ex citation at 331 nm is simila r to that of the wild-type enzyme with a well-formed emission maximu m ~385 nm (Figure 2.3D). A notable deviation in emission maximum is observe d for the S254A variant. Specifically, excitation of S254A at 331 nm results in a broader fluorescence emission band centered around 450 nm. The changes in the fluorescence emission prof ile for the alanine variant suggest that the tautomeric equilibrium betw een at least two forms of the PLP cofactor aldimine linkage with the catalyt ic lysine is perturbed. Reaction of glycine with the S254 variants. The reaction of 60 M S254 variants with increasing amounts of glycine resulted in an increased absorbance at 420 nm (Figure 2.4). A global fit of the data for the reacti on of S254A with glycin e yielded values for k1 of 0.159 0.04 s-1, k-1 of 0.072 0.001 s-1, and a KD of 6.6 0.57 mM. The fitting of the data corresponding to the reaction of the S254T variant with glycine provided parameters of k1 of 0.11 0.01 s-1, and k-1 of 0.070 0.004, and a KD of 1.5 0.39 mM. ALA binding kinetics monitored by intrinsic protein fluorescence. The observed rates of intrinsic protein fl uorescence quenching were determ ined as a function of ALA concentration, and the results are presented in Figure 2.5. The hyperbolic nature of the
69 Figure 2.4. Reaction of the S254 variants (60 M) with increasing concentrations of glycine. Data were fit to Equation 2.2 for a two-exponentia l process, yielding equilibrium and rate constants for S254A of KD = 6.6 0.57 mM, k1 = 0.159 0.04 s1, and k-1 = 0.072 0.001 s-1. The fitted constants for S254T were: KD =1.5 0.39 mM, k1 = 0.11 0.01 s-1 and k-1 = 0.070 0.004 s-1.
70 Figure 2.5. Reaction of wild-type ALAS and the S254 variants (5 M) with ALA. The observed rate constants were calculated by fitting the decrease in intrinsic protein fluorescence over time to Equation 2.3 for a single-exponential process. The resolved equilibrium and rate constants for wild-type ALAS (A) were: KD = 500 16 M, k1 = 0.120 0.015 s 1, and k-1 = 0.140 0.05 s-1. For the S254A variant (B) the constants were: KD = 855 66 M, k1 = 0.235 0.006 s 1, and k-1 = 0.29 0.02 s-1, and for the S254T variant were: KD = 832 49 M, k1 = 0.19 0.09 s-1, and k-1 = 0.057 0.007 s 1.
71 binding data indicate a two-step process, and may be ascribed to formation of a collision complex followed by the conformational ch ange associated with ALA binding (7). Significantly, for each of the three enzymes the resolved rate constants for the off rate conformational change (k-1) coincide with the kcat values determined through steady-state kinetics. The dissociation constants of the variants for ALA are increased by approximately 60% over that of the wild-type enzyme. Pre-steady-state reaction of the variant enzyme-glycine complexes with succinylCoA. ALAS catalysis involves th e sequential binding of gl ycine first (I-II), then succinyl-CoA (III-IV), followed by formation of an enol-quinonoid equilibrium (VI-VII) after the liberation of CO2 (Scheme 2.1) (7). In order to determine the microscopic rates associated with the lifetime of the quinonoid intermediate we monitored the time course of the ALAS-catalyzed reacti on under single turnover condi tions. The time courses of the absorbance change were best fit to a sequential, three-step mechanism outlined by Equation 3.1. Among all the enzymes test ed, a single step assigned to quinonoid intermediate formation, followed by a biphasic step of its decay were observed (Figure 2.6). For each enzyme, the global fit of the sp ectral data at 510 nm is shown as a solid line overlaid with the time course data at 510 nm (dots). The rate constants associated with quinonoid intermediate formation (Qf) differ between the two variants. For S254A, the value is 4.8 0.2 s-1, a rate similar to that of the wild-type enzyme (7). However, the rate value for the S254T variant was decreased, showing a 4-fold reduction with a rate of 1.4 0.3 s-1. These data suggest that the loss of the hydroxyl group at position 254 has only a modest effect on quinonoid intermediate formation, and therefore does not appear to play any obvious role in active site chemis try during catalysis. The rates assigned to
72 Figure 2.6. Reaction of wild-type ALASand S254 variant-glycine complexes with succinyl-CoA under single turnover conditions. The data for single turnover quinonoid intermediate formation and decay reaction kinetics were fitted with the SIFIT program (OLIS, Inc.). The data ( ) are overlaid with the line representing the fitted data. ( ). (A) The rate constants for the three step sequence in the wild-type enzyme were 6.0 s-1, 2.0 s-1, and 0.075 s-1. (B) The rate constants for the three steps in S254Acatalyzed reaction were 4.8 s-1, 0.8 s-1, and 0.19 s-1. (C) The rate constants for the three steps in S254T-catalyzed reaction were 1.4 s-1, 0.2 s-1, and 0.037 s-1.
73 the two-step quinonoid intermediate decay (Qd1 and Qd2) also differ among the variants. Both variants have initial qui nonoid intermediate decay rates th at are at least 2-fold lower than that of the wild-type enzyme (i.e., 0.8 s-1 and 0.25 s-1 vs. 2.0 s-1). However, the second step of quinonoid intermediate decay, which most closely approximates kcat for the overall reaction, is 40% faster for the S254A variant, in comparison to that of the wild-type enzyme. This increase d value agrees with the greater kcat that was determined from the steady-state kinetic analysis (Table 2.1). As such, the rates for S254A observed during the lifetime of the quinonoid intermedia te indicate fast transformation of the enzyme-substrate complex to the enzyme-product complex (Qf and Qd1), followed by a comparatively faster rate of product dissoci ation and the return of the conformational equilibrium to the catalytically favored open conformation (Qd2).
74Discussion The spectroscopic and kinetic properties of the S254A and S254T murine ALAS variants were examined and compared to thos e of the wild-type enzy me in an effort to better understand the role of this residue in the catalytic pro cess. Mutation of serine to alanine removes the side chain oxygen atom and eliminates the po ssibility of hydrogen bond formation with the PLP cofactor and succinyl-CoA. The co nservative S254T mutation adds a methyl group, and is predicted to allow for hydrogen bonding, although some steric constraints may be introduced due to the tight packing in this region of the active site. Comparison of the steady-state kinetic parameters of the variants to those of the wild-type enzyme determ ined under similar conditions reveals the unusual kinetic significance of S254 (Table 2.1) Point mutations typically re sult in increased Michaelis constants and decreased turnover numbers, but significantly, in the S254A variant both of these parameters are increased. The Km for succinyl-CoA is incr eased 25-fold over that of the wild-type enzyme, while kcat is increased 2-fold. It is notable that mutation of an evolutionarily conserved residu e in such a metabolically important enzyme leads to an enhanced kcat, even though the effect is not dramatic. The kcat for murine eALAS is considered to be defined by a conformational change of the enzyme accompanying ALA release (7). This is further supported here by the stopped-flow data in Figure 6, where the protein fluorescence associated ALA off rates (k-1) are indistinguishable from the steady-state kcat values (Table 2.1). These data also indicate that the rate-limiting step of the reaction cycle is unaltered in the mutated enzymes. The increase in kcat observed for the S254A varian t may be attributable to diminished stability of the closed conformati on, leading to faster reversion to the open
75 conformation and product release. In this interpretation, the large concomitant increase in Km for succinyl-CoA in the S254A variant woul d result not only from the direct effect of loss of hydrogen bonding to the substrate, but also from the less direct effect of impaired enzyme conformational responsiveness to the substrate that is believed to be required for optimal binding. The three-fold decrease in kcat without any effect on the Km for succinyl-CoA resulting from the more c onservative S254T mutation would then arise primarily from enhanced stability of the closed conformation. Both mutations substantially diminish the catalytic efficiency with succinyl-CoA (kcat/SCoA mK), but in different ways, and while the effects of th ese very conservative mutations on the steadystate kinetic parameters may appear relativel y modest, they are presumably sufficient to have metabolically harmful effects in vivo, given the strong evolutionary selection for serine at this position. The S254A mutation has significant effects on the cofactor mi croenvironment, as determined by fluorescence and CD spectrosc opies. The fluorescence spectra suggest a rather dramatic alteration in cofactor tautomeric equili bria occurs upon loss of hydrogen bonding interactions between the phenolic oxy gen of PLP and the side chain of S254 (Fig. 2.3D). The diminished enolamine tautomer emission at 386 nm in the S254A mutant is consistent with loss of hydrogen bonding at the cofactor phenolic oxygen, and an increased proportion of the cofactor maybe in the ketoenamine (35). CD spectroscopic evaluations of the conforma tional effects of th e S254A and S254T mutations in the far-UV region verify the mu tations do not significan tly alter the overall secondary structure of the enzymes (Fig. 2.3A ). Any alterations in the conformational equilibria of the active site loop, which adopts an extended conformation and is thus not
76 CD active, are not apparent in these spectra, but the visi ble CD spectra of the S254A variant does diverge substantially from those of the wild-type and the S254T variant. The S254A holoenzyme maximum at 435 nm is blue shifted towards ~420 nm, and the ratio of the mean residual ellipticity at this wavelength to the one at 330 nm was increased (Fig. 2.3B). These ellipticities ar ise from Cotton effects associated with the ketoenamine (435 nm) and enolamine (330 nm ) cofactor aldimine bond tautomers, and are indicative of the microenvironment surroundi ng this linkage between the cofactor and the active site lysine (36). Succinyl-CoA binding to the wild-type and S254T variants induce decreases in asymmetry of the cofactor, while the S 254A mutant is relatively unchanged under similar conditions (Fig. 2.3C). A logical interpre tation is that the decrease in asymmetry observed for the wild-t ype and S254T variants arises from partial conversion of the internal al dimine to free PLP aldehyde b ound at the active site, as is observed in three out of four R. capsulatus crystal structure active sites upon succinylCoA binding (8). In the crystal structures these ev ents are accompanied by transition to a closed conformation, from which it might be concluded that the S254A variant retains the internal aldimine in the presence of succi nyl-CoA, and may not be induced to adopt a closed conformation upon binding of this substrate. The quinonoid intermediate single-turnove r profiles of the two S254 variants indicate that they follow a ch emical mechanism similar to th at of the wild-type enzyme (Fig. 2.6A, B). A rapid step of AL A-bound quinonoid intermediate formation upon decarboxylation is followed by two successively slower decay steps, associated with protonation of the ALA-quinonoid intermed iate and ALA release, respectively (7). The quinonoid intermediate formation rate decreas ed 4-fold for the S254T variant, which
77 could be explained by a change in the flow of electrons from the site of bond scission throughout the cofactor, precipita ted by a shift in the conformational equilibrium towards closed. In the ALAS crystal structure, PL P is noted to change position by 15 degrees when substrate is bound (8). Changes to the stereoelectronic relationship between the cofactor and the -carbon bonds of the external aldimi ne can influence the chemical mechanism dramatically (7). Therefore, it is possibl e that the hydrogen bond between serine-254 and the phenolic oxygen of PLP may be an influential part of maintaining the angle of PLP during not only formation and d ecay of the quinonoid intermediate, but also during the complete reaction cycle. The increase observed in the second step of quinonoid intermediate decay in the S254A varian t is also consistent with the increases in kcat and the ALA off rate determined from enzyme fluorescence quenching. By utilizing the microscopic paramete rs obtained from the single turnover reactions of the variant enzyme s, coupled with the product an d substrate reaction data, we were able to model the kinetic mechanisms as shown in Figure 2. 7. Additionally, the Gibbs free energy associated with the glyc ine and ALA were calculated (Table 2.2). The kinetic simulations revealed that th e S254T mutation significantly retards the chemical mechanism. Conversely, the modeled pathway for S254A highlights the increases observed in the kinetics of the varian t. Overall, the mechanistic data for both variants support the hypothesis that the interactio n between S254 and the O3 of PLP is a limiting factor in enforcing an induced fit mechanism by coupling substrate recognition to conformational equilibria. However, how the structural differences between the variant enzymes with ligand bound accomplish this awaits three-dimensional structural information.
78 A B Figure 2.7. Kinetic mechanisms of the S254 variant enzymes. The single turnover quinonoid formation and decay reaction kinetics of the variant enzymes and the reactions with glycine and ALA and were used to model the kinetic mechanisms. The fits ((A) S254A) and ((B) S254T) were in distinguishable from those completed based on the global fit of spectral data. E, ALAS; G, gl ycine; EG, ALAS-glycine complex; SCoA, succinyl-CoA; EGSCoA, ALAS-glycine-s uccinyl-CoA complex; EQ, observable quinonoid intermediate; EALA1, ALAS-ALA in ternal aldimine w ith active site loop closed; and EALA2, ALAS-ALA internal aldimine with active site loop open. Table 2.2. Gibbs free energy associated with the wild-type ALASand S254 variant-catalyzed reactions. Enzyme GALA ( kcal / mol ) GGly ( kcal / mol ) Wild-type 4.18 3.34 S254A 4.08 3.97 S254T 4.07 2.78
79Acknowledgements This work was supported by the National Institutes of Health (grant DK63191 to GCF). References (1) Akhtar, M., Abboud, M. M., Barnard, G., Jordan, P., and Zaman, Z. (1976) Mechanism and stereochemistry of en zymic reactions in volved in porphyrin biosynthesis. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 273, 117-136. (2) May, B. K., Dogra, S. C., Sadlon, T. J., Bhasker, C. R., Cox, T. C., and Bottomley, S. S. (1995) Molecular regul ation of heme biosynthesis in higher vertebrates. Prog. Nucleic Acid Res. Mol. Biol. 51, 1-51. (3) May, A., and Bishop, D. F. (19 98) The molecular bi ology and pyridoxine responsiveness of X-linke d sideroblastic anaemia. Haematologica 83, 56-70. (4) Bottomley, S. S. (2006) Conge nital sideroblastic anemias. Curr. Hematol. Rep. 5, 41-49. (5) Shoolingin-Jordan, P. M., Al-Daihan, S., Alexeev, D., Baxter, R. L., Bottomley, S. S., Kahari, I. D., Roy, I., Sarwar, M., Sawyer, L., and Wang, S. F. (2003) 5Aminolevulinic acid synthase: mech anism, mutations and medicine. Biochim. Biophys. Acta. 1647, 361-366. (6) Bottomley, S. S. (2004) Sideroblastic anemias in Wintrobe's Clinical Hematology (Greer, J. F., J. Lukens, J.N. Rodgers, G.M. Paraskevas, R. Glader, B., Ed.) pp 1012-1033, Lippincott, Williams, & Wilkins, Philadelphia. (7) Hunter, G. A., Zhang, J., and Ferreira G. C. (2007) Transi ent kinetic studies support refinements to the chemical and kinetic mechanisms of aminolevulinate synthase. J. Biol. Chem. 282, 23025-23035. (8) Astner, I., Schulze, J. O., van den He uvel, J., Jahn, D., Schubert, W. D., and Heinz, D. W. (2005) Crystal structure of 5-aminolevulinate synthase, the first enzyme of heme biosynthesis, an d its link to XLSA in humans. EMBO J. 24, 3166-3177. (9) Eliot, A. C., and Kirsch, J. F. (2004 ) Pyridoxal phosphate en zymes: mechanistic, structural, and evolutionary considerations. Annu. Rev. Biochem. 73, 383-415. (10) Jager, J., Moser, M., Sauder, U., and Ja nsonius, J. N. (1994) Crystal structures of Escherichia coli aspartate aminotransferase in two conformations. Comparison of an unliganded open and two liganded closed forms. J. Mol. Biol. 239, 285-305. (11) Picot, D., Sandmeier, E., Thaller, C., Vincent, M. G., Christen, P., and Jansonius, J. N. (1991) The open/closed conf ormational equilibrium of aspartate aminotransferase. Studies in the crystal line state and with a fluorescent probe in solution. Eur. J. Biochem. 196, 329-341. (12) McPhalen, C. A., Vincent, M. G., Pico t, D., Jansonius, J. N., Lesk, A. M., and Chothia, C. (1992) Domain closure in mitochondrial as partate aminotransferase. J. Mol. Biol. 227, 197-213. (13) Christen, P., and Mehta, P. K. (2001) From cofactor to en zymes. The molecular evolution of pyridoxal-5'-phos phate-dependent enzymes. Chem. Rec. 1, 436-447.
80 (14) Alexander, F. W., Sandmeier, E., Mehta, P. K., and Christen, P. (1994) Evolutionary relationships among pyri doxal-5'-phosphate-dependent enzymes. Regio-specific alpha, beta and gamma families. Eur. J. Biochem. 219, 953-960. (15) Jansonius, J. N., Eichele, G., Ford, G. C., Kirsch, J. F., Picot, D., Thaller, C., Vincent, M. G., Gehring, H., and Christ en, P. (1984) Crystallographic studies on the mechanism of action of mitoc hondrial aspartate aminotransferase. Prog. Clin. Biol. Res. 144B, 195-203. (16) Jansonius, J. N., Eichele, G., Ford, G. C., Kirsch, J. F., Picot, D., Thaller, C., Vincent, M. G., Gehring, H., and Christ en, P. (1984) Three-dimensional structure of mitochondrial aspartate aminotransfera se and some functi onal derivatives: implications for its mode of action. Biochem. Soc. Trans. 12, 424-427. (17) Hunter, G. A., and Ferreira, G. C. (1999) Pre-steady-state reaction of 5aminolevulinate synthase. Evidence fo r a rate-determining product release. J. Biol. Chem. 274, 12222-12228. (18) Zhang, J., and Ferreira, G. C. (2002) Transient state kinetic investigation of 5aminolevulinate synthase reaction mechanism. J. Biol. Chem. 277, 44660-44669. (19) Gregoret, L. M., Rader, S. D., Fletterick, R. J., and Cohen, F. E. (1991) Hydrogen bonds involving sulfur atoms in proteins. Proteins 9, 99-107. (20) Rajagopal, S., and Vishveshwara, S. (2005) Short hydrogen bonds in proteins. FEBS J. 272, 1819-1832. (21) Thompson, J. D., Higgins, D. G., and Gibson, T. J. (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Research 22, 4673-80. (22) Ferreira, G. C., and Dailey, H. A. (1993) Expression of mammalian 5aminolevulinate synthase in Escherichia coli. Overproduction, purification, and characterization. J. Biol. Chem. 268, 584-590. (23) Miyazaki, K., and Takenouchi, M. (2 002) Creating random mutagenesis libraries using megaprimer PCR of whole plasmid. Biotechniques 33, 1033-1034, 10361038. (24) Laemmli, U. K. (1970) Cleavage of struct ural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680-5. (25) Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk, D. C. (1985) Measurement of protei n using bicinchoninic acid. Anal. Biochem. 150, 7685. (26) Hunter, G. A., and Ferreira, G. C. (1995) A continuous spectrophotometric assay for 5-aminolevulinate synthase th at utilizes substrate cycling. Anal. Biochem. 226, 221-224. (27) Schwede, T., Kopp, J., Guex, N., and Peitsch, M. C. (2003) SWISS-MODEL: An automated protein homology-modeling server. Nucleic Acids Res. 31, 3381-3385. (28) Guex, N., and Peitsch, M. C. (1997) SWISS-MODEL and the Swiss-PdbViewer: an environment for comparative protein modeling. Electrophoresis 18, 2714-23. (29) Chen, G. C. Y., J.T. (1977) Two-point calibration of cicular dichrometer with D10-camphosulphonic acid. Analytical Letters 10, 1195-1207.
81 (30) Provencher, S. W., and Glockner, J. (1981) Estimation of globular protein secondary structure from circular dichroism. Biochemistry 20, 33-37. (31) Tsai, M. D., Weintraub, H. J., Byrn, S. R., Chang, C., and Floss, H. G. (1978) Conformation-reactivity rela tionship for pyridoxal Schiff's bases. Rates of racemization and alpha-hydrogen exchange of the pyridoxal Schiff's bases of amino acids. Biochemistry 17, 3183-3188. (32) Durbin, J., and Watson, G. S. (1970) Te sting for serial correla tion in least squares regression. Biometrika 37, 409-414. (33) Barshop, B. A., Wrenn, R. F., and Fr ieden, C. (1983) Analysis of numerical methods for computer simulation of kine tic processes: development of KINSIM-a flexible, portable system. Anal. Biochem. 130, 134-145. (34) Kelly, S. M., and Price, N. C. (2000) The use of circular dichroism in the investigation of protein structure and function. Curr. Protein Pept. Sci. 1, 349384. (35) Tai, C. H., Rabeh, W. M., Guan, R., Schnackerz, K. D., and Cook, P. F. (2008) Role of Histidine-152 in cofactor orientat ion in the PLP-dependent O-acetylserine sulfhydrylase reaction. Arch. Biochem. Biophys. 472, 115-125. (36) Ferreira, G. C., Neame, P. J., and Dailey, H. A. (1993) Heme biosynthesis in mammalian systems: evidence of a Schiff base linkage between the pyridoxal 5'phosphate cofactor and a lysine residue in 5-am inolevulinate synthase. Protein Sci. 2, 1959-1965.
82 Chapter Three Arg-85 and Thr-430 in murine 5-aminolevu linate synthase coordinate acyl-CoAbinding and contribute to substrate specificity Abstract 5-Aminolevulinate synthase (ALAS) catal yzes the rate-limiting step of heme biosynthesis in mammals thr ough the condensation of succiny l-Coenzyme A and glycine to produce 5-aminolevulinate, Coenzyme-A (CoA) and carbon dioxide. ALAS is a member of the -oxoamine synthase family of py ridoxal 5'-phosphate (PLP)-dependent enzymes and shares high degree of structural similarity and reaction mechanism with the other members of the family. The X -ray crystal structur e of ALAS from Rhodobacter capsulatus reveals that the alkanoate component of succinyl-CoA is coordinated by a conserved arginine and a threonine. The functions of the corresponding acyl-CoAbinding residues in murine erythroid ALAS (R85 and T430) in relation to acyl-CoA binding and substrate discrimination were exam ined using site-directed mutagenesis and a series of CoA-derivatives. The catalytic efficiency of the R85L variant with octanoylCoA was 66-fold higher than that of the wild-t ype protein, supporting the proposal of this residue as key in discriminating substrate bi nding. Substitution of the acyl-CoA-binding residues with hydrophobic amino acids caused a li gand-induced negative dichroic band at 420 nm in the CD spectra, suggesting that th ese residues affect substrate-mediated
83 changes to the PLP microenvironment. Transi ent kinetic analyses of the R85K variantcatalyzed reactions confirm that this substitution decreases microscopic rates associated with formation and decay of a key reaction inte rmediate and show that the nature of the acyl-CoA tail seriously affect product binding. These resu lts show that the bifurcate interaction of the carboxylate moiety of succinyl -CoA with R85 and T430 is an important determinant in ALAS function and may play a role in substrate specificity.
84Introduction 5-Aminolevulinate synthase (ALAS; EC 126.96.36.199) is a pyridoxal 5-phosphate (PLP)-dependent enzyme consisting of tw o identical subunits, each containing one molecule of covalently bound PLP. ALAS catalyzes the Claisen-like condensation of glycine and succinyl-CoA to yield carbon dioxide (CO2), CoA, and 5-amino-4oxopentanoate (5-aminolevulinate; ALA), and represents the first step of porphyrin biosynthesis in animals, fungi, and some b acteria. The structural and mechanistic properties of ALAS are markedly similar to those of 8-amino-7-oxononanoate synthase (AONS), serine palmitoyl transferase (SPT ), and 2-amino-3-ketobutyrate-CoA ligase (KBL) (1-3). The x-ray crystal structure of the holo form of Rhodobacter capsulatus ALAS was solved at 2.1 resolution and also as enzyme-substrate complexes with either glycine (2.7 ) or succinyl-CoA (2.8 ) (4). ALAS is classified as a member of the oxoamine synthase subfamily of fold type I PLP-dependent enzyme s. AONS, SPT, and KBL are the other members and represent the closest structural relatives, with the enzymes of the subfamily sharing a C root mean square deviation of approximately 1.5 (5, 6). The reaction chemistries are also hi ghly similar, all involving small amino acids, CoA esters, and 1,3-aminoketones. AON S catalyzes the committed step in biotin biosynthesis,(7) SPT catalyzes the first step of sphingolipid biosynthesis,(8) and KBL catalyzes the degrad ation of threonine (9). Despite the remarkable structural and m echanistic similarities in this important group of enzymes the molecular mechanisms underlying substrate specificity remain largely unexplored. SPTs utilize palmitoyl-C oA as the preferred physiological substrate
85(10), however, Han et. al. have shown that the SPT of a Coccolithovirus is more active when utilizing myristoyl-CoA, a substrate si milar to palmitoyl-CoA but shorter by two carbons (11). Prior to the elucidation of the X-ray crystal structure of R. capsulatus ALAS, the bacterial enzyme-catalyzed reac tion was examined w ith non-physiological acyl-CoA derivatives as substrates (12). Results of this investigation indicate that some naturally occurring three, four and five carbon CoA thioesters can act as substrates and that both acyl chain length and hydrophilicity of the acy l-CoA substrate are important factors in determining specifici ty. The CoA substrate specificity of ALAS is of interest due to localization of the eukaryotic en zyme in the inner mitochondrial matrix. Specifically, 90% of cellular acetyl-CoA and between 92and 97% of short and long chain acyl-CoAs are located within this or ganelle, providing an abundant supply of possible alternative substrat es for meALAS. As such, promiscuous reactions with alternative CoA substrates would produce highly reactive 1-3aminoketones of unknown biological significance, that are potentially capable of dimerizing to form toxic dihydropyrazines (13, 14). Previous investigations regarding the bi nding of the amino acid substrate of ALAS and substrate specificity led to the conclusion that the ALAS active site only accommodates the smallest naturally occurring amino acid, namely glycine (15). Variants of R. sphaeroides ALAS in which the glycinebinding threonine (T83) is mutated to the subtly smaller amino acid serine show a dramatic improvement in acceptance of non-physiological amino acid substrates (15). This finding along with the crystal structures suggest that steric fact ors within the glycine-binding region of the active site are the major determinants of amino acid substrate specificity.
86 The ALAS active site is within a cleft at the subunit interface and is delimited by a -strand bent around the PLP cofactor, in whic h the pyridinium ring of the cofactor lies at the bottom of the cavity (4). Connection between the su rface of the enzyme and the active site is by an amphipath ic channel, which is occupied by succinyl-CoA in the substrate-bound structure (Figur e 3.1). Two distinct moieti es of succinyl-CoA interact with the enzyme: the solvent accessible adenosyl component and the buried succinate. The alkanoic acid moiety of succi nyl-CoA is bound to the active site via a strong hydrogen bond network that stabilizes a cl osed enzyme conformation (Lendrihas et. al., submitted) (4, 16). At the end of a hydrophobic tunnel, the guanidino group of the highly conserved R21 (R85 in murine erythroi d ALAS (meALAS)) donates a hydrogen bond to the carboxylate constituent of succinyl-CoA (Figure 3.1). Simultaneously, the hydroxyl group of the conserved T365 (T430 in meALAS), which is positioned at the apex of a conformationally dynamic active site loop, bridges both the carboxylic acid moiety of succinyl-CoA and the side chain of R21 to complete a hydrogen bonding triad (Figure 3.1) (4). Accordingly, the chemical characteris tics of the acyl-tail of the CoA substrate may be a determining factor for the enzyme in discriminating substrate entry into the active site. In this study, we investig ate the role of the conser ved R85 and T430 residues of meALAS in recognition and binding of the acy l-CoA substrate in relation to catalysis. Substitutions of the conserved residues with more hydrophobic amino acids (i.e., R85L and T430V) were introduced to examine the e ffect of hydrophobicity and steric hindrance on specificity toward the CoA-derived substr ate. Such a difference would alter the aliphaticity of the substrate-binding cleft, which, in turn, could affect the acyl chain-
87Figure 3.1 The acyl-CoA binding cleft in R. capsulatus ALAS. The ALAS dimer appears above the hydrogen bond network maintained between the alkanoic acid component of succinyl-CoA and the side ch ains of the conser ved residues (R21 and T365) is indicated by dashed yellow lines. The PLP cofactor, succinyl-CoA substrate and the corresponding R and T residues (R85 and T430) are shown in stick format.
88 binding properties of this channel. The results presented he re for the R85 and T430 variants of meALAS show th at these residues are involve d in both the orientation and binding of the succinyl-CoA substrate in th e active site and may also, following the substrate binding, assist in enzyme closure. Materials Reagents. The following reagents were purch ased from Sigma-Aldrich Chemical Company (St. Louis, MO): ampicill in, DEAE-Sephacel, Ultrogel AcA-44, mercaptoethanol, PLP, bovine serum albu min, succinyl-CoA, ALA-hydrochloride, ketoglutaric acid, -ketoglutarate dehydrogenase, HEPE S-free acid, MOPS, tricine, thiamine pyrophosphate, NAD+, and the bicinchoninic acid pr otein determination kit. Glucose, glycerol, glycine, disodium ethylenediamine tetraacetic acid dihydrate, ammonium sulfate, magnesium chloride he xahydrate, and pota ssium hydroxide were acquired from Fisher Scientific (Pittsburgh, PA). Sodium dodecyl sulfate polyacrylamide gel electrophoresis reagents we re acquired from Bio-Rad. Sal I, Blp I, Xho I, Bam HI restriction enzymes, Vent DNA Polymerase, and T4 DNA ligase were from New England Biolabs (Ipswich, MA). Oligonucleotides were synthesized by Integrated DNA Technologies (Coralville, IA). Methods Mutagenesis. The pGF23 expression plasmid enc odes the full-length sequence for the murine, mature eALAS (17). The R85L variant was ge nerated using a previously described method (18). The mutagenic primers used for the R85L mutation were 5GAC CAC ACC TAC CTT GTG TTC AAG ACT GT-3 and 5-ACA GTC TTG AAC ACA A GG TAG GTG TGG TC-3, with the intr oduced codon substitu tion underlined.
89 The PCR-generated fragment containing the R8 5L mutation was used as a megaprimer in an independent round of PCR. The PCR-genera ted fragment was sequenced to verify the presence of the desired mutations. Subse quently, the PCR product was then digested with Sal I and Bam HI and subcloned into pGF23 digest ed similarly. The site-directed mutagenesis for the T430V variant was perf ormed on the whole plasmid pGF23 using a previously described method (19). The mutagenic oligonucleo tide for T430V was: 5ATC AAC TAC CCA GTT GTG CCT CTG GGT-3, w ith the introduced codon substitution underlined. The PCR-generated DNA was sequenced between the Blp I and Bam HI restriction enzyme sites to confirm the presence of the mutation. The product was then digested with Blp I and Bam HI and subcloned into pGF 23 digested similarly. The pMAL2 expression plasmid, described above, encodes the full-length sequence for the murine, mature eALAS, with the arginine at position 85 mutated to leucine. The R85L/T430V double mutated variant was cons tructed by digesting the T430V-encoding plasmid (pTL30) with Xho I and Bam HI. This T430V mutation-encoding fragment was subcloned into pMAL2 digested similarly. Protein purification, SDS-PAGE, protein determination and steady-state analysis. Recombinant murine eALAS and the R85 and R85/T430 variants were purified from DH5 Escherichia coli bacterial cells containing the ove rexpressed protein as previously described (17). Sufficient expression of the T430V variant could not be obtained. The protein purity was over 90% judged by SDS-PAGE (20) and protein concentration was determined by the bicinchoninic acid method us ing bovine serum albumin as the standard (21). All protein concen trations are reported on the basi s of a subunit molecular weight
90 of 56 kDa. Enzymatic activity was determin ed by a continuous spectrophotometric assay at 30oC (22). Structural analyses. The protein data base files 2BWN, 2BWO, and 2BWP, corresponding to the R. capsulatus ALAS holoenzyme, succinyl-CoA bound, and glycine bound crystal structures were used as templates to model the the murine eALAS Michaelis complex structure (4). Hydrogen bond determinations were accomplished using Deepview/Swiss-PdbViewer software (23, 24). Circular dichroism spectroscopic measurements. Spectroscopic measurements were performed with enzyme that was dial yzed in 20 mM HEPES, pH 7.5 with 10% glycerol to remove free PLP. Circular dichroism (CD) spectra were obtained using an AVIV CD spectrometer calibrated for both wavelength maxi ma and signal intensity with an aqueous solution of D-10 camphorsulfonic acid (25). Protein concentrations were 100 M for each enzyme tested. The final conc entration of each CoA-derivative was 100 M, giving a 1:1 molar ratio of enzyme to ligand. At least three CD spectra were collected per experiment and averaged, using a 0.1 cm path length cuvette with a total volume of 300 l. Blank CD spectra contained all components of the solution except enzyme. CD spectra containing the enzyme sample were collected immediately after adding the enzyme. The spectra of the samp les containing enzyme were analyzed after subtracting the blank spectra. Stopped-flow spectroscopy. All of the experiments were carried out at 30C in 100 mM HEPES, pH 7.5 and 10% (v/v) glycerol. The concentration of reactants loaded into the two syringes was always 2-fold greater than that present in the cell compartment after mixing, with glycine and the enzyme pr e-incubated in one syringe and the CoA-
91 derivative in another. B ecause of the difference in Km values for the CoA-derivatives among the two enzymes tested, different CoAderivative concentrations were used to ensure the identification of a single enzyme catalyzed event. For the wild-type reaction, the final concentrations were: 120 M wild-type ALAS, and 130 mM glycine. The final concentrations of each independently exam ined CoA-derivative were: succinyl-CoA, 10 M; octanoyl-CoA, 10 M; butyryl-CoA, 20 M; -hydroxybutyryl-CoA, 30 M; glutaryl-CoA, 30 M. For the R85K-catalyzed reaction, the final concentrations were: 120 M R85K, and 130 mM glycine. The fina l concentrations of each independently examined CoA-derivative were: succinyl-CoA, 20 M; octanoyl-CoA, 10 M; butyrylCoA, 10 M; -hydroxybutyryl-CoA, 10 M; glutaryl-CoA, 20 M. Rapid scanning stopped-flow kinetic measurements were conducted using an OLIS model RSM-1000 stopped-flow spectrophotometer. The dead time of this instrument is approximately 2 ms, and the observation chamber optical path length is 4.0 mm. Scans covering the wavelength region 270-550 nm were acquired at a rate of 31, 7 or 3 scans per second in order to condense the resulting data files to a tr actable size for data fitting analysis. An external water bath was utilized to maintain constant temperature (30oC) of the syringes and observation chamber. Observed rate constants were determined by global fitting of the acquired spectral data sets, using the single value decomposition software provided by OLIS, Inc.(26) The quality of fits were judged by visual analysis of the calculated residuals in conjunction with the Durbin-Watson statistic (27). Single turnover data were interpreted using a three kinetic step mechanism as described by Equation 3.1. (Equation 3.1) D C B A k k k 3 obs 2 obs 1 obs
92 The observed rate constants were determined from at least three re plicate experiments, and the reported values represent the average and standard error of measurement for each experimental condition. Intrinsic protein fluorescence quenching. The pre-steady state kinetics of the product binding reaction of ALAS and the R85 and T430 variants were examined by measuring changes in the intrinsic protei n fluorescence intensity. An OLIS RSM-1000F rapid mixing spectrofluorimeter, equipped w ith a high-intensity xenon arc lamp, was used to follow the reaction. The enzyme and ligand in 20 mM HEPES (pH 7.5) and 10% glycerol were maintained at 30oC in separate syringes prior to their mixing in the reaction chamber. The concentrations of enzyme a nd ligand in the reaction chamber were 1/2 of those in the syringes. The intrinsic pr otein fluorescence, as measured with 5 M enzyme, was evaluated in the presence of increasing co ncentrations of the product, ALA. The excitation wavelength and the slit width were 280 and 5 mm, respectively. Scheme 3.1 illustrates the relationship between wavele ngth maximum and the dynamic process being monitored. The emitted light was filte red using a cutoff filter (WG 320; 80% transmittance at 320 nm, (Edmund Optics, Barrington, NJ)). Typically, 500 time points were collected for varying lengths of time, and three or more experiments were averaged. Each averaged data set was then fitted to Equation 3.2, using the global fitting software provided with the instrument. (Equation 3.2) 0 1) (A e A t Ft k obsobs where Fobs(t) is the observed fluorescence change (in arbitrary units) at time t, kobs1 is the observed first-order rate constant, A1 is the pre-exponen tial factor and A0 is the offset. The
93 observed rate constants were then plotted ag ainst ligand concentrati on and the data were fitted to Equation 3.2 by nonlinear regression. The rates of dissociation (koff) and association (kon) as well as the liga nd binding constants (KD) were calculated from the asymptotic maximal observed rate, the ordina te intercept, and the ligand concentration (x) in Equation 3.3 (Equation 3.3) x K x k k x fD off on ) ( Results Kinetic characterization of the R85 and R85/T430 variants. The steady-state kinetic parameters of the ALAS variants were determined and the results are summarized in Table 3.1. Wild-type ALAS was active with all of the CoA-derivatives tested. The Km for octanoyl-CoA was the lowe st with a value of 0.51 M compared to 2.9 M for succinyl-CoA. This decreased value contribute d to a catalytic efficiency value that was 2-fold higher than the reaction completed with the physiol ogical substrate succinyl-CoA. Glutaryl-CoA and -hydroxybutyryl-CoA were the least cat alytically efficient, due to 6fold increases in the Km for both substrates. Changing R85 to leucine (R85L) imparted dramatic changes with resp ect to the physiolo gical substrate succinyl-CoA. The kcat associated with the R85L-catalyzed react ion using succinyl-CoA as the substrate decreased greater than 11-fold, while the SCoA mK increased 7-fold. The 66-fold increase in catalytic efficiency toward octanoyl-CoA as well as the 4-fold increase found with butyryl-CoA for the R85L va riant highlight a shift toward acceptance of more hydrophobic CoA-derivatives within the acyl-Co A-binding cleft of this enzyme. The replacement of R85 with lysine (R85K) yiel ded a turnover number that was similar to
94 that of the wild-type enzyme. When the diffe rent acyl-CoA substrates were tested, this pattern continued with the exception of the reaction with glutaryl-CoA. This CoAderivative reacted slower than the physiological substrate succinyl-CoA as evidenced by a 21-fold decrease in kcat. Among the catalytic effici encies calculated for R85K, octanoyl-CoA yielded the highest with a value 13-fold greater than that of the same reaction containing succinyl-C oA. The steady-state data obtained from the double variant (R85L/T430V) suggest that steric hindrance as well as hydrophobicity of the active site are important determinants for preference of CoA-derivative. Affinity for octanoyl-CoA in the double variant is dimi nished, resulting in a 2-fold higher Km. Conversely, the reaction of the double variant with butyryl-CoA gave a Km value of 0.54 M, a value indistinguishable from octanoyl-CoA in the R85L variant, and 31-fold lower than octanoyl-CoA in the double variant. The double vari ant-catalyzed reaction with glutaryl-CoA, a derivative similar to su ccinyl-CoA but longer by one methylene carbon, showed undetectable activity as measured unde r the assay conditions tested. These data are shown graphically as normalized specificity constants in Figure 3.2, in which the ratio of the catalytic efficiency of each variant for a particular substrate is compared to the catalytic efficiency of the variant with succinyl-CoA.
95Table 3.1 Comparison of steady-state kine tic constants for wild-type ALAS, R85K, R85L, and R85L/T430V with CoA derivatives as substrates. Parameter Wild-Type ALAS R85K R85L R85L/T430V Succinyl-CoA as a Substrate kcat, min-1 10.0 0.2 6.4 0.2 0.94 0.10 0.11 0.03 CoA app mK,, M 2.9 0.1 12.4 0.6 20.3 1.0 9.6 0.8 kcat/CoA app mK,, min-1 M-1 3.6 0.2 0.53 0.04 0.050 0.003 0.010 0.002 Gly mK, mM 24.2 0.4 20.0 0.4 63.1 2.2 98.4 3.5 kcat/Gly mK, min-1mM-1 0.43 0.04 0.32 0.06 0.010 0.002 0.001 0.002 Octanoyl-CoA as a Substrate kcat, min-1 3.4 0.2 10.3 0.4 1.8 0.3 (1.0 0.1) x 10-3 CoA app mK,, M 0.51 0.04 1.5 0.04 0.55 0.03 17.2 0.6 kcat/CoA app mK,, min-1 M-1 6.8 0.8 7.0 0.2 3.3 0.1 (5.6 0.2) x 10-5 Gly mK, mM 17.1 0.9 25.2 1.1 55.2 5.0 74.2 3.3 kcat/Gly mK, min-1mM-1 0.14 0.01 0.52 0.02 0.030 0.005 (1.0 0.08) x 10-5 Butyryl-CoA as a Substrate kcat, min-1 6.3 0.3 10.2 0.3 2.01 0.08 0.060 0.002 CoA app mK,, M 6.1 0.09 2.7 0.1 9.3 1.0 0.54 0.03 kcat/CoA app mK,, min-1 M-1 1.0 0.1 3.7 0.4 0.21 0.04 0.03 0.002 Gly mK, mM 29.3 4.6 17.2 0.7 70.3 6.6 88.2 3.9 kcat/Gly mK, min-1mM-1 0.26 0.03 0.50 0.01 0.030 0.005 (6.1 0.8) x 10-4 -Hydroxybutyryl-Co A as a Substrate kcat, min-1 4.0 0.8 2.8 0.1 0.65 0.03 (1.0 0.2) x 10-4 CoA app mK,, M 9.8 1.0 5.5 0.2 6.1 0.8 74.2 0.6 kcat/CoA app mK,, min-1 M-1 0.41 0.04 0.51 0.04 0.11 0.02 (1.3 0.4) x 10-6 Gly mK, mM 22.1 0.8 18.2 3.2 59.1 4.7 92.2 6.0 kcat/Gly mK, min-1mM-1 0.17 0.06 0.14 0.03 0.011 0.003 (1.0 0.3) x 10-6 Glutaryl-CoA as a Substrate kcat, min-1 7.0 0.4 0.30 0.03 2.2 0.1 n/d CoA app mK,, M 17.0 1.6 7.5 0.6 30.1 1.1 n/d kcat/CoA app mK,, min-1 M-1 0.41 0.05 0.04 0.003 0.07 0.006 n/d Gly mK, mM 28.4 0.8 21.2 0.2 70.6 4.2 n/d kcat/Gly mK, min-1mM-1 0.29 0.02 0.010 0.002 (8.0 0.7) x 10-4 n/d n/d, not determined
96Circular dichroism spectroscopy. To verify whether the R85K, R85L or R85L/T430V amino acid substitu tions introduced substan tial changes in secondary structure, CD spectra in the far-UV region ( 200-270 nm) were record ed for the wild-type and variant enzymes (data not shown). All enzymes displayed similar CD spectra indicating that the introduced residue exchanges did not result in gross differences in the overall conformation of the ALAS protein. The enzyme-bound cofactor gives rise to a defined CD spectrum in the visible re gion (300-500 nm) because of anisotropic interactions between the amino acid side ch ains and the chromophore in the active site (Figure 3.3). The CD spectrum thus cont ains information about the asymmetric orientation of the bound PLP co factor in the active site, in cluding the aldimine linkage between the PLP cofactor and the active site lysine (Scheme 3.1)(28, 29). The visible CD spectra of the WT ALAS (Figure 3.3A) show th at in the presence of either succinylor octanoyl-CoA there is a positive Cotton effect. However, close insp ection of the results indicates that there is a distinct difference in the CD maximum in relation to that of the WT ALAS; the wavelength maximum for succi nyl-CoA blue shifts 15 nm to ~420 nm, while the octanoyl-CoA dichroic band at ~ 435 nm increases. The spectra observed for the R85K variant are different (Figure 3. 3B). The changes caused by succinyl-CoA binding to the R85K variant mimic those observed upon octanoyl-CoA interacting with WT ALAS, in that the spectral differen ces between the two are unremarkable. Substitutions of polar residues with non-polar amino acids yielded proteins (R85L and R85L/T430V) in which ligand binding induces changes in the PLP environment and, consequently, display CD spectra (visible regi on) that are markedly different from those of wild-type ALAS and the R85K variant. Within th e R85L enzyme, succinyl-CoA
97 binding does not cause a shif t in the microenvironment of the chromophore (Figure 3.3C). However, all of the remaining Co A-derivatives cause a negative Cotton effect, decreasing the amplitude of the spectra ~5 Steric hindrance and enhanced hydrophobicity of the acyl-CoA-binding clef t are hypothesized to be the most exaggerated in the double variant. This theory is supported by the CD data which suggest that R85L/T430V is affected most by the bi nding of octanoyland butyryl-CoA (Figure 3.3D). The positive Cotton effect observed in the spectra for the enzyme in the presence of these two hydrophobic ligands co uld suggest that the exclusi on of water and degree of active site alipha ticity are crucial components for de termining substrat e specificity. Overall, a comparison of the CD spectra of the wild-type and R85K variant enzymes with those of the hydrophobic variants (R85L and R 85L/T430V) show that th e distinct change in the microenvironment of the PLP pocket correlates well with the degree of hydrophobicity inherent to the bound ligand.
98Figure 3.2. Comparison of normalized specificity constants for murine eALAS variants with different CoA substrates. The specificity constant (kcat/CoA app mK,) with succinyl-CoA as substrate was arbitrarily define d as equal to 1.00 for each ALAS variant. Scheme 3.1. The absorbance maxima of ch emical species in the ALAS-catalyzed reaction. R=OP 2 3O ALAS Variant wtR85KR85LR85L/T430V Normalized Specificity Constant 0 5 10 15 60 65 70 Succinyl-CoA Octanoyl-CoA Butyryl-CoA Hydroxybutyryl-CoA Glutaryl-CoA
99 Figure 3.3. Visible circular dichroism spec tra of wild-type ALAS and the R85 and R85/T430 variants. (A) wild-type ALAS (B) R85K, (C) R85L, and (D) R85L/T430V. Spectra of the holoenzymes are in purple. Spectra of the holoenzymes (100 M) in the presence of 100 M CoA derivative are indicated according to the following color scheme: pink, succinyl-CoA; green, octanoyl-CoA; blue, -hydroxybutyryl-CoA; black, butyryl-CoA; red, glutaryl-CoA.
100 ALA-binding kinetics monitored by the transient intrinsic protein fluorescence. To determine whether a change in the de gree of hydrophobicity of the acyl-CoA-binding pocket could influence the product binding and concomitant quenching of the intrinsic protein fluorescence, wild-type ALAS and the enzyme variants were rapidly mixed with excess ALA and the changes in intrinsic pr otein fluorescence were monitored. The observed pseudo first-order decay rates of these kinetic tr aces were dependent on ALA concentration, and differed among the enzyme s tested (Figure 3.4) In all cases the change in observed rate as a function of ALA concentration wa s hyperbolic, indicating two binding steps. Presumably these two step s are indicative of ra pid formation of an initial collision complex followed by a slower shift to the closed conformation observed in the crystal structures of the R. capsulatus enzyme. While the resolved ALA off rates (i.e. k-1) coincide with the kcat values determined through steady-state kinetics, suggesting that a conformational change associated with ALA release defines kcat for each enzyme, the collision complex dissociation constants of the variants are increased. The resolved on and off rates for the reaction betw een ALA and wild-type ALAS were k1 = 0.120 0.015 s-1 and k-1 = 0.140 0.005 s-1, respectively, with a KD of 500 16 M. The rates for the reaction of the R85K va riant with ALA were ~2-fold lower with k1= 0.090 0.003 s-1 and k-1 0.079 0.003 s-1; however, the dissocia tion constant was 3fold higher at 1470 30 M, indicating decreased affinity for ALA. Among the more hydrophobic variants, the rates of protein quenc hing with ALA were the most rapid for R85L with on and off rates of 0.039 0.009 s-1 and 0.024 0.003 s-1, respectively, values 3-fold lower that those of the wild-type enzyme. A KD value of 1130 27 M corresponding to a 3-fold greater dissociation of ALA from the R85L variant than that of
101 wild-type ALAS was calculated. The double variant bound ALA le ss tightly, with a KD 13-fold lower than that for wild-type ALAS and an estimated value of 6710 33 M. Figure 3.4. Reaction of wild-type ALAS, R85K, R85L and R85L/T430V (5 M) with ALA. The observed rate constants were calcula ted by fitting the decrease in intrinsic protein fluorescence over time to Equation 3.2 fo r a single exponential pr ocess. (A) wildtype ALAS (B) R85K (C) R85L (D) R85L/T430V.
102 The on and off rates for R85L/T430V were also the lowest of the enzymes tested, with rates measuring k1= 0.007 0.0001 s-1, and k-1 0.005 0.001 s-1, respectively Pre-steady-state reaction of the variant enzyme-glycine comp lexes with acyl-CoAderivatives. The R85L and R85L/T430V variant enzyme-glycine complexes did not yield a measurable absorbance change at 510 nm, previously assigned to a quinonoid reaction intermediate (Scheme 3.1), when rapi dly mixed in the presence of any of the 5 acyl-CoA derivatives tested (16, 30, 31). Consequently, the investigation of the transient kinetics associated with the formation and decay of the quinonoid intermediate was based upon the reactions catalyzed by wild-type ALAS and the R85K variant, both of which demonstrate a quantifiable absorbance change at 510 nm upon the addition of the acylCoA substrates to the enzyme-glycine complexe s. The rates associated with the lifetime of the quinonoid intermediate, measured du ring the enzyme catalyzed reactions, were elucidated (Table 3.2). The absorbance change timecourses were fitted to a sequential, three-step mechanism outlined by Equati on 4.1. An initial burst of quinonoid intermediate formation, followed by a two-step decay was characteri stic of each of the enzymes tested (Figure 3.5). Of all the acyl-CoA derivatives examined, only hydroxybutyryl-CoA, when rapidly mixed with the wild-type enzyme-glycine complex, failed to produce an absorbance change at 510 nm. Overall, faster rates of quinonoid intermediate formation were observed for the wild-type enzyme, as compared to the R85K variant (Table 3.2). The rates of quinonoid formation when octanoyl-CoA was used as the substrate (2.3 s-1 and 1.9 s-1) were of the same order of magnitude as the values recorded for succinyl-CoA (6.0 s-1and 4.2 s-1) for both the wild-type and R85K
103Table 3.2. Rates of quinonoid intermed iate formation and decay under singleturnover conditions. Parameter succinyl octanoyl butyryl -hydroxybutyryl glutaryl Wild-type ALAS (s-1) (s-1) (s-1) (s-1) Qf 6.0 0.6 2.3 0.4 0.41 0.04 n/d 0.220 0.03 Qd1 2.00 0.30 0.071 0.004 0.070 0.005 n/d 0.091 0.005 Qd2 0.072 0.005 0.020 0.003 0.011 0.003 n/d 0.0010 0.0004 R85K (s-1) (s-1) (s-1) (s-1) (s-1) Qf 4.20 0.1 1.91 0.20 0.323 0.020 0.121 0.021 0.192 0.042 Qd1 1.10 0.1 0.11 0.02 0.002 0.0001 0.022 0.005 0.134 0.071 Qd2 0.050 0.004 0.070 0.003 0.041 0.007 0.013 0.001 0.0010 0.0001 n/d, not determined; Qf, quinonoid intermediate formation; Qd1, first step of quinonoid intermediate decay; Qd2, second step of quinonoid intermediate decay
104 Figure 3.5. Reaction of wild-type AL ASand R85K-glycine complexes with different CoA derivatives under single turnover conditions. The data ( / ) are overlaid with the line representing the best-fit curve ( ). The rate constants for the three step sequence corresponding to both enzymes are listed in Table 3.2. Green and red data points correspond with wild-type ALAS and the R85K variant, respectively. (A) octanoyl-CoA, (B) butyryl-C oA, (C) succinyl-CoA, (D) glutaryl-CoA, (E) hydroxybutyryl-CoA.
105 enzymes, respectively. These data support the increased catalytic efficiency observed from the experiments performed in the steadystate. Further comparison of the reaction catalyzed by wild-type ALAS with butyryl-CoA vs. glutaryl-CoA showed that quinonoid intermediate formation was accelerated 90%. A similar enhancement was observed for the R85K-catalyzed reaction which showed a 70% increase in th e rate of quinonoid intermediate formation with butyryl-CoA vs. glutaryl-CoA. The preference for butyrylCoA over glutartyl CoA suggests that in ad dition to the hydrogen bonding properties of R85 and T430, the amino acids that line th e hydrophobic tunnel leadin g to the terminal guanidino group may play a role in substrate acceptance and orientation. Curiously, only the R85K-glycine complex, when rapidly mixed with -hydroxybutyryl-CoA, gave a time-dependent absorbance change at 510 nm and rate associated with quinonoid intermediate formation (0.12 s-1). This is in stark contrast to the observations made of wild-type ALAS, where no quantifiable change with this substrate was detected. This slow rate may be explained by the mixed polarity of the substrate tail, an attribute which simultaneously imparts hydrogen bonding character, as well as aliphaticity to the acylCoA-binding cleft of the variant enzyme.
106Discussion The reactions catalyzed by the highly related members of the -oxoamine synthase subfamily of PLP-dependent enzyme s can be compared with respect to the specificity of the acyl-CoA substrate due to the elucidation of the three-dimensional structures of subfamily members together with mutagenesis, sp ectroscopic and kinetic methods (1, 4-6, 32). Both of the variant enzymes c onstructed for the arginine residue (R85L and R85K) as well as the doubly muta ted enzyme (R85L/T430V) were expressed, overproduced, and then purified as holoenzymes, indicating that cof actor binding by the apoprotein was not affected by the introduc tion of the amino acid substitutions. However, replacement of the invariant threonine residue with valine (T430V) resulted in a poorly expressed, unstable, and proteolyti cally susceptible enzyme that was never purified to homogeneity (data not shown). All of the purifiable variants were active with the physiological substrate succ inyl-CoA. Since the threonin e to valine replacement at position 430 appears to dramatically affect pr otein stability, we suggest that T430 is essential not only for optimal molecular recogn ition of succinyl-CoA, but also for stable folding. R85 may be less crucial for prope r enzyme function, a finding supported by the crystallographic data for SPT (6). Given that this enzyme lacks the arginine residue implicated in salt bridge formation with the carboxylate group of CoA substrates in the other three members of the -oxoamine synthase subfamily and utilizes palmitoyl-CoA, an acyl-CoA derivative of increased aliphaticity, it is proposed that acyl-CoA binding in ALAS may be driven by non-covalent intera ctions between the two residues and the substrate. However, considering the struct ural and mechanistic data for ALAS and SPT,
107 turnover likely remains orchestrated by amino acids that are proxim al to the site of carbon bond scission (16, 33). Comparison of the active sites of AONS a nd ALAS showed that the coordination of the acyl-CoA substrate is assisted by way of pantetheine associat ion with the enzyme face and tail interactions with the buried hydrophobic tunnel (1, 4, 6). Both R85 and T430 coordinate the carboxylate tail of the acyl-CoA substrate in meALAS. The steadystate kinetic analysis of th e variants (R85K, R85L, and R 85L/T430V) with the family of CoA-derivatives showed that the apparent Michaelis parameters (CoA app mK,) are dramatically different when compared to those of wild-type ALAS. Acyl-CoA substrates of increased hydrophobicity (e.g., octanoyland butyryl-CoA) demonstr ated greater affinity for the variants where the substituted amino acid was aliphatic in nature (R85L and R85L/T430V). The 36-fold decrease in the (CoA app mK,) for octanoyl-CoA in the R85L variant leads us to s uggest that the exclus ion of water from the acyl-CoA-binding tunnel is a determining feature of substrate binding. Further, in the double variant, an 18-fold reduction in the Michaelis cons tant for butyryl-CoA also s upports this hypothesis. The introduction of valine at position 430 w ould reduce the diameter of the hydrophobic tunnel, making steric hindrance a more signifi cant consideration for substrate binding. These differences identified between the va riant enzymes and wild-type ALAS suggest that reaction specificity is driven by the chem ical characteristics of the CoA-derived tail and the hydrogen-bonding potential of the i nvariant acyl-CoA-binding residues, a phenomenon recognized in the acyl-CoA thioesterases of the peroxisome (34, 35). The chemical characteristics of the acylCoA tail are a determining factor in the substrate specificity of another family of enzymes that utilize related substrates in
108 turnover, the crotonase family (36, 37). Among those enzymes, octanoyl-CoA has been shown to bind in a charact eristic bent conformation (36). This substrate conformation is accomplished by two structurally conserve d hydrogen bond-donating groups to the carbonyl moiety of the substrat e and through the entropical ly driven loss of water coordinated by the hydrophobic amino aci ds that line the binding cleft (36). The CD data for butyryl-CoA with the double variant in addition to the substrate configuration observed in enzymes that physio logically utilize octanoyl-Co A led us to suggest that octanoyl-CoA, which differs from butyryl-CoA by a four methylene bridge, most likely bends in the ALAS active site This hypothesis is further supported by the data obtained for both wild-type ALAS and R85L with octanoyl-CoA. For wild-type ALAS, the catalytic efficiency for the non-physiological substrate octanoyl-CoA is ~100% greater than that of succinyl-CoA (Table 3.1 and Figure 3.2). Congruently, the specificity constant for octanoyl-CoA compared to that of succinyl-CoA in th e R85L variant is 66fold higher, indicating a signi ficant change in substrate sp ecificity. Therefore, we hypothesize that octanoyl-CoA, devoid of a salt bridge to anchor with the guanidino group of R85, likely bends, excludes water fr om the active site, a nd assists in enzyme closure, a conformation postulate d to be essential for turnover (16). Interestingly, the specificity constant for the wild-type enzyme with octanoylCoA is higher than the physiol ogical substrate, succinyl-CoA, al beit at the cost of a threefold reduction in the specificity constant for the other substrate, glycine. Nevertheless, this finding and the activity of the enzyme w ith the other CoA esters tested here lead us to raise the question as to what extent ALAS may catalyze formation of 1,-3aminoketones other than ALA in vivo. This is currently unknown, but may warrant
109 further investigation. Conceivably, the poten tial toxicity, associated with the generation of other aminoketones rather than ALA, c ould be minimized through the action of a regulatory acyl-CoA-binding protein. In fact studies have demonstrated that the acylCoA binding protein binds long-ch ain acyl-CoA esters with hi gh specificity and affinity (with Kd values of 110 nM); hence by interactin g with acyl-CoA utilizing enzymes, the acyl-CoA-binding protein may provide a mechan ism for control of free acyl-CoA esters and regulation of the activity of acyl-CoA utilizing enzymes. Further substrate specificity in vivo might be enhanced via a substrate channeling mechanism involving interaction of ALAS with succinyl-CoA synthetase (38). The requirement for such a mechanism is emphasized by the evidence presented here. The chromophoric properties of PLP in ALAS provide a valuable probe for positional alterations to the amino acids that comprise the cofactor binding cleft. Since PLP is not a chiral molecule, the Cotton eff ect of PLP bound in the active site must result from certain asymmetric distortion of the PLP molecule through interaction with the enzyme. Binding of acyl-CoA substrates to ALAS (or ALAS variants) most likely induced changes, even if subtle, in the PL P-protein interaction, as reflected by the different visible CD spectra (Fig. 3). Curious ly, for each of the enzymes tested, the most noticeable spectral deviations from the spectra obtained in the absenc e of substrate were produced by CoA-derivatives with tails that matched the chemical nature of the amino acid substitution introduced in ALAS. In the R85L/T430V double va riant, binding of either octanoyland butyryl-CoA induced a ma rked positive Cotton effect (Figure 3.3). A definite interpretation for the relationship between substrate binding and induced Cotton effect of the PLP cofact or cannot be provided at this point. Perhaps the observed
110 effects could be explained, in part, by a shift in the enzyme conformational equilibrium towards the closed conformation, which has been hypothesized for the wild-type enzyme to be energetically driven by succinyl-CoA binding (Lendrihas et al., unpublished data) (16, 18). However, a decrease in active site diameter, triggered by the entropic loss of water upon the binding of a more hydrophobic substr ate, could also expl ain the change in the cofactor microenvironment as evidenced by the CD data. This scenario is supported by ligand binding to the asymmetric protein host. It is therefore premature to assign the observed CD spectral differences to a particular molecular event (e.g., a protein conformational change, a PLP re orientation or a perturbation of the electronic system of the chromophore). Nevertheless, the CD sp ectra in the UV region of all the variants were indistinguishable from that of the w ild-type enzyme (data not shown), indicating no gross alterations in secondary structure, and thus suggesting that the differences observed in the visible CD spectra, upon substrate bindin g, are confined to th e PLP-binding cleft. Although specific contributions by single amino acid substitutions cannot be easily disentangled using CD, the CD spectral differences in th e visible range among ALAS variants with the family of CoA-derivativ es of different hydrophobicities lead us to propose that interactions between key residues (e.g., R85 and T430) that bind the tail of CoA play a role in determining substrate specificity. The proposed kinetic mechanism of the AL AS-catalyzed reaction is limited by product release, or opening of the active s ite loop coincident w ith product release (16). Utilization of protein fluorescence to study the reverse ca talytic reaction, i.e., the reaction of the enzyme with the product, resolves the ALA off rate, and confirms it to be indistinguishable from kcat (16). With this in mind, we investigated whether mutations to
111 the acyl-CoA-binding residues affect the rate s of product release and/or perturb the enzyme-product equilibrium (Figure 3.4). Quen ching of the ALAS in trinsic fluorescence with ALA obeyed first-order decay kinetics. For all the variant catalyzed-reactions, rates of product release and capture are diminished. The most dramatic decrease is seen in the double variant; this enzyme exhibited a 13-fold reduction in the on and off rates (kon and koff) for the reaction with ALA. Since Co A-derivatives of in creased hydrophobicity bind with greater affinity to the non-polar variants (R85L a nd R85L/T430V), it is possible that the decreased affinity for the product, a -aminoacid, may be reversed for aminoketones of decreased polarity. In this scenario and in agreement with a mechanism in which a conformational step follows ligand bindi ng, the affinity of the aminoketones towards ALAS would not be reduced. The Kd values would either increase or remain unchanged; this hypothesis, how ever, awaits further experimentation with aminoketones of differing hydrophobicity. The three-dimensional structures of ALAS and AONS revealed a hydrogen bonding network between the in variant arginine and th reonine residues and the carboxylate moieties of acyl-CoA substrates (1, 4). Accordingly, how the use of chemically different acyl-CoA-derivatives w ould affect the transient kinetic parameters of the enzymes was evaluated. Single tu rnover reactions with a family of CoAderivatives were used to determine the ra tes of quinonoid intermediate formation and decay. The reaction catalyzed by wild-type AL AS with glutaryl-CoA as the substrate exhibited a 30-fold lower rate corres ponding to quinonoid intermediate formation vs. the rate calculated with succinyl-C oA. One possible explanation for the retarded rate is that the binding of glutaryl-CoA affects the hydrophobic acyl-Co A-binding tunnel in such a
112 way that cofactor-mediated electron transf er from the site of bond scission to the resonance stabilized pyridinium ring of PLP is perturbed. This phenonmenon is further supported by the dramatic 70-fold reduction in the second step of quinonoid intermediate decay, where the absorbance change at 510 nm appears to stabilize to a level that approaches a steady-state (Figure 3.5D). The R85K variant demonstrated a change in the first step of quinonoid intermediate decay, wi th a 10-fold lower rate for both octanoylCoA and glutaryl-CoA when compared to the physiological substr ate succinyl-CoA. When R85 is mutated to a lysine, the enzyme is chemically similar to wild-type ALAS in many respects, presumably because this conservative replacement retains the positive charge and hydrogen bonding cap abilities. In addition, in silico protonation of the lysine amino group to create the -ammonium charge center would contribute charge and polar interactions characteristic of the guanidinium group (23, 24). However, the molecular volume of the amino acid side chain is different, as is the electrostatic charge distribution. With respect to their n-alkyl moieties, the n-propylguanidine side ch ain of arginine is longer than the n-butylamine side chain of lysine by 1.6 (40). The R85K substitution could therefore accommodate the additional sp3 hybridized carbon atom present in glutaryl-CoA, allowing for a re duction in steric strain and/ or unfavorable Van der Waals interactions. Further, the increased hydr ophobic tunnel length could also assist the bending of octanoyl-CoA within the cleft, a circumstance rationalized above. In summary, the spectroscopic and kinetic studies detailed here demonstrate not only the role played by R85 a nd T430 in determining acyl-CoA substrate specificity, but also provide insight into the structurefunction relationships in ALAS and the oxoamine synthase subfamily as a whole. Although R85 and T430 recognize a part of
113 the substrate that is distal from the bulk of the ligand and the active si te, the ability of all the enzymes to turnover non-physiological substrates remains intact. Changing these residues to leucine and valine abates succinyl -CoA binding and catalytic efficiency, as well as increases the affinity of the enzyme for CoA-derivatives of greater aliphaticity. These observations indicate that a mutationmediated decrease in substrate binding energy could be accountable for the enhanced affinity measured for the hydrophobic CoA-derivatives, instead of a direct mechan istic linkage between these residues and the site of bond cleavage. Cert ainly, the conserved amino acid duo (R85 and T430) are at some distance from the PLP-binding site, and therefore the coupling of substrate binding to di-carbon cleavage in the active site presum ably involves coordinated movement of the enzyme upon acyl-CoA binding. However, further experiments to prove this await the development of tractable fluorescent probe s and the elucidation of three-dimensional crystal structures with CoA-derivatives bound.
114Acknowledgements This work was supported by the National Institutes of Health (grant DK63191 to GCF). References (1) Webster, S. P., Alexeev, D., Campopi ano, D. J., Watt, R. M., Alexeeva, M., Sawyer, L., and Baxter, R. L. (2000) Mechanism of 8-amino-7-oxononanoate synthase: spectroscopic, kinetic, and crystallog raphic studies. Biochemistry 39, 516-28. (2) Alexeev, D., Alexeeva, M., Baxter, R. L ., Campopiano, D. J., Webster, S. P., and Sawyer, L. (1998) The crys tal structure of 8-ami no-7-oxononanoate synthase: a bacterial PLP-dependent, acyl-CoA-condensing enzyme. Journal of Molecular Biology 284, 401-19. (3) Ikushiro, H., Hayashi, H., and Kaga miyama, H. (2004) Reactions of serine palmitoyltransferase with serine and mo lecular mechanisms of the actions of serine derivatives as inhibitors. Biochemistry 43, 1082-1092. (4) Astner, I., Schulze, J. O., van den He uvel, J., Jahn, D., Schubert, W. D., and Heinz, D. W. (2005) Crystal structure of 5-aminolevulinate synthase, the first enzyme of heme biosynthesis, an d its link to XLSA in humans. EMBO J. 24, 3166-3177. (5) Eliot, A. C., and Kirsch, J. F. (2004 ) Pyridoxal phosphate en zymes: mechanistic, structural, and evolutionary considerations. Annu. Rev. Biochem. 73, 383-415. (6) Yard, B. A., Carter, L. G., Johnson, K. A., Overton, I. M., Dorward, M., Liu, H., McMahon, S. A., Oke, M., Puech, D., Barton, G. J., Naismith, J. H., and Campopiano, D. J. (2007) The structure of serine palmitoyltransferase; gateway to sphingolipid biosynthesis. J. Mol. Biol. 370, 870-886. (7) Eisenberg, M. (1987) Biosynthesis of biotin and lipoic acid, Vol. 1, Washington D.C. (8) Hanada, K. (2003) Serine palmitoyltran sferase, a key enzyme of sphingolipid metabolism. Biochim. Biophys. Acta 1632, 16-30. (9) Bell, S. C., and Turner, J. M. (1976) Bacterial catabolism of threonine. Threonine degradation initiated by L-th reonine-NAD+ oxidoreductase. Biochem. J. 156, 449-458. (10) Merrill, A. H., Jr. (198 3) Characterization of serine palmitoyltransferase activity in Chinese hamster ovary cells. Biochim. Biophys. Acta 754, 284-291. (11) Han, G., Gable, K., Yan, L., Allen, M. J., Wilson, W. H., Moitra, P., Harmon, J. M., and Dunn, T. M. (2006) Expression of a novel marine viral single-chain serine palmitoyltransferase and construc tion of yeast and mammalian single-chain chimera. J. Biol. Chem. 281, 39935-39942. (12) Shoolingin-Jordan, P. M., LeLean, J. E., and Lloyd, A. J. (1997) Continuous coupled assay for 5-amin olevulinate synthase. Methods Enzymol. 281, 309-316. (13) Hunter, G. A., Rivera, E., and Fe rreira, G. C. (2005) Supraphysiological concentrations of 5-aminolevulinic acid dimerize in solution to produce
115 superoxide radical anio ns via a protonated dihydropyrazine intermediate. Arch. Biochem. Biophys. 437, 128-137. (14) Onuki, J., Teixeira, P. C., Medeiros, M. H., Dornemann, D., Douki, T., Cadet, J., and Di Mascio, P. (2002) Is 5-aminolevulinic acid involved in the hepatocellular carcinogenesis of acute intermittent porphyria? Cell Mol. Biol. (Noisy-le-grand) 48, 17-26. (15) Shoolingin-Jordan, P. M., Al-Daihan, S., Alexeev, D., Baxter, R. L., Bottomley, S. S., Kahari, I. D., Roy, I., Sarwar, M., Sawyer, L., and Wang, S. F. (2003) 5Aminolevulinic acid synthase: mech anism, mutations and medicine. Biochim. Biophys. Acta. 1647, 361-366. (16) Hunter, G. A., Zhang, J., and Ferrei ra, G. C. (2007) Tran sient kinetic studies support refinements to the chemical and kinetic mechanisms of aminolevulinate synthase. J. Biol. Chem. 282, 23025-23035. (17) Ferreira, G. C., and Dailey, H. A. (1993) Expression of mammalian 5aminolevulinate synthase in Escherichia coli. Overproduction, purification, and characterization. J. Biol. Chem. 268, 584-590. (18) Gong, J., Hunter, G. A., and Fe rreira, G. C. (1998) Aspartate-279 in aminolevulinate synthase a ffects enzyme catalysis th rough enhancing the function of the pyridoxal 5'phosphate cofactor. Biochemistry 37, 3509-17. (19) Miyazaki, K., and Takenouchi, M. (2 002) Creating random mutagenesis libraries using megaprimer PCR of whole plasmid. Biotechniques 33, 1033-1034, 10361038. (20) Laemmli, U. K. (1970) Cleavage of struct ural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680-5. (21) Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk, D. C. (1985) Measurement of protei n using bicinchoninic acid. Anal. Biochem. 150, 7685. (22) Hunter, G. A., and Ferreira, G. C. (1995) A continuous spectrophotometric assay for 5-aminolevulinate synthase th at utilizes substrate cycling. Anal. Biochem. 226, 221-224. (23) Schwede, T., Kopp, J., Guex, N., and Peitsch, M. C. (2003) SWISS-MODEL: An automated protein homology-modeling server. Nucleic Acids Res. 31, 3381-3385. (24) Guex, N., and Peitsch, M. C. (1997) SWISS-MODEL and the Swiss-PdbViewer: an environment for comparative protein modeling. Electrophoresis 18, 2714-23. (25) Chen, G. C. Y., J.T. (1977) Two-point calibration of cicular dichrometer with D10-camphosulphonic acid. Analytical Letters 10, 1195-1207. (26) Tsai, M. D., Weintraub, H. J., Byrn, S. R., Chang, C., and Floss, H. G. (1978) Conformation-reactivity rela tionship for pyridoxal Schiff's bases. Rates of racemization and alpha-hydrogen exchange of the pyridoxal Schiff's bases of amino acids. Biochemistry 17, 3183-3188. (27) Durbin, J., and Watson, G. S. (1970) Te sting for serial correla tion in least squares regression. Biometrika 37, 409-414. (28) Schnackerz, K. D., Tai, C. H., Potsch, R. K., and Cook, P. F. (1999) Substitution of pyridoxal 5'-phosphate in D-serine dehydratase from Escherichia coli by
116 cofactor analogues provides informati on on cofactor binding and catalysis. J. Biol. Chem. 274, 36935-36943. (29) Moscowitz, A. (1961) Some applications of the kronig-kramer s theorem to optical activity. Tetrahedron 13, 48-54. (30) Hunter, G. A., and Ferreira, G. C. (1999) Pre-steady-state reaction of 5aminolevulinate synthase. Evidence fo r a rate-determining product release. J. Biol. Chem. 274, 12222-12228. (31) Zhang, J., and Ferreira, G. C. (2002) Transient state kinetic investigation of 5aminolevulinate synthase reaction mechanism. J. Biol. Chem. 277, 44660-44669. (32) Schmidt, A., Sivaraman, J., Li, Y., La rocque, R., Barbosa, J. A., Smith, C., Matte, A., Schrag, J. D., and Cygler, M. (2001) Three-dimensional structure of 2-amino3-ketobutyrate CoA ligase from Escherichia coli complexed with a PLP-substrate intermediate: inferred reaction mechanism. Biochemistry 40, 5151-5160. (33) Dunathan, H. C. (1966) Conformati on and reaction specificity in pyridoxal phosphate enzymes. Proc. Natl. Acad. Sci. U S A 55, 712-716. (34) Hunt, M. C., Solaas, K., Kase, B. F., and Alexson, S. E. (2002) Characterization of an acyl-coA thioesterase that functi ons as a major regulator of peroxisomal lipid metabolism. J. Biol. Chem. 277, 1128-1138. (35) Hunt, M. C., and Alexson, S. E. (2008) Novel functions of acyl-CoA thioesterases and acyltransferases as auxiliary enzymes in peroxisomal lipid metabolism. Prog. Lipid Res. 47, 405-421. (36) Engel, C. K., Kiema, T. R., Hiltune n, J. K., and Wierenga, R. K. (1998) The Crystal Structure of Enoyl-CoA Hydr atase Complexed with Octanoyl-CoA Reveals the Structural Adaptations Re quired for Binding of a Long Chain Fatty Acid-CoA Molecule. J. Mol. Biol. 275, 859-847. (37) Engel, C. K., Mathieu, M., Zeelen, J. P., Hiltunen, J. K., and Wierenga, R. K. (1996) Crystal structure of enoyl-coenz yme A (CoA) hydratase at 2.5 angstroms resolution: a spiral fold de fines the CoA-binding pocket. Embo J 15, 5135-5145. (38) Furuyama, K., and Sassa, S. (2000) In teraction between succ inyl CoA synthetase and the heme-biosynthetic enzyme ALAS-E is disrupted in sideroblastic anemia. J. Clin. Invest. 105, 757-764. (39) Kelly, S. M., Jess, T. J., and Price, N. C. (2005) How to study proteins by circular dichroism. Biochim. Biophys. Acta. 1751, 119-139. (40) Creighton, T. R. (1983) Proteins, Structures a nd Molecular Properties, W.H. Freeman and Company, New York.
117 Chapter Four Hyperactive enzyme variants engineered by synthetically shuffling a loop motif in murine 5-aminolevulinate synthase Abstract The regulatory step of the heme biosynthe tic pathway in mammals is catalyzed by the pyridoxal 5'-phosphate -dependent enzyme, 5-aminolevul inate synthase (EC 188.8.131.52). Aminolevulinate is biosynthesized by conde nsing succinyl-CoA a nd glycine to yield coenzyme-A and carbon dioxide. A conser ved active site lid wa s shown to change conformation 3.5 between the holoenzy mic form and succinyl-CoA-bound forms of Rhodobacter capsulatus ALAS. We employed synthetic shuffling and preand steadystate kinetic analyses to determine the role of the lid motif in the ALAS-catalyzed reaction. Functional variants containing mutations to resi dues that comprise the lid (Y422-R439) were isolated based on genetic complementation in Escherichia coli strain HU227 and fluorescence microscopy. All of the positive isolates showed a spectrum of amino acid substitutions, a finding which validates our screening method. Each of the lid variants examined showed increases in kcat and catalytic efficiency with both substrates, observations which support a cruc ial role for the active site li d in product tu rnover. The single turnover reaction data for the shuffled variants reveal th at this lid has an important role quinonoid intermediate decay and product release. Energetically favorable
118 thermodynamic activation parameters for a libra ry isolate also sugge st that the entire active site lid is involve d in stabilizing the r eaction coordinate. Over all, our data support a hypothesis whereby the lid closes over the active site during catalysis; once chemistry has taken place, lid dynamics determine the rate of product release.
119Introduction Aminolevulinate (ALA) is the univers al building block of tetrapyrolle biosynthesis (1). In eucaryotes and the -subclass of purple bact eria the production of ALA is catalyzed by 5-aminolevulinat e synthase (ALAS) (EC 184.108.40.206) (2). ALAS is classified as a fold-type I pyridoxal 5phosphate (PLP)-dependent enzyme, and like the evolutionarily related transaminases (3) functions as a homodimer, with a PLP cofactor bound at each of the two active sites, which occur in clefts at the subu nit interface (4). The reaction catalyzed by ALAS is the Clai sen-like condensation of succinyl-CoenzymeA (CoA) and glycine to yield CoA, carbon dioxide (CO2) and ALA. This reaction type, coupled with the structural information about the protein, include it as a member of the oxoamine synthase subfamily of PLP-depende nt enzymes; a family which includes 7amino-8-oxononanoate synthase (AONS) (5). Genetic defects in the gene corresponding to the erythroid specific isoform of the enzyme are associated with a congenital disorder, X-linked sideroblastic anemia (XLSA) (6). X-ray crystal structures of ALAS and AONS reveal that they share similar active site geometry and proximal active site motifs (7, 8). Whereas the three dimensional structure of ALAS contains the substrate succinyl-CoA the AONS structure was cocrystallized with the product 7-amino8-oxononanoate (AON). In both cases, each enzyme has an active site lid, comprised of amino acids which stretch from one hinge of the lid to other, with the apex of the lid coordinating the ca rboxylate group of the ligand. Comparison of holoenzymic ALAS and AO NS, with the substrate and product bound forms of the enzymes indicate that bindi ng of these ligands within the active site precipitates active site lid dyna mics that may signify a change in conformation. The
120 active site lid of ALAS contai ns 18 residues; 8 of these are completely conserved, while 10 vary considerably according to nature. One corresponding residue in murine erythroid ALAS (mALAS2) that is positioned at the ap ex of the lid, T430, generates pathological affects associated with XLSA when mutated in humans (9). This residue appears crucial for catalysis and may be an important determ inant of substrate specificity (Lendrihas et. al, in press) (7). Two -sheets flank the position of the lid ( 12 and 13) in the Cterminal domain of the enzyme, where furthe r interactions between the carboxy terminal portion of the lid occur between two -helices on the adjacent subunit ( 2 and 3). While evidence is available for the role of discrete active site residues in ALASfunction, no studies have been carried out on the role of the active site lid that surrounds the site of catalysis and interacts with the acyl-CoA substrate (10-12). One glycine-rich loop identified in mALAS2 formed by residues 142-154, sandwiched between 3 and 4 was proposed to be involv ed in cofactor binding (13). Several residues of this sequence (G142, G144 and R149) did not tolerate mutation (13). However, mutations to residues N150 and I151 in the same study generated vari ants with turnover num bers greater than that of wild-type ALAS. Further, this loop is implicated in su ccinyl-CoA binding, and based on the position of these residues deep within the th ree dimensional structure, mutations are likely to acutely disrupt the mobility of the motif (7). The importance of the active site lid in the condensation mechanism of an acylCoA substrate and amino acid dono r was first identified in AONS (8). According to the X-ray crystal structures the -sheet of the C-terminal dom ain undergoes a conformational transition. This conformational change incl udes the 5.5 displacemen t of the active site lid. From the ALAS crystal structure with succinyl-CoA bound the change in position of
121 the outermost top of the lid moves inward by 3.5 when compared to the position of the lid in the holoenzymic form (Figure 4.1) (7). Accordingly, the poor electron density evidenced in the corresponding AONS motif coupled with the non-covalent interactions identified between the lid and the residues that comprise the active site indicate that these active site lids are probably disordered during unliganded conditions, a circumstance likely reversed in the presence of substrate or product. In order to examine the role of the active site lid in the ALAS-catalyzed reaction, we used synthetic shuffling to identify f unctional amino acid mutati ons and to evaluate the contribution of lid re sidues to catalysis. The reported an alysis of the cat alytic role of the active site lid in the mechanism of ALA production suggests that the ALAS chemical mechanism may have evolved so as to be limited by a conformational change. The lid presumably closes over the active site dur ing catalysis, thereby facilitating enzymemediated chemistry. Once chemistry occurs, the dynamics of the activ e site lid determine the rate of product release.
122Figure 4.1. The position of th e active site loop in the R. capsulatus ALAS crystal structure. In (A), the overlap of one monomer of holoenzymic (magenta) and succinylCoA-bound (green) ALAS from R. capsulatus. In (B) the active si te lid in the closed position (teal) is perched above the catalytic cleft of the enzyme. Succinyl-CoA and the co-factor PLP are shown as sticks. The image was constructed using Pymol and PDB entries 2BWN and 2BWO. The primary amino acid se quence of the lid is +NH3YVQAINYPTVPRGEELLRCOOA B
123Materials Reagents. DEAE-Sephacel, -mercaptoethanol, PLP, bovine serum albumin, succinyl-CoA, ALA-hydrochloride, -keto-glutarate, -ketoglutarate dehydrogenase, Bis-Tris, HEPES-free acid, MOPS, thiamin pyrophosphate, NAD+, and the bicinchoninic acid protein determination kit were purchased from Sigma-Al drich Chemical Company. Ultrogel AcA 44 was from IBF Biotechnics. Glycerol, glycine, disodium ethylenediamine tetraacetic acid dihydrate, ammonium sulfate, ascorbic acid and magnesium chloride hexahydrate were acquired from Fisher Scientific. Sodium dodecyl sulfate polyacrylamide gel electrophoresis reag ents were acquired from Bio-Rad. PD-10 columns were from Amersham Biosciences. Restriction enzymes and polymerases were from New England Biolabs. Synthetic oligonucleotides were obtained from Integrated DNA Technologies. Methods Construction of the ALAS synthetic shuffled library. The design and experimental approach for construction of the synthetic shuffled library was based on previously described methods (14-16). Codons for 10 of the 18-ami no acid loop were targeted for mutagenesis using synthetic shuffling, so that multiple amino acid variations could be introduced in these 10 positions (Table 4.1) The ALAS active site loop-encoding fragment was reconstituted from partially ove rlapping oligonucleotides using PCR, with each of the codons for the 10 positions harboring, in addition to the wild-type amino acid, alternative amino acids encoded by nucleotide degeneracies (Table 4.1). Following the assembly by PCR, the reaction product was am plified with another round of PCR having as primers oligonucleotides which annealed ag ainst the 5 and 3 ends of the generated
124 Table 4.1. Designed mutations for incorporation at indicated positions within the ALAS active site loop1. Position WT V 1 V 2 V 3 V 4 V 5 V 6 V 7 V 8 V 9 V 10 V 11 V 12 Codon 423 V2 L1 I2 M1 (M/G)TK 425 A2 S2 P2 G2 R2 W1 C1 (M/G)SS 428 Y2 F2 H2 S2 C2 I2 L2 N2 R2 (M/T)(A/K)Y 432 P3 A3 G3 T3 D2 S2 H2 N2 R1 E1 K1 (M/G)(M/G)(S/T) 433 R6 V4 G4 L4 I3 S2 K2 D2 E2 H2 N2 Q2 M1 (M/G)(A/K)N 434 G2 K1 D1 E1 N1 R1 S1 RRW 435 E1 R4 T3 A3 G3 P3 D2 S2 H2 N2 Q1 K1 (M/G)(M/G)(S/T) 436 E1 L1 Q1 V1 SWG 437 L3 R5 L3 K2 I2 Q2 M1 H1 N1 S1 M(R/T)(M/G) 438 L3 F1 YTK 1WT denotes amino acid found in mALAS2 active site loop, while V re fers to variants encoded within the library. Bold amino acid s indicate amino acids that are found in ALASs from different species. Codon indicates the nucleotide codon used to obtain the indicated mixtur e of amino acid residues. Subscripts following amino acid designations are the number of different codons for that amino acid. Nucleotide degeneracies a re represented in the IUB code: Y, C/T; M, A/C; K, G/T; R, G/A; S, C/G; W, A/T; N, A/C/G/T.
125 PCR product and covered the sequence for restri ction enzyme sites used in subsequent subcloning (i.e., Xba I and Bam HI). To minimize the wi ld-type ALAS background during the screening of the lib rary for functional ALAS variants, the synthetic shuffling library was subcloned into a mock ALAS expr ession vector. The mo ck vector contained the wild-type ALAS-encoding sequence fr om the pGF23 expression plasmid (17), with the exception of the region encoding the active site loop and flanked by the Xba I and Bam HI restriction enzyme sites, which was replaced with a non-ALAS related sequence. The primers, conditions for the annealing reacti on and PCR, and mock vector used in this study are described in supplemental experimental procedures. The ligation reactions and DNA digestions with restrict ion endonucleases were according to standard protocols in molecular biology (18). Screening of the ALAS synthetic shuffl ed library and isolation of functional ALAS variants. Library screening and selection of functional variants was accomplished by reversing the phenotype of E. coli hemA(HU227) (19, 20). HemAcells can only survive if harboring a functiona l ALAS or when ALA (or hemi n) is added to the medium (20). Electrocompetent E. coli HU227 cells were transformed with the library and plated onto LB/ampicillin medium without ALA to al low selection of the active ALAS variants as previously described (19) (Figure 4.2). To score the total number of colonies produced and assess transformation efficiency one-tenth of each transformation reaction was spread onto LB/ampicillin plates containing 10 g/ml ALA. Functional ALAS clones (i.e., isolated from the ALA minus plates) were then picked and plated onto defined MOPS medium to induc e protein overexpression (17) and screened for porphyrin overproduction using fluorescence microscopy. Briefly, the plates with the functional
126 Figure 4.2. The generation and screening of the library. A library of over 330,000 possible ALAS variants was constructed with PCR using a series of degenerate mixed base oligonucleotides. The PCR product was ligated into an expression vector. The resulting plasmids were transformed into Escherichia coli strain HU227 and plated on LB + ampicillin agar with and wit hout ALA. The colonies that grew in the absence of ALA were identified as functional variants. F unctional variants were screened for porphyrin overproduction by fluorescence microscopy. Single colony isolation and se q uencin g of p lasmid DNA Library of synthetically shuffled variants ALA auxotrophic E. coli strain Selection Functional variants subjected to fluorescence microscopy LB agar+ ALA Annealing & fill-in PCR N C Ligation of PCR product hem A-bacteriumLB agar-ALA PCR Transformation of expression p lasmids
127 ALAS clones were examined with a Nikon Eclipse E1000 fluorescence microscope (Nikon, Tokyo) fitted with either a Nikon Triple Band filter set for excitation at 385-400 nm and emission at 450-465 nm for 4 ,6-diamidino-2-phenylindole (DAPI) detection or a Nikon Cube BV2A excitati on filter set for excitation at 400440 nm and emission at 450-465 nm with a 610 nm long pass filter and a band pass filter at 550 nm 20 nm for porphyrin detection. Photographs were taken with a CCIR high performance COHU CCD camera and the images were processed with Image software Genus 2.81 from Applied Imaging. The level of por phyrin accumulated in the functional ALAS clones was compared to that of bacterial cells harboring wild-type ALAS and grown under the same experimental conditions (Figur e 4.3). Bacterial cells harboring ALAS variants, which accumulated gr eater porphyrin levels than b acterial cells with wild-type ALAS, were grown in LB/ampicillin medium in 96-well plates, and the glycerol stocks generated from overnight cultures were s ubmitted to the ICBR Genomics Core at the University of Florida for DNA sequenci ng of the correspo nding plasmids. Overexpression, purification and spectroscopi c analyses of ALAS active site loop variants. Overexpression was from the alkaline phosphatase (phoA) promoter, and the conditions for promoter induction were as previously described for mALAS2 (17). However, induction of the phoA promoter controlling the expression of the F1, SS2, F10 and H1 variants was accomplished by growing the bacterial cells harboring the expression plasmids for these variants in MOPS, a low phosphate concentration and defined medium (17), supplemented with 10mg/L ascorbic acid for 30 hours at 20oC. Purification, storage, handling, and spectroscopic analysis of the mALAS2 variants were conducted as described previously (21). Protein concentrations were determined by the
128 Figure 4.3. Differential fluorescence of ALAS variant isolates streaked on expression agar. 1. DAPI visualized cells contai ning: K313A (negative control) (A), wild-type ALAS (B), hyperactive variant SS2 (C). 2. Cells visualized for red fluorescence containing: K313A (negative control) (A), wild-type ALAS (B), hyperactive variant SS2 (C). 3. Overlay of DAPI and red fl uorescence visualized cells: K313A (negative control) (A), wild-type ALAS (B), hyperactive variant SS2 (C).
129 bicinchoninic acid method using bovine serum albumin as the standard (17). Reported enzyme concentrations are based on the monomeric molecular mass of 56 kDa (17). Protein purity was assessed using SDS-PAGE. Steady-state and pre-steady-sta te kinetic characterization of ALAS active site loop variants. ALAS steady-state activity of the ALAS active site loop variants was determined at 20C using a c ontinuous spectrophotometric as say as described previously for wild-type ALAS (22). The ALAS activity data, acq uired using a Shimadzu UV 2100 dual-beam spectrophotometer, were plotted vs. substrate concentration in which one of the substrate concentrations varied, while th e second was kept constant. The steady-state kinetic parameters were determined by fitting the data to the Michaelis-Menten equation using non-linear regression analys is software as reported in (10). The ALAS steady-state kinetic parameters of wild-t ype ALAS and the SS2 variant activity assays were also determined at 15, 25, 30 and 35 C. Rapid scanning stopped-flow measurements were performed using a model RSM100 stopped-flow spectrophotometer (OLIS, Inc.). This instrument has a dead-time of approximately 2-ms and an observation cham ber path length of 4 mm. Spectral scans covering a wavelength range of 300-510 nm were collected at a rate of 1000 scans/s and then averaged to 62 scans/s to reduce the data files to an appropria te size for global fit analyses. The temperature of the syringes and the stopped-flow cell compartment was maintained at 20oC by an external water bath. Pre-st eady-state kinetic reactions of the variant enzymes were examined under si ngle turnover conditions, using final concentrations of 60 M enzyme, 120 mM glycine and 10 M succinyl-CoA in 100 mM HEPES, pH 7.5 and 10% (v/v) gly cerol as previously described (23). Single turnover
130 ] Ligand [ ] Ligand [D max K Y Ydata were evaluated using either a three-kinetic-step, or a two-step kinetic mechanism as described by Equations 4.1 and 4.2, respectively (23). (Equation 4.1) (Equation 4.2) Observed rate constants were de termined by Robust Global Fitting of the spectral data using the single value decomposition software provided by OLIS Inc. as previously reported (21, 23). Quality of the calculated fits was judged by analysis of the calculated residuals, and the simplest mechanism that described the experimental data was used. Determination of the dissociation const ants for the binding of glycine and ALA. The equilibrium dissociation constant (KD) for the binding of glycine to the SS2 variant was determined in 20 mM HEPES (p H 7.5) and 10% glycerol at 20oC and by titrating the SS2 variant (60 M) with increasing concentrations of glycine (0.6 mM-60 mM) and monitoring the increase in absorbance at 420 nm upon formation of the external aldimine between PLP and glycine (10). The KD value was determined by fitting the data to Equation 4.3, where Y is the absorbance increase at 420 nm, Ymax is the maximum increase in absorbance, and [Ligand] is the glycine concentration, using non-linear regression analysis software. (Equation 4.3) D C B A C B A k k k k k 3 obs 2 obs 1 obs 2 obs 1 obs
131 Binding of ALA to the SS2 variant resulted in quenching of the fluorescence emission at 428 nm upon excitation at 330 nm due to formati on of an external aldimine with the PLPcofactor. Thus to determine the dissociation constant for the binding of ALA to SS2, the change in fluorescence emission at 428 nm ( exc = 330 nm) was monitored upon titration of SS2 (60 M) with increasing concentrations of ALA (0.5 mM-128 mM). The changes in fluorescence at 428 nm were plotted as a function of ALA concentration, and the KD value was determined by fitting the data to Equation 4.3, using non-linear regression analysis software. In Equation 4.3, Y is the total change in fluorescence at 428 nm, Ymax is the initial fluorescence, and [Ligand] is the ALA concentration. Determination of the thermodynamic activation parameters. The temperature dependence of the steady-state kinetic paramete rs of wild-type ALAS and the SS2 variant were examined using the same experimental conditions as described above and covering the temperature range of 15-35 C. The natural log the calculated values for the turnover numbers (lnkcat) were plotted vs. the inverse of temperature and the data were fit to the Arrhenius equation (Equation 4.4). (Equation 4.4) ) ln( 1 ) ln( A T R E ka obs where Ea is the activation energy, R is the universal gas constant, T is the absolute temperature, and A is the frequency factor. The determined activation energies were then used to calculate the thermodynamic activation parameters, H G and S as previously described (23) Data analysis for the definition of the mi nimal kinetic mechanism of a selected ALAS active site loop variant. The KinTekSim kinetic simulation software (24) was used
132 to model the single wavelength kinetic traces at 510 nm for the r eaction catalyzed by the SS2 variant and thus estimate the forward and reverse rate constants. The interior rate constants were allowed to float, while the previously determined KD values for binding of glycine and ALA to the SS2 variant and w ild-type ALAS were maintained constant (21) Results Construction of the synthetically shu ffled library and screening of functional ALAS variants. To the extent that the conserved active site lid in mALAS2 may limit catalysis, synthetic shuffling was used to ev aluate the contribution of amino acids that comprise the lid make to motif dynamics and enzyme turnover. If all 18 amino acid residues on the active site lid were mutated to all possible substitutions, there would be a total of 1820 possibilities, an intractable number of combinations for our experimental approach. Therefore, instead of a random mutagenesis approach, the number of potential mutations at a single position was limited by ut ilizing mixed base oligonucleotides that encode specific changes to the cDNA. Mi xed base oligonucleotid e primers encoding amino acids that have a precedent in nature were incorporated into the primary sequence of wild-type ALAS, from Y 422-R439 (Table 4.1). A multiple sequence alignment quantifying the naturally occurring amino acid diversity pres ent at specific positions in the lid was performed (Figure 4.4). These AL AS sequences were incorporated into the library. To preserve the functional integrity of ALAS, we always included the wild-type nucleotides in the primers. Further, the conserved residues of the active site lid were not mutated; an additional consid eration employed to reduce the possibility of recovering inactive variants. Screening of the syntheti cally shuffled library included a two-pronged approach (Figure 4.2). First, E. coli strain HU227 need aminolevulinate to thrive (25) ;
133 mutants transformed into this strain are able to provide the deplet ed metabolite required for colony formation on agar plates lacking AL A. Next, to elucidate which variants are catalyzing the production of ALA at a rate fast er than wild-type ALAS, the appearance of fluorescence associated with excess porphyrin biosynthesis was used (Figure 4.3).
134Figure 4.4. Multiple alignment of the amino acid sequences of the ALAS loop region. Active Site Loop Y422...............R439 1 ALAS_ DELLEU 486 LLLSKHGI Y V Q A IN Y PTV PRG-EELL R LAP SPHH-SPQMMEDFVE KLLAAWTEVGLPLQD -VSIAACNFCRRPVH FELMSEWERSYFGNM 573 2 ALAS_ DLLLEU 486 LLLSKHGI Y V Q A IN Y PTV PRG-EELL R LAP SPHH-SPQMMEDFVE KLLAAWTEVGLPLQD -VSIAACNFCRRPVH FELMSEWERSYFGNM 573 3 ALAS_HOMSAP 491 LLLSKHGI Y V Q A IN Y PTV PRG-EELL R LAP SPHH-SPQMMEDFVE KLLLAWTAVGLPLQD -VSVAACNFCRRPVH FELMSEWERSYFGNM 578 4 ALAS_RATNOR 491 LLLAKHSI Y V Q A IN Y PTV PRG-EELL R LAP SPHH-SPQMMENFVE KLLLAWTEVGLPLQD -VSVAACNFCRRPVH FELMSEWERSYFGNM 578 5 ALAS_RATRAT 491 LLLAKHSI Y V Q A IN Y PTV PRG-EELL R LAP SPHH-SPQMMENFVE KLLLAWTEVGLPLQD -VSVAACNFCRRPVH FELMSEWERSYFGNM 578 6 ALAS_MUSMUS 491 LLLSKHSI Y V Q A IN Y PTV PRG-EELL R LAP SPHH-SPQMMENFVE KLLLAWTEVGLPLQD -VSVAACNFCHRPVH FELMSEWERSYFGNM 578 7 ALAS_DANRER 486 ILLEKHNI Y V Q A IN Y PTV PRG-EELL R LAP SPFH-NPIMMNYFAE KLLDVWQEVGLPLNG -PAQASCTFCDRPLH FDLMSEWEKSYFGNM 573 8 ALAS_DANROS 486 ILLEKHNI Y V Q A IN Y PTV PRG-EELL R LAP SPFH-NPIMMNYFAE KLLDVWQEVGLPLNG -PAQASCTFCDRPLH FDLMSEWEKSYFGNM 573 9 ALAS_OPSTAU 486 SLLEKHNI Y V Q A IN Y PTV PRG-QELL R LAP SPHH-HPAMMEYFVD KLVEVWQEAGLLLNG -PATVSCTFCDRPLH FDLMSEWEKSYFGNM 573 10 ALAS_HOMSPN 545 ELMSRHNI Y V Q A IN Y PTV PRG-EELL R IAP TPHH-TPQMMNYFLE NLLVTWKQVGLELKP -HSSAECNFCRRPLH FEVMSEREKSYFSGL 632 11 ALAS_ DELDEL 545 ELMSRHNI Y V Q A IN Y PTV RRG-EELL R IAP TPHH-TPQMMNYFVE NLLATWKRVGLELKP -HSSAECNFCRRPLH FEVMSEREKSYFFGM 632 12 ALAS_MOUDOM 546 ELMTRHNI Y V Q A IN Y PTV PRG-EELL R IAP TPHH-TPQMMNFFVE KLLVTWKRVGLELKP -HSSAECNFCRRPLH FEVMSEREKAYFSGM 633 13 ALAS_GALVAR 540 KLMSQHSI Y V Q A IN Y PTV PRG-EELL R IAP TPHH-TPQMMSYFLE KLLATWKDVGLELKP -HSSAECNFCRRPLH FEVMSERERSYFSGM 627 14 ALAS_XENLAE 308 KLMRDYSI Y V Q A IN Y PTV PRG-EELL R IAP TPHH-NPQ-----------------------------------------------344 15 ALAS_OPSBET 532 LMMSHHNI Y V Q A IN Y PTV ARG-DELL R IAP TPHH-TPEMMKYFVD RLVQTWKEVGLELKP -HSSAECTFCQQPLH FEVMNEREKSYFSGL 619 16 ALAS_OPSPAR 532 LMMSHHNI Y V Q A IN Y PTV ARG-DELL R IAP TPHH-TPEMMKYFVD RLVQTWKEVGLELKP -HSSAECTFCQQPLH FEVMNEREKSYFSGL 619 17 ALAS_DANDAN 518 IMMSRYNI Y V Q A IN Y PTV ARG-EELL R IAP TPHH-TPQMMKYFVD KLTQTWTEVGLPLKP -HSSAECNFCRQPLH FEIMSEREKSYFSGL 605 18 ALAS_MYXGLU 565 ELMSRHNI Y V Q A IN Y PTV PRG-EEML R VVV TPHH-TPQMMQYFVE HLTNSWKDIGLNLRP -HASAECNYCKMPIH FELMSEHDQVYFDGM 652 19 ALAS_STRDRO 512 SLLEEHNI Y V Q A IN S PTV PSG-EEKL R IAP SPXH-TPDMMDRFVA SLSEVWAKSGLRFNT PICPRECEFCKNPEK FEELSSRERSFAEES 578 20 ALAS_DROMEL 463 VLIEQFGH Y L Q S IN Y PTV ARG-QEKL R LAP TPFH-TFEMMNALVT DLKKVWEMVDLSTNV PLSPNACMFCNSESC WHQDTSPDLECGIPN 529
135 21 ALAS_LIMPOL 494 ELISMHGH Y V Q A IN Y PTV PRG-EEKL R IAP TPFH-TRPMMEAFVR DLVSVWRGLKLPLRD GICEKKCEFCEKPLY FEHLESRVL----560 22 ALAS_SEPOFF 540 DLLIKHNI Y V Q A IN Y PTV ARG-EEKL R VAA TPHH-TKEMMDHFVD CVVKVWLEHGLTLNP -DSTRPAEFNVKFKK FSI----------603 23 ALAS_GLYDIB 516 ELMEEHGI Y V Q P IN Y PTV PRG-QELL R VAP TPHH-TKEMMDSFVN ATLSVFLNNNIELKS -TCGINCLYCHQPMK CEAFTNRER----586 24 ALAS_GALGAL 420 ALLEEHGL Y V Q A IN H PTV PRGQELLL R IAP TPHH-SPPMLENLAD KLSECWGAVGLPRED -PPGPSCSSCHRPLH LSLLSPLER----508 25 ALAS_SINMEL 346 LLLDNFGI Y V Q P IN Y PTV PKK-TERL R ITP TPMH-SDADIDHLVS ALHSLWSRCALARAV A--------------------------405 26 ALAS_SMRMEL 275 LLLDNFGI Y V Q P IN Y PTV PKK-TERL R ITP TPMH-SDADIDHLVS ALHSLWSRCALARAV A--------------------------334 27 ALAS_AGRTUM 364 ILLDNHGV Y V Q P IN Y PTV PRK-TERL R ITP TPLH-TDADIEQLVG ALHQLWSHCALARAV A--------------------------423 28 ALAS_AGRRAD 346 ILLDSHGV Y V Q P IN Y PTV PRK-TERL R ITP TPLH-SDADIEHLVG ALHQLWSHCALARAV A--------------------------405 29 ALAS_RHIRAD 276 ILLDSHGV Y V Q P IN Y PTV PRK-TERL R ITP TPLH-SDADIEHLVG ALHQLWSHCALARAV A--------------------------335 30 ALAS_RHOPSE 344 ELINRYGI Y V Q P IN Y PTV PRG-TERL R ITP SPQH-TDADIEHLVQ ALSEIWARVGLAKAA ---------------------------403 31 ALAS_RHOPSE 346 ALLARHAI Y V Q P IN Y PTV ARG-QERF R LTP TPFH-TTSHMEALVE ALLAVGRDLGWAMSR RAA------------------------407 32 ALAS_EUGGRA 343 LLLKRFQI Y V Q P IN Y PTV DVG-TERL R ITV SPVH-TNEHMATLIT ALLQVWEELGLPLRP PVFDTEGYPEVEAAE WLPNSAMWR----409 33 ALAS_BRAJAP 348 LLLEEHGI Y I Q P IN Y PTV AKG-SERL R ITP SPYH-DDGLIDQLAE ALLQVWDRLGLPLKQ KSLAAE---------------------409 34 ALAS_BRAJAP 275 LLLEEHGI Y I Q P IN Y PTV AKG-SERL R ITP SPYH-DDGLIDQLAE ALLQVWDRLGLPLKQ KSLAAE---------------------339 35 ALAS_BRUMEL 345 RLLEVHGI Y I Q P IN Y PTV PRG-TER LR ITP SPLH-DDKLIDGLKD ALLEVWNELGIPFAE PSAPQAANSDRIIPL MVSKAGG------425 36 ALAS_ZYMMOB 287 ILLNEYGA Y V Q P IN F PTV PRG-TERL R FTP GPTH-NEAMLRELTD SLVAIWHRLDMRFAA ---------------------------345 37 ALAS_RHOCAP 348 MLLSDYGV Y V Q P IN F PTV PRG-TERL R FTP SPVH-DLKQIDGLVH AMDLLWARCA-------------------------------401 38 ALAS_RHOCAP 278 MLLSDYGV Y V Q P IN F PTV PRG-TERL R FTP SPVH-DLKQIDGLVH AMDLLWARCA-------------------------------331 39 ALAS_PARDEN 292 MLLADFSI Y V Q P IN F PTV PRG-TERL R FTA SPVH-DPKQIDHLVK AMDSLWSQCKLNRST SAA------------------------339 40 ALAS_PARZEA 360 MLLADFSI Y V Q P IN F PTV PRG-TERL R FTA SPVH-DPKQIDHLVK AMDSLWSQCKLNRST SAA------------------------409 41 ALAS_RHOPSE 364 MLLIHFGI Y V Q P IN F PTV PRG-TERL R FTP SPVH-DSGMIDHLVK AMDVLWQHCALNRAE VVA------------------------407 42 ALAS_EMENID 510 KLLEEHGI Y V Q A IN Y PTV PRG-EERL R ITP TPGH-TQELRDHLVE AVNTVWNDLGIKRAS DWKAMGGFVGVGVEA AELENQPIWT---598 Figure 4.4. Continued
136 43 ALAS_EMENID 440 KLLEEHGI Y V Q A IN Y PTV PRG-EERL R ITP TPGH-TQELRDHLVE AVNTVWNDLGIKRAS DWKAMGGFVGVGVEA AELENQPIWTDAQLN 528 44 ALAS_ASPNID 440 KLLEEHGI Y V Q A IN Y PTV PRG-EERL R ITP TPGH-TQELRDHLVE AVNTVWNDLGIKRAS DWKAMGGFVGVGVEA AELENQPIWTDAQLN 528 45 ALAS_ASPORI 508 KLLEEHGI Y V Q A IN Y PTV PRG-EERL R ITP TPGH-IKEHRDHLVQ AVQTVWNELGIKRTS DWEAQGGFVGVGVDG AEAENQPIWNDVQLG 596 46 ALAS_NEUCRA 334 KLLNDHQI Y V Q S IN Y PTV PVG-QERL R ITP TPGH-TKQFRDHLVA ALDSIWTELGIKRTS DWAAEGGFIGVGEAE AEPVAPLWTDEQLG 421 47 ALAS_GIBFUJ 393 MLLNDYGI Y V Q A IN Y PTV PVG-QERL R VTP TPGH-IKEYRDQLVE AIDEIWTRLDIKRTS DWAAEGGFIGVGEQD -NVQEPLWTDKQLN 479 48 ALAS_YARLIP 404 LLLTKHQI Y V Q A IN F PTV PIG-QERL R VTP TPGH-HEGLCDELVA ALEDVWQELDLKRVE DWTAEGGLCGVGEGV --EVEPLWSEEQLS 489 49 ALAS_CANALB 451 LLLNKHDI Y V Q A IN F PTV PIG-EERL R ITP TPGH-GPELSKQLVE AVDSVFTELNLNRIN DWKKLGGLVGVGVEG AAKVEHIWTEEQLA 538 50 ALAS_CANALB 381 LLLNKHDI Y V Q A IN F PTV PIG-EERL R ITP TPGH-GPELSKQLVE AVDSVFTELNLNRIN DWKKLGGLVGVGVEG AAKVEHIWTEEQLA 468 51 ALAS_DEBHAN 392 LLLDKYNI Y V Q A IN F PTV PIG-QERL R ITP TPGH-GPELSNQLIG ALDSVFNELSLSRIG DWEGKGGLCGVGEPD IEPIEHIWTSEQLA 479 52 ALAS_SACCER 435 ILINKHQI Y V Q A IN F PTV ARG-TERL R ITP TPGH-TNDLSDILIN AVDDVFNELQLPRVR DWESQGGLLGVGESG FVEESNLWTSSQLS 522 53 ALAS_SACCAS 365 ILINKHQI Y V Q A IN F PTV ARG-TERL R ITP TPGH-TNDLSDILIN AVDDVFNELQLPRVR DWESQGGLLGVGESG FVEESNLWTSSQLS 452 54 ALAS_CANGLA 347 ILMEKHRI Y V Q A IN F PTV SRG-TERL R ITP TPGH-TNDLSDILIA AVDDVFNELQLPRIR DWEMQGGLLGVGDKN FVPEPNLWTEEQLS 434 55 ALAS_KLULAC 387 ILMDKHRI Y V Q A IN F PTV ARG-TERL R ITP TPGH-TNDLSDILMD ALEDVWSTLQLPRVR DWEAQGGLLGVGDPN HVPQPNLWTKDQLT 474 56 ALAS_EREGOS 373 ILMEKHRI Y V Q A IN F PTV PRG-TERL R ITP TPGH-TNDLSDVLLD AMDDVWKTLQLPRVS DWAAHGGLLGVGEPD YVPEANLWTEEQMS 460 57 ALAS_AGABIS 460 KLLSEHDI Y V Q A IN Y PTV ARG-EERL R ITV TPRH-TMEQMEGLIR SLNQVFEELNINRLS DWKLAGGRAGVGIPG AADDVQPIWTDEQIG 548 58 ALAS_AGADIV 390 KLLSEHDI Y V Q A IN Y PTV ARG-EERL R ITV TPRH-TMEQMEGLIR SLNQVFEELNINRLS DWKLAGGRAGVGIPG AADDVQPIWTDEQIG 478 59 ALAS_SCHPOM 475 SLLHDHNI Y V Q S IN F PTV SVG-TERL R ITP TPAHNTEHYVQSLTN AMNDVWSKFNINRID GWEKRGIDVGRLCKF PVLPFTTTH----558 60 ALAS_SCHMIK 405 SLLHDHNI Y V Q S IN F PTV SVG-TERL R ITP TPAHNTEHYVQSLTN AMNDVWSKFNINRID GWEKRGIDVGRLCKF PVLPFTTTH----488 61 ALAS_RICPRO 342 MLLNEYGI Y V Q H IN F PTV PRG-TERL R IIP TPAH-TDKMINDLST ALVHIFDELDIELSS AKELNKEVRLHLIA-------------414 62 ALAS_RICTYP 342 MLLNEYGI Y V Q H IN F PTV PRG-TERL R IIP TPAH-TDKMINDLST ALVHIFAELDIELSS TKELNKEVRLHLIA-------------414 63 ALAS_RICPRO 272 MLLNEYGI Y V Q H IN F PTV PRG-TERL R IIP TPAH-TDKMINDLST ALVHIFDELDIELSS AKELNKEVRLHLIA-------------344 64 ALAS_RICCON 354 MLLNEYGI Y V Q H IN F PTV PRG-TERL R IIP TPAH-TDKMINDLSV ALVQIFAELDIELSS AKELNEEVRLNLIA-------------426 65 ALAS_CHRVIO 354 RLLEEFDI Y V Q P IN Y P S V PRG-GERF R LTV GPRR-SHEEIQRFVA ALKHCLA----------------------------------Figure 4.4. Continued
137Figure 4.4. Multiple alignment of the amino acid sequences of the ALAS loop region. The amino acid sequences were obtained from public databases (NCBI) using a BLAST search and aligned using CLUSTAL W (1). The 10 positions within the 18-amino acid sequence targeted for s ynthetic shuffling are high-lighted in cyan The amino acid numbering in red refers to that of murine eryt hroid ALAS (mALAS2). Represented proteins are: ALAS_DELLEU: Delphinapterus leucas ALAS (20138447); ALAS_DLLLEU: Delphinapterus leucas cook inlet subspecies 2 (5281116); ALAS_HOMSAP, Homo sapiens ALAS erythroid (4557299); ALAS_RATNOR, Rattus norvegicus erythroid ALAS (51980582); ALAS_RATRAT, Rattus rattus erythroid ALAS (6978485); ALAS2_MUSMUS, Mus musculus erythroid ALAS (33859502); ALAS_DANRER, Danio rerio ALAS (18858263); ALAS_DANROS, Danio roseus ALAS (20138448); ALAS_OPSTAU, Opsanus tau ALAS (1170202); ALAS_ORYLAT, Oryzias latipes ALAS (49022596); ALAS_HOMSPN, Homo sapiens erythroid ALAS (4502025); ALAS_DELDEL, Delphinus delphis erythroid ALAS (20138445); ALAS_MOUDOM, Mus musculus domesticus erythroid ALAS (23956102); ALAS_GALVAR, Gallus varius erythroid ALAS (122821); ALAS_XENLAE, Xenopus laevis erythroid ALAS (44968228); ALAS_OPSBET, Opsanus beta ALAS (1170206); ALAS_OPSPAR, Opsanus pardus ALAS (532630); ALAS_DANDAN, Danio danglia ALAS (32451642); ALAS2_MYXGLU, Myxine glutinosa erythroid ALAS (4433550); ALAS1_BRALAN, Branchiostoma lanceolatum ALAS 1 (28630217); ALAS1_STRDRO, Strongylocentrotus droebachiensis ALAS 1 (4433548); ALAS1_DROMEL, Drosophila melanogaster ALAS 1 (2330591); LAS1_LIMPOL, Limulus polyphemus ALAS 1 (4433540); ALAS1_SEPOFF, Sepia officinalis ALAS 1 (4433546); ALAS2_GLYDIB, Glycera dibranchiate ALAS 2 (4433544); ALAS2_GALGAL, Gallus gallus gallus ALAS 2 (1170201); ALAS_SINMEL, Sinorhizobium meliloti 1021 ALAS (15966742); ALAS_SMRMEL, Sinorhizobium meliloti ALAS (18266808); ALAS_AGRTUM, Agrobacterium tumefaciens ALAS (889869); ALAS_AGRRAD, Agrobacterium radiobacter ALAS (95069); ALAS_AGRTUM, Agrobacterium tumefaciens ALAS (122818); ALAS_RHOPAL, Rhodopseudomonas palustris ALAS (4001678); ALAS_RHOSPO, Rhodobacter sporagenes ALAS (541302); ALAS_EUGGRA, Euglena gracilis ALAS (12620813); ALAS_BRAELK, Bradyrhizobium elkanii ALAS (66534); ALAS_BRAJAP, Bradyrhizobium japonicum ALAS (30179569); ALAS_BRUMEL, Brucella melitensis ALAS (25286547); ALAS_ZYMMOB, Zymomonas mobilis ALAS (4511998); ALAS_RHOGLU, Rhodobacter gluconicum ALAS (97435); ALAS_RHOCAP, Rhodobacter capsulatus ALAS (122828); ALAS_PARDEN, Paracoccus denitrificans ALAS (1170207); ALAS_PARZEA, Paracoccus zeaxanthinifaciens ALAS (537435); ALAS_RHOSPH, Rhodobacter sphaeroides ALAS (541301); ALAS_EMENID, Emericella nidulans ALAS (418756); ALAS_EMCNID, Emericella nidulans ALAS (585244); ALAS_ASPNID Aspergillus nidulans ALAS (40745239); ALAS_ASPORY, Aspergillus oryzae ALAS (5051989); ALAS_NEUCRA, Neurospora crassa ALAS (52782908); ALAS_GIBFUJ, Gibberella fujikuroi ALAS (15721883); ALAS_YARLIP, Yarrowia lipolytica ALAS (52782857); ALAS_CANBER, Candida berate ALAS (7493758); ALAS_CANALB, Candida albicans ALAS (10720014); ALAS_DEBHAN, Debaryomyces hansenii ALAS (52782855); ALAS_SACCER, Saccharomyces cerevisiae ALAS (6320438); ALAS_SACCAS, Saccharomyces castellii ALAS (122831); ALAS_CANGLA, Candida glabrata ALAS (52782865); ALAS_KLULAC, Kluyveromyces lactis ALAS (52788271); ALAS_EREGOS, Eremothecium gossypii ALAS (52782894); ALAS_ASPBIS, Agaricus bisporus ALAS (1679599); ALAS_AGADIV, Agaricus divoniensus ALAS (2492846); ALAS_SCHPOM, Schizosaccharomyces pombe ALAS (7492336); ALAS_SCHMIK, Schizosaccharomyces mikatae ALAS (52782853); ALAS_RICCON, Rickettsia conorii ALAS (7433712); ALAS_RICTYP, Rickettsia typhi ALAS (51474008); ALAS_RICPRO, Rickettsia prowazekii ALAS (6225494); ALAS_RICRIC, Rickettsia rickettsia ALAS (2528635); ALAS _CHRVIO, Chromobacterium violaceum ALAS (34102112). (1) Thompson, J. D., Higgins, D. G., and Gibson, T. J. (1994) Nucleic Acids Res. 22 4673-4680
138 Under these conditions, it was possible to isol ate ALAS variants with turnover numbers greater than that of wild-type ALAS. In pa nels (A1, B1 and C1) of Figure 4.3 cells are visualized with a DAPI filter, which was us ed a means of assaying for single colony isolates. The magnification for each imag e was identical, implying that colony size differed between those cells harboring a hypera ctive variant compared to wild-type and the negative control (K313A) (26) This decrease is colonial diameter is likely due to ALA toxicity (27) The contrast between those ce lls harboring the inactive ALAS (K313A) and those complemented with wild -type ALAS show clearly the lack of porphyrin-derived red fluoresce nce in the negative control (Figure 4.3 Panels A2 and B2). The dissimilarity in accumulated porphyrin levels between wild-type ALAS and the SS2 variant is highlighted by the considerably greater red fluorescence present in the library isolate compared to wild-type ALAS (Figure 2 Panels B3 and C3). In summary, comparison of accumulated porphyrin levels present in cells as visualized by fluorescence microscopy of bacterial col onies on expression agar proved a reliable method for obtaining hyperactive isolates. Qualitative analysis of the isolat ed active site lid variants. The mutations present among the active site lid varian ts is shown in Table 4.2. Functional mutations were detected at 9 of the 18 positions that comprise the lid. Based on our experimental design it can be concluded that the mutation of Asn427 to His is most likely an artifact of the primer design and PCR. Amino acid substi tutions observed among th e variants included highly disruptive changes ( e.g ., A425P, P432E, L437K, E435K) as well as more conservative variations ( e.g ., V423L, A425G, Y428H, R433K) This suggests that noncovalent forces including hydrogen bonding as well as Van der Waals and electrostatic
139 Table 4.2. Amino acids substitutions in active site lid variants. Shuffled positions are indicated in red. Wild-type ALAS Y V Q A I N Y PTV P R G E E L L R Single variants A8 T D8 Q G7 H Quadruple variant F1 KK Q Q Penta variant F10 I QN T N Hexa variants A4 I P C RK N F3 G H HN K K H1 N NI E K K Hepta variant SS2 L R E I N Q K
140 interactions may be perturbed in the vari ants. Replacements at residues toward the carboxy terminus of the lid, (P432-R439) favored conversion to basi c amino acids, likely positively charged at physiological pH, while residues that comprised the portion of the lid proximal to the amino terminus (Y422-V 431) showed less restriction toward which amino acid was permissible. Several positions showed a propensity for a particular substitution as demonstrated by the frequenc y of the observed muta tion. At position 423, valine was found substituted th ree times for leucine, indi cating that hydrophobicity may be a necessary characteristic at this position in the lid. Toward the C-terminal end of the lid, E435 was found mutated twice to both glut amine and lysine, L437 was substituted for lysine three times and glutamine twice. Th is indicates that the presence of positively charged residues in this area of the lid may facilitate lid dynamics. Steady-state kinetic an alysis of the active site lid variants. The steady-state kinetic characterization of 9 variants from the library was carried out at 20oC (Table 4.3). Compared to wild-type ALAS, all of enzy mes had higher turnover numbers. The SS2 variant showed a kcat 16-fold higher than that of the wild-type enzyme. Similarly, the kcat values for the quadruply, quintuply and hextupl y mutated F1, F10 and H1 variants were also markedly higher compared to wild-typ e ALAS. The remaining variants, including the single point variants, also showed enha nced turnover, ranging from a 50% increase observed in D8 to an 8-fold increased displayed by the A4 isolate. While the turnover number data suggest that a single amino acid substitution is enough to increase enzyme activity, it is the combination of amino acids substitutions toward the carboxy terminal region of the lid that elicit the greatest increas es in turnover number. As compared to that of wild-type ALAS, the Michaelis constant for succinyl-CoA was notably different
141 Table 4.3. Kinetic parameters for the rea ctions of hyperactive ALAS enzymes1 Steady-state parameters Pre-steady-state parameters Enzyme kcat s-1 Gly mK mM SCoA mK M kcat/Gly mK mM-1 s-1 kcat/SCoA mK M-1 s-1 Qf 2 s-1 Qd1 3 s-1 Qd2 4 s-1 WT 0.02 0.01 14 2.0 11 1.0 1.1x10-3 1.5x10-3 0.8 0.53 0.01 SS2 0.31 0.06 12 1.2 3.0 0.3 0.02 0.10 16 1.7 N/A A4 0.16 0.01 14 1.1 2.3 0.4 0.01 0.07 7.1 1.7 0.07 F3 0.20 0.01 13 1.1 2.6 0.4 0.02 0.08 7.7 1.1 0.17 H1 0.23 0.14 13 1.4 2.0 0.3 0.02 0.11 10 1.5 N/A F10 0.17 0.01 16 1.8 1.1 0.7 0.01 0.16 3.7 0.98 0.15 F1 0.20 0.02 16 1.2 2.9 0.4 0.01 0.07 4.0 1.1 N/A A8 0.07 0.01 25 3.7 2.3 0.4 2.5x10-3 0.03 4.8 0.81 0.12 D8 0.03 0.01 24 3.1 1.7 0.7 1.3x10-3 2.0x10-3 5.4 0.44 0.02 G7 0.07 0.01 15 1.7 1.5 0.1 4.4x10-3 0.05 4.4 0.87 0.12 1Enzymatic reactions monitored at 20 C; 2Rate for quinonoid intermediate formation; 3Rate for first step of qu inonoid intermediate decay; 4Rate for second step of quinonoid intermediate decay.
142 among the active site lid va riants. All of the variants show ed at least a 3-fold reduction in theSCoA mK Accordingly, due to the enhanced turnover evident among the multiply mutated variants, the catalytic efficiency w ith respect to succinyl-CoA increased no less than 10-fold. For the same reason, because the Km for glycine was relatively unaltered, the catalytic efficiency with glycine also increased among the isolated variants. The increase in affinity observed among the active site lid variants with respect to succinylCoA supports a hypothesis whereby the binding of this substrate switches a pre-existing protein conformational equi librium towards a closed, catalytically competent, conformation (Lendrihas, et. al. submitted) (28) Pre-steady-state reaction of the active site lid variant enzyme-glycine complexes with succinyl-CoA. To understand the contribution that the active site li d makes toward the reaction catalyzed by ALAS, the transient kinetic parameters of the library variants were elucidated. The formation and decay rates of a ke y step in the chemical mechanism were obtained (Table 4.3). Under single tu rnover reaction conditi ons, the lifetime of a transient quinonoid intermediate was observe d with respect to time as a change in absorbance at 510 nm. The absorbance change timecourses were fit to a sequential mechanism with either two or three-steps, eq uations 1 and 2, respect ively. An initial burst of quinonoid intermediate formation, followed by a two-step rate of decay was characteristic to 6 of the enzymes tested (F igure 4.5A-F). Compared to wild-type ALAS (0.8 s-1), the A4 and F3 multiply mutated lid varian ts, showed a 9-fold increase in the rate of quinonoid intermediate formation (7.1 s-1 and 7.7 s-1, respectively). Additionally, the rates corresponding with the first step of qui nonoid intermediate decay in these variants (1.7 s-1 and 1.1 s-1) were also increased over that of wild-type ALAS (0.53 s-1).
143 Figure 4.5. The single turnover reactions of isolated hyperactive ALAS variants. The pre-steady state kinetic parameters we re calculated under single turnover conditions (60 M enzyme, 10 M succinyl-CoA, and 120 mM glycine) at 20oC and by monitoring absorbance changes at 510 nm. The reaction catalyzed by the wild-type enzyme is characterized by a single step of quinonoid intermediate formation and 2-step process of decay. The reactions catalyzed by the F10 and H1 variants (panels G, and H, respectively) are markedly different, they follow a single
144 step of intermediate formation and single step of intermediate decay. The remaining variant catalyzed reactions: A4, D8, G7, A8, F3 and F1 (panels A-F, respectively) resemble that of the wildtype enzyme.
145 Accordingly, the second step of quinonoid intermediate decay, a step hypothesized to include the dissociation of product and ope ning of the putative active site lid was markedly increased in these va riants, with values of 0.07 s-1 and .017 s-1, rates 7and 17fold higher when compared to 0.01 s-1 for wild-type ALAS. These data support the increased catalytic efficiency observed from the experiments performed in the steadystate. The singly mutated variants A8 D8 and G7 also formed the quinonoid intermediate at least 4-fold faster wh en compared to wild-type ALAS (4.8 s-1, 5.4 s-1, 4.4 s-1, respectively). However, the first rate of biphasic quinonoid intermediate decay was similar. The rates for all three proteins di d not exceed that displayed by wild-type ALAS. This suggests that the coordinated action of mo re than one residue may be responsible for initiating active site lid dynamics, and altering the kinetic mechanism. In the remaining 3 variants, the lifetime of the transient quinonoid intermediate was dramatically different (Figure 4.5H, 4.5I and 4.6A). Instead of pr oceeding by a mechanism similar to wild-type ALAS, these enzymes appear to condense the bi phasic rate of decay into a single step. Consequently, the data are fitted to a two step sequential mechanism. SS2, H1, and F1 all form the quinonoid intermediate faster than wild-type ALAS. Most notably, SS2 does so at a rate that is 20-fold faster (16 s-1 vs.0.8 s-1 for wild-type ALAS) (Figure 4A). This group of variants also show the greatest enhancement in the rate associated with quinonoid intermediate decay. SS2, H1, F10 and F1 all display at least a 2-fold increase in the rate of quinonoid decay. However, si nce the second step of quinonoid intermediate decay is not observed among these variants it is possible that as the rate of decay becomes monophasic and shows an observed rate that is significantly greater than that measured for the slower second phase observed in wi ld-type ALAS, the true rate of
146 quinonoid intermediate decay is no longer conformationally dependent, and approximates the rate of product release. Equilibrium Constants for the SS2 variant The dissociation constants for the binding of glycine and ALA to th e SS2 variant at pH 7.5 and 20oC were used to model the kinetic mechanism (Figure 4.6B and 4. 6C). Schiff base formation between the cofactor of the SS2 variant and glycine was monitored spectroscopically at 420 nm. The glycine concentration dependent data we re fit to a standard hyperbola and the KD for glycine was found to be 4.12 0.57. This valu e is 50% lower than that of the wild-type enzyme. The increased affinity of SS2 for glycine suggests that the active site lid mutations facilitate substrate binding, a finding coincident with increased catalytic efficiency. The dissociation constant for ALA, as measured in the SS2 variant, was determined by monitoring the change in fluorescence emitted at 428 nm upon excitation at 330 nm. To determine the KD for the product, the enzyme was reacted with increasing concentrations of ALA. The dissociation c onstant for ALA was determined to be 1.82 0.19 mM, a value 62% higher compared to that of wild-type ALAS. This value, a notable decrease in affinity between the product a nd the SS2 variant may be attributable to increased active site lid flexibility, a ci rcumstance supported by the hyperactivity measured for SS2 in the stea dyand pre-steady-state. Thermodynamic properties of wild -type ALAS and the SS2 variant. The dependence of the turnover number ( kcat) on temperature was characterized over the range 288308 K for both wild-type ALAS and the SS2 variant (Figure 4.7). The kcat values were calculated using the same experimental method and data treatment as was described for the reported steady-state kinetic parame ters. The turnover numbers for each enzyme
147 Figure 4.6. The SS2 variant catalyzed reaction. The pre-steady state kinetic parameters for SS2 were calculated under single turnover conditions at 20oC and by: (A) monitoring quinonoid intermediate absorbance changes at 510 nm; (B) monitoring internal aldimine formation absorbance changes at 420 nm, and; (C) monitoring the change in fluorescence emission at 428nm upon excitation at 330 nm upon the addition of ALA.
148 were used to construct an Arrhenius plot, from which the thermodynamic activation parameters were derived (Table 4.4). Th e Arrhenius plot of the experimental data showed a linear dependence in the temperature range for both enzymes. From the slope of the straight line, an activation energy ( Ea) of 0.5 kcal/mol was calculated for the wildtype enzyme. For the SS2 enzyme, Ea was determined to be 0.1 kcal/mol a value 80% lower compared to that of wild-type ALAS This suggests that the SS2-catalyzed reaction may decrease the energy barrier for enzyme motions that are coupled to the reaction. Different slope values were calcu lated from the linear relationship between the kcat values for both enzymes and te mperature. The slope value associated with wild-type ALAS was -24, compared to -9.2 for the SS2 variant. These two dissimilar values suggest that the SS2-catalyzed reaction pro ceeds by an alternate reaction pathway, where chemistry and lid mobility determines the rate limiting step. Figure 4.7. The thermal dependence of the SS2 variant catalyzed reaction. kcat values were calculated by performing steady-stat e kinetics at different temperatures. (A) an Arrhenius plot depicting the difference between the wild-type ( ) and the SS2 variant catalyzed ( ) reactions at 288, 292, 297, 302 and 308K. Error bars are plotted over and partially obscured by the data points.
149 Table 4.4. Thermodynamic activation pa rameters of wild-type ALAS and the SS2 variant. Enzyme Slope Ea ( kcal/mol ) H ( kcal/mol ) G ( kcal/mol ) S ( kcal/molK ) SS2 -9.2 0.01 0.003 -0.58 16 -56 WTALAS -24 0.05 0.002 -0.55 17 -58 Figure 4.8. The simulated kinetic mechanism of the SS2 variant catalyzed reaction. Transition from an internal aldi mine with lysine 313 to an ex ternal aldimine with glycine is the first step. Succinyl-CoA binds second which is followed by quinonoid formation, protonation of the quino noid to yield an aldimine bound mol ecule of ALA. Finally, the release of ALA from the active site is the rate limiting step. E; SS2: G; glycine: EG; enzyme-glycine complex: SCoA; Su ccinyl-CoA: EGSCoA; ALAS-glycine-Succinyl-CoA complex: EQ; enzyme bound to prot onated quinonoid: EALA; enzyme-ALA complex. KD = 4.28 mM E+G EG+SCoA EGSCoA EALA E+ALA KD = 1.82 mM 1.7 s1 16 s1 2.2 M1 s1 1.5 s1 0.22 s1 11 s1 EQ
150Discussion Comparison of the X-ray crystal struct ures of holoenzymic ALAS and AONS with the succinyl-CoAand AON-bound struct ures, revealed the presence of a conformationally mobile active site lid. The out ermost part of the active site lid of ALAS undergoes a 3.5 change in conformation in the presence of succinyl -CoA (Figure 4.1). To address the role of this dynamic struct ure in the enzyme-catalyzed reaction we employed synthetic shuffling to alter the ami no acid composition of the active site lid in mALAS2 (Y422-R439). Our results suggest th at amino acid substitutions within the active site lid lead to altered lid dynamics, a circumstance affecting enzymatic turnover, and one that ultimately liberates the en zyme from a conformationally limited rate determining step. The approach used to evaluate the active site lid of ALAS u tilized a mutagenic technique called synthetic shuffling (14) Non-conserved amino acids that comprise the lid vis vis the multiple sequence alignment, tolerated sequence substitutions differently based on their position in the motif (Figur e 4.4 and Table 4.1). Shuffled residues proximal to the amino terminus (V423, A425, Y428) were found to be mutated a total of 12 times in the isolated variants. This is in stark contrast to the 31 mutations identified among the residues that comprise the carboxy terminal portion of the lid (P432, R433, G434, E435, L437). The imbalance in muta tional frequency evid ent between the two halves of the lid suggest that the role of th e carboxy terminal half of the lid may be less structural in nature, as demons trated by ability of drastically mutated variants such as SS2 and F1 to not only retain activity, but tur nover product at rates 10and 15-fold faster compared to wild-type ALAS. All of the si ngly mutated variants had replacements in
151 both halves of the lid (A 8, D8 and G7). While the isolat ion of G7 may be an artifact of the mutational approach, G7, coupled with A8 and D8 contained substitutions whereby non-polar amino acids were swapped with re sidues capable of forming a hydrogen bond. The A425T mutation present in the A8 variant could affect the orientation of the amino terminal portion of the lid with the two -helices ( 2 and 3) of the adjacent monomer, a scenario also possible in the multiply mutated variants SS2, A4, and F3. In fact, the contribution of these abutting helical residue s to ALAS function has been previously addressed (19) Gong et. al. found that mutating N150 to lysine, and I151 to phenylalanine and leucine, in mALAS2, increased the Vmax over that of wild-type ALAS. Indeed, the active site lid library data, coupl ed with the mutational data on residues N150 and I151 agrees with three prev ious models where specific in teractions that drive ligandinduced closure and catalysis are prop osed to be interdomain in nature (29-31) Two characteristics are shared among all of the variants in the library with respect to steady-state kinetic parameters (Table 4.3). First, all of the enzymes examined showed an increase in turnover number. Second, the catalytic efficien cy for both substrates was enhanced. These characteristics indicate th at the reaction catalyzed by ALAS may be limited by a conformational change leading to product release, a mechanism evaluated among other enzymes (32, 33) Accordingly, mutations that affect the mobility of a conformationally mobile enzyme structure lead to consequences for reactions in which the physical step of product releas e, rather than chemistry, is rate limiting. Therefore the enhanced turnover observed among the active site lid variants potentia lly validates our ALAS catalytic mechanism in which the enzy me conformation switches between closed and open to stabilize reaction intermediates and release product, respectively; a
152 mechanism recently proposed for another PLP-dependent enzyme, diaminopimelate decarboxylase (34) The majority of variants which showed an increase in kcat were found to have mutations that occur toward th e carboxy terminal porti on of the active site lid. At position 433, the A4 and F1 variants contained mutations to lysine, compounded with multiple mutations to the motif. A pr evious study investigating single point mutations to R433 (R433L a nd R433K), found that the kcat values increased 20% and 100% for the respective substitutions (11) Those data coupled w ith the findings of this study suggest that position 433 may significantly contribute to lid mobility and the dissociation of ALA from the enzyme. Ho wever, the variants that had the largest increases in turnover number a nd catalytic efficiency were t hose with mutations to both regions of the lid, e.g ., SS2 and F3. These data support the hypothesis, based on the crystal structure of R. capsulatus ALAS, that variation to both regions of the active site lid leads to relaxation of helices 2 and 3, with concomitant disruption of the active site lid interaction with the aldimine-bound produ ct, likely resulting in acceleration of the conformationally limited release of ALA. Conversely, the si ngle point mutant identified in D8 (L437Q) showed onl y a modest increase in kcat compared to wild-type ALAS. This finding implies that mutations to L437 alone are not sufficient to accelerate product turnover. Supportively, since mutations to this residue were identifi ed in concert with multiple replacements in 5 variants, we believe that the major effect of enhancing the kcat value for enzymes in the library was due to multiplex variation to the amino acid composition of the motif. With regard to the nature of the rate limiting step in variants isolated from the library, a conformational change is likely the predominant kinetic ba rrier in the single-
153 turnover reaction. That is, in a process th at can only be limited by the second phase of quinonoid intermediate decay or by the rate of product release, the latter must be the more rapid, as evidenced by steady-state turnov er numbers. The rate limiting step in 6 of the variants from the library is identified as the opening of the active site lid triggered by product formation (Figure 4.5A-F). The func tion of this change in structure likely disrupts the interaction between the carboxylate moiety of ALA, so that the aldimine bond formed between the product and PLP ca n be strongly polarized, and ultimately lysed. Additionally, lid opening enhances pr oduct dissociation (compared to wild-type ALAS, the SS2 variant showed a 4-fold increase in the ALA DK ) by potentially: uncovering ALA from within a catalytic vacuole, rest oring electron density to the site of -carbon bond scission, and minimizing contact between product and enzyme. Further, since the second rate of quinonoid interm ediate decay approximates kcat in wild-type ALAS and these 6 variants it is likely that the there is a single dominant energy barrier, and that it is thermodynamically advantageous to limit tur nover by equalizing th e energy of quinonoid intermediate decay in the en zyme-catalyzed reaction. The pre-steady-state behavior of three variants from the library differs from that observed with the wild-type and other library isolates (Figur e 4.5G, 4.5H and 4.6A). For these three library isolates (SS2, F1 and H1 ) the observed rate of quinonoid intermediate decay is condensed into a single step. The quinonoid intermediate formation rate immediately after mixing the SS2-glycine comple x with succinyl-CoA is ~20-fold faster than the corresponding ra te in wild-type ALAS, suggesting that a step afte r catalysis is only partially rate determining for this va riant under these conditions. This model is consistent with the decrease in the kcat-derived slope and Ea for this mutant when the
154 temperature is varied (Table 4.4). Togeth er with the greater dissociation constant determined for ALA with these specific mutations, the data suggest that a conformational change leading to product release may no longer be rate limiting for the SS2-catalyzed reaction. In several other enzymes where chemistry has been implicated as rate limiting (35-37) it was suggested that the effects of conformational change on kcat were the result of a thermodynamically driven commitment to cata lysis. In the three variants that appear condensed with respect to the ra te of quinonoid inte rmediate decay, Qd is greater than that of wild-type ALAS and all the active site lid variants tested (Tab le 4.3). This finding suggests that the reactions catalyzed by SS2, F1 and H1 are energetically favorable. Indeed a scenario that the increases in exothermy and positive differential activation entropy identified in the SS2 variant strongly support. Based on the kinetic simulation of the si ngle turnover data corresponding to SS2, quinonoid intermediate decay has condensed in to a single step, indicating that the opening of the active site lid to allow ALA dissociation is no longe r a kinetically relevant part of the mechanism (Figure 4.8). Thermodynamic data further support this spectroscopic observation (Figure 4.7 and Ta ble 4.4). Comparison of the wild-type ALASand SS2-catalyzed reacti ons indicates that there is a 6% difference in the Gibb's free energy of the reaction. The decrease of net energy observed for the SS2-variant catalyzed reaction, supports the loss of the kinetic step, as the excess energy required by the wild-type enzyme will have to be absorb ed from the environment, a circumstance whereby the ALAS-catalyzed reaction compensa tes by requiring an additional step. This interpretation is supported by the refined mechanism of th e ALAS-catalyzed reaction by Hunter et. al. wherein the reverse rate of en zyme-ALA1 conversion to enzyme-
155 quinonoid is greater than the fo rward rate of enzyme-ALA1 conversion to enzyme-ALA2 (28) Overall the transient kinetic data suggest that active site lid fl exibility and enzyme activity are tightly coupled a nd that both the rate limitation to the enzyme reaction and conformational motion are the same process dependent, in part, upon the spectrum of amino acids that comprise the motif. In conclusion, a mutational analysis of the active site lid of mALAS2, identified by structural investigation, rev eals a role of the lid in determining the rate limiting step of the enzyme-catalyzed reaction. Variants isolat ed from the library contained mutations to the non-conserved residues that comprise the acti ve site lid, with a marked preference for substitutions toward the carboxy terminal region of the motif. Kinetic characterization of library isolates showed that mutations to the lid result in turnover numbers and catalytic efficiencies that were always greater than those of wild -type ALAS; strongly suggesting that the active site lid is a crucial dete rminant of both substrate binding and product turnover. Additionally, in a subset of varian ts, the rate associated with product release appears independent of a conformational change. The development of tractable fluorescent probes as well as solvent viscosity studies with the variants should prove useful in determining the conformational dynamics of the ALAS-catalyzed reaction.
156Acknowledgements This work was supported by the National Institutes of Health (grant DK63191 to GCF). References (1) Jordan, P. M. (1991) Biosynthesis of Tetrapyrroles Elsevier, Amsterdam. (2) Stackebrandt, E., Murray, R. G. E. and Truper, H. G. (1988) Pr oteobacteria classis nov., a Name for the Phylogenetic Taxon Th at Includes the "Purple Bacteria and Their Relatives". Int. J. Syst. Bacteriol. 38 321-325. (3) Eliot, A. C., and Kirsch, J. F. (2004 ) Pyridoxal phosphate en zymes: mechanistic, structural, and evolutionary considerations. Annu. Rev. Biochem. 73 383-415. (4) Tan, D., and Ferreira, G. C. (1996) Active site of 5-aminolevulinate synthase resides at the subunit interface. Evidence from in vivo heterodimer formation. Biochemistry 35 8934-8941. (5) Christen, P., and Mehta, P. K. (2001) From cofactor to en zymes. The molecular evolution of pyridoxal-5'-phosphate-dependent enzymes. Chem. Rec. 1 436-447. (6) Bottomley, S. S. (2006) Conge nital sideroblastic anemias. Curr. Hematol. Rep. 5 41-49. (7) Astner, I., Schulze, J. O., van den He uvel, J., Jahn, D., Schubert, W. D., and Heinz, D. W. (2005) Crystal structure of 5-aminolevulinate synthase, the first enzyme of heme biosynthesis, an d its link to XLSA in humans. EMBO J. 24 3166-3177. (8) Webster, S. P., Alexeev, D., Campopi ano, D. J., Watt, R. M., Alexeeva, M., Sawyer, L., and Baxter, R. L. (2000) Mechanism of 8-amino-7-oxononanoate synthase: spectroscopic, kinetic, and crystallog raphic studies. Biochemistry 39 516-28. (9) Bottomley, S. S. (2004) Sideroblastic anemias in Wintrobe's Clinical Hematology (Greer, J. F., J. Lukens, J.N. Rodgers, G.M. Paraskevas, R. Glader, B., Ed.) pp 1012-1033, Lippincott, Williams, & Wilkins, Philadelphia. (10) Gong, J., Hunter, G. A., and Fe rreira, G. C. (1998) Aspartate-279 in aminolevulinate synthase affects enzyme catalysis through enhancing the function of the pyridoxal 5'phosphate cofactor. Biochemistry 37 3509-17. (11) Tan, D., Harrison, T., Hunter, G. A., a nd Ferreira, G. C. (1998) Role of arginine 439 in substrate binding of 5-aminolevulinate synthase. Biochemistry 37 14781484. (12) Turbeville, T. D., Zhang, J., Hunter, G. A., and Ferreira, G. C. (2007) Histidine 282 in 5-aminolevulinate synthase a ffects substrate binding and catalysis. Biochemistry 46 5972-5981. (13) Gong, J., Kay, C. J., Barber, M. J., and Ferreira, G. C. (1996) Mutations at a glycine loop in aminolevulinate syntha se affect pyridoxal phosphate cofactor binding and catalysis. Biochemistry 35 14109-14117. (14) Ness, J. E., Kim, S., Gottman, A., Pak, R., Krebber, A., Borchert, T. V., Govindarajan, S., Mundorff, E. C., and Minshull, J. (2002) Synthetic shuffling
157 expands functional protein diversity by allowing amino acids to recombine independently. Nat. Biotechnol. 20 1251-1255. (15) Ostermeier, M. (2003) Synthetic gene lib raries: in search of the optimal diversity. Trends Biotechnol. 21 244-247. (16) Zha, D., Eipper, A., and Reetz, M. T. (2003) Assembly of designed oligonucleotides as an efficient method fo r gene recombination: a new tool in directed evolution Chembiochem. 4 34-39. (17) Ferreira, G. C., and Dailey, H. A. (1993) Expression of mammalian 5aminolevulinate synthase in Escherichia coli Overproduction, purification, and characterization. J. Biol. Chem. 268 584-590. (18) Huala, E., Moon, A. L., and Ausube l, F. M. (1991) Aerobic inactivation of Rhizobium meliloti NifA in Escherichia co li is mediated by lon and two newly identified genes, snoB and snoC. Journal of Bacteriology 173 382-90. (19) Gong, J., and Ferreira, G. C. (1995) Aminolevulinate sy nthase: functionally important residues at a glycine loop, a putative pyridoxal phosphate cofactorbinding site. Biochemistry 34 1678-1685. (20) Li, J. M., Brathwaite, O., Cosloy, S. D., and Russell, C. S. (1989) 5Aminolevulinic acid synthesis in Escherichia coli. Journal of Bacteriology 171 2547-52. (21) Hunter, G. A., and Ferreira, G. C. (1999) Pre-steady-state reaction of 5aminolevulinate synthase. Evidence fo r a rate-determining product release. J. Biol. Chem. 274 12222-12228. (22) Hunter, G. A., and Ferreira, G. C. (1995) A continuous spectrophotometric assay for 5-aminolevulinate synthase th at utilizes substrate cycling. Anal. Biochem. 226 221-224. (23) Hansson, M. D., Karlberg, T., Rahardja, M. A., Al-Karadaghi, S., and Hansson, M. (2007) Amino acid residues His183 and Glu264 in Bacillus subtilis ferrochelatase direct and facilitate the in sertion of metal ion into protoporphyrin IX. Biochemistry 46 87-94. (24) Barshop, B. A., Wrenn, R. F., and Fr ieden, C. (1983) Analysis of numerical methods for computer simulation of kine tic processes: development of KINSIM-a flexible, portable system. Anal. Biochem. 130 134-145. (25) Nandi, D. L. (1978) Studies on de lta-aminolevulinic acid synthase of Rhodopseudomonas spheroides. Reversibility of the reaction, kinetic, spectral, and other studies related to the mechanism of action. Journal of Biological Chemistry 253 8872-7. (26) Hunter, G. A., and Ferreira, G. C. (1999) Lysine-313 of 5-aminolevulinate synthase acts as a general base during formation of the quinonoid reaction intermediates. Biochemistry 38 3711-3718. (27) Onuki, J., Rech, C. M., Medeiros, M. H ., de, A. U. G., and Di Mascio, P. (2002) Genotoxicity of 5-aminolevulinic and 4,5-dioxovaleric acids in the salmonella/microsuspension mutagenicity assay and SOS chromotest. Environ. Mol. Mutagen. 40 63-70. (28) Hunter, G. A., Zhang, J., and Ferrei ra, G. C. (2007) Tran sient kinetic studies support refinements to the chemical and kinetic mechanisms of aminolevulinate synthase. J. Biol. Chem. 282 23025-23035.
158 (29) Karpusas, M., Branchaud, B., and Re mington, S. J. (1990) Proposed mechanism for the condensation reaction of citrate s ynthase: 1.9-A structure of the ternary complex with oxaloacetate and carboxymethyl coenzyme A. Biochemistry 29 2213-2229. (30) McPhalen, C. A., Vincent, M. G., Pico t, D., Jansonius, J. N., Lesk, A. M., and Chothia, C. (1992) Domain closure in mitochondrial as partate aminotransferase. J. Mol. Biol. 227 197-213. (31) Colonna-Cesari, F., Perahia, D., Karp lus, M., Eklund, H., Braden, C. I., and Tapia, O. (1986) Interdomain motion in liver alcohol dehydrogenase. Structural and energetic analysis of the hinge bending mode. J. Biol. Chem. 261 1527315280. (32) Hanson, J. A., Duderstadt, K., Watkins, L. P., Bhattacharyya, S., Brokaw, J., Chu, J. W., and Yang, H. (2007) Illuminati ng the mechanistic roles of enzyme conformational dynamics. Proc. Natl. Acad. Sci. U S A 104 18055-18060. (33) Rozovsky, S., and McDermott, A. E. (2001) The time scale of the catalytic loop motion in triosephosphate isomerase. J. Mol. Biol. 310 259-270. (34) Hu, T., Wu, D., Chen, J., Ding, J., Ji ang, H., and Shen, X. (2008) The catalytic intermediate stabilized by a "down" active site loop for diaminopimelate decarboxylase from Helicobacter pylori Enzymatic characterization with crystal structure analysis. J. Biol. Chem. 283 21284-21293. (35) Venkitakrishnan, R. P., Zaborowski, E., McElheny, D., Benkovic, S. J., Dyson, H. J., and Wright, P. E. (2004) Conformationa l changes in the active site loops of dihydrofolate reductase duri ng the catalytic cycle. Biochemistry 43 16046-16055. (36) Brooks, H. B., and Phillips, M. A. (1997) Characteriza tion of the reaction mechanism for Trypanosoma brucei ornithine decarboxylase by multiwavelength stopped-flow spectroscopy. Biochemistry 36 15147-15155. (37) Codreanu, S. G., Ladner, J. E., Xi ao, G., Stourman, N. V., Hachey, D. L., Gilliland, G. L., and Armstrong, R. N. (2002) Local protein dynamics and catalysis: detection of segmental moti on associated with rate-limiting product release by a glutathione transferase. Biochemistry 41 15161-15172.
159 Chapter Five Summary and Conclusion The three-dimensional structures of Rhodobacter capsulatus ALAS holoenzyme and succinyl-CoA bound ALAS revealed open and closed conformational states of the enzyme, respectively (1) The presence of these two conformational forms agrees with a previous proposal, based on the transi ent kinetic characterization of the ALAS pathway, in which the enzyme undergoes a tr ansition from the open to the closed state upon succinyl-CoA binding and returns to the open conformation upon ALA release (2, 3) A closer look at the ac tive site shows that a conser ved serine residue (S189 in R. capsulatus ALAS and S254 in meALAS) changes the orientation of its hydrogenbonding pattern in a succinyl-CoA-dependent manner (1) Succinyl-CoA binding induced dramatic changes in the visible CD spectra of both the wild-type and S254A proteins. Additionally, this amino acid substitu tion elicited a 2-fold increase in enzyme activity, while simultaneously increasing the Km for succinyl-CoA 25-fold. These experimental findings coupled w ith the structural data sugges t that this residue is an important determinant in conformer equilib rium by promoting energetically favorable interactions between the chro mophore and succinyl-CoA, and by stabilizing a closed Michaelis-complex c onfiguration. To address the functionality of S254 in the ALAS-catalyzed reaction, kinetics experiments were performed on wild-type ALAS and two ALAS variants (S254A and
160 S254T). Substitution of serine with alanin e eliminates hydrogen bond formation between the amino acid and the phenolic oxygen of PLP and succinyl-CoA. Notably, steady-state kinetic parameters calculated for this variant indicate that both the turnover number and the Michaelis constant for succinyl-CoA increased. This finding implies unusual functional complexity regarding the correlation between this mutation and the enzymecatalyzed reaction. The means by which the enzyme manages to increase activity despite a reduction in affinity for the more comp lex of the two substrates may include a mechanism whereby the equilibrium is predominantly shifted toward the closed conformational state. The S254A mutation has notable effect s on the cofactor microenvironment as determined by CD spectroscopy. CD spectro scopic evaluations of the conformational effects of the S254A and S254T mutations illustrate the differences between the two amino acid substitutions. Succinyl-CoA bi nding to the wild-type and S254T variant induced changes in the CD spectra associ ated with the microenvironment of the chromophore, while CD spectra correspondin g to the S254A mutant were unchanged under similar conditions. This dissimilarity between wild-type ALAS and S254A may be the result of a partial convers ion of the internal aldimine to free PLP aldehyde present in the active site, a circumstance which is observed in three out of the four R. capsulatus crystal structure active s ites upon succinyl-CoA binding (1) In the crystal structures these events are coincident with the clos ed conformation, from which it might be concluded that the S254A variant retains th e internal aldimine in the presence of succinyl-CoA, and may not be induced to adopt a closed conformation upon binding of this substrate.
161 Pre-steady-state kinetic analyses of both the wild-type ALASand variantcatalyzed reactions show that upon decarboxylation, the ALA-bound quinonoid intermediate is formed followed by two successively slower steps in which the intermediate decays. These two steps are assigned to protonation of the ALA-quinonoid intermediate and ALA release, respectively (3) For the S254T variant, the rate of quinonoid intermediate formation decreased 4fold compared to that of wild-type enzyme. This reduction may indicate a change in the flow of electrons from the site of bond scission to the resonance stabilized carbanion, induced by a shift of the conformational equilibrium towards the closed conformation. Changes in the position of PLP observed in the three-dimensional structur es available for ALAS show a shift of 15 degrees when substrate is bound (1) These observations, coupled with experimental evidence suggesting that even subtle changes to the stereoelectronic parameters of the external aldimine and PLP modulate cataly sis, indicate that the hydrogen bonding potential of position 254 in ALAS may be an important feature in the regulation of the transition from the open to the closed conformation (4, 5) Together the data for both ALAS variants support a postulate whereby non-covalent forces between S254 and the phenolic oxygen influence an induced fit mech anism in which substrate recognition is coupled to conforma tional equilibria. ALAS is a member of the -oxoamine synthase subf amily of pyridoxal 5'phosphate (PLP)-dependent enzymes and shares a high degree of struct ural similarity and reactivity with the other members of the family (6) Despite the remarkable structural and mechanistic similarities in this im portant group of enzymes the molecular mechanisms underlying substrate specificity remain largely unexplored. The X-ray
162 crystal structure of ALAS from Rhodobacter capsulatus reveals that the alkanoate component of succinyl-CoA is coordinated by a conserved arginine and a threonine (1) The functions of the corresponding acyl-CoA-bi nding residues in murine erythroid ALAS (R85 and T430) in relation to acyl-CoA bi nding and substrate discrimination were examined using site-directed mutagenesi s and a series of CoA-derivatives. The acyl-CoA substrate binds to the en zyme through an interaction between the pantetheine moiety of CoA with the enzyme surface and via hydrogen-bonding interactions between the terminal alkanoate group and R85 and T430 (1) The steadystate kinetic analysis of th e variants (R85K, R85L, and R 85L/T430V) with the family of CoA-derivatives showed that the apparent Michaelis parameters are dramatically different when compared to those of wild-type ALAS. Acyl-CoA substrates of increased hydrophobicity ( e.g ., octanoyland butyryl-CoA) bound with higher affinity to variants where the substituted amino acid was aliphatic in nature (R85L and R85L/T430V). The 36-fold decrease in substrate bi nding for octanoyl-CoA in th e R85L variant suggests that the exclusion of water from the acyl-CoA-binding cleft is an important feature of substrate binding. It is li kely that reaction specificity is driven by the chemical characteristics of the CoA-derived ta il and the hydrogen-bonding potential of the invariant acyl-CoA-binding residues, a circ umstance which has been proposed for the acyl-CoA thioesterases of the peroxisome (7, 8) The use of chemically different acyl-CoA -derivatives affects the transient kinetic parameters of a variant enzyme (R85K) signifying potential alterations to a key mechanistic step in the ALAS-catalyzed reacti on. Single turnover reac tions with a family of CoA-derivatives were used to determine the rates of quinonoid intermediate formation
163 and decay for the R85K variant. The R85K variant-catalyzed reactio n was decelerated in the first step of quinonoid intermediate d ecay, with a 10-fold lower rate for both octanoyl-CoA and glutaryl-CoA when compared to the physiological substrate succinylCoA. When R85 is mutated to a lysine, the enzyme is chemically similar to wild-type ALAS in many respects, presumably because this conservative replacement retains the positive charge and hydrogen bonding capabilities. However, the molecular volume of the amino acid side chain is different. With respect to their n -alkyl moieties, the n propylguanidine side chain of ar ginine is longer than the n -butylamine side chain of lysine by 1.6 (9) The R85K substitution could therefore accommodate the additional sp3 hybridized carbon atom present in glutar yl-CoA, allowing for a reduction in steric strain and/or unfavorable van der Waals interactions. Both of these possibilities could be contributing factors in molecular recognition of the physiological subs trate succinyl-CoA. In all, the experimental data support multif unctional roles for these amino acids (R85 and T430) in regulating substrate specificity, and linking the bi furcate coordination of the acyl-CoA tail with the mechanistic chemistry of the active site. In X-ray crystal structures of 8-amino-7-oxononanoate synthase from E. coli and R. capsulatus ALAS, a loop, covering a conserved sequence of amino acids, was shown to migrate 3.5 and 5.5 between the holoenzym ic forms and acyl-CoA-bound forms of the two enzymes, respectively (1, 10) Comparison of holoenzymic ALAS and AONS with the substrateand product-bound forms of th e enzymes indicates that binding of these ligands within the active site precipitates movement of the loop. These structural observations suggest, but do not prove, that the dynamics of this active site lid may signify a change in conformation. In order to examine the role of this active site loop (or
164 lid) in the ALAS-catalyzed reaction, we used genetic mani pulation to identify functional amino acid mutations and to evaluate the contribution of lid residues to catalysis. The approach used to evaluate the active site lid of ALAS utilized a mutagenic technique called synthetic shuffling (11) The active site lid of ALAS contains 18 residues; 8 of these are completely conserve d, while the remaining amino acids show no pattern with respect to evolution (1) Mutations were observed throughout the entire motif. However, more mutations were found in the carboxy-terminal portion of the lid compared to the segment closer to the ami no-terminus (31 to 12, respectively). This inequality suggests that greater plasticity is associated with the Cterminal part of the loop. Each of the isolated variants was show n to have increased turnover numbers and enhanced catalytic efficiency with both subs trates. These characteristics, identified among active site lid variants, indicate th at the reaction catalyzed by ALAS may be limited by the amino acid composition of th e lid. Further, conformational changes centered in active si te loops have been repo rted to have important mechanistic roles in other enzymes (12, 13) Accordingly, mutations th at affect the mobility of a conformationally mobile enzyme structure ma y have consequences in which the physical step of product release, rather than chemistry, becomes rate limiting. Therefore the enhanced turnover observed among the active site lid variants potentia lly validates our ALAS catalytic mechanism in which the enzy me conformation switches between closed and open to stabilize reaction intermedia tes and release product, respectively, a mechanism recently proposed for another PLP-dependent enzyme, diaminopimelate decarboxylase (14)
165 The pre-steady-state behavi ors of three variants from the library (SS2, F1 and H1)differed from that observed with the wild -type enzyme and other library isolates: the two steps associated with the quinonoid interm ediate decay were condensed into a single step. For the SS2 variant-catalyzed reacti on, quinonoid intermediate formation is ~20fold faster than the corresponding rate in wi ld-type ALAS, suggesting that a step after catalysis is only partially rate -determining for this variant. Together with the greater dissociation constant of the SS2 variant fo r ALA (Table?), the data suggest that a conformational change leadi ng to product release no longer is rate-limiting for the SS2catalyzed reaction. Indeed, studies on several other enzymes where a chemical step has been implicated as rate-limiting show that the effects of conformational changes on kcat were the result of a thermodynamically driven commitment to catalysis (15-17) In fact, increases in exothermy and pos itive differential activation en tropy were also determined for the SS2 variant-catalyzed reaction. Based on the kinetic simulation of the single turnover data for the reaction catalyzed by SS2, quinonoid intermediate decay has co ndensed into a single step, indicating that the opening of the active site lid to allow ALA dissociation is no longer a kinetically relevant part of the mechanism. Thermodynamic data further support this spectroscopy-derived observation. The de crease of net energy observed for the SS2 variant-catalyzed reaction, suppor ts the loss of this kinetic step. Conversely, the excess energy required by the wild-type enzyme woul d likely have to be absorbed from the environment, a circumstance whereby the AL AS-catalyzed reaction would compensate by requiring an additional step (3) This interpretation is supported by the refined mechanism of the ALAS-cat alyzed reaction by Hunter et. al. wherein the reverse rate of
166 enzyme-ALA1 conversion to enzyme-quinonoid is greater than the forward rate of enzyme-ALA1 conversion to enzyme-ALA2 (3) Overall the transient kinetic data suggest that active site lid flexibility and enzyme activity are tigh tly coupled and that both the rate limitation to the enzyme reaction and conformational motion are the same process, dependent, in part, upon the spectrum of amino acids that comprise the motif. The kinetic parameters calculated for the lib rary isolates show that these mutations confer hyperactivity. Through these investig ations, hypotheses pertai ning to the reaction mechanism of ALAS have implications in the pathways of disease involving both iron metabolism and porphyrin biogenesis. Data presented here and co nclusions set forth regarding limits to ALA turnove r and individual steps associ ated with substrate binding and rates of reaction, specifically address f acets of XLSA and the recently discovered erythroid ALAS-relate d porphyria, X-linked do minant protoporphyria (19) Many mutations associated with diminished ALAS activity in vivo are located in the PLPbinding cleft (1, 19) Significantly, the conformationa lly responsive S254 residue forms a hydrogen bond with the phenolic oxygen of the cofactor. Enhanced turnover of ALA, and subsequent biosynthesis of porphyrins and porphyrin precursors are the foreseeable consequences of the reactions catalyzed by isolates from the active site lid library. Indeed, these circumstances resemble the pa thology that is associated with X-linked dominant protoporphyria (18) The knowledge related to ALAS from the st udies presented here contribute to the goal of therapeutic interven tion of porphyrin-accumulative diseases, like cancer. Further, biochemical developments in the field of biotechnology will also be enhanced through this research by addressing the mol ecular requirements of treatments like
167 photodynamic therapy. In conclusion, these data are likely to provi de insight into the rate-determining step of enzymes limited by a product release, or a conformational change leading to product release. References (1) Astner, I., Schulze, J. O., van den Heuvel, J., Jahn, D., Schubert, W. D., and Heinz, D. W. (2005) Crystal structure of 5-aminolevulinate synthase, the first enzyme of heme biosynthesis, and its link to XLSA in humans. Embo J. 24 31663177. (2) Hunter, G. A., and Ferreira, G. C. (1999) Pre-steady-state reaction of 5aminolevulinate synthase. Evidence fo r a rate-determining product release. J. Biol. Chem. 274 12222-12228. (3) Hunter, G. A., Zhang, J., and Ferreira, G. C. (2007) Transient kinetic studies support refinements to the chemical and kinetic mechanisms of aminolevulinate synthase. J. Biol. Chem. 282 23025-23035. (4) Turbeville, T. D., Zhang, J., Hunter, G. A., and Ferreira, G. C. (2007) Histidine 282 in 5-aminolevulinate synthase a ffects substrate binding and catalysis. Biochemistry 46 5972-5981. (5) Tai, C. H., Rabeh, W. M., Guan, R., Schnackerz, K. D., and Cook, P. F. (2008) Role of Histidine-152 in cofactor orient ation in the PLP-dependent O-acetylserine sulfhydrylase reaction. Arch. Biochem. Biophys. 472 115-125. (6) Eliot, A. C., and Kirsch, J. F. (2004) Pyridoxal phosphate enzymes: mechanistic, structural, and evolutionary considerations. Annu. Rev. Biochem. 73 383-415. (7) Hunt, M. C., and Alexson, S. E. (2008) Novel functions of acyl-CoA thioesterases and acyltransferases as auxiliary enzymes in peroxisomal lipid metabolism. Prog. Lipid Res. 47 405-421. (8) Hunt, M. C., Solaas, K., Kase, B. F., and Alexson, S. E. (2002) Characterization of an acyl-coA thioesterase that functi ons as a major regulator of peroxisomal lipid metabolism. J. Biol. Chem. 277 1128-1138. (9) Creighton, T. R. (1983) Proteins, Structures and Molecular Properties W.H. Freeman and Company, New York. (10) Webster, S. P., Alexeev, D., Campop iano, D. J., Watt, R. M., Alexeeva, M., Sawyer, L., and Baxter, R. L. (2000) Mechanism of 8-amino-7-oxononanoate synthase: spectroscopic, kinetic, and crystallogr aphic studies. Biochemistry 39 516-528. (11) Ness, J. E., Kim, S., Gottman, A., Pak, R., Krebber, A., Borchert, T. V., Govindarajan, S., Mundorff, E. C., and Mi nshull, J. (2002) Synthetic shuffling expands functional protein diversity by allowing amino acids to recombine independently. Nat. Biotechnol. 20 1251-1255. (12) Hanson, J. A., Duderstadt, K., Watkins, L. P., Bhattacharyya, S., Brokaw, J., Chu, J. W., and Yang, H. (2007) Illuminati ng the mechanistic roles of enzyme conformational dynamics. Proc. Natl. Acad. Sci. U S A 104 18055-18060.
168 (13) Rozovsky, S., and McDermott, A. E. (2001) The time scale of the catalytic loop motion in triosephosphate isomerase. J. Mol. Biol. 310 259-270. (14) Hu, T., Wu, D., Chen, J., Ding, J., Ji ang, H., and Shen, X. (2008) The catalytic intermediate stabilized by a "down" active site loop for diaminopimelate decarboxylase from Helicobacter pylori Enzymatic characteri zation with crystal structure analysis. J. Biol. Chem. 283 21284-21293. (15) Brooks, H. B., and Phillips, M. A. (1997) Characterizat ion of the reaction mechanism for Trypanosoma brucei ornithine decarboxylase by multiwavelength stopped-flow spectroscopy. Biochemistry 36 15147-15155. (16) Codreanu, S. G., Ladner, J. E., Xiao, G., Stourman, N. V., Hachey, D. L., Gilliland, G. L., and Armstrong, R. N. (2002) Local protein dynamics and catalysis: detection of segmental motion associated with rate-limiting product release by a glutathione transferase. Biochemistry 41 15161-15172. (17) Venkitakrishnan, R. P., Zaborowski, E., McElheny, D., Benkovic, S. J., Dyson, H. J., and Wright, P. E. (2004) Conformationa l changes in the ac tive site loops of dihydrofolate reductase duri ng the catalytic cycle. Biochemistry 43 16046-16055. (18) Whatley, S. D., Ducamp, S., Gouya, L., Grandchamp, B., Beaumont, C., Badminton, M. N., Elder, G. H., Holme, S. A., Anstey, A. V., Parker, M., Corrigall, A. V., Meissner, P. N., Hift, R. J., Marsden, J. T., Ma, Y., MieliVergani, G., Deybach, J. C., and Puy, H. (2008) C-terminal deletions in the ALAS2 gene lead to gain of f unction and cause X-linked dominant protoporphyria without anemia or iron overload. Am J Hum Genet 83 408-14. (19) Bottomley, S. S. (2006) C ongenital sideroblastic anemias. Curr. Hematol. Rep. 5 41-49.
169 About the Author Thomas Lendrihas received two Bachel or of Science degrees in Biology and Chemistry and Bachelor of Arts Degree in Mu sic from Eckerd College in 2002, where he was a Ford Foundation Undergraduate Research Scholar. He had a Master of Arts degree in Medical Bioethics and Humanities from Un iversity of South Florida, College of Medicine in 2007. Since 2003, he has been a graduate student in the Ph.D. program in the Department of Molecular Medicine, Co llege of Medicine, Un iversity of South Florida, Tampa, FL. In addition to his scientific accomplishments, Thomas received acclaim as a classical pianist, giving recitals and performing in concert.
xml version 1.0 encoding UTF-8 standalone no
record xmlns http:www.loc.govMARC21slim xmlns:xsi http:www.w3.org2001XMLSchema-instance xsi:schemaLocation http:www.loc.govstandardsmarcxmlschemaMARC21slim.xsd
leader ntm 2200397Ka 4500
controlfield tag 001 002324025
008 101007s2009 xx a ob 000 0 eng d
datafield ind1 8 ind2 024
subfield code a E14-SFE0003057
b L564i 2009
Investigation into the rate-determining step of mammalian heme biosynthesis :
molecular recognition and catalysis in 5-aminolevulinate synthase /
by Thomas Lendrihas.
xii, 168 leaves :
Dissertation (Ph.D.)--University of South Florida, 2009.
Includes bibliographical references.
Text (Electronic dissertation) in PDF format.
ABSTRACT: The biosynthesis of tetrapyrolles in eukaryotes and the alpha-subclass of purple photosynthetic bacteria is controlled by the pyridoxal 5-phosphate (PLP)-dependent enzyme, 5-aminolevulinate synthase (ALAS). Aminolevulinate, the universal building block of these macromolecules, is produced together with Coenzyme A (CoA) and carbon dioxide from the condensation of glycine and succinyl-CoA. The three-dimensional structures of Rhodobacter capsulatus ALAS reveal a conserved active site serine that moves to within hydrogen bonding distance of the phenolic oxygen of the PLP cofactor in the closed, substrate-bound enzyme conformation, and simultaneously to within 3-4 angstroms of the thioester sulfur atom of bound succinyl-CoA. To elucidate the role(s) this residue play(s) in enzyme activity, the equivalent serine in murineerythroid ALAS was mutated to threonine or alanine.The S254A variant was active, but both the KmSCoA and kcat values were increased, by 25- and 2-fold, respectively, suggesting the increase in turnover is independent of succinyl-CoA-binding. In contrast, substitution of S254 with threonine results in a decreased kcat, however the Km for succinyl-CoA is unaltered. Removal of the side chain hydroxyl group in the S254A variant notably changes the spectroscopic properties of the PLP cofactor and the architecture of the PLP-binding site as inferred from circular dichroism spectra. Experiments examining the rates associated with intrinsic protein fluorescence quenching of the variant enzymes in response to ALA binding show that S254 affects product dissociation. Together, the data led us to suggest that succinyl-CoA binding in concert with the hydrogen bonding state of S254 governs enzyme conformational equilibria.As a member of the alpha-oxoamine synthase family, ALAS shares a high degree of structural similarity and reaction chemistry with the other enzymes in the group. Crystallographic studies of the R. capsulatus ALAS structure show that the alkanoate component of succinyl-CoA is bound by a conserved arginine and a threonine. To examine acyl-CoA-binding and substrate discrimination in murine erythroid ALAS, the corresponding residues (R85 and T430) were mutated and a series of CoA substrate analogs were tested. The catalytic efficiency of the R85L variant with octanoyl-CoA was 66-fold higher than that calculated for the wild-type enzyme, suggesting this residue is strategic in substrate binding. Hydrophobic substitutions of the residues that coordinate acyl-CoA-binding produce ligand-induced changes in the CD spectra, indicating that these amino acids affect substrate-mediated changes to the microenvironment of the chromophore.Pre-steady-state kinetic analyses of the R85K variant-catalyzed reaction show that both the rates associated with product-binding and the parameters that define quinonoid intermediate lifetime are dependent on the chemical composition of the acyl-CoA tail. Each of the results in this study emphasizes the importance of the relationship between the bifurcate interaction of the alkanoic acid component of succinyl-CoA and the side chains of R85 and T430.From the X-ray crystal structures of Escherichia coli 8-amino-7-oxonoanoate synthase and R. capsulatus ALAS, it was inferred that a loop covering the active site moved 3-6 between the holoenzymic and acyl-CoA-bound conformations. To elucidate the role that the active site lid plays in enzyme function, we shuffled the portion of the murine erythroid ALAS cDNA corresponding to the lid sequence (Y422-R439), and isolated functional variants based on genetic complementation in an ALA-deficient strain. Variants with potentially greater enzymatic activity than the wild-type enzyme were screened for increased porphyrin overproduction. Turnover number and the catalytic efficiency of selected functional variants with both substrates were increased for each of the enzyme variants tested, suggesting that increased activity is linked to alterations of the loop motif. The results of transient kinetics experiments for three isolated variants when compared to wild-type ALAS showed notable differences in the pre-steady-state rates that define the kinetic mechanism, indicating that the rate of ALA release is not rate-limiting for these enzymes. The thermodynamic parameters for a selected variant-catalyzed reaction indicated a reduction in the amount of energy required for catalysis. This finding is consistent with the proposal that, in contrast to the wild-type ALAS reaction, a protein conformational change associated with ALA release no longer limits turnover for this variant enzyme.
Advisor: Gloria C. Ferreira, Ph.D.
X-linked sideroblastic anemia
t USF Electronic Theses and Dissertations.