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Theses and Dissertations

6-1-2009

Investigation into the rate-determining step ofmammalian heme biosynthesis: Molecularrecognition and catalysis in 5-aminolevulinatesynthaseThomas LendrihasUniversity of South Florida

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Scholar Commons CitationLendrihas, Thomas, "Investigation into the rate-determining step of mammalian heme biosynthesis: Molecular recognition andcatalysis in 5-aminolevulinate synthase" (2009). Theses and Dissertations. Paper 2059.http://scholarcommons.usf.edu/etd/2059

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, α-oxoamine synthase, transient kinetics, pyridoxal 5'-phosphate, porphyria, photodynamic therapy

© Copyright 2009, Thomas Lendrihas

Acknowledgements

I wish to express 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 support throughout the course of my graduate

work. Most of all, to Dr. Gloria C. Ferreira, I am deeply appreciative 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 Molecular Medicine, for their intellectual and

personal contributions. To Dr. Gregory A. Hunter and Dr. Tracy D. Turbeville, I am

indebted for both their scientific and emotional 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 administrative 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. Knowles, Mitchell M. McNelly, Laura Jackson

Roberts, Louis J. Smith and Thomas F. Zarella, for their enduring encouragement and

love. Finally, I wish to acknowledge my family, 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: biogenesis, 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 steady-state kinetic parameters. 66 Table 2.2. Gibb’s free energy associated with the S254 variant-catalyzed

reactions. 78 Table 3.1. Comparison of steady-state kinetic constants for wild-type ALAS,

R85K, R85L, and R85L/T430V with CoA derivatives as substrates. 95

Table 3.2. Rates of quinonoid intermediate formation and decay under single-

turnover conditions. 103 Table 4.1. Designed mutations for incorporation at indicated positions

within the ALAS active 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 parameters 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 structure 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 structures of coproporphyrinogen III

oxidase. 22 Figure 1.5. The three-dimensional structure of ferrochelatase from Homo

sapiens. 28 Figure 1.6. Enzymes in the heme degradation pathway. 32 Figure 2.1. Structural models for murine erythroid ALAS based on the R.

capsulatus crystal structures. 57 Figure 2.2. Multiple sequence alignment of phylogenetically diverse

members of the α-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 the S254 variants (60 µM) with increasing

concentrations of glycine. 69 Figure 2.5. Reaction of wild-type ALAS and the S254 variants (5 µM)

with ALA. 70 Figure 2.6. Reaction of wild-type ALAS- and 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 different 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 µM) with ALA. 101 Figure 3.5. Reaction of wild-type ALAS- and R85K-glycine complexes

with different CoA derivatives 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 screening of the library. 126 Figure 4.3. Differential fluorescence of ALAS variant 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 reactions of isolated hyperactive ALAS

variants. 141 Figure 4.6. The SS2 variant-catalyzed reaction. 147 Figure 4.7. The thermal dependence of the SS2-variant catalyzed reaction. 148 Figure 4.8. The simulated kinetic mechanism 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 synthase (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] piperazine-N’-[2-ethane sulfonic acid])

HMB Hydroxymethylbilane

HO Heme oxygenase

HRM Heme regulatory motif

INH 4-Bromo-3-(5'-carboxy-4'-chloro-2'-fluoro-phenyl)-1-methyl-5-

trifluoromethyl-pyrazol

IRE Iron response element

IRP IRE-binding protein

KBL 2-Amino-3-ketobutyrate-CoA ligase

meALAS Murine erythroid ALAS

mno montalcino (zebrafish variant displaying 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 ALAS-

catalyzed 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 tetrapyrolles in eukaryotes and the -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 murine

erythroid ALAS was mutated to threonine or alanine. The S254A variant was active, but

both the SCoAmK 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

xi

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 -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

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 Å 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.

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 transcriptional repressors such as Bach-1 (4, 5). Message

translation is highly regulated through the phosphorylation of eucaryotic translation

initiation factor 2A (eIF2α) by a number of heme-dependent kinases (6-9).

Hierarchically, among eucaryotes and the α-subclass of purple photosynthetic

bacteria, the heme biosynthetic 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 ferrous 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 regulated process involving heme oxygenase

and biliverdin reductase, which together degrade heme to carbon monoxide, iron and

bilirubin (14). Heme, present in hemoproteins, is degraded 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-limiting step of heme degradation is potently

cytoprotective (20). Several disease states are associated 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 structures provide the framework for identifying

structural components of both the molecular basis 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, the enzyme on which my theses are based.

Enzymes in the heme biosynthesis pathway

Aminolevulinate synthase

5-Aminolevulinate synthase (EC 2.3.1.37) (ALAS), is the first, and key regulatory

enzyme of heme biosynthesis in non-plant eucaryotes and the α-subclass of purple

photosynthetic bacteria (Figure 1.1) (23). ALAS catalyzes the Claisen-like condensation

3

of glycine and succinyl-Coenzyme-A to yield coenzyme-A (CoA), carbon dioxide (CO2)

and 5-aminolevulinate (ALA) (23). For the remaining monera, and in all plants, ALA is

synthesized from an alternative pathway involving the five carbon skeleton of glutamate

(24, 25). Interestingly, the photosynthetic phytoflagellate Euglena gracilis synthesizes

ALA via both pathways, with non-plastid heme synthesized from glycine or the Shemin

pathway, and chloroplast heme coming from glutamate-derived ALA (26).

The ALAS-catalyzed reaction takes place in the mitochondria of non-plant

eucaryotes (11). This reaction is tightly regulated, and the rate of reaction determines the

anabolic production of downstream metabolites in the pathway (27, 28). Mammalian

genomes contain two genes which code for two 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 X-

chromosome at band Xp11.21 (30). The nucleotide sequences of the two genes are

notably different; however, the two protein 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 weight after the presequence is clipped (33).

Heme and iron regulate both gene expression and sub-cellular localization of

ALAS. Mammalian ALAS proteins contain heme regulatory motifs (HRM) which

consist of a conserved dipeptide, Cys-Pro (32). By way of HRMs, depleted intracellular

heme pools inhibit the mitochondrial translocation of ALAS1 (34). Plentiful heme on the

other hand in differentiating erythrocytes does not contribute to

4

Figure 1.1. Enzymes and intermediates of the heme biosynthetic pathway.

Mito

chon

drio

n

Cyt

osol

5

mitochondrial import of the ALAS2 enzyme (34). Transcription of the ALAS1 gene is

upregulated by peroxisome proliferator-activated coactivator 1α (PGC-1α) (35), through

promoter-mediated interactions with both nuclear regulatory factor 1 (NRF-1) and the fox

head family member FOX01 (36). Transcription of the ALAS2 gene is regulated by

erythroid-specific factors including GATA-binding protein 1 (globin transcription factor

1), a protein which is chiefly responsible for the activation of globin production in red

cells (37). The resultant ALAS2 transcript contains a 5' iron regulatory element (IRE)

which binds with the IRE-binding 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 prosthetic group within the protein restricts

formation of the IRE-IRP-1 complex, permitting 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-3-

ketobutyrate-CoA ligase (KBL), 7-amino-8-oxononanoate synthase (AONS), and serine

palmitoyl transferase (SPT) (40-43). Based on structure and function, ALAS is classified

as a fold-type I pyridoxal-5’-phosphate (PLP)-dependent enzyme and as a member of the

α-oxoamine synthase subfamily; AONS, KBL and SPT represent the closest structural

relatives, with the enzymes of the subfamily sharing a Cα RMSD of 1.5 Å (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 liberated and the aminoketone product of the enzyme-catalyzed

reaction is further metabolized. Enzymes in the α-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 subunit, 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 homodimer, where each monomer consists

of three domains. The N-terminal domain, discrete 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 helices. The C-terminal domain independently

interacts with the N-terminal domain through three alpha-helices and with the central

core domain of the enzyme via a three-stranded, anti-parallel beta-sheet. The orientation

of the PLP cofactor can be considered to occur through interactions with three specific

protein moieties. First, the phosphate group is bound tightly via 6 hydrogen bonds

(where three are intrasubunit, and three intersubunit). Second, pi-stacking interactions

between the conjugated systems of the cofactor and a conserved active site histidine also

stabilize the position of PLP. Third, hydrogen bonding between a conserved aspartate

residue and the pyridinium nitrogen enhance the electron withdrawing properties of the

cofactor. This PLP microenvironment and adjacent C-terminal domain delimit the

substrate-binding channel, connecting the solvent exposed surface of the enzyme with the

hydrophobic core of the enzyme, where the acyl-CoA-binding cleft is located (49).

Studies focused on the role conserved amino acids play in the reaction catalyzed

by murine ALAS revealed several notable findings. First, the catalytic lysine (K313)

(ALAS2 numbering), essential for enzyme activity 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 withdrawing capacity of the PLP cofactor was also

found (51). Recognition of the carboxyl group of the 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 orientation and electronics of the PLP cofactor

(55).

The ALAS chemical mechanism (Scheme 1.1) is complex and involves: binding

of glycine (I); transaldimination with the active site lysine (K313) 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 α-amino-β-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 rate-

determining step of the ALAS-catalyzed reaction 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 reaction rates, when measured with a variety

of acyl-CoA derivatives, are comparable (56).

The proposed model of ALAS catalysis, based on kinetic data obtained in both the

steady- and 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 binding energy of succinyl-CoA is utilized to

8

R=OPO3

2-

Scheme 1.1. The chemical mechanism of ALAS. The individual steps are: binding of

glycine (I); transaldimination with the active site lysine (K313, murine erythroid ALAS

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 α-amino-β-

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).

9

favor the conversion of the population of equilibrium 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 surrounded by iron-laden mitochondria (60), and thus the

designation of ringed sideroblasts. Diminished ALAS2 activity in red blood cells is the

main reason why retained iron is a hallmark 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 symptoms including: fatigue, disorientation and both hepato- and

splenomegaly (62, 63). The toxicity of excess iron has 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 leading 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 and the succinyl-CoA synthetase reveal that abrogation of

this association may be responsible for the pathological 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 porphyrin-associated disorder when defective (69).

Now, ALAS2 gene deletions have been identified in eight families with the following

genetic manifestations c.1706-1709 delAGTG (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 protoporphyrin IX. Consequently, these variants

demonstrate the first instance of an erythroid ALAS-related porphyria, X-linked

dominant protoporphyria. With this finding, several features of the biochemical

mechanism of this mode of disease require further research. First, since the experiments

were performed on lysates of bacterial cells harboring the expression plasmid for the

truncated proteins and not with deletion-variants purified to homogeneity, the opportunity

for other proteins and metabolites affecting the reaction cannot be ruled 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. Certainly, elimination of a

protein sequence of this magnitude could potentially 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 mitochondrion of eukaryotic cells remains to

be elucidated. Once in the cytosol, 2 molecules of ALA are asymmetrically condensed

by the metalloenzyme porphobilinogen synthase (EC 4.2.1.24; PBGS or ALA

dehydratase (ALAD)) (Figure 1.1) (72). The formation of porphobilinogen (PBG), a

monopyrrole, is the first common step of tetrapyrrole biosynthesis among all living

organisms. PBGS isolated from different organisms, from bacteria to humans, share a

high degree of sequence identity (73). These enzymes are large, exhibiting homo-

octameric quaternary structure, and molecular 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 N-terminal 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 quaternary structure differently, and the result is

an oligomer of distinctive functionality (77).

Catalysis by PBGS begins with the formation of independent Schiff base bonds

between 2 conserved active site lysine residues and the substrates (78, 79). The

destination of the ALA molecule within the monopyrrole 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 acetyl component of PBG, while the propionyl-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) links with ALA in the P-site (C4) position

12

via 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. Three-dimensional structures obtained with a

series of substrate and product analogs have been completed, and the role of an active-

site 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 residues 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 oligomerization of mutated enzymes occurs in a

manner that favors the less active hexameric state. These hexamers, resulting from eight

porphyria-associated variants, may be the first example of a morpheein-based

conformational disease. Diminished enzyme activity leads to the accumulation of ALA.

13

Symptoms associated with the poor porphobilinogen production include: intermittent

acute neurovisceral attacks and a propensity toward poisoning by lead (86). Inhibition of

this enzyme by succinylacetone, a metabolite found in among individuals with hereditary

tyrosinemia type I, also causes pathological conditions similar to those 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 4.3.1.8; PBGD) or hydroxymethylbilane

synthase) (Figures 1.1 and 1.2). The physiologically relevant reaction product is the

linear tetrapyrrole hydroxymethylbilane (HMB) a.k.a. pre-uroporphyrinogen (74). HMB

is exceedingly unstable and can undergo spontaneous cyclization to form the non-

physiological isomer uroporphyrinogen I (88). PBGD sequences are highly conserved

throughout evolution and among diverse 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-dimensional structural analysis shows that the

monomeric protein is organized into three equal-sized domains. Domain I houses most

of the catalytic and substrate-binding 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 α-helices (green) and β-sheets (rust) (A). Perched

at the top of the enzyme is the unique dipyrromethane cofactor (depicted as sticks in CPK

color format) (B).

A

B

15

hexapyrrole. Subsequently, PBGD cleaves the distal tetrapyrrole and releases HMB. A

thioether linkage to an active site cysteine retains 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 leading to diminished enzyme activity (12).

Hundreds of PBGD mutations have been identified, with acute attacks of porphyria

affecting 1:100000 individuals, with presentation more common in women than men

(92). However, only recently was a poly-deletion mutant identified in exon 15 of the

PBGD gene (93). The deletion occurs in a conserved region of the protein 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 circulatory disturbances, ultimately resulting

in tachycardia (95). A majority of the mutations identified in the human PBGD gene

perturb carboxylate binding between conserved arginines and the cofactor or substrate

(96). However, recently, some of the mutations 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 fourth enzyme of the heme biosynthetic

pathway, uroporphyrinogen III synthase (EC 4.2.1.75) (UROS) (Figure 1.3). UROS

catalyzes closure of the tetrapyrrole macrocycle 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 contains two unambiguous

domains connected by a short β-ladder (yellow). Each domain is characterized by a β-

sheet core (magenta) surrounded by α-helices (teal).

17

molecular mass; however significant sequence deviations 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 crystallographically unique UROS structures

(consisting of apoenzymic, ligand-bound and product-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 unambiguous domains connected

by a short β-ladder (103). Each domain is characterized by a β-sheet core surrounded by

α-helices. The substrate binding cleft is located at the domain 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 loss of the C20 hydroxyl group and the formation

of a carbocation at C20. C16 of the D-ring is then susceptible to electrophilic attack and

the spirocyclic pyrrolenine intermediate is generated. Subsequently, azafluvene is

formed permitting cyclization and D-ring 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 structural domains of UROS are the chief

modulators of the closed enzyme conformation (104). The C and D rings demonstrate

increased flexibility, a characteristic consistent with the sterically-mediated acts of

catalytic cyclization and D ring inversion. Biological relevance of all porphyrins is

18

demarcated by asymmetry about the D-ring of tetrapyrolles. Without enzymatic

conversion, HMB spontaneously cyclizes to a toxic dead end 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 Günther disease, HMB is non-enzymatically

converted to uroporphyrinogen I and is subsequently 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 cloned into expression vectors, and the

respective proteins were purified to homogeneity and characterized (106). Kinetics

experiments indicated that most mutated enzymes had significantly decreased activity,

while others maintained reaction rates comparable to those of the wild-type enzyme.

This suggested that mechanisms besides turnover may be responsible for the pathology

observed in CEP. Located in α-helix 3, perched above the active-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 mutation-free form of the enzyme (107). The causative

agent for this pathology was faulty transcription, a defect which was linked to a mutation

19

in the erythroid-specific GATA1 transcription factor. These particular data provide

evidence that the functional responsiveness of erythroid specific promoters is different.

Uroporphyrinogen decarboxylase

The acetate side chains of uroporphyrinogen III are decarboxylated in four

successive steps by uroporphyrinogen decarboxylase (EC 4.1.1.37; UROD), leaving the

four methyl groups characteristic of the product coproporphyrinogen 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 isolated UROD proteins is low;

however common structural features have been 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 (β/α) barrel, with the active site housed at the

end of a deep cleft, delimited by the C-terminal loops of the barrel. Evolutionarily

conserved residues line the cleft, and several invariant basic residues are crucial for

binding the propionate groups of the substrate (109, 110). A dynamic active site loop

located at the head of the active site undergoes conformational 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 groups proceeds in a clockwise manner (114, 116, 117).

To accomplish this, sequential decarboxylation requires the 180o reorientation of the

intermediate, a process whereby the substrate is flipped around its C10-C20 axis.

Controversy exists as to the mechanism by which 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 collaborate during the decarboxylation of a single

uroporphyrinogen substrate. It has been proposed that the dimeric organization of UROD

protects the reactants from solvent exposure, allowing the reaction intermediates to be

passed and chemically modified between each monomer (114, 116). Further, analysis of

UROD compared with another cofactorless decarboxylase, orotidine 5'-monophosphate

decarboxylase (ODCase), indicates that a protonated basic residue assists the transition of

the polar carboxylate groups from water to the comparatively less polar hydrophobic core

by stabilizing the post-scission 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 for all three probands indicated two

novel missense mutations and one previously identified polymorphism, giving these

patients an unique and previously unidentified compound heterozygote genotype (120).

Dermatitic photosensitivity is the hallmark symptom of PCT (119); a clinical

21

manifestation which shows a marked increase in intensity among individuals with co-

morbidities 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 rings A and B to form protoporphyrinogen IX

(Figure 1.1). Coproporphyrinogen oxidase (EC 1.3.3.3; CPO) in humans is oxygen-

dependent, found in the intermembranous space of mitochondria and produces

coproporphyrinogen III, carbon dioxide 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 prosthetic groups or cofactors has not been

identified (127). The human enzyme exists as a homodimer, with a monomeric mass of

~37 kDa (Figure 1.4A) (128). CPO contains an elaborate subunit interface with 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 anti-

parallel β-sheet flanked by α-helices. The active site, delimited on both sides by the β-

sheet and helices, elegantly houses the substrate while minimizing contacts with the

solvent. At the head of the active site, an α-helix acts as a lid to modulate the solvation-

state 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 anti-

parallel β-sheet (red) flanked by α-helices (green) (A). The E. coli enzyme (PDB code:

1olt) functions as a dimer (monomer shown) (B) and contains a catalytically essential S-

adenosyl-L-methionine cofactor and a [4S-4S] cluster (C).

B

A

C

23

Oxygen-independent CPOs are remarkably 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-L-

methionine (SAM) (Figure 1.4C). As an oxidizing 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 physiological electron depository for the product of

the aerobic reaction. With respect to the mechanism of catalysis, the anaerobic

conversion of coproporphyrinogen III to protoporphyrinogen IX is only partially

understood (130, 131), although two of the steps are well documented. These steps are

the generation of the 5'-deoxyadenosyl radical 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 histidines of human CPO, suggests that

catalysis can occur despite alterations to these evolutionarily selected residues (132).

Further work by the same group using substrate analogs where the C and D rings were

modified to replace alkyl groups with the ring 13- and 17-propionate moieties led the

investigators to postulate that the propionate 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 rotates) remains to be elucidated.

Mutation in the human CPO gene are associated with hereditary 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 mapped 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 substrate and bifurcate interactions between

the carboxylate tail of the substrate and two arginine residues (136). However, only one

arginine residue (Arg401) has been reported 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 10–200 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 pathway, protoporphyrinogen oxidase (EC 1.3.3.4;

PPO) (Figure 1.1) (139). This step of the pathway constitutes a branch point whereby

protoporphyrin IX is supplied to produce chlorophyll 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 dinucleotide 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

25

tabacum (144). This enzyme exists as a loosely associated dimer with three defined

monomeric domains. The domains are responsible for FAD-, protoporphyrinogen IX-,

and membrane-binding. The active site of PPO is located between the FAD- and the

substrate-binding domains. Mutations associated with the human condition, variegate

porphyria (VP) have been mapped using the plant model with notable success (145).

A binding mechanism for the PPO-catalyzed reaction has been proposed based on

experimental results obtained with the INH (4-bromo-3-(5'-carboxy-4'-chloro-2'-fluoro-

phenyl)-1-methyl-5-trifluoromethyl-pyrazol) and acifluorfen inhibitors (144, 146). The

pyrazole ring of INH serves as an A ring model, which is coordinated in the active site by

pi-stacking interactions with a conserved phenylalanine residue. Ring B of the

macrocycle substrate is stabilized by hydrophobic interactions provided by the side

chains of two highly conserved leucine residues. Oxidation of the C20 methylene bridge

between rings A and D occurs by way of the FAD cofactor. Next, imine-enamine

tautomerizations initiate all hydride transfers from C20. The three remaining oxidation

reactions involving FAD generate three hydrogen peroxide molecules. Curiously, the

candidates for the catalytic base involved in this reaction are not evolutionarily conserved

(144). As such, a potential base which could execute proton abstraction from the

substrate is the FADH--derived peroxide anion.

PPO deficiency causes VP (147). Missense, nonsense, and 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 prevalent in South Africans of European descent

(148). A founder mutation attributed 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 abnormal gene has resulted in very high

prevalence; as many as twenty thousand South Africans may carry this gene (149). VP

presents as acute neurovisceral attacks and/or dermatitic photosensitivity. It is the

variability observed in pathological presentation 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 identifying silent carriers of mutated genes (150).

Efforts to examine the mechanism of presentation 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 human PPO mRNA was

microinjected into mno embryos. Rescue of the mno phenotype by overexpression of

human PPO suggests functional conservation of the enzyme across chordates.

Consequently, mno appears an excellent model for investigation of PPO and a valuable

tool for identification of therapeutic agents of VP. Increased plasma porphyrin in VP is

detected by monitoring fluorescence emission at 626–628 nm, upon excitation at 405 nm

(152). Other porphyrias including EPP and PCT are marked by fluorescence emission

peaks 636 nm and 618–622 nm, upon excitation 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 4.99.1.1) (FC) is the last enzyme in the heme biosynthetic

pathway, and it catalyzes the insertion 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 interact 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 three-dimensional structure 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 [2Fe-

2S] cluster. Curiously B. subtilis FC is devoid of this prosthetic group, and exists as a

monomer (160). The function of these iron-sulfur 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 β-sheet delimited by an α-helix in a β-α-β 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 interdomain in nature. Metal binding and catalysis

likely take place within this region and involve several evolutionarily conserved residues.

Sub-cellular localization studies 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 tunnel, which permits substrate association

28

Figure 1.5. The three-dimensional structure of ferrochelatase from Homo sapiens

(PBD code: 1hrk). The 2Fe-2S cluster is coordinated by four cysteine residues. This

prosthetic group is located at the subunit interface of the functional dimer.

29

and product release; a scenario mediated by the conserved hydrophobic 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 subsequent metallation with ferrous iron (164).

Distortion of the tetrapyrrole macrocycle is proposed to occur after its binding to the

active site cleft of FC (165). This step has been identified as a defining feature of

ferrochelatase-catalyzed metallation. In fact, 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 porphyrin, which in turn

affects metallation efficiency (165).

Chelatases catalyze the insertion 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 tetrapyrroles, several theories exist. 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 bacterial 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 conserved acidic amino acids comprising a π-helix.

While the function of this helix remains unclear, hypotheses regarding the regulatory

nature of this motif have been suggested (169, 172). The mechanism by which

metallation takes places requires further research, 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 oligomer with iron bound, as well as in vitro assays

proving protein-protein interactions between this protein and FC, it is proposed that

frataxin delivers ferrous iron to FC (165, 175). Certainly, the channeling of metal atoms

by way of a chaperone-ferrochelatase complex has the advantages of minimizing Fenton

chemistry between the substrate and the aqueous environment. Understanding the

relationship between iron-delivery and heme synthesis represents an intriguing future

direction of ferrochelatase-related research.

Mutations in the FC gene result in a congenital disorder called, erythropoietic

protoporphyria (EPP) (176). EPP is transmitted as an autosomal dominant trait; however

evidence suggests that a second defective copy is necessary for pathological presentation

(177, 178). Accumulation of protoporphyrin is localized to reticulocytes, and is a

significant component of bile (12). Patients with protoporphyrin-rich bile often show

cholestasis, cirrhosis and remarkably fluorescent gall stones (12). Of all the porphyrias,

EPP results in the greatest dermatitic photosensitivity. Most EPP patients function with

FCs that turnover substrate at a rate less than 20% of that of the wild-type protein (12).

While this diminished activity likely stems from mutations in the protein; a common

31

wild-type FC allelic variant has been identified 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 nature. The overproduction of protoporphyrin is

also linked to acute liver damage, highlighting 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 1.14.99.3; HO) (Figure 1.6) has 3 isoforms (14). The first,

HO-1, is a highly inducible 32-kDa protein, that catalyzes the first and rate-limiting step

in the degradation of heme from red blood 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 unresponsive 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 have physiological and

pathological functions which include protection 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 intermediates of the heme catabolic pathway.

33

and degenerative diseases (20, 184, 185). Further, the heme catabolic pathway is of

major importance to the degradation of the globins and other hemoproteins, many of

which are affective stressors of the aforementioned disordered states.

Mammalian HO-1 proteins share a high degree of sequence similarity. Among

eucaryotes, a conserved C-terminal hydrophobic tail of ~20 residues appears to function

in anchoring HO-1 to the microsomal membrane. 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 sequence. Several bacterial HOs, including that

from Neisseriae meningitides, have been crystallized and require an NADH reductase for

enzymatic activity (187). These enzymes function as water-soluble monomers of ~25

kDa. In procaryotes, HOs function to release iron to the environment, a process which

increases microbial survival and pathogenesis, and mitigates heme toxicity (187). The X-

ray 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 α-helices (189). An evolutionarily conserved histidine in the proximal helix is

the axial ligand for the substrate (189). On the opposite side, an α-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 α-meso edge of heme is pointed toward the protein interior and

is swaddled by a series of conserved hydrophobic residues. Conformational

heterogeneity is observed in the HO-1-heme complex 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 α-meso heme position is oxidized by diatomic oxygen (O2) in the presence

of NADPH to yield α-hydroxyheme. Subsequently, the α-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, the main photoreceptor for photosynthesis.

Transcriptional regulation of the HO-1 gene involves activators such as Nrf2 and

repressors such as the heme-binding protein Bach-1 (192). Both the activating and

repressing factors require heterodimerization 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 allows 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

1.3.1.24; BVR) (Figure 1.6 (193, 194), an enzyme that reduces the C10 bridge of

biliverdin. BVR is evolutionarily conserved 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 mammalian species, BVR is greater than 80%

identical, a degree of similarity bolstered 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 Src 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, NADPH is utilized in the basic range (201).

While the structural basis for the unique cofactor/pH-dependence 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 and without the cofactor bound (203). In the rat crystal,

extensive interactions between the enzyme termini occur by way of a β-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 NADH-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 would 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 interactions between the enzyme and the

cofactor. Second, substrate specificity related to succinyl-CoA is examined through

molecular interactions between two conserved residues (Arg85 and Thr430 in murine

ALAS) and the chemical nature of the acyl-CoA-derived tail. Third, the rate-determining

step of the enzyme-catalyzed reaction is addressed in the context of the conformational

mobility of an active site loop. The conclusions set forth in each chapter are related to

the advancement of knowledge regarding not only the ALAS-catalyzed reaction, but

reactions catalyzed by members of the α-oxoamine synthase subfamily and PLP-

dependent enzymes as a group.

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Chapter Two

Serine-254 enhances an induced fit mechanism in murine 5-aminolevulinate

synthase

Abstract

5-Aminolevulinate synthase (EC 2.3.1.37) (ALAS), is a homodimeric pyridoxal

5'-phosphate (PLP)-dependent enzyme and catalyzes the initial step of the heme

biosynthetic pathway in animals, fungi, and some bacteria. This reaction involves the

condensation of glycine and succinyl-Coenzyme 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, substrate-

bound enzyme conformation, and simultaneously to within 3-4 angstroms of the thioester

sulfur atom of bound succinyl-CoA. To evaluate 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 SCoAmK and kcat values are

increased, by 25- and 2-fold, respectively, suggesting unusual functional complexity. In

contrast, the S254T mutation results in a significant decrease in kcat without

altering SCoAmK . Circular dichroism spectroscopy reveals that removal of the side chain

hydroxyl group in the S254A variant dramatically alters the PLP microenvironment as

52

well as the responsiveness of this microenvironment to succinyl-CoA binding. Protein

fluorescence stopped-flow experiments confirm that the mutations differentially alter the

rates of conformational responsiveness to ALA 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 α-oxoamine synthases.

53

Introduction

5-Aminolevulinate synthase (EC 2.3.1.37; ALAS) is a homodimeric PLP-

dependent enzyme that catalyzes the first and key regulatory enzyme of the heme

biosynthetic pathway in non-plant eucaryotes and the α-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 characterized 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 PLP-

binding site (5, 6).

The ALAS chemical mechanism (Scheme 2.1) is complex and involves a high

degree of stereoelectronic control, with individual 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 α-amino-β-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) (7). This mechanistic complexity is manifested

structurally as an enzyme with an unusually 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 enzyme 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 α-oxoamine synthase subfamily, each of

which catalyze reactions between small amino acids and CoA esters to generate 1,3-

aminoketones, while also sharing 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 conformation, consistent with the induced fit

hypothesis, where electrostatic and hydrophobic interactions between the substrates,

cofactor, and amino acids comprising the active site provide the energetic impetus to

stabilize this catalytically optimal conformation (15, 16).

Prior to solution of an ALAS crystal structure, kinetic data led investigators to

propose that ALAS transitions between open and closed conformations during the

catalytic cycle (7, 17). Steady-state kinetic experiments demonstrate that the kinetic

mechanism is ordered, with glycine binding before succinyl-CoA, yet in transient kinetic

studies binding of succinyl-CoA accelerates the apparent rate at which glycine binds to

ALAS by over 250,000-fold (18). This enhancement might occur by utilization of part of

the intrinsic binding energy for succinyl-CoA to shift the enzyme conformer equilibrium

towards a closed conformation wherein transaldimination 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 closed conformations, respectively, further

supporting the hypothesis that enzyme dynamics play a crucial role during the ALAS

catalytic cycle (8). While the structure in general collapses slightly around the bound

substrates, a more conformationally mobile loop of amino acids located between two β-

sheets at the subunit interface closes directly over the channel leading approximately 20Å

down into the deeply recessed active site (Figure 2.1) (8). A conserved threonine at the

apex of the mobile loop forms a strong hydrogen bond (~2.5Å) with the carboxylate tail

of succinyl-CoA in the substrate-bound structure and appears to simultaneously provide

molecular recognition for succinyl-CoA while helping to lock this substrate 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 this serine may be an important determinant in conformer

equilibrium and catalysis by providing orientational binding energy between the cofactor

and substrate, while stabilizing a closed Michaelis complex conformation. The

conservation of this residue in ALAS and the other α-oxoamine synthases suggests an

important functionality that may be general 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 the positionally equivalent S254A and S254T

variants and investigated the effects of these mutations on the kinetic and spectroscopic

properties of the enzyme. The results support the postulate that S254 is a key

57

A B

C

multifunctional residue that couples succinyl-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 modeled by alignment of open

holoenzyme and closed glycine and succinyl-CoA bound monomeric structures. Serine-

254 is hidden by the succinyl-CoA ester in this view from the perspective 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 other structural features are for the closed

conformation. (B) Serine-254 in the open conformation. (C) Serine-254 in the closed

conformation with succinyl-CoA bound.

58

Figure 2.2. Multiple sequence alignment of phylogenetically diverse members of the

α-oxoamine synthase family in the region of murine eALAS serine-254. The amino

acid sequences were retrieved from public databases (NCBI) and aligned using

CLUSTAL W (21). The conserved serine residue is high-lighted in cyan. The amino

acid numbering in red refers to that of murine erythroid 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---------TPKIVAFETVHSMDGAICPLEELCD H. sap. AL2 NDPDHLKKLLEKSNPK---------IPKIVAFETVHSMDGAICPLEELCD H. sap. AL1 NDVSHLRELLQRSDPS---------VPKIVAFETVHSMDGAVCPLEELCD S. cer. ALA NDLNELEQLLQSYPKS---------VPKLIAFESVYSMAGSVADIEKICD R. cap. ALA NDVAHLRELIAADDPA---------APKLIAFESVYSMDGDFGPIKEICD A. nig. AON SCPRSLEDVLRREVEGDE-MVRNGKKNVFLVIESIYSMDGDIAPIREFVE A. tha. AON CDMYHLNSLLSNCKMKR----------KVVVTDSLFSMDGDFAPMEELSQ M. mar. AON NNTVDLIEIL-EKN-KN-------YENKFIVTDAVFSMDGDIAPVGELKK E. col. AON NDVTHLARLLASPCPGQ----------QMVVTEGVFSMDGDSAPLAEIQQ H. sap. KBL LDMADLEAKLQEAQKH---------RLRLVATDGAFSMDGDIAPLQEICC C. kor. KBL CDLADLEDKL-RQVHKK-------YNKILIITDGVFSMDGDIAPLDGITK E. col. KBL NDMQELEARLKEAREAG-------ARHVLIATDGVFSMDGVIANLKGVCD H. sap. SPT NNMQSLEKLLKDAIVYGQPRTRRPWKKILILVEGIYSMEGSIVRLPEVIA S. cer. SPT GDMVGLEKLIREQIVLGQPKTNRPWKKILICAEGLFSMEGTLCNLPKLVE A. tha. SPT NTPGHLEKVLKEQIAEGQPRTHRPWKKIIVVVEGIYSMEGEICHLPEIVS E. col. SPT NDAKDLERRMVRLGER--------AKEAIIIVEGIYSMLGDVAPLAEIVD * : .: : .** * : .

59

Materials

Reagents. The following were purchased from Sigma-Aldrich Chemical

Company (St. Louis, MO): ampicillin, DEAE-Sephacel, Ultrogel AcA-44, -

mercaptoethanol, PLP, bovine serum albumin, succinyl-CoA, ALA-hydrochloride, -

ketoglutaric acid, -ketoglutarate dehydrogenase, HEPES-free acid, MOPS, tricine,

thiamine pyrophosphate, NAD+, and the bicinchoninic acid protein determination kit.

Glucose, glycerol, glycine, disodium ethylenediamine tetraacetic acid dihydrate,

ammonium sulfate, magnesium chloride hexahydrate, and potassium 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 synthesized by Integrated DNA Technologies

(Coralville, IA).

Methods

Mutagenesis. The pGF23 expression plasmid encoded the full-length sequence for

the murine, mature eALAS (22). Site-directed mutagenesis for the S254A and S254T

mouse ALAS variants was performed on the whole plasmid pGF23 using a previously

described method (23). The mutagenic oligonucleotides for S254A and S254T were: 5’-

GAG ACT GTT CAT GCC ATG GAT GGT GCC-3’ and 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 presence of the intended 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 steady-state analysis.

Recombinant murine eALAS and the S254 variants were purified from DH5

Escherichia coli bacterial cells containing the overexpressed protein as previously

described (22). Purity was determined by SDS-PAGE (24) and protein concentration

determined by the bicinchoninic acid method using 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 determined 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 were collected using an AVIV CD spectrometer

calibrated for both wavelength maxima and signal intensity with an aqueous solution of

D-10 camphorsulfonic acid (29). CD spectra recorded from 190-240 nm were analyzed

by the ridge regression method using a modified version of the computer program

CONTIN developed by Provencher and Glöckner (30). Protein concentrations were 10

M and 100 M for the near and far CD spectra, respectively. 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 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 samples were collected immediately after adding the enzyme. The spectra of the

samples containing enzyme were analyzed after substracting the blank spectra. Co-factor

fluorescence spectra were collected on a Shimadzu RF-5301 PC spectrofluorophotometer

using protein concentrations of 2 μM. 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 experiments were carried out at 30°C in 100

mM HEPES, pH 7.5 and 10% (v/v) glycerol. The 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 concentrations were used to ensure the

identification of a single enzyme-catalyzed event. For the S254A-catalyzed reaction, the

final concentrations were: 120 μM S254A, 130 mM glycine, and 30 μM succinyl-CoA.

The final concentrations of the reactants for S254T were: 50 μM S254T, 130 mM

glycine, and 10 μM succinyl-CoA. 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

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 syringes and observation chamber. Specifically, spectral data

covering the 270 – 550 nm range were analyzed 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 judged 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 mechanism as described by Equation 2.1.

(Equation 2.1) D C B A

k k k 3obs

2obs

1obs

The observed rate constants were determined from at least three replicate

experiments, and the reported values represent the average and standard error of

measurement for each experimental condition. The forward and reverse rate constants

depicted in the kinetic mechanism (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 the reaction of glycine with the variant enzymes. The

reactions of the murine eALAS variants with glycine were performed using the same

instrument as was described for the single turnover reactions with the enzyme-glycine

complex and succinyl-CoA. The final enzyme concentration was 60 μM. The glycine

concentration was always at least 10-fold greater than the enzyme concentration to ensure

that pseudo-first order kinetics 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

tkit ceaA i

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 concentrations 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 nonlinear regression analysis program SigmaPlot10

(Systat, San Jose, CA)

(Equation 2.3) C B A

k k 2obs

1obs

(Equation 2.4) 11

][

][

k

SK

Skk

Dobs

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 state kinetics of the

product binding reaction of ALAS and the two serine variants were examined by

measuring changes in the intrinsic protein fluorescence intensity. An OLIS RSM-1000F

rapid mixing spectrofluorimeter, equipped with 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 and ligand in the reaction chamber were 1/2 of

64

those in the syringes. The intrinsic protein fluorescence, as measured with 5 μM enzyme,

was evaluated in the presence of increasing concentrations 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 (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 2.5, using the Global fitting software provided with the instrument.

(Equation 2.5) 01)( AeAtF tkobs

obs

where Fobs(t) is the observed fluorescence change (in arbitrary units) at time t, kobs is the

observed first-order rate constant, A1 is the pre-exponential factor and A0 is the offset. The

observed rate constants were then plotted against ligand concentration and the data were

fitted to Equation 2.6 by nonlinear regression. The rates of dissociation (koff) and

association (kon) as well as the ligand binding constants (KD) were calculated from the

asymptotic maximal observed rate, the ordinate intercept, and the ligand concentration (x)

in Equation 2.6.

(Equation 2.6) xK

xkkxf

D

offon )(

65

Results

Kinetic characterization of the S254 variants. The steady-state kinetic parameters

of the S254 variants were determined and the results are summarized in Table 2.1. The

mutation of serine-254 to alanine resulted in a kcat 2-fold higher than that 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 overall catalytic efficiency for

succinyl-CoA decreased 36-fold, while the value for glycine remained unchanged as

compared to ALAS values. The replacement of serine-254 with threonine caused a 2-

fold decrease in the wild-type kcat value. The Michaelis constants 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 active site were perturbed by the replacement of

the serine with either an alanine or a threonine. Gross structural changes evidenced by

changes in the alpha helix and beta sheet content 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 variants are negligible. Locally chiral

substructures that comprise the PLP microenvironment modulate the visible CD

characteristics of the chromophore. Spectra for the wild-type and variant enzymes, as

holo- and succinyl-CoA-bound enzymes, were collected (Figure 2.3). The spectra for

wild-type ALAS and S254T holoenzymes displayed 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 GlymK

(mM)

SCoAmK

(µM) catk

(s-1) catk / Gly

mK

(mM·s-1) catk / SCoA

mK

(µM·s-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 emission 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 μM succinyl-CoA; (D) upon excitation

of the cofactor at 331 nm.

68

(Figure 2.3B). Wild-type ALAS and variant enzymes responded differently to the

presence of 100 μM ligand (Figure 2.3C). Comparison 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 under 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 excitation at 331 nm is similar to that of the wild-type

enzyme with a well-formed emission maximum ~385 nm (Figure 2.3D). A notable

deviation in emission maximum is observed 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 profile for the alanine variant

suggest that the tautomeric equilibrium between at least two forms of the PLP cofactor

aldimine linkage with the catalytic 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 reaction of S254A with glycine 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 fluorescence quenching were determined 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 µM) with increasing concentrations of

glycine. Data were fit to Equation 2.2 for a two-exponential process, yielding

equilibrium and rate constants for S254A of KD = 6.6 ± 0.57 mM, k1 = 0.159 ± 0.04 s-

1, 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 µM)

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 µM,

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 µM,

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 µM, 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 change 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 succinyl-

CoA. ALAS catalysis involves the sequential binding of glycine 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 reaction under single turnover conditions. 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 tested, 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 spectral 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 chemistry during catalysis. The rates assigned to

72

Figure 2.6. Reaction of wild-type

ALAS- and 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 S254A-

catalyzed 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 quinonoid intermediate decay rates that 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 increased 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 intermediate indicate fast transformation of the

enzyme-substrate complex to the enzyme-product complex (Qf and Qd1), followed by a

comparatively faster rate of product dissociation and the return of the conformational

equilibrium to the catalytically favored open conformation (Qd2).

74

Discussion

The spectroscopic and kinetic properties of the S254A and S254T murine ALAS

variants were examined and compared to those of the wild-type enzyme in an effort to

better understand the role of this residue in the catalytic process. Mutation of serine to

alanine removes the side chain oxygen atom and eliminates the possibility of hydrogen

bond formation with the PLP cofactor and succinyl-CoA. The conservative 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 determined under similar conditions reveals the unusual kinetic

significance of S254 (Table 2.1). Point mutations typically result 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 increased 25-fold over that

of the wild-type enzyme, while kcat is increased 2-fold. It is notable that mutation of an

evolutionarily conserved residue 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 variant may be attributable to

diminished stability of the closed conformation, 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 would 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 conservative S254T mutation would then arise

primarily from enhanced stability of the closed conformation. Both mutations

substantially diminish the catalytic efficiency with succinyl-CoA (kcat/SCoAmK ), but in

different ways, and while the effects of these very conservative mutations on the steady-

state kinetic parameters may appear relatively 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 microenvironment, as

determined by fluorescence and CD spectroscopies. The fluorescence spectra suggest a

rather dramatic alteration in cofactor tautomeric equilibria occurs upon loss of hydrogen

bonding interactions between the phenolic oxygen 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 conformational effects of the S254A and S254T

mutations in the far-UV region verify the mutations do not significantly 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 visible 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 arise from Cotton effects associated with the

ketoenamine (435 nm) and enolamine (330 nm) cofactor aldimine bond tautomers, and

are indicative of the microenvironment surrounding 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 S254A mutant is relatively

unchanged under similar conditions (Fig. 2.3C). A logical interpretation is that the

decrease in asymmetry observed for the wild-type and S254T variants arises from partial

conversion of the internal aldimine to free PLP aldehyde bound at the active site, as is

observed in three out of four R. capsulatus crystal structure active sites upon succinyl-

CoA binding (8). In the crystal structures these events 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 succinyl-CoA, and may not be induced to adopt a

closed conformation upon binding of this substrate.

The quinonoid intermediate single-turnover profiles of the two S254 variants

indicate that they follow a chemical mechanism similar to that of the wild-type enzyme

(Fig. 2.6A, B). A rapid step of ALA-bound quinonoid intermediate formation upon

decarboxylation is followed by two successively slower decay steps, associated with

protonation of the ALA-quinonoid intermediate and ALA release, respectively (7). The

quinonoid intermediate formation rate decreased 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, precipitated by a shift in the conformational equilibrium towards

closed. In the ALAS crystal structure, PLP is noted to change position by 15 degrees

when substrate is bound (8). Changes to the stereoelectronic relationship between the

cofactor and the α-carbon bonds of the external aldimine can influence the chemical

mechanism dramatically (7). Therefore, it is possible 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 decay 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 variant is also consistent with the increases in

kcat and the ALA off rate determined from enzyme fluorescence quenching.

By utilizing the microscopic parameters obtained from the single turnover

reactions of the variant enzymes, coupled with the product and substrate reaction data, we

were able to model the kinetic mechanisms as shown in Figure 2.7. Additionally, the

Gibb’s free energy associated with the glycine and ALA were calculated (Table 2.2).

The kinetic simulations revealed that the S254T mutation significantly retards the

chemical mechanism. Conversely, the modeled pathway for S254A highlights the

increases observed in the kinetics of the variant. Overall, the mechanistic data for both

variants support the hypothesis that the interaction 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 indistinguishable from those completed based on the

global fit of spectral data. E, ALAS; G, glycine; EG, ALAS-glycine complex; SCoA,

succinyl-CoA; EGSCoA, ALAS-glycine-succinyl-CoA complex; EQ, observable

quinonoid intermediate; EALA1, ALAS-ALA internal aldimine with active site loop

closed; and EALA2, ALAS-ALA internal aldimine with active site loop open.

Table 2.2. Gibb’s free energy associated with the wild-type ALAS- and S254

variant-catalyzed reactions.

Enzyme ΔGALA

(kcal/mol) ΔGGly

(kcal/mol) Wild-type 4.18 3.34 S254A 4.08 3.97 S254T 4.07 2.78

79

Acknowledgements

This work was supported by the National Institutes of Health (grant DK63191 to GCF).

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(12) McPhalen, C. A., Vincent, M. G., Picot, D., Jansonius, J. N., Lesk, A. M., and Chothia, C. (1992) Domain closure in mitochondrial aspartate aminotransferase. J. Mol. Biol. 227, 197-213.

(13) Christen, P., and Mehta, P. K. (2001) From cofactor to enzymes. The molecular evolution of pyridoxal-5'-phosphate-dependent enzymes. Chem. Rec. 1, 436-447.

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(14) Alexander, F. W., Sandmeier, E., Mehta, P. K., and Christen, P. (1994) Evolutionary relationships among pyridoxal-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 Christen, P. (1984) Crystallographic studies on the mechanism of action of mitochondrial 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 Christen, P. (1984) Three-dimensional structure of mitochondrial aspartate aminotransferase and some functional 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 5-aminolevulinate synthase. Evidence for a rate-determining product release. J. Biol. Chem. 274, 12222-12228.

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(26) Hunter, G. A., and Ferreira, G. C. (1995) A continuous spectrophotometric assay for 5-aminolevulinate synthase that 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.

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(32) Durbin, J., and Watson, G. S. (1970) Testing for serial correlation in least squares regression. Biometrika 37, 409-414.

(33) Barshop, B. A., Wrenn, R. F., and Frieden, C. (1983) Analysis of numerical methods for computer simulation of kinetic 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, 349-384.

(35) Tai, C. H., Rabeh, W. M., Guan, R., Schnackerz, K. D., and Cook, P. F. (2008) Role of Histidine-152 in cofactor orientation 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-aminolevulinate synthase. Protein Sci. 2, 1959-1965.

82

Chapter Three

Arg-85 and Thr-430 in murine 5-aminolevulinate synthase coordinate acyl-CoA-

binding and contribute to substrate specificity

Abstract

5-Aminolevulinate synthase (ALAS) catalyzes the rate-limiting step of heme

biosynthesis in mammals through the condensation of succinyl-Coenzyme A and glycine

to produce 5-aminolevulinate, Coenzyme-A (CoA) and carbon dioxide. ALAS is a

member of the α-oxoamine synthase family of pyridoxal 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 structure 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-CoA-

binding residues in murine erythroid ALAS (R85 and T430) in relation to acyl-CoA

binding and substrate discrimination were examined using site-directed mutagenesis and

a series of CoA-derivatives. The catalytic efficiency of the R85L variant with octanoyl-

CoA was 66-fold higher than that of the wild-type protein, supporting the proposal of this

residue as key in discriminating substrate binding. Substitution of the acyl-CoA-binding

residues with hydrophobic amino acids caused a ligand-induced negative dichroic band at

420 nm in the CD spectra, suggesting that these residues affect substrate-mediated

83

changes to the PLP microenvironment. Transient kinetic analyses of the R85K variant-

catalyzed reactions confirm that this substitution decreases microscopic rates associated

with formation and decay of a key reaction intermediate and show that the nature of the

acyl-CoA tail seriously affect product binding. These results 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.

84

Introduction

5-Aminolevulinate synthase (ALAS; EC 2.3.1.37) is a pyridoxal 5’-phosphate

(PLP)-dependent enzyme consisting of two 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-4-

oxopentanoate (5-aminolevulinate; ALA), and represents the first step of porphyrin

biosynthesis in animals, fungi, and some bacteria. 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 Å resolution and also as enzyme-substrate complexes with either

glycine (2.7 Å) or succinyl-CoA (2.8 Å) (4). ALAS is classified as a member of the α-

oxoamine synthase subfamily of fold type I PLP-dependent enzymes. AONS, SPT, and

KBL are the other members and represent the closest structural relatives, with the

enzymes of the subfamily sharing a Cα root mean square deviation of approximately 1.5

Å (5, 6). The reaction chemistries are also highly similar, all involving small amino

acids, CoA esters, and 1,3-aminoketones. AONS catalyzes the committed step in biotin

biosynthesis,(7) SPT catalyzes the first step of sphingolipid biosynthesis,(8) and KBL

catalyzes the degradation of threonine (9).

Despite the remarkable structural and mechanistic similarities in this important

group of enzymes the molecular mechanisms underlying substrate specificity remain

largely unexplored. SPTs utilize palmitoyl-CoA 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 similar 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 reaction was examined with 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 acyl-CoA substrate are important

factors in determining specificity. The CoA substrate specificity of ALAS is of interest

due to localization of the eukaryotic enzyme in the inner mitochondrial matrix.

Specifically, 90% of cellular acetyl-CoA and between 92- and 97% of short and long

chain acyl-CoAs are located within this organelle, providing an abundant supply of

possible alternative substrates for meALAS. As such, promiscuous reactions with

alternative CoA substrates would produce highly reactive 1-3-aminoketones of unknown

biological significance, that are potentially capable of dimerizing to form toxic

dihydropyrazines (13, 14).

Previous investigations regarding the binding 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 glycine-binding 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 factors 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 β-strand bent around the PLP cofactor, in which the pyridinium ring of the cofactor lies

at the bottom of the cavity (4). Connection between the surface of the enzyme and the

active site is by an amphipathic channel, which is occupied by succinyl-CoA in the

substrate-bound structure (Figure 3.1). Two distinct moieties of succinyl-CoA interact

with the enzyme: the solvent accessible adenosyl component and the buried succinate.

The alkanoic acid moiety of succinyl-CoA is bound to the active site via a strong

hydrogen bond network that stabilizes a closed 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 erythroid 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 characteristics 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 investigate the role of the conserved R85 and T430 residues of

meALAS in recognition and binding of the acyl-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 effect of hydrophobicity and steric hindrance

on specificity toward the CoA-derived substrate. Such a difference would alter the

aliphaticity of the substrate-binding cleft, which, in turn, could affect the acyl chain-

87

Figure 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 chains of the conserved 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 here for the R85 and T430

variants of meALAS show that these residues are involved in both the orientation and

binding of the succinyl-CoA substrate in the active site and may also, following the

substrate binding, assist in enzyme closure.

Materials

Reagents. The following reagents were purchased from Sigma-Aldrich Chemical

Company (St. Louis, MO): ampicillin, DEAE-Sephacel, Ultrogel AcA-44, -

mercaptoethanol, PLP, bovine serum albumin, succinyl-CoA, ALA-hydrochloride, -

ketoglutaric acid, -ketoglutarate dehydrogenase, HEPES-free acid, MOPS, tricine,

thiamine pyrophosphate, NAD+, and the bicinchoninic acid protein determination kit.

Glucose, glycerol, glycine, disodium ethylenediamine tetraacetic acid dihydrate,

ammonium sulfate, magnesium chloride hexahydrate, and potassium hydroxide were

acquired from Fisher Scientific (Pittsburgh, PA). Sodium dodecyl sulfate polyacrylamide

gel electrophoresis reagents were 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 encodes the full-length sequence for

the murine, mature eALAS (17). The R85L variant was generated using a previously

described method (18). The mutagenic primers used for the R85L mutation were 5’-

GAC CAC ACC TAC CTT GTG TTC AAG ACT GT-3’ and 5’-ACA GTC TTG AAC

ACA AGG TAG GTG TGG TC-3’, with the introduced codon substitution underlined.

89

The PCR-generated fragment containing the R85L mutation was used as a megaprimer in

an independent round of PCR. The PCR-generated fragment was sequenced to verify the

presence of the desired mutations. Subsequently, the PCR product was then digested

with Sal I and Bam HI and subcloned into pGF23 digested similarly. The site-directed

mutagenesis for the T430V variant was performed on the whole plasmid pGF23 using a

previously described method (19). The mutagenic oligonucleotide for T430V was: 5’-

ATC AAC TAC CCA GTT GTG CCT CTG GGT-3’, with 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 pGF23 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 constructed 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 overexpressed 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 using bovine serum albumin as the standard

(21). All protein concentrations are reported on the basis of a subunit molecular weight

90

of 56 kDa. Enzymatic activity was determined 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 dialyzed 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 maxima and signal intensity with

an aqueous solution of D-10 camphorsulfonic acid (25). Protein concentrations were 100

μM for each enzyme tested. The final concentration 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 samples containing enzyme were analyzed after

subtracting the blank spectra.

Stopped-flow spectroscopy. All of the experiments were carried out at 30°C 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 pre-incubated in one syringe and the CoA-

91

derivative in another. Because of the difference in Km values for the CoA-derivatives

among the two enzymes tested, different CoA-derivative concentrations were used to

ensure the identification of a single enzyme catalyzed event. For the wild-type reaction,

the final concentrations were: 120 μM wild-type ALAS, and 130 mM glycine. The final

concentrations of each independently examined CoA-derivative were: succinyl-CoA, 10

μM; octanoyl-CoA, 10 μM; butyryl-CoA, 20 μM; β-hydroxybutyryl-CoA, 30 μM;

glutaryl-CoA, 30 μM. For the R85K-catalyzed reaction, the final concentrations were:

120 μM R85K, and 130 mM glycine. The final concentrations of each independently

examined CoA-derivative were: succinyl-CoA, 20 μM; octanoyl-CoA, 10 μM; butyryl-

CoA, 10 μM; β-hydroxybutyryl-CoA, 10 μM; glutaryl-CoA, 20 μM. 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 tractable 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 3obs

2obs

1obs

92

The observed rate constants were determined from at least three replicate 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 protein fluorescence intensity. An OLIS RSM-1000F

rapid mixing spectrofluorimeter, equipped with 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 and ligand in the reaction chamber were 1/2 of

those in the syringes. The intrinsic protein fluorescence, as measured with 5 μM enzyme,

was evaluated in the presence of increasing concentrations of the product, ALA. The

excitation wavelength and the slit width were 280 and 5 mm, respectively. Scheme 3.1

illustrates the relationship between wavelength maximum and the dynamic process being

monitored. The emitted light was filtered 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) 01)( AeAtF tkobs

obs

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-exponential factor and A0 is the offset. The

93

observed rate constants were then plotted against ligand concentration and the data were

fitted to Equation 3.2 by nonlinear regression. The rates of dissociation (koff) and

association (kon) as well as the ligand binding constants (KD) were calculated from the

asymptotic maximal observed rate, the ordinate intercept, and the ligand concentration (x)

in Equation 3.3

(Equation 3.3) xK

xkkxf

D

offon )(

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 lowest with a value of 0.51 μM compared to 2.9 μM for

succinyl-CoA. This decreased value contributed to a catalytic efficiency value that was

2-fold higher than the reaction completed with the physiological substrate succinyl-CoA.

Glutaryl-CoA and β-hydroxybutyryl-CoA were the least catalytically efficient, due to 6-

fold increases in the Km for both substrates. Changing R85 to leucine (R85L) imparted

dramatic changes with respect to the physiological substrate succinyl-CoA. The kcat

associated with the R85L-catalyzed reaction using succinyl-CoA as the substrate

decreased greater than 11-fold, while the SCoAmK 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 variant highlight a shift toward acceptance of more

hydrophobic CoA-derivatives within the acyl-CoA-binding cleft of this enzyme. The

replacement of R85 with lysine (R85K) yielded a turnover number that was similar to

94

that of the wild-type enzyme. When the different acyl-CoA substrates were tested, this

pattern continued with the exception of the reaction with glutaryl-CoA. This CoA-

derivative reacted slower than the physiological substrate succinyl-CoA as evidenced by

a 21-fold decrease in kcat. Among the catalytic efficiencies calculated for R85K,

octanoyl-CoA yielded the highest with a value 13-fold greater than that of the same

reaction containing succinyl-CoA. 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 diminished, resulting in a 2-fold higher Km.

Conversely, the reaction of the double variant with butyryl-CoA gave a Km value of 0.54

μM, a value indistinguishable from octanoyl-CoA in the R85L variant, and 31-fold lower

than octanoyl-CoA in the double variant. The double variant-catalyzed reaction with

glutaryl-CoA, a derivative similar to succinyl-CoA but longer by one methylene carbon,

showed undetectable activity as measured under 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.

95

Table 3.1 Comparison of steady-state kinetic 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

CoAappmK , , μM 2.9 ± 0.1 12.4 ± 0.6 20.3 ± 1.0 9.6 ± 0.8

kcat/CoA

appmK , , min-1· μM-1 3.6 ± 0.2 0.53 ± 0.04 0.050 ± 0.003 0.010 ± 0.002

GlymK , mM 24.2 ± 0.4 20.0 ± 0.4 63.1 ± 2.2 98.4 ± 3.5

kcat/GlymK , min-1·mM-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

CoAappmK , , μM 0.51 ± 0.04 1.5 ± 0.04 0.55 ± 0.03 17.2 ± 0.6

kcat/CoA

appmK , , min-1· μM-1 6.8 ± 0.8 7.0 ± 0.2 3.3 ± 0.1 (5.6 ± 0.2) x 10-5

GlymK , mM 17.1 ± 0.9 25.2 ± 1.1 55.2 ± 5.0 74.2 ± 3.3

kcat/GlymK , min-1·mM-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

CoAappmK , , μM 6.1 ± 0.09 2.7 ± 0.1 9.3 ± 1.0 0.54 ± 0.03

kcat/CoA

appmK , , min-1· μM-1 1.0 ± 0.1 3.7 ± 0.4 0.21 ± 0.04 0.03 ± 0.002

GlymK , mM 29.3 ± 4.6 17.2 ± 0.7 70.3 ± 6.6 88.2 ± 3.9

kcat/GlymK , min-1·mM-1 0.26 ± 0.03 0.50 ± 0.01 0.030 ± 0.005 (6.1 ± 0.8) x 10-4

β-Hydroxybutyryl-CoA 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

CoAappmK , , μM 9.8 ± 1.0 5.5 ± 0.2 6.1 ± 0.8 74.2 ± 0.6

kcat/CoA

appmK , , min-1· μM-1 0.41 ± 0.04 0.51 ± 0.04 0.11 ± 0.02 (1.3 ± 0.4) x 10-6

GlymK , mM 22.1 ± 0.8 18.2 ± 3.2 59.1 ± 4.7 92.2 ± 6.0

kcat/GlymK , min-1·mM-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

CoAappmK , , μM 17.0 ± 1.6 7.5 ± 0.6 30.1 ± 1.1 n/d

kcat/CoA

appmK , , min-1· μM-1 0.41 ± 0.05 0.04 ± 0.003 0.07 ± 0.006 n/d

GlymK , mM 28.4 ± 0.8 21.2 ± 0.2 70.6 ± 4.2 n/d

kcat/GlymK , min-1·mM-1 0.29 ± 0.02 0.010 ± 0.002 (8.0 ± 0.7) x 10-4 n/d

n/d, not determined

96

Circular dichroism spectroscopy. To verify whether the R85K, R85L or

R85L/T430V amino acid substitutions introduced substantial changes in secondary

structure, CD spectra in the far-UV region (200-270 nm) were recorded 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 region (300-500 nm) because of anisotropic

interactions between the amino acid side chains and the chromophore in the active site

(Figure 3.3). The CD spectrum thus contains information about the asymmetric

orientation of the bound PLP cofactor in the active site, including 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 that in the presence of either succinyl- or

octanoyl-CoA there is a positive Cotton effect. However, close inspection 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 succinyl-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 differences 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 region) that are markedly different from those

of wild-type ALAS and the R85K variant. Within the R85L enzyme, succinyl-CoA

97

binding does not cause a shift in the microenvironment of the chromophore (Figure

3.3C). However, all of the remaining CoA-derivatives cause a negative Cotton effect,

decreasing the amplitude of the spectra ~5 θ. Steric hindrance and enhanced

hydrophobicity of the acyl-CoA-binding cleft 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 binding of octanoyl- and butyryl-CoA (Figure

3.3D). The positive Cotton effect observed in the spectra for the enzyme in the presence

of these two hydrophobic ligands could suggest that the exclusion of water and degree of

active site aliphaticity are crucial components for determining substrate specificity.

Overall, a comparison of the CD spectra of the wild-type and R85K variant enzymes with

those of the hydrophobic variants (R85L and R85L/T430V) show that the distinct change

in the microenvironment of the PLP pocket correlates well with the degree of

hydrophobicity inherent to the bound ligand.

98

Figure 3.2. Comparison of normalized specificity constants for murine eALAS

variants with different CoA substrates. The specificity constant (kcat/CoA

appmK , ) with

succinyl-CoA as substrate was arbitrarily defined as equal to 1.00 for each ALAS variant.

Scheme 3.1. The absorbance maxima of chemical species in the ALAS-catalyzed

reaction.

R=OP 23O

ALAS Variantwt R85K R85L R85L/T430V

Nor

mal

ized

Spe

cifi

city

Con

stan

t

0

5

10

15

606570

Succinyl-CoAOctanoyl-CoAButyryl-CoA

Hydroxybutyryl-CoAGlutaryl-CoA

99

Figure 3.3. Visible circular dichroism spectra 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 (100μM) in the

presence of 100 μM CoA derivative are indicated according to the following color

scheme: pink, succinyl-CoA; green, octanoyl-CoA; blue, β-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 degree 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 protein fluorescence were monitored. The

observed pseudo first-order decay rates of these kinetic traces were dependent on ALA

concentration, and differed among the enzymes tested (Figure 3.4). In all cases the

change in observed rate as a function of ALA concentration was hyperbolic, indicating

two binding steps. Presumably these two steps are indicative of rapid 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 between 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 μM.

The rates for the reaction of the R85K variant 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 dissociation constant was 3-

fold higher at 1470 ± 30 µM, indicating decreased affinity for ALA. Among the more

hydrophobic variants, the rates of protein quenching 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 μM

corresponding to a 3-fold greater dissociation of ALA from the R85L variant than that of

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wild-type ALAS was calculated. The double variant bound ALA less tightly, with a KD

13-fold lower than that for wild-type ALAS and an estimated value of 6710 ± 33 μM.

Figure 3.4. Reaction of wild-type ALAS, R85K, R85L and R85L/T430V (5 µM) with

ALA. The observed rate constants were calculated by fitting the decrease in intrinsic

protein fluorescence over time to Equation 3.2 for a single exponential process. (A) wild-

type 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 complexes with acyl-CoA-

derivatives. 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 rapidly 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 acyl-

CoA substrates to the enzyme-glycine complexes. The rates associated with the lifetime

of the quinonoid intermediate, measured during the enzyme catalyzed reactions, were

elucidated (Table 3.2). The absorbance change timecourses were fitted to a sequential,

three-step mechanism outlined by Equation 4.1. An initial burst of quinonoid

intermediate formation, followed by a two-step decay was characteristic of each of the

enzymes tested (Figure 3.5). Of all the acyl-CoA derivatives examined, only β-

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

103

Table 3.2. Rates of quinonoid intermediate formation and decay under single-

turnover conditions.

Parameter succinyl octanoyl butyryl β-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 ALAS- and 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-CoA, (C) succinyl-CoA, (D) glutaryl-CoA, (E) β-

hydroxybutyryl-CoA.

105

enzymes, respectively. These data support the increased catalytic efficiency observed

from the experiments performed in the steady-state. 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 the rate of quinonoid

intermediate formation with butyryl-CoA vs. glutaryl-CoA. The preference for butyryl-

CoA over glutartyl CoA suggests that in addition to the hydrogen bonding properties of

R85 and T430, the amino acids that line the hydrophobic tunnel leading to the terminal

guanidino group may play a role in substrate acceptance and orientation. Curiously, only

the R85K-glycine complex, when rapidly mixed with β-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 acyl-

CoA-binding cleft of the variant enzyme.

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Discussion

The reactions catalyzed by the highly related members of the α-oxoamine

synthase subfamily of PLP-dependent enzymes 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, spectroscopic and kinetic

methods (1, 4-6, 32). Both of the variant enzymes constructed for the arginine residue

(R85L and R85K) as well as the doubly mutated enzyme (R85L/T430V) were expressed,

overproduced, and then purified as holoenzymes, indicating that cofactor binding by the

apoprotein was not affected by the introduction of the amino acid substitutions.

However, replacement of the invariant threonine residue with valine (T430V) resulted in

a poorly expressed, unstable, and proteolytically susceptible enzyme that was never

purified to homogeneity (data not shown). All of the purifiable variants were active with

the physiological substrate succinyl-CoA. Since the threonine to valine replacement at

position 430 appears to dramatically affect protein stability, we suggest that T430 is

essential not only for optimal molecular recognition of succinyl-CoA, but also for stable

folding. R85 may be less crucial for proper 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 α-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 interactions between the two residues and the

substrate. However, considering the structural and mechanistic data for ALAS and SPT,

107

turnover likely remains orchestrated by amino acids that are proximal to the site of α-

carbon bond scission (16, 33).

Comparison of the active sites of AONS and ALAS showed that the coordination

of the acyl-CoA substrate is assisted by way of pantetheine association 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 steady-

state kinetic analysis of the variants (R85K, R85L, and R85L/T430V) with the family of

CoA-derivatives showed that the apparent Michaelis parameters ( CoAappmK , ) are dramatically

different when compared to those of wild-type ALAS. Acyl-CoA substrates of increased

hydrophobicity (e.g., octanoyl- and butyryl-CoA) demonstrated greater affinity for the

variants where the substituted amino acid was aliphatic in nature (R85L and

R85L/T430V). The 36-fold decrease in the ( CoAappmK , ) for octanoyl-CoA in the R85L

variant leads us to suggest that the exclusion 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 constant for butyryl-CoA also supports this hypothesis. The

introduction of valine at position 430 would reduce the diameter of the hydrophobic

tunnel, making steric hindrance a more significant consideration for substrate binding.

These differences identified between the variant enzymes and wild-type ALAS suggest

that reaction specificity is driven by the chemical characteristics of the CoA-derived tail

and the hydrogen-bonding potential of the invariant acyl-CoA-binding residues, a

phenomenon recognized in the acyl-CoA thioesterases of the peroxisome (34, 35).

The chemical characteristics of the acyl-CoA 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 characteristic bent conformation (36). This substrate conformation is

accomplished by two structurally conserved hydrogen bond-donating groups to the

carbonyl moiety of the substrate and through the entropically driven loss of water

coordinated by the hydrophobic amino acids 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 physiologically utilize octanoyl-CoA 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 the R85L variant is 66-

fold higher, indicating a significant change in substrate specificity. Therefore, we

hypothesize that octanoyl-CoA, devoid of a salt bridge to anchor with the guanidino

group of R85, likely bends, excludes water from the active site, and assists in enzyme

closure, a conformation postulated to be essential for turnover (16).

Interestingly, the specificity constant for the wild-type enzyme with octanoyl-

CoA is higher than the physiological substrate, succinyl-CoA, albeit at the cost of a three-

fold reduction in the specificity constant for the other substrate, glycine. Nevertheless,

this finding and the activity of the enzyme with the other CoA esters tested here lead us

to raise the question as to what extent ALAS may catalyze formation of 1,-3-

aminoketones other than ALA in vivo. This is currently unknown, but may warrant

109

further investigation. Conceivably, the potential toxicity, associated with the generation

of other aminoketones rather than ALA, could be minimized through the action of a

regulatory acyl-CoA-binding protein. In fact, studies have demonstrated that the acyl-

CoA binding protein binds long-chain acyl-CoA esters with high specificity and affinity

(with Kd values of 1–10 nM); hence by interacting with acyl-CoA utilizing enzymes, the

acyl-CoA-binding protein may provide a mechanism 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 effect 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 PLP-protein interaction, as reflected by the

different visible CD spectra (Fig. 3). Curiously, for each of the enzymes tested, the most

noticeable spectral deviations from the spectra obtained in the absence 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 variant, binding of

either octanoyl- and butyryl-CoA induced a marked positive Cotton effect (Figure 3.3).

A definite interpretation for the relationship between substrate binding and induced

Cotton effect of the PLP cofactor 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 substrate, could also explain 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 reorientation or a perturbation of the electronic system of

the chromophore). Nevertheless, the CD spectra in the UV region of all the variants

were indistinguishable from that of the wild-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 binding, are confined to the PLP-binding cleft.

Although specific contributions by single amino acid substitutions cannot be easily

disentangled using CD, the CD spectral differences in the visible range among ALAS

variants with the family of CoA-derivatives 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 ALAS-catalyzed reaction is limited by

product release, or opening of the active site loop coincident with product release (16).

Utilization of protein fluorescence to study the reverse catalytic 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 rates of product release and/or perturb the

enzyme-product equilibrium (Figure 3.4). Quenching of the ALAS intrinsic 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 CoA-derivatives of increased hydrophobicity

bind with greater affinity to the non-polar variants (R85L and R85L/T430V), it is

possible that the decreased affinity for the product, a δ-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 binding, the affinity of the aminoketones

towards ALAS would not be reduced. The Kd values would either increase or remain

unchanged; this hypothesis, however, awaits further experimentation with aminoketones

of differing hydrophobicity.

The three-dimensional structures of ALAS and AONS revealed a hydrogen

bonding network between the invariant arginine and threonine residues and the

carboxylate moieties of acyl-CoA substrates (1, 4). Accordingly, how the use of

chemically different acyl-CoA-derivatives would affect the transient kinetic parameters

of the enzymes was evaluated. Single turnover reactions with a family of CoA-

derivatives were used to determine the rates of quinonoid intermediate formation and

decay. The reaction catalyzed by wild-type ALAS with glutaryl-CoA as the substrate

exhibited a 30-fold lower rate corresponding to quinonoid intermediate formation vs. the

rate calculated with succinyl-CoA. One possible explanation for the retarded rate is that

the binding of glutaryl-CoA affects the hydrophobic acyl-CoA-binding tunnel in such a

112

way that cofactor-mediated electron transfer 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, with a 10-fold lower rate for both octanoyl-

CoA and glutaryl-CoA when compared to the physiological substrate 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 capabilities. In addition, in silico protonation of the lysine

amino group to create the ε-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 chain of arginine is

longer than the n-butylamine side chain of lysine by 1.6 Å (40). The R85K substitution

could therefore accommodate the additional sp3 hybridized carbon atom present in

glutaryl-CoA, allowing for a reduction in steric strain and/or unfavorable Van der Waals

interactions. Further, the increased hydrophobic 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 and T430 in determining acyl-CoA substrate specificity, but

also provide insight into the structure-function relationships in ALAS and the α-

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 site, 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 mutation-mediated decrease in substrate binding

energy could be accountable for the enhanced affinity measured for the hydrophobic

CoA-derivatives, instead of a direct mechanistic linkage between these residues and the

site of bond cleavage. Certainly, 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-α-carbon cleavage in the active site presumably involves coordinated movement of

the enzyme upon acyl-CoA binding. However, further experiments to prove this await

the development of tractable fluorescent probes and the elucidation of three-dimensional

crystal structures with CoA-derivatives bound.

114

Acknowledgements

This work was supported by the National Institutes of Health (grant DK63191 to GCF).

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Chapter Four

Hyperactive enzyme variants engineered by synthetically shuffling a loop motif in

murine 5-aminolevulinate synthase

Abstract

The regulatory step of the heme biosynthetic pathway in mammals is catalyzed by

the pyridoxal 5'-phosphate-dependent enzyme, 5-aminolevulinate synthase (EC 2.3.1.37).

Aminolevulinate is biosynthesized by condensing succinyl-CoA and glycine to yield

coenzyme-A and carbon dioxide. A conserved active site lid was shown to change

conformation 3.5 Å between the holoenzymic form and succinyl-CoA-bound forms of

Rhodobacter capsulatus ALAS. We employed synthetic shuffling and pre- and steady-

state kinetic analyses to determine the role of the lid motif in the ALAS-catalyzed

reaction. Functional variants containing mutations to residues 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 crucial role for the active site lid in product turnover. The

single turnover reaction data for the shuffled variants reveal that this lid has an important

role quinonoid intermediate decay and product release. Energetically favorable

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thermodynamic activation parameters for a library isolate also suggest that the entire

active site lid is involved in stabilizing the reaction coordinate. Overall, 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.

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Introduction

Aminolevulinate (ALA) is the universal building block of tetrapyrolle

biosynthesis (1). In eucaryotes and the α-subclass of purple bacteria the production of

ALA is catalyzed by 5-aminolevulinate synthase (ALAS) (EC 2.3.1.37) (2). ALAS is

classified as a fold-type I pyridoxal 5’- phosphate (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 subunit interface (4).

The reaction catalyzed by ALAS is the Claisen-like condensation of succinyl-Coenzyme-

A (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 α-

oxoamine synthase subfamily of PLP-dependent enzymes; a family which includes 7-

amino-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 co-

crystallized with the product 7-amino-8-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 carboxylate group of the ligand.

Comparison of holoenzymic ALAS and AONS, with the substrate and product bound

forms of the enzymes indicate that binding of these ligands within the active site

precipitates active site lid dynamics that may signify a change in conformation. The

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active site lid of ALAS contains 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 apex 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 determinant of substrate specificity (Lendrihas et.

al, in press) (7). Two β-sheets flank the position of the lid (β12 and β13) in the C-

terminal domain of the enzyme, where further interactions between the carboxy terminal

portion of the lid occur between two α-helices on the adjacent subunit (α2 and α3).

While evidence is available for the role of discrete active site residues in ALAS-

function, 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 α3 and α4

was proposed to be involved 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 variants with turnover numbers greater than

that of wild-type ALAS. Further, this loop is implicated in succinyl-CoA binding, and

based on the position of these residues deep within the three 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 acyl-

CoA substrate and amino acid donor was first identified in AONS (8). According to the

X-ray crystal structures the β-sheet of the C-terminal domain undergoes a conformational

transition. This conformational change includes the 5.5 Å displacement of the active site

lid. From the ALAS crystal structure with succinyl-CoA bound the change in position of

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the outermost top of the lid moves inward by 3.5 Å 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 functional amino acid mutations and to evaluate

the contribution of lid residues to catalysis. The reported analysis of the catalytic 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 during catalysis, thereby facilitating enzyme-

mediated chemistry. Once chemistry occurs, the dynamics of the active site lid determine

the rate of product release.

122

Figure 4.1. The position of the active site loop in the R. capsulatus ALAS crystal

structure. In (A), the overlap of one monomer of holoenzymic (magenta) and succinyl-

CoA-bound (green) ALAS from R. capsulatus. In (B) the active site 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 sequence of the lid is +NH3—YVQAINYPTVPRGEELLR—COO-

A B

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Materials

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-Aldrich 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 reagents 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-amino 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 overlapping 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 amplified with another round of PCR having

as primers oligonucleotides which annealed against the 5’ and 3’ ends of the generated

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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 refers to variants encoded within the library. Bold amino acids indicate amino acids that are found in ALASs from different species. Codon indicates the nucleotide codon used to obtain the indicated mixture of amino acid residues. Subscripts following amino acid designations are the number of different codons for that amino acid. Nucleotide degeneracies are 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 restriction enzyme sites used in subsequent

subcloning (i.e., Xba I and Bam HI). To minimize the wild-type ALAS “background”

during the screening of the library for functional ALAS variants, the synthetic shuffling

library was subcloned into a mock ALAS expression vector. The mock vector contained

the wild-type ALAS-encoding sequence from 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 reaction and PCR, and mock vector used in this

study are described in supplemental experimental procedures. The ligation reactions and

DNA digestions with restriction endonucleases were according to standard protocols in

molecular biology (18).

Screening of the ALAS synthetic shuffled 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). HemA- cells can only

survive if harboring a functional ALAS or when ALA (or hemin) 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 allow 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 induce 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 without ALA. The colonies that grew in the absence of ALA

were identified as functional variants. Functional variants were screened for porphyrin

overproduction by fluorescence microscopy.

Single colony isolation and sequencing of plasmid 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

hemA-

bacterium

LB agar-- ALA

PCR

Transformation of expression plasmids

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 excitation filter set for excitation at 400–440 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 porphyrin accumulated in the functional ALAS

clones was compared to that of bacterial cells harboring wild-type ALAS and grown

under the same experimental conditions (Figure 4.3). Bacterial cells harboring ALAS

variants, which accumulated greater porphyrin levels than bacterial cells with wild-type

ALAS, were grown in LB/ampicillin medium in 96-well plates, and the glycerol stocks

generated from overnight cultures were submitted to the ICBR Genomics Core at the

University of Florida for DNA sequencing of the corresponding plasmids.

Overexpression, purification and spectroscopic 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 containing: 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 fluorescence 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-state kinetic characterization of ALAS active site loop

variants. ALAS steady-state activity of the ALAS active site loop variants was

determined at 20°C using a continuous spectrophotometric assay as described previously

for wild-type ALAS (22). The ALAS activity data, acquired using a Shimadzu UV 2100

dual-beam spectrophotometer, were plotted vs. substrate concentration in which one of

the substrate concentrations varied, while the second was kept constant. The steady-state

kinetic parameters were determined by fitting the data to the Michaelis-Menten equation

using non-linear regression analysis software as reported in (10). The ALAS steady-state

kinetic parameters of wild-type ALAS and the SS2 variant activity assays were also

determined at 15, 25, 30 and 35 ºC.

Rapid scanning stopped-flow measurements were performed using a model RSM-

100 stopped-flow spectrophotometer (OLIS, Inc.). This instrument has a dead-time of

approximately 2-ms and an observation chamber 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 appropriate 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-steady-state kinetic reactions of the

variant enzymes were examined under single turnover conditions, using final

concentrations of 60 μM enzyme, 120 mM glycine and 10 μM succinyl-CoA in 100 mM

HEPES, pH 7.5 and 10% (v/v) glycerol as previously described (23). Single turnover

130

]Ligand[

]Ligand[

D

max

K

YY

data 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 determined 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 constants 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 (pH 7.5) and 10% glycerol at 20oC and by titrating the

SS2 variant (60 μM) 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

3obs

2obs

1obs

2obs

1obs

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 formation of an external aldimine with the PLP-

cofactor. 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 μM) 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 parameters 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( ATR

Ek a

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, ΔH, ΔG and ΔS, as

previously described (23).

Data analysis for the definition of the minimal 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 reaction 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 wild-type ALAS were maintained constant (21).

Results

Construction of the synthetically shuffled 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 evaluate 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 utilizing mixed base oligonucleotides that

encode specific changes to the cDNA. Mixed base oligonucleotide primers encoding

amino acids that have a precedent in nature were incorporated into the primary sequence

of wild-type ALAS, from Y422-R439 (Table 4.1). A multiple sequence alignment

quantifying the naturally occurring amino acid diversity present at specific positions in

the lid was performed (Figure 4.4). These ALAS 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 consideration employed to reduce the possibility of recovering

inactive variants. Screening of the synthetically 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 depleted metabolite required

for colony formation on agar plates lacking ALA. Next, to elucidate which variants are

catalyzing the production of ALA at a rate faster than wild-type ALAS, the appearance of

fluorescence associated with excess porphyrin biosynthesis was used (Figure 4.3).

134

Figure 4.4. Multiple alignment of the amino acid sequences of the ALAS loop region. Active Site Loop Y422...............R439 1 ALAS_DELLEU 486 LLLSKHGIYVQAINY PTVPRG-EELLRLAP SPHH-SPQMMEDFVE KLLAAWTEVGLPLQD -VSIAACNFCRRPVH FELMSEWERSYFGNM 573 2 ALAS_DLLLEU 486 LLLSKHGIYVQAINY PTVPRG-EELLRLAP SPHH-SPQMMEDFVE KLLAAWTEVGLPLQD -VSIAACNFCRRPVH FELMSEWERSYFGNM 573 3 ALAS_HOMSAP 491 LLLSKHGIYVQAINY PTVPRG-EELLRLAP SPHH-SPQMMEDFVE KLLLAWTAVGLPLQD -VSVAACNFCRRPVH FELMSEWERSYFGNM 578 4 ALAS_RATNOR 491 LLLAKHSIYVQAINY PTVPRG-EELLRLAP SPHH-SPQMMENFVE KLLLAWTEVGLPLQD -VSVAACNFCRRPVH FELMSEWERSYFGNM 578 5 ALAS_RATRAT 491 LLLAKHSIYVQAINY PTVPRG-EELLRLAP SPHH-SPQMMENFVE KLLLAWTEVGLPLQD -VSVAACNFCRRPVH FELMSEWERSYFGNM 578 6 ALAS_MUSMUS 491 LLLSKHSIYVQAINY PTVPRG-EELLRLAP SPHH-SPQMMENFVE KLLLAWTEVGLPLQD -VSVAACNFCHRPVH FELMSEWERSYFGNM 578 7 ALAS_DANRER 486 ILLEKHNIYVQAINY PTVPRG-EELLRLAP SPFH-NPIMMNYFAE KLLDVWQEVGLPLNG -PAQASCTFCDRPLH FDLMSEWEKSYFGNM 573 8 ALAS_DANROS 486 ILLEKHNIYVQAINY PTVPRG-EELLRLAP SPFH-NPIMMNYFAE KLLDVWQEVGLPLNG -PAQASCTFCDRPLH FDLMSEWEKSYFGNM 573 9 ALAS_OPSTAU 486 SLLEKHNIYVQAINY PTVPRG-QELLRLAP SPHH-HPAMMEYFVD KLVEVWQEAGLLLNG -PATVSCTFCDRPLH FDLMSEWEKSYFGNM 573 10 ALAS_HOMSPN 545 ELMSRHNIYVQAINY PTVPRG-EELLRIAP TPHH-TPQMMNYFLE NLLVTWKQVGLELKP -HSSAECNFCRRPLH FEVMSEREKSYFSGL 632 11 ALAS_DELDEL 545 ELMSRHNIYVQAINY PTVRRG-EELLRIAP TPHH-TPQMMNYFVE NLLATWKRVGLELKP -HSSAECNFCRRPLH FEVMSEREKSYFFGM 632 12 ALAS_MOUDOM 546 ELMTRHNIYVQAINY PTVPRG-EELLRIAP TPHH-TPQMMNFFVE KLLVTWKRVGLELKP -HSSAECNFCRRPLH FEVMSEREKAYFSGM 633 13 ALAS_GALVAR 540 KLMSQHSIYVQAINY PTVPRG-EELLRIAP TPHH-TPQMMSYFLE KLLATWKDVGLELKP -HSSAECNFCRRPLH FEVMSERERSYFSGM 627 14 ALAS_XENLAE 308 KLMRDYSIYVQAINY PTVPRG-EELLRIAP TPHH-NPQ------- --------------- --------------- --------------- 344 15 ALAS_OPSBET 532 LMMSHHNIYVQAINY PTVARG-DELLRIAP TPHH-TPEMMKYFVD RLVQTWKEVGLELKP -HSSAECTFCQQPLH FEVMNEREKSYFSGL 619 16 ALAS_OPSPAR 532 LMMSHHNIYVQAINY PTVARG-DELLRIAP TPHH-TPEMMKYFVD RLVQTWKEVGLELKP -HSSAECTFCQQPLH FEVMNEREKSYFSGL 619 17 ALAS_DANDAN 518 IMMSRYNIYVQAINY PTVARG-EELLRIAP TPHH-TPQMMKYFVD KLTQTWTEVGLPLKP -HSSAECNFCRQPLH FEIMSEREKSYFSGL 605 18 ALAS_MYXGLU 565 ELMSRHNIYVQAINY PTVPRG-EEMLRVVV TPHH-TPQMMQYFVE HLTNSWKDIGLNLRP -HASAECNYCKMPIH FELMSEHDQVYFDGM 652 19 ALAS_STRDRO 512 SLLEEHNIYVQAINS PTVPSG-EEKLRIAP SPXH-TPDMMDRFVA SLSEVWAKSGLRFNT PICPRECEFCKNPEK FEELSSRERSFAEES 578 20 ALAS_DROMEL 463 VLIEQFGHYLQSINY PTVARG-QEKLRLAP TPFH-TFEMMNALVT DLKKVWEMVDLSTNV PLSPNACMFCNSESC WHQDTSPDLECGIPN 529

135

21 ALAS_LIMPOL 494 ELISMHGHYVQAINY PTVPRG-EEKLRIAP TPFH-TRPMMEAFVR DLVSVWRGLKLPLRD GICEKKCEFCEKPLY FEHLESRVL------ 560 22 ALAS_SEPOFF 540 DLLIKHNIYVQAINY PTVARG-EEKLRVAA TPHH-TKEMMDHFVD CVVKVWLEHGLTLNP -DSTRPAEFNVKFKK FSI------------ 603 23 ALAS_GLYDIB 516 ELMEEHGIYVQPINY PTVPRG-QELLRVAP TPHH-TKEMMDSFVN ATLSVFLNNNIELKS -TCGINCLYCHQPMK CEAFTNRER------ 586 24 ALAS_GALGAL 420 ALLEEHGLYVQAINH PTVPRGQELLLRIAP TPHH-SPPMLENLAD KLSECWGAVGLPRED -PPGPSCSSCHRPLH LSLLSPLER------ 508 25 ALAS_SINMEL 346 LLLDNFGIYVQPINY PTVPKK-TERLRITP TPMH-SDADIDHLVS ALHSLWSRCALARAV A-------------- --------------- 405 26 ALAS_SMRMEL 275 LLLDNFGIYVQPINY PTVPKK-TERLRITP TPMH-SDADIDHLVS ALHSLWSRCALARAV A-------------- --------------- 334 27 ALAS_AGRTUM 364 ILLDNHGVYVQPINY PTVPRK-TERLRITP TPLH-TDADIEQLVG ALHQLWSHCALARAV A-------------- --------------- 423 28 ALAS_AGRRAD 346 ILLDSHGVYVQPINY PTVPRK-TERLRITP TPLH-SDADIEHLVG ALHQLWSHCALARAV A-------------- --------------- 405 29 ALAS_RHIRAD 276 ILLDSHGVYVQPINY PTVPRK-TERLRITP TPLH-SDADIEHLVG ALHQLWSHCALARAV A-------------- --------------- 335 30 ALAS_RHOPSE 344 ELINRYGIYVQPINY PTVPRG-TERLRITP SPQH-TDADIEHLVQ ALSEIWARVGLAKAA --------------- --------------- 403 31 ALAS_RHOPSE 346 ALLARHAIYVQPINY PTVARG-QERFRLTP TPFH-TTSHMEALVE ALLAVGRDLGWAMSR RAA------------ --------------- 407 32 ALAS_EUGGRA 343 LLLKRFQIYVQPINY PTVDVG-TERLRITV SPVH-TNEHMATLIT ALLQVWEELGLPLRP PVFDTEGYPEVEAAE WLPNSAMWR------ 409 33 ALAS_BRAJAP 348 LLLEEHGIYIQPINY PTVAKG-SERLRITP SPYH-DDGLIDQLAE ALLQVWDRLGLPLKQ KSLAAE--------- --------------- 409 34 ALAS_BRAJAP 275 LLLEEHGIYIQPINY PTVAKG-SERLRITP SPYH-DDGLIDQLAE ALLQVWDRLGLPLKQ KSLAAE--------- --------------- 339 35 ALAS_BRUMEL 345 RLLEVHGIYIQPINY PTVPRG-TERLRITP SPLH-DDKLIDGLKD ALLEVWNELGIPFAE PSAPQAANSDRIIPL MVSKAGG-------- 425 36 ALAS_ZYMMOB 287 ILLNEYGAYVQPINF PTVPRG-TERLRFTP GPTH-NEAMLRELTD SLVAIWHRLDMRFAA --------------- --------------- 345 37 ALAS_RHOCAP 348 MLLSDYGVYVQPINF PTVPRG-TERLRFTP SPVH-DLKQIDGLVH AMDLLWARCA----- --------------- --------------- 401 38 ALAS_RHOCAP 278 MLLSDYGVYVQPINF PTVPRG-TERLRFTP SPVH-DLKQIDGLVH AMDLLWARCA----- --------------- --------------- 331 39 ALAS_PARDEN 292 MLLADFSIYVQPINF PTVPRG-TERLRFTA SPVH-DPKQIDHLVK AMDSLWSQCKLNRST SAA------------ --------------- 339 40 ALAS_PARZEA 360 MLLADFSIYVQPINF PTVPRG-TERLRFTA SPVH-DPKQIDHLVK AMDSLWSQCKLNRST SAA------------ --------------- 409 41 ALAS_RHOPSE 364 MLLIHFGIYVQPINF PTVPRG-TERLRFTP SPVH-DSGMIDHLVK AMDVLWQHCALNRAE VVA------------ --------------- 407 42 ALAS_EMENID 510 KLLEEHGIYVQAINY PTVPRG-EERLRITP TPGH-TQELRDHLVE AVNTVWNDLGIKRAS DWKAMGGFVGVGVEA AELENQPIWT----- 598

Figure 4.4. Continued

136

43 ALAS_EMENID 440 KLLEEHGIYVQAINY PTVPRG-EERLRITP TPGH-TQELRDHLVE AVNTVWNDLGIKRAS DWKAMGGFVGVGVEA AELENQPIWTDAQLN 528 44 ALAS_ASPNID 440 KLLEEHGIYVQAINY PTVPRG-EERLRITP TPGH-TQELRDHLVE AVNTVWNDLGIKRAS DWKAMGGFVGVGVEA AELENQPIWTDAQLN 528 45 ALAS_ASPORI 508 KLLEEHGIYVQAINY PTVPRG-EERLRITP TPGH-IKEHRDHLVQ AVQTVWNELGIKRTS DWEAQGGFVGVGVDG AEAENQPIWNDVQLG 596 46 ALAS_NEUCRA 334 KLLNDHQIYVQSINY PTVPVG-QERLRITP TPGH-TKQFRDHLVA ALDSIWTELGIKRTS DWAAEGGFIGVGEAE A-EPVAPLWTDEQLG 421 47 ALAS_GIBFUJ 393 MLLNDYGIYVQAINY PTVPVG-QERLRVTP TPGH-IKEYRDQLVE AIDEIWTRLDIKRTS DWAAEGGFIGVGEQD --NVQEPLWTDKQLN 479 48 ALAS_YARLIP 404 LLLTKHQIYVQAINF PTVPIG-QERLRVTP TPGH-HEGLCDELVA ALEDVWQELDLKRVE DWTAEGGLCGVGEGV --E-VEPLWSEEQLS 489 49 ALAS_CANALB 451 LLLNKHDIYVQAINF PTVPIG-EERLRITP TPGH-GPELSKQLVE AVDSVFTELNLNRIN DWKKLGGLVGVGVEG -AAKVEHIWTEEQLA 538 50 ALAS_CANALB 381 LLLNKHDIYVQAINF PTVPIG-EERLRITP TPGH-GPELSKQLVE AVDSVFTELNLNRIN DWKKLGGLVGVGVEG -AAKVEHIWTEEQLA 468 51 ALAS_DEBHAN 392 LLLDKYNIYVQAINF PTVPIG-QERLRITP TPGH-GPELSNQLIG ALDSVFNELSLSRIG DWEGKGGLCGVGEPD -IEPIEHIWTSEQLA 479 52 ALAS_SACCER 435 ILINKHQIYVQAINF PTVARG-TERLRITP TPGH-TNDLSDILIN AVDDVFNELQLPRVR DWESQGGLLGVGESG -FVEESNLWTSSQLS 522 53 ALAS_SACCAS 365 ILINKHQIYVQAINF PTVARG-TERLRITP TPGH-TNDLSDILIN AVDDVFNELQLPRVR DWESQGGLLGVGESG -FVEESNLWTSSQLS 452 54 ALAS_CANGLA 347 ILMEKHRIYVQAINF PTVSRG-TERLRITP TPGH-TNDLSDILIA AVDDVFNELQLPRIR DWEMQGGLLGVGDKN -FVPEPNLWTEEQLS 434 55 ALAS_KLULAC 387 ILMDKHRIYVQAINF PTVARG-TERLRITP TPGH-TNDLSDILMD ALEDVWSTLQLPRVR DWEAQGGLLGVGDPN -HVPQPNLWTKDQLT 474 56 ALAS_EREGOS 373 ILMEKHRIYVQAINF PTVPRG-TERLRITP TPGH-TNDLSDVLLD AMDDVWKTLQLPRVS DWAAHGGLLGVGEPD -YVPEANLWTEEQMS 460 57 ALAS_AGABIS 460 KLLSEHDIYVQAINY PTVARG-EERLRITV TPRH-TMEQMEGLIR SLNQVFEELNINRLS DWKLAGGRAGVGIPG AADDVQPIWTDEQIG 548 58 ALAS_AGADIV 390 KLLSEHDIYVQAINY PTVARG-EERLRITV TPRH-TMEQMEGLIR SLNQVFEELNINRLS DWKLAGGRAGVGIPG AADDVQPIWTDEQIG 478 59 ALAS_SCHPOM 475 SLLHDHNIYVQSINF PTVSVG-TERLRITP TPAHNTEHYVQSLTN AMNDVWSKFNINRID GWEKRGIDVGRLCKF PVLPFTTTH------ 558 60 ALAS_SCHMIK 405 SLLHDHNIYVQSINF PTVSVG-TERLRITP TPAHNTEHYVQSLTN AMNDVWSKFNINRID GWEKRGIDVGRLCKF PVLPFTTTH------ 488 61 ALAS_RICPRO 342 MLLNEYGIYVQHINF PTVPRG-TERLRIIP TPAH-TDKMINDLST ALVHIFDELDIELSS AKELNKEVRLHLIA- --------------- 414 62 ALAS_RICTYP 342 MLLNEYGIYVQHINF PTVPRG-TERLRIIP TPAH-TDKMINDLST ALVHIFAELDIELSS TKELNKEVRLHLIA- --------------- 414 63 ALAS_RICPRO 272 MLLNEYGIYVQHINF PTVPRG-TERLRIIP TPAH-TDKMINDLST ALVHIFDELDIELSS AKELNKEVRLHLIA- --------------- 344 64 ALAS_RICCON 354 MLLNEYGIYVQHINF PTVPRG-TERLRIIP TPAH-TDKMINDLSV ALVQIFAELDIELSS AKELNEEVRLNLIA- --------------- 426 65 ALAS_CHRVIO 354 RLLEEFDIYVQPINY PSVPRG-GERFRLTV GPRR-SHEEIQRFVA ALKHCLA-------- --------------- ---------------

Figure 4.4. Continued

137

Figure 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 synthetic shuffling are high-lighted in cyan. The amino acid numbering in red refers to that of murine erythroid 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 isolate ALAS variants with turnover numbers

greater than that of wild-type ALAS. In panels (A1, B1 and C1) of Figure 4.3 cells are

visualized with a DAPI filter, which was used a means of assaying for single colony

isolates. The magnification for each image was identical, implying that colony size

differed between those cells harboring a hyperactive 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 cells harboring the inactive ALAS

(K313A) and those complemented with wild-type ALAS show clearly the lack of

porphyrin-derived red fluorescence 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 colonies on expression agar proved a reliable

method for obtaining hyperactive isolates.

Qualitative analysis of the isolated active site lid variants. The mutations present

among the active site lid variants 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 substitutions observed among the 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 non-

covalent 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 P T V P R G E E L L RSingle variants

A8 T D8 Q G7 H

Quadruple variant

F1 K K Q Q Penta variant

F10 I Q N T N Hexa variants

A4 I P C R K N F3 G H H N K K H1 N N I E K K

Hepta variant SS2 L R E I N Q K

140

interactions may be perturbed in the variants. Replacements at residues toward the

carboxy terminus of the lid, (P432-R439) favored conversion to basic amino acids, likely

positively charged at physiological pH, while residues that comprised the portion of the

lid proximal to the amino terminus (Y422-V431) showed less restriction toward which

amino acid was permissible. Several positions showed a propensity for a particular

substitution as demonstrated by the frequency of the observed mutation. At position 423,

valine was found substituted three times for leucine, indicating 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 glutamine and lysine, L437 was substituted for

lysine three times and glutamine twice. This indicates that the presence of positively

charged residues in this area of the lid may facilitate lid dynamics.

Steady-state kinetic analysis 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 enzymes 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 hextuply mutated F1, F10 and H1 variants were

also markedly higher compared to wild-type ALAS. The remaining variants, including

the single point variants, also showed enhanced 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 increases 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 reactions of hyperactive ALAS enzymes1

Steady-state parameters Pre-steady-state

parameters

Enzyme kcat s-1

GlymK

mM

SCoAmK

μM

kcat/GlymK

mM-1 s-1

kcat/SCoAmK

μM-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 quinonoid intermediate decay; 4Rate for second step of quinonoid intermediate decay.

142

among the active site lid variants. All of the variants showed at least a 3-fold reduction in

the SCoAmK . Accordingly, due to the enhanced turnover evident among the multiply

mutated variants, the catalytic efficiency with 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 succinyl-

CoA supports a hypothesis whereby the binding of this substrate switches a pre-existing

protein conformational equilibrium 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 lid makes toward

the reaction catalyzed by ALAS, the transient kinetic parameters of the library variants

were elucidated. The formation and decay rates of a key step in the chemical mechanism

were obtained (Table 4.3). Under single turnover reaction conditions, the lifetime of a

transient quinonoid intermediate was observed 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, equations 1 and 2, respectively. An initial

burst of quinonoid intermediate formation, followed by a two-step rate of decay was

characteristic to 6 of the enzymes tested (Figure 4.5A-F). Compared to wild-type ALAS

(0.8 s-1), the A4 and F3 multiply mutated lid variants, 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 quinonoid 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 were calculated under

single turnover conditions (60μM

enzyme, 10μM 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 wild-

type enzyme.

145

Accordingly, the second step of quinonoid intermediate decay, a step hypothesized to

include the dissociation of product and opening of the putative active site lid was

markedly increased in these variants, with values of 0.07 s-1 and .017 s-1, rates 7- and 17-

fold 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 steady-

state. The singly mutated variants A8, D8 and G7 also formed the quinonoid

intermediate at least 4-fold faster when 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 did not exceed that displayed by wild-type ALAS.

This suggests that the coordinated action of more 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 proceeding by a mechanism similar to wild-type

ALAS, these enzymes appear to condense the biphasic 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, since 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 wild-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 the 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 were fit to a standard hyperbola and the KD for

glycine was found to be 4.12 ± 0.57. This value 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 constant 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 and the SS2 variant may be attributable to

increased active site lid flexibility, a circumstance supported by the hyperactivity

measured for SS2 in the steady- and 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

288–308 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 parameters. 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). The 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 wild-

type 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 calculated from the linear relationship between the

kcat values for both enzymes and temperature. 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 proceeds 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-state 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 parameters of wild-type ALAS and the SS2 variant.

Enzyme Slope Ea (kcal/mol)

H (kcal/mol)

G (kcal/mol)

S (kcal/mol·K)

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 aldimine with lysine 313 to an external aldimine with glycine

is the first step. Succinyl-CoA binds second which is followed by quinonoid formation,

protonation of the quinonoid to yield an aldimine bound molecule 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; Succinyl-CoA: EGSCoA;

ALAS-glycine-Succinyl-CoA complex: EQ; enzyme bound to protonated quinonoid:

EALA; enzyme-ALA complex.

KD = 4.28 mM

E+G EG+SCoA EGSCoA EALA E+ALA KD = 1.82 mM

1.7 s-116 s-1 2.2 μM-1· s-1

1.5 s-10.22 s-111 s-1

EQ

150

Discussion

Comparison of the X-ray crystal structures of holoenzymic ALAS and AONS

with the succinyl-CoA- and AON-bound structures, revealed the presence of a

conformationally mobile active site lid. The outermost part of the active site lid of ALAS

undergoes a 3.5 Å change in conformation in the presence of succinyl-CoA (Figure 4.1).

To address the role of this dynamic structure in the enzyme-catalyzed reaction we

employed synthetic shuffling to alter the amino acid composition of the active site lid in

mALAS2 (Y422-R439). Our results suggest that amino acid substitutions within the

active site lid lead to altered lid dynamics, a circumstance affecting enzymatic turnover,

and one that ultimately liberates the enzyme from a conformationally limited rate

determining step.

The approach used to evaluate the active site lid of ALAS utilized a mutagenic

technique called synthetic shuffling (14). Non-conserved amino acids that comprise the

lid vis á vis the multiple sequence alignment, tolerated sequence substitutions differently

based on their position in the motif (Figure 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 mutational frequency evident between the two

halves of the lid suggest that the role of the carboxy terminal half of the lid may be less

structural in nature, as demonstrated by ability of drastically mutated variants such as SS2

and F1 to not only retain activity, but turnover product at rates 10- and 15-fold faster

compared to wild-type ALAS. All of the singly mutated variants had replacements in

151

both halves of the lid (A8, D8 and G7). While the isolation 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 residues 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 α-helices (α2 and α3) 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 residues 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, coupled with the mutational data on residues N150

and I151 agrees with three previous models where specific interactions that drive ligand-

induced closure and catalysis are proposed 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 efficiency for both substrates was

enhanced. These characteristics indicate that 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 release, rather than chemistry, is rate limiting. Therefore the

enhanced turnover observed among the active site lid variants potentially validates our

ALAS catalytic mechanism in which the enzyme 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 the carboxy terminal portion 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 previous study investigating single point

mutations to R433 (R433L and R433K), found that the kcat values increased 20% and

100% for the respective substitutions (11). Those data coupled with the findings of this

study suggest that position 433 may significantly contribute to lid mobility and the

dissociation of ALA from the enzyme. However, the variants that had the largest

increases in turnover number and catalytic efficiency were those 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 α2 and α3, with concomitant disruption of the active site

lid interaction with the aldimine-bound product, likely resulting in acceleration of the

conformationally limited release of ALA. Conversely, the single point mutant identified

in D8 (L437Q) showed only 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 identified 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 barrier in the single-

153

turnover reaction. That is, in a process that 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 turnover 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 function 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 can be strongly polarized, and ultimately

lysed. Additionally, lid opening enhances product dissociation (compared to wild-type

ALAS, the SS2 variant showed a 4-fold increase in the ALADK ) by potentially: uncovering

ALA from within a catalytic vacuole, restoring electron density to the site of α-carbon

bond scission, and minimizing contact between product and enzyme. Further, since the

second rate of quinonoid intermediate 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 turnover by equalizing the energy of quinonoid

intermediate decay in the enzyme-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 (Figure 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 complex with succinyl-CoA is ~20-fold faster

than the corresponding rate in wild-type ALAS, suggesting that a step after catalysis is

only partially rate determining for this variant 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). Together 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 catalysis. In the three variants that appear

condensed with respect to the rate of quinonoid intermediate decay, Qd is greater than

that of wild-type ALAS and all the active site lid variants tested (Table 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 single turnover data corresponding to SS2,

quinonoid intermediate decay has condensed 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 (Figure 4.8). Thermodynamic data further support this

spectroscopic observation (Figure 4.7 and Table 4.4). Comparison of the wild-type

ALAS- and SS2-catalyzed reactions 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 absorbed from the environment, a circumstance

whereby the ALAS-catalyzed reaction compensates by requiring an additional step. This

interpretation is supported by the refined mechanism of the ALAS-catalyzed reaction by

Hunter et. al., wherein the reverse rate of enzyme-ALA1 conversion to enzyme-

155

quinonoid is greater than the forward rate of enzyme-ALA1 conversion to enzyme-ALA2

(28). Overall the transient kinetic data suggest that active site lid flexibility and enzyme

activity are tightly 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.

In conclusion, a mutational analysis of the active site lid of mALAS2, identified

by structural investigation, reveals a role of the lid in determining the rate limiting step of

the enzyme-catalyzed reaction. Variants isolated from the library contained mutations to

the non-conserved residues that comprise the active 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 determinant of both substrate binding and product

turnover. Additionally, in a subset of variants, 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.

156

Acknowledgements

This work was supported by the National Institutes of Health (grant DK63191 to GCF).

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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 transient kinetic characterization of the ALAS

pathway, in which the enzyme undergoes a transition 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 active site shows that a conserved serine residue (S189 in R.

capsulatus ALAS and S254 in meALAS) changes the orientation of its hydrogen-

bonding 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 substitution elicited a 2-fold increase in enzyme

activity, while simultaneously increasing the Km for succinyl-CoA 25-fold. These

experimental findings coupled with the structural data suggest that this residue is an

important determinant in conformer equilibrium by promoting energetically favorable

interactions between the chromophore and succinyl-CoA, and by stabilizing a closed

Michaelis-complex configuration.

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 alanine 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 enzyme-

catalyzed reaction. The means by which the enzyme manages to increase activity despite

a reduction in affinity for the more complex of the two substrates may include a

mechanism whereby the equilibrium is predominantly shifted toward the closed

conformational state.

The S254A mutation has notable effects on the cofactor microenvironment as

determined by CD spectroscopy. CD spectroscopic evaluations of the conformational

effects of the S254A and S254T mutations illustrate the differences between the two

amino acid substitutions. Succinyl-CoA binding to the wild-type and S254T variant

induced changes in the CD spectra associated with the microenvironment of the

chromophore, while CD spectra corresponding to the S254A mutant were unchanged

under similar conditions. This dissimilarity between wild-type ALAS and S254A may be

the result of a partial conversion 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 sites upon succinyl-CoA binding (1). In the crystal structures

these events are coincident with the closed conformation, from which it might be

concluded that the S254A variant retains the 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 ALAS- and variant-

catalyzed 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 4-fold 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 structures 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 catalysis, 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 mechanism in which substrate recognition is

coupled to conformational equilibria.

ALAS is a member of the -oxoamine synthase subfamily of pyridoxal 5'-

phosphate (PLP)-dependent enzymes and shares a high degree of structural similarity and

reactivity with the other members of the family (6). Despite the remarkable structural

and mechanistic similarities in this important 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-binding residues in murine erythroid ALAS

(R85 and T430) in relation to acyl-CoA binding and substrate discrimination were

examined using site-directed mutagenesis and a series of CoA-derivatives.

The acyl-CoA substrate binds to the enzyme 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 steady-

state kinetic analysis of the variants (R85K, R85L, and R85L/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., octanoyl- and 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 binding for octanoyl-CoA in the R85L variant suggests that

the exclusion of water from the acyl-CoA-binding cleft is an important feature of

substrate binding. It is likely that reaction specificity is driven by the chemical

characteristics of the CoA-derived tail and the hydrogen-bonding potential of the

invariant acyl-CoA-binding residues, a circumstance 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 reaction. Single turnover reactions with a family

of CoA-derivatives were used to determine the rates of quinonoid intermediate formation

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and decay for the R85K variant. The R85K variant-catalyzed reaction was decelerated in

the first step of quinonoid intermediate decay, with a 10-fold lower rate for both

octanoyl-CoA and glutaryl-CoA when compared to the physiological substrate 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 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 arginine is longer than the n-butylamine side chain of

lysine by 1.6 Å (9). The R85K substitution could therefore accommodate the additional

sp3 hybridized carbon atom present in glutaryl-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 substrate succinyl-CoA.

In all, the experimental data support multifunctional roles for these amino acids (R85 and

T430) in regulating substrate specificity, and linking the bifurcate 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 Å between the holoenzymic forms and acyl-CoA-bound forms of the

two enzymes, respectively (1, 10). Comparison of holoenzymic ALAS and AONS with

the substrate- and product-bound forms of the 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

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“lid”) in the ALAS-catalyzed reaction, we used genetic manipulation 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 conserved, 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 amino-terminus (31 to 12, respectively). This

inequality suggests that greater plasticity is associated with the C-terminal part of the

loop.

Each of the isolated variants was shown to have increased turnover numbers and

enhanced catalytic efficiency with both substrates. These characteristics, identified

among active site lid variants, indicate that the reaction catalyzed by ALAS may be

limited by the amino acid composition of the lid. Further, conformational changes

centered in active site loops have been reported to have important mechanistic roles in

other enzymes (12, 13). Accordingly, mutations that affect the mobility of a

conformationally mobile enzyme structure may 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 potentially validates our

ALAS catalytic mechanism in which the enzyme conformation switches between closed

and open to stabilize reaction intermediates and release product, respectively, a

mechanism recently proposed for another PLP-dependent enzyme, diaminopimelate

decarboxylase (14).

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The pre-steady-state behaviors 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 intermediate decay were condensed into a single

step. For the SS2 variant-catalyzed reaction, quinonoid intermediate formation is ~20-

fold faster than the corresponding rate in wild-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 for ALA (Table?), the data suggest that a

conformational change leading to product release no longer is rate-limiting for the SS2-

catalyzed 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 positive differential activation entropy 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 condensed 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 decrease of net energy observed for the SS2

variant-catalyzed reaction, supports the loss of this kinetic step. Conversely, the excess

energy required by the wild-type enzyme would likely have to be absorbed from the

environment, a circumstance whereby the ALAS-catalyzed reaction would compensate

by requiring an additional step (3). This interpretation is supported by the refined

mechanism of the ALAS-catalyzed reaction by Hunter et. al., wherein the reverse rate of

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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 tightly 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 library isolates show that these mutations

confer hyperactivity. Through these investigations, hypotheses pertaining to the reaction

mechanism of ALAS have implications in the pathways of disease involving both iron

metabolism and porphyrin biogenesis. Data presented here and conclusions set forth

regarding limits to ALA turnover and individual steps associated with substrate binding

and rates of reaction, specifically address facets of XLSA and the recently discovered

erythroid ALAS-related porphyria, X-linked dominant protoporphyria (19). Many

mutations associated with diminished ALAS activity in vivo are located in the PLP-

binding cleft (1, 19). Significantly, the conformationally 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 pathology that is associated with X-linked

dominant protoporphyria (18).

The knowledge related to ALAS from the studies presented here contribute to the

goal of therapeutic intervention of porphyrin-accumulative diseases, like cancer.

Further, biochemical developments in the field of biotechnology will also be enhanced

through this research by addressing the molecular requirements of treatments like

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photodynamic therapy. In conclusion, these data are likely to provide insight into the

rate-determining step of enzymes limited by a product release, or a conformational

change leading to product release.

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(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.

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About the Author

Thomas Lendrihas received two Bachelor of Science degrees in Biology and

Chemistry and Bachelor of Arts Degree in Music 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 University 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, College of Medicine, University of South

Florida, Tampa, FL. In addition to his scientific accomplishments, Thomas received

acclaim as a classical pianist, giving recitals and performing in concert.