Structure-Function Studies of 5-Aminolevulinic Acid (Ala) Synthases

Home , DMSO reductase, Tryptophan synthase

STRUCTURE-FUNCTION STUDIES OF 5-AMINOLEVULINIC ACID (ALA) SYNTHASES

James Chege Kaganjo

A Dissertation

Submitted to the Graduate College of Bowling Green State University in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

December 2017

Committee:

Jill Zeilstra-Ryalls, Advisor

Maria Rizzo Graduate Faculty Representative

Ray Larsen

Scott Rogers

Paul Morris

James Kaganjo

Jill Zeilstra-Ryalls, Advisor

The essential metabolite 5-aminolevulinic acid (ALA) is a precursor in the production of tetrapyrroles and 2-amino-3-hydroxycyclopent-2-en-1-one (C5N unit), a substructure of C5N- containing polyketides. In non-plant eukaryotes and Į-proteobacteria, the pyridoxal 5’-phosphate

(PLP)-dependent ALA synthase enzyme catalyzes the synthesis of ALA. While an understanding as to the distinct roles of multiple ALA synthases in animals is well developed, much less is known about the presence of more than one enzyme in bacteria. The role of HemA and HemT ALA synthase isoenzymes in Rhodobacter sphaeroides was investigated by comparing the enzymatic properties of three HemAs and two HemTs from three strains: one strain has hemA and hemT genes present and both are expressed, another has both genes but hemT is not expressed, and the third strain has the hemA gene only. Although all five enzymes had similar kinetic properties, HemA enzymes were more sensitive to hemin with a 13-fold difference in Ki value compared to HemT. HemT was found to be sensitive to oxidation suggesting that the hemT gene encodes an enzyme that is more active anaerobically, which agrees well with the maximal anaerobic-dark transcription of hemT in strain 2.4.9 (Coulianos N,

Kaganjo J, and JH Zeilstra-Ryalls. 2017. Mauscript in preparation). These properties indicate that the role of HemT is to supply ALA when inhibitory heme levels are elevated in the cell, which is thought to occur when cells undergo a transition from aerobic to anaerobic-dark respiration. iv In addition to catalyzing the ALA synthesis reaction, bifunctional ALA synthases

(cyclizing ALA synthases) in actinomycetes catalyze the cyclization of ALA-CoA to form the

C5N unit. Whether other ALA synthases whose only known function is providing ALA for tetrapyrrole biosynthesis also possess this bifunctionality is not known. This question was investigated by comparing the ALA-CoA cyclization activity of R. sphaeroides strain 2.4.9

HemA and HemT enzymes relative to that of a bona fide cyclizing ALA synthase, AcaC of

Streptomyces aizunensis strain NRLL-B-11277. Although both HemA and HemT had cyclization activity, it was only 10% and 6% that of AcaC respectively. However, the specific

ALA synthase activities of these "classical" ALA synthases were 315-fold higher than that of the

AcaC. A study of mutants in which conserved amino acid residue differences at positions 83 and

363 in each kind of enzyme were investigated revealed that they are not solely responsible for the differences in ALA synthase and ALA-CoA cyclization activities of these enzymes.

Bioinformatic analyses suggest that the differences in activities could be due to differences in conformational changes as the enzymes undergo catalysis, and active site architecture.

To my parents. vi ACKNOWLEDGMENTS

First and foremost, I would like to thank my advisor Dr. Jill Zeilstra-Ryalls for giving me the opportunity to work in her lab for the past six years. Without her support, encouragement and understanding I would not have achieved so much. I would also like to thank my committee members, Dr. Ray Larsen, Dr. Scott Rogers, Dr. Paul Morris and Dr. Maria Rizzo for their support, guidance and support. I would like to thank Dr. Wenjun Zhang and Dr. Joyce Liu from

UC Berkeley, for their work on the ALA-CoA cyclization assays, creating the HemA and AcaC mutants and providing the AcaC, and I would like to thank current and past members of the

Zeilstra lab for providing a great environment to pursue my studies and for the helpful and insightful discussions that were always useful in my research.

Finally, I would like to acknowledge my late parents for their support, sacrifice and determination to ensure I get a good education. vii

TABLE OF CONTENTS

Page

CHAPTER I. INTRODUCTION ...... 1

5-aminolevulinic acid...... ………………………………………………... 1

ALA synthesis……..………………………………………………………………… 1

Pyridoxal 5’-phosphate ...... 4

PLP-dependent enzymes……..……………………………………………………… 5

Į-oxoamine synthase family of fold type I………………………………………….. 9

ALA synthase……………………………………………...... 12

Research overview………………………………………………………………….. 15

References ...... 17

CHAPTER II. ROLE OF ALA SYNTHASE ISOENZYMES IN Rhodobacter sphaeroides…………………………………………………….………………...... ………. 25

Introduction………...……………………………………………………………….. 25

Rhodobacter sphaeroides...... 25

Tetrapyrrole biosynthesis in R. sphaeroides ...... 25

Redox sensitivity of ALA synthase ...... 28

ALA synthase isoenzymes ...... 29

Heterodimerization of HemA and HemT ...... 32

Materials and methods ...... 34

Bacterial strains, plasmids and expression of HemA and HemT ...... 34 viii

Protein purification ...... 36

Coaffinity purification of HemA and HemT heterodimer ...... 37

Ultraviolet-visible spectroscopy of HemA and HemT ...... 38

Protein concentration and analysis ...... 38

Immunoblot analysis ...... 39

InVision™ his-tag in-gel stain assay ...... 40

Chemical cleavage of HemA ...... 41

ALA synthase and 2-amino-3-ketobutyrate CoA ligase activity assays ...... 41

Hemin inhibition assays ...... 42

Alkylation of cysteine residues in HemA and HemT ...... 43

Bioinformatic analysis and protein modelling ...... 43

Docking of substrates and cofactor into the active site of HemA

and HemT protein models ...... 44

Results ...... 47

Protein expression and purification ...... 47

Catalytic properties ...... 51

Substrate preference ...... 54

Feedback inhibition by hemin ...... 55 ix

Sensitivity of HemA and HemT to oxidation ...... 60

Role of ClpX in activating HemA and HemT ...... 66

Isolation of HemA and HemT heterodimers ...... 70

Discussion ...... 76

References ...... 81

CHAPTER III. STRUCTURE-FUNCTION ANALYSIS OF THE BIFUNCTIONALITY

OF ALA SYNTHASES ...... ……………………………. 92

Introduction ...... …………………………………………………………….. 92

Materials and methods……………………………………… ..……………………. 102

Bacterial strains, plasmids and protein production ...... 102

Protein purifications, concentration determinations, SDS-PAGE

and InVision™ his-tag in-gel stain assay, and ALA synthesis activity

assays and determination of kinetic parameters ...... 103

ALA-CoA cyclization activity assays ...... 104

Bioinformatic analyses, protein modelling and structural analyses ...... 104

Results ……………………………………………………...... 107

Protein purification ...... 107 x

ALA-CoA cyclase activities of classical and cyclizing ALA synthases ...... 108

ALA synthase activities of classical and cyclizing ALA synthases ...... 112

Discussion ...... ………………………………………………….. 117

References ...... 134

CHAPTER IV. SUMMARY AND CONCLUSIONS ...... 140

References ...... ……………………………………………………………………………… 145 xi

LIST OF FIGURES

Figure Page

1 The biosynthetic pathways of ALA formation, and the products

derived from ALA ...... 3

2 Structures of vitamin B6 and pyridoxal 5’-phosphate...... 5

3 Chemical structures of the internal and external aldimines of PLP-dependent

enzymes...... 6

4 Resolved crystal structures of representative enzymes belonging to each of

the five fold types of PLP-dependent enzymes ...... 8

5 Superpositioned monomeric structures of five of the six members of the

Į-oxoamine synthase family ...... 11

6 Structure of R. capsulatus ALA synthase homodimer ...... 13

7 The active site of ALA synthase with bound succinyl-CoA and glycine-PLP ...... 14

8 The branched pathway of tetrapyrrole biosynthesis in R. sphaeroides ...... 27

9 Growth profiles of R. sphaeroides 2.4.9 wild type and hemT mutant bacteria

inoculated from aerobic-dark precultures and incubated under anaerobic dark

conditions with DMSO ...... 32

10 Multiple sequence alignment of ALA synthases from R. sphaeroides and

Rhodobacter capsulatus ...... 48 xii

11 SDS-PAGE and Invision His-Tag staining of proteins purified from

E. coli expression strains using Ni-affinity chromatography ...... 49

12 Color and UV absorbance spectra of purified HemA and HemT ...... 51

13 Percentage activity remaining of HemA and HemT enzymes in the presence

of different concentrations of hemin ...... 58

14 Characterization of the inhibition mechanism of HemA and HemT by hemin ...... 59

15 Time-dependent inhibitory effect of sulfhydryl modifying agents on

HemA and HemT ...... 61

16 Predicted structure of R. sphaeroides 2.4.9 HemT homodimer ...... 63

17 Multiple sequence alignment of ALA synthases in the region of HemT

cysteine 281 ...... 64

18 Percentage activity remaining of R. sphaeroides 2.4.9 HemT wild type and

HemT_C281P mutant enzymes in the presence of different concentrations

of hemin ...... 66

19 Multiple sequence alignment of ClpX amino acid sequences from

R. sphaeroides ATCC 17209, E. coli, D. rerio (zebra fish)

and S. cerevisiae (yeast) ...... 68

20 SDS-PAGE and Western blot analysis of differentially tagged proteins xiii

purified from E. coli overexpressing either His6-tagged HemT_C281P or

Strep-tagged HemA only using Strep-tactin and Nickel affinity chromatography .... 73

21 SDS-PAGE and Western blot analysis of differentially tagged proteins

purified from E. coli using Strep-tactin and Nickel affinity chromatography ...... 74

22 SDS-PAGE and immunoblot analysis of samples incubated with

hydroxylamine-HCl ...... 76

23 C5N unit-containing polyketides produced by actinomycetes ...... 93

24 The two biosynthetic pathways of ALA ...... 94

25 Synthesis of 2-amino-3-hydroxycyclopent-2-en-1-one (C5N unit)

in actinomycetes...... 96

26 The reaction mechanisms for ALA synthesis and ALA-CoA cyclization

by ALA synthase...... 100

27 SDS-PAG of purified proteins ...... 108

28 Percent relative cyclase activities of purified S. aizunensis wild type AcaC

and mutants AcaC _S83T, AcaC _S363T, and AcaC _S83T&S363T, and

R. sphaeroides 2.4.9 wild type HemA and mutants HemA_T83S,

HemA_T363S, and HemA_T83S&T363S, and

R. sphaeroides 2.4.9 wild type HemT ...... 109 xiv

29 Predicted structures of R. sphaeroides 2.4.9-HemA and S. aizunensis AcaC with

substrates docked in the active site ...... 111

30 Predicted structures of R. sphaeroides 2.4.9-HemA and S. aizunensis AcaC with

substrates docked in the active site ...... 114

31 Multiple amino acid sequence alignment of cyclic and classical ALA synthases,

together with partial sequences of hepta-variant mutant mouse ALAS2 with

substituted residues highlighted in cyan ...... 117

32 Solved crystal structures of R. capsulatus ALA synthase in open

(PDB ID 2BWN-light grey) and closed (PDB ID 2BWO-dark grey)

conformations superpositioned on each other ...... 123

33 Solved crystal structures of E. coli AONS superpositioned on models of the

same protein modelled using R. capsulatus ALA synthase ...... 125

34 Crystal structures of the members of Į-oxoamine synthase family

(A-AONS, B-SPT, C-KBL, D-CsqA) in their closed conformation

superpositioned with the crystal structure of R. capsulatus ALA synthase

crystal structure in closed conformation (PDB ID 2BWO) ...... 128

35 Second channel of the predicted tertiary structures of HemA (A and B)

and AcaC (C and D) ...... 132

36 The structures of heme B, heme A and bacteriochlorophyll ...... 141 xv

LIST OF TABLES

Table Page

1 Members of the Į-oxoamine synthase family ...... 10

2 Bacterial strains and plasmids ...... 44

3 Kinetic properties of R. sphaeroides HemA and HemT enzymes ...... 53

4 Members of the Į-oxoamine synthase family of fold type I PLP-dependent

enzymes ...... 54

5 Specific activities of R. sphaeroides 2.4.9 HemA and HemT using different

substrates ...... 55

6 Specific activities of R. sphaeroides 2.4.9 HemA and HemT incubated

with different concentrations of vitamin B12 ...... 57

7 Specific activities of R. sphaeroides 2.4.9 ALA synthases purified in the

presence of different reducing agents ...... 62

8 Specific activities and kinetic properties of R. sphaeroides 2.4.9 HemT

wild type and HemT_C281P mutant ALA synthases purified in the presence

of different reducing agents ...... 65

9 Specific activities of R. sphaeroides 2.4.9 HemT_C281P mutant ALA

synthases purified from clpX minus and clpX plus E. coli strains ...... 69

10 Protein yields from the coexpression system of differentially tagged HemA xvi

and HemT polypeptides ...... 72

11 Bacterial strains and plasmids ...... 105

12 Kinetic properties of R. sphaeroides 2.4.9 HemT, HemA and HemA mutant

and wild type S. aizunensis ALA synthases...... 116 xvii

LIST OF ABBREVIATIONS

Pg microgram

Pl microliter

PM micromolar

Å Angstroms

ALA 5-aminolevulinic acid

ALAS 5-aminolevulinic acid synthase

AONS 8-amino-7-oxononanoate synthase

BSA Bovine serum albumin

C5N 2-amino-3-hydroxycyclopent-2-en-1-one

CAI-1 3-aminotridecan-4-one

CoA Coenzyme A

CSS complex formation significance score

DAHP 3-Deoxy-D-arabino-heptulosonate-7-phosphate

DMSO Dimethyl sulfoxide

DTT Dithiothreitol

ES Enzyme substrate complex

ES‡ ES at the transition state xviii

GABA gamma-aminobutyric acid

GLU glutamate

GSAM glutamyl-1-semialdehyde aminotransferase

IPTG isopropyl ȕ-D-1-thiogalactopyronoside

KBL 2-amino-3-ketobutyrate-CoA ligase kDa Kilo Dalton

Ki inhibition constant

Kn Kanamycin

KnR Kanamycin resistant

LAI-1 3-hydroxypentadecan-4-one

LOMETS Local Meta-Threading-Server mg milligram mM millimolar

Ni-IDA Nickel iminodiacetic acid nm nanometer

OD optical density

PAGs polyacrylamide gels

PDB Protein Data Bank xix

PDBePISA Protein Data Bank in Europe Proteins, Interface, Structures and Assemblies

PLP pyridoxal-5-phosphate

RMSD Root-mean-square deviation

RNA Ribonucleic acid

S Substrate

SAM S-adenosylmethionine

SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis

SpR Spectinomycin resistant

SPT Serine palmitoyl transferase

StR Streptomycin resistant

TCA Trichloroacetic acid tRNA transfer RNA tRNAGLU glutamate transfer RNA

UV Ultra violet

UV-VIS ultraviolet-visible

Vmax maximum velocity

Vo initial velocity

XLEPP X-linked erythropoetic protoporphyria xx

XLSA X-linked sideroblastic anemia

MEL Murine erythroleukemia 1

CHAPTER I

INTRODUCTION

5-aminolevulinic acid

The essential metabolite 5-aminolevulinic acid (ALA) plays a role in all living organisms as a precursor in the biosynthesis of tetrapyrroles (1). These molecules play important roles in many biological processes. Examples of cyclic tetrapyrroles include heme that binds iron and is part of hemoproteins involved in oxygen and electron transport, chlorophyll

(bacteriochlorophyll) that binds magnesium and plays a role in harvesting light energy in photosynthesis, and vitamin B12 that binds cobalt and is an essential cofactor for various enzymes (2). ALA is also a precursor in the synthesis of 2-amino-3-hydroxycyclopent-2-en-1- one five membered ring, or C5N unit (3). The C5N unit is part of the structure of several secondary metabolites that are produced by some actinomycetes.

ALA synthesis

The first precursor of tetrapyrroles to be discovered was glycine in 1945, when David

Shemin ingested 15N-labeled glycine and found that it was incorporated into hemin (4). The second precursor was discovered in 1951 by Shemin and Wittenberg, who reported that a succinyl-coenzyme complex derived from the tricarboxylic acid cycle was a precursor of tetrapyrroles (5). The complex was later confirmed by Gibson et al. and Kikuchi et al. to be succinyl-CoA (6, 7). In 1953 Shemin and Russel reported that ALA could replace glycine and succinate in tetrapyrrole biosynthesis, thus establishing it as the first committed precursor (1).

The enzymatic synthesis of ALA from glycine and succinyl-CoA by ALA synthase was later described by Kikuchi et al. (7) and Gibson et al. (6) in 1958. Synthesis of ALA by ALA synthase requires pyridoxal 5’-phosphate (PLP), which is bound to the enzyme through a Schiff 2 base to a catalytic lysine residue in the active site (8). This biosynthetic pathway of ALA synthesis by ALA synthase is known as the C4 or Shemin pathway and it supplies ALA for tetrapyrrole biosynthesis in non-plant eukaryotes and Į-proteobacteria (Figure 1).

In 1974, Beale and Castelfranco reported that ALA in plants was synthesized from glutamate instead of succinyl-CoA and glycine (9). It was subsequently shown that this process required three enzymes (10-14). The first is glutamyl-tRNA synthetase, the same enzyme that is involved in protein synthesis. The other two enzymes, glutamyl-tRNA reductase and PLP- dependent glutamyl-1-semialdehyde aminotransferase (10-14), are specific to ALA formation.

This so-called C5 pathway, which is present in plants, archaea, and bacteria other than alpha- proteobacteria (15), is shown in Figure 1.

Although ALA production in most organisms is formed exclusively by one or the other of the biosynthetic pathways, some exceptions are known. In Euglena gracilis the C5 pathway is present in the chloroplasts where it supplies ALA for chlorophyll biosynthesis. The C4 pathway is present in the mitochondria of the organism, where it supplies ALA for heme biosynthesis (16,

17). The C4 and C5 pathways are also present in actinomycetes that produce C5N unit- containing polyketides (18). In these bacteria, it has been shown that the C5 pathway supplies

ALA exclusively for tetrapyrrole biosynthesis while the C4 pathway supplies ALA exclusively for production of the C5N unit that forms part of the structure of certain secondary metabolites

(18).

Glutamyl-1-semialdehyde aminotransferase (GSAM), the last enzyme in the C5 pathway that converts glutamate-1-semialdehyde into ALA, and ALA synthase are both PLP dependent.

GSAM is proposed to have existed before ALA synthase and that ALA synthase evolved from

GSAM as a means of separating ALA synthesis from protein synthesis (19). 3

Figure 1. The biosynthetic pathways of ALA formation, and the products derived from ALA.

The dedicated reactions of the C5 pathway are enclosed by a rectangle. In the first reaction 4 glutamyl-tRNAGlu is reduced to glutamate-1-semialdehyde by glutamyl-tRNA reductase. PLP- dependent glutamyl-1-semialdehyde aminotransferase then catalyzes the synthesis of ALA from glutamate-1-semialdehyde (19). The C4 pathway, in which ALA is formed through the condensation of glycine and succinyl-CoA is enclosed by an oval. This reaction is catalyzed by

PLP-dependent ALA synthase (20). The rest of the tetrapyrrole biosynthetic pathway showing the synthesis of the three major cyclic tetrapyrroles and the structures of uroporphobilinogen III, the last common precursor in the biosynthetic pathway, and protoporphyrin IX, which marks the branching point of heme and chlorophyll synthesis, are highlighted in grey. The C5N unit and the secondary metabolite asukamycin produced by Streptomyces nodosus subsp asukaensis (3) are enclosed by a rounded rectangle.

Pyridoxal 5’-phosphate

Pyridoxal 5’-phosphate (PLP) is the biologically active form of vitamin B6 (21). The aldehyde group of PLP (Figure 2) can form a Schiff base with amino acid substrates, and the electron withdrawing potential of the pyridine ring (Figure 2) moiety acts as an electron sink for substrates bound at the aldehyde position which stabilizes the quinonoid intermediate (22), These properties enable PLP to catalyze a wide range of enzymatic and non-enzymatic reactions that include transaminations, racemizations, Į-decarboxylation, Į-eliminations, aldo cleavage, ȕ and

Ȗ eliminations (22-24). 5

Figure 2. Structures of vitamin B6 and pyridoxal 5’-phosphate. The aldehyde group and the pyridine ring that endow PLP with its chemical properties are identified by rectangles.

PLP-dependent enzymes

PLP-dependent enzymes catalyze a wide range of reactions. They include oxidoreductases, isomerases, lyases, hydrolases and transferases (25). In fact, these represent five of the six general classes of enzyme-catalyzed reactions based on the enzyme commission nomenclature classification. The apo enzyme stabilizes the reaction intermediates of the PLP- catalyzed reactions, which lowers the transition energy, thereby speeding up the reactions (23,

26). The amino acid residues in the active sites of the enzymes recognize and properly orient substrates and intermediates, which facilitates breakage of the correct bonds ensuring reaction specificity. The active site of the enzyme also provides an environment that excludes water molecules, thereby minimizing PLP side reactions (23). In all PLP-dependent enzymes that catalyze reactions involving amino acids, PLP is bound in the active site to a lysine residue through the formation of an internal aldimine bond, as shown in Figure 3 (26, 27). Binding of an amino acid substrate displaces the internal aldimine to form an external aldimine (Figure 3).

Reaction specificity in PLP-dependent enzymes arises from (a) the type of bond that is broken at 6 the Į-carbon position of the amino acid substrate, and (b) differences in the covalent modifications that take place in the successive steps (28).

Figure 3. Chemical structures of the internal and external aldimines of PLP-dependent enzymes.

In the resting state, PLP is bound to the catalytic lysine residue in the active site to form an internal aldimine. Binding of substrate displaces the internal aldimine to form the external aldimine intermediate (26).

The first classification of PLP-dependent enzymes catalyzing reactions involving amino acid substrates was based on the chemical properties of the enzymes, which focused on the differences in the positions of the carbon atom that undergo covalent modification in the amino acid substrate. This classification grouped these enzymes into three classes, Į, ȕ, and Ȗ.

Members of the Į class catalyze reactions that involve covalent changes on the Į-carbon and catalyze transamination, decarboxylation, elimination and replacement reactions. The ȕ class members catalyze reactions that involve covalent changes on the ȕ-carbon. Members of this class catalyze elimination and replacement reactions. The Ȗ family members catalyze reactions that involve covalent changes on the Ȗ-carbon. Members of this class also catalyze elimination and replacement reactions (27). Although members of each class have the same chemical 7 properties, alignment of their amino acid sequences reveals that they have different evolutionary lineages (27).

As more amino acid sequences and crystal or model structures of PLP-dependent enzymes became available, PLP-dependent enzymes were reclassified based on similarities in amino acid sequences, and on secondary and tertiary structure similarities. Thus, this classification pertains to their evolutionary relationships, and defines five distinct fold types (29).

Solved crystal structures representing enzymes from each of these fold types are shown in Figure

4. Enzymes that belong to fold type I are usually active as homodimers with a few exceptions, which act as higher order oligomers. The active sites of the homodimeric enzymes are located on the subunit interface. Consequently, residues from both subunits are involved in substrate and cofactor binding. The two active sites of these enzymes function independently, although a few cases of asymmetry have been reported, including gamma-aminobutyric acid (GABA) aminotransferase (30) and glutamate-1-semialdehyde aminotransferase (31). Members of fold type II function as homodimers or oligomers as well, although the functional residues in the active sites are only contributed by one subunit (32, 33). Members of fold types III function as homodimers with each subunit folding into two distinct domains; PLP binds in a cleft located in one of these domains. Members of fold type IV also function as homodimers with each subunit folding into two domains; the active site of these enzymes is located on the domain interface.

Fold type V enzymes differ from those of the other fold types in that they use the phosphate group of PLP for catalysis rather than the pyridine ring and the aldehyde group (29, 34, 35). 8

Figure 4. Solved crystal structures of representative enzymes belonging to each of the five fold types of PLP-dependent enzymes. Aspartate amino transferase (A) is representative of fold type

I enzymes, tryptophan synthase (B) is representative of the fold type II enzymes, alanine racemase (C) is representative of fold type III enzymes, D-amino acid aminotransferase (D) is 9 representative of fold type IV enzymes, and maltodextrin phosphorylase (E) is representative of fold type V enzymes. The structures were obtained from the Protein Data Bank (PDB) and the figure was generated using PyMOL (The PyMOL Molecular Graphics System, Version 1.4.1

Schrödinger, LLC.).

Į-oxoamine synthase family of fold type I enzymes

Evolutionary differences within members of the fold type I enzymes are evident from the differences in the conformation of the N terminal part of their tertiary structures (35). Based on these differences, fold type I PLP-dependent enzymes can be further subgrouped into eight subclasses (34, 35). Members of the Į-oxoamine synthase family belong in subclass II of fold type I PLP-dependent enzymes. These enzymes catalyze reactions between a small amino acid and acyl-coenzyme A esters to form 1,3-aminoketones (25, 36). The Į-oxoamine synthase family currently consists of six members as shown in Table 1. Except for LqsA, crystal structures of the other five enzymes in this subgroup are available, and they reveal that they are very similar in structure, as shown by the superpositioning of their monomers in Figure 5.

Table 1. Members of the Į-oxoamine synthase family.

Enzyme Substrates Product Role References

ALA synthase L-glycine and 5-aminolevulinic acid Tetrapyrrole 7, 37

succinyl-CoA (ALA) biosynthesis

ALA-CoA 2-amino-3- Secondary 7, 37

hydroxycyclopent-2-en-1- metabolite

one biosynthesis

8-amino-7- L-alanine and 8-amino-7-oxononanoate Biotin 38 oxononanoate pimeloyl-CoA biosynthesis synthase (AONS)

Serine palmitoyl L-serine and 3-ketodihydrosphingosine Sphingolipid 39 transferase (SPT) palmitoyl-CoA biosynthesis

2-amino-3- L-glycine and Į-amino-ȕ-ketobutyrate Threonine 40 ketobutyrate-CoA acetyl-CoA metabolism ligase (KBL)

CqsA (S)-2- 3-aminotridecan-4-one Quorum 41

aminobutyrate/S- (CAI-1) sensing

adenosylmethionine

(SAM) and

decanoyl-CoA

LqsA Not identified 3-hydroxypentadecan-4- Quorum 42

one (LAI-1) sensing 11

Figure 5. Superpositioned monomeric structures of five of the six members of the Į-oxoamine synthase family. Shown are Rhodobacter capsulatus ALA synthase (magenta), PDB ID 2BWN

(43), Escherichia coli KBL (grey), PDB ID 1FC4 (44), E. coli AONS (cyan), PDB ID 1DJE

(45), Sphingomonas paucimobilis SPT (yellow), PDB ID 2JG2 (46) and Vibrio cholera CsqA

(orange), PDB ID 2WKA (41). Bound PLP is shown as spheres. The structures were obtained from PDB and aligned with PyMOL. The figure was generated using PyMOL (The PyMOL

Molecular Graphics System, Version 1.4.1 Schrödinger, LLC.).

The reaction mechanisms of all members of the Į-oxoamine synthase family are very similar, as outlined here (7, 44, 45, 47-52): (A) Prior to substrate binding, the enzyme is in the open conformation. Binding of the amino acid substrate leads to the formation of the external aldimine and triggers the enzyme to begin transitioning to the closed conformation. (B) The Į- proton of the amino acid substrate is abstracted to form a resonance stabilized quinonoid 12 intermediate. Subsequent binding of the acyl-CoA substrate accelerates the abstraction of the pro-R proton and also the adoption of the closed conformation of the enzyme. (C) The quinonoid intermediate attacks the acyl-CoA substrate to form a ȕ-ketoacid and subsequent loss of the CoA. (D) With the exception of KBL where the ȕ-ketoacid retains its carboxyl group, the quinonoid product is then formed by decarboxylation of the ȕ-ketoacid. (E) This quinonoid product is then protonated. (F) Finally, the product is released, which is the rate limiting step of the reaction.

ALA synthase

The polypeptides of ALA synthase homodimers have molecular masses that range from

44 to 65 kilodaltons. Eukaryotic ALA synthase genes are encoded for in the nucleus and they possess a leader peptide sequence on the N terminus that is essential for the transport of the precursor ALA synthase into the mitochondrion. Once inside, the leader sequence is cleaved to form the mature ALA synthase. The only available crystal structures for the enzyme are those of

R. capsulatus ALA synthase (43); one structure is that of PLP bound to the enzyme (PDB ID

2BWN), another is that of glycine bound to the enzyme (PDB ID 2BWP), and a third is that of enzyme complexed with succinyl-CoA and PLP (PDB ID 2BWO). As shown in Figure 6, ALA synthase has two active sites that are located on the subunit interface with amino acids from both monomers contributing to the binding of substrates (Fig 7; 43, 53). 13

Figure 6. Structure of R. capsulatus ALA synthase homodimer PDB ID 2BWO. Each polypeptide is colored differently (cyan and grey), with PLP and succinyl-CoA (shown in sticks) bound to the 2 active sites formed at the interface of the 2 polypeptides. The figure was generated using PyMOL (The PyMOL Molecular Graphics System, Version 1.4.1 Schrödinger,

LLC.).

As shown in Figure 7, PLP is bound deep inside the active site pocket, which helps in excluding water molecules, which enhances reaction specificity by eliminating the possibility of side reactions taking place. It is the amino group of glycine that forms a Schiff base with PLP, with the carboxylate portion of the cofactor bound by serine, asparagine and arginine residues through hydrogen bonds as shown on Figure 7. The adenylyl moiety of succinyl-CoA sits in a hydrophobic pocket on the surface of the enzyme and forms hydrogen bonds with serine 137, aspartate 138 and serine 139, which play a role in the specificity of the enzyme for succinyl- 14

CoA. The carboxyl group of succinate is bound deeply within the active site, in a region that has high affinity for acidic groups (43). How the enzyme harnesses the binding energy generated by succinyl-CoA binding to adopt the closed conformation is currently not known.

Figure 7. The active site of ALA synthase with bound succinyl-CoA and glycine-PLP (43).

Amino acid residues that form hydrogen bonds (black dashed lines) are shown as blue sticks with those from the second subunit shown as red sticks. The figure was generated using PyMOL

(The PyMOL Molecular Graphics System, Version 1.4.1 Schrödinger, LLC.).

Research overview

While studies continue toward further resolving mechanistic details of catalysis by ALA synthases, there are many other questions regarding the relationship between the structure and function of these enzymes. In mammals, there are two isoenzymes of ALA synthase whose roles are well-described. The housekeeping ALA synthase is expressed in all tissues, and supplies

ALA for basal heme requirements. The erythroid-specific ALA synthase is expressed in differentiating erythroid cells, where it supplies ALA for the large amount of heme required for hemoglobin synthesis (57). ALA synthase isoenzymes have also been reported in bacteria (58-

61). But their significance is yet to be resolved.

Another unresolved question pertains to the recently discovered bifunctionality of ALA synthases present in actinomycetes (62). In those species that produce secondary metabolites that contain the C5N unit as part of their structure, ALA synthase catalyzes the synthesis of ALA and, after its activation by addition of coenzyme A involving an acyl-CoA ligase, it also catalyzes the cyclization of ALA-CoA to form 2-amino-3-hydroxycyclopent-2-en-1-one five- membered rings (C5N units) that are then attached to the rest of the structure of the secondary metabolites (62). These ALA synthases are very similar to ALA synthases that are involved in

ALA synthesis for tetrapyrrole biosynthesis, which has led to the proposal that the synthases dedicated to tetrapyrrole biosynthesis can also catalyze the ALA-CoA cyclization reaction.

This dissertation describes studies on the structure-function relationships of ALA synthases that could explain (a) the role of ALA synthase isoenzymes in Rhodobacter sphaeroides, and (b) whether the bifunctionality of actinomycete ALA synthases that are involved in the synthesis of the C5N unit extends to ALA synthase enzymes whose main role is 16 to provide ALA for tetrapyrrole biosynthesis. Chapter two presents progress with respect to the former, and chapter three describes results of an investigation of the latter. 17

References

1. Shemin D, Russell CS. 1953. į-Aminolevulinic acid, itsrole in the biosynthesis of porphyrins and purines 1. J Am Chem Soc 75:4873-4874.

2. Milgrom LR. 1997. The colours of life: an introduction to the chemistry of porphyrins and related compounds. Oxford University Press, New York.

3. Nakagawa A, Wu TS, Keller PJ, Lee JP, Omura S, Floss HG. 1985. Biosynthesis of asukamycin. Formation of the 2-amino-3-hydroxycyclopent-2-enone moiety. J Chem Soc, Chem

Commun 1985(8):519-521.

4. Shemin D, Rittenberg D. 1945. The utilization of glycine for the synthesis of a porphyrin. J Biol Chem 159:567-568.

5. Shemin D, Wittenberg J. 1951. The mechanism of porphyrin formation; the role of the tricarboxylic acid cycle. J Biol Chem 192:315-334.

6. Gibson KD, Laver WG, Neuberger A. 1958. Initial stages in the biosynthesis of porphyrins. 2. The formation of delta-aminolaevulic acid from glycine and succinyl-coenzyme A by particles from chicken erythrocytes. Biochem J 70:71-81.

7. Kikuchi G, Kumar A, Talmage P, Shemin D. 1958. The enzymatic synthesis of delta- aminolevulinic acid. J Biol Chem 233:1214-1219.

8. Ferreira GC, Neame PJ, Dailey HA. 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.

9. Beale SI, Castelfranco PA. 1974. The biosynthesis of delta-aminolevulinic acid in higher plants: II. Formation of C-delta-aminolevulinic acid from labeled precursors in greening plant tissues. Plant Physiol 53:297-303. 18

10. Huang DD, Wang WY. 1986. Chlorophyll biosynthesis in Chlamydomonas starts with the formation of glutamyl-tRNA. J Biol Chem 261:13451-13455.

11. Huang DD, Wang WY, Gough SP, Kannangara CG. 1984. delta-aminolevulinic acid- synthesizing enzymes need an RNA moiety for activity. Science 225:1482-1484.

12. Avissar YJ, Beale SI. 1988. Biosynthesis of tetrapyrrole pigment precursors : formation and utilization of glutamyl-tRNA for delta-aminolevulinic acid synthesis by isolated enzyme fractions from Chlorella vulgaris. Plant Physiol 88:879-886.

13. Weinstein JD, Mayer SM, Beale SI. 1987. Formation of delta-aminolevulinic acid from glutamic acid in algal extracts : separation into an RNA and three required enzyme components by serial affinity chromatography. Plant Physiol 84:244-250.

14. Bruyant P, Kannangara CG. 1987. Biosynthesis of ǻ-aminolevulinate in greening barley leaves. VIII: Purification and characterization of the glutamate-tRNA ligase. Carlsberg

Res Commun 52:99-109.

15. Avissar YJ, Ormerod JG, Beale SI. 1989. Distribution of delta-aminolevulinic acid biosynthetic pathways among phototrophic bacterial groups. Arch Microbiol 151:513-519.

16. Mayer SM, Beale SI. 1992. Succinyl-coenzyme A synthetase and its role in 5- aminolevulinic acid biosynthesis in Euglena gracilis. Plant physiol 99:482-487.

17. Weinstein JD, Beale SI. 1983. Separate physiological roles and subcellular compartments for two tetrapyrrole biosynthetic pathways in Euglena gracilis. J Biol Chem

258:6799-6807.

18. Petrícek M, Petríceková K, Havlícek L, Felsberg J. 2006. Occurrence of two 5- amionolevulinate biosynthetic pathways in Streptomyces nodosus subsp. asukaensis is linked to the production of asukamycin. J Bacteriol 188:5113-5123. 19

19. Schulze JO, Schubert W-D, Moser J, Jahn D, Heinz DW. 2006. Evolutionary relationship between initial enzymes of tetrapyrrole biosynthesis. J Mol Biol 358:1212-1220.

20. Shemin D, Kikuchi G. 1958. Enzymatic synthesis of 5-aminolevulinic acid. Ann NY

Acad Sci 75:122-128.

21. Heyl D, Luz E, Harris SA, Folkers K. 1951. Phosphates of the vitamin B6 group. I. The structure of codecarboxylase. J Am Chem Soc 73:3430-3433.

22. Metzler DE, Ikawa M, Snell EE. 1954. A general mechanism for vitamin B6-catalyzed reactions. J Am Chem Soc 76:648-652.

23. Hayashi H, Wada H, Yoshimura T, Esaki N, Soda K. 1990. Recent topics in pyridoxal

5'-phosphate enzyme studies. Annu Rev Biochem 59:87-110.

24. Toney MD. 2005. Reaction specificity in pyridoxal phosphate enzymes. Arch Biochem

Biophys 433:279-287.

25. Percudani R, Peracchi A. 2003. A genomic overview of pyridoxal-phosphate-dependent enzymes. EMBO Rep 4:851-854.

26. Hayashi H. 1995. Pyridoxal enzymes: mechanistic diversity and uniformity. J Biochem

118:463-473.

27. Alexander FW, Sandmeier E, Mehta PK, Christen P. 1994. Evolutionary relationships among pyridoxal-5'-phosphate-dependent enzymes. Regio-specific alpha, beta and gamma families. Eur J Biochem 219:953-960.

28. John RA. 1995. Pyridoxal phosphate-dependent enzymes. Biochim Biophys Acta

1248:81-96.

29. Grishin NV, Phillips MA, Goldsmith EJ. 1995. Modeling of the spatial structure of eukaryotic ornithine decarboxylases. Protein Sci 4:1291-1304. 20

30. Churchich JE, Moses U. 1981. 4-aminobutyrate aminotransferase. The presence of nonequivalent binding sites. J Biol Chem 256:1101-1104.

31. Hennig M, Grimm B, Contestabile R, John RA, Jansonius JN. 1997. Crystal structure of glutamate-1-semialdehyde aminomutase: an Į2-dimeric vitamin B6-dependent enzyme with asymmetry in structure and active site reactivity. Proc Natl Acad Sci 94:4866-4871.

32. Hyde C, Ahmed S, Padlan E, Miles EW, Davies D. 1988. Three-dimensional structure of the tryptophan synthase alpha 2 beta 2 multienzyme complex from Salmonella typhimurium. J

Biol Chem 263:17857-17871.

33. Mehta PK, Christen P. 2009. The molecular evolution of pyridoxal-5'-phosphate- dependent. Advances in Enzymology and Related Areas of Molecular Biology, Volume 74, Part

B: Mechanism of Enzyme Action 227:129.

34. Schneider G, Kack H, Lindqvist Y. 2000. The manifold of vitamin B6 dependent enzymes. Structure 8:R1-6.

35. Käck H, Sandmark J, Gibson K, Schneider G, Lindqvist Y. 1999. Crystal structure of diaminopelargonic acid synthase: evolutionary relationships between pyridoxal-5ƍ-phosphate- dependent enzymes. J Mol Biol 291:857-876.

36. Eliot AC, Kirsch JF. 2004. Pyridoxal phosphate enzymes: mechanistic, structural, and evolutionary considerations. Annu Rev Biochem 73:383-415.

37. Zhang L, Chen J, Chen N, Sun J, Zheng P, Ma Y. 2013. Cloning of two 5- aminolevulinic acid synthase isozymes HemA and HemO from Rhodopseudomonas palustris with favorable characteristics for 5-aminolevulinic acid production. Biotechnol Lett 35:763-768. 21

38. Mann S, Ploux O. 2011. Pyridoxal-5'-phosphate-dependent enzymes involved in biotin biosynthesis: structure, reaction mechanism and inhibition. Biochim Biophys Acta 1814:1459-

1466.

39. Hanada K. 2003. Serine palmitoyltransferase, a key enzyme of sphingolipid metabolism.

Biochim Biophys Acta 1632:16-30.

40. Mukherjee JJ, Dekker EE. 1987. Purification, properties, and N-terminal amino acid sequence of homogeneous Escherichia coli 2-amino-3-ketobutyrate CoA ligase, a pyridoxal phosphate-dependent enzyme. J Biol Chem 262:14441-14447.

41. Jahan N, Potter JA, Sheikh MA, Botting CH, Shirran SL, Westwood NJ, Taylor

GL. 2009. Insights into the biosynthesis of the Vibrio cholerae major autoinducer CAI-1 from the crystal structure of the PLP-dependent enzyme CqsA. J Mol Biol 392:763-773.

42. Spirig T, Tiaden A, Kiefer P, Buchrieser C, Vorholt JA, Hilbi H. 2008. The

Legionella autoinducer synthase LqsA produces an alpha-hydroxyketone signaling molecule. J

Biol Chem 283:18113-18123.

43. Astner I, Schulze JO, van den Heuvel J, Jahn D, Schubert W-D, Heinz DW. 2005.

Crystal structure of 5-aminolevulinate synthase, the first enzyme of heme biosynthesis, and its link to XLSA in humans. EMBO J 24:3166-3177.

44. Schmidt A, Sivaraman J, Li Y, Larocque R, Barbosa JA, Smith C, Matte A, Schrag

JD, Cygler M. 2001. Three-dimensional structure of 2-amino-3-ketobutyrate CoA ligase from

Escherichia coli complexed with a PLP-substrate intermediate: inferred reaction mechanism.

Biochemistry 40:5151-5160. 22

45. Webster SP, Alexeev D, Campopiano DJ, Watt RM, Alexeeva M, Sawyer L, Baxter

RL. 2000. Mechanism of 8-amino-7-oxononanoate synthase: spectroscopic, kinetic, and crystallographic studies. Biochemistry 39:516-528.

46. Yard BA, Carter LG, Johnson KA, Overton IM, Dorward M, Liu H, McMahon SA,

Oke M, Puech D, Barton GJ, Naismith JH, Campopiano DJ. 2007. The structure of serine palmitoyltransferase; gateway to sphingolipid biosynthesis. J Mol Biol 370:870-886.

47. Ikushiro H, Fujii S, Hayashi H. 2007. Analysis of the reaction mechanism of serine palmitoyltransferase. Chem Phys Lipids 149:S63.

48. Ikushiro H, Islam MM, Okamoto A, Hoseki J, Murakawa T, Fujii S, Miyahara I,

Hayashi H. 2009. Structural insights into the enzymatic mechanism of serine palmitoyltransferase from Sphingobacterium multivorum. J Biochem 146:549-562.

49. Hunter GA, Zhang J, Ferreira GC. 2007. Transient kinetic studies support refinements to the chemical and kinetic mechanisms of aminolevulinate synthase. J Biol Chem 282:23025-

23035.

50. Zaman Z, Jordan PM, Akhtar M. 1973. Mechanism and stereochemistry of the 5- aminolaevulinate synthetase reaction. Biochem J 135:257-263.

51. Nandi DL. 1978. Studies on delta-aminolevulinic acid synthase of Rhodopseudomonas spheroides. Reversibility of the reaction, kinetic, spectral, and other studies related to the mechanism of action. J Biol Chem 253:8872-8877.

52. Kelly RC, Bolitho ME, Higgins DA, Lu W, Ng W-L, Jeffrey PD, Rabinowitz JD,

Semmelhack MF, Hughson FM, Bassler BL. 2009. The Vibrio cholerae quorum-sensing autoinducer CAI-1: analysis of the biosynthetic enzyme CqsA. Nat Chem Biol 5:891-895. 23

53. Tan D, Ferreira GC. 1996. Active site of 5-aminolevulinate synthase resides at the subunit interface. Evidence from in vivo heterodimer formation. Biochemistry 35:8934-8941.

54. Fratz EJ, Clayton J, Hunter GA, Ducamp S, Breydo L, Uversky VN, Deybach JC,

Gouya L, Puy H, Ferreira GC. 2015. Human erythroid 5-aminolevulinate synthase mutations associated with X-Linked protoporphyria disrupt the conformational equilibrium and enhance product release. Biochemistry 54:5617-5631.

55. Fratz EJ, Stojanovski BM, Ferreira GC. 2013. Toward heme: 5-aminolevulinate synthase and initiation of porphyrin synthesis in The Handbook of Porphyrin Science (Ferreira

G. C., Kadish K. M., Smith K. M., Guilard R., ed), pp. 1–78, World Scientific Publishing Co.,

New Jersey.

56. Kardon JR, Yien YY, Huston NC, Branco DS, Hildick-Smith GJ, Rhee KY, Paw

BH, Baker TA. 2015. Mitochondrial ClpX activates a key enzyme for heme biosynthesis and erythropoiesis. Cell 161:858-867.

57. Riddle RD, Yamamoto M, Engel JE. 1989. Expression of G-aminolevulinate synthase in avian cells: separate genes encode erythroid-specific and nonspecific isozymes. Proc Natl

Acad Sci USA 86:792-796.

58. Tuboi S, Kim HJ, Kikuchi G. 1970. Occurrence and properties of two types of delta- aminolevulinate synthetase in Rhodopseudomonas spheroides. Arch Biochem Biophys 138:147-

154.

59. Tai T-N, Moore MD, Kaplan S. 1988. Cloning and characterization of the 5- aminolevulinate synthase gene(s) from Rhodobacter sphaeroides. Gene 70:139-151. 24

60. Sato K, KoichiIshida K, Shirai M, Shimizu S. 2014. Occurrence and some properties of two types of į-aminolevulinic acid synthase in a facultative methylotroph,Protaminobacter ruber. Agric Biol Chem 49:3423-3428.

61. Zhang L, Chen J, Chen N, Sun J, Zheng P, Ma Y. 2013. Cloning of two 5- aminolevulinic acid synthase isozymes HemA and HemO from Rhodopseudomonas palustris with favorable characteristics for 5-aminolevulinic acid production. Biotechnol Lett 35:763-768.

62. Zhang W, Bolla ML, Kahne D, Walsh CT. 2010. A three enzyme pathway for 2- amino-3-hydroxycyclopent-2-enone formation and incorporation in natural product biosynthesis.

J Am Chem Soc 132:6402-6411.

CHAPTER II

ROLE OF ALA SYNTHASE ISOENZYMES IN Rhodobacter sphaeroides

Introduction

Rhodobacter sphaeroides. Rhodobacter sphaeroides are facultative phototrophic bacteria that belong to the Į-3 subclass of proteobacteria (1). They are well known for their metabolic flexibility. When oxygen is present, they rely upon aerobic respiration for cellular energy. When oxygen becomes limiting, they undergo morphological changes and synthesize a photosynthetic apparatus to capture light energy, if it is available, by anoxygenic photosynthesis.

When both oxygen and light are limiting, they can use anaerobic dark respiration with alternate electron acceptors such as dimethyl sulfoxide (DMSO). Finally, if external electron acceptors are absent, and there is no light, they can obtain energy through fermentation (2). The ability of

R. sphaeroides to use different types of energy metabolisms is enabled by its capacity to synthesize the tetrapyrroles vitamin B12, heme and bacteriochlorophyll.

Tetrapyrrole biosynthesis in R. sphaeroides. Tetrapyrroles are brightly colored porphyrin compounds that function as essential cofactors and pigments. The unsubstituted porphyrin (macrocyclic) structure consists of four pyrrole rings that are joined together at their Į- carbon atoms by methine bridges. Porphyrins bind a metal atom at the center of the porphyrin ring which is the main feature that gives them their biological functions. The major cyclic tetrapyrroles include heme that binds iron and is part of hemoproteins involved in oxygen and electron transport, chlorophyll (bacteriochlorophyll) that binds magnesium and plays a role in harvesting light energy in photosynthesis, and vitamin B12 that binds cobalt and is an essential cofactor for various enzymes (3). 26

The tetrapyrrole biosynthesis pathway as it takes occurs in R. sphaeroides is illustrated in

Figure 8. In this organism, 5-aminolevulinic acid (ALA), the first committed precursor in the pathway (4) is synthesized through the C4 or Shemin pathway by the decarboxylative condensation of succinyl-CoA (C4 precursor) and glycine in a reaction that is catalyzed by the

PLP-dependent enzyme ALA synthase (5). This pathway links tetrapyrrole synthesis to central carbon metabolism through the use of succinyl-CoA, and protein synthesis through glycine usage. The first branch point of the pathway occurs at the level of uroporphyrinogen III, which is, then, the last common intermediate for the three kinds of tetrapyrroles. Thereafter, one branch leads to the synthesis of vitamin B12 through multiple reactions that include the incorporation of cobalt. Other branches lead to the formation of heme and bacteriochlorophyll, which are both derived from protoporphyrin IX (6-8). Heme is synthesized by the incorporation of iron into protoporphyrin IX while bacteriochlorophyll synthesis involves a series of reactions, beginning with the incorporation of magnesium into protoporphyrin IX. 27

Figure 8. The branched pathway of tetrapyrrole biosynthesis in R. sphaeroides. Enzymes that are known to catalyze the reactions of the common tetrapyrrole biosynthesis pathway are indicated for each of the step. Vitamin B12 synthesis branches off from uroporphyrinogen III, and involves multiple additional steps (broken line), including the incorporation of cobalt. Heme and bacteriochlorophyll synthesis branches off from protoporphyrin IX. Heme is formed by incorporation of iron into protoporphyrin IX, incorporation of magnesium into protoporphyrin IX directs the tetrapyrrole toward bacteriochlorophyll formation.

The synthesis of tetrapyrroles in R. sphaeroides is highly regulated such that each type is synthesized at the appropriate overall and relative levels required to meet the metabolic needs of the cell. During aerobic growth, cells use heme in the aerobic respiratory chain, while bacteriochlorophyll synthesis is inhibited. The major factor that is responsible for this regulation is oxygen availability. When oxygen becomes limiting, heme and bacteriochlorophyll synthesis is increased in preparation for anoxygenic photosynthesis. This regulation is particularly important to limit the production of hazardous molecules such as singlet oxygen, which is promoted when bacteriochlorophyll is present under aerobic-light conditions (2). Vitamin B12 is required in trace amounts under all conditions by enzymes that catalyze a wide range of reactions in the cell involved in the synthesis of acetyl-CoA, methionine, tetrahydrofolate, deoxyribonucleotides and also bacteriochlorophyll synthesis (6, 9, 10). Hence, ALA is an essential metabolite under all conditions, and its regulated synthesis must be such that the need for all tetrapyrroles is met, regardless of the metabolic state of the cell. One of the mechanisms governing ALA production involves regulation of the first enzyme in the pathway, ALA synthase. Expression of ALA synthase genes is regulated at both the level of enzyme production

(11, 12) and the level of enzyme activity (12, 13). However, there have been no studies to examine regulation of ALA synthase activity since the discovery of more than one ALA synthase gene in these bacteria. What remains clear from those early studies is that neither bacteriochlorophyll a nor vitamin B12, the other end products of the pathway, inhibit ALA synthase activity (13).

Redox sensitivity of ALA synthase. The presence of redox sensitive disulfide compounds has been reported to increase the activity of ALA synthase (15-19). Furthermore, reduced cysteine residues that are maintained by a thioredoxin system have been shown to be 29 important for ALA synthase activity and may be involved in succinyl-CoA binding (18, 19).

Using a purification method that followed enzyme activity in different chromatography fraction peaks, Tuboi et al. (20) reported the purification of two separate chromatographic fraction peaks using a Sephadex G-200 column that had ALA synthase activity. The first fraction had high activity while the second fraction had low activity which was increased by incubating the fraction with disulfide compounds (15-17, 20). The redox sensitive fraction was reported to predominate in cells grown under semi-anaerobic conditions in the light (11). Since ALA synthase isoenzyme genes had not been reported in R. sphaeroides at that time, the protein composition of the fractions is not known and the significance of the differential redox sensitivity is unclear.

ALA synthase isoenzymes. A classic example of the possible role of isoenzymes in branched biosynthetic pathways such as the tetrapyrrole pathway in R. sphaeroides is that of the isoenzymes of aromatic amino acid biosynthesis in Escherichia coli. The common precursor in the pathway is 3-deoxy-D-arabino-heptulosonate-7-phosphate (DAHP), whose formation is catalyzed by DAHP synthases. E. coli has three such enzymes. Each of these isoenzymes is differentially feedback inhibited by tyrosine, phenylalanine, and tryptophan to achieve the balanced production of the three amino acids (21).

ALA synthase isoenzymes have been reported in animals and bacteria (20, 22-25). In animals, where the tetrapyrrole pathway is not branched as they only synthesize heme, there are two ALA synthase isoenzymes that are differentially expressed to meet the varying demand for heme in different cells (22). The two isoenzymes are nuclear-encoded and they have N-terminal sequences that target them to the mitochondria (26, 27). In humans, the gene encoding ALA synthase 1 (ALAS1) is located on chromosome 3, and it is constitutively expressed in all cells to 30 supply heme for hemoproteins such as cytochromes. The gene encoding the erythroid-specific enzyme ALA synthase 2 (ALAS2) is located on the human X chromosome. It is only expressed during erythroid differentiation in erythropoietic cells to supply heme specifically for hemoglobin synthesis. Other than the N-terminal mitochondrial targeting sequences, the two human isoenzymes are very similar to each other at the amino acid sequence level, with 74% identity and 87% similarity. Defects in ALAS2 enzyme, due to mutations in the single copy of the gene in males, manifest as X-linked sideroblastic anemia (XLSA) and X-linked erythropoetic protoporphyria (XLEPP) (28-33). Mutations have also been identified in the ALAS1 gene, however, the only phenotypic description is limited to their occurrence in tumor cells (34).

Presumably, ALAS1 is essential, and so loss-of-function genotypes would be lethal. Conversely, elevated, disregulated ALAS1 activity can lead to overproduction of porphyrins, causing a variety of neurological, digestive, and other disorders, a possibility that has led to several studies

(35, 36). Thus most of the studies of the animal ALA synthases have focused on expression (37-

42), and most enzyme kinetic studies have focused on ALAS2. As a consequence of the limited scope of these studies, it is not presently known whether the isoenzymes differ in terms of their kinetic properties, as their distinctive roles are only described with respect to their differential expression.

The significance of ALA isoenzymes in bacterial species is yet to be resolved. R. sphaeroides strain 2.4.1 was the first bacterium found to have two ALA synthase genes, hemA and hemT (23). The hemA gene is located on chromosome I while hemT is located on chromosome II (43). In that strain hemA gene is transcribed under all growth conditions and up- regulated in the absence of oxygen (43). Although all R. sphaeroides strains examined thus far have a hemA gene, not all possess a hemT gene (44). The two strains in which expression of 31 hemT has been studied to date are 2.4.1 and 2.4.9 (43, 45, 46). In R. sphaeroides 2.4.1, hemT expression has not been observed under any condition tested so far (43, 45, 46), while in R. sphaeroides 2.4.9, hemT is expressed under all conditions, and transcription is upregulated when the cells are growing under anaerobic-dark conditions in which dimethyl sulfoxide (DMSO) is provided as alternate electron acceptor (45). Unpublished results suggest that the differences in expression of hemT in the two strains are due to specialized sigma factors that are present in strain 2.4.9 but absent from strain 2.4.1, and the activity of these specialized sigma factors is governed by growth conditions, with maximal activity under anaerobic-dark with DMSO growth conditions (45, 46). However, anaerobic conditions also induce hemA transcription (45), which raises the question as to whether or not the HemA and HemT enzymes possess unique properties that require both to be expressed in this organism to support its full range of metabolic capabilities.

The importance of HemT enzyme is evident with the observation that hemT knock-out mutants of R. sphaeroides 2.4.9 have a longer lag phase compared to wild type cells when they are transitioning from aerobic-to anaerobic-dark with DMSO growth conditions (46, 47; Figure

9). To complete this transition, the cells need to assemble a new respiratory chain that delivers electrons to DMSO reductase. During the transitioning state, ALA synthesis activity would need to be increased since anaerobiosis is inducing for bacteriochlorophyll formation, while heme production must be sustained to meet the heme demand of newly forming respiratory chain. It is currently thought that up-regulation of hemA in strain 2.4.9 is not sufficient to meet this demand.

Consequently, hemT may be required to augment ALA production. In strain 2.4.1, hemA is greatly up-regulated, and so able to meet the increased ALA demand. This compensates for the absence of hemT expression. Whether these differences in transcription completely explain the 32

roles of hemA and hemT can only be determined by evaluating other possibilities; including the

possibility that these isoenzymes differ in their kinetic properties.

Figure 9. Growth profiles of R. sphaeroides 2.4.9 wild type and hemT mutant inoculated from aerobic-dark precultures and incubated under anaerobic dark conditions with DMSO. Values are presented as the mean of three replicates with standard errors of the mean indicated by error bars

(46, 47).

Heterodimerization of HemA and HemT. ALA synthases function as homodimers with two active sites that are located on the subunit interfaces, and with amino acids from both

monomers contributing to the binding of substrates (48, 49). Hence, for ALA synthase,

dimerization of the two subunits is critical for activity. Protein dimerization could also regulate

enzyme activity by acting as a regulatory mechanism that ensures sufficient protein is present in

the cell for the enzyme to dimerize (50). If the concentration is sufficient, the subunits dimerizes 33 and forms an active enzyme and if the concentration is low, dimerization will not take place and the enzyme remains inactive (50). Through intersubunit communication, dimerization can also influence enzyme activity by influencing substrate binding in the active site of each subunit through positive or negative cooperativity (50, 51). In positive cooperativity, substrate binding in the active site of one subunit leads to conformational changes that increase substrate affinity in the active site of the other subunit. In negative cooperativity, substrate binding in one active site leads to conformational changes that influences substrate binding and product release leading to low rate of catalysis in one of the active sites, as has been observed in glutamate-1-semialdehyde aminomutase (51).

Currently, the dimerization process of ALA synthase is not well understood. Since the polypeptides of ALA synthase isoenzymes are very similar, they could form heterodimeric proteins that possess enzymatic properties that are different from those of homodimeric ALA synthases thereby affecting production of ALA in the cell. R. sphaeroides HemA and HemT polypeptides are both 407 amino acids in length with predicted molecular masses of 44.62 and

44.31 kilodaltons respectively. The transcription profiles of hemA and hemT suggest that both polypeptides could co-exist in the cytoplasm (45). Whether heterodimeric ALA synthase, consisting of HemA and HemT subunits is formed under those circumstances is not known, nor what its contribution to ALA metabolism might be. Interestingly, it has been reported that during erythroid differentiation in murine erythroleukemia (MEL) cells, ALAS1 levels rapidly decrease as the ALAS2 levels increase rapidly to meet the high demand of heme in these cells

(37), suggesting that both enzymes are present for some period of time in erythroid cells, and the possible formation of ALAS1/2 heterodimers. 34

Herein, the overexpression, purification and characterization of HemA and HemT ALA synthases from three strains of R. sphaeroides is reported. HemA and HemT of R. sphaeroides could possess unique properties, which must be considered together with their differential expression in order to explain their presence in R. sphaeroides; the possibility that there are also

HemA-HemT heterodimers having properties that differ from the homodimers must also be considered.

To address these possibilities the specific aims are as follows:

1. Purify HemA and HemT, and determine their kinetic properties.

2. Investigate whether or not the HemA and HemT polypeptides can form heterodimers, and if so determine the kinetic properties of the heterodimeric enzyme.

Materials and methods

Bacterial strains, plasmids and expression of hemA, hemT and hemT_C281P.

Bacterial strains and plasmids are listed in Table 2. The expression system for the heterologous production of HemA and HemT proteins consists of the Escherichia coli strain DH5Įphe (52) and plasmids pIND4-(strain number)-hemA or pIND4-(strain number)-hemT or pIND4-2.4.9- hemT_C281P. To construct these expression vectors, synthetic polyhistidine-tagged hemA and hemT genes purchased from Integrated DNA Technologies (Coralville, IA) having NcoI recognition sites at their 5’ ends and HindIII recognition sequences at their 3’ ends were cloned into pIND4 (53), which had been treated with the same enzymes.

Expression of the hemA and hemT genes began by growing precultures prepared by inoculating Luria Bertani (LB) growth medium (54) containing 50 μg/ml kanamycin (Kn) from a frozen stock of the plasmid-bearing bacteria, which was then grown overnight at 37oC with 35 shaking at 200 rpm. LB-Kn medium was inoculated with the preculture at a volume/volume

o (v/v) ratio of 1:100 and incubated at 37 C with shaking at 200 rpm until OD660 of the culture reached approximately 0.6. Expression of polyhistidine-tagged hemA and hemT was then induced with the addition of isopropyl ȕ-D-1-thiogalactopyranoside (IPTG) (Sigma-Aldrich, St.

Louis, MO) to a final concentration of 0.4 mM, followed by an additional 2 h incubation. The cells were collected by centrifugation at 8327 × g for 15 min at 4°C in an Eppendorf 5810R centrifuge using an A-4-62 swinging bucket rotor (Westbury, NY) and the cell pellet was stored at -80°C.

Coexpression of differentially tagged (polyhistidine-tag and Strep-tag) proteins consisted of E. coli strain BW25113 (55, 56) harboring either pIND-(2.4.9-hemT_C281P-Strep)::ȍSpR/StR

R R and pBBR-(2.4.9-hemT_C281P-His6) or pIND-(2.4.9-hemA-Strep):: ȍSp /St and pBBR-(2.4.9-

R R hemT_C281P-His6) (Table 2). To construct pIND-(2.4.9-hemT_C281P-Strep)::ȍSp /St , synthetic DNA coding for (NcoI)-HemT_C281P-Strep-tag-(HindIII) was cloned into pIND4.

The plasmid was then restricted with ClaI for insertion of :SpR/StR and destruction of the KnR gene. To construct pIND-(2.4.9-hemA-Strep)::ȍ SpR/StR, synthetic DNA coding for (NcoI)-

HemA-Strep-tag-HindIII was cloned into pIND4. Then the plasmid was isolated from a dam+ strain of E. coli and restricted with ClaI for insertion of :SpR/StR and destruction of the KnR gene. Since the ClaI site internal to hemA is methylated by Dam, plasmid isolated from the dam+ strain was not restricted by the enzyme at that site. To construct pBBR-(2.4.9-hemT_C281P-

His6), a PvuII-HindIII fragment containing the gene for HemT_C281P-His6 was isolated from pIND-2.4.9-hemT_C281P and ligated with HincII-HindIII-treated pBBR1MCS2 (Table 2).

Coexpression began by preparation of a preculture by inoculating LB medium containing

50 μg/ml Kn and 50 μg/ml each of spectinomycin (Sp) and streptomycin (St) from a frozen stock 36 of the plasmid-bearing bacteria, which was then incubated overnight at 37oC with shaking at 200 rpm. The preculture was used to inoculate LB-Kn, Sp/St medium at a 1:100 v/v ratio. The culture was grown to an OD660 of 0.6 and protein expression was induced by the addition of

IPTG to a final concentration of 0.5 mM. Incubation continued overnight at 30°C with shaking at 200 rpm. The cells were collected by centrifugation at 8327 × g for 15 min at 4°C in a

Beckman J2-21 centrifuge (Beckman Coulter, Inc., Brea, CA) with a fiberlite F10BA-6x500Y rotor (Thermo Fisher Scientific Inc., Rockville, MD) and the cell pellet was stored at -80°C.

Protein purification. Cell pellets were resuspended in 2 ml wash buffer (20 mM tricine,

10% glycerol, 25 mM NaCl, 40 mM imidazole, 20 μM PLP, and 1 mM dithiothreitol (DTT), pH

7.2) with 100 μl protease inhibitor complex that has a cocktail of protease inhibitors (Sigma-

Aldrich, St. Louis, MO) that target a wide range of proteases including serine, cysteine, aspartyl- and metallo-endopeptidase proteases that may be present in the bacteria. The cells were lysed by passaging them through a French pressure cell (Thermo Fisher Scientific Inc., Milford, MA) at

700 lb/in2. The cell lysates were then incubated with DNaseI at a final concentration of 25 U/ml

(Sigma-Aldrich, St. Louis, MO) on ice for 30 min to degrade DNA that can block the nickel affinity resin. To pellet insoluble material, the lysates were centrifuged at 20,817 x g at 4°C for

15 min using an Eppendorf 5810R refrigerated centrifuge and a F-45-30-11 fixed-angle rotor

(Eppendorf North America, Hauppauge, NY). The cleared crude lysate was incubated with 1 ml

His60 Nickel superflowTM resin (Clontech Laboratories Inc., Mountain View, CA) in a closed 5 ml column placed on a 3-D rotator (Labline Scientific Instruments, Mumbai, India) for 1 h at

4°C. The column was then clamped vertically and the resin allowed to settle. To remove unbound proteins, the column was washed with 20 column volumes of wash buffer (20 mM tricine, 10% glycerol, 25 mM NaCl, 40 mM imidazole, 20 μM PLP, and 1 mM dithiothreitol 37

(DTT), which was determined to be sufficient to remove all unbound proteins by checking absorption at 280 nm. The bound proteins were then eluted using 10 column volumes of elution buffer (20 mM tricine, 10% glycerol, 25 mM NaCl, 350 mM imidazole, 20 μM PLP, and 1 mM

DTT, pH 7.2) and concentrated using Amicon Ultra-15 centrifugal filter device with a molecular mass cutoff of 10,000 daltons (Millipore, Bedford, MA) and stored at -80oC.

Coaffinity purification of HemA and HemT heterodimeric protein. Crude lysates of cells coexpressing either Strep-tagged HemT_C281P and polyhistidine-tagged HemT_C281P polypeptides or Strep-tagged HemA and polyhistidine-tagged HemT_C281P polypeptides were prepared as described above in 2 ml wash buffer (50 mM NaH2PO4.H2O, 10% glycerol (v/v) 300 mM NaCl, 1mM DTT, 20 ȝM PLP, pH 7.2). The cleared crude lysate was then loaded onto a 1 ml Strep-tactin column (Qiagen, Germantown, MD) attached to the ÄKTA purifier system (GE

Healthcare Bio-Sciences, Pittsburgh, PA) which was used to enrich for Strep-tagged proteins.

These can include, in the case of differently tagged HemT_C281P polypetides, Strep-tagged

HemT_C281P homodimers and Strep-tagged HemT_C281P and polyhistidine-tagged

HemT_C281P heterodimers. In the case of the co-expressed differentially tagged HemA and

HemT polypeptides, Strep-tagged HemA homodimers and Strep-tagged HemA and polyhistidine-tagged HemT_C281P heterodimers. The column was washed with 10 column volumes of wash buffer (50 mM NaH2PO4.H2O, 10% glycerol (v/v) 300 mM NaCl, 1mM DTT,

20 ȝM PLP, pH 7.2), to remove unbound protein. Bound protein was eluted with a 20 column volume linear gradient (0%-100%) of wash buffer mixed with elution buffer (50 mM

NaH2PO4.H2O, 10% glycerol (v/v), 300 mM NaCl, 1mM DTT, 20 ȝM PLP, 2.5 mM desthiobiotin) in 1 ml elution fractions. Absorbance was monitored at 280 nm. 38

The peak elution fractions from the Strep-tactin purification were loaded onto a His60

Nickel superflowTM resin and incubated for 1 h. Nonspecifically bound proteins that would include either Strep-tagged HemT_C281P homodimers or Strep-tagged HemA homodimers were removed by washing the column with 20 column volumes of wash buffer (20 mM tricine, 10% glycerol, 25 mM NaCl, 20 mM imidazole, 20 μM PLP, and 1 mM dithiothreitol (DTT), pH 7.2) which was enough to wash out the unbound protein. The bound proteins were then eluted using

10 column volumes of elution buffer (20 mM tricine, 10% glycerol, 25 mM NaCl, 350 mM imidazole, 20 μM PLP, and 1 mM DTT, pH 7.2) and concentrated using Amicon Ultra-15 centrifugal filter device.

Ultraviolet-visible spectroscopy of HemA and HemT. The ultraviolet-visible spectroscopic measurements of HemA and HemT were performed in the range of 250-900 nm using Hitachi U-2910 spectrophotometer to determine PLP interaction with the two proteins.

The concentration of HemA and HemT was 1.3 mg/ml in elution buffer (20 mM tricine, 10% glycerol, 25 mM NaCl, 350 mM imidazole, 20 μM PLP, and 1 mM DTT, pH 7.2).

Protein concentration and analysis. Protein concentrations were determined using the

Bio-Rad Protein Assay Reagent (Bio-Rad Laboratories, Hercules, CA) per the manufacturer's instructions. Bovine serum albumin (BSA) was used to generate the standard curves. Protein samples were mixed with sterile Milli-Q water (ultrapure water purified with Millipore Milli-Q lab water system) to a final volume of 800 μl. Then 200 μl of the BioRad reagent was added, and the solution was mixed by vortexing. The tubes were incubated at room temperature for 5 mins and the OD at 595 nm was determined using a Hitachi U-2910 spectrophotometer (Hitachi

High Technologies America Inc., Schaumburg, IL). 39

The purified proteins from the elution fractions were analyzed for purity by SDS-PAGE

(Sodium dodecyl sulfate polyacrylamide gel electrophoresis) using routine methods (57) and

12% gradient polyacrylamide tris-glycine precast gels (Invitrogen, Carlsbad, CA). Protein samples were mixed with equal volumes of 2 x concentrated Laemmli buffer (20% glycerol

(v/v), 2% SDS (w/v), 2.5% Bromophenol Blue (w/v), 5% ȕ-mercaptoethanol (v/v), Tris-base

0.125 mM; ref. 58), and then boiled for 5 min. Precision Plus ProteinTM Kaleidoscope TM standards (Bio-Rad Laboratories) were used to estimate the apparent molecular masses of proteins. ImageJ (59) was used to analyze gel images to estimate purity.

Immunoblot analysis. Following SDS-PAGE, samples were transferred onto a

BioTrace NT, nitrocellulose blotting membrane (Pall Corporation., Pensacola, FL) following a standard transfer protocol (60). For polyhistidine tag detection, membranes were washed two times with 15 ml TBS (0.15 M NaCl, 10 mM Tris-HCl, pH 7.5) for 10 min each before incubation with 15 ml blocking solution (3% BSA in TBS) for 1 h. The membranes were then washed two times with 20 ml TBSTT (0.5 M NaCl, 20 mM Tris-HCl, 0.2% Triton X-100, 0.05%

Tween-20, pH 7.5) for 10 min each, and once for 10 min with 15 ml TBS. This was followed by incubation with anti-polyhistidine monoclonal antibody (EMD Millipore Sigma, Billerica, MA) at the recommended dilution of 1:1000 for 1 h, after which the membranes were washed two times with 20 ml TBSTT and one time with 15 ml TBS, each for 10 min, and then they were incubated with a 1:30,000 dilution of goat anti-mouse IgG conjugated with alkaline phosphatase

(AP) in blocking solution as secondary antibody (Sigma-Aldrich, St. Louis, MO). The membranes were washed for 10 min each five times with 20 ml TBSTT and once with 10 ml AP buffer (100 mM Tris-HCl, 100 mM NaCl, 5 mM MgCl2, pH 9.5). Immunocomplexes were 40 detected by incubating the membrane with BCIP®/NBT Liquid Substrate System (Sigma-

Aldrich).

For Strep-tag detection, membranes were incubated with blocking solution (TBST with

5% BSA) for 1 hour at room temperature. They were then incubated with anti-Strep tag monoclonal antibody (Abcam, Cambridge, MA) as the primary antibody in blocking solution at the recommended dilution of 1:5000 overnight at 4°C. The membranes were washed three times with 10 ml TBST for 5 min each before incubation with a 1:30,000 dilution of goat anti-mouse

IgG conjugated with alkaline phosphatase (AP) as secondary antibody (Sigma-Aldrich) for 1 h at room temperature. The membrane was then washed three times again with TBST for 5 min each and one time with 10 ml AP buffer for 10 mins. Immunocomplexes were detected by incubating the membrane with BCIP®/NBT Liquid Substrate System.

InVision™ his-tag in-gel stain assay. Polyhistidine-tagged proteins were also detected using the InVision™ his-tag in-gel stain assay (Invitrogen Life Technologies, Carlsbad, CA) following the manufacturer’s protocol. After SDS-PAGE, the gel was fixed with 100 ml fixing solution (40 ml methanol, 10 ml glacial acetic acid and 50 ml Milli-Q water) for 1 h at room temperature. The gel was washed twice for 10 mins each using 100ml Milli-Q water and then incubated overnight with the ready-to-use solution of InVisionTM His-tag In-gel stain (Invitrogen

Life Technologies, Carlsbad, CA). The next day it was washed twice for 10 mins each with 100 ml 20 mM phosphate buffer pH 7.8 and imaged immediately using a UV transilluminator (UVP

LLC., Upland, CA) equipped with a Kodak EDAS 290 camera (Eastman Kodak Company,

Molecular Imaging Systems, Rochester, NY)).

Chemical cleavage of HemA. Chemical cleavage of HemA was performed using hydroxylamine-HCl (Flinn scientific Inc., Batavia, IL), which specifically targets asparagine- 41 glycine bonds (61), and so is specific for HemA as HemT lacks such a sequence. The cleavage reaction was initiated by adding 100 ul (22 uM) of protein to 1 ml of cleavage buffer (2 M hydroxylamine-HCl, 2 M guanidine-HCl (ICN Biomedicals, Inc. Aurora, OH), 0.2 M potassium carbonate; pH 9.0). The mixture was then incubated at 45°C for 4 h. Trichloroacetic acid was added at a final concentration of 5% (v/v) to stop the reaction and precipitate the protein. The solution was centrifuged for 10 min at 14000 rpm to pellet the protein, and the pellet was resuspended in Milli-Q water. Cleavage products were resolved by SDS-PAGE and, following staining with Coomassie Brilliant Blue R 250, they were evaluated for the presence of polypeptides corresponding to the apparent molecular masses of the cleavage products.

ALA synthase and 2-amino-3-ketobutyrate CoA ligase activity assays. ALA synthase activity was determined using the method of Burnham (62) as modified by Neidle and Kaplan

(43). The reaction mix was incubated at 370C for 10 mins. The total reaction volume was 102

μl, and contained 200 μM succinyl-CoA, 1.5 mM PLP, glycine mix pH 7.2 (120 mM glycine,

9.8 mM MgCl2, 20 mM tricine) and 50 μl protein sample. Reactions were stopped by the addition of 50 Pl 10% trichloroacetic acid (TCA) and centrifuged at 16,873 x g at room temperature for 2 min using an Eppendorf 5418 centrifuge and a FA-45-18-11 fixed-angle rotor to remove the precipitated protein. The amount of ALA formed during the reaction was determined by first converting it to a pyrrole compound (ALA-pyrrole; 2-methyl-3-acetyl-4-(3- propionic acid) pyrrole) through chemical condensation at 100°C with 160 mM acetylacetone in

200 Pl 1 M acetate buffer pH 4.7 for 15 min. A volume of 350 Pl of freshly prepared modified

Ehrlich’s reagent (6.7 mM p-dimethyaminobenzaldehyde, 42 ml glacial acetic acid, and 8 ml

70% perchloric acid) was then added to convert the ALA-pyrrole into a purple colored ALA- pyrrole complex that is formed when the ALA-pyrrole reacts with p- 42 dimethyaminobenzaldehyde. After a 20 min incubation at room temperature to fully allow for color development, the absorption at 556 nm was determined using the Hitachi U-2910 spectrophotometer. Enzyme activity was expressed as μmoles of ALA formed per hour, calculated using an extinction coefficient (H) for the ALA-pyrrole derivative of 68,000 M-1 cm-1

(62).

Kinetic parameters were determined by performing ALA synthase enzyme assays at varying concentrations of glycine (10 to 120 mM) or succinyl-CoA (1 to 20 μM). The data were then plotted to generate graphs of enzyme initial velocity versus substrate concentration. The Km and Vmax values were estimated by fitting the graphs to the Michaelis-Menten equation (Vo =

(Vmax * S)/ (Km + S)) using nonlinear regression with SigmaPlot 14 software (Systat Software,

San Jose, CA).

Assays of ALA synthases for 2-amino-3-ketobutyrate-CoA ligase activity were performed in the same way as ALA synthase activity assays, except acetyl-CoA replaced succinyl-CoA in the reaction mixtures.

Hemin inhibition assays. Inhibition by hemin was evaluated by assaying enzyme activity in the presence of different concentrations of hemin. A stock solution of 10 mM hemin was prepared by dissolving 0.01 moles in a solution of 0.01 M NaOH in 50% ethanol and 50%

Milli-Q water. The enzyme was pre-incubated with hemin for 5 mins, and then added to the reaction mix to start the reaction. Inhibition constants (Ki) were estimated by carrying out enzyme kinetic assays as described above in the presence of different concentrations of hemin with enzyme being added last to start the reaction. The Ki value was estimated with GraphPad

Prism 5 software using the equation for noncompetitive inhibition, Vo = Vmaxinh * S/ Km + S, in which Vmaxinh = Vmax/(1+I/Ki) and I is the hemin concentration. 43

Alkylation of cysteine residues in HemA and HemT. Alkylation of HemA and HemT cysteine residues was performed using the sulfhydryl reagents iodoacetamide (IA) and N- ethylmaleimide (NEM) as described by Bolt et al. (18). IA and NEM stock solutions of 100 mM each were prepared in 20 ȝM tricine buffer, pH 7.2. Either 2.2 ȝM of HemA or HemT protein samples were incubated with varying concentrations of IA and NEM in a final reaction volume of 500 μl at 37°C. At 5 min intervals 50 μl aliquots were removed over a total of 20 mins, and 5 mM dithiothreitol (DTT) was added to stop the alkylation reaction. The ALA synthase activity was then assayed. Protein incubated with tricine buffer only was used as a control and assumed to be 100% active.

Bioinformatic analysis and protein modelling. Predicted ALA synthase amino acid sequences were obtained from the National Center for Biotechnology Information (NCBI).

Multiple sequence alignments were generated using ClustalW version 2.0 (63) and alignments were displayed using GeneDoc Multiple Sequence Alignment Editor and Shading Utility (64).

Protein models were generated using SWISS-MODELLER (65-67). R. capsulatus

HemA crystal structures PDB IDs 2BWN (open conformation) and 2BWO (closed conformation) were used as templates to create the models. The coordinate files of the crystal structures were edited using notepad so as to have equal number of amino acid residues on each subunit in order for SWISS-MODELLER to accept the crystal structure PDB files as templates for modeling homodimers. Models were saved as PDB files and figures were generated using

PyMOL (The PyMOL Molecular Graphics System, version 1.8.0.0 Open-Source. Schrodinger,

LLC.).

Docking of substrates and cofactor into the active site of HemA and HemT protein models. Succinyl-CoA, glycine and PLP were docked to the protein models through docking by 44 superposition using PyMol. The homodimeric model structures of HemA and HemT were aligned with the crystal structures of R. capsulatus ALA synthase PDB IDs 2BWO and 2BWP such that the bound substrates and cofactors were aligned in the active sites of HemA or HemT.

The 2BWO structure has bound succinyl-CoA and PLP while the 2BWP structure has glycine that is bound to PLP. The substrates and cofactor were then extracted from their respective crystal structures using the PyMol extract object command which created new PDB files for succinyl-CoA, PLP and PLP bound to glycine. The extracted substrate and cofactor files together with the PDB file for either HemA or HemT were saved as one PDB file to yield HemA and HemT models with bound substrates and cofactor. 45

Table 2. Bacterial strains and plasmids.

Strain Relevant properties Reference

Escherichia coli

DH5Įphe F-, (ĳ80dlacZǻM15) recA1 endA1 hsdR17 52

supE44 thi-1 gyrA96 relA1 deoR ǻ(lacZYA-

argF)U169 phe::Tn10dCmR

JW0428-1 F-, ǻ(araD araB)567 ǻlacZ4787(::rrnB3) 55

ǻclpX724::kan rph-1 ǻ(rhaD-rhaB)568 hsdR514,

Ȝ-

BW25113 F-, ǻ(araD-araB)567 ǻlacZ4787(::rrnB-3) rph 1 55, 56

ǻ(rhaD-rhaB)568 hsdR514, Ȝ-

Plasmids

pIND4 IPTG-inducible expression vector, pMG160 53

replicon; KnR

pBBR1MCS2 Broad host range cloning vector; KnR 68

pIND4-2.4.9-hemA pIND4 derivative with synthetic 2.4.9-hemA gene This study

with a C-terminus polyhistidine tag and having

an NcoI recognition site at the 5’ end and HindIII

recognition site at the 3’ end; KnR

pIND4-2.4.9-hemT pIND4 derivative with synthetic 2.4.9-hemT gene This study

with a C-terminus polyhistidine tag and having

an NcoI recognition site at the 5’ end and HindIII

recognition site at the 3’ end; KnR

pIND4-2.4.1-hemA pIND4 derivative with synthetic 2.4.1-hemA gene This study

with a C-terminus polyhistidine tag and having 46

an NcoI recognition site at the 5’ end and HindIII

recognition site at the 3’ end; KnR

pIND4-2.4.1-hemT pIND4 derivative with synthetic 2.4.1-hemT gene This study

with a C-terminus polyhistidine tag and having

an NcoI recognition site at the 5’ end and HindIII

recognition site at the 3’ end; KnR

pIND4-2.4.3-hemA pIND4 derivative with synthetic 2.4.3-hemA gene This study

with a C-terminus polyhistidine tag and having

an NcoI recognition site at the 5’ end and HindIII

recognition site at the 3’ end; KnR

pIND4-2.4.9-hemT_C281P pIND4 derivative with synthetic 2.4.9- This study

hemT_C281P gene with a C-terminus

polyhistidine tag and having an NcoI recognition

site at the 5’ end and HindIII recognition site at

the 3’ end; KnR

pIND(2.4.9-hemA- pIND4 derivative with synthetic 2.4.9-hemA gene This study

R R Strep):::Sp/St with a C-terminus polyhistidine tag; Sp /St

pIND(2.4.9-hemT_C281P- pIND4 derivative with synthetic 2.4.9- This study

Strep):::SpR/StR hemT_C281P gene with a C-terminus polyhistidine tag; SpR/StR

pBBR(2.4.9-hemT_C281P- A pBBR1MCS2 derivative with synthetic 2.4.9- This study

His6) hemT_C281P gene with a C-terminus

polyhistidine tag; KnR 47

Results

Protein expression and purification. To investigate whether ALA synthases from R. sphaeroides have different enzymatic properties, and towards understanding the significance of

ALA synthase isoenzymes in this organism, enzymatic properties of ALA synthases from three wild type strains of R. sphaeroides were characterized and compared. The reason for examining the enzymes present in these different strains is because representation of the ALA synthase genes and their expression differs among them. In wild type strain 2.4.9, both hemA and hemT genes are present and expressed (45); in wild type strain 2.4.1, which has both hemA and hemT genes, hemT is transcriptionally silent (43, 45), in wild type strain 2.4.3, only the hemA gene is present.

The HemA ALA synthases in the three wild type strains 2.4.1, 2.4.3, and 2.4.9 are highly conserved, as shown in Figure 10. There are eight amino acid residue differences between 2.4.1

HemA and 2.4.3 HemA, seven amino acid residue differences between 2.4.9 HemA and 2.4.3

HemA and only one amino acid residue difference between 2.4.1 HemA and 2.4.9 HemA. The two HemT ALA synthases are also highly conserved with only four amino acid residue differences between them. HemA and HemT are also conserved relative to each other. In wild type strain 2.4.1 HemA is 55% identical and 71% similar to HemT (215/391 and 281/391); in wild type strain 2.4.9 HemA is 55% identical and 72% similar to 2.4.9 HemT (216/391 and

283/391). This degree of similarity suggests that their enzymatic properties could be similar.

2.4.1-HemA : MDYNLALDTALNRLHTEGRYRTFIDIERRKGAFPKAMWRKPDGSEKEITVWCGNDYLGMGQHPVVLGAMHEALDSTGAGSGGTRNISGTTLYHKRLEAELADLHG : 105 2.4.3-HemA : MDYNQALDTALNRLHTEGRYRTFIDIERRKGAFPKAMWRKPDGSEKEITVWCGNDYLGMGQHPVVLGAMHEALENTGAGSGGTRNISGTTLYHKRLEAELADLHG : 105 2.4.9-HemA : MDYNLALDTALNRLHTEGRYRTFIDIERRKGAFPKAMWRKPDGSEKEITVWCGNDYLGMGQHPVVLGAMHEALDSTGAGSGGTRNISGTTLYHKRLEAELADLHG : 105 R. caps-ALAS: MDYNLALDKAIQKLHDEGRYRTFIDIEREKGAFPKAQWNRPDGGKQDITVWCGNDYLGMGQHPVVLAAMHEALEAVGAGSGGTRNISGTTAYHRRLEAEIADLHG : 105 2.4.9-HemT : MEFSQHFQKLIDDMRLDGRYRTFAELERIAGEFPTALWHGPDGQARRVTVWCSNDYLGMGQNAEVLAAMHRSIDLSGAGTGGTRNISGTNRQHVALEAELADLHG : 105 2.4.1-HemT : MEFSQHFQKLIDDMRLDGRYRTFAELERIAGEFPTALWHGPDGQARRVTVWCSNDYLGMGQNAEVLAAMHRSIDLSGAGTGGTRNISGTNRQHVALEAELADLHG : 105

2.4.1-HemA : KEAALVFSSAYIANDATLSTLPQLIPGLVIVSDKLNHASMIEGIRRSGTEKHIFKHNDLDDLRRILTSIGKDRPILVAFESVYSMDGDFGRIEEICDIADEFGAL : 210 2.4.3-HemA : KEAALVFSSAYIANDATLSTLPQLIPGLVIVSDKLNHASMIEGIRRSGTEKHIFKHNDLDDLRRILSSIGKGRPILVAFESVYSMDGDFGQIKEICDIADEFGAL : 210 2.4.9-HemA : KEAALVFSSAYIANDATLSTLPQLIPGLVIVSDKLNHASMIEGIRRSGTEKHIFKHNDLDDLRRILTSIGKDRPILVAFESVYSMDGDFGRIKEICDIADEFGAL : 210 R. caps-ALAS: KEAALVFSSAYIANDATLSTLRVLFPGLIIYSDSLNHASMIEGIKRNAGPKRIFRHNDVAHLRELIAADDPAAPKLIAFESVYSMDGDFGPIKEICDIADEFGAL : 210 2.4.9-HemT : KESALIFTSGWISNLAALGTLGKVLPECAIFSDALNHNSMIEGIRRSGAERFIFRHNDPAHLDRLLSSVDPTRPKIVAFESVYSMDGDIAPIAEICDVAERHGAL : 210 2.4.1-HemT : KESALIFTSGWISNLAALGTLGKILPECAIFSDALNHNSMIEGIRRSGAERFIFHHNDPVHLDRLLSSVDPARPKIVAFESVYSMDGDIAPIAEICDVAERHGAL : 210

2.4.1-HemA : KYIDEVHAVGMYGPRGGGVAERDGLMDRIDIINGTLGKAYGVFGGYIAASSKMCDAVRSYAPGFIFSTSLPPVVAAGAAASVRHLKGD--VELREKHQTQARILK : 313 2.4.3-HemA : KYIDEVHAVGMYGPRGGGVAERDGLMDRIDIINGTLGKAYGVFGGYIAASAKMCDAVRSYAPGFIFSTSLPPVVAAGAAASVRHLKGD--VELREKHQTAAKILK : 313 2.4.9-HemA : KYIDEVHAVGMYGPRGGGVAERDGLMDRIDIINGTLGKAYGVFGGYIAASSKMCDAVRSYAPGFIFSTSLPPVVAAGAAASVRHLKGD--VELREKHQTQARILK : 313 R. caps-ALAS: TYIDEVHAVGMYGPRGAGVAERDGLMHRIDIFNGTLAKAYGVFGGYIAASAKMVDAVRSYAPGFIFSTSLPPAIAAGAQASIAFLKTAEGQKLRDAQQMHAKVLK : 315 2.4.9-HemT : TYLDEVHAVGLYGPRGGGISDRDGLADRVTIIEGTLAKAFGVMGGYVSGPSLLMDVIRSMSDSFIFTTSICPHLAAGALAAVRHVKAHP--DERRRQAENAVRLK : 313 2.4.1-HemT : TYLDEVHAVGLYGPRGGGISDRDGLADRVTIIEGTLAKAFGVMGGYVSGPSLLMDVIRSMSDSFIFTTSICPHLAAGALAAVRHVKAHP--DERRRQAENAVRLK : 313

2.4.1-HemA : MRLKGLGLPIIDHGSHIVPVHVGDPVHCKMISDMLLEHFGIYVQPINFPTVPRGTERLRFTPSPVHDSGMIDHLVKAMDVLWQHCALNRAEVVA : 407 2.4.3-HemA : MRLKGLGLPIIDHGSHIVPVHVGDPVHCKMISDMLLEHFGIYVQPINFPTVPRGTERLRFTPSPVHDSGMIDHLVKAMDVLWQHCALNRAEVVA : 407 2.4.9-HemA : MRLKGLGLPIIDHGSHIVPVHVGDPVHCKMISDMLLEHFGIYVQPINFPTVPRGTERLRFTPSPVHDSGMIDHLVKAMDVLWQHCALNRAEVVA : 407 R. caps-ALAS: MRLKALGMPIIDHGSHIVPVVIGDPVHTKAVSDMLLSDYGVYVQPINFPTVPRGTERLRFTPSPVHDLKQIDGLVHAMDLLWARCALNRAEASA : 409 2.4.9-HemT : VLLQKAGLPVLDTPSHILPVMVGEAHLCRSISEALLARHAIYVQPINYPTVARGQERLRLTPTPFHTTSHMEALVEALLAVGRDLGWAMSRRAA : 407 2.4.1-HemT : VLLQKAGLPVLDTPSHILPVMVGEAHLCRSISEALLARHAIYVQPINYPTVARGQERFRLTPTPFHTTSHMEALVEALLAVGRDLGWAMSRRAA : 407

Figure 10. Multiple sequence alignment of ALA synthases from R. sphaeroides and Rhodobacter capsulatus. Conserved residues are highlighted in grey. Residue differences among HemAs are highlighted in red while residue differences between HemTs are highlighted in cyan. Amino acid residues that form the subunit interface in R. capsulatus are highlighted in black (49). Differences between HemA and HemT in the subunit interface residues are highlighted in yellow. The sequences were retrieved from NCBI, aligned with ClustalW, and edited using GENEDOC (64). 49

All five ALA synthase genes were modified by the addition of sequences coding for a C- terminus polyhistidine (His6) tag. The recombinant proteins were overproduced in E. coli and purified using Ni-IDA affinity chromatography. The purified proteins migrated as single bands in polyacrylamide gels (PAGs) near the 50 kilodaltons (kD) marker (Figure 11), and were also detected in the same PAGs subjected to an in-gel visualization analysis for identification of polyhistidine-tagged proteins, using the InVision His-Tag stain (Invitrogen). Image analysis of the PAG using ImageJ software (59) indicated that the purity of the homogeneous proteins was greater than 90%.

Figure 11. SDS-PAGE and Invision His-Tag staining of proteins purified from E. coli expression strains using Ni-affinity chromatography. (A) Coomassie Brilliant Blue R 250- stained SDS polyacrylamide gel of purified protein samples. M: molecular mass marker, lane 1:

R. sphaeroides 2.4.1 HemA, lane 2: R. sphaeroides 2.4.3-HemA, lane 3: R. sphaeroides 2.4.9-

HemA, lane 4: R. sphaeroides 2.4.9-HemT, lane 5: R. sphaeroides 2.4.1-HemT, lane 6: R. sphaeroides 2.4.9-HemT_C281P. (B) Invision His-Tag staining of proteins in the same SDS gels as described in A. 50

The purified proteins were observed to be different in color with HemT having a light- yellow color and HemA having a more intense darker yellow-red color as shown in Figure 12A.

The yellowish color could be explained by the presence of bound pyridoxal 5’-phosphate (PLP) cofactor, which absorbs at approximately 430 nm. To examine this, ultraviolet-visible (UV-VIS) spectra of the two proteins were obtained as shown in Figure 12B. PLP bound to ALA synthase via an internal aldimine has a peak at 430 nm (69). Absorbance peaks for both proteins at 280 nm is predominantly due to absorbance by tryptophan and tyrosine. Purified HemT had an additional absorbance at 430 nm, which is indicative of PLP bound to a lysine residue via an internal aldimine. Purified HemA had additional absorbances at 356, 423, 540 and 580 nm, as shown in Fig 12B. These peaks are indicative of heme bound to protein (71) suggesting that purified HemA has heme bound. Whether HemT binds PLP poorly relative to HemA is not conclusive as the peak at 430 nm indicative of PLP binding for HemA is dominated by the peak contributed by absorbance presumably due to bound heme. Additionally, purified HemT was more unstable and came out of solution after a few minutes at room temperature.

Figure 12. Color and UV-visible absorbance spectra of purified HemA and HemT. (A) The purified HemA (1.3 mg/ml) and HemT (1.3 mg/ml) proteins are visibly different in color. HemA is more brightly colored compared to HemT, which has a pale-yellow color. (B) UV-visible spectral scans of HemA (1.3 mg/ml) and HemT (1.3 mg/ml) reveal the purified proteins have distinctive profiles. In addition to absorbances at 280 nm, HemT has an absorbance peak at 430 nm, indicating bound PLP; HemA has absorbance peaks at 356 nm, 540 nm and 580 nm. These absorbances correspond to those of hemoproteins (70), and suggests that purified HemA has bound heme.

Catalytic properties. The enzymatic properties of the purified enzymes were determined using the same conditions for all proteins. Their specific activities and kinetic parameters are summarized in Table 3. Thus, the Km and Vmax values for the enzymes were approximated using nonlinear regression and were used to calculate turnover number (kcat) and catalytic efficiency (kcat/Km) values. The specific activities of 2.4.1- and 2.4.9-HemAs were similar; HemA of 2.4.3 had approximately 2-fold higher specific activity. The specific activities 52 of the two HemT proteins were similar and were also comparable to those of the HemA enzymes. The HemT enzymes had glycine Kms that were slightly higher than those of HemAs.

Succinyl-CoA Kms and turnover number (kcat) values were similar for all the enzymes. The glycine catalytic efficiency (kcat/Km) for HemA ALA synthases was higher than that of HemTs.

Finally, the catalytic efficiencies for succinyl-CoA were the same with the exception of 2.4.1-

HemT, which was lower compared to the others. This could be due to the somewhat higher succinyl-CoA Km of that enzyme.

Collectively, the catalytic properties of the five ALA synthases indicate that there are no major differences in their enzymatic capacities. Therefore, other reasons that could explain the presence of ALA synthase isoenzymes need to be considered.

Table 3. Kinetic properties of R. sphaeroides HemA and HemT enzymes.

Property* 2.4.1-HemA 2.4.3-HemA 2.4.9-HemA 2.4.9-HemT 2.4.1-HemT

Specific activitya 68.0 ± 4.0 113.5 ± 17.7 62.9 ± 3.5 63.0 ± 6.3 54.7 ± 5.0

Gly Km (mM) 43.4 ± 6.7 26.4 ± 2.3 31.7 ± 2.8 52.5 ± 5.0 58.0 ± 6.1

S-CoA Km (μM) 17.6 ± 2.1 18.4 ± 5.6 14.6 ± 3.2 15.0 ± 3.3 28.7 ± 3.0

-1 kcat (h ) 3958.1 ± 407.2 4655.3 ± 188.5 3804.0 ± 304.3 3627.3 ± 400.6 3629.5 ± 150.3

Gly kcat/Km (mM/h) 91.8 176.3 120.0 69.1 62.6

S-CoA kcat/Km (μM/h) 224.9 253.0 260.6 241.8 126.5 aμmoles ALA/h/mg

*Values are the means of at least three replicates ± standard deviation.

Substrate preference. Members of the Į-oxoamine synthase family catalyze reactions that involve a small amino acid and acyl-CoA esters, as shown in Table 4. The ability of HemA or HemT to use substrates other than glycine and succinyl-CoA was examined. In this way, whether or not one or the other enzyme primarily functions to catalyze a reaction other than ALA formation could be evaluated.

Table 4. Members of the Į-oxoamine synthase family of fold type I PLP-dependent enzymes.

Enzyme Substrates Reference

5-aminolevulinic acid synthase L-glycine and succinyl-CoA 71

8-amino-7-oxononanoate synthase (AONS) L-alanine and pimeloyl-CoA 72

Serine palmitoyl transferase (SPT) L-serine and palmitoyl-CoA 73

2-amino-3-ketobutyrate-CoA ligase (KBL) L-glycine and acetyl-CoA 74

In ALA synthase the amino acid residue at position 83 (R. sphaeroides HemA/HemT numbering) plays an important role in recognition and specificity of the type of amino acid that is used as a substrate (49, 75, 76). Shoolingin-Jordan et al. (76) reported that a threonine at position 83 restricts the enzyme to glycine as the only amino acid that can be used as a substrate.

This has been confirmed in other reports (75). Analysis of the crystal structure of R. capsulatus

ALA synthase shows that amino acids bigger than glycine would interfere with the binding of succinyl-CoA (49). Both HemA and HemT have a threonine at position 83, which eliminates the possibility that they can catalyze the reactions catalyzed by AONS and SPT as those enzymes use amino acids bigger than glycine. However, KBL, like ALA synthase, uses glycine. Whether or not HemA or HemT in strain 2.4.9, where both enzymes are expressed, can utilize acetyl-CoA as a substrate was examined. The forward reaction catalyzed by KBL enzyme is the second in 55 the threonine utilization pathway, which is an alternate pathway for serine biosynthesis. If either

HemA or HemT showed a preference for acetyl-CoA as substrate, it would indicate that their primary role is to function as a KBL rather than an ALA synthase. While the presence of a bona fide kbl gene in R. sphaeroides makes this possibility less likely, the KBL activities of HemA and HemT were evaluated by substituting acetyl-CoA for succinyl-CoA in the reaction mixture.

At optimal concentrations of these substrates, specific activities of HemA and HemT are approximately 8-fold and 15-fold less, respectively, for acetyl-CoA compared to succinyl-CoA

(Table 5). Therefore, the primary role of both HemA and HemT is thought to be synthesis of

ALA.

Table 5. Specific activities of R. sphaeroides 2.4.9 HemA and HemT using different substrates.

Specific activity (μmoles product/h/mg)* Enzyme with succinyl-CoA (200 μM) with acetyl-CoA (200 μM)

2.4.9-HemA 62.9 ± 3.5 8.2 ± 1.0

2.4.9-HemT 63.0 ± 6.3 4.2 ± 1.7

* Values are the means of at least three replicates ± standard deviation.

Feedback inhibition by hemin. Hemin, an end product of tetrapyrrole biosynthesis, has been reported to regulate ALA synthase, the first enzyme in the pathway. The human ALA synthase genes encoding ALAS1 and ALAS2 are differentially regulated by heme at various levels. Heme inhibits the ALAS1 gene at the level of transcription (39), translation (38) and import into the mitochondria (40, 77). Transcription of the ALAS2 gene has been reported to be induced by heme in differentiating cells (37), but import into the mitochondria is not inhibited by heme (77). Whether heme differentially regulates ALAS1 and ALAS2 activities is not known 56 yet. However, a recent study has reported that heme competitively inhibits ALAS1 activity (78).

In that study, heme and protoporphyrin IX were reported to bind to the hydrophobic binding pocket of the adenylyl moiety of succinyl-CoA and a 750 nM concentration of heme caused a loss of approximately 70% of ALAS1 activity. Protoporphyrin IX was reported to be a more potent inhibitor with complete loss of activity with 225 nM protoporphyrin IX. These results are contradictory to previous reports of heme inhibition of R. sphaeroides ALA synthase (13), where heme was reported to be a noncompetitive inhibitor and protoporphyrin IX had no inhibitory effect. Further, in a related phototrophic bacterium, Rhodopseudomonas palustris, which also has ALA synthase isoenzymes, Zhang et al. (25) reported differences in hemin inhibition of the two enzymes. Addition of 2 μM hemin in the reaction mix led to a loss of 40% of activity of both enzymes. Higher concentrations of hemin continued to further reduce activity of one, but there was no change in activity of the other enzyme at concentrations up to 10 μM hemin (25).

The significance of differential inhibition of these Rps. palustris enzymes by hemin or how this might describe their roles in that bacterium is not known because the study was focused on characterizing the isoenzymes for application in ALA production, and so there is no information as to relative expression of the corresponding genes. Moreover, it is important to note that each of the Rps. palustris enzymes are no more similar to HemA than they are to HemT in amino acid sequence, and so provide no information as to how hemin would affect activity of the R. sphaeroides enzymes.

Previous studies have reported that vitamin B12 and bacteriochlorophyll a have no inhibitory effect, while hemin is a noncompetitive inhibitor of ALA synthases of R. sphaeroides

(13, 14). However, those studies predated the discovery of ALA synthase isoenzymes in R. sphaeroides, and because the protein purification protocols used followed activity, rather than 57 affinity purification of a target protein, whether the purified product was HemA, HemT or a combination of the two proteins is not known. Here, the inhibitory effects of vitamin B12 on R. sphaeroides 2.4.9 HemA and HemT was investigated. As shown on Table 6, vitamin B12 was not inhibitory toward either HemA or HemT. Therefore, they are consistent with the previous observation that this cofactor does not affect ALA synthase activity.

Table 6. Specific activities of R. sphaeroides 2.4.9 HemA and HemT incubated with different concentrations of vitamin B12.

Specific activity (μmoles ALA/h/mg)*

Vitamin B12 (ȝM) HemA HemT

0 61.7 ± 5.3 61.4 ± 4.9

0.05 61.7 ± 3.6 59.2 ± 0.9

0.1 67.4 ± 0.9 58.9 ± 3.1

1 66.2 ± 2.7 59.5 ± 4.0

2 63.6 ± 0.9 57.6 ± 0.4

5 73.7 ± 2.7 61.4 ± 1.3

10 68.7 ± 0.9 63.0 ± 2.7

* Values are the means of at least three replicates ± standard deviation.

The profiles of hemin inhibition of the affinity purified R. sphaeroides proteins are shown in Figure 13. The three HemA ALA synthase enzymes had similar inhibition profiles, as did the two HemT enzymes. However, the results indicate that HemA enzymes are more inhibited by hemin than the HemT enzymes. 58

Figure 13. Percentage activity remaining of HemA and HemT enzymes in the presence of different concentrations of hemin. The enzymes were preincubated with different concentrations of hemin for 5 mins and then added to a reaction mix containing glycine, succinyl-CoA and PLP to start the reaction. Values are the means of at least three replicates with standard deviations indicated by error bars.

The inhibition of HemA and HemT by hemin was examined further as a function of substrate concentration to determine whether it competes with glycine, succinyl-CoA or both substrates for binding. Comparisons of initial velocity against substrate concentration in the presence of increasing hemin concentrations are indicative of noncompetitive inhibition for both substrates for HemA and HemT (Figure 14). This type of inhibition was also reported by

Burnham and Lascelles for purified R. sphaeroides ALA synthase preparations whose protein composition is not known (13). 59

A B 0 PM Hemin 0 PM Hemin 0.5 PM Hemin 0.20 0.5 PM Hemin 0.25 1 PM Hemin 1 PM Hemin 0.20 0.15 moles ALA/h) moles ALA/h) 0.15 P 0.10 P 0.10 0.05 0.05

0.00 0.00 Initial velocity( 0 5 10 15 20 Initial velocity ( 0 50 100 150 succinyl-CoA (PM) Glycine (mM)

C D 0 PM Hemin 0 PM Hemin 2.5 PM Hemin 2.5 PM Hemin 0.4 0.4 5 PM Hemin 5 PM Hemin

0.3 0.3 moles ALA/h) moles ALA/h) P 0.2 P 0.2

0.1 0.1

0.0 0.0 Initial velocity ( 0 5 10 15 20 Initial velocity ( 0 50 100 150 succinyl-CoA (PM) Glycine (mM)

Figure 14. Characterization of the inhibition mechanism of HemA and HemT by hemin. (A) and (B), substrate versus initial velocity graphs in the presence of different hemin concentrations for HemA. (C) and (D), substrate versus initial velocity graphs in the presence of different hemin concentrations for HemT. Values are the means of at least three replicates with standard deviations indicated by error bars.

To determine the apparent inhibition constant (Ki) values for 2.4.9-HemA and 2.4.9-

HemT, enzyme kinetic assays were performed in the presence of different concentrations of 60

hemin. The Ki value for each was estimated with GraphPad Prism 5 software using the equation for noncompetitive inhibition, Vo = Vmaxinh * S/ Km + S. Where Vmaxinh = Vmax/(1+I/Ki) and I is hemin concentration. The apparent Ki value of 2.4.9-HemA was calculated to be 0.6 μM while that for 2.4.9-HemT was 8 μM, which represents a 13-fold difference. Thus, while their kinetic properties are very similar, HemA is more sensitive to hemin inhibition than HemT.

Sensitivity of HemA and HemT to oxidation. HemA and HemT each have five cysteines, of which three are invariant among all known ALA synthases. Cysteine residues are thought to play a role in the enzymatic activity of the ALA synthases (18), as reduced cysteine residues have been shown to increase the enzymatic activity of these enzymes (19). Further, pre- incubation of R. sphaeroides ALA synthase with succinyl-CoA has been shown to protect the enzyme from the inhibitory effects of the sulfhydryl reagents (18). As there are no cysteine residues that are directly involved in succinyl-CoA binding, the ability of succinyl-CoA to protect against alkylation could be interpreted to mean that reduced cysteine residues play a role in changes from open to closed conformations, which are known to be triggered by the binding of succinyl-CoA (49, 79, 80).

Given that the distribution of cysteines differs between HemA and HemT, the effect of modifying sulfhydryl reagents on HemA and HemT was examined to determine whether these sequence differences give rise to different properties. HemT was more sensitive to alkylation by iodoacetamide (IA) and N-ethylmaleimide (NEM) relative to HemA (Figure 15), suggesting that

HemT might be more sensitive to oxidation compared to HemA. If true, then it would be important to adequately protect this enzyme from inactivation during purification. 61

Figure 15. Time-dependent inhibitory effect of sulfhydryl modifying agents on HemA and

HemT. The enzymes were incubated with either (A) 1.5 mM iodoacetamide (IA) or (B) 0.05 mM N-ethylmaleimide (NEM) at 37oC in a final volume of 500 μl. Aliquots were taken every 5 min and assayed for enzyme activity. Values are the means of four replicates with standard deviations indicated by error bars.

When ȕ-mercaptoethanol was replaced with the stronger reducing agent dithiothreitol

(DTT) in the buffers used during purification, an approximately 6.7-fold higher specific activity was measured for HemT (Table 7). By contrast, the specific activity of HemA protein purified in the presence of DTT relative to that purified in the presence of ȕ-mercaptoethanol differed less than 1.5-fold (Table 7).

Table 7. Specific activities of R. sphaeroides 2.4.9 ALA synthases purified in the presence of different reducing agents.

Specific activity (μmoles ALA/h/ml/mg)*

Enzyme Purified with buffers having Purified with buffers having

5 mM ȕ-mercaptoethanol 1 mM DTT

2.4.9-HemT 9.6 ± 0.3 63.0 ± 6.3

2.4.9-HemA 62.9 ± 3.5 78.9 ± 8.2

*Values are the means of at least three replicates ± standard deviation.

To further investigate the sensitivity of HemT to oxidation, the location of the two cysteine residues, the proximities of cysteines 131 and 281 (R. sphaeroides HemT numbering) to each other and to other cysteines within the quaternary structure of HemT was investigated

(Figure 16) by protein modelling. Cysteine 281 was found to be located on the interface between the two subunits, and these cysteines are in close proximity; the models indicate they are separated by a distance of 8.9 Angstroms (Å). Analysis of the predicted structure of HemT using

PDBePISA (81) revealed that cysteine 281 in HemT could be involved in hydrogen bonding with valine 252 and methionine 253 of the other subunit (R. sphaeroides HemT numbering). Cysteine

281 is also located near the N terminus of an Į-helix. For Į-helices having a cysteine on their N terminus in redox responsive proteins, the Į-helix is reported to be important in maintaining the reduced state of the cysteine residues by helping lower the pKa (82, 83). All of this suggests that cysteine 281 might be responsible for HemT redox sensitivity.

Figure 16. Predicted structure of R. sphaeroides 2.4.9-HemT homodimer. The two polypeptides are colored differently (cyan and dark grey). Bound succinyl-CoA and PLP are shown as sticks in the two active sites. Cysteines are shown as blue sticks. The distance between the two cysteine 281 residues is shown as a black dotted line. The predicted hydrogen bonds of cysteine

281with valine 252 and methionine 253 are shown as dotted red lines. The figure was generated using PyMOL (The PyMOL Molecular Graphics System, Version 1.8.0.0 Schrödinger, LLC.).

Alignment of ALA synthases from different organisms showed that a proline is conserved among ALA synthases at the 281 position. Only HemTs have a cysteine residue at position 281, as shown in Figure 17.

281 Rhodobacter capsulatus : FNGTLAKAYGVFGGYIAASAKMVDAVRSYAPGFIFSTSLPPAIAAGAQASIAF 294 Rhodobacter vinaykumarii : FNGTLGKAYGVFGGYIAASAKMVDAVRSYAKGFIFSTSLPPAIAAGARASIAF 294 Pseudoruegeria sp.SK021 : VNGTLGKSYGVMGGYIATSHRMVDAIRSYAPGFIFTTSLPPAIAAGAAASIRH 294 Rhodobacter sphaeroides KD131 : INGTLGKAYGVFGGYIAASSKMCDAVRSYAPGFIFSTSLPPVVAAGAAASVRH 294 Rhodobacter sphaeroides 249 HemA : INGTLGKAYGVFGGYIAASSKMCDAVRSYAPGFIFSTSLPPVVAAGAAASVRH 294 Rhodobacter sphaeroides HemA : INGTLGKAYGVFGGYIAASSKMCDAVRSYAPGFIFSTSLPPVVAAGAAASVRH 294 Rhodobacter sphaeroides 2.4.3 HemA : INGTLGKAYGVFGGYIAASAKMCDAVRSYAPGFIFSTSLPPVVAAGAAASVRH 294 Rhodopseudomonas palustris HEMA : IEGTLAKAFGVVGGYIAGSSAVCDFVRSFASGFIFSTSPPPAVAAGALASIRH 293 Rhodobacter sphaeroides 2.4.1 HemT : IEGTLAKAFGVMGGYVSGPSLLMDVIRSMSDSFIFTTSICPHLAAGALAAVRH 294 Rhodobacter sphaeroides 2.4.9 HemT : IEGTLAKAFGVMGGYVSGPSLLMDVIRSMSDSFIFTTSICPHLAAGALAAVRH 294 Rhodobacter sphaeroides KD131 HemT : IEGTLAKAFGVMGGYVSGPSLLMDVIRSMSDSFIFTTSICPHLAAGALAAVRH 294 Rhodopseudomonas palustris HemO : MEATLAKAFGCLGGYIAGKAEVIDAVRSYAPGFIFTTALPPPICAAATAAIKH 293 Homo sapiens ALAS2 : ISGTLGKAFGCVGGYIASTRDLVDMVRSYAAGFIFTTSLPPMVLSGALESVRL 295 Homo sapiens ALAS1 : ISGTLGKAFGCVGGYIASTSSLIDTVRSYAAGFIFTTSLPPMLLAGALESVRI 295 Lachancea thermotolerans CBS 6340 : ITGTLGKSFGTVGGYVAGSLRLIDWLRSYAPGFIFTTSLPPSVMAGAAEAIRY 398 Zygosaccharomyces rouxii CBS 732 : ITGTLGKSFGTVGGYVAASQRLVDWLRSYAPGFIFTTSLPPSVMAGATEAIRY 392 Eremothecium sinecaudum : ITGTLGKSFGTVGGYIAASEKLIDWVRSYAPGFIFTTSLPPAVLAGAAEAIRY 388 Saccharomyces cerevisiae S288C : ITGTLGKSFGSVGGYVAASRKLIDWFRSFAPGFIFTTTLPPSVMAGATAAIRY 383 Streptomyces aizunensis NRRL : VMGTLAKGYGTVGGYIAGPAALVDAVRTLSRAFVFTTSLPPAVAAGALEAVRY 294 Streptomyces nodosus spp asukaensis: VMGTLAKGFGTAGGYIAGPAALIDAVRNFSRGFIFTTSIPPATAAGALAAVQH 294 Figure 17. Multiple sequence alignment of ALA synthases in the region of HemT cysteine 281

(highlighted in red). Valine 252 and methionine 253 that are predicted to form hydrogen bonds with cysteine 281 are highlighted in cyan. The amino acid sequences were retrieved from NCBI and aligned using CLUSTAL W (9).

To determine whether the cysteine at position 281 in HemT is responsible for the sensitivity of the enzyme to oxidation, a mutant protein having a proline substitution was examined. The specific enzyme activity of HemT_C281P mutant was approximately 5.1-fold that of the wild type when enzyme activity was performed in the presence of ȕ-mercaptoethanol

(Table 8). In the presence of DTT, the specific activity of the C281P mutant was approximately

1.3-fold higher than in the presence of ȕ-mercaptoethanol, the same difference as was found for

HemA purified in the presence of the two reducing agents. These results indicate that cysteine

281 could be responsible for oxidation sensitivity of HemT. As shown in Table 8, the kinetic parameters of HemT_C281P protein were the same as those of wild type HemT and the mutant had a similar hemin inhibition profile as the wild type as well (Figure 18). 65

Table 8. Specific activities and kinetic properties of R. sphaeroides 2.4.9 HemT wild type and HemT_C281P mutant ALA synthases purified in

the presence of different reducing agents.

Specific activitya Catalytic properties

Gly S-CoA 5 mM ȕ-me 1 mM DTT kcat/Km kcat/Km Gly S-CoA -1 Km (mM) Km (μM) kcat (h ) (mM/h) (μM/h)

2.4.9-HemT_WT 15.4 ± 0.4 79.1 ± 0.0 52.5 ± 5.0 15.0 ± 3.3 3627.3 ± 400.6 69.1 241.8

2.4.9-HemT_C281P 51.0 ± 2.7 64.3 ± 3.1 46.8 ± 3.6 26.1 ± 5.0 3328.0 ± 98.9 71.1 127.5

aμmoles ALA/h/mg.

Values are the means of at least three replicates ± standard deviation. 66

Figure 18. Percentage activity remaining of R. sphaeroides 2.4.9-HemT wild type and

HemT_C281P mutant enzymes in the presence of different concentrations of hemin. The enzymes were added to a reaction mix containing glycine, succinyl-CoA, PLP and different concentrations of hemin to start the reaction. Values are the means of at least three replicates with standard deviations indicated by error bars.

Role of ClpX in activating HemA and HemT. ClpX is part of a two-component ATP- dependent AAA+ protease system that is present in all organisms (84). ClpX requires ATP for its activity and forms the regulatory element of ClpXP, which recognizes and unfolds target proteins and then delivers them to the protease element ClpP for degradation (85, 86). In

Saccharomyces cerevisiae, a new role for ClpX has been reported, whereby it is involved in the activation of ALA synthase (87). It partially unfolds apo-ALA synthase, which facilitates the binding of PLP into the active site pocket thereby forming ALA synthase holoenzyme (87). The 67

ALA synthase activation activity of ClpX is thought to be conserved in eukaryotes as it was also reported to be important in activation of ALAS1 and ALAS2 in the zebra fish, Danio rerio (86).

Whether ALA synthase activation by ClpX takes place in Bacteria is not known.

ClpX proteins of S. cerevisiae and D. rerio are 39% and 54% identical to R. sphaeroides

2.4.9 ClpX, respectively (Figure 19). This similarity suggests that the ALA synthase activation activity of ClpX could also be conserved in R. sphaeroides. Furthermore, purified HemT may bind PLP poorly compared to HemA (Figure 12), which could mean ClpX mediates PLP binding

HemT. To investigate whether ClpX is involved in such activation of HemT, HemT was overexpressed in an E. coli ClpX minus strain (JW0428-1) and also the ClpX wild type E. coli parent strain (BW25113). The activities of the proteins purified from the two expression systems in the absence of PLP were compared to the activities of proteins purified from both strains in the presence of PLP.

Substrate recognition and specificity of ClpX is governed by three different loops that are present on the central pore of the ClpX ring. These loops are named pore 1 loop, pore 2 loop and

RKH loop which has a conserved RKH amino acid sequence motif (86, 88). R. sphaeroides

ClpX is 71% identical to E. coli ClpX, and the pore 1, pore 2 and RKH loop regions (88) are conserved (Fig. 19), suggesting that E. coli ClpX would recognize HemT as a substrate. 68

Rsph-2.4.9 : ------MANNTGSDSKNTLYCSFCGKSQHEVRKLIAG : 31 E. coli : ------MTDKRKDGSGKLLYCSFCGKSQHEVRKLIAG : 31 Zebra fish : MSCTCAIAARKFINSAHKGISGSRVQLLSLSRPGTREVRLARRVPVRSFSETAVYYASKDGVKDGDGGKKSVGEGSGKRTSSGNSGKGGSQLRCPKCGDPCTHVETFVSS : 110 Yeast : ------MLKSASQNFFRAYSSRIGRYAATASGKLAQ : 30

Rsph-2.4.9 : PTVFICDECVELCMDIIREETKSTG------LKSADGVPTPREICKVLDDYVIGQMHAKRVLSVAVHNHYKRLN------: 99 E. coli : PSVYICDECVDLCNDIIREEIKEVAP------HRERSALPTPHEIRNHLDDYVIGQEQAKKVLAVAVYNHYKRLRN------: 101 Zebra fish : TRFVKCEKCHHFFVVLSETDTKKSLSKDPESAAEAVKLAFQQKPPPPPKKIFAYLDKYVVGQDHAKKVLSVAVYNHYKRIYNNMPAGSRQQQVEVEKQASLTPRELLQIA : 220 Yeast : S------RLSNIPTPKALKKFLDEYIVGQEIGKKVLSVAVYNHYLRINDKQKKGELQRQRELMEREKIADDRDEPIF : 101

pore 1 loop | | Rsph-2.4.9 : ------HSSKTDIELSKSNILLIGPTGCGKTLLAQTLARILDVPFTMADATTLTEAGYVGEDVENIILKLLQASEYNVERAQRGI : 178 E. coli : ------GDTSNGVELGKSNILLIGPTGSGKTLLAETLARLLDVPFTMADATTLTEAGYVGEDVENIIQKLLQKCDYDVQKAQRGI : 180 Zebra fish : GISPHGNALGASMQQQLNQQTPPEKRGGEVLDSTHTDIKLEKSNIVLLGPTGSGKTLLAQTLAKCLDVPFAICDCTTLTQAGYVGEDIESVIAKLLQDANYVIEKAQQGI : 330 Yeast : SGNSESKAGWRNLQRQFNLAG------REVDEDLELSKSNVLVVGPSGSGKTLLATTLAKILNVPIAITDCTQLTQAGYIGEDVEVCIERLLVNAEFDVARAEKGI : 201

pore 2 loop RKH loop | | | | Rsph-2.4.9 : VYIDEVDKITRKSDNPSITRDVSGEGVQQALLKIMEGTVASVPPQGGRKHPQQ------EFLQVDTTNILFICGGAFAGLEKIIAQRGK------: 261 E. coli : VYIDEIDKISRKSDNPSITRDVSGEGVQQALLKLIEGTVAAVPPQGGRKHPQQ------EFLQVDTSKILFICGGAFAGLDKVISHRVET------: 264 Zebra fish : VFLDEVDKIG-SVPGIHQLRDVGGEGVQQGLLKLLEGTIVNVPEKNTRKLRG------ETVQVDTTNILFVASGAFNGLDRIISRRKNEKYLGFGTPSNMGKGR : 427 Yeast : IVLDEIDKLAKPAAS-IGTKDVSGEGVQQSLLKIIEGHKVEITVKRPVKHDIDGQKNQTTTKKDEVFVVDTSNILFMIMGAFVGLDKHIVKRIEDMKKIQKAGESVESSN : 310

Rsph-2.4.9 : ------GSGIGFGAEVKDP-DARGVGE---LFKELEPEDLLKFGLIPEFVGRLPVIATLTDLDEAALVTILTEPKNALVKQYQRLFEIEGVKLTFTADALTAIAKRAIKR : 361 E. coli : ------GSGIGFGATVKAKSDKASEGE---LLAQVEPEDLIKFGLIPEFIGRLPVVATLNELSEEALIQILKEPKNALTKQYQALFNLEGVDLEFRDEALDAIAKKAMAR : 365 Zebra fish : RAAAAADLANTTGGEVDAVAEIEEKDR---LLKHVEARDLIEFGMIPEFVGRLPVVVPLHSLDEETLVRILTEPRNAVVPQYQALFSMDKCELNMTPDALRAIARLALER : 534 Yeast : SKEVEKERAKKFRFSNTLEQVELDNGKKVCALDLTTPTDLVSFGLIPELIGRVPIITALQPLQRDDLFHILKEPKNALLDQYEYIFKQFGVRLCVTQKALKKVAQFALKE : 420

Rsph-2.4.9 : KTGARGLRSIMEDILLDTMFELPGLEGVEEVVVNEEAVN------SGAKPLLIYTEVTKKKDATAS------: 421 E. coli : KTGARGLRSIVEAALLDTMYDLPSMEDVEKVVIDESVID------GQSKPLLIYGKPEAQQASGE------: 424 Zebra fish : KTGARGLRSIMEKLLLDPMFEVP-HSDIVSVDVSKDVVQ------GKAPPQYIRAAAKESSEEEYDSGIEEENWTRQADAAKN----- : 610 Yeast : GTGARGLRGIMERLLLNVNYDCP-GSNIAYVLIDEATVDSLQETEHSLASQVDVKYYSGDEKDSLIRDVSEEDKKLGVMLEKELGHSANIHTPTIPKRSLT : 520

Figure 19. Multiple sequence alignment of ClpX amino acid sequences from R. sphaeroides ATCC 17209 (Rsph-2.4.9), E. coli, D. rerio (zebra fish) and S. cerevisiae (yeast). The pore 1, 2 and RKH loops that are involved in substrate specificity (86, 88) are indicated on top of the alignment. 69

If ClpX is involved in PLP loading of HemT, the enzyme purified from the wild type strain should have higher activity than the one purified from the clpX null mutant strain. As shown in Table 9, the specific activities of enzymes purified from both strains in the presence versus the absence of PLP differed less than 1.5-fold. Apparently, the protein purified from either strain had PLP bound since it was active, even in the absence of PLP in the reaction mix.

That activity improved when PLP was present in the mixture suggests even purified protein can bind PLP; even protein that had been purified from the clpX mutant in the absence of PLP had this capacity, based on the specific activity. Collectively, these results indicate that ClpX is not required for PLP loading of HemT. (Note that the more oxygen-stable C281P mutant form of

HemT was examined here.)

Table 9. Specific activities of R. sphaeroides 2.4.9 HemT_C281P mutant ALA synthases purified from clpX minus and clpX plus E. coli strains.

Specific activity of enzyme purified Specific activity of enzyme purified

from clpX minus E. colia from clpX plus E. colia

No PLP in the PLP in the No PLP in the PLP in the

reaction mix reaction mix reaction mix reaction mix

PLP in the 40.3 ± 1.4 61.8 ± 10.5 37.7 ± 1.8 62.5 ± 8.3 purification buffers

No PLP in the 21.3 ± 7.7 67.1 ± 20.5 21.0 ± 1.9 67.1 ± 11.4 purification buffers a μmoles ALA/h/mg

Values are means of at least three replicates ± standard deviation.

Isolation of HemA and HemT heterodimers. The active site of ALA synthase is located on the subunit interface, with amino acid residues from both monomers contributing to the active site (48, 49). The process of dimerization of ALA synthase is not yet described.

However, the importance of the subunit interface in dimerization of Į-oxoamine synthases has been examined by Turbeville et al. (89). They constructed genes encoding single-chain chimeric polypeptides of ALA synthase and AONS, and found that the products did not form a heterodimer; rather, the two linked monomers formed two independent homodimers of ALAS and AONS. This indicated that, although the two proteins are highly similar in terms of structure, there are key residue differences in the subunit interface that are important for dimerization such that homodimerization of ALAS and AONS is the predominant, possibly exclusive, outcome.

Analysis of the crystal structures of R. capsulatus ALA synthases (PDB IDs 2BWN and

2BWO) using PDBePISA (81) gives a complex formation significance score (CSS) of 0.96 which is an indicator of the importance of the subunit interface in dimerization. The score ranges between 0 (not essential) and 1 (essential). Therefore, the amino acid residues that are important for dimerization could be located in the subunit interface. Comparisons of the residues present on the subunit interface of HemA and HemT (Figure 10) indicate that most residues are conserved between HemA and HemT, possibly including those most critical to dimerization.

Since in strain 2.4.9 both HemA and HemT polypeptides are thought to be present in the cell at the same time, based upon hemA and hemT transcription profiles (45), it raises the question as to whether the polypeptides can form a heterodimeric enzyme having properties that differ from each homodimeric enzyme. 71

Heterodimer formation consisting of HemA and HemT polypeptides was evaluated by coexpressing Strep-tagged HemA and His6-tagged HemT_C281P polypeptides in E. coli. The procedure for the isolation of heterodimers was as follows: crude E. coli lysate that could contain both Strep-tagged HemA and His6-tagged HemT homodimers, and also Strep-tagged HemA-

His6-tagged HemT heterodimers, was loaded onto a Strep-tactin column. Bound protein (a possible mixture of Strep-tagged HemA homodimers and Strep-tagged HemA-His6-tagged

HemT heterodimers) was eluted in a linear gradient of desthiobiotin (0 to 2.5 mM). The peak eluate from the Strep-tactin column was then incubated with Nickel-IDA resin to resolve Strep- tagged HemA-His6-tagged HemT heterodimers that would bind to the resin from Strep-tagged

HemA homodimers that would not bind. Unbound protein was collected in the flow through and wash, and bound protein was eluted using imidazole. As a control that the affinity tags were not affecting heterodimerization, Strep- and His6-tagged HemT_C281P was coexpressed and purified following the same protocol. The protein yields for the elution fractions that were resolved for heterodimers was approximately 0.2% of the total affinity purified protein from the coexpression of differentially tagged HemA and HemT_281P as shown in Table 10. For the coexpression of differentially tagged HemT_C281P, the heterodimer protein yield was 0.4 % that of the total affinity purified protein. The low protein yield of the differentially tagged proteins could be due to the influence of the affinity tags or it could be other factors that are involved in the folding process of the polypeptides and dimerization process.

Table 10. Protein yields from the coexpression system of differentially tagged HemA and HemT

polypeptides.

Differentially tagged HemA and HemT coexpression. Protein yield/liter

Strep-tagged HemA 2.6 mg

His6-tagged HemT 0.8 mg

HemA (Strep-tag)/HemT (His6-tag) heterodimer 7.5 ȝg

Differentially tagged HemT_C281P coexpression

His6-tagged HemT_C281P 0.9 mg

Strep-tagged HemT_C281P 0.5 mg

HemT_C281P (His6-tag)/HemT_C281P (Strep-tag) 6.0 ȝg

To confirm the presence of Strep-tagged and His6-tagged proteins in the resolved

heterodimers, anti-Strep-tag and anti-His6-tag monoclonal antibodies were used to detect the

affinity tags. Surprisingly, while the anti-Strep tag antibody was observed to be specific for

Strep-tagged protein (Figure 20), the anti-His6 monoclonal antibody was not specific for the

His6-tagged proteins, and detected both Strep-tagged HemA and His6-tagged HemT-C281P

polypeptides (Figure 20). 73

Figure 20. SDS-PAGE and Western blot analysis of differentially tagged proteins purified from

E. coli overexpressing either His6-tagged HemT_C281P or Strep-tagged HemA. Proteins were purified using Nickel and Strep-tactin affinity chromatography, respectively. (A) Coomassie

Brilliant Blue R 250-stained SDS polyacrylamide gels of purified protein samples. Lane 1: His6- tagged HemT_C281P, lane 2: Strep-tagged HemA, M: marker, lane 3: Strep-tagged HemA, lane

4: His6-tagged HemT_C281P. (B) Immunoblots of membranes from transfer of proteins from the same PAGs as in (A) that were probed with anti-Strep-tag antibody (lanes 1 and 2) and anti-

His6-tag antibody (lanes 3 and 4).

Immunoblot analysis of the protein samples purified from the coexpression system revealed that, following the sequential steps of affinity chromatography, only Strep-tagged protein could be detected (Figure 21, lane 1). However, both affinity tags could be detected in the eluate of Strep-tagged HemA homodimer (Figure 21, lane 2). This could be due to non- specificity of the anti-His6 antibody or the possibility that the differentially tagged heterodimer was not binding to the Nickel column and was being lost in the wash together with the Strep- tagged HemA homodimer. Only the His6-tag could be detected in the eluate resolved for the

His6-tagged HemT_C281P homodimer (Figure 21 lane 3). Immunoblot analysis of the protein samples purified from the coexpression system of differentially tagged HemT_C281P revealed 74 that only the Strep tag could be detected in the eluate resolved for the heterodimer (lane 5). In lane 6 which contained sample resolved for Strep-tagged HemT_C281P homodimer, only the

Strep tag could be detected. In lane 7 which contained sample resolved for His6-tagged

HemT_C281P homodimer, only the His6-tag could be detected. This indicates that, heterodimiers of differentially tagged HemT_C281P were not being purified from the expression system.

Figure 21. SDS-PAGE and Western blot analysis of differently tagged proteins purified from E. coli using sequential Strep-tactin and Nickel affinity chromatography. (A) Coomassie Brilliant

Blue R 250-stained SDS polyacrylamide gel of purified protein samples. M: marker, lane 1: eluate from Nickel affinity column loaded with eluate from the Strep-tactin column, lane 2: wash from Nickel affinity column loaded with eluate from the Strep-tactin column, lane 3: eluate from

Nickel affinity column loaded with flow through from the Strep-tactin column, lane 4: mix of samples loaded in lanes 2 and 3, lane 5: eluate from nickel affinity column loaded with eluate from the Strep-tactin column, lane 6: wash from Nickel affinity column loaded with eluate from the Strep-tactin column, lane 7: eluate from Nickel affinity column loaded with flow through 75 from the Strep-tactin column. (B) Immunoblot probed with anti-Strep-tag antibody. (C)

Immunoblot probed with anti-His6-tag antibody.

Since the presence of heterodimers could not be confirmed on the basis of differences in molecular mass or by probing of blots with tag-specific antibodies, the ability to distinguish

HemA from HemT using differential chemical cleavage with hydroxylamine-HCl was investigated. Hydroxylamine specifically targets asparagine-glycine bonds (60). HemA has one such dipeptide, which when cleaved by hydroxylamine-HCl would yield two polypetides that have calculated molecular weights of 26.8 and 18.9 kD; HemT lacks any asparagine-glycine bonds. As shown Figure 22A, two peptides corresponding to the calculated sizes of the HemA cleavage products were observed following treatment of purified Strep-tagged HemA. Also, it is the 18.9 kD band corresponding to the C terminal portion of HemA that is detected with both of the affinity tag antibodies (Figure 22B). Since, HemA cleavage yields two bands that are not present in HemT, this indicates that hydroxylamine treatment of purified protein samples can be used to determine their polypeptide composition, and so confirm the presence of heterodimeric protein. 76

Figure 22. SDS-PAGE and immunoblot analysis of samples incubated with hydroxylamine-HCl.

(A) Coomassie Brilliant Blue R 250-stained SDS polyacrylamide gel of purified protein samples.

Lane 1: purified His6-HemT, lane 2: purified Strep-HemA. M: marker; lane 5: Strep-HemA; lane 6: His6-HemT. (B) Immunoblots probed with anti-Strep-tag antibody (Lanes 1 and 2) and anti-His6-tag antibody (lanes 5 and 6).

Discussion

To determine whether there are differences in enzymatic properties among R. sphaeroides ALA synthases and to resolve the roles of HemA and HemT isoenzymes, ALA synthases from three strains of R. sphaeroides were purified and their enzymatic properties compared. Although there are differences in the distribution and expression of hemA and hemT among the three strains, the specific ALA synthase activities and kinetic properties of HemA and

HemT enzymes from the three strains were very similar. This suggests that other differences must be considered to explain the roles of HemA and HemT isoenzymes in this bacterium. 77

Based on the substrate preference of HemA and HemT for succinyl-CoA, as opposed to acetyl-CoA from strain 2.4.9, the strain in which both enzymes are expressed, both HemA and

HemT primarily function as ALA synthases and not as other members of the Į-oxoamine synthase family. This makes sense, because R. sphaeroides has a bona fide KBL gene.

Although not directly assayed for, these ALA synthases are also not likely to function as AONSs because all three of the strains examined here require biotin. Moreover, HemA and HemT have a threonine at position 83 (R. sphaeroides HemA/HemT numbering) which restricts glycine as the only possible amino acid substrate. AONS and SPT utilize alanine and serine respectively, both of which are larger than glycine. Those amino acids would obstruct the binding of the second substrate in ALA synthases. Therefore, the evidence available to date suggests the primary function of both HemA and HemT is to synthesize ALA.

Feedback inhibition assays revealed that vitamin B12 is not inhibitory for HemA or

HemT. Lack of inhibition by vitamin B12 would make sense as it is only present in the cell in trace amounts. However, hemin was found to be inhibitory for both HemA and HemT, with

HemA enzymes being more sensitive to feedback inhibition by hemin than the HemT enzymes.

Hence, the role of HemT could be to augment ALA production when HemA activity is inhibited by elevated heme levels. Recent studies of hemA and hemT expression in R. sphaeroides strain

2.4.9 suggest that hemT transcription is important when cells are transitioning from aerobic to anaerobic dark growth with DMSO (47). Feedback inhibition of HemA by hemin becomes problematic at such a time, since the cells need to assemble a new respiratory chain that delivers electrons to DMSO reductase. Hence, HemT, which is less inhibited by hemin, would be able to supply ALA for heme production and so offset the reduction in ALA synthase activity due to feedback inhibition of HemA. The hemT gene in wild type strain 2.4.1 is silent, but hemA is 78 more strongly up-regulated under anaerobic-dark conditions in that strain than in wild type strain

2.4.9. It is thought that this difference in hemA transcription regulation bypasses the need for

HemT in strain 2.4.1. Since strain 2.4.3 is incapable of anaerobic dark-DMSO growth as it lacks a functional DMSO reductase, there may be no need for a HemT enzyme.

Purified wild type HemT enzyme was found to be more sensitive to oxidation than purified wild type HemA enzyme. Based upon the comparative properties of the wild type and proline-substituted mutant HemT enzymes, reduced cysteine at position 281 is important for

HemT activity. The residue is not predicted to be involved in substrate or cofactor binding. It is located on the subunit interface just below the PLP binding pocket. The proximity of the cysteines from both subunits suggests the possibility that, under oxidizing conditions, they could form a disulfide bond that would then restrict the conformational changes of the two subunits necessary for catalysis (49, 80, 90-92). The two subunits of ALA synthases are proposed to interact with each other to allosterically control enzyme activity; the opening of one active site influences the closure of the partner active site in the protein (92). This would presumably be restricted by the formation of the disulfide bond. Therefore, HemT would be less active under aerobic conditions, which is consistent with hemT expression data that show transcription is induced in cells cultured anaerobically in the dark (with DMSO) (45).

These findings argue that HemA can provide the cell with sufficient ALA other than when cytoplasmic heme levels are elevated. Apparently, this happens when cells are transitioning from aerobic to anaerobic respiration using DMSO as alternate electron acceptor.

The role of HemT enzyme, which is far less sensitive to inhibition by heme, although it is susceptible to inactivation in the presence of oxygen, is then to sustain or augment ALA production when HemA activity is diminished by high heme levels. This augmentation is not 79 required in strain 2.4.1 in which hemT is not expressed, presumably because the hemA gene is adequately up-regulated at the transcription level such that the diminished enzyme activity is offset by synthesizing more enzyme. Because strain 2.4.3 does not encode DMSO reductase, it cannot respire on that alternate electron acceptor. However, as suggested for strain 2.4.1, wild type 2.4.3 may have evolved to accommodate a high-heme cytoplasm in the absence of hemT through a combination of up-regulation of hemA and having a gene that encodes a HemA enzyme with nearly 2-fold higher specific activity relative to any of the other ALA synthases examined here.

Apparently, when purified from E. coli, HemA is bound by heme. The significance of this observation remains to be determined. However, since similarly purified HemT has no detectable heme bound, the higher affinity of HemA for heme suggested by the purification result would be consistent with the greater Ki value for heme of HemA versus HemT.

Since expression data indicate that HemA and HemT are both present in the cytoplasm of strain 2.4.9 when grown anaerobically in the dark with DMSO (45), it is possible that the polypeptides dimerize to form a heterodimer having unique properties. Toward investigating this possibility, Strep-tagged HemA and His6-tagged HemT_C281P were coexpressed in E. coli.

The protein yields for the resolved heterodimers of differentially tagged HemA and

HemT_C281P was only 0.2% of the total affinity purified protein. This very low yield may be due to poor binding of the heterodimer to the affinity resin; however, it supports the hypothesis that heterodimers of HemA and HemT can form. While technical difficulties prevented the use of antibodies to confirm the formation of heterodimers, the use of selective chemical cleavage has proven to be a valid alternative. 80

The differences in detectable PLP bound to purified HemA and HemT proteins (Figure

13) and the recent report by Kardon et al. (87) that ClpX is involved in PLP loading of ALA synthase in S. cerevisiae and D. rerio led to the hypothesis that ClpX could also be required in

PLP loading in HemT. However, that hypothesis was not supported as there was no difference in activity for HemT overexpressed in a ClpX-minus strain versus the ClpX-plus parent strain. 81

References

1. Woese CR, Kandler O, Wheelis ML. 1990. Towards a natural system of organisms: proposal for the domains Archaea, Bacteria, and Eucarya. Proc Natl Acad Sci U S A 87:4576-

4579.

2. Zeilstra-Ryalls JH, Kaplan S. 2004. Oxygen intervention in the regulation of gene expression: the photosynthetic bacterial paradigm. Cell Mol Life Sci 61:417-436.

3. Milgrom LR. 1997. The colours of life: an introduction to the chemistry of porphyrins and related compounds. Oxford University Press, New York.

4. Shemin D, Russell CS. 1953. į-Aminolevulinic acid, itsrole in the biosynthesis of porphyrins and purines 1. J Am Chem Soc 75:4873-4874.

5. Kikuchi G, Kumar A, Talmage P, Shemin D. 1958. The enzymatic synthesis of delta- aminolevulinic acid. J Biol Chem 233:1214-1219.

6. Warren JM, Deeryy E. 2009. Vitamin B12 (cobalamin) biosynthesis in the purple bacteria, p 81-95. In Hunter C, Daldal F, Thurnauer M, Beatty J (ed), The Purple Phototrophic

Bacteria, vol 28. Springer, Dordrecht, The Netherlands.

7. Willows RD, Kriegel AM. 2009. Biosynthesis of bacteriochlorophylls in purple bacteria., p 57-59. In Hunter C, Daldal F, Thurnauer M, Beatty J (ed), The Purple Phototrophic

Bacteria, vol 28. Springer, Dordrecht, The Netherlands.

8. Zeilstra-Ryalls JH. 2008. Chapter 39: Regulation of the tetrapyrrole biosynthetic pathway, p 777-798. In Hunter C, Daldal F, Thurnauer M, Beatty J (ed), The Purple Phototrophic

Bacteria, vol 28. Springer, Dordrecht, The Netherlands.

9. Martens J-H, Barg H, Jahn D. 2002. Microbial production of vitamin B12. Appl

Microbiol Biotechnol 58:275-285. 82

10. Roth JR, Lawrence JG, Bobik TA. 1996. Cobalamin (coenzyme B12): synthesis and biological significance. Annu Rev Microbiol 50:137-181.

11. Tuboi S, Kim HJ, Kikuchi G. 1970. Differential induction of fraction I and fraction II of delta-aminolevulinate synthetase in Rhodopseudomonas spheroides under various incubation conditions. Arch Biochem Biophys 138:155-159.

12. Lascelles J. 1960. The synthesis of enzymes concerned in bacterio-chlorophyll formation in growing cultures of Rhodopseudomonas spheroides. Journal of General Microbiology 23:487-

489.

13. Burnham BF, Lascelles J. 1963. Control of porphyrin biosynthesis through a negative feedback mechanism. Biochem J 87:462-472.

14. Warnick GR, Burnham BF. 1971. Regulation of porphyrin synthesis. Purification and characterization of G-aminolevulinic acid synthase. J Biol Chem 246:6880-6885.

15. Inoue I, Oyama H, Tuboi S. 1979. On the nature of the activating enzyme of the inactive form of delta-aminolevulinate synthetase in Rhodopseudomonas spheroides. J Biochem

86:477-482.

16. Tuboi S, Hayasaka S. 1972. Control of -aminolevulinate synthetase activity in

Rhodopseudomonas spheroides. II. Requirement of a disulfide compound for the conversion of the inactive form of fraction I to the active form. Arch Biochem Biophys 150:690-697.

17. Tuboi S, Hayasaka S. 1972. Partial purification of an enzyme which is required for the conversion of the inactive form to the active form of delta-aminolevulinate synthetase from

Rhodopseudomonas spheroides. J Biochem 72:219-222. 83

18. Bolt EL, Kryszak L, Zeilstra-Ryalls JH, Shoolingin-Jordan PM, Warren MJ. 1999.

Characterisation of the Rhodobacter sphaeroides 5-aminolevulinic acid synthase isoenzymes,

HemA and HemT, isolated from recombinant Escherichia coli. Eur J Biochem 265:290-299.

19. Clement-Metral JD. 1979. Activation of ALA synthase by reduced thioredoxin in

Rhodopseudomonas sphaeroides Y. FEBS Lett 101:116-120.

20. Tuboi S, Kim HJ, Kikuchi G. 1970. Occurrence and properties of two types of delta- aminolevulinate synthetase in Rhodopseudomonas spheroides. Arch Biochem Biophys 138:147-

154.

21. Pittard AJ. 1996. Biosynthesis of the aromatic amino acids., p 458-484. In Neidhardt

FC, Curtis III R, Ingraham JL, Lin ECC, Low KB, Magasanik B, Reznikoff WS, Riley M,

Schaechter M, Umbarger HE (ed), Escherichia coli and Salmonella Cellular and Molecular

Biology, vol 1. ASM Press, Washington DC.

22. Riddle RD, Yamamoto M, Engel JE. 1989. Expression of G-aminolevulinate synthase in avian cells: separate genes encode erythroid-specific and nonspecific isozymes. Proc Natl

Acad Sci USA 86:792-796.

23. Tai T-N, Moore MD, Kaplan S. 1988. Cloning and characterization of the 5- aminolevulinate synthase gene(s) from Rhodobacter sphaeroides. Gene 70:139-151.

24. Sato K, KoichiIshida K, Shirai M, Shimizu S. 2014. Occurrence and some properties of two types of į-aminolevulinic acid synthase in a facultative methylotroph,Protaminobacter ruber. Agric Biol Chem 49:3423-3428.

25. Zhang L, Chen J, Chen N, Sun J, Zheng P, Ma Y. 2013. Cloning of two 5- aminolevulinic acid synthase isozymes HemA and HemO from Rhodopseudomonas palustris with favorable characteristics for 5-aminolevulinic acid production. Biotechnol Lett 35:763-768. 84

26. Ferreira GC, Gong J. 1995. 5-aminolevulinate synthase and the first step of heme biosynthesis. J Bioenerg Biomembr 27:151-159.

27. Medlock AE, Dailey HA. 2009. Regulation of mammalian heme biosynthesis, p 116-

127. In Warren JM, Smith AG (ed), Tetrapyrroles. Springer.

28. Cotter PD, Baumann M, Bishop DF. 1992. Enzymatic defect in "X-linked" sideroblastic anemia: molecular evidence for erythroid delta-aminolevulinate synthase deficiency. Proc Natl Acad Sci U S A 89:4028-4032.

29. Cotter PD, Rucknagel DL, Bishop DF. 1994. X-linked sideroblastic anemia: identification of the mutation in the erythroid-specific delta-aminolevulinate synthase gene

(ALAS2) in the original family described by Cooley. Blood 84:3915-3924.

30. Whatley SD, Ducamp S, Gouya L, Grandchamp B, Beaumont C, Badminton MN,

Elder GH, Holme SA, Anstey AV, Parker M, Corrigall AV, Meissner PN, Hift RJ,

Marsden JT, Ma Y, Mieli-Vergani G, Deybach JC, Puy H. 2008. C-terminal deletions in the

ALAS2 gene lead to gain of function and cause X-linked dominant protoporphyria without anemia or iron overload. Am J Hum Genet 83:408-414.

31. Ducamp S, Schneider-Yin X, de Rooij F, Clayton J, Fratz EJ, Rudd A, Ostapowicz

G, Varigos G, Lefebvre T, Deybach JC, Gouya L, Wilson P, Ferreira GC, Minder EI, Puy

H. 2013. Molecular and functional analysis of the C-terminal region of human erythroid-specific

5-aminolevulinic synthase associated with X-linked dominant protoporphyria (XLDPP). Hum

Mol Genet 22:1280-1288.

32. Bishop DF, Tchaikovskii V, Nazarenko I, Desnick RJ. 2013. Molecular expression and characterization of erythroid-specific 5-aminolevulinate synthase gain-of-function mutations causing X-linked protoporphyria. Mol Med 19:18-25. 85

33. Fratz EJ, Clayton J, Hunter GA, Ducamp S, Breydo L, Uversky VN, Deybach JC,

34. McVean GA, Altshuler DM, Durbin RM, Abecasis GR, Bentley DR, Chakravarti A,

Clark AG, Donnelly P, Eichler EE, Flicek P, Gabriel SB, Gibbs RA, Green ED, Hurles ME,

Knoppers BM, Korbel JO, Lander ES, Lee C, Lehrach H, Mardis ER, Marth GT, McVean

GA, Nickerson DA, Schmidt JP, Sherry ST, Wang J, Wilson RK, Gibbs RA, Dinh H,

Kovar C, Lee S, Lewis L, Muzny D, Reid J, Wang M, Wang J, Fang X, Guo X, Jian M,

Jiang H, Jin X, Li G, Li J, Li Y, Li Z, Liu X, Lu Y, Ma X, Su Z, Tai S, et al. 2012. An integrated map of genetic variation from 1,092 human genomes. Nature 491:56-65.

35. Zheng J, Tian Q, Hou W, Watts JA, Schrum JW, Bonkovsky HL. 2008. Tissue- specific expression of ALA synthase-1 and heme oxygenase-1 and their expression in livers of rats chronically exposed to ethanol. FEBS Lett 582:1829-1834.

36. Tian Q, Li T, Hou W, Zheng J, Schrum LW, Bonkovsky HL. 2011. Lon peptidase 1

(LONP1)-dependent breakdown of mitochondrial 5-aminolevulinic acid synthase protein by heme in human liver cells. J Biol Chem 286:26424-26430.

37. Fujita H, Yamamoto M, Yamagami T, Hayashi N, Sassa S. 1991. Erythroleukemia differentiation. Distinctive responses of the erythroid-specific and the nonspecific delta- aminolevulinate synthase mRNA. J Biol Chem 266:17494-17502.

38. Yamamoto M, Hayashi N, Kikuchi G. 1983. Translational inhibition by heme of the synthesis of hepatic delta-aminolevulinate synthase in a cell-free system. Biochem Biophys Res

Commun 115:225-231. 86

39. Yamamoto M, Kure S, Engel JD, Hiraga K. 1988. Structure, turnover, and heme- mediated suppression of the level of mRNA encoding rat liver delta-aminolevulinate synthase. J

Biol Chem 263:15973-15979.

40. Yamauchi K, Hayashi N, Kikuchi G. 1980. Translocation of delta-aminolevulinate synthase from the cytosol to the mitochondria and its regulation by hemin in the rat liver. J Biol

Chem 255:1746-1751.

41. Yomogida K, Yamamoto M, Yamagami T, Fujita H, Hayashi N. 1993. Structure and expression of the gene encoding rat nonspecific form delta-aminolevulinate synthase. J Biochem

113:364-371.

42. Braidotti G, Borthwick IA, May BK. 1993. Identification of regulatory sequences in the gene for 5-aminolevulinate synthase from rat. J Biol Chem 268:1109-1117.

43. Neidle EL, Kaplan S. 1993. Expression of the Rhodobacter sphaeroides hemA and hemT genes, encoding two 5-aminolevulinic acid synthase isozymes. J Bacteriol 175:2292-2303.

44. Nereng KS, Kaplan S. 1999. Genomic complexity among strains of the facultative photoheterotrophic bacterium Rhodobacter sphaeroides. J Bacteriol 181:1684-1688.

45. Coulianos N. 2011. A comparison of ALA synthase gene transcription in three wild type strains of Rhodobacter sphaeroides. MS Thesis, Bowling Green State University.

46. Stoian N. 2018. Regulation of ALA synthase gene transcription in Rhodobacter sphaeroides 2.4.9. Doctoral dissertation, Bowling Green State University.

47. Coulianos N, Kaganjo J, and JH Zeilstra-Ryalls. 2017. Mauscript in preparation.

48. Tan D, Ferreira GC. 1996. Active site of 5-aminolevulinate synthase resides at the subunit interface. Evidence from in vivo heterodimer formation. Biochemistry 35:8934-8941. 87

49. Astner I, Schulze JO, van den Heuvel J, Jahn D, Schubert W-D, Heinz DW. 2005.

Crystal structure of 5-aminolevulinate synthase, the first enzyme of heme biosynthesis, and its link to XLSA in humans. EMBO J 24:3166-3177.

50. Marianayagam NJ, Sunde M, Matthews JM. 2004. The power of two: protein dimerization in biology. Trends Biochem Sci 29:618-625.

51. Stetefeld J, Jenny M, Burkhard P. 2006. Intersubunit signaling in glutamate-1- semialdehyde-aminomutase. Proc Natl Acad Sci U S A 103:13688-13693.

52. Eraso JM, Kaplan. S. 1994. prrA, a putative response regulator involved in oxygen regulation of photosynthesis gene expression in Rhodobacter sphaeroides. J Bacteriol 176:32-43.

53. Ind AC, Porter SL, Brown MT, Byles ED, de Beyer JA, Godfrey SA, Armitage JP.

2009. Inducible-expression plasmid for Rhodobacter sphaeroides and Paracoccus denitrificans.

Appl Environ Microbiol 75:6613-6615.

54. Luria SE, Burrous JW. 1957. Hybridization between Escherichia coli and Shigella. J

Bacteriol 74:461.

55. Baba T, Ara T, Hasegawa M, Takai Y, Okumura Y, Baba M, Datsenko KA, Tomita

M, Wanner BL, Mori H. 2006. Construction of Escherichia coli K-12 in-frame, single-gene knockout mutants: the Keio collection. Mol Syst Biol 2:2006 0008.

56. Datsenko KA, Wanner BL. 2000. One-step inactivation of chromosomal genes in

Escherichia coli K-12 using PCR products. Proc Natl Acad Sci U S A 97:6640-6645.

57. Schägger H. 2006. Tricine-SDS-PAGE. Nat Protoc 1:16-22.

58. Laemmli UK. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685. 88

59. Schneider CA, Rasband WS, Eliceiri KW. 2012. NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9:671-675.

60. Kurien BT, Scofield RH. 2006. Western blotting. Methods 38:283-293.

61. Bornstein P, Balian G. 1977. Cleavage at Asn-Gly bonds with hydroxylamine. Methods

Enzymol 47:132-145.

62. Burnham BF. 1970. >@G-Aminolevulinic acid synthase (Rhodopseudomonas spheroides). Methods Enzymol 17:195-204.

63. Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H,

Valentin F, Wallace IM, Wilm A, Lopez R, Thompson JD, Gibson TJ, Higgins DG. 2007.

Clustal W and Clustal X version 2.0. Bioinformatics 23:2947-2948.

64. Nicholas KB, Nicholas HB, Deerfield DW. 1997. GeneDoc: analysis and visualization of genetic variation.

65. Arnold K, Bordoli L, Kopp J, Schwede T. 2006. The SWISS-MODEL workspace: a web-based environment for protein structure homology modelling. Bioinformatics 22:195-201.

66. Biasini M, Bienert S, Waterhouse A, Arnold K, Studer G, Schmidt T, Kiefer F,

Gallo Cassarino T, Bertoni M, Bordoli L, Schwede T. 2014. SWISS-MODEL: modelling protein tertiary and quaternary structure using evolutionary information. Nucleic Acids Res

42:W252-258.

67. Bordoli L, Kiefer F, Arnold K, Benkert P, Battey J, Schwede T. 2009. Protein structure homology modeling using SWISS-MODEL workspace. Nat Protoc 4:1-13.

68. Kovach ME, Phillips RW, Elzer PH, Roop II RM, Peterson KM. 1994. pBBR1MCS:

A broad -host-range cloning vector. BioTechniques 16: 800-802. 89

69. Nandi DL. 1978. į-Aminolevulinic acid synthase of Rhodopseudomonas spheroides.

Arch Biochem Biophys 188:266-271.

70. Hira S, Tomita T, Matsui T, Igarashi K, Ikeda-Saito M. 2007. Bach1, a heme- dependent transcription factor, reveals presence of multiple heme binding sites with distinct coordination structure. IUBMB Life 59:542-551.

71. Shemin D, Kikuchi G. 1958. Enzymatic synthesis of 5-aminolevulinic acid. Ann NY

Acad Sci 75:122-128.

72. Mann S, Ploux O. 2011. Pyridoxal-5'-phosphate-dependent enzymes involved in biotin biosynthesis: structure, reaction mechanism and inhibition. Biochim Biophys Acta 1814:1459-

1466.

73. Hanada K. 2003. Serine palmitoyltransferase, a key enzyme of sphingolipid metabolism.

Biochim Biophys Acta 1632:16-30.

74. Mukherjee JJ, Dekker EE. 1987. Purification, properties, and N-terminal amino acid sequence of homogeneous Escherichia coli 2-amino-3-ketobutyrate CoA ligase, a pyridoxal phosphate-dependent enzyme. J Biol Chem 262:14441-14447.

75. Kaufholz AL, Hunter GA, Ferreira GC, Lendrihas T, Hering V, Layer G, Jahn M,

Jahn D. 2013. Aminolaevulinic acid synthase of Rhodobacter capsulatus: high-resolution kinetic investigation of the structural basis for substrate binding and catalysis. Biochem J

451:205-216.

76. Shoolingin-Jordan PM, Al-Daihan S, Alexeev D, Baxter RL, Bottomley SS, Kahari

ID, Roy I, Sarwar M, Sawyer L, Wang SF. 2003. 5-Aminolevulinic acid synthase: mechanism, mutations and medicine. Biochim Biophys Acta 1647:361-366. 90

77. Munakata H, Sun JY, Yoshida K, Nakatani T, Honda E, Hayakawa S, Furuyama K,

Hayashi N. 2004. Role of the heme regulatory motif in the heme-mediated inhibition of mitochondrial import of 5-aminolevulinate synthase. J Biochem 136:233-238.

78. Kitatsuji C, Ogura M, Uchida T, Ishimori K, Aono S. 2014. Molecular mechanism for heme-mediated inhibition of 5-aminolevulinic acid synthase 1. Bull Chem Soc of Jpn 87:997-

1004.

79. Hunter GA, Ferreira GC. 1999. Pre-steady-state reaction of 5-aminolevulinate synthase. Evidence for a rate-determining product release. J Biol Chem 274:12222-12228.

80. Zhang J, Ferreira GC. 2002. Transient state kinetic investigation of 5-aminolevulinate synthase reaction mechanism. J Biol Chem 277:44660-44669.

81. Krissinel E, Henrick K. 2007. Inference of macromolecular assemblies from crystalline state. J Mol Biol 372:774-797.

82. Kortemme T, Creighton TE. 1995. Ionisation of cysteine residues at the termini of model alpha-helical peptides. Relevance to unusual thiol pKa values in proteins of the thioredoxin family. J Mol Biol 253:799-812.

83. Roos G, Loverix S, Geerlings P. 2006. Origin of the pKa perturbation of N-terminal cysteine in alpha- and 3(10)-helices: a computational DFT study. J Phys Chem B 110:557-562.

84. Sauer RT, Baker TA. 2011. AAA+ proteases: ATP-fueled machines of protein destruction. Annu Rev Biochem 80:587-612.

85. Sauer RT, Bolon DN, Burton BM, Burton RE, Flynn JM, Grant RA, Hersch GL,

Joshi SA, Kenniston JA, Levchenko I, Neher SB, Oakes ES, Siddiqui SM, Wah DA, Baker

TA. 2004. Sculpting the proteome with AAA(+) proteases and disassembly machines. Cell

119:9-18. 91

86. Baker TA, Sauer RT. 2012. ClpXP, an ATP-powered unfolding and protein-degradation machine. Biochim Biophys Acta 1823:15-28.

87. Kardon JR, Yien YY, Huston NC, Branco DS, Hildick-Smith GJ, Rhee KY, Paw

BH, Baker TA. 2015. Mitochondrial ClpX activates a key enzyme for heme biosynthesis and erythropoiesis. Cell 161:858-867.

88. Martin A, Baker TA, Sauer RT. 2008. Diverse pore loops of the AAA+ ClpX machine mediate unassisted and adaptor-dependent recognition of ssrA-tagged substrates. Mol Cell

29:441-450.

89. Turbeville TD, Zhang J, Adams WC, Hunter GA, Ferreira GC. 2011. Functional asymmetry for the active sites of linked 5-aminolevulinate synthase and 8-amino-7- oxononanoate synthase. Arch Biochem Biophys 511:107-117.

90. Hunter GA, Zhang J, Ferreira GC. 2007. Transient kinetic studies support refinements to the chemical and kinetic mechanisms of aminolevulinate synthase. J Biol Chem 282:23025-

23035.

91. Lendrihas T, Hunter GA, Ferreira GC. 2010. Targeting the active site gate to yield hyperactive variants of 5-aminolevulinate synthase. J Biol Chem 285:13704-13711.

92. Na I, DeForte S, Stojanovski BM, Ferreira GC, Uversky VN. 2017. Molecular dynamics analysis of the structural and dynamic properties of the functionally enhanced heptavariant of mouse 5-aminolevulinate synthase. J Biomol Struct Dyn doi:10.1080/07391102.2016.1269688:1-14.

CHAPTER III

STRUCTURE-FUNCTION ANALYSIS OF THE BIFUNCTIONALITY OF

ALA SYNTHASES

Introduction

Actinomycetes are well known for their secondary metabolite production. Polyketides, which have characteristic complex carbon skeletons, are one type of secondary metabolite produced by these bacteria. These compounds confer a survival advantage to the bacteria by playing roles in self-defense, aggression or communication (1). Some polyketide metabolites contain 2-amino-3-hydroxycyclopent-2-en-1-one, also known as C5N unit, as part of their structure (Figure 23). The C5N unit-containing polyketides are otherwise quite diverse in structure, as shown in Figure 23, and are usually grouped according to similarity in overall chemical structures. These polyketides have a wide range of biologically important activities such as antiviral (2), antibacterial (3), antifungal (4), reverse transcriptase inhibitor (5) and antitumor (6) activities. The specific role of the C5N unit in C5N unit-containing polyketides is not yet known as some studies have reported that the removal of the unit led to increased activity, while others reported no change in activity (3, 7-10).

Figure 23. C5N unit-containing polyketides produced by actinomycetes. Black rectangles

identify the C5N unit substructure present in each.

Using radiolabeled feeding experiments, Nakagawa et al. (11) reported that the C5N unit is synthesized from glycine and succinate via 5-aminolevulinic acid (ALA) in Streptomyces

nodosus subsp asukaensis, which produces the C5N-containing polyketide asukamycin (11). In that study (11), the researchers also proposed that the reaction mechanism for C5N unit synthesis may be through an intramolecular cyclization reaction of ALA that would involve a PLP- dependent enzyme (11). Among organisms, the synthesis of ALA occurs through two alternative routes. One route is referred to as the C5 pathway as glutamate is the precursor, and is present in plants, archaea and most bacteria. The two reactions that are specific to the pathway are 94 illustrated in Figure 24 below. The second route is called the C4 or Shemin pathway found in animals and Į-proteobacteria. In this pathway, ALA is formed from the decarboxylative condensation of succinyl-CoA and glycine in a reaction that is catalyzed by the PLP-dependent enzyme ALA synthase (12).

Glu(UUC) CO tRNA CHO + + H2N H3N Mg2 + NADPH + NADP+ Glutamyl-tRNA reductase

COOH COOH Glutamyl-tRNAGlu Glutamate-1-semialdehyde Glutamyl-1-semialdehyde PLP aminotransferase

CO + CoASH COOH 2 H2N

COOH PLP + O O ALA synthase NH2 SCoA COOH Glycine Succinyl-CoA 5-aminolevulinic acid (ALA)

Figure 24. The two biosynthetic pathways of ALA. The dedicated reactions of the C5 pathway are shown in the top part of the figure. In the first reaction glutamyl-tRNAGlu is reduced to glutamate-1-semialdehyde by glutamyl-tRNA reductase. PLP-dependent glutamyl-1- semialdehyde aminotransferase then catalyzes the synthesis of ALA from glutamate-1- semialdehyde (13). The bottom part of the figure shows the C4 pathway in which ALA is formed through the condensation of glycine and succinyl-CoA, a reaction catalyzed by PLP- dependent ALA synthase (12). 95

Although ALA production in most organisms is formed exclusively by one or the other of the biosynthetic pathways, two pathways have been reported in Euglena gracilis. The C5 pathway is functional in the chloroplasts and is used to provide ALA for chlorophyll and protoheme synthesis, while the C4 pathway is functional in the mitochondria and supplies ALA for heme biosynthesis (14). The conclusion by Nakagawa et al. (11) that the ALA used in the synthesis of the C5N unit was formed from glycine and succinate suggested the presence of the

C4 pathway in actinomycetes as well, although these bacteria are known to produce ALA via the

C5 pathway.

Petrícek et al. (15) identified genes encoding the C4 enzyme ALA synthase, as well as the C5 enzymes, thereby confirming the presence of both C4 and C5 pathways in S. nodosus subsp. asukaensis. In that study, Petrícek et al. also found that disruption of the C5 pathway led to ALA auxotrophy, while ALA formation was sustained when the ALA synthase gene was disrupted. However, inactivation of the ALA synthase gene did lead to a lack of production of the secondary metabolite asukamycin. That the ALA synthase gene product was capable of catalyzing ALA formation was demonstrated by complementation of E. coli and Streptomyces coelicolor having mutations in their respective glutamyl tRNA-reductase gene that made the C5 pathway nonfunctional. Hence it was concluded that, in S. nodosus, the C5 pathway supplies

ALA for tetrapyrrole biosynthesis while the C4 pathway supplies ALA for asukamycin biosynthesis (15). Although ALA from the C4 pathway was known to be the precursor for the

C5N unit, the reaction mechanism for the synthesis of the C5N unit remained unsolved. It should be noted that the complementation experiments involved hyper-overexpression of the ALA synthase gene, suggesting the Streptomyces enzyme has very low activity. 96

The biosynthesis of the C5N unit was finally described by Zhang et al. (16) in

Streptomyces aizunensis NRLL-B-11277, which produces the C5N unit-containing secondary metabolite ECO-02301 (Figure 23, (16)). The biosynthetic pathway by which C5N units are formed is as follows (Figure 25): (i) ALA synthase catalyzes the synthesis of ALA from the condensation of glycine and succinyl-CoA. (ii) An acyl-CoA ligase then converts the ALA to

ALA-CoA. (iii) ALA synthase then catalyzes a second reaction in which it cyclizes the ALA-

CoA to form the C5N unit (16). The two enzymes, together with an amide synthetase that attaches the C5N unit to the rest of the structure of the polyketide, are encoded as an operon in the genome (16, 17).

COOH CO2 + CoASH H N H N 2 AMP + PPi 2 ATP + CoASH COOH PLP + O O O ALA synthase Acyl-CoA ligase NH 2 SCoA COOH O Glycine Succinyl-CoA 5-aminolevulinic acid (ALA) SCoA

ALA-CoA

ALA synthase PLP CoASH O O

NH NH2 R Amide synthetase

HO OH 2-amino-3-hydroxycyclopent-2- en-1-one(C N unit) 5

Figure 25. Synthesis of 2-amino-3-hydroxycyclopent-2-en-1-one (C5N unit) in actinomycetes.

ALA synthases found in organisms that produce the C5N unit-containing polyketides are referred to as cyclizing ALA synthases (17) to distinguish them from ALA synthases that produce ALA specifically for tetrapyrrole synthesis, which are referred to as classical ALA synthases. The discovery of the novel bifunctionality of cyclizing ALA synthase in C5N unit synthesis was the first report of the ability of any ALA synthase to catalyze the intramolecular cyclization of ALA-CoA, confirming the C5N unit synthesis hypothesis by Nakagawa et al. (11).

The cyclization reaction mechanism has been proposed to be similar to the ALA synthesis reaction mechanism, which is well described in the literature (12, 18-20). Both of the reaction mechanisms are discussed below.

The ALA synthesis reaction mechanism (Figure 26A) involves the ordered binding of substrates accompanied by conformational changes of the enzyme (21, 22). In the resting state

(open conformation), pyridoxal 5’-phosphate (PLP) is bound to the catalytic lysine residue to form an internal aldimine, Figure 26A-I (23). Glycine binding to the active site results in the breaking of the internal aldimine and consequent formation of the external aldimine between

PLP and glycine (Figure 26A-II). With the assistance of the conjugated ring system of PLP and the catalytic lysine residue, the Pro-R proton of glycine is abstracted, creating a resonance- stabilized quinonoid intermediate (Figure 26A-III). Binding of succinyl-CoA generates energy used to shift the enzyme to a closed conformation that ensures succinyl-CoA is in an optimal orientation relative to glycine in order for catalysis to take place. The enzyme-PLP-glycine carbanion attacks the carbonyl carbon atom of succinyl-CoA leading to the release of CoA and formation of the ȕ-ketoacid aldimine intermediate (Figure 26A-IV). The intermediate then undergoes a decarboxylation step that is assisted by a conserved histidine residue to form an enol intermediate (Figure 26A-V). Protonation of the enol group by the catalytic lysine residue, 98 presumably using the same proton abstracted from glycine, forms the ALA-bound external aldimine (Figure 26A-VI). The reversion of the enzyme into open conformation is accompanied by ALA release, which is the rate-limiting step.

The proposed ALA-CoA cyclization reaction mechanism is as shown in Figure 26B. In the resting state (open conformation) PLP forms an internal aldimine bond with the conserved catalytic lysine residue just as in the ALA synthesis reaction (Figure 26B-I). Binding of ALA-

CoA in the active site leads to not only the formation of the external aldimine bond with PLP

(Figure 26B-II) but also generates energy used to convert the enzyme from open to closed conformation. The Pro-R proton of carbon 5 of ALA-CoA is then abstracted with the help of

PLP and the catalytic lysine residue forming a resonance-stabilized quinonoid intermediate

(Figure 26B-III). The intermediate then nucleophilically attacks the thioester carbon of ALA-

CoA to cyclize the ALA forming the C5N unit. Reversion of the enzyme into the open conformation would be accompanied by release of the C5N unit.

While the two reaction mechanisms are very similar, there are some differences: (i) The

ALA synthesis reaction involves two substrates, glycine and succinyl-CoA. Glycine binds first to form the external aldimine, followed by succinyl-CoA binding, which triggers the conformational change. The C5N unit synthesis reaction involves one substrate, ALA-CoA, which forms the external aldimine and triggers the conformational change. (ii) The nucleophilic attack of the thioester carbon by the carbanion is external in the case of the ALA synthesis while in the case of the C5N unit formation it is internal. (iii) The ALA synthesis reaction pathway involves a decarboxylation step that is absent in the C5N unit synthesis reaction cycle.

Based on structural similarities noted between the crystal structure of the R. capsulatus classical ALA synthase and the modelled structure of S. aizunensis cyclizing ALA synthase, 99

Zhang et al. (16) proposed that classical ALA synthases could also catalyze the ALA-CoA cyclization reaction. However, since those organisms in which this second activity has been demonstrated rely upon the C5 pathway for ALA used in tetrapyrrole production (15), it is possible that there may be a specialization among the ALA synthases, such that some do, and others do not possess cyclizing activity. This possibility raises questions as to what structural features of these enzymes contribute to this specialization, and what evolutionary events lead to this outcome.

100

101

Figure 26. The reaction mechanisms for ALA synthesis and ALA-CoA cyclization by ALA synthase. (A) The ALA synthesis reaction mechanism. In the resting state PLP is bound to the catalytic lysine residue 248 (R. sphaeroides HemA numbering) (I). Glycine binding results in the formation of the external aldimine (II) that is deprotonated by the lysine residue to form the carbanion intermediate (III). The carbanion then nucleophilically attacks succinyl-CoA to release the CoA and form the ȕ-ketoacid aldimine intermediate (IV). Decarboxylation leads to the formation of an enol intermediate (V) which is reprotonated to form the ALA-bound external aldimine. ALA is then released to regenerate the internal aldimine. (B) The ALA-CoA cyclization reaction mechanism. PLP is bound to a catalytic lysine residue in the active site to form the internal aldimine (I). ALA-CoA binding leads to formation of the external aldimine (II) which is deprotonated to form the resonance stabilized quinonoid intermediate (III). The thioester carbon of ALA-CoA is then attacked by the carbanion releasing CoA and cyclizing the

ALA (IV). Release of the C5N unit regenerates the internal aldimine (16).

In a study characterizing cyclizing ALA synthases, it was reported that these enzymes had ALA synthase specific activities that are on the order of 520-folds lower than specific activities of classical ALA synthase (15, 16). Additionally, Zhang et al. reported that the catalytic efficiency of the cyclic ALA synthase was higher for the ALA-CoA cyclization reaction than the ALA synthesis reaction (16). Petrícek et al. hypothesized that the low ALA synthase activity of the cyclizing enzymes could be due to the presence of serines at positions 83 and 363

(R. sphaeroides HemA numbering), which are invariantly threonines in classical ALA synthases

(15). This hypothesis had not yet been experimentally tested. A phylogenetic study of the cyclic

ALA synthases (17) reported that the two serine residues are also conserved in all cyclic ALA synthases. 102

The threonine/serine residues at positions 83 and 363 are located in the active site of the enzymes. For classical ALA synthases, the two conserved threonine residues are involved in substrate specificity and recognition. Threonine 83 forms a hydrogen bond with the carboxylate oxygen of succinyl-CoA and plays an important role in the specificity of ALA synthase for glycine (22, 24, 25). Threonine 363 forms a hydrogen bond with the second oxygen of the carboxyl group of succinyl-CoA, and is important in recognition and stabilization of succinyl-

CoA in an optimal orientation for catalysis (22). With respect to ALA-CoA cyclizing activity, serine 83 was shown to be important in C5N unit synthesis by Rui et al. (26) who reported that S. nodosus subsp. asukaensis ALA synthase mutant protein in which serine 83 is replaced with threonine failed to produce asukamycin, but was able to complement the requirement of ALA in a Streptomyces coelicolor strain that relied upon the enzyme (hyper-overexpressed) to provide

ALA required for growth (the normal C5 pathway was nonfunctional because of a defective glutamyl tRNA-reductase gene). Whether the residues at these two positions are important in defining, or are fully responsible for distinguishing cyclizing ALA synthases from classical ALA synthases is not known.

In this study, the ALA synthase and ALA-CoA cyclase activities of two classical ALA synthases from R. sphaeroides 2.4.9 and one cyclizing ALA synthase from S. aizunensis NRRL-

B-11277 were directly compared. The roles of threonines versus serines at positions 83 and 363 in the ALA-CoA cyclizing and ALA synthesis activities were investigated.

Materials and methods

Bacterial strains, plasmids and protein production. Bacterial strains and plasmids are listed in Table 11. Production of wild type ALA synthases was as described in Chapter 2. 103

Plasmids expressing HemA_T83S, HemA_T363S and HemA_T83S/T363S mutant proteins were obtained from Joyce Liu in the lab of Wenjun Zhang (University of California, Berkeley), and they were overexpressed in E. coli TOP10 (Invitrogen). Purification of the mutant proteins followed the same protocol as described for wild type proteins in Chapter 2 using IPTG at a final concentration of 0.5 mM for induction. The S. aizunensis NRLL ALA-CoA cyclase (AcaC) expression vector used was pET30-acaC (16), and E. coli strain BL21 Gold (DE3) was used for protein production. Plasmids expressing mutant AcaC_S83T, AcaC_S363T, and

AcaC_S83T/S363T proteins were derived from pET30-acaC. All of these strains were obtained from Joyce Liu.

For production of wild type and mutant AcaC protein precultures were used to inoculate

LB medium containing 50 μg/ml kanamycin (Kn), which were incubated at 37oC with shaking at

200 rpm to an OD660 of approximately 0.5. After cooling on ice for 10 min protein expression was induced by the addition of IPTG to a final concentration of 0.12 mM and the cultures were incubated for 11 hours at 25oC. Cells were collected by centrifugation at 8327 × g for 15 min at

4oC in an Eppendorf 5810R centrifuge using an A-4-62 swinging bucket rotor and stored at -

80oC.

Protein purifications, concentration determinations, SDS-PAGE and InVision™ his- tag in-gel stain assay, and ALA synthesis activity assays and determination of kinetic parameters. All were performed as described in Chapter 2, with the exception of AcaC assays for ALA synthase activity. These were performed at 30ºC using glycine at concentrations between 0 and 1 M. 104

ALA-CoA cyclization activity assays. ALA-CoA cyclase assays of purified wild type and mutant HemA protein, and of purified wild type and mutant AcaC protein were performed by Joyce Liu in the Zhang lab following the protocol described by Zhang et al (16).

Bioinformatic analyses, protein modelling and structural analyses. Bioinformatic analyses and protein modelling followed the same procedure as described in Chapter 2.

Additionally, ALA-CoA and ALA-CoA bound with PLP were drawn using Chemsketch free edition software (ACD/Structure Elucidator, version 15.01, Advanced Chemistry Development,

Inc., Toronto, ON, Canada, www.acdlabs.com, 2015). The structures were converted into PDB files using Discovery studio 2016 (Dassault Systèmes BIOVIA, Discovery Studio Modeling

Environment, Release 2016, San Diego: Dassault Systèmes, 2016). Then ALA-CoA and ALA-

CoA-PLP ligands were docked to proteins using PyMOL.

105

Table 11. Bacterial strains and plasmids.

Strains Relevant properties Reference

Escherichia coli

DH5Įphe F–, (ĳ80dlacZǻM15) recA1 endA1 hsdR17 supE44 27

thi-1 gyrA96 relA1 deoR ǻ(lacZYA-argF) U169

phe::Tn10d; CmR.

TOP10 F–, mcrA ǻ(mrr-hsdRMS-mcrBC) ĭ80lacZǻM15 Invitrogen

ǻlacX74 recA1 araD139 ǻ(ara leu) 7697 galU

galK rpsL (StrR) endA1 nupG

BL21 Gold (DE3) F–, fhuA2 [lon] ompT gal (Ȝ DE3) [dcm] ¨hsdS Ȝ Stratagene

DE3 = Ȝ sBamHIo ¨EcoRI-B

int::(lacI::PlacUV5::T7 gene1) i21 ¨nin5

Plasmids

pIND4 IPTG-inducible expression vector, pMG160 28

replicon; KnR.

pIND4-2.4.9-hemA pIND4 derivative with synthetic 2.4.9-hemA gene This

with a C-terminus polyhistidine tag and having an study.

NcoI recognition site at the 5’ end and HindIII

recognition site at the 3’ end; KnR.

pIND4-2.4.9-hemT pIND4 derivative with synthetic 2.4.9-hemT gene This

with a C-terminus polyhistidine tag and having an study.

NcoI recognition site at the 5’ end and HindIII

recognition site at the 3’ end; KnR.

pIND4-2.4.9-hemA_T83S pIND4 derivative with synthetic 2.4.9-hemA_T83S Joyce Liu

gene with a C-terminus polyhistidine tag and having 106

an NcoI recognition site at the 5’ end and HindIII

recognition site at the 3’ end; KnR. pIND4-2.4.9-hemA_T363S pIND4 derivative with synthetic 2.4.9-hemA_T363S Joyce Liu

gene with a C-terminus polyhistidine tag and having

an NcoI recognition site at the 5’ end and HindIII

recognition site at the 3’ end; KnR. pIND4-2.4.9-hemA_T83S/T363S pIND4 derivative with synthetic 2.4.9- Joyce Liu

hemA_T83S/T363S gene with a C-terminus

polyhistidine tag and having an NcoI recognition

site at the 5’ end and HindIII recognition site at the

3’ end; KnR. pET30-acaC pET-30 Xa/LIC derivative harboring S. aizunensis (16)

NRRL B-11277 acaC gene; KnR. pET30-acaC_S83T pET-30 Xa/LIC derivative harboring S. aizunensis Joyce Liu

NRRL B-11277 acaC_T83S mutant gene; KnR. pET30-acaC_S363T pET-30 Xa/LIC derivative harboring S. aizunensis Joyce Liu

NRRL B-11277 acaC_T363S mutant gene; KnR. pET30-acaC_S83T/S363T pET-30 Xa/LIC derivative harboring S. aizunensis Joyce Liu

NRRL B-11277 acaC_T83S/T363S mutant gene;

KnR.

107

Results

To investigate whether classical ALA synthases have ALA-CoA cyclizing activity, and also to examine the role of the conserved residue differences between the cyclizing and classical enzymes at positions 83 and 363, a collaborative investigation was initiated with the lab of

Wenjun Zhang at the University of California, Berkeley, who first described the cyclizing activity of the S. aizunensis NRRL B-11277 cyclic ALA synthase (16). Two ALA synthase isoenzymes (HemA and HemT) from R. sphaeroides 2.4.9 were selected as representatives of classical ALA synthases. HemA and HemT are 55% (216/391) identical to each other with a similarity of 72% (283/391). The S. aizunensis enzyme (named AcaC for ALA-CoA cyclase) was used as a representative of cyclizing ALA synthases. AcaC is 50% (192/386) and 51%

(194/380) identical to HemA and HemT with similarities of 65% (254/386) and 70% (266/380), respectively.

Protein purification. The recombinant polyhistidine-tagged proteins were expressed in

E. coli and purified using Ni-IDA affinity chromatography. Purified wild type HemA (45.7 kDa), HemT (45.3 kDa), AcaC (49.2 kDa) and mutant HemA and AcaC proteins all migrated as single bands near the 50 kDa molecular mass marker as shown in Figure 27. The purified proteins were also detected in the same PAGs that were subjected to an in-gel visualization analysis for detection of polyhistidine-tags, using the InVision His-Tag stain (Invitrogen). Image analysis of the PAG gel using ImageJ software (29) indicated that the purities of the homogeneous proteins were greater than 90%. 108

Figure 27. SDS-PAG of purified proteins. Image of the Coomassie Brilliant Blue R 250 stained gel (A) and image of the same gel stained with Invision His-Tag stain (B). Lane 1: R. sphaeroides 2.4.9-HemT, lane 2: R. sphaeroides 2.4.9-HemA, lane 3: R. sphaeroides 2.4.9-

HemA_T83S, lane 4: R. sphaeroides 2.4.9-HemA_T363S, lane 5: R. sphaeroides 2.4.9-

HemA_T83S/T363S, M: Precision Plus Protein Kaleidoscope standards (Bio-Rad Laboratories), lane 6: S. aizunensis AcaC, lane 7: S. aizunensis AcaC_S83T, lane 8: S. aizunensis AcaC_S363T, lane 9: S. aizunensis AcaC_S83T/S363T.

ALA-CoA cyclase activities of classical and cyclizing ALA synthases. ALA-CoA cyclization activities were determined by Joyce Liu, in the lab of W. Zhang, using a previously described assay (16). As shown in Figure 28 (data provided by J. Liu), HemA and HemT were able to produce the C5N unit, supporting the hypothesis by Zhang et al. (16) that classical ALA synthases can also catalyze the ALA-CoA cyclization reaction. However, although C5N unit formation was demonstrated for both HemA and HemT, their relative cyclase activities were

10% and 6% that of AcaC, respectively. The low ALA-CoA cyclase activity of HemA and 109

HemT indicates that there are amino acid sequence differences between classical ALA synthases and cyclizing ALA synthases that are important for the ALA-CoA cyclase activity.

Figure 28. Percent relative cyclase activities of purified S. aizunensis wild type AcaC and mutants AcaC _S83T, AcaC _S363T, and AcaC _S83T&S363T, and R. sphaeroides 2.4.9 wild type HemA and mutants HemA_T83S, HemA_T363S, and HemA_T83S&T363S, and R. sphaeroides 2.4.9 wild type HemT. Relative cyclase activities were calculated by comparing activities to that of S. aizunensis wild type cyclic AcaC (arbitrarily set at 100%). Values are the means of at least three replicates with standard deviations indicated by error bars. The data are those of J. Liu (W. Zhang lab, University of California-Berkeley).

Whether the low ALA-CoA cyclase activities of HemA and HemT relative to AcaC are due to amino acid residue differences at positions 83 and 363 (R. sphaeroides HemA numbering) was investigated by studying mutant proteins having reciprocal substitutions – serines were 110 replaced with threonines in AcaC and threonines were replaced with serines in HemA. The mutant proteins docked ALA-CoA bound to PLP were modeled to examine the consequence of the mutations on the active site of each enzyme. Of note in this modelling is that the mutations do not have any effect on the active site architecture, and neither threonines nor serines form hydrogen bonds with ALA-CoA, as shown in Figure 29. If the low ALA-CoA cyclase activity was due to the threonine residues in the two positions in HemA (Figure 29A), then it may be that the methyl group of threonine, which is absent from serine, causes steric hindrances that would limit the optimal orientation of ALA-CoA. Substitution of the threonines in HemA with serines

(Figure 29B) would eliminate this steric hindrance, and so improve ALA-CoA cyclase activity.

By the same reasoning, substitution of serines with threonines in AcaC (Figure 29D) would decrease the ALA-CoA cyclase activity of the enzyme.

111

Figure 29. Predicted structures of R. sphaeroides 2.4.9-HemA and S. aizunensis AcaC with substrates docked in the active site. The threonines or serines at positions 83 and 363 are shown as red sticks with hydrogen bonds to the carboxyl group of succinyl-CoA indicated for each residue. The catalytic lysine 248 residue is shown as a blue stick. Docked ALA-CoA is shown in the active sites of R. sphaeroides 2.4.9-HemA wild type protein (A), 2.4.9-

HemA_T83S/T363S mutant protein (B), S. aizunensis AcaC wild type protein (C), and

AcaC_S83T/S363T mutant protein (D).

112

Consistent with the prediction from modelling, the cyclase activities of AcaC_S83T and

S363T were approximately 60% that of wild type AcaC, and that of the double mutant was 30%

(Figure 28). However, the abilities of the serine-substituted HemA mutant proteins to form C5N were not improved relative to the wild type protein, with the doubly-substituted protein retaining approximately 90% of the cyclizing activity. These results indicate that, while the residues at positions 83 and 363 are important for ALA-CoA cyclase activity, they are not solely responsible for distinguishing a cyclase from an ALA synthase.

ALA synthase activities of classical and cyclizing ALA synthases. ALA synthase assays were performed at an optimum temperature of 37°C for HemA and HemT and 30°C for

AcaC, and a pH optimum of 7.2 for both purified enzymes. The specific activities and kinetic parameters of HemA and HemT were obtained from Chapter two. As listed in Table 12, the specific activities for the classical ALA synthases HemA and HemT were 62.9 U/mg, 63.0

U/mg, respectively (1 unit is defined as 1 μmole ALA formed per hour). The specific activity of

AcaC was 0.2 U/mg, which is 315-fold lower than that of the R. sphaeroides enzymes. The turnover number (kcat) for AcaC was 76 and 73-fold lower than that of HemA and HemT, respectively, and AcaC exhibited low affinities for substrates compared to those of HemA and

HemT; the Km for glycine is 13 and 8-fold higher than that of HemA and HemT, respectively, while the Km for succinyl-CoA was 8-fold higher than that of both HemA and HemT. These results indicate that, although AcaC is capable of catalyzing the ALA synthase reaction, it is a far less efficient ALA synthase compared to HemA and HemT. The low ALA synthase activity measured for AcaC is in agreement with ALA synthase activities of cyclizing ALA synthases reported previously (15, 16). 113

Whether the conserved serines at positions 83 and 363 are responsible for the low ALA synthase activity of AcaC, as proposed by Petrícek et al. (15), was analyzed by examining mutant proteins in which the serines were replaced with threonines. Likewise, the corresponding threonines in HemA were substituted with serines. Modelling results (Figure 30) indicate that the hydrogen bonds between the oxygens of the carboxyl group of succinyl-CoA and threonines at positions 83 and 363 are also present when serines are present at the same positions in wild type AcaC and mutant HemA. However, the methyl groups that contribute to hydrophobic interactions that aid in optimal orientation of succinyl-CoA (22) are absent when serines are present at the two positions. Additionally, the hydrogen bond between S83 and the carboxyl group of succinyl-CoA in AcaC is predicted to be 0.7 Å longer than that of T83 of HemA. Thus, based on the models, substitution of these residues would lead to mutant AcaC proteins that are more like wild type HemA, and mutant HemA proteins would be more like wild type AcaC. The prediction is that the ALA synthase activity of the mutant AcaC proteins would improve while that of the mutant HemA proteins would worsen.

As shown in Table 12, the specific ALA synthase activity of the HemA single and double mutant proteins were approximately 4-fold less than that of wild type HemA, as expected.

However, unlike the prediction, the specific activities of AcaC_S83T Aca_S363T, and

AcaC_S83T/S363T were 9-fold, 4-fold, and 40-fold less than wild type AcaC, respectively. The very low ALA synthase activities of the AcaC mutants made it impossible to accurately determine their kinetic properties.

114

Figure 30. Predicted structures of R. sphaeroides 2.4.9-HemA and S. aizunensis AcaC with substrates docked in the active site. Threonines and serines at position 83 and 363 are shown as red sticks with hydrogen bonds to the carboxyl group of succinyl-CoA indicated for each residue. The catalytic lysine 248 residues are shown as blue sticks. Docked succinyl-CoA and glycine bound to PLP are shown in the active site of R. sphaeroides 2.4.9-HemA wild type (A),

2.4.9-HemA _T83S/T363S mutant (B), S. aizunensis AcaC wild type (C), and AcaC

_S83T/S363T mutant (D). 115

The kcat values of HemA_T83S, HemA_T363S, and HemA_T83S/T363S mutant proteins were approximately 5-fold lower than that of wild type HemA. The Km values of the mutant enzymes for glycine were lower than that of the wild type, but the Km values for succinyl-CoA were higher. The apparent increased affinity of the mutant proteins for glycine may be due to the absence of the threonine methyl group in serine that could improve the binding of glycine, as van der Waals contacts between the threonine methyl group and the alpha carbon of glycine would be missing (22). Improved glycine affinity has also been reported for a R. capsulatus ALA synthase T83S mutant protein (25). The reduced affinity for succinyl-CoA of the mutant enzymes may be due to a less than optimal orientation of succinyl-CoA in the active site, caused by the missing hydrophobic interactions contributed by the methyl groups of the threonines, and lengthening of the hydrogen bond from 2.6 Å for threonine to 3.4 Å for serine (Figure 30B).

While the serine-substituted HemA mutants have decreased turnover numbers and lower affinities for succinyl-CoA relative to wild type HemA, as is also true for wild type AcaC, their affinities for glycine were higher than AcaC. These results suggest that residues at positions 83 and 363 are important for ALA synthase activity, but they are not the sole determinants of the low ALA synthase activity exhibited by AcaC.

116

Table 12. Kinetic properties of R. sphaeroides 2.4.9 HemT, HemA and HemA mutant and wild type S. aizunensis ALA synthases.

Propertya

Gly S-CoA Gly S-CoA Enzyme Km Km kcat kcat/Km kcat/Km Specific activityb (mM) (μM) (h-1) (mM/h) (μM/h)

Wild type 2.4.9-HemA* 62.9 ± 3.5 31.7 ± 2.8 14.6 ± 3.2 3804.0 ± 304 120.0 260.6

Wild type 2.4.9-HemT* 63.0 ± 6.3 52.5 ± 5.0 15.0 ± 3.3 3627.3 ± 400 69.1 241.8

2.4.9-HemA_T83S 13.6 ± 2.5 14.0 ± 1.9 77.4 ± 16.0 728.4 ± 116.0 52.0 9.4

2.4.9-HemA_T363S 14.2 ± 2.0 11.3 ± 3.8 164.5 ± 38.1 722.0 ± 123.6 63.9 4.4

2.4.9-HemA_T83S/T363S 15.7 ± 2.1 16.7 ± 5.4 130.2 ± 25.9 787.5 ± 151.9 47.2 6.1

Wild type S. aizunensis AcaC 0.20 ± 0.1 411.9 ± 108.1 118.3 ± 51.7 49.5 ± 7.7 0.1 0.4

AcaC_S83T 0.023 ± 0.0 ND ND ND ND ND

AcaC_S363T 0.051 ± 0.0 ND ND ND ND ND

AcaC_S83T/S363T 0.005 ± 0.0 ND ND ND ND ND

ND not determined a Values are the means of at least three replicates ± standard deviation. bμmoles ALA/h/mg

*data from chapter two.

117

Discussion

In this study, the ALA synthase and ALA-CoA cyclase activities of S. aizunensis AcaC, representing cyclizing ALA synthases, and R. sphaeroides 2.4.9 HemA and HemT, representing classical ALA synthases, were compared. The results reveal that HemA and HemT can catalyze the ALA-CoA cyclization reaction, as proposed by Zhang et al. (16). Despite considerable structural similarity, however, the cyclase activities of HemA and HemT were approximately

10% that of AcaC while the specific ALA synthase activities of those enzymes were 315-fold higher than that of AcaC. Thus, these enzymes, while capable of performing both activities, appear to be specialized for one activity or the other.

Toward understanding this specialization, two conserved residues at positions 83 and 363 that are threonines in all classical ALA synthases but serines in all known cyclizing ALA were evaluated by examining mutant proteins in which threonines were replaced with serines and serines were replaced with threonines. It was thought that threonine to serine substitutions would reduce the ALA synthase activity of HemA while improving its ALA-CoA cyclizing activity, and for AcaC, the serine to threonine substitution would improve ALA synthase activity while the ALA-CoA cyclizing activity would decrease. This expectation was not borne out as

ALA-CoA cyclase activity of HemA mutant proteins and ALA synthase activity of AcaC mutant proteins were not improved relative to the respective wild type proteins. Interestingly, while single- and double serine-substituted mutant HemAs had lower ALA synthase activity as expected, their ALA-CoA cyclization activities were only approximately 10% lower than wild type HemA. Single threonine substituted AcaC proteins had decreased ALA synthase and ALA-

CoA cyclase activities, and both activities of the doubly substituted protein were even lower. It may be that the methyl group of threonine is constricting an already small active site in AcaC. 118

This would then suggest that there are differences in the active site architecture of the two enzymes that could not be observed by modeling of wild type and mutant proteins.

The reciprocal substitution results indicate that the residues at positions 83 and 363 are important for enzyme activity within the overall sequence context of each enzyme. However, they also indicate that other differences are important with respect to each of the activities as well. Whether other conserved residue differences identified from alignment of classical and cyclizing ALA synthases (Figure 31) play a role in the observed differences in activity remains to be tested. Possibilities include residues at positions 135, 145 and 149, which are located in close proximity to the hydrophobic binding pocket of the adenylyl moiety of succinyl-CoA; isoleucine 149 in the solved crystal structure of R. capsulatus ALA synthase is located at the bottom of this hydrophobic pocket (22). Differences in these residues could affect the binding of the adenyl moiety, affecting catalysis.

119

REGION 1 REGION 2 10------24 40----47 | | | | * S. xanthochromogenes ----MPRHLDYFTDQLTDEQRRTRTFLEIGRRAGNFPSAIARTA---DEDSEVSVWCSNDYLGMGQHPAVLAAVHDAVARFGAGSGGSRNIGGTN S. aizunensis NRRL-B-11277 --MNLHLESYSTGVTAKELAERRREFLEIGRRSGHFPSASARQD---GVDSQISVWCSNDYLGMGQNPQVIEAMKKTIDTHGVGSGGSRNIGGTN S. nodosus ssp. asukaensis ----MNKHLDFFAREMEEFGARRREFLEIGRRAGRFPSAVARQGQ-DGTDVEISVWCSNDYLGMGQNPFVLEAVKNAVDAFGAGSGGSRNIGGTN S. aureus SOK1/5-04 ----MTLHVDLFSQEMKEFAHHKRQFLEIGRNAGRFPSAVARQGH-DGTDVEISVWCSNDYLGMGQNPSVIEAMKEAIDAFGAGSGGSRNIGGTN S. sp. CNB091 --MNHYLELFSRKTGEGGASDAKREFLELGRLAGKFPAARSLRDG---VTSEISVWCSNDYLGMGQHPLVLAATREALDEYGAGSGGSRNIGGTN S. sp. PCS3-D2 --MNQYLELFSRHMKEGGSAEAKREFLEVSRLAGKFPAARSLRDG---IGSEISVWCSNDYLGMGQHPLVLAATREALDAYGAGSGGSRNIGGTN S. rapamycinicus NRLL5491 -MTTQYLDLFSRLTE--GSDGGKREFLEIGRLAGQFPVASVIG---VEDTSRISVWCSNDYLGMGQHPAVLEAMKEAVDEYGAGAGGSRNIGGTN S. sp. PRh5 -MTTQYLDLFSRLTE--GSDGGKREFLEIGRLAGQFPAASVSG---VEDTSRINVWCSNDYLGMGQHPAVLEAMKEAVDEYGAGAGGSRNIGGTN S. ghanaensis ATCC14672 MoeA5 MDISSSMDFFVRLARETGDRK--REFLELGRKAGRFPAASTSNG------EISIWCSNDYLGMGQHPDVLDAMKRSVDEYGGGSGGSRNTGGTN S. clavuligerus ATCC27064 --MSQYMDFFARLAHESADRK--REFLELGRIAGRFPAATTAENG------EIAVWCSNDYLGMGQHPAVLEAMHRAVDEYGAGSGGSRNIGGTH R. capsulatus SB1008 HemA ---MDYNLALDKAIQKLHDEGRYRTFIDIEREKGAFPKAQWN--RPDGGKQDITVWCGNDYLGMGQHPVVLAAMHEALEAVGAGSGGTRNISGTT R. sphaeroides 2.4.9 HemA ---MDYNQALDTALNRLHTEGRYRTFIDIERRKGAFPKAMWR--KPDGSEKEITVWCGNDYLGMGQHPVVLGAMHEALENTGAGSGGTRNISGTT Rps. palustris HemA ---MNYEAYFRRQLDGLHREGRYRVFADLERHAGSFPRATHH--RPEGA-GDVTVWCSNDYLGMGQHPAVLTAMHEALDSCGAGAGGTRNIAGTN Rps. palustris HemO ---MQYNKFFEDAVARLHDERRYRVFADLERIAGRFPYAVWH--SQTGP-RDVVIWCSNDYLGMGQHPKVVGAMVETTTRIGTGAGGTRNIAGTH R. sphaeroides 2.4.9 HemT ---MEFSQHFQKLIDDMRLDGRYRTFAELERIAGEFPTALWH--GPDGQARRVTVWCSNDYLGMGQNAEVLAAMHRSIDLSGAGTGGTRNISGTN M. musculus ALAS2 ---FGYDQFFRDKIMEKKQDHTYRVFKTVNRWANAYPFAQHFS-EASMASKDVSVWCSNDYLGISRHPRVLQAIEETLKNHGAGAGGTRNISGTS H. sapiens ALAS2 ---FSYDQFFRDKIMEKKQDHTYRVFKTVNRWADAYPFAQHFS-EASVASKDVSVWCSNDYLGMSRHPQVLQATQETLQRHGAGAGGTRNISGTS H. sapiens ALAS1 ---FQYDRFFEKKIDEKKNDHTYRVFKTVNRRAHIFPMADDYS-DSLITKKQVSVWCSNDYLGMSRHPRVCGAVMDTLKQHGAGAGGTRNISGTS | S | G | S SS | 1------24 77------89 SECOND CHANNEL SECOND CHANNEL

* * * * S. xanthochromogenes HRHVELERELADWHGKDAALLFTSGYTANDGALAVIAGNPEGTVVFSDELNHASIIDGLRRSGAQKRIFRHNDVAHLAELLAQADPEAPKLIVLE S. aizunensis NRRL-B-11277 HYHVLLEAELADLHGKEAALLFTSGYTANDGSLSVLAGTPKDTIVFSDEKNHASIIDGLRHSGAQKHIFRHNDVAHLAELLAAAPADRPKLIVLE S. nodosus ssp. asukaensis HYHVLLENELAALHGKEEALIFPSGFTANDGALTVLAGRAPGTLVFSDELNHASIIDGLRHSGAEKRIFRHNDMAHLEELLAAADPERPKLIVLE S. aureus SOK1/5-04 HYHVLLEKELAAFHGKEAALLFSSGYTANDGALSVLAGRMPGTIVYSDALNHASIIDGLRHSGAQKRIFRHNDVAHLEELIAADPADRPKLIVLE S. sp. CNB091 HYHVLLEKELADLHGKADALLFTSGYTANDGALTVLAGLPEKCIVFSDEMNHASIIDGLRHSGAEKRIFRHNDMAHLEELISATDPDRPKMVVME S. sp. PCS3-D2 HYHVLLEQELADLHGKANALLFTSGYTANDGALTVLAGRLGDCVVFSDEMNHASIIDGLRHSGAEKRIFRHNDTAHLEELISATDSDRPKMIVME S. rapamycinicus NRLL5491 HYHVALEKELSALHGKDDALLFTSGYTANDGALSVIAGRMEGCVVFSDALNHASIIDGLRHSGAQKRIFRHNDTAHLEELLAAADPDAPKLIVSE S. sp. PRh5 HCHVLLEKELSALHGKDDALLFTSGYTANDGALSVIAGRMEGCVVFSDALNHASIIDGLRHSGAQKRIFRHNDTAHLEELLAAADPDAPKLIVTE S. ghanaensis ATCC14672 MoeA5 HFHVALEREPAEPHGKEDAVLFTSGYSANEGSLSVLAGAVDDCQVFSDSANHASIIDGLRHSGARKHVFRHKDGRHLEELLAAADRDKPKFIALE S. clavuligerus ATCC27064 HHHVLLERELADLHGKEEALLFTSGYSANEGSLSVLAGRIDGCEVFSDQGNHASIIDGLRHSGARKHIFRHNDRAHLEELLAAADPDVPKLIVTE R. capsulatus SB1008 HemA AYHRRLEAEIADLHGKEAALVFSSAYIANDATLSTLRVLFPGLIIYSDSLNHASMIEGIKRNAGPKRIFRHNDVAHLRELIAADDPAAPKLIAFE R. sphaeroides 2.4.9 HemA LYHKRLEAELADLHGKEAALVFSSAYIANDATLSTLPQLIPGLVIVSDKLNHASMIEGIRRSGTEKHIFKHNDLDDLRRILSSIGKGRPILVAFE Rps. palustris HemA HYHVLLEQELAALHGKESALLFTSGYVSNWASLSTLASRMPGCVILSDELNHASMIEGIRHSRSETRIFAHNDPRDLERKLADLDPHAPKLVAFE Rps. palustris HemO HPLVQLEAEIADLHGKEAALLFTSGYVSNQTGLSTLGKLIPNCLILSDALNHNSMIEGIRQSGCERVVWRHNDTAHLEELLIAAGPDRPKLIAFE R. sphaeroides 2.4.9 HemT RQHVALEAELADLHGKESALIFTSGWISNLAALGTLGKVLPECAIFSDALNHNSMIEGIRRSGAERFIFRHNDPAHLDRLLSSVDPTRPKIVAFE M. musculus ALAS2 KFHVELEQELAELHQKDSALLFSSCFVANDSTLFTLAKLLPGCEIYSDAGNHASMIQGIRNS-AAKFVFRHNDPGHLKKLLEKSDPKTPKIVAFE H. sapiens ALAS2 KFHVELEQELAELHQKDSALLFSSCFVANDSTLFTLAKILPGCEIYSDAGNHASMIQGIRNSGAAKFVFRHNDPDHLKKLLEKSNPKIPKIVAFE H. sapiens ALAS1 KFHVDLERELADLHGKDAALLFSSCFVANDSTLFTLAKMMPGCEIYSDSGNHASMIQGIRNSRVPKYIFRHNDVSHLRELLQRSDPSVPKIVAFE PP SS PP S S S S 120

* S. xanthochromogenes SVYSMSGDVAPLAEIAELAHQYGATTFLDEVHAIGMYGPEGAGIAPREGIADRFDVIMGTLAKGLGTAGGYIAGPQDLIDAVRTLSRAFVFTTSL S. aizunensis NRRL-B-11277 SVYSMSGDIAPLAEIAELARRYDATTYIDEVHAVGMYGPQGAGIAAREGIADQFTVVMGTLAKGYGTVGGYIAGPAALVDAVRTLSRAFVFTTSL S. nodosus ssp. asukaensis SVYSMSGDIAPLAETAALARRHGATTFIDEVHAVGMYGPQGAGIAAREGIADEFTVVMGTLAKGFGTAGGYIAGPAALIDAVRNFSRGFIFTTSI S. aureus SOK1/5-04 SVYSMSGDIAPLAEIADIAKRYGASTFLDEVHAVGMYGPEGAGIAAREGIADDFTVIMGTLAKGFGTAGGYIAGPAALVDAVRSFSRSFIFTTSL S. sp. CNB091 SVYSMGGDVAPLAEIARIAREHGAMTFLDEVHAVGMYGPEGAGIAAGLGIADEFTVIMGTLAKGFGTTGGYIAGPAELVDAVRGLSRPFIFTTAL S. sp. PCS3-D2 SVYSMGGDVAPLAEIAAIARKHGAMTFLDEVHAVGMYGPQGAGIAASLGLADEFTVIMGTLAKGFGTTGGYIAGPAELVDAVRGLSRSFIFTTAL S. rapamycinicus NRLL5491 SVYSMNGDIAPLAEIADIADRHGAMTFLDEVHAVGMYGPQGAGIAAREGLADRFTVIMGTLAKGFGTNGGYIVGPAEVIEAVRMFSRSFIFTTAM S. sp. PRh5 SVYSMNGDIAPLAEIAEIAERHGAMTFLDEVHAVGMYGPQGAGIAAREGLADRFTVIMGTLAKGFGTNGGYIVGPAEVIEAVRMFSRSFIFTTAM S. ghanaensis ATCC14672 MoeA5 SVHSMRGDIALLAEIAGLAKRYGAVTFLDEVHAVGMYGPGGAGIAARDGVHCEFTVVMGTLAKAFGMTGGYVAGPAVLMDAVRARARSFVFTTAL S. clavuligerus ATCC27064 SVHSMGGDIAPLAELADLAKRYGAVTFLDEVHAVGMYGPQGAGIAAREGLASSFTVIMGTLAKGFGMTGGYVAGPAVLIDAIRAHARSFVFTTAL R. capsulatus SB1008 HemA SVYSMDGDFGPIKEICDIADEFGALTYIDEVHAVGMYGPRGAGVAERDGLMHRIDIFNGTLAKAYGVFGGYIAASAKMVDAVRSYAPGFIFSTSL R. sphaeroides 2.4.9 HemA SVYSMDGDFGQIKEICDIADEFGALKYIDEVHAVGMYGPRGGGVAERDGLMDRIDIINGTLGKAYGVFGGYIAASAKMCDAVRSYAPGFIFSTSL Rps. palustris HemA SVYSMDGDIAPIAEICDVADAHNAMTYLDEVHGVGLYGPNGGGIADREGISHRLTIIEGTLAKAFGVVGGYIAGSSAVCDFVRSFASGFIFSTSP Rps. palustris HemO SLYSMDGDIAPLSKICDLAEKYNAMTYCDEVHAVGMYGPRGAGVAERDGVMHRIDIMEATLAKAFGCLGGYIAGKAEVIDAVRSYAPGFIFTTAL R. sphaeroides 2.4.9 HemT SVYSMDGDIAPIAEICDVAERHGALTYLDEVHAVGLYGPRGGGISDRDGLADRVTIIEGTLAKAFGVMGGYVSGPSLLMDVIRSMSDSFIFTTSI M. musculus ALAS2 TVHSMDGAICPLEELCDVAHQYGALTFVDEVHAVGLYGARGAGIGERDGIMHKLDIISGTLGKAFGCVGGYIASTRDLVDMVRSYAAGFIFTTSL H. sapiens ALAS2 TVHSMDGAICPLEELCDVSHQYGALTFVDEVHAVGLYGSRGAGIGERDGIMHKIDIISGTLGKAFGCVGGYIASTRDLVDMVRSYAAGFIFTTSL H. sapiens ALAS1 TVHSMDGAVCPLEELCDVAHEFGAITFVDEVHAVGLYGARGGGIGDRDGVMPKMDIISGTLGKAFGCVGGYIASTSSLIDTVRSYAAGFIFTTSL G P PP P P* SPP REGION 3 REGION 4 (Active site loop) 316------354 358------374 | | | * | S. xanthochromogenes PPATAAGALAAVRHLRTSE--AERQSLARNAQLLHRLLDEADVPYLSSDSHIVSALVGDDALCKQASALLLDRHKIYVQAINAPSVRAGEELLRI S. aizunensis NRRL-B-11277 PPAVAAGALEAVRYLRNSD--VERKVLAENAQLLHRLLDEADIPFISPDSHIVSAFIGDDETCKQASRLLFERHGIYVQSINAPSVPLGQEILRI S. nodosus ssp. asukaensis PPATAAGALAAVQHLRASE--GERTRLAANAGLLHRLLKERDIPFVSDQSHIVSVFVGDDGLCRQASALLLERHGIYVQPINAPSVRAGEEILRV S. aureus SOK1/5-04 PPAIAAGALAGVRHLRSSD--EERDRLRDNAQLLHRLLDERNIPFVSPMSHIVSVFVGDDSLCRKASEFLLQRHGIYVQAINAPSVKAGEEILRV S. sp. CNB091 PPAVAAGALAAVRHLRTSE--EERDRLRENARLTHRLLRERGIPFLSDGSHIVSVFVGDDALARRISALLLARHGIYVQAINAPSVRVGQEILRS S. sp. PCS3-D2 PPSVVAGALAAVRHLRASD--EERDRLRENARLTHRLLTEHGIPFVSDESHIVSIFVGDDALAQRISGLLLERHGIYVQSINAPSVRAGQEILRS S. rapamycinicus NRLL5491 PPAVAAAALAAVRHLRSSE--AERERLWANAQSMHRLLKERRIPFISDLTHIVSVLVRNEALCKRMSTALLDRHGIYVQAINAPSVRAGEEILRV S. sp. PRh5 PPAVAAAALAAVRHLRSSE--AERERLWANAQLMHRLLKERRIPFISDLTHIVSVLVRNEALCKRMSTALLERHGIYVQAINAPSVRAGEEILRV S. ghanaensis ATCC14672 MoeA5 PPAVAAGALAAVRHLRGSD--EERRRPAENARLTHGLLRERDIPVLSDRSPIVPVLVGEDRMCKRMSALPLERHGAYVQAIDAPSVPAGEEILRI S. clavuligerus ATCC27064 PPSVAAGALAAVRHLRGSE--EERRRLTANASLLHGLLRERSLPFLSDQSHIVSALVGDERRAKGMSRLLLERYGIYVQAVDAPSVRVGEEILRI R. capsulatus SB1008 HemA PPAIAAGAQASIAFLKTAEGQKLRDAQQMHAKVLKMRLKALGMPIIDHGSHIVPVVIGDPVHTKAVSDMLLSDYGVYVQPINFPTVPRGTERLRF R. sphaeroides 2.4.9 HemA PPVVAAGAAASVRHLKGD--VELREKHQTAAKILKMRLKGLGLPIIDHGSHIVPVHVGDPVHCKMISDMLLEHFGIYVQPINFPTVPRGTERLRF Rps. palustris HemA PPAVAAGALASIRHLRAS--SAERERHQDRVARLRARLDQAGVAHMPNPSHIVPVMVGDAALCKQISDELISRYGIYVQPINYPTVPRGTERLRI Rps. palustris HemO PPPICAAATAAIKHLKSS--SWERERHQDRAARVKAVLNNAGIPVMPTDTHIVPVFVGDAEKCKKASDLLLEQHGIYIQPINYPTVARGLERLRI R. sphaeroides 2.4.9 HemT CPHLAAGALAAVRHVKAHP--DERRRQAENAVRLKVLLQKAGLPVLDTPSHILPVMVGEAHLCRSISEALLARHAIYVQPINYPTVARGQERLRL M. musculus ALAS2 PPMVLSGALESVRLLKGEEGQALRRAHQRNVKHMRQLLMDRGFPVIPCPSHIIPIRVGNAALNSKICDLLLSKHSIYVQAINYPTVPRGEELLRL H. sapiens ALAS2 PPMVLSGALESVRLLKGEEGQALRRAHQRNVKHMRQLLMDRGLPVIPCPSHIIPIRVGNAALNSKLCDLLLSKHGIYVQAINYPTVPRGEELLRL H. sapiens ALAS1 PPMLLAGALESVRILKSAEGRVLRRQHQRNVKLMRQMLMDAGLPVVHCPSHIIPVRVADAAKNTEVCDELMSRHNIYVQAINYPTVPRGEELLRI HEPTA-VARI | | LQAINRPTVEINQEKLR | | S G 337------352 SECOND CHANNEL 121

S. xanthochromogenes APSATHTEADVQKFAEALAGIWQELEIATASDRAAGRTSAVKELV------S. aizunensis NRRL-B-11277 APSTVHGREDVENFAEALRGIWKELNIPTATDRNWLS------S. nodosus ssp. asukaensis APSATHTTGDVEKFAEAVEGIWRDLGIPRAQERAGR------S. aureus SOK1/5-04 APSATHTSSDVEKFAEALDGIWQELDIPRTEHTRGN------S. sp. CNB091 APGAVHEPSEIEGFVTALDGIWRELGVTRAQGPHRSP------S. sp. PCS3-D2 APGAVHEPAAVAEFVAALDGIWEELGVTRAPGWRRSAVTHG------S. rapamycinicus NRLL5491 APGAVHSTSEVEMFVKALDQVWEELGGPREDAEIRLRNETHRG------S. sp. PRh5 APGAVHSTSEVEMFVKALDQVWEELGGPREDVEMRLRNGAGRG------S. ghanaensis ATCC14672 MoeA5 APSAVHETEEIHRFVDALDGIWSELGAARRV------S. clavuligerus ATCC27064 APSAVHRPEEVREFAAALDTVWTGLDGPRTTAAV------R. capsulatus SB1008 HemA TPSPVHDLKQIDGLVHAMDLLWARCALNRAEASA------R. sphaeroides 2.4.9 HemA TPSPVHDSGMIDHLVKAMDVLWQHCALNRAEVVA------Rps. palustris HemA TPSPQHTDADIEHLVQALSEIWTRVGLAKAA------Rps. palustris HemO TPSPYHDDKLIDALAEALVQVWGQLGLPLGAKAIAAE------R. sphaeroides 2.4.9 HemT TPTPFHTTSHMEALVEALLAVGRDLGWAMSRRAA------M. musculus ALAS2 APSPHHSPQMMENFVEKLLLAWTEVGLPLQDVSVAACNFCHRPVHFELMSEWERSYFGNMGPQYVTTYA H. sapiens ALAS2 APSPHHSPQMED-FVEKLLLAWTAVGLPLQDVSVAACNFCRRPVHFELMSEWERSYFGNMGPQYVTTYA H. sapiens ALAS1 APTPHHTPQMMNYFLENLLVTWKQVGLELKPHSSAECNFCRRPLHFEVMSEREKSYFSGLSKLVSAQA-

Figure 31. Multiple amino acid sequence alignment of cyclic and classical ALA synthases, together with partial sequences of heptavariant mutant mouse ALAS2 with substituted residues highlighted in cyan (30). Amino acid residues that are involved in cofactor and substrate binding (22, 23, 31) are highlighted in black and marked with a letter below the alignment representing the substrate or cofactor each residue binds to (PLP-P, glycine-G, succinyl-CoA-S). Residues of R. capsulatus spanning four flexible regions undergoing large conformational changes as the enzyme transitions from opened to closed states are shown in bold, and the R. capsulatus numbering of these residues are indicated above each block. Amino acid residue differences between classical ALA synthases and cyclic ALA synthases that are absolutely conserved among each kind of enzyme are marked with asterisks above the alignment. The residues contributing to the second channel (32) are also identified below the aligned sequences, with numbering according to R. capsulatus HemA. 122

Since other residues that are involved in cofactor and substrate binding, as identified by

Astner et al. (22), are conserved between HemA and AcaC, differences in amino acid residues not directly involved in substrate binding could be important in explaining the differences in activity between classical (HemA and HemT) and cyclizing (AcaC) ALA synthases. These residues may be important in defining, and confining the degree of protein flexibility, which has been reported to be crucial for efficient enzyme function in ALA synthases (18, 21, 22, 30, 33).

Furthermore, flexible regions in proteins that undergo conformational changes during catalysis have been reported to be important for enzyme function, as they play a role in optimal positioning of substrate and specificity (34-36). To investigate whether there are differences in protein flexibility between classical (HemA and HemT) and cyclizing (AcaC) ALA synthases, the crystal structures of R. capsulatus ALA synthase (22) and the other members of the Į- oxoamine synthase family (37-40), were examined, while also considering what is known about

ALA synthases from other studies.

The reaction mechanism of ALA synthase is reported to proceed through an induced fit mechanism whereby, in its resting state, the enzyme is in the open conformation and upon substrate binding it undergoes a conformational change into the catalytically active closed conformation (18, 21, 30, 41). After completion of the condensation reaction, the enzyme reverts to its open conformation, releasing ALA in the process. This last step is the rate limiting step in ALA synthesis (21, 22). The availability of the crystal structures of ALA synthase of R. capsulatus in open and closed conformations has enabled the identification of the flexible regions that are involved in the conformational change. These regions are all located in close proximity to the active site, as shown in Figure 32, and comprises four flexible regions made up of amino acids 10-24, 40-47, 316-354 and 358-374 (Figure 31, R. capsulatus HemA numbering). 123

Figure 32. Solved crystal structures of R. capsulatus ALA synthase in open (PDB ID 2BWN- light grey) and closed (PDB ID 2BWO-dark grey) conformations superpositioned on each other.

The RMSD score of the two structures is 0.86. The four flexible regions are colored differently.

Residues in flexible regions represented by residues 10-24 are colored red, 40-47 are colored 124 green, 316-354 are colored blue and 358-374 are colored magenta. The numbering is based on

R. capsulatus ALA synthase (A). (B) image A rotated 180º on the y-axis.

An amino acid sequence alignment of classical and cyclizing ALA synthases (Figure 31) reveals that the corresponding amino acid residues of the flexible regions in HemA and AcaC are different, supporting the hypothesis that there could be differences in protein flexibility.

However, the predicted difference in protein flexibility is not evident from the protein models, which could be due to limitations of homology modelling of flexible regions, a limitation which has been noted by others (42, 43). By way of demonstrating the problems of homology modelling of flexible regions, models of open and closed configurations of 8-amino-7- oxononanoate synthase (AONS) were generated using the crystal structures of R. capsulatus

ALA synthase as templates. Superpositioning of these models upon the solved crystal structures of AONS reveals discrepancies that are identified in Figure 33. 125

126

Figure 33. Solved crystal structures of E. coli AONS superpositioned on models of the same protein modelled using R. capsulatus ALA synthase. (A) Superpositioned crystal structures of

AONS in open (light grey) and closed conformations (dark grey) PDB ID 1DJE and 1DJ9 respectively. (B) Image of A rotated 180 degrees on the y-axis. (C) Superpositioned crystal structure of AONS in open conformation (light grey) with AONS model predicted using R. capsulatus ALA synthase in open conformation (dark grey) PDB ID 2BWN as template. The

RMSD is 1.69. (D) Image of C rotated 180 degrees on the y-axis. (E) Superpositioned crystal structure of AONS in closed conformation (light grey) with AONS model (dark grey) created using R. capsulatus ALA synthase in closed conformation PDB ID 2BWO as template. The

RMSD is 1.78. (F) Image of E rotated 180 degrees on the y-axis. The four flexible regions are colored differently. Residues in flexible regions represented by residues 10-24 are colored red,

40-47 are colored green, 316-354 are colored blue and 358-374 are colored magenta. The numbering is based on R. capsulatus ALA synthase.

To further investigate whether differences in protein flexibility between HemA and AcaC could exist, the solved crystal structures of other members of Į-oxoamine synthase family were examined. All members of the Į-oxoamine synthase family function as homodimers, are similar in structure and enzymatic mechanism, and catalyze reactions between a small amino acid and coenzyme A esters (44, 45). The induced fit mechanism is also conserved within this protein family (46), and the closed conformation is reported to be important in determining substrate specificity (44). Superpositioning of the crystal structure of R. capsulatus ALA synthase in closed conformation with the available crystal structures of other members of Į-oxoamine synthase family in their closed conformations shows that there are differences in the flexible 127 regions, especially within the flexible region encompassing residues 357-373 (R. capsulatus

ALA synthase numbering), as shown in Figure 34. These differences might be necessary in order to accommodate differences in the sizes of the substrates, especially the acyl-CoA substrate, which is responsible for inducing the conformational change from open to closed conformation that optimally orients the substrates for proper catalysis (21, 41). 128

129

Figure 34. Crystal structures of the members of Į-oxoamine synthase family (A-AONS, B-SPT,

C-KBL, D-CsqA) in their closed conformation superpositioned with the crystal structure of R. capsulatus ALA synthase crystal structure in closed conformation (PDB ID 2BWO). (A)

Superpositioned crystal structures of AONS (dark grey) PDB ID 1DJ9. (B) Image of A rotated

180 degrees on the y-axis. (C) Superpositioned crystal structures of SPT (dark grey) PDB ID

2JG2. (D) Image of C rotated 180 degrees on the y-axis. (E) Superpositioned crystal structures of KBL (dark grey) PDB ID 1FC4. (F) Image of E rotated 180 degrees on the y-axis. (G)

Superpositioned crystal structures of CsqA (dark grey) PDB ID 2WKA. (H) Image of A rotated

180 degrees on the y-axis. Residues in flexible regions represented by residues 10-24 are colored red, 40-47 are colored green, 316-354 are colored blue and 358-374 are colored magenta.

The numbering is based on R. capsulatus ALA synthase.

With respect to the ALA synthesis and ALA-CoA cyclization reactions, the acyl-CoA substrates are different in size, and they also are thought to differ in the way they are oriented in the active site, as shown in Figs. 29 and 30. These differences may require that the enzymes undergo induced conformational changes that are specific for each reaction such that the substrates are optimally oriented in the active site. It is possible that AcaC has evolved to undergo a conformational change triggered by both succinyl-CoA and ALA-CoA, which then allows it to catalyze both ALA formation and ALA-CoA cyclization reactions that are necessary to synthesize C5N units. In the case of HemA or HemT, which are dedicated ALA synthases, substrate binding causes a conformational change that is tailored towards maximal ALA synthesis in order to meet the ALA demand for tetrapyrrole biosynthesis, but which is not optimal for ALA-CoA cyclization. Recent reports by Lendrihas et al. (30) and Stovanojski et al.

(32) on analysis of the ALA synthase structure and enzyme activity offer support to the idea that 130 amino acid sequence differences in flexible regions are important in distinguishing ALA synthase and ALA-CoA cyclizing activities.

- Lendrihas et al. (30) have reported that the amino acid composition of the flexible region encompassing residues 357-373 (colored magenta in Figure 32) is important for ALA synthase activity (30). They found that murine ALAS2 with amino acid substitutions within this flexible region had increased ALA synthase activity. This flexible region is located above the active site of the enzyme, and its movement allows the opening and closing of the active site as shown in

Figure 32 (22, 30). For one mutant, having seven substitutions in this region (Figure 31), the kcat is 15-fold relative to the wild type. Molecular dynamics analysis of this heptavariant by Na et al.

(33) revealed that the active site loop of the mutant protein is less flexible, and the mutant was also found to adopt a more stable open conformation state compared to the wild type. As a consequence, it reverted back from the closed to the open conformation more quickly than the wild type protein, and so the rate of product release was higher. There are amino acid sequence differences between HemA and AcaC in this region (Fig. 31) that might account for difference in activity.

- Stovanojski et al. (32) identified a second channel separate from the one occupied by succinyl-

CoA in the solved crystal structure of R. capsulatus ALA synthase. This channel is only visible when the protein is in the open conformation, and it is believed to be important for glycine entry or ALA exit (32). The residues at positions 83 and 363, serines in AcaC and other cyclases, and threonines in HemA and other ALA synthases, are located at the bottom of this second channel, and so would play no role in the architecture of the channel that would affect its function.

However, the second channel is formed by different parts of the polypeptides of the dimeric enzyme that include residues 1-24, 77-89, and 337-352. Two of the flexible regions, 10-24 and 131

316- 354 form part of this second channel, and so, if the amino acid sequence differences between AcaC and HemA in these regions give rise to differences in protein flexibility, the function of this second channel would also be affected. Indeed, the high Km of AcaC for glycine might be due to more restricted access of glycine into the active site, and the low kcat of the enzyme caused by slow release of product. According to the protein models (Figure 35), there are no differences in the second channel which could be due to limitations of homology modelling.

132

Figure 35. Second channel of the predicted tertiary structures of HemA (A and B) and AcaC (C and D). Regions of the protein that form part of the second channel are colored differently with the region encompassing residues 1-24 colored cyan, residues 77-89 (from the second subunit) colored magenta and residues 337-352 colored red. In (A) the location of threonines 83 and 363 is shown in blue. 133

In conclusion, the results of this investigation have confirmed the hypothesis by Zhang et al. (16) that classical ALA synthases are capable of ALA-CoA cyclization to form the C5N unit.

Although important for activity in the respective enzymes, conserved sequence differences at positions 83 and 363 are not solely responsible for the low ALA synthase activity of AcaC and low ALA-CoA cyclase activity of HemA. Further analysis of the amino acid sequence composition of HemA and AcaC and the crystal structures of the members of Į-oxoamine synthase family of the fold type I PLP-dependent enzymes, based on what is known about ALA synthesis reaction mechanism suggests that differences in active site architecture, protein flexibility and active site accessibility via a proposed second channel may also be important in explaining the differences in activity between classical and cyclizing ALA synthases.

134

References

1. Hopwood DA, Sherman DH. 1990. Molecular genetics of polyketides and its comparison to fatty acid biosynthesis. Annu Rev Genet 24:37-66.

2. Shimizu K, Tamura G. 1981. Reductiomycin, a new antibiotic. I. Taxonomy, fermentation, isolation, characterization and biological activities. J Antibiot (Tokyo) 34:649-653.

3. Zeeck A, Frobel K, Heusel C, Schroder K, Thiericke R. 1987. The structure of manumycin. II. Derivatives. J Antibiot (Tokyo) 40:1541-1548.

4. McAlpine JB, Bachmann BO, Piraee M, Tremblay S, Alarco AM, Zazopoulos E,

Farnet CM. 2005. Microbial genomics as a guide to drug discovery and structural elucidation:

ECO-02301, a novel antifungal agent, as an example. J Nat Prod 68:493-496.

5. Hanajima S, Ishimaru K, Sakano K, Roy SK, Inouye Y, Nakamura S. 1985.

Inhibition of reverse transcriptase by limocrocin. J Antibiot (Tokyo) 38:803-805.

6. Nakano H, Yoshida M, Shirahata K, Ishii S, Arai Y, Morimoto M, Tomita F. 1982.

Senacarcin A, a new antitumor antibiotic produced by Streptomyces endus subsp. aureus. J

Antibiot (Tokyo) 35:760-762.

7. Hayashi K, Nakagawa M, Fujita T, Tanimori S, Nakayama M. 1994. Nisamycin, a new manumycin group antibiotic from Streptomyces sp. K106. II. Structure determination and structure-activity relationships. J Antibiot (Tokyo) 47:1110-1115.

8. Hinzen B, Raddatz S, Paulsen H, Lampe T, Schumacher A, Habich D, Hellwig V,

Benet-Buchholz J, Endermann R, Labischinski H, Brotz-Oesterhelt H. 2006. Medicinal chemistry optimization of acyldepsipeptides of the enopeptin class antibiotics. ChemMedChem

1:689-693. 135

9. Laakso JA, Mocek UM, Van Dun J, Wouters W, Janicot M. 2003. R176502, a new bafilolide metabolite with potent antiproliferative activity from a novel Micromonospora species.

J Antibiot (Tokyo) 56:909-916.

10. Werner G, Hagenmaier H, Drautz H, Baumgartner A, Zahner H. 1984. Metabolic products of microorganisms. 224. Bafilomycins, a new group of macrolide antibiotics.

Production, isolation, chemical structure and biological activity. J Antibiot (Tokyo) 37:110-117.

11. Nakagawa A, Wu TS, Keller PJ, Lee JP, Omura S, Floss HG. 1985. Biosynthesis of asukamycin. Formation of the 2-amino-3-hydroxycyclopent-2-enone moiety. J Chem Soc, Chem

Commun 1985(8):519-521.

12. Kikuchi G, Kumar A, Talmage P, Shemin D. 1958. The enzymatic synthesis of delta- aminolevulinic acid. J Biol Chem 233:1214-1219.

13. Schulze JO, Schubert W-D, Moser J, Jahn D, Heinz DW. 2006. Evolutionary relationship between initial enzymes of tetrapyrrole biosynthesis. J Mol Biol 358:1212-1220.

14. Yang H-S, Hoober JK. 1995. Divergent pathways for G-aminolevulinic acid synthesis in two species of Arthrobacter. FEMS Microbiol Lett 134:259-263.

15. Petrícek M, Petríceková K, Havlícek L, Felsberg J. 2006. Occurrence of two 5- amionolevulinate biosynthetic pathways in Streptomyces nodosus subsp. asukaensis is linked to the production of asukamycin. J Bacteriol 188:5113-5123.

16. Zhang W, Bolla ML, Kahne D, Walsh CT. 2010. A three enzyme pathway for 2- amino-3-hydroxycyclopent-2-enone formation and incorporation in natural product biosynthesis.

J Am Chem Soc 132:6402-6411. 136

17. Petrícková K, Chronáková AZ, T., Chrudimský T, Pospíšil S, Petrícek M, Krištufek

V. 2015. Evolution of cyclizing 5-aminolevulinate synthases in the biosynthesis of actinomycete secondary metabolites: outcomes for genetic screening techniques. Front Microbiol 6:814-829.

18. Hunter GA, Zhang J, Ferreira GC. 2007. Transient kinetic studies support refinements to the chemical and kinetic mechanisms of aminolevulinate synthase. J Biol Chem 282:23025-

23035.

19. Nandi DL. 1978. Studies on delta-aminolevulinic acid synthase of Rhodopseudomonas spheroides. Reversibility of the reaction, kinetic, spectral, and other studies related to the mechanism of action. J Biol Chem 253:8872-8877.

20. Zaman Z, Jordan PM, Akhtar M. 1973. Mechanism and stereochemistry of the 5- aminolaevulinate synthetase reaction. Biochem J 135:257-263.

21. Hunter GA, Ferreira GC. 1999. Pre-steady-state reaction of 5-aminolevulinate synthase. Evidence for a rate-determining product release. J Biol Chem 274:12222-12228.

22. Astner I, Schulze JO, van den Heuvel J, Jahn D, Schubert W-D, Heinz DW. 2005.

Crystal structure of 5-aminolevulinate synthase, the first enzyme of heme biosynthesis, and its link to XLSA in humans. EMBO J 24:3166-3177.

23. Ferreira GC, Neame PJ, Dailey HA. 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.

24. Shoolingin-Jordan PM, Al-Daihan S, Alexeev D, Baxter RL, Bottomley SS, Kahari

ID, Roy I, Sarwar M, Sawyer L, Wang SF. 2003. 5-Aminolevulinic acid synthase: mechanism, mutations and medicine. Biochim Biophys Acta 1647:361-366. 137

25. Kaufholz AL, Hunter GA, Ferreira GC, Lendrihas T, Hering V, Layer G, Jahn M,

Jahn D. 2013. Aminolaevulinic acid synthase of Rhodobacter capsulatus: high-resolution kinetic investigation of the structural basis for substrate binding and catalysis. Biochem J

451:205-216.

26. Rui Z, Petrícková K, Škanta F, Pospíšil S, Yang Y, Chen CY, Tsai SF, Floss HG,

Petrícek M, Yu TW. 2010. Biochemical and genetic insights into asukamycin biosynthesis. J

Bio Chem 32:24915-24924.

27. Eraso JM, Kaplan. S. 1994. prrA, a putative response regulator involved in oxygen regulation of photosynthesis gene expression in Rhodobacter sphaeroides. J Bacteriol 176:32-43.

28. Ind AC, Porter SL, Brown MT, Byles ED, de Beyer JA, Godfrey SA, Armitage JP.

2009. Inducible-expression plasmid for Rhodobacter sphaeroides and Paracoccus denitrificans.

Appl Environ Microbiol 75:6613-6615.

29. Schneider CA, Rasband WS, Eliceiri KW. 2012. NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9:671-675.

30. Lendrihas T, Hunter GA, Ferreira GC. 2010. Targeting the active site gate to yield hyperactive variants of 5-aminolevulinate synthase. J Biol Chem 285:13704-13711.

31. Hunter GA, Ferreira GC. 1999. Lysine-313 of 5-aminolevulinate synthase acts as a general base during formation of the quinonoid reaction intermediates. Biochemistry 38:3711-

3718.

32. Stojanovski BM, Hunter GA, Jahn M, Jahn D, Ferreira GC. 2014. Unstable reaction intermediates and hysteresis during the catalytic cycle of 5-aminolevulinate synthase: implications from using pseudo and alternate substrates and a promiscuous enzyme variant. J

Biol Chem 289:22915-22925. 138

33. Na I, DeForte S, Stojanovski BM, Ferreira GC, Uversky VN. 2017. Molecular dynamics analysis of the structural and dynamic properties of the functionally enhanced heptavariant of mouse 5-aminolevulinate synthase. J Biomol Struct Dyn doi:10.1080/07391102.2016.1269688.

34. Gunasekaran K, Nussinov R. 2007. How different are structurally flexible and rigid binding sites? Sequence and structural features discriminating proteins that do and do not undergo conformational change upon ligand binding. J Mol Biol 365:257-273.

35. Hammes-Schiffer S, Benkovic SJ. 2006. Relating protein motion to catalysis. Annu Rev

Biochem 75:519-541.

36. Hanson JA, Duderstadt K, Watkins LP, Bhattacharyya S, Brokaw J, Chu JW, Yang

H. 2007. Illuminating the mechanistic roles of enzyme conformational dynamics. Proc Natl Acad

Sci U S A 104:18055-18060.

37. Jahan N, Potter JA, Sheikh MA, Botting CH, Shirran SL, Westwood NJ, Taylor

GL. 2009. Insights into the biosynthesis of the Vibrio cholerae major autoinducer CAI-1 from the crystal structure of the PLP-dependent enzyme CqsA. J Mol Biol 392:763-773.

38. Schmidt A, Sivaraman J, Li Y, Larocque R, Barbosa JA, Smith C, Matte A, Schrag

JD, Cygler M. 2001. Three-dimensional structure of 2-amino-3-ketobutyrate CoA ligase from

Escherichia coli complexed with a PLP-substrate intermediate: inferred reaction mechanism.

Biochemistry 40:5151-5160.

39. Alexeev D, Alexeeva M, Baxter RL, Campopiano DJ, Webster SP, Sawyer L. 1998.

The crystal structure of 8-amino-7-oxononanoate synthase: a bacterial PLP-dependent, acyl-

CoA-condensing enzyme. J Mol Biol 284:401-419. 139

40. Yard BA, Carter LG, Johnson KA, Overton IM, Dorward M, Liu H, McMahon SA,

Oke M, Puech D, Barton GJ, Naismith JH, Campopiano DJ. 2007. The structure of serine palmitoyltransferase; gateway to sphingolipid biosynthesis. J Mol Biol 370:870-886.

41. Zhang J, Ferreira GC. 2002. Transient state kinetic investigation of 5-aminolevulinate synthase reaction mechanism. J Biol Chem 277:44660-44669.

42. Dobbins SE, Lesk VI, Sternberg MJ. 2008. Insights into protein flexibility: The relationship between normal modes and conformational change upon protein-protein docking.

Proc Natl Acad Sci U S A 105:10390-10395.

43. Sanchez R, Pieper U, Melo F, Eswar N, Marti-Renom MA, Madhusudhan MS,

Mirkovic N, Sali A. 2000. Protein structure modeling for structural genomics. Nat Struct Biol 7

Suppl:986-990.

44. Eliot AC, Kirsch JF. 2004. Pyridoxal phosphate enzymes: mechanistic, structural, and evolutionary considerations. Annu Rev Biochem 73:383-415.

45. Percudani R, Peracchi A. 2003. A genomic overview of pyridoxal-phosphate-dependent enzymes. EMBO Rep 4:851-854.

46. Lendrihas T, Hunter GA, Ferreira GC. 2010. Serine 254 enhances an induced fit mechanism in murine 5-aminolevulinate synthase. J Biol Chem 285:3351-3359.

140

CHAPTER IV

SUMMARY AND CONCLUSIONS

ALA synthase is a PLP-dependent enzyme that functions as a homodimer with two active sites located at the interface of the subunits (1, 2). While knowledge of the reaction mechanism of ALA synthase catalysis has advanced considerably, other questions about the enzyme have remained unanswered. All animals and many bacterial have more than one ALA synthase gene.

The need for multiple genes, as opposed to regulated expression of a single gene, suggests that the enzymes themselves have distinctive properties that are necessary for the organism. This question has been investigated for ALA synthase isoenzymes in Rhodobacter sphaeroides.

Three HemA and two HemT enzymes were examined. All were found to have similar specific activities and kinetic properties. However, the HemA enzymes are more sensitive to hemin inhibition than HemT. Interestingly, the UV-vis spectra of purified HemA is indicative of protein bound heme while that of HemT is only indicative of protein bound PLP. This suggests that HemA has high affinity for heme binding compared to HemT, which could further explain the difference in hemin sensitivity between the two enzymes. This differential sensitivity toward hemin inhibition suggests HemT is necessary for augmenting ALA formation under conditions where HemA activity is inhibited. In strain 2.4.9 hemT expression is upregulated under anaerobic dark growth with DMSO. Unlike aerobic respiration, which relies upon molecular components having the A form of heme, anaerobic respiration uses components with heme B. If inhibition is limited to the B form of heme, HemA would be fully active aerobically, but reduced in activity under anaerobic conditions. It may be that the presence of the farnesyl group on heme

A, which is similar to the hydrophobic tail present in bacteriochlorophyll a (Figure 36) that is 141 known to be non-inhibitory to ALA synthase activity (8), prevents binding of the heme to

HemA. This remains to be determined.

Figure 36. The structures of heme B, heme A and bacteriochlorophyll. The farnesyl group of heme A and the hydrophobic tail of bacteriochlorophyll are shown in rectangular boxes.

Purified HemT enzyme was found to be sensitive to oxidation, which was determined to be due to oxidation of cysteine 281 (R. sphaeroides HemT numbering). This raises the possibility that the enzyme is designed to function exclusively under anaerobic conditions, which would be consistent with its transcription profile; the gene is maximally expressed under anaerobic-dark conditions. Cysteine 281 from the two subunits are located at the subunit interface and are in close proximity to each other, suggesting they could form a disulfide bond under aerobic conditions that potentially limits the movement of the two subunits necessary for catalysis, leading to decreased ALA synthase activity.

Another outstanding question pertains to the recently discovered bifunctionality of certain

ALA synthases. In certain actinomycetes, the enzyme synthesizes ALA, but also catalyzes the cyclization of ALA-CoA to form C5N units that are part of the structure of certain secondary 142 metabolites (3). Whether all ALA synthases can catalyze both reactions was not yet known.

ALA synthase activities and the relative ALA-CoA cyclization activities of HemA, HemT and a bona fide bifunctional enzyme, AcaC, were examined. Although all enzymes can catalyze the

ALA-CoA cyclization reaction, the ALA-CoA cyclization activity of HemA and HemT was only

10% and 6% that of AcaC respectively, while their specific ALA synthase activities were 315- fold higher than that of AcaC. Hence, it seems that AcaC and other cyclizing ALA synthases have evolved to be better at catalyzing the ALA-CoA cyclization reaction compared to the ALA synthesis reaction, and to catalyze both reactions with efficiencies that allow them to meet the demand for C5N unit for secondary metabolite synthesis. The high catalytic efficiency (kcat/Km) of AcaC for ALA-CoA compared to succinyl-CoA as reported by Zhang et al. (9) could arise from a need for the enzyme to preferentially catalyze the ALA-CoA cyclization reaction when

ALA-CoA is available at the active site. When ALA-CoA is converted to a C5N unit the active site can bind glycine and succinyl-CoA and synthesize ALA that is then directed to the acyl-CoA ligase. This potentially allows the flow of metabolites toward the C5N unit synthesis pathway.

Although they were found to be important for activity in the case of each enzyme, reciprocal replacement of conserved amino acid residue differences at positions 83 and 363 (R. sphaeroides HemA numbering) revealed that they are not solely responsible for the low ALA synthase activity reported for AcaC or the low ALA-CoA cyclization activity of HemA. Further analysis of the available structures of other members of Į-oxoamine synthase family, in combination with protein modelling, indicated that differences in conformational changes and active site architecture could be responsible or important. These differences are currently not able to be demonstrated structurally due to the limitations of homology modelling as have been reported elsewhere (10, 11). It could be that the conformational flexibility of cyclizing ALA 143 synthases allows them to optimally catalyze the ALA-CoA cyclization reaction while at the same time, when ALA-CoA is not available, they are able to catalyze the ALA synthesis reaction.

The high ALA-CoA cyclizing efficiency of AcaC compared to the condensation of succinyl-CoA and glycine (9) suggests that there are differences in the amino acid residues that stabilize both the enzyme substrate complex (ES) and the ES at the transition state (ES‡) or residues that only stabilize the ES‡ only. These residues are known to influence the catalytic efficiency of an enzyme (12). Since the two amino acid residues that were analyzed in this study are the only conserved differences among those enzymes that are known to preferentially bind glycine and succinyl-CoA (1), other residues are likely to be important for the ALA synthesis reaction that have not been discovered yet, and these residues also influence the ALA-CoA cyclization reaction. Resolving the evolutionary changes that have led to the dual activity of cyclizing ALA synthases would be important in understanding the structure-function relationship of cyclizing ALA synthases, and in the process also help in further delineating the reaction mechanism of ALA synthases. These changes might be identified through reciprocal exchange of sections of classical and cyclizing ALA synthases. Then individual amino acid residues within the relevant sections can be targeted for further analysis. Alternatively, cyclizing ALA synthases in actinomycetes are reported to have evolved from a classical ALA synthase that was acquired through horizontal gene transfer. ALA synthase genes that are found in actinomycetes that do not produce C5N unit-containing secondary metabolites are thought to represent the original ALA synthase genes that were acquired during this horizontal gene transfer (3). Such genes might be in various stages of evolving into cyclizing ALA synthases and may be useful in identifying the important amino acid residues. Along these lines, examining their ALA synthase activities in future would be informative. 144

The exclusive supply of ALA for C5N unit synthesis by cyclizing ALA synthase raises the question of intracellular trafficking of ALA, since actinomycetes synthesize ALA for tetrapyrrole biosynthesis through the C5 pathway, which is also active in these bacteria. It may be that the ALA formed by the C5 pathway is not freely available in the cell, and is directly channeled to tetrapyrrole synthesis through a close association between glutamyl-1- semialdehyde aminotransferase and porphobilinogen synthase. Another possibility is that association between the cyclizing ALA synthase and acyl-CoA ligase enzymes is such that the acyl-CoA ligase only accepts substrate from the cyclizing ALA synthase.

145

References

1. Astner I, Schulze JO, van den Heuvel J, Jahn D, Schubert W-D, Heinz DW. 2005.

Crystal structure of 5-aminolevulinate synthase, the first enzyme of heme biosynthesis, and its link to XLSA in humans. EMBO J 24:3166-3177.

2. Tan D, Ferreira GC. 1996. Active site of 5-aminolevulinate synthase resides at the subunit interface. Evidence from in vivo heterodimer formation. Biochemistry 35:8934-8941.

3. Petrícková K, Chronáková AZ, T., Chrudimský T, Pospíšil S, Petrícek M, Krištufek

V. 2015. Evolution of cyclizing 5-aminolevulinate synthases in the biosynthesis of actinomycete secondary metabolites: outcomes for genetic screening techniques. Front Microbiol 6:814-829.

4. Tai T-N, Moore MD, Kaplan S. 1988. Cloning and characterization of the 5- aminolevulinate synthase gene(s) from Rhodobacter sphaeroides. Gene 70:139-151.

5. Riddle RD, Yamamoto M, Engel JE. 1989. Expression of G-aminolevulinate synthase in avian cells: separate genes encode erythroid-specific and nonspecific isozymes. Proc Natl

Acad Sci USA 86:792-796.

6. Nereng KS, Kaplan S. 1999. Genomic complexity among strains of the facultative photoheterotrophic bacterium Rhodobacter sphaeroides. J Bacteriol 181:1684-1688.

7. Coulianos N. 2011. A comparison of ALA synthase gene transcription in three wild type strains of Rhodobacter sphaeroides. MS Thesis, Bowling Green State University.

8. Burnham BF, Lascelles J. 1963. Control of porphyrin biosynthesis through a negative feedback mechanism. Biochem J 87:462-472.

9. Zhang W, Bolla ML, Kahne D, Walsh CT. 2010. A three enzyme pathway for 2- amino-3-hydroxycyclopent-2-enone formation and incorporation in natural product biosynthesis.

J Am Chem Soc 132:6402-6411. 146

10. Dobbins SE, Lesk VI, Sternberg MJ. 2008. Insights into protein flexibility: The relationship between normal modes and conformational change upon protein-protein docking.

Proc Natl Acad Sci U S A 105:10390-10395.

11. Sanchez R, Pieper U, Melo F, Eswar N, Marti-Renom MA, Madhusudhan MS,

Mirkovic N, Sali A. 2000. Protein structure modeling for structural genomics. Nat Struct Biol 7

Suppl:986-990.

12. Fersht AR. 1995. Enzyme Structure and Mechanism. W. H. Freeman and Company,

New York.