PURIFICATION AND CHARACTERIZATION OF SPHAEROIDES POLYHISTIDINE-TAGGED HEMA AND COMPARISON WITH PURIFIED POLYHISTIDINE- TAGGED HEMT

Xiao Xiao

A Thesis

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

MASTER OF SCIENCE

August 2013

Committee:

Dr. Jill Zeilstra-Ryalls, Ph.D., Advisor

Dr. Rogers O. Scott

Dr. Zhaohui Xu

ii

© 2013

Xiao Xiao

All Rights Reserved

iii

ABSTRACT

Jill Zeilstra-Ryalls, Ph.D, Advisor

All , molecules that include , , and , are derived from 5-aminolevulinic acid (ALA). In the purple non-sulfur

Rhodobacter sphaeroides ALA is formed by the condensation of glycine and succinyl-CoA, catalyzed by the pyridoxal-phosphate dependent enzyme ALA synthase. Two ALA synthase genes, hemA and hemT are present in R. sphaeroides wild type strain 2.4.1. When expressed, either one of the gene products can satisfy the ALA requirement of the cell. Towards understanding the presence of two ALA synthases in one organism, each enzyme should be characterized individually in order to define what is similar and different about the enzymes.

Using this information, one may be able to infer how the activities of the two ALA synthases are coordinate in R. sphaeroides. In this study, R. sphaeroides 2.4.1 recombinant polyhistidine- tagged HemA (rHemA) was affinity purified and its optimum temperature and pH, specific activity, and kinetic properties were determined. The effect of added hemin on its activity was also evaluated, as was its secondary structure composition using circular dichroism. These characteristics were then compared to those of recombinant polyhistidine-tagged HemT (rHemT).

Two major differences were noted. First, the catalytic capacity of rHemA is more than ten times greater than rHemT. Second, rHemA has a higher affinity for succinyl-CoA than rHemT. A hypothesis that could explain the significance of these differences posits that HemT is needed to reduce ALA synthesis under conditions in which succinyl-CoA and glycine are in greater demand for other than ALA formation, such as energy generation for the former and protein synthesis for the latter. Examining hemA and hemT expression under conditions that would impose such metabolic priorities would be appropriate to test this hypothesis. iv

ACKNOWLEDGEMENTS

Foremost, I would like to express my sincere appreciation to my advisor, Dr. Jill Zeilstra-

Ryalls, for the opportunity to work in her laboratory. She offered huge support on my two years master program. I learn extensive research experience and became thinking critically because of her excellent guidance. Besides, I would like to thank the rest of my thesis committee, Dr. Scott

O. Rogers and Dr. Zhaohui Xu for their warm encouragement and insightful comments. I would also thank Dr. Maneewan Suwansaard and my workmate James for their help with my experiments. v

TABLE OF CONTENTS

Page

CHAPTER I: BACKGROUND INFORMATION AND SPECIFIC AIMS ……………….. 1

Introduction……………………………………………………………………………... 1

A. ………………………………………………………… 1

B. ………………………………………………………………….……. 1

C. 5-Aminolevulinic acid (ALA) synthesis pathway…………………………..…….... 2

D. ALA synthases and its isozymes ………………………………………..…………. 4

Specific Aims…………………………………………………………………...... 5

CHAPTER II: PURIFICATION AND CHARACTERIZATION OF POLYHISTIDINE-

TAGGED HEMA …………………………………………………………………………….. 7

Introduction………………………………………….………………………………...... 7

Materials and Methods………………………………………………………………….... 7

Bacterial strains, , and growth conditions………………………………….. 7

Crude lysate preparation and rHemA affinity purification……………………………. 9

Protein concentration determinations……………………………...…………………. 10

ALA synthase activity assays……………………………….………………………... 11

Determining pH and temperature optima of purified rHemA………….…………….. 12

Hemin addition analysis………………………………………..……………………… 12

Determination of Km, kcat, Vmax values for succinyl-CoA and glycine of rHemA……. 12

SDS-PAGE, immunoblot analysis, and InvisionTM in-gel stain…………...... …. 13

Circular dichroism spectrometry and analysis……………………………………… 14

EKMaxTM treatment of rHemA…………………………………………….……..... 14 vi

Protein modeling …………………………………………………………………….. 15

Results………………………………………………….…………...... 15

Expression and purification of rHemA……………………………….…………..…... 15

Immunoblot analysis and in-gel polyhistidine staining of purified rHemA…...……… 18

Temperature and pH optima of purified rHemA…………………………….……….. 19

The effect of hemin on the ALA synthase activity of rHemA……………………… 20

Glycine and succinyl-CoA Km determinations for rHemA…………………………… 21

Circular dichroism (CD) analysis of rHemA.……………………………………...…. 22

Discussion……………………………………..……………………………………...... 23

CHAPTER III: A COMPARISON OF THE PROPERTIES OF POLYHISTIDINE-TAGGED

HEMA TO THOSE OF POLYHISTIDINE-TAGGED HEMT…………………..………….. 25

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

Differences and similarities between purified rHemA and rHemT………………...... 26

REFERENCES……………………………………………………………………………….. 30

vii

LIST OF FIGURES

FIGURES Page

1 Schematic diagram of tetrapyrrole synthesis in R. sphaeroides. ……………………... 2

2 The two biosynthesis pathways by which ALA is formed. ………………………….. 3

3 Schematic diagram (A) of the vector pIND-hemA and amino acid sequence (B)

of the product……………………………………………………….……………… 9

4 SDS-PAGE of the soluble and insoluble fractions of samples.………………………. 16

5 SDS-PAGE of samples collected during affinity purification of rHemA……………. 17

6 The amount of ALA formed per mg of protein versus assay incubation time……….. 18

7 Immunoblot, and InVisionTM in-gel staining of rHemA…………………………...… 19

8 Effect of temperature and pH on rHemA activity……...…………………………… 20

9 Effect of hemin on rHemA activity. …………………………………………………. 20

10 ALA synthase activity versus substrate concentrations, and nonlinear regression

plots of the data used to determine glycine and succinyl-CoA Kms …………………. 21

11 Preliminary circular dichroic spectrum of purified rHemA.……………………….… 22

12 Amino acid sequence alignment of R. sphaeroides 2.4.1 wild type HemA and HemT

generated using ClustalW ………………………………………………………...... 25

13 Tertiary structure model superpositions of R. sphaeroides 2.4.1 rHemA and wild type

HemA upon the solved crystal structure of ALA………….. 26

viii

LIST OF TABLES

TABLES Page

1 Purification profile of rHemA………………………..……………………………… 17

2 Kinetic parameters for rHemA .………………………………………………..….. 22

3 Predicted, calculated, or known secondary structure compositions of R. sphaeroides

rHemA, wild type HemA, and Rhodobacter capsulatus ALA synthase……………. 23

4 Comparison of the kinetic properties of rHemA and rHemT…….………………….. 27

ix

LIST OF ABBREVIATIONS

ALA 5-aminolevulinic acid

ALAS 5-aminolevulinic acid synthase

ATP adenosine triphosphate

CD circular dichroism

Cys cysteine

Da dalton

HCl hydrochloric acid

IPTG Isopropyl β-D-1-thiogalactopyranoside kDa kilodalton

Kn kanamycin

LB Luria-Bertani

M molar

ml milliliter

mM millimolar

min minute

nm nanometer

OD optical density

PLP pyridoxal 5’-phosphate

psi pounds per square inch

rHemA recombinant polyhistidine-tagged HemA

rHemT recombinant polyhistidine-tagged HemT

SDS sodium dodecyl sulfate x

TCA trichloroacetic acid

Trp tryptophan

Tyr tyrosine

β-ME β-mercaptoethanol

ε molar extinction coefficient

µg micrograms

µl microliter

1

CHAPTER I: BACKGROUND INFORMATION AND SPECIFIC AIMS

INTRODUCTION

A – Rhodobacter sphaeroides

R. sphaeroides belongs to the Class alpha- which has been isolated from

both fresh and salt water throughout the world (1, 2). These remarkably metabolically versatile

are capable of production (3), atmospheric nitrogen and carbon dioxide

fixation, and can obtain energy through aerobic respiration, or anoxygenic

(reviewed in ref. 4). Because of their versatility they have served as model

organisms in both applied and basic research. Such studies include exploring ways to exploit

their useful metabolic capabilities and investigating how such wide-ranging metabolisms are

efficiently orchestrated.

B – Tetrapyrroles

Tetrapyrroles, are composed of four rings that are connected with each other by a

methane or methylene unit. Rhodobacter sphaeroides is capable of synthesizing three kinds of

biologically relevant cyclic tetrapyrroles, heme, bacteriochlorophyll, and vitamin B12, via a branching biosynthetic pathway (reviewed in ref. 5; Fig. 1). These molecules play vital roles in various catabolic and anabolic biological processes in bacteria. Electron transferring are contained in many proteins like cytochromes that are involved in both aerobic and photosynthetic electron transport chains (reviewed in ref. 6). Bacteriochlorophyll is responsible for light-harvesting in the photosynthesis system of R. sphaeroides. Vitamin B12 is an essential

cofactor for many important anabolic enzymes, such as methionine synthase (reviewed in ref. 7).

As could be predicted from their roles in these bacteria, the production of these different 2

tetrapyrroles is tightly regulated and their relative and absolute amounts change significantly,

according to environmental conditions. For example, when light is present under anaerobic

conditions, R. sphaeroides synthesizes a large amount of bacteriochlorophyll to support

anoxygenic photosynthesis, while under aerobic conditions bacteriochlorophyll synthesis

blocked because when light is also present, it has the potential to generate toxic singlet

(5).

Figure 1. Schematic diagram of tetrapyrrole synthesis in R. sphaeroides. Highlighted according to their light absorbing properties are the biologically important endproducts of the branching pathway; heme (red), bacteriochlorophyll (green), and vitamin B12 (grey). Also highlighted (blue) is the first enzyme in all tetrapyrrole production, ALA synthase. In this organism, both HemA and HemT proteins are ALA synthases.

C – 5-Aminolevulinic acid (ALA) synthesis pathway

ALA is the precursor molecule in the synthesis of all tetrapyrroles (8). The synthesis of

ALA occurs via two completely different pathways (Fig. 2). In animals, fungi, and alphaproteobacteria, ALA is produced by the C4 pathway or Shemin pathway, while in algae, 3

archaea, , and all other bacteria, ALA synthesis is produced by the C5 pathway (8). In the

C4 (Shemin) pathway, a condensation reaction between glycine and succinyl-CoA is catalyzed

by the pyridoxal 5’-phosphate (PLP)-dependent enzyme ALA synthase. Initially, PLP is covalently bound to a specific lysine of ALA synthase. Then the amino group of glycine forms a

Schiff base linkage to the aldehyde of PLP. ALA is formed through the condensation between the PLP-glycine Schiff base complex and succinyl-CoA (reviewed in ref. 9). The C5 pathway is

composed of three reactions. The first reaction is the activation of glutamate by ligating it to

tRNAGlu, which is catalyzed by the same glutamyl-tRNA synthetase as is used in translation, and

this reaction is ATP and Mg2+ dependent. The second step is the reduction of the glutamate to a

semialdehyde, catalyzed by glutamyl-tRNA reductase. The final step is the rearrangement of the

semialdehyde to form ALA, which is catalyzed by PLP-dependent glutamate 1-semialdehyde

aminotransferase (reviewed in ref. 10). The differences in the biochemistries of these two pathways have been of interest from an evolutionary perspective. Furthermore, unlike the C4 pathway, the C5 pathway is directly linked to energy . Additionally, the metabolites from which ALA is derived in the two pathways are very different and so ALA production is connected to different processes, according to which pathway is present in the cell.

4

Figure 2. The two biosynthesis pathways by which ALA is formed. The C4 (Shemin) pathway (A) is present in animals, fungi, humans, and alphaproteobacteria, while the C5 pathway (B) is present in plants, algae, archaea, and most bacteria.

D –ALA synthases and its isozymes

Isozymes are defined as different variants of the same enzyme having identical functions

but different amino acid sequences, and they usually differ in kinetic and regulatory properties as

well (reviewed in ref. 11, 12). If one enzyme can provide enough products for an organism, it

would seem illogical for an organism to have an isozyme.

In animals, including humans, two different ALA synthase genes are present. The

presence of these isozymes in these organisms can be reasonably explained by their specific roles and different expression patterns. ALAS1 is a housekeeping gene that is ubiquitously expressed in tissues, especially in liver. ALAS1 is responsible for providing heme for respiratory cytochromes and other hemoproteins. In addition, heme can repress the expression of ALAS1

(13). ALAS2 is only expressed in erythroid cells, and its major role is to supply the heme for synthesis of hemoglobin in erythroid cells (13). The expression of ALAS2 is increased significantly during the later stage of erythropoiesis, and its expression is not inhibited by heme

(13). Therefore, the function of the two ALAS gene products in animals cannot complement each 5 other and the presence of both is necessary.

Rhodobacter sphaeroides bacteria also possess two ALA synthase isozymes, HemA and

HemT, encoded by hemA and hemT, respectively (14). However, while all of the wild type strains of R. sphaeroides examined thus far possess a hemA gene not all have a hemT gene (15,

16). This diversity in ALA synthase gene representation has created a useful experimental environment that recent studies have begun to use towards understanding the significance of having duplicate ALA synthase genes (17). Until now hemA and hemT have been investigated in

R. sphaeroides wild type strain 2.4.1. Since in that strain hemT is transcriptionally silent under all standard growth conditions, including aerobic, anaerobic-dark, phototrophic, diazotrophic, and autotrophic conditions, its role in the cell remained a mystery (18, 19, 20). Investigations of hemA and hemT in another R. sphaeroides wild type strain, 2.4.9, have revealed that, in that strain hemT is expressed, but its pattern of expression is different from hemA (17). Thus, hemA is expressed aerobically but transcription is also induced by the absence of oxygen, and this happens under anaerobic-dark conditions when cells are relying upon anaerobic respirations and also under anaerobic-light conditions when cells are growing phototrophically. However, hemT is only expressed under anaerobic-dark conditions. These studies have therefore established that in anaerobic-dark cells both HemA and HemT are present, but they do not explain their respective roles (17, N. Coulianos and J. Zeilstra-Ryalls, unpublished results).

While R. sphaeroides was among the first organisms from which ALA synthase activity was purified and examined (21), this predated knowledge of the presence of two ALA synthase genes. Further, while it is known that the strain used in that study has both hemA and hemT (15), the expression profiles of the genes in that strain has not been determined. Thus, the molecular composition of the purified ALA synthase activity is not known. A more recent study involved 6 purification of ALA synthase activity from E. coli with a plasmid expressing hemA. In that study, certain kinetic properties of the enzyme were examined, but other features of ALA synthase described in the early study were not considered, such as inhibition by hemin (22).

SPECIFIC AIMS

The ultimate goal of this work is to understand why there are two ALA synthases presented in R. sphaeroides. The hypothesis to be addressed here is that HemA and HemT have distinct characteristics that define their roles in R. sphaeroides. This study encompasses the following specific aims:

1. Purify and characterize polyhistidine-tagged R. sphaeroides 2.4.1 HemA.

2. Compare the properties of purified polyhistidine-tagged HemA to those of purified polyhistidine-tagged HemT.

7

CHAPTER II: PURIFICATION AND CHARACTERIZATION OF POLYHISTIDINE-

TAGGED HEMA

INTRODUCTION

Rhodobacter sphaeroides 2.4.1 has two ALA synthases, the products of the hemA and

hemT genes (14, 18). Understanding the presence of two ALA synthases requires that each

protein be purified and characterized. Towards that end, this chapter describes the successful

expression, purification, and characterization of recombinant polyhistidine-tagged HemA

(rHemA).

MATERIALS AND METHODS

Bacterial strains, plasmids, and growth conditions. The pIND-hemA (L. Cooper and J.

Zeilstra-Ryalls, unpublished results) expression plasmid was derived from pIND4 (23) by

inserting a hemA gene that was modified at its 5' end by the creation of an NcoI restriction

endonuclease recognition site and at its 3' end by the addition of sequences that code for an

enterokinase recognition site followed by a polyhistidine tag upstream of a HindIII recognition

sequence (Fig. 3). Plasmid pIND4 is a derivative of the pMG170 shuttle vector (24) that retains

the sequences of the Rhodobacter blasticus endogenous plasmid pMG160 and the ColE1 origin

of that vector, and so is capable of replicating in both E. coli and R. sphaeroides. Also included

from pMG170 is the selectable kanamycin resistance gene. The recombinant hemA gene from R.

sphaeroides 2.4.1 is expressed from an isopropyl β-D-1-thiogalactopyranoside (IPTG)-inducible

promoter. To ensure adequate regulation, the lacIq gene is also present on the plasmid. The host used for production of recombinant polyhistidine-tagged HemA (rHemA) was E. coli strain

DH5αphe (25). The bacteria were precultured in 5 ml LB medium (26) with kanamycin (50 8

µg/ml) overnight at 37oC with shaking (200 rpm) in an INNOVA 4200 incubator shaker (New

Brunswick Scientific, Enfield, CT). Then 1 ml of the preculture was used to inoculate 100 ml of

fresh LB with kanamycin (50 µg/ml), which was grown to an optical density (OD660 nm) of 0.6,

measured using a U-2910 spectrophotometer (HITACHI High technologies American Inc,

Schaumburg, II). IPTG (final concentration, 0.4 mM) was then added to induce expression of the hemA gene. The cells were collected after 3 hours of further incubation.

9

Figure 3. Schematic diagram (A) of the plasmid vector pIND-hemA (L. Cooper and J. Zeilstra-Ryalls, unpublished results) used for producing R. sphaeroides 2.4.1 recombinant polyhistidine-tagged HemA protein in E. coli strain DH5αphe (25), and amino acid sequence (B) of the product. Plasmid pIND-hemA is a derivative of pIND4 (23), and indicated are sequences of the endogenous plasmid from Rhodobacterblasticus pMG160 that is also capable of replicating in R. sphaeroides (24), the ColE1 origin R q of replication, a kanamycin resistance gene (Km ), the lacI gene, the engineered LacI-repressible PA1/04/03 promoter, and TL, T0 and T1 transcription terminators. Details regarding these features are provided in ref. 20. In the sequence of the plasmid product, the colored amino acids are those that were added during engineering of the hemA gene. The enterokinase EKMaxTM cleaves after the lysine residue shaded red, and the polyhistidine-tag used for affinity purification is shaded green.

Crude lysate preparation and rHemA affinity purification. 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 (Westbury, NY). The pellet was then resuspended in 2 ml lysis buffer (20

mM tricine, 10% v/v glycerol, 15 mM NaCl, 5 mM imidazole, 20 µM pyridoxal phosphate (PLP),

and 5 mM β-ME, pH 7.2) to which 0.1 ml of a proteinase inhibitor cocktail solution (4-(2-

aminoethyl) benzenesulfonyl fluoride hydrochloride, 2 mM, phosphoramidon 1 mM, bestatin

130 mM, E-64 epoxide 14 mM, leupeptin 1 mM, aprotinin 0.2 mM, pepstatin A 10 mM; Sigma-

Aldrich, Louis, MO) was added. Crude lysates were prepared by passaging the resuspended 10

cells through a French pressure cell (Thermo Scientific, Milford, MA) at 700 pounds per square

inch. The lysate was placed on ice and 30 U of DNase I (Sigma-Aldrich) per ml was added. The

mixture was incubated for 30 min at 4oC, and the insoluble material was subsequently collected by centrifugation at 20,817 × g for 15 min at 4oC in an Eppendorf 5810R centrifuge using an

F45-30-11 rotor. The supernatant was decanted into a new tube for subsequent purification of

rHemA.

The rHemA protein was affinity purified using 3 ml His60 Ni2+ Superflow Resin

(Clontech, Mountain View, CA). The resin was first loaded onto a column and equilibrated by

washing with 30 ml of lysis buffer (4oC ). Then the crude lysate was gravity-loaded onto the

resin. The protein was allowed to bind to the resin by placing the column on a Lab-Line 3-D rotator for 30 min in a 4oC. The column was then positioned and clamped vertically and the resin was allowed to settle. Unbound and nonspecifically-bound proteins were removed using wash buffer I (20 mM tricine, 10% v/v glycerol, 150 mM NaCl, 10 mM imidazole, 20 µM PLP, and 5 mM β-ME, pH 7.2). When the absorbance at 280 nm collected from the column was less than 0.01, wash buffer II (20 mM tricine, 10% v/v glycerol, 150 mM NaCl, 20 mM imidazole, 20

µM PLP, and 5 mM β-ME, pH 7.2) was applied. When the absorbance at 280 nm collected from the column was again less than 0.01, rHemA protein was eluted using elution buffer (20 mM tricine, 10% v/v glycerol, 25 mM NaCl, 350 mM imidazole, 20 µM PLP, and 5 mM β-ME, pH

7.2).

Protein concentration determinations. A spectrophotometric method described by

Greenfield (27), was used to determine the protein concentration, based on calculated extinction

coefficients and using protein fully denatured by 6 M guanidine-HCl. The molar extinction coefficients of the rHemA dimeric protein were calculated using the following equations (27, 28): 11

ε280 = 5690(nTrp) + 1280(nTyr) + 120(nCys) (1)

ε288 = 4815(nTrp) + 385(nTyr) + 75(nCys) (2)

In these equations, n is the number of each residue present in rHemA protein; Trp is tryptophan,

Tyr is tyrosine and Cys is cystine. Since the relative contribution of disulfide bonds to the

absorbance above 280 nm is only 10% that of tyrosine and 2% that of tryptophan, a zero value is

used if the number is not known (28). The calculated values of ε280 and ε288 for rHemA protein

are 67,660 and 39,050 M-1 cm-1, respectively. The rHemA dimer concentration C, in mol/l, is

then calculated from the ratio of A280/ε 280 and A288/ε288. The values should be the same which

suggest the protein was fully unfolded (27). Here, the values differed by approximately 4%.

Typically, purified protein concentrations are too low for reliably measuring the

absorbances at 280 and 288 nm. Therefore, the highly sensitive Bio-Rad Dye Reagent Kit (Bio-

Rad, Hercules, CA) was used according to the manufacturer's instructions, in most instances, and

with bovine serum albumin as a reference. These relative concentrations were converted to

absolute concentrations using a factor of 1.52, which is based on a comparison of the values

determined spectrophotometrically (averaged) versus using the Bio-Rad Dye Reagent Kit.

ALA synthase activity assays. ALA synthase activity was assayed using the method of

Burnham (21), as modified by Neidle and Kaplan (18). The reaction was started by mixing 50 µl

of purified protein to a mixture of buffer with substrates (0.28 µM PLP, 9.8 mM MgCl2, 0.98 mM Tris-HCl, 19.6 µM succinyl-CoA, and 98 mM glycine pH 7.2). The total volume of the reaction mixture was 102 µl. After the reaction mixture was incubated at 37oC, 50 µl of 10%

trichloroacetic acid (TCA) was added to terminate the reaction. The contents of the reaction tube

was then decanted into a 1.5 microfuge tube containing 200 µl of 1 M acetate buffer (pH 4.7)

and 5 µl of 99% acetylacetone (Sigma-Aldrich). The microfuge tube was incubated at 100oC for 12

15 min, during which the ALA and acetylacetone were condensed to form 2-methyl-3-acetyl-5- propionic acid pyrrole (ALA-pyrrole). The amount of product formed was determined by adding

350 µl of Ehrlich’s reagent (134.1 mM p-dimethyaminobenzaldehyde (Sigma-Aldrich), 84% v/v glacial acetic acid (EMD Chemicals, Gibbstown, NJ), and 11.2% v/v perchloric acid (GFS

Chemicals, Columbus, OH)) and incubating the tube at room temperature for 20 min. Then the absorbance at 556 nm was determined. The ALA synthase activity was deduced from the quantity of the pyrrole-Ehrlich’s complex formed, using an extinction coefficient (ε) of 0.068

µM-1cm-1 (29). The units of activity, µM ALA formed per hour, are then calculated using the

following formula (29):

OD556nm x volume of mixture in cuvette (ml) x 60 (min/h) Units = ______(3) ε (µM-1cm-1) x volume of sample (ml) x time (min) x path length (cm)

Determining pH and temperature optima of purified rHemA. The pH profile of the

purified rHemA was determined by assaying for ALA synthase activity as described above using

a 5 min incubation, except the final pH value in the reaction mixture was adjusted to 6.8, 7.2, 7.5,

7.8 and 8. The temperature profile of rHemA was obtained by assaying for ALA synthase

activity at 30oC, 37oC, 44oC and 55oC.

Hemin inhibition analysis. Hemin solutions (10 µM, 1 mM, and 10 mM) were prepared by dissolving hemin in 0.01 M NaOH and 100% ethanol (1:1, v/v). Then ALA synthase activity was assayed in reaction mixtures to which appropriate amounts of the proper hemin solutions

had been added to achieve a final hemin concentration of 0.1 µM, 1 µM, 10 µM, 50 µM and 100

µM.

Determination of Km, kcat and Vmax values for succinyl-CoA and glycine of rHemA.

To obtain data for determining the Km for glycine, ALA synthase activity was assayed in the 13

presence of different concentrations of glycine ranging from 0 to 100 mM while maintaining a

constant succinyl-CoA concentration of 19.6 µM. The data for determining the Km value for succinyl-CoA were obtained by assaying for ALA synthase activity in the presence of varying succinyl-CoA concentrations ranging from 0 to 160 µM, while maintaining a constant glycine concentration of 33.6 mM. Then nonlinear regression analysis of the data sets was performed using Sigmaplot 10.0 (Systat software, San Jose, CA) to generate graphs from which the Km values were deduced, corresponding to the concentration of each substrate at half the maximum velocity rate of the reaction. The Vmax values are directly obtained from the Km graph, and the

kcat value were calculated by using Vmax divide the molar concentration of the enzyme.

SDS-PAGE, immunoblot analysis, and InvisionTM in-gel stain. SDS-PAGE was performed using 12% polyacrylamide Tris-Glycine precast gels (Invitrogen, Carlsbad, CA).

Protein samples were prepared by mixing the samples with an equal volume of 2 x concentrated

Laemmli buffer (30), then boiling them for 5 min. The running buffer was 49.5 mM Tris-base,

0.384 M glycine, and 6.9 mM SDS, pH 8.3. Protein was stained using staining solution (45% methanol, 45% distilled water, 10% glacial acetic acid, 0.025% Coomassie Brilliant Blue R-250)

and destained in the same solution without Coomassie blue. The Bio-Rad Precision Plus

ProteinTM KaleidoscopeTM marker was used to estimate molecular masses. Protein samples were

subjected to immunoblot analysis according to standard procedures (31), using as primary

antiserum a 1:10,000 dilution of anti- HemA rabbit antiserum (22) and alkaline phosphatase-

conjugated goat anti-rabbit antisera (Sigma-Aldrich) as the secondary antibody. The detection of

immunocomplexes was achieved by using the BCIP®/NBT Liquid Substrate System (Sigma-

Aldrich).

The in-gel staining using InVisionTM (Invitrogen) stain to detect polyhistidine-tagged 14

protein was performed according to the manufacturer's instructions. Detection was by exposing

the gel to ultraviolet light using an ultraviolet transilluminator.

Circular dichroism spectrometry and analysis. Purified protein samples were

concentrated 5.2 times using filters with a molecular mass cutoff of 10,000 Da (Amicon Ultra,

Bedford, MA). The filters were loaded with sample and then centrifuged at 8327 × g for 15 min

at 4oC using a A-4-62 swinging bucket rotor in an Eppendorf 5810R centrifuge. The

concentrated protein samples were then dialyzed overnight at 4oC against 4 liters of dialysis

buffer (20 mM tricine, 5 mM NaCl, 10% glycerol (v/v), pH 7.2) to remove imidazole and PLP.

Circular dichroism (CD) spectrometry was performed using an AVIV 62DS spectrometer (AVIV

associate, Lakewood, NJ). The alpha helical content Cα, was calculated using the following

equations (32,33):

Cα1 = (-[θ]222+3,000)/39,000 (4)

Cα2 = - ([θ]208 + 4,000) / 29,000 (5)

In these equations, [θ]222 is the molar ellipticity at 222 nm (32) and [θ]208 is the molar ellipticity

at 208 nm (33).

The CD spectra were also analyzed using the DichroWeb analysis server

(http://dichroweb.cryst.bbk.ac.uk) (34, 35, 36), and K2D program was chosen for the analysis

because it does not require a reference and the CD spectra of our data fit well the range for the

program, which is from 200 to 250 nm.

EKMaxTM treatment of rHemA. EKMaxTM (Invitrogen, Carlsbad, CA) treatment was performed according to the manufacturer's instructions using 50 U of enzyme to treat 52 µg of purified protein that had been dialyzed against dialysis buffer (20 µM PLP, 20 mM tricine, 10% glycerol, 5 mM β-ME, and 10 mM calcium chloride, pH 7.2) that removed imidazole and sodium 15

chloride, which can reduce the activity of EKMaxTM (37); the calcium present in the dialysis

buffer can increase the activity of EKMaxTM (37). The EKMaxTM enzyme specifically

recognizes (Asp)4Lys and cleaves after the lysine residue (38), which should remove the polyhistidine-tag from the protein (Fig. 3). The reaction mixture was incubated at 4 oC for 16 hours.

Protein modeling. The 3-D structure of rHemA was modeled using LOMETS (Local

Meta-Threading-Server; 39). The images were captured by projections of the models using the graphics visualization tool RasMol 2.7.5.2 (40). The superimposing of the 3-D structure of rHemA with recombinant polyhistidine-tagged HemT was generated using TM align (41).

RESULTS

Expression and purification of rHemA. To estimate the optimum time following induction with IPTG at which to collect the cells, samples of induced culture were taken at 0.5, 1,

2, and 3 hours. Crude lysates were prepared, and samples of both lysate and insoluble material were examined using SDS-PAGE (Fig. 4). A prominent band of stained protein was detected at approximately 50 kDa in every sample, and was assumed to correspond to rHemA. The relative band intensity in the lysate appeared to increase over time, while at the 3 hour time point the amount of protein in the pellet seemed to be less than at the 2 hour time point. On the basis of this assessment, the length of induction that seemed to maximize yield of soluble protein was considered to be 3 h. 16

Figure 4. SDS-PAGE of the soluble and insoluble fractions of samples of E. coli strain DH5αphe with pIND-hemA that were collected at various time points following the addition of IPTG to the culture. Samples were prepared as described in the Materials and Methods. Lanes 1, 3, 5, and 7 are samples of the soluble fractions collected at 0.5, 1, 2 and 3 hours after the addition of IPTG and lanes 2, 4, 6, and 8 are samples of the insoluble fractions collected at 0.5, 1, 2, and 3 hours after the addition of IPTG. The approximate masses of the marker proteins (M) are indicated, and the protein that is thought to be rHemA is indicated by the arrow.

Using the protocol described in the Materials and Methods, the purification profile of rHemA is shown in Fig. 5 and presented in Table 1. The rHemA protein was purified more than

500-fold in the final eluate, and the typical yield from 100 ml of cell culture was 0.5 mg of protein. The purified protein remained active for approximately one week when stored at -20 oC

or -80 oC refrigerators. This improved to about 30 days when the protein was first concentrated

to 0.46 mg/ml. 17

Figure 5. SDS-PAGE of samples collected during affinity purification of rHemA. Lanes are M: molecular mass marker (sizes are as indicated), L: sample of the crude lysate, F: sample of the flow through, W1: sample collected after application of wash buffer I, W2: sample collected after application of wash buffer II, E: sample of the eluate. Each sample contained 3 µg of protein (except for the wash fractions; for which maxiumum volumes were loaded as the protein concentrations were below detection), as determined by the Bio-Rad assay. Further details are provided in the Materials and Methods.

Table 1. Purification profile of rHemA

Volume Concentration Specific activity Fraction Purification fold (ml) (mg/ml) (µmol ALA/h/mg)

Crude lysate 6 12.57 0.7 ± 0.1 1

Flow through 15 1.73 3.1 ± 0.1 4

Wash I 50 6.53×10-2 5.6 ± 0.3 7.6

Wash II 25 N.A.a N.A. N.A.

Eluate 5 1.63×10-2 410.7 ± 29.5 554

aN.A.: Not applicable; the protein concentration is below detection.

18

The linear range of the ALA synthase activity assay using rHemA was assessed by assaying at 37oC and pH 7.2. On the basis of the results (Fig. 6), all subsequent assays were

performed using a 5 min incubation time.

.016

.014

moL) .012 

.010

.008 ALA accumulation ( accumulation ALA

.006

.004 0 2 4 6 8 10 12 time (min)

Figure 6. The amount of ALA formed per mg of protein versus assay incubation time. The assay was performed as described in the Materials and Methods using 100 mM glycine and 40 µM succinyl-CoA. The points are the average of three independent assays, and the ranges are indicated by the error bars. The graph was generated using Sigmaplot software (Systat Software).

Immunoblot analysis and in-gel polyhistidine staining of purified rHemA. A

polypeptide of approximately 50 kDa molecular weight was detected by immunoblot analysis of

samples of affinity purified protein (Fig. 7). This result indicates that the purified protein is

rHemA.

Towards assessing the influence of the polyhistidine tag on the protein, its removal was

attempted by subjecting purified protein to EKMaxTM treatment, which should remove the tag.

Although the protein would still be modified in sequence relative to the wild type (Fig. 3B), the

treatment should nevertheless generate a protein whose sequence is closer to that of the wild type.

The presence of the polyhistidine tag before and after treatment was evaluated by using the 19

InVisionTM in-gel stain (Fig. 7). Since a visual comparison of the Coomassie stained gel versus

the same gel stained with the InVisionTM stain indicates that all of the protein that migrates to a

position of approximately 50 kDa is detectable by both stains, it was concluded that the

enzymatic treatment was ineffective at removing the tag. Further attempts were not made

because these optimum conditions (according to the manufacturer), while failing to remove the

tag, appeared to promote non-specific cleavage, in that lower molecular weight bands of protein

were observed using Coomassie staining.

Figure 7. Immunoblot, and InVisionTM in-gel staining of rHemA. Lane 1: immunoblot probed with rabbit anti-HemA antiserum as primary antibody (9,10), lane 2: Coomassie-stained gel after blotting, lane M: molecular mass marker (sizes are indicated), lanes 3 and 4: InvisionTM stained SDS-PAG of protein samples before (lane 3) and after (lane 4) treatment with EKMaxTM, lanes 5 and 6: Coomassie-stain of the same SDS-PAG.

Temperature and pH optima of purified rHemA. The temperature and pH profiles for rHemA protein are shown in Fig. 8. Based on the profiles, the optimum temperature for the

ALA synthase is 37 oC and the optimum pH is 7.2.

20

Figure 8. Effect of temperature and pH on rHemA activity. Shown are percent activity versus temperature (A) and pH (B). The y-axes have been truncated to provide maximum visual resolution of the values.

The effect of hemin on the ALA synthase activity of rHemA. The activity of purified rHemA was examined in the presence of varying concentrations of added hemin. The results, shown in Fig. 9, indicate that there is approximately a 25% reduction in ALA synthase activity in the presence of 100 µM hemin.

Figure 9. Effect of added hemin on rHemA activity. Shown is percent activity versus concentration of added hemin. The y-axis has been truncated to provide maximum visual resolution of the values. 21

Glycine and succinyl-CoA Km determinations for rHemA. The Km values for glycine

and succinyl-CoA were determined according to the procedures described in the Materials and

Methods section. The nonlinear regression plots of the data are presented in Fig. 10, and both

have R2 values of more than 0.99.

Figure 10. ALA synthase activity versus substrate concentrations, and nonlinear regression plots of the data used to determine glycine and succinyl-CoA Kms. ALA synthase activity versus glycine concentration for rHemA with the corresponding nonlinear regression plot (A), ALA synthase activity versus succinyl-CoA concentration for rHemA with the corresponding nonlinear regression plot (B)

Table 2 lists the Km values calculated from the nonlinear regression plots (Fig. 9). The

Vmax and Km values were deduced from the plots using the following equation:

V0= Vmax [S] / (Km + [S]) (6)

In this equation, V0 is the initial velocity and [S] is the substrate concentration.

The kcat value is calculated from the following equation:

kcat = Vmax / [S] (7)

The values are included in Table 2. Thus, the Km values for rHemA compare reasonably well to those reported for ALA synthase activity purified from E. coli expressing wild type hemA, which were 1.88 mM for glycine and 17 µM for succinyl-CoA, respectively (22). 22

Table 2. Kinetic parameters for rHemA.

Parameter Succinyl-CoA Glycine

Km 7.82±1.67 µM 8.65±1.30 mM

-1 -1 kcat 1377.7 h 635.2 h

-1 -1 kcat/Km 81.2 h µM 159.3 h mM

-1 -1 Vmax 1.02 µmol h 0.80 µmol h

Circular dichroism (CD) analysis of rHemA. Samples of purified protein were

examined by CD. However, the spectra were obtained for protein purified using a protocol that

did not include the second wash step, and so differ from the protein samples analyzed in other

parts of this study. Therefore the CD results presented here (Fig. 11) should be regarded as

tentative. (Instrumentation unavailability has prevented further CD analysis at present.)

Figure 11. Preliminary circular dichroic spectrum of purified rHemA. The graph (line) of the experimental data (filled circles) was plotted using a running average. Reconstructed data using the K2D algorithm are also shown (open circles).

The secondary structure composition of rHemA was calculated from the CD data and

using the K2D algorithm (31, 32, 33). The percent alpha helix, beta sheet, and random coil were 23

also predicted for both rHemA and the wild type protein using LOMETS (39). Table 3 lists the

values for both proteins, as well as the known secondary structure composition of R. capsulatus

ALA synthase, according to its solved crystal structure (42). The small differences in values

suggest that all three proteins are similar in structure.

Table 3. Predicted, calculated, or known secondary structure compositions of R. sphaeroides rHemA, wild type HemA, and R. capsulatus ALA synthase (42).

Calculated %s Secondary Predicted %s Known %s for Predicted %s from CD structure for wild type R. capsulatus for rHemA spectral data element HemA ALA synthase for rHemA

alpha helix 41.1 42.3 40.0 44.0

beta sheet 12.2 13.7 12.0 16.0

random coil 46.4 43.9 48.0 40.0

DISCUSSION

Active recombinant polyhistidine-tagged R. sphaeroides 2.4.1 HemA (rHemA) was

successfully affinity purified from Escherichia coli strain DH5αphe with pIND-hemA. The

optimum temperature and pH for the purified protein were determined, and the effect of added

hemin was assessed. A study of ALA synthase activity purified from R. sphaeroides reported a

pH optimum of 7.8-8.0 (21), while the optimum pH for rHemA was 7.2. Further, 100 µM hemin reduced the purified ALA synthase activity by approximately 82%, while the activity of purified rHemA was reduced by approximately 25%. The lack of knowledge of the molecular composition of the ALA synthase activity in that study suggests that these differences might be explained by the presence of both HemA and HemT in the purified protein.

More recently, ALA synthase activity was purified from E. coli expressing wild type 24

hemA (22). While that report did not include an analysis of temperature or pH optima, the

kinetic properties of the purified protein were determined. The Km values for glycine and succinyl-CoA differ by less than 10-fold compared to those of rHemA. However, the specific

activity of the recombinant protein was approximately 13-fold higher, and the turnover number

was 1.5 to 3.0-fold higher. The estimated enrichment of the wild type protein (22) was

approximately 42-fold while using affinity chromatography for purification of rHemA protein

achieved a more than 500-fold enrichment (Table 1). It is therefore possible that the differences

in kinetic properties can be accounted for by differences in protein purity.

The preliminary analysis of secondary structure composition of rHemA using CD indicates that the protein resembles that of the solved crystal structure of R. capsulatus ALA synthase. This suggests that the recombinant protein is not greatly different from the wild type protein, and in turn suggests that the results reported here for rHemA are indicative of the behavior of native HemA protein. Importantly, they are thought to be meaningful and useful in a comparison of rHemA to recombinant polyhistidine-tagged HemT.

25

CHAPTER III. A COMPARISON OF THE PROPERTIES OF POLYHISTIDINE-

TAGGED HEMA TO THOSE OF POLYHISTIDINE-TAGGED HEMT

INTRODUCTION

This chapter will focus on comparing the characteristics of recombinant polyhistidine- tagged R. sphaeroides 2.4.1 HemA (rHemA) to those of recombinant polyhistidine-tagged R.

sphaeroides 2.4.1 HemT (rHemT), which was purified and characterized in parallel with this

investgation (J. Kaganjo, MS Thesis, submitted). From an alignment of the amino acid

sequences of HemA and HemT (Fig. 12), the deduced percent identity is 53, and the percent

similarity is 71.

Figure 12. Amino acid sequence alignment of R. sphaeroides 2.4.1 wild type HemA and HemT generated using ClustalW (43). Similarities between the two proteins are shown in reverse video (black background, white font). Residues described for R. capsulatus ALA synthase (42) that are involved in PLP recognition and binding (green), glycine recognition and binding (blue), and succinyl-CoA recognition and binding (red) are highlighted.

It should be noted that, while both wild type proteins are 407 amino acid residues in 26

length, the recombinant HemA and HemT proteins are 433 and 424 amino acids in length,

respectively. However, as suggested by comparative modeling (Fig. 13 and J. Kaganjo, MS

Thesis, submitted), as well as secondary structure prediction analysis (Table 3 and J. Kaganjo,

MS Thesis, submitted), the additional sequences do not appear to significantly alter the overall

folding of either protein. Therefore, differences in properties of the recombinant proteins are

considered to be informative as to potential differences between the native ALA synthase

enzymes.

Figure 13. Tertiary structure model superpositions of R. sphaeroides 2.4.1 rHemA and wild type HemA upon the solved crystal structure of R. capsulatus ALA synthase (42). In panel A, the model of R. sphaeroides rHemA (red) is superimposed upon the solved crystal structure of R. capsulatas ALA synthase (yellow). In panel B, the model of R. sphaeroides wild type HemA (red) is superimposed upon the solved crystal structure of R. capsulatus ALA synthase (yellow). Models were generated using the LOMETs (39) and the superimposed images were generated using TM-Align (41).

Differences and similarities between purified rHemA and rHemT

The optimum temperature and pH are same for rHemA and rHemT. The effect of added hemin is also similar for both proteins; in the presence of 100 µM hemin, rHemA activity was reduced by 25% while the activity of rHemT was reduced by 30%. In neither case is activity 27 reduced to anywhere near the extent reported previously (21). Whether or not the greater purity of the recombinant proteins achieved by affinity purification versus that achieved by conventional purification processes accounts for this difference is not yet known, nor is it known whether or not the molecular composition of the ALA synthases can alter the influence of hemin on activity.

A comparison of the kinetic data for each protein (Table 4; values for rHemT were from J

Kaganjo, MS Thesis, submitted) reveals that the Km values for glycine are very similar, and the

Km for succinyl-CoA is 2-fold lower for rHemA than for rHemT. Examining the amino acid residues present in HemA and HemT at positions corresponding to the region of the protein involved recognition and binding of succinyl-CoA (42) reveals that lysine 156 in HemA is arginine 156 in HemT (Fig. 12). Although both lysine and arginine are positively charged basic amino acids, mutagenesis resulting in a residue substitution would confirm if it is the cause of the lower affinity. It would also make it possible to consider whether this difference is significant in vivo.

Table 4. Comparison of the kinetic properties of rHemA and rHemT.

Polyhistidine-tagged Polyhistidine-tagged Property HemA HemT

Specific activity (µmol ALA/h/mg) 624±45 30±4

Succinyl-CoA Km (µM) 7.8±1.7 18.0±3

Succinyl-CoA Vmax (µmol ALA/h) 1.0 2.7

Succinyl-CoA kcat (µmol/h/mg) 635.2 67.5

Succinyl-CoA kcat/Km 80.0 3.8

Glycine Km (mM) 8.7±1.3 9.0±2.0

Glycine Vmax (µmol/h) 0.80 1.15

Glycine kcat(µmol/h /mg) 1377.7 50.0

Glycine kcat/Km 153 6 28

There is greater confidence that the far more dramatic difference in specific activities

(more than 20-fold) as well as kcat values (more than 9-fold for succinyl-CoA and nearly 28-fold for glycine) are important and telling about the functioning of the proteins in the cell. Thus, one assumes that cells relying upon HemA-ALA synthase activity have the capacity to consume or divert more glycine and succinyl-CoA to ALA production than cells relying upon HemT-ALA synthase activity.

The more obvious alternative roles for the ALA precursor molecules are in protein

synthesis with respect to glycine and in central carbon metabolism with respect to succinyl-CoA.

These metabolic connectivities suggest that it would be deleterious to the cell if ALA production

so predominates as to deplete their availability for use in the other pathways. Therefore, since

ALA is essential, regardless of what other metabolism is taking place, the utility of having genes

encoding enzymes with high and low activities can be explained by a combination of the need to

sustain ALA production under all conditions in combination with the need to have the flexibility

to redirect flow of metabolites. This can be accomplished by adjusting the ALA synthase protein

composition of the cytoplasm.

Within the context of what is currently known about the expression of the hemA and hemT genes, the only situation in which HemT could be present is when HemA is present as well, and based on transcription analyses, both enzymes would be maximally present when cells are growing by anaerobic respiration using dimethyl sulfoxide as alternate electron acceptor (17).

Although the relative amounts of HemA and HemT protein present under those conditions are not yet known, predictions based on the relative amounts of transcription suggest that HemA protein would be in excess. If true, one would need to explain how it is that HemT might be important in that situation. Compared to aerobic respiration or phototrophy, anaerobic 29

respiration generates less energy for the cell, and so the importance of balancing anabolic and

catabolic metabolisms is acute. Yet, should light become available, it is also important that ALA

production be ramped up quickly in order to provide bacteriochlorophyll, for photosynthesis;

perhaps far more quickly than can be accommodated by regulation at the level of transcription.

Indeed, anaerobic hemA transcription is the same regardless of the presence or absence of light,

which evokes a post-transcriptional regulatory event for increasing ALA synthase activity. A

cytoplasm having both HemA and HemT polypeptides makes the formation of heterodimers

possible in that they are both present. If such heterodimers do form, such proteins may have

properties that differ from the HemA and HemT homodimer properties. For example, it may be

that a HemA-HemT heterodimer has lower catalytic activity than a HemA homodimer but higher catalytic activity than the HemT homodimer. If true, the regulatory process that would escalate

ALA synthase activity might be elimination of HemT, causing a shift in oligomerization equilibrium towards the more active HemA homodimer formation.

Critical evidence to substantiate these arguments is not yet available. However, it is through knowledge of the properties of the pure proteins that such a hypothesis could be conceived. Further, this knowledge is required in order to appreciate the properties of a heterodimeric protein, should its existence be demonstrated. 30

REFERENCES

1. Porter S. L., D. A. Wilkinson, E. D. Byles, G. H. Wadhams, S. Taylor, N. J. Saunders, and J. P. Armitage. 2011. Genome sequence of Rhodobacter sphaeroides strain WS8N. J.

Bacteriol. 193: 4027-4028.

2. Panwichian S., D. Kantachote, B. Wittayaweerasak, and M. Mallavarapu. 2010. Isolation of purple nonsulfur bacteria for the removal of heavy metals and sodium from contaminated shrimp ponds. Electronic J. Biotechnol. 13:3-4.

3. Hillmer P. and H. Gest. 1977. H2 metabolism in the photosynthetic bacterium

Rhodopseudomonas capsulata: H2 production by growing cultures. J. Bacteriol. 129:724–731.

4. Mackenzie C., M. Choudhary, F. W. Larimer, P. F. Predki, S. Stilwagen, J. P. Armitage,

R. D. Barber, T. J. Donohue, J. P. Hosler, J. E. Newman, J. P. Shapleigh, R. E. Sockett, J.H.

Zeilstra-Ryalls, and S. Kaplan. 2001. The home stretch, a first analysis of the nearly completed genome of Rhodobacter sphaeroides 2.4.1. Photosynth. Res. 70:19–41.

5. Lascelles J. 1978. Regulation of pyrrole synthesis, p. 795-808. In R. K. Clayton and W. R.

Sistrom (ed.), The photosynthetic bacteria. Plenum Publishing Corp., New York.

6. Frankenberg N., J. Moser, and D. Jahn. 2003. Bacterial heme biosynthesis and its biotechnological application. Appl Microbiol. Biotechnol. 63:115-27.

7. Martens J. H., H. Barg, M. J. Warren, and D. Jahn. 2002. Microbial production of vitamin

B12. Appl. Microbiol. Biotechnol. 58:275–285.

8. Jahn D., D.W. Heinz. 2009. Chapter 2: Bioynthesis of 5-Aminolevulinic Acid, p. 29-42,

Tetrapyrroles: Birth, Life and Death. Landes Bioscience and Springer Science+Business Media,

LCC.

9. Ajioka R. S., J. D. Phillips, and J. P. Kushner. 2006. Biosynthesis of heme in mammals. 31

Biochim Biophys. Acta. 1763:723–736.

10. Beale S. I. 1990. Biosynthesis of the tetrapyrrole pigment precursor, δ-aminolevulinic acid, from glutamate. Physiol. 93:1273-1279.

11. Baron D. N. 1965. Isoenzymes and their clinical application. Ann. R. Coll. Surg. Engl.

37:263–271.

12. Goodfriend T. L. and N. O. Kaplan.1965. Isoenzymes in Clinical Diagnosis. Circulation.

32:1010-1019.

13. May B. K., S. C. Dogra, T. J. Sadlon, C. R. Bhasker, T. C. Cox, and S. S. Bottomley.

1995. Molecular regulation of heme biosynthesis in higher vertebrates. Prog. Nucleic. Acid Res.

Mol. Biol. 51:1-51.

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

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

16. Choudhary M., X. Zanhua, Y. X. Fu, and S. Kaplan. 2007. Genome analyses of three strains of Rhodobacter sphaeroides: evidence of rapid evolution of chromosome II. J. Bacteriol.

189:1914-1921.

17. Coulianos N. 2011. A comparison of ALA synthase gene transcription in three wild type strains of Rhodobacter sphaeroides. (Master's Thesis), Bowling Green State University. OH,

USA.

18. Neidle E. L. and S. Kaplan. 1993. 5-Aminolevulinic acid availability and control of spectral complex formation in hemA and hemT mutants of Rhodobacter sphaeroides. J. Bacteriol.

175:2304-2313. 32

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

20. Zeilstra-Ryalls J.H. and S. Kaplan. 1995. Regulation of 5-aminolevulinic acid synthesis in Rhodobacter sphaeroides 2.4.1: the genetic basis of mutant H-5 auxotrophy. J. Bacteriol.

177:2760–2768.

21. Burnham B. F. and J. Lascelles. 1963. Control of biosynthesis through a negative-feedback mechanism. Biochem. J. 87:462-472.

22. Bolt E. L., L. Kryszak, J. Zeilstra-Ryalls, P. M. Shoolingin-Jordan, and M. J. Warren.

1999. Characterization of the Rhodobacter sphaeroides 5- aminolaevulinic acid synthase isoenzymes, HemA and HemT, isolated from recombinant Escherichia coli. Eur. J. Biochem.

265:290-299.

23. Ind A. C., S.L. Porter, M. T. Brown, E. D. Byles, J. A. de Beyer , S. A. Godfrey, and J. P.

Armitage. 2009. Inducible-Expression plasmid for Rhodobacter sphaeroides and Paracoccus denitrificans. Appl. Environ. Microbiol. 75:6613-6615.

24. Inui M., K. Nakata, J. H. Roh, A. A. Vertes, and H. Yukawa. 2003. Isolation and molecular characterization of pMG160, a mobilizable cryptic plasmid from Rhodobacter blasticus. Appl. Environ. Microbiol. 69:725-733.

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

43.

26. Luria S. E., J. W. Burrous. 1957. Hybridization between Escherichia coli and Shigella. J.

Bacteriol. 74:461-476.

27. Greenfield N. J. 2006. Using circular dichroism spectra to estimate protein secondary 33

structure. Nat. Protoc. 1:2876–2890.

28. Edelhoch H. 1967. Spectroscopic determination of tryptophan and tyrosine in proteins.

Biochemistry 6:1948–1954.

29. Burnham B. F. 1970. δ-Aminolevulinic acid synthase (from spheroides). Meth. Enzymol. 17A:195-204.

30. Laemmli U. K. 1970. Cleavage of structural proteins during the assembly of the head of T4. Nature 227:680–685.

31. Harlow E. and D. Lane. 1999. Using antibodies: a laboratory manual. Cold Spring Harbor

Laboratory Press, Cold Spring Harbor, N.Y.

32. J. A. Morrow, M. L. Segall, S. Lund-Katz, M. C. Phillips, M. Knapp, B. Rupp, and K. H.

Weisgraber. 2000. Differences in stability among the human apolipoprotein E isoforms

determined by the amino-terminal domain. Biochemistry 39:11657–11666.

33. Greenfield N. and G. D. Fasman. 1969. Computed circular dichroism spectra for the

evaluation of protein conformation. Biochemistry 8:4108–4116.

34. Lobley A., L. Whitmore, and B. A. Wallace. 2002. DICHROWEB: an interactive website for the analysis of protein secondary structure from circular dichroism spectra. Bioinformatics

18:211-212.

35. Whitmore L. and B.A. Wallace. 2004. DICHROWEB: an online server for protein

secondary structure analyses from circular dichroism spectroscopic data. Nucleic Acids Res.

32:668-673.

36. Whitmore L. and B. A. Wallace 2008. Protein secondary structure analyses from circular

dichroism spectroscopy: methods and reference databases. Biopolymers 89:392-400.

37. Baratti J., S. Maroux, and D. Louvard. 1973. Effect of ionic strength and calcium ions on 34

the activation of trypsinogen by enterokinase. A modified test for the quantitative valuation of

the enzyme. Biochem. Biophys. Acta. 321:632-638.

38. Anderson L. E., K. A. Walsh, and H. Neurath. 1977. Bovine enterokinase. purification, specificity, and some molecular properties. Biochemistry 16:3354-3360.

39. Wu S. and Y. Zhang. 2007. LOMETS: A local meta-threading-server for protein structure prediction. Nucleic Acids Res. 35:3375-3382.

40. Sayle R. A. and E. J. Milner-White. 1995. RASMOL: biomolecular graphics for all.

Trends Biochem. Sci. 20:374.

41. Zhang Y. and J. Skolnick. 2005. TM-align: a protein structure alignment algorithm based on TM-score. Nucleic Acids Res. 33:2302-2309.

42. Astner I., J. O. Schulze, J. van den Heuvel, D. Jahn, W. D. Schubert,and D. W. Heinz.

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.

43. Larkin M. A., G. Blackshields, N. P. Brown, R. Chenna, P. A. McGettigan, H.

McWilliam, F. Valentin, I. M. Wallace, A. Wilm, R. Lopez, J. D. Thompson, T. J. Gibson, and D.

G. Higgins. 2007. CLUSTALW and CLUSTAL X version 2.0. Bioinformatics 23:2947–2948.