Recycling of Vitamin B12 and NAD+ Within the Pdu Microcompartment of Salmonella Enterica Shouqiang Cheng Iowa State University

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2010 Recycling of vitamin B12 and NAD+ within the Pdu microcompartment of Salmonella enterica Shouqiang Cheng Iowa State University

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+ Recycling of vitamin B12 and NAD within the Pdu microcompartment of Salmonella enterica

Shouqiang Cheng

A dissertation submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Major: Biochemistry

Program of Study Committee: Thomas A. Bobik, Major Professor Alan DiSpirito Basil Nikolau Reuben Peters Gregory J. Phillips

Iowa State University

Ames, Iowa

2010

Table of contents

Abstract...... iv

Chapter 1. General introduction...... 1

1.1. Vitamin B12...... 1

1.1.1. Discovery and structure of vitamin B12...... 1

1.1.2. Biosynthesis of vitamin B12...... 3

1.1.3. Absorption, transport and cellular uptake of vitamin B12...... 7

1.1.4. Function of vitamin B12...... 9

1.1.5. Vitamin B12-related diseases ...... 17 1.2. Nicotinamide adenine dinucleotide (NAD+) ...... 19

1.2.1. Discovery and structure of NAD+ ...... 19

1.2.2. Biosynthesis of NAD+ ...... 19

1.2.3. Function of NAD+ ...... 21

1.2.4. NAD+-related diseases ...... 24

1.3. AdoCbl-dependent 1,2-propanediol (1,2-PD) degradation ...... 25

1.3.1. Source of 1,2-PD...... 25

1.3.2. 1,2-PD utilization ...... 26

1.3.3. The pdu locus in S. enterica ...... 27

1.3.4. Control of the pdu/cob regulon ...... 28

1.3.5. Pathogenesis...... 30

1.4. Bacterial microcompartment...... 31

1.4.1. Discovery and diversity...... 31

1.4.2. Component and architecture...... 34

1.4.3. Function and mechanism...... 38

1.5. Research overview ...... 41

1.6. Thesis organization ...... 42 Chapter 2. Characterization of the PduS cobalamin reductase of Salmonella and its role in the Pdu microcompartment...... 43

2.1. Abstract ...... 44

2.2. Introduction...... 45

2.3. Materials and methods ...... 48

iii

2.4. Results...... 53

2.5. Discussion ...... 71

2.6. Acknowledgements ...... 76 Chapter 3. PduQ is a 1-propanol dehydrogenase associated with the Pdu microcompartment of Salmonella enterica...... 78

3.1. Abstract ...... 78

3.2. Introduction...... 78

3.3. Materials and methods ...... 80

3.4. Results...... 85

3.5. Disscusion ...... 96

3.6. Acknowledgements ...... 99

Chapter 4. Conclusions ...... 100

References...... 104

Acknowledgements...... 124

Abstract

Salmonella enterica is capable of utilizing 1,2-propanediol (1,2-PD) as a sole carbon and energy source in a coenzyme B12 (Adenosylcobalamin, AdoCbl) -dependent fashion that involves a bacterial microcompartment (MCP), the Pdu MCP. Pdu MCP is a polyhedral organelle composed entirely of protein subunits and its function is to sequester the intermediate propionaldehyde formed in the first step of 1,2-PD degradation in order to mitigate its toxicity and prevent DNA damage. Several sequentially-acting metabolic enzymes, including the diol dehydratase PduCDE and the propionaldehyde dehydrogenase PduP that respectively use AdoCbl and NAD+ as cofactors, are encapsulated within the solid protein shell of the MCP. During catalysis, the adenosyl-group of AdoCbl bound to PduCDE is periodically lost due to by-reactions and is usually replaced by a hydroxo-group resulting in the formation of an inactive

OH-Cbl-enzyme complex, and the NAD+ is converted to NADH by PduP, which make it necessary to regenerate or/and replenish AdoCbl and NAD+ within the Pdu MCPs.

Recent crystallography studies suggested that some Pdu MCP shell proteins, such as

PduA, T and U, have pores that may mediate the transport of enzyme substrates/cofactors across the MCP shell. However, it’s possible that the cofactors might be regenerated in the local vicinity of the metabolic enzymes within the MCP, which could contribute to higher activity of the enzymes and thus to higher efficiency of 1,2-PD degradation.

In this work, two proteins encoded by the pdu locus consisting of genes specifically involved in 1,2-PD utilization, PduS and PduQ, were overexpressed, purified and characterized in vitro, and their in vivo fuctions were investigated as well.

Purified PduS enzyme exhibited the abilities to catalyze the two successive uni-electron reductions from cob(III)alamin to cob(I)alamin in vitro. The results indicated

that PduS is a monomer and each monomer of PduS contains one non-covalently bound

FMN and two [4Fe-4S] clusters which are oxygen-labile. Genetic studies showed that a pduS deletion decreased the growth rate of Salmonella on 1,2-PD supporting a role in cobalamin reduction in vivo. Further SDS-PAGE and Western blot of purified Pdu MCP and following MS-MS analysis demonstrated that the PduS protein is a component of the

Pdu MCP. In addition, two-hybrid experiments indicated that PduS interacts with the

PduO adenosyltransferase which is also located in the Pdu MCP and catalyzes the terminal step of AdoCbl synthesis.

Purified PduQ enzyme was identified as an oxygen-sensitive iron-dependent alcohol dehydrogenase (ADH) that catalyzes the interconversion between propionaldehyde and

1-propanol in vitro. The propionaldehyde reduction activity of PduQ was about 45 times higher than that of the 1-propanol oxidation at pH 7.0, indicating that this enzyme is more efficient for catalyzing aldehyde reduction in cells where approximatedly neutral pH is maintained. Kinetic studies indicated PduQ has a high affinity for the substrate and cofactor involved in this reduction reaction, which also suggest that the physiological function of PduQ is the reduction of aldehyde with the conversion of NADH to NAD+.

The pduQ deletion impaired growth on 1,2-PD and led to a lower cell density, indicating that PduQ was required to support the maximal rate of 1,2-PD degradation by Salmonella.

SDS-PAGE and Western blot of purified Pdu MCP and subsequent MS-MS analysis demonstrated that the PduQ protein is also associated with the Pdu MCP. Co-affinity purification illustrated a specific strong interaction between PduQ and PduP.

In conjunction with prior results, this work indicates that the Pdu MCP encapsulates a complete AdoCbl recycling system and that it is able to recycle the electron carrier NAD+ as well within the MCP lumen.

Chapter 1. General introduction

1.1. Vitamin B12

Vitamin B12 is also known as cobalamin (Cbl). In a broad sense, B12 refers to a group of cobalt-containing vitamer compounds including cyanocobalamin (CN-Cbl), hydroxocobalamin (OH-Cbl), and the two coenzyme forms of B12:

5'-deoxyadenosylcobalamin (adenosylcobalamin, AdoCbl), and methylcobalamin

(MeCbl). The term B12 may be properly used to refer to cyanocobalamin, the principal

B12 form used for foods and in nutritional supplements.

Vitamin B12 is the most chemically complex of all the vitamins. The structure of B12 is based on a corrin ring, which is similar to the porphyrin ring found in heme, chlorophyll, and cytochrome. De novo synthesis of B12 occurs only in certain bacteria and archaea but the assimilation of complex precursors is more widespread taking place in many microorganisms and in higher animals (202). Vitamin B12 is an essential cofactor for a variety of enzymes that are widely distributed among microorganisms and higher animals

(14, 203). Plants and fungi are thought to neither synthesize nor use B12 in their metabolism (70).

Vitamin B12 is naturally found in foods of animal origin including meat and milk products. Non-ruminant animals must obtain B12 in their diet.

1.1.1. Discovery and structure of vitamin B12

Vitamin B12 was discovered by accident in medicine. B12 deficiency is the cause of pernicious anemia, an anemic disease that was usually fatal and had unknown etiology when it was first described by Combe in 1824 (80). In 1925, Whipple and co-worker showed that the regeneration of red blood cells in anemic dogs induced by bleeding was stimulated by a diet containing liver (182), and in 1926 Minot and Murphy reported a

remarkable improvement in patients fed a diet of raw liver (152). This clue started the search for the “liver factor” or “anti-pernicious anemia factor”. In 1928, Edwin Cohn prepared a liver extract that was 50 to 100 times more potent than the natural liver products (56). The extract was the first workable treatment for the disease. These events in turn led to discovery of the water soluble vitamin, called vitamin B12, in the liver juice in 1948 by Karl A. Folkers (181) and E. Lester Smith (212). In 1955, the B12 structure was elucidated by X-ray crystallographic analysis in the laboratory of Dorothy Hodgkin

(102).

A B

FIG. 1.1. The structure of vitamin B12. A, Structural formula of vitamin B12; B, Stick model of vitamin

B12; C, The various ligation states of cobalamin derivatives (17).

The structure of tetrapyrrolic vitamin B12 is shown in Figure 1.1. B12 has three parts: a central ring, an upper ligand, and a nucleotide loop. The upper ligand is a cyano (CN-) group that can be replaced by a hydroxo (OH-), methyl (CH3-), or 5’-deoxyadenosyl

(Ado-) group. The central cobalt atom coordinatively binds to the four equatorial nitrogen ligands of the corrin ring. Attached to a carbon of the corrin ring is a nucleotide side chain terminated by a 5,6-dimethylbenzimidazole (DMB) group, which may act as an axial

ligand toward the cobalt atom. Cobalamin complexes in which the DMB is bound to the cobalt atom are usually named base-on forms, whereas if such coordination is absent or replaced by an exogeneous ligand the designation base-off is used (Fig. 1.1C).

The cobalt atom in cobalamins can exist under three main formal oxidation states, Co+3,

Co+2, and Co+1, which display quite different chemical properties. Roughly speaking,

Co+3 appears as an electrophile, Co+2 as a radical, and Co+1 as a nucleophile.

Oxidoreductive conversions between the three oxidation states are thus of key importance in the chemistry of vitamin B12. When the oxidation state of the cobalt atom decreases its coordination number tends to decrease accordingly.

1.1.2. Biosynthesis of vitamin B12

According to a widely accepted historical view, B12 de novo synthesis is restricted to some bacteria and archaea (202). Many animals, including humans, and protists require

B12 but apparently do not synthesize it de novo. Plants and fungi are thought to neither synthesize nor use B12 in their metabolism (70).

1.1.2.1. De novo synthesis

Vitamin B12 is synthesized de novo via two alternative routes, aerobic and anaerobic pathways that differ primarily in the early stages that lead to the contraction of the macrocycle and excision of the extruded carbon molecule and its attached methyl group

(183) (Fig. 1.2). The aerobic pathway, which has been determined experimentally in

Pseudomonas denitrificans (206), incorporates molecular oxygen into the macrocycle as a prerequisite to ring contraction. The anaerobic route, which has been most extensively studied in Salmonella enterica serovar Typhimurium LT2 (referred to hereafter as S. enterica) (189), takes advantage of a chelated cobalt ion, in the absence of oxygen, to set the stage for ring contraction.

FIG. 1.2. Biosynthesis of precorrin 6 (aerobic pathway; light arrows) and cobalt-precorrin 6

(anaerobic pathway; bold arrows) on the route from ALA to vitamin B12 (206). The two pathways are identical from ALA to precorrin 2, at which point they diverge as shown. PBG, porphobilinogen; HMB, hydroxymethylbilane; SAM, S-adenosyl-L-methionine.

The biosynthetic route to AdoCbl from its five-carbon precursor, 5-aminolaevulinic acid (ALA) may be divided into three sections: (1) the biosynthesis of uroporphyrinogen

III (UroIII) from ALA, which is shared by both pathways (Fig. 1.2); (2) the conversion of

UroIII into the ring-contracted, deacylated intermediate (precorrin 6 or cobalt-precorrin 6), wherein lies the primary differences between the two pathways (Fig. 1.2); and (3) the transformation of this intermediate to AdoCbl (Fig. 1.3).

First, UroIII, which is also an intermediate during heme, chlorophyll, F430, and siroheme synthesis, is biosynthesized through a common pathway from ALA utilizing the enzymes ALA dehydratase (HemB), porphobilinogen deaminase (HemC) and UroIII synthase (HemD) (Fig. 1.2) (206).

Next, UroIII is converted into precorrin 6 (aerobic) or cobalt-precorrin 6 (anaerobic)

(Fig. 1.2). The aerobic pathway requires five S-adenosyl-L-methionine (SAM)

-dependent methyltransferases (CobA, I, J, M, F) for the introduction of six methyl groups and is characterized by the incorporation of molecular oxygen by a monooxygenase (CobG). The anaerobic pathway also requires five methyltransferases

(CysG, CbiK or X, CbiL, H, F), and is distinguished by the early incorporation of cobalt into the macrocycle.

FIG. 1.3. Aerobic pathway from precorrin 6 to AdoCbl (206). SAM, S-adenosyl-L-methionine; Ado-, adenosyl.

Lastly, precorrin 6 or cobalt-precorrin 6 is converted into AdoCbl. The known pathway and enzymes necessary for the conversion of precorrin 6 into AdoCbl in the aerobic pathway are shown in Figure 1.3. The structures of the intermediates of the anaerobic pathway between cobalt-precorrin 6 and AdoCbl are believed to the same as those of the aerobic pathway, with the exception that, since the cobalt ion is already in place,

dihydro-precorrin 6, precorrin 8 and hydrogenobyrinic acid are replaced with the corresponding cobalt complexes. Thus, in the anaerobic pathway, hydrogenobyrinic acid is replaced by cobyrinic acid, which then is the substrate for bisamidation to afford cob(II)yrinic acid a,c-diamide. All of the ensuing intermediates are identical in the aerobic and anaerobic pathways.

1.1.2.2. Assimilation and recycling

De novo synthesis occurs only in prokaryotes but the assimilation of complex precursors is more widespread taking place in many microorganisms and in higher animals (202). A model for the assimilation of CN-Cbl and OH-Cbl to AdoCbl, based on work done in a number of laboratories, is shown in Figure 1.4. CN-Cbl is first reductively decyanated to cob(II)alamin (91, 119, 238). Next, cob(II)alamin is reduced to cob(I)alamin and ATP:cob(I)alamin adenosyltransferase (ATR) transfers a

5'-deoxyadenosyl-group from ATP to cob(I)alamin to form AdoCbl (39, 42, 111, 113, 131,

215, 221, 248). OH-Cbl assimilation occurs by a similar pathway except that the first step is reduction of OH-Cbl to cob(II)alamin by cobalamin reductase or by the reducing environment of the cell (82, 239).

FIG. 1.4. Proposed pathway from CNCbl to AdoCbl (adapted from ref. (195)).

The pathway used for the assimilation of OH-Cbl and CN-Cbl is also used for intracellular cobalamin recycling. During catalysis the adenosyl-group of AdoCbl is periodically lost due to by-reactions and is usually replaced by a hydroxo-group resulting in the formation of an inactive OH-Cbl-enzyme complex (228). Cobalamin recycling

begins with a reactivase that converts inactive OH-Cbl-enzyme complex to OH-Cbl and apoenzyme (154, 155). Next, the process described in Figure 1.4 converts OH-Cbl to

AdoCbl, which spontaneously associates with apoenzyme to form active holoenzyme

(154, 155, 228). In the organisms that have been studied, cobalamin recycling is essential, and genetic defects in this process block AdoCbl-dependent metabolism (17, 66, 113).

1.1.3. Absorption, transport and cellular uptake of vitamin B12

Cobalamins cannot be synthesized by higher organisms and must be supplied with the diet. Mammals have developed a complex pathway, sketched on the left side of Figure 1.5, for absorption, transportation and cellular uptake of cobalamin.

FIG. 1.5. Absorption, transport and cellular uptake of cobalamins in mammals (177).

This pathway involves three separate binding proteins, haptocorrin (HC), intrinsic factor (IF) and transcobalamin (TC), which form tight complexes with Cbl (4, 190). The dietary Cbl is preferentially bound to salivary HC to form the complex HC-Cbl.

Pancreatic proteases in the duodenum cleave HC, Cbl is released and then binds to IF

forming IF-Cbl (5). Inside the enterocytes, the IF is degraded and the free Cbl binds to TC.

The TC-Cbl complex is then released into plasma, where it is endocytosed by membrane receptors, R-TC-Cbl (161, 174) (Fig. 1.5, left side). Inside the target cells, TC-Cbl is degraded into lysosomes, releasing cobalamin molecules which are metabolized to the two cofactors: 5’-deoxyadenosyl-Cbl in mitochondrion and methyl-Cbl in cytosol (Fig.

1.5, right side) (13).

FIG. 1.6. Hypothetical scheme of cobalamin uptake in E. coli (177). The proteins involved in the transport and B12 cellular uptake, whose X-ray structures are available, are shown by the ribbon representation for the protein moiety and by stick and ball for cobalamin. The X-ray structures of OmpF, ExbB and ExbD have not been determined and are indicated by a colored form.

In E. coli (and other Gram-negative bacteria), a hypothetical representation of the Cbl import across the membrane (38) is sketched in Figure 1.6. The B12 transmembrane

import is accomplished by the B12-uptake (Btu) system, consisting of an outer membrane transporter BtuB (Fig. 1.6, left side) and an ATP-binding cassette (ABC) transporter,

BtuC2D2F, located in the inner membrane (Fig. 1.6, right side). It is proposed that Cbl is taken up by BtuB, located in the outer membrane, and then released in the periplasmatic space (Fig. 1.6, left side). In the periplasmatic space Cbl is bound to BtuF and the resulting BtuF-Cbl complex binds to the periplasmatic side of BtuC2D2, located in the inner membrane (Fig. 1.6, right side), and feeds Cbl to the latter. The hydrolysis of ATP, located between the two BtuD domains powers the import of Cbl in the E. coli cells.

1.1.4. Function of vitamin B12

Vitamin B12 is an indispensable organometallic cofactor associated with a variety of enzymes that are widely distributed among microorganisms and higher animals (14, 203).

Based on the reaction mechanisms, the enzymes are divided into three classes:

AdoCbl-dependent isomerases, MeCbl-dependent methyltransferases, and corrinoid-dependent reductive dehalogenases, which catalyze intramolecular rearrangement, methyl group transfer, and reductive dehalogenation, respectively.

1.1.4.1. AdoCbl-dependent intramolecular rearrangement

The isomerases are the largest subfamily of AdoCbl-dependent enzymes found in bacteria. In enteric bacteria, they play important roles the B12-dependent degradation of ethanolamine, propanediol, and glycerol (Fig. 1.7), which may be the earliest use of B12

(15, 16, 145). In these reactions, the B12-mediated rearrangement generates an aldehyde that can be oxidized and provide ATP; the oxidations can be balanced by reducing a portion of the aldehyde to an alcohol. In nonenteric bacteria, the AdoCbl -dependent amino mutases (specific for glutamate, lysine, leucine, or ornithine) catalyze mechanistically similar reactions that support fermentation of these amino acids.

FIG. 1.7. AdoB12-dependent enzymes involved in fermentative degradation in enteric bacteria (188). Only the first reaction for each pathway employs cobalamin as a cofactor.

The AdoCbl-dependent methylmalonyl-CoA mutase (MCM or MUT), which is found in both bacteria and higher animals, catalyzes the isomerization of

(R)-methylmalonyl-CoA and succinyl-CoA during the catabolism of branched-chain amino acids, thymine, uracil, cholesterol, and odd-chain fatty acids (76) (Fig. 1.8).

MUT

(R)-methylmalonyl-CoA succinyl-CoA

FIG. 1.8. Isomerization of (R)-methylmalonyl-CoA and succinyl-CoA catalyzed by methylmalonyl-CoA mutase.

In some organisms, an AdoCbl-dependent ribonucleotide reductase (class II) catalyzes

the conversion of ribonucleotides to deoxyribonucleotides (Fig. 1.9), which is fundamentally important for DNA replication and repair (18).

FIG. 1.9. The reaction catalyzed by class II ribonucleotide reductase (18).

In the above enzymes, AdoCbl acts as a free radical generator. The general catalytic mechanism is shown in Figure 1.10. The reaction begins with homolytic rupture of the

C-Co(III) bond of AdoCbl. The C and Co atoms each acquire one electron, leading to the formation of cob(II)alamin and the deoxyadenosyl radical (step 1). This is followed by hydrogen atom (H-atom) abstraction from the substrate by the deoxyadenosyl radical to generate a reactive primary substrate radical (step 2). Rearrangement generates a secondary product-centered radical (step 3). Finally, a second H-atom transfer from deoxyadenosine yields product (step 4), and recombination of the cofactor centered radical pair (step 5) completes a turnover cycle. The catalytic mechanism of class II ribonucleotide reductase is a bit different from the general mechanism described above in that a thiyl radical rather than the initially formed 5′-deoxyadenosyl radical abstracts a hydrogen atom from the substrate and reduction occurs at C2′ of the substrate in class II ribonucleotide reductases (Fig. 1.9).

1 5

2 4

FIG. 1.10. The reaction mechanism catalyzed by AdoCbl enzymes (CH2-R = Ado) (177).

1.1.4.2. MeCbl-dependent methyl group transfer The MeCbl-dependent methyltransferases play an important role in amino acid metabolism in many organisms including humans as well as in one-carbon metabolism and CO2 fixation in anaerobic microorganisms (18). As a subclass of the group transfer enzymes, MeCbl-dependent methyltransferases catalyze the movement of a methyl group from a methyl donor (X-CH3) to a methyl group acceptor (Nuc) (Fig. 1.11), where X and

Nuc are the leaving group and the nucleophile, respectively (18). In these methyltransferases, MeCbl is cleaved heterolytically and acts as a methyl group carrier.

FIG. 1.11. Methyl donors and acceptors used by the B12-dependent methyltransferases (18).

Methionine synthase (MS) from Escherichia coli (MetH) is the most extensively studied B12-dependent methyltransferase that transfers a methyl group from

5-methyltetrahydrofolate (CH3-THF) to homocysteine to form methionine and tetrahydrofolate (THF) at the final step in methionine biosynthesis (Fig. 1.12) (69, 73).

The MeCbl-dependent methionine synthase works in two steps in a ping-pong reaction

5 (73) (Fig. 1.12). First, MeCbl is formed by a methyl group transfer from N -CH3-THF with formation of MeCbl and THF. In the second step, MeCbl transfers this methyl group to homocysteine, regenerating the cofactor cobalamin and releasing the product methionine.

FIG. 1.12. Methionine synthesis catalyzed by MeB12-dependent methionine synthase (73).

In many anaerobic bacteria, methyl-corrinoids are involved in acetyl-CoA synthesis via the Wood-Ljungdahl pathway (176, 220, 243) (Fig. 1.13A). In this pathway, a methyl group is transferred from CH3-THF via a methyl-corrinoid/iron sulfur protein to

CO-dehydrogenase, which synthesizes acetyl-CoA from this methyl group, CO, and coenzyme A (77). A corrinoid plays an analogous role in the energy-yielding metabolism of acetogenic bacteria, which synthesize acetate from 2 CO2 as a means of generating a

terminal electron sink (176).

Methyl-corrinoids are also essential for formation of methane by the strictly anaerobic methane-producing archaea (220) (Fig. 1.13B). Corrinoid proteins play a role in the transfer of methyl groups from methanogenic substrates to the thiol group of the methanogen-specific cofactor, coenzyme M. Different enzymes mediate methyl transfer from alternative methanogenic substrates such as acetate (78), methylamines (44), methanol, pyruvate (33) and methyltetrahydromethanopterin (CH3-H4MPT), an intermediate of methanogenesis from formate and CO2 (169). The latter reaction is analogous to methionine synthesis in that the methyl group is transferred from an intermediate pterin to a thiol group via MeCbl by an integral membraneme thyltransferase

(169). Considerable energy is released by transfer of a methyl group to coenzyme M from

CH3-H4MPT (93, 220). The energy of the transfer to coenzyme M can be recovered by coupling the methyl transfer to extrusion of a sodium ion, which eventually leads to generation of a proton motive force (19).

FIG. 1.13. Methyltransferases involved in anaerobic CO2 fixation and energy metabolism (18). A The Wood-Ljungdahl Pathway. B MeCbl-dependent methyltransferases in methanogenesis. ACS, acetyl-CoA synthase; CODH, CO dehydrogenase; CFeSP, corrinoid iron-sulfur protein; and MeTr, AcsE-methyltransferase.

1.1.4.3. Corrinoid-dependent reductive dehalogenation

Bacterial reductive dechlorination reactions play important roles in the detoxification of chlorinated aromatic and aliphatic compounds in contaminated environments (101,

135). Both aerobic and anaerobic microorganisms can perform dehalogenation; however, anaerobic bacteria are more efficient in removing halogen atoms from polyhalogenated compounds (71).

Several anaerobic bacteria use chlorinated compounds as growth-supporting electron acceptors in their energy metabolism, coupling reductive dehalogenation to electron transport phosphorylation (103, 140). Reductive dehalogenases catalyzing the reductive dechlorination of chlorinated ethenes, phenols, or benzoates have been isolated from several species (54, 142, 150, 160, 162, 205):

R-Cl + 2e- + 2H+ → RH + HCl

There are two major classes of anaerobic dehalogenases: the heme containing

3-chlorobenzoate reductive dehalogenase from Desulfomonile tiedjei (162) and the vitamin B12-dependent enzymes such as that from Desulfitobacterium chlororespirans

(123). The B12-dependent reductive dehalogenases, most of which also contain iron-sulfur clusters, are either embedded in or attached to the membrane by a small anchoring protein.

This membrane association is important in linking the dehalogenation reaction to the electron transport pathway of anaerobic respiration. A few B12-dependent dehalogenases have been purified, including the haloalkane (perchloroethylene and trichloroethene) dehalogenases from Desulfitobacterium strain PCE-S (150) and the

3-chloro-4-hydroxy-phenylacetate dehalogenases from Desulfitobacterium hafniense (54).

These proteins contain a corrinoid, a [4Fe-4S] cluster, and a [3Fe-4S] cluster per monomeric unit. Multiple dehalogenases are evident in the genome sequence of D. hafniense strain DCB-2. Thus, the ability of certain dehalorespiring microorganisms to

metabolize various types of chlorinated organic compounds (e.g., chloroalkanes plus chlorobenzoates) is conferred by separate (but homologous) enzymes (232).

FIG. 1.14. Alternative mechanisms for corrinoid-dependent reductive dehalogenation (18).

The role of B12 in the reductive dehalogenases appears to be significantly different from those of the AdoCbl-dependent isomerases and the MeCbl-dependent methyltransferases. Mechanistic studies on the anaerobic dehalogenases have lagged behind studies on the aerobic enzymes. Two working models for the mechanisms of dehalorespiring reductive dehalogenases are shown in Figure 1.14. In Path A, an organocobalt adduct is formed as in the methyltransferases. In Path B, the corrinoid serves as an electron donor. In Path A, the halide is eliminated as the aryl-Co(III) intermediate is formed. Path B resembles the Birch reduction of hydroxylated aromatics in which a radical anion is formed. Both mechanisms assume that the corrinoid is the site of dehalogenation based on the ability of B12 to catalyze dehalogenation in solution (124) and light-reversible inhibition by propyl iodide, a characteristic of corrinoid-dependent reactions (149, 159).

1.1.5. Vitamin B12-related diseases

There are two B12-dependent reactions in the human body: methionine synthase and methylmalonyl-CoA mutase.

Methionine synthetase, a methyl transferase, is presumed to be important primarily in recycling folate and secondarily in producing methionine (3). Methionine synthase is the only enzyme present in higher animals that uses CH3-THF as a substrate. Thus, a defect in methionine synthase or B12 traps folate as CH3-THF resulting in a subsequent deficiency of THF. Since THF is an indispensible cofactor functioning in synthesis of purines and pyrimidines, the lack of this reaction underlies many aspects of human B12-deficiency disorders including pernicious anemia and megaloblastosis.

Methylmalonyl-CoA mutase may serve mainly to remove toxic products of lipid breakdown (133). The mutase or B12 deficiency results in an often fatal methylmalonic acidemia (133), and in certain neuropsychiatric symptoms. These symptoms may result from synthesis of abnormal myelin lipids in the presence of accumulated methylmalonic acid (MMA) (3, 175), a myelin (a dielectric material that forms myelin sheath around the axon of a neuron) destabilizer. Excessive MMA will prevent normal fatty acid synthesis, or it will be incorporated into fatty acid itself rather than normal malonic acid. If this abnormal fatty acid subsequently is incorporated into myelin, the resulting myelin will be too fragile, and demyelination will occur. Although the precise mechanism(s) are not known with certainty, the result is subacute combined degeneration of central nervous system and spinal cord (157).

Thus, disorders of cobalamin transport and metabolism (Fig. 1.15), whether inherited or acquired, manifest themselves as deficiencies in the activities of one or both of these enzymes and lead to accumulation in blood and urine of one or both of the substrates, homocysteine and methylmalonyl-CoA, respectively.

FIG. 1.15. Schematic view of the intracellular B12 trafficking pathway (17). The known and proposed functions of the gene products associated with the complementation groups cblA-G and mut are indicated. The question marks highlight some aspects of the trafficking pathway that are poorly characterized.

Clinical studies on patients with inborn errors of cobalamin metabolism identified eight genetic complementation groups that underlie this disease, cblA-G and mut (87, 241), and provided early insights into a complex pathway for intracellular processing of B12. From such studies, these cobalamin disorders can be divided to three subclasses: those resulting

(i) in combined defects in both the MeCbl and AdoCbl branches (cblF, cblC, and cblD),

(ii) in an isolated defect in MeCbl branch (cblG and cblE), and (iii) in an isolated defect in AdoCbl branch (cblA, cblB, and mut). To date, all eight genes responsible for the defects in intracellular cobalamin metabolism have been identified but some of their functions await elucidation: the gene LMBRD1 (cblF locus) encodes a B12 transporter;

MMACHC (cblC) encodes a decynase and dealkylase; MMADHC (cblD) probably codes for a transpoter or an adaptor protein; the cblG locus encodes methionine synthase; cblE encodes methionine synthase reductase; cblA encodes MMAA with functions of editing, gating and radical protection; cblB codes for adenosyltransferase and mut encodes methylmalonyl-CoA mutase (17).

1.2. Nicotinamide adenine dinucleotide (NAD+)

Nicotinamide adenine dinucleotide, abbreviated NAD+, is a coenzyme found in all living cells and also acts as substrate for some enzymes. NAD+ has been central to many of the greatest discoveries in the biological sciences.

1.2.1. Discovery and structure of NAD+

In the early 20th century, Arthur Harden and William Youndin reconstituted cell-free glucose fermentation with two fractions, one termed ‘zymase’ that was heat-labile and retained by dialysis, and one termed ‘cozymase’ that was heat-stable and passed through dialysis (92). Zymase was not a purified enzyme but a protein fraction that contained glycolytic enzymes. The cozymase fraction contained ATP, Mg2+ and the NAD+ coenzyme. Years later, Hans von Euler and Otto Warburg independently determined the chemical structure of NAD+ (198, 236). The compound is a dinucleotide, since it consists of two nucleotides joined through their phosphate groups, with one nucleotide containing an adenine base and the other containing nicotinamide (Fig. 1.16).

FIG. 1.16. The structure of Nicotinamide adenine dinucleotide (NAD+). A, Structural formula of NAD+; B, Stick model of NAD+.

1.2.2. Biosynthesis of NAD+

According to a widely accepted concept, a combination of de novo pathway from

amino acids and salvage pathways by recycling preformed components contributes to the biosynthesis of NAD+ (166, 187). Some NAD+ is also converted into nicotinamide adenine dinucleotide phosphate (NADP+) that has the chemistry similar to that of NAD+ but different roles in metabolism.

FIG. 1.17. NAD+ biosynthesis (de novo and salvage) pathways (adapted from references (34, 115)). Two de novo pathways, the aspartate pathway and kynurenine pathway, converge at quinolinic acid, and subsequently use three common steps to synthesize NAD+. In salvage metabolic pathways, three natural nicotinamide-ring contained compounds, nicotinic acid (Na), nicotinamide (Nam), and nicotinamide riboside (NR), and possibly nicotinic acid riboside (NaR), are recycled back to the NAD+.

1.2.2.1. De novo production

Most organisms synthesize NAD+ from simple building-blocks from the amino acids tryptophan or aspartic acid (23). The specific set of reactions differs among organisms, but a common feature is the generation of quinolinic acid from an amino acid - either tryptophan (Trp) (kynurenine pathway) in animals and some bacteria, or aspartic acid

(Asp) (aspartate pathway) in some bacteria and plants (21, 83, 116, 137) (Fig. 1.17). The quinolinic acid is converted to nicotinic acid mononucleotide (NaMN) by transfer of a

phosphoribose moiety. An adenylate moiety is then transferred to form nicotinic acid adenine dinucleotide (NaAD). Finally, the nicotinic acid moiety in NaAD is amidated to a nicotinamide moiety, forming nicotinamide adenine dinucleotide (23, 34, 115).

1.2.2.2. Salvage pathways

In an alternative fashion, more complex components of the coenzymes are taken up from food as the vitamin called niacin (B3) which is a mixture of nicotinic acid and nicotinamide. Similar compounds such as nicotinic acid (Na), nicotinamide (Nam), and nicotinamide riboside (NR) are released by reactions that break down the structure of

NAD+. These precursors are fed into the NAD+ biosynthetic pathway, shown in Figure

1.17, through adenylation and phosphoribosylation reactions that recycle them back into the active form (34, 115, 146).

Salvage pathway is essential, especially for some microorganisms such as yeast

Candida glabrata and the bacterium Haemophilus influenzae which are NAD+ auxotrophs - they cannot synthesize NAD+ de novo - but possess salvage pathways and thus are dependent on external sources of NAD+ or its precursors (141, 179).

1.2.3. Function of NAD+

NAD+ has several essential roles in all living systems. For several decades, it has been known that NAD+ acts as a coenzyme in redox reactions. In recent years, multiple enzyme-mediated, nonredox roles for NAD+ have been discovered, such as acting as a donor of ADP-ribose moieties in ADP-ribosylation reactions, as a precursor of the second messenger molecule cyclic ADP-ribose, as well as acting as a substrate for a group of enzymes called sirtuins that use NAD+ to remove acetyl groups from proteins and as a dehydrating agent for bacterial DNA ligases.

1.2.3.1. Oxidation-reduction reactions

NAD+ is classically known to play a major role as a coenzyme in numerous metabolic reactions. The main function of NAD+ in metabolism is to act as an electron carrier to transfer electrons from one molecule to another, involved in redox reactions. Such

+ + reactions (RH2 + NAD → NADH + H + R) involve the removal of two hydrogen atoms

− + from the reactant RH2, in the form of a hydride ion (H ), and a proton (H ). The proton is

+ released into solution, while the reductant RH2 is oxidized and NAD was reduced to

NADH by transfer of the hydride to the nicotinamide ring. NADH can then be used as a reducing agent to donate electrons and be re-oxidized to NAD+ (Fig. 1.18). This means the coenzyme can continuously cycle between the NAD+ and NADH forms without being consumed (170). These electron transfer reactions are the main function of NAD+ and are catalyzed by a large group of enzymes called oxidoreductases.

FIG. 1.18. NAD+ as a coenzyme for reversible hydride transfer (23).

The redox reactions are vital in all parts of metabolism, but one particularly important area where these reactions occur is in the release of energy from nutrients. Here, reduced compounds such as glucose are oxidized, thereby releasing energy. This energy is transferred to NAD+ by reduction to NADH, as part of glycolysis and the citric acid cycle, and then to respiratory chain where the energy eventually is released through oxidative phosphorylation.

1.2.3.2 Regulatory reactions

NAD+ serves as a substrate for covalent modifications of target molecules and is also

involves in signal transduction as precursors of messenger molecules (249). All known derivatives share structural similarity owing to their ADP-ribose backbone (Fig. 1.19).

Thus, NAD+-mediated signalling reactions can be referred to as ADP-ribosyl transfers, because the initial cleavage of NAM (leaving the ADP-ribosyl moiety) is common to all of them. The eventual acceptors of the transfer reaction are rather diverse, including macromolecules such as proteins, or small molecules such as water (yielding ADP-ribose), acetate [yielding OAADPr (O-acetyl ADP-ribose)] or the adenine ring leading to intramolecular cyclization into cADPr (cyclic ADP-ribose; Fig. 1.19) (170).

FIG. 1.19. Signalling derivatives of NAD+ (170).

ADP-ribosylations are catalyzed by mono-ADP-ribosyltransferases (ARTs) and poly(ADP-ribose polymerases (PARPs), which involves either the addition of a single

ADP-ribose moiety or the transferral of ADP-ribose to aceptors in long branched chains, respectively (170). Mono-ADP-ribosylation was originally identified as the mechanism of a group of bacterial toxins, notably cholera toxin, but it is also involved in normal cell signaling (24, 58, 231). The poly(ADP-ribose) structure is involved in the regulation of several cellular events and is most important in the cell nucleus, in processes such as

DNA repair and telomere maintenance (45).

NAD+ is also the precursor of cyclic ADP-ribose, which is produced from NAD+ by

ADP-ribosyl cyclases (or cADP-ribose synthases), as part of a second messenger system

(88). This molecule acts in calcium signaling by releasing calcium from intracellular stores (89). It does this by binding to and opening a class of calcium channels called ryanodine receptors, which are located in the membranes of organelles, such as the endoplasmic reticulum (90).

Sirtuins such as silent information regulator 2 (Sir2) are another class of

NAD+-consuming enzymes, which transfer an acetyl group from their substrate protein to the ADP-ribose moiety of NAD+; this cleaves the coenzyme and releases nicotinamide and O-acetyl-ADP-ribose. The sirtuins mainly seem to be involved in regulating transcription through deacetylating histones and altering nucleosome structure (27, 107).

1.2.3.3. DNA ligation

Other NAD+-dependent enzymes include bacterial DNA ligases, which join two DNA ends by using NAD+ as a substrate to donate an adenosine monophosphate (AMP) moiety to the 5' phosphate of one DNA end. This intermediate is then attacked by the 3' hydroxyl group of the other DNA end, forming a new phosphodiester bond (83, 134, 240). This contrasts with all DNA ligases identified within archaea, viruses and eukarya, which use

ATP to form DNA-AMP intermediate (134, 213, 227, 240).

1.2.4. NAD+-related diseases

Due to its ubiquitous coenzyme roles and specific signaling functions, NAD+ is indispensible in cellular metabolism and regulation. A lack of NAD+ or its precursors such as niacin and tryptophan in the diet causes the vitamin deficiency disease pellagra

(100). This high requirement for NAD+ results from the constant consumption of the molecule in reactions like histone modifications, DNA repair, and telomere maintenance in cell nucleus (23, 45), accounting for that the enzymes involved in the salvage pathways appear to be concentrated in this organelle (6).

1.3. AdoCbl-dependent 1,2-propanediol (1,2-PD) degradation

1.3.1. Source of 1,2-PD

1,2-PD is a major product of the fermentation of the sugars L-fucose and L-rhamnose

(2, 10, 12, 35, 36, 138) (Fig. 1. 20). These two methyl pentoses are metabolized in E. coli and S. enterica through inducible pathways that show a striking parallelism. Both pathways are sequentially mediated by a permease, an isomerase, a kinase, and an aldolase (2, 36). The stereochemical difference between the configurations of both sugars at carbon 2 and carbon 4 disappears with cleavage of the fuculose-1-phosphate or rhamnulose-1-phosphate by the corresponding aldolases, yielding in each case dihydroxyacetone phosphate (DHAP) and L-lactaldehyde. The two homologous sets of inducible proteins are specific for the metabolism of their corresponding sugars and are coded for by two different gene clusters.

FIG. 1.20. Pathway of L-fucose and L-rhamnose dissimilation (adapted from ref. (12, 36)).

L-lactaldehyde is a branching point in the metabolic pathway of L-fucose and

L-rhamnose utilization. Further metabolism of L-lactaldehyde follows two different pathways, depending on the availability of oxygen. Under aerobic conditions, the aldehyde is oxidized to L-lactate in an irreversible reaction catalyzed by the

NAD+-dependent lactaldehyde dehydrogenase and L-lactate is further oxidized to pyruvate by a dehydrogenase of the flavoprotein class to enter central metabolism.

Whereas under anaerobic conditions, L-lactaldehyde is reduced to L-1,2-propanediol by the enzyme propanediol oxidoreductase in a typical fermentative mechanism in which

NADH is oxidized to NAD+ (12, 35, 36).

1.3.2. 1,2-PD utilization

In E. coli, 1,2-PD is excreted to the environment as a fermentation product (55). In contrast, a number of enteric and lactic acid bacteria such as S. enterica (163) further utilize 1,2-PD in a coenzyme B12-dependent fashion.

FIG. 1.21. Anaerobic metabolism of 1,2-PD.

The first step in this degradative pathway is the AdoB12-dependent diol dehydratase, which produces propionaldehyde (Fig. 1.21). This aldehyde can be oxidized to form propionyl-CoA by propionaldehyde dehydrogenase or alternatively can be reduced to form propanol by propanol dehydrogenase. Through forming and excreting propanol, cells can balance internal redox reactions. Under anaerobic conditions, propionyl-CoA can be converted by a phosphotransacylase to propionyl-phosphate, and then by a reversible acetyl-/propionyl-kinase to propionate, producing one molecule of ATP.

According to this scheme, anaerobic metabolism of 1,2-PD could provide an electron sink

(propanol) and a source of ATP but no source of carbon since both propanol and propionate are excreted (188).

Under aerobic conditions, S. enterica can use propionyl-CoA as both a carbon and an energy source through a methycitrate pathway (105) (Fig. 1.22).

FIG. 1.22. Pathway for propionate catabolism in S. enterica (105). PrpE, propionyl-CoA synthetase; PrpC, 2-methylcitrate synthase; PrpD, 2-methylcitrate dehydratase; PrpB, 2-methylisocitrate lyase; Pps, PEP synthetase; Ppc, PEP carboxylase.

1,2-PD is a major product of the anaerobic degradation of L-rhamnose and L-fucose and is thought to be an important carbon and energy source in natural environments (138).

These two sugars are common in plant cell walls, bacterial capsules (exopolysaccharides) and the glycoconjugates of eukaryotic (intestinal epithelial) cells. So the ability to degrade 1,2-PD is thought to provide a selective advantage in anaerobic environments such as the large intestines of higher animals, sediments and the depths of soils. The utilization ability is found in a number of bacterial genera including Salmonella, Shigella,

Lactococcus, Lactobacillus, Yersinia, Listeria, and Klebsiella (53).

1.3.3. The pdu locus in S. enterica

S. enterica degrades 1,2-PD via a coenzyme B12-dependent pathway (110).

Twenty-four genes specifically involved in 1,2-PD utilization (pdu) are found in a single contiguous cluster, the pdu locus (31, 110) (Fig. 1.23). This locus consists of the pocR and pduF genes as well as the adjacent, but divergently transcribed pdu operon which includes 22 pdu genes (pduABB'CDEGHJKLMNOPQSTUVWX, pduB and pduB’ arise

from alternate start sites.). The pdu locus encodes a transcriptional regulator (30, 184), a diffusion facilitator (51), enzymes for the degradation of 1,2-PD (31, 32, 129, 139) and cobalamin metabolism (31, 74, 113, 156, 194), as well as proteins for the formation of a bacterial microcompartment (MCP) used for 1,2-PD utilization (31, 94, 95, 193). These genes for 1,2-PD utilization (pdu) were acquired together with 20 genes for AdoCbl biosynthesis by a single horizontal gene transfer event that is thought to be one of several related events important to the divergence of S. enterica and E. coli (31, 128, 189).

FIG. 1.23. The pdu locus of S. enterica.

Based on experimental evidence and/or sequence similarity the proteins encoded by the pdu locus can be categorized into six groups: PocR, transcriptional regulator (30, 184);

PduF, 1,2-PD diffusion facilitator (51); PduA, PduB, PduB’, PduJ, PduK, PduN, PduT, and PduU, MCP shell proteins (31, 62, 95); PduCDE, PduL, PduP, PduQ, PduW, 1,2-PD degradative enzymes (diol dehydratase, phosphotransacylase, propionaldehyde dehydrogenase, propanol dehydrogenase, and propionate kinase, correspondingly) (31, 32,

129, 139, 164); PduGH, PduO, PduS, and PduX, cobalamin synthesis and recycling (diol dehydratase reactivase, adenosyltransferase, cobalamin reductase, and L-threonine kinase, correspondingly) (31, 52, 74, 113, 156, 194); and PduM and PduV, unknown function - possibly an MCP shell protein and a GTPase (53).

1.3.4. Control of the pdu/cob regulon

The pdu operon maps adjacent to the cob operon which encodes B12 synthetic enzymes in S. enterica. The two operons are both induced by 1,2-PD using a single regulatory protein. Two global regulatory systems (Crp/Cya and ArcA/ArcB) affect inducibility of the cob and pdu operons (1, 7, 72). Both operons are activated aerobically and

anaerobically by Crp protein and anaerobically by ArcA protein. The control of the regulon depends on five promoters, all located in the central region between the cob and pdu operons; four of these promoters are activated by the PocR protein (50).

FIG. 1.24. Regulation of the cob/pdu regulon of S. enterica (188). Boxes enclose structural genes. Black arrows designate transcripts. Gray arrows indicate regulatory influence; dashed gray arrows indicate the proposal that a higher level of PocR protein may be required to activate these promoters.

The current model for control of this system includes the following features (Fig. 1.24):

The cob and pdu operons are transcribed divergently from promoters indicated at the far right and left in the figure. These promoters are activated whenever both the PocR protein and 1,2-PD are present. Global control of the two operons is exerted by varying the level of PocR protein. The pocR gene is transcribed by three promoters controlled both by global regulatory proteins and by autoinduction (50). The shortest transcript (from Ppoc) appears to be regulated only by Crp/cAMP. The P2 transcript clearly is autoregulated by

PocR and, in addition, requires the ArcA protein, which signals a reduced cell interior

(50). Control of the P1 promoter is less certain but may involve either the Fnr protein

(responding to a reduced cell interior) or the Crp protein (responding to a shortage of carbon and energy) in addition to the PocR activator. The involvement of these proteins in control of P1 is based heavily on the presence of appropriate binding sites upstream of that promoter.

It is proposed that the regulon has three states (188). In the off condition (during aerobic growth on glucose), all promoters are at their lowest level; PocR protein is

produced by the basal expression levels of promoters P1, P2, and Ppoc. Cells enter a standby state when they grow under any set of global conditions appropriate for induction, but without 1,2-PD. These conditions include aerobic growth on a poor carbon source and/or growth without oxygen. These conditions stimulate expression of both the PduF transporter and the PocR regulatory protein, but the cob and pdu operons remain uninduced. The standby expression of PocR can occur from the Ppoc and/or the P1 promoter without inducer. The two proteins induced in the standby state (PduF and PocR) are those needed to sense 1,2-PD. If 1,2-PD appears, the P1 and P2 promoters are induced, increasing the level of PocR protein and placing the system in the on state. The resulting high level of PocR/1,2-PD complex induces expression of the cob and pdu promoters.

These results indicate that 1,2-PD catabolism is the primary reason for de novo B12 synthesis in S. enterica (30, 189). If one includes the cob genes, S. enterica maintains nearly 50 genes primarily for the degradation of 1,2-PD. Virtually all natural Salmonella isolates tested synthesize AdoCbl de novo and degrade 1,2-PD (128).

1.3.5. Pathogenesis

The capacity for 1,2-PD degradation is found among a number of genera that inhabit the large intestines of higher animal including some pathogens. Studies conducted with two intracellular human pathogens, Salmonella and Listeria, have implicated 1,2-PD degradation in pathogenesis. In Salmonella, pdu genes are induced in host tissues and co-regulated with invasion genes by CsrA (57, 126). In addition, pdu mutants are impaired for growth in macrophages and show a virulence defect in competitive index assays with mice (98, 121). In Listeria, comparative genomics indicates a link between

1,2-PD degradation and pathogenesis, and several pdu genes are induced during intracellular growth in host cells (43, 114).

1.4. Bacterial microcompartment

Many bacteria conditionally express protein-based organelles referred to here as microcompartments (MCPs). These MCPs are thought to be involved in at least seven different metabolic processes ranging from CO2 fixation to the degradation of small organic molecules, and the number is growing (53). MCPs are very large and structurally sophisticated. They are usually about 100-150 nm in cross-section and consist of

10,000-20,000 polypeptides of 10-20 types. Their unifying feature is a solid shell constructed from proteins having bacterial microcompartment (BMC) domains that are widely distributed across the bacterial kingdom. Encased within the shell are sequentially-acting metabolic enzymes. MCPs are made completely of protein subunits

(49, 94, 104). There are no confirmed reports of RNA, DNA, or lipids functionally associated with the MCPs that have been investigated. In the cases that have been studied, the metabolic sequences associated with MCPs have toxic or volatile intermediates. It is thought that the shell of the MCP confines such intermediates, thereby enhancing metabolic efficiency and/or protecting cytoplasmic components.

1.4.1. Discovery and diversity

Carboxysomes were the first bacterial MCPs to be discovered (Fig. 1.25 A, B). They were observed more than 50 years ago as inclusion bodies with a polygonal appearance in the cytoplasm of cyanobacteria Phormidium uncinatum (68). Initially, carboxysomes were mistaken for viruses because of their resemblance to phage particles. Over the next 20 years, these inclusions, which were named polyhedral bodies in 1961 (109), were found in various other cyanobacteria (for example, Nostoc punctiforme, Synechococcus elongatus, Anabaena cylindrica and Symploca muscorum) and chemoautotrophs (for example, Halothiobacillus neapolitanus, Acidithiobacillus ferrooxidans, Nitrobacter

winogradskyi and Nitrococcus mobilis) (53, 209, 245). Polyhedral bodies were first isolated in 1973 from the chemoautotrophic bacterium H. neapolitanus by differential and sucrose step-gradient centrifugation, following rupture in a French pressure cell (210).

These bodies were surrounded by a proteinaceous envelope (the shell) and contained the enzymes ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO) and carbonic anhydrase (CA) involved in CO2 fixation; to reflect this finding, the name carboxysome was proposed (85, 210). It is now widely accepted that although carboxysomes are probably found in all cyanobacteria, they are present in only a limited number of chemoautotrophs: namely, some of the sulphur and nitrifying bacteria (for example, species of Thiobacillus, Halothiobacillus, Thiomonas, Acidithiobacillus, Nitrobacter and

Nitrosomonas) (48, 96, 245). A C D

FIG. 1.25. Electron micrographs of MCPs (100-150 nm in cross section) A. The carboxysomes (α-type) in H. neapolitanus; B. The purified carboxysomes (α-type) from H. neapolitanus. C. The Pdu MCPs in S. enterica grown on 1,2-PD; D. The purified Pdu MCPs from S. enterica.

A variety of studies indicate two main classes of carboxysomes, α and β, which vary slightly in composition. Each class is thought to have co-evolved with a different phylogenetic group of RuBisCO (8, 9). The α-carboxysome is associated with form 1A

RuBisCO which is found in chemoautotrophs and α-cyanobacteria while the

β-carboxysome is associated with form 1B RuBisCO which is found in the

β-cyanobacteria (9). Overall, α- and β-carboxysomes are thought to have similar architectural and mechanistic principles.

For many years, carboxysomes were the only known bacterial MCPs. Then, in 1994, genetic studies and sequence analyses indicated that the PduA protein (encoded by the pdu locus that is involved in B12-dependent 1,2-PD degradation by S. enterica) exhibited relatively high sequence identity to carboxysome shell protein CcmK (51). A few years later, electron microscopy showed that S. enterica conditionally formed MCPs during

B12-dependent growth on 1,2-PD (211) (Fig. 1.25 C, D). Further work indicated that seven more Pdu proteins (PduB/B’/J/K/N/T/U) are related to the shell proteins involved in the formation of carboxysomes (31), providing additional evidence for Pdu MCP formation. A simultaneous study on B12-dependent ethanolamine utilization (eut) by S. enterica indicated that five eut genes (eutK/L/M/N/S) encode homologues of the shell proteins of the carboxysomes and the Eut MCPs were observed by electron microscopy as well (40, 122).

Genomic analyses indicated that bacterial MCPs are widespread and functionally diverse. Approximately 20-25% of bacteria encode homologues of BMC-domain proteins, although none has been detected in the archaea or eukarya (29). Multiple genes for

BMC-domain proteins are often found in gene clusters that also encode MCP-associated enzymes or putative enzymes. Variation in such gene clusters suggests bacterial MCPs have diverse functions. Established MCPs include carboxysomes and the Pdu and Eut

MCPs. Several putative MCPs are inferred from gene clustering analyses: 1. A MCP associated with a pyruvate-formate lyase homolog is proposed to be involved in the production of ethanol from pyruvate (234); 2. A MCP was suggested to be involved in the oxidation of ethanol by Clostridium kluyveri (208); 3. A putative MCP in Rhodopirellula baltica SH 1 is associated with a lactate dehydrogenase homologue; 4. A putative MCP in

Carboxydothermus hydrogenoformans Z-2901 is associated with an isochorismatase-family protein; 5. A putative MCP Solibacter usitatus Ellin6076

associated with a dihydrodipicolinate synthase homologue, and the list is expected to grow as more genome sequences become available (29). Certainly, this diversity is one of the most intriguing aspects of bacterial MCPs. It raises important questions about their functional principles, the extent to which those principles are conserved, and the degree to which they can be adapted and modified (or engineered) to fulfill the needs of a particular metabolic system (53).

1.4.2. Component and architecture

For many years there was uncertainty about the overall shape of carboxysomes. Recent electron cryotomography of carboxysomes from H. neapolitanus (α-type) and

Synechococcus strain WH8102 (β-type) indicate an icosahedral or quasi-icosahedral structure (106, 200) (Fig. 1.25 B). The shape of Pdu MCPs is also roughly polyhedral, but they are more irregular than carboxysomes (Fig. 1.25 D). Eut MCPs have not been purified (117, 244), but the unisolated Eut MCPs in Salmonella cells look like carboxysomes and Pdu MCPs under the electron microscope (40).

In the lumen of the MCPs, there are several enzymes presumed to carry out required metabolic functions. For example, the carboxysomes consist mainly of RuBisCO and CA involved in CO2 fixation (85, 210), while the Pdu MCPs contain PduCDE, PduGH, PduO, and PduP, functioning in 1,2-PD degradation (94, 129).

Gene sequencing and MCP purification experiments demonstrated that MCPs are composed of a few protein components in addition to those enzymes (49, 94, 104, 173).

The most abundant nonenzymatic components purified from both carboxysomes and Pdu

MCPs are members of a conserved family of small proteins, typically around 100 (single) or 200 (tandem) amino acids in length, that define the BMC protein domain (InterPro domain IPR000249, Pfam 00936). These BMC proteins are the major constituents of

MCP shells. Furthermore, in all organisms for which genomic data have revealed a BMC protein for a presumptive MCP, multiple paralogous BMC proteins have been found. For example, CsoS1A/B/C/D are present in α-type carboxysome, CcmK/O in β-type,

PduA/B/B’/J/K/T/U for Pdu, and EutL/M/K/S for Eut. The involvement of multiple distinct BMC proteins in the shell appears to be a strongly conserved feature (20, 118).

To date, crystal structures of 13 different BMC proteins have been determined, some in multiple crystal forms and conformations. These include three (CsoS1A/C/D) from the

α-type carboxysome (120, 229, 230), two (CcmK2/4) from the β-type (118), three

(PduA/T/U) from the Pdu MCP (60, 62), four (EutK/L/M/S) from the Eut MCP (191,

224), and one (EtuB) predicted to function in an ethanol-utilizing (Etu) MCP (99). From these crystal structures, basic features of assembly and transport have been inferred.

The BMC domain adopts an α/β-fold with a central four-stranded antiparallel sheet flanked by small helices. Single BMC-domain proteins usually form flat hexamers with cyclic (C6) symmetry, or some variation on that arrangement, while the shell proteins with two tandem BMC domains form hexagonal trimers (pseudohexamers) (117) (Fig.

1.26). Numerous lines of evidence therefore show that hexamers and pseudohexamers comprise the building blocks of bacterial MCP shells (118, 229, 247). Most BMC shell proteins adopt structures in which the hexamer has a prominent bowl-shaped depression on one side, specifically the side on which both the N and C termini reside in the typical

BMC domain (244).

Several different crystal structures of BMC proteins reveal tightly packed two-dimensional layers of molecules (118, 191, 229, 230) (Fig. 1.26). The shape of the

BMC hexamers and pseudohexamers are evidently tailored for further side-by-side assembly into a mixed molecular sheet, roughly 20 Å thick on average, with a center-to-center distance between hexamers/pseudohexamers ranging from 66 to 69 Å,

suggesting that MCP shells are a mosaic of different types of BMC-domain proteins. The molecular layer is nearly solid, except for narrow pores through the centers of the hexamers/pseudohexamers and where hexamers/pseudohexamers meet at twofold and threefold axes of symmetry. Another BMC protein (EutS) is a bent hexamer that could form the edges of the shell (224). The presence of BMC shell proteins effectively defines the bacterial MCP family of organelles (244).

FIG. 1.26. The elements of the architecture of the carboxysome shell, modeled as an icosahedron assembled from hexamers (single BMC domain proteins, blue), pseudohexamers (tandem BMC domain proteins, green, gold, and pink), and pentamers (quaternary structure of Pf03319 domain in CcmL, yellow) (117).

In addition to the major (BMC-type) shell proteins, the presence of one minor protein is also conserved across MCPs and is named according to MCP type: CsoS4A/B

(α-carboxysome), CcmL (β-carboxysome), PduN, or EutN - collectively referred to as the

CcmL/EutN protein family (Pfam 03319). Proteins from the CcmL/EutN family are found encoded in operons wherever BMC genes are present (244). However, the homologous

gene products in the α-carboxysome operon (CsoS4A/B) were not detected initially in purified H. neapolitanus carboxysomes (49), but the deletion of the ccmL gene in cyanobacterial β-carboxysomes gave rise to aberrant, elongated tubular carboxysomes

(173). Crystal structures of corresponding proteins, CsoS4A and CcmL, from the two types of carboxysomes clarified their structural roles. Both proteins are pentamers whose sizes and shapes are compatible with formation of vertices in an icosahedral shell assembled mainly from BMC hexamers (222). On that basis, rough atomic models of carboxysome shells have been constructed (Fig. 1.26). Models in which the CcmL or

CsoS4 proteins occupy icosahedral vertices explain the two key observations noted above.

First, vertex proteins would be present in only 60 copies per shell, compared to a few thousand for the BMC proteins and the RuBisCO subunits, explaining difficulties in detection; later studies confirmed the expression of the csoS4A/B genes (47). Second, in the absence of pentagonal vertices, a flat layer of hexamers can roll up into a tube, whereas introducing the (Gaussian) curvature required to close the ends is problematic without pentamers (244).

In different MCPs, multiple proteins are present that are apparently neither enzymatic nor homologous to the major BMC or the minor CcmL/EutN shell protein family. Such proteins are presumed either to help organize the MCP enzymes, or to be minor structural components of the shell. There is considerable variation regarding the accessory proteins present in different MCPs (244).

Like the lipid bilayer of membrane bound-organelles, the bacterial MCP shell has conduits through which metabolites selectively pass. The canonical BMC proteins whose structures have been determined typically have pores with diameters ranging from 4 to 6

Å (118, 229, 247). These pores are likely large enough to permit the diffusion of small

− substrates and products of the known MCPs (e.g., bicarbonate (HCO3 ), 3-PGA,

1,5-RuBP, ethanolamine, and 1,2-PD). Positive electrostatic potential has been noted in the hexameric pores of the major BMC proteins of the carboxysome (223, 229). It has

− been suggested that this could confer a kinetic advantage for HCO3 diffusion into the shell. CO2 and O2, whose respective outward and inward passages are undesirable, would

− not benefit from the same electrostatic advantage as HCO3 (244).

A recent study showed the structure of a newly identified carboxysome shell protein,

CsoS1D, existing in two dramatically different conformations (120). The two conformations of CsoS1D correspond to an open and closed central pore. The closed form is essentially occluded, whereas the open form has a pore roughly 14 Å in diameter.

Likewise, EutL has been visualized in two different conformations, one that is nearly occluded (191, 224) and one that has a roughly 11 Å pore (224). In addition to offering an explanation of how larger molecules might be transported, the two conformations suggest a potential gating mechanism that could control which molecules pass through the shell

(120, 224). This would help address the question of how larger molecules might be able to pass through a shell that presents a diffusion barrier to smaller compounds (244).

A second potentially significant observation is that CsoS1D tends to form a double-disk structure via a face-to-face interaction of two trimers (or pseudohexamers). This creates a large interior cavity. In two different crystal forms, the double-disk is composed of one trimer in the open configuration and a second in the closed configuration, leading the investigators to suggest an alternating access mechanism (120). The discovery of dynamic shell proteins (CsoS1D and EutL) hints that further studies on diverse MCP proteins will reveal a multitude of varied pores and transport mechanisms (244).

1.4.3. Function and mechanism

Data have been presented for the existence of bacterial MCPs carrying out at least

seven different metabolic pathways. Three of these pathways have been explored experimentally (carboxysomes, Pdu and Eut MCPs).

Carboxysome is thought to work in conjunction with C1 transport systems as part of a bacterial CO2 concentrating mechanism (CCM) thereby enhancing carbon fixation and autotrophic growth (53, 172, 244). It is now generally understood that CA is colocalized with RuBisCO inside the carboxysome (59, 97, 171) and that this is a key mechanistic feature for enhancing CO2 fixation when the concentration of inorganic carbon is low, as it is in most natural environments. The carboxysome comprises the second part of the

− CCM, whose first part is the concentration of HCO3 inside the cell by active transport across the cytoplasmic membrane and into the cytosol (9). Current models hold that

− HCO3 then crosses the protein shell and enters the carboxysome by diffusion. In the

− lumen of the carboxysome, CA dehydrates HCO3 to CO2 to increase the level of CO2 in the local vicinity of RuBisCO, thereby countering the low catalytic efficiency and poor substrate selectivity of RuBisCO for CO2 (Fig. 1.27). The carboxysome shell is believed

− to play a role in allowing or limiting the transport of substrates and products [i.e., HCO3 ,

CO2, 3-phosphoglycerate (3-PGA), and ribulose-1,5-bisphosphate (RuBP)] (67, 118, 180,

196) and possibly O2 (144), which competes with CO2 in its reaction with RuBisCO.

FIG. 1.27. CO2 fixation by carboxysomes (adapted from ref. (244)). CA, carbonic anhydrase; RuBP, ribulose-1,5-bisphosphate, RuBisCO, ribulose-1,5-bisphosphate carboxylase/oxygenase; 3-PGA, 3-phosphoglycerate.

Pdu and Eut MCPs have a different function than carboxysomes (53, 244) (Fig 1.28).

They work to mitigate aldehyde toxicity and diffusive loss. A common feature of the

1,2-PD and ethanolamine degradative pathways is that both proceed through aldehyde intermediates (propionaldehyde and acetaldehyde, respectively); hence, it was proposed that MCPs might be formed to sequester aldehydes which were known cytotoxins (219).

Studies showed that mutants unable to form the Pdu MCPs accumulated large amounts of propionaldehyde and underwent a temporary period of growth arrest due to propionaldehyde toxicity (95, 193). In addition, propionaldehyde was found to be a mutagen and mutation frequencies were increased in MCP mutants during growth on

1,2-PD (193). The aldehyde toxicity model is further supported by the findings that polA

(DNA repair polymerase) and gsh (glutathione biosynthesis) mutants were unable to grow on ethanolamine (185, 186). However, other studies have indicated that the main role of the Eut MCP might be to prevent the loss of the volatile metabolic intermediate, acetaldehyde (167). In the case of propionaldehyde (boiling point = 48 °C), loss due to volatility appears less important than for acetaldehyde (boiling point = 20 °C) (193).

A B

FIG. 1.28. Proposed pathways for 1,2-PD (A) and ethanolamine (B) degradation by Pdu and Eut MCPs, respectively (244). PduCDE, B12-dependent diol dehydratase; PduO, adenosyltransferase; PduGH, diol dehydratase reactivase; PduP, propionaldehyde dehydrogenase; PduQ, propanol dehydrogenase; PduL, phosphotransacylase; EutBC, B12-dependent ethanolamine ammonia lyase; EutE, aldehyde dehydrogenase; EutD, phosphotransacetylase; EutG, alcohol dehydrogenase.

1.5. Research overview

Salmonella is able to utilize 1,2-PD as a sole carbon and energy source in an

AdoCbl-dependent fashion involving a bacterial MCP (Pdu MCP). Pdu MCP is composed entirely of protein subunits and its function is to confine the toxic intermediate propionaldehyde during 1,2-PD degradation. Several sequentially-acting metabolic enzymes, including the diol dehydratase PduCDE and the propionaldehyde dehydrogenase PduP that respectively use AdoCbl and NAD+ as cofactors, are encased within the solid protein shell of the MCP. Recent crystallography studies suggested that some shell proteins such as PduA, PduT and PduU, have pores that may mediate the transport of enzyme substrates/cofactors across the MCP shell. However, localized regeneration of cofactors within the MCP might be advantageous for the activity of the metabolic enzymes and the efficiency of 1,2-PD degradation. Thus, this study tests the hypothesis that AdoCbl and NAD+ are recycled entirely within the Pdu MCP.

The first part of this study investigates the bifunctional cobalamin reductase PduS. This protein was over-expressed and characterized including its cofactor requirements and kinetic properties. The effect of the pduS deletion on 1,2-PD degradation was examed through growth studies. Further SDS-PAGE and Western blot of purified Pdu MCP, and

MS-MS analysis were carried out to test whether the PduS protein is a component of the

Pdu MCP. Two-hybrid experments were conducted to detect the interaction between PduS and the PduO adenosyltransferase that is located in the Pdu MCP and catalyzes the terminal step of AdoCbl synthesis.

The second part of this study investigates the 1-pronanol dehydrogenase PduQ.

Experiments similar to those with PduS were used to characterize PduQ biochemically.

The His-tag protein affinity pull-down assays were performed to test the interaction of

PduQ and the PduP propionaldehyde dehydrogenase which is located in the Pdu MCP and converts NAD+ to NADH.

In conjunction with prior results, the results reported here demonstrate the recycling systems of cobalamin and NAD+ within the Pdu MCP.

1.6. Thesis organization

This dissertation consists of four chapters. Chapter 1 is an introductory review on

+ vitamin B12, NAD , 1,2-PD degradation, and the bacterial MCP. Chapter 2 and 3 are two papers related to the topic of the thesis about cofactor regeneration within the Pdu MCP.

The first paper has been published in the Journal of Bacteriology and the second one is to be submitted to a journal. Chapter 4 is a general concluding chapter, where the results and conclusions of the two papers are summarized. Both papers were written by me with the editorial guidance and assistance from my major professor, Dr. Thomas A. Bobik.

Chapter 2. Characterization of the PduS cobalamin reductase of

Salmonella and its role in the Pdu microcompartment

This chapter, in part or in full, is a reprint of the material as it appears in the Journal of

Bacteriology (volume 192, pp. 5071-5080, 2010 by Shouqiang Cheng and Thomas A.

Bobik).

2.1. Abstract

Salmonella enterica degrades 1,2-propanediol (1,2-PD) in a coenzyme B12

(adenosylcobalamin, AdoCbl) -dependent fashion. Salmonella obtains AdoCbl by assimilation of complex precursors such as vitamin B12 and hydroxocobalamin.

Assimilation of these compounds requires reduction of their central cobalt atom from

+3 +2 +1 Co to Co to Co followed by adenosylation to AdoCbl. In this work, the His6-tagged

PduS cobalamin reductase from S. enterica was produced at high levels in Escherichia coli, purified and characterized. The anaerobically purified enzyme reduced cob(III)alamin to cob(II)alamin at a rate of 42.3 ± 3.2 μmol min-1 mg-1, and it reduced cob(II)alamin to cob(I)alamin at a rate of 54.5 ± 4.2 nmol min-1 mg-1 protein. The apparent Km values of PduS-His6 were 10.1 ± 0.7 μM for NADH and 67.5 ± 8.2 μM for hydroxocobalamin in cob(III)alamin reduction. The apparent Km values for cob(II)alamin reduction were 27.5 ± 2.4 μM for NADH and 72.4 ± 9.5 μM for cob(II)alamin. HPLC and

MS indicated that each monomer of PduS contained one molecule of non-covalently bound FMN. Genetic studies showed that a pduS deletion decreased the growth rate of

Salmonella on 1,2-PD supporting a role in cobalamin reduction in vivo. Further studies demonstrated that the PduS protein is a component of the microcompartments used for

1,2-PD degradation (Pdu MCPs) and that it interacts with the PduO adenosyltransferase which catalyzes the terminal step of AdoCbl synthesis. These studies further characterize

PduS, an unusual MCP-associated cobalamin reductase, and in conjunction with prior results, indicate that the Pdu MCP encapsulates a complete cobalamin assimilation system.

2.2. Introduction

Coenzyme B12 (Adenosylcobalamin, AdoCbl) is an indispensable cofactor for a variety of enzymes that are widely distributed among microorganisms and higher animals (14,

203). Organisms obtain AdoCbl by de novo synthesis or by assimilation of complex precursors such as vitamin B12 (cyanocobalamin, CN-Cbl) and hydroxocobalamin

(OH-Cbl) which can be enzymatically converted to AdoCbl. De novo synthesis occurs only in prokaryotes but the assimilation of complex precursors is more widespread taking place in many microorganisms and in higher animals (202). A model for the assimilation of CN-Cbl and OH-Cbl to AdoCbl, based on work done in a number of laboratories, is shown in Figure 2.1. CN-Cbl is first reductively decyanated to cob(II)alamin (91, 119,

238). Next, cob(II)alamin is reduced to cob(I)alamin and ATP:cob(I)alamin adenosyltransferase (ATR) transfers a 5'-deoxyadenosyl-group from ATP to cob(I)alamin to form AdoCbl (39, 42, 111, 113, 131, 215, 221, 248). Studies indicate that prior to reduction cob(II)alamin binds the ATR and undergoes a transition to the 4-coordinate base-off conformer (148, 165, 214, 216, 218). Transition to the 4-coordinate state raises the midpoint potential of the cob(II)alamin/cob(I)alamin couple by about 250 mV, facilitating reduction (217). OH-Cbl assimilation occurs by a similar pathway except that the first step is reduction of OH-Cbl to cob(II)alamin by cobalamin reductase or by the reducing environment of the cell (82, 239).

apoenzyme (154, 155). Next, the process described in Figure 2.1 converts OH-Cbl to

AdoCbl which spontaneously associates with apoenzyme to form active holoenzyme (154,

155, 228). In the organisms that have been studied, cobalamin recycling is essential, and genetic defects in this process block AdoCbl-dependent metabolism (17, 66, 113).

reductive CN-Cbl decyanation ATP PPPi

? cob(II)alamin cob(I)alamin AdoCb l OH-Cbl cob(III)alamin cob(II)alamin adenosylation reduction reduction

breakdown (AdoCbl is unstable in vivo)

FIG. 2.1. Cobalamin assimilation and recycling pathway. Many organisms are able to take up CN-Cbl and OH-Cbl and convert them to the active coenzyme form, AdoCbl. This process involves reduction of the central cobalt atom of the corrin ring followed by addition of a 5’-deoxyadenosyl (Ado) group via a carbon-cobalt bond. The Ado group is unstable in vivo, and AdoCbl breaks down to form OH-Cbl. Consequently, cobalamin recycling is required for AdoCbl-dependent processes and recycling uses the same pathway that functions in the assimilation of cobalamin from the environment.

Salmonella enterica degrades 1,2-propanediol (1,2-PD) via an AdoCbl-dependent pathway (110). 1,2-PD is a major product of the anaerobic degradation of common plant sugars rhamnose and fucose and is thought to be an important carbon and energy source in natural environments (138, 163). Twenty-four genes for 1,2-PD utilization (pdu) are found in a contiguous cluster (pocR, pduF and pduABB'CDEGHJKLMNOPQSTUVWX)

(31, 110). This locus encodes enzymes for the degradation of 1,2-PD and cobalamin recycling, as well as proteins for the formation of a bacterial microcompartment (MCP)

(31). Bacterial MCPs are simple proteinaceous organelles used by diverse bacteria to optimize metabolic pathways that have toxic or volatile intermediates (29, 48, 53, 246).

They are polyhedral in shape, 100-150 nm in cross-section (about the size of a large virus), and consist of a protein shell that encapsulates sequentially-acting metabolic

enzymes. Sequence analyses indicate that MCPs are produced by 20-25% of all bacteria and function in seven or more different metabolic processes (53). The function of the Pdu

MCP is to confine the propionaldehyde formed in the first step of 1,2-PD degradation in order to mitigate its toxicity and prevent DNA damage (31, 94, 95, 193). Prior studies indicate that 1,2-PD traverses the protein shell and enters the lumen of the Pdu MCP where it is converted to propionaldehyde and then to propionyl-CoA by

AdoCbl-dependent diol dehydratase (PduCDE) and propionaldehyde dehydrogenase

(PduP) (32, 129). Propionyl-CoA then exits the MCP into the cytoplasm where it is converted to 1-propanol, propionate or enters central metabolism via the methylcitrate pathway (105, 164). The shell of the Pdu MCP is thought to limit the diffusion of propionaldehyde in order to protect cytoplasmic components from toxicity. The Pdu MCP was purified and 14 major polypeptide components were identified

(PduABB'CDEGHJKOPTU) all of which are encoded by the pdu locus (94).

PduABB'JKTU are confirmed or putative shell proteins (94, 95, 193). PduCDE and PduP catalyze the first 2 steps of 1,2-PD degradation as described above (31, 32, 94, 129). The

PduO and PduGH enzymes are used for cobalamin recycling. PduO is an adenosyltransferase (113) and PduGH is a homolog of the Klebsiella diol dehydratase

(DDH) reactivase which mediates the removal of OH-Cbl from inactive [OH-Cbl-DDH] complex (154, 155). However, a reductase which is also required for cobalamin recycling was not previously identified as a component of the Pdu MCP (94). This raises the question of how cobalamin is recycled for the AdoCbl-dependent DDH that resides within the Pdu MCP.

Prior studies indicated that the PduS enzyme (which is encoded by the pdu locus) is a cobalamin reductase (194). Very recently PduS was purified from S. enterica and shown

to be a flavoprotein that can mediate the reduction of 4-coordinate cob(II)alamin bound to

ATR but was not further characterized (147). In this study, PduS from S. enterica is purified and more extensively characterized including its cofactor requirements and kinetic properties. In addition, we show that PduS is a component of the Pdu MCP. This finding in conjunction with prior work indicates that in addition to 1,2-PD degradative enzymes the Pdu MCP encapsulates a complete cobalamin recycling system.

2.3. Materials and methods

Bacterial strains and growth conditions. The bacterial strains used in this study are listed in Table 2.1. The rich media used were Luria-Bertani/Lennox (LB) medium (Difco,

Detroit, MI) (151) and Terrific Broth (TB) (MP Biomedicals, Solon, OH) (225). The minimal medium used was no-carbon-E (NCE) medium (25, 233).

TABLE 2.1. Bacterial strains used in this study. Species and strain Genotype Source E. coli BE118 BL21(DE3) RIL/pTA925-pduO Lab collection C41(DE3) F– ompT hsdSB (rB- mB-) gal dcm (DE3) Lucigen BE1355 C41(DE3)/pET-41a This study BE1356 C41(DE3)/pET-41a-pduS This study BE1374 C41(DE3)/pET-41a-pduS-His6 This study BacterioMatch® II Δ(mcrA)183 (mcrCB-hsdSMR-mrr)173 endA1 Two-Hybrid System hisB supE44 thi-1 recA1 gyrA96 relA1 lac [F´ Stratagene Reporter Strain lacIq HIS3 aadA Kanr] S. enterica serovar Typhimurium LT2 BE1352 ΔpduS::frt This study BE287 LT2/pLAC22 Lab collection BE1353 ΔpduS::frt /pLAC22 This study BE1354 ΔpduS::frt /pLAC22-pduS This study

Chemicals and reagents. Antibiotics, OH-Cbl, CN-Cbl, AdoCbl, iodoacetate, DNase I,

FMN, FAD, and corynebacterial sarcosine oxidase were from Sigma Chemical Company

(St. Louis, MO). IPTG was from Diagnostic Chemicals Limited (Charlotteville PEI,

Canada). PfuUltra high-fidelity DNA polymerase was from Stratagene (La Jolla, CA).

Taq DNA polymerase, restriction enzymes, and T4 DNA ligase were from New England

Biolabs (Beverly, MA). Other chemicals were from Fisher Scientific (Pittsburgh, PA).

Construction of plasmids and a pduS-deletion mutant. The pduS gene was amplified by PCR (using genomic DNA of S. enterica as the template) then cloned into pET-41a vector (Novagen, Cambridge, MA) using NdeI and HindIII restriction sites incorporated into the PCR primers as previously described (192). A gene for the production of

PduS-His6 was similarly cloned with the sequence encoding the His6-tag incorporated into the reverse PCR primer. The resulting plasmids pET-41a-pduS, pET-41a-pduS-His6, as well as the pET-41a without insert were introduced into Escherichia coli C41(DE3)

(Lucigen, Middleton, WI). The native pduS gene was also cloned into pLAC22, BglII to

HindIII, for complementation experiments. The native pduS and pduO genes were cloned into both pBT and pTRG (Stratagene) from BamHI to XhoI for two-hybrid analyses. All of the above inserts were verified by DNA sequencing.

The pduS deletion mutant was constructed by a PCR-based method (65) using primers

(pduS-DKF: CCAATGCCGAAGCCATTCGGGAACTGCTGGAGGAACTGCTATAATT

GTAGGCTGGAGCTGCTTCG and pduS-DKR: CTAAAATTCCTATAGCCTGAGACA

TGGTTAACCTCTTACAGATATGAATATCCTCCTTAGTTC). These primers were designed to remove nearly the entire pduS coding sequence, but leave predicted translation signals of the adjacent genes (pduQ and pduT) intact. The presence of the pduS deletion was verified by PCR as described (65).

Growth of expression strains and purification of PduS-His6. 200 ml TB containing

25 mg/ml kanamycin was inoculated with 2 ml of BE1374 grown in similar medium and the cells were cultivated in a 1 liter baffled Erlenmeyer flask at 37 °C with shaking at 275 rpm. When the culture reached an OD600 of about 0.5, riboflavin (10 μM), ferric

ammonium citrate (50 μg/ml), L-cysteine (1 mM), and IPTG (0.1 mM) were added. The culture was incubated for additional 18 h at 30 °C and 200 rpm shaking. Cells were then harvested by centrifugation at 4 °C and 6,000 x g for 10 min and used immediately for protein purification. Cells were resuspended in buffer A (50 mM potassium phosphate, pH

7.5, 300 mM NaCl, 5 mM β-mercaptoethanol, 0.4 mM AEBSF [4-(2-aminoethyl) benzenesulfonyl fluoride-HCl]) containing 20 mM imidazole and a few crystals of bovine pancreas DNase I. Cells were broken using a French pressure cell (SLM Aminco,

Rochester, NY) at 20,000 psi. Lysates were centrifuged for 30 min at 39,000 x g then filtered with a 0.45-μm-pore cellulose-acetate filter. The filtrate (soluble fraction) was applied to a pre-equilibrated Ni-NTA column (Qiagen, Valencia, CA). The column was washed with 20 bed volumes of buffer A containing 100 mM imidazole and proteins were eluted with buffer A containing 300 mM imidazole. Control strains BE1355, BE1356 were grown in parallel with expression strain BE1374 and similar procedures were used for preparing whole-cell extracts and soluble fractions. All protein purification steps were carried out at 4 °C either under strictly anaerobic conditions with deaerated buffers inside a glove box (Coy Laboratory, Grass Lake, MI) with a nitrogen:hydrogen:CO2 (90:5:5) atmosphere, or under aerobic conditions.

SDS-PAGE and Western blots. Protein concentration was determined using Bio-Rad

(Hercules, CA) protein assay reagent with bovine serum albumin (BSA) as a standard.

SDS-PAGE was performed using Bio-Rad 12% or 10-20% gradient Tris-HCl ready gels.

Protein bands were visualized by staining with Bio-Safe Coomassie Stain (Bio-Rad). For

Western blots, the proteins on SDS-PAGE gels were transferred to polyvinylidene fluoride (PVDF) membranes and probed using SuperSignal West Pico Chemiluminescent

Substrate (Pierce, Rockford, IL) according to the manufacturer’s instructions with

primary mouse anti-PduS sera at the final concentration of 0.5 μg/ml and secondary goat anti-mouse IgG-HRP (Santa Cruz Biotechnology, Santa Cruz, CA) at 40 ng/ml.

Gel filtration. FPLC was carried out using an ÄKTA system (GE Healthcare,

Piscataway, NJ). Aerobically and anaerobically purified PduS-His6 (0.5 ml at a concentration of 20 µM) was applied to a Superdex 200 HR 10/30 column equilibrated with 50 mM sodium phosphate, pH 7.0, 150 mM NaCl under aerobic conditions. The column was eluted with that same buffer at a flow rate of 0.25 ml/min and the protein elution profile was determined by monitoring absorbance at 280 nm. Gel filtration standards from Bio-Rad were used to construct a calibration curve of log MW versus retention time.

Identification of the flavin cofactor bound to PduS-His6. Purified PduS-His6 (in

Ni-NTA elution buffer) was boiled for 10 min to release the flavin cofactor. The suspension was rapidly cooled on ice and denatured protein was removed by centrifugation at 16,000 x g for 5 min using an Eppendorf 5415D centrifuge.

Flavin-containing supernatant was filtered with a 0.22 μm Millex-GV syringe filter

(Millipore, Bedford, MA) and analyzed using a Varian ProStar HPLC system (Palo Alto,

CA), consisting of a model 230 solvent delivery module, a model 430 autosampler, a model 325 UV-visible detector and a Microsorb-MV 100-5 C18 column. The column was developed with 19 ml linear gradient from 7% to 90% methanol in 5 mM ammonium acetate pH 6.0 at a flow rate of 1 ml/min. Flavins were detected by monitoring absorbance at 450 nm. Authentic FAD and FMN were used as standards and flavins separated by HPLC were collected and analyzed by MS.

Iron and sulfide content assays. Non-heme iron assays were carried out by using the method of Fish (79). Acid-labile sulfide assays were performed with a modified Beinert’s

method (22, 41) under anaerobic conditions except that 10 μl instead of 2 μl of FeCl3 (23 mM in 1.2 M HCl) solutions were used at the last step for color development.

Cob(III)alamin and cob(II)alamin reduction assays. Enzyme assays were performed under anaerobic conditions as described previously with some modifications (194). All reactions were initiated by addition of NADH or iodoacetate to assay mixtures except where noted in the text. Cob(III)alamin reductase assays contained 50 mM CHES/NaOH

(pH 9.5), 1.6 mM KH2PO4, 0.5 mM MgCl2, 0.5 mM NADH, 0.2 mM OH-Cbl and cob(III)alamin reductase as indicated in the text. The conversion of OH-Cbl to cob(II)alamin was monitored spectrophotometrically by measuring the absorbance

-1 -1 decrease at 525 nm and quantified using Δε525 = 4.9 mM cm (194).

Cob(II)alamin reductase activity was assayed using two methods. In method one, iodoacetate was used as a chemical trap for cob(I)alamin. Iodoacetate reacts rapidly and quantitatively with cob(I)alamin to form carboxymethylcobalamin (CM-Cbl) with a

–1 –1 concomitant absorbance increase at 525 nm (Δε525 = 5.3 mM cm ) (26). Assay mixtures contained 50 mM CHES/NaOH (pH 9.5), 1.6 mM KH2PO4, 0.5 mM MgCl2, 0.5 mM

NADH, 0.2 mM cob(II)alamin, 0.5 mM iodoacetate, and cob(II)alamin reductase. The second method for measuring cob(II)alamin reduction was a coupled assay with the PduO adenosyltransferase as described in a prior report (194). Assay mixtures contained the same components as assay one except that 0.5 mM iodoacetate was replaced by 0.5 mM

ATP and 1.6 μM PduO adenosyltransferase purified from BE118 as described (111).

PduO converts ATP and cob(I)alamin to AdoCbl which occurs with an increase in

–1 –1 absorbance at 525 nm (Δε525 = 4.8 mM cm ).

Growth studies. Growth studies were performed using a Synergy HT Microplate reader (BioTek, Winooski, VT) as previously described (139). For growth of strains

carrying plasmids, media were supplemented with 100 μg/ml ampicillin. For complementation studies, IPTG was used at 10 μM to induce expression of genes cloned into pLAC22.

Pdu MCP purification. Cells were grown at 37 °C with shaking at 275 rpm to an

OD600 of 1.0-1.2 in 400 ml NCE medium supplemented with 1 mM MgSO4, 0.5% succinate, and 0.6% 1,2-PD inoculated with 2 ml of an overnight LB culture. Cells were harvested by centrifugation at 6,000 x g for 10 min at 4 °C. Then, MCPs were purified as previously described and stored at 4 °C until used (94).

MALDI-TOF MS-MS. Bands of interest were excised from SDS-PAGE gels and digested “in-gel” with trypsin. The resulting peptides were extracted and analyzed by

MALDI-TOF MS-MS using a QSTAR XL quadrupole TOF mass spectrometer (AB/MDS

Sciex, Toronto, Canada) equipped with an oMALDI ion source. CHCA was used as matrix and the mass spectrometer was operated in the positive ion mode. Mass spectra for

MS analysis were acquired over m/z 600 to 2200. After every regular MS acquisition, two

MS-MS acquisitions were performed against most intensive ions. The molecular ions were selected by information dependent acquiring program in the quadrupole analyzer and fragmented in the collision cell. All spectra were processed by MASCOT

(MatrixScience, London, UK) database search.

Two-hybrid analyses. The BacterioMatch II Two-Hybrid System (Stratagene) was used to detect the potential interactions between the PduS and PduO proteins in vivo following manufacturer’s instructions.

2.4. Results

Sequence analysis. The PduS protein of S. enterica is composed of 451 amino acid

residues. Over 100 PduS homologues present in GenBank are associated with 1,2-PD degradation based on gene proximity. PduS also has 43% sequence identity with the RnfC subunit of NADH:ubiquinone oxidoreductase (RnfABCDEG) encoded by the bacterial rnf operon (199), and the N-terminus of PduS (amino acids 1-165) has 25% amino acid identity with the Nqo1 (NDUFV1 in mammals) subunit of the bacterial and mitochondrial

NADH-quinone oxidoreductase complex I (168, 197, 204). PduS has several conserved binding sites for substrates and cofactors, including an NADH-binding motif

28 33 ( GXGX2G where X denotes any amino acid) (37), an FMN-binding motif

120 125 264 274 ( YX2G[D/E]E ) (125, 197) and two canonical [4Fe-4S] motifs ( CX2CX2CX3C

309 320 and CX2CX2CX4C ) (86, 132) (Fig. 2.2A, B). PduS also has several conserved

25 39 domains. An N-terminal glycine-rich loop ( GX2GXGGAG[F/L]P[A/T]X2K ), proposed to bind the adenosine diphosphate (ADP) moiety of NADH, is conserved among PduS,

RnfC and Nqo1 (Fig. 2.2C) (197). A non-classical Rossmann fold following the Gly-rich loop is proposed to bind FMN (197). A soluble ligand-binding β-grasp fold (SLBB) domain which belongs to a newly defined superfamily is proposed to bind cobalamin in

PduS (46). The two [4Fe-4S] motifs of PduS are found in a Fer4 domain (PF00037) (46).

PduS has a C-terminal sandwich barrel hybrid motif (SBHM). Some SBHM domains carry covalently associated ligands including biotin and lipoate, but PduS has no known covalently bound ligands (46, 108).

B C

FIG. 2.2. Sequence analyses of PduS. (A) Structure-based sequence alignment by clustal X2 and PSIPRED. α-helix and β-strand are represented by coil and arrow, respectively. CfPduS, PduS from Citrobacter freundii (GI: 171854200); Ec, Escherichia coli (GI: 157159374); Li, Listeria innocua (GI: 16800175); Ri, Roseburia inulinivorans (GI: 225376079); Se, Salmonella enterica (GI: 16765383); Ye, Yersinia enterocolitica (GI: 123442959); TaRnfC, RnfC from Thermanaerovibrio acidaminovorans (GI: 269791702). (B) The domain architecture of PduS protein (46). (C) Sequence logo of the amino acid residues (present at positions 25-39 in PduS of S. enterica) in the glycine-rich region used for nucleotide-binding. Polar residues are in green, basic in blue, acidic in red, and hydrophobic in black.

Expression and purification of PduS-His6 protein. E. coli strain BE1374 was constructed to produce high levels of C-terminal-His6-tagged PduS protein via a T7 expression system. This strain produced relatively large amounts of protein near the expected molecular mass of PduS-His6 (49.2 kDa) and the major portion of this protein was found in the soluble fraction of cell extracts (Fig. 2.3). In contrast, cell extracts from a control strain containing the expression plasmid without insert (BE1355) produced much less protein near 49.2 kDa. PduS-His6 was purified under both aerobic and

anaerobic conditions by Ni-NTA affinity chromatography. Based on SDS-PAGE anaerobically purified PduS-His6 protein was about 90% homogenous (Fig. 3, lane 4), and a similar level of purity was obtained for aerobically purified enzyme.

FIG. 2.3. SDS-PAGE analysis of the anaerobic purification of PduS-His6 from BE1374. Lane 1, protein standards; lane 2, 12 μg whole-cell extract; lane 3, 12 μg soluble fraction; lane 4, 3 μg purified

PduS-His6. The gel was 12% polyacrylamide and was stained with Coomassie.

Oligomeric state of PduS-His6 protein. Size exclusion chromatography of anaerobically purified PduS-His6 showed a major peak and a small shoulder which corresponded to a monomer with an apparent molecular mass of 49 kDa and a dimer of

98 kDa, respectively (Fig. 2.4A). Chromatography of aerobically purified PduS-His6 displayed a major peak and a larger shoulder indicating that the dimer comprised a larger proportion in aerobically purified PduS-His6 (Fig. 2.4B). The small amount of dimerization observed following aerobic purification may have resulted from the formation of intermolecular disulfide bonds perhaps involving cysteine residues freed from the iron-sulfur clusters which are oxygen-labile.

FIG. 2.4. Oligomeric state and molecular mass determination of PduS-His6 by size exclusion chromatography. PduS-His6 is predominant in monomeric fractions. (A) Profile of anaerobically purified PduS-His6. (B) Profile of aerobically purified PduS-His6.

Characterization of the flavin cofactor of PduS-His6. Aerobically Purified

PduS-His6 was yellow in color, and its UV-visible spectrum exhibited peaks at 375 and

450 nm and a shoulder at 480 nm indicative of a flavin cofactor (Fig. 2.5).

To determine the binding stoichiometry and the type of flavin (FAD or FMN), purified

PduS-His6 was denatured with heat and the released cofactors were analyzed by HPLC and MS. The flavin extracted from PduS-His6 eluted at 9.96 min by reverse phase HPLC, which was similar to the retention time for authentic FMN (9.86 min) (Fig. 2.6). In contrast, FAD eluted at 8.90 min (Fig. 2.6). In addition, further studies showed that the

flavin released from PduS-His6 co-migrated with authentic FMN by reverse phase HPLC following co-injection (Fig. 2.6).

0.6

0.5

0.4

0.3

Absorbance 0.2

0.1

0 300 350 400 450 500 550 600

Wavelength (nm)

FIG. 2.5. UV-visible spectrum of purified PduS-His6 (solid line) in comparison to that of authentic FMN (dashed line). The absorbance peaks at 375 and 450 nm with a shoulder at 480 nm are diagnostic of flavin.

FIG. 2.6. Reverse phase HPLC analyses of flavin released from PduS-His6. Flavins were detected by absorbance at 450 nm. a, authentic FAD; b, boiled authentic FAD; c, authentic FMN; d, flavin released from PduS-His6; e, authentic FMN plus flavin from PduS-His6. The absorbance profiles shown were determined by separate HPLC analyses then overlaid.

Lastly, HPLC and MS were used to confirm and quantify the flavin cofactor of

PduS-His6. The mass spectrum of the flavin released from purified PduS-His6 showed a major peak with an m/z of 455.1 corresponding to that of FMN (Fig. 2.7). Quantitation by

HPLC determined a stoichiometry of 0.93:1, (FMN:PduS-His6) indicating that PduS-His6 binds 1 molecule of FMN per monomer.

A B

FIG. 2.7. MS analyses of the flavin extracted from purified PduS-His6. (A) Authentic FMN. (B) Flavin extracted from PduS-His6 and purified by HPLC. The MS spectrum of flavin from PduS-His6 displayed a peak with an m/z of 455.1 corresponding to that of authentic FMN.

Flavin cofactors may be covalently or non-covalently bound to flavoproteins. The observation that flavin was released from PduS-His6 by heat treatment indicated non-covalent binding of FMN to PduS-His6. To further investigate, we used SDS-PAGE.

Covalently bound flavins remain protein-associated during SDS-PAGE and can be detected by fluorescence following UV illumination of gels (207). In contrast, non-covalently bound flavins are not detected. The FMN cofactor of PduS-His6 could not be detected by UV illumination of SDS-PAGE gels indicating non-covalent binding (Fig.

2.8). In contrast, the flavin of sarcosine oxidase from Corynebacterium, which is known to be covalently bound (242), co-migrated with the protein during SDS-PAGE and was

readily detected by UV illumination (Fig. 2.8). Thus, results indicate that FMN is non-covalently bound to PduS-His6.

FIG. 2.8. PduS binds FMN non-covalently. (A) SDS-PAGE of PduS-His6 (lane 2) and sarcosine oxidase from Corynebacterium sp. (lane 3). (B) Detection of flavin fluorescence in PduS-His6 (lane 4) and sarcosine oxidase from Corynebacterium sp. (lane 5) on an SDS-PAGE gel. The fluorescence of the corynebacterial sarcosine oxidase was due to the FMN covalently bound to the β subunit (SoxB).

Fluorescence of PduS-His6 following SDS-PAGE was undetectable indicating that FMN binding is non-covalent. To detect fluorescence, the SDS-PAGE gel was soaked with 7% acetic acid for 15 min followed by spraying with performic acid, and then illuminated with UV light (λ = 365 nm) (207).

Iron and sulfide contents. To test the presence of iron-sulfur clusters in PduS protein, the contents of iron and sulfide in PduS-His6 were measured. The results indicated that anaerobically purified PduS-His6 contains 7.6 ± 0.3 mol of iron and 7.1 ± 0.5 mol of acid-labile sulfide per mol of PduS-His6 monomer, which is consistent with the motif prediction result that PduS sequence contains two [4Fe-4S] binding sites. Whereas aerobically purified PduS-His6 contains 3.0 ± 0.3 mol of iron and 2.8 ± 0.3 mol of sulfide per mol of PduS monomer, suggesting that the iron-sulfur clusters of PduS are oxygen-labile.

In vitro cob(III)alamin and cob(II)alamin reductase activities of the PduS-His6 enzyme. PduS-His6 was purified using strict anaerobic conditions and tested for cob(III)alamin and cob(II)alamin reductase activities after each purification step (Table

2.2). The whole-cell extract from the PduS-His6 expression strain (BE1374) exhibited about 350-fold higher cob(III)alamin reductase activity (3.9 ± 0.3 μmol min-1 mg-1) than did extracts from the control strain carrying the expression plasmid without insert

-1 -1 (BE1355) (11.0 ± 3.1 nmol min mg ). Purification of the PduS-His6 protein by anaerobic Ni-NTA chromatography increased the cob(III)alamin reductase specific activity approximately 11-fold to 42.3 ± 3.2 μmol min-1 mg-1.

Assays showed that the PduS-His6 protein also catalyzed the reduction of cob(II)alamin to cob(I)alamin. Soluble cell extracts of expression strain BE1374 had 4.9 ± 0.5 nmol min-1 mg-1 cob(II)alamin reductase activity when iodoacetate was used to trap cob(I)alamin and 4.3 ± 0.4 nmol min-1 mg-1 in a coupled assay with the PduO adenosyltransferase (see assay procedures in the methods section). In contrast, control extracts from BE1355 lacked detectable cob(II)alamin reductase activity. Purification of the PduS-His6 protein by anaerobic Ni-NTA chromatography increased the cob(II)alamin reductase specific activity about 11-fold to 54.5 ± 4.2 nmol min-1 mg-1 by the iodoacetate assay and about 9-fold to 40.3 ± 4.1 nmol min-1 mg-1 by the PduO-linked assay.

PduS-His6 was also purified under aerobic conditions. Aerobically purified PduS-His6 retained about 40% activity for both cob(III)alamin and cob(II)alamin reduction. The reduced activity following aerobic purification was likely due to oxidative damage to catalytic sites such as the iron-sulfur centers predicted from sequence analysis (Fig. 2).

TABLE 2.2. Anaerobic purification of PduS-His6.

Total Cob(III)alamin reductase Cob(II)alamin reductase Purification step protein Specific activity Total activity Purification Specific activity Total activity Purification (mg) (μmol min-1 mg-1) (μmol min-1) fold (nmol min-1 mg-1) (nmol min-1) fold Whole-cell extract 220 3.9 ± 0.3 858 1 4.4 ± 0.4 968 1 Soluble fraction 166 4.2 ± 0.4 697 1.1 4.9 ± 0.5 813 1.1 Ni-NTA eluate 1.6 42.3 ± 3.2 67.7 10.8 54.5 ± 4.2 87.2 12.4 a0.5 mM iodoacetate was used to trap produced cob(I)alamin. 62

The C-terminal His6-tag had no obvious effect on PduS enzymatic activities based on the comparisons of the reductase activities in the whole-cell extracts and soluble fractions of BE1374 and BE1356, which respectively express PduS-His6 and wild-type PduS.

Reaction requirements. To determine the PduS reaction requirements, key assay components were individually omitted. For cob(III)alamin reduction, there was no activity in the absence of NADH or OH-Cbl. In the absence of PduS-His6, cob(III)alamin reduction occurred at a rate of 3.5 ± 0.5 nmol min-1 (more than 1100-fold slower than the

PduS-catalyzed reaction) due to chemical reduction by NADH. The PduS-His6 cob(III)alamin reductase lacked measurable activity with CN-Cbl. In the cob(II)alamin reduction assays, controls showed that no detectable CM-Cbl was formed in the absence of NADH, cob(II)alamin, PduS-His6, or iodoacetate. Similarly, in linked assays with the

PduO adenosyltransferase, no AdoCbl was measurable in the absence of NADH, cob(II)alamin, PduS-His6, ATP or PduO. Moreover, no cob(I)alamin production could be directly observed by monitoring absorbance at 388 nm (113) in the absence of trapping agents (iodoacetate or PduO and ATP).

Effects of pH, temperature, and divalent metal ions on cobalamin reductase activities of PduS-His6. To determine the effects of pH on the activity of purified

PduS-His6 cob(III)alamin and cob(II)alamin reductase activities were measured at pH values ranging from 7.0 to 10.5 with an interval of 0.25 pH units using anaerobically purified PduS-His6. Under the conditions used, the PduS-His6 cob(III)alamin reductase activity was maximal at pH 9.5, and this value was set as 100%. Sharp decreases in activity were observed at pH 9.0 and pH 10.0 (51.4 and 40.0% drop, correspondingly), and further decreases were displayed at lower and higher pH values. In contrast, pH had a relatively small effect on PduS-His6 cob(II)alamin reductase activity. The highest activity

was measured at pH 9.0-9.5 with 17% decrease at pH 8.0 and 10.0. These results suggest that an ionizable group contributes to cob(III)alamin reductase activity but not to cob(II)alamin reduction. The buffer used to test pH effects was that standard assay buffer,

CHES/NaOH.

To determine the effects of temperature on the activity of purified PduS-His6, cob(III)alamin and cob(II)alamin reductase activities were measured under standard assay conditions with the temperature set at 25, 30, 37, 42, or 50 °C. The relative cob(III)alamin reductase specific activities were as follows: 47.2% at 25 °C, 64.3% at 30 °C, 100% at 37

°C, 92.0% at 42 °C, 78.4% at 50 °C, while PduS-His6 cob(II)alamin reductase activity had a different profile: 24.1% at 25 °C, 38.9% at 30 °C, 73.9% at 37 °C, 100% at 42 °C,

80.6% at 50 °C. The different temperature profiles for the cob(III)alamin and cob(II)alamin reductase activities of PduS, together with the different pH profiles tentatively suggest their reduction may occur at different catalytic sites.

To determine the effect of divalent metal ions on PduS-His6 enzymatic activities, seven chloride salts including CaCl2, CdCl2, CoCl2, MgCl2, MnCl2, NiCl2, ZnCl2, were tested separately at 0.5 mM in the activity assays at 37 °C and pH 9.5. Mg2+ and Ca2+ ions significantly stimulated PduS-His6 cob(III)alamin reductase activity, by 1.35- and 1.5-fold respectively, compared to the activity measured with no added divalent ions. Addition of

Mn2+ ions resulted in a somewhat smaller enhancement of 1.25-fold on cob(III)alamin reductase activity. Ni2+ was slight inhibitory resulting in an 8% decrease in activity relative to the unsupplemented control. PduS-His6 cob(II)alamin reductase activity was stimulated around 1.1-fold by Ca2+, Mg2+, and Mn2+ ion, and was inhibited by Ni2+ which resulted in a 60% drop in activity. Cd2+, Co2+, and Zn2+ ions abolished both the cob(III)alamin and cob(II)alamin reductase activities of PduS-His6.

Preference for the electron donor of PduS-His6. To determine the cofactor specificity of PduS-His6 both cobalamin reductase activities were assayed in the presence of either

NADH or NADPH at 0.5 mM. Results showed that purified PduS-His6 enzyme had

16.9% relative activity for both activities with NADPH compared to NADH which was the preferred substrate.

Linearity of the reactions. The effects of PduS-His6 concentration on enzymatic activities were determined. Cob(III)alamin reduction was proportional to PduS-His6 concentration from 5 to 50 nM when 0.5 mM NADH and 0.2 mM OH-Cbl were used as substrates. Linear regression yielded an R2 value of 0.999. Cob(II)alamin reduction, measured in nanomoles of CM-Cbl generated per minute, was linear from 0.2 to 2 μM

2 PduS-His6 (R = 0.998) when the assay mixture contains 0.5 mM NADH, 0.2 mM cob(II)alamin, and 0.5 mM iodoacetate as the cob(I)alamin trapping agent.

Kinetic analysis of PduS-His6 activities. Steady-state kinetic studies for cob(III)alamin and cob(II)alamin reduction were performed using purified PduS-His6 with varied concentrations of one substrate and a fixed concentration of the other substrate. Kinetic parameters were obtained by non-linear curve fitting to the

Michaelis-Menten equation v = (Vmax [S])/(Km + [S]) using GraphPad Prism 5 Software

(GraphPad Software, San Diego, CA). For cob(III)alamin reduction, the apparent Km values for NADH and OH-Cbl were 10.1 ± 0.7 μM and 67.5 ± 8.2 μM, respectively (Fig.

–1 –1 9; Table 3). The enzyme Vmax was 43.1 ± 0.5 or 46.6 ± 1.6 μmol min mg when OH-Cbl or NADH were held at constant levels, respectively. The fixed concentrations of OH-Cbl and NADH used were 200 µM and 500 µM, respectively. 500 µm NADH is 50-fold higher than the apparent Km (98% saturating) and 200 µM OH-Cbl is 3-fold higher than the apparent Km (75% saturating) which was the highest level that could be used while

still retaining a linear response from the spectrophotometer.

FIG. 2.9. Kinetics of cob(III)alamin and cob(II)alamin reductions by purified PduS-His6. A,

Cob(III)alamin reduction assays performed with 25 nM anaerobically purified PduS-His6. B,

Cob(II)alamin reduction assays performed with 1 μM anaerobically purified PduS-His6 protein in the presence of 0.5 mM iodoacetate. When the concentration of NADH was varied, OH-Cbl or cob(II)alamin was used at 0.2 mM. When the concentration of OH-Cbl or cob(II)alamin was varied, NADH was held at 0.5 mM. The inserts show Lineweaver-Burk (double-reciprocal) plots.

TABLE 2.3. Kinetic parameters for cob(III)alamin and cob(II)alamin reduction by purified PduS-His6.

Variable K a V a k k /K Reaction m max cat cat m substrate (μM) (nmol min-1 mg-1) (s-1) (μM-1 s-1) Cob(III)alamin NADH 10.1 ± 0.7 (43.1 ± 0.5) x 103 35.3 ± 0.4 3.5 reductionb OH-Cbl 67.5 ± 8.2 (46.6 ± 1.6) x 103 38.2 ± 1.3 0.57 Cob(II)alamin NADH 27.5 ± 2.4 56.8 ± 1.1 (46.6 ± 0.9) x 10-3 1.7 x 10-3 reductionc cob(II)alamin 72.4 ± 9.5 64.7 ± 3.6 (53.0 ± 3.0) x 10-3 0.7 x 10-3 a The values of Km and Vmax were from non-linear regression using GraphPad Prism 5. b Cob(III)alamin reduction assays were performed with 25 nM anaerobically purified PduS-His6. c Cob(II)alamin reduction assays were performed with 1 μM anaerobically purified PduS-His6 protein in the presence of 0.5 mM iodoacetate.

The kinetics of cob(II)alamin reduction were determined using iodoacetate to trap cob(I)alamin. Non-linear regression indicated an apparent Km of 27.5 ± 2.4 μM and 72.4 ±

9.5 μM for NADH and cob(II)alamin, respectively (Fig. 2.9; Table 2.3). The enzyme Vmax was 56.8 ± 1.1 nmol min–1 mg–1 when the cob(II)alamin concentration was held constant and 64.7 ± 3.6 nmol min–1 mg–1 when NADH was held at a saturating level. For the above kinetic studies, NADH and cob(II)alamin were used at fixed concentrations of 500 µM and 200 µM, respectively.

Phenotype and complementation of a pduS deletion mutation. To investigate the function of PduS in vivo, we measured the growth of a pduS deletion mutant (BE1352) on

1,2-PD minimal medium.

BE1352 grew slowly (64% wild-type rate) on 1,2-PD minimal medium supplemented with saturating levels of CN-Cbl (100 nM) (Fig. 2.10A). At limiting concentrations of

CN-Cbl (20 nM), the pduS deletion mutant grew more slowly (61%) and to a lower cell density than wild-type S. enterica (Fig. 2.10B). Complementation analysis was also conducted. Production of PduS from pLAC22 fully corrected the slow growth phenotype of the ΔpduS mutant at saturating CN-Cbl (Fig. 2.10C) and limiting CN-Cbl (not shown) confirming that slow growth was due to the pduS deletion, and not due to polarity or an unknown mutation. The growth curves shown in Fig. 2.10 A-C were all repeated at least three times with triplicate cultures. The slow growth phenotype of the pduS deletion mutant supports a role for PduS in cobalamin reduction in vivo as further described in the discussion section.

FIG. 2.10. Phenotype and complementation of a pduS deletion mutation. Cells were grown in NCE minimal medium with 1,2-PD as a sole carbon and energy source. (A) The pduS deletion moderately impaired growth on 1,2-PD supplemented with 100 nM CN-Cbl. (B) The pduS deletion impaired growth and decreased cell density on 1,2-PD with limiting CN-Cbl (20 nM). (C) The pduS deletion was complemented by ectopic expression of pduS at 100 nM CN-Cbl. 10 μM IPTG was used to induce production of PduS from pLAC22.

Interaction between PduS and PduO. Prior studies suggested the PduS and PduO interact in vitro which makes sense because they catalyze sequential reactions in cobalamin recycling (194). To examine the potential interactions between PduS and PduO in vivo, two-hybrid analyses were performed. For the system used, detection of protein-protein interactions is based on transcriptional activation of the yeast HIS3 reporter gene which confers 3-amino-1,2,4-triazole (3-AT) resistance. In application, a reporter strain is co-transduced with a target and bait plasmid pair, and the number of

3-AT resistant colonies estimates the strength of the protein-protein interaction.

To test for a PduS-PduO interaction, the number of colonies on non-selective and selective (3-AT) media was counted at different dilutions after co-transformation (Table

2.4). When PduO and PduS were produced from the target (pTRG) and bait (pBT) plasmids or vice versa a substantial number of 3-AT resistant co-transformants were obtained. In contrast, very few 3-AT resistant transformants were observed in controls that lacked either pduO or pduS (Table 2.4). These results indicated that PduS interacts with PduO in vivo.

TABLE 2.4. Two-hybrid analysis of PduS-PduO interactions. Number of coloniesa Non-selective medium Selective medium (3-AT) Plasmid pair 1/100 1/1000 1/10 1/1000 pBT-LGF2/pTRG-Gal11b TNTCc ~2000 TNTC ~2000 pBT-pduS/pTRG-pduO >2000 255 >2000 152 pBT-pduO/pTRG-pduS >2000 296 >2000 129 pBT-pduS/pTRG >2000 410 0 0 pBT/pTRG-pduS >2000 564 1 0 pBT-pduO/pTRG >2000 170 2 0 pBT/pTRG-pduO >2000 400 3 0 aNumber of colonies formed following co-transformation of the reporter strain with the bait and target plasmids on selective and nonselective media at the dilution indicated. bLGF2 and Gal11 are a positive control known to strongly interact. cTNTC: Too numerous to count.

A B

FIG. 2.11. PduS is a component of the Pdu MCP. (A) 10-20% SDS-PAGE gel stained with Coomassie brilliant blue. Lane 1, molecular mass markers; lane 2, 10 μg Pdu MCPs purified from wild-type Salmonella. (B) Western blot for PduS. Lanes 1 and 2, 10 μg whole-cell extract or purified MCPs from wild-type; lanes 3 and 4, 10 μg whole-cell extract or purified MCPs from BE1352 (ΔpduS).

PduS is a component of Pdu MCP. To determine the cellular location of PduS, MCPs purified from wild-type S. enterica and pduS deletion mutant BE1352, were analyzed by

SDS-PAGE, MALDI-TOF MS-MS, and Western blot. A band near the expected molecular mass of PduS (48.4 kDa) and close to the PduP band at 49.0 kDa, was observed on an SDS-PAGE gel of MCPs purified from the wild-type strain (Fig. 2.11A lane 2).

This band was further analyzed by MALDI-TOF MS-MS. Five sequences (KSHPLIQRR,

RAIDALTPLLPDGIRL, KVDQQLMWQQAARL, RQHIGASAVANVAVGERV,

RHLIGHELSPHLLVRA) identified by MS-MS of a trypsin digest matched the PduS protein indicating it is a component of purified Pdu MCPs. In addition, Western blot with

PduS antisera detected a band near 48 kDa in MCPs purified from wild type S. enterica

(Fig. 2.11B lane 2), while no PduS band was detected in MCPs purified from the pduS

deletion mutant (Fig. 2.11B lane 4). Furthermore, when similar amounts of protein were analyzed, Western blot easily detected PduS in purified MCPs but not in crude cell extracts (Fig. 2.11B lane 2 and 1). This indicated that purified MCPs were substantially enriched in PduS. Cumulatively, these results show that PduS protein is a component of

Pdu MCPs.

2.5. Discussion

In previous studies, an E. coli strain was constructed that produced high levels of the

PduS enzyme of S. enterica (194). Crude cell extracts from this strain had substantially increased cob(III)alamin and cob(II)alamin reductase activities suggesting that the PduS enzyme played a role in cobalamin assimilation and recycling (194). Very recently, the

PduS enzyme was purified and shown to be a flavoprotein that can supply electrons for the synthesis of AdoCbl by the human and Lactobacillus ATR enzymes; however, PduS was not further characterized (147). In this report, C-terminally His6-tagged PduS protein was purified and characterized more extensively. Results showed that PduS is a flavoprotein that contains 1 FMN per monomer. Anaerobically purified PduS-His6 protein exhibited cob(III)alamin and cob(II)alamin reductase activities of 42.3 μmol min-1 mg-1 and 54.5 nmol min-1 mg-1, respectively. Although, the cob(II)alamin reductase activity is

776-fold lower than the cob(III)alamin reductase activity, it is reasonable to infer that this activity is relevant in vivo. The midpoint potentials of cob(III)alamin/cob(II)alamin (+240 mV) and cob(II)alamin/cob(I)alamin (-610 mV) (13, 136) rationally account for the incongruity between the two activities. AdoCbl is needed only in very small quantities, generally from 1-10 μM in prokaryotes (202). At least one other enzyme with a well-documented role in AdoCbl synthesis (the CobA ATR) was reported to have a specific activity of 53 nmol min-1 mg-1 (221) which is very close to the cob(II)alamin

reductase activity of PduS reported here. Moreover, the disparity in cob(III)alamin and cob(II)alamin reductase activities of PduS is similar to that reported for the CobR corrin reductase from Brucella melitensis which is unrelated in amino acid sequence to PduS but catalyzes similar reactions (127). Thus, we infer that the lower, cob(II)alamin reductase activity of PduS is within a physiologically relevant range.

Kinetic studies of purified PduS-His6 were also performed. The apparent Km values of

PduS-His6 were 67.5 μM and 10.1 μM for OH-Cbl and NADH respectively in cob(III)alamin reduction, and 72.4 μM for cob(II)alamin and 27.5 μM for NADH in cob(II)alamin reduction. Typical intracellular cobalamin levels are about 1 to 10 µM in prokaryotic organisms (202) and the levels of nicotinamide coenzymes are 1-3 mM (158).

This suggests that in vivo NADH would be saturating and cobalamin would be limiting for PduS activity; however, results presented in this report indicate that PduS is a component of the Pdu MCP and hence may reside in the MCP lumen where the local cobalamin concentration is unknown but might be elevated relative to the cytoplasm of the cell. Thus, the Km values of PduS for cobalamins might reflect its localization within the lumen of the Pdu MCP.

Several other systems capable of reducing cobalamin in vitro have been previously described (81, 82, 127, 147, 194). The reduction of cob(III)alamin to cob(II)alamin is facile and certainly occurs to some extent in the cytoplasm of prokaryotic cells (which is a strongly reducing environment) without the requirement for a specific reductase (82). In addition, several flavoproteins as well as FMNH2 and FADH2 reduce cob(III)alamin to cob(II)alamin in vitro and may contribute to cob(III)alamin reduction in vivo (82, 147,

194). The reduction of cob(II)alamin to cob(I)alamin is more difficult due to extremely low redox potential of the cob(II)alamin/cob(I)alamin couple (-610 mV) (136). Prior

+ studies showed that FADH2, FMNH2, ferredoxin NADP reductase (Fpr), flavodoxin A

(FldA), flavin:NADH reductase (Fre) as well as flavoproteins with no known involvement in B12 metabolism can provide electrons for cob(II)alamin reduction in vitro in the presence of ATR (82, 147). Results indicated that these systems specifically reduce

4-coordinate ATR-bound cob(II)alamin whose midpoint potential is about 250 mV higher than that of 5-coordinate cob(II)alamin (147, 214). In this report, we showed that purified

PduS is able to reduce both cob(III)alamin and cob(II)alamin in the absence of additional proteins. The reduction of cob(II)alamin to cob(I)alamin was detected by trapping cob(I)alamin with iodoacetate and by using a coupled assay with the PduO ATR. In both assays, the rates of cob(II)alamin reduction were similar, 54.5 and 40.3 nmol min-1 mg-1, respectively. This showed that ATR was nonessential for cob(II)alamin reduction by PduS.

This raises the question of how PduS overcomes the redox-potential-barrier for cob(II)alamin reduction. One possibility is that PduS binds cob(II)alamin and converts it to the 4-coordinate form prior to reduction.

The fact that cobalamin can be reduced by FMNH2, FADH2 and various flavoproteins raises the question of the relative contributions of these systems in vivo. To examine the role of PduS in vivo, we tested the effects of a pduS deletion mutation on

AdoCbl-dependent growth on 1,2-PD with saturating or limiting CN-Cbl (Fig. 2.10A, B).

Under both conditions, the pduS deletion impaired growth, indicating that PduS was required to support the maximal rate of 1,2-PD degradation under the conditions used.

The likely reason that the pduS deletion exhibited only a partial growth defect is that S. enterica has multiple cobalamin reducing systems as described above. A similar partial growth defect on 1,2-PD was observed following genetic deletion of the Salmonella

PduO ATR which is partially redundant with the CobA enzyme (113). Thus, results

presented here indicate that PduS provides additional cobalamin reduction capacity required for maximal growth on 1,2-PD and may also be important within the confines of the Pdu MCP as discussed below.

FIG. 2.12. Proposed model for cobalamin recycling inside the Pdu MCPs of S. enterica. Prior studies showed that AdoCbl-dependent diol dehydratase (PduCDE) resides in the lumen of the Pdu MCP. PduCDE is subject to mechanism-based inactivation in which damaged Cbl cofactor (OH-Cbl) becomes tightly enzyme bound. Diol dehydratase reactivase (PduGH) releases OH-Cbl from PduCDE. Then, OH-Cbl is reduced by PduS to cob(I)alamin and adenosylated to AdoCbl by the PduO adenosyltransferase. AdoCbl spontaneously associates with apo-PduCDE to form active holoenzyme. The recycling of OH-Cbl is required to maintain the activity of PduCDE and hence is essential for the degradation of 1,2-propanediol as a carbon and energy source. The finding that PduS is a component of the Pdu MCP suggests that cobalamin recycling can occur entirely within the MCP lumen.

Results presented here demonstrated that PduS is associated with Pdu MCPs. PduS was identified as a component of purified MCPs following SDS-PAGE (Fig. 2.11A) and

MS-MS. In addition, Western blot showed that purified MCPs were substantially enriched

in PduS which established a specific association between PduS and the Pdu MCP (Fig.

2.11B). Furthermore, two-hybrid studies indicated an in vivo interaction between PduS and PduO which was previously localized to the Pdu MCP (94). Prior proteomics studies missed PduS as an MCP component (94). This was likely due to the difficulty in separating PduS from the PduP enzyme (a major MCP component) by SDS-PAGE (Fig.

2.11A). Alternatively, PduS may have been missed due to its high pI (8.82) which would have resulted in migration to the edge of the gel or off during isoelectric focusing step of

2D electrophoresis used in prior studies (94). Thus, substantial evidence indicates PduS is a component of the Pdu MCP. This establishes that all the enzymes needed for cobalamin recycling are components of the Pdu MCP suggesting that cobalamin recycling can occur entirely within the lumen of the Pdu MCP (Fig. 2.12). The AdoCbl-dependent diol dehydratase (PduCDE) required for 1,2-PD degradation which has been localized to the lumen of the Pdu MCP is subject to mechanism-based inactivation and requires cobalamin recycling for activity (154, 155). This significantly revises our understanding of how the Pdu MCP functions.

The finding that PduS is involved in cobalamin recycling might provide an insight into cobalamin reduction in human mitochondria, where the reductive reactions needed for the synthesis of AdoCbl are not well understood. AdoCbl is an essential cofactor for the mitochondrial methylmalonyl-CoA mutase (MUT or MCM) that converts methylmalonyl-CoA to succinyl-CoA during the catabolism of branched-chain amino acids, thymine, uracil, cholesterol, and odd-chain fatty acids (76). The current view is that cobalamin, probably as cob(II)alamin, is transported into the mitochondria, where the second reduction yielding cob(I)alamin occurs followed by adenosylation by human ATR to form AdoCbl (17). However, the human mitochondrial cobalamin reductase(s) have not

been identified. An earlier study showed that human methionine synthase reductase

(hMSR) can act as the reducing enzyme in combination with hATR to generate AdoCbl from cob(II)alamin in vitro (130). Initially, this suggested that hMSR might function as the elusive cobalamin reductase in mitochondria, but, this possibility now seems unlikely since a recent report indicates that the hMSR protein is restricted to the cytosol (84). Our sequence analyses showed that the Nqo1 subunits of bovine and human mitochondrial respiratory complex I have 25% amino acid identity (44% similarity, expect = 8x10-9) to the N-terminal region (amino acids 1-165) of the PduS protein of S. enterica. PduS and

Nqo1 also share SLBB domains, (amino acids 173-221 in PduS) which have 33% amino acid identity (expect = 7x10-4) and this domain was proposed to bind cobalamin in the

PduS protein (46). Moreover, PduS and Nqo1 both contain binding sites for NADH, FMN, and an iron-sulfur cluster, involved in electron transfer (Fig. 2B). Although there is currently no evidence for a soluble ligand interacting with the SLBB domain in Nqo1, it’s possible that cobalamin might bind this domain and accept electrons from Nqo1 given the similarity between PduS and Nqo1. Hence, Nqo1 might have dual functions in electron transport for a respiratory chain and cobalamin reduction for AdoCbl synthesis, although this is speculative at this time.

2.6. Acknowledgements

This work was supported by NSF grant MCB0616008 to TAB. We are indebted to Dr.

Yasuhiro Takahashi from the Division of Life Science at Saitama University (Japan) for offering the plasmids pRKNMC, pRKISC, and pRKSUF017. We thank Dr. Siquan Luo from the Proteomics Facility at Iowa State University for aid with MS-MS analyses, Dr.

Tracie A. Bierwagen in the ISU BBMB Department for help with gel filtration, and the

ISU DNA Sequencing and Synthesis Facility for assistance with DNA analyses and the

ISU W.M. Keck Metabolomics Research Laboratory for assistance with the mass spectrometry analyses.

Chapter 3. PduQ is a 1-propanol dehydrogenase associated with the Pdu

microcompartment of Salmonella enterica

3.1. Abstract

Salmonella enterica degrades 1,2-propanediol (1,2-PD) using a bacterial microcompartment (MCP). The MCP consists of a solid protein shell encasing metabolic enzymes such as propionaldehyde dehydrogenase PduP which uses NAD+ as a cofactor.

In this work, the His6-tagged PduQ protein from S. enterica was produced at high levels in Escherichia coli, purified and characterized. PduQ was identified as an iron-dependent alcohol dehydrogenase (ADH) that reduces propionaldehyde to 1-propanol with NADH as electron donor under physiological conditions although it is able to catalyze the reverse reaction as well. Our results also demonstrated that PduQ is a component of Pdu MCPs.

Genetic studies showed that a pduQ deletion impaired the growth of Salmonella on

1,2-PD supporting a role in NAD+ regeneration within the MCP in vivo. PduQ is also shown to interact with PduP. An N-terminal signal sequence was previously shown to direct packaging PduP into Pdu MCPs. Here PduQ might use an alternative packaging strategy by which it specifically binds to PduP.

3.2. Introduction

Salmonella enterica degrades 1,2-propanediol (1,2-PD) via a coenzyme B12

(Adenosylcobalamin, AdoCbl) -dependent pathway (110). 1,2-PD is a major product of the anaerobic degradation of rhamnose and fucose that are common in plant cell walls, bacterial capsules and the glycoconjugates of eukaryotic cells. The ability to degrade

1,2-PD is thought to provide a selective advantage in anoxic environments such as the

large intestines of higher animals, sediments and the depths of soils (138, 163).

Twenty-four genes specifically involved in 1,2-PD utilization (pdu) are found in a single contiguous cluster, the pdu locus (31, 110). This locus consists of the pocR and pduF genes as well as the adjacent, but divergently transcribed pdu operon which includes 22 pdu genes (pduABB'CDEGHJKLMNOPQSTUVWX). The pdu locus encodes a transcriptional regulator (30, 184), a diffusion facilitator (51), enzymes for the degradation of 1,2-PD (31, 32, 129, 139, 164) and cobalamin metabolism (31, 74, 112,

156, 194), as well as proteins for the formation of a bacterial microcompartment (MCP) used for 1,2-PD utilization (31, 61, 94, 95, 193).

Bacterial MCPs are simple proteinaceous organelles used by diverse bacteria to optimize metabolic pathways that have toxic or volatile intermediates (28, 48, 53, 245).

They are polyhedral in shape, 100-150 nm in cross-section (about the size of a large virus), and consist of a protein shell that encapsulates sequentially-acting metabolic enzymes. Based on sequence analyses, it is estimated that MCPs are produced by 20-25% of all bacteria and function in seven or more different metabolic processes (53). The function of the Pdu MCP is to confine the propionaldehyde formed in the first step of

1,2-PD degradation in order to mitigate its toxicity and prevent DNA damage (31, 94,

193). The Pdu MCP was purified and 15 polypeptide components were identified

(PduABB'CDEGHJKOPSTU) all of which are encoded by the pdu locus (52, 94).

A prior study indicated that the pduP gene codes for a lumen propionaldehyde dehydrogenase converting 1,2-PD to propionaldehyde using NAD+ as a cofactor (129).

This raises the question of how NAD+ crosses the MCP shell or/and how NAD+ is regenerated within the Pdu MCP. Recent crystallography studies suggested that some shell proteins such as PduA, PduT and PduU, may mediate the transport of enzyme

substrates/cofactors across the MCP shell (60, 62). However, localized regeneration of these cofactors within the MCP might allow more efficient 1,2-PD degradation.

A recent study indicated that a short N-terminal sequence of some lumen enzymes such as PduP efficiently packages the proteins into the Pdu MCP (75). Nevertheless, not all of the enzymes located in the MCP possess such a terminal extension. Therefore, other mechanism(s) may be applied for protein targeting.

In this study, PduQ from S. enterica was purified and was identified as an oxidoreductase that catalyzes the interconversion between propionaldehyde and

1-propanol. The result also showed that PduQ is a component of the Pdu MCP. This finding suggests that, besides cobalamin (52), the Pdu MCP is able to recycle NAD+ internally. In addition, an alternative mechanism for targeting proteins into MCPs was proposed based on the interaction between PduQ and PduP.

3.3. Materials and methods

Bacterial strains and growth conditions. The bacterial strains used in this study are listed in Table 3.1. The rich media used were Luria-Bertani/Lennox (LB) medium (Difco,

Detroit, MI) (151) and Terrific Broth (TB) (MP Biomedicals, Solon, OH) (226). The minimal medium used was no-carbon-E (NCE) medium (25, 233).

Chemicals and reagents. Antibiotics, cyanocobalamin (CN-Cbl), DNase I, NAD+,

NADH, NADP+, and NADPH were from Sigma Chemical Company (St. Louis, MO).

IPTG was from Diagnostic Chemicals Limited (Charlotteville PEI, Canada). PfuUltra high-fidelity DNA polymerase was from Stratagene (La Jolla, CA). Taq DNA polymerase, restriction enzymes, and T4 DNA ligase were from New England Biolabs (Beverly, MA).

Other chemicals were from Fisher Scientific (Pittsburgh, PA).

TABLE 3.1. Bacterial strains used in this study. Species and strain Genotype Source E. coli BE237 BL21(DE3) RIL/pET-41a Lab collection BE272 BL21(DE3) RIL/pTA925-pduP Lab collection BE1421 BL21(DE3) RIL/pET-41a-His6-pduP Lab collection BE1422 BL21(DE3) RIL/pET-41a-pduP-His6 Lab collection BE1500 BL21(DE3) RIL/pET-41a-pduQ This work BE1501 BL21(DE3) RIL/pET-41a-His6-pduQ This work BE1502 BL21(DE3) RIL/pET-41a-pduQ-His6 This work S. enterica serovar Typhimurium LT2 BE191 ΔpduP Lab collection BE269 ΔpduP/pLAC22 Lab collection BE270 ΔpduP/pLAC22-pduP Lab collection BE287 LT2/pLAC22 Lab collection BE903 ΔpduQ::frt This work BE919 ΔpduQ::frt/pLAC22 This work BE942 ΔpduQ::frt/pLAC22-pduQ This work

Construction of plasmids and a pduQ-deletion mutant. The pduQ gene was amplified by PCR (using genomic DNA of S. enterica as the template) then cloned into pET-41a vector (Novagen, Cambridge, MA) using NdeI and HindIII restriction sites incorporated into the PCR primers as previously described (192). The genes for the production of His6-PduQ and PduQ-His6 were similarly cloned with the sequence encoding the His6-tag incorporated into either the forward or the reverse PCR primer. The resulting plasmids pET-41a-pduQ, pET-41a-His6-pduQ, as well as the pET-41a-pduQ-His6 were introduced into Escherichia coli BL21 (DE3) RIL (Stratagene).

The native pduQ gene was also cloned into pLAC22 (237) between BglII and HindIII sites for complementation experiments. Vector pLAC22 allows tight regulation of protein production by IPTG (237). All of the above inserts were verified by DNA sequencing.

The pduQ deletion mutant was constructed by a PCR-based method (64). The deletion removed nearly the entire pduQ coding sequence, but left predicted translation signals of the adjacent genes (pduP and pduS) intact. The presence of the pduQ deletion was verified by PCR as described previously (64) and by genomic DNA sequencing.

Phage P22 transduction. Transductional crosses were performed as described using

P22 HT105/1 int-210, a mutant phage that has high transducing ability (201).

Transductants were tested for phage contamination and sensitivity by streaking on green plates against P22 H5.

Growth of expression strains and purification of PduQ-His6. Four hundred ml of

TB containing 25 mg/ml kanamycin was inoculated with 2 ml of BE1502 grown in similar medium and the cells were cultivated in a 1 liter baffled Erlenmeyer flask at 37 °C with shaking at 275 rpm. When the culture reached an optical density at 600 nm (OD600) of about 0.5, 0.5 mM IPTG was added. The culture was incubated for additional 20 h at

16 °C and 250 rpm shaking. Cells were then harvested by centrifugation at 4 °C and

6,000 x g for 10 min and used immediately for protein purification under strictly anaerobic conditions or under aerobic conditions as previously described except using buffer A (50 mM potassium phosphate, pH 8.5, 300 mM NaCl, 5 mM β-mercaptoethanol,

0.4 mM AEBSF [4-(2-aminoethyl) benzenesulfonyl fluoride-HCl]) at pH 8.5 (52).

Control strains BE237, BE1500 were grown in parallel with expression strain BE1502 and similar procedures were used for preparing whole-cell extracts and soluble fractions.

SDS-PAGE and Western blots. Protein concentration was determined using Bio-Rad

(Hercules, CA) protein assay reagent with bovine serum albumin (BSA) as a standard.

SDS-PAGE was performed using Bio-Rad 12% or 10-20% gradient Tris-HCl ready gels.

Protein bands were visualized by staining with Bio-Safe Coomassie Stain (Bio-Rad). For

Western blots, the proteins on SDS-PAGE gels were transferred to polyvinylidene fluoride (PVDF) membranes and probed using SuperSignal West Pico Chemiluminescent

Substrate (Pierce, Rockford, IL) according to the manufacturer’s instructions with primary rabbit antisera against synthetic peptide ADNRISVFSEITPD (present at

positions 51-64 in PduQ of S. enterica) (Genscript, Piscataway, NJ) at the final concentration of 0.5 μg/ml and secondary goat anti-rabbit IgG-HRP (Santa Cruz

Biotechnology, Santa Cruz, CA) at 40 ng/ml.

Enzymatic activity assays. The catalytic assays were performed under anaerobic conditions. All reactions were initiated by addition of NADH or NAD+ to assay mixtures except where stated otherwise. Propionaldehyde reduction assays contained 100 mM

Tris-HCl (pH 7.0), 0.4 mM NADH, 150 mM propionaldehyde and the enzyme as indicated in the text. 1-propanol oxidation assays contained 100 mM Tris-HCl (pH 9.0), 1 mM NAD+, 500 mM 1-propanol and the enzyme. The interconversion between propionaldehyde and 1-propanol was monitored spectrophotometrically by measuring the absorbance change at 340 nm due to consumption or formation of NADH and quantified

-1 -1 using Δε340 = 6.22 mM cm .

Metal analysis. The content of Fe in the purified protein was determined by a high-resolution double-focusing inductively coupled plasma mass spectrometry (ICP-MS) using a Finnigan Element 1 (Thermo Scientific) operated at medium resolution (m/Δm =

4,000) as described in a prior report (235). Prior to metal analysis samples were applied to a Sephadex™ G-25 PD-10 desalting column (GE Healthcare, Piscataway, NJ) pre-equilibrated with MilliQ water (resistivity >18 m ) and then were eluted with MilliQ water inside an anaerobic chamber or under aerobic condition in order to eliminate reagents and metals unbound to the protein. Eluates were collected in metal-free glass tubes. Blank water samples, representing the eluates of the Ni-NTA elution buffer collected in the same volume range from a parallel PD-10 column, were also analyzed.

Ga was used as an internal standard to quantify the elemental concentrations of interest.

Growth studies. Growth studies were performed as previously described (139) with

modifications. 2 ml LB broth containing 100 μg/ml ampicillin if needed was inoculated with a fresh single colony and incubated overnight at 37 °C with shaking at 275 rpm.

Cells were collected from the overnight culture by centrifugation at 10,000 x g for 1 min.

The cell pellet was washed once and resuspended in NCE medium supplemented with

0.6% 1,2-PD, 1 mM MgSO4, 50 μΜ Ferric citrate, and 0.12 mM each of valine, isoleucine, leucine, threonine. The resulting cell suspension was used to inoculate 0.3 ml

(final volume) of similar medium containing CN-Cbl to an optical density of around 0.1 at 600 nm. Cultures were grown in 48-well flat-bottom plates (Falcon) at 37 °C with fast shaking and OD600 was measured using a Synergy HT Microplate reader (BioTek,

Winooski, VT). For complementation studies, IPTG was used at 10 μM to induce expression of genes cloned into pLAC22.

Pdu MCP purification. Cells were grown at 37 °C with shaking at 275 rpm to an

OD600 of 1.0-1.2 in 400 ml NCE medium supplemented with 1 mM MgSO4, 0.5% succinate, and 0.6% 1,2-PD inoculated with 2 ml of an overnight LB culture. For strains carrying plasmids (pLAC22 or derivatives), the medium additionally contains 100 μg/ml ampicillin and 20 μM IPTG. Cells were harvested by centrifugation and washed twice with 40 ml buffer B (50 mM Tris-HCl, pH 8.0, 500 mM KCl, 12.5 mM MgCl2, 1.5%

1,2-PD). Cells were resuspended in 25 ml buffer C [1:1.5 bufferB:B-PER II (Pierce)] containing 5 mM β-mercaptoethanol, 0.4 mM ABESF, 25 mg lysozyme, and 2 mg DNase

I and incubated at room temperature with 60 rpm shaking for 30 min and then on ice for 5 min. Cell debris was removed by centrifugation twice at 12,000 x g for 5 min at 4 ˚C and the MCPs were spun down at 20,000 x g for 20 min at 4 ˚C. The pellet was washed once in 10 ml buffer C containing 5 mM β-mercaptoethanol and 0.4 mM ABESF and resuspended in 1 ml buffer D (50 mM Tris-HCl, pH 8.0, 50 mM KCl, 5 mM MgCl2, 1%

1,2-PD) containing 5 mM β-mercaptoethanol and 0.4 mM ABESF. The remanent cell debris was removed at 12,000 x g for 1 min at 4 ˚C for three times. Purified MCPs were stored at 4 °C until used.

MALDI-TOF MS-MS. The tandem mass spectroscopy was performed as previously described (52).

His-tag protein affinity pull-down assays. The soluble cell lysates containing recombinant His-tagged protein bait were applied to the pre-equilibrated Ni-NTA column.

After washing away the unbound proteins with buffer A containing 100 mM imidazole, the soluble cell lysates containing recombinant unfused protein prey were loaded onto the column where the bait was immobilized. Following washing, the protein-protein interaction complex was eluted from the column and was detected by SDS-PAGE.

3.4. Results

Sequence analysis. The PduQ protein of S. enterica is composed of 370 amino acid residues (31). Over 100 PduQ homologues present in GenBank are associated with

1,2-PD degradation based on gene proximity. According to the amino acid sequence,

PduQ of S. enterica belongs to the family of iron-activated group III alcohol dehydrogenase (Pfam0456) (178). It has 32% sequence identity (49% overall similarity) with the 1,2-propanediol dehydrogenase from E. coli (153), and 34% identity (51% overall similarity) with 1,3-propanediol dehydrogenase from Klebsiella pneumoniae

(143). PduQ also shares 36% identity (52% overall similarity) with the Salmonella putative ethanol dehydrogenase EutG involved in ethanolamine utilization (31, 219).

PduQ possesses conserved binding sites for substrates and cofactors, including an

89 91 241 259 NAD-binding motif ( GGG ) and a motif ( GX2HX2AHX2GX5PHG where X

denotes any amino acid) with conserved Asp175 and His179 involved in Fe-binding (11,

143, 153) (Fig. 3.1). Although PduQ contains another classic NAD(P)-binding fingerprint

(13GXGXX[A/G]18) (37), previous studies indicated that this pattern was not conversed or involved in binding NAD(P) in group III alcohol dehydrogenase (63, 153).

FIG. 1. Sequence alignment by clustal X2. EcFucO, 1,2-propanediol dehydrogenase from Escherichia coli (GI 16130706); KpDhaT, 1,3-propanediol dehydrogenase from Klebsiella pneumoniae (GI 940440); SeEutG, ethanol dehydrogenase from Salmonella enterica (GI 687647); SePduQ, propanol dehydrogenase from S. enterica (GI 5069460).

Expression and purification of PduQ-His6 protein. E. coli strain BE1502 was constructed to produce high levels of C-terminal-His6-tagged PduQ protein via a T7 expression system. This strain produced relatively large amounts of protein near the expected molecular mass of PduQ-His6 (40.3 kDa) (Fig. 3.2). In contrast, cell extracts from a control strain containing the expression plasmid without insert (BE237) produced much less protein near 40.3 kDa. PduQ-His6 was purified under both aerobic and anaerobic conditions by Ni-NTA affinity chromatography. Based on SDS-PAGE anaerobically purified PduQ-His6 protein was about 95% homogenous (Fig. 3.2, lane 4), and a similar level of purity was obtained for aerobically purified enzyme.

FIG. 2. SDS-PAGE analysis of the anaerobic purification of PduQ-His6 from BE1052. Lane 1, protein standards; lane 2, 10 μg whole-cell extract; lane 3, 10 μg soluble fraction; lane 4, 2 μg purified

PduQ-His6. The gel was Bio-Rad 10-20% gradient ready gel and was stained with Coomassie.

In vitro catalytic activities of the PduQ-His6 enzyme. PduQ-His6 was purified using strictly anaerobic conditions and tested for the activities as a propionaldehyde reductase and as a propanol dehydrogenase after each purification step (Table 3.2).

The whole-cell extract from the PduQ-His6 expression strain (BE1502) exhibited about

23-fold higher propionaldehyde reductase activity (2.8 ± 0.2 μmol min-1 mg-1) than did extracts from the control strain carrying the expression plasmid without insert (BE237)

-1 -1 (0.12 ± 0.03 nmol min mg ). Purification of the PduQ-His6 protein by anaerobic

Ni-NTA chromatography increased the propionaldehyde reductase specific activity approximately 19.8-fold to 55.5 ± 4.2 μmol min-1 mg-1.

TABLE 3.2. Anaerobic purification of PduQ-His6. Total Propionaldehyde reductasea Propanol dehydrogenaseb Purification step protein Specific activity Total activity Purification Specific activity Total activity Purification (mg) (μmol min-1 mg-1) (μmol min-1) fold (μmol min-1 mg-1) (μmol min-1) fold Whole-cell extract 403.6 2.8 ± 0.2c 1130 1 0.8 ± 0.1c 322.9 1 Soluble fraction 246.4 3.1 ± 0.3c 763.8 1.1 0.9 ± 0.2c 221.8 1.1 Ni-NTA eluate 4.4 55.5 ± 4.2 244.2 19.8 5.1 ± 0.7 22.4 6.4 abAssayes were performed with 100 mM Tris-HCl at pH 7.0 and pH 9.0, respectively. cThe values of the background activities were not subtracted.

Assays showed that the PduQ-His6 protein also catalyzed the reverse reaction, oxidation of 1-propanol. Whole cell extracts of the expression strain BE1502 had 0.8 ±

0.1 μmol min-1 mg-1 1-propanol dehydrogenase activity. While control extracts from

BE237 also exhibited 1-propanol dehydrogenase activity (0.42 ± 0.08 μmol min-1 mg-1).

Purification of the PduQ-His6 protein by anaerobic Ni-NTA chromatography increased the 1-propanol dehydrogenase specific activity about 13.4-fold to 5.1 ± 0.7 μmol min-1 mg-1.

The alcohol dehydrogenase(s) in the expression host stain contributed to the values in

Table 3.2 and displayed relatively high activity and preference for 1-propanol oxidation under the tested conditions, which partially accounts for the incongruity of the purification folds in two activities of the purified protein.

PduQ-His6 was also purified under aerobic conditions. Aerobically purified PduQ-His6 retained about 20% activity, indicating that the enzyme is oxygen-labile.

The C-terminal His6-tag had no significant effect on PduQ enzymatic activities based on the comparisons of the catalytic activities in the whole-cell extracts and soluble fractions of BE1502 and BE1500, which respectively express PduQ-His6 and unfused

PduQ.

Reaction requirements. To determine the PduQ reaction requirements, key assay components were individually omitted. For propionaldehyde reduction, there was no activity in the absence of NADH, propionaldehyde, or PduQ-His6. In the propanol dehydrogenase assays, no activity was measurable in the absence of NAD+, 1-propanol, or PduQ-His6.

100

40 propionaldehyde reduction 1-propanol oxidation 20 Relative acitivity (%) 0 567891011 pH

FIG. 3.3. pH dependence of propionaldehyde reduction and 1-propanol oxidation activities catalyzed by PduQ-His6 of S. enterica. The maximal activities for propionaldehyde reduction and 1-propanol oxidation were achieved at pH 7.0 and 9.0, respectively. The assasy buffer was 100 mM Tris-HCl.

Effects of pH on PduQ-His6 activity. The pH optima for PduQ-His6 activities were determined in 100 mM Tris-HCl buffer with pH value ranging from 6.0 to 10.0. The anaerobically purified PduQ-His6 exhibited maximal activity for propionaldehyde reduction at pH 7.0 while the maximal 1-propanol dehydrogenase activity was achieved at pH 9.0 (Fig. 3.3), which is consistent with the common feature of pH preference of several alcohol dehydrogenases (178).

Preference for the coenzyme of PduQ-His6. To determine the cofactor specificity of

PduQ-His6 both catalytic activities were assayed in the presence of either NADH or

NADPH at 0.4 mM and of either NAD+ or NADP+ at 1 mM. Results showed that anaerobically purified PduQ-His6 enzyme had 13% relative activity for propionaldehyde redution with NADPH compared to NADH and 11% relative activity for 1-propanol

oxidation with NADP+ compared to NAD+, showing that PduQ has a preference for

NAD(H).

x104 25 Fe

10 Intensity (cps) Intensity 5

0 55.0 55.5 56.0 56.5 57.0 m/z

FIG. 3.4. Identification of Fe in purified PduQ-His6 using ICP-MS (cps: counts per second).

Characterization of Fe ion in PduQ-His6. Fe was detected from PduQ-His6 protein using ICP-MS (Fig. 3.4). The anaerobically purified PduQ-His6 protein contains 1.01 ±

0.04 Fe-atom per monomeric unit while the aerobically purified PduQ-His6 protein contains 0.18 ± 0.02 Fe-atom per monomeric unit, indicating that iron was lost much readily in the presence of oxygen. In addition, the PduQ catalytic activities were almost totally inhibited by 1 mM EDTA. In combination with the above results that the anaerobically purified PduQ-His6 has much higher enzymatic activity, PduQ is iron-dependent and oxygen-labile.

Linearity of the reactions. The effects of PduQ-His6 concentration on enzymatic activities were determined. Propionaldehyde reductase was proportional to PduQ-His6

concentration from 10 to 95.5 nM when 0.4 mM NADH and 150 mM propionaldehyde were used as substrates. Linear regression yielded an R2 value of 0.9981. Propanol

2 dehydrogenase was linear from 0.16 to 1.4 μM PduQ-His6 (R = 0.9994) when the assay mixture contains 1 mM NAD+ and 500 mM 1-propanol.

Kinetic analysis of PduQ-His6 activities. Steady-state kinetic studies for propionaldehyde reduction and 1-propanol oxidation were performed using anaerobically purified PduQ-His6 with varied concentrations of one substrate and a fixed concentration of the other substrate. Kinetic parameters were obtained by non-linear curve fitting to the

Michaelis-Menten equation v = Vmax [S]/(Km + [S]) using GraphPad Prism 5 Software

(GraphPad Software, San Diego, CA). For propionaldehyde reduction, the apparent Km values for NADH and propionaldehyde were 45.3 ± 5.3 μM and 16.0 ± 2.0 mM,

–1 –1 respectively (Table 3.3). The enzyme Vmax was 77.7 ± 3.0 or 81.3 ± 4.5 μmol min mg when propionaldehyde or NADH was held at 150 mM and 400 µM, respectively.

For 1-propanol oxidation, non-linear regression indicated an apparent Km of 267.5 ±

22.3 μM and 95.8 ± 9.2 mM for NAD+ and 1-propanol, respectively (Table 3.3). The

–1 –1 –1 –1 enzyme Vmax was 9.2 ± 0.7 μmol min mg or 8.4 ± 0.8 μmol min mg when the

1-propanol or NAD+ was held at fixed concentrations of 500 mM and 1 mM, respectively.

TABLE 3.3. Kinetic parameters for propionaldehyde reduction and 1-propanol oxidation by anaerobically purified PduQ-His6. K a V a k k /K Reaction Variable substrate m max cat cat m (μM) (µmol min-1 mg-1) (s-1) (μM-1 s-1) Propionaldehy NADH 45.3 ± 5.3 77.7 ± 3.0 52.2 1.15 de reductionb propionaldehyde (16.0 ± 2.0 ) x 103 81.3 ± 4.5 54.6 3.41 x 10-3 1-propanol NAD+ 267.5 ± 22.3 9.2 ± 0.7 6.18 2.31 x 10-2 oxidation c 1-propanol (95.8 ± 9.2 ) x 103 8.4 ± 0.8 5.64 5.89 x 10-5 a The values of Km and Vmax were from non-linear regression using GraphPad Prism 5. bc Assayes were performed with 100 mM Tris-HCl at pH 7.0 and pH 9.0, respectively.

Phenotype and complementation of a pduQ deletion mutation. To investigate the function of PduQ in vivo, we measured the growth of a pduQ deletion mutant (BE903) on

1,2-PD minimal medium. BE903 grew slowly and to a lower cell density than wild-type S. enterica on 1,2-PD minimal medium supplemented with either saturating or limiting levels of CN-Cbl (100 or 20 nM) (Fig. 3.5A, B). Complementation analysis was also conducted. Production of PduQ from pLAC22 fully corrected the impaired growth phenotype of the ΔpduQ mutant at saturating CN-Cbl (Fig. 3.5C) and limiting CN-Cbl

(not shown) confirming that impaired growth was due to the pduQ deletion, and not due to polarity or an unknown mutation. The growth curves shown in Figure 3.5 A-C were all repeated at least three times with triplicate cultures. The growth phenotype of the pduQ deletion mutant suggests a role for PduQ in NAD+ recycling in vivo as further described in the discussion section.

PduQ is a component of Pdu MCP. To determine the cellular location of PduQ,

MCPs purified from wild-type S. enterica and pduQ deletion mutant BE903, were analyzed by SDS-PAGE and MALDI-TOF MS-MS. A band near the expected molecular mass of PduQ (39.5 kDa) was observed on an SDS-PAGE gel of MCPs purified from the wild-type strain (Fig. 3.6 lane 2) while absent in MCPs purified from the pduQ deletion mutant (Fig. 3.6 lane 3). This band was further analyzed by MALDI-TOF MS-MS. All three sequences (MNTFSLQTRL, RFNAGVPRA, RIPAMVQAALADVTLRT) identified by MS-MS of a trypsin digest matched the PduQ protein. Western blot using the antisera against a small peptide in PduQ also detected a band neat 39 kDa from the purified MCP.

These results indicated that PduQ protein is a component of Pdu MCPs.

A 0.9 0.8 0.7 0.6 0.5

600 0.4

OD 0.3

0.2 LT2 ΔpduQ 0.1 0.0 0 5 10 15 20 25 30 35 40 Time (h) B 0.35 0.30

0.25 0.20

600 0.15 LT2 OD 0.10 ΔpduQ 0.05

0.00 0 5 10 15 20 25 30 35 40 45 Time (h) C 0.9 0.8 0.7 0.6 0.5

600 0.4 OD 0.3 LT2/pLAC22 0.2 ΔpduQ/pLAC22

0.1 ΔpduQ/pLAC22-pduQ

0.0 0 5 10 15 20 25 30 35 40 Time (h)

FIG. 3.5. Phenotype and complementation of a pduQ deletion mutation. Cells were grown in NCE minimal medium with 1,2-PD as a sole carbon and energy source. The pduQ deletion impaired growth and decreased cell density on 1,2-PD supplemented with saturating (100 nM) CN-Cbl (A) and limiting (20 nM) CN-Cbl (B). The pduQ deletion was complemented by ectopic expression of PduQ induced by 10 μM IPTG at 100 nM CN-Cbl (C).

FIG. 6. PduQ is a component of the Pdu MCP and disappears in the absence of PduP. 10-20% SDS-PAGE gel stained with Coomassie brilliant blue. Lane 1, molecular mass markers; lane 2, 10 μg Pdu MCPs purified from wild-type Salmonella; lane 3, 10 μg Pdu MCPs purified from a pduQ deletion mutant BE903; lane 4, 10 μg Pdu MCPs purified from a pduP deletion mutant BE191.

Interaction between PduQ and PduP. The Pdu MCPs purified from the pduP deletion mutant BE191 contained much less PduQ protein than those from the wild type

(Fig. 3.6 lane 4 and 2), suggesting an interaction between PduQ and PduP. To examine the potential interaction of PduQ and PduP in vitro, His-tag pull-down assays were performed.

The following SDS-PAGE demonstrated that unfused PduQ was co-purified with the N- or C-terminally His-tagged PduP and unfused PduP was co-purified with the C-terminally

His-tagged PduQ (Fig. 3.7). Control experiments showed that unfused PduQ and unfused

PduP could not bind to the Ni-NTA column. These results indicated that PduQ specifically interacts with PduP. Worthy of mention is that the unfused PduP was not co-purified with the N-terminal-His-tagged PduQ (Fig. 3.7 lane 7), suggesting that the

N-terminus of PduQ may be involved in the PduQ-PduP interaction.

FIG. 3.7. His-tag protein affinity pull-down assays. Lane 1, protein standards; lane 2, His6-PduP; lane 3,

His6-PduP+PduQ; lane 4, PduP-His6; lane 5, PduP-His6+PduQ; lane 6, His6-PduQ; lane 7,

His6-PduQ+PduP; lane 8, PduQ-His6; lane 9, PduQ-His6+PduP.

3.5. Disscusion

Alcohol dehydrogenases (ADHs) are a family of oxidoreductases that catalyze the interconversion between alcohols and aldehydes or ketones, and they are widely distributed in virtually all living organisms (178). ADHs can be divided into three classes based on their cofactor specificity: (i) NAD(P), (ii) pyrollo-quinoline quinone (PQQ), heme or cofactor F420, and (iii) flavin adenine dinucleotide (FAD). The

NAD(P)-dependent alcohol dehydrogenases are the best-studied class and are generally subdivided in three major groups: zinc-dependent long-chain ADHs (group I), zinc-independent short-chain ADHs (group II), and the iron-activated ADHs (group III)

(178).

The results presented in this report demonstrated that PduQ encoded by the pdu locus of S. enterica is an iron-dependent ADH which catalyzes the interconversion between propionaldehyde and 1-propanol. The data showed that the propionaldehyde reduction

activity was about 45 times higher than that of the 1-propanol oxidation at pH 7.0, indicating that this enzyme is more efficient for catalyzing aldehyde reduction in cells where approximatedly neutral pH is maintained. Moreover, kinetic data in Table 3.3 exhibited high catalytic efficiency for the propionaldehyde reduction and high affinity for the substrate and cofactor involved in this reaction, which also suggest that the physiological function of PduQ is the reduction of aldehyde with the consumption of

NADH to NAD+.

To examine the role of PduQ in vivo, we tested the effects of a pduQ deletion mutation on AdoCbl-dependent growth on 1,2-PD with saturating or limiting CN-Cbl (Fig. 3.5A,

B). Under both conditions, the pduQ deletion impaired growth and led to a lower cell density, indicating that PduQ was required to support the maximal rate of 1,2-PD degradation by Salmonella under the conditions used.

Within the MCP protein shell, one of the main catabolic enzymes for 1,2-PD utilization, propionaldehyde dehydrogenase PduP, catalyzes the reaction of propionaldehyde to propionyl-CoA using NAD+ as a cofactor. As a result, the electron carrier NAD+ must cross the solid protein shell from the cytosol or/and be regenerated inside the MCPs to avoid depletion of the lumenal NAD+. Recent crystallography studies suggested that some shell proteins such as PduA, PduT and PduU, may mediate the transport of enzyme substrates/cofactors across the MCP shell (60, 62). Nonetheless, localized regeneration of cofactors within the MCP might be advantageous for both the activity of the metabolic enzymes and the efficiency of 1,2-PD degradation. Our recent study indicates that cobalamin, a cofactor for another lumen enzyme diol dehydrogenase PduCDE, is recycled within the MCPs (52). In this study, SDS-PAGE of the Pdu MCPs purified from the wild type Salmonella (Fig. 3.6) and the following MS-MS analyses demonstrated that PduQ is

a component of Pdu MCPs. In conjunction with its in-vitro and in-vivo function and properties discussed above, PduQ may serve to help balance redox reactions during

1,2-PD fermentation by converting a portion of the propionaldehyde to 1-propanol and

NADH to NAD+, which support the NAD+-recycling mechanism within the MCPs (Fig.

3.8).

FIG. 3.8. Proposed function of PduQ in NAD+ regeneration inside the Pdu MCPs of S. enterica. The dashed line indicates the shell of the MCP. Prior studies showed that propionaldehyde dehydrogenase (PduP) resides in the lumen of the Pdu MCP and PduP uses NAD+ as a coenzyme. The MCP lumen enzyme PduQ helps recycle the electron carriers and balance redox reactions by converting a portion of the propionaldehyde to 1-propanol and NADH to NAD+ to avoid depleting NAD+ within the MCP lumen.

A recent study demonstrated that a short N-terminal signal peptide of some lumen enzymes such as PduP can direct packaging proteins into the Pdu MCPs through binding specific sites on the interior surface of shell protein assemblies (75). However, the MCP lumen enzyme PduQ lacks an obvious packaging signal sequence in comparison with

EutG (Fig. 3.1), a PduQ functional counterpart in Eut MCPs involved in ethanolamine utilization (219). In this report, the SDS-PAGE of the MCPs purified from the pduP deletion mutant only displayed trace amount of PduQ protein (Fig. 3.6 lane 4), and the

His-tag pull-down assays demonstrated a specific interaction between PduQ and PduP

(Fig. 3.7). These results suggested an alternative packaging strategy by which some lumen enzymes lacking N-terminal signal sequences specifically bind to the shell-immobilized proteins that mediate the targeting of these enzymes into the MCPs.

3.6. Acknowledgements

This work was supported by NSF grant MCB0956451 to TAB. We thank Megan

Mekoli and Travis Witte from Dr. Robert S. Houk group in the Department of Chemistry at Iowa State University for help with ICP-MS analyses, Siquan Luo from the ISU Protein

Facility for aid with MS-MS analyses, and the ISU DNA Sequencing and Synthesis

Facility for assistance with DNA analyses.

100

Chapter 4. Conclusions

A number of enteric and lactic acid bacteria such as S. enterica grow on

1,2-propanediol (1,2-PD) in a coenzyme B12-dependent fashion (163). 1,2-PD is a major product of the anaerobic degradation of the common plant sugars, rhamnose and fucose (2,

10, 12, 35, 36, 138). The ability to degrade this compound is thought to provide a selective advantage in anaerobic environments such as the large intestines of higher animals, sediments and the depths of soils. Prior studies have shown that 1,2-PD degradation is one of the most complex metabolic processes known. The degradation of this small molecule requires an unusual proteinous microcompartment (Pdu MCP) that sequesters a toxic metabolic intermediate propionaldehyde (31, 193). The Pdu MCP consists of a solid protein shell that encapsulates sequentially-acting metabolic enzymes, including the diol dehydratase PduCDE and the propionaldehyde dehydrogenase PduP that respectively use AdoCbl and NAD+ as cofactors. During catalysis, the adenosyl-group of AdoCbl bound to PduCDE is periodically lost due to by-reactions and is usually replaced by a hydroxo-group resulting in the formation of an inactive

OH-Cbl-enzyme complex (228), and the NAD+ is converted to NADH by PduP (129), which makes it necessary to regenerate or/and replenish AdoCbl and NAD+.

Recent crystallography studies suggested that some shell proteins such as PduA, PduT and PduU, have pores that may mediate the transport of enzyme substrates/cofactors across the MCP shell (60, 62). However, localized regeneration of these cofactors within the MCP might be advantageous for both the activity of the metabolic enzymes and the efficiency of 1,2-PD degradation.

Cobalamin recycling within the Pdu MCP

101

The first part of this study (Chapter 2) investigated the bifunctional cobalamin reductase PduS. This protein was over-expressed, purified, and characterized including its kinetic properties. The results indicated that PduS is a monomer and each monomer of

PduS contains one non-covalently bound FMN and two [4Fe-4S] clusters. Purified PduS enzyme exhibited the ability to reduce cob(III)alamin to cob(II)alamin and cob(II)alamin to cob(I)alamin in the presense of a trapping agent in vitro. Genetic studies showed that a pduS deletion decreased the growth rate of Salmonella on 1,2-PD supporting a role in cobalamin reduction in vivo. Further SDS-PAGE and Western blot of purified Pdu MCP and MS-MS analysis demonstrated that the PduS protein is a component of the Pdu MCP.

In addition, two-hybrid experiments indicated that PduS interacts with the PduO adenosyltransferase which is also located in the Pdu MCP and catalyzes the terminal step of AdoCbl synthesis.

The finding that PduS is a MCP protein, establishes that all the enzymes needed for cobalamin recycling are components of the Pdu MCP suggesting that cobalamin recycling can occur entirely within the lumen of the Pdu MCP: diol dehydratase reactivase PduGH releases OH-Cbl from PduCDE; then, OH-Cbl is reduced by PduS to cob(I)alamin and adenosylated to AdoCbl by the PduO adenosyltransferase; finally, AdoCbl spontaneously associates with apo-PduCDE to form active holoenzyme. The recycling of AdoCbl is required to maintain the activity of PduCDE and hence is essential for the degradation of

1,2-PD as a carbon and energy source.

NAD+ recyling within the Pdu MCP

The second part of this study (Chapter 3) investigated the 1-pronanol dehydrogenase

PduQ. This protein was over-expressed, purified, and characterized. PduQ was identified

102

as an iron-dependent alcohol dehydrogenase (ADH) which catalyzes the interconversion between propionaldehyde and 1-propanol. However, the propionaldehyde reduction activity of PduQ was about 45 times higher than that of the 1-propanol oxidation at pH

7.0, indicating that this enzyme is more efficient for catalyzing aldehyde reduction in cells where approximatedly neutral pH is maintained. Moreover, kinetic data exhibited high catalytic efficiency for the propionaldehyde reduction and high affinity for the substrate and cofactor involved in this reaction, which also suggest that the physiological function of PduQ is the reduction of aldehyde with the consumption of NADH to NAD+.

Silmilar to PduS, SDS-PAGE and Western blot of purified Pdu MCP and subsequent

MS-MS analysis demonstrated that the PduQ protein is also associated with the Pdu MCP.

To examine the role of PduQ in vivo, the effects of a pduQ deletion mutation on

AdoCbl-dependent growth on 1,2-PD were tested and the results showed that the pduQ deletion impaired growth and led to a lower cell density, indicating that PduQ was required to support the maximal rate of 1,2-PD degradation by Salmonella. Furthermore, the His-tag pull-down assays demonstrated a specific strong interaction between PduQ and PduP.

All these results support that the MCP lumen enzyme PduQ helps recycle the electron carriers and balance redox reactions by converting a portion of the propionaldehyde to

1-propanol and NADH to NAD+ to avoid depleting NAD+ within the MCP lumen.

A model for the Pdu MCP

In addition, the phosphotransacylase PduL is also associated with the Pdu MCPs, which was confirmed by Western blot. Thus, the Pdu MCPs encapsulate the enzymes needed to regenerate the cofactors required by other enzymes encased within the protein

103

shell although these cofators may also cross the shell. PduGH, PduO, and PduS constitute the AdoCbl recycling system; PduQ (perhaps with PduS) functions in NAD+ recycling; and PduL regenerates CoA; which are able to support the maximal degradation of 1,2-PD by Salmonella (Fig. 4.1).

PduQ NADH

+ OH O NAD O O

ATP PduCDE PduP SCoA SCoA HSCoA Pi ADP PduGH PduL Pi O Cob(III)alamin AdoCbl 2- OPO3 PduS PduO PPPi NADH + ATP NAD Cob(I)alamin O

OPO 2- 3

FIG. 4.1. The Pdu MCPs regenerate enzymatic cofactors internally. The MCP protein shell is indicated by the dashed line. The PduGH, PduO, and PduS constitute the AdoCbl recycling system; PduQ (perhaps with PduS) functions in NAD+ recycling; and PduL regenerates CoA; which are able to support the maxmal degradation of 1,2-PD by Salmonella.

104

References

1. Ailion, M., T. A. Bobik, and J. R. Roth. 1993. Two global regulatory systems (Crp and Arc) control the cobalamin/propanediol regulon of Salmonella typhimurium. J. Bacteriol. 175:7200-7208.

2. Al-Zarban, S., L. Heffernan, J. Nishitani, L. Ransone, and G. Wilcox. 1984. Positive control of the L-rhamnose genetic system in Salmonella typhimurium LT2. J. Bacteriol. 158:603-608.

3. Allen, R. H., S. P. Stabler, D. G. Savage, and J. Lindenbaum. 1993. Metabolic

abnormalities in cobalamin (vitamin B12) and folate deficiency. FASEB J. 7:1344-1353.

4. Alpers, D. H., and G. J. Russel-Jones. 1999. Intrinsic factor, haptocorrin and their receptors,

p. 411-440. In R. Banerjee (ed.), Chemistry and biochemistry of B12. John Wiley & Sons, New York, NY, USA.

5. Andersen, C. B., M. Madsen, T. Storm, S. K. Moestrup, and G. R. Andersen. 2010.

Structural basis for receptor recognition of vitamin-B12-intrinsic factor complexes. Nature 464:445-448.

6. Anderson, R. M., K. J. Bitterman, J. G. Wood, O. Medvedik, H. Cohen, S. S. Lin, J. K. Manchester, J. I. Gordon, and D. A. Sinclair. 2002. Manipulation of a nuclear NAD+ salvage pathway delays aging without altering steady-state NAD+ levels. J. Biol. Chem. 277:18881-18890.

7. Andersson, D. I. 1992. Involvement of the Arc system in redox regulation of the Cob operon in Salmonella typhimurium. Mol. Microbiol. 6:1491-1494.

8. Badger, M. R., D. Hanson, and G. D. Price. 2002. Evolution and diversity of CO2 concentrating mechanisms in cyanobacteria. Funct. Plant Biol. 29:161-173.

9. Badger, M. R., and G. D. Price. 2003. CO2 concentrating mechanisms in cyanobacteria: molecular components, their diversity and evolution. J. Exp. Bot. 54:609-622.

10. Badía, J., J. Ros, and J. Aguilar. 1985. Fermentation mechanism of fucose and rhamnose in Salmonella typhimurium and Klebsiella pneumoniae. J. Bacteriol. 161:435-437.

11. Bairoch, A. 1991. PROSITE: a dictionary of sites and patterns in proteins. Nucleic Acids Res. 19:2241-2245.

12. Baldomà, L., and J. Aguilar. 1988. Metabolism of L-fucose and L-rhamnose in Escherichia coli: aerobic-anaerobic regulation of L-lactaldehyde dissimilation. J. Bacteriol. 170:416-421.

105

13. Banerjee, R. 2006. B12 trafficking in mammals: a case for coenzyme escort service. ACS Chem. Biol. 1:149-159.

14. Banerjee, R. 1999. Chemistry and biochemistry of B12. John Wiley & Sons, New York, NY, USA.

15. Banerjee, R. 2001. Radical peregrinations catalyzed by coenzyme B12-dependent enzymes. Biochemistry 40:6191-6198.

16. Banerjee, R. 1997. The Yin-Yang of cobalamin biochemistry. Chem. Biol. 4:175-186.

17. Banerjee, R., C. Gherasim, and D. Padovani. 2009. The tinker, tailor, soldier in intracellular

B12 trafficking. Curr. Opin. Chem. Biol. 13:484-491.

18. Banerjee, R., and S. W. Ragsdale. 2003. The many faces of vitamin B12: catalysis by cobalamin-dependent enzymes. Annu. Rev. Biochem. 72:209-247.

19. Becher, B., V. Muller, and G. Gottschalk. 1992. N5-methyl-tetrahydromethanopterin:coenzyme M methyltransferase of Methanosarcina strain Go1 is an Na+-translocating membrane protein. J. Bacteriol. 174:7656-7660.

20. Beeby, M., T. A. Bobik, and T. O. Yeates. 2009. Exploiting genomic patterns to discover new supramolecular protein assemblies. Protein Sci. 18:69-79.

21. Begley, T. P., C. Kinsland, R. A. Mehl, A. Osterman, and P. Dorrestein. 2001. The biosynthesis of nicotinamide adenine dinucleotides in bacteria. Vitam. Horm. 61:103-119.

22. Beinert, H. 1983. Semi-micro methods for analysis of labile sulfide and of labile sulfide plus sulfane sulfur in unusually stable iron-sulfur proteins. Anal. Biochem. 131:373-378.

23. Belenky, P., K. L. Bogan, and C. Brenner. 2007. NAD+ metabolism in health and disease. Trends Biochem. Sci. 32:12-19.

24. Berger, F., M. H. Ramírez-Hernández, and M. Ziegler. 2004. The new life of a centenarian: signalling functions of NAD(P). Trends Biochem. Sci. 29:111-118.

25. Berkowitz, D., J. M. Hushon, H. J. Whitfield, Jr., J. Roth, and B. N. Ames. 1968. Procedure for identifying nonsense mutations. J. Bacteriol. 96:215-220.

26. Blanche, F., M. Couder, L. Debussche, D. Thibaut, B. Cameron, and J. Crouzet. 1991.

Biosynthesis of vitamin B12: stepwise amidation of carboxyl groups b, d, e, and g of cobyrinic acid a,c-diamide is catalyzed by one enzyme in Pseudomonas denitrificans. J. Bacteriol. 173:6046-6051.

27. Blander, G., and L. Guarente. 2004. The Sir2 family of protein deacetylases. Annu. Rev. Biochem. 73:417-435.

106

28. Bobik, T. A. 2006. Polyhedral organelles compartmenting bacterial metabolic processes. Appl. Microbiol. Biotechnol. 70:517-525.

29. Bobik, T. A. 2006. Polyhedral organelles compartmenting bacterial metabolic processes. Appl. Microbiol. Biotechnol. 70:517-525.

30. Bobik, T. A., M. Ailion, and J. R. Roth. 1992. A single regulatory gene integrates control of

vitamin B12 synthesis and propanediol degradation. J. Bacteriol. 174:2253-2266.

31. Bobik, T. A., G. D. Havemann, R. J. Busch, D. S. Williams, and H. C. Aldrich. 1999. The propanediol utilization (pdu) operon of Salmonella enterica serovar Typhimurium LT2 includes genes necessary for formation of polyhedral organelles involved in coenzyme

B12-dependent 1,2-propanediol degradation. J. Bacteriol. 181:5967-5975.

32. Bobik, T. A., Y. Xu, R. M. Jeter, K. E. Otto, and J. R. Roth. 1997. Propanediol utilization genes (pdu) of Salmonella typhimurium: three genes for the propanediol dehydratase. J. Bacteriol. 179:6633-6639.

33. Bock, A. K., A. Prieger-Kraft, and P. Schönheit. 1994. Pyruvate – a novel substrate for growth and methane formation in Methanosarcina barkeri. Arch. Microbiol. 161:33-46.

34. Bogan, K. L., and C. Brenner. 2008. Nicotinic acid, nicotinamide, and nicotinamide riboside: a molecular evaluation of NAD+ precursor vitamins in human nutrition. Annu. Rev. Nutr. 28:115-130.

35. Boronat, A., and J. Aguilar. 1981. Metabolism of L-fucose and L-rhamnose in Escherichia coli: differences in induction of propanediol oxidoreductase. J. Bacteriol. 147:181-185.

36. Boronat, A., and J. Aguilar. 1979. Rhamnose-induced propanediol oxidoreductase in Escherichia coli: purification, properties, and comparison with the fucose-induced enzyme. J. Bacteriol. 140:320-326.

37. Bossemeyer, D. 1994. The glycine-rich sequence of protein kinases: a multifunctional element. Trends Biochem. Sci. 19:201-205.

38. Bradbeer, C. 1999. Cobalamin transport in bacteria, p. 489-506. In R. Banerjee (ed.),

Chemistry and biochemistry of B12. John Wiley & Sons, New York, NY, USA.

39. Brady, R. O., E. G. Castanera, and H. A. Barker. 1962. The enzymatic synthesis of cobamide coenzymes. J. Biol. Chem. 237:2325-2332.

40. Brinsmade, S. R., T. Paldon, and J. C. Escalante-semerena. 2005. Minimal functions and physiological conditions required for growth of Salmonella enterica on ethanolamine in the absence of the metabolosome. J. Bacteriol. 187:8039-8046.

41. Broderick, J. B., T. F. Henshaw, J. Cheek, K. Wojtuszewski, S. R. Smith, M. R. Trojan, R.

107

M. McGhan, A. Kopf, M. Kibbey, and W. E. Broderick. 2000. Pyruvate formate-lyase-activating enzyme: strictly anaerobic isolation yields active enzyme containing a [3Fe-4S]+ cluster. Biochem. Biophys. Res. Commun. 269:451-456.

42. Buan, N. R., and J. C. Escalante-semerena. 2006. Purification and initial biochemical characterization of ATP:Cob(I)alamin adenosyltransferase (EutT) enzyme of Salmonella enterica. J. Biol. Chem. 281:16971-16977.

43. Buchrieser, C., C. Rusniok, F. Kunst, P. Cossart, and P. Glaser. 2003. Comparison of the genome sequences of Listeria monocytogenes and Listeria innocua: clues for evolution and pathogenicity. FEMS Immunol. Med. Microbiol. 35:207-213.

44. Burke, S. A., and J. A. Krzycki. 1995. Involvement of the "A" isozyme of methyltransferase II and the 29-kilodalton corrinoid protein in methanogenesis from monomethylamine. J. Bacteriol. 177:4410-4416.

45. Burkle, A. 2005. Poly(ADP-ribose). The most elaborate metabolite of NAD+. FEBS J. 272:4576-4589.

46. Burroughs, A. M., S. Balaji, L. M. Iyer, and L. Aravind. 2007. A novel superfamily containing the β-grasp fold involved in binding diverse soluble ligands. Biol. Direct. 2:4.

47. Cai, F., B. B. Menon, G. C. Cannon, K. J. Curry, J. M. Shively, and S. Heinhorst. 2009. The pentameric vertex proteins are necessary for the icosahedral carboxysome shell to

function as a CO2 leakage barrier. PLoS One 4:e7521.

48. Cannon, G. C., C. E. Bradburne, H. C. Aldrich, S. H. Baker, S. Heinhorst, and J. M. Shively. 2001. Microcompartments in prokaryotes: carboxysomes and related polyhedra. Appl. Environ. Microbiol. 67:5351-5361.

49. Cannon, G. C., and J. M. Shively. 1983. Characterization of a homogenous preparation of carboxysomes from Thiobacillus neapolitanus. Arch. Microbiol. 134:52-59.

50. Chen, P., M. Ailion, T. Bobik, G. Stormo, and J. Roth. 1995. Five promoters integrate control of the cob/pdu regulon in Salmonella typhimurium. J. Bacteriol. 177:5401-5410.

51. Chen, P., D. I. Andersson, and J. R. Roth. 1994. The control region of the pdu/cob regulon in Salmonella typhimurium. J. Bacteriol. 176:5474-5482.

52. Cheng, S., and T. A. Bobik. 2010. Characterization of the PduS cobalamin reductase of Salmonella enterica and its role in the Pdu microcompartment. J. Bacteriol. 192:5071-5080.

53. Cheng, S., Y. Liu, C. S. Crowley, T. O. Yeates, and T. A. Bobik. 2008. Bacterial microcompartments: their properties and paradoxes. Bioessays 30:1084-1095.

54. Christiansen, N., B. K. Ahring, G. Wohlfarth, and G. Diekert. 1998. Purification and

108

characterization of the 3-chloro-4-hydroxy-phenylacetate reductive dehalogenase of Desulfitobacterium hafniense. FEBS Lett. 436:159-162.

55. Cocks, G. T., T. Aguilar, and E. C. Lin. 1974. Evolution of L-1, 2-propanediol catabolism in Escherichia coli by recruitment of enzymes for L-fucose and L-lactate metabolism. J. Bacteriol. 118:83-88.

56. Cohn, E. J., G. R. Minot, G. A. Alles, and W. T. Salter. 1928. The nature of the material in liver effective in pernicious anemia. II. J. Biol. Chem. 77:325-358.

57. Conner, C. P., D. M. Heithoff, S. M. Julio, R. L. Sinsheimer, and M. J. Mahan. 1998. Differential patterns of acquired virulence genes distinguish Salmonella strains. Proc. Natl. Acad. Sci. USA 95:4641-4645.

58. Corda, D., and M. Di Girolamo. 2003. Functional aspects of protein mono-ADP-ribosylation. EMBO J. 22:1953-1858.

59. Cot, S. S., A. K. So, and G. S. Espie. 2008. A multiprotein bicarbonate dehydration complex essential to carboxysome function in cyanobacteria. J. Bacteriol. 190:936-945.

60. Crowley, C. S., D. Cascio, M. R. Sawaya, J. S. Kopstein, T. A. Bobik, and T. O. Yeates. 2010. Structural insights into the mechanisms of transport across the Salmonella enterica Pdu microcompartment shell. J. Biol. Chem. [Epub ahead of print].

61. Crowley, C. S., M. R. Sawaya, T. A. Bobik, and T. O. Yeates. 2008. Structure of the PduU shell protein from the Pdu microcompartment of Salmonella. Structure 16:1324-1332.

62. Crowley, C. S., M. R. Sawaya, T. A. Bobik, and T. O. Yeates. 2008. Structure of the PduU shell protein from the Pdu microcompartment of Salmonella. Structure 16:1324-1332.

63. Daniel, R., R. Boenigk, and G. Gottschalk. 1995. Purification of 1,3-propanediol dehydrogenase from Citrobacter freundii and cloning, sequencing, and overexpression of the corresponding gene in Escherichia coli. J. Bacteriol. 177:2151-2156.

64. Datsenko, K. A., and B. L. Wanner. 2000. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc. Natl. Acad. Sci. USA 97:6640-6645.

65. Datsenko, K. A., and B. L. Wanner. 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.

66. Dobson, C. M., T. Wai, D. Leclerc, H. Kadir, M. Narang, J. P. Lerner-Ellis, T. Hudson, D. S. Rosenblatt, and R. A. Gravel. 2002. Identification of the gene responsible for the cblB

complementation group of vitamin B12-dependent methylmalonic aciduria. Hum. Mol. Genet. 11:3361-3369.

67. Dou, Z., S. Heinhorst, E. B. Williams, C. D. Murin, J. M. Shively, and G. C. Cannon.

109

2008. CO2 fixation kinetics of Halothiobacillus neapolitanus mutant carboxysomes lacking

carbonic anhydrase suggest the shell acts as a diffusional barrier for CO2. J. Biol. Chem. 283:10377-10384.

68. Drews, G., and W. Niklowitz. 1956. Beiträge zur Cytologie der Blaualgen. II. Zentroplasma und granuläre Einschlüsse von Phormidium uncinatum. Arch. Mikrobiol. 24:147-162.

69. Drummond, J. T., and R. G. Matthews. 1993. Cobalamin-dependent and cobalamin-independent methionine synthases in Escherichia coli: two solutions to the same chemical problem. Adv. Exp. Med. Biol. 338:687-692.

70. Duda, J., Z. Pedziwilk, and K. Zodrow. 1967. Studies on the vitamin B12 content of the leguminous plants. Acta Microbiol. Pol. 6:233-238.

71. El Fantroussi, S., H. Naveau, and S. N. Agathos. 1998. Anaerobic dechlorinating bacteria. Biotechnol. Prog. 14:167-188.

72. Escalante-Semerena, J. C., and J. R. Roth. 1987. Regulation of cobalamin biosynthetic operons in Salmonella typhimurium. J. Bacteriol. 196:2251-2258.

73. Evans, J. C., D. P. Huddler, M. T. Hilgers, G. Romanchuk, R. G. Matthews, and M. L. Ludwig. 2004. Structures of the N-terminal modules imply large domain motions during catalysis by methionine synthase. Proc. Natl. Acad. Sci. USA 101:3729-3736.

74. Fan, C., and T. A. Bobik. 2008. The PduX enzyme of Salmonella enterica is an L-threonine

kinase used for coenzyme B12 synthesis. J. Biol. Chem. 283:11322-11329.

75. Fan, C., S. Cheng, Y. Liu, C. M. Escobara, C. S. Crowleyb, R. E. Jeffersonc, T. O. Yeates, and T. A. Bobik. 2010. Short N-terminal sequences package proteins into bacterial microcompartments. Proc. Natl. Acad. Sci. USA 107:7509-7514.

76. Fenton, W. A., R. A. Gravel, and D. S. Rosenblatt. 2001. Disorders of propionate and methylmalonate metabolism, p. 2165-2193. In C. R. Scriver, A. L. Beaudet, D. Valle, and W. S. Sly (ed.), The metabolic and molecular bases of inherited disease. McGraw-Hill, New York, NY, USA.

77. Ferry, J. G. 1995. CO dehydrogenase. Annu. Rev. Microbiol. 49:305-333.

78. Ferry, J. G. 1992. Methane from acetate. J. Bacteriol. 174:5489-5495.

79. Fish, W. W. 1988. Rapid colorimetric micromethod for the quantitation of complexed iron in biological samples. Methods Enzymol. 158:357-364.

80. Folkers, K. 1982. History of vitamin B12: pernicious anemia to crystalline cyanocobalamin, p.

1-15. In D. Dolphin (ed.), Vitamin B12, vol. 1. John Wiley & Sons, New York, NY, USA.

81. Fonseca, M. V., and J. C. Escalante-Semerena. 2001. An in vitro reducing system for the

110

enzymic conversion of cobalamin to adenosylcobalamin. J. Biol. Chem. 276:32101-32108.

82. Fonseca, M. V., and J. C. Escalante-Semerena. 2000. Reduction of Cob(III)alamin to Cob(II)alamin in Salmonella enterica serovar Typhimurium LT2. J. Bacteriol. 182:4304-4309.

83. Foster, J. W., and A. G. Moat. 1980. Nicotinamide adenine dinucleotide biosynthesis and pyridine nucleotide cycle metabolism in microbial systems. Microbiol. Rev. 44:83-105.

84. Froese, D. S., X. Wu, J. Zhang, R. Dumas, W. M. Schoel, M. Amrein, and R. A. Gravel. 2008. Restricted role for methionine synthase reductase defined by subcellular localization. Mol. Genet. Metab. 94:68-77.

85. Fukuzawa, H., E. Suzuki, Y. Komukai, and S. Miyachi. 1992. A gene homologous to chloroplast carbonic anhydrase (icfA) is essential to photosynthetic carbon dioxide fixation by Synechococcus PCC7942. Proc. Natl. Acad. Sci. USA 89:4437-4441.

86. George, D. G., L. T. Hunt, L.-S. L. Yeh, and W. C. Barker. 1985. New perspectives on bacterial ferredoxin evolution. J. Mol. Evol. 22:20-31.

87. Gravel, R. A., M. J. Mahoney, F. H. Ruddle, and L. E. Rosenberg. 1975. Genetic complementation in heterokaryons of human fibroblasts defective in cobalamin metabolism. Proc. Natl. Acad. Sci. USA 72:3181-3185.

88. Guse, A. H. 2004. Biochemistry, biology, and pharmacology of cyclic adenosine diphosphoribose (cADPR). Curr. Med. Chem. 11:847-855.

89. Guse, A. H. 2004. Regulation of calcium signaling by the second messenger cyclic adenosine diphosphoribose (cADPR). Curr. Mol. Med. 4:239-248.

90. Guse, A. H. 2005. Second messenger function and the structure-activity relationship of cyclic adenosine diphosphoribose (cADPR). FEBS J. 272:4590-4597.

91. Hannibal, L., J. Kim, N. E. Brasch, S. Wang, D. S. Rosenblatt, R. Banerjee, and D. W. Jacobsen. 2009. Processing of alkylcobalamins in mammalian cells: A role for the MMACHC (cblC) gene product. Mol. Genet. Metab. 97:260-266.

92. Harden, A., and W. J. Young. 1906. The alcoholic ferment of yeast-juice. Proc. R. Soc. Lond. Ser. B 77:405-420

93. Harms, U., D. S. Weiss, P. Gärtner, D. Linder, and R. K. Thauer. 1995. The energy conserving N5-methyltetrahydromethanopterin:coenzyme M methyltransferase complex from Methanobacterium thermoautotrophicum is composed of eight different subunits. Eur. J. Biochem. 228:640-648.

94. Havemann, G. D., and T. A. Bobik. 2003. Protein content of polyhedral organelles involved

in coenzyme B12-dependent degradation of 1,2-propanediol in Salmonella enterica serovar

111

Typhimurium LT2. J. Bacteriol. 185:5086-5095.

95. Havemann, G. D., E. M. Sampson, and T. A. Bobik. 2002. PduA is a shell protein of

polyhedral organelles involved in coenzyme B12-dependent degradation of 1,2-propanediol in Salmonella enterica serovar typhimurium LT2. J. Bacteriol. 184:1253-1261.

96. Heinhorst, S., G. C. Cannon, and J. M. Shively. 2006. Carboxysomes and carboxysome-like inclusions, p. 141-165. In J. M. Shively (ed.), Complex intracellular structures in prokaryotes. Springer-Verlag, Berlin, Heidelberg, Germany.

97. Heinhorst, S., E. B. Williams, F. Cai, C. D. Murin, J. M. Shively, and G. C. Cannon. 2006. Characterization of the carboxysomal carbonic anhydrase CsoSCA from Halothiobacillus neapolitanus. J. Bacteriol. 188:8087-8094.

98. Heithoff, D. M., C. P. Conner, U. Hentschel, F. Govantes, P. C. Hanna, and M. J. Mahan. 1999. Coordinate intracellular expression of Salmonella genes induced during infection. J. Bacteriol. 181:799-807.

99. Heldt, D., S. Frank, A. Seyedarabi, D. Ladikis, J. B. Parsons, M. J. Warren, and R. W. Pickersgill. 2009. Structure of a trimeric bacterial microcompartment shell protein, EtuB, associated with ethanol utilization in Clostridium kluyveri. Biochem. J. 423:199-207.

100. Henderson, L. M. 1983. Niacin. Annu. Rev. Nutr. 3:289-307.

101. Hendrickson, E. R., J. A. Payne, R. M. Young, M. G. Starr, M. P. Perry, S. Fahnestock, D. E. Ellis, and R. C. Ebersole. 2002. Molecular analysis of Dehalococcoides 16S ribosomal DNA from chloroethene-contaminated sites throughout North America and Europe. Appl. Environ. Microbiol. 68:485-495.

102. Hodgkin, D. C., J. Pickworth, J. H. Robertson, K. N. Trueblood, R. J. Prosen, and J. G.

White. 1955. The crystal structure of the hexacarboxylic acid derived from B12 and the molecular structure of the vitamin. Nature 176:325-328.

103. Holliger, C., G. Wohlfarth, and G. Diekert. 1998. Reductive dechlorination in the energy metabolism of anaerobic bacteria. FEMS Microbiol. Rev. 22:383-398.

104. Holthuijzen, Y. A., J. F. L. van Breemen, J. G. Kuenen, and W. N. Konings. 1986. Protein composition of the carboxysomes of Thiobacillus neapolitanus. Arch. Microbiol. 144:398-404.

105. Horswill, A. R., and J. C. Escalante-Semerena. 1999. Salmonella typhimurium LT2 catabolizes propionate via the 2-methylcitric acid cycle. J. Bacteriol. 181:5615-5623.

106. Iancu, C. V., H. J. Ding, D. M. Morris, D. P. Dias, A. D. Gonzales, A. Martino, and G. J. Jensen. 2007. The structure of isolated Synechococcus strain WH8102 carboxysomes as

112

revealed by electron cryotomography. J. Mol. Biol. 372:764-773.

107. Imai, S., C. M. Armstrong, M. Kaeberlein, and L. Guarente. 2000. Transcriptional silencing and longevity protein Sir2 is an NAD-dependent histone deacetylase. Nature 403:795-800.

108. Iyer, L. M., E. V. Koonin, and L. Aravind. 2003. Evolutionary connection between the catalytic subunits of DNA-dependent RNA polymerases and eukaryotic RNA-dependent RNA polymerases and the origin of RNA polymerases. BMC Struct. Biol. 3:1.

109. Jensen, T. E., and C. C. Bowen. 1961. Organization of the centroplasm in Nostoc pruniforme. Proc. Iowa Acad. Sci. 68:86-89.

110. Jeter, R. M. 1990. Cobalamin-dependent 1,2-propanediol utilization by Salmonella typhimurium. J. Gen. Microbiol. 136:887-896.

111. Johnson, C. L. V., M. L. Buszko, and T. A. Bobik. 2004. Purification and initial characterization of the Salmonella enterica PduO ATP:Cob(I)alamin adenosyltransferase. J. Bacteriol. 186:7881-7887.

112. Johnson, C. L. V., E. Pechonick, S. D. Park, G. D. Havemann, N. A. Leal, and T. A. Bobik. 2001. Functional genomic, biochemical, and genetic characterization of the Salmonella pduO gene, an ATP: cob(I)alamin adenosyltransferase gene. J. Bacteriol. 183:1577-1584.

113. Johnson, C. L. V., E. Pechonick, S. D. Park, G. D. Havemann, N. A. Leal, and T. A. Bobik. 2001. Functional genomic, biochemical, and genetic characterization of the Salmonella pduO gene, an ATP:cob(I)alamin adenosyltransferase gene. J. Bacteriol. 183:1577-1584.

114. Joseph, B., K. Przybilla, C. Stuhler, K. Schauer, J. Slaghuis, T. M. Fuchs, and W. Goebel. 2006. Identification of Listeria monocytogenes genes contributing to intracellular replication by expression profiling and mutant screening. J. Bacteriol. 188:556-568.

115. Katoh, A., and T. Hashimoto. 2004. Molecular biology of pyridine nucleotide and nicotine biosynthesis. Front. Biosci. 9:1577-1586.

116. Katoh, A., K. Uenohara, M. Akita, and T. Hashimoto. 2006. Early steps in the biosynthesis of NAD in Arabidopsis start with aspartate and occur in the plastid. Plant Physiol. 141:851-857.

117. Kerfeld, C. A., S. Heinhorst, and G. C. Cannon. 2010. Bacterial microcompartments. Annu. Rev. Microbiol. 64:391-408.

118. Kerfeld, C. A., M. R. Sawaya, S. Tanaka, C. V. Nguyen, M. Phillips, M. Beeby, and T. O. Yeates. 2005. Protein structures forming the shell of primitive bacterial organelles. Science 309:936-938.

113

119. Kim, J., C. Gherasim, and R. Banerjee. 2008. Decyanation of vitamin B12 by a trafficking chaperone. Proc. Natl. Acad. Sci. USA 105:14551-14554.

120. Klein, M. G., P. Zwart, S. C. Bagby, F. Cai, S. W. Chisholm, S. Heinhorst, G. C. Cannon, and C. A. Kerfeld. 2009. Identification and structural analysis of a novel carboxysome shell protein with implications for metabolite transport. J. Mol. Biol. 392:319-333.

121. Klumpp, J., and T. M. Fuchs. 2007. Identification of novel genes in genomic islands that contribute to Salmonella typhimurium replication in macrophages. Microbiology 153:1207-1220.

122. Kofoid, E., C. Rappleye, I. Stojiljkovic, and J. Roth. 1999. The 17-gene ethanolamine (eut) operon of Salmonella typhimurium encodes five homologues of carboxysome shell proteins. J. Bacteriol. 181:5317-5329.

123. Krasotkina, J., T. Walters, K. A. Maruya, and S. W. Ragsdale. 2001. Characterization of

the B12- and iron-sulfur-containing reductive dehalogenase from Desulfitobacterium chlororespirans. J. Biol. Chem. 276:40991-40997.

124. Krone, U. E., K. Laufer, R. K. Thauer, and H. P. Hogenkamp. 1989. Coenzyme F430 as a possible catalyst for the reductive dehalogenation of chlorinated C1 hydrocarbons in methanogenic bacteria. Biochemistry 28:10061-10065.

125. Kumagai, H., T. Fujiwara, H. Matsubara, and K. Saeki. 1997. Membrane localization, topology, and mutual stabilization of the rnfABC gene products in Rhodobacter capsulatus and implications for a new family of energy-coupling NADH oxidoreductases. Biochemistry 36:5509-5521.

126. Lawhon, S. D., J. G. Frye, M. Suyemoto, S. Porwollik, M. McClelland, and C. Altier. 2003. Global regulation by CsrA in Salmonella typhimurium. Mol. Microbiol. 48:1633-1645.

127. Lawrence, A. D., E. Deery, K. J. McLean, A. W. Munro, R. W. Pickersgill, S. E. J. Rigby, and M. J. Warren. 2008. Identification, characterization, and structure/function analysis of a corrin reductase involved in adenosylcobalamin biosynthesis. J. Biol. Chem. 283:10813-10821.

128. Lawrence, J. G., and J. R. Roth. 1996. Evolution of coenzyme B12 synthesis among enteric bacteria: evidence for loss and reacquisition of a multigene complex. Genetics 142:11-24.

129. Leal, N. A., G. D. Havemann, and T. A. Bobik. 2003. PduP is a coenzyme-a-acylating propionaldehyde dehydrogenase associated with the polyhedral bodies involved in

B12-dependent 1,2-propanediol degradation by Salmonella enterica serovar Typhimurium LT2. Arch. Microbiol. 180:353-361.

130. Leal, N. A., H. Olteanu, R. Banerjee, and T. A. Bobik. 2004. Human ATP:Cob(I)alamin

114

adenosyltransferase and its interaction with methionine synthase reductase. J. Biol. Chem. 279:47536-47542.

131. Leal, N. A., S. D. Park, P. E. Kima, and T. A. Bobik. 2003. Identification of the human and bovine ATP:Cob(I)alamin adenosyltransferase cDNAs based on complementation of a bacterial mutant. J. Biol. Chem. 278:9227-9234.

132. Leartsakulpanich, U., M. L. Antonkine, and J. G. Ferry. 2000. Site-specific mutational analysis of a novel cysteine motif proposed to ligate the 4Fe-4S cluster in the iron-sulfur flavoprotein of the thermophilic methanoarchaeon Methanosarcina thermophila. J. Bacteriol. 182:5309-5316.

133. Ledley, F. D. 1990. Perspectives on methylmalonic acidemia resulting from molecular cloning of methylmalonyl CoA mutase. Bioessays 12:335-340.

134. Lehman, I. R. 1974. DNA ligase: structure, mechanism, and function. Science 186:790-797.

135. Lendvay, J. M., F. E. Löffler, M. Dollhopf, M. R. Aiello, v. Daniels, B. Z. Fathepure, M. Gebhard, R. Heine, R. Helton, J. Shi, R. Krajmalnik-Brown, C. L. Brown, J. Major, C. L. , M. J. Barcelona, E. Petrovskis, R. Hickey, J. M. Tiedje, and P. Adriaens. 2003. Bioreactive barriers: A comparison of bioaugmentation and biostimulation for chlorinated solvent remediation. Environ. Sci. Technol. 37:1422-1431.

136. Lexa, D., and J. M. Saveant. 1983. The electrochemistry of vitamin B12. Acc. Chem. Res. 16:235-243.

137. Lima, W. C., A. M. Varani, and C. F. Menck. 2009. NAD biosynthesis evolution in bacteria: lateral gene transfer of kynurenine pathway in Xanthomonadales and Flavobacteriales. Mol. Biol. Evol. 26:399-406.

138. Lin, E. C. C. 1996. Dissimilatory pathways for sugars, polyols, and carboxylates, p. 307-342. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology, 2nd ed. American Society for Microbiology, Washington, DC, USA.

139. Liu, Y., N. A. Leal, E. M. Sampson, C. L. V. Johnson, G. D. Havemann, and T. A. Bobik.

2007. PduL is an evolutionarily distinct phosphotransacylase involved in B12-dependent 1,2-propanediol degradation by Salmonella enterica serovar typhimurium LT2. J. Bacteriol. 189:1589-1596.

140. Löffler, F. E., J. R. Cole, K. M. Ritalahti, and J. M. Tiedje. 2004. Diversity of dechlorinating bacteria, p. 53-87. In M. M. Häggblom and I. D. Bossert (ed.), Dehalogenation: microbial processes and environmental applications. Kluwer Academic, New York, NY, USA.

115

141. Ma, B., S. J. Pan, M. L. Zupancic, and B. P. Cormack. 2007. Assimilation of NAD+ precursors in Candida glabrata. Mol. Microbiol. 66:14-25.

142. Magnuson, J. K., R. V. Stern, J. M. Gossett, S. H. Zinder, and D. R. Burris. 1998. Reductive dechlorination of tetrachloroethene to ethene by a two-component enzyme pathway. Appl. Environ. Microbiol. 64:1270-1275.

143. Marçal, D., A. T. Rêgo, M. A. Carrondo, and F. J. Enguita. 2009. 1,3-Propanediol dehydrogenase from Klebsiella pneumoniae: decameric quaternary structure and possible subunit cooperativity. J. Bacteriol. 191:1143-1151.

144. Marcus, Y., J. A. Berry, and J. Pierce. 1992. Photosynthesis and photorespiration in a mutant of the cyanobacterium Synechocystis PCC 6803 lacking carboxysomes. Planta 187:511-516.

145. Marsh, E. N., and C. L. Drennan. 2001. Adenosylcobalamin-dependent isomerases: new insights into structure and mechanism. Curr. Opin. Chem. Biol. 5:499-505.

146. Mattevi, A. 2006. A close look at NAD biosynthesis. Nat. Struct. Mol. Biol. 13:563-564.

147. Mera, P. E., and J. C. Escalante-Semerena. 2010. Dihydroflavin-driven adenosylation of 4-coordinate Co(II) corrinoids: are cobalamin reductases enzymes or electron transfer proteins? J. Biol. Chem. 285:2911-2917.

148. Mera, P. E., M. St Maurice, I. Rayment, and J. C. Escalante-Semerena. 2009. Residue Phe112 of the human-type corrinoid adenosyltransferase (PduO) enzyme of Lactobacillus reuteri is critical to the formation of the four-coordinate Co(II) corrinoid substrate and to the activity of the enzyme. Biochemistry 48:3138-3145.

149. Miller, E., G. Wohlfarth, and G. Diekert. 1997. Comparative studies on tetrachloroethene reductive dechlorination mediated by Desulfitobacterium sp. strain PCE-S. Arch. Microbiol. 168:513-519.

150. Miller, E., G. Wohlfarth, and G. Diekert. 1998. Purification and characterization of the tetrachloroethene reductive dehalogenase of strain PCE-S. Arch. Microbiol. 169:497-502.

151. Miller, J. H. 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, NY.

152. Minot, G. R., and W. P. Murphy. 1926. Treatment of pernicious anemia by a special diet. J. Am. Med. Assoc. 87:470-476.

153. Montella, C., L. Bellsolell, R. Pérez-Luque, J. Badía, L. Baldoma, M. Coll, and J. Aguilar. 2005. Crystal structure of an iron-dependent group III dehydrogenase that interconverts L-lactaldehyde and L-1,2-propanediol in Escherichia coli. J. Bacteriol. 187:4957-4966.

116

154. Mori, K., T. Tobimatsu, and T. Toraya. 1997. A protein factor is essential for in situ reactivation of glycerol-inactivated adenosylcobalamin-dependent diol dehydratase. Biosci. Biotechnol. Biochem. 61:1729-1733.

155. Mori, K., and T. Toraya. 1999. Mechanism of reactivation of coenzyme B12-dependent diol dehydratase by a molecular chaperone-like reactivating factor. Biochemistry 38:13170-13178.

156. Mori, K., and T. Toraya. 1999. Mechanism of reactivation of coenzyme B12-dependent diol dehydratase by a molecular chaperone-like reactivating factor. Biochemistry 38:13170-13178.

157. Naidich, M. J., and S. U. Ho. 2005. Case 87: subacute combined degeneration. Radiology 237:101-105.

158. Neidhardt, F. C., J. Ingraham, K. Low, B. Magasanik, M. Schaechter, and H. Umbarger (ed.). 1987. Purines and pyrimidines, vol. 1. American Society for Microbiology, Washington, DC.

159. Neumann, A., G. Wohlfarth, and G. Diekert. 1995. Properties of tetrachloroethene and trichloroethene dehalogenase of Dehalospirillum multivorans. Arch. Microbiol. 163:276-281.

160. Neumann, A., G. Wohlfarth, and G. Diekert. 1996. Purification and characterization of tetrachloroethene reductive dehalogenase from Dehalospirillum multivorans. J. Biol. Chem. 271:16515-16519.

161. Nexø, E. 1998. Cobalamin binding proteins, p. 461-471. In B. Kräutler, D. Arigoni, and B. T.

Golding (ed.), Vitamin B12 and B12 proteins. Wiley-VCH, Weinheim, Germany.

162. Ni, S., J. K. Fredrickson, and L. Xun. 1995. Purification and characterization of a novel 3-chlorobenzoate-reductive dehalogenase from the cytoplasmic membrane of Desulfomonile tiedjei DCB-1. J. Bacteriol. 177:5135-5139.

163. Obradors, N., J. Badia, L. Baldomà, and J. Aguilar. 1988. Anaerobic metabolism of the L-rhamnose fermentation product 1,2-propanediol in Salmonella typhimurium. J. Bacteriol. 170:2159-2162.

164. Palacios, S., V. J. Starai, and J. C. Escalante-semerena. 2003. Propionyl coenzyme A is a common intermediate in the 1,2-propanediol and propionate catabolic pathways needed for expression of the prpBCDE operon during growth of Salmonella enterica on 1,2-propanediol. J. Bacteriol. 185:2802-2810.

165. Park, K., P. E. Mera, J. C. Escalante-Semerena, and T. C. Brunold. 2008. Kinetic and spectroscopic studies of the ATP:corrinoid adenosyltransferase PduO from Lactobacillus reuteri: substrate specificity and insights into the mechanism of Co(II)corrinoid reduction. Biochemistry 47:9007-9015.

117

166. Penfound, T., and J. W. Foster. 1996. Biosynthesis and recycling of NAD, p. 721-730. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology. American Society for Microbiology, Washington, DC, USA.

167. Penrod, J. T., and J. R. Roth. 2006. Conserving a volatile metabolite: a role for carboxysome-like organelles in Salmonella enterica. J. Bacteriol. 188:2865-2874.

168. Pilkington, S. J., J. M. Skehel, R. B. Gennis, and J. E. Walker. 1991. Relationship between mitochondrial NADH-ubiquinone reductase and a bacterial NAD-reducing hydrogenase. Biochemistry 30:2166-2175.

169. Poirot, C. M., S. M. Kengen, E. Valk, J. T. Keltjens, C. v. d. Drift, and G. D. Vogels. 1987. Formation of methylcoenzyme M from formaldehyde by cell-free extracts of Methanobacterium thermoautotrophicum. Evidence for the involvement of a corrinoid-containing methyltransferase. FEMS Microbiol. Lett. 40:7-13.

170. Pollak, N., C. Dolle, and M. Ziegler. 2007. The power to reduce: pyridine nucleotides - small molecules with a multitude of functions. Biochem. J. 402:205-218.

171. Price, G. D., and M. R. Badger. 1989. Expression of human carbonic anhydrase in the

cyanobacterium Synechococcus PCC7942 creates a high CO2-requiring phenotype: evidence

for a central role for carboxysomes in the CO2 concentrating mechanism. Plant Physiol. 91:505-513.

172. Price, G. D., M. R. Badger, F. J. Woodger, and B. M. Long. 2008. Advances in

understanding the cyanobacterial CO2-concentrating-mechanism (CCM): functional components, Ci transporters, diversity, genetic regulation and prospects for engineering into plants. J. Exp. Bot. 59:1441-1461.

173. Price, G. D., S. M. Howitt, K. Harrison, and M. R. Badger. 1993. Analysis of a genomic DNA region from the cyanobacterium Synechococcus sp. strain PCC7942 involved in carboxysome assembly and function. J. Bacteriol. 175:2871-2879.

174. Quadros, E. V., Y. Nakayama, and J. M. Sequeira. 2009. The protein and the gene encoding the receptor for the cellular uptake of transcobalamin-bound cobalamin. Blood 113:186-192.

175. Qureshi, A. A., D. S. Rosenblatt, and B. A. Cooper. 1994. Inherited disorders of cobalamin metabolism. Crit. Rev. Oncol. Hematol. 17:133-151.

176. Ragsdale, S. W. 1991. Enzymology of the acetyl-CoA pathway of CO2 fixation. Crit. Rev. Biochem. Mol. Biol. 26:261-300.

177. Randaccio, L., S. Geremia, N. Demitri, and J. Wuerges. 2010. Vitamin B12: unique

118

metalorganic compounds and the most complex vitamins. Molecules 15:3228-3259.

178. Reid, M. F., and C. A. Fewson. 1994. Molecular characterization of microbial alcohol dehydrogenases. Crit. Rev. Microbiol. 20:13-56.

179. Reidl, J., S. Schlör, A. Kraiß, J. Schmidt-Brauns, G. Kemmer, and E. Soleva. 2000. NADP and NAD utilization in Haemophilus influenzae. Mol. Microbiol. 35:1573-1581.

180. Reinhold, L., X. Zviman, and A. Kaplan. 1989. A quantitative model for carbon fluxes and photosynthesis in cyanobacteria. Plant Physiol. Biochem. 27:945-954.

181. Rickes, E. L., N. G. Brink, F. R. Koniuszy, T. R. Wood, and K. Folkers. 1948. Crystalline

vitamin B12. Science 107:396-397.

182. Robscheit-Robbins, F. S., and G. H. Whipple. 1925. Blood regeneration in severe anemia II. Favorable influence of liver, heart and skeletal muscle in diet. Am. J. Physiol. 72:408-418.

183. Roessner, C. A., P. J. Santander, and A. I. Scott. 2001. Multiple biosynthetic pathways for

vitamin B12: variations on a central theme. Vitam. Horm. 61:267-297.

184. Rondon, M. R., and J. C. Escalante-Semerena. 1992. The poc locus is required for 1,2-propanediol-dependent transcription of the cobalamin biosynthetic (cob) and propanediol utilization (pdu) genes of Salmonella typhimurium. J. Bacteriol. 174:2267-2272.

185. Rondon, M. R., A. R. Horswill, and J. C. Escalante-Semerena. 1995. DNA polymerase I function is required for the utilization of ethanolamine, 1,2-propanediol, and propionate by Salmonella typhimurium LT2. J. Bacteriol. 177:7119-7124.

186. Rondon, M. R., R. Kazmierczak, and J. C. Escalante-Semerena. 1995. Glutathione is required for maximal transcription of the cobalamin biosynthetic and 1,2-propanediol utilization (cob/pdu) regulon and for the catabolism of ethanolamine, 1,2-propanediol, and propionate in Salmonella typhimurium LT2. J. Bacteriol. 177:5434-5439.

187. Rongvaux, A., F. Andris, F. Van Gool, and O. Leo. 2003. Reconstructing eukaryotic NAD metabolism. Bioessays 25:683-690.

188. Roth, J. R., J. G. Lawrence, and T. A. Bobik. 1996. Cobalamin (coenzyme B12): synthesis and biological significance. Annu. Rev. Microbiol. 50:137-181.

189. Roth, J. R., J. G. Lawrence, M. Rubenfield, S. Kieffer-Higgins, and G. M. Church. 1993.

Characterization of the cobalamin (vitamin B12) biosynthetic genes of Salmonella typhimurium. J. Bacteriol. 175:3303-3316.

190. Rothenberg, S. P., E. V. Quadros, and A. Regec. 1999. Transcobalamin II, p. 441-473. In R.

Banerjee (ed.), Chemistry and biochemistry of B12. John Wiley & Sons, New York, NY, USA.

191. Sagermann, M., A. Ohtaki, and K. Nikolakakis. 2009. Crystal structure of the EutL shell

119

protein of the ethanolamine ammonia lyase microcompartment. Proc. Natl. Acad. Sci. USA 106:8883-8887.

192. Sambrook, J., and D. Russell. 2001. Molecular cloning: a laboratory manual, 3rd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, NY.

193. Sampson, E. M., and T. A. Bobik. 2008. Microcompartments for B12-dependent 1,2-propanediol degradation provide protection from DNA and cellular damage by a reactive metabolic intermediate. J. Bacteriol. 190:2966-2971.

194. Sampson, E. M., C. L. V. Johnson, and T. A. Bobik. 2005. Biochemical evidence that the pduS gene encodes a bifunctional cobalamin reductase. Microbiology 151:1169-1177.

195. Sampson, E. M., C. L. V. Johnson, and T. A. Bobik. 2005. Biochemical evidence that the pduS gene encodes a bifunctional cobalamin reductase. Microbiology 151:1169-1177.

196. Satoh, R., M. Himeno, and A. Wadano. 1997. Carboxysomal diffusion resistance to ribulose 1,5-bisphosphate and 3-phosphoglycerate in the cyanobacterium Synechococcus PCC7942. Plant Cell Physiol. 38:769-775.

197. Sazanov, L. A., and P. Hinchliffe. 2006. Structure of the hydrophilic domain of respiratory complex I from Thermus thermophilus. Science 311:1430-1436.

198. Schlenk, F., and H. von Euler. 1936. Cozymase. Naturwissenschaften 24:794-795.

199. Schmehl, M., A. Jahn, A. Meyer zu Vilsendorf, S. Hennecke, B. Masepohl, M. Schuppler, M. Marxer, J. Oelze, and W. Klipp. 1993. Identification of a new class of nitrogen fixation genes in Rhodobacter capsulatus: a putative membrane complex involved in electron transport to nitrogenase. Mol. Gen. Genet. 241:602-615.

200. Schmid, M. F., A. M. Paredes, H. A. Khant, F. Soyer, H. C. Aldrich, W. Chiu, and J. M. Shively. 2006. Structure of Halothiobacillus neapolitanus carboxysomes by cryo-electron tomography. J. Mol. Biol. 364:526-535.

201. Schmieger, H. 1971. A method for detection of phage mutants with altered transducing ability. Mol. Gen. Genet. 110:378-381.

202. Schneider, Z. 1987. The occurrence and distribution of corrinoids, p. 157-223. In Z.

Schneider and A. Stroiński (ed.), Comprehensive B12. Walter de Gruyter, Berlin, Germany.

203. Schneider, Z., and A. Stroiński. 1987. Comprehensive B12. Walter de Gruyter, Berlin, Germany.

204. Schuelke, M., J. Loeffen, E. Mariman, J. Smeitink, and L. van den Heuvel. 1998. Cloning of the human mitochondrial 51 kDa subunit (NDUFV1) reveals a 100% antisense homology of its 3'UTR with the 5'UTR of the gamma-interferon inducible protein (IP-30) precursor: is

120

this a link between mitochondrial myopathy and inflammation? Biochem. Biophys. Res. Commun. 245:599-606.

205. Schumacher, W., C. Holliger, A. J. Zehnder, and W. R. Hagen. 1997. Redox chemistry of cobalamin and iron-sulfur cofactors in the tetrachloroethene reductase of Dehalobacter restrictus. FEBS Lett. 409:421-425.

206. Scott, A. I., and C. A. Roessner. 2002. Biosynthesis of cobalamin (vitamin B12). Biochem. Soc. Trans. 30:613-620.

207. Scrutton, N. S. 1999. Identification of covalent flavoproteins and analysis of the covalent link. Methods Mol. Biol. 131:181-193.

208. Seedorf, H., W. F. Fricke, B. Veith, H. Bruggemann, H. Liesegang, A. Strittmatter, M. Miethke, W. Buckel, J. Hinderberger, F. Li, C. Hagemeier, R. K. Thauer, and G. Gottschalk. 2008. The genome of Clostridium kluyveri, a strict anaerobe with unique metabolic features. Proc. Natl. Acad. Sci. USA 105:2128-2133.

209. Shively, J. M. 1974. Inclusion bodies of prokaryotes. Annu. Rev. Microbiol. 28:167-187.

210. Shively, J. M., F. Ball, D. H. Brown, and R. E. Saunders. 1973. Functional organelles in prokaryotes: polyhedral inclusions (carboxysomes) of Thiobacillus neapolitanus. Science 182:584-586.

211. Shively, J. M., C. E. Bradburne, H. C. Aldrich, T. A. Bobik, J. L. Mehlman, S. Jin, and S. H. Baker. 1998. Sequence homologs of the carboxysomal polypeptide CsoS1 of the thiobacilli are present in cyanobacteria and enteric bacteria that form carboxysomes - polyhedral bodies. Can. J. Bot. 76:906-916.

212. Smith, E. L. 1948. Presence of cobalt in the anti-pernicious anaemia factor. Nature 162:144-145.

213. Sriskanda, V., Z. Kelman, J. Hurwitz, and S. Shuman. 2000. Characterization of an ATP-dependent DNA ligase from the thermophilic archaeon Methanobacterium thermoautotrophicum. Nucleic Acids Res. 28:2221-2228.

214. St Maurice, M., P. Mera, K. Park, T. C. Brunold, J. C. Escalante-Semerena, and I. Rayment. 2008. Structural characterization of a human-type corrinoid adenosyltransferase

confirms that coenzyme B12 is synthesized through a four-coordinate intermediate. Biochemistry 47:5755-5766.

215. St Maurice, M., P. E. Mera, M. P. Taranto, F. Sesma, J. C. Escalante-Semerena, and I. Rayment. 2007. Structural characterization of the active site of the PduO-type ATP:Co(I)rrinoid adenosyltransferase from Lactobacillus reuteri. J. Biol. Chem. 282:2596-2605.

121

216. Stich, T. A., N. R. Buan, and T. C. Brunold. 2004. Spectroscopic and computational studies of Co2+ corrinoids: spectral and electronic properties of the biologically relevant base-on and base-off forms of Co2+ cobalamin. J. Am. Chem. Soc. 126:9735-9749.

217. Stich, T. A., N. R. Buan, J. C. Escalante-Semerena, and T. C. Brunold. 2005. Spectroscopic and computational studies of the ATP:corrinoid adenosyltransferase (CobA) from Salmonella enterica: insights into the mechanism of adenosylcobalamin biosynthesis. J. Am. Chem. Soc. 127:8710-8719.

218. Stich, T. A., M. Yamanishi, R. Banerjee, and T. C. Brunold. 2005. Spectroscopic evidence for the formation of a four-coordinate Co2+ cobalamin species upon binding to the human ATP:cobalamin adenosyltransferase. J. Am. Chem. Soc. 127:7660-7661.

219. Stojiljkovic, I., A. J. Bäumler, and F. Heffron. 1995. Ethanolamine utilization in Salmonella typhimurium: nucleotide sequence, protein expression, and mutational analysis of the cchA cchB eutE eutJ eutG eutH gene cluster. J. Bacteriol. 177:1357-1366.

220. Stupperich, E. 1993. Recent advances in elucidation of biological corrinoid functions. FEMS Microbiol. Rev. 12:349-365.

221. Suh, S.-J., and J. C. Escalante-semerena. 1995. Purification and initial characterization of the ATP:corrinoid adenosyltransferase encoded by the cobA gene of Salmonella typhimurium. J. Bacteriol. 177:921-925.

222. Tanaka, S., C. A. Kerfeld, M. R. Sawaya, F. Cai, S. Heinhorst, G. C. Cannon, and T. O. Yeates. 2008. Atomic-level models of the bacterial carboxysome shell. Science 319:1083-1086.

223. Tanaka, S., M. R. Sawaya, M. Phillips, and T. O. Yeates. 2009. Insights from multiple structures of the shell proteins from the β-carboxysome. Protein Sci. 18:108-120.

224. Tanaka, S., M. R. Sawaya, and T. O. Yeates. 2010. Structure and mechanisms of a protein-based organelle in Escherichia coli. Science 327:81-84.

225. Tartoff, K. D., and C. A. Hobbs. 1987. Improved media for growing plasmid and cosmid clones. Bethesda Res. Lab. Focus 9:12-14.

226. Tartoff, K. D., and C. A. Hobbs. 1987. Improved media for growing plasmid and cosmid clones. Bethesda Res. Lab. Focus 9:12-14.

227. Tomkinson, A. E., and D. S. Levin. 2005. Mammalian DNA ligases. Bioessays 19:893-901.

228. Toraya, T. 2003. Radical catalysis in coenzyme B12-dependent isomerization (eliminating) reactions. Chem. Rev. 103:2095-2128.

229. Tsai, Y., M. R. Sawaya, G. C. Cannon, F. Cai, E. B. Williams, S. Heinhorst, C. A. Kerfeld,

122

and T. O. Yeates. 2007. Structural analysis of CsoS1A and the protein shell of the Halothiobacillus neapolitanus carboxysome. PLoS Biol. 5:e144.

230. Tsai, Y., M. R. Sawaya, and T. O. Yeates. 2009. Analysis of lattice-translocation disorder in the layered hexagonal structure of carboxysome shell protein CsoS1C. Acta Crystallogr. D Biol. Crystallogr. 65:980-988.

231. Ueda, K., and O. Hayaishi. 1985. ADP-ribosylation. Annu. Rev. Biochem. 54:73-100.

232. van de Pas, B. A., J. Gerritse, W. M. de Vos, G. Schraa, and A. J. Stams. 2001. Two distinct enzyme systems are responsible for tetrachloroethene and chlorophenol reductive dehalogenation in Desulfitobacterium strain PCE1. Arch. Microbiol. 176:165-169.

233. Vogel, H. J., and D. M. Bonner. 1956. Acetylornithinase of Escherichia coli: partial purification and some properties. J. Biol. Chem. 218:97-106.

234. Wackett, L. P., J. A. Frias, J. L. Seffernick, D. J. Sukovich, and S. M. Cameron. 2007. Genomic and biochemical studies demonstrating the absence of an alkane-producing phenotype in Vibrio furnissii M1. Appl. Environ. Microbiol. 73:7192-7198.

235. Wang, J., R. S. Houk, D. Dreessen, and D. R. Wiederin. 1998. Identification of inorganic elements in proteins in human serum and in DNA fragments by size exclusion chromatography and inductively coupled plasma mass spectrometry with a magnetic sector mass spectrometer. J. Am. Chem. Soc. 120:5793-5799.

236. Warburg, O., and W. Christian. 1936. Pyridin, der wasserstoffübertragende Bestandteil von Gärungsfermenten. (Pyridin-Nucleotide). Biochem. Z. 187:291-328.

237. Warren, J. W., J. R. Walker, J. R. Roth, and E. Altman. 2000. Construction and characterization of a highly regulable expression vector, pLAC11, and its multipurpose derivatives, pLAC22 and pLAC33. Plasmid 44:138-151.

238. Watanabe, F., Y. Oki, Y. Nakano, and S. Kitaoka. 1988. Occurrence and characterization of cyanocobalamin reductase (NADPH; CN-eliminating) involved in decyanation of cyanocobalamin in Euglena gracilis. J. Nutr. Sci. Vitaminol. (Tokyo) 34:1-10.

239. Watanabe, F., Y. Oki, Y. Nakano, and S. Kitaoka. 1987. Purification and characterization of aquacobalamin reductase (NADPH) from Euglena gracilis. J. Biol. Chem. 262:11514-11518.

240. Wilkinson, A., J. Day, and R. Bowater. 2001. Bacterial DNA ligases. Mol. Microbiol. 40:1241-1248.

241. Willard, H. F., I. S. Mellman, and L. E. Rosenberg. 1978. Genetic complementation among inherited deficiencies of methylmalonyl-CoA mutase activity: evidence for a new class of human cobalamin mutant. Am. J. Hum. Genet. 30:1-13.

123

242. Willie, A., D. E. Edmondson, and M. S. Jorns. 1996. Sarcosine oxidase contains a novel covalently bound FMN. Biochemistry 35:5292-5299.

243. Wood, H. G., S. W. Ragsdale, and E. Pezacka. 1986. The acetyl-CoA pathway: a newly discovered pathway of autotrophic growth. Trends Biochem. Sci. 11:14-18.

244. Yeates, T. O., C. S. Crowley, and S. Tanaka. 2010. Bacterial microcompartment organelles: protein shell structure and evolution. Annu. Rev. Biophys. 39:185-205.

245. Yeates, T. O., C. A. Kerfeld, S. Heinhorst, G. C. Cannon, and J. M. Shively. 2008. Protein-based organelles in bacteria: carboxysomes and related microcompartments. Nat. Rev. Microbiol. 6:681-691.

246. Yeates, T. O., C. A. Kerfeld, S. Heinhorst, G. C. Cannon, and J. M. Shively. 2008. Protein-based organelles in bacteria: carboxysomes and related microcompartments. Nat. Rev. Microbiol. 6:681-691.

247. Yeates, T. O., Y. Tsai, S. Tanaka, M. R. Sawaya, and C. A. Kerfeld. 2007. Self-assembly in the carboxysome: a viral capsid-like protein shell in bacterial cells. Biochem. Soc. Trans. 35:508-511.

248. Zhang, J., C. M. Dobson, X. Wu, J. Lerner-Ellis, D. S. Rosenblatt, and R. A. Gravel. 2006. Impact of cblB mutations on the function of ATP:cob(I)alamin adenosyltransferase in

disorders of vitamin B12 metabolism. Mol. Genet. Metab. 87:315-322.

249. Ziegler, M. 2000. New functions of a long-known molecule. Emerging roles of NAD in cellular signaling. Eur. J. Biochem. 267:1550-1564.

124

Acknowledgements

I would like to express my profound gratitude to my academic advisor Dr. Thomas A.

Bobik for his patient guidance, endless support, and invaluable suggestions throughout my PhD study. I am also much indebted to the rest members of my POS committee: Dr.

Alan DiSpirito, Dr. Basil Nikolau, Dr. Reuben Peters, and Dr. Gregory J. Phillips for their advice during my work and for the time and efforts in reviewing this thesis.

My sincere thanks go to Dr. Tracie A. Bierwagen in the BBMB Department at Iowa

State University for help with gel filtration, and to Megan Mekoli and Travis Witte from

Dr. Robert S. Houk group in the ISU Chemistry Department for aid with ICP-MS analyses. I am grateful to Siquan Luo from the ISU Protein Facility for assistance with

MS-MS analyses, to the ISU DNA Sequencing and Synthesis Facility for assistance with

DNA analyses, to the ISU W. M. Keck Metabolomics Research Laboratory for assistance with the mass spectrometry analyses, and to the ISU Microscopy and Nanoimaging

Facility of the Office of Biotechnology for assistance with the electron microscopy.

I would like to make a special acknowledgement to Dr. Yasuhiro Takahashi from the

Division of Life Science at Saitama University (Japan) for offering the plasmids pRKNMC, pRKISC, and pRKSUF017, which are useful for expressing iron-sulfur proteins.

It is a pleasure to convey thanks to Chenguan Fan and Sharmistha Sinha for the pleasant collaboration with them. I would also like to thank all of the members of the

Bobik group for making the lab an enjoyable working environment.

Words fail me to express my deepest appreciation and love to my parents, my sister and her family for their understanding and constant encouragement, and to my beloved

Xingqi who puts up with me through all of this.