Download Recycling of vitamin B12 and NAD+ within the Pdu

Graduate Theses and Dissertations
Graduate College
2010
Recycling of vitamin B12 and NAD+ within the
Pdu microcompartment of Salmonella enterica
Shouqiang Cheng
Iowa State University
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Recommended Citation
Cheng, Shouqiang, "Recycling of vitamin B12 and NAD+ within the Pdu microcompartment of Salmonella enterica" (2010).
Graduate Theses and Dissertations. Paper 11713.
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Recycling of vitamin B12 and NAD+ within the Pdu
microcompartment of Salmonella enterica
by
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
Copyright © Shouqiang Cheng, 2010. All rights reserved.
ii
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
iv
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
v
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.
1
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
hydroxocobalamin
vitamer
(OH-Cbl),
5'-deoxyadenosylcobalamin
compounds
and
the
including
two
(adenosylcobalamin,
cyanocobalamin
coenzyme
AdoCbl),
and
forms
(CN-Cbl),
of
B12:
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
2
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
C
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
3
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.
4
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)
5
-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,
6
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
7
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
8
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
9
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,
corrinoid-dependent
reductive
MeCbl-dependent
dehalogenases,
which
methyltransferases,
catalyze
and
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.
10
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
FIG.
1.8.
Isomerization
of
(R)-methylmalonyl-CoA
succinyl-CoA
and
succinyl-CoA
catalyzed
by
methylmalonyl-CoA mutase.
In some organisms, an AdoCbl-dependent ribonucleotide reductase (class II) catalyzes
11
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).
12
1
5
2
4
3
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).
13
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
(73) (Fig. 1.12). First, MeCbl is formed by a methyl group transfer from N5-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
14
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.
15
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
16
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).
17
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.
18
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).
19
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
20
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
21
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
22
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
23
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
24
(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).
25
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
26
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).
27
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
28
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
29
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
30
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).
31
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
32
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.
33
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
34
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
35
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,
36
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
37
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,
38
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 pseudo- hexamers). 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
39
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.
40
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.
41
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
42
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.
43
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).
44
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
Co+3 to Co+2 to Co+1 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.
45
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).
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
46
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).
CN-Cbl
reductive
decyanation
?
OH-Cbl
ATP
cob(I)alamin
cob(II)alamin
cob(III)alamin
reduction
PPPi
cob(II)alamin
reduction
adenosylation
AdoCb
l
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
47
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
48
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
E. coli
BE118
BL21(DE3) RIL/pTA925-pduO
C41(DE3)
F– ompT hsdSB (rB- mB-) gal dcm (DE3)
BE1355
C41(DE3)/pET-41a
BE1356
C41(DE3)/pET-41a-pduS
BE1374
C41(DE3)/pET-41a-pduS-His6
®
BacterioMatch
II
Δ(mcrA)183 (mcrCB-hsdSMR-mrr)173 endA1
Two-Hybrid
System
hisB supE44 thi-1 recA1 gyrA96 relA1 lac [F´
Reporter Strain
lacIq HIS3 aadA Kanr]
S. enterica serovar Typhimurium LT2
BE1352
ΔpduS::frt
BE287
LT2/pLAC22
BE1353
ΔpduS::frt /pLAC22
BE1354
ΔpduS::frt /pLAC22-pduS
Source
Lab collection
Lucigen
This study
This study
This study
Stratagene
This study
Lab collection
This study
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).
49
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
50
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
51
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
52
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
decrease at 525 nm and quantified using Δε525 = 4.9 mM-1 cm-1 (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
concomitant absorbance increase at 525 nm (Δε525 = 5.3 mM–1 cm–1) (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
absorbance at 525 nm (Δε525 = 4.8 mM–1cm–1).
Growth studies. Growth studies were performed using a Synergy HT Microplate
reader (BioTek, Winooski, VT) as previously described (139). For growth of strains
53
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
54
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
(28GXGX2G33 where X denotes any amino acid) (37), an FMN-binding motif
(120YX2G[D/E]E125) (125, 197) and two canonical [4Fe-4S] motifs (264CX2CX2CX3C274
and
309
CX2CX2CX4C320)
(86, 132) (Fig. 2.2A, B). PduS also has several conserved
domains. An N-terminal glycine-rich loop (25GX2GXGGAG[F/L]P[A/T]X2K39), 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).
55
A
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
56
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.
57
A
B
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
58
flavin released from PduS-His6 co-migrated with authentic FMN by reverse phase HPLC
following co-injection (Fig. 2.6).
0.6
Absorbance
0.5
0.4
0.3
0.2
0.1
0
300
350
400
450
500
Wavelength (nm)
550
600
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.
59
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
60
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.
61
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
(BE1355) (11.0 ± 3.1 nmol min-1 mg-1). 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.
Purification step
Whole-cell extract
Soluble fraction
Ni-NTA eluate
a
Total
protein
(mg)
220
166
1.6
Cob(III)alamin reductase
Specific activity
Total activity
Purification
fold
(μmol min-1 mg-1)
(μmol min-1)
3.9 ± 0.3
858
1
4.2 ± 0.4
697
1.1
42.3 ± 3.2
67.7
10.8
Cob(II)alamin reductase
Specific activity
Purification
Total activity
fold
(nmol min-1 mg-1)
(nmol min-1)
4.4 ± 0.4
968
1
4.9 ± 0.5
813
1.1
54.5 ± 4.2
87.2
12.4
0.5 mM iodoacetate was used to trap produced cob(I)alamin.
62
63
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
64
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.
65
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
PduS-His6 (R2 = 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.
9; Table 3). The enzyme Vmax was 43.1 ± 0.5 or 46.6 ± 1.6 μmol min–1 mg–1 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
66
still retaining a linear response from the spectrophotometer.
A
B
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.
Reaction
Cob(III)alamin
reductionb
Cob(II)alamin
reductionc
a
Kma
(μM)
10.1 ± 0.7
67.5 ± 8.2
27.5 ± 2.4
72.4 ± 9.5
Vmaxa
(nmol min-1 mg-1)
(43.1 ± 0.5) x 103
(46.6 ± 1.6) x 103
56.8 ± 1.1
64.7 ± 3.6
kcat
(s-1)
35.3 ± 0.4
38.2 ± 1.3
(46.6 ± 0.9) x 10-3
(53.0 ± 3.0) x 10-3
kcat/Km
(μM-1 s-1)
3.5
0.57
1.7 x 10-3
0.7 x 10-3
The values of Km and Vmax were from non-linear regression using GraphPad Prism 5.
b
c
Variable
substrate
NADH
OH-Cbl
NADH
cob(II)alamin
Cob(III)alamin reduction assays were performed with 25 nM anaerobically purified PduS-His6.
Cob(II)alamin reduction assays were performed with 1 μM anaerobically purified PduS-His6 protein
in the presence of 0.5 mM iodoacetate.
67
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.
68
A
B
C
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.
69
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.
Plasmid pair
pBT-LGF2/pTRG-Gal11b
pBT-pduS/pTRG-pduO
pBT-pduO/pTRG-pduS
pBT-pduS/pTRG
pBT/pTRG-pduS
pBT-pduO/pTRG
pBT/pTRG-pduO
a
Number of coloniesa
Non-selective medium
Selective medium (3-AT)
1/100
1/1000
1/10
1/1000
TNTCc
~2000
TNTC
~2000
>2000
255
>2000
152
>2000
296
>2000
129
>2000
410
0
0
>2000
564
1
0
>2000
170
2
0
>2000
400
3
0
Number 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.
b
c
LGF2 and Gal11 are a positive control known to strongly interact.
TNTC: Too numerous to count.
70
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
71
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
72
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
73
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
74
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
75
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
76
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
77
ISU W.M. Keck Metabolomics Research Laboratory for assistance with the mass
spectrometry analyses.
78
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
79
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
80
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).
81
TABLE 3.1. Bacterial strains used in this study.
Species and strain
Genotype
E. coli
BE237
BL21(DE3) RIL/pET-41a
BE272
BL21(DE3) RIL/pTA925-pduP
BE1421
BL21(DE3) RIL/pET-41a-His6-pduP
BE1422
BL21(DE3) RIL/pET-41a-pduP-His6
BE1500
BL21(DE3) RIL/pET-41a-pduQ
BE1501
BL21(DE3) RIL/pET-41a-His6-pduQ
BE1502
BL21(DE3) RIL/pET-41a-pduQ-His6
S. enterica serovar Typhimurium LT2
BE191
ΔpduP
BE269
ΔpduP/pLAC22
BE270
ΔpduP/pLAC22-pduP
BE287
LT2/pLAC22
BE903
ΔpduQ::frt
BE919
ΔpduQ::frt/pLAC22
BE942
ΔpduQ::frt/pLAC22-pduQ
Source
Lab collection
Lab collection
Lab collection
Lab collection
This work
This work
This work
Lab collection
Lab collection
Lab collection
Lab collection
This work
This work
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.
82
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
83
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
using Δε340 = 6.22 mM-1 cm-1.
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
84
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%
85
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
NAD-binding motif (89GGG91) and a motif (241GX2HX2AHX2GX5PHG259 where X
86
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.
87
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)
(0.12 ± 0.03 nmol min-1 mg-1). 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.
Purification step
Whole-cell extract
Soluble fraction
Ni-NTA eluate
ab
c
Total
protein
(mg)
403.6
246.4
4.4
Propionaldehyde reductasea
Specific activity
Total activity
Purification
(μmol min-1 mg-1)
(μmol min-1)
fold
c
2.8 ± 0.2
1130
1
3.1 ± 0.3c
763.8
1.1
55.5 ± 4.2
244.2
19.8
Propanol dehydrogenaseb
Specific activity
Total activity
(μmol min-1 mg-1)
(μmol min-1)
0.8 ± 0.1c
322.9
0.9 ± 0.2c
221.8
5.1 ± 0.7
22.4
Purification
fold
1
1.1
6.4
Assayes were performed with 100 mM Tris-HCl at pH 7.0 and pH 9.0, respectively.
The values of the background activities were not subtracted.
88
89
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.
90
Relative acitivity (%)
100
80
60
40
propionaldehyde reduction
1-propanol oxidation
20
0
5
6
7
8
9
10
11
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
91
oxidation with NADP+ compared to NAD+, showing that PduQ has a preference for
NAD(H).
4
Intensity (cps)
x10
25
Fe
20
15
10
5
0
55.0
55.5
56.0
m/z
56.5
57.0
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
92
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
dehydrogenase was linear from 0.16 to 1.4 μM PduQ-His6 (R2 = 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,
respectively (Table 3.3). The enzyme Vmax was 77.7 ± 3.0 or 81.3 ± 4.5 μmol min–1 mg–1
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
enzyme Vmax was 9.2 ± 0.7 μmol min–1 mg–1 or 8.4 ± 0.8 μmol min–1 mg–1 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.
a
Reaction
Variable substrate
Propionaldehy
de reductionb
1-propanol
oxidation c
NADH
propionaldehyde
NAD+
1-propanol
Kma
(μM)
45.3 ± 5.3
(16.0 ± 2.0 ) x 103
267.5 ± 22.3
(95.8 ± 9.2 ) x 103
Vmaxa
(µmol min-1 mg-1)
77.7 ± 3.0
81.3 ± 4.5
9.2 ± 0.7
8.4 ± 0.8
kcat
(s-1)
52.2
54.6
6.18
5.64
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.
kcat/Km
(μM-1 s-1)
1.15
3.41 x 10-3
2.31 x 10-2
5.89 x 10-5
93
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.
94
OD600
A
B
0.9
0.8
0.7
0.6
0.5
0.4
0.3
0.2
0.1
0.0
LT2
ΔpduQ
0
5
10 15 20 25 30 35 40
Time (h)
0.35
0.30
OD600
0.25
0.20
0.15
LT2
ΔpduQ
0.10
0.05
0.00
OD600
C
0.9
0.8
0.7
0.6
0.5
0.4
0.3
0.2
0.1
0.0
0
5
10 15 20 25 30 35 40 45
Time (h)
LT2/pLAC22
ΔpduQ/pLAC22
ΔpduQ/pLAC22-pduQ
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).
95
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 Nor 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.
96
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
97
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
98
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
99
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).
OH
OH
PduQ
+
O NAD
OH
ATP
NADH
PduP
PduCDE
PduL
PduGH
SCoA
SCoA
HSCoA
ADP
Pi
O
O
Pi
O
Cob(III)alamin
PduS
AdoCbl
PduO
NADH
PPPi
OPO32-
NAD+ Cob(I)alamin ATP
O
OPO32-
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
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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.