A proteomic view of the facultatively chemolithoautotrophic lifestyle of Ralstonia eutropha H16

RESEARCH ARTICLE

A proteomic view of the facultatively

chemolithoautotrophic lifestyle of Ralstonia

eutropha H16

Edward Schwartz1, Birgit Voigt2, Daniela Z .uhlke2, Anne Pohlmann1, Oliver Lenz1,Dirk Albrecht2, Alexander Schwarze1, Yvonne Kohlmann1, Cornelia Krause1,Michael Hecker2 and B .arbel Friedrich1

1 Institut f .ur Biologie, Mikrobiologie, Humboldt-Universit .at zu Berlin, Berlin, Germany2 Institut f .ur Mikrobiologie, Ernst-Moritz-Arndt-Universit .at Greifswald, Greifswald, Germany

Received: May 20, 2009

Revised: August 21, 2009

Accepted: August 26, 2009

Ralstonia eutropha H16 is an H2-oxidizing, facultative chemolithoautotroph. Using 2-DE in

conjunction with peptide mass spectrometry we have cataloged the soluble proteins of this

bacterium during growth on different substrates: (i) H2 and CO2, (ii) succinate and (iii) glycerol.

The first and second conditions represent purely lithoautotrophic and purely organoheterotrophic

nutrition, respectively. The third growth regime permits formation of the H2-oxidizing and CO2-

fixing systems concomitant to utilization of an organic substrate, thus enabling mixotrophic

growth. The latter type of nutrition is probably the relevant one with respect to the situation faced

by the organism in its natural habitats, i.e. soil and mud. Aside from the hydrogenase and Calvin-

cycle enzymes, the protein inventories of the H2-CO2- and succinate-grown cells did not reveal

major qualitative differences. The protein complement of the glycerol-grown cells resembled that

of the lithoautotrophic cells. Phosphoenolpyruvate (PEP) carboxykinase was present under all

three growth conditions, whereas PEP carboxylase was not detectable, supporting earlier findings

that PEP carboxykinase is alone responsible for the anaplerotic production of oxaloacetate from

PEP. The elevated levels of oxidative stress proteins in the glycerol-grown cells point to a

significant challenge by ROS under these conditions. The results reported here are in agreement

with earlier physiological and enzymological studies indicating that R. eutropha H16 has a

heterotrophic core metabolism onto which the functions of lithoautotrophy have been grafted.

Keywords:

Autotrophy / Hydrogenase / Lithotrophy / Microbiology / Oxidative stress /

Phosphoenolpyruvate-pyruvate-oxaloacetate node

1 Introduction

Ralstonia eutropha H16 is a strictly respiratory, facultatively

chemolithoautotrophic representative of the b-proteo-

bacteria. The soil- and mud-dwelling organism grows well

on simple organic acids and a few sugars. In the absence of

organic substrates, it thrives on H2 as sole energy source,

fixing CO2 via the Calvin–Benson–Bassham cycle. With

regard to these alternative growth modes, R. eutropha H16 is

a typical representative of the aerobic H2 oxidizers [1, 2].

Recently, the deciphering of the entire 7 416 678-bp genome

of R. eutropha H16 was completed [3, 4]. The genome

consists of three circular replicons: chromosome 1

(4.05 Mbp), chromosome 2 (2.91 Mbp) and the megaplasmid

pHG1 (0.45 Mbp). A total of 6626 coding sequences were

predicted. A disproportionately large fraction of the total

coding capacity is devoted to transport systems – another

indication that the organism in its natural habitat is

confronted with and can grow on a wide variety of

substrates.

Abbreviations: FBA, fructose-1,6-bisphosphate aldolase; FBP,

fructose-1,6-bisphosphatase; GAPDH, glyceraldehyde 3-phos-

phate dehydrogenase; GN, glycerol minimal medium; MBH,

membrane-bound hydrogenase; PEP, phosphoenolpyruvate;

PGK, phosphoglycerate kinase; PK, pyruvate kinase; RH, regula-

tory hydrogenase; SH, soluble hydrogenase; SN, succinate

minimal medium; TCA, tricarboxylic acid

Correspondence: Dr. Edward Schwartz, Institut f .ur Biologie,

Mikrobiologie, Humboldt-Universit .at zu Berlin, Chausseestr.

117, 10115 Berlin, Germany

E-mail: [email protected]

Fax: 149-3020938102

& 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com

5132 Proteomics 2009, 9, 5132–5142DOI 10.1002/pmic.200900333

Oxidation of H2 in R. eutropha H16 is catalyzed by two

NiFe hydrogenases. One of them, a dimeric membrane-

bound hydrogenase (MBH), is the first component of an

H2-dependent electron transport chain which conserves

energy during lithotrophic growth. The MBH is coupled –

both physically and electronically – to the respiratory

chain by a specialized b-type cytochrome. The hydrogenase

dimer is located on the periplasmic side of the cytoplasmic

membrane. The second energy-conserving hydrogenase is a

hexameric protein located in the cytoplasm. This

soluble hydrogenase (SH) couples H2 oxidation to the

reduction of NAD1. It consists of a dimeric hydrogenase

module, a dimeric diaphorase module and a homodimer of

a 19-kD protein responsible for the NADPH-dependent

reductive activation of the enzyme [5]. The SH supplies

the organism with reductant for, e.g. fixation of CO2, giving

R. eutropha a great advantage over lithotrophs that synthe-

size NADH via reverse electron flow. A remarkable aspect of

the hydrogenases is the complex pathway of post-transla-

tional maturation steps required for the assembly and

insertion of the bimetallic center at the active site of each

enzyme.

The MBH and SH are encoded along with accessory

proteins in two large operons on the 450-kb megaplasmid

pHG1. The expression of the hydrogenase genes is

co-ordinately controlled by an H2-dependent signal trans-

duction chain. The H2-sensing component is itself an

hydrogenase-like dimer called a regulatory hydrogenase

(RH). When H2 becomes available, the RH transmits a

signal, the nature of which is presently not well understood,

to a histidine kinase, which in turn interacts with an NtrC-

like response regulator, causing the latter to activate tran-

scription from the hydrogenase promoters [6–8].

Early enzymological studies revealed that the production

of hydrogenases in R. eutropha H16 is not strictly H2-

dependent [9]. Hydrogenase expression is tightly repressed

on succinate (SN) and pyruvate. However, during growth on

other carbon/energy sources hydrogenase expression is

derepressed to varying degrees. The degree of derepression

mediated by the different substrates is more or less corre-

lated with the growth rates they support. Thus, hydrogenase

expression is susceptible to a form of catabolite repression.

During lithoautotrophic growth, CO2 is fixed by ribulose-

1,5-bisphosphate carboxylase/oxygenase (RuBisCO). The

RuBisCO is encoded along with phosphoribulokinase and

the other Calvin-cycle enzymes in a 12-gene operon present

in two copies, one located on the megaplasmid pHG1, the

other on chromosome 2 [10, 11].

The expression patterns of the key components of litho-

trophic metabolism in R. eutropha H16, the hydrogenases

and the enzymes of the Calvin–Benson–Bassham cycle,

have been the subject of detailed investigations in the past

years, which have provided us with a basic understanding of

the underlying regulatory mechanisms [6, 9, 11–13]. Hence,

this is not the main interest of this study. Rather, our aim

was to obtain information about the global metabolic char-

acteristics correlated with the typical growth modes of

R. eutropha H16. Specifically, we cultivated R. eutropha H16

under three different regimes: lithoautotrophically on H2

and CO2, heterotrophically on glycerol (a substrate that

supports only slow growth and leads to high-level expression

of hydrogenase and Calvin-cycle enzymes) and SN (tight

repression of these enzymes). Using 2-D gels we cataloged

the proteins present in cells grown under these three

conditions.

This study was also conceived with more general ques-

tions regarding the bases of lithoautotrophic metabolism

and the underlying features of facultative and obligate

lithoautotrophy in mind. During the 1960s and 1970s, these

questions prompted numerous studies and were the subject

of vigorous debate among microbiologists [14, 15]. With the

advent of molecular genetic techniques, attention shifted to

the characterization of the specialized enzymes and proteins

involved in the various forms of lithoautotrophy. Although

great progress has been made in this area, the adaptations of

core metabolism to lithoautotrophic lifestyles remain to be

characterized [16]. The holistic approaches of the genomic

era provide tools for such studies that were unimaginable 20

years ago. With the availability of genomic sequences of

both facultative and obligate lithoautotrophs, interest in

their general metabolism is reawakening [17, 18]. This study

is a step toward understanding one such organism in the

hope that it will contribute to our knowledge of lithoauto-

trophy in general.

2 Materials and methods

2.1 Strain and culture conditions

R. eutropha H16 (DSM428, ATCC 17699) was cultivated at

301C in mineral salts medium (100 mL of culture in 500-mL

baffled flasks with shaking at 120 rpm) [19]. Lithoautotrophic

cultures were grown under an atmosphere of hydrogen,

oxygen and carbon dioxide at a ratio of 8:1:1 v/v/v. For

heterotrophic experiments the medium was supplemented

with either 0.4% w/v succinate (succinate minimal medium

(SN)) or 0.4% glycerol (glycerol minimal medium (GN)) as

carbon source. Growth was monitored by measuring

turbidity at 436 nm (Supporting Information Fig. S1).

2.2 Preparation of soluble proteins

Cells were harvested during the exponential growth phase at

OD436 5 1 by centrifugation (6000� g, 41C, 15 min), washed

twice with TEP buffer (10 mM Tris, 1 mM EDTA, pH 7.5,

0.3 mg/mL PMSF), resuspended in 0.5 mL TEP buffer, and

disrupted by four passages through a French pressure cell

(SLM, Rochester, NY, USA) at 900 psi. Crude protein

extracts were separated from cellular debris by ultra-

centrifugation (90 000� g, 41C, 30 min). The resulting

Proteomics 2009, 9, 5132–5142 5133


supernatant consisting of the soluble protein fraction was

stored at �801C until use.

2.3 2-DE, imaging and protein identification

The protein content of the soluble fraction was determined

using the RotiNanoquant Kit (Roth). In total, 500mg protein

extract were separated by 2-DE as described previosly [20]. The

IPG reswelling solution for R. eutropha H16 protein extracts

contained 2% w/v CHAPS, 8 M urea, 0,5% v/v Pharmalyte

3–10 and 13 mM DTT. Isoelectric focusing was performed

using non-linear IPGs in the pH range 3–10 (Amersham

Biosciences). 2-D gels were fixed and stained with colloidal

Coomassie Brilliant Blue G-250 (Amersham Biosciences). For

the processing of gel images and gel-based relative quantita-

tion, Decodon Delta 2D software was used. Protein identifi-

cation was performed as described by Voigt et al. [21].

Quantitative protein data presented here are the arithmetic

means of three sets of data obtained from two biological

replicates. For the quantitative comparison of expression

patterns for the three growth conditions investigated, ratios of

mean normalized spot volumes were calculated.

2.4 RT-PCR

Total RNA from lithoautotrophically and heterotrophically

grown cells was isolated with the ribopure bacteria

system (Applied Biosystems, Darmstadt) after quenching

the cells with 50% �801C methanol and stabilizing the

RNA with RNAprotect (Qiagen, Hilden). The integrity

of the RNA was checked using RNA 6000 Nano assay

chips on a Bioanalyzer 2100 (Agilent). In total, 2mg of total

RNA were reverse transcribed with the high-capacity RNA-

to-cDNA kit (Applied Biosystems). Diluted cDNA samples

were used as templates in Real-time qPCR analysis

using specific primer pairs and SYBR Green fluorescent

dye. Real-time PCR was performed using PowerSYBR

Green PCR Mastermix on a 7500 Fast PCR Cycler (Applied

Biosystems). Uniformity of the product was checked

for every PCR by the determination of a dissociation

curve. Pairs of primers with lengths of 19–21 nucleotides

were optimized for use at an annealing temperature of

58–601C. Each primer pair amplified a fragment of

150–200 bp. All primer pairs showed the same efficiency

(100710%) in a control qPCR experiment with serially

diluted cDNA templates. Relative expression ratios were

determined by the ddCt method using gyrB as a constitutive

control. Primers used for qPCR analysis were ppc 301

(50-TGTTCAACCGCATCAGCAA-30), ppc 302 (50-CAGCAA

CTCAACCTGCAAGTG-30), pepck 311 (50-CAACGCCATTC

CAGTTCAAGT-30), pepck 312 (50-CCGAGGATTTACTGCG

TCAAC-30), ppsA 321 (50-GGTGGACGCCGATGTTGT-30),

ppsA 322 (50-GGAAACCGAAGTGACCGAAGT-30), pyk1

331 (50-CAGCTCGACACCGATTTCCT-30), pyk1 332 (50-GG

TGACGGCTCATCCACAAT-30), gyrB 151 (50-GCCTGCAC-

CACCTTGTCTTC-30) and gyrB 152 (50-TGTGGAGGGAC

CTGACT-30).

2.5 Generation of an unmarked knockout mutation

A mutant of R. eutropha H16 with an in-frame lesion in the

prkA gene was obtained using a gene replacement protocol as

described previously [22]. Briefly, a segment of genomic DNA

encompassing prkA was amplified by polymerase chain

reaction using the primer pair 50-CGGACAGTACCGGTCG-

CCTGTTCATTTTCTCG-30 and 50-ACCGCGCGCTTGGCCA-

GGGCGGTGGCGGGACGACG-30. The purified amplicon

was treated with AgeI and inserted into vector plasmid

Litmus28 (NEB). The resulting plasmid was cut with BglII to

excise a 930-bp internal fragment. The prkA deletion allele

was transferred to suicide vector pLO2 [22]. The resulting

mobilizable, deletogenic plasmid was introduced into

R. eutropha H16 by conjugative transfer from the donor strain

Escherichia coli S17-1. Sucrose-resistant transconjugants

were screened by PCR for the desired deletions. This yielded

the R. eutropha derivative HFJ74 ( 5 H16 prkAD).

3 Results and discussion

3.1 General features and statistics

A total of 292 proteins were identified during 2-D gel-based

proteome analysis of three different growth modes

(Supporting Information Tables S1–S3). The protein profiles

of the three growth modes are summarized in a multi-color

protein map (Fig. 1). The false-color map differentiates

between (i) proteins up-regulated in cells growing on one of

the substrates, (ii) proteins identified in cells growing on two

substrates and (iii) constitutively expressed proteins present

under all three growth conditions. Surprisingly, 81% of all

proteins identified in the present study are encoded on

chromosome 1 versus 13% for chromosome 2. This is

remarkable since the latter represents nearly 40% of the total

coding capacity (Table 1). The underrepresentation of

proteins derived from chromosome 2 is, however, in accor-

dance with the finding, that chromosome 1 encodes key

functions of general metabolism involved in e.g. DNA repli-

cation, transcription and translation. Chromosome 2, on the

other hand, harbors genes for alternative metabolic pathways

including various pathways for the decomposition of

aromatic compounds and the utilization of alternative nitro-

gen sources. The majority of these genes are probably not

expressed under the growth conditions tested in our study.

The proteins identified in our 2-D gels were classified

according to function. The major functional groups represent

amino acid biosynthesis (19% of all identified proteins),

carbohydrate metabolism (12%) and energy metabolism

(11%) (Supporting Information Fig. S2).

5134 E. Schwartz et al. Proteomics 2009, 9, 5132–5142


3.2 Key enzymes of lithoautotrophic growth

Proteins belonging to both of the energy-conserving hydro-

genases were identified in 2-D gels of lithoautotrophically

(H2-CO2-) grown cells of R. eutropha H16 (Table 2). All five

subunits of the SH were found in the protein complement

of the H2-CO2-grown cells. The subunits HoxF, HoxU,

HoxY and HoxH were up-regulated by the factors of 9.4, 2.4,

2.6 and 11.3, respectively, compared with SN-grown cells.

The previous physiological studies on R. eutropha H16

grown in SN and H2-CO2 cultures showed an approx.

200-fold induction of both MBH and SH activities under the

latter conditions [9]. These values reflect similarly strong

induction of promoter activity of the PMBH and PSH

promoters and corresponding increases in the abundance of

the MBH and SH transcripts [12].

The fifth subunit of the SH, the HoxI protein, was

identified early on as a dominant species in SDS-PAGE gels

of lithoautotrophically grown cells of R. eutropha H16.

Under these conditions, HoxI (formerly designated

‘‘B-protein’’) represents 4% of the total protein content [23].

In this study we identified HoxI as a dominant protein

species in 2-D gels of H2-CO2-grown cells and found it to be

91-fold up-regulated with respect to SN-grown cells. The

physiological role of the highly abundant HoxI protein was

for a long-time enigmatic. Recently, it was shown that HoxI

is a subunit of the SH (two HoxI monomers/SH holo-

enyzme) and is responsible for the reductive activation of

the enzyme by NADPH [5].

Proteins of the MBH are not necessarily to be expected

in the soluble fraction. Nevertheless, one of the two

subunits of the MBH, the [Ni-Fe] center-containing HoxG,

H2-CO2 H2-CO2 + H2-CO2 + constitutive(H CO +

Chr1Glycerol

Glycerol

Succinate

Glycerol Succinate

Glycerol + Succinate

(H2-CO2 +Glycerol + Succinate)

Chr2pHG1

H2-CO2 Succinate

Figure 1. False-color map of the

proteins detected in soluble

extracts of R. eutropha H16

cells grown under three differ-

ent conditions. In the first

dimension proteins were sepa-

rated in a pH gradient in the

range 3–10. The color coding

used for the protein spots is

defined in the lower part of the

figure. Selected proteins are

labelled. The color of the label

indicates the location of the

corresponding gene in the

genome.

Table 1. Statistics for the R. eutropha H16 genome

Feature Chromosome 1 Chromosome 2 Megaplasmid pHG1 Total

Size (bp) 4 052 032 2 912 490 452 156 7 416 678Coding capacity coding sequence 3651 (55%) 2555 (39%) 420 (6%) 6626 (100%)Proteins identified 236 (81%) 37 (13%) 19 (6%) 292 (100%)

Proteomics 2009, 9, 5132–5142 5135


was present and sixfold up-regulated compared with SN-

grown cells. The small subunit (HoxK), which is anchored

in the membrane via an hydrophobic C-terminal segment,

was not detected.

In addition to the subunits of the energy-conserving

hydrogenases, the products of three other hydrogenase-

related genes were also identified in H2-CO2-grown cells.

These were the large subunit (HoxC) of the RH, the

hydrogenase accessory protein HypB2 and a maturation

protease (HoxW). The formation of active hydrogenases

requires a complex maturation apparatus composed of

multiple proteins. Since the latter proteins catalyze the

formation of active-site metallocomplexes in the maturing

hydrogenases, they are probably much less abundant than

their substrates, the hydrogenases.

Not surprisingly, hydrogenase proteins were also identi-

fied in 2-D gels of the glycerol-grown cells of R. eutrophaH16. This is in accord with the previous findings demon-

strating a 4100-fold induction of hydrogenases in glycerol

cultures [9]. The SH subunits HoxF, HoxU, HoxY, HoxH

and HoxI and the MBH protein HoxG were up-regulated by

the factors of 65.1, 6.2, 12.5, 22.3, 288.5 and 13.5, respec-

tively, relative to SN cultures.

In the absence of organic growth substrates, R. eutrophaH16 can fix CO2 by means of a RuBisCO enzyme and the

other enzymes of the Calvin–Benson–Bassham cycle [10].

Both the chromosomal and megaplasmid cbb operons

(designated cbbc and cbbp, respectively) are actively tran-

scribed during growth on H2 [11]. Like the hydrogen-

oxidizing system, the Calvin-cycle enzymes are also expres-

sed under conditions of energy limitation, e.g. during

growth on glycerol as carbon and energy source. We iden-

tified and quantified 11 Cbb proteins in our 2-D gels: CbbEp,

CbbEc, CbbFp, CbbFc, CbbPp, CbbGp, CbbGc, CbbKp,

CbbKc, CbbAp and CbbAc (Table 3 and Supporting Infor-

mation Table S1). The dual R. eutropha H16 cbb operons are

coordinately controlled by CbbR, a LysR-type activator

protein. The absence of CbbR from our 2-D gels is not

surprising, since the previous studies using reporter gene

fusions to monitor promoter activity showed that tran-

scription from the chromosomal cbbR promoter is weak and

constitutive, pointing to low levels of CbbR in the cell [11].

For all but two proteins, the induction ratio H2-CO2 versusSN is higher than glycerol versus SN. We note that this is the

opposite of the trend observed for the hydrogenase-related

proteins.

Table 2. Hydrogenase-related proteins identified in 2-D gels

Gene Locus tag Product Ratio H2-CO2/SNa) Ratio GNb)/SN

hoxG PHG002 Membrane-bound [NiFe] hydrogenase, large subunit 6.0 13.5hoxC PHG021 Regulatory [NiFe] hydrogenase, large subunit 4.0 51.0hoxF PHG088 NAD-reducing hydrogenase diaphorase moiety, large subunit 9.4 65.1hoxU PHG089 NAD-reducing hydrogenase diaphorase moiety, small subunit 2.4 6.2hoxY PHG090 NAD-reducing hydrogenase, H2ase moiety, small subunit 2.6 12.5hoxH PHG091 NAD-reducing hydrogenase, H2ase moiety, large subunit 11.3 22.3hoxW PHG092 [NiFe] hydrogenase C-terminal protease (HoxH-specific) 6.2 15.8hoxI PHG093 NAD-reducing hydrogenase subunit 91.2 288.5hypB2 PHG095 [NiFe] hydrogenase maturation protein 2.6 5.5

a) Minimal medium containing succinate.b) Minimal medium containing glycerol.

Table 3. Proteins of the Calvin cycle identified in 2-D gels


cbbEp PHG423 Ribulose-phosphate 3-epimerase 0.7 1.4cbbEc H16_B1391 Ribulose-phosphate 3-epimerase 1.8 1.1cbbFp PHG422 FBP 1.0 1.1cbbFc H16_B1390 FBP 4.3 0.5cbbPp PHG421 Phosphoribulokinase 26.7 16.0cbbGp PHG418 GAPDH 44.1 9.5cbbGc H16_B1386 GAPDH 145.9 8.5cbbKp PHG417 PGK 13.9 3.2cbbKc H16_B1385 PGK 7.7 2.9cbbAp PHG416 FBA 4.4 0.8cbbAc H16_B1384 FBA 17.3 1.6




3.3 Central metabolic pathways

The comparison of protein complements of cells grown

under lithoautotrophic and organoheterotrophic conditions

showed surprisingly little difference in central pathways.

Among the enzymes of these pathways present in our 2-D

gels, we found phosphoenolpyruvate (PEP) carboxykinase

(H16_A3711), PEP synthetase (H16_A2038) and pyruvate

kinase (PK) (H16_A0567) under all three growth conditions.

These proteins belong to a network of enzymes inter-

connecting the metabolites PEP, oxaloacetate, pyruvate and

malate, which is known as the ‘‘PEP-pyruvate-oxaloacetate

node’’ and is a major switchboard controlling the distribu-

tion of carbon among different metabolic pathways (Fig. 2)

[24]. Two other enzymes of the network responsible for

interconverting C3 and C4 intermediates, PEP carboxylase

and pyruvate carboxylase, were not detected in our 2-D gels,

although corresponding genes are present in the R. eutrophaH16 genome (H16_A2921 and H16_A1251). In many

bacteria PEP carboxylase and pyruvate carboxylase catalyze

the carboxylation of PEP and pyruvate, respectively, to

oxaloacetate and, thus, contribute to the anaplerotic regen-

eration of the key C4 intermediate. The results of our 2-D

gels are interesting in light of an earlier study by Schobert

F6P E4P

S7PS1,7BP GAP

GAP Xu5P

CbbF /CbbF /Fbp CbbA /CbbA

CbbF /CbbF

G6PPgi1/Pgi2

CbbT /CbbT /TktAA

GAPTpiAGlpK

glycerol glycerol -3-P

GlpD

DHAP

Ru1,5BP

Ru5P

DHAP

Xu5P R5P

p

F1,6BP

CbbP /CbbPCbbS L /

CbbS L

CbbE /CbbE /Rpe

CbbT /CbbT /TktA

CbbG /CbbG /GapA

CbbK /CbbK /Pgk

CbbA /CbbA

CbbA /CbbA /FbaA/FbaB

RpiA

3PGA

CbbE /CbbE /Rpe1,3PGA

pyruvate

acetyl CoA

PEP

Pgm1/Pgm2/A1100

Eno

oxalo-acetate

Pck Ppc

Pyk1/Pyk2/Pyk3

PdhA1BL

CO

CbbS L

PpsA

2PGA

Pyc

B

acetyl-CoA

citrateCisY/A1229/B0357/B0414/B2211

AcnA/AcnB

FumA/FumC

fumarate

Icd2/icd1/icd3

Mdh1/Mdh2

isocitrate

malate

oxalo-acetate

MaeA/MaeB

y

α– keto-glutarateOdhABL

SucCD

SdhABCD

succinate

succinyl-CoA

Figure 2. Central metabolic pathways of R. eutropha H16. (A) Schematic map representing the segments of metabolism relevant for this

study. Arrows indicate the entry points of the carbon sources used in our cultures. Enzyme designations are based on the gene anno-

tations of the R. eutropha H16 genome project. Proteins identified in our 2-D gels are underlined. Metabolites are given in italics. DHAP,

dihydroxyaceton phosphate; E4P, erythrose-4-phosphate; F1,6BP, fructose-1,6-bisphosphate; F6P, fructose-6-phosphate; GAP, glycer-

aldehyde-3-phosphate; G6P, glucose-6-phosphate; 1,3PGA, 1,3-diphosphoglyceric acid; 2PGA, 2-phosphoglyceric acid; 3PGA, 3-phos-

phoglyceric acid; R5P, ribose-5-phosphate; Ru1,5BP, ribulose-1,5-bisphosphate; Ru5P, ribulose-5-phosphate; S1,7BP, sedoheptulose-1,7-

bisphosphate; S7P, sedoheptulose-7-phosphate and Xu5P, xylulose-5-phosphate. (B) The PEP-pyruvate-oxaloacetate node of R. eutropha

H16. Enzymatic reactions discussed in the text are represented by solid black arrows. Other relevant reactions are indicated by grey

arrows. The values for relative transcript abundance (fold difference H2-CO2 versus SN) as determined by RT-PCR are given for four

enzymes: PEP carboxykinase (PEPCK), PEP carboxylase (PPC), PEP synthetase (PPS) and PK. The thickness of the respective arrows is

proportional to these values. The values in parentheses give the fold difference (H2-CO2 versus SN) of the corresponding proteins in 2-D

gels. PC, pyruvate carboxylase; ME, malic enzyme; MDH, malate dehydrogenase and OADC, oxaloacetate decarboxylase.

Proteomics 2009, 9, 5132–5142 5137


and Bowien [25], who measured enzyme activities of the

PEP-pyruvate-oxaloacetate node in cells of R. eutropha H16

grown under various conditions. Neither PEP carboxylase

activity nor pyruvate carboxylase activity was detectable

under any of the conditions tested, including lithoauto-

trophic growth on H2 and CO2 and organoheterotrophic

growth on SN. Schobert and Bowien concluded that in

R. eutropha H16 PEP carboxykinase is, in addition to its

activity as an oxaloacetate decarboxylase, the sole enzyme

capable of carboxylating C3 intermediates to replenish the

oxaloacetate pool.

The levels of the three enzymes of the PEP-pyruvate-

oxaloacetate node were slightly higher (�twofold) in 2-D

gels of lithoautotrophically grown cells compared with

SN-grown cells. In order to obtain more reliable information

on the expression of these important enzymes, we decided

to monitor the relative abundance of the respective tran-

scripts. Using RT-PCR we measured relative transcript

levels for four enzymes in H2-CO2- and SN-grown cells: PEP

carboxykinase, PEP carboxylase, PEP synthetase and PK.

The results of these experiments are shown in Fig. 2B. PEP

carboxykinase expression was tenfold higher in the

lithoautotrophically grown cells versus SN cultures. PK and

PEP synthetase transcripts were 4.5- and 7.6-fold more

abundant, respectively, in the H2-CO2-grown cells. PEP

carboxylase transcripts were present in cells under both

growth conditions but the levels were low. Taken together,

the above results indicate that expression of PEP carbox-

ykinase, PEP synthetase and PK is in fact elevated in cells

growing on H2 and CO2. Furthermore, they suggest

that the gene for PEP carboxylase (ppc) is transcribed,

but that an active enzyme is not formed. The reasons for

higher expression in the lithoautotrophically growing cells

are not clear.

We also identified proteins corresponding to a nearly

complete set of enzymes of gluconeogenesis (fructose-1,6-

bisphosphatase (FBP), fructose-1,6-bisphosphate aldolase

(FBA), triosephosphate isomerase, glyceraldehyde 3-phos-

phate dehydrogenase (GAPDH), phosphoglycerate kinase

(PGK), phosphoglycerate mutase, enolase and PK) in 2-D

gels of lithoautotrophically grown cells. The enzymes FBP,

FBA, GAPDH and PGK are present in multiple isoenzymes

encoded by the duplicate genes of the two cbb operons and

one or two additional chromosomal alleles not associated

with the cbb operons. Clearly, the organism must synthesize

hexoses from C3 building blocks during both lithoauto-

trophic growth and growth on glycerol or SN, requiring the

activities of some or all of the above enzymes. During

growth on SN the cbb genes are tightly repressed. With the

exception of the fbaA gene for FBA, the non-cbb alleles were

expressed at roughly the same levels in H2-CO2-grown cells

as in SN-grown cells, and thus appear to be constitutive.

Expression of the non-cbb gene for FBA is sevenfold up-

regulated. The reason for this is not apparent. During

lithoautotrophic growth, the activities of FBP, FBA, triose-

phosphate isomerase, GAPDH and PGK are required for the

regeneration of the carbon acceptor ribulose-1,5-bispho-

sphate. Under these conditions, the dual cbb operons are

fully induced, providing additional enzymatic capacity to

meet the demands of CO2 fixation. In other words, for

each of the above-named enzymes with the exception of

triosephosphate isomerase, at least three isoenzymes are

formed. In each case, at least two of them were detected in

2-D gels from lithoautotrophic cultures. Proteins corre-

sponding to the specialized enzymes of glycolysis, which in

R. eutropha H16 proceeds via the Entner–Doudoroff Path-

way, were absent from 2-D gels for our three growth

conditions. This is not surprising, since none of the

substrates used in these studies is catabolized by Entner–

Doudoroff enzymes.

Most of the tricarboxylic acid (TCA)-cycle enzymes were

resolved in the 2-D gels of lithoautotrophically grown cells.

Aconitate hydratase, isocitrate dehydrogenase, a-ketogluta-

rate dehydrogenase (subunits E1 and E3), succinyl-CoA

synthetase, succinate dehydrogenase, fumarase, malate

dehydrogenase and citrate synthase were present, although

not all of them could be quantified. This finding is in line

with the results of earlier enzymological and isotopic label-

ling studies [26–30] which demonstrated activities corre-

sponding to a complete set of TCA cycle enzymes in

lithoautotrophically grown cells of R. eutropha H16. The

comparison of H2-CO2- and SN-grown cells revealed that

two enzymes were up-regulated under lithoautotrophic

conditions: aconitate hydratase (sevenfold) and isocitrate

dehydrogenase 1 (3.5-fold).

3.4 Growth on glycerol

In general, the expression of proteins representing the

central metabolic pathways was not markedly different in

lithotrophically grown and glycerol-grown cells. Interest-

ingly, several proteins involved in the biosynthesis of

macromolecules were significantly down-regulated in the

glycerol-grown cells relative to the SN cultures (Supporting

Information Table S3). These include enzymes of pathways

for the biosynthesis of amino acids (IlvC, MetE and CysD),

ribosomal proteins (RplJ, RplI and RpsA) and a component

of the transcriptional apparatus (RpoA). This pattern of

down-regulation is symptomatic for stress responses

observed in E. coli and other bacteria [31]. The curtailment of

expression of this set of genes is triggered by various stress

conditions including nutrient shift-down and exposure to

oxygen radicals. The glycerol-grown cells of R. eutropha H16

revealed the overproduction of four proteins that are

involved in the detoxification of ROS. Catalase encoded by

the gene katG (H16_A2777) was 13.6-fold up-regulated in

glycerol-grown cells. The organic hydroperoxide resistance

protein Ohr (H16_B0157) [32], was up-regulated ninefold

relative to SN-grown cells. Two other proteins, a peroxi-

redoxin (H16_A0306) and a glutathione peroxidase

(H16_A3102), were only detected in cells growing on



glycerol. A peroxiredoxin (H16_A1460) and a superoxide

dismutase, the product of gene sodA (H16_A0610), were

present, but not significantly up-regulated. The function of

the above-named proteins is to protect the cell against ROS.

ROS arise in organisms during normal aerobic metabolism

e.g. via the reaction of O2 with flavoproteins [33]. Certain

conditions lead to increased production of superoxide and

thus increase the incidence of damage to DNA and other cell

components. This state of oxidative stress provokes the

induction of protective proteins, which in E. coli constitute

the soxRS and oxyR regulons [34]. However, some of these

proteins are also produced under other stress conditions

such as nutrient limitation. Furthermore, the response to

oxidative stress is not identical in all bacteria [35, 36]. Thus,

the diagnosis of oxidative stress based on the proteomic data

is not always clear-cut. Two lines of evidence support the

assumption that R. eutropha H16 is subject to exacerbated

oxidative stress during growth on glycerol. First, the catalase

that is up-regulated in the glycerol-grown cells of R. eutrophaH16 is a heme-containing, bifunctional catalase/peroxidase

similar to the hydroperoxidase I of E. coli. In contrast to the

hydroperoxidase II, the hydroperoxidase I is not usually

present in aerobically growing cells, but is induced when

cells are challenged with H2O2 [37]. Another hallmark of the

response to oxidative stress is a shift in the relative levels of

aconitase isoenzymes [38, 39]. Like other bacteria, R. eutro-pha H16 encodes two distinct aconitases: an oxidation-

resistant type (AcnA; H16_A2638) and an oxidation-labile

form (AcnB; H16_B0568). Although the former is present in

the same amounts in glycerol- and SN-grown cells of

R. eutropha H16, the expression of the oxidation-labile form

is curtailed tenfold during growth on glycerol.

What is the cause of elevated levels of ROS in the

glycerol-grown cells? The hydrogenases are formed during

growth on glycerol as well as during growth on H2.

However, the levels of SH under the former conditions are

significantly higher (ca. 4% of total cell protein; [23]) than

during lithoautotrophic growth. Furthermore, the glycerol

cultures are exposed to twice as much O2 as the lithoauto-

trophic cultures. It has been known for some time that

the SH produces superoxide in vitro in the presence

of O2 and electron donors, leading to its own destruction [40,

41]. The copious amounts of SH in the glycerol-grown cells

may produce enough ROS to elicit an oxidative stress

response.

3.5 Regulatory proteins

In total, 12 proteins with functions in signal transduction

and regulation were identified in the proteome of R. eutro-pha cells (Table 4). Three of these proteins, Rho, NusA and

NusG, are transcription termination factors that mediate

termination at both constitutive and variable termination

sites. TypA belongs to the family of TypA/BipA proteins that

are ribosome-binding GTPases controlling both general

housekeeping processes, such as cellular response to stress

[42], as well as strain-specific behavior, such as virulence in

Salmonella typhimurium [43] and symbiotic interaction with

plants in Sinorhizobium meliloti [44]. The proteins GreA1

and DksA1 act in concert with ppGpp and NTPs to shift the

equilibrium between free RNA polymerase and closed RNA

polymerase–promoter complex, thereby regulating the

activity of rRNA promoters, many tRNA promoters and

some mRNA promoters [45]. TypA and GreA1 are down-

regulated tenfold during growth on glycerol. One of the two

isoforms of the nitrogen regulatory protein PII encoded in

the genome was identified in the 2-D gels. This protein was

up-regulated in both the H2-CO2- and the glycerol-grown

cells.

Two response regulators and a serine/threonine protein

kinase-designated PrkA (H16_B0700) were identified in the

protein profiles (Table 4). PrkA is up-regulated twofold and

threefold in H2-CO2-grown cells and glycerol-grown cells,

respectively (ratio H2-CO2/SN: 2.0; ratio glycerol/SN: 3.0).

PrkA belongs to the family of regulators that control cellular

Table 4. Proteins with functions in transcription and regulation identified in 2-D gels


nusA H16_A2307 Transcription pausing factor L 1.2 0.2H16_A1372 H16_A1372 Response regulator, NarL-family 4.5 3.8rho H16_A2395 Transcription termination factor Rho 1.8 0.1prkA H16_B0700 Serine protein kinase 2.0 3.0H16_A0750 H16_A0750 Nitrogen regulatory protein PII 9.1 3.1hoxC PHG021 Regulatory [NiFe] hydrogenase, large subunit 4.0 51.0H16_A1463 H16_A1463 Response regulator, OmpR-family 7.1 10.6nusG H16_A3502 Transcription antitermination protein 1.1 0.3rpoA H16_A3458 DNA-directed RNA polymerase, a-subunit 0.3 0.3typA H16_A2294 GTP-binding elongation factor family protein 0.3 0.1greA1 H16_A2451 Transcription elongation factor GreA 0.0 0.1dksA1 H16_A0194 DnaK suppressor protein 1.3 0.9


Proteomics 2009, 9, 5132–5142 5139


processes by phosphorylating a target protein or proteins at

serine, threonine or tyrosine residues (serine/threonine

protein kinases). Originally discovered and characterized in

eukaryotes, recent years have seen numerous reports of

these proteins in bacteria [46–53].

With the aim of gaining new insights into regulatory

processes involved in reprogramming the metabolism of

R. eutropha H16 for the transition between lithoautotrophic

and organoheterotrophic growth, we generated a knockout

mutant for prkA. The resulting strain, designated HFJ74,

carried an unmarked in-frame deletion in the prkA gene.

The mutant was not significantly affected in lithoauto-

trophic growth on H2-CO2 or organoheterotrophic growth

on SN (data not shown). The role of the prkA gene product

remains unclear.

4 Concluding remarks

The protein inventories obtained in this study provide a

wide-angle albeit static picture of the metabolism of

R. eutropha H16 for three different growth modes. These

results offer a framework for the reassessment of the data of

earlier enzymological studies in light of the recently

published genomic sequences [3, 4]. One interesting finding

is the observation that lithoautotrophically grown cells

contain PEP carboxykinase but no pyruvate carboxylase.

This feature is not correlated with facultative lithoauto-

trophy, since in the facultative lithotroph Paracoccus versutusA2 the anaplerotic formation of oxaloacetate is catalyzed by

pyruvate carboxylase [54].

The previously published enzymological studies revealed

that lithoautotrophically growing cells of R. eutropha H16

contain the enzymatic activities corresponding to a complete

TCA cycle [29, 30]. In accordance with these findings, this

study showed that most of the proteins of TCA cycle

enzymes are detectable in the lithoautotrophically grown

cells. A hallmark of obligate lithoautotrophs is the incom-

plete or interrupted TCA cycle, which becomes a bifurcated

pathway with a biosynthetic but not a bioenergetic role.

These organisms usually lack a functional a-ketoglutarate

dehydrogenase [14, 15, 55]. This type of adaptation has been

documented for Thiobacillus denitrificans, Acidithiobacillusferrooxidans, Hydrogenobacter thermophilus TK-6, and Nitro-somonas europaea [56–59]. Facultative lithoautotrophs, on the

other hand, are able to stop the TCA cycle during the phases

of lithoautotrophic growth. This is achieved by repressing

the expression of a-ketoglutarate dehydrogenase. A repre-

sentative of this group of organisms is P. versutus A2 [54, 56].

Unlike P. versutus A2, R. eutropha H16 maintains a func-

tional TCA cycle during lithoautotrophic growth. Thus,

inactivation of the TCA cycle is not a prerequisite for

lithoautotrophic growth.

The finding that R. eutropha H16 retains a functional

TCA cycle during lithoautotrophic growth is of major

importance in the context of the mixotrophic capabilities of

this organism. Mixotrophy, i.e. the capacity to utilize inor-

ganic and organic substrates concomitantly, has been

reported for R. eutropha H16 and for some other aerobic H2

oxidizers including Aquaspirillum autotrophicum [60], for the

carboxidotroph Hydrogenophaga pseudoflava [61] and for

the facultative sulfur oxidizer P. versutus A2 [54, 62]. For the

latter three organisms there is convincing evidence that

mixotrophic growth does not involve CO2 fixation. In a

series of elegant experiments based on mutants of

R. eutropha H16 and carbon-limited chemostat cultures,

K.arst and Friedrich showed that, unlike the mixotrophic

bacteria named above, R. eutropha H16 does in fact rely on

CO2 fixation for mixotrophic growth [63]. For a chemostat

culture of R. eutropha H16 growing in the presence of H2

and limited for succinate as the sole carbon source, the

authors observed an increase in yield of 135%. Since there

was no exogenous CO2 available to the culture, but the yield

increase was largely dependent on the activity of the Calvin-

cycle enzymes, the authors concluded that the carbon

substrate was respired and the resulting CO2 then fixed and

assimilated to form cell material. A small but significant

increase in growth yield (14%) was observed when a mutant

defective for CO2 fixation was cultivated on succinate and

H2, indicating that, to a limited extent, both substrates were

used concomitantly as energy sources.

Taken together, the different lines of evidence show that

R. eutropha H16 maintains a basically heterotrophic core

metabolism during lithoautotrophic growth on H2 and CO2.

This is compatible with the notion that R. eutropha H16 is a

heterotroph, which has relatively recently acquired the

megaplasmid-encoded capacity for lithoautotrophic growth.

The authors thank Susanne Paprotny und Enrico Klotz forexpert technical assistance and Decodon GmbH (Greifswald,Germany) for providing the Delta2D software. E. S. is grateful toBotho Bowien for enlightening discussions. This work wassupported by grants from the Bundesministerium f .ur Bildungund Forschung in the framework of the program ‘‘GenomicResearch on Bacteria (GenoMik)’’ to B. F and M. H. and fromthe Fonds der chemischen Industrie to B. F.

The authors have declared no conflict of interest.

5 References

[1] Friedrich, B., Friedrich, C. G., in: Codd, G. A., Dijkhuizen, L.,

Tabita, F. R. (Eds.), Autotrophic Microbiology and One-

Carbon Metabolism, Kluwer Academic Publishers,

Dordrecht 1990, pp. 55–92.

[2] Schwartz, E., Friedrich, B., in: Dworkin, M., Falkow, S.,

Rosenberg, E., Schleifer, K. H., Stackebrandt, E. (Eds.), The

Prokaryotes, 3rd Edn. Springer, New York 2006, pp.

496–563.

[3] Schwartz, E., Henne, A., Cramm, R., Eitinger, T. et al.,

Complete nucleotide sequence of pHG1: a Ralstonia eutropha



H16 megaplasmid encoding key enzymes of H2-based

lithoautotrophy and anaerobiosis. J. Mol. Biol. 2003, 332,

369–383.

[4] Pohlmann, A., Fricke, W. F., Reinecke, F., Kusian, B. et al.,

Genome sequence of the bioplastic-producing ‘‘Knallgas’’

bacterium Ralstonia eutropha H16. Nat. Biotechnol. 2006,

24, 1257–1262.

[5] Burgdorf, T., Lenz, O., Buhrke, T., van der Linden, E. et al.,

[NiFe]-hydrogenases of Ralstonia eutropha H16: modular

enzymes for oxygen-tolerant biological hydrogen oxidation.

J. Mol. Microbiol. Biotechnol. 2005, 10, 181–196.

[6] Lenz, O., Friedrich, B., A novel multicomponent regulatory

system mediates H2 sensing in Alcaligenes eutrophus. Proc.

Nat. Acad. Sci. USA 1998, 95, 12474–12479.

[7] Friedrich, B., Buhrke, T., Burgdorf, T., Lenz, O., A hydrogen-

sensing multiprotein complex controls aerobic hydrogen

metabolism in Ralstonia eutropha. Biochem. Soc. Trans.

2005, 33, 97–101.

[8] Burgdorf, T., van der Linden, E., Bernhard, M., Yin, Q. Y.

et al., The soluble NAD1-Reducing [NiFe]-hydrogenase

from Ralstonia eutropha H16 consists of six subunits and

can be specifically activated by NADPH. J. Bacteriol. 2005,

187, 3122–3132.

[9] Friedrich, C. G., Friedrich, B., Bowien, B., Formation of

enzymes of autotrophic metabolism during heterotrophic

growth of Alcaligenes eutrophus. J. Gen. Microbiol. 1981,

122, 69–78.

[10] Bowien, B., Kusian, B., Genetics and control of CO2 assim-

ilation in the chemoautotroph Ralstonia eutropha. Arch.

Microbiol. 2002, 178, 85–93.

[11] Kusian, B., Bednarski, R., Husemann, M., Bowien, B.,

Characterization of the duplicate ribulose-1,5-bisphosphate

carboxylase genes and cbb promoters of Alcaligenes

eutrophus. J. Bacteriol. 1995, 177, 4442–4450.

[12] Schwartz, E., Gerischer, U., Friedrich, B., Transcriptional

regulation of Alcaligenes eutrophus hydrogenase genes.

J. Bacteriol. 1998, 180, 3197–3204.

[13] Schwartz, E., Buhrke, T., Gerischer, U., Friedrich, B., Positive

transcriptional feedback controls hydrogenase expression

in Alcaligenes eutrophus H16. J. Bacteriol. 1999, 181,

5684–5692.

[14] Smith, A. J., Hoare, D. S., Specialist phototrophs, litho-

trophs, and methylotrophs: a unity among a diversity of

procaryotes? Bacteriol. Rev. 1977, 41, 419–448.

[15] Matin, A., Organic nutrition of chemolithotrophic bacteria.

Annu. Rev. Microbiol. 1978, 32, 433–468.

[16] Wood, A. P., Aurikko, J. P., Kelly, D. P., A challenge for 21st

century molecular biology and biochemistry: what are the

causes of obligate autotrophy and methanotrophy? FEMS

Microbiol. Rev. 2004, 28, 335–352.

[17] Chain, P., Lamerdin, J., Larimer, F., Regala, W. et al.,

Complete genome sequence of the ammonia-oxidizing

bacterium and obligate chemolithoautotroph Nitrosomo-

nas europaea. J. Bacteriol. 2003, 185, 2759–2773.

[18] Beller, H. R., Chain, P. S., Letain, T. E., Chakicherla, A. et al.,

The genome sequence of the obligately chemolithoauto-

trophic, facultatively anaerobic bacterium Thiobacillus

denitrificans. J. Bacteriol. 2006, 188, 1473–1488.

[19] Eberz, G., Friedrich, B., Three trans-acting regulatory func-

tions control hydrogenase synthesis in Alcaligenes eutro-

phus. J. Bacteriol. 1991, 173, 1845–1854.

[20] B .uttner, K., Bernhardt, J., Scharf, C., Schmid, R. et al.,

A comprehensive two-dimensional map of cytosolic

proteins of Bacillus subtilis. Electrophoresis 2001, 22,

2908–2935.

[21] Voigt, B., Schweder, T., Sibbald, M. J., Albrecht, D. et al.,

The extracellular proteome of Bacillus licheniformis grown

in different media and under different nutrient starvation

conditions. Proteomics 2006, 6, 268–281.

[22] Lenz, O., Schwartz, E., Dernedde, J., Eitinger, M. et al., The

Alcaligenes eutrophus H16 hoxX gene participates in

hydrogenase regulation. J. Bacteriol. 1994, 176, 4385–4393.

[23] K .arst, U., Suetin, S., Friedrich, C. G., Purification and

properties of a protein linked to the soluble hydrogenase

of hydrogen-oxidizing bacteria. J. Bacteriol. 1987, 169,

2079–2085.

[24] Sauer, U., Eikmanns, B. J., The PEP-pyruvate-oxaloacetate

node as the switch point for carbon flux distribution in

bacteria. FEMS Microbiol. Rev. 2005, 29, 765–794.

[25] Schobert, P., Bowien, B., Unusual C3 and C4 metabolism in

the chemoautotroph Alcaligenes eutrophus. J. Bacteriol.

1984, 159, 167–172.

[26] Hirsch, P., Georgiev, G., Schlegel, H. G., [CO2 fixation by

knallgas bacteria. III. Autotrophic and organotrophic CO2

fixation]. Arch. Mikrobiol. 1963, 46, 79–95.

[27] Glaeser, H., Schlegel, H. G., [Synthesis of enzymes of the

tricarboxylic acid cycle in Hydrogenomonas eutropha strain

H 16]. Arch. Mikrobiol. 1972, 86, 315–325.

[28] Br .amer, C. O., Steinb .uchel, A., The malate dehydrogenase

of Ralstonia eutropha and functionality of the C3/C4 meta-

bolism in a Tn5-induced mdh mutant. FEMS Microbiol. Lett.

2002, 212, 159–164.

[29] Tr .uper, H. G., Tricarboxylic acid cycle and related enzymes

in Hydrogenomonas strain H16G1 grown in various carbon

sources. Biochim.Biophys. Acta 1965, 111, 565–568.

[30] Smith, A. J., London, J., Stanier, R. Y., Biochemical basis of

obligate autotrophy in blue-green algae and thiobacilli.

J. Bacteriol. 1967, 94, 972–983.

[31] Chang, D. E., Smalley, D. J., Conway, T., Gene expression

profiling of Escherichia coli growth transitions: an expanded

stringent response model. Mol. Microbiol. 2002, 45,

289–306.

[32] Mongkolsuk, S., Praituan, W., Loprasert, S., Fuangthong, M.

et al., Identification and characterization of a new organic

hydroperoxide resistance (ohr) gene with a novel pattern of

oxidative stress regulation from Xanthomonas campestris

pv. phaseoli. J. Bacteriol. 1998, 180, 2636–2643.

[33] Gonzalez-Flecha, B., Demple, B., Metabolic sources of

hydrogen peroxide in aerobically growing Escherichia coli.

J. Biol. Chem. 1995, 270, 13681–13687.

[34] Imlay, J. A., Cellular defenses against superoxide and

hydrogen peroxide. Annu. Rev. Biochem. 2008, 77, 755–776.

Proteomics 2009, 9, 5132–5142 5141


[35] Mostertz, J., Scharf, C., Hecker, M., Homuth, G., Tran-

scriptome and proteome analysis of Bacillus subtilis gene

expression in response to superoxide and peroxide stress.

Microbiology 2004, 150, 497–512.

[36] Wolf, C., Hochgr .afe, F., Kusch, H., Albrecht, D. et al.,

Proteomic analysis of antioxidant strategies of Staphylo-

coccus aureus: diverse responses to different oxidants.

Proteomics 2008, 8, 3139–3153.

[37] Schellhorn, H. E., Regulation of hydroperoxidase (catalase)

expression in Escherichia coli. FEMS Microbiol. Lett. 1995,

131, 113–119.

[38] Gardner, P. R., Fridovich, I., Superoxide sensitivity of the

Escherichia coli aconitase. J. Biol. Chem. 1991, 266,

19328–19333.

[39] Cunningham, L., Gruer, M. J., Guest, J. R., Transcriptional

regulation of the aconitase genes (acnA and acnB) of

Escherichia coli. Microbiology 1997, 143, 3795–3805.

[40] Schneider, K., Schlegel, H. G., Production of superoxide

radicals by soluble hydrogenase from Alcaligenes eutro-

phus H16. Biochem. J. 1981, 193, 99–107.

[41] Schlesier, M., Friedrich, B., In vivo inactivation of soluble

hydrogenase of Alcaligenes eutrophus. Arch. Microbiol.

1981, 129, 150–153.

[42] Freestone, P., Trinei, M., Clarke, S. C., Nystrom, T. et al.,

Tyrosine phosphorylation in Escherichia coli. J. Mol. Biol.

1998, 279, 1045–1051.

[43] Qi, S. Y., Li, Y., Szyroki, A., Giles, I. G. et al., Salmonella

typhimurium responses to a bactericidal protein from

human neutrophils. Mol. Microbiol. 1995, 17, 523–531.

[44] Kiss, E., Huguet, T., Poinsot, V., Batut, J., The typA gene is

required for stress adaptation as well as for symbiosis

of Sinorhizobium meliloti 1021 with certain Medicago

truncatula lines. Mol. Plant Microbe Interact. 2004, 17,

235–244.

[45] Rutherford, S. T., Lemke, J. J., Vrentas, C. E., Gaal, T. et al.,

Effects of DksA, GreA, and GreB on transcription initiation:

insights into the mechanisms of factors that bind in the

secondary channel of RNA polymerase. J. Mol. Biol. 2007,

366, 1243–1257.

[46] Deutscher, J., Saier, M. H., Jr., Ser/Thr/Tyr protein phos-

phorylation in bacteria – for long time neglected, now

well established. J. Mol. Microbiol. Biotechnol. 2005, 9,

125–131.

[47] Cozzone, A. J., Role of protein phosphorylation on serine/

threonine and tyrosine in the virulence of bacterial patho-

gens. J. Mol. Microbiol. Biotechnol. 2005, 9, 198–213.

[48] Munoz-Dorado, J., Inouye, S., Inouye, M., A gene encoding

a protein serine/threonine kinase is required for normal

development of M. xanthus, a gram-negative bacterium.

Cell 1991, 67, 995–1006.

[49] Matsumoto, A., Hong, S. K., Ishizuka, H., Horinouchi, S.

et al., Phosphorylation of the AfsR protein involved in

secondary metabolism in Streptomyces species by a

eukaryotic-type protein kinase. Gene 1994, 146, 47–56.

[50] Herro, R., Poncet, S., Cossart, P., Buchrieser, C. et al., How

seryl-phosphorylated HPr inhibits PrfA, a transcription

activator of Listeria monocytogenes virulence genes.

J. Mol. Microbiol. Biotechnol. 2005, 9, 224–234.

[51] Fischer, C., Geourjon, C., Bourson, C., Deutscher, J., Clon-

ing and characterization of the Bacillus subtilis prkA gene

encoding a novel serine protein kinase. Gene 1996, 168,

55–60.

[52] Madec, E., Laszkiewicz, A., Iwanicki, A., Obuchowski, M.

et al., Characterization of a membrane-linked Ser/Thr

protein kinase in Bacillus subtilis, implicated in develop-

mental processes. Mol. Microbiol. 2002, 46, 571–586.

[53] Kurbatov, L., Albrecht, D., Herrmann, H., Petruschka, L.,

Analysis of the proteome of Pseudomonas putida KT2440

grown on different sources of carbon and energy. Environ.

Microbiol. 2006, 8, 466–478.

[54] Smith, A. L., Kelly, D. P., Wood, A. P., Metabolism of Thio-

bacillus A2 grown under autotrophic, mixotrophic and

heterotrophic conditions in chemostat culture. J. Gen.

Microbiol. 1980, 121, 127–138.

[55] Wood, A. P., Woodall, C. A., Kelly, D. P., Halothiobacillus

neapolitanus strain OSWA isolated from ‘‘The Old Sulphur

Well’’ at Harrogate (Yorkshire, England). Syst. Appl.

Microbiol. 2005, 28, 746–748.

[56] Peeters, T. L., Liu, M. S., Aleem, M. I., The tricarboxylic acid

cycle in Thiobacillus denitrificans and Thiobacillus A2.

J. Gen. Microbiol. 1970, 64, 29–35.

[57] Ingledew, W. J., Thiobacillus ferrooxidans. The bioener-

getics of an acidophilic chemolithotroph. Biochim. Biophys.

Acta 1982, 683, 89–117.

[58] Shiba, H., Kawasumi, T., Igarashi, Y., Kodama, T. et al., The

deficient carbohydrate metabolic pathways and the

incomplete tricarboxylic acid cycle in an obligately auto-

trophic hydrogen-oxidizing bacterium. Agric. Biol. Chem.

1982, 46, 2341–2354.

[59] Hooper, A. B., Biochemical basis of obligate autotrophy in

Nitrosomonas europaea. J. Bacteriol. 1969, 97, 776–779.

[60] Fasnacht, M., Aragno, M., Mixotrophy and lithohetero-

trophy by Aquaspirillum autotrophicum. Experientia 1982,

38, 1377.

[61] Kiessling, M., Meyer, O., Profitable oxidation of carbon

monoxide or hydrogen during heterotrophic growth of

Pseudomonas carboxydoflava. FEMS Microbiol. Lett. 1982,

13, 333–338.

[62] Gottschal, J. C., Kuenen, J. G., Mixotrophic growth of

Thiobacillus A2 on acetate and thiosulfate as growth limit-

ing substrates in the chemostat. Arch. Microbiol. 1980, 126,

33–42.

[63] K .arst, U., Friedrich, C. G., Mixotrophic capabilities of Alca-

ligenes eutrophus. J. Gen. Microbiol. 1984, 130, 1987–1994.



A proteomic view of the facultatively chemolithoautotrophic lifestyle of Ralstonia eutropha H16

Documents

Transcript of A proteomic view of the facultatively chemolithoautotrophic lifestyle of Ralstonia eutropha H16