Molecular Microbiology (2009) 71(4), 876–894 !
doi:10.1111/j.1365-2958.2008.06568.x
First published online 22 December 2008
Genome-wide responses to carbonyl electrophiles in Bacillus
subtilis: control of the thiol-dependent formaldehyde
dehydrogenase AdhA and cysteine proteinase YraA by the
MerR-family regulator YraB (AdhR)
Nguyen Thi Thu Huyen,1†
Warawan Eiamphungporn,2† Ulrike Mäder,3
Manuel Liebeke,4 Michael Lalk,4 Michael Hecker,1
John D. Helmann2* and Haike Antelmann1**
1
Institute for Microbiology, 3Interfaculty Institute for
Genetics and Functional Genomics and 4Institute for
Pharmaceutical Biology, Ernst-Moritz-Arndt-University of
Greifswald, D-17487 Greifswald, Germany. 2Department
of Microbiology, Cornell University, Ithaca, NY
14853-8101, USA.
Summary
Quinones and a,b-unsaturated carbonyls are naturally occurring electrophiles that target cysteine residues via thiol-(S)-alkylation. We analysed the global
expression profile of Bacillus subtilis to the toxic carbonyls methylglyoxal (MG) and formaldehyde (FA).
Both carbonyl compounds cause a stress response
characteristic for thiol-reactive electrophiles as
revealed by the induction of the Spx, CtsR, CymR,
PerR, ArsR, CzrA, CsoR and SigmaD regulons. MG
and FA triggered also a SOS response which indicates DNA damage. Protection against FA is mediated
by both the hxlAB operon, encoding the ribulose
monophosphate pathway for FA fixation, and a thioldependent formaldehyde dehydrogenase (AdhA)
and DJ-1/PfpI-family cysteine proteinase (YraA). The
adhA–yraA operon and the yraC gene, encoding a
g-carboxymuconolactone decarboxylase, are positively regulated by the MerR-family regulator,
YraB(AdhR). AdhR binds specifically to its target
promoters which contain a 7-4-7 inverted repeat
(CTTAAAG-N4-CTTTAAG) between the -35 and -10
elements. Activation of adhA–yraA transcription by
AdhR requires the conserved Cys52 residue in vivo.
Accepted 3 December, 2008. For correspondence. *E-mail jdh9@
cornell.edu; Tel. (+1)607 2556570; Fax (+1) 607 2553904;
**E-mail: antelman@uni-greifswald.de; Tel. (+49) 3834 864237; Fax
(+49) 3834 864202. †These authors contributed equally to the work.
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd
We speculate that AdhR is redox-regulated via thiol(S)-alkylation by aldehydes and that AdhA and YraA
are specifically involved in reduction of aldehydes
and degradation or repair of damaged thiolcontaining proteins respectively.
Introduction
Reactive oxygen species (ROS) are produced as an
unavoidable consequence of the aerobic lifestyle (Imlay,
2002). The oxidants generated from incomplete O2 reduction can react with cellular components, such as amino
acids, carbohydrates or lipids, to produce a range of
unstable oxidation products that break down to diffusible
electrophiles (Marnett et al., 2003; Farmer and Davoine,
2007). These secondary oxidants include a,b-unsaturated
aldehydes, which are of particular importance as these
have two sites of reactivity, which can lead to the formation of cyclic adducts or cross-links. Thus, eukaryotic and
prokaryotic cells have evolved defence mechanisms to
detoxify ROS and reactive electrophiles and to repair the
resulting damage.
Redox-sensing transcription factors respond to
changes in the cellular redox status caused either directly
by ROS or as consequence of oxidized reactive intermediates which act as electrophiles. Such redox-sensing
proteins often have conserved cysteine residues that
serve as a primary target for oxidative or covalent
modifications. The thiol group of cysteine can undergo
either reversible modification, such as disulphide bond
formation, or irreversible modifications including oxidation
to cysteine sulphinic or sulphonic acids or thiol-(S)alkylation (Giles et al., 2003; Marnett et al., 2003; Barford,
2004; Satoh and Lipton, 2007). The reaction of electrophilic quinones, hydroquinones and a,b-unsaturated carbonyls involves a 1,4-reductive nucleophilic addition of
thiols to quinones (Michael-type addition) (O’Brian, 1991;
Monks et al., 1992; Kumagai et al., 2002; Marnett et al.,
2003). Thiol-(S)-alkylation can function in redox signalling
as first described for the Escherichia coli Ada protein,
which functions both as a DNA repair protein, by methyl
Regulation of carbonyl detoxification by AdhR 877
transfer from methylated bases and phosphates, and
as a transcription activator regulated by cysteine-Smethylation in response to environmental methylating
agents such as methyl chloride, the antibiotic streptozotocin, N-methyl-N′-nitrosourea and N-methyl-N′-nitro-Nnitrosoguanidine (Sedgwick and Vaughan, 1991; He
et al., 2005; Takinowaki et al., 2006). A mechanistically
related pathway is operative in human neuronal cells
where it regulates transcription in response to electrophiles (Satoh and Lipton, 2007). Electrophiles can protect
neurons through activation of the Keap-Nfr2 pathway
resulting in the induction of the phase-2 antioxidant
enzymes glutathione (GSH) reductase and quinone
oxidoreductase. Neuroprotective electrophiles react with
specific cysteine residues in the Keap1 protein via thiol(S)-alkylation which in turn liberates the transcription
factor Nfr2. Nfr2 translocates to the nucleus where it activates transcription of antioxidant enzymes (Holtzclaw
et al., 2004; Wakabayashi et al., 2004; Dinkova-Kostova
et al., 2005; Satoh et al., 2006; Satoh and Lipton, 2007).
We have recently discovered a redox-regulated system
in the Gram-positive bacterium Bacillus subtilis which
regulates the response to quinone-like electrophiles
and diamide (Töwe et al., 2007; Antelmann et al.,
2008; Leelakriangsak et al., 2008). Quinones, such as
menaquinone and ubiquinone, are endogenously produced electrophiles which shuttle electrons as part of the
respiratory chain (Taber, 1993; Gennis and Stewart,
1996). The menaquinone concentration in B. subtilis is
~2–5 nmol (mg membrane)-1 (Farrand and Taber, 1974).
Quinones are also abundant in soil, where they account
for a large fraction of the redox-active component of
humic substrances (Ratasuk and Nanny, 2007; Steinberg
et al., 2008) and in dissolved organic matter from a variety
of ecosystems (Nurmi and Tratnyek, 2002; Cory and McKnight, 2005). A subset of these quinones are cell permeable and model compound studies suggest that they will
likely react with thiol-containing proteins via thiol-(S)alkylation leading to protein cross-linking and aggregation
(Liebeke et al., 2008). B. subtilis expresses an adaptive
response to quinone-like electrophiles and diamide that is
mediated by two MarR-type repressors, YodB and MhqR.
These control the expression of quinone or azocompound
reductases and thiol-dependent dioxygenases that function in detoxification of these electrophiles. The MarR/
DUF24-family repressor YodB is alkylated in vitro at the
conserved Cys6 residue and this residue was essential
for repression also in vivo (Töwe et al., 2007; Leelakriangsak et al., 2008).
Besides quinone compounds, a,b-unsaturated aldehydes are highly toxic natural electrophiles which are
generated from oxidation of amino acids, lipids and carbohydrates (Marnett et al., 2003; Farmer and Davoine,
2007). Quinones and a,b-unsaturated aldehydes both
react with cysteine thiols via the Michael addition
mechanism. Methylglyoxal (MG) is the most studied toxic
natural a-oxoaldehyde and is produced from triosephosphate intermediates as a by-product of glycolysis,
and also from amino acids and acetone (Ferguson et al.,
1998; Booth et al., 2003; Kalapos, 2008). MG reacts with
nucleophilic centres of DNA, RNA and with the sidechains of the amino acids arginine, lysine and cysteine.
The reaction of MG with arginine and lysine residues
results in the formation of the advanced glycation endproducts (AGE), such as argpyrimidine or MG-crosslinked lysine dimers (Oya et al., 1999; Bourajjaj et al.,
2003). These AGE modifications are associated with
pathogenesis of various human diseases such as diabetes and cancer (Logsdon et al., 2007; Toth et al., 2007).
The major protective mechanism against MG in E. coli is
the spontaneous reaction with GSH to form the hemithioactel, followed by isomerization to S-lactoylglutathione
and conversion to D-lactate by the glyoxalase I and II
enzymes (Ferguson et al., 1998; Booth et al., 2003)
(Fig. 1A).
Formaldehyde (FA) is another highly toxic carbonyl
compound and causes protein–protein and protein–DNA
cross-links in vivo (Orlando et al., 1997). Like MG, FA
reacts as an electrophile with the side-chains of arginine
and lysine. As FA is a key intermediate in the metabolism
of C1 compounds in methanotrophic and methylotrophic
bacteria, it is ubiquitously distributed in the environment.
Thus, prokaryotes and eukaryotes have evolved conserved pathways for FA detoxification. These include
GSH-dependent formaldehyde dehydrogenases (Fdh),
which are conserved in all kingdoms of life (Harms et al.,
1996). Fdh catalyses the NAD-dependent oxidation
of S-hydroxymethylglutathione into S-formylglutathione
which is hydrolysed by a S-formylglutathione hydrolase to
GSH and formate (Jensen et al., 1998) (Fig. 1B). In addition, the ribulose monophosphate pathway (RuMP), used
for FA fixation in methylotrophs, is involved in FA detoxification in many non-methylotrophs including B. subtilis
(Yurimoto et al., 2005; Kato et al., 2006). 3-Hexulose-6phosphate synthase and 6-phospho-3-hexuloisomerase,
the key enzymes in the RuMP pathway, are encoded by
the hxlAB operon in B. subtilis. The hxlAB operon is positively regulated by the MarR/DUF24 family protein HxlR,
which is 39% identical to the quinone-responsive YodB
repressor (Yurimoto et al., 2005; Leelakriangsak et al.,
2008).
In this study, we have applied genome-wide transcriptome and proteome profiling to define the adaptive
response of B. subtilis to MG and FA. Both compounds
induced a thiol-specific electrophile stress response
superficially similar to that seen with quinones and
diamide. In addition, FA and MG strongly induced the
adhA–yraA operon encoding a conserved Fdh and a cys-
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
878 N. T. T. Huyen et al. !
A
CH3
H
CH3
C
O
C
O
+R-SH
H
CH3
Glx-I
C
O
C
OH
HC
SR
Methylglyoxal (MG)
OH
C
CH3
Glx-II
Glx
II
+H2O
O
HC
C
R-SH
SR
Hemithioacetal
OH
O
OH
S-Lactoylthiol
Lactate
B
NAD
H
H
C
O
+R-SH
H
Formaldehyde (FA)
NADH+H
AdhA
H
C
OH
RS
S
S-HydroxyHydroxy
methylthiol
H
H
C
O
RS
S
Formylthiol
S-Formylthiol
+H2O
R-SH
C
O
HO
Formate
Fig. 1. Thiol-dependent detoxification pathways for methylgloxal (A) and formaldehyde (B) in bacteria.
A. MG reacts spontaneously with GSH to form hemithioacetal, followed by isomerization to S-lactoylglutathione by the glyoxalase I enzyme.
S-lactoylglutathione is substrate for the glyoxalase II enzyme and is converted to D-lactate and free GSH. This scheme is adapted from
Ferguson et al. (1998).
B. In prokaryotes and eukaryotes FA is detoxified by GSH-dependent Fdhs (Harms et al., 1996). FA reacts spontaneously and reversibly with
GSH to form S-hydroxymethylglutathione. Fdh-like enzymes such as class III alcohol dehydrogenases catalyse the NAD-dependent oxidation
of S-hydroxymethylglutathione into S-formylglutathione (Jensen et al., 1998). This GSH thiol ester is reversibly hydrolysed by a
S-formylglutathione hydrolase to GSH and formate. This scheme is adapted from Jensen et al. (1998).
teine proteinase of the DJ1/PfpI family. This operon is
controlled by the MerR/NmlR-family regulator AdhR (formerly YraB), which contains a conserved Cys52 residue
essential for transcription activation in response to electrophilic carbonyls in vivo.
is encoded by the yurT gene in B. subtilis, it is not known
if this functions in MG detoxification.
Results and discussion
We performed transcriptome and proteome analyses in
response to 1 mM FA, 2.8 mM and 5.6 mM MG. Genes
that were >3-fold upregulated and downregulated by FA
and MG were sorted according to the known stress regulons in Tables S1 and S2. The microarray results revealed
that 1 mM FA and 5.6 mM MG caused the upregulation
of the CtsR, Spx and CymR regulons in B. subtilis. The
proteome results verfied the induction of the CtsRdependent Clp proteases ClpE, ClpC, the CymRregulated proteins CysK, YrhB, YxeP, YxeK, and the Spxdependent proteins TrxB, NfrA, YqiG and YugJ by both
carbonyl compounds (Fig. 3). The induction of the CtsR,
CymR and Spx regulons in B. subtilis was previously
shown also by quinone-like electrophiles and diamide
(Antelmann et al., 2008). Remarkedly, FA induced the
CtsR, Spx and CymR regulons at significantly higher
ratios than those observed for quinones and diamide
(Leichert et al., 2003; Tam et al., 2006; Duy et al., 2007).
Transcriptional induction of selected genes and operons
controlled by CtsR, Spx and CymR was verified using
Northern blot analyses (Fig. 4). Because of this overlapping expression profile by aldehydes, quinones and
B. subtilis tolerates high concentrations of FA and MG
Cells were grown in a minimal medium to an OD500 of 0.4
and treated with 0.5–10 mM FA and 1.4–11.2 mM MG to
define concentrations which cause growth inhibition but
are sublethal. As noted for other bacteria, B. subtilis was
able to tolerate high doses of FA: 1–2 mM FA caused a
decreased growth rate but did not affect the number of
viable counts (Fig. 2A). The survival ratio was significantly
decreased by 5 mM FA, indicating that this concentration
is toxic for B. subtilis. The tolerance level of B. subtilis to
FA is comparable to that of Pseudomonas putida, E. coli
or other bacilli (Roca et al., 2008). This is consistent with
the demonstration that B. subtilis is able to detoxify FA,
probably via the RuMP pathway encoded by the hxlAB
operon (Yurimoto et al., 2005; Kato et al., 2006). In
response to MG challenge, the growth rate of wild-type
cells was slightly reduced by concentrations of 1.4 mM
and inhibited by 2.8 mM, but without loss of viability
(Fig. 2B). Toxic effects of MG were observed after exposure to 5.6 mM MG. Although a putative methylglyoxalase
FA and MG induce the general electrophile stress
response (CtsR, Spx, CymR regulons)
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
Regulation of carbonyl detoxification by AdhR 879
A
Formaldehyde
B
Methylglyoxal
10
OD500 (nm))
OD500 (nm))
10
1
0.1
0.01
-150
co
1 mM FA
5 mM FA
-90
-30
30
90
210
Viable coun
V
nts (cfu)
Viable coun
V
nts (cfu)
108
107
106
105
co
1 mM FA
5 mM
M FA
103
102
-150
-90
-30
0.5 mM FA
2 mM FA
10 mM
M FA
30
90
0.01
-120
0
270
109
104
0.1
0.5 mM FA
2 mM FA
10 mM FA
150
150
210
270
1
co
2.8 mM MG
11.2 mM MG
-60
60
0
1.4 mM MG
5.6 mM MG
60
120
0
180
80
240
0
300
60
120
180
240
300
109
108
107
106
105
104
103
102
co
1.4 mM MG
2.8 mM MG
5.6 mM MG
11 2 mM MG
11.2
-120 -60
0
Time(min)
Time(min)
Fig. 2. Growth curves (upper panels) and survival ratios (lower panels) of B. subtilis wild-type cells in the presence of FA (left) and MG
(right). B. subtilis wild-type cells were grown in minimal medium to an OD500 of 0.4 and exposed to 0.5, 1, 2, 5 and 10 mM FA or 1.4, 2.8, 5.6
and 11.2 mM MG at the time points that were set to zero. Appropriate dilutions were plated for viable counts (cfu ml-1).
diamide, we can regard this signature as a ‘general electrophile stress-response’ in B. subtilis. Spx is a global
transcriptional regulator that controls genes that function
in maintenance of thiol-homeostasis under conditions of
electrophile stress such as the thioredoxin/thioredoxin
reductase encoding trxA and trxB genes (Zuber, 2004).
CtsR controls the Clp proteases which are involved in
repair or degradation of damaged or misfolded proteins
(Krüger and Hecker, 1998; Derré et al., 1999). The cysteine metabolism repressor CymR controls sulphur
metabolism operons and several pathways leading to cysteine formation (Even et al., 2006). Together, the induction
of the CtsR, Spx and CymR regulons by thiol-reactive
electrophiles indicates an imbalanced thiol-redox homeostasis due to depletion of low-molecular-weight (LMW)
thiol buffers such as cysteine.
FA and MG induce weakly quinone detoxification
systems (YodB, YvaP regulons)
The MarR-type repressors YodB and MhqR control
paralogous oxidoreductases and thiol-dependent dioxygenases that function in diamide and quinone detoxification in B. subtilis (Töwe et al., 2007; Antelmann et al.,
2008; Leelakriangsak et al., 2008). The yfiDE (catDE)
operon encodes another thiol-dependent catechol-2,3dioxygenase (Tam et al., 2006) that is regulated by the
YodB-paralog repressor YvaP (CatR) (H. Antelmann and
K. Kobayashi, unpubl. data). MG leads to weak upregulation of the YodB-dependent yodC, spx and azoR1
genes, the MhqR-dependent mhqA gene as well as the
yfiDE (catDE) operon. Thus, we hypothesize that electrophilic carbonyl compounds react with regulatory thiols of
several distinct redox-sensing transcription factors.
FA and MG induce the oxidative stress response
(PerR regulon)
FA and MG caused the induction of the peroxide stress
responsive PerR regulon (Fig. 4). However, in the proteome experiments we observed no induction of KatA,
AhpC and AhpF. Aldehydes caused a relatively weak
induction of the PerR regulon at the transcriptional level,
which might be not visible at the level of protein synthesis.
The induction of the PerR regulon by FA and MG could
indicate that electrophilic carbonyl compounds produce
ROS (Mongkolsuk and Helmann, 2002). MG has been
shown to provoke free radical generation involving ROS
and methylglyoxal radicals, which contribute also to MG
toxicity mechanisms in eukaryotic cells (Kalapos, 2008).
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
880 N. T. T. Huyen et al. !
168_
168 co
168_5.6 mM MG
ClpC
GroEL
NfrA
YfkO
YjbG
KatA
ClpE
YrhB YqiG
YacK YxeP
HemH
HemH
YugJ
TrxB
Cah
CysK
YfjR
YwfI
YdgI HxlA
DnaK
GroEL
Ah
F
AhpF
YxeK
YqiG
YrhB
YxeP
YugJ
CysK
ClpE
AhpF
YxeK
YacK
MrgAC
ClpC
ClpE
YjbG
KatA
168_ co
168
168_1mM FA
MrgAC
NfrA
YfkO
AhpC
TrxB
YvyD
ClpP
GrpE
YfjR
YwfI
YdgI HxlA
AzoR1
AhpC
CH3
AzoR1
H
C
O
Tpx
C
MrgA
H
C
O
MrgA
O
H
Fig. 3. Dual-channel images of the protein synthesis pattern of B. subtilis wild-type before (green image) and 10 min after the exposure to
5.6 mM MG (A, red image) or 1 mM FA (B, red image). Cytoplasmic proteins were labelled with L-[35S]methionine and separated by 2D PAGE
as described in Experimental procedures. Image analysis of the autoradiograms was performed using the Decodon Delta 2D software.
Proteins that are synthesized at increased levels in response to FA or MG stress in at least two independent experiments are indicated by
white labels. Their respective induction ratios are listed in Table S1. Spot identification was performed using MALDI-TOF-TOF mass
spectrometry from Coomassie-stained 2D gels as described in Experimental procedures.
Similarly, FA causes cross-links between proteins and
DNA, which results in the generation of secondary electrophiles which in turn could lead to ROS production
(Orlando et al., 1997). It is also possible that aldehydes
react with functionally important Cys residues in PerR
although the four Cys residues in PerR co-ordinate a
tightly bound zinc ion and are known to be generally
non-reactive with peroxides (Lee and Helmann, 2006).
FA and MG induce metal-ion efflux systems
(ArsR, CsoR, CzrA regulons)
Furthermore, several cation efflux systems were strongly
upregulated by carbonyl stress. These include the ArsRregulated arsR–yqcK–arsBC operon (Moore et al.,
2005), the CzrA-regulated czcDtrkA operon, and the
CsoR-regulated copZA operon (Smaldone and Helmann,
2007) (Fig. 4). These metal efflux systems are regulated
by metal-sensing repressors that all have exposed cysteine residues as part of their metal-binding sites, which
might be reactive with aldehydes (Morby et al., 1993;
Harvie et al., 2006; Liu et al., 2007; Chen and He,
2008).
FA and MG induce motility and chemotaxis
(SigmaD regulon)
FA and MG also lead to induction of many genes of the
SigmaD regulon that are involved in motility, chemotaxis
and cell wall turnover (Serizawa et al., 2004). These
include the flgKLM, fliDST and motAB operons for assembly of flagella and genes encoding methyl-accepting
chemotaxis proteins (mcpABC, tlpAB, yfmS, yoaH, yvaQ).
Electrophiles could act as repellents which are sensed by
methyl-accepting chemotaxis proteins to enable cells to
avoid these toxic compounds (Garrity and Ordal, 1995;
Szurmant and Ordal, 2004).
FA and MG cause DNA damage (LexA regulon)
FA and MG induced the DNA damage responsive SOS
(LexA) regulon. The SOS regulon includes for example
the uvrB, uvrC and recA genes encoding DNA repair
and recombination systems and also the yhaZ gene
encoding a DNA alkylation repair enzyme (Au et al.,
2005). DNA damage likely results from the known ability
of FA and MG to cause protein–DNA cross-links, for
example between the amino groups of lysine and
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
Regulation of carbonyl detoxification by AdhR 881
MG
FA
Co 2.8 5.5 1
CtsR
2.66
MG
2.3 kb
FA
Co 2.8 5.5 1
mM
clpE
1.82
1 82
1.52
1.82
1.52
mM
1.2 kb
1.05
hxlAB
HxlR
1.05
2.66
Spx
2.0 kb
1.82
1
52
1.52
1.05
0.7 kb
ArsR
nfrA
2.66
3.6kb
yrhE-yrhD
1.82
2.66
1.82
1.52
1.05
4.7
2.66
0.58
0.48
CymR
nfrA-ywcH
1.0 kb
cysK
2.2 kb
1.82
1.52
1 82
1.82
1.52
2.0 kb
yrhFG-yrzI
1.3 kb
1.05
arsR-yqcKarsBC
1.05
2.66
PerR
1.82
1.52
1.3 kb
katA
1.05
Fig. 4. Transcriptional analysis of selected genes that are strongly induced in the microarray analyses by FA and MG. Transcript analysis
of clpE, nfrA, cysK, arsB and katA indicates the induction of the CtsR, Spx, CymR, ArsR and PerR regulons by FA and MG, which overlaps
with the response to diamide and quinone-like electrophiles (Antelmann et al., 2008). Transcription of the HxlR-controlled hxlAB operon is
specifically induced by aldehydes. The formate dehydrogenase operon yrhED and the formate transporter operon yrhFG–yrzI respond
specifically to MG. The arrows point towards the size of the specific transcripts.
cytosine (Orlando et al., 1997; Kalapos, 2008). Similar
responses of DNA damage-inducible enzymes were also
shown in genome-wide expression profiling for FA in
P. putida (Roca et al., 2008).
MG selectively induces genes for formate metabolism
and transport
Exposure of cells to MG leads to the selective induction of
an uncharacterized formate dehydrogenase (YrhE) and a
formate transporter (YrhG) encoded by the yrhED and
yrhFG–yrzI operons respectively (Fig. 4). This response
could be caused by contamination of commercially available MG with formate (Kalapos, 1999). However, detailed
1
H nuclear magnetic resonance (1H-NMR) analysis confirmed no formate traces in MG and a composition of 88%
MG, 1.7% acetic acid, 2.1% methanol and 4.7% propanediol (data not shown).
Genes repressed by MG and FA
Genes involved in ATP generation were repressed under
both FA and MG stress conditions, perhaps as a result of
slower growth (Table S2). Exposure to MG caused downregulation of many transition phase regulons, such as
CodY, TnrA, sH, Rok and AbrB (Ratnayake-Lecamwasam
et al., 2001; Molle et al., 2003; Yoshida et al., 2003;
Hamon et al., 2004; Albano et al., 2005; Sonenshein,
2007). The Fur regulon was selectively repressed by FA
treatment for reasons that are not yet clear.
Hierarchical clustering analyses for electrophilic
compounds (aldehydes, quinones and diamide)
To visualize the overlapping expression of genes and
regulons by all thiol-reactive electrophiles, we performed
a hierarchical clustering analysis using transcriptomic
data of B. subtilis cells exposed to diamide (Leichert et al.,
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
882 N. T. T. Huyen et al. !
2003), methylhydroquinone (MHQ) (Duy et al., 2007), catechol (Tam et al., 2006), MG and FA (Fig. 5). For the
cluster analysis we chose a subset of 271 genes, which
are > 3-fold induced by any of these electrophiles. The
complete cluster shows ten major groups of genes which
are arranged as nodes with similar expression profiles.
The common induction of the general electrophile stress
responsive CtsR, Spx, CymR regulons, the metal ion
stress responsive ArsR, CzrA and CsoR regulons and the
SigmaD regulon by all thiol-reactive electrophiles is
reflected in nodes 6, 7, 2 and 3. The induction of genes
regulated by YodB, YvaP (CatR) and MhqR is more specific for quinones and diamide than for FA and MG (nodes
4 and 10). FA and MG lead to selective induction of the
HxlR, AdhR and LexA regulons (node 5). The specific
induction of the yrhED and yrhFG–yrzI operons by MG is
visualized in node 1.
FA and MG induce specific aldehyde detoxification
systems (HxlR and AdhR regulons)
Our cluster analysis revealed also a small subset of genes
that were selectively induced by both carbonyls (FA and
MG) but not by other thiol-reactive electrophiles (Fig. 5,
node 5). These include the hxlAB operon that encodes the
genes for the RuMP pathway as major route for FA detoxification in B. subtilis (Yurimoto et al., 2005). We observed
very high inductions of the hxlAB operon (20- to 140-fold)
and hxlR (3- to 10-fold) by both MG and FA (Fig. 4) and
the HxlA protein was identified also in the proteome
(Fig. 3).
In addition, the cluster analyses revealed also the
aldehyde-specific induction of the adhA gene (10- to
40-fold) (Fig. 5, node 5) encoding a NADH-dependent
alcohol dehydrogenase (Table S1). AdhA is similar to
GSH-dependent Fdhs, which are evolutionarily conserved
among prokaryotic and eukaryotic organisms (Harms
et al., 1996). The conserved S-nitrosoglutathione (GSNO)
reductase activity of Fdh-like enzymes has been first
demonstrated in E. coli, Saccharomyces cerevisiae and
mouse macrophages (Liu et al., 2001). Furthermore, Adhlike enzymes played the key role in the defence against
nitric oxide (NO) stress in Haemophilus influenzae (Kidd
et al., 2007) and in virulence in Streptococcus pneumoniae (Stroeher et al., 2007). Fdh-like enzymes that function as mycothiol-dependent NO reductases have been
also characterized in Mycobacterium smegmatis (Vogt
et al., 2003). In contrast, B. subtilis AdhA was induced
specifically by FA and MG, but not under conditions of
NO stress (Moore et al., 2004). We hypothesize that
AdhA functions in the thiol-dependent detoxification of
aldehydes. The AdhA protein usually works together with
the S-formylthiol hydrolase encoded by homologues of
the human estD gene that converts S-formylthiol to
formate (Jensen et al., 1998) (Fig. 1B). However, genes
with similarity to estD are not encoded in the B. subtilis
genome.
In addition to adhA, the upstream-encoded yraB and
yraC genes are strongly upregulated by MG and FA. The
yraC gene was removed from the SubtiList database (previous accession number BG 13778) but is still included
in our microarrays. We have reannotated the yraC gene
as a 192 bp open-reading frame encoding a putative
g-carboxymuconolactone decarboxylase (CMD) (Fig. S1A
and B). However, YraC is much smaller than other annotated CMD family enzymes. CMD domain enzymes are
involved in protocatechuate catabolism and in some bacteria a gene fusion event leads to expression of CMD with
a hydrolase involved in the same pathway (Eulberg et al.,
1998). In Methanosarcina acetivorans a CMD domain
enzyme had protein disulphide reductase activity, which
depends on a CxxC motif (Lessner and Ferry, 2007).
CMD-like domains are also found in the alkylhydroperoxide reductase AhpD of mycobacteria (Bryk et al., 2002).
The yraB gene encodes a MerR family regulator of the
NmlR subfamily that all regulate the expression of adhAlike genes (Kidd et al., 2005). As we here show that YraB
regulates induction of adhA, we rename YraB as AdhR.
NmlR of Neisseria gonorrhoeae is a redox-controlled
regulator of the GSNO reductase AdhC, a Cpx-type
ATPase, and a thioredoxin reductase and protects against
NO stress (Kidd et al., 2005). NmlR-like regulators confer
also protection against NO stress in H. influenzae and
S. pneumoniae (Kidd et al., 2007; Stroeher et al., 2007).
NmlR homologues (including AdhR) share a conserved
Cys52 residue (B. subtilis numbering) that could play a
role in sensing of NO stress or aldehydes. However, the
detailed mechanisms of transcriptional regulation for
NmlR-like regulators under conditions of NO or FA stress
have thus far not been described.
The novel MerR/NmlR-type regulator AdhR (YraB)
positively regulates the adhA–yraA operon and yraC in
response to aldehydes
The adhA gene is located upstream of yraA, which
encodes a DJ-1/PfpI-family protein, some of which may
function as cysteine proteinases. Although their functions
are not well understood, structures have been resolved
for human DJ-1, yeast Hsp31p, and four E. coli homologues YhbO, SCRP 27a, YajL and Hsp31 (Wei et al.,
2007). They all possess a conserved Cys residue in the
nucleophile elbow motif. Recently, Hsp31p has been
shown to be involved in the protection against ROS
(Skoneczna et al., 2007). Previous microarray results
revealed that yraA is very strongly upregulated by
diamide, quinones, acid, ethanol and vancomycin in a
Spx-dependent manner (Nakano et al., 2003; Thackray
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
SigmaD
YodB,YvaP,
MhqR, PerR
HxlR,
AdhR,
LexA
CymR,
Spx
CtsR,
Spx
Spx,
S
HrcA
2
3
4
5
6
7
8
MG/FA >
MHQ/Cat
5
4
1
MG >
FA/MHQ/Cat
10
6
7
Electrophile stress response
3
2
Fig. 5. Hierarchical clustering analysis of gene expression in response to electrophiles in B. subtilis. The treatment with the electrophiles includes diamide (Dia), catechol (Cat),
methylhydroquinone (MHQ), formaldehyde (FA) and methylglyoxal (MG). Genes expression data were clustered based on the induction ratios leading to ten defined groups (nodes). Nodes
enriched for genes that belong to the electrophile stress responsive regulons (CtsR, Spx, CymR, ArsR, CzrA, CsoR, SigmaD), the quinone-responsive regulons [PerR, YodB, CatR (YvaP),
MhqR] and the aldehyde-responsive regulons (HxlR, AdhR, LexA) are shown to the right of the cluster. Red indicates induction and green repression under the specific conditions of
electrophile stress (see Experimental procedures for details).
9
10 MhqR
ArsR,
CzrA,
CsoR,
SigmaD
Spx
1
MHQ/Cat >
MG/FA
Regulation of carbonyl detoxification by AdhR 883
884 N. T. T. Huyen et al. !
Fig. 6. Transcript analyses (A), transcriptional organization (B) and promoter alignments of the adhR, yraC and adhA–yraA operons which
are regulated by the MerR-type transcriptional regulator AdhR in response to carbonyl compounds.
A. For Northern blot experiments 5 mg of RNA each was isolated from the B. subtilis strains before (co) and 10 min after exposure to 1 mM
FA, 2.8 mM and 5.5 mM MG. Northern blots were hybridized with yraA, adhA or adhR-specific mRNA probes (shown underlined). The arrows
point towards the sizes of the adhA–yraA, yraA and adhR specific transcripts.
B. Gene organization of adhR, yraC and adhA–yraA. Transcriptional start sites are indicated by bent arrows.
C. The transcription start sites of the adhR, yraC and adhA–yraA specific mRNAs were determined by 5′ RACE and are indicated as +1. The
-10 and -35 promoter elements are underlined. The conserved inverted repeat sequences are shown in upper case letters.
and Moir, 2003; Jervis et al., 2007; Antelmann et al.,
2008; Eiamphungporn and Helmann, 2008).
To study the regulation and transcriptional organization
of adhA and yraA, we performed Northern blot experiments using RNA isolated from B. subtilis wild-type and
adhR mutant cells after exposure to FA and MG (Fig. 6A).
Two transcripts were detected using a yraA-specific RNA
probe, both of which were strongly elevated under conditions of aldehyde stress in the wild-type. The larger 1.8 kb
transcript corresponds to the adhA–yraA bicistronic transcript, which was also detected using an adhA specific
RNA probe. This adhA–yraA mRNA is not induced in the
adhR mutant by aldehydes confirming that AdhR activates
transcription of the adhA–yraA operon. The smaller 0.5 kb
transcript originates from a promoter that precedes yraA
and is induced in both the wild-type and adhR mutant
strain. Thus, transcription of yraA is induced monocistronically in a Spx-dependent manner and as part of the bicistronical AdhR-controlled adhA–yraA message selectively
by aldehydes. Finally, using an adhR-specific RNA probe,
the autoregulation of adhR was verified by Northern blot
analyses.
Identification of a conserved inverted repeat in the
adhA, adhR and yraC promoter regions
Next, we used in silico analyses to identify conserved
binding sites for the AdhR regulator upstream of adhA
and adhR. A conserved 7-4-7 inverted repeat (IR) with
the consensus sequence CTTAAAG-N4-CTTTAAG was
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
Regulation of carbonyl detoxification by AdhR 885
[AdhR] nM
0
5 10 20 40 80 160
MG
10 mM FA
10 mM MG
Control
0
5
10
20 40
80 160 0
5
10 20 40
80 160
Fig. 7. Binding of purified AdhR to the adhR,
yraC and adhA promoters using DNA
electrophoretic mobility shift assay. Increasing
amounts of purified AdhR protein were added
to the labelled adhR (A), yraC (B) and adhA
(C) promoter probes. Approximately 10 mM of
MG or FA was added to analyse the effect of
these compounds on the DNA binding activity
of AdhR to these target promoters. (D) AdhR
does not bind to an unrelated promoter
fragment (the yoeB promoter region).
FA
0
5 10
0
5 10
160 160 160 160 160 160
mM
[AdhR] nM
identified in the promoter regions of adhA and adhR. This
consensus was used to search the genome sequence of
B. subtilis using the search pattern function of the SubtiList database. In addition to the adhA and adhR promoters, this conserved IR sequence was detected upstream
of yraC (Fig. 6B). Transcription of yraC was also induced
under conditions of aldehyde stress as revealed by the
microarray data.
The transcription start sites of adhA, adhR and yraC
were mapped using 5′ RACE to positions 34, 42–43
and 82–83 nt upstream of their respective start codons
(Fig. 6B). The -10 and -35 elements of all three promoter
regions were separated by 19 bp consistent with the elongated spacer regions typical for target promoters for MerR
family regulators (Brown et al., 2003; Hobman et al.,
2005). The conserved IR sequence overlapped slightly
the -35 promoter region. These conserved binding sites
and the MerR-type promoter architecture suggest that
adhR, yraC and adhA–yraA constitute the aldehydeinducible AdhR regulon of B. subtilis.
AdhR binds specifically to the adhA, adhR and yraC
promoter regions
Electrophoretic mobility shift assays (EMSAs) were performed using purified His-tagged AdhR protein and adhA,
adhR and yraC promoter probes. AdhR bound with high
affinity to all three target promoter regions, but not to a
control DNA fragment (Fig. 7). Like other MerR family
members, AdhR binds to its target operator sequences
both with and without specific inducer and addition of
MG or FA had little effect on the AdhR DNA binding
activity.
Transcriptional activation of the adhA–yraA operon
requires the conserved Cys52 of AdhR
All AdhR homologues share a conserved Cys52 residue
(Kidd et al., 2005). To analyse the role of Cys52 in regulation of adhA–yraA transcription by AdhR, we constructed adhR mutant strains that ectopically expressed
a functional FLAG-tagged AdhR or the AdhRC52A
mutant protein using a xylose-inducible promoter (Pxyl)
and an integrational plasmid vector (Bhavsar et al.,
2001). The adhR and adhRC52A complemented DadhR
mutant strains were grown in minimal medium and
expression from the Pxyl promoter was induced by addition of 2% xylose (Fig. 8A). As activation of adhA–yraA
transcription requires aldehydes, cells were treated with
2.8 mM MG or 1 mM FA for 10 min after xylose addition.
The isolated RNA was subjected to adhA-specific Northern blot analyses. The results showed that adhA–yraA
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
886 N. T. T. Huyen et al. !
adhA-yraA
B
Fig. 8. The conserved Cys52 of AdhR is essential for activation of
adhA–yraA transcription by aldehydes.
A. For adhA-specific Northern blot analysis, RNA was isolated from
strains before (co) and after treatment with 2% xylose for 20 min
and subsequently with 2.8 mM MG or 1 mM FA for 10 min.
B. Western blot analysis was performed using Anti-FLAG antiserum
to confirm expression of FLAG–AdhR and FLAG–AdhRC52A
proteins.
transcription is similar strongly activated by MG and FA
in wild-type cells and in the adhR–FLAG-complemented
adhR mutant strain (Fig. 8A). However, no expression
of adhA–yraA occurred in the adhRC52A mutant in
response to aldehyde treatment. Western blot analyses
confirmed that FLAG–AdhR and FLAG–AdhRC52A proteins are produced in similar amounts in the adhR
and adhRC52A-complemented DadhR mutant strains
(Fig. 8B). These results indicate that Cys52 is required
for activation of transcription by AdhR, presumably
because aldehydes modify this conserved Cys residue
via thiol-(S)-alkylation to allosterically activate AdhR.
In contrast to AdhR, the MerR/NmlR regulator of
N. gonorrhoeae acts as a transcriptional repressor of
adhC transcription under normal conditions (Kidd et al.,
2005). Repression by NmlR requires Zn(II) and transcription of adhC is derepressed by diamide. Mutation of any of
the four Cys residues in NmlR resulted in constitutive
repression and lack of activation of adhC transcription
(Kidd et al., 2005).
1 mM FA
0.6
0.4
wt
∆hxlR
yfkM
∆yraAy
0.2
adhA
∆a
Previous analyses showed that the DhxlR deletion mutant
was sensitive to FA exposure (Yurimoto et al., 2005). To
determine to which extent the RuMP pathway and the FA
dehydrogenase pathway confer resistance to carbonyl
compounds, growth experiments were performed using
DhxlR, DadhR, DyraA and DadhA mutants. All mutant
strains showed the same growth rates like the wild-type in
the absence of FA and MG challenge (data not shown).
However, the growth rates of the DhxlR and DadhA
mutants were significantly reduced in the presence of 0.5
and 1 mM FA compared with wild-type cells (Fig. 9). This
indicates that both FA detoxification systems contribute to
FA resistance of B. subtilis. However, neither of the tested
MG
0.8
wt
The DhxlR, DadhA and DyraAyfkM mutants are sensitive
to FA stress
growth rate (h-1)
0.5 mM FA
1
wt
1.8 kb
A
yfkM
∆yraAy
Co MG FA Co MG FA Co MG FA Co MG FA
mutants showed an increased sensitivity to methylglyoxal
(data not shown).
The adhB gene encodes an adhA-paralogous alcohol
dehydrogenase which is part of the sG–controlled yraGF–
adhB–yraED operon (Steil et al., 2005) and also induced
by FA and MG (3- to 8-fold) (Table S1). However, no
growth differences were detected for the DadhADadhB
double mutant compared with the DadhA single mutant in
the presence of FA and MG (data not shown). Thus, adhB
does not contribute significantly to aldehyde resistance of
growing B. subtilis cells.
The growth rate of the DyraA mutant was also similar
to that of the wild-type after exposure to FA (data not
shown). As yraA displays sequence similarity to the
sB-dependent yfkM gene, we investigated the growth
phenotype of the DyraADyfkM double mutant. The
DyraADyfkM double mutant showed a reduced growth
rate after treatment with FA and MG compared with the
wild-type and to the yraA single mutant (Fig. 9). Thus,
both putative DJ-1/PfpI-family cysteine proteinases can
replace each other in protection against reactive aldehydes, presumably by repair or degradation of proteins
with specific thiol-modifications. However, according to
previous studies, YraA had no detectable protease or
chaperone activity (Thackray and Moir, 2003). In ongoing
studies, we are using proteomic approaches and radioactively labelled electrophiles to identify possible substrates for the YraA and YfkM proteases in B. subtilis.
Together, our results show that B. subtilis encodes
multiple systems involved in resistance to FA: the HxlRregulated RuMP pathway and the AdhR-controlled thioldependent AdhA and candidate cysteine proteinases
YraA and YfkM.
∆hxlR
∆adhR
∆adhR
adhRC52A
yfkM
∆yraAy
∆adhR
adhR
adhA
∆a
CU1065
Fig. 9. The DadhA and DhxlR mutants are sensitive to FA stress
and the DyraADyfkM mutant is sensitive to FA and MG. B. subtilis
wild-type (wt), DadhA, DyraADyfkM and DhxlR mutant strains were
grown in minimal medium to an OD500 of 0.4 and treated with 0.5
and 1 mM FA or 1.4 mM MG. The growth rates of the strains after
exposure to FA and MG are shown.
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
Regulation of carbonyl detoxification by AdhR 887
A
C
B
cysteine
1.2
10
10
1
1
OD500(nm)
ratio
1
0.8
0.6
0.4
control
FA 2 mM
FA 2 mM + Cys 1 mM
FA 2 mM
M+C
M
Cys 2 mM
FA 2 mM + Cys 5 mM
0.1
0.2
MG 5.5 mM
M
MG 2.8 mM
M
FA 1.0 mM
M
FA 0.5 mM
M
co
0.01
-120 -60
0
60
120 180
240 300
control
MG 3 mM
MG 3 mM + Cys 3 mM
MG 3 mM + Cys 6 mM
0.1
0.01
Time (min)
-120 -60
0
60
120 180 240 300
Time (min)
Fig. 10. The LMW thiol cysteine protects against aldehyde toxicity in B. subtilis.
A. Changes in the level of cysteine in the metabolome were measured using GC-MS in B. subtilis cellular extracts after exposure to FA and
MG as described in Experimental procedures. Average values and standard deviations are quantified from at least three independent
cultivation experiments and the ratios are related to the untreated control.
B., C. B. subtilis wild-type cells were grown in minimal medium to an OD of 0.4 and different concentrations of extracellular cysteine were
added prior to exposure of FA (B) or MG (C). These experiments indicate that 1 mM cysteine was able to protect against 2 mM FA and 3 mM
cysteine titrates 3 mM MG which restored the growth.
The LMW thiol cysteine functions in detoxification of
FA and MG
In previous studies we found that cysteine functions in
detoxification of quinone-like electrophiles (Liebeke et al.,
2008). The results of this study also suggest that cysteine
is an important target for toxic aldehydes. We measured
the intracellular concentration of cysteine in the metabolome after exposure to FA and MG using GC/MS
analyses. Levels of cysteine were decreased about fivefold after exposure to 0.5–1 mM FA and 5.5 mM MG in
B. subtilis (Fig. 10A). In contrast to quinone treatment,
other metabolites were not changed by FA and MG
exposure.
To determine if extracellular cysteine can protect cells
against aldehyde toxicity, we pretreated cells with cysteine prior to FA and MG challenge and monitored
growth (Fig. 10B and C). The results showed that wildtype cells pretreated with 1–2 mM cysteine were protected against the toxicity of 2 mM FA and growth was
restored (Fig. 10B). Also, pretreatment of cells with
3 mM cysteine suppressed the growth inhibitory effect of
3 mM MG (Fig. 10C). These experiments showed that
equimolar amounts of cysteine are able to titrate toxic
aldehydes, consistent with the notion that electrophilic
aldehydes target cysteine. The microarray data showed
the strong upregulation of an uptake system for cystine
perhaps to replenish the intracellular thiol pools.
However, pretreatment of the cells with cystine was not
protective against subsequent FA challenge (data not
shown). This might indicate that aldehydes react with the
added thiol compounds outside the cell. By using
1
H-NMR analysis we confirmed that FA reacts with cys-
teine in vitro resulting in a thiazolidine carboxylic acid
(Fig. S2A). The reaction of MG with cysteine leads to
generation of the acetylated derivative of the thiazolidine
carboxylic acid (Fig. S2B).
The transcriptomic results revealed DNA damage by
aldehydes, which indicates that aldehydes also cause
cross-links between the side-chains of lysine or arginine
and nucleic acids. However, unlike Cys, pretreatment of
cells with arginine and lysine did not restore growth after
addition of toxic MG or FA concentrations (data not
shown). These results suggest that Cys is the strongest
nucleophile, which is targeted in vivo by aldehydes.
Conclusion
In this study we have analysed the global response of
B. subtilis to the electrophilic carbonyls FA and MG. The
global expression profiles suggest that carbonyls act, like
diamide and quinones, as thiol-reactive electrophiles,
which most probably target cysteine residues via the thiol(S)-alkylation chemistry. Thus, cysteine is also one of the
first barriers for electrophilic aldehydes as supported
by protection experiments with extracellular cysteine.
However, in contrast to quinones and diamide, carbonyl
electrophiles display also a unique genetic response,
which is governed by the HxlR and AdhR regulators. Both
the HxlR and AdhR regulons confer resistance to FA.
These pathways and protection mechanisms against FA
exposure that were predicted and demonstrated in this
study are summarized in a schema in Fig. 11.
AdhR is further characterized here as novel regulator
of the MerR/NmlR-family, which positively controls a
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
888 N. T. T. Huyen et al. !
Fig. 11. Protective mechanisms against formaldehyde toxicity in B. subtilis. Exposure of B. subtilis to FA induces general electrophile-stress
responsive regulons (CymR, Spx, CtsR), the peroxide specific PerR regulon, metal ion-efflux regulons (ArsR, CsoR, CzrA), the DNA damage
inducible LexA regulon and aldehyde detoxification regulons (AdhR, HxlR). Pathways that have been demonstrated in this study are shown
with solid arrows and predicted pathways are displayed with dashed arrays. The AdhR and HxlR regulons for specific detoxification are
marked in yellow and the other FA-inducible stress regulons are marked in red.
(1). FA reacts with low-molecular-weight thiols (R-SH) to S-hydroxymethylthiol, which is converted to S-formylthiol by the thiol-dependent
AdhR-controlled AdhA and further by an unknown hydrolase to formate.
(2). S-formylthiols could be also exported via the metal ion efflux systems such as ArsBC, CopA, CzcD, which are upregulated by FA.
(3). In addition, FA is detoxified via the RuMP pathway by HxlA generating hexulose-6-phosphate (H6P) and HxlB producing
fructose-6-phosphate (F6P).
(4). The reaction of FA with thiol buffers (e.g. cysteine) results in depletion of reduced LMW thiols and an imbalanced thiol-redox homeostasis.
This leads to derepression of the CymR regulon to increase cysteine biosynthesis.
(5). Depletion of reduced thiol buffers also causes induction of the Spx regulon to induce thiol-disulphide reducing systems (TrxAB) and
restore the thiol-redox balance.
(6). FA causes DNA–protein cross-links resulting in DNA damage and induction of the LexA regulon to repair DNA damages.
(7). FA reacts also with protein thiols leading to protein cross-links or S-hydroxymethyl modifications in proteins. These modified or aggregated
proteins are proteolytically degraded by the CtsR-regulated ClpCP machinery and repaired or degraded by the YraA cysteine proteinase.
(8). The reaction of FA with DNA and proteins could generate also secondary reactive intermediates that in turn could produce ROS which
lead to the induction of PerR-regulated oxidative stress defence enzymes.
conserved thiol-dependent Fdh (AdhA) and the putative
cysteine proteinase YraA, both of which protect against
FA toxicity. Transcriptional regulation by AdhR requires
the conserved Cys52 residue which we hypothesize
is alkylated by electrophilic carbonyl compounds
leading to transcriptional activation of the AdhR-dependent
adhA–yraA, yraC and adhR operons. Future studies are
directed to define the nature of the post-translational thiolmodifications, which are caused by aldehydes in AdhR as
well as in cellular proteins in vivo. These might involve
S-hydroxymethylcysteine modifications or cross-links
between different thiol-containing proteins by FA which
remains to be elucidated.
Experimental procedures
Bacterial strains and growth conditions
The bacterial strains used were B. subtilis 168 (trpC2) and
CU1065 (trpC2), DadhA (trpC2,adhA::mls r), DadhADadhB
DyraA
(trpC2,yraA::cm r),
(trpC2,adhA::mls radhB::spc r),
DadhR
DyraADyfkM
(trpC2,yraA::cm ryfkM::mls r),
(trpC2,adhR::km r), which are described below. B. subtilis
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
Regulation of carbonyl detoxification by AdhR 889
strains were cultivated under vigorous agitation at 37°C in
Belitsky minimal medium described previously (Stülke et al.,
1993). E. coli strains were grown in LB for DNA manipulation.
The antibiotics were used at the following concentrations:
1 mg ml-1 erythromycin, 25 mg ml-1 lincomycin, 5 mg ml-1
chloramphenicol and 10 mg ml-1 kanamycin. Methylglyoxal
was purchased from Sigma Aldrich and FA was ordered from
Roth.
Gene deletions were generated using long-flankinghomology polymerase chain reaction (LFH-PCR) as previously described (Mascher et al., 2003). Primers used to
amplify the up- and down-fragments are listed in Table S3.
Fragments were amplified using Pfu DNA polymerase (Stratagene) and were joined using Expand Long Template PCR
System (Roche). Five microlitres of the LFH-PCR product
was introduced into the desired strain by transformation. Integration and deletion of the gene were confirmed by PCR.
Construction of adhR mutant containing adhR–FLAG
and adhRC52A–FLAG under PxylA control was performed
using the pSWEET plasmid (a derivative of pDG364 containing PxylA promoter and chloramphenicol resistance gene)
(Bhavsar et al., 2001). To generate pSWEET containing
adhR–FLAG, adhR (including the predicted RBS site
upstream of adhR) was fused in frame with a sequence
encoding the FLAG epitope tag and amplified by PCR using
primers 4151/4152 as listed in Table S3. The resulting fragment was digested with PacI and BamHI and cloned into
pSWEET that was digested with the same enzymes, placing
adhR–FLAG downstream of the PxylA promoter. The
pSWEET containing adhRC52A–FLAG was generated by
PCR mutagenesis. Using pSWEET containing adhR–FLAG
as a template, first-round PCR was performed in two separate reactions with primers 4151/4175 and 4152/4170
(Table S3). The PCR products were joined by a second round
of PCR using primers 4151/4152. The product was digested
with PacI and BamHI and cloned into pSWEET that was
digested with the same enzymes. The resulting plasmids
were verified by DNA sequencing then digested with PstI and
transformed into adhR::Km strain where the fragments integrated into the amyE locus. The resulting strains were
selected by KmrCmr and confirmed by PCR.
Proteome analysis and mass spectrometry
Cells grown in minimal medium to an OD500 of 0.4 were
pulse-labelled for 5 min each with 5 mCi of L-[35S]methionine
per ml before (control) and 10 min after exposure to 1 mM
FA, 2.8 mM or 5.6 mM MG. Preparation of cytoplasmic
L-[35S]methionine-labelled proteins and separation by twodimensional gel electrophoresis (2D-PAGE) using the immobilized pH gradients (IPG) in the pH range 4–7 was perfomed
as described (Tam et al., 2006). The quantitative image
analysis was performed with the DECODON Delta 2D software (http://www.decodon.com). Proteins showing an induction of at least twofold during the 5 min of L-[35S]methionine
pulse in two independent experiments were considered as
significantly induced. For identification of the proteins by
mass spectrometry, non-radioactive protein samples of
200 mg were separated by 2D-PAGE and the 2D gels were
stained with Colloidal Coomassie brilliant blue (Amersham
Biosciences). Spot cutting, tryptic digestion of the proteins
and spotting of the resulting peptides onto the MALDI-targets
(Voyager DE-STR, PerSeptive Biosystems) were performed
using the Ettan Spot Handling Workstation (AmershamBiosciences, Uppsala, Sweden) as described previously
(Eymann et al., 2004). The MALDI-TOF-TOF measurement
of spotted peptide solutions was carried out on a ProteomeAnalyzer 4800 (Applied Biosystems, Foster City, CA, USA) as
described previously (Eymann et al., 2004).
Transcriptome analysis
For microarray analysis, B. subtilis wild-type cells were grown
in minimal medium to OD500 of 0.4 and harvested before
and after exposure to 1 mM FA, 2.8 mM and 5.6 mM MG
respectively. Total RNA was isolated by the acid phenol
method as described (Majumdar et al., 1991). Generation of
fluorescence-labelled cDNA and hybridization with B. subtilis
whole-genome microarrays (Eurogentec) was performed as
described previously (Jürgen et al., 2005). Genes showing
induction or repression ratios of at least threefold in three
independent experiments were considered as significantly
induced. The averages ratios and standard deviations for all
induced or repressed genes are calculated from three independent transcriptome experiments after 10 min of exposure
to 2.8 and 5.6 mM MG or 1 mM FA and listed in Table S4 and
S5 (http://microbio1.biologie.uni-greifswald.de/publications.
html). All microarray datasets are available in the GEO database under accession numbers [GSM350786-GSM350794].
Hierarchical clustering analysis
Clustering of gene expression profiles in response to different
electrophiles was performed using Cluster 3.0 (de Hoon
et al., 2004). The transcriptome datasets were derived either
from previous publications or this study and included log2fold expression changes 10 min after exposure of B. subtilis
to the electrophilic compounds diamide (1 mM) (Leichert
et al., 2003), MHQ (0.33 mM) (Duy et al., 2007), catechol
(2.4 mM) (Tam et al., 2006), FA (1 mM) and MG (5.6 mM).
The complete dataset of all induced genes under these different conditions of electrophile treatment is listed and classified into regulons in Table S6 (http://microbio1.biologie.unigreifswald.de/publications.html). After hierarchical clustering,
the output was visualized using TreeView (Eisen et al., 1998).
For the clustering as shown in Fig. 5 we have used 271
selected genes that are induced by any of these electrophilic
compounds in B. subtilis [e.g. CtsR, CymR, Spx, PerR, ArsR,
CsoR, CzrA, YodB, YvaP (CatR), MhqR, LexA, SigmaD,
AdhR and HxlR regulons].
Northern blot experiments
Northern blot analyses were performed as described
(Wetzstein et al., 1992) using RNA isolated from B. subtilis
wild-type cells before (control) and 10 min after the treatment
with 1 mM FA, 2.8 mM and 5.6 mM MG respectively. Hybridizations specific for adhA, yraA, adhR, clpE, nfrA, cysK, katA,
yqcK, hxlA, yrhE and yrhG were performed with the
digoxigenin-labelled RNA probes synthesized in vitro using
T7 RNA polymerase from T7 promoter containing internal
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
890 N. T. T. Huyen et al. !
PCR products of the respective genes using the primer sets
listed in Table S3.
Western blot analysis
Anti-FLAG polyclonal rabbit antiserums (SIGMA) were used
at a dilution of 1:3000 to analyse the cytoplasmic protein
amounts of FLAG–AdhR and FLAG–AdhRC52A proteins produced in the AdhR and AdhRC52A complemented adhR
mutant strains after induction with 2% Xylose. Fifty micrograms of each protein sample was loaded onto a 12% SDSPAGE gel and the Western blot analysis was performed as
described previously (Liebeke et al., 2008).
5! RACE-PCR
The transcription start sites of adhR, yraC and adhA genes
were mapped. Total RNA was prepared from B. subtilis
CU1065 grown aerobically in LB medium with and without the
addition of 2 mM methylglyoxal. The cell cultures were grown
to an OD600 of 0.4 and split into two flasks with equal volume,
methylglyoxal was added to one flask to a final concentration
of 2 mM and cells were harvested 20 min after treatment. The
RNeasy mini kit (Qiagen) was used as a method to extract
total RNA from the cell lysates. The RNA concentration
was quantified by NanoDrop spectrophotometer (Nanodrop
Tech., Welmington, DE). The 2 mg of total RNA was used
as a template for reverse transcription using MultiscribeTM
Reverse transcriptase (Taqman, Roche) and a gene-specific
primer (GSP1). The resulting cDNA was purified by gel
extraction purification column (Qiagen) and added poly(dC)
tail at their 3′-ends with terminal deoxynucleotidyl transferase
(New England Biolabs). The resulting cDNA was amplified by
PCR using the poly-dG primer to anneal at the poly(dC) tail
and a second gene-specific primer (GSP2), complementary
to a region upstream of the GSP1 primer. PCR products were
separated by gel electrophoresis and sequenced.
Expression and purification of the AdhR protein
AdhR protein was purified using PrepEaseTM His-Tagged
High Yield purification Resin (USB). The adhR gene was PCR
amplified from B. subtilis chromosomal DNA with oligonucleotides, designed to engineer an NdeI site upstream and a
HindIII site downstream of the adhR gene. The PCR product
was cloned into pET16b (Novagen) via the NdeI and HindIII
sites. The sequence of adhR in pET16 was verified by DNA
sequencing. E. coli BL21/DE3 containing this plasmid was
grown to mid-logarithmic phase at 37°C in 1 l of LB medium
and 100 mg of ampicillin per ml. Isopropylthiogalactopyranoside (IPTG) was added to 1 mM and cells were harvested
after further incubation for 2 h. After centrifugation, the pellet
was resuspended in 5 ml LEW (Lysis-Equilibration-Wash)
buffer (50 mM NaH2PO4, 300 mM NaCl pH 8.0) and
sonicated. After centrifugation, the supernatant was transferred to a clean tube and then incubated for 15 min at room
temperature (RT) with 1 g PrepEase Ni2+-IDA affinity resin
(USB). The protein-bound resin was transferred to a column
and washed extensively with LEW buffer. AdhR was eluted by
adding 10 ml of 50 mM NaH2PO4, 300 mM NaCl and 250 mM
imidazole pH 8.0 to a column. Fractions of 1 ml were collected and tested for protein content using Protein Assay Dye
(Bio-Rad) before pooling for subsequent purification. The
protein was concentrated by ammonium sulphate precipitation and further purified by chromatography on an FPLC
Superdex-200 column in 10 mM Tris-HCl pH 8.0, 50 mM
NaCl, 1 mM DTT, 0.1 mM EDTA and 10% glycerol buffer
followed by dialysis into 10 mM Tris-HCl pH 8.0, 50 mM
NaCl, 1 mM DTT, 0.1 mM EDTA and 50% glycerol buffer and
stored at -80°C.
DNA electrophoretic mobility shift assays
DNA fragments containing the promoter regions of adhR,
yraC, adhA and control yoeB were generated by PCR.
Approximately 50 ng of purified PCR products were endlabelled using T4 polynucleotide kinase (New England
Biolabs) and 50 mCi of [g-32P]-ATP. The labelled DNA probes
were purified by gel extraction purification column (Qiagen)
and 5000 c.p.m. of each probe was incubated with different
amount of purified AdhR protein for 5 min at RT in EMSAbinding buffer (20 mM Tris-HCl pH 8.0, 50 mM NaCl, 5%
glycerol, 5 mg ml-1 Salmon sperm DNA and 50 mg ml-1 BSA).
The binding reactions were incubated in the presence of MG
or FA (final concentration 10 mM) for 20 min at RT. Samples
were separated by 6% native polyacrylamide gel electrophoresis in Tris-borate buffer at pH 8.0 on ice and constant
voltage (100 V) for 2 h. Gels were dried and the radiolabelled
bands were visualized using phosphoimaging.
Determination of the cysteine content in the
metabolome of B. subtilis using GC/MS
Bacillus subtilis cells were grown in minimal medium to an
OD500 of 0.4 and treated with FA (0.5–1 mM) and MG (2.8 and
5.5 mM) for 30 min. Cells were rapidly collected by the fast
filtration method and intracellular metabolites were analysed
using gas chromatography/mass spectrometry (GC/MS) as
described previously (Liebeke et al., 2008).
H-nuclear magnetic resonance-spectroscopy
1
A solution of FA or MG was incubated with equimolar cysteine
for 10 min at 25°C in a 1H-NMR tube (Norell ST-500).
1
H-NMR spectra were recorded on a Bruker 600.27 MHz
Avance-II spectrometer equipped with a BACS-60 sample
changer and controlled with TOPSPIN 2.0 (Bruker, Rheinstetten, Germany). Bruker Icon 1H-NMR software was used
to automate the 1H-NMR data collection. 1H-NMR spectra
were measured with a standard Bruker pulse sequence (noesygppr), solvent presaturation, a sweep width of 12365 Hz,
64K data points at 298.5K. A total of 2 dummy scans and 256
scans were used to obtain the 1H-NMR spectra. All peak
positions were measured relative to the TMSP (1 mM sodium
3-trimethylsilyl-[2,2,3,3-D4]-1-propionic acid) reference peak
set to d = 0.0 p.p.m. AMIX® Bruker Biospin was used for compound identification by matching the obtained spectra with a
1
H-NMR spectra databank.
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 71, 876–894
Regulation of carbonyl detoxification by AdhR 891
Acknowledgements
We thank the Decodon company for support with the
Decodon Delta 2D software, Sebastian Grund for excellent
technical assistance and Dirk Albrecht for mass spectrometry
analysis. We thank Dr Ahmed Gaballa for first noting the
conserved binding sites associated with the AdhR regulon.
We are grateful to Kazuo Kobayashi for providing the hxlR
mutant strain. This work was supported by grants from the
Deutsche Forschungsgemeinschaft, the Bundesministerium
für Bildung und Forschung (BACELL-SysMo 031397A), the
Fonds der Chemischen Industrie, the Bildungsministerium of
the country Mecklenburg-Vorpommern and European Union
grants BACELL-Health (LSHG-CT-2004-503468), BACELLBaSysBio (LSHG-CT-2006-037469) to M.H., and from the
National Science Foundation (grant MCB-0640616) to J.D.H.
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