Báo cáo khoa học: Electron-transfer subunits of the NiFe hydrogenases in Thiocapsa roseopersicina BBS pptx

Electron-transfer subunits of the NiFe hydrogenases in
Thiocapsa roseopersicina BBS
Lı
´via
S. Pala
´
gyi-Me
´
sza
´
ros
1
, Judit Maro
´
ti
2
,Do
´
ra Latinovics
1
,Tı
´mea
Balogh
1
,E
´
va Klement
3
,
Katalin F. Medzihradszky
3

,Ga
´
bor Ra
´
khely
1,2
and Korne
´
l L. Kova
´
cs
1,2
1 Department of Biotechnology, University of Szeged, Hungary
2 Institute of Biophysics, Biological Research Centre, Hungarian Academy of Sciences, Szeged, Hungary
3 Proteomics Research Group, Biological Research Centre, Hungarian Academy of Sciences, Szeged, Hungary
Hydrogenases are metalloenzymes that catalyse the
reversible oxidation of molecular hydrogen according
to the reaction: H
2
M 2H
+
+2e
)
. They can catalyse
the reaction in both directions in vitro, but usually
either evolve or oxidize (take up) H
2
in vivo. The
hydrogenases can be classified according to the metal
content of their active centre: NiFe, FeFe or Fe

hydrogenases [1]. The core of an NiFe hydrogenase
consists of a small electron-transfer subunit and a
large catalytic subunit. Additional proteins are required
for post-translational maturation of the hydrogenase
polypeptides and for connection of the core dimer to
other bioenergetic ⁄ redox processes of the cells. These
accessory hydrogenase-related proteins typically partici-
pate in metallocentre assembly and the transcriptional
regulation of the hydrogenases, and some seem to have
an electron-transfer function [1,2]. The accessory genes
are often located in the close vicinity of hydrogenase
structural genes, but may also be found scattered in
the genome. Numerous microorganisms contain more
Keywords
electron transfer; haem, cytochrome b;
iron–sulfur protein; NiFe hydrogenase;
Thiocapsa roseopersicina
Correspondence
K. L. Kova
´
cs, Department of Biotechnology,
University of Szeged, H-6726 Szeged,
Ko
¨
ze
´
pfasor 52, Hungary
Fax: +36 62 544352
Tel: +36 62 544351
E-mail:

(Received 31 August 2008, revised 6
October 2008, accepted 29 October 2008)
doi:10.1111/j.1742-4658.2008.06770.x
Thiocapsa roseopersicina BBS contains at least three different active NiFe
hydrogenases: two membrane-bound enzymes and one apparently localized
in the cytoplasm. In addition to the small and large structural subunits,
additional proteins are usually associated with the NiFe hydrogenases, con-
necting their activity to other redox processes in the cells. The operon of
the membrane-associated hydrogenase, HynSL, has an unusual gene
arrangement: between the genes coding for the large and small subunits,
there are two open reading frames, namely isp1 and isp2. Isp1 is a b-type
haem-containing transmembrane protein, whereas Isp2 displays marked
sequence similarity to the heterodisulfide reductases. The other membrane-
bound (Hup) NiFe hydrogenase contains the hupC gene, which codes for a
cytochrome b-type protein that probably plays a role in electron transport.
The operon of the NAD
+
-reducing Hox hydrogenase contains a hoxE
gene. In addition to the hydrogenase and diaphorase parts of the complex,
the fifth HoxE subunit may serve as a third redox gate of this enzyme. The
physiological functions of these putative electron-mediating subunits were
studied by disruption of their genes. The deletion of some accessory pro-
teins dramatically reduced the in vivo activities of the hydrogenases,
although they were fully active in vitro. The absence of HupC resulted in a
decrease in HupSL activity in the membrane, but removal of the Isp1 and
Isp2 proteins did not have any significant effect on the location of HynSL
activity. Through the use of a tagged HoxE protein, the whole Hox
hydrogenase pentamer could be purified as an intact complex.
Abbreviation
tat, twin arginine transport.

164 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS
than one hydrogenase. Each enzyme has a specific phys-
iological function, e.g. NAD
+
reduction, electron
removal, H
2
recycling for energy conservation, etc. [3].
The electrons derived from H
2
oxidation are used for
the reduction of the central quinone pool or terminal
electron acceptors, such as fumarate, NO
3
)
or SO
4
2)
.It
is noteworthy that, in spite of their specific expression
and physiological role, one enzyme can take over the
function of another to some extent [4].
Thiocapsa roseopersicina BBS belongs to the family
of purple sulfur photosynthetic bacteria, the Chromati-
aceae [5]. During anoxygenic photosynthesis, this bac-
terium requires reduced sulfur compounds (e.g. S
2)
,S
0
or S

2
O
3
2)
) as electron sources for CO
2
fixation.
T. roseopersicina produces at least three NiFe hydro-
genases (Hyn, Hup and Hox) and contains the genes
of the so-called regulatory hydrogenase (HupUV) [6].
However, their physiological roles are still unclear.
Both the HynS and HupS subunits have a ‘tat’-type
(‘twin arginine transport’) signal sequence; they are
therefore transported through the membrane by the
‘tat’ system [7] and are anchored to the membrane on
the periplasmic side. The Hox enzyme has no signal
for transport across the membrane. Hyn hydrogenase
(formerly Hyd [8]) is a membrane-bound bidirectional
enzyme which has remarkable stability under extreme
conditions; it is extracted from the photosynthetic
membrane as the catalytically active HynSL dimer [9].
The gene arrangement of the hyn operon is unusual:
the genes of the small and large subunits are separated
by a 2-kbp intergenic region. In this section, two open
reading frames, isp1 and isp2, have been recognized
[8]. The putative Isp1 and Isp2 gene products exhibit
remarkable similarity to the DsrK and DsrM subunits,
respectively, of the dissimilatory sulfite reductase com-
plex [10]. Isp1 harbours few transmembrane domains,
and a putative b-type haem-binding site has been pre-

dicted by in silico analysis. In contrast, the putative
Isp2 is a cytoplasmic enzyme resembling the hetero-
disulfide reductases [8]. Similar gene structures can be
found in only a few bacteria, e.g. in Chromatium
vinosum [10] (Accession No. U84760), Aquifex aeolicus
[11], Aquifex pyrophilus [12] and an Archaeon, Acidi-
anus ambivalens [13], but their physiological role has
not been clarified so far.
The other membrane-bound hydrogenase of
T. roseopersicina, HupSL, is encoded in the hupSLCD-
HIR operon [14]. It belongs to the group of uptake
NiFe hydrogenases which recycle H
2
produced by the
nitrogenase complex [15]. As a consequence of the
periplasmic location of the NiFe hydrogenases [16], H
2
oxidation leads to the formation of a proton gradient
which is used for ATP synthesis [9]. Next to the hupSL
genes encoding for the small and large hydrogenase
subunits, the operon contains the hupC gene. In Wolli-
nella succinogenes, strong evidence has been provided
that HupC, a b-type cytochrome [1], can transfer
electrons from the NiFe hydrogenase to the quinones
[17]. Hence, HupC may link the electron transfer from
Hup hydrogenases to the quinone pool.
The third (Hox) hydrogenase has been partially
purified from the soluble fraction of the cells [18]. The
genomic structure of the hox operon suggests a hetero-
pentameric enzyme (HoxEFUYH). The HoxFU

subunits are usually the NAD
+
-reducing part of the
complex, and the HoxYH subunits are responsible for
hydrogenase activity [19]. Recently, a similar enzyme
has been purified and partially characterized from a
closely related strain, Allochromatim vinosum [20]. Hox
hydrogenases are composed of at least four subunits;
the HoxYH and HoxFU dimers form the hydrogenase
and diaphorase catalytic cores, respectively [19]. In sev-
eral cases, additional subunits have also been identi-
fied. In the Hox enzyme of Ralstonia eutropha (which
was purified as a heterotetrameric enzyme for many
years), a new subunit was discovered, and the compo-
sition HoxFUYHI
2
was suggested [21]. In cyanobacte-
ria and the phototrophic bacteria T. roseopersicina and
A. vinosum, the heterotetrameric Hox enzyme is sup-
plemented by a HoxE subunit, which is unrelated to
the HoxI protein [18,20,22]. In T. roseopersicina, it has
been shown previously that in-frame deletion of the
hoxE gene impairs Hox activity in vivo, although the
remaining part of the complex (HoxFUYH) still shows
unaltered H
2
-dependent NAD
+
-reducing activity
in vitro [18]. However, the roles of HoxE and the

Hox complex are still not fully understood.
In this article, we show that the various hydrogenas-
es use distinct electron-transfer subunits and routes.
Deletion of the HupC, Isp1,2 and HoxE proteins
clearly reveals their physiological relationships to their
respective hydrogenases. Affinity purification of the
HoxE-tagged protein under mild conditions confirms
the heteropentameric structure of this complex.
Results
Isp1 and Isp2 are expressed proteins
The in silico analysis of the intergenic region of the
hynS and hynL genes indicated two open reading
frames. It has been established that the hynS-isp1-isp2-
hynL region is cotranscribed [23]. In order to confirm
that isp1 and isp2 are really coding regions, the hynS-
isp1-isp2-hynL* genes were cloned behind a T7
promoter (see Experimental procedures). The genes of
L. S. Pala
´
gyi-Me
´
sza
´
ros et al. Electron-transfer subunits of NiFe hydrogenases
FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS 165
the construct were expressed in the Escherichia coli
BL21(DE3) host, and the bands corresponding to the
calculated molecular masses of Isp1 (24.6 kDa) and
Isp2 (48.4 kDa) could be clearly identified (data not
shown). The small and large subunits were also

detected. This means that all the translational signals
necessary for the expression of the HynSL and Isp
subunits are functionally present in the construct, and
are recognized by the translational apparatus of
E. coli. The coexpression of the Hyn and Isp subunits
suggests that they probably form a functional
complex.
Isp1 and Isp2 are required for the in vivo function
of Hyn hydrogenase
The solubilized and purified Hyn hydrogenase con-
tained only the HynSL subunits [23]. The role of the
Isp proteins in T. roseopersicina is unknown, but com-
putational analysis has shown that Isp1 is a b-type
haem-containing transmembrane electron carrier,
whereas Isp2 seems to be a redox Fe–S-containing pro-
tein. If these subunits are involved in the electron flow
from ⁄ to the hydrogenase, their removal would abolish
the hydrogenase activity in vivo, where the endogenous
electron donors ⁄ acceptors must be used.
Therefore, a double isp1-isp2 in-frame mutant was
constructed in the T. roseopersicina GB2131 (DhoxH,
DhupSL) strain (ISP12M, see Experimental proce-
dures). The hydrogenase activities were measured both
in vivo (without the addition of an artificial electron
carrier) and in vitro (in the presence of redox viologen
dyes). The data in Table 1 unequivocally prove that
the in vivo H
2
-producing activity of the isp1,2 mutant
strain is completely lost and the in vivo H

2
uptake
activity is dramatically decreased relative to the control
GB2131 (DhoxH, DhupSL) strain containing all the
functional gene products of the Hyn operon. A single
Isp1 in-frame deletion mutant was also constructed
(ISP1M). Mutation of the Isp1 protein brings about
the same phenotype as the deletion of both Isp1 and
Isp2 (Table 1). Some remaining in vivo H
2
uptake
activity of the Hyn hydrogenase can be detected in
both mutants, which suggests an alternative, less effec-
tive electron-transfer pathway.
In the in vitro measurements, in which benzyl-violo-
gen was used as an artificial electron acceptor, the H
2
uptake activity was not influenced by the lack of Isp1
or Isp1,2 proteins (Table 1). On the one hand, these
and the in silico results confirm that the Isp proteins
play an essential role in the H
2
reduction and oxida-
tion ability of Hyn hydrogenase in its natural environ-
ment, but the lack of these subunits has no effect on
the hydrogenase activity in the artificial assay. On the
other hand, this also means that the Isp1,2 proteins do
not affect the post-translational maturation and
expression level of the Hyn enzyme. A trivial rationali-
zation of these observations is that the lack of Isp1 or

Isp1,2 proteins results in blockade of the electron flow
from ⁄ to Hyn hydrogenase under physiological condi-
tions.
As the computational analysis implies that Isp1 is
an integrated membrane protein, it is plausible to
assume that the HynSL dimer is anchored to the mem-
brane through the Isp1 protein. Accordingly, we inves-
tigated the localization of Hyn hydrogenase in the Isp
mutant strains. Unexpectedly, the in vitro H
2
uptake
measurements on the various cellular fractions indi-
cated that a similar proportion of Hyn hydrogenase
remained in the membrane fraction in the presence
and absence of the Isp proteins (Table 2). This is
surprising, as our protein purification experiments
demonstrated that the HynSL subunits are only loosely
associated with the membrane and can be easily
Table 1. Activities of Hyn hydrogenase in vivo and in vitro in the
presence and absence of the Isp1 and Isp2 proteins. The results
are given as percentages of the level for GB2131. The cultures
were grown on Pfennig’s medium with 4 gÆL
)1
of Na
2
S
2
O
3
. The val-

ues are normalized to bacteriochlorophyll content. The GB112131
strain (DhupSL, DhoxH, DhynS-isp1-isp2-hynL) and the M539 strain
(hypF mutant) containing no active NiFe hydrogenase served as
negative controls.
Strain
Relative H
2
production
in vivo
Relative
H
2
uptake
in vivo
Relative
H
2
uptake
in vitro
GB2131
(DhupSL, DhoxH)
100 ± 10.2 100 ± 13.7 100 ± 10.0
ISP1M
(DhupSL, DhoxH, Disp1)
0.00 29.5 ± 9.8 100.4 ± 9.2
ISP12M t
(DhupSL, DhoxH, Disp12)
0.0 35.6 ± 5.9 116.2 ± 8.3
Table 2. Location of Hyn hydrogenase with and without the Isp1,2
proteins (see description in Table 1).

Strain
Relative uptake activity
in vitro
Membrane
fraction
Soluble
fraction
GB2131 (DhupSL, DhoxH) 100 ± 23.2 100 ± 7.7
ISP1M (DhupSL, DhoxH, Disp1) 106.2 ± 33.4 102.7 ± 4.0
ISP12M (DhupSL, DhoxH, Disp12) 112.4 ± 0.3 113.9 ± 6.8
Electron-transfer subunits of NiFe hydrogenases L. S. Pala
´
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´
sza
´
ros et al.
166 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS
washed off [23]. The strength of the HynSL–membrane
interaction apparently does not depend on the presence
or absence of the Isp1 protein.
The expression of the Hup enzyme depends on
the thiosulfate content of the medium
With a view to examining the function of the HupC
protein in T. roseopersicina,aDhynSL, DhoxH
(GB1131) strain was created (see Experimental proce-
dures). This strain is suitable for the measurement of
Hup hydrogenase activity alone, without the contribu-
tions of Hyn and Hox hydrogenases. However, under
standard growth conditions, i.e. in the presence of

4gÆL
)1
Na
2
S
2
O
3
, only very low HupSL hydrogenase
activity was detected in the DhynSL, DhoxH (GB1131)
strain. It was postulated that this concentration of
thiosulfate resulted in a redox potential in the cells,
which downregulated the activity of HupSL hydro-
genase (as an uptake, electron-donating enzyme). To
test this hypothesis, the expression level and in vitro
activity of the Hup hydrogenase were measured in cells
grown in the presence of various amounts of thiosul-
fate. The data in Table 3 clearly illustrate that the
lower the thiosulfate concentration in the medium, the
higher the Hup hydrogenase activity both in vivo and
in vitro. The effects of the thiosulfate content on the
expression level of the hupSL genes were additionally
monitored by quantitative RT-PCR. The data in
Table 4 reveal that a decrease in the thiosulfate con-
tent of the medium from 4 to 2 gÆL
)1
resulted in a dra-
matic (> 16-fold) increase in the hupSL mRNA level.
These data suggest that, when Hup is the only active
hydrogenase in the cell, its activity strongly depends

on the thiosulfate content of the medium, and changes
in the activity primarily correlate with the expression
level of the enzyme. Hence, as a practical consequence,
the subsequent experiments on Hup activity were per-
formed with samples grown in the presence of 2 gÆL
)1
thiosulfate.
HupC is an electron-transfer subunit of Hup
hydrogenase
To establish the function of HupC, its gene was
deleted in-frame in the DhynSL, DhoxH (GB1131)
strain, and the HupSL activities were compared both
in vivo and in vitro.
The in vivo H
2
uptake activity of Hup hydrogenase
was substantially decreased in the DhupC (HCMG4)
strain. At the same time, the in vitro activity was twice
as high as that of the strain harbouring HupC
(Table 5). A comparison of the hupSL mRNA levels
of the cells containing or lacking the hupC gene per-
ceptibly revealed that a loss of the hupC gene had a
positive effect on the transcription level of the hupSL
genes (Table 4). To check that the effect was really
linked to the loss of HupC, a complementation experi-
ment was performed by introducing an expression
Table 3. In vivo and in vitro H
2
uptake activities of the GB1131
(DhynSL, DhoxH) strain grown photoautotrophically (Pfennig’s) at

various Na
2
S
2
O
3
concentrations. The results are given as percent-
ages of that for the sample grown with 1 gÆL
)1
of Na
2
S
2
O
3
.
Concentration of
Na
2
S
2
O
3
(gÆL
)1
)
Relative H
2
uptake activity
In vivo In vitro

4 0.0 0.0
2 45.0 ± 2.6 83.6 ± 34.2
1 100.0 ± 5.6 100.0 ± 11.1
Table 4. Relative mRNA levels of the hup operon in the presence
(GB1131) and absence (HCMG4) of the hupC gene at various
Na
2
S
2
O
3
concentrations. The cultures were grown on Pfennig’s
medium with 2 or 4 gÆL
)1
of Na
2
S
2
O
3
. The mRNA levels were
determined by quantitative RT-PCR and the results are given as
percentages of the level for GB1131. The values are normalized to
the total RNA content.
Strain
4gÆL
)1
Na
2
S

2
O
3
2gÆL
)1
Na
2
S
2
O
3
GB1131
(DhynS-isp1-isp2-hynL, DhoxH)
100.0 ± 0.0 1650.0 ± 44.5
HCMG4
(DhupC, DhynS-isp1-isp2-hynL,
DhoxH)
300.0 ± 20.0 2700.0 ± 102.8
Table 5. Activities of Hup hydrogenase in vivo and in vitro in the
presence (GB1131, pMHE6C HCMG4) and absence (HCMG4) of the
HupC protein. The cultures were grown on Pfennig’s medium with
2gÆL
)1
of Na
2
S
2
O
3
. The hydrogenase activity values are normalized

to the bacteriochlorophyll content. The results are given as a percent-
age of the level for GB1131. The GB112131 strain (DhupSL, DhoxH,
DhynS-isp1-isp2-hynL) and the M539 strain (hypF mutant) containing
no active NiFe hydrogenase served as negative controls.
Strain
Relative H
2
uptake activity
In vivo In vitro
GB1131 (DhynS-isp1-isp2-hynL,
DhoxH)
100.0 ± 2.6 100.0 ± 14.5
HCMG4 (DhupC,
DhynS-isp1-isp2-hynL, DhoxH)
40.4 ± 5.5 198.9 ± 5.5
pMHE6C HCMG4 (DhupC,
DhynS-isp1-isp2-hynL,
DhoxH, pMHE6C)
68.3 ± 10.3 231.2 ± 33.5
L. S. Pala
´
gyi-Me
´
sza
´
ros et al. Electron-transfer subunits of NiFe hydrogenases
FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS 167
cassette containing the hupC gene driven by the crt
promoter (see Experimental procedures). Table 5
shows that the plasmid-borne HupC (pMHE6C ⁄

HCMG4 in Table 5) partially ( 50%) restored the
Hup hydrogenase activity in vivo.
It is plausible to assume that HupC serves as a
membrane anchor for Hup hydrogenase [24]. In con-
trast with the findings on Hyn hydrogenase, the lack
of HupC substantially reduced the Hup hydrogenase
activity in the membrane fraction, i.e. 98% of the
activity was lost in the hupC deletion mutant. The
hydrogenase activity in the soluble fraction was also
decreased; therefore, it is unlikely that HupSL was
released from the membrane and accumulated in the
cytoplasm. The lower total activity in the HupC-minus
cell fractions might be explained by the lower stability
of the HupSL enzyme in the absence of HupC in the
disrupted and fractionated cells relative to the wild-
type (Table 6). It is noteworthy that the HupSL activ-
ity was significantly higher in the soluble than in the
membrane fraction in the pMHE6C ⁄ HCMG4 (HupC
complementing) strain (Table 6). The plasmid-borne
HupC could possibly restore the stability of HupSL,
although the majority of the activity remained in the
soluble fraction.
These data suggest that HupC has no role in the
maturation process of HupSL hydrogenase, but influ-
ences the in vivo activity and the expression level of the
HupSL enzyme. Taken together with the findings of
computational analysis, the HupC protein serves an
electron-transport role in T. roseopersicina and proba-
bly forms a functional complex with the small and
large hydrogenase subunits in vivo.

Purification of Hox hydrogenase
In T. roseopersicina, the cytoplasmic Hox hydrogenase
is coded by the hoxEFUYH operon. The enzyme
contains hydrogenase (HoxYH) and diaphorase (Ho-
xEFU) subunits [18]. The diaphorase subunits of the
Hox-type hydrogenases exhibit significant sequence
similarities to three subunits (NuoEFG) of
NADH:ubiquinone oxidoreductase [18,25]. The
in-frame deletion of the hoxE gene led to the complete
loss of Hox activity in vivo, whereas the enzyme was
fully active in vitro [18]. This suggests that HoxE may
function in vivo as an electron-transfer protein. Thus,
HoxE would offer a third channel for the electrons in
addition to the hydrogenase and diaphorase catalytic
centres. To test whether the HoxE protein forms a
functional complex with the HoxFUYH subunits, its
FLAG-tagged form was expressed from a pMHE6
expression vector [26] under the control of the
T. roseopersicina crt promoter (pMHE6HoxE Table 7).
HoxE was purified by affinity chromatography via the
FLAG-tag under very mild conditions in order to pre-
serve the protein–protein interactions (see Experimen-
tal procedures). The proteins eluted from the affinity
column were separated on SDS-polyacrylamide gel and
analysed by MALDI-TOF-MS. Each subunit of the
HoxEFUYH enzyme complex was easily identified,
indicating that HoxE is physically associated with the
other (HoxFUYH) subunits (Fig. 1).
Discussion
Hydrogenases are widespread in the microbial world.

The actively expressed hydrogenases must have a dedi-
cated physiological role within the cells. For the in vivo
function, the catalytic dimers of NiFe hydrogenases
must be connected to other oxidoreductases directly or
via electron-transfer subunits. In this study, attempts
were made to identify the redox partners and the elec-
tron-channelling subunits of all three hydrogenases in
the cells.
In T. roseopersicina, there are at least three NiFe
hydrogenases (HynSL, HupSL and HoxYH) with
distinct properties and different functions. The HypF
accessory protein is required for the maturation of
every NiFe hydrogenase, and disruption of the hypF
gene therefore results in the hydrogenase-minus pheno-
type [27]. However, the hydrogenase-less cells showed
virtually identical growth properties as the wild-type
under standard growth conditions. Special growth con-
ditions, i.e. photoautotrophic in the presence of H
2
and only 0.005% Na
2
S, were identified, in which the
presence of each hydrogenase was important, including
the Hup enzyme being essential for H
2
-dependent
growth (data not shown). This indicates that the Hup
enzyme has a direct connection to the central redox,
i.e. quinone, pool. Nonetheless, the real redox partners
Table 6. Location of HupSL hydrogenase with (GB1131, pMHE6C

HCMG4) and without (HCMG4) the HupC subunit (see description
in Table 5).
Strain
In vitro relative uptake activity
Membrane
fraction
Soluble
fraction
GB1131 (DhynS-isp1-isp2-hynL,
DhoxH)
100.0 ± 12.9 36.0 ± 8.5
HCMG4 (DhupC,
DhynS-isp1-isp2-hynL, DhoxH)
1.8 ± 0.24 11.3 ± 1.8
pMHE6C HCMG4 (DhupC,
DhynS-isp1-isp2-hynL,
DhoxH, pMHE6C)
47.3 ± 0.9 115.0 ± 7.1
Electron-transfer subunits of NiFe hydrogenases L. S. Pala
´
gyi-Me
´
sza
´
ros et al.
168 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS
and ⁄ or electron channels of each hydrogenase remain
poorly understood.
The genes coding for the HynSL enzyme are sepa-
rated by two open reading frames, which have been

shown to code for real proteins, Isp1 and Isp2. Both
proteins have been demonstrated to be important for
the function of the HynSL enzyme in vivo, but neither
for its in vitro activity or expression. Therefore, they
probably play an electron-transfer role from ⁄ to the
Hyn enzyme. The heterodisulfide reductase homologue
Isp2 is probably an oxidoreductase; its redox substrate
still remains to be identified. We conclude that the
Hyn enzyme is indirectly linked to the central
redox ⁄ bioenergetic processes via the Isp1,2 proteins
and an unknown redox substrate.
A direct coupling of the HupC protein to the uptake
HupSL hydrogenase was demonstrated in this study.
Deletion of the hupC gene resulted in reduced and
enhanced activities of HupSL in vivo and in vitro,
respectively. As HupC is supposed to react with qui-
nones directly [17], the reduced in vivo activity stems
from the obstruction of the electron flow from the
hydrogenase. Consequently, HupC is suggested to be
the third subunit of the Hup complex, catalysing the
H
2
-dependent reduction of quinones. This is in line
with the observation that HupSL hydrogenase is
essential for H
2
-dependent growth under the above-
mentioned growth conditions.
The expression level of the HupSL enzyme was
upregulated both by disrupting the HupC subunit and

by decreasing the thiosulfate content of the medium. It
is assumed that both processes lead to a more oxidized
quinone pool, as the disrupted HupC cannot transfer
the electrons from HupSL, and thiosulfate serves as
reducing power for the photosynthetic carbon fixation
via the central quinone pool [28]. The redox status of
the quinone pool may influence HupSL expression: the
increased electron requirement is reflected in a higher
expression level of the electron-donating Hup hydro-
genase.
Interestingly, removal of the transmembrane elec-
tron-transfer subunits of the Hyn and Hup enzymes
gave rise to distinct effects on the locations of their
corresponding hydrogenases. The lack of Isp1 did not
change the membrane association of the Hyn enzyme,
whereas the elimination of HupC led to detachment of
Table 7. Strains and plasmids. Indicated strains and plasmids are from Stratagene, La Jolla, CA, USA.
Strain or plasmid Relevant genotype or phenotype
Reference
or source
Thiocapsa roseopersicina
BBS Wild-type [5]
GB2131 hupSLD::Gm, hoxHD::Er [18]
GB1121 hynSLD_::Smr, hupSLD_::Gmr [18]
GB1131 hynSLD_::Smr, hoxHD::Er This work
GB112131 hynSLD::Smr, hupSLD::Gm, hoxHD::Er [18]
M539 hypFD::Km [27]
ISP1M hupSLD::Gm, hoxHD::Er, isp1D This work
ISP12M hupSLD ::Gm, hoxHD::Er, D isp1, D isp2 This work
HCMG4 hynSLD_::Smr, hoxHD::Er, DhupC This work

pMHE6C ⁄ HCMG4 hynSLD_::Smr, hoxHD::Er, DhupC, pMHE6C This work
pMHE6HoxE ⁄ GB2131 hupSLD::Gm, hoxHD::Er, pMHE6HoxE This work
Escherichia coli
XL1-Blue MRF
¢ D (mcrA)183, D (mcrCB-hsdSMR-mrr)173, endA1, supE44, thi-1, recA1, gyrA96, relA1
lac [F¢ proAB lacIqZDM15 Tn10 (Tet
r
)]
c
Stratagene
BL21 (DE3) F
)
ompT gal dcm lon hsdS
B
(r
B
)
m
B
)
) k(DE3) [lacI lacUV5-T7 gene 1 ind1 sam7 nin5] Stratagene
Plasmids
pBluescript SK+ Amp
r
, cloning vector, ColE1 ori Stratagene
pK18mobSacB Km
r
sacB RP4 oriT ColE1 ori [36]
pUC19 Amp
r

, cloning vector, ColE1 ori [37]
pMHE6crtKm Km
r
, mob
+
, expression vector containing the promoter region of crtD gene [26]
pMHE6C Km
r
, mob
+
, hupC gene cloned after the crtD gene This work
pMHE6HoxE Km
r
, mob
+
, hoxE gene cloned after the crtD gene This work
pISP1M Km
r
, in-frame up- and downstream homologous regions of isp1 in pK18mobsacB This work
pISP12M Km
r
, in-frame up- and downstream homologous regions of isp1-isp2 in pK18mobsacB This work
pHCD2 Km
r
, in-frame up- and downstream homologous regions of hupC in pK18mobsacB This work
pTSH2 ⁄ 8 Cosmid clone containing hyp operon [8]
L. S. Pala
´
gyi-Me
´

sza
´
ros et al. Electron-transfer subunits of NiFe hydrogenases
FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS 169
the HupSL enzyme from the membrane. The absence
of HupC also resulted in a destabilization of HupSL.
A similar phenomenon has been described for HupC
in Rhodobacter capsulatus [24]. In T. roseopersicina, the
plasmid-borne HupC restored the stability and mem-
brane association of HupSL, but a significant amount
of enzyme remained in the soluble fraction. Controver-
sial data have been published in the literature with
regard to the membrane anchoring role of HupC.
Deletion of HoxW, the HupC homologous protein in
R. eutropha, resulted in detachment of the hydrogenase
from the membrane [29]. In contrast in Pseudomo-
nas hydrogenovora, the location of the HupSL dimer in
the hupC mutant strain did not change [30].
It has been shown previously that HoxE is required
for the in vivo function of the Hox hydrogenase [18].
Here, we have demonstrated that HoxE fulfills this
role in association with the other subunits of the Hox
hydrogenase. Purification of the affinity-tagged HoxE
under mild conditions resulted in copurification of the
four other (HoxFUYH) subunits. We observed that
the hydrogenase dimer dissociated from the HoxEFU
trimer relatively easily (data not shown). A similar
finding has been published recently for the A. vinosum
Hox hydrogenase [20]. This means that the enzyme com-
plex has three gates for electron flow: one for H

2
oxid-
ation ⁄ proton reduction, one for the NAD
+
⁄ NADH
redox reaction and one functioning as an electron
channel via the HoxE subunit. This makes the potential
physiological function of this hydrogenase more
complex, as the Hox hydrogenase has a potential to be
associated with various metabolic pathways involving
redox changes.
In cyanobacteria, the Hox hydrogenase was initially
suggested to have a relationship to the respiratory
complex [25], but evidence challenging this idea was
later published [31]. A valve role of the Hox enzyme
was suggested for the low-potential electrons generated
during photosynthesis [32]. The three gates for electron
flow are in line with the valve hypothesis. However,
depending on the sulfur source, the Hox enzyme is
able to produce H
2
either under illumination or in the
dark [33], and thus its physiological function cannot
be restricted to photosynthetic electron flow, but the
respiratory and fermentative processes should also be
considered.
Experimental procedures
Bacterial strains and plasmids
The strains and plasmids are listed in Table 7. The T. roseo-
persicina strains were grown photoautotrophically in

Pfennig’s medium under anaerobic conditions in liquid
cultures with continuous illumination at 27–30 °C for
4–5 days [27]. The acetate-supplemented (2 gÆL
)1
) plates
were solidified with 7 gÆL
)1
of Phytagel (Sigma, St Louis,
MO, USA) [34]. The plates were incubated in anaerobic jars
by means of the AnaeroCult (Merck, Darmstadt, Germany)
system for 2 weeks. The E. coli strains were maintained on
LB-agar plates. Antibiotics were used in the following con-
centrations (mgÆL
)1
): for E. coli, ampicillin (100), kanamycin
(25), tetracyclin (20); for T. roseopersicina, kanamycin (25),
streptomycin (5) and gentamycin (5).
Expression of the hynS-isp1-isp2-hynL* genes
of T. roseopersicina in E. coli using the T7
promoter
⁄
RNS polymerase system
The hynS-isp1-isp2-hynL* gene products were produced
from pTSH2 ⁄ 8 [8] in the E. coli BL21(DE3) strain. This
construct contains the native promoter ⁄ regulatory region
and, additionally, the complete hynS, isp1 and isp2 genes
and truncated hynL (denoted by *). The incomplete hynL
did not interfere with the outcome of the experiments.
Expression of the genes was induced by isopropyl thio-b-d-
P

T7
P
crtD
RBS
HoxE
FLAG-StrepII
kDa
120
100
321M
85
70
60
50
40
HoxF
HoxU
HoxE
HoxE-TAG
HoxH
30
25
20
Phenylalanyl t-RNA synthetase β subunit
Phenylalanyl t-RNA synthetase α subuni
t
Fig. 1. The HoxFUYH subunits copurifiy with the tagged HoxE sub-
unit during the course of affinity chromatography. (A) Scheme of
the cassette used to express tagged HoxE in T. roseopersicina
(P

crtD
, carotenoid promoter; P
T7
, T7 promoter; RBS, ribosome-bind-
ing site; M, marker). (B) Protein patterns of the elution fractions
separated by SDS-PAGE. The soluble fraction of T. roseopersicina
cells expressing tagged HoxE was loaded onto an Anti-Flag affinity
column, the resin was washed, and the bound proteins were eluted
three times by Flag peptide (for details, see Experimental proce-
dures) (1, 2, 3 indicate the elution fractions). The bordered bands
were cut and analysed by mass spectrometry.
Electron-transfer subunits of NiFe hydrogenases L. S. Pala
´
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´
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ros et al.
170 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS
galactoside, and monitored by the incorporation of
l-[
35
S]methionine into the proteins synthesized [35]. The
samples were separated in an SDS-polyacrylamide gel and
analysed by a Phosphor Imager (Phosphor Imager 445
SI, Molecular Dynamics, Uppsala, Sweden).
Conjugation
Conjugation was carried out as described previously [27].
Deletion of the isp1,2 genes
The in-frame deletion constructs were derived from the

pK18mobsacB vector [36]. The upstream region of the
isp1,2 genes was amplified with the otsh14r (5¢-GAT
CGCGATATTGAACATC-3¢) and trhydo3 (5¢-CATA
TGGCTGCCCGTAACCCCACTGAT-3¢) primers. The
product was cloned into the polished BamHI site of pUC19
[37], yielding pUNSBamHI.
To clone the downstream region, another PCR was per-
formed with the isp1o7 (5¢-TCGCACGCTGGTACAA
CGGG-3¢) and isp2o2 (5¢-ACCAGGTGCTCGGCGAT
CAT-3¢) primers. This fragment was cloned into the XbaI-
digested and blunted pUNSBamHI vector (pUS2). The
2502 bp EcoRI fragment of pUS2 was ligated with the
EcoRI fragment of pK18mobSacB, yielding pISP12M.
The plasmid was transformed into the E. coli S17-1(kpir)
strain, and then conjugated into the T. roseopersicina
GB2131 strain as described previously [27]. The single
recombinants selected through their kanamycin resistance
were grown in liquid medium. The double recombinants
were selected on 3% sucrose-containing plates. The
sucrose-resistant and kanamycin-sensitive colonies were
selected, and the genotype was confirmed by Southern
blotting and hybridization (ISP12M).
Deletion of the isp1 gene
The upstream homologous region was taken from the
pUNSBamHI vector. The downstream homologous region
was amplified with the isp1o8 (5¢-AGCTGACGCACATCT
TCACG-3¢) and isp2o7 (5¢-GGTGAGACCGACCACCG
GGA-3¢) primers. The product was cloned into the BamHI-
cleaved and polished pUNSBamHI construct (pUS3). The
EcoRI fragment of pUS3 was cloned into pK18mobSacB

(pISM1 ⁄ 3). The construct was conjugated into the GB2131
strain and the double recombinants were selected as
described below.
Deletion of the hupC gene
For deletion of the hupC gene, the pHCD1 and pHCD2
in-frame deletion constructs were created as follows. The
upstream region of hupC was amplified with the ohup20
(5¢-CGAGCAGGCCAAGTATTC-3¢) and ohup19 (5¢-TGT
TGGTCAGGCGGATCT-3¢) primers, and the 836 bp PCR
product was cloned into the SmaI-digested pK18mobsacB
(pHCD1). The downstream region was amplified with the
ohup21 (5¢-GGCGGATGTTCAAGGACG-3¢) and ohup22
(5¢-TCGACCACGACACTGAAG-3¢) primers. The 800 bp
fragment obtained was cloned into the PstI-digested
polished pHCD1 (pHCD2). This construct was conjugated
into the T. roseopersicina GB1131 strain, yielding the
HCMG4 strain. The double recombinants were selected
and the genotypes were confirmed as described above.
Construction of HupC-expressing plasmid
The hupC gene was amplified with the ohupc1 (5¢-CATAT
GTCGCGAGCTGCGTCGCG-3¢) and ohupc2 (5¢-AAGCT
TTGGCCGATCGTCCTTGAACAT-3¢) primers containing
NdeI and HindIII recognition sites. The 777 bp PCR prod-
uct was inserted into the EcoRV-digested pBluescripSK+
(pBtC). The 777 bp NdeI-HindIII-digested fragment was
ligated into the corresponding sites of pMHE6crtKm [26],
resulting in pMHE6C.
RNA isolation
For RNA isolation, T. roseopersicina was grown in 60 mL
of liquid medium in a hypovial to A

600 nm
= 1–1.5; 15 mL
of culture was centrifuged at 15 000 g for 2 min, the pellet
was suspended in 300 lL of SET buffer [20% sucrose,
50 mm EDTA (pH 8.0) and 50 mm Tris ⁄ HCl (pH 8.0)]
and 300 lL of SDS buffer was added [20% SDS, 1%
(NH
4
)
2
SO
4
, pH 4.8]; 500 lL of saturated NaCl was
added next, the sample was centrifuged at 20 000 g for
10 min and the clear supernatant was transferred into a
new tube. 2-Propanol (70% of the total volume of the
supernatant) was added to the solution and the mixture
was centrifuged at 20 000 g for 20 min. The pellet was
washed twice with 1 mL of 70% ethanol. The dried pellet
was suspended in 20 lL of diethylpyrocarbonate-treated
water.
DNase I treatment
DNase treatment took place in the presence of 10· reaction
buffer with MgCl
2
(Fermentas, Burlington, Canada) and
DNase I (RNase-free, Fermentas) at 37 °C for 1 h. The
reaction was inactivated by heat at 65 °C for 10 min in the
presence of EDTA (Fermentas).
Reverse transcription and quantitative real-time

PCR
For reverse transcription, the OmniscriptÒ Reverse Trans-
criptase Kit (Qiagen, Hilden, Germany) was used according
L. S. Pala
´
gyi-Me
´
sza
´
ros et al. Electron-transfer subunits of NiFe hydrogenases
FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS 171
to the manufacturer’s instructions. One microgram of
DNase I-treated total RNA was added to a master mix
[10· buffer RT, dNTP mix (0.5 mm of each dNTP), reverse
primer (0.2 lm), RNase inhibitor (10 units ⁄ reaction),
Omniscript Reverse Transcriptase (2 units ⁄ reaction) in
diethylpyrocarbonate-treated water] on ice in a final volume
of 20 lL. The reaction mixture was then incubated at
37 °C for 60 min.
Reverse transcription was initiated from the huprto2
(5¢-CGCTTGAGCCGATTCTGAACAT-3¢) primer specific
for the hupL gene. The cDNA produced during reverse
transcription was used as a template for quantitative PCR,
which was performed using the ohupSRT1 (5¢-GGA
CAAGGGCAGCTTCTATCA-3¢) and ohupSRT2 (5¢-CG
CATTGGCCTCGATACC-3¢) primers located in the hupS
gene. PCR was carried out and the products were measured
with an Applied Biosystems (Foster City, CA, USA) 7500
real-time PCR instrument. PCR was performed in a total
volume of 25 lL, including 1 lL of cDNA, 12.5 lLof

Power SYBR Green PCR Master Mix (Applied Bio-
systems), forward and reverse primers (12.5 pmol of each)
and 9 lL of nuclease-free water. The following programme
was applied: 95 °C for 10 min; 95 °C for 15 s and 60 °C
for 1 min for 40 cycles; 95 °C for 15 s; 60 °C for 1 min;
95 °C for 15 s; 60 °C for 15 s. A calibration curve was gen-
erated using sixfold dilutions of pKK48 plasmid DNA
(containing the sequence of the hupS gene) in the 100 to
0.001 ngÆlL
)1
concentration range.
Activity measurements
The hydrogenase activities of the various mutants were
measured both in vivo and in vitro. In all experiments, the
HypF mutant (lacking any NiFe hydrogenase activity) and
the GB112131 strain (DhoxH, DhupSL, DhynS-isp1-isp2-
hynL) were used as negative controls.
In vitro H
2
uptake activity measurements
The samples were suspended in 2 mL of 20 mm potassium
phosphate buffer containing 0.4 mm of oxidized benzyl-
viologen. The cuvettes were closed with SubaSeal rubber
stoppers. The gas phase was flushed with H
2
and the H
2
uptake activity was measured spectrophotometrically at
600 nm and 60 °C.
In vivo hydrogen evolution activity

measurements
Cultures (60 mL) were grown in 100 mL hypovials; the gas
phase was then flushed with N
2
after inoculation and the
H
2
produced was measured gas chromatographically [27]
on day 6.
In vivo H
2
-uptake activity measurements
Medium (60 mL) was inoculated into 100 mL hypovials;
the gas phase was flushed with N
2
and 5 mL of pure H
2
was injected into the bottles. The cultures were grown
under illumination and the H
2
content of the gas phase was
measured gas chromatographically on day 6.
Preparation of membrane and soluble fractions
of T. roseopersicina
The membrane fractions were prepared from 50 and
110 mL cultures for Hyn and Hup measurements, respec-
tively. The cells were harvested by centrifugation at 7000 g,
suspended in 1 mL of 20 mm potassium phosphate buffer
(pH 7.0) and broken by sonication [Bandelin Sonopuls
(Berlin, Germany) HD2070 ultrasonic homogenizer; at 85%

amplitude six times for 10 s]. The broken cells were centri-
fuged at 15 000 g for 10 min. The debris (sulfur globules
and intact cells) was discarded and the supernatant was
centrifuged at 100 000 g for 1.5 h. The pellet was washed,
resuspended in 800 lL of potassium phosphate buffer
(pH 7.0) and used as membrane fraction. The supernatant
was regarded as the soluble fraction.
Measurements of bacteriochlorophyll content
The bacteriochlorophyll content was estimated using a
methanol extraction procedure, as described previously [38].
The absorption of the samples was measured at 772 nm;
the extinction coefficient was 8.41 g
)1
ÆLÆcm
)1
. The in vivo
and in vitro activities were normalized to the bacteriochlo-
rophyll content of the samples.
Construction of the double-tagged hoxE gene
For the construction of an expression system capable of
producing the HoxE protein of T. roseopersicina fused with
tandem FLAG-tag-Strep-tag II at the C-terminus, a 501-bp
fragment was amplified from the pTCB4 ⁄ 2 clone [8] using
the TCHO32 (5¢-CATATGAGTCTGCAGCAAGCCA-3¢)
and TCHO33 (5¢-AAGCTTGGTCAGCTCCTCGAGC-3¢)
primers and cloned into the SmaI site of pBluescript SK+
(pBtHoxE). The 494 bp NdeI-HindIII fragment of
pBtHoxE was ligated into the NdeI-HindIII-digested
pMHE6crtKm vector (pMHE6HoxE). The construct was
confirmed by sequencing and conjugated into the T. roseo-

persicina GB1121 strain.
Purification of Hox hydrogenase
Four grams of cell paste from a GB1121 ⁄ pMHE6HoxE-
Km culture were suspended in 5 mL of NaCl ⁄ Tris [50 mm
Electron-transfer subunits of NiFe hydrogenases L. S. Pala
´
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´
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´
ros et al.
172 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS
Tris (pH 7.4) and 150 mm NaCl]. The sample was sonicated
with a Bandelin Sonopuls HD2070 ultrasonic homogenizer
(at medium mode, amplitude 2.4 times for 10 s). The cell
debris and sulfur crystals were removed by centrifugation
(27 000 g, 10 min). The supernatant was incubated with
300 lL of ANTI-FLAG M2 affinity resin (Sigma) at 4 °C
for 2 h with gentle shaking. The matrices were washed
seven times with 1.5 mL of NaCl ⁄ Tris. For elution, the
slurry was washed twice with 100 lL and once with 50 lL
of NaCl ⁄ Tris with FLAG-peptide (200 lgÆmL
)1
). Aliquots
were collected and the samples were analysed by SDS-
PAGE.
SDS-PAGE and protein staining
SDS-PAGE and silver staining of proteins were performed
as described by Ausubel et al. [39].
Identification of proteins by MALDI-TOF-MS

Coomassie blue-stained gel bands were cut out and analy-
sed by MALDI-TOF-MS, as described previously [26].
Bioinformatics tools
Protein sequences in the various databases were compared
with the blast (P, X) programs (.
gov), the peptide mass fingerprints and the power spectral
density spectra; a database search was performed using the
National Center for Biotechnology Information protein
database with Protein Prospector MS-Fit and MS-Tag,
respectively ( />Acknowledgements
The contribution of Drs B. D. Fodor and A
´
. T. Kov-
a
´
cs in the early phase of this work is gratefully
acknowledged. This work was supported by EU pro-
jects HyVolution FP6-IP-SES6 019825 and FP7 Col-
laborative Project SOLAR-H2 FP7-Energy-212508,
and by domestic funds (GOP-2007-1.1.2, Asbo
´
th-
DAMEC-2007 ⁄ 09, Baross OMFB-00265 ⁄ 2007 and
KN-RET-07 ⁄ 2005).
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