Báo cáo khóa học: Comparison of native and recombinant chlorite dismutase from Ideonella dechloratans pot

Comparison of native and recombinant chlorite dismutase
from
Ideonella dechloratans
Helena Danielsson Thorell
1
, Natascha H. Beyer
2
, Niels H. H. Heegaard
2
, Marcus O
¨
hman
1
and Thomas Nilsson
1
1
Department of Chemistry, Karlstad University, Sweden;
2
Department of Autoimmunology, Statens Serum Institut, Copenhagen,
Denmark
A detailed comparison between native chlorite dismutase
from Ideonella dechloratans, and the recombinant version
of the protein produced in Escherichia coli, suggests the
presence of a covalent modification in the native enzyme.
Although t he native and r ecombinant N- and C-terminal
sequences are identical, the enzymes display different
electrophoretic mobilities, and produce different p eptide
maps upon digestion with trypsin and s eparation of
fragments using capillary electrophoresis. Comparison of
MALDI m ass s pectra of tryptic peptides from the native
and recombinant enzymes suggests two locations for

modification in the native protein. Mass spectrometric
analysis of isolated peptides from a tryptic digest of the
native enzyme identifies a possible cross-linked dipeptide,
suggesting an intrachain cross-link in the parent protein.
Spectrophotometric titration of the native enzyme in the
denatured state reveals two titrating components absorb-
ing at 295 nm, suggesting the presence of about one
tyrosine residue per subunit with an anomalously low
pK
a
. The EPR spectrum for the recombinant enzyme is
different from that of the native enzyme, and contains a
substantial contribution of a low-spin species with the
characteristics of bis-histidine coordination. These results
are discussed in terms of a covalent cross-link b etween a
histidine and a tyrosine sidechain, similar to those found
in other heme enzymes operating under highly oxidizing
conditions.
Keywords: chlorate; chlorite dismutase; recombinant chlorite
dismutase; post-translational modification.
Chlorate- and perchlorate-respiring bacteria have attracted
interest due to their potential use in the treatment of soil and
water contaminated by oxyanions of c hlorine. Perchlorate,
chlorate, and chlorite are recognized as potential health and
environmental hazards [1–3]. In general, these compounds
are not formed naturally. Rather, their appearance in the
natural e nvironment is due to their manufacture and use as
bleaching agents, disinfectants, h erbicides, and components
of explosives and rocket propellants [4–8]. The m icrobial
decomposition of oxochlorates is important in the treatment

of pulp bleaching efflu ents [9], as well as in the degradation
of oxochlorates released into the environment b y o ther
routes [10]. Despite the fact that oxochlorates are not
formed naturally, chlorate-respiring bacteria are quite
ubiquitous [11,12].
Ideonella dec hloratans is a well-characterized species
capable of chlorate respiration [13]. Chlorate is first
converted to chlorite by a periplasmic chlorate reductase
[14]. In the second step, chlorite i s decomposed to chloride
and molecular oxygen by chlorite dismutase [15]. The
presence of chlorite dismutase i s a prerequisite for b acterial
growth as chlorite is toxic due to its high reactivity. The
oxygen produced is utilized by a c ytochrome c oxidase [13].
Chlorite dismutase h as been purified, i nitially from strain
GR-1 [16,17], and subsequently from strain CKB [18], and
from I. dechloratans [15]. Chlorite dismutases isolated from
the different species appear quite similar, being homotetra-
meric heme proteins with molecular masses around
100 k Da. T he gene encoding chlorite dismutase has been
cloned and sequen ced from two d ifferent species, I. dechlo-
ratans [19] and Dechloromonas agitata [20]. The latter
reference a lso describes a homologous gene in the genome
of Magnetospirillum magnetotacticum, but in this case no
expression of chlorite dismutase has been observed.
The I. dechloratans chlorite dismutase gene has been
expressed in Escherichia coli, and the r esulting recombinant
enzyme has b een partial ly characterized [19]. In the present
study, we present a more detailed characterization of
recombinant chlorite dismutase, a nd a c omparison with
thenativeenzyme.Ourresultssuggestthepresenceof

a post-translational modification, possibly an intrachain
covalent cross- link, in the enzyme p roduced in the n atural
host.
Materials and methods
Protein purification
Native chlorite dismutase was purified from I. dechloratans
(ATCC 51718) a s previously described [15]. Recombinant
chlorite dismutase was expressed and purified from E. coli
as described i n [19], except that the cells were homogenized
by a B ead-Beater (Biospec Products, B artlesville, USA).
Correspondence to T. Nilsson, Karlstad University, Department of
Chemistry, SE 651 88 Karlstad, Sweden. Fax: + 4 6 54 7001457,
Tel.: + 46 54 7001776, E-mail:
(Received 6 May 2004, revised 8 July 2004, accepted 14 July 2004)
Eur. J. Biochem. 271, 3539–3546 (2004) Ó FEBS 2004 doi:10.1111/j.1432-1033.2004.04290.x
Protein purity was estima ted by SDS/PAGE, and p rotein
concentration w as determined by the b icinchoninic acid
protein assay (Pierce Biotechnology, Rockford, IL, USA).
Peptide mapping by capillary electrophoresis
Recombinant and native chlorite dismutases were trans-
ferred from a SDS/PAGE gel to a polyvinylidene difluoride
membrane (ProBlott, Applied B iosystems, Stockholm,
Sweden) by electroblotting. After staining with Ponceau S,
appropriate portions were cut from the membrane and
treated with trypsin (1 : 10, 37 °C, 20 h) as described in [21].
Separation of the t ryptic fragments was performed u sing a
polydimethyl acrylamide coated fused silica capillary as
described in [22].
Peptide mass mapping
For in-gel d igestion and sample p reparation, gel plugs from

SDS/PAGE stained with Coomassie brilliant blue were
excised. In-gel digestion was carried out according to the
protocol for silver stained bands in [23] and modified as
described in [24]. Micropurification was performed accord-
ing to Ku ssmann et al. [25] and Gobom et al. [26]. Samples
were eluted directly onto a polished steel target plate with
0.8 lL a-cyano-4-hydroxycinnamic acid, 6 mgÆmL
)1
in
0.1% trifluoro acetic a cid, 30% methanol, and 30% aceto-
nitrile (premade from Agilent Technologies, Palo Alto,
USA), and left to a ir-dry.
For p eptide separation by RP-HPLC, the purified native
enzyme was also digested by trypsin in solution. The
peptides were separated b y HPLC and peak fractions were
analyzed by MALDI-MS. Native chlorite dismutase (20 lL
at 7 mgÆmL
)1
) was precipitated with 3 lL trichloroacetic
acid (100%), left 30 min on ice, and centrifuged at 10 000 g,
15 min. The precipitate was washed with cold a cetone,
vortexed and centrifuged at 10 000 g,15minandthen
resuspended in 20 lLof8
M
urea in 0.4
M
NH
4
HCO
3

,
pH 8. Water was added to a volume of 80 lL. Trypsin
(4 lg) was added, and the sample was incubated with
shaking at 37 °C, 52 h in a n Eppendorf Thermomixer. The
digest was fractioned on a Vyd ac C18 peptide column, with
a gradient of 3–97% buffer A (70% acetonitrile in 0.1%
trifluoroacetic acid, v/v), 1 mLÆmin
)1
, over 1 h. Fractions
were collected manually, subsequently dried i n a speed-vac
and resuspended in 10 lL of 0.1% (v/v) trifluoroacetic acid.
One microliter was applied to the polished steel target
(Scout 384) with 0.5 lL a-cyano-4-hydroxycinnamic acid
(Agilent) and allowed to dry (dried droplet).
Peptide mass s pectra were recorded on a Bruker
UltraFlex TOF reflector mass spectrometer (Bruker Dal-
tonics, Bremen, Germany), equipped with a nitrogen laser
(k ¼ 337 nm). The spectra were recorded in the positive
mode, using the reflector mass analyzer. Calibration was
initially performed b y external c alibration using t he
Bruker Peptide Standard. Whenever possible, internal
mass calibration was subsequently carried out on the
in-gel digestion spectra using the porcine trypsin auto-
digestion products (m/z 841 .502 and 2210.096). Data
analysis was c arried out by
M
/
Z
)
FREEWARE

, e dition
2001.08.14 (Proteomics, N ew York, NY, USA). Database
searches were carried out using
PROFOUND
(Proteomics),
searching
NCBINR
, version 2002/11/27. All chemicals were
analytical grade.
In silico enzymatic digestion of the protein sequence and
calculation of t he mass of each peptide was carried out by
MS
-
DIGEST
, ProteinProspector ( />package).
C-Terminal sequencing
A C-terminal sequencer Procise 495 C (Applied Biosystems)
was used for C-terminal sequencing of the native enzyme.
Spectrophotometric titration
Native chlorite dismutase, 6 l
M
(monomer), was diluted in
6
M
guanidinium chloride, 10 m
M
borate, 10 m
M
Tris/HCl,
pH 6. Aliquots of 1

M
sodium hydroxide w ere added to the
solution. At each pH value, the U V/visible spectrum was
recorded using a Shimadzu U V2101 spectrophotometer.
Fitting of theoretical titration curves to data was carried out
using I
GOR
(Wavemetrics, Portland, OR, USA).
Electron paramagnetic resonance (EPR) spectroscopy
EPR spectra were acquired on a Bruker ER-200D-SCR
spectrometer equipped with an Oxford Instruments ESR-9
helium cryostat. The concentrations of species giving rise to
high- a nd low-spin signals were estimated as described in
[27] and [28], respectively.
Results
Electrophoresis of proteins and tryptic peptides
We have previously reported different electrophoretic
mobilities for the native and recombinant c hlorite dismu-
tases when examined by SDS/PAGE [ 19]. The recombinant
enzyme migrates with a mobility close to that predicted by
the amino acid sequence (corresponding to a molecular
mass of 28 kDa), whereas the native enzyme migrates faster
(corresponding to a m olecular mass of 25 kDa). The
molecular mass, calculated from the DNA sequence, of
the mature prote in is 27.8 kDa. The recombinant protein
contains an extra N-terminal methionine and its predicted
molecular mass is 27.9 kDa. As we have suggested [19], a
possible explanation of the different mobilities is post-
translational processing of chlorite dismutase in I. dechlo-
ratans. Proteolytic processing at the N-terminus, however,

can be excluded from the N-terminal sequencing reported in
our earlier work [15]. In the present work, the C-terminal
sequence was also investigated, and found to be that
predicted from the gene (see below). These results exclude
proteolytic processing as an explanation for the different
mobilities of the native and recombinant enzymes.
To investigate other covalent modifications that could
affect the electrophoretic mobility, tryptic peptide maps of
native and recombinant enzymes were prepared. During
the course of this work we found that the recombinant
enzyme was less stable than the native enzyme during the
latter stages of the purification procedure, and was only
possible obtain in about 70% purity. Tryptic digests were
therefore prepared from proteins blotted from SDS gels to
3540 H. Danielsson Thorell et al. (Eur. J. Biochem. 271) Ó FEBS 2004
polyvinylidene difluoride membranes. The digests were
analyzed by capillary e lectrophoresis using a coated
capillary. F igure 1 shows electropherograms o f tryptic
digests from the native and r ecombinant enzymes. Migra-
tion times in this type of analysis are prone to variability
[29], but most of the peptide peaks seen in the electro-
pherogram of the native enzyme are also found in that
obtained from the recombinant enzyme. However, there
are clear differences, particularly at later migration times
(marked in the figure), which are not due to migration
time shifts. Thus, two peaks (denoted by arrows in Fig. 1)
in the e lectropherogram of the n ative enzyme are missing
in the electropherogram of the recombinant enzyme.
There are also two peaks in the electropherogram of the
recombinant enzyme, which do not appear to have

counterparts in the native enzyme. Our finding that
different peptide maps are obtained from the native and
recombinant enzymes suggests a difference between their
covalent structures. A lthough t he nature of such a
difference cannot be inferred from these results, we note
that anomalously high electrophoretic mobilities in SDS/
PAGE analyses have been observed in p roteins containing
covalent cross-links, such as disulfide bonds [30,31] or
cross-links caused by oxidative c oupling of sidechains
[32,33]. The higher electrophoretic mobility in these
proteins is probably due to the smaller hydrodynamic
radius caused by the cross-link.
Mass spectrometry
Detailed investigations of possible differences between
native and recombinant chlorite dismutase covalent
structure were carried out using MALDI-TOF mass
spectrometry. Tryptic peptide mass maps of the native
enzyme, from in-gel digestion and digestion in solution,
were analyzed. Masses covering most of the predicted
amino acid sequence of t he enzyme could be i dentified in
these s pectra, when a llowing four missed cleavages in the
tryptic in silico digestion. The sequence coverage based
on the mass spectra, and on C-terminal sequencing
of the n ative enzyme, is shown in Fig. 2A. Four
fragments, corresponding to HK(52–53), RK(180–181),
VPENKYHVR(215–223) and T(242) (bold) were not
covered.
To compare the native and recombinant enzymes,
peptides were generated by in-gel trypsin digestion and
subject to mass analysis using as above. For the native

enzyme, we obtained basically the s ame sequence coverage
as above. The recombinant enzyme produced, however, a
prominent peak at a mass of 1571.7, which is completely
absent in the native enzyme. A comparison of the mass
spectra obtained from the native and r ecombinant enzymes
is shown in Fig. 3. Analysis of the sequence reveals the
fragment HKEKVIVDAYLTR(52–64) (Fig. 2B) as the
probable origin of t his p eak. This fragment i ncludes
HK(52–53), which is missing in the sequence coverage of
the native enzyme. This result implicates HK(52–53) as a
possible l ocation for a covalent modification. The fragment
VPENKYHVR(215–223), also missing in the m ass spectra,
is another possible location. In the mass spectrum of
recombinant enzyme, VPENK(215–219) was absent,
whereas YHVR(220–223) was observed as a part of
fragment (220–241).
To identify modified fragments, t ryptic peptides from the
native enzyme were separated by HPLC and individually
analyzed by MS. Matching sequence coverage was obtained
after a nalysis of the mass spectra of the individual peptide
fractions. One peptide fraction from t he chromatographic
separation produced a mass spectrum containing a peak
(m/z ¼ 1679.8) (Fig. 4), corresponding to the sum (minus
one hydrogen) of the fragments containing HKEK(52–55)
and VPENKYHVR(215–223) (Fig. 2C). We could not,
however, detect a fragment at m/z ¼ 1426 corresponding to
fragment (52–53) combined wit h fragment (215–223).
Localization of a modification to fragment (52–53) is
therefore t entative.
Fig. 1. Separation of tryptic peptides of native

(A) and re combinant (B) c hlorite dismutase by
capillary e lectrophoresis with a polydimethyl
acrylamide-coated fused silica capillary.
Dashed arrows indicate correspondence, and
solid arrows denote peaks that d o not h ave
counterparts in the other e lectro pherogram.
Ó FEBS 2004 Native and recombinant chlorite dismutase (Eur. J. Biochem. 271) 3541
Spectrophotometric titration of the tyrosines in native
chlorite dismutase
Several of the covalent cross-links observed earlier [ 34]
include tyrosine sidechains, and we note the presence of
tyrosine in the VPENKYHVR fragment (215–223) that was
indicated by t he mass spectrometry analyses to be involved
in a cross-link. Cappuccio et al. [35] and McCauley et al.
[36] showed, using spectrophotometric titrations of model
compounds, t hat a cross-linked t yrosine has lowered pK
a
value of the phenolic proton. To investigate the possibility
of tyrosine sidechains with an anomalously low pK
a
value i n
chlorite dismutase, spectrophotometric t itration of the
native enzyme, completely denatured in 6
M
guanidinium
chloride, was carried out. Figure 5 shows the absorbance at
295 n m (the absorption m aximum of t he tyrosinate ion [ 37]
as a function of pH. A curve fit of a single titration curve
(Fig. 5 A) did not yield a satisfactory fit, suggesting the
presence of more than one titrating component. This is not

expected when the enzyme is completely denatured, as all
tyrosines should b e in t he same chemical environment. A
curve fit with two titrating components gave a better fit
(Fig. 5 B). The major component, accounting for 92% of the
total amplitude, t itrated with a pK
a
value of 1 0.15 ± 0.0 3,
in accordance with the pK
a
value of 10.1 for tyrosine [35].
For the minor component, accounting for 8% of the total
amplitude, a pK
a
value o f 8 .35 ± 0.3 w as found. This is
similar to the value found for the histidine methyl ester
derivative studied in [35]. Chlorite dismutase contains 12
tyrosine residues per subunit. We note that the fraction of
the minor component corresponds to about one of the 12
tyrosines titrating with the lower pKa value.
EPR
The EPR spectrum o f the recombinant c hlorite dismutase
at pH 7 is shown in Fig. 6. In contrast to the EPR
spectrum o f the native enzyme at neutral pH (trace A; see
Fig. 3. Mass analyses of tryptic peptides from
native (A) and recombinant (B) chlorite
dismutase. Only the 1558–1615 mass range is
shown.
Fig. 2. Sequences for the complete p rotein and for detected fragments
of chlorite dismutase. (A) The native chlorite dismutase amino acid
sequence with the coverage obtained by u sing MALD I-MS. The bold

and italic sequences were not detected. The sequence in it alics i s t hat
obtained in C-terminal sequencing. (B) The calculated monoisotopic
mass [MH]
+
of the peptides a re shown. (C) The monoisotopic size of
the peptides tha t would result from h istidine–tyrosine covalent linkage
(1679.92 D a).
3542 H. Danielsson Thorell et al. (Eur. J. Biochem. 271) Ó FEBS 2004
also [15]), which contains only high-spin heme, the
spectrum of the recombinant enzyme (trace B) is a
mixture of contributions from high- and low-spin species.
The high-spin heme component in the spectrum consists
of both a rhombic and an axial species with a total
concentration of 58 l
M
. The majority of the high-spin
heme has the characteristics of a rhombically distorted
heme. From th e spectrum, the g-values 6.31 and 5.47 are
obtained. Essentially the same g-values are found in the
EPR spectrum o f native chlorite dismutase at neutral pH.
A minor part of the high-spin heme is axial with a g-value
at 5.9. This axial high-spin heme is not found in the
spectrum of the native enzyme from I. dechloratans but a
similar component was observed the in EPR spectra of
chlorite dismutase from strain GR-1 recorded at neutral
pH [38]. For the l ow-spin component, the, g-values at
3.04, 2.25, and 1.52 are obtained. The integrated ampli-
tude for this signal corresponds to a concentration of
43 l
M

, which is little less than half of the total heme
concentration.
Fig. 4. MALDI mass spectrum of at HPLC
fraction of tryptic digest of nativ e chlorite
dismutase. The 1679.8 Da mass fra gment is
denoted by an arrow. The inset shows an
expanded view of the 1600–1700 m ass range.
Fig. 5. Spectrophotometric titration of tyrosine
residues in the denatured na tive chlorite dismu-
tase. The titration was monito red at 295 nm at
which only tyrosinate absorbs. The protein
contains 1 2 tyrosine residues per s ubunit. (A)
The solid line is the result of curve fitting with
A
tot
¼ 0.216 and pK
a
¼ 10.1. (B) The solid
line is the result of curve fitting with A
tot1
¼
0.206, pK
a1
¼ 10.15, A
tot2
¼ 0.017, pK
a2
¼
8.35.
Ó FEBS 2004 Native and recombinant chlorite dismutase (Eur. J. Biochem. 271) 3543

Discussion
The detailed characterization of recombinant I. dechlora-
tans chlorite dismutase, and comparison with the native
enzyme carried out here, suggest the presence of a covalent
modification in chlorite dismutase produced in the n atural
host, but not in the recombinant version of the enzyme.
Comparison of mass spectra for tryptic peptides obtained
from the native a nd recombinant e nzymes suggest HK( 52–
53) and YHVR(220–223) as sites of modification. Further-
more, a fragment, i solated by HPLC, in the tryptic digest
of the native enzyme could be identifi ed as a possible
product of cross-linking between HKEK(52–55) and
VPENKYHVR(215–223) (Fig. 2C). Cross-linking is an
attractive candidate for covalent modification, as it would
account also for the higher electrophoretic mobility (due to
the smaller hydrodynamic radius) observed for the native
enzyme, and for the different peptide maps observed after
tryptic cleavage and separation by capillary electrophoresis.
The nonenzymatic formation of covalently or oxida-
tively modified amino acids has been demonstrated
[39–41], an d s everal cases of c ross-links inc luding histidine
and tyrosine residues in oxidative enzymes have been
reported recently [34]. The crystal structure of galactose
oxidase revealed that t he enzyme contained a modified
active site tyrosine covalently cross-linked to a cysteine at
the ortho-position to t he phenolic oxygen [42]. M ore
recently, cytochrome c oxidase has also been found to
contain a modified tyrosine, with the crystal structures
showing a covalent link between the active site tyrosine
(at the ortho-position) to the imidazole N

e
of a histidine
[43,44]. A different type of histidine–tyrosine cross-link
was discovered in the crystal structure of catalase HPII
from E. coli [45,46]. In t his case, a covalent bond joins C
b
of the essential t yrosine and one of the imidazole nitrogens
(N
e
) of a histidine. These observations, together with the
presence of a t yrosine residue in the fragment suggested to
contain the cross-link (Fig. 2C), prompted us to investi-
gate the presence of m odified tyrosines. The results
obtained from spectrophotometric titration of the native
enzyme denatured in g uanidinium chloride suggest that
about one of the 1 2 tyrosines in chlorite d ismutase t itrates
with an anomalously low p K
a
value. The pK
a
value
obtained (8.3) is similar to that found for a model
histidine-phenol compound, 1-o-phenol(acetyl)histidine
methyl ester [35], and suggests the presence of a modified
tyrosine in chlorite dismutase. The result of the spectro-
photometric titration, t ogether with the mass spectrometric
data implicating the tyrosine-containing fragment VPEN-
KYHVR(215–223) as a p art of a cross-link, is consistent
the participation of tyrosine in cross-linking. From the low
pK

a
value of 8.3 found in the spectrophotometric titration,
the catalase HPII variant of cross-link is less likely as
substitution at the C
b
is not expected to affect the phenolic
pK
a
value.
An histidine–tyrosine bond may be somewhat labile [45]
and t his, in addition to ionic suppression, could explain the
rather low y ield of the dipeptide fragment mass in the MS
analyses. T he fragmented dipeptide would not necessarily
yield its const ituent two try ptic frag ment p eptide ma sses as
fragmentation may involve various parts of t he molecule
and sidechains may be derivatized. Also, the small mole-
cular mass part of the dipeptide would be prone to be
obscured in the area of the mass spectrum dominated by
signals from matrix components.
The environment of t he heme group in the recombinant
enzyme was investigated using EPR s pectroscopy. In
contrast to what is observed i n the native enzyme, the
EPR spectrum sho ws the p resence of s everal species. The
major components are a high-spin species with a spectrum
similar to that observed in the native enzyme, a nd a low-
spin species. An earlier characteriz ation using optical
spectroscopy [19] also revealed two components, one with
a native-like spectrum and one with absorption maxima at
405 and 525 nm in the oxidized state. The later species
could not be reduced by dithionite. T his species is

probably the same as the one displaying the low-spin
EPR signal. The g-values of t his component a re different
from those found for the low-spin component in the EPR
spectrum of native chlorite dismutase at high pH (2.56,
2.19, and 1.87) [15], and they are also distinct f rom those
found in other hydroxide-coordinated systems [47]. The
g-values are more similar t o those observed f or bis-
histidine coordinated heme [47]. Moreover, a similar EPR
spectrum was observed i n [38] after addition of imidazole
to chlorite dismutase from GR-1. Therefore, the heme
group is p robably coordinated by two histidine s idechains
in the low-spin component of the recombinant chlorite
dismutase. These results suggest a difference in structure of
the heme pocket in the native and recombinant e nzymes,
with a histidine sidechain being more accessible for heme
coordination from the distal side in the recombinant
enzyme. The difference between the heme environmen ts in
the native and recombinant enzymes is probably due to
structural differences caused b y covalent cross-linking.
Fig. 6. EPR spectra of native and of recombinant chlorite d ismutase at
neutral pH. (A) Native chlorite dismutase. (B) Recombinant chlorite
dismutase. Prote in concentrations we re about 100 l
M
(hem). EPR
conditions: temperature 10 K; microwave power, 2 mW; mic rowave
frequency, 9.449 GHz; modulation amplitude, 20 G.
3544 H. Danielsson Thorell et al. (Eur. J. Biochem. 271) Ó FEBS 2004
As discussed above, the histidine residue in fragment (52–
53) could b e involved in cross-linking, and an interestin g
possibility is that t his residue is available for coordination

in the recombinant enzyme where a cross-link is not
present.
Cross-links involving oxidatively coupled sidechains have
been found in enzymes operating under highly oxidizing
condition, and have been suggested to originate from
radicals formed in the reaction of the heme group with
oxidants. In cytochrome c oxidase, a tyrosyl radical is
formed during the reaction of the mixed-valence state of the
enzyme with oxygen [ 48]. MacMillan et al.[49]have
reported the EPR signal of a radical generated in cyto-
chrome c oxidase. This signal was proposed to originate
from the cross-linked tyrosine. In catalase, the compound I,
containing Fe(IV) and a porphyrin radical is produced after
the r eaction with one equivalent of hydrogen peroxide. For
catalase HPII, which contains a histidine–tyrosine cross-
link, it has been proposed that compound I is the species in
which the post-translational modification takes place
[45,50,51]. Although the catalytic mechanism of chlorite
dismutase is not known, the formation o f similar interme-
diates appears likely, given t he nature of the reactant. The
reaction of chlorite with other heme enzymes, horseradish
peroxidase and chloroperoxidase, has been shown to
produce the highly oxidized compound I [52]. M oreover,
a radical signal is present in the EPR s pectrum of chlorite
dismutase f rom strain GR-1 [38]. The formation of a c ross-
link i n chlorite dismutase by oxidative coupling, similar to
the mechanisms suggested for cytochrome c oxidase
[41,48,49] and catalase HPII [45,50,51], therefore appears
possible.
Cross-linking is expected to increase the stability of a

protein, and it absence in the recombinant enzyme could
account for the lower stability during t he latter stages of
its purification. The catalytic properties of the recombinant
enzyme are, however, similar t o those of t he native
enzyme, suggesting cross-linking is not important for
catalysis. This would be similar to cytoch rome c oxidase,
where the histidine–tyrosine cross-link has been suggested
to play role in preserving the binuclear site architecture
[40,41,53,54].
In conclusion, our comparison between the n ative and
recombinant I. dechloratans chlorite dis mutase suggests
that the enzyme produced in the natural host contains a
covalent modification, probably an intrachain cross-link
involving a residue in the 52–55 region and a residue in
the 215–223 region. A tyrosine–histidine cross-link
appears possible, and could account for EPR differences
between the native and recombinant enz ymes as well as
the spectrophotometric titration of the native enzyme.
More work is, however, needed to establish the nature of
the m odification.
Acknowledgements
We thank Roland Aasa ( Chalmers University of Technology, Sweden)
for recording the EPR spectrum and for h elpful suggestions regarding
its interpretation. We also thank Annika Norin and Ella Cederlun d
(Karolinska i nstitutet, Sweden) for C-terminal amino a cid sequencing
of the n ative enz yme, and Justyna M. Czarna for help with the mass
spectrometric analyses.
References
1. Rosemarin, A., Mattson, J., Lehtinen, K J., Notini, M. &
Nyle

´
n, E. (1986) Effects of pulp mill chlorate on Fucus vesiculosus
– a summar y of projects. Ophelia Suppl. 4, 219–224.
2. Urbansky, E.T. (1998) Perchlorate chemistry: implications for
analysis and remediation. Bioremediation J. 2, 81–95.
3. Renner, R. (2003) Environmental health: academy to mediate
debate over ro cket-fu el contaminants. Science 299, 1829.
4. A
˚
slander, A. (1928) Experiments on the eradication of canada
thistle, Cirsum arve nse, with c hlorates and other herbicides.
J. Agric. Res. 36, 915–934.
5. Germga
˚
rd, U., Teder, A. & Tormund, D. (1981) Chlorate for-
mation during chlorine dioxid e bleac hing of softwoo d kraft pu lp.
Pap. Puu 63, 1 27–133.
6. Rosemarin, A., Lehtinen, K J., Notini, M . & Mattson, J. (1994)
Effects of pulp mill chlorate on baltic sea algae. Environ. Pollut. 85 ,
3–13.
7. Herman, D.C. & Frankenberger, W.T.J. (1999) Bacterial reduc-
tion of p erchlorate a nd nitrate i n w at er. J. Environ. Qual. 28, 1018–
1024.
8. Hogue, C. (2003) Rocket-fueled river. Chem. Eng. News 81, 37–46.
9. van Wijk, D.J., Kroon, S.G.M. & Garttener-Arends, I.C.M.
(1998) Toxicity of chlorate and chlorite to selected species of algae,
bacteria, and fungi. Ecotoxicol. Environ. Safety 40 , 206–211.
10. Logan, B.E. (1998) A review of chlorate- and perchlorate-respiring
microorganisms. Bioremediation J. 2, 69–79.
11. O’Connor, S.M. & Coates, J.D. (2002) Universal immunoprobe

for (per)chlo rate-reducing b acteria. Appl. Environ. Microbiol. 68,
3108–3113.
12. Lovley, D.R. & Coates, J.D. ( 2000) Novel forms of anaerobic
respiration of environmental relevance. Curr. Opin. Microbiol. 3,
252–256.
13. Malmqvist, A
˚
., Welander, T., Moore, E., Ternstro
¨
m, A., Molin,
G. & Stenstro
¨
m, I. (1994) Ideonella dechloratans Generalnov.,
sp.nov., a new bacterium capable of growing anaerobically with
chlorate as an electron acceptor. System. Appl. Microbiol. 17,
58–64.
14. DanielssonThorell,H.,Stenklo,K.,Karlsson,J.&Nilsson,T.
(2003) A gene cluster for chlorate metabolism in Ideone lla
dechloratans. Appl. E nviron. Microbiol. 69, 5585–5592.
15. Stenklo, K., Danielsson T horell, H., B ergius, H., Aasa, R . &
Nilsson, T. (2001) Chlorite dismutase from Ideonella dechloratans.
J. Biol. Inorg. Chem. 6, 601–607.
16. Rikken, G.B., Kroon, A.G. & van Ginkel, C.G. (1996) Trans-
formation o f (per)chlorate into chloride by a newly isolated bac-
terium: red uction and dismutation. Appl. Microbiol. Biotechnol.
45, 420–426.
17. Kengen, S.W., Rikken, G.B., Hagen, W.R., van Ginkel, C.G. &
Stams, A.J. (1999) Purification and characterization of (per)-
chlorate reductase from the chlorate-respiring strain GR-1.
J. Bacteriol. 181 , 6706–6711.

18. Coates, J.D., Michaelidou, U., Bruce, R.A., O’Connor, S.M.,
Crespi, J.N. & Achenbach, L.A. (1999) Ubiquity a nd diversity of
dissimilatory (per)chlorate-reducing bacteria. Appl. Environ.
Microbiol. 65, 5234–5241.
19. Danielsson Thorell, H., Karlsson, J., Portelius, E. & Nilsson, T.
(2002) Cloning, characterisation, and expression of a novel gene
encoding chlorite dismutase from Ideonella dechloratans. Biochim.
Biophys. Acta 1577, 4 45–451.
20. Bender, K .S., O’Connor, S .M., Chakraborty, R ., Coates, J .D. &
Achenbach, L.A. (2002) Sequencing and transcriptional analysis
of the chlorite dismutase gene of Dechloromonas agitata and its u se
as a metabolic probe. Appl. Environ. Microbiol. 68, 4820–4826.
21. Walker, J.M. (1998) Protein Protocols on CD-ROM.Humana
Press Inc., Totowa, N J, USA.
Ó FEBS 2004 Native and recombinant chlorite dismutase (Eur. J. Biochem. 271) 3545
22. Wan, H., O
¨
hman, M. & Blomberg, L.G. (2001) Bonded
dimethylacrylamide as a p ermanent coating for capillary electro-
phoresis. J. Chr omatogr. A 924, 59–70.
23. Shevchenko,A.,Wilm,M.,Vorm,O.&Mann,M.(1996)Mass
spectrometric sequencing of proteins silver-stained polyacrylamide
gels. Anal. Chem. 68, 850 –858.
24. Jensen, O.N., Larsen, M.R. & Roepsto rff, P. (1998) Mass spec-
trometric identification and microcharacterization of proteins
from electrophoretic gels: strategies and applications. Proteins
Suppl. 2, 74–89.
25. Kussmann, M., Lassing, U., Sturmer, C.A., Przybylski, M. &
Roepstorff, P. (1997) M atrix-assisted laser desorption/ionization
mass spectrometric peptide mapping of th e neural cell adhesion

protein n eurolin purified by sodium dodecyl sulfate polyacryl-
amide gel electrophoresis or acid ic pre cipitation. J. Mass Spec-
trom. 32, 4 83–493.
26. Gobom, J., Nordhoff, E., Mirgorodskaya, E., Ekman, R. &
Roepstorff, P. (1999) Sample purification a nd preparation tech-
nique b ased on nano-scale reverse d-phase columns for the sensi-
tive analysis of complex peptide mixtures by matrix-assisted laser
desorption/ionization mass spectrometry. J. Mass Spe ctrom. 34,
105–116.
27. Aasa, R., Albracht, P.J., Falk, K .E., Lanne, B. & Va
¨
nnga
˚
rd, T.
(1976) EPR signals from cytochrome c oxidase. Biochim. Biophys.
Acta 422, 2 60–272.
28. Aasa, R. & Va
¨
nnga
˚
rd,T.(1975)EPRsignalandintensityand
powder shapes: a reexamination. J. M agn. Reson. 19, 308–315.
29. Grossman, P.D. & Colburn, J.C. (1992) Capillary Electrophoresis.
Academic Press, Inc, San D iego, CA.
30. Selimova, L.M., Zaides, V.M. & Zhdanov, V.M. (1982) Disulfide
bonding in influenza virus proteins as revealed by polyacrylamide
gel electrophoresis. J. Virol. 44, 450–457.
31. Shvetsov, A., Musib, R., Phillips, M., Rubenstein, P.A. & Reisler,
E. (2002) Locking the hydrophobic l oop 262–274 to G-actin sur-
face by a disulfide bridge prevents filament formation. Biochem-

istry 41, 10787–10793.
32. Baron, A.J., Stevens, C., Wilmot, C., Seneviratne, K.D., Blakeley, V.,
Dooley, D.M., Phillips, S.E., K nowles, P.F. & McPherson, M.J.
(1994) Structure and mechanism o f galactose oxidase: the free
radical site. J. Biol. Chem. 269, 25095–25105.
33. Whittaker, M.M. & Whittaker, J.W. (2003) Cu(I)-dependent
biogenesis of the galactose oxidase redox cofactor. Biol. Chem.
278, 22090–220101.
34. Okeley,N.M.&vanderDonk,W.A.(2000)Novelcofactorsvia
post-translational modifications of enzyme active sites. Chem.
Biol. 7, R159–R171.
35. Cappuccio, J.A., Ayala, I., Elliott, G.I., Szundi, I., Lewis, J.,
Konopelski, J .P., Barry, B.A. & Einarsdottir, O. (2002) Modeling
the active s ite of cytochrome o xidase: synthesis a nd characteriza-
tion of a cross-linked histidine-p henol. J. Am. Chem. Soc. 124,
1750–1760.
36. McCauley,K.M.,Vrtis,J.M.,Dupont,J.&vanderDonk,W.A.
(2000) Insights into the functional role o f t he tyrosine-histidin e
linkage in cytochrome c oxidase. J. Am. C hem. Soc. 122, 2403–
2404.
37. Tanford, C., H auenstein, J.D. & Rands, D .G. (1956) Phenolic
hydroxyl ionization in proteins II ribonuclease. J. Am. Chem. Soc.
77, 6409–6410.
38. Hagedoorn, P.L., D e G eus, D.C. & Hagen, W.R. ( 2002) S pec-
troscopic ch aracterization a nd ligand-bindin g p roperties of
chlorite dismutase from the chlorate respiring bacterial strain
GR-1. Eur. J. Biochem. 269, 4905–4911.
39. Dooley, D.M. (1999) Structure and biogenesis of topaquinone and
related cofactors. J. Biol. Inorg. Chem. 4, 1–11.
40. Rogers, M.S. & Dooley, D.M. (2001) Posttranslationally modified

tyrosines f rom galactose oxidase and c ytochrome c o xidase. Adv.
Protein Chem. 58, 387–436.
41. Rogers, M.S. & Dooley, D.M. (2003) Copper-tyrosyl radical
enzymes. Curr. Opin. Chem. Biol. 7, 189–196.
42. Ito, N., P hillips, S.E., St evens, C., Ogel, Z.B., M cPherson, M.J.,
Keen, J.N., Yadav, K.D. & Knowles, P.F. (1991) Novel thioether
bond revealed by a 1.7 A
˚
crystal structure of galactose oxidase.
Nature 350, 87–90.
43. Ostermeier, C ., Harrenga, A., Ermler, U. & Michel, H. (1997)
Structure at 2.7 A
˚
resolution of the Paracoccus denitrificans two-
subunit cytoc hrome c oxidase c omplexed with an antib ody FV
fragment. Proc. Natl Acad. Sci. USA 94, 10547–10553.
44. Yoshikawa, S., S hinzawa-Itoh, K ., Nakash ima, R., Yaono, R.,
Yamashita, E., Inoue, N., Yao, M., Fei, M.J., Libeu, C.P.,
Mizushima, T., Yamaguchi, H., Tomizaki, T. & Tsukihara, T.
(1998) Redox-c ouple d cryst al stru ctural ch ang es in bovine hear t
cytochrome c oxidase. Science 280, 1723–1729.
45. Bravo, J., Fita, I., Ferrer, J.C., Ens, W., Hillar, A. , Switala, J. &
Loewen, P.C. (1997) Identification of a novel bond b etween a
histidine and the essential tyrosine in catalase HPII of Escherichia
coli. Protein Sci. 6, 1016–1023.
46. Bravo, J., Mate, M.J., Schneider, T., Switala , J., Wilson, K.,
Loewen, P.C. & Fita, I. (1999) Struc ture of catalase HPII from
Escherichia coli at 1 .9 A
˚
resolution. Proteins 34 , 155–166.

47. Gadsby, P.M.A. & Thomson, A.J. (1990) Assignment of the axial
ligands of ferric io n in low-sp in h emoproteins by near-infrared
magnetic circular dichroism and electron paramagnetic resonance
spectroscopy. J. Am. Chem. Soc. 112, 5003–5011.
48. Proshlyakov, D.A., Pressler, M.A. & Babcock, G.T. (1998)
Dioxygen activation and bond cleavage by m ixed-valence cyto-
chrome c oxidase. Proc. N atl Acad. Sci. USA 95 , 8020–8025.
49. MacMillan, F., Kannt, A ., Behr, J., Prisner, T. & M ichel, H.
(1999) Direct evidence for a tyrosine radical in the reaction of
cytochrome c oxidase with hydrogen peroxide. Biochemistry 38,
9179–9184.
50. Mate, M.J., Sevinc, M.S., Hu, B., Bujons, J., Bravo, J., Switala, J.,
Ens, W., Loewen, P.C. & Fita, I. (1999 ) Mutants that alter the
covalent structure of c atalase hydroperoxidase II from Escherichia
coli. J. Biol. C hem. 274, 2 7717–27725.
51. Melik-Adamyan,W .,Bravo,J., Carpena,X.,Switala,J .,Mate,M.J.,
Fita,I.&Loewen,P.C.(2001)Substrate flow in catalases deduced
from the crystal structures of active site variants o f HPII from
Escherichia coli. Proteins 44, 270–281.
52. Hollenberg, P.F., Rand-Meir, T. & Hager, L.P. (1974) The r eac-
tion of chlorite with horseradish peroxidase and chloroperoxidase:
enzymatic chlorination and spectral intermediates. J. Biol. C hem.
249, 5816–5825.
53. Das, T.K ., Pec oraro, C., Tomson, F.L., Gennis, R.B. & Rousseau, D.L.
( 1998) The post-translational modification in cyto chrome c oxidase
is required to establish a functional environme nt of the catalytic
site. Biochemistry 37, 14471–14476.
54. Pinakoulaki,E.,Pfitzner,U.,Ludwig,B.&Varotsis,C.(2002)
The role o f the cross-link H is-Tyr in the functional properties of
the binuclear center in cytochrome c oxidase. J. Biol. Chem. 277,

13563–13568.
3546 H. Danielsson Thorell et al. (Eur. J. Biochem. 271) Ó FEBS 2004