Tài liệu Báo cáo khoa học: Diol dehydratase-reactivating factor is a reactivase – evidence for multiple turnovers and subunit swapping with diol dehydratase pdf

Diol dehydratase-reactivating factor is a
reactivase – evidence for multiple turnovers and subunit
swapping with diol dehydratase
Koichi Mori, Yasuhiro Hosokawa, Toshiyuki Yoshinaga and Tetsuo Toraya
Department of Bioscience and Biotechnology, Graduate School of Natural Science and Technology, Okayama University, Japan

Keywords
adenosylcobalamin; coenzyme B12; diol
dehydratase; diol dehydratase-reactivating
factor; reactivase
Correspondence
T. Toraya, Department of Bioscience and
Biotechnology, Graduate School of Natural
Science and Technology, Okayama
University, Tsushima-naka, Kita-ku,
Okayama, 700-8530, Japan
Fax: +81 86 251 8264
Tel: +81 86 251 8194
E-mail:
(Received 26 July 2010, revised
17 September 2010, accepted 1 October
2010)
doi:10.1111/j.1742-4658.2010.07898.x

Adenosylcobalamin-dependent diol dehydratase (DD) undergoes suicide
inactivation by glycerol, one of its physiological substrates, resulting in the
irreversible cleavage of the coenzyme Co–C bond. The damaged cofactor
remains tightly bound to the active site. The DD-reactivating factor reactivates the inactivated holoenzyme in the presence of ATP and Mg2+ by
mediating the exchange of the tightly bound damaged cofactor for free
intact coenzyme. In this study, we demonstrated that this reactivating
factor mediates the cobalamin exchange not stoichiometrically but catalytically in the presence of ATP and Mg2+. Therefore, we concluded that the

reactivating factor is a sort of enzyme. It can be designated DD reactivase.
The reactivase showed broad specificity for nucleoside triphosphates in the
activation of the enzymcyanocobalamin complex. This result is consistent
with the lack of specific interaction with the adenine ring of ADP in the
crystal structure of the reactivase. The specificities of the reactivase for
divalent metal ions were also not strict. DD formed 1 : 1 and 1 : 2 complexes with the reactivase in the presence of ADP and Mg2+. Upon complex formation, one b subunit was released from the (ab)2 tetramer of the
reactivase. This result, together with the similarity in amino acid sequences
and folds between the DD b subunit and the reactivase b subunit, suggests
that subunit displacement or swapping takes place upon formation of the
enzymreactivase complex. This would result in the dissociation of
the damaged cofactor from the inactivated holoenzyme, as suggested by
the crystal structures of the reactivase and DD.
Structured digital abstract
l
MINT-7997177: Reactivase alpha (uniprotkb:O68195), Reactivase beta (uniprotkb:O68196),
Diol Dehydratase gamma (uniprotkb:Q59472), Diol Dehydratase beta (uniprotkb:Q59471) and
Diol Dehydratase alpha (uniprotkb:Q59470) physically interact (MI:0915) by comigration in
non denaturing gel electrophoresis (MI:0404)
l
MINT-7997157: Diol Dehydratase alpha (uniprotkb:Q59470), Diol Dehydratase beta (uniprotkb:Q59471), Diol Dehydratase gamma (uniprotkb:Q59472), Reactivase beta (uniprotkb:
O68196) and Reactivase alpha (uniprotkb:O68195) physically interact (MI:0915) by molecular
sieving (MI:0071)

Abbreviations
AdePeCbl, adeninylpentylcobalamin; AdoCbl, adenosylcobalamin or coenzyme B12; CN-Cbl, cyanocobalamin; DD, diol dehydratase.

FEBS Journal 277 (2010) 4931–4943 ª 2010 The Authors Journal compilation ª 2010 FEBS

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Reactivase for coenzyme B12-dependent enzyme

K. Mori et al.

Introduction
Adenosylcobalamin (AdoCbl)-dependent enzymes catalyze chemically difficult reactions by the use of highly
reactive radicals. The homolytic cleavage of the Co–C
bond of the coenzyme forms a Co(II) species and an
adenosyl radical, which triggers the reactions [1].
Although enzymes generally deal with highly reactive
intermediates by ‘negative catalysis’ [2], cobalamin
enzymes tend to undergo mechanism-based inactivation because of the involvement of highly reactive radical intermediates during catalysis [3]. Diol dehydratase
(DD) (EC 4.2.1.28) catalyzes the AdoCbl-dependent
conversion of 1,2-propanediol, glycerol and 1,2-ethanediol to the corresponding aldehydes [4,5]. Its physiological substrates are 1,2-diols, such as 1,2-propanediol
[6,7], but it functionally substitutes for glycerol dehydratase (EC 4.2.1.30), an isofunctional enzyme of DD,
in the anaerobic dissimilation of glycerol by Klebsiella oxytoca and some other bacteria that lack glycerol
dehydratase [8,9]. Despite their roles, both enzymes
undergo mechanism-based inactivation by glycerol
[5,10–12], accompanying the irreversible cleavage of
the Co–C bond of the enzyme-bound coenzyme. The
damaged cofactor thus formed remains tightly bound
to the apoenzyme and is not displaced by intact AdoCbl, resulting in the irreversible inactivation of the
enzyme.
This apparent inconsistency was resolved by our
finding of the rapid reactivation of glycerol-inactivated
enzymes in permeabilized Klebsiella pneumoniae and
K. oxytoca cells (in situ) [13,14]. Specific protein factors
that are responsible for the reactivation of the
inactivated holoenzymes of DD [15–17] and glycerol

dehydratase [18–20] were found, and designated
DD-reactivating factor and glycerol dehydratasereactivating factor, respectively. They reactivated the
O2-inactivated [16,19,20] and 3-butene-1,2-diol-inactivated [21,22] holoenzymes as well. We demonstrated
that these factors reactivate the inactivated holoenzymes by a molecular chaperone-like mechanism
(Fig. 1) [16,17,19,23]. Salient features are as follows.
The reactivating factor binds ATP and hydrolyzes it to
ADP by its own weak ATPase activity. The resulting
ADP-bound form of the reactivating factor has a high
affinity for the enzyme, and interacts with the inactivated holoenzyme to form a tight apoenzymreactivating factor complex, with the concomitant release of the
damaged cofactor. The reactivating factor reverts to a
low-affinity form through the replacement of bound
ADP by free ATP, resulting in the dissociation of the
apoenzymreactivating factor complex into apoenzyme
and the reactivating factor. Active holoenzyme is then
4932

Fig. 1. Mechanism of the reactivation of inactivated holoenzymes
by reactivating factors. E, DD or glycerol dehydratase; RF, DD-reactivating factor or glycerol dehydratase-reactivating factor; X-Cbl, a
damaged cofactor; AdoH, 5¢-deoxyadenosine.

reconstituted from apoenzyme and free AdoCbl. DD
does not form a complex with the reactivating factor
while it exists as an active holoenzyme. The glycerol
dehydratase-reactivating factor reactivates the inactivated hologlycerol dehydratase in a similar manner.
Both dehydratase-reactivating factors exist as a2b2
heterotetramers [a, DdrA or GdrA (DhaF); b, DdrB or
GdrB (DhaG)] [16,19,20]. Liao et al. reported the
crystal structure of the nucleotide-free form of glycerol
dehydratase-reactivating factor [24]. Independently, we
solved the crystal structures of the DD-reactivating

factor in both the ADP-bound and nucleotide-free
forms [25]. The structures of both reactivating factors
are similar. Their a subunits have a structural feature
common to the ATPase domains of actin superfamily
proteins, including Hsp70 molecular chaperones.
Interestingly, their b subunits have similar folds to the
b subunits of diol and glycerol dehydratases. Such
structural characteristics provide important clues to
help solve the mechanisms of action of these reactivating factors – that is, subunit swapping might occur.
However, no biochemical evidence for this has been
obtained so far. A similar reactivating factor for
ethanolamine ammonia lyase has been reported [26].
It has also been reported that a protein named E2
activates lysine-5,6-aminomutase in an ATP-dependent
manner, although its exact function is not yet known
[27].
In this study, we examined whether and how the
complexes between DD and its reactivating factor
are formed. Specificities of the reactivating factor for
nucleotides and divalent cations were also investigated. In addition, it was determined whether the
reactivating factor-mediated cobalamin release is
catalytic.

FEBS Journal 277 (2010) 4931–4943 ª 2010 The Authors Journal compilation ª 2010 FEBS


Reactivase for coenzyme B12-dependent enzyme

K. Mori et al.

Results

A

Evidence for multiple turnovers of
DD-reactivating factor in cobalamin exchange
In a previous article, we reported the number of the
reactivating factor-mediated reactivations of DD
during the dehydration of glycerol [16]. The number of
reactivations per molecule of DD was calculated to be
approximately six under conditions where the reactivating factor was added to a 10-fold molar excess relative to the enzyme. This indicates that the enzyme
undergoes multiple inactivation–reactivation cycles. On
the other hand, the maximum number of reactivations
per molecule of the reactivating factor was observed to
be approximately two, at a molar ratio of the reactivating factor to the enzyme of 0.5. As the reactivating
factor exists as a dimer of ab heterodimers, i.e. (ab)2,
it remained unclear whether the reactivating factormediated reactivation of inactivated holoenzymes is
catalytic or stoichiometric. It is experimentally not
possible to demonstrate multiple turnovers for the
reactivating factor in this reactivation assay, probably
because of the inhibition of the holoenzyme by accumulated 3-hydroxypropionaldehyde.
To avoid this difficulty, we examined whether the
reactivating factor can mediate multiple turnovers of
the replacement of tightly bound cyanocobalamin
(CN-Cbl) (an inactive coenzyme analog lacking the adenine ring in the upper axial ligand; a model of damaged
cofactors) for free adeninylpentylcobalamin (AdePeCbl)
(an inactive coenzyme analog containing the adenine
ring in the upper axial ligand; a model of intact coenzyme, AdoCbl) in the presence of ATP and Mg2+ at a
molar ratio of the reactivating factor to the enzyme of
0.1 (Fig. 2). The spectrum of the enzyme obtained after

the removal of unbound cobalamins indicated that the
enzyme-bound CN-Cbl was replaced by AdePeCbl in a
manner dependent on both the reactivating factor and
ATP ⁄ Mg2+ (Fig. 2A,B). Figure 2C shows the time
course of the exchange of enzyme-bound CN-Cbl for
AdePeCbl in the presence of the reactivating factor and
ATP ⁄ Mg2+. About 70% of the enzyme-bound CN-Cbl
was replaced with AdePeCbl within 1 h, and the
replacement was almost complete within 4 h. The total
amount of enzyme-bound cobalamin remained almost
constant (1.9–2.1 molỈmol)1) during incubation. It is
thus evident that the reactivating factor mediates the
exchange of enzyme-bound CN-Cbl for free AdePeCbl
for a 10-fold molar excess of the enzyme under the
conditions employed. This strongly suggests that
the reactivation of the inactivated holoenzymes by the
factor is not stoichiometric but catalytic. Hence, the

B

C

Fig. 2. Evidence for the catalytic turnover of the DD-reactivating
factor. (A) The reactivating factor-mediated replacement of enzymebound CN-Cbl with free AdePeCbl in the presence of ATP and
Mg2+ was analyzed by the spectral change of enzyme-bound cobalamin. A 10-fold excess of the enzymCN-Cbl complex over the
reactivating factor was used. Experimental details are described in
the text. After removal of unbound cobalamin at 0 min (thick solid
line), 30 min (thin solid line), 60 min (thin long-dashed line),
120 min (thin short-dashed line), 240 min (thin dotted line) and
360 min (thick dotted line) of incubation, the spectrum of enzymebound cobalamin was measured. Inset: spectra of enzyme-bound

CN-Cbl (solid line) and AdePeCbl (dotted line). (B) Experimental conditions were the same as in (A), except that spectra were taken
after 360 min of incubation in the absence of ATP and Mg2+ (thick
solid line) or without the reactivating factor in the presence of ATP
and Mg2+ (thick dotted line). (C) Time course of the reactivating factor-mediated exchange of enzyme-bound CN-Cbl for AdePeCbl. The
extent of exchange was determined from the change in absorbance at 364 nm. The total amount of enzyme-bound cobalamin
was determined spectrophotometrically after conversion to the
dicyano form. Inset: a semilogarithmic plot.

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Reactivase for coenzyme B12-dependent enzyme

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DD-reactivating factor can be designated DD reactivase
as well. The cobalamin exchange occurs through the
intermediary formation of apoenzyme [17,19]. As AdePeCbl binding to apoenzyme takes place much faster
than CN-Cbl release, the rate of CN-Cbl replacement
can be considered to be the rate of CN-Cbl release. The
inset of Fig. 2C indicates that the rate of CN-Cbl
replacement (release) follows pseudo-first-order kinetics, and the rate constant of the reactivase in cobalamin
release (kcat,cbl-release) for CN-Cbl was calculated to be
0.27 min)1 at 37 °C from the initial rate.
The time course of the reactivation of glycerol-inactivated holoenzyme by the reactivase in the presence of
ATP and Mg2+ at a molar ratio of the reactivase to the
enzyme of 0.1 is shown in Fig. 3. From the initial rate,
the rate constant of the reactivase in the reactivation

(kcat,react) was calculated to be 0.071 ± 0.008 min)1 at
37 °C. Considering that the enzyme contains two cobalamin-binding sites in the (abc)2 dimer, it can be
assumed that the reactivase mediates the exchange of
enzyme-bound damaged cofactor for intact AdoCbl
with a rate constant (kcat,cbl-release) of 0.14 min)1.
Kinetic parameters of the reactivase for ATP in
DD (re)activation and ATP hydrolysis
Kinetic constants for ATP in the reactivation of glycerol-inactivated holoenzyme and the activation of the

enzymCN-Cbl complex by the reactivase were measured (Table 1). Km values for ATP in the reactivation
and the activation were essentially the same:
6.9 ± 0.4 mm and 6.8 ± 1.6 mm, respectively. This is
reasonable, because these two events are different
aspects of the same phenomenon [14,16]. Km values for
the ATPase activity were also measured in the presence
and absence of equimolar apoenzyme (Table 1). The Km
for ATP in the ATPase activity in the absence of enzyme
was 61 ± 14 lm, i.e. two orders of magnitude smaller
than that in the DD (re)activation. Moreover, the Km
for ATP in the ATPase activity was essentially not
affected by the presence of enzyme (67 ± 17 lm).
The kcat in the ATPase activity was estimated to be
1.4 ± 0.1 min)1 in the absence of apoenzyme. The
ATPase activity was slightly inhibited by the presence of
apoenzyme (kcat = 1.2 ± 0.1 min)1). These values are
in good agreement with those previously reported [17].
Nucleotide and divalent cation specificities of
the reactivase
The specificities of the reactivase for nucleotides in the
activation of the enzymCN-Cbl complex were studied

in the presence of AdoCbl and Mg2+ (Table 2).
Although ATP was most effective (40% as compared
with the apoenzyme control), CTP, UTP and GTP
showed comparable efficiencies (82%, 75% and 55%
relative to ATP, respectively). Moreover, 2¢-dATP and
3¢-dATP were 55% and 63% as effective as ATP.
The efficiencies of various divalent metal ions for the
activation of the enzymCN-Cbl complex by the
reactivase were also examined at 3 mm (Table 2). The
Table 1. Kinetic parameters of the reactivase for ATP.

Km for
ATP (mM)
Reactivationa
Activationa
ATPase (+DD)b
ATPase ()DD)b
Fig. 3. Time course of the reactivation of the glycerol-inactivated
holoenzyme by DD reactivase. The glycerol-inactivated holoenzyme
formed as described in the text was subjected to ultrafiltration on a
Microcon YM-10 microconcentrator (Millipore). To a concentrated
protein fraction containing 1.2 nmol of glycerol-inactivated holoenzyme, we added 2.3 M 1,2-propanediol, 38 lM AdoCbl, 19 mM ATP
and 19 mM MgCl2 in 0.02 M potassium phosphate buffer (pH 8.0)
without or with 0.12 nmol of reactivase to a total volume of
160 lL. After incubation at 37 °C for the indicated time periods,
20 lL aliquots were withdrawn, and the amount of DD reactivated
was measured by the 3-methyl-2-benzothiazolinone hydrazone
method [33] after appropriate dilution.

4934

6.9
6.8
0.067
0.061

±
±
±
±

0.4
1.6
0.017
0.014

Vmax (lmol
propionaldehyde
formed in 10 min)

kcat (min)1)

2.2 ± 0.6
17 ± 2
1.2 ± 0.1
1.4 ± 0.1

a
The glycerol-inactivated holoenzyme (31 pmol) or the enzymCNCbl complex (53 pmol) was incubated at 37 °C for 10 min with
0.21 nmol of reactivase in 70 lL of 0.02 M potassium phosphate buffer (pH 8.0) containing 0.6 M 1,2-propanediol, 0.01 M KCl, and 21 lM

AdoCbl, with 0–40 mM each of ATP and MgCl2. The reaction was
terminated by addition of 70 lL of 0.1 M potassium citrate buffer
(pH 3.6). The amount of propionaldehyde formed was determined as
described in the text after appropriate dilution. b The reactivase
(0.22 nmol) was incubated at 37 °C for 5 min with or without apoenzyme (5.0 units, 0.20 nmol) in 50 lL of 0.01 M potassium phosphate
buffer (pH 8.0) containing 0.3–10 mM each of [32P]ATP[cP] and
MgCl2. ATPase activity was measured as described in the text.

FEBS Journal 277 (2010) 4931–4943 ª 2010 The Authors Journal compilation ª 2010 FEBS


Reactivase for coenzyme B12-dependent enzyme

K. Mori et al.

Table 2. Nucleotide and divalent cation specificities of the reactivase for the activation of the enzymCN-Cbl complex. The enzymCN-Cbl
complex (DDỈCN-Cbl) (61 pmol) was incubated at 37 °C for 10 min with and without 0.30 nmol of reactivase in 50 lL of 0.02 M potassium
phosphate buffer (pH 8.0) containing 21 lM AdoCbl and 1.2 M 1,2-propanediol in the presence and absence of 3 mM ATP (or an indicated
nucleotide) and 3 mM MgCl2 (or a chloride salt of the indicated divalent metal ions). The reaction was terminated by addition of 50 lL of
0.1 M potassium citrate buffer (pH 3.6). The amount of propionaldehyde formed was determined as described in the text after approriate
dilution.
Propionaldehyde
formed
Run no.

Enzyme

Reactivase

Nucleotide

Metal ion

1

apoDD
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
apoDD
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl
DDặCN-Cbl

)
+
+
+
+
+
+

+
)
+
+
+
+
+
+
+
+

)
)
ATP
GTP
CTP
UTP
2Â-dATP
3Â-dATP
)
ATP
ATP
ATP
ATP
ATP
ATP
ATP
ATP

)

)
Mg2+
Mg2+
Mg2+
Mg2+
Mg2+
Mg2+
)
)
Mg2+
Ca2+
Cr2+a
Mn2+
Co2+
Ni2+
Cu2+

13.4
0.1
5.3
3.0
4.4
4.0
2.9
3.4
13.7
0.0
6.0
0.1
0.1

11.1
5.8
2.9
0.1

(%)

(lmol)

2

a

1.6
0.1
1.2
0.4
1.0
0.8
0.6
0.1
1.8
0.0
0.2
0.1
0.1
0.4
0.1
0.7
0.0

Relative
activity (%)

100
1
40
22
33
30
22
25

100
0
44
1
1
81
42
21
1

)
)
100
57
83
75
55
64
)
)
100
2
2
185
97
48
2

Cr2+ added might be oxidized to Cr3+ by air in the reaction mixture.

divalent cations tested did not inhibit the DD activity at
3 mm in the standard assay conditions (data not shown).
Mn2+ was most effective for the activation (81% as
compared with the apoenzyme control; 185% relative to
Mg2+). Co2+ and Ni2+ were also effective (97% and
48%, respectively, relative to Mg2+), whereas Ca2+,
Cr2+ (Cr3+) and Cu2+ were not (< 2% relative to
Mg2+). Mg2+ was used routinely in the (re)activation
assay, because it is a physiological divalent metal ion.
The specificity of the reactivase for divalent metal ions
in ATP hydrolysis was also measured (Table 3).
Although the reactivase hydrolyzed ATP to some extent
even in the absence of divalent metal ions, Mg2+,
Mn2+, Co2+ and Ni2+ enhanced the ATPase activity
by 3.7–4.5 fold at 3 mm. On the other hand, Ca2+, Cr2+
(Cr3+) and Cu2+ had little or no effect on the ATPase
activity as compared with the control without divalent
metal ions. Irrespective of the presence of divalent
cations, ATP was hydrolyzed to ADP + Pi by the reactivase (data not shown).
Analysis of complex formation between DD and
its reactivase by gel filtration
Apoenzyme was incubated with the reactivase in the
presence of ADP or ATP and Mg2+, and then subjected

Table 3. Divalent cation specificity of the reactivase for the ATPase
activity. The reactivase (0.22 nmol) was incubated at 37 °C for
30 min with 3 mM [32P]ATP[cP] in 50 lL of 0.01 M potassium phosphate buffer (pH 8.0) in the presence and absence of the indicated
divalent metal chloride (3 mM). The ATPase activity was measured
as described in the text.

Metal ion

ATPase
activity (min)1)

Relative
activity (%)

Mg2+
Ca2+
Cr2+a
Mn2+
Co2+
Ni2+
Cu2+
–

1.99
0.52
0.38
2.23
2.14
2.42
0.80
0.54

100
26
19
112

107
122
40
27

±
±
±
±
±
±
±
±

0.04
0.01
0.01
0.01
0.04
0.09
0.05
0.02

a

Cr2+ added might be oxidized to Cr3+ by air in the reaction
mixture.

to gel filtration on a Superose 6 column that had
been preliminarily equilibrated with nucleotide ⁄ Mg2+containing buffer (Fig. 4). In the presence of ATP, the

enzyme and the reactivase eluted separately at their
respective retention times. In contrast, a peak of the free
reactivase decreased and a new peak appeared in the
presence of ADP. The latter peak was eluted with a

FEBS Journal 277 (2010) 4931–4943 ª 2010 The Authors Journal compilation ª 2010 FEBS

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Reactivase for coenzyme B12-dependent enzyme

K. Mori et al.

A

B

Fig. 4. Analysis of interaction between DD and its reactivase by
gel filtration column chromatography. Experimental details are
described in the text. 1, apoenzyme + reactivase (+ADP); 2, apoenzyme (+ADP); 3, reactivase (+ADP); 4, apoenzyme + reactivase
(+ATP). Positions of the free apoenzyme (DD), reactivase (DD-R)
and the enzymreactivase complex(es) (DDỈDD-R) are indicated on
the tops of chromatograms.

66 K–

C

retention time that was slightly shorter than that of the

free enzyme and much shorter than that of the reactivase. When this peak was subjected to SDS ⁄ PAGE, the
peak comprised all of the subunits from the enzyme
(a, b, and c) and the reactivase (a and b) (data not
shown). It was thus evident that this peak contained the
enzymreactivase complex(es) in addition to a small
amount of the free enzyme. In fact, when this peak was
analyzed by nondenaturing PAGE, two bands of the
enzymreactivase complexes described below were
detected (data not shown). A similar peak containing
the enzymreactivase complexes was observed in the
absence of nucleotide as well, although the content of
complexes was rather lower than in the presence of
ADP.
Characterization of the enzymreactivase
complexes by nondenaturing PAGE
When the enzymCN-Cbl complex was incubated with
the reactivase in the presence of ADP and Mg2+ and
followed by nondenaturing PAGE, the bands of the
enzyme and the reactivase were markedly reduced
in density, and two bands of the enzymreactivase
complexes (upper and lower) appeared above them
(Fig. 5A, lane c). When apoenzyme was used instead of
the enzymCN-Cbl complex, both bands of the
enzymreactivase complexes were observed in the
presence of ADP (Fig. 5A, lane e). The abundance of
4936

66 K–
45 K–
36 K–

29 K–
24 K–
20 K–
14 K–

Fig. 5. Analysis of the enzymreactivase complex by PAGE. Experimental details are described in the text. (A) Nondenaturing PAGE
in the presence of ADP and Mg2+. Lanes: (a) apoenzyme; (b) reactivase; (c) enzymCN-Cbl complex + reactivase (+ADP); (d) enzymCN-Cbl complex (+ADP); (e) apoenzyme + reactivase (+ADP);
(f) apoenzyme (+ADP); (g) reactivase (+ADP). +ADP indicates that
samples were incubated with ADP ⁄ MgCl2. (B, C) Bands i–vi in (A)
were excised and subjected to SDS ⁄ PAGE on 6% (B) and 14% (C)
gels. Lanes i–vi correspond to bands i–vi in (A). Lane D: purified
enzyme. Lane R: purified reactivase. Positions of DD and its subunits (aD, bD and cD) and the reactivase (DD-R) and its subunits (aR
and bR) are indicated on the right of the gels. In (C), to improve visibility of the bands of small subunits, especially bR, three excised
pieces of the same bands from nondenaturing PAGE were subjected together to SDS ⁄ PAGE. This resulted in saturation of the
bands of large subunits, i.e. aD and aR. Densitometric analysis was
carried out with other gels in which such saturation did not occur
(not shown).

the upper band relative to the lower band of the
enzymreactivase complex formed from apoenzyme
was significantly larger than that formed from the

FEBS Journal 277 (2010) 4931–4943 ª 2010 The Authors Journal compilation ª 2010 FEBS


Reactivase for coenzyme B12-dependent enzyme

K. Mori et al.

enzymCN-Cbl complex. These results are consistent

with previous data [17]. Three bands were clearly seen in
the lanes containing the free reactivase, i.e. two adjacent
thick bands and one thin band on the front line, irrespective of the presence of enzyme (Fig. 5A, lanes b, c,
e, and g). To determine their subunit compositions,
bands i–vi were excised and subjected to SDS ⁄ PAGE
on 6% and 14% gels (Fig. 5B,C), followed by densitometric analyses. Both band i and band vi of the
enzymreactivase complexes comprised all of the
subunits from the enzyme (a, b, and c) and the reactivase (a and b) (Fig. 5B,C, lanes i and vi). Subunit
compositions of the bands were the same when either
the enzymCN-Cbl complex or apoenzyme was used
(data not shown). If the a, b and c subunits of the
enzyme are abbreviated as aD, bD, and cD, respectively,
and the a and b subunits of the reactivase are abbreviated as aR and bR, respectively, molar ratios of aD, bD,
cD, aR and bR in bands i and vi were determined to be
about 2 : 2 : 2 : 2 : 1 and 1 : 1 : 1 : 2 : 1, respectively,
by densitometric analysis. Therefore, it was demonstrated that bands i and vi are (aDbDcD)2Ỉ(aRỈaRbR) and
(aDbDcD)2Ỉ(aRỈaRbR)2 complexes, respectively. We
named the former the enzymreactivase (1 : 1) complex
and the latter the enzymreactivase (1 : 2) complex.

When bands ii and iii of Fig. 5A, i.e. two thick bands of
the reactivase, were subjected to SDS ⁄ PAGE, they contained both aR and bR, although the ratios of subunits
were different (Fig. 5C, lanes ii and iii). Densitometric
analysis indicated that molar ratios of aR to bR for
bands ii and iii were approximately 2 : 1 and 1 : 1,
respectively. These results indicated that the upper and
lower bands of the reactivase correspond to the aRỈaRbR
and (aRbR)2 complexes, respectively. The thin band on
the front line of nondenaturing PAGE (Fig. 5A, band
iv) contained only bR (Fig. 5C, lane iv). It possibly

represents a monomer of bR, as there is no direct
interaction between two adjacent bR subunits in the
crystal structure of the (aRbR)2 tetramer of the reactivase [25]. In the absence of ADP ⁄ Mg2+ or in the
presence of ADP but in the absence of Mg2+, the
enzymreactivase complex was formed in small
amounts from apoenzyme and the reactivase and not at
all from the enzymCN-Cbl complex and the reactivase
(data not shown).
Affinity of the reactivase for DD
Figure 6 shows the dependence of enzymreactivase
complex formation on reactivase concentration. The

A

Fig. 6. Dependence of complex formation
on reactivase concentration at a fixed
enzyme concentration. (A) Apoenzyme +
reactivase (left, none; right, +ADP).
(B) EnzymCN-Cbl complex + reactivase
(left, none; right, +ADP). Experimental conditions were similar to those for nondenaturing
PAGE in Fig. 5, except that the reactivase
and enzyme concentrations were varied and
fixed (1 lM), respectively. The number on the
top of each lane indicates the reactivase
concentration (lM). Lane R: reactivase 2 lM.
Positions of the enzyme, reactivase and the
enzymreactivase complexes are indicated
on the right of the gels: (i) enzymreactivase
(1 : 2) complex; (ii) enzymreactivase (1 : 1)
complex; (iii) enzyme; (iv) reactivase

(aRỈaRbR); (v) reactivase [(aRbR)2]; (vi) small
subunit of the reactivase (bR).

B

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K. Mori et al.

enzymreactivase (1 : 1) complex (band ii) was formed
even at the lowest concentration of reactivase tested
(0.25 lm) in the presence of ADP and Mg2+. In the
absence of ADP and Mg2+, it was observable at
‡ 0.5 lm reactivase. Similarly, the enzymreactivase
(1 : 2) complex (band i) appeared clearly at 1 lm reactivase in the presence of ADP and Mg2+, whereas it was
observed at ‡ 2 lm reactivase in the absence of ADP
and Mg2+. Moreover, although the enzymreactivase
(1 : 2) species was the only enzymreactivase complex
observed at ‡ 4 lm reactivase in the presence of ADP
and Mg2+, some enzymreactivase (1 : 1) complex
remained even at the highest concentration of reactivase
tested (20 lm) in the absence of ADP and Mg2+. The
apparent KD values of the reactivase for formation of
the enzymreactivase complex were 0.4 lm and 3 lm in
the presence and absence of ADP and Mg2+, respectively. When similar experiments were carried out with

the enzymCN-Cbl complex in place of apoenzyme,
essentially no complex formation was observed, even at
20 lm reactivase, in the absence of ADP and Mg2+. In
contrast, in the presence of ADP and Mg2+, the reactivase formed complexes with DD, accompanying the
release of tightly bound CN-Cbl from the enzyme [17].
However, the enzymreactivase (1 : 1) complex
remained at 20 lm reactivase. The apparent KD of the
reactivase was 0.7 lm.

Discussion
In the present study, we demonstrated the multiple
turnovers of the DD-reactivating factor in the in vitro
activation of the inactive enzymCN-Cbl complex,
and thus redesignated the reactivating factor [16,17]
DD reactivase. This is reasonable in vivo from the
viewpoint of the cellular economy of energy. If the
reactivating factor could not mediate the multiple
exchanges in the reactivation of inactivated holoenzymes, its presence would not be advantageous to the
bacterial cells. We have previously demonstrated that
the hydrolysis of ATP by the reactivating factor is
catalytic [17]. Moreover, the reactivation observed in
permeabilized cells of K. oxytoca seems to be catalytic
[13,14], although molar ratios of the reactivating factor
to the enzyme remain obscure. We previously failed to
demonstrate the multiple turnovers of the reactivase in
the reactivation of inactivated holoenzymes. One possible reason for this difficulty might be the accelerated
rate of inactivation of the enzyme with b-hydroxypropionaldehyde accumulating to an extremely high
concentration (20–180 mm), as the reactivation was
monitored by product formation from reactivated
holoenzyme at high concentrations of the enzyme and

4938

the reactivase. It would be easier to demonstrate the
multiple turnovers of DD reactivase and glycerol
dehydratase reactivase in the in situ reactivation,
because the reactivation takes place in toluene-treated
cells, where local concentrations of the enzyme and the
reactivase are high enough for reactivation, and an
inhibitory product, b-hydropropionaldehyde, diffuses
away.
From the initial rate of exchange of enzyme-bound
CN-Cbl for AdePeCbl, the rate constant of the reactivase in cobalamin release (kcat,cbl-release) for CN-Cbl
was calculated to be 0.27 min)1 at 37 °C. From the
initial rate of reactivation of the glycerol-inactivated
holoenzyme, the rate constant of the reactivase in
the reactivation (kcat,react) was calculated to be
0.071 ± 0.008 min)1 at 37 °C. Considering that the
enzyme contains two cobalamin-binding sites in the
(abc)2 dimer, it can be assumed that the reactivase
mediates the exchange of enzyme-bound damaged
cofactor for intact AdoCbl with a rate constant
(kcat,cbl-release) of 0.14 min)1. This value is about half of
the above-mentioned kcat,cbl-release for CN-Cbl release.
This difference might be attributable to the difference
in release rate between CN-Cbl and the damaged
cofactor. Another possible explanation is that the
enzyme activity of the reconstituted abcỈAdoCbl
complex in one trimer might be affected by the neighboring trimer of the same enzyme molecule, i.e. by the
presence of the damaged cofactor or AdoCbl and their
absence. The kcat of the reactivase in ATP hydrolysis

in the presence of enzyme (1.2 min)1) was slightly
smaller than that in its absence (1.4 min)1). It was in
the same range as the rate constant of the enzyme in
the suicide inactivation with glycerol (1.3 min)1) [17],
but about five-fold and 10-fold larger than the rate
constants for the release of CN-Cbl (0.27 min)1) and
the damaged cofactor (0.14 min)1), respectively.
Therefore, ATP hydrolysis and cobalamin release or
reactivation might be not very tightly coupled. The
reactivation of the inactivated holoenzymes by the
reactivase seems to be physiologically relevant, because
kcat,cbl-release is much larger than the rate constant for
bacterial growth on glycerol.
The reactivase exhibited broad specificities for nucleotides and divalent metal cations, both of which are
absolutely required for the in vitro activation of the
enzymCN-Cbl complex. We have previously reported
similar specificities in the in situ reactivation of the
glycerol-inactivated hologlycerol dehydratase with
K. pneumoniae cells [13]. It was established that the
reactivase-mediated reactivation of the inactivated
holoenzymes with ATP and Mg2+ takes place in two
steps: (a) ADP-dependent cobalamin release with

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K. Mori et al.

concomitant formation of the apoenzymreactivase
complex; and (b) ATP-dependent dissociation of the

complex to apoenzyme and the reactivase [17]. ATP
plays dual roles, i.e. as a precursor of ADP in the first
step, and an effector to change the reactivase to a form
with low affinity for the enzyme. The nucleotides used
in this study (ATP, GTP, CTP, UTP, 2¢-dATP, and 3¢dATP) were effective in overall activation, although
the efficiencies were somewhat different. This suggests
that these nucleotides are also effective in both steps.
We determined the crystal structure of the reactivase
in the ADP-bound and nucleotide-free forms [25].
ADP is bound to the ATPase domain, a core domain
of the a subunit. This domain shares common
structural features with the ATPase domain of actin
superfamily proteins, including Hsp70 molecular chaperones. The reactivase binds ADP without specific
interactions with the adenine ring through hydrogen
bonding or base stacking. The broad specificity of the
reactivase for the base moiety is thus consistent with
its crystal structure. The O2¢ atom of ADP is hydrogen
bonded to the –COO) group of Glua459 and the e-NH2
group of Lysa462. These hydrogen bonds exist in the
interaction between Hsc70 and ADP as well. The residue corresponding to Glua459 of the DD reactivase is
Alaa461 in glycerol dehydratase reactivase. Furthermore, 2¢-dATP retained half of the efficacy of ATP in
the activation of the enzymCN-Cbl complex. It was
therefore concluded that these hydrogen bonds are not
essential for (re)activation. Similarly, 3¢-dATP retained
half of the efficacy of ATP in the activation. In the crystal structure of the reactivase, no amino acids were
found to be hydrogen bonded to O3¢ of ADP. Thus, no
requirement for the 3¢-OH group seems to be reasonable
from its crystal structure.
The reactivase has two distinct divalent metal ionbinding sites in the ab heterodimeric unit [25]. One of
them is present in the interface between the a and

b subunits. This metal ion is coordinated by four
amino acids (Aspa166, Aspa183, Thra105, and
Glub31), all of which are completely conserved in both
reactivases for diol and glycerol dehydratases. These
coordinations are maintained in the reactivase, irrespective of the ADP binding. The crystal structure of
the DD reactivase suggested that this metal ion is
Mg2+ in the ADP-bound form, whereas it is Ca2+ in
the nucleotide-free form. It might be possible that
Mg2+ occupies this site in vivo and is replaced by
Ca2+ in the purification of the reactivase by hydroxyapatite column chromatography [25]. Liao et al. also
reported this metal ion to be Ca2+ in the nucleotidefree form of glycerol dehydratase reactivase that is
crystallized in the presence of Ca2+ [24]. In the case of

Reactivase for coenzyme B12-dependent enzyme

the ADP-bound form, Ca2+ in this site may be
replaced by Mg2+ upon incubation of the reactivase
with ADP and Mg2+. The other divalent metal ion is
Mg2+, which interacts with the b-phosphate group of
ADP in the nucleotide-binding site of the a subunit.
This Mg2+ was not found in the nucleotide-free form.
Mg2+, Mn2+, Co2+ and Ni2+ enhanced the ATPase
activity of the reactivase, although the reactivase can
hydrolyze ATP to ADP even without divalent metal
ions. These metal ions were effective in the activation
of the enzymCN-Cbl complex by the reactivase in the
presence of ATP, although relative efficiencies were
not always correlated. On the other hand, the reactivase was unable to activate the enzymCN-Cbl complex even in the presence of ATP with Ca2+, Cr2+or
Cu2+ or without divalent metal ions. These metal ions
had little or no enhancing effect on the ATPase activity of the reactivase. Thus, the reactivase-mediated

activation of the enzymCN-Cbl complex absolutely
requires the hydrolysis of ATP in the presence of divalent metal ions. The reactivase does not form the
enzymreactivase complexes from the enzymCN-Cbl
complex in the presence of ADP without divalent
cations. These results suggest that the binding of ADP
alone to the ATPase domain of the reactivase a subunit is not sufficient to cause a conformational change
of the enzyme, resulting in the release of adenine-lacking cobalamins, such as CN-Cbl and damaged cofactor. The fact that the relative efficiencies of metal ions
for the reactivation are not always correlated with the
ATPase activity of the reactivase might be attributable
to the different characteristics of binding of these divalent ions to the other metal ion-binding site in the
interface between the a and b subunits, although the
binding specificity of this site for metal ions remains
unclear.
We have previously demonstrated the formation of
the enzymreactivase complex from apoenzyme and
the reactivase in the presence of ADP ⁄ Mg2+ or in the
absence of nucleotide, although the exact subunit compositions of the resulting complexes remained unclear.
Our present study indicated that two kinds of complexes with different subunit compositions were
formed. These complexes contain the enzyme and the
reactivase in 1 : 1 and 1 : 2 molar ratios, and release
of the reactivase b subunit was observed upon complex formation (Fig. 7A). These results constitute clear
evidence for the displacement of the reactivase b subunit by the enzyme b subunit (subunit swapping) upon
formation of the complex between the enzyme and the
reactivase. At present, it is not clear which complex is
involved or whether both complexes are involved in
the reactivation. The dissociation of (aRbR)2 into

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4939

Reactivase for coenzyme B12-dependent enzyme

K. Mori et al.

cofactor to pass through it. Intact cofactor, an adenine-containing cobalamin, is not released from the
enzyme by the reactivase. One reason might be its
larger size, and the other possible reason might be that
the additional interaction between its adenine moiety
and the enzymes’s adenine-binding pocket stabilizes
the interaction between the enzyme a and b subunits.
In contrast, even in the absence of ADP, the reactivase
forms the enzymreactivase complex with apoenzyme.
However, it does not release the damaged cofactor
from the inactivated holoenzymes under this condition. This may be because the steric repulsion is less
or is canceled by the conformational flexibility in the
absence of ADP ⁄ Mg2+. In order to prove or disprove
these predictions, we have to await the structural
analysis of a real enzymreactivase complex.

A

B

Experimental procedures
Fig. 7. Subunit swapping between DD and the reactivase (DD-R)
(A) and the existence of a cavity between DD a (pink) and b (green)
subunits (B). a, aD (pink) or aR (light blue) subunit; b, bD (green) or
bR (orange) subunit; c, cD subunit (dark blue).

aRỈaRbR and bR in the presence of ADP and Mg2+
was observed even without the enzyme. The crystal
structure of the reactivase also suggested that the
interactions between the reactivase a and b subunits
are weakened at least partially by the ADP binding
[25]. The space that is opened by the dissociation of
the reactivase b subunit would most likely be occupied
by the enzyme b subunit, as these subunits have similar folds [25,28]. The docking model of the aDbDcDỈaR
complex indicates that marked steric repulsion is
induced between the enzyme a subunit and the reactivase a subunit in the complex [25]. The amino acid
side chains that come closer than the van der Waals
contact in the modeled structure would push each
other aside and result in tilting of the enzyme a subunit with respect to the enzyme b subunit. Thus, it
would lead to the release of the damaged cofactor, an
adenine-lacking cobalamin, from the enzyme, because
cobalamin is bound between the a and b subunits of
the enzyme. The crystal structure of DD revealed that,
like ethanolamine ammonia lyase [29], the enzyme has
˚
˚
a cavity  5 A in height and  15 A in width between
the a and b subunits (Fig. 7B). The tilting of the
a subunit with respect to the b subunit upon subunit
˚
swapping is estimated to be  6 A, based on the mod˚
eled complex, forming a cavity  11 A in height. The
size of this cavity is comparable with that of adeninelacking cobalamins, and thus allows the damaged
4940

Materials
Crystalline AdoCbl was a gift from Eisai (Tokyo, Japan).
CN-Cbl was obtained from Glaxo Research Laboratories
(Greenford, UK). AdePeCbl was synthesized according to
published procedures [30]. [32P]ATP[cP] was obtained from
PerkinElmer (Waltham, MA, USA). 2¢-DeoxyATP and
3¢-deoxyATP were obtained from Sigma-Aldrich (St Louis,
MO, USA). All other chemicals were commercial products
of the highest grade available and were used without
further purification. K. oxytoca recombinant DD and its
reactivase were purified to homogeneity from overexpressing Escherichia coli JM109 harboring expression plasmid
pUSI2E(DD) [31] and E. coli JM109 or B834 harboring
expression plasmid pUSI2ENd(6 ⁄ 5b) [16,32], respectively,
as reported previously.

Enzyme and protein assays
The amount of aldehydic products formed by DD was
determined by the 3-methyl-2-benzothiazolinone hydrazone
method [33]. One unit of the enzyme is defined as the
amount of enzyme activity that catalyzes the formation of
1 lmol of propionaldehyde per minute at 37 °C. The
reactivation of the glycerol-inactivated holoenzyme and the
activation of the enzymCN-Cbl complex by the reactivase
were assayed with 1,2-propanediol as substrate in the
presence of 21 lm AdoCbl and appropriate concentrations
of ATP and MgCl2. In some experiments, ATP and MgCl2
were replaced with other nucleotides and chloride salts of
divalent metal cations, respectively. The protein concentrations of the purified enzyme and reactivase were determined
by measuring the absorbance at 280 nm, based on the
method of Gill and von Hippel [34], as described previously

[16].

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K. Mori et al.

Glycerol-inactivated holoenzyme and
enzymCN-Cbl complex
The glycerol-inactivated holoenzyme was formed by incubation of apoenzyme (70 units, 2.8 nmol) with 38 lm AdoCbl
at 37 °C for 30 min in 1.7 mL of 0.05 m potassium phosphate buffer (pH 8.0) containing 19% glycerol. Glycerol
and excess AdoCbl were removed by dialysis at 4 °C for
49 h against 600 volumes of 0.05 m potassium phosphate
buffer (pH 8.0) containing 2% 1,2-propanediol with a
buffer change. The enzymCN-Cbl complex was prepared
by incubation of apoenzyme (30–45 units, 1.2–1.8 nmol)
with 13 lm CN-Cbl at 37 °C for 30 min in 540 lL of
0.05 m potassium phosphate buffer (pH 8.0) containing 2%
1,2-propanediol and 0.6–1% Brij35.

ATPase activity
The ATP-hydrolyzing activity of the reactivase was assayed
by the release of [32P]Pi from [32P]ATP[cP], as described previously [17], with minor modifications. Appropriate concentrations of [32P]ATP[cP] ( 6 · 103 d.p.m. per nmol) and
MgCl2 were incubated at 37 °C for appropriate times with
the reactivase, in the presence and absence of apoenzyme, in
0.01 m potassium phosphate buffer (pH 8.0) in a total volume of 50 lL. After termination of the reaction by addition
of 0.45 mL of an ice-cold suspension of 6% (w ⁄ v) charcoal
in 50 mm NaH2PO4 and mixing vigorously for 10 min, the
charcoal was removed by centrifugation at 16 000 g for
5 min. The amount of radioactivity in 0.2 mL of the supernatant was determined by liquid scintillation counting, and

ATPase activity was obtained by subtracting the radioactivity of a minus reactivase control. In some experiments,
MgCl2 was replaced with other divalent metal chlorides.

Reactivase-mediated exchange of enzyme-bound
CN-Cbl for AdePeCbl
The enzymCN-Cbl complex was formed by incubation of
30 units of apoenzyme (1.2 nmol) with 80 lm CN-Cbl at
37 °C for 60 min in 75 lL of 0.04 m potassium phosphate
buffer (pH 8.0) containing 2% 1,2-propanediol and 0.5%
Brij35. To the resulting mixture were added 19 lg of reactivase (0.12 nmol), together with 40 lm AdePeCbl, 20 mm
ATP and 20 mm MgCl2 in 0.03 m potassium phosphate
buffer (pH 8.0) containing 1% 1,2-propanediol and 0.1%
Brij35, in a total volume of 150 lL. After incubation at
37 °C for appropriate periods, the exchange reaction was
terminated by addition of 150 lL of 0.01 m potassium
phosphate buffer (pH 8.0) containing 2% 1,2-propanediol,
0.2% Brij35, and 60 mm EDTA. The resulting mixture was
then subjected to ultrafiltration on a Microcon YM-10
microconcentrator (Millipore, Billerica, MA, USA) to
remove unbound cobalamins. The protein fraction retained
on the filter was washed twice by the addition of 150 lL of

Reactivase for coenzyme B12-dependent enzyme

0.01 m potassium phosphate buffer (pH 8.0) containing 2%
1,2-propanediol, 0.2% Brij35, and 10 mm EDTA, and this
was followed by ultrafiltration. The spectrum was measured
after addition of 150 lL of 0.01 m potassium phosphate
buffer (pH 8.0) containing 2% 1,2-propanediol, 0.2%
Brij35 and 10 mm EDTA to the protein fraction. Experiments without the reactivase or ATP ⁄ MgCl2 were also performed as controls. Similar experiments without cobalamins

were also carried out, for correction of spectra. After spectral measurement, 200 lL of the mixture was incubated at
37 °C for 10 min with 0.06 m citric acid and 5 m guanidine
hydrochloride to denature the proteins. After neutralization, 4 mg of KCN was added, and the resulting mixture
(440 lL) was illuminated for 10 min on ice with a 250 W
tungsten light bulb from a distance of 10 cm to convert cobalamins to a dicyano form. The total amount of enzymebound cobalamin was calculated from the absorbance at
368 nm for dicyanocobalamin (eM, 368, 30.4 · 103 m)1Ỉcm)1)
[35].

Analysis of the enzymreactivase complex by gel
filtration
Complex formation between DD and the reactivase was
analyzed by gel filtration on a Superose 6 (10 ⁄ 300 GL)
column, using an FPLC system (GE Healthcare, Little
Chalfont, UK). In the presence and absence of 21 mm
adenine nucleotide (ATP or ADP) and 21 mm MgCl2, apoenzyme (12 units, 0.48 nmol) were incubated at 37 °C for
60 min with 0.27 mg (1.7 nmol) of reactivase in 170 lL of
0.04 m potassium phosphate buffer (pH 8.0) containing
1.4% 1,2-propanediol and 0.7% Brij35. The resulting mixture was applied to a column that had been equilibrated
with 0.05 m potassium phosphate buffer (pH 8.0) containing 2% 1,2-propanediol and 0.5% Brij35 with or without
the corresponding adenine nucleotide (ADP or ATP) and
MgCl2 (1 mm each). The column was developed with the
same buffer at a flow rate of 0.4 mLỈmin)1. The enzyme
and the reactivase alone were also applied under the same
conditions as controls. The elution of proteins was monitored by the absorbance at 280 nm.

Analysis of the enzymreactivase complexes by
PAGE
The reactivase (12 lg, 76 pmol) was incubated with 10 mm
dithiothreitol at 30 °C for 30 min in 5 lL of 0.01 m potassium phosphate buffer (pH 8.0). ADP and MgCl2 (10 mm
each) were added to the resulting mixture in a total volume

of 6 lL, and the mixture was incubated at 20 °C for
20 min to form the ADP-bound form of the reactivase.
The enzymCN-Cbl complex was formed by the incubation
of apoenzyme (0.38 units, 15 pmol) with 15 lm CN-Cbl
at 30 °C for 30 min in 6 lL of 0.01 m potassium phosphate buffer (pH 8.0) containing 2% 1,2-propanediol and

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1% Brij35. Six microliters of the mixture containing the
reactivasADP complex were added to 6 lL of the mixture
containing the enzymCN-Cbl complex. After 1.5 h at
30 °C, the mixture was subjected to PAGE on a 5% gel
under nondenaturing conditions, as described by Davis
[36], with some modifications: 0.1 m 1,2-propanediol, 1 mm
ADP, 1 mm MgCl2 and 5 mm dithiothreitol were added to
the gels, and 0.1 m 1,2-propanediol, 1 mm ADP and 1 mm
Mg(OCOCH3)2 were added to the electrode buffer. The
enzymCN-Cbl complex, apoenzyme and the reactivasADP complex alone were also subjected to electrophoresis
under the same conditions. After electrophoresis, protein
was stained with Coomassie Brilliant Blue R-250. Bands
were excised from the gel, soaked in water, and equilibrated
with the SDS-containing sample buffer. Then excised gels
were subjected to SDS ⁄ PAGE on 14% and 6% gels under

the conditions described by Laemmli [37]. Protein was
stained again with Coomassie Brilliant Blue R-250. Densitometric analysis of gels was performed with a Printgraph AE-6911CX system (ATTO, Tokyo, Japan) and
nih-image, version 1.6.3 (National Institutes of Health).

Model figure
Figure 7B was generated with chimera [38], using the atomic
coordinates for the enzymCN-Cbl complex (Protein Data
Bank accession code: 1EGM).

Acknowledgements
This work was supported in part by Grants-in-Aid for
Scientific Research [(B) 13480195 and 17370038 and
Priority Areas 753 and 513 to T. Toraya], a Grant-inAid for Young Scientists [(B) 18770111 to K. Mori]
from the Japan Society for Promotion of Science and
the Ministry of Education, Culture, Sports, Science and
Technology, Japan, and a Grant in Aid for Natural
Sciences Research (to T. Toraya) from the Asahi Glass
Foundation, Tokyo, Japan. We thank Y. Kurimoto for
her assistance with manuscript preparation.

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