| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
J. Biol. Chem., Vol. 276, Issue 43, 39919-39925, October 26, 2001
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
From the Laboratory of Plant Molecular Biology, Graduate School of
Agricultural Sciences, Nagoya University, Nagoya 464-8601, Japan
Received for publication, April 20, 2001, and in revised form, June 25, 2001
Fatty acid synthesis in pea chloroplasts is
regulated by light/dark. The regulatory enzyme acetyl-CoA
carboxylase is modulated by light/dark, presumably under redox
regulation. Acetyl-CoA carboxylase is a multienzyme complex composed of
biotin carboxylase and carboxyltransferase (CT). To demonstrate the
redox regulation of CT, composed of the nuclear-encoded The carboxylation of acetyl-CoA by acetyl-CoA carboxylase
(ACCase,1 EC 6.4.1.2) is the
first committed step of fatty acid synthesis. ACCase is the enzyme
considered most responsible for the primary regulation of fatty acid
synthesis. Most plants, with the exception of the grass family
(Poaceae), have two types of ACCase: the heteromeric, prokaryotic form
in plastids and the homodimeric, eukaryotic form in cytosol (1, 2). The
grass family has the homodimeric form in both plastids and cytosol (3,
4). In plants, de novo fatty acid biosynthesis takes place
in the chloroplasts, and the heteromeric ACCase, except in the grass
family, is the regulatory enzyme involved. The heteromeric ACCase is a
multi-enzyme complex composed of four subunits, namely biotin
carboxylase, the biotin carboxyl carrier protein, and the
carboxyltransferase (CT) made up of the In pea leaves, fatty acid synthesis is modulated by light/dark,
presumably by regulation of the plastidic ACCase (5). We previously
proposed that the redox cascade is involved in the light activation of
pea plastidic ACCase (6) and that CT, but not biotin carboxylase, is
redox-regulated (7). The activity of several chloroplast enzymes
involved in the Calvin cycle is regulated by reversible
disulfide-dithiol interchange (8). During photosynthetic electron
transport in the light, covalent redox modification mediated by the
ferredoxin-thioredoxin signal transduction pathway leads to the
reductive light activation of several stromal target enzymes. Thus, the
ferredoxin-thioredoxin pathway links light and the Calvin cycle. We
have proposed that this pathway also links light and fatty acid
synthesis (6). To demonstrate the proposition, it is necessary to
identify the target cysteine residues responsive to redox.
Previously we showed that pea recombinant CT expressed in
Escherichia coli has properties similar to those of
authentic CT and is redox-regulated (9). In this study, using the
recombinant enzyme, we designed experiments to determine the cysteine
residues involved in redox regulation by site-directed mutagenesis
and to determine the role of two subunits. There are 2 and 11 cysteines in the Cloning of cDNAs for the Deleted accA and accD--
RNA
editing of the accD transcript encoding the
PCR was performed for 35 cycles (10 s at 98 °C, 1 min at 55 °C,
and 3 min at 72 °C) using Pyrobest DNA polymerase (Takara) or for 35 cycles (15 s at 94 °C, 30 s at 55 °C, and 3 min at 65 °C)
using KOD Plus (Toyobo). The final PCR products were purified and
inserted into the EcoRV site of the pZErO-2 vector.
The inserted sequences of the resultant plasmids were determined by the
dideoxy chain termination method using an automatic DNA sequencer
(model LIC-400, Li-Cor, Inc.).
Site-directed Mutagenesis of Cysteine by
Alanine--
Site-directed mutagenesis of Construction of Expression Vector--
The expression vector for
the deletion mutant pHis Expression and Purification of Recombinant Proteins--
After
transformation of the E. coli BL21 (DE3) strain (Novagen)
with the expression vectors, cells were grown in an LB medium, and the
proteins were induced and purified according to the method described
(10). From the cell pellets of a 1-liter culture, about 3 mg of
recombinant protein was obtained. The recombinant proteins of the
full-length or the deleted wild-type CT were stable and stored at
Gel Filtration--
The deleted wild-type CT was dialyzed
against the buffer containing 50 mM Tricine-KOH (pH 8), 150 mM NaCl, 1 mM EDTA, and 0.5 mM DTT.
Fifty µl of the dialysate (285 µg) was separated with a Superdex
200 column using a SMART system (Amersham Pharmacia Biotech).
Immunoblotting--
Immunoblot analysis was done as described
previously (10). The antibodies against the Measurement of CT Activity--
CT activity was measured by the
reverse reaction method as described elsewhere (10). The carboxyl
transfer from [2-14C]malonyl-CoA to biotin methyl ester
was measured. The resultant [14C]acetyl-CoA was
evaporated, and the residual [2-14C]malonyl-CoA was
determined. A CT fraction in a total of 50 µl of reaction mixture
(100 mM Tricine-KOH (pH 8), 10 mM biotin methyl ester, 125 µM malonyl-CoA (0.64 kBq), 37.5 µg of bovine
serum albumin) was incubated at 30 °C. After 0 and 20 min of
incubation, 20 µl of the mixture was transferred to a tube containing
5 µl of 6 N HCl to terminate the reaction. The mixture
was heated to dryness at 98 °C for 20 min in a draft chamber,
dissolved in 25 µl of H2O, and applied to Whatman 3MM
paper (1 × 1 cm). The difference between radioactivity at time
zero and after 20 min of incubation was designated as CT activity.
Specific activity is expressed as µmol/min/mg of protein.
Intermolecular Disulfide Bonds--
Proteins eluted from a
nickel column (diluted to 1 mg/ml) were alkylated by incubating with 50 mM iodoacetic acid (Sigma) for 10 min at 37 °C. The
alkylated proteins were treated with an SDS sample loading buffer in
the presence and absence of 720 mM 2-mercaptoethanol. The
proteins (about 0.5 µg) were run on SDS-PAGE (7.5 or 10% gel for the
deletion mutant and 5% gel for the full-length enzyme) and analyzed on
immunoblots probed with either the anti- Extracts from Plants Incubated in Light or Dark--
Pea
seedlings (Pisum sativum cv. Alaska) were grown in a 16 h light/8 h dark cycle at 23-25 °C. Plants (8 days old) were kept in darkness for 16 h (dark-adapted plants) and were then exposed to 2 h of white light at 60 µE/s/m2 (light-adapted
plants). From the dark- or light-adapted plants, intact chloroplasts
were isolated and ruptured as described previously (7). All procedures
for dark-adapted plants were performed in a dark room under a dim green
safety light. The soluble proteins from the lysed extracts were
precipitated by ammonium sulfate and stored at Cysteine Residues in Pea Pea CT and its recombinant enzyme were redox-regulated in
vitro (6, 10), suggesting the presence of redox-sensitive
cysteines in the There are two cysteines at positions 247 and 267 in the pea
Thiol-Disulfide Exchange between Nuclear-encoded and
Chloroplast-encoded Subunits of Pea Acetyl-CoA Carboxylase*
ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
and the
chloroplast-encoded 
subunits, we identified the cysteine residues
involved in such regulation. We expressed the recombinant CT in
Escherichia coli and found that the partly deleted CT was,
like the full-length CT, sensitive to a redox state. Site-directed
mutagenesis of the deleted CT showed that replacement by alanine of the
cysteine residue 267 in the polypeptide or 442 in the polypeptide resulted in redox-insensitive CT and broke the
intermolecular disulfide bond between the and polypeptides. Similar results were confirmed in the full-length CT. These results indicate that the two cysteines in recombinant CT are involved in redox
regulation by intermolecular disulfide-dithiol exchange between the and subunits. Immunoblots of extract from plants incubated in the
light or dark supported that such a disulfide-dithiol exchange is
relevant in vivo. A covalent bond between a nuclear-encoded polypeptide and a chloroplast-encoded polypeptide probably regulates the enzyme activity in response to light.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
and subunits (1, 2). In
these subunits, the subunit is encoded by the chloroplast genome,
and the other three are encoded by the nuclear genome.
and polypeptides, respectively. We identified
one cysteine in the subunit and one cysteine in the subunit
involved in redox regulation and found an intermolecular disulfide bond
between the nuclear-encoded subunit and the chloroplast-encoded
subunit. These results observed in recombinant enzyme were consistent
with the results of in vivo experiments. Probably such a
disulfide bond is redox-regulated in response to light.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
polypeptide
is required for functional CT (9). The accD cDNA, but
not the accD gene, was used for cloning. The cDNAs
encoding the full-length and polypeptides in the pZErO-2
vector (Invitrogen) prepared previously (10) were used as templates.
The deleted accA (
accA) cDNA encoding
N-terminal 372 amino acid residues starting from the accurate
N-terminal sequence lacking a transit peptide (, Fig.
1A) was amplified using forward primer PA01
(5'-TCATATGAAGTTGAGGAAGGTGAAGAGG-3') and reverse primer
PA03
(5'-CCGCTCGAGTTCTCCTTACCTCTTTTTCGTGTTGAC-3'). PA01 and PA03 have NdeI and XhoI restriction
sites (underlined), respectively. PA03 has a ribosome binding site
(italics) and a stop codon (bold). The deleted accD
(accD) cDNA encoding C-terminal 371 amino acid
residues () starting from amino acid position 220 was amplified
using forward primer PD03
(5'-CCGCTCGAGATGATGGAAAAATTAGCTCGTTTA-3') and reverse
primer PD02
(5'-CCGCTCAGCTTAAGTCAAAGGAACGAAACC-3'). PD03 and
PD02 have XhoI and BlpI restriction sites,
respectively, and a stop codon. The deleted accD cDNA
encoding C-terminal 333 amino acid residues from 258 to 590, and
lacking a domain
CX2CX15CX2C in the , was amplified using forward primer PD04
(5'-CCGCTCGAGATGAGCAGTTCAGATAGAATCGAC-3') and reverse
primer PD02.
accA and
accD cDNAs was introduced by PCR. The accA
and accD cDNAs in the pZErO-2 vector (10) were
used as templates for the first and second PCRs, except for mutagenesis
at Cys-230 or Cys-233. In the first PCR, the forward primer,
PA01 for accA or PD01
(5'-CCGCTCGAGATGATAAATGAAGACCCATCTAG-3') for
accD and reverse primers containing the desired mutation
("Reverse primer" in Table I) were used. In the second PCR, the
forward primers ("Forward primer" in Table I), the compliment
sequence of the reverse primer in Table I, and the reverse primer, PA02 (5'-CCGCTCGAGTTCTCCTTAAGAGAAGTTGCGATTTACACCAAC-3')
for accA and PD02 for accD were used. The
first and second PCR products were purified by electrophoresis on 1%
agarose gel and subsequent extraction using the Sephaglas Band Prep kit
(Amersham Pharmacia Biotech). The mixture containing the first and
second PCR products was used as a template for the final PCR. The final
PCR was performed using PA01 and PA03 for accA cDNA
mutation and using PD03 and PD02 for accD cDNA
mutation. To prepare a mutation at Cys-230 or Cys-233 in , the
first PCR was done by forward primer containing parts of PD03 sequences
(Table I) and by reverse primer PD02 using the accD cDNA in the pZErO-2 vector as a
template. The second PCR was done by forward primer PD03 and reverse
primer PD02 using the first PCR products as a template. For preparation
of the mutants of full-length cDNA, the final PCR was performed
using PA01 and PA02 for accA and using PD01 and PD02 for
accD. PCR, cloning, and sequence confirmation were performed
as described for the deleted cDNAs.
Primers for site-directed mutagenesis
AD (Fig. 2A) was constructed
as described for the full-length CT (10). For coexpression of the
and polypeptides to produce the CT complex in E. coli, accA and accD cDNAs were
bicistronically inserted into the pET-19b vector containing a
decahistidine tag (His tag), which allows a fused protein to be
purified by a nickel column. First, the accD cDNA was
ligated to the XhoI/BlpI site of pET-19b, and
then the accA cDNA was ligated to the
XhoI/NdeI sites of the pET-19b vector carrying
the accD fragment. In the resultant pHisAD, the His
tag was fused to the N terminus of the polypeptide. The His tag
was not removed from the recombinant throughout the experiments.
80° C for characterization. The mutant proteins eluted from a
nickel column with a solution containing 150 mM imidazole,
500 mM NaCl, and 20 mM Tris-HCl (pH 7.9) were
unstable and were immediately used for the enzyme assay and
SDS-PAGE.
polypeptide were also
prepared using the method described (10). The antibodies against the
polypeptide were prepared using an oligopeptide containing 15 amino
acid residues from 399 to 413 of polypeptide as described elsewhere
(11).
polypeptide or the anti-
polypeptide IgG.
80 °C until use.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
and Polypeptides
and polypeptides consisting of 825 and 590 amino acid residues, respectively. Fig.
1A shows schematically the
position of cysteines deduced from pea accA and
accD cDNAs and the conserved domain in E. coli. N-terminal 310 amino acids of the pea polypeptide were
conserved in E. coli with 81% similarity (47% identity), and C-terminal 278 amino acids of the pea polypeptide, excluding the reiterated sequence, were conserved in E. coli with 78%
similarity (39% identity). The other domains absent from E. coli counterparts are not always conserved among plant
proteins.
View larger version (53K):
[in a new window]
Fig. 1.
Cysteine residues in the and polypeptides. A,
schematic representation of the full-length ( and
) and deleted (, , and
*) polypeptides. The numbers are the amino
acid positions of each full-length polypeptide. The positions of
cysteines are marked by asterisks. B,
sequence alignments of the regions designated a,
b, and c in A. The numbers
are the amino acid positions of the full-length pea polypeptides.
Black arrowhead, cysteine residues in pea; white
arrowhead, possible initiation methionine in and *;
asterisk, cysteines in
CX2CX12-15CX2C.
Amino acid residues identical in one-half or more of the pea sequences
are shaded black, and conservative substitutions
are shaded gray.
polypeptide. Alignment shows that the sequences around these cysteines
are highly conserved (Fig.
2B); Cys-247 is found in higher plants, and Cys-267 is found in all sequences available. In pea
polypeptides, there are 11 cysteines. Five of them in the
N-terminal domain are not conserved among plant proteins (data not
shown), but six cysteines in the C-terminal domain are considerably conserved. Four of them, at positions 230, 233, 249, and 252, are found
in all the counterparts available and form a motif, CX2CX12-15CX2C.
Cysteine at position 442 is found only in pea, and cysteine at 466 is
found in all photosynthetic organisms available. Taking into account
the redox-insensitive properties of E. coli CT, such a
sequence comparison does not directly predict the redox-sensitive
cysteines in both the and polypeptides.
View larger version (25K):
[in a new window]
Fig. 2.
Expression of the deleted CTs and their redox
sensitivity. A, expression vector. A ribosomal site
(rbc) was inserted. B, partial purification.
Proteins from a nickel column eluate were separated on SDS-PAGE. 5 µg
of protein was used for staining. For immunoblotting, 0.5 µg of
proteins was used. C, gel filtration. Proteins (285 µg)
from a nickel column eluate were analyzed with a Superdex 200 column.
The fractions of CT activity were electrophoresed, stained with
Coomassie Brilliant Blue, or probed with anti-CT (Anti-) or
anti-CT (Anti-) IgGs. D, effect of DTT (
) and
2-mercaptoethanol (-SH, 
) on the deleted CT. The
activity of the eluate from a nickel column was preincubated with the
indicated concentration of thiol for 10 min at 30 °C and measured.
The activity was relative to that at 8 mM DTT. Results are
the means of two independent measurements. Different preparations
showed almost the same profiles. For comparison, the activity of
full-length CT (dotted lines, right scale) is
shown (10). The specific activities of the deleted CT and the
full-length CT at 8 mM DTT were 0.366 and 0.151 unit,
respectively.
Requirements of the Deleted CT for DTT
Expression--
There are 13 possible cysteines involved in redox
regulation (Fig. 1A), and we attempted to reduce them. To
investigate whether the domains not conserved in E. coli,
namely, the C-terminal half in the subunit and the N-terminal half
in the subunit, are necessary for redox regulation, we prepared
enzymes in which these sequences were deleted, using an expression
plasmid, pHisAD (Fig. 2A), and examined the redox sensitivity.
The eluate partially purified by a nickel column showed redox-sensitive
CT activity. The specific activity was 0.366 unit. The eluate exhibited
three major bands of 52, 45, and 44 kDa by Coomassie Brilliant Blue
staining (Fig. 2B). A band of 44 kDa reacted with antibodies
against the polypeptide, agreed with the calculated molecular mass,
44.3 kDa, and was identified as His tag . Both 52- and 45-kDa
bands reacted with antibodies against the polypeptide.
Determination of the N-terminal sequence of the 52- and 45-kDa bands
revealed that the 52-kDa band corresponded to the protein
starting from the predicted first methionine at position 220 (Fig.
1B-b). The N-terminal sequence of the 45-kDa polypeptide started from serine at position 259, suggesting that the
third methionine at position 258 was recognized as another starting
point. The calculated molecular masses of the and a polypeptide
starting from serine 259 (*) were 41.7 and 37 kDa, respectively,
and the observed size was about 8-10 kDa larger than the calculated
value. Such a shift in molecular mass on SDS-PAGE was observed for the
full-length subunit and was attributable to the intrinsic amino
acid sequence (11). Usually the 52-kDa band was denser than that of the
45 kDa, and was the major product.
To estimate the molecular size of the deleted CT, the partially
purified eluate was separated by gel filtration in the presence of DTT.
The peak CT activity appeared in fraction 16. Coomassie Brilliant Blue
staining and immunoblotting of separated fractions revealed a peak of
His tag , , and * polypeptides at fraction 16. A peak
composed of only the His tag polypeptide was not found, unlike
the expression of full-length polypeptides (10), suggesting that most
of the expressed His tag formed a complex with or
*. The molecular size of the deletion mutant at fraction 16 estimated from the standard proteins was about 250 kDa and was
approximately one-half the size of the full-length CT, 490 kDa. The
calculated molecular mass of (His tag
)22 was also approximately one-half
that of the full-length CT, (His tag )22, and the molecular composition of the deleted CT was probably (His tag
)22 or (His tag
)2*2.
Active Form and Domain Required for Catalytic Activity--
Two
forms of the deleted CT were obtained by expression of pHisAD. To
determine the form(s) having CT activity, we constructed another
plasmid to express and * polypeptides starting from the
methionine at position 258. The eluate obtained by expression of this
plasmid showed no CT activity. This result indicates that the
measurable activity by expression of pHisAD is the complex of His
tag with but not that of * polypeptides. The
deleted sequences from 220 to 257, which contain a
CX2CX12-15CX2C motif conserved among all the polypeptides, are required for catalytic activity but not for association with the polypeptide. This is not a case in which a zinc finger motif is usually required for
complex formation. It is likely that the
CX2CX12-15CX2C motif is important for catalytic activity.
Redox Sensitivity--
We examined the sensitivity of the deleted
CT partially purified to the redox state and compared it with that of
the full-length CT. In this experiment, the effect of a known reductant
of S-S bonds, DTT, was tested. DTT activated the enzyme in a profile similar to that of the full-length enzyme (Fig. 2D). The
monothiol 2-mercaptoethanol did not activate the enzyme efficiently.
These results indicate that the deletion mutant was sensitive to the redox state, like the full-length CT (9), and that the redox-sensitive cysteines were located in the and polypeptides. Thus, the deleted sequences, the C-terminal half of the polypeptide, and the
N-terminal half of the polypeptide are not necessary for redox
regulation and may have some other functions. The number of possible
cysteines was reduced from 13 to 8.
Requirements of Site-directed Mutants for DTT
To identify the redox-sensitive cysteines in the deleted CT, a set of mutants, each with a single substitution of cysteine to alanine, was constructed, and the requirement of each mutant CT for DTT was tested. There were no significant differences in the efficiency of expression and purification of all mutants compared with the deleted wild-type enzyme. However, solubility differed with different mutant enzymes. Most mutant enzymes were precipitated under low ionic strength or by freezing. Therefore, each enzyme activity was determined using freshly eluted fraction from nickel column.
The relative specific activity of mutants in either the presence or
absence of 8 mM DTT was determined (Table
II). The deleted CT is redox-sensitive,
and if a cysteine involved in redox regulation is replaced, the
resultant mutant enzyme is expected to be converted to a
redox-insensitive form. We surveyed such an enzyme active in the
absence of DTT. In the presence of DTT, the mutant proteins retained
varying levels of activity. Substitutions of Cys-267 in and
Cys-466 in had modest effects, retaining more than 50% of the
wild-type activity. Other mutant enzymes were more impaired, retaining
only 0-30% of the wild-type activity, suggesting that alanine
replacements of cysteine residues does not satisfy the structural
requirements at these sites for enzyme folding and the substrate
binding. In particular, Cys-233 or -252 in a CX2CX12-15CX2C
motif is probably required for catalytic activity. In the absence of
DTT, only two mutants, the substitution of Cys-267 in the and
the substitution of Cys-442 in the polypeptides, were active and
their activities were almost equal to those occurring in the presence
of DTT. These results indicate that Cys-267 in and Cys-442 in
are required for redox regulation.
|
To confirm the role of the two cysteines at the same position in the
full-length CT, we generated mutant enzymes with a substitution of
cysteine by alanine and measured their activity (Table II). Although
the CT activity was low, measurable activity was found in the absence
of DTT, indicating that Cys-267 in the subunit or Cys-442 in the
subunit is required for redox sensitivity and that the redox
sensitivity of the full-length enzyme is probably mediated exclusively
through these two cysteines. The molecular size of the deleted CT is
about half that of the full-length CT, and the specific activity of the
deleted CT corrected for the difference of molecular size was
comparable with that of the full-length. The extra sequences, the
C-terminal region of the polypeptide and the N-terminal region of
the polypeptide (Fig. 1A), do not play an important role
for exhibition of the enzyme activity. The low activity of these
mutants suggests that the replacement of the cysteine by alanine may
bring about inappropriate conformation for catalytic activity, probably
by interacting with the extra sequences of the full-length CT. The
interaction, which does not occur in the wild-type full-length enzyme,
is somehow harmful for the expression of the full activity by
preventing efficient substrate binding or subunit-subunit interaction.
Intermolecular Disulfide Bond
Each subunit had only one cysteine responsive to redox, suggesting
that an intermolecular but not an intramolecular disulfide bond is
formed under nonreduced conditions. The molecular composition of CT is
probably 22, and there are possible
intermolecular disulfide bonds between homodimers or heterodimers. To
determine which type of S-S bond was formed, SDS-PAGE analysis of
several recombinant enzymes was performed in the presence or absence of the reducing agent 2-mercaptoethanol. To prevent rearrangement of
disulfide bonds during treatment with a sample loading buffer, SH
groups were alkylated with iodoacetic acid. The modified proteins were
denatured and separated by SDS-PAGE in the presence or absence of
2-mercaptoethanol and probed with antibodies against the or polypeptides (Fig. 3).
|
When the deleted wild-type enzyme was analyzed, the band of His tag
(44 kDa) observed in the presence of 2-mercaptoethanol was
partly shifted to about 125 kDa in its absence (Fig. 3a). The bands of (52 kDa) and * (45 kDa) were also shifted to about 125 kDa. These results suggest that both the His tag and
polypeptides formed intermolecular S-S bonds, probably between
His tag and or *. The apparent molecular mass, about 125 kDa, was larger than the sum of the two bands, probably because of the slow migration of this alkylated and S-S-bonded polypeptide.
To further confirm the S-S bond between the two molecules, a CT enzyme
consisting of a full-length and was expressed, and the
intermolecular S-S bond was analyzed (Fig. 3b). In this enzyme, the polypeptide is twice as large as , and we
expected shifted bands larger than 125 kDa, which would react with both anti- and anti- polypeptide antibodies. The number of cysteine residues was the same as that of the deleted wild-type CT, and its
redox sensitivity was similar to that of the wild type (data not
shown). In the absence of 2-mercaptoethanol, both anti- and anti-
polypeptide IgG reacted with the two shifted bands at ~200 kDa,
supporting the idea that the polypeptide formed an S-S bond with
or * under nonreducing conditions, although the apparent
molecular size of the shifted bands was again larger than the sum of
the two polypeptides. These results support the idea that the shifted
bands are heterodimers of His tags and or * and
suggest that the S-S bond is formed between Cys-267 in and Cys-442
in or *.
To confirm the presence of the S-S bond in full-length wild-type CT,
we analyzed the CT under nonreducing conditions (Fig. 3c).
The shifted bands of about 230 kDa were observed in both and polypeptides, supporting the idea that the S-S bond was formed in
full-length CT as expected. The shifted minor bands probably resulted
from the polypeptide, starting from the second and third methionine
of the polypeptide. The remaining polypeptide was the
polypeptide expressed in excess not associated with the polypeptide
(10).
If the disulfide bond is formed between Cys-267 in the and
Cys-442 in the or *, mutation of these cysteines breaks the disulfide bond and results in the loss of the shifted band under
nonreducing conditions. We analyzed the mutants and found that the
molecular size of the polypeptides is the same in the presence and
absence of 2-mercaptoethanol. We lost the shifted heterodimer bands of
His tags and or * under nonreducing conditions
(Fig. 3, d and e). The results indicate that each
cysteine is necessary for an intermolecular S-S bond and that a
critical disulfide bond is formed between Cys-267 in the subunit
and Cys-442 in the subunit.
Occurrence of Disulfide Form of the Enzyme in Vivo
We have shown previously that ACCase from light-adapted plants is
in a much more reduced form than that from dark-adapted plants and that
light-dependent reduction of ACCase occurs in vivo (7). To ensure that the results observed in the recombinant enzyme (Fig. 3) were applied in vivo, we examined disulfide
bridge formation in the enzyme by immunoblots of extracts from plants in light or dark (Fig. 4). The shifted
bands of about 230 kDa were observed in both and polypeptides
in the absence of 2-mercaptoethanol, suggesting that both the and
polypeptides formed a S-S bond in vivo. The disulfide
form of about 230 kDa was more abundant in the dark-adapted plants than
in the light-adapted plants; the reverse was found for thiol forms of
about 98 kDa for polypeptide and 90 kDa for polypeptide,
suggesting that a light-dependent reduction of the
disulfide bond indeed occurred in vivo. Probably the
thiol-disulfide exchange observed in vitro is relevant
in vivo, although we cannot completely exclude the
possibility that a different set of cysteine residues is involved in
the S-S bond in vivo.
|
| |
DISCUSSION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
|
|---|
The experiments reported here show that the two cysteines, Cys-267
in the polypeptide and Cys-442 in the polypeptide, are
required for redox regulation of the recombinant CT. The observation appears to be relevant in vivo, and the two cysteines are
likely involved in the reductive activation of CT, and therefore in
that of ACCase, by disulfide-dithiol interchange. In the light,
electrons from photosystem I are shuttled through the electron
transport chain to ferredoxin and are transferred to thioredoxins by
ferredoxin-thioredoxin reductase and then to ACCase, reducing the
disulfide bonds in the CT and resulting in the light activation of
fatty acid synthesis.
All of the polypeptides encoded by the chloroplast genome form a
functional complex with polypeptides encoded by the nuclear genome. For
example, ribulose-bisphosphate carboxylase, RNA polymerase, Clp
protease, and ATP synthase are composed of chloroplast-encoded and
nuclear-encoded polypeptides (12). In these enzymes, the catalytic or
structural polypeptides are encoded by the chloroplast genome, and the
regulatory polypeptides are encoded by the nuclear genome. The same may
be true for pea CT, although further experiments are needed to
characterize the precise function of the two subunits. The
chloroplast-encoded polypeptide has catalytic and regulatory functions, and the nuclear-encoded polypeptide has a regulatory function. To date, we do not know the molecular interaction between catalytic and regulatory polypeptides. A new finding obtained in our
experiments indicates that molecular interaction of the two
polypeptides is characterized as follows: the nuclear-encoded polypeptide forms a covalent bond with the chloroplast-encoded polypeptide and thereby changes enzyme activity.
There are several chloroplast enzymes activated by thioredoxin (13),
including fructose-1,6-bisphosphatase,
sedoheptulose-1,7-bisphosphatase, phosphoribulokinase, ATP synthase,
NADP-dependent malate dehydrogenase, glucose-6-phosphate
dehydrogenase, and ribulose-bisphosphate-carboxylase activase. The
cysteine residues involved in redox regulation are identified by
site-directed mutagenesis, and some consensus motifs among plants have
been found for each enzyme. However, a consensus motif among enzymes
has not yet been found. A comparison of the primary sequences of these
enzymes with redox-insensitive cytosolic isoforms or with the
redox-insensitive counterpart from different organisms reveals that the
redox-sensitive cysteines are located on extra loops or extensions
(14). Light-regulated NADP-dependent malate dehydrogenase
possesses sequence extensions at both the N- and C-terminal ends
compared with the constitutive active counterpart and two cysteines at
the N terminus and two cysteines at the C terminus are involved in
redox regulation (15, 16). Chloroplast fructose bisphosphatase
possesses an insertion containing three cysteines compared with the
constitutive active cytosolic enzyme and two of them are involved in
redox regulation (17, 18). In contrast to these enzymes, different
features are found for pea ACCase. First, the molecular structure of
pea chloroplast ACCase differs considerably from
redox-insensitive cytosolic ACCase (1, 2), and we cannot propose
a possible origin of redox-sensitive cysteines, although there are some
similarities in their CT domains. Second, the regulatory cysteine in
the polypeptide is not located in the plant-specific extra domains
but in domains conserved among organisms, reflecting the differences
from other chloroplast enzymes in terms of the evolutionary origin of
redox sensitivity. Third, intermolecular disulfide bonds among
different polypeptides are involved in redox regulation, whereas
intramolecular disulfide bonds are involved in the other chloroplast enzymes.
Available sequence data show that the Cys-267 in the polypeptide is
conserved among organisms but the Cys-442 in the polypeptide is
found only in pea. Other organisms have a serine at this position. Replacement of Cys-442 by serine resulted in CT with very low activity
(data not shown) suggesting that Cys-442 is important for pea CT. At
present, we cannot propose a possible consensus sequence of CT for
redox regulation. There is a possibility that this type of redox
regulation is unique to the pea, although spinach (19) and tobacco
ACCases2 are redox-regulated,
and another set of cysteines may be responsible in other species. In
the case of sedoheptulose bisphosphatase, the redox-sensitive cysteines
of Chlamydomonas reinhardtii were predicted to be different
from those of plants (20).
Replacement of Cys-247 or -267 in the polypeptide by serine
resulted in mutants of low activity, less than 3% of the deleted wild-type (data not shown), whereas the corresponding replacements by
alanine residues allowed the more active CT, about 30-80% of the
wild-type (Table II). Serine residues at these positions are not
compatible with the formation of a functional complex of CT activity,
but alanine replacement of Cys-247 or Cys-267 satisfies the structural
requirement at these sites for CT folding and stability. Similar
results were obtained for replacement of the cysteine by serine in the
polypeptide. Serine is a hydrophilic amino acid, and alanine, like
cysteine, is hydrophobic. It is likely that the hydrophobicity of
cysteine plays an important role in the functional folding of CT. Such
results were reported for rhodopsin (21) and small soluble proteins
such as trypsin inhibitor (22) and lysozyme (23).
A conserved motif,
CX2CX12-15CX2C,
forms a putative zinc finger motif. The deletion of this domain
suggests that this motif is required for catalytic activity, probably
binding metal ions. A preliminary experiment to identify the metal ion
by flame reaction showed the presence of Zn but not of
Fe.3 Further experiments are
needed in order to understand the precise roles of the and subunits.
| |
ACKNOWLEDGEMENTS |
|---|
We thank H. Mori for analysis of protein sequences, H. Iguchi for analysis of the intermolecular disulfide bond, and M. Hirose and Y. Nagano for discussions.
| |
FOOTNOTES |
|---|
* This work was supported by Grants-in-aid 10460147 for Scientific Research and 11151216 on Priority Areas from the Japanese Ministry of Education, Science, Sports, and Culture and by Grants JSPS-RIFT 96L006012 and JSPS-RFTF 97R16001 from the Japan Society for the Promotion of Science, Research for the Future Program.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed: Laboratory of Plant
Molecular Biology, Graduate School of Agricultural Sciences, Nagoya
University, Nagoya 464-8601, Japan. Tel.: 81-52-789-4165; Fax:
81-52-789-296; E-mail: sasaki@agr.nagoya-u.ac.jp.
Published, JBC Papers in Press, August 23, 2001, DOI 10.1074/jbc.M103525200
2 Y. Madoka, and Y. Sasaki, unpublished observation.
3 A. Kozaki, K. Mayumi, and Y. Sasaki, unpublished observation.
| |
ABBREVIATIONS |
|---|
The abbreviations used are: ACCase, acetyl-CoA carboxylase; CT, carboxyltransferase; DTT, dithiothreitol; His tag, decahistidine tag; PAGE, polyacrylamide gel electrophoresis; PCR, polymerase chain reaction.
| |
REFERENCES |
|---|
|
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
|
|---|
| 1. | Sasaki, Y., Konishi, T., and Nagano, Y. (1995) Plant Physiol. 108, 445-449[CrossRef][Medline] [Order article via Infotrieve] |
| 2. | Ohlrogge, J., and Browse, J. (1995) Plant Cell 7, 957-970[CrossRef][Medline] [Order article via Infotrieve] |
| 3. |
Konishi, T.,
and Sasaki, Y.
(1994)
Proc. Natl. Acad. Sci. U. S. A.
91,
3598-3601 |
| 4. |
Konishi, T.,
Shinohara, K.,
Yamada, K.,
and Sasaki, Y.
(1996)
Plant Cell Physiol.
37,
117-122 |
| 5. | Sauer, A., and Heise, K.-P. (1984) Z. Naturforsch. Sect. C Biocsci 39, 268-275 |
| 6. |
Sasaki, Y.,
Kozaki, A.,
and Hatano, M.
(1997)
Proc. Natl. Acad. Sci. U. S. A
94,
11096-11101 |
| 7. | Kozaki, A., and Sasaki, Y. (1999) Biochem. J. 339, 541-546 |
| 8. | Buchanan, B. B. (1980) Annu. Rev. Plant Physiol. 31, 341-374 |
| 9. |
Sasaki, Y.,
Kozaki, A.,
Ohmori, A.,
Iguchi, H.,
and Nagano, Y.
(2001)
J. Biol. Chem.
276,
3937-3940 |
| 10. |
Kozaki, A.,
Kamada, K.,
Nagano, Y.,
Iguchi, H.,
and Sasaki, Y.
(2000)
J. Biol. Chem.
275,
10702-10708 |
| 11. |
Sasaki, Y.,
Hakamada, K.,
Suama, Y.,
Nagano, Y.,
Furusawa, I.,
and Matsuno, R.
(1993)
J. Biol. Chem.
268,
25118-25123 |
| 12. |
Martin, W.,
and Herrmann, R. G.
(1998)
Plant Physiol.
118,
9-17 |
| 13. | Schürmann, P., and Jacquot, J.-P. (2000) Annu Rev. Plant Physiol. Plant Mol. Biol. 51, 371-400[CrossRef] |
| 14. | Rulland, E., and Miginiac-Maslow, M. (1999) Trends Plant Sci. 4, 136-141[CrossRef][Medline] [Order article via Infotrieve] |
| 15. |
Issakidis, R.,
Miginiac-Maslow, M.,
Decottignies, P.,
Jacquot, J.-P.,
Cretin, C.,
and Gadal, P.
(1992)
J. Biol. Chem.
267,
21577-21588 |
| 16. |
Issakidis, E.,
Saarinen, M.,
Decottignies, P.,
Jacquot, J.-P.,
Cretin, C.,
Gadal, P.,
and Migniac-Maslow, M.
(1994)
J. Biol. Chem.
269,
3511-3517 |
| 17. | Jacquot, J.-P., Lopez-Jaramillo, J., Chueca, A., Cherfils, J., Lemaire, S., Chedozeau, B., Miginiac-Maslow, M., Decottignies, P., Wolosiuk, R., and Lopez-George, J. (1995) Eur. J. Biochem. 229, 675-681[Medline] [Order article via Infotrieve] |
| 18. | Jacquot, J.-P., Lopez-Jaramillo, J., Migniac-Maslow, M., Lemaire, S., Cherfils, J., Chueca, A., and Lopez-Jaramillo, J. (1997) FEBS Lett. 401, 143-147[CrossRef][Medline] [Order article via Infotrieve] |
| 19. | Hunter, S. C., and Ohlrogge, J. B. (1998) Arch. Biochem. Biophys. 359, 170-178[CrossRef][Medline] [Order article via Infotrieve] |
| 20. | Anderson, L. E., Huppe, H. C., Li, A. D., and Stevens, F. J. (1996) Plant J. 10, 553-560[CrossRef][Medline] [Order article via Infotrieve] |
| 21. |
Karnik, S. S.,
Sakmar, T. P.,
Chen, H.-B.,
and Khorana, H. G.
(1988)
Proc. Natl. Acad. Sci. U. S. A.
85,
8459-8463 |
| 22. |
Staley, J. S.,
and Kim, P.
(1992)
Proc. Natl. Acad. Sci. U. S. A.
89,
1519-1523 |
| 23. |
Inaka, K.,
Taniyama, Y.,
Kikuchi, M.,
Morikawa, K.,
and Matsushima, M.
(1991)
J. Biol. Chem.
266,
12599-12603 |
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati What's this?
This article has been cited by other articles:
|
|
 |
|
  L. Michelet, M. Zaffagnini, H. Vanacker, P. Le Marechal, C. Marchand, M. Schroda, S. D. Lemaire, and P. Decottignies In Vivo Targets of S-Thiolation in Chlamydomonas reinhardtii J. Biol. Chem., August 1, 2008; 283(31): 21571 - 21578. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. Belkebir, R. D. Paepe, A. Tremolieres, F. Aid, and G. Benhassaine-Kesri Sethoxydim affects lipid synthesis and acetyl-CoA carboxylase activity in soybean J. Exp. Bot., November 1, 2006; 57(14): 3553 - 3562. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Hisabori, S. Hara, T. Fujii, D. Yamazaki, N. Hosoya-Matsuda, and K. Motohashi Thioredoxin affinity chromatography: a useful method for further understanding the thioredoxin network J. Exp. Bot., June 1, 2005; 56(416): 1463 - 1468. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
P. Geigenberger, A. Kolbe, and A. Tiessen Redox regulation of carbon storage and partitioning in response to light and sugars J. Exp. Bot., June 1, 2005; 56(416): 1469 - 1479. [Abstract] [Full Text] [PDF]
|
|
| ||||||||
