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Identification of the 7-Hydroxymethyl Chlorophyll a Reductase
of the Chlorophyll Cycle in Arabidopsis
W
Miki Meguro, Hisashi Ito,1 Atsushi Takabayashi, Ryouichi Tanaka, and Ayumi Tanaka
Institute of Low Temperature Science, Hokkaido University, Sapporo 060-0819, Japan
The interconversion of chlorophyll a and chlorophyll b, referred to as the chlorophyll cycle, plays a crucial role in the
processes of greening, acclimation to light intensity, and senescence. The chlorophyll cycle consists of three reactions: the
conversions of chlorophyll a to chlorophyll b by chlorophyllide a oxygenase, chlorophyll b to 7-hydroxymethyl chlorophyll a
by chlorophyll b reductase, and 7-hydroxymethyl chlorophyll a to chlorophyll a by 7-hydroxymethyl chlorophyll a reductase.
We identified 7-hydroxymethyl chlorophyll a reductase, which is the last remaining unidentified enzyme of the chlorophyll
cycle, from Arabidopsis thaliana by genetic and biochemical methods. Recombinant 7-hydroxymethyl chlorophyll a
reductase converted 7-hydroxymethyl chlorophyll a to chlorophyll a using ferredoxin. Both sequence and biochemical
analyses showed that 7-hydroxymethyl chlorophyll a reductase contains flavin adenine dinucleotide and an iron-sulfur
center. In addition, a phylogenetic analysis elucidated the evolution of 7-hydroxymethyl chlorophyll a reductase from divinyl
chlorophyllide vinyl reductase. A mutant lacking 7-hydroxymethyl chlorophyll a reductase was found to accumulate
7-hydroxymethyl chlorophyll a and pheophorbide a. Furthermore, this accumulation of pheophorbide a in the mutant was
rescued by the inactivation of the chlorophyll b reductase gene. The downregulation of pheophorbide a oxygenase activity
is discussed in relation to 7-hydroxymethyl chlorophyll a accumulation.
INTRODUCTION
In land plants and cyanobacteria, chlorophyll is synthesized from
Glu (Tanaka and Tanaka, 2006; Tanaka and Tanaka, 2007) and
exists as chlorophyll-protein complexes (Barber et al., 2000).
During greening, newly synthesized chlorophyll assembles with
various proteins to form the chlorophyll-protein complexes of the
photosystems (Shimada et al., 1990), which harvest and transfer
light energy and drive the electron transport that is indispensable
to photosynthesis (Green and Durnford, 1996; Fromme et al.,
2003). Because chlorophyll is a potentially dangerous molecule
that generates reactive oxygen species (op den Camp et al.,
2003), the chlorophyll that is released from the complex during
senescence is converted to safe molecules of nonfluorescent
chlorophyll catabolites (Hörtensteiner, 2006). When chlorophyll
metabolism is not well regulated, and intermediate molecules
accumulate (Papenbrock et al., 2000; Meskauskiene et al.,
2001), necrotic lesions appear on leaves (Pruzinská et al.,
2003; Hirashima et al., 2009). By contrast, when the chlorophyll
supply is limited, photosynthesis activity becomes low, resulting
in the retardation of plant growth (Liu et al., 2004). Therefore, it is
important that the synthesis and degradation of chlorophyll be
strictly regulated during both greening and senescence.
In order for the plant to achieve a desirable chlorophyll a/b
ratio, a portion of newly synthesized chlorophyll a or chlorophyl1 Address
correspondence to [email protected].
The author responsible for distribution of materials integral to the
findings presented in this article in accordance with the policy described
in the Instructions for Authors (www.plantcell.org) is: Hisashi Ito
([email protected]).
W
Online version contains Web-only data.
www.plantcell.org/cgi/doi/10.1105/tpc.111.089714
lide a is converted to chlorophyll b or chlorophyllide b by
chlorophyllide a oxygenase (CAO) (Tanaka et al., 1998; Espineda
et al., 1999). When the plant needs to decrease the level of
chlorophyll b, chlorophyll b is converted to chlorophyll a by
chlorophyll b reductase (CBR) (Scheumann et al., 1996) and
7-hydroxymethyl chlorophyll a (HMChl) reductase (HCAR) (Ito
et al., 1996; Scheumann et al., 1998). This interconversion pathway between chlorophyll a and chlorophyll b is referred to as
the chlorophyll cycle (Rüdiger, 2002). However, the chlorophyll
cycle entails more than just the interconversion of chlorophyll a
and chlorophyll b; it also plays an essential role in the formation
and degradation of light-harvesting chlorophyll a/b–protein
complexes (LHC). The level of LHC has been shown to be greatly
reduced in chlorophyll b–less mutants (Murray and Kohorn,
1991), which have a defect in CAO (Oster et al., 2000); by
contrast, the formation of LHC of photosystem II (LHCII) has been
reported to be accelerated in CAO-overexpressing plants
(Tanaka et al., 1999, 2001; Tanaka and Tanaka, 2005). Two
CBR genes, NOL and NYC1, have been identified using the staygreen mutant of rice (Oryza sativa) (Kusaba et al., 2007). The
degradation of the LHC was found to be almost completely
inhibited, but other chlorophyll a–protein complexes were degraded in cbr mutants (Kusaba et al., 2007). It has been reported
that when trimeric LHCII was incubated with recombinant CBR
(NOL), chlorophyll b in LHCII was converted to HMChl, and all of
the chlorophyll, including chlorophyll a, was released from LHCII,
indicating that CBR participates in the first step of LHC degradation (Horie et al., 2009). The chlorophyll cycle plays a crucial
role in the processes of greening, acclimation to light intensity,
seed formation, and senescence, where LHCII is actively synthesized or degraded. Therefore, the elucidation of the properties and the regulation of the enzymes for the chlorophyll cycle
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are indispensable for an understanding of various physiological
processes.
In the chlorophyll cycle, CAO and CBR have been identified by
mutant screening of Chlamydomonas reinhardtii (Tanaka et al.,
1998) and rice (Kusaba et al., 2007), respectively, and their
physiological functions, regulation, and distribution in photosynthetic organisms have been elucidated. However, HCAR has not
yet been identified. HCAR catalyzes the reduction of a hydroxymethyl group to a methyl group, a process in which the substitution of a hydroxyl (OH) with a hydrogen (H) occurs. The
reduction of an OH group is a chemically difficult reaction; therefore, chlorophyll b–to–chlorophyll a conversion had not been
considered to occur prior to the discovery of chlorophyll b to
chlorophyll a conversion within isolated plastids (Ito et al., 1993).
Although enzymatic examples of this reaction are rare, a similar
reaction has been reported and well studied with ribonucleotide
reductase, which catalyzes the substitution of the 29-OH group of
a ribonucleotide with a hydrogen (Bollinger et al., 2008). This
reaction occurs via a free radical mechanism. However, the
mechanism of the substitution of the OH with a hydrogen by
HCAR might be different from that by ribonucleotide reductase
because the OH exists on the pyrrole ring of HMChl. Thus,
enzymatic information is indispensable for an understanding of
this difficult reaction.
In this study, we obtained an Arabidopsis thaliana mutant that
accumulated HMChl, a substrate of HCAR. This mutant was
impaired in a putative iron-sulfur flavoprotein, which has a high
sequence similarity to divinyl chlorophyll vinyl reductase (DVR) of
Synechocystis PCC6803. The recombinant protein expressed in
Escherichia coli converted HMChl to chlorophyll a, a reaction that
used ferredoxin as a reducing equivalent. We emphasize that this
difficult reaction involving the substitution of an OH with an H was
achieved by a single enzyme, the flavoprotein now referred to as
HCAR. We also found accumulation of pheophorbide a, a degradation product of chlorophyll, in the hcar mutant, and further
discuss the evolution of HCAR.
RESULTS
Arabidopsis Homolog of Cyanobacterial DVR
Figure 1 shows the chlorophyll metabolic pathway, including the
later steps of chlorophyll a synthesis, the chlorophyll cycle, and
the initial steps of chlorophyll degradation. Divinyl chlorophyllide
a is converted to monovinyl chlorophyllide a and is then phytylated to chlorophyll a, which is converted to chlorophyll b by CAO.
In the degradation processes, chlorophyll b is converted to
chlorophyll a before its degradation and is subsequently dechelated and dephytylated to form pheophorbide a.
DVR is responsible for the reduction of a vinyl group to an ethyl
group in the chlorophyll synthesis pathway, and the Arabidopsis
DVR (At-DVR) is encoded by AT5G18660 (Nagata et al., 2005).
Recently, a cyanobacterial DVR (Sy-DVR) has been identified
from Synechocystis PCC6803 (Islam et al., 2008; Ito et al., 2008).
The Sy-DVR protein encoded by slr1923 is the only protein
responsible for the reduction of the 8-vinyl group. Interestingly,
Sy-DVR demonstrates no sequence similarity to At-DVR, and
Synechocystis PCC6803 has no genes showing homology to AtDVR. However, the Arabidopsis gene AT1G04620 was found to
be homologous to Sy-DVR. Figure 2 shows the alignment of the
amino acid sequences of Sy-DVR and AT1G04620. These two
proteins have a high sequence similarity (54% identity), and both
exhibit predicted binding motifs for a flavin and an iron-sulfur
cluster. In spite of this high sequence similarity, it appears that
AT1G04620 cannot substitute for AT5G18660 as mutants in
AT5G18660 have no monovinyl chlorophyll (Nagata et al., 2005).
The extended N-terminal region found in AT1G04620 was
predicted to be a chloroplast transit peptide (Emanuelsson et al.,
1999). Chloroplast localization of the protein was confirmed by
the expression of AT1G04620 fused to green fluorescence
protein (GFP) (see Supplemental Figure 1 online). GFP fluorescence colocalized with chlorophyll fluorescence, indicating chloroplast localization of AT1G04620.
Phenotype of the hcar Mutant
The high sequence similarity of AT1G04620 to Sy-DVR of the
chlorophyll synthesis pathway suggested the participation of the
AT1G04620 gene product in chlorophyll metabolism. To gain
insight into the function of the AT1G04620 gene product, we
analyzed two T-DNA insertion lines, SALK_018790C (hcar-1) and
CS908281 (hcar-2). We referred to this mutant as the hcar
mutant, as we found that the AT1G04620 gene product catalyzes
the conversion of HMChl to chlorophyll a in this study as it is
described hereafter. To confirm the absence of the AT1G04620
gene product in the hcar-1 mutant, we performed an immunoblot
analysis using an antibody against the protein. No signal was
obtained with protein extracts from the mutant (see Supplemental Figure 2 online), indicating that the mutant completely lacked
the gene product. However, we could not detect any difference in
the visible phenotype of green plants between the hcar-1 mutant
and the wild type (Figure 3). Interestingly, the mutant exhibited
the stay-green phenotype when the plants were transferred to
the darkness: the mutant remained green during dark incubation,
whereas in the wild type, the leaves turned yellow after 5 d of dark
incubation. To quantify the changes in chlorophyll content, we
measured chlorophyll levels during dark incubation (see Supplemental Figure 3 online). Decrease in chlorophyll content was
suppressed in both hcar mutants.
A closer examination of the mutant revealed that some leaves
were wilted after dark incubation. To quantify the cell death
phenotype of the hcar-1 mutant, electrolyte leakage was measured in the hcar-1 mutant during dark incubation (see Supplemental Figure 4 online). A large increase in electrolyte leakage
was observed in hcar-1 leaves, indicating that hcar-1 leaf cells
lost membrane integrity during dark incubation. This phenotype
was similar to the pao mutant, which has been shown to accumulate a large amount of pheophorbide a (Hirashima et al.,
2009).
Mutants of the enzymes for chlorophyll metabolism are known
to accumulate intermediate molecules of chlorophyll synthesis or
degradation. To investigate the effect of the mutation in the
AT1G04620 locus on chlorophyll metabolism, chlorophyll and its
precursors were extracted from green leaves of the wild type and
hcar mutants and subjected to HPLC (Figure 4). We found that
7-Hydroxymethyl Chlorophyll a Reductase
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Figure 1. Chlorophyll Metabolic Pathway in Land Plants.
Land plants possess two CBR isozymes, NYC1 and NOL. The metal-chelating substance (MCS) has not yet been identified. CS, chlorophyll synthase;
PPH, pheophytinase; PAO, pheophorbide a oxygenase.
the levels of chlorophyll a and chlorophyll b were similar between
the wild type and the mutants. Although we could not find any
visual differences between the green leaves, HMChl, an intermediate molecule between chlorophyll a and chlorophyll b, was
found in hcar mutants (Figures 4B and 4C). When senescent
leaves were examined by HPLC, an accumulation of pheophorbide a was found in addition to HMChl (Figures 4E and 4F).
HMChl and pheophorbide a were either not found or found at low
levels in wild-type leaves (Figures 4A and 4D).
Next, the levels of HMChl (Figure 5A) and pheophorbide a
(Figure 5B) in hcar-1 were quantified. The levels of HMChl were
between 30 and 60 nmol/g fresh weight, which corresponds to
;2% of the total chlorophyll. Furthermore, the level of pheophorbide a in hcar-1 was comparable to the level that has been
reported in an antisense pao mutant with an impaired PaO
previously reported as As-ACD1 (Tanaka et al., 2003). In addition, we found that pheophorbide a did not accumulate before
the dark incubation in the wild type, pao, or hcar-1 (Figure 5B); a
low level of pheophorbide a accumulated after 3 d of dark
incubation and drastically increased after 5 d in both of the
mutants. The level of pheophorbide a in hcar-1 corresponded to
;60% of that in the pao mutant after 5 d of dark incubation.
These results suggest that HCAR and PaO were both downregulated in hcar-1. Although HMChl did not accumulate in the
wild type during the early phase of dark incubation, substantial
amounts of HMChl accumulated after 5 d in the dark. The levels
of HMChl in the wild type varied among experiments. Unidentified environmental or developmental changes might induce the
accumulation of HMChl in the wild type.
Because the accumulation of pheophorbide a indicated an
impairment of PaO in the hcar-1 mutant, we examined the PaO
protein level by immunoblotting (see Supplemental Figure 2
online). The PaO level was similar to that of the wild type,
indicating that HCAR did not affect the expression of PaO. Next,
we examined the effect of HMChl accumulation on PaO activity
using the hcar-1 nyc1 nol triple mutant, in which HMChl is not
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Figure 2. Multiple Amino Acid Sequence Alignment of AT1G04620, Slr1923, and FpoF
AT1G04620 homologous sequences were aligned using ClustalW, as implemented in BioEdit. Conserved residues are shaded with black. Filled circles
represent the Cys residues that are predicted to coordinate the iron-sulfur cluster. Open circles represent the conserved sequence motif in FADcontaining proteins (KxxxxxGxG) (Dym and Eisenberg, 2001). FpoF does not contain this motif in the corresponding region. FpoF might contain an
unidentified FAD binding motif.
produced due to a lack of CBRs (Figure 5A). HMChl accumulated
in the hcar-1 mutant but not in the hcar-1 nyc1 nol triple mutant. A
large amount of pheophorbide a accumulated in the hcar-1 mutant, and small amount of pheophorbide a accumulated in the
wild type. Interestingly, the triple mutant did not accumulate
pheophorbide a even after 5 d of dark incubation (Figure 5B),
indicating a strong correlation between HMChl and pheophorbide a accumulation. HMChl is also an intermediate molecule of
chlorophyll b synthesis that is catalyzed by CAO. However,
HMChl was at a very low level or not detected in the hcar-1 nyc1
nol triple mutant, indicating that HMChl was produced from
chlorophyll b by CBR in the hcar-1 mutant.
amined the effect of the mutation in HCAR on the conversion of
chlorophyll b to chlorophyll a in greening tissues. Previously, we
reported that when greening cucumber (Cucumis sativus) cotyledons were exposed to light for several hours and then incubated in the dark, chlorophyll a increased with a concomitant
decrease in chlorophyll b without a significant change in the total
chlorophyll content (Tanaka et al., 1991). Considering that
Effects of the Mutation in HCAR on the Conversion of
Chlorophyll b to Chlorophyll a in Greening Cotyledons
The accumulation of HMChl in hcar suggested the involvement of
HCAR in the conversion of chlorophyll b to chlorophyll a. It has
been reported that the conversion of chlorophyll b to chlorophyll
a is high during both greening (Ito and Tanaka, 1996) and
senescence (Scheumann et al., 1999). Therefore, we next ex-
Figure 3. The Visible Phenotype of the Wild Type, pao, and hcar-1 during
Dark Incubation.
Wild-type (WT), pao, and hcar-1 plants were grown for 4 weeks under
continuous light and then incubated in complete darkness at 248C for the
indicated number of days.
7-Hydroxymethyl Chlorophyll a Reductase
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nant AT1G04620 protein in E. coli and determined whether it could
convert HMChl to chlorophyll a. The His-tagged protein lacking the
transit peptide was purified from E. coli as a single band (see
Supplemental Figure 5 online), as revealed by SDS-PAGE analysis, which showed 48 kD that corresponded to the expected size
of 50.9 kD (AT1G04620 lacking the transit sequence). The substrate, 7-hydroxymethyl chlorophyllide a (HMChlide), was incubated with the recombinant protein in the presence of ferredoxin,
ferredoxin NADPH oxidoreductase (FNR), and NADPH, and the
reaction products were examined by HPLC. After incubation, we
found that chlorophyllide a was formed with a concomitant decrease in HMChlide (Figure 7), indicating that AT1G04620 had
catalytic activity. Furthermore, the requirement for ferredoxin in
this reaction is consistent with the previously reported results
obtained with isolated plastids (Scheumann et al., 1998). The
Figure 4. The HPLC Profile of Chlorophyll Accumulated in the Wild Type
and hcar Mutants.
Seedlings were grown for 4 weeks under continuous light. Chlorophyll was
extracted from the wild type ([A] and [D]), hcar-1 ([B] and [E]), and hcar-2
([C] and [F]) before ([A] to [C]) and after ([D] to [F]) dark incubation for 5 d.
Chl a, chlorophyll a; Chl b, chlorophyll b; Pheide a, pheophorbide a.
chlorophyll is not newly synthesized in the dark in angiosperm
species, it was concluded that the increase in chlorophyll a and
decrease in chlorophyll b was caused by the conversion of
chlorophyll b to chlorophyll a during the dark incubation. When
we performed the same experiments with greening Arabidopsis
seedlings (Figure 6), both wild-type and hcar-1 mutant etiolated
seedlings that were exposed to light for 4 h accumulated chlorophyll a and chlorophyll b. When the wild-type seedlings were
incubated in the dark for 24 h, chlorophyll a increased with a
decrease in chlorophyll b (Figure 6), which is consistent with our
results obtained with cucumber cotyledons. This clearly shows
that the conversion of chlorophyll b to chlorophyll a actively
occurred in the Arabidopsis seedlings. When the same experiment was performed with the mutant, chlorophyll b decreased as
in wild type, but there was no concomitant chlorophyll a increase
(Figure 6). Interestingly, a large amount of HMChl accumulated
after the dark incubation, indicating that the mutation did not
seriously affect the conversion of chlorophyll b to HMChl but
almost completely inhibited the conversion of HMChl to chlorophyll a. This clearly shows that HCAR is involved in the conversion of HMChl to chlorophyll a.
Measurement of HCAR Activity of the AT1G04620
Recombinant Protein Expressed in E. coli
The accumulation of HMChl in greening cotyledons and green
leaves of the hcar mutant impaired in AT1G04620 suggested that
AT1G04620 is responsible for the conversion of HMChl to chlorophyll a. To examine this possibility, we expressed the recombi-
Figure 5. Chlorophyll Accumulation in the Wild Type and Mutants.
The wild type (WT), pao, and the hcar-1 and hcar-1 nyc1 nol mutants
were grown for 4 weeks under continuous light and then incubated in
complete darkness at 248C for the indicated number of days. HMChl (A)
and pheophorbide a (B) contents are indicated per gram fresh weight
(FW) of leaf. The error bars represent the SD of three samples from a
single harvest.
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Figure 6. Changes in the Chlorophyll Content during the Dark Incubation
of Greening Seedlings.
Wild-type and hcar-1 seeds were germinated for 3 d under dark conditions and transferred to light conditions (100 mmol m 2 s 1) at 248C for
4 h (0 hD). The seedlings were then transferred to complete dark
conditions for 24 h (24 hD). Chlorophylls were extracted before and after
the dark incubation from the seedlings and were subjected to HPLC
analysis. The error bars represent the SD for three samples. Black,
chlorophyll a; gray, chlorophyll b; white, HMChl.
natural substrate of HCAR should be HMChl instead of HMChlide
because NOL, which produces the substrate of HCAR, converts
chlorophyll b to HMChl and because HMChl, not HMChlide, accumulated in the hcar mutant. Accordingly, we used HMChl as the
substrate instead of HMChlide for the enzymatic assay. An E. coli
cell extract was used for this experiment because the cell extract
exhibited a higher activity than purified HCAR probably because
HCAR was partly inactivated during the purification procedures.
Although the level of chlorophyll a was low, HMChl was converted
to chlorophyll a by the recombinant protein (Figure 7). These
results indicate that the AT1G04620 product converted a hydroxymethyl group on the pyrrole ring to a methyl group using reduced
ferredoxin. Based on these experiments, AT1G04620 was named
HCAR.
We found that the four conserved Cys residues for the ironsulfur cluster binding motif were conserved in all of the HCARs of
green plants and in Sy-DVR (Figure 2). However, we could not
determine the presence of an iron-sulfur cluster merely by absorbance spectra. Thus, we measured the iron content of HCAR by a
colorimetric method and found that the iron content was 0.85
atoms per HCAR protein. It is well known that cofactors are often
lost during heterologous expression. Indeed, the expression of
iron-sulfur cluster-containing proteins has been reported to be
difficult, and a significant amount of expressed proteins have been
found to lack their iron-sulfur cluster (Oppenheimer et al., 2010).
Assuming that some portion of recombinant HCAR was denatured
and had lost the iron, holo-HCAR might potentially have more iron
per protein moiety. Both sequence analysis and iron determination
suggested the presence of an iron-sulfur cluster in HCAR, though
we have no information concerning the type or stoichiometry of
iron binding. Further information would be obtained by x-ray
crystallographic analysis.
Phylogenetic Analysis of HCAR
As described in Figure 2, HCAR has a high sequence similarity to
DVR of Synechocystis PCC 6803. These types of DVRs were
found in other cyanobacteria and in photosynthetic bacteria,
including purple bacteria and green sulfur bacteria. Figure 10
shows the phylogenetic tree of HCAR and DVR using F420H2
dehydrogenase as an outgroup. HCARs were found in both
The Redox Cofactors of HCAR
The amino acid alignment predicted binding motifs for both a
flavin (Dym and Eisenberg, 2001) and an iron-sulfur cluster
(Figure 2). As these two redox cofactors were expected to play
an important role in the enzymatic reaction, we examined the
presence of these factors by biochemical methods. Spectral
analysis of purified HCAR showed a peak at 450 nm, which is
typical of an oxidized flavin (Figure 8). The absorption peak at 450
nm disappeared when the flavin was reduced with dithionite,
indicating that HCAR contained a flavin molecule that may
function in catalysis. To identify which flavin species was present, we employed HPLC analysis. Figure 9 shows the HPLC
profile of authentic flavin adenine dinucleotide (FAD) and flavin
mononucleotide (FMN), which were clearly separated into two
peaks. Next, the flavin molecule was isolated from purified HCAR
and subjected to HPLC. Only one peak corresponding to FAD
was detected, indicating that the HCAR enzyme contains FAD.
Figure 7. The Conversion of HMChl(ide) to Chlorophyll(ide) a by HCAR.
Purified recombinant HCAR was incubated with HMChlide without any
reductant (A) or with NADPH, FNR, and ferredoxin (B). After incubation, the
pigments were extracted from the mixture and analyzed by HPLC.
HMChl was incubated with lysed E. coli harboring the empty vector (C) or
expressing recombinant HCAR (D), in the presence of NADPH, FNR, and
ferredoxin. After incubation, the pigments were extracted from the mixture
and analyzed by HPLC. Chlide a, chlorophyllide a; Chl a, chlorophyll a.
7-Hydroxymethyl Chlorophyll a Reductase
Figure 8. The Absorbance Spectra of the Oxidized and Reduced HCAR.
Purified HCAR (1 mg/mL, solid line) expressed in E. coli was reduced by
the addition of dithionite (10 mM, dashed line).
green plants and green algae. In this tree, HCAR branched off
within the DVR cluster and was most closely related to diatom
DVR phylogenetically, suggesting that HCAR branched off from
DVR at the eukaryotic stage. These results indicate that HCAR of
the chlorophyll cycle evolved from DVR of the bacterial chlorophyll synthesis pathway.
DISCUSSION
The Reduction of HMChl to Chlorophyll a by HCAR
It has been reported that the F subunit of F420H2 dehydrogenase
of Methanolobus tindarius (Fpo) is an iron-sulfur flavoprotein
(Deppenmeier, 2004) and has sequence similarity to HCAR. In
this study, we succeeded in detecting FAD in recombinant HCAR
that was reduced by sodium dithionite. The binding motif for an
iron-sulfur center was conserved at the N-terminal region of
F420H2 dehydrogenase (Johnson and Mukhopadhyay, 2005) and
HCAR, and iron was detected in recombinant HCAR. From these
results, we concluded that HCAR contains a flavin and an ironsulfur cluster as redox cofactors. However, the route of electron
transfer from ferredoxin to HMChl in HCAR is uncertain at
present. In the F subunit of the Fpo complex, the two-electron
carrier F420H2 reduces FAD, from which an electron is transferred
to an iron-sulfur cluster (Kulkarni et al., 2009); the iron-sulfur
cluster of the F subunit then transfers an electron to the BCDI
subunit of Fpo. The FrhB subunit of the F420-reducing hydrogenase complex also has weak homology to HCAR, and it has been
reported to contain FAD but lack the iron-sulfur cluster (Johnson
and Mukhopadhyay, 2005). FrhB passes the electron of F420H2
to the iron-sulfur cluster of the FrhG subunit in the complex.
Accordingly, the sequence similarity between HCAR, FpoF, and
FrhB suggests an electron transfer from FAD to the iron-sulfur
cluster in HCAR. However, this hypothesis is not necessarily
accepted because HCAR receives an electron from a oneelectron carrier (ferredoxin), yet FpoF and FrhB receive electrons
from a two-electron carrier (F420H2). Another proposal is that
ferredoxin first reduces the iron-sulfur cluster and then an elec-
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tron is transferred to FAD, which stores two electrons and
reduces HMChl to chlorophyll a. This idea is consistent with
the reported electron transfer in ferredoxin-dependent Glu synthase, an iron-sulfur flavoprotein, in which flavin is reduced by
ferredoxin via the iron-sulfur cluster (Vanoni et al., 2005). Further
study will be required to elucidate the electron transfer in HCAR.
The reductive elimination of a hydroxyl group is a difficult
reaction due to the strong carbon-oxygen bond. This is the
reason why enzymatic examples for this reaction are rarely found
in the literature. A well-studied example is ribonucleotide reductase, which catalyzes the replacement of the ribose 29-OH in
ribonucleotides with hydrogen (Bollinger et al., 2008); a free
radical is an essential component for this replacement. However,
a catalytic mechanism for the replacement of an OH by a hydrogen on a pyrrole ring has not yet been elucidated. Based on
the finding that the conversion of chlorophyll b to chlorophyll a
occurs in vivo in the presence of reduced ferredoxin and ATP,
Folly and Engel (1999) have proposed a reaction mechanism for
the conversion of HMChl to chlorophyll a. These authors hypothesized that the unique electron arrangement of the cyclic 18p electron porphyrin system facilitates the elimination of water
and the stabilization of the carbocation, but this process demands an activator, such as ATP. In this report, we showed that
ATP was not required for the substitution of the OH with hydrogen in the reaction catalyzed by HCAR, which has a high
sequence similarity to the DVR enzyme that catalyzes the reduction of –CH=CH2 to –CH2–CH3. Thus, a portion of the reaction
mechanism of HCAR may be similar to the DVR reaction. Based
on these findings, we propose the following reaction mechanism
for HCAR, as described below (see Supplemental Figure 6
online). The first step is the production of a hydroxymethyl cation. The loss of a water molecule and the addition of an electron
yields a methylene radical, which is converted to a methylene
anion or a methyl radical and then to a methyl group.
Figure 9. HPLC Analysis of the Released Flavin from HCAR.
Authentic FAD and FMN (A) and flavin extracted from purified HCAR (B)
were analyzed by HPLC. The flavin was released from recombinant HCAR by
heat treatment.
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Figure 10. Phylogenetic Tree of HCAR.
Neighbor-joining tree constructed with the translated sequence of HCAR and DVR. Bootstrap values for each clade are indicated on each node, and the
scale bar indicates the number of amino acid substitutions per site. The F subunit of Methanosaeta F420H2 dehydrogenase was used as an outgroup.
The alignment used for this analysis is available as Supplemental Data Set 1 online.
Clearly, the exact reaction mechanism may be elucidated by
an electron transfer route and the crystal structure of HCAR.
The Regulation of PaO Activity by HCAR
The hcar-1 mutant carries a T-DNA in an exon, and HCAR was
below the level detectable by immunoblotting, indicating that the
mutant completely lacked HCAR activity; however, the level of
HMChl was also low. One possible reason for this may be the
conversion of HMChl to other molecules, such as 7-hydroxymehyl
pheophorbide a and HMChlide, although these molecules were
not detected by HPLC. The same phenomenon has been reported
with the Arabidopsis pheophytinase mutant (Schelbert et al.,
2009). In this mutant, the conversion from pheophytin to pheophorbide was reported to be completely blocked, yet the level of
pheophytin was low. Therefore, it might be reasonable to speculate that the chlorophyll degradation pathway has another route,
in addition to the major route, due to the wide substrate specificity.
In the chlorophyll cycle, the first reaction, the conversion of
chlorophyll b to HMChl, is catalyzed by CBR; the second re-
action, the conversion of HMChl to chlorophyll a, is catalyzed by
HCAR (Figure 1). In Arabidopsis, there are two CBR genes, NOL
and NYC1 (Horie et al., 2009). These two CBRs have a high
sequence similarity but are differentially expressed during development in Arabidopsis. HCAR is encoded by a single gene
and is constitutively expressed. It is expected that HCAR and
NOL should be regulated to convert chlorophyll b to chlorophyll a
efficiently during greening; by contrast, HCAR and NYC1 should
be coordinated during senescence. However, HCAR does not
affect the activity of CBR because the conversion of chlorophyll b
to HMChl occurred in the greening cotyledons of the hcar
mutant, as well as in the wild type. The proposed independent
regulation of these two enzymes would be different from PaO
and red chlorophyll catabolite reductase (Pruzinská et al., 2003).
By contrast, the mutation in HCAR induced the accumulation of
pheophorbide a, a degradation product of chlorophyll. The level of
accumulated pheophorbide a in the hcar-1 mutant was almost
60% of what has been reported for the pao mutant (Tanaka et al.,
2003), indicating that PAO activity was severely downregulated in
the hcar mutant. Immunoblot analysis clearly showed that the PaO
7-Hydroxymethyl Chlorophyll a Reductase
protein level in the hcar-1 mutant was almost the same as in the
wild type, indicating that PaO catalytic activity was downregulated
in the hcar mutant. One possible hypothesis is that PaO is active
only when HCAR is present, for example, due to a direct interaction between the two enzymes. However, this idea is not consistent with the report that PaO is active without HCAR in in vitro
experiments (Pruzinská et al., 2003). A second hypothesis is the
inhibition of PaO activity via HMChl, which accumulated in the hcar
mutant. This hypothesis is supported by the results that pheophorbide a did not accumulate in the hcar-1 nyc1 nol triple mutant,
which also did not accumulate HMChl; it is also consistent with in
vitro experiments that suggested that pheophorbide b was a
competitor of PaO, and FCC production by PaO was inhibited by
pheophorbide b (Hörtensteiner et al., 1995). However, this hypothesis cannot be fully accepted because PaO activity was not
completely inhibited by pheophorbide b in in vitro experiments; by
contrast, PaO seemed to be severely inhibited in the hcar mutant.
We suggest that it would be difficult for HMChl to be a competitive
inhibitor of PaO because HMChl retains the phytyl chain, making it
difficult to access PaO, which resides in the chloroplast envelope.
Although HMChl might not directly regulate PaO activity, it is
possible to speculate that HMChl or its degradation products is
involved in modulation of PaO activity through an unidentified
mechanism. This idea is supported by the finding that both HMChl
and pheophorbide a accumulated in the Arabidopsis hmc1 mutant, which has a defect in the Nap1 gene (Nagane et al., 2010).
The Distribution and Evolution of HCAR
DVR has been identified from Synechocystis PCC6803 by a
bioinformatics method (Ito et al., 2008). This DVR is found to be
present in other cyanobacteria, except for marine types of Synechococcus and Prochlorococcus, some photosynthetic bacterial
groups, and diatoms. Interestingly, the phylogenetic tree of HCAR
and DVR clearly showed that HCAR appeared within the DVR clade,
indicating that HCAR evolved from DVR (Figure 10). This phylogenetic tree strongly suggests that the ancestor of chlorophytes and
chromophytes should have both types of DVRs (At-DVR and SyDVR) and that Sy-DVR evolved to HCAR in the green lineage. Thus,
a chlorophyll biosynthesis enzyme may have evolved to catalyze
other steps of chlorophyll metabolism. Although the phylogenetic
relationship is not clear, sequence similarities are found between the
different enzymes for tetrapyrrole metabolism. PaO and CAO
belong to the same Rieske-non heme iron oxygenase family (Gray
et al., 2002) and have sequence similarity. The amino acid sequences of red chlorophyll catabolite reductase, which reduces the
linear tetrapyrrole of the red chlorophyll catabolite, were shown to
have a low degree of sequence similarity to ferredoxin-dependent
bilin reductases (Sugishima et al., 2009). This phenomenon is
reasonable because the substrates (linear or open tetrapyrroles)
have common structure, and the reaction mechanisms are partly
shared. In addition to chlorophyll a, there are many chlorophyll
species, such as chlorophyll b, chlorophyll c, chlorophyll d, and
chlorophyll f (Chen et al., 2010). It might be possible that changes in
the catalytic properties of chlorophyll biosynthetic enzymes contribute to the diversity of chlorophyll species.
HCAR has been found only in green algae and land plants,
such as Arabidopsis, mosses, and prasinophytes. Therefore, it
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may be concluded that HCAR was acquired in the green lineage.
The distribution of CBR is consistent with that of HCAR, which
suggests a coevolution of HCAR and CBR. However, not all of
the chlorophyll b–containing organisms have necessarily retained the chlorophyll cycle. In the cyanobacterial lineage,
Prochlorococcus has chlorophyll b, but it does not contain genes
for CBR, HCAR, or cyanobacterial DVR in its genome. Thus, the
appearance of the chlorophyll cycle seems to be consistent with
that of LHC in green lineages. This is reasonable because the
chlorophyll cycle regulates the formation and degradation of
the LHC.
With this study, all of the enzymes responsible for the chlorophyll cycle are now identified. This progress will enable the
detailed molecular analysis of the regulation and function of the
chlorophyll cycle in various physiological processes, such as
greening, acclimation, and senescence.
METHODS
Plant Material and Growth Conditions
Arabidopsis thaliana (Columbia ecotype) was grown for 4 to 6 weeks in a
chamber equipped with white fluorescent lamps under continuous illumination at a light intensity of 100 mmol m–2 s–1 at 248C. The plants were
subsequently transferred to darkness for 3 to 9 d at 248C for the chlorophyll
degradation experiments. The leaves were harvested at the times indicated
in Results. For the greening experiments, seedlings were grown on agar
plates containing half-strength diluted Murashige and Skoog medium for 3
d in the dark. Etiolated seedlings were illuminated (100 mmol m–2 s–1) for 4 h
and incubated in the dark for an additional 24 h. The T-DNA insertion
mutants, lacking either AT1G04620 (SALK_018790C, hcar-1; CS908281,
hcar-2), AT4G13250 (SALK_091664, nyc1), or AT5G04900 (AL759262, nol),
were obtained from the ABRC (Ohio State University) and GABI-Kat,
respectively. Every mutant was crossed, and the triple mutant was identified by PCR-based genotyping. Production of the pao mutant that
overexpressed antisense RNA for the PaO gene was described in our
previous report (Tanaka et al., 2003).
Pigment Analysis
Leaves were weighed and pulverized in acetone using a ShakeMaster
grinding apparatus (BioMedical Science), and the extracts were centrifuged for 15 min at 22,000g. The pigments were separated on a Symmetry
C8 column (150 3 4.6 mm; Waters), according to a method reported
previously (Zapata et al., 2000). The elution profiles were monitored by
measuring the absorbance at 653 nm (SPD-M10A; Shimadzu), and the
pigments were identified by their retention times and spectral patterns.
Pigment quantification was performed using the area of the peaks.
Enzymatic Assay
The coding region of HCAR lacking its transit peptide was amplified by
PCR (PrimeSTAR Max DNA polymerase; Takara) using primers
59-GGATCCTCCGTCGTTAACTCTTCTTC-39 (the underlined region is
an engineered BamHI site) and 59-CTCGAGTTTCTTGGAGAGCATTTTAT-39 (the underlined region is an engineered XhoI site) and cloned into
the BamHI/XhoI sites of pET-30a(+) (Novagen). The recombinant protein
obtained from this plasmid possesses His tags at the both the N and C
termini. This recombinant protein did not exhibit enzymatic activity, and it
was expected that the His tag at the N terminus disturbed the HCAR
activity. Therefore, the His tag at the N terminus was removed by PCR
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using the primers 59-ATGTCCGTCGTTAACTCTTCTTC-39 and 59-ATGTATATCTCCTTCTTAAA-39 to amplify the plasmid DNA. The PCR products were phosphorylated by T4 polynucleotide kinase (Takara), and
self-ligation was performed. The expression plasmid was introduced into
Escherichia coli BL21 cells. The transformed cells were grown at 378C
with 200 mL Luria-Bertani medium containing kanamycin (50 mg/mL) and
100 mM of ammonium ferric citrate until the optical density at 600 nm
reached at 1.3. The expression of the HCAR gene was induced with 0.2
mM isopropyl-b-D-thiogalactopyranoside for 3 h. After incubation, the
culture was harvested by centrifugation, and the collected cells were
resuspended in buffer (5 mM imidazole, 500 mM NaCl, and 20 mM TrisHCl, pH 7.9) and disrupted by sonication. The recombinant HCAR
contained in the soluble fraction was purified using a nickel column (His
bind kit; Novagen); a desalting column (Mini Trap G-25; GE Healthcare)
was used to remove the imidazole from the purified HCAR fraction.
HMChl was obtained by reducing chlorophyll b with 100 mM NaBH4 in
methanol, whereas HMChlide was obtained from HMChl by hydrolysis
with recombinant chlorophyllase (Tsuchiya et al., 1999).
Purified HCAR was diluted to a concentration of 160 mg/mL in buffer (50
mM Tris-HCl, pH 7.5, and 100 mM NaCl) containing 0.9 mg/mL Glc, 0.4
mg/mL Glc oxidase (Nacalai), and 0.04 mg/mL catalase (Sigma-Aldrich)
to reduce the O2 level in the reaction mixture. Fifty microliters of the
mixture was used for the reaction. HMChlide was dissolved in acetone to
a concentration of 0.15 mM, and 1 mL of HMChlide solution, 1 mL of 50
mM NADPH, 1 mL of spinach (Spinacia oleracea) FNR (0.1 mg/mL; SigmaAldrich) and 1 mL of spinach ferredoxin (2.2 mg/mL; Sigma-Aldrich) were
added to the reaction mixture.
When HMChl was used as the substrate, HCAR-expressing E. coli cells
were lysed with 4 mL of BugBuster (Novagen), and 50 mL of the lysate and
1mL of HMChl (0.09 mM) dissolved in acetone were incubated with the an
above-described reductant. The mixtures were incubated for 15 min at
378C, after which, 200 mL of acetone was added. After centrifugation at
22,000g for 10 min, the supernatant containing the chlorophyll derivatives
was analyzed by HPLC as described above.
Spectral Measurement of HCAR
The absorbance spectrum of the purified HCAR was measured at a
concentration of 1 mg/mL (U-331 spectrophotometer; Hitachi). For the
reduction of HCAR, dithionite was added to a concentration of 10 mM.
Determination of the Flavin Species in HCAR
Recombinant Arabidopsis HCAR protein that was overexpressed in E.
coli and then purified as described above was incubated at 1008C for 7
min to release the flavin. After centrifugation at 22,000g for 10 min, the
supernatant (containing the flavin) and FAD and FMN standards (SigmaAldrich) were analyzed by HPLC. An aliquot of the sample was subjected
to a ODS column (6 3 150 mm, Shim-Pack ODS; Shimadzu) equilibrated
with 5 mM ammonium acetate, pH 6.0 (buffer A). The column was eluted
with a linear gradient of 0 to 30% buffer B (methanol containing 5 mM
ammonium acetate, pH 6.0) for 9 min and linear gradient of 100% buffer B
for 1 min at a flow rate of 0.75 mL/min. The effluent from the column was
monitored at the wavelength of 450 nm (Stuehr et al., 1991).
Quantification of Iron in HCAR
The iron content was determined colorimetrically using bathophenanthroline as an iron-chelating agent. Three hundred microliters of the eluate
from the nickel column that contains recombinant HCAR (see above) was
incubation with 150 mL of 1 N HCl at 958C for 2 min. To remove the
insoluble material, 150 mL of trichloroacetic acid (100% [w/v]) was added.
After centrifugation, 150 mL of ascorbic acid (0.16 mg/mL) was added to
300 mL of the supernatant. Bathophenanthroline solution (150 mL of 0.67
mg/mL) and 150 mL of sodium acetate solution (300 mg/mL) were added
to the mixture. The samples were incubated at room temperature for 5
min, and the absorbance at 535 nm was measured. The concentration of
ferrous ion was calculated using an iron standard solution (Wako).
Sequence Analysis
The database search was performed using the National Center for Biotechnology Information database (http://www.ncbi.nlm.nih.gov/blast). Amino
acid sequences were aligned using the ClustalW program (Thompson
et al., 1994) in the BioEdit program (http://www.mbio.ncsu.edu/BioEdit/
bioedit.html).
Phylogenetic Analysis
For the phylogenetic analysis, the protein sequences were aligned using
MEGA 4 software (Tamura et al., 2007), and the midpoint-rooted neighborjoining tree was generated using the same software with the following
parameters: Poisson model, uniform rates, complete deletion, and bootstrap (1000 replicates).
Plasmid Construction and Plant Transformation
To express HCAR-GFP fusion proteins, we modified the pGreenII MH
binary vector that was constructed in our previous study (Yamasato et al.,
2005). The vector contained the cauliflower mosaic virus 35S promoter,
the tobacco mosaic virus v sequence, the GFP (S65T) sequence, and the
nopaline terminator in the backbone of pGreenII-0029 (Hellens et al.,
2000). The coding region of HCAR was amplified by PCR using the
primers 59- GTCGACATGATTACTGTCGTCACCTC- 39 (the underlined
section is an engineered SalI site) and 59- CCATGGATTTCTTGGAGAGCATTTTAT-39 (the underlined section is an engineered NcoI site). The
GFP (S65T) gene in this plasmid was fused with the HCAR gene at the SalI
and NcoI sites. The plasmids were subsequently transformed into an
Agrobacterium tumefaciens (strain GV2260), and wild-type Arabidopsis
was transformed.
Analysis of GFP Expression by Confocal Laser Scanning Microscopy
Fluorescence images were recorded on a C1si Spectral Imaging confocal
laser scanning microscopy system with an ECLIPSE 80i microscope
(Nikon). The microscope was equipped with a Nikon Plan Apo VC 3100
objective. An argon laser was used to generate an excitation source at
488 nm, and GFP fluorescence was recorded at 500 to 550 nm. A blue
diode laser was used to generate an excitation source at 405 nm, and
chlorophyll fluorescence was recorded at 650 to 680 nm. Images were
processed with EZ-C1 Viewer 3.20 (Nikon).
Immunoblot Analysis
Total protein was extracted from leaves by grinding with extraction buffer
(50 mM Tris, pH 6.8, 2 mM EDTA, 10% [w/v] glycerol, 2% [w/v] SDS, and
6% [v/v] 2-mercaptoethanol). Homogenates were centrifuged at 22,000g
for 10 min. The supernatants were separated by 10% SDS-PAGE, and the
resolved proteins were transferred onto a Hybond-P membrane (GE
Healthcare). HCAR protein was detected with an anti-HCAR rabbit
primary antiserum (diluted 1:10,000) that was raised against the recombinant Arabidopsis HCAR polypeptide used for the enzymatic assay. PaO
protein was detected as previously reported (Hirashima et al., 2009). Antirabbit IgG linked to horseradish peroxidase (GE Healthcare) was used as
a secondary antibody. The horseradish peroxidase activity was detected
using the ECL Plus protein gel blotting detection system (GE Healthcare)
following the manufacturer’s protocol.
7-Hydroxymethyl Chlorophyll a Reductase
Accession Numbers
The HCAR homologous protein amino acid sequence data from this article
can be found in the GenBank/EMBL data libraries under the following
accession numbers: Anabaena variabilis ATCC 29413, YP_324708; Arabidopsis, NP_171956; Chlamydomonas reinhardtii, XP_001699546; Chlorobium phaeobacteroides DSM 266, YP_910679; Chloroflexus aurantiacus
J-10-fl, YP_001636150; Gloeobacter violaceus PCC 7421, NP_923824;
Methanosaeta thermophila PT, YP_842613; Micromonas sp RCC299,
XP_002503439; Oryza sativa japonica Group, CAE01504; Ostreococcus
lucimarinus CCE9901, XP_001416225; Ostreococcus tauri, XP_003080144;
Physcomitrella patens subsp patens, XP_001770443; Populus trichocarpa,
XP_002315601; Rhodopseudomonas palustris BisA53, YP_780232; R. palustris BisB5, YP_570899; Synechococcus elongatus PCC 6301, YP_170905;
Synechococcus sp WH 5701, ZP_01086192; Synechococcus sp WH 7805,
ZP_01124436; Synechocystis sp PCC 6803, NP_441896; Thalassiosira
pseudonana CCMP1335, XP_002288079; Thermosynechococcus elongatus
BP-1, NP_682635; and Vitis vinifera, XP_002285592.
Supplemental Data
The following materials are available in the online version of this article.
Supplemental Figure 1. HCAR Accumulation in the Chloroplast.
Supplemental Figure 2. Immunoblot Analysis of HCAR and PaO in
Wild-Type and hcar-1 Plants.
Supplemental Figure 3. Chlorophyll Retention during Dark Incubation.
Supplemental Figure 4. Electrolyte Leakage of the Leaves during
Dark Incubation.
Supplemental Figure 5. SDS-PAGE Analysis of Purified HCAR.
Supplemental Figure 6. Hypothesized Reaction Pathway of Hydroxymethyl Reduction.
Supplemental Data Set 1. Alignment of HCAR Homolog for the
Construction of Phylogenetic Tree in Figure 10.
ACKNOWLEDGMENTS
We thank Hitoshi Tamiaki for valuable discussion and Sachiko Tanaka
for technical assistance. This work was supported by a Grant-in-Aid for
Scientific Research (No. 21370014 to A.T. and No. 68700307 to R.T.)
from the Ministry of Education, Culture, Sports, Science and Technology
of Japan.
AUTHOR CONTRIBUTIONS
M.M., H.I., and A.Tanaka designed the research. M.M. and H.I. performed
research. M.M., H.I., A.Takabayashi, R.T., and A.Tanaka analyzed data.
H.I. and A.Tanaka wrote the article.
Received July 26, 2011; revised August 23, 2011; accepted September 2,
2011; published September 20, 2011.
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Identification of the 7-Hydroxymethyl Chlorophyll a Reductase of the Chlorophyll Cycle in
Arabidopsis
Miki Meguro, Hisashi Ito, Atsushi Takabayashi, Ryouichi Tanaka and Ayumi Tanaka
Plant Cell; originally published online September 20, 2011;
DOI 10.1105/tpc.111.089714
This information is current as of June 18, 2017
Supplemental Data
/content/suppl/2011/09/09/tpc.111.089714.DC1.html
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