From the Université Joseph Fourier et CNRS, UMR
5575, BP53, CERMO, F-38041 Grenoble cedex 9, France, the
§ Lehrstuhl für Pflanzenphysiologie,
Ruhr-Universität Bochum, Universitätsstraße 150, D-44801
Bochum, and the ¶ Lehrstuhl für Pflanzenphysiologie,
Universität Bayreuth, Universitätsstraße 30,
D-95447 Bayreuth, Germany
Received for publication, September 23, 2002, and in revised form, October 15, 2002
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ABSTRACT |
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We recently put forth a model of a
protochlorophyllide (Pchlide) light-harvesting complex operative
during angiosperm seedling de-etiolation (Reinbothe, C., Lebedev, N.,
and Reinbothe, S. (1999) Nature 397, 80-84). This model,
which was based on in vitro reconstitution experiments with
zinc analogs of Pchlide a and Pchlide b and the two NADPH:protochlorophyllide oxidoreductases (PORs), PORA and PORB, of
barley, predicted a 5-fold excess of Pchlide b,
relative to Pchlide a, in the prolamellar body of
etioplasts. Recent work (Scheumann, V., Klement, H., Helfrich, M.,
Oster, U., Schoch, S., and Rüdiger, W. (1999) FEBS
Lett. 445, 445-448), however, contradicted this model and
reported that Pchlide b would not be present in etiolated
plants. Here we demonstrate that Pchlide b is an abundant
pigment in barley etioplasts but is rather metabolically unstable. It
is rapidly converted to Pchlide a by virtue of 7-formyl reductase activity, an enzyme that had previously been implicated in
the chlorophyll (Chl) b to Chl a reaction
cycle. Our findings suggest that etiolated plants make use of 7-formyl
reductase to fine tune the levels of Pchlide b and Pchlide
a and thereby may regulate the steady-state level of
light-harvesting POR-Pchlide complex.
Angiosperms have developed sophisticated mechanisms to harvest
sunlight and to convert this into various physiological responses (1).
They make use of various photoreceptors, such as the red/far red
light-absorbing phytochromes, the blue light-absorbing cryptochromes, and the blue light-absorbing phototropins, to adapt to different light
qualities and quantities and to sense the direction and duration of
incident light (for a review, see Ref. 2). All of these photoreceptors
are chromoproteins, which undergo characteristic spectral changes upon illumination.
Another blue and red light-absorbing protein-pigment complex is the
protochlorophyllide
(Pchlide)1 holochrome (3). It
is localized in the prolamellar body of etioplasts. These plastids form
when angiosperms germinate in darkness. The entire developmental
process of seedling germination leading to prolamellar bodies is termed
skotomorphogenesis or etiolation (1). In plants displaying an hypogeic
type of germination, it takes place underneath the soil. The newborn
seedlings then utilize all nutrient reserves contained in the seed to
bring the cotyledons above the soil.
Previous work has shown that the Pchlide holochrome is a higher
molecular mass complex of about 600 kDa (3). More recent work suggested
that it may be composed of galacto- and sulfolipids (4), Pchlide (5,
6), and an enzyme called the NADPH:Pchlide oxidoreductase (POR; EC
1.3.33.1) (7, 8). It was discovered that two distinct forms of POR
exist in barley etioplasts, called PORA and PORB (9, 10). Moreover, two
species of Pchlide have been distinguished in isolated prolamellar
bodies by low temperature in situ fluorescence measurements:
Pchlide 628/632 (the first number indicates the absorption maximum, the
second the respective fluorescence emission maximum at the chosen
excitation wavelength) and Pchlide 650/657 (for a review, see Ref. 11).
Whereas the former remained quantitatively unchanged upon illumination
with a single, 1-ms flash of white light, the latter was readily
converted to Chlide 684/690. This differential behavior led scientists
to name Pchlide 628/632 photoinactive and Pchlide 650/657 photoactive (summarized in Ref. 11). Both before and after flash light
illumination, energy transfer was observed, taking place from
photoinactive Pchlide to photoactive Pchlide in etiolated plants and
from photoinactive Pchlide to Chlide in preflashed plants (12-17).
To resolve all of these puzzling previous observations, we proposed a
model of a "light-harvesting POR-Pchlide" complex, named LHPP (18).
Based on in vitro reconstitution experiments with synthetic
zinc analogs of Pchlide, we put forth the idea that LHPP may be
composed of 5 PORA-Pchlide b-NADPH ternary complexes and 1 PORB-Pchlide a-NADPH ternary complex embedded into the lipid bilayers of the prolamellar body of etioplasts (18) (see also Ref. 19
for a summary).
A particularly important question that had thus far remained unanswered
was whether Pchlide b implicit in the LHPP model would be
present in etiolated barley plants. Whereas previous work had indicated
that Pchlide b is present in green plants (20), no comparable study had thus far been available reporting the
identification of Pchlide b in etiolated plants, where the
pigment, according to our in vitro reconstitution
experiments (18), should be found in maximum levels. In a recent paper,
Scheumann et al. (21) even generally questioned the
existence of Pchlide b, but at the same time demonstrated
that barley etioplasts rapidly convert exogenously added zinc
protopheophorbide b (ZnPPb) to ZnPPa.
This prompted us to conclude that Chl(ide) b reductase,
presumably responsible for this conversion (22-26), could also
metabolize the endogenously occurring Pchlide b to Pchlide
a.
In the present study, we readdressed the experimental design of
Scheumann et al. (21). We demonstrate that Chl(ide)
b reductase (which may alternatively be named 7-formyl
reductase; see below) is indeed able to convert Pchlide b to
Pchlide a in situ. This reaction already occurs upon plastid
lysis and subsequent detergent solubilization of isolated prolamellar
bodies. Both experimental steps lead to a denaturation of the
prolamellar body and the release of the PORA and make Pchlide
b readily accessible to 7-formyl reductase. Our results
provoke the idea that 7-formyl reductase may be involved in fine tuning
the levels of Pchlide b and Pchlide a in etioplasts.
Pigments--
All glassware used throughout this study was
pretreated with diethyl pyrocarbonate (DEP). This compound had
previously been shown to inhibit bacterial and plant Rieske-type
oxygenases (27, 28), to which Chlide a oxygenase and related
enzymes belong (29, 30). To block this activity seemed particularly
necessary, in order to allow accurate determination of Pchlide
a and Pchlide b levels, respectively.
Pigments were extracted from intact barley etioplasts as described
herein and in Ref. 31. Separation by HPLC was performed on a C18
reverse phase silica gel column (Macherey-Nagel Co., 250 × 4.6 mm, Nucleosil ODS 5 µm) as described in Ref. 32. Either a step
gradient was used, starting with 34% 25 mM aqueous
ammonium acetate, 15% acetone, and 51% methanol (buffer A),
increasing to 16% H2O, 60% acetone, and 24% methanol
within 20 min (buffer B), and finally to 100% acetone another 4 min
later, or linear gradients from buffer A to buffer B. Absorbance
measurements were made at 455 nm, which corresponds to the Soret band
of Pchlide b, to detect and quantify Pchlide a
and Pchlide b levels. At this wavelength, the extinction
coefficients of Pchlide b and Pchlide a are
5-fold different (21). As internal standards, we used synthetic
Pchlides a and b, which were prepared from
Chlides a and b with an excess of
2,3-dichloro-5,6-dicyanobenzoquinone as described in Refs. 21 and 32.
At a flow rate of 1 ml/min, Pchlide a has a retention time
of ~15 min, and Pchlide b has a retention time of ~12.5
min. For simultaneous separation of Pchlide a and Pchlide
b and their reduced products (i.e. Chlides
a and b, respectively), a C30 reverse phase
column (250 × 4.6 mm, 5 µm; YMC Inc., Willmington, NC) (33) was
used. HPLC was performed in a Varian ProStar model 410 apparatus, a
ProStar model 240 pump, and a ProStar 330 photodiode array detector,
essentially as described in Ref. 33 (see accompanying paper (31) for
details). In some experiments, a combination of octadecyl silica and
poly(ethylene) powder media was used, likewise allowing separation of
both porphyrins and chlorins in the same HPLC run (34).
Chemical synthesis of ZnPPa and ZnPPb and their
binding to isolated prolamellar body membranes was performed as
described in Ref. 32. 7-Hydroxy-Pchlide a was synthesized as
described herein. Liquid secondary ion mass spectrometry was performed
in an m-nitrobenzyl alcohol matrix with a Finnigan model
MAT900 and a cesium gun (20 kV, 1 mA) according to Schoch
et al. (32).
Preparation and Solubilization of Prolamellar
Bodies--
Etioplasts were prepared from etiolated barley plants by
Percoll density gradient centrifugation as described previously (35, 36). For low temperature analyses at 77 K (see below) and pigment measurements (see above), the etioplast suspension was directly used.
Plastids to be lysed and further subfractionated were diluted with the
buffer described by Li et al. (37). After sedimentation, the
latter plastids were fractionated into prolamellar bodies, prothylakoids, and stroma, on discontinuous step gradients of sucrose
(38). n-Octyl- Protein and Pigment Analyses--
Etioplasts prepared on a
Percoll gradient were extracted with an 100-fold excess of either 100%
acetone containing 0.1% DEP or only 80% acetone lacking DEP. By
analogy, plastid subfractions obtained as described above were treated
identically. Protein was recovered by centrifugation, washed several
times with ethanol and ether, and run by SDS-PAGE on 10-20% (w/v)
polyacrylamide gradients, whereas pigments found in the corresponding
supernatant fractions were directly used for fluorescence measurements
(see below).
Immunodecoration of electrophoretically resolved proteins was performed
using an ECL Western blotting analysis system (Amersham Biosciences)
and an anti-POR-specific antiserum (9).
Spectroscopic Analyses--
Low temperature fluorescence
measurements were performed at 77 K at an excitation wavelength of 440 nm (39), in a spectrometer LS50B (PerkinElmer Life Sciences).
Previous studies had shown that barley and cucumber etioplasts
contain enzyme activity that converts Chl b to Chl
a (22). By analogy, this enzyme activity was found to also
convert the nonesterified precursor of Chl b, Chlide
b, as well as pyrochlorophyllide b and the
magnesium-free pheophorbide b into the respective Chl a and 7-hydroxy compounds (23-26).
To test whether this enzyme activity, which we tentatively named
7-formyl reductase to indicate this broad substrate specificity, would
also be able to convert Pchlide b to Pchlide a in
situ, we followed the experimental design of Scheumann et
al. (21). Briefly, intact barley etioplasts were isolated on a
Percoll gradient, sedimented by centrifugation, lysed, and incubated
with ZnPPb, the zinc analog of Pchlide b (32). As
a control, barley etioplasts were left intact (26) and incubated
identically. Mock incubations lacking ZnPPb were conducted
in parallel.
Two different pigment extraction procedures were used. In the first
case, we used an almost pure, nonaqueous solution of acetone containing
0.1% DEP, which was used in order to block the potential generation of
Pchlide b by virtue of the previously identified Chlide
a oxygenase and related enzymes (27-30). This solution is referred to as 100% acetone throughout the rest of the paper. In the
second case, an aqueous, non-DEP-supplemented solution containing only
80% acetone was used, as reported in Ref. 21. Pigment analyses were
made by HPLC, using a photodiode array detector. As shown previously
(21, 32), this allowed simultaneous separation and identification of
pigments during the actual HPLC run.
When etiolated leaf material was extracted with 100% acetone, HPLC
analyses revealed the existence of three main porphyrin species,
eluting at 11 min (peak 1), 12.5 min (peak 2), and 15 min (peak 3),
respectively (Fig. 1A).
Depending on the steepness of the solvent gradient and actual flow
rate, some variations in the retention times were seen (see also Fig.
4). The relative peak intensities, however, were maintained. This at
first glance indicated that the pigments contained in peak 2 were more
abundant than those resolved in peaks 1 and 3 (Fig. 1A). By
contrast, when replicate etioplast samples were extracted with only
80% acetone (21), the level of pigment in peak 3 seemed to largely
exceed those contained in the other fractions (Fig. 1B).
Only traces of peak 2 were seen, and peak 1 remained even below the
level of detection.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
-D-glucoside treatment of
isolated prolamellar bodies was performed as described in Ref. 21.
After solubilization, the assay mixtures were centrifuged, and the
resulting supernatant and membrane fractions were analyzed separately
as specified herein.
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Detection of pigments in etiolated barley
plants. Pigments were extracted from dark-grown barley plants
either with 100% acetone containing 0.1% (v/v) DEP (A) or
an aqueous, non-DEP-supplemented 80% (v/v) solution of acetone
(B). The extracts were separated by HPLC, and porphyrins
were identified by absorbance measurements at 455 nm (see "Materials
and Methods").
Absorbance profiles of the resolved pigments are shown in Fig.
2. They demonstrated that the pigments
contained in peak 2 had a main absorption maximum at 448 nm and two
lower maxima at 578 and 622 nm, respectively (Fig. 2B).
These corresponded to values reported previously for Pchlide
b: the so-called Soret band (448 nm), the Qx
band (578 nm), and the Qy band (622 nm) (21). The
Qx band had a higher absorbance than the Qy
band, which is a typical feature of all investigated pigments of the proto b series (21, 32).
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For the pigments contained in peak 3, a main absorption maximum at 438 nm and a second, lower band at 628 nm were observed (Fig. 2C). These maxima are characteristics of Pchlide a (5, 7). The absorption spectrum of pigments contained in peak 1 was similar to that of the pigments resolved in peak 3; however, the minor peak seemed slightly blue-shifted (Fig. 2A), suggesting the presence of 7-OH-Pchlide a (21).
To prove the identity of the various compounds, synthetic standards were prepared. 7-Hydroxy-Chl a, Chl b, and Chl a were used as educts in a combined enzymatic and chemical procedure (21, 32). In the first step, the phytol chain was removed by the chlorophyllase reaction (40). In the second step, the double bond in ring D of the macrocycle was reestablished by chemical dehydrogenation with 2,3-dichloro-5,6-dicyanobenzoquinone (21, 32).
Fig. 3A shows absorption
spectra of Chl a, Chl b, and 7-hydroxy-Chl
a. It became apparent that the absorption maxima and the shapes of the curves were identical to those known from the literature (e.g. Refs. 24 and 26). After hydrolysis of the pigments by virtue of the chlorophyllase reaction, giving rise to Chlide
a, Chlide b, and 7-hydroxy-Chlide a,
respectively, basically the same spectra were obtained (Fig.
3B), which is in agreement with previous findings that the
phytol chain in the esterified pigments does not affect their
absorption properties as compared with the nonesterified pigments
(e.g. Refs. 21 and 32). Upon oxidation with
2,3-dichloro-5,6-dicyanobenzoquinone, striking changes occurred in the
absorption properties of all three compounds, however. For Chlide
b and Chlide a, spectra were obtained (Fig.
3C) that were indistinguishable from those shown in Fig. 2,
B and C, respectively, indicating the production
of Pchlide b and Pchlide a. The spectrum of
7-hydroxy-Pchlide a was very similar to that of Pchlide
a but was slightly blue-shifted in the red region of the
spectrum (Fig. 3C versus Fig. 2A).
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Mass spectrometry was used to further characterize the chemically prepared and natural compounds. Table I shows that the molecular ion peaks at m/z 612.4 ± 0.2, 626.9 ± 0.3, and 628.4 ± 0.2 were indistinguishable for the natural and synthetic pigments. According to previous work, they correspond to 7-hydroxy-Pchlide a (m/z 628.4), Pchlide b (m/z 626.9), and Pchlide a (m/z 612.4) (21, 32). These results thus ultimately confirmed the presence of 7-OH-Pchlide a, Pchlide b, and Pchlide a in etiolated barley leaves.
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Also with isolated, intact barley etioplasts, 7-OH-Pchlide
a, Pchlide b, and Pchlide a were
readily detectable (Fig. 4A). Careful quantitative pigment measurements showed that Pchlide b (Fig. 4A, peak 2) was
~4-5-fold more abundant in concentration than Pchlide a
(Fig. 4A, peak 3). Again, substantial
amounts of 7-hydroxy-Pchlide a accumulated (Fig.
4A, peak 1).
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The results presented thus far tempted us to conclude that a major part
of Pchlide b originally present in barley etioplasts may be
converted to Pchlide a via 7-hydroxy-Pchlide a.
We hypothesized that this conversion could be due to 7-formyl reductase
activity. To test this idea, conversion of the exogenously administered model substrate ZnPPb was studied in subsequent experiments.
Percoll-purified intact etioplasts were allowed to break during the
incubation with ZnPPb. Before testing the conversion of the
exogenously added ZnPPb, we quantified endogenous pigments
extractable with practically pure, nonaqueous acetone (see above) after
plastid lysis. HPLC analyses revealed that already during plastid
breakage, a massive pigment conversion occurred (Fig. 4B).
Both the relative decrease in Pchlide b and
7-hydroxy-Pchlide a levels and the simultaneous increase in
the amount of Pchlide a are clearly indicative of such a
pigment conversion. This conversion was further pronounced when
reisolated prolamellar bodies obtained from lysed etioplasts were
solubilized with n-octyl--D-glucoside, a
detergent that had frequently been used in previous studies
(e.g. Refs. 21 and 34) (Fig. 4C).
Low temperature in situ fluorescence measurements were
performed at 77 K (18, 39) in order to analyze the functional state of
the different porphyrin pigments. Fig. 5
shows that the intensity of Pchlide F650/657, the predominant
fluorescence peak of intact prolamellar bodies of etioplasts (see
Introduction), was drastically reduced upon etioplast lysis and
subsequent membrane solubilization (Fig. 5, dashed and
dotted lines, respectively, versus
solid line). With
n-octyl--D-glucoside-solubilized membranes,
in fact, no Pchlide F650/657 could be traced. Instead, Pchlide F628/632
became the prevalent spectral pigment species (Fig. 5,
dotted line).
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We next analyzed the effect of externally added ZnPPb. We
assumed that the pigment, if applied in excess, should be able
to compete out Pchlide b to Pchlide a
conversion. Percoll-purified intact etioplasts were supplemented with
ZnPPb and then lysed hypotonically, and the membranes were
sedimented and solubilized with
n-octyl--D-glucoside. Binding of
ZnPPb, as well as that of ZnPPa used as a
control, to the solubilized membranes was tested in two different ways.
We first performed low temperature spectroscopic measurements at 77 K. These showed that almost indistinguishable levels of ZnPPa
and ZnPPb were bound to the membranes (Fig.
6). Interestingly, in neither case was a
long wavelength pigment species restored which emitted at 657 nm
(Pchlide F650/657) (Fig. 6).
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As a second method, we quantified ZnPPa and ZnPPb
binding by HPLC (26). However, also by these pigment measurements, no difference in ZnPPa and ZnPPb binding could be
seen (Fig. 7, Binding, compare
columns 1 and 2).
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Conversion of bound pigments was tested in a subsequent experiment. Detergent-treated membranes were subjected to a prolonged dark incubation, and pigment conversion was analyzed by fluorescence and HPLC measurements. Again, no change could be seen (Fig. 7, Conversion, columns 7 and 8 versus columns 3-6), demonstrating that either pigment was stable and not converted into other compounds. The addition of stromal extract to the ZnPPb-containing assays (Fig. 7, Conversion, columns 7 and 8 versus columns 3-6) and/or NADPH, glucose 6-phosphate, glucose-6-phosphate-dehydrogenase, ferredoxin-NADPH oxidoreductase plus ferredoxin, which had collectively been used to restore Chl(ide) b reductase activity in previous studies (23, 26), proved unsuccessful in our experiments (Fig. 7, columns 9 and 10).
The finding that the
n-octyl--D-glucoside-solubilized membranes
bound approximately the same levels of ZnPPb and
ZnPPa (Figs. 5 and 6) at first glance seemed to contradict
the LHPP model (18). According to this model, at least a 50-fold
difference in ZnPPb binding and a 2-fold difference in
ZnPPa binding should have been seen, reflecting the 5-fold
higher abundance of the PORA as compared with that of the PORB and
their ~10-fold different substrate specificities (18). Whereas PORA
expressed in vitro has been shown to bind 10-fold higher
levels of ZnPPb as compared with ZnPPa, PORB
displayed a 10-fold greater specificity for ZnPPa and bound
only little ZnPPb (18, 31).
An explanation for this apparent paradox could be that the PORA was denatured, was partially degraded, or had become soluble upon etioplast lysis and/or membrane solubilization, including respective centrifugation steps. To follow the fate of the PORA and PORB in the different fractions, we consequently performed Western blot analyses. For comparison, we preflashed isolated, intact etioplasts before analysis and subfractionation with a saturating 1-ms flash of white light, which had previously been used to induce the disintegration of the prolamellar body (13).
Fig. 8A shows that with the
dark-incubated, nonflashed samples, both the PORA and PORB proteins
were detectable in the intact etioplasts and in the respective sediment
fraction obtained after plastid lysis and centrifugation. Upon
solubilization of the sedimented membranes, the picture then changed.
Only the PORB was retained in the sediment fraction, whereas the PORA
was almost quantitatively released into the respective supernatant
(Fig. 8A). With the preflashed sample, this release was
already detectable upon plastid lysis. Then almost all PORA was found
in the supernatant fraction and only traces remained bound to the
resedimented membranes (Fig. 8A). Upon detergent treatment,
this remainder was released into the supernatant obtained after
centrifugation of the assays.
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The various findings reported thus far implied that Pchlide b to Pchlide a conversion may be related to the release of the PORA from the prolamellar body, either artificially as a result of detergent solubilization of the isolated membranes or, more naturally, as part of the light-induced disintegration of these structures. Because the latter process should allow PORA, which is per se a Pchlide-reducing enzyme (10), to regain its activity, we quantified the level of Chlide b and Chlide a as well as Pchlide b and Pchlide a in the various fractions highlighted in Fig. 8A.
Fig. 8B shows a representative HPLC chromatogram. Table II summarizes the results. They confirmed that 7-formyl reductase is present both in the flashed and nonflashed etioplast samples. In the latter, it was rapidly activated upon plastid lysis and membrane solubilization and converted practically all of the preexisting Pchlide b to Pchlide a. In the preflashed sample, 7-formyl reductase was active as well, but it did not seem to gain its full activity. In the supernatant of preflashed, lysed plastids, only one-third of the total pigment was accounted for by Pchlide a. The remainder was present as Pchlide b. In the respective pellet fraction, we recovered ~3-fold lower levels of Pchlide b relative to Pchlide a and found significant levels of Chlide a (Table II).
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Upon membrane solubilization, only Pchlide a could be
detected in the resedimented membranes of the preflashed etioplasts (Table II). Remarkably, this fraction did not contain either Chlide b or Chlide a (Table II); nor were we able to
trace any Pchlide b. In the respective supernatant, the only
detectable pigment was Chlide a (Table II). This suggested
that residual Pchlide b present in the pellet of the lysed
etioplasts had been converted to Pchlide a.
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DISCUSSION |
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In the present study, we readdressed the previously published plastid work-up and pigment extraction procedure of Scheumann et al. (21), which involves hypotonic plastid lysis, the sedimentation of the prolamellar body, and subsequent membrane solubilization. Finally, the solubilized membranes were sedimented, and the obtained pellet and supernatant fractions, respectively, were characterized further.
Our results demonstrate that plastid lysis and membrane solubilization collectively lead to the denaturation of the prolamellar body, the release of the PORA, and the conversion of most, if not all, of the total Pchlide b to Pchlide a. The latter reaction was presumably catalyzed by 7-formyl reductase. This enzyme had originally been implicated in the Chl b to Chl a reaction cycle of chloroplasts during photosynthetic acclimation and leaf senescence (26). But it is also highly active in etioplasts, as shown in this and previous studies (22-26). We assume that 7-formyl reductase may be involved in fine tuning the amounts of Pchlide b and Pchlide a and thereby could regulate the steady-state level of LHPP in etioplasts.
Flash light illumination of intact etioplasts, which has for a long time been known to induce the disintegration of the prolamellar body (13), caused the simultaneous release of the PORA and Pchlide b from the inner plastid membranes. But it did not induce the immediate enzymatic reduction of Pchlide b to Chlide b or the quantitative transformation of Pchlide b to Pchlide a. These important results lend more, although indirect, support to our previous conclusion that the PORA remains, in the first place, catalytically inactive as a Pchlide b-reducing enzyme (18). Moreover, they demonstrate that 7-formyl reductase activity is partially suppressed during the light-induced transformation of etioplasts into chloroplasts. Although we do not yet know the reasons for this effect, we hypothesize that 7-formyl reductase may be involved in controlling the rate of PORA-driven Chlide b synthesis in illuminated plants. It is tempting to speculate that PORA-derived Chlide b could play regulatory roles for the establishment of the light-harvesting structures in etiolated plants at the beginning of illumination, whereas Chl(ide) b synthesized by virtue of Chlide a oxygenase (29, 30, 41) in light-adapted plants could serve housekeeping functions during photosynthesis (for reviews, see Refs. 42 and 43). According to recent work (44), Chlide a oxygenase is well able to accept both Pchlide a and Chlide a as substrate, although with different apparent affinities, but its expression in etiolated, illuminated, and light-adapted plants and its localization have not yet been examined.
In addition to these aspects, the results presented in this study
answer the long lasting question of whether or not Pchlide b
is occurring in etiolated plants (18-21, 45). Our findings show that
Pchlide b is present in barley etioplasts and indeed accounts to amounts well compatible with the LHPP model (18, 19).
According to this model, Pchlide b was supposed to be
~4-5-fold more abundant than Pchlide a. However, given
that Pchlide b is metabolically unstable, the pigment rather
easily escapes the detection. This could explain why previous pigment
extraction procedures failed to detect the pigment (21). Further work
is needed to see whether there is a Pchlide b/Pchlide
a interconversion cycle similar to that reported previously
for Chl a and Chl b (29).
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ACKNOWLEDGEMENTS |
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This work was performed in part in the Department of Prof. Dr. E. W. Weiler at the Institute for Plant Physiology, Ruhr-Universität Bochum, Bochum, Germany. We are grateful to Prof. Weiler for stimulating interest and continuous support of the work. We thank Dr. M. Kuntz (Centre National de la Recherche Scientifique (CNRS), Grenoble, France) for critical reading of the manuscript and help with the HPLC.
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FOOTNOTES |
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* 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: Lehrstuhl
für Pflanzenphysiologie, Universität Bayreuth,
Universitätsstr. 30, 95447 Bayreuth, Germany. Tel.:
49-921-55-26-27; Fax: 49-921-75-77-442; E-mail:
christiane.reinbothe@uni-bayreuth.de.
Published, JBC Papers in Press, October 24, 2002, DOI 10.1074/jbc.M209737200
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ABBREVIATIONS |
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The abbreviations used are: Pchlide, protochlorophyllide; Chlide, chlorophyllide; Chl, chlorophyll; DEP, diethyl pyrocarbonate; HPLC, high performance liquid chromatography; LHPP, light-harvesting POR-Pchlide complex; POR, NADPH:protochlorophyllide oxidoreductase; ZnPP, zinc protopheophorbide..
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REFERENCES |
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