From the Lehrstuhl für Pflanzenphysiologie,
Universität Bayreuth, Universitätsstraße 30, D-95447
Bayreuth, Germany, the ¶ Lehrstuhl für Pflanzenphysiologie,
Ruhr-Universität Bochum, Universitätsstraße 150, D-44801
Bochum, Germany, and the
Université Joseph Fourier et
CNRS, UMR 5575, BP53, CERMO, F-38041 Grenoble cedex 9, France
Received for publication, September 23, 2002, and in revised form, October 15, 2002
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ABSTRACT |
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NADPH:protochlorophyllide oxidoreductase (POR; EC
1.1.33.1) is a key enzyme for the light-induced greening of
angiosperms. In barley, two POR proteins exist, termed PORA and PORB.
These have previously been proposed to form higher molecular weight light-harvesting complexes in the prolamellar body of etioplasts (Reinbothe, C., Lebedev, N., and Reinbothe, S. (1999)
Nature 397, 80-84). Here we report the in
vitro reconstitution of such complexes from chemically
synthesized protochlorophyllides (Pchlides) a and
b and galacto- and sulfolipids. Low temperature (77 K)
fluorescence measurements revealed that the reconstituted,
lipid-containing complex displayed the same characteristics of
photoactive Pchlide 650/657 as the presumed native complex in the
prolamellar body. Moreover, Pchlide F650/657 was converted to
chlorophyllide (Chlide) 684/690 upon illumination of the reconstituted
complex with a 1-ms flash of white light. Identification and
quantification of acetone-extractable pigments revealed that only the
PORB-bound Pchlide a had been photoactive and was converted
to Chlide a, whereas Pchlide b bound to the
PORA remained photoinactive. Nondenaturing PAGE of the reconstituted
Pchlide a/b-containing complex further demonstrated a size similar to that of the presumed native complex in vivo, suggesting that both complexes may be identical.
NADPH:protochlorophyllide oxidoreductase
(POR)1 is a key enzyme for
the light-induced greening of etiolated angiosperm plants. It catalyzes
the only known light-dependent step of chlorophyll biosynthesis, the reduction of protochlorophyllide (Pchlide) to chlorophyllide (Chlide) (1-3). In barley, two POR proteins have been
identified, termed PORA and PORB (4). Both are light- and
NADPH-dependent enzymes, which differ remarkably in their expression patterns during plant development. PORA appears only transiently in dark-grown seedlings, whereas PORB is expressed in
etiolated, illuminated, and light-adapted plants (4). The partial
overlap in expression suggests that both PORA and PORB may be needed
for efficient seedling de-etiolation. We proposed that in the
prolamellar body of etioplasts, the PORA and PORB may cooperate in
terms of a novel "light-harvesting POR-Pchlide a/b" complex termed LHPP (5).
In vitro reconstitution experiments with
synthetic zinc analogs of Pchlide a and Pchlide
b, termed zinc protopheophorbides (ZnPP) a and
b, respectively, indeed supported such a model.
PORA-ZnPPb-NADPH and PORB-ZnPPa-NADPH ternary
complexes were found to form oligomers (5). We observed that light,
which was absorbed by ZnPPb, was transferred onto
ZnPPa (5). This, by virtue of PORB, was reduced to zinc
pheophorbide a (5), the zinc analog of Chlide a
(6). The existence of analogous higher molecular weight light
harvesting structures in vivo was inferred from previously
reported energy transfer reactions, taking place from so-called
photoinactive Pchlide to photoactive Pchlide and from photoinactive
Pchlide to Chlide, in prolamellar bodies before and after flash light illumination (7-12).
Previous critiques questioned the existence of a Pchlide
a/b-containing light-harvesting complex in
vivo, based on the following main arguments (13). First, previous
work seemed to indicate a lack of Pchlide b in etiolated
plants (14). Second, respective in vitro reconstitution
experiments had thus far not been presented for Pchlide a
and Pchlide b, which, according to the LHPP model (5),
should be cognate substrates of the PORB and PORA, respectively. Third,
neither the reconstituted nor the presumed authentic complex had been
resolved under native conditions as higher molecular weight,
lipid-containing structures.
In the present study, we addressed these important questions and
performed in vitro reconstitution experiments. We
demonstrate that the PORA and PORB display the same stringent substrate
specificities for Pchlide a and Pchlide b as
those reported previously for ZnPPa and ZnPPb
(5). We further show that reconstituted PORA-Pchlide b-NADPH
and PORB-Pchlide a-NADPH ternary complexes establish higher molecular mass structures, the spectroscopic and physicochemical properties of which are very close, in most aspects even
indistinguishable, from those of the presumed native complex.
Cloning Procedures--
Double-stranded DNAs encoding the mature
parts of the PORA and PORB proteins of barley were generated by a
polymerase chain reaction-based approach (15). The following primer
pairs were used: Primer 1 (5'-AACTGCAGATGGGCAAGAAGACGCTGCGGCAG-3') plus 2 (5'-AACTGCAGGGTGGATCATAGTCCGACGAGCTT-3'), and primers 3 (5'-AACTGCAGATGGGCAAGAAGACTGTCCGCACG-3') plus 4 (5'-AACTGCAGTGATCATGCGAGCCCGACGAGCTT-3'), as well as cDNA clones A7 (16) and L2 (4), respectively, as templates. After subcloning
into the PstI site of pUC19 (New England Biolabs), the DNAs
for PORA and PORB were cut out with BamHI and
HindIII and inserted into identically treated pSP64 vectors
(Promega) (17). The identity of the different clones was confirmed by DNA sequencing, using a T7 DNA sequencing kit (Promega) and the gel
system described in Ref. 18.
Preparation of Pigments--
Chemical synthesis, purification,
and characterization of zinc- and magnesium-Pchlides a and
b were performed as described in Refs. 6 and 14. HPLC was
carried out either on a C18 reverse phase silica gel column (250 × 4.6 mm, Nucleosil ODS 5 µm; Macherey-Nagel Co.) (6) or a C30
reverse phase column (250 × 4.6 mm, 5 µm; YMC Inc., Wilmington,
NC) (19), using a Varian ProStar model 410 apparatus, a ProStar model
240 pump, and a ProStar 330 photodiode array detector. In some
experiments, a C18 reverse phase silica gel column (250 × 4.6 mm,
Hypersil ODS 5 µm; HyPurityTM), a Dynamax absorbance
detector model UV-1, and a Dynamax SD-200 pump were used.
In Vitro Transcription/Translation and Reconstitution of
POR-Pigment Complexes--
Radiolabeled PORA and PORB molecules were
synthesized by coupled in vitro transcription/translation
(20) of the recombinant clones specified above and purified as
described previously (3). Equal amounts of the PORA and PORB, as
determined by counting their radioactivities and correcting the rates
of incorporation for the different methionine contents (21), were
supplemented with NADPH (0.5 mM final concentration) and
synthetic Pchlide a, Pchlide b, ZnPPa,
or ZnPPb. In all cases, 10 µM final porphyrin concentrations were used. After a 15-min incubation in the dark, the
assay mixtures were subjected to gel filtration on Sephadex G15
equilibrated in assay buffer (22). Enzyme-pigment complexes eluted with
the flow-through were extracted with acetone (see below), and pigments
were quantified in a spectrometer LS50B (PerkinElmer Life Sciences)
(23).
For the reconstitution of LHPP, equimolar amounts of the reconstituted
PORA-Pchlide b-NADPH and PORB-Pchlide a-NADPH
ternary complexes were incubated in the dark, as described previously (5). Then the resulting high molecular weight complexes were separated
from free, nonassembled POR-pigment-NADPH complexes by gel filtration
on Sephadex G100 (5) or Superose 6 (Amersham Biosciences). Fractions
containing PORA-PORB supracomplexes were identified by radioactivity
measurements, pooled and in turn supplemented with a mixture of
galacto- and sulfolipids containing monogalactosyl diacylglycerol,
digalactosyl diacylglycerol, and sulfoquinovosyl diacylglycerol
(58:36:6 mol %; see Ref. 5). The sample was then cooled to Protein Analyses--
Three different methods were employed to
prepare and analyze POR-pigment complexes, that of Ryberg and Sundqvist
(26), that of Klement et al. (27), and a modified version of
that of Gerhardt and Heldt (28).
In the first case, etioplasts were isolated from 5-day-old dark-grown
barley plants by differential centrifugation (for details, see Ref.
26). One aliquot of the final etioplast suspension was extracted with
an excess of 100% acetone containing 0.1% (v/v) diethyl
pyrocarbonate, and protein was recovered by centrifugation and
subsequently used for Western blot analysis (29), using an antiserum
against the PORA of barley (16). In three additional samples, the
plastids were lysed hypotonically in a buffer containing 1 mM MgCl2, 1 mM EDTA, 20 mM TES, 10 mM HEPES, pH 7.2, and etioplast inner membranes comprising prolamellar bodies and prothylakoids prepared by homogenization in a glass homogenizer. All samples were
then centrifuged at 7,700 × g for 15 min. For one
sample, proteins found in the resulting pellet and supernatant,
respectively, were extracted with acetone (see above), sedimented, and
used for subsequent Western blot analysis (see above). In the case of
the second sample, the obtained pellet was resuspended in the buffer
described before but containing 50% sucrose and sonicated three times
for 5 s each with a Branson Sonifier (Danbury, CT) (microtip,
medium tune), and the homogenate was placed at the bottom of a
continuous 10-50% (w/w) sucrose gradient. In case of the third
sample, essentially the same procedure was followed, except for the
fact that the buffer used for ultrasonication lacked sucrose and that
the etioplast inner membranes were loaded from the top onto the gradient.
After centrifugation of the gradients at 25,400 × g
for 2 h, several different bands were seen. For the bottom-loaded
gradient, these corresponded to buoyant densities of ~1.17 g
cm
According to Klement et al. (27), etioplasts were prepared
from etiolated barley plants by Percoll density gradient
centrifugation, retrieved, diluted, and resedimented by centrifugation.
The resulting etioplast pellet was resuspended in a buffer containing
50 mM Tricine/KOH, pH 7.2, 20% glycerol, and four equal
parts were loaded onto sucrose buffer containing 0,5 M
sucrose, 1 mM MgCl2, 1 mM EDTA, 10 mM Tricine, 10 mM HEPES, pH 7.2. After a step
of ultracentrifugation at 80,000 × g for 20 min, the
resulting pellet was resuspended in the same buffer as described before
but containing 2.5 mM
n-octyl-
As the third method, we adapted the nonaqueous fractionation technique
of Gerhardt and Heldt (28) to isolated intact etioplasts. Briefly,
etioplasts were isolated by differential centrifugation and Percoll
density gradient centrifugation and further purified on cushions of
Percoll as described (30). Each plastid sample was divided into two
equal parts, of which one was exposed to a single flash of white light,
whereas the other was kept in darkness. Etioplasts were then
resedimented and immediately quenched and ground under liquid nitrogen.
The etioplast powder was then lyophilized at Electrophoresis--
SDS-PAGE was performed in 10-20% (w/v)
gradients of polyacrylamide as described (31). Nondenaturing,
analytical PAGE was performed in 3-mm-thick 7.5% (w/v) polyacrylamide
gels, and the gels were run using a discontinuous buffer system
(25).
Our previous reconstitution experiments had shown that the PORA
and PORB proteins of barley are able to form higher molecular weight
light-harvesting complexes if complexed with ZnPPb and ZnPPa, respectively, plus NADPH (5). As a first step to
establish such complexes also with their presumed natural substrates,
we synthesized Pchlides a and b chemically (6,
14). The isolated and purified pigments are characterized in an
accompanying paper (32). They were added to PORA and PORB polypeptides,
which had been synthesized from corresponding cDNA clones by
coupled in vitro transcription/translation (20). Pigment
binding was tested in the following manner. Different amounts of the
PORA and PORB were added to an excess of NADPH and isolated pigment,
incubated for 15 min in darkness, and subsequently separated from the
excess of nonbound pigments by gel filtration on Sephadex G15 (22). POR-pigment complexes running in the flow-through were then extracted with 100% acetone containing 0.1% (v/v) diethyl pyrocarbonate, and
pigments were identified and quantified by HPLC and room temperature absorbance and fluorescence measurements.
Fig. 1 shows a plot of the amount of
Pchlide a and Pchlide b bound to the PORA or PORB
versus the enzyme concentrations. From the linear
relationships, it turned out that 1 µg of the PORA bound ~34.11 ng
of Pchlide b and 31.95 ng of ZnPPb but only 3.5 ng of Pchlide a and 3.28 ng of ZnPPa,
respectively. This corresponded to 27.72 pmol of PORA, 54.3 pmol of
Pchlide b (ZnPPb), and 5.7 pmol of Pchlide
a (ZnPPa), respectively, and suggested that there were 1:2 versus 1:0.2 stoichiometries of PORA to pigment in
the recovered PORA-Pchlide (ZnPP) complexes. For the PORB, just the opposite binding preferences were seen: 1 µg (26.26 pmol) of the PORB
bound ~33.2 ng of Pchlide a (31.1 ng of
ZnPPa)(each corresponding to ~54.2 pmol of the pigment)
and only 3.2 ng of Pchlide b (3.02 ng of ZnPPb)
(each corresponding to ~5.1 pmol of the pigment).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
196 °C
and analyzed by fluorescence emission measurements at an excitation
wavelength of 440 nm. For flash light experiments, the sample was
warmed to about
25 °C (24), exposed to a single, 1-ms flash of
white light, and immediately dipped into liquid nitrogen. Then the
spectroscopic measurements were repeated. For the experiment described
in the legend to Fig. 8, two parallel samples were prepared, of which
one was exposed to flash light as above, whereas the other was kept in
darkness before nondenaturing electrophoresis (25).
3 and ~1.21 g cm
3, respectively. In
case of the top-loaded gradient, the lighter band was also obtained,
but not the heavier. Instead a novel, diffuse band spreading over at
least three fractions (designated T1-3 in Fig. 6) was
recovered in the uppermost parts of the gradient. According to Ryberg
and Sundqvist (26), band T1 as well as the band at 1.21 g
cm
3 should represent prothylakoids, whereas the 1.17 g cm
3 band should be identical with prolamellar bodies.
All of the different bands were retrieved from the gradients, diluted
4-fold with the buffer described above, and finally centrifuged at
42,500 × g for 2 h. Proteins found in the
resulting pellet fractions were then extracted with acetone (see above)
and analyzed by Western blotting (29).
-D-glucoside and 5% glycerol. After
gentle shaking for 30 min, the assays were recentrifuged as described
previously, and the obtained pellet was treated with the same detergent
buffer, but containing 15 mM
n-octyl-
-D-glucoside and 30% glycerol. The
suspension was again gently shaken for 30 min and subjected to
ultracentrifugation at 200,000 g for 20 min. The resulting
supernatant was loaded onto a column (9 × 32 mm) of
DEAE-cellulose (Sigma) equilibrated with a buffer containing 5 mM n-octyl-
-D-glucoside, 10%
glycerol, 0.3 mM MgCl2, 0.3 mM
EDTA, 10 mM Tricine, 10 mM HEPES, pH 7.2. Protein was eluted from the column by applying 12 ml of a gradient consisting of 5-20 mM
n-octyl-
-D-glucoside dissolved in
equilibration buffer. Fractions of 0.5 ml were taken and analyzed for
the presence of POR by Western blotting with the PORA antiserum
described before.
50 °C. About 200-300
mg of the dry plastid material were transferred at
35 °C into a
mixture of heptane/carbon tetrachloride
(C7H16/CCl4 66:34 (v/v), density
1.28 g/cm3). The suspension was in turn ultrasonicated at
70 °C with 10 5-s pulses in a Branson Sonifier (see above) and
poured through a layer of quartz wool contained in a filter to remove
any remaining coarse material. The flow-through was diluted 3-fold with
heptane and centrifuged for 2 min at 3000 × g. The
clear supernatant was discarded, and the sediment was resuspended in 3 ml of a CCl4/C7H16 mixture of the
same density as that described before. Two 200-µl aliquots were
withdrawn for determination of Pchlide a and Pchlide b levels by HPLC and enzyme activities (see below). The
remainder was loaded onto a freshly prepared, exponential 1.28-1.50
g/cm3 density gradient of
CCl4/C7H16. After centrifugation at
25,000 × g for 2.5 h, during which time the
material distributed isopycnically in the gradient, 1.2-ml fractions
were removed, starting from the top of the gradient, and subsequently
divided into three equal portions. One-third was used for determination
of marker enzymes (NADPH:glycerinaldehyde-phosphate dehydrogenase,
plastid stroma; phosphoenolpyruvate carboxylase, cytosol;
-mannosidase, vacuole), the second was used for SDS-PAGE, and the
third portion was used for assay of Pchlide a and Pchlide
b levels. All three divided portions and the two aliquots
taken from the original sample (see above) were diluted 3-fold with
heptane and centrifuged for 8 min at 18,000 × g. The
supernatant was discarded, except for the last 200 µl, which were
used to resuspend the sediment by swirling it with a calcined quartz.
All samples were then dried for 18 h in a desiccator and then
processed for electrophoresis, enzyme assays, or pigment measurements.
All manipulations were performed under a dim green light. Moreover, any
step that was expected to potentially lead to trapping of condensing
water vapor was carefully avoided (28).
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Pigment binding characteristics of the PORA
(A) and PORB (B). PORA and PORB
proteins were produced by coupled in vitro
transcription/translation of respective recombinant clones and
purified, and 1, 0.2, 0.1, and 0.02 POR protein equivalents were
subsequently incubated for 15 min in the dark with Pchlide a
( ), Pchlide b (
), ZnPPa
(
), and ZnPPb (
), respectively. After a
step of gel filtration on Sephadex G15, POR-bound pigments were
extracted with a solution of practically pure acetone containing 0.1%
(v/v) diethyl pyrocarbonate. Pigments were identified and quantified by
either HPLC analyses and subsequent absorbance measurements or by
fluorescence spectroscopy, taking into account previously
published absorption and emission coefficients of isolated
pigments (6, 14). The plots show the amounts of PORA-bound and
PORB-bound pigments versus the PORA and PORB protein
concentrations in the assays. 1 POR protein equivalent corresponded to
2.78 pmol of the PORA and 2.64 pmol of the PORB, respectively.
Given that the PORA and PORB displayed the same stringent substrate specificities for Pchlides a and b as those reported previously for ZnPPa and ZnPPb (5), we next established reaction conditions that would allow the generation of higher molecular weight Pchlide a/b-POR light-harvesting complexes. Equimolar amounts of reconstituted PORA-Pchlide b-NADPH and PORB-Pchlide a-NADPH ternary complexes were mixed and, after a 15-min incubation in darkness, subjected to a further step of gel filtration on Sephadex G100 (5) or size-fractionated on a Superose 6 column (see "Materials and Methods").
Fig. 2A shows a size
fractionation on Superose 6. It revealed that the pigment-complexed
PORA and PORB indeed gave rise to a higher molecular weight complex.
Its size of ~480 kDa was similar to that of the so-called Pchlide
holochrome of bean (33). Free, nonassembled PORA- and PORB-pigment
ternary complexes were eluted at much later time points (Fig.
2A).
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To determine the stoichiometry of the PORA and PORB in the recovered supracomplexes, fractions containing the different PORs were pooled (see Fig. 2A) and separated by SDS-PAGE, and the gel was subjected to autoradiography. This experiment revealed that in the recovered higher molecular weight complex about five PORA-Pchlide b-NADPH ternary complexes interacted with just one PORB-Pchlide a-NADPH complex (Fig. 2B, fraction 5).
The oligomeric PORA-PORB protein complex contained in fraction 5 was subsequently supplemented with a lipid mixture containing monogalactosyl diacylglycerol, digalactosyl diacylglycerol, and sulfoquinovosyl diacylglycerol (58:36:6 mol %), which had been prepared from pigment-free prolamellar bodies of barley etioplasts (5). Then low temperature measurements were performed at 77 K, in a PerkinElmer Life Sciences spectrometer LS50B (23).
Fig. 2C (solid line) demonstrates that two fluorescence peaks could be seen at an excitation wavelength of 440 nm: one at 657 nm and the other at 632 nm. Because these two peaks corresponded to photoactive Pchlide 650/657 and photoinactive Pchlide F628/632, known from the prolamellar body of etioplasts (e.g. Ref. 5), we exposed the lipid-containing complex to a saturating 1-ms flash of white light. As shown in Fig. 2C (dashed line), this gave rise to the quantitative conversion of Pchlide F650/657 to Chlide 684/690.
We next extracted pigments from the flashed sample with acetone (see
above) and ran HPLC analyses, as described in Ref. 14. Separation on a
C18 column is shown in Fig.
3A. As demonstrated in an
accompanying paper (32), the pigment peak eluting at ~12.5 min is
identical with Pchlide b, whereas the peak eluting at ~15 min is identical with Pchlide a (Fig. 3A,
panel a). Upon flash light illumination, a novel
peak appeared at ~14 min (Fig. 3A, panel
b). Based on our previous in vitro reconstitution
experiments, we assumed that this peak might be due to Chlide
a. To demonstrate this, another type of HPLC analysis was
performed. Taking into account a recent paper of Fraser et
al. (19), a C30 column was used. Synthetic Chlides a
and b were prepared by the chlorophyllase reaction (34) and
used as standards. Fig. 3B (panel c)
shows that Chlide a and Chlide b were well
resolved on the C30 column and also separated from Chl a and
Chl b. When the pigments, eluting at ~14 min on the C18
column (see Fig. 3A), were applied to the C30 column, the
only detectable pigment was Chlide a (Fig. 3B, panel d). Thus, only Chlide a had been
produced upon flash light illumination of the reconstituted complex
(Table I). By contrast, Pchlide
b, which was ~5-fold more abundant than Pchlide
a, did not seem to be photoconvertible at all under the
tested conditions. Indeed, no Chlide b was formed (Table
I).
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An explanation for these findings could be that the PORA was per
se inactive with Pchlide b and thus unable to convert
the pigment to Chlide b. If so, no Chlide b
should be produced also in situ. To test this hypothesis, we
first analyzed pigments that were formed in isolated prolamellar bodies
upon flash light illumination by HPLC. Fig.
4 shows a representative separation of
pigments before (A) and after (B) a saturating
1-ms flash of white light. In addition to Pchlide b and
Pchlide a, eluting at 12.5 and 15 min, respectively (Fig.
4A), 7-hydroxy-Pchlide a could also be detected
(peak 1) (for details, see Ref. 32). Upon
flashing the sample, a novel pigment peak appeared (Fig. 4B,
peak 4), the retention time and absorption
properties of which were indistinguishable from those of Chlide
a identified previously (data not shown, but see Fig. 3). At
the same time, increasing amounts of Pchlide a
(peak 3) were detectable. As shown in an
accompanying paper (32), barley etioplasts contain an enzyme called
7-formyl reductase that converts Pchlide b to Pchlide
a via 7-hydroxy-Pchlide a (see also Ref. 14).
Upon resolution of pigments contained in peak 4 by subsequent HPLC on a C30 column, indeed only Chlide a and no Chlide b was observed (data not shown).
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As a second approach, the in vitro synthesized PORA was incubated with ZnPPb (which is chemically more inert than Pchlide b) and separated from nonbound pigment by gel filtration, and POR-pigment complexes eluted with the flow-through exposed to white light. Parallel samples were kept in darkness. As a control, the PORB was used.
Fig. 5 shows representative room
temperature fluorescence emission spectra of PORA- and PORB-bound
pigments after their extraction with acetone. They revealed that the
PORA indeed converted ZnPPb to zinc pheophorbide
b (Fig. 5B). By analogy, also ZnPPa was converted to zinc pheophorbide a (Fig. 5A). Also with the
PORB, the same principal results were obtained (Fig. 5, C
and D, respectively). For either POR protein, a strict
correlation was observed between the amounts of products formed and
bound substrates, regardless of whether ZnPPa and
b2 or Pchlides a and b had
been used (Fig. 5).
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All results presented thus far implied that the reconstituted, lipid-containing Pchlide a/b-POR protein complex may be identical with the presumed native complex in the prolamellar body of etioplasts. However, except for the spectroscopic data, no other line of evidence seemed to exist to support this notion. We consequently sought to identify the native complex by classical biochemical approaches and to compare its properties with those of the reconstituted complex.
Ryberg and Sundqvist (26) had shown that isolated etioplast inner
membranes from wheat can be resolved into different subfractions, designated prolamellar bodies and prothylakoids, respectively, based on
their different buoyant densities in sucrose gradients. We readdressed
this previous work for barley etioplasts and analyzed the abundance of
the PORA and PORB after various steps of the plastid work-up procedure
(see "Materials and Methods"). As shown in Fig.
6, already during the very first step of
isolation of the so-called etioplast inner membranes (presumed to
comprise prolamellar bodies and prothylakoids) (26), a major part of the PORA became soluble. The same effect was seen for oat and wheat
etioplasts, which were analyzed in parallel (data not given). In all
cases, subsequent steps of prolamellar body and prothylakoid separation
turned out to correlate with a further solubilization of the PORA. This
is shown for barley in Fig. 6. With etioplast inner membranes, which
had been diluted with a buffer lacking any additives for membrane
stabilization, the PORA was quantitatively released and then
co-migrated in the uppermost fractions of a top-loaded 10-50% (w/w)
sucrose density gradient. With barley etioplast inner membranes, which
had been sonicated in a buffer containing 50% sucrose and loaded from
the bottom, both the PORA and PORB appeared to be recovered in the
lower parts of the gradient, containing prolamellar bodies. However,
the approximately equimolar amounts of the PORA and PORB, which were at
variance with their original stoichiometries in intact etioplasts, and
the fact that both POR proteins migrated to slightly different
positions in the gradient argued against working further with these
samples.
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In a recent paper, Klement et al. (27) reported the isolation of a pigment-free POR protein from oat. Such a preparation seemed particularly interesting to us because it would allow testing of the pigment binding properties of the PORA and PORB in a more natural environment than in the in vitro system. In principal, the method of Klement et al. (27) employs differential detergent solubilization of POR from the prolamellar body.
When we reproduced the published protocol and followed the fate of the
PORA and PORB by Western blotting (Fig.
7), again drastic losses of the PORA
during successive steps of the membrane preparation and solubilization
procedure became apparent. In the final supernatant, approximately
equal levels of the PORA and PORB were seen, which were at variance
with the determined 5:1 protein stoichiometry in intact etioplasts.
Moreover, subsequent chromatography on DEAE-cellulose gave rise to at
least three different POR protein bands, representing the PORA, the
PORB, and a slightly smaller degradation product (Fig. 7). To what
extent these proteins would contribute to pigment binding could not be
estimated.
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Given this uncertainty and the fact that neither the method of Ryberg and Sundqvist (26) nor that of Klement et al. (27) allowed the recovery of an intact PORA-PORB protein complex, we sought alternative methods. Taking into account a paper of Gerhardt and Heldt (28) on enzyme and metabolite measurements in different subcellular compartments, we adapted nonaqueous protein and pigment extraction and fractionation to isolated etioplasts. The method employed is based on the fact that the proteins and metabolites of a given compartment aggregate together upon lyophilization. Because each compartment has a characteristic protein, lipid, carbohydrate, and ion complement, different fractions are obtained in nonaqueous gradients of carbon tetrachloride/heptane. These and an original etioplast sample were analyzed with respect to the PORA and PORB protein abundances as well as Pchlide a and Pchlide b levels. Moreover, we used nondenaturing, analytical PAGE (25) to directly visualize POR-pigment complexes in etioplasts prior to fractionation.
Table II shows the determined PORA and PORB as well as Pchlide a and Pchlide b levels. It turned out that PORA is ~4.2-fold more abundant in amount than PORB. Quantification of pigments showed that etioplasts contain an ~4.5-fold excess of Pchlide b relative to Pchlide a. Of the total Pchlide, only 18% was photoreducible. This photoconvertible Pchlide turned out to be identical with Pchlide a (Table II).
|
Fig. 8A (panel
a, lane D) shows a nondenaturing,
analytical PAGE of the presumed natural POR-pigment complex. Based on
the red light-induced autofluorescence of Pchlide F650/657, this
complex could directly be visualized by fluorography. Western blot
analyses confirmed that it contained POR (Fig. 8A,
panel b, lane D). Flash light illumination and subsequent mild detergent treatment in the
presence of 0.2% (v/v) Triton X-100 prior to electrophoresis dissociated this total POR into two subfractions (Fig. 8A,
FL). Upon scaling up the procedure 1000-fold, these could be
identified as PORA and PORB by protein sequencing (data not shown).
Their approximate stoichiometry was similar to that determined from the
carbon tetrachloride/heptane gradients (Table II) and also matched that
expected from our previous in vitro reconstitution experiments (Fig. 2). Indeed, when the in vitro
reconstituted, lipid-containing complex was subjected to nondenaturing
PAGE, a similar, although slightly smaller, complex could be seen (Fig. 8B). This complex contained both PORA and PORB and displayed
the same type of autofluorescence as the presumed native complex. Moreover, it was rapidly dissociated into the two POR proteins upon
flash light illumination.
|
An HPLC analyses of pigments reextracted from the electrophoretically
resolved native POR-pigment complex is shown in Fig. 9. It demonstrated that the complex
contained both Pchlide b and Pchlide a (Fig.
9A, peak 2 and 3,
respectively). In addition, substantial amounts of 7-hydroxy-Pchlide
a could also be seen (Fig. 9A, peak
1). Upon flash light illumination, correlating with the
disintegration of the complex (see Fig. 8A), only Chlide a was produced (Fig. 9B, peak
4). It co-migrated with the PORB protein band (Fig.
9D). Protochlorophyllide b, by contrast, remained quantitatively unchanged (Fig. 9, compare B
versus A) and co-migrated with the PORA protein
band (Fig. 9C). This result not only confirmed the
previously determined substrate specificities but also that only
PORB's bound pigment (i.e. Pchlide a) had been
converted to Chlide a. The bulk of the pigment,
corresponding to Pchlide b, remained
nonphotoconvertible.
|
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DISCUSSION |
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In the present study, we addressed three different questions. First, would the PORA and PORB display the same stringent substrate specificities for their presumed natural substrates (Pchlide b and Pchlide a, respectively) as reported previously for their zinc counterparts (5)? Second, would the resulting PORA-Pchlide b-NADPH and PORB-Pchlide a-NADPH ternary complexes be able to establish higher molecular weight light-harvesting structures with galacto- and sulfolipids, as proposed previously (5)? Third, would similar, Pchlide b-containing complexes exist in vivo?
The answers to all of these questions were positive. We were able to demonstrate that PORA binds ~10-fold higher amounts of Pchlide b (ZnPPb) relative to Pchlide a (ZnPPa). PORB, by contrast, was specific for Pchlide a (ZnPPa) and bound ~10-fold lower levels of Pchlide b (ZnPPb). Either POR protein likewise converted these compounds into their respective products in vitro. However, if PORA-Pchlide b-NADPH and PORB-Pchlide a-NADPH ternary complexes were mixed and reconstituted to higher molecular weight complexes, only the PORB remained active. In the presence of galacto- and sulfolipids, the reconstituted Pchlide a/b-POR complex displayed the features of Pchlide F650/657. This Pchlide F650/657 was converted to Chlide F684/690 upon flash light illumination. Indistinguishable spectral pigment species and pigment conversions have been described for isolated prolamellar body membranes of etioplasts (see Introduction). Moreover, we were able to resolve the lipid-containing structure both from the prolamellar body and after in vitro reconstitution into similar higher molecular weight complexes under native conditions. Based on all of these findings, we conclude that the reconstituted and analyzed authentic complexes may be structurally and functionally identical.
How is LHPP made in vivo? A key aspect related to this question refers to the origin of Pchlide b. The existence of this pigment has long been a matter of dispute (see literature cited in Ref. 14). As shown in this and an accompanying paper (32), the pigment is present in etiolated barley plants but is rapidly converted to Pchlide a if no precautions are taken. 7-Formyl reductase presumably responsible for this conversion is highly active in barley etioplasts. It was for a long time implicated in Chl b to Chl a conversion, but it appears that 7-formyl reductase plays a more general role in fine tuning the levels of both porphyrins and chlorins in dark-grown and illuminated plants (14, 32).
Enzymes, which may synthesize Pchlide b, have not been identified. The most likely candidates are proteins that could display (P)Chlide a oxygenase activity. Previous work has shown that there is a family of related proteins, which may exhibit such an activity (35-41). Tanaka et al. (36) cloned a Chlamydomonas reinhardtii cDNA for a putative Chlide a oxygenase. Later studies by Espineda et al. (37) and Tomitani et al. (38) showed that highly related Chlide a oxygenase sequences also occur in Arabidopsis thaliana and other plant species. The Arabidopsis protein was expressed in bacteria and suggested to display Chlide a, but not Pchlide a, oxygenase activity (39). Recent work by Xu et al. (40, 41), however, highlighted that heterologous expression of the Arabidopsis Chlide a oxygenase in cyanobacteria leads to Pchl(ide) b and Chl(ide) b accumulation. Although this demonstrates that Chlide a oxygenase is well able to bind and convert Pchlide a to Pchlide b, it is not yet known whether Chlide a oxygenase is expressed in etiolated plants. If this does not occur, another enzyme should exist that drives Pchlide b synthesis.
Recent, yet unpublished work for barley shows that there is indeed a
protein related to Chlide a oxygenase (for sequence
comparisons, see Refs. 35 and 36), which is able to convert Pchlide
a to Pchlide b. It is part of the
substrate-dependent import machinery in the plastid
envelope through which the cytosolic precursor of the PORA is imported
into the organelle (21, 22, 30). We were able to demonstrate that
Pchlide a, formed upon feeding isolated plastids the Pchlide
precursor 5-aminolevulinic acid, is converted to Pchlide b.
Concomitantly, the envelope-bound PORA precursor was chased into the
plastids and processed to mature size. These findings imply that
Pchlide b synthesis is directly coupled to the import step
and that the novel Pchlide a oxygenase is located in the
plastid envelope. Consistent with such an idea are also previous
findings that isolated envelope membranes of spinach chloroplasts
contain Pchlide (42, 43). Work is in progress to further characterize
the novel Pchlide a oxygenase.
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ACKNOWLEDGEMENTS |
---|
This work was inaugurated in the Department of Prof. Dr. K. Apel (Institute for Plant Sciences, Swiss Federal Institute of Technology, Zurich, Switzerland); pursued in the laboratory of Prof. Dr. R. Mache (Université Joseph Fourier and CNRS, Grenoble, France); and completed in the Department of Prof. Dr. E. W. Weiler (Institute for Plant Physiology, Ruhr-Universität Bochum, Bochum, Germany). We are grateful to K. Apel for allowing some of the initial experiments to be performed in his laboratory and to R. Mache and E. Weiler for stimulating interest and continuous support of the work. We thank Dr. M. Kuntz (CNRS, Grenoble, France) for expert help with the HPLC as well as critical reading of the manuscript.
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FOOTNOTES |
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* This work was supported by Deutsche Forschungsgemeinschaft Grant RE1465/1-1,1-2 (to C. R.).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.M209738200
2 C. Reinbothe, F. Buhr, S. Pollmann, and S. Reinbothe, unpublished data.
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ABBREVIATIONS |
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The abbreviations used are: POR, NADPH:protochlorophyllide oxidoreductase; Chl, chlorophyll; Chlide, chlorophyllide; HPLC, high performance liquid chromatography; LHPP, light-harvesting POR-Pchlide complex; Pchlide, protochlorophyllide; ZnPP, zinc protopheophorbide; TES, N-tris(hydroxymethyl-2-aminomethanesulfonic acid; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Griffiths, W. T. (1975) Biochem. J. 152, 623-635[Medline] [Order article via Infotrieve] |
2. | Griffiths, W. T. (1978) Biochem. J. 174, 681-692[Medline] [Order article via Infotrieve] |
3. | Apel, K., Santel, H.-J., Redlinger, T. E., and Falk, H. (1980) Eur. J. Biochem. 111, 251-258[Abstract] |
4. | Holtorf, H., Reinbothe, S., Reinbothe, C., Bereza, B., and Apel, K. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 3254-3258[Abstract] |
5. | Reinbothe, C., Lebedev, N., and Reinbothe, S. (1999) Nature 397, 80-84[CrossRef] |
6. | Schoch, S., Helfrich, M., Wiktorsson, B., Sundqvist, C., Rüdiger, W., and Ryberg, M. (1995) Eur. J. Biochem. 229, 291-298[Abstract] |
7. | Smith, J. H. C., and Benitez, A. (1954) Plant Physiol. 29, 135-143 |
8. | Kahn, A., Boardman, N. K., and Thorne, S. W. (1970) J. Mol. Biol. 48, 85-101[Medline] [Order article via Infotrieve] |
9. | Mathis, P., and Sauer, K. (1972) Biochim. Biophys. Acta 267, 498-511[Medline] [Order article via Infotrieve] |
10. | Vaughan, G. D., and Sauer, K. (1974) Biochim. Biophys. Acta 347, 383-394[Medline] [Order article via Infotrieve] |
11. | Ignatov, N. V., and Litvin, F. F. (1981) Biofizika 26, 664-668[Medline] [Order article via Infotrieve] |
12. | Fradkin, L. I., Domanskaya, I. N., Radyuk, M. S., Domanskii, V. P., and Kolyago, V. M. (1993) Photosynthetica 29, 227-234 |
13. | Armstrong, G. A., Apel, K., and Rüdiger, W. (2000) Trends Plant Sci. 5, 40-44[CrossRef][Medline] [Order article via Infotrieve] |
14. | Scheumann, V., Klement, H., Helfrich, M., Oster, U., Schoch, S., and Rüdiger, W. (1999) FEBS Lett. 445, 445-448[CrossRef][Medline] [Order article via Infotrieve] |
15. | Innis, M. A., Gelfand, D. H., Sninsky, J. J., and White, T. J. (1990) PCR Protocols , Academic Press, Inc., San Diego, CA |
16. | Schulz, R., Steinmüller, K., Klaas, M., Forreiter, C., Rasmussen, S., Hiller, C., and Apel, K. (1989) Mol. Gen. Genet. 217, 355-361[Medline] [Order article via Infotrieve] |
17. | Maniatis, T., Fritsch, E. F., and Sambrook, J. (1982) Molecular Cloning: A Laboratory Manual , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY |
18. | Sanger, F., Nickler, S., and Coulsen, A. R. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 5463-5467[Abstract] |
19. | Fraser, P. D., Pinto, E. M. S., Holloway, D. E., and Bramley, P. M. (2000) Plant J. 24, 551-555[CrossRef][Medline] [Order article via Infotrieve] |
20. | Krieg, P. A., and Melton, D. A. (1984) Nucleic Acids Res. 12, 7057-7070[Abstract] |
21. |
Reinbothe, C.,
Lebedev, N.,
Apel, K.,
and Reinbothe, S.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
8890-8894 |
22. | Reinbothe, S., Reinbothe, C., Runge, S., and Apel, K. (1995) J. Cell Biol. 129, 299-308[Abstract] |
23. |
Lebedev, N.,
van Cleve, B.,
Armstrong, G. A.,
and Apel, K.
(1995)
Plant Cell
7,
2081-2090 |
24. |
Lebedev, N.,
and Timko, M. P.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
9954-9959 |
25. | Reinbothe, S., Nelles, A., and Parthier, B. (1991) Eur. J. Biochem. 198, 365-373[Abstract] |
26. | Ryberg, M., and Sundqvist, C. (1982) Physiol. Plant. 56, 125-132 |
27. |
Klement, H.,
Helfrich, M.,
Oster, U.,
Schoch, S.,
and Rüdiger, W.
(1999)
Eur. J. Biochem.
265,
862-874 |
28. | Gerhardt, R., and Heldt, H. W. (1984) Plant Physiol. 75, 542-547 |
29. | Towbin, H., Staehlin, T., and Gordon, J. (1979) Proc. Natl. Acad. Sci. U. S. A. 76, 4350-4354[Abstract] |
30. |
Reinbothe, S.,
Runge, S.,
Reinbothe, C.,
van Cleve, B.,
and Apel, K.
(1995)
Plant Cell
7,
161-172 |
31. | Scharf, K.-D., and Nover, L. (1982) Cell 30, 427-437[Medline] [Order article via Infotrieve] |
32. | Reinbothe, S., Pollmann, S. & Reinbothe, C. (2003) J. Biol. Chem. 800-806 |
33. | Boardman, N. K. (1962) Biochim. Biophys. Acta 62, 63-79[CrossRef][Medline] [Order article via Infotrieve] |
34. | McFeeters, R. F., Chichester, C. O., and Whitaker, J. R. (1971) Plant Physiol. 47, 609-618 |
35. |
Caliebe, A.,
Grimm, R.,
Kaiser, G.,
Lübeck, J.,
Soll, J.,
and Heins, L.
(1997)
EMBO J.
16,
7342-7350 |
36. |
Tanaka, A.,
Ito, H.,
Tanaka, R.,
Tanaka, N. K.,
Yoshida, K.,
and Okada, K.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
12719-12723 |
37. |
Espineda, C. E.,
Linford, A. S.,
and Devine, D. A.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
10507-10511 |
38. | Tomitani, A., Okada, K., Miyashita, H., Matthijs, C. P., Ohno, T., and Tanaka, A. (1999) Nature 400, 159-162[CrossRef][Medline] [Order article via Infotrieve] |
39. | Oster, U., Tanaka, R., Tanaka, A., and Rüdiger, W. (2000) Plant J. 21, 305-310[CrossRef][Medline] [Order article via Infotrieve] |
40. |
Xu, H.,
Vavilin, D.,
and Vermaas, W.
(2002)
J. Biol. Chem.
277,
42726-42732 |
41. |
Xu, H.,
Vavilin, D.,
and Vermaas, W.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
14168-14173 |
42. |
Joyard, J.,
Block, M.,
Pinaeu, B.,
Albrieux, C.,
and Douce, R.
(1990)
J. Biol. Chem.
265,
21820-21827 |
43. |
Pineau, B.,
Dubertret, G.,
Joyard, J.,
and Douce, R.
(1986)
J. Biol. Chem.
261,
9210-9215 |