Photosynthesis Research 64: 63–76, 2000. © 2000 Kluwer Academic Publishers. Printed in the Netherlands.
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Regular paper
Localization of NADPH-protochlorophyllide reductase in plastids of barley at different greening stages Xavier Barth´elemy1 , Gw´enaëlle Bouvier1 , Alfons Radunz2 , Sarah Docquier3 , Georg H. Schmid2 & Fabrice Franck1 1 Laboratory
of Photobiology, Institute of Plant Biology B22, University of Li´ege, Sart-Tilman, B-4000 Li´ege, Belgium; 2 Lehrstuhl Zellphysiologie, Fakultät für Biologie, Universität Bielefeld, D-4800 Bielefeld 1, Germany; 3 Laboratory of Plant Morphology, Institute of Plant Biology B22, University of Li´ ege, Sart-Tilman, B-4000 Li´ege, Belgium; ∗ Author for correspondence (e-mail:
[email protected]) Received 14 September 1999; accepted in revised form 4 April 2000
Key words: chlorophyll synthesis, chloroplast, envelope membrane, greening, NADPH-protochlorophyllide oxidoreductase, protochlorophyllide, thylakoid
Abstract The localization of protochorophyllide (Pchlide) and of NADPH-protochlorophyllide oxidoreductase (POR, EC 1.6.99.1) within (etio)chloroplasts has been investigated at selected stages of greening of barley seedlings. Pchlide pigment and POR protein contents were evaluated in different plastid membrane fractions by fluorescence spectroscopy and immunoblot analysis using a monospecific polyclonal antibody raised against the purified enzyme. Fluorescence analysis showed the presence of Pchlide in both the envelope and thylakoid membranes. During greening, the Pchlide content, expressed on a total protein basis, decreased in thylakoid membranes, whereas it increased in the envelope membranes. POR proteins were detected mainly in thylakoid membranes at early greening stages. In contrast, the weak amount of POR proteins was associated more specifically with envelope membranes of mature chloroplasts. Whatever the greening stage, thylakoid-bound Pchlide and POR proteins were more abundant in the thylakoid regions which remained unsolubilized after mild Triton treatment used as standard procedure to prepare PS II particles. This suggests the preferential association of Pchlide and POR to the appressed regions of thylakoids. Abbreviations: Chl – chlorophyll; Chlide – chlorophyllide a; EDTA – ethylen diamine tetraacetic acid; HEPES – 4-(2-hydroxyethyl)-1-piperazinethanesulfonic acid; Pchlide – protochlorophyllide a; POR – NADPHprotochlorophyllide oxidoreductase; PS II – Photosystem II; SDS – sodium dodecyl sulfate Introduction Meristematic cells of higher plant seedlings contain small organelles, bounded by a double membrane envelope, from which all other plastids originate. The proplastids are precursors of chloroplasts that develop under normal daylight (Mullet 1988). In complete darkness, proplastids develop into etioplasts. These organelles lack Chl and Chl-binding proteins. Their internal membrane system is composed of networks of tubular membranes, called prolamellar bodies, and
prothylakoids. When etioplasts are exposed to light, drastic changes occur in their structural and functional organization, including the breakdown of prolamellar bodies and the development of photosynthetically active thylakoid membranes typical of chloroplasts (Kahn 1968; Henningsen and Boynton 1974; Robertson and Laetsch 1974; reviewed by Ryberg et al. 1993). NADPH-protochlorophyllide oxidoreductase (POR; EC1.6.99.1) catalyzes the only strictly light-dependent step in Chl synthesis of higher plants: The reduction of protochlorophyllide a (Pchlide) to chlorophyl-
64 lide a (Chlide) (Griffiths 1978, reviewed by Fujita 1996; Lebedev and Timko 1998; Beale 1999). Phototransformable NADPH–POR–Pchlide complexes accumulate in internal membranes of etioplasts during growth in darkness. Their concentration rapidly decreases upon exposure to light. When Chl accumulation reaches its maximum rate during illumination of dark-grown seedlings, the enzyme activity drops to low levels and only weak amounts of POR proteins are detected (Kay and Griffiths 1983). However, small amounts of NADPH–POR–Pchlide complexes are continuously regenerated and phototransformed in the light, which explains the fast Chl accumulation (Griffiths et al. 1985; Franck and Strzalka 1992). In barley and in Arabidopsis, two distinct isoenzymes of POR have been identified (Holtorf et al. 1995; Armstrong et al. 1995). Although the activities of the two enzymes are similar, their expression during chloroplast differenciation upon illumination of etiolated seedling is distinct. In contrast to the light-regulated POR enzyme studied so far (PORA), the second POR protein (PORB) is constitutively expressed and becomes the only POR protein after a few hours of greening (reviewed by Reinbothe et al. 1996). POR are membrane-bound proteins. In etioplasts of dark-grown plants, most of the POR proteins have been found in the prolamellar body (Ryberg and Sundqvist 1982; Shaw et al. 1985) and POR appeared as a critical determinant of prolamellar body development in darkness (Sperling et al. 1998). On the basis of immunogold labeling results, Ryberg and Dehesh (1986) suggested that the phototransformation of Pchlide to Chlide initiates a translocation of POR from the disrupted prolamellar bodies to the prothylakoids. In mature spinach chloroplasts, Pchlide, Chlide pigments and POR-immunoreactive protein have been reported on the outer surface of the outer envelope membrane (Pineau et al. 1986; Joyard et al. 1990). Photoreduction of Pchlide has been also shown on that membrane (Pineau et al. 1993). The later steps of Chl synthesis, such as esterification of Chlide a with phytol or earlier alcohol precursors, are supposed to take place in the thylakoid membranes (Block et al. 1980). Some results are not fully consistent with a restricted localization of POR to the chloroplast envelope. Fradkin et al. (1981a) detected Pchlide in thylakoid sub-membrane particles obtained from digitonintreated mature barley chloroplasts. Moreover, excitation energy transfer from Pchlide to Chl was demonstrated in mature chloroplasts (Fradkin et al. 1981b), implying a close spatial relationship between these
two pigments. The efficiency of this energy transfer has been found to increase during continuous illumination of dark-grown barley seedlings (Shlyk et al. 1984). These findings suggest that in chloroplasts POR is not only associated to the envelope membrane, but also to thylakoids. Due to their low concentration in mature leaves, the distribution of POR proteins in plastid membranes is difficult to detect by immunogold labelling in ultrathin sections. Previous studies using this method have, however, indicated that low amounts of POR were associated with thylakoids of mature barley chloroplasts (Dehesh et al. 1987). In redarkened plastids of greening pea leaves, Pchlide pigment was detected by fluorescence spectrsocopy and found to be mainly associated to a polyribosomerich fraction of primary thylakoids (Lebedev et al. 1990). Import studies have also shown that POR proteins can be directed to the thylakoids of chloroplasts (reviewed in Lebedev and Timko 1998). When dark-grown seedlings are illuminated, Chl is synthesized at high rates during the first hours of illumination. Therefore, greening plants provide a suitable experimental system to investigate POR localization in relation with Chl synthesis. In the present study, we have isolated different membrane systems of plastids at selected greening stages and we have investigated the distribution of POR proteins and of Pchlide pigments among envelope membranes and in different thylakoid fractions obtained upon partial solubilization of the purified thylakoids by the detergent Triton X100.
Materials and methods Plant material Seedlings of barley (Hordeum vulgare cv. Pavilion) were grown in darkness on vermiculite and tap water at 23 ◦ C for 7 days. Continuous greening of barley was performed under fluorescent tubes at 15 W m−2 . Green control leaves were grown under the same light intensity and temperature for 7 days with 14–10 h light-dark cycles. Transmission electron microscopy After 7 days of growing, the first leaves of the seedlings were 9–10 cm long. A 5 mm section was excised at distances of 2 and 4 cm from the top of five leaves at different greening stages. The different pieces of leaves were fixed in 2% potassium permanganate in
65 200 mM sodium cacodylate buffer (pH 7.4) for 45 min at room temperature. After dehydratation in a graded alcohol series, the pieces of leaves were embedded in Epon. Ultrathin transversal sections (approximately 80–90 nm, pale silver interference color) were made in each sample with a Sorvall MT-2 ultra-microtome using a Diatome diamond knife. The sections were collected on 200 mesh copper grids and subsequently contrasted for 3 min with 4.6% uranyl acetate solution in 50% ethanol. The sections were finally examined with a Zeiss EM 900 electron microscope at 80KV. Isolation of purified envelope and thylakoid membranes from (etio)chloroplasts Leaves (40 g) were mixed in a Multimix mixer during 10 s with 250 ml of 25 mM HEPES–NaOH buffer (pH 7.5) containing 0.4 M sucrose, 5 mM NaCl, 5 mM MgCl2 and 1 mM EDTA. The homogenate was filtered through four layers of cheesecloth and one layer of nylon mesh (32 µm pore size). To remove cell debris, the filtrate was centrifuged at 500 g for 10 min. The supernatant was then centrifuged at 2000 g for 15 min. The pellet was resuspended in the same buffer and the (etio)chloroplasts were purified by discontinuous sucrose density gradient centrifugation according to Ikeuchi and Murakami (1982). Osmotic lysis of these intact plastids was done by suspending them for few min at 0 ◦ C in hypotonic 25 mM HEPES–NaOH buffer (pH 7.5) containing 5 mM NaCl, 5 mM MgCl2 and 1 mM EDTA. This step was followed by sedimentation in a discontinuous sucrose gradient as described by Douce et al. (1973) leading to the simultaneous recovery of the two chloroplast membrane fractions, envelope and thylakoids, but also the soluble fraction, the stroma. Figure 1 shows the successive steps of this procedure and the subsequent separation of PS II-enriched membranes. Preparation of PS II enriched membranes O2 -evolving Triton PS II particles were isolated according to Berthold et al. (1981) with some modifications.The thylakoid membranes isolated by the method of Douce et al. (1973) were washed with 40 mM Mes– NaOH buffer (pH 6.0), containing 0.4 M sucrose, 15 mM NaCl, 5 mM MgCl2 , 1 mM MnCl2 and 1 mM EDTA. They were resuspended at a final concentration of 2 mg per ml and treated with Triton X-100 (15 mg Triton X100 per mg Chl, during 15 min for the green leaves extracts; or 5 mg Triton X100 per mg Chl, during 10–15 min for the greening leaves extracts).
After incubation, the suspension was centrifuged for 30 min at 35 000 g. As judged by SDS–PAGE (data not shown), the resulting pellet contained a high amount of PS II proteins, whereas the supernatant contained solubilized membrane material enriched in PS I. Fluorescence spectroscopy Fluorescence emission and excitation spectra were recorded at 77 K using a Perkin-Elmer LS50-B fluorimeter (Buckinghamshire, UK) equipped with a red-sensitive photomultiplier, or with an optical multichannel analyzer (OMA II, EG & G Princeton Applied Research, Princeton, New Jersey). Spectra were corrected for wavelength-dependent sensitivity of the detector. SDS-PAGE Membrane samples were solubilized and heated to 95 for 3 min with the SDS-reducing buffer containing 2% SDS, 0.1% bromophenol blue, 1% mercaptoethanol, 10% glycerol, 4.5 M urea and 62.5 mM Tris–HCl (pH 6.8). SDS–PAGE was carried out in a Biometra cooled minigel system according to Laemmli (1970). A 6% staking polyacrylamide gel and a separating gel with a 8–20% linear polyacrylamide gradient (of 90:1 acrylamide/ bis-acrylamide) were used. The gels were polymerized in presence of 4.5 M urea. An equal protein amount was loaded in the different wells. For Western blot experiments, the gels were run in duplicate. Only one gel was stained with Coomassie Blue for protein analyses.
◦C
Western blot analysis Polypeptides separated by SDS–PAGE of the duplicate gels were electroblotted to a nitrocellulose membrane (HATF/08250, Millipore, Bruxelles, Belgium) essentially according to Towbin et al. (1979), using a semi-dry transfer unit (SemiPhor TE77, Hoefer, SF, USA). As transfer buffer, a solution containing 24 mM Tris–HCl (pH 8.4), 192 mM glycine and 20% methanol was used. The transfer was performed at room temperature with 0.8 mA/cm2 constant power. Specific polypeptides were detected on the nitrocellulose by using antisera containing polyclonal antibodies against POR from barley (see below), D1 from oat (Kruse et al. 1993) and CF1 from spinach (Radunz and Schmid 1989), followed by a goat anti-rabbit IgG(H+L) horseradich peroxidase conjugate (Gamma,
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Figure 1. Schematic outline of the procedure for preparation of different membrane materials from (etio)chloroplasts.
Liège, Belgium) and HRP color development reagent (Bio-Rad, Richmond, California). POR antibody preparation Prolamellar bodies were isolated according to Ryberg and Sundqvist (1982) with only minor modifications, and were used for POR isolation. For the first purification step, the prolamellar bodies were suspended in a 25 mM HEPES–NaOH buffer (pH 7.5) containing 0.5 M sucrose, 5 mM NaCl, 5 mM MgCl2 and 1 mM EDTA. 2.5% (W/V) decylmaltoside and 2.5% (W/V) octylglucoside were added to the solution and incubated on ice for 15 min. The sample was then centrifuged at 4 ◦ C, 15 000 g for 15 min. The supernatant was collected and the proteins were concentrated by acetone precipitation. Two volumes of precooled (−18 ◦ C) acetone were added. After a few minutes, a centrifugation was applied at 4 ◦ C, 10 000 g for 30 min. The protein pellet was resuspended and solubilized with the SDS-reducing buffer (see below). Before electrophoresis the proteins were heated at 95 ◦ C for 3 min, then cooled on ice and centrifuged at 15 000 g to remove insoluble material. Preparative SDS–PAGE electro-elution (Prep Cell: model 491, Bio Rad, Richmond, California) was applied as final purification step using the discontinuous buffer system of Laemmli (1970). A 5.5 cm high 12% separating gel and a 1 cm high 4% stacking gel (of 45:1 acrylamide/bis-acrylamide) were polymerized in the tube of the apparatus. The power supply was adjusted to 30 mA constant current and SDS running
buffer (50 mM Tris, 384 mM glycine, 0.1% SDS) was pumped through the elution chamber at a rate of 0.6 ml/ min. The elution chamber outlet was connected to a fraction collector and 3 ml fractions were collected. In order to locate the fractions containing the POR protein, 50 µl of each fraction were analyzed by SDS–PAGE electrophoresis. The best fractions with respect to purity of the POR protein were pooled and concentrated using acetone precipitation. After centrifugation, the final pellet (± 500 µg) was solubilized in a phosphate buffer (pH 7.0) before injection into rabbits. Isoelectric focusing Preparative isoelectric complexes solubilized with the Rotofor Cell mond, California) as Kurdziel et al. 1997).
focusing of pigment–protein from etioplasts was realized Instruments (Bio Rad, Richalready described (Mysliwa-
Protein determination The total protein content of each subfraction was determined according to Bearden (1978) using a bovine serum albumin standard. Measurements of pigment concentration Total pigments were extracted in 3 ml of 80% acetone/ 20% water (v/v) from 100 µl of membranes suspensions. Insoluble material was removed by centrifugation and the Chl a and b concentration of the
67 supernatants were measured by fluorometry according to Meister (1992). For PChlide concentration measurements, Chls and carotenoids were first removed by transfer to petroleum ether (1 ml on 3 ml of acetone extract) according to Selstam and Widell (1986). Room temperature fluorescence emission spectra of the pigments in the acetone phase were recorded with the LS 50-B spectrophotometer set at an excitation wavelength of 436 nm. The excitation and emission slit widths were 10 and 5 nm, respectively. The total PChlide content of each sample was calculated from the fluorescence emission intensity at 634 nm. Calibration had been previously performed with PChlide extracted by the same way from barley etioplasts, using an absorption coefficient of 30.4 mM−1 cm−1 in the red absorption band (Brouers and Michel-Wolwertz 1983).
Results Preliminary observations on the plastid ultrastructure and on the nature of different (etio) chloroplast membrane fractions used in this study Plastids prepared at three different greening stages were compared in this study. Etiolated leaves collected after 5 or 12 h of continuous illumination were selected to provide etiochloroplasts in which Chl accumulates at fast and slower rate, respectively. Mature leaves from photoperiodically grown plants were used as green control. Intact plastids were extracted according to Ikeuchi and Murakami (1982) and their different membrane systems separated according to Douce et al. (1973). Whole thylakoids were further fractionated by partial solubilization by Triton X100 according to Berthold et al. (1981). Because greening induces marked changes in the shape, size and internal membrane organization of plastids (for review, see Ryberg et al. 1993), it was important to analyse plastid ultrastructure at the greening stages selected here for the preparation of different membrane fractions and to verify that the procedures used for isolating intact plastids and for separating their different membranes were adapted to the plant material in our conditions. Ultrathin plastid sections were examined by electron microscopy in order to probe changes in thylakoid organization during greening in our conditions. Electron micrographs in Figure 2 are representative of the development state of plastids located at a 4–4.5 cm distance from leaf tip after increasing times of continuous
illumination. After 3 h, remnants of prolamellar bodies were still observed and the newly formed thylakoids showed numerous perforations and protuberances but also short appressed regions. Appressed thylakoids were clearly visible after 5 h of greening. The number of appressed thylakoids in stacked regions and the relative length of appressed versus non-appressed thylakoids increased with further greening, in agreement with previous reports (Henningsen and Boynton 1974; Robertson and Laetsch 1974). Sections prepared at a distance of 2–2.5 cm from the leaf tip showed the same features, with the only noticeable difference that prolamellar body structures were only seldom observed after 3 h of greening. The intactness of the isolated plastids was verified by phase-contrast microscopy observations and further separation of envelope and thylakoid membranes was checked by SDS–PAGE (data not shown). Polypeptides of 54, 36, 30 and 14 kDa always dominated in the envelope fraction. The adapted method of Berthold et al. (1981) for PS II particles preparation from whole thylakoids yielded a pelletable sub-thylakoid fraction enriched in characteristic PS II polypeptides CP43, CP47 and LHCII. Already after 5 h of greening, the treatment of purified thylakoids by Triton X100 resulted in the recovery of a pelletable fraction. It is well-established that with fully green leaves this fraction, usually referred to as ‘PS II particle’ preparation, originates from the appressed thylakoids (Vallon et al. 1986). Since thylakoid appression was already well-advanced after 5 h of greening (Figure 2) it is highly probable that at this early greening stage the pelletable fraction obtained after Triton X100 treatment also originates from appressed thylakoids. In the following, this fraction will be referred to as ‘PS II thylakoids’ and the supernatant containing the remaining thylakoid material in solubilized form will be referred to as ‘Triton-solubilized thylakoids’. 77 K fluorescence properties and Pchlide content of intact plastids, of isolated envelope membranes and of thylakoid membrane fractions The 77 K fluorescence emission spectra of intact etiochloroplasts prepared after 5 or 12 h of greening revealed the normal Chl emission components of green leaves (Figure 3A): The 685 and 695 nm bands, corresponding to pigment–protein complexes of the PS II core antenna, and the broad 740 nm band corresponding to PS I pigment–protein complexes (Briantais et al. 1986). In addition to these Chl bands, a weak 635
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Figure 2. Plastid sections from primary leaves of 7-day-old dark-grown barley seedlings exposed to continuous illumination (as indicated). These plastid sections were issued from leaf pieces excised at 4 cm from the leaf tip. After an illumination of 3 h, primary lamellar layers show very short regions of thylakoid appression (indicated by arrows) and remnants of prolamellar bodies (Pb). After 5 h of illumination, thylakoids show larger and more frequent appressed regions and some well-defined grana (G).
nm band was detected. This band is due to Pchlide under nonphotoactive form (Virgin 1981). Its intensity decreased during greening but it was still detected in intact, mature chloroplasts (Figure 3B) in agreement with previous reports (Lebedev et al. 1985).
77 K fluorescence spectra of plastid envelopes, PS II thylakoids and Triton-solubilized thylakoids were compared, using an equal Chl concentration of 10 µg ml−1 . Substantial differences were found in the Chl region between the different membrane materials. This
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Figure 3. Comparison of the 77 K fluorescence emission spectra of etiochloroplasts isolated from 7-day-old dark-grown barley seedlings exposed to continuous illumination during 5 h (——), 12 h (-·-·-·) and from mature chloroplasts (· · · · · · ). The amount of plastids was adjusted to a Chl concentration of 10 µg/ml in each case. Excitation wavelength : 440 nm. The Pchlide region was magnified in the lower panel.
was true for the three developmental stages investigated. Figure 4 (A, B) compares the 77 K fluorescence spectra found in the Chl region of the different membrane fractions isolated from 5 h etiochloplasts and from mature chloroplasts. The two characteristic PS II fluorescence bands at 685 and 695 nm (shoulder) were clearly detected in the PS II thylakoid fractions. The 695 nm shoulder was lower or absent in Tritonsolubilized thylakoids. In mature chloroplasts, the 740 nm emission band of PS I was almost restricted to Triton-solubilized thylakoids. Envelope membranes showed a single Chl emission band of weak intensity at 683–685 nm. According to Pineau et al. (1986, 1993), this emission of chloroplast envelope arises from a mixture of Chl a and Chlide. In addition to Chl(ide) emission bands, weak Pchlide emission bands were also observed around 635 nm in the different membrane fractions. When using high magnification in this region (Figures 5A, B), this Pchlide emission appeared higher in envel-
Figure 4. Comparison of 77 K fluorescence emission spectra of membrane fractions isolated from etiochloroplasts of dark-grown barley seedlings exposed to continuous illumination during 5 h (A) and from mature barley chloroplasts (B). Envelope membranes (——), PS II thylakoids (· · · · · · ) and Triton-solubilized thylakoid subfractions (-·-·-·-·). The amount of samples was adjusted to give a Chl concentration of 10 µg/ml in each case. Excitation wavelength: 440 nm.
ope membranes than thylakoid membrane fractions. This was true at both developmental stages. However, the relative Pchlide emission intensity in envelope, when compared to the two thylakoid fractions, was much larger in mature chloroplasts than in 5-h illuminated etiochloroplasts. Additionally, when comparing the two thylakoid fractions, it appeared that Pchlide fluorescence was significantly more intense in PS II thylakoids than in Triton-solubilized thylakoids at both developmental stages. Results obtained with 12
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Figure 6. Comparison of the excitation spectrum of Pchlide fluorescence emission at 635 nm in 80% acetone extracts of whole thylakoids isolated from etiochloroplasts of dark-grown barley seedlings exposed to continuous illumination during 5 h (——), 12 h (- - - - -) and from mature chloroplasts (· · · · · · ).
Figure 5. Magnified view of the Pchlide region in the 77 K fluorescence emission spectra of Figure 4. The spectra of envelope membranes was divided by 10 to facilitate presentation.
h etiochloroplasts were intermediate between those obtained with 5 h etiochloroplasts and mature chloroplasts (data not shown). Some variations in position and width of the Pchlide emission band were observed in different membrane fractions. These differences were not further investigated, but they indicate some heterogeneity in the physico-chemical state of Pchlide. The Pchlide fluorescence found here around 630–635 nm may partly result from disaggregation of photoactive NADPH– POR–Pchlide complexes having an emission band at 655 nm. These complexes have been detected in intact leaves during grening (Franck and Strzalka 1992). However, a 655 nm emission band was not observed here after plastid isolation, even when adding NADPH during the isolation process (data not shown). Because quantitative measurements of Pchlide concentration cannot be achieved on the basis of the above 77 K fluorescence data, we next extracted Pchlide from the membrane fractions and we quantified their Pchlide concentration by measuring the fluorescence intensity at 634 nm under 436 nm excitation of the extracts at room temperature. In this condition, the fluorescence originated from Pchlide, and not from other Chl precursors. This is highlighted in Figure 6, which shows that the excitation spectra of the 634
nm fluorescence from the extracts of whole thylakoids have their maximum at 438 nm, irrespective of the greening stage. This wavelength is typical of mixtures of Pchlide pigments in monovinyl and divinyl forms, which have close excitation maxima in the Soret region at room temperature (Lebedev et al. 1985). A shoulder around 460 nm, which became more pronounced with greening, was due to a slight overlap effect with the fluorescence of Chl b (some amount of which remained in the acetone phase after transfer of the bulk of Chl to petroleum ether). On the basis of the excitation spectrum of pure Chl b (data not shown), we estimated that this effect caused only minor deviations in Pchlide quantitation from the 634 nm fluorescence intensity under 436 nm excitation. No indication of the presence of other pigments contributing to the 634 nm fluorescence was found from the excitation spectra of any fraction obtained in this work. Pchlide concentrations in the different membrane fractions are compared in Figure 7. Whereas 77 K fluorescence spectra were compared on a total Chl basis (in order to maintain comparable optical conditions), Pchlide concentrations in Figure 7 are expressed on a total protein concentration basis to facilitate the comparison with the POR distribution by immunoblot experiments (see next section). The Pchlide content of whole thylakoids of 5 h etiochloroplasts was about 10 times higher than the one of the envelope fraction (Figure 7, lanes a and b). This contrasted with the pigment distribution between these two fractions found in 12 h etiochloroplast and ma-
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Figure 7. Pchlide/protein ratios (ng mg−1 ) in the different plastid membrane subfractions of etiochloroplasts isolated from dark-grown barley seedlings exposed to continuous illumination during 5 h, 12 h and from mature chloroplasts. (a) whole thylakoids, (b) envelope membranes, (c) Triton-solubilized thylakoid sub-fraction, (d) PSII thylakoid sub-fraction.
ture plastid. In those cases, envelope membranes were strongly enriched in Pchlide, compared to whole thylakoids. During greening, the amount of Pchlide, expressed on a total protein basis, decreased in whole thylakoid membranes whereas it increased in envelope membranes. The relatively high 77 K Pchlide fluorescence of envelope membranes mentioned above for 5 h etiochloroplasts (see Figure 5A) did not correlate with a high amount of Pchlide in that fraction. This resulted from the fact that the Chl/protein ratio was about 1300 times lower in envelope membranes than in thylakoids of etiochloroplasts. Consequently, the relative Pchlide fluorescence was much higher on a Chl basis than on a protein basis. A more detailed analysis of Pchlide distribution in thylakoid membranes was made by comparing the Pchlide contents of ‘PS II thylakoid’ and ‘Tritonsolubilized thylakoid’ fractions resulting from partial solubilization with Triton X100 (Figure 7, lanes c and d). For the three selected greening stages, the PS IIthylakoid pellet was enriched in Pchlide, compared to the Triton-solubilized membranes of the supernatant. The ratio of Pchlide concentrations in PS II-thylakoids versus Triton-solubilized thylakoids increased from around 1.25 to more than 2. This trend was very reproducible. Immunochemical analyses of the different (etio)chloroplast subfractions To investigate the correlation between Pchlide pigment and POR proteins abundance in different membrane fractions, we have used a monospecific poly-
clonal antibody raised against this enzyme. The specificity of this antibody for POR proteins was first analyzed. A major polypeptide of 36 kDa and another one of slightly higher molecular mass (approx. 38 kDa) crossreacted with the antiserum. The concentration of these polypeptides (on total protein basis) rapidly decreased during the first 4 h of greening (Figure 8). After 6 h of illumination, only the band around 38 kDa could be observed when increasing the protein concentration (data not shown). The above data suggest that the two immunoreactive polypeptides are the two closely related PORA and B proteins identified in barley (Holtorf et al. 1995). Moreover, we found that the two reactive polypeptides of 36 and 38 kDa had the same pI of ± 5.2. This was shown by Western blot experiments against leaf proteins separated by isoelectric focusing (data not shown). This pI value is similar to the one found for POR from dark-grown wheat (Wiktorsson et al. 1992). Due to their close apparent molecular weight, the two immunoreactive POR polypeptides were not always resolved in Western blots. In the following analysis, we refer to POR without distinction between PORA and PORB. The POR antibody was used in Western blots of different (etio)chloroplasts fractions (envelope membrane, whole thylakoids, PS II-thylakoids and Tritonsolubilized thylakoids). To better characterize the different membrane fractions, we used additional antibodies raised against two polypeptides localized exclusively in thylakoid membranes: One raised against D1 polypeptide (32 kDa) of the PS II reaction center and another one raised against the CF1 55 kDa polypeptide of the coupling factor. As shown in the Western blots of Figure 9, the D1 and the CF1 polypeptides were never detected in envelope membranes. This result practically excludes a contamination of the envelope membrane fraction with thylakoid membranes by the method used in this study. When comparing the PS II-thylakoid and Triton-solubilized thylakoid fractions, it appeared that in mature chloroplasts (Figure 9C), CF1 polypeptide was almost absent from PS II thylakoids. This result confirms previous immunocytochemical studies on spinach PS II membranes (Vallon et al. 1986) and confirms that our PS II thylakoid fraction from green barley originates from non-appressed thylakoids, which are known to be practically devoided of coupling factor proteins (Vallon et al. 1986; reviewed in Staehelin and Van der Staay 1996). At earlier stages of greening (Figures 9A, B), however, no clear-cut partition of CF1 between the two thylakoid sub-fractions
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Figure 8. Western blots of POR proteins at different times of continuous greening of dark-grown barley leaves (indicated on top in hours). A total protein amount of 5 µg was loaded on each slot.
was found. These results may arise from a changing polypeptide distribution in different thylakoid regions during the biogenesis of the photosynthetic apparatus. The D1 polypeptide of PS II reaction center gave rise to an intense immunochemical reaction in the PS IIthylakoid subfraction at early greening stages (Figure 9A) when the ratio of reaction center proteins–lightharvesting complexes is high (Mysliwa Kurdziel et al. 1997). In mature chloroplasts, D1 is expected to be associated preferentially, but not exclusively, in the appressed thylakoids (Vallon et al. 1986). From the low intensity bands found here with the two sub-fractions of mature thylakoids (Figure 9C), it was difficult to appreciate D1 distribution between these samples, but it was clearly detected in both of them. The results of Western blots with the PORantibody indicated a continuous change in POR protein distribution among different membrane materials extracted from 5-h, 12-h and green (etio)chloroplasts (Figure 9). When comparing whole thylakoids to envelope membranes (lanes 1 and 2), it was apparent that POR distribution changed in favor of envelope membranes as a function of the extent of greening. On the other hand, POR appeared always predominant in PS II thylakoids compared to Triton-solubilized thylakoids (lanes 4 and 3). During the period of fast Chl accumulation (5 h of greening), POR was predominant in PS II-thylakoids, but significant amounts were detected also in Triton-solubilized thylakoids. In mature chloroplasts, it was not detected in that fraction. These changes in POR distribution among different plastid membrane fractions were in general qualitative agreement with those concerning the Pchlide pigment (see previous section and Figure 7). In the experiment of Figure 9, however, POR was not detected in Triton-solubilized thylakoids of mature chloroplasts, although small amounts of Pchlide
were measured in this fraction (Figure 7). We repeated Western blots with an increased (25 X) concentration of loaded proteins but were still unsuccessful in detecting POR in this fraction (data not shown). We cannot exclude that the detergent solubilization step artefactually caused the recovery of minor pigment amounts in this fraction, which would explain that Pchlide pigment could be detected despite the absence of POR.
Discussion In this study, we have separated envelope membranes and thylakoids of plastids at different developmental stages and analyzed their Pchlide and POR contents. The good separation of these fractions in both etiochloroplasts and chloroplasts was first attested by their different polypeptide contents and was confirmed by typical differences in 77 K fluorescence spectra. Western blots of D1 and CF1 polypeptides also indicated the absence of significant contamination of envelope membranes by thylakoid material. The detection of significant levels of POR proteins and Pchlide in the envelope membrane confirms earlier studies on mature chloroplasts (Joyard et al. 1990; Pineau et al. 1993). POR is a nuclear-encoded protein synthesized in the cytosol. The precursor, pPOR (approximately 44 kDa) is taken up into plastids where it is processed to its mature size (Teakle and Griffiths 1993; reviewed in Lebedev and Timko 1998). Moreover, there are indications that a great part of the Chl biosynthetic chain, from protoporphyrinogen IX to Pchlide, is located in the chloroplast envelope (Pineau et al. 1986, 1993; Joyard et al. 1990; Matringe et al. 1992). The enzymes which catalyze the
73
Figure 9. Western blot analysis of the different membrane subfractions of etiochloroplasts isolated from dark-grown barley seedlings exposed to continuous illumination during 5 h (A) or 12 h (B) and from mature chloroplasts (C). Lane 1: whole thylakoid membrane fractions. Lane 2: envelope membrane fractions. Lane 3: Triton-solubilized thylakoids. Lane 4: PS II-thylakoids. A total protein amount of 10 µg has been loaded on each slot. Protein blots have been probed with our polyclonal antibody raised against POR (36–38 kDa) and with polyclonal antibodies against D1 (32 kDa) and CF1 (55 kDa).
74 next steps (esterification of Chlide a with phytol or alcohol precursors as well as the final step of Chl a oxygenation to Chl b) are supposed to take place in the thylakoid membranes (Block et al. 1980; Soll et al. 1983; Porra 1997). According to our results, POR and Pchlide contents of envelope membranes are very low on a total protein basis at early greening stages (i.e. after 5 h of continuous illumination of etiolated leaves) in contrast to mature chloroplasts. In this study, POR and Pchlide were also detected in thylakoid membranes of both etiochloroplasts and mature chloroplasts. Its high relative abundance there at early greening stages may be functionally related to the assembly of the core complexes of both photosystems since Chl has been shown to be involved in the stabilization of plastid-encoded pigment–protein complexes (Klein et al. 1988; Kim et al. 1994). In greening and in mature green leaves investigated here, thylakoid-bound POR and Pchlide were preferentially associated with the pelletable thylakoid fraction which was recovered after partial Triton X100 solubilization (the PS II-thylakoid fraction), rather than with the supernatant (the Triton-solubilized thylakoid fraction). To appreciate the meaning of this result, it is necessary to consider the origin of these fractions. For fully green leaves, partial solubilization yields membrane fragments originating from appressed thylakoids which, at that stage, are almost devoided of coupling factor and PS I complexes (Vallon et al. 1986; Staehelin and Van der Staay 1996). We believe that in greening leaves pelletable membrane material after partial solubilization originates also from less exposed appressed membranes, since thylakoid appression was observed in ultrathin sections of plastids already after 5 h of greening. On the other hand, the presence of LHC II proteins (which correlates with thylakoid appresion) was demonstrated earlier in the same material already after 3–4 h of greening (Sigrist and Staehelin 1994; Mysliwa-Kurdziel et al. 1997). The 77 K fluorescence spectra and Western blots with D1 antibody provide evidence of PS II enrichment in these membrane fragments also. The Triton-solubilized membranes should mainly originate from non-appressed thylakoids, but may also contain some solubilized material from appressed regions. On the basis of this interpretation, our results indicate that at the earliest greening stage investigated here (5 h), the thylakoid-bound POR is associated preferentially to appressed thylakoids, but that significant amounts also occur in non-appressed thylakoids. The latter conclusion is in line with a previous study on
greening pea, in which Pchlide and monomeric Chl species were detected by fluorescence spectroscopy in an electron-dense fraction of primary thylakoids (Lebedev et al. 1990). In mature leaves, our study indicates that thylakoid-bound POR is exclusively located in appressed thylakoids. This, in turn, suggests a specific role of POR in the appressed regions of the photosynthetic membranes. In this context, it has been recently observed that an inhibition of Chl synthesis rapidly causes an inhibition of PS II activities and a loss of PS II components in mature green leaves illuminated by moderated light (Feierabend and Dehne 1996). The rapid turn-over of PS II components, including Chl, may require the presence of part of the Chl biosynthetic system, including POR, in the appressed thylakoids. Several studies have shown that POR becomes susceptible to proteolytic attack shortly after exposure of etiolated plants to light (Kay and Griffiths 1983; Häuser et al. 1984; Reinbothe et al. 1995). A light-induced nuclear-encoded protease, lacking in etioplasts of barley, has been suggested to degrade the POR–Chlide complexes. This protease has been principally localized in the stroma of chloroplasts (Reinbothe et al. 1995). This can provide an explanation for the preferential localization of Pchlide pigment and POR in appressed thylakoids, which should be less exposed than non-appressed thylakoids to the protease. It is important to note that our polyclonal antibody prepared against purified POR from etioplasts showed high specificity against POR, but reacted with two distinct polypeptides of around 38 and 36 kDa at very early stages of greening in continuous light. These two polypeptides might correspond to the two PORA and PORB isoenzymes identified in barley and in Arabidopsis (Holtorf et al. 1995; Armstrong et al. 1995; reviewed in Reinbothe et al. 1996). However, due to the difficulty of always clearly separating these two close bands and because it is impossible at present to unambigously ascribe them to the two POR gene products, we cannot discuss their respective distribution in different plastid membranes on the basis of our results. Finally, it is remarkable that differences in Pchlide and POR contents of the different membrane fractions from (etio-)chloroplasts correlate well to each other, at least qualitatively. This suggests that most Pchlide is originally bound to the enzyme. The association of Pchlide with POR has been shown to provide protection against photo-oxidative damage (Sperling et al.
75 1997) and it is likely that this effect is exerted during the whole greening process.
Acknowledgements This work was financed by grant no. 2.4597.99 of the Belgian National Funds of Scientific Research. Xavier Barthélemy and Gwénaëlle Bouvier thank the FRIA for the award of a PhD fellowship.
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