The Thylakoid Lumen of Chloroplasts - The Journal of Biological ...

1 downloads 1 Views 158KB Size Report
The chloroplast compartment enclosed by the thyla- koid membrane, the “lumen,” is poorly characterized. The major aims of this work were to design a ...

THE JOURNAL OF BIOLOGICAL CHEMISTRY © 1998 by The American Society for Biochemistry and Molecular Biology, Inc.

Vol. 273, No. 12, Issue of March 20, pp. 6710 –6716, 1998 Printed in U.S.A.

The Thylakoid Lumen of Chloroplasts ISOLATION AND CHARACTERIZATION* (Received for publication, September 9, 1997, and in revised form, January 7, 1998)

Thomas Kieselbach‡, Åsa Hagman, Bertil Andersson, and Wolfgang P. Schro¨der§ From the Department of Biochemistry, Arrhenius Laboratories for Natural Sciences, Stockholm University, S-106 91 Stockholm, Sweden

The chloroplast compartment enclosed by the thylakoid membrane, the “lumen,” is poorly characterized. The major aims of this work were to design a procedure for the isolation of the thylakoid lumen which could be generally used to characterize lumenal proteins. The preparation was a stepwise procedure in which thylakoid membranes were isolated from intact chloroplasts. Loosely associated thylakoid surface proteins were removed, and following Yeda press fragmentation the lumenal content was recovered in the supernatant following centrifugation. The purity and yield of lumenal proteins were determined using appropriate marker proteins specific for the different chloroplast compartments. Quantitative immunoblot analyses showed that the recovery of soluble lumenal proteins was 60 – 65% (as judged by the presence of plastocyanin), whereas contamination with stromal enzymes was less than 1% (ribulose-bisphosphate carboxylase) and negligible for thylakoid integral membrane proteins (D1 protein). Approximately 25 polypeptides were recovered in the lumenal fraction, of which several were identified for the first time. Enzymatic measurements and/or amino-terminal sequencing revealed the presence of proteolytic activities, violaxanthin de-epoxidase, polyphenol oxidase, peroxidase, as well as a novel prolyl cis/trans-isomerase.

The chloroplast is the photosynthetic organelle of green algae and higher plants. The chloroplast architecture comprises an envelope membrane, which encloses the soluble stroma as well as the highly specialized thylakoid membrane. The stromal compartment contains mainly the components of the Calvin cycle, which are required for the fixation of carbon dioxide. The thylakoids, on the other hand, carry out the light reactions of photosynthesis leading to the production of NADPH and ATP. The thylakoid membrane has a characteristic flat shape and is differentiated into appressed grana stacks and single non-appressed stroma-exposed lamellae. The inner surface of the thylakoid membrane encloses a narrow, continuous compartment, the lumen (1, 2). Electron microscopy studies of spinach thylakoids have suggested that the lumen is a densely packed space (3). No isolation method has so far been available for obtaining a high yield of pure thylakoid lumen. Thus, the

* This work was supported in part by grants from the Swedish Forestry and Agriculture Research Council, the Swedish Natural Science Research Council, and the Carl Trygger Foundation. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. ‡ Recipient of a guest research grant by The Swedish Institute. § To whom correspondence should be addressed: Dept. of Biochemistry, Arrhenius Laboratories, Stockholm University, S-106 91 Stockholm, Sweden. Tel.: 46-8-164392; Fax: 46-8-153679; E-mail: [email protected] biokemi.su.se.

present knowledge of the lumen from a compositional and functional point of view is fragmentary and is gathered from several independent approaches, addressing only single aspects of this compartment. By developing a technique for obtaining inside-out thylakoids, the investigation of the membrane surface of the lumenal side became possible (4). This work contributed to the discovery of the extrinsic proteins PsbO, PsbP, and PsbQ (5, 6) that bind to the lumenal side of photosystem II and are thought to stabilize the water oxidizing complex (7, 8). More recent studies have shown that these subunits of photosystem II occur also as soluble lumenal proteins (9). This pool of unassembled PsbO, PsbQ, and PsbP was resistant to proteolytic degradation and was capable of assembling into photosystem II (10). Furthermore, it was found that during photoinhibitory conditions the extrinsic proteins were released from the membrane into the lumen (11, 12). Other important components of the thylakoid lumen are plastocyanin, the primary electron donor of photosystem I (13, 14), and PsaN, a photosystem I subunit that is extrinsically bound to the lumenal side of the thylakoid membrane (15). Recent investigations have revealed that polyphenol oxidases (16, 17) and violaxanthin de-epoxidase are also present in the thylakoid lumen (18). Furthermore, the carboxyl-terminal processing protease for the D1 protein (19, 20) and a processing protease for plastocyanin (21) were found on the lumenal surface of the thylakoids, whereas chaperones may be located in the lumen (22). So far all lumenal proteins have been found to be nuclearencoded and synthesized as precursors in the cytoplasm. These precursor proteins have characteristic amino-terminal bipartite transit peptides, which direct their import into the chloroplast stroma and across the thylakoid membrane into the lumen (23–25). On the basis of this property, bipartite transit peptides have become typical markers for lumenal proteins. However, not all chloroplast proteins encoded with such presequences are routed into the lumenal space. The PsbW protein and CFoII, for instance, are synthesized with bipartite transit peptides but have been shown to be integral proteins of the thylakoid membrane (26 –28). In this study we have developed a procedure by which a lumenal fraction can be isolated in a highly pure form from spinach thylakoids. We have carried out the first systematic characterization of this compartment, and we show that the thylakoid lumen contains a high concentration of proteins, among which at least 25 distinct polypeptides can be identified. Several have been characterized in terms of enzymatic activity or amino-terminal sequence. EXPERIMENTAL PROCEDURES

Plant Material—Spinach (Spinacia oleracea) was grown hydroponically for 6 weeks with alternating periods of 10 h light and 14 h darkness.

6710

This paper is available on line at http://www.jbc.org

Thylakoid Lumen of Chloroplasts Isolation of the Thylakoid Lumenal Content—The general approach applied consists of three principal steps as follows: (i), preparation of chloroplasts, (ii) purification of carefully washed thylakoids, and (iii) rupture of thylakoids by a Yeda press and isolation of the lumenal content (Fig. 1). Spinach leaves (200 g) were blended twice for 5–10 s in 330 mM sorbitol, 50 mM Hepes-KOH (pH 7.8), 10 mM KCl, 1 mM EDTA, 0.15% (w/v) bovine serum albumin, 4 mM sodium ascorbate, and 7 mM cysteine. The resulting slurry was filtered through four layers of nylon mesh (20 mm), and the filtrate was centrifuged 1 min at 1000 3 g. The pellets were resuspended in 330 mM sorbitol, 50 mM Hepes-KOH (pH 7.8), 10 mM KCl, centrifuged for 1 min at 1000 3 g, and resuspended in 12–18 ml of the same buffer. The yield was 60 –70% intact chloroplasts containing 50 – 60 mg of chlorophyll. The chloroplasts were diluted with 10 mM sodium pyrophosphate buffer (pH 7.8) to a final chlorophyll concentration of 0.2 mg ml21 and resuspended in a glass homogenizer. The thylakoids were collected by centrifugation for 5 min at 7500 3 g and sequentially washed in the same way twice with each of the following buffers: I, 10 mM sodium pyrophosphate (pH 7.8) to remove the soluble stromal proteins; II, 2 mM Tricine (pH 7.8), 300 mM sucrose to partially remove ATP synthase, the membrane-attached fraction of Rubisco,1 and unidentified extrinsic thylakoid membrane proteins; III, 30 mM sodium phosphate (pH 7.8), 50 mM NaCl, 5 mM MgCl2, 100 mM sucrose (fragmentation buffer) to equilibrate the thylakoids for Yeda press fragmentation. The thylakoid pellets were suspended in a small volume of fragmentation buffer to a concentration of 3.5– 4.5 mg of chlorophyll ml21 (total yield: 25–30 mg of chlorophyll). The washed thylakoids were then passed once through a Yeda press at a nitrogen pressure of 10 megapascals and centrifuged for 1 h at 200,000 3 g and 2 °C. The supernatant was separated from the pellet and centrifuged a second time under the same conditions to spin down residual membrane particles. The entire isolation procedure was performed on ice, and the chloroplasts and thylakoid membranes were purified under green light. The lumenal fraction was either used directly or stored in liquid nitrogen. Thermolysin Treatment of Thylakoids—The washed thylakoids (2 mg of chlorophyll ml21) were incubated in the presence of 10 mM thermolysin (0.4 mg ml21) for 2 min on ice in 100 mM sucrose, 30 mM Hepes (pH 7.8), 50 mM NaCl, 5 mM MgCl2, and 2 mM CaCl2. The digestion was stopped by adding EDTA to final concentration of 50 mM, and the thylakoids were washed twice with 100 mM sucrose, 30 mM Hepes (pH 7.8), 50 mM NaCl, and 50 mM EDTA. These conditions were balanced to degrade the major part of peripheral proteins on the stromal side of the thylakoid membrane, without degrading too much of the integral membrane proteins. Harsher treatments were found to lead to leaky membranes followed by a loss of especially the lumenal part of the exposed stromal lamellae. Measurements of Chlorophyll, Protein, and Oxygen Evolution—Chlorophyll concentrations were measured as described (29). Determination of soluble proteins was carried out according to Ref. 30 and that of membrane proteins was performed essentially as described (31). The standard used was bovine serum albumin. Oxygen evolution activities and intactness of the chloroplasts were measured with a Clarke-type electrode at 20 °C using potassium hexacyanoferrate(III) as the electron acceptor (32). Electrophoretic and Immunoblot Analysis—SDS-PAGE was performed by the method described in Ref. 33 in slab gels containing 18% (w/v) polyacrylamide and 2 M urea. Determination of molecular masses were performed by using the low molecular weight electrophoresis calibration kit from Pharmacia Biotech Inc. For immunoblotting analyses proteins were transferred onto a polyvinylidene difluoride membrane in a semidry electroblotter system (Millipore). The antisera used were raised in rabbits against the following spinach proteins: phosphoribulokinase (K.-H. Su¨ss, Institute for Plant Genetics and Crop Plant Research, Gatersleben); violaxanthin de-epoxidase (H.-E. Åkerlund, University of Lund); plastocyanin (P.-Å. Albertsson, University of Lund); PsbO, PsbP, PsbQ, the lumen fraction, and Rubisco (our own production). The immunoreactivity was detected using goat anti-rabbit IgG-conjugated horseradish peroxidase in combination with an enhanced chemiluminescence detection. Quantification was performed using a Fast Scan Personal Densitometer and the ImageQuant software

1 The abbreviations used are: Rubisco, ribulose-bisphosphate carboxylase; AC, accession number; Chl, chlorophyll; FNR, ferredoxin-NADP1 reductase; LHCII, light harvesting complex II; PAGE, polyacrylamide gel electrophoresis; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl] glycine.

6711

from Molecular Dynamics. To be able to compare directly the results for the chlorophyll-free lumenal fractions with those of the chloroplast and thylakoids, a chlorophyll equivalent was used for the lumenal fraction. It was calculated as follows: [Lumeneq] 5 [Chl]thyl 3 Vthyl/Vlumen. However, the amount of plastocyanin and ferredoxin-NADP reductase (FNR) in the lumenal fraction was determined from the difference between the content in the washed thylakoids and their residual membrane fragments. Enzymatic Assays—Mitochondrial cytochrome c oxidase was assayed according to Ref. 34 and malate dehydrogenase according to the Worthington manual.2 As controls mitochondria from potato and spinach were used (kindly provided by P. Pavlov and X. Zhang, Stockholm University). Catalase activity was determined by oxygen evolution in the presence of 60 mM hydrogen peroxide (32). Phosphoribulokinase was assayed according to Ref. 35. To prevent oxidative inactivation of this enzyme during the lumenal isolation, all buffers contained 5 mM dithiothreitol. Diphenolase activity of polyphenol oxidase was determined at pH 4.6 using the substrate 3,4-dihydroxyphenylpropionic acid (36). Peroxidase activity was measured using the substrate 2,29-azinobis(3ethylbenzthiazoline-6-sulfonic acid) as described by Sigma.3 Protease activity was tested using a dye-linked peptide (PepTag, Promega) and the in vitro translated, [35S]methionine-labeled b-subunit of mitochondrial ATP synthase of Nicotiana plumbaginifolia (37) as artificial substrates. The activity of violaxanthin de-epoxidase was determined as in Ref. 38. All enzymatic activities were measured on freshly prepared samples (except those for violaxanthin de-epoxidase) at 25 °C in the presence of saturating substrate concentrations. Assay for ATP and ATPase Activity—ATP was detected using the firefly luciferase system (39) and the ATP monitoring reagent from BioOrbit. ATPase activity was tested according to Ref. 40. Amino-terminal Protein Sequence Analysis—Proteins were sequenced from polyvinylidene difluoride membrane following resolution by SDS-PAGE essentially as described (41). The amino-terminal sequence analyses were performed by P. I. Ohlsson (University of Umeå) using an Applied Biosystems pulsed liquid phase sequenator (ABS 477A). Searches in the data bases of EMBL and SwissProt as well as sequence alignments were carried out using the Wisconsin GCG software (42). RESULTS

Preparation and Characterization of the Thylakoid Lumen— The purpose of the present study was to obtain a preparation of thylakoid lumen in a yield and purity sufficiently high to make it generally useful for characterizing this chloroplast compartment. The developed method, as outlined in the scheme of Fig. 1, starts with the isolation of intact spinach chloroplasts. The chloroplasts obtained were normally 60 –70% intact and had an oxygen evolving activity of 140 –185 mmol of O2 mg Chl21 h21. In the next step the chloroplasts were disrupted by osmotic shock, and the thylakoids were collected and purified by several washing steps. Soluble stromal proteins were effectively removed by 10 mM sodium pyrophosphate (pH 7.8), and 2 mM Tricine (pH 7.8) containing 300 mM sucrose was used to remove ATP synthase, membrane-attached Rubisco, and other unidentified peripheral thylakoid proteins. Finally, the thylakoids were equilibrated in fragmentation buffer; these thylakoids retained an oxygen evolution rate of 90 –165 mmol of O2 mg Chl21 h21. The washed thylakoids were then fragmented by a Yeda press, and the released lumenal content was separated from the membrane fragments by two ultracentrifugation steps. The final fraction was free of chlorophyll and had a protein concentration of 0.3– 0.5 mg ml21, giving a total yield of 2–3 mg of protein from 200 g of spinach leaves. The separation during the course of the preparation was followed by electrophoretic analysis of the polypeptide pattern of the starting chloroplasts, the stromal fraction, the washed thylakoids, and the final lumenal preparation (Fig. 2). The dominant bands in the isolated chloroplasts (lane 1) were the 2 Available on-line at the following address: http//:www. worthington-biochem.com/manual/manIndex.html. 3 Available on-line at the following address: http://www.sigma.sial. com.

6712

Thylakoid Lumen of Chloroplasts

FIG. 1. Scheme for preparation of the thylakoid lumen from spinach chloroplasts.

large and small subunits of Rubisco and the LHCII. The two subunits of Rubisco and other soluble proteins were found in the stromal fraction (lane 2). The membrane-bound LHCII and other integral membrane proteins were recovered in the thylakoid fraction (lane 3). Finally, lane 4 shows the soluble lumenal proteins released after the disruption of the washed thylakoids and removal of the membrane fragments. As summarized in Table I, there were more than 25 polypeptides in the purified lumenal fraction. Furthermore, the polypeptide pattern (Fig. 2, lane 4) is clearly different from that of the other chloroplast subfractions. These polypeptides range in molecular mass from 14 to 100 kDa. There are 5 dominant, 10 to 12 medium abundance, and 8 to 10 low abundance polypeptides. Silver staining the polypeptides led to the detection of five additional bands in the molecular mass range of 20 – 45 kDa. Furthermore, an investigation of the low molecular mass polypeptides of the lumenal fraction by Tricine/SDSPAGE (43) revealed five additional low abundance polypeptides in the range of 6 –14 kDa (not shown). Following this general description of the protein content of the isolated lumenal fraction, several different approaches were used (summarized in Table I) to identify the individual polypeptides. Initially three of the four major polypeptides (Fig. 2, lane 4) at positions 32.5, 25, and 18 kDa were immunologically identified as the extrinsic proteins PsbO, PsbP, and PsbQ

of photosystem II. This is consistent with the observation that a soluble unassembled pool of these proteins occurs in the thylakoid lumen (9). The fourth major protein of the lumenal fraction, with the apparent molecular mass of 15 kDa, was identified by immunoblotting as plastocyanin. The proteins of the lumenal fraction were systematically studied by amino-terminal protein sequencing. This confirmed the identification of plastocyanin, PsbO, PsbP, and PsbQ. Polyphenol oxidase was identified as a polypeptide with an apparent molecular mass of 64 kDa, and four new polypeptides were discovered at 40, 29, 17.4, and 16.5 kDa (see Tables I and IV). Finally, ferredoxin NADP1 reductase (FNR) was found to migrate at apparent molecular masses of 38 and 37.5 kDa. The detection of FNR, which is located on the stromal side of the thylakoid surface, prompted a more detailed analysis of contamination in the isolated lumenal fraction. Therefore, the washed thylakoids were treated by limited proteolysis with thermolysin prior to the Yeda press fragmentation. Lumenal proteins, unlike the contamination, should be protected against such proteolytic degradation. The conditions of proteolysis were designed to ensure degradation of most of the peripheral protein on the stromal side of the thylakoid membrane and to minimize membrane rupture. More extensive proteolysis could cause disruption of the thylakoid membrane, as was previously reported (44). The intensity of the polypeptide bands of the lumenal fraction following SDS-PAGE before (Fig. 2, lane 4) and after (Fig. 2, lane 5) thermolysin treatment was compared by densitometric analysis of the Coomassie Blue-stained SDS gels using the PsbO protein as an internal reference protein. The data obtained from three different preparations demonstrated that most of the polypeptides of the lumenal fraction were not degraded, arguing that these proteins were located within the thylakoid lumen. Only in two cases were polypeptides significantly degraded (Fig. 2). A 40-kDa protein decreased in abundance by approximately 60%, whereas the 38kDa FNR (Fig. 2) was degraded to 80 – 85%, which is consistent with its location at the stromal surface of the thylakoid membranes (1, 2). The FNR may have been sheared off the outer thylakoid surface during the Yeda press treatment, thereby ending up with the lumenal proteins in the supernatant following centrifugation. In another test for purity an immunoblot analysis was performed to quantify marker proteins associated with specific chloroplast compartments: plastocyanin as a soluble lumenal protein; PsbO as a lumenal extrinsic thylakoid-bound protein; the D1 reaction center protein of photosystem II as an integral membrane protein; the FNR for the outer thylakoid surface; Rubisco as the major stromal protein, and phosphoribulokinase as typical soluble stromal enzyme. As presented in Table II, 60 – 65% of the total amount of plastocyanin and 10% that of the extrinsic PsbO protein were recovered in the lumenal fraction. Furthermore, the lumenal fraction was free of D1 protein demonstrating the absence of contaminating thylakoid membranes. In addition, less than 1% of the total amount of the large subunit of Rubisco was present in the lumenal fraction. The major contamination was FNR. Between 10 and 40% of this enzyme was found in the lumen; this represents approximately 10% of the total protein content of the lumenal fraction. As an additional indication of stromal contaminations as well as to find possible extra chloroplast contaminants, we assayed for a number of different marker enzymes. These marker activities included phosphoribulokinase from the chloroplast stroma, catalase from the cytoplasm, peroxisomes, and vacuole, and the mitochondrial enzymes cytochrome c oxidase (inner membrane) and malate dehydrogenase (matrix). As shown in Table III, the contamination of the isolated lumenal

Thylakoid Lumen of Chloroplasts

6713

FIG. 2. SDS-PAGE stained with Coomassie Brilliant Blue for polypeptide analyses of the isolated lumenal fraction and comparative analyses with other chloroplast subfractions. Lane 1, chloroplasts (10 mg of Chl); lane 2, stromal fraction (40 mg of protein); lane 3, thylakoids (10 mg of Chl); lane 4, lumenal fraction (40 mg of protein); lane 5, lumenal fraction (40 mg of protein) after thermolysin treatment. The bands of plastocyanin, the extrinsic subunits of photosystem II, LHCII, and Rubisco were identified by immunoblotting and/or amino-terminal sequencing (data not shown).

TABLE I Polypeptides of the isolated thylakoid lumen The apparent molecular masses were determined by SDS-PAGE from the Coomassie blue-stained proteins of the lumenal fraction. The application of silver staining led to the detection of additional bands at the apparent molecular masses of 19.5, 30.5, 30, 42, and 43 kDa (data not shown). Apparent molecular mass

Identity

Method of identification

kDa

100 64.5 57.5 54.0 40.0 38.0 and 37.5 32.5

? Polyphenol oxidase ? Large subunit of Rubisco and ? New protein (see Table IV) Ferredoxin NADP1 reductase PsbO protein

29.0 and 28.5 28.0 27.5 25.0

New protein (see Table IV) ? ? PsbP protein

24.0 23.0 21.0 20.0 18.5 18.0

? ? ? ? ? PsbQ protein

17.6 17.4 17.2 16.5 15.5 15.0

? New protein (see Table IV) ? New protein (see Table IV) ? Plastocyanin

14.0 6–14

? Five protein bands

Protein sequencing Immunoblotting Protein sequencing Protein sequencing Immunoblotting, protein sequencing Protein sequencing Immunoblotting, protein sequencing

Immunoblotting, protein sequencing Protein sequencing Protein sequencing Immunoblotting, protein sequencing Tricine/SDS-PAGE

fraction by phosphoribulokinase was only 0.3% of the total activity of the isolated chloroplasts and that of catalase was even lower. The activity of cytochrome c oxidase in the chloroplasts was 0.007 mmol of oxidized cytochrome c min21 mg protein21 and that of spinach mitochondria was 8 mmol of oxidized cytochrome c min21 mg protein21. Furthermore, the activity of malate dehydrogenase was 0.5 mmol of oxidized NADH min21 mg protein21 and that of the spinach mitochondria was 330 mmol of oxidized NADH min21 mg protein21.

TABLE II Immunological analyses of chloroplast subcompartment marker enzymes The fractions of the lumenal preparation were analyzed by immunoblotting for the following: (i) plastocyanin and the PsbO protein from the thylakoid lumen; (ii) the integral D1 protein and the peripheral ferredoxin NADP1 reductase (FNR) from the thylakoid membrane, and (iii) the stromal enzymes Rubisco and phosphoribulokinase. The amounts of these proteins in the chloroplasts were arbitrarily set at 100% and their relative amounts in the other fractions determined. The samples tested from the chloroplasts, the washed thylakoids, and their residual membrane fragments contained 2–5 mg of chlorophyll and those from the lumenal fractions 2–20 mg of protein. Chloroplasts

Washed thylakoids

Lumenal fraction

Residual thylakoid fragments

%

Thylakoid lumen Plastocyanin PsbO protein Thylakoid membrane D1 protein Ferredoxin-NADR1 reductase Chloroplast stroma Rubisco (large subunit) Phosphoribulokinase

100 100

60–75 80–90

60–65 10

2–10 70–80

100 100

100–110 40–60

0 10–40

100–105 5–30

100 100

8–20 ,1 Under detection limit

NDa ND

ND, not determined.

Based on these values the contamination of the lumenal fraction by soluble mitochondrial proteins was estimated to be lower than 0.2%. The activity of cytochrome c oxidase was not detectable in the lumenal fraction. Enzyme Activities of the Thylakoid Lumen—Earlier work had suggested that polyphenol oxidase (17) and violaxanthin de-epoxidase (18) are located in the lumen of the spinach thylakoids. Therefore, we analyzed the isolated lumenal fraction for the presence of these two enzymes. The specific activity of polyphenol oxidase increased from the chloroplasts to the washed thylakoids and attained a maximum in the lumenal fraction (Table III). The fraction of the polyphenol oxidase activity that remained with the thylakoid fragments was 8% of that present in the lumenal fraction. The presence of polyphenol oxidase in the lumenal fraction was also confirmed by amino-terminal protein sequencing of the polypeptide with an apparent molecular mass of 64.5 kDa (Table I).

6714

Thylakoid Lumen of Chloroplasts

TABLE III Activities of marker enzymes The fractions of the lumenal preparation were screened for the activities of marker enzymes. The specific activities for the lumenal extract were determined using an equivalent chlorophyll concentration as described under “Experimental Procedures.” ND, not determined. Chloroplasts

Thylakoid lumen Polyphenol oxidasea Violaxanthin De-epoxidaseb Chloroplast stroma Phosphoribulokinasec Mitochondria and cytoplasm Cytochrome c oxidased Cytochrome c oxidasee Malate dehydrogenasef Malate dehydrogenaseg Catalasei

Washed thylakoids

Lumenal fraction

Residual thylakoid fragments

0.01 6 0.01 ND

0.09 6 0.05 ND

0.25 6 0.05 18 6 1.8

0.02 6 0.005 ND

761

0.07 6 0.001

0.024 6 0.002

ND

0.06 6 0.005 0.007 6 0.0006 0.18 6 0.02 ND 341 6 240

0.02 6 0.007 0.003 6 0.001 0.14 6 0.002 ND 14 6 7

0 0 0.05 6 0.04 0.5 6 0.3 0.7 6 0.2

ND ND ND ND ND

mmol oxidized 3,4-dihydroxyphenylpropionic acid min21 mg Chl21. mmol of violaxanthin min21 g protein21. c mmol oxidized NADPH min21 mg Chl21. d mmol oxidized cytochrome c min21 mg Chl21. e mmol oxidized cytochrome c min21 mg protein21. f mmol oxidized NADH min21 mg Chl21. g mmol oxidized NADH min21 mg protein21. h mmol O2 mg h21 Chl21. a b

The specific activity of violaxanthin de-epoxidase in the lumenal fraction was 18 mmol of violaxanthin/g of protein21 min21 (Table III), which corresponds to 2–5% of the value reported for the purified spinach enzyme (38). In addition, the immunoblot shown in Fig. 3 reveals that violaxanthin de-epoxidase was highly enriched in the lumenal fraction as compared with the original thylakoid preparation. In searching for other lumenal enzymes, we considered the observation that photosystem II can produce hydrogen peroxide (45), and we therefore tested the lumenal fraction for peroxidase activity. Surprisingly, 60% of the total peroxidase activity of the thylakoids was recovered in the lumenal fraction, which contrasts with earlier work (46) that suggested that peroxidases are not present in this chloroplast compartment. The lumen-associated peroxidase activity was not affected by limited proteolysis of the thylakoids. Finally, the turnover of the D1 protein of photosystem II during photoinhibition involves proteolytic cleavages in both stromal and lumenal loops (47). In addition, processing of imported precursor proteins and the carboxyl-terminal extension of the D1 protein must occur in the lumen (25, 20). The lumenal preparation was therefore tested for protease activities. The overall proteolytic specific activity in the chloroplast stroma and thylakoids was approximately equal, whereas the proteolytic activity in the lumenal fraction was only 10% that in the thylakoids. The pH optimum of the lumenal proteolytic activity was between pH 7 and pH 8. However, in contrast to the thylakoids this activity was still detectable in the pH range of 4 – 6. Metalloprotease inhibitors were found to inhibit this protease activity by 30%. Moreover, the addition of ATP did not stimulate the proteolytic activity. These results corroborate the presence of lumen-located proteases. The lumenal fraction was also tested for the presence of ATP and ATPase activity. The ATP concentration was found to be lower than 100 nM, and no significant ATPase activity could be detected. Identification of Unknown Lumenal Proteins via Amino-terminal Sequencing—To find new lumenal proteins, amino-terminal sequencing of polypeptides from the lumenal fraction was performed after their separation by SDS-PAGE. This approach led to the identification of four polypeptides of the apparent molecular masses of 40.0, 29.0, 17.4, and 16.5 kDa (Table IV). The 29-kDa protein shows no homology to any known sequence in the SwissProt and EMBL data bases. However, the

FIG. 3. Immunoblot analysis of the thylakoids and the lumenal fraction from a lumenal preparation for violaxanthin de-epoxidase. Lane 1, thylakoids (20 mg of Chl), and lane 2, lumenal fraction (20 mg of protein). TABLE IV Amino-terminal sequences of four unknown proteins from the thylakoid lumenal fraction kDa

1

40.0 29.0 17.4 16.5

VLISGPXIKD ADLIQRRQRS ANQRLPPLSN APLEDEDDLE

30

PEALLRYALP EFQSDIKGIL DPKRKE.... LLEKVKRDRK

IDNKAIREVQ YTVIKKNPDL .......... KRLERQGAI.

amino-terminal sequence of the 16.5-kDa protein was homologous to partial protein sequences encoded by Arabidopsis thaliana clones 250E4T7 (AC W43350) and 77E7T7 (AC T45153). The cDNAs of both clones overlapped partially and were very similar. Fig. 4A shows an alignment of the polypeptide with the translation product (g1327818) of clone 250E4T7. The sequence from A. thaliana contains parts of the presequence and the amino terminus. Furthermore, the presequence reveals features of a bipartite transit peptide with a hydrophobic core and the motif AXA at the putative processing site (24). As shown in Fig. 4B, the amino-terminal sequence of the 17.4-kDa protein was 68% homologous to a protein encoded by a cDNA fragment from A. thaliana (clone 104N18T7, AC T21992). This sequence, also of unknown function, contains a part of a bipartite transit peptide. The motif VIA at the putative processing site is analogous to the corresponding motif VLA in the precursor of the PsbQ protein from spinach (AC P12301). Furthermore, this motif is flanked by a hydrophobic region that is typical of thylakoid transfer domains present on transit peptides of lumenal proteins. As shown in Fig. 4C, the amino-terminal sequence of the 40.0-kDa protein was homologous to the putative amino terminus of a hypothetical protein g1001111 from the cyanobacterium Synechocystis sp. PCC6803 (AC D64001). Similar to the

Thylakoid Lumen of Chloroplasts

6715

FIG. 4. Alignment of the amino-terminal sequences of the 16.5-kDa protein (A), the 17.4-kDa protein (B), and the 40-kDa protein (C) with homologous sequences of A. thaliana and Synechocystis sp. PCC6803. The prepeptides of these sequences show similar features as the bipartite transit peptides of lumenal chloroplast proteins as a hydrophobic core region and the motif AXA at the putative processing site (underlined). In case of the 17.4-kDa protein the putative processing site reveals the motif VIA, which is very similar to the processing site VLA of the PsbQ protein from spinach (AC P12301). Positive charged residues are marked with a plus sign.

Arabidopsis sequences, this protein also contains a transit peptide with a hydrophobic core region and AXA motif at the putative processing site. Interestingly, the carboxyl-terminal 191 residues of hypothetical protein g1001111 is highly homologous to 184 residues of the carboxyl terminus of peptidylprolyl cis/trans-isomerase B (AC D90900, g1651784) from this cyanobacterium (not shown). This suggested that the 40-kDa protein from spinach and the Synechocystis protein g1001111 were functionally related to peptidyl cis/trans-isomerase. Recently, the spinach 40-kDa protein was isolated, and its corresponding cDNA was cloned and sequenced (AC Y12071).4 The deduced gene product is a unique high molecular mass immunophilin protein that is located in the thylakoid lumen. This prediction was biochemically confirmed through direct enzymatic analysis of the cis/trans protein isomerization (rotamase) activity of the purified spinach 40-kDa protein4 that is typical for immunophilins (48). In addition, the lumenal fraction from our work was found to have a significant rotamase activity.5 We have not been able to successfully sequence the N termini of the low abundance lumenal proteins. DISCUSSION

The thylakoid lumen represents a continuous exoplasmic cellular space that is poorly characterized compared with the other chloroplast compartments. Interest in the lumenal side of the thylakoid membrane has mainly focused on electron transport events associated with the inner membrane surface. Indirectly, the thylakoid lumen has also been considered as essential for the generation of the trans-thylakoid proton gradient that drives ATP synthesis and for the anion and cation currents established by ion channels in the thylakoid membranes (49, 50). More recently, with an increased understanding of 4 Fugolsi, H., Vener, A. V., Altschied, L., Herrmann, R. G., and Andersson, B. (1998) EMBO J., in press. 5 A. V. Vener, unpublished data.

biosynthesis, regulation, and stress protection of the photosynthetic apparatus, the requirement for auxiliary enzymes (51) in the lumenal space has become more apparent. The present isolation of a thylakoid lumenal fraction gives high yield and low degree contamination, thereby providing a new tool for biochemical analysis of this chloroplast compartment. Based upon a volume to chlorophyll ratio for thylakoids of 3.3 ml per mg of chlorophyll (52) and the yield of 75–120 mg of lumenal protein per mg of chlorophyll in the starting thylakoid material, the protein concentration in the lumenal space is estimated to be higher than 20 mg ml21. Thus, soluble proteins in the lumen are at micromolar to millimolar concentrations, which is similar to that of the chloroplast stroma. Hence, the thylakoid lumen is comprised of a densely packed core of soluble components, as suggested from electron microscopy studies (3). The number of resolved polypeptides in the isolated lumenal fraction was approximately 25, of which 15 remain to be functionally identified. In contrast to what is observed for the thylakoid proteins only a few of these are of low molecular mass. The purity of the isolated lumenal fraction was very high as judged by its low contamination of stromal and thylakoid integral membrane proteins. The one notable exception was FNR, which is functionally associated with the outer thylakoid surface and which could not be removed by various pre-washes. Moreover, FNR is a notoriously “sticky” protein that contaminates various protein preparations including those of cytochrome b/f (53), PsbO (54), and PsbS (55). The PsbO, PsbP, and PsbQ proteins and plastocyanin were found to be the major proteins present in the isolated lumenal fraction. The presence of extrinsic PsbO, PsbP, and PsbQ polypeptides is consistent with the previous observation of an unassembled, stable, lumenal pool of these polypeptides (9, 10), particularly under photoinhibitory conditions (11, 12). The amount of soluble PsbO in the isolated lumenal fraction was

6716

Thylakoid Lumen of Chloroplasts

10% relative to the total content of chloroplasts. This agrees with the finding that 8% of the PsbO of spinach occurs in an unassembled state in the thylakoid lumen (9). The identification of violaxanthin de-expoxidase and polyphenol oxidase as soluble polypeptides of the lumen corroborates previous work suggesting the occurrence of these enzymes in this compartment (16 –18). Furthermore, the presence of a peroxidase in the lumen may be important for detoxification of hydrogen peroxide produced by photosystem II upon illumination (45). The function of lumenal polyphenol oxidase is not understood. Since chloroplasts from peas lack this enzyme (16), it is not likely to play a role crucial for photosynthesis. The polyphenol oxidase might be transported into the lumen where it would be stored safely separated from its substrate in the vacuole. The stability of the three unassembled extrinsic proteins of photosystem II brings up the question of proteolytic activities in the thylakoid lumen. The present analyses as well as previous studies (56, 57) clearly reveal the presence of proteases in this compartment. However, the protease activity of the lumenal fraction was relatively low, only 10% of that found in the thylakoid fraction, suggesting that most of lumenal protease activity is bound to the inner thylakoid surface. The 16.5-kDa lumenal protein is highly homologous to the deduced sequence of the A. thaliana clone 250E4T7 (g1327818), which shows a typical bipartite transit peptide. However, the three arginine residues close to the putative processing site (Fig. 4) represent an unknown motif for previously determined bipartite transit peptides of chloroplast proteins. It would be of interest to determine the specific import pathway by which this protein is transported from the cytoplasm into the thylakoid lumen. The possibility of lumenal chaperones (22) as well as experimental indications for phosphorylation of lumenal proteins (58) have been reported. This work does not support this view since we could not show the presence of ATP or demonstrate ATPase activity in the lumen. The presence of nucleotides in the thylakoid lumen will require more detailed studies. Acknowledgments—We thank Drs. P.-Å. Albertsson and H.-E. Åkerlund (University of Lund) and Dr. K.-H. Su¨ss (Institute for Plant Genetics and Crop Plant Research, Gatersleben) for the generous supply of antibodies. We greatly appreciate the excellent technical assistance of Ann-Christine Holmstro¨m. REFERENCES 1. Andersson, B., and Barber, J. (1994) Adv. Mol. Cell Biol. 10, 1–53 2. Hall, D. O., and Rao, K. K. (1994) Photosynthesis, 5th Ed., Cambridge University Press, Cambridge, UK 3. Weibull, C., and Albertsson, P.-Å. (1988) J. Ultrastruct. Mol. Struct. Res. 100, 55–59 4. Andersson, B., and Åkerlund, H.-E. (1978) Biochim. Biophys. Acta 503, 462– 472 5. Åkerlund, H.-E., and Jansson, C. (1981) FEBS Lett. 124, 229 –232 6. Åkerlund, H.-E., Jansson, C., and Andersson, B. (1982) Biochim. Biophys. Acta 681, 1–10 7. Murata, N., and Miyao, M. (1985) Trends Biochem. Sci. 10, 122–124 8. Vermaas, W. F. J., Styring, S., Schro¨der, W. P., and Andersson, B. (1993) Photosynth. Res. 38, 249 –263 9. Ettinger, W. F., and Theg, S. M. (1991) J. Cell Biol. 115, 321–328 10. Hashimoto, A., Yamamoto, Y., and Theg, S. M. (1996) FEBS Lett. 391, 29 –34 11. Hundal, T., Virgin, I., Styring, S., and Andersson, B. (1990) Biochim. Biophys. Acta 1017, 235–241 12. Eisenberg-Domovich, Y., Oelmu¨ller, R., Herrmann, R. G., and Ohad, I. (1995)

J. Biol. Chem. 270, 30181–30186 13. Haehnel, W., Berzborn, R. J., and Andersson, B. (1981) Biochim. Biophys. Acta 637, 389 –399 14. Haehnel, W. (1984) Annu. Rev. Plant. Physiol. 35, 659 – 693 15. He, W.-Z., and Malkin, R. (1992) FEBS Lett. 308, 298 –300 16. Sommer, A., Ne’eman, E., Steffens, J. C., Mayer, A. M., and Harel, E. (1994) Plant Physiol. 105, 1301–1311 17. Sokolenko, A., Fulgosi, H., Gal, A., Altschmied, L., Ohad, I., and Herrmann, R. G. (1995) FEBS Lett. 371, 176 –180 18. Hager, H., and Holocher, K. (1994) Planta 192, 581–589 19. Inagaki, N., Mori, H., Fujita, S., Yamamoto, Y., and Satoh, K. (1995) in Photosynthesis: From Light to Biosphere (Mathis, P., ed) Vol. 3, pp. 783–786, Kluwer Academic Publishers Group, Drodrecht, Netherlands 20. Oelmu¨ller, R., Herrmann, R. G., and Pakrasi, H. B. (1996) J. Biol. Chem. 271, 21848 –21852 21. Kirwin, P. M., Elderfield, P. D., Williams, R. S., and Robinson, C. (1988) J. Biol. Chem. 263, 18128 –18132 22. Schlicher, T., and Soll, J. (1996) FEBS Lett. 379, 302–304 23. von Heijne, G., Steppuhn, J., and Herrmann, R. G. (1989) Eur. J. Biochem. 180, 535–545 24. Robinson, C., and Klo¨sgen, R. B. (1994) Plant Mol. Biol. 26, 15–24 25. Robinson, C., and Knott, T. G. (1996) in Molecular Genetics of Photosynthesis (Andersson, B., Salter, A. H., and Barber, J., eds) pp. 145–159, IRL Press at Oxford University Press, Oxford 26. Michl, D., Robinson, C., Shackleton, J. B., Herrmann, R. G., and Klo¨sgen, R. B. (1994) EMBO J. 13, 1370 –1317 27. Lorkovic, Z. J., Schro¨der, W. P., Pakrasi, H. B., Irrgang, K.-D., Herrmann, R. G., and Oelmu¨ller, R. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 8930 – 8934 28. Shi, L.-X., Schro¨der, W. P. (1997) Photosynth. Res. 53, 45–53 29. Porra, R. J., Thompson, W. A., and Kriedemann, P. E. (1989) Biochim. Biophys. Acta 975, 384 –394 30. Bradford, M. M. (1976) Anal. Biochem. 72, 248 –254 31. Markwell, M. A. K., Haas, S. M., Bieber, L. L., and Tolbert, N. E. (1978) Anal. Biochem. 87, 206 –210 32. Leegood, R. C., and Malkin, R. (1986) in Photosynthesis Energy Transduction: A Practical Approach (Hipkins, M. F., and Baker, N. R., eds) pp. 9 –26, IRL Press at Oxford University Press, Oxford 33. Laemmli, U. K. (1970) Nature 227, 680 – 685 34. Errede, E., Kamen, M. D., and Hatefi, Y. (1978) Methods Enzymol. 53, 45– 46 35. Porter, M. A., Milanez, S., Stringer, C. D., and Hartman, F. C. (1986) Arch. Biochem. Biophys. 245, 14 –23 36. Espı´n, J. C., Morales, M., Varo´n, R., Tudela, J., and Garcı´a-Ca´novas, F. (1995) Anal. Biochem. 231, 237–246 37. Boutry, M., and Chua, N.-H. (1985) EMBO J. 4, 2159 –2165 38. Arvidsson, P.-O., Bratt, C. E., Carlsson, M., and Åkerlund, H.-E. (1996) Photosynth. Res. 46, 141–149 39. Lundin, A., Rickardsson, A., and Thore, A. (1976) Anal. Biochem. 75, 611– 620 40. Fromme, P., and Gra¨ber, P. (1990) Biochim. Biophys. Acta 1016, 29 – 42 41. Matsudaira, P. (1987) J. Biol. Chem. 262, 10035–10038 42. Genetics Computer Group (1996) Program Manual for the Wisconsin Package, Version 9, Madison, WI 43. Scha¨gger, H., and von Jagow, G. (1987) Anal. Biochem. 166, 368 –379 44. Carter, D. P., and Staehlin, A. (1980) Arch. Biochem. Biophys. 2, 364 –373 45. Schro¨der, W. P., and Åkerlund, H.-E. (1986) Biochim. Biophys. Acta 848, 359 –363 46. Hayakawa, T., Kanematsu, S., and Asada, K. (1984) Plant Cell Physiol. 25, 883– 889 47. Barber, J., and Andersson, B. (1992) Trends Biochem. Sci. 17, 61– 66 48. Fischer, G., Wittmann-Liebold, B., Lang, K., Kiefhaber, T., and Schmid, F. X. (1989) Nature 337, 476 – 478 49. Pottosin, I. I., and Scho¨nknecht, G. (1995) J. Membr. Biol. 148, 143–156 50. Pottosin, I. I., and Scho¨nknecht, G. (1996) J. Membr. Biol. 152, 223–233 51. Andersson, B. (1992) in Trends in Photosynthesis Research (Barber, J., Guerrero, M. G., and Medrano, H., eds) pp. 71– 86, Intercept Ltd., Andover, UK 52. Heldt, H. W., Werdan, K., Milovancev, M., and Geller, G. (1973) Biochim. Biophys. Acta 314, 224 –241 53. Clark, R. D., and Hind, G. (1983) J. Biol. Chem. 258, 10348 –10354 54. Andersson, B., Jansson, C., Ljungberg, U., and Åkerlund, H.-E. (1985) in Molecular Biology of the Photosynthetic Apparatus (Arntzen, C., Bogorad, L., Bonitz, S., and Steinback, K., eds) pp. 21–31, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY 55. Funk, C., Schro¨der, W. P., Napiwotzki, A., Tjus, S. E., Renger, G., and Andersson, B. (1995) Biochemistry 34, 11133–11141 56. Kuwabara, T., and Hashimoto, Y. (1990) Plant Cell Physiol. 31, 581–589 57. Sokolenko, A., Altschmied, L., and Herrmann, R. G. (1997) Plant Physiol. 115, 827– 832 58. Gal, A., Zer, H., and Ohad, I. (1997) Physiol. Plant. 100, 869 – 885

Suggest Documents