Tubular Crystals of a Photosystem II Core Complex

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J. Mol. Biol. (1996) 259, 241–248

Tubular Crystals of a Photosystem II Core Complex Georgios Tsiotis1*, Thomas Walz1, Aspasia Spyridaki2, Ariel Lustig3 Andreas Engel1 and Demetrios Ghanotakis2 1

M. Mu¨ller Institute for Microscopical Structure Biology, 3Department of Biophysical Chemistry Biozentrum, University of Basel, Klingelbergstr. 70 CH-4056 Basel, Switzerland

2

Department of Chemistry University of Crete, P.O. Box 1470, 71409 Iraklion, Crete Greece

An oxygen evolving photosystem II core complex containing all three extrinsic proteins (33, 23, 17 kDa) was isolated from spinach and reconstituted into tubular two-dimensional crystals of 72.9 nm diameter and 1-2 micrometers length. While the 17 and 23 kDa polypeptides were lost during crystallization, the extrinsic 33 kDa protein was retained. The optical spectrum of the crystallized core was characteristic of an intact PSII core complex. Immunoelectron microscopy revealed that the lumenal surface of the PSII complex was exposed at the outside of the cylindrical tubes. The projection of the complex was determined from flattened tubular crystals by negative stain electron microscopy and image analysis to 2.0 nm resolution. Rhombic unit cells (a = 16.2 nm, b = 13.7 nm; g = 142.4°) contained one PSII complex.

7 1996 Academic Press Limited

*Corresponding author

Keywords: immuno-gold labeling; 2D crystals; photosystem II; oxygen evolving complex; 33 kDa protein

Introduction Photosynthetic electron transfer in plants involves photosystems I and II (PSI, PSII), which act together with cytochrome b6 f to transfer electrons from water to NADP+. PSII from higher plants contains more than 22 polypeptides (Masojidek et al., 1987) some of which have a distal antenna function. At least nine subunits, the psbA-psbF and psbO-psbQ gene products, are necessary for plastoquinone reduction and oxygen evolution. The membrane-integral core complex consists of D1 and D2 (psbA and psbD gene products), a heterodimeric cytochrome b559 (psbE and psbF gene products) and two Chla-binding inner light-harvesting antenna proteins CP47 and CP43 (psbB and psbC gene product; Bricker, 1990). A set of hydrophilic proteins (psbO-psbQ gene products) which have apparent molecular masses of 33, 23 and 17 kDa form the oxygen evolving complex (OEC), and are associated with the lumenal surface of the PSII complex (Andersson & Akerlund, 1987). The 33 kDa extrinsic protein stabilizes the Mn cluster of the OEC. The function of the other two extrinsic proteins is closely related Abbreviations used: Chla, chlorophyll a; LHC, light-harvesting complex; CP, chlorophyll proteins; OTG, octyl-b-thioglucoside; PSII, photosystem II; DMPC, dimyristoyl phosphatidyl choline; OEC, oxygen evolving complex; MES, (2-[N-morpholino]ethanesulfonic acid); 2D, 3D, two- and three-dimensional; STEM, scanning transmission electron microscopy. 0022–2836/96/220241–08 $18.00/0

to the unique requirement of Ca2+ and Cl− for O2 production. Several other subunits such as the psbH, psbJ and psbL gene products have been identified in PSII core complexes from higher plants but their function is not known (Ikeuchi & Inoue, 1986; Koike & Inoue, 1985; Koike et al., 1989). A series of integral membrane proteins which bind Chla/b and carotenoids are associated with PSII and form the light-harvesting complex (LHC) (for a review see Jansson, 1994). The major component, LHC-II, consists of several proteins in the range of 24 to 29 kDa and is considered to bind up to 50% of the total Chl present in thylakoid membranes. Recently the structure of LHC-II was ˚ resolution by electron determinated to a 3.4 A crystallography (Ku¨hlbrandt et al., 1994). Three minor pigment-proteins CP29, CP26 and CP24 (each binding a few percent of the total Chl) have been characterized and it was proposed that they function as linker proteins between the PSII core complex and LHC-II (Bassi & Dainese, 1992). Several structural studies of either PSII crystals or isolated PSII complexes have been performed. Electron microscopy and single-particles imageaveraging of isolated PSII complexes from spinach allowed the localization of the 33 kDa protein and the LHC-II complex on the PSII core complex (Boekema et al., 1995). Small 2D crystals of a core complex comprised of the CP47, D1, D2 and cytochrome b559 , have provided a resolution of 2.5 nm, but no information regarding the structure of the entire PSII complex (Dekker et al., 1990). 7 1996 Academic Press Limited

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Tubular Crystals of Photosystem II Core Complex

Recently, low resolution 3D structures of PSII complexes which had been depleted either of the antenna proteins or of the antenna proteins and the OEC polypeptides, were reported (Santini et al., 1994). The structural analysis showed a dimeric organization of the PSII complex. In contrast, an analysis of the 3D structure of PSII obtained from small crystals containing the OEC polypeptides and antenna proteins and from small crystals which had been depleted of the OEC polypeptides, suggested that PSII is monomeric (Ford et al., 1995, Holzenburg et al., 1993). In this paper we report the reconstitution of tubular crystals of a PSII oxygen evolving complex. The crystals were characterized by various techniques. Image analysis of the tubes enabled us to derive a map of the structure in projection at 2.0 nm resolution. Mass measurement of the crystals in combination with analytical ultracentrifugation indicated that the unit cell contains one PSII complex.

Results After one to five days the PSII complex reconstituted with exogenous DMPC showed membranous structures with prominent, regularly arranged particles when examined by negative stain electron microscopy. The preparations were highly enriched in tubes (approximately 80% of the sample, see Figure 1), but a few vesicles were also present. Tubes grown under these conditions measured 73(25) nm (n = 76) in diameter and exhibited a variable length of 1–2 mm. At higher magnification the helical packing of the particles in the tubes became distinct (Figure 1 inset). The polypeptide composition of these crystals was examined by SDS-PAGE. As shown in Figure 2A, the solubilized isolated PSII core complex consists of the CP47, CP43, D1, D2, CP29, CP26 polypeptides and four proteins migrating below 10 kDa. Further, all three OEC proteins (17, 23 and 33 kDa) were present in the complex (lane 1). Lane 2 shows the protein pattern of the PSII crystals which apparently lacked the 17 and 23 kDa polypeptides. Nevertheless, the room temperature absorption spectrum of the 2D crystals exhibited a profile characteristic of intact PSII complexes (Figure 2B), as specifically suggested by the 678 nm peak (Newell et al., 1988). The broad shoulder at 470 to 490 nm is due to carotenoids, whereas the peak at 438 arises primarily from Chla. The lack of a strong shoulder at 650 nm shows that there is very little Chlb present. Negatively stained flattened tubular PSII crystals displayed a distinct periodic pattern arising from the two superimposed collapsed layers (Figure 3A). The calculated power spectrum (Figure 3B) therefore represents the superposition of two separate diffraction patterns. To distinguish between the two sets of diffraction spots, reconstituted PSII tubes were freeze dried and

Figure 1. Negatively stained tubular crystals of the purified PSII core complexes recorded at low magnification. While the tubes show a rather fixed diameter, they vary strongly in length. At higher magnification the helicaly packed protein particles become distinct (inset). Scale bars represent 500 nm and 100 nm in the inset.

unidirectionally shadowed with platinum-iridiumcarbon (Figure 3C). This method reveals only the top layer which allowed the lattice parameters of the tubular crystals to be defined unambiguously from the diffraction pattern shown in Figure 3D. In Figure 3E the top lattice is marked on the left side and the bottom lattice on the right side. From a series of 30 micrographs of negatively stained specimens the best images were selected by their optical diffraction pattern for image processing. Top and bottom layers of the collapsed PSII tubes were processed independently. The four best correlation averages, each comprised of approximately 100 unit cells, all emerged from the top layer and were added to yield a projection map at 2.0 nm resolution (Figure 4A), as established from the diffraction orders above background. The unit cell had a rhombic shape (a = 16.2 nm; b = 13.7 nm; g = 142.4) and housed two elongated domains separated by a central depression. Although the central cavity was open on one side but closed on the other by a connecting bridge the pseudo 2-fold symmetry was distinct. Both major protrusions

Tubular Crystals of Photosystem II Core Complex

243

Figure 2. A, Characterization of the polypeptide composition by SDS-gel electrophoresis of purified PSII complexes (lane 1) and of the tubular 2-D crystals (lane 2). B, Room temperature absorption spectra of the isolated PSII complexes (dotted line) and of the PSII complex crystals obtained (continuous line).

exhibited a tripartite structure, whereas the minor protrusions were elongated. The same general features were also observed with the single particle

averages (Figure 4B) calculated from images of solubilized, negatively stained PSII complexes recorded with the STEM (Figure 4D), and single

Figure 3. Image analysis of the tubular PSII crystals. Straight areas of the negatively stained tubes (A) were selected for image processing. The obtained power spectrum (B) is difficult to index as both layers of the collapsed tube contribute to the diffraction pattern and produce two overlaying sets of spots. To solve this problem, tubes were freeze-dried and unidirectionally shadowed with a platinum/iridium alloy (C). The power spectrum of the shadowed sample (D) shows only diffraction spots deriving from the top layer of the tube and therefore the reciprocal lattice could be determined unambiguously (marked in black). As all micrographs were digitized in the same way, the diffraction spots seen in the power spectrum from negatively stained tubes could also be indexed. In E the reciprocal lattices are marked that define the positions of the diffraction spots seen in B: the lattice in the left half derives from the top layer of the tube while the lattice on the right half originates from the bottom layer. Scale bars represent 100 nm in A, and (20 nm)−1 in B. The arrow in C indicates the shadowing direction.

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Tubular Crystals of Photosystem II Core Complex

Figure 4. A, Projection map of negatively stained tubular PSII crystals. The unit cell (a = 16.2 nm; b = 13.2 nm; g = 142.4°) contains a monomer. B, Averaged projection of PSII complex. Sum of 68 top view particles. C, Averaged projection of the PSII complexes from the non-ordered crystals. Sum of 250 particles. Bar represents 5 nm in A to C. D, Electron micrographs of negatively stained PSII complexes. Bar represents 25 nm. E, Non-well ordered tubular crystals of PSII complexes. Bar represents 50 nm.

particles averages (Figure 4C) from disordered crystals grown by dialysis (Figure 4E). Mass measurement and analytical ultracentrifugation were carried out to determine the unit cell stoichiometry. Mass measurements with the STEM yielded a mass/area histogram with a distinct peak at 8.83(20.26) kDa/nm2 (n = 414) for air-dried flattened tubes (Figure 5A). Therefore, each unit cell comprising an area of 136 nm2 has a mass of 600 kDa which includes protein and surrounding lipids. The density of the tubes was measured by analytical ultracentrifugation in RDDG (Figure 5B), as well as a static gradient (Figure 5C) to determine the lipid-protein ratio. The density of the tubular crystals was found to be 1.19 g/ml. Assuming a density of 1.33 g/ml for proteins and 1.03 g/ml for the DMPC (Lustig et al., 1991), we calculate that the protein represented 53% of the mass, i.e. 318 kDa. To identify the orientation of the PSII complex in the membrane, immunogold labeling was performed with two different gold conjugated antibodies. One antibody was raised against the lumenal 33 kDa protein of the OEC while the second one was specific for the amino acid residues, 230 to 235 of the D2 protein. The antibody against the 33 kDa polypeptide labeled the tubular crystals (Figure 6A), as well as the vesicles (data not shown).

In comparison, the antibody against D2 did not label the tubes (Figure 6B), whereas it labeled the vesicles (data not shown. This observation indicates that the

Figure 5. Mass analysis of tubular crystals by scanning transmission electron microscopy (A). The banded membrane containing regular arrays of PSII complex (vertical broad line) r = 1.194 g/ml in a Nicodenz RDDG in a 4 mm RDDG-BC cell at 8000 rpm, 22°C. Photo taken after 40 minutes at a = 80° schlieren angle (B). The banded membrane containing regular arrays of PSII complex (vertical broad line) r = 1.184 g/ml in a static Nicodenz DG in a SS 1.5 mm Alum cell at 34,000 rpm, 22°C. Photo taken after 48 hours, a = 70° (C).

Tubular Crystals of Photosystem II Core Complex

Figure 6. On-grid immunogold labeling of PSII crystals. Detection of the outside of the tubular crystals was achieved by a gold-coupled anti-33 antibody (A). A gold-coupled anti-D2 antibody did not react to the tube (B). Scale bar represent 100 nm.

lumenal side of the complex faces the outside of the tubes.

Discussion A recent report suggests the PSII complex to be incorporated as dimer in a large submolecular unit comprised of PSII and LHCII (Boekema et al., 1995). However, when solubilized with dodecyl-b-maltoside or octyl-b-glucoside the PSII complex exists as a monomer with a weak 2-fold symmetry when viewed along the normal to the membrane. In this projection the PSII monomer has a length of 13 nm and a width of 9 nm (Boekema et al., 1995). A variety of ordered PSII arrays have been described, all induced by Triton X-100 treatment of chloroplast membranes. Reported unit cell dimensions and shapes are a = 53 nm, b = 53 nm, g = 90° (Santini et al., 1994); a = 26.7 nm, b = 17.8 nm, g = 90° (Bassi et al., 1989); a = 18.9 nm; b = 16.8 nm, g = 91° (Holzenburg et al., 1993); a = 11.5 nm, b = 16.1 nm, g = 75° (Lyon et al., 1993). According to SDS-PAGE all these crystals exhibited the full set of PSII polypeptides with the exception of the extrinsic proteins 33, 23 and 17 kDa which were either lost during the crystallization (Lyon et al., 1993), or specifically removed by various salt- and Tris-washing procedures (Bassi et al., 1989; Holzenburg et al., 1994). Images of negatively stained crystals reported by Bassi et al., and those of the crystal described by Holzenburg et al., have a similar appearance, but these authors come to different conclusion concerning the unit cell stoichiometry. In addition, the averages calculated from negatively

245 stained crystals show different morphologies (Bassi et al., 1989; Dekker et al., 1990; Ford et al., 1995; Holzenburg et al., 1993) with little similarity to the projections of the solubilized monomeric PSII complex (Boekema et al., 1995). In this paper we present tubular crystals reconstituted from solubilized PSII monomers in the presence of DMPC by a simple dilution procedure. SDS-PAGE showed that the 23 kDa and 17 kDa polypeptides were the only polypeptides lost during crystallization, while the optical spectrum was identical to that of the native PSII complex. These 2D crystals had lattice vectors a = 16.2 nm, b = 13.7 nm, g = 142.4°, and folded into cylinders of 73(25) nm in diameter. Their morphology is similar to that of the tubular crystals described by Lyon et al. (1993). However, the lattice dimensions of our crystals are apparently too small to provide space for one PSII dimer per unit cell. We therefore used the STEM to determine the mass per area of the collapsed tubes, and the ultracentrifuge to determine their density. From these data, we calculate that the unit cell contained a protein mass of 318 kDa. This is close to the mass of one PSII complex which amounts to 359.5 kDa without the 17 kDa and 23 kDa OEC proteins. In addition, the stoichiometry of one PSII complex per unit cell was corroborated by the similar dimensions and morphologies of the unit cell and the averaged projection of solubilized complexes. This finding contradicts the dimeric unit cell reported by Lyon et al. (1993). Considering the similarity of the unit cell morphology, it is likely that their crystals are identical to those reported here. The intrinsic pseudo 2-fold symmetry of the PSII complex, and the assumption of a molecular mass of 200 kDa for one PSII complex without the OEC proteins may have promoted a misinterpretation of the PSII stoichiometry in the unit cell. The unit cell stoichiometry of the 2D PSII long crystals has been controversial. Santini et al. (1994) assigned their low resolution 3D structure to a dimeric complex. In contrast, a 3D map exhibiting similar features has been interpreted as a monomeric PSII complex (Holzenburg et al., 1993). The experimental evidence discussed in our work may help to resolve this controversy. The sequence homology of the D1 and D2 polypeptides (Trebst, 1986) as the pronounced similarity in the hydrophobicity profiles of the CP43 and CP47 subunits (Bricker, 1990) together with the current consensus of a one-to-one stoicheometry of these proteins (Satoh, 1985) support the idea of a PSII complex with a pseudo 2-fold symmetry. According to their sequence, D1 and D2 are expected to possess rather short hydrophilic loops connecting the transmembrane spans, whereas CP43 and CP47 are expected to have significant hydrophilic domains (Bricker, 1990). Considering the peripheral position of stain excluding protrusions in our projections maps, we speculate that the CP43 and CP47 are located at the periphery of the complex. This interpretation is in line with the

246 conclusion of Ford et al. (1995) from their 1.8 nm 3D map of the Tris-washed PSII complex. Several studies have shown that the OEC proteins are at the lumenal surface of the thylakoid membranes (Seibert et al., 1987). From the amino acid sequence and membrane folding homology of the D2 polypeptide and the M subunit of the bacterial reaction center, it has been postulated that the loop between helices IV and V is exposed to the stromal membrane surface, with the amino acid residues 230 to 235 forming a part of this loop (Michel & Deisenhofer, 1988; Trebst, 1986). The antibody against the 33 kDa protein reacted with both tubes and vesicles whereas the antibody against D2 reacted with vesicles but not with tubes. This suggests that the lumenal surface of the PSII complex is at the outside of the cylindrical tubes, and that the 33 kDa protein is bound to the tubular crystals. A stoichiometry of one copy of each of the OEC proteins per PSII complex has previously been suggested based on crosslinking experiments and Fourier difference analysis of 2D crystals of PSII and Tris-treated PSII crystals (Enami et al., 1991; Holzenburg et al., 1994). In addition, a tentative location of the 33 kDa protein has been proposed (Boekema et al., 1995). The projection of our tubes showed two very similar protrusions which we interpret as the hydrophilic lumenal protrusions of the CP43 and CP47 which both have an estimated mass of about 20 kDa (Bricker, 1990). SDS-PAGE and the immunogold labeling experiments provide solid evidence that the 33 kDa protein was present. If only one copy of the 33 kDa protein had been bound per reaction center, we should have observed a significant asymmetry. On the other hand, previous experiments have shown that there are two 33 kDa proteins per reaction center (Xu & Bricker, 1993) which would be consistent with our structural observations. To resolve this question and to localize the subunits with better accuracy higher resolution micrographs are required. The tubular crystals presented in this work may become a solid basis for such a structural analysis.

Materials and Methods Isolation and crystallization of a photosystem II complex The PSII complex was isolated as described (Mishra & Ghanotakis, 1994). The complex was then resuspended in 50 mM MES (pH 6), 0.4 M sucrose, 10 mM NaCl, 0.4% octyl-b-thioglucoside (OTG) and mixed with dimyristoyl phosphatidyl choline (DMPC) which had been solubilized in 1.3% OTG. The lipid concentration was varied to yield a lipid/protein ratio ranging between 0.1 and 1. The resulting solution was diluted with 50 mM 2-Hepes over a ×4 range and incubated at temperatures of 6 to 22°C. Solutions were monitored by withdrawing 3 ml samples every eight hours and examining the contents by electron microscopy using negative staining. Tubes appeared reproducibly within these temperature/dilution ranges after a period of one to five days. To obtain vesicles,

Tubular Crystals of Photosystem II Core Complex

dialysis experiments were carried out using the same conditions, Hoenger et al. (1990).

Fixed beam electron microscopy Specimens were adsorbed to glow-discharged, carboncoated Parlodion films that had been mounted on electron microscope grids. After washing in double-distilled water, the membranes were negatively stained with 0.75% (w/v) uranyl formate. Alternatively, adsorbed tubes were quickly frozen in liquid nitrogen, freeze-dried at −80°C in a Balzers BAF 300 and unidirectionally shadowed with platinum-iridium-carbon at an elevation angle of 45° and 80 Hz. The samples were observed in a Hitachi H-7000 electron microscope operated at 100 kV acceleration voltage and 50,000× magnification. Images were recorded on Kodak SO-163 films without pre-irradiation, at a dose of typically 2000 electrons/nm2.

Scanning transmission electron microscopy Samples were adsorbed to glow discharged thin carbon films supported by fenestrated films on gold coated copper grids. After washing with water, samples were stained with uranyl formate (0.75%) and subsequently placed in a pretreatment vacuum chamber directly connected to a Vacuum Generators HB5 STEM. Elastic annular dark field images were recorded at 100 kV acceleration voltage and doses between 2000 and 4000 electrons/nm2. For mass measurement tubular crystals were adsorbed to thin carbon film supported by fenestraded film, washed with double-destilled water, and air-dried. Elastic dark field images were recorded in STEM at doses of 306(20.27) electrons/nm2. The mass/area value of circular areas of 1475 nm2 were evaluated as described by Mu¨ller et al. (1992).

Image analysis Micrographs of 2D crystals were inspected in an optical diffractometer and selected according to the alignment of the microscope and the observed crystal order. Suitable areas were digitized as described (Ford et al., 1990) using an Eikonix 850 CCD camera at a pixel size corresponding to 0.64 nm on the sample. The averaged PSII projection was calculated using the SEMPER image processing system. A rapid evaluation of crystal quality and unit cell morphology was achieved by Fourier peak filtration (Aebi et al., 1973). Residual lattice disorder was eliminated by correlation averaging (Saxton & Baumeister, 1982). For single particle averaging STEM dark field images of well preserved particles sampled at 0.35 nm were selected interactively, using the SEMPER image processing (Saxton et al., 1979) system installed on a VAX 3100 work station. 387 particles were selected for the PSII complex comprising 64 × 64 pixels. In the first step, the images were aligned with correlation techniques, starting with a single projection as first reference. Intermediate references were constructed from the sums of the best particles as estimated from their correlation with the reference. In the next step, over 60% of the aligned particles were submitted to multivariate statistical analysis, and automatic classification (Frank et al., 1987; van Heel, 1984). The largest class was then used as reference for further refinement.

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Immune electron microscopy Colloidal gold particles, 08 nm in diameter, were prepared by reduction of tetrachloroauric acid with sodium citrate in the presence of tannic acid (Slot & Geuze, 1985). Antibodies against the 33 kDa and D2 proteins were conjugated to the colloid particles as described (Baschong & Wrigley, 1990). After conjugation, the complexes were dialyzed against phosphate-buffered saline (PBS) and centrifuged at 45,000 g for 15 minutes. The soft pellet was resuspended in the crystallization buffer. For immunogold labeling, the gold-conjugate was added to the reconstituted membranes in various ratios. After incubation for one hour at 8°C in the dark, the mixture was centrifuged for five minutes in an Eppendorf microfuge at 5000 g and then washed with the crystallization buffer. After two washing steps, samples were negatively stained with uranyl formate and electron microscopy was carried out as described above.

Spectroscopic methods UV-visible absorbance spectra of the PSII complex and reconstituted PSII were recorded with a Zeiss UMPS-80 single beam microspectralphotometer which was coupled to a Hewlett Packard 3100 computer.

Analytical ultracentrifugation The density of the lipid/PSII complex has been determinated with the model E (Beckman) analytical ultracentrifuge equipped with a schlieren optical system by the rapid dynamical density gradient (RDDG) method (Lustig et al., 1991), and using a static gradient. For RDDG a Nicodenz (Nyegaard, Oslo, Norway)/Hepes buffer solution r = 1.231 g/ml was underlayered under a r = 1.146 g/ml solution containing the reconstituted complex in a 4 mm SBDS-BC cell at 22°C, 8000 rpm. Static density gradient centrifugation was carried out in a 1.5 mm SS 4° Alum cell at 22°C, 34,000 rpm, using an initial density of Nicodenz buffer solution of r = 1.202 g/ml.

Other methods SDS gel electrophoresis experiments were carried out as described by Ikeuchi & Inoue (1988). Protein was determined using the BCA protein assay (Pierce), and chlorophyll concentration was measured following Arnon (1949).

Acknowledgements The authors thank G. Fritzsch for help with the microphotometer measurements, N. Pante for the advice regarding immunogold labeling techniques and S. Mu¨ller for reading the manuscript. The project was supported by the M. E. Mu¨ller-Foundation of Switzerland, the Department of Education of the Kanton Basel-Stadt. A.E. was supported by a grant of the Swiss National Foundation for Scientific Research (grant 31-42435.94) and D.F.G. was supported by a grant from the Greek Secretariat for Research and Technology.

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Edited by R. Huber (Received 6 November 1995; received in revised form 16 February 1996; accepted 25 March 1996)

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