J. Biosci., Vol. 15, Number 2, June 1990, pp. 93–98. © Printed in India.

Lamellar dispersion and phase separation of chloroplast membrane lipids by negative staining electron microscopy R. C. YASHROY Biology Division, Carleton University and National Research Council, Ottawa, Canada Present address: Biophysics, Electron Microscopy and Instrumentation Section A. N. Division (Bldg.), Indian Veterinary Research Institute, Izatnagar, Bareilly 243 122, India MS received 28 February 1990 Abstract. Aqueous dispersions of lipids isolated from spinach chloroplast membranes were studied by electron microscopy after negative staining with phosphotungstic acid. Influence of low temperature (5°C for 24 h) was also investigated. It was observed that when contacted with water, these lipids, as such, formed multilamellar structures. Upon sonication, these multilamellar structures gave rise to a clear suspension of unilamellar vesicles varying in size (diameter) between 250 and 750 Å. When samples of sonicated unilamellar vesicles were stored at 5°C for 24 h or more, they revealed a variety of lipid aggregates including liposomes, cylindrical rods (about 100 Å wide and up to 3600 Å long), and spherical micellar structures (100-200 Å in diameter)—thus indicating phase separation of lipids.

Keywords. Chloroplast; lipids; membranes; electron microscopy.

Introduction Chloroplast membranes have an unusual lipid composition which includes chlorophylls, carotenoids, sterols, quinones, phospholipids and glycolipids (Benson, 1966, 1971). Unlike most other membranes in the biological world, phospholipids constitute only a minor fraction (about 10%) of the total chloroplast membrane lipids (Kates, 1970). Monogalactosyl-diacyl-diglycerol (MGDG), which constitutes the largest lipid component in the chloroplast membranes, does not form a lamellar structure; but instead it forms a reversed hexagonal liquid-crystalline phase. However, digalactosyl-diacyl-diglycerol (DGDG) forms an aqueous bilayer structure (Larsson and Puang-Ngern, 1979; Murphy, 1986). Larsson and PuangNgern (1979) assumed the lipid organization in thylakoid membranes to be a bilayer and opined against inverted-bilayer structure as proposed by Kreutz (1966) from X-ray diffraction studies. The present work was undertaken with the view to investigate the modalities as to how a variety of diverse lipids which constitute the chloroplast membranes may co-exist all together in aqueous lipid dispersions and how they undergo phase separation due to the effect of low temperature. Materials and methods Isolation of chloroplast membranes Chloroplasts were isolated from chilled spinach leaves (without mid-ribs) by the Abbreviations used: MGDG, Monogalactosyl-diacyl-diglycerol; DGDG, digalactosyl-diacyl-diglycerol.

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method of Arnon et al. (1956) and suspended in 0·2 Μ sodium chloride containing, 5 mM magnesium chloride. The preparation was sonicated for 2 min at 5°C under a stream of nitrogen gas (to prevent air-oxidation of lipids) and then centrifuged at 1000 g (0°C) for 7 min to obtain the membrane pellet. Extraction of membrane lipids Lipids were extracted from the membrane-pellet following the procedure of Bligh and Dyer (1959). All solvents were bubbled with nitrogen gas before use and the routine drying and concentration of lipids was also accomplished under a stream of nitrogen gas. The final lipid-extract was dissolved in a mixture of chloroform and methanol (2:1 v/v), and stored at 10°C after sealing under an atmosphere of nitrogen gas. Preparation of aqueous lipid dispersions For preparation of aqueous lipid dispersions, an aliquot of lipid-extract was completely dried under a stream of nitrogen gas and then under vacuum (1-2 h), and was subsequently vortexed with a known volume (normally 1-2 ml) of distilled water for about 15 min at room temperature in a glass tube sealed under an atmosphere of nitrogen gas. To prepare sonicated lipid-water dispersions, the vortexed aqueous lipiddispersions were sonicated for about 10 min at 20°C in glass tubes sealed under an atmosphere of nitrogen gas, so as to obtain a clear (green-coloured) solution. Electron microscopy The aqueous lipid dispersions (sonicated or unsonicated) were stained with phosphotungstic acid by the method described by Lucy and Glauert (1964). Satisfactory results were also obtained by first drying (under nitrogen gas) a thin film of dilute lipid-extract (in chloroform-methanol 2:1 v/v) laid on a forvar-coated copper grid and then dipping it in distilled water for 2 min. The water-washed grid was then stained by placing a drop of 1% phosphotungstic acid on its coated-side for 2 min. Excess stain was soaked away by touching a filter paper strip. The grid was then dried under a stream or nitrogen gas and examined under Siemens Elmiskop I electron microscope at an instrument magnification of × 45,000. Results Figure 1 shows the vortexed aqueous lipid dispersions prepared from the spinach chloroplast membranes. A typical multilamellar (liposomal) structure of these dispersions is clearly noticeable. Each lamella corresponds to a lipid bilayer structure. Rare spherical structures (arrow) about 100 A in diameter are also seen in these preparations which correspond to spherical micellar structures. When the liposomal suspensions are sonicated, a clear green-coloured solution results. On examination under the electron microscope after negative staining, it

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Figure 1. Electron micrograph of aqueous dispersion of lipids extracted from spinach chloroplast membranes after staining with phosphotungstic acid. Multilamellar (liposomal) structures are observable in the whole view. A few spherical structures (arrow-head) about 100 Å in diameter are also sparsely observable which are interpreted as spherical micelles (× 225,000).

Figure 2. Electron micrograph of sonicated aqueous dispersion of lipids extracted from spinach chloroplast membranes after staining with phosphotungstic acid. Unilamellar microvesicles of diameter varying from 250-750 Å are observable (× 225,000).

reveals unilamellar vesicles (figure 2) with diameter ranging between 250 and 750 Å. When samples of sonicated lipid-microvesicles (figure 2) are stored at temperature of 5°C for 24 h or more, a totally different picture appears under the

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electron microscope. Figure 3 shows such an electron micrograph wherein a variety of lipid aggregates are seen. Structures like multilamellar liposomes cylindrical rods

Figure 3. Electron micrograph of sonicated aqueous dispersion of lipids extracted from spinach chloroplast membranes after storage at low temperature (5°C for 24 h) and staining with phosphotungstic acid. Different lipid aggregates observable are (a) multilamellar vesicles, (b) spherical micelles of diameter ranging between 100-200 Å and (c) bunches of cylindrical rods with thickness of about 100 Å and length upto 3600 Å, besides (d) original (sonicated) unilamellar micro-vesicles (× 157,500).

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Lipids of chloroplast membranes and micelle-like spheres, besides the pre-existing unilamellar vesicles indentifiable. The cylindrical rods have a diameter of approximately 100 Å.

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Discussion Chloroplast membranes contain a diverse variety of lipids and only a few of them are known to form a lipid bilayer structure (Murphy, 1986). The present studies (see figures 1, 2) reveal that many of the constituent lipids form non-bilayer lipid aggregates in isolation, but when they are mixed together, they form a fairly stable multibilayer structure (figure 1) which, upon sonication gives rise to unilamellar lipid micro-vesicles with diameter ranging between 250–750 Å. Such multilamellar aqueous lipid-dispersions and unilamellar microvesicles are known to represent a lipid-bilayer structure as has been revealed by studies on lipids from many other diverse sources (Kimelberg, 1977; Huang, 1969; Bangham et al., 1974). These observations also support the commonly accepted assumption that the lipid organisation in the chloroplast membranes, should be in the form of a bilayer. The chloroplast membranes and their extracted lipid-water dispersions show a high lipidfluidity leading to the detection of sharp carbon-13 nuclear magnetic resonances arising from lipid fatty-acyl carbons (YashRoy 1987a, b). They also reveal gel-toliquid crystalline phase transition around 18°C (YashRoy, 1990). When samples of sonicated unilamellar vesicles (figure 2) were kept at temperature of 5°C for 24 h or more, marked changes were observed (figure 3) in the lipid organisation. Multilamellar liposomal-vesicles (figure 3a) were observed which were apparently formed by 'fusion' of unilamellar vesicles. In addition, structures like cylinderical rods (figure 3b) and spherical micelles (figure 3c) were also seen. It is clear that prolonged exposure to low-temperature of samples of these unilamellar vesicles results in the phase separation of the constituent lipids. Polarizing microscope and low-angle X-ray diffraction studies (Larsson and PuangNgern, 1979) of isolated chloroplast membrane-lipids taken individually, showed that when dispersed in water, the lipid, DGDG formed a lamellar phase. The lipid, sulphoquinovsyl-diacyl-glycerol is also known to form a lamellar phase with water (Murphy, 1986). On the other hand, the lipid, MGDG formed a reversed hexagonal liquid-crystalline phase with water. The latter phase seemingly appears as cylindrical rods under the electron microscope (figure 3). The spherical micelle-like structures (figure 3c) may well represent aggregates of molecules like chlorophyll, sterols etc. The present investigation thus provides evidence that the diverse lipids of chloroplast membrane when dispersed in water all-together, form the lamellar (bilayer) structure under normal conditions. The lamellar structure is fairly stable. Under conditions of prolonged exposure to low-temperature (5°C), these lipid-water dispersions undergo phase separation giving rise to different kinds of lipidaggregates (lamellar, micellar and cylindrical rods). Acknowledgement The author is thankful to the Canadian Fellowship Committee for financial support.

Commonwealth

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References Arnon, D. I., Allen, M. B. and Whatley, F. R. (1956) Biochim. Biophys. Acta, 20, 449. Bangham A. D., Hill, Μ. W. and Miller, Ν. G. A. (1974) in Methods in membrane biology (ed. E. D. Koran) (New York: Plenum) vol. 1, p. 1 Benson, A. A. (1966) J. Am, Oil Chem. Soc., 43, 265. Benson A. A. (1971) in Structure and function of chloroplasts (ed. M. Gibbs) (Berlin, New York: SpringerVerlag) p. 129. Bligh, E. C. and Dyer, W. J. (1959) Can. J. Biochem. Physiol.,37, 911 Huang, C-H. (1969) Biochemistry, 6, 344. Kates, M. (1970) Adv. Lipid Res., 8, 225. Kimelberg H. K. (1977) in Dynamic aspects of cell surface organisation. Cell surface reviews (eds G. Poste and G L. Nicolson) (Amsterdam, New York: North Holland) p. 205. Kreutz, W. (1966) In Biochemistry of chloroplasts (ed. T. W. Goodwin) (London, New York: Academic press vol. 1, p. 83. Larsson, K. and Puang-Ngern, S. (1979) in Advances in biochemistry and physiology of plant lipids (eds L.–A. Appelqurst and C. Liljenberg) (New York: Elsevier/North Holland Biomedical Press) p. 27. Lucy, J. A. and Glauert, A. M. (1964) J. Mol. Biol., 8, 727 Murphy, D. J. (1986) Biochim. Biophys. Acta, 864, 33, YashRoy, R. C. (1987a) Indian J. Biochem. Biophys., 24, 177. YashRoy, R. C. (1987b) J. Biochem. Biophys. Methods, 15, 229. YashRoy, R. C. (1990) J. Biochem. Biophys. Methods, (in press).