J. Biosci., Vol. 15, Number 4, December 1990, pp. 281-288. © Printed in India.
Magnetic resonance studies of dynamic organisation of lipids in chloro-
R. C. YASHROY*
Biology Department, Carleton University and National Research Council, Ottawa,
*Present address: Electron Microscopy and Instrumentation Section, A. N. Division
Buildings, Indian Veterinary Research Institute, Izatnagar 243 122, India
MS received 20 July 1990
Abstract. Spinach chloroplast membranes and aqueous dispersions of their extracted
lipids have been studied by spin label (stearic acid) electron spin resonance and carbon-13
nuclear magnetic resonance techniques. Combined with electron microscope studies, first
systematic evidence is found for the existence of a dynamic lipid-bilayer structure in the
Keywords. Chloroplast; membranes; lipids; nuclear magnetic resonance; electron spin
It is generally believed that the thylakoid membranes of the plant chloroplasts have
a fluid lipid-bilayer structure interacting with membranous proteins (Curatolo,
1987). Curatolo (1987), however has credited this viewpoint to Quinn and Williams
(1983), who in turn have stated that it is difficult to reconcile this view due to the
fact that the predominant membrane lipid, monogalactosyl diacylglycerol (MGDG)
normally takes up hexagonal-II structure when dispersed alone in water and
furthermore it forms inverted lipid micellar structure when dispersed together-with
the second most abundant lipid, digalactosyl diacylglycerol (DGDG), of the
Unlike most other biological membranes which are rich in phospholipids, the
chloroplast thylakoid membranes contain little phospholipid (Kates et al., 1970).
Despite the fact that MGDG, the major constituent lipid of chloroplast membranes,
does not form an aqueous lipid-bilayer structure (Larsson and Puang-Ngern, 1979),
the other lipid constituents such as DGDG, sulphaquinovosyldiacylglycerol and
phospholipids may well be responsible for the formation of a lipid-bilayer structure
which can serve as a suitable matrix of the thylakoid membrane in which the
proteins are embedded (Murphy, 1986; Van Gurp et al. 1988; Anderson, 1975).
Absence of studies on dynamic state of lipid molecules in membranes led Weier and
Benson (1966) to propose a lipid-nonbilayer structure for thylakoid membranes.
The present investigation, therefore, was undertaken to systematically study the
dynamic organisation of the chloroplast photosynthetic membranes employing spin
label electron spin resonance (ESR) and carbon-13 nuclear magnetic resonance (13C-
NMR) spectroscopy. Together-with earlier studies (YashRoy, 1980, 1990a),
convincing evidence has been gathered suggesting the existence of a lipid-bilayer
structure in the chloroplast membranes.
Abbreviations used: MGDG, Monogalactosyl diacylglycerol; DGDG, digalactosyl diacylglycerol; ESR,
electron spin resonance; 13C-NMR, carbon-13 nuclear magnetic resonance; TMS, tetramethylsilane.
Materials and methods
The chloroplast membranes prepared (Arnon et al., 1956) from fresh spinach leaves
were vortexed, under dark conditions, with a dried film of stearic acid spin label for
a satisfactory labelling of the membranes in about 20 min. An aliquot of lipids
extracted (Nichols, 1963) from the chloroplast membranes (finally dissolved in
chloroform) was mixed with the spin label (dissolved in chloroform). The mixture
was dried under a stream of nitrogen gas and subsequently vortexed with water for
about 20 min under dark conditions for use as model membranes in the ESR
studies. The molar ratio of chlorophyll (Vernon, 1960)-to-spin label was kept 20:1
for all ESR preparations. Vortexed lipid-water dispersions made from the lipid
extract of the chloroplast membranes were also studied by EM after negative
staining (Lucy and Glauert, 1964; YashRoy, 1990a).
Spin labels (figure 1) employed for these studies contained a nitroxide (doxyl)
Figure 1. Diagramamatic representation of stearic acid labels showing location of the
nitroxyl (doxyl) moiety at 5, 7, 9, 12, 13, 14 and 16 carbons with respect to carboxyl group.
Chloroplast membrane lipid organisation 283
moiety at 5,7,9,12,13,14, or 16th carbon from the carboxyl group of the stearic acid
molecule. All ESR spectra were taken under dark conditions at 37°C on the Varian
E-9 spectrometer equipped with variable temperature (±0·2°C) control unit. The
magnetic field used was 3232 G with scan time of 4 min, microwave power of 5 mW,
scan range of 100 G, time constant of 0·3 or 1·0 s and frequency of 9 GHz. In many
instances, the maxima and minima of the hyperfine splittings were resolved by
increasing the receiver gain about 10-fold. The rotational correlation times and
order parameters for ESR studies were also calculated (Cannon et al., 1975).
Natural abundance proton-decoupled 13C-NMR spectra of chloroplast membranes
(YashRoy 1990b) (chlorophyll content, 5mg/ml) suspended in deuterated water con-
taining 0·2 Μ NaCl and 5 mM MgCl2 and sonicated lipid-D2O dispersions were
obtained with Varian XL-100 or Bruker CXP-300 FT NMR spectrometer equipped
with a variable temperature (±0·2°C) control unit. The spectra were collected with
field frequency lock on deuterium of solvent or external fluorine; the latter was found
specially useful for viscous membranes samples. An external tetramethylsilane
(TMS) was used as the standard reference. The radiofrequency pulse angle of 45°
and acquisition time of 0·4 s were used to obtain the Fourier transform 13C-NMR
spectra of chloroplast membranes and sonicated lipid microvesicles. For lipids
dissolved in organic solvents such as deutrated chloroform (CDC13) or methanol
(CDOD), the radiofrequency pulse angle used was 22·5° and the acquisition time
was 0·8 to 1·0 s.
Results and discussion
Inspection of ESR spectra (figure 2, top-to-bottom) of the spin labelled chloroplast
membranes reveals changes in line-shape, line-width and hyperfine splittings,
signifying gradual increase in degree of sub-molecular motions of lipid environment
from 5-doxyl to the 16-doxyl spin label positions. For example, 5-line spectra of the
membrane-incorporated 5,7 and 9-doxyl labels gradually turn into 3-line spectra for
12 to 16-doxyl labels. Also, the line-widths (note the central line) become
increasingly narrow in going from 7 to 16-doxyl spin labelled membranes. These
variations in the spectra demonstrate existence of a fluidity of flexibility gradient in
terms of increasing degree of motions in the segments of the fatty-acyl chains from
near lipid headgroups to the terminal (fatty-acyl) methyl groups in the chloroplast
Model membranes prepared by aqueous dispersion of lipids extracted from the
chloroplast membranes, also reveal a similar flexibility gradient. Figure 3 shows the
variations in linewidth, Wo (G) of the central spectral line of the ESR spectra of the
spin-labelled chloroplast membranes and model membranes made from their lipid-
extract. The observation of exceedingly shorter linewidths in the order of 9,12,14
and 16-doxyl positions of the labels incorporated in the two membrane systems
signifies the existence of progressively increasing rates of segmental motions
towards the terminal methyl groups of the lipid fatty-acyl chains. The similarity of
this flexibility gradient between the chloroplast membranes and model membranes
made from their lipid-extract indicates a similarity in the organization of the lipids
in these two membrane systems. Interestingly, both in chloroplast membranes and
their extracted aqueous lipid dispersions there is no significant decrease in
linewidth, Wo from 5 to 9-doxyl positions signifying persistence of a high degree of
Figure 2. 9-GHz ESR spectra of the chloroplast membranes labelled with (from top to
bottom) 5, 7, 9, 12, 13, 14 and 16-doxyl stearic acid spin labels. All the spectra were taken at
370C, in dark with the chloroplast membranes suspended in 0·2 Μ NaCl containing 5mM
MgCl2. The spectrometer settings were: magnetic field, 3232 G; time constant, 1 s; scan
time, 4 min; microwave power, 10 mw; and modulation amplitude of 1 G.
motional restriction of fatty-acyl chains in this region of the two membranes, This
characteristic differentiates the flexibility gradient of a lipid bilayer from that of
hexagonal phase, Hll as inferable from the observations made by Sternin et al,
A flexibility gradient such as this observed here is known to exist in other
biological (excluding chloroplast) membranes viz., mitochondrial (Vignais et al.,
1975), sarcoplasmic reticular (Seelig and Hasseilbach, 1971) and plasma (Rottem
et al, 1970) membranes in addition to many model lipid bilayer membrane systems
(McConnell and McFarland, 1972). This has been explained as due to the packing
of the lipid molecules in a bilayer, with the lipid headgroups anchored at the
lipid-water interface (McConnell and McFarland, 1972). Figure 4 shows
multilamellar structures formed by the aqueous lipid-extract of the chloroplast
membranes. These structures correspond to the multi-bilayers (liposomes) formed
by many aqueous lipid systems that constitute the basis of the lipid-bilayer model
for the biological membranes (Bangham, 1968; Mühlethaler et al, 1965; YashRoy,
Chloroplast membrane lipid organisation 285
Figure 3. Variation of linewidth, Wo in G of the central line of the ESR spectra recorded
in dark from stearic acid spin labelled chloroplast membranes (▲) and their extracted
aqueous lipid dispersions (●) with the position of the doxyl moiety (5, 7, 9, 12, 13, 14 and
16) on the stearic acid spin label (SASL).
Figure 4. Election micrograph of the aqueous dispersion of the lipids extracted from the
chloroplast membranes after staining with phosphotungstic acid (× 190,000).
The above observations of the flexibility gradient are also supported by
NMR studies of the chloroplast membranes and their sonicated aqueous lipid
dispersions. Majority of
membranes can be assigned to carbons of linolenic acid (YashRoy, 1987a,b),
Figure 5 summarizes the observations on the NMR line-widths of resonances
13C-NMR resonances arising from the chloroplast
Figure 5. An overall view of the selected resonances of
extracted from chloroplast membranes. The left hand side shows a sketch of a lipid
molecule having a globular headgroup (hg) and single (for clarity) extended fatty acyl chain
with one carbonyl group and 3 double bonds. (A) Lipids in deuterated methanol, depicting
(top-to-bottom) largely unaggregated (lipid) forms (top) and
13C-NMR spectra of lipids
corresponding lipid headgroups, fatty acyl carbonyl, –(CH2)n, –C=C– and terminal
methyl groups. (B) Lipids in deuterated chloroform and (C) lipids dispersed in deuterated
water. Note the changes in linewidths from top-to-bottom of each of the lipid preparations
(A), (B) and (C).
Chloroplast membrane lipid organisation 287
corresponding to various segments of fatty-acyl chains e.g, those located proximal
to lipid head-groups, fatty-acyl methyl terminals and intermediate positions. The
left-hand-side shows the model diagram of a lipid molecule with a ‘globular’
headgroup and having a single (for clarity) fatty-acyl chain containing a C=O
group and 3 double bonds (representing linolenic fatty-acyl chain found most
abundantly in chloroplast membranes). Figure 5A represents some select 13C-NMR
resonances arising from the extracted lipids (of chloroplast membranes) dissolved in
CD3OD wherein these lipids, by and large, do not form any specific aggregates.
This is clearly evident from sharp line-widths of all the
head-group, C=O group, fatty-acyl chain (segment) or terminal methyl group.
When the same lipids are dissolved in CDC13, they form aggregates with
headgroups packed-in and tails fanning-but 'freely'. This is revealed by the
NMR linewidths (figure 5B) which clearly show broadening of headgroup and
C=Ο group resonances and yet retaining narrowness of fatty-acyl chain
resonances. When dispersed in D2O as unilamellar sonicated microvesicles
(YashRoy 1990a) (figure 5C), these lipids show variedly restricted movement
consistent with a flexibility gradient, similar to that observed by 2H-NMR and spin
label ESR studies of lipid-bilayer systems (Sternin et al, 1988). The line-broadening
effect is maximum on headgroups and C=Ο groups and somewhat less on the
-(CH2)n and HC = CH segments and the least on terminal-CH3 group of the fatty-
acyl chains. A similar order of restricted mobility of carbons from headgroups towards
the terminal methyl groups of the fatty-acyl chains is noticeable from
spectrum of chloroplast membranes (figure 6). In essence, the fact that the electron
Figure 6. 75-MHz natural abundance
membranes at 30°C. The membranes were suspended in 0·2 Μ sodium chloride containing
5 mM magnesium chloride in deutrated water. Chemical shift in parts permillion (ppm) in
reference to external TMS (not shown).
microscopically observable liposomal multibilayer structures (figure 4) formed by
aqueous dispersion of the lipids extracted from the chloroplast membranes depict a
flexibility gradient closely parelleling the flexibility gradient revealed by the native
chloroplast membranes (especially notable from ESR spectral line-widths, as in
figure 3) strongly suggests that the dynamic organisation of lipids in the chloroplast
membranes is largely bilayer.
13C-NMR spectrum of spinach chloroplast
288 Download full-text
The author thanks the Canadian Commonwealth Scholarship and Fellowship
Committee for financial support.
Anderson, J. Μ. (3975) Biochim. Biophys. Acta, 416, 191.
Arnon, D. I., Allen, M. B. and Whutley, F. R. (1956) Biochim. Biophys. Acta, 20, 449.
Bangham, A. D. (1968) Prog. Biophys. Mol. Biol., 18, 29.
Cannon, Β., Polnaszek, C. F., Butler, K. W., Ericksson, L. E. G. and Smith, I. C. P. (1975) Arch.
Biochem. Biophys., 167, 505.
Curatolo, W. (1987) Biochim. Biophys. Acta, 906, 137.
Kates, M., Paoletti, R. and Kritchevsky, D. (1970) Adv. Lipid. Res., 8, 225.
Larsson, K. and Puang-Ngern, S. (1979) in Advances lipid research (eds L.A. Appelquist and
C. Liljenberg) (Amsterdam: Elsvier/North Holland Biomedical Press) p. 27.
Lucy, J. A. and Glauert, A. M. (1964) J. Mol. Biol., 8, 727.
McConnell, Η. Μ. and McFarland, Β. G. (1972) Ann. Ν. Υ. Acad. Sci., 195, 207.
Mühlethaler, K„ Moor, H. and Szarkowski, J. W. (1965) Planta, 67, 305.
Murphy, D. J. (1986) Biochim. Biophys. Acta, 864, 33.
Nichols, Β. W. (1963) Biochim. Biophys. Acta, 70, 417.
Quin, P. J. and Williams, W. P. (1983) Biochim. Biophys. Acta, 737, 223.
Rottem, S., Hubbel, W. L., Hayiluk, L. and McConnell, M. M. (1970) Biochim. Biophys. Acta, 219, 104.
Seelig, J. and Hasselbach, W. (1971) Eur. J. Biochem., 21, 17.
Sternin, E., Fine, Β., Bloom, Μ., Tilock, C. P. S., Wong, Κ. F. and Cullis, P. R. (1988) Biophys. J., 54,
Van Gurp, M., Van Ginkel, G. and Levine, Y. K. (1988) Biochim. Biophys. Acta, 938, 71.
Vernon, L. P. (1960) Anal. Chem., 32, 1144.
Vignais, P. M., Devaux, P. and Colbeau, A. (1975) in Biomembranes-lipids, proteins and receptors (eds
R. M. Burton and L. Packer) (Missouri: BI Science Publications) p. 318.
Weier, T. E. and Benson, AAA. (1966) in Biochemistry of chloroplasts (ed. T. W. Goodwin) (New York,
London: Academic Press) vol. 1, p, 91.
YashRoy, R. C. (1980) Fluidity and lipid protein interactions in the chloroplast membranes: A. magnetic
resonance study; Ph. D. thesis, Carleton University, Ottawa, Canada
YashRoy, R. C. (1987a) Indian J. Biochem. Biophys., 24, 177.
YashRoy, R. C. (1987b) J. Biochem. Biophys. Methods, 15, 229,
YashRoy, R. C. (1990a) J. Biosci., 15, 93.
YashRoy, R. C. (1990b) J. Biochem. Biophys. Methods, 20, 353.