Changes in membrane lipid structure of illuminated chloroplasts— Studies with spin-labeled and freeze-fractured membranes

ARCHIVES OF BIOCHEMISTRY AND BIOPHYSICS 160, 90 99

(1974)

Changes in Membrane Lipid Structure of Illuminated Chloroplasts-Studies with Spin-Labeled and Freeze-Fractured Membranes ~ J. T O R R E S - P E R E I R A f R. M E H L H O R N , A. D. K E I T H ,

AND

LESTER PACKER a

Department of Physiology-Anatomy, University of California, Berkeley, California 94720 and Department of Biophysics, Pennsylvania State University, University Park, Pennsylvania 16802 Received August 27, 1973 A spin-labeled hydrocarbon molecule having a lipid-water solubility suchthat both polar and apolar signals could be observed in chloroplast suspensions was used to monitor effects of light excitation. In glutaraldehyde-fixed chloroplasts, effects of light-induced swelling and shrinking were ascribed to a lipid structural change comparable to a temperature shift of 2~ C. Under similar conditions freeze-fracture electron microscopy revealed changes in average sizes and clustering of membrane particles. Hence, the particle-free areas of the membrane interior, presumed to be the lipid domains, undergo changes in organization when chloroplasts are illuminated. The chloroplast structure has unique structural and functional features in that photoexcitation readily produces a variety of coincident structural and functional changes in the polylamellar thylakoid membrane array. T h a t the general functional states of chloroplasts under dark or light conditions relate to tile structural properties has been demonstrated by light-scattering and lighttransmission studies (1, 2), electron micrographs taken from chloroplast samples glutaraldehyde-fixed under light or dark conditions (3, 4), ANS 4 fluorescence under comi 1 The authors thank A. Bearden for the use o his EPR spectrometer and Ann Y.-I. Wang for assistance in the freeze-fracture electron microscopy experiments. This research was supported by the National Science Foundation (GB 20951), the Atomic Energy Commission and by a fellowship (to J.T.-P.) from the University of Luanda, Angola, Portugal. 2 On leave from the Department of Biology, Faculty of Sciences, University of Luanda, Angola, Portugal. a Please address reprint requests to Lester Packer, Department of Physiology-Anatomy,University of California, Berkeley, California 94720. 4 Abbreviations used: ANS, anilinonaphthalene sulfonic acid; PMS, phenazine methosulphate; 6Nll, 2,2-dimethyl-5,5-dipentyl-N-oxyloxazoli-

parable conditions (4), the distribution of membrane particles seen in freeze-fractured membranes taken from samples under dark and light conditions (5), and a report employing lipid-soluble spin labels (6). Although the above studies establish that structural changes in the chloroplasts at the organelle and membrane level are induced by light stimulation, they do not establish the extent of involvement of membrane lipids in the structural and functional changes in grana membranes. I t was previously shown that a lightdependent reduction of the nitroxide 2,2, 5,5 - tetramethyl - 3 - carbamid - pyrroline 1-N-oxyl occurred in whole edls of Chlamydomonas reinhardti Dang (7). This reduction was reversible, and dark conditions restored the spin signal. The present investigation was designed to reveal whether lipid physical-state changes accompany illumination of chloroplasts. Evidine; TES, N-tris (hydroxymethyl) methyl-2amino-ethane sulfonic acid; TEMPOL, 2,2,6,6tetramethylpiperidine-N-oxyl; A12NS, acid form of 12-nitroxide stearate; BSA, bovine serunl albumin ; A4NS, acid form of 4-nitroxide stearate ; FCCP, carbonyl cyanide p-trifluoromethoxyphenyl-hydrazone. 90

Copyright 9 1974 by Academic Press, Inc. All rights of reproduction in any form reserved.

MEMBRANE STRUCTURE OF CHLOROPLASTS de nc e for l i g h t - i n d u c e d s t r u c t u r a l r e o r g a n i z a t i o n of lipids in t h e m e m b r a n e s was obt a i n e d b y studies e m p l o y i n g h y d r o c a r b o n spin labels an d is s u p p o r t e d b y freeze-fracture electron microscopy revealing microm o r p h o l o g i c a l changes w h i c h are c o i n c i d e n t w i t h some p h y s i o l o g i c a l alterations. MATERIALS AND METHODS

Preparations. Class II chloroplasts (without outer membranes) were prepared from spinach (Spinacia oleracea L.) leaves, as previously described (4), and washed twice in the isolation medium by centrifugation at 1500g $or 6 min. Chlorophyll concentrations were determined as previously reported (8). Assay conditions. Experiments with the abovementioned type of preparation were carried out at 25~ C in a Japan Electron Optics Laboratory X-brand E P R spectrometer, model JES-ME-IX, equipped with a variable temperature-control unit, with an approximate temperature accuracy of =t=0.5~ C. The reaction mixture contained 0.1 M NaC1 or NaAe, 50 ~M PMSL and chloroplasts (1000 ~g chlorophyll/ml), pH 6.5. Spin-label concentrations are given in figure legends, unless otherwise stated, and were introduced into the thylakoid membranes in ethanol, so that the final ethanol concentration was always less than 1%. The concentration of ethanol used had no detectable effects in chloroplasts. Red actinic light was 600-700 nm (Eastman Kodak interference filter No. 2412). Light intensity was adjusted to 225 ftcandles using a Gossen 7.65-1287 photometer. Light was transmitted to the capillary tube of the E P R spectrometer by quartz fiber optics. Chloroplasts were also fixed with glutaraldehyde under conditions leading to shrinkage or swelling of the organelles so that after fixation no further volume changes were detected. Chloroplasts equivalent to 1 mg of chlorophyll were suspended in either 0.1 M NaAe or NaC1 plus 50 ~M pyocyanine perchlorate, pH 6.8 (total volume of the suspension, 3 ml). The suspension was stirred and kept at 25~ by using a thermostatic device. Illumination with white light at a saturating intensity was performed during 5 min and fixation was done with 500 mM glutaraldehyde (Polysciences-electron microscope grade, pH 7.0) for 2 min. Glutaraldehyde was immediately removed from the chloroplast suspension by eentrifugation and the chloroplasts were washed twice by centrifugation at 1500g for 6 rain and resuspended in ~he isolation medium. The same procedure was employed for control chloroplasts kept in the dark. Spin-label experiments were subsequently conducted in the dark so that light-induced spin

91

reduction could safely be ruled out as contributing to partitioning changes. Centrifugation experiments of spin-labeled chloroplasts showed that the hydrophobic signal remained with the pellet and is associated with the chloroplast lipids while the polar signal remained with the aqueous phase and is not bound to the chloroplasts. For this reason the degree of partitioning is quite sensitive to the concentration of chloroplast lipids in the suspension. Accordingly, the chlorophyll concentration, assumed to be proportional to the lipid concentration, was carefully controlled in comparisons of partitioning involving different chloroplast treatments. Proton transport and resulting gradients of H + activity across membranes are now recognized to be integrally related to phosphorylation and ion transport in chloroplasts (9). To estimate energy coupling in chloroplast membranes when the spin label 6 N l P was present, measurements of pH changes in the external medium using a glass electrode (10), the light-induced quenching of the intramembrane pH indicator atebrin (11, 12), and ferricyanide reduction (13) have been employed. With these methods we have evaluated the effect of the spin label 6Nll on energy-coupling parameters. Light-induced pH changes were detected with a pH-recording device as previously described (4). Cyclic photophosphorylation was assayed at 20~ C and pH 7.7, using 5-ml reaction mixtures containing 250 mM sucrose, 10 m~ NaC1, 3 mM MgC12,2 mM KH~PO4, 100 ~M ADP, 6 ~ pyocyanine, chloroplasts equivalent to 146 tLg of chlorophyll, and several 6Nll concentrations. The initial rapid burst of H + uptake was ignored in the calculations (14, 15). In the absence of phosphorylation conditions, proton movements were assayed at 20~ C and pH 6.3, using 5-ml reaction mixtures consisting of 100 mM NaC1, 6 ~ pyocyanine, chloroplasts equivalent to 90 ~g of chlorophyll, and different 6 N l l concentrations. Calibrations were made with standard HC1. Atebrin fluorescence changes were investigated using a front-face fluorescence speetrophotometer (12) with a thermostated eylindricM glass cuvette constructed with a single flat window, on which the ultraviolet excitation light (420 nm) was collimated (angle 60~ Fluorescence emission was detected at the surface of the same window, using emission filters Corning 3-72 and 9782, and a green filter (480-605 nm). The euvette was illuminated by a cooled quartz-iodine lamp (Westinghouse 45 W), whose light was filtered by a Corning CS1-69 heat filter and a red filter cutting off below 63O nm. This chloroplast excitation light was passed through a mechanical shutter. The contents of the euvette were stirred on the top by a

92

TORRES-PEREIRA ET AL.

Teflon stirrer and additions were made with roplast suspensions with 29.2 t~g ehloromicroliter syringes. The device included a d.c. phyll/ml, showed that at 6.7 and 13.5 ua.t amplifier and a galvanometric recorder (Texas 6 N l l , the rate of A T P synthesis was, respecInstruments Incorporated, Recti/Riter 0-10). tively, 47.61 and 24.04 ~moles of A T P proAtebrin fluorescence changes were assayed at 20~C, duced/mg chlorophyll/hr, as compared to a using 1.5-ml reaction mixtures containing 200 mM sucrose, 20 mM NaC1, 10 mM TES 4 pH 8.0, control of 93.30. Atebrin fluorescence changes, studied in 4 ~Mpyocyanine, 2 ~ atebrin, chloroplasts equivalent to 36 t~g of chlorophyll, and several 6Nll 24 ~g chlorophyll/ml mixtures, showed that at 22.3 and 44.6 ~,sr 6 N l l , the half-times of concentrations. Ferricyanide reduction was monitored at 420 fluorescence quenching and recovery, and nm and room temperature using a Beckman DB the percentage of total quenching were, respeetrophotometer connected to a Honeywell spectively, 7.0 see, 21.0 see, and 88.3 %, and Electronik 19 recorder. The 3-ml reaction mixtures 14.5 see, 9.5 see, and 53.0%, relative to a contained 100 mM sucrose, 5 mM MgCI~ , 10 mM 4.0 see, 34.0 see, and 92.5 % control. TES pH 8.3,250 ~Mpotassium ferricyanide, chloroHill reaction activity, as measured by plasts equivalent to 10 tLg of chlorophyll, and differricyanide reduction in 3.34 ~g ehloroferent 6Nll concentrations. Chloroplasts were phyll/ml chloroplast suspensions, showed illuminated by red light (600-700 nm, 250 ftat 33.5 and 67 ~ 6 N l l , respectively, 41.18 candles). Freeze-fracture electron microscopy. Class II and 20.53 tmloles ferricyanide reduced/rag chloroplasts (0.34 mg ehlorophyll/ml) in 0.1 M chlorophyll/hr, with reference to a 82.37 Na-aeetate plus 50 t~M pyocyaniae perchlorate, at control. pH 6.8 and 25~ C, were illuminated with white Summarizing these data, bulk pH changes, light at a saturating intensity during 5 rain. A photophosohorylation, fluorescence change drop of the sample was rapidly frozen in Freon 22 kinetics, and ferricyanide reduction were under illumination and replicas obtained accord- significantly affected (approximately ~ of ingtoWrigglesworth et al. (16) on a Balzers freezeetching apparatus. A similar procedure was used control activity) by 0.75, 0.23, 0.92, and for control samples kept under dark conditions. A 10.0 ~moles of 6 N l l per ~mole of chlorophyll. stage which accepts four different samples was Actual spin-label concentrations in the used. Specimens were fractured at -110 ~ C and chloroplasts would be lower in the control immediately replicated. Replicas were examined experiments than implied by the above using a Siemens Elmiscope Ia electron microscope. figures because one expects an increase in All photographs presented are positives, i.e., the aqueous population of a partitioning label were processed so as to show shadows in white. at the higher water to lipid ratios. NevertheShadow direction is indicated by arrows on the less, under our E P R experimental conditions, photographs. it could be safely assumed that the spin label did not significantly affect the above RESULTS mentioned chloroplast functions. T h a t is, Effect of 6 N l l on Functional Parameters our spin-labeling experiments with F C C P 4 Controls with 6 N l l were conducted to were conducted at a 6 N l l concentration evaluate its effect on certain chloroplast of 0.04 ~mole of 6 N l l per #mole of chlofunctional parameters, prior to E P R ex- rophyll and the F C C P effect was quite periments. Unfortunately, it is not techni- clear. However, it is apparent that 6N 11 is a cally feasible to use in these controls chlo- membrane perturber, perhaps an inhibitor. rophyll concentrations as high as those used Reduction of Spin Labels by Chloroplasts in the E P R experiments. The addition of a variety of spin labels Bulk p H changes, detected in chloroplast suspensions with 18 ~g chlorophyll/ml, to chloroplast preparations under dark conshowed that at 13.5 and 40.0 u~ 6 N l l , the ditions results in a stable spin signal with no extent of the proton changes was, respec- detectable loss of signal over a period of 30 tively, 0.42 and 0.21 #eq H + / m g chlorophyll, min or more. Light exposure of the chlororelative to a 0.81 peq H + / m g chlorophyll plasts results in relatively rapid spin loss. control. A return to dark conditions results in a Phosphorylation experiments, using chlo- return of spin at about ~ the spin-loss rate.

MEMBRANE STRUCTURE OF CHLOROPLASTS ]?his observation was also made several years ago by Weaver and Chon and was attributed to a light-dependent reversible chemical reduction of the paramagnetie nitroxyl group going to a diamagnetic hydroxyl amine (7). The exact kinetics depend on the spin-label structure, spinlabel localization, concentration of spin label and chloroplast membranes, temperature, and other specific conditions used. A typical ease with the concentration of 6Nll held at 3 X 10-a ~ results in about 30 % loss of signal in about 10 min. Return of the signal in the dark requires about, 40 min. Spin labels are sensitive to chemical reduction by a variety of hydrogen donors such as ascorbate and sulfhydryl groups. Reduction of TEMPOL 4 by aseorbate in water and the subsequent recovery of reduced TEMPOL by ether extraction yields a reduced product that is reoxidized when added to dark-adapted chloroplasts. Light stimulation of the preparation causes the oxidation process to accelerate. The addition of competitive electron carriers, such as pyoeyanine or PMS, in small concentrations, results in acceleration of the light-dependent reduction of 6Nll and of the subsequent recovery of spin in the dark. Site of spin label reduction. In studies of partitioning changes the possibility exists that polar and hydrocarbon signals will be reduced to a different extent. To assess the importance of such differential reduction we have studied reduction of labels localized in different domains of the chloroplast prepa-. ration. The spin-labeled fatty acid, A12NS 4, forms a strongly intereaeting complex with BSAt at a molar ratio of one and at this molar ratio does not exchange with the chloroplasts. The spectrum of this complex is characteristic of a highly immobilized nitroxide molecule. This signal is readily reduced by aseorbate in water. When BSA labeled with A12NS in this manner is added to a chloroplast, suspension, no signal reduction is observed under illumination; however, label reduction is readily achieved with the addition of PMS. This reduction is reversible in the dark. Further studies of nitroxide reduction sites were conducted in chloroplasts with-

93

out BSA using A12NS and A4NS 4(having the nitroxide group at carbon 4 of stearic acid). These same spin-labeled fatty acids have been used in a variety of studies employing different biomembranes and model membrane systems which have established that they orient normal to the bilayer or merebrane surface with the carboxyl group localized at the hydrated polar:hydrocarbon interface (17). Rates of reduction in the absence of PMS were slightly different, A4NS being reduced about 30 % more quickly than A12NS. These data suggest that the spin-label reduction occurs within the chloroplast membranes, perhaps at the polar-apolar interface; however, the eyelie cofaetor PMS, as seen with BSA, extends this domain of reduction to the exterior aqueous phase of the chloroplasts.

Line-shape changes in unfixed chloroplasts. ESR spectra of chloroplasts containing 3 • 10-5 ~I 6 N l l are shown in Fig. 1. Spectral features arising from the label in polar and hydrocarbon environments are partially resolved in the high field line and are designated by h - l P and h-lH, respectively. Measurements of these parameters were used to monitor line-shape changes resulting from various treatments of the chloroplasts. The population of 6Nll dissolved in the membrane hydrocarbon regions yields three differentially broadened lines having the general properties of line broadening due to restriction of spin-label motion. There may also be line-broadening contributions from heterogeneity effects when some part of the spin-label population is exposed to zones of intermediate polarity thereby giving rise to some mixed g values and hyperfine coupling values. The possibility of some spin labelspin label interactions is also not ruled out. Moreover, the polar and apolar absorption lines overlap so that quantification of the label concentration in these two domains is not possible. In view of these considerations the high field line shape may vary as a result of spin-signal reduction alone and changes in the relative values of h-lH and h-lP do not necessarily imply that the relative solubility of the spin label in polar and hydrophobic regions has changed. Line-shape changes coneomitant with sig-

94

TORRES-PEREIRA ET AL.

FIG. 1. Electron spin resonance spectra of 3 X 10-5 M 6Nll spin label dispersed in chloroplasts fixed with glutaraldehyde while suspended in sodium acetate medium. Upper and lower spectra resulted from fixation carried out under illumination and in the dark, respectively. In this case, the spin label was introduced after fixation, but similar results are obtained when it is introduced before fixation. Other conditions of the experiment are described in the text. nal loss of 6 N l l , probably due to reduction, can be monitored in terms of the ratio h - l P / h - l H plotted against a measure of the spin-label concentration in one of the environments, e.g., polar or apolar. Small fluctuations in apparent signal intensities between different samples were unavoidable in these experiments. T o allow for this possibility, line-shape changes occurring during the course of light-induced spin reduction are plotted as h - l P / h - l H versus log h-lP. The high field aqueous component of 6 N l l is the best resolved of any line element, has a constant line width (W-1P) under the conditions used, is localized in the aqueous medium, and, consequently, h - l P / i n s t r u ment gain is a sensitive measure of the concentration of 6 N l l in a given volume of aqueous medium. Scale factors between spectra of different samples will thus produce only horizontal shifts of reduction curves without affecting their shapes. 9 Upon illumination of chloroplasts by red light, h - l P decreases, and there is a slight increase of h - l H . These spectral changes are stabilized after a period of about 3 min.

The time dependence of line shape changes of control and F C C P uncoupled chloroplasts is compared in Fig. 2. Two points are noteworthy: (a) the rate and extent of reduction are greater in the uncoupled chloroplasts, and (b) the reduction curves are similar in shape. I t appears that the lipid solubility of the spin label is not significantly affected b y this uncoupler during light excitation.

Partitioning Changes in Fixed Chloroplasts In glutaraldehyde-fixed mitochondria where no spin reduction occurs, the h - l P / h - l H ratio depends on the oscillatory state of the mitochondria at the time of fixation (18). Similarly, in glutaraldehyde fixed chloroplasts spin reduction in the dark does not occur over a period of hours. Thus, spectra of chloroplasts subjected to different treatments can be compared directly. Glutaraldehyde is assumed to immobilize amine containing proteins and lipids through crosslinking reac.tions between primarily aqueous amines. Since chloroplasts are reported to conrain little or no phosphatidyl ethanolamine (19) it was assumed that glutaraldehyde did

MEMBRANE STRUCTURE OF CHLOROPLASTS

% Z

CONTROL- ~ /

r

////

95

," / ,tD, ~I~ O~ D2

/ u-

k ///

I0

L,

+50 F~MFCCP Lz

2

/

P,

~ I~ k~

P,

0

9 ,6

I L7

f'.8

rl.9

I 2.0

Lo9 Polar Peok (hp)

FIG. 2. Line-shape modifications obtained during 60-sec intervals with chloroplasts suspended in sodium chloride medium plus or minus FCCP. 6Nll initial bulk concentration was 4 X 10-5 M. Spectra were recorded according to the sequence: Do, initial dark condition; L~, L~, L~, successive l-rain scans with a continuous red light intensity of 225 ft-c; D~ , D~, D~, D~, Ds , successive scans made in the dark, after the red light was turned off. (9 Control; (Q) plus 30 tLMFCCP. Other conditions as described in the text. -0.200 -

-0.400 3 6 . 0 % = o. ]~ -o~o0

27,0 o t " ~,.o..\

.~

-0,800

14

x

\ -I .000

N

%%

I 1,0~ i

-

3200

I

l

3300

I

I

3400 I l T x IO3

1

I

3500

FIG. 3. Arrhenius plot of 6Nll partitioning changes in glutaraldehyde-fixed chloroplasts suspended in sodium acetate medium. 6Nll was 3 X 10-5 M. Other conditions as described in the text. not affect lipids directly. However, membrane lipoprotein complexes generally manifest strong interactions and it seems plausible t h a t protein immobilization involves a

concomitant stabilization of associated lipid structure. Results of experiments with fixed chloroplasts are set out in Table I. Slight changes in spin-label partitioning result from experimental conditions which produce swelling (NaC1) and shrinking (NaAc) of the thylakoids (1, 2, 9). To assess the quantitative significance of partitioning changes observed in the glutaraldehyde-fixed chloroplasts we related our data to lipid structural changes resulting from a change of temperature. I n Fig. 3 partitioning changes of 6 N l l in glutaraldehyde-fixed chloroplasts are displayed against t e m p e r a t u r e on an Arrhenius plot. T h e discontinuity observed at 18 ~ C is similar to lipid-phase changes observed in other m e m b r a n e systems b y spin-labeling techniques (20). All other experiments described herein were carried out at 25 ~ C. At this temperature, changes in partitioning observed with 6 N l l in glutaraldehyde-fixed swollen and contracted chloroplasts are equivalent to a t e m p e r a t u r e increment of about 2~ Although temperature-dependent

TORRES-PEREIRA ET AL.

96

TABLE I LIGHT-INDUCED CHANGES IN CHLOROPLAST VOLUME AND IN THE PARTITIONING OF

6Nll IN

CHLOROPLAST MEMBRANES a

Chloroplast volumeb Sample

Chlorophylt (pg/ml)

1 2 3

700 1600 1800

Packed volume (%) 6Nll (~M)

NaAc dark

NaAc light

NaCI dark

NaCI light

3.5 11.5 12.0

1.5 3.5 3.2

3.7 13.5

4.8 17.0

6Nll partitioning Sample

h-1 HC/h-IP

Chlorophyll (~g/ml)

6Nll

700 1600 1800

15 30 30

NaAc dark

NaAc light

NaC1 dark

NaC1 light

0.480 • 0.0101 0.540 4. 0.0201 0.525 4. 0.0451 0.420 4- 0.03c 0.430 :t: 0.0201 0.485 4- 0.015 0.565~ 0.005 0.485 4- 0.02~ 0.515 • 0.0351 0.575 4- 0.0351

Chloroplasts were fixed with glutaraldehyde under conditions leading to shrinkage or swelling, a~ described in Materials and Methods--Assay Conditions. b Chlorocrit measurements were performed in samples centrifuged at ll,000g for 3 rain. and illumination-induced structural changes m a y be qualitatively different, this provides some estimate of the order of magnitude of the structural effects.

Freeze-Fracture Electron Microscopy Fracture faces of chloroplast thylakoid membranes were examined plus and minus light in the same manner as the spin-label analysis. Characteristic fracture faces are revealed in Fig. 4 according to the designations of Goodenaugh and Staehelin (21) which are comparable to other designations of Branton and P a r k (22). A fracture face containing numerous small particles (about 4000-5000/t~m 2) is easily recognized although these are difficult to analyze accurately with regard to size, density, and distribution. The B faces contain a smaller n u m b e r of larger particles and, therefore, facilitate an analysis of particle size and density in the stacked (B,) and unstacked (Bu) m e m b r a n e areas. Chloroplasts illuminated in the presence of a weak acid anion solution undergo a decrease in volume as shown in Table I. Under these conditions an approximate 25 %

thinning of a 'fused-pair" of membrane,~ occurs (4). Figure 4 and Table I I show that the particle density of the B s face analyzed over the entire area of the Bs face, does not change; however, there is a slight change ir the relative area of the m e m b r a n e covered b y particles. There is a clustering or aggregation of particles and in the areas where this occurs, particle densities increase 70 % upon illumination from 2000 to 3400/um 2. Concomitantly, the apparent size determined by approximate linear measurements of the particles decreases from 160 to 110 A. This estimated 32 % decrease in particle sizes is an approximation because analyses of theii dimensions do not take into account certain error due to the deposition of the thin layer of platinum which increases the shadow size. The data suggest t h a t the organization but not the size of particlefree areas, which are presumably the lipid areas, undergoes appreciable change upon illumination. DISCUSSION This investigation has established that light-induced changes in the organization of

MEMBRANE STRUCTURE OF CHLOROPLASTS

FIG. 4. Fracture faces of thylako]d membranes obtained from chloroplasts suspended in sodium acetate solutions m~nus und plus light. Other conditions are described in the text.

97

98

TORRES-PEREIRA ET AL.

membrane proteins and lipids accompany illumination of chloroplasts. A highly schematic diagram illustrating a possible explanation of spin-label partitioning and the examination of membrane particles seen by freeze fracture is shown in Fig. 5. Upon illumination in the presence of a weak acid anion solution, a reorganization of membrane particles, i.e., proteins or lipoproteins, occurs. There is a clustering of particles with an inTABLE II ]AGtVI-I,:DUCED CHANGES OF PARTICLE D I S T R I BU]IO'~ ON THE B s FRACTURE FACE OF C t t l , 0 R O P L A S T GRANA MEMBRANES a

Particle density/ tim 2

Dark Whole area Light W h o l e area Cluster area

Average particle diameter

(~_)

Area covered by particles

(%)

~000 •

100 160 •

1( 40.0 4- 4.0

2100 • 3400 •

100 [10 • 200 110 •

1( 2~.0 • 10132.3 •

3.0 4.0

These d a t a a r e from d a r k and light samples o b t a i n e d u n d e r conditions described in M a t e r i a l and M e t h o d s - - F r e e z e F r a c t u r e E l e c t r o n Microscopy. DARK- Na Acetate

W~ter. . . .

~

.....

crease in areas which are particle free. Undel these conditions there is also an increased solubility of our hydrocarbon spin-label. The spin label and freeze-fracture data presented here are consistent with lightinduced fluorescence changes of chlorophyll and ANS (4, 9, 23), photoactivated light scattering reflecting gross swelling or contraction (1, 2, 9), electron microscopic (3, 4), and X-ray data (24) showing light-induced membrane-thickness changes. The present data, however, establish that the membrane lipid matrix undergoes physical state changes with certain physiological alterations. Despite the evident large light-induced changes in molecular organization of the membrane revealed by conventional and freeze-fracture electron microscopy, and the response of external and internal fluorescent probes, the partitioning of lipids like 6 N l l is relatively unchanged. These results are interpreted to indicate that appreciable changes in protein aggregation do not result in similar amplitude changes [11 parts of the membrane into which 6Nll partitions. Experiments dealing with spin-labeled chloroplasts previously reported by Tzapin et al. (6), although interesting, are not directly comparable with results presented

& ~

.....

.embr~

.......

0-:-=g

~

........

a4

..... Polar Interfaces

.... Particleaggregation

----f~

solulbility of smallmolecules Modified lipid

Fro. 5. An i n t e r p r e t a t i o n of spin-label p a r t i t i o n i n g changes and r e o r g a n i z a t i o n of memb r a n e particles in chloroplasts upon i l l u m i n a t i o n i n a sodium acetate solution.

MEMBRANE STRUCTURE OF CHLOROPLASTS

10. CHANCE, B., Am) NIeH1MURA, M. (1967) in Methods in Enzymology (Estabrook, R. W., and Pullman, iV[. E., eds.), Vol. 10, p. 641, Academic Press, New York. 11. KRAAYENHOF, R. (1970) Fed. Eur. Bioehem. Soc. Letters 6, 161-165. 12. KRAAYENHOF,R., IZAWA,S., AND CHANCE,B. (1972) Planl Physiol. 50, 713 718. 13. WHATLEY, F. R., AND LOSADA,M. (1964) in Photophysiology (Giese, A. C., ed.), Vol. 1, p. 111, Academic Press, New York. 14. SCHWARTZ, M. (1968) Nature (London) 219, 915~19. 15. WEST, J., ANn PACKE~, L. (1970) Bioenergetics 1, 405-412. 16. WRIGGLESWORTH, J. M., PACKER, L., AND BRANTO~, D. (1970) Biochim. Biophys. Acta 205, 125-135. REFERENCES 17. MEHLHORN,R. J., AND KEITH, A. D. (1972) in DEAMER, D. W., CROFTS, A. R., AND PACKER, Membrane Molecular Biology (Fox, C. F., L. (1967) Biochim. Biophys. Acta 131, 81-96. and Keith, A. D., eds.), p. 192, Sinauer CROFTS, A. R., DEAMER, D. W., AND PACKER, Associates, Inc., Stamford, Connecticut. L. (1967) Biochim. Biophys. Acta 131, 97- 18. TINBERG, H. M., PACKER, L., AND KEIT~, 118. A. D. (1972) Biochim. Biophys. Acta283, 193MURAKAMI, S., AND PACKER, L. (1970) Plant 205. Physiol. 45, 289-299. 19. ALLEN, C. F., GooD, P., TROSPER, T., AND MURAKAMI, S., AND PACKER, L. (1970) J. PARK, R. B. (1972) Biochem. Biophys. Res. Cell Biol. 47, 332-351. Commun. 48, 907-913. WANe, A.Y.-I., AND PACKER, L. (1973) Bio- 20. RAISON, J. K., LYONS, J. M., MEHLHORN, chim. Biophys. Acta 305, 488-492. R. J., AND KEITH, A. D. (1971) J. Biol. TZAPIN, A. I., MOLOTKOVSKY, Y. G., GOLDChem. 246, 4036-4040. FIELD, M. G., AND DZJUBENKO, V. S. (1971) 21. GOODENOUGH,V. W., AND STAEHELIN, A. L. Eur. J. Biochem. 20, 218-224. (1971) J. Cell Biol. 48, 594-619. WEAVER, E. C., AND CHON, ~I. P. (1966) 22. BRANTON,D., AND PARK, R. (1967) J. UltraScience 153, 301-303. struct. Res. 19, 283-303. KIRK, J. T. O. (1968) Planta 78, 201-207. 23. MURAKAMI, S., AND P~CKER, L. (1971) Arch. Biochem. Biophys. 146, 337-347. MURAKAMI, S., TORREs-PEREIRA, J., AND PACKER, L. (1973) in Bioenergetics of Photo- 24. SADLER, D. M., LEFORT-TRAN, M., AND POUPmLE, M. (1973) Biochim. Biophys. synthesis (Govindjee, ed.), in press, AcaActa 298, 620-629. demic Press, New York.

here for the folloMng reasons: T h e cited reports have a different plant source for chloroplast e x t r a e t i o n - - V i c i a faba L. was used; spin-label p u r i t y was in question; the spin label used showed appreciable water solubility in our hands and is readily hydrolyzed to eaprylic acid and T E M P O L b y some biological preparations; inspection of their published spectra indicates excessive line broadening b y i n s t r u m e n t amplitude m o d u l a t i o n or local concentration aggregates; spectral measurements t a k e n from their spectra indicate unusually low membrane lipid viscosity, and chloroplast coneentrations are n o t controlled.

1. 2.

3.

4. 5. 6.

7.

8. 9.

99