Feb 17, 1982 - The light activation mechanism of the latent H+-ATPase was investi- gated in .... activity when present inthe light stage in untreated chloroplasts.
Plant Physiol. (1982) 70, 87-91 0032-0889/82/70/0087/05/$00.50/0
Activation and Deactivation of H+-ATPase in Intact Chloroplasts' Received for publication November 17, 1981 and in revised form February 17, 1982
YOSEPHA SHAHAK Biochemistry Department, Weizmann Institute of Science, Rehovot 76 100, Israel purified (18) as well as membrane-bound CF1 (20). The results described in this communication for intact chloroplasts support the idea of a PSI electron acceptor participating in the activation of the ATPase and further suggest that the deactivation of the enzyme involves oxidation reactions.
ABSTRACT The light activation mechanism of the latent H+-ATPase was investigated in intact spinach (Spinacia oleraea, Hybrid 424) chloroplasts. The following observations were made. (a) Photosystem I electron acceptors such as methyl viologen, nitrite, oxaloacetate, etc., inhibit the light activation of the enzyme. (b) The electron transfer inhibitor 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) fully inhibits the process. (c) Ascorbate plus diaminodurene or dithionite can restore light activation in DCMUpoisoned chloroplasts. (d) The activated state of the enzyme decays rather slowly (within a few minutes) after illumination of the intact chloroplasts. (e) The rate of dark decay is accelerated by oxidants (H202 or ferricyanide) and slowed down by dithiothreitol. It is suggested that the physiological mechanism for regulation of the H+-ATPase involves oxidation and reduction reactions in a manner which resembles the regulation of the light-activated carbon cycle enzymes.
MATERIALS AND METHODS
Spinach (Spinacia oleracea, Hybrid 424, Ferry Morse Seed Co., Mountain View, CA) was grown in a growth room under 10 h 26°C day and 14 h 18°C night cycles. A combination of fluorescent and incandescent light yielded around 4,500 lux of white light. Intact chloroplasts were isolated from leaves of 1- to 2-month-old plants, 1 to 2 h after the night period. Isolation procedure was as described previously (24). Intactness ranged between 60 and 75% as assayed by ferricyanide reduction before and after osmotic shock (16). CO2 fixation rates as measured by an 02 electrode in a medium containing 0.35 sorbitol, 50 mm Na Tricine (pH 8), 0.25 mM Pi, 10 mm NaHCO3, 1,000 units/ml catalase, and 25 ytg Chl/ ml, ranged between 50 and 70 timol/mg Chl h. The ATPase activity of intact chloroplasts was routinely activated by 3 min of illumination with 100,000 lux white light intensity in a thermostated bath at 23°C (unless otherwise stated). The activation medium was the same as the CO2 fixation medium except for the omission of NaHCO3 and the higher Chl concentration (0.35 mg/ml). For ATPase activity assay chloroplasts were osmotically shocked right after illumination (unless otherwise indicated) by a 5- or 10-fold dilution in a medium containing 10 mm Na Tricine (pH 8), 2 mm MgCl2, and 2 mm ATP (containing 1-4 x 105 cpm/ml y-labeled [32P]ATP). The reaction was terminated by addition of 5% TCA plus 0.4 mm Pi and centrifugztion. For other reaction conditions, see "Results." The release of 32Pi was assayed according to Avron (2) as follows: 0.5-ml aliquots of the supernatant were thoroughly mixed with 0.4 ml of 5% ammonium molybdate in 4 N HCI (w/v) and 2.0 ml of a 1:1 mixture of isobutanol and benzene. The tubes were centrifuged for 30 s in a table centrifuge, and l-ml aliquots of the organic phase were taken for counting in a Scintillation counter. y-Labeled [32P]ATP was synthesized by photophosphorylation and purified according to Magnusson et al. (17). 32Pi content ofthe [32P]ATP thus prepared was about 0.25%. -
The chloroplast ATP synthase catalyzes the formation of ATP in the light in an essentially irreversible manner. The enzyme has been found, though, to possess a latent ATPase activity. Exposure of the hydrolytic activity requires a special treatment of the chloroplasts, the most common one being preillumination in the presence of an electron carrier and a dithiol reagent. Activation of the ATPase was suggested to involve two principal steps: a lightinduced, AjiH, 2 mediated conformational change followed by an interaction of the modified site with the thiol reagent (for review, see Ref. 3). Most of the studies on ATPase activity of the ATP synthase were done in broken, washed (class C) chloroplasts, and their relevance to the in vivo mechanism is not obvious, since some properties of the enzyme might alter during breakage and washing, or interaction with stroma components might be lost. A few recent reports indicate that in intact (class A) chloroplasts, where the active part of the enzyme (CF1) is surrounded by the native stromal content, the ATPase is indeed activated by light. In this case, no addition of electron carriers or thiol reagents is required (12, 19, 20, 23). Buchanan et al. (4, 5, 18) and Mills et aL (20) suggested that the activation mechanism for the hydrolytic activity of the ATP synthase in vivo involves thioredoxin, a dithiol protein which is found in the chloroplast stroma and undergoes photoreduction by PSI via ferredoxin-thioredoxin reductase. This was based on studies of the effect of isolated thioredoxin on ATPase activity of the
RESULTS Effect of Electron Acceptors on Light Activation. The herbicide MV is a low potential, autooxidizable electron acceptor which effectively competes with ferredoxin for PSI reducing equivalents. Figure 1 demonstrates its effect on light activation of the ATPase. Intact chloroplasts were preilluminated for the indicated period and then osmotically shocked and assayed for ATPase activity in the dark. The presence of MV during light activation clearly inhibited ATPase activity (Fig. 1). When added after illumination, it had no effect (not shown). Under similar conditions, the addition of MV increased the formation of transthylakoid ApH in the light, as measured by 9-aminoacridine fluorescence quenching (not
' Supported by the Israel Academy of Sciences and Humanities-Basic Research Foundation. 2 Abbreviations: transmembranal electrochemical proton gradient; PGA, 3-phosphoglycerate; MV, methyl viologen; OAA, oxaloacetate; DAD, diaminodurene; DCCD, dicyclohexylcarbodiimide; ApH, transmembranal pH gradient; CF,, chloroplast coupling factor.
AiH',
87
Plant Physiol. Vol. 70, 1982
SHAHAK
88
shown; also, cf. Ref. 11). Other, naturally occurring, electron acceptors were also found to inhibit the activation stage of the ATPase. The effect of oxaloacetic acid, nitrite, bicarbonate, and PGA is summarized in Table 1. Although the effect of these compounds was smaller than MV, it was consistent, with nitrite being more potent than OAA, PGA, or HCO3 None of these inhibited when present in the dark stage only. All were added at a saturating concentration for electron transport. As is also evident from Figure I and Table I (and was also reported in Ref. 19), osmotically shocked chloroplasts hydrolyze ATP without a preillumination step at about one-third the rate obtained after activation. This dark activity was neither inhibited by energy transfer inhibitors (Dio-9 in Table I and DCCD in Ref. .
40
control
16-
E cr
30 _X
tM
°0 20
no
illumination
E
:I
I
4 5 3 illumination, min FIG. 1. Effect of MV on light activation of ATPase. Chloroplasts were illuminated in activation mixture as described in "Materials and Methods." At the indicated time, 100-,Il samples were osmotically shocked and assayed for dark ATPase activity for 10 min. MV concentration was 50 ,LM where added.
2
0
19) nor stimulated by uncouplers (Table I). It might originate in the nonlatent, Mg-specific ATPase located in the envelope membranes (21). The addition of an uncoupler (0.3-0.4 mm NH4C1) to the assay medium increased the relative activity due to the lightactivated enzyme and was therefore used in all the following experiments. The uncoupler concentration was chosen to yield a linear rate during the assay time. With higher NH4Cl concentrations, the initial rate of ATP hydrolysis was faster as in Carmeli (6), but linearity was not maintained (not shown). The relative inhibition by OAA, KNO2, or MV was the same when assayed in the presence or absence of NH4Cl (Table I). Activation by PSI. Activation of the ATPase can be mediated by PSI activity alone. As demonstrated in Table IIA, the PSII electron transfer inhibitor DCMU prevented light activation while having no effect when added after illumination. Reduced DAD, an electron donor to PSI (26), fully restored light activation in DCMU-poisoned intact chloroplasts. In the absence of ascorbate, DAD restored only 60% of the activity. The dark ATPase activity (of chloroplasts preincubated in dark instead of light) was essentially insensitive to ascorbate plus DAD (not shown). Dithionite was reported before to restore PSI activity in DCMUinhibited intact chloroplasts (8). It was also found here to restore partially the light activation of the ATPase (Table IIB). In this case, however, dithionite by itself inhibited about 50% of ATPase activity when present in the light stage in untreated chloroplasts. Dark Decay of the Activated State. Dark decay of the activated state was found to be accelerated by oxidizing agents. In Figure 2, intact chloroplasts were illuminated, then transferred to darkness and aliquots were osmotically shocked and assayed for ATPase activity after different dark intervals. Light-induced activity decayed to half the initial activity (t6/2) within about 100 s dark time. The addition of the oxidant H202 at the beginning of the dark period markedly accelerated the decay. About 0.5 mm H202 reduced tl/2 by 4-fold. The concentration dependence of the H202
effect is shown in Figure 3. In this experiment, the dark interval between light activation and ATPase activity assay was kept constant (2 min) while the H202 concentration varied. When present in the assay medium only, (Fig. 3, upper trace) the oxidant
Table I. Effect of Electron Acceptors on Light Activation of A TPase Chloroplasts were light-activated as described in "Materials and Methods" except for the indicated additions, then broken by five times dilution in a medium containing 20 mm Na Tricine (pH 8) and 2 mM MgCl2. After 15 s, 2 mM [32PJATP was added without (a) or with (b) 0.4 mM NH4Cl. Reaction was stopped after 10 min (a) or 3 min (b). Dark controls were preincubated for 3 min in the dark instead of light. Rates are given in jmol/mg Chl. h. ATPase Activity Addition to Activation +NH4CI (b) -NH4CI (a) rate L-D % No L-Da rate A None OAA, 5mM KNO2, 5 mM MV, 50 yiM Dark control Dark control + Dio-9h
39.3 35.0 30.5 24.8 11.2
28.1 23.8 19.3 13.6 0
100 85 69 48
None
36.1
26.1
100
B 20.4 30.4 NaHCO3, 10 mM 22.0 32.0 PGA, 3 mM 0 10.0 Dark control a L - D, light minus dark activity. b Dio-9 concentration in the assay medium was 6 jig/ml.
78 84
105.5 94.3
74.2 60.7 12.7 9.6
92.8 81.6 61.5 48.0 0
100 88 66 52
H+-ATPase REGULATION IN INTACT CHLOROPLASTS Table II. Activation of H+ A TPase by PSI Activation and assay conditions were as described for Table I (b). Concentrations of the compounds added to light activation (or assay) stage in experiment A were: DCMU, 20 (4) /tM; ascorbate (Asc), 10 (2) mM; DAD, 100 (20) ,M. In experiment B: DCMU, 15 (3.75) fLM; Na2S204, 10 (2) mm. DAD and dithionite were prepared just before use. Addition to ActivaATPase Activity Addition to Assay tion ,umol/mg Chl- h %
75
1004 0
DCMU DCMU + Asc DCMU + Asc + DAD
79.7 5.0 6.7 80.4 78.4 73.2 78.4 80.3
100 6 8 101 98 92 98 101
76.4
96
95.9 9.3 33.4 48.4 61.0
100 10 35 50 64
B None DCMU DCMU + S2042-
S204 2DCMU + S2042-
added after dark interval
80
0U
60
0
added before dark interval
40 0
A
None DCMU DCMU + Asc DCMU + Asc + DAD Asc Asc + DAD
w
89
20
0~
F
0L 0
1000
500 (100)
(0)
(200)
H202 . tM FIG. 3. Concentration dependence of the acceleration of dark decay by H202. Chloroplasts were light-activated as in Figure 2, darkened for 2 min, then broken by five times dilution and assayed for ATPase activity in the presence of 0.3 mm NH4Cl. H202 was added at the end of the illumination (-) or together with I32PIATP mixture (0). The abscissa indicates the H202 concentration during the dark decay (open numbers) or in assay medium (numbers in parentheses). The control rate was 33.5 ,umol ATP hydrolyzed/mg Chl * h (corrected for dark activity). The H202 concentration was determined in an 02 electrode by addition of catalse.
I
I
I
I
0.6
08
1.0
I
80
control (a) 0
:E
E
60
E
c0
Ca -o
E
N
401
)) 2
0O
ct
0
a) .5
E
+dark interval ( b)
-0
20)
0
0~
H-
no
preillumination(c)
UL
Dark interval, min FIG. 2. Effect of H202 and DTT on dark decay of the activated state of the ATPase. Intact chloroplasts were isolated without Na ascorbate. Light activation was as described in "Materials and Methods" except for the absence of catalase in the activation mixture. At the end of the illumination, 10 mm DTT or 0.5 mm H202 were injected (where indicated), and the test tube was transferred to darkness. After the indicated dark time, 100-1l samples were thoroughly mixed with 0.8 ml of 20 mm Na Tricine (pH 8) and 3 mM MgCl2. Assay was started 5 s later by addition of 2 mm [32PJATP and 0.3 mm NH4Cl, and stopped after 4 min by TCA + Pi as described in "Materials and Methods." Dark control (dashed line) was preincubated for 3 min in the dark instead of light. Numbers in brackets are the time for half decay in s.
had only a small effect. However, its addition at the end of the preillumination period caused deactivation, with 0.25 mm H202
giving 50% inhibition (Fig. 3, lower trace). The chloroplast envelope is highly impermeable to ferricyanide (16). However, in chloroplasts which were osmotically shocked at
0
0.2
0.4
K3Fe (CN)6,mM FIG. 4. Effect of ferricyanide on the ATPase dark decay in osmotically shocked chloroplasts. Intact chloroplasts (isolated without ascorbate) were illuminated for 3 min as described in "Materials and Methods," then osmotically shocked in the dark by 10-fold dilution. In a, dilution medium contained 10 mm Na Tricine (pH 8), 3 mM MgCl2, mm Pi (pH 8), ferricyanide at the indicated concentration, 2 mm ATP (containing 7 x 105 cpm [32P]ATP, and 0.3 mm NH4C1. The assay was stopped after 3 min by 5% TCA. In b, chloroplasts were treated as in a, except for the omission of ATP and NH4Cl from the dilution medium. A mixture of [32P]ATP and NH4Cl was added 1.5 min after illumination (final concentration, same as in a) and TCA after 3 more min. c was the same as b, except for preincubation in the dark instead of light.
the end of the light activation, ferricyanide markedly increased the decay. Concentration dependence of this effect is given in Figure 4 which shows that light-induced ATPase activity decayed to half the original activity during a 90-s dark period in the
90
SHAHAK
Plant Physiol. Vol. 70, 1982
absence of ferricyanide. Its presence at 0.5 to 1.0 mm concentration A control mechanism which involves an activation process is during this period was sufficient to fully inhibit the activity. In expected to also involve a deactivation reaction which would intact chloroplasts, the dark decay was essentially insensitive to convert the ATPase back to its latent state. The dark decay of ferricyanide, unless the latter was added together with the perme- ATPase in broken chloroplasts is increased by ADP and slowed able mediator DAD. In the latter case, the deactivation was much down by Pi (7). In intact chloroplasts, where both ATP, ADP, and accelerated (data not shown). Pi are part of the stromal content (for review, see Ref. 15), the The thiol reductant DTT was found to slow down the dark
decay of the activated state. DTT (10 mM) increased the tl/2 of the decay by 2.4-fold (Fig. 2). When assayed immediately following light activation, ATPase activity was the same in the presence or absence of DTT (time-0 in Fig. 2), suggesting that the endogenous thiol compound is sufficient for maximal activation under the experimental conditions used. DISCUSSION In the work described here, the effect of electron carriers, oxidants and reductants on the light activation and dark deactivation of intact chloroplast H+-ATPase was investigated. Mg2+dependent ATPase activity was assayed in osmotically shocked chloroplasts which had been preilluminated while intact. The addition of PSI electron acceptors was found to inhibit the light activation process. Physiological as well as nonphysiological, envelope-permeable electron acceptors had an inhibitory effect only when present during preillumination (Fig. I; Table I). The reduction of these acceptors by illuminated intact chloroplasts is known to be coupled to the formation of a transthylakoid pH gradient (I 1, 14). The activation of ATPase in broken chloroplasts requires both the formation of ApH and modification of the membrane bound CF1 by a thiol reagent (3). By analogy, the inhibition of the activation in intact chloroplasts might be due to an effect on the reduction of a native thiol reagent since ApH is not decreased but rather increased by the electron acceptors. Moreover, the degree of inhibition correlates well with the site of electron acceptance relative to the site of ferredoxin. The best inhibitor was MV, followed by N02-, OAA, PGA, and HCO3 (Table I). MV accepts electrons before ferredoxin and would thus most efficiently inhibit electron flow to other acceptors. Nitrite reductase takes electrons from reduced ferredoxin. The reduction of the other acceptors by ferredoxin involves more enzymic steps and might thus be expected to be less inhibitory. The following scheme shows electron transfer routes to the electron acceptors, (Th, thioredoxin). OAA or PGA or
HCO3
NADP
thioredoxin
Fd-Th reductose
Fd-NADP reductase
nitrite reductase
Fd MV
PS I
N02
half-time for dark decay is rather slow, about 100 s (Fig. 2). Halftimes for dark decay of 3 or 7 min have been reported for ATPase activity of intact chloroplasts as assayed by ATP-induced ApH formation (19) or reverse electron transport (23), respectively. Therefore, inactivation of the ATPase by tight binding of ADP to CF1 which occurs within a few s after illumination (25) may not play an important role in this respect in vivo. Alternatively, it is suggested here that the latent and activated states of the enzyme are controlled by oxidation-reduction reactions as indicated by the stimulation of dark decay by H202 and ferricyanide and inhibition by DTT (Figs. 2-4). A similar control mechanism has been described for light-activated stromal enzymes involved in C02 fixation, like fructose 1,6-bisphosphatase, phosphoribulokinase, NADP-malate dehydrogenase, etc. They were all suggested by Buchanan et al. (4, 5) to be activated by reduced thioredoxin and inactivated by either soluble or membrane-bound oxidants or by a yet unknown mechanism (5, 27). H202 is a physiological oxidant which is generated in the light and reversibly inhibits C02 fixation by intact chloroplasts (9). More specifically, it inhibits enzymes which require reduced sulfhydryl groups for activity, including fructose or sedoheptulose bisphosphatase and phosphoribulokinase (10, 13). Thus, even though the carbon-cycle enzymes mentioned are considered to be soluble while CF, is a membrane bound enzyme, they all seem to share common mechanisms of activation and deactivation. A somewhat different mechanism for the light activation of the carbon-cycle has been suggested to include membrane-bound light effect modulators (LEMs) which might accept electrons at different sites following PSI (1). The data presented here do not exclude the possibility that such modulation might participate also in the activation of the H+-ATPase in vivo, although the ease of removal of the ATPase activator (19, 20) favors the involvement of soluble factors. Another aspect related to this work is worth mentioning. The chemiosmotic hypothesis postulates the existence of two separate proton pumps in the thylakoid membrane (as well as in other energy conserving systems): the electron transfer chain and the ATP synthase complex. These are linked to each other via the common proton pool. The data presented here suggest that the electron transport system also interacts with the ATP synthase through oxidgtion-reduction reactions at the level of the acceptor side of PSI., Indeed, electron microscopy indicates that PSI reaction centers and the ATP synthase complex are located close to each other in the stroma lamellae as well as in the grana margins. Moreover, immunological studies suggest that the NADP reductase is located in depressions formed by the knobs of CF1 (for a comprehensive review, see Ref. 22). Ferredoxin, thioredoxin reductase, and thioredoxin might form loosely bound complexes in those depressions, to be readily available for the activation of the ATPase when required.
Acknowledgments-Drs. M. Avron and U. Pick are acknowledged for helpful Further support for the essential role of PSI in the light acti- discussions, and Dr. G. Hind for reviewing the manuscript. vation process comes from the restoration of the activation by LITERATURE CITED reduced DAD or sodium dithionite in DCMU-poisoned intact chloroplasts (Table II). The restoration by dithionite might be the 1. ANDERSON LE, M AVRON 1976 Light modulation of enzyme activity in chlororesult of a direct reduction of the physiological thiol activator plasts. Generation of membrane-bound vicinal-dithiol groups by photosynthetic electron transport. Plant Physiol 57: 209-213 and/or restoration of cyclic electron flow which would also proby Swiss chard chloroplasts. Biochim vide AiH' (8). The results are compatible with those described by 2. AVRON M 1960 40:Photophosphorylation Biophys Acta 257-272 Mills et al. (20) for DCMU-treated broken chloroplasts supplied 3. BAKKER-GRUNWALD T 1977 ATPase. In A Trebst, M Avron, eds, Encyclopedia with ferredoxin, thioredoxin reductase, and reduced dichlorophenof Plant Physiology-Photosynthesis I, Vol 5. Springer-Verlag, Berlin, pp 369-373 olindophenol under anaerobic conditions.
H+-ATPase REGULATION IN INTACT CHLOROPLASTS 4. BUCHANAN BB 1980 Role of light in the regulation of chloroplast enzymes. Annu Rev Plant Physiol 31: 341-374 5. BUCHANAN BB, RA WOLOSIUK, P SCHURMANN 1979 Thioredoxin and enzyme regulation. Trends Biochem Sci 4: 93-96 6. CARMELI C 1969 Properties of ATPase in chloroplasts. Biochim Biophys Acta 189: 256-266 7. CARMELI C, Y LIFSHITZ 1972 Effect of Pi and ADP on ATPase activity in chloroplasts. Biochim Biophys Acta 267: 86-95 8. CROWTHER D, JD MILLS, G HIND 1979 Protonmotive cyclic electron flow around Photosystem I in intact chloroplasts. FEBS Lett 98: 386-390 9. EGNEUS H, U HEBER, U MATTHIESEN, M KiRK 1975 Reduction of oxygen by the electron transport chain of chloroplasts during assimilation of carbon dioxide. Biochim Biophys Acta 408: 252-268 10. HELDT HW, CJ CHON, R McC LILLEY, AR PORTIS 1977 The role of fructose and sedoheptulosebisphosphatase in the control of CO2 fixation. Evidence from the effect of Mg" concentration, pH and H202. In DO Hall, J Coombs, TW Goodwin, eds, Proceedings of the 4th International Congress on Photosynthesis. Biochemical Society, London, pp 469-478 11. HIND G, D CROWTHER, Y SHAHAK, ER SLOVACEK 1981 The function and mechanism of cyclic electron transport. In G Akoyunoglou, ed, Photosynthesis-Proceedings of the 5th International Congress on Photosyntheis, Vol 2. Balaban International Science, Philadelphia, pp 87-97 12. INOUE Y, Y KOBAYASHI, K SHIBATA, U HEBER 1978 Synthesis and hydrolysis of ATP by intact chloroplasts under flash illumination and in darkness. Biochim Biophys Acta 504: 142-152 13. KAISER WM 1979 Reversible inhibition of the Calvin cycle and activation of oxidative pentose phosphate cycle in isolated intact chloroplasts by hydrogen peroxide. Planta 145: 377-382 14. KOBAYASHI Y, Y INOUE, K SHIBATA, U HEBER 1979 Control of electron flow in intact chloroplasts by the intrathylakoid pH, not by the phosphorylation potential. Planta 146: 481-486 15. KRAUSE GH, U HEBER 1976 Energetics of intact chloroplasts. In J Barber, ed,
16. 17. 18. 19. 20. 21.
22. 23. 24. 25. 26.
27.
91
The Intact Chloroplasts. Elsevier/North Holland Biomedical Press, Amsterdam, pp 171-214 LILLEY R McC, MP FITZGERALD, KG RIENITs, DA WALKER 1975 Criteria of intactness and the photosynthetic activity of spinach chloroplast preparations. New Phytol 75: 1-10 MAGNUSSON RP, AR PORTIS, RE MCCARTY 1976 Quantitative, analytical separation of adenine nucleotides by column chromatography of polyethyleneimine-coated cellulose. Anal Biochem 72: 653-657 McKINNEY DA, BB BUCHANAN, RA WOLOSIUK 1979 Association of a thioredoxin-like protein with chloroplast coupling factor (CF1). Biochem Biophys Res Commun 86: 1178-1184 MILLS JD, G HIND 1979 Light induced Mg2" ATPase activity of coupling factor in intact chloroplasts. Biochim Biophys Acta 547: 455-462 MILLS JD, P MITCHELL, P SCHURMANN 1980 Modulation of coupling factor ATPase activity in intact chloroplasts. The role of thioredoxin. FEBS Lett 112: 173-177 POINCELOT RP, PR DAY 1974 An improved method for the isolation of spinach chloroplast envelope membrane. Plant Physiol 54: 780-783 SANE PV 1977 Topology of the thylakoid membrane of the chloroplast. In A Trebst, M Avron, eds, Encyclopedia of Plant Physiology-Photosynthesis I, Vol 5. Springer-Verlag, Berlin, pp 522-542 SCHREIBER U 1980 Light-activated ATPase and ATP-driven reverse electron transport in intact chloroplasts. FEBS Lett 122: 121-124 SHAHAK Y, D CROWTHER, G HIND 1980 Endogenous cyclic electron transport in broken chloroplasts. FEBS Lett 114: 73-78 SHOSHAN V, BR SELMAN 1979 The relationship between light-induced adenine nucleotide exchange and ATPase activity in chloroplast thylakoid membranes. J Biol Chem 254: 8801-8807 TREBST A, E PISTORIUS 1965 Zum Mechanismus des photosynthetischen Elektronentransportes in isolierten Chloroplasten. Substituierte p-Phenylenediamine als Elektronendonatoren (II). Z Naturforsch 20b: 143-144 WOLOSIUK RA, BB BUCHANAN 1977 Thioredoxin and glutathione regulate photosynthesis in chloroplasts. Nature (Lond) 266: 565-567
