energy conversion in the functional membrane of … … · energy conversion in the functional...

355

Biochimiea et Bioph),sica Acta, 505 (1979) 355 -427 ©Elsevier/North-Holland Biomedical Press

BBA 86054

ENERGY CONVERSION IN THE FUNCTIONAL MEMBRANE OF PHOTOSYNTHESIS .

ANALYSIS BY LIGHT PULSE AND E L E C T R I C PULSE METHODS

THE CENTRAL ROLE OF THE ELECTRIC FIELD

H.T. WITT

Max- Volmer-lnstitut far Physikalische Chemie und Molekularbiologie, Teehnische Universitdt Berlin, Strasse des 1 7. Juni 135, 1 Berlin 12 {Germany}

(Received September 6th, 1978)

Contents

I. Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357

II.

III.

The overall concept . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 358

The methodological advance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 A. Excitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 B. Characterization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 C. Registration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 360 D. Information . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 360

IV.

V.

Energy migration in the antennae pigment systems . . . . . . . . . . . . . . . . . . . . . . . 362 A. Collecting energy migration 362 B. Dissipating energy migration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362

Photoreactions of the chlorophylls . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 A. Chlorophyll a I (P-700) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364

1. A dimer as reaction center of antennae system I . . . . . . . . . . . . . . . . . . . . . 364 2. Photooxidation and vectorial electron ejection . . . . . . . . . . . . . . . . . . . . . . 364 3. The phases of rereduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

Abbreviations: AA, Field-indicating absorption change and electrochromic shift, respectively; BChl, bacteriochlorophyll; Car, carotenoid; CCCP, carbonylcyanide m-chlorophenylhydrazone; CF~, coupling factor of the ATPase; Chl, chlorophyll; Chl-a, chlorophyll a; Chl-b, chlorophyll b; Chl-a I (P- 700), chlorophyll a I (reaction center of antennae system I); Clil-ali (P-680), chlorophyll alI (reaction center of antennae system II); Cyt, cytochrome; DCMU, 3-(3,4-dichlorophenyl)-l,l-dimethylurea; DCIP, 2,6-dichlorophenol indophenol; A4), electrical potential difference; a ~ , surface potential difference; F, electric field strength; G, free energy; i, ion current; ill+, H ÷ current; iK+, K + current;f, current density; PC, plastocyanine; PMS, N-methylphenazonium sulfate; PQ(1), plastoquinone, primary electron-acceptor of Chl-all (==-X-320); PQ(2), plastoquinone, connector between PQ(1) and PQ pool; PQ, plastoquinone, member of the PQ pool; r, half-life and half-rise time, respectively; X-320, plastoquinone, primary electron-acceptor of Chl-all (=-PQ(1); 1 (...), single turnover conditions; ss(...), steady state in saturating permanent light; S, enzyme system for the cleavage of H20.

356

B. C h l o r o p h y l l a t l (P-680) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

1. React ion center of antennae system II . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

2. Photooxida t ion and vectorial electron ejection . . . . . . . . . . . . . . . . . . . . . . 365

3. The phases of rereduct ion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 366

C. Coupling of chlorophyl l a I with chlorophyl l a II . . . . . . . . . . . . . . . . . . . . . . . 367 1. Stoichiometr ics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367

VI. Electron transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367

A. Primary acceptor of ch lo rophy l l a l : ferredoxin . . . . . . . . . . . . . . . . . . . . . . . . 368

B. Donor site of chlorophyl l ai : p lastocyanine . . . . . . . . . . . . . . . . . . . . . . . . . . 368

C. Primary electron acceptor of chlorophyl l a l l : p las toquinone X-320 . . . . . . . . . . . . 368

D. Donor site of chlorophyl l a l i : H 20 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 369

E. Link between chlorophyl l a I and chlorophyl l a l l : p las toquinone pool . . . . . . . . . . 370 F. The electron transport chain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371

VII. Electric field generat ion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373

A. Detect ion by electrochromism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373

B. Kinetics of the electrochromic changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374

C. Mapping of the versati l i ty of informat ion obtainable by the electrochromic method . . 374 1. Discriminat ion between e lect rochromism and other phenomena . . . . . . . . . . . . 375

D. Properties of e lectrochrolnism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376

E. Proof of the elctrochromic method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377

1. a. Spectroscopic evidence - analysis of the shape . . . . . . . . . . . . . . . . . . . . . 377

b. Spectroscopic evidence comparison with model systems . . . . . . . . . . . . . . 377 2. Kinetic evidence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378

3. Electrical evidence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378

F. Orientat ion, polar izat ion and local izat ion of the field . . . . . . . . . . . . . . . . . . . . 379 G. Linear indicat ion of the field . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 380

H. Calibrat ion of the field . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 381

I. Funct ional unit of the electric field . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 384

K. Generat ion of the field by electron ejection from the excited chlorophylls a I and a l l . . 384 L. Decay of the field by ion fluxes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 384

M.Elec t rochromic measurements in the steady state . . . . . . . . . . . . . . . . . . . . . . 385

1. Total electrical potent ia l difference . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 385 2. Surface potent ia l difference . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 386 3. Electrochemical pro ton potent ia l . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 387

N. Electrochromic measurements with artificial probes; applicat ion to further systems;

other techniques for electric measurements . . . . . . . . . . . . . . . . . . . . . . . . . . 388

VIII. Molecular organization based on analysis of the electric field . . . . . . . . . . . . . . . . . 389

A. Asymmetr ic membrane architecture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 389

B. Local izat ion of the primary acceptors and donors . . . . . . . . . . . . . . . . . . . . . . 390 C. Local izat ion of the chlorophyl l react ion centers . . . . . . . . . . . . . . . . . . . . . . . 391

D. Organizat ion of plas toquinone as a pool and strand . . . . . . . . . . . . . . . . . . . . . 391 E. Localizat ion of the cleavage of water . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391

F. Or ienta t ion of antennae chlorophylls and carotenoids . . . . . . . . . . . . . . . . . . . . 392 G. Complexa t ion of antennae chlorophylls with carotenoids . . . . . . . . . . . . . . . . . . 393 H. Phase t ransi t ion and conformat ional changes . . . . . . . . . . . . . . . . . . . . . . . . . 395

IX. Proton t ranslocat ion - subsequence of the field generat ion . . . . . . . . . . . . . . . . . . 396 A. External pro ton uptake and internal release . . . . . . . . . . . . . . . . . . . . . . . . . 396 B. Stoichiometr ics of H÷/e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397

1. Two sites of external pro ton uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397 2. Two sites of internal pro ton release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397

3. The mechanist ic meaning of the H+/e relat ionship . . . . . . . . . . . . . . . . . . . . 398 C. Kinetics of external pro ton uptake and internal release . . . . . . . . . . . . . . . . . . . 399 D. Evolut ion of the proton gradient ApH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400

357

E. The proton potential as control of the electron transfer . . . . . . . . . . . . . . . . . . . 401 F. The plastoquinone pool as possible hydrogen pump . . . . . . . . . . . . . . . . . . . . . 402

1. Variation of the activity of the plastoquinone hydrogen pump . . . . . . . . . . . . . 404

X. Vectorial pathway of e, H ÷ and H in the zigzag scheme . . . . . . . . . . . . . . . . . . . . . 404

Phosphorylation in the light-induced field and proton gradient . . . . . . . . . . . . . . . . 405 A. Acceleration of the field decay during phosphorylation . . . . . . . . . . . . . . . . . . . 406 B. Retardation of the field decay with removal of F1-ATPase . . . . . . . . . . . . . . . . . 407 C. Uncoupling of phosphorylation by a competitive pathway for the field decay . . . . . . 407 D. Functional unit of phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 408 E. Uncoupling of phosphorylation by a competitive pathway for the ApH decay . . . . . . 409 F. Competition between basal and phosphorylating pathways of H ÷ . . . . . . . . . . . . . 410

1. Dependence on A¢ and apH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 410 2. Stoichiometrics of H+/ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 410 3. Apparent thresholds and interchangeability of Aq~ and apH . . . . . . . . . . . . . . . 411

G. Proton energy and energy requirement for ATP formation . . . . . . . . . . . . . . . . . 411

XI.

XII. Phosphorylation induced by an external electric field . . . . . . . . . . . . . . . . . . . . . . 413 A. Principle of the external field method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 413 B. Yield of ATP formation in an external field . . . . . . . . . . . . . . . . . . . . . . . . . 414 C. Conformational change and turnover time of the ATPase . . . . . . . . . . . . . . . . . . 414 D. Gating of the ATPase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 415

XIII. Data for discrimination between the different hypotheses on the mechanism of phosphorylation in photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Coupling between the electron transport chain and tile ATPase . . . . . . . . . . . . . . B. Meehanistics within the ATPase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

XIC. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

418 418 420

420

421

421

I. Preface

The pulse m e t h o d s w i th wh ich this review is c o n c e r n e d have been deve loped for

p h o t o s y n t h e s i s since 1955. (1) M m o s t all p r i m a r y processes are observab le on ly b y these

m e t h o d s . (2) Several o t h e r basic events have been observed first b y the use o f these tech-

n iques . (3) The m e t h o d s c o n t r i b u t e , f u r t h e r m o r e , to the s t u d y o f n u m e r o u s par t ia l reac-

t ions in the overall process . In t o t o , the sub jec t s ana lysed in this way cover the ' essent ia ls ' necessary to evaluate the f r a m e w o r k o f the mo lecu l a r c o n c e p t o f p h o t o s y n t h e s i s .

The top ics to wh ich these t e c h n i q u e s were appl ied b y us are those w h i c h are set o u t in

the Con ten t s . These are c o n n e c t e d m a i n l y w i th the phys i cochemica l f o u n d a t i o n s o f p h o t o -

syn thes i s o f green p lan ts . In pa r t these have been reviewed in Refs. 1 , 2 and 3. The ex ten-

sions wi th in the last years are m a i n l y the sub jec t o f this review. Fo r the pu rpose o f

p re sen t ing a se l f -con ta ined repor t , p rev ious results are m e n t i o n e d br ief ly .

The sequence o f the topics covered is ident ica l to the sequence o f the r eac t ion

pa t t e rns . The review star ts w i th energy mig ra t ion in the p i cosecond range and ends w i th

the f o r m a t i o n o f N A D P H and A T P in the mi l l i second d o m a i n .

358

Some of these studies are based on biological and biochemical results; others have been corroborated or extended by biochemical research work.

No emphasis has been laid on any particular item within this review. However, separation of charges and the electric field generation, respectively, emerge as the primary act of photosynthesis. All further reactions are subsequent to this process. The field is engaged also at the end of the overall processes in the formation of ATP. The field represents, furthermore, the vectorial character of tile molecular machinery and gave first evidence of the asymmetrical distribution of the molecules within the membrane. Furthemlore, versatility of information obtainable from thorough analysis of the properties of tile field is immense. Thus the electric field runs through photosynthesis like a red thread. This is the reason for the choice of the subtitle.

Considerable attention has been paid to detailed subdivision of the various topics. I have attempted to arrange these and their subdivisions in a logical sequence, hoping that the list of contents might thereby serve also as an abstract.

Results reported by other workers which are related to the work reviewed here have been described and discussed recently in several excellent multiauthor books and reviews. For a comprehensive discussion on these results I refer the reader to this literature as mentioned in the following pages.

!I. The overall concept

Photosynthesis in green plants takes place in subcelhilar organelles, the chloroplasts. The inner system of the chloroplasts contains about 1000 small compartments, the so-called thylakoids. Thylakoids are disk-shaped vesicles with a diameter of about 5000 A. (see Fig. 1). Several thylakoids can be arranged in a super-unit through interconnection.

For literature on subjects covered in this section, see recently published books on photosynthesis [4-6] .

In one thykaloid membrane about l0 s pigment molecules are embedded. The pigments are Chl-a and -b and carotenoid (Car). Light energy absorbed in these so-called antennae pigments is channeled by a 'collecting' type of energy migration to the photoactive centers. Superfluous energy is channeled out of the systems via a 'dissipating' type of energy migration (see Fig. 1, top).

The energy of excited chlorophyll centers is used for an electron transfer from H20 via various carriers to the Cmal acceptor NADP ÷. This transfer starts with an electron shift from the donors at the inside to the acceptors at the outside. Thereby a field is set up across the membrane (see Fig. 1, center). About 200 such electrons transport chains are embedded in one thykaloid membrane, i.e., each chain is surrounded by about 500 antennae pigment molecules (see Fig. 3).

Protolytic reaction with the separated charges induce a proton translocation and circulation, respectively (see Fig. 1, bottom). The potential of the protons is used for phosphorylation, i.e., for the synthesis of ATP from ADP + P.

With the reducing power of NADPH and the energy stored in ATP, absorbed CO2 can be reduced in the Calvin cycle into sugar and all the other energy-rich natural compounds.

It is the aim of this review to present results on which this concept is based and to outline experiments which give evidence on details of the molecular machinery by which the light energy is converted into the chemical energy of NADPH and ATP.

hv

collecting and . . . . . . ___~'" 105antennae dissipating / / ~.- . . . . pigments per energy migration ~ ~ ~ ~ ~ ~ " ~ - ' = - ~ ~ thyldkoid

359

coupling of NADP',~ ~ .13"'"'" ~ ' ~ 200 electron 2 chlorophyll ..t~l" ~ transport chains centers ~ ~ per thylakoid vectorial t ~1~ - ~ electron Chl QI Chl Oil H20 transfer

Proton circulation

Phospho- rylation

t /2 NADPH H ° / AOP ATP

%1/2 H H* 1//, 2 H* 2

/ 1 thylakoid

out

17oA

Fig. 1. Simplified scheme of the reaction sequences in the funct ional membrane o f photosynthes i s (see Section 1I).

III. The methodological advance

In contrast to photophysics and photochemistry in vitro, the field of photosynthesis is less accessible for investigation into reaction mechanism because only a very small proportion of pigments (1°/0o) is engaged in photoreactions. Furthermore, the signals induced by the pulses are extremely small because the analysis has been made predominantly under single turnover conditions where insight into the relations between the complex reaction patterns is optimal.

Therefore, based on the light flash method introduced by Norrish and Porter [6a], since 1954, kinetic methods have, been developed in our laboratory for a high time resolution together with a high signal-to-noise ratio (S/N) [7-9].

IliA. Excitation For one turnover without recycling, the pulse must be shorter than the duration of the

event under consideration. This requirement was realized by using normal flashes (10/~s), ultrashort flashes (1/as) and laser giant pulses (10 ns). Recently, the photosynthetic membrane has been energized with external electric field pulses instead of light pulses. Equipment has been developed in which these different types of pulse can be run periodically (0.1-1000 Hz). This is a necessity for application of the sampling technique discussed below.

IIIB. Characterization With respect to the indication of the events, the following physical changes connected

with the various reactions have been used.

360

(I) Intrinsic absorption changes. These are due to the change of optical properties of the compounds engaged in the reaction.

(2) Intrinsic absorption shifts. These are based on the Stark effect, i.e., on the shift of tile absorption bands of any membrane pigment which is exposed to an intrinsic electric field generated within the membrane. These optical changes are also called 'electrochromic changes'.

(3) Extrinsic absorption and fluorescence changes. These are caused by artificial probes added to the reaction mixture, for instance pit indicators, which respond to change in proton concentrations.

(4) Electrostatic changes. A change of the intrinsic electric field can be transmitted under special conditions to macroscopic electrodes in the form of an electrostatic induction.

(5) Chemical changes. The pulse studies have been supplemented by the measurement of chemical changes, i.e. of products such as Oz, ATP and adenosine nucleotides in general, which indicate the actions of electron transport, phosphorylation and conformational changes.

IIIC. Registration For the observation of the small single turnover signals within short times, a sampling

technique was introduced [8,9]. The development of appropriate devices for the storage of signals made it possible to increase the signal-to-noise ratio, S/N, with the number, n, of signals averaged such that SIN ~ ~ The extremely high sensitivity of this sampling technique enables the detection of very small signals. In combination with periodic pulse excitation (see above) it was possible to bring the time resolution for biological events from seconds down to 10 ps [8]. Only in isolated reaction centers of bacteria from which the bulk of pigments have been separated and which are 'optically clear', are available signals of such a size that the time range can be extended into the picosecond domain [ 10]

II1D. Information The signals induced by excitation of the photosynthetic apparatus with the different

types of pulse outlined above have been investigated on various physical and chemical modifications of the system. In this way, under single turnover conditions, the following information has been obtained (see Fig. 2).

(1) Intrinsic absorption changes (1955) gave information on: (a) Energy migration; (b) Light reactions; and (c) Electron transfers.

(2) Intrinsic absorption shifts and electrochromic changes, respectively, (1967) provided results on:

(a) Electric events in general; (b) Molecular organization; (c) Vectorial pathways; and (d) Phosphorylation.

(3) Extrinsic absorption and fluorescence changes (1969) provided information on: (a) Proton translocations.

(4) Electrostatic changes (1972) supported some conclusions on electric events which were derived from the electrochromic changes.

(5) Energization with external electric pulses (1975) revealed information on: (a) Properties of the ATPase complex: (b) Discrimination between different hypotheses on the mechanism of phosphoryla-

lation in photosynthesis.

361

B

_ _ _

• / , - ~ * • ~ Car

i ~'.~,,ip."~",~ /*90 .60oc q60oC i 430 460 o o2atm O 2

400 500 600 700nm

1A ('i....-V.....~ q Light reaction, s

decoy : IOtas, i u , l / c , 22!10 "3 '[~'/0 0 )(PT00) 20ms

cT) q- 0 {.1

"5

o

F

°t ~r~~"---~. .~, . - c

v I/ I i,

_16.10-3 i~/682 L

400 500 600 70Onto

D 32.,~%..

1 !10 l. - " " . /°-° -x. .--\ ! . ," 5, O~ /

, ~ , s

280 3~Onm - - J

C~l-~ a l " r tse~ 20ns

Chl-o I decay :f31..ts), 351.ts,

(P680) 2001.t s

Electron transfers

(X-320}

PQ- r,se -" 20ns ~ decay =0.6 rrs

PQ

E i "\

/ "\ ? " \ /

.~1!10-3 • . "...L263

240 280 32Ohm

pQ= r ise -" 0 6 m s l

I t decoy : 2 0 m s PQ

I Electric events

F 1llo .3 j \ " ........ .~'.'., ~,f~v" cu-a __., c h t-a"

r i s e ~< 20 ns

, J , decay ,~ 2 0 m s

~00 500 600 700nrr

wavelength

Fig. 2. Intrinsic transient difference spectra which indicate different events of the mechanism of photosynthesis (see subsection IIID). References: A, ] 6; B, 31 ;C, 51 ; D, 69, 71 a; E, 69; F, 90.

362

IV. Energy migration in the antennae pigment systems

IVA. Collecting energy migratkm In photosynthesis the molecular machinery must be adaptable to dim as well as to

bright light. At low intensity the very few quanta excite tile correspondingly small number of reaction centers with high probability because energy transfer between the bulk pigments channels the energy, probably via the singlet states, from Car to Chl and from Chl to Chl within approx. 10 -13 s. In this way, also, the photoactive centers are 'visited' after approx. 10 -11 s, whereby tile energy is trapped. This phenomenon was discovered as early as 1932-1936 [11,12].

One can distinguish between two different pigment systems I and II (see Fig. 3). Each of them channels energy to one of the two chlorophyll reaction centers which operate in an electron transport chain (The arguments are outlined in SectionV).

The mechanism of the moving of excitons within the two different antennae systems to the traps is discussed in the review, Ref. 13. The mechanism of the distribution of the excitons from one antennae system to the other via a light harvesting pigment complex ('spill-over') is reviewed in Ref. 14.

IVB. Dissipating energy migration From our results obtained in 1961 [15,15a] and from those extended in 1969 evidence

was produced for the existence of another type of energy migration in photosynthesis [ 16-18]. At high light intensities, superfluous energy stays in the excited Chl states. The energy must be dissipated quickly because it is known that excited Chl is irreversibly destroyed by photooxidation with the omnipresent 02. Energy dissipation may occur by re-emission as fluorescence or by non-radiative transitions.

An additional 'valve reaction' is probably energy transfer and migration after inter- system crossing via triplet states in the opposite direction from Chl to Car (see Fig. 3). Because excited Car is not irreversibly destroyed by photooxidation with Oz, in contrast to excited Chl, this process can be used to drain offharmful photooxidation energy from Chl. This T-T energy transfer and migration is obviously 'invented' by nature for high light intensity conditions to protect antennae chlorophyll from photooxidation. Tile transfer can be regarded as the counterpart to the S-S energy transfer and migration 'invented' for lower intensities.

The existence of this 'triplet valve' is based on eight results on the ab;orption changes occurring with the formation of the triplet states of carotenoids (Car T) [16,17]. (1) These absorption changes occur in less than 20 ns and decay within 3 ~s (see Fig. 2A). (2) Three bleachings in tile absorption bands of Car at 430, 460 and 490 nm and no bleaching in the red, where Chl absorbs, indicate a Car reaction. (3) The rise of absorption at the position 520 nm, at which the absorption of Car T is observed in vitro indicates the generation of Car T in vivo. (4). This interpretation is supported by reversibility at --160°C and by tile fact that tile changes at 520 nm can be completely quenched by paramagnetic gases such as 02 and restored in diamagnetic gases such as N2 [15a]. (5) The quenching by paramagnetic gases is due to an accelerated decay of Car T [19]. (6)Because tile spectral changes can be induced with red light which excites only Chl, energy transfer and migration from Chl to Car must take place. (7) That this pathway from Chl to Car is used only for superfluous light energy follows from the observation that with increasing light intensity the formation of Car T starts only when photosynthesis begins to reach satura-

363

hv I ~500. hv D

/~ ~ ~ ?-',;~" ~ ' J ~ N collecting / ~ I ~ ; . ' , " ~ L _ l l I \ s-sEoer~y / V[_.~-,.:~,..:.: / k__.Jj ~ r ~ migrotion

\ ^ r-n f + J ! / dissipating J ~ L@ ~---~ / T-'I Energy

I I P,gmentsystem [ Pigmentsystem ~.

/ / . / NADP'* ~ - - 7 - - - C h l - a I . . . . . pQ ------ Chl-a.g / H2 0

'hq '- '~/ '~1 Cars It S • s I ~ ' - ~ S CN-Q CN-a I-- collecting } "~* {Chi-b) -. I ~ antennae I S-S Energy

' ~ " -- s/, I---pJgments ' migration ~'----~Chl-a'/'~/ , "" }

. / / ' ~ , , , ~ 1" [ ~ / / - - - - - "L, Energy

~ a r . 3 g . s Carl. (Chl-b T) 4----chl-2 J / ~ migration

Fig. 3. Collecting and dissipating energy migrations in the antennae pigments of photosynthesis (see subsections IVA and B) [651.

tion [16]. (8) In chloroplasts, about 20% of the superfluous energy is dissipated via this 'triplet valve' [20].

From these and other experiments we concluded that some specific organization of Car and Chl in the thylakoid membrane enables in bright light dissipation of quanta by T-T transfer and migration. The mechanism explains also the long known observation that chlorophyll is irreversibly oxidized in systems deficient in carotenoids [38]. Recently, this type of energy migration was established to exist also in photosynthetic bacteria [48,49] but with a much lower yield (1%) than in chloroplasts (20%).

V. Photoreactions of the chlorophylls

Far red light (700-730 nm) produces a poor electron transfer. The yield is, however, enhanced to normal efficiency when shorter wavelength light (less than 700 rim) is added. In 1958, Emerson [21] proposed from this 'two-color' effect that two antennae pigment systems are engaged in the electron transfer: one with a long wavelength photoactive center (less than 730 nm) and one with a shorter wavelength center (less than 700 rim) [22] see (Fig. 3, top).

364

Hill and Bendall [23] assumed in 1960 that the postulated two photoactive centers may be arranged in series within the electron transfer chain and that the coupling should be effected by cytochromes (but see subsections VIC, E). In an energy diagram, this concept results in the so-called Z scheme (see Fig. 5, bottom, p. 372).

In 1961, evidence for the coupling of two centers in series was given by spectroscopic phenomena observed by Kok, Duysens and Witt and their co-workers. In these experiments it was shown that an intermediate reaction of photosynthesis followed by absorption changes shows antagonistic behavior whether light of less than 700 nm or greater than 700 nm be used. In this way it was shown that oxidation of electron carriers such as Chl-al, Cyt-f and PQ is supported by far-red light (greater than 700 nm) but their reduction is supported by shorter wavelength light (less than 700 nm) [24-27] . Furthermore, it was demonstrated that absorption at 515 nm and fluorescence at 685 nm is small in far-red light, but enhanced in red light [26,28].

The photoactive chlorophylls within the two antennae pigment systems, which drive the whole photosynthetic machinery, have been elaborated by the pulse techniques as recounted below.

VA. Chlorophyll ai (P- 700) VA-1. A dimer as reaction center o f antennae system L Absorption changes at 700 nm

were discovered in 1957 by Kok and attributed to the reaction of a pigment, P-700 [29]. In vivo, a further bleaching at 438 nm was observed by Rumberg and Witt [30] and, later, a splitting in a double band at 682 and 700 nm [31 ]. The difference spectrum is shown in Fig. 2B. It closely resembles that observed by Seifert and Witt for the oxidation of a chlorophyll a in butanol [32]. An additional small band was observed at 810 nm [33].

The attribution of the spectrum to one and the same pigment, chlorophyll a, was established by identical kinetic behavior upon variation of external conditions [30]. Because the pigment can be excited with far-red (730 nm) light [29] it must be the reaction center of the long wavelength antennae system I, and it is therefore designated chlorophyll a~.

From the double band noted above we concluded that two Chl-a I molecules are combined in one reaction center with an oblique structure which would cause symmetric exciton splitting of the energy levels of the twin [31 ]. Further evidence for the existence of the dimer has been obtained from ESR measurements [35] and circular dichroism analysis of the double band [36]. Model studies on dinrers are reviewed in Ref. 37. In vitro, the redox potentials of dimers are only slightly different (70 mV) from that of the monomeric chlorophylls. The advantage of a dimeric reaction center, therefore, must be motivated by reasons which are, to date, not known.

VA-2. Photooxidation and vectorial electron ejection. The reaction of Chl-ai is extremely fast and takes place within 20 ns. This process represents a primary event [17]o Evidence for this follows also from our observation that Chl-a I oxidation can be stimulated in the light even at -150°C [39].

Three experiments give evidence that Chl-al is oxidized in light. (1) Potassium ferricyanide oxidizes Chl-al in the dark. Because the light-induced absorption change of Chl- al disappears under these conditions, it follows that Chl-at is oxidized in light [40]. The redox potential has been estimated as +0.43 and +0.45 V [40,41]. Values between 0.4 and 0.5 V are discussed in Ref. 44. (2) With reduced phenazonium methosulphate (PMS-) the recovery of Chl-ai absorption changes is strongly accelerated, i.e., Chl,a I is reduced in the dark by PMS- and, therefore, oxidized in the light [30]. (3) The expected radical

365

behavior of Chl-al was established by ESR measurements [42]. These measurements were refined. The kinetics of the ESR signal correspond to those of Chl-al absorption changes [33,44].

Because Chl-al is present as a dimer (see above) the unpaired electron of the radical cation must be delocalized over two Chl-a[, i.e., it exists as (Chl-a~)~.

The mechanism of the photooxidat ion of Chl-ai is by no means comparable with chlorophyll oxidations in solution. This became obvious with our finding that the electrons released from Chl-a~ are shifted across the membrane (see subsection VIIK). The electrons are vectorially ejected in less than 20 ns from the excited chlorophylls at the inner side of the membrane to the outside. It is evident, therefore, that a highly organized chlorophyll center must be responsible for this extraordinary feature. Perhaps the properties of the dimeric Chl-a I have an essential function in this respect. Probably also complexation with proteins plays an important role. Details on the structure of this center have not as yet been elucidated. For further remarks on this item see subsection VB-2.

VA-3. The phases of rereduetion. We found from the time course of the absorption change that Chl-a~ is reduced in three phases: 10 /.ts, 200 /xs [45,46] and 20 ms [30] due to the redox states of the electron donors (see Fig. 2B). The donor for the 10/Js kinetics is probably Cyt-f [45,73]. The 200/xs reaction is due to the oxidation of plasto- c}/anine [73] and the 20 ms period is caused by the oxidation of plastohydroquinone via PC [47]. These redox reactions are outlined in more detail in the section on electron transport (see Section VI).

VB. Chlorophyll-all (,°-680) The photoreaction center of the antennae system II is of special interest because this

reaction is the driving force for the cleavage of water and evolution of oxygen and thereby the energy source for almost all living systems.

VB-1. Reaction center o f antennae system II. D6ring, Witt and co-workers [50--52] observed absorption changes which have been attributed to the reaction center of system II. The difference spectrum is shown in Fig. 2C [51 ]. It closely resembles that for oxidation of chlorophyll a, apart from a blue shift [32]. The same characteristic kinetic behavior at all wavelengths in dependence of different parameters indicates that the spectrum is due to one substance [51]. It can be excited only with light of less than 700 nm. Far-red light (728 nm) which activates predominantly Chl-a I does not affect this pigment. Therefore, it must be the center of the short wavelength antennae system I1 and has been designated chlorophyll arl. The location at tile site of the H20 oxidation is supported by the fact that the Chl-all reaction is inhibited by DCMU under repetitive excitation conditions in the same way as the cleavage of H20, while the reaction of Chl- a~ is not at all inhibited in the presence of system I electron donors [51 ].

Conclusive indication for the existence of Chl-aii as a dimer does not exist (see subsection VB-2), in contrast to the case of Chl-a~.

A small absorption change observed in the infrared at 825 nm is also assumed to hldi- cate the formation of the Chl-a~i cation [53].

VB-2. Photooxidation and vectorial electron e/ection. Because a product of the reaction of Chl-ail , the electric field, is generated in less than 20 ns (see subsection VIIIB), we concluded that the reaction of Chl-a H in the light takes place also in less than 20 ns [2]. This fast response indicates a primary event. The response of Chl-all at cryogenic temperatures supports this conclusion [54]. However, whether under these conditions the reaction is reversible or not, is controversial (see Ref. 55).

366

Chl-all may be oxidized or reduced in the light. However, reduction arid formation of Chl-a~i, respectively, is in contrast to the following results. We have shown that a special plastoquinone, X-320, accepts an electron from Chl,aii in less than 1 ~ (see subsection VIC). I fChl~ i is the donor of X-320, Chl-aii should be reoxidized within less than 1 ~ . The absorption change indicates, however, that the reaction time of Chl-all takes up to 200/as (see subsection VB-3). Therefore, Chl-ati must be oxidized in the light [56]. This is in accordance with the fact that the plastoquinone anion, X-320, is a primary product (see subsection VIC).

Support for oxidation in the light is given furthermore by the similarity of the difference spectrum with the spectrum of the photooxidation of Chl-a in vitro [32]. A flash- activated reversible ESR signal was also attributed to the existence of a radical cation of Chl-all [57]. Its kinetics correspond to those of the Chl-ail absorption changes [124]. The midpoint potential is presumably greater than 0.8 V because Chl-a~i finally oxidizes water (see subsection VID).

Electron spin resonance signals from non-oxygen-evolving preparations treated with detergents show a narrowing of the linewidth of the spectrum which should indicate the existence of a dimer [58]. However, these conclusions are not free from ambiguities: (a) because the measurements were carried out under non-physiological conditions where no cleavage of H20 takes place; and (b) because narrowing of the linewidth per se is not conclusive - it occurs also in the case of monomers in suitable solvents.

The photooxidation mechanism of Chl~ii is, as in the case of Chl-a I, completely different from that of chlorophylls in solution. We have shown also in this case that Chl- ali undergoes a vectorial redox reaction, i.e., one electron of Chl-a~i is vectorially ejected in less than 20 ns across the membrane towards the primary acceptor PQ(1) (-=X-320) (see subsection VIIK). How this transfer takes place is a fundamental but open question. Quantum mechanic 'tunnels' or electron 'pipelines' via overlapping n-electron systems of chlorophylls or carotenoids may bridge the gap between Chl-ai,ii and their primary acceptors. Perhaps also intermediate electron carriers with certain differences in midpoint potential and appropriate distance within the membrane may support the vectorial transfer and the stabilization of the charge separation.

Possible mechanisms of vectorial electron tunneling have been discussed by Kuhn [65b] and Hopfield [65c].

VB-3. The phases of rereduction. The time course of the absorption change of Chl-aii indicates that Chl-a~t is reduced at least in three phases: approx. 3/as [59], 35/as [60] and 200/~s [51 ] (see Fig. 2C). A further very fast phase (much shorter than 1/as) might exist because the time resolution of tiffs measurement was restricted to the/as range. Be- cause of the short lifetime of the signals of Chl-aii in the red, interference with the million- times greater chlorophyll fluorescence takes place. This problem has been overcome with a light beam modulated at the very high frequency of 14 MHz, which allows the elimina- tion of the fluorescence [52a].

The times of rereduction probably reflect different redox reactions within that enzyme, S, which is responsible for the cleavage of water. As outlined in subsection VID, S has to accumulate stepwise four oxidizing equivalents by four turnovers of Chl-a H before two H20 are split into one 02, four e- and four H ÷ [62]. It might be that the kinetics observed at Chl-aH, ~n, reflect the turnover times, Tn, of the conversion of these states S~-$4 into each other (see Fig. 4b and discussion in subsection VID). That in principle the kinetic pattern of Chl-aii depends on the redox state of the enzyme system, S, has been shown in Ref. 66.

367

Refined studies indicate that the reduction kinetics of Chl-a~t depend strongly on the t f concentration in the inner phase of the thylakoids but only slightly on the H ÷ concentration of the outer phase. (This was shown for the 35/~s phase [63].) The dependence of the Chl-a~i reduction on pHin supports our conclusion that Chl-all is probably located towards the inner side of the membrane (see subsection VIIIC).

For further details on Chl-atl see reviews in Refs. 55 and 64.

VC. Coupling of chlorophyll ai with chlorophyll ali Upon addition of DCMU the reversible absorption changes of Chl-a I and Chl-ali dis-

appear [51]. Under these conditions the artificial electron donor PMS- can, however, supply Chl-a~ with electrons, i.e., Chl-al is reactivated by PMS- but not Chl-aii. These results indicate that under normal conditions Chl-aii supports Chl-ai with electrons, i.e., both are coupled in series [51 ].

Assuming the same extinction coefficient for Chl-aii as for Chl-ai, it can be estimated from the absorption changes that about one molecule Chl-aii is active per electron chain [56,60]. This means that the ratio Chl-al : Chl-an is about 1 : 1, a reasonable value when both centers are coupled in series.

VC-1. Stoichiometrics. However, counting quantitatively the Chl-a I and Chl-all molecules which are acting together in the linear electron transport chain, it follows for isolated spinach chloroplasts that Chl-ai1 is in abundance. The ratio is Chl-at : Chl-all = 0 .85 :1 [74].

Furthermore, it was established that 25% of the total amount of Chl-ai is not engaged in the linear electron transport chain but is functionally isolated [74]. Perhaps, this excess Chl-a I (25%) is responsible for a cyclic pathway of electrons (see subsection VIF) and is topologically separated from the linear electron chain as already proposed in Ref. 88.

Chl-ai and Chl-aii are probably not different in their chemical constitution but only in their special environment. A functional and antennae pigment-depleted Chl-ai or Chl- aii center has not been isolated to date.

With respect to the question of whether a singlet or a triplet state preceeds the photooxidation of the Chl-ai and Chl-aii, it appears that in vivo the singlet states are involved [71b1.

VI. Electron transfer *

VIA. Pn'mary electron acceptor of chlorophyll ai: ferredoxin The nature of the primary acceptor is controversial. Absorption changes at 430 nln of

a pigment (P-430) have been attributed to an iron-sulfur protein and bound ferredoxin, respectively [33]. A midpoint potential of -730 mV was observed [65a]. From ESR experiments it is proposed that another substance, X, is the primary acceptor acting between Chl-a I and P-430. There is no evidence that X is a pheophytin. For a detailed discussion see Ref. 44.

Between the final electron acceptors, NADP ÷ and Chl-al, two other electron carriers are discussed, ferredoxin, Fd, and ferredoxin-NADP ÷ reductase, (a flavoprotein containing FAD) [67,68].

* (see Fig. 5, page 372).

368

VIB. Donor site of chlorophyll al: plastoeyanine Plastocyanine - a copper containing protein - was isolated by Katoh et al. [72]. It is

a two-electron acceptor with a midpoint potential of +370 inV. From flash-induced absorption changes it results that PC is reduced in 20 ms simulta-

neously with the oxidation of pQz-. PC is oxidized in 200/is [73] simultaneously with one of the reduction phases of Chl-a~ [45]. PC is, therefore, an electron donor for Chl- a~. That PC is the main donor is strengthened by the ratio of Chl-a I : PC = 1 : 1 [73] which indicates that Chl-ai + PC corresponds to a capacity of three electrons. This capacity satisfies that of System I which we estimated in an independent way to be three [47].

On the other hand, Cyt-f, which was originally discussed to be the electron donor for Chl-al, acts probably only on a bypass of the linear electron transport. Cyt-fis reduced in 20 ms and oxidized in approx. 10 ~ts [45,73]. The I0 ~ts time corresponds to one of the three reduction times of Chl-al (see subsection VA-3). However, only 15% of Chl-a~ are reduced within this time via Cyt-f. This is also understandable from the deficiency of Cyt- f: the ratio of the active reactants is only Chl-a I : Cyt-f= 1 : 0.4 [46]. The role of Cyt-f parallel to PC is as yet unknown. For further discussion on PC and Cyt-f see reviews in Refs. 72 and 75 and recent data in Ref. 53a.

VIC. Primary electron acceptor of chlorophyll ali: plastoquinone X-320 The difference spectrum of a reactant X-320 is depicted in Fig. 2D, p. 361 [69,71a]. It

was shown that X-320 is a plastoquinone molecule, PQ(1) in a special environment, alternating between the fully oxidized and semiquinone anion state. It is functioning as a primary acceptor of Chl-ali and as connector molecule, PQ(I) between Chl-all and the pool of plastoquinone (see Fig. 5, p. 372).

The conclusions are based on the following seven results [47,69,70]. (1) X-320 is formed rapidly, in less than 1/~s [71a]. (2) It can be generated even at

-160°C [70]. (3) X-320 is formed by excitation of Chl-aii but not of Chl-a I. (4) The ratio X-320 : Chl-aH is 1 : I [69]. (5) The kinetic behavior of X-320 in the presence of DCMU is in accordance with the properties which are expected for the primary electron acceptor of Chl-ali [70]. (6) The decay time of X-320 (0.6 ms) corresponds to the rise time of the formation of plastohydroquinone [47] (see Fig. 2D, E). (7) Tile spectrum of X-320 corresponds to that of PQ- which has been produced artificially in butanol [71,71d].

ESR studies on PQ(-1) have failed to detect such signals. This may be due to unusual broadening of the ESR spectrum through interaction with other paramagnetic centers [44].

The fluorescence quencher, Q, which is assumed to be identical with the primary electron-acceptor, corresponds in its kinetics to that of X-320 [64]. The midpoint potential is about +0.04 V. A pheophytin (midpoint potential -0 .6 V in vitro) as electron carrier between Chl-alx and X-320 has not been observed to date.

Absorption changes at 550 nm of the so-called C-550 component are probably due to a shift of the absorption band of chromophoric molecules in response to the generation of X-320 [55]. C-550 serves, therefore, only as an indicator and is not identical with an electron acceptor as was originally supposed [55].

Evidence for a further acceptor in a sidepath at the level of Chi-an, the role of which is not clear, is reviewed in Ref. 71c. The finding of a second midpoint potential of ~0.24 V may be related to this result [113].

369

VID. Donor site of chlorophyll ail: H20 The ultimate electron donor of Chl-a~i is H20. With the oxidation of H20 , liberation

of 02 and H ÷ takes place. The oxidation is catalyzed by an enzyme system, S. Its turnover time per electron has been estimated by the double pulse method in the following way.

(1) The passage time necessary before 02 evolution from H20 can take place a second time is approx. 0.6 ms [76] (see Fig. 4a). (2) The time for an electron transfer from Chl- at[ into the PQ pool also takes 0.6 ms. This follows from the passage time necessary before the charge separation and the reduction of the PQ pool can be generated a second time (see Fig. 4a) [76,181]. On the other hand, the reoxidation time of PQ(l) (X-320) has been estimated independently as 0.6 ms [47] (see Fig. 2D). This means that within the electron transfer sequence

[PQ pool + - - - - PQ(1) + - ' - " Chl'all ~- S-H20]

the reoxidation of PQ(1) is rate limiting. Therefore, the turnover time for the cleavage of H20 might be in the order of 0.6 ms. Such a value has also been found by another method, discussed in Ref. 62.

It has been shown by Kok and Joliot [62] that S has to accumulate stepwise four oxidizing equivalents (four positive holes) by four turnovers of Chl-an before two H20 molecules are split into one 02, four electrons and four protons. It has been postulated in a formal model that these steps represent four states of c+ c2+ _~ c3+ ~ ~4+ Ol ~ 02 - - 03 ~ 04 --~ So, whereby the transitions from $44 + into the initial state, So, is librating O~ in the dark (see Fig. 4b). It is open to debate at which state the uptake of H20 takes place. Also the nature of the S states is unknown except for the involvement of four manganese molecules (reviewed in Refs. 62 and 77).

(1) The oxidation times, T1-T4, of the four S states should be in the order of the H20 oxidation, i.e., 0.6 ms (see above). They have been estimated indirectly and are not known precisely. The values range between approx. 200 tzs and 1 ms [62,77]. Reduction

ii/ , -! J=: 0 • I

-~ Ca. E

_ o

lime t d between two single turnover flashes

Fig. 4. a. Recovery time of 02 evolution, reduction of the PQ pool and charge separation (meas,~ red through the voltage za4~) as a function of the time, t d, between two l]ashes [76,181 ].

out , l . h ~

/ poo, Chiiall 1

,.... ................... J--L.. _

in I I/2 H20 • 20ns

t X-320(-PQ-)

? 1/2 H20

0.6ms

.2.hVjj

~; / p°°' Chl;Oll; 1#,02 IH*

370

hv I hv I hv [ hv I

.,. I Oh{ell Chlo. H [Chlali Chla;, Ij ChlQII ChlQ.II Ch[o,i Chln..lll Chloll

/ I ÷ \ 2 T1 ,,-/~@e- S ; ~ . _ . ÷ ~ , / ' e - / * \ / e-fl

~ S 0

, , , lq 2 H 2 0 [ (H*) i (H*) I (H*) I (H*)

I I I I ] l • I, I

f I I t I

02 tl t2 t3 t4

reduction times of ChlaTi

Tn ox ida t ion

t imes of Sn

tn r eleo, se t imes

of protons

Fig. 4. b. Possible correlation between the reaction times of Chl-ail, splitting enzyme system S, T n (subsection VID) and the release of H ÷ On the stoichiometries of H ÷ see the end of this subsection.

rn, (subsection VB-3) the water from H20 , tn, (subsection IXC).

of the states $2 and $3 can be accelerated by the addition of chemicals such as thiophenes and CCCP [78]. In this way the lifetime of the charge accumulation can be shortened 100-fold. This effect is an advantage in several approaches [78a]. The mode of action of these agents is not known.

(2) The different reduction times, %, of Chl-a~i (approx. 3 #s, 35/as and 200/~s) (see subsection VB-3) may be correlated with the release of the electrons from the S states but this has not as yet been shown directly fm any of these values.

(3) There exists a third set of reaction times accompanied with the water oxidation. These are the times, tn, for the release of the four protons from two H20. The tentative values are t2 ~ 100/Is, t3 ~ 300-600/~s and t4 ~ 1 ms [188] (see subsection IXC). The sequence of the release of the protons in respect to the four oxidizing states So-$4 is not known with certainty. It has been reported for O, 1, 1,2 H ÷ [78b] and 1,0, 1,2 H ÷ [78c,78d].

A more thorough analysis of the interrelations between Tn, rn and t n may give a deeper insight into the mechanistic details of the enzyme system S (see Fig. 4b).

The possible chemical meaning of the S states has been outlined in a model by Renger [79]. In this scheme the highly reactive intermediates are assumed to be stabilized at groups containing manganese as central ion.

Other components such as chloride [79a], cytochrome b-559 [79b] and quinones [79c] have been inferred to be involved with the process of water oxidation. However, the functional role of these compounds still remains an open question.

VIE. Link between chlorophyll ai and chlorophyll all: plastoquinone pool Plastoquinone has been shown to be a component of the electron transport in photo-

synthesis [80]. The midpoint potential is +0.06 V at pH 8. With our finding of the transient absorption change of PQ in the ultraviolet (see

Fig. 2E) the estimation of the functional meaning of PQ was possible [81,69,47]. The following rdne results demonstrate the unique function of PQ as (a) electron link

between Chl-al and Chl-all, (b) electron buffer in the form of a pool, (c) interconnector of several electron chains in form of a strand, (d) hydrogen pump, and (e) regulator of the electron transfer rate.

371

(1) PQ is reduced by exciting Chl-aii. pQz- is finally reoxidized by Chl-a~. It is, therefore, located between Chl-a I and Chl-ali (Fig. 5). (2) The extent of the absorption change indicates that PQ exists as a pool of about seven molecules between both centers [86a,69]. Maximally four can be reduced in saturating light, i.e., because of being divalent, PQ exists as a dynamic storage device for eight electrons. A pool was observed also by the size of the 02 gush after the onset of 02 evolution [82] and the kinetics of the fluorescence induction [83]. (3) The connector between the PQ pool and Chl-aH is the semiquinone anion PQ(-D (=-)(-320) which was already discussed as the primary acceptor of Chl-all in subsection VIC. (4) For the mechanism of the generation of pQ2- from PQ(1) it was postulated that two PQ(I)ions (X-320)of two electron chains dismutate into pQ2- and PQ [47]:

PI Q *- PIQl +- 2 hv - (1) PiQl -Ch l - an

pQZ- PQ1 PQl-Chl-al l

Recently, evidence was given that an additional two-electron acceptor molecule is located between the PQ(1) molecule (X-320) and the PQ pool [84,85]. It is very likely that this molecule is also a PQ molecule, PQ(2). The reaction sequence suggested is:

PQ~2-) " PQ(1) ~ PQ(-2) " PQ(-I) ~ PQ(-2) " PQo) +- PQ(2) "PQfi) ~ PQ(2) "PQo) (2)

(5) The PQ molecules in the pool are reduced into a hydroquinone in 0.6 ms and reoxidized at 20°C in 20 ms [47]. The reoxidation within 20 ms has been. identified as the rate limiting reaction of photosynthesis [2]. (6) The PQ pools of the least ten electron chains are interconnected with each other in the form of a strand and chain cross-connection, respectively. This was concluded in principle for the following reason. If most (e.g. 90%) of the Chl-aii centers are blocked by DCMU, all Chl-a 1 centers can nevertheless carry out redox reactions because the remaining Chl-a n (10%) can supply, after 10 turnovers, all Chl-a I centers with electrons from H20 via the assumed electron guiding PQ strand. For details see Ref. 86.

(7) The PQ pool probably works as a pump for bound hydrogen across the membrane [ 190]. (8) PQ operates, furthermore, as a regulator of the electron transfer rate. The regulator and pump are discussed separately in detail in subsections IXE and IXF.

VIF. The electron transfer chain The electron carriers discussed above represent together a linear sequence of transfers

of electrons as outlined in the chain in the upper part of Fig. 5. The characteristic absorption changes of its individual components are listed in the bottom part of the upper figure. Transfer times between the carriers are depicted in the top. On the basis of the light energy absorbed by Chl-ai and Chl-ali and the approximate midpoint potentials of the carriers, one can construct an energy diagram for the electron transfer as is depicted in the lower part of Fig. 5. According to thermodynamics, under natural conditions about 70% is available as free energy [13a]. It is obvious that there nevertheless remains available energy of about 0.3 eV from hp I and 0.4 eV from hPii . This energy could be primarily stored if the electron transfer occurred vectorially from H20 at the inside of the membrane to NADP + at the outside of the membrane. Such a charge transfer would produce a transmembrane electric field. This has been measured as outlined in Sec- tion VII. In fact, from the proof of such a field the vectorial charge separation has been concluded.

Several investigations have been devoted to the analysis of the cytochrome f ' s (see

372

- 2On:

N A D PLFF~ -D- x '' /~C

( I

I I

L I I I I

/ /

/ ~ 3 S

682.700nm

10ps ~_20ns 200ps 20ms 0,6ms 20ms

X-320 [

I I I I I

415 431 563nm

\

hv u

=lps 35ps

200ps 0.6 ms

H*

I 407 320nm 425 \

I ~09 \ 555nm 439 435

60Ohm 263nm 559nm 682nm

Volt

- -1.0

---~--

NADP" 0

Cht-o I'

- . 1 . 0

hVl X-320:

PQ pool ~ . . . ~ - -

~ . ~ l c y t - f)

C h l - a I (PT00)

Chl-a~

hVll

H20

Chl-all (P680)

Fig. 5. Top. Electron transfer chain with transfer times and optical data of the electron carriers [2]. Bottom. Energy diagram of the electron transfer and midpoint potentials (see subsection VIF).

373

subsection VIB) and b's. As yet no agreement has been reached on its function and location within the linear electron transport chain. Speculations on a possible participation in the H20 oxidation and in the so-called Q-cycle are mentioned in subsection VID and subsection IXF-1, respectively.

The outlined electron pathway from HzO to NADP ÷ is termed 'linear electron transport'. It has been shown that electrons can be recirculated from Chl-a I (via a Cyt-b) back to Chl-a I. This transfer is termed a 'cyclic electron transport chain' [4,5]. This cyclic pathway may be topologically separated from the linear one (see subsection VC-1).

VII. Electric field generation

VIIA. Detection by electrochromism Experimental evidence was given in 1967 by Junge and Witt [89] for the existence of

electric fields across the functional membrane of photosynthesis first reported in Ref. 1. The electric field was measured by so-called 'field-indicating absorption changes', &A.

Electric fields can induce (1) physical changes of the absorption spectra of the pigments in the membrane through 'electrochromism' or (2) chemical changes between the pigments in the membrane which change the absorption spectra of the pigments, e.g. through a displacement of a dissociation equilibrium. It has been shown that the under- lying mechanism is due to 'electrochromism' (see subsection VILE).

Fig. 6 shows in a simplifie~l form the principle of electrochromism (a refined descrip- tion is presented below). Absorption of light transforms a pigment molecule from its ground state to an excited state. If an electric field is applied and if one state has a different dipole moment or polarizability from the other, the energy difference between these states depends on the magnitude of the electric field. Thus, the absorption band of such a molecule is shifted. This band shift causes 'field-indicating absorption changes', AA. Because the lifetimes of absorption changes due to events other than electrical fields are not shortened by substances like ionophores (which increase the membrane permeability), one can discriminate between field-indicating absorption changes and optical changes arising from other events.

The spectra of the absorption changes whose decay can be accelerated by ionophores are presented in the upper part of Fig. 7. On the left-hand side field-indicating absorption changes of chloroplasts from green plants are shown [90] and on the right-hand side, those from of chromatophores from photosynthetic bacteria [91]. Because the absorption bands of all pigments present in the membrane respond to the electric field, these spectra are rather complicated compared to the schematic one shown in Fig. 6.

without with electric field

0

A

l w i t h o u t - ~ - w i t h electr¢ / j \ ,.,o

t, ig. 6. Simplified scheme of the mechan i sm of electrochromism.

C

~ C ~._o .c ~.

~ m ~ g

AA

/

374

A I / I

10-3

0

2~

m

S,~r.',, 7,~. ,,ght ~,,,,,,:,d

.r \ o,, ~,,,or.,,,,°,,,,

~-" ..... - i ,~ , L -

i i ...... 20°5

~ decay lOrns- ls 653nm

t ' ~ total eLectricolly induced on carol i , chlorophyll and carotenoid

eno d mult i layer s

- 7 ~- - ~ . . . . . 7J. 7

V / chlorophyll b

t I I I 400 500 600 700 n m

w a v e l e n g t h

/1, electr ical ly ~nduced on

lutem m o n o l a y e r s

4O0 ~

wovelength

Fig. 7. Top. Spectra of the field-indicating absorption change induced by light on chloroplasts and chromatophores of bacteria [90,91 ] (subsection VIIA). Bottom. Spectra of the absorption changes of Chl-a, Chl-b and Car multilayers in a microcapacitor induced by an electric field in the dark [ 102 104] (see subsection VIIE).

VllB. Kinetics of the electrochromic changes The field-indicating absorption changes are characterized by a rise time shorter than

20 ns [92] and a decay time in the range of 10 ms-1 s depending on the membrane permeability and pH gradient, respectively (see subsection VIIL). The time course is shown in Fig. 8, left. Recently, it was reported that the rise time in isolated chloroplasts is biphasic [93]. However, reinvestigation has not confirmed this result. It was found that the effect was due to double hits of the reaction centers [94]. *

It has been demonstrated: (1) The fast rise of the changes in the light indicates the formation of an electrical field and electric potential difference, 2x4~, across the thylakoid membrane. (2) The decay in the dark indicates the breakdown of the field by ion fluxes. (3) The indication is based on electrochromic changes of the bulk pigments in the thylakoid membrane. These statements are illustrated in Fig. 8, bot tom right. Support for the scheme is given in the following sections. An electrical analog is depicted in Fig. 8, top right. The light reaction, which shifts electrons across the membrane, is symbolized by a generator which charges the membrane capacitor. The electrochromic effect of the bulk pigments (Chl-a, -b and Car) indicates the electrical potential changes, which are symbolized by a voltmeter. The discharge of the membrane capacitor occurs by ion fluxes through different channels. These are symbolized as resistors which are specified in the following sections. For reviews see Refs. 2, 3 and 34a-c .

VIIC. Mapping of the versatility of information obtainable by the electrochromic method As outlined in detail below, the electrochromic change has the properties that:

* According to a new technique developed in this laboratory, the rise time of the field generation is less than 3 ns (Trissel, H.-W. and Gr~iber, P., unpublished results).

375

8 u

.c_

/ 20ns

I I 2 0 0 m s

t i m e

[ Phospho- Voltmeter hv Basal r yioting Ar tificical

J ~ f lux flux flux ~ - - a, ~ " . *VMC,

c ,-o . . . . . . . . . . . . . . . . . . . . . . . . .................. ................ ! ..............

Car-~l n ~ ' ' J ' ) K+H + H + K +

ner v current

Fig. 8. Left. Time course of the field-indicating absorption changes [51. Right. Scheme of the electric events on the thylakoid membrane of chloroplasts with the electrical analog [2] (see subsection VIIB).

(1) It represents an intrinsic molecular voltmeter and ammeter respectively; (2) The response is prompt; (3) The voltage and current are indicated linearly; (4) The signals can be calibrated in terms of absolute values.

With this 'instrument' the following far-reaching information has been obtained, and this is discussed in the next sections:

(5) A light-induced electric field is set up across the membrane; (6) The field is generated by a vectorial shift of electrons from the inside to the out-

side of the membrane; (7) The electron shift is initiated by the ejection of electrons from the excited Chl-al

and -all; (8) The charge separation gives evidence for asymmetric arrangement of the electron

carriers within the membrane; (9) The two 'generators', Chl.a I and Chl-ali, are located towards the inside; the

primary electron acceptors are located towards the outside; (10) A vectorial pathway of electrons, H ÷ and bound hydrogen, in the form of a zig-

zag scheme, has been evaluated. More sophisticated analysis of the electrochromic changes gave information on the fol-

lowing topics: (11) Location and orientation of the bulk chlorophylls; (12) Complexation of bulk chlorophylls with carotenoids; (13) Phase transition and conformational changes within the membrane; (14) Generation of surface potential differences; (15) Coupling of the field decay with phosphorylation; (16) Utilization of the field energy for phosphorylation; (17) Discrimination between the different hypotheses on the mechanism of phos-

phorylation. These many types of information obtained by the electrochromic method can be obtained in principle also for other biological membranes or interfaces in general. If they are not pigmented, they can be stained with dyes (for details see subsection VIIN).

VIIC-1. Discrimination between electrochromism and other phenomena. The maximal electrochromic changes occur at approx. 515 nm (see Fig. 7). Therefore, analysis of the field-indicating absorption changes are carried out mostly at this wavelength. However,

376

measurements at 515 nm may be confused with changes at 515 nm due to other events with different kinetics and different spectral changes in the blue and red.

(1) Fast changes at 515 nm (rise less than 20 ns, decay 3/as) are due to the formation of triplet states of carotenoids (see subsection IVB).

(2) Slow changes at 515 nm (in the range of seconds) may be due to oxidation of Car caused by pH changes in the inner space of the thylakoids [95].

(3) Further slow optical changes at 515 nm (in the range of seconds to minutes) are due to light scattering caused by structural changes induced through pHin changes [105].

In view of this situation, it should be pointed out, therefore, that conclusions drawn from slow absorption changes at 515 nm can lead to misinterpretations if careful, thorough analysis is not carried out. However, most of the basic events of photosynthesis take place within 10 ms, (20°C) or less. Therefore, we could restrict the following conclusions on the analysis of the fast electrochromic absorption changes which can be recognized unambigiously as such. This is the case because the field-indicating changes can be separated kinetically from others in flash studies and, furthermore, because they can be tested by the response to ionophores (see subsection VILE-2).

Apparent contradictions such as: (1) anomalous electrochromic behavior of nmtants; (2) decrease of the electrochromism at low temperature; (3)shifts due to solvatochromism; and (4) increase of electrochromism due to chlorophyll oxidation, have been explained quantitatively by the complexation of chlorophylls with carotenoids. This is outlined in detail in subsection VIIIG.

VIID. Properties of electrochromism In liquid solution, three effects contribute to electrochromism [96,97] : (1) a change of

the angle between the molecular transition dipole moment and the electric vector of the light wave by orientation of the molecular dipole in the electrical field ('orientation effect'); (2) direct influence of the electric field on the transition dipole moment, resulting in a change of the shape of the absorption band ('transition moment effect') and (3) a shift of the absorption band by the electric field ('band shift effect').

In lipid layers the electrochromic spectra can be explained in a first approximation by the band shift effect alone. However, if the permanent dipole moments of the molecules have no preferential orientation relative to the electric field, the band shifts of the different molecules will go in opposite directions, so that on the whole a 'band broadening effect' results [98,99].

The band shift effect is based on the potential energy of permanent and induced molecular dipole moments in the electric field. Due to a difference between the permanent dipole moments, /ag and /ae, and between the polarizabilities, ag and %, of the ground and excited states, the field causes a change of the difference between the ground and excited energy level of

h a y = (pie - ~lg) f 1 O/g)F2 ~ ( ~ e - (3 )

where

(Ill e - - [ l g ) F = ]].i e Jig] • f " cos 0' -+

is the scalar product of the dipole moment difference and the field strength. ( a e agUE 2 is the product of the induced dipole moment difference ( a e -- a g ) F and the vector F. In the case of carotenoids, this is given approximately by Aall "/72 . cos20 where Aall is the polarizability difference parallel to the long axis of the molecule, and 0 is the angle

377

between the molecular axis and the field direction [100]. (Note: 0' in Eqn. 3 is different from 0 .)

A shift of the whole absorption band by Av (which is always small as compared to the width of the band) yields a change of the molar absorption coefficient, e, at a fixed frequency of

Ae= ,Se (--Av) ] (62e~ (--Au) 2 + ( ) (4) (~-v) v + ~ \~-~-f F ""

Inserting Eqn. 3 and introducing the wavenumber ~ = v/c, one obtains approximately

A e - II*e --l~gl cos 0 • + - - - - - " COS20 "F 2 he' ~-~u] F 2 hc V

1 llJe-lagl 2 2 / 6 2 e ] + h2c2 cos 20' (5) 2 • F ~ 5 } V

The first term depends linearly and the second and third term quadratically on the field strength. This has also been shown experimentally [ 101 ].

If the molecules have a permanent dipole moment difference with a fixed preferential orientation relative to the field (so that the average of cos 0 ' is other than zero), the first term in Eqn. 5 is predominant at field changes in the order o f F = 1 • l0 s V/cm.

In this case: ( I ) the spectrum of absorption changes should follow the first derivative of absorption bands (SA/6V)F; (2) the shape of the spectrum should not depend on the field strength; (3) the magnitude of the absorption changes should be approximately proportional to the field strength.

VILE. Proof o f the electrochromic method I~7IE-l.a. Spectroscopic evidence - analysis o f the shape. The properties 1 to 3

formulated at the end of the last subsection (VIID) correspond closely to those of the field-indicating spectrum in Fig. 7, top, left [90] :

(1) In the red region the peaks at 648 and 660 nm are located antisymmetrically to a center at 653 rim. This shift is similar to the derivative of Chl-b band at 653 nm. The peaks at 668 and 680 nm are located antisymmetrically to a center at 673 rim. This shift is similar to the derivative of a Chl-a band at 673 nm. In the blue spectral range the interpretation is complicated due to the superposition of Chl-a, Chl-b and Car shifts (each carotenoid has three absorption bands in the blue).

(2) The shape of the spectrum in Fig. 7 is independent of the field strength, i.e. the location of the maxima does not shift with light intensity changes (unpublished results).

(3) The amplitude of the spectrum is proportional to the field strength (see subsection VIIG).

VIIE-I.b. Spectroscopic evidence - comparison with model systems. To substantiate the conclusion that the spectral changes in Fig. 7 are field-indicating and due to electrochromism, multilayers of lipids with incorporated molecules of Chl-a, Chl-b and Car were built up on glass slides. These layers were exposed in the dark between two electrodes to electrical fields of up to 1 • 10 6 V / c m and the resulting absorption changes were measured [ 102-104]. The superposition of these electrically induced changes is depicted in Fig. 7, bottom, left. The agreement with the light-induced spectrum in chloroplasts in Fig. 7, top, left, is good. Only, in the red spectral region the spectrum in chloroplasts is much more structured. This is due to the fact that different aggregates of Chl-a bulk pigments

378

are present in vivo; on the multilayer, however, only one Chl-a type is present. For an optimal fit a nrean angle of 0 ~ 74 ° between the long axes of the bulk carotenoid molecules and the normal to the membrane has to be assumed [104].

The method of measuring electric events by field-indicating absorption changes in green plants has been extended by Jackson and Crofts [91] to photosynthetic bacteria. The corresponding absorption changes are shown in Fig. 7, top, right. The spectrum has been attributed to changes of carotenoids. The fast rise of these changes is less than 1/as [91,106] as in the case of chloroplasts. The light-induced spectrum has been mimicked by exposing the membrane to a diffusion potential [91]. To substantiate the conclusion that the spectral changes in Fig. 7, top, right are band shifts of carotenoids in a field and are due to electrochromism, the multilayer technique was applied in this case also. Layers of lipids incorporated with molecules of carotenoids (lutein) were built up on glass slides. When these are exposed to an electrical field, absorption changes result as depicted in Fig. 7, bottom, right [103]. The agreement with Fig. 7, top, right is very good. In this case, no parameter had to be varied for an optimal fit (in chloroplasts the mean orientation of the carotenoids was assumed to be 74°). This agreement without any parameter variation justifies the application of the multilayer technique to the three- component system in chloroplasts as shown in Fig. 7, left.

VIIE-2. Kinetic evidence. If the interpretation in Fig. 7 is correct, it should be possible to change the field decay and the attributed absorption change into a faster one by increasing ion fluxes across the membrane. This corresponds to a shunt parallel to the basal ion flux (see artificial channel in Fig. 8). Such channels have been realized by treat- ment of the membrane with ionophores as Gramicidin D (GmcD) [89] which increases the permeability specifically for alkali ions. If the decay rate ( i / r ) reflects the discharge of a loaded membrane by ions and if one type of ion, e.g. K ÷, is in excess, it is expected that:

1/~-(+CmcD) = const. CK+" CGmcD + 1/~-(-CmcD) (6)

This relationship has been verified [89]. Under optimal physiological conditions the ions which are responsible for the decay rate, 1/r(-GmcD), are predominately protons (see subsection VIIL).

VIIE-3. Electrical evidence. When in a chloroplast suspension two macroscopic electrodes are located at a distance of d ~ 1 cm near the top and bottom of a cuvette and a non-saturating flash is fired through one electrode from the top, each thylakoid becomes to a small extent asymmetrically charged (see Fig. 9, left). This is because in the upper part of each thylakoid more light is absorbed than in the lower part. The corresponding potential difference

A~top -- A~)bottom = A A ~ (7)

between the upper and lower part induces a signal at the electrodes [107]. Fowler and Kok observed in this way that the signal is diminished by one-half when one of the two light reactions is blocked. This confirms our earlier result that each light reaction contributes one-half to the potential (see subsection VIIK). The electrically measured signal decays in 10/~s. The signal measured by the field-indicating absorption changes, 2xO, however, decays in approx. 100 ms, i.e. 1 " 104 times slower. This discrepancy was explained by Witt and Zickler [108] as follows. The electrically measured decay is caused by the equilibration of the asymmetrically charged thylakoid via ion fluxes in the water phase parallel to the plane of the membrane (rij ~ 10 Vs). However, the voltage measured by the

379

T ~ Icm

voltmeter

A thyla

<~+ J¢ /

k0id B

Q.

'ID

1 o

0.5

E difference of the electrical potent~l difference AA~(elec.) measured by electrodes

• |

• t / I /

I I 0.5 1

Fig. 9." Left. Scheme of the electrostatic measuring technique [107,108]. Right. Fie/d-indicating absorption change, zXq~, as a function of the electric response of macroscopic electrodes, A ~ , when thylakoids are excited by non-saturating flash-light [65,181] (see subsection VIIE-3).

field-indicating changes decays by ion flux perpendicularly through the lipid phase of the membrane (r± ~ 100 ms), i.e. rtl < < r±. To be able to measure r± by the electrodes, it is necessary to prevent the equilibration via the membrane surface in 10/as. This holds if ~'ll > r±. A drastic increase of rlf has been realized by increasing the viscosity of the aqueous phase from 1 to 103 cP through addition of 3 M sucrose, r3 has been decreased by addition of the ionophore Valinomycin. Under these conditions, zll > r±, the electrically measured time course of AAq~ is indeed in fair agreement with that of A4~ measured by the field-indicating absorption changes [108].

The kinetic agreement provides further support for the view that the 'field-indicating absorption changes' indicate electrical potential differences.

In Fig. 9, right, we compare A~bop t measured by the field-indicating absorption changes with AA~e I measured by the electrodes. The signals are proportional to each other. Because small values of AACel are proportional to the total voltage across the membrane, it can be concluded that the field-indicating absorption change responds linearly to the electric potential [ 108,181 ]. This supports our previous results discussed in subsection VIIG.

VIIF. Orientation, polarization and localization of the field 1. The electrodes can respond only to electron shifts perpendicular to the membrane

surface (see A in Fig. 9, left) and not to shifts in the plane (see B in Fig. 9, left). The observed response, therefore, indicates that the field-indicating absorption changes are due to field changes oriented with one component perpendicular to the plane of the membrane. This supports the conclusion drawn from the response of Aq~(t) to ionophores (see subsection VIIE-2).

2. If the outside of the membrane is charged negatively and the inside positively, then in the experiment of Fig. 9 the upper part of the membrane is more negative than the

380

lower one. In this case a negative charge should be induced on the upper electrode. This was observed by a corresponding response of the macroscopic voltmeter in Fig. 9. There- fore, the light-induced field is polarized from a positive inside of the membrane to a negative outside. This is in agreement with the observed passive efflux of cations and passive influx of anions [ 109,110].

3. In subsections VIIK and VII1C it will be explained that with the photooxidation of the two photoactive chlorophylls, one electron each of Chl-al and Ctrl-atI is shifted rapidly (in less than 20 ns) from the inside of the membrane to the outside. Therefore the field generated simultaneously with this charge separation is first localized as a dipole field.

4. In a second stage this localized field is delocalized over the thylakoid membrane by ion redistribution within the inner and outer aqueous phase of the menrbrane.

Evidence for the existence of a delocalized field follows from the fact that only one pore per thylakoid (set up by Gramicidin D) discharges the membrane by recombination of all the charges via this single pore (see subsection VII.l). The time for the conversion of the localized field into a delocalized one can be calculated from the time, r,, which is necessary for the equilibration of the non-equilibrated charge distribution set up by a non-saturating flash (see Fig. 9). Under physiological conditions this time is approx. 10/is (see subsection VIIE-3). The equilibration takes place at 200 active centers. The time, r, of the delocalization between two centers is therefore approx, r ~ 0.1/is. The delocalization by ion distribution can be blocked by cooling. This is outlined in subsection VIIIG, paragraph 4, p. 395.

VIIG. Linear indication of the field Different observations indicate that the extent of the field-indicating absorption

change, AA, at all wavelengths is linearly proportional to the electric potential, A¢. (1) In subsection VIIE-3 this was shown by the proportionality of AA and Aq~op t respectively, with the electrostatic induction 2xA¢ on electrodes. (2) In Fig. 4a, subsection VID, this is demonstrated through the parallelism between &4 and the amount of electrons translocated from H20 to the PQ pool. (3) In subsection VIIK it is concluded from the observation that ALl as well as the translocated charges are doubled if, after one light reaction, both are excited. (4) In subsection VIIH experiments are cited which indicate that AA responds linearly to the diffusion potential set up across the membranes [91]. (5) Further support was given by comparing charge collection and field-indicating absorption changes at low temperatures [ 11 l ].

The linearity is the basis for the calibration of &A in absolute electric values (see subsection VIIH). The linearity is, furthermore, of importance for conclusions on the molecular arrangement between chlorophylls and carotenoids within the membrane (see below).

According to Eqn. 5 in subsection VIID, a linear relationship is expected only for those pigment molecules which have a permanent dipole moment difference, A~, between the excited and ground states. A/l does exist for the chlorophylls but not for the carotenoids. Therefore, the absorption changes of carotenoids should be proportional to the square of the field strength (see Eqn. 5). However, the field-indicating absorption change at 515 nm, which has been attributed to the carotenoids (see Fig. 7), responds linearly to the field strength (see above).

Furthermore, the extent of the change at 5 l 5 nm is more than ten times larger than that expected according to the second term in Eqn. 5. Sununarizing, one can say that the linearization and amplification of the observable absorption changes of the carotenoids

381

in vivo correspond to the qualities of the first term in Eqn. 5. This contradiction was explained by the assumption that the carotenoids in vivo are

exposed not only to the light-induced field, F, but additionally to a strong permanent field, Fp, perpendicular to the membrane [102,103]. In this case the absorption, A, is expected from Eqn. 5 to be

A =A o +b(Fp +F) z (8)

where A 0 is the absorption without any field. The proportionality factor b depends on the polarizability difference and on the first derivative of the absorption band (see Eqn. 5). The field-induced absorption change z£4 = A(F) - A (F = 0) is given by

~A = 2bFpF + bF 2 (9)

If Fp is large compared with F (2 • l0 s V/cm, see subsection VIIH), the term bb a can be neglected. This is the case if

Fp >~ 2" 106 V/cm (10)

Then, with 2bFp = const.

&A v const. F (11)

In this way the linearity between &A and F is formally explained. However, we are left with the essential question of according to which mechanism the carotenoids can be exposed to a permanent field. It has been shown that this is possible if the carotenoids are complexed with a polarizing molecule. Evidence is given that this molecule is a chlorophyll. Thus, the observed linearization and amplification leads finally to detailed insight into the molecular organization of the membrane. This is outlined in subsection VIIIG.

VIIH. Calibration o f the field The field-indicating absorption changes have the specification of a molecular volt-

meter. In subsection VIIG it is shown that the field-indicating absorption changes, &A, depend linearly on the field strength and thereby also linearly on the transmembrane voltage, Aq~, i.e.

A~ = Q/C=a • &A (12)

where Q = charges translocated across the membrane during the absorption changes (~A), C = capacity of the membrane, and a = proportionality factor. Because the derivative of the absorption changes with respect to time indicates the current, i, arising from the charges, the former also have the specification of a molecular ammeter

i = C. d(A4))/dt = C" a . d(~L4)/dt (13)

Absolute data for A4~ and i have been obtained in three different ways: 1. In our first approach the number of charges which are translocated across the mem-

brane in one turnover was used for calibration. In subsection IXB-3 it is shown that two elementary charges, 2e, per electron chain are translocated. One electron chain covers an area So "~ 1 • l0 s (A) 2. For the thickness (l) of the isolating layer of the membrane we use l ~ 30 A. With the effective dielectric constant of phospholipid membranes, D ~ 2, it follows that in one turnover for the initial potential difference [112]:

1A~=a" lZ£4 Q - 2 e ' l - ~ 50 mV ( 1 4 ) C S o ' D ' D o

382

and

1 F ~ 2 • 10 s V/cm

With Eqns. 12 and 13 it follows that with arbitrary values of ZKzl :

50 mV • kA electric potential difference - 1 ~ z 1 - a •

a = 50 mV/lZ£4 and

(15)

(16)

1/dr 50mY d(AA) current density j = ~ - . . . . . . . (17)

cnl 2 AA dt

The onset of the field in less than 20 ns amounts to a current density inside the field generator of j ~ 1 A/cm 2. The field decay in 20 ms amounts to a current density across the membrane of]" ~ 1/aA/cm =.

2. In a second approach, diffusion potentials have been set up by Jackson and Crofts across the membrane of chromatophores of bacteria. This has been done by salt jumps in the presence of ion translocators which increase the membrane permeability specifically for cations or anions. The resulting absorption changes have been calibrated in volts by assuming that the Nernst equation is valid [91] (C = salt concentration of monovalent ions):

Ac~ = (RT/~7) In Cin/Cout (18)

In comparison with the extent of the light-induced absorption changes, electric potentials have been estimated which are comparable with those obtained for chloroplasts according to the method mentioned above. The authors have concluded from their experiment that, in bacteria also, the field-indicating absorption changes, zX/I, depend linearly on the electrical potential change, Aq~ (see subsection VIIG).

Corresponding salt jump tests with chloroplasts are not unequivocal [1 l 5]. The optical changes observed were caused mainly by light scattering and so forth [116]. But recently in new experiments light scattering effects have been eliminated [117]. Under such conditions the salt pump method indicates that in a single turnover flash the field-indicating absorption changes in chloroplasts correspond to a value of

1 A ( / ) = 15-35 mV (19)

3. In a further approach we used the ionophore Alamethicin for the calibration of the absorption change [118]. Alamethicin is known to form voltage dependent pores in black lipid membranes [119]. Incorporation of Alamethicin in thylakoid membranes leads to a sharp increase of the transmembrane current above a critical voltage, Aq~c, which indicates a gating. Above A~bc Alamethicin is drawn by electric forces into the membrane, forming channels of varying sizes for ions, especially for K +. Below Aq5 c the current drops back to the slow basal one (see Fig. 10) [118]. The relation between Alamethicin concentration and the critical voltage, AqSc, found in black lipid membranes [120] has been used to calibrate the field-indicating absorption change in absolute transmembrane electric potentials [ 118] :

1 a q ~ c 1 - a ~ c 2 = ~60 mV" log c2 (20)

C1

383

E

1.0 0 . 8

i 0./,

c~ 0.2 c c~ u

"1o c -o 0.1 (b

~Ac1 h A o

÷ Alamethicin

C1:10"6/ml //

° " " ° " ~ . ~ ' . ~ o _...." I I

5815m-V- , . . . . . &A,~; 2

&Ao

*Alamethicin

C2=lO'Sg/ml

i

~ o .,.,,,.. ~ ° ~ o ~

I I I f f I I I 0 100 200 H 0 100 200

time m s

Fig. 10. Field-!ndicating absorption change in a single turnover flash as a function of the concentration of a voltage depending ionophore (Alamethicin) [ 118] (see subsection Villi).

13 is known only for artificial lipid layers. With 13 = 0.7M3.9 we obtained, according to Fig. 10:

~A~b ~ 105--135 mV (21)

This value might be overestimated. 4. In subsection VI1N, other techniques are discussed for measuring transmembrane

electric potentials. These methods are, however, slow and not applicable for conclusions on single turnover condit ions with the exception of a microelectrode method. In this lat ter case a value of 1A~ ~ 40 mV [121] was obtained.

The values available by the methods discussed are listed in Table I. Because of the divergences of the results, we use in the following as approximation

IA~) ~ 50 mV (22)

For the electric potential difference in the steady state see subsection VilM.

TABLE I

ELECTRICAL POTENTIAL DIFFERENCE AT SINGLE TURNOVER CONDITIONS

Method: ! 2 3 4

l ~ ( r n V ) 50 15 35 105 135 40

384

VII.I. Functional unit of the electric field The acceleration of the electrical field decay by ionophores has been discussed in sub-

section VIIE-2. The decay can be achieved with extremely low concentration of the ionophore Gramicidin D (see Fig. 23, top, left, p. 408). Only one pore of Gramicidin D per 1 " 10 s Chl molecules or per one thylakoid, respectively, causes more than 50% acceleration of the field decay [89]. The result of this 'titration experiment' is evidence for the assumption that the functional unit of the electrical events is one thylakoid or a larger unit, i.e. several thylakoids interconnected. This indicates that the electric field is a collective property of a closed vesicle. This conclusion is supported additionally by the fact that over the whole range of Gramicidin D concentration used, the electron transfer measured via the 02 evolution is not influenced at all (see Fig. 23, top, p. 408).

The thylakoid vesicle must be also the 'unit' for protolytic reactions and the generation of a pH gradient, because these processes represent a subsequent stage of the charge separation (see Section IX), i.e.

A~ unit = A pH unit = 1 vesicle (23)

It would be of great interest to know whether the collapse of the field-indicating absorption changes by one channel per vesicle is also valid when, instead of the thylakoids, vesicles with a size of quite different order are used, e.g. the chromatophores of photosynthetic bacteria. Indeed, the collapse of the field-indicating absorption changes in the chromatophores starts at a value of one Valinomycin per 3 - 103 BChl molecules [122] (Fig. 23, bottom, left). This number of molecules is spread over the area of a vesicle with a diameter of about 500 A, which is the size of a chromatophore [122].

VIIK. Generation of the field by electron ejection from the excited chlorophyll a~ and a{i When both light reactions are excited in one turnover flash the field-indicating changes

have a magnitude as shown in Fig. 18a, p. 398. When, however, the Chl-al reaction is chemically eliminated with DCII" + ferricyanide, the amplitude of the absorption changes diminishes by about one-half [112]. When, on the other hand, the contribution of Chl- ali is prevented by DCMU, the amplitude of the absorption changes again diminishes by one-half (Fig. 18a, p. 398). This indicates that, in a single turnover, one-half of the field or potential change is set up at each light reaction center:

h p I ~ 1/2 • 1A~b and hvIi ~ 1/2 • 1A~ (24)

This conclusion has been supported additionally by measuring the action spectra of the absorption changes at 515 nm. Two spectra have been obtained which correspond to the action spectra of Chl-a I and Chl-ali [112].

VIIL. Decay of the field by ion fluxes In isolated chloroplasts the field decay in the dark does not coincide with the efflux of

protons [ 183]. At low internal H ÷ concentration this is due to better conductivity of the membrane for other ions such as Mg 2+, K ÷, CI-, etc. However, with decreasing internal pHin from 8 to 5 and increasing ApH, respectively, the rate of the field decay increases 20-fold [123]. Similar results have been obtained with Chlorella cells (Gr~iber, P. and Witt, H.T. unpublished results) (see Fig. 11, left). (The different ApH values were established by preillumination of various intensities). The results indicate that under these conditions it is the proton which contributes predominantly to the discharge of the field. At ApH 3 the field decay and proton effiux occur in approx. 10 ms.

E

o .Q o 5

o ~t' ,y, Lt~

0 05 1 15s t ime

e~

3 8 5

'k

10. 25

s gs ,o

~ c ~ .':2 26

m

"5 T , , J , , , i 0 600 1600 2"-OOms

ttme

Fig. 11. Left. Decay time of the field-indicating absorption change as a function of the ApH value across the thylakoid membrane of chloreIla cells (Gr~/ber, P. and Witt, H.T., unpublished results). Right. Time course of the field- and pH-indicating absorption change on chromatophores of bacteria (see subsection VIIL) [125]. (Left and right at single turnover conditions.)

7,6 ~_

5,0 0

D

In chromatophores of photosynthetic bacteria the field decay by H ÷ has been shown directly by comparing the decay of the field-indicating absorption change with pH- indicating absorption change [125] (see Fig. 11, right). The decay of the field coincides approximately with the effiux of H ÷.

Refined analysis of the different pathways of the proton fluxes across the membrane - a basal and a phosphorylating one - are outlined in subsection XIF.

VIIM. Electrochromic measurements in the steady state VIIM-1. Total electric potential difference. After the onset of steady-state light, the

electrochromic changes increase up to a factor of four in relation to excitation with a single turnover flash: z~xAmax/IZ~ ~ 4 [126b] i.e., the maximum voltage amounts to A~' = 200 mV. This increase is due to the filling up of the PQ pool with eight electrons [47]. This potential drives cations outwards and anions inwards (see subsection VIIF). Protolytic reactions with the separated charges result in an active inward translocation of H + and decrease of pHin (see Section IX). The decreasing pHin , on the other hand, increases the field-driven H ÷ efflux (see Fig. 11, left). In the steady state, the H + effiux fully compensates the active H + influx. The initial high potential, AqT, decreases with the internal acidification towards a relative low steady-state value, SSA4). This is caused by the increase of the H + efflux (see subsection VIIL) and by the retardation of the electron transport (see subsection IXE), both due to internal acidification. In the case of steady- state light on isolated chloroplasts, the electrochromic absorption changes, SSAA, may be subject to error because of slow optical changes which are not caused by the field (see subsection VIIC-1). Such effects have been exlcuded, however, in measurements with Chlorella cells through a special calibration technique [114]. In this way one obtains in saturating light SSAA/azL4 ~ 2. According to Eqn. 16 this corresponds to a total

electric potential difference (ss) ~ 100 mV (25)

A similar value was obtained from delayed light emission (see subsection VIIN). On the other hand, studies of ion distribution and with microelectrodes result in a value in the order to 10 mV (see subsection VIIN). Rumberg [34] explained this discrepancy in the steady state as follows. Intrinsic probes such as electrochromism and delayed light emission respond to potential differences between the inner and outer membrane surface

386

(100 mV), whereas extrinsic probes such as ion distribution and microelectrodes respond to potential differences between the inner and outer aqueous bulk phases (10 mV). This means that in the steady state electrochromism responds to an additional potential, SSA~ + SSA4, while the ion distribution method is sensitive to SSAq5 only.

a • SS~A = SSA4~ + SSA4 ~ 100 mV (26)

The additional potential was proposed to be a surface potential difference, SSA4,between the inner and outer surface of the membrane. SSA4 is probably generated by the acidification of the inner thylakoid space (see next paragraph and Fig. 12a).

VIIM-2. Surface potential difference. Biological membranes are loaded with surface charges. The thylakoid membrane bears at the outside negative surface charges at pH 7 [126]. The corresponding counter ions are solvated in the water phase. This gives rise to a surface potential, 4, between the surface and the solution. 4 depends on the ion concentration, c. According to Gouy-Chapman, the relation between the surface charge density, a, 4 and c is

s inh(~ ' 4/2 RT) = const, o/x/~ (27)

if monovalent ions are present only. If we assume in the dark under symmetric conditions pHin = pHou t = 8, the value 4out - 4 i n = A~j across the membrane is zero (see Fig. 12a, '1 ').

In the light, charge separation takes place and Aq~' is set up between the inner and outer phases of the membrane (Fig. 12a, '2'). In subsequent protolytic reactions the pHin value is decreased. In saturating steady-state light, pHin decreases from 8 to 5. Thereby large amounts of H + are translocated inward and are bound to buffering groups of the inner surface (see subsection IXD). Thereby the inner surface charge density, o, can change to a more positive value (see Fig. 12a, '3'). This may result in the formation of a surface potential difference, SSA4, within the membrane. In this case, in toto in the light a potential difference, SSA4~ + SSA4 is present across the membrane (see Fig. 12a, '4'). SSA4 may play an important regulatory role in respect to ion translocation [187].

SSA4 was estimated in two ways. (1) A~ was first estimated by Rumberg and Muhle from Oin and Oout according to

Eqn. 27 [127]. It resulted that under saturating steady-state light conditions and c 1 • 1 0 -2 M the extent is

SSA4 ~ 80--90 mV (27a)

(2) In a further approach A 4 was estimated from the field-indicating absorption changes. That part of the electrochromic changes, SSAA, which is due to SSA4 (see Eqn. 26) should depend on the ion concentration, c (see Eqn. 27). If, on the other hand, the dependence of ssA4~ on c can be neglected, the plot of SSAA as a function of c should reveal SSA 4. Assuming for isolated chloroplasts a total transmembrane voltage of 100 mV as in the case of Chlorella (see Eqn. 26) one obtains under saturating steady-state light at c -~ 10 -a - 10 .2 M as a preliminary result (Ref. 128 and Tiemann, R., Zickler, A. and Witt, H.T., unpublished results):

ssA4~ ~ 20 mV SSA~j ~ 80 mV (28)

~SA~j is in fair agreement with the value of Rumberg and Muhle [127] (see Eqn. 27a). In single turnover where the A~b creating ApH value is negligible (see subsection IXD) it is

1Aq~ ~ 50 mV XA4 ~ 0 (28a)

387

in

pH8

AqJ=0

out

pH8 pH8

hv

E}I i -

i - i -

B

pH8

ApH

3

/.C / =

÷ / /

pH5

m T . . . . . . . . . _ ~-'~ ssA~, T /

pH8

Fig. 12. a. Extent of the electric potential, A4~, (due to charge separation) and the surface potential, A@, (due to acidification) during the onset of steady state light across a thylakoid membrane (see subsection VIIM). The pH values indicated are those of the aqueous bulk phases. For the pH values at the boundaries see text in subsection VIIM-3.

Summarizing, it results that in the steady state two different types of electric potential, SSA¢ and SSAff, are responsible for the electric potential difference, ssA¢ has to be considered if one regards the bulk phases, and SSAq~ + SSAff if one regards the boundaries

of the membrane (see next subsection VIIM-3). V11M-3. Electrochemical proton potential. The electrochemical potential difference of

the proton defined between the outer and inner bulk phases is in the steady-state (see

(Fig. 12.b):

SSAGH÷ (bulk) = 9 " SSA~b + 2.3 RT SSA pH (bulk) (29)

with SSA¢ ~ 20 mV (see Eqn. 28) and SSApH (bulk) ~ 3.3 (see Eqn. 36, p. 401).

Because in each phase SSG is constant from the bulk up to the boundary of the membrane and this independent of change of the electric and chemical component within one

phase, it follows

SSA GH+ (bulk) =- ssA GH+ (boundaries) (29a)

The electrochemical potential difference of the protons defined between the outer and

inner boundaries (see Fig. 12. b):

SSA GH+ (boundaries) = ~'" (SSA~b + SSAff) + 2.3 "RT ssA pH (boundaries) (30)

S~A4) + ssA~b ~ 100 mV (see Eqn. 26)

388

mV

-- 180 o el

0 .o 120

z , - .m,.., u

60

0 c- o e- U

ApH

between "1- ~'~,, boundories O.

z 2

betweenXX~ SSA~.SSA~ :IO0~V ~-~ 1 bulks J ~ _ ~ u

ssAq) = 20mV ;" -4 5s k-- 0

time

between bulks ~

SSApH : 3.3

/ / ' \ ] / " beiween SSA-S - 2 /" boundaries V

-4 ss k--

Fig. 12. b. Proposed time course of the electric potential A~, A~ k and Aptf between tile inner and outer aqueous bulk phases (solid lines) and between the inner and outer boundaries of the membrane (dotted lines) [127]. Excitation with steady state light of saturating intensities. Details see subsection VIIM-2 and VIIM-3.

The value of SSApH(boundaries) can be calculated from Eqn. 29a. It results that

SS6pH(boundaries) ~2 (30a)

The development of the surface potential difference, A~, is compensated by a decrease of the pH gradient between the boundaries of the membrane (see Fig. 12b). Energetic considerations on the proton potential between the inner and outer spaces of the thylakoid membrane can be calculated, therefore, either with the values of the bulk phases (see Eqn. 29) or with the values adjusted at the boundaries (see Eqns. 30 and 30a). For a corresponding application see subsection XIG.

VIIN. Electrochromic measurements with artificial probes; application to further systems; other techniques for electric measurements

The electrochromic method is useful also for the analysis of electric phenomena in other biological membranes or interfaces in general. So far, the method has been extended to photosynthetic bacteria [91]. If the systems are not pigmented they can be stained with dyes. In this way we have shown that, in principle, e.g. with the artificial rhodamine b, the method is also applicable [ 130]. Safranine has been introduced as probe for membranes of mitochondria [133]. Corresponding probes could be valuable also for 'prompt' electric measurement on vesicles of membranes from nerves. However, in the case of artificial dyes one must take care over possible artefacts due to additional optical changes caused, for example, by photochemical reactions of these dyes. Also, nonphysio- logical interactions of the artificial dye with the membrane have to be considered. The probe merocyanine [131] used in chromatophores of bacteria is obviously not useful for measurements: The rise of the absorption change of merocyanine in bacteria has a time lapse of 50 ms after the rise of the field-indicated intrinsic carotenoid absorption change. The lapse was explained by the assumption that the carotenoids respond only to local fields and that the delocalization takes 50 ms. However, we have shown that the delocalization takes only 0.1/as (see subsection VIIF). Furthermore, at cooling to -35°C the absorption changes of merocyanine vanish whereas the electrochromic changes of the

389

intrinsic carotenoids as well as the electron transfer are present in full [1321. It may be that a reorientation of merocyanine in the field takes place lasting approx. 50 ms, whereby the time response may be delayed.

In comparison with other methods (see below) the electrochromic method with intrinsic pigments has the advantages of (a) its unrestricted time resolution, (b) 'broad- casting' the electric events from an undisturbed biological system and (c) its relative tech- nical simplicity in respect to the variety of information obtainable (see subsection VIIC). For measurements of very slow potential changes however, one has to eliminate other slow optical changes (taking seconds or more) such as scattering, etc., as has been pointed out in subsection VIIC-1.

(1) Indication of slow electrical potential changes has been dervied from the intrinsic changes in delayed light emission caused by the recombination of oxidized chlorophyll with negative charges [134,135]. It was deduced that the steady-state potential is 7 5 - 105 mV [134a] (similar to Eqn. 26).

(2) Ion distribution evaluated by the Nernst equation has been used for estimation of potential difference [136,137]. For chloroplast in the steady state the voltage calculated between the bulks is ss/xq5 ~ 10 mV (similar to Eqn. 28).

(3) Potential changes in mitochondria and chromatophores of bacteria have been measured by observation of transmembrane fluxes of synthetic ions driven by the intrinsic field [138].

(4) By inserting microelectrodes into stacks of thylakoids it was assumed that the measured voltage indicates the potential span between the inner and outer bulk of the thylakoids, although the tip radius of an electrode is of the size of one thylakoid [139]. The measured voltage induced by single turnover flashes is 1A4)~ 40 mV, similar to Eqn. 28a. The voltage measured under steady-state conditions is SSAq~ ~ 10 mV (similar to Eqn. 28). Nevertheless, this value has to be questioned because the ion-conducting hole produced by the injection probably acts like a shunt under steady-state conditions, which must falsify the results.

VIII. Molecular organization based on analysis of the electric field

This section is restricted to information on the membrane architecture based on a refined analysis of the properties of the electric field. Only the organization of the plastoquinones in VIIID has been concluded from other data.

VIIIA. Asymmetric membrane architecture The generation of an electric field is a vectorial event which usually can be achieved

only by asymmetric organization. Therefore, evidence that an electric field is set up across the thylakoid membrane, was the first evidence for anisotropic arrangement of the electron carriers within the membrane. The more refined analysis of the properties of the field and the events induced by it provides at some points quite detailed information on the location of the molecules and the special arrangement of the components within the membrane.

The results were confirmed and extended by immunological studies and measurements with artificial lipophilic and lipophobic electron carriers. This is reviewed in detail in Ref. 140.

390

hv I hv l 1 NADPH H ÷ ~ ) A D P ATP ' p

N A D P . \ H ÷ ) H + ¢) l ~ ] PMout I I I

) _ ~ e - t ~ ~ H* & ( ~ ) + & , l

Chl-al K',,t'tipool Chl-all . I_ (

/ thylokoid H 1 , H "].,, ,', ~i, ,,i.l. ( membrane ~, 4u2 ~, 2n2u / I"'"n ~

Fig. 13. Zigzag scheme of the vectorial pathways of electrons, protons and hydrogens within the molecular machinery of photosynthesis derived from pulse spectroscopic studies [1,2]. The time sequence of the reaction patterns has been evaluated as follows. 1. Excitation of Chl-a I and Chl-all. 2. Vectorial ejection of electrons from Chl-a I and Chl-ali at the membrane inside to the outside and electric field generation. 3. Oxidation of H20 , reduction of NADP +, reduction of PQ and reoxidation. 4. Proton translocation into the inner phase through protolytic reactions with the charges at the outer and inner surface of the membrane. 5. Discharging of the energized state through efflux of protons. 6. Formation of free ATP from ADP + P by the energy released with the efflux of H ÷ via the ATP- synthetase. (See Sections VIII and X.)

VIIIB. Localization of the primary aceeptors and donors The polarity of the transmembrane field which is set up in less than 20 ns provides

evidence concerning the direction of the primary charge separation. Because we detected the negative charges on the outside of the membrane and the positive charges on the inside of the membrane (see subsection VIIE-3), the primary electron acceptors are

sol

sol

outside

_L iptds -- Z

-nnnn- Chl-a UUUUz insi de

NADP÷ H*

~ -6 IPlastoqu'nones ' ATPase ] _ _ ' X- 320

Cyt-

H* H ÷ H *

Fig. 14. Preliminary topography of the molecular machinery of photosynthesis based on functional experiments. The two black 'trunks' symbolize the two photoactive centers, consisting of Chl-a I and Chl-all which probably are complexed with proteins. The porphyrin rings are located towards the inner surface [65] (see Section VIII).

391

located at the outer surface and the primary donors at the inner one. The number of charges separated per single turnover is two (see subsection IXB-3)

and this indicates that two primary acceptors and two donors are present per electron chain.

One primary acceptor (at System II) was identified as a special plastoquinone, PQ(1) (X-320) (see subsection VIC). It was shown that a proteinaceous shield, which can be removed by trypsin, is rendering PQ(I), inaccessible to artificial electron acceptors such as ferricyanide [141]. The primary acceptor (at System I) is probably a special ferredoxin (see subsection VIA).

The two primary donors, located at the inside, are Chl-a I and Chl-a n (see next paragraph). Again, the donor for Chl-ai is PC. Antibody studies are in accordance with the location of PC at the inside of the membrane [142], but see Ref. 129. The donor for Chl-a n is, again, the H20 splitting enzyme system, S (see subsection VID). Different studies are in accordance concerning the location of S at the inside (see subsection VIIIE).

VllIC. Localization o f the chlorophyll reaction centers The onset of the electric field, i.e. the negative charging at the membrane outside and

the positive at the inside, takes place in less than 20 ns (see subsection VIIB). On the other hand, the positive charging of the porphyrin ring, i.e. its photooxidation, also takes place in less than 20 ns (see subsection VA-2). If one assumes that the field generation and photooxidation of the chlorophylls occur simultaneously, this leads to the conclusion that the positive charges at the membrane inside are identical to the positive charges of the chlorophylls. But this means that the porphyrin rings of Chl-a I as well as those of Chl-ait are located towards the inner surface of the membrane [56,143]. This was recently supported by the PHin dependency of the Chl-aii reaction (see subsection VB-3).

Absorption changes induced by polarized light flash reflects linear dichroism of the bands of Chl-a I, suggesting that the orientation of the porphyrin ring is parallel to the plane of the membrane [144,145]. Chl-ai is probably an oblique dimer (see subsection VA-1); in this case the suggested orientation would be the average of both molecules.

Measurements on the polarization of the absorption band of Chl-aiI suggest that the 'red' transition moment is predominantly parallel to the surface of the membrane [146].

For selective information on the orientation by the polarized light technique see Ref. 147.

VIIID. Organization o f plastoquinone as a pool and strand Plastoquinone exists as a pool of about 4 - 7 molecules within one electron transfer

chain (see subsection VIE). From this result and the conclusion that the pool operates probably as a transmembrane H-pump (see subsection IXF), it is very likely that the pool is functionally located between the outside and the inside of the membrane. This may be realized by a corresponding static arrangement or by dynamic tumbling, as lipid-soluble molecules, between the external and internal faces of the membrane (see subsection IXF).

The pools of different electron chains are combined with each other in form of a strand and chain cross-connection, respectively (see subsection VIE). In this way at least 10 chains are electronically and protonically interconnected.

VIIIE. Localization o f the cleavage o f water (1) Chl-aii is located towards the inside of the membrane (see subsection VIIIC). The

terminal electron donor for Chl-a~i is H20. From this we conclude that the cleavage of

392

H20 is probably also located at the inner surface [89]. This argument can be strengthened. The time for the cleavage of H20 is approx. 0.6 ms (see subsection VID). If the cleavage were to take place at the outer surface, the electron librated from H20 would have to cross the membrane from the outside to Chl-a~i at the inside in approx. 0.6 ms. In this case the transmembrane field generated at Chl-aii should be quenched within approx. 0.6 ms. This is not the case. The field decays in 10 ms or more (see subsection VIIB).

(2) The cleavage of H20 is coupled with the liberation of H +. The H + release from H20 can be observed at the membrane exterior only if the membrane is made permeable for H ÷. This is understandable only if the cleavage occurs at the inside [174, 175].

(3) In a single turnover two H ÷ are released at the membrane inside. This was measured with pH indicators located in the internal thylakoid space [176,177] (see subsection IXB-2). Because one H ÷ is attributable to the oxidation of plastohydroquinone, the other must be released from H20. This again supports evidence for the cleavage of HzO at the inside.

(4) The cleavage of H20 can be inactivated by an external pH of 9 if this pH can be equilibrated with the internal space. This also indicates that the H20 splitting center is exposed to the inside [148].

VIIIF. Orientation o f the antennae chlorophylls and carotenoids The coincidence between the electrochromic spectra in vitro and in vivo in Fig. 7 can

be achieved if the linear (not the quadratic) proportions of the electrochromic absorption changes of the chlorophylls in vitro are used for the superposition. However, a term which depends linearly on the electric field strength can be obtained only if the molecular dipole moment differences have a strong preferential orientation relative to the electric field so that the average of cos 0 in Eqn. 5 over all molecules is other than zero. Hence, it follows that the bulk chlorophylls are asymmetrically arranged in the membrane [149].

The orientation of the bulk chlorophylls with respect to the plane of the thylakoid membrane (see Fig. 15A) has been evaluated in the following way [104]. From the surface-pressure/area diagram of chlorophyll a and b on a water surface it can be concluded that Chl-a and -b are anchored in the water surface with the polar C = O group as indicated in Fig. 15, C, D. This is also the orientation on a plane of a capacitor if the orientation remains preserved after the transfer of the layer from the water to the slide. At these orientations a field, F , has been applied to the capacitor as indicated and electrochromic absorption changes of Chl-a and Chl-b registered. To obtain agreement with the electrochromic absorption changes in vivo, the sign of the linear electrochromic change of Chl-b in vitro in the blue region has to be changed but not the sign of the red region. Therefore it can be concluded that the blue vector of the dipole moment difference o f Chl-b in vivo has a component opposite to the electric field (see Fig. 15A). Regarding Chl-a the sign of the in vitro spectrum in the blue and the red region has not to be changed to match the spectrum in vivo. Therefore the vectors of Chl-a in vitro should have the same orientation with respect to the field as in vivo (see Fig. 15A).

The extent of the electrochromic changes caused by the carotenoids depends on the angle, 0, between the long axes of these molecules and the field. For an optimal fit with the electrochromic changes in vivo one has to assume 0 ~ 74 ° (see subsection VIIE-lb). Because the field is perpendicular to the membrane, the inclination angle of the carotenoids in the membrane is c~ ~ 16 °, i.e. on the whole, the molecules lie rather flat. This is in agreement with the results of linear dichroism measurements. They show that

393

in vi tro )

> chlorophyll b ) >

>

...---"~'plane of the capacitor "0 =(plane of the water surface}

D

chlorophyll a /

-(plane of the water surface)

C

in viva

chlorophyl[ b and

chlorophyll o

outside

insbde f

plane of the membrane

A outside

L u t e ~

~nslcle

B Fig. 15. Left; C, D. Orientation of the chlorophylls in vitro on a water surface and plane of a capacitor, respectively [104]. Right, A. Orientation of the antennae chlorophylls in the thylakoid membrane [104]. The 'red' and 'blue' arrows indicate the vector of the dipole moment difference. Right, B. Complex formation of antennae carotenoids (lutein) with chlorophylls in the thylakoid membrane (see subsection VIIIF) [153]. c~ ~ 16 ° in respect to the plane of the membrane.

caro tenoids are preferent ia l ly parallel to the membrane [150] . Such results were con-

f i rmed by polar ized f luorescence studies [ 151 ].

VIIIG. Complexation of antennae chlorophylls with carotenoids Because the e lec t rochromic caro tenoid changes in vivo respond l inearly to the

l ight- induced field, F , and because they are amplif ied, i t was necessary to pos tu la te that

394

the carotenoids are exposed to a permanent field of Fp >~ 2 - 106 V/cnl perpendicular to the membrane (see subsection VIIG). It is proposed that Fp may be realized by a complex formation of the carotenoids with polarizing molecules. Such a molecular complex can be regarded as a new molecule having a permanent dipole moment. As a model for this, it was shown that a carboxylic group, inserted asymmetrically into a carotenoid molecule, acts like a local electric field, b] parallel to the molecular long axis. Since F I has an optimal direction to the molecule, it can be smaller than the effective Fp by a factor 1/cos 0, i.e. Fp = Fl/COS 0. With 0 = 74 ° (or c~ = 16 °) a local field o f f I >~ 5.5 • 10 s V/cm would correspond to the value of Fp >~ 2 • 106 V/cm oriented perpendicular to the membrane [152].

As possible candidates for the role of polarizing complex partners of the carotenoids, the chlorophylls have been investigated. It was shown by electrochromic measurements on thin capacitors in vitro that the carotenoid lutein forms asymmetrical complexes with Chl-a as well as with Chl-b. One OH group of lutein is attached to the Mg atom of Chl, which attracts negative charges from the lutein [ 153] (see Fig. 15B).

That such complexes do exist in vivo and that they are responsible for the field- indicating absorption changes of the carotenoids is supported by the following six pieces o f evidence.

(1) Lutein-Chl complexes indeed show linearization and amplification (35-fold) in an applied field F as observed for the electrochromic changes of carotenoids in vivo (see subsection VIIG). This is demonstrated in Fig. 16.

(2) By the complex formation the absorption spectrum of lutein is shifted to longer wavelengths (solvatochromism). The maximum of the electrochromic changes of the

AA 1

lutein- / ' ~ chlorophyll-o-

10-3 complex linear

oc: ~ / ~ 3 \\ dependency

c

450 55Ohm ..o 0 o~ F 1 = 2.105 Vlcm

5 =1/3 F~ 2.10-4

quadratic lutein dependency

0 4~0 - - v SSOn~ wavelength

Fig. 16. Absorption change of a lutein-chlorophyll a complex (top) and lutein alone (bottom) in a microcapacitor induced by an electric field (see subsection VIIIG). In the indicated range between 470-550 nm, Chl-a does not contribute to electrochromic changes. Therefore, both changes (top and bottom) are due to the lutein only. (K.-U. Sewe, unpublished).

395

lutein-Chl-b complex in vitro is located at 517 [153]. This suggests that this special complex accounts for the main part of the field-indicating absorption change in vivo which is located at this wavelength (see Fig. 7, top, left).

(3) The maximum of the electrochromic changes of the lutein-Chl-a complex in vitro is located at 512 nm [153]. Hence, it can be expected that in Chl-b-lacking mutants the maximum of the electrochromic absorption change should appear at 512 rim. This has just been observed for a pea mutant and a barley mutant (Ref. 155 and Heber, U., unpublished).

(4) Since Chl-b is located in the regions of the membrane that belong to Photosystem II [156] the main part of the field-indicating absorption change at 517 nm, &A s17, which is due to the lutein-Chl-b complex, can be observed only if the field is present in the regions of Photosystem II. If, however, Chl-ail is blocked, the field which is generated by Chl-al oxidation can be indicated by the lutein-Chl-b complex in Photosystem II only after this has been delocalized by ion redistributions. However, if this delocalization is hindered by cooling, disappearance of changes at 517 nm can be expected. Such a phenomenon has indeed been observed [ 157].

(5) If only a certain proportion of the carotenoid molecules is distinguished by complex formation from the bulk carotenoids, the electrochromic spectrum of only these carotenoids is characterized by the first derivative of the absorption spectrum (see subsection VIID). However, this proportion has undergone a permanent solvatochromic shift (see above). Such a shift is much greater than the electrochromic shifts induced by the light- induced electric field. This solves the apparent contradiction observed in Refs. 159 and 160 that the field-indicating absorption change of a mutant containing only one type of carotenoids exhibits a wavelength difference of 3.5 nm which is much larger as compared to the first derivative of the absorption spectrum of the bulk carotenoid [158]. A model of two different pools of carotenoids, one of which is electrochromically sensitive and shifted to longer wavelengths, was proposed also from the analysis of field-indicating absorption changes at low temperatures [ 161 ].

(6) The concept of solvatochromic shifts of carotenoids complexed with chlorophylls is, furthermore, supported by the findings that the absorption spectrum of carotenoids in Rhodopseudomonas spheroMes can be shifted additionally (2 rim) to longer wavelengths by chemical oxidation of the bulk bacteriochlorophyll in the dark [162]. This can be explained by an increase of the electron-attracting forces of the oxidized bacteriochlorophyll on the carotenoid [158]. In this way also the results reported in R6fs. 163 and 164 can be explained.

VIIIH. Phase transition and conforrnational changes If a phase transition occurs within the lipids of the membrane, it is expected that the

activation energy of the basal ion current across the membrane is influenced by the degree of order within the membrane. The ion current can be monitored by the rate constant of the decay of the electrochromic absorption change. This constant, l / r , plotted versus the reciprocal temperature should reveal a break at the temperature where such a phase transition occurs. In this way a phase transition in the thylakoid membrane has been observed by us at about 18°C (see Fig. 17) [165]. This temperature in accordance with the result from NMR measurements [165a]. The application of this simple method may support forthcoming studies on the state of the membrane.

If conformational changes of the membrane are associated with changes, z2xD, in the overall dielectric constant, D, of the membrane, this effect should be monitored by an

396

-~ s4 J

~- 20 ¢ -

~ 10 ¢ - e -

~ U

>., c:~ 2

o ,-- I

2

L,-

1 9 i i i i

30 21 13 5°C 0 \

0

0

"-, ~18°C

° N \ i J I I I

3.2 3.L, 3.6.10 3 K -1

reciprocal absolute temperature

Fig. 17. Rate of the field-indicating absorption change as a function of the reciprocal absolute temperature [ 165 ] (see subsection XIIIH).

additional field change, AF = - 2 e - A D / S "Do" D 2 (derived from Eqn. 14). In Chlorella cells such additional absorption changes at 515 nm have been observed [168] and interpreted in this way [2]. In bacteria, changes at 515 nm have been interpreted as indicating conformational changes of the ATPase [167]. It is very likely that these contribute to a change of the overall D value for the membrane which results in a AF change as explained above.

IX. Proton translocation - subsequence of the field generation

IXA. External proton uptake and internal release A pH increase in suspensions of illuminated chloroplasts was discovered by Jagendorf

and Hind [169]. In continuous light, amounts of about 300 H ÷ per electron chain are translocated into the inner phase of the chloroplasts [170]. This H ÷ uptake is not stopped by electrostatic repulsion forces because the uptake is accompanied by the transfer of a nearly equivalent number of other ions (e.g. C1- and Mg 2+) [171]. The uptake can be increased several-fold by addition of permeable bases, probably through additional buffering of the internal space. From this result it was concluded that the H ÷ uptake is due to translocation across a membrane and not due to binding at the outer surface. From morphological studies it is likely that the H ÷ uptake within the chloroplast takes place by the thylakoids. But experimental evidence for this assumption was given only by ' t i tration' o f the functional unit of the H ÷ translocation (see subsection VII.I).

According to a hypothesis of Mitchell [172] one mechanism of the H ÷ translocation could be that one H ÷ is taken up by the reduced PQ and one by the proton-binding terminal electron acceptor at the outer surface of the membrane. Furthermore, one H ÷

397

is released with H~O oxidation and one with the oxidation of 1/2 PQH2 at the inside of the membrane (see Fig. 13, p. 390).

IXB. Stoichiometries o f H+/e The mechanistic correlation between electron transfer, field generation and H ÷ trans-

location necessitates the knowledge of the number of H ÷ ions translocated per electron transfer and also of the number of sites of interaction. In continuous light such H÷/e measurements are complicated because it is difficult to estimate the rate of the H ÷ translocation and electron transfer simultaneously with any great accuracy. Therefore, the results obtained differ as H÷/e = 2 - 6 . Jagendorf [173] has discussed the inaccuracy of results with values higher than H÷/e = 2.

To avoid the complications outlined we introduced the H*/e estimation in a single turnover flash. Under these conditions only one electron is transferred and no measurement of the rate of electron transfer is necessary. Using pH indicators and the repetitive pulse technique, the H ÷ transfer coupled with this one-electron transfer can be analyzed with great accuracy. Possible side effects of the indicators due to binding changes, redox reactions, photoreactions, distribution changes between the inner and outer space, etc. were eliminated by subtracting signals which are insensitive to permeable and.non-permeable buffers, respectively. In this way the following results were obtained.

IXB-1. Two sites o f external proton uptake. With different proton-binding terminal electron acceptors there results a ratio of

(H+/e)out ~ 2 (31 )

By blocking Chl-all but not Chl-a I tile ratio is decreased to H*/e ~ 1 (see Fig. 18a). This indicates that at each of the two light reaction centers one proton is taken up at the outside under single turnover conditions [112].

hv I 1 + Hout hVll 1 + ~ Hout (32)

The first H ÷ was attributed to H + uptake by the terminal electron acceptor, the second H ÷ to uptake by reduced plastoquinone, PQ(2). (PQ(I) is not protonated. This was concluded from the independence of the oxidation kinetics of PQ]) on pHout [74].) PQ(2) is protonated at the hydroquinone level, i.e. at PQ22-. If potassium ferricyanide (low concentration) is used as terminal acceptor, one obtains H+/e ~ 1 because reduced ferricyanide does not bind a H ÷ and, therefore, only one H + uptake by PQ is possible [174]. If however, ferricyanide (high concentration) acts as electron acceptor of PQ [166], H+/e = 0 because in this case the electrons are accepted by ferricyanide before reduced PQ can be protonated [ 174].

IXB-2. Two sites o f internal proton release. The H÷/e ratio with respect to the internal phase was recorded indirectly by measuring the pH value of the external phase after leaking out the protons released in the inner phase [ 174,175]. H+/e was measured directly with indicators which respond in the inner phase, such as aminoacridine [176] and neutral red [ 177]. In both cases is is found that

(H+/e)in ~- 2 (33)

Discrimination between the two H ÷ was again achieved by blocking Chl-ali. In this case the ratio is decreased to H+/e = 1 (see Fig. 18a). This means that at each light reaction center one H + is released at the inside under single turnover conditions [176,177]:

hv I ~ 1 H + hVll -- 1 H + (34)

398

Q Chl-al÷ii

i 0 5

CN-o I

I ]2s Voltoge

0 5ms

Chl~l.ll

. c . 9

I I I ~ Z 0

Chl_al H* e-chain

:i:t oo oo,. 1 outside

0

< H* < Chl-aT÷iT Chl-a I

. . . . . . l ___io ~ Proton release '~ =u 0.5 I inside

= ~' I ~ 2

Time Fig. 18. a. Extent of the transmembrane voltage, external proton uptake and internal proton release as a function of the excitation of Chl-a I and Chl-ali, respectively (see subsections VIIK and IXB) [112,176]. (For the absolute time course of the H + uptake and release see next subsection IXC.)

One H ÷ is attributed to the oxidation of 1/2 H20, the other to the oxidation of 1/2 plastohydroquinone (see Fig. 13).

Fowler and Kok [179] reported on H*/e = 3 4 at low flash frequencies using the glass electrode technique. However, their high values were criticized and explained by Saphon and Crofts [180] as an over-estimation of the buffer capacity. For discussion on the so-called Q-cycle model see subsection IXF-2.

IXB-3. The mechanistic meaning of the 14*/e relationship. If the H ÷ translocation is the result of protolytic reactions with the charges separated across the membrane, it can be concluded from Eqns. 3 1 - 3 3 that the potential change is due to a separation of two elementary charges per electron chain. This would be in accordance with the photooxidation of two chlorophylls (see subsections VA and VB). This value has been used for calibration of the transmembrane voltage 1Aq) in subsection VIIH.

Moreover, the result of Eqns. 32 and 34 indicates that the potential change in a single turnover flash is due to a separation of one elementary charge at each light reaction. The field-indicating absorption change participates also in equal proportions at each light reaction (see Fig. 18a). This implies again a linear relationship between field-indicating absorption change, zXA, and potential changes A~ besides the other evidence outlined in in subsection VIIG.

399

IXC. Kinetics o f external proton uptake and internal release The measurement of pH changes induced by single turnovers in combination with the

repetitive technique not only has the advantage that the estimation of the rate of the electron transfer can be circumvented (see subsection IXB) but implies also the possibility of measuring the kinetics of the uptake and release of each H ÷.

(1) For the external proton uptake, rise times on stripped chloroplasts of 20-70 ms [183,112] were observed. When they are broken and isolated in a density gradient the response time is 8ms [112]. This decrease may be due to a change flora a stacked arrangement of the thylakoids to isolated ones. In this case the indicator may be less apart from the surface of the thylakoid membrane. A remaining diffusion layer can be eliminated by sand grindings and uncoupling agents. Under these conditions at light reaction center I the time for the H ÷ upake matches the time for the reduction of the terminal electron acceptor (approx. 1 ms) [184]. At light reaction center II the H + uptake at PQ(2) takes 2 ms [ 184,185] and is obviously truly delayed against the reduction of PQ(2) which occurs in 0.6 ms because PQ(-1) is oxidized in this time [47]. PQ(I) (-X-320) is not protonated (see subsection XIB-1 and Ref. 154).

(2) With respect to the internal phase it is expected that the time of proton release from plastoquinone occurs simultaneously with the oxidation time of PQH2, which is 20 ms [47]. Indeed, such a slow component has been measured in Fig. 18b, left [177].

The time of the release of the proton accompanied by the oxidation of 1/2 H20 should be as fast as the turnover time of the cleavage of H20, or faster. The turnover time of this cleavage is approx. 0.6 ms (see subsection VID). The time measured for the H ÷ release from H20 is about 0.3 ms [177] (see Fig. 18b, right). This value is in fair agreement with the expected one. A refined analysis of this phase results in the individual times, tn of 0.1 ms, 0.3-0.6 ms and 1 ms [188] due to the release o fH ÷ from four different S-states (see subsection VID).

IXD. Evolution of the proton gradient zSpH The initiating processes of the ApH evolution and their time sequence at the area of

light reaction lI are depicted in Fig. 19.

°"

=. ~ {,.16 ~ .g

C ..,_ "T o

. c m I i I

0 sores 0 sores o • - t i m e

Chl-a MI Chl'a[l

proton releose

i n s i d e

Fig. 18. b. Time course of the proton release inside the thylakoid space as a function of the activation of Chl-a I and Chl-aii. Left. H + release due to the PQII oxidation and cleavage of H20. Right. l{+release due to the cleavage of H20 only { 1771 (see subsection IXC).

400

KCl

KCI 1"120 KO~ KC~ OH- .2o ~ E:l .2o

o /f' .............. ' 0ro,0 ,, I '00.,o t ',, L C ~ : , , , ~ , . , , . j . . . . . . . . . . . I ~

Kct ½H:,O I H" I AJPase HC~ KCl KCI L i H*basa I

localized delocolized delocalized field field field (K+)

i +proton gradient I i I

energizotion energizotion of the of the

membrane aqueous phases J h' I t

( I ) z1Ons . ,- ( 2 1 ~0.1~.s (3 ) ~Irns . ( 4 } - t O m s ' - ( I )

Fig. 19. Evolution of the different phases of energization on the thylakoid membrane at the area of light reaction center II (see subsection IXD),

(a) The light-induced charge separation within the membrane results firstly in a generation of a localized field and voltage 2xqSl,,~ , respectively. The localized field is generated in less than 20 ns (see subsection VIIB). In this stage the membrane is energized through the conversion of light energy into electric energy AGel (F • A~b) and redox energy 2xG• (stored in Chl-a{1 and PQ(-i)). 2xGa is converted in the following steps via protolytic reactions finally into AG~ and the energy of a H ÷ gradient (2.3 - R T . ApH).

(b) First, however, the field is delocalized through ion redistributions. This occurs at physiological KCI concentrations (1 • 10 -3 M) within 0.1 /as (see subsection VIIF). At this stage energization is due to a delocalized field, Ag~deloc, between the inner and outer aqueous bulk phases.

(c) In a subsequent protolytic reaction the external uptake of H ÷ (or release of OH-) and the internal H + release (due to the H20 oxidation) results in an excess of one negative ion in the outer phase and one positive ion in the inner water phase. Those ions which are in abundance are now attached to the membrane by Coulomb forces, e.g. CI- and K ÷. Thus, the H + uptake and H + release which correspond to a H* inward translocation do not discharge the field across the membrane. The replacement of the primary localized charges within the membrane by ions of tire outer and inner water phase at the membrane is tile stage at which ApH is generated and tile water bulk phases are energized by A~b and ApH. The time for the ApH evolution at the area of light reaction II is in the range of 1 ms and determined by the H ÷ uptake and release, as outlined in the preceding section. A corresponding development (1) (2)--(3)--(4) takes place at the area of reaction center I.

(d) In the last stage the K* and H + ions -- displayed to the potential span - are driven outwards. At low ApH values the effiux of K + and other counter ions predominate. At higher ApH the H + efflux outruns the other ion fluxes and takes place in the order of 10 ms (see subsection VIIL).

In a single turnover the 'ApH value is small and, therefore, difficult to measure with precision. The range is, however, in the order of:

~ApH <_ 0.1 (35)

401

In steady state light at saturating intensities ApH between the bulk phases amounts to

SSApH (bulk) ~ 3.3 (36)

This difference is due to a pHin decrease from pHin = 8 to pHin ~ 5. The pHin value was first estimated by Rumberg and Siggel through a quantitative analysis of tile rate of oxidation of PQH2 and reduction of Chl-al, respectively, as a function of the internal acidification [189]. In this method it was assumed (a) that the rate of the electron transfer from PQH2 to Chl-a~ is controlled mainly by the pHin value (for a restriction of this assumption see subsection IXE) and (b) that pHin equilibrates with the pHout by addition of gramicidin D. Under these conditions the electron transfer rate could be calibrated in dependence of PHin = pHou t by changing pHou t in the presence of Gramicidin D.

Different other methods like distribution of weak acids, response of permeable pH indicators, etc. indicate a ApH between 2 .5-4 units. For reviews see Refs. 171 and 192.

As has been pointed out in subsection VIIM-3, the pH gradient at the boundaries of the membranes amounts to

SSApH (boundaries) ~ 2 (see Eqn. 30a)

The large difference in the extent of 2xpH on one hand between the bulk phases (ApH'3.3) and on the other hand between the boundaries (ApH 2) indicates that estima- tions of ApH values can lead to misinterpretation if a precise specification is not made as to at which locus within the phase the indicators are situated. This difficulty may be also the reason for the large divergences of the ApH values reported in the literature (see above).

Considering (a) the maximal amount of translocated H + ions (300 H + per electron chain or 60 000 H + per thykaloid), (b) the value of ApH ~ 3 and (c) the internal volume of the thylakoid (50 1 per tool chlorophyll) it can be estimated that only 60 H + per thylakoid are able to move freely, i.e. 99.9% must be bound to buffering groups at the inner surface of the thylakoid membrane.

IXE. The proton potential as control o f the electron transfer Because the electron transfer is coupled with the proton translocation, one expects a

dependency of the electron trans(er rate on pHin as well as on pHou t. It was shown that the rate limiting step in electron flow is the turnover of PQ [2]. Because this reaction is also a protolytic one, probably PQ is the site of control by pHin and pHout.

Regarding first the dependence on pHin, it was observed that if the pftin is changed from 8 to 5, the electron transfer rate is decreased six-fold [189]. The reason is probably the back pressure of Hin on the oxidation of PQH2.

This explanation is supported by the following results. (1) It was shown that the oxidation time of PQH2 depends strongly on pHin [189]. (2) It was furthermore demonstrated that the kinetics of the internal H* release at the PQ site parallels the kinetics of PQH2 oxidation when the latter is changed 20-fold [190] (Fig. 20, center). It is probably not the direct oxidation of PQH2 but the commutation of PQIt2 and PQ to PQH which is limiting the oxidation. This was concluded from the result that the rate of PQH2 oxidation is proportional to PQH2 × PQ [47].

Regarding the dependence on pHout, the external H ÷ uptake at the second PQ~-2) on the outside (not PQ(-o see subsection IXB-1) could bottleneck the turnover of PQ if the pHout is relatively high. Therefore, in general at a constant ApH an optimal pHin and PHout should be required for an optimal turnover of PQ and electron transfer, respec-

402

tively. The experimental result that a mean value of pHin + pHou t governs the electron transfer rate [191] has been interpreted in this way.

A control of the electron transfer via the second term of the H ÷ potential, i.e., SSA4~, is, of course, also expected. However, in the steady state the latter is small (see Eqn. 28).

IXF. The plastoquinone pool as possible hydrogen pump As outlined in subsection XIF, the inward H ÷ translocation is followed by a H + effiux

via basal and ATP-forming pathways. In toto these movements correspond to a H ÷ circulation. This cycle is closed, however, only if between Systems I and II a hydrogen (H ÷ plus e-) is translocated from the outside to the inside (see Fig. 20). Based on protolytic properties of redox reactions in quinone systems, the PQ pool was inferred to be a candidate for a 'pump' for hydrogen (H ÷ + e-) from the membrane exterior to the interior [172]. Recently it was discussed that a pump not coupled to redox reactions may be responsible for this H transfer [192]. Such a pump may operate as in the manner of the light-driven H ÷ pump of bacteriorhodopsin (see below).

However, that the PQ pool might operate as a hydrogen pump driven by redox reactions is supported by the following five pieces of evidence.

(1) The number of active PQ molecules is about four times larger than that of the other members of the electron chain (see subsection VIE). This makes the PQ pool suitable to span the membrane from the outside to the inside. That the pool may be arranged in this way is supported by the following observations.

(2) The primary acceptor of light reaction center II located at the membrane outside is a special PQ(1) molecule (X-320) (see subsection VIC). Obviously, also at least one PQ molecule is located at the inside because the time of reoxidation of PQH2, 20 ms [47], corresponds to the time of the H ÷ release at the inside [177] (see subsection IXC)I

(3) The kinetics of the electron release from PQH2 can be changed drastically by corresponding variation of the turnover of Chl-ai with appropriate far red light intensities. Recently, we have shown that the kinetics of the internal H ÷ release parallel the kinetics of the electron release from PQH2 over a range of 20-fold variation [190] (see Fig. 20, center).

(4) The number of protons, H~ut, taken up at System II from the outer phase increases in parallel with the number of electrons taken up by the PQ pool [194] (see Fig. 20, bottom, left).

(5) Recently, it has also been possible to show that the number of the internal protons released, Hi*n, (corrected for the H ÷ release due to the oxidation of H20) increases in parallel with the nubmer of electrons released by the PQ pool [ 190] (see Fig. 20, bottom, right).

With respect to the mechanism of the hydrogen pumping through the PQ pool, two possibilities can be considered: (a) a diffusion type of mechanism and (b) a hopping type of mechanism. In the diffusion type of mechanism each individual molecule (oxidized or reduced) of the PQ pool is assumed to be able to move statistically between the inside and the outside of the membrane. The hopping mechanism envisages the PQ pool as a lattice-like structure. In this case the transport occurs after PQ collision by a statistical hopping of an electron and proton from one PQ to another. If by chance via mechanism a or b H (i.e., PQH2) is at the membrane interior next to the oxidized plastocyanine PC 2÷, the electron from PQH2 is trapped at this place, whereas the remaining proton is released into the inner space due to its much higher affinity to the internal water phase than to the hydrophobic part of the membrane lipids. The internal H + release into the inner water

403

out hv I hv n r t ) - ~ I ) "I

H + H + )

/ . . . . ~ • PQI PQ - \ ~h°~'~et/fl ....... Cht-al I'-~I,~.PC++ 20m-'~s I/z PQH2-'-'-'///I/Chl-a;l I-;I.,.,.,,,., .... Jl L,..,.~

' / in H+ 4 2 it+ /2 H20

Cx

"5 c

~6

5S'|

2

05

e

0.2 I ~ I 0.2 05 2

rate of reoxidation o f \PO pool

/

I

5s-1

/ / i

/ " / i

electron uplake b~ PQ electron chain

~r

o ~ 4

2

0 0

/ / i

/

electron release from PQ pool electron chain

Fig. 20. Top. Scheme of the proton and hydrogen translocations in the thylakoid membrane. Center. Rate of the internal H ÷ release as a function of the rate of the reoxidation of the PQ pool [190]. Bottom. External proton uptake and internal release as a function of the electron uptake and release by the PQ pool (see subsection IXF) [194,190].

404

phase occurs very probably via protonizable heads of lipids acting as buffer groups at tile inner face of the membrane [ 195}.

For both models one has to assume that due to the long hydrophobic side-chain of PQ, the PQ molecules are soluble within the lipids of the thylakoid membrane. This is supported by the observation that in artificial bilayers of lipids plastoquinones are indeed soluble [186].

It is of interest to compare the mecllanism of the hydrogen pumping PQ pool with the light-driven proton pump realized by bacteriorhodopsin in the membrane of halobacteria (for review see Ref. 196). The essential difference between both pumps is the following. The PQ pump translocates a neutral atom (H*+ e-) in the dark; the rhodopsin pump, however, translocates a charged atom (H +) in the light. The PQ pump works mechanisti- cally relatively simply (see above) because the energy requiring step takes place by a preceding reduction of PQ at the outside (via electron ejection from Chl-a~I) and by an oxidation via PC at the inside (see Fig. 20). This means that the overall process of H ÷ pumping is subdivided into two sequences: (a) a vectorial electron transfer at the Chl-an reaction center from inside to outside; and (b) the vectorial H transfer via the PQ pool from outside to inside.

On the other hand, the rhodopsin pump translocates a proton, H +. Therefore, this translocation must be coupled with the energy requiring act, i.e. with generation of an electric field and ApH formation. Because such a pump has to perform all elementary steps within one highly specialized protein - and therefore, regarded as a system "beautiful for its simplicity" - the reaction cycle is expected to be rather complex. It is, therefore, not surprising that such a pump includes a photoreaction, a conformational change, and a sequence of as yet not well-known different H + transfer reactions within the enzyme.

IXF-1. Variation o f the activity of the plastoquinone hydrogen pump. The linear transport of one electron is coupled in respect to the PQ-pool with the ratio of H÷/e = 1. This was shown in the single turnover experiment in subsection IXB-1. Also in the flash regime experiment in Fig. 20, bottom the ratio is H+/e = 1 at pHou t = 8 and with saturating flashes. Recently, it has been discussed that with the linear transfer of one electron, PQ may transfer two H*, as postulated by Mitchell in the Q-cycle model [182[ and its varia- tions discussed in Ref. 192. In this case the ratio would be W/e = 2. This might be possible if an extra electron, ee×, is taken up and released by PQ in addition to the electron supplied to PQ via the linear electron flow. This might be realized if the rate of eex (bound, e.g., to cytochrome b's) is shuttled in phase with the rate of the linear electron transfer, thereby transferring simultaneously a second H + via PQ. Indeed, under continuous light excitation (but not under single turnover conditions) a variation of the H÷/e ratio between 1 and 2 has been found depending indeed on an appropriate electron transfer rate and H + concentration [179,198a]. In this case under optimal conditions the PQ- pool might act maximally even as a double pump.

For a thorough discussion on H ÷ translocation in chloroplasts see Refs. 192 and 193.

X. Vectorial pathway of e, H ÷ and H in the zigzag scheme

The above results and their interrelationships, derived so far from pulse spectroscopic studies, lead (a) to a zigzag scheme with vectorial pathways of electrons, protons and hydrogen atoms (see Fig. l 3) and (b) to a preliminary picture of the molecular organization of the essential building blocks of the molecular machinery with the membrane (see Fig. 14, p. 390). The sequence of the reaction patterns has been evaluated as follows.

405

1. Absorption of light in the antennae pigments. 2. Excitation of photoactive Chl-a I

and Chl-ail via energy migration in the antennae pigments which channels the quanta towards the two reaction centers. 3. Vectorial photoejection of electrons from Chl-a~ and Chl,ali located at the inside of the membrane towards the electron acceptors located at the outside. Simultaneously a localized field and electric potential difference A~bto c is generated. At this stage the energization of the membrane takes place. 4. Delocaliza- tion of the field and Aq~, respectively, through ion redistribution and energization of the aqueous bulk phases. 5. External H ÷ uptake (or OH- release) at the electron acceptors and internal H ÷ release at the electron donors through protolytic reactions. 6. Liberations of H ÷ and OH- from tile membrane through replacement by ions of the outer and inner water phases. This is the stage of the generation of a pH gradient, ApH, and the energization of the aqueous bulk phases by A~b and ApH. 7. Subjection of the protons to the potential span of A~b and ApH and discharging of the energized state through efflux of protons. 8. The energy released with this H ÷ flux via the ATP synthetase is coupled with the formation of ATP. This is outlined in the following section (XI). 9. In toto the H ÷ movement corresponds to a H ÷ circulation. This cycle is complete if a H (H ÷ + e-) is translocated from the outside to the inside. The PQ pool probably operates as such a pump for hydrogen (see subsection IXF).

XI. Phosphorylation with the light-induced electric field and proton gradient

According to the hypothesis of Mitchell [172] the electrochemical proton potential can be used for ATP synthesis from ADP + Pi. This should be realized by the potential- driven proton efflux via a special membrane-bound enzyme, the ATP synthetase (see Figs. 13 and 14). In contrast, other hypotheses of phosphorylation assume energy transport between the electron chain and the ATPase either via unknown chemical intermediates [199] or via protein-protein interactions [200-202].

The electrochromic method developed since 1967 turned out to be a very useful instrument to explore experimentally the mechanism of phosphorylation and to discriminate between the validity of the different theoretical concepts. Up to this point phosphorylation was not included in the reactions described above because of omission of the necessary reagents ADP + P. With respect to the field decay, we have discussed so far: (a) undifferentiated pathways for H ÷ and other ions across the membrane (see subsection VIlL) and (~) artificial pathways set up by ionophores for special ions (see subsection VIIB). According to the Mitchell hypothesis there should exist during phosphorylation (7) a pathway for H ÷ via the ATPase complex (see Figs. 8, 13 and 14).

With respect to clarification of the mechanism of phosphorylation, the methodical advance since 1966 has been as follows.

1. The starting point of the development was the detection of an 'intermediate' of phosphorylation which could be measured by absorption changes at 515 nm. This was concluded from the parallelism between the amount of ATP formation and the extent of the absorption changes at 515 nm when both are varied strongly by chemical interven- tions. Under these conditions the electron transfer is, however, not changed [203,204]. The changes at 515 nm are those which have been recognized as field-indicating (see Section VII).

2. Next, the behavior of the field-indicating absorption change was analysed in the presence of ADP + P. The decay is strongly accelerated, obviously due to H + efflux via the ATPase pathway.

406

3. Tile responsibility of the ATPase pathway for the acceleration has been checked by removal and reconstitution of the F1 coupling factor of the ATPase in the presence of ADP + P.

4. By setting up competitive pathways for the field decay and for the ApH decay, the dependence of phosphorylation on A~ and ApH was analyzed under single turnover conditions.

5. With the examination of the competition between the basal and phosphorylating H ÷ effiux as a function of A¢ and ApH, attempts have been made to explain apparent threshold effects and divergences of the H+/ATP ratio, and Hp/ATP = n has been evaluated.

6. With the evaluated extent of Aq~, 2xpH and n, the proton energy was estimated and compared with the energy requirement for ATP formation.

7. As a consequence of these results, attempts were made to carry out phosphorylation with an external electric field in the dark. This method turned out to be very useful for gaining further information on conformational change, turnover time and gating of the ATPase, etc.

XIA. Acceleration of the field decay during phosphorylation Under conditions in which the potential difference decreases slowly in seconds due to

low basal permeability of the membrane, field-indicating absorption changes at 515 nm on chloroplasts are observed as shown in Fig. 21, left. Under phosphorylating conditions the decay time, r, decreases 2- to 5-fold [205,206]. The acceleration is reversed by addition of the inhibitor phloricin [205] or antibodies against the ATPase [209a]. Also in chromatophores of bacteria [125] and in whole chlorella cells [208] an acceleration was demonstrated (see Fig. 21, center and right). In algae a wild type was compared with a non-phosphorylating mutant.

At continuous illumination the effect of the field indicating absorption change is greatly decreased under phosphorylating conditions in chloroplasts [211a] and bacterial chromatophores [211b]. Both effects probably reflect the same phenomena as reported above, i.e. a decrease of the decay time of the field under conditions of phosphorylation.

The extra ion flux indicated by the acceleration of the field decay under phosphorylating conditions is obviously generated by the opening of the ATPase for H +. This is proved in the following subsection XIB.

c

a

-~o

chloroplosts chromatophores chtoret ta

+ ADP

I I 0 0.25s

O.F 0 0.Ss

time

n o n - phosphor yloting

0.5 1.0 s

Fig. 21. Acceleration (stimulation) of the field decay during phosphorylation on chloroplasts, chromatophores and Chlorella (see subsection XIA) [205,206,l 25,208].

407

X1B. Retardation of the field decay with removal of the F1ATPase If the stimulated field decay and the extra proton flux, respectively, are due to the

extra conductance of an opened ATPase for H +, the stimulation should be inhibited when the coupling factor, FI, of the ATPase (which enables the synthesis) is removed. A basic requirement for such a proof is that with the removal of F~ the basal field decay be not affected. This is the case for chromatophores of bacteria [125], in contrast to chloroplasts [209]. Therefore, chromatophores are a proper object with which to check the statement. Resolved chromatophores show no stimulated field decay when ADP + P is added. The capacity for stimulation by ADP + P was, however, completely restored after incubation with Fv This is in parallel with the total recovery of the phosphorylating activity [ 125] (see Fig. 22). This indicates additionally that the F1 reconstitution results in the native configuration.

XIC. Uncoupling of phosphorylation by a competitive pathway for the field decay The results in subsections IXA and 1XB imply that phosphorylation should be

decreased if the H ÷ flux via the ATPase pathway is in competition with other ions (e.g. K*) channeled through a bypass set up by ionophores. In this case, ApH remains unchanged for driving protons via the ATPase pathway but A~ does not. Indeed, parallel to the acceleration of the field decay induced with the ionophore Valinomycin [206] or Gramicidin D [ 123] the synthesis of ATP is decreased. Gramicidin D is much more effective than Valinomycin.

The effect of Gramicidin D is shown in Fig. 23, top. A parallelism between the acceleration of the field decay and the decrease of ATP formation is observed [ 123]. This is also the case with Valinomycin in chromatophores of photosynthetic bacteria (see Fig. 23, bottom) [122].

In steady-state light, Valinomycin does not inhibit phosphorylation [206,210]. This is expected because under this condition the fluxes of counter-ions are equilibrated across the membrane, i.e., the fluxes of ions other than H ÷ are independent of the membrane permeability.

- f 0 'a') ":- o ~ 100 Control

_o.~ / 0 0.5 1.0

Control " ~ I00 • . / > ,

g

"~ 50

0

115 0 I 05

Coupl ing factor extract added ,I~

f r o - -

I I 1.0 I 5

Fig. 22. Acceleration (stimulation) of the field decay and the rate of phosphorylat ion on chromatophores of bacteria after incubation with different amounts of the coupling factor F 1 (see subsection XIB) [ 125 ]. * (mg protein/mg bacteriochlorophyll).

408

XID. Functional unit of phosphorylation An impor tan t da tum which can be obta ined from the results in Fig. 23, top, is the

following. The m i n i m u m of Gramicidin D which acts as a bypass for the field decay via the ATPase acts also as a deactivator of ATP format ion. This number is one Gramicidin D pore per 1 - l0 s chlorophylls and per thylakoid, respectively, (One pore consists of two

Gramicidin D molecules.) Therefore, the funct ional uni t of the electric events is the same as for phosphoryla t ion,

i.e., one thylakoid or a larger uni t , e.g., several thylakoids with in terconnect ion . Because the un i t of the electric events is also that of the protolyt ic event (see subsect ion VIII) , it

is:

Aq5 uni t = ApH uni t = ATP uni t = 1 vesicle. (37)

200msl

0

0.1 0.5 I 5 • II , ,

=-....~... o2 ~ ~ 02

u

• ~,

12 a_

II A ~ 0 0 10 -9 10-8M

0.1 0.5 1 5 , II , , , t""h . ~ ~ . °z

0 10 "9 10.'8M

concentration of Gromicidin (GmcD)

or

GmcD thylakoid

10(3

tJ

0

E

o

- q

Chromotophores

=ql I J

10-9 I0-8

100

~. so O _

10-7M 0

_~0,1 Vm~ ' 47-103 BCh[

"~ Gr

Vrnc

chromatophore

0.5 1 5 10 J w w

AT

I A

10-9 10-8 ,=

10-7M concentration of VaUnomycin (Vmc)

Fig. 23. Decay time of the field (ApH = constant) and the yield of ATP formation as a function of the concentration of added ionophores. Top. Thylakoids of chloroplasts with Gramicidin D I123]. Bottom. Chromatophores of bacteria with Valinomycin (see subsection XIC and D) [ 122].

409

When instead of the thylakoids, vesicles with a volume of quite different order are regarded, e.g. the chromatophores of photosynthetic bacteria which are 100-fold smaller, again the minimum number which acts as a bypass for the field decay acts also as a deactivator of phosphorylation. 20% are deactivated by about one Valinomycin per approx. 5 • 103 bacteriochlorophylls. The latter are spread over an area which is the size of one chromatophore, see Fig. 23, bottom [122]. The result in both cases indicates that a closed vesicle in general is the functional unit for the electric, protolytic and phosphorylating events.

In the case of thylakoids, the electron transfer was measured simultaneously with the field decay and ATP formation through 02 evolution. In the range ofGramicidin D concentration where the drastic collapse of the field decay as well as of ATP formation takes place, no influence on the transport of electrons is observed (see Fig. 23, top). This supports the evidence that a whole closed membrane is the 'site of operation' of the electric events and thereby also of phosphorylation, but is not the domain of an electron transport chain as supposed in other hypotheses.

XIE. Uncoupling of phosphorylation by a competitive pathway for the ApH decay Phosphorylation should be decreased if the H + flux be channeled via an artificial path-

way.instead of via the ATPase pathway. Nigericin has been shown to catalyze a neutral exchange of K*/I-I + across the membrane. In the presence of this substance the decay of ApH is accelerated. In this case Aq~, but not ApH, should remain unchanged for driving protons via the ATPase pathway. Unfortunately, this is not the case in chloroplast of spinach. However, in chromatophores of bacteria in the presence of the nigericin-type ionophore, dianemycin, the decay time of k¢ is unchanged (see Fig. 24, top). On the other hand, dianemycin accelerates the decay of ApH (Fig. 24, center). Parallel to the acceleration of this decay, the synthesis of ATP is decreased [125] (Fig. 24, bottom). When at high concentration all protons are channeled through the bypass, synthesis of ATP stops.

Decay tiptoe of the 500 f field indicating 3o0 ~- absorption change [ (% control) tO0~

10Q

Decoy time of the pH indicating 50 absorption change ('/° control)

0

& 100 Stimulation of the fast phase of the field indicating absorption change 5o ['/. control)

o--// . . . . .

-+-,. ,

~-fl-~ ~ ,= ATP

' 11,o'2_ - ' ~ 0 100 Dianemycm

102rT

Fig. 24. Decay time of the H ÷ concentration (Aq~ = constant) compared with the yield of ATP formation (measured by the field stimulation) as a function of Dianemycin (see subsection XIE) [125]. (Subject: chromatophores of bacteria).

410

)(IF. Competition between the basal and phosphorylating pathways of H + XIF-1. Dependence on A¢ and ~ H . In continuous light it is not possible to vary

independently the values of A¢ and ApH through the change of the intensity• If, however, the steady state (ss) is established with periodic light flash of different frequency and intensity the averaged magnitudes of Aq~ and ApH can be varied independently. With this technique it is possible to examine the basal and phosphorylating proton efflux as a function of A4~ and ApH [211].

At A@ = const, the proton flux, H~, via the basal pathway depends linearly on the ratio of the H ÷ concentration (Fig. 25B), i.e., the exponent is one:

~ I ~ + + I " + + + "f ( 3 8 ) (Hin /Uot l t ) H p ~ ( H i n / H o u t )

• ÷

The proton flux via the ATPase pathway, Hp, depends non-linearly on this ratio (7 = 2.3-2.6) (Fig• 25A). Similar exponents, 7 = 1.0 and 2.0, respectively, for the two fluxes have been obtained first with a different method, but supposing A~ = 0 [212]. In Ref. 213 7 = 3 was found but identified directly with H~/ATP.

• ÷

The total flux ILl ÷ = I~I~ + Hp in dependence of ApH is depicted in Fig. 25C. At high electrochemical potential nearly all protons are channeled through the ATPase pathway

• • +

(ILl ÷ ~ I~I~) whereas at low potential the basal flux outruns the phosphorylatlng one (H Hb.

This result explains phenomena such as (a) the unusual divergence of the stoichiometrics of H+/ATP reported in the literature, (b) the so-called threshold effects and (c) the interchangeability Aq5 and ApH (see below).

XIF-2. Stoichiometrics of H+/ATP. The H+/ATP ratio follows from Fig. 25 and corresponding curves at Aq5 = 75 and 125 mV by dividing the total flux ILl + with the rate of ATP

- + •

formation ATP, i.e., H+/ATP = (I~I~ + Hb)/ATP. The result versus ApH is shown in Fig. 26, left. Fig. 26, right shows a plot of H+/ATP vs. A¢.

The degree of coupling between proton flux and ATP synthesis, i.e. H+/ATP, depends strongly on Aq5 and ApH. The effect is explained by the results of Fig• 25C, i.e., by the

i i

.g_ =

I1. 0.3

e~

A , B

0-3 Mol ATP 14ol Chb s /

a\ PHou ~ 7,8,9 o /

A@~ = 50mY / °c, 13 o/

~ o /- D

o

/°

i i 20 2.5

ApH

S o ~

3 0 ; 1 0 " 3

1 0 P H ° u t 8

• ' ~ A¢~=S0mV

c 3 o

~ o

i I 2.0 2.5

ApH

i 3.0 3.0

c

20110-3 Mot w ,', r Mol Chl.s / /

i I

10 /// / ,,;

5 / / / / .'-~/,'/// l /

I /// z's

ApH

Fig. 25. Phosphorylating proton flux, tt~, (A) and basal, HE, (B) as a function of Apt! (Aq~ = constant = 50 mV) (see subsection XIF-1) [211].

411

~-~ zo

3O or

2O

H* ATP

A¢I= 50mV ~j

2 ~

curve calculated

\ A*3:125mV ~ ' ' i ' l ~ ~

1.5 2.0 2.5

5

1.5

3,0

A p H

,~TP

-#

"pH 2.0 \=.

ATP

-.\\o

- - A ~ i i S0 75 100 125rnV

e lect r ica l potential difference A ¢

20

15

10

5

~5 - - 0

Fig. 26. H*]ATP as a function of Z~pH and A~, respectively. The dependencies are a consequence of the results of Fig. 25 (see subsection XIF-2) [211 ].

dependence of the basal and phosphorylating flux on different exponential powers. The "4- "4- true value, i.e., Hp/ATP is obtained at high A(b and ApH where H b < < Hp. In this range

we obtained [211]:

n ~ H ; / A T P ~ 2.5 (39)

i.e. five protons have to drop through the potential span via the ATPase pathway to generate two ATP molecules.

The number of ions which pass across the ATPase coupled pathway can be obtained also from the difference of the field decay with and without phosphorylation (see Fig. 21). The conclusion was that the synthesis of one ATP molecule is coupled to the field-driven flux of about three H ÷, i.e. n = Hp/ATP ~ 3 [206]. With other methods, values between 2 - 4 have been measured. For a thorough discussion see Refs. 192 and 215.

XIIF-3. Apparent thresholds and interehangeability of Aq) and ApH. In Fig. 27 (taken from the same type of measurements as that used for Fig. 25) the number of ATP molecules per flash and electron chain is plotted as a function of ApH and A~, respectively. At low potentials the yield increases non-linearly, suggesting a 'threshold'. The threshold disappears at high ApH and Aq~, respectively. The threshold is an apparent one. It is simply explained, again by the dependency of I:t~ and I:Ip on different exponential powers. At low ApH (or Aq~) it is H~ ~ 1~ and the ATP yield is zero (see Fig. 25). The range of 'inversion' (I:I~ ~ t:ffp) must be responsible for the threshold. A first report on a threshold of Aq~ was originally interpreted as an electric triggering of the ATPase [206] which is different from the explanation in this paragraph.

1. The results in Figs. 26 and 27 demonstrate at least qualitatively that with respect to phosphorylation £xq~ and ApH are interchangeable. 2. This was also demonstrated by superimposing a diffusion potential onto a pH gradient which by itself was insufficient to phosphorylate [216]. 3. This is furthermore shown by the evidence that phosphorylation is possible with ,54 only or ApH only (see Section XII).

XIG. Proton energy and energy requirement for A TP forrnation According to the results in this report and to those of several other laboratories [215]

412

1.5

1.0

= 0.5

I1.

• ',e3 = l~W-JmV

PHout8 /

A /

&e =50mV /

_._I..1

1.5 2,0 2.5 3.0 ApH

t5J

i

tOF

esl-

oe

translocated H + per f lash and electron chain

1 2 3 t, 5 , ! ¢ i

pU .a /XpH =2.77/ / ApH=2.6 "OUlU o / / &pH=2t.

/ / / / / / /

/ / / / 50 16o 1543nr~/

electrical potential difference &@

Fig. 27. Thresholds and yield of ATP formation as a function of ApH and A0, respectively. The dependencies are a consequence of the results of Fig. 25 (see subsection XIF-3) [2111.

the energized state for phosphorylation is the electrochemical potential of H +. In this case we have to consider, according to Mitchell [ 172], the following relations.

In saturating steady state light it holds

n " SSAGH* ~ SSAGp (40)

SSAGH+ = free energy of the protons (see Eqns. 29 and 30, respectively)

SSAGp = free energy of the phosphorylation = AG O' + RT" In SS(ATP/ADP • P) (41)

From the concentrations of ADP, P and ATP in this state [217] and from the revised AG O' value [218] it is found that

SSAGp = 13.5 kcal/mol (42)

Eqn. 40 is satisfied according to Eqns. 20, 30 and 41, for instance, with the three sets of data listed in Table II. We can consider, thereby, either the values between the bulk

TABLE I1

DIFFERENT SETS OF DATA RESULTING IN >->13.5 kcal/mol ACCORDING TO THE EQNS. 29 AND 30, RESPECTIVELY

A(n = 2) B(n = 2.5) C01 = 3)

Between ss,xo = 90 mV SSApH = bulk 3.3 phases

Between SSAq~ + ssA¢ = SSApH =

boundaries 170 mV 2.0

ssAck = 32 mV SSApH = ss&0 = 1 mV SSAptt =

3.3 3.2

ss&0 + SSA¢ = SSApH = ssA<p + SSA~0 SSAp H =

92 mV 2.3 61 mV 2.2

413

phases or, on the other hand, those between the boundaries (see subsection VIIM-3). The values of B and C in Table II are in fair accordance with those which have been measured (see subsections VIIM-2,3 and XIF-2). Because these values are not, however, established beyond doubt, we can conclude at the present state only that the energetic consideration is not in contradiction with the mechanism proposed. However, regarding the findings discussed in this present section and taking into account the arguments in subsection XIIIA, evidence is advanced in favour of the chemiosmotic mechanism.

XII. Phosphorylation induced by an external electric field

In the light-induced phosphorylation the protons are displayed to an electric term (~r. AO) plus a chemical one (2.3 • RT. ApH). Jagendorf and Uribe have demonstrated in their fundamental experiment that phosphorylation is possible in the dark if an artificial pH gradient alone be set up across the thylakoid membrane [219]; for details see Ref. 215. To prove that ATP can be generated already by the precursor of the pH gradient, i.e. by the transmembrane electric field only, we have replaced the intrinsic electric generators by an external electric generator. In this way transmembrane electric potentials can be induced in the dark (see next subsection XIIA). Under these conditions a pH gradient between the internal and external aqueous bulk phase is absent but phosphorylation takes place with yields comparable with those in the light [220,221 ].

Phosphorylation by an artificial electric potential set up with a diffusion potential across the membrane gave yields of ATP which are extremely small [227a].

The external field method is valuable - also for more sophisticated problems, besides phosphorylation, which cannot be answered under conditions of light excitation; for instance:

(1) The ATP synthesis or the conformational changes in dependency on the electric potential can be resolved with this method (see below). (By excitation with light the result would be falsified through the interference with the pH gradient which is generated at higher intensities.)

(2) The action of the AYPase at ApH = 0, as a function of pH (pHout = pHin), can be measured also in this way and should give new information on the mechanism within the ATPase.

(3) Furthermore, the interference of the light-induced electric potential with the potential induced simultaneously with the external field may give information in which way the electrons are shifted vectorially across the membrane, etc.

XIIA. Principle of the external field method Electric fields can be induced at vesicles from an external source as shown schemati-

cally in Fig. 28 [220,221]. In the left-hand scheme, A, two macroscopic electrodes with a distance of 1 mm are depicted. An aqueous solution is placed between these electrodes and 200 V are applied. If we imagine within the solution a sphere of water with a diameter which is one thousand times smaller than the distance between the electrodes, the voltage across this sphere is, of course, one thousand times smaller than the total voltage, i.e. 200 inV. If we regard a sphere of lipid instead of the sphere of water, as shown in the center, B, of Fig. 28, the non-conducting lipid gives rise to a charge accumulation in the water at the outer surface of the sphere. Therefore, the voltage across the sphere is higher and can be calculated to be at the poles 300 inV. If we replace the inner space of the

414

A 200 V

B C

,I

' I ' ~ 1 0 - ~

H20

- i : Q ~

i

1

[H2- O . I

- ~ 4 " I

Fig. 28. Principle of tile external electric field method (see subsection XIlA) [220,221].

sphere of lipid by water, this corresponds to a vesicle with a non-conducting membrane. This is shown in Fig. 28C. The voltage across the whole sphere is, of course, still 300 mV but the distribution of counter ions inside splits up the voltage into two parts: 150 mV across the membrane at the left half of the vesicle and 150 mV across the right half. The generated transmembrane field has at one half of the vesicle the same direction as the light-induced field, i,e. positive inside and negative outside. In the scheme this is the left- hand half. For this reason the yield of one external voltage pulse can be compared with the yield of one light pulse if the former one is related to the double amount of chloroplast and chlorophyll, respectively. External fields have been used also for stimulation of delayed light emission on chloroplasts (227b,c).

XIlB. Yield o f ATP formation in an external field Under the described conditions ATP is generated in the dark as shown in Fig. 29,

left. The amount increases linearly with the number of external voltage pulses. In Fig. 29, right, this yield is compared with the amount of ATP generated in saturating light pulses of the same duration. Because the yield is nearly the same we conclude that phosphorylation generated by light is equivalent to phosphorylation in the dark generated by an external electric field [220]. Electron transport inhibitors like DCMU have no influence on the ATP yield induced by external voltage pulses. But inhibitors of the ATPase like phlorizin inhibit completely the ATP synthesis in the external voltage pulse. These results indicate that the presence of an electric field alone is sufficient for synthesis of ATP. The protons driven by this field may be 'taken' not from the inner aqueous phase but primarily from the inner membrane surface. Whether the postulated proton well in coupling factor of the ATPase converts the field into a local pH difference, is an open question.

XIIC. Conformational change and turnover time o f the A TPase When the membrane is energized, conformational changes take place at the chloroplast

ATPase which has been shown first be Ryrie and Jagendorf [233] (for reviews see

415

"O _=

¢-.

O3

0_ I-- <

O E (D

Mol ATP 8"}0"3 Mo[ Ch1.2

.j'" . /

0 / * _ * . . . . . . . . 5 10

number of external voltage pulses

, , f

O3

O

C

0 E o

Mol ATP 8 10-3 Mol Chl

D

/ • t

number of light pulses

Fig. 29. Yield of ATP formation as a function of the number of external voltage pulses in the dark and as a function of the number of saturating light pulses. Pulse duration in both cases 30 ms (see subsection XIIB) [220,221 ].

Ref. 222). We asked in the following for the relation between ATP synthesis and conformational changes. The latter were measured according to a method evaluated by Slater [223], Boyer [224] and Strotmann et al. [224a]: The ATPase contains tightly bound adenosine nucleotides (AMP, ADP and ATP). These are released only after energization of the membrane. It is assumed that this is caused by a conformational change of the ATPase. Thus, the amount of nucleotides released is proportional to the number of ATPases which have changed their conformation. Those ATPases are called 'active' ones.

In our experiment, excitation was induced by the external field instead of by light for the following reasons.

In the light AO and ApH depend in a different way on the light intensity and neither A~ nor ApH depends linearly on the intensity. However, the transmembrane voltage induced by an external voltage pulse is directly proportional to the external voltage. Such linearity is necessary for the analysis in question.

In Fig. 30, right, the amount of nucleotides released and ATPases which change their conformation increase non-linearly as a function of the external electric field strength and the transmembrane electric potential induced. Under the same conditions the amount of ATP generated has been measured in Fig. 30, left. About six times more ATP is generated than nucleotides released and ATPases have changed their conformation [225]. This discrepancy can be explained by the assumption that each active ATPase generates about six ATP, i.e. each active ATPase carries out six turnovers during the pulse duration of 30 ms. Then the turnover time of the ATPase should be 5 ms. Because the ATP/AdN* ratio is nearly constant at different electric potentials, it can be concluded, furthermore, that the turnover time is independent of the electric potential across the membrane (see Fig. 31, top) [2251.

XIID. Gating of the A TPase For the synthesis of one ATP molecule, nH÷ must be translocated via the ATPase (see

subsection XIF-2). It is very likely that these protons pass only those ATPases which change their conformation, thereby opening a gate for protons. In Fig. 31 the results of

416

Q)

o

c

n <

"6 C :3 0 E

potent ia l difference 0 0 ~100 I I. _ 1 10 "/~ Mol ADN 6 10 /~ MOl ATP

Mol Ch{ MOl Ch l

o U 0 100

t r a n s m e r n b r a n e electr ic ~100 ~200mV "~

. /

pulse durat ion : 30ms t - :3 0

5;0 lO00'V/cm OE

05

-200mY

i /

f ~•J, pulse dura t ion = 30ms

100 500 100~0 V/cm

ex te rna l e lectr ic f ield s t rength

Fig. 30. A m o u n t of ATP generated (left) and a m o u n t o f adenosine nucleotides released (right) as a funct ion o f the t ransmembrane electric potential difference induced by an external electric field in the dark (see subsect ion XIIC) [ 225 }.

Fig. 30 are replotted (solid lines) but extrapolated to higher potentials (broken lines). At the top the turnover time of the ATPase is shown in dependence on A~. The linear extra- polation to higher voltages is in accordance with independent results (squares) calculated from measurements of Smith and Bayer [226]. At the bottom the fraction of 'active'

~ -~/ 550 n • ,-7 C

2 3 " 6 0 • ' ~ "< t-, 0 100 200 mV

o OL I - -

u

~0.5 0

C .9

02

F-

1.0 10 Ul / :

/ ! /

/ J I t

/ /

/ /

7" J

0 ~ "- o"~- ' '~° S, ' ' -~ ~

0 100 200 rnV e lec t r ic potential d i f f e r e n c e

! ~05

I

I lo.2

¢1

O

O

O 1 3

1> --4 "13 ¢3 b~

?, o

Fig. 31. Turnover t ime of the ATPase (top) and fraction o f active ATPases (bot tom) as a funct ion of the t r ansmembrane electric potential difference induced by an external electric field in the dark (see subsect ions XIIC, D) [ 225 ].

417

ATPases, i.e. those which change their conformation, is depicted. It is probable that the extrapolated curve has to reach unity at high potentials. The square results from measurements at high values of ApH 4.4 [226]. These results show that the chloroplast ATPase can work as an electrically gated enzyme for translocation of protons. The number of 'active' enzymes, i.e. number of open channels, depends off the electric potential. The turnover rate and proton current density, respectively, is, however, independent of the potential.

An 'electrically gated ATPase' was discussed by Junge [214,214a]. In this case the increase of phosphorylation with A¢ [206] above a critical value (threshold)was explained theoretically by a decrease of the turnover time. This assumption is different from the experimental results presented in this chapter. It should be noted that the fraction of 'active' ATPases is not fixed to a special ensemble but migrates statistically between all ATPases within 3 s [225]. The averaged rate of ATP synthesis, ATP, is:

(A) ~ ~ ~ . n(a¢) -No (43) TO

To = turnover time = constant, 7?" No = active ATPases. The dependence of ATP on A¢ is due to the dependency of the active fraction 7? on A¢. In Fig. 32, left, this result is depicted in a simplified scheme. The circles represent the ATPases (total number No). The length of the arrows indicate the turnover rate of the active ones. For comparison the other possible extreme is depicted in Fig. 32, right. All ATPases, No, are active but the turnover rate, 1/r, depends on A¢.

1 (B) ATP ~ "No (44) T(A¢)

The results discussed above show that mechanism A is realized. Mechanism A has, inter

A

t I 0 0 0 0 0\0 o 0

low A¢~

high A~

B

low A~

high / ~

Fig. 32. Two mechanisms - A and B - of the cooperat ion be tween the turnover t ime and the number o f active ATPases at low and high t r ansmembrane electric potential difference, A4~. Mechanism A is realized according to subsect ion XIID [ 221,225 ].

418

alia, the following advantage. The time for gathering n protons in order to synthesize one ATP is minimal and constant and thereby independent of the light intensity. This is not realized by the mechanism B and, therefore, gives rise to problems at low intensities.

The conformational change and activation of the ATPase may be due to indirect action of the electric field in producing a local proton gradient across the ATPase. How- ever, direct interaction of the field with electric properties of the ATP also could be discussed.

XIlI. Data for discrimination between the different hypotheses on the mechanism of phosphorylation in photosynthesis

For an understanding of phosphorylation two major mechanisms have to be described. (1) The transfer of energy from the electron transport chain to the ATP synthesizing enzyme, the ATP synthetase; (2) The use of the transmitted energy through individual reactions within the ATPase which lead to the formation of free ATP.

XIIIA. Coupling between the electron transport chain and the A TPase In respect to the energy transfer, three hypotheses were presented: ( l ) the chemical

[199]; (2) the conformational [200-202]; and (3) the chemiosmotic [172]. The chemical and conformational theories assume within localized domains a direct energetic interaction between the electron transport chain and the ATPase either through an energy- rich chemical intermediate or through an energetically conformational change transduced by protein-protein interactions. Both types of transmitter are symbolized in a and fi by a squiggle (~). In order to account for the existence of the field and A~, respectively, one has, therefore, to regard in the two hypotheses the existence of A~b as acting in a sidepath. Neither scheme, a and/3, is realistic according to the following results.

e --> (~) ~ ATP I

LX¢

e -~ (~) -~ ATP

A¢ (9)

-+ A4~ --~ ATP (3')

(a) A~b in a sidepath at the level of the electron-transfer is in contradiction to the fact that the acceleration of the field decay can be realized, either mainly by a H ÷ efflux, i l l + ,

(see subection VIIL) or mainly by a K ÷ efflux, iK ÷, (see subsection VIIE-2). According to the scheme, ATP formation should be independent of the type of acceleration. But phosphorylation is inhibited under conditions of a K ÷ efflux. Even the slightest change from iH÷ towards iK+ deactivates ATP formation (one potassium channel per thylakoid (see subsection XID)).

(~) A~b in a sidepath at the level of the squiggle (4) is in contradiction to the fact the squiggle must be formed in much less than 20 ns because A~ generation has been measured to take place in less than 20 ns (see subsection VIIB). The synthesis of (~) in much less than 20 ns is very unlikely, especially the synthesis of an energy-rich chemical com- pound. Also, a conformational change transmitted by protein-protein interactions cannot take place within the nanosecond range. Furthermore, A~ is a result of photooxidation of the chlorophylls (see subsection VIIK) and not a consequence of an intermediate, (~).

419

Because phosphorylation takes place by excitation with external electric fields and because under these conditions no electron transfer from H20 to NADP ÷ takes place, the result indicates that ATP formation is possible without any electron transfer and, therefore, also possible without the postulated subsequent reactions, i.e. the formation of an intermediate ( - ) as shown in ct and/3.

(7) On the other hand, the results obtained in photosynthesis by the pulse technique presented in this report and the results obtained through several other methods are in favor of a coupling of the electron transfer and phosphorylation by the H ÷ potential, i.e. by 2x4 and &pH, respectively:

(1) The evidence for a transmembrane electric field and Axe, respectively, as well as for a &pH is a 'pro' for discussing the hypothesis at all.

The electric field is the very basic element of the mechanism. The delocalized ApH formation, induced by the charge separation, is a subsequent event but not a prerequisite for phosphorylation (see Section XII).

(2) The field decay caused by efflux of protons is accelerated during phosphorylation (see subsection XIA).

(3) The acceleration disappears with the removal of the coupling factor of the ATPase and reappears in full with its reconstitution (shown on chromatophores from bacteria, see subsection X1B).

• (4) The proton flux via the ATPase pathway is proportional to the rate of phosphorylation. (In our hands the proportional factor is n ~ 2.5, i.e. about five protons have to drop through the potential span to generate two ATP's) (see subsection XIF-2).

(5) The field decay via artificial pathways set up by ionophores competitively inhibits ATP generation (see subsection XIC).

(6) Already a single potassium channel per thylakoid acts as a bypass for the intrinsic field decay as well as a deactivator of ATP formation, i.e. the functional unit is one vesicle and not the domain of an electron transfer chain (see subsection XID).

(7) Phosphorylation is possible in full in an artificial H ÷ potential set up by an external electric field (see subsection XIIB).

(8) The &pH decay is accelerated during ph0sphorylation [178]. (9) The &pH decay via artificial pathways set up by uncouplers competitively inhibits

ATP generation (see subsection XIE). (10) Phosphorylation is possible in an artificial H ÷ potential set up by an external pH

gradient [219]. (11) In reconstituted systems phosphorylation takes place if a pit gradient is set up by

the proton pumping bacteriorhodopsin [230]. (12) Conversely the supply of ATP generates a H ÷ potential via the reversal of the

mechanism of ATP synthesis, i.e. via ATP hydrolysis [231 ]. (13) The free energy of the protons estimated from the electric and osmotic values of

the bulk phase or from the corresponding values at the boundaries (see Table II in subsection XIG) satisfies with n = 2.5-3 the free energy requirement (13.5 kcal) for ATP formation. However, precise measurements are required to improve the energy balance argument beyond doubt (see subsection XIG).

Williams [227,228] suggested a local proton potential between the membrane and the outer water phase, i.e. his concept is restricted to two phases. Furthermore, these protons are regarded as anhydrous, operating in a local domain between the electron transport chain and the ATPase. This mechanism can be criticized in that the domain of an electron transport chain is not the functional unit of phosphorylation but a whole vesicle (see sub-

420

section XID). Also, experiments of Oft et al. [229] cannot be used as an argument for the suggestion of Williams. Ort et al. have concluded that a time lag in the rise of ATP formation is shorter than the rise of Aq5 and ApH, respectively. Howevei, A~b has not been measured within this intermediate state. It was assumed only from the addition of Valinomycin + KC1 that A4~ might be zero within this pre-steady state. But this is correct only under single turnover conditions where the field is quenched by the charge translocation via Valinomycin. However, in the steady state Valinomycin has no influence on the field and on phosphorylation at all (see subsection X1C) and, probably, only very little in the pre-steady state.

XIIIB. Mechanistics within the A TPase Although as yet the pulse methods are not related to results on this topic, for com-

pleteness, the main results should be mentioned in brief. The protons driven by A~ and ApH through the ATPase subsequently can induce

several individual reactions before ATP is synthesized and released into the solution. The ATPase has been shown by Racker et al. [232] to be composed of five subunits.

Ryrie and Jagendorf [233] demonstrated that energization of the membrane is accompanied by conformational changes of the ATPase which permit interaction with ligands. Boyer and Slater [234,235] suggest that the free energy of the proton potential is used for an energy-requiring conformational change which releases bound ATP at one site and bind ADP + P at another site in a mode favoring bound ATP formation. Mitchell [236] proposed direct utilization of the proton energy for combining P with ADP at the active site of the ATPase. Other intermediate reactions may be of importance on the way to the synthesis of ATP. However, at the present time no prefererence for any one of the discussed mechanisms is given. An answer is in the domain of future work. In this respect the exact stoichiometry of Hp/ATP, the structure-function relationship, the characterization of binding sites, the interpretation of the conformational changes - energetic or catalytic - etc., are important questions for further efforts. For a more comprehensive discussion on this subject see Refs. 237,238.

X1V, Conclusions

The framework of the basic concept of the molecular mechanism of photosynthesis obtained from the results of the pulse methods and all the other techniques now applied to this field of research is evident. The site of the operation is a 'two-dimensional-type reaction vessel', the membrane. Thereby the reactants are concentrated and exposed to fast turnover which can compete even with large influxes of quanta. The compartmenta- tion of the membrane into closed vesicles, i.e. into an 'interior-exterior' system enables asymmetrical arrangement of the reactants. This allows vectorial reactions and the establishment of a storage device. Such constructions are used also in vision, respiration, nerves, etc.

The extraordinary specificity of the reactions within such a site of operation, especially in photosynthesis, is characterized by: (1) the regulation of the utilization of quanta via two types of energy migration; (2) the cooperation of two photoreaction centers, Chl-a I and Chl-aH, for the transfer of one electron from H20 to NADP+; (3) vectorial electron ejection out of the excited state of Chl-a I and Ch[-aii and electric field generation, respectively; (4) protolytic reactions and pumping of H; (5) the proton potential as energy transmitter; (6) the use of H20 as electron source; and (7) ATP formation in the ATP synthesizing enzyme.

421

Each of these events per se is an unique ' invention' by nature, and is in part applied in molecular machineries o f other life cycles. To support the identification of these events and to help to clarify the mode of the coupling of these processes was the purpose of the efforts described in this report.

Although progress has been made, it is evident that only a framework of the principle has been reached. Important questions are open. (1) The mechanistic details of the vectorial electron ejection across the membrane and the stabilization of the primary charge separation are not understood. (2) The molecular reactions which lead to the cleavage of H20 are still a mystery. (3) The detailed architecture o f the membrane and the role of the motion of reactants within the lipid layers are not known. (4) The function of the cytochromes is not established. (5) The mechanism by which ATP is synthesized within the ATP synthetase must also be left to future work.

Research in this field is in a state of rapid advance. It is hoped that the usefulness of the pulse methods and the different modes of their application outlined in this essay can help to answer some of these questions also in forthcoming studies on this unique molecular machinery which is the most important energy converter on earth and thereby the very basis of life.

Acknowledgements

I should like to express my gratitude to all members of the Max-Volmer-lnstitut who contr ibuted to the results described in this review.

I am most grateful to Dr. P. Gr~ber, Prof. W. Junge, Dr. R. Reich, Dr. G. Renger, Professor B. Rumberg and Dipl.-Phys. E. Schlodder for studying the manuscript and for helpful discussion. Many thanks are due to Miss D. DiFiore and Mrs. A Schulze for tech- nical assistance. To Mrs. Ch. Proll I am greatly indebted for writing and correcting the manuscript through the different stages of planning.

Our studies, discussed in this essay, have been supported by grants of the Deutsche Forschungsgemeinschaft, by the Commission of the European Communities, by ERP Sonderverm6gen and by the Stiftung Volkswagenwerk.

References

1 Witt, H.T. (1967) in Fast Reactions and Primary Processes in Chemical Kinetics (Nobel Sympo- sium V) (Cleasson, S., ed.), pp. 261-316, Almqvist and Wiksell, Stockholm; lnterscience Publ., New York

2 Witt, H.T. (1971) Q. Rev. Biophys. 4, 365-477 3 Witt, H.T. (1975) in Bioenergetics of Photosynthesis (Govindjee, ed.), pp. 493 554, Academic

Press, New York 4 Govindjee fed.) (1975) Bioenergetics of Photosynthesis - Cell Biology, pp. 1--698, Academic

Press, New York 5 Trebst, A. and Avron, M. (eds.) (1977) Photosynthesis I - Photosynthetic Electron Transport

and Photophosphorylation, pp. 1-730, Springer-Verlag, Berlin 6 Barber, J. (ed.) (1977) Primary Processes of Photosynthesis - Topics in Pliotos~ nthesis, Vol. 2,

pp. 1-516, Elsevier, Amsterdam 6a Norrish, R.G. and Porter, G. (1949) Nature 162,658 -661 7 Witt, H.T. (1955) Naturwiss. 42, 72-73 8 Witt, H.T. (1967) in Fast Reactions and Primary Processes in Chemical Kinetics (Nobel Sympo-

sium V) (Cleasson, S., ed.), pp. 81-97, Almqvist and Wiksell, Stockholm; Interscience Publ., New York

9 Rdppel, H. and Witt, H.T. (1970) Methods Enzymol. 16,316-380

422

10 Kaufmann, K.J,, Dutton, P.L., Netzel, T.L., Leigh, J.S. and Rentzipis, P.M. (1975) Science 188, 1301-1304

11 Emerson, R. and Arnold, W. (1932) J. Gen. Physiol. 16,191 12 Gaffron, H. and Wohl, K. (1936) Naturwiss. 24, 81 13 Knox, R.S. (1977) in Bioenergctics of Photosynthesis (Govindjee, ed.), pp. 55-97 Academic

Press, New York 13a Knox, P,.S. (1977) in Primary Processes of Photosynthesis Topics in l'laotosynthcsis, Vol. 2

(Barber, J., ed.), pp. 56 -93 , Elsevier/North-ttolland, Amsterdam 14 Williams, W.P. (1977) in Primary Processes of Photosynthesis - Topics in Photosynthesis, Vol. 2

(Barber, J., ed.), pp. 99 147 EAsevier/North-Holland, Amsterdam 15 Moraw, R. and Witt, H.T. (1961) Z. Phys. Chem., Neue Folge 29, 1 - d 2 15a Zieger, G., MLiller, A. and Witt, H.T. (1961) Z. Phys. Chem., Neue I.'olge 29, 13 24 16 Wolff, Ch. and Witt, H.T. (1969) Z. Naturforsch. 24b, 1031-1037 17 Witt, K. and Wolff, Ch. (1970) Z. Naturforsch. 25b, 387-388 18 Buchwald, H-E. and Wolff, Ch. (1971) Z. Naturforsch. 26b, 51 -53 19 Mathis, P. and Galmiche, C.R. (1967) Acad. Sc. Paris 264, 1903-1906 20 Wolff, Ch. (1972) Biophys. Congr. Abstr., Moscow, 351 352 21 Emerson, R. (1958) Annu. Rev. Plant Physiol. 9, 1 22 Emerson, R. and Rabinowitch, E. (1960) Plant Physiol. 35 ,411-485 23 Hill, R. and Bendall, D.S. (1960) Nature 186,136 24 Kok, B. and Hoch, G. (1961) in Light and Life (McElroy, W.D. and Glass, B., eds.), pp. 397-

416, John Hopkins Press, Baltimore 25 Duysens, L.N.M., Amesz, J. and Kamp, B.M. (1961) Nature 190 ,510-511 26 Witt, H.T., Mtiller, A. and Rumberg, B. (1961) Nature 191 ,194-195 27 Rumberg, B., Smidt-Mende, P., Weikard, J. and Witt, H.T. (1963) in Photosynthetic Mechanism

of Green Plants, Acad. Sci. Res. Counc. Washington, Publ. 1145, pp. 18-34 28 Duysens, L.N.M. and Sweers, H.E. (1963) in Studies of Microalgae and Photosynthetic Bacteria

(Jap. Soc. Plant Physiol., ed.), pp. 353-372 , University of Tokyo Press, Tokyo 29 Kok, B. (1957) Acta Bot. Neerl. 6 , 3 1 6 - 3 3 6 30 Rumberg, B. and Witt, H.T. (1964) Z. Naturoforsch. 19b, 693- 707 31 D6ring, G., Bailey, J.L., Kreutz, W. and Witt, H.T. (1968) Naturwiss. 55,219--224 32 Seifert, K. and Witt, H.T. (1969) in Progress in Photosynthesis Research, Vol. !I (Metzner, H.,

ed.), pp. 750-756 , Int. Union Biol. Sci., Ttibingen 33 Ke, B. (1973) Biochim. Biohpys. Acta 301, 1 -33 34 Rumberg, B. (1977) in Encyclopoedia of Plant Physiology, New Series, Vol. 5, Photosynthesis I

(Trebst, A. and Avron, M., eds.), pp. 405-415 , Springer-Verlag, Berlin 34a Junge, W. (1977) Annu. Rev. Plant Physiol. 2 8 , 5 0 3 - 5 3 6 34b Junge, W. (1977) in Encyclopoedia of Plant Physiology, New Series, Vol. 5, Photosynthesis

(Trebst, A. and Avron, M., eds.), pp. 59 93, Springer-Vcrlag, Berlin 35 Norris, J.R., Uphaus, R.A., Crespi, H.L., Katz, J.J. (1971) Proc. Natl. Acad. Sci. U.S. 6 8 , 6 2 5 -

628 36 Philipson, K., Sato, V. and Sauer, K. (1972) Biochemistry 11 ,4591-4595 37 Seely, G.R. (1977) in Primary Processes of Photosynthesis - Topics in Photosynthesis, Vol. 2

(Barber, J , ed.), pp. 55 -97 , Elsevier/North-Holland, Amsterdam 38 Griffiths, M., Sistrom, W.R. and Cohen-Ba zire, G. (1955) Nature 176, 1211 - 1214 39 Miiller, A. and Witt, H.T. (1961) Nature 189 ,944-945 40 Kok, B. (196t) Biochirn. Biophys. Acta 48,527 532 41 Rumberg, B. (1964) Z. Naturforsch. 19b, 707 716 42 Beinert, H. and Kok, B, (1963) in Photosynthetic Mechanism of Green Plants (Jagendorf, A.T.,

ed.), pp. 131 - 137, Natl. Aead. Sci. Res. Coun., Washington, D.C., Publ. 1145 43 Warden, J.T. and Bolton, J.T. (1972) J. Am. Chem. Soc, 94, 4352-4353 44 Evans, M.C.W. (1977) in Primary Processes of Photosynthesis Topics in Photosynthesis, Vol. 2

(Barber, J., cd.), pp. 465 492, Elsevier/North-Holland, Amsterdam 45 Haehnel, W., D6ring, G. and Witt, H.T. (1971) Z, Naturforsch. 26b, 1171-1174 46 Haehnel, W. (1973) Biochim. Biophys. Acta 305,618 631 47 Stiehl, H.H. and Witt, H.T. (1969) Z. Naturforsch. 24b, 1588 1598

423

48 Renger, G. and Wolff, Ch. (1977) Biochim. Biophys. Acta 460, 4 7 - 5 8 49 Monger, T.G., Cogdell, R.J. and Parson, W.W. (1976) Biochim. Biophys. Acta 449, 136-153 50 D6ring, G., Stiehl, H.H. and Witt, H.T, (1967) Z. Naturforsch. 22b, 639-644 51 D6ring, G., Rengcr, G., Vater, J. and Witt, H.T. (1969) Z. Naturforsch. 24b, 1139-1143 52 D6ring, G. and Witt, H.T. (1972) in Proc. 2nd Int. Congr. Photosynthesis Res., Stresa 1971

(Forti, G., Avron, M. and Me!andri , A., eds.), pp. 39 45, Junk, Den Haag 52a Buchwald, 1t.-I.;. (1972) Z. Anal. Chem. 261 ,286 - 290 53 ttavemann, J. and Mathis, P. (1976) Biochim. Biophys. Acta 440, 346 355 53a Bougcs-Boquet, B. (1977) Biochim. Biophys. Acta 462,362 370 54 Ke, B., Satu, S., Shaw, E. and Beinert, H. (1974) Biochim. Biophys. Acta 347, 36 48 55 Knaff, D.B. (1977) Photochem. Photobiol. 26,327 340 56 Wolff, Ch., G1//ser, M. and Witt, H.T. (1975) Proc. 3rd. Int. Congr. Photosynthesis, Rehovot, Israel

(Avron, M., ed.), pp. 295-305, Elsevier, Amsterdam 57 Malkin, R. and Bacrden, A.J. (1975) Biochim. Biophys. Acta 396,250 259 58 Van Gorkom, H.J., Pulles, M.P.J. and Wcssels, J.S.C. (1975) Biochim. Biophys. Acta 4 0 8 , 3 3 1 -

339 59 Renger, G., Eckert, H.-J. and Buchwald, H.-E. (1978) EEBS Lett. 90, 10-14 60 Gl/iser, M., Wolff, Ch., Buchwald, H.-E. and Witt, H.T. (1974) FEBS Lett. 42, 81-85 61 Malkin, S. (1977) in Primary Processes of Photosynthesis - Topics in Photosynthesis, Vol. 2

(Barber, J., ed.), pp. 349-431, Elsevier/North-Holland, Amstcrdam 62 Joliot, P. and Kok, B. (1975) in Bioencrgetics of Photosynthesis (Govindjee, ed.), pp. 388-412,

Academic Press, New York 63 Renger, G., Gl~iser, M. and Buchwald, H.-E, (1977) Biochim. Biophys. Acta 461 ,392 -402 64 Amesz, J. and Duysens, L.N.M. (1977) in Primary Processes of Photosynthesis - Topics in Photo-

synthesis, Vol. 2 (Barber, ed.), pp. 149-185, Elsevier/North-Holland, Amsterdam 65 Witt, H.T. (1976) in Excited States of Biological Molecules (J.B. Birks, ed.), pp. 245 261, Wiley

Interscience, London 65a Demeter, S. and Kc, B. (1977) Biochim. Biophys. Acta 4 6 2 , 7 7 0 - 7 7 4 65b Kuhn, H. (1972) Chem. Phys. Lipids 8 , 4 0 1 - 4 0 4 65c Hopfield, J.J. (1974) Proc. Natl. Acad. Sci. U.S. 71, 3640-3644 66 Gl/iser, M., Wolff, Ch. and Rengcr, G. (1976) Z. Naturforsch. 31c, 712-721 67 Forti, G. (1977)in Photosynthesis I - Photosynthetic Electron Transport and Photophosphoryla-

tion (Trebst, A. and Avron, M., eds.), pp. 222-226, Springer-Verlag, Berlin 68 Hall, D.O. and Rao, K.K. (1977) in Photosynthesis ! Photosynthetic Electron Transport and

Photophosphorylation (Trebst, A. and Avron, M., cds.), pp. 206 216, Springer-Verlag, Berlin 69 Stiehl, H.H. and Witt, H.T. (1968) Z. Naturforsch. 23b, 220-224 70 Witt, K. (1973) FEBS Lett. 38, 116-118 71 Bensasson, R. and Land, E.I. (1973) Biochim. Biophys. Acta 325 ,175-181 71a Renger, G. and Wolff, Ch. (1976) Biochim. Biophys. Acta 4 2 3 , 6 1 0 - 6 1 4 71b Dismukcs, Ch.G., McGuire, A., Blankenship, R. and Sauer, K. (1978) Biophys. J. 21,239 256 71c Diner, B.A. (1977) in Proc. 4th Int. Congr. Photosynthesis, Reading, U.K. (Hall, D.O., Coombs,

J. and Goodwin, T.W., eds.), pp. 359 372, Biochemical Society, London 71d Van Gorkum (1974) Biochim. Biophys. Acta 347 ,439 -442 72 Katoh, S. (1977) in Photosynthesis I - Photosynthetic Electron Transport and Photophos-

phorylation (Trebst, A. and Avron, M., cds.), pp. 247-252, Springcr-Vcrlag, Berlin 73 Haehnel, W. (1977) Biochim. Biophys. Acta 459 ,418-441 74 Haehnel, W. (1976) Biochim, Biophys. Acta 440, 506-521 75 Cramer, W.A. and Whitmarsh, J. (1977) Annu. Rev. Plant Physiol. 28, 133-172 76 Vater, J., Rcnger, G., Stiehl, H.H. and Witt, H.T. (1968) Naturwiss. 5 5 , 2 2 0 - 2 2 4 77 Radmer, R. and Cheniae, G. (1977) in Primary Processes of Photosynthesis - Topics in Photo-

synthesis, Vol. 2 (Barber, J., ed.), pp. 303-348, Elsevier/North-Holland, Amsterdam 78 Renger, G., Bouges-Bocquet, B. and Delosme, R. (1973) Biochim. Biophys. Acta 292, 796 807 78a Rcnger, G., Bouges-Boquct, B. and Buchel, K.H. (1973) J. Bioenerg. 4,491 505 78b Junge, W., Rengcr, G. and Ausl~/nder, W. (1977) FI'~BS Lett. 79, 155-159 78c Saphon, S. and Crofts, A.R. (1977) Z. Naturforsch. 32c, 617-626 78d Fowler, Ch. (1977) Biochim. Biophys. Acta 462,414 -421 79 Renger, G. (1977) FEBS lett. 81,223 228

424

79a Kelley, P.M. and Izawa, S. (1978) Biochim. Biophys. Acta 502,198 -2 t0 79b ltorton, P. and Croze, E. (1977) Biochim. Biophys. Acta 462, 86-101 79c Sadewater, D.A. and Dilley, R.A. (1978) Biochim. Biophys. Acta 501,208 216 80 Bishop, N.I. (1959) Biochem. Bull. N.Y. 45, 1696 1702 81 Klingenberg, M., Miiller, A. Schmidt-Mende, P. and Witt, tI.T. (1962) Nature 194,379 380 82 Joliot, P. (1965) Biochim. Biophys. Acta 102, 116 134 83 Malkin, S. and Kok, B. (1966) Biochim. Biophys. Acta 126,413 432 84 Bouges-Bocquet, B. (1973) Biochim. Biophys. Acta 314 ,250 -254 85 Vclthuys, B.R. and Amcsz, J. (1974) Biochim. Biophys. Acta 333, 85 94 86 Siggel, U., Renger, G., Stiehl, H.H. and Rumberg, B. (1972) Biochim. Biophys. Acta 256,328

335 86a Rumberg, B., Sehmidt-Mende, P., Weikard, J. and Witt, H.T. (1963) Natl. Acad. Sci., Natl. Res.

Coun., Washington, D.C., Publication 1145 87 Duysens, L.N.M. (1959) Brookhavcn Symp. Biol. 11, 10-25 88 Witt, H.T., Miiller, A. and Rumberg, B. (1963) Nature 197,987 991 89 Junge, W. and Witt, H.T. (1968) Z. Naturforsch. 23b, 244-254 90 Emrich, ll.M., Jungc, W. and Witt, H.T. (1969) Z. Naturforsch. 24b, 1144-1146 91 Jackson, J.B.and Crofts, A.R. (1969) FEBS Lett. 4, 185 92 Wolff, Ch., Buchwald, H.-E., Rfippel, H. and Witt. H.T. (1969) Z. Naturforsch. 24b, 1038-1041 93 Joliot, P., Delosmc, R. and Joliot, A. (1977) Biochim. Biophys. Acta 459, 47 57 94 Delosme, R., Zickler, A. and Joliot, P. (1978) Biochim. Biophys. Acta 504, 165-174 95 Hager, A. (1969) Planta 89,224 243 96 Labhart, H. (1961) Helv. Chim. Acta 4 4 , 4 4 7 - 4 5 6 97 Liptay, W. (1969) Angew. Chem. 81,195 231 98 Reich, R. and Schmidt, S. (1972) Ber. Bunsenges. Physik. Chem. 76 ,589 -598 99 Schmidt, S. and Reich, R. (1972) Ber. Bunsenges. Physik. Chem. 76,599 602

100 Schmidt, S. and Reich, R. (1972) Bet. Bunsenges. Physik. Chem. 76, 1202-1208 101 Schmidt, S., Reich, R. and Witt, H.T. (1969) Z. Naturforsch. 24b, 1428 1431 102 Schmidt, S., Reich, R. and Witt, H.T. (1971) Naturwiss. 58,414 103 Schmidt, S., Reich, R. and Witt, H.T. (1972) Proc. 2rid Int. Congr. Photosynthesis Res., Stresa

(Forti, G., Avron, M. and Melandri, A., eds.), Vol. 2, pp. 1087-1095, Junk, Den Haag 104 Reich, R., Scheerer, R., Sewe, K.-U. and Witt, H.T. (1976) Biochim. Biophys. Acta 449, 285 -

294 105 Thorne, S.W., Horvath, G., Kahn, A. and Boardman, K. (1975) Proc. Natl. Acad. Sci. U.S. 72,

3858-3862 106 Baltschcffsky, M. (1969) Arch. Biochiem. Biophys. 130, 646-652 107 Fowler, Ch.F. and Kok, B. (1974) Biochim. Biophys. Acta 357 ,299-307 108 Witt, H.T. and Zickler, A. (1973) FEBS Lett. 3 7 , 3 0 7 - 3 1 0 109 Dillcy, R.A. and Vernon, L.P. (1965) Arch. Biochcm. Biophys. 111 ,365-375 110 Schr6der, H., Muhle, H. and Rumberg, B. (1971) Proc. 2nd Int. Congr. Photosynthesis, Stresa

(Forti, G., Avron, M. and Melandri, A., eds.), Vol. 2, pp. 919-930, Junk, Den Haag 111 Amesz, J. and Groth, B.G. (1976) Biochim. Biophys. Acta 440,301 313 112 Schliephake, H., Junge, W. and Witt, H.T. (1968) Z. Naturforsch. 23b, 1571-1578 113 Malkin, R. (1978) FEBS Lett. 8 7 , 3 2 9 - 3 3 3 114 GrSbcr, P. and Witt, H.T. (1974) Biochim. Biophys. Acta 333, 389-392 115 Strichartz, G.R. and Chance, B. (1972) Biochim. Biophys. Acta 256, 71 84 116 Gross, E.L. and Libbey, J.W. (1972) Arch. Biochem. Biophys. 153 ,457-467 117 Schapendonk, A.H.C.M. and Vredcnberg, W.J. (1977) Biochim. Biophys. Acta 462, 613-621 118 Zickler, A., Witt, H.T. and Boheim, G. (1976) FEBS Lett. 66, 142 148 119 Mueller, P. and Rudin, D.D. (1968) Nature 217 ,713 -719 120 Eizenberg, M., Hall, J.E. and Mead, C.A. (1973) J. Membrane Biol. 14, 143-176 121 Bulychev, A.A. and Vredenberg, W.J. (1976) Biochim. Biophys. Acta 423 ,548 -556 122 Saphon, S., Jackson, J.B., Lerbs, V. and Witt, H.T. (1975) Biochim. Biophys. Acta 408, 58 66 123 Bocck, M. and Witt, H.T. (1972) Proc. 2nd Int. Cong. Photosynthesis Res. Stresa 1971 (Forti,

G., Avron, M. and Melandri, A , eds.), pp. 903-911, Junk, Den Haag 124 Goldfield, M.G., Halilov, R.I., Hangulov, S.V., Kononenko, A.A. and Knox, P.P. (1978) Biochem.

Biophys. Res. Comm. 85, 1199- 1203

425

125 Saphon, S., Jackson, J.B. and Witt, H.T. (1975) Biochim. Biophys. Acta 408, 67 82 126 Nobel, P.S. and Mel, H.C. (1966) Arch. Biochem. Biophys. 113,695 702 126a Bose, S. andHoch, G.l:,.(1978) Z. Naturforsch. 33c, 105- 107 126h Witt, 1t.T. (1955) Z. Elektroehem. 59,981-986 126c Witt, H.T., Rumberg, B., Sehmidt-Mende, P., Siggcl, U., Skcrra, B., Vater, 1. and Weikard, J

(1965) Angew. Chem. (Int. Edn.) 4, 799 819 127 Rumberg, B. and Muhle, H. (1976) Bioelectrochem. Bioenerg. 3,393- 403 128 Zickler, A. (1979) Thesis, Berlin 129 B6ger, P. (1978) in Photosynthesis '77. Proc. IV Int. Congr. Photosynthesis (Hall, D.O., Coombs,

J. and Goodwin, T.W., eds.), pp. 755 764, The Biochemical Society, London 130 Emrich, E.M., Junge, W. and Witt, H.T. (1969) Naturwiss. 56, 514-515 131 Chance, B. and Baltscheffsky, M. (1975) in Biomembranes (Eisenberg, H., Katchalski-Katzir, E.

and Manson, L.A., eds.), Vol. 7, pp. 33-55, Plenum Press, New York 132 De Grooth, B.G. and Amesz, J. (1977) Biochim. Biophys. Acta 462, 237 246 133 Akerman, K.E. and Wikstr6m, M.K.F. (1976) FEBS Lett. 68, 191 197 134 Barber, J. and Kraan, G.P.B. (1970) Bioehim, Biophys. Acta 197, 49 59 134a Barber, J. (1972) FEBS Lett. 20,251-254 135 Wraight, C.A. and Crofts, A.R. (1971) Eur. J. Biochem. 19, 386-397 136 Rottenberg, H., Grunwald, T. and Avron, M. (1972) Eur. J. Biochem. 25, 54-63 137 Schr6der, H., Muhle, M. and Rumberg, B. (1972) Proc. 2nd Int. CongL Photosynthesis Res.

(Forti, G., Avron, M. and Melandri, A., eds.), pp. 919-930, Junk, Den Haag 138 Bakecva, L.E., Grinius, L.L., Jasaitis, A.A,, Kuliene, V.V., Levitsky, D.O., Liberman, E.A.,

Severina, L.I. and Skulachev, V.P. (1970) Biochim. Biophys. Acta 216, 13 -21 139 Bulychev, A.A. and Vredenberg, W.J. (1976) Biochim. Biophys. Acta 423,548 556 140 Trebst, A. (1974) Annu. Rev. Plant Physiol. 25,423-458 141 Renger, G. (1976) Biochim, Biophys. Acta 440, 287-300 142 Hauska, G.A., McCarty, R.E., Berzborn, R.J. and Racker, E. (1971) J. Biol. Chem. 246, 3524 143 Junge, W. (1974) Proc. 3rd Int. Congr. Photosynthesis Res. Rehovot, Israel (Avron, M., ed.)

pp. 273-286, Elsevier, Amsterdam 144 Junge, W. and Eckhof, A. (1974) Bioehim. Biophys. Acta 357,103-117 145 Breton, J. (1976) Biochim. Biophys. Acta 459, 66--75 146 Mathis, P., Breton, J., Vermeglio, A. and Yates, M. (1976) FEBS Lett. 63, 171 173 147 Gregory, R.P.F. (1977) in Primary Processes of Photosynthesis - T o p i c s in Photosynthesis,

Vol. 2 (Barber, J., ed), pp. 465-492, Elsevier/North-Holland, Amsterdam 148 Reimer, S. and Trebst, A. (1975) Biochem. Phys. Pflanz. 168,225 149 Reich, R. and Scheerer, R. (1976) BeE Bunsenges. Physik. Chem. 80, 542-547 150 Breton, J., Michel-Villaz, M. Paillotin, G. (1973) Biochim. Biophys. Acta 314, 42-56 151 Bccker, J., Breton, J., Gcacintov, N.E. and Trentacosti, T. (1976) Biochim. Biophys. Acta 440,

531 544 152 Reich, R. and Sewe, K.-U. (1977) Photochem. Photobiol. 26, 11-17 153 Sewe, K.-U. and Reich, R. (1977) Z. Naturforsch. 32c, 161-171 154 Renger, G. and Tiemann, R. (1979) Biochim. Biophys. Acta 545,316-324 155 Hildreth, W.W. (1970) Arch. Biochem. Biophys. 139, 1-8 156 Thomas, J.B. (1972) Proc. 2nd Int. Congr. Photosynthesis Res., Stresa 1971 (Forti, G., Avron, M.

and Melandri, A., eds.), Vol. 2, p. 1509, Junk, Den Haag 157 Conjeaud, H., Michel-Villaz, M., Vermeglio, A. and Mathis, P. (1976) FEBS Lctt. 71, 138-141 158 Sewc, K.-U. and Reich, R. (1977) FEBS Lett. 80, 30-34 159 Crofts, A.R., Prince, R.C., Hohnes, N.G. and Crowther, D. (1974) Proc. 3rd Int. Congr. Photo-

synthesis (Avron, M., ed.), pp. 1131 - 1146, Elsevier, Amsterdam 160 Holmes, N.G. and Crofts, A.R. (1977) Biochim. Biophys. Acta 459,492-505 161 De Grooth, B.G. and Amcsz, J. (1977) Biochim. Biophys. Acta 462,237-246 162 Okada, M. and Takamiya, A. (1970) Plant Cell Physiol. 11,713-721 163 Heathcote, P., Vermeglio, A. and Clayton, R.K. (1977) Biochim. Biophys. Acta 461,358-364 164 Cogdell, R.J., Celis, S., Celis, A. and Crofts, A.R. (1977) in 4th Int. Congr. Photosynthesis,

Reading, U.K. (Hall, D.O., Coombs, J. and Goodwin, T.W., eds.), Biochemical Society, London 165 Gr~ber, P. and Witt, H.T. (1974) Proc. 3rd Int. Congr. Photosynthesis, Rehovot, Israel (M. Avron,

ed.), pp. 951 956, Elsevier, Amsterdam

426

165a Torres-Pereira, J., Mehlhorn, R., Keith, A.D. and Packer, L. (1974) Arch. Biochem. Biophys 160, 9 0 - 9 9

166 Schmidt-Mende, P. and Witt, H.T. (1968) Z. Naturforsch. 23b, 228 235 167 Baltscheffsky, M. (1977) in Living Systems as Energy Converters (Buyer, R., Allen, M.J. and

Massue, J.P., eds.), pp. 199 207, North-Holland, Amsterdam 168 Witt, H.T. and Moraw, R. (1959) Z. Physik. Chem. Neue Folge 20, 253 282 169 Jagendorf, A.T. and Hind, G. (1963) in Photosynthetic Mechanism of Green Plants, Natl. Acad.

Sei. Res. Coun., Washington, D.C. Publ. 1145, pp. 599 -610 170 Neumann, J. and Jagendorf, A.T. (1964) Arch. Biochem. Biophys. 107, 1 0 9 - t 19 171 Rottenberg, H. (1977) in Photosynthesis 1 - Photosynthetic Electron Transport and Photophos-

phorylation (Trebst, A. and Avron, M., eds.), pp. 338-349, Springer-Verlag, Berlin 172 Mitchell, P. (1966) Biol. Rev. 4 1 , 4 4 5 - 5 0 2 173 Jagendorf, A.T. (1977) in Photosynthesis I - Photosynthetic Electron Transport and Photophos-

phorylation (Trebst, A. and Avron, M., eds.), pp. 307 337, Springer-Verlag, Berlin 174 Junge, W. and Ausl~inder, W. (1974) Biochim. Biophys. Acta 333, 5 9 - 7 0 175 Fowler, C.F. and Kok, B. (1974) Biochim. Biophys. Acta 357,308 318 176 Gr/iber, P. and Witt, H.T. (1975) FEBS Lett. 59, 59 -70 177 Ausl/inder, W. and Junge, W. (1975) FEBS Lett. 59 ,310 315 178 Rumbcrg, B. and Siggel, U. (1969) Naturwiss. 56, 130 132 178a Pick, U., Rottenberg, H. and Avron, M. (1973) Ft'BS Lett 32, 91 94

179 Fowler, C.F. and Kok, B. (1976) Biochim. Biophys. Acta 423,510 523 180 Saphon, S. and Crofts, A.R. (1977) Z. Naturforsch. 32c, 810-816 181 Witt, H.T. and Zickler, A. (1974) FEBS Lett. 39,205 208 182 Mitchell, P. (1975) FEBS Lett. 59, 137--139 183 GriJnhagen, H.H. and Witt, H.T. (1970) Z. Naturforsch. 25b, 373 380 184 Ausl~nder, W. and Junge, W. (1974) Biochim. Biophys. Acta 357,285 298 185 Haehnel, W. (1976) Biochim. Biophys. Acta 4 2 3 , 4 9 9 - 5 0 9 186 Hauska, G. (1977) in Bioenergeties of Membranes (Packer, L., Papageorgiou, G.C. and Trebst, A.,

cds.) pp. 177 187, Elsevier/North-Holland Biomedical Press, Amsterdam 187 McLaughlin, S. (1977) in Current Topics in Membranes and Transport (Knotter, F. and Kleinzel-

ler, A., eds.), pp. 71 144, Vol. 5, Academic Press, New York 188 Auslblndcr, W. and Junge, W. (1978) in The Proton and Calcium Pumps (G.F. Azzone et al.,

eds.) pp. 31-44 , Elsevier/North-Holland Biomedical Press, Amsterdam 189 Rumberg, B. and Siggel, U. (1969) Naturwiss. 56, 130 132 190 Tiemann, R., Renger, G., Gr/iber, P. and Witt, H.T. (1979) Biochim. Biophys. Acta, in press 191 Bambcrger, E.S., Rottcnberg, H. and Avron, M. (1973) Eur. J. Biochem. 34 ,557 -563 192 Hauska, G. and Trebst, A. (1977) in Current Topics in Bioenergetics, Vol. 6 (Sanadi, D.R., ed.),

pp. 151-220, Academic Press, New York 193 Reeves, S.G. and Hall, D.O. (1978) Biochim. Biophys. Acta 463 ,275 -297 194 Reinwald, E., Stiehl, H.H. and Rumberg, B. (1968) Z. Naturforsch. 23b, 1616 1617 195 Siggel, U. (1976) Bioelcctrochem. Bioenerg. 3 ,302 318 196 Oesterhelt, D. (1976) Angew. Chem. 88, 16 -24 197 Witt, H.T. and Moraw, R. (1959) Z. Physik. Chem. Neue Folge 20, 253 282 198 Joliet, P. and Delosme, R. (1974) Biochim. Biophys. Acta 357,267 284 198a Rathenow, M. and Rumberg, B. (1978) Biophysiktagung Ulm, Abstr. F11 199 Slater, E.C. (1953)Nature 172, 975 982 200 Boyer, P.D. (1967) Curt. Topics Bioenerg. 2, 99- 149 201 Green, D.E., Asai, J., Harris, R.A. and Penniston, J.T. (1968) Arch. Biochem. Biophys. 125,

684 705 202 Slater, E.C. (1972) in Mitocbondria: Biogenesis and Bioenergetics Biomcmbrancs: Molecular

Arrangements and Transport Mechanisms (S.G. Van den Bergh et al., eds.), pp. 133 146, Elsevier, Amsterdam

203 Witt, H.T., D6ring, G., Rumberg, B., Schmidt-Mende, P., Siggel, U. and Stiehl, H.H. (1966) Brookhaven Symp. Biol. No. 19, pp. 160-168, Upton, New York

204 Rumberg, B., Schmidt-Mende, P., Siggel, U. and Witt, tt.T. (1966) Angew. Chem. 5 ,550 551 205 Rumberg, B. and Siggel, U. (1968) Z. Naturforsch. 23b, 239 244 206 Junge, W., Rumberg, B. and Schr6der, H. (1970) Fur. J. Biochem. 14, 575 581

427

207 Baltscheffsky, M. (1969) Arch. Biochem. Biophys. 130, 646 208 Joliot, P. and Delosme, R. (1974) Biochim. Biophys. Acta 357,267-284 209 Schmid, R. and Junge, W. (1975) Biochim. Biophys. Acta 394, 76-92 209a Schmid, R., Shavit, N. and Junge, W. (1976) Biochim. Biophys. Acta 430, 145-153 210 Jackson, 1 .B., Crofts, A.R. and yon Stedingk, L.-V. (1968) Eur. J. Biochem. 6, 41-54 211 Gr~iber, P. and Witt, H.T. (1976) Biochim. Biophys. Acta 423, 211a Baltschcffsky, M. and Hall, D.O. (1974) FEBS Lett. 39 ,345-348 211 b Baltschcffsky, M. (1971 ) in Encrgy Transd uction in Respiration and Photosynthesis, (Quagliariello,

E., Papa, S. and Rossi, C.S., eds.), p. 639, Adriatica Editrice, Bari 212 Rumberg, B. and Schr6der, H. (1974) Proc. VI Int. Congr. Photobiol., Frankfurt 1972 (G.O.

Schcnck, ed.), paper No. 036, Max-Placnk-lnst., Miilheim 213 Portis, A.R. and McCarty, R.E. (1974) J. Biol. Chem. 249, 6250-6254 214 Junge, W. (1970) Eur. J. Biochem. 14, 582-592 214a Junge, W. (1972) Proc. 2nd Int. Congr. Photosynthesis Res., Stresa 1971 (Forti, G., Avron, M.

and Melandri, A., eds.), pp. 1065-1074, Junk, Den Haag 215 Jagendorf , A.T. (1975) in Bioenergetics of Photosynthesis (Govindjee, ed.), pp. 414 494,

Academic Press, New York 216 Schuldiner, S., Rottenberg, H. and Avron, M. (1973) Eur. J. Biochem. 39,455-462 217 Kraayenhof, R. (1969) Biochim. Biophys. Acta 180, 213 215 218 Rosing, J. and Slater, E.C. (1972) Biochim. Biophys. Acta 267,275-290 219 Jagendorf, A.T. and Uribe, E. (1966) Proc. Natl. Acad. Sci. U.S. 50, 170-177 220 Witt, H.T., Schlodder, E. and Gr~iber, P. (1976) FEBS Lett. 69, 272-276 221 Witt, H.T., Schlodder, E, and GrSber, P. (1977) in Bioenergetics of Membrane~ (Packer, L.,

Papagcorgiou, G.C. and Trebst, A., eds.), pp. 447-457, Elsevier/North-Holland Biomedical Press, Amsterdam

222 Kraayenhof, R. (1977) in Photosynthesis I - Photosynthetic Electron Transport and Photophos- phorylation (Trebst, A. and Avron, M., eds.), pp. 423-428, Springer-Verlag, Berlin

223 Harris, D.A. and Slater, E.C. (1975) Biochim. Biophys. Acta 387,335-348 224 Boyer, P.D., Smith, D.G., Rosing, J. and Kayalar, C. (1975) in Electron Transfer Chains and

Oxidative Phosphorylation (Quagliariello, E., Papa, S., Palmieri, F., Slater, E.C. and Silibrandi, N., eds.), pp. 361-372, North-Holland Publ. Co., Amsterdam

224a Strotmann, H., Bickel, S. and Huchzermeyer, B. (1976) FEBS Lett. 61 ,194-198 225 Gr~iber, P., Schlodder, E. and Witt, H.T. (1977) Biochim. Biophys. Acta 461,426-440 226 Smith, D.J. and Boyer, P.D. (1976) Proc. Natl. Acad. Sci. U.S. 73, 4314-4318 227 Williams, R.J.P. (1975) in Electron Transfer Chains and Oxidative Phosphorylation (Quagliariello,

E., Papa, S., Palmieri, F., Slater, E.C. and Siliprandi, N., eds.), pp. 417 422, North-Holland Publ. Co., Amsterdam

227a Uribe, E.(1973) FEBSLett. 36,143-147 228 Williams, R.J.P. (1976) Trends Biochem. Sci. N 222-224 228a Arnold. W. and Azzi, J. (1971) Photochem. Photobiol. 14,233 240 228b Ellenson, J.L. and Sauer, K. (1976) Photochem. Photobiol. 23, 113-123 229 Ort, D.R., Dilley, R.A. and Good, N.E. (1976) Biochim. Biophys. Acta 449,108 124 230 Racker, E. and Stoeckenius, W. (1974) J. Biol. Chem. 249,662 231 Carmeli, C. (1970) FEBS Lett. 7,297 -300 232 McCarty, R.I'. and Racker, E. (1966) Brookhaven Symp. Biol. 19, 202-214 233 Ryrie, I.J. and Jagendorf, A.T. (1971) J. Biol. Chem. 246, 3771-3774 234 Boyer, P.D., Cross, R.L. and Momscn, W. (1973) Proc. Natl. Acad. Sci. U.S. 70, 2837-2839 235 Harris, D.A. and Slater, E.C. (1975) Biochim. Biophys. Acta 387,335-348 236 Mitchell, P. (1974) FEBS Lett. 43 ,189-194 237 Boyer, P.D., Chance, B., Ernster, L., Mitchell, P., Racker, E. and Slater, E.C. (1977) Annu. Rev.

Biochem. 46,955-1026 238 Harris, D.A. (1978) Biochim. Biophys. Acta 463,245-273

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