Photosynthesis Research 24: 75-80, 1990. © 1990 Kluwer Academic Publishers. Printed in the Netherlands.

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Photosynthetic formation of inorganic pyrophosphate in phototrophic bacteria Beston F. Nore I, Pgd Nyr6n, Gaza F. Salih & Ake Strid Department of Biochemistry, Arrhenius Laboratories for .Natural Sciences', University of Stockholm, S-106 91 Stockholm, Sweden. t To whom correspondence should be addressed Received 21 February 1989; accepted in revised form 28 September 1989

Key words: ATP synthesis; flash-induced; photophosphorylation; pyrophosphate synthesis; Rb. capsutatus; Rps. blastica; Rps. viridis Abstract

In this paper we report studies on photosynthetic formation of inorganic pyrophosphate (PP0 in three phototrophic bacteria. Formation of PP~ was found in chromatophores from Rhodopseudomonas vMdis but not in chromatophores from Rhodopseudomonas blastica and Rhodobacter capsufatus. The maximal rate-of PPi synthesis in Rps. viridis was 0.15#tool PPi formed/(min,/~mol Bacteriochlorophytl) at 23°C. The synthesis of PP~ was inhibited by electron transport inhibitors, uncouplers and fluoride, but was insensitive to oligomycin and venturicidin. The steady state rate of PP~ synthesis under continuous illumination was about 15% of the steady-state rate of ATP synthesis. The synthesis of PP~ after short light flashes was also studied. The yield of PPi after a single lms flash was equivalent to approximately 1 #tool PPi/500#mol Bacteriochlorophyll. In Rps. vMdis chromatophores, PPi was also found to induce a membrane potential, which was sensitive to carbonyl cyanide p-trifluoromethoxyphenylhydrazone and NaF.

Abbreviations; BChl-Bacteriochlorophyll, FoFi-ATPase-Membrane bound proton translocating ATP synthase, FCCP-Carbonyl cyanide p-trifluoromethoxyphenylhydrazone, H+-PPase-Membrane bound proton translocating PPi synthase, TPP + - Tetraphenyl phosphonium ion, TPB--Tetraphenyl boron ion, A ~ - Transmembrane electrical potential difference.

Introduction

Many investigations on the photophosphorylation of ADP to ATP in phototrophic bacteria have been published. In contrast there are only a few reports on photophosphorylation of P~into inorganic pyrophosphate (PPi) [1, 2, 9, 18]. All these studies were performed upon chromatophores isolated from the phototrophic non-sulphur purle bacterium Rhodospiriltum rubrum. The synthesis of PPi in R. rubrum is catalyzed by a reversible membrane bound proton translocating PPase (H+-PPase). This energy transducing enzyme converts chemical energy from the hydrolysis of PPj to an electrochemical gradient of protons

across the cytoplasmic membrane. The catalytic portion of the H ~-PPase is located at the cytoplasmic surface of the plasma membrane of the cells, but in Ribi press vesicles (chromatophores) the active site is located at the outer surfiace. Intact cells pump protons outwards via the H + -PPase during PPi hydrolysis, whereas chromatophores pump protons inwards, creating a proton motive force (acidic and positive inside). The proton motive force generated by light, induces PP~ synthesis by chromatophores if Pi is present in the medium. PPi synthesis can be achieved in both continuous [18] and intermittent light [19]. It is also possible to drive PPi synthesis by the reverse transhydrogenase reaction [15] and by artificially imposed ion poten-

76 tials [24]. It is mainly the rate constants of PP~ synthesis, not of PPi hydrolysis, which are influenced by the electrical potential across the membrane [25]. The H + -PPase has been solubilized and purified from R. rubrum chromatophores [16]. Despite the fact that photosynthetic PP~ formation only has been found in R. rubrum there might as well be other phototrophic bacteria capable of catalyzing PP~ synthesis. A membrane bound PPase has been found in Rhodopseudomonas palustris [14, 21], and there are indications of membrane bound PPases in Rhodopseudomonas viridis [10], in Rhodobacter sphaeroides [23] and in Chromatium vinosum [13]. The reason why, until now, PP~ synthesis coupled to electron transport has been more closely examined only in R. rubrum might be the lack of a method for continuous monitoring of PP~ synthesis. In this paper we examine the possibility of PP~ synthesis in chromatophores from Rps. viridis, Rps. blastica and Rb. capsulatus.

addition of thiamin pyrophosphate hydrochloride (1 mg/1). BChl was measured by extraction with acetone/methanol (7:2) using e790nm= 75mM -t * cm- 1 [7], •767nm ~--" 76 mM-~ • cm-1 [8] and eV72nrn = 75mM -1 * cm -1 [6] for Rps. viridis, Rps. blastica and Rb. capsulatus, respectively. Cells were harvested and chromatophores were prepared as in Ref. 22, except that cells from Rps. viridis and Rps. blastica were passed twice through the Ribi Cell fractionator.

Assay of A TPase and PPase activity The method used was a modification, previously described [18], of the Rathbun method [20]. The assay mixture contained 0.1 M glycylglycine (pH 7.4), 0.75raM MgClz, 0.5mM ATP or PPi, and chromatophores corresponding to 1.4 #M BChl.

Measurement of continuous photophosphorylation Materials and methods

Purified luciferase (EC 1.13.12.7), D-luciferin and L-luciferin were obtained from LKB Wallac (Turku, Finland). Venturicidin was obtained from BDH Chemicals Ltd (Poole, England). Myxothiazol was purchased from Boehringer (Mannheim, FRG). TPP+C1 - and Na+TPB - were from Aldrich Chemical Co. Ltd (Steinheim, FRG). Other materials were of reagent grade and obtained from commercial sources.

Preparation of chromatophores Rps. viridis (DSM 133) was grown anaerobically in light at 30°C in the medium described by Bose et al. [3] with the addition of nicotinic acid (1 rag/l), yeast extract (0.3 g/l), thiamine pyrophosphate hydrochloride (1 mg/1), 3mM sodium acetate, and 17raM glutamate. Batch cultures of Rps. blastica (strain NC1B 11576) were grown anaerobically in light at 30°C in the same medium [3] with the addition of yeast extract (1 g/l), pyruvate (0.5g/l) but only half the amount of malate. Rb. capsulatus (strain N22) was grown anaerobically in light at 30°C in the medium [3] with the

Continuous monitoring of PP~ synthesis was carried out as described earlier [17], with some minor modifications. All experiments were performed at 23°C. The standard assay volume was 1 ml and contained the following components: 0.1 M glycylglycine (pH 7.75); 2raM ethylenediamine tetraacetic acid; 10raM magnesium acetate; 0.1% bovine serum albumin; l mM 1,4-dithioerythritol; 20/~g D-luciferin; 0.4 #g L-luciferin; 0.4 mg polyvinylpyrrolidone 360,000; 5#M adenosine 5'phosphosulphate; 0.3 U ATP-sulphurylase; 1 mM NaPi (PPi free) (pH 7.5); 0.1 mM sodium succinate; 1 #M P~, PS-di(adenosine-5') pentaphosphate; 1 #g oligomycin; purified luciferase (for the amount see Ref. 17) and chromatophores corresponding to BChl concentrations of between 0.5 and 2.3 #M. The ATP synthesis assay was performed in the same manner as the PP~ determination with the exception that the standard assay volume contained 50#M ADP. Adenosine-5'-phosphosulphate and ATP-sulphurylase were omitted.

Measurement of flash-induced PPI and A TP synthesis The ATP and PPi were determined as described

77

earlier [19]. A flash unit emitting flashes of 1 ms duration was used. The assay was the same as above for measurement of continuous ATP and PPi syntheses, except that chromatophores corresponding to a BChl concentration of 2.5 #M were used.

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The relative A~ (interior positive) was monitored by using the TPB ion selective electrode as described by Casadio et al. [4]. The sensor membrane containing TPB- as anion exchanger was made as described by Kamo et al. [11]. The experiments were carried out at 23°C. The 2ml assay buffer contained 1 #M TPB ; 0.1 #M TPP + (to increase the permeability of the chromatophore membrane for TPB-); 50mM Na-glycylglycine (pH 7.5); 0.1 mM Na-succinate; 25mM NaC1 and 5mM MgC12. Chromatophores were added to a final concentration of 9#M BChl. The reaction was started either by illumination or by addition of 0.5 mM PP~.

Results

o 20

PPi

1

2

Time

3

(min)

Fig. 1. A typical trace from the measurements of photophosphorylation of P~ to PPi during continuous illumination of Rps. viridis chromatophores. Experimental conditions were as described under Materials and Methods. The following additions were made: 20 pmol PPi, 10 #mol fluoride. The BChl concentration was 0.75#M.

ed by the uncoupler FCCP, the electron transport inhibitor myxothiazol and the PPase inhibitor fluoride, but not by the FoF1-ATPase inhibitors oligomycin and venturicidin.

Synthesis of PPi under continuous illumination The monitoring of PPi synthesis in continuously illuminated Rps. viridis chromatophores is shown in Fig. 1. In order to eliminate phosphorylation of endogenous ADP by the FoF1-ATPase and the adenylate kinase activity, oligomycin and pi,ps_ di(adenosine-5') pentaphosphate were included. The steady-state rate of PPi synthesis under continuous illumination was 0.15pmol PPi/ (rain • #mol BChl). This rate is about 15% of the steady-state rate of ATP synthesis under the same conditions. We could not detect any PP~ synthesis activity at all with chromatophores from Rps. blastica and Rb. capsulatus. The absence of PP~ synthesis activity was not due to badly coupled particles since a substantial ATP synthesis rate was obtained, 2.8 #tool ATP/(min • #tool BChl) for Rps. blastica and 2.9 #tool ATP/min • #mol BChl) for Rb. capsulatus. The results are in accordance with Ref. [26] and [5] respectively. Table 1 gives a summary of the effects of different compounds tested on the PP~ synthesis. The PP~ synthesis was inhibit-

Synthesis of PPi and ATP after light flashes As demonstrated in Fig. 2, the chromatophores of Rps. viridis were also competent in catalyzing a flash-stimulated PPi synthesis. The amount of PPi formed after a 1 ms light flash was approximately equivalent to 1/~mol PPi/500#mol BChl. This value can be compared with the yield of ATP after one flash, which was 1 #tool ATP/1000 #tool BChl. These ratios were the same as have been observed earlier for R. rubrum. However, in R. rubrum the yield of PPi after a short flash was higher than in Rps. viridis [19].

Hydrolysis of PPi and A TP The Rps. viridis chromatophores, but not the Rps. blastica or the Rb. capsulatus chromatophores, catalyzed the hydrolysis of PP~. The rate of PPi hydrolysis was 1.0 #tool PPj hydrolyzed/(min • #tool

78 Table 1. The effect of various agents on photosynthetic PPi formation in Rps. viridis Additions

#mol PPi formed/ (min * ~mol BChl)

Rate of PPi formed (% of control)

Control Oligomycin (1 #g/ml) Venturicidin (1 #g/ml) FCCP (1 #M) Myxothiazol (1 #M) Na-Fluoride (10 mM)

0.15 0.15 0.15 0.05 0.0 0.0

100 100 100 33 0 0

PPi formation in the presence and absence of these agents was measured as described in the Materials and Methods. The BChl concentration was 1.2 #M.

BChl) (Table 2). The rates of ATP and PPi hydrolysis were stimulated in Rps. viridis chromatophores only to a very small extent (5-15%) in the presence of I # M FCCP, compared with 2- and 7-fold at their best in R. rubrum for PP~ and ATP hydrolysis respectively (Table 2).

extent, was observed after addition of 10 mM of the PPase inhibitor fluoride (not shown). The PPiinduced potential was similar to the ATP-induced membrane potential (not shown).

Discussion

PPi-induced membrane potential generation (AtP) In Fig. 3 the PPcinduced response of a TPB- ion selective electrode in Rps. viridis chromatophores is shown. The addition of 0.5 mM PPi to a suspension containing 9 #M BChl induced a membrane potential, which corresponds to a signal by the electrode of about 10 mV (arbitrary units). The PPi-induced membrane potential could be abolished by addition of 1 #M FCCP. The same effect, but to a lower 100

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Fig. 2. A typical trace from the measurements of PP~ formation after a single 1 ms flash in Rps. viridis chromatophores. The experimental conditions were as described under Materials and Methods. The following additions were made: I pmol PP~. The BChl concentration was 3.75 #M.

In Table 2 are summarized the results of the experiments on four different phototrophic bacteria taken from this and earlier studies [18]. Of the four phototrophic bacteria studied, only R. rubrum and Rps. viridis catalyzed photophosphorylation of Pi to PPi. Both the highest PPi synthesis and the highest PP~ hydrolyzing activity were obtained with R. rubrum chromatophores. No PPi hydrolyzing or synthesizing activity was observed in chromatophores from Rps. blastiea or Rb. capsulatus. The large difference in PPi synthesis activity between R. rubrum and Rps. viridis can be explained mainly by a difference in coupling capacity. This is indicated by the low uncoupler stimulation of both the PPase and the ATPase activities in Rps. viridis chromatophores and also by the low ATP synthesis activity (0.5 #tool ATP formed/(min • #tool BChl) in Rps. viridis (see also Ref. [12]) compared to 10#tool ATP formed/(min • #tool BChl) in R. rubrum. The planer structure of the intracytoplasmic membranes of Rps. viridis is hardly compatible with the formation of well coupled vesicles. This fact explains the differences in rate observed between the membrane preparations from the two bacteria. The present results show that the phototrophic bacterium Rps. viridis contains a membrane bound PPase, which is able to couple the energy liberated from light driven electron transport to synthesis of PPi. Both continuous illumination and short light flashes are possible energy sources for driving PP~

79 Table 2. The rate of PP~ and ATP synthesis, and PPi and ATP hydrolysis in four phototrophic bacteria

Activity

Bacteria studied

PPi sYnthesis~ ATP synthesis PPi hydroly sisc : Control + 2#M FCCP ATP hydrolysisC: Control + 1/~M FCCP

Rps. viridis

R. rubrum b

Rps. blastica

Rb. capsulatus

0.1 0.5

1.2 10.0

0.0 2.8

0.0 2.9

1.0 1.2

6.0 12.0

0.0 0.0

0.0 0.0

1.36 1.50

1.0 8.0

0.8 1.4

n.d. n.d.

See the Materials and Methods for experimental details. "#mol PPi or ATP formed/(min *#mol BChl). b Ref. [18]. C#mol PP~ or ATP hydrolyzed/(min • # mol BChI).

synthesis. The hydrolysis of PP~ is coupled to proton pumping over the membrane, as indicated by the generation of a membrane potential. We have shown in this study that R. rubrum is not unique among phototrophic bacteria in being able to couple light driven electron flow to the synthesis of PP~. Why some, and not all, phototrophic bacteria contain a membrane bound PPase is not easily explained.

Acknowledgements This work was partly supported as a grant by the

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Fig. 3. A typical trace from the measurements of PPi-induced generation of A ~ in Rps. viridis chromatophores. The experimental conditions were as described under Materials and Methods. The BChl concentration was 9/~M. In the assay medium, 0.5 #mol PP~ was added.

Swedish Natural Science Research Council (NFR) to Prof. M. Baltscheffsky and to P.N.. We are grateful to Miss Yoko Sakai for measuring the: membrane potentials, Dr. Baz Jackson for the Rb. capsulatus culture (strain N22) and Dr. Gunnar Falk for the Rps. blastica culture.

References 1. Baltscheffsky, H. and yon Stedingk, L.-V. (1966) Biochem. Biophys. Res. Commun. 22, 722-728. 2. Baltscheffsky, H., von Stedingk, L.-V., Heldt, H.-W. and Klingenberg, M. (1966) Science 153, 1120-1122. 3. Bose, S.K., Gest, H., and Ormerod, J.G. (1961) J. Biol. Chem. 236, PC 13-14. 4. Casadio, R., Venturoli, G. and Melanderi, B.A. (1981) Photobiochem. Photobiophys. 2, 245-253. 5. Clark, A.J., Cotton, N.P.J. and Jackson, J.B. (1983) Biochim. Biophys. Acta 723, 440-453. 6. Clayton, R.K. (1963) Biochim. Biophys. Acta 75, 312-323. 7. Clayton, R.K. (1963) in Bacterial Photosynthesis (Gest, H., San Pietro, A., and Vernon, L.P., eds.), p. 495-500, Antioch Press, Yellow Springs, Ohio. 8. Clayton, R.K. (1966) Photochem. Photobiol. 5, 669-677. 9. Guillory, R.J. and Fisher, R.R. (1972) Biochem. J. 129, 471-481. 10. Jones, O.T.G. and Saunders, V.A. (1972) Biochim. Biophys. Acta 275, 427-436. 1I. Kamo, N., Muratsugu, M., Hongoh, R. and Kobatake, Y. (1979) J. Membr. Biol. 49, 105-121. 12. Kerber, N.L., Pucheu, N.L. and Garcia, A.F. (1977) FEBS Lett. 80, 49-52. 13. Knaff, D.B. and Carr, J.W. (1979) Arch. Biochem. Biophys. 193, 379-384. I4. Knobloch, K. (1975). Z. Naturforsch. 30c, 342-348. 15. Nore, B.F., Husain, I., Nyr6n, P. and Baltscheffsky, M. (1986) FEBS Lett. 200, 133-138. 16. Nyr6n, P., Hajnal, K. and Baltscheffsky, M. (1984) Biochim. Biophys. Acta 766, 630-635. 17. Nyr6n, P. and Lundin, A. (1985) Anal. Biochem. 151, 504-509.

8O 18. Nyr6n, P., Nore, B.F. and Baltscheffsky, M. (1986) Biochim. Biophys. Acta 851,276-282. 19. Nyr6n, P., Nore, B.F. and Baltscheffsky, M. (1986) Photobiochem. Photobiophys. I1, 189-196. 20. Rathbun, W.B. and Betlach, V.M. (1969) Anal. Biochem. 28, 436-445. 21. Schwarm, H.-M., Vigenschow, H. and Knobloch, K. (1986) Biol. Chem. Hoppe-Seyler 367, 127-133.

22. Shakhov, Yu.A., Nyr6n, P. and Baltscheffsky, M. (1982) FEBS Lett. 146, 177-180. 23. Sherman, L.A. and Clayton, R.K. (1972) FEBS Lett. 22, 127-132. 24. Strid, A., Karlsson, I.-M. and Baltscheffsky, M. (1987) FEBS Lett. 224, 348-352. 25. Strid, A., Nyr+n, P., Boork, J. and Baltscheffsky, M. (1986) FEBS Lett. 196, 337-340. 26. Zannoni, D. (1984) Arch. Microbiol. 140, 15-20.

Photosynthetic formation of inorganic pyrophosphate in phototrophic bacteria.

In this paper we report studies on photosynthetic formation of inorganic pyrophosphate (PPi) in three phototrophic bacteria. Formation of PPi was foun...
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