Photosynthesis Research 39: 137-147, 1994. © 1994KluwerAcademicPublishers. Printedin the Netherlands. Regular paper

Membrane barriers and Mehler-peroxidase reaction limit the ascorbate available for violaxanthin de-epoxidase activity in intact chloroplasts Christian Neubauer & Harry Y. Yamamoto* Department of Plant Molecular Physiology, 3190 Maile Way, St. John 503, University of Hawaii at Manoa, Honolulu HI 96822, USA; *Author for correspondence Received 15 July 1993; acceptedin revised form 7 October 1993

Key words: non-radiative energy dissipation, non-photochemical quenching, ascorbate peroxidase, hydrogen peroxide, Photosystem II Abstract

The presence of an acidic lumen and the xanthophylls, zeaxanthin and antheraxanthin, are minimal requirements for induction of non-radiative dissipation of energy in the pigment bed of Photosystem II. We recently reported that ascorbate, which is required for formation for these xanthophylls, also can mediate the needed lumen acidity through the Mehler-peroxidase reaction [Neubauer and Yamamoto (1992) Plant Physiol 99: 1354-1361]. It is demonstrated that in non-CO2-fixing intact chloroplasts and thylakoids of Lactuca sativa, L. c.v. Romaine, the ascorbate available to support de-epoxidase activity is influenced by membrane barriers and the ascorbate-consuming Mehler-peroxidase reaction. In intact chloroplasts, this results in biphasic kinetic behavior for light-induced de-epoxidation. The initial relatively high activity is due to ascorbate preloaded into the thylakoid before light-induction and the terminal low activity due to limiting ascorbate from the effects of chloroplast membranes barriers and a light-dependent process. A five-fold difference between the initial and final activities was observed for light-induced de-epoxidation in chloroplasts pre-incubated with 120 mM ascorbate for 40 min. The light-dependent activity is ascribed to the competitive use of ascorbic acid by ascorbate peroxidase in the Mehler-peroxidase reaction. Thus, stimulating ascorbic peroxidase with H20 2 transiently inhibited de-epoxidase activity and concomitantly increased photochemical~ quenching. Also, the effects inhibiting ascorbate peroxidase with KCN, and the Kra values for~ascorbate peroxidase and violaxanthin de-epoxidase of 0.36 and 3.1 mM, respectively, support this conclusion. These results indicate that regulation of xanthophyll-dependent non-radiative energy dissipation in the pigment bed of Photosystem II is modulated not only by lumen acidification but also by ascorbate availability. Abbreviations: A P O - a s c o r b a t e peroxidase; M P - M e h l e r ascorbate-peroxidase; NIG-nigericin; NPQ - non-photochemical quenching; F o - dark fluorescence; F - fluorescence at any time; F M - maximal fluorescence of the (dark) non-energized state; F~ - maximal fluorescence of the energized state; qp - coefficient for photochemical fluorescence quenching; VDE - violaxanthin de-epoxidase; k - firstorder rate constant for violaxanthin de-epoxidase activity

Introduction

Dissipation of light energy as heat in the pigment bed appears to protect Photosystem II by down-

regulating quantum efficiency. This dissipation of energy, measured experimentally as rapidly-reversible non-photochemical fluorescence quenching (NPQ), is induced when plants are exposed

138 to more light than can be used by CO 2 fixation. The capacity for NPQ varies among plants, being generally higher in sun-tolerant plants than in shade plants (Demmig-Adams et al. 1989). The mechanism and factors influencing NPQ induction are of interest. It has been shown repeatedly that rapidly-reversible NPQ, variously termed qE or SVE depending on how it is quantified (Schreiber et al. 1986, Gilmore and Yamamoto 1991b), is dependent on the presence of an acidic lumen (Briantais et al. 1979). Because of the ApH dependency, this NPQ is also termed 'high-energy' fluorescence quenching. The role of lumen acidity in NPQ, however, is complex and not yet completely understood. There appears to be a threshold ApH for NPQ (Noctor and Horton 1990) that varies with the xanthophyll composition (Gilmore and Yamamoto 1993). Furthermore, it appears from the effects of antimycin that low lumen pH and the presence of de-epoxidized xanthophylls are not final determinants of NPQ (Oxborough and Horton 1987, Gilmore and Yamamoto 1990). Aggregation of LHC II has been implicated in NPQ but also recently questioned (Horton et al. 1991, Huner et al. 1993). The efflux of Mg 2+ or other cations may be required (Gilmore and Yamamoto 1992). Numerous in vivo and in vitro studies have shown that the amount of NPQ is related directly to zeaxanthin concentration (reviewed in Demmig-Adams and Adams 1992a). Recently, antheraxanthin has been shown to have a similar effect as zeaxanthin (Gilmore and Yamamoto 1993). Antheraxanthin increases NPQ and its presence apparently explains NPQ that previously appeared to be unrelated to zeaxanthin (Demmig-Adams et al. 1989, Gilmore and Yamamoto 1991b). Zeaxanthin and antheraxanthin are formed by de-epoxidation of violaxanthin (Yamamoto et al. 1962) under high light (Sapozhnikov 1957) or when CO 2 fixation becomes limiting (Siefermann 1972). Re-epoxidation occurs under darkness or under reduced light (Sapozhnikov 1957). After extended stress treatments, zeaxanthin and antheraxanthin may remain even overnight (Demmig et al. 1988, Demmig-Adams and Adams 1992b). About 80% of violaxanthin is de-epoxidized in intact chloroplasts (Siefermann

and Yamamoto 1974b, Pfiindel and Dilley 1992). In intact leaves up to 90% conversion has been observed at noon (Demmig-Adams and Adams 1992b). The fraction that is available for deepoxidation varies with light intensity, growth conditions (Siefermann and Yamamoto 1974a, Thayer and Bj6rkman 1990) and long periods of reaction (Pf/indel and Dilley 1993). However, even under light-saturating conditions, where the maximum amount of violaxanthin is available, its concentration in situ is not saturating for deepoxidase activity. Accordingly, the kinetics of de-epoxidation is first-order with respect to violaxanthin. Temporal effects on de-epoxidase activity, therefore, must be evaluated by the first-order rate constant (k) so as to be independent of decreasing substrate concentration. Violaxanthin de-epoxidase activity requires an acidic pH (optimally 5.2) and the presence of ascorbate in both thylakoids and in purified enzyme reactions (reviewed in Yamamoto 1985). In isolated chloroplasts, the required acidic conditions can be met by the light-induced proton pump mediated either by artificial electron acceptors or by the Mehler-peroxidase reaction (Yamamoto et al. 1972, Neubauer and Yamamoto 1992). In the latter, oxygen is the electron acceptor, and the resulting superoxide radical is dismutated by superoxide dismutase to hydrogen peroxide, which is then reduced by ascorbate peroxidase (reviewed in Asada and Takahashi 1987). Relatively little attention has been given to the ascorbate requirement for violaxanthin de-epoxidase activity. Ascorbate appears to be specific for the de-epoxidase since no alternative electron donor has been found for chloroplast reactions and it is sufficient for high de-epoxidase activity in thylakoid-free reactions (Yamamoto 1985). Chloroplasts contain 15-25 mM ascorbate which represents 30-40% of the total ascorbate present in spinach leaves (Foyer et al. 1983, Gillham and Dodge 1986). Ascorbate can accumulate up to 50-100 mM during the summer period (see references in Aristarkhov et al. 1988). It is transported from extra plastid sources across the envelope by a specific translocator and then diffuses into the lumen (Anderson et al. 1983, Beck et al. 1983). Ascorbate permeability through chloroplast membranes is slow but suffi-

139 cient to allow equilibration of ascorbate between external medium and lumen in 15 to 20 min (Aristarkhov et al. 1988, Foyer et al. 1990). Also, monodehydroascorbate and dehydroascorbate in the stroma can be reduced to ascorbate by monodehydroascorbate reductase and dehydroascorbate reductase, respectively (Asada and Takahashi 1987). In addition, the monodehydroascorbate radical can be reduced by PSI without participation of the reductase (Miyake and Asada 1992). Siefermann and Yamamoto (1974b) have demonstrated that de-epoxidation can be supported by dehydroascorbate in a ferredoxin-dependent light reaction. In isolated chloroplasts, light-induced de-epoxidation in the presence of ascorbate was biphasic whereas the ascorbate-induced reaction in the dark at pH 5.2 was monophasic (Siefermann and Yamamoto 1974a). These differing kinetics were interpreted as due to loading of the thylakoids with ascorbate. The possibility of a light-dependent effect on biphasic kinetics, however, was not excluded. Herein, we have investigated the biphasic phenomenon for evidence of the effects of membrane barriers and the possible involvement of the light-dependent, ascorbate-consuming Mehler-peroxidase reaction. The results indicate that for light-induced conditions, the initial high activity is from ascorbate preloaded into chloroplasts and the following low activity from limiting ascorbate due to membrane barriers and the competitive use of ascorbate by the Mehler-peroxidase reaction.

supplemented with 30 or 120 mM ascorbate. All reactions were at 30/xg chlorophyll ml -~ and 25 °C. The samples were stirred continuously in an open 1 × 1 cm glass cuvette, which allowed the study of the effect of peroxide-addition and excluded a possible decrease of the oxygen concentration in the medium due to the autoxidation of ascorbate. Ribulosebisphosphate carboxylase/oxygenase activity was inhibited with 5 mM iodoacetamide (IAA) added 3 min before illumination. This concentration of IAA had no effect on APO activity, as assessed by its effect on the hydrogen peroxide-induced photochemical fluorescence quenching of intact, uncoupled chloroplasts (Neubauer and Schreiber 1989).

Chlorophyll fluorescence and AAsos s4o absorbance changes

Materials and methods

Chlorophyll fluorescence and absorbance changes related to violaxanthin de-epoxidation were measured as described earlier (Gilmore and Yamamoto 1991b, Neubauer and Yamamoto 1992). Zeaxanthin and antheraxanthin were analyzed chromatographically as described by Gilmore and Yamamoto (1991a). If not stated otherwise, the light intensities measured at the cuvette were 200 ixmol m -2 s -t and 2500/xmol m - 2 s -t for the actinic and saturating light, respectively. Photochemical fluorescence quenching is expressed as quenching coefficient, qp (Schreiber et al. 1986). Zeaxanthin formation was induced in the dark by adding ascorbate to intact or broken chloroplasts in pH 5.2 reaction medium containing 0.25 M citrate buffer and 0.5/xM nigericin.

Chloroplast isolation and experimental conditions

Results

Intact, class A chloroplasts were isolated from dark-adapted lettuce (Lactuca sativa L. cv. Romaine) as described by Neubauer and Yamamoto (1992). Freshly broken, class-D chloroplasts were prepared by homogenizing 2.5ml intact chloroplasts (30/~g chlorophyll m1-1) for 10 s in a 10 ml ground-glass tissue homogenizer in reaction medium (0.3M sorbitol, l mM MgCI 2, 2 mM EDTA, 0.5 mM KH2PO4, 30 mM KCI, 1 mM MnCI2, 50raM HEPES; pH7.6),

The 505-nm absorbance change reflects a decrease in violaxanthin or an increase in zeaxanthin plus antheraxanthin (Siefermann and Yamamoto 1974a). Since the antheraxanthin concentration is low under the conditions reported here, the absorbance change reflects essentially zeaxanthin formation. Violaxanthin de-epoxidase activity can be evaluated as either the initial rate of the absorbance change or as the first-order rate constant (k, min -1) with

140 respect to violaxanthin. The latter allows following activity temporally under conditions of limiting substrate (Siefermann and Yamamoto 1974a, Pfiindel and Dilley 1993). The effects of ascorbate concentration, preincubation time and chloroplast intactness on violaxanthin de-epoxidase activity under light and dark conditions are summarized in Table 1. Figure 1 shows first-order plots for typical light (condition 2; curve A) and dark (condition 7; curve B) reactions in Table 1. As observed earlier by Siefermann and Yamamoto (1974a), light-induced de-epoxidation in intact chloroplasts, pre-incubated with ascorbate, showed two first-order phases, the relative activity changing from high to low (conditions 1-4, Fig. 1, curve A). Increasing ascorbate concentration and preincubation time increased initial activity but had little effect on final activity. The ratio of initial to final activity increased from 1.18 for condition 1 to 5.05 for condition 4. In the former, chloroplasts were preincubated for 10 min with 30 mM ascorbate and in the latter, for 40min with 120 mM ascorbate. The difference in initial and final activity was less for broken chloroplasts (conditions 5-6) in part due to a higher final activity in broken chloroplasts than in intact chloroplasts. The effects of ascorbate concentration and pre-incubation time on the initial activity is consistent with an effect of preloading of the lumen as suggested earlier (Siefermann

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Fig. 1. Typical first-order plots of the 505 nm absorbance change in intact chloroplasts for light-initiated reaction at p H 7 . 0 (A) and ascorbate-initiated dark reaction at p H 5.2 (B). For the former, chloroplasts were incubated with 30 mM for 40 min before onset of light at time zero. The dark reaction at p H 5.2 was initiated by adding 30 mM ascorbate at time zero. First-order rate constants (k) were calculated from the slopes of the first 2 min (A = 0.457 m i n - l ; B = 0.141 min 1) and from the final linear rates between 5-10 min (A = 0.197 min-1; B = 0.290 min -1) of reaction. AA~05 and AAts05 reflect the maximal absorbance change and the change at any given time, respectively. AA~05 obtained in the light was used to calculate the dark activity. For further conditions see Fig. 2 and 'Materials and methods'.

and Yamamoto 1974a). However, loading of ascorbate into a special thylakoid domain that is accessible to violaxanthin de-epoxidase rather than the lumen is also possible. For dark-induced de-epoxidation in intact chloroplasts, ascorbate was not preloaded and de-epoxidation was initiated instead by adding

Table 1. The effects of ascorbate concentration and preincubation time on de-epoxidase activity. Violaxanthin de-epoxidase activity is expressed as the first-order rate constant k (min-1). Initial rates (ki) were calculated from the first 2 min of de-epoxidation and final rates (kt) from 10 to 15 min of the reaction. Total reaction time was at least 18 min. Intact (class A) or broken (class D) chloroplasts were pre-incubated with either 30 or 120 mM ascorbate for 10 and 40 min before onset of actinic illumination. The data are mean v a l u e s+- S.D. from 3 (conditions 2-10) or i0 (condition 1) experiments using different chloroplast preparations. For further conditions see Fig. 2 and 'Materials and methods' Condition

Preincubation (min)

Class

k i (min- 1)

k t (min- 1)

ki/k t

Light 1) 30 mM ASC 2) 30 mM ASC 3) 120 m M ASC 4) 120 m M ASC 5) 30 m M ASC 6) 120 mM ASC

10 40 10 40 10 10

A A A A D D

0.271 0.473 1.077 1.156 0.456 0.705

--- 0.020 --- 0.083 -+ 0.079 -+ 0.091 - 0.085 -+ 0.100

0.230 0.219 0.238 0.229 0.299 0.282

-+ 0.039 -+ 0.067 -+ 0.044 - 0.046 -+ 0.048 --- 0.062

1.18 2.16 4.53 5.05 1.53 2.50

Dark, p H 5.2 7) 30 mM ASC 8) 120 m M ASC 9) 30 mM ASC 10) 120 mM ASC

0 0 0 0

A A D D

0.142 0.171 0.314 0.327

--- 0.034 -+ 0.056 -+ 0.061 --- 0.083

0.298 0.307 0.314 0.327

- 0.049 --- 0.081 --- 0.064 -+ 0.073

0.48 0.56 1.00 1.00

141 ascorbate. In contrast with light-induced reactions, the initial activity was slower than the final activity (conditions 7-8; Fig. 1, curve B), presumably reflecting the slow diffusion of ascorbate across membrane barriers. In broken chloroplasts (conditions 9-10), de-epoxidation was monophasic. The final rates under dark-reaction conditions were similar, within experimental error, to final rates of light-induced reactions in broken chloroplasts (conditions 5-6) and were comparable to values reported by Siefermann and Yamamoto (1974a) for the light reaction and the long-term experiments by Pffindel and Dilley (1993) for dark reactions. Importantly, these rates are significantly higher than for light-induced de-epoxidation in intact chloroplasts (conditions 1-4). While membrane barriers to ascorbate diffusion are apparent, there appears to be an additional light-dependent factor that partially inhibits de-epoxidase activity. The Mehler-peroxidase reaction is a light-dependent activity that consumes ascorbate. The intact chloroplasts used in this study were isolated by a method that preserves ascorbate peroxidase, a key enzyme of the Mehler-peroxidase reaction while omitting catalase activity, by removal of the peroxisomes during isolation (Nakano and Asada 1980, Neubauer and Schreiber 1989). The hypothesis was tested that the Mehler-peroxidase reaction inhibits de-epoxidase activity by competitively consuming ascorbate. Hydrogen peroxide was added to the intact chloroplasts to oxidize ascorbate by the action of pfascorbate peroxidase, and de-epoxidation and photochemical fluorescence quenching, qv, were followed. As shown by Schreiber and Neubauer (1990), H202-induced stimulation of qp can be used to measure ascorbate-peroxidase activity. The actual Hill oxidant for the observed photochemical fluorescence quenching is the monodehydroascorbate radical formed by the enzymatic H 2 0 2 reduction (Miyake and Asada 1992). Figure 2 shows the effect of H 2 0 2 o n the kinetics of light-induced room-temperature fluorescence and 505-nm absorbance, in the presence of 30 mM. The fluorescence pattern of the control (upper left curve) is typical for non-COE-fixing intact chloroplasts (Neubauer and Yamamoto 1992). Maximum fluorescence (F~) decreased gradually to a steady-state level, reflecting de-

Control "E

0.5 mM H202

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-

ON

NIG

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ON 1"t202

NIG

Fig. 2. Effect of H202 on fluorescence and light-induced zeaxanthin de-epoxidation, in intact non-CO2-flxing lettuce chloroplasts. Chlorophyll fluorescence and 505 nm-absorbance changes were measured simultaneously. Intact percollchloroplasts were illuminated in the presence of 5 mM IAA. The dark-level fluorescence yield, Fo, is reflected by the dotted line. Variable (F), and maximal fluorescence of the non-energized (FM) and energized (F~) state are labeled as indicated. Hydrogen peroxide (0.5 raM) was added about 2.3 rain after onset of actinic light. Nigericin (NIG, 10 ~ M ) was added after 13 rain of illumination. Chloroplasts were pre-incubated with 30 mM ascorbate for 10 rain before onset of actinic light. This condition gives minimal differences in the kinetics between the initial and later phases of deepoxidation (see Table 1). Continuous red, actinic light (200/zmol m 2 s-l) was switched on at the open arrow. For other conditions see 'Materials and methods'.

velopment of NPQ. Most of this quenching was reversed by nigericin and was therefore ApH dependent. When H 2 0 2 w a s added (upper right curve), variable fluorescence (F) as well as F~ decreased rapidly. This sudden stimulation of NPQ was due to the known effect of Photosystem I acceptors decreasing fluorescence yield and was not due to ApH stimulation (data not shown). Photochemical fluorescence quenching, qp, calculated 5 s before and 15 s after H 2 0 2 addition, increased from 0.15 to 0.52. After 12 min, no difference was evident between the peroxide-treated sample and control. These effects are presumably due to transient H20 2induced stimulation of ascorbate oxidation by ascorbate-peroxidase activity (Neubauer and Schreiber 1989). The 505-nm change was also typical for the control (Fig. 2, lower left curve)

142 and as expected, was not reversed by nigericin (Yamamoto et al. 1972). Addition of H 2 0 2 (lower right curve) transiently inhibited AAs05 but had almost no effect on the maximal change and therefore on-de-epoxidation. Chromatographic analysis showed that the final zeaxanthin concentration in the peroxide-treated sample was 95% of the control (data not shown). No significant 505-nm absorbance or fluorescence changes were induced in the absence of ascorbate, demonstrating yet again that ascorbate is required for de-epoxidation and xanthophyll-related non-radiative energy dissipation. To further investigate the peroxide effect on violaxanthin de-epoxidation and photochemical fluorescence quenching, the data of Fig. 2 were replotted on a semi-log scale as first-order kinetics of the 505-nm absorbance change (Fig. 3A) and on a linear scale for the photochemical 50 "o x o

10

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fluorescence quenching (Fig. 3B). The reaction conditions were the same as in condition 1, Table 1 that induced only a small difference in the rates of the two phases. The de-epoxidation rates were 0.260min -1 and 0.230min -1 for the initial and final rates, respectively. Addition of 0.5 mM hydrogen peroxide totally inhibited deepoxidase activity for about two minutes after which activity gradually returned to the control rate. H 2 0 2 addition also strongly increased photochemical quenching coincident with inhibition of de-epoxidation. H 2 0 2 at 10/~M had similar effects on AAs05 and qp as did 0.5 mM except for a shorter duration of the transient (data not shown). These effects are consistent with stimulation of linear electron flow mediated by ascorbate peroxidase-dependent H 2 0 2 reduction (Asada and Takahashi 1987, Neubauer and Schreiber 1989). The relationship between ascorbate peroxidase and violaxanthin de-epoxidase activity during the transient period in Fig. 2 was compared by plotting the first-order rate constant (k) for deepoxidation against qp (Fig. 4). The de-epoxidation rate constants for the transition period were estimated from slopes of tangent lines applied to the curve. The peroxide-induced qp correlated directly with the decrease in violaxanthin deepoxidase activity down to low qp. This implies that ascorbate consumption by ascorbate peroxidase-dependent H 2 0 2 reduction may affect violaxanthin de-epoxidation even at hydrogen

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Fig. 3. Effect of hydrogen peroxide on (A) the 505 nm absorbance change plotted as first-order kinetics and on (B) photochemical fluorescence quenching, qr plotted on a linear scale. The H~O2-data were derived from the experiment shown in Fig. 2. (A) The open triangles show the 505 nm absorbance change in the presence of 30 mM ascorbate, incubated for 10 min, and the closed circles effect of peroxide addition on violaxanthin de-epoxidation, under the same conditions. (B) The open triangles and the closed circles show the qp changes for the same conditions as in (A).

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Fig. 4. The correspondence of qr and the first-order rate constants (k) for de-epoxidation in the presence of H 2 0 2 and light. The k values were estimated as the slopes of tangent lines applied to the first-order rate plot in Fig. 3A. The first-order rate constants are plotted against peroxide-induced qp from Fig. 3B for equivalent times.

143 peroxide concentrations below 10/xM (see above). The transient inhibition of de-epoxidase activity by H20 2 was not due to a direct and reversible effect of H20 2 because 1 mM H20 2 did not inhibit the activity of the isolated enzyme (data not shown). Table 2 compares the effects of H20 2 on deepoxidation under light and dark-induced conditions. In light, H202 totally inhibited violaxanthin de-epoxidation in the presence of 30 mM but not 120 mM ascorbate (conditions 1 and 2). For intact chlorplasts the effect of H20 2 under dark conditions at pH5.2 was 20% less than under light conditions (conditions 5 and 7). In broken chloroplasts and in the dark, inhibition was only 25% and 16% in the presence of 30 and 120 mM ascorbate, respectively (conditions 8-9). Independent of chloroplast intactness, inhibition of de-epoxidation induced by H20: was transient when added in the light or in the dark at pH 5.2 (not shown). The generally lower peroxide effect at pH 5.2, especially in broken chloroplasts, can be explained by decreased ascorbate peroxidase activity in broken chloroplasts due to enzyme dilution (Miyake and Asada 1992) and the suboptimal reaction pH for ascorbate peroxidase (Asada and Takahashi 1987). Table 2 also shows the effects of KCN on

light-induced de-epoxidase activity and the inhibitory effect of H20 2 addition. KCN is an effective inhibitor of ascorbate peroxidase (Nakano and Asada 1981) but has no direct effect on violaxanthin de-epoxidase in intact chloroplasts (compare conditions 5 and 6, Table 2) or of the isolated enzyme (data not shown). Its influence on the peroxide effect can be mostly, if not entirely, attributed to inhibition of ascorbate peroxidase activity. KCN at 1 mM inhibits violaxanthin de-epoxidation indirectly by inhibiting ascorbate peroxidase and thus the Mehler-peroxidase-mediated ApH (Neubauer and Yamamoto 1992). Because KCN can affect ApH, this inhibitor cannot be used to test possible competition between ascorbate peroxidase and violaxanthin de-epoxidase. Low KCN concentrations (30 and 50/~M) partly inhibited de-epoxidase activity in the controls but also partly reduced the inhibitory effect of H20 2 on de-epoxidation (compare Table 1, condition 1 and Table 2, conditions 3-4). This contrasting effect can be interpreted in terms of the previously mentioned inhibition of Mehler-peroxidase-dependent ApH in the control and the reduced consumption of ascorbate by ascorbate peroxidase in the H20 2treated sample. Under dark-induced de-epoxidation at pH5.2, inhibiting ascorbate peroxidase

Table 2. Effects of H20 2 and KCN on de-epoxidation rate. Hydrogen peroxide (0.5 mM) was added when about 50% of the maximal light-induced 505 nm absorbance change had occurred. Rate constants (k) of de-epoxidation were measured for 2 min after adding HzO 2. Control values were measured at equivalent times. Chloroplasts were pre-incubated for 40 min with ascorbate for conditions 1 and 2. In the KCN reactions (conditions 3 and 4), ascorbate (30 mM) was added 10 min before onset of actinic light. The dark reaction in the presence of i mM KCN (condition 6) was induced upon addition of 30 mM ascorbate. The data are values for 3 (conditions 2, 5, 7 and 8) and 8 (condition 1) measurements of different chloroplast batches. For n = 2 (conditions 3, 4 and 6), the measured k values are given. For other conditions see Table 1 Condition

Class

k (min 1)

% Inhibition

Control

plus H20 z

A A A A

0.394 --- 0.069 1.156 +- 0.091 0.279; 0.320 0.220; 0.348

0.000 -+ 0.000 0.328 +- 0.110 0.023; 0.045 0.066; 0.089

i00 71 91; 86 70; 74

A A

0.298 - 0.049 0.304; 0.276

0.054 -+ 0.010 0.304; 0.276

81 0; 0

A D D

0.307 +- 0.081 0.314 -+ 0.064 0.327 ---0.073

0.122 -+ 0.081 0.236 +- 0.064 0.275 -+ 0.069

60 25 16

Light 1) 30 mM ASC 2) 120 mM ASC 3) 30 tzM KCN 4) 50 ~ M KCN

Dark, pH 5.2 5) 30 mM ASC 6) 30 mM ASC + 1 mM KCN

KCN 7) 120 mM ASC 8) 30 mM ASC 9) 120 mM ASC

144 totally with 1 mM KCN completely suppressed the effect of added H202 on violaxanthin deepoxidation (condition 6). In this case inhibition of ascorbate peroxidase obviously could not influence the required pH but did prevent potential competition for ascorbate by Mehler-peroxidase activity. The affinities of ascorbate peroxidase and violaxan~in de-epoxidase for ascorbate were evaluated by the effects of ascorbate concentrations on light-induced H202-dependent qp (Fig. 5) and dark-induced de-epoxidation (Fig. 6). Figure 5A shows that at 0.1 mM ascorbate, ascorbate-peroxidase induced qp was 80% maximal. Even without externally added ascorbate, peroxide-induced qp was about 12% of that present at 30mM ascorbate. This stimulation was possibly due to the internal pool of ascor-

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Fig. 6. The effects of ascorbate concentrations on darkinduced de-epoxidase at pH5.2 (A) and the corresponding Eadie-Hofstee plot (B). Violaxanthin de-epoxidase activity was initiated by adding ascorbate to the intact chloroplasts. The activity was calculated from the maximal rate of the 505 nm absorbance change, using the difference extinction coefficient for de-epoxidation of 63 mM-1 cm-1 (Yamamoto 1985). The enzyme unit (U) for violaxanthin de-epoxidase activity is defined as 1/~mol violaxanthin de-epoxidized min-1. The KM calculated from the slope of the Eadie-Hofstee plot is 3.1 mM ascorbate.

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qp

Fig. 5. The effects of ascorbate concentrations on ascorbate peroxidase activity in intact chloroplasts (A) and the corresponding Eadie-Hofstee plot (B). Ascorbate peroxidase activity was measured as qa, induced on addition of 0.5 mM H202, 2 min after onset of actinic red light (90/.~M m -2 s -1) and in the presence of 0.5/xM nigericin. The choroplasts were incubated for 5 min with ascorbate before illumimnation to allow equilibration between stroma and external medium. The K M calculated from the slope of the EadieHofstee plot is 0.36 mM ascorbate.

bate. Figure 5B shows the Eadie-Hofstee plot of the ascorbate-dependent qp in the presence of 0.5 mM H 2 0 2. The K M was 0.36 mM ascorbate, in agreement with values reported by Groden and Beck (1979) and Chen and Asada (1989). The K M was independent of light intensity but decreased to 0.24mM ascorbate when 50/xM H 2 0 2 was used (data not shown). Figure 6A shows the effect of ascorbate concentration on de-epoxidase activity. At 3 mM ascorbate activity was 50% and at 0.1 mM almost no activity was observed. The K M from Fig. 6B was 3.1 mM ascorbate for violaxanthin de-epoxidase activity. Purified violaxanthin de-epoxidase has a K M for ascorbate of 6.5 mM (Rockholm and Yamamoto, personal communication).

145 Discussion

We recently showed that the ascorbate-dependent Mehler-peroxidase reaction can mediate the photosynthetic proton pump sufficiently to induce de-epoxidation and non-photochemical fluorescence quenching (Neubauer and Yamamoto 1992). The electron flow induced by the Mehlerperoxidase reaction produces no net change in molecular oxygen (Asada and Badger 1984) and therefore the electron transport is pseudocyclic. In isolated chloroplasts, this system explains the light-induced de-epoxidation and non-photochemical fluorescence quenching that takes place in the absence of CO 2 fixation or added electron acceptors (Neubauer and Yamamoto 1992). Ascorbate therefore has at least two functions in xanthophyll-related fluorescence quenching; it is directly required for de-epoxidase activity and supports, through the Mehler-peroxidase reaction, acidification of the lumen. The biphasic property of de-epoxidation introduces yet another confounding effect of ascorbate. The fact that the initial phase is affected by both asorbate preincubation time and concentration (Table 1, conditions 1-4), and is less apparent in broken chloroplasts, strongly suggest that diffusion pressure of ascorbate through membrane barriers (mainly the envelope membrane) influence this phase. De-epoxidation activity during the slow second phase was strongly influenced by light and membrane intactness, being slowest in the presence of light and in intact chloroplasts. This light effect appears to be through induction of Mehler-peroxidase reaction which effectively serves as an ascorbate sink. Ascorbate peroxidase is partly membrane bound (Miyake and Asada 1992). This strategic location may cause a greater decrease of ascorbate concentration near the membrane-bulk interface and a corresponding greater effect than expected from bulk-phase changes. Hydrogen peroxide-addition stimulated consumption of ascorbate by ascorbate peroxidase activity (even with 10/~M H202) and resulted in total inhibition of de-epoxidation presumably until H20 2 was completely depleted (Fig. 2). In chloroplasts, H20 2 and consequently, monodehydroascorbate, are produced at rates of 120/zM s -1 and 240/xM s -x , respectively (Asada

and Takahashi 1987, Asada 1992). The reduction of monodehydroascorbate by PSI and the monodehydroascorbate reductase is very effective (Asada and Takahashi 1987, Asada 1992). A substantial hpH is generated (Neubauer and Yamamoto 1992) even though the ascorbate pool is maintained largely in the reduced form (90 to 95%) during the course of light transitions and high light treatment (Foyer et al. 1989). However, terminal violaxanthin de-epoxidation was significantly lower under light conditions compared to dark reactions at pH 5.2 (Table 1). We assume that de-epoxidase activity does not operate maximally relative to the apparent stromal ascorbate concentration when hpH is formed solely by Mehler-peroxidase-dependent electron transport. Although not yet shown directly, the in vitro evidence suggests the Mehler-peroxidase-reaction may influence de-epoxidation in vivo. Varying half times in the range of 1.5 to 10 min were observed consistently for in vivo deepoxidation (Demmig-Adams 1990). The Oz-dependent electron flow can proceed at significant rates even under maximal CO 2 fixation (Robinson 1988) provided ascorbate peroxidase is not inactivated by the presence of DTF (Gerbling et al. 1984, Chen and Asada 1989) or peroxide is not decomposed by the presence of catalase. Due to the strategic location of the membranebound ascorbate-peroxidase (Miyake and Asada 1992) the local ascorbate concentration might decrease significantly to affect ascorbate availability for violaxanthin de-epoxidation without a large change in bulk-phase ascorbate concentration. Foyer et al. (1990) proposed that energy consumption by PS II is regulated at least in part by ascorbate which crosses the thylakoid membrane after its reduction in the stroma by PSI activity. We propose that this putative regulation by lumenal ascorbate is due to its stimulation of de-epoxidation. If lumen acidity is derived from the Mehler-peroxidase reaction, ascorbate concentration on both sides of the thylakoid is important for regulation. Ascorbate on the stroma side affects the Mehler-peroxidase reaction due to its need for H20 z reduction by ascorbate peroxidase and ascorbate on the lumen side stimulates de-epoxidation. The result is an acidic lumen, zeaxanthin and antheraxanthin

146 formation and decrease in the intrinsic quantum yield of Photosystem II by non-radiative dissipation of energy. Although violaxanthin de-epoxidation occurs diurnally in plants that are under no apparent environmental stress, high zeaxanthin levels are induced in plants under various stresses (Demmig-Adams and Adams 1992a). Under high light and extended stress conditions, where a decrease in Photosystem II quantum yield would be desirable, stress-induced increase of the total ascotbate pool (Gillham and Dodge 1987, Sch6ner et al. 1989) would favor induction of rapid deepoxidation and the related non-radiative energy dissipation. Under this condition, the lumen could become loaded with sufficient ascorbate in the dark to achieve complete de-epoxidation without interference from the Mehler-peroxidase reaction or membrane diffusion barriers. There is evidence that following extended stress conditions, zeaxanthin and antheraxanthin may remain in leaves even overnight (Demmig et al. 1988 Demmig-Adams and Adams 1992b). Because of the residual de-epoxidized pigments, competition for available ascorbate would not have as great an effect on subsequent light treatments. Leaves with higher dark levels of zeaxanthin and antheraxanthin would be better poised for photoprotection by non-radiative energy dissipation. Overall, these results show that ascorbate plays a pivotal role for modulating Photosystem II down-regulation by its linkages to violaxanthin de-epoxidase activity and the Mehler-peroxidase-mediated ApH.

Acknowledgements This work was supported by the Deutsche Forschungsgemeinschaft (Research Fellowship (C.N.), SFB 176 and 251) and by a US Department of Agriculture National Research Initiative Competitive Research Grant 90-37280-5594. We wish to thank David Rockholm for providing the isolated violaxanthin de-epoxidase and Narendranath Mohanty for critical comments.

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