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In ‘Beijing 24’ peach [Prunus persica (L.) Batch] trees, a series of source leaves with differing levels of end products were created by retaining fruit (“+fruit”), removing fruit (“−fruit”), or reducing the light period. To alter the light period, leaves were covered with a bag made of brown inner paper and outer silver paper, which was then removed at different times the next day. The highest level of end products were obtained by fruit removal, while reducing the light period resulted in a lower level than “+fruit.” Net photosynthetic rate (Pn) and stomatal conductance (gs) decreased, but leaf temperatures (Tleaf) increased, following an increase in end product levels in leaves. After the “−fruit” treatment, reduced Pn was correlated with lower gs, and Tleaf increase was concomitant with decreases in maximal quantum yield of photosystem II (Fv/Fm), actual photochemical efficiency of photosystem II (ΦPSII), and photochemical quenching, and with an increase in nonphotochemical quenching. However, there were no significant differences in chlorophyll fluorescence between “+fruit” and the two treatments reducing the light period. The ΦPSII decreased following an increase in foliar sorbitol level, and it linearly decreased as sucrose and starch increased. Although fruit removal resulted in a significant accumulation of sucrose, sorbitol, and starch in leaves throughout the day, the extractable activities of several important enzymes involved in carbohydrate leaf storage and translocation did not decrease. Therefore, instead of feedback regulation by the accumulation of end products in source leaves, a high Tleaf induced by decreased stomatal aperture may play a key role in regulation of photosynthesis by limiting the photochemical efficiency of the PSII reaction centers under high levels of the end products in peach leaves.
Crop yield and fruit quality in fruit trees are highly dependent on efficient capture of solar energy and subsequent allocation of photoassimilate. Source-sink relationships are important factors influencing these allocation patterns. Fruit constitute the main sink for photoassimilates during the fruit growth period. Removing or retaining fruit has often been used in studies of plant source-sink relationships (Syvertsen et al., 2003; Vaast et al., 2005). Reduced photosynthetic rates under low sink demand have often resulted in end product accumulation. This accumulation of end products has been often cited as the reason for the decline of Pn (Iglesias et al., 2002; Paul and Foyer, 2001; Wu et al., 2008; Zhou and Quebedeaux, 2003). This influence of end product accumulation has been considered a direct feedback mechanism regulating photosynthesis (Paul and Pellny, 2003). However, this conclusion is controversial (Eliezer and Huber, 1992; Stitt, 1991). For example, photosynthesis was not influenced in some species, although there was accumulation of end products in source leaves in response to low sink demand (Lunn and Hatch, 1995; Nautiyal et al., 1999; Roper et al., 1988). In addition, some recent studies with peach (Li et al., 2007) and coffee (Coffea arabica L.) trees (DaMatta et al., 2008) showed that low sink demand created by removing fruit reduced Pn with accumulation of the end products, but there was no influence on in vitro activity of key enzymes in the biosynthetic pathways of the end products. However, there have been few studies on the influence of end product accumulation on photosynthetic efficiency and activities of related enzymes.
Source-sink relationships can be also modified by reducing or prolonging the light period. Reducing the light period decreased end product amount in source leaves, and this was similar to the effect of increasing sink demand (Li et al., 2007). By removing fruit and reducing the light period, respectively, it may be possible to create a range of source leaves with differing end product levels. This would be a valuable method for investigating the possible feedback effect of end product levels on photosynthetic efficiency and the relationship between the end products and activities of related enzymes.
A positive linear relationship was observed between stomatal conductance (gs) and Pn in previous studies, showing that stomatal closure was an important factor in the decline in photosynthesis with low sink demand (Cheng et al., 2008; Li et al., 2001). Increased Tleaf following a reduction in stomatal aperture has been proposed as a critical factor in regulating photosynthesis under low sink demand in fruit trees (Li et al., 2005). These changes in Tleaf have also been related to reduced Fv/Fm (Jifon and Syvertsen, 2003). This article presents the results of studies on the short-term effect of end product accumulation on peach leaf Pn, focusing on ΦPSII and the activities of related enzymes. Also, the responses of Pn, gs, and ΦPSII to end product levels in leaves were analyzed.
The experiment was conducted in Pinggu District, Beijing, China, during the 2006 growing season. The peach cultivar used was Beijing 24, which matures during early September. Trees, grafted on wild peach rootstocks, were planted 2 m apart within rows and 5 m apart between rows in Spring 1998. They were trained to a “Y” training system and were pruned by the long pruning method in winter (Li et al., 1994).
During the final stage of rapid fruit growth (on 22 Aug. 2006), 1-year-old shoots located on the southwest and southeast sides of 15 to 20 trees in the outer part of the canopy were used as the units for source-sink manipulation. These previous-season shoots were 1.4 to 1.7 m aboveground, and were selected according to their uniformity in length (40–50 cm) and growth status (at least one new shoot > 30 cm).
Each 1-year-old shoot, supporting one fruit and one new shoot of 10 fully developed leaves retained by topping and removal of smaller basal leaves, was considered a plot. The export of assimilates was strictly controlled by girdling the base and top parts of the previous-season shoots to control the exchange of assimilates between treated and nonexperimental parts of the tree.
Four treatments were applied in this study: 1) “+fruit” treatment (control), 2) “−fruit” treatment, 3) a shoot supporting a fruit, covered with a bag made of brown inner paper and outer silver paper in the late afternoon of 22 Aug. with the bags removed at 0930 hr (“0930 hr bag removal”) on 23 Aug., and 4) a shoot supporting a fruit, covered with the same bag as used in the previous treatment, but with the bags removed at 1130 hr (“1130 hr bag removal”) on 23 Aug. A total of five plots for each treatment was used for the measurements of photosynthetic parameters and chlorophyll fluorescence, and 15 to 20 plots were used for the measurements of sugar content and enzyme activities.
Photosynthetic gas exchange and chlorophyll fluorescence response of five leaves per treatment (one leaf per replicate) were measured between 0700 and 1700 hr on an hourly basis on 23 Aug. 2006, the day after initiating source-sink manipulation. For the “0930 hr bag removal” and “1130 hr bag removal” treatments, measurements were made 30 min after bag removal to allow for light adaptation by the leaves.
Pn was measured with a portable photosynthesis system (LI-6400; LI-COR, Lincoln, NE) and photosynthetically active radiation (PAR), gs, transpiration rate (E), intercellular carbon dioxide concentration (Ci), and Tleaf were obtained simultaneously when Pn was measured.
Chlorophyll fluorescence parameters were measured with a plant efficiency analyzer [PEA (Hansatech, Norfolk, UK)] and a fluorescence monitoring system (FMS-2; Hansatech). After a 30-min dark adaptation period, minimum fluorescence (F0), maximum fluorescence (Fm), the steady-state fluorescence (Fs), and Fv/Fm, were measured by the PEA. The PEA was also used to estimate the predawn value of Fm. The FMS-2 was used to determine the maximum fluorescence level in the light-adapted state (Fm′), the minimal fluorescence level in the light-adapted state (F0′), and the efficiency of excitation capture by open PSII reaction centers (Fv′/Fm′).
The following calculations were made: (1) photochemical quenching (qP) = (Fm′ − Fs)/(Fm′ – F0′), and nonphotochemical quenching (NPQ) = (Fm/Fm′) – 1; (2) ΦPSII = (Fm′ − Fs)/Fm′ (Genty et al., 1989).
Leaves from five plots of each treatment were harvested every 2 h between 0700 and 1700 hr (for the bagging treatments, leaves were sampled starting at 1000 or 1200 hr, respectively), and the midrib of each leaf was removed. One half of the leaf was used for carbohydrate analysis and the other half was used for enzyme extraction and biochemical analyses. Leaf samples were frozen immediately in liquid nitrogen and were kept at −80 °C until analysis.
For soluble carbohydrate extraction, about 1.5 g of fresh weight of leaves was ground to a fine powder in liquid nitrogen using a mortar and pestle. The powder was extracted three consecutive times in 10 mL of 80% ethanol in a water bath at 80 °C for 10 min each time, and solutions were centrifuged (Sigma Laborzentrifugen, Osterode, Germany) at 19,118 gn for 10 min after each extraction. The combined ethanol supernatants for each sample were evaporated to dryness in a water bath at 85 °C, 1.5 mL of double distilled water was added, and samples were placed in a rotary shaker (THZ-C-1; Taicang Instrument, Jiangsu, China) for 1 h (4 °C). The solution was then centrifuged at 19,118 gn for 10 min, the supernatant was passed though a SEP-C18 cartridge (Supelclean ENVI C18 SPE; Supelco, Bellefonte, PA) and filtered through a 0.22-μm filter, and the filtered solution was taken for the measurement of soluble carbohydrates.
Sorbitol, glucose, fructose, and sucrose were separated by high-performance liquid chromatography [HPLC (Dionex, Sunnyvale, CA)], and they were detected using a Shodex RI-101 refractive index detector with reference cell maintained at 40 °C (Dionex). A Transgenomic CARB Sep Coregel 87C column (300 × 7.8 mm i.d., 10 μm particle size; Dionex) with a guard column cartridge was used. The column was maintained at 85 °C with a TCC-100 thermostatted column compartment (Dionex). Degassed, distilled, deionized water was used at a flow-rate of 0.6 mL·min−1 as the mobile phase. The injection volume was 10 μL. The Chromeleon chromatography data system (Dionex) was used to integrate peak areas according to external standard solution calibrations (sugar reagents were purchased from Sigma-Aldrich, St. Louis). Starch remaining in the tissue was extracted and measured according to Li and Li (2005).
The enzyme extracts were obtained as follows: 1.0 g of frozen leaf tissue was ground to a fine powder in liquid nitrogen with quartz sand. Extraction buffer was added to the ground sample, and the slurry was ground to mix thoroughly. The extraction buffer contained 50 mmol·L−1 N-2-hydroxyethylpiperazine-N-ethane-sulphonicacid [Hepes-NaOH (pH 7.5, Sigma-Aldrich)], 10 mmol·L−1 magnesium chloride hexahydrate [MgCl2 (Sigma-Aldrich)], 1.0 mmol·L−1 disodium ethylenediamine tetraacetate [EDTANa2 (Sigma-Aldrich)], 2.5 mmol·L−1 dithiothreitol [DTT (Sigma-Aldrich)], 0.05% t-octylphenoxypolyethoxyethanol (Triton X-100, Sigma-Aldrich), and 0.1% bovine serum albumin [BSA (Sigma-Aldrich)]. The homogenate was filtered through four layers of gauze, and the filtrate was centrifuged at 13,000 gn for 10 min at 2 °C. The supernatant was dialyzed immediately by a 10-fold volume of diluted extraction buffer (minus Triton X-100) for 20 h at 4 °C, and the dialyzate was changed one time. In these analyses, leaves were not allowed to thaw before grinding in extraction buffer, and particular care was taken to work rapidly once the material had been taken up in buffer at 4 °C. The enzyme extracts after dialysis were used for all enzyme assays.
Aldose-6-phosphate reductase [A6PR (EC 1.1.1.200)] activity was assayed in the direction of synthesis following the oxidation of nicotinamide adenine dinucleotide phosphate [NADPH (reduced form, Sigma-Aldrich)] in the presence of glucose-6-P at 340 nm, as described by Merlo and Passera (1991). The reaction mixture contained 0.1 mol·L−1 Tris (hydroxymethyl) aminomethane [Tris-HCl (pH 8.8, Sigma-Aldrich)], 0.1 mmol·L−1 NADPH, 50 mmol·L−1 glucose-6-P, and 50 μL of extract. One unit of A6PR is defined as the amount of enzyme catalyzing the oxidation of 1.0 μmol·min−1 NADPH at 25 °C under standard assay conditions. The activity of sorbitol dehydrogenase [SDH (EC 1.1.1.14)] was assayed by following the reduction of nicotinamide adenine dinucleotide [NAD+ (Sigma-Aldrich)] in the presence of D-sorbitol at 340 nm, according to the method described by Negm and Loescher (1979). The reaction mixture contained 0.1 mol·L−1 Tris-HCl (pH 9.0), 1.0 mmol·L−1 NAD+, 0.5 mol·L−1 D-sorbitol, and 0.1 mL of extract. Sorbitol and glucose-6-phosphate [G-6-P (Sigma-Aldrich)], dissolved in the same buffer as used in the assay, were used to start the reactions.
Sucrose-phosphate synthase [SPS (EC 2.4.1.14)] activity was determined by measuring the conversion rate of fructose-6-phosphate [F-6-P (Sigma-Aldrich)] to sucrose-phosphate (S-6-P; Sigma-Aldrich), according to the method described by Huber and Israel (1982) with some changes. The assay mixtures (140 μL) contained 50 mmol·L−1 Hepes-NaOH (pH 7.5), 10 mmol·L−1 MgCl2, 5 mmol·L−1 sodium fluoride (NaF), 25 mmol·L−1 uridine 5′-diphosphoglucose disodium salt [UDPG (Sigma-Aldrich)], 10 mmol·L−1 F-6-P, and 80 μL of extract. The reaction was stopped after 40 min at 25 °C by adding 140 μL of 1.0 mol·L−1 NaOH. Unreacted F-6-P was destroyed by placing the tubes in boiling water for 10 min. After cooling, 0.5 mL of 0.1% resorcinol in 95% ethanol and 1.5 mL of 30% HCl were added, and the tubes were incubated at 80 °C for 8 min and A520 was measured.
ADP glucose-pyrophosphorylase [ADPGPPase (EC 2.7.7.27)] activity was determined by measuring pyrophosphate-dependent glucose-1-phosphate formation from adenosine 5′-diphosphoglucose disodium (ADPG; Sigma-Aldrich) at 25 °C in a total of 1 mL of solution, according to the procedures described by Rufty et al. (1983). The reaction was started by addition of ADPG to a final concentration of 5 mmol·L−1 and production of NADPH was monitored at 340 nm.
The activities of acid invertase [AI (EC 3.2.1.26)] and neutral invertase [NI (EC 3.2.1.26)] were assayed in a reaction mixture (1 mL) containing 0.1 mol·L−1 Na-acetate (pH 4.8), 0.1 mol·L−1 sucrose, and 0.2 mL of extract. After incubation of the mixture for 40 min at 37 °C, the reactions were stopped by addition of 1 mL of dinitrosalicylic acid reagent (DNS; Beijing Chemical Reagent Corporation, Beijing, China), and reducing sugars released from sucrose were determined according to the method of Li and Li (2005). Total amylase activity [(EC 3.1.1.1) and (EC 3.1.1.2)] was assayed in a reaction mixture (1 mL) containing 0.1 mol·L−1 Na-acetate (pH 6.5), 1.5 mmol·L−1 NaF, 5 mmol·L−1 Ca (NO3)2, 0.5% soluble starch, and 0.2 mL of extract. After incubation at 30 °C for 40 min, the reaction was ended, and the release of reducing groups was determined as in the invertase assay. In invertase and amylase assays, blanks containing reaction mixtures were incubated with DNS.
A completely randomized design was used with five replicates for each treatment. The data were subjected to analysis of variance (ANOVA), and means were compared by Duncan's new multiple range test or t test (when there were only two groups) using SPSS (version 13.0 for Windows; SPSS, Chicago). Each value of the mean and standard error in the figures represents five replicates of each treatment. Unless otherwise indicated, significant differences among means are given at P < 0.05.
Fruit removal significantly reduced Pn from 1000 to 1500 hr and gs from 9000 to 1600 hr, respectively, compared with the “+ fruit” and after removing the bags covering leaves (Fig. 1, A and B). Moreover, the two “bag removal” treatments resulted in significantly higher Pn and gs from 1300 to 1500 hr than “+ fruit.” A significantly lower E was observed at 0900 hr and 1200 to 1600 hr in the “−fruit” treatment than the other three treatments. However, there were no significant differences in E between “+ fruit” and the two “bag removal” treatments (Fig. 1C).
Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 134, 6; 10.21273/JASHS.134.6.667
Opposite to the effect on Pn, gs and E, fruit removal significantly increased Tleaf from 1100 to 1500 hr in general compared with the other three treatments. Moreover, a lower Tleaf was observed for “bag removal” treatments at 1100, 1300, and 1500 hr than “+ fruit” (Fig. 1D).
Ci changed slowly from 0700 to 1300 hr except for the “−fruit” treatment. For the “−fruit” treatment, the highest Ci appeared at 1400 hr and then decreased gradually. However, no significant differences in Ci were found among the treatments until 1400 hr. The Ci of the “−fruit” treatment was significantly higher than those of other treatments at 1400 hr, but was lower than other treatments at 1600 and 1700 hr (Fig. 1E).
The “−fruit” treatment significantly decreased Fv/Fm (Fig. 2A) from 1100 or 1200 hr to the end of the day, ΦPSII (Fig. 2B) at 0900, 1100, and 1400 to 1600 hr, qP (Fig. 2C) at 1000 to 1400 hr, and Fv′ /Fm′ at 1400 hr (Fig. 2D), but significantly increased F0 (Fig. 2E) from 1100 hr to the end of the day and NPQ (Fig. 2F) at 1300 to 1700 hr compared with the other three treatments. However, except for ΦPSII and qP, the other chlorophyll fluorescence parameters showed no response in general to “+fruit” and the two “bag removal” treatments. ΦPSII and qP of the “+fruit” treatment were significantly higher than that of the “930 bag removal” treatment at 1100 hr and 1000 to 1100 hr, respectively (Fig. 2, E and F).
Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 134, 6; 10.21273/JASHS.134.6.667
Fruit removal and “bag removal” treatments altered the content of soluble sugars and starch in leaves (Fig. 3). The “−fruit” generally resulted in the highest foliar content of soluble sugars and starch among the treatments, and the contents of soluble sugars and starch in “+fruit” leaves were higher than those in the two ‘bag removal’ treatments. Moreover, the later time of bag removal lowered the content of soluble sugars and starch more than the earlier removal time. The differences in sorbitol and starch content among the treatments remained unchanged throughout the day generally (Fig. 3, A and C). As regards sucrose, fructose and glucose content in the leaves, the magnitude of the differences among treatments sharply decreased after 1300 or 1500 hr (Fig. 3, B, D, and E).
Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 134, 6; 10.21273/JASHS.134.6.667
Compared with “+fruit”, “−fruit” significantly increased A6PR and SDH activities at 1100 hr and from 0900 to 1100 hr, respectively (Fig. 4). However, “bag removal” treatments did not alter A6PR and SDH activities. As regards the activities of the other related enzymes of carbohydrate metabolism, no obvious effect of the “−fruit” and “bag removal” treatments were observed (data not shown).
Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 134, 6; 10.21273/JASHS.134.6.667
Pn increased with increased Tleaf until about 30 °C, remained high between 30 and 34 °C, and then sharply decreased when Tleaf continued to increase (Fig. 5A). Moreover, when Tleaf was greater than a critical temperature of about 34 °C, Pn was significantly correlated with gs (r = 0.79***) (Fig. 5B).
Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 134, 6; 10.21273/JASHS.134.6.667
There were significant relationships between Pn and foliar amounts of end products including sorbitol, sucrose, and starch (Fig. 6). Pn decreased following an increase in sorbitol, sucrose, and starch content, indicating that the Pn change was somehow affected by their accumulation. The accumulation of the end products was also correlated to gs (Fig. 7). A high content of sorbitol, sucrose, and starch in leaves was related to lower gs, implied that high content of end products changed gs, presumably by altering osmotic pressure in gourd cell or signal transduction (Fig. 7, A–C), and then lower gs may have resulted in Pn decrease. However, significant relationships were only found between gs and foliar content in starch.
Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 134, 6; 10.21273/JASHS.134.6.667
Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 134, 6; 10.21273/JASHS.134.6.667
The response of ΦPSII to end product amount is shown in Fig. 8. When foliar sorbitol content increased, ΦPSII decreased (Fig. 8A). ΦPSII also decreased linearly with the sucrose increase (Fig. 8B). However, a relationship between ΦPSII and starch content was not apparent (Fig. 8C).
Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 134, 6; 10.21273/JASHS.134.6.667
Opposite to ΦPSII, NPQ increased following an increase in foliar sorbitol content (Fig. 9A), and NPQ was significantly correlated with sucrose and starch content (Fig. 9, B and C).
Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 134, 6; 10.21273/JASHS.134.6.667
Sorbitol, sucrose, and starch are the end products of photosynthesis in many Rosaceae species (Loescher, 1987). In this study, fruit removal resulted in accumulation of end products in leaves, while reducing the light period (“bag removal” treatments) decreased end product levels (Fig. 3). The short-term effects of weakening sink strength by fruit removal in the present study significantly decreased Pn. By contrast, removal of the bags covering the leaves had an effect similar to increasing sink strength, resulting in increased Pn (Fig. 1A). Those effects of varying sink demand on Pn, gs, and Tleaf were similar to those reported in peach trees by Li et al. (2007) and Wu et al. (2008).
Pn was related to the foliar amounts of the three end products in this study, decreasing as foliar end product levels increased (Fig. 6). Accumulation of end products in leaves is usually cited as the principal reason for lower photosynthesis under reduced sink demand, as a direct feedback mechanism (Iglesias et al., 2002; Layne and Flore, 1995). Consistent with this hypothesis, photosynthesis and the activity of related key biosynthetic enzymes should be depressed by end product accumulation with low sink demand (“−fruit”) or enhanced when end products are low (“bag removal”). However, the present results indicate that key extractable enzyme activities did not change correspondingly. These enzyme activities did not decrease, and A6PR and SDH even increased at certain times during the day with low sink demand and high end product content (Fig. 4). In prior studies of peach (Li et al., 2007) and coffee (DaMatta et al., 2008), differences in Pn also seemed unlikely to have been caused by photochemical impairments or a direct end product-mediated feedback down-regulation of photosynthesis. Given that in vitro enzyme activities were not influenced by foliar end product content, the modified photosynthetic rates created by source-sink manipulation may therefore be regulated by a mechanism other than direct feedback effects on related biosynthetic enzymes.
There was a positive linear relationship between gs and Pn in peach in previous studies (DeJong, 1986; Li et al., 2005, 2007; Tan and Buttery, 1986), as in the present study (Fig. 5B). Li et al. (2001) suggested that stomatal aperture may be the trigger or promoter and leaf temperature the critical factor regulating photosynthesis when modifying source-sink relationships. In the present study, the different sink demands affected gs and E (Fig. 1, B and C). gs was negatively correlated with foliar end product content, especially that of starch (Fig. 7, A–C). Low gs decreases leaf transpiration and therefore increases leaf temperature, which would influence photosynthesis (Li et al., 2001; Li et al., 2005). In the “−fruit” treatment, the high Tleaf was above the optimal level for the photosynthesis (Fig. 5A). High Tleaf results in damage to the photosynthesis apparatus and cell structure by the generation and accumulation of reactive oxygen species such as hydrogen peroxide (H2O2) and malondialdehyde (MDA) (Duan et al., 2008). Damage to the photosynthetic apparatus and cell structure was confirmed by chlorophyll fluorescence results. Low sink demand by fruit removal increased F0 throughout the day when compared with the “+fruit” and “bag removal” treatments (Fig. 2E), indicating damage to the PSII reaction center complex. Fv/Fm represents electron transport efficiency and photochemical efficiency in the PSII reaction center. Layne and Flore (1995) reported that continuous lighting significantly reduced Fv and Fv/Fm within 1 d in sour cherry (Prunus cerasus L.). The decline in Fv/Fm observed in “−fruit” leaves (Fig. 2A) was presumably related to increased dissipation of excess energy.
Pn was significantly correlated with gs (Fig. 5B), and the latter was correlated with end product levels (Fig. 7), indicating that end product accumulation may affect Pn via altering gs. In addition, end product accumulation may also affect photochemical efficiency. ΦPSII decreased as end products increased (Fig. 8), while NPQ increased (Fig. 9). High Pn at low end product content could be explained by high ΦPSII and low NPQ. ΦPSII is the product of qP and Fv′/Fm′ in open PSII centers (Genty et al., 1989), so increased ΦPSII could be due to an increase in photochemical quenching and Fv′/Fm′. NPQ has usually been used as an indicator of the appearance in the photosynthetic apparatus of mechanisms to prevent overexcitation of reaction centers (Ivanov and Edwards, 2000). Leaves with lower end product levels had higher photosynthetic capacity, so they use a bigger fraction of the absorbed light in electron transport. As a result, there was less thermal dissipation of excitation energy.
In conclusion, accumulation of end products in leaves decreased Pn, gs, and E, but increased Tleaf, in peach leaves. However, high foliar end product content had no direct feedback effect on the activities of related enzymes of carbon metabolism in vitro. Thus, changes in Pn in response to accumulation of end products may result from a negative effect on photosynthetic efficiency. Stomatal aperture may start to close as the initial response and thus affect Pn. Accumulated soluble sugar and starch in leaves may change osmotic pressure of the guard cells, resulting in such a gs change. End products may also act as signals, with crosstalk with phytohormones such as abscisic acid (ABA) (Gibson, 2005; Lloyd and Zakhleniuk, 2004; Roitsch, 1999). End product accumulation may also affect photochemical efficiency, changing F0, Fv/Fm, and ΦPSII. However, it will be necessary to focus on the mechanism of end product alteration of stomatal movement for understanding the effect on Pn. Also, the possibility that accumulation of end products may down-regulate photosynthesis in peach leaves by deactivating some of the key enzymes in carbon fixation, such as Rubisco, should be studied in the future.
Contributor Notes
The research was supported by the National Natural Science Foundation of China (No. 30370986) and by the Science Foundation of the President of the Chinese Academy of Sciences. We thank Prof. Douglas D. Archbold, Univ. of Kentucky, and Prof. W. Loescher, Michigan State Univ. for critical review of the manuscript.
We thank Yanfeng Xing (Director, Fruit Office, Pinggu District, Beijing, China) for the use of the experimental orchard facilities.
Corresponding author. E-mail: shhli@wbgcas.cn
