Planta (2010) 232:523–531 DOI 10.1007/s00425-010-1193-y
ORIGINAL ARTICLE
Sinks for photosynthetic electron flow in green petioles and pedicels of Zantedeschia aethiopica: evidence for innately high photorespiration and cyclic electron flow rates Charilaos Yiotis • Yiannis Manetas
Received: 26 March 2010 / Accepted: 6 May 2010 / Published online: 21 May 2010 Ó Springer-Verlag 2010
Abstract A combination of gas exchange and various chlorophyll fluorescence measurements under varying O2 and CO2 partial pressures were used to characterize photosynthesis in green, stomata-bearing petioles of Zantedeschia aethiopica (calla lily) while corresponding leaves served as controls. Compared to leaves, petioles displayed considerably lower CO2 assimilation rates, limited by both stomatal and mesophyll components. Further analysis of mesophyll limitations indicated lower carboxylating efficiencies and insufficient RuBP regeneration but almost similar rates of linear electron transport. Accordingly, higher oxygenation/carboxylation ratios were assumed for petioles and confirmed by experiments under non-photorespiratory conditions. Higher photorespiration rates in petioles were accompanied by higher cyclic electron flow around PSI, the latter being possibly linked to limitations in electron transport from intermediate electron carriers to end acceptors and low contents of PSI. Based on chlorophyll fluorescence methods, similar conclusions can be drawn for green pedicels, although gas exchange in these organs could not be applied due to their bulky size. Since our test plants were not subjected to stress we argue that higher photorespiration and cyclic electron flow rates are innate attributes of photosynthesis in stalks of calla lily. Active nitrogen metabolism may be inferred, while Electronic supplementary material The online version of this article (doi:10.1007/s00425-010-1193-y) contains supplementary material, which is available to authorized users. C. Yiotis (&) Y. Manetas (&) Laboratory of Plant Physiology, Department of Biology, University of Patras, 265-00 Patras, Greece e-mail:
[email protected] Y. Manetas e-mail:
[email protected]
increased cyclic electron flow may provide the additional ATP required for the enhanced photorespiratory activity in petiole and pedicel chloroplasts and/or the decarboxylation of malate ascending from roots. Keywords Stem photosynthesis CO2 assimilation PSI Photorespiration Cyclic electron flow
Introduction Although leaves are the main sites for efficient photosynthetic assimilation of CO2, functional chloroplasts also abound in many other organs having quite different primary functions (Aschan and Pfanz 2003). We may distinguish between those organs whose chloroplasts are restricted behind stomatal-less periderms (i.e., twigs) or pericarps with very few stomata and those possessing abundant stomata (i.e., green stems) (Nilsen 1995). In the first case, the high diffusive resistance to gas exchange, in conjunction with the high heterotrophic/autotrophic cell ratios, restores an internal aerial environment quite different from that of leaves. Thus, internal CO2 partial pressures may be extremely high (up to 20%), while O2 may fall to hypoxic levels (Pfanz et al. 2002; Borisjuk and Rolletschek 2009). Hence, photosynthesis in these bulky organs is believed to serve in the re-fixation of respiratory CO2 and/ or in the alleviation of the consequences of hypoxia (Pfanz et al. 2002; Borisjuk and Rolletschek 2009). Interest in the photosynthetic attributes of peridermal stems and fruits has been recently revived, in an attempt to elucidate whether partial reactions of photosynthesis are adjusted to the particular internal environment and the metabolic demands of these organs. Both the light reactions (Hetherington et al. 1998; Manetas 2004; Kotakis et al. 2006; Ivanov et al.
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2006; Filippou et al. 2007; Kalachanis and Manetas 2010) and biochemical reactions of photosynthesis (Xu et al. 1997; Alessio et al. 2005; Borisjuk et al. 2005; Ivanov et al. 2006; Wittmann et al. 2006; Berveiller and Damesin 2008) were investigated. The case of green stems has been mainly studied in xerophytic perennials growing in permanently or periodically dry habitats (Osmond et al. 1987; Comstock and Ehleringer 1990; Nilsen and Sharifi 1994; Yiotis et al. 2008). Since these plants are drought-deciduous with usually small and short-lived leaves, the aim of the corresponding investigations was to assess the contribution of stems to the plant’s total carbon budget. Leaf type photosynthesis assimilating atmospheric CO2 in chlorenchymatous cells similar to palisade and/or spongy mesophyll cells was found (Gibson 1983; Nilsen 1995; Yiotis et al. 2006). However, green, non-foliar organs are not a peculiarity of xerophytes, but the norm in most plant species. Detailed research in the photosynthetic properties of such organs has been neglected, although their contribution to the total photosynthetic area may be substantial and their carbon and energy metabolism may be particular. It has been reported, for example, that green petioles of tobacco and Arabidopsis may engage the decarboxylating part of C4 metabolism (Hibberd and Quick 2002; Brown et al. 2010) and use organic acids coming up with the transpiration stream (Ben Zioni et al. 1971) as an additional carbon source for their photosynthesis. We argue that such an activity would set specific metabolic demands and may require quantitative and/or qualitative adjustments in the photosynthetic attributes of such organs. To that aim, gas exchange and chlorophyll fluorescence methods both in light- and darkadapted material were used to assess possible limitations in photosystems’ function, electron flow and CO2 assimilation in the sizable petioles and pedicels of Zantedeschia aethiopica, using corresponding leaves as controls.
Materials and methods Plant material Zantedeschia aethiopica (L) Spreng. (Araceae) was grown from root cuttings obtained from a single, healthy plant. The resulting 15 individuals were grown in 15-l pots, containing a mixture of compost, clay and grains of pumice enriched with fair turf, in a natural-lit glasshouse and were amply irrigated. Mature plants were used after 6 months of growth. Gas exchange For CO2 assimilation rates, a gas analyzer (Li-6400, Li-cor, Lincoln, Ne, USA) equipped with the standard Li-6400 leaf
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chamber, a LED light source (6400-02B, with both red and blue LEDs) and a CO2-regulating device (6400-01 CO2 Injector System) was used. Only intact and attached to the plant leaves and petioles were measured, since the size of pedicels precluded credible measurements due to considerable gas leakage. In all cases, temperature within the chamber was kept at 22°C. The following protocol was applied. The plant material was enclosed in the chamber and illuminated at 800 lmol m-2 s-1 under 400 ppm CO2 up to full photosynthetic induction, as judged from three consecutive stable readings of CO2 assimilation (A) and stomatal conductance (gs) rates, usually within 30 min. A versus intercellular CO2 (Ci) concentrations curves were then assessed by adjusting the chamber atmospheric CO2 (CA) from 0 to 2,000 ppm in 11 ascending steps, each with 3 min duration. The system was then left to equilibrate again at 2,000 lmol m-2 s-1 and 400 ppm CO2 and light saturation curves of A were obtained by adjusting the light level from 2,000 to 0 lmol m-2 s-1, in 10 descending steps, each with 3 min duration. In all cases, the duration of the steps was long enough to obtain stable A readings. A versus Ci curves were used to calculate the carboxylation efficiency (CE, as the initial slope of the CO2limiting phase of the A vs. Ci curve) and the relative stomatal limitations as I ¼ ðA0 AÞ=A0 , where A0 and A are the CO2 assimilation rates obtained in the absence and presence of stomatal limitations, i.e., at Ci and CA equaling 400 ppm, respectively (Farquhar et al. 1980). The maximum quantum yield for CO2 assimilation was computed from the slope of the initial, light-limiting phase of the A versus photon flux curve. Modulated chlorophyll fluorescence in light-adapted plant material The material used for the CO2 assimilation measurements (i.e., leaves and petioles) plus pedicels was subsequently inserted into the leaf clip of a pulse-amplitude modulated fluorimeter (Mini-PAM, Walz, Effeltrich, Germany). The instrument provides a weak red LED measuring beam (\0.05 lmol m-2 s-1) plus a white light halogen source for saturation pulses (8,000 lmol m-2 s-1, 0.8 s). An external halogen source provided actinic white light. Incident photon flux was adjusted through a trimmer and measured on line with a small quantum sensor located on the leaf clip. Actinic light was step-wise decreasing from 1,600 to 50 lmol m-2 s-1 while saturation pulses were superimposed every 30 s. A 3 min duration of each actinic step was enough to obtain stable readings of PSII photo 0 chemical efficiency, which was calculated as Fm F0 0 0 :Fm according to Maxwell and Johnson (2000). F0 is the Fm 0 fluorescence yield just before the saturation pulse and Fm the maximum fluorescence yield obtained during the pulse.
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Apparent linear electron transport rate (ETR) was computed according to Genty et al. (1989) as ETR ¼ 0 0 Fm F 0 Fm PAR Absorptance 0:5; where PAR is the incident photosynthetically active radiation and 0.5 a correction factor assuming equal distribution of energy absorption by the two photosystems. Sample absorptance was obtained in separate measurements with an imagingPAM system (see below). Since we aimed to compare ETR obtained from fluorescence data and A obtained from gas exchange, care was taken to keep corresponding experimental conditions as close as possible. Hence, measurements with the MiniPAM were performed in a ventilated room keeping temperature at 22°C and CO2 and H2O vapor pressures close to ambient. In addition, both types of measurements were performed with fully photosynthetically induced material in order to ensure maximum stomatal conductance. The O2 dependence of apparent linear electron transport rate was assessed in separate experiments by using leaf discs and cut segments from petioles and pedicels. To this purpose, the plant material was placed on top of moistened filter paper in a home-made, parallelepiped glass cuvette. A gas mixing pump (G400, Qubit Systems Inc., Kingston, Canada) and pure gas cylinders (Air Liquide, Athens, Greece) were used to produce the required gas mixture which, after passing through a humidifier, was led to the cuvette at a flow rate of 1 l min-1. Relative humidity ([95%), temperature (22°C) and O2 levels were monitored by inserting a Vaisala (HM 34, Vaisala Inc., Woburn, Ma, USA) humidity cup and thermocouple and the PSt1 oxygen microsensor in the outlet part of the cuvette. The cuvette was appropriately attached to an imaging-PAM system (Walz, Effeltrich, Germany) equipped with a bank of blue LEDs providing measuring, actinic and saturation pulse light and a CCD camera capturing fluorescence. The signal used was integrated over the whole sample area, avoiding the edges which were unavoidably wounded by cutting. Before each series of measurements the instrument probes the sample reflectance in the red (650 nm) and infrared (780 nm) band. A built-in equation is then used for the relative estimation of sample absortivity as 1 ðR650 = R780 Þ; to be used in the calculation of ETR both in the imaging-PAM and the Mini-PAM measurements. A white filter paper equally reflecting in the two spectral bands was used for system calibration. After absortivity measurements, the samples were illuminated with 400 lmol m-2 s-1 actinic irradiances, during which saturating pulses (2,400 lmol m-2 s-1, 0.8 s) were superimposed every 60 s so as to close all PSII reaction centers. ETR was calculated as described above in the experiments with the Mini-PAM. The experiment started by pumping ambient air concentration to the samples (O2, 21%; CO2, 0.04%; N2, 78.96%), followed by step-wise decrease in O2 to 0%
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and back to ambient air, while N2 was used as a buffer (i.e., CO2 was kept constant). Each step lasted typically for 5–6 min and during the second half of this period ETR values were constant. The relative magnitude of cyclic electron flow around PSI was assessed through the post-illumination transient increase in fluorescence yield after turning off actinic light (Mano et al. 1995; Munne´-Bosch et al. 2005). For that purpose, plants were dark-adapted for 1 h and the plant material to be measured (leaf, petiole or pedicel) was inserted in the leaf clip of the Mini-PAM fluorimeter and illuminated for 3 min at 1,000 lmol m-2 s-1 by the external white light source. Upon light/dark transition, the fluorescence yield dropped suddenly to a value approach0 ing F0 (hereby defined as F0 ) and subsequently displayed a transient increase to a peak value (hereby defined as FP0 ) within the next 60 s. This transient, post-illumination rise in apparent F0 is due to dark reduction of plastoquinone by stromal reductants and it is considered as a measure of the potential for cyclic electron flow. The relative magnitude 0 of this fluorescence rise is given as F0P F0 F0P (Kotakis et al. 2006). Prompt chlorophyll fluorescence measurements in dark-adapted plant material Fast chlorophyll fluorescence transients of intact darkadapted (overnight) samples were captured by a Hansatech (Handy-PEA, Hansatech Instruments Ltd, Kings’ Lynn, Norfolk, UK) fluorimeter. For excitation, a bank of three red (peak at 650 nm) LED’s provided 3,000 lmol m-2 s-1 at sample level and fluorescence was recorded from 10 ls to 1 s with data acquisition rates 105, 104, 103, 102 and 10 readings in the time intervals of 10–300 ls, 0.3–3 ms, 3–30 ms, 30–300 ms and 0.3–1 s, respectively. Cardinal points in the fluorescence versus time curve used for further calculation of biophysical parameters were the following: maximal fluorescence intensity (Fm); fluorescence intensity at 20 ls, considered as the first credible measurement (F0); fluorescence intensity at 300 ls (F300ls), needed for the calculation of the initial slope (M0) of the relative variable fluorescence versus time curve; fluorescence intensity at 2 and 30 ms, i.e., at the J and I steps, respectively (FJ and FI). These primary data were used to derive the following parameters according to the JIP-test (Strasser et al. 2004), as extended to analyze also events in or around PSI (Jiang et al. 2008; Tsimilli-Michael and Strasser 2008; Oukarroum et al. 2009). a.
The photosynthetic efficiencies at the onset of illumination, i.e., the maximum quantum yield of primary photochemistry uPo ¼ TR0 =ABS ¼ ðFm F0 Þ=Fm ¼ Fv =Fm (where TR and ABS denote the trapped and
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Table 1 Chlorophyll content (mg cm-2), pigment ratios (mg mg-1) and absorptance of leaves, petioles and pedicels of Z. aethiopica Leaves Chl a ? b Chl a/Chl b
Petioles
34.2 ± 4.5a 3.26 ± 0.11
Pedicels
24.3 ± 3.9b a
41.6 ± 6.9c b
2.91 ± 0.08c
2.72 ± 0.18 a
Total carotenoids/Chl a ? b
0.208 ± 0.010
Absorptance
0.808 ± 0.013a
ab
0.197 ± 0.018
0.186 ± 0.007b
0.834 ± 0.022b
0.840 ± 0.020b
Values are means ± SD from 10 (chlorophyll content and pigment ratios) and 20 (absorptance) independent measurements. Different letters for each parameter indicate significant differences (p \ 0.05) between leaves, petioles and pedicels
absorbed excitation energy fluxes); the efficiency to conserve trapped excitation energy as redox energy (i.e., electron transfer, ET) wEo ¼ET0 =TR0 ; the quantum yield of electron transfer to intermediate electron carriers uEo ¼ET0 =ABS ¼ uPo wEo ; the efficiency of electron transfer between intermediate carriers to the reduction of end electron acceptors (RE) of PSI, dRo ¼RE0 =ET0 ; and the quantum yield of reduction of end electron acceptors of PSI, uRo ¼ uPo wEo dRo : b. The specific flux for absorption per active (i.e., QAreducing) center, ABS/RC, reflecting the relative antenna size. c. The relative amplitude of the I–P phase, 1 -VI, reflecting the content of PSI reaction centers. In addition, the area over the O–P transient, normalized over the variable fluorescence, Sm ¼ Area=ðFm =F0 Þ; was also calculated as a measure of the relative pool size of total electron carriers. The formulas used for the calculation of the above parameters are given as supplementary electronic material (Online resource 1). Other methods For pigment extraction, the hot dimethylsulfoxide (DMSO) method (Wittmann et al. 2001) was used, as conventional grinding in a mortar was difficult for petioles and pedicels. After clearing the extracts by centrifugation, photosynthetic pigments were determined in the supernatant after scanning in a Shimadzu UV-160A spectrophotometer and using the equations of Wellburn (1994). Internal O2 levels in petioles and pedicels were recorded both under darkness and illumination by inserting a calibrated 140 lm fiber optic microsensor (type PSt1) of a Microx TX3 oxygen meter (PreScens, Regensburg, Germany) into the organs according to Rolletschek et al. (2009). Statistics When needed, significance of differences in the measured parameters between leaves, petioles and pedicels were
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assessed by one-way analysis of variance (ANOVA) (SPSS 15.00 statistical package). The number of independent measurements in each case is given in the legends of figures and tables.
Results Conventional microscopic observations indicated the presence of stomata in both petioles and pedicels, yet at half the stomatal frequencies of leaves. Abundant aerenchyma was also evident (not shown). Compared to leaves, area-based chlorophyll concentrations were lower in petioles and higher in pedicels while chl a/chl b ratios were lower in petioles and pedicels. The carotenoid/chlorophyll ratio was slightly lower only in pedicels (Table 1). Light absorptance in petioles and pedicels was comparable to that of leaves (Table 1). Net CO2 assimilation rates (A) were considerably higher in leaves at both limiting and saturating light levels, while dark respiration rates were higher in petioles (Fig. 1a). Pedicels were not measured since their size did not allow credible gas exchange measurements. Leaf stomatal conductance increased with light intensity while that of petioles was considerably lower and light-independent (Fig. 1b). Intercellular CO2 (Ci) concentrations were higher in petioles at low light but almost similar at high light (Fig. 1c). Corresponding A versus Ci curves obtained at 800 lmol m-2 s-1 PAR displayed considerably lower initial slopes and finally attained levels in petioles, indicating both mesophyll and stomatal limitations for CO2 assimilation (Fig. 2). Hence, as shown in Table 2, parameter I, denoting stomatal limitations, is almost 50% higher in petioles, while carboxylation efficiency, calculated from the initial slope, was fourfold higher in leaves. We may note that apart from stomatal and Rubisco activity/content limitations, a reduced capacity to use the incident irradiance may also be deduced from the considerably lower CO2 assimilation rates in petioles at saturating PAR and Ci. Mesophyll limitations, however, could not be ascribed to a lower linear electron transport rate (ETR) in petioles, as this was almost similar to that of leaves (Fig. 3). The same was true for pedicel linear ETR. Hence, the discrepancy
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a
b
Fig. 2 Response of the rate of photosynthesis (A) to intercellular concentration of CO2 (Ci) in leaves (dark circles) and petioles (open rectangles) at 800 lmol m-2 s-1 irradiance. The curves were fitted using Hill 1 model
c Table 2 Calculated values for relative stomatal limitations (I), carboxylation efficiency (CE) and maximum apparent quantum yield for CO2 assimilation in leaves and petioles of Z. aethiopica Parameters
Leaves
Petioles
I
0.252 ± 0.018
a
0.371 ± 0.079b
CE
0.074 ± 0.004a
0.018 ± 0.006b
a
0.017 ± 0.005b
Max apparent quantum yield
0.047 ± 0.005
Values are means ± SD from 6 and 8 independent measurements for leaves and petioles, respectively. Different letters for each parameter indicate significant differences (p \ 0.05) between leaves and petioles Fig. 1 Light response curves of net photosynthesis (A), stomatal conductance (gs) and intercellular CO2 (Ci) in leaves (dark circles) and petioles (open rectangles). Data are means ± SD from 6 (leaves) and 8 (petioles) independent measurements
between comparable ETR’s, but considerably different A, points to higher photorespiration rates in petioles (and probably pedicels). Notwithstanding that the assumed higher oxygenation/carboxylation ratios in petioles occur at similar intercellular CO2 (see Fig. 1c) and O2 levels (0.20 and 0.21 mol mol-1 O2 for light-adapted petioles and pedicels are not shown). The assumed higher photorespiration rates in petioles and pedicels were further examined by applying chlorophyll fluorescence-based ETR measurement in discs of leaves and cut segments of petioles and pedicels in the presence of varying partial pressures of O2 and normal CO2. As shown in Fig. 4, ETR was progressively decreased with decreasing O2, yet the decrease was more steep in petioles and even steeper in pedicels. When the oxygenation reaction was completely suppressed in the absence of O2, ETR’s were reduced by 31, 52 and 96% in
leaves, petioles and pedicels, respectively. Hence, the higher photorespiration rates in non-foliar organs were confirmed. Cyclic electron flow activity was also higher in petioles and pedicels (Fig. 5). In fact, leaves displayed negligible post-illumination fluorescence yield increase, while that of petioles and pedicels was considerable. Fast chlorophyll fluorescence rise curves from all organs displayed the typical O–J–I–P transients when plotted on a logarithmic time scale, yet curves from petioles and pedicels lie higher than those of leaves (Fig. 6a), indicating that at each time point the fraction of ‘‘closed’’ PSII centers was higher in these organs. The differences were maximized at the low (i.e., microseconds) and high (i.e., milliseconds) part of the curve and more specifically in the K-band (around 300–600 ls) and the I-step (30 ms). This is better shown in Fig. 6b, where the F versus t kinetics for leaves is subtracted from that of petioles and pedicels. Hence, the most profound qualitative differences were the higher initial slope of the curve in the O–J phase and the decrease in
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Fig. 3 Light response curves of the linear electron transport rate (ETR) in leaves (dark circles), petioles (open rectangles) and pedicels (open triangles). Data are means ± SD from 6 (leaves) and 8 (petioles) independent measurements
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Fig. 5 Relative cyclic electron flow around PSI [(FP0 - F00 )/FP0 ], assessed from the post-illumination fluorescence yield increase, in leaves, petioles and pedicels. Data are means ± SD from 15 independent measurements. Different letters denote statistically significant differences (p \ 0.05)
a
b
Fig. 4 Fluctuation of the normalized electron transport rate (ETR) of leaves (dark circles), petioles (open rectangles) and pedicels (open triangles) relatively to the atmospheric concentration of O2. All ETR values of each organ are normalized at the corresponding max ETR value under ambient conditions (O2, 21%; CO2, 0.04%). Data are means ± SD from 12 independent measurements
the relative amplitude of the I–P phase in petioles and pedicels. Results of a numerical analysis of the O–J–I–P transient are shown in Table 3. Relative antenna size per active (QA-reducing) PSII centers (as reflected in the parameter ABS/RC) was higher in petioles and pedicels while relative pool sizes of total electron carriers (as shown by the parameter Sm) were lower. This indicates a more ‘‘shadeacclimated’’ character in petioles and pedicels, probably linked to their more perpendicular position in relation to leaf laminae. This is also supported by the lower chl a/chl b
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Fig. 6 Fast chlorophyll fluorescence transients (O–J–I–P) from leaves (dark circles), petioles (open rectangles) and pedicels (open triangles) of Z. aethiopica given on a logarithmic time scale and expressed either as relative variable fluorescence (Vt, i.e. double normalized at the F0 and FP points), or after subtraction of the petiole and pedicel transients from that of the leaves (DVt). Each point represents the mean from 15 leaves, petioles or pedicels
ratios in petioles and pedicels (Table 1). Maximum quantum yield of photon trapping of PSII (as given by uPo, equivalent to Fv/Fm) was high and in all organs.
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Table 3 Numerical values for quantum yields and flux ratios (uPo, uEo, uRo, wEo, dRo), specific flux for absorption per QA-reducing PSII center (ABS/RC, reflecting the relative antenna size), relative pool size of total electron carriers (Sm) and the relative amplitude of the I–P phase (1 - VI, reflecting the content of PSI reaction centers) Parameters
Leaves
Petioles
uPo
0.80 ± 0.02a
0.81 ± 0.02a
0.78 ± 0.01b
uEo
a
b
0.35 ± 0.02b
b
0.40 ± 0.03
0.37 ± 0.03
uRo
0.20 ± 0.02
0.14 ± 0.01
0.14 ± 0.01b
wEo
0.49 ± 0.03a
0.46 ± 0.03b
0.45 ± 0.02b
dRo
a
b
0.40 ± 0.02c
b
0.18 ± 0.01c
1 - VI Sm ABS/RC
a
Pedicels
0.49 ± 0.02
a
0.37 ± 0.02
0.24 ± 0.02
0.17 ± 0.01
a
b
36.0 ± 3.3
26.0 ± 1.7c
22.9 ± 2.1 a
2.17 ± 0.23
b
2.63 ± 0.26
2.81 ± 0.20c
Values are means ± SD from 15 independent measurements. Different letters for each parameter indicate significant differences (p \ 0.05) between leaves, petioles and pedicels
Corresponding quantum yield of electron flow from PSII to intermediate e- carriers (uEo) was slightly (yet significantly) lower in petioles and pedicels, while the quantum yield of reduction of end acceptors of PSI (uRo) was considerably lower, indicating progressively increasing limitations in linear electron flow along PSI. This is also evident in the parameters wEo and dRo, being the probabilities of electron transport from reduced QA to intermediate carriers (wEo) and from intermediate carriers to end acceptors of PSI (dRo). Thus, wEo was slightly (i.e., 6–8%) lower in petioles and pedicels, while corresponding decreases in dRo were considerable (24–18%). Hence, limitations in linear electron flow at time zero (i.e., in darkadapted samples) are more pronounced in PSI. Concomitant to that may be the finding that the relative amplitude of the I–P phase (1 - VI in Table 3) is considerably lower (29–25%) in petioles and pedicels. 1 - VI has been recently linked to the content of PSI reaction centers (Oukarroum et al. 2009).
Discussion It is evident from the results of this investigation that petioles and pedicels of Z. aethiopica display high rates of photorespiration at the expense of CO2 assimilation. In leaves, photorespiration has been implicated in stress resistance (Niyogi 2000). For example, under water stressdependent stomatal closure, carbon assimilation is limited by the availability of CO2 and, as a result, the electron transport system is over-reduced and may lead to photoinhibitory damage. Photorespiration, in this case, may function as an alternative sink of excess electrons (Stuhlfauth et al. 1990; Valentini et al. 1995). Apparently,
the distribution of electrons between CO2 assimilation and photorespiration is governed by the catalytic properties of Rubisco and the CO2/O2 ratio at the vicinity of the enzyme. In our case, however, no appreciable differences in the CO2/O2 ratios in the intercellular spaces of leaves, petioles and pedicels were evident. Accordingly, we may assume a reduced Rubisco specificity factor and/or a reduced mesophyll conductance to CO2 diffusion. Our results do not allow us to discriminate between the two possibilities. We may note, however, that although species-specific variations in Rubisco specificity factor have been reported (Delgado et al. 1995; Galme´s et al. 2006), it is not known whether similar variations do exist between organs of the same species. We may also note that the similarity in the intercellular CO2 concentration between leaves and petioles occurs in spite of the much lower stomatal conductance in the latter. This can be linked to the low carboxylation efficiency in petioles (Table 2). Alternatively, an enrichment of petiole intercellular spaces with CO2 diffused from the root and/or the leaf through the abundant aerenchyma could be inferred. Although photorespiration can serve as an alternative electron sink under adversity, a possible function in the absence of stress is seldom discussed. An implication in amino acid metabolism has been theoretically assumed (Heldt 2005) and documented with the use of appropriate Arabidopsis mutants (Rachmilevitch et al. 2004). We may assume a similar function in photosynthetic non-foliar organs as well. It is also evident in the present investigation that, apart from photorespiration, petioles and pedicels also display active cyclic electron flow around PSI (Fig. 5). Physiological functions of cyclic PSI electron flow are, again, linked to stress. Firstly, cyclic flow contributes to nonphotochemical energy quenching when the absorbed photon energy is in excess of what is needed for CO2 assimilation (Niyogi 2000). This can be applied in our case, as CO2 assimilation rate in petioles is very low while their absorptance is equally high to that of leaves (Fig. 1a; Table 1). Equal absorptance in the presence of unequal chlorophyll concentrations (see Table 1) are not contradictory since the dependence of absorptance on chlorophyll concentration is very weak, at least for the range of concentrations found in our test organs (Dima et al. 2006). The second proposed function for cyclic electron flow is linked to photorespiration. Under photorespiratory conditions, the ATP/NADPH ratio required to drive CO2 assimilation is increased (von Caemmerer 2000). Since linear electron flow produces both ATP and NADPH while cyclic electron flow produces only ATP (Bukhov and Carpentier 2004), a diversion of electron flow from PSI back to intermediate carriers would provide
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the additional ATP required. In our case, however, CO2 assimilation rates in petioles are already low as a result of decreased Rubisco activity and/or content (Figs. 1, 2; Table 2). Yet, we may add that although the overall energetic requirements of the photorespiratory cycle are only slightly higher compared to the Calvin cycle (Edwards and Walker 1983), that part of photorespiration occurring in the chloroplast (i.e., regeneration of PGA from glycerate) is considerably more energy-consuming. Alternatively (or in addition), ATP demand in petioles (and possibly pedicels) would increase if malate coming up from roots with the transpiration stream (Ben Zioni et al. 1971) is actively decarboxylated in petioles. This has been proved in leaf petioles of tobacco and Arabidopsis, which also contain levels of decarboxylating enzymes approaching those of C4-leaves (Hibberd and Quick 2002; Brown et al. 2010). Depending on the decarboxylating enzyme used, either ATP is consumed or NAD(P)H is produced (Edwards and Walker 1983). Hence, an active cyclic flow would act as an electron valve restoring the ATP/NADPH ratio. A similar function has been proposed for fruits and peridermal stems (Kotakis et al. 2006; Kalachanis and Manetas 2010) suffering from hypoxia (Pfanz et al. 2002) which suppresses mitochondrial respiration (Geigenberger 2003). The results from the analysis of the fast chlorophyll fluorescence rise curves confirm the absence of limitations in the linear electron flow along PSII and the presence of corresponding limitations in the linear flow along PSI. They also provide a possible mechanistic base for the diversion of electrons to the cyclic flow. Petioles and pedicels (this investigation), fruits (Kalachanis and Manetas 2010) and peridermal stems (Kotakis et al. 2006) display characteristic similarities in the O–J–I–P fluorescence rise curves indicative of low PSI contents and low probabilities for electron flow from reduced intermediate carriers to end PSI acceptors. These limitations in linear electron flow along PSI may facilitate the diversion of electrons back to intermediate carriers. In conclusion, the link between increased photorespiration and cyclic PSI electron flow already observed in leaves under adversity (Makino et al. 2002) could be an innate attribute of petioles under normal conditions, serving particular functions like an active amino acid metabolism and/ or malate decarboxylation.
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