JXB Advance Access originally published online on July 2, 2004
Journal of Experimental Botany 2004 55(403):1687-1696; doi:10.1093/jxb/erh190
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RESEARCH PAPER |
Internal recycling of respiratory CO2 in pods of chickpea (Cicer arietinum L.): the role of pod wall, seed coat, and embryo
1CSIRO Plant Industry, GPO Box 1600, Canberra ACT 2601, Australia
2CSIRO Plant Industry, Private Bag No. 5, Wembley WA 6913, Australia
* To whom correspondence should be addressed. Fax: +61 2 6246 5000. E-mail: robert.furbank{at}csiro.au
Received 5 February 2004; Accepted 19 April 2004
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Abstract |
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It has previously been proposed that respiratory CO2 released from the embryo in grain legume pods is refixed by a layer of cells on the inner pod wall. In chickpea this refixation process is thought to be of significance to the seed carbon budget, particularly under drought. In this study it is reported that the excised embryo, seed coat, and pod wall in chickpea are all photosynthetically competent, but the pod wall alone is capable of net O2 evolution over and above respiration. The predominant role of the pod wall in refixation is supported by measurements of fixation of isotopically labelled CO2, which show that more than 80% of CO2 is fixed by this tissue when provided to the pod interior. Chlorophyll concentrations are of the same order for embryo, seed coat, and pod wall tissues in younger pods on both an area and a fresh weight basis, but decline differentially with development from 12–30 d after podding. Imaging of chlorophyll distribution in the pod wall suggests that less than 15% of chloroplasts are located in the inner layer of cells thought to refix CO2 in legumes; this would be sufficient to refix less than 40% of respired CO2. It is concluded that while all tissues of the pod are capable of refixing respiratory carbon, the entire pod wall is responsible for the majority of this process, rather than a specialized layer of cells on the inner epidermis. The role of this fixed carbon in the pod for reallocation to the seed is discussed
Key words: Chickpea, photosynthesis, pod wall, respiration, seed coat
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Introduction |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
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Photosynthesis by reproductive structures of both dicots and monocots has been a controversial research area for some years. In cereals, some authors claim that up to 76% of grain carbon can be derived from ear photosynthesis while others stress the role of ear photosynthesis in refixing respiratory carbon (Araus et al., 1993
, and references therein). In dicots there is considerable interspecific variation in the photosynthetic capacity of reproductive structures. In oilseed rape (Brassica napus L.) pods, for example, rates of net CO2 fixation can be as high as 35% that found in leaves on an area basis and equivalent on a chlorophyll basis (King et al., 1998). In legumes, net pod photosynthesis is generally lower and, in chickpea, rates are generally less than 5% of that in the subtending leaf (Leport et al., 1999; Ma et al., 2001). However, in dicots, the role of photosynthesis in reproductive structures to reduce respiratory CO2 loss has also been emphasized (Flinn et al., 1977; Atkins et al., 1977; King et al., 1998; Ma et al., 2001). There has also been considerable controversy over the pathway of photosynthesis in these tissues of both cereals and dicots, with frequent claims of C4 photosynthesis and refutation of this (Imaizumi et al., 1997; Bort et al., 1996; Singal et al., 1987; King et al.,1998).
There is now a body of evidence in a range of legumes that the pod structure is capable of refixing a substantial proportion of respiratory CO2 generated during the light (Flinn et al., 1977; Sambo et al., 1977; Sheoran et al., 1987; Leport et al., 1999; Ma et al., 2001). Recently, there has also been a focus on the potential importance of this refixation process under terminal drought (Ma et al., 2001). There is limited information, except in field pea, as to the specific tissue responsible for this refixation, the developmental regulation of this process or the specialization of the pod structure to optimize this process. Here, an analysis of the contribution to respiratory CO2 refixation by various tissues in the pod of chickpea (Cicer arietinum L.) is presented both at the tissue and cellular level and the implications for the physiological role of photosynthesis in this reproductive structure are discussed.
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Materials and methods |
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Growth of plant material
Chickpea plants (Cicer arietinum L. cv. Garnet) were grown singly in 25 cm diameter pots in a compost mix in a naturally lit Canberra glasshouse during the summer under 21/14 °C day/night temperatures and watered daily to maintain soil moisture close to field capacity. Pods were tagged immediately after fertilization for the determination of developmental age.
Harvesting of pod material
Three stages of development were chosen for the experiments described here: full pod elongation with seeds in the early seed development/cell division phase (12–15 days after podding, DAP), seeds in the late cell division/early storage product accumulation phase (22–24 DAP) and, finally, pods where seeds were in the late storage product accumulation phase, predesiccation (28–30 DAP). These stages are shown in Fig. 1, which also illustrates the relative sizes of the embryo, seed coat, pod wall, and pod gas space at these developmental ages. Podding occurred approximately 6 d after flowering in these experiments.
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Oxygen exchange and chlorophyll fluorescence measurements
Pods of various ages were removed from the plant at the beginning of the photoperiod and immediately dissected for measurements of oxygen exchange at a range of light intensities on excised tissue. Pods were cut along the major vascular bundle and the seed removed. Pod walls from half of the fruit were placed in a leaf-disc electrode (Hansatech, Kings Lynn, Norfolk, UK) as described in Delieu and Walker (1981)
for polarographic determination of O2 evolution in the presence of a water-saturated gas phase containing 1% CO2 provided from a bicarbonate buffer. Pod tissue was kept at 25 °C by circulating water. Illumination was provided by a 150 W quartz halogen slide projector lamp attenuated by neutral density filters. Seed coats and embryos were assayed in a similar fashion using seeds dissected into halves and placed in the cuvette with the outer surfaces facing the light source. Chlorophyll fluorescence was measured in 22–24 DAP intact pods, or intact seeds excised from them, including seed coats. The same cuvette was used as for O2 exchange measurements and chlorophyll fluorescence was detected using the Waltz PAM 101 pulse-modulated detection system (Walz-Effeltricht, Germany) described by Schreiber et al. (1986). In these experiments the pod tissue was illuminated from a halogen lamp (Schott KL 1500, Schott, Germany) using a quadrifurcated fibre-optic light guide providing a PAR of 400 µmol quanta m–2 s–1 at the leaf surface. Pod tissue was illuminated in various gas phases until a steady fluorescence yield was obtained (15–20 min). Where DCMU was added, it was painted with a brush onto the surface of the detached pod 5 min before illumination at a concentration of 100 µM in 98% ethanol. Ethanol alone had no effect on chlorophyll fluorescence. Electron transport rates were calculated according to Genty et al. (1989), assuming 50% of the absorbed photons were utilized by photosystem 2. Absorption of pod tissues was taken to be 1% transmission, ignoring reflectance, as the organs were too small to measure reliably in an Ulbricht sphere. Chlorophyll was measured in methanol extracts according to Porra et al. (1989). Pod wall and seed coat transmission of photosynthetically active radiation (PAR) was measured using a Li-Cor quantum probe sensor (Li-Cor, Lincoln, Nebraska, USA) and the illumination system described above. The area of individual pod tissues was determined with a leaf area meter (Li-Cor, Lincoln, Nebraska, USA).
13CO2 labelling
Feeding of 13CO2 to intact, attached chickpea pods and subsequent analysis of isotopic content of individual tissues was carried out as described in Ma et al. (2001). Attached pods in the glasshouse (in the light at 09.00–10.00 h) were injected with 0.5 ml of 13CO2 (99.9 atom%), placed into the space between the pod wall and the seed coat using a syringe. Pods were then harvested 1 h and 24 h after labelling, dissected into embryo, seed coat, and pod wall then freeze-dried and analysed by mass spectrometry. The 1 h period of exposure was the shortest which could be used to minimize the translocation of fixed carbon while ensuring a reliable measurement of 13C incorporation (data not shown).
14CO2 leak rate determination
Intact attached pods were injected with 0.5 ml 14CO2 (Amersham Life Sciences, Amersham, UK; 2.5 µmol; 20x106 dpm µmol–1), using the same procedures as for 13CO2 feeding. Attached pods were placed in air in a sealed 25 ml cylindrical Perspex cuvette illuminated at 1000 µmol quanta m–2 s–1 by a 150 W quartz projector lamp. The gas phase was circulated through a closed system by an aquarium pump, bubbling through a solution of 30% v/v hyamine (methylbenzethonium hydroxide; Sigma–Aldrich) in methanol to trap the 14CO2 released from the pod. Samples of hyamine were removed at regular intervals and the radioactivity determined by scintillation counting (see Hatch et al., 1995, for the principle of measurement). The entire procedure was carried out at 25 °C. After 1 h in the light, the pods were removed and frozen in liquid nitrogen. The pod wall, seed coat, and embryo were ground separately in liquid nitrogen then extracted with 70% ethanol and water at 80 °C for the determination of total radioactivity incorporated into the soluble and insoluble fractions (Lunn and Hatch, 1995).
Microscopy
Pod wall, seed coat, and seed sections (150–250 µm), from fresh glasshouse-grown material harvested mid-photoperiod, were cut using a sledge microtome. Sections were examined for chlorophyll fluorescence using a Leica DMR epifluorescence microscope (Leica Microsystems, Sydney, Australia) fitted with a long-pass filter for excitation wavelengths 450–490 nm, a 510 nm dichroic mirror, and for detection a 515 nm long-pass filter to collect both green and red fluorescence simultaneously. Images were recorded digitally. Sections were also iodine-stained for starch visualization (Lunn and Furbank, 1997) and examined under transmitted light with the same microscope system. Sections of the entire seed were cut by hand and examined for chlorophyll distribution with a Leica laser scanning confocal microscope, exciting with a 633 nm red HeNe laser, and collecting emission between 660 and 720 nm. Quantification of chlorophyll fluorescence images was done using the Imagequant software package (Amersham Biosciences, Amersham, UK).
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Results |
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Chlorophyll concentration in the pod wall, embryo, and seed coat was measured together with the light transmission characteristics of the pod wall and seed coat at three stages of development (Fig. 2A–E). At 12–15 DAP, chlorophyll concentration was highest in all tissues. The pod wall and the seed coat contained the highest absolute chlorophyll contents at the two earliest stages, similar on a fresh weight basis in both tissues but higher in the seed coat on a surface area basis. By 28–30 DAP, chlorophyll concentration had declined by more than 80% in seed coats and 60–70% in pod walls. By contrast, while embryo chlorophyll on an area basis declined between 12 and 22 DAP, it remained constant between the two later stages. Interestingly, seed coat transmission increased with development, which would potentially allow greater light utilization by the embryo (Fig. 2E). Pod wall light transmission remained relatively constant with development, despite changing chlorophyll concentration, suggesting that other cellular constituents play a major role in light absorption by this tissue.
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While the above data are indicative of bulk chlorophyll concentration, they do not reflect the cellular localization of chloroplasts or any morphological specialization which may be present in the tissues of the pod. Figures 3 and 4 address this issue by microscopic analysis of chlorophyll fluorescence and starch distribution in sections of the pod wall (Fig. 3) and embryo and seed coat (Fig. 4). Figure 3 shows a transverse section through the pod wall of a 22–24 DAP pod either proximal (A, B) or distal (C, D) to the major vascular bundle. Figure 3A and C show light micrographs after iodine staining for starch while Fig. 3B and D show fresh sequential sections examined using fluorescence to image chlorophyll distribution. Note the heavily lignified schlerenchyma cells (labelled schl) in a band close to the inner pod wall. Starch staining was confined to the cells at the outside of this layer despite the presence of a layer of chloroplast-containing cells on the inner pod wall (B, D). Chloroplasts were more abundant on the inner side of the pod wall distal (D) rather than proximal (B) to the vascular bundle. Figure 3E shows a quantification of the chlorophyll fluorescence across the area of the pod wall section shown in (D), indicated by the line. In this representative section, 87% of the chlorophyll was located in the outer wall region, including the schlerenchyma layer. In four other sections, this varied from 85% to 90% (data not shown). Figure 4A and B show similar sections from the seed coat of seeds 22–24 DAP, either iodine-stained (A) or using fluorescence (B). Starch is restricted principally to a heavily stained area immediately below the epidermis or an area of the outer seed coat adjacent to the inner seed coat (A). Unlike the pod wall, there is no distinct localization of chlorophyll across the section, but there appears to be a higher chlorophyll concentration in the area below the epidermis, heavily stained for starch (A, B). Figure 4C shows a laser confocal microscope image of a thick fresh section of seed and seed coat to determine if chlorophyll distribution in the embryo was localized to any specific cells. Consistent with Fig. 2, fluorescence intensity was lower in the embryo than the adjacent seed coat, and there appeared to be no specialized localization of chloroplasts within the embryo.
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To estimate the relative photosynthetic capacity of the component tissues of the pod, the response of net O2 exchange of pod wall (A), seed coat (B), and embryo (C) to light intensity at three stages of development is shown in Fig. 5. Measurements were made under saturating CO2, commensurate with concentrations measured in the gas phase of the chickpea pod space (Ma et al., 2001
). Only the pod wall was capable of net O2 evolution at physiological light intensities. In both pod wall and seed coat, photosynthetic rate increased initially during development then declined with developmental age but was always saturated at light intensities of approximately 1000 µmol quanta m–2 s–1. On an area basis, while incapable of net photosynthesis due to high rates of respiration, the seed coat of the two earlier developmental stages was capable of higher rates of gross photosynthesis than the pod wall. Interestingly, while unresponsive to light at the two earlier developmental stages, embryo photosynthesis was just detectable at the 28–30 DAP stage. By contrast, both pod wall and seed coat gross photosynthetic capacity were highest at 22–24 DAP, falling to very low levels at 28–30 DAP, however, the respiration rate in these tissues also progressively declined in the later stages of development.
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Using the pod and seed coat light transmission data of Fig. 2, the surface area of pod wall, seed coat, and embryo and interpolating from the response of O2 exchange on an area basis to irradiance (Fig. 5), it is possible to estimate the relative photosynthetic contributions of the individual pod tissues at physiologically relevant irradiation. This estimation is shown in Fig. 6 for the three developmental stages at an incident PAR of 1500 µmol quanta m–2 s–1 at the pod surface. Clearly, the pod wall alone would be capable of net O2 evolution and, presumably, net CO2 fixation in the intact pod under these conditions. Net pod photosynthesis, calculated from the sum of the O2 exchange rates per organ, was positive at the two earlier stages, but at 28–30 DAP, net respiration would be predicted for an intact pod. This is in fact, similar to the observations of Ma et al. (2001)
, where net photosynthesis in young pods, measured using CO2 exchange, was observed to decline with age and water stress.
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An alternative method of estimating photosynthetic electron transport rate, which is not complicated by respiration or measurements of gas exchange, is pulse-modulated measurement of photosystem 2 chlorophyll fluorescence (Schreiber et al., 1986
). This technique allows the calculation of the electron transport rate and equivalent rates of gross O2 evolution, providing the light intensity during measurement and the absorption characteristics of the tissue are accurately known (Genty et al., 1989). Thus, in tissue exclusively fixing respired or internally generated CO2, but not carrying out net gas exchange, photosynthetic capacity can still be estimated (Maxwell et al., 1998). Table 1 shows the result of such an experiment carried out with intact pods or excised seeds (22–24 DAP) exposed to saturating irradiance either in air, 5% CO2 or CO2-free air. Assuming that four electrons are required to be transported through photosystem II per O2 evolved, the calculated values of gross O2 evolution by the intact pod correspond well with the O2 exchange measurements made with isolated pod walls, when respiration rate in the dark is added to net O2 evolution at the appropriate irradiance (Fig. 5A). Notably, when DCMU [3-(3,4-dichlorophenyl)1,1-dimethylurea], an inhibitor of photosystem 2 electron transport, was painted on the pod, electron transport was reduced by 90%. The residual electron transport was due to small amounts of fluorescence emanating from within the pod, as determined by measuring the pod wall following excision of the seed after treatment (data not shown). The excised seed, including the seed coat, in this experiment was capable of more than 50% of the electron transport rate of the pod wall in 5% CO2, when illuminated at the same irradiance. A second pod of similar developmental age was measured under the three gas phases to determine if pod photosynthesis was largely dependent on externally supplied CO2 or if photosynthesis was already saturated by internal respiratory CO2. Clearly the latter is the case, as photosynthetic electron transport did not respond to either high CO2 or a total lack of externally supplied CO2. Similarly, the excised seed (including seed coat) was unresponsive to externally supplied CO2. In this experiment, the rates of seed electron transport were similar to those seen for the pod, commensurate with the gross O2 evolution rates calculated from Fig. 5.
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As an independent measure of the capacity for the pod wall to fix respiratory CO2, intact pods (22–24 DAP), still attached to the plant, were exposed to 13CO2, injected into the pod space, and, after 1 h of exposure under saturating illumination, tissue was excised and analysed for distribution of 13C (Fig. 7). Consistent with the other measurements made here, the majority of CO2 was fixed by the pod wall (approximately 80%) with the seed coat and embryo together contributing less than 20%. After 24 h, around 30% of the fixed carbon had been remobilized from the pod wall to the embryo while there was no net translocation of carbon from the seed coat during this period.
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It has been suggested that for the efficient capture of released respiratory CO2 in legume pods, a substantial diffusion barrier exists within the pod wall (Atkins et al., 1977
). To test this for chickpea, the pod space of an attached pod (22–24 DAP) was injected with 14CO2 of known specific activity (as done for 13C in Fig. 7) and the efflux of radiolabelled carbon into CO2-free air was followed by trapping released CO2 in a closed chamber (a representative time-course of efflux is shown in Fig. 8). This experiment determines the maximum rate of CO2 efflux across the pod wall under a large CO2 gradient. The rate of CO2 efflux from the pod was linear over a period of 1 h (Fig. 8). This was repeated for six pods of varying age from 15–28 DAP with similar results (data not shown). Leakage rates were remarkably reproducible, ranging from 115 dpm min–1 to 160 dpm min–1. From the specific activity of the 14CO2 supplied and the pod surface area, this equates to an efflux rate of 0.24–0.34 nmol CO2 m–2 s–1. Analysis of pod tissues following labelling confirmed the conclusion that the pod wall was the major photosynthetic tissue refixing CO2 with 76–98% of labelled carbon appearing in the pod wall 1 h after labelling of the six pods of varying age (data not shown).
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Discussion |
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There have been many reports in the literature of the photosynthetic characteristics of legume pods (Crookston et al., 1974
; Flinn et al., 1977; Sambo et al., 1977; Atkins and Flinn, 1978; Sheoran et al., 1987; Leport et al., 1999; Ma et al., 2001) and of Brassica species (King et al., 1998, and references therein). It has frequently been observed that the net photosynthetic gas exchange of intact legume pods is minimal (see Leport et al., 1999; Ma et al., 2001, and references therein). Stomatal density of the pod wall epidermis in legumes is less than one-third that of leaves (Ma et al., 2001, and references therein) and in chickpea, net CO2 exchange rates are at best 5% of that in the subtending leaf (Ma et al., 2001). While these measurements of whole pods tend to diminish the importance of pod photosynthesis in seed filling, they do not adequately address the respiratory CO2 refixation capacity of the pod. Respiration rates in the light are often of the same magnitude as photosynthetic capacity, making interpretation of such data difficult. This problem of measuring net fluxes may be further complicated by inhibitory effects of light and high CO2 on respiration (Haupt-Herting et al., 2001; Pinelli and Loreto, 2003), although such effects have only been reported in leaves to date. Recent work using gas exchange and stable isotope labelling has indicated that the pod wall is a major organ of refixation in chickpea pods (Ma et al., 2001) and is not affected by water deficit in the remainder of the plant. The stable isotope data in that work, however, was complicated by the long period after labelling before the tissue was measured (24 h) and the resultant possibility for remobilization of fixed carbon. Respiratory CO2 is also released predominantly within the embryo, beneath the seed coat, while in labelling experiments, CO2 is supplied outside the seed coat in the pod space. Thus, it is important to examine the individual photosynthetic capacity of tissues within the pod to assist with the interpretation of such data. This proved to be pivotal in Brassica napus, where contrary to expectation, the embryo itself was the major site of respiratory CO2 refixation and the seed coat an effective barrier to CO2 diffusion (King et al., 1998).
The examination of individual tissues of the pod carried out here (Figs 2, 5, 6, 7) supports the hypothesis that the pod wall is the major organ of respiratory CO2 refixation in chickpea (Ma et al., 2001, and references therein). While chlorophyll concentration on a fresh weight basis is comparable between pod wall and seed coat, the fresh weight distribution between tissues within the pod, the reduced PAR through light absorption by the pod, and the relatively high respiration rate of the seed coat and embryo result in net negative contributions from cells of the seed (Figs 5, 6). It is worthy of note, however, that the seed coat is almost capable of refixing its entire respiratory CO2 production, while the embryo, unlike in Brassica napus (King et al., 1998), is never close to positive carbon balance in the light (Fig. 4). While the measurements made above estimate photosynthetic capacity/photosynthetic electron transport capacity, they do not directly measure the contribution of individual tissues to the fixation of CO2 from the pod space, as electron acceptors other than photosynthesis (such as in the use of NADPH for storage product synthesis) may play a role in reproductive structures (as postulated for Arabidopsis and Brassica napus by Ruuska et al., 2002). The conclusions from the O2 exchange data are, however, supported by the isotopic labelling experiments (Figs 7, 8) in which up to 80% of pod-space-derived CO2 was refixed by the pod wall, rather than the seed tissues. This approach is, however, also imperfect, as CO2 was injected into the pod space whereas the majority of respiratory CO2 release is normally occurring within the embryo, tissue difficult to inject with labelled carbon.
Microscopic examination of the chlorophyll localization in seeds and pod walls revealed several interesting observations (Figs 3, 4). The embryo chlorophyll was low and evenly distributed throughout the cotyledons with no evidence for specialized localization (Fig. 3C). The seed coat, by contrast, contained a layer of chloroplast-rich cells directly below the epidermis (Fig. 3B). This layer of cells is also rich in starch which is predominantly localized to this layer and a layer between the inner and outer seed coats (Fig. 3A). Seed coat starch in legumes such as pea (Pisum sativum) is thought to be a temporary carbon reserve for seed-filling which is remobilized later in development (Rochat and Butin, 1992). However, it is not clear in this case whether the starch in these outer layers of the seed coat is derived from de novo fixed carbon or translocated photoassimilate. The mechanism whereby the seed coat partitions starch between these cell types, which all contain plastids, is unknown.
Of particular relevance to the role of the pod wall in respiratory CO2 refixation are the data shown in Fig. 2. As seen in pea (Atkins et al., 1977), cells of the inner epidermis of the pod wall are rich in chloroplasts (Fig. 2D, E). However, in the majority of the pod wall tissue, i.e. that distal to the major vascular bundle, these cells contain no more than 15% of the pod wall chlorophyll and even less in the regions proximal to the vascular bundle (Fig. 3B, D, E). This observation is important because, based on the photosynthetic capacities and respiration rates determined in Figs 5 and 6, 15% of pod chlorophyll would allow this inner layer of cells to refix less than 40% of total respired carbon. Since the intact pod at the earlier stages shows small, but significant, rates of net photosynthetic gas exchange (Ma et al., 2001; this study), it is likely that the outer pod wall contributes significantly to this refixation process, contrary to reports for pea (Atkins et al., 1977). It is also worthy of note that the inner half of the pod wall of Phaseolus vulgaris (bean) contains no chloroplasts at all (Crookston et al., 1974), suggesting that there may be considerable interspecific variation in the pod structure of legumes and sites of CO2 refixation.
The structure of the chickpea pod wall also raises questions as to the relative role of the inner and outer layers of chloroplast-containing cells. As in pea (Atkins et al., 1977), in chickpea there is a heavily thickened layer of cells present in the mesocarp between the inner epidermis of the pod wall and the outer photosynthetic layers (the yellow/green fluorescent band in Figs 3B and D marked schl). It has been proposed that this layer provides a diffusion barrier to the efflux of respiratory CO2 that is trapped and refixed by the inner epidermal chloroplasts (Atkins et al., 1977; Ma et al., 2001). The presence of a substantial barrier to CO2 diffusion by the pod wall is evident from the high CO2 concentration generated in the chickpea pod in the dark (Ma et al., 2001) and by the slow efflux of CO2 from the pod after the 13CO2 and 14CO2 feeding experiments (Figs 7, 8). From Figure 7 and the specific activity of the 14CO2 provided, it can be calculated that CO2 efflux rates in the light vary from 0.24 nmol CO2 m–2 s–1 to 0.34 nmol CO2 m–2 s–1. From the respiration rates measured during the 13C labelling of intact pods (Fig. 6), a pod wall conductance to CO2 can also be calculated. Using this method, a similar calculated flux of 0.6 nmol CO2 m–2 s–1 is obtained (data not shown). If gross photosynthetic CO2 fixation rates are of the order of 4–5 µmol m–2 s–1 (Ma et al., 2001; this study), this potential leak rate represents only a tiny fraction of the photosynthetic capacity of the pod, and suggests an extremely tight and efficient refixation mechanism. By contrast, cuticular conductance in leaves with stomata closed has been estimated to be 100–400 µmol m–2 s–1 (Boyer et al., 1997), more than four orders of magnitude higher than the values obtained here for chickpea pods under the physiological CO2 gradient. In C4 leaves, the bundle sheath is a compartment specialized for the retention of CO2 and leakage of CO2 from these specialized cells, measured using a comparable technique, averaged around 16% of net photosynthetic rate (Hatch et al., 1995). Recently, the resistance of the bundle sheath compartment was also calculated from electron micrographs and component cellular resistances (von Caemmerer and Furbank, 2003) yielding values of 100–300 m2 s mmol–1 (or conductances of around 3–10 µmol m–2 s–1). Once again, even this conductance value is much higher than found here for the chickpea pod wall. However, surprisingly, when the thickness of the pod wall is taken into account, the reason for the low values obtained here becomes apparent. While the average thickness of the C4 bundle sheath is around 20 µm (and much of this is not traversed by CO2 during diffusion), the average thickness of the pod wall in the current experiments was 300–400 µm (Fig. 1). Assuming diffusivity and effective porosity values for cytoplasm used previously (von Caemmerer and Furbank, 2003), and a 300 µm diffusion path length, the calculated conductance to CO2 is 0.027 nmol m–2 s–1. When multiplied by a CO2 gradient of 30 (1% CO2 inside, 0.03% outside) this equates to a flux of approximately 0.8 nmol m–2 s–1. This value is very similar to the measured values in this study. Thus, a specialized schlerenchyma or cuticle may not be required to retain CO2 within the chickpea pod; a large diffusion path-length is sufficient to make the structure essentially ‘CO2 tight’. A similar conclusion was reached by von Caemmerer and Furbank (2003) concerning the relative roles of cell wall suberization and organelle location/cell size in providing resistance to CO2 diffusion in the C4 system.
The CO2 leakage measurements made here are also consistent with the data of Table 1. Photosynthetic electron transport in intact pods was found to be insensitive to the external gas phase (from zero CO2 to 5% CO2). From the current study it is difficult to determine the individual contributions of pod thickness, the cuticle, and the schlerenchyma layer in the pod mesocarp to the diffusion resistance of the pod wall. Interestingly, Ma et al. (2001) reported that pod CO2 fixation responded linearly to pod space CO2 concentrations from zero to 2%, far in excess of what would be required within the intercellular space of a leaf to saturate photosynthesis. This is also consistent with the outer mesocarp chloroplasts contributing to a large degree to respiratory CO2 refixation and that pod space CO2 must diffuse through a substantial barrier to reach this photosynthetic tissue.
It has been proposed (Ma et al., 2001) that if the inner epidermal cells of chickpea pod wall are responsible for the refixation of respiratory CO2, then this process may continue under drought, despite the drop in water potential known to occur in the outer cells of the pod wall (Shackel and Turner, 2000). As the seed coat and embryo appear to be somewhat insulated from this drop in water potential (Shackel and Turner, 2000), then the inner pod wall tissues may also maintain a higher water potential. The current results, however, cast some doubt on this hypothesis because the majority of CO2 fixation in the pod wall appears to be in the outer mesocarp cells which are subjected to low water potentials similar to those found in leaves during drought stress (Shackel and Turner, 2000). In light of the chloroplast localization seen here in the pod wall and the substantial CO2 diffusion resistance provided by the thickness of this wall, it is tempting to suggest that the schlerenchyma layer is solely a barrier to water movement in order to retain moisture around the seed during drought conditions.
The micrographs shown in Fig. 3A and C, where starch was stained with iodine, show that in the chickpea pod wall, starch is only produced in the mesocarp on the outside of the schlerenchyma layer. This starch showed little evidence of diurnal turnover (data not shown). The inner epidermal cells contained no starch and presumably produced only sucrose, which could then be translocated to the seed. These observations suggest that carbon fixed by these inner cells represents the rapidly remobilized labelled carbon seen in the 13C fixation experiment (Fig. 7). In 24 h, about 25% of carbon fixed by the pod wall was translocated to the seed. This is certainly similar to the proportion of respiratory carbon which the inner epidermal layer could potentially fix. Interestingly, in a previous study Ma et al. (2001) showed that, even after 6 d, only another 13% of carbon was lost from the pod wall. Taken together, these data suggest that at least a proportion of carbon fixed by the outer pod wall may act as a longer term carbohydrate reserve for remobilization during seed filling. This is consistent with changes in dry weight of the pod wall, especially under water deficits (Davies et al., 1999) and observations made in Vigna radiata (mungbean) where starch is stored during early development of the pod and remobilized later in seed filling (Chopra et al., 2000). This storage reserve would also be analogous to the starch stored in the seed coat of legumes earlier in development (Rochet and Butin, 1992) and polysaccharides stored in the stems of cereals which are later remobilized for grain filling (Wardlaw and Willenbrink, 2000, and references therein).
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Acknowledgements |
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The authors wish to thank Dirk Buessis, Walter Tate, Christiane Ludwig, and Renee Buck for growing and tagging plant material and for assistance with 13C labelling.
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References |
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