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Dep. of Crop and Soil Sciences, Cornell Univ., 521 Bradfield Hall, Ithaca, NY 14853
* Corresponding author (TLS1{at}cornell.edu)
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ABSTRACT |
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 TOPNOTES ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Abbreviations: ABA, abscisic acid • DAP, days after pollination • ELISA, enzyme linked immunosorbant assay • FW, fresh weight • gs, and stomatal CO2 diffusive conductance • PFD, photon flux density • Pn, net photosynthetic CO2 exchange rates
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INTRODUCTION |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Photosynthate supply has substantial impact on kernel set, as indicated by studies involving altered illumination levels (Singh and Jenner, 1984; Setter and Flannigan, 1989; Andrade et al., 1999), elevated atmospheric CO2 (Mitchell et al., 1996), and culture with various levels of exogenous sugars (Singh and Jenner, 1984; Lee et al., 1989). Furthermore, studies of water deficit indicate that the magnitude of loss in kernel set is correlated with the extent of loss in photosynthesis (Schussler and Westgate, 1991a, b) and photosynthate influx into kernels (Schussler and Westgate, 1995), and that kernel set can be partially recovered with stem infusion of supplemental sucrose (Zinselmeier et al., 1995a; Zinselmeier et al., 1999). Nevertheless, in these studies the loss in kernel set was not fully attributed to carbohydrate deprivation caused by stress (Schussler and Westgate, 1991a,b, 1994, 1995; Zinselmeier et al., 1995a). Thus, additional factors may be involved.
Hormones are potential signals which could regulate development in response to stress. In response to high temperature stress during the post-pollination phase, maize and wheat (Triticum aestivum L.) kernels accumulate considerably less zeatin and zeatin riboside, a possible contributor to their high rate of abortion (Cheikh and Jones, 1994; Banowetz et al., 1999). Exogenously applied cytokinin has been shown to increase kernel set in maize (Dietrich et al., 1995). Although studies have not been reported on the possible involvement of altered cytokinin levels in relation to loss of kernel set during water deficit or photosynthate deprivation, such effects are plausible given the rapid rise in cytokinin concentrations during the post-pollination phase (Lur and Setter, 1993; Dietrich et al., 1995), and the substantial inhibition of cell division in response to water and light deprivation (Ober et al., 1991; Artlip et al., 1995; Cheng and Lur, 1996).
Several lines of evidence indicate that ABA plays a role in the loss of kernel set in response to water deficit at the post-pollination phase of development in maize (Ober et al., 1991; Artlip et al., 1995) and wheat (Westgate et al., 1996). ABA accumulates in endosperms or whole kernels in response to water deprivation (Ober et al., 1991; Westgate et al., 1996) and kernel set is lost in response to exogenously supplied ABA (Myers et al., 1990; Mambelli and Setter, 1998). In addition, the rise in ABA levels in maize are specific to the kernels in apical ear regions (Ober et al., 1991), which are most prone to loss in kernel set (Artlip et al., 1995).
Studies by Ober and Setter (1992) indicated that the source of stress-induced ABA that accumulates in endosperms during their period of cell division is the maternal tissues, not the filial tissues (endosperm and embryo). The impact of stress on ABA levels in maternal tissues of maize reproductive organs, however, has not been reported.
In the current studies, we compared plant response to light deprivation, which was expected to limit photosynthate supply, with response to water deficit, which limits photosynthesis but also may exert additional stress effects. Our objective was to test the possible involvement of cytokinins, ABA, and sugar depletion in altering kernel set. We subjected maize plants to discrete 5-d periods of water and light deprivation at both pre- and post-pollination stages, and analyzed hormone levels and nonstructural carbohydrates in two components of reproductive organs in apical and basal ear regions, differing in kernel set.
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MATERIALS AND METHODS |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Pots (10 L) contained peat, vermiculite and perlite (1:1:1 v/v/v) with 6 g of pulverized limestone, 35 g of CaSO4, 42 g of powdered FeSO4, 1 g of fritted trace elements (Peters FTE 555, Scotts Co., Marysville OH), and 3 g of wetting agent (AquaGro G, Aquatrols, Pennsauken, NJ). An automated system watered pots with a nutrient solution containing 0.6 g L-1 of Peters 15-16-17 (Scotts Co., Marysville OH) at 2-h intervals during the day. Watering was sufficient to leach excess nutrient salts.
In Exp. I, plants were sown on seven dates, creating seven replicate batches, with flowering dates from 13 March to 8 June, 1997. In Exp. II, plants were sown on five dates, creating five replicate batches, with flowering dates from June to August, 1998. For each batch, plants were assigned randomly to the developmental stage at which treatments were to occur (pre- or post-pollination), then plants were assigned randomly to control, shade, or water deficit treatments. There were four plants in each stage x treatment combination of each batch. Two of the plants were chosen randomly for tissue sampling on the last day of treatment. The remaining two plants were returned to control conditions and harvested at maturity to measure kernel yield per plant.
In both Exp. I and II, plants used for post-pollination treatments were pollinated 4 to 5 d after silk emergence. Plants used for pre-pollination treatments were pollinated on the day treatments were terminated. In both cases, ears were bagged prior to silking, fresh pollen was collected from neighboring well-watered sibs and pollen was applied manually to silks.
Water and Shade Treatments
We began water deficit treatments on the first day of silk emergence or at 1 d after pollination (DAP) for pre-pollination and post-pollination treatments, respectively. On these dates, each plant/pot system was fully watered, allowed to drain freely, and initial weights were obtained. At this point, irrigation was withheld, and plants were allowed to deplete water until each plant/pot system was <50% of initial weight (usually attained at 2 d after withholding irrigation). Then, pots were irrigated daily at 0800 h to return weights to the 50% gravimetric set point. Visible signs of stress (leaf rolling and glaucous leaves) began at 1.5 to 2 d after withholding water; lower leaf necrosis occurred at later stages of stress.
Shade treatments were begun 1 d after the start of water deficit treatments (1 d after silk emergence and 2 DAP for pre- and post-pollination stages, respectively) to permit water deficit treatments to deplete soil water and approach a stressful condition at the time shade treatments were begun. For treatments imposed at the pre-pollination stage, tissues were sampled at 6 d after silk emergence and for those imposed at the post-pollination stage, tissues were sampled at 7 DAP. Thus, both shade and water deficit will be hereafter referred to as 5-d treatments.
Photosynthesis and Stomatal Conductance
Net photosynthetic CO2 exchange rates (Pn) and stomatal CO2 diffusive conductance (gs) were measured in Exp. II on the first leaf above the node of primary ear insertion. Data were obtained with a portable closed gas exchange system (LI-6200, LI-COR Inc., Lincoln, NE) equipped with a 0.25-L leaf cuvette (12-cm2 exposed area). Data for shaded plants were obtained with ambient shade illumination. In control and water deficit treatments, when ambient illumination was <1000 µmol photons m-2 s-1, light for gas exchange measurements was supplemented with an LED red light source (Quantum Devices, Lincoln, NE) that provided 1200 µmol photons m-2 s-1 during measurements. Measurements were made between 1100 and 1400 h on the days indicated in figure legends. Four plants representing each treatment combination were sampled in each of the five replicate blocks of the study.
Kernel Sampling
At 0800 to 1000 h on the final date of treatments, tissues were sampled from apical regions (Exp. I) or both apical and basal regions (Exp. II) of the ears. For the pre- and post-pollination stages, basal samples were obtained from sites 33% of the distance from the base to apex of the ear. For the pre-pollination stage, apical samples were from sites about 10 to 15% of the distance from apex to base, while for the post-pollination stage, apical samples were from sites in the 5th to 7th ring of fertilized kernels counting from the apex of the ear.
At the pre-pollination stage, florets were excised at the pedicel and frozen in liquid N2. Then, the underlying region of cob (rachis) containing the internal ring of dense vascular tissue (hereafter referred to as vascular cob) was excised with a razor blade and frozen in liquid N2. At the post-pollination stage, pericarp and nucellus were excised at a point 33% distal from the kernel tip and this portion of the pericarp/nucellus was discarded, then endosperm and nucellus tissues were scooped out from the remaining portion of the kernel and frozen in liquid N2. The underlying pedicel tissues, including the vascular and placental tissues forming the cup-shaped structure at the endosperm base, were excised and frozen in liquid N2.
Tissues were weighed, then hormones and sugars were extracted in a 10:1 ratio (v/w) of extraction solvent (0.8 m3 methanol/m3 aqueous solution) to tissue fresh weight. Hormone and sugar concentrations were expressed on a fresh rather than a dry weight basis because sugars and hormones are expected to exert effects in proportion to their concentration in cellular solution. Tissue water contents were about 850 to 900 g/kg at the water potentials to which kernels were subjected in these studies (Ober et al., 1991). Tissues were macerated with a pestle, stored at -18°C for 
1 wk, and centrifuged prior to further use of extracts.
Sugars Assay
The concentrations of glucose, fructose, and sucrose were determined by an enzyme-coupled-reaction assay based on hexokinase and glucose-6-phosphate dehydrogenase (Cairns, 1987). Phosphogluco-isomerase and invertase were used to convert fructose and sucrose stoichiometrically to glucose for subsequent assay. All enzymes and reagents were from Sigma Chemical Co (St. Louis, MO). Extracts were evaporated to dryness at 24°C and assayed directly along with sucrose, fructose and glucose calibration standards spanning the useable range from 0 to 10 µg per well. Starch was assayed in insoluble debris following three extractions of sugars with methanol. Water was added and starch was hydrated by heating to 121°C for 10 min in an autoclave. After cooling, starch was completely hydrolyzed to glucose with amyloglucosidase (#208469, Gibco BRL) as described by Ober et al. (1991), and aliquots were assayed for glucose as described above.
ABA Chromatography
A 100-µL aliquot of each extract was dried in vacuo at <24°C, then redissolved in 100 µL of Solvent I ([0.2 m3 methanol + 0.01 m3 glacial acetic acid]/m3 aqueous solution). ABA was separated with reverse-phase chromatography on columns packed with 15 mg of 40-µm diameter C18-silica material (J.T. Baker Chemicals, Phillipsburg, NJ). Solvents were eluted by centrifuging at about 3000 g in a swinging-bucket rotor containing 1.7-mL conical collection tubes. Columns were pre-equilibrated with Solvent I, then samples were loaded in 100 µL, contaminants were eluted with 400 µL of Solvent I and discarded, then ABA was eluted with 100 µL of Solvent II ([0.55 m3 methanol + 0.01 m3 glacial acetic acid]/m3 aqueous solution). Radiolabelled tracer ABA indicated greater than 90% recovery of ABA with this procedure. ABA fractions were dried in vacuo at <24°C.
ABA Assay
ABA fractions from C18 chromatography were assayed for ABA by indirect enzyme linked immunosorbant assay (ELISA) in a method similar to that described by Walker-Simmons (1987) and Ober et al. (1991), with modification as described below. Briefly, plates (Costar High Binding #3366, Corning Inc., Corning, NY) were coated overnight with 1.4 µg of ABA-4'-bovine serum albumin conjugate in 200 µL of 50 mM NaHCO3 buffer, pH 9.6, containing 0.2 g L-1 NaN3 at 5°C. Plates were washed four times with TBST wash buffer, which contained Tris-buffered saline (TBS; 50 mM Tris-hydroxymethyl-aminomethane, pH 7.5, 1 mM MgCl2, 10 mM NaCl, and 0.2 g L-1 NaN3) to which 1 g L-1 Tween-20 (#P-7949, Sigma Chemical Co.) was added. ABA eluates from C18 chromatography were redissolved in 100 µL of TBST. Samples were then incubated with primary antibody with the following in each well: 90 µL of TBSA (TBS with 1 g/kg bovine serum albumin [A-8022, Sigma Chem. Co.]), 2 µL of C18 eluate, and 100 µL of TBSA containing 0.2 µg of anti-ABA monoclonal antibody (clone #15-I-C5, Mertens, Deus et al., 1983; currently available from Agdia Inc., Elkhart, IN). On each plate, a duplicate set of (+)ABA standards (Sigma Chemical Co.) containing a 1:2 dilution series of 12 values from 12.3 to 0.006 pmol/well served as a calibration curve. The antibody was added last to all wells on a plate. After incubation overnight at 5°C, plates were washed four times with TBST and 200 µL of secondary antibody solution containing 10 nL of anti-mouse IgG-alkaline phosphatase conjugate (#A-3562, Sigma Chemical Co.) in TBSA was added per well. After incubation overnight at 4°C, plates were washed four times with TBST and 0.2 mg p-nitrophenyl phosphate (PNPP) was added in 200 µL of buffer containing 0.9 M diethanolamine, pH 9.8, and 3 mM MgCl2. Plates were incubated for about 1 h at 24°C and absorbance at 405 nm was read with a plate reader (model 750, Cambridge Technology, Watertown, MA). (+)ABA content was determined by calculations based on (+)ABA calibration standards and a logit transformation of data.
Cytokinin Chromatography
In Exp. II, cytokinin samples were partially purified prior to ELISA with two types of chromatography: immunoaffinity and reverse phase C18. Immunoaffinity columns were packed with 80 µL of acrylic beads (O-7628, Sigma Chem. Co.) to which affinity-purified anti-zeatin antibody was conjugated by the oxirane reaction. The anti-zeatin antibody was a polyclonal antibody made in rabbit (Oryctolagus cuniculus L.) that was specific for zeatin and zeatin-containing molecules [zeatin riboside (ZR), dihydroxyzeatin, dihydroxyzeatin riboside]; its production and characterization was described by Lur and Setter (1993). This antibody was affinity-purified on a column packed with poly-L-lysine conjugated agarose beads (P-1666, Sigma Chem. Co.) to which zeatin riboside was immobilized by the IO4 reaction (MacDonald and Morris, 1985). Rabbit serum was loaded onto the ZR-immobilized column, contaminants were removed with 20 bed volume washes each of 0.5 M NaCl and 10 mM Tris-HCl buffer (pH 7.5), and anti-zeatin antibody was eluted with glycine buffer (100 mM, pH 2.5), triethylamine buffer (100 mM, pH 11.5) and methanol. These antibody fractions were combined and freeze dried prior to conjugation to acrylic beads as described above. Zeatin and related compounds in plant extracts were separated on the anti-zeatin immunoaffinity columns as follows. For each sample, 400 µL of extract was dried in vacuo at <24°C. Samples were spiked with 42 kBq of [3H]dihydrozeatin (Amersham Co., Arlington Heights, IL, HPLC-purified just prior to use), dissolved in 75 µL of TBST and loaded onto 80-µL-bed immunoaffinity columns that had been pre-equilibrated with TBST. Samples were loaded slowly (1–2 min) by gravity, contaminants were washed out with 300 µL TBST, zeatin-containing compounds were eluted with 200 µL of methanol, and the zeatin fraction was dried in vacuo. Recovery of zeatin compounds averaged >90% on the basis of analysis of radioactivity in non-zeatin fractions; reported data are corrected for losses in recovery.
C18 separation of zeatin-related compounds used the same system described above for ABA. Samples were loaded in 75 µL of Solvent A (10 mM tri-ethylamine-acetate, pH 3.4, in water), zeatin was eluted with 125 µL of Solvent B (0.24 m3 methanol:0.80 m3 Solvent A [v/v]), and zeatin-riboside was eluted with 200 µL of Solvent C (0.30 m3 methanol:0.70 m3 Solvent A [v/v]).
Zeatin/ZR Assay
An indirect competitive ELISA was used for zeatin and zeatin riboside fractions after C18 chromatography. Fractions were dried in vacuo at <24°C, then redissolved in 100 µL of TBST. ELISA plates (Costar High Binding #3366) were coated overnight at 5°C with 4 ng of zeatin-riboside-bovine serum albumin conjugate (MacDonald and Morris, 1985) in 200 µL of 50 mM NaHCO3 buffer, pH 9.6, containing 0.2 g L-1 NaN3. Plates were washed four times with TBST, and the following were added to each well: 10 to 60 µL of dried C18 eluate redissolved in TBST, sufficient additional TBST to bring to 100 µL, and 100 µL of TBSA containing 1.4 µg of anti-trans-zeatin riboside monoclonal antibody (clone #J3-I-B3, Eberle et al., 1986; currently available from Agdia Inc., Elkhart, IN). On each plate, a duplicate set of zeatin-riboside (Sigma Chem. Co.) standards containing a 1:2 dilution series of 12 values from 20 to 0.01 pmol/well served as a calibration curve. The antibody was added last to all wells on a plate. After incubation overnight at 5°C, secondary antibody and PNPP color development steps were followed as described above for ABA ELISA. The assay gave a sigmoidal standard curve with an inflection point at 0.3 pmol, and a useable range from 0.04 to 5 pmol/well.
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RESULTS |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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0.05) increased in vascular cobs and endosperm/nucellus in response to stress. Concomitantly, starch concentrations declined in florets, endosperm/nucellus, and pedicels in response to stress (Fig. 2a,c,d). Thus, water deficit increased partitioning into sugars relative to starch reserves.
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0.05) from controls at 3 to 5 d after terminating the treatments (Fig. 4a,c). Stomatal conductance (gs) paralleled Pn throughout the treatment and recovery phases of shade and water deficit treatments (Fig. 4b,d). Thus, the shade and water deficit treatments exerted a similar effect on photosynthesis at both pre- and post-pollination stages.
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DISCUSSION |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Both water deficit and shade decreased photosynthesis to a similar extent during 5-d stress periods, and photosynthesis recovered similarly after stress was relieved (Fig. 4). It has been suggested that shade as well as water deficit might exert their influence by altering photosynthate status of the affected tissues. Infusion of sucrose solution into maize stem partially ameliorated the negative impact of water deficit on kernel set (Zinselmeier et al., 1999; Zinselmeier et al., 1995a), consistent with the postulated role of photosynthate supply in regulating kernel set during drought episodes.
The current studies indicate, however, that if photosynthate status is the common underlying factor by which kernel set responds to shade and water deficit, the process is more complex than simple depletion of sugar in the affected tissues. In response to water deficit, sucrose concentrations increased in the reproductive tissues (florets at the pre-pollination stage and endosperm/nucellus at the post-pollination stage) while hexose and starch concentrations were either unchanged (pre-pollination florets) or increased (post-pollination endosperm/nucellus) (Fig. 2, 5, and 6). Furthermore, in highly vascularized plant parts (vascular cobs at the pre-pollination stage and pedicel at the post-pollination stage), water deficit either had no effect or increased sugar concentrations. In contrast, hexose concentrations in florets, vascular cobs and endosperm/nucellus decreased in response to shade. Furthermore, the effects on sugar concentration were not any greater in apical than in basal ear regions.
Thus, the current data are contrary to the outcome predicted if sugar concentration were the sensed property that initiated kernel abortion. Similarly, in previous studies of maize, pre-pollination water deficit had no effect or slightly increased sugar concentrations in unpollinated ovary tissues (Schussler and Westgate, 1991b, 1994; Schussler and Westgate, 1995; Zinselmeier et al., 1995b), while post-pollination water deficit did not substantially affect sugar concentrations in developing endosperms (Ober et al., 1991).
In contrast to the lack of treatment effect on sugar concentrations per gram fresh weight (FW), the increment of nonstructural carbohydrate accumulation was substantially decreased by shade and water deficit at both pre- and post-pollination stages (Fig. 7). These data agree with studies of Schussler and Westgate (1995) who suggested that kernel set depends on the rate of photosynthate flux into ovaries rather than on concentration per se. If regulation based on photosynthate flux is involved, it apparently involves coordinate suppression of fresh weight growth such that sugar concentrations are maintained (Fig. 2, 5, 6). Setter (1993) suggested that a possible mechanism of sensing sugar flux would be via a system that responds to changes in the concentration of a rapidly turning over pool of carbohydrate-pathway intermediates, such as UDP-glucose, in conjunction with continuous carbohydrate use in respiration and other consumptive pathways. Indeed, Zinselmeier et al. (1999) observed that water deficit decreased maize ovary UDP-glucose, and the levels partially recovered in response to exogenous sucrose feeding. This model appears to have potential validity in the post-pollination stage, where effects on carbohydrate flux were greater in apical than basal tissues (Fig. 7), corresponding to the greater loss of kernel set in apical kernels (Fig. 1). Also, it follows from the observation that after relief from stress, basal kernels recover, whereas apical kernels lag and fail to recover endosperm cell division and further development (Ober et al., 1991; Artlip et al., 1995). It is unclear, however, how such a model could explain the current observation that at the pre-pollination stage the flux of carbohydrate is drastically decreased in both apical and basal florets, while kernel set is mostly lost in the apical florets (Fig. 1, 7).
Cytokinin
In the current work, we obtained a composite estimate of the zeatin family of compounds. It is possible that such an estimate could mask changes in an individual zeatin compound. In many previous studies, however, treatments affected the whole family of interconvertable zeatin compounds (Redig et al., 1996). Previous studies have indicated that cytokinin concentrations decrease substantially in in vitro cultured maize kernels subjected to heat stress (35°C) at the early post-pollination stage, corresponding to stress effects on endosperm cell division (Cheikh and Jones, 1994). Exogenously applied cytokinin improves maize kernel set (Dietrich et al., 1995). Other stresses, including water deprivation and salinity in a variety of plant systems, have provided contradictory evidence on the impact of stress on endogenous cytokinin levels (Hare et al., 1997). In the current study, shade did not affect zeatin-like cytokinin concentrations, while water deficit had inconsistent effects (Fig. 9). At the pre-pollination stage, water deficit increased cytokinins in apical florets, whereas at the post-pollination stage it decreased cytokinins in apical and basal pedicels.
Zeatin-like cytokinin levels at the pre-pollination stage were generally about 10-fold lower than at the post-pollination stage, consistent with observations in developing wheat carpels (Lee et al., 1988). Thus, the sites of zeatin-like cytokinin synthesis and flux, as well as regulatory roles, probably differ at these contrasting developmental stages, and accordingly, the impact of stress on their concentrations differ.
Abscisic Acid
At both the pre- and post-pollination stages, water deficit substantially increased ABA concentrations (Fig. 8). Previous studies had similarly shown that water deficit elevates ABA levels in maize endosperm at the post-pollination stage (Ober et al., 1991; Myers et al., 1992) and in wheat spikelets at the pre-anthesis stage (Westgate et al., 1996). The present data also show that water deficit substantially elevates ABA concentrations in plant parts containing a high proportion of phloem tissue: vascular cob and pedicel (Fig. 8). Previous study showed that in plants subjected to water stress, ABA accumulating in endosperms at early post-pollination stages was of maternal origin and that phloem transport from leaves to kernels was a possible source of such ABA (Ober and Setter, 1992). The current observations are consistent with long distance phloem transport of ABA into reproductive tissues, although they do not rule out local ABA synthesis in the vascular cob and pedicel. The latter possibility is apparently limited, however, as in vitro cultured maize kernels accumulate only modest amounts of ABA in response to low water potential treatments (-2.0 MPa) applied via culture media (Myers et al., 1992).
Surprisingly, ABA concentrations in reproductive tissues also increased in response to shade treatments (Fig. 8). Few studies have examined ABA in response to shade or other light treatments. Leaves of wheat (Nan et al., 1999), tomato (Lycopersicon esculentum Mill.) (Basiouny et al., 1994), and strawberry (Fragaria grandiflora Duch.) (Kubik and Michalczuk, 1993) accumulated ABA in response to low-light treatments, and shade increased ABA concentration in phloem exudates from strawberry leaves (Kubik and Michalczuk,, 1993).
It is possible that the shade-induced increase in ABA is an outcome from complex, interacting regulatory systems. Young and Gallie (2000) have shown that regulation of programmed cell death during maize endosperm development involves interactions between ABA and ethylene signaling pathways. In some plant systems, such interactions appear to involve ABA stimulation of ethylene accumulation (Gomez-Cadenas et al., 2000). Indeed, Cheng and Lur (1996) showed that in response to shade during pollination and early kernel development, maize kernel tissues substantially accumulated 1-aminocyclopropane-1-carboxylate synthase (ACC synthase), a key enzyme in ethylene biosynthesis, and ethylene. Moreover, in citrus (Citrus reshni Hort.), photosynthate deprivation due to leaf defoliation increased ABA levels in fruit, and this in turn elevated ACC and ethylene levels, which led to fruit abscission (Gomez-Cadenas et al., 2000).
Responses Specific to the Apical Zone
Water deficit at the post-pollination stage increased ABA substantially more in tissues of apical than basal kernels (Fig. 8), corresponding with loss in kernel set (Fig. 1). Previous studies had shown that water deficit elevates ABA concentrations in endosperms at 7 to 13 DAP to a greater extent in apical than basal kernels (Ober et al., 1991). The current studies extend these findings to show that pedicel, as well as endosperm/nucellus, exhibited apical specificity in ABA accumulation, and that shade increased pedicel ABA concentrations to a greater extent in apical than basal kernels.
In contrast to the post-pollination stage, at the pre-pollination stage, ABA concentration tended to be only slightly higher, and not statistically higher (P 0.05) in apical than basal kernels. For both ABA and photosynthate flux (Fig. 8), stress had striking apical specificity at the post- but not the pre-pollination stage. Thus, to explain the apical specificity in kernel set at the pre-pollination stage, additional information is needed. A contributing factor may be the more advanced developmental stage of basal than apical florets, which may permit the basal florets to better resume growth after water or light deprivation is relieved. Our current studies are addressing this possibility with an examination of development during recovery from treatments.
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ACKNOWLEDGMENTS |
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NOTES |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Received for publication September 5, 2000.
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REFERENCES |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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) and water deficit (
) treatments during pre-pollination (upper panels) or post-pollination (lower panels) developmental stages in Exp. II. Net photosynthetic CO2 exchange rate (Pn; left panels) and stomatal CO2 conductance (gs; right panels) of 3 to 5 replicates (mean ± SEM) are shown. Data are plotted with respect to the days from treatment termination (DFTT). On the days of treatment termination (0 DFTT), plants were re-watered or removed from shade at 0800 and gas exchange data was acquired at 1100 to 1400 h.
