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Plant Physiol. (1999) 121: 25-36
Starch and the Control of Kernel Number in Maize at
Low Water
Potentials1
Christopher Zinselmeier2,
Byeong-Ryong Jeong3, and
John S. Boyer*
College of Marine Studies and College of Agriculture and Natural
Resources, University of Delaware, Lewes, Delaware 19958
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ABSTRACT |
After
reproduction is initiated in plants, subsequent reproductive
development is sometimes interrupted, which decreases the final number
of seeds and fruits. We subjected maize (Zea mays L.) to
low water potentials ( w) that frequently cause this kind of failure. We observed metabolite pools and enzyme activities in the
developing ovaries while we manipulated the sugar stream by feeding
sucrose (Suc) to the stems. Low w imposed for 5 d around pollination allowed embryos to form, but abortion occurred and
kernel number decreased markedly. The ovary contained starch that
nearly disappeared during this abortion. Analyses showed that all of
the intermediates in starch synthesis were depleted. However, when
labeled Suc was fed to the stems, label arrived at the ovaries. Solute
accumulated and caused osmotic adjustment. Suc accumulated, but other
intermediates did not, showing that a partial block in starch synthesis
occurred at the first step in Suc utilization. This step was mediated
by invertase, which had low activity. Because of the block, Suc feeding
only partially prevented starch disappearance and abortion. These
results indicate that young embryos abort when the sugar stream is
interrupted sufficiently to deplete starch during early ovary
development, and this abortion results in a loss of mature seeds and
fruits. At low w, maintaining the sugar stream partially
prevented the abortion, but invertase regulated the synthesis of ovary
starch and partially prevented full recovery.
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INTRODUCTION |
In plants, reproduction involves intense biosynthetic activity.
Anthesis is generally rapid, and large amounts of photosynthate are
deposited in the developing reproductive structures. An example is
maize (Zea mays L.), which deposits about one-half of its
aboveground dry mass in the mature kernels (McPherson and Boyer, 1977 ;
Jurgens et al., 1978). This deposition depends on the number of
reproductive structures, which is initially determined by the
conversion of vegetative shoot apices to reproductive ones. Photoperiod
and other triggers are signals for the conversion, but the succeeding environment can also have an effect by changing floral development, altering pollination, or preventing fruit growth. The mechanisms for
the latter changes are generally unknown, but have a large impact on
agriculture because they decrease the number of seeds and fruits.
In maize, these later effects occur frequently and losses in kernel
development are especially important. Factors such as low water
potential (w) can arrest ovary growth,
especially if young embryos abort (Westgate and Boyer, 1986). The
abortion blocks kernel development despite the successful completion of
all of the earlier steps in reproduction, and kernel number is
permanently reduced. Many of the embryos can be rescued by feeding Suc
to the stem in substrate quantities at the time of low
w (Boyle et al., 1991b; Zinselmeier et al.,
1995a). The Suc must be fed in a quantity that replaces the
photosynthetic products missing because of inhibited photosynthesis
(Boyle et al., 1991b; Zinselmeier et al., 1995a). However, full
recovery has not yet been achieved by this method. Low
w affects many enzymes and metabolic
activities (Kramer and Boyer, 1995), and it is possible that Suc cannot
be fully utilized by the embryos. Because so many factors can change, those that limit embryo development remain unidentified.
Little is known about the fate of incoming carbon during the early
phases of reproductive development. Starch was correlated with early
pod development in soybean (Fader and Koller, 1985), and was responsive
to low w (Zinselmeier et al., 1995a, 1995b) and Suc feeding in maize (Zinselmeier et al., 1995a), but its location
and involvement in ovary development appear otherwise unexplored.
Schussler and Westgate (1991) showed that sugars are absorbed less
rapidly in embryos at low w than in controls,
and Zinselmeier et al. (1995b) showed that invertase loses activity, but other activities were not explored. To identify rate-limiting enzyme steps in metabolism, it is necessary to alter the metabolite flow and determine where the flow might be restricted. Mutations that
alter enzyme activities and therefore carbon flow have been used
successfully to study starch metabolism during late reproduction in
maize (Shannon and Garwood, 1984; Caspar, 1994), but no mutations are
available for comparable studies during early reproduction around the
time of pollination.
To alter carbon flow by a different means, we adapted the method of
Boyle et al. (1991b) and Zinselmeier et al. (1995a) for feeding Suc to
the intact plant. Supplying Suc increased the carbon flowing through
pathways otherwise depleted by the effects of low
w. It was sufficient to observe only whether
intermediary metabolites accumulated or were depleted in these pools. A
pool showing accumulation was considered to be upstream of the
rate-limiting step, and pools showing depletion were considered to be
downstream. The enzyme mediating the transition step was considered to
be rate-limiting and thus to regulate flow through the metabolic pathway. Compared with the mutant approach, this method did not otherwise manipulate enzyme activity, but it had the advantage that
specific changes could be made in intact plants, thus permitting ready
interpretation.
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MATERIALS AND METHODS |
Growth Conditions
Maize (Zea mays L. cv Pioneer Hi-Bred 3732) plants were
grown in 22-L pots containing 11 kg of Evesboro loamy sand, loamy substratum (coated, mesic, Typic Quartzipsamments) amended by mixing
peat:sand:soil in volumes of 1:1:1, and dolomitic limestone (275 g per
pot) to adjust the soil pH to 6.9. Ten seeds were planted in each pot
and the soil mix was saturated with a nutrient solution (Hoagland and
Arnon, 1950) and allowed to drain. The solution consisted of 4 mM KNO3, 6 mM
Ca(NO3)2·4H2O,
2 mM
MgSO4·7H2O, 2 mM
KH2PO4, 0.5 µM
CuSO4·5H2O, 10 µM
MnSO4·H2O, 2 µM
ZnSO4·7H2O, 25 µM
H3BO3, 0.5 µM
H2MoO4, and 50 µM iron citrate. The pots were placed in a
controlled-environment chamber with day/night temperatures and RH of
30°C/20°C ± 1°C and 40%/95% ± 5%, respectively.
Cool-white fluorescent lamps provided a 14-h photoperiod with an
irradiance of 850 to 1,000 µmol PAR m 2
s1 throughout the day at the top of the canopy.
After two-and-a-half weeks, seedlings were thinned to two plants per
pot and supplied with the same nutrient solution until drainage. One
week later, seedlings were thinned to one plant per pot and were
similarly supplied with nutrient solution twice weekly. Supplemental
KNO3 (12 mM) was supplied during
rapid stem elongation (21-50 d after planting). All nutrient additions
were terminated at silking, and water was supplied as required.
Planting density was approximately six plants per square meter, which
is equivalent to field densities for this genotype.
Low w Treatments
Plants were maintained at high w, except
for a period of 6 d during flowering (Fig.
1). When silks first appeared (d  5 from pollination), the soil was brought to field capacity by supplying 4 L
of water to each pot. Water was then withheld and was not fully
resupplied until d 1. The silks fully emerged and the plants were
pollinated on d 0. By rewatering on d 1, there was 1 d between pollination and rewatering to allow time for fertilization to occur
when w was at its lowest. To minimize leaf
senescence at low w, a small amount of water
was supplied on d 1 and d 0 to replace the water lost the previous
day (100-150 mL d1). For the high
w controls, the plants were maintained by
watering to drainage daily throughout the flowering period (about 1,500 mL d1).
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| Figure 1.
Schedule of watering and infusions for the low
w treatment. Water was withheld at the times shown by
the black bar. Infusions were made at the end of the dark periods, as
shown by the white bar. Arrowheads 1 through 5 on the plant show
infusion sites at each internode and correspond with arrowheads 1 through 5 on the white bar. A single infusion was made at sites 1 and
2. Two infusions were made at sites 3 to 5 to ensure uptake of 6.8 g of Suc.
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Net photosynthesis was measured on the unshaded third leaf from the top
of the plant, usually 3 h into the photoperiod, using a clamp-on
cuvette in a closed system (Sheoran and Boyer, 1989). The
w was determined on the same leaf using
isopiestic thermocouple psychrometry according to the method of Boyer
(1995). The w was also measured in the
ovaries, which we defined as the lower, swollen part of the pistillate
flower after the stigma and style were removed, and the glumes and
lemmae were detached where possible (Kiesselbach, 1949). For the
measurement, five to six pistils were excised from the ear, the silks
and other structures were removed quickly, the ovaries were placed in a
psychrometer cup, and the w was measured in an
isopiestic thermocouple psychrometer (Boyer, 1995). After this
measurement, the ovaries were removed from the cup, placed in a syringe
barrel, frozen, thawed, and the cell solution was extracted by pressing
on the syringe plunger. The osmotic potential
(s) of the solution was determined in the isopiestic psychrometer according to the method of Boyer (1995). The
turgor pressure (p) was determined with the
following equation: p = w s.
Stem Infusions
Suc solution from sugarcane (0.438 M,
s = 1.1 MPa) was infused into some of the
stems at low w using the method of Boyle et
al. (1991a), except that the infusions were begun in the dark at the
end of the night period, and the cavity was quickly filled with
deionized water and sealed with a pre-assembled rubber
septum/needle/syringe that acted as a reservoir. The Suc solution was
decanted into the reservoir, the reservoir was covered with the rubber
plunger of the syringe, and the air was vented using a 20-gauge needle. Infusions were usually installed within 30 s and sterile
techniques were used when possible. There was a single infusion site
per internode on d 4 and d 3, and two infusion sites per internode on d 2 to d 0 (Fig. 1). The ear internode and four additional internodes (one above and three below the ear internode) were used for
the infusions. Each day, 45 mL of infusion medium (6.8 g of Suc) was
supplied to every infused plant. This provided sufficient Suc to
completely replace that normally produced by photosynthesis (Jurgens,
et al., 1978; Westgate and Boyer, 1985a). Plants at high
w were not infused, because a previous study
(Zinselmeier et al., 1995a) found no effect of infusion at high
w.
Pollination
Tassels were excised from each plant 1 to 2 d before the
first silks appeared and stored at 4°C in a refrigerator with their bases in a nutrient solution (30 mM Suc, 0.02 M
KNO3, 0.02 M NH4NO3, 1.25 mM KH2PO4, 3.0 mM
CaCl2·2H2O, and 1.5 mM
MgSO4·7H2O). The evening before pollination, tassels
were brought to room temperature, and pollen was collected the next
morning by shaking the tassels in a bag. The plants were immediately
hand-pollinated by inverting the bag over the silks and shaking.
Metabolite Assays
Ovary samples were obtained from plants in each treatment and were
immediately frozen in liquid N2, freeze-dried,
and ground to a fine powder using a mortar and pestle. Suc and reducing
sugars were analyzed on powder samples of 50 mg dry weight according to
the method of Singletary and Below (1990) with slight modifications. Suc and reducing sugars were extracted three times using 80% (v/v) ethanol at 60°C. The supernatants were pooled and evaporated to dryness using forced air at 40°C. Suc and reducing sugars were resolubilized in 1 to 3 mL of water, sonicated, and mixed thoroughly. An aliquot was assayed in triplicate using the method of Handel (1968)
for Suc and the method of Nelson (1944) for reducing sugars. Starch was
analyzed in the insoluble pellet using a modified procedure from Hanft
and Jones (1986). The pellet was washed with water, boiled to
gelatinize the starch, and allowed to cool to room temperature. The
starch was enzymatically degraded using a mixture of amyloglucosidase (2 mg mL1) and amylase (1 mg
mL1) in 50 mM sodium acetate
buffer (pH 4.5). Samples were incubated at 55°C for 2 h. An
aliquot was removed and reducing sugars were quantified in triplicate
according to the method of Nelson (1944).
Phosphorylated intermediates were extracted by the method of Jelitto et
al. (1992) with a few modifications. Powder samples of 50 mg dry weight
were transferred to 1.5-mL microcentrifuge tubes and 0.5 mL of 10%
(v/v) TCA was added to each. After leaving the mixture on ice for
1 h, the tubes were centrifuged at 1,300g for 5 min,
and the supernatant was collected. The pellet was resuspended twice in
0.3 mL of water, centrifuged, and the supernatant collected and
combined with the first supernatant. The pellet was discarded. The
extract was transferred to a 10-mL glass test tube and 1 mL of
ethylether was added to extract the lipids. After vortexing, the
mixture was centrifuged at 1,500g and the ethylether phase was discarded. The TCA extract was washed three more times with ethylether. After the final wash, the TCA extract was carefully transferred to microcentrifuge tubes and the volume was reduced to 0.6 mL in a freeze drier (Speed-Vac, Savant Instruments, Holbrook, NY). The extract was mixed with 5 mg of activated charcoal prewashed with water and centrifuged for 30 s at full speed in a
microcentrifuge. The charcoal-treated extract was used for measuring
UDP-Glc by the method of Keppler and Decker (1984); Glc-6-P, Glc-1-P,
Fru-6-P, and 3-phosphoglyceric acid by the method of Stitt et al.
(1989); and Pi by the method of Taussky and Shorr (1953). The
extraction method and stability of metabolites during extraction was
checked by measuring metabolites added to the homogenized tissue before extracting with TCA. The recovery for UDP-Glc was 79%, for Glc-6-P 85%, for Glc-1-P 76%, for Fru-6-P 98%, and for 3-phosphoglyceric acid 72%.
Enzyme Assays
All enzyme assays were optimized for substrate concentration, pH,
and assay duration with the ovary extracts. Crude enzyme extracts were
made from samples of 100 mg dry weight from the freeze-dried ovaries by
grinding in a mortar and pestle in 4 mL of soluble extraction buffer
containing 50 mM HEPES-NaOH (pH 7.2), 5.0 mM
MgCl2, 15% (v/v) ethylene glycol, 1.0 mM EDTA, and 1.0 mM DTT (1 M NaCl
also was added when extracting insoluble acid invertase). Samples were
centrifuged at 14,000g for 4 min to pellet insoluble
material, the soluble protein fraction was decanted, and the remaining
pellet was washed three times with the soluble extraction buffer prior
to repeating the extraction with the high-salt buffer for insoluble
invertase. All extracts were desalted according to the micro-desalting
procedure of Helmerhorst and Stokes (1980) by applying 450 µL of
extract to a 3-mL column containing Sephadex G-25 and centrifuging at
750g for 2 min.
Acid invertase activity was divided into soluble and insoluble forms
according to the method of Doehlert and Felker (1987). Activity was
determined by mixing 100 µL of desalted extract, 200 mM
sodium acetate (pH 4.8), and 100 mM Suc (1 mL final assay volume) and incubating at 30°C for 30 min. The reaction was
terminated by adding 1 mL of Nelson's no. 1 reagent, and reducing
sugars were quantified according to the method of Nelson (1944) using Glc as a standard (50-200 µg mL1).
Debranching enzyme (pullulanase or limit dextrinase) was
determined according to the method of Doehlert and Kuo (1990) by mixing
100 µL of desalted extract, HEPES-NaOH (pH 7.2), 5.0 mM
MgCl2, and 2.5% (w/v) pullulan (1 mL final assay
volume), and incubating at 37°C for 60 min. The reaction was
terminated by adding 3 mL of a dinitrosalicylic acid (DNS) solution.
Reducing power was quantified using the DNS assay with maltotriose as a
standard (0.5-3.5 mg mL1).  -Glucosidase
activity was determined according to the method of Okita et al. (1979)
by mixing 100 µL of desalted extract, 50 mM citrate-HCl
(pH 6.0), and 10 mM maltose (1 mL final assay volume), and
incubating at 37°C for 60 min. The reaction was terminated by adding
3 mL of DNS solution. Reducing power was quantified using the DNS assay
with Glc as a standard (1-4 mg mL1). Total
amylase activity was determined according to the method of Doehlert and
Kuo (1990) by mixing 100 µL of desalted extract, 50 mM
citrate-HCl (pH 6.0), and 2.5% (w/v) boiled soluble potato starch (1 mL final assay volume) and incubating at 37°C for 60 min. The
reaction was terminated by adding 3 mL of DNS solution. Reducing power
was quantified with the DNS assay using maltose as a standard (1-4 mg
mL1).
Comparing Enzyme Activities and Metabolite Contents
To conduct these experiments, it was necessary to express the
metabolite contents and enzyme activities in a form that would allow a
comparison between treatments. Expressing them per unit of dry weight
was unsuitable as a basis of comparison because ovary sugars and starch
accounted for 75% of the ovary dry weight at pollination, and these
intermediates varied markedly with w. Ovary
fresh weight was similarly unsatisfactory because water was 90% of the
fresh weight and varied with w. Therefore, all measurements were expressed on a per-ovary basis.
Isotope Labeling
To determine whether the carbon in the Suc fed to the stems was
transported to the ovaries, we fed Suc from sugar beet instead of
sugarcane. The sugar beet Suc had been synthesized by C-3
photosynthesis and was impoverished in 13C, but
the dry matter of the maize ovaries had been synthesized in the parent
plant by C-4 photosynthesis and did not show this impoverishment. The
13C was defined by the  13C ratio = 1,000 × (Rsample Rstandard)/ (Rstandard),
where R is 13C/12C and the standard
is Pee Dee Belemnite. The 13C ratio for the sugar beet
Suc was 25 and for the ovaries was 12.
Therefore, transport of infused Suc carbon to the ovaries could be
detected by a 13C ratio below 12.
Sugar beet Suc was obtained from a market and fed to stems as above.
Individual ovaries were sampled from developing ears, freeze-dried,
weighed, sealed in aluminum tinfoil cups, and the
13C ratio of the dry matter was determined in
a ratio mass spectrometer after combustion to
CO2. It should be noted that the arrival of sugar
beet carbon at the ovaries diluted the maize dry mass already present,
and the experiment thus detected the arrival of substrate quantities of
Suc carbon.
Microscopy
Ovaries were removed from developing ears and sectioned in the
midplane parallel to the long axis of the ear. The sections were
stained with I2-KI solution (0.20% [w/w]
I2 and 0.53% [w/w] KI), briefly rinsed with
deionized water, and viewed under a dissecting microscope.
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RESULTS |
Ovary Starch Dynamics
Starch occupied the ovary tissues around the nucellus (ovary wall)
and surrounded the vascular tissues at the base of the ovary before the
young pistils were pollinated (Fig. 2A).
There was no starch in the nucellus. As the ovary enlarged during and after pollination, starch remained in the same tissues (Fig. 2, B-D).
By d 10, the nucellus and starch-containing tissues in the wall were
pushed to the periphery of the ovary by the developing endosperm (Fig.
2, C and D). At d 12, starch began to be deposited in the endosperm,
and endosperm activity began to dominate the biosynthetic activity of
the ovary (Fig. 2, D and E). The final starch content of the mature
kernel was large compared with that in the ovary at the time of
pollination (Fig. 2F).
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| Figure 2.
Starch location and amount in maize ovaries and
developing kernels. A, Ovary on d 5 before pollination. Starch
(region stained black) is located at the ovary base around the phloem
and in the ovary tissues around the nucellus (ovary wall). The nucellus
occupies the center of the ovary and is starchless. The embryo sac is
not present in this section. Silk normally attached to the ovary apex
has been removed. W, Ovary wall; N, nucellus, V, vein. B, Ovary at
pollination on d 0. The location of starch is similar to that in A. Magnification is the same as in A. C, Ovary on d 10 after pollination.
Endosperm occupies most of the ovary interior, and the nucellus is at
the periphery, inside the starch-containing ovary wall. D, Ovary on d
12. Starch deposition is beginning in the endosperm. The nucellus is at
the periphery, inside the starch-containing ovary wall. Magnification
is the same as in C. E, Ovary on d 23. Starch is being rapidly
deposited in the endosperm. The nucellus and ovary wall are forming the
outer covering of the caryopsis. F, Starch content of the ovaries as
they develop into mature caryopses. Bars in A, C, and E indicate 1 mm
for all micrographs.
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When we subjected plants to the low w
treatment starting on d 5 (Fig. 1), the leaf
w decreased (Table
I). By d 0, it reached about 1.3 MPa
and net photosynthesis decreased to 26% to 34% of the control rate
(Table I). The plants were hand-pollinated on that day. After allowing
1 d for fertilization, the plants were rewatered at the end of d
1, and w and photosynthesis returned to
control levels by the next day (d 2, data not shown; for examples in
similar experiments, see Boyle et al., 1991b; Zinselmeier et al.,
1995a). Using light microscopy, Westgate and Boyer (1986) showed that
embryos were present but were aborted by this treatment. We judged
whether embryos developed according to the number of kernels that
developed. The short exposure to low w
decreased the number of kernels to 7.4% of the control at high
w (Table I) and thus disrupted embryo
development.
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Table I.
Kernel number, leaf w, and net
photosynthesis in maize deprived of water around the time of
pollination
w and photosynthesis were measured on the day of
pollination (d 0 in Fig. 1) on the third leaf at the top of the plant
3 h into the photoperiod. Kernel number was determined at
maturity. Adequate water was supplied to controls, and water was
withheld from low w plants for 5 d prior to the
measurements (see Fig. 1). Data are means ± SE of
four to six plants.
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Suc was fed to some of the stems starting on d 4 and continuing until
the day of pollination (d 0). With Suc infusion, embryo development
continued in 62% of the ovaries (Table I). Sufficient Suc was fed to
replace the photosynthate not being produced by the leaves. The leaves
normally produced 5 g of dry mass per day at pollination (Jurgens
et al., 1978), and 6.8 g was the amount fed. The feeding during
low w allowed embryo development to continue, but the major part of kernel development occurred afterward, when photosynthesis had returned to control levels. Thus, the feeding partially prevented the disruption of embryo development by low w, and the parent plant continued to grow the
kernel after water was resupplied.
The low leaf w caused the ovaries to lose
their starch (Fig. 3). Starch had
disappeared from the basal tissue and most of the ovary wall by d 0 and
2 of the low w treatment (Fig. 3, I and J),
while the controls showed a normal starch distribution (Fig. 3, C and
D). Feeding Suc to the stems during the low w treatment partly maintained the starch in the ovaries (Fig. 3, F and
G). Quantitative analyses confirmed the starch depletion at low
w and the partial maintenance by Suc feeding
(Fig. 3K). The starch depletion was reflected in a low accumulation of
ovary dry matter that was partly reversed by Suc feeding (Fig.
4).
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| Figure 3.
Starch location and amount in maize ovaries around
the time of pollination, showing the effects of Suc fed to the stems at
low w. A, Ovary on d 5 before pollination. Bar
indicates 1 mm for all micrographs. B, Ovary on d 2 before
pollination. C, Ovary on d 0 at pollination. D, Ovary on d 2 after
pollination. Note that starch (region stained black) is present at the
ovary base around the phloem and in the ovary tissues around the
nucellus. E through G, Same as B through D except water was withheld on
d 5 before pollination and resupplied on d 1 after pollination, and
stems were fed with Suc solution starting on d 4. Starch location is
unchanged from B through D. H through J, Same as E through G except no
Suc was fed to the stems. In I and J, starch has nearly disappeared
from the ovary base around the phloem and the ovary tissues around the
nucellus. K, Starch contents of ovaries in micrographs. The black bar
on the x axis shows time when water was withheld from
the soil and the white bar shows time when Suc was infused into the
stems, as in Figure 1. Data are means ± SD of six
plants.  , Control;  , low w + Suc infusion;  ,
low w.
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| Figure 4.
Ovary dry mass showing effects of Suc fed to the
stems at low w. The black bar on the x
axis shows time when water was withheld from the soil and the white bar
shows time when Suc was infused into the stems, as in Figures 1 and 3.
Data are means ± SD for six plants. , Control;
, low w; , low w + Suc infusion.
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Instead of feeding Suc from a C-4 plant (sugarcane) to the maize stems
(also C-4), we fed C-3 Suc from a C-3 plant (sugar beet) in order to
determine whether the natural label in the C-3 sugar reached the
ovaries. The lower 13C ratio of the Suc from
sugar beet caused the 13C ratio to decrease in
the ovaries (Fig. 5). The decrease could be observed at the first sampling in fed plants at low
w. Similarly feeding controls also decreased
the 13C ratio. Without Suc infusion, the
13C ratio did not decrease at low
w or in the controls, indicating that the
label in the stem-fed Suc was being rapidly transported to the ovaries.
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| Figure 5.
Ovary 13C ratios
showing effects of sugar beet Suc fed to the stems at low
w around the time of pollination. The black bar on the
x axis shows time when water was withheld from the soil
and the white bar shows time when Suc was infused into the stems, as in
Figures 1 and 3. Data are means ± SD for three
plants. , High w;  , high w + sugar
beet Suc; , low w; , low w + sugar
beet Suc.
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Suc feeding did not change leaf w or
photosynthesis (Table I), and it also had no effect on ovary
w (Fig. 6A).
However, it altered the components of ovary w.
There was only a slight decrease in ovary s in
the unfed plants, but a large decrease in ovary
s in the fed plants (Fig. 6B). As a result,
the ovary p decreased substantially at low
w but was essentially maintained in the fed
plants (Fig. 6C). The difference in s of the
fed and unfed plants showed that stem feeding increased the solute
concentration in the ovaries, and p improved
as a result.
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| Figure 6.
Effects of withholding water and feeding Suc
to the stems around the time of pollination on w (A),
s (B), and p (C). On the x
axis, the black bar shows time when water was withheld from the soil
and the white bar shows time when Suc was infused into the stems, as in
Figures 1 and 3. Data are means ± SD for six plants.
, Control; , low w; , low w + Suc infusion.
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Regulation of Starch Biosynthesis within the Ovaries
During the early development of maize ovaries, Suc metabolism
starts with acid invertase (Shannon and Dougherty, 1972; ap Rees, 1984;
Doehlert and Felker, 1987; Xu et al., 1995), and mutants of this enzyme
block much of the development (Miller and Chourey, 1992; Cheng et al.,
1996). Ovary Suc levels decreased at low w. When we fed Suc to the stems, the ovary Suc increased to control levels
(Fig. 7A). The activities of soluble and
insoluble acid invertases were markedly decreased at low ovary
w compared with the controls, and Suc feeding
caused a slight recovery (Fig. 7, B and C). The reducing sugars were
decreased at low w, and Suc feeding also
caused a partial recovery (Fig. 7D). The reducing sugars are the
product of the invertase reaction, and their lack of full recovery with
stem feeding was apparent when contrasted with the recovery of
substrate Suc (compare Fig. 7, A and D, on d 0 and 2).
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| Figure 7.
Ovary Suc content, acid invertase activity, and
reducing sugar content showing effects of withholding water and feeding
Suc to the stems around the time of pollination. A, Suc (invertase
substrate). B, Soluble acid invertase activity. C, Insoluble acid
invertase activity. D, Reducing sugars (invertase product). The black
bar on the x axis shows time when water was withheld
from the soil and the white bar shows time when Suc was infused into
the stems, as in Figures 1 and 3. Data are means ± SD
for six plants. , Control; , low w; , low
w + Suc infusion.
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In all of the succeeding pools of phosphorylated intermediates for
starch biosynthesis (Glc-6-P, Fru-6-P, and Glc-1-P), a similar pattern
of depletion at low w and partial recovery
with Suc infusion was observed (Fig. 8).
For UDP-Glc, which is interconvertible to ADP-Glc (the likely substrate
for starch biosynthesis), there was a marked depletion at low
w (Fig. 9A) and
only a partial recovery when Suc was fed. As an internal control, we
measured the pool of Pi. Because the pool was large and mostly
inorganic, it would be expected to be less affected than the pools of
phosphorylated intermediates. Figure 9B shows that both the low
w treatment and Suc feeding had only a slight
effect.
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| Figure 8.
Ovary contents of Glc-6-P (A), Fru-6-P (B), and
Glc-1-P (C) showing effects of withholding water and feeding Suc to the
stems around the time of pollination. The black bar on the
x axis shows time when water was withheld from the soil
and the white bar shows time when Suc was infused into the stems, as in
Figures 1 and 3. Data are means ± SD for six plants.
, Control; , low w; , low w + Suc infusion.
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| Figure 9.
Ovary contents of UDP-Glc (A), and Pi (B) showing
effects of withholding water and feeding Suc to the stems around the
time of pollination. The black bar on the x axis shows
time when water was withheld from the soil and the white bar shows time
when Suc was infused into the stems, as in Figures 1 and 3. Data are
means ± SD for six plants. , Control; , low
w; , low w + Suc infusion.
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Regardless of the availability of intermediates in starch biosynthesis,
the concentration of phosphate and certain phosphorylated intermediates
can affect starch biosynthesis. In leaf chloroplasts, starch can
accumulate and is regulated by the concentration of 3-phosphoglyceric
acid, which stimulates synthesis, and Pi, which inhibits it
(Plaxton and Preiss, 1987; Preiss, 1988). We analyzed 3-phosphoglyceric acid and expressed the concentration
according to the total water content of the ovaries (Fig.
10A). The concentration was low and
became somewhat lower at low w, and was fully
restored by feeding Suc to the stems. On the other hand, the Pi
concentration expressed similarly was 6 to 8 mM in the
controls, which is in the regulatory range (Fig. 10B). The
concentration increased to about 11 mM as the ovaries
dehydrated. The concentration returned partially to control
levels when Suc was fed to the stems.
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| Figure 10.
Ovary concentrations of 3-phosphoglyceric
acid (A) and Pi (B) showing effects of withholding water and
feeding Suc to the stems around the time of pollination. Concentrations
are based on the total water content of the ovaries. The black bar on
the x axis shows time when water was withheld from the
soil and the white bar shows time when Suc was infused into the stems,
as in Figures 1 and 3. Data are means ± SD for six
plants. , Control; , low w; , low
w + Suc infusion.
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Starch Breakdown within the Ovaries
The total amylase activity in the ovaries was large (Fig.
11). Because the assays indicated the
maximum velocity for the enzyme, they do not represent in vivo
activity, but the measured activity was sufficient to break down the
ovary starch pool in about 20 min and thus was consistent with the
starch breakdown we observed in vivo. The activity was moderately
decreased at low w, and there was a slight
recovery when Suc was fed. Activities of debranching enzyme and
-glucosidase were below the limits of reliable detection (data not
shown).
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| Figure 11.
Amylase activity in ovaries showing effects of
withholding water and feeding Suc to the stems around the time of
pollination. The black bar on the x axis shows time when
water was withheld from the soil and the white bar shows time when Suc
was infused into the stems, as in Figures 1 and 3. Data are means ± SD for six plants. , Control; , low
w; , low w + Suc infusion.
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DISCUSSION |
Ovary Starch Dynamics
Starch was a major constituent of maize ovaries before pollination
and therefore before any endosperm was present. It was readily
mobilized, in contrast to endosperm starch, which does not turn over
until the kernel matures, dehydrates, and resumes development during
germination. The ovary pool nearly disappeared when
w became low enough to inhibit photosynthesis,
whereupon embryo development was irreversibly arrested. The
disappearance of starch indicated the end of sugar availability.
Because this was lethal, the starch disappearance was the central event
controlling embryo development and thus kernel number.
Figure 12 shows the likely path for Suc
delivery and starch mobilization when photosynthesis was inhibited
at low w. Sugars were delivered to the base of
the ovary, where they were incorporated into expanding ovary structures
or starch around the veins and nucellar tissue. When photosynthesis was
inhibited, the starch broke down and allowed ovary development to
continue with the released sugars. When the starch was depleted, the
embryos aborted and ovary development ceased.
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| Figure 12.
Dynamics of ovary starch leading to embryo
abortion at low w. A, Suc is delivered to the tissues
below the ovary (large arrow) and either used in developing structures
or stored as starch around the veins and in the ovary tissues around
the nucellus (small arrows). B, With inhibited photosynthesis, Suc
delivery is curtailed ( ), starch is mobilized (small
arrows), and ovary development continues. C, When the starch is
depleted, ovary development ceases irreversibly. The final kernel
number is diminished accordingly.
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Feeding substrate quantities of Suc to the stems of the parent plant
prevented some of the starch disappearance and some of the arrest in
embryo development. However, we were unable to rescue all of the
embryos, indicating that other factors also were important. One of
these factors could have been transport, but labeled Suc was delivered
from the stems to the ovaries and resulted in substantial increases in
solute concentrations inside the ovaries, including Suc. Therefore, an
internal block in starch biosynthesis was more likely to have been the
cause. The sugar and phosphorylated intermediates likely to be
involved are shown in Figure 13, and
all of them were depleted at low w.
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| Figure 13.
Likely pathway of starch biosynthesis in maize
ovaries around the time of pollination. Bold print shows that Suc
accumulated with Suc feeding at low w. Light print shows
that all other metabolites were depleted despite Suc feeding at low
w. The depletion indicates that a block occurs at the
first step in the pathway (acid invertase).
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Feeding Suc to the stems returned Suc in the ovaries to control levels,
but did not fully return the other pools (Fig. 13). As a consequence,
carbon flow in the ovaries was restricted at the first step of the
path, impoverishing all of the downstream pools. This block thus
regulates starch synthesis to a considerable degree. The first step is
mediated by invertase (Shannon and Dougherty, 1972; ap Rees, 1984;
Doehlert and Felker, 1987; Miller and Chourey, 1992; Xu et al., 1995;
Cheng et al., 1996), and its low activity at low
w supports the concept that it restricted
carbon flow.
These results indicate that the young ovaries were markedly dependent
on carbon flow, and a short time without carbon was lethal. Kernel
number at maturity was determined by this flow. In maize grown under
our conditions, stored reserves were sometimes low and were not drawn
upon when young ovaries were developing at low
w (Westgate and Boyer, 1985a). Decreasing
photosynthesis thus had an immediate inhibitory effect on the sugar
stream to the ovaries (Westgate and Boyer, 1986; Boyle et al., 1991b;
Schussler and Westgate, 1995). When the decreased sugar stream was
coupled with decreased carbon processing inside the ovaries, kernel
number was markedly decreased. Later in kernel development, sugar
reserves were abundant in the stems and leaves of the parent plant, and the sugars were mobilized to support kernel development when
photosynthesis was inhibited (McPherson and Boyer, 1977; Jurgens et
al., 1978; Westgate and Boyer, 1985a). At that time, kernel number was
unaffected and the kernels matured with a smaller size because of the
diminished amount of total carbon. This difference in reserve
mobilization appears to explain why the kernel number is determined
early but size is determined later in reproductive development.
The delivery of Suc was the key to these findings. The rapidity of
uptake of the fed solution suggests that absorption was by the exposed
xylem vessels, with the leaves as the first destination. Suc was
probably loaded into the phloem by the leaves and delivered to other
parts of the plant. It was essential to feed quantities of Suc
comparable to those produced by photosynthesis in the whole parent
plant. In earlier experiments, we varied the amount of fed Suc but
failed to rescue the developing kernels with lesser amounts (data not
reported). Assuming 500 ovaries per plant and a weight gain of 1 mg in
each ovary because of Suc feeding, the amount of Suc needed to account
for Suc delivery to the ovaries was only 0.5 g. About 6.8 g
of Suc was fed each day to the plant. We assume that such large
quantities were required because the phloem delivered Suc to various
tissues, and the ovaries were only one of the beneficiaries.
Although label in fed Suc was delivered to the ovaries and there was
elevated Suc in the ovary tissues, we do not know whether the label was
delivered to the same sites that it would be during normal
photosynthesis. The analyses required the ovaries to be removed from
the ear, and some of the basal tissue inevitably remained attached to
the ovary. Because Suc was delivered to the base of the ovaries by the
phloem, which is not directly attached to the ovary interior, it is
possible that some of the delivered Suc remained in the basal tissues.
Schussler and Westgate (1991) showed that ovaries isolated from maize
ears at low w absorbed Suc less rapidly than
controls. Some of the Suc could have arrived at the phloem-unloading
sites and been detected in the label and Suc assays, but may not have
been delivered to particular ovary tissues where invertase activity was
controlled.
Water Relations and Starch Regulation
The Suc feeding had no effect on the w of
the ovaries or leaves, probably because the s
of the fed Suc (1.1 MPa) was so similar to the
w of these organs (1 and 1.3 MPa in
ovaries and leaves, respectively, at pollination). Also, the volume of fed solution was small (45 mL d1) compared with
water flux through the plant at low w (150 mL d1). Boyle et al. (1991b) and Zinselmeier et
al. (1995a) found no effect of the fed solution on leaf
w or photosynthesis with similar treatments.
Nevertheless, there was a marked effect on the components of
w. The s of the
ovaries decreased more at low w when Suc was fed than when it was not. For each ovary, about 0.2 mg of new Suc and
0.3 mg of new reducing sugar accumulated because of the feeding and was
present in about 20 mg of the total ovary water. These new solutes
represent a calculated change in s of 0.31 MPa, and the feeding-induced change that was measured was about 0.35
MPa. This osmotic adjustment (Meyer and Boyer, 1972; Morgan, 1984) was
thus dependent on Suc feeding. In effect, we were able to "turn on"
osmotic adjustment in the ovaries by feeding Suc to the stems. The
ovaries were more turgid as a result. Whether the higher turgidity
caused more growth is uncertain, but the accumulation of these large
amounts of sugars during osmotic adjustment indicated that additional
substrate was present for ovary metabolism. In agreement with these
experiments, Westgate and Boyer (1985b) could not find osmotic
adjustment in developing silks of pistillate florets from unfed plants
at low w.
Because the ovaries of unfed plants showed little osmotic adjustment
and had low turgor, their water content was diminished at low
w. This had the effect of concentrating solute
in the cells. One consequence was an increase in the concentration of Pi when ovaries had low w (even though the
content of Pi was similar for the treatments). While the cellular
distribution of Pi was not investigated, most of it was likely to have
been in the vacuoles of the ovary cells, and its inhibitory action
takes place in the plastids. Nevertheless, the high overall
concentration of Pi at low w is consistent
with a possible decrease in starch biosynthesis by feedback inhibition
similar to that described by Plaxton and Priess (1987) and Priess
(1988). The effect was partially reversed when Suc was fed at low
w and thus was consistent with the reversal of
starch depletion in these plants.
Regulation of Kernel Number
The mechanisms controlling kernel number are of considerable
interest. In their extensive review of how water affects crop reproduction, Salter and Goode (1967) concluded that water supply is
the most important to reproduction during the early stages, around
pollination and early fruit formation, when seed and fruit number are
determined. However, they cited little information about the mechanisms
of the losses. It is often proposed that low w
at these stages can prevent pollination or cause pollen sterility in
maize (Lonnquist and Jugenheimer, 1943; Herrero and Johnson, 1981;
Bassetti and Westgate, 1993). Although this clearly can occur (for
review, see Saini, 1997), it was not a factor here. We pollinated by
hand to ensure that pollination occurred. We showed that pollen
collected from plants at low w was viable and
could maintain viability when it was highly desiccated (Westgate and
Boyer, 1986). We pollinated when w was at its
lowest and dehydration of the plant was the most severe. Zygotes could
be seen in micrographs of the arrested ovaries (Westgate and Boyer, 1986). Therefore, pollination and fertilization were not factors, and
the block in kernel development occurred afterward, while the zygotes
were developing.
These results indicate that pollination and fertilization were less
affected than embryo and ovary development as the sugar stream
diminished at low w around the time of
pollination. It seems that mature pollen may have had its own reserves,
as observed by Sheoran and Saini (1996) in rice, and that reserves were
present in the embryo sac, although we did not explore either of these possibilities. If so, the success of fertilization may have depended on
these local reserves, while the susceptibility of embryo development may have depended on starch reserves around the nucellus and in the
ovary basal tissues.
Other factors may also contribute to the control of kernel number. Low
w causes increased ABA concentrations in
plants. Saini (1997) pointed out that infusing ABA into stems of barley
caused pollen abortion, and Morgan (1980) showed that spraying ABA on wheat plants had a similar effect. It is possible that ABA causes stomatal closure and decreases photosynthesis, and if this is the case,
then ABA might have its influence through an effect on carbon
availability similar to that seen here.
| |
FOOTNOTES |
1
This work was supported by the U.S. Department
of Agriculture National Research Initiative Competitive Grants Program
(grant nos. 91-37100-6617 and 94-37100-0753).
2
Present address: Pioneer Hi-Bred International,
7300 NW 62nd Avenue, Johnston, IA 50131-1004.
3
Present address: Department of Agronomy, Taegu
University, 15 Naeri, Jinyang, Kyongsan, Kyongbuk, Korea 713-714.
*
Corresponding author; e-mail boyer{at}udel.edu; fax
302-645-4007.
Received February 1, 1999;
accepted May 26, 1999.
| |
ACKNOWLEDGMENTS |
We thank Dr. An-Ching Tang for help with some of the artwork,
Larry Giles for measuring 13C ratios,
Elizabeth Oelke for a gift of sugar beet Suc, and Dr. George Singletary
for suggestions about the storage of maize pollen.
| |
LITERATURE CITED |
ap Rees T
(1984)
Sucrose metabolism.
In
DH Lewis,
eds, Storage Carbohydrates in Vascular Plants.
Cambridge University Press, London, pp 53-73
Bassetti P,
Westgate M
(1993)
Water deficit affects receptivity of maize silks.
Crop Sci
33:
279-282
[Abstract/Free Full Text]
Boyer JS
(1995)
Measuring the Water Status of Plants and Soils.
Academic Press, San Diego
Boyle MG,
Boyer JS,
Morgan PW
(1991a)
Stem infusion of maize plants.
Crop Sci
31:
1241-1245
[Abstract/Free Full Text]
Boyle MG,
Boyer JS,
Morgan PW
(1991b)
Stem infusion of liquid culture medium prevents reproductive failure of maize at low water potentials.
Crop Sci
31:
1246-1252
[Abstract/Free Full Text]
Caspar T
(1994)
Genetic dissection of the biosynthesis, degradation, and biological functions of starch.
In
EM Meyerowitz,
CR Somerville,
eds, Arabidopsis.
Cold Spring Harbor Laboratory, Cold Spring Harbor, NY, pp 913-936
Cheng WH,
Taliercio EW,
Chourey PS
(1996)
The miniature1 seed locus of maize encodes a cell wall invertase required for normal development of endosperm and maternal cells in the pedicel.
Plant Cell
8:
971-983
[Abstract]
Doehlert DC,
Felker FC
(1987)
Characterization and distribution of invertase activity in developing maize (Zea mays L.) kernels.
Physiol Plant
70:
51-57
Doehlert DC,
Kuo TM
(1990)
Sugar metabolism in developing maize kernels of starch deficient endosperm mutants of maize.
Plant Physiol
92:
990-994
[Abstract/Free Full Text]
Fader GM,
Koller HR
(1985)
Seed growth rate and carbohydrate pool sizes of the soybean fruit.
Plant Physiol
79:
663-666
[Abstract/Free Full Text]
Handel EV
(1968)
Direct microdetermination of sucrose.
Anal Biochem
22:
280-283
[CrossRef][ISI][Medline]
Hanft JM,
Jones RJ
(1986)
Kernel abortion in maize. I. Carbohydrate concentration patterns and acid invertase activity of maize kernels induced to abort in vitro.
Plant Physiol
81:
503-510
[Abstract/Free Full Text]
Helmerhorst E,
Stokes GB
(1980)
Microcentrifuge desalting: a rapid, quantitative method for desalting small amounts of protein.
Anal Biochem
104:
130-135
[CrossRef][ISI][Medline]
Herrero MP,
Johnson RR
(1981)
Drought stress and its effect on maize reproductive systems.
Crop Sci
21:
105-110
Hoagland DR,
Arnon DI
(1950)
The water culture method for growing plants without soil.
Calif Agric Exp Stn Circ
347:
1-32
Jelitto T,
Sonnewald U,
Willmitzer L,
Hajirezeai M,
Stitt M
(1992)
Inorganic pyrophosphate content and metabolites in potato and tobacco plants expressing E. coli pyrophosphatase in their cytosol.
Planta
188:
238-244
[CrossRef][ISI]
Jurgens SK,
Johnson RR,
Boyer JS
(1978)
Dry matter production and translocation in maize subjected to drought during grain fill.
Agron J
70:
678-682
[Abstract/Free Full Text]
Keppler D, Decker K (1984) Uridine 5 -diphosphoglucose and uridine
5-diphosphogalactose. In HU Bermeyer, ed, Methods of
Enzymatic Analysis, Vol VII: Metabolites 2. Verlag Chemie, Weinheim,
Germany, pp 524-530
Kiesselbach TA (1949) The Structure and Reproduction of Corn.
University of Nebraska Press, Lincoln
Kramer PJ,
Boyer JS
(1995)
Water Relations of Plants and Soils.
Academic Press, San Diego
Lonnquist JH,
Jugenheimer RW
(1943)
Factors affecting the success of pollination in corn.
J Am Soc Agron
35:
923-933
McPherson HG,
Boyer JS
(1977)
Regulation of grain yield by photosynthesis in maize subjected to a water deficiency.
Agron J
69:
714-718
[Abstract/Free Full Text]
Meyer RF,
Boyer JS
(1972)
Sensitivity of cell division and cell elongation to low water potentials in soybean hypocotyls.
Planta
108:
77-87
[CrossRef][ISI]
Miller ME,
Chourey PS
(1992)
. The maize invertase-deficient miniature-1 seed mutation is associated with aberrant pedicel and endosperm development.
Plant Cell
4:
297-305
[Abstract/Free Full Text]
Morgan JM
(1980)
Possible role of abscisic acid in reducing seed set in water-stressed wheat plants.
Nature
289:
655-657
Morgan JM
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Abstract
Introduction