Download Callose Deposition Is Responsible for Apoplastic

Plant Physiol. (1998) 118: 83–90
Callose Deposition Is Responsible for Apoplastic
Semipermeability of the Endosperm Envelope of
Muskmelon Seeds1
Kyu-Ock Yim and Kent J. Bradford*
Department of Vegetable Crops, University of California, Davis, California 95616–8631
Semipermeable cell walls or apoplastic “membranes” have been
hypothesized to be present in various plant tissues. Although often
associated with suberized or lignified walls, the wall component
that confers osmotic semipermeability is not known. In muskmelon
(Cucumis melo L.) seeds, a thin, membranous endosperm completely encloses the embryo, creating a semipermeable apoplastic
envelope. When dead muskmelon seeds are allowed to imbibe,
solutes leaking from the embryo are retained within the envelope,
resulting in osmotic water uptake and swelling called osmotic distention (OD). The endosperm envelope of muskmelon seeds stained
with aniline blue, which is specific for callose (b-1,3-glucan). Outside of the aniline-blue-stained layer was a Sudan III- and IVstaining (lipid-containing) layer. In young developing seeds 25 d
after anthesis (DAA) that did not exhibit OD, the lipid layer was
already present but callose had not been deposited. At 35 DAA,
callose was detected as distinct vesicles or globules in the endosperm envelope. A thick callose layer was evident at 40 DAA,
coinciding with development of the capacity for OD. Removal of
the outer lipid layer by brief chloroform treatment resulted in more
rapid water uptake by both viable and nonviable (boiled) seeds, but
did not affect semipermeability of the endosperm envelope. The
aniline-blue-staining layer was digested by b-1,3-glucanase, and
these envelopes lost OD. Thus, apoplastic semipermeability of the
muskmelon endosperm envelope is dependent on the deposition of
a thick callose-containing layer outside of the endosperm cell walls.
The presence of semipermeable apoplastic “membranes”
has been reported in sugarcane stems (Welbaum et al.,
1992), developing seeds (Bradford, 1994), and roots
(Steudle and Peterson, 1998). For example, sugarcane stems
accumulate high apoplastic Suc concentrations but the xylem stream within the vascular bundles is virtually free of
solutes (Welbaum and Meinzer, 1990). Welbaum et al.
(1992) demonstrated that a semipermeable apoplastic barrier exists between the vascular bundles and storage parenchyma apoplast. Solutes and water are transported into
developing seeds primarily through the phloem, and high
apoplastic solute concentrations are involved in maintaining phloem unloading in sink tissues (Wolswinkel, 1992).
At the same time, excess water delivered via the phloem is
1
This work was supported by the U.S. Department of Agriculture Binational Agricultural Research and Development Fund
(grant no. US-2422-94).
* Corresponding author; e-mail [email protected]; fax
1–530 –752– 4554.
recycled back to the mother plant via an apoplastic pathway (Oparka and Gates, 1981; Peoples et al., 1985). Strategically located semipermeable apoplastic membranes may
prevent solute movement by mass flow back to the mother
plant, retaining solutes within the unloading regions of
developing seeds (Bradford, 1994). The tetrazolium ion,
which is used for vital staining, does not penetrate into
most grass seeds (Brown, 1907) or through the inner coat of
watermelon (Thornton, 1968), tomato, or pepper seeds
(Beresniewicz et al., 1995b). Similarly, the lanthanum ion, a
water-soluble heavy metal, is accumulated at the inner
seed coat adjacent to the endosperm in tomato and pepper
(Beresniewicz et al., 1995b; Taylor et al., 1997). In an examination of seeds of 500 species from more than 40 families,
semipermeability was very common, except in the Leguminosae and some genera of Cistaceae and Cruciferae
(Gola, 1905, cited by Kotowski, 1927). The Casparian strip
in the root endodermis has been considered to be essentially impermeable to both water and solutes, but Steudle
and Peterson (1998) have proposed that a semipermeable
Casparian strip is required to explain water and solute
movement through roots.
Suberized or lignified walls are often associated with
apoplastic permeability barriers (O’Brien and Carr, 1970;
Cochran, 1983; Welbaum and Bradford, 1990; Jacobsen et
al., 1992; Welbaum et al., 1992). Suberin has been suggested
as the semipermeable material in seeds of corn (Johann,
1942), Johnsongrass (Harrington and Crocker, 1923), tomato, and pepper (Beresniewicz et al., 1995a), whereas
cutin may serve this role in leek seeds (Taylor et al., 1997).
In pollen grains, Heslop-Harrison (1964) suggested that
callose in the pollen mother cells acts as a “molecular
sieve” to isolate the pollen cells from maternal compounds.
Nonetheless, although many histochemical studies have
attempted to relate the composition of semipermeable
cell walls to their function, there is no direct demonstration that specific wall components are responsible for
semipermeability.
In muskmelon (Cucumis melo L.) seeds the embryo is
completely enclosed by a membranous envelope that has
been described as consisting of a layer of endosperm cells
and several layers of thick-walled perisperm cells (originating from the nucellus); it was therefore termed the
“perisperm envelope” (Singh, 1953; Welbaum and BradAbbreviations: DAA, days after anthesis; OD, osmotic distention; SEM, scanning electron microscopy.
83
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84
Yim and Bradford
ford, 1990). However, as will be shown here, the envelope
surrounding the muskmelon embryo is composed entirely
of a single layer of endosperm cells covered by a thick,
noncellular external layer. We will therefore refer to this
tissue as the “endosperm envelope.” When dead muskmelon seeds are allowed to imbibe, solute leakage from the
embryo generates an osmotic gradient across the semipermeable endosperm envelope, resulting in water uptake and
swelling, known as OD (Welbaum and Bradford, 1990).
The intact endosperm envelope can shrink and swell repeatedly in response to external osmotic conditions without significant loss of solutes (Welbaum and Bradford,
1990). A semipermeable envelope is present in many species of Cucurbitaceae and Compositae (e.g. lettuce), as
shown by the development of OD in hydrated dead seeds
(Hill and Taylor, 1989).
In muskmelon, endosperm envelope semipermeability
was maintained after various treatments that killed the
embryos (freezing and thawing the hydrated seeds, boiling, autoclaving, aging, and 100% methanol), whereas the
semipermeability was lost after strong acid or alkaline
treatments (Welbaum and Bradford, 1990). A Sudan IVstaining (lipid-containing) layer was detected on the outer
surface of the endosperm envelope and was assumed to be
involved in semipermeability, in analogy with the presumed role of suberized cell walls in other tissues (Welbaum and Bradford, 1990). However, no direct evidence
was available to determine which component of the endosperm envelope is responsible for semipermeability. We
demonstrate here that an extracellular layer composed primarily of callose is entirely responsible for the semipermeable properties of the muskmelon endosperm envelope.
MATERIALS AND METHODS
Muskmelons (Cucumis melo L. cv Top Mark) were field
grown at the University of California (Davis) in 1995 and
1996, and seeds were harvested, dried, and stored at 220°C
(Welbaum and Bradford, 1988). For the seed-development
study, flowers were tagged at anthesis and harvested at 5-d
intervals from 25 to 60 DAA.
Anatomy
Intact seeds were hand sectioned, and decoated seeds
were embedded in paraffin (Jensen, 1962) and thin sectioned (10 mm) with a microtome. Decoating was done
manually with forceps without damaging the endosperm
envelope or embryo. Sections on glass slides were stained
with 0.05% aniline blue in 0.1 m phosphate buffer (pH 8.2)
or a Sudan III and IV mixture (equal volumes of saturated
solutions in 70% ethanol) for approximately 5 min and
observed using light microscopy. To decrease unspecific
staining by aniline blue, toluidine blue O (0.5% in 0.1 m
phosphate buffer, pH 7.0) was used after aniline blue staining. To confirm the specificity of the blue staining for
callose, mature seeds were hand sectioned, stained with
0.001% synthetic aniline blue fluorochrome (Sirofluor, Biosupplies Australia, Parkville, Australia) for 20 min, and
observed with a fluorescence microscope (Axiovert 100,
Plant Physiol. Vol. 118, 1998
Zeiss) with a fluorescein isothiocyanate filter (excitation,
470 nm; emission, 515 nm; beam splitter, 505DCLP).
For SEM, seeds were freeze-dried for 3 d and mounted
on aluminum stubs. For cross-sectional views, endosperm
envelopes fractured during freeze-drying were positioned
with a fracture plane on an aluminum stub. Samples were
sputter coated with 30-nm gold (SEM coating system, BioRad) and observed with a scanning electron microscope
(model DS 130, International Scientific Instruments, Top
Con Technologies, Inc., Paramus, NJ) at 10 kV.
OD and Water-Imbibition Kinetics
To induce OD, decoated seeds were killed by boiling
them in water for 3 min, and were then incubated on
water-saturated blotter paper. To observe the imbibition
kinetics, viable or boiled decoated seeds with or without
chloroform treatment (see below) were incubated on watersaturated blotter paper in Petri dishes at 30°C. At frequent
intervals, seeds were briefly blotted on lint-free tissues for
30 s, weighed, and returned to the Petri dish. Observations
were terminated when live seeds completed germination
(radicle emergence).
Chloroform and Enzyme Treatment
To remove the outer lipid-containing layer, decoated
seeds were dipped in chloroform for 3 min, rinsed in water,
and incubated on water-saturated germination paper overnight. To treat with b-1,3-glucanase, decoated seeds exhibiting OD (with or without chloroform treatment) were
incubated on germination paper saturated with 3 3 1024
units of endo-b-1,3-glucanase (purified from Helix pomatia;
Fluka, Buchs, Switzerland) in 0.1 m citrate-phosphate
buffer (pH 5.5). After the seeds had lost OD, they were
examined using light microscopy or SEM as described
above.
RESULTS
Anatomy of the Endosperm Envelope
Hand-sectioned muskmelon seeds showed dark-blue
staining (aniline blue) of the endosperm envelope below
the spongy tissue of the seed coat (Fig. 1A, arrow). Thin
sections of decoated, paraffin-embedded seeds showed
specific gray-blue staining with aniline blue associated
with the endosperm envelope outside of a single layer of
rectangular endosperm cells (Fig. 1B). The specificity of the
blue staining for callose observed by light microscopy was
confirmed by fluorescence microscopy using synthetic aniline blue fluorochrome (Stone et al., 1984). The endosperm
envelope showed yellow fluorescence only when stained
with this fluorochrome (Fig. 1C, arrow), whereas the testa
and thin lipid layer of the endosperm envelope showed
autofluorescence in the presence or absence of the dye
(data not shown). Only triploid nuclei were found when
the nDNA contents of entire envelope tissues were analyzed using flow cytometry, demonstrating that all living
cells originated from the endosperm (data not shown).
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Callose and Apoplastic Semipermeability
85
Figure 1. A, Cross-section of a dry muskmelon seed stained with 0.05% aniline blue and 0.5% toluidine blue O. The testa
(TE) is at the top, covering a spongy layer. The endosperm envelope (arrow) below the spongy layer shows an outer
blue-stained layer. The bar represents 10 mm. B, Thin section of paraffin-embedded, decoated dry muskmelon seed stained
with 0.05% aniline blue and a saturated mixture of Sudan III and IV. An aniline blue-staining layer (gray-blue) is adjacent
to a layer of rectangular endosperm cells (EN), and a Sudan-staining layer (orange-red) is present on top of the anilineblue-staining layer. The bar represents 10 mm. C, Similar to B, but stained with synthesized aniline blue fluorochrome and
viewed by fluorescence microscopy with a fluorescein isothiocyanate filter. The bright-yellow fluorescent band (arrow)
identifies the location of callose in the endosperm envelope. The testa (TE) shows strong autofluorescence that was not
dependent on the presence of the fluorochrome (data not shown). The bar represents 10 mm. D, Surface of the endosperm
envelope of muskmelon seeds with OD viewed by SEM. The bar represents 10 mm. E, Freeze-fractured cross-section of the
muskmelon endosperm envelope viewed by SEM. Above the inner rectangular endosperm cells (EN) is a globular, callosic
layer having a “foamy” appearance (CA). Some crushed cells (CC) appear to be present in the outer lipid-containing layer
(LL). The bar represents 2 mm. F, Removal of the lipid-containing layer by chloroform treatment. Decoated muskmelon seeds
were dipped in chloroform for 3 min and hand sectioned. Sections were stained with aniline blue and Sudan III and IV. The
Sudan-staining layer was removed but the aniline-blue-staining layer was intact after chloroform treatment (compare with
Fig. 2D). The bar represents 10 mm. G, Freeze-fractured cross-section of the chloroform-treated muskmelon endosperm
envelope viewed by SEM. The globular layer remained but the waxy outer layer was removed by the chloroform treatment.
The bar represents 2 mm.
When viewed using SEM, the surface of the endosperm
envelope was smooth and waxy in appearance without
obvious cellular structure (Fig. 1D), in contrast to the cellular outlines clearly visible in back-illuminated lightmicroscope surface views (Welbaum and Bradford, 1990).
In cross-view (Fig. 1E), the outer lipid layer appeared to
consist of plate-like or waxy sheets and possibly one or two
layers of crushed cells. Beneath this was a thick layer
composed of globules that had a “foamy” appearance and
no cellular compartmentation or contents. Below the globular layer was a single layer of rectangular endosperm
cells.
Because the outermost layer of the envelope stained
orange-red with Sudan III and IV, indicating a lipid-
containing component (Fig. 1B), we tested whether this
lipid layer could be removed by dipping decoated seeds in
chloroform. This treatment removed the Sudan-staining
layer (Fig. 1F, arrow; compare with Fig. 2D, arrow), but did
not affect aniline blue staining of the callose layer (Fig. 1F).
This was also evident in SEM, in which the chloroform
treatment removed the outer layers and exposed the globular, foam-like layer covering the endosperm cells (Fig. 1G;
compare surface with that in Fig. 1D). Because suberin is
not soluble in chloroform, the Sudan-staining material is
apparently not composed of suberin. Thus, the endosperm
envelope of muskmelon contains a single line of endosperm cells covered by a thick, globular callosic layer,
which in turn is covered by what appears to be a waxy
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86
Yim and Bradford
Plant Physiol. Vol. 118, 1998
Figure 2. A, Cross-section of developing muskmelon seeds at 25 DAA. The seeds were decoated carefully, sectioned, and stained with
aniline blue and Sudan III and IV. Only Sudan
staining (arrow) was detected on the endosperm
envelope. B, Freeze-fractured cross-section of
endosperm envelope 25 DAA viewed by SEM.
Only the plate-like or waxy appearance was
evident; the globular layer present in mature
envelopes was absent. C, Cross-section of developing muskmelon seeds at 35 DAA. Decoated seeds were hand sectioned and stained
with aniline blue. Aniline-blue-staining vesicles
were observed in the endosperm envelope.
These vesicles were not present in other developmental stages. D, Cross-section of mature
muskmelon seeds (55 DAA). Decoated seeds
were hand sectioned and stained with aniline
blue and Sudan III and IV. The orange Sudanstaining lipid layer (arrow) was observed on top
of the thick, aniline-blue-staining layer. The bar
in each panel represents 5 mm.
outer coating and some crushed cell remnants possibly
derived from the perisperm or testa.
Relationship between Semipermeability and Callose
Deposition in Developing Muskmelon Seeds
Early in development (25 DAA), muskmelon seeds are
not capable of OD, but their endosperm envelopes become
semipermeable at approximately 40 DAA (Welbaum and
Bradford, 1990; data not shown). However, endosperm
envelopes at 25 DAA can be stained with Sudan III and IV
(Fig. 2A, arrow) but not with aniline blue (Fig. 2A). Using
SEM, the globular layer was not observed at 25 DAA (Fig.
2B). At 30 DAA, the endosperm envelope did not stain with
aniline blue (data not shown), but at 35 DAA, aniline
blue-staining vesicles were evident (Fig. 2C). By 40 DAA, a
thick aniline blue-staining callosic layer was present (data
not shown), coinciding with the acquisition of semipermeability, and at 50 DAA, virtually all seeds exhibited both
semipermeability and the anatomical characteristics of mature seeds (Fig. 2D).
Role of the Lipid Layer in Water Absorption
and Semipermeability
Decoated muskmelon seeds showed a typical triphasic
imbibition time course, with an initial phase of rapid water
uptake (0–12 h) followed by a plateau phase of relatively
constant water content (12–18 h) (Fig. 3A). Radicle emergence occurred between 18 and 22 h (Fig. 3B), and was
accompanied by additional water uptake associated with
embryo growth (Fig. 3A). Boiled seeds exhibited identical
initial water- absorption kinetics, but did not attain a
water-content plateau; instead, they continued to absorb
water and became osmotically distended (Fig. 3). Removal
of the outer lipid layer from viable decoated seeds by
dipping them in chloroform hastened the initial uptake of
water and subsequent radicle emergence (Fig. 1, F and G),
but did not affect the plateau water content (Fig. 3). When
boiled seeds were treated with chloroform, water absorption was even more rapid and OD was achieved within 4 to
6 h, 12 h earlier than for boiled seeds with the lipid layer
present (Fig. 3). Thus, the outer lipid layer slows the rate of
initial water uptake, but is not required for semipermeability of the endosperm envelope.
Role of the Callose Layer in Semipermeability
Callose (b-1,3-glucan) is hydrolyzed by endo-b-1,3glucanase, so seeds exhibiting OD were incubated in commercially purified enzyme to determine whether the callose layer would be degraded and whether this would
affect semipermeability. Decoated melon seeds were boiled
and incubated on water-saturated blotting paper to induce
OD, which was maintained for at least 7 d regardless of
whether the outer lipid layer was present (Fig. 4). When
allowed to imbibe on solutions containing b-1,3-glucanase,
seeds began to lose OD after 5 d, and by 7 d of incubation,
only 20% of the seeds maintained semipermeability (Fig. 4).
A chloroform dip before b-1,3-glucanase treatment accel-
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Callose and Apoplastic Semipermeability
Figure 3. A, Water-absorption kinetics of muskmelon seeds. Decoated seeds were incubated on water-saturated blotters at 30°C with
or without chloroform treatment (3-min dip) and/or boiling. Dotted
lines for control and chloroform-treated seeds indicate germinated
seeds. F, Control seeds; f, chloroform-treated seeds; E, boiled
seeds; M, boiled and chloroform-treated seeds. The bars represent 6
SE (n 5 20). B, Percentages of germination (control [F] and
chloroform-treated [f]) and OD (boiled [E] and boiled and
chloroform-treated [M]) of decoated muskmelon seeds.
erated the loss of OD (Fig. 4), suggesting that the lipid layer
restricts access by the enzyme to the callose layer.
Endosperm envelopes from seeds that had been treated
with both chloroform and b-1,3-glucanase and had lost OD
did not stain with aniline blue or Sudan III and IV (Fig. 5A;
compare with Fig. 2D). The b-1,3-glucanase treatment digested the globular layer outside of the endosperm cells,
leaving only a thin, porous network in the surface view
(Fig. 5B). A fractured cross-section of the b-1,3-glucanasetreated endosperm envelope showed that the callose layer
had essentially disappeared (Fig. 5C, arrow), leaving only a
few isolated regions where the globular material could still
be found (Fig. 5C, CA). The effects of the enzyme treatment
on OD (Fig. 4) and on callose degradation (compare Fig. 1,
A, B, E, and G, with Fig. 5, A–C) leave little doubt that the
callose layer is responsible for the semipermeability of the
endosperm envelope.
DISCUSSION
Although it was hypothesized long ago that callose may
act as a “molecular filter” in plants by altering the gelfiltration properties of the cell wall (Heslop-Harrison,
1964), no direct in vivo evidence to support this hypothesis
87
has been reported. Extracted callose has a high waterholding capacity (Barskaya and Balina, 1971; Vithanage et
al., 1980), and low permeability to small molecules of
water-swollen callose (Eschrich and Eschrich, 1964, cited
by Stone and Clarke, 1992). Here we present evidence that
a thick deposit of callose covering the endosperm envelope
of muskmelon seeds serves as a semipermeable molecular
filter that readily allows movement of water but not of
solutes.
The thick outer wall of the muskmelon endosperm envelope can be stained with both aniline blue dye and
aniline blue fluorochrome (Fig. 1, A–C, F), which are specific for b-1,3-glucans, and is also virtually completely
digested by b-1,3-glucanase (Fig. 5, A–C). Thus, the thick
globular layer of the outer endosperm wall (Fig. 1, E and G)
is composed largely of callose, although the presence of
other components in addition to callose cannot be excluded. The b-1,3-glucanase treatment also causes loss of
OD (Fig. 4), providing direct evidence that the callose layer
is responsible for semipermeability. This conclusion is further supported by the simultaneous deposition of callose,
the development of semipermeability (Welbaum and Bradford, 1990), and the increase in the energy required to
penetrate the endosperm envelope (Oluoch, 1996) at
around 40 DAA during seed development.
Callose generally occurs in plant cells as a component of
specialized wall or wall-associated structures at certain
stages of growth. Callose has been detected in or on cell
walls of various tissues, including cell plates, cotton seed
fibers, pollen grain cell walls, innermost pollen tube walls,
endosperms, sieve plates, and abscission zones (Esau, 1948;
Currier, 1957; Heslop-Harrison, 1964; Scott et al., 1967;
Morrison and O’Brien, 1976; Waterkeyn, 1981; Stone and
Figure 4. Loss of OD after b-1,3-glucanase treatment. Decoated
muskmelon seeds were killed in boiling water for 3 min and incubated overnight on blotters saturated with distilled water to induce
OD. For chloroform treatments, boiled seeds with OD were dipped
into a chloroform solution for 3 min and rinsed with water. Seeds in
buffer (M) or in buffer after chloroform treatments (E) maintained OD
for at least 7 d. The presence of b-1,3-glucanase in the buffer (L)
initiated the loss of OD at 5 d of incubation or after only 1 d in seeds
pretreated in chloroform (Œ). The bars represent 6 SE from three
independent experiments of 20 seeds each.
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88
Yim and Bradford
Figure 5. A, Cross-section of chloroform- and b-1,3-glucanasetreated muskmelon seeds. Seeds with OD were treated with chloroform and incubated for 36 h on blotters saturated with solution
containing b-1,3-glucanase. Seeds that lost OD were hand sectioned
and stained with aniline blue and Sudan III and IV. The Sudanstaining lipid layer and the aniline-blue-staining callose layer were
removed (arrow; compare with Fig. 2D). B, Surface of chloroformand b-1,3-glucanase-treated muskmelon endosperm envelope
viewed by SEM (compare with Fig. 1G). C, Freeze-fractured crosssection of chloroform- and b-1,3-glucanase-treated muskmelon endosperm envelope viewed by SEM. The cross-section shows that the
globular layer is removed (arrow; compare with Fig. 1G), leaving
only a few remnants in some areas (CA). The bar in each panel
presents 5 mm.
Clarke, 1992). Callose deposition is also induced locally by
wounding, stress, and fungal or viral infection (Currier,
1957; Esau and Cronshaw, 1967; Coffey, 1976). Despite the
widespread occurrence of callose, its general function is not
well understood (for review, see Stone and Clarke, 1992). It
may serve as a matrix for deposition of other cell wall
materials, as in developing cell plates and sieve-plate
pores; as a cell wall-strengthening material, as in cotton
seed hairs (Maltby et al., 1979) and pollen; as a sealing or
plugging material at the plasma membrane of pit fields,
plasmodesmata, and sieve-plate pores (Eschrich, 1975); as a
mechanical obstruction to growth of fungal hyphae; or as a
special permeability barrier, as in pollen mother cell walls
and muskmelon endosperm envelopes (Heslop-Harrison,
1964; this report). In addition, the degree of polymerization, age, and thickness of the deposits may vary the physical properties of callose (Stone and Clarke, 1992). For
Plant Physiol. Vol. 118, 1998
example, callose deposits in clover seed coats apparently
prevent water absorption (Bhalla and Slattery, 1984), in
contrast to the high water permeability of the muskmelon
endosperm envelope (Fig. 3A).
Lipid-containing or suberized cell walls (stained with
Sudan dye) are frequently associated with semipermeable
regions (Fig. 1B; Harrington and Crocker, 1923; Johann,
1942; O’Brien and Carr, 1970; Cochran, 1983; Welbaum and
Bradford, 1990; Jacobsen et al., 1992; Welbaum et al., 1992;
Beresniewicz et al., 1995a). However, the lipid-containing
outer layer of the melon endosperm envelope slows water
uptake but is not responsible for semipermeability (Fig. 3).
Whether callose is also associated with other “suberized”
apoplastic membranes, such as the Casparian strip of the
root endodermis or the sheaths of sugarcane vascular bundles, is unknown. Cucumber, zucchini, watermelon, and
barley seeds, all of which exhibit semipermeability, also
have a thick aniline-blue-staining layer inside of the seed
coat (data not shown). However, some seeds have semipermeable layers that do not contain callose (Beresniewicz
et al., 1995a; K.-O. Yim and K.J. Bradford, unpublished
results for lettuce endosperm envelopes). Because embryos
often leak solutes upon initial imbibition (Simon and Mills,
1983), a semipermeable envelope would prevent loss of
solutes to the environment until the embryo is capable of
reabsorbing them before initiation of radicle growth. There
are many additional locations in the plant where the ability
to restrict solute movement in the apoplast while permitting water continuity and flow would be advantageous,
particularly in developing seeds (Bradford, 1994), in the
root (Steudle and Peterson, 1998), and in the vascular system (Canny, 1995).
The large amounts of callose present and its specific
deposition only on the outer side of the endosperm envelope (Fig. 1, B, C, and E) raise intriguing questions about
the mechanism of callose synthesis in these cells. Synthesis
of callose via a well-characterized plasma membranebound callose synthase is usually activated temporally by
specific signals such as wounding, infection, or other
stresses (for review, see Delmer and Amor, 1995). The
plasma membrane callose synthase is activated by micromolar concentrations of Ca21 and b-glucoside (Hayashi et
al., 1987), suggesting that transient increases in Ca21 concentration caused by cell perturbation, such as by microbe
infection or wounding, induce callose deposition outside of
the plasma membrane. Although less well characterized,
there is some evidence for a Golgi-vesicle-mediated
callose-synthesis system in developmentally regulated
callose-rich tissues such as pollen tube walls, pollen tube
plugs, and developing cell plates. In pollen tubes callose
synthase activity was associated with Golgi vesicles (Helsper et al., 1977) and showed little dependence on Ca21
(Schlüpmann et al., 1993; Li et al., 1997). Callose was detected by immunogold labeling in Golgi vesicles, which
were concentrated at the tips of germinating pollen tubes of
camellia (Hasegawa et al., 1996). However, Li et al. (1997)
proposed that inactive callose synthase is secreted in vesicles at the tips of tobacco pollen tubes and is activated
upon insertion into the plasma membrane. When callose
accumulation occurred at about 35 DAA in the muskmelon
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Callose and Apoplastic Semipermeability
endosperm envelope, discrete vesicles or globules were
stained with aniline blue (Fig. 2C), consistent with the
globular appearance of the callose layer (Fig. 1, E and G).
However, more detailed studies of the callose synthesis
and deposition process in muskmelon endosperm envelopes are required to determine the mechanism by which
the callose layer is formed.
In conclusion, we have demonstrated that the semipermeability of the muskmelon endosperm envelope is caused
by a callose-containing layer deposited outside of the outer
walls of the endosperm cells. A chloroform-soluble waxy
layer outside of the callosic layer delays water uptake but
is not required for semipermeability. These results directly
confirm, for the first time to our knowledge, the longstanding hypothesis that callose may act as a molecular
sieve. They also show that the presence of Sudan-stainable
material does not in itself imply impermeability to water.
The existence of semipermeable apoplasts resulting from the
deposition of callose (or other materials) in the cell wall
provides options for controlling water and solute transfer in
many parts of the plant and at critical stages of development.
ACKNOWLEDGMENTS
The authors express their appreciation to Dr. Deborah Delmer
(University of California, Davis) for critical reading and constructive comments on the manuscript and for technical assistance with
the fluorescence microscopy. The assistance of Dr. Sunitha Gurusinghe with the flow cytometry of endosperm nuclei is gratefully
acknowledged.
Received February 17, 1998; accepted June 14, 1998
Copyright Clearance Center: 0032–0889/98/118/0083/08.
LITERATURE CITED
Barskaya EI, Balina NV (1971) The role of callose in plant anthers.
Fiziol Rast 18: 716–721
Beresniewicz MM, Taylor AG, Goffinet MC, Koeller WD (1995a)
Chemical nature of a semipermeable layer in seed coats of leek,
onion (Liliaceae), tomato and pepper (Solanaceae). Seed Sci
Technol 23: 135–145
Beresniewicz MM, Taylor AG, Goffinet MC, Terhune BT (1995b)
Characterization and location of a semipermeable layer in seed
coats of leek and onion (Liliaceae), tomato and pepper (Solanaceae). Seed Sci Technol 23: 123–134
Bhalla PL, Slattery HD (1984) Callose deposits make clover seeds
impermeable to water. Ann Bot 53: 125–128
Bradford KJ (1994) Water stress and the water relations of seed
development: a critical review. Crop Sci 34: 1–11
Brown R (1907) On the existence of a semi-permeable membrane
enclosing the seeds of some of the Gramineae. Ann Bot 21: 79–87
Canny MJ (1995) Apoplastic water and solute movement: new
rules for an old space. Annu Rev Plant Physiol Plant Mol Biol 46:
215–236
Cochran MP (1983) Morphology of the crease region in relation to
assimilate uptake and water loss during caryopsis development
in barley and wheat. Aust J Plant Physiol 10: 473–491
Coffey MD (1976) Flax rust resistance involving the K gene: an
ultrastructural survey. Can J Bot 54: 1443–1457
Currier HB (1957) Callose substance in plant cells. Am J Bot 44:
478–488
Delmer DP, Amor Y (1995) Cellulose biosynthesis. Plant Cell 7:
987–1000
89
Esau K (1948) Phloem structure in the grapevine, and its seasonal
changes. Hilgardia 18: 217–296
Esau K, Cronshaw J (1967) Relation of tobacco mosaic virus to the
host cells. J Cell Biol 33: 665–678
Eschrich W (1975) Sealing systems in phloem. In MH Zimmermann, JA Milburn, eds, Encyclopedia of Plant Physiology, Vol 1:
Transport in Plants. I. Phloem Transport. Springer-Verlag, Berlin, pp 39–56
Eschrich W, Eschrich B (1964) Das verhalten isolierter callose
gegenüber wässrigen lösungen. Ber Dtsch Bot Ges 77: 329–331
Gola G (1905) Richerche sui rapporti tra i tegumenti seminali e le
soluzioni saline. Annali di Botanica 3: 59–100
Harrington GT, Crocker W (1923) Structure, physical characteristics, and composition of the pericarp and integument of
Johnson grass seed in relation to its physiology. J Agric Res 23:
193–222
Hasegawa Y, Nakamura S, Nakamura N (1996) Immunocytochemical localization of callose in the germinated pollen of
Camellia japonica. Protoplasma 194: 133–139
Hayashi T, Read SM, Bussell J, Thelen M, Lin FC, Brown RM,
Delmer DP (1987) UDP-glucose:(1,3)-b-glucan synthases from
mung bean and cotton. Differential effects of Ca21 and Mg21 on
enzyme properties and on macromolecular structure of the glucan product. Plant Physiol 83: 1054–1062
Helsper JPFG, Veerkamp JH, Sassen MMA (1977) b-Glucan synthetase activity in Golgi vesicles of Petunia hybrida. Planta 133:
303–308
Heslop-Harrison J (1964) Cell walls, cell membranes and protoplasmic connections during meiosis and pollen development. In
HF Linskens, ed, Pollen Physiology and Fertilization. NorthHolland, Amsterdam, pp 29–47
Hill HJ, Taylor AG (1989) Relationship between viability, endosperm integrity and imbibed lettuce seed density and leakage.
HortScience 24: 814–816
Jacobsen JS, Fisher DG, Maretzki A, Moore PH (1992) Anatomy
of sugarcane stem in relation to phloem unloading and sucrose
storage. Bot Acta 18: 959–969
Jensen WA (1962) Botanical Histochemistry: Principles and Practice. WH Freeman, San Francisco, CA, pp 35–99
Johann H (1942) Origin of the suberized semipermeable membrane in the caryopsis of maize. J Agric Res 64: 275–281
Kotowski F (1927) Semipermeability of seed coverings and stimulation of seeds. Plant Physiol 2: 177–186
Li H, Bacic A, Read SM (1997) Activation of pollen tube callose
synthase by detergents. Evidence for different mechanisms of
action. Plant Physiol 114: 1255–1265
Maltby D, Carpita NC, Montezinos D, Kulow C, Delmer DP
(1979) b-1,3-Glucan in developing cotton fibers. Structure, localization and relationship of synthesis to that of secondary wall
cellulose. Plant Physiol 63: 1158–1164
Morrison IN, O’Brien TP (1976) Cytokinesis in the developing
wheat grain: division with and without a phragmoplast. Planta
130: 57–67
O’Brien TP, Carr DJ (1970) A suberized layer in the cell walls of
the bundle sheath of grasses. Aust J Biol Sci 23: 275–287
Oluoch MO (1996) Effects of priming and stage of development on
vigor and longevity of muskmelon (Cucumis melo L.) seeds. PhD
dissertation. Virginia Polytechnic Institute and State University,
Blacksburg, Virginia
Oparka KJ, Gates P (1981) Transport of assimilates in the developing caryopsis of rice (Oryza sativa L.): the pathways of water
and assimilated carbon. Planta 152: 388–396
Peoples MB, Pate JS, Atkins CA, Murray DR (1985) Economy of
water, carbon and nitrogen in the developing cowpea fruit. Plant
Physiol 77: 142–147
Schlüpmann H, Bacic A, Read SM (1993) A novel callose synthase
from pollen tubes of Nicotiana. Planta 191: 470–481
Scott PC, Miller LW, Webster BD, Leopold AC (1967) Structural
changes during bean leaf abscission. Am J Bot 54: 730–734
Simon EW, Mills LK (1983) Imbibition, leakage and membranes.
In C Nozzolillo, PJ Lea, FA Loewus, eds, Mobilization of Reserves in Germination. Plenum Press, New York, pp 9–27
Downloaded from on June 16, 2017 - Published by www.plantphysiol.org
Copyright © 1998 American Society of Plant Biologists. All rights reserved.
90
Yim and Bradford
Singh B (1953) Studies on the structure and development of seeds
of Cucurbitaceae. Phytomorphology 3: 224–239
Steudle E, Peterson CA (1998) How does water get through roots?
J Exp Bot 49: 775–788
Stone BA, Clarke AE (1992) Chemistry and Biology of (133)-bGlucans. La Trobe University Press, Victoria, Australia, pp
355–429
Stone BA, Evans NA, Bonig I, Clarke AE (1984) The application of
Sirofluor, a chemically defined fluorochrome from aniline blue
for the histochemical detection of callose. Protoplasma 122:
191–195
Taylor AG, Beresniewicz MM, Goffinet MC (1997) Semipermeable layer in seeds. In EM Black, AJ Murdoch, TD Hong, eds,
Basic and Applied Aspects of Seed Biology. Kluwer Academic
Publishers, Dordrecht, The Netherlands, pp 429–436
Thornton ML (1968) Seed dormancy in watermelon Citrullus vulgaris Shrad. Proceedings of the Association of Official Seed
Analysts 58: 80–84
Vithanage HIMV, Gleeson PA, Clarke AE (1980) The nature of
callose produced during self-pollination in Secale cereale. Planta
148: 498–509
Plant Physiol. Vol. 118, 1998
Waterkeyn L (1981) Cytochemical localization and function of the
3-linked glucan callose in the developing cotton fibre cell wall.
Protoplasma 106: 49–67
Welbaum GE, Bradford KJ (1988) Water relations of seed development and germination in muskmelon (Cucumis melo L.) I.
Water relations of seed and fruit development. Plant Physiol 86:
406–411
Welbaum GE, Bradford KJ (1990) Water relations of seed development and germination in muskmelon (Cucumis melo L.) IV.
Characteristics of the perisperm during seed development. Plant
Physiol 92: 1038–1045
Welbaum GE, Meinzer FC (1990) Compartmentation of solutes
and water in developing sugarcane stalk tissue. Plant Physiol 93:
1147–1153
Welbaum GE, Meinzer FC, Grayson RL, Thornham KD (1992)
Evidence for and consequences of a barrier to solute diffusion
between the apoplast and vascular bundles in sugarcane stalk
tissue. Aust J Plant Physiol 19: 611–623
Wolswinkel P (1992) Transport of nutrients into developing seeds:
a review of physiological mechanisms. Seed Sci Res 2: 59–73
Downloaded from on June 16, 2017 - Published by www.plantphysiol.org
Copyright © 1998 American Society of Plant Biologists. All rights reserved.