Annals of Botany 77 : 105–122, 1996
Structure and Macromolecular Composition of the Seed Coat of the Musaceae
P E T E R G R A V EN*†, C H R I S G. D E K O S T E R†, J A A P J. B O O N† and F E R R Y B O U M AN*
* Hugo de Vries-Laboratory, UniŠersity of Amsterdam, Kruislaan 318, 1098 SM Amsterdam and † FOM Institute
of Atomic and Molecular Physics, Unit for Mass Spectrometry of Macromolecular Systems, Kruislaan 407,
1098 SJ Amsterdam, The Netherlands
Received : 20 June 1995
Accepted : 30 August 1995
Mature seed coats of representatives of all three genera of Musaceae were analysed for macromolecular composition
with various mass spectrometric techniques and compared with scanning electron microscopy and light microscopy
in combination with histochemical techniques. Mass spectrometric techniques are more sensitive and more specific
in identifying macromolecular compounds than histochemical methods. The macromolecular ‘ fingerprint ’ of the seed
coats of Musaceae showed unique components of aromatic phenols. The seed coat structure of all three genera is
homogeneous within the Musaceae. It is characteristic at the family level and most complex within the Zingiberales.
Very remarkable are the separation of the outer cell walls from the exotestal layer, exposing a secondary surface with
silica crystals, and the relatively thick mesotesta which protects the seed, e.g. against the biting forces and passage
through the digestive tracts of dispersing agents. Germination takes place with an operculum and is facilitated by a
predetermined rupture layer in the micropylar collar. The musaceaous seed presents a good example of the solution
of conflicting demands of protection and germination.
# 1996 Annals of Botany Company
Key words : Musaceae, Musa, Ensete, Musella, seed coat, pyrolysis (gas chromatography) mass spectrometry,
histochemistry, anatomy, macromolecules, silica, lignin, cellulose, vegetable polyphenols, operculum, germination.
INTRODUCTION
Hardly anything is known about the chemistry of seed
coats. The biochemical and histochemical literature on
seeds is almost fully focused on the composition of storage
material in embryo and endosperm. The functional interpretation of the seed coat is an almost unexplored field of
research, because of the lack of reliable data on the presence
and distribution of macromolecular components. Currently
it is possible to describe the anatomy in more detail with
microscopical and histochemical techniques and to study
the macromolecular composition of the testal layers of the
mature seed with mass spectrometrical techniques.
In source pyrolysis mass spectrometry (PyMS) is a relative
simple method to obtain information from plant tissue at
the molecular level (Boon, 1989). The PyMS analysis
provides a chemical ‘ fingerprint ’ of the seed coat. Chemical
identification of individual pyrolysis products is performed
by using pyrolysis gas chromatography (PyGC) combined
with mass spectrometry (PyGCMS).
The order Zingiberales is a well-defined and coherent
taxon. Within this order there are questions about the
relation of seed coat structure and chemistry to germination.
The Musaceae represents the most basal lineage in the order
(Manchester and Kress, 1993). This basal position of the
Musaceae, its special type of germination and its welldifferentiated seed coat forms an interesting subject to study
the relation of structure, macromolecular composition and
functions of the testal layers, and moreover to produce a
new data set relevant for macrosystematic research.
In most recent classifications the family of the Musaceae,
0305-7364}96}020105­18 $12.00}0
to which the bananas belong, there are three genera : Musa
L., with about 35 species, Ensete Bruce., with three species
and a more obscure genus Musella (Fr.) C. Y. Wu with one
species. The acceptance of the Musella as a distinct genus is
only provisional (Manchester and Kress, 1993). A number
of genera formerly mostly included in the Musaceae are
nowadays placed in the separated families Strelitziaceae,
Heliconiaceae and Lowiaceae (Nakai, 1941 ; Tomlinson,
1962 ; Cronquist, 1981 ; Dahlgren and Rasmussen, 1983).
Each taxon possesses a comparable range of anatomical
variation which overlaps to a minimum with other groups
within the Zingiberales (Tomlinson, 1962). The family is
tropical-subtropical Asian (Musa, Musella) and African
(Ensete), but Musa-cultivars are nowadays cultivated
troughout the tropical regions of the world.
Most edible cultivated clones are seedless whereas other
plants, wild or cultivated, produce one to many seeds.
Breeding programs are hindered by the low seed set, as well
as by slow and non-uniform germination (Shepherd, 1954 ;
Stotzky, Cox and Goose, 1962). The ovary of Musa is three
locular, contains numerous ovules, and develops into a
longish berry. The fruit of Musa contains a higher number
of seeds than that of Ensete. The seeds rarely attain 1 cm in
diameter, whereas the seeds of Ensete are mostly larger, up
to 1±8 cm, although there may be some overlap in size. The
shape of the seeds varies greatly, is often irregularly and
sharply angular, either rarely globose or smooth. The
variability in shape of the pale or dark brown to black seeds
is due to compression between neighbouring seeds
(Simmonds, 1960). The embryological literature has been
summarized by Davis (1966), that on seed anatomy by
# 1996 Annals of Botany Company
106
GraŠen et al.—Composition of Musaceaous Seed Coats
Netolitzky (1926) and Takhtajan (1985). In earlier studies
McGahan (1961 a) described the anatomy of the seed of
Musa balbisiana and Bouharmont (1963) the development
of the ovule to mature seed in Musa acuminata.
The ovule is anatropous, bitegmic, and crassinucellar,
with a multi-layered outer integument and a thin inner
integument. The micropylar part of the seed coat develops
into an operculum or seed lid. This operculum is surrounded
by a characteristic micropylar collar. During germination
the seed lid is displaced by the elongating radical-hypocotyl
axis (McGahan, 1961 b). The mature seed contains mainly
endosperm, which is copious, starchy and mealy, with large,
eccentric, compound starch grains, and some perisperm
(Cronquist, 1981). The endosperm is initially nuclear,
becoming cellular later, and extends during its development
at the expense of the perisperm. The embryo remains
relatively small, is situated under the operculum and is
partly enclosed by the micropylar collar. In the ovular stage
a trichomatous aril develops out of the funicle surrounding
the seed coat (Netolitzky, 1926 ; Friedrich and Strauch,
1975). According to Friedrich and Strauch (1975) the aril
becomes reduced and only some small vestiges can be found
near the funicle on the mature seed.
Seeds of Ensete can be distinguished from those of Musa
by a rimmed hilar depression that is nearly as wide as the
seed itself. Seeds of Musella resemble those of Musa, being
distinguished from Ensete by the lack of a wide hilar
depression and rim (Manchester and Kress, 1993). Most
species of Musa have a rough verrucate seed coat, whereas
the seed coat of Musella and Ensete is smooth.
In this study data from scanning electron microscopy
(SEM), light microscopy (LM), histochemistry and mass
spectrometry are compared, to acquire insight into the
functions and chemical composition of specialized testal
layers.
MATERIALS AND METHODS
Plant material
Mature seeds of Musa acuminata Colla, Musa balbisiana
Colla, Musa mannii H. Wendland, Musa paradisiaca L.,
Musa rosacea Jacquin, Musa textilis Ne! e, Musa Šelutina H.
Wendland & Drude, Musa sp., Ensete Šentricosum
(Welwitsch) Cheesman, Ensete glaucum (Roxb.) Cheesman,
and Musella lasiocarpa (Fr.) C. Y. Wu used in this study
were collected via botanical gardens all over the world. The
seeds were stored dry at room temperature in closed plastic
containers.
Germination
Seeds of Musa Šelutina and Musa rosacea were soaked in
water for 24 h, and sown in petri dishes on wet filter paper
and in soil under greenhouse conditions.
with a hexane}dichloromethane mixture (1 : 1 v}v), after 3 h
it was centrifuged, the pellet was suspended in deionized
water, dried and extracted with an ethanol}acetone mixture
(1 : 1 v}v) for another 3 h, and finally washed several times
in deionized water. The homogenized and extracted seed
coat material was suspended in a few drops of water.
Transmethylation
For methylation of phenolic hydroxyl and carboxyl
groups, a small droplet of 2±5 % (w}v) tetramethylammonium (TMAH) was added to the wet sample on the
wire of the direct insertion probe prior to PyMS and
PyGCMS experiments.
Silylation
An off-line pyrolysate was prepared by placing a few
drops of an aqueous suspension of extracted seed coat on a
wire with a Curie point of 510 °C. The wire was dried under
rotation in vacuum. After drying the wire was placed in a
glass liner and the glass liner was flushed for a minute with
argon, directly followed by pyrolysis at atmospheric
pressure. The procedure was repeated several times with a
new wire in the same glass liner and the final condensate
that formed on the inner wall of the glass liner was collected.
Silylation of the off-line pyrolysate was performed by
adding 70 µl of BSTFA}TMSCL [N.O.bis(TMS)
trifluoroamide}trimethylchlorosilane] in 235 µl pyridine to
the sample in the dark. After 24 h the pyridine was
evaporated by flushing with nitrogen and the reaction
product was resuspended in dichloromethane and analysed
with GCMS.
Acid transesterification
For hydrolysis a few milligrams of the extracted seed coat
were placed in a tube with a screw top and 12  sulphuric
acid (approx. 2 ml) was added. The tube was flushed with
nitrogen before sealing and the sample was incubated at
room temperature for 3 h. The acid was diluted to 1  and
the sample was further hydrolysed at 100 °C overnight. The
sample was then centrifuged and the residue was hydrolysed
again with 0±25  sulphuric acid. The tube was flushed with
nitrogen and incubated at 100 °C overnight. The solid
residue was separated from the liquid hydrolysate by
centrifugation and washed several times with deionized
water and dried overnight over phosphorus pentoxide in a
vacuum dessicator at 50 °C (Pastorava, Oudemans and
Boon, 1993).
Ethyl esters were prepared by using approx. 1 mg of the
extracted sample in a plastic tube with a screw top with a
20 µl H SO }ethanol solution (1±25 µl H SO in 1 ml
# %
# %
ethanol). The tube was sealed with a Teflon tape and
incubated at 100 °C for 4 h.
Sample preparation for mass spectrometry
Samples of seed coats of Musaceae were collected and
homogenized in a few drops of water in a small glass mortar
with a glass pestle. The homogenized seed coat was extracted
Direct temperature resolŠed or pyrolysis mass spectrometry
A small drop of a suspension of the unextracted or
homogenized and extracted seed coat was placed on a
GraŠen et al.—Composition of Musaceaous Seed Coats
107
F. 1. A, Mature seed of Musa balbisiana (¬10). B, Musa sp. showing, partly removed, the thin brittle remains of the aril (a), the outer epidermis
(oe) of the outer integument and crystalline silica bodies (si). Irregular radial divisions arise at the outer layer of the mesotesta (arrowhead), which
gives the seed a rough appearance in some species (¬90). C–E, Musa balbisiana ; The mesotesta (me) has developed into 20–25 layers of sclerotic
cells. C, The cells of the four to five uppermost layers are more thickened and less elongated (¬140). D, Backscatter image showing the silica
bodies as a thin layer on top of the mesotesta (¬135). E, Sclerotic cells with pits at the lower part of the mesotesta (¬1010). F, The inner epidermis
of the testa develops into an endotesta (en) consisting of U-shaped thickened cells. The tegmen (te) consists of two layers of tangentially elongated
cells. A thick cuticle bounds the inner surface of the tegmen and has a tegmic and nucellar origin (Musa balbisiana) (¬610). G, In the mature
seeds the perisperm (pe) is completely compressed and only the cell walls remain (Musa mannii) (¬1380). H, The tegmic cells are filled with a
spongy like structure (Musa balbisiana) (¬2690).
108
GraŠen et al.—Composition of Musaceaous Seed Coats
F. 2. For legend see facing page.
GraŠen et al.—Composition of Musaceaous Seed Coats
Pt}Rh (9 : 1) wire of the insertion probe for in-source
pyrolysis mass spectrometry (PyMS) on a JEOL DX-303
double focusing mass spectrometer. The wire was inserted
into the evacuated ion source, after drying the sample. The
ion source temperature was 180 °C. The wire was resistively
heated at 16 °C s−" to a final temperature of 800 °C.
Compounds, formed upon desorptionor pyrolysis, were
ionised by 16 eV electron impact (EI) or ammonia chemicalionization (CI) PyMS. The scan cycle time was 1 s and the
selected massrange was m}z 20–1000.
Curie-point pyrolysis gas chromatography mass
spectrometry
Curie-point pyrolysis gas chromatography mass spectrometry under EI conditions was conducted on a Finnigan
INCOS 50 quadrupole mass spectrometer connected to a
HP 5890 series II gas chromatograph. About 5 µg of a
suspension of a sample was applied to a ferromagnetic wire
with a Curie-point temperature of 610 °C. The pyrolysis
products were flushed towards a 25 m CPSILS-5 CB fused
silica capillary column (i.d. 0±32 mm ; film thickness 1±2 µm)
using He as carrier gas (flow 30 cm s−"). The GC oven was
kept at 30 °C during pyrolysis and was subsequently
programmed to 320 °C at a rate of 6 °C min−". The
compounds were ionised at 70 eV electron impact energy.
The scan cycle time was 0±5 s with a selected mass range of
m}z 20–500.
On column injection gas chromatography mass
spectrometry
On column injection gas chromatography mass spectrometry (GCMS) was performed using a HRGC MEGA 2
series FISONS instruments gas chromatograph which was
connected to a Jeol DX-303 double focusing mass spectrometer. The column used was a 25 m SIMDIST (i.d.
0±32 mm, film thickness 0±15 µm). Helium was used as
carrier gas (flow 30 cm s−"). The GC oven was kept at 30 °C
and subsequently heated to 375 °C at a rate of 6 °C min−".
Compounds were ionized at 70 eV electron impact energy,
at a source temperature of 180 °C. The scan cycle time was
0±8 s over a mass range of m}z 20–500.
Scanning electron microscopy
Seeds were sectioned transversely and fixed in FAA
(formalin, ethanol, acetic acid), dehydrated in acetone,
incubated in tetramethylsilane and air dried (Dey et al.,
1989). The seeds were coated with a thin layer of
109
gold}palladium and examined with an ISI-DS130 Dual
Stage electron microscope (9 kV) fitted with an ISIRobinson Detector RBSE 130R (19 kV) for backscatter
imaging.
Mature seeds of Musa balbisiana, Musa paradisiaca,
Musa textilis and Musa Šelutina were examined for the
presence and localization of silica in the seed coat. The seeds
were sectioned transversely, fixed on aluminium stubs
cemented with carbon cement and air dried under low
relative humidity at room temperature. The specimens were
carbon coated and examined in a Cambridge Stereoscan 150
(at 20 kV) fitted with a backscatter detector (KE Developments, Cambridge, UK) and a LINK energy-dispersive Xray analyser [Department of Molecular Cell Biology
(EMSA) of the University of Utrecht, The Netherlands].
The presence of silica was detected with the help of the
diagrams of the X-ray spectra. The specific peak (keV, 1±74)
for silica in the spectrum was isolated and the spectrum
pulses were simultaneous with the scanning of the sample
carried back so that an element distribution image of silica
could be created (X-ray mapping).
Light microscopy and histochemistry
Hand-cut sections were stained for vegetable polyphenols
(‘ tannins ’) with ferric chloride, vanillin}HCl and the nitroso
reaction, for lipids with Magdala red, Nile blue sulphate
method, for cuticle (neutral fats and fatty acids) with Sudan
IV, for ‘ cutin ’ with azure B method and auramine O
method, for suberin with phosphate buffer method and
cyanin method, for lignin with sodium hypochlorite}HCl,
phloroglucinol}HCl and lignin pink, for callose with soda
method, Corallin and aniline blue , for pectin with ruthenium
red, and for cellulose with chlor-zinc-iodide (Gahan, 1984 ;
Krisnamurthy, 1988). Sections for staining with ruthenium
red and for the cyanin}potassium method were also
pretreated with Eau de Javelle (potassium hypochlorite).
Samples for semi-thin sectioning were fixed in FAA or
Craf III, or were softened in 10 % ammonia (1 h at 50 °C)
or a mix of glycerol and Aerosol OT in water (Schmid and
Turner, 1977 ; Setoguchi, Tobe and Ohba, 1992), dehydrated
in a n-butyl alcohol series and embedded in glycerol
methylacrylate polymer (Feder and O’Brien, 1968).
RESULTS
Mature seed coat anatomy and histochemistry
In mature seed coats the cells are difficult to interpret with
light microscopy due to the extreme hardness of the tissue.
This has hampered the observations on the mature seed
F. 2. A, Each exotestal cell contains a multi-angular crystalline silica body (Musa sp.) (¬500). B–C, The exotesta easily releases and while
separating, the tips of the silica bodies break off and stick to the membrane (Musa sp.) (B, ¬220 ; C, ¬665). D, The silica bodies are situated
against the inner periclinal wall of the epidermal cells (Musa sp.) (¬270). E, The tip of the silica bodies breaks off after the release of the outer
periclinal wall and the radial walls (Musa sp.) (¬1010). F, Backscatter image of the silica bodies of Musa balbisiana (¬605). G, Backscatter image
of the seed coat of Musa balbisiana with partly released epidermis and remains of the aril (¬75). H–I, Backscatter image and bright dot mapping
on silica in the seed coat of Musa paradisiaca (H, I ¬165). J, Backscatter image of released epidermal wall of Musa balbisiana and silica bodies
which are situated on the inner periclinal wall (¬300). K–L, Backscatter and bright dot mapping view on the inner side of the separated epidermal
wall, showing the broken tips of the silica bodies and the silica encrusted walls and membrane of the exotesta of Musa balbisiana (K, L ¬630).
110
GraŠen et al.—Composition of Musaceaous Seed Coats
hi
mi
op
se
ab
mc
em
ii
oi
end
nu
pt
cd
F. 3. Schematic drawing of a longitudinal section of the Musa seed coat (¬30). ab, abscission layer ; cd, chalazal disk ; em, embryo ; end,
endosperm ; hi, hilum ; ii, inner integument ; mc, micropylar collar ; mi, micropyle ; nu, nucellar tissue or perisperm ; oi, outer integument ; op,
operculum ; pt, parenchymatic tissue between chalaza and nucellus ; se, silicified exotesta.
coats of Musaceae. Softening of the seeds facilitates
sectioning of the material, but it releases some of the
components in the seed coat (e.g. silica). Most stains used
were negative or aspecific.
The seed coat anatomy of all three genera in Musaceae
was practically homogeneous therefore the structure of the
Musa sp. only is described here.
During the development of the seed the aril jellifies and
dries out. In the mature seed (Fig. 1 A) the remains of the
aril adhere closely to the seed coat (Fig. 1 B). The aril
surrounds the ripe seed partly as a thin brittle layer and, if
not released, swells up during germination.
The cells of the outer epidermis of the outer integument
are radially stretched and vary in length (65–190 µm). The
epidermis is covered with a thin cuticle which stains
positively with sudan IV, azure B and auramine O. This
indicates ‘ cutin ’ and fatty acids in the cuticle. Each
epidermal cell contains a multi angular crystalline silica
body, which is situated against the inner periclinal wall (Figs
1 D, 2 A, G–J). The walls of the epidermal cells remain
relatively thin and are encrusted with a net-like silica
structure. The outer periclinal and radial walls easily
separate from the seed coat (Fig. 2 G, J). During the release,
the tips of the silica bodies may break off and stick to the
membrane (Fig. 2 B–C, J–L). The main part of the silica
body remains attached to the seed coat after release of the
epidermal wall layer (Fig. 2 A, D–F).
The middle layers of the outer integument have developed
into 20–25 layers (180–350 µm) of sclerotic cells (Fig.
1 C–E). These vary greatly in shape (length up to 300 µm ;
width about 10 µm), size and orientation. The secondary
cell wall contains phenolic compounds in the S2 layer,
which gave a very weak response with the nitroso reaction
and the vanillin}HCl test. These compounds increase in
relative amount from the inner to the outer side of the
mesotesta. The phenolic compounds are more concentrated
in the four to five uppermost cell layers of which the cell
walls are more thickened and the cells are less elongated.
The histochemical tests on lignin were mostly negative, only
in very thin sections a light pink colouration could be seen
in the S1 layer of the secondary wall, using lignin pink.
In some species locally irregular periclinal divisions lead
to protrusions on the outer layer of the mesotesta, which are
the cause of a rough surface on the seed of most Musa
species (Fig. 1 A–B). The innermost layer of the outer
integument develops into an endotesta consisting of Ushaped thickened cells (Fig. 1 F). Testa and tegmen are not
separated by a distinct cuticular layer. The tegmen consists
of two layers of longitudinally elongated cells (Fig. 1 F–G).
The inner wall of the outer layer and the inner and outer
F. 4. A, An abscission layer (ab) has developed through the outer integument and delimitates the operculum (op) (Musa balbisiana) (¬40) B,
The tegmen and nucellus (nu) rupture just beneath the abscission layer (Musa sp.) (¬100). C–D, The operculum consists of three distinct layers.
The raphal zone is followed by a parenchymatic tannin zone and a sclerenchymatic zone (Musa sp.) (C ¬60, D ¬30). E, The silicified vascular
bundle ends in the chalazal disk (cd) consisting of thin walled cells (Musa paradisiaca) (¬55). F, The powdery endosperm (end) forms a thick
layer along the inner side of the seed (Musa balbisiana) (¬705).
GraŠen et al.—Composition of Musaceaous Seed Coats
F. 4. For legend see facing page.
111
112
GraŠen et al.—Composition of Musaceaous Seed Coats
T     1. List of a number of compound classes and the
pyrolysis mass spectrometric and PyGCMS techniques by
which they can detected
PyMS
EI
Polysaccharide
Protein*
Lignin
Vegetable
polyphenol
(tannin)
Cutin
Phenolic acids
Lipids
(x)
x
x
x
(x)
x
PyGCMS
NH TMAH
$
CI
EI
x
x
(x)
x
(x)
EI
TMAH
EI
GCMS
EI
(x)
x
x
x
(x)
x
(x)
(x)
x
x
(x)
x†
x
* Poor response.
† After acid transesterification.
walls of the innermost layer are thickened. The tegmic cells
are filled with a spongy-like structure of unknown composition (Fig. 1 H). A thick cuticle, which strongly stains
with Sudan IV and Nile blue, is grafted to the inner surface
of the tegmen. This cuticle partly originates from the inner
layer of the inner integument and partly has a nucellar
origin (Fig. 1 F–G).
In the mature seed the perisperm is completely compressed
and only the cell walls remain (Fig. 1 G). The powdery
endosperm (Fig. 4 F) of the mature seed forms a thick layer
along the inner side of the seed and leaves a cavity in the
middle of the seed (Fig. 3).
At the micropylar side a typical micropylar collar is
formed by a local ingrowth of the outer integument into the
inner integument and the nucellus (Figs 3 and 4 A). An
abscission layer continuous with the micropylar collar has
developed through the outer integument and envelops the
operculum (Fig. 4 A). The operculum (width 1–1±5 mm) is
mainly formed by the exostomal part at the outer integument
and hilar}raphal tissue (Fig. 4 D). The operculum consists
of three distinct zones (Fig. 4 B–D). The raphal top zone is
followed by a parenchymatic ‘ tannin ’ zone and a sclerenchymatic zone. The hilum and micropyle are situated at the
top of the operculum (Fig. 3). The silicified vascular bundle
is widely branched through the top zone of the operculum,
running through the raphal mesotesta and ending in the
chalazal disk (Figs 3 and 4 E). During germination the
operculum is easily pushed out by the embryonic root, due
to the abscission layer. Removing the operculum does not
result in faster or higher percentage of germination. The
F. 5. Pyrolysis (EI) mass spectrum of the non-extracted seed coat of Musa Šelutina showing hydroxymethoxybenzyl cation (m}z 137) as major
fragment ion. D, guaiacyl lignin ; E, syringyl lignin ; _, polysaccharides ; ­, lipids ; q : Musaceae characteristics ; f, ferulic acid ; pc, p-coumaric
acid ; t, vegetable polyphenol markers.
GraŠen et al.—Composition of Musaceaous Seed Coats
113
×3
576
100
602
550
618
704
620
572
632
Relative abundance
691
592
732
648 662 676
0
500
550
600
650
700
750
950
1000
×3
100
882
857
830
798
911
0
750
800
850
900
m/z
F. 6. Lipid fraction of pyrolysis (EI) mass spectrum of the total seed coat of Musa Šelutina. ­, di- and triglycerides ; ', waxes.
tegmen and nucellus rupture also just beneath the abscission
layer in the outer integument (Fig. 4 B–C). The epidermal
cells at the nucellar apex are radially elongated and fill the
bottom of the micropylar channel (Fig. 3).
The inner and outer integument are discontinuous at the
chalazal side. At this place a central disk has developed
consisting of thin-walled cells (Fig. 4 E). In the developing
seed these cells are filled with small reddish brown vacuoles
which fuse later on to a single vacuole. According to White
(1928), McGahan (1961 a) and Grootjen (1981, unpubl. res.)
those cells are filled with a gelatinous substance. The
contents become brittle in the mature seed. On top of the
114
GraŠen et al.—Composition of Musaceaous Seed Coats
TIC
180
G
210
S
576
Dg
676
Wx
10
20
30
40
Scans
50
60
70
F. 7. Total ion current and mass chromatograms of the total seed
coat of Musa Šelutina. The ions of the mass chromatogram comprise
markers of several compound classes as guaiacyl lignin (G), syringyl
lignin (S), diglycerides (Dg) and waxes (Wx).
chalazal disk, between the insertion of the inner integument,
a parenchymatic tissue is situated.
Macromolecules
No single technique exists to identify all macromolecular
components at once. The evidence for the presence of the
macromolecular components has to come from a combination of separation and ionisation methods. Table 1 lists
a number of component classes which can be recognized
with direct mass spectrometric and PyGCMS techniques.
By addition of TMAH as a reagent it is possible to directly
transesterify ester bound components which facilitates their
detection by mass spectrometry.
Figure 5 shows the overall spectrum of the non-extracted
seed coat of Musa. This spectrum contains markers of a
volatile fraction and a non-extractable part of the polymer
system. All the three genera of the Musaceae gave the same
characteristic chemical ‘ fingerprint ’ of their seed coat. A
lipid fraction of fatty acids (m}z 256, C : ; 262, C : [M") #
"' !
H O] ; 264, C : [M-H O] ; 280, C : ; 282, C : ; 284, C : ),
#
") #
") "
") !
#
") "
and as shown in Fig. 6, diglyceridic fragments of triglycerides
(e.g. m}z 550, C , [M-H O] ; 576, C , [M-H O] ; 602,
#
"' ")
#
"' "'
C , [M-H O]), triglycerides (e.g. m}z 830, C , , [M#
"' "' ")
") ")
H O] ; 857, C , , [M-H O] ; 882, C , , [M-H O]) and
#
") ") ")
#
#
"' ") ")
wax esters (m}z 592­n.28), are thermally desorbed in an
early stage of the temperature ramp (Fig. 7), whereas the
biopolymeric materials are pyrolysed at higher
temperatures. The presence of C and C hydroxy fatty
"'
")
acids is confirmed with Py-TMAH-EI-GCMS and PyTMAH-(ammonia)CI-GMS (not shown), indicating the
presence of cutin (Boon, unpubl. res.).
Evidence for lignin in the mass spectrum of the nonextracted seed coat sample of Musaceae can be seen as the
monomeric pyrolysis products at m}z 124, 138, 150, 152,
164, 166, 178, 180 and pyrolysis products of dimers at m}z
272, 328, 340, 358 of guiacyl lignin. A low abundance of
syringyl lignin, indicated by m}z 154, 167, 168, 180, 182,
194, 196, 208 and 210, is present in the mass spectrum.
Furthermore the spectrum yields peaks at m}z 137, 200,
338, 356, 374 which are prominent for the Musaceae seed
coats. The lignin pyrolysis products and the prominent
peaks for the Musaceae are not extractable and are part of
the polymer system of the Musaceae seed coat, as shown in
Fig. 8. The presence of these compounds is also confirmed
by PyGCMS as shown in Fig. 9 and Table 2.
The spectra of the extracted and non-extracted seed coat
show also pyrolysis products indicating the presence of
polysaccharides as shown by m}z 43, 57, 60, 73, 126 and 144
for hexose sugars and m}z 58, 85 and 114 for pentose sugars
(Pouwels, Eijkel and Boon, 1989). The presence of polysaccharides is confirmed by ammonia chemical ionization.
Figure 10 shows the pyrolysis ammonia chemical ionization
mass spectrum of the extracted seed coat of Musa. This
spectrum contains mass peaks for ammoniated hexose sugar
monomers (m}z 134, 144, 162, 180, 222) which are prominent
for cellulose and ammoniated pentose sugar monomers
(m}z 132, 150) and dimers (m}z 264, 282), which are
prominent for hemi-cellulose (Pouwels and Boon, 1990).
The abundance of dimeric and trimeric sugars is low due to
the presence of anorganic matter as indicated by the relative
high m}z 134 (Scheijen and Boon, 1989). The ammonia CI
data also confirm the presence of guaiacyl (m}z 163) and
syringyl (m}z 193) lignin (Hage, Mulder and Boon, 1993 ;
Hage, Weeding and Boon, 1995).
In the mass spectrum (Figs 5, 8) of the extracted and nonextracted seed coat samples mass peaks are observed for
phenolic compounds (m}z 94, 110, 124), which amongst
others can indicate the presence of vegetable polyphenols.
These compounds are also components of the polymeric
system and appear at high temperatures in the mass
chromatogram as shown in Fig. 7. The presence of these
compounds is confirmed with Py (EI) GCMS.
Monocotyledonous plants usually have relatively high
m}z 120 and 150 for p-coumaric acid and ferulic acids either
present ester bound to the polysaccharide fraction or ether
bound to the true lignin structure (Boon, 1989). Also the
extracted seed coat of the monocotyledonous Musaceae
contains relative high amounts of these phenolic acids. The
Py-EI-MS (Fig. 8) shows peaks indicating ester bound (m}z
120) and ether bound (m}z 164) p-coumaric acid, and ester
bound (m}z 150) and ether bound (m}z 194) ferulic acid.
Also the ammonia CI spectrum indicates the presence of pcoumaric acid and ferulic acid as indicated by the
pseudomolecular ions m}z 121 (vinylphenol) and m}z 151
(vinylguaiacol). The PyGCMS of the TMAH treated
samples of the seed coat confirm the presence of these
compounds as shown in Fig. 11. In this figure the mass
chromatograms of the masses m}z 192 and m}z 222 indicate
p-coumaric acid and ferulic acid, respectively (Mulder,
Hage and Boon, 1992).
The Musaceae seed coat contains a number of characteristic compounds. The Py-EI-MS shows a peak at m}z
194, which is still present after extraction. This peak is
identified with Py-EI-GCMS as a caustic secondary metabolite zingirone. However, the most remarkable peaks are
aromatic compounds which all have the fragment ion m}z
137 (hydroxymethoxybenzyl cation). Additional mass
spectrometric techniques did not provide sufficient information to identify these compounds.
GraŠen et al.—Composition of Musaceaous Seed Coats
115
×2
46
100
pc
120
pc
61
137
164
85
58
151
180
114
73
98
126
t
200
Relative abundance
t
0
40
60
100
80
120
140
160
180
200
×2
100
224
338
244
272
356
374
302
256
0
220
240
260
280
320
300
340
360
380
m/z
F. 8. Pyrolysis (EI) mass spectrum of the seed coat of Musa Šelutina after extraction with hexane}dichloromethane and acetone}ethanol. D,
guaiacyl lignin ; E, syringyl lignin ; _, polysaccharides ; u, lipids ; q, Musaceae characteristics ; f, ferulic acid ; pc, p-coumaric acid ; t, vegetable
polyphenol markers.
DISCUSSION
The main structure of the seed coats of the Musaceae taxa
that have been examined are similar. As mentioned before,
vestiges of the aril are still present on the mature seed,
although it is easily released, especially when the aril and
exotesta cells are dried out and compressed. Musaceae have
fleshy fruits and are predominantly zoochorously dispersed.
It seems that the aril has lost its function during evolution
and probably the fruit has taken over the nutritive function
for the dispersal of the seeds by animals (Friedrich and
Strauch, 1975).
Very characteristic in Musaceae is the separation of the
outer cell wall layer from the silicified exotesta. The silica on
116
GraŠen et al.—Composition of Musaceaous Seed Coats
31
100
29
46
34
54
Relative intensity
27
50
23
30
36
32
9
53
26
1
50
28
35
7
8
4
2 5
6
3
33
18
13 14
1112 15
17
10
16
21
19
20
40 4345 47
3839
37
22
24
25
41
42 44
51
48 52
49
55
56
71
62
74
76 78
57
61
58
5960
69
64
63 65
66
67
68
70 72
73
79
0
1000
10:58
2000
21:55
3000
32:52
4000
43:49
scan number
retention time
F. 9. Total ion chromatogram of a pyrolysis (EI) gaschromatograph mass spectrometric run of a solvent extracted seed coat of Musa Šelutina.
the exposed surface of the mature seed coat of the Musaceae
may function as a mechanical barrier against environmental
forces, pathogens, chewing insects, other predators, and
passing alimentary tracts of dispersers. In combination with
vegetable polyphenols, which are located more to the inner
side of the seed coat, in the mesotesta and in the tegmen, it
can be a sufficient barrier against fungi. In other monocotyledons with silicified seed coats more often the silica is
located in the endotesta. In Commelinaceae the siliciferous
endotestal layer splits along its radial walls as a result of
which the exo- and mesotestal layers of the seed become
separated. Netolitzky (1926) mentioned the occurrence of
siliciferous endotestas in Commelinaceae, Zingiberaceae
and of a siliciferous exotesta in Musaceae, and suggested a
protection of embryo and endosperm as one of the main
functions.
The silica layer may help to prevent the seed coat from
oppression and shrinking during germination. The role of
silica in the cell wall seems to be analogous to that of lignin
as a compression resisting element, i.e. more rigid replacement for the water between the microfibrils and other
carbohydrate components of the wall of non-lignified cells
(Raven, 1983). Furthermore, the silica can also restrict
water loss by hydration of the silica gel (Kaufman et al.,
1981 ; Yoshida, Ohnishi and Kitagishi, 1962).
The thick cuticle at the innermost side of the seed coat
may function as a last barrier to water diffusion in
combination with outer layers. The tightly packed, thick
walled cells of the mesotesta are probably impermeable for
water by the presence of water repellent substances, such as
lignin, deposited in the polymeric network of the cell walls.
Also the unknown phenolic compounds may contribute to
a tight polymeric network. The vegetable polyphenolic
substance in the tegmen probably hampers the diffusion of
oxygen through the seed coat (Guan, 1991). In addition, the
other possible openings at the micropylar side and the
chalazal side (chalazal disk) of the seed coat are blocked.
The germination of Musa is variable and relatively
difficult under artificial conditions. Stotzky et al. (1962)
scarified the lateral side of Musa balbisiana seeds, which
raised the germination percentage up to 80 % and shortened
the time to germinate from 3–6 weeks to 6–10 d. Probably
water entry and gas exchange are made possible by to the
interruption of the silica layer, cuticle, and mesotesta.
The hard seed coat of Musa leads to conflicting demands
of protection and germination. Hard seed coats offer an
efficient protection during maturation, dispersal and dormancy ; however, it hampers germination, because the
embryo needs strong forces to rupture the seed coat.
Germination lids with predetermined rupture layers facilitate germination. In Musaceae germination takes place
by squeezing out the operculum as a result of the elongating
radical-hypocotyl axis, without rupturing the seed coat
(McGahan, 1961 b). Removal of the operculum of the
GraŠen et al.—Composition of Musaceaous Seed Coats
117
T     2. PyGCMS data on Musa velutina seed coat. Identified and unknown peaks from a chromatogram after extraction.
From eŠery peak the molecular ion is giŠen (M+d) ; PS, Polysaccharide ; L, Lignin ; Lp, Lipid ; n, aromatic phenols of unknown
structure with fragment ion m}z 137
Peak
no.
Compound
Origin
Scan
M+d
120
162
226
235
252
262
291
302
321
409
491
500
526
531
553
591
628
637
718
789
872
958
1038
1128
1212
1263
1280
1469
1527
1550
1578
1711
1740
1781
1837
1923
2031
2047
2062
2094
2101
2080–
2175
2172
2229
2246
2294
2383
44
72
86
72
82
82
60
70
74
96
84
74
92
102
84
96
82
96
116
84
98
110
94
112
108
108
124
122
138
110
120
152
124
150
154
152
168
164
178
166
178
162
2391
2397
2482
2518
2518
2534
182
194
194
196
178
202
2653
2679
192
200
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
carbon dioxide
propanedialdehyde
2,3-butanedione
2-butanone
2-methylfuran
3-methylfuran
acetic acid
2-butanol cis}trans
hydroxypropane
2,5-dimethylfuran
but-3-enal-2-one
3-hydroxypropanal
unknown
pyruvic acid
(2H)-furan-3-one
3-furaldehyde
4,2-pentadienal
2-furaldehyde
1-(acetyloxy) propane-2-one
(5H)-furan-2-one
2,3-dihydro-5-methylfuran-2-one
5-methyl-2-furaldehyde
phenol
2-hydroxy-3-methyl-2-cyclopentene-1-one
4-methyl-phenol
2-methyl-phenol (o-cresol)
guaiacol
2-ethylphenol
4-methylguaiacol
1,2 dihydroxybenzene
4-vinylphenol
guaiacylethane (¯4-ethylguaiacol)
methyl-1,2-dihydroxybenzene
4-vinylguaiacol
syringol
guaiacolaldehyde
4-methylsyringol
4-(trans-2-propenyl)guaiacol
4-butene guaiacol
4-acetyl guaiacol
methyl 2-isopropylbenzoate
levoglucosan
43
44
45
46
47
52
methyl α-methyl cinnamate
unknown lignin component
4-vinylsyringol
methyl β-methyl cinnamate
2-butanone, 4-(4 hydroxy-3 methoxy
phenyl ; zingirone
syringylaldehyde
4-(prop-cis-2-enyl-syringol
4-(prop-trans-2-enyl-) syringol
4-acetyl syringol
4-(prop-2-enol) guaiacol
unknown
53
54
unknown
unknown
55
unknown
*
2688
202
56
unknown
*
2720
202
57
phtalaat
artefact
2750
(149)
48
49
50
51
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
PS
L
L
L
L
L
L
L
L
L
L
L
L
L
PS
L
L
L
L
L
L
L
*
Remarks
m}z 65, 77, 92
pc
f
176
178
180
176
194
m}z 137, 159, 169, 187,
202
m}z 89, 117, 145, 177, 192
naphthalene skeleton, m}z
128, 139, 157, 185, 200
m}z 137, 159, 169, 187,
202
m}z 137, 159, 169, 187,
202
118
GraŠen et al.—Composition of Musaceaous Seed Coats
T     2. (cont.)
Peak
no.
Compound
Origin
Scan
M+d
58
unknown
*
2773
220
59
unknown
*
2796
220
60
61
62
phtalaat
C : fatty acid
"' !
unknown
artefact
Lp
2888
2920
3028
(149)
256
228
63
64
65
66
67
C : fatty acid
") #
unknown
C : fatty acid
") "
C : fatty acid
") !
unknown
Lp
*
Lp
Lp
*
3155
3161
3167
3200
3552
280
264
282
284
288
68
69
70
phtalaat
3-methoxy-4,4«-hydroxy stilbene
unknown
artefact
L
*
3665
3752
3893
(149)
272
338
71
unknown
*
3902
336
72
unknown
*
3945
338
73
unknown
*
3955
338
74
unknown
*
4026
350
75
unknown
*
4062
342
76
unknown
*
4111
358
77
unknown
*
4137
334
78
79
unknown
unknown
*
*
4150
4333
340
340
Musaceae seeds permitted water to reach the embryo, but
did not result in a higher percentage of germination. This
indicates that water entering the seed is unable to hydrate
the embryo. So Musaceae also have an embryo imposed
dormancy.
The mass spectrometric ‘ fingerprint ’ of the macromolecular composition of Musaceae shows some unique
and characteristic mass peaks. Although no structure can be
assigned, these mass peaks seem to be specific for Musaceae,
and will be very useful as an additional data set for
macrosystematic research of Angiosperm families (Graven,
unpubl. res.). Further studies are in progress to obtain more
information about the chemical structure of these unknown
Musaceae components.
With the phloroglucinol}HCl test or other tests no lignin
components could be detected, although the test with lignin
pink gave the indication of the presence of lignin in the S1
layer of the secondary wall of the mesotesta. These findings
are supported by the results of McGahan (1961 a), who also
obtained negative results for lignin when routine tests for
lignin were used. Also Stover and Simmonds (1987)
Remarks
m}z 137, 169, 187, 202,
220
m}z 137, 151, 162, 202,
220
m}z 77, 105, 123, 151, 185,
228
m}z 137, 151, 216, 264
m}z 137, 149, 225, 241,
257, 273, 288
m}z 137, 141, 169, 183,
201, 214, 338
m}z 137, 152, 169, 181,
197, 211, 275, 303, 321,
336
m}z 137, 152, 169, 181,
197, 211, 275, 303, 321,
336
m}z 137, 141, 169, 183,
201, 214, 310, 338
m}z 137, 141, 169, 195,
200, 310, 338, 350
m}z 137, 151, 163, 281,
342
m}z 137, 151, 163, 179,
190, 222, 340, 358
m}z 137, 151, 163, 175,
188, 202, 231, 270, 274,
281, 303, 317, 334
m}z 137, 151, 203, 340
m}z 137, 151, 163, 175,
190, 203, 340
described the seed coat as not lignified. The PyMS fingerprint
of Musaceae indicated the presence of guaiacyl and syringyl
lignin components. This observation was confirmed with Py
(EI) GCMS.
Harris and Hartley (1976) stated that cell walls containing
ferulic acid bound to polysaccharides give a negative
phloroglucinol}HCl test for lignin. The macromolecular
data of Musaceae show that the seed coat indeed contains
ferulic acid, but is also lignified. A combination of several
components in the cell wall is probably the reason why most
stains are negative.
Monocotyledonous plants usually have relatively high
m}z 120 and 150 from p-coumaric and ferulic acids, either
present ester bound to the polysaccharide fractions or ether
bound in the true lignin structure (Boon, 1989). Ferulic acid
bound to the primary wall has been recorded in nearly half
of the monocotyledon families and in about 10 families of
the dicotyledons only (Harris and Hartley, 1980).
Many studies have shown that phenolic acids are able to
affect germination, but usually via the germination rate
(time needed for germination) rather than via the total seed
GraŠen et al.—Composition of Musaceaous Seed Coats
119
180
100
134
pc
151
150
pc
121
132
162
193
144
98
224
222
Relative abundance
0
80
60
100
120
140
160
180
200
220
×5
100
242
339
356
391
279
264
282
376
300
0
240
260
280
300
340
320
360
380
400
m/z
F. 10. Pyrolysis (ammonia CI) mass spectrum of the extracted seed coat of Musa Šelutina. D, Guaiacyl lignin ; E, syringyl lignin ; _,
polysaccharides ; q, Musaceae characteristics ; U, phenolic compounds ; f, ferulic acid ; pc, p-coumaric acid.
germination (Kuiters, 1990). Seeds of Musa Šelutina and
Musa rosacea germinated relatively easily in 3–8 weeks.
Within the Musa seed coat the phenolic acids seem to be
genuine structural components of the cell walls, not affecting
germination.
The use of histochemical methods for the detection of
lipids shows the limitation of this approach (Gahan, 1984).
There is no specific functional group in lipids for
attachement of staining material. If a lipid is attached to
other macromolecular components or even not attached but
covered by other macromolecules, the result of a staining
will be negative. The presence of the insoluble cutin polymer
in the cuticle is not specifically detectable with histochemical
methods. The PyMS and PyGCMS data still contained C
"'
and C fatty acids after extraction of lipids, which can
")
indicate the presence of insoluble polymeric cutin. The
120
GraŠen et al.—Composition of Musaceaous Seed Coats
OCH3
C
C
1000
OCH3
C
O
222
OCH3
C
C
O
192
C OCH3
OCH3
TIC
2000
3000
4000
Scans
F. 11. Py (EI) GCMS partial mass chromatogram of the TMAH treated samples of the extracted seed coat of Musa Šelutina for m}z 192
(p-coumaric acid) and m}z 222 (ferulic acid).
evidence for the presence of cutin hydroxy fatty acids was
given by using methylated samples of the extracted seed
coat using Py(EI)GCMS and Py(NH -CI)GCMS.
$
Waxes are also associated with the polymeric material in
the cuticle to provide an effective diffusion barrier
(Kolattukudy, Espelie and Soliday, 1981). This may explain
the relatively high contribution of wax esters in the mass
spectrum of the Musa seed coat. The cuticle between tegmen
and the remains of the nucellus of the Musaceae have
several functions as a last barrier against water loss, ion
diffusion and several pathogens.
The mass spectrometric techniques are sensitive and
specific methods in identifying macromolecular compounds.
It is not always possible to locate the different compounds
in the differentiated tissues of the seed coat, unless these can
be separated from each other and analysed separately. The
isolation of the different seed coat sections by hand is still
the limiting factor. Histochemical methods are mainly
useful in localising certain compounds. However, it has
been shown that especially in mature seed coats the stains
are not always succesful in attaching a specific compound.
Many pyrolysis products were identified using PyGCMS,
confirming the general findings of the PyMS spectra.
Caustic products are widely distributed within the family
Zingiberaceae. Members of the family Zingiberaceae have
attracted continuous phytochemical interest due to their
considerable importance as natural spices or as medicinal
plants (Pandji et al., 1993). In literature a lot of rhizomes
and seeds are described as tasting caustic (Hegnauer,
1962–1990). Taxa of Zingiberaceae have been studied for
insecticidal activity (Grainge and Ahmed, 1988 ; Pandji et
al., 1993). The identification of zingirone in the Musa seed
coat confirms the relationship of the Musaceae with
Zingiberaceae. Zingirone was first described in the rhizome
of Zingiber officinale. The presence of products as zingirone
in the seed coat of Musa is another defence line against
specific chewing insects and other predators.
The thermal stability of condensed vegetable polyphenols
is high. The occurrence of the components 1,2dihydroxybenzene
(m}z
110)
and
methyl-1,2dihydroxybenzene (m}z 124) indicates the presence of
proanthocyanidins. Lignins have been reported to contain
only negligible amounts of catechol (Galletti and Reeves,
1992). And so it is reasonable to use catechol as a diagnostic
fragment for condensed vegetable polyphenols (Galletti and
Reeves, 1992). Vegetable polyphenols are extremely complex
compounds. They may be highly specific in at least some
and probably most of their possible interactions, via an
enormous potential for stereo specificity (Zucker, 1983).
Within the seed vegetable polyphenols have several mixed
functions, involving a defence mechanism against living
plant enemies, a delay in decomposition when the seed is a
component of the seed bank, and influencing dormancy in
seeds.
The real cause of dormancy of Cornus seeds is due to a
high level of vegetable polyphenols in the seed coat
restricting the embryonal development (Guan, 1990).
Proanthocyanidins are present in most families of the
monocotyledons. According to Dahlgren, Clifford and Yeo
(1985) cells with vegetable polyphenols of the Musaceae
may contain proanthocyanidins. Ellis, Foo and Porter
(1983) have described the occurrence of proanthocyanidins
GraŠen et al.—Composition of Musaceaous Seed Coats
in the fruit skin of Musa sapientum. Within the line of this
study no specific information on vegetable polyphenolic
structures has been found.
Many monocotyledons have anatomically relatively
simple seed coats. Endotestal seeds as found in Dioscoreales,
Zingiberales, Commelinales and others may be original for
monocotyledons. The seed coat structure of the Musaceae is
characteristic at the family level and the most complex one
within the Zingiberales. Very remarkable are the micropylar
collar, the relative thick mesotesta, the unique macromolecular ‘ fingerprint ’ with the typical Musaceae phenolic
compounds of still unknown composition, and the breaking
of the cell walls in the exotestal layer resulting in a surface
with silica crystals. More often the endotesta is silicified, as
in Commelinaceae, Strelitziaceae, Rapateceae, and
Marantaceae. In Commelinaceae also a silicified surface
remains due to a breaking of the endotesta. The Zingiberales
is one of the most indisputable natural suprafamilial groups
(Dahlgren et al., 1985). The seed coat structure of the
Musaceae is characteristic and the most complex one of the
Zingiberales, whereas that of the Cannaceae is considered as
the most derived one within this order. The rather complex
seed coat is characteristic for this family and seems to be
unique if compared to other families of the Zingiberales and
to other monocotyledons.
A C K N O W L E D G E M E N TS
The authors wish to thank J. B. M. Pureveen, N. Devente,
J. Dahmen, F. D. Boesewinkel and M. M. Mulder for
technical assistance. Special thanks are for B. Ursem of the
Hortus Botanicus in Amsterdam for obtaining most of the
seed material and W. J. Kress for sending seeds of the genus
Musella.
The investigations were supported by the Life Science
Foundation (SLW), which is subsidised by the Netherlands
Organisation for Scientific Research (NWO).
LITERATURE CITED
Boon JJ. 1989. An introduction to pyrolysis mass spectrometry of
lignocellulosic material : case studies on barley straw, corn stem
and Agropyron. In : Chesson A, Ørskov ER, eds. Physicochemical
characterization of plant residues for industrial and feed use.
London : Elsevier Applied Science, 25–45.
Bouharmont J. 1963. Evolution de l’ovule fe! conde! chez Musa acuminata
Colla subsp. burmannica Simmonds. La Cellule 63 : 261–279.
Cronquist A. 1981. An integrated system of classification of flowering
plants. New York : Columbia University Press.
Dahlgren RMT, Clifford HT, Yeo PF. 1985. The families of the
monocotyledons : Structure, eŠolution and taxonomy. Berlin :
Springer Verlag.
Dahlgren RMT, Rasmussen FN. 1983. Monocotyledon evolution.
Characters and phylogenetic estimation. In : Hecht MK, Wallace
B, Prance GT, eds. EŠolutionary botany Vol. 16. New York :
Plenum Publishing Corporation.
Davis GL. 1966. Systematic embryology of the Angiosperms. New York,
John Wiley & Sons, Inc.
Dey SUD, Basu Baul TS, Roy B, Dey D. 1989. A new rapid method of
air drying for SEM using tetramethylsilane. Journal of Microscopy
156 : 259–261.
Ellis CJ, Foo LY, Porter LJ. 1983. Enantiomerism : A characteristic of
the proanthocyanidin chemistry of the Monocotyledonae. Phytochemistry 22 : 483–487.
121
Feder N, O’Brien TP. 1968. Plant microtechnique : some principles and
new methods. American Journal of Botany 55 : 123–142.
Friedrich WC, Strauch F. 1975. Der Arillus der Gattung Musa.
Botaniska Notiser 128 : 339–349.
Gahan PB. 1984. Plant histochemistry and cytochemistry. London :
Academic Press.
Galletti GC, Reeves JB. 1992. Pyrolysis}gas chromatography}iontrapdetection of polyphenols (vegetable tannins) : preliminary
results. Organic Mass Spectrometry 27 : 226–230.
Grainge M, Ahmed S. 1988. Handbook of plants with pest-control
properties. New York : John Wiley.
Guan KL. 1990. Studies on dormancy of Cornus officinalis seed.
International Conference on Seed Science and Technology, 12–15
March 1990, Hangzhou, China, 52.
Guan KL. 1991. Studies on complex dormancy of Cornus officinalis
seed. Chinese Journal of Botany 3 : 145–150.
Hage ERE van der, Mulder MM, Boon JJ. 1993. Structural characterization of lignin polymers by temperature-resolved in-source
pyrolysis-mass spectrometry and Curie-point pyrolysis-gas
chromatography}mass spectrometry. Journal of Analytical and
Applied Pyrolysis 25 : 149–183.
Hage ERE van der, Weeding TL, Boon JJ. 1995. Ammonia chemical
ionization mass spectrometry of substituted phenylpropanoids
and phenylalkyl phenyl ethers. Journal of Mass Spectrometry 30 :
541–548.
Harris PJ, Hartley RD. 1976. Detection of bound ferulic acid in cell
walls of the Graminae by ultraviolet fluorescence microscopy.
Nature 259 : 508–510.
Harris PJ, Hartley RD. 1980. Phenolic constituents of the cell walls
monocotyledons. Biochemical Systematics Ecology 8 : 153–160.
Hegnauer R. 1962–1990. Chemotaxonomy der Pflanzen Vol. 1–9. Basel :
Birkha$ user Verlag.
Kaufman PB, Dayanandan P, Takeoka Y, Bigelow WC, Jones JD, Her
R. 1981. Silica in shoots of higher plants. In : Simpson TL , Volcani
BE, eds. Silicon and siliceous structures in biological systems.
Berlin : Springer Verlag, 409–449.
Kolattukudy PE, Espelie KE, Soliday CL. 1981. Hydrophobic layers
attached to cell walls, cutin, suberin and associated waxes.
Encyclopaedia of Plant Physiology, New Series 13B : 225–254.
Krisnamurthy KV. 1988. Methods in plant histochemistry. Chetput,
Madras : S. Viswanathan (Printers & Publishers) Private Limited.
Kuiters AT. 1990. Role of the phenolic substances from decomposing
forest litter in plant soil interactions. Acta Botanica Neerlandica
39 : 329–348.
Manchester SR, Kress WJ. 1993. Fossil bananas (Musaceae) : Ensete
oregonense sp. nov. from the Eocene of western North America
and its phytogeographic significance. American Journal of Botany
80 : 1264–1272.
McGahan MW. 1961 a. Studies on the seed of banana. I. Anatomy of
seed and embryo of Musa balbisiana. American Journal of Botany
48 : 230–238.
McGahan MW. 1961 b. Studies on the seeds of banana. II. The
anatomy and morphology of the seedling of Musa balbisiana.
American Journal of Botany 48 : 630–637.
Mulder MM, Hage ERE van der, Boon JJ. 1992. Analytical in source
pyrolytic methylation electron impact mass spectrometry of
phenolic acids in biological matrices. Phytochemical Analysis 3 :
165–172.
Nakai T. 1941. Notulae ad Plantas Asiae Orientalis (XVI). Journal of
Japanese Botany 17 : 189–203.
Netolitzky F. 1926. Anatomie der Angiospermen-Samen. Handbuch der
Pflanzen Anatomie Band X (K. Linsbauer). Berlin : Gebru$ der
Borntraeger.
Pandji C, Grimm C, Wray V, Witte L, Proksch P. 1993. Insecticidal
constituents from four species of the Zingiberaceae. Phytochemistry 34 : 415–419.
Pastorava I, Oudemans TFM, Boon JJ. 1993. Experimental polysaccharide chars and their ‘ fingerprints ’ in charred archaeological
food residues. Journal of Analytical and Applied Pyrolysis 25 :
63–75.
Pouwels AD, Boon JJ. 1990. Analysis of beech wood samples, its milled
122
GraŠen et al.—Composition of Musaceaous Seed Coats
wood lignin and polysaccharide fractions by Curie-point and
platinum filament pyrolysis mass spectrometry. Journal of Analytical and Applied Pyrolysis 17 : 97–126.
Pouwels AD, Eijkel GB, Boon JJ. 1989. Curie-point pyrolysis-capillary
gas chromatography-high-resolution mass spectrometry of microcrystalline cellulose. Journal of Analytical and Applied Pyrolysis
14 : 237–280.
Raven JA. 1983. The transport and function of silicon in plants.
Biological ReŠiew 58 : 179–207.
Scheijen MA, Boon JJ. 1989. Loss of oligomer information in pyrolysisdesorption}chemical-ionization mass spectrometry of amylose
due to the influence of potassium hydroxide. Rapid Communication
Mass Spectrometry 7 : 238–240.
Schmid R, Turner MD. 1977. Contrad 70, an effective softener of the
herbarium material for anatomical study. Taxon 26 : 551–552.
Setoguchi H, Tobe H, Ohba H. 1992. Seed coat anatomy of Crossostylis
(Rhizophoraceae) : its evolutionary and systematic implications.
Botanical Magazine Tokyo 105 : 625–638.
Shepherd K. 1954. Seed fertility of ‘ Gros Michel ’ bananas. Journal of
Horticultural Science 29 : 1–11.
Simmonds NW. 1960. Notes on banana taxonomy. I Two new species
of Musa. Kew Bulletin 14 : 198–212.
Stotzky G, Cox EA, Goose RD. 1962. Seed germination studies in
Musa. I Scarification and aseptic germination of Musa balbisiana.
American Journal of Botany 49 : 515–520.
Stover RH, Simmonds NW. 1987. Bananas. Tropical Agricultural Series
third ed. London : Longman Scientific & Technical.
Takhtajan A . 1985. Anatomia seminum comparatiŠa. Tomus 1 Liliopsida
seu monocotyledones. Leninpoli : Nauka.
Tomlinson PB. 1962. Phylogeny of the Scitaminae. Morphological and
anatomical considerations. EŠolution 16 : 192–213.
White PR. 1928. Studies on the bananas. An investigation of the floral
morphology and cytology of certain types of the genus Musa L.
Zeitschrift fuX r Zellforschung und Mikroskopische Anatomie 7 :
673–735.
Yoshida S, Ohnishi Y, Kitagishi K. 1962. Histochemistry of silicon in
rice plant. I A new method for determining the localization of
silicon within plant tissues. Soil Science and Plant Nutrition 8 :
30–35.
Zucker WV. 1983. Tannins : does structure determine function ? An
ecological perspective. The American Naturalist 121 : 335–365.