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2007 SOUTHEASTERN NATURALIST 6(1):47–66
Anatomy of the Laminar Organs of Commelina erecta
Roland R. Dute1,*, Brian E. Jackson1, Ryan D. Adkins1,
and Debbie R. Folkerts1
Abstract - A study was undertaken to compare the anatomy of the laminar floral parts
with that of the spathes and leaves of Commelina erecta L. Each flower has two types
of petals and two types of sepals. In contrast to the other organs, the petals have a
completely open venation system whose vein endings consist solely of modified
bundle-sheath cells. Bundle sheaths of leaves and spathes, but not the floral organs,
contain sclerified cells for support. The high density of hook-shaped trichomes on the
outer surface of the spathe and of glandular microhairs on the inner surface might
indicate protective and secretory functions, respectively. Anomalous stomatal apparatuses
are more common on floral organs than on spathes or leaves. Leaves and spathes
appear to have a more detailed developmental program than sepals and petals.
Commelin (erect dayflower), with 170 species worldwide, is the largest
genus within the Commelinaceae. Nine species are found in the US and only
three of them are native (Faden 1993, 2000). Commelina erecta L. is the most
widespread of the native species (Faden 1993). Its range includes much of the
East Coast, Southeast, and southern Midwest (Faden 2000). The flowers have
been thoroughly described for taxonomic purposes as having two different
types of petals (two that are large, blue, and clawed; one that is small and
colorless) and two different sizes of sepals (Brashier 1966, Pennell 1916,
Radford et al. 1968). Thus, along with the leaves and spathes, which enclose the
inflorescences, there are six different laminar organs in C. erecta. Since floral
organs represent modified leaves (Weberling 1992), we decided to undertake a
comparative anatomical study of the various laminar organs of C. erecta, with
emphasis on vasculature and arrangement of stomatal apparatuses.
Materials and Methods
Specimens of vegetative and floral organs were collected from the property
of the senior author in Lee County, AL. Some specimens were viewed in
the living condition using bright-field microscopy. This technique was especially
effective for studying the floral organs that were only a few cell layers in
thickness. Other material was fixed in FAA (90 ml 50% ethanol, 5 ml glacial
acetic acid, 5 ml formalin; Johansen 1940), then cleared in 50% ethanol and
stored in the same fluid for an extended period of time. This material was
viewed directly either using Nomarski optics (McCrone et al. 1978) or a bright
1Department of Biological Sciences, 101 Life Sciences Building, Auburn University,
AL 36849. *Corresponding author - firstname.lastname@example.org.
48 Southeastern Naturalist Vol. 6, No. 1
field microscope following staining. Stain recipes included: 1) 1% safranin in
50% ethanol, 2) 0.5% aqueous toluidine blue O (TBO) in 0.02 M sodium
benzoate buffer, 3) 2% ferrous sulphate in acidified formalin for tannin
identification (Ruzin 1999), 4) 0.1% aniline blue in 0.1 N K3PO4 for callose
identification (Martin 1959), and 5) 1% I2KI for starch identification. Callose
distribution was detected using a fluorescence microscope.
Material was preserved and embedded in plastic resin in two different
ways. The first method involved fixation in 3% glutaraldehyde in 0.05 M
potassium phosphate buffer (pH 6.8). Dehydration to 95% ethanol was
followed by infiltration and embedment in JB-4 resin (Polysciences, Inc.,
Warrington, PA). Alternatively, some material was preserved in 3% buffered
glutaraldehyde and, after washing in buffer, postfixed in 1% buffered
OsO4. The tissue was next dehydrated in an alcohol/acetone series and
embedded in Spurr’s resin (Spurr 1969).
Resin-embedded material was sectioned (6-m thick sections for JB-4
material; 2-m for Spurr material) using a Porter-Blum MT-2B ultramicrotome.
The resulting sections were heat-fixed to glass microscope slides and
stained with TBO. The stained sections were then covered with Permount
mounting medium and a glass coverslip.
Photographs of slide material were made using either a 35-mm camera
attached to a compound microscope with Ektachrome 64T color slide film,
or a Nikon D70 Digital Camera attached to the microscope.
A dissecting microscope was used to observe both living and fixed and
cleared material via both bright-field and dark-field options. Material was
photographed with a Nikon D70 Digital Camera.
Air-dried leaves and spathes were ashed and the material viewed with
both bright-field and scanning electron microscopy (SEM) in order to investigate
mineral inclusions. Elemental analysis was accomplished with an
energy dispersive spectrometer (EDS) attachment to the same microscope
(Postek et al. 1980.)
Supplementary material was fixed in buffered glutaraldehyde, dehydrated
in ethanol followed by acetone, and dried by the critical-point method using
liquid CO2 (Postek et al. 1980). Dried material was attached to aluminum
stubs with double-stick carbon tape. Specimens were coated with a goldpalladium
vapor and viewed at 15 kV accelerating voltage using a Zeiss 940
Digital Scanning Microscope.
A voucher specimen was deposited in the Auburn University Herbarium
Results and Discussion
Flowers of C. erecta arise from within a folded leaflike spathe that
contains mucilage (Fig. 1). The leaflike nature of the spathe is evident once
the organ is removed from the plant and unfolded (Fig. 2), although its shape
differs from the lanceolate leaves. Unfolding the spathe requires some tear2007
R.R. Dute, B.E. Jackson, R.D. Adkins, and D.R. Folkerts 49
Figures 1–4. 1) Habit view of C. erecta flower and its spathe. MS = medial fertile
stamen, P = posterior petal, PS = proximal fertile stamens, S = spathe, and ST =
staminode. Scale bar = 10 mm. 2) Unfolded spathe showing its foliar outline. Scale
bar = 5 mm. 3) Size comparison of large and small petal. Both organs have been
cleared and then stained with safranin. Scale = 5 mm. 4) A composite photograph of
three sepals—two large, one small—viewed by dark-field microscopy. Note the three
main veins in each organ. Scale bar = 1 mm.
50 Southeastern Naturalist Vol. 6, No. 1
ing as the proximal margins are fused. The calyx consists of three sepals
(Fig. 4); the two anterior ones are fused into a cup, the posterior one is free,
smaller, and of a different shape. The corolla has two large, posterior, blue
petals with claws (Figs. 1, 3). The anterior petal is much smaller, colorless to
nearly so, and wedge-shaped. The difference between the two petal types is
apparent in Figure 3. The androecium consists of three proximal fertile
stamens and three distal staminodes (Fig. 1). The anther of the shorter,
median fertile stamen is of a different shape from the others. It is referred to
as the feeding anther because it is a primary attractant for insects (McCollum
et al. 1984). The gynoecium consists of three fused carpels.
The population of C. erecta produced flowers from mid-June through
early September. Three to five flowers extend (one at a time) from each
enfolding spathe. Flower extension and anthesis from a given spathe varies
from every other day to every fourth day. The period of anthesis for a given
flower is brief; flowers are about one-third open an hour before sunrise, but
are completely closed and wilted by about an hour after noon (R.R. Dute,
Comparison of sectional views of the various organs shows consistent
differences. Figure 5 shows a transection of a vegetative leaf. Anatomically,
this organ is dorsiventral with a single, well-developed layer of palisade
cells above a considerable volume of irregular, loosely packed, spongy
parenchyma cells. Both cell types are chlorophyllous. In contrast, a sectional
view of the spathe, the other photosynthetic organ, shows only a homogeneous,
chlorophyllous mesophyll (Fig. 6). The mesophyll region of the floral
organs also is homogeneous, but largely achlorophyllous (Fig. 7). Although
a faint green color can sometimes be seen in these other organs, for example
in the small sepal, the number of chloroplasts involved is miniscule in
comparison to the leaves and spathes and cannot be observed by microscopy
other than in the guard cells of the epidermis.
All organs have a considerable amount of intercellular space within the
mesophyll; however, the limb of the large petal seems to be the most
aerenchymatous of the organs (Fig. 8). The contrast in tissue density to that
of the other organs, including that of the claw of the large petal (Fig. 9), is
considerable. Added to the lack of density is the poor staining contrast of the
limb tissues due to very thin cell walls. Adequate preservation of the limb
tissues of the large petal proved difficult and could only be approached
through glutaraldehyde fixation and plastic embedment. Even so, considerable
cell shrinkage occurred.
Epidermal cells of the leaf are large and occupy a significant portion of the
cross-sectional area of the leaf (41%, range 31–50%, N = 10; Fig. 5). These
large epidermal cells are a typical feature of the Commelinaceae in general
(Tomlinson 1966) and of Commelina in particular (Brückner 1926, Preston
1898). It has been hypothesized that these enlarged cells function in water
storage (q.v. discussion in Tomlinson 1966). The ratio of epidermal area to the
2007 R.R. Dute, B.E. Jackson, R.D. Adkins, and D.R. Folkerts 51
entire cross-sectional area of the organ is significantly greater (P = 0.0027) in
the spathe (62%, range 44–75%, N = 16; Fig. 6). Enlarged size of the
epidermal cells does not extend to the guard or subsidiary cells (Figs. 5, 6).
Both leaves and spathes have the same arrangement of veins consisting
of parallel longitudinal veins connected by narrower diameter transverse
veins (Tomlinson 1969) or commissural bundles (Esau 1977) to give a
closed venation system (Fig. 10). The venation patterns of large and small
Figure 5–9. 5) Transection of vegetative leaf. AB = abaxial (lower) epidermis, AD =
adaxial (upper) epidermis, G = guard cell, P = palisade parenchyma, S = subsidiary cells,
SP = spongy parenchyma, and T = trichome. Scale bar = 50 m. 6) Transection of spathe.
The chloroplasts are visible as spots in the mesophyll (M). Note that the mesophyll is
undifferentiated unlike that of the leaf. AD = adaxial epidermis, AB = abaxial epidermis,
and G = guard cells. Scale bar = 50 m. 7) Transection of small sepal. The mesophyll
(M) is undifferentiated. Scale bar = 100 m. 8) Transection of the limb of the large petal.
The mesophyll (M) cells are separated by large intercellular spaces. The arrows indicate
attachment of the mesophyll cells to the bundle sheath of a vascular bundle. Scale bar =
25 m. 9) A transection of the claw of the large petal. The cells of the mesophyll are
more tightly packed than those of the limb. Scale bar = 50 m.
52 Southeastern Naturalist Vol. 6, No. 1
sepals are similar and consist of three main veins which not only are
interconnected by small veins but which fuse distally (Fig. 4). In addition,
there are branch veins in the large sepals that end blindly (Fig. 4). In contrast
to the organs mentioned thus far, venation of the limb of the large petal is of
a dichotomous, open type (Fig. 11). Venation of the small petal is also open,
but of a simpler architecture and with fewer veins than its larger counterpart.
Although both sepals and petals have three major veins, the origin of these
Figure 10–15. 10) Closed system of veins in spathe. C = commissural vein, and L =
longitudinal bundles. Scale bar = 0.5 mm. 11) Dichotomous open venation of large
petal (limb). Scale bar =1 mm. 12) Vein end in large petal. The arrow indicates
termination of the xylem tissue. Bundle-sheath cells extend beyond this point as a
branched vein prolongation. Scale bar = 50 m. 13) Bright-field microscopic image
of the edge of a petal showing a detail of a branched vein prolongation in a living
petal. The arrows denote projections extending from the modified bundle-sheath
cells. Scale bar = 25 m. 14) Portion of vein prolongation as seen with Nomarski
optics. The arrow indicates a nucleus. Scale bar = 25 m. 15) A vein prolongation
branching from a vein. This vein is located in the center of the limb away from the
petal’s edge. Scale bar = 25 m.
2007 R.R. Dute, B.E. Jackson, R.D. Adkins, and D.R. Folkerts 53
veins from the receptacular stele differs. Namely, the primary laterals of the
sepals share a vascular origin with the main veins of the petals, whereas the
median veins of the sepals have a separate vascular origin (Hardy and
Stevenson 2000a, b).
Detailed study of vein endings from cleared specimens of large petals
indicates that “vein prolongations,” consisting of modified bundle-sheath
cells, extend beyond the xylem and phloem for a considerable distance
(Fig. 12). The branching or “digitation” of these vein prolongations is not
uncommon (Figs. 12, 13). The parenchymatous nature of these cells is
indicated by their possession of nuclei and cytoplasm (Fig. 14). Similar vein
prolongations can occasionally be found as minor branches from a vein at
other locations in the petal (Fig. 15). Vein prolongations occur in small
petals as well, but they lack the complexity (branching) of those found in
Confirmation of the structure of vein endings in large petals comes from
serial cross sections beginning at the petal’s edge. Only bundle-sheath cells
of the vein prolongation are initially present (Fig. 16), followed by the
appearance of a sieve element (Fig. 17), in turn followed by a tracheary
element (Fig. 18). One can see the densely cytoplasmic nature of the bundlesheath
cells relative to other cells in the petal.
As the petal’s margin is approached, the number of cell layers decreases,
and near a vein’s terminus, the vascular bundle is often surrounded by only
cells of the two epidermal surfaces (Figs. 16–18). These epidermal cells are
attached to the bundle-sheath cells by lobes (Figs. 16–18). These lobes of the
epidermal cells probably result from restricted areas of cell wall growth as
the petal expands. Localization of callose in the walls of vein prolongations
is depicted using aniline blue fluorescence in Figure 19. Sites of fluorescence
probably represent clusters of plasmodesmata connecting bundlesheath
cells with epidermal lobes (Currier 1957). In addition to epidermal
lobes, cells of the vein prolongations themselves possess tiny projections
from their surfaces (Fig. 13). The function of these projections is unknown.
In more proximal parts of the petal, the organ is thicker and there are
more cell layers (referred to as the mesophyll, Fig. 8). In such instances,
the mesophyll cells are attached to the bundle sheaths by lobes (Fig. 8,
Bundle-sheath cells of the large petals contain large multigrained amyloplasts
(Fig. 20) that give a distinct positive reaction to iodine, and that, in
living cells, can be seen to move in response to cytoplasmic streaming. Size
and number of starch grains show a gradient in the bundle-sheath cells from
most numerous and largest in the proximal parts of the petal to absent near
There are two ways to view the vasculature of the large petals—ontogenetically
and functionally. According to Weberling (1992), the open venation
of perianth parts is the result of the reduced number of cell layers formed at the
margin of the organ as it continues to divide. In other words, a certain amount
54 Southeastern Naturalist Vol. 6, No. 1
of petal thickness is necessary for the veins to join into a closed system. We
hypothesize that the vein prolongations occur because there are not enough
layers of cells near the margin to support the formation of xylem and phloem
tissues. Thus, the vein prolongations represent a relictual vascular tissue that
distributes water and food to the margin.
2007 R.R. Dute, B.E. Jackson, R.D. Adkins, and D.R. Folkerts 55
The importance of vascular bundle sheaths in transport of substances to
and from vascular tissues is well-known (Ding et al. 1988; Evert et al. 1977,
1996a). The large petals and the other floral parts clearly function as sinks
and import food and water. We hypothesize that movement of nutrients from
bundle sheath cells to epidermal and/or mesophyll cells is symplastic via
intercellular connections involving the cell lobes.
Cellular structure of the bundle sheath varies within and among organs.
In general, sheath cells are elongate and appear circular when viewed in
cross section. Bundle-sheath cells are highly cytoplasmic (in contrast to the
surrounding mesophyll cells) and often contain large starch grains, especially
in the proximal region of a given organ. In the parallel, elongate
vascular bundles of leaves (except for the mid-vein), the bundle sheath has
fibrous cells with more or less thickened (sclerified), lignified secondary
walls where the sheath adjoins the phloem of the bundle (Fig. 21). Lignification
of the walls of these cells is indicated by the blue-green hue imparted by
TBO staining (O’Brien et al. 1964). In some instances, it appears as if the
sheath is double at the fiber site, with the inner row consisting of fibers and
the outer row consisting of either parenchyma cells or fibers (Figs. 21, 22).
The remainder of the sheath’s circumference consists of parenchyma cells.
In vascular bundles from smaller diameter veins, the sclerified cells are still
present but with thinner secondary walls. Commissural bundles at right
angles to and connecting with the elongate vascular bundles possess a
single-layered bundle sheath consisting only of parenchyma cells (Fig. 23).
The architecture of the bundle sheath is similar in spathes, where the elongate
vascular bundles have a sheath that has sclerified cells only opposite the
phloem. The bundle sheath of the spathe is uniseriate. Here again, the sheath
of commissural bundles is strictly parenchymatous. Tomlinson (1969) notes
that for leaves of the Commelinaceae “the sheath (is) often completed either
both above and below or only below by thick-walled, collenchymatous or
even fibrous cells” and that transverse (commissural) veins are “sheathed by
thin-walled elongated cells.”
Commelina erecta is a C3 monocot (Illinois Plant Information Network
2002), and interesting similarities can be found by comparing the anatomy of
the bundle sheath of leaf and spathe with the bundle sheath in leaf veins of
those C3 species (grasses) that have been intensively studied. Sheaths of
longitudinal veins of C3 grasses consist of two concentric layers; an outer
parenchyma sheath and an inner so-called “mestome” sheath whose walls are
Figures 16–19 (opposite page). Sequential cross sections through the petal beginning
near the margin and moving proximally. Initially, only cells of the vein prolongation
(VP) are present (16), followed by an organized bundle sheath (B) containing a single
sieve element (SE) (17). Further back from the margin, the first tracheary element (T)
of the xylem appears within the vascular bundle (18). Extensions from the epidermal
cells to the bundle sheath/vein prolongations are indicated by unlabeled arrows in
Figs. 16–18. Scale bar for Figures 16–18 = 20 m. 19) Fluorescence microscopy of
branched vein prolongation showing sites of callose deposition. Scale bar = 25 m.
56 Southeastern Naturalist Vol. 6, No. 1
asymmetrically thickened (O’Brien and Carr 1970). In barley, the thickness of
mestome sheath walls is correlated with vein diameter, with increasing thickness
from small to medium to large (Evert et al. 1996b). The walls of the
mestome sheath cells contain suberin layers (Evert et al. 1996b; O’Brien and
Carr 1970), a feature thought to restrict movement of photoassimilates from
mesophyll to bundle-sheath cells to a symplastic pathway (Evert et al. 1996b).
Experimental evidence in wheat shows that photoassimilate first enters the
smaller longitudinal veins where it is passed by transverse (commissural)
veins to the large diameter longitudinal veins for export from the leaf (Altus
and Canny 1982).
Figures 20–24. 20) Multigrained amyloplasts (S) in bundle sheath (B) of branching
vein in large petal. T = tracheary elements. Scale bar = 25 m. 21) Cross section of
longitudinal vein from leaf. The bundle sheath is a double layer (B1, B2) only adjacent
to the phloem. The inner of these two layers (B2) is sclerified. B = bundle sheath, P =
phloem tissue, X = xylem tissue. Scale bar = 10 m. 22) Another cross section of a
longitudinal leaf vein where both bundle-sheath layers opposite the phloem are
sclerified. Scale bar = 10 m. 23) Commissural bundle of leaf; leaf bundle sheath (B)
is entirely parenchymatous. Scale bar = 10 m. 24) Parenchymatous bundle sheath
(B) in vein of small petal. Scale bar = 10 m.
2007 R.R. Dute, B.E. Jackson, R.D. Adkins, and D.R. Folkerts 57
The thickened bundle-sheath cells of C. erecta do not qualify as part of a
mestome sheath because they have evenly thickened walls. Also, with few
exceptions, thickened cells are exclusively located opposite the phloem. We
hypothesize that the thick-walled cells of the vascular bundles function in
support. Brückner (1926) also refers to sickle-shaped fiber masses associated
with sieve elements in leaves of the Commelinaceae. According to him, these
fibers serve for support. Thus, in C. erecta, commissural veins would provide
little in the way of support because they consist exclusively of parenchyma
cells. The support hypothesis is strengthened by the absence of thick-walled
sheath cells in the temporary floral organs, which we hypothesize are supported
only by turgor pressure. In these cases, the sheath is a more-or-less
distinct layer of parenchyma cells (Fig. 24).
According to Tomlinson (1966, 1969), hairs on vegetative organs of the
Commelinaceae can be divided into the following: glandular microhairs
(clavate trichomes) (Fig. 25) and macrohairs. In the latter group, the types
found on Commelina are two-celled prickle hairs (Fig. 26), hook hairs (Fig.
27), and uniseriate hairs of differing numbers of cells (Fig. 27—essentially
elongate prickle hairs). EDS analysis of ashed leaves and spathes shows distal
cells of hook and uniseriate hairs to be silicified. The response of prickle hairs
to polarized light indicates possible silicification of their walls as well.
Two features of trichome distribution in C. erecta are worthy of comment
(Table 1). The first feature is the large number of hook hairs on the
abaxial surface of the spathe and their absence from the adaxial surface (Fig.
28). In contrast, the latter surface consists almost entirely of glandular
microhairs (Fig. 29). The physical barrier provided by hook-shaped trichomes
to insects is often lethal (q.v. Gilbert 1971 for a classic example),
and the protection provided to the enclosed inflorescence makes perfect
sense. Glandular microhairs of the adaxial (enclosed) surface of the spathe,
in contrast, might be responsible for the mucilaginous substance that occupies
the cavity formed by the folded spathe. Three-celled glandular hairs are
implicated in mucilage secretion in Tradescantia zebrina Heynh. (inchplant)
(Commelinaceae) (Thaler et al. 2001, as Zebrina pendula Schnizl.).
Table 1. Distribution of trichomes on the surface of various organs. ab = abaxial surface, ad =
adaxial surface, and r = rare.
Microhairs Prickle hairs Hook hairs Uniseriate hairs
Large petal (ad and ab)
Small petal (ad)
Small petal (ab) Y Y (r)
Large sepal (ad) Y cluster Y (r)
Large sepal (ab) Y Y (r)
Small sepal (ad) Y cluster
Small sepal (ab) Y cluster
Spathe (ad) Y Y
Spathe (ab) Y Y Y
Leaf blade (ad) Y Y Y
Leaf blade (ab) Y Y Y
Leaf margin Y
58 Southeastern Naturalist Vol. 6, No. 1
The second feature of note is the clustering of glandular microhairs at the
termination of the main veins of large sepals (adaxial surface) and small
sepal (both surfaces) (Fig. 30). In such instances, the nature of the secretory
material, if any exists, is unknown, and according to numerous studies
(summarized in Fahn 2000), the substance could be one of many different
chemical compounds. We would hypothesize, however, that the secreted
material is water soluble and is transported in the xylem to the vein ending
(in either its final form or as a precursor substance) and transferred to the
cells of the trichomes for secretion.
Figures 25–30. Trichomes. 25) Glandular microhairs on abaxial surface of small
petal. Scale bar = 50 m. 26) Two-celled prickle hair on leaf margin. Scale bar = 25
m. 27) Hook hairs (H) and uniseriate hairs (U) on the abaxial leaf surface. Scale bar
=100 m . 28) Hairs on abaxial (outer) surface of spathe (SEM). Scale bar = 100 m.
29) Glandular microhairs on adaxial (inner) side of spathe (SEM). Scale bar = 200
m. 30) Cluster of glandular microhairs on abaxial surface of small sepal. Scale bar =
2007 R.R. Dute, B.E. Jackson, R.D. Adkins, and D.R. Folkerts 59
Stebbins and Jain (1960) have given a detailed description of the structure
and development of the stomatal apparatus in leaves of Commelina
communis L. (Asiatic dayflower), and Kaushik (1971) has provided information
on the structure of the stomatal apparatus in five other species of
Commelina. According to Stebbins and Jain (1960), the mature apparatus
consists of the two guard cells whose aperture is “oriented parallel to the
long axis of the leaf.” The guard cells, in turn, are surrounded by six
subsidiary cells—a terminal subsidiary cell proximal (basal) to the guard
cell pair, one distal (above) to the pair, and a pair of subsidiary cells
(laterals) on either flank of the guard cells. The situation is the same for
leaves of C. erecta (Fig. 31). In contrast, Preston (1898) considers the
number of subsidiary cells in C. nudiflora (in old literature = C. diffusa
Burm. f.) or C. communis (identity uncertain) to be four, but this observation
was based only on cross-sectioned leaves and spathes.
In the present study, the structure of the stomatal apparatus was investigated
using both regular bright-field microscopy (Fig. 31) and Nomarski
optics (Fig. 32). The latter technique, in particular, enables one to clearly
distinguish cell walls and nuclei as well as the numerous, distinct plastids
that are a feature of the guard cells.
All laminar organs investigated in this study have stomatal apparatuses,
although the frequency, arrangement, and structure of these features vary
among organ types, and indeed, vary between adaxial and abaxial epidermal
surfaces of a given organ. Stomata are densely distributed over the abaxial
(lower) epidermis of the leaf, and while still common, are less densely
distributed on the adaxial (upper) epidermal surface, typically in the costal
regions. Stomata are densely distributed over the abaxial (outer) surface of
the spathe, but, with the exception of a ring of stomata just inside the rim
of the spathe’s opening (over the marginal vein), are absent from the adaxial
(inner surface). Few to no stomata exist on the adaxial surface of large
sepals, but they are more common on the abaxial surface. In the latter case,
most stomata are clustered over and around the vascular nexus formed by the
anastomosis of the three main veins at the distal portion of the organ. A few
stomata are also present on the costal region of the three main veins proximal
to the nexus. The situation regarding the small sepal is similar to that of
the large sepals, with more stomata on abaxial than adaxial surface. For
example, counts of two adaxial surfaces gave numbers of only three and
seven, whereas numbers of stomata on the corresponding abaxial surfaces
were 34 and 30. On the abaxial surface, stomata are clustered over and
around the vein nexus, with decreasing numbers located over the three main
veins proximal to said nexus. The small petal, in contrast to the other organs,
possesses a higher frequency of stomata on the adaxial surface (in a band
from base to tip), whereas the abaxial epidermis has very few. No stomata
have been found on the abaxial surface of the large petal, but a few scattered
ones are present on the adaxial surface. Perhaps the near absence of stomata
60 Southeastern Naturalist Vol. 6, No. 1
Figures 31–36. Stomata. Figs. 31, 35 = bright-field; Figs. 32–34, 36 = Nomarski. 31)
Typical stomatal apparatus on the upper leaf surface. Numbers 1 and 2 indicate
terminal subsidiary cells and 3 thru 6 represent pairs of flanking subsidiary cells. G =
guard cells. Scale bar = 50 m. 32) Stomatal apparatus (on the abaxial surface of a
small sepal) missing both terminal subsidiary cells (at X) and one lateral subsidiary
cell (at Y). The partition at Z is just a fold in the tissue and not a cell wall. Scale bar
= 25 m. 33) A doublet stomatal apparatus on the abaxial surface of a small sepal.
The Xs indicate three lateral subsidiary cells on one flank of the doublet. Scale bar =
25 m. 34) The adaxial surface of a small sepal with a guard cell pair, but no
subsidiary cells. The pavement cells flanking the guard cells are labeled with Ps.
Scale bar = 25 m. 35) A triplet stomatal apparatus on the abaxial side of a large
sepal. Scale bar = 25 m. 36) A doublet on the abaxial surface of a large sepal where
the guard cell pairs point in different directions. Scale bar = 25 m.
2007 R.R. Dute, B.E. Jackson, R.D. Adkins, and D.R. Folkerts 61
on the large petals is associated with the absence of a photosynthetic mesophyll
as well as the extreme sensitivity of these organs to water loss.
Stebbins and Jain (1960) show for foliar stomata of C. communis that the
first set of lateral subsidiaries form by asymmetric divisions of neighboring
cells that border the guard cell mother cell. These divisions are followed by
formation of the terminal subsidiary cells, and by asymmetric divisions of
neighboring epidermal cells. Finally, the second set of lateral subsidiary
cells arises from divisions of the first-formed lateral subsidiary cells at about
the time the guard mother cell divides to form two guard cells. The end result
is a stomatal apparatus consisting of a guard cell pair surrounded by six
The basic architecture (Fig. 31) of the stomatal apparatus is consistent on
both leaf and spathe surfaces, with few exceptions (notably on the adaxial
leaf surface). In the floral organs, however, stomatal abnormalities are
frequent in number and diverse in nature. On the abaxial surfaces of large
and small sepals, there are many cases where one or both terminal subsidiary
cells are absent (Fig. 32), where a lateral subsidiary cell does not divide (Fig.
32), or where there are extra lateral subsidiaries (Fig. 33). The stomatal
apparatuses of both petal types and of the adaxial surfaces of large and small
sepals lack subsidiary cells entirely. In such instances, the regular epidermal
cells (pavement cells) bordering the lateral sides of the guard cells assume
the shape of butterfly wings (Fig. 34) or have no particular shape.
Another modification of the basic stomatal architecture is the manufacture
of doublets (Fig. 33) or even triplets (Fig. 35) where more than one pair
of guard cells are in direct contact with one another. The guard cell pairs
involved can all have their long axes parallel to the long axis of the organ
(Figs. 33, 35) or one guard cell pair can form its longitudinal axis at a slight
angle to the organ’s longitudinal axis (Fig. 36). Where subsidiary cells are
involved with doublets, all guard cell pairs can share the same lateral
subsidiary cells (Fig. 33) or have separate ones (Fig. 36). Doublets and
triplets are not uncommon on the surfaces of floral organs. For example, on
the abaxial surface of one large sepal, the guard cell pairs were grouped in
the following arrangements: 25 singlets, five doublets, and one triplet.
Modifications of the architecture of the stomatal apparatus are not
restricted to C. erecta in the Commelinaceae. Kaushik (1971) frequently
observed abnormalities of the stomatal apparatus in leaves of five
Commelina species, including missing or extra subsidiary cells and contiguous
stomata. Stebbins and Jain (1960) found complexes in the leaf
epidermis of C. communis where a terminal subsidiary cell was absent.
Drawert (1941) found many aberrant examples in leaf sheaths of Tradescantia
virginiana L. (as T. virginica (sic), Virginia spiderwort). Normally,
the stomatal apparatuses in this species possess four subsidiary cells, but
examples with only two lateral or two lateral plus one terminal were
observed. Often the number of subsidiary cells was determined by the
number of pavement epidermal cells adjacent to the guard cell initial
62 Southeastern Naturalist Vol. 6, No. 1
Figures 37–40. 37) Raphide canal
(C) found in a large sepal
using Nomarski optics. N =
nucleus, and R = raphides. Scale
bar = 25 m. 38) Location of
crystals (light spots) in a leaf
cross section as visualized using
polarizing microscopy. Note the
occurrence of large crystals in
the lower epidermis (AB). The
sclerified walls of the bundle
sheath cells are evident (B).
Scale bar = 100 m. 39) A tannin
idioblast in a large petal.
Scale bar = 25 m. 40) Tannin
idioblasts in a small petal. Scale
bar = 100 m.
2007 R.R. Dute, B.E. Jackson, R.D. Adkins, and D.R. Folkerts 63
(guard cell mother cell). Other aberrations are mentioned as well. Both
Drawert (1941) and Stebbins and Jain (1960) comment on the migration of
nuclei of the adjacent epidermal cells toward the common wall with the
guard cell initial. Drawert speaks of this process in reference to the diffusion
of division hormones emanating from the guard cell initial. Stebbins
and Jain show nuclear migration to be a step in the polarization of the
cytoplasm, ultimately leading to the asymmetric division of an epidermal
cell into a small subsidiary cell and a larger (pavement) epidermal cell.
In a recent review article on stomatal development in Arabidopsis
thaliana (L.) Heynhold (mouse-ear cress, Brassicaceae), Nadeau and Sack
(2003) stress the importance of “intercellular signaling rather than mitosis
allocated factors” in controlling division orientation in guard mother cells.
They also discuss a mutant (too many mouths [tmm]) in which many guard
cell pairs develop in contact with one another. Unfortunately, since
Arabidopsis produces no subsidiary cells and is a dicotyledon, we do not
know how applicable it would be to the study of C. erecta. Nevertheless,
some form of communication involving signals as well as membrane receptors
would be expected in C. erecta.
Why the architecture of the stomatal apparatus in C. erecta exists in such
diversity in the laminar floral organs is unknown. Neither is the functional
status of the guard cells known, although examples of open stomata have
been observed in living specimens of small petals. Perhaps the transient
nature of these organs releases them from the normal constraints on stomatal
structure and function. Developmental and molecular studies of these structures
can shed further light on these questions.
Other anatomical features
Clusters of calcium oxalate crystals are known to occur in elongate
cells arranged in series in leaves and stems of the Commelinaceae
(Brückner 1926, Tomlinson 1969). Such raphide-canals occur not only in
the vegetative leaves of C. erecta, but also in all organs examined in this
study (Table 2, Fig. 37).
Both Brückner (1926) and Tomlinson (1969) refer to the presence of
crystals elsewhere in leaf cross sections. Brückner noted in C. communis
massive crystals of varying shape in the lower epidermis of leaves.
Table 2. The distribution of some of the common anatomical features in the laminar organs of C.
erecta. The pigment of the large petal is blue and is confined to the vacuoles of the epidermal
cells, whereas that of the other organs is chlorophyll present in the chloroplasts of the mesophyll.
Y = present, S = slight pigment, and N = not present.
Pigment Stomata Raphides Other crystals Tannin tubes
Large petal Y Y Y Y
Small petal N Y Y Y
Large sepal S Y Y Y Y
Small sepal S Y Y Y Y
Spathe Y Y Y Y Y
Leaf Y Y Y Y Y
64 Southeastern Naturalist Vol. 6, No. 1
Tomlinson noted that “large rhombohedral crystals are otherwise common,
especially in the hypodermis and epidermis of the lamina.” This information
is true also for C. erecta leaves, where polarizing microscopy indicates
crystal size and distribution (Fig. 38). The relatively large size of crystals in
the lower epidermis is quite obvious. However, crystals are absent from the
guard cells and inner lateral subsidiary cells. The large crystals tend to be
elongate, sometimes lozenge-shaped. X-ray analysis of ashed material indicates
all crystals to consist of calcium salts. Very small crystals (crystal sand
and small prisms) can be located in spathes and sepals with some difficulty.
Identification is made easier by using polarizing microscopy on ashed
spathes, where the mineral content is concentrated. No crystals (other than
raphides) have been observed with certainty in the petals, but none of this
material was ashed.
Tomlinson (1969) mentions the presence of tannin in palisade tissue of
leaves in the Commelinaceae. Indeed, tannin idioblasts are common in the
mesophyll of all organs investigated in the present study. When stained
with a mixture of glacial acetic acid, formalin, and FeSO4, these cells
developed a blue precipitate indicative of tannins (Ruzin 1999). However,
these same cells also take up other stains such as safranin, and thus might
contain a mixture of ergastic materials. The shape of the idioblasts varies
from that of a mesophyll cell (typical of the large petal; Fig. 39), to
elongate tubes (e.g., the small petal in Fig. 40) to even small spheres (as
found in vegetative leaves). Some organs can contain tannin idioblasts of
more than one shape.
In conclusion, although similarities exist, the floral organs are of a
simpler structure than the leaves and, to a lesser extent, the spathes. Lax
developmental constraints or abbreviated ontogenetic sequences in petals
and sepals appear responsible for the numerous stomatal aberrations as well
as for vein prolongations (in the large petals) and absence of sclerified
sheath cells. These features in turn are correlated with the transient nature of
the reproductive organs. It would be interesting to search for similar aberrations
in other genera of the Commelinaceae as well as to investigate the
ontogeny of said structures during petal and sepal development.
The authors wish to thank Dr. Christine Sundermann for the use of her Nomarski
microscope. We also wish to thank the reviewers and Dr. Robert Faden, the Guest
Editor, for improving the quality of this manuscript.
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