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Toxicon 54 (2009) 197–207
Contents lists available at ScienceDirect
Toxicon
journal homepage: www.elsevier.com/locate/toxicon
Parotoid macroglands in toad (Rhinella jimi): Their structure and
functioning in passive defence
Carlos Jared a, *, Marta M. Antoniazzi a, Amarildo E.C. Jordão b, José Roberto M.C. Silva b,
Hartmut Greven c, Miguel T. Rodrigues d
a
Laboratório de Biologia Celular, Instituto Butantan, Av. Vital Brasil 1500, 05503-900, São Paulo, Brazil
Departamento de Histologia e Embriologia, Instituto de Ciências Biomédicas, Universidade de São Paulo, São Paulo, Brazil
c
Institut für Zoomorphologie und Zellbiologie der Heinrich-Heine-Universität, Düsseldorf, Germany
d
Departamento de Zoologia, Instituto de Biociências, Universidade de São Paulo, São Paulo, Brazil
b
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 7 January 2009
Received in revised form 27 March 2009
Accepted 30 March 2009
Available online 15 April 2009
When toads (Rhinella) are threatened they inflate their lungs and tilt the body towards the
predator, exposing their parotoid macroglands. Venom discharge, however, needs
a mechanical pressure onto the parotoids exerted by the bite of the predator. The structure
of Rhinella jimi parotoids was described before and after manual compression onto
the macroglands mimicking a predator attack. Parotoids are formed by honeycomb-like
collagenous alveoli. Each alveolus contains a syncytial gland enveloped by a myoepithelium and is provided with a duct surrounded by differentiated glands. The epithelium
lining the duct is very thick and practically obstructs the ductal lumen, leaving only
a narrow slit in the centre. After mechanical compression the venom is expelled as a thin
jet and the venom glands are entirely emptied. The force applied by a bite of a potential
predator may increase alveolar pressure, forcing the venom to be expelled as a thin jet
through the narrow ductal slit. We suggest that the mechanism for venom discharge
within all bufonids is possibly similar to that described herein for Rhinella jimi and that
parotoids should be considered as cutaneous organs separate from the rest of the skin
specially evolved for an efficient passive defence.
Ó 2009 Elsevier Ltd. All rights reserved.
Keywords:
Amphibia
Bufonidae
Rhinella jimi
Parotoid
Macrogland
Skin glands
Venom release
Passive defence
1. Introduction
Amphibian skin glands are a synapomorphy of the
group and are usually present in great numbers on the
whole body of all species. Among amphibian skin glands
the most conspicuous are undoubtedly the parotoids.
Parotoids (for etymology and the recommendation to use
parotoid instead of ‘‘paratoid’’ or ‘‘parotid’’ see Cannon and
Palkuti, 1976; Tyler et al., 2001) are multiglandular structures, which can be found as paired protuberances, postorbital in position, in a variety of Anura (Wilber and Carroll,
* Corresponding author. Tel.:/fax: þ55 11 37267222x2234.
E-mail addresses: [email protected], [email protected] (C. Jared).
0041-0101/$ – see front matter Ó 2009 Elsevier Ltd. All rights reserved.
doi:10.1016/j.toxicon.2009.03.029
1940; Lutz, 1971; Toledo et al., 1992) and Urodela (Phisalix,
1922; Luther, 1971; Brodie and Smatresk, 1990; Toledo and
Jared, 1995). Because parotoids consist of an aggregation of
numerous secretory units, they have been named macroglands to differentiate them from the common mucous
and granular glands of the remaining skin (see Toledo and
Jared, 1995).
There seems to be no doubt that parotoids as well as the
granular glands distributed over the body of a toad, from
which the parotoids are derived (Toledo et al., 1993; Toledo
and Jared, 1995), are involved in chemical defence
including defence against microorganisms (Tempone et al.,
2008). Behavioural studies in both Anura and Urodela, have
demonstrated that many species show characteristic
defence behaviour when threatened. Urodela, for example,
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position their body, tilting it toward the source of contact
(for review see Brodie, 1983) and passively release unpalatable or toxic cutaneous glandular contents onto the body
surface, rather than actively spraying them at predators
(e.g., Brodie, 1983; Brodie and Smatresk, 1990). Anura,
especially toads, inflate their lungs and also assume special
postures to present the parotoids to the source of danger,
but seem to spray their venom only in response to physical
pressure (e.g., Hinsche, 1928; Toledo and Jared, 1995).
Secretions of the granular cutaneous glands (and parotoids) may be very toxic to animals (Lutz, 1966; Toledo and
Jared, 1989a, 1995; Barthalmus, 1989; Clark, 1997). In
bufonids they contain steroids such as bufogenines and
bufotoxins that, when in contact with the buccal mucosa
of many vertebrates, especially of snakes, have cardioacceloratory properties increasing the strength of the
heart beat and decreasing heart rate (Habermehl, 1981).
The venom can also exert a marked effect as a local
anaesthetic (Habermehl, 1981). In the case of a bite, after
venom ingestion, the potential predator can show intense
salivation and excitation, paralysis, trembling and convulsions, often leading to death (Vital Brazil and Vellard, 1925,
1926; Garrett and Boyer, 1993; Pineau and Romanoff, 1995;
Sakate and Lucas de Oliveira, 2000; Sonne et al., 2008).
Anuran granular or venom glands are syncytial in nature
(Fox, 1986; Delfino, 1991; Terreni et al., 2003). They are
irregularly distributed over the whole body surface, but are
found also in aggregated forms in certain parts of the body
such as the parotoid and the paracnemic macroglands of
bufonids (Hostetler and Cannon, 1974; Toledo and Villa,
1987; Toledo et al., 1992) or the lumbar macroglands of
some leptodactylids (Toledo and Jared, 1989b, 1995; Toledo
et al., 1996; Lenzi-Mattos et al., 2005). Studies on the
structure and function of such aggregations are limited.
Specifically in toads, there are some articles that describe
their secretory syncytia, the secretion granules and the
myoepithelial layer (Hostetler and Cannon, 1974; Cannon
and Hostetler, 1976; Toledo et al., 1992; Barthalmus, 1994;
Almeida et al., 2007). In addition, a special vascularization
net irrigating the toad parotoids has been described in
detail (Hutchinson and Savitzky, 2004). On the other hand,
the biochemistry and biological effects of toad skin,
including the parotoids, are relatively well studied (Pasquarelli et al., 1987; Rossi et al., 1997; Maciel et al., 2003;
Shimada et al., 2006; Tempone et al., 2008).
In the present article we describe the parotoids of the
toad Rhinella jimi focusing on the structure of their tissues
before and after mechanical pressure. Our results show that
the ducts of the individual granular glands are lined by
thick differentiated epithelia constituting a plug that
hinders the passage of secretion. Further we suggest that
the special arrangement of tissues around the glands and
inside the ducts, together with pressure applied on the
macroglands, e.g., by the experimenter or a predator, lead
to venom discharge in the form of strong jets, characterizing a peculiar type of passive defence in toads.
2. Material and methods
Rhinella jimi (Fig. 1) is a Brazilian toad restricted to the
semiarid Caatinga of northeastern Brazil and previously
confused with Rhinella schneideri (Stevaux, 2002). It was
originally named as Bufo jimi (Stevaux, 2002) and renamed
as Rhinella jimi (Chaparro et al., 2007). Six female adult
specimens were collected at Fazenda São Miguel, district of
Angicos, State of Rio Grande do Norte (5 390 4300 S, 36 360
1800 W), Brazil and brought to the vivarium of the Cell
Biology Laboratory of Instituto Butantan, São Paulo.
To mimic the bite of a possible predator (e.g., some
snakes, mammals and birds of prey), a large and constant
pressure was applied to the right parotoids of the six live
individuals by squeezing them laterally between the thumb
and the forefinger. This process led to the expulsion of
venom (see Fig. 1(5)). After that, all the animals were
sacrificed with an overdose of Thionenbutal and
compressed and non-compressed parotoids were removed
for histological study.
2.1. Anatomy
For this study the parotoids were removed from existing
specimens from the herpetological collection of the Instituto Butantan, fixed in 10% formalin and kept in 70%
ethanol. The parotoids were divided on a frontal plane into
two pieces. The alveolar content was carefully removed
using a toothpick to expose the internal macrogland
framework. Pore disposition was examined by direct
observation of the parotoid surface or by light microscopy
using whole mount preparation of the parotoid skin surface
flattened on a glass slide.
2.2. Histology
The parotoids removed from the six collected individuals were divided transversely to the gland larger axis into
four pieces and fixed in 4% formaldehyde, 1% glutaraldehyde, buffered in 0.1 M phosphate buffer, pH 7.2 (McDowell
and Trump, 1976) for 4 days. The pieces were transversely
oriented and embedded in paraffin, sectioned 4 mm thick,
stained with Haematoxylin–Eosin and Mallory trichrome
for general observations, and submitted to Bromophenol
Blue, and PAS combined with Alcian Blue pH 2.5 (Bancroft
and Steven, 1990), for detection of proteins in general and
mucosubstances, respectively. Picrosirius staining followed
by polarized microscopy was used in a few sections for the
analysis of collagen fibres (Junqueira et al., 1979). Micrographs were taken with an Olympus BX60 light microscope
connected to Olympus PM-C35DX photo equipment or
with an Olympus BX51 light microscope equipped with
a digital camera and with the software Image-Pro Express
(Media Cybernetics).
2.3. Transmission electron microscopy
Small cubes (8 mm3) of parotoid superficial tissue containing single parotoid pores were fixed in a mixture of
2.5% glutaraldehyde and 2% paraformaldehyde in 0.1 M
cacodylate buffer, pH 7.2, post-fixed in 1% osmium
tetroxide, dehydrated and embedded in Spurr resin.
Samples were oriented to obtain transverse ultrathin
sections of the pores. Sections were cut in a Sorvall MT6000
ultramicrotome, stained with uranyl acetate and lead
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C. Jared et al. / Toxicon 54 (2009) 197–207
citrate (Reynolds, 1963) and examined in a LEO 906E
transmission electron microscope.
3. Results
3.1. Anatomy of the parotoid glands and venom discharge
induced by external pressure
The parotoids are conspicuous glandular structures,
which are easily dissected and removed from the body
when the dorsal skin around them is cut. Externally the
parotoid skin seems similar to the rest of the dorsal skin,
except for the presence of a large number of well defined
pores (Fig. 1). When the macroglands are horizontally cut
into two pieces and the pasty secretion is removed,
a honeycomb-like structure is revealed (Fig. 1(2,3)),
composed of many alveoli. Transverse histological section
through the parotoid shows that each alveolus contains
a very large bottle-shaped gland, full of secretion and
closed on the top by a thick ductal epithelium apparently
forming a plug (Fig. 1(4)). Serial sections, however, reveal
that there is a very narrow ductal lumen forming a slit (Figs.
1(4) and 3(14)) through which the secretion can pass when
the gland is squeezed. Since the parotoid forms a convex
structure, the bottle-shaped glands are larger and more
elongated in the central region and gradually decrease in
size toward its borders. The pores on the surface of the
parotoids measure from 0.5 to 1 mm, depending on the
individual, and appear as circular depressions, almost
entirely obstructed internally by solid tissue, except for the
small slit in the centre. Each pore is externally surrounded
by smaller pores forming a special pore pattern on the
parotoid surface (Figs. 1(6) and Fig. 3(16)).
A constant manual pressure applied to the parotoids
forces the venom to be discharged only from the pores
corresponding to the glands which were effectively
squeezed between the fingers. The venom is expelled in the
form of thin jets (Fig. 1(5)). It is sticky and has a colour
varying from white to yellowish.
3.2. Histology and histochemistry
The dorsal skin has an irregular surface and is composed
of the epidermis containing around six cellular layers, and
the dermis with the stratum spongiosum, where mucous
and granular glands are observed, and the stratum compactum, mainly constituted by thick collagen fibres
(Fig. 2(7)). The mucous glands are arranged just below the
epidermis and have a typical acinar form, with large lumina
and thin ducts opening in the skin surface. The secretory
epithelium reacts differentially to PAS and Alcian Blue
corresponding to the presence of two types of secretory
cells (Fig. 2(13)) but it does not react to Bromophenol Blue
(Fig. 2(10)). The granular glands are larger than the mucous
glands and consist of a syncytium with no lumen (Fig.
2(7,9)) and filled with secretion positive to Bromophenol
Blue (Fig. 2(10)) and negative to PAS and Alcian Blue
(Fig. 2(11)). Both types of gland are encased by myoepithelia and are connected to the skin surface through
epithelial ducts. A continuous basophilic calcified dermal
layer runs parallel to the epidermis, between the two
199
dermal strata, underlining the glands (Fig. 2(7)). The
unpigmented ventral skin shows a flat surface, a much
thinner stratum compactum and a small number of glands,
mainly mucous glands (Fig. 2(8)).
The histological analysis of the parotoids show that they
are thickened portions of the dorsal skin in which, besides
the presence of mucous and regular granular glands, very
large bottle-shaped syncytial granular glands are accumulated in the stratum compactum. Each gland is connected to
the exterior through a narrow aperture (slit) in the centre
of the ductal lining epithelia. In the parotoid the differentiation of the two dermal strata (spongiosum and compactum) is not as clear as in the rest of the dorsal skin.
However, similarly to the dorsal skin, the mucous and
regular granular glands are located just under the
epidermis (Fig. 2(9)). Moreover, epidermis and dermis
are traversed by the large ducts connecting the surface to
the large syncytial granular glands. These ducts are surrounded by differentiated glands exclusively present in the
parotoid (Fig. 2(9–11)). Differently from the dorsal skin, the
calcified dermal layer in the parotoid is much thicker and
spread among the connective tissue fibres, forming an
irregular band about 150 mm thick below the epidermis
(but above the bottle-shaped granular glands), in which the
mucous glands are immersed (Fig. 2(9)).
The bottle-shaped syncytial granular glands of the
parotoid are filled with a network of secretion products
(Figs. 2(9) and 4(22,23)). This secretion is very reactive to
Alcian Blue and negative to Bromophenol Blue, differently
from the granular glands of the dorsal skin (compare Fig.
2(10,11)). The syncytial nuclei are arranged in a single
peripheral row. Each gland is enveloped by a monolayer of
thin myoepithelial cells, with flat and elongated nuclei
(Fig. 4(23)).
The large ducts of the bottle-shaped glands are surrounded by a number of glands clearly different from
mucous and granular glands both in size and histochemistry (compare Fig. 2(9,10), and Fig. 2(12,13)). In histological
transverse sections, these glands appear circularly arranged
around the main duct (Fig. 3(15)), forming a rosette-like
arrangement. In whole mount preparations the small pores
of these glands can be seen surrounding the main pore of
the large syncytial granular gland (Fig. 3(16)). Although
similar to the common mucous glands in terms of their
typical acinar structure and their positive reaction to PAS
and Alcian Blue, these differentiated glands are much
larger, with diameters around 370 mm, whereas the
common mucous glands have diameters around 100 mm
(Fig. 2(9,10)). Also, the acini of the differentiated glands are
formed by high prismatic cells, full of a homogeneous
secretion very reactive to Bromophenol Blue (Fig. 2(10))
and to PAS (Fig. 2(11,12)), and sparse cells, very positive to
Alcian Blue, pH 2.5, located mainly in the apical secretory
epithelium (Fig. 2(12)). Occasionally the lumen contains
secretion.
Longitudinal serial sections reveal that bundles of
collagen fibres surround each duct, forming a distinct
arrangement from the rest of the stratum compactum
(Fig. 3(14)). At about 500 mm from the skin surface, the duct
lining epithelium is thickened and practically obstructs the
ductal lumen, leaving only a narrow slit (measuring about
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201
Fig. 2. (7) Rhinella jimi dorsal skin. The dermis shows many mucous (*) and granular (g) glands. The calcified dermal layer (ca) is present dividing the stratum
spongiosum (ss) from the stratum compactum (sc). E, epidermis; v, blood vessels. Paraffin, HE. (8) Rhinella jimi ventral skin. When compared to the dorsal skin, it
is much thinner and with a small number of glands, which are mainly mucous (*). The dermis is well vascularized, with many blood vessels (v) under the
epidermis (E). sc, stratum compactum; ss, stratum spongiosum. Paraffin, HE. (9) Rhinella jimi parotoid macrogland. Apical portion of the macrogland where four
glandular types are distinguish: common mucous gland (*), common granular gland (g), differentiated gland (dg) and parotoid granular gland (G). Note that the
differentiated cells are much larger than the mucous cells. A thick calcified layer (ca) is seen in the upper dermis (D) underlining the epidermis (E). A small part of
the ductal epithelium (d) is observed in the parotoid granular gland. Paraffin, HE. (10) Rhinella jimi parotoid macrogland. Equivalent section to Fig. 2(9) stained
with Bromophenol Blue. Mucous glands (*) and parotoid granular gland (G) are negative, the common granular gland (g) is positive and the differentiated glands
(dg) are highly positive to the method. E, epidermis; D, dermis, d, ductal epithelium. Paraffin. (11–13) Rhinella jimi parotoid macrogland. Comparison of the four
gland types present in the macrogland by PAS-AB (pH 2.5). The parotoid granular gland (G) is positive to Alcian Blue, while the common granular glands (g) are
negative. The cells of the mucous glands (*) are mainly positive to Alcian Blue (arrows, Fig. 2(13)) while in the differentiated glands (dg) most cells are positive to
PAS, with a few cells positive to Alcian Blue (arrows, Fig. 2(12)). E, epidermis; D, dermis; d, duct; s, secretion. Paraffin.
Fig. 1. (1) Female Rhinella jimi exhibiting the right parotoid macrogland localized at the postorbital region. The arrows point to the glandular pores. (2) A parotoid
sectioned according to a frontal plane, from which the venom was withdrawn. Notice the alveolar, honeycomb-like internal structure. (3) Higher magnification of
the alveoli showing the walls (arrows) and floors (asterisks). (4) Longitudinal section of two parotoid bottle-shaped glands (G). The arrows point to the pores
obstructed by a thick epithelium. D, dermis. Paraffin, HE. (5) Venom jets squirting from the pores, after parotoid manual compression. (6) View of the pores on the
parotoid surface. Note the small slit (large arrows) in the duct centre. Each pore is surrounded by smaller pores (small arrows).
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Fig. 3. (14) Rhinella jimi parotoid macrogland. Histological longitudinal section of a duct obstructed by the ductal epithelium (d) and surrounded by the
differentiated glands (asterisks) and wrapped by distinct bundles of collagen fibres (arrowheads). Note the thin canal (small arrow) running in the central portion
of the duct. The larger arrow point to the external side of the pore. ca, calcified dermal layer; D, dermis; E, epidermis. Paraffin, HE. (15) Rhinella jimi parotoid
macrogland. Histological transverse section of the duct surrounded by specialized mucous glands (asterisks), forming a rosette. E, epidermis. Paraffin, HE. (16)
Rhinella jimi parotoid macrogland. Whole mount preparation of the parotoid epidermis in the region of the pore. Most of the duct volume is obstructed by the
plug, leaving only a central pore (central arrow). Circling disposed arrows point to the differentiated glands around the central pore. (17) Rhinella jimi parotoid
macrogland. Larger magnification of the duct epithelial cells, showing difference in shape between the peripheral and central cells. Paraffin, HE. (18) Rhinella jimi
parotoid macrogland. The duct epithelial cells are positive to the PAS method. (19 and 20) Rhinella jimi parotoid macrogland. Transmission electron micrographs
of the epidermal cells composing the thick duct lining, which are voluminous at the periphery (Fig. 3(19)), becoming flatter towards the lumen, with slender
cytoplasm processes (asterisks) (Fig. 3(20)). All cells show electron transparent cytoplasm inclusions (i). Nu, nuclei.
40 mm) in the centre. Longitudinal sections through the
epithelium reveal that the cells arranged towards the
ductal slit are flatter than the basal cells (Fig. 3(17)). In
addition, the whole ductal epithelium is quite reactive to
PAS (Fig. 3(18)). The transmission electron microscopy of
these cells shows several cytoplasm inclusions containing
material of medium electron density (Fig. 3(19,20)). The
connections among the central flatter cells consist of very
loose interdigitations (Fig. 3(20)).
The connective tissue surrounding the large syncytial
glands is divided into two strata, a loose and richly
vascularized stratum, closely adjacent to the gland
myoepithelia, and a dense stratum (Fig. 4(21,22)). The
dense stratum is continuous with the superficial stratum
compactum, and envelopes each gland, structuring the
honeycomb-like framework of the parotoid (see Fig. 1(2,3)).
The parotoid base is flat and constituted of the same dense
connective tissue.
After parotoid compression, conspicuous structural
changes are observed in the alveoli affected by squeezing:
they are empty and almost completely free of secretion. The
periphery, where the syncytial nuclei are located, collapses
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203
Fig. 4. (21) Rhinella jimi parotoid macrogland. Two granular parotoid glands, one non-compressed filled with secretory product (G) and another compressed,
with the syncytium (sy) completely collapsed in the centre, appearing in orange. Note the difference in volume between the loose connective tissue (lt) in both
glands. The dense connective tissue (dt) is unchanged. Blood vessels (arrows). Paraffin, Mallory trichrome staining. (22) Rhinella jimi parotoid macrogland.
Connective tissue between two parotoid granular glands (G). The loose (lt) and dense (dt) connective tissues are clearly distinguish. Blood vessels (arrows) are
abundant in the loose connective tissue. Paraffin, HE. (23) Rhinella jimi parotoid macrogland. Part of the syncytium (sy) of a non-compressed alveolus, sheathed
by the myoepithelial layer (my) and full of granular secretory product (G) . The arrow points to a blood vessel. Paraffin, HE. (24) Rhinella jimi parotoid macrogland.
A compressed parotoid gland, equivalent to the one observed in Fig. 4(21), with the syncytium (sy) completely collapsed in the center. The arrows point to blood
vessels. lt, loose connective tissue, dt, dense connective tissue. Paraffin, HE. (25) Higher magnification of Fig. 4(24), where rests of the secretory product are seen
inside the syncytium, which is enveloped by the myoepithelial layer (my). lt, loose connective tissue, dt, dense connective tissue. Paraffin, HE.
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Fig. 5. (26 and 27) Rhinella jimi parotoid macrogland. Connective tissue between two non-compressed parotoid granular glands (G) in non-polarized (Fig. 5(26))
and polarized (Fig. 5(27)) microscopy. The loose connective tissue (arrows) is compressed towards the dense connective tissue (dt). Paraffin, Picrosirius staining.
(28 and 29) Rhinella jimi parotoid macrogland. Connective tissue between two compressed parotoid granular glands (G) in non-polarized (Fig. 5(28)) and
polarized (Fig. 5(29)) microscopy. The loose connective tissue (lt) is expanded towards the dense connective tissue (dt). Paraffin, Picrosirius staining.
together with the surrounding myoepithelium: both
(syncytium and myoepithelium) are now seen as a wrinkled structure in the centre of the alveolus (compare full
and empty alveoli in Fig. 4(21)). The loose vascularized
connective tissue, adjacent to the myoepithelium, appears
expanded occupying a larger space between the collapsed
gland and the dense connective tissue, which is unchanged
(Fig. 4(21,24,25)). The comparison of a gland full of secretion with a squeezed gland in Picrosirius stained sections
viewed under polarized microscopy shows the difference
between non-expanded and expanded loose connective
tissue surrounding each of the wrinkled syncytia (compare
Fig. 5(26,27) with Fig. 5(28,29)).
Serial histological sections of the compressed glands did
not show any significant modification of the ducts after
secretion release except for a discrete widening of the
ductal slit and the presence of secretion in the ductal
lumen.
4. Discussion
Rhinella jimi has a wide distribution in Brazilian Caatinga. Rhinella schneideri, which is a close relative of R. jimi
(probably consisting of a complex of species), lives in the
central Brazilian Cerrados (dry open woodlands). Both
species, despite the presence of the paracnemic macroglands (characteristic of the Rhinella schneideri group) in
their hind limbs, comprise large individuals, up to 300 mm
in adult females. One of the diagnostic features which
enables the recognition of Rhinella jimi as a distinctive
species in the Rhinella schneideri group is the presence of
macroglands also in the forelimbs (Stevaux, 2002). This
characteristic has already been described by (Toledo and
Jared, 1995, Fig. 2(13)) who referred to such glandular
accumulations as radioulnar macroglands, in order to
distinguish them from the paracnemic macroglands,
previously studied by Toledo and Villa (1987). Different
from other anurans inhabiting open and dry regions,
Rhinella jimi and Rhinella schneideri remain completely
exposed to the environment for long periods, without
running the apparent risk of desiccation, a fact that is
probably associated with the presence of a calcified dermal
layer (Elkan, 1976; Toledo and Jared, 1993b), covering the
entire body, including the parotoids. In both species parotoid macroglands are very prominent, a fact commented on
by Lutz (1925) in the original description of Rhinella
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schneideri. At that time he qualified those macroglands as
‘‘enormous’’ although referring to them as ‘‘parotids’’.
Besides their large size, they occupy a prominent position
in relation to the body and, considering the combination of
defensive behaviours usually found in these toads, it seems
evident that they depend on the parotoid macroglands in
their defence.
Rhinella jimi when threatened usually exhibits a stereotyped defensive behaviour, characterized by the inflating
of the lungs and the assumption of a stiffened and voluminous form. This behavioural posture is often followed by
the head-butting behaviour, in which the animal tilts the
body towards the threatening agent, exposing one of the
parotoids (Toledo and Jared, 1995, referred to as Bufo paracnemis). Head-butting seems to be an important part of
the toad defensive strategy. The act of exposing the parotoid certainly increases the chances of the animal having its
venom release system triggered when the predator bites it.
Our observations strongly suggest that jets of venom are
only discharged from the parotoids after a considerable
mechanical pressure. In fact, spontaneous discharge of jets
from the parotoids has not been reported in Anura and has
rarely been in Urodela (Brodie, 1983; Brodie and Smatresk,
1990). In toads, however, pressure seems to be necessarily
exerted by an external force which in nature must correspond to the attack or biting of a predator. Such external
pressure was mimicked herein by manual squeezing.
Toads seem to have a graded sequence of defensive
behaviours: first inflating the lung and lifting the body,
which is followed by spraying the venom, if the predator
should attack and/or bite his victim. Even if the toad is
killed, the defensive strategy may be efficient at the level
of species defence, since the predator may be able to
associate the stereotyped defence behaviour with the
distasteful or harmful venom when grasping another toad.
However, this system seems not to have been explored in
detail in toads and depends on the predator’s capabilities,
as many animals are known to have found ways to
avoid poison. Experiments in captivity in which frogs
(Eupemphix nattereri, formerly Physalaemus nattereri)
were offered to the procyonid Nasua nasua (coati) showed
that this mammal rejected the first frog but could ingest
the second after vigorously rolling and rubbing it against
the soil (Sazima and Caramaschi, 1986). We have many
times observed in the field half-eaten dead toads where
the head and most of the skin were discarded by the
predator. In urodelans, chickens learn avoidance of the
efts of the salamander Notophthalmus viridescens (Brandon et al., 1979) and a hedgehog avoided Pleurodeles waltl
after biting a specimen several times on the tail (Nowak
and Brodie, 1978).
Although the external pressure is probably the most
important factor for parotoid venom release, lung inflation
must be significant in the defence of Rhinella jimi since,
besides making the animal appear larger, the lung pressure
against the body is possibly carried over to the parotoid
floor and transferred to the parotoid bottle-shaped glands.
The state of turgidity thus produced and the probable aid of
the individual glandular myoepithelial layer make it such
that, by external pressure, the secretion is expelled through
the duct slit causing venom jets to squirt out.
205
The dense connective tissue surrounding each bottleshaped gland forms a resistant framework responsible for
the shape maintenance of the parotoid, which appears
macroscopically unaltered even after venom release. On
the other hand, the loose connective tissue seems to form
part of the secretion release system. Also, the loose tissue
must have a significant role in syncytium and myoepithelial
maintenance, since it is highly vascularized and must be
involved in gland nutrition and transport of precursor
molecules for venom synthesis (Hostetler and Cannon,
1974; Cannon and Hostetler, 1976; Erspamer, 1994;
Hutchinson and Savitzky, 2004). Immediately after the
explosive emptying, the syncytium is totally collapsed and
the space between the dense connective tissue and the
myoepithelium is filled by the obviously expanded loose
connective tissue. Simultaneously with the syncytium
secretion refilling, the loose connective tissue is gradually
pushed towards the dense connective tissue walls.
The ultrastructure of compressed and non-compressed
parotoids in Rhinella icterica (formerly Bufo ictericus),
suggests that the alveolus, when full of venom, is under
a constant internal pressure, since organelles are crowded
in a small cytoplasm volume. After discharge, however, the
organelles are easily recognized and distributed in a larger
volume of cytoplasm (Toledo et al., 1992). Our histological
results strongly indicate that similar features must occur in
the parotoid alveoli of Rhinella jimi. When the alveoli are
full, it is most probable that the normal pressure of the
venom on the loose connective tissue maintains a condition of relative internal turgidity. The moment Rhinella jimi
feels threatened its defence mechanism of inflating its
lungs may contribute to the increase of the internal pressure of each alveolus. Since the collagen surrounding the
alveolus and the duct appears very resistant, when an
external force is applied to the parotoid, e.g., by a bite, the
individual alveolar pressure is increased to a threshold
level, forcing the venom to eject through the narrow slit by
compressing the ductal epithelial lining cells. The ductal
thick epithelium could be compared in this context with
a small nozzle. Only the alveoli directly compressed cause
the venom discharge, liberating simultaneous multidirected ‘‘shots’’ inside the predator’s mouth. This diffuse
mechanism of venom elimination in a spray form can
reinforce the envenomation through breathing mucosa.
The parotoid alveoli which were not directly compressed
remain practically intact and ready to be triggered in the
event of a new attack.
This putative mechanism does not contradict the fact
that granular glands of the body skin and also the parotoid
glands to some extent may discharge their secretion by
contraction of the myoepithelium. Both mechanisms may
complement each other at the moment the animal is
threatened. Contraction of the myoepithelium is triggered
by an adrenergic mechanism evoked by orthosympathetic
stimulation (Holmes and Balls, 1978; Delfino et al, 1982;
Nosi et al., 2002). In addition, the myoepithelial cells in
macroglands may have the additional function of homogenizing the voluminous gland secretion by constant gentle
contractions.
The thick epithelium present in the parotoid ducts,
although previously shown elsewhere (Lobo, 2005; Lobo
Author's personal copy
206
C. Jared et al. / Toxicon 54 (2009) 197–207
et al., 2005; Jared et al., 2007) was herein described in
detail, with suppositions of its role in venom release. In the
same way, although the large differentiated glands in the
parotoids and their special arrangement around each pore
of the bottle-shaped glands have already been shown
(Lobo, 2005; Lobo et al., 2005; Jared et al., 2007), they were
herein described in more detail. The function of these
differentiated glands is entirely unknown as yet, but based
on their specific location, one should expect their secretions to have a role in venom release or in making part of
the venom itself. In fact, Almeida et al. (2007) reported that
in Rhinella icterica, the parotoid venom is composed of
a mixture of products released by different gland types.
Almeida et al. (2007), however, did not make a distinction
between the common mucous cells and the herein named
differentiated glands, which they called mixed glands due
to the protein and mucous nature of their secretory
product. We have shown the same type of histochemical
results in R. jimi. Also, Almeida et al. (2007) did not notice
the clear association of these mixed glands with the large
parotoid ducts which have already been described by Lobo
(2005), Lobo et al. (2005) and Jared et al. (2007).
Another morphological aspect of R. jimi skin deserving
attention is the difference between the calcified dermal
layer of the dorsal skin and of the parotoid macrogland
skin. In the parotoid this layer is much thicker and is
located more superficially, just below the epidermis and
above the bottle-shaped glands. Considering anuran
integument in general, this observation seems quite
unexpected since the calcified layer is usually localized
below the glands, even when they are large (personal
observations). This unusual more external position of the
calcified dermal layer in the parotoid probably confers to
the macrogland a more intense superficial mechanical
resistance, and may help, at the moment of a bite, to
canalize the internal venom pressure towards the ducts,
forcing the venom to be released through the ductal slits.
Association of the calcified dermal layer distribution
pattern with the mechanism of gland discharge has already
been proposed for Physalaemus nattereri (presently
Eupemphix nattereri) (Lenzi-Mattos et al., 2005), whose
posterior dorsal skin is provided with a pair of black and
circular macroglands resembling two black eyes. These
macroglands appeared to be the only region of the frog’s
integument where the calcified dermal layer is absent,
probably facilitating the venom to be expelled when the
attack of a predator is directed to the macroglandular
region (Lenzi-Mattos et al., 2005).
Besides R. jimi, we have examined the parotoid of
many different species of Rhinella (R. schneideri, R. icterica,
R. margaritifera, R. crucifer and R. granulosa). In all the
studied species the morphological and histochemical
patterns of the parotoid macroglands were very similar to
what we have described for R. jimi, including the duct
arrangement, the differentiated glands around the ducts
and the location of the calcified dermal layer. This similarity
is a strong indication that the structure of the parotoid
macrogland herein described may constitute a basal character within the genus.
Based on the fact that parotoid macroglands are
common to the whole genus Rhinella, and that the general
morphology of these structures is quite similar in many
toad species, we suggest that the mechanism for venom
release within the genus is possibly similar to that we have
described for Rhinella jimi. In addition, supported by the
structural complexity herein shown, the parotoid macroglands can no longer simply be considered as mere gland
aggregations but should be regarded as highly differentiated structures, characteristic of bufonids, forming cutaneous organs separate from the rest of the skin, which were
specially evolved for an efficient passive defence.
Acknowledgements
This work was made possible due to the help and
kindness of Mr Francisco de Assis Rodrigues, manager of
the Fazenda São Miguel (District of Angicos, Rio Grande do
Norte State). Thanks are due to CNPq (307247/2007-4 for
CJ, and 302212/82-5 for MTR), FAPESP and Fundação
Butantan. We would like to thank Prof. Reynaldo C. de
Toledo for important insights regarding the manuscript.
We also thank Simone G.S. Jared and Maria Helena Ferreira
for their efficient technical assistance. Specimens were
collected under 172/1999 and 193/2001 IBAMA permits.
Conflict of interest
The authors have no conflicts of interest.
References
Almeida, P.G., Felsemburgh, F.A., Azevedo, R.A., Brito-Gitirana, L., 2007.
Morphological re-evaluation of the parotoid glands of Bufo ictericus
(Amphibia, Anura, Bufonidae). Contrib. Zool 76, 145–152.
Bancroft, J.B., Steven, A., 1990. Theory and Practice of Histological Techniques. Churchill Livingstone, Edinburgh.
Barthalmus, G.T., 1989. Neuroleptic modulation of oral dyskinesias
induced in snakes by Xenopus skin mucus. Pharmacol. Biochem.
Behav 30, 957–959.
Barthalmus, G.T., 1994. Biological roles of amphibian skin secretions. In:
Heathole, H., Barthalmus, G.T., Heatwole, A.Y. (Eds.), Amphibian
biology – The integument. Surrey Beatty & Sons, Chipping Norton, pp.
382–410.
Brandon, R.A., Labanick, G.M., Huheey, J.E., 1979. Learned avoidance of
brown efts, Notophthalmus viridescens louisianensis (Amphibia, Urodela, Salamandridae), by chickens. J. Herpetol 13, 171–176.
Brodie Jr., E.D., 1983. Salamander antipredator postures. Copeia 1977,
523–535.
Brodie Jr., E.D., Smatresk, N.L., 1990. The antipredator arsenal of fire
salamanders: spraying of secretions from highly pressurized dorsal
skin glands. Herpetologica 46, 1–7.
Cannon, M.S., Hostetler, J.R., 1976. The anatomy of the parotoid gland
in Bufonidae with some histochemical findings. II. Bufo alvarius.
J. Morphol 148, 137–160.
Cannon, M.S., Palkuti, G.A., 1976. The ‘parotoid’ gland of Bufonidae. Toxicon 14, 149–151.
Chaparro, J.C., Pramuk, J.B., Gluesenkamp, A.G., 2007. A new species of
arboreal Rhinella (Anura: Bufonidae) from cloud forest of southeastern Peru. Herpetologica 63 (2), 203–212.
Clark, B.T., 1997. The natural history of amphibian skin secretions, their
normal functioning and potential medical applications. Biol. Rev. 72,
365–379.
Delfino, G., 1991. Ultrastructural aspects of venom secretion in anuran
cutaneous glands. In: Tu, A.T., Dekker, M. (Eds.), Reptile Venoms and
Toxins. Handbook of Natural Toxins, vol. 5. Marcel Dekker Inc., New
York, pp. 777–802.
Delfino, G., Amerini, S., Mugelli, A., 1982. In vitro studies on the ‘‘venom’’
emission from the skin of Bombina variegata pachypus (Bonaparte)
(Amphibia, Anura, Discoglossidae). Cell Biol. Int. Rep 6, 843–850.
Author's personal copy
C. Jared et al. / Toxicon 54 (2009) 197–207
Elkan, E., 1976. Ground substance: an anuran defense against desiccation.
In: Lofts, B. (Ed.), Physiology of the Amphibia, vol. 3. Academic Press,
London, pp. 101–111.
Erspamer, V., 1994. Bioactive secretions of the amphibian integument. In:
Heathole, H., Barthalmus, G.T., Heatwole, A.Y. (Eds.), Amphibian
Biology – The Integument. Surrey Beatty & Sons, Chipping Norton, pp.
176–350.
Fox, H., 1986. Dermal glands. In: Bereiter Hahn, J., Matoltsy, A.G.,
Richards, K.S. (Eds.), Biology of the Integument, vol. 2. Vertebrates.
Springer, Berlin, pp. 116–135.
Garrett, C.M., Boyer, D.M., 1993. Bufo marinus (Cane Toad). Predation.
Herpetol. Rev. 24, 148.
Habermehl, G., 1981. Venomous Animals and their Toxins. Springer Verlag, Berlin.
Hinsche, G., 1928. Kampfreaktionen bei einheimischen. Anuren. Biol. Zbl
48, 577–617.
Holmes, C., Balls, M., 1978. Invitro studies on the control of myoepithelial
cell contraction in the granular glands of Xenopus laevis skin. Gen.
Comp. Endocrinol 36, 255–263.
Hostetler, J.R., Cannon, M.S., 1974. The anatomy of the parotoid gland in
Bufonidae with some histochemical findings. I, Bufo marinus. J. Morphol 142, 225–240.
Hutchinson, D.A., Savitzky, A.H., 2004. Vasculature of the parotoid glands
of four species of toads (Bufonidae: Bufo). J. Morphol 260, 247–254.
Jared, C., Antoniazzi, M.M., Jordão, A.E.C., Silva, J.R.M.C., Vasconcellos, T.P.,
Zanotti, A.P., Rodrigues, M.T., 2007. Morphology of the parotoid
macroglands in the toad Chaunus jimi and the mechanism of venom
expulsion. Annals of the Third Congress, Brazilian Society of Herpetology, Belém, Brazil.
Junqueira, L.C.U., Bignolas, G., Brentani, R.R., 1979. Picrosirius staining plus
polarization microscopy, a specific method for collagen detection in
tissue sections. Histochem. J. 11, 447–455.
Lenzi-Mattos, R., Antoniazzi, M.M., Haddad, C.B., Tambourgi, D.V.,
Rodrigues, M.T., Jared, C., 2005. The inguinal macroglands of the frog
Physalaemus nattereri (Leptodactylidae): structure, toxic secretion
and relationship with deimatic behaviour. J. Zool 266, 385–394.
Lobo, S, 2005. Macroglândulas parotóides de sapos (Anura: Bufonidae):
um estudo integrativo. Dissertação (Mestrado em Morfologia),
Departamento de Morfologia, Universidade Federal de São Paulo, São
Paulo.
Lobo, S., Piffer, T.R.O., Antoniazzi, M.M., Carneiro, S.M., Jared, C., 2005.
Morphophysiology of the paratoid macroglands in toads (Amphibia,
Anura, Bufonidae). Mem. Inst. Butantan 61, 84.
Luther, W., 1971. Distribution, biology and classification of salamanders.
In: Bücherl, W., Buckley, E.E. (Eds.), Venomous Animals and their
Venoms, vol. 2. Academic Press Inc., Boston, pp. 557–568.
Lutz, A., 1925. Batraciens du Brésil. C.R. Séances Soc. Biol. Paris 93,
211–214.
Lutz, B., 1966. Biological significance of cutaneous secretions in toads and
frogs. Mem. Inst. Butantan 33, 55–59.
Lutz, B., 1971. Venomous toads and frogs. In: Bücherl, W., Buckley, E.E.
(Eds.), Venomous Animals and their Venoms, vol. 2. Academic Press,
Boston, pp. 423–473.
Maciel, N.M., Schwartz, C.A., Pires, O.R., Sebben, A., Castro, M.S., Sousa, M.
V., Fontes, W., Schwartz, E.N.F., 2003. Composition of indolealkylamines of Bufo rubescens cutaneous secretions compared to six other
Brazilian bufonids with phylogenetic implications. Comp. Biochem.
Physiol 134, 641–649.
McDowell, E.M., Trump, B.F., 1976. Histologic fixatives suitable for diagnostic
light and electron microscopy. Arch. Pathol. Lab. Med 100, 405–414.
Nosi, D., Terreni, A., Alvarez, B.B., Delfino, G., 2002. Serous gland polymorphism in the skin of Phyllomedusa hypochondrialis azurea
(Anura, Hylidae): response by different gland types to norepinephrine
stimulation. Zoomorphology 121, 139–148.
207
Nowak, R.T., Brodie, E.D., 1978. Rib penetration and associated antipredator adaptations in the salamander Pleurodeles waltl (Salamandridae).
Copeia 1978, 424–429.
Pasquarelli, P., Mendes, E.G., Sawaya, P., 1987. The action of parotoid
venom on the heart of the toad (Bufo ictericus ictericus Spix 1824) and
its effects on the inhibition caused by vagal stimulation. Comp. Biochem. Physiol 87, 393–399.
Phisalix, M., 1922. Animaux Venimeux et Venins. Masson & Co., Paris.
Pineau, X., Romanoff, C., 1995. Envenomation of domestic carnivorous.
Rec. Méd. Vét 171, 182–192.
Reynolds, E.S., 1963. The use of lead citrate at high pH as an electronopaque stain in electron microscopy. J. Cell. Biol. 17, 208–212.
Rossi, M.H., Blumenthal, E.E.A., Jared, C., 1997. Bufadienolides from the
venom of Bufo paracnemis (Amphibia, Anura, Bufonidae). An. Assoc.
Bras. Quı́m 46, 21–26.
Sakate, M., Lucas de Oliveira, P.C., 2000. Toad envenoming in dogs: effects
and treatment. J. Venom. Anim. Toxins 6, 1–9.
Sazima, I., Caramaschi, U., 1986. Descrição de Physalaemus deimaticus,
sp. n., e observações sobre comportamento deimático em Physalaemus nattereri (Steindachner) – Anura. Leptodactylidae. Rev. Biol.
13, 91–101.
Shimada, K., Miyashiro, Y., Nishio, T., 2006. Characterization of in vitro
metabolites of toad venom using high-performance liquid chromatography and liquid chromatography–mass spectrometry. Biomed.
Chromatogr 20, 1321–1327.
Sonne, L., Rozza, D.B., Wolffenbüttel, A.N., Meirelles, A.E.W.B., Pedroso, P.M.O.,
Oliveira, E.C., Driemeier, D., 2008. Intoxicação por veneno de sapo em um
canino. Cienc. Rural 38, 1787–1789.
Stevaux, M.N., 2002. A new species of Bufo Laurenti (Anura, Bufonidae)
from Northeastern Brazil. Rev. Bras. Zool 19, 235–242.
Tempone, A., Pimenta, D., Lebrun, I., Sartorelli, P., Taniwaki, N., Andrade Jr., H.,
Antoniazzi, M.M., Jared, C., 2008. Antileishmanial and antitrypanosomal
activity of bufadienolides isolated from the toad Rhinella jimi parotoid
macrogland secretion. Toxicon 52, 13–21.
Terreni, A., Nosi, D., Greven, H., Delfino, G., 2003. Development of serous
cutaneous glands in Scinax nasica (Anura, Hylidae): patterns of
poisons biosynthesis and maturation with larval glands in specimens
of other families. Tissue Cell 35, 274–287.
Toledo, R.C., Jared, C., 1989a. Considerações sobre o veneno de anfı́bios.
Cienc. Cult 41, 250–258.
Toledo, R.C., Jared, C., 1989b. Estudo histológico das glândulas lombares
de Pleurodema thaul (Amphibia, Anura, Leptodactylidae). Rev. Bras.
Biol. 49, 421–428.
Toledo, R.C., Jared, C., 1993. The calcified dermal layer in anurans. Comp.
Biochem. Physiol 104, 443–448.
Toledo, R.C., Jared, C., 1995. Cutaneous granular glands and amphibian
venoms. Comp. Biochem. Physiol 111, 1–29.
Toledo, R.C., Villa, N., 1987. Estudo histológico das glândulas tibiais de Bufo
paracnemis (Amphibia, Anura, Bufonidae). Rev. Bras. Biol. 47, 257–264.
Toledo, R.C., Jared, C., Brunner, A., 1992. Morphology of the large granular
alveoli of toad (Bufo ictericus) parotoid glands before and after
compression. Toxicon 30, 745–753.
Toledo, R.C., Jared, C., Brunner, A., 1996. The lumbar glands of the frog
Pleurodema thaul (Amphibia, Anura, Leptodactylidae): an ultrastructural study. Rev. Bras. Biol. 56, 451–457.
Tyler, M.J., Burton, T., Bauer, A.M., 2001. Parotid or parotoid: on the
nomenclature of an amphibian skin gland. Herpetol. Rev. 32, 79–81.
Vital Brazil, O., Vellard, J., 1925. Contribuição ao estudo do veneno de
batrachios do gênero Bufo. Braz. Med 2, 175–180.
Vital Brazil, O., Vellard, J., 1926. Contribuição ao estudo do veneno de
batrachios. Mem. Inst. Butantan 3, 7–70.
Wilber, C.G., Carroll, P.L., 1940. Studies on the histology of the glands in
the skin of Anura. I. The parotoid gland of Bufo americanus Holbrook.
Trans. Am. Microsc. Soc 59, 123–128.