Toxicon 57 (2011) 512–524
Contents lists available at ScienceDirect
Toxicon
journal homepage: www.elsevier.com/locate/toxicon
Review
On the venom system of centipedes (Chilopoda), a neglected group
of venomous animals
Eivind A.B. Undheim*, Glenn F. King
Institute for Molecular Bioscience, The University of Queensland, St. Lucia, Queensland 4072, Australia
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 2 September 2010
Received in revised form 8 December 2010
Accepted 11 January 2011
Available online 19 January 2011
Centipedes are among the oldest extant terrestrial arthropods and are an ecologically
important group of soil and leaf litter predators. Despite their abundance and frequent,
often painful, encounters with humans, little is known about the venom and venom
apparatus of centipedes, although it is apparent that these are both quite different from
other venomous lineages. The venom gland can be regarded as an invaginated cuticle and
epidermis, consisting of numerous epithelial secretory units each with its own unique
valve-like excretory system. The venom contains several different enzymes, but is strikingly different to most other arthropods in that metalloproteases appear to be important.
Myotoxic, cardiotoxic, and neurotoxic activities have been described, most of which have
been attributed to high molecular weight proteins. Neurotoxic activities are also unusual in
that G-protein coupled receptors often seem to be involved, either directly as targets of
neurotoxins or indirectly by activating endogenous agonists. These relatively slow
responses may be complemented by the rapid effects caused by histamines present in the
venom and from endogenous release of histamines induced by venom cytotoxins. The
differences probably reflect the ancient and independent evolutionary history of
the centipede venom system, although they may also be somewhat exaggerated by the
paucity of information available on this largely neglected group.
Ó 2011 Elsevier Ltd. All rights reserved.
Keywords:
Centipedes
Chilopoda
Venom
Venom gland
1. Introduction
Class Chilopoda, or centipedes, represents one of the four
major myriapod lineages (Arthropoda; Myriapoda). They
are present on every continent except Antarctica and are an
important group of terrestrial predatory arthropods. There
are about 3300 species worldwide within five extant orders:
Scutigeromorpha, Lithobiomorpha, Craterostigmomorpha,
Geophilomorpha, and Scolopendromorpha (Fig. 1). Several
morphological characters unite the members of Chilopoda,
including a single pair of antennae, three pairs of mouthparts, a posterior genital opening (opisthogoneate), and 15
to 191 trunk-segments each bearing a single pair of walking
* Corresponding author. Tel.: þ61 7 3346 2322; fax: þ61 7 3346 2101.
E-mail addresses: [email protected], eivindandreas@gmail.
com (E.A.B. Undheim).
0041-0101/$ – see front matter Ó 2011 Elsevier Ltd. All rights reserved.
doi:10.1016/j.toxicon.2011.01.004
legs (Edgecombe and Giribet, 2007; Lewis, 1981). However,
the most obvious, and infamous, shared derived character of
centipedes is the modification of the first pair of walking
legs into venomous appendages often called poison claws,
forcipules, toxicognaths, or maxillipeds.
The maxillipeds are shaped like a set of piercing forceps,
each consisting of four segments: a large trochanteroprefemur, two short segments (tibia and tarsus), and
a powerful apical claw termed the tarsungula. The outer
surface of each claw has several pits with protruding pegs
similar to those involved in chemoreception in insects,
which may be used for tasting prey, stimulating the
secretion of venom by sensing penetration of prey, or both
(Jangi and Dass, 1977; Ménez et al., 1990). Venom is
secreted through a pore located on the outer curvature near
the tip of each claw, which again is connected to each
maxilliped’s venom gland through a chitinous venom duct.
E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
513
centipedes. This review, which aims to gather the widely
scattered literature on the venom and venom apparatus of
centipedes, reveals that centipedes represent a venomous
arthropod lineage which is very different from those
already described.
2. Centipede phylogeny
Fig. 1. A scolopendromorph commonly found in Australia, Ethmostigmus
rubripes (Scolopendridae).
Centipedes are thought to have split from the remaining
myriapods about 440 million years ago (mya) (Murienne
et al., 2010). The oldest recognizable order from the fossil
record is the Scutigeromorpha, of which fossilized legs
belonging to a Crussolum sp. have been found from the late
Silurian almost 420 mya (Shear and Edgecombe, 2009). The
earliest fossilized maxillipeds, from the early Devonian about
400 mya, belonged to another scutigeromorph of the genus
Crussolum and are very similar to those of the modern scutigeromorph Scutigera coleoptrata (Anderson and Trewin,
2003). The remaining orders probably all arose well before
this, however, with the last two orders to diverge being
Scolopendromorpha and Geophilomorpha which split just
over 410 mya (Murienne et al., 2010). This is similar to the
divergence of scorpions, the earliest fossils of which date back
to the mid-Silurian 428 mya (Dunlop, 2010). The centipede
venom apparatus thus represents one of the oldest extant
venom systems known among terrestrial animals.
Unlike scorpions, however, whose venom system is
among the most extensively studied of any animal, relatively little is known about the ancient venom system of
For practical purposes, the phylogenies of the chilopod
orders are briefly described here (for a review on the
phylogeny and evolutionary biology of centipedes see
Edgecombe and Giribet, 2007). As summarized in Fig. 2, the
Scutigeromorpha forms a sisterclade (sub-class Notostigmophora) to the remaining orders (sub-class Pleurostigmophora) based on a suite of traits including compound
eyes and dorsal spiracles. Lithobiomorpha forms the basal
outgroup within the Pleurostigmophora, and is followed by
the previously monotypic Craterostigmomorpha, which
like the lithobiomorphs also has anamorphic development,
leaving Scolopendromorpha and Geophilomorpha as the
sole epimorphic orders (clade Epimorpha). The higherlevel systematics of each order is relatively uncontroversial
and is summarized at the bottom of Fig. 2. Briefly, the
orders of Chilopoda contain one to three families each with
the exception of Geophilomorpha, which contains 14
families. 13 of these families are often grouped in a clade
termed Adesmata; all members of this clade have ventral
defensive glands and a variable number of segments within
species (Edgecombe and Giribet, 2007).
3. The venom gland
3.1. General ultrastructure
The venom glands of Chilopoda can be regarded as
invaginations of the cuticle containing a high number of
epidermal sub-glands, with each sub-gland, or secretory unit,
consisting of four different cell types (Hilken et al., 2005;
Fig. 2. Conceptual diagram of the phylogeny of centipedes (Chilopoda) based on the work by Edgecombe and Giribet (2004, 2007), Edgecombe and Koch (2008),
and Koch et al. (2009). Note that 13 of the 14 families within Geophilomorpha are often grouped in a clade termed Adesmata.
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Fig. 3. The centipede venom gland based on Ethmostigmus rubripes (A–C) and Lithobius forficatus (D). (A) Placement of the venom gland (vg) in E. rubripes
maxilliped, which spans from the tarsungula (t) to the proximal region of the trochanteroprefemur (tr) near the coxosternum (cs). (B) Longitudinal section of the
same venom gland showing the venom duct (vd), eccentric perforated lumen (l) and secretory epithelium (se) surrounded by the peripheral muscle layer (pm)
and basal lamina (bl). (C) Same gland again viewed in cross-section showing the radial muscle fibers (rm) and secretory units (su, framed). (D) A schematic
representation of the secretory unit, which consists of a chitinised epithelial cell (ec), canal cell (cc), intermediate cell (ic), and secretory cell (sc) containing
secretory granules (sg) released into the secretory body (sb) by exocytosis. The nozzle-like structure can be seen in the atrium of the epithelial cell. Diagrams are
adapted from Ménez et al. (1990) (A, C) and Rosenberg and Hilken (2006) (D).
Lewis, 1981; Rosenberg and Hilken, 2006) (Fig. 3). The lumen
of the venom gland, often referred to as the poison calyx, is
continuous with the venom duct and is lined with a single
layer of epithelial cells with a well-developed chitinous
cuticle, each of which encompasses an atrium forming a pore
in the lumen wall (Antoniazzi et al., 2009; Dass and Jangi,
1978; Ménez et al., 1990; Nagpal and Kanwar, 1981;
Rosenberg and Hilken, 2006). Connected proximally to each
perforate epithelial canal cell is a larger and more complex
canal cell that encloses a secretory duct and, along with
a smaller intermediary cell, a large non-cellular secretory
body (Rosenberg and Hilken, 2006). Lateral to the secretory
body are one or two large secretory cells that are connected to
the secretory body through the intermediary cell (Rosenberg
and Hilken, 2006). The nucleus of each secretory cell is
located in the wider peripheral (basal) region of the cell,
which is also rich in rough endoplasmic reticulum and
mitochondria (Antoniazzi et al., 2009). In this region of the
secretory cells the venom can be observed as dense secretory
granules 24–1600 mm in diameter, which in the case of
Otostigmus and Ethmostigmus (Scolopendridae: Otostigminae) often contain rod-like structures (Antoniazzi et al., 2009;
Ménez et al., 1990). As maturation of the venom occurs, the
contents of each granule become more amorphous and
transparent, and are eventually discharged into the secretory
body through exocytosis (Antoniazzi et al., 2009; Ménez et al.,
1990; Rosenberg and Hilken, 2006).
It has been suggested that venom secretion is holocrine
based on the presence of degenerate secretory cells
(Cornwall, 1916; Dass and Jangi, 1978; Rosenberg and
Hilken, 2006). However, no evidence of holocrine secretion was found by neither Antoniazzi et al. (2009), Jarrar
(2010), or Ménez et al. (1990). Rather, venom has been
described as secreted from the secretory cells by exocytosis
from both Lithobiomorpha (Rosenberg and Hilken, 2006)
and Scolopendromorpha (Ménez et al., 1990), although
only the former presented morphological data to support
this. It therefore seems that the presence of degenerating
secretory cells has perhaps been wrongfully interpreted as
evidence for holocrine secretion, and that the mode of
secretion is actually merocrine. Constriction of the gland is
a dramatic process that is likely to generate trauma to the
cuticle-lacking secretory cells, some which may consequently degenerate and be replaced by division and
differentiation of the intermediate cells (Dass and Jangi,
1978; Rosenberg and Hilken, 2006). In scorpions, for
example, cells do occasionally degrade and nuclear cell
products can be found in the venom despite their apocrine
mode of venom secretion (Hjelle, 1990).
Although it has been suggested that the venom-gland
lumen is the primary location for venom storage based on
the presence of venom-like components (Antoniazzi et al.,
2009; Ménez et al., 1990), these compounds are likely the
remains left after venom expulsion. Instead, it appears that
venom is stored in the secretory body of each secretory
unit, described as “extracellular space” by Rosenberg and
Hilken (2006), which probably corresponds to the “bag of
secretion” described in earlier studies (Cornwall, 1916).
Interestingly, a thin fold of the cuticle of the canal cell
protrudes deeply into the atrium of the epithelial cell,
forming a one-way nozzle (Antoniazzi et al., 2009; Dass and
Jangi, 1978; Rosenberg and Hilken, 2006) (Fig. 3D). Proximal to this nozzle, a dense sphincter-like structure is
present in the cell wall of the canal cell, which is thought to
be under neuro-musculatory control and responsible for
regulating the secretion of venom into the lumen
E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
(Antoniazzi et al., 2009; Dass and Jangi, 1978; Rosenberg
and Hilken, 2006). As the secretory bodies are in effect
branches of the lumen, the nozzle-like structures may also
provide a physical barrier to pathogenic agents that may
enter through the open venom duct.
In scolopendromorphs, septa of radial striated muscle
fibers are interspersed between the secretory units, and
are connected to the lumen wall in one end and the
peripheral muscle fibers in the other (Antoniazzi et al.,
2009; Dass and Jangi, 1978; Ménez et al., 1990) (Fig. 3C).
The peripheral muscle layer is composed of both longitudinal and circular striated muscle fibers, which are
finally enclosed by a layer of connective tissue often
referred to as the basal lamina (Antoniazzi et al., 2009;
Dass and Jangi, 1978; Ménez et al., 1990). A striated
peripheral muscle layer also surrounds the venom glands
of geophilomorphs (Duboscq, 1898). It is thought that
these muscles are responsible for contraction and
constriction of the gland during venom ejection (Dass and
Jangi, 1978; Ménez et al., 1990). However, neither the
peripheral nor radial muscle fibers were observed in
Lithobius forficatus (Rosenberg and Hilken, 2006), which
may suggest a different method of gland constriction in
more basal chilopods. Depending on the size and position
of the venom gland, the surrounding musculature may be
responsible for venom ejection by squeezing the gland
against the cuticle in a similar fashion to that of scorpions
(Hjelle, 1990). This remains speculative, however, as the
venom glands of only a few species have been described
in detail, none of which belong to either Scutigeromorpha
or Craterostigmomorpha.
3.2. Shape, size and placement
The size, shape and placement of centipede venom
glands are variable both within and between orders. In the
three basal orders Scutigeromorpha, Lithobiomorpha and
Craterostigmomorpha, the elongate venom gland is situated distally in the maxilliped, generally spanning from the
distal segment (tarsungula) to the distal half of the trochanteroprefemur along the outer curvature of the maxilliped (Chao and Chang, 2006; Edgecombe and Giribet,
2008; Edgecombe and Barrow, 2007; Rosenberg and
Hilken, 2006). The lumen, or calyx, is generally more or
less cylindrical and is restricted to the distal 10% of the
gland (Chao and Chang, 2006; Rosenberg and Hilken,
2006), a trait that seems to be plesiomorphic within the
Chilopoda.
Also in Geophilomorpha are the glands lacking
a chitinous lumen for most of the length (Duboscq, 1898). In
some (e.g. schendylids and mecistocephalids), the gland
contains only a simple cylindrical lumen much like that of
the more basal centipede orders (Foddai et al., 2003;
Hoffman and Pereira, 1991). In others such as geophilids,
however, numerous chitinous tubes radiate from the lumen
into the gland in a bush- or lampbrush-like arrangement
(Foddai and Minelli, 1999). The glands are generally cylindrical in shape (Bonato et al., 2006) and contained in the
trochanteroprefemur, although there are some fascinating
exceptions: In members of Marsikomerus, a genus in the
family Schendylidae, the entire venom apparatus is
515
contained within the tarsungula and includes a very small
venom gland (Hoffman and Pereira, 1991). In Henia (Chaetechelyne) vesuviana, a dignathodontid (Dignathodontidae),
the venom glands are located in the trunk, between the
12th and 18th segments (Duboscq, 1898), whereas in the
related Aphilodon angustatus (Aphilodontidae), the glands
are placed even further back into the trunk, between the
15th and 23rd segments, with one gland in front of
the other and each gland occupying most of the volume of
the three segments it spans (Pereira et al., 2007).
In the Scolopendromorpha, the relative length of the
lumen of the venom gland differs between the families
(Edgecombe and Koch, 2008; Lewis, 1981, 2002). In Cryptopidae, the general shape of the gland and lumen appears
to be similar to that of the Lithobiomorpha (Chao and
Chang, 2006; Lewis, 2002). The size and shape of the
venom gland in cryptopids varies considerably, ranging
from comparatively large and pear-shaped (Antoniazzi
et al., 2009; Duboscq, 1898) to just a few glandular cells
at the terminal end of the venom duct in Cryptops daszaki
(Lewis, 2002). In Scolopendridae, however, the lumen
spans nearly the entire length of the elongate cylindrical
venom gland (Chao and Chang, 2006) (Fig. 3B). This may
allow greater control over the secretion of venom components, although differential synthesis of toxins has not been
demonstrated in centipedes. The glands generally span
along the outer curvature of the trochanteroprefemur of
each maxilliped (Antoniazzi et al., 2009; Chao and Chang,
2006; Ménez et al., 1990; Nagpal and Kanwar, 1981),
although exceptions include Asanada socotrana and
Arthrorhabdus formosus where the glands extend into the
posterior part of the maxilliped coxosternite (Edgecombe
and Koch, 2008).
Like scolopendrids, the venom glands of some scolopocryptopids contain a lumen of almost equal length to the
gland (Chao and Chang, 2006; Snodgrass, 1965). It seems,
however, that this feature, along with the shape and size of
the gland, varies considerably even within genera.
Anatomical drawings of Scolopocryptops sexspinosa (prev.
Otocryptops Crabill, 1953), for example, depicts the venom
glands as ranging from the posterior part of the maxilliped
coxosternite to the distal end of the trochanteroprefemur
with the lumen spanning the length of the gland (Snodgrass,
1965). In S. rubiginosus and S. capillipedatus, the gland and
lumen reach the entire length of the trochanteroprefemur in
a serpentine fashion, terminating near the coxosternal joint
(Chao and Chang, 2006). In contrast, the lumen of the venom
glands in Newportia longitarsis and the troglobitic Thalketops
grallatrix are rather short and contained only in the distal
part of the trochanteroprefemur, but are relatively cylindrical and elongate compared to the almost globular lumen
of the closely related Kethops euterpe (Crabill, 1960;
Edgecombe and Koch, 2008). Although no glands were
described in these studies, the short and wide lumens
suggest that the length of the lumen in relation to the venom
gland is rather like that found in the other chilopod orders.
The basal position of the species with relatively short
lumens within Scolopocryptopidae (Edgecombe and Koch,
2008) suggests that an elongation of the venom gland
lumen is a derived feature that has arisen at least twice in
the Scolopendromorpha.
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4. Clinical importance
Centipede stings are fairly frequent, usually nondramatic events largely caused by scolopendromorphs,
most of which are of such benign nature that they are never
reported to hospitals or physicians (Balit et al., 2004; Lin
et al., 1995; Serinken et al., 2005; Vazirianzadeh et al.,
2007). In Brazil, about five percent of envenomations
reported to the Vital Brazil Hospital of the Butantan Institute were due to centipedes, none of which resulted in
death or subsequent complications (Knysak et al., 1998;
Medeiros et al., 2008). Although several fatalities have
been attributed to centipedes, most of these appear to
result not from the venom but rather secondary infections
that are commonly associated with centipede stings (Blay,
1955; Marsh, 1957; Uzel et al., 2009; Veraldi et al., 2010);
such infections may result in serious complications and in
very rare cases death (e.g., by necrotizing fasciitis)
(Serinken et al., 2005). There may also be instances where
a centipede is interpreted as a plausible cause of death due
to its presence on or near the corpse, despite a total absence
of evidence for its actual involvement in the mortality (e.g.
some citations of the case reported by Harada et al. 1999).
Langley (2005) and Langley and Morrow (1997) found that
centipedes were responsible for six deaths in the USA
between 1979 and 2001, although no observations were
given as to the actual causes of death. In comparison, they
also reported a total of 1060 fatalities due to wasps, bees
and hornets from the same time period. Thus, the only
apparently well documented mortalities from a centipede
sting are that of a 7 year-old girl in the Philippines that was
stung on the head, presumably by S. subspinipes, and died
29 h later (Pineda, 1923 cited in e.g. Remington, 1950), a 21
year-old female stung by a centipede in Thailand (Balit
et al., 2004), and an army officer from Rodrigues that
accidently drank a small centipede and was stung in the
back of the throat probably resulting in death by asphyxiation (Lewis et al., 2010).
Several centipede species are capable of inflicting severe
symptoms in humans (Supplementary Table), including
myocardial ischemia and infarction (Ozsarac et al., 2004;
Yildiz et al., 2006), hemoglobinuria and hematuria
(Vazirianzadeh et al., 2007), hemorrhage (Knysak et al.,1998),
and rhabdomyolysis (Logan and Ogden, 1985; Rouvin et al.,
2008). Several allergy-like complications have been recorded after centipede envenomations, such as severe itching
(Bush et al., 2001; Mumcuoglu and Leibovici,1989), fever and
chills (Lin et al., 1995), general rash (Supakthanasiri et al.,
2004), eosinophilic cellulitis (Friedman et al., 1998), and
anaphylaxis (Supakthanasiri et al., 2004). A Type IV allergic
reaction triggered by centipede venom was the presumed
cause of death of a 21 year-old female (Balit et al., 2004). Most
centipede envenomations, however, produce transient
symptoms that can last a few days but do not require acute
medical treatment (Bush et al., 2001; Lin et al., 1995; Mohri
et al., 1991; Remington, 1950).
Virtually all envenomations by centipedes are characterized by an instant, local, burning pain that ranges in
intensity from excruciating to mild and in some cases
radiates or spreads to other parts of the victim’s body (e.g.,
see Baerg, 1924, 1925, Balit et al., 2004, Medeiros et al.,
2008, Remington, 1950, Sutherland and Tibballs, 2001).
The level of pain appears to differ between taxa. Cormocephalus spp. typically cause only mild pain whereas Ethmostigmus spp. and larger Scolopendra spp. usually cause
severe pain, although this could merely reflect the size of
specimens and hence amount of venom injected. After
pain, localized swelling and erythema are the most
common symptoms of centipede envenomations, although
it seems that swelling is more common in cases involving
otostigmine scolopendrids such as Otostigmus spp. and
Ethmostigmus spp. (Balit et al., 2004; Knysak et al., 1998;
Medeiros et al., 2008). There are also several other relatively frequent complications associated with stings, such
as localized necrosis, paresthesia, lethargia, hot sensation
around the sting site, dizziness, headache and nausea (Bush
et al., 2001; Coffin, 1919; Sutherland and Tibballs, 2001).
Thus, while generally of a relatively benign nature, centipede envenomations are still able to produce trauma,
particularly in cases involving allergies and small children.
5. Effect of centipede venom on animals
While centipedes cannot be said to be lethal to humans,
they are capable of killing relatively large mammals, such as
in the case reported by McKeown (1930) where a 4 year-old
bull terrier died from the sting of a 15 cm Ethmostigmus
rubripes. Large scolopendrid species are known to feed on
vertebrates, including bats (Molinari et al., 2005), rats (Clark,
1979), amphibians (Carpenter and Gillingham, 1984; Forti
et al., 2007) and reptiles (Easterla, 1975), although the
toxicity towards mammals differs between taxa. Bücherl
(Bücherl 1946 in Minelli 1978) found that the LD50 in adult
mice with body weights about 20 g varied tenfold between
scolopendromorphs; the median intramuscular LD50 in
terms of number of individual glands per mouse ranged from
0.012 to 0.16 in equally sized Otostigmus scabricauda (Scolopendridae; Otostigminae) and Scolopocryptops ferrugineus
(Scolopocryptopidae), respectively. The corresponding
values for Cryptops iheringi (Cryptopidae), Scolopendra viridicornis and S. subspinipes (Scolopendridae; Scolopendrinae)
were 0.34, 0.25, and 1.2 glands respectively, although both
specimens of Scolopendra were more than twice the size of
the remaining species. In E. rubripes (Otostigminae), injecting
venom gland extract into the body cavities of juvenile mice
with body weights of about 3.4–4.2 g gave an LD100 of
0.01 0.02 glands/gram mouse (Ménez et al., 1990). Venoms
of S. viridicornis, Otostigmus pradoi and C. iheringi also induced
pain in mice even with very small doses of venom (0.9 mg)
(Malta et al., 2008). Centipede venoms also have potent
insecticidal properties, similar to or exceeding that of many
spiders (Quistad et al., 1992). When Ménez et al. (1990)
injected the equivalent of 0.3 glands from E. rubripes into
the abdomen of Locusta migratoria, for example, death
occurred within a few seconds, whereas the LD100 in the
moth Helicoverpa punctigera for the same venom was determined to be 0.02 0.02 glands/gram.
6. Venom composition
The long list of symptoms and complications induced
by centipede envenomations suggests that the venom
E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
comprises a diverse cocktail of toxins. In their comparative
electrophoretic analysis of three scolopendromorph venoms obtained by electrostimulation, including one cryptopid and two scolopendrid species, Malta et al. (2008)
showed that all venoms contained proteins ranging in size
from about 7 kDa to almost 200 kDa, that several of these
probably were polymeric proteins, and that all venoms
exhibited a range of toxic activities. Rates et al. (2007) found
62 and 65 protein species by 2D chromatographic analysis of
S. viridicornis nigra and S. angulata venom, respectively, with
molecular masses ranging from 1.3 to 22.6 kDa. N-terminal
sequencing of 13 and 11 of these protein species from S.
v. nigra and S. angulata, respectively, yielded a total of 10
protein families, nine of which did not show significant
similarities to protein sequences in the UniProt database.
Although the work of Rates et al. (2007) and Malta et al.
(2008) are two of a very limited number of modern studies
exploring the toxinological diversity of centipede venoms
obtained by milking by electrostimulation of living specimens, several toxins and toxic activities have been described
from the venom milked by electrostimulation and venom
gland extractions from various species since the first
demonstration of the insecticidal properties of centipede
venom by Jules MacLeod in 1878 (Lewis,1981). These consist
of both non-protein and protein compounds, of which the
latter can be further divided into enzymes and those
without enzymatic activity (Table 1).
517
6.1. Enzymes
6.1.1. Esterases
Various non-specific esterases have been detected from
the maxilliped extract of Scolopendra morsitans, including
four a-naphthyl acetate positive esterases, three acid
phosphatases and four alkaline phosphatases (Kass and
Batsakis, 1979; Mohamed et al., 1983). Non-specific esterases are found in the venoms of a diverse set of taxa,
including spiders (Heitz and Norment,1974; Rodrigues et al.,
2006), snakes (Sulkowski et al., 1963; Tu and Chua, 1966)
and octopus (Undheim et al., 2010). Although their
venomous function is uncertain, they are thought to perhaps
play a part in the release of endogenous purines during
envenomation, which then act as multitoxins causing
a multitude of pharmacological effects including immobilization through hypotension (Aird, 2002; Dhananjaya and
D’Souza, 2010). In hymenopterans, acid phosphatase also
acts as an allergen (Lima and Brochetto-Braga, 2003).
Another type of esterase that is perhaps more interesting in the context of venoms are type A2 phospholipases
(PLA2). PLA2 activity has been described from the milked
venoms of both the scolopendrid subfamilies Otostigminae
(O. pradoi) and Scolopendrinae (S. viridicornis and S. viridis)
(González-Morales et al., 2009; Malta et al., 2008).
However, the only characterized centipede venom PLA2 is
Scol/Pla from Mexican specimens of S. viridis described by
Table 1
Toxins and toxic activities found in centipedes.
Toxin type/activity
Enzymes
Acid phosphatases
Alkaline phosphatases
Phospholipase A2
General esterases
Hyaluronidases
Metalloproteases
Non-metalloproteases
Non-enzymatic proteins
Cardiotoxins
CRISPs
Species
Molecular weight
Reference
Scolopendra morsitans
S. morsitans
Otostigmus pradoi
S. viridicornis
S. viridis
S. morsitans
O. pradoi
S. viridicornis
S. morsitans
Cryptops iheringi
O. pradoi
S. viridicornis
C. iheringi
O. pradoi
2 slow, 1 fast band on PAGE
3 slow, 1 med band on PAGE
(Mohamed et al., 1983)
(Mohamed et al., 1983)
(Malta et al., 2008)
(Malta et al., 2008)
(González-Morales et al., 2009)
(Mohamed et al., 1983)
(Malta et al., 2008)
(Malta et al., 2008)
(Mohamed et al., 1983)
(Malta et al., 2008)
(Malta et al., 2008)
(Malta et al., 2008)
(Malta et al., 2008)
(Malta et al., 2008)
S. subspinipes
S. angulata
S. viridicornis
Disintegrins
S. viridicornis
Haemolysins
O. pradoi
S. subspinipes
S. viridicornis
Myotoxins
C. iheringi
O. pradoi
S. viridicornis
Neurotoxins
Scolopendra sp. (Brazil)
Scolopendra sp. (Mexico)
S. morsitans
S. viridicornis
Other non-peptidic active components
Histamine
S. subspinipes
Serotonin
S. morsitans
S. viridicornis
13.75 kDa
4 slow
32, 43–67 kDa
40–66 kDa
2 slow, 1 fast band on page
25, 44, 121 kDa
15, 23, 35 kDa
22, 42 kDa
25 kDa
23, 35 kDa
60 kDa
20’566 Da
20’417 Da
2594, 3017 Da
>80 kDa
1456–7360 Da
(Gomes et al., 1983)
(Rates et al., 2007)
(Rates et al., 2007)
(Bhagirath et al., 2006)
(Malta et al., 2008)
(Peng et al., 2009)
(Malta et al., 2008)
(Malta et al., 2008)
(Malta et al., 2008)
(Malta et al., 2008)
(Stankiewicz et al., 1999)
(Gutiérrez et al., 2003)
(Mohamed et al., 1980)
(Rates et al., 2007)
(Gomes et al., 1982a; Gomes et al., 1983)
(Mohamed et al., 1980)
(Welsh and Batty, 1963)
518
E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
Gonzalez-Morales et al. (2009). This PLA2 has five disulfide
bonds and a molecular weight of 13.75 kDa, which is typical
for venom PLA2, and appears to have high enzymatic
activity. Phylogenetic analysis shows that Scol/Pla clusters
with human group X PLA2 (Schaloske and Dennis, 2006),
although in a deeply split clade. Scol/Pla nevertheless
seems to be more similar to Group II PLA2 of snake venoms
rather than the Group III PLA2 found in hymenopteran and
scorpion venoms (Xin et al., 2009), which is not surprising
considering the time since divergence and consequent
analogous development of the venom systems.
Malta et al. (2008) were unable to detect PLA2 activity in
the venom of the cryptopid C. iheringi, suggesting perhaps
that PLA2 may not be ubiquitous in centipede venoms, but
rather independent recruitments in a few extant families,
which seems to be the case in spiders (Nagaraju et al., 2006;
Usmanov and Nuritova, 1994; Vassilevski et al., 2009).
However, the activities detected in O. pradoi and S. viridicornis were also very weak (Malta et al., 2008), and
Gutiérrez et al. (2003) failed to detect any phospholipase
activity in the venoms of Scolopendra sp. collected from the
same region as the S. viridis specimens from which Scol/Pla
was isolated (González-Morales et al., 2009). However, one
cannot exclude the possibility of an ancient recruitment of
PLA2s into centipede venoms followed by structural or
functional modifications leading to reduced or perhaps
even non-existent enzymatic activity, such as the myotoxic
[Lys49]PLA2s found in the venom of pitvipers (Viperidae:
Crotalinae) (Lomonte et al., 2003).
6.1.2. Proteases
Several proteases have been recorded from the venoms
of both scolopendrid and cryptopid scolopendromorphs.
Mohamed et al. (1983) detected at least three different
amino acid naphthylamidase-type aminopeptidases from
the maxilliped extract of S. morsitans, while Said (1958)
demonstrated the presence of both endo- and exopeptidases, including fibrinolytic but not fibrinogenolytic
activity, from the venom of the same species. In S. viridicornis two gel bands at about 22 and 44 kDa showed
caseinolytic and gelatinolytic, and fibrinogenolytic activities, respectively (Malta et al., 2008). In the otostigmine
O. pradoi, at least three gelatinolytic and caseinolytic
proteases were detected with molecular weights of about
15, 23, and 35 kDa, while in C. iheringi (Cryptopidae) three
gel bands with similar activities corresponding to 25, 44,
and 121 kDa were found (Malta et al., 2008). Most of the
activities found by Malta et al. (2008) were blocked by 1,10phenanthroline, suggesting that, unlike most arachnids
(Ma et al., 2009; Vassilevski et al., 2009), centipede venom
proteases are mainly metalloproteases.
Venom metalloproteases are an important constituent of
viperid snake venoms where they commonly act as
hemorrhagic factors by degrading extracellular matrix and
preventing blood clot formation (Gutiérrez and Rucavado,
2000). Although mainly associated with snake venoms,
metalloproteases have also been described from venoms of
the araneomorph spiders (Nagaraju et al., 2007) (TrevisanSilva et al., 2010), where they also appear to act primarily
as haemorrhagins. Said (1958) found that S. morsitans
maxilliped extract had fibrinolytic activity while Malta et al.
(2008) reported that O. pradoi and S. viridicornis venoms
both showed weak fibrinogenolytic activity. There are also
reports of anticoagulant activity in the venom of S. subspinipes (Bush et al., 2001). However, based on a skin test,
Malta et al. (2008) reported that only S. viridicornis exhibited
hemorrhagic activity, which was still very low compared to
that of Bothrops jararaca (Viperidae: Crotalinae). While local
hemorrhage has been reported from Cryptops sp. envenomation (Knysak et al., 1998), the results obtained by Malta
et al. (2008) suggest that centipede venom metalloproteases do not primarily function as haemorrhagins.
Metalloproteases are also involved in skin damage, edema,
blister formation, myonecrosis and inflammation, which is
consistent with several of the recurrent symptoms associated
with centipede bites (see Section 4 and Supplementary Table)
(Gutiérrez and Rucavado, 2000). The inflammatory effect
induced by B. jararaca venom, for example, has been shown to
be mainly due to the activity of metalloproteases (Zychar et al.,
2010). In addition, a non-hemorrhagic metalloprotease with
strong myotoxicity has been described from the venom of
Macrovipera lebetina (Viperidae: Viperinae) (Hamza et al.,
2010), suggesting that the hyperalgesic, high myotoxic and
high oedematogenic activities exhibited by the scolopendromorph venom (Malta et al., 2008) may at least partially be
due to the activities of metalloproteases.
Bhagirath et al. (2006) found disintegrin activity in
venom milked by electrostimulation of S. subspinipes
collected in Manipur, India. Disintegrins exert their toxic
effect by inhibiting platelet aggregation and, although they
are not enzymes per se, they are activated by cleavage of
the N-terminal domains of PII class metalloproteases
(Calvete et al., 2003). Although the anticoagulatory or
haemorrhagic effects of S. subspinipes venom have not been
determined, Bush et al. (2001) reported that blood “oozed
out” of the puncture marks for several minutes after being
stung by an adult S. subspinipes, suggesting that such
activities may be present.
Although the majority of proteolytic activity in centipede venoms seems to stem from metalloproteases, gelatinolytic activity from non-metalloproteases, most likely
serine proteases (Matsui et al., 2000), have also been
detected in the venoms of O. pradoi and C. iheringi (Malta
et al., 2008). These activities were weak, suggesting that
they may play a role in toxin activation by cleaving
precursor peptides to create one or more mature toxins,
rather than, for example, impeding the victim’s ability to
maintain hemostasis. Although no anti- or procoagulant
activity was detected by Malta et al. (2008) it is interesting
to note the experiment conducted by Briot (1904) where he
injected maxilliped extract from S. morsitans into the vein
behind the ear of a 2 kg rabbit; this resulted in death within
in 1 min and an autopsy revealed congealed blood in the
atrium, portal vein, and vena cava. However, no in-depth
study of centipede venom has ever been conducted to
investigate the diversity and function of its proteases.
6.1.3. Hyaluronidases
Hyaluronidases are found in most animal venoms, where
they are often regarded as “spreading factors” that increase
the pathological impact of venom components such
as haemorrhagins and neurotoxins (Girish et al., 2004;
E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
Kuhn-Nentwig et al., 2004; Long-Rowe and Burnett, 1994).
Hyaluronidase activity has been found in the venom of
scolopendrid centipedes, where strong activity was detected
in gel bands corresponding to masses of 40–66 kDa in S.
viridicornis and O. pradoi as well as an additional band of
weak activity at about 32 kDa in O. pradoi (Malta et al.,
2008). However, little activity was detected in the venom
of C. iheringi (Malta et al., 2008).
6.2. Non-enzymatic proteins
6.2.1. Myotoxins
As mentioned, the crude venoms of scolopendromorph
centipedes were shown to be highly myotoxic in that at
least as much creatine kinase could be detected from
muscle incubated with centipede venom as with the positive control, Bothrops jararacussu venom (Malta et al.,
2008). While metalloproteases may explain some of this
activity, myotoxins also include other types of venom
components, including myotoxic PLA2s and Low Molecular
Weight Myotoxins (LMWM) (Gutiérrez and Rucavado,
2000). For example, in most Bothrops spp., including B.
jararacussu, the observed myotoxicity has largely been
attributed to myotoxic PLA2s with varying degrees of
enzymatic activity (Gutiérrez and Lomonte, 1995). Unfortunately, the only PLA2 described from Chilopoda, Scol/Pla
(see Section 6.1.1), has not been assayed for any toxic
activities apart from its phospholipolytic activity.
A third possibility for the cause of the observed myotoxicity is the presence of LMWM. These may or may not
cause myonecrosis through breakdown of the sarcolemma
and a consequent increase in serum creatine kinase
(Fletcher et al., 1996). LMWMs are found in the venom of
vipers but snake-like LMWM, such as crotamines, do not
cause elevated levels of serum creatine kinase (Mebs et al.,
1983) and are thus unable to explain the strong release of
creatine kinase detected by Malta et al. (2008) in their
myotoxicity assay. Although the myonecrotic activity of
bee-venom melittin does cause a release of creatine kinase
into the serum, LMWMs also usually rapidly induce paralysis of skeletal muscle by interacting with voltage gated
sodium (NaV) channels on skeletal muscle sarcolemma to
cause depolarization and contraction (Lima and BrochettoBraga, 2003; Oguiura et al., 2005). Gomes et al. (1982b)
found that the venom of S. subspinipes is unable to cause
contractions in skeletal muscle, and thus it is unlikely that
much of centipede venom myotoxicity is due to LMWM.
While it seems that the myotoxicity induced by centipede venom is mainly restricted to localized tissue necrosis
in humans (see Section 4), this may not be the case in
smaller animals including centipede prey and predators.
Rhabdomyolysis and consequent myoglobinuria and renal
failure resulting from centipede envenomations has been
reported in humans (Logan and Ogden, 1985; Rouvin et al.,
2008) as have cases of proteinuria (Hasan and Hassan,
2005; Vazirianzadeh et al., 2007), and systemic myotoxicity could explain the prolonged time before death of the
dog in the case reported by McKeown (1930). With
decreasing body size of the victim, and thus increased
relative amount of venom, centipede venom myotoxins
519
may induce an effect of ecologically significant magnitude
within an ecologically relevant timeframe.
6.2.2. Cardiotoxins
Cardiotoxic effects have been demonstrated using
maxilliped extracts from S. morsitans and S. subspinipes
(Gomes et al., 1982b; Gomes et al., 1983; Mohamed et al.,
1980). Gomes et al. (1983) isolated a thermolabile, acidic,
cardiotoxic protein with a molecular weight of 60 kDa from S.
subspinipes which largely caused the same cardiovascular
effects as the maxilliped extract (Gomes et al.,1982b) and had
an intravenous LD50 in male mice of 41.7 mg/kg. They suggested that the toxin be referred to as “Toxin-S” until its
chemical nature is determined. The effect of Toxin-S on isolated hearts from toad, rat, guinea pig, and rabbit was found
be concentration dependent: at low concentrations (2 ng/ml)
a transient positive inotropic effect was observed in all hearts,
while at higher concentrations (mg/ml) the toxin caused a 50%
decrease in contractile amplitude in toad heart and irreversible cardiac arrest in the remaining hearts within 1 min
(Gomes et al.,1983). The latter concentration of the toxin also
caused a gradual negative inotropic effect on isolated guinea
pig auricle. In all cases the heart rate remained unaffected. In
mammals, Toxin-S also caused vasoconstriction, increased
capillary permeability, slow smooth muscle contraction, and
hypertension, but not skeletal muscle contractions or
hemolysis (Gomes et al., 1983).
In contrast to the effects observed using S. subspinipes,
Mohamed et al. (1980) found that application of very low
concentrations of maxilliped extract from S. morsitans
caused a triphasic response in the amplitude of rabbit
ileum rhythmic contractions consisting of an initial phase
of slight inhibition, a short phase of stimulation that was
prevented by lowering the buffer calcium concentration,
and finally a prolonged phase of gradual inhibition.
Mohamed et al. (1980) suggested that the increased ileum
contractions were due to the presence of serotonin based
on inhibition by cyproheptadine; however, in addition to
blocking the effect of serotonin, cyproheptadine is also an
antihistamine and calcium channel blocker (Stone et al.,
1961; Winquist et al., 1984). The S. morsitans maxilliped
extract also affected isolated toad heart in a different
manner to that of S. subspinipes, occasionally causing an
elevated heart rate soon after application of the extract
(Gomes et al., 1982b; Mohamed et al., 1980).
The toxins responsible for the cardiotoxicities observed
in S. morsitans and S. subspinipes appear to act on different
receptor types, indicating there could be substantial variation between taxa. Unfortunately, no information is
available on the cardiovascular effects of venoms from
species other than the two mentioned here. However,
cardiotoxins are probably also present in other species of at
least Scolopendra, as cardiovascular-related symptoms have
been reported from the stings of S. heros (Bush et al., 2001),
two unidentified Turkish centipedes (Ozsarac et al., 2004;
Yildiz et al., 2006), and one unidentified Taiwanese centipede (Chaou et al., 2009).
6.2.3. Neurotoxins
Neurotoxic proteins and peptides are arguably the most
important components of arthropod venoms as they
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E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
provide an efficient means of rapidly paralyzing prey
without expending much venom. While the relative
importance of neurotoxins in centipedes remains
unknown, such components are nevertheless present. For
example, centipede venom is known to cause instant rigid
paralysis in envenomated houseflies, cockroaches and
crickets (e.g. Rates et al., 2007). When Rates et al. (2007)
separated the venom of S. viridicornis using reverse-phase
HPLC, six out of 13 fractions showed various activities
including agitation, rapid leg and wing movement, and
salivation in the housefly Musca domestica. No fractions
were lethal, although one fraction with peptides ranging in
size from 1455 to 7360 Da caused reversible paralysis and
loss of motor control. In the same study, Rates et al. (2007)
found two proteins, each from the venom from S. angulata
and S. viridicornis, that had low but significant N-terminal
sequence homologies to the cysteine-rich secretory protein
(CRISP) allergen V from wasp venom and Tripurin from the
snake Trimeresurus purpureomaculatus (Viperidae; Crotalinae). The specific functions of these toxins is not known,
although CRISP in general appear to block voltage gated
calcium (CaV) channels and Kþ channels to induce smooth
muscle paralysis (Fry, 2005).
The first direct effects of centipede venoms on the insect
nervous system were described by Stankiewicz et al.
(1999), who isolated a neurotoxic high-molecular weight
fraction (>80 kDa) from the venom of an unspecified
species of Scolopendra from Brazil, which was obtained by
electrostimulation. This fraction, termed SC1, was shown to
cause an increase in leak current in giant axons of the
American cockroach Periplaneta americana, possibly due to
the formation or opening of non-specific pores in the
membrane by components in SC1 (Stankiewicz et al., 1999).
Further purification of SC1 by ion exchange chromatography abolished the activity, suggesting that the proteins
involved are either unstable or act synergistically. Although
venom PLA2 or phospholipase D activating peptides were
suggested as possible mediators of the membrane instability, these are usually stable structures and have
a molecular weight well below 80 kDa.
Using central cholinergic synapses from P. americana,
untransfected Xenopus oocytes, and Xenopus oocytes
expressing Drosophila muscarinic receptor 1, Stankiewicz
et al. (1999) showed that SC1 contains muscarinic
receptor agonists. This supports the contention of
Mohamed et al. (1980) that muscarinic agonists are present
in scolopendrid venoms, but also raises the question as to
what these agonists are. Very few muscarinic toxins have
been described; most are from snake venoms and belong to
the three-finger toxin (3-FTX) or PLA2 protein families
(Servent and Fruchart-Gaillard, 2009). Again, however, the
large size of the components in SC1 and their unstable
nature is atypical for most members of these families
(Servent and Fruchart-Gaillard, 2009; Stankiewicz et al.,
1999).
Gutiérrez et al. (2003) found that a fraction from venom
milked from a Mexican Scolopendra sp., when applied to
the ventral abdominal ganglia of the freshwater crayfish
Cambarellus cambarellus, caused release of the neurotransmitters g-aminobutyric acid (GABA) and glutamate. In
contrast with Stankiewicz et al. (1999), Gutiérrez et al.
(2003) found that this release was probably mediated via
increased membrane permeability to Naþ, as it was prevented by terodotoxin (a blocker of NaV channels), excessive extracellular Kþ, and an absence of extracellular Naþ.
This release of neurotransmitter was also observed by
Stankiewicz et al. (1999), who described an initial brief
depolarization nearly identical to that of acetylcholine
before the slower muscarinic response caused by SC1 set in.
However, the participation of Ca2þ channels could not be
excluded, and the precise mechanism involved in neurotransmitter release remains to be determined.
Stankiewicz et al. (1999) also found that SC1 agonized
a second G-protein-coupled receptor (GPCR), namely the
lysophosphatidic acid (LPA) receptor, which is a highly
unusual target for venom toxins. It has been shown,
however, that both PLA2 and Loxosceles phospholipase D
are able to produce LPA through the lysis of lysophosphatidylcholine (Lee and Lynch, 2005; Noguchi et al., 2009),
meaning that LPA-receptor activation could also be
a secondary event due to the release of endogenous LPA.
Interestingly, LPA and LPA-receptors are associated with
numerous cellular and physiological responses, including
Ca2þ mobilization in cells and the onset of peripheral pain
(Noguchi et al., 2009).
6.2.4. Cytotoxins
Venom cytotoxic peptides are generally of low molecular weight, lack cysteines, are highly cationic, and typically
have an isoelectric point (pI) > 8.75 (Kuhn-Nentwig, 2003).
Toxicity towards erythrocytes or various cultured cells has
been demonstrated in venoms from O. pradoi (Malta et al.,
2008), Scolopendra sp. (Cohen and Quistad, 1998), S. multidens (Peng et al., 2009), S. s. mutilans (Peng et al., 2009), S.
viridicornis (Malta et al., 2008) and C. iheringi (Malta et al.,
2008), and the maxilliped extraction of S. subspinipes deehani (Gomes et al., 1982b). It is noteworthy as part of this
list that although Cornwall (1916) is frequently cited as
demonstrating the hemolytic property of venom from
Ethmostigmus spinosus, the extract used in this study did
not contain venom glands nor any part of the maxillipeds.
While the observed cytotoxic activity from centipede
venoms may be partially, if not mostly, due to myotoxins
such as various PLA2, some of this activity is probably due
to the action of cytotoxic peptides whose primary function
may in fact be antimicrobial, that is, to prevent the establishment of pathogens in the otherwise benign growth
environment presented by the venom duct and gland
lumen.
Three so-called antimicrobial peptides have been
described from centipede venom, all from S. subspinipes
mutilans milked by electrostimulation. The first of these
was scolopendrin I, a 4498 Da peptide which was shown to
kill several species of Gram-positive and Gram-negative
bacteria as well as fungi, an activity that was unaffected by
pre-incubation at 100 C (Wenhua et al., 2006). Scolopendrin I showed no hemolytic or agglutination activity
towards mouse erythrocytes, perhaps suggesting it functions only as an antimicrobial agent or that any other
activities were abolished by the purification steps, which
included incubation at 100 C (Wenhua et al., 2006). The
sequence and structure of scolopendrin I remains to be
E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
determined, however, as does its mechanism of antimicrobial activity.
The remaining two antimicrobial peptides, scolopin-1
and scolopin-2, were described by Peng et al. (2009) as
21- and 25-residue peptides, respectively. Like most
antimicrobial peptides, scolopin-1 and -2 are cationic,
amphipathic, contain no cysteines, and have a pI > 9
(Kuhn-Nentwig, 2003; Peng et al., 2009). Both scolopin-1
and -2 potently kill fungi as well as Gram-positive and
Gram-negative bacteria, including antibiotic-resistant
strains of Staphylococcus aureus, Escherichia coli, and
Bacillus dysenteriae. Unlike scolopendrin I, they also show
moderate hemolytic activity (Peng et al., 2009; Wenhua
et al., 2006). More interestingly, however, scolopin-1
and -2 induce release of histamine from mast cells (Peng
et al., 2009). Thus, like many other “antimicrobial”
peptides, scolopin-1 and -2 might be involved in the
general toxic effect of the venom through release of
endogenous histamines (Kuhn-Nentwig, 2003).
6.3. Non-peptidic venom components
In addition to the peptidic components mentioned
above, a number of non-peptidic components have been
detected in centipede venoms. Mohamed et al. (1983)
reported lipoproteins, phospholipids, cholesterol, free
fatty acids, triglycerides, cholesterol esters, and squalene
from the venom of S. morsitans. These components,
however, were most likely cellular products and an artifact
of the venom extraction method, which consisted of
grinding entire maxillipeds with sand and mixing with
water (Mohamed et al., 1983). More interestingly, 5hydroxytryptamine (5-HT or serotonin) and histamine have
been described from centipede venoms, which are well
known algesics that cause instant sharp pain (Chahl and
Kirk, 1975).
Serotonin has been described in the milked venom of S.
viridicornis (Welsh and Batty, 1963) where its function was
proposed, along with histamine, acetylcholine and kinins,
to be defensive due to its ability to cause immediate pain
and to facilitate venom spreading via increased membrane
permeability. Serotonin was also proposed to be present in
S. morsitans venom as the venom induced hyperglycaemia
and liver glycogenolysis analogous to the effects caused by
serotonin when triggering the release of epinephrine from
the adrenal medulla (Mohamed et al., 1980). In addition to
causing pain, serotonin also causes an increase in vascular
permeability, which probably complements the effects of
venom cardiotoxins.
Like serotonin, histamine has also been described from
the venom system of Scolopendridae, including maxilliped
extracts from S. subspinipes (Mohamed et al., 1980). It is
likely that histamine is commonly involved in centipede
envenomations as treatment of patients with antihistamines usually results in immediate improvement
(Medeiros et al., 2008; Supakthanasiri et al., 2004). Histamine, like serotonin, causes an instant sharp pain and
increased vascular permeability (Chahl and Kirk, 1975). It is
also oedematogenic and is involved in immunological and
inflammatory responses, resulting in general swelling,
severe itching, general rash, and swollen lymph nodes, all
521
of which are fairly common symptoms following centipede
envenomation (Coffin, 1919; Lin et al., 1995; Rouvin et al.,
2008; Supakthanasiri et al., 2004).
However, the levels of histamine (Gomes et al., 1982a)
and serotonin (Welsh and Batty, 1963) in centipede venom
are probably not sufficient to cause the severe effects
attributed to them. It is therefore likely that most centipede
venoms contain histamine- and serotonin-releasing agents
such as scolopin-1 and -2 that cause release of endogenous
stores in envenomated individuals through degranulation
of mast cells (see Section 6.2.4), which is the case for both
hymenopteran (Lima and Brochetto-Braga, 2003) and
arachnid (Liu et al., 2007) venoms.
6.4. Function of centipede venom
The immobilizing components of centipede venoms
appear to fall into two functional categories: (i) nonpeptidic components and neurotransmitter-releasing
peptide neurotoxins producing a nearly immediate, transient effect; (ii) larger proteins acting via slower neurotoxic
and myotoxic pathways. Unlike arachnids, the initial rapid
paralytic response seems not to be caused by the action of
ion-channel neurotoxins, but rather an endogenous release
of histamines and neurotransmitters inducing immediate
hypotension, pain, and rigid muscle paralysis in insects.
This may then overlap with the onset of slower effects that
are caused by agonizing GPCRs, such as severe radiating
pain, rhabdomyolysis, and cardiac arrest. Considering the
prevalence of these neurotransmitters and GPCRs in the
animal kingdom, as well as the age of these neuronal
signaling mechanisms, this may represent an ancient
mechanism for venomous prey subduction (Hoyle, 2011).
This could have been retained due to the very generalist
diet of scolopendrid species, which includes a wide variety
of invertebrate and, in larger species, vertebrate prey. The
presence of a tetrodotoxin-sensitive NaV channel agonist or
Ca2þ pore forming toxin, however, indicates that non-GPCR
toxins are also present in centipede venoms. It will be
interesting to examine whether centipedes that prey
predominantly on arthropod prey, such as scutigeromorphs, possess the same apparently biphasic venom
function.
7. Conclusion
It appears that the centipede venom apparatus differs
substantially from that of other arthropods in both functional morphology and venom pharmacology, reflecting
the ancient divergence of centipedes. This may, however,
be an artifact of the paucity of information available on
centipede venoms, particularly with regard to the neurotoxic venom components. Indeed, when the taxonomic
distribution and coverage is considered, the large information void on the venom of this ecologically important
and publicly well-known group of animals becomes
evident. It will be interesting to follow the outcomes of
future studies, which hopefully will reveal more details
about the prevalence of neurotoxins in centipede venoms
as well as more information on the venom systems of nonscolopendromorph taxa.
522
E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
Acknowledgments
We acknowledge financial support from The University of Queensland (International Postgraduate Research
Scholarship, UQ Centennial Scholarship, and UQ Advantage Top-Up Scholarship to EABU). Many thanks to David
Morgenstern for sharing his knowledge on the structure
and function of arachnid venom glands and venom
components.
Ethical Statement
The authors declare that this manuscript complies with
the Elsevier Ethical Guidelines for Journal Publication and
acknowledge the Duties of Authors described therein.
Funding source
International Postgraduate Research Scholarship (IPRS),
University of Queensland Centennial Scholarship (UQCent),
University of Queensland Advantage Top-Up Scholarship
(UQAdv).
Conflict of Interest
The authors declare that there are no conflicts of interest.
Appendix. Supplementary data.
Supplementary data related to this article can be found
online at doi:10.1016/j.toxicon.2011.01.004.
References
Aird, S.D., 2002. Ophidian envenomation strategies and the role of
purines. Toxicon 40, 335–393.
Anderson, L.I., Trewin, N.H., 2003. An early Devonian arthropod fauna
from the Windyfield cherts, Aberdeenshire, Scotland. Palaeontology
46, 467–509.
Antoniazzi, M.M., Pedroso, C.M., Knysak, I., Martins, R., Guizze, S.P.G.,
Jared, C., Barbaro, K.C., 2009. Comparative morphological study of the
venom glands of the centipede Cryptops iheringi, Otostigmus pradoi
and Scolopendra viridicornis. Toxicon 53, 367–374.
Bücherl, W., 1946. Açâo do veneno dos Escolopendromorfos do Brasil
sobre alguns animais de laboratorio. Mem. Inst. Butantan 19, 181–197.
Baerg, W.J., 1924. The effect of the venom of some supposedly poisonous
arthropods. Ann. Entomol. Soc. Am. 17, 343–352.
Baerg, W.J., 1925. The effect of the venom of some supposedly poisonous
arthropods of the canal zone. Ann. Entomol. Soc. Am. 18, 471–478.
Balit, C.R., Harvey, M.S., Waldock, J.M., Isbister, G.K., 2004. Prospective
study of centipede bites in Australia. Clin. Toxicol 42, 41–48.
Bhagirath, T., Chingtham, B., Mohen, Y., 2006. Venom of a hill centipede
Scolopendra viridicornis inhibits growth of human breast tumor in
mice. Indian J. Pharmacol 38, 291.
Blay, E.R., 1955. Treatment of centipede bites. Br. Med. J. 2, 1619.
Bonato, L., Barber, A., Minelli, A., 2006. The European centipedes hitherto
referred to Eurygeophilus, Mesogeophilus, and Chalandea (Chilopoda,
Geophilomorpha): taxonomy, distribution, and geographical variation
in segment number. J. Nat. Hist. 40, 415–438.
Briot, A., 1904. Sur les venin des Scolopendres. C. R. Seances Soc. Biol. Fil
57, 476–477.
Bush, S.P., King, B.O., Noris, R.L., Stockwell, S.A., 2001. Centipede envenomation. Wilderness Environ. Med. 12, 93–99.
Calvete, J.J., Moreno-Murciano, M.P., Theakston, R.D.G., Kisiel, D.G.,
Marcinkiewicz, C., 2003. Snake venom disintegrins: novel dimeric
disintegrins and structural diversification by disulphide bond engineering. Biochem. J. 372, 725–734.
Carpenter, C.C., Gillingham, J.C., 1984. Giant centipede (Scolopendra
alternans) attacks marine toad (Bufo marinus). Caribb. J. Sci. 20, 71–72.
Chahl, L.A., Kirk, E.J., 1975. Toxins which produce pain. Pain 1, 3–49.
Chao, J.L., Chang, H.W., 2006. Variation of the poison duct in Chilopoda
centipedes from Taiwan. Nor. J. Entomol. 53, 139–151.
Chaou, C.-H., Chen, C.-K., Chen, J.-C., Chiu, T.-F., Lin, C.-C., 2009. Comparisons of ice packs, hot water immersion, and analgesia injection for
the treatment of centipede envenomations in Taiwan. Clin. Toxicol.
47, 659–662.
Clark, D.B., 1979. A centipede preying on a nestling Rice Rat (Oryzomys
bauri). J. Mammology 60, 654.
Coffin, S.W., 1919. Notes on a case of centipede bite. Lancet 193, 1117–1118.
Cohen, E., Quistad, G.B., 1998. Cytotoxic effects of arthropod venoms on
various cultured cells. Toxicon 36, 353–358.
Cornwall, J.W., 1916. Some centipedes and their venom. Indian J. Med. Res.
3, 541–557.
Crabill, R.E.J., 1953. Concerning a new genus, Dinocryptops, and the
nomenclatorial status of Otocryptops and Scolopocryptops (Chilopoda:
Scolopendromorpha: Cryptopidae). Entomol. News 64, 96.
Crabill, R.E.J., 1960. A new American genus of cryptopid centipede, with
an annotated key to the scolopendromorph genera from America
north of Mexico. Proc. United States Natl. Mus 111, 1–15.
Dass, C.M.S., Jangi, B.S., 1978. Ultrastructural organization of the poison
gland of the centipede Scolopendra morsitans Linn. Indian J. Exp. Biol.
16, 748–757.
Dhananjaya, B.L., D’Souza, C.J.M., 2010. The pharmacological role of
nucleotidases in snake venoms. Cell Biochem. Funct 28, 171–177.
Duboscq, O., 1898. Recherches sur les chilopodes. Arch. Zool. Exp. Gen 6,
481–650.
Dunlop, J.A., 2010. Geological history and phylogeny of Chelicerata.
Arthropod Struct. Dev. 39, 124–142.
Easterla, D.A., 1975. Giant desert centipede preys upon snake. Southwest.
Nat. 20, 411.
Edgecombe, G.D., Barrow, L., 2007. A new genus of scutigerid centipedes
(Chilopoda) from Western Australia, with new characters for morphological phylogenetics of Scutigeromorpha. Zootaxa 1409, 23–50.
Edgecombe, G.D., Giribet, G., 2004. Adding mitochondrial sequence data
(16S rRNA and cytochrome c oxidase subunit I) to the phylogeny of
centipedes (Myriapoda: Chilopoda): an analysis of morphology and
four molecular loci. J. Zoolog. Syst. Evol. Res. 42, 89–134.
Edgecombe, G.D., Giribet, G., 2007. Evolutionary biology of centipedes
(Myriapoda: Chilopoda). Annu. Rev. Entomol. 52, 151.
Edgecombe, G., Giribet, G., 2008. A New Zealand species of the transTasman centipede order Craterostigmomorpha (Arthropoda: Chilopoda) corroborated by molecular evidence. Invertebr. Syst 22, 1–15.
Edgecombe, G.D., Koch, M., 2008. Phylogeny of scolopendromorph
centipedes (Chilopoda): morphological analysis featuring characters
from the peristomatic area. Cladistics 24, 872–901.
Fletcher, J.E., Hubert, M., Wieland, S.J., Gong, Q.-H., Jiang, M.-S., 1996.
Similarities and differences in mechanisms of cardiotoxins, melittin
and other myotoxins. Toxicon 34, 1301–1311.
Foddai, D., Minelli, A., 1999. A troglomorphic geophilomorph centipede
from southern France (Chilopoda: Geophilomorpha: Geophilidae). J.
Nat. Hist. 33, 267–287.
Foddai, D., Bonato, L., Pereira, L.A., Minelli, A., 2003. Phylogeny and
systematics of the Arrupinae (Chilopoda: Geophilomorpha: Mecistocephalidae) with the description of a new dwarfed species. J. Nat.
Hist. 37, 1247–1267.
Forti, L.R., Fischer, H.Z., Encarnação, L.C., 2007. Treefrog Dendropsophus
elegans (Wied-Neuwied, 1824)(Anura: Hylidae) as a meal to Otostigmus tibialis Brölemann, 1902 (Chilopoda: Scolopendridae) in
the Tropical Rainforest in southeastern Brazil. Braz. J. Biol. 67,
583–584.
Friedman, I.S., Phelps, R.G., Baral, J., Sapadin, A.N., 1998. Wells’ syndrome
triggered by centipede bite. Int. J. Dermatol. 37, 602–605.
Fry, B.G., 2005. From genome to "venome": molecular origin and evolution
of the snake venom proteome inferred from phylogenetic analysis of
toxin sequences and related body proteins. Genome Res. 15, 403–420.
Girish, K.S., Shashidharamurthy, R., Nagaraju, S., Gowda, T.V., Kemparaju, K.,
2004. Isolation and characterization of hyaluronidase a "spreading
factor" from Indian cobra (Naja naja) venom. Biochimie 86, 193–202.
Gomes, A., Datta, A., Sarangi, B., Kar, P.K., Lahiri, S.C., 1982a. Occurrence of
histamine and histamine release by centipede venom. Indian J. Med.
Res. 76, 888–891.
Gomes, A., Datta, A., Sarangi, B., Kar, P.K., Lahiri, S.C., 1982b. Pharmacodynamics of venom of the centipede Scolopendra subspinipes dehaani
Brandt. Indian J. Exp. Biol. 20, 615–618.
Gomes, A., Datta, A., Sarangi, B., Kar, P.K., Lahiri, S.C., 1983. Isolation,
purification and pharmacodynamics of a toxin from the venom of the
E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
centipede Scolopendra subspinipes dehaani Brandt. Indian J. Exp. Biol.
21, 203–207.
González-Morales, L., Diego-GarcÌa, E., Segovia, L., Gutiérrez, M.d.C.,
Possani, L.D., 2009. Venom from the centipede Scolopendra viridis say:
purification, gene cloning and phylogenetic analysis of a phospholipase A2. Toxicon 54, 8–15.
Gutiérrez, J.M., Lomonte, B., 1995. Phospholipase A2 myotoxins from
Bothrops snake venoms. Toxicon 33, 1405–1424.
Gutiérrez, J.M., Rucavado, A., 2000. Snake venom metalloproteinases: their
role in the pathogenesis of local tissue damage. Biochimie 82, 841–850.
Gutiérrez, M.d.C., Abarca, C., Possani, L.D., 2003. A toxic fraction from
scolopendra venom increases the basal release of neurotransmitters
in the ventral ganglia of crustaceans. Comp. Biochem. Physiol. C
Toxicol. Pharmacol 135, 205–214.
Hamza, L., Girardi, T., Castelli, S., Gargioli, C., Cannata, S., Patamia, M.,
Luly, P., Laraba-Djebari, F., Petruzzelli, R., Rufini, S., 2010. Isolation and
characterization of a myotoxin from the venom of Macrovipera lebetina transmediterranea. Toxicon 56, 381–390.
Harada, K., Asa, K., Imachi, T., Yamaguchi, Y., Yoshida, K., 1999. Centipede
inflicted postmortem injury. J. Forensic Sci. 44, 849–850.
Hasan, S., Hassan, K., 2005. Proteinuria associated with centipede bite.
Pediatr. Nephrol. 20, 550–551.
Heitz, J.R., Norment, B.R., 1974. Characteristics of an alkaline phosphatase
activity in brown recluse venom. Toxicon 12, 181–187.
Hilken, G., Rosenberg, J., Brockmann, C., 2005. Ultrastructure of the epidermal
maxilla II-gland of Scutigera coleoptrata (Chilopoda, Notostigmophora)
and the ground pattern of epidermal gland organs in Myriapoda. J.
Morphol. 264, 53–61.
Hjelle, J.T., 1990. Anatomy and morphology. In: Polis, G.A. (Ed.),
The Biology of Scorpions. Stanford University Press, Stanford, pp.
54–56.
Hoffman, R.L., Pereira, L.A., 1991. Systematics and biogeography of Marsikomerus Attems, 1938, a misunderstood genus of centipedes (Geophilomorpha: Schendylidae). Insecta Mundi 5, 45–60.
Hoyle, C.H.V., 2011. Evolution of neuronal signalling: Transmitters and
receptors. Auton. Neurosci., in press, doi:10.1016/j.autneu.2010.05.007.
Jangi, B.S., Dass, C.M.S., 1977. Chemoreceptive function of the poison fang in
the centipede Scolopendra morsitans L. Indian J. Exp. Biol. 15, 803–804.
Jarrar, B.M., 2010. Morphology, histology and histochemistry of the
venom apparatus of the centipede, Scolopendra valida (Chilopoda,
Scolopendridae). Int. J. Morphol. 28, 19–25.
Kass, L., Batsakis, J.G., 1979. Cytochemistry of esterases. Crit. Rev. Clin. Lab.
Sci. 10, 205–223.
Knysak, I., Martins, R., Bertim, C.R., 1998. Epidemiological aspects of
centipede (Scolopendromorphae: Chilopoda) bites registered in
greater S. Paulo, SP, Brazil. Rev. Saude Publica 32, 514–518.
Koch, M., Pärschke, S., Edgecombe, G.D., 2009. Phylogenetic implications
of gizzard morphology in scolopendromorph centipedes (Chilopoda).
Zool. Scr. 38, 269–288.
Kuhn-Nentwig, L., 2003. Antimicrobial and cytolytic peptides of
venomous arthropods. Cell. Mol. Life Sci. 60, 2651–2668.
Kuhn-Nentwig, L., Schaller, J., Nentwig, W., 2004. Biochemistry, toxicology
and ecology of the venom of the spider Cupiennius salei (Ctenidae).
Toxicon 43, 543–553.
Langley, R.L., 2005. Animal-Related fatalities in the United States - An
Update. Wilderness Environ. Med 16, 67–74.
Langley, R.L., Morrow, W.E., 1997. Deaths resulting from animal attacks in
the United States. Wilderness Environ. Med 8, 8–16.
Lee, S., Lynch, K.R., 2005. Brown recluse spider (Loxosceles reclusa)
venom phospholipase D (PLD) generates lysophosphatidic acid (LPA).
Biochem. J. 391, 317–323.
Lewis, J.G.E., 1981. The Biology of Centipedes. Cambridge University Press,
Cambridge.
Lewis, J.G.E., 2002. The scolopendromorph centipedes of Mauritius and
Rodrigues and their adjacent islets (Chilopoda: Scolopendromorpha).
J. Nat. Hist. 36, 79–106.
Lewis, J.G.E., Daszak, P., Jones, C.G., Cottingham, J.D., Wenman, E.,
Maljkovic, A., 2010. Field observations on three scolopendrid centipedes from Mauritius and Rodrigues (Indian Ocean) (Chilopoda:
Scolopendromorpha). Int. J. Myriapodol 3, 123–137.
Lima, P.R.d., Brochetto-Braga, M.R., 2003. Hymenoptera venom review
focusing on Apis mellifera. J. Venom. Anim. Toxins Incl. Trop. Dis. 9,
149–162.
Lin, T.J., Yang, C.C., Yang, G.Y., Ger, J., Tsai, W.J., Deng, J.F., 1995. Features of
centipede bites in Taiwan. Trop. Geogr. Med. 47, 300.
Liu, T., Bai, Z.-T., Pang, X.-Y., Chai, Z.-F., Jiang, F., Ji, Y.-H., 2007. Degranulation of mast cells and histamine release involved in rat pain-related
behaviors and edema induced by scorpion Buthus martensi Karch
venom. Eur. J. Pharmacol. 575, 46–56.
523
Logan, J.L., Ogden, D.A., 1985. Rhabdomyolysis and acute renal failure
following the bite of the giant desert centipede Scolopendra heros.
West. J. Med. 142, 549.
Lomonte, B., Angulo, Y., Calderón, L., 2003. An overview of lysine-49
phospholipase A2 myotoxins from crotalid snake venoms and
their structural determinants of myotoxic action. Toxicon 42,
885–901.
Long-Rowe, K.O., Burnett, J.W., 1994. Characteristics of hyaluronidase and
hemolytic activity in fishing tentacle nematocyst venom of Chrysaora
quinquecirrha. Toxicon 32, 165–174.
Ménez, A., Zimmerman, K., Zimmerman, S., Heatwole, H., 1990. Venom
apparatus and toxicity of the centipede Ethmostigmus rubripes (Chilopoda, Scolopendridae). J. Morphol. 206, 303–312.
Ma, Y., Zhao, R., He, Y., Li, S., Liu, J., Wu, Y., Cao, Z., Li, W., 2009. Transcriptome analysis of the venom gland of the scorpion Scorpiops
jendeki: Implication for the evolution of the scorpion venom arsenal.
BMC Genomics 10, 290.
Malta, M.B., Lira, M.S., Soares, S.L., Rocha, G.C., Knysak, I., Martins, R.,
Guizze, S.P.G., Santoro, M.L., Barbaro, K.C., 2008. Toxic activities of
Brazilian centipede venoms. Toxicon 52, 255–263.
Marsh, F., 1957. Centipede bites. Br. Med. J. 2, 825.
Matsui, T., Fujimura, Y., Titani, K., 2000. Snake venom proteases affecting
hemostasis and thrombosis. Biochim. Biophys. Acta – Prot. Struct.
Mol. Enzymol 1477, 146–156.
McKeown, K.C., 1930. Centipedes and centipede bites. Aust. Mus. Mag 4,
59–60.
Mebs, D., Ehrenfeld, M., Samejima, Y., 1983. Local necrotizing effect of
snake venoms on skin and muscle: relationship to serum creatine
kinase. Toxicon 21, 393–404.
Medeiros, C.R., Susaki, T.T., Knysak, I., Cardoso, J.L.C., Málaque, C.M.S.,
Fan, H.W., Santoro, M.L., França, F.O.S., Barbaro, K.C., 2008. Epidemiologic and clinical survey of victims of centipede stings admitted to
Hospital Vital Brazil (São Paulo, Brazil). Toxicon 52, 606–610.
Minelli, A., 1978. Secretions of centipedes. In: Bettini, S. (Ed.), Handbook
of Experimental Pharmacology: Arthropod Venoms. Springer–Verlag,
Berlin.
Mohamed, A.H., Zaid, E., El-Beih, N.M., Abd El-Aal, A., 1980. Effects of an
extract from the centipede Scolopendra morsitans on intestine,
uterus and heart contractions and on blod glucose and liver and
muscle glycogen levels. Toxicon 18, 581–589.
Mohamed, A.H., Abu-Sinna, G., El-Shabaka, H.A., Abd El-Aal, A., 1983.
Proteins, lipids, lipoproteins and some enzyme characterizations of
the venom extract from the centipede Scolopendra morsitans. Toxicon
21, 371–377.
Mohri, S., Sugiyama, A., Saito, K., Nakajima, H., 1991. Centipede bites in
Japan. Cutis 47, 189.
Molinari, J., Gutierrez, E.E., De Ascencao, A.A., Nassar, J.M., Arends, A.,
Marquez, R.J., 2005. Predation by giant centipedes, Scolopendra
gigantea, on three species of bats in a Venezuelan cave. Caribb. J. Sci.
41, 340–346.
Mumcuoglu, K.Y., Leibovici, V., 1989. Centipede (Scolopendra) bite: a case
report. Isr. J. Med. Sci. 25, 47.
Murienne, J., Edgecombe, G.D., Giribet, G., 2010. Including secondary
structure, fossils and molecular dating in the centipede tree of life.
Mol. Phylogenet. Evol. 57, 301–313.
Nagaraju, S., Mahadeswaraswamy, Y.H., Girish, K.S., Kemparaju, K., 2006.
Venom from spiders of the genus Hippasa: biochemical and pharmacological studies. Comp. Biochem. Physiol. C Toxicol. Pharmacol
144, 1–9.
Nagaraju, S., Girish, K.S., Fox, J.W., Kemparaju, K., 2007. “Partitagin”
a hemorrhagic metalloprotease from Hippasa partita spider venom:
role in tissue necrosis. Biochimie 89, 1322–1331.
Nagpal, N., Kanwar, U., 1981. The poison gland in the centipede Otostigmus
ceylonicus; Morphology and cytochemistry. Toxicon 19, 898–902.
Noguchi, K., Herr, D., Mutoh, T., Chun, J., 2009. Lysophosphatidic acid
(LPA) and its receptors. Curr. Opin. Pharmacol. 9, 15–23.
Oguiura, N., Boni-Mitake, M., R$dis-Baptista, G., 2005. New view on
crotamine, a small basic polypeptide myotoxin from South American
rattlesnake venom. Toxicon 46, 363–370.
Ozsarac, M., Karcioglu, O., Ayrik, C., Somuncu, F., Gumrukcu, S., 2004.
Acute coronary ischemia following centipede envenomation: case
report and review of the literature. Wilderness Environ. Med 15,
109–112.
Peng, K., Kong, Y., Zhai, L., Wu, X., Jia, P., Liu, J., Yu, H., 2009. Two novel
antimicrobial peptides from centipede venoms. Toxicon 55,
274–279.
Pereira, L.A., Uliana, M., Minelli, A., 2007. Geophilomorph centipedes
(Chilopoda) from termite mounds in the northern Pantanal wetland
of Mato Grosso, Brazil. Stud. Neotrop. Fauna Environ. 42, 33–48.
524
E.A.B. Undheim, G.F. King / Toxicon 57 (2011) 512–524
Pineda, E.V., 1923. A fatal case of centipede bite. J. Philippine Is. Med.
Assoc. 3, 59–63.
Quistad, G.B., Dennis, P.A., Skinner, W.S., 1992. Insecticidal activity of
spider (Araneae), centipede (Chilopoda), scorpion (Scorpionidae), and
snake (Serpentes) venoms. J. Econ. Entomol. 85, 33–39.
Rates, B., Bemquerer, M.P., Richardson, M., Borges, M.H., Morales, R.A.V.,
De Lima, M.E., Pimenta, A., 2007. Venomic analyses of Scolopendra
viridicornis nigra and Scolopendra angulata (Centipede, Scolopendromorpha): shedding light on venoms from a neglected group.
Toxicon 49, 810–826.
Remington, C.L., 1950. The bite and habits of a giant centipede (Scolopendra
subspinipes) in the Philippine Islands. Am. J. Trop. Med. Hyg 1, 453.
Rodrigues, M.C.A., Guimarães, L.H.S., Liberato, J.L., Polizeli, M.d.L.T.d.M.,
dos Santos, W.F., 2006. Acid and alkaline phosphatase activities of
a fraction isolated from Parawixia bistriata spider venom. Toxicon 47,
854–858.
Rosenberg, J., Hilken, G., 2006. Fine structural organization of the poison
gland of Lithobius forficatus (Chilopoda, Lithobiomorpha). Nor. J.
Entomol. 53, 119–127.
Rouvin, B., Seck, M., N’diaye, M., Diatta, B., Saissy, J.M., 2008. Morsure de
scolopendre chez une drépanocytaire hétérozygote au Sénégal. Med.
Trop. 68, 647–648.
Said, E., 1958. On the peptidases and esterases in the venom of Buthus quinquestriatus and Scolopendra morsitans. Proc. Egypt. Acad. Sci. 13, 76–84.
Schaloske, R.H., Dennis, E.A., 2006. The phospholipase A2 superfamily and
its group numbering system. Biochim. Biophys. Acta Mol. Cell Biol.
Lipids 1761, 1246–1259.
Serinken, M., Erdur, B., Sener, S., Kabay, B., Cevik, A., 2005. A case of mortal
necrotizing fasciitis of the trunk resulting from a centipede (Scolopendra moritans) bssite. Internet J. Emerg. Med 2, 1–8.
Servent, D., Fruchart-Gaillard, C., 2009. Muscarinic toxins: tools for the
study of the pharmacological and functional properties of muscarinic
receptors. J. Neurochem., 1193–1202.
Shear, W.A., Edgecombe, G.D., 2009. The geological record and phylogeny
of the Myriapoda. Arthropod Struct. Dev. 39, 174–190.
Snodgrass, R.E., 1965. Arthropod Anatomy. Hafner Publishing Company,
New York. 193–224.
Stankiewicz, M., Hamon, A., Benkhalifa, R., Kadziela, W., Hue, B., Lucas, S.,
Mebs, D., Pelhate, M., 1999. Effects of a centipede venom fraction on
insect nervous system, a native Xenopus oocyte receptor and on an
expressed Drosophila muscarinic receptor. Toxicon 37, 1431–1445.
Stone, C.A., Wenger, H.C., Ludden, C.T., Stavorski, J.M., Ross, C.A., 1961.
Antiserotonin-antihistaminic properties of cyproheptadine. J. Pharmacol. Exp. Ther. 131, 73.
Sulkowski, E., Björk, W., Laskowski, M., 1963. A specific and nonspecific
alkaline monophosphatase in the venom of Bothrops atrox and their
occurrence in the purified venom phosphodiesterase. J. Biol. Chem.
238, 2477–2486.
Supakthanasiri, P., Ruxrungtham, K., Klaewsongkram, J., Chantaphakul, H.,
2004. Anaphylaxis to centipede bite. J. Allergy Clinical Immunology
113, S244.
Sutherland, S.K., Tibballs, J., 2001. Australian Animal Toxins: the Creatures,
Their Toxins and Care of the Poisoned Patient, second ed. Oxford
University Press, Melbourne.
Trevisan-Silva, D., Gremski, L.H., Chaim, O.M., da Silveira, R.B.,
Meissner, G.O., Mangili, O.C., Barbaro, K.C., Gremski, W., Veiga, S.S.,
Senff-Ribeiro, A., 2010. Astacin-like metalloproteases are a gene
family of toxins present in the venom of different species of the
brown spider (genus Loxosceles). Biochimie 92, 21–32.
Tu, A.T., Chua, A., 1966. Acid and alkaline phosphomonoesterase activities
in snake venoms. Comp. Biochem. Physiol. 17, 297–307.
Undheim, E.A.B., Georgieva, D.N., Thoen, H.H., Norman, J.A., Mork, J.,
Betzel, C., Fry, B.G., 2010. Venom on ice: first insights into Antarctic
octopus venoms. Toxicon 56, 897–913.
Usmanov, P.B., Nuritova, F.A., 1994. The anticoagulant action of phospholipase A from Eresus niger spider venom. Toxicon 32, 625–628.
Uzel, A.P., Steinmann, G., Bertino, R., Korsaga, A., 2009. Dermohypodermite bactérienne et phlegmon du membre supérieur par morsure
de scolopendre: à propos de deux cas. Chir. Main 28, 322–325.
Vassilevski, A., Kozlov, S., Grishin, E., 2009. Molecular diversity of spider
venom. Biochem. Mosc 74, 1505–1534.
Vazirianzadeh, B., Rahmanei, A.H., Moravvej, S.A., 2007. Two cases of
chilopoda (centipede) biting in human from Ahwaz, Iran. Pak. J. Med.
Sci. Q. 23, 956.
Veraldi, S., Chiaratti, A., Sica, L., 2010. Centipede bite: a case report. Arch.
Dermatol. 146, 807–808.
Welsh, J.H., Batty, C.S., 1963. 5-Hydroxytryptamine content of some
arthropod venoms and venom-containing parts. Toxicon 1, 165–170.
Wenhua, R., Shuangquan, Z., Daxiang, S., Kaiya, Z., Guang, Y., 2006.
Induction, purification and characterization of an antibacterial
peptide scolopendrin I from the venom of centipede Scolopendra
subspinipes mutilans. Indian J. Biochem. Biophys 43, 88–93.
Winquist, R.J., Siegl, P.K., Baskin, E.P., Bohn, D.L., Morgan, G., Wallace, A.A.,
1984. Calcium entry blocker activity of cyproheptadine in isolated
cardiovascular preparations. J. Pharmacol. Exp. Ther. 230, 103.
Xin, Y., Choo, Y.M., Hu, Z., Lee, K.S., Yoon, H.J., Cui, Z., Sohn, H.D., Jin, B.R.,
2009. Molecular cloning and characterization of a venom phospholipase A2 from the bumblebee Bombus ignitus. Comp. Biochem.
Physiol. B, Biochem. Mol. Biol. 154, 195–202.
Yildiz, A., Biçeroglu, S., Yakut, N., Bilir, C., Akdemir, R., Akilli, A., 2006.
Acute myocardial infarction in a young man caused by centipede
sting. Emerg. Med. J. 23, e30.
Zychar, B.C., Dale, C.S., Demarchi, D.S., Gonçalves, L.R.C., 2010. Contribution of metalloproteases, serine proteases and phospholipases A2 to
the inflammatory reaction induced by Bothrops jararaca crude venom
in mice. Toxicon 55, 227–234.