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Morphology, Histology and Serotonin Immunoreactivity on Salivary Glands of Stick Insect, Phobaeticus serratipes (Phasmida: Phasmatidae)


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The salivary gland plays a significant role in physiological processes in insects including food lubrication, extra-oral digestion and enzyme secretion. This study was conducted to describe the morphology and histology of salivary glands of stick insect, Phobaeticus serratipes (Phasmida: Phasmatidae). The observation on gross morphology of salivary glands was photographed by using DSLR Canon EOS 6D camera attached to a stereo microscope. The histological study of the salivary glands involves special staining procedures of periodic Schiff's acid reagent and Alcian blue method. The immunohistochemical study of the biogenic amine serotonin distribution was observed under fluorescence microscope Zeiss AxiocamMRm Apotome.2. Results showed that the salivary glands were the acinar type that consists of two cells, parietal cell and zymogenic cell. The serotonin immunoreactivity of the salivary glands which located on the nerve fibers and might act as a neurotransmitter.
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Volume 14, Number 1,March 2021
ISSN 1995-6673
Pages 11 - 15
Jordan Journal of Biological Sciences
Morphology, Histology and Serotonin Immunoreactivity on
Salivary Glands of Stick Insect, Phobaeticus serratipes
(Phasmida: Phasmatidae)
Wan Nurul ‘Ain, W.M.N and Nurul Wahida Othman*
Centre for Insect Systematics, Department of Biology and Biotechnology, Faculty of Science and Technology, Universiti Kebangsaan
Malaysia, 43600 Bangi, Selangor, Malaysia.
Received November 11, 2019; Revised Feb16, 2020; Accepted June 1, 2020
The salivary gland plays a significant role in physiological processes in insects including food lubrication, extra-oral
digestion and enzyme secretion. This study was conducted to describe the morphology and histology of salivary glands of
stick insect, Phobaeticus serratipes (Phasmida: Phasmatidae). The observation on gross morphology of salivary glands was
photographed by using DSLR Canon EOS 6D camera attached to a stereo microscope. The histological study of the salivary
glands involves special staining procedures of periodic Schiff’s acid reagent and Alcian blue method. The
immunohistochemical study of the biogenic amine serotonin distribution was observed under fluorescence microscope Zeiss
AxiocamMRm Apotome.2. Results showed that the salivary glands were the acinar type that consists of two cells, parietal
cell and zymogenic cell. The serotonin immunoreactivity of the salivary glands which located on the nerve fibers and might
act as a neurotransmitter.
Keywords: Morphology, histology, serotonin, Phobaeticus serratipes, salivary gland
* Corresponding author e-mail:
1. Introduction
Phasmatodea insects are herbivores that use their
bodies’ similarities to twigs, leaves, branches and lichen as
an advantage for camouflaging themselves with vegetation
(Bedford, 1978). Phasmatodea consists of 3000 species
that are divided into 3 Families and 500 Genera (Whiting
et al., 2003) including the largest insect, Phobaeticus
chani Bragg, with the length of the female can reach up to
567 mm (Hennemen and Conle, 2008). They belong to the
monophyletic group within the Orthopteroidea that is
similar with Orthoptera, Blattaria, Dermaptera,
Dictyoptera, Grylloblattodea and Mantophasmatodea
(Flook and Rowel, 1998). The earliest described stick
insects from West Malaysia were Marmessoidea rosea
(Fabricius) in 1793 and Heteropteryx dilatata (Parkinson)
in 1798 (Seow-Choen, 2005). To date, only five families
of stick insects were recorded in Malaysia. They are
Heteronemiidae, Phasmatidae, Aschiphasmatidae,
Bacillidae and Phylliidae.
The salivary gland of insects is the gland associated
with the nutrients intake where secretion is usually
involved in the digestion and lubrication of food (Ali,
1997). There are two main types of the salivary glands of
insects which are acinar as in the locust and cockroaches
and tubular as in blowfly and Lepidoptera (Ali, 1997). The
general function of salivary secretions is digestion but they
may perform additional functions in some insects. For
example, the saliva of locusts and cockroaches contains
digestive enzymes (Gardiner, 1972; Kendall, 1969), the
saliva of mosquitoes contains anticoagulants and irritants
(Gardiner, 1972; Ribeiro, 1992) and labial on Lepidoptera
can produce silk (Kafatos, 1968).
Salivation can be controlled either by direct nervous
innervation or via neurohormone. Biogenic amines are one
of the mediators involved in the control of insect
salivation. They also act as neurotransmitters,
neuromodulators or neurohormones in the nervous system
and various peripheral organs of vertebrates or
invertebrates (Baumann et al., 2003; Blenau and Baumann,
2001; Evans, 1980; Roeder, 1994). Serotonin, dopamine,
octopamine and tyrosine hydroxylase are some of the
biogenic amines. Salivary glands can be innervated from
several sources. Most insects have salivary nerve which
projects from suboesophageal ganglion but some species
like Periplaneta americana and Rhodnius prolixus are also
equipped with a salivary nerve that projects from
stomatogastric nervous system (Baptist, 1941; Davis,
1985). Dopamine and serotonin emulate an important role
in the control of salivary glands in most insects (Baines
and Tyrer, 1989; Berridge, 1970; House, 1973). Salivary
nerves of stick insects receive axonal projection from
salivary nerve 1 (SN1) and salivary nerve 2 (SN2) in the
subesophageal ganglion (Ali and Orchard, 1996).
Immunohistochemistry shows that dopamine presents in
SN1 whereas serotonin presents in SN2.
Not much research has been done on the salivary
glands of stick insects. In this study, we report on the
morphology and histology of the salivary gland of
© 2021 Jordan Journal of Biological Sciences. All rights reserved - Volume 14, Number 1
Phobaeticus serratipes. We present a detailed description
of the salivary gland using stereo microscope and light
microscope. We also present results of the location of
serotonin in the salivary gland. Stick insects are
hypothesized to have an acinar type of salivary glands
based on their feeding behaviour with chewing mouthpart.
Salivary glands of insects have at least two different types
of secretion cells such as parietal cells and zymogenic cells
(Beams and King, 1932). The serotonin might be present
on the nerve of the salivary gland as it is known as a
monoamine neurotransmitter.
2. Material and Methods
2.1. Samples preparation
Samples were collected from Langkawi Island, Kedah,
Fraser’s Hill, Pahang and Gunung Ledang, Johor. Overall,
30 fresh samples were used in this study.
2.2. Gross Morphology of Salivary Glands
All stick insects were weighed before dissection.
Solutions of methylene blue were injected at the jointed
segment between the legs and abdomen and also between
the head and thorax of the stick insects. Samples were left
for an hour at room temperature. Dissections were done in
phosphate-buffered saline (PBS) and images of salivary
glands in-situ and ex-situ were captured using a DSLR
Canon EOS 6D camera attached to a stereo microscope.
2.3. Tissue sections
The salivary glands were fixed in the formalin solution
for 2-4 hours. Then, the formalin solution was removed by
washing in 70% ethanol. Next, the glands were dehydrated
through a series of ethanol (50%, 90%, 100%) for an hour
each. Tissue was left in sub-Xylene for an hour, infiltrated
with wax (3x at 58ºC) and embedded. Tissue was
sectioned (3-5µm) using Leica RM2245 microtome. The
slides containing tissues were stained using Alcian blue
staining followed by periodic acid-Schiff's reagent (PAS).
Images of the stained sections were observed under the
light microscope (Zeiss Axio Scope) with iSolutionLite
2.4. Tissue sections immunofluorescence
The serotonin detection was performed in the tissue
sections and whole mounts salivary gland tissue (Wan
Nurul ‘Ain and Nurul Wahida, 2015). Briefly, slides of
tissue sections were rehydrated through a series of
solutions (xylene and ethanol (2x100%, 95%, 70%). Then,
the slides were further rehydrated with phosphate-buffered
saline (PBS) for 10 minutes. The excess wash buffer was
drained. Slides were partially dried after removal from
PBS except for the tissue sections. After that, 2 drops of
pre-blocking agent PBT (PBS of 50ml +0.2% bovine
serum albumin, 25ul +0.1% TritonX-100, 5ul) was used to
cover tissue sections and left for 20 minutes and then
tapped off and wiped away. Tissue sections were covered
with diluted primary antibody (anti serotonin) or negative
control (PBT). The primary antibody was diluted 1 in
1000(1µl in 1ml of PBT + 1% normal goat serum, 100ul)
(PBT+N). Slides were incubated overnight at 4ºC. Slides
were rinsed with PBS to wash off excess serum and
drained. Tissues were then covered with PBT plus normal
goat serum (PBT+N). Then, the tissues were incubated
overnight (4 °C) with secondary antibody conjugated to
Dylight (dilution 1:300). The antibody was washed with
PBT (30 min, 4x) before a complete inversion in PBS and
further washing with series of ethanol (100%, 95%, 70%,
50%) to rehydrate it. The slides were mounted in a mixture
of 50% glycerol and PBS and covered with a coverslip.
The slides were then dried out on a slide warmer overnight
and observed under a fluorescence microscope (Zeiss
AxiocamMRm Apotome.2) with ZenPro2012 software and
Olympus FSX100 microscope.
2.5. Whole mount tissue immunofluorescense
Fresh salivary glands were fixed in 4%
paraformaldehyde in PBS (18 hours at 4ºC). After washing
it with PBS, the tissue was permeabilized by exposing it to
methanol (5 min, 70% MeOH in PBS, 60min, 100%
MeOH and 5min, 70% MeOH in PBS) before washing
with PBS (5min, 2x). The tissues were incubated for
30min in 100 mL of PBT+N (PBT + 5% normal goat
serum) before being processed with 100 mL of diluted
(1:1000) primary antibody serotonin and incubated
overnight at 4ºC. The antibodies were washed off by
multiple rinses with PBS (5min, 3x) and PBT (45min, 2x).
Then, secondary antibody Dylight (1:300) was added to
each vial, and the tissues were incubated at 4ºC overnight.
The tissues were washed in PBT (5min, 3x) before a
complete inversion in PBT for 2 hours. The tissues were
cleared in a mixture of 50% glycerol and 50% PBS
overnight before mounting on slides. The control samples
were processed with similar procedure but with absence of
the primary antibody. The distribution of serotonin was
observed under fluorescence microscope (Zeiss Axiocam
MRm Apotome.2) with ZenPro2012 software and
Olympus FSX100 microscope.
3. Result and Discussion
3.1. General morphology of salivary glands
The salivary glands of Phoebaeticus serratipes consist
of cluster of small globular types of acini. This acinar type
of salivary glands had also been reported in other species
of stick insect such as Carausius morosus (Asimakopoulos
and Orchard, 1998) and other solid feeder insects such as
grasshopper, Gastrimargus musicus (Nurul-Wahida and
Cooper, 2014) and cockroach, Periplaneta americana (Just
and Walz, 1996). Based on the gross morphology of the
salivary glands that had been studied, the salivary glands
of P. serratipes are paired glands that can be found at both
sides of the lateral prothorax and extended to the
metathorax (Figure1).
© 2021 Jordan Journal of Biological Sciences. All rights reserved - Volume 14, Number 1 13
Figure 1. Image of in-situ (red arrows) along the prothorax to
The size and distribution of salivary glands are
different between the male and female stick insects due to
the difference in the size of their body. The size of female
is bigger than the male stick insect. The process of saliva
secretion occurred at the glands of the globular acinar. The
transparent and fine asinus duct canal acts as the connector
between all the acini. This canal has an outlet from each of
the acinus globules where it channels out the saliva from
the acinus to the lower part of the labium of the mouthpart.
The saliva will be collected in the collecting ducts before it
is secreted. The collecting ducts from both sides of
salivary glands will be fused at the head capsule and
opened to become a salivary cup at the labium (Kendall,
1969), thus forming the main duct (Figure 2). The saliva is
secreted from the main duct.
Figure 2. Ex-situ of acinar salivary glands of P. serratipes. md
(main duct); cd (collecting duct).
Based on ex-situ and in-situ observation, no secretory
gland was detected in P. serratipes. This secretory gland
can be found in other species such as Asceles glaber
(Dossey et al., 2012) and Oreophoetes peruana (Eisner et
al., 1997).
3.2. Histology of salivary glands
The acinar glands of P. serratipes consist of two types
of cells, parietal and zymogenic (Figure 3). Each acinus
cell is covered by a basal membrane on the outside. Basal
membrane mould acinus cell a round shape which later
forms acinus globule.
Figure 3. Tissue section of P. serratipes salivary gland. Tissue
was stained with Alcian blue and PAS reagent. A (Acini); Bm
(basal membrane); Pa (parietal cell); z (zymogenic cell).
Parietal cells are cone-shaped and have wide basal
connected with basal membrane cells. They are located
between zymogenic cells and extended towards the centre
of each acinus. The nucleus of the parietal cell is big and
oval-shaped. It is located at the centre of the cell. In
contrast, zymogenic cells have an irregular shape. The
basal of the cell is smaller compared to parietal cells. The
nucleus is small in size and present on the side of each
3.3. Serotonin distributions on salivary glands
The serotonin on the salivary glands of P. serratipes can
be seen clearly at the axons along the ducts to the acini
globules and the nerve fibers in the acini (Figure 4).
Figure 4. The serotonin distributions on the nerve fibers of P.
serratipes salivary glands (yellow arrows).
Moreover, for the cross-section of the tissues, the
serotonin was distributed on both cells in the salivary
glands, parietal cells and zymogen cells (Figure 5). This
result was supported by Nurul-Wahida and Cooper (2014)
who reported the presence of serotonin on both parietal
and zymogenic cells of yellow-winged grasshopper,
Gastrimargus musicus. Serotonin is absent on the salivary
gland of controlled stick insect (Figure 6).
The presence of serotonin will produce saliva with high
protein content (Just and Walz, 1996). Electrical
innervation towards the nerves or glands of the salivary
ducts that superfusion with dopamine and serotonin will
stimulate the secretion of saliva (Just and Walz, 1996).
© 2021 Jordan Journal of Biological Sciences. All rights reserved - Volume 14, Number 1
Liquid secretion rate is controlled by peripheral cell (p-
cell) at the base of each acinar globule and involved in the
transportation of water and electrolytes as in cockroaches,
Periplaneta americana (Kessel and Beams, 1963;
Sutherland and Chillseyzn, 1968). Central cell, also known
as c-cell, will react with serotonergic innervation and
supply the proteinaceous components to the saliva (Just
and Walz, 1994, 1996; Walz et al., 2006). The parietal
cells of stick insects have similar function and morphology
as the peripheral cells or p-cells of the cockroaches,
whereas the zymogenic cells are similar to the central cells
or c-cells.
Figures 5-6. (5) Serotonin distributions on the cross section of
salivary glands tissues of P. serratipes (yellow arrows). (6) No
serotonin-like immunoreactive process on salivary glands of
control stick insect.
4. Conclusion
It can be concluded that the serotonin in P. serratipes
plays a role as a neurotransmitter that is similarly
described in the Periplaneta americana (Ali, 1997; Ali and
Orchard, 1995) due to its presence on the nerve fibers of
the salivary glands. Besides, the distribution of serotonin
on both parietal and zymogenic cells suggests that the
serotonin also innervates the production of proteinaceous
and non-proteinaceous saliva for this species.
The authors would like to thank Universiti Kebangsaan
Malaysia and Ministry of Higher Education of Malaysia
(MOHE) for the facilities and grant provided (Young
Researcher Encouragement Grant GGPM 2013 089
and FRGS/1/2015/WAB13/UKM/02/01), Nazca Scientific
and Olympus from Universiti Putra Malaysia for the
fluorescence microscope facility. We also would like to
express our gratitude to Dr. Azman Sulaiman for helping
us with the samples.
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Conference Paper
Full-text available
This study was conducted to identify the morphology of digestive tract of stick insect, Phobaeticus serratipes (Phasmida: Phasmatidae) as well as its histological structure. The distribution of biogenic amine, serotonin on the structure was also determined. Samplings for this study were conducted in highland areas in Langkawi Island, Kedah, Fraser Hill, Pahang and Mount of Ledang, Johor. The in-situ and ex-situ observation on gross morphology of digestive tract was done using stereo microscope Stemi v10 that was connected with DSLR Canon EOS 6D camera. The histological study of the alimentary tract involves special staining procedures of Periodic Acid Schiff's reagent and Alcian blue method. Serotonin distribution was conducted using immunohistochemical method with Dylight as secondary antibody. The serotonin distribution was observed under fluorescence microscope Zeiss Axiocam MRm Apotome.2 with ZenPro 2012 software and also Olympus FSX100 microscope. Digestive tract morphology and histology showed that P. serratipes foregut is the longest section consists of the oral cavity, pharynx, esophagus, crop and proventriculus. Gastric caeca and ventriculus embodied the midgut area. Midgut having columnar epithelial cells as the major cells that serve its purposes in nutrient absorptions and enzymatic metabolisms. The hindgut was the shortest part of the digestive tract that consists of Malpighian tubules, ileum, colon, rectum and anus. The serotonin of digestive tract were scattered and disorganized and mostly found on the muscle cells.
Full-text available
The family Phasmatidae Gray, 1835 is reviewed and the subfamily Phasmatinae shown to be polyphyletic. Based on features of the exosceleton of the insects, egg-morphology and copulation habits a new arrangement of Phasmatidae is proposed. The monophyly of Lanceocercata Bradler, 2001 is confirmed but this name shown to be a synonym of Phasmatidae, hence Lanceocercata is here referred to as Phasmatidae sensu stricto. Six subfamilies belong in Phasmatidae sensu stricto all of which share several common and supposedly apomorphic characters: Phasmatinae, Tropidoderinae, Extatosomatinae (stat. nov.), Xeroderinae, Pachymorphinae and “Platycraninae”. The other two subfamilies contained in Phasmatidae sensu Bradley & Galil, 1977 (Eurycanthinae and Cladomorphinae) are not cosely related and here regarded as subfamilies of Phasmatidae sensu lato. The subfamily Phasmatinae sensu Bradley & Galil, 1977 is shown to be polyphyletic. The two tribes Pharnaciini and Clitumnini (= Baculini Günther, 1953) are removed from Phasmatinae and shown to be closely related to each other. They are transferred to the here established subfamily Clitumninae, a subordinate clade of Phasmatidae sensu lato. The subfamily Lonchodinae is closely related to Clitumninae, hence removed from Diapheromeridae and transferred to Phasmatidae sensu lato. The tribes Achriopterini and Stephanacridini (formerly in Phasmatinae) are shown to be not closely related to either Phasmatinae sensu stricto, Clitumninae or Lonchodinae, and provisionally must be treated as tribes of Phasmatidae sensu lato (incerte sedis). A re-arrangement of Phasmatidae sensu stricto is proposed along with determinating keys to all subfamilies and their tribes. The subfamilies Phasmatinae, Tropidoderinae and Extatosomatinae stat. nov. are re-described and discussed in detail. Full lists of genera are provided for each tribe. Only three of seven tribes formerly in Phasmatinae remain in the subfamily, this is Phasmatini, Acanthomimini and Acanthoxylini. The subfamily Tropidoderinae contains three tribes: Tropidoderini, Monandropterini and Gigantophasmatini trib. nov. The tribe Extatosomatini Clark-Sellick, 1997 is removed from Tropidoderinae and raised to subfamily level (Extatosomatinae stat. nov.). Several genera are transferred to other tribes or subfamilies. Didymuria Kirby, 1904 is removed from Tropidoderini, since it differs by having a closed internal micropylar plate in the eggs (open in all Tropidoderini). It here remains as a genus incerte sedis of Tropidoderinae and its systematic position clearly deserves further clarification. Gigantophasma Sharp, 1898 from the Loyalty Islands is removed from Pharnaciini, and becomes the type genus of the tribe Gigantophasmatini trib. nov.. Anophelepis Westwood, 1859 is removed from “Platycraninae” and shown to belong in Phasmatinae: Acanthomimini. The two Australian genera Arphax Stål, 1875, and Vasilissa Kirby, 1896 are removed from Acanthoxylini and provisionally transferred to Acanthomimini, but their position remains as yet debatable. Echetlus Stål, 1875 is misplaced in “Platycraninae” and shown to be a likely member of Phasmatinae. The two Brazilian species Echetlus evoneobertii Zompro & Adis, 2001 and Echetlus fulgens Zompro, 2004b are obviously misplaced and belong in the New World Diapheromeridae: Diapheromerinae: Diapheromerini. The subfamily Pachymorphinae is briefly discussed and considered polyphyletic. Three genera of Pachymorphinae: Gratidiini Bragg, 1995 (Parapachymorpha Brunner v. Wattenwyl, 1893 and Cnipsomorpha Hennemann et al., 2008) are transferred to Clitumninae: Medaurini trib. nov. The genus Gongylopus Brunner v. Wattenwyl, 1907 is transferred from Pachymorphinae: Gratidiini to Clitumninae: Clitumnini. The subfamily Xeroderinae is briefly discussed and shown likely to be polyphyletic, due to it contains two fundamentally different types of genitalia in the males. Only the genera Xeroderus Gray, 1835 and perhaps Epicharmus Stål, 1875 clearly belong in Phasmatidae sensu stricto. Both, the Pachymorphinae and Xeroderinae certainly deserve more detailed investigation to clarify their systematic positions with confirmation. Two generic groups are recognized within Clitumnini (subfamily Clitumninae). Due to differing by genital features and egg-morphology Medaura Stål, 1875 and Medauroidea Zompro, 2000 are removed from Clitumnini and transferred to the newly described Medaurini trib. nov.. The new tribe furthermore contains two genera formerly included in Pachymorphinae: Gratidiini and transferred here, Cnipsomorpha Hennemann et al., 2008 and Parapachymorpha Brunner v. Wattenwyl, 1893. Phryganistria Stål, 1875 is removed from Clitumnini and transferred to Pharnaciini. Nesiophasma Günther, 1934 is shown to belong in the tribe Stephanacridini. The Australasian subfamily Lonchodinae Brunner v. Wattenwyl, 1893 has formerly been included in Diapheromeridae Zompro, 2001 (= Heteronemiidae by Bradley & Galil, 1977). However, numerous features of the genitalia and egg morphology show close relation to the Oriental subfamily Clitumninae instead. Thus, Lonchodinae is here transferred to the family Phasmatidae (sensu lato). Within Lonchodinae the new tribe Neohiraseini trib. nov. is recognized and contains the five genera formerly placed in the “Neohirasea-complex” of that subfamily, namely Andropromachus Carl, 1913, Neohirasea Rehn, 1904, Pseudocentema Chen, He & Li, 2002, Qiongphasma Chen, He & Li, 2002 and Spinohirasea Zompro, 2001. It differs from all other Lonchodinae (= tribe Lonchodini) by the well developed vomer of males and the lack of a capitulum in the eggs. The genus Cladomimus Carl, 1915 was previously misplaced in Clitumninae: Pharnaciini and is here transferred to Lonchodinae: Lonchodini. It appears to be close to the Australian Hyrtacus Stål, 1875. Leprocaulinus Uvarov, 1940 and Phenacocephalus Werner, 1930 are removed from the subfamily Necrosciinae and transferred to Lonchodinae: Lonchodini. Extensive research on the genera which belong to the tribe Pharnaciini Günther, 1953 and taking features of the genital exosceleton and egg-morphology into account, has shown this tribe to be polyphyletic. Based on such features two generic groups are easily recognized within Pharnaciini sensu Günther, 1953. Males of the first group have a longitudinally split anal segment, which consists of two separate, more or less elongate semi-tergites and forms a clasping apparatus, the vomer is strongly reduced or lacking, the profemora have a prominent, lamellate medioventral carina which is strongly displaced towards the anteroventral carina and the eggs have an open internal micropylar plate with a clear median line. Only the genera falling into this group remain in Pharnaciini. Males of the second group in contrast have an anal segment which is not split, but possess a clearly visible, well sclerotised, triangular or hook-like external vomer, an indistinct medioventral carina on the profemora and eggs with a closed internal micropylar plate. Most of the genera which fall into the second group are here transferred to the tribe Stephanacridini Günther, 1953, this is Diagoras Stål, 1877b, Eucarcharus Brunner v. Wattenwyl, 1907, Phasmotaenia Návas, 1907 and Sadyattes Stål, 1875. A detailed discussion of the differences between Pharnaciini and Stephanacridini is provided along with distinguishing keys, illustrations and maps showing the distinct geographic distributions. The five genera that belong in Pharnaciini are: Baculonistria gen. nov., Pharnacia Stål, 1877a, Phobaeticus Brunner v. Wattenwyl, 1907 (= Baculolonga Hennemann & Conle, 1997a, = Lobophasma Günther, 1934b syn. nov. , = Nearchus Redtenbacher, 1908 syn. nov. ), Tirachoidea Brunner v. Wattenwyl, 1893 stat. rev. and Phryganistria Stål, 1875. Pharnacia annulata Redtenbacher, 1908 and Pharnacia enganensis Redtenbacher, 1908 were misplaced in Pharnacia Stål, 1877 (tribe Pharnaciini) and are transferred to the genus Sadyattes Stål, 1875 (tribe Stephanacridini, comb. nov.). Phobaeticus kuehni Brunner v. Wattenwyl, 1907 is removed from Phobaeticus Brunner v. Wattenwyl, 1907 (Phasmatinae: Pharnaciini) and shown to belong in Nesiophasma Günther, 1934c (tribe Stephanacridini, comb. nov.). Phobaeticus incertus Brunner v. Wattenwyl, 1907 (= Nearchus grubaueri Redtenbacher, 1908 syn. nov.) is unlikely to belong in Pharnaciini and here only retained in the original genus Phobaeticus Brunner v. Wattenwyl, 1907 with doubt, it may belong in Nesiophasma Günther, 1934c (tribe Stephanacridini). Based on a total of almost 700 examined specimens, the Oriental tribe Pharnaciini Günther, 1953 is revised at the species level. The new genus Baculonistria gen. nov. (Type species Baculonistria alba (Chen & He, 1990) comb. nov.), is described to contain three species from Central and Eastern China. Tirachoidea Brunner v. Wattenwyl, 1893 was erroneously synonymised with Pharnacia Stål, 1877 and is here re-established as a valid genus (stat. rev.). All five genera are re-diagnosed and differentiated, their systematic position within Pharnaciini discussed, and complete synonymic and species-listings as well as distribution maps and determination keys to the insects and eggs are provided. Detailed descriptions, diagnoses, synonymic listings, illustrations, material listings, distribution maps and measurements are provided for all 40 valid species. The type material of a further two species appears to be lost. Seven new species are described: Pharnacia borneensis spec. nov. from Borneo; Pharnacia palawanica spec. nov. from Palawan, Phobaeticus mucrospinosus spec. nov. from Sumatra, Phobaeticus palawanensis spec. nov. from Palawan, Tirachoidea herberti spec. nov. from Borneo, Tirachoidea siamensis spec. nov. from Thailand and S-Vietnam and Phobaeticus chani Bragg spec. nov. from Borneo. Phobaeticus chani Bragg spec. nov. is the world’s longest known insect with a maximum body length of 357 mm and an overall length of 567 mm in the female. Twelve new synonymies were discovered: Bactridium grande Rehn, 1920 = Phobaeticus serratipes (Gray, 1835) syn. nov.; Pharnacia rigida Redtenbacher, 1908 = Phobaeticus sumatranus Brunner v. Wattenwyl, 1907, syn. nov.; Clitumnus irregularis Brunner v. Wattenwyl, 1907 = Phibalosoma tirachus Westwood, 1859, syn. nov.; Pharnacia magdiwang Lit & Eusebio, 2008 = Pharnacia ponderosa Stål, 1877 syn. nov.; Pharnacia spectabilis Redtenbacher, 1908 = Phibalosoma hypharpax Westwood, 1859, syn. nov.; Pharnacia semilunaris Redtenbacher, 1908 = Eucarcharus inversus Brunner v. Wattenwyl, 1907, syn. nov.; Pharnacia chiniensis Seow-Choen, 1998c = Pharnacia biceps Redtenbacher, 1908, syn. nov.; Nearchus grubaueri Redtenbacher, 1908 = Phobaeticus incertus Brunner v. Wattenwyl, 1907, syn. nov.; Phibalosoma maximum Bates, 1865 = Cladoxerus serratipes Gray, 1835, syn. nov.; Phobaeticus lambirica Seow-Choen, 1998a = Eucarcharus rex Günther, 1928, syn. nov.; Phobaeticus sichuanensis Cai & Liu, 1993 = Baculum album Chen & He, 1990, syn. nov. and Phobaeticus beccarianus Brunner v. Wattenwyl, 1907 is shown to represent the previously unknown female of Phobaeticus sobrinus Brunner v. Wattenwyl, 1907 (syn. nov.) Lectotypes are designated for: Nearchus redtenbacheri Dohrn, 1910, Pharnacia biceps Redtenbacher, 1908, Pharnacia ingens Redtenbacher, 1908, Pharnacia heros Redtenbacher, 1908, Phibalosoma westwoodi Wood-Mason, 1875, Phobaeticus sinetyi Brunner v. Wattenwyl, 1907, and Phobaeticus sumatranus Brunner v. Wattenwyl, 1907. A neotype is designated for Nearchus maximus Redtenbacher, 1908 and Phobaeticus magnus nom. nov. introduced as a replacement name for Nearchus maximus Redtenbacher, which is a junior homonym of Phibalosoma maximum Bates, 1865. The previously unknown males of Pharnacia heros Redtenbacher, 1908, Phobaeticus ingens (Redtenbacher, 1908), Tirachoidea jianfenglingensis (Bi, 1994), Pharnacia sumatrana (Brunner v. Wattenwyl, 1907), Phryganistria fruhstorferi (Brunner v. Wattenwyl, 1907) and Tirachoidea westwoodii (Wood-Mason, 1875) as well as the females of Pharnacia ponderosa Stål, 1877a and Pharnacia tirachus (Westwood, 1859) are described and illustrated for the first time. A brief description on the basis of colour photos of the so far unknown male of Pharnacia kalag Zompro, 2005 are presented. Detailed descriptions and illustrations are provided for the eggs of 24 species. The eggs of the following 18 species are described and illustrated for the first time: Pharnacia borneensis spec. nov., Pharnacia palawanica spec. nov., Pharnacia ponderosa Stål, 1877a, Pharnacia sumatrana (Brunner v. Wattenwyl, 1907), Pharnacia tirachus (Westwood, 1859), Phobaeticus hypharpax (Westwood, 1859), Phobaeticus chani Bragg spec. nov., Phobaeticus incertus Brunner v. Wattenwyl, 1907, Phobaeticus magnus nom. nov., Phobaeticus philippinicus (Hennemann & Conle, 1997a), Phobaeticus sinetyi Brunner v. Wattenwyl, 1907, Phryganistria grandis Rehn, 1906, Phryganistria virgea (Westwood, 1848), Tirachoidea biceps (Redtenbacher, 1908), Tirachoidea herberti spec. nov., Tirachoidea jianfenglingensis (Bi, 1994) and Tirachoidea siamensis spec. nov.. Several species were originally placed in or subsequently transferred into wrong genera by various authors. Consequently, numerous taxa are here transferred or re-transferred to other genera, which results in 22 new or revised combinations or status of genera and species (comb. nov. / stat. rev. / stat. nov.). A list of the taxonomic changes made in this revision is provided in the summary (see 9.2), which in all lists 70 nomenclatural changes.
Immunohistochemistry in Carausius morosus reveals that the salivary glands are innervated by two paired neurons, the SN1 and SN2, located in the suboesophageal ganglion. The SN1 stain for tyrosine hydroxylase-like immunoreactivity, while the SN2 stain for serotonin-like immunoreactivity. Tyrosine hydroxylase is the rate limiting enzyme in the formation of catecholamines, and immunoreactivity to it is indicative of dopamine presence in insects (Evens 1980, and Ali and Orchard 1996). Both neurons project axons through the salivary nerve, and branch over the acini. Immunohistochemistry and biocytin filling shows that the salivary ducts are targets for branches from the salivary nerve. Dopamine and serotonin cause a dose-dependant increase in cyclic AMP levels in the salivary glands, indicating cyclic AMP may play a role as a second messenger. Increases in cyclic AMP induced by dopamine and serotonin, can be inhibited by vertebrate dopaminergic and serotonergic receptor antagonists respectivly. The rank order of potency of dopaminergic antagonists (based on IC(50) values) of SCH-23390 > flupenthixol > chlorpromazine > butaclamol, suggests the presence of receptors similar to vertebrate D(1)-like receptors. The rank order of potency of serotonergic receptor antagonists of spiperone > ketanserin > mianserin > cyproheptadine, suggests the presence of receptors similar to vertebrate 5HT(2) receptors. Electrical stimulation of the salivary nerve also elevates cyclic AMP levels in the salivary glands, an elevation that can be partially inhibited by 0.1 mM SCH-23390 and cyproheptadine.
Yellow-winged grasshoppers (Gastrimargus musicus) were captured in the field to examine the morphology and amine immunohistochemistry of their salivary glands. Fifty-eight grasshoppers were collected, with only five being males. Eight of 53 female grasshoppers had food in their crop, and the salivary glands of those insects were significantly heavier than those of grasshoppers without food in their crops. The salivary gland of the yellow-winged grasshopper was an acinar-type gland, similar to gland descriptions for other Orthoptera. The primary secretory part of acini of each gland is composed of zymogen and parietal cells. Staining patterns indicated that serotonin and dopamine could act as neurotransmitters and/or neurohormones to stimulate the glands. The pattern of staining of serotonin in the salivary gland suggested that serotonin stimulates both zymogen and parietal cells. Only the parietal cells were positively stained with dopamine. Comparing staining of glands of grasshoppers with food in their crop with the glands of grasshoppers with empty crops suggested a reduction in staining for serotonin in the latter. The differential staining pattern suggests that these amines have different roles in the salivary gland of G. musicus. The lack of difference in structure but increased mass with feeding suggests that all glands were active, but that secretion was actively occurring only in animals with the heavier glands.
The salivary glands of the locust Schistocerca gregaria are influenced by at least two nerves. The suboesophageal salivary nerve (nerve 7b) is excitatory eliciting copious secretion when active. The prothoracic posterior transverse nerve is also capable of evoking increases in secretion, but only if the innervation from the salivary nerve is present. This is, in part, because activity in the transverse nerve influences the firing of the two suboesophageal salivary neurones that have their axons in the salivary nerve. The effect of the salivary nerve is mimicked by both 5-HT and dopamine, whereas the action of the transverse nerve on the glands is mimicked by the peptides YGGFMRFamide and YGGFLRFamide.
The labial gland of adult saturniids produces a voluminous liquid at about the time of ecdysis, in response to either confinement of the moth or physostigmine injection. This liquid normally facilitates escape from the cocoon. During secretion the gland accumulates K+ and HCOS−from the blood, and Rb+ if it is available, but excludes rather efficiently Na+, Li+, Sr2+, Ca2+, Mg2+, phosphate and other small molecular weight components of blood. Cl− is approximately equally distributed in blood and secretion. The pH of the secretion is approximately 8·5, as compared with 6·5 for blood. The secretion is in osmotic equilibrium with blood, even after alteration of blood osmotic pressure by injection of hypertonic or hypotonic solutions. For both Cl− and Na+ the rate of entry into the secretion is proportional to concentration in blood; i.e. the efficiency of exclusion is characteristic for each ion, within a wide range of blood concentrations. Secretion is accompanied by a potential difference across the secretory cell (ca. + 25 mV. lumen positive). Secretion is abolished by dinitrophenol, but not by ouabain or acetazoleimide. The results are best explained in terms of a respiration-mediated active accumulation of blood K+, in exchange for H+, in the basal region of the cells; this is presumably followed by release of the K+ into the lumen, with concomitant reabsorption of H+ from the secretion. The alkaline secretion then traps blood CO2 by converting it to HCO3−. Water follows osmotically, and other ions enter passively, at a rate determined by the corresponding permeability of the cells.