|
| |
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online May 2, 2008
Journal of Experimental Biology 211, 1645-1656 (2008)
Published by The Company of Biologists 2008
doi: 10.1242/jeb.014472
Synaptic transmission in neurons that express the Drosophila atypical soluble guanylyl cyclases, Gyc-89Da and Gyc-89Db, is necessary for the successful completion of larval and adult ecdysis
Department of Integrative Biosciences, Oregon Health and Science University, 611 SW Campus Drive, Portland, OR 97239, USA
* Author for correspondence (e-mail: mortonda{at}ohsu.edu)
Accepted 14 March 2008
 |
Summary |
|---|
 TOP Summary INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
|
INTRODUCTION |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
; Morton and Hudson,
2002). The synthesis of cGMP is catalyzed by guanylyl cyclases
that can be classified as either receptor guanylyl cyclases, which are
integral membrane proteins, or soluble guanylyl cyclases, which are localized
to the cytoplasm (Lucas et al.,
2000). Soluble guanylyl cyclases can be further subdivided into
conventional soluble guanylyl cyclases, which are the targets of the gaseous
messenger nitric oxide (NO) (Lucas et al.,
2000), and the atypical soluble guanylyl cyclases that are poorly
activated by NO (Morton,
2004a) and have been proposed to act as oxygen sensors
(Gray et al., 2004). The
Drosophila genome contains five genes that code for subunits of
soluble guanylyl cyclases (sGCs) (Morton,
2004a). Two of these, Gyc
-99B and
Gycβ-100B, form the conventional NO-sensitive sGC and
the other three genes code for atypical subunits that form oxygen-sensitive
sGCs (Morton, 2004b). We have
previously shown that the atypical sGC subunits, Gyc-88E, Gyc-89Da
and Gyc-89Db, are expressed in a subset of central and chemosensory
peripheral neurons (Langlais et al.,
2004; Morton et al.,
2005a). Based on the expression pattern and biochemical properties
of these subunits, we predicted that the sGCs formed from the atypical
subunits would be involved in olfaction, gustation and/or oxygen sensation
(Vermehren et al., 2006).
In this study we began to examine the function of the neurons that express
the atypical sGCs Gyc-89Da and Gyc-89Db. Using the predicted promoter regions
of these genes we generated flies that express the yeast transcription factor
GAL4 (Brand and Perrimon, 1993)
under the control of the promoters for each GC subunit. We then used these
flies to express the light chain of tetanus toxin (TNT) under the control of
an upstream activation sequence (UAS) that is regulated by GAL4 to block
synaptic transmission in the cells that express Gyc-89Da and Gyc-89Db. The
results show that at least some of these neurons are required for the
successful initiation and completion of larval and adult ecdysis.
|
MATERIALS AND METHODS |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
) unless
otherwise indicated.
Cloning Gyc-89Da and Gyc-89Db promoter regions
The Gyc-89Da promoter (p89Da) lines were generated by cloning the
1.9 kb sequence upstream of the translational start site of Gyc-89Da
that included the 5' UTR and first intron. The 1931 bp PCR fragment was
amplified from genomic DNA using the primers
5'-GGGTTCTTTTTTGTAGAGAAA-3' and
5'-GGTTTATGTTTCTGAAAAAATG-3' and Taq DNA polymerase
(Invitrogen, Carlsbad, CA, USA). The PCR fragment was gel purified and
subcloned directly into the Topo-TA vector (Invitrogen). The Gyc-89Da
promoter fragment was then excised with BamHI and NotI
restriction enzymes (Invitrogen) and subcloned into the P-element vectors
pPTGAL (Sharma et al., 2002)
and green fluorescent protein (GFP) pGreen Pelican
(Barolo et al., 2000). The
Gyc-89Db promoter (p89Db) lines were generated in a similar manner by
cloning the 3398 bp sequence 5' of Gyc-89Db, using the
primers 5'-GCACCTGTGGCTCTCTTA-3' and
5'-GATGGGGCAGGATGTGAA-3'. The PCR fragment was then subcloned into
the pPTGAL and GFP vectors. DNA for microinjection was prepared by
ultracentrifugation using a CsCl gradient. About 200–400
wy1;; embryos were co-injected with 0.2 µg
µl–1 of helper plasmid (
25.7 
2–3 wc)
and 1 µg µl–1 of either the GFP or GAL4 constructs in
0.85 mmol l–1 NaH2PO4, 9.15 mmol
l–1 Na2HPO4, 5 mmol
l–1 KCl buffer, and maintained at 18°C until eclosion.
Eleven insertion lines were recovered for p89Da-GAL4, two for
p89Da-GFP, four for p89Db-GAL4 and two for
p89Db-GFP. Chromosomal locations of the P-element insertions were
determined by standard crossing methods to a ubiquitous balancer line
(w;CyO/KrIf-1;Tm6B,Tb+/D1).
In situ hybridization
In situ hybridization of embryos using digoxygenin-labeled probes
was carried out as previously described
(Langlais et al., 2004).
Confocal microscopy
Tissue or whole larvae were mounted in aqueous mounting media (gel mount,
Sigma, St Louis, MO, USA) and viewed on a Nikon Eclipse E800 fluorescence
microscope equipped with argon and helium/neon lasers (Radiance 2100, Bio-Rad,
Hercules, CA, USA) to excite the GFP and dsRed proteins (emission filters
500–530 nm and 570–650 nm, respectively). Usually, a stack of 15
sections was taken (z-axis step size, 2.5 µm) using Laser Sharp
2000 software (Bio-Rad) and image processing was carried out using ImageJ
(http://rsb.info.nih.gov/ij/).
Identification of neurons expressing GFP was made based on a combination of
their morphologies and their relative position.
Mortality/viability analysis
The mortality of developing animals expressing the active or inactive forms
of TNT was determined as follows. To determine the hatching success rate,
flies were allowed to lay eggs on standard fruit plates
(Sullivan et al., 2000)
overnight at 25°C. Embryos were collected and arranged in rows on a new
fruit plate and allowed to hatch for 36 h at 25°C. Hatched and un-hatched
embryos were then counted. To examine the survival of first instar larvae to
the end of the third instar stage and into the pupal stage, a known number of
first instar larvae were placed in a standard food vial or on a custom-made
food disc in a plastic 8.5 cm Petri dish, with a few granules of dried yeast.
The food disc was prepared by melting standard fly food, pouring it onto a
large glass plate and compressing it between a second glass plate separated by
3 mm spacers. When solidified, 5.6 cm diameter discs were then cut from this
sheet. Larvae placed in vials or on discs were kept at 25°C until
pupariation and counted. The developmental stage and location of any dead
larvae were recorded. To examine defects during eclosion, developing adults
were collected from vials at stage P15 and lined up on a strip of double-sided
clear tape on a glass slide. Just prior to eclosion, the operculum was
manually removed from each pupa, and the pre-eclosion staging markers
(Kimura and Truman, 1990;
McNabb et al., 1997) were
observed through a dissecting microscope. At least 15 experimental and control
animals from three separate crosses were examined for pre-eclosion
defects.
GAL80ts experiments
The temperature-sensitive inhibitor of GAL4, GAL80ts
(Zeidler et al., 2004), was
used in some experiments to control temporal expression of TNT. Using standard
crossing schemes (Greenspan,
1997), a homozygous fly line was created that contained GAL80ts
driven by the tubulin promoter (tubP-GAL80ts) on chromosome 2 and the
p89Da-GAL4 insertion on chromosome 3. These flies were then crossed
with UAS-TNTa, UAS-TNTi or UAS-dsRed flies, resulting in offspring that
contained tubP-GAL4ts, p89Da-GAL4 and UAS-TNTa, UAS-TNTi or
UAS-dsRed. These crosses were carried out at 18°C, the permissive
temperature for GAL80ts. At 18°C, GAL80ts inhibits the action of GAL4.
Experimental vials, as described in Results, were switched at specific stages
to an incubator set at 30°C, the non-permissive temperature for GAL80ts
(Zeidler et al., 2004) that
allows GAL4 to function and activate transcription of UAS-controlled
transgenes.
|
RESULTS |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
;
Morton et al., 2005a). By
examining the number and position of these cells we concluded that these three
genes were co-expressed in many of the peripheral sensory neurons
(Langlais et al., 2004;
Morton et al., 2005a). To
better define these co-expression patterns, we used upstream regions of each
gene to drive expression of GFP in transgenic flies (designated
p89Da-GFP and p89Db-GFP). The portions of genomic DNA
5' to Gyc-89Da and Gyc-89Db (1.9 and 3.4 kb,
respectively) were sufficient to drive GFP expression in embryos in a pattern
that closely resembled that seen with in situ hybridization
(Fig. 1). By contrast, however,
a 5 kb length of genomic DNA 5' to Gyc-88E was insufficient to
drive expression of GFP in any cells. Peripheral neurons that expressed GFP in
p89Da-GFP and p89Db-GFP larvae included a small subset of cells in the dorsal
and terminal ganglia (Fig.
1G–I), which innervate the olfactory and gustatory organs,
respectively (Stocker, 1994;
Heimbeck et al., 1999),
neurons innervating external sensilla along the lateral body wall
(Fig. 1D–F) and neurons
that innervate the terminal sensory organs
(Fig. 1J–L)
(Langlais et al., 2004;
Morton et al., 2005a). These
expression patterns closely matched those seen using in situ
hybridization in late embryos (Langlais et
al., 2004) (Fig.
1).
|
). In our previous studies using a combination of in
situ hybridization and immunocytochemistry with a neuronal marker in
embryos, we showed that Gyc-89Da and Gyc-89Db were expressed
in a single les and a single ves cell in the thoracic segments and
Gyc-89Db was expressed in a tracheal dendritic neuron (td cell) in
the first two abdominal segments only
(Langlais et al., 2004;
Morton et al., 2005a). Direct
visualization of GFP expression driven by the two promoters also labeled these
cells. Two cells on each side of the larvae were observed in each of the
thoracic segments of p89Da-GFP and p89Db-GFP larvae
(Table 1). One of these cells
was located in a more lateral position and the other in a more ventral
position corresponding to the positions of the les and ves cells,
respectively. The td neuron that was identified in the first two abdominal
segments of embryos using in situ hybridization of Gyc-89Db
(Langlais et al., 2004) was
also observed in p89Db-GFP larvae with the exception that these cells
were also seen in abdominal segment 3
(Table 1). In the terminal
abdominal segments there are seven sensory organs on either side in the larvae
(Stocker, 1994), and in
situ hybridizations in embryos showed that a single neuron innervating
six of these structures expressed Gyc-89Da and Gyc-89Db
(Langlais et al., 2004;
Morton et al., 2005a). Direct
visualization of GFP in p89Da-GFP and p89Db-GFP larvae also revealed these
neurons, and we were able to detect a single neuron in each of the seven
sensory organs (Table 1). These
experiments show that there is a very good match between expression of
Gyc-89Da and Gyc-89Db determined by in situ
hybridization and expression of GFP driven by portions of 5' genomic
DNA.
|
Our previous studies (Langlais et al.,
2004; Morton et al.,
2005a) used cell counts to determine the overlap in the expression
patterns of the three atypical sGC subunits. These experiments indicated that
in the majority of peripheral cells Gyc-89Da and Gyc-89Db
were co-expressed with Gyc-88E and suggested that in many cells all
three subunits were co-expressed (Langlais
et al., 2004; Morton et al.,
2005a). To determine the co-expression patterns of
Gyc-89Da and Gyc-89Db more accurately, we generated
transgenic flies that expressed the yeast transcription factor GAL4 under the
control of the upstream regions of these genes (designated p89Da-GAL4
and p89Db-GAL4) and crossed them with flies that expressed red
fluorescent protein (dsRed) under UAS control. Examination of larvae from
these crosses showed the same number of peripheral neurons as seen with the
p89Da-GFP and p89Db-GFP lines, respectively
(Table 1). We then generated
larvae that contained three transgenes: p89Da-GFP, p89Db-GAL4 and
UAS-dsRed, and examined the neurons under both green and red
fluorescence. In larvae expressing p89Da-GFP, p89Db-GAL4 and
UAS-dsRed we consistently observed four neurons that expressed
Gyc-89Da and four neurons that expressed Gyc-89Db in the
dorsal ganglion, but none of these neurons co-expressed the two subunit genes
(Fig. 2A). Similarly, in the
terminal ganglion, three to four neurons expressed Gyc-89Da and three
to four neurons expressed Gyc-89Db, but we never observed neurons in
the terminal ganglion that expressed both subunits
(Fig. 2A). In addition to the
neurons in the dorsal and terminal ganglia within the larval head, we
consistently observed co-expression in two neurons that appeared to be part of
the monoscolopidial chordotonal organs
(Fig. 2A), which lie adjacent
to the terminal ganglia (Campos-Ortega and
Hartenstein, 1997). Expression of Gyc-89Db alone was also
seen in two to three multidendritic arborization neurons
(Grueber et al., 2002) in the
head region.
|
In contrast to the terminal and dorsal ganglia, co-expression of
Gyc-89Da and Gyc-89Db was seen in
peripheral neurons in the thoracic segments. Both cells in all three thoracic
segments co-expressed Gyc-89Da and Gyc-89Db
(Fig. 2B,
Table 1). The td neuron in the
abdominal segments, however, only expressed Gyc-89Db and did
not show Gyc-89Da expression (Fig.
2B, Table 1). In
the caudal segments, Gyc-89Da and Gyc-89Db
were co-expressed in neurons that innervate peg sensilla on the sensory cones
(Fig. 2C). There are seven
sensory cones on each side of the larvae, which have been described as
possible chemosensory detectors (Stocker,
1994). Each sensory cone appears to be innervated by one to two
neurons and only a single neuron in each cone co-expressed the two subunit
genes. A neurite was clearly visible extending to the peg sensilla at the tip
of each cone (Fig. 2C,
inset).
Gyc-89Da and Gyc-89Db were also expressed in the larval CNS, but no co-expression of the two subunits was detected in central neurons (Fig. 2D). Numerous cells were labeled in the brain lobes and ventral nerve cord, with a larger number of cells that expressed Gyc-89Db compared with Gyc-89Da observed. The only cells in which Gyc-89Da and Gyc-89Db were co-localized were in the ring gland (neurosecretory organ), where several intrinsic secretory cells of the corpora cardiaca expressed both subunits (data not shown).
|
). TNT
abolishes transmitter release by cleaving the synaptic vesicle protein
synaptobrevin (Schiavo et al.,
1992; Link et al.,
1992). Identification of behaviors that are disrupted when TNT is
expressed in subsets of neurons in Drosophila provides a simple tool
for examining the function of these neurons
(Sweeney et al., 1995). When
we expressed the active form of TNT (TNTa) in the neurons that express
Gyc-89Da and Gyc-89Db the animals failed to develop into
adult flies and died at three specific developmental stages associated with
the molt (Fig. 3). As a control
we also used a fly line that expressed an inactive form of TNT (TNTi) under
UAS control (Sweeney et al.,
1995). About 90% of eggs generated from these crosses survived to
adult (Fig. 3). For those
neurons that expressed Gyc-89Da, expression of TNT caused slightly
reduced hatching rates (84.9±1.8% compared with 92.3±0.4% for
expression of TNTi, P<0.05). The majority of the hatchlings
survived to the end of the third instar and about 50% died at the beginning of
metamorphosis, at about the time of pupariation. All of the animals that
successfully pupated died just prior to adult eclosion (emergence from the
pupal cuticle). By contrast, expression of TNT in the
Gyc-89Db-expressing neurons caused all the animals to die at the
first–second larval transition (Fig.
3).
Death of p89Da-GAL4 x UAS-TNTa larvae during third instar possibly due to drowning
To determine whether there were general behavioral deficits during larval
stages, we tested the third instar p89Da-GAL4 x UAS-TNTa larvae for
their general locomotion on 1% agarose plates
(Osborne et al., 1997) and
observed no significant differences between those expressing TNTa and those
expressing TNTi in distance traveled (2.37±0.26 and 1.99±0.16 cm
min–1, respectively: N=9 for each genotype,
Student's t-test P=0.2372). Although locomotion and growth
appeared normal, we found substantial numbers of late third instar larvae that
expressed TNTa in the Gyc-89Da neurons deeply buried in the food, whereas most
of those that expressed TNTi had all crawled up the wall of the vial and
formed puparia. We previously suggested that the Drosophila atypical
sGCs act as neuronal O2 sensors because they are activated by
reduced O2 concentrations
(Morton, 2004b). In addition,
a related sGC in C. elegans (GCY-35) has been shown to mediate
O2-sensitive behaviors (Gray et
al., 2004). This suggested that larvae with inactive Gyc-89Da
neurons might not be able to sense that they were deep in the food and
suffocating. To prevent the TNTa-expressing animals from burying deeply into
the food, Petri dishes containing a thin layer of food (
3 mm thick) were
used instead of the normal food vials (3 cm depth of food). Under these
conditions, no significant differences in the number of animals that
successfully pupated were observed between those expressing TNTa and those
expressing TNTi (Fig. 4).
|
Neurons that express Gyc-89Da are necessary for expression of the adult eclosion motor program
Almost all (98.2±0.8%) of the p89Da-GAL4 x UAS-TNTa larvae
that successfully formed puparia failed to eclose as adult flies regardless of
whether they were raised on thin discs of food or in normal vials. To
investigate more closely the causes of the failure to eclose, we examined
animals during the final few hours before eclosion for the appearance of
developmental markers that precede eclosion
(Kimura and Truman, 1990).
Animals were examined under a dissecting microscope and the opercula removed
to facilitate detection of the markers. About 8 h prior to eclosion the head
of the developing adult is smooth, indicating that molting fluid is still
present between the pupal and adult cuticle. About 3 h prior to eclosion the
head has a grainy appearance, which indicates that molting fluid absorption is
complete. The tracheal system and the space between the adult and pupal
cuticles fill with air about 1 h before eclosion leading to a whitish
appearance of the head. This is followed at about 40 min before eclosion by
the extension of the ptilinum from the front of the head, which ruptures the
thin pupal cuticle. All of these stages occurred in the proper order in
p89Da-GAL4 x UAS-TNTa and p89Da-GAL4 x UAS-TNTi animals. Some
animals were also removed from the pupal cases at the onset of tracheal air
filling and examined under a microscope to determine the extent of tracheal
air filling throughout the body, which appeared normal and was comparable to
that of similarly staged control animals.
About 40 min after ptilinum extension, the p89Da-GAL4 x UAS-TNTi
control animals began inflating their heads and pushed forward, which enabled
the animals to rupture and push open the operculum. After several strong head
inflations (10–20 s), the p89Da-GAL4 x UAS-TNTi control animals
began the stereotyped abdominal, posterior-directed peristalsis movements
(eclosion behavior) leading to complete emergence from the puparium. The
eclosion of these control animals is comparable to that of wild-type Canton-S
animals (McNabb et al., 1997).
The animals that expressed the active form of TNT (p89Da-GAL4 x
UAS-TNTa) also began head inflations about 40 min after ptilinum
extension, but the magnitude of the inflations appeared much weaker. These
weaker head inflations rarely succeeded in completely opening the operculum,
when it was left intact. Several head inflations/deflations occurred and the
animal did push forward noticeably, but the abdominal peristalsis component of
the eclosion motor program was never initiated in these animals.
Interestingly, head inflations/deflations occurred for up to 4 more hours.
During this time, starting about an hour after the first head inflations, the
antennae occasionally twitched, and the abdomen sporadically contracted
several times a minute, but these movements did not resemble the coordinated
series of peristaltic contractions seen in control animals. Even when the
operculum was removed just prior to the expected time of eclosion, the animals
expressing TNTa in the Gyc-89Da neurons never succeeded in escaping from the
puparium.
After wild-type flies have eclosed they undergo wing inflation and cuticle
tanning (see Ewer and Reynolds,
2002). To determine whether animals that expressed TNTa in the
Gyc-89Da neurons completed these processes in the absence of eclosion, we
removed them from their puparia and observed them for up to 2 h. These animals
remained immobile, never showed any wing inflation and after 2 h still had
un-tanned cuticle. By contrast, animals that expressed TNTi inflated their
wings and tanned their cuticles within 30 min of eclosion.
Neurons that express Gyc-89Db are necessary for larval ecdysis
In contrast to the p89Da-GAL4 x UAS-TNTa progeny, the majority of
which survived to the third instar, when TNTa was expressed in the cells that
express Gyc-89Db, all the larvae died before they reached the second
instar (Fig. 3). Fewer of the
p89Db-GAL4 x UAS-TNTa progeny hatched compared with controls expressing
TNTi (85.3±1.7% vs 92.6±1.3%, Student's
t-test, P<0.05) and the newly hatched larvae expressing
TNTa appeared to become increasingly sluggish after a day of feeding on the
surface of the food. Although the larvae appeared morphologically normal they
all failed to reach the second larval instar. About 80% of the p89Db-GAL4
x UAS-TNTa larvae that hatched survived through the feeding phase and
began the developmental events that lead to ecdysis (83.2±1.4% compared
with 97.4±0.3% TNTi controls, Student's t-test,
P<0.05). Of the 16.8% of larvae that died during the feeding
phase, most were found buried under the food (63% buried vs 37% on
top). This appeared to be similar to the third instar larvae expressing TNTa
in the Gyc-89Da neurons, many of which appeared to drown in their food,
possibly because they failed to detect that they had become anoxic.
There are several morphological markers that have been described that
precede larval ecdysis in Drosophila
(Park et al., 2002;
Clark et al., 2004). About 1 h
prior to the first larval ecdysis the new mouth parts of the second instar
larvae become visible, a stage referred to as double vertical plates [dVP
(Park et al., 2002)]. This is
followed about 10 min later by the collapse of the old tracheal lining and
almost immediately by the trachea filling with air. Shortly after tracheal air
filling the pre-ecdysis and ecdysis behaviors commence
(Park et al., 2002). Over 99%
of the p89Db-GAL4 x UAS-TNTa larvae that survived past the first instar
feeding phase were found with transparent fluid-filled double trachea and dVP.
No animals were found with air-filled tracheae or that initiated pre-ecdysis
or ecdysis peristaltic movements. We presume that the larvae found with dVP
and transparent, fluid-filled trachea probably died from suffocation as they
were unable to air fill their tracheae.
Gyc-89Da and Gyc-89Db neurons do not contain EH or CCAP
The endocrinology underlying the initiation of ecdysis and eclosion in
Drosophila has been extensively studied and is known to be regulated
by the release and action of at least three peptide hormones: ecdysis
triggering hormone (ETH), eclosion hormone (EH) and crustacean cardiac
acceleratory peptide (CCAP) (McNabb et
al., 1997; Park et al.,
2002; Clark et al.,
2004). Deletion of the gene encoding these peptides or elimination
of the cells that secrete the peptide lead to similar phenotypes to those
described above for expression of TNTa in the cells that express
Gyc-89Da and Gyc-89Db. For example, deletion of the
ETH gene leads to lethality at the first larval ecdysis with most of
the animals dying after tracheal inflation and some animals showing some
`ecdysis-like' movements, although none initiated pre-ecdysis movements
(Park et al., 2002). Killing
the EH cells leads to larval death, with most of the larvae dying with
fluid-filled tracheae, although the timing of larval pre-ecdysis and ecdysis
behaviors appeared unaffected (McNabb et
al., 1997; Clark et al.,
2004). Furthermore, EH cell knockouts led to the disruption of
adult eclosion, notably with a longer period between ptilinum extension and
eclosion (McNabb et al.,
1997). It is possible that the effects of TNTa expression in the
Gyc-89Da and Gyc-89Db neurons were due to a failure of the release of some or
all of these peptides. The simplest explanation for this would be if TNTa was
expressed in the peptidergic cells directly preventing peptide release. ETH is
expressed in the epitracheal gland located along the major tracheal branches
(O'Brien and Taghert, 1998;
Park et al., 2002). No
expression of GFP or dsRed was seen in these cells but expression was seen in
the brain and ventral nerve cord in locations that might correspond to the EH
and/or CCAP cells (Fig. 2)
(Langlais et al., 2004).
|
) or the
upstream region of CCAP coupled to GAL4
(Park et al., 2003).
Fig. 5 shows images of the CNS
from these larvae. EH is expressed in a single pair of cells in the ventral
midline of the brain (Horodyski et al.,
1993). Fig. 5A,B
clearly shows that there was no overlap in the expression of dsRed driven by
the upstream region of EH with GFP driven by the promoter of either
Gyc-89Da or Gyc-89Db. CCAP is expressed in a row of paired
cells along the lateral edges of the ventral nerve cord
(Park et al., 2003) and again
there was no overlap with the expression of Gyc-89Da or
Gyc-89Db (Fig.
5C,D).
The row of cells lying adjacent to the CCAP cells appears to be in a
similar position to peptidergic neurons that express the transcription factor
`dimmed' (Hewes et al., 2003).
To determine whether any of the Gyc-89Da- or
Gyc-89Db-expressing cells also expressed dimmed, we generated larvae
that contained the dimmed-GAL4, UAS-dsRed and p89Da-GFP or
p89Db-GFP transgenes. Examination of the nervous tissue of these
larvae showed that paired cells in the ventral nerve cord co-expressed
Gyc-89Da and dimmed (Fig.
5E). These cells appeared to be in a similar position to the
dorsal furin-containing neurons (d1–d11) described by Hewes et al.
(Hewes et al., 2003). These
furin-expressing cells also express the transcription factor `apterous' and it
has been suggested that neuropeptides from these cells are involved in the
control of larval ecdysis (Park et al.,
2004). We also tested whether there was overlap in the expression
patterns of the Gyc-89Da or Gyc-89Db cells using an apterous enhancer trap
line (Park et al., 2004).
Examination of the larval CNS from animals bearing UAS-dsRed, ap-GAL4
and p89Da-GFP or p89Db-GAL4 showed no overlap in expression
of either Gyc-89Da or Gyc-89Db and apterous (data
not shown). It therefore appears that the subset of Gyc-89Da neurons
that are dimmed positive are apterous negative.
Reduced levels of cGMP in Gyc-89Da and Gyc-89Db neurons has no effect on larval ecdysis or eclosion
Several studies have shown that cGMP mediates the action of EH and although
the identity of the guanylyl cyclase that is activated is not known, there is
evidence that it is an atypical sGC (see
Morton and Simpson, 2002).
Because we were specifically targeting neurons that expressed this subclass of
sGC, it was possible that cGMP in these neurons was required for, or
modulated, ecdysis and eclosion. To test for this we expressed a cGMP-specific
phosphodiesterase (bovine PDE5) in the Gyc-89Da and Gyc-89Db neurons using a
UAS-bPDE5 fly line (Broderick et al.,
2004). Progeny of these crosses all developed and eclosed normally
with no differences in either the time taken for development or the number of
animals that successfully eclosed compared with either of the parental lines
(data not shown). We also closely observed the final stages of adult eclosion
as described above and could detect no differences in the time taken for any
of the stages prior to eclosion (data not shown).
Expression of TNTa in the Gyc-89Da neurons is necessary early in adult development to prevent eclosion
The action of TNTa in the Gyc-89Da neurons to prevent eclosion could be due
to its acute action at the time of eclosion or it could be due to disruption
of an earlier developmental event that is subsequently required for eclosion.
To distinguish between these two possibilities we co-expressed GAL80ts, a
temperature-sensitive inhibitor of GAL4, with p89Da-GAL4 and
UAS-TNTa to control the temporal expression of TNTa. The yeast GAL80
protein blocks GAL4 activity by binding to its transcriptional activation
domain, a function that is retained when it is expressed in
Drosophila (Yocum and Johnston,
1984; Zeidler et al.,
2004). A temperature-sensitive version of GAL80 (GAL80ts) was
created by inserting a temperature-controlled yeast intein into a crucial
region of the GAL80 peptide and placing this construct under the control of a
ubiquitous tubulin promoter (tubP-GAL80ts)
(McGuire et al., 2003). We
then generated flies that were homozygous for p89Da-GAL4 and
tubP-GAL80ts, which were then crossed with UAS-TNTa and UAS-TNTi
flies. At the permissive temperature (18°C), GAL80ts inhibits GAL4 action,
preventing TNT expression, while at 30°C GAL80ts is inactive, allowing
GAL4 to drive expression of TNT. To test the effectiveness of this system we
first crossed tubP-GAL80ts;p89Da-GAL4 flies to UAS-dsRed flies and raised the
offspring at either 18°C or 30°C. The larvae raised at 30°C
displayed robust dsRed expression whereas larvae raised at 18°C displayed
no dsRed expression in the CNS or PNS, but did retain some expression in the
salivary glands, indicating that GAL80ts function was mostly inhibited at this
temperature (data not shown). To confirm that we could use this system to
regulate TNT expression during adult development we transferred flies
containing tubP-GAL80ts;p89Da-GAL4 and UAS-dsRed transgenes
from 18°C to 30°C about mid-way through adult development (96 h at
18°C) and examined expression of dsRed in the brain at various times after
temperature shift. Examples of these experiments are shown in
Fig. 6. No expression was seen
in the absence of a temperature shift (Fig.
6A), low levels of expression were seen 7 h after the temperature
shift (Fig. 6B) and robust
expression was seen 16 h after the shift to 30°C
(Fig. 6C).
|
Expression of TNTa in the Gyc-89Db neurons early in adult development also prevents eclosion
Because TNTa expression in Gyc-89Db neurons blocked the first larval
ecdysis it was not possible to determine whether Gyc-89Db neurons were
required for all ecdyses. To test this we again used the temperature-sensitive
GAL80. Flies were generated that were homozygous for both p89Db-GAL4
and tubP-GAL80ts and these were crossed with both UAS-TNTa and
UAS-TNTi. The progeny were maintained at 18°C and then shifted to 30°C
at various times after WP and the number that successfully eclosed was noted.
These data are also shown in Fig.
6D. The results were very similar as those found with Gyc-89Da.
When the temperature shift occurred early in adult development, eclosion was
subsequently blocked whereas shifting at later stages had little effect on
eclosion rates. Interestingly, the critical time in development was also
similar to that found for expression of TNTa in the Gyc-89Da neurons. When TNT
was expressed earlier than 48 h after WP, the majority of the animals failed
to eclose.
We also shifted the p89Db-GAL4;tubP-GAL80ts;UAS-TNTa and UAS-TNTi animals
from 18°C to 30°C prior to the WP stage to determine whether they
could successfully form puparia. Animals expressing UAS-TNTa that were shifted
during the first instar failed to form puparia (2.7% success) whereas those
shifted during the second and third instar were more successful (70.5% and
93.8%, respectively). Soon after puparium formation, Drosophila
larvae undergo a process known as head eversion, which appears to be the
behavior equivalent to pupal ecdysis in other holometabolous insects
(Zdárek and Friedman,
1986). All of the larvae containing
p89Db-GAL4;tubP-GAL80ts and UAS-TNTa that were shifted to
30°C during the second and third instar that successfully formed puparia
also successfully completed head eversion (data not shown). Thus neurons that
express Gyc-89Db are required for larval and adult ecdyses, but not
head eversion.
|
DISCUSSION |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
;
Morton et al., 2005a). In the
present study we extended these observations by generating fly lines that
expressed either GFP or GAL4 under the control of the upstream regions for two
of the three atypical sGCs, Gyc-89Da and Gyc-89Db. We first
confirmed that the expression patterns generated by the predicted promoter
regions for each of the genes matched the expression patterns revealed by
in situ hybridization. For the peripheral sensory neurons, we
compared the number of cells labeled and their positions in the embryonic
in situ preparations with third instar larvae expressing GFP under
the direct control of the predicted promoter regions for each gene. In
general, the number and position of the cells labeled with either technique
matched well. These included neurons innervating the dorsal and terminal
organs, lateral and ventral external sensilla and the terminal sensory cones.
These data strongly suggested that the sequences of DNA 5' to the
Gyc-89Da and Gyc-89Db genes that we used in our constructs
were sufficient to mimic the expression patterns of these genes.
In the earlier in situ hybridization studies, we used the
locations of the cells and cell counts to conclude that in most, if not all,
of the peripheral sensory neurons all three of the atypical sGCs were
co-expressed in the same cells (Langlais
et al., 2004; Morton et al.,
2005a). Using the GFP and GAL4 reporter lines we now show more
clearly the extent of the overlap between Gyc-89Da and
Gyc-89Db expression and in contrast to the in situ
hybridization data, the present study shows relatively little overlap in the
expression pattern of these two subunits. There are two primary reasons for
the apparent overlap of expression seen using in situ hybridization.
Firstly, the sequence similarity between Gyc-89Da and
Gyc-89Db makes it difficult to generate gene-specific probes to each
gene, possibly resulting in some non-specific labeling. Secondly, most of the
cell counts made with the simultaneous application of two probes used a probe
to Gyc-88E in combination with either Gyc-89Da or
Gyc-89Db. The cell counts were similar in both cases to the cell
counts using single probes (Langlais et
al., 2004) suggesting that all three genes were co-expressed in
the same cells. If we had carried out cell counts combining short,
gene-specific probes to both Gyc-89Da and Gyc-89Db, we might
have been able to see more cells than with either probe alone, which would
have suggested a lack of co-expression. The results from the present study
using expression of fluorescent proteins showed that in the olfactory and
gustatory neurons there is no overlap in the expression patterns, with each
subunit expressed in about four neurons in each ganglion. Along the lateral
body wall each of the single neurons innervating external sensilla
co-expressed Gyc-89Da and Gyc-89Db whereas the tracheal
dendritic neurons only expressed Gyc-89Db. The terminal sensory cones
showed a similar pattern to that seen with in situ hybridization
– a single neuron in each cone expressed both atypical sGCs. In the CNS
it had not previously been possible to examine co-expression patterns because
of the number cells and lack of morphological markers. The reporter constructs
show clearly that Gyc-89Da and Gyc-89Db are not co-expressed
in any CNS neurons. The only locations where they are co-localized are in some
of the intrinsic cells of the corpora cardiaca. Because Gyc-89Da and Gyc-89Db
are only enzymatically active when co-expressed with the third atypical sGC,
Gyc-88E (Langlais et al.,
2004; Morton et al.,
2005a), we assume that Gyc-88E is also expressed in all
the central and peripheral cells that express Gyc-89Da and
Gyc-89Db. The in situ hybridization experiments support
this, but as we have not been able to define the promoter region for
Gyc-88E or generate antisera against Gyc-88E, we have not been able
to show this directly.
The identity of the Gyc-89Da and Gyc-89Db cells in the CNS are unknown at
this time. We tried a variety of markers that appeared to have similar
expression patterns but were unable to see any overlap. These include cells
that contain the neuropeptides EH and CCAP and the transcription factor
apterous. The transcription factor dimmed is expressed in a wide variety of
peptidergic neurons (Hewes et al.,
2003) and showed co-expression with a small subset of neurons
expressing Gyc-89Da but had no overlap in the CNS with
Gyc-89Db. This suggests that this subset of Gyc-89Da neurons is
peptidergic, but the identity of the peptide is unknown.
Functions of the Gyc-89Da and Gyc-89Db neurons
We used the neurotoxin TNT to determine the function of the neurons that
express Gyc-89Da and Gyc-89Db. TNT blocks synaptic
transmission by cleaving the synaptic vesicle protein synaptobrevin
(Schiavo et al., 1992;
Link et al., 1992) and
expression of TNT in specific neurons in Drosophila has helped define
neuronal networks involved in a variety of behaviors
(Martin et al., 2002). When we
expressed TNT in either the Gyc-89Da or Gyc-89Db neurons we found that the
animals died at three specific stages (Fig.
3). TNT expression in Gyc-89Da neurons caused about half of the
animals to die at the late larval stage and the remainder of the animals to
die just prior to the onset of the eclosion motor program. The lethality of
third instar larvae was associated with burrowing into the food and could be
rescued by allowing the animals to develop in a thin layer of food. We have
shown that the activity of the atypical sGCs is regulated by oxygen and have
suggested that they could act as oxygen sensors
(Morton, 2004b). If the
Gyc-89Da neurons are excited by reduced oxygen levels and activate
hypoxia-associated behaviors, when synaptic transmission of these neurons is
blocked by TNT the animals would no longer respond to hypoxia. One response of
larvae to hypoxia is to withdraw from the food and activate searching
behaviors (Wingrove and O'Farrell,
1999). Our results are consistent with the Gyc-89Da neurons either
detecting hypoxia or activating the hypoxia escape behavior. If synaptic
transmission is blocked in these neurons the animals would fail to respond to
hypoxia, continue to burrow into the food and suffocate. Most of the larval
lethality that we observed was late in the third instar, presumably because at
this time locomotion increases as the larvae crawl out of their food to seek a
location to pupate. Wingrove and O'Farrell
(Wingrove and O'Farrell, 1999)
have shown that the hypoxia escape response is probably mediated by a
cGMP/protein kinase G (PKG) pathway and we have preliminary data that show
that cGMP in the Gyc-89Da neurons is necessary for a normal hypoxia escape
response (Morton et al.,
2005b).
The other major period of lethality for animals when TNT was expressed in
the Gyc-89Da neurons was just prior to adult eclosion. All of the stages that
precede the onset of the eclosion motor program occurred normally and in the
appropriate order until the final stage – head expansion. This time
corresponds closely to the time at which defects in eclosion were first
detected in flies lacking the neurons that contain EH
(McNabb et al., 1997), a
neuropeptide known to trigger ecdysis and eclosion in most, if not all,
insects (Truman et al., 1981).
The simplest interpretation of our findings is that Gyc-89Da is
expressed in EH neurons and expression of TNT in these cells prevented EH
release. This is not the case as there was no overlap in EH and
Gyc-89Da expression patterns. Cells that are immediately upstream and
downstream of EH in the pathway that regulates eclosion are the Inka cells of
the epitracheal glands and the CCAP neurons (see
Ewer and Reynolds, 2002;
Clark et al., 2004). No overlap
in the expression pattern of Gyc-89Da with either of these cells was
seen. It is possible that the Gyc-89Da neurons act further upstream of the EH
cells, although the Gyc-89Da neurons must affect a larger number of neurons
associated with eclosion than the EH cells as the phenotype of TNT expression
in the Gyc-89Da neurons is much more severe than the EH cell deletion. Almost
all the p89Da-TNTa animals failed to eclose whereas about a third of the flies
lacking EH cells complete development and eclose
(McNabb et al., 1997). As with
the EH cell deletions, expression of TNT in the Gyc-89Da neurons prevented
normal tanning and wing expansion.
Expression of TNT in the Gyc-89Db neurons also phenocopied the effects of
eliminating a neuropeptide associated with eclosion. ETH acts both upstream
and downstream of EH as part of a positive feedback loop
(Clark et al., 2004) and
mutations that eliminate the ETH gene cause lethality at the first
larval ecdysis (Park et al.,
2002). The timing of death in ETH–/– larvae
(Park et al., 2002) is very
similar to that of larvae expressing TNT in the Gyc-89Db neurons – just
prior to air filling of the tracheae. In wild-type larvae it only takes about
2 min between tracheal collapse and air filling of the tracheae
(Park et al., 2002;
Clark et al., 2004).
Interestingly, in the EH cell deletions, many of the larvae also failed to air
fill their tracheae and this was a major cause of lethality
(McNabb et al., 1997). Thus
elimination of either ETH or EH results in a similar phenotype to blocking
neurotransmission in the Gyc-89Db neurons. As described above for the Gyc-89Da
neurons, a simple explanation for this is that TNT is expressed in ETH, EH or
CCAP cells preventing the release of the peptides. This is not the case as we
saw no overlap in expression patterns with any of these peptides and
Gyc-89Db. Additional peptides believed to be involved with activation
of the ecdysis behavior program include peptides in the apterous cells
(Park et al., 2004), corazonin
(Kim et al., 2004) and the
products of the hugin gene (Meng
et al., 2002). Although we saw no overlap with apterous it is
possible that the release of other peptides is prevented by expression of TNT
in the Gyc-89Da or Gyc-89Db neurons. Although TNT blocks synaptic transmission
at conventional synapses in Drosophila, there is little experimental
support that in Drosophila TNT will block peptide release
(Martin et al., 2002).
Another possibility is that synaptic transmission in Gyc-89Db neurons is
directly required for successful air filling of tracheae. The physiological
basis for air filling the tracheae at each molt is not known although the ETH
neuropeptide is involved (Park et al.,
2002). Gyc-89Db is expressed (in the absence of
Gyc-89Da) in sensory neurons that are closely associated with
tracheae (Fig. 2B). If these
neurons signal to the CNS when apolysis has progressed sufficiently to trigger
air filling, failure of synaptic transmission in these cells could account for
the inability of the animal to successfully fill its tracheae with air.
Effects of temporal control of TNT expression
The data discussed above support the model that synaptic transmission in
neurons that express Gyc-89Da and Gyc-89Db is required for
either the activation or the generation of the eclosion and ecdysis motor
programs. This model predicted that TNTa expression in these neurons just
prior to ecdysis or eclosion is sufficient to block the behavior. Our
experiments using GAL80ts to control the temporal expression of TNTa showed
that this was not the case. TNTa expression about 100 h before eclosion is
required to block eclosion. This delay is not explained by the time it takes
for the inhibition of GAL4 activation to be removed. McGuire et al.
(McGuire et al., 2003) found
that 30 min after the temperature shift a reporter gene was detectable and
half-maximal expression was seen after 3 h. Our data showed that expression of
the red fluorescent protein dsRed was detectable 7 h after the temperature
shift and was robust 16 h later.
Our findings suggest that TNT expression probably blocks a developmental
event that is required for the subsequent expression of eclosion behaviors.
Previous studies have also shown developmental effects of TNT expression in
Drosophila neurons. In the embryonic neuromuscular system, expression
of TNT in motor neurons resulted in the expected blockage of neuromuscular
synaptic transmission and also reduced synaptic input to the motor neurons
(Baines et al., 1999). In a
detailed analysis of the effects of TNT expression in the developing optic
lobe, Hiesinger et al. (Hiesinger et al.,
1999) found that cell survival, axonal pathfinding and target
recognition were unaffected by TNT expression, but morphological alterations
in photoreceptor terminals was observed between 25% and 50% of adult
development. This time corresponds to the window of sensitivity that we
observed for the blockage of adult eclosion. This suggests that expression of
TNT in the Gyc-89Da and Gyc-89Db neurons disrupts their connectivity and leads
to a blockade in eclosion. It is interesting that the Gyc-89Da neurons are not
required for larval ecdysis, whereas the Gyc-89Db neurons are, but for adult
eclosion TNT expression in either population of cells blocked the behavior. In
the larval CNS there was very little overlap of expression, and similarly in
the adult CNS few, if any, neurons co-expressed Gyc-89Da and
Gyc-89Db.
In summary, by using targeted expression of TNT in neurons that express two atypical soluble GCs, we have shown that synaptic transmission in neurons that express Gyc-89Da is required for adult eclosion and neurons that express Gyc-89Db are required for larval ecdysis and adult eclosion. By contrast, neither of these populations of neurons is required for head eversion. The use of the driver lines also enabled us to more completely map the expression patterns of these two GC subunits and we found some co-expression in peripheral sensory neurons, but almost no overlap in CNS expression.
|
References |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
Baines, R. A., Robinson, S. G., Fujioka, M., Jaynes, J. B. and Bate, M. (1999). Postsynaptic expression of tetanus toxin light chain blocks synaptogenesis in Drosophila. Curr. Biol. 9,1267 -1270.[CrossRef][Medline]
Barolo, S., Carver, L. A. and Posakony, J. W. (2000). GFP and b-Galactosidase transformation vectors for promoter/enhancer analysis in Drosophila.BioTechniques 29,727 -732.
Bodmer, R., Barbel, S., Shepard, S., Jack, J. W., Jan, L. Y. and Jan, Y. N. (1987). Transformation of sensory organs by mutations of the cut locus of D. melanogaster. Cell 51,293 -307.[CrossRef][Medline]
Brand, A. H. and Perrimon, N. (1993). Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development 118,401 -415.[Abstract]
Broderick, K. E., Kean, L., Dow, J. A., Pyne, N. J. and Davies,
S. A. (2004). Ectopic expression of bovine type 5
phosphodiesterase confers a renal phenotype in Drosophila. J. Biol.
Chem. 279,8159
-8168.
Campos-Ortega, J. and Hartenstein, V. (1997).Embryonic Development of Drosophila melanogaster (2nd edn) . Berlin: Springer-Verlag.
Clark, A. C., del Campo, M. L. and Ewer, J.
(2004). Neuroendocrine control of larval ecdysis behavior in
Drosophila: complex regulation by partially redundant neuropeptides.
J. Neurosci. 24,4283
-4292.
Ewer, J. and Reynolds, S. (2002). Neuropeptide control of molting in insects. Horm. Brain Behav. 3, 1-92.
Gray, J. M., Karow, D. S., Lu, H., Chang, A. J., Chang, J. S., Ellis, R. E., Martletta, M. A. and Bargmann, C. I. (2004). Oxygen sensation and social feeding mediated by a C. elegans guanylate cyclase homologue. Nature 430,317 -322.[CrossRef][Medline]
Greenspan, R. J. (1997). Fly Pushing. Plainview: Cold Spring Harbor Laboratory Press.
Grueber, W. B., Jan, L. Y. and Jan, Y. N. (2002). Tiling of the Drosophila epidermis by multidendritic sensory neurons. Development 129,2867 -2878.[Medline]
Heimbeck, G., Bugnon, V., Gendre, N., Habelin, C. and Stocker,
R. F. (1999). Smell and taste perception in Drosophila
melanogaster larva: toxin expression studies in chemosensory neurons.
J. Neurosci. 19,6599
-6609.
Hewes, R. S., Park, D., Gauthier, S. A., Schaefer, A. and
Taghert, P. H. (2003). The bHLH protein Dimmed controls
neuroendocrine cell differentiation in Drosophila.Development 130,1771
-1781.
Hiesinger, P. R., Reiter, C., Schau, H. and Fischbach, K. F.
(1999). Neuropil pattern formation and regulation of cell
adhesion molecules in Drosophila optic lobe development depends on
synaptobrevin. J. Neurosci.
19,7548
-7556.
Horodyski, F. M., Ewer, J., Riddiford, L. M. and Truman, J. W. (1993). Isolation, characterization and expression of the eclosion hormone gene in Drosophila melanogaster. Eur. J. Biochem. 215,221 -228.[Medline]
Kim, Y.-J., Spalovská-Valachová, I., Cho, K.-H.,
Zitnanova, I., Park, Y., Adams, M. E. and 
itnan, D.
(2004). Corazonin receptor signaling in ecdysis initiation.
Proc. Natl. Acad. Sci. USA
101,6704
-6709.
Kimura, K. and Truman, J. W. (1990). Postmetamorphic cell death in the nervous and muscular systems of Drosophila melanogaster. J. Neurosci. 10,403 -411.[Abstract]
Langlais, K. K., Stewart, J. A. and Morton, D. B.
(2004). Preliminary characterization of two atypical sGC in the
central and peripheral nervous system of Drosophila melanogaster.J. Exp. Biol. 207,2323
-2338.
Link, E., Edelman, L., Chou, J. H., Binz, T., Yamasaki, S., Eisel, U., Baumert, M., Südhof, T. C., Niemann, H. and Jahn, R. (1992). Tetanus toxin action: inhibition of neurotransmitter release linked to synaptobrevin proteolysis. Biochem. Biophys. Res. Commun. 189,1017 -1023.[CrossRef][Medline]
Lucas, K. A., Pitari, G. M., Kazerounian, S., Ruiz-Stewart, I.,
Park, J., Schulz, S., Chepenik, K. P. and Waldman, S. A.
(2000). Guanylyl cyclases and signaling by cyclic GMP.
Pharmacol. Rev. 52,375
-413.
Martin, J. R., Keller, A. and Sweeney, S. T. (2002). Targeted expression of tetanus toxin: a new tool to study the neurobiology of behavior. Adv. Genet. 47, 1-47.[Medline]
McGuire, S. E., Le, P. T., Osborn, A. J., Matsumoto, K. and
Davis, R. L. (2003). Spatiotemporal rescue of memory function
in Drosophila. Science
302,1765
-1768.
McNabb, S. L., Baker, J. D., Agapite, J., Steller, H., Riddiford, L. M. and Truman, J. W. (1997). Disruption of a behavioral sequence by targeted death of peptidergic neurons in Drosophila. Neuron 19,813 -823.[CrossRef][Medline]
Meng, X., Wahlstrom, G., Immonen, T., Kolmer, M., Tirronen, M., Predel, R., Kalkkinen, N., Heino, T., Sariola, H. and Roos, C. (2002). The Drosophila hugin gene codes for myostimulatory and ecdysis-modifying neuropeptides. Mech. Dev. 117,5 -13.[CrossRef][Medline]
Morton, D. B. (2004a). Invertebrates yield a plethora of atypical guanylyl cyclases. Mol. Neurol. 29, 97-115.[CrossRef]
Morton, D. B. (2004b). Atypical sGCs in
Drosophila can function as molecular oxygen sensors. J.
Biol. Chem. 279,50651
-50653.
Morton, D. B. and Hudson, M. L. (2002). Cyclic GMP regulation and function in insects. Adv. Insect Physiol. 29,1 -54.[CrossRef]
Morton, D. B. and Simpson, P. J. (2002). Cellular signaling in eclosion hormone action. J. Insect Physiol. 48,1 -13.[Medline]
Morton, D. B., Langlais, K. K., Steward, J. A. and Vermehren, A. (2005a). Comparison of the properties of the five sGC subunits in Drosophila melanogaster. J. Insect Sci. 5, 12.[Medline]
Morton, D. B., Langlais, K. K., Steward, J. A. and Vermehren, A. (2005b). Atypical sGCs in Drosophila as possible oxygen sensors. 2005 abstract viewer/itinerary planner, program 613.7. Washington, DC: Society for Neuroscience. http://www.stn.org/index.cfm?pagename=abstracts_ampublications.
O'Brien, M. A. and Taghert, P. H. (1998). A peritracheal neuropeptides system in insects: release of myomodulin-like peptides at ecdysis. J. Exp. Biol. 201,193 -209.[Abstract]
Osborne, K. A., Robichon, A., Burgess, E., Butland, S., Shaw, R.
A., Coulthard, A., Pereira, H. S., Greenspan, R. J. and Sokolowski, M. B.
(1997). Natural behavior polymorphism due to a cGMP-dependent
protein kinase of Drosophila. Science
277,834
-836.
Park, D., Han, M., Kim, Y. C., Han, K. A. and Taghert, P. H. (2004). Ap-let neurons – a peptidergic circuit potentially controlling ecdysial behavior in Drosophila. Dev. Biol. 269,95 -108.[CrossRef][Medline]
Park, J. H., Schroeder, A. J., Helfrich-Forster, C., Jackson, F.
R. and Ewer, J. (2003). Targeted ablation of CCAP
neuropeptide-containing neurons of Drosophila causes specific defects
in execution and circadian timing of ecdysis behavior.
Development 130,2645
-2656.
Park, Y., Filippov, V., Gill, S. S. and Adams, M. E. (2002). Deletion of the ecdysis-triggering hormone gene leads to lethal ecdysis deficiency. Development 129,493 -503.[Medline]
Schiavo, G., Benfenati, F., Poulain, B., Rossetto, O., DeLaurento, P. P., DasGupta, B. R. and Montecucco, C. (1992). Tetanus and botulinum-B neurotoxins block neurotransmitter release by proteolytic cleavage of synaptobrevin. Nature 359,832 -835.[CrossRef][Medline]
Sharma, Y., Cheung, U., Larsen, E. W. and Eberl, D. F. (2002). PPTGAL, a convenient Gal4 P-element vector for testing expression of enhancer fragments in Drosophila.Genesis 34,115 -118.[CrossRef][Medline]
Stocker, R. F. (1994). The organization of the chemosensory system in Drosophila melanogaster: a review. Cell Tissue Res. 275,3 -26.[CrossRef][Medline]
Sullivan, W., Ashburner, M. and Hawler, R. S. (2000). Drosophila Protocols. New York: Cold Spring Harbor Laboratory Press.
Sweeney, S. T., Kendal, B., Keane, J., Niemann, H. and O'Kane, C. J. (1995). Targeted expression of tetanus toxin light chain in Drosophila specifically eliminates synaptic transmission and causes behavioral effects. Neuron 14,341 -351.[CrossRef][Medline]
Truman, J. W., Taghert, P. H., Copenhaver, P. F., Tublitz, N. J. and Schwartz, L. M. (1981). Eclosion hormone may control all ecdyses in insects. Nature 291, 70-71.[CrossRef]
Vermehren, A., Langlais, K. K. and Morton, D. B. (2006). Oxygen-sensitive guanylyl cyclases in insects and their potential roles in oxygen detection and in feeding behaviors. J. Insect Physiol. 52,340 -348.[CrossRef][Medline]
Wingrove, J. A. and O'Farrell, P. H. (1999). Nitric oxide contributes to behavioral, cellular, and developmental responses to low oxygen in Drosophila. Cell 98,105 -114.[CrossRef][Medline]
Yocum, R. R. and Johnston, M. (1984). Molecular cloning of the GAL80 gene from Saccharomyces cerevisiae and characterization of a gal80 deletion. Gene 32, 75-82.[CrossRef][Medline]
Zdárek, J. and Friedman, S. (1986). Pupal ecdysis in flies: mechanisms of evagination of the head and expansion of the thoracic appendages. J. Insect Physiol. 32,917 -923.[CrossRef]
Zeidler, M. P., Tan, C., Bellaiche, Y., Cherry, S., Hader, S., Gayko, U. and Perrimon, N. (2004). Temperature-sensitive control of protein activity by conditionally splicing inteins. Nat. Biotechnol. 22,871 -876.[CrossRef][Medline]
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati 
Twitter What's this?
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
