Conservation of capa peptide-induced nitric oxide signalling in Diptera
1 Institute of Biomedical and Life Sciences, Division of Molecular Genetics,
University of Glasgow, Glasgow G11 6NU, UK
2 Centre for Tropical Veterinary Medicine, Royal School of Veterinary
Studies, University of Edinburgh, Edinburgh EH9 1QH, UK
* Author for correspondence (e-mail: s.a.davies{at}bio.gla.ac.uk)
Accepted 23 August 2004
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Summary |
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Key words: Malpighian tubule, fluid transport, mosquito, tsetse, capa receptor, NOS/cGMP
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Introduction |
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Drosophila melanogaster Malpighian tubules are now accepted as a
genetic model of transporting epithelia
(Dow and Davies, 2003). In the
development of this renal model, different techniques have been developed to
assess tubule function: fluid transport rates
(Dow et al., 1994a
),
electrophysiological responses (Davies et
al., 1995
), ion transport
(Dow, 1999
) and calcium
signalling using aequorin transgenes
(Rosay et al., 1997
). This
battery of physiological assays, in combination with the powerful genetic
tools associated with Drosophila, has allowed rapid, organotypic
analysis of the cell-specific control of tubule function
(Dow and Davies, 2003
). Given
the conserved and critical role of the tubule in insect life, findings from
the Drosophila tubule may usefully be applied to those insect species
with less developed genomic resources but greater economic or medical
significance (Dow and Davies,
2003
). In particular, findings from Drosophila might be
useful in studies of other Diptera; for example, Aedes, Anopheles and
Glossina.
Diuresis in Drosophila tubules has been shown to be directly
stimulated by exogenous guanosine 3', 5'-cyclic monophosphate
(cGMP), which enters tubule cells via a cyclic nucleotide transporter
(Riegel et al., 1998), and by
nitric oxide (NO; Dow et al.,
1994b
). NO/cGMP signalling is compartmentalised to principal cells
in the main, fluid-secreting segment of tubules, containing the electrogenic
vacuolar H+-ATPase (V-ATPase) pump
(Dow, 1999
), which energises
fluid transport. Furthermore, electrophysiological studies suggest that cGMP
signalling modulates V-ATPase activity
(Davies et al., 1995
),
suggesting that cGMP signalling may regulate ion transport in tubules.
NO/cGMP signalling is also activated by a nitridergic family of
neuropeptides, capa, which comprise the only known insect NO/cGMP-mobilising
peptides. Capa-1 and capa-2 are encoded by the capa gene in
Drosophila (Kean et al.,
2002). Both capa-1 and capa-2, as well as the closely related
Manduca sexta CAP2b, induce diuresis and stimulate NO/cGMP
signalling and intracellular calcium increases in Drosophila tubule
principal cells (Davies et al.,
1995
,
1997
;
Kean et al., 2002
;
Rosay et al., 1997
). To date,
secretion by Aedes and Anopheles tubules has not been shown
to be stimulated by NO or cGMP, although the A. stephensi gene
encoding nitric oxide synthase (NOS) has been cloned
(Luckhart et al., 1998
); also,
A. gambiae tubules have been shown to express NOS transcripts
(Dimopoulos et al., 1998
). All
known insect NOS-encoding genes are very similar
(Davies, 2000
), resulting in
virtually identical sequences for NOS protein; as such, conservation of
function at the physiological level may be anticipated. Recently, data mining
of the A. gambiae genome has identified capa peptides in this species
(Riehle et al., 2002
).
Although capa-like signalling beyond the Diptera can be inferred from the
existence of the cardinal CAP2b in the Lepidoptera, other reports
have suggested that cGMP is antidiuretic in other insect orders, for example
Hemiptera (Quinlan et al.,
1997
) and Coleoptera (Eigenheer et al.,
2002
,
2003
), or that
CAP2b is without effect in Orthoptera
(Coast, 2001b
). It is thus of
great interest to assess the phylogenetic scope of the highly unusual
autocrine capa/NOS/NO/cGMP signalling model beyond Drosophila.
Furthermore, the application of knowledge of tubule function in D.
melanogaster to those of insect disease vectors will advance
understanding of tubule physiology in the context of specific cell types and
tubule regions in these animals.
Our results show that, whereas all insect tubules so far studied contain NOS (and thus have the machinery to respond to capa), only the dipteran species studied show functional responses. The scope of action of this peptide may thus be general within, but limited to, certain endopterygote orders.
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Materials and methods |
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Aedes aegypti
These were obtained as non-infective, sugar-water-fed adults from a colony
maintained by Professor E. Devaney, University of Glasgow. Female animals were
used upon receipt.
Anopheles stephensi and Anopheles gambiae
Non-infective, sugar-water-fed, adults were provided as a kind gift of Dr
L. Ranford-Cartwright, University of Glasgow. Female animals were used upon
receipt. If mosquitoes were not used immediately, they were maintained over a
12 h:12 h photoperiod at 55% humidity at 22°C, on 5% sucrose (v/v)
solution ad libitum for a maximum of 3 days before use in
experiments.
Glossina morsitans
Non-infective adults were provided by Dr S. Welburn, University of
Edinburgh, and by Professor D. Barry, University of Glasgow. Animals were used
immediately upon receipt.
Schistocerca gregaria
These were obtained from Bugs Direct (Well Cottages, Devon, UK) and either
used immediately or maintained on grass over a 12 h:12 h photoperiod at 55%
humidity at 22°C for a maximum of 34 days. All insects were
cold-anaesthetised and decapitated prior to dissection to isolate intact
tubules.
Peptides
Capa neuropeptides used in this study are shown in
Table 1. Of the
Drosophila capas, capa-1 (GANMGLYAFPRVamide) was used here, because
of its identical mode of action to, but slightly greater potency than, capa-2
(Kean et al., 2002). Both
A. gambiae capa peptides were synthesised: QGLVPFPRVamide
(AngCAPA-QGL) and GPTVGLFAFPRVamide (AngCAPA-GPT). All
peptides were synthesised by Invitrogen Corp. (Renfrew, UK). A.
gambiae capa peptides were identified by data mining the A.
gambiae genome. While this study was in progress, identical sequences for
Anopheles capa peptides were published elsewhere
(Riehle et al., 2002
).
|
Reverse-transcription (RT)-PCR for capa receptor
Analysis of capa receptor expression in dipteran tubules was carried out by
RT-PCR according to standard protocols
(Dow et al., 1994b) from cDNA
templates prepared from Drosophila melanogaster, Anopheles stephensi
and Anopheles gambiae tubules. The capa receptor has been identified
in D. melanogaster; searching the A. gambiae genome reveals
a possible candidate for the Anopheles capa receptor.
For each cDNA preparation, 20 tubules were dissected, poly(A)+ RNA extracted (Dynal mRNA direct kit; Dynal Biotech UK, Wirral, UK) and reverse transcribed with Superscript Plus (Gibco BRL, Invitrogen Ltd, Paisley, Renfrewshire, UK). 1 µl of the reverse transcription reaction was used as a template for PCR, containing the following gene-specific primer pairs: Drosophila capaR Forward, 5'-GCGGCCGCCTAAAATGAATTCATCGACCG-3'; Drosophila capaR Reverse, 5'-GTCTAGAGCCTCGTGCTTAAATACAAG-3'; putative A. gambiae capaR Forward, 5'-TGTTGACCGTGTTGAAGTGTTGC-3'; putative A. gambiae capaR Reverse, 5'-CTGTTCTTTGCCTTTCCAATGCTC-3'. Additionally, to control against genomic contamination in cDNA preps, primers that had been designed around intron/exon boundaries of the capa receptor gene were used. Use of such primers verified the cDNA quality used in PCR reactions. Further controls were performed that included nonreverse transcribed template (i.e. no cDNA).
PCR cycle conditions for reactions with Drosophila cDNA template were as follows: 93°C (3 min), 36 cycles of [93°C (30 s), 54.3°C (30 s), 72°C (1 min)] and 72°C (1 min).Conditions were similar for A. gambiae and A. stephensi cDNA templates except that the annealing temperature used was 59°C. PCR products obtained from such RT-PCR experiments were cloned using the Invitrogen Topoisomerase (TOPO TA Cloning) system (Renfrew, Scotland). Cloned plasmids were purified using Qiagen kits (Crawley, UK) and sequenced to confirm their identity.
Very few A. gambiae were available for study; thus, for all following experiments, A. stephensi was used.
Immunocytochemistry
Immunocytochemistry to fixed, intact tubules from all insect species was
performed using a universal anti-NOS (anti-uNOS) antibody according to
previously published protocols (MacPherson
et al., 2001), as described in the legend to
Fig. 2. The anti-uNOS antibody
is an affinity-purified rabbit universal anti-NOS antibody, used at 1:100
dilution and specified for Drosophila use (anti-uNOS; PA1-039;
Affinity BioReagents, via Cambridge BioScience, Cambridge, UK). This
antibody is directed against an epitope that is closely conserved in
mammalian, insect and even crustacean NOS peptides
(Table 2).
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This antibody has been used previously in Drosophila
(Broderick et al., 2003;
Dow and Davies, 2001
;
Gibbs and Truman, 1998
).
Specificity of this antibody has been demonstrated by immunoblotting DNOS
(Drosophila nitric oxide synthase) protein expression (
150 kDa
protein) in tubules from dNOS transgenic lines; overexpression of
DNOS results in increased protein by western analysis, which correlates with
increased NOS enzyme activity by direct assays. Also, this antibody has
successfully been used for immunocytochemistry in both wild-type and
dNOS transgenic Drosophila tubules and in eye tissue
(Broderick et al., 2003
;
Dow and Davies, 2001
;
Gibbs and Truman, 1998
).
Staining was visualised using a fluorescein-labelled goat anti-rabbit antibody
(Diagnostics Scotland, Edinburgh, UK), used at 1:250 dilution. In order to
visualise principal cell nuclei, tubules were counterstained in 1 µg
ml-1 4', 6'-diamidino-2-phenylindole hydrochloride
(DAPI; Sigma Aldrich, Gillingham, UK) for 2.5 min
(Broderick et al., 2004
).
Stained tubules were mounted in VectaShield (Vector Labs, Peterborough, UK).
Staining in whole-mount tubules was detected by immunofluorescence using an
Axiocam imaging system (Zeiss, Welwyn Garden City, UK).
NADPH diaphorase assay for NOS activity
An assay for NOS-associated NADPH diaphorase activity in
Drosophila tubule extracts
(Broderick et al., 2003;
Kean et al., 2002
) was
modified for analysis in 96-well plates. Intact tubules were dissected from
animals (lines as described in legend to
Fig. 3). For each species,
either six tubules (Drosophila), five tubules (Aedes and
Anopheles) or two tubules (Glossina and
Schistocerca) were used. For each sample, tubules were placed in 93
µl of 50 mmol l-1 Tris HCl, pH 7.4, 1% Triton X100 and 5 µl
of 10 mmol l-1 XTT
[2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide],
sodium salt in 96-well plates. Samples were incubated at 25°C for 20 min.
For peptide stimulations, either 1 µl of each peptide (D.
melanogaster capa-1, AngCAPA-QGL or AngCAPA-GPT to a
final concentration of 10-7 mol l-1) or 1 µl
phosphate-buffered saline (PBS; control) were added for a further 10 min. To
each sample, either 1 µl of 100 mmol l-1 NADPH or 1 µl PBS
(for no-substrate controls) was added, and samples were incubated at 25°C
for 7 min. For each species, replicate samples were prepared under the
following conditions: tubules only, tubules + NADPH, tubules + peptide,
tubules + peptide + NADPH. Samples were homogenised and colorimetric analysis
performed for all samples by spectrophotometry at 450 nm (Berthold Mithras
plate reader; Berthold Technologies, Redbourn, UK). Blanks were prepared from
incubation buffer. Blank samples, `tubules only' samples and `tubules +
peptide' samples gave very similar readings. The overall mean of readings for
blanks, `tubules only' samples and `tubules + peptide' samples within each
experiment was subtracted from results of `tubules + NADPH' and `tubules +
peptide + NADPH', respectively. In order to normalise data for all species,
results were expressed as % increase over unstimulated tubules (±
S.E.M.; N=46); i.e. (corrected values for `tubules
+ peptide + NADPH' minus mean of corrected values for `tubules + NADPH'/mean
of corrected values for `tubules + NADPH')x100%.
|
To further verify that stimulated NADPH diaphorase activity was due to NOS activation, we utilised the NOS inhibitor NG-nitro-L-arginine-methyl ester (L-NAME; Calbiochem, Beeston, UK) in the assays above. For these samples, L-NAME was added to samples prepared as described above, at a concentration of 2 µmol l-1 for 20 min prior to stimulation with capa peptides. Results were expressed as % change (± S.E.M.; N=4) of both peptide-stimulated samples and capa + L-NAME samples compared with controls (samples without either capa or L-NAME).
Tubule cGMP assays
Cyclic GMP levels were measured in pooled samples of tubules dissected from
insects, as detailed in Fig. 4
legend, by radioimmunoassay (Amersham Biotrak Amerlex M; Amersham Biosciences,
Chalfont St Giles, UK), as previously described
(Dow et al., 1994b). Tubules
were pre-incubated with the cGMP-specific phosphodiesterase inhibitor
Zaprinast (Calbiochem) at 10-5 mol l-1 for 10 min. For
peptide stimulations, either 1 µl of each peptide (D. melanogaster
capa-1, AngCAPA-QGL or AngCAPA-GPT to a final concentration
of 10-7 mol l-1) or 1 µl PBS (control) were added for
a further 10 min. Incubations were terminated with ice-cold ethanol and
homogenised. The ethanol was evaporated and samples were resuspended in 0.05
mol l-1 sodium acetate buffer (Amersham Biosciences) and processed
for cGMP content according to manufacturer's protocol. Data were normalised
across insect species by expressing results as fmol cGMP µg-1
protein (± S.E.M.; N=46). Protein
concentrations were determined by Bradford assay.
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Fluid secretion assays
Tubule secretion was measured according to standard procedures. Intact
Malpighian tubules were isolated into 9 µl drops of a freshly prepared
mixture of Schneider's medium (Gibco BRL, Invitrogen Ltd) and
Drosophila saline (1:1, v/v) under liquid paraffin, and fluid
secretion rates measured in tubules as detailed elsewhere
(Dow et al., 1994a). Briefly,
one end of the tubule was wrapped around a metal pin and the rest of the
tubule bathed on the saline drop. A nick was made near the ureter, a drop of
secreted fluid collected every 10 min, and the diameter measured using an
eyepiece micrometer. The volume of each droplet was calculated as
4/3
r3, where r is the radius of the droplet,
and secretion rates plotted against time. Secretion was measured under basal
conditions to establish a steady rate of secretion prior to stimulation with
peptide(s).
For other insect species, procedures were as for Drosophila, with appropriate modifications to accommodate the widely differing sizes of the tubules. However, apart from Drosophila, all insect tubules were left in the saline bubble in the paraffin dish for at least 15 min after dissection, then wound around the pin and left for another 15 min. Tubules were then nicked to allow bubbles to form; experimental readings commenced 10 min after this. Basal rates were measured for 30 min prior to stimulation with peptides (D. melanogaster capa-1, AngCAPA-QGL or AngCAPA-GPT).
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Results |
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A gene encoding a Drosophila capa receptor has been identified and
functionally characterised (Iversen et
al., 2002; Park et al.,
2002
). There is a single clear homologue for the CAPA-R in the
published Anopheles genome (BlastP; P=5x10-90). This
is encoded by a gene with GenBank no. XP_312952. At present, there are no
compelling matches in available Aedes or other insect sequences; the
nearest match in the Aedes genome has been annotated as a 5-HT7
receptor. Nonetheless, if a CAPA-R homologue were found to be expressed in the
tubule of another insect, it would strengthen the case for functional
conservation of capa signalling. By RT-PCR with intron-spanning primers, it
was possible to show that the Anopheles homologue was indeed
expressed in the Anopheles tubule
(Fig. 1), although the
non-degenerate Anopheles primers did not identify a match in the
Aedes tubule.
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NOS immunoreactivity in tubules across species
Capa acts on Drosophila tubules to activate NOS via
intracellular calcium: any nitridergic action of capa in other insects would
thus require the presence of NOS in tubules. Accordingly, the distribution of
NOS in the Malpighian tubules of other species was investigated by
immunocytochemistry for NOS (Fig.
2). A universal anti-NOS antibody was utilized for these
experiments; this antibody has been previously shown to be specific for
Drosophila NOS both in the eye
(Gibbs and Truman, 1998) and
in tubules by immunocytochemistry
(Broderick et al., 2003
;
Dow and Davies, 2001
) and by
western blotting (Broderick et al.,
2003
). Previous work has shown that NOS is expressed in only
principal cells of D. melanogaster tubules
(Broderick et al., 2003
;
Davies, 2000
). Here, we show
clear NOS immunoreactivity only in the main, fluid-transporting segment of the
tubule (Fig. 2Aii, region
marked `m').
In mosquito tubules, NOS immunoreactivity is observed only in the cytoplasm
of principal cells (examples of unstained stellate cells marked by arrows in
Fig. 2Bii,iii,Cii,iii). In both
A. aegypti and A. stephensi, counterstaining of cell nuclei
with DAPI shows the smaller nuclei of the stellate cells (arrows in
Fig. 2Biv,Civ), as in
Drosophila (Broderick et al.,
2004), of which the cytoplasm remains unstained. In contrast to
Drosophila, however, the entire length of the tubule is stained using
anti-NOS antibody in both mosquito species.
Fig. 2Dii,iii shows NOS
immunoreactivity in G. morsitans tubules. Interestingly, in this
dipteran species, staining with the anti-NOS antibody appears in all cells.
Close inspection of DAPI-stained G. morsitans tubules
(Fig. 2Div) does not reveal
cell nuclei of different sizes, as in D. melanogaster
(Broderick et al., 2004),
A. gambiae or A. stephensi
(Fig. 2Biv,Civ); it appears
that, unlike other Diptera, Glossina does not have obvious stellate
cells. Furthermore, staining is also observed throughout the tubule, rather
than merely in the main segment.
In the orthopteran out-group, S. gregaria, high background staining is observed in the control tubules (Fig. 2Ei). However, increased staining is observed with the anti-NOS antibody throughout the tubule (Fig. 2Eii) at the membrane and in the cytoplasm, suggesting that expression of NOS occurs in these tubules. This is consistent with a previous report of NAPDH diaphorase activity in orthopteran tubules (Locusta migratoria; M. Elphick, personal communication).
It is thus clear that all the insects studied have at least some of the machinery (NOS) to produce a nitridergic response to capa.
Capa peptides elevate NADPH diaphorase activity in dipteran tubules
NADPH diaphorase staining is an obligate correlate of NOS activity, both in
vertebrates and in insects (Elphick,
1997; Davies,
2000
). We have previously adapted this assay for measurements
in vitro (Kean et al.,
2002
), allowing quantification of NOS-associated NADPH diaphorase
activity, which accurately reflects NOS activity
(Broderick et al., 2003
).
Results in Fig. 3A show that
tubules stimulated with all capa peptides tested (i.e. D.
melanogaster capa-1, AngCAPA-QGL and AngCAPA-GPT)
increase NADPH diaphorase activity across the Diptera. Interestingly, capa-1
is at least as effective as, if not better than, the A. gambiae
peptides in raising NOS activity, at least at the concentration tested, which
was based on the maximum response of D. melanogaster tubules to
capa-1 as shown in previous work (Kean et
al., 2002). By contrast, although S. gregaria tubules
both contain NOS immunoreactivity and display similar resting levels of NADPH
diaphorase activity to dipteran tubules (results not shown), none of the capa
peptides tested elevated NADPH diaphorase activity in this orthopteran
species. In each case, L-NAME inhibited the increase in NADPH
diaphorase activity to control (unstimulated) levels, confirming the
association between increased NADPH diaphorase and NOS activation in these
species (Elphick, 1997
).
Capa peptides elevate cGMP in dipteran tubules
Previous work has shown that NO release, induced by M. sexta and
D. melanogaster capa peptides, increases cGMP content in
Drosophila tubules (Davies et al.,
1995; Kean et al.,
2002
). Radioimmunoassay for cGMP content showed that all three
capa peptides stimulated an increase in cGMP content in (only) dipteran
tubules (Fig. 4). G.
morsitans tubules were the most responsive to both AngCAPA-QGL
and AngCAPA-GPT. Also, while capa-1 was the most effective at
increasing tubule NOS activity (Fig.
3), this was not the case for the cGMP assay
(Fig. 4). Finally, capa
peptides do not increase cGMP in S. gregaria tubules [data in fmol
cGMP µg-1 protein (± S.E.M.; N=4):
unstimulated tubules: 0.097±0.003; capa-1 stimulated tubules,
0.093±0.008; AngCAPA-QGL, 0.096±0.008;
AngCAPA-GPT, 0.106±0.003].
Activation of NO/cGMP signalling by capa peptides increases fluid secretion
Previous work has shown that fluid secretion is potently stimulated by
capa-1 in D. melanogaster (Kean
et al., 2002); Fig.
5 shows such stimulation of fluid transport by D.
melanogaster tubules with capa-1 at an EC50 value of between
10-7 and 10-8 mol l-1. Capa-1 also stimulates
fluid transport by A. aegypti, A. stephensi and G. morsitans
tubules. However, still higher rates of secretion occur at very high
concentrations of peptide, between 10-3 (A. stephensi) and
10-4 mol l-1 (A. aegypti). Furthermore, G.
morsitans tubules are only stimulated to 50% over basal levels at all
concentrations of capa-1. In S. gregaria, capa-1 has either no
significant effect on secretion or is inhibitory (10-5,
10-6, 10-8 mol l-1). Similarly, capa-1 does
not stimulate fluid secretion by tubules from the dictyopteran roach
Periplaneta americana (data not shown). Thus, of the species sampled
to date, the stimulatory effects of capa-1 on tubules are confined to the
Diptera.
|
Figs 6, 7 show the first demonstration of the physiological effects of A. gambiae capa peptides on tubule fluid secretion in both mosquito and other Diptera. Drosophila tubule secretion is stimulated in a dose-dependent manner in response to AngCAPA-QGL, with an apparent EC50 of 10-5 mol l-1 (Fig. 6). All other dipteran tubules tested also respond to AngCAPA-QGL and are more sensitive to the peptide compared with Drosophila tubules, especially at low concentrations [10-6, 10-7, 10-8 mol l-1 (G. morsitans)]. Apart from at 10-3 mol l-1, tubules from A. aegypti and A. stephensi show similar responses at all concentrations tested. Also, G. morsitans tubules show a similar pattern of response to both mosquito species. At 10-7 mol l-1 AngCAPA-QGL, stimulation of secretion rates in A. aegypti, A. stephensi and G. morsitans is identical. By contrast, tubule secretion rates in S. gregaria tubules are not significantly altered at any concentration of AngCAPA-QGL.
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Similarly to capa-1 and AngCAPA-QGL, Drosophila tubules
respond to all concentrations of AngCAPA-GPT tested, although are
most responsive at concentrations of 10-5 mol l-1,
with the maximal response occurring at 10-3 mol l-1
(Fig. 7). However, responses of
all other dipteran tubules tested are similar at 10-5 and
10-6 mol l-1. Maximal response of A. aegypti
tubules occurs at 10-5 mol l-1, of A. stephensi
tubules at 10-6 mol l-1 and of G. morsitans
tubules at 10-7 mol l-1 AngCAPA-GPT.
Interestingly, no stimulation of secretion was observed with A.
aegypti tubules at 10-7 mol l-1. Note also that the
secretion response of these tubules to 10-6 mol l-1
AngCAPA-GPT is very low; these results are reproducible
(N>30). As with the other capa peptides, no significant response
is obtained from S. gregaria tubules.
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Discussion |
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In this work, we show that NOS immunoreactivity is observed in principal
cells throughout A. aegypti, A. stephensi and G. morsitans
tubules. By contrast, immunoreactivity for NOS is observed in all tubule cells
in S. gregaria. For these experiments, an anti-NOS antibody to an
epitope contained in all insect NOS sequences known to date was used
(Table 2). Although we cannot
assert that this antibody is specific to NOS alone, it faithfully reports
increased NOS expression via an inducible NOS transgene in
Drosophila tubules (Broderick et
al., 2003) and is consistent with other measures of NOS activity
reported here.
We also demonstrate that D. melanogaster and A. gambiae
capa peptides all stimulate NOS activity, increase cGMP production and elicit
an increase in fluid secretion rates in several dipteran species. Thus, this
suggests that not only are conserved features of the capa peptide sequences
functionally important but that conservation of the sequence and function of
the capa receptors must also exist within the Diptera. In particular, we have
identified a likely Anopheles homologue of the Drosophila
CAPA-R, which is abundantly expressed in Anopheles tubule.
Importantly, none of the capa peptides tested activate NO/cGMP signalling or
elevate fluid secretion in S. gregaria. Indeed, capa-1 may be
anti-diuretic at some concentrations (Fig.
5), although this is not linked to an increase in cGMP content
(Fig. 4). The data are
supported by work on L. migratoria, which shows that M.
sexta CAP2b does not affect fluid secretion by these tubules
(Coast, 2001b; see
Wegener et al., 2002
). We have
thus demonstrated, for the first time, physiological roles for A.
gambiae capa peptides and that capa-stimulated fluid secretion is
confined to a range of dipteran insects. We have also measured
neuropeptide-stimulated secretion rates in G. morsitans tubules for
the first time. Measurement of fluid secretion in the tsetse fly was first
published nearly 30 years ago (Gee,
1976a
,b
).
More recent work has re-visited cAMP-stimulated fluid secretion by G.
morsitans tubules (Isaacson and
Nicolson, 1994
). However, our recent development of
Glossina tubule physiology will allow study of a critical tissue in a
disease vector. The demonstration of conservation of capa signalling in
medically important insect vectors suggests new possibilities for novel
insecticide targets for pest control.
Importantly, we extend the phylogenetic scope of diuretic cGMP signalling
beyond Drosophila. It is apparent that cGMP can act as an
anti-diuretic signal in some insects. For example, in T. molitor, two
anti-diuretic hormones that act via cGMP have been isolated
(Eigenheer et al., 2002,
2003
). However, the existence
of anti-diuretic, cGMP-mobilising hormones in some insects need not point to a
universal mode of action by cGMP in insect tubules. Rather, this suggests a
critical distinction in the use of cGMP by different animals and, more than
that, a relevant role of cell or tissue concentration of cGMP in
physiology.
Locust tubules contain NOS but do not respond to capa. This result does
not, however, rule out nitridergic signalling in nondipteran tubules.
NOS-encoding genes have been characterized from multiple orders of insect
(Davies, 2000), and all contain
well-conserved calmodulin-binding domains, implying that, like
Drosophila NOS, they are calcium/calmodulin regulated. It is thus
probable that any neuropeptide that elevates calcium in Schistocerca
(or indeed any insect) tubule will activate NOS to generate NO. The capa
peptides perform such a role in Diptera, but our evidence suggests that they
do not in Orthoptera. Consistent with this argument, calcium has been shown to
be important in L. migratoria tubule stimulation by a partially
purified hormone, and cGMP has been shown to be diuretic
(Morgan and Mordue, 1985
). Of
course, the generation of NO in a tissue does not imply that it will be sensed
by soluble guanylate cyclase in the same tissue. In the future, it will be of
interest to follow the phylogenetic distribution of NO-sensing in insect
tubules, in particular those from nondipteran species, including orthopteran
insects.
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Acknowledgments |
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