Skip to main content

Thank you for visiting You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

CAPA neuropeptides and their receptor form an anti-diuretic hormone signaling system in the human disease vector, Aedes aegypti


Insect CAPA neuropeptides are homologs of mammalian neuromedin U and are known to influence ion and water balance by regulating the activity of the Malpighian ‘renal’ tubules (MTs). Several diuretic hormones are known to increase primary fluid and ion secretion by insect MTs and, in adult female mosquitoes, a calcitonin-related peptide (DH31) called mosquito natriuretic peptide, increases sodium secretion to compensate for the excess salt load acquired during blood-feeding. An endogenous mosquito anti-diuretic hormone was recently described, having potent inhibitory activity against select diuretic hormones, including DH31. Herein, we functionally deorphanized, both in vitro and in vivo, a mosquito anti-diuretic hormone receptor (AedaeADHr) with expression analysis indicating highest enrichment in the MTs where it is localized within principal cells. Characterization using a heterologous in vitro system demonstrated the receptor was highly sensitive to mosquito CAPA neuropeptides while in vivo, AedaeADHr knockdown abolished CAPA-induced anti-diuretic control of DH31-stimulated MTs. CAPA neuropeptides are produced within a pair of neurosecretory cells in each of the abdominal ganglia, whose axonal projections innervate the abdominal neurohaemal organs, where these neurohormones are released into circulation. Lastly, pharmacological inhibition of nitric oxide synthase (NOS) and protein kinase G (PKG) signaling eliminated anti-diuretic activity of CAPA, highlighting the role of the second messenger cGMP and NOS/PKG in this anti-diuretic signaling pathway.


Neuropeptides are central regulators of behaviours and control a plethora of physiological processes in all eukaryotic organisms. Insects, like many other animals, contain a comprehensive repertoire of neuropeptides along with their cognate receptors, which are essential for controlling complex biological phenomena including circadian rhythms, diapause, development, reproduction, pheromone biosynthesis, metabolism, circulation, stress, as well as hydromineral balance1,2,3,4,5,6,7,8,9,10. Insects have a high surface area to volume ratio, which has implications for their ability to maintain levels of water and ions within a normal homeostatic range. In order to ensure their survival, most insects have a relatively ‘simple’ excretory system comprised of the Malpighian ‘renal’ tubules (MTs) and hindgut (ileum and rectum). The MTs produce the primary urine acting to clear the haemolymph of excess ions, metabolites and toxins while the hindgut generally functions in reabsorptive processes eliminating any unintentional loss of essential ions and amino acids11,12. The insect excretory system is under complex control, which may include direct innervation and regulation by neurotransmitters such as proctolin, as observed in the hindgut of many insects13,14. The excretory system in insects is also under the control by various circulating hormones15,16, which is the sole mechanism of extrinsic control in the non-innervated MTs, while endocrine factors may also influence the hindgut4.

The overwhelming majority of studies investigating regulators of the insect excretory system have focused on diuretic regulators of the MTs17,18,19,20,21,22,23,24,25, with only a few studies characterizing factors responsible for controlling reabsorptive processes across hindgut epithelia12,26,27,28,29,30. In addition, a few anti-diuretic factors that inhibit primary urine secretion by the insect MTs have also been reported31,32,33,34,35,36, acting to counter the activity of the diuretic hormones that increase ion and water secretion rates. We recently identified an endogenous anti-diuretic hormone in the disease-vector mosquito, Aedes aegypti, that strongly inhibits select diuretic factors including the mosquito natriuretic peptide (a calcitonin-related diuretic hormone)37, which is critical for the post-prandial sodium-rich diuresis that follows blood gorging by adult females22. Similarly, anti-diuretic activity of CAPA neuropeptides has been reported earlier in larval A. aegypti36 as well as in other insects31,38,39,40,41,42, with signaling involving cGMP as a second messenger31,37,40,42,43. In addition to their clear anti-diuretic roles, CAPA peptides have also been linked to desiccation, where desiccation stress in Drosophila melanogaster leads to upregulation of capa mRNA, which is suggested to elevate CAPA levels in the CNS44. In many insects, CAPA peptides act through a conserved nitridergic signaling pathway leading to increased fluid secretion by MTs24,44. The mosquito anti-diuretic hormone is a member of the CAPA peptide family, which along with other insect PRXamide peptides, share homology to the vertebrate neuromedin U peptides45. CAPA neuropeptides are most abundant in specialized neurosecretory ventral abdominal (Va) neurons46,47,48,49 of the abdominal ganglia (or in the analogous neuromeres in insects with fused abdominal ganglia)50,51 and stored within abdominal perivisceral organs52,53,54,55, which are major neurohaemal organs facilitating neurohormone release into circulation for delivery to target organs expressing receptors.

In the present study, we utilized a combination of molecular tools, heterologous functional assays, physiological bioassays and reverse genetics techniques to identify and unravel the functional role of an anti-diuretic hormone receptor in the disease-vector mosquito, A. aegypti. Our data provides further evidence that mosquito CAPA neuropeptides, together with their cognate receptor identified herein, function in a neuroendocrine system halting the stimulatory activity of diuretic hormones that, if left unregulated, may compromise ion and water homeostasis in this important anthropophilic mosquito.


Anti-diuretic hormone receptor identification and sequence analysis

The complete CAPA receptor in A. aegypti was identified and found to be 3461 bp with an open reading frame of 2139 bp encoding a receptor protein of 712 residues. The 5′ and 3′ untranslated regions were comprised of 899 bp and 423 bp, respectively (Fig. S1A). The gene structure model revealed the cloned cDNA mapped to eleven exons spanning a genomic region of over 351 Kb, with the start codon positioned within the third exon and the translation termination (stop) codon located in the eleventh exon, which also contains the predicted polyadenylation signal at nucleotide position 3405–3410 (Fig. S1B). The deduced protein sequence encodes a receptor protein that displays the prototypical features of rhodopsin receptor-like (family A) GPCRs56,57,58, including the highly conserved tryptophan residue in the first extracellular loop involved in receptor trafficking, the D/E-R-Y/F motif at the border between the third transmembrane domain and second intracellular loop along with the NSxxNPxxY motif found within the seventh transmembrane domain (Fig. S1A). Phylogenetic analysis using maximum likelihood methods revealed the deduced receptor protein sequence shares greatest evolutionary relationship with the orthologous CAPA receptor proteins identified or predicted in other dipterans organisms, including for example the fruit fly, non-biting midges, house fly, blow fly along with the more closely-related sister mosquito species (Fig. S2).

Functional ligand-receptor interaction heterologous assay

The endogenous peptidergic ligands for the cloned anti-diuretic hormone receptor were identified using a heterologous functional assay using CHO-K1 cells stably expressing a bioluminescent calcium sensor, aequorin59,60. The receptor was activated by all endogenously expressed peptides encoded by the CAPA gene in A. aegypti (Fig. 1A), including two CAPA peptides (periviscerokinins) and a pyrokinin 1-related peptide. Notably however, the pyrokinin 1-related peptide displayed very poor activity compared to the two CAPA peptides, which were the most potent ligands with half maximal effective concentrations in the low nanomolar range (EC50 = 5.62–6.76 nM), whereas a significantly higher concentration of pyrokinin-1 was needed to achieve even low level CAPAr activation. Several other endogenous mosquito peptides as well as additional insect peptides belonging to distinct peptide families were tested and displayed no detectable activity over background levels of luminescence (Fig. 1B). Controls where the CHO-K1-aeq cells were transfected with empty pcDNA3.1+ vector showed no detectable luminescence response (data not shown) to any of the peptides used in this study, confirming the calcium-based luminescence signal was a result of CAPA neuropeptide ligands activating the transiently expressed A. aegypti CAPA receptor.

Figure 1

CAPA neuropeptide (anti-diuretic hormone) receptor (CAPAr) functional deorphanization using a heterologous assay. (A) Normalized dose-response curve demonstrating specificity of CAPAr functional activation by CAPA gene-derived neuropeptides. (B) Raw luminescent response following application of each CAPA gene-derived neuropeptide and representative neuropeptides belonging to several insect families, each tested at 10 μM. For peptide sequence information and species origin, see Table S3. Only CAPA gene-derived neuropeptides resulted in a significant luminescent response relative to BSA control (vehicle). At this saturating dose, no difference in response was observed between the two endogenous CAPA neuropeptides, AedaeCAPA1 and AedaeCAPA2; AedaePK1, demonstrated a significantly lower luminescent response (only ~20% activity compared to either CAPA peptide), but nonetheless this response was significantly higher compared to all other tested peptides that were identical to background luminescent responses obtained with vehicle control (BSA). Different letters denote bars that are significantly different from one another as determined by one-way ANOVA and Tukey’s multiple comparison post-hoc test (p < 0.01). Data represent the mean ± standard error (n = 3).

CAPAr transcript profile and cell-specific localization

We determined the developmental expression profile of the A. aegypti CAPA receptor (CAPAr) transcript in each post-embryonic developmental stage of the mosquito. Over the four larval stages and pupal stage of development, the CAPAr transcript level remained unchanged (Fig. 2A); however, in adults, CAPAr transcript levels were significantly higher in adult male mosquitoes compared to adult female, pupal stage and first instar larval mosquitoes (Fig. 2A). To confirm sites of biological action of the anti-diuretic hormones in vivo, we determined the CAPAr expression profile in adult A. aegypti, examining several tissues/organs in adult male and female mosquitoes. In males, CAPAr transcript was detected in reproductive tissues, head, carcass (i.e. the headless mosquito excluding the alimentary canal and reproductive tissues), and midgut while low background levels were found in the hindgut (Fig. 2B). Notably, CAPAr transcript was observed in the Malpighian ‘renal’ tubules (MTs) where expression was significantly enriched by ~150-fold compared to all other tissues/organs examined (Fig. 2B). A similar expression profile was observed in female mosquitoes with CAPAr transcript present in head, carcass, and midgut, while low background levels were detected in hindgut and reproductive organs. Similar to males, CAPAr was significantly enriched in the MTs of females relative to all other examined tissues/organs by nearly 150-fold (Fig. 2B). Using fluorescent in situ hybridization, the CAPAr transcript was localized specifically to principal cells of the MTs and absent in stellate cells (Fig. 2C). Specificity of CAPAr transcript localization was confirmed using sense control probe, with no signal detected in any cell type of the MTs (Fig. 2D).

Figure 2

Expression analysis of CAPAr transcript in the mosquito, A. aegypti. (A) Ontogenic expression profile of CAPAr transcript over post-embryonic stages of the A. aegypti mosquito shown relative to transcript levels in 1st instar larvae. (B) Spatial expression is analyzed in various tissues/organs from four-day old adult females, with transcript abundance shown relative to levels in the male midgut. (C) Cell-specific expression of CAPAr mRNA in principal cells (arrows) of MTs from adult female A. aegypti detected using an anti-sense probe, with no detection in the stellate cells (arrowheads). (D) No signal was detected in preparations hybridized with control CAPAr sense probe. All images acquired using identical microscope settings; scale bars in (C,D) are 100 μm.

CAPA transcript and mature neuropeptide immunolocalization within the abdominal ganglia

CAPA-like immunoreactivity was localized within all six of the abdominal ganglia, including the terminal ganglion. Specifically, each abdominal ganglion contains a pair of ventrally-localized strongly immunoreactive neurosecretory cells (Fig. 3A). Axonal projections from these CAPA-like immunoreactive neurosecretory cells emanate dorsally and anteriorly within each ganglion, exiting via the median nerve (Fig. 3B,C), with immunoreactive projections innervating the perivisceral organs (Fig. 3D), which are the primary neurohaemal release sites in the ventral nerve cord facilitating neurohormone delivery into the insect haemolymph61,62. Validation that these immunoreactive neurosecretory cells in the abdominal ganglia were indeed CAPA-producing neurons was established by co-localization of CAPA transcript with CAPA-like immunoreactivity. Weakly staining CAPA-like immunoreactive cells were also observed in other regions of the central nervous system, including the brain, suboesophageal ganglion and thoracic ganglia (Fig. S3); however, CAPA transcript was significantly enriched (~140-fold) only within the abdominal ganglia but not in other regions of the nervous system (Fig. S4). Within the abdominal ganglia, CAPA transcript co-localized within each pair of strongly staining CAPA-like immunoreactive neurosecretory cells (Fig. 3F–H). Preparations treated with CAPA transcript sense probes did not detect any cells in the abdominal ganglia nor anywhere else in the central nervous system.

Figure 3

Mapping of anti-diuretic hormone in the abdominal ganglia of the central nervous system and associated neurohaemal organs in adult A. aegypti. (A) Immunohistochemical distribution of CAPA neuropeptides in the abdominal ganglia (AG); specifically, a pair of highly immunoreactive neurosecretory cells within AG1–2 (A), AG3–4 (A’) and AG4–6 (A”). Higher magnification of AG3 (B) and AG4 (C) demonstrating CAPA immunoreactivity within large ventrally-positioned neurosecretory cells with axonal projections emanating dorsally within the ganglia and projecting anteriorly into the median nerve. (D) CAPA immunoreactivity in abdominal preparations with dorsal cuticle removed leaving the ventrally-localized AG within the abdominal segment showing immunoreactive processes innervating the abdominal neurohaemal (perivisceral) organs. CAPA transcript localization by fluorescent in situ hybridization revealing pairs of neurons within each AG including AG1 (E), AG2 (E’), AG3–4 (E”) and AG5 and the terminal abdominal ganglion (TAG; E”’). Co-localization of CAPA immunoreactivity (F) and CAPA transcript (G) was verified in all abdominal ganglia with representative preparation in (H) showing transcript and immunoreactivity co-detection and overlap. Scale bars: 200 µm (A,D), 100 µm (B,C,E) and 50 µm (F,H).

CAPAr knockdown abolishes anti-diuretic hormone activity

To confirm that the anti-diuretic hormone action of the CAPA neuropeptides36,37 are mediated through this specific receptor expressed within the principal cells of the MTs, one day-old female A. aegypti were injected with dsCAPAR to knockdown CAPAr transcript levels. Relative to control mosquitoes injected with dsARG (encoded ampicillin-resistance gene cloned from standard plasmid vector), CAPAr transcript was significantly reduced by ~75% in four-day old females (Fig. 4A). With significant knockdown verified in four-day old adult mosquito samples from the same experimental cohort, standard Ramsay assay was conducted as previously described37 on dsRNA-treated females to examine whether the anti-diuretic hormone activity of a CAPA anti-diuretic hormone was compromised. Our results confirmed that the CAPA neuropeptide, specifically AedaeCAPA-1, had no inhibitory activity against DH31-stimulated MTs in CAPAr knockdown females (Fig. 4B). In contrast, AedaeCAPA-1 potently inhibited DH31-stimulated fluid secretion by MTs in dsARG-treated control female mosquitoes (Fig. 4B).

Figure 4

RNA interference (RNAi) of CAPAr abolishes anti-diuretic activity of CAPA neuropeptide on adult female A. aegypti MTs. (A) Verification of significant knockdown (>75%) of CAPAr transcript in MTs of four-day old adult female A. aegypti by RNAi achieved through injection of dsCAPAr on day one post-eclosion. (B) Functional consequences of CAPAr knockdown demonstrating loss of anti-diuretic hormone activity by AedaeCAPA-1 against DromeDH31-stimulated fluid secretion by MTs. In (A), knockdown of CAPAr transcript was analyzed by one-tailed t-test (* denotes significant knockdown, p < 0.01). In (B), fluid secretion rates by MTs presented as mean ± SEM and analyzed by one-way ANOVA and Tukey’s multiple comparison post-test, where different letters denote treatments that are significantly different (p < 0.05, n = 14–33).

Effect of pharmacological blockade on the inhibition of fluid secretion

To further understand the anti-diuretic signaling pathway involving the CAPA neuroendocrine system, pharmacological blockers, including inhibitors of NOS (L-NAME) and PKG (KT5823), were tested against diuretic hormone-stimulated MTs alone and together with either AedaeCAPA-1 or cGMP. In DH31-stimulated MTs, L-NAME had no influence on the inhibitory effect of cGMP whereas the inhibitory effect of AedaeCAPA-1 was abolished (Fig. 5A). In 5HT-stimulated MTs, the results indicate that neither AedaeCAPA-1 nor cGMP inhibition are influenced by L-NAME (Fig. 5B). Application of KT5823 abolished the inhibitory effect of both cGMP and AedaeCAPA-1 in DH31-stimulated MTs (Fig. 6A). Similarly, the inhibitory activity of AedaeCAPA-1 and cGMP on 5HT-stimulated tubules was abolished when treated with KT5823 (Fig. 6B). Collectively, these results indicate that AedaeCAPA-1 inhibits select diuretic factors acting on the principal cells and involves NO and cGMP as a second messenger in DH31-stimulatd tubules, whereas cGMP, but not NO, is critical in the anti-diuretic activity of AedaeCAPA-1 on 5HT-stimulated MTs.

Figure 5

Effect of a nitric oxide synthase (NOS) inhibitor (L-NAME) on the anti-diuretic activity of AedaeCAPA-1 and cGMP in DromeDH31-stimulated A. aegypti MTs. The NOS inhibitor, L-NAME, was applied against (A) DromeDH31- and (B) 5HT-stimulated MTs alone or in the presence of either AedaeCAPA-1 or cGMP. Secretion rates are presented as mean ± SEM, n = 17–22. Columns that are significantly different from unstimulated controls are denoted with a distinct letter, as determined by a one-way ANOVA and Bonferroni post-test (p < 0.05).

Figure 6

Effect of a protein kinase G (PKG) inhibitor (KT5823) on the anti-diuretic activity of AedaeCAPA-1 and cGMP in DromeDH31-stimulated A. aegypti MTs. The PKG inhibitor, KT5823, was applied against (A) DromeDH31- and (B) 5HT-stimulated MTs alone or in the presence of either AedaeCAPA-1 or cGMP. Secretion rates are presented as mean ± SEM, n = 16–25. Columns that are significantly different from unstimulated controls are denoted with a distinct letter, as determined by a one-way ANOVA and Bonferroni post-test (p < 0.05).


Like many animals, insects must regulate the ionic and osmotic levels of their internal environment to ensure homeostatic levels of water and electrolytes are maintained. This is critical not only for challenges linked to feeding, including the intake of too much or too little water and/or ions, but is also important for daily exchange of these elements with the environment through other routes such as waste elimination or water loss during respiration. The insect excretory system acts to maintain hydromineral balance of the haemolymph by either increasing the removal of water and/or ions in excess or the recycling of these same elements when in short supply. The insect Malpighian ‘renal’ tubules (MTs) play a key role as the organ responsible for primary urine production, which can then be modified by downstream elements of the excretory system such as the hindgut4. The MTs are the chief iono- and osmoregulatory organ and are under rigorous control by neuroendocrine factors, including both diuretic hormones (DH) and anti-diuretic hormones (ADH), which regulate transepithelial movement of ions and osmotically obliged water. These hormones consist of a variety of peptides as well as other neurochemicals produced by neurosecretory cells in the brain and ventral nerve cord63,64. Classically, DHs stimulate primary urine secretion by the MTs, whereas ADHs increase fluid reabsorption from the hindgut15,65. However, countless studies in diverse insect species have established that ADHs can also act on the MTs to reduce primary urine secretion31,36,37,38,40,66. CAPA neuropeptides have been demonstrated to display potent anti-diuretic effects in a variety of insects38,39,42,67, including A. aegypti mosquitoes36,37, while they have been shown to function as DHs and ADHs in D. melanogaster40,68,69,70.

The current study provides definitive evidence supporting the importance of this anti-diuretic hormone system in the disease vector mosquito, A. aegypti, by characterization and functional deorphanization of an anti-diuretic hormone receptor that is highly enriched in the MTs and demonstrates high selectivity for the mosquito CAPA neuropeptides. Previous studies have functionally deorphanized a number of CAPA receptor orthologs in other insects including dipterans44,71,72,73, lepidopterans74, coleopterans75, hemipterans76, as well as in the southern cattle tick77. Here, we have functionally validated the specific ligands of the elusive A. aegypti CAPA receptor demonstrating that two of the peptides encoded by the mosquito CAPA gene50, Aedae-CAPA1 and –CAPA2, potently activate this receptor leading to calcium signaling that elicits a bioluminescent response. While none of the other tested ligands representing multiple insect peptide families were active on the mosquito CAPA receptor, the third peptide encoded by the CAPA gene, Aedae-PK1, had low agonist activity with a potency of over five orders of magnitude lower compared to the canonical CAPA ligands. Aedae-PK1 is a member of the pyrokinin-1 family of peptides that contain the GXWFGPRL-NH2 (where normally X = V, M or L) consensus C-terminal sequence and recently a revised tryptopyrokinin nomenclature has been adopted to differentiate these neuropeptides from distinct pyrokinin families78. In agreement with our findings, a subset of previous studies on insect CAPA receptor orthologs have shown minor responsiveness to tryptopyrokinin ligands, with high doses eliciting low level CAPA receptor activation74,75,76. Interestingly, this minor promiscuousness has not been observed for other dipteran CAPA receptors characterized previously71,72,73.

Members of the insect CAPA neuropeptide family are often also referred to as periviscerokinins due to their myotropic activity on visceral muscle and their source of release from the segmental abdominal neurohaemal organs known as perivisceral/perisympathetic organs31,51,75. Herein, we have immunolocalized CAPA neuropeptides within a pair of ventral neurosecretory cells within each of the six abdominal ganglia, whose axonal projections extend dorsally and anteriorly exiting each abdominal ganglion via the median nerve. CAPA immunoreactivity extends towards and is localized to the abdominal neurohaemal organs, the perivisceral organs, where these neuropeptides can be released into the haemolymph to elicit their neurohormonal actions on target sites expressing the CAPA receptor. The CAPA transcript was highly enriched within the abdominal ganglia of adult mosquitoes, confirming the transcript encoding the anti-diuretic hormone prepropeptide colocalized to these same neurosecretory cells. In support of these findings, peptidomic approaches using MALDI-TOF mass spectrometry have previously provided evidence for the presence of putative CAPA neuropeptides within isolated abdominal ganglia, including the terminal ganglion, from adult A. aegypti50. Collectively, these findings establish that the transcript and the mature peptide are present within the adult mosquito abdominal ganglia with the neurohormones being released into the insect circulatory system to act upon target tissues. Lastly, considering the low level CAPA transcript and immunoreactivity detected in other regions of the nervous system indicates that the abdominal ganglia, and their associated neurohaemal organs, are the primary source of the anti-diuretic hormone in adult A. aegypti. This also corroborates earlier peptidomic studies indicating the absence CAPA peptides, or differential processing of the CAPA precursor, in other regions of the nervous system aside from the abdominal ganglia where these neuropeptides are highly abundant50,79,80.

Having established the origin of the CAPA neuropeptide anti-diuretic hormones and their potent activity on the heterologously expressed CAPA receptor (CAPA-R), we next aimed to confirm the expression profile of the transcript encoding CAPA-R. Expression of CAPAr was observed in all post-embryonic ontogenic stages with significant enrichment in adult male mosquitoes, compared to females. Although the biological relevance of this differential expression remains unclear, this may relate to the sexual size dimorphism between adult male and female A. aegypti81, with the smaller males being inherently more susceptible to desiccation stress due to their higher surface area to volume ratio. In other insects, CAPAr transcript expression has been observed throughout most post-embryonic developmental stages72,75,82. The MTs are composed of two cell types forming a simple epithelium; large principal cells and thin stellate cells63. Principal cells facilitate the active transport of cations (Na+ and K+) into the lumen of the MTs from the haemolymph, while the stellate cells facilitate the transepithelial secretion of Cl, the predominant inorganic anion83. In adult stages, expression analysis of CAPAr verified significant enrichment of this receptor in the MTs in both male and female mosquitoes. Furthermore, cell-specific expression mapping confirmed that the CAPAr transcript is restricted to the principal cells of the MTs and absent in the smaller stellate cells. In other insects, CAPAr transcript has been detected in various regions of the alimentary canal75,76,84, including the principal cells of the MTs where this receptor is exclusively expressed in the fruit fly44. All in all, these observations are in line with physiological roles established for CAPA neuropeptides, which have been shown to modulate rates of fluid secretion by MTs in various insects36,37,41,62,85,86. In dipterans, these effects are mediated via action on the principal cells acting through a second messenger cascade involving calcium, nitric oxide and cGMP signaling24,87.

We next examined whether normal anti-diuretic hormone signaling, which requires the neuronally-derived CAPA peptide hormones activating their receptor expressed in the principal cells of the MTs, could be impeded by using RNA interference against the CAPAr transcript. One-day old mosquitoes were injected with CAPAr-targeted dsRNA resulting in knockdown at four-day old, where CAPAr transcript was significantly reduced. We examined whether CAPAr knockdown females retained sensitivity to CAPA peptides, which have been shown to inhibit fluid secretion by MTs by select diuretic hormones37. Indeed, CAPAr knockdown abolished the anti-diuretic activity of a CAPA neuropeptide against MTs stimulated with DromeDH31, an analog of mosquito natriuretic peptide. Collectively, through RNAi-mediated knockdown, these findings confirm that mosquito anti-diuretic hormones, which belong to the CAPA peptide family, are produced in pairs of neurosecretory cells in each of the abdominal ganglia whereby they are released through the neurohaemal organs and influence the MTs by acting on their receptor expressed within the principal cells of this organ. Further, the results confirm that sustained anti-diuretic hormone signaling, which requires the steady state expression of ligand and receptor, is necessary for facilitating the anti-diuretic control of the MTs.

In D. melanogaster and other dipterans, CAPA peptides have been shown to stimulate the nitric oxide (NO)/cGMP signaling pathway to induce diuresis87. When released, CAPA peptides bind to GPCRs found in principal (type I) cells of MTs, increasing Ca2+ levels in the cell through activation of L-type voltage gated calcium channels88. The influx of Ca2+ through these channels activates NOS, causing the production of NO, which subsequently activates guanylate cyclase to increase levels of cGMP in the MTs44. Ultimately, the activation of the NO/cGMP pathway stimulates the apical V-type H+-ATPase (proton pump), to increase fluid secretion in D. melanogaster. In the mosquito A. aegypti, CAPA peptides lead to activation of PKG, via elevated levels of cGMP36 and exogenous cGMP considerably inhibits fluid secretion rate37. Here, we sought to establish the roles of NO, cGMP and PKG on the anti-diuretic effects of CAPA peptides on adult mosquito MTs. Inhibitory doses of cGMP and a CAPA neuropeptide, namely AedaeCAPA-1, were treated with a NOS inhibitor, L-NAME, and a PKG inhibitor, KT5823. These investigations established that L-NAME did not alter the inhibitory effects of exogenous cGMP since this drug inhibits NOS, which is upstream of cGMP and, as a result, inhibition of DH31-stimulated secretion was unaffected. Contrastingly, AedaeCAPA-1 mediated inhibition of DH31-stimulated MTs was mitigated in the presence of L-NAME, reducing the anti-diuretic effects observed with AedaeCAPA-1. Comparatively, these findings are similar but are not identical to the effects of the PKG inhibitor, KT5823, which abolished the anti-diuretic activity of both AedaeCAPA-1 and cGMP, resulting in normal DH31-induced diuresis. Similar results were observed in 5HT-stimulted MTs with one exception; AedaeCAPA-1 inhibition appeared to be independent of NOS since L-NAME had no influence on the anti-diuretic activity of AedaeCAPA-1 in 5HT-stimulated MTs. Interestingly, the inhibition of both DH31- and 5HT stimulated diuresis by AedaeCAPA-1 and cGMP were sensitive to the PKG inhibitor, KT5823, which indicates that while some differences in signaling associated with inhibition of different diuretic hormones may occur, these inhibitory pathways likely converge and involve cGMP activating protein kinase G. Taken together, the findings in this study provide definitive evidence that CAPA peptides are anti-diuretic hormones in the mosquito A. aegypti, which inhibit fluid secretion of adult mosquito MTs through a signaling cascade involving the NOS/cGMP/PKG pathway. Further studies are necessary in mosquitoes as well as other insects to elucidate the differential regulation by DHs and ADHs given ample data supporting that cGMP and related effectors can be both stimulatory24,44,68,89 and inhibitory31,32,36,37,43,81,90 in their control on insect MTs. In conclusion, we have established an anti-diuretic hormone system in the adult mosquito A. aegypti providing evidence of a neural-renal axis whereby the neuropeptidergic anti-diuretic hormone is released by the abdominal segmental neurohaemal organs and subsequently targets their cognate receptor expressed within the principal cells of the MTs to counteract the activity of a subset of mosquito diuretic hormones. Fine-tuning of stimulatory and inhibitory hormones controlling the insect excretory system is of utmost importance to ensure overall organismal homeostasis to combat variable environmental conditions or feeding-related states that could perturb hydromineral balance if left unregulated.

Materials and Methods

Animals and dissections

Various stages of A. aegypti (Liverpool strain) were obtained from a laboratory colony maintained as described previously91. All mosquitoes were raised under a 12:12 light-dark cycle regime. Whole insects at each post-embryonic stage were used for examining developmental expression profiles and dissected tissues and organs were isolated from adults of each sex that were four-days post-eclosion. Adults were immobilized with brief exposure to carbon dioxide and then dissected to isolate individual organs using fine forceps (Fine Science Tools, North Vancouver, British Columbia, Canada) under nuclease-free Dulbecco’s phosphate-buffered saline (DPBS) at room temperature (RT).


The dissected tissues/organs were fixed overnight at 4 °C with 4% paraformaldehyde prepared in DPBS and were then washed several times with DPBS to remove fixative. The tissues were subsequently permeabilized in 4% Triton X-100, 10% normal sheep serum (NSS) and 2% bovine serum albumin (BSA) prepared in DPBS and incubated for 1 hour at RT on a rocking platform and then washed several times with DPBS to remove any traces of the permeabilization solution. The primary antibody was prepared using a custom affinity-purified rabbit polyclonal antibody (Genscript, Piscataway, NJ) produced against Rhodnius prolixus RhoprCAPA-2 (EGGFISFPRV-NH2; a kind gift from Prof. Ian Orchard, University of Toronto), which was diluted in 0.4% Triton X-100 containing 2% NSS and 2% BSA in DPBS. The stock antibody was diluted 1:1000 for stand-alone immunohistochemistry; however, when fluorescence in situ hybridization (FISH) preceded immunohistochemistry, the antibody was diluted 1:500 in the aforementioned solution. Tissues were incubated in the primary antibody solution for 48 hours at 4 °C on a rocking platform, and no primary controls were incubated in the same solution of 0.4% Triton X-100 containing 2% BSA and 2% NSS in DPBS, but lacking primary antibody. After the primary antibody incubation, tissues were washed three times for one hour each with DPBS at RT. The secondary antibody solution was prepared using either FITC-conjugated sheep anti-rabbit immunoglobulin G (Jackson ImmunoResearch Laboratories, West Grove, PA) or Alexa Fluor 488-conjugated cross-adsorbed goat anti-rabbit immunoglobulin G (Life Technologies, Burlington, ON) diluted 1:200 in DPBS containing 10% NSS. The tissues were incubated in the secondary antibody solution overnight at 4 °C on a rocking platform and were then washed with DPBS several times at RT. Tissues were mounted in ProLong Diamond Antifade Mountant containing DAPI (Molecular Probes, Eugene, OR) onto microscope slides and analyzed using a Lumen Dynamics XCite™ 120Q Nikon fluorescence microscope (Nikon, Mississauga, ON, Canada) or EVOS FL Auto Live-Cell Imaging System (Life Technologies, Burlington, ON).

Determination of the complete cDNA of an A. aegypti anti-diuretic hormone receptor

The Anopheles gambiae CAPA receptor identified previously71 was used as a query for Megablast screening of the A. aegypti genomic scaffold database available locally on a lab computer running Geneious® 6.1.8 (Biomatters Ltd, Auckland, New Zealand) and the highest scoring hits (mapping within supercontig1.1) were assembled and predicted introns were excised. Using the Primer3 module in Geneious® 6.1.8, gene-specific oligonucleotides targeting this region (see Table S1) were designed to amplify this predicted partial fragment using Q5 High Fidelity DNA Polymerase (New England Biolabs, Whitby, On) with whole adult female A. aegypti cDNA as template. The PCR product was purified, A-tailed, cloned into pGEM-T vector (Promega, Madison, WI, USA) and nucleotide sequence was confirmed by Sanger sequencing (Center for Applied Genomics, Hospital for Sick Children, Toronto, ON). After successful validation of the cloned partial sequence, primers were designed (as described above) to perform 5′ and 3′ rapid amplification of cDNA ends (RACE)-PCR utilizing the Clontech SMARTer 5′/3′ RACE Kit (Takara BIO USA Inc, CA, USA) as recently described60. To facilitate cloning of amplicons, the linker sequence GATTACGCCAAGCTT, which overlaps with the pRACE vector provided in the RACE kit, was added to the 5′ ends of the gene-specific primers (Table S1). First-strand cDNA synthesis was prepared using 1 μg total RNA from whole adult female mosquitoes using the 3′ CDS primer (provided in the kit) and a gene-specific reverse primer to generate template cDNA for 5′ RACE. Nested PCR reactions utilized gene-specific forward (3′ RACE) and reverse (5′ RACE) primers (see Table S1) and a universal primer mix (UPM) to amplify the complete cDNA encoding A. aegypti CAPAr with optimal cycling parameters determined empirically. Specifically, for both 5′ and 3′ RACE this included an initial denaturation at 94 °C for 1 min, followed by 40 cycles of 30 s at 94 °C, 30 s at 68 °C, and 3 min at 72 °C to amplify PCR products using SeqAmp DNA Polymerase. Following three rounds of nested PCR amplification using gene-specific primers, the amplicons were separated on a 1% agarose gel, extracted and cloned into the linearized pRACE vector. Plasmid DNA was isolated using a Monarch plasmid miniprep kit (New England Biolabs, Whitby, ON) and several clones were sent for sequencing for sequence validation. Finally, primers were designed at the 5′ and 3′ ends of the complete cDNA sequence (including UTRs) and used for final PCR amplification of the full receptor cDNA with Q5 High Fidelity DNA polymerase to confirm base pair accuracy.

Heterologous receptor functional activation bioluminescence assay

The open reading frame of the cloned A. aegypti CAPAr was inserted into pcDNA3.1+ mammalian expression vector following procedures described previously60,92,93. Using a recombinant CHO-K1 cell line stably expressing aequorin59, A. aegypti CAPAr was transiently expressed following growth and transfection conditions as reported recently60. Cells were harvested for the functional assay at 48 hours post-transfection by detaching cells from the culture flask using 5 mM EDTA in Dulbecco’s PBS (DPBS; Wisent Corp., St. Bruno, QC) and later cells were resuspended at a concentration of 106–107 cells/mL in assay media and incubated with coelenterazine h as described previously93. Prior to running the functional assay, cells were diluted 10-fold in assay media and left to incubate for one additional hour. Several endogenous as well as other insect neuropeptides representing a variety of neuropeptide families (see Table S2) were tested by preparing serial dilutions of each peptide in assay media. All peptides were commercially synthesized at a purity of >90% (Genscript, Piscataway, NJ) and 1 mM stock solutions were prepared by dissolving 1 mg of each peptide in water or DMSO as appropriate based on specific peptide characteristics. Recombinant CHO-K1 cells expressing the A. aegypti CAPAr were loaded into each well of multi-well plate using an automated injector module linked to a Synergy 2 Multi Mode Microplate Reader (BioTek, Winooski, VT) which measured kinetic luminescent response from each well for 20 sec immediately following cell loading onto the different peptides at various doses. Data was compiled in Microsoft Excel and analyzed in GraphPad Prism 8.0 (GraphPad Software, San Diego, CA).

RNA probe template preparation

To obtain a template for synthesizing DIG-labelled RNA probes for use in FISH, a 373 bp fragment of the A. aegypti CAPA partial mRNA (GenBank Accession: XM_001650839) previously described50 and a 743 bp product of the anti-diuretic hormone receptor identified herein with primers designed (see Table S3) using the Primer3 plugin in Geneious® 6.1.8 (Biomatters Ltd., Auckland, New Zealand) were amplified using standard Taq DNA Polymerase (New England Biolabs, Whitby, ON) following manufacturer- recommended conditions. PCR products were column-purified with PureLink Quick PCR Purification Kit (Life Technologies, Burlington, ON) and amplified in a subsequent PCR reaction to generate cDNA products with incorporated T7 promoter sequence (see Table S1) to facilitate in vitro RNA synthesis of anti-sense or sense probes. The final purified PCR products for use as templates for RNA probe synthesis were quantified on a SYNERGY 2 Microplate reader (Biotek, Winooski, VT).

Digoxigenin (DIG)-labelled RNA probe synthesis

PCR templates generated as described above (see Table S3) were used for in vitro transcription reactions using the HiScribe T7 RNA Synthesis Kit (New England Biolabs, Whitby, ON) following the recommended conditions when using modified nucleotides. Digoxigenin-labelled UTP was supplemented in a 35:65 ratio (DIG-UTP to standard UTP) either as a separate analog (digoxigenin-11-UTP) or in a pre-mixed 10× DIG-RNA labelling mix (Sigma-Aldrich, Oakville, ON). Template DNA was removed following treatment with RNase-free DNase I (New England Biolabs, Whitby, ON) and an aliquot of the synthesized RNA probes were then visually assessed using standard agarose gel electrophoresis and quantified on a SYNERGY 2 Microplate reader (Biotek, Winooski, VT).

Fluorescence in situ Hybridization (FISH)

An optimized FISH procedure based on a protocol described previously for R. prolixus94,95 was utilized involving peroxidase-mediated tyramide signal amplification to localize cells expressing either the CAPA peptide mRNA or the anti-diuretic hormone receptor (CAPAr) mRNA. Tissues/organs were dissected under nuclease-free Dulbecco’s phosphate-buffered saline (DPBS; Wisent, St. Bruno, QC) and were immediately placed in microcentrifuge tubes containing freshly-prepared fixation solution (4% paraformaldehyde prepared in DPBS) and fixed for 1–2 hours at RT or overnight at 4 °C on a rocker. Tissues/organs were subsequently washed five times with 0.1% Tween-20 in DPBS (PBT) and treated with 1% H2O2 (diluted in DPBS) for 10–30 minutes at RT to quench endogenous peroxidase activity. Tissues/organs were then incubated in 4% Triton X-100 (Sigma Aldrich, Oakville, ON) in PBT for 1 hour at RT to permeabilize the tissues and then washed with copious PBT. A secondary fixation of the tissues/organs was performed for 20 minutes in 4% paraformaldehyde in DPBS and then washed using PBT to remove all traces of fixative. The tissues/organs were then rinsed in a 1:1 mixture of PBT-RNA hybridization solution (50% formamide, 5× SSC, 0.1 mg/mL heparin, 0.1 mg/mL sonicated salmon sperm DNA and 0.1% Tween-20) which was then replaced with RT RNA hybridization that had been prepared earlier by denaturing in a boiling water bath for five minutes and subsequently cooled on ice for five minutes. The samples were then incubated at 56 °C for 1–2 hours, which served as the pre-hybridization treatment. During the pre-hybridization incubation, labelled RNA probe (anti-sense for experimental or sense for control) was added to pre-boiled RNA hybridization solution (2–4 ng/uL final concentration) and this mixture was heated at 80 °C for 3 minutes to denature the single-stranded RNA probes and then cooled on ice for 5 minutes. The samples were then incubated overnight in this hybridization solution containing the DIG-labelled RNA probe at 56 °C. The following day, samples were washed twice with fresh hybridization solution (minus probe) and subsequently with 3:1, 1:1 and 1:3 (vol/vol) mixtures of hybridization solution-PBT (all pre-warmed to 56 °C). The tissues were subsequently washed with PBT pre-warmed to 56 °C and in the final wash step were left to equilibrate to RT. Next, to reduce non-specific staining, samples were blocked with PBTB (DPBS, 0.1% Tween-20, 1% Molecular Probes block reagent; Invitrogen, Carlsbad, CA) for one hour. Tissues/organs were then incubated with a mouse anti-DIG biotin-conjugated antibody (Jackson ImmunoResearch Laboratories, West Grove, PA) diluted 1:400 and incubated for 1.5 hrs at RT on a rocker in the dark. The antibody solution was then removed and tissues were subjected to several washes in PBTB over the course of one hour. Tissues/organs were then incubated with horseradish peroxidase-streptavidin conjugate (Molecular Probes, Eugene, OR) diluted 1:100 in PBTB for 1 hour and the tissues were once again washed with PBTB several times over the course of an hour. Finally, prior to treatment with tyramide solution for the signal amplification of the target mRNA transcripts, samples were washed twice with PBT and once with DPBS. Afterwards, a tyramide solution was prepared consisting of Alexa Fluor 568 (or Alexa Fluor 647) tyramide dye in amplification buffer containing 0.015% H2O2. After experimenting with various dilutions of the labeled tyramide, a 1:100 and 1:500 dilution of tyramide dye gave optimal results with minimal background staining for the ganglia and MTs, respectively. After the last DPBS wash was removed from the tissues/organs, the tyramide solution was added and the tissues were incubated in the dark for 1 hour on a rocker at RT. The tyramide solution was then removed and the samples were washed with DPBS several times over the course of an hour. The tissues/organs were stored in DPBS overnight at 4 °C and then mounted on cover slips with mounting media comprised of DPBS with 50% glycerol containing 4 μg/mL 4ʹ,6- diamidino-2-phenylindole dihydrochloride (DAPI). For preparations involving transcript and neuropeptide co-detection in the nervous system, following the tyramide treatment, neural tissues were washed several times with DPBS and then incubated with primary antibody following the immunohistochemistry protocol described above. Tissues/organs were analyzed using a Lumen Dynamics XCite™ 120Q fluorescence microscope (Nikon, Mississauga, ON, Canada) or EVOS FL Auto Live-Cell Imaging System (Life Technologies, Burlington, ON).

Synthesis of dsRNA for RNA interference and RT-qPCR

Double-stranded RNA (dsRNA) was synthesized and column-purified using the MEGAscript® RNAi Kit (Invitrogen, Carlsbad, CA) following the recommended protocol using primers for dsCAPAr synthesis (see Table S3) and primers as reported previously for dsARG96, which is an ampicillin resistance gene cloned from standard sequencing plasmid (pGEM T-Easy) that served as a negative control. A Nanoject Nanoliter Injector (Drummond Scientific, Broomall, PA) was used to inject one-day old female mosquitoes with 1 μg (in ~140 nL) of either dsCAPAR or dsARG. After injection, mosquitoes were recovered in a photo period-, temperature- and humidity-controlled incubator. Total RNA was then isolated from four-day old whole female mosquitoes injected with dsCAPAr or dsARG using the Monarch Total RNA Miniprep Kit (New England Biolabs, Whitby, ON, Canada) and used as template (500 ng) for cDNA synthesis using the iScript™ Reverse Transcription Supermix (Bio-Rad, Mississauga, ON, Canada) following recommended guidelines diluting cDNA ten-fold prior to quantitative RT-PCR. AedaeCAPAr and AedaeCAPA transcript levels were quantified using gene-specific primers that were positioned on different exons (see Table S3) and PowerUP™ SYBR® Green Master Mix (Applied Biosystems, Carlsbad, CA, United States) and measured on a StepOnePlus Real-Time PCR System (Applied Biosystems, Carlsbad, CA, United States) following conditions described previously92. A similar procedure for cDNA synthesis and transcript quantification as outlined above was followed for total RNA isolated from each post-embryonic developmental stage and tissues/organs dissected from adult stage mosquitoes. Relative expression levels were determined using the ΔΔCt method and were normalized to the geometric mean of rp49 and rps18 reference genes, which were previously characterized and determined as optimal endogenous controls30. Measurements were taken from three biological replicates, all of which included three technical replicates per reaction and a no-template negative control.

Malpighian tubule fluid secretion assay

In order to determine fluid secretion rates, a modified Ramsay secretion assay85 was performed on isolated MTs of 3–6 day old adult female A. aegypti, as reported recently37. Tissue dissections were performed under physiological saline prepared as described previously86 diluted 1:1 with Schneider’s Insect Medium (Sigma-Aldrich, Oakville, ON). Individual MTs were removed and transferred to a Sylgard-lined Petri dish containing 20 μl saline bathing droplets immersed in hydrated mineral oil to prevent evaporation. The proximal end of the MT was removed from the bathing saline and wrapped around a Minuten pin to allow for secretion measurements. Dosages of 25 nmol l−1 DromeDH3122 or 100 nmol l−1 5HT97,98 alone or in combination with 1 fmol l−1 AedaeCAPA-137 were applied to the isolated MTs as previously described37. Dosage of AedaeCAPA-1 was based on a dose-response curve of AedaeCAPA-1 against DH31-stimulated tubules (Fig. S5), and the selected dose used to assess pharmacological blockade of prospective signaling partners was within the range of effective titres of AedaeCAPA-1. To investigate the effects of the pharmacological blockers, a nitric oxide synthase (NOS) inhibitor, Nω-Nitro-L-arginine methyl ester hydrochloride (L-NAME), and protein kinase G (PKG) inhibitor, KT5823, were used against 5-HT- and DH31-stimulated MTs. Dosages of 2 µmol l−1 L-NAME (manufacturer’s recommended dose) and 5 µmol l−1 KT582336 were applied to the MTs. The inhibitors were treated in conjunction with 1 fmol l−1 AedaeCAPA-1 and/or 100 nmol l−1 cyclic guanosine monophosphate, 8 bromo-cGMP (cGMP)37 (Sigma-Aldrich, Oakville, ON, Canada). Unstimulated controls consisted of tubules bathed in physiological saline with no diuretic application. Following a 60-minute incubation period, the size of the secreted droplet was measured using an eyepiece micrometer and fluid secretion rate (FSR) was calculated as described previously21.

Data availability

All data generated or analysed during this study are included in this manuscript [and its Supplementary information files].


  1. 1.

    Schoofs, L., De Loof, A. & Van Hiel, M. B. Neuropeptides as regulators of behavior in insects. Annu. Rev. Entomol. 62, 35–52 (2017).

    CAS  PubMed  Article  Google Scholar 

  2. 2.

    He, Q., Wu, B., Price, J. & Zhao, Z. Circadian rhythm neuropeptides in Drosophila: Signals for normal circadian function and circadian neurodegenerative disease. Int. J. Mol. Sci. 18, 886 (2017).

    PubMed Central  Article  CAS  PubMed  Google Scholar 

  3. 3.

    Terhzaz, S. et al. Insect capa neuropeptides impact desiccation and cold tolerance. Proc. Natl. Acad. Sci. USA 112, 2882–2887 (2015).

    ADS  CAS  PubMed  Article  Google Scholar 

  4. 4.

    Coast, G. M., Orchard, I., Phillips, J. E. & Schooley, D. A. Insect diuretic and antidiuretic hormones. Adv. Insect Phys. 29, 279–409 (2002).

    CAS  Article  Google Scholar 

  5. 5.

    Hillyer, J. F. Insect heart rhythmicity is modulated by evolutionarily conserved neuropeptides and neurotransmitters. Curr. Opin. Insect Sci. 29, 41–48 (2018).

    PubMed  Article  Google Scholar 

  6. 6.

    Gäde, G. Regulation of intermediary metabolism and water balance of insects by neuropeptides. Annu. Rev. Entomol. 49, 93–113 (2004).

    PubMed  Article  CAS  Google Scholar 

  7. 7.

    Nässel, D. R. & Winther, Å. M. E. Drosophila neuropeptides in regulation of physiology and behavior. Prog. Neurobiol. 92, 42–104 (2010).

    PubMed  Article  CAS  Google Scholar 

  8. 8.

    Raikhel, A. S. S., Brown, M. R. R. & Belles, X. Hormonal control of reproductive processes. Compr. Mol. Insect Sci. 3, 433–491 (2005).

    CAS  Article  Google Scholar 

  9. 9.

    Van Wielendaele, P., Badisco, L. & Vanden Broeck, J. Neuropeptidergic regulation of reproduction in insects. Gen. Comp. Endocrinol. 188, 23–34 (2013).

    PubMed  Article  CAS  Google Scholar 

  10. 10.

    Rafaeli, A. Pheromone biosynthesis activating neuropeptide (PBAN): regulatory role and mode of action. Gen. Comp. Endocrinol. 162, 69–78 (2009).

    CAS  PubMed  Article  Google Scholar 

  11. 11.

    Phillips, J. E. et al. Some major transport mechanisms of insect absorptive epithelia. Comp. Biochem. Physiol. A Comp Physiol 90, 643–650 (1988).

    CAS  PubMed  Article  Google Scholar 

  12. 12.

    Phillips, J. E. et al. Mechanisms and control of reabsorption in insect hindgut. Adv. Insect Phys. 19, 330–422 (1986).

    Google Scholar 

  13. 13.

    Cantera, R. & Nässel, D. R. Dual peptidergic innervation of the blowfly hindgut: a light- and electron microscopic study of FMRFamide and proctolin immunoreactive fibers. Comp. Biochem. Physiol. C. 99, 517–25 (1991).

    CAS  PubMed  Article  Google Scholar 

  14. 14.

    Steele, R. W., Lange, A. B., Orchard, I. & Starratt, A. N. Comparison of the myotropic activity of position-2 modified analogues of proctolin on the hindgut of Periplaneta americana and the oviduct of Locusta migratoria. J. Insect Physiol. 43, 931–938 (1997).

    PubMed  Article  Google Scholar 

  15. 15.

    Coast, G. The endocrine control of salt balance in insects. Gen. Comp. Endocrinol. 152, 332–338 (2007).

    CAS  PubMed  Article  Google Scholar 

  16. 16.

    O’Donnell, M. & Spring, J. Modes of control of insect Malpighian tubules: synergism, antagonism, cooperation and autonomous regulation. J. Insect Physiol. 46, 107–117 (2000).

    PubMed  Article  Google Scholar 

  17. 17.

    Baldwin, D. C., Schegg, K. M., Furuya, K., Lehmberg, E. & Schooley, D. A. Isolation and identification of a diuretic hormone from Zootermopsis nevadensis. Peptides 22, 147–152 (2001).

    CAS  PubMed  Article  Google Scholar 

  18. 18.

    Lehmberg, E. et al. Identification of a diuretic hormone of Locusta migratoria. Biochem. Biophys. Res. Commun. 179, 1036–1041 (1991).

    CAS  PubMed  Article  Google Scholar 

  19. 19.

    Furuya, K. et al. Cockroach diuretic hormones: characterization of a calcitonin-like peptide in insects. Proc. Natl Acad. Sci. USA 97, 6469–6474 (2000).

    ADS  CAS  PubMed  Article  Google Scholar 

  20. 20.

    Te Brugge, V., Paluzzi, J.-P., Schooley, D. A. & Orchard, I. Identification of the elusive peptidergic diuretic hormone in the blood-feeding bug Rhodnius prolixus: a CRF-related peptide. J. Exp. Biol. 214, 371–381 (2011).

    Article  CAS  Google Scholar 

  21. 21.

    Donini, A., O’Donnell, M. J. & Orchard, I. Differential actions of diuretic factors on the Malpighian tubules of Rhodnius prolixus. J. Exp. Biol. 211, 42–48 (2008).

    CAS  PubMed  Article  Google Scholar 

  22. 22.

    Coast, G. M., Garside, C., Webster, S. G., Schegg, K. M. & Schooley, D. A. Mosquito natriuretic peptide identified as a calcitonin-like diuretic hormone in Anopheles gambiae (Giles). J. Exp. Biol. 208, 3281–3291 (2005).

    CAS  PubMed  Article  Google Scholar 

  23. 23.

    Maddrell, S. H., Herman, W. S., Mooney, R. L. & Overton, J. A. 5-Hydroxytryptamine: a second diuretic hormone in Rhodnius prolixus. J. Exp. Biol. 156, 557–566 (1991).

    CAS  PubMed  Google Scholar 

  24. 24.

    Davies, S. A. et al. Signaling by Drosophila capa neuropeptides. Gen. Comp. Endocrinol. 188, 60–66 (2013).

    CAS  PubMed  Article  Google Scholar 

  25. 25.

    Cabrero, P. et al. The Dh gene of Drosophila melanogaster encodes a diuretic peptide that acts through cyclic AMP. J. Exp. Biol. 205, 3799–3807 (2002).

    CAS  PubMed  Google Scholar 

  26. 26.

    Audsley, N. & Phillips, J. E. Stimulants of ileal salt transport in neuroendocrine system of the desert locust. Gen. Comp. Endocrinol. 80, 127–137 (1990).

    CAS  PubMed  Article  Google Scholar 

  27. 27.

    Audsley, N., McIntosh, C. & Phillips, J. E. Actions of ion-transport peptide from locust corpus cardiacum on several hindgut transport processes. J. Exp. Biol. 173, 275–288 (1992).

    CAS  Google Scholar 

  28. 28.

    Audsley, N., Meredith, J. & Phillips, J. E. Haemolymph levels of Schistocerca gregaria ion transport peptide and ion transport-like peptide. Physiol. Entomol. 31, 154–163 (2006).

    CAS  Article  Google Scholar 

  29. 29.

    Audsley, N., Jensen, D. & Schooley, D. A. Signal transduction for Schistocerca gregaria ion transport peptide is mediated via both cyclic AMP and cyclic GMP. Peptides 41, 74–80 (2013).

    CAS  PubMed  Article  Google Scholar 

  30. 30.

    Paluzzi, J.-P., Vanderveken, M. & O’Donnell, M. J. The heterodimeric glycoprotein hormone, GPA2/GPB5, regulates ion transport across the hindgut of the adult mosquito, Aedes aegypti. PLoS One 9, e86386 (2014).

    ADS  PubMed  PubMed Central  Article  CAS  Google Scholar 

  31. 31.

    Paluzzi, J.-P. & Orchard, I. Distribution, activity and evidence for the release of an anti-diuretic peptide in the kissing bug Rhodnius prolixus. J. Exp. Biol. 209, 907–15 (2006).

    CAS  PubMed  Article  Google Scholar 

  32. 32.

    Massaro, R. C. et al. The mechanism of action of the antidiuretic peptide Tenmo ADFa in Malpighian tubules of Aedes aegypti. J. Exp. Biol. 207, 2877–2888 (2004).

    CAS  PubMed  Article  Google Scholar 

  33. 33.

    Laenen, B., De Decker, N., Steels, P., Van Kerkhove, E. & Nicolson, S. An antidiuretic factor in the forest ant: purification and physiological effects on the Malpighian tubules. J. Insect Physiol. 47, 185–193 (2001).

    CAS  PubMed  Article  Google Scholar 

  34. 34.

    Lavigne, C., Embleton, J., Audy, P., King, R. R. & Pelletier, Y. Partial purification of a novel insect antidiuretic factor from the Colorado potato beetle, Leptinotarsa decemlineata (Say) (Coleoptera: Chrysomelidae), which acts on Malpighian tubules. Insect Biochem. Mol. Biol. 31, 339–347 (2001).

    CAS  PubMed  Article  Google Scholar 

  35. 35.

    Eigenheer, R. A., Nicolson, S. W., Schegg, K. M., Hull, J. J. & Schooley, D. A. Identification of a potent antidiuretic factor acting on beetle Malpighian tubules. Proc. Natl Acad. Sci. USA 99, 84–89 (2002).

    ADS  CAS  PubMed  Article  Google Scholar 

  36. 36.

    Ionescu, A. & Donini, A. AedesCAPA-PVK-1 displays diuretic and dose dependent antidiuretic potential in the larval mosquito Aedes aegypti (Liverpool). J. Insect Physiol. 58, 1299–1306 (2012).

    CAS  PubMed  Article  Google Scholar 

  37. 37.

    Sajadi, F., Curcuruto, C., Al Dhaheri, A. & Paluzzi, J.-P. Anti-diuretic action of a CAPA neuropeptide against a subset of diuretic hormones in the disease vector Aedes aegypti. J. Exp. Biol. 221, (2018).

  38. 38.

    Quinlan, M. C., Tublitz, N. J. & O’Donnell, M. J. Anti-diuresis in the blood-feeding insect Rhodnius prolixus Stål: the peptide CAP2b and cyclic GMP inhibit Malpighian tubule fluid secretion. J. Exp. Biol. 200, 2363–2367 (1997).

    CAS  PubMed  Google Scholar 

  39. 39.

    Coast, G. M., Nachman, R. J. & Lopez, J. The control of Malpighian tubule secretion in a predacious hemipteran insect, the spined soldier bug Podisus maculiventris (Heteroptera, Pentatomidae). Peptides 32, 493–499 (2011).

    CAS  PubMed  Article  Google Scholar 

  40. 40.

    Rodan, A. R., Baum, M. & Huang, C.-L. The Drosophila NKCC Ncc69 is required for normal renal tubule function. Am. J. Physiol. Physiol. 303, C883–C894 (2012).

    CAS  Article  Google Scholar 

  41. 41.

    Coast, G. M. et al. Neurohormones implicated in the control of Malpighian tubule secretion in plant sucking heteropterans: The stink bugs Acrosternum hilare and Nezara viridula. Peptides 31, 468–473 (2010).

    CAS  PubMed  Article  Google Scholar 

  42. 42.

    Wiehart, U. I. M., Nicolson, S. W., Eigenheer, R. A. & Schooley, D. A. Antagonistic control of fluid secretion by the Malpighian tubules of Tenebrio molitor: effects of diuretic and antidiuretic peptides and their second messengers. J. Exp. Biol. 205, 493–501 (2002).

    CAS  PubMed  Google Scholar 

  43. 43.

    Quinlan, M. C. & O’Donnell, M. J. Anti-diuresis in the blood-feeding insect Rhodnius prolixus Stål: antagonistic actions of cAMP and cGMP and the role of organic acid transport. J. Insect Physiol. 44, 561–568 (1998).

    CAS  PubMed  Article  Google Scholar 

  44. 44.

    Terhzaz, S. et al. Mechanism and function of Drosophila capa GPCR: a desiccation stress-responsive receptor with functional homology to human neuromedinU receptor. PLoS One 7, e29897 (2012).

    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

  45. 45.

    Jurenka, R. The PRXamide neuropeptide signalling system. Conserved in animals. Adv. Insect Phys. 49, 123–170 (2015).

    Article  Google Scholar 

  46. 46.

    Gabilondo, H. et al. A targeted genetic screen identifies crucial players in the specification of the Drosophila abdominal Capaergic neurons. Mech. Dev. 128, 208–21 (2011).

    CAS  PubMed  Article  Google Scholar 

  47. 47.

    Gabilondo, H. et al. Segmentally homologous neurons acquire two different terminal neuropeptidergic fates in the Drosophila nervous system. PLoS One 13, e0194281 (2018).

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  48. 48.

    Santos, J. G., Pollák, E., Rexer, K.-H., Molnár, L. & Wegener, C. Morphology and metamorphosis of the peptidergic Va neurons and the median nerve system of the fruit fly, Drosophila melanogaster. Cell Tissue Res. 326, 187–199 (2006).

    CAS  PubMed  Article  Google Scholar 

  49. 49.

    Suska, A., Miguel-Aliaga, I. & Thor, S. Segment-specific generation of Drosophila Capability neuropeptide neurons by multi-faceted Hox cues. Dev. Biol. 353, 72–80 (2011).

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  50. 50.

    Predel, R. et al. Neuropeptidomics of the mosquito Aedes aegypti. J. Proteome Res. 9, 2006–2015 (2010).

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  51. 51.

    Predel, R. & Wegener, C. Biology of the CAPA peptides in insects. Cell. Mol. Life Sci. 63, 2477–2490 (2006).

    CAS  PubMed  Article  Google Scholar 

  52. 52.

    Eckert, M., Herbert, Z., Pollak, E., Molnar, L. & Predel, R. Identical cellular distribution of all abundant neuropeptides in the major abdominal neurohemal system of an insect (Periplaneta americana). J. Comp. Neurol. 452, 264–275 (2002).

    CAS  PubMed  Article  Google Scholar 

  53. 53.

    Pollak, E., Eckert, M., Molnar, L. & Predel, R. Differential sorting and packaging of capa-gene related products in an insect. J. Comp. Neurol. 481, 84–95 (2005).

    CAS  PubMed  Article  Google Scholar 

  54. 54.

    Wegener, C., Linde, D. & Eckert, M. Periviscerokinins in cockroaches: release, localization, and taxon-specific action on the hyperneural muscle. Gen. Comp. Endocrinol. 121, 1–12 (2001).

    CAS  PubMed  Article  Google Scholar 

  55. 55.

    Tublitz, N. J. & Truman, J. W. Identification of neurones containing cardioacceleratory peptides (CAPs) in the ventral nerve cord of the tobacco hawkmoth, Manduca sexta. J. Exp. Biol. 116, 395–410 (1985).

    CAS  PubMed  Google Scholar 

  56. 56.

    Schiöth, H. B. & Fredriksson, R. The GRAFS classification system of G-protein coupled receptors in comparative perspective. Gen. Comp. Endocrinol. 142, 94–101 (2005).

    PubMed  Article  CAS  Google Scholar 

  57. 57.

    Fredriksson, R., Lagerström, M. C., Lundin, L.-G. & Schiöth, H. B. The G protein-coupled receptors in the human genome form five main families. Phylogenetic analysis, paralogon groups, and fingerprints. Mol. Pharmacol. 63, 1256–1272 (2003).

    CAS  PubMed  Article  Google Scholar 

  58. 58.

    Rizzo, M. J., Evans, J. P., Burt, M., Saunders, C. J. & Johnson, E. C. Unexpected role of a conserved domain in the first extracellular loop in G protein-coupled receptor trafficking. Biochem. Biophys. Res. Commun. 503, 1919–1926 (2018).

    CAS  PubMed  Article  Google Scholar 

  59. 59.

    Paluzzi, J.-P. et al. Investigation of the potential involvement of eicosanoid metabolites in anti-diuretic hormone signaling in Rhodnius prolixus. Peptides 34, 127–34 (2012).

    CAS  PubMed  Article  Google Scholar 

  60. 60.

    Wahedi, A. & Paluzzi, J.-P. Molecular identification, transcript expression, and functional deorphanization of the adipokinetic hormone/corazonin-related peptide receptor in the disease vector, Aedes aegypti. Sci. Rep. 8, 2146 (2018).

    ADS  PubMed  PubMed Central  Article  CAS  Google Scholar 

  61. 61.

    Raabe, M. Synthesis and Release Sites of Neurohormones. in Recent Developments in Insect Neurohormones 1–68. (Springer US, 1989).

    Google Scholar 

  62. 62.

    Raabe, M., Cazal, M., Chalaye, D. & de Bessé, N. Action cardioaccélératrice des organes neurohémaux périsympathiques ventraux des quelques insectes. C. R. Acad. Sci. Hebd. Seances Acad. Sci. D. 263, 2002–2005 (1966).

    CAS  PubMed  Google Scholar 

  63. 63.

    Beyenbach, K. W. Transport mechanisms of diuresis in Malpighian tubules of insects. J. Exp. Biol. 206, 3845–3856 (2003).

    CAS  PubMed  Article  Google Scholar 

  64. 64.

    Coast, G. M. Neuroendocrine control of ionic homeostasis in blood-sucking insects. J. Exp. Biol. 212, 378–386 (2009).

    CAS  PubMed  Article  Google Scholar 

  65. 65.

    Phillips, J. Comparative physiology of insect renal function. Am. J. Physiol. 241, R241–57 (1981).

    CAS  PubMed  Google Scholar 

  66. 66.

    Eigenheer, R. A. et al. Isolation, identification and localization of a second beetle antidiuretic peptide. Peptides 24, 27–34 (2003).

    CAS  PubMed  Article  Google Scholar 

  67. 67.

    Paluzzi, J. P., Naikkhwah, W. & O’Donnell, M. J. Natriuresis and diuretic hormone synergism in R. prolixus upper Malpighian tubules is inhibited by the anti-diuretic hormone, RhoprCAPA-α2. J. Insect Physiol. 58, 534–542 (2012).

    CAS  PubMed  Article  Google Scholar 

  68. 68.

    Davies, S. A. et al. Neuropeptide stimulation of the nitric oxide signaling pathway in Drosophila melanogaster Malpighian tubules. Am. J. Physiol. 273, R823–7 (1997).

    ADS  CAS  PubMed  Google Scholar 

  69. 69.

    MacMillan, H. A. et al. Anti-diuretic activity of a CAPA neuropeptide can compromise Drosophila chill tolerance. J. Exp. Biol. (2018).

    PubMed  Article  Google Scholar 

  70. 70.

    Kean, L. et al. Two nitridergic peptides are encoded by the gene capability in Drosophila melanogaster. Am. J. Physiol. Integr. Comp. Physiol. 282, R1297–R1307 (2002).

    CAS  Article  Google Scholar 

  71. 71.

    Olsen, S. S., Cazzamali, G., Williamson, M., Grimmelikhuijzen, C. J. P. & Hauser, F. Identification of one capa and two pyrokinin receptors from the malaria mosquito Anopheles gambiae. Biochem. Biophys. Res. Commun. 362, 245–51 (2007).

    CAS  PubMed  Article  Google Scholar 

  72. 72.

    Iversen, A., Cazzamali, G., Williamson, M., Hauser, F. & Grimmelikhuijzen, C. J. Molecular cloning and functional expression of a Drosophila receptor for the neuropeptides capa-1 and -2. Biochem. Biophys. Res. Commun. 299, 628–633 (2002).

    CAS  PubMed  Article  Google Scholar 

  73. 73.

    Park, Y., Kim, Y.-J. J. Y.-J. & Adams, M. E. Identification of G protein-coupled receptors for Drosophila PRXamide peptides, CCAP, corazonin, and AKH supports a theory of ligand-receptor coevolution. Proc. Natl Acad. Sci. USA 99, 11423–11428 (2002).

    ADS  CAS  PubMed  Article  Google Scholar 

  74. 74.

    Shen, Z. et al. BNGR-A25L and -A27 are two functional G protein–coupled receptors for CAPA periviscerokinin neuropeptides in the silkworm Bombyx mori. J. Biol. Chem. 292, 16554–16570 (2017).

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  75. 75.

    Jiang, H., Wei, Z., Nachman, R. J., Adams, M. E. & Park, Y. Functional phylogenetics reveals contributions of pleiotropic peptide action to ligand-receptor coevolution. Sci. Rep. 4, 6800 (2014).

    ADS  CAS  PubMed  PubMed Central  Article  Google Scholar 

  76. 76.

    Paluzzi, J. P., Park, Y., Nachman, R. J. & Orchard, I. Isolation, expression analysis, and functional characterization of the first antidiuretic hormone receptor in insects. Proc. Natl. Acad. Sci. USA 107, (2010).

  77. 77.

    Yang, Y., Bajracharya, P., Castillo, P., Nachman, R. J. & Pietrantonio, P. V. Molecular and functional characterization of the first tick CAP2b (periviscerokinin) receptor from Rhipicephalus (Boophilus) microplus (Acari: Ixodidae). Gen. Comp. Endocrinol. 194, (2013).

  78. 78.

    Veenstra, J. A. The contribution of the genomes of a termite and a locust to our understanding of insect neuropeptides and neurohormones. Front. Physiol. 5, 454 (2014).

    PubMed  PubMed Central  Article  Google Scholar 

  79. 79.

    Predel, R. et al. Peptidomics of CNS-associated neurohemal systems of adult Drosophila melanogaster: a mass spectrometric survey of peptides from individual flies. J. Comp. Neurol. 474, 379–392 (2004).

    CAS  PubMed  Article  Google Scholar 

  80. 80.

    Wegener, C., Reinl, T., Jänsch, L. & Predel, R. Direct mass spectrometric peptide profiling and fragmentation of larval peptide hormone release sites in Drosophila melanogaster reveals tagma-specific peptide expression and differential processing. J. Neurochem. 96, 1362–1374 (2006).

    CAS  PubMed  Article  Google Scholar 

  81. 81.

    Wormington, J. D. & Juliano, S. A. Sexually dimorphic body size and development time plasticity in Aedes mosquitoes (Diptera: Culicidae). Evol. Ecol. Res. 16, 223–234 (2014).

    PubMed  PubMed Central  Google Scholar 

  82. 82.

    Graveley, B. R. et al. The developmental transcriptome of Drosophila melanogaster. Nat. 471, 473–479 (2011).

    ADS  CAS  Article  Google Scholar 

  83. 83.

    O’Donnell, M. J., Dow, J. A., Huesmann, G. R., Tublitz, N. J. & Maddrell, S. H. Separate control of anion and cation transport in Malpighian tubules of Drosophila melanogaster. J. Exp. Biol. 199, 1163–1175 (1996).

    PubMed  Google Scholar 

  84. 84.

    Chintapalli, V. R., Wang, J. & Dow, J. A. Using FlyAtlas to identify better Drosophila melanogaster models of human disease. Nat. Genet. 39, 715–720 (2007).

    CAS  PubMed  Article  Google Scholar 

  85. 85.

    Ramsay, J. A. Active transport of water by the Malpighian tubules of the stick insect, Dixippus morosus (Orthoptera, Phasmidae). J. Exp. Biol. 31, 104–113 (1954).

    CAS  Google Scholar 

  86. 86.

    Petzel, D. H., Berg, M. M. & Beyenbach, K. W. Hormone-controlled cAMP-mediated fluid secretion in yellow-fever mosquito. Am. J. Physiol. 253, R701–R711 (1987).

    CAS  PubMed  Google Scholar 

  87. 87.

    Pollock, V. P. et al. Conservation of capa peptide-induced nitric oxide signalling in Diptera. J. Exp. Biol. 207, 4135–4145 (2004).

    CAS  PubMed  Article  Google Scholar 

  88. 88.

    MacPherson, M. R. et al. L-type calcium channels regulate epithelial fluid transport in Drosophila melanogaster. Am. J. Physiol. Physiol. 280, C394–C407 (2001).

    CAS  Article  Google Scholar 

  89. 89.

    Davies, S. A. et al. CAP2b, a cardioacceleratory peptide, is present in Drosophila and stimulates tubule fluid secretion via cGMP. Am. J. Physiol. 269, R1321–6 (1995).

    CAS  PubMed  Google Scholar 

  90. 90.

    Ruka, K. A., Miller, A. P. & Blumenthal, E. M. Inhibition of diuretic stimulation of an insect secretory epithelium by a cGMP-dependent protein kinase. Am. J. Physiol. Ren. Physiol 304, F1210–6 (2013).

    CAS  Article  Google Scholar 

  91. 91.

    Rocco, D. A., Kim, D. H. & Paluzzi, J.-P. Immunohistochemical mapping and transcript expression of the GPA2/GPB5 receptor in tissues of the adult mosquito, Aedes aegypti. Cell Tissue Res. 369, 313–330 (2017).

    CAS  PubMed  Article  Google Scholar 

  92. 92.

    Gondalia, K., Qudrat, A., Bruno, B., Fleites Medina, J. & Paluzzi, J. P. Identification and functional characterization of a pyrokinin neuropeptide receptor in the Lyme disease vector, Ixodes scapularis. Peptides 86, 42–54 (2016).

    CAS  PubMed  Article  Google Scholar 

  93. 93.

    Oryan, A., Wahedi, A. & Paluzzi, J.-P. V. Functional characterization and quantitative expression analysis of two GnRH-related peptide receptors in the mosquito. Aedes aegypti. Biochem. Biophys. Res. Commun. 497, 550–557 (2018).

    CAS  PubMed  Article  Google Scholar 

  94. 94.

    Paluzzi, J. P., Russell, W. K., Nachman, R. J. & Orchard, I. Isolation, cloning, and expression mapping of a gene encoding an antidiuretic hormone and other CAPA-related peptides in the disease vector. Rhodnius prolixus. Endocrinol. 149, 4638–4646 (2008).

    CAS  Article  Google Scholar 

  95. 95.

    Paluzzi, J.-P. & Orchard, I. A second gene encodes the anti-diuretic hormone in the insect, Rhodnius prolixus. Mol. Cell Endocrinol. 317, 53–63 (2010).

    CAS  PubMed  Article  Google Scholar 

  96. 96.

    Durant, A. C., Chasiotis, H., Misyura, L. & Donini, A. Aedes aegypti Rhesus glycoproteins contribute to ammonia excretion by larval anal papillae. J. Exp. Biol. 220, 588–596 (2017).

    PubMed  Article  Google Scholar 

  97. 97.

    Clark, T. M. & Bradley, T. J. Additive effects of 5-HT and diuretic peptide on Aedes Malpighian tubule fluid secretion. Comp. Biochem. Physiol. - A Mol. Integr. Physiol. 119, 599–605 (1998).

    Article  Google Scholar 

  98. 98.

    Veenstra, J. A. Effects of 5-hydroxytryptamine on the Malpighian tubules of Aedes aegypti. J. Insect Physiol. 34, 299–304 (1988).

    CAS  Article  Google Scholar 

Download references


The authors are greatly appreciative of the CAPA antibody used in this study that was provided by Prof. Ian Orchard (University of Toronto Mississauga). This research was funded by a Natural Sciences and Engineering Research Council of Canada (NSERC) Discovery Grant and an Ontario Ministry of Research, Innovation, and Science Early Researcher Award to J-PP. FS received an NSERC CGS-M and Vernon Stong Scholarship in support of her graduate research. AL received an NSERC CGS-M Scholarship.

Author information




F.S. and J.-P.P. conceived the study. F.S. wrote the initial draft manuscript. F.S. collected the majority of the data while A.W., A.L., L.T.B., A.M., A.U. and C.P. contributed some data for a subset of the experiments. All authors contributed towards the editorial review and finalization of the manuscript granting approval to submit the study for publication.

Corresponding author

Correspondence to Jean-Paul V. Paluzzi.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Sajadi, F., Uyuklu, A., Paputsis, C. et al. CAPA neuropeptides and their receptor form an anti-diuretic hormone signaling system in the human disease vector, Aedes aegypti. Sci Rep 10, 1755 (2020).

Download citation

Further reading


By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.


Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing