Expression profile of genes encoding allatoregulatory neuropeptides in females of the spider Parasteatoda tepidariorum (Araneae, Theridiidae)

Authors: Marta Katarzyna Sawadro aff001;  Agata Wanda Bednarek aff001;  Agnieszka Ewa Molenda aff001;  Agnieszka Izabela Babczyńska aff001
Authors place of work: Department of Animal Physiology and Ecotoxicology, University of Silesia in Katowice, Bankowa, Katowice, Poland aff001
Published in the journal: PLoS ONE 14(9)
Category: Research Article
doi: 10.1371/journal.pone.0222274


Allatoregulatory neuropeptides are multifunctional proteins that take part in the synthesis and secretion of juvenile hormones. In insects, allatostatins are inhibitors of juvenile hormone biosynthesis in the corpora allata while allatotropins, act as stimulators. By quantitative real-time PCR, we analyzed the gene expression of allatostatin A (PtASTA), allatostatin B (PtASTB), allatostatin C (PtASTC), allatotropin (PtAT) and their receptors (PtASTA-R, PtASTB-R, PtASTC-R, PtAT-R) in various tissues in different age groups of female spiders. In the presented manuscript, the presence of allatostatin A, allatostatin C, and allatotropin are reported in females of the spider P. tepidariorum. The obtained results indicated substantial differences in gene expression levels for allatoregulatory neuropeptides and their receptors in the different tissues. Additionally, the gene expression levels also varied depending on the female age. Strong expression was observed coinciding with sexual maturation in the neuroendocrine and nervous system, and to a lower extent in the digestive tissues and ovaries. Reverse trends were observed for the expression of genes encoding the receptors of these neuropeptides. In conclusion, our study is the first hint that the site of synthesis and secretion is fulfilled by similar structures as observed in other arthropods. In addition, the results of the analysis of spider physiology give evidence that the general functions like regulation of the juvenile hormone synthesis, regulation of the digestive tract and ovaries action, control of vitellogenesis process by the neuropeptides seem to be conserved among arthropods and are the milestone to future functional studies.


Biology and life sciences – Biochemistry – Neurochemistry – Neurochemicals – Neuropeptides – Hormones – Peptide hormones – Neuroscience – Genetics – Gene expression – Anatomy – Nervous system – Reproductive system – Ovaries – Organisms – Eukaryota – Animals – Invertebrates – Arthropoda – Arachnida – Spiders – Insects – Medicine and health sciences – Research and analysis methods – Database and informatics methods – Bioinformatics – Sequence analysis – Sequence motif analysis


Neuropeptides represent the largest class of signal compounds, which also include steroids and sesquiterpenoids. Based on protein sequence similarity and main physiological functions, insect neuropeptides are grouped into families. One of the largest neuropeptide family comprise the allatoregulatory neuropeptides [1, 2, 3].

Allatoregulatory neuropeptides are multifunctional proteins with conserved protein sequences [4]. They are commonly found in insects from which they were isolated for the first time. They have also been identified in various groups of arthropods as well as in non-arthropod groups such as annelids and mollusks: Cancer borealis [5], Carcinus maenas [6], Triops newberryi [7], Penaeus monodon [8], Homarus americanus [9], Deroceras reticulatum [10, 11, 12]. Numerous manuscripts report demonstrating allatoregulatory reactivity in basal metazoans [13]. In addition to conservation of protein sequence, this further indicates, that allatoregulatory signaling might be conserved among metazoans, and thus it might be an ancestral mechanism. Insects are often regarded as a reference group for studies of allatoregulatory peptides in other groups of arthropods due to in insects allatoregulatory neuropeptides have been described in most detail so far. Based on these studies in insects, one of the primary roles of allatoregulatory neuropeptides is the regulation of the juvenile hormones (JHs) biosynthesis. JHs in insects (and methyl farnesoate–a standard product in JH synthesis in insects, serve different functions in arthropods [14, 15]) control crucial processes such as molting, metamorphosis, ovary development, vitellogenesis, mating behavior, and oviposition. Recent studies also confirmed the role of JHs in morphogenesis and caste determination in social insects [16]. Therefore, allatoregulatory neuropeptides indirectly affect these JH-dependent processes.

Allatostatins (ASTs) have an inhibitory effect, whereas allatotropins (ATs) stimulate biosynthesis and release of the JHs by the corpora allata (CA) [2, 17, 18, 19, 20]. ASTs are a superfamily of invertebrate neuropeptides that were initially defined by their action as inhibitors of JH biosynthesis in vivo. To date, more than 60 types of ASTs have been isolated and characterized from a variety of insect species. These peptides can be classified into three groups: the FGL-allatostatin (A type), the W(X)6W allatostatin (B type), and the lepidopteran (Manduca sexta) allatostatin (C type) [19] while allatotropins are not divided into subgroups. They were first isolated from M. sexta (Manse-AT), and their sequence (GFKNVEMMTARGF-NH2) was confirmed based on cDNA and was characterized in other insects like Spodoptera frugiperda as well as Aedes aegypti [11].

Among arachnids, ASTs have been identified in ticks. A-type ASTs were detected in the synganglion of Dermacentor variabilis [21], whereas AST C (Manse-AST) was described in the central nervous system of Ixodes scapularis [22]. Despite the detection of these peptides in ticks, their role has not been revealed. Furthermore, there are only a few publications that are based on the immunohistochemical data, AST A protein localization in the central nervous system of the model spider species Cupiennius salei [23]. However, the quoted data do not provide information on the role of ASTs and AT in spiders (compare with [11]).

In insects, in which allatoregulatory neuropeptides are relatively well-known in comparison with other arthropods, neuropeptides are produced and secreted in the central nervous system by neurosecretory cells and interneurons [2, 17, 18, 19, 20]. The major neurosecretory systems of insects are located behind the central nervous system and include corpora cardiaca (CC) and CA glands together with their connections. The CC, are neurohaemal organs and are among the most essential structures of the neuroendocrine system of insects. They stores and release the neuropeptides which are synthesized in the neuroendocrine cells of the brain. This importance is further highlighted by the ability of the CC glands to produce certain peptides from specific neurosecretory cells themselves. Apart from the CC, CA glands are also responsible for releasing the neuropeptides in the neuroendocrine system in insects, but their primary function is the synthesis of the juvenile hormones [2].

The neuroendocrine system of insects regulates most critical metabolic, behavioral, homeostatic, developmental, and reproductive processes [1, 24], which are widely studied in spiders because they are among the most abundant invertebrate predators in terrestrial ecosystems, agroecosystems, and woodlands [25, 26, 27]. Moreover, spiders are crucial in the development of efficient, sustainable, low-input agricultural systems [28]. However, how such important spider behaviors as web building, molting, sexual maturation, and complex behaviors like parental care are controlled on the neurohormonal level is still insufficiently studied. Unfortunately, knowledge about the function of the neuroendocrine and nervous systems of spiders is still rudimentary. Many of the quoted publications refer to research conducted in the 1960s [29, 30, 31, 32, 33].

It seems that the neuroendocrine and nervous systems of spiders may be the site of neuropeptide synthesis. Bonaric [32] described the neuroendocrine complex (Schneider I organs + neurohemal organs) as the CC homolog and Schneider II organs as the structure homologous to insect CA. Loesel et al. [23] also confirmed that in addition to the neuroendocrine system in C. salei, other nervous neurosecretory cells might be responsible for the secretion of AST. In addition, the neurohemal organ (tropfenkomplex) was identified as another secretory organ, which stores substances produced in the central nervous system and transported through the Schneider I organ [32]. Since neuropeptides are neurotransmitters, it can be assumed that they are secreted into the hemolymph by exocytosis and are transported to target cells where they bind to specific receptors [34]. Therefore, to analyze the function of allatoregulatory neuropeptides, it is thus first necessary to identify their tissue of synthesis as well as receptor location.

The main aims of this study, which is the first attempt to investigate spiders allatoregulatory neuropeptides, were to (i) detect the presence of these substances in the model spider species Parasteatoda tepidariorum C. L. Koch, 1841 (Araneae, Theridiidae) and to (ii) identify the tissues where the synthesis takes place and target sites of their action by the determination of expression of genes encoding allatostatin A (AST A), allatostatin B (AST B), allatostatin C (AST C) and allatotropin (AT) as well as their receptors in various tissues of female spiders. In addition, age-dependent expression rates were investigated.

Material and methods

Spider breeding

P. tepidariorum females were obtained, from laboratory-bred strains of the Department of Animal Physiology and Ecotoxicology, University of Silesia. The animals were bred at 25 ± 1°C at 70% relative humidity under L: 16 h, D: 8 h photoperiodic cycle. They were fed with laboratory cultured Drosophila melanogaster or D. hydei and were irrigated regularly.

For the needs of the presented study, the following ontogenetic stages of P. tepidariorum females were selected: the penultimate nymphal stage (35th day of life), the last nymphal instar (38th day of life), mature females (40th day of life), females after mating (43rd day of life) and females after oviposition (47th day of life). The spider development time was counted from leaving the cocoon, according to Miyashita [35] and behavioral observations.

Sample preparation

Five different tissues and organs of P. tepidariorum–the midgut glands with midgut (MG), neuroendocrine and nervous system (NS), ovaries (OV), hindgut (HG), and integument (INT)–were used for the analysis. Moreover, the entire body of adult spiders (EB) was used as a biological material. The number of tissues/organs per sample was determined during a pilot study and varied depending on the size and age of spiders (Table 1). The preliminary results were based on the analysis of the weight of tissues needed to obtain the proper concentration of total RNA, which was used to prepare a cDNA template in the reverse transcription process. The required weight of tissues/organs is correlated with the number of dissected individuals. Due to the efficiency of reverse transcription (80%), a minimum of 700 ng/μl of total RNA had to be isolated. For example, in the case of females in the penultimate nymphal stage (day 35 of life), due to the small size of the ovaries, it was necessary to dissect the ovaries from 20 spiders to obtain 700 ng/μl of total RNA (Table 1).

Tab. 1.

The number of tissues/organs selected for detection of allatoregulatory neuropeptides in the spider P. tepidariorum depending on their stages of ontogenesis.

<h2>The number of tissues/organs selected for detection of allatoregulatory neuropeptides in the spider <i>P</i>. <i>tepidariorum</i> depending on their stages of ontogenesis.</h2>

The tissues and organs were dissected on ice in sterile phosphate-buffered saline (PBS, 137 mM NaCl, 10 mM phosphate buffer (K2HPO4, KH2PO4), 2.7 mM KCl, pH 7.4). All tissues and organs were immediately frozen in liquid nitrogen and stored at -70°C until use. All samples were performed in six replicates.

In silico search

AST A, AST B, AST C, and AT and their receptor protein sequences of different insects (see S1 Table for species and accession number from the NCBI protein database [36]) were used to search homologous proteins in P. tepidariorum. The IAAA00000000.1 (TSA 25-JUL-2015) version of the current Transcriptome Shotgun Assembly database was searched with default parameters of tblastn [37] and the candidates with based on the sequence homology criterion that was proposed by Pearson [38] were identified.

The transcript sequences that were selected this way have been searched for the presence of ORFs and then transcribed into protein sequences using the Geneious (ver. 9.1.2) software. These sequences were used to confirm the identity of the putative neuropeptides and neuropeptide receptor genes in P. tepidariorum with gene sequences of other arthropods. For this purpose, the Geneious, Clustal Omega and InterProScan [39] software and algorithms were used. Primers for the further analysis of the section of putative genes encoding the neuropeptides and neuropeptide receptors in P. tepidariorum were designed using the Geneious software (Table 2).

Tab. 2.

Forward and reverse sequences for the PtASTA, PtASTA-R, PtASTB, PtASTB-R, PtASTC, PtASTC-R, PtAT, PtAT-R and PtRP49 primers.

<h2>Forward and reverse sequences for the Pt<i>ASTA</i>, Pt<i>ASTA-R</i>, Pt<i>ASTB</i>, Pt<i>ASTB-R</i>, Pt<i>ASTC</i>, Pt<i>ASTC-R</i>, Pt<i>AT</i>, Pt<i>AT-R</i> and Pt<i>RP49</i> primers.</h2>

Possible gene duplication was checked by performing multiple sequence alignment of chosen sequences in Clustal Omega and their pairwise comparison in BLAST software. The criterion of duplicated gene proposed by Ouedraogo et al. [40] was used. Phylogenetic tree was constructed with MEGA X [41] using the Neighbor-Joining method [42] with a bootstrap of 2000 replicates [43] The tree is drawn to scale, with branch lengths in the same units as those of the evolutionary distances used to infer the phylogenetic tree (S1S13 Figs)).

The bioinformatics analysis of the P. tepidariorum genome has indicated that some of the tested neuropeptides and their receptors have paralogs. Therefore, the data about gene duplication have not been placed in the manuscript and have been only used in the primer designing.

RNA isolation and cDNA preparation

Total RNA was isolated from the tissues/organs using TRIzol Reagent (Invitrogen, Carlsbad, CA, USA) according to the standard method [44]. Remaining genomic DNA contamination was removed using the TURBO DNA-free kit (Ambion, Austin, TX, USA) according to the manufacturer’s protocol. Quality of RNA was assessed for using a NanoDrop 2000 spectrometer (Thermo Scientific, Wilmington, USA). Aliquots of 1 μg of total RNA were retrotranscripted using the Reverse Transcription System (Promega, Madison, WI, USA) and random primers according to the recommendations of the manufacturer. The reverse transcription products were then diluted with 80 μl of nuclear-free water. Prepared cDNA was used as the qPCR template.

Quantitative real-time PCR (qPCR)

Relative expression of selected genes was quantified by real-time PCR performed in the LightCycler 480 System (Roche, Basel, Switzerland) using SYBR Green Select Master Mix (2x) (Applied Biosystems, Foster City, CA, USA). Reactions for each of the six biological replicates in all age groups were run in triplicates in a 15 μl volume. The reaction master mixes were prepared using SYBR Green Master Mix 2x (7.5 μl), 20 mM forward and reverse primer (0.48 μl), cDNA template (3 μl), and nuclear-free water (4.02 μl) in 96-well white plates (Roche, Basel, Switzerland). All qPCR reactions were performed under the following conditions: 3 minutes for the pre-PCR denaturation and polymerase activation step at 95°C, followed by 40 cycles of 15 s denaturation at 95°C, a 20 s hybridization step at 57°C and a 45 s elongation step at 72°C. The quantification and calculation of the gene expression levels were performed according to the standard curve method [45]. Standard curves for every gene of five dilution series (from 1010 to 106 copies of DNA molecules) were constructed from purified cDNA (using the QIAquick PCR Purification Kit, following the manufacturer’s protocol). Melting curve analyses were performed to confirm the amplification of a single PCR product. The thermal profile for the melting curve determination began with incubation of 1 min at 60°C with a gradual increase in temperature (1°C/15 s) to 95°C, during which time changes in fluorescence were monitored. No-template control tubes, containing water instead of template mRNA, were run under the same conditions for each gene. Real-time PCR efficiencies were calculated using the standard curve methods and normalized to the expression of the housekeeping gene ribosomal protein 49 (PtRP49) for each sample. Preliminary tests (not shown) of the most commonly used references genes (RP49, RP18, α-actin, β-actin, 3-phosphate dehydrogenase; [46]) revealed that only the PtRP49 gene was stably expressed in various tissues, regardless of the stage of spider ontogenesis.


The results are reported as mean values ± SD. Normality was checked using the Kolmogorov-Smirnov test. The data were tested for homogeneity of variance using Levene’s test of equality of error variances. Whenever a significant effect was observed, the Tukey multiple comparison test was used for a post hoc one-way analysis of variance (ANOVA). Analysis of variance for the level of genes encoding allatoregulatory neuropeptides (PtASTA, PtASTB, PtASTC, PtAT) and their receptors (PtASTA-R, PtASTB-R, PtASTC-R, PtAT-R) at different stages of ontogenesis of P. tepidariorum females was performed using two-way ANOVA with the experimental groups (various tissues and bands) and the stages of life as the sources of differences. Results with p≤ 0.05 were considered to be significant. The data were analyzed using GraphPad Prism ver. 6.

Results and discussion

Expression of genes encoding allatoregulatory neuropeptides and their receptors was visualized by a two-step real-time polymerase chain reaction (qPCR) method with cDNA as a template extracted from tissues/organs and from the entire body of P. tepidariorum females in various stages of ontogenesis. Proper products, corresponding to the expected fragments, were amplified for PtASTA, PtASTC, and PtAT (Fig 1C). Among all tissues/ organs of the P. tepidariorum females analyzed in the experiment, no amplification was observed for genes encoding AST B and its receptor. The correct sequence of the allatostatin B receptor in the transcriptome of the P. tepidariorum spider has not been demonstrated by in silico analysis. The nucleotide sequences most closely related to ASTB-R do not contain TM1-TM7 conserved transmembrane domains (S7 Fig). It seems that AST B does not belong to the physiological neurosecretions of P. tepidariorum, and it does not play a role in the regulation of their physiological processes. On the other hand, this may also indicate a gene mutation resulting in a significant change of the nucleotide sequence in the genome. So far, among all allatostatins, peptides belonging to allatostatin B have been characterized and isolated only in five species of insects: Locusta migratoria [47], M. sexta [48], Carausius morosus [49], Gryllus bimaculatus [50] and D. melanogaster [51]. In contrast, allatostatin A and allatostatin C are widely distributed in various groups of insects. This may indicate a limited occurrence of this peptide in arthropods. Moreover, the potency of the B type allatostatins is approximately 50% lower than that of the A type allatostatins in G. bimaculatus. Lorenz et al. [52], as well as Wang [51], proved that the activity of AST B is limited to insects from the family of crickets (Gryllidae). These peptides isolated from C. morosus inhibited CA activity only in crickets but did not affect the CA of the C. morosus itself. Therefore, it can be assumed that limited occurrence of allatostatin B in insects and its limited activity may explain the possible lack of allatostatin B in P. tepidariorum females.

<h2>The relative expression level of genes encoding allatoregulatory neuropeptides in <i>Parasteatoda tepidariorum</i> females.</h2>
Fig. 1.

The relative expression level of genes encoding allatoregulatory neuropeptides in Parasteatoda tepidariorum females.

Changes of the expression level of allatostatin A (PtASTA), allatostatin C (PtASTC), allatotropin (PtAT) (A) and their receptors’ genes (PtASTA-R, PtASTC-R, PtAT-R) (B) during the ontogenesis stages in the entire body of P. tepidariorum females. Differential expression of genes encoding allatoregulatory neuropeptides and their receptors in female (C) NS–neuroendocrine and nervous system, OV–ovaries, MG–midgut glands with midgut, HG–hindgut, Vgs–vitellogenesis process, Ma–mating, Ovi–oviposition.

The obtained results confirmed the presence of allatoregulatory neuropeptides in P. tepidariorum. They revealed substantial differences in genes encoding allatoregulatory neuropeptides expression levels and their receptors in the different tissues, which may indicate the diverse production of neuropeptides and their subsequent action in the target tissues. The level of gene expression also varied depending on the age of individuals. However, for all transcripts excluding the entire body (Fig 1A and 1B), the highest expression level coincides with the beginning of sexual maturation at the 40th day of life.

It is clear that there might be more paralogs or that the genes observed are most likely homologs of the insect neuropeptides, but that a detailed phylogenetic analysis was not performed. It seems, that to confirm the presence of paralogs it would be necessary to perform phylogenetic analysis based on the newly available genome described by Posnien et al. [53] or Schwager et al. [54].

Allatoregulatory neuropeptide expression in various tissues of mature females

Among all tissues/organs of the female P. tepidariorum analyzed in the experiment, proper products of amplification for PtASTA, PtASTC, PtAT, and their receptors were observed. The highest level of expression of AST A, AST C, and AT genes was recorded in the neuroendocrine and nervous systems. Moreover, the same level of expression for the gene encoding AST C was observed in the hindgut (Fig 2), whereas the maximum expression level of receptors was found in the hindgut for PtASTA-R and PtAT-R as well as in the ovaries for PtASTC-R (Fig 2). In case of integument, no products of the amplification of all analyzed genes was obtained.

<h2>Tissue-specific expression of allatoregulatory neuropeptides.</h2>
Fig. 2.

Tissue-specific expression of allatoregulatory neuropeptides.

Relative expression of allatostatin A, allatostatin C, allatotropin and their receptors of P. tepidariorum females at the 40th day of ontogenesis (the day of sexual maturity) [mean ± SD]. NS–neuroendocrine and nervous system, OV–ovaries, MG–midgut glands with midgut, INT–integument, HG–hindgut (HG). Different letters indicate statistically significant differences: a-d differences between levels of expression of genes encoding allatoregulatory neuropeptides (PtASTA, PtASTC, PtAT) in tested tissues, A-D differences between levels of expression of genes encoding allatoregulatory neuropeptide receptors (PtASTA-R, PtASTC-R, PtAT-R) in tested tissues. Asterisks indicate statistically significant differences between levels of expression of gene encoding allatoregulatory neuropeptide and its receptor in the same tissue. Tukey's multiple comparison test, p≤0.05.

For all allatoregulatory neuropeptides significant differences in gene expression levels were found compared to their receptor expression. In the neuroendocrine and nervous systems, the relative level of PtASTA was significantly higher than the PtASTA-R level (Fig 2). The same correlation was observed for the gene encoding AST C and its receptor (Fig 2). In the remaining tissues, the inversed tendency was observed, where the expression of PtASTA-R gene was significantly higher than the expression of the gene encoding PtASTA. The most significant difference in the expression of the neuropeptide and its receptor was observed in the hindgut, where the expression of PtASTA-R was 20-fold higher than that of PtASTA (Fig 2). Analysis of the expression of PtASTC and PtASTC-R genes revealed that the relative level of PtASTC expression was also higher than the level of PtASTC-R in the hindgut. Furthermore, comparison of expression of the gene encoding the receptor of AT was higher than the level of PtAT in all tissues/organs excluding neuroendocrine and nervous systems, where a significant difference between the level of both transcripts was not observed (Fig 2).

The highest expression of genes encoding PtASTA, PtASTC, and PtAT in the neuroendocrine and nervous systems in comparison to other tissues of P. tepidariorum confirms the presence of neurosecretory cells, constituting the main site of synthesis and secretion of allatoregulatory neuropeptides. Our results are the first evidence consistently indicating the neuroendocrine and nervous system as a place of synthesis of allatoregulatory neuropeptides in spiders and confirming the assumptions described before by Bonaric [32]. Moreover, these results coincide with the data published for C. salei, where the presence of allatostatin A in the arcuate body of the central nervous system by immunochemistry method has been demonstrated [23]. This clearly confirmed the presence of allatostatins in the nervous system of spiders.

In addition, to the neuroendocrine and nervous systems, high differences of the transcripts levels have been demonstrated in the different tissues. The expression of genes encoding AST A, AST C, and AT was primarily confirmed in the ovaries, midgut glands with midgut and in the hindgut. In a study conducted by Stay et al. [55] in Diploptera punctata as well as by Witek and Hoffmann [56] in Gryllus bimaculatus, it was demonstrated that allatoregulatory neuropeptides can also be secreted by the ovaries, which would confirm the high expression of genes encoding these neuropeptides obtained in the ovaries.

The results of the expression of genes encoding the allatoregulatory neuropeptide receptors also indicate considerable variation according to tissue and ontogenesis stage. The expression of genes encoding the receptors of allatoregulatory neuropeptides was also demonstrated in the neuroendocrine and nervous systems, but its level was lower than in the other tissues. As explained earlier, the tissue referred to as the neuroendocrine and nervous systems contained all the structures of the neuroendocrine system described by Bonaric [29, 30, 31, 32] as well as by Bonaric and Juberthie [33]. In the aforementioned manuscripts, the neuroendocrine and nervous systems were selected as the site of JH synthesis and/or its analog as well as other substances belonging to the neurosecretions of spiders. The presence of allatoregulatory neuropeptide receptors in this complex of structures indicates the allatoregulatory control of the functioning of the neuroendocrine and nervous systems. We can assume that these neuropeptides influence the secretion of other neurotransmitters through neurosecretory cells of the neuroendocrine and nervous systems [1, 20, 57, 58]. To date, the expression of genes encoding allatoregulatory neuropeptide receptors in the nervous system of insects has been reported in A. aegypti [59], D. melanogaster [60] or Apis mellifera [61]. Similar results obtained in insects and for P. tepidariorum suggest that the allatoregulatory neuropeptides in spiders may play the primary control function of JH secretion. Additionally, it has been previously shown that AST can affect visual information processing [62], learning and memory [61]. The multitude of nervous system functions makes it difficult to deduce, based only on the expression of the studied transcripts.

Analysis of the expression of PtASTA-R and PtAT-R genes confirmed their highest level in the hindgut, while the maximum expression level of the gene encoding the AST C receptor (PtASTC-R) was observed in the ovaries. The strong expression of analyzed genes in the hindgut may indicate the use of these neuropeptides for the regulation of gastrointestinal function as well as in intestinal peristaltic contractions, as in insects [58, 63, 64, 65, 66, 67]. Furthermore, it has also been shown that allatoregulatory neuropeptides control the secretion of digestive enzymes and stimulate their activity [68, 69, 70]. It cannot be excluded that similar mechanisms may occur in spiders. This conclusion is confirmed by the high expression of these transcripts in the midgut glands with midgut responsible for the production and secretion of digestive enzymes in spiders. The obtained results may also indicate the participation of allatoregulatory neuropeptides in other processes that take place in the midgut glands and midgut: energy metabolism processes, protein synthesis, and mobilization of spare substances. Protein synthesis in midgut glands and the midgut is also associated with the initiation of the vitellogenesis process. Hitherto, there is only one publication reporting the effect of AST A on reducing the production of vitellogenins in the fat body, as well as the inhibition of secretion of these proteins to hemolymph in B. germanica [71].

Proper products, corresponding to the expected fragments, amplified for genes encoding receptors of allatoregulatory neuropeptides in the ovaries suggest that they may regulate oocyte development, vitellogenesis, and choriogenesis, which has been confirmed by Woodhead et al. [72] in D. punctata, and the biosynthesis of ecdysone in the ovaries presented by Lorenz [73] in G. bimaculatus and Lorenz et al. [74] in Blaptica dubia.

Results from this study may confirm that in P. tepidariorum females the digestive tract and ovaries are the main tissues under allatoregulatory control. The process of vitellogenesis in P. tepidariorum is controlled on various levels (the gene expression, protein synthesis, protein deposition in ovaries) by allatoregulatory neuropeptides, indicated by allatotropin and allatostatin injections [75]. The allatoregulatory neuropeptides regulate the spider P. tepidariorum metabolism by inducing changes in the concentration of glycogen, lipids, and proteins in the midgut glands with midgut [76].

Age-dependent allatoregulatory neuropeptide expression

The expression profile of genes encoding allatoregulatory neuropeptides and their receptors was dependent on the stage of ontogenesis of P. tepidariorum females. Analysis of the entire body of P. tepidariorum females revealed that the expression of PtAT, PtASTA-R, and PtASTC-R was stable in most cases, throughout their life, except the 35th day of ontogenesis when the PtASTA-R expression was significantly higher than the level in the other stages (Fig 1A and 1B). In females at the last nymphal stage (38th day of life) and 3 days after copulation (43rd day of life), the level of PtASTA and PtASTC expression was significantly lower compared to the earlier stages of ontogenesis. Compared to other genes encoding allatoregulatory neuropeptides, a lower level of expression for the PtAT gene in the entire body of P. tepidariorum females was confirmed (Fig 1A).

Expression of all transcripts in selected tissues or organs was also age-dependent (Fig 1C). The same relation of changes in the expression level as in the entire body was confirmed. The maximum level of expression of every gene was observed on the day of sexual maturity (40th day of ontogenesis) in all tested tissues, except for the expression of PtASTA in the hindgut, where the highest expression level was observed in the penultimate nymphal stage. The highest expression of transcripts on sexual maturation day may indicate an increase in neuropeptide production. Such dependence may indicate the role of these substances in physiological processes that take place very intensively during that period. Among them, vitellogenesis is the essential process; it begins in spiders in the last nymphal stage [77] and intensifies significantly when sexual maturity is reached. Because vitellogenesis requires high energy expenditure, the high level of expression of transcripts in the midgut glands with midgut may also indicate their influence on the regulation of energy mobilization from the energy storage compounds.

Moreover, on the 38th and 43rd days of life, the relative expression levels of PtASTA, PtASTA-R, PtASTC, and PtASTC-R were lower than in the earlier stages of ontogenesis. In the neuroendocrine and nervous systems as well as in the midgut glands with midgut the expression levels of PtASTA, PtASTC and PtASTA-R transcripts on the 38th day were twice as low as on the 35th day, whereas in the ovaries the expression level of PtASTC was 11-fold lower. Inhibition of the expression of genes encoding ASTs at this stage of ontogenesis may indicate a reduction in the rate of physiological processes associated with the preparation of the body to achieve sexual maturity. The PtAT gene expression pattern was different from genes encoding ASTs; PtAT gene expression was not observed until the 40th day of individual development. The effect of AT on the secretion of JH and/or its analog, indispensable in development and larval growth, confirmed in insects may explain the lack of PtAT gene expression on days 35 and 38 of ontogenesis. These results may confirm that the synthesis of AT in P. tepidariorum females does not occur in the nymphal stages. In insects, it has been proved that larval-larval molting and metamorphosis can only occur as a result of the significant reduction of the JH level, which inhibits both of these processes [78, 79]. Based on the main AT function in insects, which is the stimulation of JH secretion, it may be relatively easy to explain the lack of its expression in the nymphal stages. A high level of JH due to the presence of allatotropin would prevent the molting and growing of spiders. This is also confirmed by the high level of allatotropin after the last molt (the day of sexual maturity) and the level of gene expression encoding enzyme of the juvenile hormone biosynthetic pathway (PtCYP15A1 enzyme responsible for catalyzing the epoxidation of methyl farnesoate to the juvenile hormone) in the same ontogenesis stages of P. tepidariorum. Bednarek et al. [80] confirmed the lack of PtCYP15A1 transcript in nymphal stages and high level of this product on the day of sexual maturity.

It was also observed that on day 43 of life the expression in the neuroendocrine and nervous systems was 7-fold lower in the case of genes encoding AST A and nearly 6-fold lower for the AST A receptor gene in relation to the day of sexual maturity. The changes in PtASTC and PtASTC-R expression levels were similar, but the differences were much more extensive. On the 43rd day of ontogenesis, a more significant difference in the obtained results was confirmed. In the midgut glands with midgut and hindgut, the expression of PtASTC was 14-fold lower with respect to the results from the 40th day of ontogenesis, whereas expression of the PtASTC-R gene in the midgut glands with midgut at the 43rd day of development was 20-fold lower compared to the previous stage of ontogenesis. Again, higher levels of expression of the PtASTA, PtASTA-R, PtASTC, and PtASTC-R transcripts were confirmed in the neuroendocrine and nervous systems and the ovaries in females after the formation of the first cocoons (47th day). In other tissues, neither transcript was expressed during the day of ontogenesis. The overlap of results obtained for genes encoding the allatoregulatory neuropeptides and results for genes encoding their receptors confirm the increased use of AST A and AST C during egg laying. The high expression of the PtASTA and PtASTC genes in the neuroendocrine and nervous system confirms their increased synthesis, while the highest expression of PtASTA-R and PtASTC-R genes in ovaries indicate their role in the regulation of the ovaries’ action. Garside et al. [81] reported that in D. punctata the expression of AST A (Dipp-AST A) was deficient in the ovaries during the initiation of vitellogenesis (analogous results on the 38th day of spider ontogenesis) and increased until spermatophore deposition. Immediately after this process, the level of Dipp-AST A was lower, and after laying eggs, it increased rapidly. However, Woodhead et al. [72] reported that the level of AST in D. punctata ovaries was increasing from 3 days after pupation, while after spermatophore deposition, AST A content rapidly increased, with the maximum level on the day of egg laying. The authors noted that the rapid increase in the level of ASTs in the ovaries is correlated with the process of choriogenesis.

In contrast, the expression profiles for PtAT and PtAT-R genes were different. No expression in the nymphal stages (35th and 38th) was detected, although, after sexual maturity, expression rates were significantly higher than PtASTA and PtASTC (Fig 1C). In all tissues, PtAT gene expression decreased gradually from days 40 to 47 of spider development. The increased expression of both the PtASTA and PtASTA-R genes in the nymphal stages and the confirmed AST A activity in relation to the synthesis of JH indirectly confirms the occurrence of JH and/or its analog in P. tepidariorum. The secretion of AST A in the nymphal stages and the lack of AT secretion may indicate inhibition of JH synthesis and/or its analog, which enabled the proper course of molting. Such mechanisms of action of allatoregulatory neuropeptides have been confirmed in insects [58, 78, 79].

To conclude, the results demonstrated in this paper confirm the presence of allatoregulatory neuropeptides (AST A, AST C, and AT) in females of the spider P. tepidariorum, which complements the current knowledge of both the physiology of spiders and the occurrence of allatoregulatory neuropeptides. Moreover, the neuroendocrine and nervous systems have been reported to be the main sites of synthesis of these compounds, whereas the digestive tract and ovaries were primarily affected by the allatoregulatory control. The highest levels of all tested neuropeptides were observed on the 40th day of females’ lives. This indicates increased production of these proteins and their increased use during the physiological processes that took place on that day, like vitellogenesis, oocyte development, or mating. These results are a starting point for future functional studies, which will advance our understanding of the role of neuropeptides in the regulation of spider physiology.

Supporting information

S1 Fig [a]

S2 Fig [a]

S3 Fig [a]

S4 Fig [a]

S5 Fig [a]

S6 Fig [tif]
Multiple alignment of allatostatin A receptor from and other arthropods.

S7 Fig [tif]
Multiple alignment of allatostatin B receptor from and other arthropods.

S8 Fig [tif]
Multiple alignment of allatostatin C receptor from and other arthropods.

S9 Fig [tif]
Multiple alignment of allatotropin receptor from and other arthropods.

S10 Fig [tif]
Multiple alignment of allatostatin A precursors from and other arthropods.

S11 Fig [tif]
Multiple alignment of allatostatin B precursors from and other arthropods.

S12 Fig [tif]
Multiple alignment of allatostatin C precursors from and other arthropods.

S13 Fig [tif]
Multiple alignment of allatotropin precursors from and other arthropods.

S1 File [pdf]
The real-time PCR results.

S1 Table [pdf]


1. Marciniak P, Szymczak M, Rosiński G. Hormony peptydowe owadów–przegląd najważniejszych rodzin. Post Biol Komórki. 2011;38: 43–63.

2. Sarkar NRS, Tobe SS, Orchard I. The distribution and effects of Dippu-allatostatin-like peptides in the blood-feeding bug, Rhodnius prolixus. Peptides. 2003;24: 1553–1563. doi: 10.1016/j.peptides.2003.07.015 14706534

3. Nassel DR. Neuropeptides in the nervous system of Drosophila and other insects: multiple roles as neuromodulators and neurohormones. Prog Neurobiol. 2002;68: 1–84. 12427481

4. Dickinson PS, Wiwatpanit T, Gabranski ER, Ackerman RJ, Stevens JS, Cashman CR, et al.Identification of SYWKQCAFNAVSCFamide: a broadly conserved crustacean C-type allatostatin-like peptide with both neuromodulatory and cardioactive properties. J Experimental Biology. 2009;212: 1140–1152.

5. Christie AE, Pascual MG. Peptidergic signaling in the crab Cancer borealis: Tapping the power of transcriptomics for neuropeptidome expansion. Gen Comp Endocrino. 2016;237: 53–67.

6. Christie AE. Expansion of the neuropeptidome of the globally invasive marine crab Carcinus maenas. Gen Comp Endocrino. 2016;235: 150–169.

7. Christie AE, Pascual MG, Yu A. Peptidergic signaling in the tadpole shrimp Triops newberryi: A potential model for investigating the roles played by peptide paracrines/hormones in adaptation to environmental change. Mar Genomics. 2018;39: 45–63. doi: 10.1016/j.margen.2018.01.005 29526397

8. Duve H, Johnsen A, Scott AG, Thorpe A. Allatostatins of the tiger prawn, Penaeus monodon (Crustacea: Penaeidea). Peptides. 2002;23: 1039–1051. doi: 10.1016/s0196-9781(02)00035-9 12126730

9. Christie AE, Miller A, Fernandez R, Dickinson ES, Jordan A, Kohn J, et al. Non-amidated and amidated members of the C-type allatostatin (AST-C) family are differentially distributed in the stomatogastric nervous system of the American lobster, Homarus americanus. Invert Neurosci. 2018;18: 2. doi: 10.1007/s10158-018-0206-6 29332202

10. Ahn SJ, Martin R, Rao S, Choi MY. Neuropeptides predicted from the transcriptome analysis of the gray garden slug Deroceras reticulatum. Peptides. 2017;93: 51–65. doi: 10.1016/j.peptides.2017.05.005 28502716

11. Sawadro M, Bednarek A, Babczyńska A. The current state of knowledge on the neuroactive compounds that affect the development, mating and reproduction of spiders (Araneae) compared to insects. Invert Neurosci. 2017;17: 4. doi: 10.1007/s10158-017-0197-8 28421370

12. Smart D, Johnston CF, Curry WJ, Williamson R, Maule AG, Skuce PJ, et al. Peptides related to the Diploptera punctata allatostatins in nonarthropod invertebrates: An immunocytochemical survey. J Comp Neurol. 1994; 347: 426–432. doi: 10.1002/cne.903470308 7822491

13. Alzugaray ME, Hernández-Martínez S, Ronderos JR. Somatostatin signaling system as an ancestral mechanism: Myoregulatory activity of an Allatostatin-C peptide in Hydra. Peptides. 2016;82: 67–75. doi: 10.1016/j.peptides.2016.05.011 27288244

14. Laufer H, Borst D, Baker FC, Reuter CC, Tsai LW, Schooley DA, et al. Identification of a juvenile hormone–like compound in a crustacean. Science. 1987;235: 202–205. doi: 10.1126/science.235.4785.202 17778635

15. Laufer H, Landau M, Borst D, Homola E. The synthesis and regulation of methyl farnesoate, a new juvenile hormone for crustacean reproduction. Adv Inver Reprod. 1986;4: 135–143.

16. Gonçalves SC, Carvalho HF, Hartfelder K. Juvenile hormone signaling in insect oogenesis. Curr Opin Insect Sci. 2019;31: 43–48. doi: 10.1016/j.cois.2018.07.010 31109672

17. Harshini S, Nachman RJ, Sreekumar S. Inhibition of digestive enzyme release by neuropeptides in larvae of Opisina arenosella (Lepidoptera: Cryptophasidae). Comp Biochem Physiol. B. 2002;132: 353–358. 12031460

18. Marciniak P, Rosiński G. Aktualny stan badań nad neuropeptydami miotropowymi owadów: tachykininy, sulfakininy i FMRFa-pokrewne peptydy. Post Biol Komórki. 2007;34: 241–249.

19. Bendena WG. Neuropeptide physiology in Insects. Neuropeptide systems as targets for parasite and pest control. Adv Exp Med Biol. 2010;692: 166–191. doi: 10.1007/978-1-4419-6902-6_9 21189679

20. Gade G, Hoffmann KH. Neuropeptides regulating development and reproduction in insects. Physiol Entomol. 2005;30: 103–121.

21. Zhu XX, Oliver JH. Cockroach allatostatin–like immunoreactivity in the synganglion of the American dog tick Dermacentor variabilis (Acari: Ixodidae). Exp Appl Acarol. 2001;25: 1005–1013. 12465854

22. Neupert S, Russell WK, Predel R, Russell DH, Strey OF, Teel PD, et al. The neuropeptidomics of Ixodes scapularis synganglion. J Proteome Res. 2009;72: 1040–1045.

23. Loesel R, Seyfarth EA, Bräunig P, Agricola HJ. Neuroarchitecture of the arcuate body in the brain of the spider Cupiennius salei (Araneae, Chelicerata) revealed by allatostatin–, proctolin–, and CCAP–immunocytochemistry and its evolutionary implications. Arthropod Struct Dev. 2011;40: 210–220. doi: 10.1016/j.asd.2011.01.002 21256976

24. De Loof A, Hoffmann KH. Neuropeptides in insect development and reproduction. Arch Insect Biochem Physiol. 2001;47: 127–128. doi: 10.1002/arch.1043 11418930

25. Bednarek A, Sawadro M, Babczyńska A. Modulation of the response to stress factors of Xerolycosa nemoralis (Lycosidae) spiders living in contaminated environments. Ecotoxicol Environ Saf. 2016;131: 1–6. doi: 10.1016/j.ecoenv.2016.04.027 27162128

26. Turnbull AL. 1973. Ecology of the true spiders (Araneomorphae). Annu Rev Entomol. 1973;18: 305–348.

27. Wise DH. Spiders in Ecological Webs. Cambridge University Press, Cambridge, UK. 1993.

28. Ekschmitt K, Wolters V, Weber M. Spiders, carabids, and staphylinids: the ecological potential of predatory macroarthropods. In: Benckiser G. editor. Fauna in Soil Ecosystems., New York: Marcel Dekker; 1997. p. 307–362.

29. Bonaric JC. Contribution a l'etude de la biofogie de developpement chez l'araignee Pisaura mirabilis (Clerck., 1758). Approche physiologique des phenomenes de mue et de diapause hivemale. Montpellier, PhD. 1980.

30. Bonaric JC. Effets des ecdysones et de l'hormone juvénile sur ladurée du cycle de mue chez l'araignée Pisaura mirabilis (Araneae, Pisauridae). Rev Arachn. 1979;2: 205–207.

31. Bonaric JC. Juberthie UEC. Ultrastructure of the Retrocerebral Neuroendocrine Complex in Pisaura mirabilis Cl. (Araneae, Pisauridae). Zool Jahrb. Abt allg Zool Physiol Tiere. 1983;87: 55–64.

32. Bonaric JC. Le systeme neuroendocrine retrocerebral des Araignees: structure et function. In: Ruzicka V. editor. Proceedings of the 15th European Colloquium of Arachnology. Ceske Budejovice: Institute of Entomology; 1995. p. 27–34.

33. Bonaric JC. Moulting Hormones. In: Nentwig W. editor. Ecophysiology of spiders. Berlin: Springer-Verlag; 1986. p. 111–118.

34. Bonaric JC, Emerit M, Legendre R. Le complexe neuroendocrine rétrocérébral et la «glande de mue» de Filistata insidiatrix Forskôl (Araneae, Filistatidae). Rev Arch. 5;1984: 301–310.

35. Miyashita K. Development and egg sac production of Achaearanea tepidariorum (CL Koch) (Araneae, Theridiidae) under long and short photoperiods. J Arachn. 1987;15: 51–58.

36. NCBI Protein database in National Library of Medicine (US), 2017. National Center for Biotechnology Information [Internet]. Available: Accesses: 20 May 2017.

37. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. Basic local alignment search tool. J Mol Biol. 1990;215: 403–410. doi: 10.1016/S0022-2836(05)80360-2 2231712

38. Pearson WR. An Introduction to Sequence and Series. Int J Res. 2014;1: 1286–1292.

39. Jones P, Binns D, Chang H-Y, Fraser M, Weizhong LW, McAnulla C, et al. InterProScan 5: genome-scale protein function classification. Bioinformatics. 2014;30: 1236–1240. doi: 10.1093/bioinformatics/btu031 24451626

40. Ouedraogo M, Bettembourg C, Bretaudeau A, Sallou O, Diot C, Demeure O, et al. 2012.The duplicated genes database: identification and functional annotation of co-localised duplicated genes across genomes. PLoS One. 2012;7(11):e50653. doi: 10.1371/journal.pone.0050653 23209799

41. Kumar S, Stecher G, Li M, Knyaz C, Tamura K. MEGA X: Molecular Evolutionary Genetics Analysis across computing platforms. Mol Biol Evol. 2018;35: 1547–1549. doi: 10.1093/molbev/msy096 29722887

42. Wong ML, Medrano JF. Real-time PCR for mRNA quantitation. BioTechniques. 2005;39: 1–11.

43. Saitou N, Nei M. The neighbor-joining method: A new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4: 406–425. doi: 10.1093/oxfordjournals.molbev.a040454 3447015

44. Chomczyński P, Sacchi N. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-choloroform extraction. Anal Biochem. 1987;162: 156–159. doi: 10.1006/abio.1987.9999 2440339

45. Bookout AL, Cummins CL, Mangelsdorf DJ. High-throughput real-time quantitative reverse transcription PCR. In: Ausubel FM, Brent R, Kingston RE, Moore DD, Seidman JG, Smith JA, et al., editors. Current Protocols in Molecular Biology. Michigan: Wiley; 2005. p. 2363–2383.

46. Felsenstein J. Confidence limits on phylogenies: An approach using the bootstrap. Evolution. 1985;39: 783–791. doi: 10.1111/j.1558-5646.1985.tb00420.x 28561359

47. Schoofs L, Holman GM, Hayes TK, Nachman RJ, De Loof A. Isolation, identification and synthesis of locusta-myoinhibiting peptide (Lom-MIP), a novel biologically active neuropeptide from Locusta migratoria. Regul Pept. 1991;36: 111–119. doi: 10.1016/0167-0115(91)90199-q 1796179

48. Blackburn MB, Wagner RM, Kochansky JP, Harrison DJ, Thomas-Laemont P, Raina AK. The identification of two myoinhibitory peptides, with sequence similarities to the galanins, isolated from the ventral nerve cord of Manduca sexta. Regul Pept. 1995;57: 213–219. doi: 10.1016/0167-0115(95)00034-9 7480870

49. Lorenz MW, Kellner R, Hoffmann KH, Gade G. Identification of multiple peptide homologous to cockroach and cricket allatostatins in the stick insect, Carausius morosus. Insect Biochem Mol Biol. 2000;30: 711–718. 10876114

50. Williamson M, Lenz C, Winther AME, Nassel DR, Grimmelikhuijzen CJP. Molecular cloning, genomic organization, and expression of a B–type (cricket–type) allatostatin preprohormone from Drosophila melanogaster. Biochem Biophys Res Commun. 2001;281: 544–550. doi: 10.1006/bbrc.2001.4402 11181081

51. Wang J. Isolation and characterization of the B–type allatostatin gene of Gryllus bimaculatus de Geer (Ensifera, Gryllidae). PhD. 2004.

52. Lorenz MW, Gäde G, Hoffmann KH. Interspecific actions of allatostatins. Mitt Dtsch Ges Allg Angew Entomol. 1997;11: 549–553.

53. Posnien N, Zeng V, Schwager EE, Pechmann M, Hilbrant M, Keefe JD, et al. A comprehensive reference transcriptome resource for the common house spider Parasteatoda tepidariorum. PLoS One. 2014;9(8):e104885. doi: 10.1371/journal.pone.0104885 25118601

54. Schwager EE, Sharma PP, Clarke T, Leite DJ, Wierschin T, Pechmann M, et al. The house spider genome reveals an ancient whole-genome duplication during arachnid evolution. BMC Biol. 2017;15: 62. doi: 10.1186/s12915-017-0399-x 28756775

55. Stay B, Sereg Bachmann JA, Stoltzmann CA, Fairbairn SE, Yu CG, Tobe SS. Factors affecting allatostatin release in a cockroach (Diploptera punctata): Nerve section, juvenile hormone analog and ovary. J Insect Physiol. 1994;50: 365–372.

56. Witek GK, Hoffmann H. Immunological evidence for FGLamide- and w2w9-allatostatins in the ovary of Gryllus bimaculatus (Ensifera, Gryllidae). Physiol Entomol. 2001;26: 49–57.

57. Abdel-Latief M, Hoffmann KH, Functional activity of allatotropin and allatostatin in the pupal stage of a holometablous insect, Tribolium castaneum (Coleoptera, Tenebrionidae). Peptides. 2014;53: 172–184. doi: 10.1016/j.peptides.2013.10.007 24140809

58. Yamanaka N, Yamamoto S, Zitnan D, Watanabe K1, Kawada T, Satake H, et al. Neuropeptide Receptor Transcriptome Reveals Unidentified Neuroendocrine Pathways. PLoS ONE. 2008;3(8): e3048. doi: 10.1371/journal.pone.0003048 18725956

59. Nouzova M, Rivera-Perez C, Noriega FG. Allatostatin-C reversibly blocks the transport of citrate out of the mitochondria and inhibits juvenile hormone synthesis in mosquitoes. Insect Biochem Mol Biol. 2015;57: 20–26. doi: 10.1016/j.ibmb.2014.12.003 25500428

60. Lenz C, Williamson M, Grimmelikhuijzen CJ. Molecular cloning and genomic organization of a second probable allatostatin receptor from Drosophila melanogaster. Biochem Biophys Res Commun. 2000;273: 571–577. doi: 10.1006/bbrc.2000.2964 10873647

61. Urlacher E, Soustelle L, Parmentier ML, Verlinden H, Gherardi MJ, Fourmy D, et al. Honey Bee Allatostatins Target Galanin/ Somatostatin-Like Receptors and Modulate Learning: A Conserved Function? PLoS ONE. 2016;11(1): e0146248. doi: 10.1371/journal.pone.0146248 26741132

62. Kreienkamp HJ, Larusson HJ, Witte I, Roeder T, Birgül N, Honck HH, et al. Functional annotation of two orphan G-proteincoupled receptors, Drostar1 and -2, from Drosophila melanogaster and their ligands by reverse pharmacology. J Biol Chem. 2002;277: 39937–39943. doi: 10.1074/jbc.M206931200 12167655

63. Lange AB, Chan KK, Stay B. 1993. Effect of allatostatin and proctolin on antennal pulsatile organ and hindgut muscle in the cockroach, Diploptera punctata. Arch Insect Biochem Physiol. 1993;24: 79–92. doi: 10.1002/arch.940240203 7902139

64. Yu CG, Hayes TK, Strey A, Bendena WG, Tobe SS. Identification and partial characterization of receptors for allatostatins in brain and corpora allata of the cockroach Diploptera punctata using a binding assay and photoaffinity labeling. Regul Pept. 1995;57: 347–358. doi: 10.1016/0167-0115(95)00048-G 7480884

65. Secher T, Lenz C, Cazzamali G, Sorensen G, Williamson M, Hansen GN, et al. Molecular cloning of a functional allatostatin gut/brain receptor and an allatostatin preprohormone from the silkworm Bombyx mori. J Biol Chem. 2001;276: 47052–47060. doi: 10.1074/jbc.M106675200 11590150

66. Meyering-Vos M, Merz S, Sertkol M, Hoffmann KH. Functional analysis of the allatostatin-A type gene in the cricket Gryllus bimaculatus and the armyworm Spodoptera frugiperda. Insect Biochem Mol Biol. 2006;36: 492–504. doi: 10.1016/j.ibmb.2006.03.008 16731345

67. Lungchukiet P, Donly BC, Zhang J, Tobe SS, Bendena WG. Molecular cloning and characterization of an allatostatin-like receptor in the cockroach Diploptera punctata. Peptides. 2008;29: 276–285. doi: 10.1016/j.peptides.2007.10.029 18237821

68. Duve H, Wren P, Thorpe A. Innervation of the foregut of the cockroach Leucophaea maderae and inhibition of spontaneous contractile activity by allatostatin neuropeptides. Physiol Entomol. 1995;20: 33–44.

69. Aguilar R., Maestro JL, Vilaplana L, Pascual N, Piulachs MD, Belles X. Allatostatin gene expression in brain and midgut, and activity of synthetic allatostatins on feeding-related processes in the cockroach Blattella germanica. Regul Pept. 2003;115: 171–177. doi: 10.1016/s0167-0115(03)00165-4 14556958

70. Fuse M, Zhang JR, Partridge E, Nachman RJ, Orchard I, Bendena WG, et al. Effects of an allatostatin and a myosuppressin on midgut carbohydrate enzyme activity in the cockroach Diploptera punctata. Peptides. 1999;20: 1285–1293. doi: 10.1016/s0196-9781(99)00133-3 10612442

71. Martin D, Piulachs MD, Bell X. Inhibition of vitellogenin production by allatostatin in the German cockroach. Mol Cell Endocrinol. 1996;121: 191–196. doi: 10.1016/0303-7207(96)03864-6 8892320

72. Woodhead AP, Thompson ME, Chan KK, Stay B. Allatostatin in ovaries, oviducts, and young embryos in the cockroach Diploptera punctata. J Insect Physiol. 2003;49: 1103–1114. 14624882

73. Lorenz MW. Neuropeptides regulating developmental, reproductive, and metabolic events in crickets: structures and modes of action. J Insect Biotechnol Sericol. 2001;70: 69–93.

74. Lorenz JI, Lorenz MW, Hoffmann KH. Regulators of ovarian ecdysteroid release display opposite effects in the cricket Gryllus bimaculatus and the cockroach Blaptica dubia. Mitt Dtsch Ges Allg Angew Entomol. 2004;14: 447–450.

75. Sawadro M, Bednarek A, Molenda A, Babczyńska A. Allatoregulatory neuropeptides role in vitellogenesis process of Parasteatoda tepidariorum (Araneae, Theridiidae) spider females. Conference paper. 31st European Congress of Arachnology. 2018.

76. Sawadro M, Bednarek A, Molenda A, Babczyńska A, Metabolizm energetyczny samic Parasteatoda tepidariorum (Araneae, Theridiidae) po iniekcji neuropeptydów allatoregulujących. Conference paper. VII Ogólnopolska Konferencja Młodych Naukowców–ARTHROPOD. 2018.

77. Bednarek A, Sawadro M, Nicewicz Ł, Babczyńska A. Vitellogenins in the spider Parasteatoda tepidariorum–expression profile and putative hormonal regulation of vitellogenesis. BMC Dev Biol. 2019;19: 4. doi: 10.1186/s12861-019-0184-x 30849941

78. Riddiford LM. Cellular and molecular actions of juvenile hormone I. General considerations and premetamorphic actions. Adv In Insect Phys. 1994;24: 213–227.

79. Smykal V, Daimon T, Kayukawa T, Takaki K, Shinoda T, Jindra M. Importance of juvenile hormone signaling arises with competence of insect larvae to metamorphose. Dev Biol. 2014;390: 221–230. doi: 10.1016/j.ydbio.2014.03.006 24662045

80. Bednarek A, Sawadro M, Nicewicz Ł, Babczyńska A. Ekspresja genów kodujących enzym epox CYP15A1 oraz receptory hormonów juwenilnych w jajnikach samic pająka Parasteatoda tepidariorum (Araneae, Theridiidae). Conference paper. VII Ogólnopolska Konferencja Młodych Naukowców–ARTHROPOD. 2018.

81. Garside CS, Koladich PM, Bendena WG, Tobe SS. Expression of allatostatin in the oviducts of the cockroach Diploptera punctata. Insect Biochem Mol Biol. 2002;32: 1089–1099. 12213245

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