- Research article
- Open Access
Neuromuscular organization and aminergic modulation of contractions in the Drosophila ovary
© Middleton et al; licensee BioMed Central Ltd. 2006
Received: 23 January 2006
Accepted: 12 June 2006
Published: 12 June 2006
The processes by which eggs develop in the insect ovary are well characterized. Despite a large number of Drosophila mutants that cannot lay eggs, the way that the egg is moved along the reproductive tract from ovary to uterus is less well understood. We remedy this with an integrative study on the reproductive tract muscles (anatomy, innervation, contractions, aminergic modulation) in female flies.
Each ovary, consisting of 15–20 ovarioles, is surrounded by a contractile meshwork, the peritoneal sheath. Individual ovarioles are contained within a contractile epithelial sheath. Both sheaths contain striated muscle fibres. The oviduct and uterine walls contain a circular striated muscle layer. No longitudinal muscle fibres are seen.
Neurons that innervate the peritoneal sheath and lateral oviduct have many varicosities and terminate in swellings just outside the muscles of the peritoneal sheath. They all express tyrosine decarboxylase (required for tyramine and octopamine synthesis) and Drosophila vesicular monoamine transporter (DVMAT). No fibres innervate the ovarioles. The common oviduct and uterus are innervated by two classes of neurons, one with similar morphology to those of the peritoneal sheath and another with repeated branches and axon endings similar to type I neuromuscular junctions.
In isolated genital tracts from 3- and 7-day old flies, each ovariole contracts irregularly (12.5 ± 6.4 contractions/minute; mean ± 95% confidence interval). Peritoneal sheath contractions (5.7 ± 1.6 contractions/minute) move over the ovary, from tip to base or vice versa, propagating down the oviduct. Rhythmical spermathecal rotations (1.5 ± 0.29 contractions/minute) also occur. Each genital tract organ exhibits its own endogenous myogenic rhythm.
The amplitude of contractions of the peritoneal sheath increase in octopamine (100 nM, 81% P < 0.02) but 1 μM tyramine has no effect. Neither affects the frequency of peritoneal sheath contractions.
The muscle fibres of the reproductive tract are circular and have complex bursting myogenic rhythms under octopaminergic neuromodulation. We propose a new model of tissue-specific actions of octopamine, in which strengthening of peritoneal sheath contractions, coupled with relaxation of the oviduct, eases ovulation. This model accounts for reduced ovulation in flies with mutations in the octopaminergic system.
The amines octopamine and tyramine are apparently ubiquitous in arthropods, with neuromodulatory roles in a wide range of behaviours including jumping, flight, courtship and even social interactions (see [1, 2] for reviews). Although cellular actions of these neuromodulators have been described, in both the CNS and PNS, their exact roles in behaviour have resisted pharmacological approaches largely because of cross-talk between members of the amine receptor family.
Recently, the sequenced Drosophila genome, together with genetic knockouts of synthetic enzymes or receptors, have provided new tools to address the roles of these neuromodulators. These include two fly mutants in which the neurogenic amines octopamine or tyramine are not synthesized [3, 4] and others in which one class of octopamine or tyramine receptors is deficient [5, 6]. Surprisingly, these knockouts survive and appeared to behave "normally", but when fly behaviour was examined in detail, jumping ability was reduced by one-third  and egg-laying performance was dramatically reduced, if not abolished, suggesting that a detailed understanding of the neuromuscular basis of egg-laying would provide an excellent system for gaining an understanding of amine action.
The neuromuscular organization of egg-laying in Drosophila and its neuro-hormonal regulation remain poorly understood. This is surprising since many mutant lines are known in which egg-laying is reduced. Further, it is clear that the female fly controls oviposition to ensure the eggs are laid on a suitable substrate . We are now able to advance this as we show below that the isolated Drosophila female reproductive tract maintains activity in vitro. This allows us to separate out aminergic modulation on the reproductive tract musculature from aminergic effects on the CNS, in ways not previously possible.
Each ovariole is surrounded by an epithelial sheath (Fig. 3A,B, first described in Drosophila , that contains bands of circular muscles that run around the ovarioles. There are no muscle fibres running parallel to the long axis of the ovarioles.
In Drosophila, the ovarioles are held together to form two ovaries by an enveloping peritoneal sheath that is a network of muscle fibres (Fig. 3A,B) forming an open mesh around the ovary. Scanning electron micrographs  show the presence of large gaps (evident in Fig. 3A) in the peritoneal sheath below which the epithelial sheaths of the ovarioles can be seen.
EM sections cut longitudinally to myofibres of the peritoneal sheath (Fig. 3C) or of the two neighbouring epithelial sheaths (Fig. 3D) show the rather irregular interdigitation of the thick and thin filaments and the presence of perforated Z-discs. These are both characteristic of muscles that are supercontractile (i.e. they contract to less than 50% of their resting length when the thick filaments go through the Z-discs ). Supercontractile muscles occur in the insect viscera and in the larval body wall musculature . This is the first indication that these peritoneal sheath muscle fibres may be supercontractile, but this is not surprising given the large volume changes that occur during post-eclosional maturation of the ovary and oogenesis.
The muscles in the walls of the common (Fig. 4A–C) and lateral oviducts (not shown) are indistinguishable. Confocal fluorescence microscopy (Fig. 4C) shows that they are striated muscles consisting of an ordered array of circular myofibres forming an almost continuous sheet around their respective lumens. Occasional twists and splits are seen in the pattern, especially around the dark "holes" that may contain the nuclei. Ultrastructurally (Fig. 4A,B) the oviduct myofibrils are significantly thicker in cross section than those in the peritoneal and epithelial sheaths around the ovariole, but are structurally identical (Fig. 3). The perforated Z-discs are very clear (Fig. 4B). Infrequently isolated areas are seen in which thick and thin filaments (myofilaments) are cut in cross-section. This may reflect the observation that not all the myofibrils within a fibre show exactly the same orientation (Fig. 4C). Although myofilaments may occasionally deviate from the fibre's long axis, the overall structure shown by confocal fluorescence microscopy did not reveal any fibres or myofibrils running longitudinally in the wall of the oviduct. The oviduct lumen (Fig. 4A) is lined by an epithelial layer (EL) that shows convoluted intracellular membranous structures and extensive microvilli on the apical surface. It seems likely that these are involved in the transport of ions and various molecules  to produce oviduct secretions that facilitate egg movement. The epithelial layer has separated from the muscle layer in this sample, which may indicate that the two layers are not strongly adherent in vivo. The circular myofibrils and muscle layer of the uterus (Fig 4D,E) are much more substantial than those of the oviducts and there is no evidence of longitudinal myofibres. Within a single myofibre, neighbouring myofibrils often appear to be in approximate register (Fig. 4E) and the myofibres seem to have tapered ends. It is not known whether these represent attachment sites through which the muscle forces are applied to the uterus or whether this occurs along the length of the myofibres. Ultrastructural studies show (Fig 4D) the sarcomeric structure of the myofibrils, again with perforated Z-discs. The muscle layer is exterior to the epithelial lining of the uterine lumen. The cells in the uterine epithelium, like those of the oviduct (Fig 4A), show a considerable amount of convoluted membranous structures consistent with secretory activity.
In all the regions of the female reproductive tract examined, we found no evidence for longitudinal muscle fibres, though occasional twisting, oblique or longitudinal myo fibrils were seen in the oviduct. Our confocal and ultrastructural observations show that the fibres are striated and their myofibrils have perforated Z-discs indicating that all the muscles are supercontractile .
The extrinsic and intrinsic muscles of the uterus are innervated by a second pair of nerves, branching from the abdominal median nerve trunk. This pair of nerves (AbNvUt, Fig. 1) innervates both the extrinsic muscles and the circular myofibres (data not shown). These nerves have small boutons, closely apposed to larger, round blebs of Shaker-fluorescence, suggesting type I-like terminals. One branch projects to the inner layers of the uterus and shows our dTdc2 marker.
Additional File 1: Video micrograph of the reproductive tract of a 7 day old female fly, showing one ovary, ovarioles, oviduct and spermatheca. During the video, note the contractions of the peritoneal sheath, ovarioles and spermatheca. This video shows 25 s of the data analysed in Fig. 7, resampled at 1.5 frames/second. QuickTime format video (created with MS-Windows v6.50; other formats of this video available online ). (MOV 932 KB)
Each ovariole moves independently within the peritoneal sheath, with the mean rate being 12.5 ± 6.4 contractions/minute (mean ± 95% confidence interval; mode 10 contractions/minute; 17 ovarioles measured in 6 preparations). There is a very diverse pattern of contractions: some move almost continuously, with up to 53 contractions/minute (Fig. 7A pink trace), others with regular bouts of activity and others remaining largely quiescent (light blue trace, 5 of the 17 ovarioles showed less than 2 contractions/minute). As shown by the traces in Fig. 7(A,B), different ovarioles with a single ovary behave quite differently
The largest recorded movements are those of the spermathecae. These are on the termini of a long thin duct, which contracts or rotates the spermatheca, so that its movements are always very large. In 7-day-old flies these show regular contractions (1.5 ± 0.29/minute, N = 6) with individual contractions lasting 5–6s.
The Fast Fourier Transform (FFT, Fig. 7B) reveals that the spermathecae have the greatest power density at 2.8 contractions/minute, owing to their freedom to move with large amplitude as their ducts contract and rotate. The spermatheca FFT has prominent sidebands, as would be expected from their tendency to remain contracted for 5–6s. The oviduct (and the peritoneal sheath) share many peaks with the spermatheca, as they are tightly mechanically coupled in this isolated preparation. The peritoneal sheath contractions show a series of small peaks in the spectrum from 10 to 40 contractions/minute, reflecting endogenous peritoneal sheath contractions, some shared with the oviduct. The ovarioles have a different peak power density, in the range 3 to 16 contractions/minute.
We have begun our pharmacological analysis by examining the response of the peritoneal sheath to octopamine and tyramine because the contractions of the peritoneal sheath are an essential first step in the movement of the egg along the reproductive tract. We used the ovary and oviduct preparation to avoid contamination of the traces by spermathecal or uterine contractions. The mean duration of the samples was 145 ± 8 s.
Muscle movements of the genital tract
The female reproductive tract is a muscular system, in which contractions are linked to movement of the eggs down the tract from ovarioles via the oviducts to the uterus and finally oviposition of the fertilised eggs. We have described circular fibres in the muscle layers of tubular tissues of the Drosophila reproductive tract (ovarioles, oviduct and uterus). In the walls of the larval and adult gut and in some other insect duct systems, the layers of circular muscles are frequently opposed by longitudinal muscles . Longitudinal fibres were apparently detected during development of the Drosophila ovary , and have been found, along with circular myofibres, in the ovaries of a number of Lepidoptera including the silk moth Hyalophora cecropia , the flour moth Ephestia kühniella , the sugar cane borer Diatraea saccharalis  and the butterfly Calpodes ethlius . However, we have found no evidence for the presence of longitudinal fibres in the female Drosophila genital tract using electron microscopy, immunocytochemistry or directed GFP expression. This is consistent with the movements measured in our ovariograms, where peristaltic waves, but not shortening, were observed. The Drosophila ovary is covered by a peritoneal sheath, which as noted by others [9, 10] contains a network of muscle fibres. Similar sheaths have been reported from other Diptera  and may perform a similar function to that of the longitudinal fibres described in other insects; the Lepidopteran ovaries lack peritoneal sheaths. The neural co-ordination of muscle contractions in the Drosophila peritoneal sheath and various tubular muscles are likely to be important for egg development and oviposition. These contractions may also be important for moving the haemolymph around the reproductive tract, especially within the ovary during the energetically and nutritionally demanding process of oogenesis.
Our anti-HRP immunostaining showed that the nerves to the ovary run along the peritoneal sheath network, outside the muscle layer rather than among its myofibrils. We have detected no direct innervation of the myofibres of the ovariolar epithelial sheath. Thus it seems likely that the contractions of the ovariole sheath are controlled by a (local) hormonal mechanism. This would fit with the persistence of spontaneous myogenic contractile activity in the isolated ovariole.
The peritoneal sheath and lateral oviduct have a common pattern of innervation, in which the nerve fibres make oval varicosities, both along their length and at their endings. This is much closer to the type II neuromuscular junction boutons described in adult thoracic muscles  than to the type I. We propose that they are modulatory terminals: a suggestion supported by the lack of the Shaker channels, which are usually seen opposite type I terminals , as is glutamate receptor immunostaining. The lack of Shaker and glutamate fluorescence was not due to poor staining as the common oviduct and uterus of the same specimens consistently showed them at similar confocal settings. We have direct evidence that these nerves are all aminergic using two markers for tyramine and octopamine synthesising neurons. First, we used a brighter GFP to extend previous data , showing the presence of dTdc2 GFP marker on the peritoneal sheath  by demonstrating that this marker highlights not just some but all the nerves running over the surface of the peritoneal sheath. As tyrosine decarboxylase (TDC) is essential for the production of both tyramine and octopamine the implication is that these are all tyramine and/or octopamine releasing neurons. Secondly, we find that all the terminals are immunostained with the Drosophila vesicular monoamine transporter (DVMAT) antiserum . There is a difference in intracellular location of dTdc2 GFP and DVMAT staining: the dTdc2 GFP co-localises with HRP marker along the length of the neurons, while the DVMAT is only seen at and between varicosities. The ovarial innervation was detected with an antiserum to tyramine β-hydroxylase, the enzyme that converts tyramine to octopamine . An antiserum to the OAMB octopamine receptor shows staining in the lateral and common oviducts, in the peritoneal sheath and possibly in the ovarioles . Taken together these observations further support the contention that these neurons release octopamine and or tyramine at the peritoneal sheath and our ovariograms confirm that octopamine increases peritoneal sheath contractions. Although the fibres seen by these approaches are the same as we find by anti-HRP staining, it remains possible that not all the neurons use tyramine/octopamine, and that a proportion of the neurons use other transmitters or use a co-transmitter (e.g. a peptide) to modulate the activity of the ovary. Anatomically similar endings have been described on the hindgut and shown to be proctolin-immunoreactive .
The common oviduct and uterus show a second pattern of innervation, where nerves, ending as round boutons, on the muscle fibre layers were common. Nerve and muscle are arranged systematically, with nerve branches running parallel to the circular muscle fibres. Here we found glutamate receptors (GluRA; though other subtypes may be present). Our detection of these receptors and of the post-synaptic Shaker channel under the blebs, both typical of adult type I neuromuscular junctions , coupled with synaptotagmin immunostaining of the fine endings in the muscle layer , suggests conventional neuromuscular junctions rather than those associated with neurohormonal release. In the oviduct, we also observed modulatory endings very similar to those found on the peritoneal sheath, in agreement with the dTdc2 or tyramine β-hydroxylase staining [4, 24]. Modulatory endings in the luminal layers of the uterine muscle have not been reported previously.
Observations of movement in isolated ovaries has been reported previously using casual observations or videotape [16, 21]. Our technique to quantify the movement of the Drosophila reproductive tract shows persistent, steady recordings in HL-3 saline. Discrimination of movement of different parts of the reproductive tract shows that the ovarioles, the peritoneal sheath of the ovary, oviduct and spermatheca all generate independent contractions. In each organ, waves of contraction come in bursts, separated by longer quiescent periods. These have all been recorded in isolation from the CNS and so are myogenic in origin. This is confirmed by the persistence of ovariolar rhythms in the isolated ovariole, and by the continued contraction of the peritoneal sheath in the isolated ovary. Myogenic rhythms have been recorded from a range of insect visceral tissues, including the gut and reproductive organs. In the gut, different regions display independent rhythms, where bath-applied amines and peptides modulate large changes in the amplitude and frequency of contractions [27, 28].
Our data indicate that octopamine significantly increases the amplitude of the peritoneal sheath contractions. In locusts (Locusta migratoria), octopamine reduces the spontaneous contractions of the oviduct [29, 30]. It diminishes both the frequency of the myogenic rhythm and basal tone. In addition, octopamine reduces the proctolin-induced contractions of the oviduct, with 50% block between 10-6 and 10-7 M. Similar data come from a dipteran, the stable fly (Stomoxys), in which octopamine also causes a reduction in the spontaneous and proctolin-induced contractions of the oviduct .
Our data have led us to a new model in which tissue-specific differences are important. Octopamine increases the strength of contractions of the peritoneal sheath, which will facilitate ovulation of the egg. At the same time, we expect octopamine to relax the oviduct, so that the egg can move into it more easily. Thus, one neuromodulator acts on two neighbouring zones with opposing effects to enhance reproductive success. This hypothesis is in accord with the locust oviduct data, and also with Stomoxys, where 10–100 nM octopamine increases the amplitude of ovarian contractions but relaxes the oviduct [21, 31].
This model resolves a paradox. Previous authors found it hard to reconcile observations that flies lacking octopamine (owing to the M18 null mutation in the tyramine β-hydroxylase gene) or with a null mutation in the Oamb type of octopamine receptor lay fewer eggs than wild type [6, 24] with the fact that octopamine reduces reproductive tract contractions. Our observations provide a simple explanation for the loss of egg-laying in octopamine-free flies: we suggest that octopamine is likely to be released during egg-laying behaviour to enhance the strength of ovarian contractions and to relax the oviduct and so speed the egg along the reproductive tract. Octopamine may also affect the spermatheca; in locusts the spermathecal contractions were increased by octopamine , and the oviduct relaxed .
Finally, we showed that tyramine had no effect on the amplitude of the contractions. This is different from the response of the locust oviduct, where tyramine had a very similar inhibitory effect to octopamine on the contraction frequency and basal tone . In locust, the effect of octopamine was mediated by cAMP, but the tyramine response did not depend on a cAMP pathway except at very high (10-4 M) concentrations. This suggests a difference in receptor activation.
In flies, several "octopamine" receptors have been reported. One sensitive to both tyramine and octopamine (TyrR = hono) [34, 35] is unlikely to be the receptor on the peritoneal sheath, which was only sensitive to octopamine. At least four receptors (Oamb and the DmOctβ receptors [36–38]) are two orders of magnitude more sensitive to octopamine than tyramine, in accord with our pharmacology, and so may be present on the peritoneal sheath. Although Oamb has been localised on the oviduct , the receptor on the peritoneal sheath awaits identification.
The reproductive tract of the female fly has only circular muscles and no longitudinal muscles. They are supercontractile, which is appropriate for a system that has to distend when the egg passes through. The muscles have complex, bursting myogenic rhythms, with independent oscillators in the ovarioles, in the peritoneal sheath around the ovary and in the spermatheca. The ovary is innervated solely by modulatory neurons, which are tyraminergic and/or octopaminergic. The oviduct and uterus are innervated both by similar aminergic fibres and by glutamatergic neurons with endings similar to type I neuromuscular junction. Octopamine, but not tyramine, modulates the peritoneal sheath around the ovary, increasing the strength of the contraction, and we propose it also relaxes the oviduct. This double effect eases the ovulation of the egg from the ovary to the oviduct.
Flies were maintained at 25°C on standard yeast-sugar-agar medium. Wild type was Canton-S. Some observations used flies from the Wee-P26 stock , which carries a green fluorescent protein (GFP) coding sequence inserted into the myosin heavy chain gene, Mhc. We used a stock in which the CD8-GFP-Shaker protein is driven by a Mhc gene promoter . This expresses Shaker-GFP in the post-synaptic densities at the type I neuromuscular junctions of the nerves ending on the muscles. To visualise the synapses of neurons expressing tyrosine decarboxylase 2 (TDC2), we used the dTdc2-GAL4 construct  to drive expression of UAS-n-syb-spH, an enhanced GFP fused to neuronal synaptobrevin .
Immunocytochemistry and microscopy
Reproductive tracts from 7-day old females were fixed at room temperature for 30 min in 4% paraformaldehyde in phosphate buffered saline (PBS) or for 3 minutes in Bouin's solution (Sigma-Aldrich, UK), washed twice with PBS and incubated in two changes of 0.5% Triton-X100 in PBS for 30 minutes. After further PBS washes, specimens were treated with antibodies or phalloidin-TRITC. Some treatments were applied sequentially to provide a double label. Dilutions and washes were made in PBS.
Actin was labelled by incubation in 50 nM TRITC-labelled phalloidin (Sigma-Aldrich) for 2 h at room temperature. Myosin was labelled by incubation in rabbit anti-myosin heavy chain antibody (1:1000)  overnight at 4°C, followed, after washing, by a further 3 h incubation at room temperature in FITC-conjugated goat anti-rabbit IgG antibody (1:250) (Sigma-Aldrich). Vesicular monoamine transporter was labelled by incubation in rabbit anti-Drosophila vesicular monoamine transporter antibody (anti-DVMAT-A, 1:500; ) overnight at 4°C followed by washing and incubation in FITC-conjugated goat-anti rabbit IgG antibody as described above. Glutamate receptors were labelled as described  by incubation with mouse anti-glutamate receptor (dGluRIIA, 1:5) , overnight at 4°C, followed, after washing, by a further 3 h incubation at room temperature in FITC-conjugated goat anti-mouse IgG antibody (1:250) (Sigma-Aldrich). Nerve fibres were labelled by incubation in Cy3-conjugated anti-horseradish peroxidase antibody (anti-HRP, 1:1000, Jackson ImmunoResearch, West Grove, PA, USA) for 2 h at room temperature.
After labelling, all specimens were washed in PBS and mounted in Vectashield (Vector Laboratories, Burlingame, CA, USA). Confocal images were obtained with a Zeiss LSM510 META mounted on a Zeiss Axioplan 2 M microscope.
Samples of female reproductive tract were processed for electron microscopy as described for flight muscle .
Contractions of the isolated reproductive tract were recorded using video-microscopy followed by computer analysis of the movements from frame to frame. This approach produces an ovariogram in which the contractions at several sites on the same preparation can be recorded and compared.
Flies were transferred to vials within 24 hours of hatching and used on day 3 or day 7. Each vial contained both males and females, so our data are assumed to be from fertilised females. Complete female genital tracts were dissected into cold calcium-free HL-3 medium . In most experiments, the spermathecae and seminal receptacle were removed to avoid the large mechanical coupling between spermatheca and oviduct movements (Figs. 10, 11). In some experiments oviducts were also removed to facilitate ovariogram recordings from isolated ovaries (Fig. 8).
Preparations were mounted in fresh HL-3 saline (Ca concentration 1 mM) at 22°C on cavity microscope slides for 20 minutes before observations began. Data were obtained using a Nikon Labophot microscope with a 4× objective through a 5× photographic eyepiece with a Panasonic WV-CL10 camera. The video recordings were captured directly to a 1 MHz PC equipped with a Hauppauge Win-TV capture card (44805), using the VirtualDub 1.5.3 software package and the Huffyuv lossless compression algorithm (Huffyuv Codec) . Resolution was set at 640 × 480 pixels, capturing at 2 or 15 frames/second, average recording time 145 ± 8 s.
Video was analysed using a custom program (avi_line, ). In this, the mouse was used to overlay lines on the video frames, so that each line crossed the light/dark boundary between the preparation and the background. For each frame, the distance of the light/dark interface from the start of the line was determined, along with the mean squared difference in intensity between successive frames of the pixels along the line (Fig. 2). This records the displacement in the plane of focus, but any movement in the vertical direction is not measured. Data were saved in a Microsoft Excel format and mean peak height and intervals between peaks calculated. The Fast Fourier Transforms (FFT) were calculated by exporting the position of the light/dark interface into the fft program from Physionet .
In a few experiments, ovarioles from a single ovary were teased apart and video recorded as above but using a 10× phase contrast objective (Fig. 9).
After the initial 20 minute acclimatisation, the activity of the isolated reproductive tract was recorded for 2–3 minutes. The solution was then changed for one containing octopamine or tyramine and a further 2–3 minutes recorded. The control HL-3 solution was then applied for another 2–3 minutes recording. Because of disk space limitations, we were unable to record for longer, and this may affect the accuracy of our measurements of the frequency of contraction. Amines and HL-3 saline components were from Sigma-Aldrich, UK.
For each preparation, the mean amplitude and interval between contractions were calculated. Differences between these means were tested in Minitab, using Mann-Whitney and Student's t-tests. All data are reported as mean ± 95% confidence interval.
We are grateful to The Wellcome Trust for their support (KP). We are grateful to Graeme Davis, Shannon Cole, Jay Hirsh and Corey Goodman for fly stocks. We also thank David Krantz, Dan Kiehart and Sandy Bernstein for antibodies. The glutamate receptor antibody was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by The University of Iowa, Department of Biological Sciences, Iowa City, IA 52242.
- Roeder T, Seifert M, Kahler C, Gewecke M: Tyramine and octopamine: Antagonistic modulators of behavior and metabolism. Arch Insect Biochem Physiol. 2003, 54: 1-13. 10.1002/arch.10102.View ArticlePubMedGoogle Scholar
- Roeder T: Octopamine in invertebrates. Prog Neurobiol. 1999, 59: 533-561. 10.1016/S0301-0082(99)00016-7.View ArticlePubMedGoogle Scholar
- Monastirioti M, Linn CE, White K: Characterization of Drosophila tyramine beta-hydroxylase gene and isolation of mutant flies lacking octopamine. J Neurosci. 1996, 16: 3900-3911.PubMedGoogle Scholar
- Cole SH, Carney GE, McClung CA, Willard SS, Taylor BJ, Hirsh J: Two functional but noncomplementing Drosophila tyrosine decarboxylase genes: distinct roles for neural tyramine and octopamine in female fertility. J Biol Chem. 2005, 280: 14948-14955. 10.1074/jbc.M414197200.View ArticlePubMedGoogle Scholar
- Kutsukake M, Komatsu A, Yamamoto D, Ishiwa-Chigusa S: A tyramine receptor gene mutation causes a defective olfactory behavior in Drosophila melanogaster. Gene. 2000, 245: 31-42. 10.1016/S0378-1119(99)00569-7.View ArticlePubMedGoogle Scholar
- Lee HG, Seong CS, Kim YC, Davis RL, Han KA: Octopamine receptor OAMB is required for ovulation in Drosophila melanogaster. Dev Biol. 2003, 264: 179-190. 10.1016/j.ydbio.2003.07.018.View ArticlePubMedGoogle Scholar
- Zumstein N, Forman O, Nongthomba U, Sparrow JC, Elliott CJH: Distance and force production during jumping in wild type and mutant Drosophila melanogaster. J Exp Biol. 2004, 207: 3515-3522. 10.1242/jeb.01181.View ArticlePubMedGoogle Scholar
- Grossfield J: Non-sexual behavior of Drosophila. The Genetics and Biology of Drosophila. Edited by: Ashburner M, Wright TRF. 1978, London: Academic Press, 1-126.Google Scholar
- King RC: Ovarian development in Drosophila melanogaster. 1970, London: Academic pressGoogle Scholar
- Mahowald AP, Kambysellis MP: Oogenesis. The Genetics and Biology of Drosophila vol 2d. Edited by: Ashburner M, Wright TRF. 1980, London: Academic PressGoogle Scholar
- Crossley AC: The morphology and development of the Drosophila muscular system. The Genetics and Biology of Drosophila. Edited by: Ashburner M, Wright TRF. 1978, London: Academic PressGoogle Scholar
- Cook BJ, Pryor NW: Structural characterization of peripheral nerve cells and nerve-muscle junctions of the oviduct of stable fly (Diptera: Muscidae). J med Ent. 1996, 33: 496-503.View ArticleGoogle Scholar
- Hertweck H: Anatomie und Variabität des Nervensystems und der Sinnesorgane von Drosophila melanogaster (Meigen). Z wiss Zool. 1931, 139: 559-663.Google Scholar
- Bellen HJ, Budnik V: The neuromuscular junction. Drosophila protocols. Edited by: Sullivan W, Ashburner M, Hawley RS. 2000, Cold Spring Harbor, New York: Cold Spring Harbour Press, 175-199.Google Scholar
- Rivlin PK, St Clair RM, Vilinsky I, Deitcher DL: Morphology and molecular organization of the adult neuromuscular junction of Drosophila. J Comp Neurol. 2004, 468: 596-613. 10.1002/cne.10977.View ArticlePubMedGoogle Scholar
- Büning J: The Insect Ovary. 1994, London: Chapman & HallView ArticleGoogle Scholar
- King RC, Agarwal SK: Oogenesis in Hyalophora cecropia. Growth. 1965, 29: 17-83.PubMedGoogle Scholar
- Cruickshank WJ: The ultrastructure and functions of the ovariole sheath and tunica propria in the flour moth. J Insect Physiol. 1973, 19: 577-587. 10.1016/0022-1910(73)90067-X.View ArticleGoogle Scholar
- dos Santos DC, Gregorio EA: Ultrastructure of the ovariole sheath in Diatraea saccharalis (Lepidoptera: Pyralidae). Biocell. 2002, 26: 229-235.PubMedGoogle Scholar
- Griffith CM, Lai-Fook J: The ovaries and changes in their structural components at the end of vitellogenesis and during vitelline membrane formation in the butterfly, Calpodes. Tiss Cell. 1986, 18: 575-588. 10.1016/0040-8166(86)90022-4.View ArticleGoogle Scholar
- Cook BJ, Peterson T: Ovarian muscularis of the stable fly Stomoxys calcitrans – Its structural, motile, and pharmacological properties. Arch Insect Biochem Physiol. 1989, 12: 15-30. 10.1002/arch.940120103.View ArticleGoogle Scholar
- Zito K, Fetter RD, Goodman CS, Isacoff EY: Synaptic Clustering of Fasciclin II and Shaker: Essential Targeting Sequences and Role of Dlg. Neuron. 1997, 19: 1007-1016. 10.1016/S0896-6273(00)80393-1.View ArticlePubMedGoogle Scholar
- Greer CL, Grygoruk A, Patton DE, Ley B, Romero-Calderon R, Chang HY, Houshyar R, Bainton RJ, Diantonio A, Krantz , et al: A splice variant of the Drosophila vesicular monoamine transporter contains a conserved trafficking domain and functions in the storage of dopamine, serotonin, and octopamine. J Neurobiol. 2005, 64: 239-258. 10.1002/neu.20146.View ArticlePubMedGoogle Scholar
- Monastirioti M: Distinct octopamine cell population residing in the CNS abdominal ganglion controls ovulation in Drosophila melanogaster. Dev Biol. 2003, 264: 38-49. 10.1016/j.ydbio.2003.07.019.View ArticlePubMedGoogle Scholar
- Anderson MS, Halpern ME, Keshishian H: Identification of the neuropeptide transmitter proctolin in Drosophila larvae: characterization of muscle fiber-specific neuromuscular endings. J Neurosci. 1988, 8: 242-255.PubMedGoogle Scholar
- Finley KD, Edeen PT, Foss M, Gross E, Ghbeish N, Palmer RH, Taylor BJ, McKeown M: dissatisfaction e ncodes a tailless-like nuclear receptor expressed in a subset of CNS neurons controlling Drosophila sexual behavior. Neuron. 1998, 21: 1363-1374. 10.1016/S0896-6273(00)80655-8.View ArticlePubMedGoogle Scholar
- Nassel DR: Peptidergic neurohormonal control systems in invertebrates. Curr Opin Neurobiol. 1996, 6: 842-850. 10.1016/S0959-4388(96)80036-5.View ArticlePubMedGoogle Scholar
- Hertel W, Pass G: An evolutionary treatment of the morphology and physiology of circulatory organs in insects. Comp Biochem Physiol A. 2002, 133: 555-575. 10.1016/S1095-6433(02)00251-9.View ArticleGoogle Scholar
- Lange AB, Tsang PKC: Biochemical and physiological effects of octopamine and selected octopamine agonists on the oviducts of Locusta migratoria. J Insect Physiol. 1993, 39: 393-400. 10.1016/0022-1910(93)90027-O.View ArticleGoogle Scholar
- Nykamp DA, Lange AB: Interaction between octopamine and proctolin on the oviducts of Locusta migratoria. J Insect Physiol. 2000, 46: 809-816. 10.1016/S0022-1910(99)00170-5.View ArticlePubMedGoogle Scholar
- Cook BJ, Wagner RM: Some pharmacological properties of the oviduct muscularis of the stable fly, Stomoxys calcitrans. Comp Biochem Physiol C. 1992, 102: 273-280. 10.1016/0742-8413(92)90111-J.View ArticlePubMedGoogle Scholar
- Clark J, Lange AB: Octopamine modulates spermathecal muscle contractions in Locusta migratoria. J Comp Physiol A Neuroethol Sens Neural Behav Physiol. 2003, 189: 105-114.PubMedGoogle Scholar
- Donini A, Lange AB: Evidence for a possible neurotransmitter/neuromodulator role of tyramine on the locust oviducts. J Insect Physiol. 2004, 50: 351-361. 10.1016/j.jinsphys.2004.02.005.View ArticlePubMedGoogle Scholar
- Saudou F, Amlaiky N, Plassat JL, Borrelli E, Hen R: Cloning and characterization of a Drosophila tyramine receptor. EMBO J. 1990, 9: 3611-3617.PubMed CentralPubMedGoogle Scholar
- Reale V, Hannan F, Midgley JM, Evans PD: The expression of a cloned Drosophila octopamine/tyramine receptor in Xenopus oocytes. Brain Research. 1997, 769: 309-320. 10.1016/S0006-8993(97)00723-3.View ArticlePubMedGoogle Scholar
- Han KA, Millar NS, Davis RL: A novel octopamine receptor with preferential expression in Drosophila mushroom bodies. J Neurosci. 1998, 18: 3650-3658.PubMedGoogle Scholar
- Balfanz S, Strunker T, Frings S, Baumann A: A family of octapamine receptors that specifically induce cyclic AMP production or Ca2+ release in Drosophila melanogaster. J Neurochem. 2005, 93: 440-451. 10.1111/j.1471-4159.2005.03034.x.View ArticlePubMedGoogle Scholar
- Maqueira B, Chatwin H, Evans PD: Identification and characterization of a novel family of Drosophila β-adrenergic-like octopamine G-protein coupled receptors. J Neurochem. 2005, 94: 547-560. 10.1111/j.1471-4159.2005.03251.x.View ArticlePubMedGoogle Scholar
- Clyne PJ, Brotman JS, Sweeney ST, Davis G: Green fluorescent protein tagging Drosophila proteins at their native genomic loci with small P elements. Genetics. 2003, 165: 1433-1441.PubMed CentralPubMedGoogle Scholar
- Poskanzer KE, Marek KW, Sweeney ST, Davis GW: Synaptotagmin I is necessary for compensatory synaptic vesicle endocytosis in vivo. Nature. 2003, 426: 559-563. 10.1038/nature02184.View ArticlePubMedGoogle Scholar
- Kiehart DP, Feghali R: Cytoplasmic myosin from Drosophila melanogaster. J Cell Biol. 1986, 103: 1517-1525. 10.1083/jcb.103.4.1517.View ArticlePubMedGoogle Scholar
- Renden RB, Broadie K: Mutation and activation of Gα s similarly alters pre- and postsynaptic mechanisms modulating neurotransmission. J Neurophysiol. 2003, 89: 2620-2638. 10.1152/jn.01072.2002.View ArticlePubMedGoogle Scholar
- Schuster CM, Ultsch A, Schloss P, Cox JA, Schmitt B, Betz H: Molecular cloning of an invertebrate glutamate receptor subunit expressed in Drosophila muscle. Science. 1991, 254: 112-114.View ArticlePubMedGoogle Scholar
- Kronert WA, O'Donnell PT, Fieck A, Lawn A, Vigoreaux JO, Sparrow JC, Bernstein SI: Defects in the Drosophila myosin rod permit sarcomere assembly but cause flight muscle degeneration. J mol Biol. 1995, 249: 111-125. 10.1006/jmbi.1995.0283.View ArticlePubMedGoogle Scholar
- Stewart BA, Atwood HL, Renger JJ, Wang J, Wu CF: Improved stability of Drosophila larval neuromuscular preparations in haemolymph-like physiological solutions. J comp Physiol A. 1994, 175: 179-191. 10.1007/BF00215114.View ArticlePubMedGoogle Scholar
- Huffyuv. [http://neuron2.net/www.math.berkeley.edu/benrg/huffyuv.html]
- AviLine. [http://biolpc22.york.ac.uk/avianal/avi_line/]
- Goldberger AL, Amaral LAN, Glass L, Hausdorff JM, Ivanov PC, Mark RG, Mietus JE, Moody GB, Peng CK, Stanley HE: PhysioBank, PhysioToolkit, and PhysioNet : Components of a New Research Resource for Complex Physiologic Signals. Circulation. 2000, 101: e215-e222.View ArticlePubMedGoogle Scholar
- Miller A: The internal anatomy and histology of imago of Drosophila melanogaster. Biology of Drosophila. Edited by: Demerec M. 1950, New York: Wiley, 424-442.Google Scholar
- Fly Ovary. [http://biolpc22.york.ac.uk/drosophila/ovary/]
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