Maturation of muscle properties and its hormonal control in an adult insect
1 Abteilung Neurobiologie, Universität Ulm, Albert-Einstein-Allee 11, D-89069 Ulm, Germany and
2 Institut für Zoologie und Anthropologie der Universität Göttingen, Berlinerstraße 28, D-37073 Göttingen, Germany
*e-mail: uwe.rose{at}biologie.uni-ulm.de
Accepted July 13, 2001
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: juvenile hormone, muscle properties, reproductive development, oviposition, insect, Locusta migratoria, development.
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Juvenile hormone, first discovered by Wigglesworth (Wigglesworth, 1934; Wigglesworth, 1936), plays a crucial role in the development and reproduction of insects. During larval and pupal development, JH acts in concert with 20-hydroxyecdysone (20E) to govern metamorphosis by inhibiting the development of adult characters (Riddiford, 1985). However, considerable information has accumulated suggesting that the original role of JH may have been the regulation of reproduction (Sehnal et al., 1996). Many insects use JH to stimulate or depress reproductive development, although the extent and timing of JH action vary considerably (for reviews, see Wyatt and Davey, 1996; Wyatt, 1997).
Reproduction in insects is often accompanied by the expression of specific behaviour that is, in many cases, directly or indirectly regulated by JH. Generally, the effects of JH are diverse, but they predominantly involve courtship behaviour, female sexual receptivity or egg-laying behaviour (Barth and Lester, 1973; Strong and Amerasinghe, 1977; Renucci et al., 1992). One of the most intriguing studies described a direct effect of JH on the neuronal elements responsible for the phonotactic response in female Acheta domesticus (Stout et al., 1993). This effect is mediated by JH via gene regulation (Stout et al., 1992). In the same species, Cayre et al. (Cayre et al., 1994) showed that neurogenesis in the mushroom bodies of adult crickets is stimulated by JH, and this might be the basis for hormonal control of oviposition behaviour. Males are less often dependent on the action of JH, although a few studies provide clear evidence for effects of JH in males. Odhiambo (Odhiambo, 1966) reported depressed locomotory activity in male locusts after allatectomy. After re-implantation of the corpora allata (CA), normal activity was restored. In the black cut worm Agrotis ipsilon, JH clearly affects male responsiveness to the female sex pheromone (Gadenne et al., 1993).
Analysis of the mechanisms of hormone action requires a detailed knowledge of the developmental changes mediated by hormones. From previous studies, two primary mechanisms of JH action have emerged. First, JH has been shown to bind directly to the membrane and to mediate its effects within minutes without the need for gene transcription (Sevala and Davey, 1989; Yamamoto et al., 1988). The second major mechanism involves gene transcription. Here, JH is thought to modulate gene transcription after it has penetrated the cells and after binding to proteins or nuclei (Jones, 1995; Dubrovsky et al., 2000; Davey, 2000). Several genes have been cloned whose expression is clearly regulated by JH (Wyatt and Davey, 1996). However, the search for a JH receptor has so far been unsuccessful, although the nuclear receptor ultraspiracle, which binds JH with specificity, has been proposed as a good candidate (Jones and Sharp, 1997).
In some insect species, the flight muscles undergo degeneration as a response to high levels of JH (Tanaka, 1994; Davis, 1975; Borden and Slater, 1968; Stegwee et al., 1963). During reproduction, they are no longer needed and now serve to liberate nutrients. In contrast, the Colorado potato beetle (Leptinotarsa decemlineata) undergoes a reproductive diapause that results from CA inactivity leading to low levels of JH. During diapause, the flight muscles undergo reversible degeneration (de Kort, 1990; Pener, 1992). However, no direct hormonal regulation of growth and functional maturation of the flight muscles has been demonstrated (Finlayson, 1975).
To determine whether JH has the potential to alter the functional properties of muscle fibres, we investigated locust abdominal longitudinal muscles. These multifunctional muscles are involved in ventilation (Hustert, 1975), flight steering (Baader, 1991) and oviposition (Vincent, 1975; Jorgensen and Rice, 1983a; Rose et al., 2000). During oviposition, females use their abdomen as an ovipositor; they extend their segments in a telescopic manner to up to six times its normal length. During this highly coordinated behaviour (Rose et al., 2000), telescopic extension mainly involves abdominal segments 47 in which the longitudinal muscles are required to follow and tolerate lengthening (superextension). In an ultrastructural study, Jorgensen and Rice (Jorgensen and Rice, 1983a) demonstrated pronounced differences between the longitudinal muscles present in abdominal segments 47 (oviposition segments) and those in the more anterior segments 13 (non-oviposition segments). One of the unique features of muscles from oviposition segments is their ability to fragment their Z-lines into so-called Z-bodies. Since the ability to tolerate extension is closely related to the sexual maturation of female locusts, we investigated whether this ability develops during adult life or is already present at adult emergence. We also explored the possible gender- and/or segment-specificity of muscle properties and whether JH is involved in the regulation of these properties. Our results indicate that, during reproductive development, the properties of the longitudinal muscle of females are subject to change, possibly to adapt them for oviposition behaviour. These properties are segment- and gender-specific and are controlled by JH, as indicated by experiments examining the inhibition of JH release and JH replacement injections.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Electrophysiology
Extracellular recordings from peripheral nerves and stimulation of motor axons were obtained using suction or monopolar hook electrodes. Muscle contractions were evoked by stimulating (0.5 ms, 0.55 V) the motor axons. To monitor stimulation-evoked action potentials, the terminal branch innervating the muscle was recorded en passant.
For two-electrode current-clamp recordings, muscles were isolated and mounted with the internal side up in a recording chamber. The internal side of longitudinal muscles is directed towards the gut, whereas the external side is adjacent to the body wall. The preparation was continuously perfused (1 ml min1) with saline (see above). Current and voltage electrodes were made from thin-walled borosilicate glass (Clark Electromedical Instruments, UK) and filled with 2 mol l1 potassium acetate containing 20 mmol l1 potassium chloride (resistance 1020 M). The current electrode was inserted in the middle of the fibre, whereas the voltage electrode was placed half-way between the end of the fibre and the current electrode. After impalement of muscle fibres, electrodes were allowed to seal for 15 min. A Ag/AgCl bath electrode served as potential reference. Only fibres from the internal layer were recorded. The signals were controlled and amplified by a two-electrode clamp amplifier (Turbo TEC01C, NPI Electronics, Tamm, Germany) connected to a personal computer. Current injection and acquisition of signals were controlled by a computer program (cell works, NPI electronics, Tamm, Germany). Acquired signals were digitised (sampling rate 10 kHz) and stored on hard disk for subsequent evaluation (Cell Works Reader, NPI Electronics, Tamm, Germany; PlotIt, Scientific Programming Enterprises).
Hormonal treatment
To inactivate the CA chemically, female locusts were treated once at the time of adult emergence by topical administration of precocene I (7-methoxy-2,2-dimethyl-3-chromene;, Sigma, 500 µg dissolved in 15 µl of acetone) onto the dorsal neck fold. Control animals were treated with acetone only (N=15). Approximately 95 % of precocene-treated animals survived the treatment and, at maturation, showed clear signs of CA degeneration (undeveloped oocytes, the cuticle remained light-coloured, no hypertrophy of the abdominal muscles was apparent). Some of the precocene-treated animals were also treated with JH [7.5 µg JH III (Sigma, 75 %) in 5 µl 70 % ethanol] or ethanol alone (N=10, control). JH III has been identified as the prevailing hormone during the gonotrophic cycle of locusts (Rembold, 1981). JH was first injected into the abdomen on day 5. Subsequent injections were made on days 8 and 11 to ensure that a sufficent titre of JH was present. This pattern of injection was chosen after preliminary experiments with a single injection or a smaller amount of JH did not induce signs of normal maturation (development of oocytes, cuticle darkly coloured). Although we have not determined the resulting level of juvenile hormone in the haemolymph, the amount injected was within or below the range of concentrations of JH or its analogue methoprene used in other studies [JH (Tawfik et al., 1997); methoprene (Dhadialla and Wyatt, 1983)]. The survival rate of JH-treated animals was approximately 70 %.
Tension recordings
Tension recordings were made from homologous dorsal longitudinal muscles of the sixth abdominal segment (see Fig. 1A, M214, Fig. 1B) or the third segment (M169). At its anterior insertion, the muscle was fixed to the Petri dish with insect pins. The posterior side was attached to the lever arm of a force transducer (Fort-10, World Precisions Instruments) mounted on a micromanipulator. This was achieved by clamping the piece of cuticle on which the muscle fibres insert to a small clamp made of stainless steel. This arrangement allowed precise lengthening of the muscle during the experiments without influencing muscle contractions. After each series of contractions, the preparation was repeatedly superfused with aerated saline. Between a series of contractions, the muscle was left unexcited for at least 5 min to recover from contractions and to adapt to length changes. Tension recordings were approximately isometric (deflection of the transducer tongue: 6 µm per 1 mN). The response of the transducer was linear over the range used in the experiments and was calibrated after each experiment.
|
Cross sections of muscle fibres and measurement of cross-sectional area
In preliminary experiments, we noted considerable hypertrophy of muscle fibres in female locusts during the first 2 weeks of adult life. To examine and quantify this observation, we measured the cross-sectional area of muscle fibres from females in abdominal segments 3 (M169) and 6 (M214). Freshly isolated muscles were pinned in a Petri dish lined with Sylgard and stretched to their resting length (3.5 mm for untreated females more than 18 days old; 2 mm for all other groups). The muscles were than fixed in 2 % glutaraldehyde, post-fixed with 2 % osmium tetroxide and subsequently dehydrated and embedded in Epon 812 (Fluka). Cross sections (0.5 and 1 µm) from the middle region of the muscle were cut on an ultramicrotome, mounted on slides and stained with Methylene Blue. The sections were then examined under a bright-field microscope. From each section, a digital picture was taken with the aid of a CCD camera (Sony ICX038AK, resolution 752x582 pixels). The mean cross-sectional area of a single fibre was determined by calculating the cross-sectional area of the entire muscle (Sigmascan Pro 5.0) and dividing by the number of fibres.
Statistical evaluation
Data are expressed as means and their standard errors (S.E.M.). Statistical significance was determined using non-parametric (MannWhitney U-test, KruskalWallis analysis of variance, ANOVA, on ranks) or, when criteria were met, parametric (one-way ANOVA) analysis. Post-hoc tests were employed for multiple comparisons (Dunns method, Tukey test). The significance level was set to P<0.05.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The length/tension relationship of longitudinal muscle 214 from female locusts less than 5 days old (Fig. 2A, N=10) revealed a peak at approximately 2 mm. Further lengthening resulted in a rapid decline in tension and eventually in breakage of muscles fibres (Fig. 2A, arrow). In all experiments, muscles from immature females did not tolerate lengths exceeding 4 mm. In contrast, muscles of females more than 18 days old exerted their maximum tension between 2.5 and 3.5 mm and tolerated stretching of more than 8 mm. At this length, twitch tension was still 35 % of the maximum tension (Fig. 2A, N=9). In four experiments, we stretched muscles up to 12 mm. In these experiments, the muscles showed no signs of damage and still generated considerable twitch tension (approximately 10 % of maximum). Our own observations suggest that the normal physiological range of longitudinal muscles is between 1 and 4 mm, and this is supported by the work of Jorgensen and Rice (Jorgensen and Rice, 1983b).
|
A comparison of the maximum tension exerted by the longitudinal muscles of females less than 5 days old and females more than 18 days old showed significant differences. Both twitch tension and tetanic tension (5, 10, 20 and 50 Hz) were approximately twice as high in muscles of females more than 18 days old (N=9) compared with females less than 5 days old (Fig. 2C; P<0.05, N=11). It was also evident that, during a single twitch, both the contraction time (the time from the beginning to the peak of tension) and the half-relaxation time were significantly increased in mature females (Table 1; P<0.05, N=6), and these increases were probably responsible for the decrease in the stimulation frequency required to elicit a smooth tetanus (Fig. 2D).
|
Approximately 90 % of the females treated with precocene on the day after adult emergence showed clear signs of impaired maturation (undeveloped oocytes, cuticle remained light coloured, no hypertrophy of abdominal muscles). Muscle 214 of mature precocene-treated animals had a length/tension curve that showed a steep increase to a maximum tension at approximately 2 mm (N=10). Additional extension resulted in a rapid decline in tension and eventual rupture of muscles fibres at 4.5 mm (Fig. 3A). The curve therefore resembled the length/tension relationship obtained from untreated females less than 5 days old (compare with Fig. 2A). Control animals (only acetone applied) showed no signs of impaired maturation (15 out of 15 locusts). Females (>18 days old, N=6) injected with both precocene and JH had well-developed oocytes and a darkly coloured cuticle that was most obvious at the site of JH injection. Muscle 214 of these animals tolerated extensions up to a length of 8 mm and above (some were extended up to 10 mm with no signs of muscle fibre damage). At approximately 8 mm, the muscles generated approximately 30 % of their maximum tension (Fig. 3A, precocene+JH). Control animals (precocene+ethanol) did not differ from animals treated with precocene alone with respect to their maturational status (10 out of 10 locusts, data not shown).
|
The measurement of maximum tension revealed a pronounced difference between the two groups. Muscles from females treated with precocene+JH (N=6) exerted tension that was 2.53.0 times greater than that of females treated with precocene alone (Fig. 3C, N=10). A comparison of the twitch kinetics revealed significantly increased twitch contraction times in precocene+JH-treated animals (Table 1, Fig. 3D; P<0.05, N=6). Although consistent differences were also apparent for the half-relaxation time, the values were not statistically different (P>0.05, N=6).
Because the effects of CA inactivation were obvious in females treated with precocene alone (cuticle coloration, undeveloped oocytes, massive fat body), but not in females treated with precocene+ JH (precocene might have failed to inactivate the CA, which would have been indistinguishable from the case in which precocene inactivated the CA and JH reversed this effect), we compared the morphology of the CA with that of non-treated females more than 18 days old (control, Fig. 3E, N=2). Females treated with precocene+JH were expected to have CA showing a noticeable atrophy (Pener et al., 1978). Freshly dissected CA from females more than 18 days old (precocene+JH) had atrophied dramatically compared with those of control animals (Fig. 3E), providing further evidence that precocene was indeed effective in females treated with precocene+JH.
Comparing the cross-sectional area of muscle fibres from females less than 5 days old with that of females more than 18 days old, we observed a pronounced hypertrophy (Fig. 4A,B, right-hand panel). The mean cross-sectional area of muscle fibres from females more than 18 days old was considerably greater (1213±119.3 µm2, N=9) than that of muscles of females less than 5 days old (285±35.3 µm2, N=7; P<0.05). The muscles of females treated with precocene hypertrophied to a lesser extent. These muscle fibres had a mean cross-sectional area of 697.2±61.8 µm2 (N=9), which was still significantly different from that of untreated females (P<0.05). Mean fibre numbers were not significant different for M214 (<5 days old, 68±5; >18 days old, 73±2; >18 days old, precocene-treated, 71±3) and M169 (<5 days old, 48±2; >18 days old, 42±3; 18 days old, precocene-treated, 42±2).
|
|
|
Segment-specificity of muscle properties
The toleration of extensive lengthening and the concomitant muscle properties were measured as described above for muscle 214 of the sixth abdominal segment, which is involved in telescopic extension during oviposition. To investigate whether similar properties and their changes can be found in homologous muscles of non-oviposition segments, we measured the properties of muscle 169 in the third abdominal segment (M169, non-oviposition segment) before and after locusts underwent reproductive development.
The length/tension relationship of muscle 169 was similar in females less than 5 days old (N=6) and in females more than 18 days old (Fig. 7A, N=9). Muscles from neither developmental stage tolerated extension of more than 44.5 mm length (Fig. 7A, arrows). At this length, twitch tension was almost negligible. The passive tension exerted by the muscles was consistently higher in females more than 18 days old (N=5) than in females less than 5 days old (Fig. 7B, N=6). A similar relationship was obtained for the maximum tension. Mean values were higher in females more than 18 days old (N=5) than in females less than 5 days old (Fig. 7C, N=6), although the difference was not significant (P>0.05).
|
The cross-sectional area of muscle 169 was found to increase in females more than 18 days old (Fig. 4A,B, left-hand panel). The mean cross-sectional area of muscles from females less than 5 days old was 380.3±35.7 µm2 (N=6), but increased to 750±76.5 µm2 (N=6) in females more than 18 days old. These values were significantly different (P<0.05). Females treated with precocene had cross-sectional areas that were similar to those of untreated females (618±30 µm2, N=8, Fig. 4A,B; P>0.05).
Muscle 169 was also able to generate action potentials, although we never observed spontaneous contractions of single muscle fibres. Potentials recorded from fibres of females less than 5 days old had a low threshold (approximately 45 mV), and the peak amplitude was 20 to 10 mV (Fig. 5C, N=3). After the females had attained maturity, the threshold for action potential generation increased to approximately 40 mV, and the peak amplitude was approximately 10 mV (Fig. 5C, N=3). The width of the action potential was considerably greater in mature females compared with females less than 5 days old (Fig. 5D). Muscle fibres from females treated with precocene were also able to generate action potentials (N=2). Their threshold was comparable with that of untreated females more than 18 days old. The peak amplitude was around 0 mV, and the width of the potentials was within the range of that of females less than 5 days old (Fig. 5D). The powerful afterhyperpolarisation that was characteristic of the fibres of muscle 214 (see Fig. 5A) was not evident in muscle 169 (N=3). The effect of Cd2+ on the generation of action potentials in M169 was similar to that on muscle 214: Cd2+ (50 µmol l1) blocked action potentials in all stages (N=2; data not shown).
Gender-specificity of muscle properties
To investigate whether the properties of muscle from males changed during reproductive development, we measured the contraction properties of muscle 214 from immature (<5 days old) and mature (>18 days old) male locusts.
Longitudinal muscles 214 of males were not able to tolerate stretch of more than 4 mm at either stage (Fig. 8A, arrows; <5 days old, N=9; >18 days old, N=7). Furthermore, the length/tension curves were nearly identical, with muscles exerting very little twitch tension at lengths between 3.5 and 4 mm.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The development of specific muscle properties during maturation depends on JH, as indicated by the experiments in which we chemically inactivated the CA with precocene. Despite its name, JH has been shown to have multiple effects in adult insects [for reviews, see (Wyatt, 1997; Wyatt and Davey, 1996)]. Most of these involve degeneration and regeneration of flight muscle (Chudacova and Gutmann, 1978; Stegwee et al., 1963). Our results, however, suggest various changes in contractile and membrane properties. These changes are correlated with maturation and are a prerequisite for successful oviposition behaviour.
Although the results of this study suggest an important role for JH, we have to be cautious in the interpretation of our results since we have not measured the effective concentration of JH in precocene-treated animals or those additionally injected with JH. In female locusts (Schistocerca gregaria, Locusta migratoria), the JH titre increased significantly during maturation, starting at the seventh day after adult emergence (Rembold, 1981; Tawfik et al., 2000). In male Schistocerca gregaria, JH titres are comparatively low, but increase gradually within the first 30 days after the final moult (Tawfik et al., 2000). It is possible that JH titres, rather than the simple presence or absence of JH, affected our results. For example, some active and passive properties (length/tension curve, time to peak contraction, twitch tetanus fusion frequency) of females treated with precocene+JH (Fig. 3) apparently differ from those of normal control females (Fig. 2). This might be due to inappropriate JH titres leading to impaired synchronisation of maturational events. It has been assumed that JH generally mediates its various effects through different pathways specified by different concentrations (Gäde et al., 1997). However, our experiments strongly suggest that at least the acquisition of specific muscle properties depends on the existence of JH.
During maturation, the muscle fibres apparently increased their cross-sectional area. This hypertrophy also results in the ability of muscle 214 to exert a dramatically increased maximum tension after completion of maturation. Similar processes have been reported during moulting of crustaceans. Here, the growth of the myofibrils from leg muscles may occur by addition of thick and thin filaments and by longitudinal myofibrillar splitting (El Haj et al., 1984). These processes seemed to be under the control of ecdysteroids, as suggested by elevated levels of RNA synthesis in muscles after ecdysteroid administration (Whiteley et al., 1992). Interestingly, polyamines, which are known to play a fundamental role in tissue growth and development [for a review, see (Morgan, 1999)], are involved in JH-dependent oviposition behaviour in crickets (Cayre et al., 1996) and in the mitogenic action of JH on adult insect neuroblasts (Cayre et al., 1997). Thus, polyamines might be involved in the hypertrophy and/or expression of longitudinal muscle properties of locusts, possibly by influencing the transcriptional or translational stages of protein synthesis or by acting as an intracellular messenger (Morgan, 1999).
The kinetics of twitch contraction determined for M214 changed significantly during the reproductive development of females (Table 1). Precocene treatment reduced contraction and half-relaxation times, whereas injection of JH caused an increase. In contrast, contraction and half-relaxation times did not increase in M169. The twitch contraction times determined for the asynchronous dorsoventral flight muscle of the bumblebee Bombus terrestris (58 ms) (Josephson and Ellington, 1997) and for the tymbal sound-producing muscle of cicadas (107 ms) (Josephson and Young, 1981) were less than or within the range of values determined in the present study, but we are not aware of quantitative data for twitch kinetics that are directly comparable with those obtained in the present study.
The kinetics of insect muscle contraction are determined mainly by the muscle fibre architecture (type of muscle fibres and their composition). Müller et al. (Müller et al., 1992) demonstrated a direct correlation between fibre type and contraction speed. Rapidly contracting fibres (fast-type) had a high mATPase activity, whereas slow fibre types exhibited a low mATPase activity. Thus, the changes in twitch kinetics in M214 during maturation might be due to changes in fibre type expression. In addition, the modulatory action of proctolin or octopamine might play a role. Octopamine increases the amplitude of neurally evoked contractions and speeds up the relaxation of skeletal muscles (OShea and Evans, 1979), and a neurone has been identified, which presumably releases octopamine, that supplies the longitudinal muscles in locusts (Ferber and Pflüger, 1990). Proctolin has also been shown to modulate the contraction of skeletal muscles [for a review, see (Orchard et al., 1989)]. Proctolin appears to be co-localised in motoneurons with the excitatory transmitter glutamate (Usherwood and Cull-Candy, 1975; OShea et al., 1985; Worden et al., 1985). The amplitude and kinetics of the contractions measured in this study might therefore be affected by the individual history of modulation or by maturation-dependent differences in the responsiveness of muscle fibres to modulatory substances such as proctolin or octopamine.
With the exception of action potential generation, JH seems to affect predominantly the longitudinal muscles in oviposition segments of females, since the maturation of muscle properties was less pronounced in muscles from non-oviposition segments (M169). However, a slight, but consistent, increase in the maximum force (Fig. 7C) and a significant hypertrophy (Fig. 4A,B) indicated that these muscle were also subject to maturational changes. Similar changes occurred during the maturation of muscle 214 in males. In M214 of precocene-treated females, the properties differed from those of females less than 5 days old (compare Fig. 2 and Fig. 3). These results suggest a developmental process that applies qualitatively to all longitudinal muscles under investigation but is quantitatively increased by JH in muscles involved in oviposition. This hypothesis is further supported by the proposal that the original role of JH may have been the regulation of reproduction (Sehnal et al., 1996; Tobe and Bendena, 1999). Medawar (Medawar, 1953) concludes that endocrine evolution is not an evolution of hormones, but an evolution of the uses to which they are put. Along this line of argument, we assume that the evolutionary pressure for the protection and survival of offspring led to the ability of specific intersegmental muscle to superextend. Since this ability is not needed before the time of egg-laying, it might have become regulated by JH. The specificity of JH for longitudinal muscles in ovipositional segments might be controlled through differential hormone receptor expression, although no JH receptor has so far been identified. Furthermore, we do not know whether JH acts via a direct or an indirect pathway.
The visual observation that muscle fibres from mature females contract spontaneously after mechanical stimulation (stretch or release) led us to investigate the underlying conductances. All muscle fibres investigated generated action potentials upon current injection. The action potentials were dependent on Ca2+, as demonstrated in experiments in which Cd2+ was added to or Ca2+ was omitted from the saline. There is good evidence that Ca2+ is the principal inward charge carrier for action potential generation in insect muscle fibres (Washio, 1972; Yamamoto et al., 1978; Ashcroft and Stanfield, 1982). Blocking of voltage-dependent Na+ channels by TTX had almost no effect on the shape of action potentials. Similar TTX-insensitive potentials were found in a tonic muscle fibre bundle of the locust hindleg (Burns and Usherwood, 1978). The sensitivity of action potentials to nifedipine suggests the involvement of an L-type Ca2+ channel. L-type Ca2+ channels, sensitive to dihydropyridines, have been characterized in vertebrates and invertebrates (Catterall et al., 1988, Triggle, 1990; Erxleben and Rathmayer, 1997; Gielow et al., 1995). However, we cannot exclude the participation of other types of Ca2+ channel in the generation of action potentials. Gielow et al. (Gielow et al., 1995) characterised two voltage-dependent Ca2+ currents (L- and T-type) in the body wall muscles of larval Drosophila melanogaster. These Ca2+ channels activate between 40 and 30 mV, which is within the range of activation observed in our experiments.
The observed differences in the amplitude, activation, width and afterhyperpolarisation of action potentials are related to the presence or absence of JH. The differences may be due to a differential expression of ion channels. In the nervous system, JH has been shown to influence the properties of identified neurons in the auditory system of crickets, in which the expression of the nicotinic acetylcholine receptor gene in an auditory neuron was increased in the presence of JH (Stout et al., 1992; Stout et al., 1993). This led to an improved directionality and decreased the threshold for phonotaxis of females more than 18 days old (Koudele et al., 1987; Walikonis et al., 1991). It is also likely that the hypertrophy of muscle fibres influenced the properties of the action potentials. When fibres increase their volume, the surface-to-volume ratio decreases. This slows the kinetics of underlying conductances (Hille, 1992) which should, in turn, reduce the amplitude of the action potential. However, our results suggest the opposite. It is possible that the increase in the surface area of the muscle fibres was independent of the volume, as suggested by the work of Jorgensen and Rice (Jorgensen and Rice, 1983a). They found highly corrugated sarcolemma and basal lamina in abdominal longitudinal muscles of mature female locusts. To gain further insight into the maturation of muscle properties, we have recently started to voltage-clamp longitudinal muscle fibres.
The reduced threshold of action potentials during reproductive development led us to assume a specific role for these potentials during oviposition. This role might be related (i) to a general activation (contraction) of muscle fibres by action potentials or (ii) to the Ca2+ influx into the muscle fibres. When fibres were stretched, twitches were visible as a result of the generation of action potentials. This activation of muscle fibres might prevent myosin and actin filaments from sliding apart when the muscle is stretched during oviposition and, thus, direct the mechanical forces to the Z-lines to enable their fragmentation. A similar mechanism was suggested by the work of Jorgensen and Rice (Jorgensen and Rice, 1983a). In addition, the influx of Ca2+ into the muscle fibres could activate a second-messenger system that might lead to the activation of enzymes or of additional intracellular events. The Ca2+-activated protease calpain, for example, digests kettin, a Z-line protein, and disrupts the Z-discs of striated muscle in insect flight muscle (Lakey et al., 1993). This process leads eventually to the disassembly of myofibrils. Similar events might be involved during superextension of longitudinal muscles.
The development and specificity of muscle fibre properties make this system well suited for further investigations of the cellular and possibly molecular changes underlying the action of JH. By comparing muscles from oviposition segments with homologous muscles from non-oviposition segments, we hope to gain further insight into how adaptive behaviour is hormonally controlled and achieved.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Ashcroft, F. M. and Stanfield, P. R. (1982). Calcium and potassium currents in muscle fibres of an insect (Carausius morosus). J. Physiol., Lond. 323, 93115.[Abstract]
Baader, A. (1991). The contribution of some neck and abdominal motoneurones in locust (Locusta migratoria) steering reactions. J. Insect Physiol. 37, 689697.
Barth, R. H. and Lester, L. J. (1973). Neuro-hormonal control of sexual behaviour in insects. Annu. Rev. Ent. 18, 445472.[Medline]
Bass, A. H. (1986). A hormone-sensitive communication system in an electric fish. J. Neurobiol. 17, 131156.[Medline]
Borden, J. H. and Slater, C. E. (1968). Induction of flight muscle degeneration by synthetic juvenile hormone in Ips confusus (Coleoptera: Scolytidae). Z. Vergl. Physiol. 61, 366368.
Bowers, W. S., Ohta, T., Cleere, J. S. and Marsella, P. A. (1976). Discovery of insect anti-juvenile hormone in plants. Science 193, 542574.[Medline]
Bullard, B. and Leonard, K. (1996). Modular proteins of insect muscle. Adv. Biophys. 33, 211222.[Medline]
Burns, M. D. and Usherwood, P. N. R. (1978). Mechanical properties of locust extensor tibiae muscles. Comp. Biochem. Physiol. 61A, 8595.
Catterall, W. A., Seagar, M. J. and Takahashi, M. (1988). Molecular properties of dihydropyridine-sensitive calcium channels in skeletal muscle. J. Biol. Chem. 263, 35353538.
Cayre, M., Strambi, C., Charpin, P., Augier, R., Renucci, M. and Strambi, A. (1996). Inhibition of polyamine biosynthesis alters oviposition behaviour in female crickets. Behav. Neurosci. 110, 11171125.[Medline]
Cayre, M., Strambi, C., Charpin, P., Augier, R. and Strambi, A. (1997). Specific requirement of putrescine for the mitogenic action of juvenile hormone on adult insect neuroblasts. Proc. Natl. Acad. Sci. USA 94, 82388242.
Cayre, M., Strambi, C. and Strambi, A. (1994). Neurogenesis in an adult insect brain and its hormonal control. Nature 368, 5759.
Chudacova, I. and Gutmann, E. (1978). Developmental changes of succinate dehydrogenase, ATPase and acid phosphatase activity in flight muscle of the normal and allatectomized adult cricket. Zool. Jb. Physiol. 82, 115.
Csernoch, L., Szentesi, P., Sárközi, S., Szegedi, C., Jona, I. and Kovács, L. (1999). Effects of tetracaine on sarcoplasmic calcium release in mammalian skeletal muscle fibres. J. Physiol., Lond. 515, 843857.
Davey, K. G. (2000). The modes of action of juvenile hormones: some questions we ought to ask. Insect Biochem. Mol. Biol. 30, 663669.[Medline]
Davis, N. T. (1975). Hormonal control of flight muscle histolysis in Dysdercus fulvoniger. Ann. Ent. Soc. Am. 68, 710714.
de Kort, C. A. D. (1990). Thirty-five years of diapause research with the Colorado potato beetle. Ent. Exp. Appl. 56, 113.
Dhadialla, T. S. and Wyatt, G. R. (1983). Juvenile hormone-dependent vitellogenin synthesis in Locusta migratoria fat body: inducibility related to sex and stage. Dev. Biol. 96, 436444.[Medline]
Dubrovsky, E. B., Dubrovskaya, V. A., Bilderback, A. L. and Berger, E. M. (2000). The isolation of two juvenile hormone-inducible genes in Drosophila melanogaster. Dev. Biol. 224, 486495.[Medline]
El Haj, A. J., Govind, C. K. and Houlihan, D. F. (1984). Growth of lobster leg muscle fibres over intermolt and molt. J. Crust. Biol. 4, 536545.
Erxleben, C. and Rathmayer, W. (1997). A dihydropyridine-sensitive voltage-dependent calcium channel in the sarcolemmal membrane of crustacean muscle. J. Gen. Physiol. 109, 313326.
Everts, M. E. (1996). Effects of thyroid hormones on contractility and cation transport in skeletal muscle. Acta Physiol. Scand. 156, 325333.[Medline]
Ferber, M. and Pflüger, H. J. (1990). Bilaterally projecting neurones in pregenital abdominal ganglia of the locust: Anatomy and peripheral targets. J. Comp. Neurol. 302, 447460.[Medline]
Finlayson, L. H. (1975). Development and degeneration. In Insect Muscle (ed. P. N. R. Usherwood), pp. 75149. London: Academic Press.
Gäde, G., Hoffmann, K. H. and Spring, J. H. (1997). Hormonal regulation in insects: facts, gaps and future directions. Physiol. Rev. 77, 9631032.
Gadenne, C., Renou, M. and Sreng, L. (1993). Hormonal control of pheromone responsiveness in the male black cutworm Agrotis ipsilon. Experientia 49, 721724.
Gielow, M. L., Gu, G. G. and Singh, S. (1995). Resolution and pharmacological analysis of the voltage-dependent calcium channels of Drosophila larval muscles. J. Neurosci. 15, 60856093.[Abstract]
Hardie, J. (1975). The tension/length relationship of an insect (Calliphora erythrocephala) supercontracting muscle. Experientia 32, 714716.
Hille, B. (1992). Ionic Channels of Excitable Membranes. Second edition. Sunderland, Massachusetts: Sinauer.
Hustert, R. (1974). Morphologie und Atembewegungen des 5. Abdominalsegments von Locusta migratoria migratorioides. Zool. Jb. Physiol. 78, 157174.
Hustert, R. (1975). Neuromuscular coordination and propioceptive control of rhythmical abdominal ventilation in intact Locusta migratoria migratorioides. J. Comp. Physiol. A 97, 159179.
Ishii, N. and Takahashi, K. (1982). Lengthtension relation of single smooth muscle cells isolated from the pedal retractor muscle of Mytilus edulis. J. Muscle Res. Cell Motil. 3, 2538.[Medline]
Jones, G. (1995). Molecular mechanisms of action of juvenile hormone. Annu. Rev. Ent. 40, 147169.[Medline]
Jones, G. and Sharp, P. A. (1997). Ultraspiracle: An invertebrate nuclear receptor for juvenile hormone. Proc. Natl. Acad. Sci. USA 94, 1349913503.
Jorgensen, W. K. and Rice, M. J. (1983a). Superextension and supercontraction in locust ovipositor muscles. J. Insect Physiol. 29, 437448.
Jorgensen, W. K. and Rice, M. J. (1983b). Morphology of a very extensible insect muscle. Tissue & Cell 15, 639644.[Medline]
Josephson, R. K. and Ellington, C. P. (1997). Power output from a flight muscle of the bumblebee Bombus terrestris. I. Some features of the dorso-ventral flight muscle. J. Exp. Biol. 200, 12151226.
Josephson, R. K. and Young, D. (1981). Synchronous and asynchronous muscles in cicadas. J. Exp. Biol. 91, 219237.
Kelly, D. B. (1986). Neuroeffectors for vocalisation in Xenopus laevis: hormonal regulation of sexual dimorphism. J. Neurobiol. 17, 231248.[Medline]
Koudele, K., Stout, J. F. and Reichert, D. (1987). Factors which influence female crickets (Acheta domesticus) phonotactic and sexual responsiveness to males. Physiol. Ent. 12, 6780.
Lakey, A., Labeit, S., Gautel, M., Ferguson, C., Barlow, D. P., Leonard, K. and Bullard, B. (1993). Kettin, a large modular protein in the Z-disc of insect muscles. EMBO J. 12, 28632871.[Abstract]
Manabe, T., Kawamura, Y., Higuchi, H., Kimura, S. and Maruyama, K. (1993). Connectin, giant elastic protein, in giant sarcomeres of crayfish claw muscle. J. Muscle Res. Cell Motil. 14, 654665.[Medline]
Maruyama, K. (1999). Comparative aspects of muscle elastic proteins. Rev. Physiol. Biochem. Pharmac. 138, 114.[Medline]
Medawar, P. (1953). Some immunological and endocrinological problems raised by the evolution of viviparity in vertebrates. Symp. Soc. Exp. Biol. Med. 7, 320338.
Miller, J. B. (1974). The lengthtension relationship of the dorsal longitudinal muscle of a leech. J. Exp. Biol. 62, 4353.[Abstract]
Morgan, D. M. L. (1999). Polyamines. Mol. Biotech. 11, 229250.[Medline]
Müller, A. R., Wolf, H., Galler, S. and Rathmayer, W. (1992). Correlation of electrophysiological, histochemical and mechanical properties in fibres of the coxa rotator muscle of the locust, Locusta migratoria. J. Comp. Physiol. B 162, 515.
Odhiambo, T. R. (1966). The metabolic effects of the corpus allatum hormone in the male desert locust. II. Spontaneous locomotor activity. J. Exp. Biol. 45, 5163.[Medline]
Orchard, I., Belanger, J. H. and Lange, A. (1989). Proctolin: a review with emphasis on insects. J. Neurobiol. 20, 470496.[Medline]
OShea, M., Adams, M. E., Bishop, C., Witten, J. and Worden, M. K. (1985). Model peptidergic systems at the insect neuromuscular junction. Peptides 6 (Suppl. 3), 417424.[Medline]
OShea, M. and Evans, P. D. (1979). Potentiation of neuromuscular transmission by an octopaminergic neurone in the locust. J. Exp. Biol. 79, 169190.
Pener, M. P. (1992). Environmental cues, endocrine factors and reproductive diapause in male insects. Chronobiol. Int. 9, 102113.[Medline]
Pener, M. P., Orshan, L. and De Wilde, J. (1978). Precocene II causes atrophy of corpora allata in Locusta migratoria. Nature 272, 350353.
Rand, M. N. and Breedlove, S. M. (1995). Androgen alters the dendritic arbors of SNB motoneurones by acting upon their target muscles. J. Neurosci. 15, 44084416.[Abstract]
Rembold, H. (1981). Modulation of JH III-titre during the gonotrophic cycle of Locusta migratoria, measured by gas chromatography-selected ion monitoring mass spectrometry. In Juvenile Hormone Biochemistry, Action, Agonism and Antagonism (ed G. E. Pratt and G. T. Brooks), pp. 1121. Amsterdam, New York, Oxford: Elsevier/North Holland.
Renucci, M., Cherkaoui, L., Rage, P., Augier, R. and Strambi, A. (1992). Juvenile hormone exerts a primer effect on oviposition behaviour in Acheta domesticus. In Insect Juvenile Hormone Research, Fundamental and Applied Approaches (ed. B. Mauchamp, F. Couillaud and J. C. Baehr), pp. 147163. Paris: INRA.
Riddiford, L. M. (1985). Hormone action at the cellular level. In Comprehensive Insect Physiology, Biochemistry and Pharmacology, vol. 8 (ed. G. A. Kerkut and L. I. Gilbert), pp. 3784. Frankfurt: Plenum Press.
Rose, U., Seebohm, G. and Hustert, R. (2000). The role of internal pressure and muscle activation during locust oviposition. J. Insect Physiol. 46, 6980.[Medline]
Sacca, L., Cittadini, A. and Fazio, S. (1994). Growth hormone and the heart. Endocrine Rev. 15, 555573.[Abstract]
Sehnal, F., Svacha, P. and Zrzavy, J. (1996). Evolution of insect metamorphosis. In Metamorphosis, Postembryonic Reprogramming of Gene Expression in Amphibian and Insect Cells (ed. L. I. Gilbert, J. R. Tata and B. G. Atkinson), pp. 358. New York, London: Academic Press.
Sevala, V. L. and Davey, K. G. (1989). Action of juvenile hormone on the follicle cells of Rhodinus prolixus: evidence for a novel regulatory mechanism involving protein kinase C. Experientia 45, 355356.
Snodgrass, R. E. (1935). The abdominal mechanisms of a grasshopper. Smithsonian Misc. Collns. 94, 187.
Stegwee, D., Kimmel, E. C., DeBoer, J. A. and Henstra, S. (1963). Hormonal control of reversible degeneration of flight muscle in the Colorado potato beetle, Leptinotarsa decemlineata Say (Coleoptera). J. Cell Biol. 19, 519527.
Stout, J., Hao, H., Atkins, G., Stiedl, O., Ramseier, J., Coburn, P., Hayes, V., Henley, J. and Kim, P. (1993). JHIII regulates phonotaxis in crickets by controlling expression of nicotinic receptors in auditory interneurons. In Insect Neurochemistry and Neurophysiology (ed. M. Loeb), pp. 343346. Boca Raton, FL: CRC Press.
Stout, J., Hayes, V., Zacharias, D., Henley, J., Stumpner, A., Hao, J. and Atkins, G. (1992). Juvenile hormone controls phonotactic responsiveness of female crickets by genetic regulation of the response properties of identified auditory interneurons. In Insect Juvenile Hormone Research: Fundamental and Applied Approaches (ed. B. Mauchamp, F. Couillaud and J. C. Baehr), pp. 265283. Paris: IRNA.
Strong, L. and Amerasinghe, F. P. (1977). Allatectomy and sexual receptivity in females of Schistocerca gregaria. J. Insect Physiol. 23, 131135.[Medline]
Tanaka, S. (1994). Endocrine control of ovarian development and flight muscle histolysis in a wing dimorphic cricket, Modicogryllus confirmatus. J. Insect Physiol. 40, 483490.
Tawfik, A. I., Osir, E. O., Hassanall, A. and Ismail, S. H. (1997). Effects of juvenile hormone treatment on phase changes and pheromone production in the desert locust, Schistocerca gregaria (Forskål) (Orthoptera: Acrididae). J. Insect Physiol. 43, 11771182.[Medline]
Tawfik, A. I., Treiblmayr, K., Hassanali, A. and Osir, E. O. (2000). Time-course of haemolymph juvenile hormone titres in solitarius and gregarius adult of Schistocerca gregaria and their relation to pheromone emission, CA volumetric changes and oocyte growth. J. Insect Physiol. 46, 11431150.[Medline]
Tobe, S. S. and Bendena, W. G. (1999). The regulation of juvenile hormone production in arthropods: Functional and evolutionary perspectives. In Annals of the New York Academy of Sciences, Neuropeptides: Structure and Function in Biology and Behaviour (ed. C. A. Sandman, F. L. Strand, B. Beckwith, B. M. Chronwall, F. W. Flynn and R. J. Nachman), pp. 300310.
Triggle, D. J. (1990). Calcium channel and calcium channel antagonists. Can. J. Physiol. Pharmac. 68, 14741481.[Medline]
Usherwood, P. N. R. and Cull-Candy, S. G. (1975). Pharmacology of somatic nervemuscle synapses. In Insect Muscle (ed. P. N. R. Usherwood), pp. 207280. London: Academic Press.
Venable, J. H. (1966). Morphology of the cells of normal, testosterone-deprived and testosterone-stimulated levator ani muscles. Am. J. Anat. 119, 271302.[Medline]
Vincent, J. F. V. (1975). How does the female locust dig her oviposition hole? J. Ent. 50, 175181.
Walikonis, R., Schoun, D., Zacharias, D., Henley, J., Coburn, P. and Stout, F. (1991). Attractiveness of the male Acheta domesticus calling song to females. J. Comp. Physiol. A 169, 751764.[Medline]
Washio, H. (1972). The ionic requirement for the initiation of action potentials in insect muscle fibres. J. Gen. Physiol. 59, 121134.
Weeks, J. C. and Truman, J. W. (1986). Steroid control of neuron and muscle development during the metamorphosis of an insect. J. Neurobiol. 17, 249267.[Medline]
Whiteley, N. M., Taylor, E. W. and El Haj, A. J. (1992). Actin gene expression during muscle growth in Carcinus maenas. J. Exp. Biol. 167, 277284.
Wigglesworth, V. B. (1934). The physiology of ecdysis in Rhodinus prolixus. II. Factors controlling moulting and metamorphosis. Q. J. Microsc. Sci. 77, 191222.
Wigglesworth, V. B. (1936). The function of the corpus allatum in the growth and reproduction of Rhodinus prolixus (Hemiptera). Q. J. Microsc. Sci. 79, 91121.
Worden, M. K., Witten, J. L. and OShea, M. (1985). Proctolin is a co-transmitter for SETi motoneuron. Soc. Neurosci. Abstr. 11, 327.
Wyatt, G. R. (1997). Juvenile hormone in insect reproduction a paradox? Eur. J. Ent. 94, 323333.
Wyatt, G. R. and Davey, K. G. (1996). Cellular and molecular actions of juvenile hormone. II. Roles of juvenile hormone in adult insects. Adv. Insect Physiol. 26, 1155.
Yamamoto, D., Fukami, J. and Washio, H. (1978). Ca-electrogenesis in mealworm muscle: A voltage clamp study. Experientia 34, 16031604.
Yamamoto, K., Chadarevian, A. and Pellegrini, M. (1988). Juvenile hormone action mediated in male accessory glands of Drosophila by calcium and kinase C. Science 239, 916919.[Medline]