EXCITATORY AND INHIBITORY ROLES OF CENTRAL GANGLIA IN INITIATION

The initiation of behaviour at appropriate times dependson the selective activation of certain neural networks andinhibition of others. The unique properties of these circuits,including their responsiveness to excitatory or inhibitoryinfluences, are determined by precise patterns of geneexpression. The relationships between gene expression andbehaviour can be conveniently examined in instances wherethey play critical roles in developmental and reproductivefunctions. For example, the initiation of insect behaviours suchas wandering and ecdysis is preceded by intense bouts ofgene expression under the control of steroid hormones(ecdysteroids), resulting in the production of specific receptors,the synthesis of signalling molecules and the precise timing oftheir release (Thummel, 1995; Hewes and Truman, 1994;Zitnan et al., 1999).

Behaviours associated with shedding of the old cuticle arecrucial for the successful development of the tobacco hornwormManduca sexta. This behavioural sequence, consisting of pre-ecdysis I, pre-ecdysis II and ecdysis, is initiated by the directaction of peptide hormones from endocrine Inka cells on thecentral nervous system (CNS) (Zitnan et al., 1996). During the1–2 days preceding larval ecdysis of M. sexta, a pulse ofecdysteroids controls the expression of specific genes both in

Inka cells and in the CNS necessary for the subsequent initiationof these behaviours (Zitnan et al., 1999). Rising ecdysteroidlevels induce ecdysis-triggering hormone (ETH) geneexpression in Inka cells, while a decline in ecdysteroid levelsregulates the late transcriptional activity required for the releaseof pre-ecdysis-triggering hormone (PETH) and ETH derivedfrom this gene (Zitnan et al., 1999). Meanwhile, elevatedecdysteroid levels induce CNS sensitivity to these peptidehormones (Zitnan et al., 1999) and the fall in ecdysteroid levelsinduces increased excitability of the brain VM neurons followedby the release of eclosion hormone (EH) (Hewes and Truman,1994). Blood-borne EH causes the release of PETH and ETHfrom Inka cells (Ewer et al., 1997; Kingan et al., 1997), whichin turn act back on the CNS (Zitnan et al., 1999). Low levels ofPETH and ETH subsequently initiate pre-ecdysis I and II.Increased ETH concentrations in the blood lead to activation ofthe ecdysis neuronal network (Zitnan et al., 1996, 1999), butecdysis behaviour is initiated after central release of crustaceancardioactive peptide (CCAP) from this network (Gammie andTruman, 1997). Each motor unit of the ecdysis behaviouralsequence is driven by a specific central pattern generator (Weeksand Truman, 1984a; Novicki and Weeks, 1995). It is thereforeimportant in the context of the needs of the animal that selective

1329The Journal of Experimental Biology 203, 1329–1340 (2000)Printed in Great Britain © The Company of Biologists Limited 2000JEB2592

Insects shed their old cuticle by performing the ecdysisbehavioural sequence. To activate each subunit of this setof programmed behaviours in Manduca sexta, specificcentral ganglia are targeted by pre-ecdysis-triggering(PETH) and ecdysis-triggering (ETH) hormones secretedfrom Inka cells. PETH and ETH act on each abdominalganglion to initiate, within a few minutes, pre-ecdysis I andII, respectively. Shortly thereafter, ETH targets thetritocerebrum and suboesophageal ganglion to activate theecdysis neural network in abdominal ganglia through theelevation of cyclic GMP (cGMP) levels. However, the onsetof ecdysis behaviour is delayed by inhibitory factor(s) fromthe cephalic and thoracic ganglia. The switch from pre-ecdysis to ecdysis is controlled by an independent clock ineach abdominal ganglion and is considerably accelerated

after removal of the head and thorax. Eclosion hormone(EH) appears to be one of the central signals inducingelevation of cGMP levels and ecdysis, but these actions arequite variable and usually restricted to anterior ganglia.EH treatment of desheathed ganglia also elicits strongproduction of cGMP in intact ganglia, suggesting that thisinduction occurs via the release of additional downstreamfactors. Our data suggest that the initiation of pre-ecdysisand the transition to ecdysis are regulated by stimulatoryand inhibitory factors released within the central nervoussystem after the initial actions of PETH and ETH.

Key words: ecdysis, insect, tobacco hornworm, Manduca sexta,eclosion hormone, ecdysis-triggering hormone, pre-ecdysis-triggering hormone, cyclic GMP.

Summary

Introduction

EXCITATORY AND INHIBITORY ROLES OF CENTRAL GANGLIA IN INITIATION OFTHE INSECT ECDYSIS BEHAVIOURAL SEQUENCE

DUSAN ZITNAN1,2,* AND MICHAEL E. ADAMS2

1Institute of Zoology, Slovak Academy of Sciences, Dúbravská Cesta 9, 84206 Bratislava, Slovakia and 2Departmentsof Entomology and Neuroscience, 5419 Boyce Hall, University of California, Riverside, CA 92521, USA

*e-mail: [email protected]

Accepted 7 February; published on WWW 23 March 2000

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activation and inhibition of these different motor units isprovided at the appropriate time and in the appropriate order.

In this paper, we identify specific target ganglia for PETHand ETH, and provide new insights into the mechanismsunderlying the switch from pre-ecdysis to ecdysis. We havefound that the ecdysis neuronal network is activated by ETHearly in pre-ecdysis, but the onset of ecdysis is delayed byinhibition from cephalic and thoracic ganglia. The eventualswitch to ecdysis is controlled by an independent clock in eachabdominal ganglion, which relieves the inhibition and initiatesecdysis. Our data show that PETH and ETH induce a series ofdownstream events within the CNS, leading to the subsequentactivation of distinct central pattern generators and theperformance of the ecdysis behavioural sequence at a specifictime. This system provides an ideal opportunity to studymechanisms underlying a centrally patterned series ofmovements, since each behaviour can be hormonally inducedand analyzed in animals in vivo and in the isolated CNS invitro.

Materials and methodsBioassays and surgical procedures

In vivo and in vitro bioassays using intact, ablated ortransected central nervous sytems were carried out todetermine the target sites for PETH and ETH in pharate fifth-instar larvae of Manduca sexta (L.) 6–8 h before ecdysis. Priorto each surgical procedure, larvae were anaesthetized withCO2. For the CNS transections, a small incision was madethrough the ventral cuticle between abdominal segments 4 and5 or 6 and 7, and the connectives between the respectiveganglia were severed using forceps. For the ablation orremoval of the brain, a small incision was made using sharpethanol-sterilized forceps into the frontal part of the head, andthe entire brain or proto- and deutocerebrum was cut off usingmicroscissors. Selected groups of these larvae were injectedwith 50 pmol of one peptide, or injected with one peptidefollowed by treatment with the other peptide. Different phasesof the induced ecdysis behavioural sequence were observedunder a stereomicroscope and recorded on videotape. To studythe inhibitory role of the cephalic and thoracic segments indelaying the onset of ecdysis, larvae were injected with50 pmol of ETH and ligated behind the head or the first twoabdominal segments. The anterior part was cut off usingscissors 10–30 min into pre-ecdysis.

For in vitro studies, the isolated CNS was transectedbetween abdominal ganglia 1 and 2 (AG1–2) or eachabdominal ganglion was individually isolated before treatmentwith PETH or ETH (100 nmol l−1). Alternatively, nerve cordswere treated with these peptides and transected 10–30 min afterthe initiation of pre-ecdysis. Burst patterns triggered by thesepeptides were detected by extracellular recordings with suctionelectrodes as described by Zitnan et al. (1996, 1999).

To compare different functions of peptides released by Inkacells or the CNS during ecdysis, the desheathed entire CNS orisolated chain of AG1–8 was treated with 100 nmol l−1 of ETH,

EH or CCAP in vitro, and induced burst patterns were recordedfrom dorsal nerves of the abdominal ganglia using suctionelectrodes. In some cases, only the anterior ganglia(suboesophageal ganglion, SG, and thoracic ganglia 1–3,TG1–3) or posterior CNS (AG1–8) were desheathed on thedorsal side and the remaining ganglia were left intact. Thesenerve cords were then treated with EH (100 nmol l−1) todetermine its ability to induce elevations in cGMP levelsand/or ecdysis bursts in intact non-desheathed ganglia.

Immunohistochemistry

For immunohistochemical detection of cGMP in the CNS,we followed modified procedures described previously (Eweret al., 1994; Gammie and Truman, 1999). Using this simplifiedprocedure, no methanol or collagenase treatment to allowantibodies to penetrate the CNS was necessary. Briefly, theCNS was fixed overnight in 4 % paraformaldehyde, washed inphosphate-buffered saline containing 0.5 % Triton X-100(PBST) and incubated for 2 days with sheep or rabbitantiserum to cGMP (dilution 1:10 000 and 1:2000,respectively) obtained from Drs J. De Vente and H. M. W.Steinbush. Specific binding of these antisera was detected byovernight incubation with horseradish peroxidase (HRP)-labelled donkey anti-sheep IgG (Jackson ImmunoresearchLabs, West Grove, PA, USA) or HRP-labelled goat anti-rabbitIgG (American Qualex, San Clemente, CA, USA). HRP wasstained with diaminobenzidine (Sigma, St Louis, MO, USA)and hydrogen peroxide, and nerve cords were mounted inglycerol.

Values are presented as means ± S.D.

ResultsPETH and ETH target each abdominal ganglion to activate

pre-ecdysis I and II

PETH and ETH induce the two distinct behavioural parts ofpre-ecdysis. PETH elicits pre-ecdysis I, which is characterizedby strong dorsoventral contractions, while ETH induces pre-ecdysis II, which consists of posterioventral and prolegcontractions (Zitnan et al., 1999).

To identify the target ganglia for each peptide, we observedthe effects of nerve cord transection on pre-ecdysis I and IIcontractions in larvae injected with PETH or ETH. Followingsurgical transection of the nerve cord between AG4 and AG5(N=10) or AG6 and AG7 (N=10), animals were injected withPETH (50 pmol). Within 4–8 min, the peptide induced strongand regular dorsoventral contractions typical of pre-ecdysis Ionly in segments posterior to the cut (Fig. 1A). Abdominalsegments anterior to the transection displayed strong, butprolonged and irregular, dorsoventral contractions (3–25 s ofcontraction and 10–30 s of relaxation). This behaviour endedwithin 35–50 min, at which time abdominal segments anteriorto nerve transection remained in permanent dorsoventralcontraction, while the posterior abdominal segmentsmaintaining a connection to the terminal abdominal ganglion(TAG, fused AG7,8) were relaxed. These experiments showed

D. ZITNAN AND M. E. ADAMS

1331Excitation and inhibition control ecdysis onset

that PETH targets all the abdominal ganglia, but thatcoordinated dorsoventral movements depend on pacemakeractivity in the TAG.

Subsequent injection of the same larvae with ETH (50 pmol)induced normal pre-ecdysis II behaviour within 8–12 min. Thisconsisted of strong posterioventral and proleg contractionsboth anterior and posterior to the transections (Fig. 1B; N=20).All animals (N=20), switched to typical ecdysis peristalticcontractions 25–36 min later, but only in segments anterior tothe cut (Fig. 1C). The transition to ecdysis behaviour thereforerequires a connection between the abdominal ganglia and theanterior CNS.

To identify specific ganglia targeted by PETH, we examinedthe effects of nerve cord transection on bursting in the dorsalnerves of the abdominal ganglia in vitro. In the isolated intactCNS, application of PETH (100 nmol l−1) induced rhythmic,synchronous bursting motor patterns typical of pre-ecdysis Iwithin 5–8 min (Fig. 2A; N=10). Transection of the abdominalconnectives led to irregular, long-duration bursting of isolatedganglia, whereas typical pre-ecdysis I bursts persisted in theTAG (N=8). Similarly, in nerve cords transected prior topeptide treatment, PETH always induced normal pre-ecdysis Ibursting in the TAG, while individually isolated AG3–6

showed only irregular, prolonged bursting (Fig. 2B, N=10).This pattern was similar to the long dorsoventral contractionsof transected larvae observed in vivo (Fig. 1A). Theseexperiments showed that all the abdominal ganglia respond toPETH application, but that interneurons projecting from theTAG determine the patterns of pre-ecdysis I bursting in theentire abdominal chain. The pacemaker interneurons providingthis input have been identified as IN-402 (Novicki and Weeks,1995).

Similar experiments were performed to ascertain whichganglia are targeted by ETH for the initiation of pre-ecdysis II.Exposure of isolated AG1–6 to ETH (100 nmol l−1) led tostrong bursting in the ventral nerves (Fig. 2C; N=12)corresponding to posterioventral and proleg contractions.However, unlike the experiments described above with PETH,typical pre-ecdysis II bursts were induced in individuallyisolated abdominal ganglia (N=8) after the transection of allconnectives (Fig. 2D). Therefore, each abdominal ganglionappears to contain the entire neural circuitry necessary for pre-ecdysis II and is not dependent on pacemaker interneuronsprojecting from distant ganglia. These results also showed thatthe target sites for ETH are located in each abdominalganglion, which is consistent with in vivo experimentsdescribed above.

Activation of the ecdysis circuitry requires cephalic ganglia

While low concentrations of Inka cell peptides trigger pre-ecdysis I and II, higher levels of ETH are necessary to triggerthe switch from pre-ecdysis to ecdysis (Zitnan et al., 1999). Aprevious report showed that the brain plays an essential role inthe transition to ecdysis (Novicki and Weeks, 1996). Byperforming experiments in vivo and in vitro, we have foundthat both the brain and the SG are important in ETH-inducedecdysis.

We examined the roles of the cephalic ganglia in theactivation of ecdysis by removing the brain of the larvae or bysevering the connectives behind the SG prior to injection of thepeptide. Most debrained animals (17 out of 20; 85 %) failed tomake the transition to ecdysis following ETH injection(50 pmol), and transection of the connectives posterior to theSG prevented ecdysis in all larvae (N=10). To determine theareas of the brain involved in the activation of ecdysis, partialablations were performed. When the protocerebrum anddeutocerebrum were ablated, leaving one or both tritocerebrallobes intact, most animals (10 out of 14; 71 %) initiated ecdysisbehaviour at the expected time. These experiments show thatthe tritocerebrum and SG are critical in facilitating thetransition to ecdysis in vivo.

We also examined the involvement of the anterior gangliain the transition to ecdysis in isolated nerve cords. In theseexperiments, the connectives were transected posterior to thebrain, SG or TG3 prior to exposure to ETH (100 nmol l−1). In25 out of 26 cases, brainless nerve cords switched from pre-ecdysis to ecdysis after the normal interval 41.64±4.2 min (Fig.3A). However, nerve cords transected posterior to the SGshowed ecdysis in only two out of eight experiments (in 45 and

A

B

C

PETH; pre-ecdysis I

ETH; pre-ecdysis II

ETH; ecdysis

Fig. 1. Effects of transection of the abdominal connectives (betweenabdominal ganglia 4 and 5) on pre-ecdysis I, pre-ecdysis II andecdysis behaviours. (A) Injection of 50 pmol of pre-ecdysis-triggering hormone (PETH) into a pharate larva induced normalrhythmic pre-ecdysis I contractions of the dorsal and ventral (D-V)muscles (dark areas) posterior to the cut (indicated by a scissorsymbol), while only prolonged and irregular dorsoventralcontractions (dotted areas) occurred in the anterior abdomen. (B) Subsequent injection of 50 pmol of ecdysis-triggering hormone(ETH) induced characteristic pre-ecdysis II contractions of the entireventral abdomen (dark areas). Approximately 30 min later, theanimal switched to ecdysis peristaltic contractions (C), whichoccurred only anterior to the cut (dark areas).

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49 min). Removal of TG1–3, leaving intact a chain of AG1–8,prevented the transition to ecdysis in all experiments (N=8).These results suggest that the action of ETH on the SG, and insome cases on TG1–3, is sufficient to activate the ecdysiscircuitry in vitro. Experiments with individually isolatedabdominal ganglia showed that motor units driving ecdysis arepresent in each abdominal ganglion (Fig. 3B; see below).

Timing of ETH-induced elevation of cGMP levels

The activation of the ecdysis circuitry is associated with theelevation of cGMP levels in a network of 27/704 neurons(Ewer et al., 1994; Ewer and Truman, 1997). Since PETH andETH activate different phases of the ecdysis behaviouralsequence (Zitnan et al., 1996, 1999), we examined the timingof elevation of cGMP levels in the CNS under naturalconditions and after exposure to these peptides in vivo and invitro.

Under natural conditions, we observed cGMPimmunoreactivity in CNS ganglia 6–8 h before ecdysis, wellbefore the release of peptides from the Inka cell (N=17).Notably, cGMP levels were elevated in the SG, TG1–3 andAG8, but not in AG1–7 (Fig. 4A). During the first 30 min ofpre-ecdysis, cGMP staining in neurons and axons of the SGand TG1–3 clearly increased (N=9). At approximately 30 mininto pre-ecdysis, cGMP staining appeared in the 27/704neurons of AG1–7 (N=10). At approximately 45 min into pre-ecdysis and lasting until ecdysis onset, strong cGMPimmunoreactivity was observed in the entire 27/704 networkand weaker staining was detected in the abdominalneurosecretory cells L2,3 that produce diuretic peptides andkinins (N=16; Chen et al., 1994).

Pre-ecdysis I induced by PETH in pharate larvae (50 pmol;N=8) or in isolated nerve cords (100 nmol l−1; N=7) was notassociated with an elevation in cGMP level. However,


B PETH; pre-ecdysis I of isolated ganglia

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A PETH; pre-ecdysis I of intact CNS

D ETH; pre-ecdysis II of isolated ganglia

C ETH; pre-ecdysis II of intact CNS

Fig. 2. Effects of the isolation ofindividual ganglia on pre-ecdysisI and pre-ecdysis II burstsinduced by pre-ecdysis-triggeringhormone (PETH) and ecdysis-triggering hormone (ETH) invitro. (A) In the intact nerve cord,PETH (100 nmol l−1) inducednormal synchronous pre-ecdysis Ibursts in the dorsal nerves ofabdominal ganglia (AG) 5–7. (B) Singly isolated AG5 andAG6 showed only prolongednon-synchronous bursts, butthe normal pre-ecdysis I burstpattern was recorded in the AG7 neuromere of theterminal abdominal ganglion. (C) In the intact nerve cord, ETH (100 nmol l−1) triggeredsynchronous pre-ecdysis II burstsin the ventral nerves of AG4–6.(D) Very similar, but lesssynchronized, bursts wererecorded in singly isolatedganglia. CNS, central nervoussystem; D, dorsal; V, ventral.


application of 50 pmol of ETH in vivo (N=8) or of 100 nmol l−1

ETH in vitro (N=7) induced strong cGMP immunoreactivityin all the neurons and axons described above during thefirst 10–15 min of pre-ecdysis (Fig. 4B). This strongimmunoreactivity persisted until the onset of ecdysis (N=10).Similar staining occurred after ETH treatment of debrainednerve cords (N=10; Fig. 4C), but not in isolated abdomens(N=8) or in the isolated AG1–8 in vitro (N=7). These resultsshow that the action of ETH on the SG is sufficient to inducea complete elevation of cGMP levels followed by ecdysis.

We also tested the cGMP response to ETH (50 pmol)injection in larvae that had had their abdomen ligated 10 mininto pre-ecdysis. The CNS of these ligated abdomens wasdissected out and processed for cGMP immunoreactivity justafter the onset of ecdysis, which was considerably accelerated,as described below. In six out of 11 experiments, all theabdominal ganglia showed strong elevation of cGMP levels in27/704 neurons and axons, but the remaining five abdominalnerve cords showed weaker staining confined to the cellbodies. These results indicate that the weaker elevation ofcGMP levels in the abdominal ganglia may be sufficient forthe activation and onset of ecdysis.

Inhibitory factors from the cephalic and thoracic gangliadelay the onset of ecdysis

Injection of pharate larvae with ETH (50 pmol) leads topre-ecdysis within 5–8 min, and ecdysis behaviour followsafter a very consistent delay (41±2.6 min) (N=15; Fig. 5;Table 1). Despite this delay, we found that ETH actuallyactivates the ecdysis neuronal network during early pre-ecdysis, but that the onset of edysis is delayed by inhibitionfrom the cephalic and thoracic segments. Removal ofinhibition by ablation of these segments greatly reduces thedelay in the onset of edysis.

We demonstrated this by injecting larvae with ETH(50 pmol), then removing the head by neck ligation (appliedbetween thoracic segments 1 and 2) at various intervals

following the initiation of pre-ecdysis. When neck ligation wasapplied 30 min into pre-ecdysis, a premature switch to ecdysisoccurred within 10–20 s in all larvae (N=10; Table 1). Thismanipulation thus reduced the delay to the onset of edysis byapproximately 10 min, suggesting that inhibition emanates atleast partly from the cephalic ganglia. If we applied neckligation earlier, at 20–25 min into pre-ecdysis, the larvaeimmediately switched to ecdysis for approximately 1–4 min,but then reverted to weak pre-ecdysis-like movements in 11out of 15 experiments. The remaining four larvae in thisexperimental group as well as eight additional animals neck-ligated 15 min into pre-ecdysis failed to continue in normalpre-ecdysis and did not initiate ecdysis (Table 1). However, ifa second ligation (applied between abdominal segments 1 and2) followed 5–15 min later, immediate and strong ecdysismovements occurred in the isolated abdomens. These resultssuggest that inhibition from the thoracic ganglia alsocontributes to the delay in the onset of ecdysis.

The role of thoracic inhibition was further examined byperforming ligations between abdominal segments 1 and 2 atvarious times after the initiation of pre-ecdysis. Abdominalligation 20–30 min into pre-ecdysis (N=11) caused a prematureswitch to ecdysis behaviour within 1–2 min (Table 1). Earlierabdominal ligations also caused premature ecdysis, but withsome delay. Animals ligated 15 min into pre-ecdysis switchedto ecdysis within 1–9 min (N=12), while ligation at 10 min intopre-ecdysis led to ecdysis within 5–14 min (N=19). Theseanimals therefore performed pre-ecdysis for only 15–24 min(18±4.1 min) before the initiation of ecdysis contractions,which is much shorter than the approximately 40 min intervalobserved consistently in intact larvae injected with ETH(Fig. 5). Animals ligated 5 min into pre-ecdysis (N=10)invariably failed to perform ecdysis (Table 1). From theseresults, we conclude that injected ETH activates the ecdysisneuronal network in the abdominal ganglia during the first10–15 min of pre-ecdysis, which correlates with theappearance of cGMP immunoreactivity in AG1–7 (Fig. 4B).

B ETH; ecdysis of isolated ganglia

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A ETH; ecdysis of debrained CNS

Fig. 3. Activation of ecdysis by ecdysis-triggeringhormone (ETH) in the debrained central nervoussystem (CNS) and singly isolated ganglia. (A) ETH (100 nmol l−1) treatment of the debrainednerve cord triggered normal ecdysis bursts at theexpected time (approximately 42 min after theinitiation of pre-ecdysis). (B) Isolation ofindividual abdominal ganglia (AG4–6) 10 mininto pre-ecdysis accelerated the onset of edysis.Ecdysis occurred in each isolated ganglionapproximately 35 min into pre-ecdysis. Note thatthe pattern of ecdysis bursts in the dorsal nerveswas altered by transection. D, dorsal; V, ventral.

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However, inhibition from both cephalic and thoracic segmentsdelays the onset of edysis.

We also determined the delay from ecdysis activation to itsonset during natural behaviour. Under our laboratoryconditions, larvae initiated natural ecdysis 60–70 min after theinitiation of pre-ecdysis (N=9; Table 1; Fig. 5). If abdominalligation was applied 35–40 min into pre-ecdysis (N=8), aswitch to ecdysis occurred within 1–16 min (Table 1; Fig. 5).Thus, these larvae switched to ecdysis 35–53 min into pre-ecdysis (40±7.5 min), which was approximately 15–30 min

earlier than in intact control larvae. Ecdysis in all ligated larvaealways started with complete proleg retraction and anteriorperistaltic movements of abdominal segment 3, followed bystrong ventral contractions and peristaltic movements ofsegments 2 and 1. Posterior segments (4–8) were subsequentlyrecruited in this behaviour. Larvae ligated 15 and 30 min intopre-ecdysis failed to show ecdysis (N=9).


Fig. 5. Effects of abdominal ligation on the onset of natural andecdysis-triggering hormone (ETH)-induced ecdysis. Under naturalconditions, larvae show pre-ecdysis I and II behaviours forapproximately 65 min and then switch to ecdysis, which lasts forapproximately 10 min. Ecdysis contractions stop when the cuticle hasbeen completely shed. Abdominal ligation (arrow) 35 min into pre-ecdysis triggers ecdysis 1–16 min later. Larvae injected with ETHshow pre-ecdysis for approximately 40 min and then switch toecdysis. Abdominal ligation (arrow) 10 min into pre-ecdysis greatlyaccelerates the onset of edysis, which occurs 5–14 min later. Sinceligated and ETH-injected larvae are not able to shed their skin,ecdysis contractions could last for up to 1 h (stippled red bar). Notethat ETH injection induces both pre-ecdysis I and II because theaction of ETH on the central nervous system causes the release ofeclosion hormone, which induces the secretion of endogenous pre-ecdysis-triggering hormone (PETH) and ETH from Inka cells (fordetails, see Zitnan et al., 1999).

ETH

ETH

Natural

Natural

0

Pre-ecdysis I Pre-ecdysis IILigation

Ligation

Ecdysis

10 20 30 40 50 60 70

Time (min)

Table 1. Time of the onset of natural or ETH-induced ecdysisin larvae ligated at different time into pre-ecdysis

Ecdysis onsetLigation time (min) N

ETH Control (no ligation) 37–46 15ETH Neck ligation (30 min) 30 10 ETH Neck ligation (20–25 min) Weak ecdysis 11ETH Neck ligation (15–20 min) No ecdysis 12ETH A1–2 ligation (20–30 min) 20–30 11ETH A1–2 ligation (10–15 min) 15–24 31ETH A1–2 ligation (5 min) No ecdysis 10Natural Control (no ligation) 60–70 9Natural A1–2 ligation (35–40 min) 35–53 8Natural A1–2 ligation (30 min) No ecdysis 9

ETH, ecdysis-triggering hormone; A1–2, ligation betweenabdominal segments 1 and 2.

Ligation time and ecdysis onset are referenced to the initiation ofpre-ecdysis.

Fig. 4. Ecdysis-triggering hormone (ETH)-induced cGMPimmunoreactivity in the central nervous system. (A) Untreated control larvae 6 h before ecdysis show weak cGMPimmunoreactivity in only two neurons in the suboesophagealganglion (SG), in thoracic ganglia (TG) 1–3 and in abdominalganglion (AG) 8 of the terminal abdominal ganglion (TAG). (B) ETH (50 pmol) injection induced, in 10–15 min, strong elevationin cGMP levels in the 27/704 network of the entire ventral nervecord. (C) A similar strong cGMP response was induced by ETH (100 nmol l−1) in the debrained central nervous system. Note that, inaddition to cells 27/704, these nerve cords showed cGMP staining inthe L2,3 neurons of most abdominal ganglia (arrows). Scale bar,150 µm.


We examined the role of inhibition in delaying ETH-induced ecdysis using the isolated CNS. Similar to in vivoexperiments, the isolated entire CNS exposed to ETH(100–150 nmol l−1) switches to ecdysis 38–47 min after theinitiation of pre-ecdysis (42.3±2.9 min; N=17). This delaywas reduced (35.2±2.5 min) upon nerve cord transection10–20 min into pre-ecdysis, applied either between AG1 andAG2 (N=18) or between each abdominal ganglion (N=13;Fig. 3B). If transections were performed 5 min into pre-ecdysis, all preparations failed to exhibit ecdysis (N=8). Asin ligated larvae, transected abdominal nerve cords andindividually isolated abdominal ganglia initiated ecdysisbursts in anterior ganglia (AG3,4) and within several minutesthese bursts subsequently occurred in more posterior ganglia(AG5–8).

These experiments suggest that ETH activates anindependent clock in each abdominal ganglion responsible forthe initiation of ecdysis at the appropriate time even if theseganglia are individually isolated as early as 10 min into pre-ecdysis (Fig. 3B). Therefore, the neurons controlling ecdysisare activated by ETH approximately 10 min after the onset ofpre-ecdysis, and the CNS is capable of the switch to ecdysis5 min later. However, this transition is delayed by inhibitoryinput from the cephalic and thoracic ganglia. The effect ofremoving these ganglia in vitro was less pronounced than invivo. This difference could be due to the lack of sensory inputto the isolated CNS.

Comparison of the actions of ETH, EH and CCAP on theisolated CNS

EH and CCAP have been implicated as downstream signalsreleased upon ETH action on the CNS (Gammie and Truman,1997, 1999). To clarify the respective roles of ETH, EH andCCAP in the regulation of ecdysis, we used the sameconcentration of these peptides (100 nmol l−1) to compare theireffects on the activation and onset of ecdysis in the desheathedentire CNS and AG1–8 in vitro. Since manual desheathing ofthe entire CNS caused spontaneous pre-ecdysis and ecdysisbursts in three cases, the activity of all nerve cords wasobserved for the first 10–15 min before application of peptide.

Nerve cords displaying spontaneous pre-ecdysis bursting werediscarded.

ETH invariably induced strong pre-ecdysis and ecdysisbursts in all desheathed CNS preparations (N=10). Thelatency to onset of these burst patterns was shorter thanobserved in the intact non-desheathed CNS; pre-ecdysisoccurred in 1–6 min (4.2±1.8 min) followed by ecdysis burstsin 28–36 min (32.4±2.8 min). Characteristic ecdysis burstswere recorded in all ganglia tested (AG3–7), and strongelevation of cGMP levels was observed in the entire networkof neurons and axons described above. However, in nervecords consisting of only AG1–8, ETH failed to induce ecdysisbursts and elevation of cGMP levels during 60–90 min ofrecording (N=7).

Exposure of the entire desheathed CNS to EH induced anelevation in cGMP levels and ecdysis bursts, but theseresponses were quite variable and usually confined to theanterior ganglia (Figs 6, 7). Ecdysis bursts occurred in eightout of 11 experiments within 20–52 min of EH application(30.6±11.9 min). However, only three nerve cords showedthese bursts in all abdominal ganglia (Fig. 6A), while fiveCNS preparations produced ecdysis only in AG3 or AG3,4(Fig. 6B) and three CNS preparations completely failed toinitiate ecdysis. In the three nerve cords showing ecdysisbursting in all abdominal ganglia, cGMP staining was strongin all neurons 27/704 and their axons (Fig. 7A). In othernerve cords, strong elevation in cGMP levels was observedin the SG and TG1–3, but weak or very weak responses weredetected in the abdominal ganglia. Thus, despite exposure ofthe entire desheathed CNS to EH, strong cGMP staining wasusually restricted to the anterior CNS.

Experiments with non-desheathed nerve cords confirmedprevious observations (Gammie and Truman, 1999) that EHdoes not penetrate the neural sheath and fails to induce anyactivity in the intact CNS within 60 min of incubation (N=8).Therefore, we desheathed only certain anterior or posteriorganglia to determine whether the action of EH on desheathedganglia induced an elevation in cGMP levels and ecdysis inintact ganglia. When desheathing was performed only inAG1–8, ecdysis occurred in six out of 11 experiments, but the

B

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AG7D

A

Fig. 6. Variable response of the isolated desheathedcentral nervous system to eclosion hormone (EH). (A) EH (100 nmol l−1) treatment of the desheathedentire nerve cord induced, in approximately 30 min,ecdysis bursts in all ganglia. (B) A chain of abdominalganglia showed ecdysis bursts only in abdominalganglion 3 (AG3), while more posterior ganglia (AG5and AG7) continued to display pre-ecdysis-like bursts.Most abdominal ganglia showed this bursting patternand failed to switch to ecdysis bursts. D, dorsal.

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onset of the behaviour was extremely variable (range22–95 min; 53.6±25.5 min). Similar to experiments describedabove, only three nerve cords showed strong ecdysis bursts andcGMP staining in all abdominal ganglia. In the remaining threenerve cords, ecdysis was recorded only in AG1–4, and fiveCNS preparations failed to initiate ecdysis during 60–90 minof recording. These nerve cords showed only weak to veryweak cGMP immunoreactivity in the abdominal ganglia.

Surprisingly, all SG and TG1–3 displayed strong cGMPstaining even if their neural sheath had not been removed(Fig. 7B).

If the brain, SG and TG1–3 were desheathed leavingAG1–8 intact (N=8), EH failed to activate ecdysis burstsduring 60–90 min, but strong cGMP staining was observed in27/704 neurons of all non-desheathed abdominal ganglia(Fig. 7B). Extended incubation of one of these nerve cordsfor approximately 2 h eventually resulted in ecdysis bursts inAG3,4. These experiments showed that the action of EH ondesheathed ganglia may result in the activation of ecdysis inintact ganglia through an elevation in cGMP levels, but implythat this activation is indirect and probably requires somedownstream signalling events.

Treatment of the desheathed abdominal nerve cord (AG1–8)with EH resulted in weak synchronous bursts similar to pre-ecdysis I within 1–5 min, followed by ecdysis bursts 10–73 minlater (39.2±18.1 min) in 15 out of 18 experiments. However,only five of these nerve cords showed ecdysis bursts in allabdominal ganglia; in 10 nerve cords, ecdysis was recordedonly in AG3 or AG3,4, while posterior ganglia showed pre-ecdysis-like synchronous bursts (Fig. 6B). In three cases, noecdysis activity was observed after 80 min of exposure. Onlyfive nerve cords showed strong cGMP staining in all abdominalganglia; in the remaining experiments, the cGMP response wasstronger in anterior ganglia (mostly in AG1,2) and weak orundetectable in the more posterior abdominal ganglia(Fig. 7C).

An earlier study suggested that central release of CCAPfrom abdominal neurons 27/704 provides an immediatesignal for the initiation and performance of ecdysis behaviour(Gammie and Truman, 1997). However, we observed thatCCAP treatment of the entire desheathed CNS did not inducetypical ecdysis bursts. Instead, the abdominal ganglia of thesenerve cords showed prolonged synchronous bursts orirregular bursting patterns (Fig. 8A). Subsequent removal ofthe brain, SG, TG1 and TG2 had no effect, but when theconnectives between AG1 and AG2 were severed after10–20 min of exposure to CCAP, typical ecdysis motor burstsoccurred immediately in all the abdominal ganglia posteriorto the cut in eight out of 10 experiments (Fig. 8A).Interestingly, in the nerve cord containing TG3 and AG1–8,CCAP induced ecdysis bursts in AG1–3, mixed pre-ecdysis-like and ecdysis bursts in AG4–6 and pre-ecdysis-like burstsin the AG7 (Fig. 8B). This indicates that inhibitory factor(s)in the SG-TG1–3 suppress the ecdysis circuitry even after therelease of CCAP. Indeed, CCAP treatment of isolated AG2–8induced strong and typical ecdysis bursts in 3–6 min (N=10).CCAP never induced an elevation in cGMP levels.

DiscussionCentral nervous system elements responding to PETH and

ETHThe initiation of each phase of the ecdysis behavioural

sequence requires the action of PETH and ETH on specific


Fig. 7. Eclosion hormone (EH)-induced elevation in cGMP levels(immunoreactive staining) in the central nervous system. (A) Strongelevation in cGMP levels in the entire 27/704 neuronal network ofthe desheathed nerve cord, which showed ecdysis bursts in allabdominal ganglia (AG). (B) EH treatment of a desheathed AG1-terminal abdominal ganglion (TAG) induced strong cGMPimmunoreactivity in intact suboesophageal ganglion (SG) andthoracic ganglia 1–3 (TG1–3). Similarly, the action of EH ondesheathed SG-TG1–3 induced strong cGMP staining in the intact,non-desheathed AG1-TAG. Note that an elevation in cGMP levelswas also detected in L2,3 neurons (arrows). (C) EH induced variablecGMP responses in the desheathed chain of AG1-TAG, whichshowed ecdysis only in anterior abdominal ganglia. Stronger cGMPstaining was detected only in AG1, while more posterior gangliaexhibited very weak immunoreactivity. Scale bar, 150 µm.


central ganglia. Novicki and Weeks (1993, 1995) showed thatthe primary pacemaker for the pre-ecdysis I motor patternresides in the TAG, but our experiments indicate that allabdominal ganglia respond to PETH exposure. Isolation ofanterior abdominal ganglia from the TAG led to prolongeddorsoventral contractions in vivo or increased uncoordinatedbursting activity in dorsal nerves of isolated abdominal gangliain vitro. Therefore, motoneurons MN2,3 causing thesecontractions (Miles and Weeks, 1991; Novicki and Weeks,1995) may be controlled in part by PETH-activated pre-motorinterneurons in each abdominal ganglion. Nevertheless, theprimary interneurons IN-402 residing in the TAG are necessaryfor the normal rhythm to occur (Novicki and Weeks, 1995). Incontrast, each abdominal ganglion appears to contain the entireensemble of ETH-sensitive circuitry necessary for theperformance of pre-ecdysis II since individually isolatedabdominal ganglia produce bursts in ventral nerves similar induration and frequency to those in the intact nerve cord. Weconclude that all abdominal ganglia respond to PETH and ETHand, therefore, presumably contain receptors for both peptides.

Our data showed the necessary role of cephalic ganglia inthe ETH-mediated activation of the ecdysis network. Theimportance of the brain for ecdysis was reported previously byNovicki and Weeks (1996), who found that most debrainedanimals were incapable of switching to ecdysis followinginjection of EH. More recent work has demonstrated that EHacts initially through the release of ETH from Inka cells, andthat the resulting high levels of ETH in the haemolymph triggerecdysis through direct action on the CNS (Zitnan et al., 1996;Ewer et al., 1997; Kingan et al., 1997). Our in vivo experiments

show that the brain tritocerebral lobes and SG are required forETH-induced ecdysis behaviour. The failure of most debrainedlarvae to initiate ecdysis could be explained by the activity ofsome peripheral factor, which may suppress the initiation ofecdysis in vivo. However, experiments on isolated, debrainednerve cords showed that the action of ETH on the SG issufficient to induce ecdysis in virtually all cases. These dataimply that ETH may trigger ecdysis by acting on downstreamelements located in the SG in the absence of the EH-producingVM neurons. These hypothetical structures and factors remainto be identified.

Role of ETH, EH and cGMP in the the activation of ecdysis

We found that ETH (100 nmol l−1) invariably induces strongcGMP staining in a subset of neurons in isolated intact ordebrained nerve cords, and that this is coupled to thesubsequent occurrence of ecdysis. In contrast, application ofEH (100 nmol l−1) to the desheathed CNS induces quitevariable elevations in cGMP levels and ecdysis. Anteriorganglia usually show stronger staining than posterior ganglia,and no augmentation of these responses is obtained followingexposure to even higher levels of EH (200–300 nmol l−1; N=6;D. Zitnan, unpublished observations). Independent experimentsshowed that approximately the same concentrations of high-performance liquid chromatography (HPLC)-purified nativeEH (100 nmol l−1) evoked ecdysis bursts only in seven out of18 preparations and a similarly weak cGMP response inisolated desheathed abdominal nerve cords (Gammie andTruman, 1999). In contrast, we found that prolonged action ofEH on desheathed ganglia induces an elevation in cGMP levels

B

AG3D

AG5D

AG7D

AG1D

10 s

10 s

AG3D

AG5D

AG7D

A

Fig. 8. Actions of crustacean cardioactive peptide (CCAP) on the isolated central nervous system. (A) CCAP treatment of the entire desheathedcentral nervous system induced prolonged synchronous bursts or irregular bursting patterns in abdominal ganglia (AG). Transection of theconnectives between AG1 and AG2 immediately resulted in normal ecdysis bursts in AG3–7. (B) A desheathed nerve cord containing the thirdthoracic ganglion (TG3) and the complete chain from AG1 to the terminal abdominal ganglion (TAG) showed ecdysis bursts (arrowheads) inAG1–3, mixed ecdysis (arrowheads) and pre-ecdysis-like bursts in AG4–6, and pre-ecdysis-like bursts in AG7. These results demonstrate theinhibitory influence of the cephalic and thoracic ganglia on ecdysis induced by CCAP. D, dorsal.

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in intact ganglia. These data suggest that, although EH is ableto induce limited cGMP production and ecdysis bursts, itsaction on CCAP neurons is not direct and probably requiressome additional downstream signalling events.

We propose that ETH acts on multiple targets in the cephalicganglia. In addition to possible effects on the VM cells, itsprimary targets are probably other neurons and structures inthe tritocerebrum and SG, which induce a cGMP productionand activate ecdysis neurons in all the abdominal ganglia.

Role of inhibition in the transition to ecdysis

We found that the cephalic and thoracic ganglia provide aninhibitory influence on the abdominal ecdysis network thatdelays the initiation of this behaviour in pharate larvae.Although our studies indicate that inhibitory input from the SGand TG1–3 probably delays the onset of edysis, the neuronsresponsible for this function remain to be identified. Cells27/704 in the SG and in TG1–3 produce cGMP 6–8 h prior tothe onset of the ecdysis behavioural sequence, but do not showCCAP immunoreactivity (Davis et al., 1993; Ewer et al.,1994). It is possible that these cells produce an inhibitory factordelaying the release of CCAP from 27/704 homologues inAG1–7, which show cGMP staining much later, after theinitiation of pre-ecdysis. Interneurons 704 in the SG andTG1–3 are particularly good candidates for this role, since theiraxons project posteriorly throughout the entire nerve cord andmake contact with each pair of 27/704 cells in the abdominalganglia (Ewer et al., 1994; Klukas et al., 1996). This may bea mechanism whereby subsequent elevations in cGMP levelfirst activate inhibitory neurons in the SG and TG1–3, whichdelays the release of CCAP from later-activated ecdysisneurons in AG1–7.

Our ligation experiments provided evidence that removal ofthe head and thorax during early pre-ecdysis greatly acceleratesthe onset of edysis. This effect was much more pronounced invivo than in isolated nerve cords. We suggest that thisdifference may be explained by the absence of peripheralsensory input into the isolated nervous system, which may bean important signal for the switch from pre-ecdysis to ecdysis.Upon the attainment of critical internal pressures and completeseparation of the old and new cuticles, the sensory input arisingfrom the body wall and gut may relieve the inhibition and sendstimulatory commands to the ecdysis network in the abdominalganglia to release CCAP. Our experiments further showed thatCCAP does not trigger typical ecdysis bursts in the entiredesheathed CNS. Normal ecdysis activity in the abdominalganglia occurs after the inhibitory ganglia have been removedby transection. Therefore, the switch from pre-ecdysis toecdysis probably requires actions that regulate both theremoval of inhibition in the cephalic and thoracic ganglia andthe release of CCAP in all abdominal ganglia. This mechanismof transition to ecdysis is not limited to the larval stage, sincedecapitation accelerated the onset of eclosion in pharate adultsof M. sexta (Ewer and Truman, 1997) and Drosophilamelanogaster (Baker et al., 1999).

Isolation of individual abdominal ganglia 10–15 min into

pre-ecdysis leads to the initiation of ecdysis approximately20 min later. These data indicate that some independentendogenous clock within each abdominal ganglion regulatesthe timing of the onset of edysis. ETH and EH are apparentlyinvolved in the activation of the ecdysis network in theabdominal ganglia and perhaps in the stimulation of inhibitoryneurons in the SG and TG1–3, but the mechanisms regulatingthe switch from pre-ecdysis to ecdysis are unknown.

Model for PETH and ETH action on the CNS

We propose a model for activation of pre-ecdysis I, pre-ecdysis II and ecdysis by Inka cell and CNS peptides (Fig. 9).The ecdysis behavioural sequence is initiated by the release oflow levels of EH from the proctodeal nerves into thehaemolymph, which induce PETH and ETH secretion fromInka cells. PETH acts on all abdominal ganglia to activate theneural circuitry specific for pre-ecdysis I (Fig. 9). Severalneurons controlling this behaviour have been identified:interneurons IN-402 located in the TAG control rhythmicbursting of paired motoneurons MN2,3 in all abdominalganglia and corresponding dorsoventral contractions (Milesand Weeks, 1991; Novicki and Weeks, 1993, 1995). Pre-ecdysis II (Fig. 9) begins 15–20 min later as ETH activatesadditional neurons in each abdominal ganglion to producestrong posterioventral and proleg contractions. Each ganglionregulates this behaviour independently upon exposure to ETH.However, the function and identity of most of the neurons andall the neurotransmitters involved in the regulation of pre-ecdysis I and II remain to be determined.

Activation of the ecdysis network is accomplished by theaction of ETH on the cephalic ganglia, which induces cGMPsynthesis in CCAP-producing neurons 27/704 in AG1–7. Theprotocerebral VM neurons containing EH appear to increasethe excitability of this network through an elevation in cGMPlevels (Ewer et al., 1997; Gammie and Truman, 1999), but theyare not necessary in this process. Our data showed that ETHrequires the tritocerebrum and/or SG to activate the ecdysiscircuitry during the early phases of pre-ecdysis. However, theonset of edysis is delayed by the inhibitory activity of aputative ecdysiostatic factor from the SG and TG1–3 (Fig. 9).This factor may be produced by 27/704 homologues in theseganglia, which do not produce CCAP and show elevatedcGMP levels hours before the onset of the behaviour. Theswitch from pre-ecdysis to ecdysis is regulated by anindependent process in each abdominal ganglion and consistsof two steps: (i) the removal of inhibition from the SG andTG1–3; and (ii) the release of CCAP from abdominal neurons.Centrally released CCAP controls the initiation andperformance of the ecdysis motor program (Gammie andTruman, 1997). This neuropeptide may modulate the activityof MN2,3, ISMM, PPR and APR motoneurons (Fig. 9), whichcontrol muscle contractions specific for ecdysis movements(Weeks and Truman, 1984a,b). Perhaps interneurons identifiedas excitors of the APR (Sandstrom and Weeks, 1991) may alsobe important for regulation of this motoneuron activity duringecdysis. After the old cuticle has been shed, sensory input



probably reactivates inhibitory neurons to stop ecdysismovements.

The discovery and functional analysis of Inka cells resultedin the identification of two peptide hormones (PETH and ETH)that orchestrate the ecdysis behavioural sequence.Physiological experiments with these peptides led to precisedefinitions of pre-ecdysis I, II and ecdysis and to thelocalization of the regions of the CNS required for theactivation and inhibition of these motor programs. Theidentification of ETH-related peptides and correspondinggenes, which control the ecdysis behavioural sequence inBombyx mori and Drosophila melanogaster (Adams andZitnan, 1997; Park et al., 1999; D. Zitnan, L. Hollar, I.Zitnanová, P. Takác, M. E. Adams, in preparation), indicatesthat the model described here may be applicable to otherinsects and perhaps to most arthropods.

Many stereotypical behavioural sequences occur in otherinvertebrate and vertebrate animals, e.g. egg-laying inmolluscs, mating and courtship rituals in fishes and nest-

building in birds (Camhi, 1984; Scheller et al., 1984). Theextent to which peptide hormones are involved in the initiationand coordination of such behaviours is not completelyunderstood. The experimental system described here couldadvance our understanding of the mechanistic principlesgoverning general aspects of animal behaviour.

We thank Inka Zitnanová and Dr Peter Takác for criticalreading of the manuscript and helpful comments. This workwas supported by grants from the National Institute of Health(AI 40555), the National Science Foundation (IBN 9514678)and Vedecká Grantová Agentúra (95/5305/800).

ReferencesAdams, M. E. and Zitnan, D. (1997). Identification of ecdysis-

triggering hormone in the silkworm Bombyx mori. Biochem.Biophys. Res. Commun. 230, 188–191.

Baker, J. D., McNabb, S. L. and Truman, J. W. (1999). The

Pre-ecdysis I Pre-ecdysis II Ecdysis

TAG

AG1–6

SG/TG

Brain VM VM

MN2,3 MN2,3

27/704

27/704

ETH

PPRAPR

PPRAPR

Ecdysisactivation

Ecdysisinhibition

ETHdepletion

cGMPelevation

PETH

EH

EHdepletion

IN-402

ISMM

Fig. 9. Model for pre-ecdysis-triggering hormone (PETH) and ecdysis-triggering hormone (ETH) activation of the central nervous systemnetworks controlling different phases of the ecdysis behavioural sequence. When the animal is prepared to ecdyse, blood-borne eclosionhormone (EH) causes the release of PETH and ETH from endocrine Inka cells (red), which act on abdominal ganglia 1–8 (AG1–8) to initiatepre-ecdysis I and II, respectively. During the early phases of pre-ecdysis, rising ETH levels act on different targets in the brain andsuboesophageal ganglion (SG) to activate the ecdysis network. However, inhibitory input from the cephalic and thoracic ganglia (TG) delaysthe onset of edysis. An independent clock in each abdominal ganglion removes the inhibition and induces the central release of crustaceancardioactive peptide (CCAP), which triggers ecdysis (see text for further details). TAG, terminal abdominal ganglion (fused AG7,8); VM,ventromedial neurosecretory cells producing EH; 27/704, network of neurosecretory cells 27 and interneurons 704 producing CCAP andcGMP; IN-402, interneurons 402; MN2,3, motoneurons 2, 3; PPR, principal planta retractor; APR, accessory planta retractor; ISMM,intersegmental muscle motoneurons. Stippled neurons are hypothetical. All neurosecretory cells are labelled in blue, interneurons in red andmotoneurons in green.

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hormonal coordination of behaviour and physiology at adultecdysis in Drosophila melanogaster. J. Exp. Biol. 202, 3037–3048.

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