1 Department of Biochemistry and Cellular and Molecular Biology, University of
Tennessee, Knoxville, TN, and Department of Biology, Brandeis University,
Waltham, MA, USA
2 Department of Neuroscience, Tufts University School of Medicine, Boston, MA,
USA
3 Zoological Institüt, University of Tübingen and Regensburg,
Germany
4 Cornell University, Entomology Department, Ithaca, NY, USA
Author for correspondence (e-mail:
je24{at}cornell.edu)
Accepted 24 February 2003
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SUMMARY |
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Key words: Molting, Neurohormone, Behavior, Pupation, Eclosion, Drosophila melanogaster
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INTRODUCTION |
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While molting (the production of the new cuticle) is regulated by the
ecdysteroid class of steroid hormones, the timing as well as the execution of
ecdysis behavior is controlled by the neuropeptides, Eclosion hormone (EH),
Ecdysis triggering hormone (ETH, and associated Pre-ecdysis triggering
hormone, PETH), and Crustacean cardioactive peptide (CCAP) (reviewed by
Ewer and Reynolds, 2002). Of
these, CCAP is believed to be the neuropeptide that turns on the ecdysis motor
program. In addition to a role in the execution of ecdysis, strong
circumstantial evidence suggests that CCAP may be one of the factors that
regulate the circadian timing of adult ecdysis (eclosion). For example, the
LARK RNA-binding protein has been implicated in the circadian control of
Drosophila eclosion (Newby and
Jackson, 1993
), and it is localized preferentially in the
cytoplasm of CCAP neurons (McNeil et al.,
1998
; Zhang et al.,
2000
).
Although our model for the hormonal control of ecdysis is consistent with
most of the available data, a number of observations suggest that the control
of this behavior occurs via a more complicated mechanism. For instance, adult
ecdysis still occurs in Drosophila lacking EH neurons
(McNabb et al., 1997).
Likewise, although the genetic deletion of the gene encoding ETH causes most
animals to die at the first larval ecdysis, many of these animals still
display ecdysis-like behavior at the end of this molt
(Park et al., 2002a
).
The complex phenotypes of these variants also raises the possibility that the role of CCAP in the control of ecdysis may not be as simple as currently proposed. Here we have used Drosophila to investigate the roles of CCAP in the control and circadian regulation of ecdysis. We find that the genetic ablation of the CCAP neurons causes defects at ecdysis. However, the type of defects observed at the ecdyses to different developmental stages as well as the severity of these defects suggest that the role of CCAP in the control of ecdysis varies during postembryonic development. In addition, although populations of flies lacking CCAP neurons exhibited a circadian rhythmicity of eclosion, the daily timing of eclosion events was abnormal in these animals, implying a modulatory role for CCAP in the circadian control of this behavior.
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MATERIALS AND METHODS |
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Generation of CCAP-GAL4 driver
We used the GAL4 system (Brand and
Perrimon, 1993) to drive gene expression in CCAP neurons. DNA
immediately 5' of the DmCCAP coding region and extending from
-516 to +39 bp was obtained by PCR using wild-type genomic DNA as a
template. The PCR products were inserted into the pPTGAL transformation vector
(Sharma et al., 2002
). The
recombinant DmCCAP promoter-GAL4 fusion (CCAP-GAL4)
construct was introduced into the germline using standard methods. Several
independent transformant lines were obtained.
Immunohistochemistry
Immunohistochemistry was performed using standard techniques (cf.
Ewer and Truman, 1996).
Primary antibodies used were rabbit anti-CCAP (used at 1:5,000; a kind gift
from Dr Hans-Jürgen Agricola, U. Jena, Germany), rabbit anti-ETH1 (used
at 1:2000; a kind gift from M. Adams and Y. Park), and mouse
anti-ß-galactosidase (anti-ß-gal; used at 1:2000; Promega).
Secondary antibodies were obtained from Jackson Immunoresearch and Molecular
Probes. Fluorescent preparations were viewed under a conventional fluorescent
microscope as well as under a confocal microscope (Biorad MRC600 with Zeiss
Axiovert inverted microscope, or a Leica DMR system).
Quantitation of immunolabeling
Fluorescently labeled tissues to be quantitated were all processed and
stained in parallel and under the same conditions. In order to quantitate CCAP
immunostaining, Z-series of confocal sections were collected at non-saturated
settings, then collapsed keeping the maximum intensity pixels. These images
were then analyzed using NIH Image. First, the background signal was
subtracted and the resulting image was smoothened. A threshold was then set
such that only the intensely stained varicosities were visible, and the number
of varicosities was counted using the same threshold for all preparations. A
`Varicosity index' was defined, based on the number of varicosities per unit
of axon length. The intensity of ETH-IR was scored qualitatively by assigning
a subjective score of 3 (strong staining) to 0 (no staining) to Inka cells.
The person scoring the preparations did not know the timepoints at which the
tissues had been fixed.
In situ hybridization
RNA in situ hybridization was carried out using standard methods (cf.
Patel, 1996). CCAP RNA probes
were labeled with DIG and visualized using either NBT/BCIP (blue reaction
product) or Fast Red (fluorescent red label; Sigma Chemical Co.). Tissues
labeled for both CCAP RNA and CCAP-IR were processed sequentially, first for
RNA in situ hybridization and reacted with NBT/BCIP, and then processed for
CCAP-IR and reacted using DAB and H2O2.
Fly strains and genetics
A UAS-lacZ line that produced cytoplasmic ß-gal expression
was obtained from the Bloomington Drosophila stock center; the
UAS-rpr strain used was obtained from H. Steller. Since driving
lacZ expression in the CCAP neurons produced ß-gal
immunoreactivity (IR) that was stronger than was the normal CCAP-IR, we
used ß-gal as an independent marker for the presence of the CCAP neurons.
Thus, for experiments involving the targeted ablation of CCAP neurons, we used
flies bearing both UAS-rpr and UAS-lacZ inserts, which were
generated by standard recombination, and are referred to as UAS-rpr +
lacZ. Targeted ablation of CCAP neurons was produced by crossing
CCAP-GAL4 flies (males were typically used) to flies bearing
UAS-rpr + lacZ. Flies used in the experiments shown in
Fig. 7A were generated by
crossing UAS-GFP flies (carrying a P{UAS-GFP.S65T} insert) to
tim-GAL4 flies (Kaneko and Hall,
2000); pdf-GAL4 transgenics have been described
previously (Park et al.,
2000
); UAS-shibirets transgenics were kindly
provided by Toshi Kitamoto (Beckman Research Institute of the City of Hope).
All flies were raised at 25°C on standard fly food under a 12 hour:12 hour
light:dark regime unless otherwise indicated.
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Pupal ecdysis
First instar larvae were collected as described above, placed in vials
containing standard fly medium, and transferred to 20°C. Animals that had
recently pupariated were examined, and those containing a bubble in mid-region
of the puparium [late stage P4(i)
(Bainbridge and Bownes, 1981)]
were selected, placed on their side on a microscope slide, and filmed at room
temperature (approx. 22°C) under dim transmitted light using a Leica DMLB
microscope (10x magnification). One experimental and one control animal
was filmed simultaneously, at one-sixth the normal speed.
Adult ecdysis (eclosion)
Pharate adults that had reached the `grainy' stage [approx. 3 hours before
eclosion (Kimura and Truman,
1990)] were placed on a microscope slide, the operculum carefully
removed in order to more clearly visualize the head movements, and the slide
placed in a humidified chamber. For consistency, only females were used.
Eclosion behavior was recorded at room temperature under a Leica dissecting
microscope at 1/6 the normal speed.
Analysis of eclosion rhythms
Crosses consisting of at least 20 males and 20 virgin females were set up
in culture bottles. As control populations, either y w; CCAP-GAL4 or
UAS-rpr homozygous flies were crossed to w1118
flies. Because of the low percentage of surviving CCAP KO adults, 20-35
bottles were set up for this genotype, whereas between eight and 10 bottles
were used for each control cross. Cultures were reared at 25°C for 5-7
days and then shifted to 18°C for the remainder of development. Developing
progeny were entrained for at least 5 days to a 12 hours light:12 hours dark
(LD 12:12) lighting schedule. Once adult eclosion commenced, bottles were
cleared every 2 hours over a 48-hour period and newly emerged adults were
counted. After this 48-hour LD collection was completed, the lights were
turned off and the populations were allowed to free run in constant darkness
(DD). After 4-5 days in DD, eclosion was then monitored (in DD) for 24 or 48
hours.
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RESULTS |
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Defining the 5' regulatory region of the CCAP gene
In order to establish that the CCAP-GAL4 transgene accurately
reproduced the expression of the DmCCAP gene, we first used it to
drive expression of the reporter lacZ, and compared the spatial
expression of the reporter to that of CCAP. In Drosophila the CCAP
peptide is consistently expressed in 2 pairs of neurons in the brain, 5 pairs
in the subesophageal ganglion, 1-2 pairs in at least 8 ganglia of the ventral
nervous system (vns) (Fig.
2A,E), as well as
in 2 pairs of strongly immunoreactive descending axons, one lateral and one
medial (see Ewer and Truman,
1996). We found no evidence of changes in the number of neurons
that expressed CCAP-IR during postembryonic development except following adult
eclosion, when there is a precipitous decrease in the number of CCAP neurons
due to their elimination by programmed cell death
(Draizen et al., 1999
). Thus,
unlike the situation in Manduca
(Davis et al., 1993
;
Loi et al., 2001
), no CCAP
immunoreactive neurons appear to be added to the pattern that is established
by the 1st instar larval stage.
We found that the GAL4 fusions bearing the -516 to +39 bp fragment of 5' DmCCAP DNA faithfully reproduced the temporal and spatial pattern of DmCCAP expression. Thus, in all cases examined, neurons that were CCAP immunoreactive were also ß-gal immunoreactive, and vice versa. The stages examined included 1st instar (0- to 2-hour, 6- to 8-hour and 21- to 24-hour-old 1st instars), mid- and late-2nd instar, pharate and wandering 3rd instar larvae, pharate pupae, pharate adults, and 6-day-old adults (late 2nd instar: Fig. 2E; other stages not shown; n>10 for each stage). All three independent transformant lines bearing this construct showed indistinguishable patterns of expression. All experiments reported here were carried out using line no. 16, hereafter referred to as CCAP-GAL4.
Targeted ablation of CCAP neurons
To produce animals lacking CCAP neurons, we drove expression of the cell
death gene reaper (rpr)
(White et al., 1994;
White et al., 1996
) in these
neurons, using the CCAP-GAL4 transgenic strain. A similar approach
has been successfully used to study the function of other Drosophila
neuropeptides and hormones (e.g. McNabb et
al., 1997
; Renn et al.,
1999
; Rulifson et al.,
2002
).
To investigate the consequences of loss of CCAP neurons on larval ecdysis, CCAP neurons had to be absent, at the latest, prior to the last larval ecdysis, that from 2nd to 3rd larval instar. To determine the extent to which targeted expression of rpr in the CCAP neurons caused their ablation prior to the end of the 2nd instar molt, we dissected the CNSs from mid-2nd instar progeny of a CCAP-GAL4 x UAS-rpr + lacZ cross, and processed them simultaneously for CCAP- and ß-gal-IR (the cytoplasmic lacZ reporter used acting as an independent and robust marker for CCAP neuronal cell bodies and processes, see Materials and Methods). In control CCAP-GAL4 x UAS-lacZ animals, CCAP- and ß-gal-IR was detectable in two pairs of neurons in the brain, around 15 pairs in the vns (average: 32.2±0.7 neurons; n=11), as well as in strongly immunoreactive descending axons (Fig. 2E). In contrast, out of 32 CCAP-GAL4 x UAS-rpr + lacZ CNSs processed, 29 had no detectable CCAP- or ß-gal-immunoreactive neurons or processes (Fig. 2F), while the remaining three CNSs had only one weakly stained neuron each but no visible stained axonal processes (not shown). Thus, the targeted expression of rpr using the CCAP-GAL4 driver produced late 2nd instar larvae that are probably entirely devoid of CCAP function.
In certain experiments that examined post-larval ecdyses, animals were transferred to 20°C after collection as first instar larvae and raised at this temperature until pupation or eclosion (see below). At this lower temperature the vast majority of the CNSs were also mostly devoid of CCAP neurons by the end of the 3rd instar (at wandering). Thus, of 22 CNSs examined at this time, 15 showed no CCAP- or ß-gal-immunoreactive neurons or processes, while four, two and one CNSs had one, two and four weakly stained neurons, respectively, and none of these CNSs had visible immunoreactive processes (not shown). When the CNS of animals raised using the eclosion rhythm paradigm (25°C to 18°C; see Materials and Methods) was processed for CCAP- and ß-gal-IR immediately after adult eclosion, 25 of 28 CNSs showed no immunoreactivity, while two and one CNSs had one and two weakly staining neurons, respectively, lacking visible processes (not shown).
Behavioral and developmental defects caused by the targeted ablation
of CCAP neurons
Larval ecdysis
In the moth Manduca sexta, addition of CCAP to an isolated larval
abdominal CNS turns on the ecdysis motor program
(Gammie and Truman, 1997b;
Zitnan and Adams, 2000
). This,
and other evidence (reviewed by Ewer and
Reynolds, 2002
), strongly implicates the CCAP neuropeptide in the
control of ecdysis behavior in this moth. In the CNS of Drosophila
larvae, CCAP-IR decreases shortly before the onset of larval ecdysis (A. C.
Clark, M. del Campo and J.E., unpublished data), suggesting that CCAP is
similarly important for the control of ecdysis in this insect.
To investigate directly the role of CCAP in larval ecdysis, we characterized animals lacking the CCAP neuronal population. Surprisingly, we found that genetic ablation of the CCAP neurons was not lethal during the larval stages. Indeed, the survival rate of CCAP KO from 1st instar to the end of the 3rd (last) instar was indistinguishable from that of the control population (97% vs. 95%, respectively; n=400 for each group). This indicates that CCAP is not essential for viability during (at least) the latter part of the 2nd larval intermolt period and the entire 3rd larval instar. Most significantly, animals lacking CCAP neurons were able to shed their old cuticle at the end of molt to the 3rd instar. Independent studies show that animals homozygous for small chromosomal deletions including CCAP (and 14 other genes) survive until the 3rd instar (J.E., unpublished data). Thus, survival of CCAP KO larvae until this stage is not due to persisting (but immunohistochemically undetectable) CCAP peptide.
To determine whether ecdysis behavior was normal in KO animals, we examined
the sequence and timing of ecdysis behavior from the 2nd to the 3rd instar.
Markers for the completion of a larval molt have been described previously
(Park et al., 2002a) (see
Fig. 3 and
Table 1). The earliest obvious
marker for the impending ecdysis is the appearance of pigmentation in the
mouth plates of the future 3rd instar (double mouth plates stage; DMP), which
occurs about 30 minutes before ecdysis. Approximately 16 minutes after the DMP
stage, air enters the new trachea, which is followed shortly by the onset of
the preparatory behavior called pre-ecdysis (see
Park et al., 2002a
).
Approximately 15 minutes after air filling, pre-ecdysis stops and the animal
executes a characteristic `biting' behavior during which it appears to be
attempting to tear the anterior region of the old cuticle. This period is then
followed by the onset of ecdysis proper, which is characterized by vigorous
peristaltic waves sweeping along the animal in a posterior-to-anterior
direction. Typically after three to four waves, the anterior cuticle breaks,
freeing the 3rd instar of its 2nd instar cuticle. After a period of a few
minutes the animal resumes feeding and locomotory behavior.
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Additionally, the subsequent events that led to cuticle shedding took approx. three times longer in CCAP KO larvae than in controls (Fig. 3A,Bb, Table 1). Both the biting period, which occurs between the end of pre-ecdysis and ecdysis onset, and the duration of ecdysis itself, were significantly extended in CCAP KO animals. Interestingly, CCAP KO larvae exhibited anterior to posterior peristaltic waves interspersed with the typically occurring posterior to anterior waves, a behavior never observed in control animals (Table 1). Because the waves moving in the anterior to posterior direction do not aid in breaking the old cuticle, the time to successful shedding of the old cuticle was lengthened.
These results reveal that the ablation of CCAP neurons is associated with defects that are strictly confined to the execution of ecdysis itself. Thus, while the entire duration of the period between DMP and ecdysis was increased by only 14%, from the normal 31.9±1.0 minutes (n=8) to 36.3±0.9 minutes (n=8), the timing and organization of ecdysis behavior itself was quite severely disrupted in the KO animals.
Pupal ecdysis
In higher Diptera such as Drosophila, pupal ecdysis (pupation)
corresponds to the behavior referred to as `head eversion'
(Zdárek and Friedman,
1986). During head eversion, the brain, which in the larva is
located behind the mouthparts, is pushed anteriorly to become positioned in
front of the thorax and the mouthparts. At the same time, the appendages,
which were formed from the eversion of the imaginal discs at pupariation, are
extended to attain their final size and shape.
In contrast to the situation observed in the larva, most CCAP KO animals died during the pupal stage. Furthermore, the appearance of KO animals at the end of pupation (Fig. 4A,B) and of metamorphosis (Fig. 4E-G) suggests that the primary cause of their death was a specific failure of pupal ecdysis. Indeed, in KO pupae and pharate adults, the head was located much more anteriorly than normal (Fig. 4A,G) or was only partially everted (Fig. 4E,F); the larval tracheae were not completely shed (Fig. 4B); and the appendages were not properly extended, resulting in a pharate adult that had abnormally short wings and legs (compare Fig. 4 E-G with 4H; Table 2).
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As shown in Fig. 4I and Table 2, CCAP KO animals initiated normal pre-ecdysis behavior (for instance the frequency of abdominal `sweeps' was the same as in controls). However, this behavior lasted significantly longer than in controls and was not followed by head eversion. Instead, abdominal pre-ecdysis movements eventually ceased during a final retraction (Fig. 4A) and were followed by a period that resembled the postecdysis period seen in the control (but which was significantly shorter in CCAP KO animals; Table 2).
Although the KO pre-pupae all lacked CCAP neurons, the morphological phenotype seen at the end of metamorphosis was somewhat variable, with, for instance, a variable amount of the adult head visible at the end of adult development (Fig. 4E-G, Table 2). However, all flies showed shortened appendages (Fig. 4E-G, Table 2), and all animals whose pupation behavior we observed in detail showed no pupal ecdysis (n=10). The basis for this variable phenotype is currently unknown,
If head eversion is stimulated by CCAP, the neuropeptide should be released
at this time. As shown in Fig.
5, a substantial decrease in CCAP-IR was indeed detected following
pupation in descending CCAP immunopositive axons. The slight increase in the
number of CCAP-immunoreactive varicosities that is apparent at the start of
pre-ecdysis is a reflection of a subtle fragmentation in the pattern of
CCAP-IR that is seen at this time, and may be the first sign that CCAP has
started to be released. The ETH peptides are known to be essential for larval
pre-ecdysis in Drosophila (Park
et al., 2002a), and the drop in ETH-IR that was observed at the
onset of pupal pre-ecdysis (Fig.
5F) suggests that these peptides may also control this behavior at
pupation.
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Adult phenotype
The phenotypes of adult CCAP KO flies suggest that CCAP neurons play some
role in post-eclosion events. KO adults do not inflate their wings, and their
cuticle appears to remain soft and untanned, as evidenced by the dimpling that
is observed on the dorsal thorax at sites of thoracic muscle insertion
(Fig. 6C). The defect in wing
expansion may be due, in part, to the failure in wing extension at the time of
pupation (see above). The tanning defect of the KO flies may occur because a
subset of the CCAP neurons expresses the gene encoding the tanning hormone,
bursicon (E. Dewey and H. W. Honegger, personal communication).
In another experiment, we employed the CCAP-GAL4 driver to
overexpress a temperature-sensitive form of shibire (shi;
the fly dynamin homolog) in the CCAP cell population (using a
UAS-shits transgene). When reared at 29°C, progeny
carrying both the CCAP-GAL4 and UAS-shits
transgenes exhibited defects in wing expansion (80-100% of the
populations), whereas control progeny (with only one transgene) had normal
wings (data not shown). At 25°C, both types of progeny had normal wings,
indicative of a temperature-sensitive effect.
Eclosion rhythms in the absence of CCAP neurons
In Drosophila, a circadian clock controls the timing of adult
emergence, with most adults eclosing between subjective dawn and late
subjective morning (Saunders,
1982). Although much is known about the circadian clock mechanism
(reviewed by Allada et al.,
2001
; Young and Kay,
2001
), comparatively little is known about how the clock regulates
the expression of overt rhythmicity (reviewed by
Jackson et al., 2001
;
Taghert, 2001
;
Wang and Sehgal, 2002
;
Park, 2002
). The
co-localization of LARK and CCAP (McNeil
et al., 1998
; Zhang et al.,
2000
) suggests that CCAP neurons could mediate the circadian
control of ecdysis, independent of its possible role in the execution of the
behavior itself. To determine whether the clock directly regulates the CCAP
cells, we examined the relative locations of the clock and CCAP neuronal
populations in larval and pharate adult brains. This was accomplished by
examining CCAP immunoreactivity in brains expressing green fluorescent protein
(GFP) in the clock cell population. Using a timeless-GAL4 driver
(Kaneko and Hall, 2000
) we
observed that projections from the TIM-containing DN2 neurons
(Kaneko et al., 1997
)
overlapped with CCAP-immunoreactive synaptic endings in the dorsal aspect of
the larval and pharate adult brains (Fig.
7A arrow, and data not shown). Interestingly, DN2 neurons are
postulated to be targets of the pigment dispersing factor (PDF)-containing
small ventral lateral neurons (LNv), and they have been implicated
in the circadian control of locomotor activity
(Helfrich-Förster et al.,
2000
). In a separate experiment using a pdf-GAL4 driver
(see Park et al., 2000
), we
demonstrated overlap between the processes of CCAP neurons and those of
tritocerebral PDF neurons (Fig.
7B, arrow). The latter population arises post-embryonically at the
mid-pupal stage, and it has been suggested that it might be involved with the
circadian control of adult eclosion
(Helfrich-Förster,
1997
).
To examine circadian rhythms of eclosion, CCAP KO and control animals were reared under conditions that produced the maximal number of pharate adults (see above and Materials and Methods), and then adult emergence was scored at two-hour intervals over the course of several days, both under a light:dark cycle (LD 12:12) and in constant darkness (DD). In three separate experiments using CCAP-GAL4 line no. 16 (Fig. 7C,D) and in two separate experiments using the independent transgenic line no. 9 (not shown), a clear rhythmicity was observed under both LD and DD conditions, with most of the animals eclosing in the dawn-early morning (or subjective dawn-early morning) interval (LD, Fig. 7C; DD, Fig. 7D). Nevertheless, there were differences between the eclosion profiles of KO and control populations. Most notably, the temporal gate of eclosion was lengthened in KO animals, with significant emergence occurring in the late night/predawn period (Fig. 7C,D; black bars). Coupled with this wider eclosion `gate', we also observed a significant diminution in the amplitude of the eclosion burst that occurs immediately following lights-on (Fig. 7C), which in control populations constitutes approximately 40% of the flies that emerge on any given day. No consistent difference in the peak time of eclosion was observed between KO and control populations in LD or DD conditions.
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DISCUSSION |
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Compensatory mechanisms in the neural bases of ecdysis
Although CCAP neurons are essential for pupation, the ecdysis motor program
of both larval and adult KO animals appears qualitatively normal, implicating
additional mechanisms in the control of these behaviors. CCAP may play a minor
role at these times or, alternatively, other neuropeptides may compensate for
the loss of CCAP. Irrespective of the exact mechanism, our results strongly
suggest that other pathways, independent or compensatory, exist, which control
the expression of these motor programs. To date the only gene that is known to
be essential for ecdysis is the ETH gene, and flies carrying the null
ETH alleles die at the first larval ecdysis
(Park et al., 2002a). However,
the ETH peptides are believed to act upstream of CCAP (reviewed by
Ewer and Reynolds, 2002
), and
in Drosophila ETH is released before CCAP at larval (M. del Campo, A.
Clark and J.E., unpublished data) and pupal (this work) ecdysis, consistent
with this hypothesis. Thus, it is unlikely that the ETH peptides act in
parallel with CCAP or can compensate for it absence. In addition, our findings
that the lack of CCAP does not cause larval lethality argue against a simple
linear pathway in which the essential function of ETH is to cause release of
CCAP leading to the initiation of the ecdysis motor program. EH is also
believed to act upstream of CCAP (reviewed by
Ewer and Reynolds, 2002
).
However, the exact role of EH in the control of ecdysis is currently unclear,
as EH KO animals are usually able to ecdyse, although their behavior is
somewhat disorganized (McNabb et al.,
1997
). An examination of the ecdysis of animals lacking both EH
and CCAP neurons compared with that of CCAP KO and EH KO animals will reveal
the extent to which EH can compensate for the lack of CCAP, and vice
versa.
Bases for the changing roles for CCAP during postembryonic
development
In addition to compensatory mechanisms, other mechanisms may contribute to
the varying importance of CCAP at different ecdyses. For instance, subsets of
CCAP neurons may participate at some ecdyses but not at others. In the
abdominal CNS of the Manduca for example, 2 pairs of
CCAP-immunoreactive neurons up-regulate the second messenger cGMP at larval
ecdysis, whereas only one pair does so at pupal and adult ecdysis
(Ewer and Truman, 1997). Since
this cGMP response likely increases the excitability of the CCAP neurons [it
is known to do so for the thoracic set
(Gammie and Truman, 1997a
)],
this change in the pattern of cGMP expression could change the relative
participation of the different CCAP neurons at each ecdysis. It is not known
if this sort of mechanism applies to Drosophila, since no cGMP
response is detected in CCAP neurons at any ecdysis in this species
(Ewer and Truman, 1996
;
Baker et al., 1999
).
Nevertheless, the differential activation of a subset of peptidergic neurons
at different times in development could provide a mechanism for modifying the
extent of the participation of these neurons in different behavioral or
developmental contexts. Alternatively, the role of CCAP may change during
postembryonic development because of changes in the expression of CCAP
receptors. Although the CCAP receptor has not been conclusively identified
(but see Park et al., 2002b
),
the completion of the Drosophila genome sequence and its subsequent
analyses has produced a list of potential candidates
(Brody and Cravchik, 2000
;
Hewes and Taghert, 2001
).
Other roles for CCAP at ecdysis
The most dramatic feature of KO animals at adult eclosion is not in the
expression of the ecdysis motor program itself, but a function that may be
cardioactive in nature. It may be that the CCAP neurons are important for
increasing hemolymph pressure, and CCAP is known to be a cardioactive peptide
in insects (see Dircksen,
1998) including Drosophila
(Nichols et al., 1999
), and to
be released at eclosion in Manduca
(Tublitz and Truman, 1985
).
Alternatively, the defect may be in fluid homeostasis. In crabs, for instance,
the shedding of the old carapace is preceded by a massive release of
hyperglycemic hormone (HH) which causes a swelling of the body via an
anti-diuretic mechanism (Chung et al.,
1999
). CCAP is also released at this time
(Phlippen et al., 2000
) and
could regulate HH release. Regardless of the bases for the defects observed in
eclosing KO animals, their phenotype suggests that maintaining a high internal
body pressure is critical for adult eclosion, and implicates the CCAP neurons
in this process.
Role of CCAP in the circadian timing of adult eclosion
Features of lark gene expression in the CCAP neurons, as well as
the potential for synaptic contact between CCAP and clock neurons suggests
that CCAP may play a role in mediating the circadian control of adult
eclosion. Although the rhythmic eclosion profile of CCAP KO populations shows
that CCAP neurons are not essential for the circadian gating of eclosion, the
distribution of eclosion events in this population indicates that these
neurons modulate the gating process. This modulation may occur via a direct
connection with clock neurons or other peptidergic neurons (e.g., those
expressing PDF), and the anatomy of CCAP neurons in the brain is consistent
with this hypothesis. The robust circadian rhythmicity of CCAP KO populations
indicates that there are multiple (and potentially redundant) cellular
pathways mediating the output of the clock.
Several lines of evidence suggest that CCAP neurons mediate the effects of
light on eclosion, indirectly via the EH neurons. In Manduca, strong
circumstantial evidence suggests that CCAP acts downstream of EH (reviewed by
Ewer and Reynolds, 2002). In
Drosophila, CCAP release occurs after EH release at larval ecdysis
(A. C. Clark, M. del Campo, and J. Ewer, unpublished data), suggesting that
the same relationship may exist in the fly. Importantly, EH KO and CCAP KO
animals both show an altered response to the light-on signal
(McNabb et al., 1997
) (this
paper), and recent evidence suggests that light can cause a premature release
of EH (S. McNabb and J. W. Truman, personal communication). Thus, it is
possible that certain CCAP neurons mediate the light-on response that is
channeled through the EH neurons.
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ACKNOWLEDGMENTS |
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We thank Anthony Clark and Landrey Milton for help in analyzing videos of eclosion and pupation, respectively. We thank Ken Kemphues for the use of the time-lapse video recorder, Hans Agricola for anti-CCAP, and Michael Adams and Yooseong Park for anti-ETH1. We are grateful to Toshi Kitamoto and Hermann Steller for UAS-shibirets and UAS-rpr transgenics, respectively, to the EDGP for DNA cosmid clones, and to the Bloomington stock center for general stocks. We are also indebted to Gyunghee Lee for assistance of germline transformation. We appreciate comments on the manuscript from Jeff Hall, and insights into the nature of the pupal lethality phenotype of CCAP KO animals from Carl Thummel. This research was supported by the University of Tennessee New Investigator Award and by NRSA-MH11946 and MH63823 to J.H.P. and by NS-44232 to J.C. Hall, Brandeis University. A.J.S. was supported by NIH NRSA MH12283. F.R.J. was supported by NIH RO1 HL59873, C.H.-F. was supported by DFG Fo207/7-3, and J.E. was supported by USDA NRICGP 2001-35302-101040.
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Footnotes |
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