|
|
|
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online 3 May 2006
doi: 10.1242/dev.02376
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Department of Biochemistry and Cellular and Molecular Biology, University of Tennessee, Knoxville, TN 37996, USA.
* Author for correspondence (e-mail: jhpark{at}utk.edu)
Accepted 24 March 2006
 |
SUMMARY |
|---|
 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Key words: Corazonin, Programmed cell death, Metamorphosis, Ecdysone receptor, Reaper, Drosophila
|
INTRODUCTION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
; Levine et al.,
1995; Consoulas et al.,
2000; Tissot and Stocker,
2000). During metamorphosis, larval neurons face mainly two
different fates: remodeling or programmed cell death (PCD). The former is a
recycling process of persistent neurons and is characterized primarily by
significant modifications in synaptic architectures, resulting from withdrawal
of larval-specific connections, followed by reconnection with new targets
(e.g. Kraft et al., 1998;
Lee et al., 2000b). By
comparison, other neurons are scheduled to die via developmentally regulated
genetic programs, as their functions are no longer required for ensuing life
stages.
Several studies have implicated ecdysone as a central endocrine regulator
that initiates genetic programs orchestrating overall reorganization processes
of the insect nervous system during metamorphosis
(Truman et al., 1993). For
example, genetic analyses have shown that ecdysone receptor activities are
essential for cell-autonomous remodeling of mushroom body 
-neurons and
SCP-immunoreactive (IR) neurosecretory cells
(Schubiger et al., 1998;
Schubiger et al., 2003;
Lee et al., 2000b). In
addition to the remodeling, ecdysone has been shown to cause apoptosis of
obsolete larval neurons. In a moth, identified motoneurons innervating larval
proleg muscles are degenerated in response to a prepupal ecdysone surge
(Weeks, 2003). Treatment of
isolated motoneurons with ecdysone in vitro also causes the death, suggesting
that ecdysone directly induces a cell-autonomous death program
(Streichert et al., 1997).
Although underlying molecular mechanisms for post-embryonic neuronal PCD
are largely unknown, extensive genetic studies have identified key molecular
players that either enhance or suppress PCD occurring in developing
Drosophila embryos and compound eyes (reviewed by
Cashio et al., 2005;
Kornbluth and White, 2005).
Apoptotic death is a direct consequence of massive destruction of cellular
components mediated by specialized proteolytic enzymes: caspases (reviewed by
Salvesen and Abrams, 2004). In
living cells, the caspases are inactivated and/or degraded by the action of
Drosophila inhibitor of apoptotic protein 1 (DIAP1) through forming a
complex with caspases (Meier et al.,
2000; Muro et al.,
2002; Wilson et al.,
2002). When cells are challenged with death stimuli, death
activators (collectively referred to as RHG) encoded by reaper
(rpr), head involution defective (hid; W
– FlyBase) and grim, which are defined by deficiency
Df(3L)H99 (White et al.,
1994; Grether et al.,
1995; Chen et al.,
1996), bind to DIAP1, liberating caspases from the DIAP1-caspase
complexes, resulting in the activation of caspases
(Wang et al., 1999;
Goyal et al., 2000). Physical
interactions between RHG proteins and DIAP1 are also known to downregulate
DIAP1 levels via ubiquitin-mediated self-degradation of DIAP1, which further
ensures an irreversible death pathway
(Ryoo et al., 2002;
Yoo et al., 2002). Products of
two additional pro-apoptotic genes, sickle and jafrac2, are
also implicated as DIAP1 antagonists
(Christich et al., 2002;
Srinivasula et al., 2002;
Tenev et al., 2002;
Wing et al., 2002).
Fundamental molecular cell death mechanisms just described appear to be
conserved in the tissues that require ecdysone for their death at a precise
developmental stage. Larval salivary glands and midgut are degenerated in
response to an ecdysone stimulus during metamorphosis in which ecdysone
activates transcription of rpr and hid directly, or
indirectly via ecdysone-responsive transcription factors such as Broad
Complex (BR-C), E74 and E93
(Jiang et al., 2000;
Lee et al., 2002). Premature
destruction of these tissues throughout larval growth is prevented by DIAP1,
until ecdysone-induced RPR or HID proteins overcome the inhibitory action of
DIAP1 (Yin and Thummel,
2004).
During prepupal stage, a number of unwanted larval neurons are removed from
the CNS in Drosophila. Neuronal degeneration is particularly
prominent in the abdominal ganglion, leading to significant shrinkage of this
neuropil (Truman, 1990;
Truman et al., 1993). Although
ecdysone is an important developmental cue for the PCD of specific motoneurons
in a moth (Weeks, 2003),
ecdysone functions in the PCD of obsolete larval neurons in the CNS are
largely unexplored. Here, we show that peptidergic neurons expressing
Corazonin (Crz) in the ventral nerve cord (vCrz) are
programmed to die during early metamorphosis in Drosophila. Our data
further suggest that activated ecdysone signaling induces rpr
expression, and subsequent activation of caspases does not involve
diap1 functions.
|
MATERIALS AND METHODS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
; Lee and Luo,
1999).
Double homozygous lines carrying both Crz-gal4 and
UAS-lacZ (or UAS-mCD8-GFP) transgenes were
generated by genetic crosses. These flies were crossed to the following
UAS responders to produce various types of transgenic manipulations:
UAS-EcR-A, UAS-EcR-B1 and
UAS-EcR-B2 to overexpress specific Ecdysone Receptor (EcR)
isoforms (Lee et al., 2000b);
UAS-EcR-B1F645A and
UAS-EcR-B1W650A to express dominant-negative (DN)
forms of the EcR-B1 (Cherbas et al.,
2003); UAS-diap1
(Hay et al., 1995) to produce
inhibitors of apoptosis; and symUAS-diap1RNAi to knockdown
diap1 mRNA (Huh et al.,
2004). In some experiments, a double homozygous Crz-gal4,
UAS-p35 line was used for inhibition of caspases
(Hay et al., 1995).
For EcR-B-null mutant, EcR31/CyO,
y+ was crossed to EcR99/CyO,
y+, from which F1 larvae lacking y+ marker
(EcR31/EcR99) were collected and
dissected after completion of apolysis as described
(Schubiger et al., 1998;
Schubiger et al., 2003). To
rescue the EcR-B mutation, y w;
EcR31/CyO, y+;
UAS-EcR-B1 (or B2) were mated with y w;
EcR99/CyO, y+;
Crz-gal4 to obtain
EcR31/EcR99;
UAS-EcR-B1 (or B2)/Crz-gal4. The
EcR-A-null (EcR112/EcRM554fs)
and EcR-B1-null
(EcRQ50st/EcR31) mutants were produced
as described previously (Bender et al.,
1997; Carney et al.,
2004).
The following deficiencies were used to generate rpr- and
hid-null mutations: Df(3L)XR38, Df(3L)H99 and
Df(3L)X14 (for short, XR38, H99, and X14,
respectively). Homozygous deletion of the rpr locus was obtained by
combining H99 with XR38 in trans as described (cf.
Peterson et al., 2002). Flies
lacking hid functions were produced by transallelic combination of
H99 (or X14) with hid05014 allele
(Grether et al., 1995).
Although most of the hid mutants are embryonic lethal, a few escapers
enabled us to assess vCrz neuronal death in these mutants
(Peterson et al., 2002).
Null mutations of caspase-encoding dronc (Nc –
FlyBase) gene were obtained by transallelic combinations of
droncI24 (or droncI29) with
dronc51 (Chew et al.,
2004; Xu et al.,
2005). Two thread (th, diap1) alleles,
th4 (loss-of-function) and thSL
(gain-of-function), were used in some experiments
(Lisi et al., 2000). The
mutant alleles used in this study are summarized in
Table 1.
|
) was
amplified by PCR, and subcloned into pBluescript at the SmaI site,
from which an XbaI-EcoRI fragment was inserted into pPTGAL
vector (Sharma et al., 2002).
For rpr-gal4 construct, previously defined 1.3 kb rpr
upstream sequence (Jiang et al.,
2000) was inserted into pPTGAL at BglII/EcoRI
sites. Each resulting vector was mixed with pUChs
2-3 helper
plasmid (Laski et al., 1986)
and injected into the y w or w1118 embryos for
germ-line transformation (Rubin and
Spradling, 1982). Crz-gal4 (S2b, T2a) and
rpr-gal4 (III) lines were used in this study.
X-gal histochemistry and immunohistochemistry (IHC)
To detect ß-galactosidase expression, CNSs were fixed in 0.2%
glutaraldehyde, washed in PBS and then incubated in
5-bromo-4-chloro-3-indolyl-ß-D-galactoside (X-gal) solution at 37°C
overnight. The tissues were rinsed in PBS, dehydrated in ethanol and mounted
in glycerol (e.g. Park et al.,
2000).
Whole-mount Crz immunohistochemistry was performed as described previously
(Choi et al., 2005). The
anti-Crz was previously referred to as anti-CAP, which was raised against
Crz-Associated Peptide within the precursor
(Choi et al., 2005). To
co-localize EcR-immunoreactivity (IRy) in Crz neurons, rabbit anti-Crz and
mouse monoclonal anti-EcR-A (15G1a) or anti-EcR-B1 (AD4.4)
(Talbot et al., 1993) were
simultaneously applied to the CNSs. The primary antibodies were detected by
FITC- or TRITC-conjugated secondary antibodies (Jackson ImmunoResearch) at
1:200 dilution. The specimens were then cleared in 60% glycerol and mounted in
Vectashield medium (Vector Laboratory). Fluorescent signals were acquired by
Leica TCS confocal microscopy or Olympus BX61 connected with CC-12 camera.
TUNEL (terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling) assay
TUNEL was performed to detect DNA fragmentation in doomed vCrz neurons
using a commercial kit (Deadend Fluorometric TUNEL system, Promega). CNSs were
fixed in 4% paraformaldehyde in PBS for 30 minutes at room temperature, washed
in PBS containing 1% triton X-100, and then incubated in a reaction containing
fluorescein-12-dUTP and terminal deoxynucleotidyl transferase at 37°C for
1 hour. Following termination of the reaction by washing tissues with 2x
SSC, the tissues were subjected to Crz immunohistochemistry. Both TUNEL- and
Crz-IR signals were visualized by epi-fluorescence.
Double labeling by in situ hybridization and immunohistochemistry
Chromogenic in situ hybridization of Crz mRNA was performed using
digoxigenin (dig)-labeled cRNA probe as described
(Lee et al., 2000a).
Fluorescent in situ hybridization using TSA (Tyramide signal amplification)
system was employed to detect rpr transcripts within Crz neurons
(e.g. Zaidi et al., 2000). For
this, CNS tissues were first hybridized with dig-labeled rpr cRNA
probe, and then the tissues were incubated with anti-Crz (1:150,000) at
4°C overnight. After rinsing, FITC-conjugated secondary antibody was added
to the tissues for 2 hours to obtain Crz-IRy. During the final 30 minutes of
the incubation period, horseradish peroxidase-conjugated anti-dig was added to
the incubation. Finally, Cy3-labeled tyramide substrate was added to produce
in situ hybridization signals as recommended (Renaissance TSA Fluorescence
Systems, Perkin Elmer).
|
RESULTS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). In third-instar larval CNSs, Crz gene
products are detected in three pairs of DL neurons and in a pair of DM neurons
in the brain, and in eight pairs of symmetrically positioned neurons (vCrz) in
the ventral nerve cord (VNC) (Fig.
1A,C) (Choi et al.,
2005). In the CNSs taken from pupae aged for 
36 hours after
puparium formation (APF), Crz transcripts are found in 6-8 cells per
hemi-brain (Fig. 1B), but none
in the VNC. When immunohistochemistry was performed at an earlier stage (12-24
hours APF), protocerebral Crz-IR patterns remain unchanged from their larval
patterns (arrowheads in Fig.
1D), while no vCrz neurons are visible (arrow in
Fig. 1D). These temporal
expression data suggest that adult-like protocerebral Crz neurons have been
established between 24-36 hours APF, while vCrz expression disappeared within
12-24 hours APF.
The loss of vCrz expression could be due to programmed death of these
neurons. We tested this possibility by transgenic rescue of cell death using
gal4/UAS system (Phelps and
Brand, 1998). As a preliminary step, we tested fidelity of the
Crz-gal4 driver. As shown in
Fig. 1E,
Crz-gal4-driven GFP signals are completely overlapped with Crz-IRy in
third-instar larval CNS. lacZ-reporter expression also faithfully
recapitulates endogenous Crz expression in all life stages
(Fig. 2; data not shown),
suggesting that the 1.2 kb upstream sequence contains cis-elements
necessary for appropriate control of spatial and developmental Crz
expression.
Using Crz-gal4 driver, we induced expression of a caspase
inhibitor, p35 (Hay et al.,
1995), to test whether p35 can prevent cell death,
thereby maintaining Crz expression in pupal VNC. Thus, a
Crz-gal4, UAS-p35 double homozygous strain was crossed to the
UAS-lacZ, and then the progeny were processed for X-gal
histochemistry. Remarkably, all vCrz neurons marked by lacZ
expression are persistently present even at 24 hours APF
(Fig. 2B, compare with 2A).
These data strongly support that the loss of vCrz expression during
pupal development is due to caspase-dependent cell death, not due to
transcriptional silencing of the Crz gene.
Developmental timing of vCrz neuronal death
To determine developmental clock of the vCrz neuronal death,
lacZ-reported Crz expression was examined in various
prepupal stages. Elimination of vCrz neurons occurs progressively over a
2- to 6-hour period after pupariation
(Fig. 2C-F). At 2 hours APF,
one or two cells are undetectable (Fig.
2D), suggesting that execution of the death has begun in some
cells at around this stage. At 3 hours APF, 40% of the neurons are lost
in a random fashion, and staining in the projections are fainter and
discontinuous, leaving broken line-like appearance (arrow in
Fig. 2E). This perhaps reflects
phagocytic elimination of non-functional cellular components possibly by glial
cells (e.g. Sonnenfeld and Jacobs,
1995; Watts et al.,
2004). At 6 hours APF, most of X-gal signals are undetectable,
except for a few faint dots (arrow in Fig.
2F). This type of signals, which most probably reflects residual
ß-galactosidase activity, was seen in some tissues taken even after 12
hours APF (arrowhead in Fig.
4A).
|
|
; Enari et al.,
1998). To determine whether such a biochemical event occurs in
doomed vCrz neurons, we performed TUNEL to detect fragmented DNAs
(Prochazkova et al., 2003). As
our foregoing data imply that pro-apoptotic machinery comes into play almost
immediately after pupariation (Fig.
2), 0- to 3-hour-old prepupal CNSs were processed for TUNEL,
followed by Crz immunohistochemistry. As a result, TUNEL signals are detected
in 4% of total vCrz neurons examined
(Fig. 3). Lack of the signals
in other vCrz neurons may be due to unsynchronized onset of the death or
inefficient labeling reaction for whole-mounted tissues. Nevertheless, these
findings, together with the caspase-dependent cell death suggest that vCrz
neuronal death occurs in an apoptotic fashion; however, precise determination
of the type of cell death (apoptotic versus autophagic) requires electron
microscopic identification of specific cytological markers
(Baehrecke, 2003).
Ecdysone signaling initiates the death program in vCrz neurons
As the beginning of vCrz death is approximately coincident with a surge of
ecdysone at pupariation (Riddiford,
1993), and the ecdysone is a key endocrine signal orchestrating
overall metamorphic reformation of the CNS in insects including
Drosophila (Weeks,
2003), ecdysone could be a developmental cue, activating genetic
death program in doomed vCrz neurons. It is well documented that ecdysone
signal is transduced by a heterodimeric receptor complex consisting of EcR and
ultraspiracle (reviewed by King-Jones and
Thummel, 2005). The EcR-encoding gene produces three isoforms (A,
B1 and B2) via usage of different promoters and alternative splicing, which
share common C-terminal ligand and DNA-binding domains but variable N termini
(Talbot et al., 1993).
Recently, Cherbas et al. (Cherbas et
al., 2003) developed two dominant-negative (DN) forms of EcR-B1
(B1F645A and B1W650A) that lack transcriptional
activator functions, but still retain DNA-binding ability. Ectopic expression
of these mutant variants effectively blocked ecdysone-led physiological
processes, perhaps via competitively inhibiting wild-type receptor functions
(Cherbas et al., 2003). This
prompted us to test whether ectopic EcR-B1DN expression interferes
with normal vCrz-cell death. For this, progeny from
UAS-lacZ; Crz-gal4 x
UAS-EcR-B1F645A (or UAS-EcR-B1W650A)
were subjected to X-gal histochemistry. Intriguingly, all of vCrz neurons are
detectable at 12 hours APF (compare Fig.
4B,C with 4A), showing that both EcR-B1DNs are
(equally) capable of blocking vCrz PCD. The results strongly support our
hypothesis that EcR-mediated signaling plays a decisive role in the initiation
of death pathway within vCrz neurons.
EcR-B isoforms are major players for vCrz PCD
Distinct temporal expression profiles of each EcR-isoform suggest that each
isoform exhibits developmental stage-specific functions for adult CNS
formation during metamorphosis (Truman et
al., 1994). For example, EcR-B-isoforms are involved in
modifications of some larval neurons during early metamorphosis
(Lee et al., 2000b;
Schubiger et al., 2003). To
determine types of EcR involved in vCrz-cell death, we examined
isoform-specific immunoreactivity in vCrz neurons, except for the EcR-B2
because of lack of available antibody. Consistent with a previous report
(Truman et al., 1994),
widespread EcR-B1-IRy was observed in white prepupal CNS
(Fig. 4D), whereas EcR-A-IRy
was almost undetectable (data not shown). Furthermore, EcR-B1-IRy was evident
in the nuclei of vCrz neurons (Fig.
4D), implicating a role of the EcR-B1 as a signal transducer for
the PCD of vCrz.
To gain more insight into isoform-specific functions, we examined Crz-IRy
in various EcR loss-of-function mutants. In animals heterozygous for
the EcR-null mutant allele (EcRM554fs) that lacks
one copy of all EcR isoforms (Bender et
al., 1997), Crz immunohistochemistry results indicated normal cell
death, suggesting that a half dose of the EcR is sufficient for mediating
vCrz-cell death (data not shown, n>5). Consistent with the absence
of EcR-A-IRy, an EcR-A-null mutation
(EcR112/EcRM554fs) did not affect
normal vCrz PCD (compare Fig. 4F with
4E). Therefore, we conclude that EcR-A isoform is not a signal
transducer for this type of cellular event.
As documented previously, animals carrying a null mutation for both EcR-B1
and EcR-B2 were generated by a trans-heterozygous combination of
EcR31 and EcR99 alleles
(Schubiger et al., 1998).
Although most of mutant animals are developmentally arrested during the course
of larval growth, a few escapers develop into prepupae
(Schubiger et al., 2003),
thereby permitting us to assess vCrz-cell death in this genetic
background. Remarkably, Crz immunohistochemistry revealed 12
vCrz neurons at stages when vCrz neurons were normally absent (compare
Fig. 4G and
Fig. 5A with
Fig. 4E), suggesting that the
death pathway in the majority of vCrz neurons is unable to proceed in the
absence of EcR-B receptors. Therefore, our histological and genetic evidence
together demonstrates that EcR-B-isoforms play significant roles for vCrz-cell
death.
|
).
Surprisingly, complete lack of EcR-B1 showed no deficit in vCrz PCD
(Fig. 4H). This result may
suggest that only EcR-B2 is necessary for the death. Alternatively, EcR-B1 and
EcR-B2 may have redundant functions, so that either isoform alone is
sufficient for normal PCD.
To address this issue, it would be necessary to analyze EcR-B2
specific mutants. As such mutants are unavailable, we employed transgenic
rescue of the EcR-B mutant. For this attempt, we generated y
w; EcR31/CyO, y+;
UAS-EcR-B1 (or EcR-B2) and y w;
EcR99/CyO, y+; Crz-gal4 strains. These
lines were crossed with each other and F1-larvae carrying y marker
[i.e. EcR31/EcR99;
Crz-gal4/UAS-EcR-B1 (or B2)] were collected and dissected
after complete apolysis (separation of the old cuticle from the underlying
epidermis) for Crz immunohistochemistry. As individual CNS morphology was
variable depending on their developmental progress, the CNSs were classified
as `early', `intermediate' or `late' phenotypes (cf.
Schubiger et al., 2003). In
the `early' group, CNSs retain white prepupa-like morphology in which the
optic lobes are not yet extended and subesophageal ganglia are broadly
attached to the VNC. In the `late' group, optic lobes are highly developed,
and areas between the subesophageal ganglia and VNC begin to constrict; this
is nearly equivalent to wild-type at 12 hours APF
(Truman et al., 1993). In the
`intermediate' group, CNSs show various levels of development between `early'
and `late' groups.
|
Reaper (rpr) is a proapoptotic executor of vCrz PCD
The pro-apoptotic genes rpr, hid and grim, defined by
H99 deletion, are well-known death activators in Drosophila
(White et al., 1994). As
rpr and hid have been shown to promote ecdysone-mediated PCD
of the salivary glands and midgut (Yin and
Thummel, 2004), we hypothesize that one or both of these genes
might induce vCrz PCD.
The PCD of vCrz neurons was unaffected by heterozygous H99
deletion, suggesting that one copy of the wild-type rpr, hid and
grim alleles is sufficient for inducing vCrz-cell death
(Fig. 6A). Heterozygosity for
XR38 deletion, which removes rpr and sickle
(Peterson et al., 2002;
Wing et al., 2002), also did
not affect normal PCD (Fig.
6B). Intriguingly however, approximately seven pairs of vCrz
neurons survived in a rpr-null mutant (XR38/H99)
(Fig. 6C, n>5), and
a similar result was obtained by Crz-gal4-mediated GFP
expression in the same mutant background (n=4, data not shown). These
data strongly support a crucial role that rpr plays in the
destruction of vCrz neurons.
|
; Peterson et al.,
2002) showed normal PCD (Fig.
6D, and data not shown), suggesting that hid is not a
death promoter for vCrz neurons. Owing to a lack of specific mutants, possible
roles for grim were not tested. However, as grim in situ
hybridization signals were hardly detectable in larval as well as early
prepupal vCrz neurons in wild type (data not shown), we suppose that
grim might not be involved in vCrz PCD. To determine cell-autonomous roles of rpr, we investigated whether rpr is expressed in vCrz neurons by in situ hybridization. To validate our rpr cRNA probe, in situ hybridization was performed on the CNSs taken from the progeny of [Crz-gal4, UAS-p35 x UAS-rpr] cross. In this genetic context, rpr transcripts are overproduced in Crz neurons but rpr-induced death is suppressed by co-expression of the p35. As a result, we detected faint but distinct rpr in situ hybridization signals seemingly in vCrz neurons in the third-instar larva (arrowheads in Fig. 6E). Except for these cells, in situ hybridization signals were nearly blank in the entire larval CNS, suggesting that rpr is transcriptionally silent in most cells of this tissue. Having validated the rpr cRNA probe, we extended in situ hybridization to wild-type prepupal CNS. At 1.5 hours APF, numerous rpr in situ hybridization signals were revealed particularly in abdominal ganglionic region (circle in Fig. 6F), while the sense probe did not produce any signals (data not shown). Together these results suggest that rpr becomes transcriptionally active soon after pupariation, thereby inducing death of numerous obsolete larval neurons in this CNS region.
We applied rpr in situ hybridization and Crz immunohistochemistry
simultaneously to see if endogenous rpr mRNAs are also expressed in
vCrz neurons prior to death. At 1.5 hours APF, rpr in situ
hybridization signals were detectable in 10% of vCrz neurons (17 out of
177) examined (Fig. 6G).
Unsynchronized or transient expression of rpr may explain such a
minor proportion of rpr-positive vCrz neurons.
To detect rpr expression in a more sensitive manner, we generated
rpr-gal4 transgenic flies. Previously, the 1.3 kb
rpr upstream sequence was shown to contain an ecdysone response
element (EcRE) and to be sufficient for ecdysone-dependent rpr
expression in salivary glands (Jiang et
al., 2000). When F1 progeny from rpr-gal4 x
UAS-mCD8-GFP cross were processed for Crz immunohistochemistry, 17
out of 80 vCrz neurons (21%) produced GFP signals at 1 hour APF
(Fig. 6H). To further
corroborate this result, p35 expression was induced by
rpr-gal4. We reasoned that if rpr is expressed in vCrz
neurons, then rpr-gal4-induced p35 would prevent vCrz PCD
(compare with Fig. 2B).
Remarkably, Crz immunohistochemistry revealed 10 neurons even at 7-9
hours APF (Fig. 6I). Incomplete
rescue of the death by p35 is somewhat similar to the results from
rpr-null mutation (cf. Fig.
6C). These data overall support the model in which EcR-B-induced
rpr expression is a primary cause for vCrz PCD.
|
). This could
be due to anti-apoptotic factors that counteract the pro-apoptotic machinery
in larval vCrz neurons. The best candidate for this role is diap1,
the anti-apoptotic activity of which comes from inhibition of caspase activity
(reviewed by Bergmann et al.,
2003). If diap1 is important for the survival of vCrz
neurons during larval growth, then these larval neurons would contain high
levels of DIAP1. Our immunohistochemistry, however, did not detect any
DIAP1-IRy within larval vCrz neurons of wild type (data not shown,
n=12). Despite this, as undetectable levels of DIAP1 still could be
sufficient for this role, we attempted to knockdown diap1 transcript
levels via RNA interference (RNAi). For this, progeny from
symUAS-diap1RNAi x
UAS-mCD8-GFP; Crz-gal4 cross was examined for GFP
signals. Such a transgenic manipulation was intended to produce
double-stranded RNAs from complementary RNAs symmetrically transcribed from
the diap1 cDNA, which then mediate degradation of diap1 mRNA
specifically in Crz neurons (Giordano et
al., 2002; Huh et al.,
2004). Our data showed that GFP expression at 1 hour APF in
diap1RNAi was essentially the same as that in wild type
(compare Fig. 7B with 7A),
indicating that diap1-knockdown did not cause premature cell death.
Similar results were obtained when two copies of the Crz-gal4
transgene were employed to increase diap1RNAi dose or by
heat-shock-induced diap1RNAi (data not shown) (cf.
Yin and Thummel, 2004).
As the negative results could be due to the lack of effective
diap1RNAi, we checked the validity of
symUAS-diap1RNAi in salivary gland PCD. In doing so, the
34B-gal4; UAS-GFP (salivary gland-specific driver)
(Yin and Thummel, 2004) was
crossed to the symUAS-diap1RNAi, and then GFP signals were
examined in white prepupa. In contrast to intense signals seen in control,
such signals were significantly reduced by diap1RNAi
expression, reflecting premature death of this tissue
(Fig. 7C). This is consistent
with a previous report employing heat shock-diap1RNAi
(Yin and Thummel, 2004), thus
validating symUAS-diap1RNAi.
Genetic reduction of diap1 levels in half also did not accelerate
death of vCrz neurons, as mutants heterozygous for th4 or
Df(3L)brm11 alleles showed normally scheduled death of vCrz neurons
(data not shown), although these alleles dominantly exacerbate
rpr-induced death of eyes (e.g.
Lisi et al., 2000). These
overall results support that RPR-mediated caspase activation does not involve
the role of RPR as a DIAP1 antagonist.
As the pro-apoptotic action of RPR can be nullified by DIAP1 (e.g.
Lisi et al., 2000), we tested
whether gain-of-diap1 functions can suppress vCrz PCD. This was
accomplished in two ways: one is thSL allele that was
shown to suppress rpr-induced photoreceptor cell death
(Lisi et al., 2000); the other
is from transgenic diap1 expression in Crz neurons
(Crz-gal4/UAS-diap1). Surprisingly, neither thSL
nor ectopic diap1 expression suppressed vCrz PCD
(Fig. 7D,E). Similar results
were obtained from ectopic diap2 expression (data not shown). The
negative results from ectopic diap1 expression is unlikely to be due
to the lack of active DIAP1, as the same transgenic construct rescued the PCD
of salivary glands (Fig.
7F).
|
;
Muro et al., 2002;
Wilson et al., 2002), the lack
of DIAP1 function may suggest that DRONC is not responsible for vCrz death.
Contradictory to this prediction, vCrz PCD was significantly delayed in
dronc-null mutants (dronc51/droncI24)
(Chew et al., 2004;
Xu et al., 2005).
Approximately 12 neurons still survived at 7 hours APF, while the number was
reduced tô8 at 16 hours APF
(Fig. 7G), and none at 48 hours
APF (data not shown). Comparable delay of the death was observed in another
dronc-null mutant (dronc51/droncI29)
(data not shown), suggesting that DRONC is a caspase that executes vCrz-cell
death. However, the delay of death in the absence of DRONC indicates that
activities from other caspases are also required for timely execution of vCrz
PCD.
Activation of DRONC may require the adaptor protein DARK, a fly homolog of
vertebrate Apaf1 (Rodriguez et al.,
1999). If so, then lack of DARK may phenocopy dronc-null
mutants. Remarkably, Crz-IR patterns in a homozygous dark-null mutant
(darkCD4) were comparable with those of dronc
mutants (compare Fig. 7H with
7G). These data suggest that an apoptosome consisting of DRONC and
DARK is an essential component for vCrz PCD.
|
DISCUSSION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). As for the latter, a group of 300 neurons (termed type
II), which had been characterized by high levels of EcR-A expression
throughout the latter half of pupal development and a subset of neurons
expressing CCAP neuropeptide in the VNC, undergo PCD in response to the fall
in ecdysone levels (Robinow et al.,
1993; Robinow et al.,
1997; Draizen et al.,
1999). Such hormonal change triggers accumulation of rpr
and grim transcripts (Robinow et
al., 1997; Draizen et al.,
1999); at least for CCAP neurons, rpr was verified to be
a death activator (Peterson et al.,
2002). It is, however, not known how the fall in ecdysone levels
is signaled to activate rpr expression and whether rpr
promotes the death by antagonizing DIAP1 functions. In contrast to the post-eclosion neurons just described, vCrz neurons are removed via PCD soon after the onset of metamorphosis (this study). Although we have no direct evidence for ecdysone as a death signal for this event, our data suggest that activation of EcR-B in response to a surge of ecdysone at pupariation might be a key upstream molecular event that, in turn, stimulates an irreversible death pathway in which rpr plays a crucial role. Therefore, these two comparative model systems show how ecdysone regulates the PCD of distinct neuronal groups at different developmental stages.
Interestingly, EcR-B receptors are also major signal transducers for
remodeling of persistent larval neurons during early metamorphosis. For
example, SCP-IR neurons and mushroom body -neurons initially lose their
neurites at this stage (Schubiger et al.,
1998; Schubiger et al.,
2003; Lee et al.,
2000b). Genetic ablations of EcR-B prevent such
processes, and the mutant phenotype is rescued by transgenic expression of
EcR-B isoforms. Therefore, EcR-B-mediated signal transduction probably
controls remodeling of persistent neurons, as well as PCD of obsolete neurons
during this crucial developmental period. These observations then raise the
important issue of how distinct neuronal fates (remodeling versus death) are
determined in response to the same hormonal stimulus and receptor types
involved. Perhaps activated EcR-B in persistent neurons might silence the
death pathway or turn on anti-apoptotic activities.
Apoptotic pathways downstream of activated EcR
In the case of ecdysone-triggered salivary gland and midgut degeneration,
northern blotting has shown that rpr and hid are
transcriptionally induced just prior to death
(Jiang et al., 1997;
Jiang et al., 2000). Such
induction requires BR-C functions, as rpr and hid expression
in these tissues is impaired by rbp5 and
2Bc2 mutant alleles of the BR-C locus
(Jiang et al., 2000;
Lee et al., 2002). Ecdysone
also directly activates rpr transcription in the salivary glands via
an interaction between activated EcR and its consensus binding sequence (EcRE)
within 1.3 kb upstream of the rpr
(Jiang et al., 2000). Our
various findings support rpr as an intracellular death promoter for
vCrz PCD. Although we do not have definitive evidence for direct activation of
rpr by ecdysone, as the 1.3 kb rpr promoter apparently
drives p35 expression in vCrz neurons
(Fig. 6I), we favor the
hypothesis that activated EcR-B directly induces rpr transcription in
the doomed vCrz neurons. Upregulation of rpr, however, does not seem
to require BR-C, as we found normal vCrz PCD in the BR-C mutants
(data not shown).
Another downstream target of the ecdysone signaling is DRONC. In vitro
treatment of salivary glands and midgut with ecdysone induces dronc
expression (Dorstyn et al.,
1999), perhaps through a direct interaction between activated
EcR-B1 and a consensus EcRE found in the dronc promoter
(Cakouros et al., 2002;
Cakouros et al., 2004).
Upregulation of dronc transcription may be important for supplying
doomed cells with a sufficient amount of DRONC in order to conduct massive
cellular destruction in response to death signals. Further analysis will be
necessary to determine if this also occurs in doomed vCrz neurons.
Involvement of RPR and caspases in the vCrz PCD raises another fundamental
issue of how RPR leads to caspase activation. According to a current model,
interactions between RPR and DIAP1 antagonize the inhibitory action of DIAP1,
resulting in the accumulation of free active caspases (reviewed by
Bergmann et al., 2003). By
contrast, we did not find any evidence for DIAP1 as a survival factor for vCrz
neurons. Although other DIAP1-like proteins in Drosophila (DIAP2,
BRUCE and DETERIN) are possibly functional in this system, we speculate that
this might not be the case. This is because failure of ectopic diap1
expression to block vCrz PCD suggests that other DIAP family members would not
be effective in inhibiting vCrz-cell death, as anti-apoptotic functions of
these proteins are mediated by consensus BIR domains
(Bergmann et al., 2003). Thus,
we propose that RPR-mediated caspase activation occurs independently of DIAP1
in vCrz neurons.
RPR perhaps mediates an assembly of the apoptosome, as both DRONC and DARK
– two essential components of the apoptosome – are necessary for
vCrz PCD (Fig. 7G,H). In
vertebrate cells, the formation of the apoptosome triggered by the release of
cytochrome C (cytC) from mitochondria is an essential step towards caspase
activation (Liu et al., 1996;
Cain et al., 1999). Although
such molecular events are not evident in flies yet, a recent report showed
that cytC-d is able to activate caspases during spermatogenesis
(Arama et al., 2006). In this
regard, it will be interesting to determine roles of cytC as an upstream
regulator for caspase activation for vCrz PCD.
|
ACKNOWLEDGMENTS |
|---|
|
REFERENCES |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Arama, E., Bader, M., Srivastava, M., Bergmann, A. and Steller, H. (2006). The two Drosophila cytochrome C proteins can function in both respiration and caspase activation. EMBO J. 25,232 -243.[CrossRef][Medline]
Baehrecke, E. H. (2003). Autophagic programmed cell death in Drosophila. Cell Death Differ. 10,940 -945.[CrossRef][Medline]
Belyaeva, E. S., Aizenzon, M. G., Semeshin, V. F., Kiss, I. I., Koczka, K., Baritcheva, E. M., Gorelova, T. D. and Zhimulev, I. F. (1980). Cytogenetic analysis of the 2B3-4—2B11 region of the X-chromosome of Drosophila melanogaster. I. Cytology of the region and mutant complementation groups. Chromosoma 81, 281-306[CrossRef][Medline]
Bender, M., Imam, F. B., Talbot, W. S., Ganetzky, B. and Hogness, D. S. (1997). Drosophila ecdysone receptor mutations reveal functional differences among receptor isoforms. Cell 91,777 -788.[CrossRef][Medline]
Bergmann, A., Yang, A. Y. and Srivastava, M. (2003). Regulators of IAP function: coming to grips with the grim reaper. Curr. Opin. Cell Biol. 15,717 -724.[CrossRef][Medline]
Brizuela, B. J., Elfring, L., Ballard, J., Tamkun, J. W. and Kennison, J. A. (1994). Genetic analysis of the brahma gene of Drosophila melanogaster and polytene chromosome subdivisions 72AB. Genetics 137,803 -813.[Abstract]
Cain, K., Brown, D. G., Langlais, C. and Cohen, G. M.
(1999). Caspase activation involves the formation of the aposome,
a large (700 kDa) caspase-activating complex. J. Biol.
Chem. 274,22686
-22692.
Cakouros, D., Daish, T., Martin, D., Baehrecke, E. H. and Kumar,
S. (2002). Ecdysone-induced expression of the caspase DRONC
during hormone-dependent programmed cell death in Drosophila is
regulated by Broad-Complex. J. Cell Biol.
157,985
-995.
Cakouros, D., Daish, T. J. and Kumar, S.
(2004). Ecdysone receptor directly binds the promoter of the
Drosophila caspase dronc, regulating its expression in
specific tissues. J. Cell Biol.
165,631
-640.
Carney, G. E., Robertson, A., Davis, M. B. and Bender, M. (2004). Creation of EcR isoform-specific mutations in Drosophila melanogaster via local P element transposition, imprecise P element excision, and male recombination. Mol. Genet. Genomics 271,282 -290.[CrossRef][Medline]
Cashio, P., Lee, T. V. and Bergmann, A. (2005). Genetic control of programmed cell death in Drosophila melanogaster.Semin. Cell Dev. Biol. 16,225 -235.[CrossRef][Medline]
Chen, P., Nordstrom, W., Gish, B. and Abrams, J. M.
(1996). grim, a novel cell death gene in Drosophila.Genes Dev. 10,1773
-1782.
Cherbas, L., Hu, X., Zhimulev, I., Belyaeva, E. and Cherbas,
P. (2003). EcR isoforms in Drosophila: testing
tissue-specific requirements by targeted blockade and rescue.
Development 130,271
-284.
Chew, S. K., Akdemir, F., Chen, P., Lu, W.-J., Mills, K., Daish, T., Kumar, S., Rodriguez, A. and Abrams, J. M. (2004). The apical caspase dronc governs programmed and unprogrammed cell death in Drosophila. Dev. Cell 7, 897-907.[CrossRef][Medline]
Choi, Y. J., Lee, G., Hall, J. C. and Park, J. H. (2005). Comparative analysis of Corazonin-encoding genes (Crz's) in Drosophila species and functional insights into Crz-expressing neurons. J. Comp. Neurol. 482,372 -385.[CrossRef][Medline]
Christich, A., Kauppila, S., Chen, P., Sogame, N., Ho, S.-I. and Abrams, J. M. (2002). The damage-responsive Drosophila gene sickle encodes a novel IAP binding protein similar to but distinct from reaper, grim, and hid. Curr. Biol. 12,137 -140.[CrossRef][Medline]
Consoulas, C., Duch, C., Bayline, R. J. and Levine, R. B. (2000). Behavioral transformations during metamorphosis: remodeling of neural and motor systems. Brain Res. Bull. 53,571 -583.[CrossRef][Medline]
Dorstyn, L., Colussi, P. A., Quinn, L. M., Richardson, H. and
Kumar, S. (1999). DRONC, and ecdysone-inducible
Drosophila caspase. Proc. Natl. Acad. Sci.
USA 96,4307
-4312.
Draizen, T. A., Ewer, J. and Robinow, S. (1999). Genetic and hormonal regulation of the death of peptidergic neurons in the Drosophila central nervous system. J. Neurobiol. 38,455 -465.[CrossRef][Medline]
Enari, M., Sakahira, H., Yokoyama, H., Okawa, H., Iwamatsu, A. and Nagata, S. (1998). A caspase-activated DNase that degrades DNA during apoptosis, and its inhibitor ICAD. Nature 391,43 -50.[CrossRef][Medline]
Giordano, E., Rendina, R., Peluso, I. and Furia, M.
(2002). RNAi triggered by symmetrically transcribed transgenes in
Drosophila melanogaster. Genetics
160,637
-648.
Goyal, L., McCall, K., Agapite, J., Hartwieg, E. and Steller, H. (2000). Induction of apoptosis by Drosophila reaper, hid, and grim through inhibition of IAP function. EMBO J. 19,589 -597.[CrossRef][Medline]
Grether, M. E., Abrams, J. M., Agapite, J., White, K. and
Steller, H. (1995). The head involution defective
gene of Drosophila melanogaster functions in programmed cell death.
Genes Dev. 9,1694
-1708.
Hay, B. A., Wasserman, D. A. and Rubin, G. M. (1995). Drosophila homologs of baculovirus inhibitor of apoptosis proteins function to block cell death. Cell 83,1253 -1262.[CrossRef][Medline]
Huh, J. R., Guo, M. and Hay, B. A. (2004). Compensatory Proliferation induced by cell death in the Drosophila wing disc requires activity of the apical cell death caspase Dronc in a nonapoptotic role. Curr. Biol. 14,1262 -1266.[CrossRef][Medline]
Jiang, C., Baehrecke, E. and Thummel, C. (1997). Steroid regulated programmed cell death during Drosophila metamorphosis. Development 124,4673 -4683.[Abstract]
Jiang, C., Lamblin, A.-F. J., Steller, H. and Thummel, C. S. (2000). A steroid-triggered transcription hierarchy controls salivary gland cell death during Drosophila metamorphosis. Mol. Cell 5,445 -455.[CrossRef][Medline]
King-Jones, K. and Thummel, C. S. (2005). Nuclear receptors: a perspective from Drosophila. Nat. Rev. Genet. 6,311 -323.[CrossRef][Medline]
Kornbluth, S. and White, K. (2005). Apoptosis
in Drosophila: neither fish nor fowl (nor man, nor worm).
J. Cell Sci. 118,1779
-1787.
Kraft, R., Levine, R. B. and Restifo, L. L.
(1998). The steroid hormone 20-hydroxyecdysone enhances neurite
growth of Drosophila mushroom body neurons isolated during
metamorphosis. J. Neurosci.
18,8886
-8899.
Laski, F. A., Rio, D. C. and Rubin, G. M. (1986). Tissue specificity of Drosophila P element transposition is regulated at the level of mRNA splicing. Cell 44, 7-19.[CrossRef][Medline]
Lee, C.-Y., Cooksey, B. A. K. and Baehrecke, E. H. (2002). Steroid regulation of midgut cell death during Drosophila development. Dev. Biol. 250,101 -111.[CrossRef][Medline]
Lee, G., Foss, M., Goodwin, S. F., Carlo, T., Taylor, B. J. and Hall, J. C. (2000a). Spatial, temporal, and sexually dimorphic expression patterns of the fruitless gene in the Drosophila central nervous system. J. Neurobiol. 43,404 -426.[CrossRef][Medline]
Lee, T. and Luo, L. (1999). Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis. Neuron 22,451 -461.[CrossRef][Medline]
Lee, T., Marticke, S., Sung, C., Robinow, S. and Luo, L. (2000b). Cell-autonomous requirement of the USP/EcR-B ecdysone receptor for mushroom body neuronal remodeling in Drosophila.Neuron 28,807 -818.[CrossRef][Medline]
Levine, R. B., Morton, D. B. and Restifo, L. L. (1995). Remodeling of the insect nervous system. Curr. Opin. Neurobiol. 5,28 -35.[CrossRef][Medline]
Lisi, S., Mazzon, I. and White, K. (2000).
Diverse domains of THREAD/DIAP1 are required to inhibit apoptosis induced by
REAPER and HID in Drosophila. Genetics
154,669
-678.
Liu, X., Kim, C. N., Yang, J., Jemmerson, R. and Wang, X. (1996). Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell 86,147 -157.[CrossRef][Medline]
Meier, P., Silke, J., Leevers, S. J. and Evan, G. I. (2000). The Drosophila caspase DRONC is regulated by DIAP1. EMBO J. 19,598 -611.[CrossRef][Medline]
Muro, I., Hay, B. A. and Clem, R. J. (2002).
The Drosophila DIAP1 protein is required to prevent accumulation of a
continuously generated, processed form of the apical caspase DRONC.
J. Biol. Chem. 277,49644
-49650.
Park, J. H., Helfrich-Förster, C., Lee, G., Li, L.,
Rosbash, M. and Hall, J. C. (2000). Differential regulation
of circadian pacemaker output by separate clock genes in Drosophila.Proc. Natl. Acad. Sci. USA
97,3608
-3613.
Peterson, C., Carney, G. E., Taylor, B. J. and White, K. (2002). reaper is required for neuroblast apoptosis during Drosophila development. Development 129,1467 -1476.[Medline]
Phelps, C. B. and Brand, A. H. (1998). Ectopic gene expression in Drosophila using GAL4 system. Methods 14,367 -379.[CrossRef][Medline]
Prochazkova, J., Kylarova, D., Vranka, P. and Lichnovsky, V. (2003). Comparative study of apoptosis-detecting techniques: TUNEL, apostatin, and lamin B. BioTechniques 35,528 -533.[Medline]
Riddiford, L. M. (1993). Hormones and Drosophila development. In Development of Drosophila Melanogaster (ed. M. Bate and A. M. Arias), pp.899 -939. New York: Cold Spring Harbor Laboratory Press.
