|
|
|
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online 14 February 2007
doi: 10.1242/dev.02804
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Department of Zoology and Animal Biology, University of Geneva, Sciences III, 30 Quai Ernest Ansermet, CH-1211 Geneva 4, Switzerland.
Author for correspondence (e-mail:
brigitte.galliot{at}zoo.unige.ch)
Accepted 10 January 2007
 |
SUMMARY |
|---|
 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Key words: Cnidarian, Evolution, Apical patterning, Regeneration, Neurogenesis, Neuronal progenitors, Interstitial stem cells, RNA interference, ParaHox gene, ß-Tubulin
|
INTRODUCTION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
),
neuromuscular differentiation
(Miljkovic-Licina et al.,
2004; Seipel and Schmid,
2005) and regeneration
(Holstein et al., 2003;
Galliot et al., 2006). Hydra
polyps display a radial symmetry with an apex or head, centered on a mouth
opening surrounded by tentacles that catch and ingest prey, and a basal disk
at the opposite extremity. Throughout the animal, the body wall consists of
two cell layers, the ectoderm and the endoderm, separated by an extracellular
matrix called mesoglea. These two cell layers are made up of myoepithelial
cells and interstitial stem cells, which provide precursors for gland cells,
neurons, nematocytes and germ cells.
The hydra nervous system is organized as a nerve net that extends
throughout the animal and is made up of two cell lineages: the sensory
mechanoreceptor cells, named nematocytes, and the neurons, with typical
synapses (Westfall, 1996).
Those two cell types follow distinct differentiation pathways
(Bode, 1996). Nematoblasts
undergo several synchronous divisions, forming syncitial cell clusters in the
ectoderm of the body column, before differentiating a typical capsule, the
nematocyst, and migrating toward the tentacles. By contrast, neuronal
precursors follow a more direct differentiation pathway, responding to local
cues along the body axis (Fujisawa,
1989). Moreover, differentiated neurons constantly change their
phenotype as they get displaced toward the extremities
(Bode, 1992). Thus, although
hydra anatomy appears very simple, highly dynamic processes are required for
its maintenance. Besides homeostasis, morphogenesis takes place in sexual and
asexual contexts, such as budding, regeneration and reaggregation. Head
regeneration, which leads to the replacement of the missing part after 2 days,
relies on the setting up of an organizer activity at the regenerating tip
(MacWilliams, 1983). This
requires the sequential activation of a specific set of `early' genes within
the endodermal cells of the regenerating tip, followed 16-20 hours
post-amputation (hpa) by the activation of the `early-late' genes at the head
patterning stage (Galliot et al.,
2006), when an intense cell proliferation takes place in the
regenerating tip (Holstein et al.,
1991).
According to several independent datasets, neurons are thought to play a
minor role in de novo head patterning: chimera experiments demonstrated the
primary role of myoepithelial cells in budding rate and regenerative capacity,
whereas nerve-free hydra display amazing budding and regenerative abilities
(Fujisawa and Sugiyama, 1978;
Marcum and Campbell, 1978).
More recently the systematic screening of hydra peptides with morphogenetic
function identified mostly epitheliopeptides
(Fujisawa, 2003). However,
neurons were also shown to produce morphogenetic peptides involved in head
differentiation (Schaller et al.,
1989; Javois and
Frazier-Edwards, 1991) and interstitial cells (i-cells) can
regulate the morphogenetic potential of the myoepithelial cells
(Sugiyama and Wanek, 1993).
Therefore, the interplay between interstitial and myoepithelial cells in
homeostatic and developmental contexts appears essential but largely
unknown.
In this work, we have investigated the putative role of neurogenesis in
head regeneration by dissecting the cellular and developmental regulation of
cnox-2, the hydra gsx homolog gene. Cnidarian
gsx/ind-related genes were identified in numerous cnidarian species,
and their expression patterns suggest an ancestral gsx/cnox-2
function in neurogenesis and oral patterning. The cnox-2/anthox2
genes are activated during apical patterning in hydra
(Schummer et al., 1992;
Gauchat et al., 2000), display
an apical expression in the Hydractinia gastrozooid polyps
(Cartwright et al., 1999) and
the sea anemone Nematostella juvenile polyps
(Finnerty et al., 2003). In
the Nematostella planula larva, anthox2 transcripts are
localized at the posterior pole, which provides the future oral pole of the
polyp. In the early Podocoryne larva, the gsx transcripts,
initially localized in the anterior endoderm, extends toward the posterior
pole (Yanze et al., 2001).
However, in the coral Acropora developing larva, cnox-2
expression was detected in neurons of the ectodermal central region of the
body axis and rarely in the oral region
(Hayward et al., 2001),
suggesting some variability in the developmental but not the cellular
regulation of the cnidarian gsx/cnox-2 gene family. Nevertheless, in
hydra, contradicting data were published as immunochemistry analyses showed a
predominant cnox-2 expression in the myoepithelial cells of the body column
and a repression during head formation
(Shenk et al., 1993a;
Shenk et al., 1993b).
Therefore a gsx/cnox-2 consensus function remains questionable in
cnidarians (Schierwater et al.,
2002). We show here that cnox-2 expression in hydra is
indeed restricted to the nervous system in both homeostatic and developmental
contexts. By analysing the proliferation rate of cnox-2+
cells, we identified a subpopulation of interstitial stem cells that behave as
bipotent neuronal progenitors, giving rise to apical neurons and nematoblasts.
Functional assays using RNAi in wild-type hydra indicate that those
cnox-2+ progenitors promote apical neurogenesis and head
patterning during head regeneration.
|
MATERIALS AND METHODS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). sf-1 mutants were kept for 2 days at 28°C to
eliminate i-cells, and they were subsequently amputated and maintained at
28°C, whereas control sf-1 hydra were kept at 18°C. Bisection
was mid-gastric in all regeneration experiments.
Bromodeoxyuridine labeling coupled to whole-mount in situ hybridization
Intact hydra (Hm) were continuously incubated with
bromodeoxyuridine (BrdU) 5 mmol/l (Sigma) for 2, 24 or 48 hours, immediately
fixed, processed for cnox-2 whole-mount in situ hybridization
(WM-ISH) and detected for BrdU as in Gauchat et al.
(Gauchat et al., 2004). For
regeneration experiments, intact hydra were incubated for 4 hours in BrdU,
washed, immediately bisected and left in Hydra medium. cnox-2 Hv cDNA
was amplified using M13 forward-20 and reverse primers or restricted at the
EcoRI site. cnox-2 riboprobes were either DIG- or
fluorescein-labeled. Imaging was performed on Axioplan2 and LSM 510 META
confocal microscopes (Zeiss).
Double-labeling WM-ISH with tyramide detection
After hybridization to the mixed DIG-labeled hyZic and
fluorescein-labeled cnox-2 riboprobes, the samples were first
incubated in the mouse anti-FITC antibody (1:100, Sigma) and detected with the
TSA-Kit#2 (Alexa 488, Invitrogen or Molecular Probes). For the second probe a
sheep anti-DIG antibody (1:400, Roche) was used, followed by the detection
with an anti-sheep HRP (1:100, Sigma) and the TSA-Kit#41 (Alexa 555). The
tyramide labeling time was 18 minutes. Samples were washed 3x 10 minutes
in PBS, mounted in Mowiol and pictured either at the Zeiss Axioplan2
microscope using a FITC filter or at the Leica SP2 confocal.
Immunohistochemistry with tyramide detection
Standard immunohistochemistry was performed on either freshly fixed or in
situ processed whole-mount animals. Samples were washed in PBS for 3x 10
minutes, treated with 3% H2O2/PBS for 1 hour, blocked in
2% BSA and incubated in the mouse monoclonal ß-tubulin antibody (1:1500,
Sigma clone 2-28-33) ON at 4°C. Subsequently the animals were washed for
3x 10 minutes in PBS and incubated in the anti-mouse HRP antibody
(1:100) for 3 hours at room temperature. The samples were detected with the
TSA-Kit#2 as above.
Double-stranded RNA interference
The cnox-2 Hv 795 bp PCR product amplified with the
cx2-hv-Xmn5' (GAACCTCTTCTTAAAACGAGTTAG) and the cx2-hv-16HB3
(GTAGGGGAATAGCTATATCCTTTCTTA) primers was inserted into the
SmaI-digested double T7 vector pPD129.36 (L4440, Fire's laboratory).
Double-stranded RNAs (dsRNAs) were produced in HT115(DE3) bacterial strain
(Timmons and Fire, 1998).
Fifty intact Hv hydra per condition starved for 2 days were given
every other day up to nine times grinded agarose that contained either no
bacteria, or bacteria having produced control dsRNAs (transformed with the
L4440 vector) or dsRNAs corresponding to cnox-2 or Kazal1
(Chera et al., 2006). The day
following the last dsRNA exposure, samples were processed for RT-PCR or
WM-ISH, or bisected for regeneration experiments.
Semi-quantitative RT-PCR analysis
mRNA was prepared from intact hydra (Hv Basel) or upper halves
collected immediately post-amputation with the QuickPrep micro-mRNA
Purification Kit (Amersham) and resuspended in 10 µl H2O. mRNA
(50 ng) was used for Sensiscript Reverse Transcription (Qiagen) and specific
cDNAs were PCR amplified for up to 24 cycles with gene-specific primers
(sequences on request).
|
RESULTS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
To verify that cnox-2 expression was restricted to i-cell
lineages, we tested the sf-1 mutants that lose their i-cells after a
2 day temperature shift but maintain their myoepithelial cells intact
(Marcum et al., 1980). At
permissive temperature, cnox-2 expression was as in wild type
(Fig. 1G,H), whereas at
restrictive temperature cnox-2 expression was abolished
(Fig. 1I,J), confirming that
cnox-2 expression is absent from the myoepithelial cell lineages. In
addition, in wild-type hydra, none of the other i-cell derivatives, including
gland cells (not shown), testes and ovaries
(Fig. 1K-N) expressed
cnox-2. In female hydra, the region facing the ovary was
cnox-2 negative (Fig.
1K,L), probably because of the recruitment of i-cells as nurse
cells by the mature oocyte. Therefore cnox-2 expression appears
restricted to two cell lineages among the i-cells: neuronal at the apical pole
and mechanoreceptor along the body column.
cnox-2+ i-cells are cycling cells
Single or twin i-cells form a heterogeneous cell population that includes
self-renewing stem cells and precursors to a variety of cell types including
gametes, gland cells, neuronal cells and nematoblasts
(Holstein and David, 1990a).
As cnox-2 transcripts were detected neither in gland cells nor in
gametes, the cnox-2+ i-cells may represent three distinct
populations: self-renewing interstitial stem cells, neuronal precursors and
nematoblast precursors. The cycling activity of cnox-2+
cells was measured through continuous BrdU-labeling
(Fig. 2): in body column over
90% cnox-2+ cells were BrdU-positive after 2 hours
(Fig. 2A,D,G), reaching 100%
after 24 hours (Fig. 2B,E,G).
Given that S-phase lasts about 12 hours in all i-cells
(Campbell and David, 1974) and
that a 15 minute incubation is sufficient to detect BrdU-incorporation (data
not shown), we calculated that the gastric cnox-2+ cells
would traverse the cell cycle in about 16 hours if cells are not
synchronized.
|
Starvation affects cnox-2 expression in the body column but not in the apex
Hydra starvation leads to dramatic cellular rearrangements in both
epithelial (Bosch and David,
1984; Holstein et al.,
1991) and interstitial
(Gauchat et al., 2004) cell
compartments. In case of cnox-2+ cell clusters, their
distribution was not significantly affected over the first 3 days, except for
a slight decrease in the single i-cell compartment
(Fig. 1G). However, when
starvation was prolonged over 10 days, a drastic reduction in the number of
gastric cnox-2+ cells was recorded
(Fig. 3G), whereas the number
of apical cnox-2+ cells did not vary
(Fig. 3F). Hence starvation
drastically alters the production and/or survival of
cnox-2+ nematoblasts, but not of the apical
cnox-2+ lineage.
Efficient cnox-2 downregulation through RNAi
We next tested cnox-2 function through RNAi loss-of-function
assay. Kazal1 was selected as control gene, as its expression is
restricted to a distinct i-cell lineage, the gland cells
(Chera et al., 2006). Moreover,
cnox-2 and Kazal1 did not show any epistatic relationships,
as evidenced by their conserved respective expressions in
Kazal1(-) and cnox-2(-) hydra
(Fig. 3B,
Fig. 4A,B). cnox-2
silencing was effective and specific, as shown by RT-PCR and WM-ISH assays:
hydra exposed repeatedly to cnox-2 dsRNAs exhibited a drastic
reduction in cnox-2 transcripts levels, whereas actin,
Kazal1 and CREB levels remained unaffected
(Fig. 3A,
Fig. 4B). When whole hydra were
compared to upper halves (enriched in apical tissue), cnox-2
silencing appeared stronger in the former ones, even though the control level
of cnox-2 expression was significantly lower in whole hydra than in
apically enriched tissue (Fig.
3A, lanes 1,3). Therefore, the decrease in cnox-2 levels
in the body column probably corresponded to the addition of two distinct
effects: the specific cnox-2 RNAi knockdown and the starvation effect
described above, leading to a drastic decrease in nematoblast production.
WM-ISH confirmed the progressive disappearance of cnox-2+
cells with the successive dsRNA exposures, becoming very rare after five
feedings in gastric (Fig. 3C)
and apical (Fig. 3D,E) regions.
The number of cnox-2+ apical cells was significantly
reduced after five exposures, by 55 and 38% when compared to mock-silenced and
Kazal1(-) hydra, respectively
(Fig. 3F). In body column, a
single dsRNA exposure led to a 48% decrease in cnox-2+
cell number (Fig. 3G, lanes
1-3). However, after five exposures, the 12 day starvation effect was
dramatic, and the difference between control and cnox-2 RNAi hydra
was no longer significant (Fig.
3G, lanes 4-6). This loss of function obviously affected hydra
homeostasis, as cnox-2(-) hydra progressively became
significantly smaller than Kazal1(-) or control hydra
(Fig. 3B,C): about twice as
short as control hydra after nine exposures (hydra exposed 9x to
cnox-2 dsRNAs: 0.44 mm ± 0.18; to control dsRNAs: 0.85 mm
± 0.28, n=10).
|
) that stains all types of neurons, nematocytes and some
i-cells, but not nematoblasts or myoepithelial cells
(Fig. 1D,E, and see Fig. S1G,H
in the supplementary material). This staining showed a highly organized ANS,
with parallel processes of the sensory neurons along the hypostome (the domed
region from the mouth opening down to the tentacle zone) providing an
inverted-basket aspect (Fig.
3H,I, arrows, and see Fig. S2 and Movie S1 in the supplementary
material). On sagittal views, the multipolar neurons were visible, spread
along meridian lines that converge toward the mouth opening
(Fig. 3H). After five
cnox-2 dsRNA exposures, the ANS was dramatically reduced, restricted
to its most apical part, whereas the sub-tentacle zone became nerve-free. The
density of sensory neurons was decreased, the systematic parallel orientation
of their processes was altered and their connections to the multipolar neurons
were no longer visible (Fig.
3J,L, and see Fig. S2 in the supplementary material). After nine
exposures, apical neurons were no longer detected
(Fig. 3N), indicating that
cnox-2 expression is required to maintain apical neurogenesis and ANS
organization.
cnox-2 is an upstream regulatory gene in the nematocyte differentiation pathway
Previous cellular expression analyses identified several candidate genes as
regulators of the nematocyte differentiation pathway. Those genes display
cellular expression patterns with striking differences when considering the
proliferation rate, the cluster size and the presence of a differentiating
capsule. cnox-2 is expressed in precursors and at early proliferative
stages but is repressed as soon as nematoblasts differentiate (this work);
hyZic is also expressed at early stages
(Lindgens et al., 2004), but
with a lower representation among precursors (6% for hyZic, over 20%
for cnox-2) and a lower BrdU-labeling index after a short incubation
than cnox-2 (40 versus 95%). Therefore, hyZic is probably
turned on at a slightly later stage than cnox-2 and appears to be a
candidate cnox-2 target gene. As additional regulators, the
hyCOUP-TF orphan receptor was proposed to repress proliferation in
nematoblast clusters, switching them to differentiation
(Gauchat et al., 2004),
whereas the achaete-scute homolog CnASH is expressed in
non-proliferative differentiating nematocytes
(Lindgens et al., 2004).
Finally, the transcription factor CREB is expressed at all proliferative
stages of this pathway (Chera et al.,
2007).
Cellular expression analysis showed that cnox-2 and hyZic transcripts indeed colocalize in most i-cells and nematoblast clusters (Fig. 4A). Moreover, in cnox-2(-) hydra, hyZic and CnASH transcripts were undetectable, hyCOUP-TF was upregulated and CREB was unaffected (Fig. 4B, left). These results support an upstream role for cnox-2 (Fig. 4C).
cnox-2 regulates apical neurogenesis
As well as the nematocyte pathway, hyCOUP-TF, CnASH and
CREB are also expressed in the neuronal lineage
(Gauchat et al., 2004;
Hayakawa et al., 2004;
Chera et al., 2007), whereas
other genes display a neuronal-specific expression, such as the PRD-class
prdl-a (Gauchat et al.,
1998) and gsc (Broun
et al., 1999), the ANTP-class msh
(Miljkovic-Licina et al.,
2004) and the RFamide neuropeptide genes
(Mitgutsch et al., 1999).
Their RT-PCR expression analysis in cnox-2(-) and control
hydra showed a significant downregulation of prdl-a, gsc and
RFamide-B, whereas hyCOUP-TF was found to be upregulated
(Fig. 4B, right), indicating
that cnox-2 contributes to the regulation of those neuronal apical
genes. Moreover, msh expression in the gastric region was slightly
decreased, suggesting that gastric cnox-2+ i-cells also
provide neuronal precursors in this region.
Concomittant de novo neurogenesis and cnox-2 upregulation during apical patterning
De novo neurogenesis, restricted to head-regenerating tips and occurring at
a time depending on the bisection level, early after decapitation and
`early-late' after gastric bisection, was previously reported
(Venugopal and David, 1981).
To monitor the cellular remodeling occurring in head-regenerating halves, we
used the anti-ß-tubulin staining and observed the complete disappearance
of any neurons from head-regenerating tips for the first 16 hpa
(Fig. 5A, and see Fig. S3 in
the supplementary material). Thereafter, pairs of i-cells appeared
(Fig. 5A, arrows), becoming
more and more numerous, until a neuronal network was detected again when
tentacle buds (TBs) emerged.
|
), was specific to the head-regeneration process, as it
was not detected in foot-regenerating tips whatever the time point
(Fig. 5B,C, arrowheads). By
contrast, the cnox-2+ neuronal density in the heads of
foot-regenerating halves remained stable all through the regeneration process
(Fig. 5C, right panel).
Along the body column a simultaneous decrease in the number of
cnox-2+ cells in both head- and foot-regenerating halves
was observed at 24 and 36 hpa, lowered by 40-50% of the normal level in
undisturbed polyps (Fig. 5C,H).
This downregulation affected proliferating nematoblasts but not single
i-cells, those actually showing a 2-fold increase of their density at 16 hpa
(Fig. 5I). Therefore this
reduction in gastric cnox-2+ cells probably corresponds to
the cell death of differentiating nematoblasts linked to the amputation stress
as previously reported (Fujisawa and
David, 1984), a scenario supported by the transient relative
increase in single cnox-2+ i-cells.
|
), corresponds to the onset of the early-late transition
during regeneration. Similarly to head-regenerating tips, those
cnox-2+ cells were identified as dividing i-cells,
neuronal precursors and differentiated neurons, first broadly distributed,
becoming restricted to the tentacle zone and the basis of the hypostome at
stage 7-8. Therefore, during budding and regeneration, cnox-2
exhibits a similar spatiotemporal regulation in the presumptive head region;
the initiation of its expression in the neuronal cell lineage precedes by 20
hours the emergence of the TBs. However, during budding,
cnox-2+ expression in the body column of the growing bud
was not altered (Fig. 5G),
indicating that the negative `post-amputation' gastric regulation does not
take place here.
De novo differentiation of cnox-2 neurons in head-regenerating tips
To trace back the origin of the cnox-2+ neurons
detected in head-regeneration tips, hydra were BrdU-labeled for 4 hours before
bisection and BrdU+/cnox-2+ cells were analysed
during early-late regeneration (Fig.
6). From 20 to 44 hpa, the density of BrdU+ cells
progressively increased in regenerating tips
(Fig. 6A-C), including
BrdU+/cnox-2+ cells surrounding the newly
formed mouth opening (Fig.
6B,C). Interestingly most cnox-2+ cells were
BrdU+, corresponding to pairs of i-cells
(Fig. 6D,F) and differentiated
neurons (Fig. 6E). These pairs
of i-cells, either asymmetric (Fig.
6D) or symmetric (Fig.
6F), frequently exhibited elongated processes, indicating that
neuronal differentiation was taking place, resulting in mature neurons that
were rare at 20 hpa but frequent at 44 hpa, still exhibiting some BrdU
labeling. In a few BrdU+ pairs of differentiating neurons, one of
the two cells did not express cnox-2 (not shown). These data indicate
that the cnox-2+ neurons detected at the early-late phase
of head regeneration result from a differentiation process of
cnox-2+ neuronal precursors, rather than induction of
cnox-2 expression in mature neurons that would migrate and express
cnox-2 once they have reached the regenerating tip.
The blockade in head regeneration in the nerve-free sf-1 mutant correlates with the lack of cnox-2 expression
sf-1 hydra maintained at a permissive temperature regenerated
their head efficiently, although delayed by 60 hours when compared with
wild-type hydra (Fig. 7A). At a
restrictive temperature, regeneration was poorly efficient, as 45% hydra had
not regenerated their head after 6 days and subsequently died
(Fig. 7A). To test a possible
correlation between the level of cnox-2 expression and the efficiency
of the head regeneration process, hydra were collected at different time
points and sorted out according to both their cnox-2+
apical cell number (null, low: fewer than 20, normal: more than 20) and their
phenotype (Fig. 7B). The
phenotypes were identified as Stage 1, `ball-shape' with no obvious apical
pole; Stage 2, `elongated' with clearly distinguishable basal and apical
poles; Stage 3, `TB1' when the TBs have just emerged; Stage 4, `TB2' when more
than two TBs elongate; Stage 5, `fully head-regenerated', similar to the adult
head (Fig. 7C-H).
|
By contrast, the cnox-2 expression pattern in regenerating sf-1 hydra maintained at a permissive temperature was similar to that recorded in wild-type hydra: numerous cnox-2 apical cells at the elongated shape stage (Fig. 7I) and a dense cnox-2+ neuronal network at later stages (Fig. 7J,K). Along the body column, cnox-2+ cell clusters became visible first in the aboral region (Fig. 7I) and then homogenously distributed along the axis (Fig. 7K). These data suggest a correlation between the level of cnox-2 apical expression and the efficiency of the head-regeneration process, even though in a few cases (7%), a complete head regeneration process occurred when cnox-2 expression was lacking or only transient.
Knocking down cnox-2 expression prevents de novo neurogenesis and apical patterning during head regeneration
To test the cnox-2 function during head regeneration, hydra were
exposed to dsRNAs repeatedly and bisected. Emergence of TBs was delayed by 42
hours in those hydra when compared with controls and at 39 hpa, the ANS was
not formed (Fig. 8A,I,J). The
number of cnox-2+ apical cells detected in
head-regenerating tips of Kazal1(-) hydra increased over
time in a similar way to non-treated hydra
(Fig. 8B,D,F). By contrast, at
30 hpa twice as few cnox-2+ apical cells were detected in
cnox-2(-) hydra (Fig.
8E, arrow) as in Kazal1(-) or control hydra (55
and 53%, respectively, Fig.
8B). At 48 hpa, two distinct phenotypes were observed: strong
(Fig. 8G) when the animal size
was small, the regeneration stacked as evidenced by the absence of TBs
(arrowheads in Fig. 8F) and the
number of cnox-2+ apical cells low
(Fig. 8B); weak when the animal
size was larger (although smaller than in Kazal1(-) hydra),
the appearance of TBs only delayed (Fig.
8H, arrowhead) and the number of cnox-2+
neurons closer to that observed in control hydra (35 per tip,
Fig. 8B, black bar). Thus a
clear correlation between the level of cnox-2+ apical
expression and the efficiency in de novo head formation was observed in
cnox-2 dsRNA-treated hydra.
|
|
DISCUSSION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
), also
share a common set of evolutionarily-conserved regulatory genes, which, in
addition to the ParaHox gsx/cnox-2 gene (this work), includes the
hyCOUP-TF nuclear orphan receptor, the PRD-CLASS homeogene
prdl-b (Gauchat et al.,
2004) and the Achaete-scute homolog CnASH
(Hayakawa et al., 2004). None
of these four genes were found expressed in any other derivatives of the
interstitial stem cells or in myoepithelial cells. Among the
hyCOUP-TF+ and prdl-b+ cells, single
i-cells were not observed and pairs of dividing i-cells were rare
(Gauchat et al., 2004).
Furthermore, CnASH is a marker for non-proliferating differentiating
nematoblasts (Lindgens et al.,
2004) and differentiating sensory neurons
(Hayakawa et al., 2004).
Therefore, cnox-2 occupies a unique position among the putative
regulators of the hydra nervous system, as it identifies a highly
proliferative subset of i-cells, the progeny of which are restricted to
neuronal and mechanosensory cell fates. Similarly, a subpopulation of i-cells
was shown to be restricted to a gametic fate
(Littefield, 1991;
Nishimiya-Fujisawa and Sugiyama,
1993), but this is the first report concerning the somatic
lineages. Previous studies showed that neuronal precursors from the peduncle
could actually provide nematocyte precursors after transplantation into a
gastric environment (Holstein and David,
1990b), suggesting that these two cell lineages indeed share
common specific precursors. In the absence of one of the criteria that define
stem cells, i.e. the evidence that these cells can self-renew
(Weissman et al., 2001), we
propose to consider the cnox-2+ i-cells as progenitors to
neurons and mechanosensory cells (Fig.
4C). Interestingly, in mice, the gsx homologs
Gsh1 and Gsh2 appear to control the size and the identity of
neuronal progenitor pools (Toresson and
Campbell, 2001; Yun et al.,
2003).
cnox-2 is an upstream regulator of neurogenesis and nematocyte differentiation
Although genes expressed in neurons are readily refractory to RNAi
(Tavernarakis et al., 2000),
the feeding RNAi strategy we applied for the first time in hydra to silence a
neurally expressed gene, indeed induced specific effects, including a drastic
disorganization of the ANS and a deficient head-regeneration process. In both
contexts, intact or regenerating hydra, the number of
cnox-2+ cells (i-cells, apical neurons, nematoblasts) was
significantly reduced after repeated exposures to cnox-2 dsRNA.
Moreover, the prdl-a, gsc, RFamide-B and RFamide-C that are
specifically expressed in apical neurons, were strongly downregulated.
Interestingly, the role of cnox-2 in neurogenesis also probably
extends to the body column, as the expression of the non-apical neuronal gene
msh was altered in cnox-2(-) hydra. However, in the
body column, we did not identify cnox-2+ neurons, suggesting that in
this region cnox-2 function is restricted to neuronal progenitors,
whereas in the apex cnox-2 would be required at least at two distinct
levels, to commit interstitial cells to the neuronal fate and to promote the
differentiation of multipolar neurons.
|
) and is
upregulated in cnox-2(-) hydra. As previously proposed,
hyZic might regulate CnASH positively in differentiating
nematocytes (Lindgens et al.,
2004). At least two genes expressed in this pathway were not
affected: CREB and NOWA, a structural gene involved in
nematocyst differentiation (Engel et al.,
2002). CREB could act upstream of cnox-2 in this
cell lineage. However, as CREB is strongly expressed in i-cells, proliferating
nematoblasts, differentiated neurons and myoepithelial cell lineages
(Kaloulis et al., 2004;
Chera et al., 2007), its
regulation within a specific cell lineage cannot be uncovered in RT-PCR assays
and requires further analyses. Finally the stable low level of NOWA
transcripts suggests that a longer period of cnox-2 silencing might
be required before differentiated nematocytes are affected.
|
). As
cnox-2 expression in the presumptive head region resumes after 16
hpa, cnox-2 belongs to the `early-late' genes. Previous hydroxurea
experiments showed that cells in S-phase at the time of amputation provide
newly differentiated neurons in head-regenerating tips
(Venugopal and David, 1981;
Holstein et al., 1986). The
analysis of the BrdU+/cnox-2+ cells confirms
this result: i-cells in S-phase at amputation time undergo mitosis and
neuronal differentiation 20 to 30 hours later in head-regenerating tips. As
those tips contain very few BrdU+ cells immediately after
amputation (Holstein et al.,
1991) (S.C. and L.G., unpublished), we anticipate that those cells
first migrate toward the tip and become cnox-2+ upon
exposure to head-specific differentiation signal(s). This scenario fits with
the injury effects previously described: injury promotes the migration of
i-cells (Fujisawa et al.,
1990), which is enhanced by the signals released by the
head-regenerating tips (Teragawa and Bode,
1991). Those signals also drive the terminal differentiation of
neuronal precursors within about 18 hours
(Venugopal and David, 1981;
Holstein et al., 1986). Hence,
cnox-2 appears to be a target gene for neurogenic signals during head
formation. A similar continuous phenotypic maturation of neurons occurs from
the gastric region to the basal extremity, accelerated upon amputation
(Technau and Holstein, 1996),
suggesting that de novo neurogenesis rather than phenotypic conversion of
existing neurons (Bode, 1992)
is the predominant mechanism during regeneration.
i-cell proliferation and de novo apical neurogenesis support head patterning in wild-type hydra
In two distinct contexts in which cnox-2 expression was reduced or
undetectable, i.e. RNAi-silenced wild-type hydra and sf-1 mutants,
head regeneration was dramatically altered. As cnox-2 supports
proliferation of i-cells and de novo neurogenesis at the early-late stage,
these results support the importance of these two cellular processes in
head-regenerating tips for head patterning, and the possible contribution of
neuronal cells as previously proposed
(Schaller et al., 1989).
Nevertheless, in a few sf-1 hydra in which the number of
cnox-2+ cells was low or null, the head regeneration
process was advanced or complete. We anticipate that such animals maintained
for several days at restrictive temperature might have transiently expressed
cnox-2, allowing the head formation process to proceed, subsequently
losing cnox-2 expression and/or cnox-2+ cells.
Alternatively a nerve-independent regeneration process might have taken over
in these animals. Similarly, in cnox-2(-) hydra that had
ultimately regenerated their head, either cnox-2 silencing was
transient and cnox-2 expression reestablished at the time of head
formation, or a cnox-2-independent alternative mechanism was
activated. Recently, a similar correlation between the disappearance of
Dickkopf+ gland cells and the blockade in head regeneration in
sf-1 hydra was reported (Guder et
al., 2006), suggesting that gland cells, besides their immediate
cytoprotective effect (Chera et al.,
2006), also participate in the head regeneration process.
The nerve-dependence of regeneration across evolution
Comparative analysis of the regeneration processes in urodeles and hydra
imposed opposed views concerning their respective nerve-dependence: in
urodeles nerve-dependence is complete, as neurotrophic factors are required
for the proliferation of blastema cells
(Singer, 1974), whereas in
hydra, nerve-dependence is dispensable, as nerve-free animals regenerate
(Marcum and Campbell, 1978;
Sugiyama and Fujisawa, 1978).
However, in urodeles the nerve-dependence is linked to the preexisting
homeostatic conditions, as limbs that developed in the absence of nerves
(aneurogenic limbs) are able to regenerate in the absence of any neuronal
support (Brockes, 1987;
Tassava and Olsen-Winner,
2003), mimicking the nerve-free hydra situation. In such hydra,
the genetic program at work in epithelial cells is not yet known, but is
likely to be different from that at work in wild-type hydra
(Schaller et al., 1980;
Hornberger and Hassel, 1997)
(S.C., unpublished). In the head-regeneration deficient mutant
reg-16, removal of i-cells rescues head regeneration, highlighting
the morphogenetic consequences of a misregulation between i-cells and
myoepithelial cells (Sugiyama and Wanek,
1993). We propose that in the wild-type context, at the stage of
head formation, hydra makes use of cell proliferation and de novo
neurogenesis, both requiring cnox-2 activity. This sequence of events would be
the most efficient and the fastest way to achieve head regeneration. In the
absence of one of these components, i-cell proliferation, neuronal
differentiation and cnox-2 activation, other routes can be taken,
although much slower and much less efficient.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/134/6/1191/DC1
|
ACKNOWLEDGMENTS |
|---|
|
Footnotes |
|---|
|
REFERENCES |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Bode, H. R. (1992). Continuous conversion of neuron phenotype in hydra. Trends Genet. 8, 279-284.[Medline]
Bode, H. R. (1996). The interstitial cell lineage of hydra: a stem cell system that arose early in evolution. J. Cell Sci. 109,1155 -1164.[Medline]
Bosch, T. C. and David, C. N. (1984). Growth regulation in Hydra: relationship between epithelial cell cycle length and growth rate. Dev. Biol. 104,161 -171.[CrossRef][Medline]
Brockes, J. P. (1987). The nerve dependence of
amphibian limb regeneration. J. Exp. Biol.
132, 79-91.
Broun, M., Sokol, S. and Bode, H. R. (1999). Cngsc, a homologue of goosecoid, participates in the patterning of the head, and is expressed in the organizer region of Hydra. Development 126,5245 -5254.[Abstract]
Campbell, R. D. and David, C. N. (1974). Cell
cycle kinetics and development of Hydra attenuata. II. Interstitial cells.
J. Cell Sci. 16,349
-358.
Cartwright, P., Bowsher, J. and Buss, L. W.
(1999). Expression of a Hox gene, Cnox-2, and the division of
labor in a colonial hydroid. Proc. Natl. Acad. Sci.
USA 96,2183
-2186.
Chera, S., de Rosa, R., Miljkovic-Licina, M., Dobretz, K.,
Ghila, L., Kaloulis, K. and Galliot, B. (2006). Silencing of
the hydra serine protease inhibitor Kazal1 gene mimics the human Spink1
pancreatic phenotype. J. Cell Sci.
119,846
-857.
Chera, S., Kaloulis, K. and Galliot, B. (2007). The cAMP Response Element Binding Protein (CREB) as an integrative HUB selector in metazoans: clues from the hydra model system. Biosystems 87,191 -203.[CrossRef][Medline]
Engel, U., Ozbek, S., Streitwolf-Engel, R., Petri, B.,
Lottspeich, F., Holstein, T. W., Oezbek, S. and Engel, R.
(2002). Nowa, a novel protein with minicollagen Cys-rich domains,
is involved in nematocyst formation in Hydra. J. Cell
Sci. 115,3923
-3934.
Finnerty, J. R., Paulson, D., Burton, P., Pang, K. and Martindale, M. Q. (2003). Early evolution of a homeobox gene: the parahox gene Gsx in the Cnidaria and the Bilateria. Evol. Dev. 5,331 -345.[CrossRef][Medline]
Fujisawa, T. (1989). Role of interstitial cell migration in generating position-dependent patterns of nerve cell differentiation in Hydra. Dev. Biol. 133, 77-82.[CrossRef][Medline]
Fujisawa, T. (2003). Hydra regeneration and epitheliopeptides. Dev. Dyn. 226,182 -189.[CrossRef][Medline]
Fujisawa, T. and Sugiyama, T. (1978). Genetic analysis of developmental mechanisms in Hydra. IV. Characterization of a nematocyst-deficient strain. J. Cell Sci. 30,175 -185.[Abstract]
Fujisawa, T. and David, C. N. (1984). Loss of differentiating nematocytes induced by regeneration and wound healing in Hydra. J. Cell Sci. 68,243 -255.[Abstract]
Fujisawa, T., David, C. N. and Bosch, T. C. (1990). Transplantation stimulates interstitial cell migration in hydra. Dev. Biol. 138,509 -512.[CrossRef][Medline]
Galliot, B. and Miller, D. (2000). Origin of anterior patterning. How old is our head? Trends Genet. 16,1 -5.[Medline]
Galliot, B. and Schmid, V. (2002). Cnidarians as a model system for understanding evolution and regeneration. Int. J. Dev. Biol. 46,39 -48.[Medline]
Galliot, B., Miljkovic-Licina, M. and Chera, S. (2006). Hydra: a niche for cell and developmental plasticity. Semin. Cell Dev. Biol. 17,492 -502.[CrossRef][Medline]
Gauchat, D., Kreger, S., Holstein, T. and Galliot, B. (1998). prdl-a, a gene marker for hydra apical differentiation related to triploblastic paired-like head-specific genes. Development 125,1637 -1645.[Abstract]
Gauchat, D., Mazet, F., Berney, C., Schummer, M., Kreger, S.,
Pawlowski, J. and Galliot, B. (2000). Evolution of Antp-class
genes and differential expression of Hydra Hox/paraHox genes in anterior
patterning. Proc. Natl. Acad. Sci. USA
97,4493
-4498.
Gauchat, D., Escriva, H., Miljkovic-Licina, M., Chera, S., Langlois, M. C., Begue, A., Laudet, V. and Galliot, B. (2004). The orphan COUP-TF nuclear receptors are markers for neurogenesis from cnidarians to vertebrates. Dev. Biol. 275,104 -123.[CrossRef][Medline]
Guder, C., Pinho, S., Nacak, T. G., Schmidt, H. A., Hobmayer,
B., Niehrs, C. and Holstein, T. W. (2006). An ancient
Wnt-Dickkopf antagonism in Hydra. Development
133,901
-911.
Hayakawa, E., Fujisawa, C. and Fujisawa, T. (2004). Involvement of Hydra achaete-scute gene CnASH in the differentiation pathway of sensory neurons in the tentacles. Dev. Genes Evol. 214,486 -492.[Medline]
Hayward, D. C., Catmull, J., Reece-Hoyes, J. S., Berghammer, H., Dodd, H., Hann, S. J., Miller, D. J. and Ball, E. E. (2001). Gene structure and larval expression of cnox-2Am from the coral Acropora millepora. Dev. Genes Evol. 211, 10-19.[CrossRef][Medline]
Holstein, T. W. and David, C. N. (1990a). Cell cycle length, cell size, and proliferation rate in hydra stem cells. Dev. Biol. 142,392 -400.[CrossRef][Medline]
Holstein, T. W. and David, C. N. (1990b). Putative intermediates in the nerve cell differentiation pathway in hydra have properties of multipotent stem cells. Dev. Biol. 142,401 -405.[CrossRef][Medline]
Holstein, T., Schaller, C. H. and David, C. N. (1986). Nerve cell differentiation in Hydra requires two signals. Dev. Biol. 115,9 -17.
Holstein, T. W., Hobmayer, E. and David, C. N. (1991). Pattern of epithelial cell cycling in hydra. Dev. Biol. 148,602 -611.[CrossRef][Medline]
Holstein, T. W., Hobmayer, E. and Technau, U. (2003). Cnidarians: an evolutionarily conserved model system for regeneration? Dev. Dyn. 226,257 -267.[CrossRef][Medline]
Hornberger, M. R. and Hassel, M. (1997). Expression of HvRACK1, a member of the RACK1 subfamily of regulatory WD40 proteins in Hydra vulgaris, is coordinated between epithelial and interstitial cells in a position-dependent manner. Dev. Genes Evol. 206,435 -446.
Javois, L. C. and Frazier-Edwards, A. M. (1991). Simultaneous effects of head activator on the dynamics of apical and basal regeneration in Hydra vulgaris (formerly Hydra attenuata). Dev. Biol. 144,78 -85.[CrossRef][Medline]
Kaloulis, K., Chera, S., Hassel, M., Gauchat, D. and Galliot,
B. (2004). Reactivation of developmental programs: The
cAMP-response element-binding protein pathway is involved in hydra head
regeneration. Proc. Natl. Acad. Sci. USA
101,2363
-2368.
Lindgens, D., Holstein, T. W. and Technau, U.
(2004). Hyzic, the Hydra homolog of the zic/odd-paired gene, is
involved in the early specification of the sensory nematocytes.
Development 131,191
-201.
Littefield, C. L. (1991). Cell lineages in Hydra: isolation and characterization of an interstitial stem cell restricted to egg production in Hydra oligactis. Dev. Biol. 143,378 -388.[CrossRef][Medline]
MacWilliams, H. K. (1983). Hydra transplantation phenomena and the mechanism of Hydra head regeneration. II. Properties of the head activation. Dev. Biol. 96,239 -257.[CrossRef][Medline]
Marcum, B. A. and Campbell, R. D. (1978). Development of Hydra lacking nerve and interstitial cells. J. Cell Sci. 29,17 -33.[Abstract]
Marcum, B. A., Fujisawa, T. and Sugiyama, T. (1980). A mutant hydra strain (sf-1) containing temperature-sensitive interstitial cells. In Developmental and Cellular Biology of Coelenterates (ed. P. Tardent and R. Tardent), pp. 429-434. Amsterdam: Elsevier/North Holland.
Miljkovic-Licina, M., Gauchat, D. and Galliot, B. (2004). Neuronal evolution: analysis of regulatory genes in a first-evolved nervous system, the hydra nervous system. Biosystems 76,75 -87.[CrossRef][Medline]
Mitgutsch, C., Hauser, F. and Grimmelikhuijzen, C. J. (1999). Expression and developmental regulation of the Hydra-RFamide and Hydra-LWamide preprohormone genes in Hydra: evidence for transient phases of head formation. Dev. Biol. 207,189 -203.[CrossRef][Medline]
Nishimiya-Fujisawa, C. and Sugiyama, T. (1993). Genetic analysis of developmental mechanisms in hydra. XX. Cloning of interstitial stem cells restricted to the sperm differentiation pathway in Hydra magnipapillata. Dev. Biol. 157, 1-9.[CrossRef][Medline]
Otto, J. J. and Campbell, R. D. (1977). Budding in Hydra attenuata: bud stages and fate map. J. Exp. Zool. 200,417 -428.[CrossRef][Medline]
Schaller, H. C., Rau, T. and Bode, H. (1980). Epithelial cells in nerve-free hydra produce morphogenetic substances. Nature 283,589 -591.[CrossRef][Medline]
Schaller, H. C., Hoffmeister, S. A. and Dubel, S. (1989). Role of the neuropeptide head activator for growth and development in hydra and mammals. Development 107,99 -107.
Schierwater, B., Dellaporta, S. and DeSalle, R. (2002). Is the evolution of Cnox-2 Hox/ParaHox genes "multicolored" and "polygenealogical?" Mol. Phylogenet. Evol. 24,374 -378.[CrossRef][Medline]
Schummer, M., Scheurlen, I., Schaller, C. and Galliot, B. (1992). HOM/HOX homeobox genes are present in hydra (Chlorohydra viridissima) and are differentially expressed during regeneration. EMBO J. 11,1815 -1823.[Medline]
Seipel, K. and Schmid, V. (2005). Evolution of striated muscle: jellyfish and the origin of triploblasty. Dev. Biol. 282,14 -26.[CrossRef][Medline]
Shenk, M. A., Bode, H. R. and Steele, R. E. (1993a). Expression of Cnox-2, a HOM/HOX homeobox gene in hydra, is correlated with axial pattern formation. Development 117,657 -667.[Abstract]
Shenk, M. A., Gee, L., Steele, R. E. and Bode, H. R. (1993b). Expression of Cnox-2, a HOM/HOX gene, is suppressed during head formation in hydra. Dev. Biol. 160,108 -118.[CrossRef][Medline]
Siddiqui, S. S., Aamodt, E., Rastinejad, F. and Culotti, J. (1989). Anti-tubulin monoclonal antibodies that bind to specific neurons in Caenorhabditis elegans. J. Neurosci. 9,2963 -2972.[Abstract]
Singer, M. (1974). Neurotrophic control of limb regeneration in the newt. Ann. N.Y. Acad. Sci. 228,308 -322.[Medline]
Sugiyama, T. and Fujisawa, T. (1978). Genetic analysis of developmental mechanisms in Hydra. II. Isolation and characterization of an interstitial cell-deficient strain. J. Cell Sci. 29,35 -52.[Abstract]
Sugiyama, T. and Wanek, N. (1993). Genetic analysis of developmental mechanisms in hydra. XXI. Enhancement of regeneration in a regeneration-deficient mutant strain by the elimination of the interstitial cell lineage. Dev. Biol. 160, 64-72.[CrossRef][Medline]
Tassava, R. A. and Olsen-Winner, C. L. (2003). Responses to amputation of denervated ambystoma limbs containing aneurogenic limb grafts. J. Exp. Zoolog. A Comp. Exp. Biol. 297, 64-79.[Medline]
Tavernarakis, N., Wang, S. L., Dorovkov, M., Ryazanov, A. and Driscoll, M. (2000). Heritable and inducible genetic interference by double-stranded RNA encoded by transgenes. Nat. Genet. 24,180 -183.[CrossRef][Medline]
Technau, U. and Holstein, T. W. (1996). Phenotypic maturation of neurons and continuous precursor migration in the formation of the peduncle nerve net in Hydra. Dev. Biol. 177,599 -615.[CrossRef][Medline]
Teragawa, C. K. and Bode, H. R. (1991). A head signal influences apical migration of interstitial cells in Hydra vulgaris. Dev. Biol. 147,293 -302.[CrossRef][Medline]
Timmons, L. and Fire, A. (1998). Specific interference by ingested dsRNA. Nature 395, 854.[CrossRef][Medline]
Toresson, H. and Campbell, K. (2001). A role
for Gsh1 in the developing striatum and olfactory bulb of Gsh2 mutant mice.
Development 128,4769
-4780.
Venugopal, G. and David, C. N. (1981). Nerve commitment in Hydra. I. Role of morphogenetic signals. Dev. Biol. 83,353 -360.[CrossRef][Medline]
Weissman, I. L., Anderson, D. J. and Gage, F. (2001). Stem and progenitor cells: origins, phenotypes, lineage commitments, and transdifferentiations. Annu. Rev. Cell Dev. Biol. 17,387 -403.[CrossRef][Medline]
Westfall, J. A. (1996). Ultrastructure of synapses in the first-evolved nervous systems. J. Neurocytol. 25,735 -746.[CrossRef][Medline]
Yanze, N., Spring, J., Schmidli, C. and Schmid, V. (2001). Conservation of Hox/ParaHox-related genes in the early development of a cnidarian. Dev. Biol. 236, 89-98.[CrossRef][Medline]
Yun, K., Garel, S., Fischman, S. and Rubenstein, J. L. (2003). Patterning of the lateral ganglionic eminence by the Gsh1 and Gsh2 homeobox genes regulates striatal and olfactory bulb histogenesis and the growth of axons through the basal ganglia. J. Comp. Neurol. 461,151 -165.[CrossRef][Medline]
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati 
Twitter What's this?
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
