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First published online January 13, 2009
doi: 10.1242/10.1242/dev.026955
Division of Basic Sciences, Fred Hutchinson Cancer Research Center, 1100 Fairview Ave N., Seattle, WA 98109, USA.
* Author for correspondence (e-mail: bedgar{at}fhcrc.org)
Accepted 24 November 2008
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SUMMARY |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Key words: EGFR, Adult midgut progenitor (AMP), Intestinal stem cell (ISC)
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INTRODUCTION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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). Similarly, multiple signaling pathways, including the EGF,
Wingless (Wnt), Dpp (TGFβ), Notch and Hedgehog pathways, are involved in
the embryonic development of the Drosophila midgut and mammalian
intestine (Sancho et al.,
2004). In both systems, cross-talk between mesodermal cells and
endoderm-derived epithelial cells in the gut primordium plays important roles
during embryonic gut development
(Stainier, 2005;
Szuts et al., 1998).
Starting from embryonic development stage 11, the Drosophila
midgut epithelium consists of two distinct cell populations: differentiating
midgut epithelial cells (larval enterocytes, ECs) and undifferentiated adult
midgut progenitors (AMPs, also referred to as midgut histoblast islets or
midgut imaginal islets) (Hartenstein et
al., 1992). In Drosophila embryos, AMPs can be marked by
expression of asense or by one of several lacZ- or
Gal4-expressing enhancer-trap insertions
(Brand et al., 1993;
Hartenstein et al., 1992;
Hartenstein and Jan, 1992).
AMPs first appear as spindle-shaped cells localized to the apical surface of
the midgut epithelium, but later migrate to the basal surface of the
epithelium where they remain throughout larval development
(Hartenstein and Jan, 1992;
Technau and Campos-Ortega,
1986). Notch signaling has been shown to be involved in the
development of Drosophila AMPs. In Notch mutant embryos, the
number of AMPs in the midgut rudiment is strongly increased at the expense of
differentiated larval ECs (Hartenstein et
al., 1992). During larval development, the ECs grow in both size
and ploidy by undergoing several endocycles, reaching 64C (DNA content) by the
wandering L3 stage (Lamb,
1982). The AMPs remain diploid throughout larval development and
appear as scattered islets of cells (hence the term `midgut histoblast
islets') in late-stage larval midguts. During pupal development, the ECs
histolyze and a new adult midgut epithelium forms from the AMPs
(Bender et al., 1997;
Jiang et al., 1997;
Li and White, 2003). Similar
midgut progenitor cells have also been found in other insect species
(Corley and Lavine, 2006).
Recently, the adult Drosophila midgut has been shown to undergo
dynamic self-renewal, a process similar to that found in the mammalian
intestine/colon. Fly and mammalian gut homeostasis are both powered by
intestinal stem cells (ISCs), and Notch signaling plays similar roles in
regulating their differentiation into mature gut cells
(Fre et al., 2005;
Micchelli and Perrimon, 2006;
Ohlstein and Spradling, 2006;
Ohlstein and Spradling, 2007;
van Es et al., 2005). Thus,
the Drosophila midgut may serve as a model to study gut homeostasis
and the development of cancers, such as colorectal carcinoma, that are
directly associated with this dynamic process in humans.
Here we describe the development of the AMPs in Drosophila larvae and pupae. We discovered that Drosophila AMPs divide extensively throughout larval development, and that their proliferation can be separated into two distinct phases. During early larval stages, the AMPs divide and disperse to form islets throughout the midgut, but during late larval development the dividing AMPs are contained within these islets. Furthermore, our study revealed that Drosophila EGFR signaling is both necessary and sufficient to induce the proliferation of AMPs during larval development.
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MATERIALS AND METHODS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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TOP, UAS-SEM, UAS-RafDN, UAS-Mkp3, UAS-sSpi,
UAS-sKrn, UAS-Krn, UAS-grk
TC and
UAS-Vn1.2. UAS-RNAi transgenes were obtained from the Bloomington
Stock Center (Bloomington, IN, USA), the National Institute of Genetics Fly
Stock Center (NIG, Japan) or the Vienna Drosophila RNAi Center (VDRC,
Austria). According to information from NIG and VDRC, all the RNAi lines used
are specific to the genes targeted (NIG,
http://www.shigen.nig.ac.jp/fly/nigfly/index.jsp;
VDRC,
http://stockcenter.vdrc.at/control/main).
Mutants
FRT42D Egfrf1, FRT42D Egfr[CO], FRT82B
Ras1c40b, spiA14 FRT40A,
FRT42D shot[65-2], FRT42D shot[V104], vnP1749
FRT80B, rhodel1 FRT80B, Krn27-3-4, vnP1749,
vn
7, stet871 and
ru1 were used (see FlyBase for further information:
http://flybase.org).
Gal4/lacZ reporters
esgGal4NP7397, spiGal4NP0261,
MyoIAGal4NP0001 (NIG, Japan), rholacZAA69,
rholacZX81, howGal424B and
esglacZK00606 were used (Bloomington Stock Center).
Lineage analysis
MARCM lineage analysis
Newly hatched first instar [24 hours after egg deposition (AED)] or
mid-third instar (96 hours AED) larvae of the correct genotype were heat
shocked for 45 minutes at 37°C. The midguts were then dissected from
wandering L3 larvae (120 hours AED) and analyzed.
Flp/Gal4 lineage analysis
Newly hatched first instar larvae (24 hours AED) of the correct genotype
were heat shocked for 20 minutes at 37°C to induce clones and then
dissected at various developmental stages and analyzed.
Enhancer traps
P-element enhancer traps with midgut expression were obtained from
several sources, including FlyView (University of Münster, Germany;
http://flyview.uni-muenster.de)
and GETDB (Gal4 Enhancer-Trap Insertion Database, NIG, Japan). We identified a
number of enhancer traps showing reporter expression specifically in the AMPs,
including one insertion in spi (NP0261) and several insertions in
esg (NP0726, 7397 and 7399). esgGal4NP7397-driven
GFP expression was used to mark the AMPs. We also identified an enhancer trap
in brush border Myosin IA (MyoIAGal4, NP0001) that drives
GFP expression specifically in midgut ECs
(Morgan et al., 1995).
Ectopic gene expression
We generated inducible AMP-, EC- and VM-specific expression systems
(esgGal4ts, MyoIAGal4ts and
howGal4ts) by combining esgGal4NP7397,
MyoIAGal4NP0001 or howGal424B
(Hartenstein and Jan, 1992)
with ubiquitously expressed temperature-sensitive alleles of the Gal4
inhibitor, Gal80 (tubGal80ts; Bloomington Stock Center)
and UAS-GFP.
Quantification of AMP clusters
We counted AMPs or AMP clusters marked by esgGal4-driven GFP
expression throughout the entire midgut during larval and pupal development.
UAS-GFP, UAS-sSpi, UAS-sKrn or UAS-Krn were induced in the
AMPs starting from first instar larvae (24 hours AED) using the
esgGal4ts system and the midguts were dissected from
wandering L3 larvae and the number of AMP clusters counted.
Generation of mutant AMP clones
Clones of AMPs homozygous for Egfrf1, Egfr[CO],
Ras1c40b, spiA14,
vnP1749, rhodel1, shot[V104] or
shot[65-2] were generated using the MARCM system
(Lee and Luo, 2001). First
instar larvae (24 hours AED) of the correct genotype were heat shocked for 45
minutes at 37°C to induce clones. Larvae were then dissected at 120 hours
AED. The number of GFP-positive clusters in each clone was quantified; in most
cases, clones from at least ten midguts were counted.
RNA in situ hybridization and immunofluorescence
RNA in situ hybridization was performed as described
(O'Neill and Bier, 1994).
Rabbit anti-dpERK (Cell Signaling) was used to detect MAP kinase activity in
the midgut. Anti-Delta and anti-Prospero were obtained from the Developmental
Studies Hybridoma Bank and used to mark ISCs and enteroendocrine cells in the
midgut. Rabbit anti-β-galactosidase (Cappel) was used to identify the
esg-positive cells in an esglacZ background. Rabbit
anti-phospho-histone H3 (PH3, Upstate) was used to identify dividing
cells.
Quantitative real-time PCR (qRT-PCR)
We used qRT-PCR to quantify levels of vn mRNA from midgut cDNA.
For mRpL30 (reference gene) primers see Buttitta et al.
(Buttitta et al., 2007); for
vn: vn 5' primer,
5'-TCACACATTTAGTGGTGGAAG-3'; vn 3' primer,
5'-TCACACATTTAGTGGTGGAAG-3'. The relative expression of
vn was analyzed on the Bio-Rad iQ5 system.
Sectioning
Wandering L3 midguts were dissected in PBS and fixed in half-strength
Karnovsky's fixative. Following dehydration, the tissues were embedded in Epon
and sectioned at 1 µm. The sections were stained with Toluidine Blue.
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RESULTS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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). When combined with UAS-GFP, esgGal4
enhancer trap NP7397 drove GFP expression specifically in the larval AMPs
(Fig. 1). Similar esg
enhancer traps have been used to mark the adult ISCs and their daughter
enteroblasts (Micchelli and Perrimon,
2006).
GFP expression driven by esgGal4 was detected in the AMPs
scattered throughout the midgut of newly hatched larvae (24 hours AED)
(Fig. 1A, arrows). AMPs
appeared as small diploid cells, and were easily distinguishable from the
large polyploid midgut enterocytes (ECs). The number of GFP-positive AMPs
increased during early larval development (24-72 hours AED)
(Fig. 1A,B); however, they
remained dispersed. Cell contacts between paired AMPs were readily observed in
the early larval midgut (Fig.
1B, inset) and are likely to represent two daughter AMPs from the
previous division migrating away from each other. By mid-third instar (96
hours AED), AMPs formed discrete 2- to 3-cell clusters
(Fig. 1C), suggesting that they
proliferate within individual islets instead of migrating away from each
other. The AMPs continued to proliferate within these clusters
(Fig. 1D), undergoing several
rounds of rapid proliferation to enlarge each cluster to 8-30 cells by the
onset of metamorphosis [0 hours after pupae formation (APF), 
130 hours
AED] (Fig. 1E).
These results do not support the idea that Drosophila AMPs are
quiescent during larval development
(Bodenstein, 1994). Instead, we
observed that the AMPs proliferate extensively during larval development,
resulting in large increases in both the number (early larval stages) and size
(late larval stages) of the AMP clusters. To further document this process, we
analyzed AMP lineages by positively marking individual AMPs with GFP using the
MARCM system (Lee and Luo,
2001). When clones were induced in first or second instar (24-48
hours AED), they all contained multiple AMP clusters by the wandering L3 stage
(120 hours AED), and all cells in any GFP-positive cluster were GFP-positive
(Fig. 2A-A'';
Fig. 4A). However, when clones
were induced in mid-third instar (96 hours AED), clusters mosaic for GFP were
observed by the wandering L3 stage (120 hours AED)
(Fig. 2B-B''). These
results confirmed that the AMPs switch to proliferating within islets to form
clusters by mid-third instar. To quantify the number of divisions during the
early proliferative phase, we counted the number of the marked AMP clusters
encompassed by each clone. When induced in the newly hatched first instar
larvae (24 hours AED), the clones contained, on average, 7.5 GFP-positive
clusters at the wandering L3 stage (120 hours AED) (see Table S1 in the
supplementary material). This suggests that the AMPs divide about four times
during the early larval stages (note that only half of all the clusters
generated by each AMP were marked in the MARCM system). Since no mosaic AMP
clusters were found in the late larval midgut when clones were induced at
first or second instar, we propose that the majority, if not all, of the early
larval AMPs disperse after each cell division. We then counted the number of
cells in each cluster at white prepupa formation (0 hours APF), when most of
the AMP clusters have stopped proliferating. Each AMP cluster contained 8 to
greater than 30 cells. This indicates that the AMPs divide an additional three
to five times within a cluster, after the clusters are established. In total,
the AMPs appear to divide seven to ten times throughout larval
development.
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). The midgut continued to contract and shorten, and by 12
hours APF it became a sac-like structure containing histolyzing larval
epithelium inside the newly formed midgut
(Juhasz and Neufeld, 2008).
Visceral muscles undergo a process termed `de-differentiation', in which the
muscle fibers histolyze; however, the muscle cells themselves do not die and
will redifferentiate to form the adult midgut VM during late metamorphosis
(Klapper, 2000). During early
metamorphosis (0-24 hours APF), the majority of cells in the newly formed
midgut epithelium gradually lost esgGal4-driven GFP expression, with
the exception of a few scattered cells that maintained strong GFP expression
(Fig. 1F-I, asterisks). By 24
hours APF, these GFP-positive cells became basally localized in the new
epithelium (Fig. 1J,
asterisks). As described below, we believe that these cells are the future
adult midgut ISCs (see Discussion).
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;
Ohlstein and Spradling, 2006),
and that this transition occurs in the pupa. This was further supported by our
observation that AMP clones induced in the larva persisted in the adult midgut
for at least 2 months (data not shown).
EGFR signaling stimulates AMP proliferation
Using the esgGal4ts system, we manipulated the activity
of several known Drosophila signaling pathways specifically in the
AMPs. Our tests included Wingless, Dpp, Hedgehog, Notch and EGFR signaling
components (see Table S2 in the supplementary material). Activation of
EGFR/RAS/MAPK signaling in the AMPs was able to drive their overproliferation
during larval development. Compared with control midguts
(Fig. 3A), in which the AMPs
appeared as 2- to 3-cell clusters by mid-third instar (96 hours AED), the
induction of activated Ras (Ras oncogene at 85D - FlyBase)
(RasV12) in the AMPs led to the formation of much larger
AMP clusters (Fig. 3B-D).
Ectopic expression of RasV12 in the AMPs throughout larval
development resulted, by the wandering stage (120 hours AED), in a midgut
comprising mostly esg-positive AMPs
(Fig. 3F) in which the
intestinal lumen was occluded. By contrast, wild-type AMPs appeared as basally
localized cell clusters in the midgut epithelium
(Fig. 3E). The following
evidence suggested that EGFR signaling promoted AMP proliferation through
activating the MAPK pathway. First, induction of
RasV12S35, which preferentially activates the MAPK
pathway, drove similar ectopic proliferation of the AMPs as did
RasV12 (see Table S2 in the supplementary material),
whereas expression of RasV12G37, which preferentially
activates the Phosphotidylinositol 3 kinase (PI3K) or Ral guanine nucleotide
exchange factor 2 (RalGDS) pathway (Karim
and Rubin, 1998; Prober and
Edgar, 2002), had little effect on their proliferation (see Table
S2 in the supplementary material). Second, ectopic expression of
Dp110 (Pi3K92E - FlyBase; PI3K) had no detectable effect on
AMP proliferation (see Table S2 in the supplementary material). Third,
increased proliferation of the AMPs was observed when activated Egfr
(TOP) (Queenan et al.,
1997), gain-of-function Raf (Rafgof)
(Brand and Perrimon, 1994) or
activated MAPK [sevenmaker (sem); rolled - FlyBase]
(Martin-Blanco, 1998) was
induced in these cells (see Table S2 in the supplementary material). Fourth,
expression of a dominant-negative form of Raf
(RafDN) (Roch et al.,
1998) together with RasV12 gave a phenotype
similar to that of RafDN alone (see Fig. S2E,F in the
supplementary material), and thus Raf is epistatic to Ras in
regulating AMP proliferation. Fifth, expression of Mkp3, a negative
regulator of MAPK (Rintelen et al.,
2003), did not affect AMP proliferation (see Fig. S2G in the
supplementary material). Interestingly, however, Mkp3 did
significantly suppress the AMP overproliferation phenotype induced by
RasV12 expression (see Fig. S2H in the supplementary
material; compare with Fig.
3D).
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In further tests we generated AMP clones defective in EGFR signaling using
the MARCM system. AMPs mutant for Egfr (Egfr[CO])
or Ras (Ras1c40b) did
not proliferate during larval development
(Fig. 4; see Table S1 in the
supplementary material). Instead of forming multiple GFP-positive clusters as
in controls (Fig. 4A), these
mutant clones appeared as single GFP-positive cells
(Fig. 4B,C). We conclude that
EGFR/RAS/MAPK signaling is required for AMP proliferation during both early
and late larval development.
Drosophila MAPK is activated in the AMPs
To examine whether downstream components of EGFR signaling are activated in
the AMPs, we stained the larval midgut with antibodies against
diphospho-extracellular signal-regulated kinase (dpERK; Rolled - FlyBase), the
level of which is a direct measurement of the activated form of
Drosophila MAPK (Gabay et al.,
1997). dpERK staining was indeed detected in the AMP clusters, but
not in the larval gut epithelial cells
(Fig. 5A-A''), indicating
activation of MAPK in these cells. This result is consistent with our genetic
results and supports the notion that EGFR signaling regulates AMP
proliferation through activating the MAPK pathway.
Next, we examined the expression patterns of several EGFR ligands in the larval midgut. We identified a Gal4 enhancer trap in spi (NP0261, see Materials and methods) that drove UAS-GFP expression specifically in the AMPs (Fig. 5B-B''). RNA in situ hybridization confirmed that spi was specifically expressed in the AMP clusters (Fig. 5D,D'). Krn was also specifically expressed in the AMPs as shown by RNA in situ hybridization (Fig. 5E,E').
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TC), a strong EGFR ligand,
the function of which is believed to be exclusively in female oogenesis
(Nilson and Schupbach, 1999),
did not affect the proliferation of the AMPs (see Table S2 in the
supplementary material). However, induction of activated (secreted)
spitz or Keren (sSpi or sKrn), two other
strong EGFR activating ligands (Reich and
Shilo, 2002; Schweitzer et
al., 1995), promoted extensive overproliferation of the AMPs
(Fig. 6A-C; see Table S2 in the
supplementary material). Furthermore, induction of wild-type Krn,
which requires cleavage by rhomboid family proteases to become fully active,
similarly promoted AMP proliferation (Fig.
6D-D'; see Table S2 in the supplementary material). In
addition, induction of sSpi or sKrn limited the dispersal of
the AMPs, thus reducing the number of the clusters in the wandering L3 larval
midguts (Fig. 6E). Induction of
the weak EGFR ligand vein (vn)
(Schnepp et al., 1998) with
esgGal4ts, however, had little effect on AMP proliferation
(see Table S2 in the supplementary material). To determine whether spi or Krn are required for AMP proliferation, we downregulated the levels of these EGFR ligands in the AMPs by RNAi. Ectopic expression of UAS-RNAi directed at spi and/or Krn using esgGal4 had no effect on AMP proliferation (see Table S2 in the supplementary material). Consistent with this, the proliferation of the AMPs in Krn27-3-4 (null allele) mutant larvae was normal (see Table S1 in the supplementary material). The same was also found for spiA14 mutant AMP clones generated in a Krn27-3-4 homozygous mutant background (see Table S1 in the supplementary material). The proliferation of mutant AMPs lacking rhomboid (rhodel1, null allele) or spi (spiA14, null allele) function, generated using the MARCM system, was also normal (see Table S1 in the supplementary material). We examined lacZ expression from two rholacZ reporters in the larval midgut (rhoAA69 and rhoX81) and found that neither were expressed in the AMPs. Since the Drosophila genome encodes multiple rhomboid-like genes, we also generated MARCM clones in the mutant background of rho-2 (stet871) and rho-3 (ru1). These mutant clones also contained normal numbers of AMP clusters (see Table S1 in the supplementary material). These results suggest that either multiple, redundant rhomboid-like genes are utilized in the AMPs, or (less likely) that rhomboid-like function is dispensable in the larval midgut. Furthermore, we conclude that spi and Krn are likely to be dispensable for AMP proliferation (see Discussion).
Surprisingly, in several vn mutants (vnP1749/P1749,
vn7/7
and vnP1749/7), few
AMP clusters were found in the late larval midgut
(Fig. 7B,C), whereas the AMPs
in wild-type controls formed many large clusters
(Fig. 7A). This suggests that
vn is required for normal AMP development. To further study the
function of vn in AMP development, we carried out lineage analysis of
the AMPs in the vn mutant animals. We induced GFP-marked AMP clones
in first instar larvae using the Flp/Gal4 system. Compared with
control midguts, which contained on average 15.2 marked AMP clusters per clone
(n=50 clones) (see Fig. S3A-A'' in the supplementary material),
we consistently observed only a single GFP-positive AMP cluster in the midguts
of weak vn mutants
(vn7/P1749; animals of this
genotype are not developmentally delayed during larval development and most
die as pharate adults) (see Fig. S3B-B'' in the supplementary material).
Furthermore, we counted the number of esg-positive cells (marked by
esglacZ) in newly hatched larval midguts. Control larval midguts
(vnP1749/+) contained on average 121 AMPs per
gut, whereas there were on average 137 AMPs per midgut in
vn7/P1749
mutants (ten midguts for each genotype were scored). This indicates that the
reduction in the number of AMP clusters in the late larval midgut of
vn7/P1749 mutants was not due
to the production of fewer AMPs during embryogenesis. Taken together, these
results suggest that the proliferation of AMPs during the early larval stages
is completely inhibited in vn mutants. However, the size of the few
remaining AMP clusters in the
vn7/P1749 mutant midguts was
relatively normal (see Fig. S3B-B'' in the supplementary material),
suggesting that the late phase of AMP proliferation is largely unaffected in
vn mutants. We speculate that the reason vn becomes
dispensable for AMP proliferation during late larval development is that
Krn and spi expression in the AMPs supplies a redundant
function.
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). The Drosophila midgut VM comprises an outer layer
of 21 longitudinal muscle strips and an inner layer of circular muscle that
forms four distinctive rows (Klapper,
2000), as revealed by a muscle-specific driver,
howGal424B, which drives UAS-GFP expression in
both types of VM cells (Fig.
5C') (Brand and Perrimon,
1993). vnlacZ expression was stronger in the inner,
circular VM cells than in the outer, longitudinal muscle
(Fig. 5C). To test the importance of Vn signaling from VM, we specifically depleted vn from VM cells by expressing UAS-Vn RNAi using an inducible muscle-specific driver, howGal4ts. Induction of vn RNAi throughout larval development (24-120 hours AED) resulted in late larval midguts with very few AMP clusters, as in vn mutants (Fig. 7D-D''). However, induction of vn RNAi starting at early third instar (72 hours AED) had no effect on the AMPs (Fig. 7E-E''). Furthermore, induction of UAS-Vn RNAi in the AMPs or in the larval ECs using the esgGal4ts or MyoIAGal4ts (MyoIA is also known as Myo31DF - FlyBase) drivers had no effect on AMP proliferation (see Fig. S4A,B in the supplementary material), suggesting that the principal source of Vn is VM. This was confirmed by quantitative real-time PCR showing that induction of UAS-Vn RNAi in VM significantly reduced vn mRNA levels in whole midguts, whereas induction of vn RNAi in ECs or AMPs did not (Fig. 7G). In further tests, we attempted to rescue the AMP phenotype of vnP1749 mutants by expressing UAS-Vn in AMPs, ECs or VM, using the esgGal4ts, MyoIAGal4ts or howGal4ts drivers. Induction of UAS-Vn in the AMPs or VM completely rescued the phenotype of vnP1749 mutants (Fig. 7F-F''; see Fig. S5A-A'' in the supplementary material). Induction of vn in the larval ECs, which constitute the bulk of the midgut mass, not only rescued the proliferative defects of the AMPs, but also caused ectopic AMP proliferation (see Fig. S5B-B'' in the supplementary material). We conclude that Vn, expressed in VM, is the principal mitogen for AMPs during early larval development. Later, autocrine Spi and Krn might complement this function.
This scenario, in which VM-derived Vn activates EGFR signaling in AMPs, is
reminiscent of the role of Vn in muscle/tendon development during
embryogenesis. In this case, muscle-derived Vn is specifically concentrated on
tendon cells and activates EGFR there
(Strumpf and Volk, 1998). The
concentration of Vn is highly dependent upon the activity of the short
stop (shot, also called kakapo) gene in the tendon
cells (Strumpf and Volk,
1998). We tested whether shot is also required for
VM-derived Vn to activate EGFR signaling in the AMPs by quantifying AMP
clusters in shot mutant MARCM clones. These clones all contained
normal numbers of AMP clusters (see Table S1 in the supplementary material),
and thus the role of shot in transducing the Vn signal is
uncertain.
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DISCUSSION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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). This is the
case for several other larval progenitor/imaginal cell types, such as the
abdominal histoblasts and cells in the salivary gland, foregut and hindgut
imaginal rings (Bodenstein,
1994). More recent studies have suggested that AMP proliferation
might precede the onset of metamorphosis
(Hall and Thummel, 1998;
Jiang et al., 1997;
Li and White, 2003). However,
these studies did not report the extensive proliferation of the AMPs that we
describe here, and failed to recognize the early larval proliferative phase
when the AMPs divide and disperse (Figs
1 and
8). The extensive proliferation
of the AMPs is similar to that of the larval imaginal disc cells, which also
proliferate throughout larval development, dividing about ten times. Lineage analysis revealed that the proliferation of the Drosophila AMPs occurs in two distinct phases (Fig. 8). In early larvae, the AMPs divide and disperse throughout the midgut to form individual islets. During later larval development, the AMPs continue to divide but do so within these islets, forming large cell clusters. We speculate that in the early larva, secretion of Vn from the midgut visceral muscle (VM) cells results in low-level activation of EGFR signaling in the AMPs, which is sufficient for their proliferation and might also promote their dispersal. We did not observe any proliferation defects in AMPs defective in shot function, suggesting that the mechanism of EGFR activation used by tendon cells during muscle/tendon development is probably not the same as in the larval midgut. Specifically, it is unlikely that the Shot-mediated concentration of Vn on AMPs activates EGFR signaling in the AMPs during early larval development. Consistent with this, we only observed dpERK staining in AMP clusters (Fig. 5A-A'') and not in the isolated AMPs present at early larval stages (24-72 hours AED; data not shown).
The mechanisms that regulate the transition between these two proliferation
phases remain unclear. We observed fewer AMP clusters when sSpi,
sKrn, TOP (activated Egfr) or
RasV12 were induced in the AMPs starting from early larval
stages (Fig. 6E; see Table S2
in the supplementary material), suggesting that EGFR signaling, in addition to
its crucial role as an AMP mitogen, might also play a role in AMP cluster
formation. In the late larval midgut (96-120 hours AED), high-level EGFR
activation, resulting from expression of spi and Krn in the
AMPs themselves, might not only promote AMP proliferation, but might also
suppress AMP dispersal and thus promote formation of the AMP clusters. How the
timing and location of Spi- or Krn-mediated EGFR activation are regulated
during larval development is also unclear. We note, however, that the
pro-ligand form of Krn acted similarly to sKrn
(Fig. 6), and that we failed to
uncover any functions for the Rho-like gene products that regulate Spi and Krn
function by proteolytic cleavage in other tissues (see Tables S1 and S2 in the
supplementary material). This suggests that the localized expression of these
ligands in the AMP clusters might be the critical parameter that controls
their effects. Consistent with this, Rho-independent cleavage and function of
Krn have been documented (Reich and Shilo,
2002).
|
).
RasV12-expressing clones generated in the wing imaginal
disc are round (Prober and Edgar,
2002), much like the AMP clusters described here, owing to
increased adhesive junctions. In developing Drosophila trachea, EGFR
activity upregulates shg expression to maintain epithelial integrity
in the elongating tracheal tubes (Cela and
Llimargas, 2006). In the eye, EGFR activity leads to increased
levels of Shg and adhesion between photoreceptors
(Brown et al., 2006;
Mirkovic and Mlodzik, 2006).
Given these precedents, it seems reasonable to suggest that high-level EGFR
activity in the AMP islets upregulates Shg and promotes the homotypic adhesion
of the AMPs. Alternatively, changes in the differentiated cells of the midgut
epithelium might promote AMP clustering. In either case, the dispersal of
early AMPs and subsequent formation of late AMP clusters facilitate the
formation of the adult midgut epithelium during metamorphosis.
AMPs give rise to adult intestinal stem cells during metamorphosis
Our study confirms previous reports that Drosophila AMPs replace
larval midgut epithelial cells to form the adult midgut epithelium during
metamorphosis (Figs 1 and
8)
(Bender et al., 1997;
Jiang et al., 1997;
Li and White, 2003).
Furthermore, we show that the majority of AMPs lose esgGal4-driven
GFP expression as they differentiate to form the new adult midgut epithelium
(Fig. 1F-J). These cells lacked
Prospero, which marks enteroendocrine cells in both the larval and adult
midgut (Micchelli and Perrimon,
2006; Ohlstein and Spradling,
2006). They went through several rounds of endoreplication during
late pupal development (not shown), and thus probably all differentiated into
adult enterocytes (ECs). During early metamorphosis, some cells in the new
midgut epithelium remained small and diploid and maintained strong
esgGal4 expression (Fig.
1I,J; Fig. 8). For
several reasons, we believe that these esg-positive cells are the
future adult intestinal stem cells (ISCs). First, esgGal4 expression
marks AMPs, including adult ISCs and enteroblasts
(Micchelli and Perrimon,
2006). Second, mitoses in the adult midgut are only observed in
ISCs (Micchelli and Perrimon,
2006; Ohlstein and Spradling,
2006), and we observed mitoses only in the esg-positive
cells during metamorphosis (see Fig. S1 in the supplementary material). Third,
esg-positive cells migrated to the basal side of the midgut
epithelium (Fig. 1J), the
location of adult ISCs (Micchelli and
Perrimon, 2006; Ohlstein and
Spradling, 2006). Fourth, AMP clones generated during early larval
development contained just a few esg-positive cells when the new
adult midgut first formed (24 hours APF) (see Fig. S1C-C'' in the
supplementary material), but when such clones were scored in newly eclosed
adults, they contained large numbers of ECs, as well as cells positive for the
enteroendocrine marker Prospero and the ISC marker Delta
(Fig. 2D,E). This suggests that
a small fraction of AMPs differentiate into adult ISCs. However,
esg-positive cells in the new pupal midgut lacked Delta expression
until eclosion (Fig.
2E-E'''; data not shown), suggesting that they are probably
not mature adult ISCs.
|
) and
ISCs in adult midgut homeostasis
(Micchelli and Perrimon, 2006;
Ohlstein and Spradling, 2006;
Ohlstein and Spradling, 2007),
we expect that Notch might also function to specify adult ISCs during
metamorphosis.
Implications for EGFR/RAS signaling in insect midgut development
EGFR signaling is both required and sufficient to promote AMP proliferation
(Figs 3,
4,
6 and
7; see Fig. S2 in the
supplementary material). Hyperactivation of EGFR signaling, such as by
expression of activated Ras (RasV12), promoted
massive AMP overproliferation and generated hyperplastic midguts that were
clearly dysfunctional (Fig.
3F). On the other hand, inhibiting EGFR/RAS/MAPK signaling
dramatically reduced AMP proliferation
(Fig. 4; see Fig. S2 in the
supplementary material). Furthermore, the ability of EGFR signaling to induce
ectopic AMP proliferation is almost unique. With the exception of larval
hemocytes (Zettervall et al.,
2004), activated EGFR signaling does not promote cell
proliferation in the imaginal discs, salivary gland imaginal rings, abdominal
histoblasts, foregut and hindgut imaginal rings. This suggests that the
regulation of AMP proliferation is different from that in other imaginal
cells.
Regulation of AMP proliferation by non-epithelial muscle cells
Despite the obvious differences between adult ISCs and their larval
progenitors, the AMPs, there are also similarities. First, when the new adult
midgut epithelium forms, larval AMPs give rise to the new adult midgut
including the adult ISCs. Many genes, such as esg, that are
specifically expressed in the larval AMPs are also expressed in the adult ISCs
(our unpublished data). Second, the structure of the midgut epithelium with
basal AMPs or ISCs is similar in larval and adult stages. Third, vn
expression in larval VM persists in the adult midgut (our unpublished data),
suggesting that Vn from the adult VM might also regulate the ISCs.
In two Drosophila stem cell models, the testis and ovary, stem
cells reside in special niches comprising other supporting cell types. These
niches maintain the stem cells and provide them with proliferative cues
(Ohlstein et al., 2004). For
example, in the testis, germ stem cells attach to the niche that comprises cap
cells. The cap cells release Jak/Stat and BMP ligands [Upd (Os) and Gbb/Dpp],
which maintain the stem cells and induce their proliferation. Whether
Drosophila ISCs utilize supporting cells that constitute a niche
remains unclear. Here we show that multiple EGFR ligands are involved in the
regulation of Drosophila AMP proliferation. During early larval
development, the midgut VM expresses the EGFR ligand vn
(Fig. 5C-C''), which is
required for AMP proliferation (Fig.
7; see Fig. S3B-B'' in the supplementary material). Thus, the
early AMPs might be considered to require a niche comprising non-epithelial
VM. Later in larval development, however, the AMPs express two other EGFR
ligands, spi and Krn
(Fig. 5E,F), which are capable
of autonomously promoting their proliferation
(Fig. 6) and may render
vn dispensable (Fig.
7E,E'; see Fig. S3B-B'' in the supplementary material).
We found, however, that depleting spi and Krn in the AMPs
did not affect AMP proliferation, suggesting that vn or another
trigger of EGFR/RAS/MAPK activity might complement spi and
Krn in late-stage larvae.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/136/3/483/DC1
|
Footnotes |
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REFERENCES |
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|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
|---|
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]
Bodenstein, D. (1994). The postembryonic
development of Drosophila. In Biology of Drosophila
(ed. M. Demerec), pp. 275-367. Cold Spring Harbor,
NY: Cold Spring Harbor Laboratory Press.
Brand, A. H. and Perrimon, N. (1993). Targeted
gene expression as a means of altering cell fates and generating dominant
phenotypes. Development
118,401
-415.[Abstract]
Brand, A. H. and Perrimon, N. (1994). Raf acts
downstream of the EGF receptor to determine dorsoventral polarity during
Drosophila oogenesis. Genes Dev.
8, 629-639.
Brand, M., Jarman, A. P., Jan, L. Y. and Jan, Y. N.
(1993). asense is a Drosophila neural precursor gene and is
capable of initiating sense organ formation.
Development 119,1
-17.[Abstract]
Brown, K. E., Baonza, A. and Freeman, M.
(2006). Epithelial cell adhesion in the developing Drosophila
retina is regulated by Atonal and the EGF receptor pathway. Dev.
Biol. 300,710
-721.[CrossRef][Medline]
Buttitta, L. A., Katzaroff, A. J., Perez, C. L., de la Cruz, A.
and Edgar, B. A. (2007). A double-assurance mechanism
controls cell cycle exit upon terminal differentiation in Drosophila.
Dev. Cell 12,631
-643.[CrossRef][Medline]
Cela, C. and Llimargas, M. (2006). Egfr is
essential for maintaining epithelial integrity during tracheal remodelling in
Drosophila. Development
133,3115
-3125.
Corley, L. S. and Lavine, M. D. (2006). A
review of insect stem cell types. Semin. Cell Dev.
Biol. 17,510
-517.[CrossRef][Medline]
Fre, S., Huyghe, M., Mourikis, P., Robine, S., Louvard, D. and
Artavanis-Tsakonas, S. (2005). Notch signals control the fate
of immature progenitor cells in the intestine. Nature
435,964
-968.[CrossRef][Medline]
Gabay, L., Seger, R. and Shilo, B. Z. (1997).
MAP kinase in situ activation atlas during Drosophila embryogenesis.
Development 124,3535
-3541.[Abstract]
Hall, B. L. and Thummel, C. S. (1998). The RXR
homolog ultraspiracle is an essential component of the Drosophila ecdysone
receptor. Development
125,4709
-4717.[Abstract]
Hartenstein, A. Y., Rugendorff, A., Tepass, U. and Hartenstein,
V. (1992). The function of the neurogenic genes during
epithelial development in the Drosophila embryo.
Development 116,1203
-1220.[Abstract]
Hartenstein, V. and Jan, Y. N. (1992). Studying
Drosophila embryogenesis with P-lacZ enhancer trap lines. Dev.
Genes Evol. 201,194
-220.
Hayashi, S., Hirose, S., Metcalfe, T. and Shirras, A. D.
(1993). Control of imaginal cell development by the escargot gene
of Drosophila. Development
118,105
-115.[Abstract]
Jiang, C., Baehrecke, E. H. and Thummel, C. S.
(1997). Steroid regulated programmed cell death during Drosophila
metamorphosis. Development
124,4673
-4683.[Abstract]
Juhasz, G. and Neufeld, T. P. (2008).
Drosophila Atg7: required for stress resistance, longevity and neuronal
homeostasis, but not for metamorphosis. Autophagy
4, 357-358.[Medline]
Karim, F. D. and Rubin, G. M. (1998). Ectopic
expression of activated Ras1 induces hyperplastic growth and increased cell
death in Drosophila imaginal tissues. Development
125, 1-9.[Abstract]
Kiger, A. A., White-Cooper, H. and Fuller, M. T.
(2000). Somatic support cells restrict germline stem cell
self-renewal and promote differentiation. Nature
407,750
-754.[CrossRef][Medline]
Klapper, R. (2000). The longitudinal visceral
musculature of Drosophila melanogaster persists through metamorphosis.
Mech. Dev. 95,47
-54.[CrossRef][Medline]
Lamb, M. J. (1982). The DNA content of polytene
nuclei in midgut and malpighian tubule cells of adult Drosophila melanogaster.
Roux's Arch. Dev. Biol.
191,381
-384.[CrossRef]
Lee, T. and Luo, L. (2001). Mosaic analysis
with a repressible cell marker (MARCM) for Drosophila neural development.
Trends Neurosci. 24,251
-254.[CrossRef][Medline]
Li, T. R. and White, K. P. (2003).
Tissue-specific gene expression and ecdysone-regulated genomic networks in
Drosophila. Dev. Cell 5,59
-72.[CrossRef][Medline]
Martin-Blanco, E. (1998). Regulatory control of
signal transduction during morphogenesis in Drosophila. Int. J.
Dev. Biol. 42,363
-368.[Medline]
Micchelli, C. A. and Perrimon, N. (2006).
Evidence that stem cells reside in the adult Drosophila midgut epithelium.
Nature 439,475
-479.[CrossRef][Medline]
Mirkovic, I. and Mlodzik, M. (2006).
Cooperative activities of drosophila DE-cadherin and DN-cadherin regulate the
cell motility process of ommatidial rotation.
Development 133,3283
-3293.
Morgan, N. S., Heintzelman, M. B. and Mooseker, M. S.
(1995). Characterization of myosin-IA and myosin-IB, two
unconventional myosins associated with the Drosophila brush border
cytoskeleton. Dev. Biol.
172, 51-71.[CrossRef][Medline]
Nilson, L. A. and Schupbach, T. (1999). EGF
receptor signaling in Drosophila oogenesis. Curr. Top. Dev.
Biol. 44,203
-243.[Medline]
O'Keefe, D. D., Prober, D. A., Moyle, P. S., Rickoll, W. L. and
Edgar, B. A. (2007). Egfr/Ras signaling regulates
DE-cadherin/Shotgun localization to control vein morphogenesis in the
Drosophila wing. Dev. Biol.
311, 25-39.[CrossRef][Medline]
O'Neill, J. W. and Bier, E. (1994).
Double-label in situ hybridization using biotin and digoxigenin-tagged RNA
probes. Biotechniques
17, 870,
874-875.
Ohlstein, B. and Spradling, A. (2006). The
adult Drosophila posterior midgut is maintained by pluripotent stem cells.
Nature 439,470
-474.[CrossRef][Medline]
Ohlstein, B. and Spradling, A. (2007).
Multipotent Drosophila intestinal stem cells specify daughter cell fates by
differential notch signaling. Science
315,988
-992.
Ohlstein, B., Kai, T., Decotto, E. and Spradling, A.
(2004). The stem cell niche: theme and variations.
Curr. Opin. Cell Biol.
16,693
-699.[CrossRef][Medline]
Prober, D. A. and Edgar, B. A. (2002).
Interactions between Ras1, dMyc, and dPI3K signaling in the developing
Drosophila wing. Genes Dev.
16,2286
-2299.
Queenan, A. M., Ghabrial, A. and Schupbach, T.
(1997). Ectopic activation of torpedo/Egfr, a Drosophila receptor
tyrosine kinase, dorsalizes both the eggshell and the embryo.
Development 124,3871
-3880.[Abstract]
Reich, A. and Shilo, B. Z. (2002). Keren, a new
ligand of the Drosophila epidermal growth factor receptor, undergoes two modes
of cleavage. EMBO J. 21,4287
-4296.[CrossRef][Medline]
Rintelen, F., Hafen, E. and Nairz, K. (2003).
The Drosophila dual-specificity ERK phosphatase DMKP3 cooperates with the ERK
tyrosine phosphatase PTP-ER. Development
130,3479
-3490.
Roch, F., Baonza, A., Martin-Blanco, E. and Garcia-Bellido,
A. (1998). Genetic interactions and cell behaviour in
blistered mutants during proliferation and differentiation of the Drosophila
wing. Development 125,1823
-1832.[Abstract]
Sancho, E., Batlle, E. and Clevers, H. (2004).
Signaling pathways in intestinal development and cancer. Annu. Rev.
Cell Dev. Biol. 20,695
-723.[CrossRef][Medline]
Schnepp, B., Donaldson, T., Grumbling, G., Ostrowski, S.,
Schweitzer, R., Shilo, B. Z. and Simcox, A. (1998). EGF
domain swap converts a drosophila EGF receptor activator into an inhibitor.
Genes Dev. 12,908
-913.
Schweitzer, R., Shaharabany, M., Seger, R. and Shilo, B. Z.
(1995). Secreted Spitz triggers the DER signaling pathway and is
a limiting component in embryonic ventral ectoderm determination.
Genes Dev. 9,1518
-1529.
Stainier, D. Y. (2005). No organ left behind:
tales of gut development and evolution. Science
307,1902
-1904.
Strumpf, D. and Volk, T. (1998). Kakapo, a
novel cytoskeletal-associated protein is essential for the restricted
localization of the neuregulin-like factor, vein, at the muscle-tendon
junction site. J. Cell Biol.
143,1259
-1270.
Szuts, D., Eresh, S. and Bienz, M. (1998).
Functional intertwining of Dpp and EGFR signaling during Drosophila endoderm
induction. Genes Dev.
12,2022
-2035.
Technau, G. M. and Campos-Ortega, J. A. (1986).
Lineage analysis of transplanted individual cells in embryos of Drosophila
melanogaster. III. Commitment and proliferative capabilities of pole cells and
midgut progenitors. Roux's Arch. Dev. Biol.
195,389
-398.[CrossRef]
van Es, J. H., van Gijn, M. E., Riccio, O., van den Born, M.,
Vooijs, M., Begthel, H., Cozijnsen, M., Robine, S., Winton, D. J., Radtke, F.
et al. (2005). Notch/gamma-secretase inhibition turns
proliferative cells in intestinal crypts and adenomas into goblet cells.
Nature 435,959
-963.[CrossRef][Medline]
Zettervall, C. J., Anderl, I., Williams, M. J., Palmer, R.,
Kurucz, E., Ando, I. and Hultmark, D. (2004). A directed
screen for genes involved in Drosophila blood cell activation.
Proc. Natl. Acad. Sci. USA
101,14192
-14197.
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