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First published online 11 February 2009
doi: 10.1242/dev.027466
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1 Department of Genetics, Harvard Medical School, 77 Avenue Louis Pasteur,
Boston, MA 02115, USA.
2 Howard Hughes Medical Institute, Harvard Medical School, 77 Avenue Louis
Pasteur, Boston, MA 02115, USA.
* Author for correspondence (e-mail: fdemontis{at}genetics.med.harvard.edu)
Accepted 23 January 2009
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SUMMARY |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Key words: Foxo, InR/Tor signaling, Myc, Muscle growth, Endoreplication, Body size, Feeding behavior
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INTRODUCTION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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), but an understanding of their inter-regulation
remains elusive (Hafen and Stocker,
2003). The Insulin receptor/Target of rapamycin (InR/Tor) pathway
regulates cell size and number and hence organ and body growth across
evolution (Hafen and Stocker,
2003). Upon Insulin/IGF binding to the Insulin and IGF receptors,
a signaling cascade of phosphorylation and docking events, antagonized by
Pten, results in the activation of the Ser/Thr kinase Akt. Akt then controls
cell survival, cell cycle, cell growth and metabolism through phosphorylation
of a number of key substrates, including the Tsc1-Tsc2 complex, and the
transcription factors of the Forkhead box O (FoxO) family. Phosphorylation and
inhibition of the Tsc1-Tsc2 complex, which has an inhibitory effect on Tor,
promotes protein synthesis (Manning and
Cantley, 2007). By phosphorylating and sequestering Foxo in the
cytoplasm, Akt further promotes cell growth and cell cycle progression
(Accili and Arden, 2004;
Greer and Brunet, 2008;
Puig and Tjian, 2006).
Similar to InR/Tor signaling, Myc has an evolutionarily conserved function
in promoting cell growth and proliferation
(de la Cova and Johnston,
2006). Myc regulates gene expression by binding to Enhancer box
sequences (E-boxes) in promoter regions of target genes, with its dimerization
partner Max, but also independently
(Steiger et al., 2008). Max
also dimerizes with itself and with members of the Mad/Mnt family, opposing
Myc-Max transcriptional activity
(Eisenman, 2001;
Gallant, 2006;
Grandori et al., 2000).
Although InR/Tor signaling, Foxo and Myc have been causally associated with
the growth of most cell types across species, how organ and body growth are in
turn determined is still unclear. Possibly, body size is decided by
stereotypical responses of each organ to growth factors, which in turn
regulate InR/Tor signaling and Myc activity. Alternatively, InR signaling in
some sensor tissues might have a pivotal role in modifying body growth in
response to the nutritional status of the organism. Consistent with this
model, InR/Tor signaling in the Drosophila fat body, which
corresponds to human liver and adipose tissue, and in endocrine glands
regulates the growth of other unrelated tissues and, consequently, of the
entire body, by modulating the actions of anabolic hormones
(Edgar, 2006). However, it is
unknown whether other tissues and mechanisms might contribute to the systemic
regulation of growth.
Muscles have important metabolic functions, undergo dramatic growth during development, and are continually remodeled throughout life. Despite their importance, it is unclear how muscle growth occurs and whether it contributes to the overall control of body size.
In vertebrates, several stimuli, including those activating InR/Tor
signaling and Myc, promote hypertrophy of skeletal muscles and cardiomyocytes
by inducing protein synthesis (Glass,
2003b). Conversely, inhibition of InR signaling, which results in
Foxo activation, promotes protein degradation and muscle atrophy
(Sandri et al., 2004;
Stitt et al., 2004). Other
processes, in particular an increase in DNA content, either by increasing the
number of nuclei or their ploidy, may be involved in muscle growth
(Brodsky and Uryvaeva, 1977;
Conlon and Raff, 1999).
Consistently, satellite cells fuse to pre-existing skeletal muscles,
increasing the number of nuclei and supporting hypertrophy
(Buckingham, 2006). Further,
cardiomyocytes increase their nuclear ploidy during the reparative growth that
follows an ischemic injury (Herget et al.,
1997; Meckert et al.,
2005). However, it is unknown whether nuclear ploidy can sustain
muscle growth, whether InR/Foxo signaling and Myc regulate these events, and
whether they crosstalk during muscle growth. Studies in epithelial and
hematopoietic cells have suggested that Myc might act either upstream
(Bouchard et al., 2007),
downstream (Bouchard et al.,
2004), or in parallel with Foxo
(Prober and Edgar, 2002).
Thus, the interplay of InR/Foxo signaling and Myc might rely on the specific
cellular context and needs to be analyzed in vivo to identify physiologically
relevant interactions.
Here, we have used Drosophila muscles to investigate: (1) how InR (Insulin-like receptor)/Tor signaling, Foxo and dMyc (Diminutive) interact in vivo during muscle growth; (2) whether they regulate the nuclear ploidy of muscle cells; (3) whether this is important for cell growth; and (4) whether muscle mass can in turn influence body size.
The Drosophila larval body wall muscles are skeletal muscles, each
comprising a single, multinucleated syncytial cell (myofiber) that arises from
the fusion of precursor cells (founder cells and fusion-competent myoblasts)
during embryonic development. Different degrees of cell fusion account for
different numbers of nuclei that are contained within distinct muscle cells
(Bate et al., 1999;
Beckett and Baylies, 2006).
During larval development, body wall muscles (see
Fig. 1A) grow dramatically in
5 days, via sarcomere assembly and the addition of novel myofibrils,
while the number of nuclei remains constant
(Bai et al., 2007;
Haas, 1950). Muscle growth may
also rely on an increase in nuclear ploidy, as previously observed for other
Drosophila tissues (Edgar and
Orr-Weaver, 2001; Maines et
al., 2004), via endoreplication (or endocycle), a modified cell
cycle in which DNA replication is not accompanied by mitosis but rather by
multiple G–S and S–G transitions
(Edgar and Orr-Weaver,
2001).
Here, we find that dMyc and InR/Foxo signaling are key regulators of endoreplication that is necessary, but not sufficient, for muscle growth. Foxo has a pivotal role in this process by regulating dmyc expression and activity downstream of InR signaling. The functional interaction of the transcription factors Foxo and dMyc controls the final muscle mass, which in turn influences body size by regulating larval feeding behavior.
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MATERIALS AND METHODS |
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SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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);
UAS-InR; UAS-Pten; UAS-Tsc1, UAS-Tsc2
(Potter et al., 2001);
UAS-InR DN (Bloomington #8253); UAS-dmyc, UAS-dmyc [second
transgene, tr2 (Orian et al.,
2007)], UAS-CycE (Bloomington #4781); UAS-dmnt
(Loo et al., 2005);
chico1/CyO, act-GFP; dm4/FM7i, act-GFP
(Pierce et al., 2008);
Dmef2-Gal4 (Ranganayakulu et al.,
1996); Mhc-Gal4 (Schuster et
al., 1996); UAS-Dcr-2
(Dietzl et al., 2007);
UAS-dmyc hp (CG10798, VDRC #2947); UAS-Akt1 hp
(CG4006, DRSC TR00202A.1); UAS-InR hairpin (hp)
[CG18402, DRSC TR00693A.1; courtesy of Dr Jianquan Ni
(Ni et al., 2008)];
Mhc-GFP (Wee-P26) (Clyne
et al., 2003); and UAS-H2B-CFP (from Dr Shu Kondo,
Harvard Medical School, Boston, MA, USA).
The PG157-Gal4 line is a lethal insertion at position 12F7 that drives high
transgene expression in ventral internal 1 muscle (VI1, also known as muscle
31 of abdominal segment 1) and muscles of the thoracic segment (see Fig. S5 in
the supplementary material). PG157-Gal4 does not drive transgene expression in
ventral longitudinal 3 and 4 muscles (VL3 and VL4, also known as muscles 6 and
7). Dmef2-Gal4 and Mhc-Gal4 drive transgene expression in all body wall
muscles, but not in other endoreplicating tissues (see Fig. S3 in the
supplementary material). For transgene expression with the Gal4-UAS system
(Brand and Perrimon, 1993),
flies were reared at 25°C (Dmef2-Gal4) or 31°C (Mhc-Gal4). Flies were
reared at 29°C for hairpin expression and at 22°C in
Fig. 1.
Body size analysis
For analysis of body weight, groups of L3 wandering larvae were weighed on
an analytical balance and the average body weight calculated. Larval staging
was supported by analysis of mouth hook morphology. Larval and pupal length
and diameter were measured manually using AxioVision v4.5 software (Zeiss).
Larval and pupal volumes were calculated assuming a prolate spheroid geometry.
For analysis of internal organs, dissected organs were stained in a
micro-chamber with the lipophilic dye FM4-64 [Molecular Probes
(Demontis and Dahmann, 2007)].
Images were acquired with an epifluorescence microscope (Zeiss). Adult flies
were analyzed according to Colombani et al.
(Colombani et al., 2005),
using the Measure Tools of the AxioVision software. Larval feeding behavior
was estimated as described previously (Wu
et al., 2005).
Histology, laser-scanning confocal microscopy and image analysis
Larvae were dissected in ice-cold Ca2+-free saline buffer (128
mM NaCl, 2 mM KCl, 4 mM MgCl2, 1 mM EGTA, 35 mM sucrose, 5 mM HEPES
pH 7.2) using dissection chambers (Budnik
et al., 2006). Body wall muscles were fixed for 20-30 minutes in
Ca2+-free saline buffer containing 4% paraformaldehyde and 0.1%
Triton X-100. After washing, body wall muscles were incubated for 10 hours
with DAPI (4',6-diamidino-2-phenylindole, 1 µg/ml) and Alexa633- or
Alexa488-conjugated phalloidin (1:100) to visualize nuclei and F-actin,
respectively. To examine biogenesis of nucleoli, an anti-Fibrillarin antibody
[EnCore Biotechnology #MCA-38F3 (Grewal et
al., 2005)] was applied (1:100), followed by incubation with
Alexa546-conjugated secondary antibodies (Molecular Probes). Muscles VL3 and
VL4 of abdominal segments 2-5 were imaged using a Leica TCS SP2 confocal
laser-scanning microscope. Confocal images were analyzed using the Measure
Tools of the AxioVision software. Statistical analysis was performed using
Student's t-test and Excel (Microsoft).
Luciferase assays, RNAi treatment and plasmid DNAs
For Luciferase assays, 15x104 S2R+ cells/cm2
were seeded in Schneider's medium (Gibco) containing 10% FCS, and transfected
one day later using the Qiagen Effectene Transfection Kit. An
actin-Renilla Luciferase reporter was co-transfected as a
normalization control.
Double-stranded RNA (dsRNA) synthesis and RNAi treatment were performed according to the DRSC protocols (http://flyRNAi.org), using amplicons DRSC34258 (dmyc) and DRSC31746 (foxo). RNAi treatment was performed for 3 days. foxo and dmyc expression were induced 24 hours prior to Luciferase assay by addition of CuSO4 directly to the culture medium to a final concentration of 500 µM. Serum starvation was also performed for 24 hours. The Luciferase assay was performed in quadruplicate using the Dual-Glo Luciferase Assay (Promega) according to the manufacturer's instructions. Luciferase activity refers to the ratio of firefly to Renilla Luciferase luminescence.
Plasmids used are pMT-foxo
(Puig et al., 2003),
pMT-dmyc (Orian et al.,
2003), actin-Renilla Luciferase, CG4364,
CG5033 and CG5033 
E-box Luciferase reporters
(Hulf et al., 2005).
Quantitative real-time RT-PCR
Total RNA was prepared from L3 wandering larvae using Trizol (Invitrogen),
followed by RNA cleanup with the RNAeasy Kit (Qiagen). The RNA QuantiTect
Reverse Transcription Kit (Qiagen) was used for cDNA synthesis, and
quantitative real-time PCR was performed with the QuantiTect SYBR Green PCR
Kit (Qiagen). 
Tub84B was used as normalization reference.
Relative quantitation of mRNA levels was calculated using the comparative
CT method.
Immunoprecipitation and immunoblotting
For immunoprecipitation, S2R+ cells were washed with ice-cold PBS, lysed
with lysis buffer (20 mM Tris pH 7.6, 150 mM NaCl, 2 mM EDTA, 10% glycerol, 1%
Triton X-100, 1 mM DTT, 1 mM PMSF and Protease inhibitors), and centrifuged at
10,000 g for 10 minutes at 4°C. Equal amounts of
supernatant were incubated with a monoclonal mouse anti-dMyc antibody
(Prober and Edgar, 2000), and
subsequently with an appropriate amount of protein A-agarose bead slurry
(Amersham) in lysis buffer. Immunoprecipitates were washed three times with
lysis buffer, boiled in sample buffer, resolved on 10% SDS-PAGE gels and
transferred to nitrocellulose membranes. Western blotting was performed with a
rabbit anti-Foxo (Puig and Tjian,
2005) or, after extensive membrane washing, with a rabbit
anti-dMyc antiserum (Maines et al.,
2004), and subsequently with anti-rabbit HRP-conjugated secondary
antibodies (Amersham). Western blot and densitometric analysis were performed
as previously described (Iurlaro et al.,
2004; Schlichting et al.,
2006).
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RESULTS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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)
(Fig. 1A; note that in
Drosophila larvae, one muscle is composed of a single multinucleated
cell). Because the DNA content of a cell has been directly correlated with its
size in a number of systems (Brodsky and
Uryvaeva, 1977; Conlon and
Raff, 1999), we have analyzed whether the differences observed in
muscle sizes correlate with either the number of nuclei or their level of
endoreplication (Edgar and Orr-Weaver,
2001).
We focused our analysis on two body wall muscles, VL3 and VL4, of third
instar (L3) wandering larvae, as they are easy to analyze and have distinct
sizes (Fig. 1A-C). VL3 muscles
possess twice the number of nuclei as VL4
(Fig. 1D), which correlates
with an approximate doubling in size (Fig.
1E) and a similar myofiber area/nucleus ratio
(Fig. 1F). Further, the nuclear
area (Fig. 1G) and the
intensity of DAPI staining (Fig.
1H), which are indicators of nuclear ploidy
(Maines et al., 2004;
Ohlstein and Spradling, 2006;
Sato et al., 2008;
Shcherbata et al., 2004;
Sun and Deng, 2007), did not
significantly differ, suggesting that the amount of endoreplication in VL3 and
VL4 nuclei is similar.
We next analyzed whether the ploidy of body wall muscle nuclei, as
indicated by nuclear size and DNA content, correlates with muscle growth
during larval development, similar to other Drosophila tissues
(Edgar and Orr-Weaver, 2001;
Maines et al., 2004). By
scoring muscle size (Fig. 1I)
and nuclear size (Fig. 1J) at
various developmental stages, a high degree of correlation between these
parameters and the intensity of DAPI staining was observed
(Fig. 1K). To test whether
endoreplication is necessary for growth, we overexpressed in muscles
Cyclin E, which has been shown to block endoreplication when present
at constant, but not oscillating, levels
(Lilly and Spradling, 1996).
Dmef2 (Mef2)-Gal4 UAS-CycE animals showed a severe
reduction in nuclear size, intensity of DAPI staining and muscle size,
demonstrating that muscle growth depends on endoreplication (see Fig. S1 in
the supplementary material). Altogether, these results reveal that the number
of nuclei is tightly coupled to the differential size of muscles, and that
increasing nuclear ploidy is required for the overall growth of the
muscles.
Inhibition of InR signaling in muscles regulates muscle growth, body size and the size of unrelated tissues
Since Insulin signaling is a major, evolutionarily conserved regulator of
cell size (Hafen and Stocker,
2003), cell cycle progression and endoreplication
(Burgering, 2008;
Ho et al., 2008), we tested
whether this pathway affects muscle growth. To modulate Insulin signaling in
muscles, three different approaches were used. First, muscles of wild-type
larvae were compared with those from larvae homozygous or heterozygous mutant
for chico [also known as Insulin receptor substrate
(IRS)] (see Fig. S1 in the supplementary material). Second, the
levels of InR and Akt (Akt1) were reduced via RNAi knockdown in muscles using
the Gal4-UAS system and Dmef2-Gal4 (see Fig. S2 in the supplementary
material), which drives transgene expression in muscles but not in other
endoreplicating tissues (see Fig. S3 in the supplementary material). Third, we
targeted the expression of activators (InR)
(Fig. 2B) and inhibitors of InR
signaling in muscles, including a dominant-negative form of InR
(InR DN) (Fig. 2C). In
all cases, inhibition of InR signaling resulted in decreased nuclear and
muscle size, as outlined by DAPI and phalloidin staining, respectively.
Conversely, overexpression of wild-type InR resulted in a significant
increase in myofiber size and nuclear size
(Fig. 2B), suggesting that InR
signaling controls muscle growth in part by modulating endoreplication.
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To test whether the size of larval organs and tissues other than muscles are affected when InR signaling is repressed using the Dmef2-Gal4 and Mhc-Gal4 muscle drivers, we examined their size in L3 wandering larvae following staining with the lipophilic dye FM4-64 to outline their dimensions. In addition to a reduction in muscle size (Fig. 2C), the size of other endoreplicating organs, such as the salivary glands, gut, fat body and epidermis, was decreased (Fig. 2G; see Fig. S8 in the supplementary material; data not shown). However, the size of non-endoreplicating tissues, including the brain, wing and eye-antennal imaginal discs was less affected. Further, upon activation of InR signaling in muscles, an increase in muscle size (Fig. 2B) was accompanied by a parallel increase in the size of most other tissues (see Fig. S3 in the supplementary material; data not shown).
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Reduction of muscle size by InR signaling non-autonomously regulates the size of other organs and affects feeding behavior
To characterize how changes in InR signaling in muscles affect the size of
other tissues, we analyzed the morphological changes induced in fat body and
salivary glands following Pten overexpression in muscles. Strikingly,
reduction of Insulin signaling in muscles was accompanied by a reduction of
cell size in endoreplicating tissues, via lipid remobilization in fat body
cells (Fig. 3A,A'), and
activation of catabolic programs possibly related to autophagy in salivary
glands (Fig. 3B,B').
Because lipid remobilization and activation of catabolic programs in
endoreplicating tissues are common events in response to improper feeding
behavior and metabolic regulation
(Colombani et al., 2003), we
tested whether feeding was affected in larvae with either repressed or
activated InR signaling in muscles. Feeding behavior is under strict control
in Drosophila and other organisms, as nutrient uptake is crucial for
appropriate developmental growth (Saper et
al., 2002). To monitor feeding activity, the number of mouth hook
contractions, which has been shown to be an indicator of this behavior
(Wu et al., 2005), was scored.
Interestingly, overexpression of the InR antagonists Pten, Tsc1 and
Tsc2 (gigas) or of foxo in muscles resulted in a
significant decrease in larval feeding, whereas InR overexpression
promoted this behavior (Fig.
3C). Thus, we propose that the levels of InR signaling in muscles
somehow modulate larval feeding behavior, which in turn influences body size
and tissue growth.
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Although most manipulations of InR signaling during larval muscle growth result in pupal lethality, we recovered Dmef2-Gal4 UAS-InR and Dmef2-Gal4 UAS-InR DN adult flies, in which InR signaling was activated and inhibited, respectively. As expected, whereas activation of InR during muscle growth resulted in larger flies, smaller flies arose upon inhibition of this pathway (Fig. 4A). To test whether developmental muscle growth regulates, in turn, the size of body parts in adults, we scored the weight, eye size, abdomen length and wing area of these recovered flies. As expected, all these parameters were respectively either increased or decreased upon activation or inhibition of InR signaling in muscles. Changes in tissue and whole-body size occurred by modulating cell size, as observed in the wings of adult flies, whereas cell number barely varied (Fig. 4B). Thus, several tissues, deriving from both endoreplicating and non-endoreplicating tissues, are affected to different extents upon developmental modulation of muscle mass.
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; Pierce et al.,
2004). To test whether dMyc plays a role in muscle growth, we
modulated its function in several ways. First, muscles from wild-type larvae
were compared with those from larvae that were homozygous or heterozygous
mutant for dmyc (see Fig. S1 in the supplementary material). Second,
levels of dMyc were reduced via RNAi knockdown in muscles (see Fig. S2 in the
supplementary material). Third, we targeted expression of dmyc in
muscles (Fig. 5E), as well as
the expression of its inhibitor dmnt (Mnt) (see Fig. S4 in
the supplementary material). In all cases, inhibition of dMyc activity
resulted in smaller muscles and nuclei and decreased body size. dMyc
overexpression was associated with an increase in nuclear size and DAPI
staining that was, however, accompanied by only a slight increase in muscle
size (Fig. 5E,H-K). Thus,
during muscle growth, dMyc is both necessary and sufficient to regulate
endoreplication. However, although dMyc and endoreplication are necessary,
they are not sufficient to sustain extensive growth.
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),
overexpression of foxo in muscles resulted in a significant, 2.5-fold
reduction in dmyc transcript levels
(Fig. 5A').
Because the regulation of dmyc gene expression by InR/Foxo might
only in part account for the regulation of dMyc activity, we tested whether
InR and Tor signaling regulate dMyc protein function, as estimated by their
ability to control dMyc-driven phenotypes in muscles. When either
Pten, or Tsc1 and Tsc2 were co-expressed together
with dmyc, dMyc activity was inhibited, resulting in defective
myofiber growth and endoreplication (Fig.
5F,G), similar to the expression of Pten
(Fig. 5C) or Tsc1 and
Tsc2 alone (Fig. 5D).
Consistent with being regulated by InR signaling, foxo overexpression
also impaired dMyc activity (see Fig. S4 in the supplementary material;
Fig. 5H-K). Quantification of
muscle phenotypes indicated that significant changes in myofiber area
(Fig. 5I), nuclear size
(Fig. 5J) and the intensity of
DAPI staining (Fig. 5K) occur
in concert, without any change in the number of nuclei
(Fig. 5H). Thus, maximal dMyc
activity relies on optimal InR/Tor signaling and inhibition of Foxo activity.
Furthermore, and contrary to previous analyses in fat body cells
(Pierce et al., 2004), dMyc
overexpression in muscles promoted endoreplication without a proportional
increase in cell size.
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Foxo inhibits dMyc transcriptional activity
dMyc acts primarily via inducing a transcriptional response, suggesting
that InR signaling might regulate dMyc function by modulating its
transcriptional activity. Similar to Pten and Tsc, Foxo can regulate dMyc
protein function in vivo (see Fig. S4 in the supplementary material;
Fig. 5H-K). To test whether
Foxo inhibits dMyc by regulating its transcriptional activity, Luciferase
assays were performed using CG4364 and CG5033
transcriptional reporters, previously described to be directly regulated by
dMyc (Hulf et al., 2005) but
not directly regulated by Foxo. The CG5033 E-box reporter is
devoid of E-boxes, the dMyc-responsive sequences, and is therefore refractory
to dMyc transcriptional regulation.
In S2R+ cells, Luciferase activity of CG4364 and CG5033
reporters was detected in response to endogenous dMyc and was increased by
overexpression of dmyc (pMT-dmyc) (for characterization of
overexpression, see Fig. S7 in the supplementary material). However,
overexpression of wild-type foxo (pMT-foxo) (see Fig. S7 in
the supplementary material) or serum starvation, which activates endogenous
Foxo, decreased the Luciferase activity of the CG4364 and
CG5033 reporters (Fig.
6G) and suppressed the transcriptional response induced by
dmyc overexpression. Further, without E-boxes, no substantial
Luciferase activity was detected, indicating that it depends on dMyc.
Consistently, RNAi treatment against dmyc and foxo
respectively attenuated and increased Luciferase activity of the
CG4364 and CG5033 reporters
(Fig. 6H; see Fig. S7 in the
supplementary material), but not of the CG5033 E-box reporter.
The increase in Luciferase activity upon foxo RNAi is likely to
reflect its ability to inhibit both dmyc gene expression and dMyc
protein function. However, upon RNAi treatment of foxo and
dmyc, no increase in Luciferase activity was observed
(Fig. 6H), further confirming
that Foxo regulates transcription of the CG4364 and CG5033
reporters via dMyc. Therefore, Foxo tightly controls dMyc function by
modulating its expression (Fig.
5) and its transcriptional activity.
dMyc primes muscle growth via nucleolus biogenesis and expression of growth-promoting genes
Although dMyc promotes endoreplication, no substantial muscle growth
results. To further dissect the role of dMyc in muscle growth, we tested
whether dMyc induces (1) nucleolus biogenesis, which is necessary for rRNA
transcription and ribosome subunit assembly
(Prieto and McStay, 2005), and
(2) the expression of genes required for protein translation; both events are
necessary for cell growth. By staining with an anti-Fibrillarin antiserum,
which outlines nucleoli (Grewal et al.,
2005), we observed a dramatic increase in the size of the
nucleolus upon dmyc overexpression with the PG157-Gal4 driver, in
comparison with controls (Fig.
7A,B). Further, dmyc overexpression with Dmef2-Gal4
increased, to different extents (Fig.
7C), the expression of some dMyc target genes
(Grewal et al., 2005;
Hulf et al., 2005) that are
involved in rRNA processing (Nop60B), ribosome assembly and
biogenesis (CG1381, CG5033, Surf6), and translational control
(eIF6), but not cell proliferation (CG4364). InR
overexpression promoted a modest increase in the expression of dMyc target
genes involved in growth (CG1381, CG5033, eIF6, Surf6). Thus, dMyc
primes muscles for growth by promoting endoreplication, nucleolus biogenesis
and the expression of some genes necessary for protein translation. However,
gene expression programs governed by dMyc require concomitant InR/Tor
signaling to drive substantial muscle growth
(Fig. 8).
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DISCUSSION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Foxo regulates endoreplication and dMyc transcriptional activity
We found that dMyc, as well as activation of InR signaling, can promote
endoreplication in muscles, whereas Foxo and inhibitors of dMyc and of InR/Tor
have the opposite effect. dMyc is likely to regulate the expression of genes
required for multiple G–S and S–G transitions during
endoreplication (Edgar and Orr-Weaver,
2001; Lilly and Duronio,
2005), similar to vertebrate Myc, which regulates key cell-cycle
regulators including cyclin D2, cyclin E, and the cyclin kinase inhibitors p21
and p27 (Cdkn1a and Cdkn1b, respectively)
(Grandori et al., 2000).
Indeed, aberrant levels of Cyclin E (Lilly
and Spradling, 1996) block muscle growth (see Fig. S1 in the
supplementary material), indicating that proper muscle growth requires tight
control of the expression and activity of endoreplication genes. Further, we
found that endoreplication is also modulated by Foxo, which is activated in
conditions of nutrient starvation, impaired InR/Tor signaling and by other
cell stressors (Arden, 2008).
Foxo presumably regulates cell cycle progression at least in part by
modulating the expression of evolutionarily conserved Foxo/Myc-target genes,
such as dacapo (the Drosophila p21/p27 homolog) and
Cyclin E, that regulate the G1–S transition, as previously
reported in mammalian systems (Grandori et
al., 2000; Salih and Brunet,
2008). Interestingly, Foxo and Myc might control different steps
in the activation of common target genes
(Bouchard et al., 2004).
In addition, we found that active Foxo can also inhibit dMyc protein
activity and regulates dmyc gene expression. Mechanistically, Foxo
could influence dMyc activity in several ways. First, it might physically
interact with dMyc, although we found no evidence to support this notion (see
Fig. S7 in the supplementary material). Second, Foxo could regulate the
expression of genes that target dMyc for proteasomal degradation, including
several ubiquitin E3 ligases that are induced by Foxo during muscle atrophy in
mice and humans (Sandri et al.,
2004; Stitt et al.,
2004). However, by analyzing dMyc protein levels by western blot,
we did not detect significant dMyc protein instability upon Foxo
overexpression (see Fig. S7 in the supplementary material). Third, Foxo might
promote the expression of transcriptional regulators that oppose dMyc
function, including Mad/Mnt (Delpuech et
al., 2007), although no substantial increase in dmnt mRNA
levels was detected upon Foxo activation in muscles (not shown). Possibly, the
expression of other dMyc regulators might be affected by Foxo. Future
experiments will be needed to dissect the Foxo-dMyc interaction.
Finally, by manipulating muscle growth and/or endoreplication, we found
that in muscles the ratio of cell size to nuclear size is not constant, and
increased nuclear size and DNA content, indicative of ploidy, is necessary but
not sufficient to drive growth. Usually, an increase in cell size is matched
by an increase in nuclear size (Neumann
and Nurse, 2007), which commonly parallels increases in nuclear
ploidy (Maines et al., 2004;
Ohlstein and Spradling, 2006;
Sato et al., 2008;
Shcherbata et al., 2004;
Sun and Deng, 2007). However,
our findings indicate that in muscles, dMyc-driven variation in nuclear size
and ploidy is permissive but not sufficient for substantial growth, even in
the presence of increased biogenesis of nucleoli and expression of genes
involved in protein translation. This is different from fat body cells, in
which dmyc overexpression induces endoreplication and proportional
cell growth (Pierce et al.,
2004). Thus, additional instructive signals, possibly modulating
protein synthesis, mitochondriogenesis, ribosome biogenesis
(Teleman et al., 2008),
sarcomere assembly (Bai et al.,
2007; Haas, 1950),
and other anabolic responses must be concomitantly received to promote maximal
muscle growth. Therefore, increases in cell size and nuclear ploidy are
surprisingly uncoupled during muscle growth.
Muscle size regulates systemic growth
Little is known about the mechanisms that control and coordinate cell,
organ and body size (Edgar,
2006; Mirth and Riddiford,
2007), and in particular how muscle growth is matched with the
growth of other tissues and of the entire organism. We found that inhibition
of InR/Tor signaling and dMyc activity in muscles impairs, in addition to
muscle mass, the size of the entire body and of other internal organs.
Similarly, overexpression of Cyclin E in muscles also resulted in
autonomous and systemic growth defects (see Figs S1 and S8 in the
supplementary material), indicating that, at least in some cases, modulation
of muscle growth by means independent from InR signaling can be sensed
systemically. In the larva, endoreplicating tissues and organs (gut, salivary
glands, epidermis, fat body) were severely affected, whereas
non-endoreplicating tissues (brain and imaginal discs) were less affected,
indicating distinct tissue responsiveness to this regulation. Similarly,
inhibition of Tor signaling in the fat body also primarily affects the size of
endoreplicating tissues (Britton et al.,
2002; Colombani et al.,
2003).
Non-autonomous regulation of tissue size may rely on humoral factors (e.g.
hormone-binding proteins, hormones, metabolites) produced by muscles in
response to achieving a certain mass
(Gamer et al., 2003). However,
alternative models are possible. In particular, we observed decreased and
increased larval feeding, respectively, upon inhibition and activation of InR
signaling in muscles. This whole-organism behavioral adaptation is possibly
due to decreased and increased efficiency of smaller and bigger muscles,
respectively, and to regulated expression of neuropeptides that hormonally
control feeding behavior. As a consequence of the regulation of feeding
behavior, nutrient uptake is decreased and larval growth is blocked in the
cells of endoreplicating tissues, which are extremely sensitive to poor
nutritional conditions, and to a lesser extent in non-endoreplicating tissues,
which are more resistant to limited nutritional supply
(Bradley and Leevers, 2003;
Colombani et al., 2003). In
turn, increased or decreased size of non-muscle tissues arise as a consequence
of abnormal feeding. Thus, muscle size coordinates with the size of other
organs and of the entire body, at least in part via a systemic, behavioral
response. Distinct tissues are differently sensitive to this regulation,
resulting in altered body proportions.
Drosophila as a disease model of muscle atrophy and hypertrophy
Understanding the mechanisms regulating muscle mass is of special interest
because they underline the etiology of several human diseases
(Glass, 2003a). Directly
relevant to our studies, both MYC and InR (INSR) signaling have been found to
regulate muscle growth and maintenance in humans
(Sandri et al., 2004;
Southgate et al., 2007;
Stitt et al., 2004;
Zhong et al., 2006). Further,
muscle atrophy is triggered by FOXO activation in several pathological
conditions (Glass, 2003b;
Sandri et al., 2004;
Stitt et al., 2004). In
addition, MYC function has been implicated in heart hypertrophy
(Bello Roufai et al., 2007;
Xiao et al., 2001;
Zhong et al., 2006), a process
that is conversely regulated by FOXO
(Evans-Anderson et al., 2008;
Skurk et al., 2005).
Our findings that Foxo functionally antagonizes dMyc during the growth of
Drosophila muscles suggest that these factors might also interact
similarly in humans. Consistent with this hypothesis, FOXO and MYC regulate,
in opposite fashions, the atrophic and hypertrophic programs in human skeletal
muscles and cardiomyocytes, and display complementary gene expression and
activity in these contexts (Lecker et al.,
2004; Mahoney et al.,
2008; Sandri et al.,
2004; Spruill et al.,
2008; Stitt et al.,
2004).
Finally, our finding that during larval development, inhibition of InR
signaling in muscles has profound systemic effects might also reflect
physiological conditions found in humans. Indeed, defective responsiveness of
muscles to Insulin during type II diabetes has autonomous effects on muscle
maintenance that are associated with systemic effects on the metabolism of the
entire organism, contributing to the improper control of glycemia and the
development of metabolic syndrome (Wells
et al., 2008). Here, we have identified feeding behavior as part
of the systemic response that in Drosophila senses perturbations in
muscle mass. These findings might help further elucidate the signals involved
in metabolic and growth homeostasis, which may be conserved across
evolution.
|
Footnotes |
|---|
Supplementary material for this article is available at http://dev.biologists.org/cgi/content/full/136/6/983/DC1
|
REFERENCES |
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|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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