Growth hormone promotes skeletal muscle cell fusion independent of insulin-like growth factor 1 up-regulation

As the body weight of GHR−/− mice was reduced, the reduction of muscle mass was evaluated by weighing two distinct hind limb muscles of female and male mice and normalizing to total body weight (Table 1, which is published as supporting information on the PNAS web site). The absolute muscle weight and the muscle/body weight of soleus and tibialis anterior were reduced in GHR−/− mice of both genders as compared with wild type. Thus, the disproportionate reduction in muscle mass is not only a consequence of the global dwarfism of the GHR−/− mice and suggests a specific action of GH as a muscle growth factor.

To study muscle fiber composition and morphometric characteristics, we performed immunohistological analysis of transversal cross sections from female soleus muscles with anti-myosin heavy chains (MyHCs) antibodies. Consistent with the reduction of muscle mass, the CSA of the entire GHR−/− soleus was reduced as compared with wild type (Fig. 6A, which is published as supporting information on the PNAS web site). Because muscle mass is determined by the number and size of individual fibers, we first counted the total number of soleus myofibers, which is set during embryonic development, and we observed no difference between the numbers of wild-type and mutant myofibers (Fig. 1B). These data, combined with the onset of growth retardation at the second week after birth (16), indicate that GH is not involved in the control of muscle growth at the embryonic and perinatal stages.

Muscles are composed of different fiber types that vary in their contractile and metabolic status. Slow myofibers express type I MyHC and are oxidative, whereas fast myofibers express type II MyHC and are glycolytic. In addition, hybrid fibers expressing MyHC type I and II are also found. In GHR−/− soleus, we measured a 26% decrease of type I fiber number and a 16% increase of type II fibers (Fig. 1C), indicating that the lack of GHR signaling causes a switch from type I and hybrid fibers to type II fibers. Consistent with these data, overexpression of human GH (hGH) in transgenic mice led to an increased percentage of type I fibers, confirming that GH has a positive role in type I fiber specification (17). Next, we measured the CSA of both fiber types in soleus. GHR deletion decreased type I and type II fiber size by 36% and 40%, respectively (Fig. 1 A, B, and D). Similar reductions in CSA were measured in male GHR−/− soleus and in GHR−/− tibialis anterior from both genders (Fig. 6B). Taken together, these data show that the reduced mass of GHR−/− muscles is due to smaller individual myofiber size rather than to a change in their number. Thus, GHR deletion specifically affects postnatal myofiber growth and specification, whereas myofiber formation during embryogenesis is unchanged.

The dwarf phenotype of GHR−/− mice is associated with low circulating IGF-1 levels, a defect that may lead to growth retardation (16). To distinguish between humoral or cell-autonomous control of muscle size in GHR−/− mice, we established primary muscle cell cultures from wild-type and GHR−/− mice. Myoblast cultures of both genotypes expanded similarly in mitogen-rich medium as assessed by cell counting and by FACS analysis (data not shown and Fig. 2A). Next, we assessed GH-stimulated cell-cycle progression by FACS analysis of cell-cycle phase distribution and by BrdU incorporation. In starvation conditions the percentage of cells in S-phase was largely reduced as compared with complete medium conditions (Fig. 2A). Of note, GH concentrations ranging from 10 to 1,000 ng/ml failed to induce S-phase entry of wild-type cells as compared with starved conditions (Fig. 2A, Fig. 7, which is published as supporting information on the PNAS web site, and data not shown). In contrast, serum treatment stimulated both wild-type and GHR−/− cell-cycle progression. To control whether the lack of GH-induced proliferation was not due to the absence of GHR expression and signaling, we checked GHR expression and phosphorylation of Janus kinase 2 (Jak2), the tyrosine kinase associated to GHR. GHR expression and GH-induced Jak2 phosphorylation were detected in wild-type cells, indicating that the first steps of GH signaling were intact (Fig. 2B). These data show that wild-type myoblasts express functional GHR receptors, although their stimulation by GH does not favor cell-cycle progression.

To measure cell size, the parameter of forward scatter height was determined by flow cytometry. Because cell size also depends on DNA content, forward scatter height was measured in G₁-phase gated cells, and similar values were obtained for myoblasts of both genotypes (Fig. 7B). Thus, the growth defect of GHR−/− muscles is not accompanied by impaired myoblast cell growth and proliferative capacity.

To determine whether the size of differentiated cells was affected by GHR deletion, we switched myoblasts to mitogen-poor differentiation medium (DM) and followed their differentiation. Forty-eight hours after mitogen removal, both wild-type and GHR−/− cells differentiated to multinucleated myotubes (Fig. 3A). The kinetics of muscle differentiation were followed by the expression of appropriate molecular markers. The expression of myogenin and the myofibrillar proteins α-skeletal actin, MyHC, and troponin T showed similar onset in GHR−/− and wild-type cells (Fig. 8A, which is published as supporting information on the PNAS web site). In addition, we did not observe an effect of GH treatment on wild-type cell differentiation (data not shown). Altogether, these observations suggest that GH signaling does not affect muscle cell differentiation.

Strikingly, 48 h after differentiation GHR−/− cells formed myotubes that were 25% smaller than wild type (Fig. 3A). To further assess the implication of GH in myotube size control, cells were induced to differentiate in the presence of GH, and myotube diameter was measured after 48 h. GH treatment increased the myotube size by 20% in wild-type cells and had no effect in GHR−/− cells (Fig. 3A). The differences in size due to GHR deletion or GH stimulation were comparable at 48 h, 72 h, and 96 h after switching to DM, indicating that GH signaling controls myotube size at the early steps of differentiation during myotube formation (Fig. 8B). To demonstrate that the absence of GHR was the primary cause of the small size of GHR−/− myotubes, we overexpressed murine GHR or β-gal control in GHR−/− cells by using adenoviral vectors. Cells were transduced at the onset of differentiation and analyzed 48 h later (Fig. 3B). The transduction of GHR−/− cells with GHR adenovirus restored the expression of GHR in a dose-dependent manner as assessed by RT-PCR analysis. Importantly, GHR expression significantly enhanced the GHR−/− myotube size. These data show that GH controls myotube size in a cell-autonomous manner during the differentiation of muscle cells.

The reduction of myofiber size could be due to an impairment of myoblast cell fusion and/or a reduction of the cytoplasmic domain regulated by one myonucleus. Fusion consists of two distinct phases: myoblast/myoblast fusion to form nascent myotubes and subsequent myoblast/myotubes fusion, resulting in a rapid accretion of size. The efficiency of the first phase can be evaluated by measuring the fusion index, which represents the proportion of the total cell population that has fused. After 48 h in DM, the fusion index did not differ as a function of the genotype or the stimulation by GH (Fig. 4A), suggesting that GHR signaling does not control nascent myotube formation.

To determine whether GH increased muscle cell size by recruiting new nuclei to existing myotubes, the number of nuclei per myotube was determined in wild-type and GHR−/− cultures. The nuclear number and the percentage of myotubes having five or more nuclei were diminished in GHR−/− myotubes as compared with wild type (Fig. 4B and Fig. 9A, which is published as supporting information on the PNAS web site). In addition, GH significantly increased the nuclear number in wild-type and not in GHR−/− cells. Of note, 48 h after differentiation the fusion process was completed, because the myotube nuclear number from both genotypes did not vary at later time points (Fig. 8B). To rule out the possibility that the increased myonuclear number was due to an effect of GH on cell proliferation/survival in DM, the number of nuclei per square millimeter was assessed after 48 h in DM (Fig. 9B). No difference in the DNA content was observed between GHR−/− and wild-type cells treated or not with GH. These data, together with the lack of GH-induced proliferation, suggest a specific role of GH in enhancing cell fusion rather than controlling the number of nuclei available for fusion. To further examine the kinetics of GH-stimulated cell fusion, cells were treated with GH at different stages of differentiation. Cells were treated once at the onset of differentiation (time 0) or at 24 h, when cells begin to fuse and nascent myotubes appear (time 24). In both cases, myotube size was analyzed at 48 h in DM. GH did not have a hypertrophic effect when added at the onset of differentiation (Fig. 4C). However, when administered at 24 h, GH increased the size of myotubes by 25%. Administration of GH at 24 h alone was sufficient to stimulate growth as much as the daily administration at time 0 and 24 h (0 + 24). Thus, GH acts when nascent myotubes are already formed to allow an increase in muscle cell size by enhancing muscle cell fusion.

The calcineurin/NFATc2 pathway has been implicated in myotube cell size control by regulating the second phase of fusion in part through the up-regulation of IL4 gene expression (13). Because the timing of GH action on cell fusion parallels NFATc2 activation (18, 19), we investigated the implication of NFATc2 and IL4 in the GH hypertrophic action. NFATc2−/− and IL4−/− cells were differentiated in the presence of GH, and myotube size was measured after 48 h. As already reported, at the basal state NFATc2−/− and IL4−/− myotubes were smaller than wild type (Fig. 4D) (13). Strikingly, GH treatment had no effect on NFATc2−/− myotube size. The complete resistance of NFATc2−/− cells to the hypertrophic action of GH was not due to the lack of expression of GHR mRNAs in those cells as assessed by RT-PCR analysis (Fig. 4D). Interestingly, the size of IL4−/− myotubes increased upon GH addition, indicating that this downstream target of NFATc2 is not involved in GH-induced cell fusion. These data suggest that NFATc2 activity is required for the growth-promoting action of GH. NFATc2 could be a direct target of the GHR signal transduction pathway or may belong to a parallel pathway that is permissive for cell fusion.

To investigate whether the reduced nuclear number of GHR−/− myotubes was also observed in mouse muscles, we counted the number of nuclei inside the sarcolemma of soleus and tibialis anterior transversal sections. A 21% decrease in myonuclear number was observed in both GHR−/− muscles as compared with wild type (Fig. 9C). These data suggest that the defect of muscle cell fusion contributes to the impaired muscle growth of GHR−/− mice.

The action of GH on muscle cell size could be mediated by IGF-1, a potent muscle growth factor whose expression is induced by GH in different experimental models. To discriminate between direct or IGF-1-mediated hypertrophic effects of GH, we first estimated the levels of IGF-1 mRNA in wild-type cells by semiquantitative RT-PCR during differentiation and GH stimulation. In the IGF-1 gene two distinct promoters drive IGF-1 transcripts. Class 1 transcripts contain exon 1 and are mainly expressed in skeletal muscle, whereas class 2 transcripts contain exon 2 and are expressed in both liver and muscle (5). Because both classes of transcripts may be regulated by GH (20), we analyzed the expression of IGF-1 mRNA isoforms containing exon 1, exon 2, or the common exons 3 and 4. As reported, the expression of all IGF-1 transcripts reached a peak 24 h after the onset of differentiation (Fig. 5A) (21). Importantly, GH stimulation did not further increase IGF-1 transcript levels at both differentiation time 24 and 48 h, indicating that the GH action on myotube hypertrophy does not correlate with IGF-1 levels. Shorter incubation times (1, 2, or 4 h) of wild-type cells with doses of GH ranging from 30 to 600 ng/ml also failed to affect IGF-1 expression (data not shown). This finding differs from what was reported in the C2C12 cell line, where GH increases the expression of IGF-1 (6, 7). The transcription factor Stat5 has been shown to participate in the control of IGF-1 expression by GH (20, 22). In wild-type myotubes we failed to detect GH-induced phosphorylation of Stat5, and adenoviral overexpression of a Stat5 dominant-negative protein (20) had no effect on the basal and GH-stimulated myotube size, indicating a marginal role of Stat5 in mediating GH action in our muscle cultures (data not shown). Moreover, we did not note any significant difference in IGF-1 levels between wild-type and GHR−/− cells (Fig. 5B). Finally, induction of GHR expression in GHR−/− cells did not affect IGF-1 expression (Fig. 5B) but enhanced myotube size (Fig. 3B). Taken together, these data suggest that in our cellular system there is no regulation of IGF-1 expression by GH and no correlation between GH-regulated myotube size and IGF-1 levels.

If the reduced GHR−/− myotube growth was due to the absence of a secreted factor, such as IGF-1 or others, we would expect that the conditioned medium (CM) from wild-type cells would rescue GHR−/− myotube size. To test this possibility, CM were collected from wild-type or GHR−/− cells and added to GHR−/− cells during differentiation. After 48 h of differentiation the myotube size was analyzed and compared with GHR−/− cells in DM. The CM from untreated or GH-stimulated wild-type cells did not affect GHR−/− cell size (Fig. 5C). This finding suggests that GHR−/− cells are not defective for the production of a secreted factor controlling myotube growth. Of note, wild-type CM did not affect wild-type myotube size, whereas CM from cells treated with exogenous IGF-1 rescued GHR−/− myotube size and exerted a hypertrophic action on wild-type cells. These data indicate that this assay is scoring the lack or presence of secreted factors rather than small variations in their amount. To evaluate the activity of CM, we tested whether it was able to reverse the small size of NFATc2−/− cells, which results from their defect in producing IL4 (13). When NFATc2−/− cells were cultured in wild-type CM they formed large myotubes similar to wild-type cells. Importantly, CM from GHR−/− cells also rescued NFATc2−/− cell size, indicating that GHR−/− cells do not produce inhibitory factors precluding growth. In conclusion, CM rescues fusion of the NFATc2−/− cells but not the GHR−/− cells. These data do not support the defective production of a secreted factor in the control of GHR−/− cell size.

To study the relationship between the growth-promoting actions of GH and IGF-1, we analyzed whether their effects were additive or mediated by a common mechanism. IGF-1 stimulated an 20% increase in myotube diameter of wild-type cells, indicating that cells are responsive to an increase in the extracellular concentration of IGF-1 (Fig. 5D). The effect of IGF-1 in GHR−/− cells was comparable to the one in wild-type cells, demonstrating that mutant cells are not refractory to this growth stimulus. Strikingly, when wild-type cells were stimulated with the combination of GH and IGF-1, myotube size increased in an additive manner as compared with cells treated with GH or IGF-1 alone. Finally, to determine whether NFATc2 activity was required for IGF-1 hypertrophic action, NFATc2−/− cells were treated with IGF-1. Interestingly, IGF-1 induced a 20% increase in NFATc2−/− myotube size and myonuclear number comparable with that observed for wild-type cells, whereas GH had no effect (Fig. 5D and Fig. 10, which is published as supporting information on the PNAS web site). This finding suggests that GH and IGF-1 use different signaling pathways to induce fusion and growth. In conclusion, our data suggest that GH promotes a cell-autonomous increase in myotube cell size that occurs through a facilitation of cell fusion and is independent of local IGF-1 up-regulation.

Wild-type and GHR−/− littermates were bred from heterozygous mice of the genetic background Sv129Ola, and offspring were genotyped as described (16). Mice were handled in accordance with institutional animal care policies. Histological immunostaining were performed as described in Supporting Text, which is published as supporting information on the PNAS web site.

Primary cultures were derived from gastrocnemius and tibialis anterior muscles of 4-week-old mice as described (ref. 15 and Supporting Test). BrdU incorporation and FACS analysis were performed as described in Supporting Text.

To differentiate muscle cells, 20,000 myoblasts per square centimeter were plated on Matrigel-coated dishes in DMEM/Ham F12/2% horse serum (DM). To measure the hypertrophic response, cells were incubated with 600 ng/ml hGH and/or 250 ng/ml IGF-1-R3 (Sigma) in DM. Hormones were added daily. After 48 h, the diameter of at least 400 myotubes was measured. The nuclear number was determined after staining with Hoechst 33258. The fusion index was determined by dividing the number of nuclei within multinucleated myotubes by the total number of nuclei analyzed.

To evaluate the effect of CM, culture medium was collected from muscle cells after 24 and 48 h in DM with or without 250 ng/ml IGF-1-R3 or 600 ng/ml hGH. Myoblasts were seeded in CM, which was replaced after 24 h. Myotube diameter was analyzed after 48 h.

Adenovirus vectors containing either β-galactosidase (β-galAdlox) or the murine GHR cDNA (GHRAdlox) were used (31). For the infection, myoblasts were infected with 30, 100, or 300 multiplicities of infection in DM for 1.5 h. After washing, cells were incubated in DM and analyzed 48 h later.

Cell lysates were prepared as described (15). For immunoprecipitation studies, cells were starved overnight and stimulated with 300 ng/ml hGH for 5 min. After cell lysis, 500 μg of total protein extract was incubated overnight with protein A Sepharose and anti-Jak2 antibodies (Upstate Biotechnology, Lake Placid, NY). Protein extracts were analyzed by immunoblotting by using the indicated primary antibodies (Supporting Text).

RNA was isolated by using TRIzol Reagent (Life Technologies). Reverse transcription was performed from 2 μg of total RNA with Moloney murine leukemia virus reverse transcriptase (Life Technologies) and poly dT primers (Amersham Pharmacia). Amplifications were done specific primers as described in Supporting Text.