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Published as doi: 10.1096/fj.06-7070com.
 (The FASEB Journal. 2007;21:3338-3345.)
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Inhibition of myoblast migration by prostacyclin is associated with enhanced cell fusion

Brenda A. Bondesen*, Kristen A. Jones*, Wayne C. Glasgow{dagger} and Grace K. Pavlath*,1

* Department of Pharmacology, Emory University, Atlanta, Georgia, USA; and

{dagger} Division of Basic Medical Sciences, Mercer University School of Medicine, Macon, Georgia, USA

1Correspondence: Emory University School of Medicine, Department of Pharmacology, Rm. 5027, O.W. Rollins Research Bldg., Atlanta, GA 30322, USA. E-mail: gpavlat{at}emory.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Satellite cells are stem cells that are critical for the formation and growth of skeletal muscle during myogenesis. To differentiate and fuse, proliferating satellite cells or myoblasts must migrate and establish stable cell-cell contacts. However, the factors that regulate myoblast migration and fusion are not understood completely. We have identified PGI2 as a novel regulator of myogenesis in vitro. PGI2 is a member of the family of prostaglandins (PG), autocrine/paracrine signaling molecules synthesized via the cyclooxygenase-1 and -2 pathways. Primary mouse muscle cells both secrete PGI2 and express the PGI2 receptor, IP, at various stages of myogenesis. Using genetic and pharmacological approaches, we show that PGI2 is a negative regulator of myoblast migration that also enhances cell fusion. Thus, PGI2 may act as a "brake" on migrating cells to facilitate cell-cell contact and fusion. Together, our results highlight the importance of the balance between positive and negative regulators in cell migration and myogenesis. This work may have implications for migration of other populations of adult stem cells and/or cells that undergo fusion.—Bondesen, B. A., Jones, K. A., Glasgow, W. C., Pavlath, G. K. Inhibition of myoblast migration by prostacyclin is associated with enhanced cell fusion.


Key Words: prostaglandins • myogenesis • muscle repair


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
SKELETAL MUSCLE GROWTH AND REPAIR depend on myogenesis, an ordered series of cellular events comprising the proliferation, migration, differentiation, and fusion of muscle precursor cells called myoblasts. This process facilitates the formation and growth of myofibers in vivo, as in regeneration following muscle injury, and the formation of multinucleated myotubes in vitro. The addition of myonuclei to growing myofibers leads to increased protein synthesis and cell size. Greater understanding of the molecular regulation of myogenesis is critical for the development of therapies to treat the loss of muscle mass associated with muscle injuries, degenerative disorders, and aging.

One class of molecules implicated in various stages of myogenesis is prostaglandins (PG), autocrine/paracrine signaling molecules derived from arachidonic acid (AA). PG synthesis involves the phospholipase A2-dependent release of AA from membrane phospholipids, conversion of free AA to the common PG precursor PGH2, and subsequent isomerization of PGH2 to one of five bioactive PG (PGE2, PGF2 {alpha}, PGI2, PGD2, and TXA2) by specific PG synthases (1) . The conversion of AA to PGH2 is the rate-limiting step in PG synthesis and is catalyzed by one of two isoforms of cyclooxygenase (COX), COX-1, or COX-2. PG bind to specific G-protein-coupled receptors and elicit diverse physiological effects, including changes in intracellular Ca2+ and cAMP, which are often cell type specific (1) .

PG, which are synthesized by muscle cells (2 3 4 5) , regulate myoblast proliferation (2 , 6) , differentiation (7) , and fusion (8 9 10 11 12) in vitro. PGF2{alpha} regulates the fusion of myoblasts with existing myotubes in vitro (10) and has been implicated in the regulation of protein synthesis both in vivo and in vitro (3 , 4 , 13 , 14) . In contrast, PGE2 has been implicated in muscle protein degradation in certain types of muscle repair (15 16 17 18 19) . Together, these studies identify PG as an important class of myogenic regulators.

PGI2, or prostacyclin, is a potent vasodilator and anticlotting agent that acts through the IP receptor, a G-protein-coupled receptor, to increase intracellular cAMP (1) . PGI2 is the most abundant PG released by excised whole muscles (16 , 20 , 21) . Although one report implicated PGI2 in myofiber formation within developing chick muscle (22) , the potential role of PGI2 in myogenesis is virtually unknown. Here we show that isolated muscle cells secrete PGI2 and express IP receptor mRNA at various stages of myogenesis in vitro. IP–/– myoblasts exhibited enhanced motility but impaired differentiation and fusion. Conversely, addition of the stable PGI2 analog iloprost to wild-type (WT) myoblasts decreased cell migration but enhanced cell fusion. Together, our results identify PGI2 as a novel regulator of myogenesis, potentially acting as a "brake" to facilitate cell-cell contact and subsequent fusion.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell culture
Primary myoblast cultures were prepared from the hind limb muscles of adult Balb/c mice (for IP RT-PCR) or C57B6;129P2 mice (for HPLC) based on methods already described except for omission of the Percoll gradient (23 , 24) . For fusion, migration, and 5-bromo-3'-deoxyuridine (BrdU) assays, myoblasts were isolated from the hind limb muscles of adult WT and IP–/– mice (25) . Myoblasts were maintained as described (10) and culture purity (>96% muscle cells) was determined by MyoD immunostaining. To induce differentiation/fusion, cells were seeded in growth medium (GM: Ham’s F10, 20% fetal bovine serum, 5 ng/ml bFGF, 100 U/ml penicillin G, and 100 µg/ml streptomycin) onto plates coated with entactin-collagen-laminin (ECL, Upstate Biotechnology, Lake Placid, NY, USA), allowed to adhere for ~1 h, then switched to differentiation media (DM: DMEM, 100 U/ml penicillin G, 100 µg/ml streptomycin, and either 1% insulin-transferrin-selenium-A supplement (ITS, Invitrogen, Carlsbad, CA, USA) or 2% horse serum). For fixation, cells were incubated in 3.7% formaldehyde for 10 min.

For migration experiments, conditioned media (CM) was used to induce migration. To prepare CM, myoblasts were incubated in ITS differentiation medium (referred to as control medium) for 24 h. The media, which had been enriched or "conditioned" with secreted factors, was then collected, filtered (0.45 µm), flash frozen, and stored at –80°C until use.

Measurement of muscle cell 3H-prostaglandin production by HPLC
C57BL/6;129P2 myoblasts were seeded onto three ECL-coated 100 mm dishes (1.2x106 cells/dish). Cells were either maintained as myoblasts or allowed to differentiate for 24 or 48 h. Media were then switched to 10 ml serum-free DMEM (F10 for myoblasts) containing 5 µCi 3H-arachidonic acid (98.6 Ci/mmol; Perkin Elmer, Norwalk, CT, USA) and 3 µg/ml unlabeled AA (BIOMOL Research Laboratories, Plymouth Meeting, PA, USA) for 4 h. The calcium ionophore A23187 (0.5 µg/ml, Sigma) was then added to the media for an additional 30 min to enhance AA release from cell membranes. Media were collected, combined with 10 ml 100% methanol and 1 ml glacial acetic acid, and stored at –20°C prior to analysis.

HPLC analysis was based on methods described earlier (26) . Briefly, AA metabolites were extracted from media samples using C18 PrepSep columns (Fisher, Pittsburgh, PA, USA) according to the manufacturer’s instructions and reconstituted in 750 µl 50% methanol. For analysis, 250 µl of each sample was used for reverse-phase HPLC using a C18-Ultrasphere column as the stationary phase. Metabolites were resolved using a mobile phase of 55–75% methanol (pH 4.0) in a stepwise gradient at a flow rate of 1.1 ml/min. PG products were identified by comparing the retention time of radiolabeled peaks to that of authentic PG standards.

RNA isolation and RT-PCR analysis
Balb/c myoblasts were seeded onto 6-well plates (2x105 cells/well) and allowed to differentiate for 8, 16, 24, 36, or 42 h. Total RNA was isolated at each time point using Trizol Reagent (Life Technologies, Gaithersburg, MD, USA) according to the manufacturer’s protocol. Myoblast RNA was collected after 24 h in GM. Total RNA (2.5 µg) was reversetranscribed using random hexamers and M-MLV reverse transcriptase (Invitrogen). Reactions were incubated at 25°C for 10 min, at 42°C for 50 min, then at 72°C for 10 min to inactivate the reverse transcriptase. cDNA (2 µl) was amplified in a 50 µl reaction containing 5 µM IP receptor primers (F: 5'-CATGGCTCGTTTGTACCGACCTC-3' and R: 5'-CAGAGGCACAGCAGTCAATGGTG-3', (27) or QuantumRNA 18S primers (Ambion, Austin, TX, USA) as a control. Kidney cDNA was used as a positive control for IP expression. Amplification cycles comprised 95°C for 5 min, 30 cycles of 95°C, 62°C, and 72°C for 30 s each, then 72°C for 5 min. PCR products were separated by electrophoresis through a 1% agarose gel and visualized by ethidium bromide staining.

Differentiation and fusion assays
WT and IP–/– myoblasts were seeded onto 6-well plates (2x105 cells per well) and allowed to differentiate for 18, 24, or 42 h before fixation. Cultures were immunostained for embryonic myosin heavy chain (EMyHC, F1.652, neat hybridoma supernatant, Developmental Studies Hybridoma Bank, Iowa City, IA, USA) as described previously (28) . Using a Zeiss Axiovert microscope equipped with a video camera and Scion Image software (version 1.63), the total number of nuclei and myotubes was counted in 10 random fields (>500 total nuclei) per condition. The fusion index (percentage of nuclei within myotubes) and differentiation index (percentage of nuclei within myotubes and EMyHC+ mononucleated cells) were determined.

BrdU assay
WT and IP–/– myoblasts were seeded onto 6-well plates (2x105 cells/well) and allowed to differentiate for 4, 8, 12, or 16 h. Myoblasts were maintained in GM and fixed 2 h after plating. Cells were incubated with 25 µM BrdU (Sigma, St. Louis, MO, USA) for 1 h prior to fixation. BrdU+ cells were quantified by immunostaining as described (24) except that cells were incubated in 1N HCl for 30 min prior to blocking. Using a Zeiss Axiovert microscope equipped with a video camera and Scion Image software (version 1.63), the percentage of BrdU+ nuclei in 10 random fields (>350 total nuclei) was determined for each condition.

PGI synthase (PGIS) immunostaining
C57BL/6;129P2 myoblasts were seeded onto 6-well plates (2x105 cells/well) and fixed immediately upon adhesion. After incubation in block buffer (TNB buffer, Perkin Elmer) for 1 h, cells were incubated with an antibody against PGIS (1:200; Abcam, Cambridge, MA, USA) in PBS overnight at 4°C. Antibody binding was visualized using the tyramide amplification system (Perkin Elmer) as described for COX-2 (29) except for the concentrations of biotin-conjugated donkey-anti-rabbit (1:800, Jackson ImmunoResearch Laboratories Inc., West Grove, PA, USA) and fluorescein tyramide (1:400). No staining was observed when the PGIS antibody was replaced with control rabbit IgG (Jackson Immunoresearch).

Cell migration assays
Migration of WT and IP–/– myoblasts was quantified using time-lapse photography as described previously (30) . Briefly, myoblasts were seeded on 35 mm plates (2x105 cells/plate) and incubated in DM for 30 min. Cultures were then supplemented with HEPES (25 mM) and transferred to a microscope stage heated to 37°C. Images were recorded every 6 min for 1 h using an Axiovert 200M microscope with a 0.3 NA 10x Plan-Neofluar objective (Carl Ziess Microimaging, Inc., Thornwood, NY, USA). Cell velocities (µm/h) were determined by tracking the paths of individual cells using ImageJ software (version 1.36b). For each genotype, the paths of 19–20 cells from three independent isolates were analyzed for a total of 58 cell paths.

Cell migration was also analyzed using a 96-well Boyden chamber assay with a polycarbonate filter (8 µM pores; Neuro Probe, K.U. Leuven, Belgium) based on described methods, with some modifications (31) . In a Boyden chamber, cell migration is measured as the number of cells that migrate from the top side of the filter through the pores to the bottom side in response to media in the lower chamber. Briefly, the lower wells of each chamber were loaded with 395 µl of control medium (to evaluate basal migration) or CM (to induce migration). Myoblasts (7.5x104) in 200 µl control medium were loaded into each upper chamber and incubated at 37°C. In some experiments, cells were loaded in control medium containing iloprost (0.01 or 0.1 µM) or vehicle (0.2% ethanol). All conditions were tested in triplicate wells. After 5 h, the filter was wiped clean of nonmigrated cells, fixed in 100% methanol for 5 min, and incubated in Gill-2 hematoxylin (Thermo Electron Corp., Pittsburg, PA, USA) for 6–12 h to stain migrated cells. The total number of cells in 10 fields (25x objective, Ziess Axiovert microscope) per well was determined.

Statistics
To determine significance between two groups, comparisons were made using the Student’s t test. Data from multiple groups with one variable parameter were analyzed by 1-way ANOVA, followed by the Newman-Keuls post-test using GraphPad Prism version 4.0a (GraphPad Software). Data from multiple groups with two variable parameters were analyzed by 2-way ANOVA, followed by the Newman-Keuls post-test using SigmaStat 2.03 (SPSS). For all statistical tests, P < 0.05 was accepted for significance.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Muscle cells synthesize essential components of the prostacyclin signaling pathway
Many studies have demonstrated that PGI2 is the most abundant PG product released by excised fetal and adult muscles (16 , 20 , 21) . However, whether components of the prostacyclin signaling pathway are present within muscle cells themselves is unknown. We first used RT-PCR to determine whether isolated muscle cells express IP receptor mRNA. As shown in Fig. 1 A, primary mouse muscle cells expressed IP receptor mRNA at various stages of myogenesis, with a decrease in IP mRNA levels during later stages. We then used immunostaining to demonstrate that myoblasts express PGI synthase (PGIS, Fig. 1B ). Finally, reverse-phase HPLC was used to measure PG secretion into culture media by proliferating myoblasts and during early (24 h) and late (48 h) stages of differentiation/fusion. As shown in Fig. 1C , muscle cells synthesize PGI2, PGE2, and PGF2{alpha} throughout myogenesis, with PGI2 (measured as 6-keto PGF1{alpha}) being the predominant product. These results demonstrate that muscle cells secrete PGI2 and express the PGI2 receptor, IP, suggesting that muscle cells can respond to PGI2 via both autocrine and paracrine mechanisms.


Figure 1
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  		Figure 1. Primary muscle cells express the IP receptor and secrete PGI2. A) IP receptor mRNA was observed throughout myogenesis but decreased during later stages of fusion. 18S rRNA was used as an internal control and kidney mRNA was used as a positive control for IP receptor expression (+). Data are representative of three independent experiments. Phase-contrast images of muscle cells at the corresponding times during myogenesis are shown (bar=100 µm). B) Fluorescent and phase-contrast images of primary myoblasts immunostained for PGI synthase (PGIS, green). Nuclei were counterstained with DAPI (blue). No staining was observed when a rabbit IgG control antibody was used in place of the primary antibody (IgG). Bar = 50 µm. C) To quantify PG production during myogenesis, the release of 3H-PG into culture media was measured by reverse-phase HPLC. PGI2, along with smaller amounts of PGE2 and PGF2{alpha}, was secreted by proliferating myoblasts and during early (24 h) and late (48 h) stages of myoblast differentiation/fusion. The stable PGI2 metabolite 6-keto PGF1{alpha} was used as a measure of PGI2 production. Similar results were obtained in two independent experiments.

IP–/– myoblasts exhibit decreased differentiation and fusion
The decrease in IP receptor mRNA levels observed during late stages of myogenesis suggests that IP may regulate early stages of myoblast differentiation and/or fusion. To determine whether the IP receptor regulates myoblast fusion, WT and IP–/– myoblasts were differentiated for 18 or 24 h. As shown in Fig. 2 A, B, the fusion index of IP–/– cells was decreased by 37% at 24 h relative to WT. IP–/– fusion was also decreased at 18 h, but this difference did not achieve statistical significance. Similarly, IP–/– myoblasts formed fewer myotubes even when differentiated for longer periods (Fig. 2C ). However, the increase in IP–/– myotube number over time was parallel to that of WT myotubes, suggesting that the initial formation of myotubes (i.e., myoblast-myoblast fusion) was impaired, but additional myoblast-myotube fusion proceeded normally.


Figure 2
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  		Figure 2. Impairment of fusion in IP–/– myoblasts. A) Representative phase-contrast images of WT and IP–/– myotubes (24 h) immunostained for EMyHC. Bar = 60 µm. B) The fusion index of IP–/– muscle cells was decreased by 37% relative to WT after 24 h in DM. C) IP–/– myoblasts formed fewer myotubes than WT at all time points examined. Data are mean ± SE, n = 4 independent cell isolates for each genotype. *P < 0.05.

Decreased myoblast fusion may indicate a defect in the ability of cells to fuse but can also result from decreased myoblast differentiation. Therefore, we examined WT and IP–/– myoblast differentiation by immunostaining for EMyHC. As shown in Fig. 3 A, the differentiation index was decreased by 18% in IP–/– cells at 18 h. At 24 and 42 h, however, WT and IP–/– differentiation did not differ, suggesting that the defect in differentiation was transient. Given that PGI2 has been implicated in cell cycle withdrawal of other cell types (32 33 34) , we used BrdU labeling to determine whether the delayed differentiation of IP–/– muscle cells was associated with impaired cell cycle withdrawal. In contrast to differentiation, the percentage of BrdU+ cells did not differ between WT and IP–/– cells (Fig. 3B ). Together, these results suggest that IP receptor signaling regulates myoblast differentiation independently of cell cycle withdrawal.


Figure 3
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  		Figure 3. IP–/– myoblasts exhibit impaired differentiation but normal cell cycle withdrawal. A) Differentiation was quantified by immunostaining for EMyHC. IP–/– differentiation was decreased 18% after 18 h in DM but did not differ significantly from that of WT muscle cells at later time points. B) Cell cycle withdrawal was analyzed using BrdU labeling. The percentage of BrdU+ cells did not differ between WT and IP–/– myoblasts at various time points during early differentiation. Data are mean ± SE, n = 4 independent cell isolates for each genotype. Mb = myoblasts in GM.

Enhanced motility of IP–/– myoblasts
Evidence suggests that cell-cell contact positively regulates myoblast differentiation in vitro (35 , 36) . As cell-cell contact depends on cell migration, alterations in cell migration may adversely affect myoblast differentiation and fusion. We hypothesized that decreased migration of IP–/– myoblasts may be one mechanism leading to decreased differentiation and fusion. To test this, we used time-lapse microscopy to track the migratory paths of individual WT and IP–/– myoblasts within the first hour in DM. Surprisingly, IP–/– cells migrated farther from their point of origin than WT, and a greater proportion of IP–/– cells migrated at greater velocities than WT (Fig. 4 A, B). In addition, the mean IP–/– cell velocity increased 35% over WT (Fig. 4C ). Similarly, IP–/– cell migration was increased 2.8-fold over WT under basal conditions in a Boyden chamber assay (Fig. 4D ). We then used CM, which contains chemotactic factors (30) , in the bottom chamber to evaluate the response of WT and IP–/– cells to a chemotactic gradient. As shown in Fig. 5 A, the same total number of WT and IP–/– muscle cells migrated in response to CM. However, this result reflects a 10-fold increase over basal in WT cells and only a 4-fold increase in IP–/– cells. Together, these results suggest that IP–/– myoblasts exhibit increased intrinsic motility but may have an impaired ability to sense and/or respond to chemotactic gradients.


Figure 4
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  		Figure 4. Enhanced motility of IP–/– myoblasts. A) Graphical representation of the migratory paths of WT (+/+) and IP–/– (–/–) myoblasts over 1 h, with each cell’s point of origin set to (0,0). The paths of 30 cells (10 cells from each of 3 independent isolates) are shown for each genotype. B) Frequency distribution of the velocities of WT and IP–/– myoblasts. C) The average velocity of IP–/– myoblasts was 35% greater than that of WT cells. Data are mean ± SE, n = 58 total cells from 3 independent isolates for each genotype. *P < 0.05. D) Migration of WT and IP–/– myoblasts was analyzed in a Boyden chamber. Under basal conditions (i.e., in the absence of a chemotactic gradient, with ITS in both top and bottom chambers), 2.8-fold more IP–/– cells migrated through the porous membrane than WT. Data are mean ± SE, n=3–4 independent cell isolates for each genotype. *P < 0.05.


Figure 5
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  		Figure 5. The stable PGI2 analog iloprost attenuates myoblast migration and enhances fusion. A) CM increased WT and IP–/– migration in a Boyden chamber assay. B) CM-induced migration of WT muscle cells was attenuated 23% by iloprost (ILO, 0.1 µM). Data are mean ± SE, n = 4 independent cell isolates. *P < 0.05. C) Representative phase-contrast images of WT myotubes (24 h) treated with either vehicle (V) or 0.1 µM iloprost (ILO). Bar = 60 µm. D) Iloprost (0.01 and 0.1 µM) increased the fusion index of WT myoblasts at 24 h. E) Iloprost had no effect on fusion of IP–/– myoblasts. Data are mean ± SE, n = 3 independent cell isolates. *P < 0.05.

Iloprost decreases myoblast chemotaxis but enhances fusion
To examine further the role of the IP receptor in migration and fusion, we determined the effect of iloprost, a stable PGI2 analog, on WT myoblast migration and fusion. Iloprost (0.1 µM) inhibited CM-induced migration of WT muscle cells by 23% (Fig. 5B ). Together, our results from pharmacologic and genetic studies suggest that signaling through the IP receptor negatively regulates myoblast motility. In light of the decreased fusion of IP–/– muscle cells, we hypothesized that iloprost would enhance fusion. As shown in Fig. 5C, D , the addition of 0.01 or 0.1 µM iloprost to fusing cultures for 24 h increased the fusion of WT myoblasts by 23% and 29%, respectively. The fusion of IP–/– myoblasts was unaffected by iloprost, suggesting that this drug is acting specifically through the IP receptor (Fig. 5E ). Together, these results implicate PGI2 as both a negative regulator of myoblast motility and a positive regulator of myoblast fusion, suggesting a potential inverse correlation between cell motility and fusion.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In this investigation, we show that PGI2 has both antimigratory and promyogenic effects on myoblasts. Cell migration is essential for muscle development, during which muscle precursor cells migrate from the somites to the limb musculature, and for adult muscle repair, when myoblasts migrate and fuse with each other or existing myofibers to rebuild muscle architecture. Several factors have been identified as positive regulators of myoblast migration, including growth factors, chemokines, cytokines, and extracellular matrix proteins. Hepatocyte growth factor (HGF), which binds to c-Met, is a promitogenic factor for myoblasts in vitro (37) and regulates myoblast migration both in vitro (38 , 39) and during development (40) . Other promigratory factors that have been studied extensively include basic fibroblast growth factor (bFGF), leukemia inhibitory factor (LIF), transforming growth factor-ß (TGFß), and insulin-like growth factor-1 (IGF-1) (38 , 39 , 41 , 42) .

In contrast to promigratory factors, little is known about antimigratory factors in myogenesis. PGI2 is one of only three antimigratory factors for myoblasts identified to date. Decorin-null C2C12 myoblasts exhibited decreased differentiation and a wider dispersion pattern than control C2C12 myoblasts both in vitro and after transplantation into developing chick limbs, which was attributed to alterations in migration (43) . Reintroduction of decorin expression into decorin-null myoblasts via adenoviral infection restored both migration and differentiation to WT levels, suggesting that myoblast differentiation and migration are closely linked processes. A second antimigratory factor, sphingosine-1-phosphate (S1P), was recently identified. In a Boyden chamber model, S1P impaired the migration of C2C12 myoblasts and abrogated the response to IGF-1, a known myoblast chemoattractant (44) . The antimigratory effect of S1P was attributed to S1P binding to the G-protein-coupled receptor (GPCR) S1P2, leading to an increase in intracellular calcium (44) . S1P treatment also enhanced differentiation and myotube formation in C2C12 cultures, although no quantitative studies were performed (45) . The antimigratory, promyogenic effects of S1P are similar to those of iloprost treatment described in the present study. Our identification of PGI2 as a negative modulator of myoblast migration further highlights the importance of both positive and negative factors during myogenesis, the coordination of which warrants further examination.

PG have been implicated in various stages of myogenesis in vitro, including myoblast proliferation (2 , 6) , differentiation (7) , and fusion (8 9 10 11 12) . However, a role for PG in muscle cell migration has not been demonstrated before. We have shown that PGF2{alpha} regulates the fusion of myoblasts with nascent myotubes in vitro (10) . In contrast to PGF2{alpha}, our results suggest that PGI2 regulates initial myotube formation via myoblast-myoblast fusion and does not appear to affect later myoblast-myotube fusion. Together, these results highlight the importance of the balance between different PGs acting at distinct stages of myogenesis.

Several studies support the role of PGI2 as an antimigratory factor. In a Boyden chamber model, iloprost inhibited the migration of neutrophils (46) , vascular smooth muscle cells (47) , vascular endothelial cells (48) , and fibroblasts (49) . PGI2 may modulate cell migration directly by altering adhesion molecule expression and/or cytoskeletal rearrangement, which could adversely affect cell adhesion to the extracellular matrix (ECM) or cell motility. For example, iloprost inhibited the migration of human aortic smooth muscle cells by inducing disassembly of actin filaments and focal adhesions (50) . Moreover, treatment of human umbilical vein endothelial cells with PGI2 increased cell-cell contacts and cell adhesion to VE-cadherin-coated plates, which was attributed to increased VE-cadherin expression and increased cortical actin rearrangement (51) . In contrast, Lindemann et al. showed that iloprost decreased the adhesion of human neutrophils to an endothelial cell monolayer in vitro and also decreased neutrophil chemotaxis toward endothelial cells in a Boyden chamber model (48) . However, these effects were independent of changes in adhesion molecule expression. Together, these results suggest that PGI2 can modulate cell adhesion and migration via adhesion molecule-dependent and -independent mechanisms.

Our results implicate PGI2 in both myoblast migration and fusion. The enhanced migration of IP–/– muscle cells correlated with decreased fusion, whereas the PGI2 analog iloprost decreased migration but enhanced fusion. However, whether enhanced fusion of PGI2-treated cells is merely a consequence of changes in cell migration or a direct result of PGI2 signaling is unclear. Prior to this study, the potential role of PGI2 in cell fusion had not been examined. One mechanism by which PGI2 may regulate cell fusion directly is the alteration of cell-cell adhesion via changes in adhesion molecules. Alternatively, PGI2-dependent changes in cell migration may modulate fusion indirectly by affecting myoblast differentiation. Recent evidence suggests that the initiation of myoblast differentiation requires cell-cell contact (36 , 52) , which depends on cell-cell and cell-extracellular matrix adhesions as well as cytoskeletal rearrangements that regulate cell migration (53) . Thus, increased motility of IP–/– muscle cells may decrease the frequency of stable cell-cell contacts, leading to the decreases observed in differentiation and subsequent fusion. During myogenesis, PGI2 may act as a "brake" to promote fusion by decreasing motility and therefore increasing the probability of stable cell-cell contacts. However, iloprost treatment enhanced myoblast fusion (Fig. 5C, D ) but had no effect on differentiation (unpublished observations), suggesting that PGI2 may regulate myoblast fusion independently of effects on differentiation.

In summary, myogenesis is influenced by both positive and negative regulators of cell migration. The net balance between these two classes of migratory regulators would be critical for myogenesis. Thus, cell migration during differentiation and fusion may be modulated not only by increasing positive regulators but also by decreasing antimigratory factors. The notion that migration can be enhanced by mitigating antimigratory signals may have implications for cell-based therapies for muscular dystrophies and other stem cell transplantation therapies. Currently, cell transplantation is complicated by inefficient migration of donor cells into and within muscle tissue. Recently, Hill et al. showed that codelivery of the promigratory factors HGF and FGF2 enhanced the migration of transplanted myoblasts into host muscle tissue (54) . In addition, pretreatment of mesangioblasts with promigratory cytokines increased engraftment and gene delivery to dystrophic muscles (55) . Our results suggest that mitigating antimigratory signals in muscle tissue may have similar benefits for enhancing donor cell migration. Our identification of PGI2 as a myogenic regulator provides further insight into the mechanisms that govern muscle cell migration and fusion, and may shed light on common pathways that regulate fusion in other cell types, inflammation, and stem cell homing.


   ACKNOWLEDGMENTS
 
We thank Dr. Garret FitzGerald for his generous gift of WT and IP–/– mouse muscle tissue. This work was supported by National Institutes of Health Grants AR47314, AR48884, AR052730, and AR051372 to G.K.P. B.A.B. was supported by American Heart Association predoctoral fellowship 0415084B.

Received for publication November 3, 2006. Accepted for publication April 12, 2007.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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