Abstract
Satellite cells are the myogenic precursors in postnatal muscle and are situated beneath the myofiber basement membrane. We previously showed that fibroblast growth factor 2 (FGF2, basic FGF) stimulates a greater number of satellite cells to enter the cell cycle but does not modify the overall schedule of a short proliferative phase and a rapid transition to the differentiated state as the satellite cells undergo myogenesis in isolated myofibers. In this study we investigated whether other members of the FGF family can maintain the proliferative state of the satellite cells in rat myofiber cultures. We show that FGF1, FGF4, and FGF6 (as well as hepatocyte growth factor, HGF) enhance satellite cell proliferation to a similar degree as that seen with FGF2, whereas FGF5 and FGF7 are ineffective. None of the growth factors prolongs the proliferative phase or delays the transition of the satellite cells to the differentiating, myogenin+ state. However, FGF6 retards the rapid exit of the cells from the myogenin+ state that routinely occurs in myofiber cultures. To determine which of the above growth factors might be involved in regulating satellite cells in vivo, we examined their mRNA expression patterns in cultured rat myofibers using RT-PCR. The expression of all growth factors, excluding FGF4, was confirmed. Only FGF6 was expressed at a higher level in the isolated myofibers and not in the connective tissue cells surrounding the myofibers or in satellite cells dissociated away from the muscle. By Western blot analysis, we also demonstrated the presence of FGF6 protein in the skeletal musle tissue. Our studies therefore suggest that the myofibers serve as the main source for the muscle FGF6 in vivo. We also used RT-PCR to analyze the expression patterns of the four tyrosine kinase FGF receptors (FGFR1-FGFR4) and of the HGF receptor (c-met) in the myofiber cultures. Depending on the time in culture, expression of all receptors was detected, with FGFR2 and FGFR3 expressed only at a low level. Only FGFR4 was expressed at a higher level in the myofibers but not the connective tissue cell cultures. FGFR4 was also expressed at a higher level in satellite cells compared to the nonmyogenic cells when the two cell populations were released from the muscle tissue and fractionated by Percoll density centrifugation. The unique localization patterns of FGF6 and FGFR4 may reflect specific roles for these members of the FGF signaling complex during myogenesis in adult skeletal muscle. (J Histochem Cytochem 48:1079–1096, 2000)
Keywords
Satellite cells are the myogenic precursors in postnatal and adult muscle. At least some of the satellite cells are mitotically active in the growing muscle, contributing myonuclei to the enlarging fibers. As muscle matures, the addition of myofiber nuclei ceases and the satellite cells become mitotically quiescent. Recruitment of these quiescent precursors occurs in response to overt muscle injury and to more subtle stresses such as stretch, exercise, and muscle hypertrophy. Depending on the degree of injury, the satellite cell progeny may fuse with existing myofibers for localized repair or may fuse with each other to form new fibers when the damage is more extensive (reviewed in Grounds and Yablonka–Reuveni 1993; Schultz and McCormick 1994; Bischoff 1994; Yablonka–Reuveni 1995).
Satellite cells undergoing proliferation and differentiation express the skeletal muscle-specific transcription factors Myf5, MyoD, myogenin, and MRF4. These four myogenic regulatory factors (MRFs) are first expressed during early embryogenesis, when they are involved in the specification of the myogenic lineage (Ludolph and Konieczny 1995; Arnold and Winter 1998). The activated satellite cells initially express Myf5 and MyoD and, correlating with cell cycle withdrawal and differentiation, express myogenin and MRF4 (Grounds et al. 1992; Smith et al. 1994; Yablonka-Reuveni and Rivera 1994; Cornelison and Wold 1997; Cooper et al. 1999). Differentiating satellite cells also express at least one member of the myocyte enhancer factor 2 (MEF2) family (Yablonka-Reuveni et al. 1999a). The MEF2s and MRFs act cooperatively during the activation of muscle-specific genes (Molkentin et al. 1995; Black and Olson 1998).
Cell culture studies have revealed many growth factors and cytokines which can affect proliferation and differentiation of satellite cells (reviewed in Grounds and Yablonka-Reuveni 1993). Among these factors are members of the fibroblast growth factor (FGF) family, whose effects on myogenesis of satellite cells have been investigated extensively both in primary satellite cell cultures and in cell lines derived from satellite cells (Allen et al. 1984; Clegg et al. 1987; Olwin and Rapraeger 1991; Smith and Schofield 1994; Pizette et al. 1996; Yoshida et al. 1996; Yablonka–Reuveni and Rivera 1997a). The detection of various FGFs in muscle tissue and in myogenic precursors further suggests that the FGFs may affect the cells involved in myogenesis via a paracrine or an autocrine mechanism (Groux-Muscatelli et al. 1990; Garrett and Anderson 1995; Lefaucheur and Sebille 1995; Hannon et al. 1996; Floss et al. 1997).
To gain insight into the regulation of satellite cells, we have been studying the dynamics of these cells in cultures of isolated myofibers. In such cultures the satellite cells remain situated in their native position under the basement membrane of the myofibers (Bischoff 1986; Yablonka-Reuveni and Rivera 1994; Yablonka–Reuveni et al. 1999a,b). The dynamics of satellite cells in the isolated myofiber system may model myogenesis in an environment in which the muscle structure is preserved, such as during muscle growth and hypertrophy. The more commonly used culture system of isolated satellite cells may more closely model events after overt muscle trauma where new myofibers are formed.
Analyzing adult rat myofibers, we previously showed that satellite cells first become positive for PCNA and MyoD on their activation and entry to the cell cycle. After a short proliferative period, the cells become negative for PCNA and MyoD and enter the differentiated, myogenin+ state (Yablonka-Reuveni and Rivera 1994, 1997b). FGF2 (basic FGF) causes a greater number of satellite cells to enter the cell cycle, enhancing the number of PCNA+/MyoD+ satellite cells in myofiber cultures. However, the mitogenic influence of FGF2 is limited to the initial days in culture and the schedule of transit from proliferation to differentiation is not changed in the presence of this mitogen (Yablonka–Reuveni and Rivera 1994, 1997b; Yablonka-Reuveni et al. 1999b).
The short-term effect of FGF2 seen in myofiber cultures was unexpected in view of the commonly held convention that FGF can maintain ongoing proliferation of myoblasts (Clegg et al. 1987). This promoted the present study on whether other members of the FGF family can support satellite cell proliferation in myofiber cultures for a longer length of time than FGF2. To date, over 20 FGF and FGF-related genes have been discovered in mammals (reviewed in Goldfarb 1997; Szebenyi and Fallon 1999). In this study we focused specifically on FGF1-FGF7 (excluding FGF3) because these FGFs were shown to be expressed in the muscle and/or in myogenic cultures (Anderson et al. 1991; Haub and Goldfarb 1991; Niswander and Martin 1992; Oliver et al. 1992; Hughes et al. 1993; deLapeyriere et al. 1993; Mason et al. 1994; Garrett and Anderson 1995; Hannon et al. 1996; Floss et al. 1997; Moscoso et al. 1998). We also investigated whether these specific members of the FGF family are indeed expressed in myofiber cultures and in cultures of isolated satellite cells.
A second aspect of the study was to analyze the expression patterns of the FGF receptors (FGFRs) during satellite cell myogenesis. The FGFs transduce their signals to the cell through transmembrane tyrosine kinase receptors (FGFRs), for which four distinct genes have been discovered (FGFR1-FGFR4). Further receptor diversity through alternative splicing of pre-mRNAs has been documented for FGFR1-FGFR3. Each FGFR variant has a distinct, although sometimes overlapping, profile of affinities to different FGFs (Ornitz et al. 1996; reviewed in Szebenyi and Fallon 1999). Therefore, it is conceivable that specific FGFs may exert unique influences on cells and tissues, depending on which FGFRs they interact with. In the case of skeletal muscle, the myogenic and nonmyogenic cells might express common and distinct FGFRs, possibly resulting in overlapping and unique effects of FGFs.
Along with the analysis of the FGF-FGFRs we also conducted studies on the effect and expression of hepatocyte growth factor (HGF) and its transmembrane receptor (c-met) during myogenesis of satellite cells. The c-met receptor was shown to be expressed by proliferating satellite cells, and it has been suggested that HGF has a unique role during recruitment of quiescent satellite cells, shortening the time the cells spend before entering the cell cycle (Allen et al. 1995; Cornelison and Wold 1997; Tatsumi et al. 1998).
Collectively, the present studies on the response of the satellite cells to the different growth factors, along with the gene expression analysis, were expected to enable the determination of which of these growth-promoting agents might play an actual physiological role in the muscle.
Materials and Methods
Animals
Male Sprague-Dawley rats (B & K Universal; Kent, WA) were used throughout the study. Unless otherwise noted, the rats were 8–10 weeks old (young adults).
Isolation and Culture of Muscle Fibers
The procedures involved in the isolation and culture of the myofibers have been previously published (Yablonka–Reuveni and Rivera 1994; Yablonka–Reuveni et al. 1999b). In brief, myofibers were isolated from the flexor digitorum brevis (FDB) muscle of the rat hindfeet employing collagenase and using one or two rats per each preparation. The final fiber preparation was dispensed into Vitrogen-coated 35-mm culture plates and the cultures received 1 ml of basal medium [MEM containing 20% controlled process serum replacement (CPSR2; Sigma, St Louis, MO)], 1% horse serum (Sigma), and streptomycin and penicillin each at 102 U/ml). Growth factors (at concentrations described in Results) with or without heparin (at 10 μg/ml) were added to the medium when the cultures were established and were replaced along with the medium every 24 hr. Human recombinant FGF2 was kindly provided by Dr. S. Hauschka (University of Washington). All other growth factors (recombinant human) were from R & D Systems (Minneapolis, MN). Heparin was from Sigma.
Isolation and Culture of Muscle Connective Tissue Cells
The connective tissue surrounding the myofibers is routinely separated from the myofibers and discarded after the collagenase digestion of the FDB muscle (Yablonka-Reuveni et al. 1999b). This connective tissue material was transferred into MEM containing 10% horse serum and was used for preparation of connective tissue cell cultures. Some myofibers were carried over but most of them were removed with a micropipet. We confirmed by light microscopy and immunostaining that this connective tissue material contains individual cells (fibroblast-like cells, macrophages) and capillaries. The connective tissue suspension was triturated with a 20-gauge needle to release single cells and to damage leftover myofibers. The cells were collected by low-speed centrifugation, resuspended in basal medium, and cultured in Vitrogen-coated 35-mm culture plates under identical conditions as the myofibers. FGF2 was added to the basal medium at 2 ng/ml to enhance cell proliferation. Typically, connective tissue cells from one rat were cultured into 7–10 plates.
Isolation and Culture of Satellite Cells
To focus specifically on satellite cells apart from the whole myofiber, we isolated satellite cells from the combined extensor digitorum longus (EDL) and tibialis anterior (TA) muscles of the rat hindlimbs by Pronase digestion (Bischoff 1997). These muscles were used because they provide more satellite cells than the FDB muscle. The hindlimb EDL/TA muscles from one rat were used in each isolation. The muscles were cleaned of their connective tissue and tendons, minced, and digested with 0.1% Pronase (45 PUK/ml, product #53702; Calbiochem, La Jolla, CA) in MEM for 1 hr at 37C/5% CO2. The digested muscle was transferred into MEM containing 10% horse serum and triturated vigorously to release single cells. The cell suspension was then filtered through a double-layered lens paper to remove debris. The cells were collected by low-speed centrifugation, resuspended in 1.5 ml of MEM, and either cultured under standard conditions (1–2 × 105 cells per 35-mm plate coated with 2% gelatin; growth medium consists of DMEM containing 25% fetal bovine serum, 10% horse serum, and 1% chicken embryo extract, streptomycin and penicillin each at 102 U/ml) or fractionated by Percoll density centrifugation to enrich for nonmyogenic and myogenic cells (Yablonka–Reuveni and Nameroff 1987; Yablonka-Reuveni et al. 1988). In this study we used a five-step Percoll gradient as shown in Figure 1, adapting the centrifugation conditions described by Bischoff (1997). The fractions collected after centrifugation are indicated by the brackets at right in Figure 1. The cells recovered from these fractions were either harvested immediately for total RNA isolation or cultured at 5–10 × 104 cells per 35-mm culture plate using the same conditions described above for the nonfractionated cells.
Immunolabeling and Counting of Satellite Cells in Myofiber Cultures
Satellite cells in methanol-fixed myofiber cultures were monitored by immunofluorescence as previously described (Yablonka–Reuveni and Rivera 1994; Yablonka–Reuveni et al. 1999b). Cultures were reacted with a mouse monoclonal antibody against PCNA (MAb 19F4; Boehringer Mannheim, Indianapolis, IN) or against myogenin (MAb F5D; developed and kindly provided by Dr. W. Wright, University of Texas) and counterstained with DAPI to visualize all nuclei. The secondary antibody was a fluorescein-conjugated goat anti-mouse IgG. We showed that the PCNA+ or myogenin+ nuclei are within fiber-associated cells and are not myofiber nuclei (Yablonka-Reuveni et al. 1999b) and that the PCNA immunostaining provides a reliable means for monitoring proliferating satellite cells in myofiber cultures (Yablonka-Reuveni and Rivera 1997b).
Duplicate culture plates were used at each time point for cell quantification. Positive cells from 30 random fibers per plate were counted, averaged for the duplicate plates, and eventually expressed per 10 fibers. Error bars represent the range of the results within the duplicate plates. Analyses were typically repeated three to five times.
Schematic representation of the Percoll density fractionation of cells isolated from the combined EDL/TA muscles of rat hindlimbs. Cells were released from the Pronase-digested muscles by vigorous triturations. The final cell suspension (in 1.5 ml of MEM) was layered on top of a five-step Percoll gradient. The concentrations of the different Percoll steps are provided along with their volumes, which are shown at left. After centrifugation, the fractions were pooled from the top with a Pasteur pipet, as indicated at right by brackets. The top fraction contained a large amount of debris and was routinely discarded. The other seven fractions, designated by % Percoll as shown at right of the brackets, were saved for further cell analysis. The bottom fraction was enriched for red blood cells, which pelleted at the bottom of the test tube. The number of cells recovered from the different Percoll fractions varied, with the 40% and 40–55% fractions containing the highest cell number. Regardless of this difference in cell numbers, the cultures from the various Percoll fractions were initiated at a similar starting density of 5 × 104 to 1 × 105 cells per 35-mm plate.
Monitoring Myogenic Cells in Primary Cultures of Muscle-dissociated Cells
Methanol-fixed cultures of muscle-dissociated cells were analyzed by single and double immunofluorescence combined with DAPI staining of nuclei, using published procedures (Yablonka-Reuveni et al. 1999a). Cultures were reacted with a mouse MAb against MyoD (Pharmingen, San Diego, CA; MAb 5.8A) (Dias et al. 1992) or the anti-myogenin described above, followed by fluorescein-conjugated goat anti-mouse IgG. Alternatively, cultures were reacted via double immunofluorescence with the rabbit polyclonal antibody against MEF2A (Santa Cruz Biotechnologies; Santa Cruz, CA) in combination with the anti-MyoD, the anti-myogenin, or the MAb MF20, which recognizes all isoforms of sarcomeric myosin heavy chain (Developmental Studies Hybridoma Bank, University of Iowa) (Bader et al. 1982). Secondary antibodies were rhodamine-conjugated goat anti-rabbit IgG and fluorescein-conjugated goat anti-mouse IgG. All antibodies used have been previously described in greater detail (Yablonka-Reuveni and Rivera 1997a; Yablonka–Reuveni et al. 1999a).
Apoptosis Analysis
The possible involvement of programmed cell death (apoptosis) in the myofiber cultures was investigated using the TdT-mediated dUTP nick end-labeling (TUNEL) method, which visualizes apoptotic nuclei (Gavrieli et al. 1992). The in situ cell death reaction kit was from Boerhinger Mannheim. As a positive control, cultures were treated with DNase I (10 U/ml) for 15 min at room temperature before the TUNEL reaction to induce strand brakes. As a negative control, cultures were reacted with the TUNEL reaction omitting the terminal transferase. After the TUNEL reaction, the cultures were immunostained with a rabbit polyclonal antibody against the mitogen-activated protein kinases ERK1/ERK2, which facilitates the tracing of fiber-associated satellite cells (Yablonka-Reuveni et al. 1999a,b).
Analysis of Gene Expression by RT-PCR
Total RNA was isolated using the acid guanidinium-thiocyanate-phenol-chloroform method (Chomczynski and Sacchi 1987). The amount of total RNA was evaluated by spectro-photometry and the integrity was checked by gel electrophoresis. The reverse transcription was performed according to the manual of Promega (Madison, WI), using 500 ng total RNA, 1 mM dNTPs (Promega), 19 ng/μl oligo (dT)15 (Gibco BRL; Rockville, MD), 10 U ribonuclease inhibitor, and 50 U M-MLV reverse transcriptase (Promega) in a total volume of 50 μl. The synthesized first-strand cDNA was then diluted with an equal volume of 1 × RT buffer containing 1 mM dNTPs. The PCR was carried out according to the protocol of Qiagen (Valencia, CA), using 2 μl cDNA in a total volume of 20 μl. The final concentrations of the PCR components were 0.2 mM dNTPs, 1.8 mM MgCl2, 0.4 μM of each sense and antisense primer (custom ordered from Gibco BRL), and 0.5 U HotStar Taq DNA polymerase (Qiagen). The PCR cycling program consisted of 45 sec at 94C, 1 min at 65C, and 2 min at 72C. A final extension step was carried out for 10 min at 72C. The PCR products were separated on 1.5% agarose gels containing ethidium bromide and were visualized on a UV transilluminator.
Figure 2 summarizes the primers used for the study, along with the corresponding RT-PCR products of several rat tissues. Primers were designed to the rat gene sequence with the exception of the primers for Myf5, FGF6, and FGFR3, which were based on the mouse sequence because the rat sequence was not available (Graves and Yablonka–Reuveni in press). The same expression patterns were seen for these three genes in parallel rat and mouse studies (data not shown). Primer pairs for the FGF receptors were designed to recognize the splice forms within each receptor gene. The Myf5 and H-ras primers were the same as described by Smith et al. (1994). The RT-PCR product from the rat FGF6 was sequenced to verify its homology to the published mouse sequence. We concluded that the 152-bp product is 98% homologous to the corresponding mouse sequence with no mismatches in the primer regions.
The analysis of glyceraldehyde-3-phosphate dehydrogenase (GAPDH), ribosomal protein S6 (rpS6), or H-ras was used as a measure for ubiquitous gene expression in the various experiments. The primers for MyoD, FGFR4, and H-ras span an intron smaller than 1000 bp and the PCR products of these genes were used as control for the absence of genomic DNA. As a further control, we carried out the RT-PCR with H-ras primers at a high cycle (38) omitting the reverse transcriptase.
Primers used in RT-PCR.
The PCR products of total RNA isolated from muscle (FDB and EDL/TA) and nonmuscle tissues (brain and liver) that are shown in Figure 2 serve as indicators for the relative level of gene expression in the myofibers and the satellite cell cultures detailed in Results. For example, compared to brain and liver, isolated myofibers express a very low level of FGFR2 and FGFR3 but a significantly higher level of FGF6.
Immunoblotting Analysis of FGF6 Protein
Various tissues collected from 8-week-old rats were frozen in liquid nitrogen, pulverized, and homogenized in 5 volumes of ice-cold buffer. The skeletal muscle tissue was homogenized in a buffer consisting of 100 mM KCl, 20 mM Tris-HCl (pH 7.5), 1 mM DTT, and the protease inhibitors leupeptin (Boehringer Mannheim), soybean trypsin inhibitor (Worthington Biochemical; Freehold, NJ), and aprotinin (Sigma), each at 10 μg/ml, and Pefablock (Boehringer Mannheim) at 240 μg/ml. The homogenate was centrifuged at 16,000 × g for 20 min and the resulting pellet and supernatant fractions were both reacted with Laemmli SDS sample buffer (1 × final concentration; Laemmli 1970). All other tissues were homogenized directly in the Laemmli SDS sample buffer containing the protease inhibitors described above. The final tissue extracts were heated at 95C for 5 min and 50 μg total protein per sample was separated on a 12% SDS-polyacrylamide gel and blotted onto a nitrocellulose membrane. The blot was stained with 0.1% Ponceau S solution (Sigma) to confirm the adequate transfer of the proteins and blocked overnight with 5% milk powder and 2% horse serum in Tris-buffered saline (TBS; 150 mM NaCl, 20 mM Tris, pH 7.8). The blot was then reacted for 1.5 hr with an affinity-purified goat polyclonal antibody raised against recombinant human FGF6 expressed in E. coli (product #AF238, R & D Systems; antibody diluted to a final concentration of 0.2 μg/ml in TBS containing 1% BSA and 2% horse serum). The blot was washed in TBS containing 0.1% Tween-20 and incubated for 1 hr with an HRP-conjugated donkey anti-goat IgG (Santa Cruz Biotechnologies; diluted 1:20,000 in the same buffer used to dilute the primary antibody). Final detection of the reactive bands was performed using the enhanced chemiluminescence reagents (ECL; Amersham, Arlington Heights, IL). Under the above conditions, the antibody against FGF6 reacted intensely with recombinant human FGF6 (see Results) and no crossreactivity was observed with recombinant human FGF1, FGF2, FGF4, FGF5, and FGF7 (data not shown).
Results
The Effects of FGFs and HGF on Satellite Cells in Isolated Myofibers
The effects of different FGFs and of HGF on enhancing the number of satellite cells in myofiber cultures was monitored by quantifying the numbers of PCNA+ and myogenin+ nuclei. The results of the investigation are shown in Figure 3. Immunostaining micrographs demonstrating that the PCNA+ and myogenin+ nuclei are within fiber-associated cells whose cytoplasm can be traced with an antibody against the mitogen-activated protein kinases ERK1/ERK2 have been published (Yablonka-Reuveni et al. 1999b). This immunocytochemical approach has provided a reliable means for monitoring the overall number of satellite cells because the cells do not fuse with their associated myofibers (Bischoff 1986, 1990; Yablonka-Reuveni and Rivera 1994). The growth factors were tested with or without heparin, whose presence can influence the mitogenic activity of certain FGFs (Spivak-Kroizman et al. 1994). The concentration of heparin (10 μg/ml) was based on the supplier's recommendation for optimal FGF1 activity.
The effect of FGF1 on the number of PCNA+ cells was examined at concentrations ranging from 0.1 to 5 ng/ml. The maximal increase in PCNA+ cells was observed at 2 ng/ml FGF1 with heparin (Figure 3A). No further increase in PCNA+ cells was seen at higher FGF1 concentrations. Heparin added to the basal medium alone (without FGF1) suppressed the number of PCNA+ cells (Figure 3B). Heparin suppressed cell proliferation even when FGF1 (or the other effective FGFs described below) was added at a low level of 0.1 ng/ml. Such a low level of FGF was sufficient to promote some proliferation in the absence of heparin. These findings indicate that the ratio between the concentration of heparin and the FGFs is critical for achieving maximal proliferation of satellite cells in myofiber cultures.
Quantification by immunofluorescence of PCNA+ and myogenin+ cells in cultured myofibers isolated from 8–10-week-old rats and maintained in basal medium with different growth factors and heparin. The concentrations used for the various additives are indicated in each panel. Heparin (hep) was added at 10 μg/ml. For each time point, cells were scored as the number of positive nuclei on individual fibers, analyzing 30 fibers per plate. Total positives were then averaged for two parallel plates and the final values in the figure are expressed per 10 fibers. Error bars show the range of the results for the parallel plates.
The effect of FGF2 (2 ng/ml) on the number of PCNA+ satellite cells was minimally enhanced, if at all, by heparin (Figure 3C). We previously found that this concentration of FGF2 produces a maximal effect on the number of PCNA+ satellite cells in myofiber cultures. A higher concentration of FGF2 (up to 10 ng/ml) did not lead to a further increase.
FGF4 (2 ng/ml) also enhanced the number of PCNA+ satellite cells in myofiber cultures. This is shown in Figure 3D, together with the results of parallel plates receiving FGF1 (with heparin) or FGF2. A higher FGF4 concentration (up to 10 ng/ml) did not support a further increase in the number of PCNA+ cells. The effect of FGF4 was not influenced by the addition of heparin (data not shown). Figure 3D also shows that the schedule of transition from the PCNA+ to the myogenin+ state follows a similar pattern in the presence of FGF1, FGF2, or FGF4. The enhancement of PCNA+ cells by these three FGFs is reflected by a similar increase in the number of myogenin+ cells.
The effect of FGF6 on the number of PCNA+ cells was tested at a concentration range between 0.1 and 10 ng/ml. The results obtained with FGF6 at 5 ng/ml (with or without heparin) are shown in Figure 3E. This concentration of FGF6 supported the maximal number of PCNA+ cells. The effect of FGF6 was only minimally (but consistently) enhanced by heparin on culture Days 2 and 3. The results obtained with parallel cultures receiving FGF2 at 2 ng/ml are also included in Figure 3E. FGF6 and FGF2 supported a similar increase in the number of PCNA+ cells.
The effect of HGF (20 ng/ml) on the number of PCNA+ cells is shown in Figure 3F. HGF at 20 and 40 ng/ml led to similar results, whereas HGF at 5 ng/ml supported a slightly smaller number of PCNA+ cells (data not shown). Heparin did not enhance the number of PCNA+ cells beyond that supported by HGF alone (not shown). As seen in Figure 3F, HGF (at 20 ng/ml) and FGF2 (at 2 ng/ml) enhanced the number of PCNA+ cells to a similar degree. Figure 3G shows that these concentrations of HGF and FGF2 also led to a similar increase in the number of myogenin+ cells. The addition of HGF (20 ng/ml) and FGF2 (2 ng/ml) together did not yield a further increase in the number of satellite cells beyond that shown in Figure 3F for each growth factor alone (not shown). Collectively, HGF (like FGF1, FGF2, and FGF4) did not modify the schedule of transition from the PCNA+ to the myogenin+ state.
The effect of FGF6 on the number of myogenin+ cells was examined using 5 ng/ml FGF6 plus heparin (the condition that supported the maximal number of PCNA+ cells). The results are shown in Figure 3H, along with control cultures receiving no additives or supplemented with HGF (20 ng/ml). Typically, the number of myogenin+ cells sharply declines between culture Days 3 and 4 in myofiber cultures receiving basal medium only and in cultures receiving FGF1, FGF2, FGF4, or HGF (Figures 3C, 3D, and 3G; also detailed in our previous publications). As seen in Figure 3H, FGF6 exerted a unique effect on the number of myogenin+ cells, slowing the rapid decline observed in the controls. The addition of FGF6 and HGF together yielded the same kinetics of myogenin+ cells as shown in Figure 3H for FGF6 alone (data not shown).
The effects of FGF5 and FGF7 on the number of PCNA+ cells in myofiber cultures were also examined. Myofibers in basal medium with or without FGF2 were used as controls in these experiments. FGF5 was first tested at a concentration range of 2–75 ng/ml and then at 100–200 ng/ml. FGF7 was tested at 10–50 ng/ml. Neither FGF5 nor FGF7 (with or without heparin) was found to enhance the number of PCNA+ cells above that seen in basal medium (not shown).
We further investigated whether the decline in the myogenin+ cells seen by Day 4 in myofiber cultures could be due to apoptosis and whether FGF6 may slow this decline by preventing apoptosis. Cultures were collected every 24 hr for 6 days, starting at time 0, and were analyzed for the presence of apoptotic nuclei using the TUNEL reaction. We concluded that regardless of the absence/presence of FGF2 or FGF6, neither the nuclei of the intact myofibers nor the nuclei of the fiber-associated satellite cells underwent programmed cell death (data not shown).
Expression of FGFs and Their Receptors in Myofiber Cultures
We asked whether the muscle fibers (with their associated satellite cells) express specific FGF or FGFR genes in comparison to the connective tissue cells surrounding the myofibers. Myofibers and muscle connective tissue cells were cultured in parallel under the same conditions used for the studies on the effects of the growth factors. FGF2 (2 ng/ml) was added to the basal medium to support proliferation. Total RNA was isolated from the cultures at different time points and was analyzed by RT-PCR (Figure 4). The RT-PCR products of each gene are shown in paired horizontal lanes; the upper lane shows the analysis of the myofibers (mf) and the lower lane shows the analysis of the connective tissue cells (ctc). Gene expression in the myofibers reflects the overall contribution of both the myofibers and the satellite cells.
As shown in Figure 4, MyoD, Myf5, myogenin, and MRF4 were detected only in the myofiber cul tures. Their expression was upregulated by the first culture day. The highest expression of MyoD seen on Day 1 preceded the highest expression of myogenin seen on Day 3. This pattern agrees with the temporal expression of the MyoD and myogenin proteins by the fiber-associated satellite cells. However, we cannot exclude that at least some, if not most, of the MRF gene expression is contributed by the myofibers themselves. The muscle regulatory/structural genes muscle LIM protein (MLP) (Arber et al. 1994; Schneider et al. 1999), troponin I fast, and troponin I slow (TnI fast, TnI slow; Krishan and Dhoot 1996) were also detected only in the myofiber cultures, with levels declining after the first day in culture.
RT-PCR analysis of gene expression in cultures of myofibers and connective tissue cells isolated from the FDB muscle of the hindfeet of 8-week-old rats. For each gene, the results are provided in paired lanes. The upper lane shows the results for myofibers (mf) and the lower lane for the connective tissue cells (ctc). The myofibers and the connective tissue cells used for RNA isolation were cultured in parallel in 35-mm plates coated with Vitrogen and using basal medium containing 2 ng/ml FGF2. Day 0 cultures were collected within the first hour and the later time points were collected at 24-hr intervals.
Depending on the time in culture, expression of FGF1, FGF2, FGF5, FGF7, and HGF was displayed by both the myofibers and connective tissue cells (Figure 4). FGF4 was not analyzed because we could not detect it in preliminary studies. FGF4 was also not detected in the mouse myogenic line MM14 (Hannon et al. 1996), and its expression has been thought to be limited to the embryo (Szebenyi and Fallon 1999). FGF6 was uniquely expressed in the myofiber cultures and not in the connective tissue cells at the PCR cycle number used in the study. The expression pattern of FGF6 was similar to that of MLP, TnI fast, and TnI slow, with levels of all genes declining after the first day in culture (Figure 4). We have not yet investigated the basis for this decline, but in view of the apoptosis studies mentioned above it is unlikely to be due to myofiber death.
Analysis of the FGF and HGF receptor genes showed a unique expression of FGFR4 by the myofibers only. All other receptor genes were expressed by both the cultured myofibers and the connective tissue cells (Figure 4). The expression of all receptor genes, excluding FGFR3, was upregulated in the myofibers by the first culture day. The expression level of FGFR2 and FGFR3 in the myofiber cultures was distinctly low compared to that of brain or liver (shown in Figure 2). The muscle tissue also expressed a very low level of FGFR2 and FGFR3 (Figure 2). The expression level of FGFR2 was far higher in the connective tissue cells than in the myofiber cultures (Figure 4), suggesting that the myofiber FGFR2 might be contributed by remaining connective tissue cells.
The expression of H-ras in myofibers was similar at all time points (Figure 4). In the connective tissue cultures, H-ras was expressed at a lower level at time 0, like the other control genes GAPDH and rpS6 (data not shown). This might be due to the reduced cell activity immediately after the cell isolation. This reduced activity in time 0 connective tissue cells has no impact on the conclusions of the study.
In the investigation shown in Figure 4, the skeletal muscle genes as well as the FGFR4 gene were expressed by the myofibers but not by the connective tissue cells. The connective tissue cells expressed a trace level of these genes when the PCR was carried out at 40 cycles (data not shown). The source of these trace signals is probably the remnant myofibers (and/or infrequent satellite cells) trapped with the connective tissue cells. A trace of FGF6 expression was also seen in the connective tissue cells at a high PCR cycle number. We also investigated whether the medium (basal medium vs the growth medium used for muscle-dissociated cells) or the matrix (Vitrogen vs gelatin) can affect the expression of FGF6 and FGFR4. The same relative higher expression levels in myofibers compared to connective tissue cells were detected for these two genes regardless of the medium or matrix used (data not shown). We also analyzed whether the expression levels of the growth factors and their receptors are changed in myofibers from 1-year-old rats. We concluded that the expression patterns of these genes in the older rats were similar to those shown in Figure 4 for myofibers from 8-week-old rats (data not shown).
Detection of FGF6 Protein in Muscle Tissue
The effect of FGF6 during myogenesis of satellite cells, along with its unique pattern of RNA expression in the myofiber system, led us to further investigate the presence of FGF6 protein in the muscle tissue. Although mRNA transcripts for FGF6 have been detected in regenerating skeletal muscle (Floss et al. 1997), there are no reports on the expression of the protein itself. The present analysis focused on verifying that FGF6 protein is present in the muscle. FGF6 is a secreted protein, and its analysis in myofiber cultures would require a comprehensive investigation of the culture medium and the matrix.
Figure 5 depicts a Western blot analysis of several rat tissues along with recombinant human FGF6 (rhFGF6). Although Lane 1 shows the results with 1 ng rhFGF6, we concluded that the assay allows the detection of as little as 0.05 ng rhFGF6. Fractionation of the FDB muscle extract to soluble and particulate material, which was performed because of the toughness of the tissue, was found to provide a means for FGF6 enrichment. FGF6 was detected in the particulate fraction (Figure 5, Lane 2) but not in the soluble material (data not shown). Testis extract (Figure 5, Lane 4) was included as a positive control for rodent FGF6 on the basis of information provided by Santa Cruz Biotechnologies about FGF6 expression in mouse. The arrows at the sides of the figure point to the migration position of rhFGF6 (left arrow) and rat FGF6-like protein (right arrow). The predicted molecular weight of the rhFGF6 is 19 kD. However, according to the supplier, the rhFGF6 is prone to both N- and C-terminal truncation and contains several proteins ranging from 14 to 16 kD. The molecular weight of the rat FGF6-like protein was found to be about 20 kD. This is in agreement with the 18-kD molecular weight reported for mitogenically active, transgene FGF6 expressed in Chinese hamster ovary cells (Asada et al. 1999).
Western blotting analysis of FGF6 protein in rat tissue extracts. About 50 μg protein was analyzed for each tissue sample. Tissue extracts along with rhFGF6 were separated on a 12% polyacrylamide gel and blotted onto nitrocellulose membrane. The blot was reacted with an affinity-purified antibody raised in goat against rhFGF6, and the secondary antibody was a peroxidase-conjugated donkey anti-goat IgG. Final detection of the positive bands was done with the enhanced chemiluminescence (ECL) reaction. Lane 1, 1 ng of rhFGF6; Lane 2, the particulate fraction of FDB muscle extract; Lanes 3 and 4, extracts of brain and testis, respectively. Arrows at left and right point to the migration positions of rhFGF6 (left) and rat FGF6-immunoreactive protein (right).
Distribution of Myogenic Cells in Cultures of Percoll-isolated Cells
To focus specifically on FGF-FGFR gene expression in the satellite cells themselves apart from the myofibers, we isolated satellite cells from the rat EDL/TA muscles. The cells were further subjected to Percoll density centrifugation to enrich for nonmyogenic and myogenic cell populations typically present together in preparations of muscle-dissociated cells (Yablonka–Reuveni and Nameroff 1987; Morgan 1988; Yablonka–Reuveni et al. 1988; Bischoff 1997).
The myogenic potential of the EDL/TA cells was first examined using preparations that were not fractionated by Percoll. Figure 6 depicts immunofluorescence micrographs of Day 5 (Figure 6A) and Day 10 (Figure 6B) cultures of nonfractionated cells reacted with the antibody against myogenin. The frequency of the myogenin+ cells was less than 5% by culture Day 5 but increased to 60–80% by culture Day 10. Multi-nucleated myotubes containing myogenin+ nuclei were also seen at this late stage. MyoD+ cells were present in these cultures by culture Day 1, reaching a frequency of about 50–60% by Day 5. A parallel RT-PCR analysis revealed the expression of MyoD by Day 1 and of myogenin by Day 2 in culture (data not shown).
To determine the distribution of myogenic cells along the Percoll fractions, cultures of Percoll-isolated cells were analyzed by double immunofluorescence with the antibodies against MyoD and MEF2A (both antibodies react with cell nuclei; Yablonka-Reuveni and Rivera 1997a; Yablonka-Reuveni et al. 1999a). The MyoD+/MEF2A− phenotype is displayed by mono-nucleated cells and represents cells at an earlier stage in the myogenic program. The MyoD±/MEF2A+ phenotype is displayed by both mononucleated and multi-nucleated cells and represents cells at a later stage in the myogenic program (Yablonka-Reuveni and Rivera 1997a; Yablonka-Reuveni et al. 1999a). These MEF2A+ cells are typically also positive for myogenin and sarcomeric myosin (Figures 7A and 7B). Table 1 and Figures 7C-7E show a MyoD±/MEF2A± distribution analysis of Percoll-isolated cells cultured for 8 days. This extended time in culture provided sufficient time for cell differentiation despite variations in growth rates between cultures from different fractions (see Table 1 for the comparison of total cell numbers on Day 8). As described in Materials and Methods, the initial plating density was similar for the different fractions.
Immunofluorescent micrographs of myogenin+ cells in cultures of muscle-dissociated cells. The secondary antibody was a fluorescein-conjugated goat anti-mouse IgG. Cultures were counterstained with the nuclear stain DAPI. Cells were isolated from the EDL/TA muscles and cultured before the Percoll isolation step. Cultures were maintained in a serum-rich growth medium consisting of DMEM with 25% fetal bovine serum, 10% horse serum, and 1% chicken embryo extract. (
The data in Table 1 demonstrate that the overall frequency of myogenic cells is significantly reduced in the cultures from 25–35% and 35% Percoll fractions and is the highest in the culture from the 55% fraction. Table 1 further shows that the emergence of the MEF2A+ phenotype is delayed in cultures from the two bottom Percoll fractions compared to cultures from the three middle fractions. This distinction in the emergence of the differentiated MEF2A+ cells might be due to variations in cell densities by culture Day 8 (summarized in Table 1). Nevertheless, it has permitted the comparison of gene expression by cells at different stages of the myogenic program (see below). Figure 7 further demonstrates that the MyoD+ cells in the bottom 55–70% fraction (Figure 7E) are larger than the MyoD+ cells in the other Percoll fractions (Figure 7D). The nature of the cells contributing these larger MyoD+ cells has not been yet resolved. Additional Percoll fractionation studies resulted in cell distributions similar to those shown in Table 1, with the exception that the frequency of myogenic cells in the top fraction ranged between 4 and 12% by culture Day 8 (not shown). This range is probably due to variations in defining the border between the 25–35% and 35% Percoll fractions after centrifugation.
The MyoD−/MEF2A− cells in the cultures of the various Percoll fractions do not necessarily represent one common nonmyogenic cell phenotype. These negative cells might consist of authentic connective tissue cells along with fibroblast-like cells that have been derived in the culture from myogenic cells. Cells negative for myogenic traits can even be detected in myogenic clones derived from single myogenic progenitors (Yablonka–Reuveni and Nameroff 1987; Yablonka–Reuveni and Rivera 1997a). Immunotracing of rat myoblasts by their expression of desmin (Allen et al. 1991) was avoided in the present study because vascular smooth muscle cells, which are likely to be present in the cultures, also express desmin (Yablonka–Reuveni et al. 1998).
Immunofluorescent micrographs of cultures of muscle-dissociated cells reacted with a polyclonal antibody against MEF2A in combination with different MAbs. The secondary antibodies were rhodamine-conjugated goat anti-rabbit IgG and fluorescein-conjugated goat anti-mouse IgG. All cultures were maintained in a serum-rich growth medium as in Figure 6. (
Distribution of MyoD+ and MEF2A+ cells in 8-day cultures of muscle-dissociated cells fractionated by Percoll
Arbitrary microscopic fields were monitored using a x40 objective.
Values are shown to the first decimal only.
Expression of the FGFs and Their Receptors in Cultures of Percoll-isolated Cells
RT-PCR analysis of the EDL/TA muscle cells cultured before Percoll fractionation demonstrated the expression of FGFs, HGF, and their receptors at various time points (data not shown). To determine more specifically the relative expression of these genes by the different cell populations present in the muscle-dissociated cells, we analyzed gene expression in Percoll-isolated cells. Figure 8 shows RT-PCR analysis of Percoll-isolated cells that were harvested after 5 days in culture. Cells from the bottom Percoll fraction were not included in the present study. Collective analysis of the seven skeletal muscle genes (MyoD, Myf5, myo-genin, MRF4, MLP, TnI fast, TnI slow) demonstrates that the cultures recovered from the 40% and 40–45% fractions are in a more advance stage of the myo-genic program, while the 35–40% and 55% fractions are lagging somewhat behind.
In contrast to the skeletal muscle genes, the expression levels of FGF1, FGF2, FGF7, and HGF decline in the cultures from the denser Percoll fractions, which raises the possibility that the nonmyogenic cells express these genes at higher levels. Compared to myofiber cultures (Figure 4), we could not detect FGF6 with 34 PCR cycles in any of the Percoll fractions. Some FGF6 expression was detected in the cultures from all the Percoll fractions at 38 PCR cycles (data not shown). This indicates that FGF6 is expressed at a far lower level by the myogenic and nonmyogenic cells compared to the myofibers. FGF5 was also barely detectable in these cultures of Percoll-isolated cells, even at 40 PCR cycles (data not shown).
As shown in Figure 8, the expression pattern of FGFR4 differs markedly from that of the other FGFRs and correlates with the degree of myogenicity of the cultures from the different Percoll fractions. In fact, FGFR4 expression level correlates well with the expression of the “early” myogenic markers MyoD and Myf5. The three genes are co-expressed even in fractions that do not yet display increased levels of the “late” differentiation-linked genes. In contrast, FGFR1 expression is reduced as the Percoll concentration increases, suggesting that FGFR1 is elevated in the non-myogenic cells. The expression of FGFR2 and FGFR3 also does not correlate with the degree of myogenicity, and their detection requires a greater number of PCR cycles compared to the other receptors. The expression level of c-met was also increased in the more myogenic fractions (i.e., 40%, 40–55%, and 55% fractions). However, the cultures derived from the top three Percoll fractions (i.e., 25–35%, 35%, and 35–40%) did not show a distinct gradual increase in the level of c-met, although the degree of myogenicity gradually increased along the gradient (Figure 8).
RT-PCR analysis of gene expression in Day 5 cultures of Percoll-fractionated cells isolated from the EDL/TA muscles. Cells recovered from the different fractions were cultured in gelatin-coated 35-mm plates using a serum-rich growth medium consisting of DMEM with 25% fetal bovine serum, 10% horse serum, and 1% chicken embryo extract. Cells were harvested for total RNA isolation after 5 days in culture. The 50–70% fraction was not examined.
Figure 9 shows RT-PCR analysis of Percoll-isolated cells harvested immediately after their isolation (Day 0) and after 2 and 4 days in culture. Only the “interface” Percoll fractions were examined. The expression profile of almost all the genes analyzed at Days 2 and 4 was similar to that of Day 5 discussed above. The expression of MyoD in the Day 2 and 4 cultures of the 25–35% fraction is in agreement with the range of myogenic cells seen in this fraction using immunofluorescence, as discussed above. FGF2, FGF7, and HGF were expressed at higher levels by cells from the least dense Percoll fractions (25–35% and 35–40%), with the expression of HGF first seen only by culture Day 4. The detection of FGF5 and FGF6 required a high PCR cycle as in Day 5 cultures (data not shown). Similar to Day 5 cultures, the distribution of both FGFR4 and c-met (but not the other receptor genes) followed a pattern of increased expression in the more myogenic fractions. This was especially apparent for FGFR4.
Myogenesis was also evident in the Day 0 cells. However, as shown in Figure 9, expression of the housekeeping genes rpS6 and GAPDH varied among the different fractions. This is likely due to the differences in the activity of the cells at the time of their isolation. The expression of the growth factor genes was also significantly reduced in the Day 0 cells compared to the Day 2 and 4 cultures. Nevertheless, the Day 0 cells recovered from the 40–55% fraction expressed the four MRFs along with rpS6 and GAPDH, suggesting that at least some of the cells in this fraction were transcriptionally active and underwent myogenesis at the time of cell isolation. Expression of FGFR1, FGFR3, FGFR4, and c-met was also detected in this fraction. Myf5 expression in the Day 0 cells recovered from the 35–40% and 55–70% fractions indicates the presence of myogenic cells in these fractions as well. The nature of the strong MRF4 signal in the 55–70% fraction has not been further investigated. It might be related to the large MyoD+ cells present in this fraction, as discussed above. The distribution of FGFR4 expression among the Day 0 Percoll fractions is similar to that of Myf5, suggesting that FGFR4 expression follows the degree of expression of the early myogenic gene, as discussed above.
RT-PCR analysis of gene expression in cultures of EDL/TA cells fractionated by Percoll density centrifugation and maintained in culture for various lengths of time. Day 0 cells were processed for total RNA isolation immediately after recovery from the Percoll fractions. Day 2 and Day 4 cultures were maintained in the serum-rich medium described in Figure 8. Total RNA for the analysis of Day 0 and Day 2 cultures was pooled from several isolations. Only the fractions recovered from the interfaces between the Percoll layers were examined.
As shown above, FGF5 is expressed in myofiber/connective tissue cultures (prepared from the FDB muscle) but is barely detected in cultures of the isolated satellite cells (prepared from the EDL/TA muscles). A further comparison of isolated satellite cell cultures from the two sources demonstrated similar mRNA expression patterns for the various genes discussed above, except for FGF5. FGF5 was again expressed at a relatively high level only in the cultures from the FDB muscle but not in cultures released from the EDL/TA muscles (data not shown).
Discussion
The investigations described here were carried out to begin defining the FGF signaling pathways that act during myogenesis of skeletal muscle satellite cells. We have taken a comprehensive approach of analyzing the effects of exogenously added growth factors on myogenesis of satellite cells in combination with exploring the gene expression patterns of these growth factors and their receptors in isolated myofibers and satellite cells. The analyses show unique expression patterns for FGF6 and FGFR4 and therefore may hint at specific regulatory roles of these members of the FGF signaling complex during myogenesis in postnatal muscle. We also attempted to conduct parallel immunocytochemical studies to localize the FGF/FGFR proteins themselves in isolated myofibers and satellite cell cultures. However, the various available antibodies did not produce satisfactory signals for immunocy-tochemistry or did not discriminate among different FGFs/FGFRs.
FGFs and HGF Promote the Proliferation of Satellite Cells to a Similar Extent
Rat satellite cells undergoing proliferation and differentiation in myofiber cultures can be traced by immunostaining of their nuclei with antibodies against PCNA and myogenin (Yablonka-Reuveni et al. 1999b). This immunofluorescence approach was used in the present study to analyze the effect of several FGFs and HGF on myogenesis of satellite cells in myofiber cultures from young adult rats. We found that FGF1, FGF2, FGF4, FGF6, and HGF (but not FGF5 and FGF7) were all capable of enhancing the number of proliferating satellite cells to a similar degree. In all cases, the effective growth factors enhanced by about twofold the peak number of PCNA+ cells. However, the schedule of transition of the cells into the myogenin+ state was not modified by any of the growth factors examined. In all cases, the enhanced number of PCNA+ satellite cells supported by the FGFs or HGF was reflected in a similar maximal number of myogenin+ cells. Therefore, the FGFs cannot suppress differentiation of satellite cells in isolated myofibers. This finding, which is in contrast to the commonly accepted view that the FGFs can delay differentiation of myo-blasts, was also observed in cultures of mouse myofibers (Yablonka-Reuveni et al. 1999a).
Sheehan and Allen (1999) reported on the proliferative effect of different FGFs in primary cultures of rat satellite cells dissociated away from the muscle. The conclusions of that study regarding the effective FGFs are in agreement with the conclusions of our present study. However, our study focuses specifically on satellite cells situated in their native position. The interpretation of growth factor effects in primary cultures can be complicated by the presence of nonmyogenic cells, which may also respond to the mitogens. Earlier studies by Allen and colleagues also suggested that HGF (but not FGF2) can shorten the lag time spent by quiescent satellite cells before they enter DNA synthesis (Johnson and Allen 1993; Allen et al. 1995; Tatsumi et al. 1998). However, our studies of satellite cells in myofibers from 9-month-old rats revealed a similar schedule of cell cycle entry and proliferation in response to FGF2 and HGF (Yablonka-Reuveni et al. 1999b). The present study with myofibers from 8–10-week-old rats further demonstrates that satellite cells follow a similar schedule of proliferation in response to HGF or the different FGFs. Hence, the satellite cells express both the FGF and the HGF signaling cascade when they enter the first round of cell proliferation in isolated myofibers.
The Effect and Site of Expression of FGF6 During Myogenesis of Satellite Cells
The mRNA expression studies indicated that FGF6 was expressed in myofiber cultures when the satellite cells enter proliferation. The presence of FGF6 protein in the muscle was also demonstrated by a sensitive Western blotting analysis. This correlation between the FGF6 expression when satellite cells initiate proliferation and the FGF6 effect on enhancing the number of proliferating satellite cells suggests that FGF6 is involved in vivo in the proliferation of satellite cells. Whereas proliferation of satellite cells was also enhanced by other FGFs, only FGF6 was able to slow the rapid decline in myogenin+ cells. By counterstaining the cytoplasm of the fiber-associated satellite cells, we showed that the routinely observed decline in myogenin+ cells is due to transient expression of myogenin but that the cells themselves remain attached to the myofibers (Yablonka-Reuveni et al. 1999b). Moreover, among the different growth factors examined, only FGF6 was uniquely expressed at a higher level by the isolated myofibers compared to the nonmyogenic cells surrounding the myofibers. The mRNA expression studies also indicated that among the different growth factors examined in myofiber cultures, only the level of FGF6 was reduced with time in culture. This decline in FGF6 expression, combined with the slower decline in the number of myogenin+ cells when FGF6 was added, has suggested a possible role for FGF6 in maintaining the myogenin+ state of the satellite cells. Our studies therefore support a dual role for FGF6 during both the proliferative and the differentiative state of satellite cells.
The unique effect of FGF6 in retarding the decline in myogenin protein expression by satellite cells may reflect a physiological role for FGF6 in regulating the differentiated state of the satellite cells and possibly of the myofibers themselves. Myogenin is a muscle-specific transcription factor for various muscle differentiation target genes (Molkentin et al. 1995). Therefore, the ability of FGF6 to prolong the myogenin+ state might eventually modulate the myogenic differentiation program by enhancing the expression of muscle structural genes. This, in turn, may support the ongoing maintenance of the muscle. The contribution of FGF6 by the myofibers themselves may allow FGF6 to be in close association with the satellite cells, efficiently enabling their continuous participation in the maintenance of healthy muscle. The muscle endures daily subtle injuries as a result of its normal activities, and the satellite cells are likely to be continuously involved in myofiber repair.
A number of published studies provide circumstantial evidence for the possible role of FGF6 in the muscle. First, although several FGFs are expressed in the developing muscle, only the expression of FGF6 is restricted to the muscle itself during early embryogenesis (Haub and Goldfarb 1991; Niswander and Martin 1992; deLa-peyriere et al. 1993; Mason et al. 1994; for summaries see Coulier et al. 1994; Floss et al. 1997). In addition, the onset of FGF6 expression during muscle development was delayed in embryos of MRF-deficient mutant mice (Patapoutian et al. 1995; Grass et al. 1996). Moreover, the prolonged phase of muscle fiber hyperplasia in postlarval fish is accompanied by the lasting expression of FGF6 to the adult stage (Rescan 1998). These findings suggest that FGF6 may participate in the continuous generation of muscle during embryogenesis and postnatal growth. Second, RNA expression studies revealed that the level of FGF6 was increased during muscle regeneration and that muscle regeneration was retarded in the FGF6−/− mutant mouse (Floss et al. 1997). Third, FGF6 was found to upregulate the expression of several differentiation markers when given at a low concentration to cultures of mouse-derived C2 cells; a higher concentration of FGF6 enhanced proliferation of these cells (Pizette et al. 1996).
The distinction of the present study is that it provides a direct insight into the site of expression and the effect of FGF6 in the adult muscle. Future studies on how FGF6 affects satellite cell proliferation and how it may regulate the expression of the differentiation-specific genes can be important for enhancing muscle maintenance. A direct effect of exogenous FGF6 on the myofiber itself would require the presence of appropriate transmembrane receptors at the myofiber surface. Obviously, an mRNA expression analysis such as that reported in the present study cannot distinguish between the contribution of the satellite cells vs the contribution of the myofibers, and can only provide preliminary evidence about the possible presence of FGFRs. We attempted to use commercial antibodies against FGFRs, especially antibodies against FGFR1 and FGFR4, to further approach the subject by immunocytochemistry. However, a comprehensive study of various anti-FGFR antibodies by multiple approaches did not confirm their suitability for immunolocalization studies of rodent cells. Interestingly, an immunohistochemical study of human muscle that utilized human-specific FGFR antibodies documented the expression of FGFR1 and FGFR4 in the plasma membrane of embryonic but not adult myofibers (Sogos et al. 1998). Therefore, the myofibers themselves might undergo modulations that lead to the expression of FGFRs. In the present rat myofiber analysis, such modulations in FGFR levels might be due to the lack of innervation, reduced stretch and activity, or to other factors contributed by the exposure to culture conditions. Changes in FGFR1 and FGFR4 expression levels in skeletal muscle subjected to chronic electrical stimulation have also been interpreted as modulations in FGFRs in the myofibers themselves (Düsterhöft et al. 1999).
Unique Expression and Possible Role of FGFR4 During Myogenesis of Satellite Cells
The mRNA expression studies conducted with isolated myofibers and isolated satellite cells revealed a unique expression pattern for FGFR4 in skeletal muscle. FGFR4 was expressed at relatively higher levels in intact myofibers and in the satellite cells isolated from the muscle compared to the muscle connective tissue cells. A higher level of FGFR4 expression was detected in satellite cells, regardless of the time spent in culture or the degree of differentiation. This suggests that FGFR4 might be expressed by proliferating and differentiating satellite cells. Expression of FGFR1, FGFR2, and FGFR3 was also seen in the myofiber cultures and in cultures of isolated satellite cells. However, only FGFR1 was readily detectable by RT-PCR, whereas the detection of FGFR2 and FGFR3 required a high number of PCR cycles. The analysis of gene expression in enriched myogenic and nonmyogenic cell populations isolated by Percoll density centrifugation has indicated that only FGFR4 expression correlates with the degree of myogenicity. Furthermore, FGF1, FGF2, FGF4, and FGF6 (but not FGF5 and FGF7) enhanced satellite cell proliferation in the myofiber studies. This characteristic profile of effective and ineffective FGFs fits well with the ligand affinities of FGFR4 established by Ornitz et al. (1996). Obviously, we cannot exclude the importance of the other FGFRs detected in our studies, because a combination of these different receptors can also yield a similar response to the FGFs as that seen in the present study (Ornitz et al. 1996).
Previous studies detected the expression of FGFR1 and FGFR4 in developing human and mouse muscles (Partanen et al. 1991; Stark et al. 1991). FREK, which is believed (but not proved) to be the avian equivalent of mammalian FGFR4, was also detected in the developing muscle and in cultures of satellite cells (Halevy et al. 1994; Marcelle et al. 1995). Recent publications have reported that FGFR1 and FGFR4 are expressed in cultures of the mouse myogenic line C2 (Pizette et al. 1996) and in primary cultures of muscle-dissociated rat satellite cells (Düsterhöft et al. 1999; Sheehan and Allen 1999). However, our study contributes new and unique information regarding the localization and expression of FGFR4 in relationship to the myogenic program, distinguishing gene expression patterns between myogenic and nonmyogenic cells, both of which are typically present in primary cultures of satellite cells. We detected the same profile of FGFR expression as reported in the present rat study in our studies of satellite cells from various mouse strains. FGFR4 was uniquely expressed by myofibers and satellite cells, FGFR1 was ubiquitously expressed by myogenic and nonmyogenic cells, and FGFR2 and FGFR3 were expressed at a far lower level than FGFR1 and FGFR4 (unpublished results).
It is notable that the mouse MM14 myogenic cells, which have been used extensively to study the role of FGFs during myogenesis, express only FGFR1 but not the other FGFRs (Templeton and Hauschka 1992; Kudla et al. 1998). The capacity of the MM14 cells to remain proliferative is highly dependent on the addition of FGF2 (Clegg et al. 1987). Our studies, which have indicated that FGFR4 is expressed during myo-genesis, also demonstrated that satellite cells rapidly proceed from proliferation to differentiation and that FGF2 (or other FGFs) cannot suppress this transition. It is therefore possible that myoblasts lacking FGFR4 may be regulated differently by FGFs than myoblasts expressing FGFR4. This suggestion is supported by studies of rat L6 myoblasts that were transfected with FGFR1 and FGFR4 (Vainikka et al. 1994; Wang et al. 1994; Shaoul et al. 1995). The FGFR1-expressing cells showed a far more robust activation of the ERK1/ERK2 mitogen-activated protein kinase signaling cascade compared to the FGFR4-expressing cells. The observed differences might be due to variations in the heparin sulfate proteoglycans required for binding of the FGF ligand to the different receptors (Bonneh–Barkay et al. 1997; Kan et al. 1999; reviewed in Szebenyi and Fallon 1999). In the absence of the appropriate heparin sulfate proteoglycans, the transfected FGFR4 might have not been fully activated. To bypass this potential difficulty, a more recent comparison of FGFR signaling has used chimeric receptors in which the external portion of the FGFR was replaced with the external domain of the PDGF receptor (Raffioni et al. 1999). The interaction of the PDGF with its receptor does not require heparan sulfate proteoglycans or other co-factors. Despite this approach, FGFR4 still elicits only a very weak activation of ERK1/ERK2 compared to FGFR1 or FGFR3 (Raffioni et al. 1999). The possibility of unique signaling by FGFR4 has also been supported by the observation that an 85-kD serine kinase was found associated only with FGFR4 (Vainikka et al. 1996).
We propose that in the satellite cell system FGFR1 may regulate ongoing proliferation of myoblasts, acting in a similar manner to its action in other cell systems. In contrast, FGFR4 might be involved in a myo-genic-specific pathway. This proposal is supported by our recent studies with satellite cells from the MyoD−/– mouse, in which we demonstrate that the onset of myogenic differentiation and FGFR4 gene expression is similarly delayed (unpublished results). Studies on the FGFR4−/– mutant mouse demonstrated that FGFR4 is dispensable during embryonic development and that the FGFR4−/– mice are healthy and fertile (Weinstein et al. 1998). This finding by itself cannot reduce the importance of FGFR4 in the adult skeletal muscle because the action of a different FGFR may substitute for the absence of FGFR4 (Weinstein et al. 1998). Similar to the FGFR4−/– mice, the MyoD−/– and FGF6−/– mutant mice also do not show an unusual phenotype and can reach the adult stage. Nevertheless, in both mutants the muscles regenerate with decreased efficiency compared to wild-type mice (Megeney et al. 1996; Floss et al. 1997). In addition, we concluded that the transition of the satellite cells from proliferation to differentiation is suppressed in myofiber cultures from the MyoD−/– mouse (Yablonka-Reuveni et al. 1999a). Therefore, in vivo muscle injury studies of the FGFR4−/– mouse and analysis of satellite cells in FGFR4−/– myofibers may generate new insight into the role of FGFR4 in the adult muscle.
In conclusion, our present study provides evidence for the unique expression patterns of FGF6 and FGFR4 during myogenesis in postnatal muscle. To focus further on the functional role of FGF6 and FGFR4 in adult skeletal muscle, we have initiated studies on satellite cells from mutant mice lacking these two components of the FGF-FGFR system. The methods developed in the present study for localizing gene expression in myofibers and muscle-dissociated cells can now be utilized for analysis of other growth factors and cytokines that have been considered to be candidate regulators of myogenesis in the postnatal muscle.
Footnotes
Acknowledgments
Supported in parts by grants to ZY-R from the Cooperative State Research, Education, and Extension Service, U.S. Department of Agriculture (agreement no. 95-37206-2356 and 99-35206-7934) and by the National Institutes of Health (AG13798).
We thank Dr David Graves for designing many of the PCR primers used in the study and for his input during the preliminary stage of the investigation.