ARTICLE

Vascular Smooth Muscle Cells Spontaneously Adopt a Skeletal Muscle Phenotype: A Unique Myf5^-/MyoD⁺ Myogenic Program

Correspondence to: Zipora Yablonka–Reuveni, Dept. of Biological Structure, Box 357420, School of Medicine, U. of Washington, Seattle, WA 98195. E-mail: reuveni@u.washington.edu

	Summary

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Smooth and skeletal muscle tissues are composed of distinct cell types that express related but distinct isoforms of the structural genes used for contraction. These two muscle cell types are also believed to have distinct embryological origins. Nevertheless, the phenomenon of a phenotypic switch from smooth to skeletal muscle has been demonstrated in several in vivo studies. This switch has been minimally analyzed at the cellular level, and the mechanism driving it is unknown. We used immunofluorescence and RT-PCR to demonstrate the expression of the skeletal muscle-specific regulatory genes MyoD and myogenin, and of several skeletal muscle-specific structural genes in cultures of the established rat smooth muscle cell lines PAC1, A10, and A7r5. The skeletal muscle regulatory gene Myf5 was not detected in these three cell lines. We further isolated clonal sublines from PAC1 cultures that homogeneously express smooth muscle characteristics at low density and undergo a coordinated increase in skeletal muscle-specific gene expression at high density. In some of these PAC1 sublines, this process culminates in the high-frequency formation of myotubes. As in the PAC1 parental line, Myf5 was not expressed in the PAC1 sublines. We show that the PAC1 sublines that undergo a more robust transition into the skeletal muscle phenotype also express significantly higher levels of the insulin-like growth factor (IGF1 and IGF2) genes and of FGF receptor 4 (FGFR4) gene. Our results suggest that MyoD expression in itself is not a sufficient condition to promote a coordinated program of skeletal myogenesis in the smooth muscle cells. Insulin administered at a high concentration to PAC1 cell populations with a poor capacity to undergo skeletal muscle differentiation enhances the number of cells displaying the skeletal muscle differentiated phenotype. The findings raise the possibility that the IGF signaling system is involved in the phenotypic switch from smooth to skeletal muscle. The gene expression program described here can now be used to investigate the mechanisms that may underlie the propensity of certain smooth muscle cells to adopt a skeletal muscle identity. (J Histochem Cytochem 48:1173–1193, 2000)

Key Words: smooth muscle, skeletal muscle, Myf5, MyoD, myogenin, FGFR4, IGF1, IGF2

	Introduction

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Smooth and skeletal muscle cells are two of the cell types used by the vertebrate body to achieve mechanical contraction. Vascular smooth muscle cells provide structural support and help regulate internal pressure within the blood vessels of vertebrates. This cell type has been characterized by the expression of various smooth muscle isoforms of structural genes involved in contraction, including smooth muscle myosin heavy chains (smMHCs), smooth muscle calponin (smCalponin), smooth muscle -actin (sm--actin), smooth muscle myosin light chain kinase (smMLCK), and -tropomyosin (Fisher and Ikebe 1995 ; Owens 1995 ; Yablonka-Reuveni et al. 1998 ). Skeletal muscle cells express their own distinct isoforms of many of these genes, e.g., skeletal muscle myosin heavy chain, sarcomeric actin, skeletal muscle myosin light chain kinase (Bucher et al. 1988 ; Sutherland et al. 1993 ). Certain structural genes, such as the intermediate filament protein desmin, are expressed in both smooth and skeletal muscle cells (Lazarides 1982 ; Yablonka-Reuveni et al. 1998 ).

Although the structural gene isoforms are for the most part characteristic of each muscle cell type, some smooth muscle cell isoforms have been detected in developing or regenerating skeletal muscle cells (e.g., smMLCK, Dalla Libera et al. 1997 ; SM22, Li et al. 1996 ; sm--actin, Woodcock-Mitchell et al. 1988 ; Sawtell and Lessard 1989 ; Yablonka-Reuveni and Rivera 1994 ). In addition, sm--actin has been seen in the developing heart (Ruzicka and Schwartz 1988 ) and in various non-muscle cell types both in vivo and in culture (e.g., Leavitt et al. 1985 ; Jahoda et al. 1991 ). Therefore, the expression of sm--actin by itself is not a sufficient condition to warrant a smooth muscle cell identity. In addition, the set of smooth muscle structural gene isoforms being expressed by a given smooth muscle cell can change as a function of its environment (Owens 1995 ). Because of the variability in the expression of structural genes in smooth muscle cells, it has been generally held that for a cell to be characterized as a differentiated smooth muscle cell, one must demonstrate the expression of multiple smooth muscle-associated structural gene isoforms (Owens 1995 ). Importantly, the expression of smMHC is restricted to smooth muscle cells and has not been detected in other developing or adult tissues (Blank et al. 1995 , and references therein).

Skeletal muscle cells express a family of myogenic regulatory factors (MRFs) consisting of MyoD, Myf5, myogenin, and MRF4. The MRFs are involved in the specification of the skeletal myogenic lineage during early embryogenesis and in the control of the skeletal muscle differentiated phenotype. MyoD and Myf5 are expressed earlier during skeletal muscle development, followed by myogenin and MRF4 (reviewed in Ludolph and Konieczny 1995 ; Yun and Wold 1996 ). In cultures of skeletal muscle myoblasts, the expression of MyoD and Myf5 is seen first in proliferating cells, preceding the expression of the differentiation-linked factors myogenin and MRF4 (Yablonka-Reuveni et al. 1999a , and references therein). The MRFs have not been described for other muscle cell types, despite much examination (van Neck et al. 1993 ; Blank et al. 1995 ). A number of other transcription factors whose expression is not exclusive to skeletal muscle have been shown to influence skeletal muscle determination and differentiation, among them members of the MEF2, Id, and pax families (reviewed in Ludolph and Konieczny 1995 ; Yun and Wold 1996 ; Borycki and Emerson 1997 ; Firulli and Olson 1997 ). MEF2 family members appear to act synergistically with the MRFs to positively regulate skeletal muscle genes (Olson et al. 1995 ; Firulli and Olson 1997 ). Members of the Id family of transcription factors have been shown to compete with and antagonize the activity of the MRFs in skeletal muscle cells (Benezra et al. 1990 ; Melnikova and Christy 1996 ), but not to interfere with smooth muscle cell gene expression (Kemp et al. 1995 ). Pax3 has been shown to act upstream of MyoD in vivo and to contribute to the establishment of myogenic identity (Tajbakhsh et al. 1997 ; Borycki and Emerson 1997 ).

Although the smooth and skeletal muscle cells share some of the molecular underpinnings of their contractile roles, they are believed to have different developmental origins. The majority of skeletal muscle arises from discrete bodies of mesoderm sequestered and determined early in development (Christ and Ordahl 1995 ). The origin of smooth muscle is less defined. Many of the smooth muscle cells found in blood vessels are believed to arise from local mesoderm underlying the developing vasculature (Hungerford et al. 1996 ). A distinct set of vascular smooth muscle cells found in selected vessels, particularly the arteries arising from the embryonic aortic arches, have been shown in chicken to derive from the neural crest (Le Lievre and Le Douarin 1975 ; Yablonka-Reuveni et al. 1995 ). An endothelial cell origin for some vascular smooth muscle cells has also been proposed (DeRuiter et al. 1997 ).

Although smooth muscle cells and skeletal muscle cells are largely distinct in their structure, function, and developmental origin, cases have been described in which smooth muscle cells acquire a skeletal muscle phenotype. Patapoutian et al. 1995a demonstrated that a subpopulation of smooth muscle cells in the developing esophagus make such a smooth-to-skeletal muscle transition. This phenomenon has been further analyzed by Kablar et al. 2000 , who suggested that the transition is initiated by and dependent on the expression of myogenic regulatory factor Myf5. Similar smooth-to-skeletal muscle transitions have been reported to occur in the developing iris (Volpe et al. 1993 ; Link and Nishi 1998 ) and in the developing urethral sphincter (Borirakchanyavat et al. 1997 ). Myofibroblasts, which are smooth muscle-related cells present in various tissues (Serini and Gabbiani 1999 ), were also shown to express some of the skeletal muscle regulatory and structural proteins in vivo and in culture (Mayer and Leinwand 1997 ). In all of the above cases, the mechanisms driving the dual muscle identity and the extent of phenotypic penetrance are unclear.

The experiments detailed here demonstrate a change from smooth-to-skeletal muscle character occurring in three smooth muscle cell lines: PAC1, A10, and A7r5. Rothman et al. 1992 established the PAC1 line from pulmonary artery. The PAC1 line has been shown to maintain the capacity to express a repertoire of smooth muscle characteristic genes, including sm--actin, smMHC, myosin regulatory light chain, the smooth muscle isoform of -tropomyosin, and the surface receptors for angiotensin II, norepinephrine, and -thrombin. It has also been shown to direct the smooth muscle-specific splicing of transgenes and to exhibit normal morphology and smooth muscle myosin heavy chain expression at passage levels of up to 100 (Rothman et al. 1992 ). This cell line was further characterized by Firulli et al. 1998 , who demonstrated that PAC1 had a near-euploid chromosome set and expressed the smooth muscle characteristic genes sm--actin, smMHC, smCalponin, SM22, and tropoelastin. We obtained this cell line to serve as a negative control for immunofluorescence experiments conducted on primary skeletal muscle satellite cells, and quite unexpectedly found that cultures of PAC1 cells exhibited positive immunofluorescence for the MRFs MyoD and myogenin and formed myotubes. The A10 and A7r5 cell lines were established from rat thoracic aorta by Kimes and Brandt 1976 and have been used extensively as in vitro models of smooth muscle. The smooth muscle properties of these two lines have been further characterized more recently by Rao et al. 1997 and Firulli et al. 1998 . We detected MRF expression in cultures of these A10 and A7r5 cells as well, although to a much lesser degree than in the PAC1 cultures. A common feature of all three cell lines was the absence of Myf5. A further in-depth analysis of the PAC1 cells led us to establish in the present study a detailed description of a regulatory and structural gene expression program associated with the phenotypic transition from smooth to skeletal muscle.

	Materials and Methods

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Source of Cell Lines and Culture Conditions
The following smooth muscle, skeletal muscle and non-muscle rat cells were used.

PAC1 and PAC1R Cells. The PAC1 rat smooth muscle cell line was originally established from the pulmonary artery of adult Sprague–Dawley rats as described by Rothman et al. 1992 , and was obtained from two sources. A higher-passage culture of these cells was obtained from the laboratory of Dr. Stephen Schwartz at the University of Washington (PAC1 cells of a similar source have been described in Firulli et al. 1998 ). A low-passage culture of the PAC1 line, at passage 50, was kindly provided by the originator of this cell line, Dr. Abraham Rothman (University of California, San Diego). We designated these PAC1 cells as PAC1R. PAC1R cells were used at passage levels of 51 to 54. Subclones of the higher passage PAC1 cell line were generated in our laboratory by the technique of limiting dilution. The cells were cultured in 24-well plates at a density of 0–1 cell per well (i.e., the cells were diluted to a density of 1 cell per 1 ml medium and each well received 0.5 ml of the diluted cell suspension). Wells were monitored via microscopy and those displaying a single clone per well were further used for clonal expansion. The different clones were named with a letter or number suffix (e.g., PAC1a or PAC12).

A7r5 and A10 Cells. The smooth muscle cell lines A7r5 and A10 were originally derived from the thoracic aortas of embryonic BDIX rats as described by Kimes and Brandt 1976 , and were obtained from the American Tissue Type Collection (ATCC; Rockville, MD).

WKY3m22 Cells. The rat WKY3m22 cell isolate was derived from the thoracic aorta smooth muscle of a 3-month old Wistar–Kyoto rat (Gordon et al. 1986 ) and was obtained from the laboratory of Dr. Stephen Schwartz at the University of Washington.

ClB Cells. The ClB smooth muscle cells were isolated in the authors' laboratory from the pulmonary artery smooth muscle layer of an 8-week old Sprague–Dawley rat (B&K Universal; Kent, WA), using a modification of the method described by Gordon et al. 1986 . The pulmonary artery, cleaned of accessory tissue, was digested with shaking for 20 min at 37C in a solution containing: 0.5 mg/ml elastase (Sigma; St. Louis, MO), 1.5 mg/ml type I collagenase (Sigma), 0.5 mg/ml soybean trypsin inhibitor (Sigma), and 2.0 mg/ml bovine serum albumin (ICN; Costa Mesa, CA) in DMEM. The digested vessel was longitudinally cut, the inner smooth muscle layer was stripped from the outer connective tissue layers using forceps, minced, and then digested with shaking for 2 additional hr at 37C in the above enzyme solution. The tissue was then further dissociated by trituration and the released cells were washed and cultured in 100-mm plates using DMEM containing 20% fetal bovine serum. The culture that grew from this procedure was passaged once and then subjected to limiting dilution cloning. ClB (Clone B) cells, expanded to passage 4–7 in DMEM containing 10% fetal bovine serum, were eventually used in the present study.

L6 and L8 Skeletal Muscle Myoblasts. The L6 and L8 lines were originated by Dr. David Yaffe from the thigh muscle of newborn Wistar rats (Yaffe 1968 ; Yaffe and Saxel 1977a ) and were obtained from ATCC.

Rat2 Embryonic Fibroblasts. The rat embryonic fibroblast line Rat2, derived from the Rat1 cell line, also called F2408, and originally established from Fisher rat embryos (Topp 1981 , and references therein), was a gift from Dr. Daniel Bowen–Pope at the University of Washington.

NRK52e Rat Kidney Epithelial Cells. The NRK52e rat kidney epithelial cell line (De Larco and Todaro 1978 ) was also obtained from the laboratory of Dr. Daniel Bowen–Pope.

All cell types were grown in Dulbecco's modified Eagle's medium (DMEM; Gibco-BRL, Gaithersberg, MD) with 10% fetal bovine serum (Sigma) and antibiotics, with the exception that the A10 cells received 20% fetal bovine serum. Cells were plated for immunocytochemical studies at a density of 5000–10,000 cells/cm² in 35-mm tissue culture dishes and for RT-PCR analysis at a density of 20,000 cells/cm² in 60-mm tissue culture dishes.

When the effect of insulin was examined, the cells were cultured in DMEM with 10% fetal bovine serum in 35-mm plates as above until reaching about 90–95% confluency. The medium was then replaced with DMEM containing 10% fetal bovine serum or 2% horse serum (Sigma), with or without insulin (bovine; Sigma, added at 1 µM). The four different media were replaced every 2–3 days.

Immunocytochemistry
Single and double immunolabeling of methanol-fixed cultures were performed using a previously described indirect immunofluorescence protocol (Yablonka-Reuveni et al. 1999a ). The reactivity of the mouse monoclonal antibodies (MAbs) was monitored with fluorescein-conjugated goat anti-mouse IgG and the reactivity of the rabbit polyclonal antibodies was monitored with a rhodamine-conjugated goat anti-rabbit IgG. The secondary antibodies were from Organon-Technika Cappel (Downingtown, PA). After exposure to the secondary antibodies, the cultures were counterstained with DAPI to monitor cell nuclei.

The following primary antibodies were used.

Anti-MyoD. The mouse MAb 5.8A, made against murine MyoD (Dias et al. 1992 ), was kindly provided by its originators, Drs. P. Houghton and P. Dias (St. Jude Childrens' Research Hospital, Memphis, TN). The rabbit polyclonal antibody against rodent MyoD, whose characterization was described in Yablonka-Reuveni and Rivera 1994 , was developed and kindly provided by Dr. S. Alema (Institute of Cell Biology, CNR; Rome, Italy). The two antibodies costained the same MyoD⁺ cells in myogenic cultures from rat and mouse skeletal muscle and did not react with myoblasts from the MyoD null mouse (Yablonka-Reuveni and Rivera 1994 ; Yablonka-Reuveni et al. 1999a , Yablonka-Reuveni et al. 1999b ).

Anti-myogenin. The hybridoma producing the mouse MAb F5D made against rodent myogenin was developed and kindly supplied by Dr. W. Wright (University of Texas Southwestern Medical Center, Dallas, TX). The use of this antibody in immunofluorescence analyses of rat and mice myoblasts has been described (Yablonka-Reuveni and Rivera 1994 ; Yablonka-Reuveni et al. 1999a , Yablonka-Reuveni et al. 1999b ).

Anti-MEF2A. The C-21 rabbit polyclonal antibody against human MEF2A was purchased from Santa Cruz Biotechnology (Santa Cruz, CA). The use of this antibody to detect MEF2A in extracts of C2 cultures via immunoblotting was previously described by Molkentin et al. 1996 . Our cell culture studies of mouse myoblasts showed that this antibody stains nuclei of mononucleated cells after their expression of myogenin and all nuclei within myotubes (Yablonka-Reuveni and Rivera 1997 ; Yablonka-Reuveni et al. 1999a ). Similar results were obtained with myogenic cultures from adult rat muscle (Kastner et al. 2000 ).

Anti-sarcomeric Myosin. The mouse MAb MF20, raised against chicken sarcomeric myosin, was developed by Bader et al. 1982 . The antibody recognizes all isoforms of sarcomeric myosin heavy chain from multiple species, including rodents. The hybridoma supernatant was obtained from the Developmental Studies Hybridoma Bank (University of Iowa; Iowa City, IA).

Anti-smooth Muscle Calponin (smCalponin). The hCP mouse MAb prepared against human uterus smooth muscle calponin was obtained from Sigma. This MAb recognizes smooth muscle calponin from a variety of mammals and does not crossreact with skeletal muscle, cardiac muscle, or non-muscle forms of calponin.

Anti-smooth Muscle Myosin Light Chain Kinase (smMLCK). The K36 mouse MAb, made against chicken gizzard smooth muscle myosin light chain kinase, was obtained from Sigma. This MAb selectively recognizes the smooth muscle form of this kinase and has been used to distinguish between smooth and skeletal muscle cells of developing rodents (Patapoutian et al. 1995a ). We previously described the use of this antibody to detect chicken aortic smooth muscle cells (Yablonka-Reuveni et al. 1998 ).

Anti-smooth Muscle Myosin Heavy Chain (smMHC). A rabbit polyclonal antibody, made against the 204-kD isoform of bovine aortic smooth muscle myosin heavy chain, was kindly provided by the originators, Drs. C. Kelley and R. Adelstein (Kelley et al. 1992 ). This antibody recognizes smooth muscle myosin heavy chains from various species. We previously described the use of this antibody to detect chicken aortic smooth muscle cells (Yablonka-Reuveni et al. 1998 ).

Anti-smooth Muscle -Actin (sm--actin). The mouse MAb against sm--actin (Sigma; clone 1A4) was originally developed by Skalli et al. 1986 and has been shown to detect sm--actin in various species, including rodents (Yablonka-Reuveni and Rivera 1994 ; Yablonka-Reuveni et al. 1998 ).

Anti-desmin. The D3 mouse MAb against chicken desmin was developed by Danto and Fischman 1984 and was obtained from the Developmental Studies Hybridoma Bank. This antibody was shown to react with smooth and skeletal muscle cells from avian and rodent species (Allen et al. 1991 ; Yablonka-Reuveni et al. 1998 , Yablonka-Reuveni et al. 1999a ).

RT-PCR
RNA was isolated from cell cultures at the indicated day after plating using TriZOL reagent (Gibco BRL; Gaithersburg, MD) according to the manufacturer's instructions. After isolation the RNA was digested with DNase I (RQ1 DNase; Promega, Madison, WI) to eliminate trace amounts of genomic DNA. The treatment was with 1 unit DNase per 10 µg of RNA, in a buffer containing 40 mM Tris, pH 7.9, 10 mM NaCl, 6 mM MgCl₂, 10 mM CaCl₂, and 0.1 U/µl RNasin (Promega) for 30 min at 37C, followed by phenol/chloroform extraction to remove the enzyme.

Reverse transcription of the RNAs was done using Maloney murine leukemia virus reverse transcriptase (MMLV-RT; Promega) and Oligo dT₁₅ obtained as a custom primer from Gibco BRL. Two µg of RNA was reverse-transcribed in a 100-µl reaction containing 50 mM Tris-HCl, pH 8.3, 75 mM KCl, 3 mM MgCl₂, 10 mM dithiothreitol, 1 mM dNTPs, 19 µg/ml Oligo dT₁₅. After a denaturation step (95C, 2 min), this reaction was brought to 42C, 400 units of MMLV-RT was added, and the reactions were carried out for 1 hr at 42C, then heated to 90C for 10 min and stored at -20C until used in a PCR.

PCRs were performed in a Perkin–Elmer 9700 thermocylcer (Perkin–Elmer Applied Biosystems; Foster City, CA). Each 20-µl reaction received 2 µl of a reverse transcription reaction and contained the following reagents in the concentrations/amounts listed (i.e., these are the final concentrations including whatever the reverse transcription reaction contributed): 10 mM Tris-HCl, pH 8.3, 50 mM KCl, 2 mM MgCl₂, 300 µM dNTPs, 400 nM each primer, and 1 unit of enzyme (AmpliTAQ Gold; Perkin-Elmer). The standard PCR profile included a preincubation step, one cycle at 95C for 12 min, followed by 20–40 cycles of 95C for 45 sec, 65C for 59 sec, 72C for 2 min, and finishing with one cycle at 72C for 8 min. For the genes pax3 and smMLCK, an annealing temperature of 62C was used rather than 65C. Sixteen µl of the PCR was run on 1.2% agarose gels containing 0.5 µg/ml ethidium bromide (Gibco BRL) and viewed and photographed on a UV transilluminator.

The primers used in this study are shown in Fig 1. Most of these were designed de novo on the basis of published sequence information using the Entrez Browser program and with the help of the programs Primer and Tm. The primers were further evaluated with the help of the program Amplify (Engels 1993 ). In most cases, but especially in cases in which both smooth and skeletal muscle isoforms of a particular gene were known to exist, e.g., MLCK, MHC, and calponin, it was possible with the Amplify program to design primers to amplify the specific isoform desired and to yield no PCR products from known closely related genes. Primers were designed to the rat gene sequence when this was possible. In the cases of Myf5 and pax3, the primers were designed to the analogous mouse sequence. In the case of smMLCK, primers were designed to a region of consensus in the rabbit and human smMLCK genes. Three sets of primers were designed on the basis of previous work. The IGF2 primers were a variation on the primers used by Hannon et al. 1992 . The Myf5 primers were the same as used by Smith et al. 1994 . The Id-1 forward primer was the same as, and the reverse primer similar to, that used by Patapoutian et al. 1995b . All primers were purchased from Gibco BRL.

View larger version (46K): [in this window] [in a new window]   Figure 1. Summary of PCR primers used in the study.

	Results

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Culture Conditions That Elicit Morphological Change in PAC1 Cells
During preliminary studies we detected myotube-like structures in crowded PAC1 cultures. These myotube-like structures, although at times containing many nuclei, were not always readily apparent because of the high density of the single cells also present in the culture. When we employed immunofluorescence with an antibody against sarcomeric myosin, these myotubes were easily detected (see below). These multinucleated myotubes were not observed in sparse cultures. As indicated in Materials and Methods, our subsequent studies were conducted using plating densities of 5000–10,000/cm² for immunofluorescence studies and 20,000 cells/cm² for RT-PCR, and maintaining the cells in DMEM containing 10% fetal bovine serum. Under these conditions, myotubes appeared at 6–8 and 4–5 days after plating, respectively. Unless otherwise noted, all studies detailed below were conducted using these conditions.

Immunofluorescence Analysis of Smooth and Skeletal Muscle Protein Expression in PAC1 Cells
We first examined the PAC1 cells for their expression of a variety of smooth muscle associated proteins to verify their original characterization as a smooth muscle cell line that retains its capacity to differentiate in vitro (Rothman et al. 1992 ). Fig 2 shows the results of an immunofluorescence analysis with antibodies against various known smooth muscle proteins. The first four panels (Fig 2A–2D) show immunostaining of Day 4 cultures for smCalponin, smMLCK, desmin, and sm--actin. Fig 2E shows the sm--actin expression in a 10-day culture, demonstrating the appearance of myotube-like structures in the later PAC1 cultures and the expression of this protein in these myotubes.

View larger version (134K): [in this window] [in a new window]   Figure 2. Immunofluorescent micrographs of PAC1 cultures reacted with antibodies against various smooth and skeletal muscle proteins. (A–C) Day 4 cultures reacted with the MAbs against smCalponin, smMLCK, and desmin, respectively. (D) Day 4 and Day 10 cultures reacted with the MAb against sm--actin; the Day 10 culture demonstrates myotube-like structures. (F,F',F'') A Day 6 culture reacted with the monoclonal and polyclonal antibodies against MyoD (MyoD and MyoD(p), respectively; parallel arrows point out the position of the same cell's nucleus co-stained by the two antibodies. (G,G',G'') A Day 2 culture reacted with the MAb against myogenin and the polyclonal antibody against MyoD; parallel arrows point out the position of the same cell's nucleus co-stained by the two antibodies. (H,H',H'') A Day 10 culture reacted with the MAb against myogenin and the polyclonal antibody against MyoD; parallel arrows point out the same nucleus within a myotube. (I,I',I'') A Day 10 culture reacted with the MAb against sarcomeric myosin and the polyclonal antibody against MEF2A; parallel arrows point out a myotube with a myosin-positive cytoplasm and MEF2A-positive nuclei. In all cases, reactivity with the MAbs was detected with fluorescein-labeled goat anti-mouse IgG and reactivity with the polyclonal antibody was detected with rhodamine-labeled goat anti-rabbit IgG. Bar = 34 µm.

The bottom four panels of Fig 2 depict double immunofluorescent micrographs that show the expression of several skeletal muscle-specific proteins in PAC1 cells. Fig 2F shows the immunostaining of a Day 6 culture with mono- and polyclonal antibodies against MyoD. Most of the cells in this field are MyoD⁺ and co-stained with both antibodies. Fig 2G shows the co-expression of MyoD and myogenin in the same PAC1 cells in a Day 2 culture. Fig 2H shows a Day 10 culture containing multinucleated myotubes positive for both myogenin and MyoD. Many more myogenin⁺ cells were seen at the later time point. Fig 2I shows the expression of sarcomeric myosin in the same cells expressing the transcription factor MEF2A. The expression of myogenin, MEF2A, and sarcomeric myosin in the myotube-like structures is a strong indication that some of cells in the PAC1 cultures have adopted a differentiated skeletal muscle cell identity (Olson et al. 1995 ). A summary of the proteins detected in the PAC1 cultures using indirect immunofluorescence is shown in Table 1.

We next examined, via immunofluorescence, cultures of PAC1 cells obtained from the originator, Dr. A. Rothman (referred to as PAC1R), to see if the tendency of PAC1 cells to adopt a skeletal muscle phenotype was a general characteristic of this cell line. A summary of the PAC1R cell analysis is shown in Table 1 and examples of immunostained cultures are provided in Fig 3. Micrographs in Fig 3 were taken at a lower magnification than that used in Fig 2. As in PAC1 cells, many of the PAC1R cells were positive for smCalponin, desmin, smMLCK, and sm--actin. In addition, the PAC1R cultures contained a significant proportion of cells expressing MyoD, although at somewhat reduced numbers compared to the PAC1 cultures. Myogenin⁺ cells were also detected in PAC1R, but their frequency was far lower compared to PAC1 cultures. Many fields in the PAC1R cultures displayed no myogenin expression and a typical positive field contained only one or two myogenin⁺ cells. In extremely rare cases, we detected in the PAC1R cultures clusters of elongated myotube-like cells that stained positive for both sarcomeric myosin and MEF2A (myosin stain is shown in Fig 3F) or for myogenin and MEF2A (data not shown). In general, far less than 0.1% of the PAC1R cells expressed sarcomeric myosin.

View larger version (72K): [in this window] [in a new window]   Figure 3. Immunofluorescent micrographs of PAC1R cultures reacted with MAbs against various smooth muscle and skeletal muscle proteins. (A,A') A Day 3 culture stained with the anti-smCalponin. (B,B') A Day 3 culture stained with the anti-desmin. (C,C') A Day 3 culture stained with the anti-smMLCK. (D,D') A Day 7 culture stained with the anti-MyoD. (E) A Day 4 culture stained with the anti-myogenin. (F,F') A Day 13 culture stained with the antibody against sarcomeric myosin. For each panel, the position of the same cell detected by immunostaining and by DAPI is pointed out by arrows. The secondary antibody used for all panels was fluorescein-labeled goat anti-mouse IgG. Bar = 76 µm.

Collectively, the lower-passage cultures (PAC1R) and the higher-passage cultures (PAC1) of the pulmonary artery-derived cell line express the same repertoire of smooth muscle-specific proteins. In addition, both PAC1R and PAC1 cultures contain cells exhibiting skeletal muscle-characteristic proteins. However, the frequency of cells exhibiting the skeletal muscle differentiated phenotype is far reduced in the PAC1R cells compared to the PAC1 cells.

Expression of Smooth and Skeletal Muscle Differentiation-associated Genes in Smooth Muscle Cell Cultures
We used RT-PCR to compare smooth and skeletal muscle gene expression changes that take place during the PAC1 and PAC1R differentiation to those in established skeletal muscle cell lines (L6 and L8) and in non-myogenic lines (Rat2 and NRK52e). ClB cells were included in the comparison as a low-passage pulmonary smooth muscle line originated in our laboratory. This analysis was done on RNA taken from cells at 1, 5, and 10 days after plating, and the results are shown in Fig 4. The expression of smooth and skeletal muscle proteins in the different cell lines was also compared by immunofluorescene and is summarized in Table 1. In the RT-PCR analysis shown in Fig 4 (as in all subsequent RT-PCR analyses), the cycle number was chosen such that the best range of expression was observable among the various cell lines examined. Under these conditions, the level of some PCR products was saturated for certain cell types or time points. Lower PCR cycles allowed the comparison of the level of the more saturated PCR products. The last row in Fig 4 and in all of the other RT-PCR figures shows the expression of the ribosomal protein rpS6 in the various cell cultures. We used rpS6 expression as an indication of the quality of the RNA samples being used in the RT-PCR reactions and as a measure of the total reaction-to-reaction variation. The ribosomal protein genes are generally held not to be transcriptionally regulated in vertebrate cells (Jacobs et al. 1985 ), and this was demonstrated by us for the rpS6 gene in the cell types and under the conditions used in this study (data not shown).

View larger version (78K): [in this window] [in a new window]   Figure 4. RT-PCR analysis of skeletal and smooth muscle-specific gene expression in the PAC1 and ClB smooth muscle cell lines, the L6 and L8 skeletal myoblast cell lines, and the Rat2 and NRK52e non-myogenic cell lines. Cultures were harvested for RNA at the indicated times. RT-PCR was carried out to the indicated cycle numbers and the PCR products were resolved on agarose gels containing ethidum bromide. The resultant bands are presented as negative images of the original gels. The bands in a given row were from the same PCR run and used cDNAs generated in parallel reactions performed at the same time. The bands in a given column were made from the same cDNA reaction. The RT-PCR analysis of the ribosomal protein S6 (rpS6) housekeeping gene was included to ensure parallelism in reaction loading and execution. For each gene, the PCR cycle number used for the data shown in the figure was chosen such that the best range of expression was observable among the various cell lines examined.

As shown in Fig 4, PAC1R, PAC1, and ClB cells (but not the other lines examined) express the three smooth muscle genes: smMLCK, smMHC, and smCalponin. Consistently, a high number of PCR cycles was required to detect smMLCK even in cultures in which all cells were positive for smMLCK by immunofluorescence (data not shown). A trace of smMLCK was also detected in some of the non-smooth muscle lines at this PCR sensitivity.

As shown in Fig 4, PAC1R and PAC1 cultures gave robust MyoD bands with a tendency towards decreasing intensity with time. This trend was even more prominent at lower cycle numbers (data not shown). The ClB smooth muscle cells and the kidney epithelial NRK52e cells showed no expression of MyoD. Despite their skeletal muscle origin, the myogenic lines L6 and L8 expressed a far lower level of MyoD than the PAC1R and PAC1 cells. The L6 and L8 lines were previously reported not to express MyoD at all (Braun et al. 1989 ). Nevertheless, this low-level MyoD expression seen in the present study was also confirmed by immunofluorescence. As shown in Fig 5, infrequent (and often weakly stained) MyoD⁺ cells were detected in L6 cultures and a similar finding was made with L8 cultures (data not shown). The Rat2 fibroblasts, which were included as a non-myogenic control, also gave a strong MyoD signal by RT-PCR (Fig 4). This MyoD expression by the Rat2 cells, which do not have an obvious linkage with skeletal myogenesis (Magun and Bowden 1979 ; Leavitt et al. 1985 ), was also verified using immunofluorescence (Fig 5). Almost all Rat2 nuclei were intensely positive for MyoD at all times analyzed.

View larger version (118K): [in this window] [in a new window]   Figure 5. Immunofluorescent micrographs of L6 cultures (A,A') and Rat2 cultures (B,B') reacted with the MAb against MyoD. The secondary antibody was fluorescein-labeled goat anti-mouse IgG. Day 5 cultures were used for the L6 cells and Day 2 cultures were used for the Rat2 cells. In each panel, arrows point to the position of the same cell in the parallel MyoD and DAPI micrographs. Bar = 34 µm.

Myogenin was also expressed by the PAC1R and PAC1 cultures. In agreement with the immunofluorescence studies, the PAC1 cells expressed a far higher level of myogenin compared to the PAC1R cells. This level of myogenin expression the PAC1 cells was similar to the maximal level of myogenin seen in the skeletal myoblast lines L6 and L8 cells. The Rat2 cell line, which displayed robust MyoD expression but whose cultures were never seen to develop myotubes, gave only a trace band for myogenin. Immunofluorescence experiments confirmed the RT-PCR data with the Rat2 cells, showing myogenin expression in extremely rare cases (far less than 0.1% of cells). Neither the NRK52e nor the ClB cells showed detectable myogenin expression by RT-PCR or immunofluorescence (Fig 4; Table 1). MRF4 expression was also increased in the PAC1 cultures compared to PAC1R cultures. However, the intensity of the MRF4 signal produced by PAC1 cells was far reduced compared to that displayed by the differentiating L6 and L8 myoblasts. A faint MRF4 product was also detected in Rat2 and NRK52e cultures.

The Myf5 expression pattern was striking (Fig 4). Only L6 and L8 cells displayed Myf5 expression. This robust Myf5 expression in the L6 and L8 cultures was also demonstrated with an antibody against Myf5 (Yablonka-Reuveni et al. 1999a ). At 38 PCR cycles a trace of Myf5 expression was seen in the Rat2 and NRK52e cultures, but the PAC1, PAC1R, and ClB cultures did not display Myf5 expression even under these conditions (data shown later in Results).

The expression of the transcription factors Id-1 and pax3 is depicted next in Fig 4. The Id-1 gene product was displayed by all cell lines analyzed, showing a similar pattern of decreasing expression with days in culture (only one time point is shown in Fig 4 for Rat 2 and NRK52e cells). Different from Id-1, the Id-3 gene was expressed at similar levels by the cells at the time points shown in Fig 4 (data not shown). Pax3 expression was displayed by PAC1R cells at all times in culture and was rapidly decreasing in cultured PAC1 cells. This expression was far reduced compared to the level of pax3 in L6, L8 and NRK52e cells. Collectively, the Id-1, Id-3, and pax3 genes did not show explicit expression patterns that might be associated with the appearance of the skeletal muscle phenotype in the PAC1 smooth muscle cells.

The structural genes slow-twitch muscle troponin I (TnIsl), fast-twitch muscle troponin I (TnIfst) and muscle lim protein (MLP), which are associated with the differentiated state of skeletal muscle and myofiber specification (Bucher et al. 1988 ; Arber et al. 1994 ; Schneider et al. 1999 ), were expressed at a similar level in PAC1, L6, and L8 cells (Fig 4). The PAC1R cultures failed to show expression of these genes at this level of PCR sensitivity. This fitted well with our observation that only very infrequent cells in PAC1R cultures expressed sarcomeric myosin (Table 1). The expression of both insulin-like growth factors (IGF1 and IGF2) in PAC1 cultures also resembled their expression pattern in L6 and L8 cultures (Fig 4). At the PCR cycle levels used, PAC1R showed undetectable IGF1 expression and a trace of IGF2 expression at later times. ClB displayed a low level expression of IGF1 and no IGF2.

Our ongoing studies have linked the expression of fibroblast growth factor receptor 4 (FGFR4) and differentiation of skeletal muscle myoblasts. This led us to analyze FGFR4 gene expression in the present analysis. As shown in Fig 4, the expression of FGFR4 in PAC1 cells parallels the expression of IGF1 and IGF2 in a similar manner to the expression of these three genes in L6 and L8 cultures. FGFR4 is also expressed in PAC1R cultures but not in the other cell types examined. It is noteworthy that the L6 cells are widely held to lack FGF receptors based on FGF binding studies (Dell and Williams 1992 ). The different conclusions are likely due to the difference in the sensitivity of the FGFR detection means.

In summary, the study shown in Fig 4 indicates that the skeletal myogenic differentiation program seen in the rat smooth muscle cell line PAC1 shares many common features with that seen in the established rat skeletal muscle lines L6 and L8. Nevertheless, the PAC1 cells were never found to express Myf5. The PAC1R cells, the predecessors of the PAC1 cells, expressed MyoD and, to a lesser degree, myogenin, but did not display the expression of many of the genes linked to skeletal muscle differentiation.

Sublines Created from PAC1 Cultures Exhibit Variable Penetrance into the Skeletal Myogenic Phenotype
We undertook to clone out sublines from the PAC1 cultures to obtain homogeneous cell populations for further studies on the smooth-to-skeletal muscle phenotypic switch. Thirteen sublines produced by limiting dilution cloning were screened using immunofluorescence staining for myogenin, and we found that all expressed myogenin to various degrees. Some clonal cultures, such as PAC1i and PAC16, displayed many myogenin⁺ cells even during the initial days in culture. Other clonal cultures, such as PAC1a, contained only a small number of myogenin⁺ cells, which became more apparent during later days in culture. The frequency and size of the multinucleated myotubes also varied among the sublines, with the highest morphological differentiation seen in the PAC1i cells. The various degrees of expression of the skeletal muscle phenotype in the PAC1 sublines were an inherited property of the cells which was maintained even after prolonged storage in liquid nitrogen. This permitted the expansion of the subline cell stocks for the subsequent studies described below.

Fig 6 shows an RT-PCR analysis done on RNA from Day 5 cultures of the 13 PAC1 sublines along with the control lines PAC1R, L8, NRK52e, and ClB. The RT-PCR results verified the initial immunofluorescence characterization. On culture Day 5, all sublines except for PAC1a expressed MyoD and myogenin. Experiments conducted at the cycle numbers shown in Fig 6 revealed a wide range in the level of MyoD and myogenin expression among the sublines. A lower PCR cycle further revealed that the highest myogenin expression was in the PAC1i and PAC16 sublines. In all sublines, the expression level of myogenin nearly paralleled MyoD. Sublines that expressed more or less myogenin also expressed more or less MyoD. The three markers of skeletal muscle terminal differentiation, TnIsl, TnIfst, and MLP, are also shown in Fig 6. The expression level of these three genes in the individual PAC1 sublines followed a parallel pattern. Sublines that expressed more or less TnIsl also expressed more or less TnIfst and MLP. The PAC1i and PAC16 sublines (and to a slightly lesser extent the PAC1c line) expressed more of these genes than did the other sublines.

View larger version (66K): [in this window] [in a new window]   Figure 6. RT-PCR analysis of skeletal and smooth muscle-specific gene expression in clonal sublines of the PAC1 smooth muscle cell line. PAC1 sublines were generated by limiting dilution cloning and labeled a–k, 2–6. The sublines g, 3, and 5 did not survive serial passage and are not included. PAC1R is the parental cell line that gave rise to PAC1 and the sublines. The L8, NRK52e, and ClB cell lines were included as examples of skeletal muscle, non-muscle, and normal smooth muscle cells, respectively. Cultures were harvested for RNA on Day 5. RT-PCR was performed to the indicated cycle number. All other details are as in Fig 4. For each gene, the PCR cycle number used for the data shown in the figure was chosen such that the best range of expression was observable among the various cell lines examined.

The expression patterns of IGF1, IGF2, and FGFR4 in the clonal PAC1 sublines shown in Fig 6 are in agreement with the expression of these genes in the parental PAC1 line. The expression of IGF1 in the PAC1 sublines followed the same pattern as the skeletal muscle differentiation markers. The highest levels of IGF1 expression was seen in PAC1i and PAC16 sublines, followed by PAC1c cells. The second group of clonal cultures, PAC1f, PAC1h, and PAC12, which display a reduced expression of the differentiation markers TnIsl, TnIfst, and MLP, demonstrate a similar reduced expression of IGF1. The sublines that showed expression of TnIsl, TnIfst, and MLP (PAC1 Clones c, f, h, i, 2, and 6) showed the expression of IGF2, and FGFR4 in addition to IGF1.

The smooth muscle-specific genes smMLCK, smMHC, and smCalponin were expressed to some degree in all of the PAC1 sublines (Fig 6), supporting the idea that these clonal populations retained smooth muscle cell identity. A reciprocal trend was seen in the sublines' expression of smMHC and skeletal muscle structural genes. Sublines i and 6, which expressed the highest levels of TnIsl, TnIfst, and MLP, showed the least expression of smMHC.

Analysis of Id-I and pax3 expression, also shown in Fig 6, did not reveal any obvious correlation between the levels of these two genes and the skeletal muscle-specific genes among the PAC1 sublines. It is possible that the expression of Id-1 and pax3 was downregulated at different rates in the different PAC1 sublines and that the summation of these different patterns resulted in the overall downregulation of these genes in the parental PAC1 line (Fig 4).

PAC1i Cells Have a High Capacity to Undergo Skeletal Muscle Differentiation
The analysis shown in Fig 6, combined with observations on the degree of morphological skeletal muscle differentiation undergone by the PAC1 sublines, indicated that the PAC1i subline was exhibiting the greatest degree of skeletal muscle differentiation. Fig 7 shows phase-contrast micrographs of PAC1R and PAC1i cells at low and high densities. The low-density appearance of the PAC1i cells seen in Fig 7C is identical to that of low-density PAC1R cells shown in Fig 7A. The high-density PAC1i culture shown in Fig 7D was generated by allowing the culture to grow for 8 days beyond that shown in Fig 7C. The Day 10 PAC1i cells formed extensive arrays of multinucleated myotubes that accounted for most of the cells on the dish. This multinucleated array was positive for sarcomeric myosin, as determined by immunofluorescence with the MF20 antibody (data not shown). Fig 7B shows the high-density appearance of the PAC1R cells cultured for 6 days. This culture is devoid of multinucleated myotubes. The absence of myotubes was also confirmed in Day 10 PAC1R cultures (data not shown). Fig 7E and Fig 7F show immunofluorescence micrographs of Day 4 and Day 10 cultures of PAC1i cells stained for myogenin (a higher and a lower magnification for culture Days 4 and 10, respectively). Comparison of the pattern of myogenin expression with the corresponding pattern of DAPI nuclear staining (Fig 7E' and 7F') shows that a very high percentage of the PAC1i cells express myogenin and are fused in large myotube-like syncytia. Smaller myotubes containing circular clusters of nuclei were already seen in earlier days in culture.

View larger version (141K): [in this window] [in a new window]   Figure 7. Phase-contrast micrographs of PAC1 (A,B) and PAC1i (C,D) life cultures, and immunofluorescent micrographs of PAC1i cultures reacted with the anti-myogenin antibody (E,E',F,F'). The secondary antibody used for the immunofluorescence analysis was fluorescein-labeled goat anti-mouse IgG. (A,B) Day 2 and Day 6 cultures of PAC1R cells demonstrating the absence of visible myotubes in both sparser and more crowded cultures. (C,D) Day 2 and Day 10 cultures of PAC1i cells demonstrating the appearance of elaborated network of myotubes in the more crowded cultures. (E,F) Immunofluorescent micrographs of Day 4 and Day 10 cultures of PAC1i cells reacted with the anti-myogenin, respectively. Almost all nuclei in PAC1i cultures, whether in mononucleated cells (E) or multinucleated myotubes (F), are positive for myogenin. Bars: A–D = 140 µm; E = 45 µm; F = 76 µm.

Fig 8 shows a comparison of the time course of gene expression in the PAC1i subline and in the skeletal myoblast line L6. The general trend towards differentiation was similar in the PAC1i and L6 cultures, as demonstrated by the expression pattern of the skeletal muscle structural genes. The expression patterns of all genes analyzed in the PAC1i cultures are in agreement with the results shown in Fig 4 for the parental PAC1 line, with the exception of IGF1, whose expression is increased in the clonal subline. The comparison of IGF1 and IGF2 expression patterns in PAC1i cells demonstrated a strong induction of IGF1 with time in culture, paralleling the induction pattern of the skeletal muscle differentiation markers. Differently from IGF1, IGF2 was expressed in the PAC1i cultures at all time points studied, and its expression was only moderately upregulated with time in culture. In the study shown in Fig 8 (similar to the study shown in Fig 4), MRF4 expression level is far higher in the L6 cells compared to the PAC1 cells. It is further notable that the increase in MRF4 expression precedes the increase in myogenin expression. This observation contrasts with studies suggesting that MRF4 expression is linked to differentiation and is typically displayed coincidentally with or shortly after myogenin (Hinterberger et al. 1991 ; Cornelison and Wold 1997 ). In view of the recent findings that MyoD and MRF4 have overlapping functions (Rawls et al. 1998 ), it is possible that the temporal expression of MRF4 may differ for different myogenic cells and in different culture models.

View larger version (81K): [in this window] [in a new window]   Figure 8. Time course of expression of skeletal and smooth muscle-specific genes in PAC1i and L6 cell cultures. Cultures were harvested for RNA at the indicated times after plating. RT-PCR was performed to the indicated cycle number. RT-PCR analysis and data collating were conducted as described in Fig 4. For each gene, the PCR cycle number used for the data shown in the figure was chosen such that the best range of expression was observable among the two cell lines at the different time points examined.

Co-expression of Smooth and Skeletal Muscle Proteins in Individual PAC1 Cells
Double immunofluorescence analyses of PAC1R, PAC1, and PAC1i cells for the expression of proteins specific to the smooth and skeletal muscle cell phenotypes were carried out to determine whether the smooth and skeletal muscle genes were co-expressed in individual cells. The results of this study are shown in Fig 9. Fig 9A shows double staining of Day 4 PAC1R cells for smCalponin and MyoD. Fig 9B similarly demonstrates the co-expression of smMLCK and MyoD in single cells in Day 5 PAC1i cultures. Fig 9C shows double immunofluorescence staining of cells in a Day 3 PAC1 culture with antibodies specific for sarcomeric and smooth muscle myosins. In each case, arrows indicate single cells that stain for both the smooth and skeletal muscle-specific proteins. An arrowhead in Fig 9C indicates a cell that is stained only for a smooth muscle protein. In all cases, the number of cells in a given culture exhibiting double staining for smooth and skeletal muscle specific proteins was only a small proportion of the total cells, suggesting that perhaps this stage was an unstable intermediate between the two differentiated phenotypes.

View larger version (132K): [in this window] [in a new window]   Figure 9. Double immunostaining for smooth and skeletal muscle proteins in different PAC1 populations. Secondary antibodies were fluorescein-labeled goat anti-mouse IgG (left column) and rhodamine-labeled goat anti-rabbit IgG (middle column). (A,A',A'') A Day 4 culture of PAC1R cells reacted with anti-smCalponin and the anti-MyoD; arrows point to the position of cells that are positive for the two antibodies. (B,B',B'') A Day 5 culture of PAC1i cells reacted with anti-smMLCK and the anti-MyoD; arrows point to the position of cells that are positive for the two antibodies. (C,C',C'') A Day 3 culture of PAC1 cells reacted with anti-sarcomeric myosin (MF20) and the anti-smooth muscle myosin (sm); parallel arrows point to the position of a cell that is positive for both myosins; parallel arrowheads point to a cell that is positive only for smooth muscle myosin. (D,D',D'' and E,E',E'') Day 12 cultures of PAC1 cells reacted with anti-smMLCK and anti-MEF2A. (D) Parallel arrowheads point out to a large myotube that is negative for smMLCK and is identified by the multiple nuclei positive for MEF2A. (E) Parallel arrowheads point to the position of a small binucleated myotube whose cytoplasm is negative for smMLCK and whose nuclei are positive for MEF2A; parallel arrows point to a small binucleated myotube positive for both smMLCK and MEF2A. Bar = 34 µm.

Fig 9D and Fig 9E show the double immunofluorescence staining of Day 12 cultures of PAC1 cells for smMLCK along with the transcription factor MEF2A, which is included to mark the myotubes. Fig 9D shows a field of cells with distinct subpopulations exhibiting either smMLCK cytoplasmic staining or MEF2A-labeled nuclei in myotubes. The large MEF2A⁺ myotube marked by an arrowhead is clearly negative for smMLCK. Fig 9E contains a binucleated cell that stains positive for both smMLCK and MEF2A and other cells (an example is indicated with an arrowhead) that stain only for MEF2A. Our preliminary surveys indicated that whereas smaller myotubes in these cultures are often positive for the smooth muscle-associated proteins such as smCalponin and smMLCK, the expression of these proteins in the more extensive myotubes seen at later times was below the level of detection by this technique.

Vascular Smooth Muscle Cell Lines A7r5 and A10 Exhibit Skeletal Muscle Gene Expression
We next examined the two widely used rat aortic smooth muscle cell lines A7r5 and A10 for the possible expression of skeletal muscle differentiation genes. Fig 10 depicts the immunofluorescence analysis of the A7r5 cell line for smooth and skeletal myogenic protein expression. Fig 10A shows that many cells in Day 1 A7r5 cultures were positive for smCalponin. Fig 10B–10D show the staining at Day 5 for the skeletal muscle proteins: sarcomeric myosin (Fig 10B), MyoD (Fig 10C), and myogenin (Fig 10D). No large myotubes were observed in the A7r5 cultures, but elongated binucleated cells were infrequently seen that stained brightly for sarcomeric myosin. An example of this is shown in Fig 10E for a Day 11 culture. The frequency of cells in the A7r5 cultures expressing skeletal muscle characteristic proteins was far less than in PAC1 cultures (no more than 5% of the cells). By immunofluorescence, the A10 cell line was positive for smCalponin, MyoD, myogenin, and sarcomeric myosin, similar in pattern and frequency to the A7r5 cell line (data not shown). In agreement with a previous report (Gallagher et al. 1995 ), the A10 cultures contained significantly more smMLCK⁺ cells than the A7r5 cultures. The immunofluorescence results with the A7r5 and A10 lines are summarized in Table 1. Our RT-PCR studies further demonstrated that the A7r5 cells expressed a far higher level of smMHC than the A10 cells (data not shown) and that both cell lines expressed comparable levels of smCalponin (Fig 11).

View larger version (81K): [in this window] [in a new window]   Figure 10. Immunofluorescence micrographs of A7r5 cultures reacted with various MAbs. The secondary antibody was fluorescein-labeled goat anti-mouse IgG. (A,A') A Day 1 culture reacted with the anti-smCalponin. (B,B') A Day 5 culture reacted with the anti-sarcomeric myosin (MF20). (C,C') A Day 5 culture reacted with the MAb against MyoD. (D,D') A Day 5 culture reacted with anti-myogenin. (E,E') A Day 11 culture reacted with anti-sarcomeric myosin (MF20). For all panels, arrows point to the position of the same cell in the parallel antibody and DAPI staining micrographs. Bars: A,B = 76 µm; C–E = 34 µm.

View larger version (32K): [in this window] [in a new window]   Figure 11. RT-PCR cycle titration analysis of skeletal and smooth muscle-specific gene expression in smooth and skeletal muscle cell lines. Cells were plated at 20,000 cells/cm2 and harvested for RNA at Day 5 after plating. RT-PCR was performed to the indicated cycle number and the PCR products were analyzed and compiled as described in Fig 4.

The smooth muscle cells WKY3m22 were also analyzed by immunofluorescene and were found negative for all skeletal muscle proteins examined (Table 1). The WKY3m22 cells were derived from the thoracic aorta and have been used by several laboratories as a model for adult aortic smooth muscle cells (Gordon et al. 1986 ; Firulli et al. 1998 ). However, the present examination shows that only a small number of these cells are positive for the smooth muscle proteins smMLCK and smCalponin (Table 1).

The level of skeletal muscle gene expression in the different smooth muscle lines was additionally compared by an RT-PCR cycle titration analysis, as shown in Fig 11. The study was conducted using total RNA from Day 5 cultures. Except for the L6 cells, no cell line displayed Myf5 expression at any level of RT-PCR amplification. Neither the ClB nor the WKY3m22 cells showed expression of the MRFs or the skeletal muscle structural genes at any PCR cycle level, whereas the A7r5, A10, and PAC1R lines showed weak expression of these genes.

Effect of Insulin on Skeletal Muscle Differentiation in Vascular Smooth Muscle Cultures
The studies described above showed a correlation between the degree of skeletal muscle differentiation and the level of expression of the IGFs in PAC1 parental cultures and sublines. Furthermore, it is well established that endogenously produced IGF1 and IGF2 can exert a strong positive effect on skeletal muscle differentiation (Florini et al. 1991 ; Montarras et al. 1996 ; reviewed in Florini et al. 1995 ). Hence, the correlation between IGF1/IGF2 expression and skeletal muscle gene expression in the PAC1 smooth muscle cells could hint at the possible involvement of the IGFs in the smooth-to-skeletal muscle phenotypic switch displayed by the PAC1 cells. To further investigate this possibility, we asked whether a high concentration of insulin (1 µM) can facilitate the emergence of the skeletal muscle differentiated phenotype in the PAC1 smooth muscle cells. Insulin administered to skeletal myogenic cultures at such a concentration can activate the IGF signaling cascades by binding to the IGF receptors, whereas at a lower physiological concentration insulin is believed to act by binding to the insulin receptors (Ewton and Florini 1981 ).

The effect of insulin on differentiation was examined in selective PAC1 sublines showing higher and lower degrees of skeletal muscle differentiation. The parental PAC1R cells, along with ClB and Rat2 cells, were examined as well. Parallel cultures were initiated in 35-mm plates at 10,000 cells/cm² using the standard growth medium. When the cultures were almost confluent the medium was changed to DMEM-based media containing 10% fetal bovine serum (FBS) ± insulin or 2% horse serum (HS) ± insulin. Dishes from the four treatment groups were analyzed after 3–8 days in the test medium for the frequency of myogenin⁺ cells, employing immunofluorescence of DAPI-stained cultures. The results, which are summarized in Table 2, show that the frequency of myogenin⁺ cells was enhanced by insulin in cultures of both PAC1R cells and the PAC1a subline. The effect of insulin was seen regardless of whether the cells were maintained in 10% FBS or 2% HS. Differentiation in PAC1J cells and in PAC16 cells was dependent on the presence of insulin when cells were maintained in 2% HS but not when cells were maintained in 10% FBS. The frequency of myogenin⁺ cells in the PAC1i cultures was independent of insulin regardless of the serum conditions. We noted, however, that the size of the myotubes in PAC1i cultures was enlarged earlier when insulin was present (data not shown). This phenomenon is reminiscent of recent reports on the regulation of myotube hypertrophy by IGF1 in cultures of established skeletal muscle cell lines (Musaro et al. 1999 ). ClB cells did not show myogenin⁺ cells under any culture conditions, whereas extremely infrequent myogenin⁺ cells were seen in the Rat2 cultures at a late time point.

	Discussion

Top Summary Introduction Materials and Methods Results Discussion Literature Cited

In this study we have shown that cultures of three smooth muscle cell lines, PAC1, A7r5, and A10, contain cells that have begun to express a skeletal muscle cell phenotype. In the case of the PAC1 cells, several lines of evidence indicate that this phenomenon does not represent a simple contamination by skeletal muscle cells from an external source. First, all PAC1 sublines underwent skeletal muscle differentiation and also displayed expression of the smooth muscle-associated genes smMLCK, smMHC, and smCalponin. Second, two stocks of PAC1 cells (PAC1 and PAC1R) that had been obtained from different laboratories showed both skeletal and smooth muscle gene expression. Third, the double immunofluorescence data demonstrated that individual cells were expressing both smooth and skeletal muscle-specific genes. Most importantly, the absence of Myf5 expression uniquely distinguishes the different smooth muscle cell lines we have analyzed from the widely used rodent skeletal muscle lines that do express Myf5.

Whereas the skeletal muscle phenotype was characterized in the different smooth muscle cell populations by the expression of a common set of genes, the various cell lines and sublines expressed the skeletal muscle phenotype to a variable degree. The converted cells in the A7r5 and A10 populations were never seen to make large multinucleated myotubes, but instead remained mononuclear or made binucleated cells. Although a higher percentage of cells in PAC1R cultures expressed MyoD than did cells in A7r5 and A10 cultures (Table 1), the PAC1R cultures exhibited far less expression of the genes associated with skeletal muscle differentiation: sarcomeric myosin (Table 1), troponin I, and MLP (Fig 11). The clonal sublines generated from the high-passage PAC1 cultures also exhibited a wide range of capacities to express skeletal muscle genes. Certain sublines showed only weak expression of many of the myogenic genes and a poor capacity to form myotubes. In contrast, the PAC1i subline exhibited a pronounced phenotypic change over just a few days in culture. PAC1i cultures reliably produced myotubes at high frequency and exhibited an expression pattern of various skeletal muscle-associated genes. The differences in the penetrance of the various subpopulations of smooth muscle cells into the full myogenic phenotype might represent discrete stages in the transition from a smooth to the skeletal muscle identity.

A dual smooth/skeletal muscle program was previously identified in the B3CH1 cell line. This line was derived from a mouse brain tumor and its cell origin is unknown. Nevertheless, on the basis of their expression of sm--actin along with several other characteristics, the B3CH1 cells have been considered to be smooth muscle cells (Schubert et al. 1974 ; Strauch and Rubinstein 1984 ). These BC3H1 cells were also found to express several skeletal muscle-specific traits, and therefore have also been thought to be related to skeletal myoblasts, although the cells do not undergo a full differentiation and do not fuse into myotubes (Braun et al. 1989 ; Block and Miller 1992 , and references therein). As detailed in the Introduction, sm--actin is expressed by various cells, skeletal myoblasts included, and therefore the expression of this actin by itself is insufficient to attribute a smooth muscle nature to the B3CH1 cells. Unlike the B3CH1 cells, which were isolated from a mouse brain tumor and whose actual origin is unknown, the PAC1, A7r5, and A10 cells were isolated from vascular smooth muscle and express multiple smooth muscle genes. Furthermore, the expression of MyoD but not Myf5 by the three vascular-derived smooth muscle lines analyzed in the present study distinguishes the cells from the B3CH1 cells, which express Myf5 but not MyoD (Braun et al. 1989 ; Block and Miller 1992 ).

MyoD Expression by Itself is Insufficient for the Promotion of the Skeletal Muscle Phenotype in Vascular Smooth Muscle Cells
MyoD was among the genes showing differential expression in the different smooth muscle lines and within the PAC1 sublines. In most but not all cases, the level of MyoD expression was generally a good predictor of the level of expression of downstream skeletal muscle genes such as myogenin and troponin I. Hence, a simple model explaining the expression of skeletal muscle genes in smooth muscle cell lines would postulate that the MyoD gene became transcriptionally active in the cultured cells and that this "leaky" expression of MyoD caused the subsequent change in phenotype. Indeed, enforced expression of MyoD is able to foster a skeletal muscle phenotype in many but not all non-muscle cell types (Weintraub et al. 1989 ; Schafer et al. 1990 ; Choi et al. 1990 ). However, we demonstrated exceptions to the MyoD level predicting the extent of terminal differentiation found in the PAC1 sublines and other cells. As summarized in Table 1 and Fig 11, PAC1R smooth muscle cells and even Rat2 embryonic fibroblasts expressed more MyoD than did A7r5 cultures, yet the A7r5 cultures expressed more sarcomeric myosin and troponin I. The Rat2 expression of MyoD without a subsequent expression of skeletal muscle differentiation markers underscores the concept that a high level of MyoD expression is not necessarily a sufficient condition to promote a fully differentiated skeletal muscle phenotype. The inability of Rat2 cells to enter the myogenin⁺ differentiated state despite MyoD expression has been also observed by George-Weinstein et al. 1998 . In the case of P19 cells, it has been demonstrated that cell–cell interactions are required to enable MyoD-expressing cells to undergo myogenic differentiation (Armour et al. 1999 ). However, in our studies even the most crowded PAC1R or Rat2 cells did not demonstrate the expression of skeletal muscle differentiation genes under standard culture conditions, regardless of high-level expression of MyoD. The studies by Schafer et al. 1990 proposed that additional regulators present in some but not all cell types may be necessary for activation of a skeletal muscle-specific program in MRF-expressing cells.

Investigating the different PAC1 cultures, we showed that the degree of skeletal muscle differentiation correlates with the increased expression of the IGFs and the growth factor receptor FGFR4. We also demonstrated that expression of IGF2 precedes expression of IGF1. Hence, IGF2 might be involved in the initiation of the skeletal myogenic program, and IGF1 expression might be the result of skeletal muscle differentiation in the PAC1 cultures. It is notable that Florini et al. 1991 demonstrated that the autocrine production of IGF2 supports spontaneous differentiation in a rodent myogenic cell line, and Montarras et al. 1996 provided a precedent for a positive effect of IGF2 on MyoD expression during skeletal myogenesis, even at the proliferating myoblast stage. A model involving the IGFs in the smooth-to-skeletal muscle transition is attractive because localized synthesis of IGFs has been broadly implicated in skeletal muscle growth, hypertrophy and regeneration (reviewed in Florini et al. 1995 ; Musaro et al. 1999 ).

The present study on the effect of insulin in PAC1 cultures provides further support for the possible involvement of the IGF signaling system during the smooth-to-skeletal muscle switch. We showed that different PAC1 populations possess a capacity to switch into the myogenin⁺ state, depending on the environment they are in. This switch in PAC1R and PAC1a was highly dependent on the addition of insulin, possibly because the cells made very little of the IGFs. The switch to the myogenin⁺ state in PAC1J and PAC16 cells required insulin only when the cells were maintained in low serum. Therefore, the PAC1J and PAC16 cells might produce (or receive from the medium) sufficient IGFs in the high-serum conditions but not in the low-serum conditions. The PAC1i cells probably produced sufficient IGFs under both serum conditions. Alternatively, different levels of the IGF receptors among the various sublines could also account for some of the observations, but that aspect has not been yet investigated. PAC1R and PAC1a cells might express only a low level of the receptors, and therefore a high concentration of insulin would be required for activating the IGF receptors. PAC1J and PAC16 cultures might express a lower level of the receptors when maintained in the low serum and therefore would need the addition of insulin at a high level to initiate IGF signaling and enhance skeletal muscle differentiation in the low-serum conditions only. PAC1i cells might express a sufficient level of the IGF receptors along with a sufficient level of the IGFs in both high and low serum.

The role of FGFR4 in skeletal myogenesis is under investigation in our laboratory. FGFR4 expression in adult myoblasts (satellite cells) correlates well with the progression of the cells through the MyoD and myogenin compartments (Kastner et al. 2000 ). In addition, primary cultures of myoblasts from mice lacking MyoD display a delayed expression of FGFR4, correlating with a delayed expression of differentiation-linked genes and fusion into myotubes (unpublished results). In the present study, the cell lines that did not display the smooth-to-skeletal muscle switch did not express FGFR4. The Rat2 cells that expressed MyoD but not FGFR4 did not display the skeletal muscle phenotype. In contrast, the PAC1R line that expressed MyoD and had some potential to undergo skeletal muscle differentiation that could be enhanced in the presence of insulin expressed FGFR4. It is therefore possible that the capacity to express FGFR4 is involved in the capacity of the smooth muscle cells to undergo skeletal myogenesis.

Myf5 Is Not Expressed During the Smooth-to-skeletal Muscle Transition
One of the more notable characteristics of our data was the observation that Myf5 expression was not activated in any smooth muscle cell lines or even in the PAC1i subline, which showed the most robust phenotypic conversion. Primary cultures of skeletal muscle myoblasts typically express both MyoD and Myf5, the latter being expressed at various levels depending on the source of the cells (reviewed in Yablonka-Reuveni et al. 1999a ). In skeletal muscle-derived myogenic cell lines established spontaneously (i.e., without the inclusion of drugs in the culture medium or the forced expression of regulatory genes, e.g., C2 mouse myoblasts and L8 rat myoblasts; Yaffe and Saxel 1977a , Yaffe and Saxel 1977b ), MyoD is expressed at various levels and Myf5 is always present (Braun et al. 1989 ; Yablonka-Reuveni and Rivera 1997 ; Kitzmann et al. 1998 ; and the present study). Nevertheless, in vivo gene inactivation studies have established that skeletal myogenesis can progress in the absence of Myf5 as long as MyoD is present (Rudnicki et al. 1993 ).

During development, it is believed that MyoD and Myf5 are initially induced in defined and separate sets of skeletal muscle cell precursors (Braun and Arnold 1996 ; Cossu et al. 1996 ). MyoD-expressing muscle cell precursors in the somites are believed to be induced by factors in the dorsal ectoderm, whereas a Myf5-dependent lineage of muscle cell precursors are believed to be induced by the neural tube, notochord, and ventral structures (Cossu et al. 1996 ). Therefore, the embryonic precursor cells that gave rise to the A7r5, A10, and PAC1 cell lines might have been exposed to a MyoD-inducing field that had disposed them to a latent MyoD⁺/Myf5^- myogenic identity. This could be a special case of what Cossu has termed "unorthodox myogenesis" (Cossu 1997 ). Alternatively, the lack of Myf5 expression in the vascular smooth muscle cell lines might reflect an inherent difference in the inducibility of the MyoD and Myf5 genes. For example, it has been noted that ectopic expression of MyoD or the other MRFs in 10T1/2 cells induced the expression of endogenous MyoD, myogenin, and MRF4 but not the endogenous Myf5 gene (Aurade et al. 1994 ).

Possible In Vivo Role of the Smooth-to-skeletal Muscle Switch
The fact that not all of the smooth muscle cell populations analyzed in the present study possess the capacity to undergo skeletal muscle differentiation suggests that skeletal myogenic potential is acquired in culture, potentially by initiating MyoD expression and activating the IGF signaling system. Nevertheless, the expression of skeletal muscle genes or the appearance of the myogenic phenotype has been noted in several developing smooth muscle tissues, including the esophagus, the iris, and the urethral sphincter (Volpe et al. 1993 ; Patapoutian et al. 1995a ; Borirakchanyavat et al. 1997 ; Kablar et al. 2000 ). Therefore, the capacity of the smooth muscle cells to undergo skeletal muscle differentiation might be imprinted in the vascular smooth muscle cells in vivo. Skeletal muscle genes are also expressed in the thymus (Grounds et al. 1992 ; Wong et al. 1999 ) and the pineal organ (Watanabe et al. 1992 ), although in these cases the cell type contributing the skeletal muscle phenotype has not been established. Vascular smooth muscle cells have been shown to undergo a phenotypic transition into the osteoblastic cell lineage (Balica et al. 1997 ). Pericytes, smooth muscle-like cells associated with all vascular capillary and post-capillary venules, have been reported to differentiate into a variety of additional cell types, including osteoblasts, chondrocytes, adipocytes, and phagocytes (reviewed in Hirschi and D'Amore 1996 ; Doherty and Canfield 1999 ). Various observations indicate that the primordial endothelium can recruit undifferentiated mesenchymal cells and direct their differentiation into pericytes in microvessels and smooth muscle cells in large vessels (reviewed in Hirschi and D'Amore 1996 ). This origin from undifferentiated mesenchymal cells may enable pericytes and smooth muscle cells to maintain a multipotent cell potential, one of which might be the skeletal muscle phenotype. Indeed, a recent study has reported on the isolation of skeletal muscle myoblasts from the developing dorsal aorta. These aorta-derived myoblasts can participate in postnatal muscle growth and development (De Angelis et al. 1999 ). This study further proposed that at least some of the adult skeletal muscle myoblasts might derive from such an aortic source rather than from the somites.

We have begun to explore the possible occurrence of a smooth-to-skeletal muscle phenotype within the elaborated skeletal muscle vasculature as an alternative source for skeletal muscle myoblasts in vivo. Such a source may provide a means to produce skeletal muscle myoblasts after a massive injury, when there is a demand for a large and rapid supply of myoblasts for tissue repair (reviewed in Grounds and Yablonka-Reuveni 1993 ). We have recently isolated a cell phenotype from the adult rat skeletal muscle that expresses smooth muscle structural genes in the same pattern as seen in ClB smooth muscle cells and that also expresses MyoD. Other markers of skeletal muscle differentiation were not detected. These cells responded mitogenically to both PDGF-BB and FGF2, and expressed a far lower level of IGF1 and FGFR4 compared to the classical adult skeletal muscle precursors (i.e., satellite cells) under the same culture conditions. Although the source of this unique cell phenotype in the muscle tissue has not yet been determined, its morphology and growth pattern in culture resembles that of vascular smooth muscle cells (unpublished results). Recent cell transplantation studies have determined that bone marrow cells and mesenchymal stem cells from various non-muscle tissues possess the capacity to participate in skeletal muscle regeneration (reviewed in Bianco and Cossu 1999 ; Seale and Rudnicki 2000 ). Whether these cells are derived from the smooth muscle of the vasculature, myofibroblasts, or other cellular sources awaits further investigation.

In conclusion, the present cell culture study provides evidence for a phenotypic transition from smooth to skeletal muscle and a detailed examination of the gene expression program associated with this transition. The observations made in the present study can now be used to begin investigating mechanisms that underlie the emergence of the skeletal muscle phenotype in various tissues and organs that do not have an obvious developmental linkage to the skeletal muscle lineage.

	Acknowledgments

Supported in part by grants to Z.Y.-R. from the National Institutes of Health (AG13798) and from the Cooperative State Research Service–US Department of Agriculture (agreement no. 95-37206-2356).

We thank Priscilla Natanson for excellent technical support. We also thank Stefanie Kästner for helpful comments on the RT-PCR analysis and Dr D. Han for important input during the preliminary stage of the investigation. We are grateful to the following investigators who kindly provided valuable reagents: Dr A. Rothman (PAC1R cells); Drs D. Han and S. Schwartz (WKY3m22 and PAC1 cells); Drs R. Seifert and D. Bowen–Pope (Rat2 and NRK52e cells); Drs R. Adelstein and C. Kelley (anti-smMHC); Dr S. Alemá (anti-MyoD, polyclonal); Drs P. Houghton and P. Dias (anti-MyoD, monoclonal); and Dr W. Wright (anti-myogenin).

Received for publication March 15, 2000; accepted May 17, 2000.

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