Reagents were purchased from Sigma-Aldrich (Milan, Italy), unless otherwise stated. Cell culture materials were purchased from Jet-Biofil (Carlo Erba Reagents-Dasit Group, Cornaredo, Italy).

Human RD (eRMS) and RH30 (aRMS) cells were purchased from the European Collection of Cell Cultures (ECACC; Salisbury, UK). The primary tumor mouse cultures, U57810 (eRMS) and U23674 (aRMS), were established from transgenic mice (41). In particular, eRMS mouse models were generated by crossing p53- or Ptch1-deficient conditional mice with Myf6-Cre mice to achieve the deletion of p53 with or without concurrent Ptch1 deletion in differentiating Myf6-positive myoblasts. The Myf6Cre/p53^−/− mouse strain was characterized by the highest percentage of eRMS (41). aRMS mouse models were characterized by knockout alleles of p53 or INK4a/ARF locus with concomitant Pax3-FOXO1 knock-in allele, which were restricted to differentiating Myf6-positive myoblasts, resulting in the Myf6Cre/Pax3-FOXO1/p53^−/− mouse strain (41).

Cells were maintained at 37°C and 5% CO₂ in a humidified incubator and cultured in a growth medium (GM) comprising high-glucose Dulbecco’s modified Eagle’s medium (DMEM) (D6429) supplemented with 10% fetal bovine serum (FBS) (FA30A15101; Carlo Erba, Milan, Italy) in the presence of 100 µg/ml penicillin/streptomycin (A5955) and 1% L-glutamine (G7513) (only for RH30 cells). To induce myodifferentiation, 80% confluent cells were switched to differentiation medium (DM) comprising DMEM supplemented with 2% horse serum (H1270).

Cells were treated with different concentrations (0.01, 0.1, 1, and 2 mM) of melatonin (461326) or vehicle alone [dimethylsulfoxide (DMSO)] (D5879). The cells were also treated with melatonin in the absence or presence of chemotherapeutic drugs such as doxorubicin (0.15 ng/ml) (D1515) and cisplatin (2 µg/ml) (P4394), which were previously diluted in H₂O and DMSO, respectively.

eRMS and aRMS cells were seeded in 24-well plates at a density of 10×10³ and 15×10³, respectively. After 24, 48 and 72 h of melatonin treatment, the cells were harvested, fixed in paraformaldehyde (F8775) and stained for 10 min with crystal violet (C0075) solution [0.2% in phosphate-buffered saline (PBS) (D8537) with 20% methanol (32213)]. The samples were then collected in 600 µl of SDS (74255) solution (1% in PBS) and absorbance of the total homogenates, as measured by reading the plate at 540 nm emission wavelengths, was proportional to the amount of viable and proliferating cells that incorporated the crystal violet. In addition, the cell proliferation was expressed as the growth rate, which was calculated using Microsoft Excel 2010 software. The results were representative of at least three independent experiments.

A neutral red assay was employed to determine the percentage of viable cells that incorporated the neutral red dye in lysosomes, as initially described by Borenfreund and Puerner (42), a protocol subsequently modified by Repetto et al (43). Briefly, the cells were seeded in 96-well plates at a density of 1.5×10³. After 24, 48 and 72 h of melatonin treatment, the cells were incubated for 2 h with neutral red dye (40 µg/ml) (N7005) dissolved in DMEM with 5% FBS. After washing the cells with PBS, 150 µl of neutral red destaining solution [50% ethanol (02860), 49% deionized water, and 1% glacial acetic acid (100015N; BDH Laboratory Supplies, Dawsonville, GA, USA)] was added, followed by gentle agitation for 10 min, until complete dissolution was achieved. Absorbance was then measured by reading the plate at 540 nm emission wavelengths. The results were analyzed using Microsoft Excel 2010 software and presented as the percentage of control values. Images of cell viability assays showed representative results of at least three independent experiments.

Protein homogenates were obtained by harvesting cells in a cold RIPA lysis buffer, comprising 20 mM Tris-HCl (ph 7.6) (T1503), 1% Nonidet P40 (NP40), 0.5% sodium deoxycholate (D6750), 0.1% SDS (74255), 50 mM NaCl (S7653) and a cocktail of protease inhibitors (1836153; Roche, Milan, Italy) plus phosphatase inhibitors [1 mM Na₃VO₄ (S6508) and 4 mM NaF (S7920)]. The protein concentration was calculated by a Bradford reagent (B6916) assay and an equal amount of protein samples was separated by SDS-PAGE under reducing conditions and transferred to polyvinylidene fluoride membranes (P2938). Incubation with specific primary antibodies was followed by peroxidase-conjugated secondary antibodies (goat polyclonal anti-mouse IgG-HRP sc-2005; from Santa Cruz Biotechnology, Inc., Dallas, TX, USA; donkey polyclonal anti-rabbit IgG no. 31458; from Thermo Scientific, Erembodegem, Belgium) and the resulting immune complexes were visualized using the enhanced chemiluminescence reagent (STS-E 500; GeneSpin, Milan, Italy). Immune-reactive bands were quantified using densitometry analyses (Software Gel-Pro Analyzer, version 4).

The antibodies were purchased from Santa Cruz Biotechnology, Inc., unless otherwise stated. The primary antibodies used were: mouse monoclonal anti-myosin heavy chain, (sc-32732; 1:1,000 dilution); mouse monoclonal anti-caveolin-3 (610420; 1:1,000 dilution; BD, Buccinasco, Italy); rabbit polyclonal anti-caspase-3 (H-277) (sc-7148; 1:500 dilution; Cell Signaling, Milan, Italy); rabbit polyclonal anti-Bax (sc-526; 1:500 dilution); rabbit polyclonal anti-Bcl-2 (sc-492; 1:500 dilution); and mouse monoclonal anti-α-tubulin (T5168; 1:10,000 dilution; Sigma-Aldrich).

RH30 cells were cultured and treated directly on coverslips in Petri dishes. After washing with 0.1 M phosphate buffer, adherent and suspended cells were fixed with 2.5% glutaraldehyde (G5882) in 0.1 M phosphate buffer for 1 h. The suspended cells adhered to polylysine-coated coverslips. The samples were post-fixed with 1% osmium tetroxide (OsO₄) (O021; Strumenti, Roma, Italy) in 0.1 M phosphate buffer for 1 h. After alcohol dehydration, the samples were critical point dried, gold sputtered and observed using a Philips 515 scanning electron microscope (FEI, Italy) (44).

RH30-treated cells were washed and fixed with 2.5% glutaraldehyde (G5882) in 0.1 M phosphate buffer for 15 min. The cells were scraped and centrifuged at 300 × g for 10 min. The pellets were fixed in 2.5% glutaraldehyde for an additional 30 min. The suspended cells were collected in Eppendorf, centrifuged and fixed for 45 min in glutaraldehyde. The samples were post-fixed in 1% OsO₄ (O021) for 1 h, alcohol dehydrated and embedded in araldite (02860) (45). Thin sections were stained with uranyl acetate and lead citrate and analyzed using a Philips CM10 transmission electron microscope (FEI).

Adherent cells were cultured and treated directly on coverslips in Petri dishes. The suspended cells were collected in Eppendorf, fixed in 4% paraformaldehyde (F8775) for 30 min and then plated on polylysine-coated coverslips. The cells were then fixed in 4% paraformaldehyde in PBS (ph 7.4) for 30 min and washed twice using PBS. The cells were then pre-treated with RNase A (10 µg/ml) (12091-021) in PBS for 30 min and exposed to an equal mixture of propidium iodide (PI; 1 µg/ml) (P3566) and acridine orange (AO; 1 µg/ml) (A3568) (all from Life Technologies, Monza, Italy) in PBS at room temperature in the dark for 10 min.

AO and PI are intercalating fluorochromes that emit green and red fluorescence, respectively, when they are bound to DNA. Only AO diffuses through the plasma membrane of both viable and early apoptotic cells. Viable cells exhibit a green nucleus with intact structure, while apoptotic cells exhibit a bright-green nucleus, with condensation of chromatin. PI only enters late apoptotic and necrotic cells, resulting in double staining with AO and PI (45). The samples were observed with a Leica TCS-SP5 CLSM connected to a DMI 6000 CS inverted microscope (Leica Microsystems CMS Gmbh; AO and PI excitation were at 488 and 500 nm, respectively, and their emission signals were detected at 617 and 525 nm, respectively).

The differences between the groups were analyzed by the unpaired Student’s t-test and one-way ANOVA test (with Dunnet’s post-hoc test), using Prism 4 software for Windows (GraphPad Software, San Diego, CA, USA). P<0.05 was considered to indicate a significant result.

We evaluated whether melatonin administration would influence the cell growth of the human RMS cell lines (i.e. embryonal RD and alveolar RH30) and primary mouse tumor cultures (i.e., embryonal U57810 and alveolar U23674). For this purpose, the proliferation of cells that received melatonin once was determined over a time-course of 72 h by means of crystal violet assay. Treating different lines with increasing concentrations of melatonin, ranging from 0.01 to 2 mM, led to a significant impairment of cell proliferation starting from a dose of 1 mM in comparison to vehicle-treated cells, as indicated after calculation of the growth rate (Fig. 1A). To determine whether the melatonin effects were attributable to the inhibition of cell proliferation rather than impaired cell viability, we performed the neutral red assay using the cell lines under the same experimental conditions. As shown in Fig. 1B, 72 h of exposure with a concentration of 1 or 2 mM triggered the loss of ~50% of RMS cells in comparison to vehicle-treated cells, indicating that melatonin has cytotoxic effects. This latter result was confirmed by immunoblotting analysis of Bax and Bcl-2 expression, two proteins that can be either pro- or anti-apoptotic, respectively (46–48).

As shown in Fig. 2, treatment of RD and RH30 cells with 1 mM melatonin promoted an increase in the pro-apoptotic Bax expression, while the expression levels of anti-apoptotic Bcl-2 were downregulated in comparison to those in untreated cells. These results indicated that melatonin, not only behaves as a cytostatic factor on RMS cell growth, but also impairs the survival of different RMS lines by triggering an apoptotic program. Thus, at the ultrastructural level, the pro-apoptotic effects of melatonin, RH30 cells treated or untreated were analyzed by electronic and confocal microscopy.

Control cells showed an obvious healthy morphology characterized by the presence of intact subcellular structures, as observed by means of SEM (Fig. 3A) and TEM (Fig. 3B and C), respectively. In addition, AO/PI double staining showed a uniform green labeling suggestive of cellular healthy structures (Fig. 3D). After 1 mM melatonin for 24 h a heterogeneous situation developed: some cells maintained good cell viability similar to the control condition, while other cells showed a round apoptotic-like morphology (Fig. 3E and F). In particular, TEM analyses revealed some cells with an intense chromatin condensation, a typical apoptotic pattern (Fig. 3G). At the confocal microscopy level some cells appeared rounded and early apoptotic features were evident (Fig. 3H). After 48 h, melatonin-treated cells were almost all detached showing a round apoptotic morphology while only a small number of adherent cells exhibited an atrophic behavior, due to cytoplasm shrinkage (Fig. 3I). In addition, rounded cells suggestive of necrotic features also appeared (Fig. 3J). These cells were characterized by typical apoptotic features, including the presence of chromatin condensation, cytoplasm vacuolization and secondary necrosis as confirmed by TEM analysis (Fig. 3K). Consistent with this, an increased number of apoptotic cells showed orange areas due to PI permeability suggestive of cells in late apoptosis (Fig. 3L). After 72 h, the melatonin-treated cells observed at the SEM level exhibited apoptotic and necrotic features, being completely detached and showing a rounded morphology, with disruption of cell membranes in those that were necrotic (Fig. 3M and N). As shown by TEM, necrotic cells were characterized by cytoplasmic vacuolization due to membrane disruption and loss of cell components during the necrotic process (Fig. 3Q). The few adherent apoptotic cells exhibited large orange areas (Fig. 3P, inset) as observed during late apoptosis, whereas suspended cells showed bright-green nuclei predictive of apoptotic bodies (Fig. 3P). RH30 cells treated with 2 mM melatonin were characterized by round, blebbed cells after 24 and 48 h (Fig. 3Q and R, respectively). TEM analysis at 48 h of treatment highlighted dark areas predictive of DNA condensation and cytoplasmic vacuolization (Fig. 3S). Using AO and PI double-staining, melatonin-treated cells showed orange staining at the nuclei due to late apoptosis and necrosis already after 24 h (Fig. 3T), as also observed after 48 h (data not shown).

Forced differentiation of tumor cells induced by anticancer agents has been widely exploited to limit the growth of tumor masses. In this regard, melatonin has been hypothesized to promote a differentiated phenotype in some tumors, such as gastric (49) and prostate cancer (50). Thus, to understand whether melatonin influences the myogenic differentiation of RMS, we employed human RD cells, which commonly exhibit a consistent myogenic potential in comparison to alveolar RH30 cells. For RD cells, the DM in the absence or presence of melatonin was replaced daily using different concentrations of melatonin. The extent of myogenic differentiation reached by cells in the different conditions was measured by immunoblotting analysis of markers that are normally increased during the differentiation of myoblasts, including caveolin-3 (Cav-3) and myosin heavy chain (MHC). Melatonin treatment led to a dose-dependent impairment of myogenic differentiation, since Cav-3 and MHC levels were reduced in comparison to the controls. In particular, melatonin at a concentration of 1 mM completely abolished the myogenic differentiation in RMS cells, as both Cav-3 and MHC levels were undetectable (Fig. 4A). We also administered melatonin to RD cells after which they were differentiated for 4 days and then analyzed for caspase-3 proteolytic activation, which is commonly utilized as a readout of the cell apoptotic program. As observed by immunoblotting analysis, melatonin-treated differentiated RD cells were characterized by increased levels of caspase-3 cleaved fragments (~19 and 17 kDa) in comparison to vehicle-treated cells after treatment for 24 and 48 h (Fig. 4B). These experiments demonstrated that melatonin has no positive effects on RMS differentiation, but behaves as a cytotoxic drug by triggering a caspase-dependent apoptosis.

Previous findings have suggested that combination therapies including melatonin and conventional cancer drugs enhance success by increasing drug efficacy while reducing their side effects. In most clinical trials where melatonin was used in conjunction with chemotherapeutic drugs, improved overall survival and patient conditions were observed (51,52). This suggests that melatonin enhances the efficacy of chemotherapy and reduces side effects (53,54). A combination of melatonin and doxorubicin was reported to enhance the growth inhibitory effect and induction of apoptosis in human hepatoma cells in comparison to melatonin or doxorubicin used alone (55). To verify the combined effects of melatonin and chemotherapy drugs on RMS chemoresistance, RD and RH30 cells were concurrently treated with 1 mM melatonin and 0.15 ng/ml doxorubicin or 2 µg/ml cisplatin for up to 48 h. As shown in Fig. 5, the two chemotherapies were effective in reducing the proliferation of RMS cell lines in comparison to the controls, as observed after 48 h of treatment. In addition, the concomitant treatment of melatonin with doxorubicin for up to 48 h produced a synergistic inhibitory effect (Fig. 5A). Similar results were obtained following treatment with melatonin and cisplatin (Fig. 5B). These results demonstrate a possible role of melatonin in improving the chemotherapy efficacy.