Nonpulsed Sinusoidal Electromagnetic Fields as a Noninvasive Strategy in Bone Repair: The Effect on Human Mesenchymal Stem Cell Osteogenic Differentiation

Abstract

In vivo control of osteoblast differentiation is an important process needed to maintain the continuous supply of mature osteoblast cells for growth, repair, and remodeling of bones. The regulation of this process has also an important and significant impact on the clinical strategies and future applications of cell therapy. In this article, we studied the effect of nonpulsed sinusoidal electromagnetic field radiation tuned at calcium-ion cyclotron frequency of 50 Hz exposure treatment for bone differentiation of human mesenchymal stem cells (hMSCs) alone or in synergy with dexamethasone, their canonical chemical differentiation agent. Five days of continuous exposure to calcium-ion cyclotron resonance affect hMSC proliferation, morphology, and cytoskeletal actin reorganization. By quantitative real-time polymerase chain reaction, we also observed an increase of osteoblast differentiation marker expression such as Runx2, alkaline phosphatase (ALP), osteocalcin (OC), and osteopontin (OPN) together with the osteoprotegerin mRNA modulation. Moreover, in these cells, the increase of the protein expression of OPN and ALP was also demonstrated. These results demonstrate bone commitment of hMSCs through a noninvasive and biocompatible differentiating physical agent treatment and highlight possible applications in new regenerative medicine protocols.

Introduction

The continuous remodeling of bone occurs through a dynamic process mediated by two key regulators, hematopoietic-derived osteoclasts and mesenchymal-derived osteoblasts.^1–3 Indeed, bone mass in adults is maintained by a local balance between osteoclastic bone resorption and osteoblastic activity mediated through various signaling molecules, such as morphogens, hormones, growth factors, cytokines, matrix proteins, and transcription factors.^4–8

Committed preosteoblasts derived from mesenchymal stem cells (MSCs) reside near the bone surface and when differentiated into mature osteoblasts, they undergo a phenotypic change, showing a large nucleus, an enlarged Golgi, and an extensive endoplasmic reticulum that support the secretion of bone matrix proteins such as type I collagen.⁹ Subsequently, the cells become terminally differentiated into osteocytes that provide mechanical support and regulate mineral deposition.

The osteoblast differentiation can be induced by glucocorticoid treatments and, in particular, dexamethasone was often used as their chemical differentiating agent. Glucocorticoids are valuable drugs for the treatment of bone diseases, but may also exert serious side effects¹⁰ with prolonged use and frequently result in bone loss, high fracture risk, and osteoporosis.^11,12

Osteogenesis is a multistep series of events modulated by an integrated cascade of gene expression characterized by three orderly stages: (1) proliferation; (2) extracellular matrix deposition and maturation; and (3) mineralization of the bone extracellular matrix.¹³ These phases are accompanied by the activation of transcription factors, among which Runx2 and specific genes associated with osteoblast phenotypes, such as alkaline phosphatase (ALP), osteocalcin (OC), and osteopontin (OPN).

ALP is considered an early differentiation marker expressed at high levels at the end of the proliferative period and during the extracellular matrix deposition and maturation, while OPN the principal bone phosphorylated glycoprotein and OC a small highly conserved molecule are expressed during the last phase of osteogenesis, associated with matrix mineralization of the bone.¹⁴

It was reported that pulsed electromagnetic field (PEMF) stimulation is able to induce the acceleration of bone formation, the proliferation of osteoblasts,^15–18 and the downregulation of osteoclast bone resorption ability.^19,20

In vivo exposure to these fields can also improve bone mineral density (BMD) and slow down the bone resorption process.^21–24 In recent years, PEMFs have been widely used in osteoporosis treatments too,^25,26 as demonstrated in the randomized controlled trials where most patients showed an improvement of BMD, vitamin D status, muscle strength, and in walking speed after such treatment.^27,28

PEMFs, apart from enhancing osteogenesis, also promote bone healing in patients with bone nonunions, improve postsurgical healing in spinal fusion conditions, and alleviate spine pain in patients who have multiple vertebral deformities and severe chronic back pain.^29–33

Considering these findings as a starting point and based on our experience with nonpulsed sinusoidal EMFs,^34–39 in this study, we evaluated if also nonionizing radiation treatment is able to affect osteogenesis and, in particular, commit human MSCs (hMSCs) to trigger bone-specific differentiation markers and progress them toward a bone phenotype.

This study has the scope of improving the knowledge on the biological effects of EMF exposure, opening up to regenerative medicine applications to be used as a tool for innovative therapeutic approaches.

Materials and Methods

Ethics statement and cell culture

According to the guidelines of the Ethics Committee of the FBF S. Peter Hospital, human umbilical cord blood samples were harvested from term deliveries at the time of birth with the mothers' written informed consent and processed immediately.

hMSCs were isolated by centrifugation using the Ficoll-Paque density gradient separation medium (Amersham). Briefly, about 50 mL of umbilical cord blood was diluted with three volume of phosphate-buffered saline (PBS) and layered upon a Ficoll-Paque gradient, in a ratio of one part of Ficoll-Paque and three parts of blood. After centrifugation, the interface layer of mononucleated cells was carefully removed, washed three times, and seeded on plastic Petri dishes in a culture medium containing Dulbecco's modified Eagle's medium/F12, 10% fetal bovine serum, penicillin (100 U/mL), and streptomycin (100 μg/mL).

After 1 week, nonadherent cells were gently rinsed off by changing the medium and when the attached hMSCs reached 80–90% confluence, they were recovered using 0.05% Trypsin-EDTA and plated on Petri dishes at a density of 10×10³ cells/cm²; the medium was subsequently changed every 3 days. Cells were characterized by flow cytometry revealing the presence of mesenchymal markers (CD73, CD105, CD90, CD44; ImmunoTools) and the absence of hematopoietic markers (CD34, CD45; ImmunoTools) on the hMSC membrane.

In our protocol, isolated hMSCs were cultured and studied for three, four, and five generations here named 3-, 4-, and 5-week-old cells. Subsequently, to promote osteogenic differentiation, 3- and 5-week old cells were grown for 5 days in the following four different conditions: control untreated cells, exposed to non-PEMF cells (50 Hz, 1 mT), dexamethasone-treated cells (100 nM), and exposed to non-PEMF cells in synergy with dexamethasone treatment.

In addition, in this article, we studied the irreversible (IRV) effect induced by non-PEMFs on the exposed hMSCs. The same four above-treated samples were further grown for another 3 days in the absence of EMF exposure to study the IRV effect.

Exposure system: description and characterization

The equipment for EMF production (solenoid) is set up in the mu-metal shielded room located in our laboratories and described in our previous articles.^36,40 Briefly, the apparatus includes a cellular incubator made of polymethylmethacrylate, a diamagnetic material that has a relative permeability less than 1, where temperature (37°C±0.1°C), atmosphere composition (5% CO₂), and humidity regulation are provided, continuously controlled, and recorded by a laboratory view program (control system). The main body of this home produced solenoid is a 5-mm-thick polyvinyl chloride cylinder, with a 33 cm diameter and 3.3 m length. It is made of 3300 turns of 1 mm diameter copper wire. It is driven by three amplifiers and a signal generator that creates static and alternated current for EMF production. A magnetic field with frequencies of 0.01 Hz to 1 kHz is produced by the equipment and a magnetic flux density the magnitude of which spans between 10 nT and 1 mT; the maximum output voltage of the generator feeding the solenoid is 33 mV root mean square (RMS). The unexposed cell experiments were run in a normal cell incubator ThermoForma 3111 (Thermo Scientific).

Ca²⁺-ion cyclotron resonance exposure parameters

The exposure parameters were calculated based on the following Lorentz's equation: \documentclass{aastex}\usepackage{amsbsy}\usepackage{amsfonts}\usepackage{amssymb}\usepackage{bm}\usepackage{mathrsfs}\usepackage{pifont}\usepackage{stmaryrd}\usepackage{textcomp}\usepackage{portland, xspace}\usepackage{amsmath, amsxtra}\pagestyle{empty}\DeclareMathSizes{10}{9}{7}{6}\begin{document} \begin{align*} f = \frac {q \mid {\bf B} _ {{\bf DC}} \mid} {2 \pi m} \end{align*} \end{document}

where q and m are, respectively, the ion's charge and mass, |B_DC| is the flux density of the applied static MF, and f is the frequency of the superimposed EMF.³⁴ We applied a low EMF intensity (in the range of mT) turned at resonance condition where there is maximum energy transfer, which enables us to see a biological effect. Under these conditions, the amount of heating due to the Joule effect was negligible, and all the effects on the cells reported are related to the calcium-ion cyclotron resonance (Ca²⁺-ICR) exposure. In our study, 3- and 5-week-old hMSCs were continuously exposed, up to 5 days, to a static MF (66 μT) and an alternating sinusoidal EMF (1 mT RMS of intensity) at 50 Hz, matching the cyclotron frequency corresponding to the charge/mass ratio of calcium ion (ICR). The exposure parameters were chosen according to the results of our previous experiments, which show a differentiation effect on human cardiac stem cells, mouse skeletal muscle cells, and NT2 embryonal carcinoma cells.^35,36,40 All experiments have been performed under single-blind conditions with numbered samples.

Cell proliferation assay

The 3-, 4-, and 5-week-old cells were cultured in the four different conditions reported above: control untreated cells, exposed cells, dexamethasone-treated cells, and exposed cells in synergy with dexamethasone treatment. To study cell proliferation, an immunohistochemical method, the anti 5-bromo-2-deoxyuridine (BrdU) antibody (Cell Proliferation Kit; Roche Diagnostic GmbH), was used.

Cells (3×10³ cells/cm²) were seeded in a 96-well plastic plate and cultured in the four above-mentioned conditions; a cell proliferation assay was performed after 72 h. BrdU (10 μM) was added to cells during the last 18 h of culture, after the cells were fixed for 30 min and a solution containing the anti-BrdU antibody was added to the cells and left for 30 min at 37°C. One hundred microliters of 2, 2′-Azino-di-3-ethylbenzthiazoline sulfonate⁴¹ (ATBS) was then added and incubated for 30 min. The cell growth rate was determined by colorimetric assay measuring the absorbencies of samples in an ELISA reader at 405 nm.

Scanning electron microscopy (SEM)

The 3-week-old hMSCs were cultured for 5 days in the four different conditions reported above, after the cells were washed in PBS and fixed with 2.5% glutaraldehyde in 0.1 M Millonig's PBS for 1 h at 4°C. After three washes in the same buffer, the samples were postfixed in 1% OsO₄, dehydrated through a graded acetone series, and critical-point dried with CO₂ in a Balzers CPD 030 critical-point drier. Specimens were coated with gold in a Balzers SCD 050 sputter instrument and observed on a Cambridge S240 scanning electron microscope.

Confocal microscopy

Three-week-old control, exposed, dexamethasone, and dexamethasone-exposed hMSCs were cultured on 0.01% poly-lysine-treated glass coverslips for 5 days. Cells were then fixed in 4% paraformaldehyde for 10 min, rinsed twice with PBS, permeabilized with PBS containing 1% bovine serum albumin (PBS/BSA) and 0.2% Triton X-100 for 5 min, and rinsed again in PBS. Cells were then incubated with FITC-phalloidin (10 mg/mL) for 1 h at room temperature in the dark and finally rinsed three times in PBS/BSA, followed by two washes with PBS. Coverslips were assembled, cell-side down, on a microscope slide with 0.625% N-propylgalate in PBS glycerol 1:1. The coverslip sandwich was sealed to prevent exposure to air and to exclude and prevent the crystal formation of H₂O. Actin was labeled with FITC-phalloidin (Sigma Chemical) using the procedure of Bellomo.⁴² The fluorescence was then analyzed using a LEICA TCS 4D Confocal Microscope supplemented with an Argon Krypton laser and equipped with 40×1.00–0.5 and 100×1.3–0.6 oil immersion lenses.

Immunofluorescence analysis

Three-week-old control, exposed, dexamethasone, and dexamethasone-exposed hMSCs were cultured on 0.01% poly-lysine-treated glass coverslips for 5 days.

The cells were fixed in 4% paraformaldehyde at 4°C for 10 min, washed twice in Ca²⁺/Mg²⁺-free PBS, and permeated at room temperature for 15 min with 0.1% Triton X-100, 1% BSA (Sigma-Aldrich). hMSCs were incubated with the primary antibody OPN (1:200) (Chemicon) in a blocking buffer for 1 h. After washing in PBS, containing 0.1% Triton X-100 and 1% BSA, the cells were incubated with the anti-rat secondary antibody (1:100; Chemicon). Cells were washed three times with PBS, stained for nuclei localization with Hoechst 33342 (trihydrochloride-trihydrate), and examined. The samples were tested by indirect immunofluorescence analysis for the presence of the OPN protein. Fluorescence measurements were obtained using an inverted microscope (Olympus IX51; RT Slider SPOT-Diagnostic Instruments) equipped with a 40×objective and with a cooled CCD camera (Spot RT Slider, acquisition rate five frames per second, full frame; Diagnostic Instruments). No significant fluorescent signal was detectable with the secondary antibody alone. Densitometric analysis of the OPN immunofluorescence was performed with ImageJ Software.

Real-time quantitative reverse transcriptase–polymerase chain reaction analysis

The total RNA was extracted from 3- and 5-week-old hMSCs cultured in the four above-mentioned conditions for 5 days using the TRIzol Reagent (Life Technologies) according to the manufacturer's instructions. One microgram of total RNA was used to synthesize first-strand cDNA with random primers using 100 U of ImProm-II™ RT-PCR kit (Promega) according to the manufacturer. The reaction was also carried out in the absence of reverse transcriptase to check for genomic DNA amplification.

The quantification of all gene transcripts was carried out by real-time quantitative reverse transcriptase–polymerase chain reaction (RT-PCR). Experiments were conducted to contrast relative levels of each transcript and endogenous control GAPDH in every sample. The data were analyzed using the equation described by Livak,⁴³ as follows:

Amount of target-2^−ΔΔCt. \documentclass{aastex}\usepackage{amsbsy}\usepackage{amsfonts}\usepackage{amssymb}\usepackage{bm}\usepackage{mathrsfs}\usepackage{pifont}\usepackage{stmaryrd}\usepackage{textcomp}\usepackage{portland, xspace}\usepackage{amsmath, amsxtra}\pagestyle{empty}\DeclareMathSizes{10}{9}{7}{6}\begin{document} \begin{align*}\Delta {\rm Ct} & = ({\rm average \ target \ Ct} - {\rm average} \ GAPDH \ {\rm Ct}) \\ \Delta \Delta {\rm Ct} & = ({\rm average} \ \Delta {\rm Ct \ treated \ sample} \\& \quad - {\rm average} \ \Delta {\rm Ct \ untreated \ sample}) \end{align*} \end{document}

Before using the ΔΔCt method for quantification, we performed a validation experiment to demonstrate that the efficiency of target genes and reference GAPDH was equal. Real-time PCR was conducted using SYBR Green I Mastermix (Applied Biosystems) using an ABI PRISM™ 7000 Sequence Detection System. Each reaction was run in triplicate and contained 0.5 μL of cDNA template along with 250 nM primers in a final reaction volume of 25 μL. The specific primers used are shown in Table 1. Cycling parameters were 50°C for 2 min, 95°C for 10 min to activate DNA polymerase, and then 40 cycles of 95°C for 15 s and 60°C for 1 min. Melting curves were performed using Dissociation Curves software (Applied Biosystems) to ensure that only a single product was amplified. As negative controls, tubes in which RNA or RT was omitted during the RT reaction were always prepared and run.

Table 1.

Primers Used for Real-Time Reverse Transcriptase–Polymerase Chain Reaction

Target	Primer sequence	Annealing temperature (°C)
ALP	5′-caatgagggcaccgtggg-3′	60
	5′-tcgtggtggtcacaatgcc-3′
OC	5′-gcagcgaggtagtgaagag-3′	60
	5′-gaaagccgatgtggtcagc-3′
OPN	5′-gtgtggtttatggactgagg-3′	60
	5′-acggggatggccttgtatg-3′
OPG	5′-agtgtagagaggataaaacgg-3′	60
	5′-gaaggtgaggttagcatgtc-3′
Runx2	5′-catcatctctgccccctct-3′	60
	5′-actcttgcctcgtccactc-3′
GAPDH	5′-catcatctctgccccctct-3′	60
	5′-caaagttgtcatggatgacct-3′

ALP, alkaline phosphatase; OC, osteocalcin; OPG, osteoprotegerin; OPN, osteopontin.

ALP histochemistry analyses

Three-week-old control, exposed, dexamethasone, and dexamethasone-exposed hMSCs were cultured for 5 days. After the treatments, cells were fixed in 2% buffered formalin for 15 min at 4°C, washed three time with PBS, and incubated for 30 min in Tris buffer (0.2 M, pH 8.3) with Naphthol AS-MX phosphate (Sigma) as a substrate and Fast Blue (Sigma) for staining.^44,45 The ALP-positive cells, stained in blue/purple, were examined with inverted optical microscopy for determining the number of positive cells (Olympus IX51; RT Slider SPOT Diagnostic Instruments) equipped with a 20× objective and with a cooled CCD camera (Spot RT Slider; Diagnostic Instruments). For each experiment, a minimum of five ranges of vision were analyzed and the cells counted. The experiments were repeated thrice.

Statistics analysis

The statistical analysis of the data was performed by using a Student's t-test, with p<0.05 as the minimum level of significance.

Results

Effect of Ca²⁺-ICR frequency on hMSC growth

The effect of 50 Hz, 1 mT nonpulsed sinusoidal EMF exposure on hMSC growth was investigated by the BrdU incorporation assay. The 3-, 4-, and 5-week-old hMSCs were studied, and the findings of cell growth at the third day of the week considered for all different treatments (control, exposed, dexamethasone, and dexamethasone-exposed cells) were reported. A statistically significant difference of BrdU cellular uptake was observed between control and exposed cells and between dexamethasone-exposed cells and dexamethasone alone (Fig. 1).

FIG. 1.

Human mesenchymal stem cell (hMSC) proliferation was analyzed by 5-bromo-2-deoxyuridine (BrdU) incorporation assay and measured in an ELISA reader at 405 nm. We performed a cell proliferation analysis in the exposed cells (ex) compared with the control ones (ctr) and in the dexamethasone-treated cells in synergy with exposure (ex+dx) compared with the dexamethasone (dx) alone in 3-, 4-, and 5-week-old cells. Statistical evaluation of the data was assessed by using a Student's t-test with p<0.05 as the minimum level of significance. Data are shown as mean SD. Asterisk identifies statistical significance (p<0.05).

In particular, the exposed cells showed a decrease of cell proliferation at week 3 and 4 and a proliferation increase at week 5 compared to the control ones. A statistically significant decrease in cell proliferation in the sample treated with dexamethasone in synergy with the exposure was also observed compared to the sample treated with dexamethasone alone (Fig. 1).

Effect of Ca²⁺-ICR frequency on hMSC shape and actin organization

The 3-week-old hMSCs exposed for 5 days showed a drastic change in cellular morphology similar to the dexamethasone-treated cells observed through scanning electron microscopy (SEM) analysis (Fig. 2A–C). The exposed cells appeared larger with a polygonal-like shape when compared to the enlonged spindle-shaped control ones (Fig. 2A, C). This effect increased further in the exposed mesenchymal cell sample when treated contemporarily with the dexamethasone treatment (Fig. 2D).

FIG. 2.

Effect of Ca²⁺-ion cyclotron resonance (ICR) exposure on hMSC morphology by scanning electron microscopy analysis. Control cells (A), dexamethasone-treated cells (B), exposed cells (C), and dexamethasone-treated cells in synergy with exposure (D).

The morphology changes were confirmed through the cytoskeletal actin filament rearrangement (Fig. 3). Confocal microscopy analysis of 3-week-old hMSCs exposed for 5 days highlighted an actin reorganization with a widespread network in the cellular body similar to the dexamethasone-treated cells (Fig. 3B, C), whereas the control cells show a periphery distribution of the actin filaments around the cell membrane (Fig. 3A). Moreover, when the exposed cells were treated together with dexamethasone, we observed a further increase in this effect (Fig. 3D).

FIG. 3.

Effect of Ca²⁺-ICR exposure on the actin distribution in hMSCs by confocal microscopy analysis. Phalloidin fluorescent staining (red) of the control cells (A), the dexamethasone-treated cells (B), the exposed cells (C), and the dexamethasone-treated cells in synergy with exposure (D). Color images available online at www.liebertpub.com/tec

Effect of Ca²⁺-ICR frequency on hMSC osteoblast differentiation marker expression

The expression of the Runx2 transcription factor and the osteoblast differentiation markers, such as ALP, OC, and OPN, was examined in 3- and 5-week-old hMSCs exposed for 3 and 5 days (Fig. 4).

FIG. 4.

Effect of Ca²⁺-ICR exposure on the mRNA expression of osteoblast differentiation markers in hMSCs. Real-time polymerase chain reaction (PCR) analysis of Runx2, alkaline phosphatase (ALP), osteopontin (OPN), osteocalcin (OC) mRNAs in 3- and 5-week-old cells exposed for 3 and 5 days (left side A, B, C, D). Real-time PCR analysis of Runx2, ALP, OPN, OC, mRNAs in 3- and 5-week-old cells exposed for 3 and 5 days and then grown for another 3 days in the absence of nonpulsed sinusoidal electromagnetic field (EMF) (irreversible [IRV] effect) (right side A, B, C, D). Statistical evaluation of the data was assessed by using a Student's t-test with p<0.05 as the minimum level of significance. Data are shown as mean SD. Asterisk identifies statistical significance (p<0.05).

The 3-week-old cells exposed for 3 and 5 days showed a statistically significant increase of the early markers at 3 days and of the late markers at 5 days, including also the transcription factor Runx2 (which is not yet downregulated), compared with control cells (Fig. 4A). The 5-week-old cells exposed for 3 and 5 days showed a statistically significant increase of the early markers at 3 days and of the late markers at 5 days, instead the transcription factor Runx2 starts to downregulate at 5 days of exposure compared to control cells (Fig. 4C).

In the 3-week-old cells treated in synergy with dexamethasone, there is a statistically significant increase of the early markers at 3 days and of the late markers at 5 days, including the transcription factor Runx2, compared to the dexamethasone alone-treated sample (Fig. 4B).

In the 5-week-old cells treated in synergy with dexamethasone, we report a statistically significant increase of the early markers at 3 days and of the late markers at 5 days except for the transcription factor Runx2, which starts its statistical significant downregulation at 5 days of exposure, compared to the dexamethasone alone-treated sample (Fig. 4D).

The upregulation of osteoblast differentiation markers, with a downregulation of Runx2, was also reported in the absence of the EMF for the next 3 days observed (IRV effect; Fig. 4A–D right side), highlighting that the differentiating effect induced by the nonpulsed sinusoidal EMF, at this time and in these cells is an IRV process.

Effect of Ca²⁺-ICR frequency on hMSC ALP protein expression

The increase of ALP expression was also confirmed at the protein level by histochemistry analysis (Fig. 5). The 3-week-old hMSCs exposed for 5 days (Fig. 5C) showed a statistically significant increase of the ALP protein activity compared with the control ones (Fig. 5A). This effect plus an increase of the cellular body size was also observed in the dexamethasone alone-treated cells (Fig. 5B) and enhanced in the exposed cells treated together with dexamethasone (Fig. 5D). In particular, in this last sample, the ALP-positive cells appear larger with a polygonal-like shape, confirming the SEM analysis results (Fig. 2B–D). The percentage of ALP-positive cells was calculated, as reported in Figure 5E.

FIG. 5.

Effect of Ca²⁺-ICR exposure on ALP protein activity in hMSCs by histochemistry analysis of control cells (A), dexamethasone-treated cells (B), exposed cells (C), and dexamethasone-treated cells in synergy with exposure (D; ×20 objective). (E) The percentage of the ALP-positive cells. Data are shown as mean SD. Asterisk identifies statistical significance (p<0.05) compared to control cells. Color images available online at www.liebertpub.com/tec

Effect of Ca²⁺-ICR frequency on hMSC OPN protein expression

The 3-week-old hMSCs exposed for 5 days (Fig. 6C) showed a statistically significant increase of the OPN protein expression compared with the control ones by immunofluorescence analysis (Fig. 6A). This effect was potentiated in the exposed sample treated together with dexamethasone (Fig. 6D) and in the dexamethasone alone one (Fig. 6B). These results were also quantified by densitogram analysis (Fig. 6E). In particular, in the control sample, the OPN protein fluorescence was not revealed, whereas in the exposed hMSCs, it was highlighted instead and appeared diffusely distributed in the cellular body with more intensity in the perinuclear region. This effect increased in the exposed sample treated together with dexamethasone where the OPN expression appeared concentrated around the nucleus and in the cytoplasm zone similarly to the dexamethasone alone-treated sample.

FIG. 6.

Effect of Ca²⁺-ICR exposure on OPN protein expression in hMSCs by indirect immunofluorescence analysis. OPN fluorescent staining (green) of control cells (A), dexamethasone-treated cells (B), exposed cells (C), and dexamethasone-treated cells in synergy with exposure (D). Nuclei are counterstained with Hoechst (blue; ×40 objective). (E) Densitogram analysis. Quantification of OPN fluorescence intensity by ImageJ Software. Data are shown as mean SD. Asterisk identifies statistical significance (p<0.05) compared to the control cells. Color images available online at www.liebertpub.com/tec

Effect of Ca²⁺-ICR frequency on hMSC osteoprotegerin expression

The osteoclastogenesis inhibitory factor (osteoprotegerin [OPG]) mRNA expression was studied by real-time PCR analysis. In our experiments, the 3- and 5-week-old cell samples treated for 5 days with dexamethasone alone or in synergy with nonpulsed sinusoidal EMF exposure both showed a completely nullified OPG mRNA expression (Fig. 7A, B). More importantly, compared to the treatments above, the exposed 3- and 5-week-old cells, instead showed an upregulation of the OPG mRNA expression (Fig. 7A, B), which was also maintained when the cells were grown further in the absence of the nonpulsed sinusoidal EMF exposure (IRV samples; Fig. 7A, B).

FIG. 7.

Effect of Ca²⁺-ICR exposure on the osteoprotegerin (OPG) mRNA expression in hMSCs. Real-time PCR analysis of the OPG mRNA expression in 3- and 5-week-old cells exposed for 5 days (left side A, B). Real-time PCR analysis of OPG, mRNA expression in 3- and 5-week-old cells exposed for 5 days and then grown for another 3 days in the absence of nonpulsed EMF (IRV effect) (right side A, B). Statistical evaluation of the data was assessed by using a Student's t-test with p<0.05 as the minimum level of significance. Data are shown as mean SD. Asterisk identified statistical significance (p<0.05).

Discussion

After the Food and Drug Administration approved PEMFs as a safe and efficient treatment for osteoporosis, nonunion fractures, and congenital pseudoarthrosis,^46–51 a rising interest toward this treatment enhanced the applications of different ranges and types of electric and EMFs (pulsed and nonpulsed) for the therapeutic treatment of bone disorders.

PEMFs have also been used for nonspinal fusion postsurgical healing treatments to improve bone healing in patients with fractures and also to prevent osteoporosis.^{29,32,33,51,52}

In the case of nonpulsed sinusoidal EMFs, they affect biological systems and processes, including cell differentiation, as reported and widely studied by international research groups.^{35,36,53–56} Presently, it is difficult to establish the possible biological mechanisms involved in the interaction effect of these EMFs, which still result under debate.

One of the reasons for this discussion is the lack of reproducibility of results due to the difficulty of having a precise EMF characterization of the fields used worldwide.

Several mechanisms have been developed to explain the biological effect of ICR exposure⁵⁷ and we report them in this study for a wider view. Some of these theories are more appealing to biochemists⁵⁸ while others to biophysics.^34,59,60

Liboff, in 1985, considered the transport of ions in membrane-bound channels under the action of a specific cyclotron frequency, suggesting a mechanism similar to the helical motion of charged particles in free space under the influence of the Lorentz force.³⁴ Instead, the Lednev theory⁵⁸ reports that during the exposure to EMFs, the reorientation of the membrane phospholipids could deform the embedded ion channels altering their dynamics. In this theory, an ion in its protein-binding site is considered as a dipole when exposed to its ICR frequency, the energy from the exposure is then transferred to the dipole, and as a result, the ion is released in the solution. The most recent approach to the ICR mechanism question has been advanced by Del Giudice et al.⁶⁰ who has framed the problem in terms of quantum electrodynamics.

To develop further knowledge on the use of non-PEMFs for therapeutic treatments, in our article, we studied the osteoblast differentiation effects on hMSCs induced by 50 Hz, 1 mT nonpulsed sinusoidal EMF exposure. In this study, we studied and reported that this treatment is able to induce osteoblast cell morphology changes and cytoskeletal reorganization, a decrease in cell proliferation, and an increase in cell differentiation, together with cell osteoclastogenesis inhibition.

Cytoskeletal remodeling is crucial for myoblast cell differentiation, as a matter of fact it generates a mechanical tension in the plasma membrane, activating stretch-activated channels and triggering Ca²⁺-dependent signals.⁶¹ This process was also reported during hMSC osteoblast commitment,^62,63 induced by the dexamethasone treatment.⁶⁴

In our experiments, nonpulsed sinusoidal EMF exposure is able to modify hMSC morphology (Fig. 2), changing it from the undifferentiated fibroblast-like phenotype to a larger and polygonal-like shape phenotype, typical of the differentiated stage.

Furthermore, in the exposed cells, this physical agent can induce a cytoskeletal actin filament rearrangement, similar to the one found in the dexamethasone-treated cells (Fig. 3) reaching its peak effect in the cells exposed and treated in synergy with the chemical agent. A combination between the physical and the chemical agents' treatment here suggests a potentiated effect due to the synergy between each other.

We also observed a statistically significant early decrease of the proliferation rate in the exposed hMSCs (Fig. 1), followed by a subsequent increase in a later time when compared with the unexposed cells, which showed a physiological constant decrease of cell proliferation like in the cells treated with dexamethasone alone or in synergy with nonpulsed sinusoidal EMF.

Having reconfirmed the results, widely found in literature, on the decrease of cell proliferation during the dexamethasone differentiating treatment, we furthermore demonstrated a restart of the proliferation trend in exposed hMSCs at 5 weeks. These findings suggest that the differentiating physical treatment is less invasive than the chemical one, because in the long term, it does not lead to a decrease of the proliferation rate and hypothetically, the osteoprogenitor cell number, which is considered one of the causes of osteoporosis fragilization.¹¹

The osteoblast commitment of exposed MSCs was confirmed by Runx2, ALP, OPN, and OC mRNA expression level analysis.

The nonpulsed sinusoidal EMF exposure is able to induce a statistically significant increase of the Runx2 transcription factor and the early and late osteoblast differentiation markers in exposed cells, when compared with the control ones. This also happens in the exposed samples treated in synergy with dexamethasone compared to the dexamethasone treatment alone. The increase of the Runx2 expression at an early time is important for directing MSCs to the osteoblast lineage and this expression has to be completely downregulated when the cells reach a fully mature osteoblast phenotype.⁶⁵ Accordingly, in our cellular system, the Runx2 mRNA expression showed an upregulation in the exposed 3-week-old cells treated for 3 and 5 days (early time) followed by a downregulation in the exposed 5-week-old cells treated for 5 days (late time).

We also proved the differentiating capability of 50 Hz, 1 mT nonpulsed sinusoidal EMF exposure in hMSCs by the study of the OPG gene expression. This marker is strongly downregulated in the samples treated with dexamethasone alone or in synergy with nonpulsed sinusoidal EMF compared to the slight, but important upregulation resulting in the exposed cells.

Bone construction is the result of dynamic interaction between bone-forming osteoblasts and bone-resorbing osteoclasts.

The dexamethasone treatment acts, on one hand, promoting the differentiation of immature osteoprogenitor cells into osteoblasts⁶⁶ and, on the other, by the increase the osteoclastogenesis⁶⁷ mediated by a decrease of the OPG expression.⁶⁸ This balance between bone formation and resorption is initially well counterweighed, but the continuous treatment with dexamethasone leads to a depletion of the osteoprogenitor pool and a decrease in bone formation in the longer term bringing to a glucocorticoid-induced osteoporosis effect.⁶⁹

In our study, the osteoblast differentiation induced in hMSCs by non-PEMFs preserves the OPG gene expression and allows the equilibrium between the osteoclastogenesis and the osteogenesis processes thus preventing bone loss, which in the long term, instead, is a side effect of the dexamethasone treatment.

The constant mRNA expression of the osteoblast's differentiation markers in the exposed hMSC grown further for another 3 days in the absence of the nonpulsed sinusoidal EMF was observed and reached its maximum IRV effect in the exposed sample treated in synergy with dexamethasone (Fig. 4).

The results on the permanent differentiating effect of resonant EMF (turned to Ca²⁺-ICR) together with an upregulation of the ALP and OPN expressions at the protein level, demonstrate a stable and lasting differentiating effect of the physical agent treatment on the hMSCs.

The results of this article open up to the possibility of using a resonant EMF to induce a well-balanced osteoblast commitment in clinical therapy for osteogenetic disease treatments.

The use of this noninvasive strategy could improve the endogenous osteogenic response, representing a new opportunity to reduce healing times and therefore the recovery of patients.

Footnotes

Acknowledgments

This work was supported by the contract grant sponsor INAIL (contract B n.22/DIPIA). The authors thank Antonella Greco for copyediting/proofreading the English in this article.

Disclosure Statement

No competing financial interests exist.

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Nonpulsed Sinusoidal Electromagnetic Fields as a Noninvasive Strategy in Bone Repair: The Effect on Human Mesenchymal Stem Cell Osteogenic Differentiation

Abstract

Introduction

Materials and Methods

Ethics statement and cell culture

Exposure system: description and characterization

Ca2+-ion cyclotron resonance exposure parameters

Cell proliferation assay

Scanning electron microscopy (SEM)

Confocal microscopy

Immunofluorescence analysis

Real-time quantitative reverse transcriptase–polymerase chain reaction analysis

ALP histochemistry analyses

Statistics analysis

Results

Effect of Ca2+-ICR frequency on hMSC growth

Effect of Ca2+-ICR frequency on hMSC shape and actin organization

Effect of Ca2+-ICR frequency on hMSC osteoblast differentiation marker expression

Effect of Ca2+-ICR frequency on hMSC ALP protein expression

Effect of Ca2+-ICR frequency on hMSC OPN protein expression

Effect of Ca2+-ICR frequency on hMSC osteoprotegerin expression

Discussion

Footnotes

Acknowledgments

Disclosure Statement

References

Ca²⁺-ion cyclotron resonance exposure parameters

Effect of Ca²⁺-ICR frequency on hMSC growth

Effect of Ca²⁺-ICR frequency on hMSC shape and actin organization

Effect of Ca²⁺-ICR frequency on hMSC osteoblast differentiation marker expression

Effect of Ca²⁺-ICR frequency on hMSC ALP protein expression

Effect of Ca²⁺-ICR frequency on hMSC OPN protein expression

Effect of Ca²⁺-ICR frequency on hMSC osteoprotegerin expression