TY - JOUR
AU - He,, Rongxin
AB - Background Electromagnetic fields (EMFs) used in stem-cell tissue engineering can help elucidate their biological principles. Objective The aim of this study was to investigate the effects of low-intensity EMFs on cell proliferation, differentiation, and cycle in mouse bone marrow stromal cells (BMSCs) and the in vivo effects of EMFs on BMSC. Methods Harvested BMSCs were cultured for 3 generations and divided into 4 groups. The methylthiotetrazole (MTT) assay was used to evaluate cell proliferation, and alkaline phosphatase activity was measured via a colorimetric assay on the 3rd, 7th, and 10th days. Changes in cell cycle also were analyzed on the 7th day, and bone nodule formation was analyzed on the 12th day. Additionally, the expression of the collagen I gene was examined by reverse transcription-polymerase chain reaction (RT-PCR) on the 10th day. The BMSCs of the irradiated group and the control group were transplanted into cortical bone of different mice femurs separately, with poly(lactic-co-glycolic acid) (PLGA) serving as a scaffold. After 4 and 8 weeks, bone the bone specimens of mice were sliced and stained by hematoxylin and eosin separately. Results The results showed that EMFs (0.5 mT, 50 Hz) accelerated cellular proliferation, enhanced cellular differentiation, and increased the percentage of cells in the G2/M+S (postsynthetic gap 2 period/mitotic phase + S phase) of the stimulation. The EMF-exposed groups had significantly higher collagen I messenger RNA levels than the control group. The EMF + osteogenic medium–treated group readily formed bone nodules. Hematoxylin and eosin staining showed a clear flaking of bone tissue in the irradiated group. Conclusion Irradiation of BMSCs with low-intensity EMFs (0.5 mT, 50 Hz) increased cell proliferation and induced cell differentiation. The results of this study did not establish a stricter animal model for studying osteogenesis, and only short-term results were investigated. Further study of the mechanism of EMF is needed. Physical therapy often is advocated to treat a variety of bone diseases, as it optimizes the quality of life for people limited by functional loss caused by disease or trauma. Electromagnetic fields (EMFs) are natural physical fields. Their discovery has led to public interest as well as fear regarding their potential harmful effects. In particular, the use of EMFs in physical therapy has proven effective in animal experiments and clinical observations. They are noninvasive and compliant, have direct dose-response effects, and are widely used in clinical physical therapy as a progress marker. Further exploration of EMFs is needed to promote their use in physical therapy in various clinical departments. Electromagnetic fields have been used widely in the treatment of orthopedic conditions and diseases, including fractures, bone nonunion, osteoporosis, osteoarthritis, congenital bone defects, fixed joint failure, transplantation, and bone osteonecrosis. Eyres et al1 examined the effects of pulsed electromagnetic fields (PEMFs) on bone formation and disuse osteoporosis sustained during limb lengthening in a double-blind study. Stimulation by PEMFs prevented bone loss adjacent to the distraction gap. In a double-blind study of the effects of PEMFs in 30 people undergoing hip revision, Dallari et al2 found that PEMF treatment aided clinical recovery and bone-stock restoration. Furthermore, clinical studies have shown that the treatment was effective, although the investigators failed to examine the mechanism of the EMFs.2,3 The mechanism underlying promotion of bone proliferation and differentiation by EMF and the experimental parameters remain unknown. Data show that EMF stimulation can increase chondrocyte proliferation in vitro,4 augment proteoglycan synthesis in full-thickness cartilage explants,5 prevent the catabolic effects of IL-1 on the extracellular matrix,6 and upregulate transforming growth factor β (TGFβ) superfamily gene regulation in vivo.7 However, there is little consensus on the optimal EMF treatment conditions. The effects of the width, intensity, and frequency of magnetic field application have been assessed in different cell types at different cell densities for different therapeutic purposes. Furthermore, the dose at which EMF exposure is useful versus harmful has not been defined. The debate on whether EMFs can affect biological systems or cause long-term health effects has not been resolved.8 These clinical research and laboratory issues have not been clarified. However, regenerative medicine is gaining importance in medical treatments. Here, we postulate that, as a part of physical therapy, EMFs used in stem-cell-tissue engineering can help elucidate the biological principles of EMFs and increase the clinical value of tissue engineering. However, several issues must be clarified: (1) whether EMFs are toxic to bone marrow stromal cells (BMSCs); (2) what, if any, changes in the biological characteristics of BMSCs occur with extended irradiation; and (3) whether irradiated BMSCs can maintain the biological characteristics of the original cells after transplantation into animals. This study focused on resolving these issues. The International Commission on Non-ionizing Radiation Protection (ICNIRP) has determined that EMF exposure of ≤0.1 mT has no biological effects.9,10 We examined whether an EMF intensity of 0.5 mT was effective in inducing BMSC-osteoblast differentiation. Previous experiments have shown that 50-Hz EMFs promoted cell proliferation, with no toxic effects.11,12 Moreover, the exposure system for EMF is compatible with alternating currents, which is 50 Hz; therefore, we used a frequency of 50 Hz. As BMSCs can differentiate into osteoblasts and exhibit osteogenic activity, they are useful in bone tissue engineering. Therefore, we measured BMSC proliferation and differentiation by exposing them to EMF levels of 0.5 mT at 50 Hz. Materials and Method Reagents The BMSCs were maintained in DMEM-LG medium (Hyclone Laboratories Inc, Logan, Utah) with 10% fetal bovine serum (Gibco, Life Technologies Corporation, Grand Island, New York). Dexamethasone, vitamin C, β-sodium glycerophosphate, methylthiotetrazole (MTT), and the alkaline phosphatase (ALP) assay kit were purchased from Sigma-Aldrich Company (St. Louis, Missouri). Trypsin was purchased from Amresco (Solon, Ohio). The enzyme-labeled meter was a product of Bio-TEK (Winooski, Vermont). Poly(lactic-co-glycolic acid) (PLGA) was provided by the School of Polymer Science and Engineering, Zhejiang University. Animals The experiments were performed on 8 Sprague-Dawley mice with initial weights of approximately 320 g. However, male and female patients received EMF in clinical treatment. In order to avoid the in vivo result bias caused by using all male or all female mice, we chose 4 male and 4 female mice as the experimental object. Mice were obtained from the Center of Experimental Animals of Zhejiang University. The mice were aged 4 months at the beginning of the study and were fed with standard pelleted food. All of the 8 mice received the same surgery (the process is described in detail in the “Determination of the Effects of Callus Formation in Mice After In Vitro EMF Stimulation” section). The surgery was performed in a sterile environment. The animals were kept in a 14/10-hour light/dark environment at a constant temperature of 22±3°C and 45%±10% humidity. Two additional Sprague-Dawley mice (4 weeks of age) were used for separating and culturing BMSCs. All animal experiments were approved by the Zhejiang University Animal Ethics Committee. Instruments The sXcELF exposure system for electromagnetic radiation in complex environments is a product of the IT'IS Foundation (Zurich, Switzerland) and was supplied by the Bioelectromagnetics Laboratory of Zhejiang Province, Zhejiang University School of Medicine. Helmholtz coils generated the magnetic fields. The principle of the exposure system for electromagnetic radiation is described in Figure 1. Figure 1 Open in new tabDownload slide The electromagnetic field stimulation system. The magnetic fields were generated by Helmholtz coils. When the alternating current ran through the Helmholtz coils, uniform electromagnetic fields were generated in space. The cells were put in the middle area. There was a magnetic flux detector on the bottom, which was used to measure the intensity of the electromagnetic fields. The whole set was installed in a thermo incubator where the temperature was maintained at 37°C and carbon dioxide was maintained at 5% automatically. A cooling device was integrated in the incubator to maintain the temperature. The thermo incubator was connected to a computer, which detected the temperature and controlled the intensity of the electromagnetic fields. Figure 1 Open in new tabDownload slide The electromagnetic field stimulation system. The magnetic fields were generated by Helmholtz coils. When the alternating current ran through the Helmholtz coils, uniform electromagnetic fields were generated in space. The cells were put in the middle area. There was a magnetic flux detector on the bottom, which was used to measure the intensity of the electromagnetic fields. The whole set was installed in a thermo incubator where the temperature was maintained at 37°C and carbon dioxide was maintained at 5% automatically. A cooling device was integrated in the incubator to maintain the temperature. The thermo incubator was connected to a computer, which detected the temperature and controlled the intensity of the electromagnetic fields. Cell Culture The Sprague-Dawley mice (4 weeks of age) were killed by cervical vertebra dislocation. The mice then were immersed into a 75% alcohol solution for 30 minutes. The bilateral femurs and tibias were cut, and the soft tissues were removed. The medullary cavity of the bone was washed with DMEM-LG medium containing 10% fetal bovine serum. To isolate BMSCs, Percoll (Sigma-Aldrich Company) density separation of mouse MSCs was used. Next, 1.073 g/mL Percoll was added, and the suspension was centrifuged at 2,100 rpm for 30 minutes. The middle layer (containing mononuclear cells) was removed, washed twice in phosphate-buffered saline, and inoculated into DMEM-LG medium. After 1 day, the medium was changed to remove nonadherent cells. Isolated BMSCs were cultured under humidity saturation in an incubator at 37°C and 5% carbon dioxide. The culture medium was first changed on the third day after isolation. To maintain the culture, the medium subsequently was changed every 3 to 4 days. When BMSCs grew to 80% confluence, cells were lightly digested with trypsin and split at a ratio of 1:2. After splitting, the cells were called a new generation. Counted this way, BMSCs were allowed to propagate for 3 generations and were divided into 4 experimental groups when the cells were observed to adhere to the culture flask.12 Grouping and Designing Third-generation BMSCs were divided into 4 groups. Cells from groups A and C were cultured in medium with 10% fetal bovine serum. Group A served as the control group, and group C was exposed to EMFs of 0.5 mT at 50 Hz. Groups B and D were exposed to osteogenesis-inducing medium containing 10% fetal bovine serum, 10 mmol/L 7-sodium glycerophosphate, 10 mmol/L dexamethasone, and 50 mg/L vitamin C.13 The cells in group B were not exposed to EMFs. Cells in group D were exposed to EMFs of the same intensity as those in group C, and the experimental duration was 8 hours every day. At other times, the cells from the 4 groups were kept in the same cultured cube. The experiment period was 12 days. The proliferation and differentiation of BMSCs are apparent in the early stage of cell breeding. Thus, the 3rd day was chosen as the early time point. The 10th day was the later time point. The 7th day was the transition stage at which the BMSCs began changing to osteoplasts. We used flow cytometry to detect the cell cycle phases on the 7th day. Collagen I is the product of mature osteoplasts. Thus, reverse transcription-polymerase chain reaction (RT-PCR) was used on the 10th day. Cellular Proliferation In order to evaluate the effect of the 0.5-mT-intensity EMF stimulation on BMSC cellular proliferation, an MTT assay was performed on days 3, 7, and 10 post-culture. Accordingly, 3,000 cells/well were placed in a 96-well dish. In each well, 20 μL of the MTT solution (0.5%) was added, and cells were allowed to incubate with the MTT solution for 4 hours. The supernatant was subsequently discarded, and 150 μL of dimethyl sulfoxide was added to each well. Low-speed oscillation for 10 minutes was used to fully dissolve resulting crystals. The optical density (OD) values for each well were measured at an absorbance of 570 nm, with an empty well serving as the control. For observation of cell morphology, photographs were taken for each group on the fifth day post-culture. All experiments were repeated 3 times. Cellular Cycle We used flow cytometry to detect the cell cycle phases (G0/G1, S, and G2/M). Groups A, B, C, and D were cultured for 7 days under the conditions described above. Cells were detached from 25-cm2 tissue culture flasks using 10 mM of trypsin-EDTA and 1 mL of 0.25% trypsin, as indicated previously. Next, 1 × 106cells from each group were fixed in 1 mL of ice-cold 70% ethanol for 30 minutes. After centrifugation (1,000 rpm for 5 minutes), cells were resuspended in 980 mL of phosphate-buffered saline containing 50 mg/mL of the DNA-specific fluorescent probe propidium iodide and 10 mg/mL of ribonuclease and were incubated on ice for 1 hour. Fluorescence was analyzed with a FACScan flow cytometer (BD FACSCanto II, BD Biosciences, San Jose, California) equipped with a 15-mW, 488-nm, air-cooled argon ion laser. For detection of propidium iodide, fluorescence emissions were collected after passage through a 570-nm band-pass filter. Doublets and aggregates were gated out of analysis on the dot-plot, which consisted of the propidium iodide fluorescence peak versus the propidium iodide fluorescence area. A minimum of 10,000 events were acquired in list mode using the BD FACSDiva software (BD Biosciences). Forward and side scatters were collected as linear signals, and fluorescence emissions were collected on a linear scale. Cell cycle analysis was conducted using the SFit model included in the BD FACSDiva software. All experiments were repeated 3 times. The proliferation indexes for the BMSCs in all groups were analyzed statistically, as follows: Proliferation Index = [(S+ G2/M)/(G0/G1+S+G2/M)] × 100%. Determination of Cellular Differentiation In order to assess the effects of 0.5-mT-intensity EMF stimulation on the cell differentiation of BMSCs, we cultured the cells from each of the 4 groups in DMEM-LG medium containing 10% fetal bovine serum. The ALP activity was measured on day 7 post-culture using the instructions provided in the ALP kit. The OD values for each well were measured at an absorbance of 490 nm, with an empty well serving as the control. Statistical analysis was performed on the resulting OD values for each of the 4 groups. Alizarin Red S staining was used to determine the level of mineralization present in each group on day 12 post-culture. A representative image from each group was selected, and photographs were taken under an inverted microscope. RT-PCR We used RT-PCR to detect the expression of the collagen I gene. Collagen I is the body's major structural protein for connective tissue and is a major component of the extracellular matrix, providing a scaffold for bone mineralization.14 Collagen controls the formation and expression of cell morphology, cell differentiation, and a variety of other biological functions.15 Therefore, collagen I messenger RNA (mRNA) expression can serve as a marker for bone formation and remodeling. Cells from each group (A, B, C, and D) were cultured for 10 days, and the total RNA from each group was extracted using a Trizol solution (Life Technologies Corporation). A UV spectrophotometer (Eppendorf, Hauppauge, New York) was used to determine RNA sample concentration and purity. For real-time polymerase chain reaction (PCR), in accordance with the requirements of the reverse transcription kit used for this study, the complementary DNA samples were preserved at −20°C. For primer design and synthesis, fluorescent primers were designed by Primer Premier 6.0 (Premier Company, Canada) and Beacon software (Bio-Rad Laboratories, Hercules, California) based on the GenBank gene sequence. Primers were synthesized by Gene Tech Biotechnology (Shanghai) Co Ltd (Shanghai, China) as follows: Rat-18S(M11188): 5′ GAATTCCCAGTAAGTGCGGGTCATA3′,5′CGAGGGCCTCACTAAACCATC3′;RatCoI(NM053304):5′TGGCCAAGAAGACATCCCTGAAGT3′,5′ACATCAGGTTTCCACGTCTCACCA 3′. For analysis of real-time PCR results, the iQTM5 multi-instrument real time quantitative PCR detection system was used (Bio-Rad Corporation, Hercules, California). The 2-ΔCT method was used to calculate differences in RNA expression between the experimental groups and the control group. Determination of the Effects of Callus Formation in Mice After In Vitro EMF Stimulation In order to detect the effects of 0.5-mT-intensity EMFs on BMSCs in the living animals' bodies, 2-mm-diameter holes were drilled in the cortical bone of the right and left femurs in 8 mice using a dental drill. The 8 mice were divided into 2 categories. Each category included 2 male and 2 female mice. Before the in vivo experiment, EMF-exposed cells were exposed at 7 days post-culture; control cells were maintained in DMEM-LG medium for 7 days. Then the 2 different kinds of BMSCs were transplanted to PLGA scaffolds of 2 mm diameter and 2 mm height. The scaffolds were subsequently inserted into the drilled holes in the cortical bone of the right and left mice femurs separately. In each mouse, the untreated control cells with PLGA were inserted into the left femurs, and EMF-exposed cells with PLGA were inserted into the right femurs. All of the mice received the same surgery on the same day and in the same sterile environment. Before and after the surgery, the animals were kept in a 14/10-hour light/dark environment at a constant temperature of 22±3°C and humidity of 45%±10%. The 2 categories were sacrificed after 4 and 8 weeks separately, and the collected specimens were sliced and stained with hematoxylin and eosin. Data Analysis The data from each group were expressed as averages (±SD). The differences among various groups were evaluated using the analysis of variance method, and P values less than .05 were considered significant. All data were statistically analyzed with SPSS software, version 18.0 (SPSS Inc, Chicago, Illinois). Role of the Funding Source Funding support for this study was provided by the Department of Orthopedic Surgery, Second Affiliated Hospital, School of Medicine, Zhejiang University. Results Cell Proliferation Cell growth and morphology were observed daily under an inverted microscope. Four hours after inoculation, third-passage cells had adhered completely and began to proliferate and form colonies. After 24 hours, the EMF-stimulated BMSCs gradually began to proliferate at an increased rate and became triangular, polygonal, and scale-shaped. With increasing time of incubation, cells converged into a cobblestone pattern and eventually overlapped one another. Our analysis revealed that the cell densities of groups C and D were significantly higher than those of groups A and B. Groups C and D exhibited a more mature cell morphology than did groups A and B (eFigure). Figures 2 and 3 detail the significant differences in growth rate observed between the EMF-stimulated groups and the control group. After a 72-hour irradiation, the BMSCs were observed to proliferate more rapidly than those exposed to osteogenesis-inducing medium (P=.0312). Compared with the control, a higher proliferation rate also was observed upon treatment with osteogenesis-inducing medium (P=.0158); however, the proliferation rate of group D (EMF and osteogenesis-inducing medium) did not exceed that of group C (EMF alone), as the presence of dexamethasone in the medium stimulated the direct differentiation of stem cells into osteoblasts. In previous studies, dexamethasone strongly inhibited DNA (48-hour treatment) but stimulated ALP.16–18 Thus, the effects of the combined treatment on proliferation were weakened. The effect of dexamethasone is discussed later. Figure 2 Open in new tabDownload slide The methylthiotetrazole's optical density (OD) values (570 nm) of the 4 groups of mouse bone marrow stromal cells on the 3rd, 7th, and 10th days of the culture. After the intervention of electromagnetic fields (EMFs), the OD values of the EMF-exposed groups were significantly higher than that of the control group (P<.05). The OD values of the EMF-exposed groups were significantly higher than that of the group exposed to osteogenesis-inducing medium only (P<.05).The OD values of the group exposed to EMFs combined with osteogenesis-inducing medium was significantly higher than that of the group exposed to osteogenesis-inducing medium only (P<.05). Figure 2 Open in new tabDownload slide The methylthiotetrazole's optical density (OD) values (570 nm) of the 4 groups of mouse bone marrow stromal cells on the 3rd, 7th, and 10th days of the culture. After the intervention of electromagnetic fields (EMFs), the OD values of the EMF-exposed groups were significantly higher than that of the control group (P<.05). The OD values of the EMF-exposed groups were significantly higher than that of the group exposed to osteogenesis-inducing medium only (P<.05).The OD values of the group exposed to EMFs combined with osteogenesis-inducing medium was significantly higher than that of the group exposed to osteogenesis-inducing medium only (P<.05). Figure 3 Open in new tabDownload slide The alkaline phosphatase (ALP)'s optical density (OD) values (490 nm) of the 4 groups of mouse bone marrow stromal cells (BMSCs) on the 3rd, 7th, and 10th days of the culture. Different groups of BMSCs were cultured in different mediums, the ALP content of the cells in the group C (electromagnetic fields [EMFs] only) was higher than that of group A (control group) and group B (osteogenesis-inducing medium only) (P<.05). The ALP content of the cells in groups C and D (EMFs combined with osteogenesis-inducing medium) were significantly higher than that of group B (P<.05). Figure 3 Open in new tabDownload slide The alkaline phosphatase (ALP)'s optical density (OD) values (490 nm) of the 4 groups of mouse bone marrow stromal cells (BMSCs) on the 3rd, 7th, and 10th days of the culture. Different groups of BMSCs were cultured in different mediums, the ALP content of the cells in the group C (electromagnetic fields [EMFs] only) was higher than that of group A (control group) and group B (osteogenesis-inducing medium only) (P<.05). The ALP content of the cells in groups C and D (EMFs combined with osteogenesis-inducing medium) were significantly higher than that of group B (P<.05). Cell Differentiation In general, ALP activity increased around day 5 in all groups. The ALP activity also increased with prolonged stimulation (EMF or osteogenic medium). The EMF-stimulated groups (C and D) had significantly higher ALP activity at each time point compared with to the control group and the group exposed to osteogenic medium alone (groups A and B, respectively). The ALP activity in the group exposed to osteogenic medium was higher than that of the control group (P=.010). The effect of electromagnetic fields on ALP activity was more significant than that of osteogenic medium treatment. The ALP activity of group D (EMF + osteogenic medium) was significantly higher than that of group C (EMF stimulation only, P=.02). Group D exhibited the highest ALP activity (Fig. 3). The 4 groups were stained with Alizarin Red S on day 12 after treatment. The staining pattern shown in Figure 4 suggested the presence of mineral salt (calcium) deposits. The degree of calcification suggested BMSCs had undergone differentiation, with group D exhibiting the most obvious osteogenic differentiation. Figure 4 Open in new tabDownload slide Alizarin Red S-stained mineralization on the 12th day post-culture for (A) group A cells, (B) group B cells, (C) group C cells, and (D) group D cells. Alizarin Red S staining showed mineral deposits and integrating into round or oval calcified nodules (indicated by arrows). In the same range of observation, the nodules induced by electromagnetic fields (EMFs) (groups C and D) were significantly larger than those in the control group (group A). The effect of EMFs was direct viewing by Alizarin Red S staining. Figure 4 Open in new tabDownload slide Alizarin Red S-stained mineralization on the 12th day post-culture for (A) group A cells, (B) group B cells, (C) group C cells, and (D) group D cells. Alizarin Red S staining showed mineral deposits and integrating into round or oval calcified nodules (indicated by arrows). In the same range of observation, the nodules induced by electromagnetic fields (EMFs) (groups C and D) were significantly larger than those in the control group (group A). The effect of EMFs was direct viewing by Alizarin Red S staining. Cell Cycle As observed by flow cytometry, the EMF-stimulated samples did not contain abnormal DNA. The results of our cell cycle experiments are shown in Figure 5. The EMF stimulation resulted in changes in the cell cycle of BMSCs, with each treatment group exhibiting different outcomes. Compared with the control group (group A: 7.05%±0.37%), the percentage of cells in the G2/M+S phase increased significantly in the group exposed to EMFs (group C: 13.98%±0.23%) (P=.012). The percentage of cells in EFM + osteogenic medium (group D: 20.23%±0.28%) in the G2/M+S phase was significantly higher than that of cells exposed to osteogenic medium alone (group B: 12.41%±0.17%) (P=.006) and the control group (P=.000). Figure 5 Open in new tabDownload slide The Proliferation Index provided a detailed statement about the cell cycle phases. Proliferation Index = [(S + G2/M)/(GO/G1 + S + G2/M)] × 100 %. We found that exposure to electromagnetic fields (EMFs) significantly increased proliferation bone marrow stromal cells (BMSCs) compared with the group exposed to osteogenesis-inducing medium only and the control group, as indicated by the increased percentage of cells in the S and G2 phases of the cell cycle (P<.05). Exposure to EMFs significantly increased the proportion of cells in the mitotic phase, resulting in an enhanced rate of cell proliferation. Additionally, the Proliferation Index value of group C was not higher than that of group D at 7 days poststimulation. Figure 5 Open in new tabDownload slide The Proliferation Index provided a detailed statement about the cell cycle phases. Proliferation Index = [(S + G2/M)/(GO/G1 + S + G2/M)] × 100 %. We found that exposure to electromagnetic fields (EMFs) significantly increased proliferation bone marrow stromal cells (BMSCs) compared with the group exposed to osteogenesis-inducing medium only and the control group, as indicated by the increased percentage of cells in the S and G2 phases of the cell cycle (P<.05). Exposure to EMFs significantly increased the proportion of cells in the mitotic phase, resulting in an enhanced rate of cell proliferation. Additionally, the Proliferation Index value of group C was not higher than that of group D at 7 days poststimulation. RT-PCR The relative expression of the collagen I gene was determined after 10 days in culture. The EMF-exposed groups had significantly higher collagen I mRNA levels than the control group (P=.000). Combined treatment with osteogenic medium and EMF stimulation resulted in significantly higher collagen I expression than was observed for the group treated only with osteogenic medium and the control group (P=.000, Table). Table Relative Expression of Collagen I in Bone Marrow Stromal Cells From Sprague–Dawley Mice (2Δct×106) Groupa . Average . SD . A 30.45 4.51 B 62.36 6.00 C 102.22b 9.15 D 135.68b,c 14.66 Groupa . Average . SD . A 30.45 4.51 B 62.36 6.00 C 102.22b 9.15 D 135.68b,c 14.66 a Third-generation bone marrow stromal cells were divided into 4 groups. Cells from groups A and C were cultured in medium with 10% fetal bovine serum. Group A served as the control group, and group C was exposed to electromagnetic fields (EMFs) of 0.5 mT at 50 Hz. Groups B and D were exposed to osteogenesis-inducing medium containing 10% fetal bovine serum, 10 mmol/L 7-sodium glycerophosphate, 10 mmol/L dexamethasone, and 50 mg/L vitamin C). The cells in group B were not exposed to EMFs. Cells in group D were exposed to EMFs of the same intensity as those in group C, and the experimental duration was 8 hours every day. b Compared with the control group (P<.05). c Compared with the group induced by osteogenic medium (P<.05). Open in new tab Table Relative Expression of Collagen I in Bone Marrow Stromal Cells From Sprague–Dawley Mice (2Δct×106) Groupa . Average . SD . A 30.45 4.51 B 62.36 6.00 C 102.22b 9.15 D 135.68b,c 14.66 Groupa . Average . SD . A 30.45 4.51 B 62.36 6.00 C 102.22b 9.15 D 135.68b,c 14.66 a Third-generation bone marrow stromal cells were divided into 4 groups. Cells from groups A and C were cultured in medium with 10% fetal bovine serum. Group A served as the control group, and group C was exposed to electromagnetic fields (EMFs) of 0.5 mT at 50 Hz. Groups B and D were exposed to osteogenesis-inducing medium containing 10% fetal bovine serum, 10 mmol/L 7-sodium glycerophosphate, 10 mmol/L dexamethasone, and 50 mg/L vitamin C). The cells in group B were not exposed to EMFs. Cells in group D were exposed to EMFs of the same intensity as those in group C, and the experimental duration was 8 hours every day. b Compared with the control group (P<.05). c Compared with the group induced by osteogenic medium (P<.05). Open in new tab Animal Experiments As shown in Figure 6, both the EMF-stimulated and control groups did not show full decomposition of the PLGA scaffold; however, significant formation of new bone was observed around the edges of the scaffold in each group. The callus present in the control group had not developed into a film, whereas examination of the irradiated group revealed a scattered sheet with a small callus. After 8 weeks, the PLGA scaffold was still not completely degraded; however, the calluses present in each group began to interconnect. Compared with the control group, the EMF-irradiated group appeared to contain flaked bone tissue near the edges of the PLGA scaffold, and the callus arrangement appeared chaotic and plastic. Figure 6 Open in new tabDownload slide The results of the animal experiment: specimens were sliced and stained with hematoxylin and eosin. Double arrows indicate new-generation bone callus. Overlapping arrows indicate the materials of poly(lactic-co-glycolic acid) (PLGA). Single arrows indicate the original bone tissue. Images 1 and 2 show the results of the fourth week. Image 1 shows the results for the control group, in which the bone marrow stromal cells (BMSCs) did not receive any treatment. The BMSCs in image 2 were irradiated for 7 days. Images 3 and 4 show the results of the eighth week. Image 3 shows the results for the control group, in which the BMSCs did not receive any treatment. The BMSCs in image 4 were irradiated for 7 days. Significant formation of new bone was observed in each group. Callus formation was more significant in the EMF-treated group than in the untreated group, particularly at the 8-week time point. Figure 6 Open in new tabDownload slide The results of the animal experiment: specimens were sliced and stained with hematoxylin and eosin. Double arrows indicate new-generation bone callus. Overlapping arrows indicate the materials of poly(lactic-co-glycolic acid) (PLGA). Single arrows indicate the original bone tissue. Images 1 and 2 show the results of the fourth week. Image 1 shows the results for the control group, in which the bone marrow stromal cells (BMSCs) did not receive any treatment. The BMSCs in image 2 were irradiated for 7 days. Images 3 and 4 show the results of the eighth week. Image 3 shows the results for the control group, in which the BMSCs did not receive any treatment. The BMSCs in image 4 were irradiated for 7 days. Significant formation of new bone was observed in each group. Callus formation was more significant in the EMF-treated group than in the untreated group, particularly at the 8-week time point. Discussion In addition to investigating the effects of EMF-treated BMSCs in maintaining significant differentiation potential and high proliferation capability, we examined their effect on cell-cycle changes. Significantly higher levels of the collagen I gene were expressed compared with untreated cells at the middle to late stages of the osteogenic culture period. The effects of the in vivo EMF treatment on osteogenesis were observed 4 and 8 weeks after implantation. There was significantly more callus formation in the EMF-treated group than in the untreated group, especially after 8 weeks. The results indicated that the use of EMF at 0.5 mT and 50 Hz on BMSCs was safe. With extended irradiation time, EMFs (0.5 mT, 50 Hz) increased cell proliferation and induced cell differentiation. After transplantation into animals, the irradiated BMSCs exhibited more apparent biological characteristics of the original cells than did the control group. Used as a noninvasive physical therapy for delayed bone-fracture healing, EMF treatment has had substantial effects in orthopedic surgery; however, it remains controversial and is not established as a standard treatment. Electrotherapy has both positive and negative effects.17–19 Due to the lack of standards for equipment and associated treatment conditions, it is difficult to compare clinical and experimental results. Additionally, identifying the experimental parameters likely to have the greatest beneficial effects in physical therapy has been problematic. Zhang et al20 reported that rectangular EMF treatment of osteoblasts increased cellular proliferation and decreased ALP activity. Triangular EMFs had an accelerative effect on cellular mineralized nodules. Sinusoidal EMF treatment of osteoblasts decreased cellular proliferation, increased ALP activity, and suppressed mineralized nodule formation. The frequency and peak magnetic flux used in the experiments were 15 Hz and 5 mT, respectively.20 In a study by Sun et al,21 the growth rates in the exponential phase of the treatment groups were comparable to those of the control group, and 20% to 60% higher cell densities were achieved in the PEMF group during the exponential cell expansion stage. The PEMF consisted of 4.5-millisecond bursts repeating at 15 Hz and 20 pulses per burst. Sun et al, therefore, suggested that enhanced cell proliferation with PEMF exposure might have resulted from shortening the lag phase. Tsai et al22 showed that the higher-density group had more cells than the lower-density group, especially at days 3 and 5, although the number of cells appeared to decrease at day 7 when exposed to PEMF stimulation; repetitive, single, quasi-rectangular pulses with a pulse duration of 300 milliseconds were used at 7.5 Hz with exposure for 2 hours per day. In the PEMF-treated group with a low initial seeding density, the runt-related transscription factor 2/core-binding factor subunit alpha-1 (Runx2/Cbfa1) mRNA levels were significantly higher at day 7 and lower at day 10. There was no significant difference in osteogenic gene expression in PEMF-treated versus untreated cultures with high initial cell densities.22 Based on the literature, it appears that the differences in cell proliferation and differentiation potential under different EMF treatments might have resulted from altered cell metabolism progression caused by different types or numbers of cytokine receptors or secretions, such as collagen, expressed on BMSCs at different osteogenic stages. Differences in experimental conditions affect the expression of certain osteogenesis-related genes that enhance osteogenesis during BMSC osteogenic differentiation. The healing capacity of EMF depends on the operating parameters and exposure protocols, as the intensity, pulse duration, and exposure time can contribute to different results. The coherent vibration of the electric charge can cause irregular gating of electrosensitive channels on the plasma membrane and disrupt cellular electrochemical balance and function.23 Because the mechanism of EMF stimulation is unclear, it is necessary to discuss the postulated mechanisms at cellular, subcellular, and molecular levels. Additionally, an evaluation of the efficacy of EMF stimulation for healing requires precisely defined measurements and computations for multiple parameters, including amplitude, field gradients, and duration of exposure.24 Further methodical, randomized controlled research is needed. Resolution of the significant disparities among clinical targets, types of electrical stimulation, and clinical outcomes is needed.25 Reports on the effects of EMF stimulation on cellular proliferation and differentiation have been contradictory. Fitzsimmons et al26 hypothesized that the ability of EMF stimulation to increase bone formation depended on and was mediated by the increased number of cells associated with the stimulation of cellular proliferation. By contrast, Diniz et al27 did not agree with this hypothesis, as PEMF stimulation accelerated osteoblast proliferation only in the early stages of culture and did not increase the total number of cells in the later stages. In that study, there was a significant difference in MTT between the control and EMF-stimulated groups until the end of the experiment. The increase in ALP activity appeared to be an effect of EMF stimulation on bone-like cells. Our investigations demonstrated that EMF stimulation significantly increased ALP activity in BMSCs.22,28 These experiments were conducted with rats under 2 treatment protocols, EMF-exposed and general cells, simultaneously. Combined with tissue engineering, this test model showed positive effects of EMF in repairing bone fractures. It is hoped that there will be further progress in regenerative medicine and physical therapeutics in the near future.29 There are no similar in vivo studies in the literature. In vitro experiments have shown that dexamethasone present in an osteogenic induction medium can direct BMSC differentiation into osteoblasts. Dexamethasone is important for promoting the expression of a variety of mature osteoblast markers at the transcriptional level.30 Kasperk et al31 found that dexamethasone influenced the activation of proliferation-related oncogenes such as c-Jun, resulting in the inhibition of cell replication. Suarez et al32 showed that dexamethasone activated the parathyroid hormone signaling system, resulting in increases in bone morphogenetic protein (BMP). Mikami et al33 suggested that BMP-2 and dexamethasone synergistically increased ALP levels in mouse C3H10T1/2 pluripotent stem cells. They might act in the same pathway or at the same stage of differentiation. Taken together, the literature and our results indicate that dexamethasone promotes ALP activity and bone nodule formation and directs proliferative osteoprogenitor cells toward terminal maturation. However, dexamethasone has side effects.34,35 Glucocorticoid can inhibit bone formation by suppressing osteoblast proliferation and resorption by suppressing osteoblast precursor recruitment.36,37 Compared with osteogenesis-inducing medium, EMF had satisfactory effects on differentiation potential and proliferation capability. We found that EMF exposure significantly increased BMSC proliferation compared with exposure to osteogenic medium or no exposure, as indicated by the increased percentage of cells in the S and G2 phases of the cell cycle. Additionally, the propidium iodide value of group C was no greater than that of group D at 7 days after stimulation. This finding might have been due to growth limitations in 6-well plates; the proliferation rate of group C probably was inhibited by cell-cell contact. Alternatively, we might have missed the most significant period of proliferation for group C. As Santini et al38 demonstrated, after 7 days of growth, the percentage of cells in the G0/G1, S, and G2/M phases of MG-63 control cells differed significantly from those of PEMF-exposed cells, with the value reaching 91.9%±3.8%. Collectively, the empirical evidence suggests that EMF exposure significantly increases the proportion of cells in the mitotic phase, enhancing the rate of cell proliferation. Because bone tissue–related cells, cytology, and microenvironment contribute to bone regeneration and healing, using animal models is essential for understanding repair processes and assessing the value of new therapies. In this study, we did not replicate the in vitro conditions for treatment with osteogenic medium and could not establish a stricter model for studying osteogenesis. However, we are establishing a more consistent animal model to examine bone repair in the second phase of the project. In animal experiments, mice are the main experimental species, so discrepancies are inevitable.21,39 However, a move toward in vivo experiments involving larger mammals should provide more conclusive results. Therefore, investigations using animals such as rabbits or sheep are needed. Clinically, the experimental results must be confirmed in human clinical trials. Regarding physical therapy, Dallari et al2 published the first randomized, prospective, double-blind study examining whether PEMF stimulation fostered bone healing and reduced the time to functional recovery. A positive effect of PEMF stimulation on bone mineral density and functional recovery of patients were observed. The treatment had no associated negative side effects. Aaron et al40 reported that EMF treatment was associated with decreased pain and improved function, effectively improving the symptoms of patients with bone defects. In 2009, Doorgakant et al41 reported the use of a PEMF bone-stimulation device to treat a patient with a tibial periprosthetic fracture following revision knee arthroplasty. Additionally, the effect of PEMF was investigated in 30 individuals undergoing hip revision.2 That study showed that PEMF treatment assisted in clinical recovery and bone-stock restoration. These reports suggested that low-intensity EMF affects the biomechanical and chemical properties of animal bones, especially cortical bone quality and bone strength. Therefore, EMF can affect the intrinsic properties of bone structure by stimulating inactive bone cells responsible for bone remodeling. Our experimental results have provided further rationale for the use of low-intensity EMFs. They are safe and aid clinical recovery and bone restoration. All data support the view that magnetic fields can be applied to stimulate cell growth in bone-like structures in vitro and in vivo. Furthermore, a 0.5-mT, 50-Hz intensity is ideal for in vivo use of EMFs for bone disorders. One limitation of these studies is that only short-term results have been investigated. Peter et al39 reported that differences between treated and control rats became evident after 3 months, and significant bone loss of up to 14% occurred during the first 3 months after total hip arthroplasty. Long-term follow-up studies are essential to clarify the effects of EMFs. The findings will contribute to biomaterial production designed for clinical applications. In conclusion, EMF exposure induced ALP secretion, which is important in bone formation, and increased collagen I gene expression. Furthermore, the percentage of cells in the G2/M+S phase increased significantly in the group exposed to EMFs, suggesting that EMF exposure increased DNA synthesis and replication, promoting mitotic proliferation, and that exposure to EMFs should promote bone formation. Additionally, morphological and structural results demonstrated that EMF use had a positive therapeutic effect, with enhanced early osseointegration in trabecular bone and a greater degree of bone mineralization and maturation. However, these results were insufficient to elucidate the mechanism of EMFs fully. Our observations of the in vivo and in vitro effects of EMF on bone regeneration provide the groundwork for developing new physical therapeutic techniques in orthopedics, which will provide the basis for further studies of the biological effects of EMFs. " Funding support for this study was provided by the Department of Orthopedic Surgery, Second Affiliated Hospital, School of Medicine, Zhejiang Univeristy. 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Dr He provided project management. © 2012 American Physical Therapy Association
TI - Effects of Low-Intensity Electromagnetic Fields on the Proliferation and Differentiation of Cultured Mouse Bone Marrow Stromal Cells
JF - Physical Therapy
DO - 10.2522/ptj.20110224
DA - 2012-09-01
UR - https://www.deepdyve.com/lp/oxford-university-press/effects-of-low-intensity-electromagnetic-fields-on-the-proliferation-VEnbWxcDS3
SP - 1208
VL - 92
IS - 9
DP - DeepDyve
ER -