Stem Cells

Stem Cell Res Ther. 2018 May 21;9(1):143. doi: 10.1186/s13287-018-0883-4.

Extremely low frequency electromagnetic fields promote mesenchymal stem cell migration by increasing intracellular Ca2+ and activating the FAK/Rho GTPases signaling pathways in vitro.

Zhang Y1, Yan J1, Xu H1, Yang Y1, Li W1, Wu H2, Liu C3.

Author information

1
Department of Orthopedics, Tongji Hospital, Tongji Medical College, Huazhong University of Science and Technology, Jiefang Avenue 1095, Wuhan, 430030, China.
2
Department of Orthopedics, Tongji Hospital, Tongji Medical College, Huazhong University of Science and Technology, Jiefang Avenue 1095, Wuhan, 430030, China. wuhua360@aliyun.com.
3
Department of Orthopedics, Tongji Hospital, Tongji Medical College, Huazhong University of Science and Technology, Jiefang Avenue 1095, Wuhan, 430030, China. liu.chaoxu@tjh.tjmu.edu.cn.

Abstract

BACKGROUND:

The ability of mesenchymal stem cells (MSCs) to migrate to the desired tissues or lesions is crucial for stem cell-based regenerative medicine and tissue engineering. Optimal therapeutics for promoting MSC migration are expected to become an effective means for tissue regeneration. Electromagnetic fields (EMF), as a noninvasive therapy, can cause a lot of biological changes in MSCs. However, whether EMF can promote MSC migration has not yet been reported.

METHODS:

We evaluated the effects of EMF on cell migration in human bone marrow-derived MSCs. With the use of Helmholtz coils and an EMF stimulator, 7.5, 15, 30, 50, and 70 Hz/1 mT EMF was generated. Additionally, we employed the L-type calcium channel blocker verapamil and the focal adhesion kinase (FAK) inhibitor PF-573228 to investigate the role of intracellular calcium content, cell adhesion proteins, and the Rho GTPase protein family (RhoA, Rac1, and Cdc42) in EMF-mediated MSC migration. Cell adhesion proteins (FAK, talin, and vinculin) were detected by Western blot analysis. The Rho GTPase protein family activities were assessed by G-LISA, and F-actin levels, which reflect actin cytoskeletal organization, were detected using immunofluorescence.

RESULTS:

All the 7.5, 15, 30, 50, and 70 Hz/1 mT EMF promoted MSC migration. EMF increased MSC migration in an intracellular calcium-dependent manner. Notably, EMF-enhanced migration was mediated by FAK activation, which was critical for the formation of focal contacts, as evidenced by increased talin and vinculin expression. Moreover, RhoA, Rac1, and Cdc42 were activated by FAK to increase cytoskeletal organization, thus promoting cell contraction.

CONCLUSIONS:

EMF promoted MSC migration by increasing intracellular calcium and activating the FAK/Rho GTPase signaling pathways. This study provides insights into the mechanisms of MSC migration and will enable the rational design of targeted therapies to improve MSC engraftment.

KEYWORDS:

Cell migration; Electromagnetic fields; Focal adhesion kinase; Intracellular Ca2+?; Rho GTPase protein famil

Int J Mol Sci. 2018 Mar 27;19(4). pii: E994. doi: 10.3390/ijms19040994.

Co-Culture with Human Osteoblasts and Exposure to Extremely Low Frequency Pulsed Electromagnetic Fields Improve Osteogenic Differentiation of Human Adipose-Derived Mesenchymal Stem Cells.

Ehnert S1, van Griensven M2, Unger M3, Scheffler H4, Falldorf K5, Fentz AK6, Seeliger C7, Schröter S8, Nussler AK9, Balmayor ER10.

Author information

1
Siegfried Weller Institute for Trauma Research, Eberhard-Karls-Universität Tübingen, 72076 Tübingen, Germany. sabrina.ehnert@med.uni-tuebingen.de.
2
Experimental Trauma Surgery, Klinikum rechts der Isar, Technical University of Munich, 81675 München, Germany. martijn.vangriensven@tum.de.
3
Experimental Trauma Surgery, Klinikum rechts der Isar, Technical University of Munich, 81675 München, Germany. marina.unger@tum.de.
4
Siegfried Weller Institute for Trauma Research, Eberhard-Karls-Universität Tübingen, 72076 Tübingen, Germany. hscheffler@bgu-tuebingen.de.
5
Sachtleben GmbH, 20251 Hamburg, Germany. falldorf@citresearch.de.
6
Sachtleben GmbH, 20251 Hamburg, Germany. fentz@citresearch.de.
7
Experimental Trauma Surgery, Klinikum rechts der Isar, Technical University of Munich, 81675 München, Germany. claudine.seeliger@tum.de.
8
Siegfried Weller Institute for Trauma Research, Eberhard-Karls-Universität Tübingen, 72076 Tübingen, Germany. sschroeter@bgu-tuebingen.de.
9
Siegfried Weller Institute for Trauma Research, Eberhard-Karls-Universität Tübingen, 72076 Tübingen, Germany. andreas.nuessler@med.uni-tuebingen.de.
10
Experimental Trauma Surgery, Klinikum rechts der Isar, Technical University of Munich, 81675 München, Germany. elizabeth.rosado-balmayor@tum.de.

Abstract

Human adipose-derived mesenchymal stem cells (Ad-MSCs) have been proposed as suitable option for cell-based therapies to support bone regeneration. In the bone environment, Ad-MSCs will receive stimuli from resident cells that may favor their osteogenic differentiation. There is recent evidence that this process can be further improved by extremely low frequency pulsed electromagnetic fields (ELF-PEMFs). Thus, the project aimed at (i) investigating whether co-culture conditions of human osteoblasts (OBs) and Ad-MSCs have an impact on their proliferation and osteogenic differentiation; (ii) whether this effect can be further improved by repetitive exposure to two specific ELF-PEMFs (16 and 26 Hz); (iii) and the effect of these ELF-PEMFs on human osteoclasts (OCs). Osteogenic differentiation was improved by co-culturing OBs and Ad-MSCs when compared to the individual mono-cultures. An OB to Ad-MSC ratio of 3:1 had best effects on total protein content, alkaline phosphatase (AP) activity, and matrix mineralization. Osteogenic differentiation was further improved by both ELF-PEMFs investigated. Interestingly, only repetitive exposure to 26 Hz ELF-PEMF increased Trap5B activity in OCs. Considering this result, a treatment with gradually increasing frequency might be of interest, as the lower frequency (16 Hz) could enhance bone formation, while the higher frequency (26 Hz) could enhance bone remodeling.

KEYWORDS:

extremely low frequency pulsed electromagnetic fields (ELF-PEMF); primary human adipose-derived mesenchymal stem cells (Ad-MSCs); primary human osteoblasts (OBs); primary human osteoclasts (OCs)

Logo of scirep

About Editorial Board For Authors Scientific Reports
Sci Rep. 2018; 8: 5108.
Published online 2018 Mar 23. doi:  10.1038/s41598-018-23499-9
PMCID: PMC5865106
PMID: 29572540

Pulsed electromagnetic fields increase osteogenetic commitment of MSCs via the mTOR pathway in TNF-? mediated inflammatory conditions: an in-vitro study

Letizia Ferroni,1 Chiara Gardin,1 Oleg Dolkart,corresponding author2 Moshe Salai,2 Shlomo Barak,3 Adriano Piattelli,4 Hadar Amir-Barak,5and Barbara Zavan1
1Department of Biomedical Sciences, University of Padova, Via G. Colombo 3, 35100 Padova, Italy
2Division of Orthopaedic Surgery, Tel Aviv Sourasky Medical Center, Tel Aviv University Sackler Faculty of Medicine, Tel Aviv, Israel
3Private Practice, Tel Aviv, Israel
4Department of Medical, Oral, and Biotechnological Sciences, University of Chieti-Pescara, Chieti, Italy
5Department of Internal Medicine E, Tel Aviv Sourasky Medical Center, Tel Aviv University Sackler Faculty of Medicine, Tel Aviv, Israel
Oleg Dolkart, moc.liamg@otraklod.
corresponding authorCorresponding author.
Author information ? Article notes ? Copyright and License information ? Disclaimer
Received 2017 Jul 11; Accepted 2018 Mar 14.

Abstract

Pulsed electromagnetic fields (PEMFs) have been considered a potential treatment modality for fracture healing, however, the mechanism of their action remains unclear. Mammalian target of rapamycin (mTOR) signaling may affect osteoblast proliferation and differentiation. This study aimed to assess the osteogenic differentiation of mesenchymal stem cells (MSCs) under PEMF stimulation and the potential involvement of mTOR signaling pathway in this process. PEMFs were generated by a novel miniaturized electromagnetic device. Potential changes in the expression of mTOR pathway components, including receptors, ligands and nuclear target genes, and their correlation with osteogenic markers and transcription factors were analyzed. Involvement of the mTOR pathway in osteogenesis was also studied in the presence of proinflammatory mediators. PEMF exposure increased cell proliferation and adhesion and the osteogenic commitment of MSCs even in inflammatory conditions. Osteogenic-related genes were over-expressed following PEMF treatment. Our results confirm that PEMFs contribute to activation of the mTOR pathway via upregulation of the proteins AKT, MAPP kinase, and RRAGA, suggesting that activation of the mTOR pathway is required for PEMF-stimulated osteogenic differentiation. Our findings provide insights into how PEMFs influence osteogenic differentiation in normal and inflammatory environments.

Introduction

Pulsed electromagnetic fields (PEMFs) have long been known to accelerate fracture repair. Exposure to PEMFs has been shown to affect cell proliferation and differentiation by influencing multiple metabolic pathways, depending upon lineage and maturation stage. In the osteoblast lineage, PEMFs contribute to bone formation induced by a demineralized bone matrix and stimulate fracture healing, probably through the action of progenitors that are already committed towards bone. Data on the mechanism of action of PEMFs and the potential involvement of specific signal transduction pathways are, however, scarce. It has been reported that PEMFs increase the activity of certain kinases belonging to known intracellular signaling pathways, such as the protein kinase A (PKA) and the MAPK ERK1/2,, and that they modulate anti-inflammatory effects by increasing the quantity of the adenosine receptors A2A. PEMFs stimulation also upregulates BMP2 expression in association with increased differentiation in mesenchymal stem cells (MSCs),.

Dental implants and total joint replacements are surgical procedures that involve the implantation of permanent biomaterials. An increasing number of these procedures has been extended to younger and middle-age patients, making long-standing biocompatibility, robustness and functionality crucial requirements for these implants. Despite many recent advances, revision surgeries of the implants continue to be a major concern due to the tissue response induced by implanted biomaterials, as well as the potential for loosening and periprosthetic osteolysis which remain significant challenges.

The basis of recent insights into osseointegration range from the pure bone healing that takes place around the implant to an immune-mediated foreign body reaction. That reaction involves a sequence of events, including protein adsorption on the surface of the implant, activation of complement and the coagulation system, recruitment of monocyte/macrophages and MSCs, activation and differentiation of these cells into functional macrophages, osteoclasts, and osteoblasts, respectively, and the formation of biological attachments between implant and new bone. The continued release of wear debris from the implants and the potential evolving infection during the lifespan of the implant might induce peri-implant inflammation, resulting in peri-implant osteolysis, aseptic loosening and subsequent implant failure necessitating further surgical intervention.

Serine/threonine kinase mammalian target of rapamycin (mTOR) has been shown to play an important role in osteoclast differentiation. It is activated by macrophage colony-stimulating factor, and its inhibition leads to decreased osteoclastogenesis,. Furthermore, mTOR expression levels are higher at the earlier stages of osteoclastogenesis and decrease at the later stages of osteoclast formation. mTOR exists in cells as part of two complexes: complex 1 (mTORC1) and complex 2 (mTORC2). mTORC1 is activated by amino acids, growth factors, oxygen, inflammation, and Wnt signaling. mTORC1 is also a negative regulator of autophagy, a lysosomal degradation process responsible for the removal of long-lived proteins and damaged organelles,. It has also been confirmed that the mTOR signaling pathway was involved in the regulation of apoptosis and autophagy in MSCs, and that its inhibition is able to attenuate age-related changes in MSCs.

This study aimed to assess the potential involvement of the mTOR signaling pathway in the osteogenic differentiation of MSCs, the cells naturally involved in bone repair processes, under stimulation with PEMFs. To this end, we analyzed potential changes in the expression of mTOR signaling pathway components, including receptors, ligands and nuclear target genes, and their correlation with osteogenic markers and transcription factors. PEMFs were generated using a miniaturized electromagnetic device (MED) (Magdent Ltd., Tel Aviv, Israel) that is used successfully to stimulate implant osseointegration in the clinical setting and in vivo to. The involvement of mTOR pathway in osteogenesis was also studied in the presence of proinflammatory mediators.

Results

Proliferation

The biocompatibility of the surface was evaluated by MTT testing for measuring mitochondria activity as well as by evaluating cell numbers. Figure 1A displays the results of MTT testing conducted in normal conditions and in the presence of proinflammatory cytokines. Mitochondrial activity increased over time in both the control and PEMF groups. The presence of inflammatory cytokines caused a well-defined decrease in MTT values. The same pattern of increased cell proliferation was demonstrated by monitoring the cell numbers (Fig. 1B). Specifically, fewer cells were found in inflammatory conditions. Moreover, PEMF treatment was able to increase cell proliferation in both conditions. The proliferation rate was significantly higher in the PEMF group compared to the controls, even in an inflammatory environment.

An external file that holds a picture, illustration, etc. Object name is 41598_2018_23499_Fig1_HTML.jpg

MSCs subjected to PEMF irradiation in the presence of proinflammatory cytokines for 30 days. (A) MTT proliferation assay. Results are expressed as mean?±?SD of at least 3 independent experiments, *p?<?0.05. (B) DNA content quantification. Results are expressed as mean?±?SD of at least 3 independent experiments, *p?<?0.05.

Morphology and cell adhesion properties

Morphologic analyses of MSCs were performed. Phalloidin-labeled F-actin (red), DAPI nuclear staining (blue) and overlaid fluorescent image of immunostained cellular components (merged) for the MSCs of the control and PEMF-treated groups are seen in Fig. 2. As shown in Fig. 2, the cells were able to attach to the implant surface in both the PEMF and control groups. The number of cells present on the implant surface with PEMF was clearly higher compared to the number of cells in the control group.

An external file that holds a picture, illustration, etc. Object name is 41598_2018_23499_Fig2_HTML.jpg

Morphologic analyses of MSCs. Phalloidin-labeled F-actin (red), DAPI nuclear staining (blue) and overlaid fluorescent image of immunostained cellular components (merged) for the MSCs of the control and PEMF-treated groups. After 7 days of culture, the cells were well-colonized throughout the implant surface, demonstrating a star-like shape associated with osteoblastic features. The cells were also able to spread after 7 days. PEMF irradiation resulted in a greater number of cells that were attached to the surfaces.

Cell adhesion properties were assessed by the analyses of gene expression of molecules involved on hyaluronan synthesis (HAS1), including receptor for extracellular hyaluronic acid molecules (CD44), integrin (ITGA1, 2, 3, 4), and cell adhesion molecules of the cadherine family, such as NCAM, VCAM, and PCAM (Fig. 3). The results are reported in all the graphs as an increase of gene expression value in samples of cells cultured in control conditions compared to cells exposed to PEMFs. PEMFs generated by MED were able to induce an increase in the expression of all these molecules, thereby confirming that they may enhance the adhesion properties of the cells. The presence of inflammatory stimuli (Fig. 3B) resulted in a reduction of cell adhesion, however, the presence of a PEMF significantly increased the expression of the integrin and cadherin receptors, thus potentially improving the ability of the cell to attach to the surface.

An external file that holds a picture, illustration, etc. Object name is 41598_2018_23499_Fig3_HTML.jpg

Analyses of cell adhesion properties in normal conditions (A) and in the presence of inflammation (B) were conducted by searching for the expression of molecules involved in hyaluronian synthesis (HAS1), i.e., extracellular receptor for hyaluronic acid (CD44), integrin (ITGA1, 2, 3, 4), and cadherin family cell adhesion molecules (NCAM; VCAM; PCAM). The results are reported as an increase in the gene expression value in samples of cells cultured on implants with MED device compared to the same gene expression obtained in normal conditions.

Osteogenic process

Real-time PCR for principal osteogenic markers, such as Runx, osteopontin, osteonectin, osteocalcin, collagen type I, wnt, foxO, ALP, BMP2, and BMP7 was performed in order to evaluate the commitment of MSCs onto osteoblastic phenotypes. The cells were cultured in the presence (Fig. 4A) and in the absence of inflammatory conditions (Fig. 4B) in order to compare the variations obtained in the control group with those obtained in the PEMF group. As illustrated in Fig. 4, in all the conditions an increase in expression of all osteogenic markers was noticed, confirming that the presence of PEMF exerts a positive effect on this process even in the presence of inflammatory cytokines. This commitment was confirmed by quantified ALP activity when MSCs were cultured in both the control and PEMF groups in the presence and absence of inflammatory stimuli (Fig. 5). Additionally, PEMFs were also able to induce a positive effect on the osteogenic process. It was clear that MSCs were also able to produce higher values of ALP in the presence of inflammatory cytokines as well. There was a significant, time-dependent ALP activity for cells grown under PEMF treatment, demonstrating the promotion of the crystallization of hydroxyapatites, a typical feature of pre-osteoblastic cells.

An external file that holds a picture, illustration, etc. Object name is 41598_2018_23499_Fig4_HTML.jpg

Real-time PCR for principal osteogenic markers, such as Runx, osteopontin, osteonectin, osteocalcin, collagen type I, wnt, foxO, ALP, BMP2, and BMP7 was performed in order to evaluate the commitment of stem cells onto an osteoblastic phenotype. The cells were cultured in the (A) presence and (B) absence of inflammatory conditions, and the variations obtained in normal implants versus implants?+?MED were compared.

An external file that holds a picture, illustration, etc. Object name is 41598_2018_23499_Fig5_HTML.jpg

Quantification of intracellular ALP activity (expressed as U/mL) in MSC exposed to PEMFs and in non-exposed MSC in the presence and absence of an inflammatory environment at 15 and 30 days. Results are expressed as mean?±?SD of at least 3 independent experiments, *p?<?0.05. *; **p?=?0.01; ***p?=?0.001.

mTOR pathway

In order to test if PEMF is able to excerpt its osteogenic properties thought mTOR pathway we used rapamicin to verify following hypothesis:

  1. rapamicin is able to reduce the osteogenic properties in absence of PEMF(control);
  2. The exposure to PEMF in presence of rapamicin could restore the osteogenic commitment of MSCs.

The osteogenic properties of MSCs seeded in the osteogenic medium have been evaluated as their ability to produce a mineralized extracellular matrix by means the ARS test. Figure 6 reports the staining on implant (A); on the medium (B) and the quantification of ARS staining (C). The osteogenic potential is related to the ability to produce a mineralized matrix. Higher values of mineralization are represented by a greater values of the red staining (Fig. 6A,B). Spectroscopy was used to assess these parameters. The quantification of the osteogenic potential is reported in Fig. 6C. It is well evident that both in normal condition (passive implant) and in presence of PEMF (active implant) a decent quantity of ARS is detectable at the time frame of 7 to 14?days. When Rapamicin was added, a well-defined decline was noticed, predominantly at 14 days in passive condition. On the contrary, in presence of PEMF, Rapamicin was not able to inhibit the process and mineralization of the extracellular environment was demonstrated.

An external file that holds a picture, illustration, etc. Object name is 41598_2018_23499_Fig6_HTML.jpg

The osteogenic properties of MSCs seeded in the osteogenic medium have been evaluated as their ability to produce a mineralized extracellular matrix by means the ARS test. staining on implant (A); on the medium (B) and the quantification of ARS staining (C). Results are expressed as mean?±?SD of at least 3 independent experiments, **p?=?0.01.

Similar phenomenon was observed at gene expression level as well. In Figs 7 and ?and88 we report the gene expression of markers related to mTOR pathway evaluated at 14 day on MSCs cultures seeded in osteogenic medium with rapamicin, with or without PEMF treatment (passive VS active implant). Results have been grouped in correlation to their involvement in mTOR pathway: mTOR1 Complexes; mTOR2 Complexes; mTOR Upstream Regulators – negative regulation; mTOR Upstream Regulators – positive regulation; mTOR Downstream Regulators – negative regulation; mTOR Downstream Regulators – positive regulation. The results were analyzed and are presented as the ratio between: active implant in osteogenic medium?+?rapamicin with the active implant in osteogenic medium without rapamicin; passive implant in osteogenic medium?+?rapamicin with the passive implant in osteogenic medium without rapamicin. Value comprises from ?2 to +2 are related to no significant variation. All genes related to the ratio in presence of a PEMF (active implant) are from ?2 to +2, indicating that no difference occurs in co-presence of rapamicine and PEMF. On the contrary in absence of PEMF defined up or dowregulation to gene related to mTOR1 involved mostly in Adipogenic commitment rather than osteogenic commitment of MSCs were demonstrated.

An external file that holds a picture, illustration, etc. Object name is 41598_2018_23499_Fig7_HTML.jpg

Gene expression of mTOR activity: (A) positive regulator, (B) negative regulator. (C) downstream effector: positive regulation, and (D) downstream effector: negative regulation.

An external file that holds a picture, illustration, etc. Object name is 41598_2018_23499_Fig8_HTML.jpg

Real-time PCR analysis of mTOR pathway markers. Gene expression levels of the selected markers are reported as ration of MSC coltured on active implants in presence of osteogenic medium and Rapamicin implants with passive implants in presence of osteogenic medium and Rapamicin. Results are expressed as mean?±?SD of at least 3 independent experiments, **p?=?0.01.

In order to highlight genes responsive to the PEMF stimulus we analyzed also the results related to the ratio of gene expression of active implant in presence of osteogenic medium plus rapamicin with the passive implant in osteogenic medium?+?rapamicine (Fig. 7).

As reported in Fig. 8, a significant difference was found in presence of PEMF and is related to:

  • •Decrease in RICTOR (receptor for mTOR2)
  • •Decrease in Protein phosphatase 2, regulatory subunit B, beta (PPP2R2B) involved on mTOR2 pathway
  • •Decrease in PKC protein (involved on Adipogenesis)
  • •Increase on VEGF (involved on angiogenesis)
  • •Decrease in Upstream regulator of negative mTOR regulator:

Protein kinase, AMP-activated, beta 1 non-catalytic subunit (PRKAB1)

Protein kinase, AMP-activated, beta 2 non-catalytic subunit (PRKAB2)

Protein kinase, AMP-activated, gamma 3 non-catalytic subunit (PRKAG3)

  • •Decrease in downstream stream regulator of negative mTOR regulator:

Calcium binding protein 39-like (CAB39L)

DNA-damage-inducible transcript 4 (DDIT4)

DNA-damage-inducible transcript 4-like (DDIT4L)

STE20-related kinase adaptor beta (STRADB)

The results indicated that PEMFs enhance mTOR signaling by inducing an increase in the value of its related proteins, such as AKT, MAPP kinase, and RRAGA. Additionally, a significant increase in Rho family of GTPases was detected. Rho family members play crucial roles in mechanical signal transduction and promote the differentiation of MSCs into osteoblasts.

Interleukin expression

MSCs were treated with PEMF in the presence of inflammatory cytokines as well as in the presence of PEMF. The results of their effect on inflammatory/anti-inflammatory activities of a PEMF are shown in Fig. 9, and they indicate that the presence of a PEMF induced a significant increase of in vitro expression of IL-10 (that exerts anti-inflammatory activity). Conversely, there was a reduction of expression of pro-inflammatory cytokines, such as IL-1, following PEMF treatment. There was no significant difference in expression of the other selected cytokines.

An external file that holds a picture, illustration, etc. Object name is 41598_2018_23499_Fig9_HTML.jpg

MSC were treated with inflammatory cytokines in the presence and absence of PEMFs. The results of the effect on inflammatory/anti-inflammatory activities of the active implants on MSC indicate a significant increase of in vitro expression of IL-10 (that exerts anti-inflammatory activity) in the presence of PEMFs generated by the MED device. Conversely, there is a reduction of expression of inflammatory cytokines, such as IL-1, in the presence of PEMFs. No significant difference in the expression of the other tested cytokines is evident.

Discussion

The principal results of the present study revealed several novel findings regarding the events involved in the stimulation of the osteogenic differentiation of MSCs induced by PEMFs. They identified a significant role of mTOR signaling during the differentiation driven by PEMF stimulation in an osteogenic microenvironment. Additionally, PEMFs were able to preserve the proliferation rate of MSCs in inflammatory conditions equal to that in a normal environment. MED-induced PEMF treatment resulted in an immunomodulatory effect in MSCs as expressed by increased IL-10 secretion. We found that PEMF stimulation of MSC proliferation mainly affected cell cycle regulation, cell structure, extracellular matrix, and some growth receptors involved in kinase pathways.

The osteointegration process begins with an inflammatory stage followed by the migration of MSCs. One of the major goals of dental, orthopedic and maxillofacial surgery is to achieve good and rapid osteointegration between implants and bone. The main research strategies to reduce implant failure aim at improving biomaterial characteristics, or stimulating bone endogenous repair, through a careful assessment of both processes by means of in vitro and in vivo experimental models before any application in humans. It had been reported that MED-generated PEMFs stimulated early bone formation around dental implants, already resulting in higher peri-implant bone-implant contact and bone mass after only 2 weeks, which suggests an acceleration of the osseointegration process by more than 3-fold. However, the exact biologic mechanism of the influence of PEMFs on bone regeneration remains to be elucidated. A recent study by Ferroni et al. concluded that PEMFs affect the osteogenic differentiation of MSCs only if they are pre-committed, and that this therapy can be an appropriate candidate for the treatment of conditions requiring an acceleration of the repair process.

We raised two major questions concerning the PEMF-related mechanism in the current study. First, we looked into the effects of PEMFs on MSCs in an inflammatory environment with regard to the ability of the cells to proliferate and adapt to the immunomodulatory changes. Understanding the mechanism of the implant’s integration, particularly the inflammatory response, is relevant for finding new treatment modalities to optimize the osteointegration and subsequent stability of the implants, which have implications in dentistry and orthopedic surgery. In this study, we added pro- and anti-inflammatory cytokines, which simulate the kinetics of their expression during early stages of implant integration in vivo, and investigated their effects on the proliferation and osteogenic differentiation of MSCs under PEMF irradiation. The proliferative capacity of MSCs is highly relevant for tissue repair. Cytokines are known to affect proliferation of different cell types. Therefore, we first analyzed the effect of selected cytokines on MSC proliferation. To the best of our knowledge, no previous study had assessed the influence of PEMF irradiation on the production of cytokines in MSC cultures. There are published data on the post-irradiation release of cytokines in mature osteoblasts and in osteoclast-like cells. In both of those studies, ELISA was used for quantification and demonstrated an increase of TNF-a, IL-1b and PG-E2 in relation both to the recruitment of the osteoclast-like cells and to the intensity of the electrical field. The current study demonstrated the ability of MED-generated PEMFs to alter the immuno-modulative activity properties of MSCs. A significant elevation in anti-inflammatory cytokines, such as IL-10, was clearly present when MSCs were seeded on implants. IL-10 acted as an anti-inflammatory substance by inhibiting the synthesis of proinflammatory cytokines, and its up-regulation in MSCs may counteract the detrimental proinflammatory effects.

Second, we examined the effects of PEMFs on the mTOR signaling pathway, and the results confirmed that PEMFs in the presence of an inflammatory environment positively affected MSC commitment into an osteoblastic phenotype through the mTOR pathway. In in vivo model demonstrated that the IGF-1 released from the bone matrix during bone remodeling stimulated osteoblastic differentiation of recruited MSCs by activation of Akt/mTOR. It had been reported that the presence of a good bone-like extracellular matrix was able to maintain bone mass by activation of mTOR in mesenchymal stem cells. We now demonstrated that PEMF irradiation positively stimulated mTOR signaling, thus increasing the osteoblastic commitment of MSCs in the presence of inflammatory stimuli as well. This commitment could also be induced by increased integrin expression, such as ?(4)?(1) integrin that has a high affinity for bone and improves the homing of MSCs to bone, thus promoting osteoblast differentiation and bone formation. mTOR is a central molecule in the regulation of cell growth in a wide variety of cells including osteoblasts, adipocytes, and myocytes. mTOR interacts with several proteins to form two distinct complexes named mTOR complex 1 (mTORC1) and 2 (mTORC2) which differ in their unique components, Raptor and Rictor. Upstream regulation and downstream products of mTORC1 are much more investigated than that of mTORC2. Though it is widely believed that the inhibition of mTOR signaling can promote osteoblastic differentiation, this issue is still controversial. While rapamycin primarily inhibits mTORC1, prolonged exposure can also disrupt mTORC2 function. This fact makes difficult the data interpretation regarding the role played by mTORC1 and mTORC2 in osteogenesis. Martin SK et al. demonstrated that using Cre-mediated gene deletion in well established in-vitro differentiation assays, have shown that mTORC1 and mTORC2 have distinct roles in MSCs fate determination. In agreement with previous studies,, blockade of Raptor in MSCs resultedin reduced adipogenic potential. Under osteoinductive conditions however, Raptor blockade promoted osteogenic differentiation. In current study we demonstrated that in osteogenic medium rapamicin is able to significantely reduce the mineralization of extracellular matrix. However, PEMF treatment is able to abolish this event, ensuring a good mineralization of extracellular environment. In light of these findings, we can assume that in presence of PEMF, the effect of rapamicin on osteoblasts behavior could be the opposit. Gene expression of 84 markers associated with mTOR pathway confirmed that no notable change in gene expression ocurred following rapamicin treatment coadministered with PEMF. While comparing gene expression under rapamicin treatment with PEMF to passive implant, reduction in mTOR2 pathway related genes was found. Namely, we found a reduction in Rictor expression that is associated to an adipogenic commitment of MSCs; and a decrease in several markers associated to a negative regulation of mTOR in both downstream and upstream levels. The most important changes are related to PKC? that, as we have previously demonstrated is strongly related to the adipogenic commitment of MSCs. We showed that PKC? recruits the 66-kD proapoptotic isoform of Shc (p66Shc) to act as oxidoreductase within mitochondria and in triggering a feed-forward cycle of ROS production, eventually leading to cell death. The same players may come together in a radically different context, i.e., the production of cellular signals linking hyperglicemia to the regulation of a transdifferentiation scheme of stem cells residing in adipose tissues. Moreover a downregulation of genes related to adipofunction such as PRKAG3 involved on insulin signalling is well evident. Finally, a significant increase in VEGF gene was demonstrated. These data confirm the ability of PEMF to promote angiogenesis, that is cruicial during tissue regeneration as we have previously demonstrated in wound healing processes,.

The differentiations of MSCs into the osteoblastic or adipogenic lineages are inter-dependent process: molecular components promoting one cell fate inhibit the mechanisms leading the differentiation of the alternative lineage. Interestingly, inducers of differentiation along one lineage often inhibit differentiation along the other. Our results suggest that in presence of osteogenic medium, PEMFs are able to induce osteogenic commitment of MSCs blocking the pathway of adipogenesis via mTOR related proteins.

This study reaults are in a line and comparable with a several previusly published papers. Ardeshirylajimi et al. investigated the the influence of prolonged pulsed extremely low frequency electromagnetic field on the osteogenic potential of cultured induced pluripotent stem cells. They concluded that combination of osteogenic medium and pulsed extremely low frequency electromagnetic field can be a great enhancement for bone differentiation of stem cells and appropriate candidate the management of bone defects and patients suffering from osteoporosis. A recently published paper by Arjmand et al. investigated the osteoinductive potential of PEMF in combination with Poly(caprolactone) (PCL) nanofibrous scaffold. Their results confirmed that the effects of PEMF on the osteogenic differentiation of ADSCs are very similar to these of osteogenic medium. They concluded that due to the immunological concerns regarding the application of bioactive molecules for tissue engineering, PEMF could be a good alternative for osteogenic medium. Additional recent article by Ardeshirylajimi et al. demonstrated that PEMF alone can induce osteogenic differentiation, but this capability was significantly increased when used in combination with electrospun polycaprolactone nanofibers. In addition, simultaneous use of osteogenic medium, PEMF and electrospun nanofibers resulted in increased osteogenic differentiation potential of induced pluripotent stem cells.

This study has several limitations, including its in vitro nature. Furthermore, the cells were grown in a monolayer, which does not accurately reflect in vivo conditions. The primary human cell cultures, however, can serve as a relevant model for examining the effects of PEMFs on bone cell physiology. The modulation of bone cell proliferation markers observed in this study have implications with regard to the immediate effects of PEMFs on bone formation and healing, as well as possible long-term implications for PEMF treatment.

In summary, the findings of the present study revealed that MED-generated PEMFs stimulate osteogenic differentiation and the maturation of the adipose tissue-derived MSCs via activation of the mTOR pathways. We also demonstrated that PEMF exposure increased cell proliferation, adhesion and the osteogenic commitment of MSCs, even in inflammatory conditions. We showed that PEMFs increased the expression of anti-inflammatory cytokines, such as IL-10, and reduced the expression of the pro-inflammatory cytokine IL-1. MSCs provided not only cell sources for connective tissues, but also had a significant influence on the immune response. Further studies are required to investigate the precise mechanisms by which mTOR signaling pathways are influenced and to discover other potential pathways involved in the PEMF-induced osteogenic effects.

Methods

PEMF exposure

The miniaturized electromagnetic device (MED) (Magdent Ltd., Tel Aviv, Israel) was the generator used to stimulate the cells. In the clinical setting, MED technology is used to actively stimulate osteogenesis and osseointegration. The MED was used with a Classix Dental Implant (3.3?mm 10?mm?L Non Touch Prime, Cortex Ltd., Shlomi, Israel). The cells were irradiated continuously for 30 days with the MED inside the incubator and under the same conditions of temperature, humidity and CO2 concentration as non PEMF irradiated cells which served as the controls.

Cell culture

MSCs were extracted from human adipose tissues of 5 healthy women and 5 healthy men (age 21–36 years, body mass index 30–38) who were undergoing cosmetic surgery procedures, following the guidelines of the University of Padova’s Plastic Surgery Clinic. The adipose tissues were digested with 0.075% collagenase (type 1?A; Sigma Aldrich, Italia) in a modified Krebs-Ringer buffer [125?mM NaCl, 5?mM KCl, 1?mM Na3PO4, 1?mM MgSO4, 5.5?mM glucose, and 20?mM HEPES (pH 7.4)] for 60?min at 37?°C, followed by 10?min with 0.25% trypsin. Floating adipocytes were discarded, and cells from the stromal-vascular fraction were pelleted, rinsed with media, and centrifuged, after which a red cell lysis step in NH4Cl was run for 10?min at room temperature. The resulting viable cells were counted using the trypan blue exclusion assay and seeded at a density of 106 cells per cm² for in vitro expansion in Dulbecco’s modified Eagle’s medium (DMEM, SIGMA Aldrich Italia) supplemented with 10% fetal calf serum and 1% penicillin/streptomycin. For treatment in inflammatory conditions, the cells were treated for 24?h with 0.1?mg/mL?1 of tumor necrosis factor-alpha (Celbio). TNF-? concentration used in the study is higher than in physiologic conditions. However, the aforementioned concentration was chosen based on the previously published papers in order to achieve effects in in-vitro studies,.

DNA content

DNA content was determined using a DNeasy kit (Qiagen, Hilden, Germany) to isolate total DNA from cell cultures following the manufacturer’s protocol for tissue isolation, using overnight incubation in proteinase K (Qiagen). DNA concentration was detected by measuring the absorbance at 260?nm in a spectrophotometer. The cell number was then determined from a standard curve (microgram DNA vs. cell number) generated by DNA extraction from the counted cells. The standard curve was linear over the tested range of 5–80?µg DNA (r?=?0.99).

MTT assay

To determine the proliferation rate of cell growth on titanium disks with or without treatment, a methyl thiazolyl-tetrazolium (MTT)-based cytotoxicity assay was performed according to the method of Denizot and Lang with minor modifications. The test is based on mitochondria viability, i.e., only functional mitochondria can oxidize an MTT solution, giving a typical blue-violet endproduct. After harvesting the culture medium, the cells were incubated for 3?h at 37?°C in 1?mL 0.5?mg/mL MTT solution prepared in phosphate buffered saline (PBS) solution. After removal of the MTT solution by pipette, 0.5?mL 10% dimethyl sulfoxide in isopropanol (iDMSO) was added for 30?min at 37?°C. For each sample, absorbance values at 570?nm were recorded in duplicate on 200??L aliquots deposited in 96-well plates using a multilabel plate reader (Victor 3 Perkin Elmer, Milano, Italy). All samples were examined after 15 and 30 days of culture.

RNA extraction and first-strand cDNA synthesis

RNase-Free DNase Set (Qiagen) from implants were cultured with adipose tissue derived mesenchymal stem cells for 15 and 25 days. The RNA quality and concentration of the samples were measured using a NanoDropTM ND-1000 Spectrophotometer (Thermo Scientific). For the first-strand cDNA synthesis, 200?ng of total RNA of each sample was reverse transcribed with M-MLV Reverse Transcriptase (Invitrogen), following the manufacturer’s protocol.

Real-time PCR

Human primers were selected for each target gene with Primer 3 software (Table 1). Real-time PCRs were carried out using the designed primers at a concentration of 300?nM and FastStart SYBR Green Master (Roche) on a Rotor-Gene 3000 (Corbett Research, Sydney, Australia). Real-time PCR was performed also according to the user’s manual for the Human mTOR signaling Profiler PCR Array (SABiosciences, Frederick, MD, USA) and using RT2 SYBR Green ROX FAST Master Mix (Qiagen). The data were analyzed using Excel-based PCR Array Data Analysis Templates (SABiosciences). The thermal cycling conditions were as follows: 15?min denaturation at 95?°C, followed by 40 cycles of 15?s denaturation at 95?°C, annealing for 30?s at 60?°C, and 20?s elongation at 72?°C. Differences in gene expression were evaluated by the 2??Ct method, using MSCs cultured in the presence and absence of inflammatory cytokines and in the presence and absence of PEMFs. Values were normalized to the expression of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) internal reference whose abundance did not change under our experimental conditions. Experiments were performed with 3 different cell preparations and repeated at least 3 times.

Table 1

List of gene related to mTOR pathway analized by RT PCR.

Description Gene
mTOR1 Complexes: MTOR associated protein, LST8 homolog (S. cerevisiae) MLST8
Mechanistic target of rapamycin (serine/threonine kinase) MTOR
Regulatory associated protein of MTOR, complex 1 RPTOR
mTOR2 Complexes: Mitogen-activated protein kinase associated protein 1 MAPKAP1
RPTOR independent companion of MTOR, complex 2 RICTOR
mTOR Upstream Regulators negative regulation: Eukaryotic translation initiation factor 4E binding protein 1 EIF4EBP1
Eukaryotic translation initiation factor 4E binding protein 2 EIF4EBP2
Protein phosphatase 2, catalytic subunit, alpha isozyme PPP2CA
Protein phosphatase 2, regulatory subunit B, beta PPP2R2B
Protein phosphatase 2?A activator, regulatory subunit 4 PPP2R4
Tumor protein p53 TP53
Unc-51-like kinase 1 (C. elegans) ULK1
Unc-51-like kinase 2 (C. elegans) ULK2
mTOR Upstream Regulators positive regulation: Cell division cycle 42 (GTP binding protein, 25?kDa) CDC42
Conserved helix-loop-helix ubiquitous kinase CHUK
Eukaryotic translation initiation factor 4B EIF4B
Eukaryotic translation initiation factor 4E EIF4E
Glycogen synthase kinase 3 beta GSK3B
Hypoxia inducible factor 1, alpha subunit (basic helix-loop-helix transcription factor) HIF1A
Heat shock 70?kDa protein 4 HSPA4
Integrin-linked kinase ILK
Myosin IC MYO1C
Protein kinase C, alpha PRKCA
Protein kinase C, beta PRKCB
Protein kinase C, epsilon PRKCE
Protein kinase C, gamma PRKCG
Ras homolog gene family, member A RHOA
Ribosomal protein S6 RPS6
Ribosomal protein S6 kinase, 70?kDa, polypeptide 1 RPS6KB1
Ribosomal protein S6 kinase, 70?kDa, polypeptide 2 RPS6KB2
Serum/glucocorticoid regulated kinase 1 SGK1
Vascular endothelial growth factor A VEGFA
Vascular endothelial growth factor B VEGFB
Vascular endothelial growth factor C VEGFC
mTOR Downstream Effectors negative regulation: AKT1 substrate 1 (proline-rich) AKT1S1
Calcium binding protein 39 CAB39
Calcium binding protein 39?L CAB39L
DNA-damage-inducible transcript 4 DDIT4
DNA-damage-inducible transcript 4-like DDIT4L
DEP domain containing MTOR-interacting protein DEPTOR
FK506 binding protein 1?A, 12?kDa FKBP1A
FK506 binding protein 8, 38?kDa FKBP8
Insulin-like growth factor binding protein 3 IGFBP3
Protein kinase, AMP-activated, alpha 1 catalytic subunit PRKAA1
Protein kinase, AMP-activated, alpha 2 catalytic subunit PRKAA2
Protein kinase, AMP-activated, beta 1 non-catalytic subunit PRKAB1
Protein kinase, AMP-activated, beta 2 non-catalytic subunit PRKAB2
Protein kinase, AMP-activated, gamma 1 non-catalytic subunit PRKAG1
Protein kinase, AMP-activated, gamma 2 non-catalytic subunit PRKAG2
Protein kinase, AMP-activated, gamma 3 non-catalytic subunit PRKAG3
Phosphatase and tensin homolog PTEN
Serine/threonine kinase 11 STK11
STE20-related kinase adaptor beta STRADB
Tuberous sclerosis 1 TSC1
Tuberous sclerosis 2 TSC2
Tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein, theta polypeptide YWHAQ
mTOR Downstream Effectors positive regulation: V-akt murine thymoma viral oncogene homolog 1 AKT1
V-akt murine thymoma viral oncogene homolog 2 AKT2
V-akt murine thymoma viral oncogene homolog 3 (protein kinase B, gamma) AKT3
V-Ha-ras Harvey rat sarcoma viral oncogene homolog HRAS
Insulin-like growth factor 1 (somatomedin C) IGF1
Inhibitor of kappa light polypeptide gene enhancer in B-cells, kinase beta IKBKB
Insulin INS
Insulin receptor INSR
Insulin receptor substrate 1 IRS1
Mitogen-activated protein kinase 1 MAPK1
Mitogen-activated protein kinase 3 MAPK3
3-phosphoinositide dependent protein kinase-1 PDPK1
Phosphoinositide-3-kinase, class 3 PIK3C3
Phosphoinositide-3-kinase, catalytic, alpha polypeptide PIK3CA
Phosphoinositide-3-kinase, catalytic, beta polypeptide PIK3CB
Phosphoinositide-3-kinase, catalytic, delta polypeptide PIK3CD
Phosphoinositide-3-kinase, catalytic, gamma polypeptide PIK3CG
Phospholipase D1, phosphatidylcholine-specific PLD1
Phospholipase D2 PLD2
Ras homolog enriched in brain RHEB
Ribosomal protein S6 kinase, 90?kDa, polypeptide 1 RPS6KA1
Ribosomal protein S6 kinase, 90?kDa, polypeptide 2 RPS6KA2
Ribosomal protein S6 kinase, 90?kDa, polypeptide 5 RPS6KA5
Ras-related GTP binding A RRAGA
Ras-related GTP binding B RRAGB
Ras-related GTP binding C RRAGC
Ras-related GTP binding D RRAGD
TEL2, telomere maintenance 2, homolog (S. cerevisiae) TELO2

Real-time PCR – mTOR

Total RNA was extracted using an RNeasy Lipid Tissue kit (Qiagen), including DNase digestion with the RNase-Free DNase. Set (Qiagen), from the mTOR signalling RT2 profiler PCR Array (gene analized are reported on Table 1). In total, 800?ng of RNA was reverse-transcribed using an RT2 First Strand kit (Qiagen). Real-time PCR was performed according to the user’s manual for the Human mTOR signalling RT2 profiler PCR Array (SABiosciences, Frederick, MD, USA) and using RT2 SYBR Green ROX FAST Master Mix (Qiagen). Thermal cycling and fluorescence detection were performed using a Rotor-Gene Q 100 (Qiagen). The data were analyzed using Excel-based PCR Array Data Analysis Templates (SABiosciences).

Alizarin Red S staining

The extracellular mineral deposits were detected by Alizarin Red S staining. Cells were fixed in 4% paraformaldehyde (Sigma-Aldrich) in PBS for 10?min at room temperature. Cells were stained adding 40?mM freshly Alizarin Red S Solution (pH 4.2) for 10?min at room temperature with gentle shaking. Cells were washed with ddH2O, then photographed by an optical microscope. Alizarin Red S stained area were quantified from microscope images of three independent experiments using ImageJ software (NIH, Bethesda, MD, USA).

ALP activity measurements

Alkaline phosphatase (ALP) activity was measured for up to 20 days of cell culture in order to evaluate the initial differentiation of Adipose Tissue Derived Mesenchymal Stem cells into preosteoblasts. Abcam’s alkaline phosphates kit (colorimetric) was used to detect the intracellular and extracellular ALP activities. The kit uses p-nitrophenyl phosphate (pNPP) as a phosphatase substrate, which is adsorbed at 405?nm when dephosphorylated by ALP. In accordance with the manufacturer’s protocol, the culture medium from each sample group was collected and pooled. At the same time, the cells were washed with PBS and then homogenized with ALP assay buffer (a total of 300??L for each group) and centrifuged at 13,000?rpm for 3?min to remove insoluble material. Different volumes of samples (medium and cells) were then added into 96-well plates, bringing the total volume in each well up to 80??L with assay buffer. In addition, 80??L fresh medium was utilized as sample background control. Thereafter, 50??L 5mMpNPP solution was added to each well containing test samples and background control and incubated for 60?min at 25?°C while shielding the plate from light. A standard curve of 0, 4, 6, 12, 16, and 20?nmol/well was generated from 1?mM pNPP standard solution, bringing the final volume to 120??L with assay buffer. All reactions were then stopped by adding 20??L of stop solution into each standard and sample reaction, except the sample background control reaction. Optical density was read at 405?nm in a microplate reader (Victor). The results were normalized by subtracting the value derived from the zero standards from all standards, samples and sample background control. The pNP standard curve was plotted to identify the pNP concentration in each sample. ALP activity of the test samples was calculated as follows:

ALP activity (U/ml) = A/V/T

where: A is the amount of pNP generated by samples (in ?mol), V is the amount of sample added in the assay well (in mL), and T is the reaction time (in minutes).

Immunofluorescence

Cells were fixed in 4% paraformaldehyde in PBS for 10?min and then incubated in 2% bovine serum albumin (BSA, Sigma-Aldrich) in PBS for 30?min at room temperature. They were then incubated with primary antibodies in 2% BSA solution in a humidified chamber for 12?h at 4?°C. The rabbit polyclonal antihuman phalloidine antibody (Millipore Corporation, MA, USA) was the primary antibody. Immunofluorescence staining was performed using the secondary antibody DyLight 549-labeled anti-rabbit IgG (H?+?L) (KPL, Gaithersburg, MD, USA) diluted 1/1000 in 2% BSA for 1?h at room temperature. Nuclear staining was performed with 2??g/mL Hoechst H33342 (Sigma-Aldrich) solution for 2?min.

Statistical analysis

One-way analysis of variance (ANOVA) was used for data analyses. Levene’s test was used to demonstrate the equal variances of the variables. Repeated measures ANOVA with a post-hoc analysis using Bonferroni’s multiple comparison was performed. T-tests were used to determine significant differences (p?<?0.05). Repeatability was calculated as the standard deviation of the difference between measurements. All testing was performed using SPSS 16.0 software (SPSS Inc., Chicago, IL, USA) (license of the University of Padua, Italy).

Acknowledgements

Research support (including PEMF generting devices and partial support in lab supplies) was provided by Mgdent Ltd.

Author Contributions

B.Z., A.P. and H.A.B. conceived and designed the experiments; L.F., O.D. and C.G. performed the experiments; B.Z., S.B. and C.G. analyzed the data; B.Z. and H.A.B. contributed reagents/materials/analysis tools; B.Z., L.F., M.S. and O.D. wrote the paper.

Notes

Competing Interests

O.D.- payed consultant of the Magdent ltd. Company. S.B.- Co-founder of Magdent ltd.

Footnotes

Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

1. Fu YC, et al. A novel single pulsed electromagnetic field stimulates osteogenesis of bone marrow mesenchymal stem cells and bone repair. PLoS One. 2014;9:e91581. doi: 10.1371/journal.pone.0091581.[PMC free article] [PubMed] [Cross Ref]
2. Petecchia L, et al. Electro-magnetic field promotes osteogenic differentiation of BM-hMSCs through a selective action on Ca(2+)-related mechanisms. Sci Rep. 2015;5:13856. doi: 10.1038/srep13856.[PMC free article] [PubMed] [Cross Ref]
3. Song M, et al. The effect of electromagnetic fields on the proliferation and the osteogenic or adipogenic differentiation of mesenchymal stem cells modulated by dexamethasone. Bioelectromagnetics. 2014;35:479–490. doi: 10.1002/bem.21867. [PubMed] [Cross Ref]
4. Yong Y, Ming ZD, Feng L, Chun ZW, Hua W. Electromagnetic fields promote osteogenesis of rat mesenchymal stem cells through the PKA and ERK1/2 pathways. J Tissue Eng Regen Med. 2016;10:E537–E545. doi: 10.1002/term.1864. [PubMed] [Cross Ref]
5. Ongaro A, et al. Pulsed electromagnetic fields stimulate osteogenic differentiation in human bone marrow and adipose tissue derived mesenchymal stem cells. Bioelectromagnetics. 2014;35:426–436. doi: 10.1002/bem.21862. [PubMed] [Cross Ref]
6. Kim MO, Jung H, Kim SC, Park JK, Seo YK. Electromagnetic fields and nanomagnetic particles increase the osteogenic differentiation of human bone marrow-derived mesenchymal stem cells. Int J Mol Med. 2015;35:153–160. doi: 10.3892/ijmm.2014.1978. [PubMed] [Cross Ref]
7. Lin CC, Lin RW, Chang CW, Wang GJ, Lai KA. Single-pulsed electromagnetic field therapy increases osteogenic differentiation through Wnt signaling pathway and sclerostin downregulation. Bioelectromagnetics. 2015;36:494–505. doi: 10.1002/bem.21933. [PubMed] [Cross Ref]
8. Purdue PE, Koulouvaris P, Nestor BJ, Sculco TP. The central role of wear debris in periprosthetic osteolysis. HSS J. 2006;2:102–113. doi: 10.1007/s11420-006-9003-6. [PMC free article] [PubMed][Cross Ref]
9. Anderson JM, Rodriguez A, Chang DT. Foreign body reaction to biomaterials. Semin Immunol. 2008;20:86–100. doi: 10.1016/j.smim.2007.11.004. [PMC free article] [PubMed] [Cross Ref]
10. Branemark R, Branemark PI, Rydevik B, Myers RR. Osseointegration in skeletal reconstruction and rehabilitation: a review. J Rehabil Res Dev. 2001;38:175–181. [PubMed]
11. Trindade R, Albrektsson T, Tengvall P, Wennerberg A. Foreign Body Reaction to Biomaterials: On Mechanisms for Buildup and Breakdown of Osseointegration. Clin Implant Dent Relat Res. 2016;18:192–203. doi: 10.1111/cid.12274. [PubMed] [Cross Ref]
12. Sundfeldt M, Carlsson LV, Johansson CB, Thomsen P, Gretzer C. Aseptic loosening, not only a question of wear: a review of different theories. Acta Orthop. 2006;77:177–197. doi: 10.1080/17453670610045902. [PubMed] [Cross Ref]
13. Glantschnig H, Fisher JE, Wesolowski G, Rodan GA, Reszka AA. M-CSF, TNFalpha and RANK ligand promote osteoclast survival by signaling through mTOR/S6 kinase. Cell Death Differ. 2003;10:1165–1177. doi: 10.1038/sj.cdd.4401285. [PubMed] [Cross Ref]
14. Indo Y, et al. Metabolic regulation of osteoclast differentiation and function. J Bone Miner Res. 2013;28:2392–2399. doi: 10.1002/jbmr.1976. [PubMed] [Cross Ref]
15. Laplante M, Sabatini DM. Regulation of mTORC1 and its impact on gene expression at a glance. J Cell Sci. 2013;126:1713–1719. doi: 10.1242/jcs.125773. [PMC free article] [PubMed] [Cross Ref]
16. Klionsky DJ, et al. Guidelines for the use and interpretation of assays for monitoring autophagy. Autophagy. 2012;8:445–544. doi: 10.4161/auto.19496. [PMC free article] [PubMed] [Cross Ref]
17. Mizushima N, Komatsu M. Autophagy: renovation of cells and tissues. Cell. 2011;147:728–741. doi: 10.1016/j.cell.2011.10.026. [PubMed] [Cross Ref]
18. Gharibi B, Farzadi S, Ghuman M, Hughes FJ. Inhibition of Akt/mTOR attenuates age-related changes in mesenchymal stem cells. Stem Cells. 2014;32:2256–2266. doi: 10.1002/stem.1709. [PubMed][Cross Ref]
19. Barak S, et al. A new device for improving dental implants anchorage: a histological and micro-computed tomography study in the rabbit. Clin Oral Implants Res. 2016;27:935–942. doi: 10.1111/clr.12661. [PubMed] [Cross Ref]
20. Ferroni L, et al. Pulsed magnetic therapy increases osteogenic differentiation of mesenchymal stem cells only if they are pre-committed. Life Sci. 2016;152:44–51. doi: 10.1016/j.lfs.2016.03.020. [PubMed][Cross Ref]
21. Deshpande S, et al. Reconciling the effects of inflammatory cytokines on mesenchymal cell osteogenic differentiation. J Surg Res. 2013;185:278–285. doi: 10.1016/j.jss.2013.06.063. [PMC free article][PubMed] [Cross Ref]
22. Li JK, Lin JC, Liu HC, Chang WH. Cytokine release from osteoblasts in response to different intensities of pulsed electromagnetic field stimulation. Electromagn Biol Med. 2007;26:153–165. doi: 10.1080/15368370701572837. [PubMed] [Cross Ref]
23. Chang K, Chang WH, Wu ML, Shih C. Effects of different intensities of extremely low frequency pulsed electromagnetic fields on formation of osteoclast-like cells. Bioelectromagnetics. 2003;24:431–439. doi: 10.1002/bem.10118. [PubMed] [Cross Ref]
24. Xian L, et al. Matrix IGF-1 maintains bone mass by activation of mTOR in mesenchymal stem cells. Nat Med. 2012;18:1095–1101. doi: 10.1038/nm.2793. [PMC free article] [PubMed] [Cross Ref]
25. Sarbassov DD, et al. Prolonged rapamycin treatment inhibits mTORC2 assembly and Akt/PKB. Mol Cell. 2006;22:159–168. doi: 10.1016/j.molcel.2006.03.029. [PubMed] [Cross Ref]
26. Martin SK, et al. Brief report: the differential roles of mTORC1 and mTORC2 in mesenchymal stem cell differentiation. Stem Cells. 2015;33:1359–1365. doi: 10.1002/stem.1931. [PubMed] [Cross Ref]
27. Xiang X, Zhao J, Xu G, Li Y, Zhang W. mTOR and the differentiation of mesenchymal stem cells. Acta Biochim Biophys Sin (Shanghai) 2011;43:501–510. doi: 10.1093/abbs/gmr041. [PubMed] [Cross Ref]
28. Aguiari P, et al. High glucose induces adipogenic differentiation of muscle-derived stem cells. Proc Natl Acad Sci USA. 2008;105:1226–1231. doi: 10.1073/pnas.0711402105. [PMC free article] [PubMed][Cross Ref]
29. Pavan C, et al. Weight gain related to treatment with atypical antipsychotics is due to activation of PKC-beta. Pharmacogenomics J. 2010;10:408–417. doi: 10.1038/tpj.2009.67. [PubMed] [Cross Ref]
30. Pinton P, Pavan C, Zavan B. PKC-beta activation and pharmacologically induced weight gain during antipsychotic treatment. Pharmacogenomics. 2011;12:453–455. doi: 10.2217/pgs.11.25. [PubMed][Cross Ref]
31. Rimessi A, et al. Protein Kinase C beta: a New Target Therapy to Prevent the Long-Term Atypical Antipsychotic-Induced Weight Gain. Neuropsychopharmacology. 2017;42:1491–1501. doi: 10.1038/npp.2017.20. [PMC free article] [PubMed] [Cross Ref]
32. Ferroni L, et al. Treatment by Therapeutic Magnetic Resonance (TMR) increases fibroblastic activity and keratinocyte differentiation in an in vitro model of 3D artificial skin. J Tissue Eng Regen Med. 2017;11:1332–1342. doi: 10.1002/term.2031. [PubMed] [Cross Ref]
33. Ferroni L, et al. Treatment of diabetic foot ulcers with Therapeutic Magnetic Resonance (TMR(R)) improves the quality of granulation tissue. Eur J Histochem. 2017;61:2800. doi: 10.4081/ejh.2017.2800.[PMC free article] [PubMed] [Cross Ref]
34. Chen Q, et al. Fate decision of mesenchymal stem cells: adipocytes or osteoblasts? Cell Death Differ. 2016;23:1128–1139. doi: 10.1038/cdd.2015.168. [PMC free article] [PubMed] [Cross Ref]
35. Ardeshirylajimi A, Soleimani M. Enhanced growth and osteogenic differentiation of Induced Pluripotent Stem cells by Extremely Low-Frequency Electromagnetic Field. Cell Mol Biol (Noisy-le-grand) 2015;61:36–41. [PubMed]
36. Arjmand M, Ardeshirylajimi A, Maghsoudi H, Azadian E. Osteogenic differentiation potential of mesenchymal stem cells cultured on nanofibrous scaffold improved in the presence of pulsed electromagnetic field. J Cell Physiol. 2018;233:1061–1070. doi: 10.1002/jcp.25962. [PubMed] [Cross Ref]
37. Ardeshirylajimi A, Khojasteh A. Synergism of Electrospun Nanofibers and Pulsed Electromagnetic Field on Osteogenic Differentiation of Induced Pluripotent Stem Cells. ASAIO J. 2018;64:253–260. doi: 10.1097/MAT.0000000000000631. [PubMed] [Cross Ref]
38. Bonora M, et al. Tumor necrosis factor-alpha impairs oligodendroglial differentiation through a mitochondria-dependent process. Cell Death Differ. 2014;21:1198–1208. doi: 10.1038/cdd.2014.35.[PMC free article] [PubMed] [Cross Ref]
39. Brun P, et al. In vitro response of osteoarthritic chondrocytes and fibroblast-like synoviocytes to a 500-730 kDa hyaluronan amide derivative. J Biomed Mater Res B Appl Biomater. 2012;100:2073–2081. doi: 10.1002/jbm.b.32771. [PubMed] [Cross Ref]
40. Denizot F, Lang R. Rapid colorimetric assay for cell growth and survival. Modifications to the tetrazolium dye procedure giving improved sensitivity and reliability. J Immunol Methods. 1986;89:271–277. doi: 10.1016/0022-1759(86)90368-6. [PubMed] [Cross Ref]
Neural Regen Res. 2018 Jan;13(1):145-153. doi: 10.4103/1673-5374.224383.

Low-frequency pulsed electromagnetic field pretreated bone marrow-derived mesenchymal stem cells promote the regeneration of crush-injured rat mental nerve.

Seo N1, Lee SH2, Ju KW2, Woo J3, Kim B4, Kim S5, Jahng JW6, Lee JH7.

Author information

1
Department of Oral and Maxillofacial Surgery, Graduate School of Dentistry, Seoul National University; Dental Research Institute, Seoul National University, Seoul, South Korea.
2
Department of Oral and Maxillofacial Surgery, Seoul National University Dental Hospital; Dental Research Institute, Seoul National University, Seoul, South Korea.
3
Department of Oral and Maxillofacial Surgery, Seoul National University Dental Hospital, Seoul, South Korea.
4
Clinical Translational Research Center for Dental Science (CTRC), Seoul National University Dental Hospital, Seoul, South Korea.
5
Department of Oral and Maxillofacial Surgery, Graduate School of Dentistry, Seoul National University; Department of Oral and Maxillofacial Surgery, Seoul National University Dental Hospital, Seoul, South Korea.
6
Dental Research Institute, Seoul National University, Seoul, South Korea.
7
Department of Oral and Maxillofacial Surgery, Graduate School of Dentistry, Seoul National University; Department of Oral and Maxillofacial Surgery, Seoul National University Dental Hospital; Dental Research Institute, Seoul National University; Clinical Translational Research Center for Dental Science (CTRC), Seoul National University Dental Hospital, Seoul, South Korea.

Abstract

Bone marrow-derived mesenchymal stem cells (BMSCs) have been shown to promote the regeneration of injured peripheral nerves. Pulsed electromagnetic field (PEMF) reportedly promotes the proliferation and neuronal differentiation of BMSCs. Low-frequency PEMF can induce the neuronal differentiation of BMSCs in the absence of nerve growth factors. This study was designed to investigate the effects of low-frequency PEMF pretreatment on the proliferation and function of BMSCs and the effects of low-frequency PEMF pre-treated BMSCs on the regeneration of injured peripheral nerve using in vitro and in vivo experiments. In in vitro experiments, quantitative DNA analysis was performed to determine the proliferation of BMSCs, and reverse transcription-polymerase chain reaction was performed to detect S100 (Schwann cell marker), glial fibrillary acidic protein (astrocyte marker), and brain-derived neurotrophic factor and nerve growth factor (neurotrophic factors) mRNA expression. In the in vivo experiments, rat models of crush-injured mental nerve established using clamp method were randomly injected with low-frequency PEMF pretreated BMSCs, unpretreated BMSCs or PBS at the injury site (1 × 106 cells). DiI-labeled BMSCs injected at the injury site were counted under the fluorescence microscope to determine cell survival. One or two weeks after cell injection, functional recovery of the injured nerve was assessed using the sensory test with von Frey filaments. Two weeks after cell injection, axonal regeneration was evaluated using histomorphometric analysis and retrograde labeling of trigeminal ganglion neurons. In vitro experiment results revealed that low-frequency PEMF pretreated BMSCs proliferated faster and had greater mRNA expression of growth factors than unpretreated BMSCs. In vivo experiment results revealed that compared with injection of unpretreated BMSCs, injection of low-frequency PEMF pretreated BMSCs led to higher myelinated axon count and axon density and more DiI-labeled neurons in the trigeminal ganglia, contributing to rapider functional recovery of injured mental nerve. These findings suggest that low-frequency PEMF pretreatment is a promising approach to enhance the efficacy of cell therapy for peripheral nerve injury repair.

KEYWORDS:

crush-injured mental nerve; low-frequency pulsed electromagnetic field; mesenchymal stem cells; nerve regeneration; peripheral nerve injury

Logo of scirep

About Editorial Board For Authors Scientific Reports
Sci Rep. 2017; 7: 9421.
Published online 2017 Aug 25. doi:  10.1038/s41598-017-09892-w
PMCID: PMC5572790
PMID: 28842627

Enhancement of mesenchymal stem cell chondrogenesis with short-term low intensity pulsed electromagnetic fields

Dinesh Parate,1 Alfredo Franco-Obregón,corresponding author2,3 Jürg Fröhlich,2,4 Christian Beyer,4 Azlina A. Abbas,5 Tunku Kamarul,5James H. P. Hui,corresponding author1,6 and Zheng Yangcorresponding author1,6
1Department of Orthopaedic Surgery, Yong Loo Lin School of Medicine, National University of Singapore, NUHS Tower Block, Level 11, 1E Kent Ridge Road, Singapore, 119288 Singapore
2Department of Surgery, Yong Loo Lin School of Medicine, National University of Singapore, NUHS Tower Block, Level 8, IE Kent Ridge Road, Singapore, 119228 Singapore
3BioIonic Currents Electromagnetic Pulsing Systems Laboratory, BICEPS, National University of Singapore, MD6, 14 medical Drive, #14-01, Singapore, 117599 Singapore
4Institute for Electromagnetic Fields, Swiss Federal Institute of Technology (ETH), Rämistrasse 101, 8092 Zurich, Switzerland
5Tissue Engineering Group (TEG), National Orthopaedic Centre of Excellence for Research and Learning (NOCERAL), Department of Orthopaedic Surgery, Faculty of Medicine, University of Malaya, Pantai Valley, Kuala Lumpur, 50603 Malaysia
6Tissue Engineering Program, Life Sciences Institute, National University of Singapore, DSO (Kent Ridge) Building, #04-01, 27 Medical Drive, Singapore, 117510 Singapore
Alfredo Franco-Obregón, gs.ude.sun@farus.
Contributor Information.
corresponding authorCorresponding author.
Author information ? Article notes ? Copyright and License information ? Disclaimer
Received 2017 Apr 13; Accepted 2017 Jul 28.

Abstract

Pulse electromagnetic fields (PEMFs) have been shown to recruit calcium-signaling cascades common to chondrogenesis. Here we document the effects of specified PEMF parameters over mesenchymal stem cells (MSC) chondrogenic differentiation. MSCs undergoing chondrogenesis are preferentially responsive to an electromagnetic efficacy window defined by field amplitude, duration and frequency of exposure. Contrary to conventional practice of administering prolonged and repetitive exposures to PEMFs, optimal chondrogenic outcome is achieved in response to brief (10?minutes), low intensity (2?mT) exposure to 6?ms bursts of magnetic pulses, at 15?Hz, administered only once at the onset of chondrogenic induction. By contrast, repeated exposures diminished chondrogenic outcome and could be attributed to calcium entry after the initial induction. Transient receptor potential (TRP) channels appear to mediate these aspects of PEMF stimulation, serving as a conduit for extracellular calcium. Preventing calcium entry during the repeated PEMF exposure with the co-administration of EGTA or TRP channel antagonists precluded the inhibition of differentiation. This study highlights the intricacies of calcium homeostasis during early chondrogenesis and the constraints that are placed on PEMF-based therapeutic strategies aimed at promoting MSC chondrogenesis. The demonstrated efficacy of our optimized PEMF regimens has clear clinical implications for future regenerative strategies for cartilage.

Introduction

Articular cartilage is an avascular tissue with low potential for self-repair. When left untreated, lesions of the articular cartilage can lead to osteoarthritis. The success of any technology aimed at repairing chondral defects will thus be based on its ability to produce tissues that most closely recapitulate the mechanical and biochemical properties of native cartilage. To this end many technologies have been advanced yet, none are without drawbacks. The ‘microfracture’ technique is commonly plagued by the formation of fibro-cartilaginous tissue of low dexterity. Autologous chondrocytes implantation and osteochondral autograft transplantation are limited by scarce cartilage production, low proliferative capacity of chondrocytes, chondrocyte de-differentiation and complications due to donor site morbidity. Stem cell-based approaches are also being actively pursued in hopes of improved outcome. Mesenchymal stem cells (MSCs) support chondrogenic differentiation and are an attractive cell source for cartilage tissue engineering. However, the neocartilage formed by conventional MSC-based repair methodologies commonly contain a mixture of fibro- and hyaline cartilage that do not achieve the biochemical, mechanical or functional properties of native cartilage.

MSCs can be differentiated along different cell lineages of mesodermal origin including osteoblasts, chondrocytes, skeletal myocytes or visceral stromal cells. Chondrogenic induction of MSCs entails proliferation, condensation, differentiation and maturation, necessitating endogenous transcriptional and developmental regulators, cell-cell and cell-matrix interactions that, in turn, are modulated by environmental stimuli including mechanical forces, temperature and oxygen levels. A common objective is to recreate as closely as possible the in vivo environmental conditions in vitro so that the rate and quality of chondrogenic development is enhanced and the functionality of the repaired tissue improved. To this end, various environmental stimuli such as hypoxia, mechanical, electric and electromagnetic stimulation are currently being explored.

Mechanical stimulation can be applied in a semi-controlled manner with the use of bioreactors designed to impart shear, compression, tension, or pressure on developing tissues. Appropriately applied mechanical stimulation positively influences MSC-induced chondrogenic differentiation, ECM deposition and the mechanical properties of the generated cartilage. At the cellular level the transduction of mechanical signals (mechanotransduction) involves their conversion into biochemical responses, often with the assistance of mechanosensitive calcium channels. Electromagnetic field (EMF)-stimulation has been shown to promote cell differentiation via the modulation of extracellular calcium entry via plasma membrane-embedded cation channels, raising the intriguing possibility that EMFs may be recruiting related pathways.

Studies examining time-variant or pulsing electromagnetic fields (PEMFs) have alluded to a benefit over articular chondrocytes or cartilaginous tissue in vitro, particularly with reference to chondrocyte proliferation, extracellular matrix (ECM) deposition, secretory activity and inflammatory status. Studies have also examined the effects of PEMF-treatment over the chondrogenic differentiation of stem cells derived from bone marrow, adipose, umbilical cord Wharton jelly, synovial fluid or peripheral blood sources. The reported consequences of PEMF-stimulation over chondrogenesis, however, are largely inconsistent. Some studies report modest enhancements in the gene expression of Sox9, aggrecan, type II collagen (Col 2) as well as deposition of sulfated glycosaminoglycan (sGAG), typically on the order of 2-folds, whereas other studies show little to no effect. On the extreme end of the spectrum, Wang et al. reported inhibition of both Sox9 and Col 2 expression concomitant with induction of hypertrophy and mineralization in response to exposures of 3?h per day at an amplitude of 1?mT. Obvious differences in stimulation protocols likely underlie reported discrepancies. Existing EMF studies have typically employed exposure durations between 30?minutes to 8?h per day and more consistently in the low milli Tesla amplitude range (3–5?mT). Empirical determination of the appropriate exposure and signal parameters for a specific biological response and given tissue are essential as there are indications that cell responses to magnetic fields obey an electromagnetic efficacy window defined by a specific combination of frequency, amplitude and time of exposure that gives rise to optimum cell response. Here, we systematically characterized the effects of PEMF exposure over MSC chondrogenic differentiation by varying the field amplitude, exposure duration and dosage with an emphasis on determining the briefest and lowest amplitude electromagnetic exposure to render a developmental outcome. Given that both mechanical stimuli and calcium entry influences chondrogenic differentiation, we investigated the ability of PEMF exposure to influence calcium homeostasis during early induction of MSCs into the chondrogenic lineage, in particular that attributed to the Transient Receptor Potential (TRP) family of cation-permeable channels, which has been broadly implicated in cellular mechanotransduction. We show that brief and single exposures to low amplitude PEMFs were most effective at stimulating MSC chondrogenesis. Our results also implicate the involvement of calcium influx and the mechanosensitive TRP channels, TRPC1 and TRPV4, in the chondrogenic development stimulated by targeted PEMF exposure.

Results

Effect of PEMF intensities and exposure durations on MSC chondrogenesis

We first sought to determine the magnetic field amplitude and duration of exposure at which MSCs undergoing chondrogenic induction are most responsive using starting conditions preliminarily tested in MSCs for chondrogenic regeneration. MSC pellets in chondrogenic differentiation medium were subjected to single exposures to PEMFs of 10?min duration at intensities ranging in amplitudes between 0–4?mT (Fig. 1A), then subjected to exposure durations between 5 and 60?min at 2?mT intensity (Fig. 1B), applied on the first day of chondrogenic induction. RNA analysis monitoring MSC chondrogenic progression at 7 days post-induction showed greatest increases in response to 10?min exposures applied at an amplitude of 2?mT as evidenced by enhancements in Sox9, aggrecan and Col 2 mRNA expression. By contrast, lower (1?mT) or higher (>3?mT) amplitude of PEMFs (Fig. 1A), or briefer (5?min) or longer (>20?min) durations of exposure (Fig. 1B), resulted in overall smaller effect sizes. The same EMF efficacy window translated to the expression of cartilaginous ECM macromolecular proteins (Fig. 1C). In response to 2?mT amplitude pulsing, a 3-fold increase in Col 2 protein was detected 21 days after chondrogenic induction, whereas no increase was detected with exposure to 3?mT. Moreover, a 2-fold increase in sGAG was detected in response to exposure to 2?mT PEMFs, whereas 3?mT PEMFs produced a significantly smaller increase. The relative ineffectiveness of prolonged exposure to PEMFs was also corroborated at the protein level. Sixty min exposures to 2?mT PEMFs did not elicit significant increases in Col 2 formation than 10?min exposures (Fig. 1C). With reference to sGAG production, PEMF amplitudes greater than 2?mT, or exposure durations of one hour, produced inferior results to 10?min exposures at 2?mT.

An external file that holds a picture, illustration, etc. Object name is 41598_2017_9892_Fig1_HTML.jpg

Effects of PEMF amplitude (A) and exposure duration (B) on MSC chondrogenesis. Real-time PCR analysis of cartilaginous markers expression after 7 days of differentiation was normalized to GAPDH and presented as fold-changes relative to levels in undifferentiated MSC. (C) Quantification of cartilaginous extracellular matrix macromolecules (Col 2 and sGAG) generated after 21 days of chondrogenic differentiation of MSC subjected to distinct PEMF parameters. All data shown are mean?±?SD, n?=?6 from 2 independent experiments. *Denotes significant increase, or decrease, compared to non-PEMF control. #Denotes significant decrease compared to 2?mT (A), or 10?min (B) PEMF exposure.

Dosage effects of PEMFs over MSC chondrogenesis

We next investigated the effect of repetitive exposures to PEMFs. MSCs were exposed to PEMFs at 2?mT for 10?min/day once, twice or thrice on days 1, 2 and 4 following chondrogenic induction (Fig. 2A). RNA analysis after 7 days of differentiation showed that a single exposure produced the greatest and most consistent increase in the expression of chondrogenic markers (Fig. 2A). In another series of experiments, MSCs pellets were exposed once on the first day of chondrogenic induction, or weekly for 3 consecutive weeks (Fig. 2B). RNA analysis after 21 days of chondrogenic differentiation showed that a single exposure to 2?mT PEMFs for 10?min given on day 1 of induction gave the greatest and most consistent increase in expression of chondrogenic markers relative to no exposure (0?mT) (Fig. 2B). Three weekly exposures either rendered no additional benefit (Col 2) or gave similar results to control (0?mT) (Sox9 and aggrecan). Moreover, the amount of ECM produced was inversely related to the total number of exposures. Single exposures produced >2-folds and ~1-fold increases in Col 2 and sGAG, respectively (Fig. 2C), whereas triple weekly exposures for three weeks (9 total exposures) completely precluded an increase Col 2 and sGAG formation. The change in the amount of DNA across samples varied less than 0.2-fold, although reaching significances at 2?mT, indicating that cell proliferation was only modestly affected within our pellet culture system (Fig. 2C).

An external file that holds a picture, illustration, etc. Object name is 41598_2017_9892_Fig2_HTML.jpg

Dosage effects of PEMFs over MSC chondrogenesis. (A) MSCs were exposed once (1x), twice (2x) or thrice (3x) per week. (B) MSCs were subjected to either a single exposure on day 1 of chondrogenic induction (1x) or once per week for 3 weeks (3x). Real-time PCR analysis of cartilaginous markers expression at 7 (A) or 21 days (B) after the induction of differentiation was normalized to GAPDH and presented as fold-changes relative to level in undifferentiated MSCs. (C) Quantification of cartilaginous ECM macromolecules generated during chondrogenic differentiation of MSCs in response to distinct PEMF dosing as indicated. MSC pellets were subjected to either a single PEMF exposure given on day 1 of chondrogenic induction (1x) once per week for 3 weeks (3x), or thrice weekly for 3 weeks (9x). Data represents the mean?±?SD, n?=?6 from 2 independent experiments. *Denotes significant increase compare to non-PEMF (0?mT) control. #Denotes significant decrease compared to single PEMF (1x) exposure.

Effect of PEMF treatment to deposition of ECM

ECM deposition in response to PEMF-exposure was also analyzed using Safranin O staining for proteoglycan and immunohistochemical staining for type II collagen (Fig. 3). Stained images of day 21 samples showed an enhanced deposition of proteoglycan and type II collagen in samples exposed only once to 2?mT for 10?min as compared to control (0?mT). By contrast, MSC samples exposed for longer (60?min), to greater amplitude (3?mT) or repeatedly (3x, 9x) yielded comparable, or inferior, ECM deposition to control.

An external file that holds a picture, illustration, etc. Object name is 41598_2017_9892_Fig3_HTML.jpg

Histological analysis of pellets exposed to PEMFs of distinct amplitude, duration and dosage. Pellets were harvested at day 21, sectioned and subjected to Safranin O or type II collagen immunohistochemistry staining. Images presented were represenation of n?=?3, taken at 100× magnification.

Ca2+ entry pathways implicated in transducing the effects of PEMFs over MSC differentiation

To investigate whether PEMF-stimulated MSC chondrogenesis was depended on calcium influx, EGTA (2?mM) was co-administered to the culture medium during PEMF exposure and summarily replaced afterwards with age-matched chondrogenic control media. RNA analysis at day 7 showed that the inclusion of EGTA significantly decreased the mRNA expression of Sox9, Col 2 and aggrecan in PEMF-treated samples (Fig. 4A), indicating that PEMF-exposure stimulates calcium influx. Conversely, transiently supplementing the differentiation medium with elevated extracellular Ca2+ (5?mM CaCl2) enhanced the mRNA expression of Sox9, Col 2 and aggrecan in otherwise non-exposed samples, and moreover, accentuated chondrogenic gene expression in PEMF-treated samples. These results corroborate that calcium influx is part of the upstream signalling cascade recruited by PEMFs contributing to chondrogenic induction.

An external file that holds a picture, illustration, etc. Object name is 41598_2017_9892_Fig4_HTML.jpg

Investigation of calcium entry pathways implicated in the PEMF-effect. (A) Involvement of Ca2+ influx in mediating the effects of PEMF-induced MSC chondrogenic differentiation. MSCs were exposed for 10?min at 2?mT alone (control, white bars), or in the presence of 2?mM EGTA (dark grey bars) or 5?mM CaCl2 (hatched bars) transiently added to the culture media. EGTA and CaCl2 were included to the bathing media 10?min before exposure and replaced with age-matched media control cultures 10?min after exposure. (B) Involvement of candidate calcium channels in mediating the effect of PEMs over MSC chondrogenic differentiation. Control MSC chondrogenic differentiation medium (white bars) was supplemented with Nifedipine (1?µM, light grey bars), Ruthenium Red (RR, 10?µM, black bars), or 2-APB (100?µM, dark grey bars) 10?min before exposure and replaced with age-matched media control cultures 10?min after exposure. Real-time PCR analysis was performed on day 7 of differentiation. Data represent the means?±?SD, n?=?6 from 2 independent experiments. *Denotes significant increase, or decrease, compared to non-PEMF (0?mT) control. #Denotes significant decrease relativeto 2?mT PEMF treatment.

To reveal the Ca2+ influx pathway recruited by PEMFs, we pharmacologically dissected the contribution of candidate channels utilizing 2-APB (100?µM) or Ruthenium Red (RR, 10?µM) as TRPC or TRPV cation channel antagonists, respectively, or Nifedipine (1?µM), as a dihydropyridine-sensitive, L-type voltage-gated calcium channel (VGCC) antagonist. Calcium channel antagonists were included into the differentiation medium 10?min before exposure to PEMFs and removed immediately afterwards with age-matched control chondrogenic media. Both 2-APB and Ruthenium Red completely inhibited the PEMF-triggered up-regulation of chondrogenic genes, whereas Nifedipine had no significant inhibitory effect (Fig. 4B). Chondrogenic inhibition by 2-APB and Ruthenium Red was also observed in non-exposed samples, indicating that TRPC- and TRPV-mediated calcium entry are similarly involved in constitutive chondrogenesis upon induction. By contrast, VGCC-mediated Ca2+ entry does not appear to play a predominant role in the early induction of chondrogenesis.

We next investigated the expression profiles of TRP channels (TRPC1, TRPC6, TRPV1, TRPV4, TRPV6) previously implicated in chondrogenesis and correlated these to our PEMF-induced chondrogenic responses. Amongst the panel of candidate TRP channels, the expression of TRPC1 and TRPV4 most closely correlated with our delineated magnetic efficacy window governing chondrogenesis with reference to PEMF amplitude, duration and dosage (Fig. 5 and Suppl. Figures 1 and 2). These results corroborate an involvement of TRPC1 and TRPV4 in the PEMF-induced enhancement of chondrogenic differentiation of MSC we observed.

An external file that holds a picture, illustration, etc. Object name is 41598_2017_9892_Fig5_HTML.jpg

Expression profiles of TRPC1 and TRPV4 in response to determined PEMF efficacy window regulating MSC chondrogenesis. Real-time PCR analysis of TRPC1 and V4 exposed to different (A) intensities, (B) durations, and (C) dosages of PEMFs. Data represent the means?±?SD, n?=?6 from 2 independent experiments. *Denotes significant increase, or decrease, compared to non-PEMF (0?mT) control. #Denotes significant decrease relative to 2 mT (A), 10?min (B), or single (1x, C) PEMF treatment.

Effect of recurring calcium influx on MSCchondrogenesis

We next investigated whether calcium entry, particularly that via TRPchannels underlies the inhibitory effect observed with repeated PEMF exposures. MSCs were exposed once, twice or thrice to 2?mT PEMFs for 10?min or, alternatively, exposed for 10?min to aged-matched control differentiation media containing elevated extracellular calcium (5?mM CaCl2) in lieu of PEMF exposure. RNA analysis at day 7 showed that MSCs treated once with PEMFs, or transiently administered elevated calcium, on day 1 exhibited enhanced chondrogenesis to comparable levels. By contrast, subsequent exposures to elevated calcium, on days 2 and 4 suppressed chondrogenesis mirrored the effect of multiple exposures to PEMFs (Fig. 6A and Suppl. Fig. 3A).

An external file that holds a picture, illustration, etc. Object name is 41598_2017_9892_Fig6_HTML.jpg

(A) MSC chondrogenic differentiation in response to multiple exposures to PEMFs or exogenously elevated calcium. MSCs were subjected to either PEMF stimulation alone (white bars) or with transient supplementation of CaCl2 alone (5?mM; hatched bars), once (1x), twice (2x) or thrice (3x) in a week. Dotted lines refers to expression level of non-treated controls. Real-time PCR analysis was performed on day 7 of chondrogenic differentiation. *Denotes significant increase relative to non-PEMF (0?mT) control. # and +denote significant differences relative to respective single (1x) exposure (white and hatched bars, respectively). P?=?PEMF treatment, Ca?=?CaCl2 supplementation. (B) MSC chondrogenic differentiation in response to multiple exposures to PEMFs alone (white bars) or in combination with calcium chelator (EGTA) or TRP channel antagonists. EGTA (2?mM; dark grey bars; “E”), Ruthenium Red (10?µM; RR, black bars; “R”) or 2-APB (100?µM; light grey bars; “C”) was added to the MSC differentiation medium during PEMF expoure applied once (1x), twice (2) or thrice (3x) per week. EGTA, RR and 2-APB were included 10?min before exposure and replaced with media harvested from age-matched chondrogenic control cultures 10?min after exposure. *Denotes significant increase compare to non-PEMF (0?mT) control. #Denotes significant decrease compared to single PEMF exposure (1x). +Denotes significant difference compared to respective PEMF control (white bar). P?=?PEMF treatment, E?=?EGTA, R?=?Ruthenium Red (RR), C?=?2-APB. Data shown are means?±?SD, n?=?6 from 2 independent experiments.

Analogously, precluding calcium entry (with EGTA) also exhibited dichotomous effects if applied during the first versus the second or third exposition to PEMFs, although in opposite direction to that observed with calcium administration or PEMFs. Whereas EGTA added during the initial exposure to PEMFs (1x) prevented PEMF-induced chondrogenesis, EGTA applied during the second or third exposure partially counteracted the inhibition of differentiation exerted by serial PEMF exposure (Fig. 6B and Suppl. Fig. 3B). Notably, impeding calcium entry with transient application of EGTA during both the second and third PEMF exposure was capable of almost completely reversing the inhibition of chondrogenesis observed with repeated PEMF exposures, suggesting that PEMFs are activating disparately functioning calcium mechanisms at early (day 1) and later stages (>day 2) of chondrogenic-induction that confer opposite effects over chondrogenesis. In contrast to the beneficial effect of calcium influx induced by PEMF at the initial stage of chondrogenesis, subsequent induction of calcium influx by repeated pulsing at later stages of chondrogenesis was suppressive of MSC chondrogenesis. The contribution of TRPC- and TRPV-mediated calcium entry to the chondrogenic-inhibition observed with repeated PEMF exposures was investigated by co-administering 2-APB (100?µM) or Ruthenium Red (RR, 10?µM), respectively, during PEMF exposure. As observed with transient EGTA application, antagonism of TRPC1/V4-mediated calcium entry during the first exposition to PEMFs was strongly inhibitory of PEMF-induced chondrogenesis, whereasTRPC1/V4 antagonism during subsequent PEMF expositions was somewhat less protective than EGTA over differentiation (Fig. 6B), implicating other yet to be determined calcium pathways in the later calcium-dependent inhibitory phase of chondrogenic progression. Notably, Ruthenium Red (TRPV4 antagonist) was capable of reverting the inhibition of differentiation and expression of TRPC1 expression in response to repeated PEMFing, whereas 2-APB (TRPC antagonist) was unable to revert the inhibition of differentiation and TRPV4 expression in response to repeated PEMFing, suggesting that TRPV4-mediated calcium entry antagonizes TRPC1 expression leading up to differentiation suppression. The dichotomous effects of precluding calcium entry by EGTA, Ruthenium Red or 2-APB when applied during the first, or the second and third, exposition to PEMFs was corroborated at the protein level. Preventing calcium entry during the initial exposure to PEMFs (1x) prevented PEMF-induced cartilaginous Col 2 and sGAG formation, while blocking calcium entry at later exposures counteracted the inhibition of differentiation exerted by serial PEMF exposure (Fig. 7). Voltage-gated L-type calcium channels, on the other hand, do not appear to be strongly implicated in the response as the expression of its subunits (CACNA1C and CACNA2D1) was not perturbed by PEMF or calcium treatment (Suppl. Fig. 3).

An external file that holds a picture, illustration, etc. Object name is 41598_2017_9892_Fig7_HTML.jpg

Quantification of cartilaginous ECM macromolecules generated by chondrogenically differentiated MSC in response to single or three weekly PEMF exposures alone (white bars) or in combination with calcium chelator (EGTA) or TRP channel antagonists as indicated. EGTA (2?mM; dark grey bars; “E”), Ruthenium Red (10?µM; RR, black bars; “R”) and 2-APB (100?µM; light grey bars; “C”) were included once during single PEMF exposures, or twice during the second and third PEMF exposure. EGTA, RR and 2-APB were added 10?min before exposure and replaced with media harvested from age-matched chondrogenic control cultures 10?min after exposure. *Denotes significant increase compare to non-PEMF (0?mT) control. #Denotes significant decrease compared to single PEMF exposure (1x). +Denotes significant difference compared to respective PEMF control (white bar). P?=?PEMF treatment, E?=?EGTA, R?=?Ruthenium Red (RR), C?=?2-APB. Data represent means?±?SD, n?=?3.

Discussion

Pulsed electromagnetic fields (PEMFs) have been demonstrated to be influential in numerous biological functions including progenitor cell fate determination and differentiation. PEMF-based therapies have been previously shown to enhance chondrocyte and cartilage explant anabolism while also limiting the catabolic consequences of inflammatory cytokines. PEMF exposure has been also reported to enhance the chondrogenic induction of stem cells. Nevertheless, inconsistent and conflicting results plague the scientific literature in this area of study, with PEMF exposures typically being applied on the order of hours per day for several days or weeks at a time. Here we report a high-efficacy of unprecedentedly brief (10?min applied once) PEMF exposure at inducing MSC chondrogenesis. We consistently detected increases in Sox9, Col 2 and aggrecan mRNA (>2-folds) in response to lone exposure to 2?mT PEMFs applied at the commencement of induction for only 10?min (Fig. 1). These increments in mRNA later translated into increased chondrogenic ECM protein formation (>2-fold) after 21 days of differentiation. By contrast, stimulation with greater amplitudes (>3?mT), longer exposures (>20?min) or more frequently (>2x/week) rendered no additional benefit, or was even less effective at promoting chondrogenesis at both the gene and protein levels. Although higher PEMF amplitudes and longer duration exposures were capable of augmenting aggrecan mRNA expression and macromolecular sGAG formation, the levels achieved were no better than those from samples treated only once with 2?mT PEMFs for 10?min. Col 2 expression was especially susceptible to overstimulation, being negatively impacted by exposures >2?mT or longer than 10?minutes. To the best of our knowledge, all published EMF studies examining chondrogenesis have employed exposure durations between 30?min to 8?h. For instance, Mayer-Wagner et al. using PEMF of 15?Hz, 5?mT, exposed MSCs undergoing chondrogenesis for 45?min every 8?h for a total of 21 days and observed less than a 2-fold increase in type II collagen expression, with no detected effect on Sox9 or aggrecan expression. Wang et al. using 1, 2, and 5?mT PEMFs at the frequency of 75?Hz exposed MSCs for 3?h per day for 4 weeks and instead observed a loss of cartilaginous phenotype associated with increased cartilage-specific extracellular matrix degradation in the later stage of chondrogenic differentiation.

Given that most conventional PEMF exposure paradigms employ a multiple exposure strategy and have reported positive chondrogenic outcome, we sought to determine the minimal number of exposures necessary to promote chondrogenesis (Fig. 2). We found that exposing MSCs once at the commencement of chondrogenic differentiation (1x) was necessary and sufficient to induce chondrogenic gene expression (Fig. 2A), which was sustainable for up to 21 days post chondrogenic-induction (Fig. 2B). The superior effect of a single pulse was also confirmed at the level of sGAG and Col 2 protein deposition (Figs 2C and ?and3).3). Indeed, in response to 3 exposures per week (10?min pulsing) for 3 consecutive weeks (9x treatments), ECM deposition was unchanged, or even inhibited, relative to unexposed samples. Ours is likely the first report to demonstrate an effectiveness of lone, 10?min, low amplitude PEMF exposures over MSC chondrogenesis, while concomitantly demonstrating the counter productivity of prolonged or repeated exposures. The possibility that prolonged or repeated PEMF exposures were merely cytotoxic, rather than truly inhibitory to chondrogenesis, was ruled out by our finding that total DNA content across all treatments was largely unchanged, despite lower Col 2 yield. In addition, the amount of sGAG was either unchanged or higher than that in control non-pulsed samples, further indicating that prolonged/repeated PEMF exposure did not adversely influence cell viability. Finally, the PEMF paradigm demonstrated here to best promote chondrogenesis (2?mT applied once for 10?min) did not alter the expression of osteogenic genes, Runx2 and ALP (Suppl. Fig. 4). Provocatively, osteogenic markers did increase following 20?min exposure to 2?mT PEMFs, thereby substantiating our assertion that reduced chondrogenic expression is not a reflection of cell death, but likely deferred chondrogenesis towards osteogenesis. Our demonstration of the high efficacy of brief and early PEMF exposure might thus help explain the existing inconsistencies and the relatively weaker responses previously reported.

Chondrogenesis is known to be modulated by calcium signaling cascades of specific temporal sensitivity,. The dependence of chondrogenesis on extracellular Ca2+ was first alluded to with the demonstration that elevated extracellular Ca2+ promoted chondrogenic differentiation in chick limb bud-derived cultures. Moreover, Sox9, the master transcription factor of chondrogenesis, is subject to Ca2+-calmodulin regulation. Elevation in cytoplasmic calcium downstream of calcium influx has been demonstrated in response to electric field (EF) or EMF stimulation during MSC-derived osteogenesis or chondrogenesis. We show that MSC chondrogenesis depends on the presence of extracellular Ca2+, whereby a transient (10?min) elevation of extracellular Ca2+ or brief (10?min) exposure to PEMFs (Figs 4Aand ?and6A)6A) enhanced MSC chondrogenic differentiation in an additive manner. Previous studies have also revealed that chondrogenesis is positively responsive to intracellular Ca2+ within a tightly controlled concentration window. A 1.25-fold increase in cytosolic Ca2+ concentration was shown to promote differentiation, whereas a moderately greater increase (1.5-fold) negatively influenced in vitrochondrogenesis. It is thus feasible that high amplitude or prolonged PEMF exposures elevate cytoplasmic calcium levels beyond the beneficial threshold for MSC chondrogenic differentiation. It is also well documented that the spatial and temporal patterns of intracellular free Ca2+ concentration play important roles in the regulation of various cellular processes, governed not only by absolute Ca2+ level, but also by periodic oscillatory changes of cytosolic Ca2+ concentration. MSCs undergoing chondrogenesis increase their frequency of Ca2+oscillations (waves) in the early stages of differentiation, coinciding with the initial period of cellular condensation during the first 2–4 days. Conversely, sustained elevations of extracellular calcium inhibit chondrogenesis, demonstrating a temporal requirement for calcium. Here we show that transient pulsing with elevated calcium recapitulates the temporal characteristic of the inhibitory actions of repeated PEMF exposures (Fig. 6A). Moreover, preventing calcium entry (with EGTA) during repeated PEMF exposure precludes the inhibition (Figs 6B and ?and7),7), defining a developmental change in calcium-sensitivity following calcium-dependent initiation of chondrogenesis. In this respect, single brief exposition to PEMFs defined by a specified electromagnetic window applied during the early stages of MSC chondrogenesis may be sufficient to provide the correct catalytic rise in intracellular Ca2+ to optimally promote the initiation of chondrogenesis. Conversely, higher exposure intensities or multiple exposures could result in excessive or sustained calcium influx that may instead disrupts or interrupts MSC-induced chondrogenesis, respectively.

Ca2+ influx via membrane-associated cation channels is a key event in initiating chondrogenesis, that can be potentially mediated by either TRP channels and/or voltage-gated calcium channels (VGCC). The transient receptor potential (TRP) channels are a diverse and widely distributed family of cation channel broadly implicated in cellular mechanotransduction. The TRPC and TRPV subfamilies have been broadly implicated in calcium homeostasis, ascribed mechanically-mediated gating, as well as implicated in the developmental programs of diverse mechanosensitive tissues. Previous studies have shown that blocking TRPV4 during the initial stages of induction inhibited chondrogenesis,. TRPV4-mediated Ca2+ signaling is also a positive regulator of Sox9 and as such, has been shown to promote chondrogenesis and in transducing the mechanical signals that support cartilage extracellular matrix maintenance and joint health. TRPC1 is expressed during early chondrocyte expansion, as well as being involved in the proliferation of mesenchymal stem cells. We detected time, intensity, and PEMF dosage-dependent up-regulations of both TRPV4 and TRPC1 that closely correlated with the PEMF-induced expression pattern of chondrogenic markers (Fig. 5). Blocking TRPC1 and TRPV4 channels with 2-APB and Ruthenium Red, respectively, in the early stage of differentiation effectively inhibited chondrogenesis, implicating these TRP channels in the initiation of chondrogenesis, and indicating that PEMFs recruit the activity of these channels to enhance chondrogenesis. Notably, blocking calcium-permeation through TRPV4 channels reverses the inhibition on chondrogenic differentiation and TRPC1 expression during repeated PEMF exposure, whereas blocking TRPC1 channels was unable to revert the inhibition on differentiation and expression of TRPV4 in response to repeated PEMF exposure, suggesting that TRPV4-mediated calcium entry antagonizes TRPC1 expression and is an essential step in initiating differentiation (Figs 6B and ?and7).7). TRPV4-mediated calcium entry may thus increase after the induction of differentiation (>2 days) serving to curtail TRPC1 expression and thereby promote differentiation by inhibiting TRPC1-medited proliferation (Suppl. Fig. 2).

An involvement of voltage-gated calcium channels was more difficult to establish. A predominant role for L-type VGCCs (CACNA1, CACNA2D1) in transducing PEMF’s effects was not supported given that a chondrogenically-effective dose of, Nifedipine, a L-type VGCC antagonist, had no significant effect on the PEMF-induced upregulation of MSC chondrogenesis (Fig. 4B). Moreover, the expression level of the L-type channel was not correlated with changes in calcium (Suppl. Fig. 3). Ca2+ influx via the low-threshold T-type VGCC had been previously implicated in tracheal chondrogenesis. The expression of T-type VGCC (CACNA1H) was induced by lone early exposure to PEMFs or transient calcium administration, and was suppressed by repeated exposures to PEMF or extracellular calcium, mirroring the expression pattern of chondrogenic markers under identical conditions (Suppl. Fig. 3A). The induction of the T-type calcium channel in response to PEMF/calcium exposure more likely reflects chondrogenic differentiation, rather than a fully determinant role in PEMF-induced chondrogenesis, as its expression was in the majority of conditions unchanged (relative to control) by removal of extracellular calcium during PEMF exposure (Suppl. Fig. 3B). Our strongest data support the interpretation that TRPC1 and TRPV4 play a more predominant role, although not necessarily exclusive, in transducing the chondrogenic effects of PEMFs. Further work will require to fully disentangle the intricasy of calcium homeostasis during the chondrogenic developmental process.

In summary, we have provided comprehensive characterization of the effects of PEMFs over MSC chondrogenic differentiation. MSCs undergoing chondrogenic induction are preferentially responsive to a well-defined window of PEMF stimulation of particular amplitude (2?mT), duration (10?min) and dosage (once on day 1 induction). By contrast, treatment with higher amplitude PEMFs, longer exposure durations or repeated expositions, as are more common in the field, are generally counterproductive, helping explain the lack of resolution in the field. Our results indicate that PEMFs mediate their effect by activating calcium influx through mechanosensitive calcium TRP channels. The unprecedented efficacy of our low amplitude, exceptionally brief and non-invasive PEMF-exposure protocol over MSC chondrogenesis has broad clinical and practical implications for the ultimate translation of related PEMF-based therapeutic strategies for stem cell-based cartilage regeneration.

Methods

Human bone marrow MSCs culture and chondrogenic differentiation

Primary human mesenchymal stem cells (MSCs) were purchased from RoosterBio Inc. (Frederick, MD), supplied at passage 3. The MSCs was further expanded in MSC High Performance Media (RoosterBio Inc.) at 37?°C in 5% CO2 atmosphere. The expanded MSCs were used at passage 5–6. Chondrogenic differentiation of MSCs was induced through 3D pellet culture as previously described. Briefly, 2.5?×?105 cells were centrifuged to form pellets and cultured in a chondrogenic differentiation medium containing high glucose DMEM supplemented with 4?mM proline, 50?µg/mL ascorbic acid, 1% ITS-Premix (Becton-Dickinson, San Jose, CA), 1?mM sodium pyruvate, and 10?7?M dexamethasone (Sigma-Aldrich, St Louis, MO), in the absence of antibiotics, for up to 7 or 21 days in the presence of 10 ng/mL of transforming growth factor-?3 (TGF?3; R&D Systems, Minneapolis, MN). To investigate an involvement of calcium influx or of calcium channels in transmitting the effects of PEMFs, cells were pre-incubated in chondrogenic media supplemented with elevated calcium, EGTA or particular calcium channel antagonist for 10?minutes prior to pulsing. Ten minutes after exposure to PEMFs the supplemented chondrogenic media was replaced with age-matched chondrogenic media (0?mT) cultures. To attenuate extracellular calcium influx, 2?mM ethylene-bis(oxyethylenenitrilo) tetraacetic acid (EGTA; Sigma) was added to the bathing media as noted. To promote extracellular Ca2+ influx the bathing media was supplemented with 5?mM CaCl2 (Sigma). To block calcium permeation through dihydropyridine-sensitive, L-type voltage-gated calcium channel (VGCC), Nifedipine (1?µM, Sigma) was added to the bathing media. 2-aminoethoxydiphenyl borate (2-APB, 100?µM, Sigma) and Ruthenium Red (10?µM, Merck Millipore) were administered as indicated to block calcium entry via TRPC and TRPV channels, respectively. Aminoglycoside antibiotics such as streptomycin were excluded in all MSC expansion and chondrogenic differentiation media to avoid interference with mechanosensitive ion channels.

PEMF Exposure system

The ELF-PEMF (extremely low frequency – pulsed magnetic field) delivery system has been described previously. For the purposes of this study a barrage of magnetic pulses of 6 ms duration was applied at a repetition rate of 15?Hz and at flux densities between 1–4?mT. Each 6 ms burst consisted of a series of 20 consecutive asymmetric pulses of 150?µs on and off duration with an approximate rise time of 17?T/s. The background magnetic flux density measured in the chamber was below 1?µT between 0?Hz to 5?kHz. The coil size, position and individual number of windings were numerically optimized by a CST low frequency solver for low field non-uniformity over a wide frequency range taking into consideration the shielding capacity of the µ-metallic chassis. The measured field non-uniformity did not exceed 4% within the uniform exposure region of the coils.

PEMF treatment

To investigate the optimum dosage of PEMF, MSCs in a 3D pellet culture were exposed to PEMFs of different exposure durations, dosage and the magnetic flux amplitude. MSCs were subjected to PEMFs of 1–4?mT amplitude with exposure times ranging between 5 to 60?min on the day of chondrogenic induction, applied once or multiple times as indicated in the respective figure legend. Cell pellets to be treated once with PEMFs (1x) were exposed on first day of chondrogenic induction. Two scenarios of multiple exposures were administrated (Fig. 2). Firstly, multiple exposures were administrated during the course of a week; double exposures (2x) were applied on days 1 and 2; triple exposures (3x) on days 1, 2 and 4. Alternatively, multiple exposures were applied on a once a week basis, for up to three week. Non-exposed (control) cells were placed within the PEMF device without current flux to produce a magnetic field to ensure that all cells were subject to the same climatic and mechanical conditions.

Real time PCR analysis

Chondrogenic cell pellets were digested in 0.25% Type II collagenase (Gibco, Life Technologies) followed by centrifugation. Total RNA was extracted using the RNeasy® Mini Kit (Qiagen, Germany). Reverse transcription was performed with 100 ng total RNA using iScript™ cDNA synthesis kit (Bio-Rad, USA). Real-time PCR was conducted using the SYBR®green assay on ABI 7500 Real-Time PCR System (Applied Biosystems, Life Technologies, USA). Real-time PCR program was set at 95?°C for 10?min, followed by 40 cycles of amplifications, consisting of a 15?s denaturation at 95?°C and a 1?min extension step at 60?°C. Primer sequences used in this study were according to previous publication and presented as Supplementary Table 1. The level of expression of the target gene, normalized to GAPDH, was then calculated using the 2???Ct formula with reference to the undifferentiated MSC. Results were averaged from triplicate samples of two independent experiments.

ECM and DNA quantification

Samples harvested were digested with 10?mg/mL of pepsin in 0.05?M acetic acid at 4?°C, followed by digestion with elastase (1?mg/mL). A Blyscan sulfated glycosaminoglycan (sGAG) assay kit (Biocolor Ltd., Newtownabbey, Ireland) was used to quantify sGAG deposition according to manufacturer’s protocol. Absorbance was measured at 656?nm and sGAG concentration was extrapolated from a standard curve generated using a sGAG standard. Type II Collagen (Col 2) content was measured using a captured enzyme-linked immunosorbent assay (Chondrex, Redmond, WA). Absorbance at 490?nm was measured and the concentration of Col 2 was extrapolated from a standard curve generated using a Col 2 standard. Values for sGAG and Col 2 content obtained were normalized to the total DNA content of respective samples, measured using Picogreen dsDNA assay (Molecular Probes, OR, USA). Quadruplicates of each group were analyzed from two independent experiments.

Histological and immunohistochemical evaluation

Samples were fixed in formalin, dehydrated, paraffin embedded, and cut into sections of 5?µm. For Safranin-O staining, the sections were incubated in hematoxylin (Sigma-Aldrich), washed and stained with fast green (Sigma-Aldrich), before staining with Safranin-O solution (AcrosOrganics). For immunohistochemistry, ultra-vision detection kit (Thermo scientific) was used. Endogenous peroxidase in the sections was first blocked with hydrogen peroxide before pepsin treatment for 20?min. Samples were treated with monoclonal antibodies of collagen type II (Clone 6B3; Chemicon Inc.) followed by incubation with biotinylated goat anti-mouse (Lab Vision Corporation). A mouse IgG isotype (Zymed Laboratories Inc.) was used as control for immunohistochemistry studies.

Statistical analysis

All experiments were performed in biological replicates (n?=?3 or 4) and results reported as mean?±?standard deviation (SD). Statistical analysis was carried out by Students t-test for comparison between two groups using the Microsoft Excel software. The level of significance was set at p?<?0.05. All quantitative data reported here were averaged from at least two independent experiments.

 

Electronic supplementary material

Acknowledgements

The study was supported by University of Malaya HIR-MoE Grant (Reference number – UM.C/625/1/HIR/MOHE/MED/32 account number – H20001-E000071) and Singapore-MIT Alliance for Research and Technology (SMART) Foundation (ING14085-BIO). Dinesh Parate was supported by NUS Research scholarship.

Author Contributions

D.P. performed experiments, analyzed data and drafted the manuscript. A.F.O., J.F. and C.B. provided technological expertise and contributed to the fabrication the PEMF facility. A.A.A., T.K., J.H.P.H. and A.F.O. provided funding and critical reading of the manuscript. A.F.O. and Z.Y. designed the study, analyzed data and provided critical revision of the manuscript. All authors reviewed the manuscript.

Notes

Competing Interests

The authors declare that they have no competing interests.

Footnotes

Electronic supplementary material

Supplementary information accompanies this paper at doi:10.1038/s41598-017-09892-w

Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Contributor Information

Alfredo Franco-Obregón, gs.ude.sun@farus.

James H. P. Hui, gs.ude.shun@iuh_semaj.

Zheng Yang, gs.ude.sun@zyisl.

References

1. Poole, A. R. et al. Composition and structure of articular cartilage: a template for tissue repair. Clin Orthop Relat Res S26–33 (2001). [PubMed]
2. Goldring MB, Goldring SR. Osteoarthritis. J Cell Physiol. 2007;213:626–34. doi: 10.1002/jcp.21258.[PubMed] [Cross Ref]
3. Ge Z, et al. Osteoarthritis and therapy. Arthritis Rheum. 2006;55:493–500. doi: 10.1002/art.21994.[PubMed] [Cross Ref]
4. Buckwalter JA, Mankin HJ. Articular cartilage: degeneration and osteoarthritis, repair, regeneration, and transplantation. Instr Course Lect. 1998;47:487–504. [PubMed]
5. Vasara, A.I. et al. Indentation stiffness of repair tissue after autologous chondrocyte transplantation. Clin Orthop Relat Res, 233–42 (2005). [PubMed]
6. Wakitani S, et al. Mesenchymal cell-based repair of large, full-thickness defects of articular cartilage. J Bone Joint Surg Am. 1994;76:579–92. doi: 10.2106/00004623-199404000-00013. [PubMed] [Cross Ref]
7. Wakitani S, et al. Human autologous culture expanded bone marrow mesenchymal cell transplantation for repair of cartilage defects in osteoarthritic knees. Osteoarthritis Cartilage. 2002;10:199–206. doi: 10.1053/joca.2001.0504. [PubMed] [Cross Ref]
8. Kuroda R, et al. Treatment of a full-thickness articular cartilage defect in the femoral condyle of an athlete with autologous bone-marrow stromal cells. Osteoarthritis Cartilage. 2007;15:226–31. doi: 10.1016/j.joca.2006.08.008. [PubMed] [Cross Ref]
9. Pittenger MF, et al. Multilineage potential of adult human mesenchymal stem cells. Science. 1999;284:143–7. doi: 10.1126/science.284.5411.143. [PubMed] [Cross Ref]
10. Goldring MB, Tsuchimochi K, Ijiri K. The control of chondrogenesis. J Cell Biochem. 2006;97:33–44. doi: 10.1002/jcb.20652. [PubMed] [Cross Ref]
11. Mahmoudifar N, Doran PM. Chondrogenesis and cartilage tissue engineering: the longer road to technology development. Trends Biotechnol. 2012;30:166–76. doi: 10.1016/j.tibtech.2011.09.002.[PubMed] [Cross Ref]
12. Woods A, Wang G, Beier F. Regulation of chondrocyte differentiation by the actin cytoskeleton and adhesive interactions. J Cell Physiol. 2007;213:1–8. doi: 10.1002/jcp.21110. [PubMed] [Cross Ref]
13. DeLise AM, Fischer L, Tuan RS. Cellular interactions and signaling in cartilage development. Osteoarthritis Cartilage. 2000;8:309–34. doi: 10.1053/joca.1999.0306. [PubMed] [Cross Ref]
14. Toh WS, Spector M, Lee EH, Cao T. Biomaterial-mediated delivery of microenvironmental cues for repair and regeneration of articular cartilage. Mol Pharm. 2011;8:994–1001. doi: 10.1021/mp100437a.[PubMed] [Cross Ref]
15. Klein TJ, Malda J, Sah RL, Hutmacher DW. Tissue engineering of articular cartilage with biomimetic zones. Tissue Eng Part B Rev. 2009;15:143–57. doi: 10.1089/ten.teb.2008.0563. [PMC free article][PubMed] [Cross Ref]
16. Raghothaman D, et al. Engineering cell matrix interactions in assembled polyelectrolyte fiber hydrogels for mesenchymal stem cell chondrogenesis. Biomaterials. 2014;35:2607–16. doi: 10.1016/j.biomaterials.2013.12.008. [PubMed] [Cross Ref]
17. Responte DJ, Lee JK, Hu JC, Athanasiou KA. Biomechanics-driven chondrogenesis: from embryo to adult. FASEB J. 2012;26:3614–24. doi: 10.1096/fj.12-207241. [PMC free article] [PubMed] [Cross Ref]
18. Mouw JK, Connelly JT, Wilson CG, Michael KE, Levenston ME. Dynamic compression regulates the expression and synthesis of chondrocyte-specific matrix molecules in bone marrow stromal cells. Stem Cells. 2007;25:655–63. doi: 10.1634/stemcells.2006-0435. [PubMed] [Cross Ref]
19. Grad S, Eglin D, Alini M, Stoddart MJ. Physical stimulation of chondrogenic cells in vitro: a review. Clin Orthop Relat Res. 2011;469:2764–72. doi: 10.1007/s11999-011-1819-9. [PMC free article] [PubMed][Cross Ref]
20. Cheing GL, Li X, Huang L, Kwan RL, Cheung KK. Pulsed electromagnetic fields (PEMF) promote early wound healing and myofibroblast proliferation in diabetic rats. Bioelectromagnetics. 2014;35:161–9. doi: 10.1002/bem.21832. [PubMed] [Cross Ref]
21. Steward AJ, Kelly DJ, Wagner DR. The role of calcium signalling in the chondrogenic response of mesenchymal stem cells to hydrostatic pressure. Eur Cell Mater. 2014;28:358–71. doi: 10.22203/eCM.v028a25. [PubMed] [Cross Ref]
22. Pingguan-Murphy B, El-Azzeh M, Bader DL, Knight MM. Cyclic compression of chondrocytes modulates a purinergic calcium signalling pathway in a strain rate- and frequency-dependent manner. J Cell Physiol. 2006;209:389–97. doi: 10.1002/jcp.20747. [PubMed] [Cross Ref]
23. Eijkelkamp N, Quick K, Wood JN. Transient receptor potential channels and mechanosensation. Annu Rev Neurosci. 2013;36:519–46. doi: 10.1146/annurev-neuro-062012-170412. [PubMed] [Cross Ref]
24. Matta C, Zakany R. Calcium signalling in chondrogenesis: implications for cartilage repair. Front Biosci (Schol Ed) 2013;5:305–24. doi: 10.2741/S374. [PubMed] [Cross Ref]
25. Pall ML. Electromagnetic fields act via activation of voltage-gated calcium channels to produce beneficial or adverse effects. J Cell Mol Med. 2013;17:958–65. doi: 10.1111/jcmm.12088.[PMC free article] [PubMed] [Cross Ref]
26. Ross CL, et al. The effect of low-frequency electromagnetic field on human bone marrow stem/progenitor cell differentiation. Stem Cell Res. 2015;15:96–108. doi: 10.1016/j.scr.2015.04.009.[PMC free article] [PubMed] [Cross Ref]
27. Petecchia L, et al. Electro-magnetic field promotes osteogenic differentiation of BM-hMSCs through a selective action on Ca(2+)-related mechanisms. Sci Rep. 2015;5 doi: 10.1038/srep13856.[PMC free article] [PubMed] [Cross Ref]
28. Schmidt-Rohlfing B, Silny J, Woodruff S, Gavenis K. Effects of pulsed and sinusoid electromagnetic fields on human chondrocytes cultivated in a collagen matrix. Rheumatol Int. 2008;28:971–7. doi: 10.1007/s00296-008-0565-0. [PubMed] [Cross Ref]
29. Vincenzi F, et al. Pulsed electromagnetic fields increased the anti-inflammatory effect of A(2)A and A(3) adenosine receptors in human T/C-28a2 chondrocytes and hFOB 1.19 osteoblasts. PLoS One. 2013;8doi: 10.1371/journal.pone.0065561. [PMC free article] [PubMed] [Cross Ref]
30. De Mattei M, et al. Effects of pulsed electromagnetic fields on human articular chondrocyte proliferation. Connect Tissue Res. 2001;42:269–79. doi: 10.3109/03008200109016841. [PubMed][Cross Ref]
31. Sakai A, Suzuki K, Nakamura T, Norimura T, Tsuchiya T. Effects of pulsing electromagnetic fields on cultured cartilage cells. Int Orthop. 1991;15:341–6. doi: 10.1007/BF00186874. [PubMed] [Cross Ref]
32. Hilz FM, et al. Influence of extremely low frequency, low energy electromagnetic fields and combined mechanical stimulation on chondrocytes in 3-D constructs for cartilage tissue engineering. Bioelectromagnetics. 2014;35:116–28. doi: 10.1002/bem.21822. [PubMed] [Cross Ref]
33. Boopalan PR, Arumugam S, Livingston A, Mohanty M, Chittaranjan S. Pulsed electromagnetic field therapy results in healing of full thickness articular cartilage defect. Int Orthop. 2011;35:143–8. doi: 10.1007/s00264-010-0994-8. [PMC free article] [PubMed] [Cross Ref]
34. Mueller MB, Tuan RS. Functional characterization of hypertrophy in chondrogenesis of human mesenchymal stem cells. Arthritis Rheum. 2008;58:1377–88. doi: 10.1002/art.23370. [PMC free article][PubMed] [Cross Ref]
35. Chen CH, et al. Electromagnetic fields enhance chondrogenesis of human adipose-derived stem cells in a chondrogenic microenvironment in vitroJ Appl Physiol (1985) 2013;114:647–55. doi: 10.1152/japplphysiol.01216.2012. [PubMed] [Cross Ref]
36. Lim, H. L. et al. Dynamic Electromechanical Hydrogel Matrices for Stem Cell Culture. Adv Funct Mater21 (2011). [PMC free article] [PubMed]
37. Wang J, et al. Pulsed electromagnetic field may accelerate in vitro endochondral ossification. Bioelectromagnetics. 2015;36:35–44. doi: 10.1002/bem.21882. [PubMed] [Cross Ref]
38. Mayer-Wagner S, et al. Effects of low frequency electromagnetic fields on the chondrogenic differentiation of human mesenchymal stem cells. Bioelectromagnetics. 2011;32:283–90. doi: 10.1002/bem.20633. [PubMed] [Cross Ref]
39. Esposito M, et al. Differentiation of human umbilical cord-derived mesenchymal stem cells, WJ-MSCs, into chondrogenic cells in the presence of pulsed electromagnetic fields. In Vivo. 2013;27:495–500.[PubMed]
40. Ongaro A, et al. Electromagnetic fields counteract IL-1beta activity during chondrogenesis of bovine mesenchymal stem cells. J Tissue Eng Regen Med. 2015;9:E229–38. doi: 10.1002/term.1671. [PubMed][Cross Ref]
41. Spaas JH, et al. Chondrogenic Priming at Reduced Cell Density Enhances Cartilage Adhesion of Equine Allogeneic MSCs – a Loading Sensitive Phenomenon in an Organ Culture Study with 180 Explants. Cell Physiol Biochem. 2015;37:651–65. doi: 10.1159/000430384. [PubMed] [Cross Ref]
42. Berg H. Problems of weak electromagnetic field effects in cell biology. Bioelectrochem Bioenerg. 1999;48:355–60. doi: 10.1016/S0302-4598(99)00012-4. [PubMed] [Cross Ref]
43. Crocetti S, et al. Low intensity and frequency pulsed electromagnetic fields selectively impair breast cancer cell viability. PLoS One. 2013;8 doi: 10.1371/journal.pone.0072944. [PMC free article] [PubMed][Cross Ref]
44. O’Conor CJ, Case N, Guilak F. Mechanical regulation of chondrogenesis. Stem Cell Res Ther. 2013;4doi: 10.1186/scrt211. [PMC free article] [PubMed] [Cross Ref]
45. Sun LY, et al. Effect of pulsed electromagnetic field on the proliferation and differentiation potential of human bone marrow mesenchymal stem cells. Bioelectromagnetics. 2009;30:251–60. doi: 10.1002/bem.20472. [PubMed] [Cross Ref]
46. Fodor J, et al. Store-operated calcium entry and calcium influx via voltage-operated calcium channels regulate intracellular calcium oscillations in chondrogenic cells. Cell Calcium. 2013;54:1–16. doi: 10.1016/j.ceca.2013.03.003. [PubMed] [Cross Ref]
47. San Antonio JD, Tuan RS. Chondrogenesis of limb bud mesenchyme in vitro: stimulation by cations. Dev Biol. 1986;115:313–24. doi: 10.1016/0012-1606(86)90252-6. [PubMed] [Cross Ref]
48. Matta C, et al. Cytosolic free Ca2+ concentration exhibits a characteristic temporal pattern during in vitro cartilage differentiation: a possible regulatory role of calcineurin in Ca-signalling of chondrogenic cells. Cell Calcium. 2008;44:310–23. doi: 10.1016/j.ceca.2007.12.010. [PubMed] [Cross Ref]
49. Argentaro A, et al. A SOX9 defect of calmodulin-dependent nuclear import in campomelic dysplasia/autosomal sex reversal. J Biol Chem. 2003;278:33839–47. doi: 10.1074/jbc.M302078200.[PubMed] [Cross Ref]
50. McCullen SD, et al. Application of low-frequency alternating current electric fields via interdigitated electrodes: effects on cellular viability, cytoplasmic calcium, and osteogenic differentiation of human adipose-derived stem cells. Tissue Eng Part C Methods. 2010;16:1377–86. doi: 10.1089/ten.tec.2009.0751.[PMC free article] [PubMed] [Cross Ref]
51. Xu J, Wang W, Clark CC, Brighton CT. Signal transduction in electrically stimulated articular chondrocytes involves translocation of extracellular calcium through voltage-gated channels. Osteoarthritis Cartilage. 2009;17:397–405. doi: 10.1016/j.joca.2008.07.001. [PubMed] [Cross Ref]
52. Kawano S, et al. Characterization of Ca(2+) signaling pathways in human mesenchymal stem cells. Cell Calcium. 2002;32:165–74. doi: 10.1016/S0143416002001240. [PubMed] [Cross Ref]
53. Curtis TM, Scholfield CN. Nifedipine blocks Ca2+ store refilling through a pathway not involving L-type Ca2+ channels in rabbit arteriolar smooth muscle. J Physiol. 2001;532:609–23. doi: 10.1111/j.1469-7793.2001.0609e.x. [PMC free article] [PubMed] [Cross Ref]
54. Wen L, et al. L-type calcium channels play a crucial role in the proliferation and osteogenic differentiation of bone marrow mesenchymal stem cells. Biochem Biophys Res Commun. 2012;424:439–45. doi: 10.1016/j.bbrc.2012.06.128. [PubMed] [Cross Ref]
55. Zhang J, Li M, Kang ET, Neoh KG. Electrical stimulation of adipose-derived mesenchymal stem cells in conductive scaffolds and the roles of voltage-gated ion channels. Acta Biomater. 2016;32:46–56. doi: 10.1016/j.actbio.2015.12.024. [PubMed] [Cross Ref]
56. Lin SS, et al. Cav3.2 T-type calcium channel is required for the NFAT-dependent Sox9 expression in tracheal cartilage. Proc Natl Acad Sci USA. 2014;111:E1990–8. doi: 10.1073/pnas.1323112111.[PMC free article] [PubMed] [Cross Ref]
57. Guilak F, Leddy HA, Liedtke W. Transient receptor potential vanilloid 4: The sixth sense of the musculoskeletal system? Ann N Y Acad Sci. 2010;1192:404–9. doi: 10.1111/j.1749-6632.2010.05389.x.[PMC free article] [PubMed] [Cross Ref]
58. Maroto R, et al. TRPC1 forms the stretch-activated cation channel in vertebrate cells. Nat Cell Biol. 2005;7:179–85. doi: 10.1038/ncb1218. [PubMed] [Cross Ref]
59. Eleswarapu SV, Athanasiou KA. TRPV4 channel activation improves the tensile properties of self-assembled articular cartilage constructs. Acta Biomater. 2013;9:5554–61. doi: 10.1016/j.actbio.2012.10.031. [PMC free article] [PubMed] [Cross Ref]
60. Phan MN, et al. Functional characterization of TRPV4 as an osmotically sensitive ion channel in porcine articular chondrocytes. Arthritis Rheum. 2009;60:3028–37. doi: 10.1002/art.24799.[PMC free article] [PubMed] [Cross Ref]
61. Hung CT. Transient receptor potential vanilloid 4 channel as an important modulator of chondrocyte mechanotransduction of osmotic loading. Arthritis Rheum. 2010;62:2850–1. doi: 10.1002/art.27617.[PubMed] [Cross Ref]
62. Kurth F, et al. Transient receptor potential vanilloid 2-mediated shear-stress responses in C2C12 myoblasts are regulated by serum and extracellular matrix. FASEB J. 2015;29:4726–37. doi: 10.1096/fj.15-275396. [PubMed] [Cross Ref]
63. Clapham DE, Runnels LW, Strubing C. The TRP ion channel family. Nat Rev Neurosci. 2001;2:387–96. doi: 10.1038/35077544. [PubMed] [Cross Ref]
64. Somogyi CS, et al. Polymodal Transient Receptor Potential Vanilloid (TRPV) Ion Channels in Chondrogenic Cells. Int J Mol Sci. 2015;16:18412–38. doi: 10.3390/ijms160818412. [PMC free article][PubMed] [Cross Ref]
65. Muramatsu S, et al. Functional gene screening system identified TRPV4 as a regulator of chondrogenic differentiation. J Biol Chem. 2007;282:32158–67. doi: 10.1074/jbc.M706158200. [PubMed] [Cross Ref]
66. Gavenis K, et al. Expression of ion channels of the TRP family in articular chondrocytes from osteoarthritic patients: changes between native and in vitro propagated chondrocytes. Mol Cell Biochem. 2009;321:135–43. doi: 10.1007/s11010-008-9927-x. [PubMed] [Cross Ref]
67. Torossian F, Bisson A, Vannier JP, Boyer O, Lamacz M. TRPC expression in mesenchymal stem cells. Cell Mol Biol Lett. 2010;15:600–10. doi: 10.2478/s11658-010-0031-3. [PubMed] [Cross Ref]
68. Lievremont JP, Bird GS, Putney JW., Jr. Mechanism of inhibition of TRPC cation channels by 2-aminoethoxydiphenylborane. Mol Pharmacol. 2005;68:758–62. [PubMed]
69. Yang Z, et al. Improved mesenchymal stem cells attachment and in vitro cartilage tissue formation on chitosan-modified poly(L-lactide-co-epsilon-caprolactone) scaffold. Tissue Eng Part A. 2012;18:242–51. doi: 10.1089/ten.tea.2011.0315. [PMC free article] [PubMed] [Cross Ref]
70. Winegar BD, Haws CM, Lansman JB. Subconductance block of single mechanosensitive ion channels in skeletal muscle fibers by aminoglycoside antibiotics. J Gen Physiol. 1996;107:433–43. doi: 10.1085/jgp.107.3.433. [PMC free article] [PubMed] [Cross Ref]
ASAIO J. 2018 Mar/Apr;64(2):253-260. doi: 10.1097/MAT.0000000000000631.

Synergism of Electrospun Nanofibers and Pulsed Electromagnetic Field on Osteogenic Differentiation of Induced Pluripotent Stem Cells.

Ardeshirylajimi A1, Khojasteh A.

Author information

1
From the Department of Tissue Engineering and Applied Cell Sciences, School of Advanced Technologies in Medicine, Shahid Beheshti University of Medical Sciences, Tehran, Iran.

Abstract

According to the current therapies failure for bone fractures and lesions, tissue engineering showed a great potential to help solve these challenges. Because the use of growth factors is very limited in the clinic, it could be very useful that could be introducing an alternative to it. Extremely low frequency pulsed electromagnetic fields (PEMF, 1 mT, 50 Hz) were used for achieving this aim. The PEMF potential in combination with electrospun polycaprolactone (PCL) nanofibers was used to investigate the osteogenic potential of human induced pluripotent stem cells (iPSCs). Several relevant osteogenic markers, such as Alizarin red staining, alkaline phosphatase activity, calcium content, gene expression, and immunocytochemistry, were used to evaluate osteoinductivity of PEMF. Results were shown that PEMF alone can induce osteogenic differentiation, but this capability increased when used in combination with PCL nanofibers significantly. In addition, simultaneous use of osteogenic medium, PEMF and PCL surprisingly increased osteogenic differentiation potential of iPSCs. According to the results, PEMF alone, iPSCs-seeded PCL, and both of them could be considered as a promising candidate for use in bone tissue engineering applications.

J Physiol Pharmacol. 2017 Apr;68(2):253-264.

Changes in viability of rat adipose-derived stem cells isolated from abdominal/perinuclear adipose tissue stimulated with pulsed electromagnetic field.

Baranowska A1, Skowron B1, Nowak B2, Ciesielczyk K1, Guzdek P3, Gil K1, Kaszuba-Zwoinska J4.

Author information

1
Department of Pathophysiology, Jagiellonian University Medical College, Cracow, Poland.
2
Department of Immunology, Jagiellonian University Medical College, Cracow, Poland.
3
Institute of Electron Technology, Cracow, Poland.
4
Department of Pathophysiology, Jagiellonian University Medical College, Cracow, Poland. jkaszuba@cm-uj.krakow.pl.

Abstract

Previous experiments demonstrated that low-frequency electromagnetic field (LF-EMF) may activate cellular death pathways in proliferating cells. Therefore, we hypothesized that LF-EMF may also influence viability of highly proliferating undifferentiated adipose-derived stem cells. Obesity is classified as a civilization disease; its etiopathogenesis is presumed to include both genetic predisposition and influence of modified environmental factors, such as unbalanced diet with excess calories and/or too low physical activity. Obesity may lead to a number of metabolic disorders, including type 2 diabetes mellitus, cardiovascular diseases (associated with atherosclerosis) related to primary hypertension and ischemic heart disease, myocardial infarction and other complications. The aim of this study was to verify if LF-EMF alters viability parameters of adipose-derived stem cells (ADSCs) isolated from rats, cultured in vitro and exposed to pulsed electromagnetic field (PEMF; 7 Hz, 30 mT). ADSCs were obtained from healthy rats and animals with experimentally-induced obesity, both males and females, pups and adults. The animals were fed with chow with either low (LF diet) or high fat content (HF diet) for 21 days. Then, ADSCs were isolated from extracted adipose tissue and used to establish cell cultures. ADSCs from the first passage were exposed to PEMF three times, 4 hours per exposure, at 24-h intervals (experimentally developed protocol of PEMF stimulation). 24 hours after the last exposure to PEMF, viability parameters of ADSCs were analyzed by flow cytometry (FCM). The study demonstrated that LF diet exerted a protective effect on PEMF-exposed ADSCs, especially in the case of male and female pups. In turn, the proportion of early apoptotic cells in PEMF-treated ADSC cultures from adult female rats maintained on HF diet turned out to be significantly higher than in other experimental groups.

Logo of sci

Stem Cells International
Stem Cells Int. 2017; 2017: 2450327.
Published online 2017 Apr 23. doi:  10.1155/2017/2450327
PMCID: PMC5420424
PMID: 28512472

Pulsed Electromagnetic Field Regulates MicroRNA 21 Expression to Activate TGF-? Signaling in Human Bone Marrow Stromal Cells to Enhance Osteoblast Differentiation

Nagarajan Selvamurugan, 1 Zhiming He, 2 Daniel Rifkin, 3 Branka Dabovic, 3 and Nicola C. Partridge 2 , *
Author information ? Article notes ? Copyright and License information ? Disclaimer

Abstract

Pulsed electromagnetic fields (PEMFs) have been documented to promote bone fracture healing in nonunions and increase lumbar spinal fusion rates. However, the molecular mechanisms by which PEMF stimulates differentiation of human bone marrow stromal cells (hBMSCs) into osteoblasts are not well understood. In this study the PEMF effects on hBMSCs were studied by microarray analysis. PEMF stimulation of hBMSCs’ cell numbers mainly affected genes of cell cycle regulation, cell structure, and growth receptors or kinase pathways. In the differentiation and mineralization stages, PEMF regulated preosteoblast gene expression and notably, the transforming growth factor-beta (TGF-?) signaling pathway and microRNA 21 (miR21) were most highly regulated. PEMF stimulated activation of Smad2 and miR21-5p expression in differentiated osteoblasts, and TGF-? signaling was essential for PEMF stimulation of alkaline phosphatase mRNA expression. Smad7, an antagonist of the TGF-? signaling pathway, was found to be miR21-5p’s putative target gene and PEMF caused a decrease in Smad7 expression. Expression of Runx2 was increased by PEMF treatment and the miR21-5p inhibitor prevented the PEMF stimulation of Runx2 expression in differentiating cells. Thus, PEMF could mediate its effects on bone metabolism by activation of the TGF-? signaling pathway and stimulation of expression of miR21-5p in hBMSCs.

1. Introduction

Abundant reports describe the effects of electricity on bone growth and fracture repair, and a variety of pulsed electromagnetic field (PEMF) devices have been developed to produce electromagnetic fields at the fracture site. These widespread PEMF devices utilize noninvasive inductive coupling and can be used along with every method of fracture fixation []. The stimulation of bone at the fracture site by the introduction of electromagnetic fields may be similar to the resulting stimulation from mechanical loading []. The beneficial therapeutic effects of such selected low energy, time varying PEMF promote fracture healing in nonunions [], increase lumbar spinal fusion rates [], and have been found to affect bone metabolism by decreasing bone resorption and increasing bone formation []. PEMFs have also been reported to stimulate the synthesis of extracellular matrix (ECM) proteins [] and may also affect several membrane receptors including those for parathyroid hormone, low density lipoprotein, insulin-like growth factor-2, insulin, and calcitonin []. Several growth factors such as bone morphogenetic proteins 2 and 4 (BMP-2, BMP-4) and transforming growth factor-beta (TGF-?) have been reported to be secreted from osteoblasts upon PEMF treatment []. It has been shown that electromagnetic stimulation could raise net Ca2+ flux in human osteoblast-like cells, and the increase in the cytosolic Ca2+ concentration could initiate activation of signaling pathways resulting in regulation of expression of bone matrix genes []. Accelerated osteogenesis has been found in bone marrow-derived mesenchymal stem cells by PEMF treatment [] and this promotion of ECM deposition was more efficient compared with adipose-tissue mesenchymal stem cells [].

Previously we have reported that both BMP-2 and PEMF (Spinal-Stim® by Orthofix, Inc., Lewisville, TX) separately stimulated proliferation of rat primary calvarial osteoblastic cells and stimulated expression of early osteoblast differentiation genes in culture []. In this study, we investigated the effects of PEMF (Cervical-Stim® by Orthofix, Inc., Lewisville, TX) on human bone marrow stromal cells (hBMSCs) proliferating and differentiated to osteoblastic cells. In addition, the underlying molecular mechanisms by which PEMF stimulates differentiation of hBMSCs into osteoblasts are not well understood. Thus, we also aimed to investigate the PEMF effects on proliferation, differentiation, and mineralization of hBMSCs by Affymetrix microarray analysis. The TGF-? signaling pathway and microRNA 21 (miR21) were most highly regulated by PEMF. Thus, in this study we systematically investigated the mechanism of action of PEMF effects on osteogenesis via TGF-? and miR21 using hBMSCs.

2. Materials and Methods

2.1. Cell Culture

Fresh human bone marrows from 21–68-year-old women were used. These were either purchased from Lonza (Walkersville, MD) or left over tissue from surgical procedures at New York University Hospital for Joint Diseases. Since these were deidentified, this is not considered Human Subjects Research by the New York University School of Medicine Institutional Review Board. In both cases, the bone marrows were freshly collected, never frozen, and immediately diluted 1?:?1 in Hank’s Balanced Salt Solution (HBSS; GIBCO Laboratories, Grand Island, NY) containing 20?IU/mL of sodium heparin (Sigma Chemical Co., St. Louis, MO). The diluted bone marrow was layered over an equal volume of Ficoll-Paque Plus (GE Healthcare, Piscataway, NJ) and centrifuged at 400g for 40?min at 18°C. The mononuclear cells at the interface layer were collected, washed three times with HBSS, resuspended and seeded into a tissue culture flask, and incubated at 37°C in the presence of 5% CO2 overnight. The next day, nonadherent cells were removed from the culture flask. Adherent cells (BMSCs) were grown to confluence then placed in 6-well plates at 6.4 × 104?cells/well for exposure to PEMF or control. All cells were incubated at 37°C in the presence of 5% CO2. The medium used for culturing these cells was ?-MEM (Corning, Tewksbury, MA) containing 15% fetal bovine serum (FBS; GIBCO, Grand Island, NY) and Penicillin-Streptomycin (GIBCO, Grand Island, NY).

2.2. PEMF Exposure

The PEMF was generated as previously described [] but was set to have similar waveform characteristics to a commercial, clinically approved proprietary device (Cervical-Stim by Orthofix Inc., Lewisville, TX). Cervical-Stim is the only device approved by the FDA for cervical fusion use and has been reported to be safe and effective []. The specific differences from our previous publication [] were a burst frequency of 15?Hz and a burst period of 67?ms. The induced magnetic field was vertical relative to the surface of the plates. The PEMF waveform was routinely checked for its consistency using a field probe and oscilloscope. The first PEMF exposure was initiated 24?h after seeding cells in wells (day 1) and continued through the entire experiment. Control plates were placed in an identical incubator on Plexiglas shelves. The CO2 concentration, humidity, and temperature of the control and treatment incubators (upper and lower chambers of the same double incubator) were identical and were not affected by the PEMF.

2.3. Cell Number

Cells were grown in normal growth medium and were trypsinized, resuspended, and counted using a hemocytometer when they reached 70–80% confluence on day 10 or 20 of culture, respectively, for the BMSCs from the younger (21–30) women versus those from the 31–65-year-old women.

2.4. Osteoblast Differentiation

Human BMSCs were seeded at 6.4 × 104 cells/well in 6-well cell culture plates and cultured for 10 days or 20 days in normal cell culture medium (?MEM + 15% FBS + 1% Penn/Strep) before they reached confluence. They were then cultured for an additional 13 (differentiation) or 23 (mineralization) days in osteogenic medium [normal growth medium supplemented with 10?4?M L-ascorbic acid, 10?8?M dexamethasone, and 1.80?mM potassium phosphate monobasic (Sigma, St. Louis, MO)]. The medium was changed three times/week.

2.5. Von Kossa Staining

For Von Kossa staining, 6 replicates of BMSCs were treated with PEMF or control daily from day 1 of culture. On day 23, 33, or 43, the cells were fixed with 95% ethanol for 15?min at 37°C, then rinsed and rehydrated through 80%, 50%, and 20% ethanol and then water, and incubated with 5% silver nitrate solution for 1?h at 37°C. The cells were rinsed with water, exposed to UV light for 10?min, and photographed. Von Kossa staining was analyzed by computer based morphometry (ImageJ: NIH, Bethesda, Maryland).

2.6. Extracellular Regulated Kinases Activation and Western Blot Analyses

Human BMSCs treated with control or PEMF for 5 and 10 days in the proliferation phase were washed with cold phosphate buffered saline (PBS) and lysed in Cell Lysis Buffer (Invitrogen, Grand Island, NY) containing protease and phosphatase inhibitor cocktails (Sigma). Cell lysates were centrifuged at 10,000?rpm for 10 minutes at 4°C and supernatants were saved and used for Western blot analysis. Twenty ?g of total cell protein was loaded per well and separated on 4–15% Mini-Protean TGX precast gels (Bio-Rad, Hercules, CA), followed by transferring to nitrocellulose membranes (Bio-Rad, Hercules, CA). The membranes were blocked and incubated with primary rabbit antibodies (Phospho-p44/42 MAPK (Erk1/2) (Thr202/Tyr204), p44/42 MAPK (Erk1/2) (137F5), or Cdk2 (sc-163; Cyclin dependent kinase 2, loading control)) overnight at 4°C. The membranes were then probed with secondary antibody conjugated with horseradish peroxidase. Finally, the bands were visualized by adding Super Signal West Dura Extended Duration Substrate (Thermo Scientific, Pittsburgh, PA) according to the manufacturer’s instructions. The primary antibodies to total ERKs and phosphorylated ERKs were obtained from Cell Signaling Technology (Danvers, MA), while the antibody to Cdk2 was obtained from Santa Cruz Biotechnology, Inc. (Dallas, TX). The secondary antibody (goat-anti-rabbit) conjugated with horseradish peroxidase (HRP) was obtained from Santa Cruz Biotechnology. Results were captured and quantitated by ChemiDoc XRS+ software (Bio-Rad, Hercules, CA). Both the Phospho-ERK1/2 and total ERK 1/2 were normalized to Cdk2 and then expressed as a percent of the values obtained in untreated control cells.

2.7. Microarray Assays

Human BMSCs of a 27-year-old healthy female donor were used for microarray experiments. Only hBMSCs expanded from the second to sixth passages were used for the experiments. PEMF treatment (Cervical-Stim) was initiated 24?h after hBMSCs were seeded, with 4?h daily exposure every day throughout the experimental period. Quadruplicate cell samples from both PEMF-treated and control groups were collected simultaneously at time points of hBMSC proliferation, osteoblast differentiation, and mineralization phases. Total RNA was isolated from cells by using TRIzol reagent (Thermo Scientific, Pittsburgh, PA) and then purified with RNeasy mini kit from Qiagen (Valencia, CA). Prior to microarray analysis, the RNA integrity was assessed by Agilent 2100 Bioanalyzer (Santa Clara, CA) and the best quality triplicate samples were chosen for the subsequent analyses. Microarrays and data analyses with Affymetrix Human U133 plus 2.0 Gene Chips (Santa Clara, CA) were performed at University of Medicine and Dentistry of New Jersey Genome Center according to the manufacturer’s instructions. In the case of gene expression where it was significantly found to be above 1.5-fold after PEMF treatment, gene ontology analysis was carried out by DAVID Bioinformatics Resources 6.7 software (NIAID, NIH).

2.8. Real-Time RT-PCR

Total RNA was isolated from cells using the total RNA isolation kit from Qiagen (Valencia, CA). For determination of expression of genes other than miR21-5p, 100?ng of total RNA from each sample was used for cDNA synthesis using TaqMan Reverse Transcription Reagents (Roche, Indianapolis, IN). Quantitative (q)PCR reactions were performed according to the real-time thermocycler machine (Realplex) manufacturer’s instructions (Eppendorf, Hauppauge, NY), which allowed real-time quantitative detection of the PCR products by measuring the increase in SYBR green fluorescence caused by binding of SYBR green to double-stranded DNA. The Power SYBR green master mix kit for PCR reactions was purchased from Invitrogen. The qPCR was performed in triplicate with reaction conditions of 95°C, 10?min, 1 cycle; 95°C, 15?sec; and 58.5°C, 1?min, for 40 cycles. Gene expression was analyzed with threshold cycle (CT) values averaged from triplicate samples and normalized to their CT values of housekeeping gene RPL13A. Primers were designed by NCBI primer Blast software. Table 1 lists the human-specific primers used for PCR amplification.

Table 1

Primers used in this study.

Gene name Forward primer (5? > 3?) Reverse primer (5? > 3?)
ALP TGGACGGCCTGGACCTCGTT AGGGTCAGGAGTTCCGTGCG
COL1A1 GGAGGCACGCGGAGTGTGAG CCTCTTGGCCGTGCGTCAGG
Osteocalcin GAGCCCCAGTCCCCTACCCG GACACCCTAGACCGGGCCGT
FOSB GCGCCGGGAACGAAATAAAC TTCGTAGGGGATCTTGCAGC
LEPR GTGGGGCTATTGGACTGACT CTTTGAGAGTCCAGCAGGCA
TBRG1 GCTAGATTCCTAGAGGCCCG GGCATCGGATCCTAAGTCGG
FBN2 CTTTAGGCCGGTTATGCAACG AATAAGCCCTTCGTCGGCTC
SOX11 TTGGAAGCGGAGAGCAACCT TGCGTTCGATCTTGGACCAT
CTNNA1 GGCAGCCAAAAGACAACAGG GGCCTTATAGGCTGCGACAT
AKT3 CTCTATTATTTGGGCTGAGTCATCA CCCCTCTTCTGAACCCAACC
CXCL12 GACAAGTGTGCATTGACCCG TGTAAGGGTTCCTCAGGCGT
THBS1 CCTCTACTCCGGACGCAC GCCCCGGTGAGTTCAAAGAT
COL5A1 CGGGGACTATGACTACGTGC CTCCAAGTCATCCGCACCTT
GPC4 CAGAGGTCCAGGTTGACACC TCGGCTTTCTCATTGGCACT
MMP16 TGCGGAACGGAGCAGTATTT TGTGCTTGTGCTGCCATTTC
TGFB2 CCCCGGAGGTGATTTCCATC AACTGGGCAGACAGTTTCGG
CDH11 CCCAGTACACGTTGATGGCT ACGTTCCCACATTGGACCTC
SPP1 GCCTCCTAGGCATCACCTG CTTACTTGGAAGGGTCTGTGGG
IL8 GGTGCAGTTTTGCCAAGGAG TTCCTTGGGGTCCAGACAGA
RPL13A AAGTACCAGGCAGTGACAG CCTGTTTCCGTAGCCTCATG
hsa-miR-21-5p UAGCUUAUCAGACUGAUGUUGA

For miR21-5p and snoR10-1, the reagents and primer sets for RT-qPCR were purchased from Qiagen. One ug of total RNA was reverse-transcribed into cDNA using the miScript II kit with miScript HiSpec Buffer according to the manufacturer’s instructions. The cDNA was then diluted 10 times and utilized as a template to amplify miR21-5p and snoR10-1 with the miScript SYBR Green PCR kit using the appropriate primers. snoR10-1 was used as normalizing gene control. The qPCR was performed in triplicate with reaction conditions of 95°C 15?min for Taq DNA polymerase activation, 94°C 15?sec denaturation, 55°C 30?sec annealing, and 70°C 30?sec extension for 40 cycles. Gene expression results of miR21-5p from either control or PEMF-treated groups were normalized to their relative snoR10-1 results.

2.9. TGF-? Signaling

Human BMSCs were cultured and treated with control or PEMF as described above. For TGF-? and BMP signaling assays, osteoblasts were treated with PEMF at days 23 and 33 and were also treated with TGF-?2 (R&D System, Minneapolis, MN) as a positive control for the TGF-? pathway. The day before assay, the cells were starved overnight (0.1% FBS medium) to reduce endogenous signaling activity. At day 23 at the same time as PEMF exposure started, 5?ng/mL TGF-?2 was added to the medium of positive control wells. Cell lysates from different groups were collected at 0, 2, and 4?h time points after treatment to examine Smad2, Smad3, and Smad1/5/8 protein phosphorylation by Western blot analysis as described above. Phospho-Smad2 (Ser465/467, 138D4)/Smad2 (D43B4), phospho-Smad3 (Ser423/425, C25A9)/Smad3 (C67H9), and phospho-Smad1/5/8/Smad1/5 antibodies were obtained from Cell Signaling Technology (Danvers, MA). In TGF-? neutralization experiments, 30?ug/mL normal rabbit IgG or TGF-? pan antibody (R&D System, Minneapolis, MN) was added to osteogenic medium during the entire differentiation period. At day 23, two non-PEMF-treated cell groups were also included with 5?ng/mL of TGF-?2 as positive controls. After 2?h of PEMF exposure, all sample groups were collected for Western blots and RT-qPCR assays.

2.10. Transient Transfection

Cells were seeded in growth medium in 6-well plates at a density of 105?cells/well on the day before the transfection. miR-21 is now referred as miR-21-5p, based on the latest miRBase release (V.21). miR21-5p inhibitor (Applied Biosystems: 4464084) designed to bind with endogenous miR21, when introduced into cells, inhibits its activities. miR21-5p mimic (Applied Biosystems: 4464066) was designed to be similar to that of endogenous miR21. A negative control miRNA (Applied Biosystems: 4464076) was included in the study. The X-treme Gene transfection reagent obtained from Roche, USA, was mixed with 50?nM of negative control miRNA, miR21 mimic, or miR21 inhibitor, and transient transfection was carried out [] for 3 or 6 days along with PEMF treatment every day for 4?h.

2.11. Statistical Analysis

Statistical analysis was done by one-way ANOVA, Student’s t-test, or Wilcoxon Ranking. Significant difference is p < 0.05. All data are shown as mean ± standard deviation with n as indicated.

3. Results

3.1. PEMF Effects on Proliferation and Differentiation of Human BMSCs from Subjects of Different Ages

We previously reported that PEMF generated by Spinal-Stim stimulated cell proliferation and expression of early differentiation marker genes in rat primary calvarial osteoblastic cultures []. In the present study we used PEMF (Cervical-Stim) to determine its effect on osteoblasts using human bone marrow cells. PEMF significantly stimulated the cell number of preosteoblasts from BMSCs of young women (21–30 years old) while not stimulating those of BMSCs from 31–65-year-old women (Figures 1(a) and 1(b)). It should also be noted that the hBMSCs from aged individuals (58, 59, and 65 years old) also required much longer time (20 days) to approach a similar cell culture density to those from the younger women. Since PEMF had an effect on preosteoblastic cell number from the younger women and cell proliferation involves activation of intracellular signaling pathways, especially extracellular regulated kinases (ERKs), we determined activation of these enzymes by PEMF. As shown in Figure 2(a), PEMF increased ERK activation (phosphorylation) after 15?min on day 5 in BMSCs from a 24-year-old woman. A similar effect was also found in hBMSCs from other younger female subjects (24- and 27-year-old women’s cells) and the quantitative analysis of ERK activation (phosphorylated ERKs) from the three individuals after normalization to total ERKs confirmed the above result (Figure 2(b)). There was no significant activation of ERKs from any of these cells on the 10th day of PEMF treatment (Figures 2(c) and 2(d)).

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.001.jpg

Effect of PEMF on hBMSC preosteoblastic cell number. (a) Human preosteoblasts derived from bone marrow stromal cells of 21–36-year-old women were treated with PEMF for 10 days; cells from 58-, 59-, and 65-year-old women were treated with PEMF for 20 days. Cell number/well was calculated using a hemocytometer (n = 3–6 wells). (b) Aggregation of the data into the two age groups, n = 5-6. The statistical p value for the younger versus older samples is shown using Student’s t-test analysis.

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.002.jpg

Effect of PEMF on ERK activation. Human BMSCs from two different 24-year-old women and a 27-year-old woman were subjected to 4?h daily PEMF treatment for either 4 days or 9 days. On the 5th (a) or 10th day (c), their cells were treated with PEMF for different time periods as indicated and whole cell lysates were obtained and subjected to Western blot analyses; cells from a 24-year-old woman are shown as an example. ((b) day 5, (d) day 10) The quantitation of activated or phosphorylated ERKs for cells from 2 separate 24-year-old women and a 27-year-old woman was determined by normalization of phosphorylated ERKs to total ERKs after normalization to Cdk2 as a loading control and expressed as a percent of untreated control cells. The results are shown for the cells of the 3 individuals. ? indicates significant increase compared to control. # indicates significant increase compared to 0 times of PEMF on the 5th day. The p value ? 0.05 is considered as significant using one-way ANOVA.

To determine the role played by PEMF in osteoblast differentiation and mineralization of hBMSCs, experiments were carried out at molecular and cellular levels. At the molecular level, the mRNA expression of alkaline phosphatase (ALP), type I collagen (COL1A1), and osteocalcin (OC), which are known osteoblast differentiation and mineralization marker genes, was determined using qRT-PCR analysis. PEMF significantly increased mRNA expression of ALP and Col1 but not OC in BMSCs that had been allowed to proliferate, differentiate, and mineralize (Figure 3(a)). We next determined the effect of PEMF on mineralization in BMSCs by Von Kossa staining (Figures 3(b) and 3(c)). PEMF significantly stimulated mineralization of BMSCs in the mineralization phase and did not in the differentiation phase.

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.003.jpg

Effect of PEMF on expression of osteoblastic marker genes and mineralization in hBMSCs. Differentiating preosteoblasts from 24–68-year-old women were grown in the presence of osteoblast differentiation medium after confluence was reached and were treated with PEMF for 33 days or 43 days (59–68-year-old samples) of culture. (a) Total RNA was isolated and subjected to qRT-PCR using specific primers for human ALP, type I collagen, OC, and RPL13A. n = 9. (b) Cells were then subjected to Von Kossa staining and the mineralized calcium deposits were quantified. n = 9. Statistical analyses were conducted using Wilcoxon signed rank test. The p value ? 0.05 is considered as significant compared with the controls. (c) An example of Von Kossa staining and mineralized calcium deposits for hBMSCs of a 24-year-old female after 33 days of osteogenic culture in the presence or absence of daily Cervical-Stim PEMF.

3.2. PEMF Regulation of Genes during hBMSC Proliferation by Microarray Analysis

A sample from a young individual was used for microarray analyses because PEMF significantly enhanced cell growth for young individuals compared to old individuals as shown in Figure 1. For assessment of the effect of PEMF on gene expression during hBMSC proliferation, on the 5th day of PEMF treatment 2?h after initiating the PEMF signal (pilot studies had shown significant PEMF stimulation of Cyclin gene expression at this time and day, data not shown), total RNA was isolated and used for the subsequent test with Affymetrix Human U133 plus 2.0 Gene Chips. After identifying significantly regulated genes, gene ontology analyses were performed by DAVID Bioinformatics Resources 6.7 software. The results indicated that PEMF stimulation of proliferating hBMSCs mainly affected genes of cell cycle regulation, cell structure, extracellular matrix (ECM), and some growth receptors or kinase pathways. There were a total of 114 known genes upregulated and 17 known genes downregulated at this time point (partially listed in Table 2). We have also included the decrease in fibrillin 2, even though it was not ?1.5-fold, since this sequesters members of the TGF-? family and is the subject of our later research in this report.

Table 2

Genes regulated by PEMF during hBMSCs proliferation by microarray analysis. Cells were from a normal 27-year-old female. Total RNA was isolated at day 5 after 2?h of PEMF treatment and used for microarray assays as described in Materials and Methods. Analysis by Student’s t-test.

Gene symbol Gene title Fold-change
(Avg PEMF versus avg controls)
p
Cell adhesion and binding and cytoskeletal and structural proteins
MMP1 Matrix metallopeptidase 1 (interstitial collagenase) 4.57 2.16E ? 03
PRC1 Protein regulator of cytokinesis 1 3.22 4.34E ? 04
CCBE1 Collagen and calcium binding EGF domains 1 2.97 1.32E ? 04
CLDN1 Claudin 1 2.71 2.26E ? 04
CENPK Centromere protein K 2.40 5.36E ? 03
GAS2L3 Growth arrest-specific 2 like 3 2.39 4.74E ? 03
CLDN11 Claudin 11 2.31 5.59E ? 04
NUSAP1 Nucleolar and spindle associated protein 1 2.16 5.04E ? 03
COL15A1 Collagen, type XV, alpha 1 2.12 1.64E ? 04
HAPLN1 Hyaluronan and proteoglycan link protein 1 2.03 1.37E ? 04
IBSP Integrin-binding sialoprotein 1.97 2.17E ? 03
FBN2 Fibrillin 2 ?1.12 1.98E ? 02
COL14A1 Collagen, type XIV, alpha 1 ?2.30 1.56E ? 02
MMP12 Matrix metallopeptidase 12 (macrophage elastase) ?3.27 1.19E ? 04
MGP Matrix Gla protein ?3.98 2.80E ? 06

p53 signaling pathway, apoptosis, and survival antiapoptotic TNFs/NF-kB/IAP pathway
BIRC5 Baculoviral IAP repeat containing 5 3.30 3.60E ? 06
GTSE1 G-2 and S-phase expressed 1 2.48 1.02E ? 03
SESN3 sestrin 3 ?2.47 3.72E ? 07

Cell cycle role of APC ?(anaphase-promoting complex) in cell cycle regulation, cell cycle/checkpoint control
CDK1 Cyclin-dependent kinase 1 5.07 1.01E ? 03
CDC20 Cell division cycle 20 homolog (S. cerevisiae) 3.00 3.32E ? 04
CCNB2 Cyclin B2 2.41 1.83E ? 03
NDC80 NDC80 kinetochore complex component homolog (S. cerevisiae) 2.38 7.59E ? 04
TYMS Thymidylate synthetase 2.36 9.43E ? 05
CCNB1 Cyclin B1 2.35 2.17E ? 03
NEK2 NIMA- (never in mitosis gene a-) related kinase 2 2.12 2.19E ? 02
CCNA2 Cyclin A2 1.95 6.42E ? 03
TTK TTK protein kinase 1.95 1.77E ? 02

Akt signaling
CCL2 Chemokine (C-C motif) ligand 2 2.83 6.76E ? 05
CSF2RB Colony stimulating factor 2 receptor, beta, low-affinity 2.31 2.04E ? 03

Other receptor, kinase, and regulator
CDKN3 Cyclin-dependent kinase inhibitor 3 2.45 2.32E ? 04
PDGFRA Platelet-derived growth factor receptor, alpha polypeptide 2.26 2.52E ? 05
CTSC Cathepsin C 2.14 5.32E ? 04
LEPR Leptin receptor ?2.19 1.46E ? 04
FGFR2 Fibroblast growth factor receptor 2 ?2.04 8.83E ? 05

3.3. PEMF Regulation of Genes in Differentiated and Mineralized hBMSCs by Microarray Analysis

In the differentiation (day 23) and mineralization stages (day 33) after daily 4?h PEMF treatment, a total of 37 (partially listed in Table 3) and 173 (partially listed in Table 4) known genes, respectively, were identified as significantly regulated. In these two stages, PEMF regulated preosteoblast gene expression and most genes were downregulated including transcriptional regulators, metabolism, proteases, and regulators and also cell adhesion and binding proteins and cytoskeletal and structural proteins. Changes in gene transcription of candidate genes chosen from microarray analyses were verified and confirmed by RT-qPCR on RNA from differentiated hBMSCs from 3 females, aged 24, 27, and 31 years (Table 5). Notably, the TGF-? signaling pathway seems to be most highly regulated by PEMF. In particular, RT-qPCR showed that fibrillin 2 (FBN2) was significantly decreased in expression by 65 ± 14%, while TGF-?2 mRNA significantly increased to 155 ± 44% and TGF-? regulator 1 (TBRG1) mRNA significantly increased to 143 ± 23%, relative to controls. In contrast, in mineralizing cells (Table 6), there was no decrease in FBN2 expression and a lesser significant increase in TGF-?2. It appears that PEMF stimulated a number of components of the TGF-? pathway in differentiating and mineralizing osteoblasts. It is notable that no components of the BMP pathway were seen to be regulated.

Table 3

Genes regulated by PEMF in differentiating hBMSCs. Cells were from a normal 27-year-old female. Total RNA was isolated at day 23 of PEMF treatment. Analysis by Student’s t-test.

Gene symbol Gene title Fold-change
(Avg PEMF versus avg controls)
p
Transcriptional regulator, RNA metabolism, and RNA transport
SPEN Spen homolog, transcriptional regulator (Drosophila) ?1.74 2.68E ? 02
FOXO3, FOXO3B Forkhead box O3; forkhead box O3B pseudogene ?1.87 3.04E ? 02
MIR21 MicroRNA 21 1.61 2.92E ? 02

Metabolic process
AKT3 v-akt murine thymoma viral oncogene homolog 3 ?1.58 2.39E ? 02

Growth factor and regulator
TBRG1 Transforming growth factor beta regulator 1 1.72 9.85E ? 03

Receptor
LEPR Leptin receptor 1.55 5.20E ? 03

Cell adhesion, motility, and cytoskeletal
ARPC5 Actin related protein 2/3 complex, subunit 5, 16?kDa 1.50 1.93E ? 02
FBN2 Fibrillin 2 ?1.45 1.47E ? 02

Signaling transduction, pathway
THBS1 Thrombospondin 1 ?1.24 1.33E ? 02

Table 4

Genes regulated by PEMF in mineralizing hBMSCs. Cells were from a normal 27-year-old female. Total RNA was isolated at day 33 of PEMF treatment. Analysis by Student’s t-test.

Gene symbol Gene title Fold-change
(Avg PEMF versus avg controls)
p
Cell adhesion, motility, and cytoskeletal
COL1A2 Collagen, type I, alpha 2 ?1.60 1.81E ? 02
COL3A1 Collagen, type III, alpha 1 ?1.61 9.49E ? 03
FN1 Fibronectin 1 ?1.93 2.31E ? 04
FBN2 Fibrillin 2 1.38 2.49E ? 02
VIM Vimentin ?1.67 1.39E ? 02

Transcriptional regulator, RNA metabolism, and RNA transport
MIR21 MicroRNA 21 ?2.16 1.28E ? 02
HNRNPA1 LOC728844 Heterogeneous nuclear ribonucleoprotein ?1.91 1.41E ? 02

Cell cycle, cell growth, and apoptosis
CCNL1 Cyclin L1 ?1.79 1.65E ? 03
CCNL2 Cyclin L2 ?2.07 3.18E ? 03

Hormone, growth factor, and cytokine
CXCL12 Chemokine (CXC motif) ligand 12 stromal cell-derived factor 1 1.60 3.83E ? 03
IL15 Interleukin 15 ?1.55 4.70E ? 03
IL8 Interleukin 8 ?2.00 3.71E ? 02
TBRG1 Transforming growth factor beta regulator 1 ?2.35 8.96E ? 03
TGFB2 Transforming growth factor, beta 2 1.39 2.75E ? 02

Metabolic process
INSIG1 Insulin induced gene 1 1.52 2.36E ? 02

Signaling transduction, pathway
DAB2 Disabled homolog 2, mitogen-responsive phosphoprotein (Drosophila) ?1.62 3.65E ? 02
THBS1 thrombospondin 1 1.52 3.67E ? 03
TIFA TRAF-interacting protein with forkhead-associated domain ?1.51 3.02E ? 03

Protease and regulator
SERPINE1 Serpin peptidase inhibitor, clade E member 1 ?2.04 2.81E ? 03
BAG2 BCL2-associated athanogene 2 ?1.61 1.14E ? 03

Table 5

Real-time RT-PCR of three different female donor samples’ hBMSCs, aged 24, 27, and 31 years in the differentiation stage. Analysis by Student’s t-test.

Gene symbol Gene title Average PEMF/control % p
COL1A1 Collagen, type I, alpha 1 133 ± 24% 3.90E ? 02
COL5A1 Collagen, type V, alpha 1 136 ± 25% 3.44E ? 02
CTNNA1 Catenin (cadherin-associated protein), alpha 1, 102?kDa 124 ± 3% 8.27E ? 05
FOSB FBJ murine osteosarcoma viral oncogene homolog B 127 ± 33% 1.11E ? 01
SOX11 SRY- (sex determining region Y-) box 11 138 ± 24% 2.59E ? 02
SPP1 Secreted phosphoprotein 1 131 ± 41% 1.31E ? 01
TGFB2 Transforming growth factor, beta 2 155 ± 44% 4.87E ? 02
TBRG1 Transforming growth factor beta regulator 1 143 ± 23% 1.65E ? 02
AKT3 v-akt murine thymoma viral oncogene homolog 3 (protein kinase B, gamma) 74 ± 15% 2.06E ? 02
FBN2 Fibrillin 2 35 ± 14% 7.38E ? 04
IL8 Interleukin 8 56 ± 35% 4.87E ? 02

Table 6

Real-time RT-PCR analysis of three different female donor samples’ hBMSCs, aged 24, 27, and 31 years in the mineralization stage. Analysis by Student’s t-test.

Gene symbol Gene title Average PEMF/control % p
CDH11 Cadherin 11, type 2, OB-cadherin (osteoblast) 130 ± 36% 1.10E ? 01
COL1A1 Collagen, type I, alpha 1 144 ± 47% 9.20E ? 02
CXCL12 Chemokine (C-X-C motif) ligand 12 146 ± 20% 8.48E ? 03
FBN2 Fibrillin 2 149 ± 54% 9.82E ? 02
FOSB FBJ murine osteosarcoma viral oncogene homolog B 165 ± 27% 6.97E ? 03
GPC4 Glypican 4 128 ± 25% 6.23E ? 02
IL8 Interleukin 8 162 ± 68% 9.49E ? 02
LEPR Leptin receptor 130 ± 19% 2.53E ? 02
MMP16 Matrix metallopeptidase 16 (membrane-inserted) 135 ± 68% 2.07E ? 01
SOX11 SRY- (sex determining region Y-) box 11 137 ± 37% 7.60E ? 02
SPP1 Secreted phosphoprotein 1 132 ± 52% 1.74E ? 01
TGFB2 Transforming growth factor, beta 2 128 ± 21% 3.99E ? 02
TBRG1 Transforming growth factor beta regulator 1 113 ± 14% 9.71E ? 02
THBS1 Thrombospondin 1 142 ± 58% 1.37E ? 01

3.4. PEMF Activation of TGF-? Signaling via Smad2 in Differentiated and Mineralizing Osteoblasts

To validate the PEMF effect on activation of the TGF-? signaling pathway, hBMSCs were subjected to differentiation (day 23) and mineralization (day 33) as described. During differentiation and mineralization, the cells were continuously treated with PEMF for 4?h each day. At days 23 and 33, cells were subjected to control, TGF-?2, or PEMF treatments for 0, 2, and 4?h. TGF-?2 was used as a positive control for activation of TGF-? signaling. Whole cell lysates were prepared and subjected to Western blot analyses using the antibodies for phosphorylated and total Smad2. The results show that PEMF stimulated activation of Smad2 by increased phosphorylation at day 23 in differentiated osteoblasts (Figure 4(a)) and less at day 33 in mineralizing osteoblasts (Figure 4(b)). To determine the specificity of activation of the TGF-?signaling by PEMF, osteoblasts were pretreated with pan-TGF-? antibody before PEMF treatment. The results show that the PEMF-stimulated Smad2 activation in differentiated osteoblasts (day 23) was blocked when cells were pretreated with pan-TGF-? antibody (Figure 4(c)). Since a recent paper has described a different PEMF signal as acting through the BMP pathway on rat calvarial osteoblasts [], we examined whether Smad1/5/8 was phosphorylated in response to the Cervical-Stim signal in hBMSCs (Figure 4(d)). We were unable to observe any stimulation of this pathway, in contrast to the activation of the Smad2 pathway, even though the strong positive control, TGF-?2, slightly stimulated Smad1/5/8 phosphorylation, as has been observed by others [].

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.004.jpg

PEMF resulted in activation of the TGF-? signaling pathway in human osteoblastic cells during differentiation and mineralization. (a) Whole cell lysates after PEMF treatment of hBMSCs of a 24-year-old female at day 23 (differentiation) and (b) at day 33 (mineralization) were subjected to Western blot analysis using the antibodies as indicated for Smad2 and Cdk2. TGF-?2 (5?ng/mL) was added to control (non-PEMF-treated) cells on days 23 and 33 as positive controls. (c) The pan-TGF-? neutralizing antibody (30?ug/mL) was added to the osteogenic medium of hBMSCs from a 24-year-old female during the entire differentiation period and lysates were prepared on day 23 of PEMF treatment, 2?h after PEMF was started or TGF-?2 was added and subjected to Western blot analysis. TGF-?2 (5?ng/mL) was added to control (non-PEMF-treated) cells on day 23 as a positive control. Cdk2 was used as loading control. (d) The same lysates were subjected to Western blot analysis for phosphorylation of Smad1/5/8 as indicated.

3.5. PEMF Stimulates Osteoblast Marker Gene Expression by Activation of the TGF-? Signaling Pathway

To determine if TGF-? signaling is responsible for the PEMF effect on expression of osteoblast differentiation marker genes such as ALP and type I collagen, this pathway was inhibited and RNA collected from differentiated osteoblasts at day 23 and subjected to RT-qPCR analysis. We found that PEMF significantly stimulated mRNA expression of ALP (Figure 5(a)) and type I collagen (Figure 5(b)) in differentiated osteoblasts. When cells were pretreated with pan-TGF-? antibody, PEMF stimulation of expression of these genes was significantly decreased (Figures 5(a) and 5(b)). Thus, this result indicates that the osteogenic effect of Cervical-Stim PEMF on hBMSCs is mediated via the TGF-? signaling pathway.

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.005.jpg

PEMF resulted in stimulation of expression of osteoblast differentiation marker genes via the TGF-? signaling pathway. Differentiated human osteoblasts derived from hBMSCs from a 30-year-old female were used. Total RNA was isolated after incubation with IgG or pan-TGF-? antibody (Pan-Anti) and treatment with control (Ctr) or PEMF and subjected to RT-qPCR using the primers for (a) ALP and (b) collagen 1A1 genes. RPL13 was used to normalize gene expression. n = 3. ? indicates significant increase compared with control IgG. # indicates significant decrease compared to all groups with ALP mRNA expression; ## indicates significant decrease compared to control or PEMF treatment with IgG incubation with collagen 1A1 mRNA expression; analysis by one-way ANOVA.

3.6. PEMF Stimulation of miR21-5p Expression in Differentiating Osteoblasts

MicroRNAs are considered to be regulators of osteogenesis and bone formation. The microarray analysis of hBMSCs subjected to differentiation at day 23 identified the stimulation of expression of miR21 (Table 3). To verify this, total RNA was obtained with differentiated hBMSCs from females aged 24 × 2, 27, 29, and 30 (young individuals) and 31, 36, 58, and 68 (older individuals) years and subjected to RT-qPCR. The result shows that the expression of miR21-5p was 155% increased in cells from the younger women but not significantly increased in cells from the older individuals after PEMF treatment (Figure 6).

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.006.jpg

PEMF stimulated expression of miR21-5p in differentiated human osteoblasts. Total RNAs from control or PEMF-treated hBMSCs of females (24 × 2, 27, 29, and 30 years old, n = 5) at day 23 of differentiation or (31, 36, 58, and 68 years old) at day 23 or 33 of differentiation were isolated and subjected to RT-qPCR using the miScript II kit with miScript HiSpec Buffer and miScript SYBR Green PCR Kit. snoR10-1 was used to normalize miR21-5p expression and the expression is shown as a percentage of the relevant control samples. ?indicates significant increase compared to control using one-way ANOVA.

3.7. PEMF and miR21-5p Stimulation of Osteoblast Differentiation Marker Gene Expression

It is evident that PEMF stimulated miR21-5p expression in differentiated osteoblasts from younger individuals (Figure 6) which strongly suggested a role for miR21-5p in promotion of osteoblast differentiation. To determine this role, hBMSCs were transiently transfected with negative control miRNA or miR21-5p mimic for 3 days and concurrently subjected to 4?h PEMF treatment every day for 6 days. Total RNA was isolated and subjected to RT-qPCR analysis. When cells were treated with PEMF, there was significantly increased ALP mRNA expression. The miR21-5p mimic alone had no effect but together with PEMF treatment caused a significant increase in ALP mRNA expression compared with PEMF treatment alone (Figure 7(a)). With type I collagen mRNA expression, no significant effect was seen with respect to PEMF, miR21-5p mimic, or both treatments under these conditions (Figure 7(b)).

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.007.jpg

PEMF resulted in stimulation of expression of ALP mRNA and its effect was further enhanced by miR21-5p. Human BMSCs from a 31-year-old female were transiently transfected with 50?nM of negative control miRNA or miR-21-5p mimic for 72?h in osteogenic medium and PEMF treatment was carried out for 4?h each day for a total of 6 days. Total RNA was isolated and RT-qPCR was carried out using the primers for ALP (a) and collagen 1A1 (b) genes. Expression of the mRNAs is shown relative to the RPL13 gene. n = 3. ?? indicates significant increase compared to negative control miRNA transfection. # indicates significant increase compared to all treatments. Analysis by one-way ANOVA.

3.8. PEMF Regulation of Smad7 via miR21-5p in Differentiating Osteoblasts

In silico analysis (http://www.microrna.org/microrna/home.do) was used to identify the putative target genes of miR21-5p for its functional importance towards osteogenic commitment. Among them some antagonistic effectors of osteogenesis such as Smad7, Smurf1, and Crim1 were found. The 3?UTR regions of Smad7, Smurf1, and Crim1 held at least 6-nt perfect complementarities to the miR21-5p seed region (Figure 8(a)). To validate these putative target genes of miR21-5p, hBMSCs were transiently transfected with either negative control miRNA or miR21-5p inhibitor and concurrently treated with PEMF for 4?h each day for 3 days. To determine the expression level of these target genes, total RNA was isolated, followed by RT-qPCR analysis. There was no significant change in mRNA expression of Smurf2 (Figure 8(b)) and Crim1 (Figure 8(c)) in the cells in the presence of PEMF treatment, miR21-5p inhibitor, or both. In the case of Smad7, there was a significant decrease in its mRNA expression after PEMF treatment, and inclusion of miR21-5p inhibitor reversed the PEMF effect resulting in increased Smad7 mRNA expression (Figure 8(d)). From these results we suggest that Smad7, an antagonist of TGF-?signaling, is likely to be miR21-5p’s target gene and PEMF downregulates its mRNA expression via miR21-5p in differentiating osteoblasts. In fact, at least two groups have shown that the 3?-UTR of Smad7 is, indeed, a target for miR21-5p, resulting in a decrease in Smad7 protein levels [].

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.008.jpg

Putative target genes of miR21-5p and PEMF decreases Smad7 mRNA through miR21-5p. (a) The putative target region analysis was performed for Smurf2, Crim1, and Smad7 mRNAs 3? UTR by miR21-5p seed sequence. ((b)–(d)) Human BMSCs from a 27-year-old female were transiently transfected with 50?nM of negative control miRNA or miR21-5p inhibitor for 72?h in osteogenic medium and PEMF treatment was carried out concurrently for 4?h each day for 3 days. Total RNA was isolated and RT-qPCR was carried out using the primers for (a) Smurf2, (b) Crim1, and (c) Smad7 genes. Expression of mRNAs is shown relative to that of the RPL13 gene. n = 3. ? indicates significant increase compared to negative control miRNA transfection or PEMF treatment with negative control miRNA transfection. # indicates significant decrease compared to PEMF treatment with miR21-5p inhibitor transfection. Analysis by one-way ANOVA.

3.9. PEMF Regulation of Runx2 Expression via miR21-5p and Smad7 in Differentiating Osteoblasts

Since Runx2 is required for osteoblast differentiation and PEMF stimulated expression of osteoblast differentiation marker genes (Figure 3), we next examined the PEMF stimulation of expression of Runx2 in differentiating hBMSCs and the role played by miR21-5p. Human BMSCs were transiently transfected with either negative control miRNA or miR-21-5p inhibitor, followed by PEMF treatment. Total RNA was isolated and subjected to RT-qPCR analysis. The result showed that there was a significant increase in expression of Runx2 mRNA in response to PEMF treatment and this effect was blocked by miR21-5p inhibitor in differentiating osteoblasts (Figure 9). From these results, we suggest that PEMF promotes its osteogenic effect via stimulation of miR21-5p expression and activation of TGF-? signaling in hBMSCs. A figure summarizing that the mechanisms we conclude are involved in PEMF stimulation of BMSCs and osteoblast differentiation is shown in Figure 10.

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.009.jpg

PEMF stimulated Runx2 expression and its effect was downregulated by miR21-5p inhibitor. Human BMSCs from a 27-year-old female were transiently transfected with 50?nM of negative control miRNA or miR21-5p inhibitor for 3 days in osteogenic medium and PEMF treatment was carried out concurrently for 4?h each day for 3 days. Total RNA was isolated and RT-qPCR was carried out using the primers for Runx2. Expression of Runx2 mRNA is shown relative to that of the RPL13 gene. n = 3. ? indicates significant decrease compared to negative control miRNA transfection or PEMF treatment with negative control miRNA transfection. # indicates significant increase compared to control treatment with negative control miRNA transfection. Analysis by one-way ANOVA.

An external file that holds a picture, illustration, etc. Object name is SCI2017-2450327.010.jpg

Schema of the mechanisms involved in PEMF stimulation of BMSC proliferation and osteoblast differentiation. The magnetic field (B) is thought to elicit Eddy Currents that act on BMSCs and cause activation of ERKs that are then involved in increased BMSC proliferation. After the BMSCs reach confluence and they are switched to differentiation medium, the magnetic field (B) and the resultant Eddy Currents cause a decrease in fibrillin 2 expression and an increase in TGF-?2 and miR21-5p expression. The decrease in fibrillin 2 would lead to an increase in the amount of available TGF-?2. The increase in miR21-5p appears to cause a decrease in inhibitory Smad7 expression, thus, enhancing TGF-?2 activation of Smad2 with resulting increase in Runx2, collagen I, and alkaline phosphatase expression in the cultures, that is, increased osteoblast differentiation.

4. Discussion

Numerous studies have shown that mechanical stimulation of bone progenitors including ultrasound [], mechanical strain [], and compression as well as shear forces has a stimulatory effect on bone progenitors involved in bone healing of critical size defects and nonunions in vivo. A broad set of investigations has aimed to unravel potential underlying molecular mechanisms and growth factor pathways involved with sophisticated in vitro methods []. A number of mechanisms have been proposed by which mechanical cues on different physical scales and identities can incorporate into growth factor signaling []. In particular, the major TGF-? growth factor superfamily of ligands (including TGF-? 1 and 2 as well as BMPs) and their downstream signaling via Smad2/3 and Smad1/5/8 transcription factors, respectively [], appears to be affected by mechanical stimulation in a diverse set of cells, with the majority of research focussing on bone progenitors, for example, BMSCs, osteoblasts, osteocytes, and chondrocytes. This is a large and ongoing field of study.

The molecular mechanisms responsible for the effect of PEMF on bone formation [] have not been completely elucidated. We found that PEMF promoted preosteoblast proliferation from hBMSCs from individuals up to age 30, but not older individuals, and stimulated differentiation marker gene expression of mineralizing hBMSCs of all ages. To dissect the mechanisms, PEMF effects on proliferation, differentiation, and mineralization of hBMSCs were examined by Affymetrix microarray analyses. We found that PEMF stimulation of hBMSC proliferation mainly affected genes of cell cycle regulation, cell structure, ECM, and some growth receptors or kinase pathways (Table 2). At the cellular and molecular levels, PEMF has been reported to promote the synthesis of ECM proteins and exert a direct effect on the production of proteins that regulate gene transcription. PEMF may affect several membrane receptors and stimulate osteoblasts to secrete several growth factors such as BMP-2 and BMP-4 and TGF-?. PEMF has been reported to affect osteoblast cellular proliferation and differentiation of bone cells in vitro by enhancing DNA synthesis [], increasing the expression of bone marker genes during differentiation and mineralization [], and enhancing calcified matrix production. Several experimental studies also demonstrated that PEMF stimulation could potently promote osteogenesis and enhance bone mineralization both in vivo and in vitro [].

The microarray data for PEMF regulation of differentiation and mineralization of hBMSCs showed regulation of transcriptional regulators, metabolism, proteases, cytokines and growth factors, and also cell adhesion and binding proteins and cytoskeletal and structural proteins (Tables ?(Tables33 and ?and4).4). Identifying the signaling pathways and their associated regulatory mechanisms of PEMF action on osteogenesis might further promote its use in clinical applications. Thus, PEMF regulated preosteoblast gene expression during the differentiation and mineralization stages, and candidate genes chosen from microarray analyses were confirmed by RT-qPCR (Tables ?(Tables55 and ?and6).6). Notably, the TGF-? signaling pathway and miR21 seem to be most highly regulated by PEMF. Thus, in the present study, we systematically investigated the mechanism of action of PEMF effects on osteogenesis via activation of TGF-? signaling and miR21-5p expression using hBMSCs.

The TGF-?/BMP signaling pathway plays a fundamental role in the regulation of bone organogenesis through the activation of receptor serine/threonine kinases. Perturbations of TGF-?/BMP activity are almost invariably linked to a wide variety of clinical outcomes including skeletal anomalies []. Phosphorylation of TGF-? (I/II) or BMP receptors activates intracellular downstream Smads, the transducer of TGF-?/BMP signals. In our studies, PEMF (Cervical-Stim) treatment activated only the Smad2 signaling component in differentiated hBMSCs (Figure 4) and activation of this signaling pathway appeared to be essential for PEMF stimulation of early osteoblast differentiation marker genes such as ALP and type I collagen (Figure 5). It is notable that it did not appear to activate the BMP pathway through Smad1/5/8 phosphorylation. The TGF-?/BMP signaling effect may be complex and highly time- and space-specific during skeletal development and bone formation. Very recently, Xie et al. [] have described a different PEMF signal as operating through the BMP receptor on the primary cilium of rat calvarial osteoblasts in culture. Our accumulated data do not indicate that the BMP pathway is involved in the signaling mechanism of either Spinal-Stim or Cervical-Stim but we cannot rule out that it may have a role if investigated further. This signaling cascade can be modulated by various factors and other pathways []. Activation of Wnt/Lrp5/?-catenin or calcium-related mechanisms by PEMF treatment for osteogenic activity have also been reported [].

Osteoblast differentiation is tightly controlled by several regulators including miRNAs [] that can regulate expression of genes during differentiation of MSCs towards osteoblasts, resulting in the osteogenic lineage. Differential expression of miRNAs could be responsible for activation of several signaling pathways such as TGF-?/BMP, Wnt/?-catenin, and transcription factors []. PEMF stimulated miR21-5p expression in differentiated hBMSCs from younger females (Figure 6) suggesting one of the ways PEMF mediates its osteogenic effect on these cells is via miR21-5p. MicroRNA 21 was one of the first miRNAs detected in the human genome and it was found to be overexpressed in several types of cancer tissues []. A role for miR21 in cell proliferation and apoptosis has been reported []. With regard to the regulation of bone formation, a number of miRNAs are expressed in the developing skeletal system and miRNA-dependent modulation of gene function can alter skeletal phenotypes across individuals and also within the same individual over time []. MicroRNAs might have direct or indirect effects for their regulatory functions in osteoblast differentiation.

To study the functional role of miR21-5p during osteoblast differentiation by PEMF treatment, it was necessary to alter its endogenous expression/activity. Overexpression of miR21-5p (mimic) in differentiated hBMSCs had no effect on mRNA expression of ALP and type I collagen (Figure 7) but required PEMF to have an enhanced effect on ALP mRNA expression which suggests that PEMF could also involve other pathways and molecules in addition to miR21-5p for its osteogenic effects in these cells. The putative targets of miR21-5p can be classified according to their negative contribution in osteogenic differentiation or positive contribution to other lineages using online software. Among them are some key regulators or antagonistic effectors of osteogenesis such as Smad7, Smurf2, and Crim1 and these genes are well documented in their antagonistic roles in osteogenesis []. Expression of the putative target genes in the presence of the miR21-5p inhibitor showed a significant increase in Smad7 mRNA expression in differentiated hBMSCs (Figure 8). The inhibitory Smads (Smad6, Smad7) potentially act as suppressors of bone formation. While Smad7 inhibits TGF-?/BMP signaling, Smad6 is less effective in inhibiting TGF-?signaling. It has been reported that Smad7 can inhibit ALP activity and suppress type I collagen mRNA and protein levels []. MicroRNA 21 has been shown to be a key regulator of TGF-? signaling [] and Smad7 was found to be one of its target genes []. Other target genes such as PTEN and STAT3 have also been reported for miR21 []. Based on our results (Figures ?(Figures77 and ?and8),8), we suggest that Smad7 is a target gene for miR21-5p during PEMF regulation of osteoblast differentiation.

Since PEMF stimulates miR21-5p expression in differentiated hBMSCs (Figure 6) and miR21-5p targets Smad7 (Figure 8(d)), the PEMF action on osteogenesis via miR21-5p and Smad7 was further investigated. Runx2 is essential for the commitment of multipotent mesenchymal cells to the osteoblastic lineage. In general, Runx2 activity can be altered by its interacting proteins and/or posttranslational modifications []. The steady-state protein level of Runx2 can be regulated by E3 ubiquitin ligases, Smurf1 and Smurf2, and it has been reported that the degradation of endogenous Runx2 can be blocked by a proteasomal inhibitor or by Smurf2 siRNA []. PEMF stimulated Runx2 mRNA in differentiated hBMSCs, and miR21-5p inhibitor prevented the PEMF stimulation of Runx2 expression (Figure 9). It has already been reported that Smad7 interacts with Smurf2 but it does not interact with Runx2 []. Hence, targeting Smad7 through miR21-5p by PEMF could possibly decrease the Smad7-dependent Smurf2 activity, resulting in stabilization of Runx2 protein, and feedback to increased transcription of Runx2 in differentiated hBMSCs. A figure summarizing that the mechanisms we conclude are involved in PEMF stimulation of BMSCs and osteoblast differentiation is shown in Figure 10. We can only speculate as to how PEMF regulates miR21-5p, but others have shown that this microRNA is regulated by transcriptional mechanisms, such as by myocardin-related transcription factor-A (58) or by STAT3 (59), and such mechanisms could possibly be implicated in PEMF’s actions.

5. Conclusions

Our results show that PEMF significantly stimulated the cell number of preosteoblasts from BMSCs of young women while not stimulating those from women older than 30. We also showed that PEMF regulates a range of genes in hBMSCs to stimulate their proliferation, differentiation, and mineralization. Our further investigation suggests a novel regulatory mechanism of PEMF action during differentiation and mineralization of hBMSCs by activation of the TGF-? signaling pathway. PEMF appears to activate this pathway in hBMSCs of younger women by inhibiting Smad7 expression through miR21-5p and in turn PEMF controls the function of Runx2 resulting in promotion of its osteogenic effect.

Acknowledgments

This work was supported by a research contract to Dr. Partridge from Orthofix, Inc., Lewisville, TX, USA. Dr. Partridge also received honoraria and travel support from Orthofix, Inc. Dr. Selvamurugan and Mr. He received travel support from Orthofix, Inc. Drs. Rifkin and Dabovic received a subcontract from Dr. Partridge’s Orthofix research contract. Drs. James T. Ryaby, Nianli Zhang, and Erik I. Waldorff of Orthofix, Inc., critically reviewed the paper.

Conflicts of Interest

The authors declare that they have no conflicts of interest.

Authors’ Contributions

Nagarajan Selvamurugan and Zhiming He contributed equally to the work; Nagarajan Selvamurugan drafted the paper; Nagarajan Selvamurugan, Zhiming He, Daniel Rifkin, and Branka Dabovic contributed to the research design, acquisition, analysis, and interpretation of data; Nicola C. Partridge designed the research, contributed to the analysis and interpretation of the data, and critically revised the paper. All authors have read and approved the final submitted manuscript.

References

1. Victoria G., Petrisor B., Drew B., Dick D. Bone stimulation for fracture healing: what?s all the fuss? Indian Journal of Orthopaedics2009;43(2):117–120. doi: 10.4103/0019-5413.50844. [PMC free article][PubMed] [Cross Ref]
2. Kooistra B. W., Jain A., Hanson B. Electrical stimulation: nonunions. Indian Journal of Orthopaedics2009;43(2):149–155. doi: 10.4103/0019-5413.50849. [PMC free article] [PubMed] [Cross Ref]
3. Garland D. E., Moses B., Salyer W. Long-term follow-up of fracture nonunions treated with PEMFs. Contemporary Orthopaedics1991;22(3):295–302. [PubMed]
4. Linovitz R. J., Pathria M., Bernhardt M., et al. Combined magnetic fields accelerate and increase spine fusion: a double-blind, randomized, placebo controlled study. Spine2002;27(13):1383–1389. doi: 10.1097/00007632-200207010-00002. [PubMed] [Cross Ref]
5. Mooney V. A randomized double-blind prospective study of the efficacy of pulsed electromagnetic fields for interbody lumbar fusions. Spine1990;15(7):708–712. doi: 10.1097/00007632-199007000-00016.[PubMed] [Cross Ref]
6. Bodamyali T., Bhatt B., Hughes F. J., et al. Pulsed electromagnetic fields simultaneously induce osteogenesis and upregulate transcription of bone morphogenetic proteins 2 and 4 in rat osteoblasts in vitro. Biochemical and Biophysical Research Communications1998;250(2):458–461. doi: 10.1006/bbrc.1998.9243. [PubMed] [Cross Ref]
7. Selvamurugan N., Kwok S., Vasilov A., Jefcoat S. C., Partridge N. C. Effects of BMP-2 and pulsed electromagnetic field (PEMF) on rat primary osteoblastic cell proliferation and gene expression. Journal of Orthopaedic Research2007;25(9):1213–1220. doi: 10.1002/jor.20409. [PubMed] [Cross Ref]
8. Partridge N. C., He Z., Selvamurugan N. Microarray analysis of pulsed electromagnetic field (PEMF) stimulatory effects on human bone marrow stromal cells. Journal of Bone and Mineral Research2014;29(supplement 1):p. 432.
9. Aaron R. K., Ciombor D. M., Simon B. J. Treatment of nonunions with electric and electromagnetic fields. Clinical Orthopaedics and Related Research2004;(419):21–29. [PubMed]
10. Ciombor D. M., Aaron R. K. The role of electrical stimulation in bone repair. Foot and Ankle Clinics2005;10(4):579–593. doi: 10.1016/j.fcl.2005.06.006. [PubMed] [Cross Ref]
11. Kuzyk P. R. T., Schemitsch E. H. The science of electrical stimulation therapy for fracture healing. Indian Journal of Orthopaedics2009;43(2):127–131. doi: 10.4103/0019-5413.50846. [PMC free article][PubMed] [Cross Ref]
12. Fitzsimmons R. J., Ryaby J. T., Magee F. P., Baylink D. J. Combined magnetic fields increased net calcium flux in bone cells. Calcified Tissue International1994;55(5):376–380. doi: 10.1007/BF00299318.[PubMed] [Cross Ref]
13. Pavalko F. M., Norvell S. M., Burr D. B., Turner C. H., Duncan R. L., Bidwell J. P. A model for mechanotransduction in bone cells: the load-bearing mechanosomes. Journal of Cellular Biochemistry2003;88(1):104–112. doi: 10.1002/jcb.10284. [PubMed] [Cross Ref]
14. Tsai M.-T., Chang W. H.-S., Chang K., Hou R.-J., Wu T.-W. Pulsed electromagnetic fields affect osteoblast proliferation and differentiation in bone tissue engineering. Bioelectromagnetics2007;28(7):519–528. doi: 10.1002/bem.20336. [PubMed] [Cross Ref]
15. Ceccarelli G., Bloise N., Mantelli M., et al. A comparative analysis of the in vitro effects of pulsed electromagnetic field treatment on osteogenic differentiation of two different mesenchymal cell lineages. BioResearch Open Access2013;2(4):283–294. doi: 10.1089/biores.2013.0016. [PMC free article][PubMed] [Cross Ref]
16. Foley K. T., Mroz T. E., Arnold P. M., et al. Randomized, prospective, and controlled clinical trial of pulsed electromagnetic field stimulation for cervical fusion. Spine Journal2008;8(3):436–442. doi: 10.1016/j.spinee.2007.06.006. [PubMed] [Cross Ref]
17. Vimalraj S., Partridge N. C., Selvamurugan N. A positive role of microRNA-15b on regulation of osteoblast differentiation. Journal of Cellular Physiology2014;229(9):1236–1244. doi: 10.1002/jcp.24557. [PMC free article] [PubMed] [Cross Ref]
18. Xie Y. F., Shi W. G., Zhou J., et al. Pulsed electromagnetic fields stimulate osteogenic differentiation and maturation of osteoblasts by upregulating the expression of BMPRII localized at the base of primary cilium. Bone2016;93:22–32. doi: 10.1016/j.bone.2016.09.008. [PubMed] [Cross Ref]
19. Wrighton K. H., Lin X., Yu P. B., Feng X.-H. Transforming growth factor ? can stimulate Smad1 phosphorylation independently of bone morphogenic protein receptors. Journal of Biological Chemistry2009;284(15):9755–9763. doi: 10.1074/jbc.m809223200. [PMC free article] [PubMed] [Cross Ref]
20. Daly A. C., Randall R. A., Hill C. S. Transforming growth factor ?-induced Smad1/5 phosphorylation in epithelial cells is mediated by novel receptor complexes and is essential for anchorage-independent growth. Molecular and Cellular Biology2008;28(22):6889–6902. doi: 10.1128/MCB.01192-08.[PMC free article] [PubMed] [Cross Ref]
21. Lin L., Gan H., Zhang H., et al. MicroRNA-21 inhibits SMAD7 expression through a target sequence in the 3? untranslated region and inhibits proliferation of renal tubular epithelial cells. Molecular Medicine Reports2014;10(2):707–712. doi: 10.3892/mmr.2014.2312. [PubMed] [Cross Ref]
22. Li Q., Zhang D., Wang Y., et al. MiR-21/Smad 7 signaling determines TGF-?1-induced CAF formation. Scientific Reports2013;3, article 2038 doi: 10.1038/srep02038. [PMC free article] [PubMed][Cross Ref]
23. Padilla F., Puts R., Vico L., Guignandon A., Raum K. Stimulation of bone repair with ultrasound. Advances in Experimental Medicine and Biology2016;880:385–427. doi: 10.1007/978-3-319-22536-4_21.[PubMed] [Cross Ref]
24. Chen J. C., Hoey D. A., Chua M., Bellon R., Jacobs C. R. Mechanical signals promote osteogenic fate through a primary cilia-mediated mechanism. FASEB Journal2016;30(4):1504–1511. doi: 10.1096/fj.15-276402. [PMC free article] [PubMed] [Cross Ref]
25. Rosa N., Simoes R., Magalhães F. D., Marques A. T. From mechanical stimulus to bone formation: a review. Medical Engineering and Physics2015;37(8):719–728. doi: 10.1016/j.medengphy.2015.05.015.[PubMed] [Cross Ref]
26. Gao J., Fu S., Zeng Z., et al. Cyclic stretch promotes osteogenesis-related gene expression in osteoblast-like cells through a cofilin-associated mechanism. Molecular Medicine Reports2016;14(1):218–224. doi: 10.3892/mmr.2016.5239. [PMC free article] [PubMed] [Cross Ref]
27. Deng M., Liu P., Xiao H., et al. Improving the osteogenic efficacy of BMP2 with mechano growth factor by regulating the signaling events in BMP pathway. Cell and Tissue Research2015;361(3):723–731. doi: 10.1007/s00441-015-2154-3. [PubMed] [Cross Ref]
28. Rahman M. S., Akhtar N., Jamil H. M., Banik R. S., Asaduzzaman S. M. TGF-?/BMP signaling and other molecular events: regulation of osteoblastogenesis and bone formation. Bone Research2015;3 doi: 10.1038/boneres.2015.5.15005 [PMC free article] [PubMed] [Cross Ref]
29. Chen G., Deng C., Li Y.-P. TGF-? and BMP signaling in osteoblast differentiation and bone formation. International Journal of Biological Sciences2012;8(2):272–288. doi: 10.7150/ijbs.2929.[PMC free article] [PubMed] [Cross Ref]
30. Fu Y.-C., Lin C.-C., Chang J.-K., et al. A novel single pulsed electromagnetic field stimulates osteogenesis of bone marrow mesenchymal stem cells and bone repair. PLOS ONE2014;9(3) doi: 10.1371/journal.pone.0091581.e91581 [PMC free article] [PubMed] [Cross Ref]
31. De Mattei M., Caruso A., Traina G. C., Pezzetti F., Baroni T., Sollazzo V. Correlation between pulsed electromagnetic fields exposure time and cell proliferation increase in human osteosarcoma cell lines and human normal osteoblast cells in vitro. Bioelectromagnetics1999;20(3):177–182. [PubMed]
32. Chang K., Chang W. H.-S. Pulsed electromagnetic fields prevent osteoporosis in an ovariectomized female rat model: a prostaglandin E2-associated process. Bioelectromagnetics2003;24(3):189–198. doi: 10.1002/bem.10078. [PubMed] [Cross Ref]
33. Jing D., Cai J., Shen G., et al. The preventive effects of pulsed electromagnetic fields on diabetic bone loss in streptozotocin-treated rats. Osteoporosis International2011;22(6):1885–1895. doi: 10.1007/s00198-010-1447-3. [PubMed] [Cross Ref]
34. Jing D., Shen G., Huang J., et al. Circadian rhythm affects the preventive role of pulsed electromagnetic fields on ovariectomy-induced osteoporosis in rats. Bone2010;46(2):487–495. doi: 10.1016/j.bone.2009.09.021. [PubMed] [Cross Ref]
35. Guo L., Zhao R. C. H., Wu Y. The role of microRNAs in self-renewal and differentiation of mesenchymal stem cells. Experimental Hematology2011;39(6):608–616. doi: 10.1016/j.exphem.2011.01.011. [PubMed] [Cross Ref]
36. Pan A., Chang L., Nguyen A., James A. W. A review of hedgehog signaling in cranial bone development. Frontiers in Physiology2013;4, article 61 doi: 10.3389/fphys.2013.00061.[PMC free article] [PubMed] [Cross Ref]
37. Jing D., Li F., Jiang M., et al. Pulsed electromagnetic fields improve bone microstructure and strength in ovariectomized rats through a Wnt/Lrp5/?-catenin signaling-associated mechanism. PLoS ONE2013;8(11) doi: 10.1371/journal.pone.0079377.e79377 [PMC free article] [PubMed] [Cross Ref]
38. Petecchia L., Sbrana F., Utzeri R., et al. Electro-magnetic field promotes osteogenic differentiation of BM-hMSCs through a selective action on Ca2+-related mechanisms. Scientific Reports2015;5, article 13856 doi: 10.1038/srep13856. [PMC free article] [PubMed] [Cross Ref]
39. Vimalraj S., Selvamurugan N. MicroRNAs expression and their regulatory networks during mesenchymal stem cells differentiation toward osteoblasts. International Journal of Biological Macromolecules2014;66:194–202. doi: 10.1016/j.ijbiomac.2014.02.030. [PubMed] [Cross Ref]
40. Li H., Yang F., Wang Z., Fu Q., Liang A. MicroRNA-21 promotes osteogenic differentiation by targeting small mothers against decapentaplegic 7. Molecular Medicine Reports2015;12(1):1561–1567. doi: 10.3892/mmr.2015.3497. [PubMed] [Cross Ref]
41. Schramke V., Allshire R. Hairpin RNAs and retrotransposon LTRs effect RNAi and chromatin-based gene silencing. Science2003;301(5636):1069–1074. doi: 10.1126/science.1086870. [PubMed] [Cross Ref]
42. Ono M., Yamada K., Avolio F., Afzal V., Bensaddek D., Lamond A. I. Targeted knock-down of miR21 primary transcripts using snoMEN vectors induces apoptosis in human cancer cell lines. PLoS ONE2015;10(9) doi: 10.1371/journal.pone.0138668.e0138668 [PMC free article] [PubMed] [Cross Ref]
43. Jiao G., Pan B., Zhou Z., Zhou L., Li Z., Zhang Z. MicroRNA-21 regulates cell proliferation and apoptosis in H2O2-stimulated rat spinal cord neurons. Molecular Medicine Reports2015;12(5):7011–7016. doi: 10.3892/mmr.2015.4265. [PubMed] [Cross Ref]
44. He X., Eberhart J. K., Postlethwait J. H. MicroRNAs and micromanaging the skeleton in disease, development and evolution. Journal of Cellular and Molecular Medicine2009;13(4):606–618. doi: 10.1111/j.1582-4934.2009.00696.x. [PMC free article] [PubMed] [Cross Ref]
45. He L., Yang N., Isales C. M., Shi X.-M. Glucocorticoid-induced leucine zipper (GILZ) antagonizes TNF-? inhibition of mesenchymal stem cell osteogenic differentiation. PLoS ONE2012;7(3) doi: 10.1371/journal.pone.0031717.e31717 [PMC free article] [PubMed] [Cross Ref]
46. Yano M., Inoue Y., Tobimatsu T., et al. Smad7 inhibits differentiation and mineralization of mouse osteoblastic cells. Endocrine Journal2012;59(8):653–662. doi: 10.1507/endocrj.EJ12-0022. [PubMed][Cross Ref]
47. Jeong Kim Y., Jin Hwang S., Chan Bae Y., Sup Jung J. MiR-21 regulates adipogenic differentiation through the modulation of TGF-? signaling in mesenchymal stem cells derived from human adipose tissue. Stem Cells2009;27(12):3093–3102. doi: 10.1002/stem.235. [PubMed] [Cross Ref]
48. Zhang B. G., Li J. F., Yu B. Q., Zhu Z. G., Liu B. Y., Yan M. microRNA-21 promotes tumor proliferation and invasion in gastric cancer by targeting PTEN. Oncology Reports2012;27(4):1019–1026. doi: 10.3892/or.2012.1645. [PMC free article] [PubMed] [Cross Ref]
49. Kim Y. J., Hwang S. H., Cho H. H., Shin K. K., Bae Y. C., Jung J. S. MicroRNA 21 regulates the proliferation of human adipose tissue-derived mesenchymal stem cells and high-fat diet-induced obesity alters microRNA 21 expression in white adipose tissues. Journal of Cellular Physiology2012;227(1):183–193. doi: 10.1002/jcp.22716. [PubMed] [Cross Ref]
50. Jonason J. H., Xiao G., Zhang M., Xing L., Chen D. Post-translational regulation of Runx2 in bone and cartilage. Journal of Dental Research2009;88(8):693–703. doi: 10.1177/0022034509341629.[PMC free article] [PubMed] [Cross Ref]
51. Selvamurugan N., Shimizu E., Lee M., Liu T., Li H., Partridge N. C. Identification and characterization of Runx2 phosphorylation sites involved in matrix metalloproteinase-13 promoter activation. FEBS Letters2009;583(7):1141–1146. doi: 10.1016/j.febslet.2009.02.040. [PMC free article] [PubMed] [Cross Ref]
52. Choi Y. H., Kim Y.-J., Jeong H. M., Jin Y.-H., Yeo C.-Y., Lee K. Y. Akt enhances Runx2 protein stability by regulating Smurf2 function during osteoblast differentiation. FEBS Journal2014;281(16):3656–3666. doi: 10.1111/febs.12887. [PubMed] [Cross Ref]
53. Ge C., Cawthorn W. P., Li Y., Zhao G., Macdougald O. A., Franceschi R. T. Reciprocal control of osteogenic and adipogenic differentiation by ERK/MAP kinase phosphorylation of Runx2 and PPAR?transcription factors. Journal of Cellular Physiology2016;231(3):587–596. doi: 10.1002/jcp.25102.[PMC free article] [PubMed] [Cross Ref]
54. Wang C. Y., Yang S. F., Wang Z., et al. PCAF acetylates Runx2 and promotes osteoblast differentiation. Journal of Bone and Mineral Metabolism2013;31(4):381–389. doi: 10.1007/s00774-013-0428-y.[PubMed] [Cross Ref]
55. Vishal M., Ajeetha R., Keerthana R., Selvamurugan N. Regulation of Runx2 by histone deacetylases in bone. Current Protein & Peptide Science2016;17(4):343–351. doi: 10.2174/1389203716666150623104017. [PubMed] [Cross Ref]
56. Kaneki H., Guo R., Chen D., et al. Tumor necrosis factor promotes Runx2 degradation through up-regulation of Smurf1 and Smurf2 in osteoblasts. Journal of Biological Chemistry2006;281(7):4326–4333. doi: 10.1074/jbc.M509430200. [PMC free article] [PubMed] [Cross Ref]