Static Magnetic Fields Enhance Skeletal

Muscle Differentiation In Vitro by Improving

Myoblast Alignment

Dario Coletti,1,2 Laura Teodori,3* Maria C. Albertini,4 Marco Rocchi,5

Alessandro Pristera,1,6 Massimo Fini,7 Mario Molinaro,1,2 Sergio Adamo1,2

� AbstractStatic magnetic field (SMF) interacts with mammal skeletal muscle; however, SMFeffects on skeletal muscle cells are poorly investigated. The myogenic cell line L6, anin vitro model of muscle development, was used to investigate the effect of a 80 6 mTSMF generated by a custom-made magnet. SMF promoted myogenic cell differentia-tion and hypertrophy, i.e., increased accumulation of actin and myosin and formationof large multinucleated myotubes. The elevated number of nuclei per myotube wasderived from increased cell fusion efficiency, with no changes in cell proliferationupon SMF exposure. No alterations in myogenin expression, a modulator of myogen-esis, occurred upon SMF exposure. SMF induced cells to align in parallel bundles, anorientation conserved throughout differentiation. SMF stimulated formation of actinstress-fiber like structures. SMF rescued muscle differentiation in the presence ofTNF, a muscle differentiation inhibitor. We believe this is the first report showing thatSMF promotes myogenic differentiation and cell alignment, in the absence ofany invasive manipulation. SMF-enhanced parallel orientation of myotubes is relevantto tissue engineering of a highly organized tissue such as skeletal muscle. SMF rescueof muscle differentiation in the presence of TNF may have important therapeuticimplications. ' 2007 International Society for Analytical Cytology

� Key termsskeletal muscle differentiation; muscle homeostasis; static magnetic field; tissue engi-neering; confocal microscopy; flow cytometry; cell culture

FEW studies exist on the effects of static magnetic fields (SMFs) on living cell and tis-

sues, compared with those of extremely low-frequency magnetic fields. Most of these

studies dealt with the potentially genotoxic or oncogenic effects SMF may play on eu-

karyotic cells and have basically excluded these possibilities [reviewed by Miyakoshi

(1)]. We have showed that 6 mT SMFs, beside not inducing cell damage, reduce the

extent of damage-induced apoptosis in glioblastoma cells (2).

In spite of contrasting evidence that SMF can provide therapeutically useful mus-

culoskeletal pain relief (3,4), both electromagnetic and magnetic fields have been widely

used in clinical practice regarding the skeletal muscular apparatus [reviewed by Pilla

(5)]. Interestingly, magnetotherapy has been reported to yield better results when com-

bined with kinesitherapy treatment, for instance aimed to counteract quadriceps muscle

atrophy in humans (6). The direct widespread use of SMF for therapeutic application

regarding the musculoskeletal apparatus is in sharp contrast with the paucity of data

regarding the mechanisms underlying the effects of SMF on skeletal musculature.

As an alternative to therapy, skeletal muscle tissue engineering is an emerging

approach to provide functional and aesthetic restoration of lost muscle tissue. Among

the major challenges to be addressed before reaching in vitro preassembled tissue trans-

Cytometry Part A � 71A: 846�856, 2007

1Department of Histology and MedicalEmbryology, Sapienza University, Rome,Italy2Interuniversity Institute of Myology3Biotec-Med, ENEA-Casaccia, Rome,Italy4Institute of Biological Chemistry,University of Urbino, Urbino, Italy5Institute of Biomathematics, Universityof Urbino, Urbino, Italy6Pharmacology Department, UniversityCollege London, London, United Kingdom7Department of Medical Sciences, Centrefor Clinical and Basic Research, IRCCSSan Raffaele, Roma, Italy

Received 2 March 2007; RevisionReceived 14 May 2007; Accepted 4 July2007

Grant sponsor: Sapienza University(Progetti di Ateneo); Grant number:C26A055711; Grant sponsor: AFM; Grantnumber: 11788; Grant sponsor: ASI –OSMA 2006; Grant sponsor: SapienzaUniversity; Grant number: Grandiattrezzature 2003.

*Correspondence to: Laura Teodori,Biotec-med, ENEA-Casaccia, ViaAnguillarese, Roma, Italy.

Email: teodori@casaccia.enea.it

Published online 10 August 2007 inWiley InterScience (www.interscience.wiley.com)

DOI: 10.1002/cyto.a.20447

© 2007 International Society forAnalytical Cytology

Original Article

fer in animals (7) is the induction of terminal differentiation

in 3D structures capable of contraction in response to electri-

cal stimulation. In vivo skeletal muscle is composed of bun-

dles of highly oriented and dense muscle fibers, multinu-

cleated syncytia derived from fusion of hundreds of myo-

blasts. The muscle fibers in native skeletal muscle are closely

packed together in an extracellular three-dimensional matrix

to form a highly organized tissue. Therefore, key features of

the muscle tissue are high cell density and precise cellular

orientation to generate longitudinal contraction. The impor-

tance of controlling cell orientation during muscle formation

arises from studies showing how the orientation of nanofi-

bers or micropatterned molds, used as a substratum for cell

adhesion, can influence myogenic cell behavior. In particular,

Huang et al. showed that the parallel orientation of nano-

fibers used as scaffold for myogenic cell cultures promotes

skeletal morphogenesis in parallel oriented myotubes of

increased length in respect to controls (8). In this context,

developing new tools to control cell orientation and behavior

are of great relevance.

SMF does affect cell shape and orientation. Strong (8 T)

SMF stimulates bone formation and regulates its orientation,

both in vitro and in vivo, without affecting cell proliferation

(9). Biological effects of 6 mT SMF (a moderate intensity in

respect to the examples mentioned earlier) on cell cultures,

such as modifications of cell shape, surface, and cytoskeleton,

have been recently summarized (1,10). Orientation of both

adherent and nonadherent cells, such as erythrocytes, sperm,

glioblastoma, and fibroblasts, can be affected by exposure to

SMF, with effects varying accordingly to the cell type and in-

tensity of the MF (1). Consequently, the deposition and orien-

tation of the extracellular matrix can also be affected by SMF,

as in the case of collagen fibrils produced by SMF-oriented

fibroblasts (11).

There are few, scattered studies on the effects SMF plays

on skeletal muscle, describing a wide range of processes from

molecular activities to organ physiology. In vitro experiments

have shown that dc-generated SMFs in the range of 0–200 mTaccelerate Ca21/calmodulin-dependent myosin light chain

phosphorylation (12). The effects of 20 mT SMF on specific

ion transporters resulted in altered bioelectrical and biome-

chanical properties of rat diaphragm muscle (13). Stronger

(650 mT) SMFs affected the functional properties of bullfrog

neuromuscular preparations (14). In addition, the introduc-

tion of magnetic microparticles into cultured myoblasts, fol-

lowed by exposure of the cells to a 50 mT SMF, promoted

myotube formation (15).

Here we show that exposure to 80 mT SMF stimulates

myogenic differentiation in the absence of other manipula-

tions, by affecting myoblast orientation through remodeling

of stress fiber-like actin filaments.

MATERIALS AND METHODS

Cell Cultures

L6 cells of the E9 subclone (16–18) were seeded at the

density of 15,000/cm2 in DMEM supplemented with 50 mg/ml

gentamicin and 10% heat-inactivated foetal bovine serum

(GM, Sigma-Aldrich, St. Louis, MO). To induce differentia-

tion, the cells were shifted to a medium containing 1% heat-

inactivated fetal bovine serum (DM). The cells were always

cultured in 35-mm plastic Petri dishes, either directly on plas-

tic or on gelatin-coated coverslips (for the confocal experi-

ments only). The cell cultures were stopped at various time-

points. The time-points are expressed in days of culture (d)

elapsed at the time of stop. They include 24 h in GM (1 day)

followed or not by up to 4 days in DM (2d – 5d), as indicated

in the figure (for time-course experiments) or in the figure

legend.

SMF Exposure

SMF of known flux densities were produced by custom-

made 4 cm 3 4 cm Neodymium magnetic plaques. The mag-

netic plaques (one per Petri dish) placed underneath the dish

at 1-mm distance from the cell monolayer in a custom-made

device, which kept them suspended in the incubator at >10 cm

of distance from any metallic surface to avoid distortion of

the field. Up to three magnetic plaques (one for each Petri

dish) were simultaneously used in each independent replicate,

placed at >10 cm of distance from each other (at which dis-

tance the magnetic field of the magnets is undetectable). The

position of the samples inside the incubator was randomly

chosen. The fields were axial, with the magnetic North (the

pole of the magnet which would be repulsed by the Earth’s

North magnetic pole) vector crossing the cells. Magnitude and

homogeneity of the magnetic field of each plaque set at 80 65 mT (1 mT 5 10 G) were checked by means of a gaussmeter

(Hall-effect Gaussmeter, GM04 Hirst Magnetic Instruments,

UK). The latter was also used to measure the magnetic field

inside the incubator (0.5 6 0.02 mT), which must be consid-

ered as a background for all the cell cultures used in this study,

including controls, and added to the 80 mTmagnetic field for

the exposed cells. Magnets do not produce temperature varia-

tion. SMF exposure started immediately after cell seeding and

continued throughout the duration of the experiments. Con-

trol samples (three for each independent replicate) were

simultaneously grown and induced to differentiate in identical

experimental conditions, with the exception of the presence of

the magnets (i.e. in the same incubator as the cell cultures

placed onto the magnetic plaques, but placed at >10 cm dis-

tance from any magnet).

Cytofluorescence Experiments

To obtain micrographs of different cell compartments

and structures, the cells were fixed with 4% formaldehyde buf-

fered solution for 10 min at RT, washed twice in PBS, and

permeabilized with 0.1% Triton X-100 buffered solution for

15 min at RT. Aspecific binding sites were blocked by incubation

with 1% BSA for 30 min, followed by

Actin staining: By either Phalloidin-Alexa 546 (collected

by the red fluorescence filter, Molecular Probes, Eugene, OR)

or FITC-conjugated Phalloidin (collected by the green fluores-

cence filter, Molecular Probes) diluted 1:50 with 1% BSA in

PBS for 20 min at RT;

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ORIGINAL ARTICLE

Nuclei staining: By TO-PRO (Molecular Probes) diluted

1:2,000 in PBS for 10 min at RT or 0.5 mg/ml Hoechst 33342

(Sigma) in PBS for 2 min at RT, or 2 mg/l DAPI (2) (Serva,

Heidelberg, Germany), as indicated in figure legends;

Myogenin staining: By F5D (hybridoma supernatant, Hy-

bridoma Bank, Iowa University, IO) diluted 1:50 with 1% BSA

in PBS for 60 min at RT, immunolocalized by an anti-mouse

Alexa 546-conjugated secondary antibody diluted 1:100 with

1% BSA in PBS for 45 min; cell shape was highlighted by

FITC-conjugated phalloidin staining. Cytoplasm staining was

performed on ethanol-fixed cells incubated with a 10 mg/l Sul-

fur-Rhodamin solution (SR 101, Serva). Photomicrographs

were obtained with a Carl Zeiss Axioplan fluorescence micro-

scope equipped with a CoolSnap cf CCD camera. Confocal

images were obtained with a Leica laser scanning microscope

TCS SP2 and acquired with Leica Confocal Software (Leica,

GE).

Flow Cytometry

The cellular monolayers were dispersed by incubation

with 0.05% trypsin for 5 min, followed by enzyme inactiva-

tion, and fixed in 70% ethanol. An aliquot of cells suspension

was then stained with 2 mg/l DAPI (Serva), analyzed as

described elsewhere (2). DNA content was measured with a

Partec PAS III flow cytometer (PARTEC, Muenster Germany).

May-Grumwald and Giemsa Staining

May-Grumwald and Giemsa staining was performed in

sequence as described elsewhere (19). Both fusion efficiency

and myotube size distribution were measured on May-Grum-

wald and Giemsa stained cultures, after 5 days of culture (24 h

in GM followed by 96 h in DM, the latter completely replaced

once). Myoblast fusion was evaluated as follows: cells were

considered fused when at least three nuclei were present in

each myotube. The ratio between the nuclei in the myotubes

and the total number of nuclei per microscope field (at least

10 randomly selected fields/dish) was expressed as the percent-

age of fusion. To evaluate hypertrophy effects, the size of the

myotubes was measured as follows: fused cells were counted,

grouped according to the number of nuclei in each myotube,

and expressed as percent of total.

Myoblast orientation was measured on May-Grumwald

and Giemsa stained cultures after 1 day (24 h in GM) or

3 days (24 h in GM followed by 48 h in DM) of culture.

Morphometric Analysis

For actin quantitative analysis, images of 10 randomly

chosen microscope fields were acquired at the confocal

microscope. Postprocessing of the image was performed using

Adobe Photoshop and Scion Image Softwares (the latter, a

software originally developed with the name of NIH Image

at NIH, Bethesda, is freely downloadable at http://www.

scioncorp.com/pages/scion_image_windows.htm). Modifica-

tions were the same for all images and consisted in a conver-

sion to gray scale, followed by the measure of the mean fluo-

rescence intensity calculated on the whole field. The values

were normalized over the number of nuclei in the field. The

values deriving from the 10 sampled fields were averaged and

the result was considered representative of one sample. At least

four independent replicates were performed in triplicate (i.e.

three Petri dishes) for each time-point and averaged. The

values of actin content were expressed as percent of control

(i.e. the mean actin content of the samples cultured for 2 days

in the absence of SMF) to normalize variability among experi-

ments.

Western Blot Analysis

Triplicate experiments were pooled to obtain a sufficient

amount of cell protein extracts for WB analysis. Cells were

scraped and homogenized in 0.1 ml RIPA buffer (10 mM Tris-

HCl, pH 7.5, 10 mM EDTA, 0.5 M NaCl, 0.5% NaDoc, 1%

NP40) supplemented with Protease Inhibitor Cocktail (Roche

Boehringer, Mannheim, Germany) by passage through a 16 G

needle. Protein content was measured using the BCA Protein

Assay (Pierce, Rockford, IL). Proteins (10 mg) were separated

by SDS-PAGE and transferred electrophoretically to nitrocel-

lulose membrane (Amersham, Piscataway, NJ). Nonspecific

binding was blocked in TBST containing 1% bovine serum al-

bumin (Sigma) (BB) for 1 h and then probed with MF20 anti-

body against myosin heavy chain (MHC; Hybridoma Bank)

diluted 1:50 in BB. After washing in TBST, blots were in-

cubated with goat anti-mouse HRP-conjugated (Biorad, Her-

cules, CA) diluted 1:20,000 in BB and antibody binding was

detected using Super Signal West Pico Chemiluminescent Sub-

strate (Pierce).

Cell Direction Measurement and Statistical Analysis

Microscope images of May-Grumwald and Giemsa

stained cells were used to study cell direction. A segment

crossing the largest part of each cell was added and used to an-

alyze the distribution of cell direction by circular statistics, as

previously described (20). Such technique is recently gaining

attention for analyzing periodic variables such as angle distri-

bution. The Rayleigh test is used to determine whether the

population from which the sample is drawn differs signifi-

cantly from isotropy (also defined as stochastic distribution),

thus providing evidence of whether statistical prove of aniso-

tropy (i.e. one-sidedness) exists. The Rayleigh test uses the

mean vector length r as a statistical test; r can range from 0 (in

case of a perfect isotropy, i.e., of existence of multiple cell

directions) to 1 (in case of a perfect anisotropy, i.e., the exis-

tence of a preferred-aligned-cell direction). Additional details

on this approach have been recently described (20). The cutoff

between anisotropy (aligned) and isotropy (stochastic) cell

direction distribution is fixed at Rayleigh test; P 5 0.05: below

being aligned distribution, above being stochastic distribution.

To evaluate whether it was possible to merge images of

cells coming from different experiments developed in the

same condition, the Kuiper test (an extension of Kolmogorov–

Smirnov test to circular data) was performed as previously

described (20). Kuiper’s test is used to test whether two sam-

ples differ significantly from each other in their circular distri-

butions. It works by comparing the sample distribution func-

tions. The maximum of the difference between the sample

848 Enhanced Muscle Differentiation by Magnetic Field

ORIGINAL ARTICLE

distribution functions is used as a statistic test. The larger the

deviation between these distribution functions, the more likely

a significant difference occurs between the samples. Five inde-

pendent experiments for each item were performed, and for

each image the number of oriented bars inserted was at least

50, with a total of over 250 cells analyzed for each exposure

condition.

ANOVA was used to compare cell fusion index means

among different treatments, and Tukey LSD was used for post

hoc analysis

RESULTS

SMF Promotes Myogenic Differentiation

and Hypertrophy and Counteracts TNF

Effects on Myogenesis

The conventional approach to induce myogenic cell lines

to differentiate consists in lowering the concentration of the

serum in the culture medium (thus called differentiation

medium, DM). We compared this standard procedure with

the exposure to a 80 mT SMF for the whole culture time,

using L6 myogenic cells as a model of myogenic differentia-

tion. An hallmark of myogenic differentiation is cell fusion

into multinucleated myotubes. About half (45% 6 6.7%) of

the cells cultured in DM were induced to differentiate and

fused into myotubes, characterized by a highly organized actin

cytoskeleton and at least three nuclei in the same cytoplasm

(Figs. 1A and 1B). A significant increase in the number of cells

fused into multinucleated myotubes (65% 6 3.6%) was

observed in cells cultured in DM and simultaneously exposed

to SMF throughout the differentiation process (Figs. 1A and

1B). The increased fusion efficiency observed upon cell expo-

sure to SMF was associated to a striking hypertrophic effect,

highlighted by the formation of myotubes containing more

than 100 nuclei, which did not occur in the absence of SMF

(Fig. 1B, graph). In addition SMF induced a significant

decrease in the smallest class of myotubes, and a general shift

of myotube size toward classes of myotubes containing several

dozens of nuclei, poorly represented in the control cultures

(Fig. 1B left panel: 2SMF). We noted that only a specific

orientation of the SMF was effective in inducing the effects

described earlier (Fig. 1B), that is the use of the magnet with

the North magnetic pole (the pole that would be repulsed by

the earth’s north magnetic pole) oriented upward, toward the

cell culture (Fig. 1B, + !SMF). This shows that SMF effects on

muscle differentiation and hypertrophy are highly specific

and, therefore, the SMF oriented with the North magnetic

pole up are referred to as SMF throughout this paper.

We verified whether the increased fusion index and hy-

pertrophy could be due to increased cell number because of

proliferation in the presence of SMF. SMF did not modify the

distribution of cell populations in the different phases of the

cell cycle, as shown by cytofluorometric analysis of the DNA

content of cultures in proliferation medium exposed to SMF

as compared to the control (Fig. 1C). These data suggest that

the SMF effect is purely promyogenic and occurs in the ab-

sence of a proliferative effect.

Since the activation of pathways leading to hypertrophy is

considered as a promising tool to counteract cytokine-induced

muscle atrophy and wastage, we tested the effect of SMF

exposure in the presence of TNF. To address this issue, L6 cells

were exposed or not to SMF since seeding, and TNF, or buffer,

treatment was started 2 h later and continued throughout the

culture. Figure 2A (left panels) depicts the well-known TNF

negative effects on myogenic cell cultures (21,22) and recapi-

tulated in our experimental conditions: few, atrophic myo-

tubes were observed in addition to a majority of elongated,

but unfused myoblasts when the cell cultures were treated

with TNF. In contrast, SMF determined an increased recruit-

ment of nuclei (i.e. cells) to myotube formation (Fig. 2A, right

upper panel). In the presence of TNF, SMF effect was domi-

nant over TNF negative effect on myotube formation (Fig. 2A,

right lower panel). The effects of the treatments mentioned

earlier were quantified by measuring the percentage of fusion

as an index of myogenic differentiation. ANOVA showed that

the treatments significantly affected the percentage of fusion

of myogenic cultures (F5 19.223; df5 3; P\ 0.0001). In par-

ticular, TNF inhibited myogenic differentiation (P \ 0.05 by

Tukey HSD Test vs. untreated cultures), while SMF promoted

myogenic differentiation (P \ 0.01 by Tukey HSD Test vs.

untreated cultures). Importantly, in the presence of TNF, SMF

rescued the efficiency of myoblast fusion to the levels of cul-

tures exposed to SMF alone (P\ 0.01 by Tukey HSD Test vs.

untreated cultures).

Among the morphological and molecular hallmarks of

hypertrophic myotubes, nuclear rings and actin accumulation

have been reported (23). Actin content of the cell cultures (red

staining) is depicted in Figure 2C, where TNF is shown to

depress this phenomenon (lower left panel). In contrast,

SMF greatly enhances the formation of actin-rich myotubes,

both in the absence and in the presence of TNF (Fig. 2C,

right panels). In agreement, the presence of nuclear rings in

the myotubes is detectable in cultures exposed to SMF,

regardless of the presence of TNF (Fig. 2A). Thus, SMF not

only enhances muscle differentiation and hypertrophy but

also removes a block due to negative signals for myogenesis

(TNF).

SMF Promotes MHC Accumulation Without Altering

Myogenic Gene Expression

To investigate molecular mechanisms underlying the

effects of SMF on myogenic differentiation, we monitored the

expression of the transcription factor myogenin in the early

phases of differentiation. Upon differentiation, myogenin

appeared expressed and correctly localized in the nucleus of

both unexposed and SMF-exposed cell cultures (Fig. 3A). No

differences in the number of myogenin-expressing cells were

apparent between the two conditions, as opposed to the signif-

icant differences in terms of differentiation index (based on

the morphological feature of multinucleated myotubes, Figs. 1

and 2). The latter found a molecular confirmation in the

enhanced accumulation of the late differentiation marker

MHC in cells exposed to SMF as compared to the control

(Fig. 3B).

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ORIGINAL ARTICLE

Figure 1. SMF enhances muscle differentiation and hypertrophy. (A) L6 cells were cultured in the absence or in the continuous pre-

sence of a 80 mT SMF, as described in Materials and Methods. The cytoplasm and the cytoskeleton of the myotubes are highlighted by

Sulphur-Rodamine (red) and FITC-conjugated phalloidin (green), respectively. Nuclei are DAPI-stained (blue). The merged images

show that exposure to SMF (right panel) induces formation of hypertrophic myotubes characterized by the presence of nuclear rings,

elevated cytoplasm/nuclei ratio, and an elevated number of nuclei participating to the syncytia. (B) Representative photomicrographs

of May-Grumwald and Giemsa-stained L6 cell cultures. The cells were cultured for 5 days in the absence of SMF (upper left panel), in

the continuous presence of SMF oriented with the North magnetic pole toward the cell monolayer (1 › SMF, lower left panel), or in thecontinuous presence of SMF with the opposite orientation (1 fl SMF, upper right panel). The number of nuclei per myotube in the cul-ture conditions mentioned earlier was evaluated as follows: myotubes (i.e. structures containing at least three nuclei in the same cyto-

plasm) were counted, grouped according to the number of nuclei per myotube, and expressed as percent of the total number of myo-

tubes analyzed (graph embedded in the figure). The application of a SMF with the North magnetic pole facing toward the cells specifi-

cally induces hypertrophy of the myotubes and significantly reduces the presence of the smallest class myotubes. Shown are the

means 6 SEM from four independent replicates, *P\ 0.05, by Student’s t test vs. not exposed to SMF. (C) Fluorometric analysis ofDNA content by PI staining in myoblasts cultured for 1 day in the absence and presence of 80 mT SMF. The inset table shows the mean

cell cycle phase frequency 6 SEM of three different experiments. The SMF does not affect cell cycle, as shown by evaluation of the %

of cells in G1, S, or G2 phase.

850 Enhanced Muscle Differentiation by Magnetic Field

ORIGINAL ARTICLE

SMF Promotes Myoblast Alignment

Cell–cell alignment and contact precedes and affects the

formation of multinucleated syncytia during muscle differen-

tiation, both in vitro and in vivo (24). Therefore, we

performed a Rayleigh test to evaluate the degree of isotropy of

the cell orientation and its dependence on both culture time

and exposure to SMF (Fig. 4A). Single-cell orientation (high-

lighted by a bar) is modified when cells are cultured in DM for

2 additional days after the first day of culture in GM (Fig. 4A,

upper panels). SMF exerts a potent effect on cell alignment,

both at early (1 day) and late (3 days) cell culture times (Fig.

4A, lower panels). As demonstrated by two-way ANOVA, both

culture time and SMF exposure significantly affect cell orienta-

tion, and these two variables do interact (Tables 1 and 2).

Thus SMF promotes the alignment of myoblasts in a time-

dependent fashion.

A pivotal role in controlling cell orientation and motility

is played by the actin cytoskeleton (25). We therefore followed

cell shape and orientation changes by labeling stress fiber-like

actin filaments in myogenic cells by FITC-phalloidin staining.

As shown by the time course in Figure 4B, myogenic cells have

the tendency to contact each other by forming bundles that

ultimately fuse into multinucleated syncytia, which show a

seemingly stochastic orientation. In contrast, SMF induces an

Figure 3. SMF differentially affects early and late myogenic differ-

entiation markers. (A) Immunofluorescence analysis for myo-

genin in control and SMF-exposed cell cultures after 2 days of cul-

ture. Myogenin (left) is expressed in both conditions in myoblasts,

in which nuclei are highlighted by Hoechst staining (right). SMF

does not seem to increase the number of myogenin-expressing

cells. (B) Western Blot analysis of myosin heavy chain (MHC) in

muscle cells cultured for 5 days in the absence or in the con-

tinuous presence of a 80 mT SMF, as described in Materials and

Methods. A portion of the Ponceau Red-stained membrane is

shown as a loading control. MHC content is increased in cell cul-

tures exposed to SMF, when compared with the control.

Figure 2. SMF rescues differentiation in the presence of TNF. (A)

Representative photomicrographs of May-Grumwald and

Giemsa-stained cell cultures. L6 cells were exposed or not to 80

mT SMF since seeding, and 5 mg/ml TNF, or buffer, treatment wasstarted 2 h later and continued throughout the culture, as indi-

cated, for 5 days. (B) Differentiation is quantitatively expressed as

% of fusion, i.e. the percent of cells sharing the same cytoplasm

(considering only myotubes with at least three nuclei) over the

total number of cells. While TNF inhibits muscle differentiation,

SMF increases the differentiation efficiency of myogenic cell cul-

tures, both in the absence and in the presence of TNF. Mean 6SEM are shown, from at least three independent replicates per-

formed in triplicate (i.e. three Petri dishes). *P \ 0.05, by Stu-

dent’s t test vs. not exposed to SMF. (C) Same cultures as men-tioned earlier, at higher magnification. The cytoskeleton is here

labeled by TRITC-conjugated phalloidin (red) while the nuclei are

Hoechst-stained (blue). SMF exposure promotes actin accumula-

tion in the myotubes, both in the absence and presence of TNF.

Cytometry Part A � 71A: 846�856, 2007 851

ORIGINAL ARTICLE

early and efficient alignment among the differentiating myo-

blasts, leading to ordered populations of myotubes with a par-

allel orientation (Fig. 4B).

SMF Enhances Actin Accumulation and Stress-Like

Fiber Formation

To analyze in detail the regulation of actin cytoskeleton

by SMF, we performed a confocal microscope analysis, which

allowed quantification of the fluorometric signal obtained by

TRITC-labeled phalloidin staining of the actin filaments. We

noted that SMF exerts a precocious effect on stress fiber-like

actin filaments: in comparison to control cells, actin filaments

appeared more pronounced and elongated in cells exposed to

SMF already after 1 day of culture (Fig. 5A). With an addi-

tional day of culture, control myoblasts showed a more evi-

dent labeling for actin, but the cell shape and size did not

Figure 4. MF affects cell orientation. (A) Morphological analysis of L6 cells cultured for 1 and 3 days, in the absence or in the continuous

presence of SMF. Cells were stained with May-Grumwald and Giemsa, and the major axis of each cell identified with a bar. The bar orien-

tation was used for Rayleigh test as described in Material and Methods. The representative photomicrographs show that both time and

SMF induce cell alignment, and that their effects are additive. (B) Cells cultured as mentioned earlier from 1 to 5 days were stained with

FITC-conjugated phalloidin staining (green). Nuclei were counterstained with TO-PRO (blue). SMF induces parallel orientation of both

cytoskeletal filaments and differentiating cells, ultimately resulting in oriented myotubes.

852 Enhanced Muscle Differentiation by Magnetic Field

ORIGINAL ARTICLE

resemble that of cells exposed to SMF, either for 1 or 2 days.

Myoblasts cultured for 2 days upon SMF exposure appeared

elongated, of larger size, and aligned, conditions associated to

an evident actin cytoskeleton (Fig. 5A).

To quantify the actin content of the cells in the different

culture conditions, we recorded the fluorescence intensity in

randomly chosen fields and normalized these values for cell

number. We noted that, with the progression of the culture

time (i.e. differentiation), myogenic cells accumulate actin and

that SMF exposure significantly increases actin content at

all the time-point examined (Fig. 5B). These data suggested

that the cells accumulated more actin in the course of differen-

tiation when exposed to SMF.

A qualitative analysis of actin filaments at single cell level

was performed on randomly chosen single cells from the dif-

ferent culture conditions, stained with TRITC-phalloidin.

Analysis of representative photomicrographs was performed

along a line (a and b) perpendicular to the main axis of the

cell (Fig. 5C, upper panels). The fluorescence intensity

recorded along this line (reported in Figure 5C, lower panels)

highlighted the presence of actin stress fiber-like filaments as

fluorescence peaks. The fixed image section thickness of the

confocal images excluded any variation of signal due to altered

cell thickness. The signal distribution corresponding to single

intracellular actin stress fiber-like filaments yielded higher and

wider peaks in cells exposed to SMF as compared to the

threshold level in the control (Fig. 5C, lower panels).

DISCUSSION

Animals have evolved in Earth’s natural magnetic field,

which ranges from 0.03 to 0.07 mT, depending on the loca-

tion. Besides, we are exposed to SMFs of residential and

anthropic origin. Such environmental exposure ranges from

intensities considered relatively low (e.g. 0.02 mT under direct

current transmission lines) to very high (e.g. up to 4,000 mT

during magnetic resonance imaging) in respect to Earth’s

background. Relatively few studies have focused on the inter-

action between SMF and living cells.

Skeletal muscle can be a target of SMF in vivo. SMF

increases norepinephrine levels in the gastrocnemius of rats

(26) and stimulate Na1-K1 ATPase and Ca21 ATPase activ-

ities (13). In contrast, no information is available on the effect

SMF exerts on skeletal muscle cells in vitro, besides a single

report where a MF is shown to accelerate myogenic differen-

tiation (15). In the latter, magnetic microparticles were intro-

duced into the cytoplasm of cultured myoblasts and the cells

were cultured in a 50 mT SMF. The authors reported that dif-

ferentiation marker expression and differentiated morphology

were detectable at earlier time-points in the cells containing

the magnetic microparticles and exposed to the magnetic field

(as opposed not only to control cultures but also to cultures

not pretreated with the microparticles and exposed to the

magnetic field). The same group also reported that magnetic

microparticles introduced into osteoblasts make this cell type

to differentiate in response to a magnetic field (27). They

reported that the cells appear elongated along the axis of the

magnetic poles and that the physical stress by magnetic force

seems to accelerate the differentiation of these cells (27). In

contrast, here we show for the first time, to our knowledge,

that SMF per se promotes differentiation of myogenic cells, in

the absence of invasive and irreversible manipulations (such as

introducing magnetic particles in the cells to ‘‘sensitize’’ them

to the magnetic field). It is important to mention that myo-

blasts are affected by an orthogonal magnetic field (with the

North pole of the magnet facing toward the cells) and they do

not align along the lines of the magnetic field. This would sug-

gest that ‘‘native’’ myoblasts respond to SMF exposure differ-

ently from cells inoculated with magnetic microparticles. It is

worth mentioning, though, that the direction of the magnetic

field is important, since we did not observe the responses

described here when the axis of the magnetic field was differ-

ently oriented. This phenomenon suggests specific effects on

cell behavior by SMF, depending on its vectorial dimension.

We do not know the mechanisms underlying this specific cel-

lular response. In addition, it will be necessary to determine

the range of static magnetic flux densities that are effective in

this phenomenon. To this purpose, a full control over mag-

netic field density and homogeneity is a critical issue. Magnets

to generate SMF of different magnetic flux densities are under

development.

The fact the myoblasts are sensitive to SMF without addi-

tional procedure and differentiate into hypertrophic myotubes

Table 1. Explorative analysis by Rayleigh test

TIME 1SMF 2SMF

1 day n 5 756 n 5 799

r 5 0.061 r5 0.023

P 5 0.062 P 5 0.661

2 days n 5 860 n 5 812

r 5 0.078 r5 0.021

P 5 0.005 P 5 0.702

Single cells were cultured for 1 and 2 days, corresponding to

1 day in GM (1 d) and 1 day in GM followed by 1 day in DM (2 d),

in the absence or presence of SMF. These objects were identified

and used for Rayleigh test to screen for orientation as described

in Material and Methods. The mean vector length (r) can range

from 0 (in case of a perfect isotropy, i.e., of existence of multiple

cell directions) to 1 (in case of a perfect anisotropy, i.e., the exis-

tence of a preferred-aligned-cell direction). n 5 number of ana-

lyzed cells; r 5 mean vector length (statistic of Rayleigh test); P 5observed significance (P-value).

Table 2. ANOVA table

SOURCE OF VARIATION df F P

Due to treatment 1 8.821 0.003

Due to time 1 1.473 0.225

Due to interaction treatment3 time 1 9.565 0.002

Residual 3,223

Total 3,226

The values resulting from statistical analysis in Table 1 are

shown. df 5 degrees of freedom; F 5 ANOVA value; P 5 observed

significance (P-value).

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ORIGINAL ARTICLE

makes this approach intriguing for tissue engineering applica-

tions. One of the main goals for in vitro assembly of tissues

and organs is to obtain large amounts of the differentiated

tissue of interests. An additional task consists in finding

strategies to ameliorate both alignment and fusion of myogenic

cells, ultimately leading to myotubes more closely resembling

the elevated organization of skeletal muscle fibers in vivo (8).

In our experimental system, the final outputs of the exposure

Figure 5. SMF promotes cellular accumulation of actin. (A) L6 myoblasts were cultured on glass coverslips for 1 and 2 days in the absence

or in the continuous presence of SMF. Actin stress-fiber like structures were stained by Alexa 564-conjugated phalloidin (red). Nuclei were

counterstained with TO-PRO (blue). (B) Quantitative analysis of the fluorescence signal obtained in the conditions mentioned earlier was

performed as described in Materials and Methods. The total fluorescence actin staining upon the treatments indicated was averaged, nor-

malized for cell number and expressed as percent of control (% of ctrl, i.e. samples cultured for 2 days in the absence of SMF). SMF expo-

sure enhances actin accumulation during myogenic cell culture. The values represent the mean 6 SEM, from at least three independent

replicates performed in triplicate (i.e. three Petri dishes). *P\ 0.05 by Student’s t test. (C) Qualitative analysis of actin filament staining,performed by plot profile analysis of the TRITC-phalloidin signal in single cells grown for 1 day in the absence or in the presence of SMF.

Fluorescence staining of actin (upper panels) was measured along the axis represented by the red dotted line (a–b). The correspondingvalues of fluorescence intensity (in arbitrary units: a.u.) plotted against the photomicrograph width are reported (lower panels). The red

arrow points to the edge of the cell in both upper and lower panels. An arbitrary threshold to cutoff low actin fluorescence against high

actin fluorescence is highlighted by the dashed/dotted line. As shown by the fluorescence signal increase above the threshold, SMF expo-

sure enhances actin accumulation at single cell level, resulting in the formation of thicker and longer actin stress-fiber like structures with

respect to the control.

854 Enhanced Muscle Differentiation by Magnetic Field

ORIGINAL ARTICLE

of myogenic cell cultures to SMF field appear to be (1) the

production of myotubes with a higher efficiency, i.e. an

increase of about 50% in the number of cells recruited into

myotubes (Figs. 2A and 2B); (2) augmented myotube size, i.e.

hypertrophy (Figs. 1A and 1B); (3) increased production of

muscle proteins, i.e. actin (Fig. 2B); MHC (Fig. 3B); (4)

improved orientation of myotubes in parallel bundles (Fig. 4B).

Based on the aforementioned facts, myogenic cell culture

exposure to SMF may be seen as a tool to amplify the produc-

tion of highly oriented, hypertrophic myotubes, that would

represent the basic units of an in vitro assembled functional

muscle.

Cell cycle analysis showed superimposable distributions

of DNA content in control versus SMF-treated cell popula-

tions, suggesting that there are no major differences in the

proliferation rate in the two conditions. Thus, SMF effects on

terminal differentiation do not seem to depend on increased

cell density, a condition which favors fusion. In general, SMF

effects on cell proliferation appear modest as reviewed by Dini

and Abbro (10) and depending on the intensity of the mag-

netic stimulus: for instance, osteosarcoma cells proliferation

inversely correlates with the increase of the field (up to

10 mT) to which they are exposed (28).

Stimulation of muscle hypertrophy is currently seen as a

promising tool to counteract muscle atrophy in pathological

conditions, such as cachexia (29). Indeed cachexia is a severe

muscle-wasting syndrome associated to many chronic disease

states, and a therapy to treat cachexia is still lacking (30,31).

TNF plays a pivotal role in mediating cachexia, and strategies

against TNF holds promise to cope with this syndrome (32).

We have shown that TNF inhibits both muscle differentiation

in vitro and muscle regeneration after experimentally induced

muscle damage (21,22,33). Based on the aforementioned facts,

we used the model of cultured myoblasts exposed to TNF to

demonstrate that SMF exposure promotes myogenic differen-

tiation and rescues myogenic differentiation in the presence of

TNF (Fig. 2). This finding is relevant insomuch as SMF

appears to be able to sustain myogenesis in an environment

which mimics pathological conditions.

We noted that in DM the vast majority of the cells express

myogenin, one of the master gene controlling myogenic differ-

entiation in these cells, both in control and SMF-exposed cells

(Fig. 3A). Besides, myogenin correctly localizes in the nucleus,

a hallmark of its activation. This suggests that epigenetic

mechanisms triggered by SMF exposure account for the hyper-

trophic phenotype observed in SMF-treated cell culture (Figs. 1

and 2).

The changes in gene expression and cell morphology that

occur during myogenic differentiation must be coordinated

with each other in a spatiotemporal fashion; one way that this

might occur is through regulation of these processes by cell–

cell adhesion and resultant signaling (24). In vitro skeletal

muscle differentiation involves myoblast alignment, elonga-

tion, and cell–cell adhesion before plasma membrane fusion

(34). We thus quantified the isotropy of the cell cultures in

terms of cell orientation under SMF exposure (Fig. 4, and

Tables 1 and 2). The results indicate that there is a significant

effect of SMF in increasing the orientation (anisotropy) of the

cells, and that this enhancement is influenced by a time effect,

becoming more evident at 3 days of culture. A significant ho-

mogeneity of cell orientation upon SMF results in the forma-

tion of more aligned bundles of cells, which ultimately fuse

more efficiently forming larger myotubes (Fig. 4B).

The actin cytoskeleton plays a critical role in determining

cell shape and orientation. In particular, during myogenesis

stress fiber-like structures are considered to act as template for

the assembly of myofibrils (35). The importance of these

structures is highlighted by the fact that in L6 cells phospholi-

pase D-mediated remodeling of actin cytoskeleton is involved

in myogenic differentiation (36). Electromagnetic fields induce

changes in lipid second messengers, such as diacyglycerol and

phosphatidic acid, which are involved in phospholipase D-

mediated signaling (37). Confocal analysis of myoblasts cul-

tured in control conditions or upon SMF exposure reveals a

profound effect of SMF on actin stress fiber-like structures:

the actin-bound phalloidin staining appears stronger and

thicker in cells exposed to SFM, both in GM and in DM, sug-

gesting that SMF promotes the assembly of bigger filaments

(Figs. 5A and 5B). Morphometric analysis of the fluorescence

signal shows that the cellular actin content is increased upon

differentiation and that SMF stimulates this accumulation

(Fig. 5B). It is intriguing that the morphological changes fol-

lowing SMF exposure do not develop along an axis corre-

sponding to the lines of the magnetic field, rather they are

manifested in an orthogonal direction. We reported a different

behavior of glioblastoma cells in response to SMF exposure

(20). This cell type orientates its cytoskeleton and cellular

morphology along the axis corresponding to the lines of the

magnetic field. Myoblasts and glia cells display very different

morphologies, likely resulting in a molecular asymmetry

which may be differentially affected by SMF. The differential

response by other cell types to SMF is not in contrast with the

hypothesis that the magnetic field affects actin polymerization

in stress-like fibers in an indirect fashion. A possible candidate

to mediate SMF signaling to actin remodeling and differentia-

tion in L6 cells is phospholipase D. Another candidate intra-

cellular effector of muscle cell hypertrophy response to SMF is

calcineurin, since we have already reported that in glioblas-

toma cells SMF induce an increase of intracellular calcium

concentration (2). Current investigations are aimed to address

this issue, and they will be reported in a future publication

dedicated to signal transduction pathways activated by SMF

exposure.

ACKNOWLEDGMENTS

Dario Coletti has been awarded an ISAC Scholarship and

this paper has been performed in the frame of the ISAC men-

torship program with Laura Teodori as a mentor. The expert

technical assistance of Ms. Carla Ramina, Immacolata Senni,

and Stefania De Grossi for confocal microscopy imaging is

gratefully acknowledged. The anti-myogenin antibody, devel-

oped by Dr. W.E. Wright, and the anti-MHC antibody, devel-

oped by Dr. D.A. Fishman, were obtained from the Develop-

mental Studies Hybridoma Bank, developed under the

Cytometry Part A � 71A: 846�856, 2007 855

ORIGINAL ARTICLE

auspices of the NICHD and maintained by the University of

Iowa, Department of Biological Sciences, Iowa City, IA 52242.

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