Mouse Myofiber Cultures281
From: Methods in Molecular Biology, vol. 290: Basic Cell Culture Protocols, Third Edition
Edited by: C. D. Helgason and C. L. Miller © Humana Press Inc., Totowa, NJ
Isolation and Culture of Skeletal Muscle Myofibers
as a Means to Analyze Satellite Cells
Gabi Shefer and Zipora Yablonka-Reuveni
Myofibers are the functional contractile units of skeletal muscle. Mononuclear satel-
lite cells located between the basal lamina and the plasmalemma of the myofiber are the
primary source of myogenic precursor cells in postnatal muscle. This chapter describes
protocols used in our laboratory for isolation, culturing, and immunostaining of single
myofibers from mouse skeletal muscle. The isolated myofibers are intact and retain their
associated satellite cells underneath the basal lamina. The first protocol discusses
myofiber isolation from the flexor digitorum brevis (FDB) muscle. Myofibers are cul-
tured in dishes coated with Vitrogen collagen, and satellite cells remain associated with
the myofibers undergoing proliferation and differentiation on the myofiber surface.
The second protocol discusses the isolation of longer myofibers from the extensor
digitorum longus (EDL). Different from the FDB myofibers, the longer EDL myofibers
tend to tangle and break when cultured together; therefore, EDL myofibers are cultured
individually. These myofibers are cultured in dishes coated with Matrigel. The satellite
cells initially remain associated with the myofiber and later migrate away to its vicinity,
resulting in extensive cell proliferation and differentiation. These protocols allow stud-
ies on the interplay between the myofiber and its associated satellite cells.
Key Words: Satellite cells; skeletal muscle; myofiber isolation; single myofiber cul-
ture; flexor digitorum brevis; extensor digitorum longus; mouse; Vitrogen collagen;
Myofibers are the functional contractile units of skeletal muscle. Although
they are established during embryogenesis by fusion of myoblasts into
myotubes, processes involved in their growth and repair continue throughout
life. The development of myofibers and their regenerative potential depends
on the availability of myogenic precursor cells. Mononuclear satellite cells
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282 Shefer and Yablonka-Reuveni
located between the basal lamina and the plasmalemma of the myofiber are
classically considered to be the myogenic precursors in postnatal muscle (1,2).
Although in a growing muscle, at least some of the satellite cells are proliferat-
ing and adding myonuclei to the enlarging muscle fibers, in a normal adult
muscle, most satellite cells are quiescent. However, in response to a variety of
conditions, ranging from increased muscle utilization to muscle injury, quies-
cent satellite cells can enter the cell cycle, replicate, and fuse into existing
myofibers or form new myofibers (reviewed in ref. 2). The cascade of cellular
and molecular events controlling satellite cell myogenesis is therefore of inter-
est for understanding the mechanisms of muscle maintenance during the life-
span, as well as for developing strategies to enhance muscle repair after severe
trauma or during myopathic diseases.
Two main in vitro strategies have been employed in the study of satellite
cells: (i) myogenic cultures prepared from mononucleated cells dissociated
from the whole muscle (i.e., primary myogenic cultures) and (ii) cultures of
isolated myofibers where the satellite cells remain in their in situ position
underneath the myofiber basal lamina. Protocols for obtaining primary myo-
genic cultures aim at releasing as many satellite cells as possible from the entire
muscle. Steps of mincing, enzymatic digestion, and repetitive trituration of
the muscle are required for breaking down the connective tissue network and
myofibers in order to release the satellite cells from the muscle bulk. These
steps are followed by procedures aimed at removing tissue debris and reducing
the contribution of nonmyogenic cells typically present in primary isolates of
myogenic cells (3–5). In contrast, approaches for isolating myofibers aim at
releasing intact myofibers that retain the satellite cells in their native position
underneath the basal lamina (4). The first method, based on breakage of the
myofibers and release of satellite cells, provides a means for studying param-
eters affecting the progeny of satellite cells as they proliferate, differentiate,
and fuse into myotubes. The second method, based on isolating intact
myofibers, allows studying satellite cells in their in situ position as well as
studying their progeny after migrating from the myofibers.
This chapter describes the two approaches used in our laboratory for isola-
tion and culture of single myofibers from mouse skeletal muscle. One approach,
first introduced by Bekoff and Betz (6) and further developed by Bischoff (7,8),
has been adopted by us for studies of satellite cells in isolated myofibers from
both rat (4,9) and mouse (10). In this case, myofibers are isolated from the
flexor digitorum brevis (FDB) muscles of the hind feet, and multiple myofibers
are typically cultured together. The FDB has been used as donor muscle
because it consists of short myofibers that do not tangle (and consequently
break) when cultured together. A second approach, introduced by Rosenblatt
and colleagues (11,12), is suitable for the isolation of longer myofibers from a
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Mouse Myofiber Cultures283
variety of limb muscles (e.g., extensor digitorum longus [EDL], tibialis ante-
rior [TA]) (11,13,14). Different from the FDB myofibers, the longer myofibers
tend to tangle and break if cultured together. Hence, typically when working
with muscles such as EDL or TA, the isolated myofibers are cultured individu-
ally. In both approaches, the culture dishes are coated with commercially avail-
able matrixes that facilitate rapid and firm adherence of the myofibers to the
Table 1 compares in brief the two approaches for myofiber isolation and
the specific use of each procedure. Both protocols for myofiber isolation can
produce high yields of intact myofibers retaining their satellite cells under-
neath the basal lamina. However, delicate handling of the donor muscle only
at the tendons throughout harvesting and processing, the type and specific
source of the digesting enzyme, the length of the enzymatic digestion period,
and the degree of trituration of the digested muscle are all important factors
that should be well controlled during the isolation procedure. Myofibers that
are damaged in the course of the isolation procedure will not survive and can
be easily distinguished from the intact myofibers because they typically
Protocols for immunocytochemical analysis of satellite cells and their prog-
eny in cultures of FDB and EDL myofibers are also included in the chapter.
Representative micrographs of FDB and EDL myofiber cultures are shown
in Figs. 1 and (FDB, panels A–D) and Fig. 2 (EDL, panels A–D).
2.1. General Comments
1. As a general rule, only sterile materials and supplies are to be used. All solutions,
unless otherwise noted, are sterilized by filtering through 0.22-µm filters,
all glassware and dissection tools are sterilized by autoclaving, and all cell-cul-
turing steps are performed using sterile techniques.
2. The cultures are maintained at 37.5°C and 5% CO2 in a humidified tissue culture
3. All culture media are stored at 4°C and used within 3 wk of preparation.
4. Before starting isolation, the tissue culture medium is prewarmed to 37°C and
then held at room temperature throughout the procedures (do not leave medium
at 37°C for an extended period of time). Before transferring solutions/media into
the tissue culture hood, spray the glass/plastic containers with 70% ethanol.
5. The quantities of glassware, media, and reagents as well as the time intervals for
enzymatic digestion described in this chapter are appropriate for the isolation
of myofibers from one adult mouse of the age and strain detailed in Subheading
2.4. Adjustments are needed when isolating myofibers from younger/older mice,
other mouse strains, mutant mice, or other laboratory rodents such as rats.
6. Muscles used for preparing isolated myofibers are harvested from the hind limbs.
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284Shefer and Yablonka-Reuveni
resembling regular primary cultures
Characteristics of Myofiber Cultures From FDB and EDL Muscles of Adult Mice
Flexor digitorum brevis (FDB)
Extensor digitorum longus (EDL)
Relative myofiber length
Number of fibers per culture dish
Typical tissue culture dish
24-Well multiwell dish
Thick, gel-like layer of native collagen type I
Thin coating of diluted, growth-factor-
prepared from bovine dermal collagen
reduced Matrigel. Matrigel is a
[Vitrogen, Cohesion Technologies (9,10,15)]
basement membrane preparation
(see Note 1).
isolated from a mouse tumor
(BD Biosciences) (11) (see Note 2).
Dulbecco’s modified Eagle’s medium (DMEM)-
DMEM-based, serum rich/mitogen rich;
based, mitogen-depleted serum;
medium can be modified to a
specific exogenous growth factors are added
serum poor/mitogen-poor one to allow
to study their effect on satellite cell activation,
analysis of satellite cell activation
proliferation, and differentiation (9,10,15).
Satellite cell profile after culturing
Satellite cells remain at the surface of the parent
Satellite cells emigrate from the parent
myofiber as they proliferate and differentiate.
myofiber and undergo multiple rounds
Satellite cells undergo a limited number
of proliferation, giving rise to an
of proliferative cycles and rapidly differentiate
elaborate network of myotubes,
without fusing with the parent myofiber.
of cells dissociated from whole muscle.
Cultures can model in vivo behavior
Cultures can model events after muscle
of satellite cells in intact fibers during growth
trauma where new myofibers are formed.
and routine muscle utilization.
Cultures typically have been maintained
Cultures typically have been maintained short-term
long term and employed in studies
and employed for studies on recruitment
of myogenic cells, progeny of satellite
of satellite cells into the cell cycle.
cells that emigrate from the myofiber
Steps of proliferation and differentiation
to the myofiber surrounding (11).
are highly synchronous (9,10).
Cultures can also be used for analysis
Cultures can be further used to study cells
of molecular and cellular events
emigrating from the myofibers as described
associated with the first round
for the EDL fiber cultures.
of satellite cell proliferation,
as in FDB cultures (16).
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Mouse Myofiber Cultures285
Fig. 1. Phase and immunofluorescent micrographs of an isolated FDB myofiber
with associated satellite cells undergoing myogenesis. Myofibers were isolated from a
3-mo-old mouse and cultured in 35-mm tissue culture dishes coated with isotonic
Vitrogen collagen. Cultures were maintained for 4 d in basal medium containing
fibroblast growth factor 2 (FGF2, 2 ng/mL) and fixed with methanol as described in
Subheading 184.108.40.206. The culture shown in this figure was reacted via double immuno-
fluorescence with a monoclonal antibody against myogenin that stains the nuclei of
myogenic cells that have entered the differentiated step of myogenesis (panel C) and
a polyclonal antibody against ERK1/ERK2 mitogen-activated protein kinases
(MAPKs), which stains the cytoplasm of all fiber-associated cells (panel D). Reactiv-
ity with the monoclonal and polyclonal antibodies was traced with a fluorescein- and
rhodamine-labeled secondary antibody, respectively. Parallel phase image (panel A)
and DAPI staining image (panel D; both myofiber nuclei and satellite cell nuclei are
stained) are shown as well. Arrows in parallel panels point to the location of the same
cell. Additional immunopositive cells present on the myofiber are not shown, as not
all positive nuclei or cells on the fibers are in the same focal plane. All micrographs
were taken with a ×40 objective. Additional details regarding the source of the
antibodies and the rationale of using these antibodies are provided in our previous
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286 Shefer and Yablonka-Reuveni
Fig. 2. Phase micrographs of EDL myofibers depicting the temporal development
of myogenic cultures from cells emanating from individual myofibers. Myofibers were
isolated from 3-mo-old mice and cultured individually in 24-well multiwell tissue cul-
ture dishes coated with Matrigel. Cultures were maintained in serum-rich/mitogen-
rich growth medium and fixed with paraformaldehyde, as described in Subheading
220.127.116.11. Satellite cells begin to emigrate from the myofiber within the first day in cul-
ture and continue to emigrate during subsequent days. Satellite cells that have
emigrated from the myofibers proliferate, differentiate, and fuse into myotubes, estab-
lishing a dense myogenic culture. Satellite cells remained attached to the muscle fiber
during the first hours after culturing (panel A). Nineteen hours after culturing, two to
three cells detached from the fiber but remained in close proximity to the fiber (panel B).
Four days following culturing, more cells are seen in the vicinity of the myofibers
(only four cells shown in panel C). By d 7, progeny of satellite cells that emigrated
from the myofiber have established a culture containing mostly proliferating myo-
blasts and some myotubes (panel D). Micrographs in panels A–C were taken with a
×40 objective to show details of the few cells that emigrated from the myofiber,
whereas the micrograph in panel D was taken with a ×10 objective to show the estab-
lishment of a dense myogenic culture.
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Mouse Myofiber Cultures 287
2.2. General Equipment
The following facilities are required for the cultures described in this chapter:
1. Standard humidified tissue culture incubator (37.5°C, 5% CO2 in air).
2. Tissue culture hood.
3. Phase-contrast microscope.
4. Stereo dissecting microscope with transmitted light base (microscope is placed
inside a tissue culture hood).
5. Low-speed agitator placed in the tissue culture incubator (Labline Instruments,
Inc., model no. 1304). The agitator is used for gently agitating the muscle during
enzymatic digestion; a shaking water bath set at 37°C can be used instead of the
6. Bunsen or alcohol burner inside the tissue culture hood.
7. Water bath (37°C).
8. Hair trimmer (optional, for shaving hair from the hind limbs prior to muscle
2.3. Surgical Tools
1. Straight operating scissors: V. Mueller, fine-tipped, Sharp/Sharp stainless steel,
165 (6.5-in.) (VWR Scientific Inc., cat. no. 25601-142), for delicate cutting and
2. Dissecting scissors: stainless steel, 140-mm (5.5-in.) length. Both blades blunt
(VWR Scientific Inc., cat. no. 25877-103), protects the surrounding tissue from
any unwanted nicks.
3. Dressing forceps: V. Mueller, serrated, stainless steel, rounded points, 140-mm
(5.5-in.) length (VWR Scientific Inc., cat. no. 25601-072).
4. Two, very fine-point forceps: extrafine tips, smooth spring action, stainless steel.
Straight, 110 mm (4.5 in.; VWR Scientific Inc., cat. no. 25607-856).
5. Microscissors, Vannas scissors: 8 cm long, straight 5-mm blades, 0.1-mm tips
(World Precision Instruments, cat. no. 14003).
6. Scalpel handle: size 3 for blades 10–15 (Bard-Parker, cat. no. 371030) and sterile
blade (no. 10; Bard-Parker, cat. no. 371110).
7. Placement instrument (VWR Scientific Inc., cat. no. 1790-034)
8. Two straight, 5-in. hemostatic forceps (VWR Scientific Inc., cat. no. 25607-302).
9. Dissecting board.
C57BL/6 mice, 2–5 mo-old, maintained according to institutional animal
care regulations. Various other mouse strains have been used in our studies
following the myofiber isolation procedures described in this chapter.
The information in this subsection is provided to assist in the identification
and isolation of the FDB and EDL muscles.
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288 Shefer and Yablonka-Reuveni
2.5.1. Flexor Digitorum Brevis
The FDB is a superficial, multipennate, broad, and thin muscle of the foot
and paw (8,17); it arises from the tendon of the plantaris as three slender
muscles converging into long tendons. At the base of the first phalanx, it divides
into two, passes around the tendon of the flexor hallucis longus obliquely across
the dorsum of the foot, and ends as the tendons insert into the second phalanx
of the second through the fifth digits. As the FDB contracts, digits 2–5 are
flexed. For additional details about the anatomy of the FDB muscle, see Note 3.
2.5.2. Extensor Digitorum Longus
The EDL muscle is situated at the lateral part of the hind limb running from
the knee to the ankle, extending to the second to fifth digits (17). The EDL
actually consists of four combined muscle bellies and their tendons; the bellies
arise from the lateral condyle of the tibia and the front edge of the fibula (two
tendons at the origin of the muscle). The tendons lie close to each other and
appear as one glistening white tendon that continues down to the surface of the
ankle. At the ankle joint, it separates to four tendons, each attached to one of
the second to fifth digits. As the EDL contracts, the four digits are extended.
For additional details about the anatomy of the EDL muscle, see Note 3.
2.6. Plastic and Glassware for Myofiber Isolation and Culture
2.6.1. FDB Myofiber Isolation and Culture
1. Standard 9-in. Pasteur pipets (VWR Scientific Products, cat. no. 0035904).
2. Standard 5-in. sterile glass Pasteur pipets (VWR Scientific Products, cat. no.
3. Wide-mouth pipets prepared from the standard 5-in. Pasteur pipets. Cut the tip of
a pipet about 3-in. from its narrow end using a file or a diamond knife. Shake the
pipet to remove any glass fragments. Use flame in hood to fire-polish the distal
ends of all Pasteur pipets listed in items 1–3 to smoothen sharp edges that can
4. Syringe filters, 0.2 and 0.45 µm (Millex-GS, Millipore, cat. no. SLGS0250S and
SLHA0250S, respectively), and 10-cm3 syringes.
5. Sterile conical tubes, 15 and 50 mL (BD Biosciences/Falcon, cat. no. 352098 and
6. Three glass Corex tubes, 15 mL (Sorvall centrifuge tubes; or alternatively 15-mL
bicarbonate Sorvall tubes).
7. Wide-bore 100-µL micropipet tips. Trim 100-µL tips 3 mm from the end to mini-
mize myofiber shearing when transferring or dispensing FDB myofibers.
8. Tissue culture dishes, 35 mm (Corning Incorporated, cat. no. 430165).
9. Two L-shaped bent pipet spreaders prepared from standard 9-in. Pasteur pipets.
Use flame to first seal the distal end, then flame about 3⁄4 in. from the sealed end
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Mouse Myofiber Cultures 289
until the pipet starts to bend. The bent pipets are used to spread the coating solu-
tion on the tissue culture dishes. Spreaders should be prepared in advance and
allowed to cool before use.
2.6.2. EDL Myofiber Isolation and Culture
1. Standard 9-in. and 5-in. sterile Pasteur pipets, syringe filters and conical tubes
listed and treated as described in items 1–6 in Subheading 2.6.1.
2. Three gradually narrower-bore pipets prepared from standard 5-in. Pasteur pipets.
Use a file or a diamond knife to prepare a set of pipets with bore diameter of
approx 2.5, 2, and 1 mm. Shake the pipet to remove any glass fragments and fire-
polish the sharp ends. These pipets are used to triturate the digested muscle in
order to release single myofibers.
3. Six plastic Petri dishes, 60 × 15 mm (Becton Dickinson Biosciences, Falcon,
cat. no. 351007).
4. Twenty four-well Falcon multiwell tissue culture dish (Becton Dickinson Bio-
sciences, cat. no. 353047) (see Note 4).
2.7. Media and Cell Culture Reagents
2.7.1. FDB Myofiber Isolation and Culture
1. DMEM (Dulbeco’s modified Eagle’s medium; high glucose, with L-glutamine,
with 110 mg/L sodium pyruvate, with piridoxine hydrochloride (Gibco–
Invitrogen Life Technologies, cat. no. 11995065) supplemented with 50 U/mL
penicillin and 50 mg/mL streptomycin (Gibco–Invitrogen, cat. no. 15140-122).
2. Horse serum (HS); standard, not heat inactivated (HyClone, cat. no. SH30074.03);
stored at –20°C (see Note 5).
3. HS, 20 mL, freshly filtered (on the day of use) through a 0.45-µm filter.
4. DMEM, 100 mL, containing 10% filtered HS. All Pasteur pipets and micropipet
tips are preflushed with DMEM containing 10% HS to prevent sticking of
myofibers during manipulation.
5. Controlled Process Serum Replacement (CPSR) (Sigma–Aldrich, stored at –20°C).
Alternative serum replacement products (e.g., Sigma–Aldrich, cat. no. S9388)
(18) can also be used depending on experimental requirements (see Note 6).
6. FDB myofiber culture medium is made up of DMEM (supplemented with anti-
biotics), 20% CPSR and 1% HS.
7. Vitrogen collagen in solution (Cohesion Technologies, cat. no. FXP-019) for
coating 35-mm tissue culture dishes. Vitrogen collagen in solution is the recom-
mended product and the use of collagen from other companies would require
prescreening to ensure compatibility (see Note 1).
8. 7X DMEM made from powder DMEM (1-L package; Sigma–Aldrich, cat. no.
D3656); used to prepare isotonic Vitrogen collagen (see Note 1).
9. Collagenase (type I, Sigma–Aldrich, cat. no. C-0130) used for muscle digestion
as described in Subheading 3.
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2.7.2. EDL Myofiber Isolation and Culture
1. DMEM and HS as listed and prepared in items 1–4 in Subheading 2.7.1.
2. Fetal bovine serum (FBS; standard, not heat inactivated) (Sigma–Aldrich, cat. no.
F-2442; stored at –20°C) (see Note 7).
3. Chicken embryo extract (CEE) (Gibco–Invitrogen, cat. no. 16460024),
stored at –20°C; or, as in our studies, prepared by the investigator (see Notes 8
4. EDL myofiber culture medium made up of DMEM (supplemented with antibiot-
ics), 20% FBS, 10% HS, and 1% CEE.
5. Matrigel (see Note 2) for coating 24-well multiwell dishes. Matrigel can be pur-
chased in its standard format (BD Biosciences, cat. no. 354234) or in its growth-
factor-reduced format (BD Biosciences, cat. no. 354230). In our studies, we use
the growth-factor-reduced format.
6. Collagenase, as listed in item 9 in Subheading 2.7.1.
2.8. Reagents and Solutions
for Fixing and Immunostaining Myofiber Cultures
2.8.1. FDB Myofiber Cultures
1. Prefixation rinse solution: DMEM as in item 1 in Subheading 2.7.1.
2. Fixative: ice-cold 100% methanol (see Note 10).
3. Rinse solution: Tris-buffered saline (TBS); 0.05 M Tris-HCl, 0.15 M NaCl,
pH 7.4 (see Note 11).
4. Detergent: Tween-20 (Sigma, cat. no. P1379).
5. Detergent solution: TBS containing 0.05% Tween-20 (TBS-TW20).
6. Blocking reagent: normal goat serum (Sigma–Aldrich, cat. no. G9023).
7. Blocking solution: TBS containing 1% normal goat serum (TBS-NGS).
8. Mounting medium: Vectashield (Vector Laboratories, Inc., Burlingham; cat. no.
9. Cover glass, 22 mm2 (Corning Labware and Equipment, cat. no. 48371-045).
2.8.2. EDL Myofiber Cultures
1. Fixative: 4% paraformaldehyde containing 0.03 M sucrose (see Notes 12
2. Rinse solution: TBS as in item 3 in Subheading 2.8.1.
3. Detergents: Triton X-100 (Sigma, cat. no. T6878); Tween-20 as in item 4 in
4. Detergent solution: TBS containing 0.5% Triton X-100 (TBS-TRX100); TBS-
TW20 as in item 5 in Subheading 2.8.1.
5. Blocking reagent and solution: same as items 6 and 7 in Subheading 2.8.1.
6. Mounting medium: same as item 8 in Subheading 2.8.1.
7. Microcover glass (VWR Scientific Inc., cat. no. 12CIR-1).
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Mouse Myofiber Cultures 291
3.1. Isolation of Single Myofibers From the Flexor Digitorum Brevis Muscle
3.1.1. Initial Steps Prior to Harvesting
the Muscle and Preparation of Digestive Enzyme
1. Add 3 mL of DMEM to six 35-mm tissue culture dishes and place the dishes in
the tissue culture incubator until muscle dissection begins.
2. Add 3 mL of DMEM containing 10% HS to three 35-mm tissue culture dishes
and place them in the tissue culture incubator until needed for the isolated single
3. Add 6 mg of collagenase type I to 3 mL of DMEM in order to prepare 0.2% (w/v)
collagenase type I solution. Use a 0.22-µm filter attached to a 10-cm3 syringe to
filter the collagenase solution into a 35-mm tissue culture dish (see Note 14).
3.1.2. Dissection of FDB Muscle
1. Euthanize one mouse according to institute regulations.
2. Shave the hind limbs and spray them lightly with 70% ethanol.
3. Secure the mouse, lying on its back, to the dissecting board by pinning down the
forelimb diagonally across from the limb being dissected.
4. Use a scalpel to cut through the skin all around and just above the ankle (after this
initial circular cut, the skin below resembles a sock).
5. Cut the skin in a straight line along the center of the ventral part of the foot
almost all the way to the digits (the cut as viewed from the front of the foot
should resemble a “T” shape).
6. Clamp a hemostatic forceps to one of the upper corners of the cut tissue (at the
junction of the circular and longitudinal cuts), shifting the skin away from the foot.
7. Hold the scalpel with its blade parallel to the longitudinal axis of the partially
exposed muscle and carefully cut away the connective tissue. Be especially care-
ful not to cut into the muscle tissue at the back of the leg, as the FDB is the most
superficial muscle of the back of the foot.
8. Clamp the second hemostat to the other corner of the cut tissue and repeat step 7.
9. When the skin is completely cut away from the foot, the FDB should be exposed
all the way to the tendons reaching the digits.
10. Turn the mouse over so that it lies on its stomach, and identify the FDB. During
the next steps of the surgery, to avoid blood cell contamination of the myofiber
preparation, be careful not to injure the small medial plantar artery that supplies
blood to the FDB. This artery passes along the medial part of the sole of the foot
and branches into the digits.
11. Carefully run the tip of the scalpel along each side of the FDB to disrupt the
connective tissue holding the muscle in place.
12. When the FDB is separated from the surrounding muscles, insert the tip of the
placement instrument underneath the FDB and gently lift the muscle so that the
flat side of the scalpel can be inserted horizontally underneath it.
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292 Shefer and Yablonka-Reuveni
13. With the blade of the scalpel underneath the muscle, running horizontal and par-
allel to the muscle, cut away the underlying connective tissue. It is best to cut
toward the heel and only lift that portion of the muscle directly over the scalpel.
14. Cut underneath the tendon to separate the muscle and a large portion of its tendon
from the heel bone.
15. Clamp the freed tendon as far as possible from the muscle tissue with a hemostat.
16. Use the hemostatic forceps to gently lift the FDB away from the leg. Use the
scalpel, running parallel to the muscle, to cut through the connective tissue while
holding the FDB down.
17. Continue cutting through the connective tissue until the tendons that connect the
FDB muscle to the digits have been exposed. When about half the length of the
three tendons has been exposed, cut the tendons and release the entire muscle from
the leg. The fourth small lateral tendon (attached to the fifth digit) and its attached
myofibers can be trimmed off.
18. Retrieve from the incubator three 35-mm tissue culture dishes containing DMEM
and place them close to the dissection area.
19. Place the harvested FDB in one of the 35-mm tissue culture dishes.
20. For harvesting the FDB from the other hind foot, repeat steps 11–17 and place
the muscle in a second 35-mm tissue culture dish.
21. Place the 35-mm tissue culture dishes, one at a time, under the stereo dissecting
22. Use fine-point forceps to pull the connective tissue perpendicular to the line of
the muscle and use scissors to cut it off.
23. Once the muscle is clean, shorten the tendons but do not cut off all of them.
24. Use a wide-bore Pasteur pipet to transfer the cleaned muscle to another
35-mm tissue culture dish containing DMEM.
25. Repeat steps 21–24 to clean the second FDB muscle.
3.1.3. Enzymatic Digestion
1. Transfer the two cleaned FDB muscles to a 35-mm tissue culture dish with the
0.2% collagenase I solution.
2. Place this 35-mm tissue culture dish on the low-speed agitator inside the tissue
culture incubator to allow gentle and continuous collagenase digestion for 2.5 h
(see Notes 14 and 15).
3. At the end of the digestion period, transfer each muscle to a 35-mm tissue culture
dish containing 10% HS.
3.1.4. Separation of the Three Tendons and Release of Myofibers
All Pasteur pipets used are preflushed with 10% HS as described in item 4
in Subheading 2.7.1.
1. Place one muscle at the time under the stereo dissecting microscope.
2. Identify the two grooves running between the three tendons separating the middle
from the two lateral tendons.
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Mouse Myofiber Cultures293
3. Being careful not to touch the muscle, insert the tip of a forceps into one of the
grooves, and, by securing the connective tissue between the tendons to the dish,
hold the muscle in place.
4. Use another pair of forceps to gently pull the connective tissue that holds the
tendons and their attached muscle tissue together.
5. Continue removing the connective tissue until the lateral tendons are separated
from the middle tendon and its attached myofibers.
6. Holding the muscle only at its tendons, transfer the muscle preparation to a
35-mm dish containing 3 mL of 10% HS.
7. While grasping one end of the middle tendon with a pair of forceps, use a second
pair of forceps to grip its surrounding connective tissue sheath and pull gently.
If the sheath does not come off easily, use fine-point forceps to pull the connec-
tive tissue perpendicular to the line of the muscle and cut it off.
8. Repeat steps 1–7 with the second FDB muscle until all six tendons and their
attached myofibers are in the 35-mm tissue culture dish containing 10% HS.
9. For one tendon at a time: hold one end of the tendon with a pair of forceps
and with the tip of a second pair gently separate the myofibers from the ten-
don. The liberation of the myofibers from the two lateral tendons should be easy;
the middle tendon requires patience because the myofibers are attached to it
10. Use a wide-bore Pasteur pipet to gently triturate the clumps of myofibers until
they disengage into single myofibers.
11. Remaining clumps should be transferred to another 35-mm tissue culture dish
containing 10% HS and further triturated until disengaged into single myofibers.
12. Set the stereo dissecting microscope magnification so that the small pieces of
connective tissue floating around in the suspension are visible and use a pair of
forceps to pick them out. Continue until the myofiber suspension is clean of con-
nective tissue debris.
13. Triturate the myofiber suspension 10 more times using a 9-in. Pasteur pipet with
a fire-polished tip to further separate small clumps of myofibers.
3.1.5. Further Purification of FDB Myofibers
1. Add 10 mL of 10% HS to each of the three glass Corex tubes.
2. Using the trimmed 100-µL pipet tip, transfer the myofiber suspension to the top
of the 10% HS column in the first Corex tube. Allow the myofibers to settle
(at 1g) through the HS column for 15 min at room temperature (see Note 16).
This step is important for purifying the myofibers from free mononucleated cells,
debris, and occasional broken myofibers.
3. As soon as the myofibers are settled, aspirate about 11 mL of the supernatant
(leaving about 1–1.5 mL). Triturate the myofiber suspension gently with a 5-in.
fire-polished Pasteur pipet and transfer the suspension to the next Corex tube as
described in step 2.
4. Allow myofibers to settle and transfer the myofiber suspension to the third Corex
tube as in steps 2 and 3.
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294 Shefer and Yablonka-Reuveni
5. Allow myofibers to settle and harvest the final myofiber suspension. Following
the third purification, the residual volume of medium to be left with the myofiber
suspension depends on the number of culture dishes and the desired myofiber
number per dish. Typically in our studies, the volume of the final myofiber sus-
pension is 300 µL, which is sufficient for culturing four to six dishes.
3.1.6. Preparation of Isotonic Vitrogen Collagen
Isotonic Vitrogen collagen can be prepared during the settling of myofibers.
The isotonic mixture should be kept on ice. Stock Vitrogen is an acidic solu-
tion, and when made isotonic, it gels rapidly if not maintained at 4°C (see Note 1).
1. Place Vitrogen collagen stock bottle, 7X DMEM, and one l5-mL conical tube
2. On ice: Add 1 vol of 7X DMEM and 6 vol of Vitrogen to the 15-mL conical tube
and mix gently. Calculate the volume of stock Vitrogen needed for the experi-
ment based on using 120 µL isotonic Vitrogen collagen to coat each 35-mm tis-
sue culture dish. Use pH paper strips to ensure a neutral pH of the Vitrogen
collagen in DMEM solution. The pH of this solution rises slightly after coating
the culture dish. If the pH remains acidic after coating a test dish, add 1–2 drops
of 1 M NaOH to the Vitrogen collagen in DMEM solution.
3.1.7. Coating Culture Dishes
With Isotonic Vitrogen Collagen and Myofiber Culturing
1. On ice: Transfer 120 µL of isotonic Vitrogen collagen to the center of a
35-mm culture dish and immediately use the L-shaped spreader to coat the
2. Gently swirl the myofiber suspension (in the 15-mL tube) for even distribution of
myofibers throughout the residual medium.
3. Remove one culture dish at a time from ice to allow rapid warming to room
4. Use a wide-bore 100-µL micropipet tip to dispense about 50 µL of the myofiber
suspension per each culture dish.
5. Gently swirl the culture dish to allow even distribution of the myofibers.
6. Repeat steps 2–5, one dish at a time, for additional culture dishes.
7. Transfer dishes to the tissue culture incubator for a minimum of 20–30 min to
allow the formation of Vitrogen collagen matrix and the adherence of the
myofibers to the matrix.
8. Remove dishes from the incubator. Gently add 1 mL of myofiber culture medium
to each dish without agitating the myofibers and return dishes to the incubator.
When the effect of growth factors on satellite cell proliferation/differentiation is
investigated, parallel cultures are maintained in myofiber culture medium with/with-
out additives and the medium is replaced every 24 h to ensure that growth factors
do not become rate limiting. Except for harvesting myofiber cultures for early
time points, cultures should be left undisturbed for the initial 18 h to allow good
adherence of myofibers to matrix.
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Mouse Myofiber Cultures295
3.2. Isolation of Single Myofibers From the EDL Muscle
3.2.1. Procedures Prior to Muscle Harvesting
18.104.22.168. PREPARATION OF MATRIGEL WORKING MIXTURE
AND COATING TISSUE CULTURE DISHES WITH MATRIGEL
1. Thaw the required amount of stock Matrigel by placing frozen aliquots on ice for
approx 20 min. When diluted into the final working solution, a 200-µL stock
aliquot should be sufficient for coating at least four 24-well multiwell dishes.
2. Prechill a 15-mL conical tube on ice and transfer the thawed Matrigel into the
tube. Add ice-cold DMEM to dilute the Matrigel to a final concentration of 1 mg/mL.
3. Place 500 µL of diluted Matrigel solution in the center of each of the 24 wells,
using a glass 1-mL pipet.
4. Swirl the 24-well multiwell dish to allow even coating of the wells.
5. Allow the Matrigel-coated dish to sit at room temperature for 5–10 min in the
tissue culture hood.
6. Transfer excess Matrigel solution from the wells back to the original tube with diluted
Matrigel that is kept on ice. Use this Matrigel solution to coat additional dishes
within the next 2 h. Do not keep diluted Matrigel for reuse on subsequent days.
7. Incubate the Matrigel-coated multiwell dishes in the tissue culture incubator until
the end of the enzymatic digestion period, but for at least 30 min.
22.214.171.124. COATING GLASSWARE AND PLASTICWARE DISHES
WITH HS FOR THE INITIAL STEPS OF MYOFIBER ISOLATION
1. Coat six plastic Petri dishes with undiluted filtered HS, prepared as described in
item 3 in Subheading 2.7.1. Transfer 1 mL of HS to each Petri dish and swirl the
dish to coat evenly.
2. Allow the dishes to sit with HS solution for 5 min at room temperature; then,
aspirate the HS and add 7 mL of DMEM to each Petri dish.
3. Incubate Petri dishes in the tissue culture incubator until needed following muscle
4. Coat the fire-polished Pasteur pipets, prepared as described in items 1 and 2 in
Subheading 2.6.2., with HS by passing 10% HS solution through the pipets sev-
126.96.36.199. PREPARATION OF THE DIGESTING ENZYME SOLUTION
Prepare 0.2% (w/v) collagenase type I solution in 3 mL of DMEM and filter
the solution into a 35-mm tissue culture dish using a 0.22-µm syringe filter
(see Note 14).
3.2.2. Dissection of EDL Muscle
1. Euthanize one mouse according to institute regulations.
2. Shave the hind limbs and spray them lightly with 70% ethanol.
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296 Shefer and Yablonka-Reuveni
3. Secure the mouse, lying on its back, to the dissecting board by pinning down the
hind limb to be operated on and the diagonal forelimb.
4. Use the straight rounded-tip scissors to cut through the skin, opening a small
incision above the knee.
5. Holding the skin with fine forceps, insert the rounded-tip scissors beneath the
incision, and carefully open the scissors to loosen the skin from the underlying
6. Extend the incision to a point just in front of the digits.
7. Loosen the skin as you go, being careful not to cut the underlying muscles or
8. Cut and remove the skin from the knee to the paw.
9. Identify the two tendons at the origin of the EDL.
10. Use microscissors to cut these tendons as far as possible from the muscle itself.
11. Identify the four tendons at the insertion of the EDL, each extending to one of the
digits but not the toe.
12. Use the microscissors to cut all four tendons.
13. Using fine forceps, gently pull the portion of the tendon before its division (to the
four tendons) until the four tendons slide from the paw up to the ankle.
14. Grasp the four tendons and carefully pull them in order to remove the EDL
15. The EDL should slide underneath the TA muscle and should pull out easily. It is
very important not to apply any force; if the muscle does not slide out easily,
one or both tendons at the origin of the muscle might still be attached to the bone.
In that case, identify the attached tendon and cut it.
16. The muscle should only be handled by its tendons to prevent damage to the
myofibers. Be careful not to injure the anterior tibial artery that supplies blood
to the EDL, to avoid blood cell contamination of the myofiber preparation.
In the upper third of its course, this artery lies between the TA and EDL
muscles (very close to the origin of the EDL muscle); in the middle third,
it lies between the TA and extensor hallucis longus. The lower third of the
artery starts at the ankle, crossing from the lateral to the medial side, lying
between the tendon of the extensor hallucis longus and the first tendon of the
insertion of EDL muscle.
3.2.3. Enzymatic Digestion
1. Holding the muscle by its four tendons, transfer it to the 35-mm tissue culture
dish containing 0.2% collagenase I solution.
2. Place the dish on the low-speed agitator located inside the incubator. Allow gentle
and continuous agitation for collagenase digestion for 60 min (see Notes 14 and 15).
3.2.4. Liberation of Single Myofibers From Muscle Bulk
Use a stereo dissecting microscope (placed inside tissue culture hood)
throughout the procedure.
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Mouse Myofiber Cultures297
1. Inspect the muscle under the stereo dissecting microscope to make sure that the
myofibers are loosened from the muscle bulk. If the myofibers are not loosened,
continue enzymatic digestion for another 10 min and check again.
2. Retrieve two Petri dishes containing 7 mL of DMEM from the incubator. Use the
widest-bore Pasteur pipet to transfer the muscle from the collagenase solution to
the DMEM to rinse away the collagenase.
3. Transfer the muscle to the second Petri dish for further dilution of any possible
4. Use another wide-bore pipet (diameter: approx 2 mm) to triturate the muscle
along its length. This orientation of the EDL muscle during triturations is critical
to prevent myofiber breakage.
5. When single myofibers are liberated from the muscle, its diameter decreases.
Therefore, use a narrower-bore pipet for subsequent triturations.
6. When 20–30 viable single myofibers are released, transfer the muscle bulk to
another DMEM-containing Petri dish and place the dish with the single myofibers
in the tissue culture incubator. The transfer of the muscle bulk to a second dish
ensures that the already released myofibers do not break during subsequent
7. Repeat trituration and transfer muscle bulk to new Petri dishes until the desired
number of viable isolated myofibers is acquired.
3.2.5. Culturing Single Myofibers in 24-Well Multiwell Dishes
1. Transfer a Matrigel-coated 24-well multiwell dish from the incubator to the tis-
sue culture hood and open its lid to allow moisture, generated during the incuba-
tion period, to evaporate.
2. Bring two Petri dishes containing single myofibers to the tissue culture hood.
3. Use a 9-in. glass Pasteur pipet to lift one myofiber, with minimal residual
medium, from the suspension and gently release the myofiber in the center of a
well. Alternate between the two myofiber-containing dishes in order to have both
early and late isolated single myofibers in each 24-well multiwell dish.
4. After myofibers are dispensed to all 24 wells, look under the stereo dissecting
microscope and make sure that indeed there is a myofiber in each well. This step
is necessary because occasionally myofibers adhere to the Pasteur pipet and are
not released to the well.
5. If needed, add a myofiber to any empty well.
6. Approximately 10 min after distributing myofibers, slowly add 500 µL of warm
culturing medium to each well, avoiding myofiber agitation.
7. Transfer the 24-well multiwell dish to the tissue culture incubator for a minimum
of 18 h (overnight).
8. Repeat steps 1–7 until the required number of cultured myofibers is reached.
9. An additional 500 µL of fresh culturing medium is provided to each well 1 wk
after culturing the myofibers. Then, to replenish the medium, every 3 d about
500 µL of the medium is aspirated and 500 µL of fresh medium is added.
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3.3. Immunolabeling of Satellite Cells in FDB and EDL Myofiber Cultures
This subsection details current protocols used in our laboratory to fix
myofiber cultures for immunofluorescence studies of satellite cells. FDB
myofiber cultures are typically fixed with ice-cold methanol (the preferred fixa-
tive when working with Vitrogen-collagen-coated dishes). EDL myofiber cul-
tures are typically fixed with paraformaldehyde warmed to 37°C. These
protocols allow recovery of intact myofibers at the end of the fixation proce-
dure. It should be noted that the ideal fixatives for FDB or EDL myofiber cul-
tures are not necessarily the optimal fixatives for antigen detection. Thus, when
analyzing single myofibers via immunofluorescence, fixatives should be opti-
mized for both preserving the myofibers and the antigens being analyzed.
Fixation protocols described in this subsection are also appropriate for
detecting proliferating satellite cells in single myofibers by autoradiography
following labeling with 3H-thymidine (7,19) or when analyzing proliferation
using bromodeoxyuridine (16,18).
3.3.1. Protocols for Fixing and Immunofluorescent Staining
of Isolated Single Myofiber Cultures
188.8.131.52. FDB MYOFIBER CULTURES
1. Warm DMEM in a water bath set at 37°C.
2. Rinse cultures with warm DMEM three times. Following the final rinse, add 1 mL
ice-cold 100% methanol to each 35-mm tissue culture dish and transfer the dishes
to 4°C for 10 min.
3. Return dishes to room temperature, aspirate the methanol, and allow the dishes to
air-dry for 10–15 min in the tissue culture hood (see Note 17).
4. Add 1.5 mL of blocking solution (TBS-NGS) to each culture dish to block non-
specific antibody binding.
5. Cultures are then kept at 4°C for overnight or longer.
6. Dilute the appropriate primary antibody in the blocking solution.
7. Rinse the cultures three times with TBS-TW20.
8. Aspirate the final TBS-TW20 rinse and add 100 µL of the primary antibody solu-
tion for 1 h at room temperature followed by an overnight incubation at 4°C in a
humidified chamber (see Notes 18 and 19).
9. Dilute the appropriate secondary antibody in the blocking solution.
10. Rinse cultures with TBS-TW20 three times.
11. Aspirate the final TBS-TW20 rinse and add 100 µL of the diluted secondary
antibody for 1–2 h at room temperature.
12. Aspirate the secondary antibody and wash three times with TBS-TW20.
13. For nuclear visualization, add 100 µL of DAPI solution (4',6-diamidino-2-
phenylindole, dihydrochloride; Sigma-Aldrich, cat. no. D8417; stock concentra-
tion 10 mg/mL, working concentration 1 µg/mL diluted in TBS-NGS prior to
use) for 30 min at room temperature (see Note 20).
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Mouse Myofiber Cultures299
14. Rinse the cultures twice with TBS-TW20 followed by a final rinse with TBS.
15. Aspirate the TBS and mount in Vectashield mounting medium (1 drop at the
center of each culture dish or well) and cover with a cover slip. Mounting medium
prevents the stained cultures from drying and retards fading of the immunofluo-
184.108.40.206. EDL MYOFIBER CULTURES
1. Warm the needed volume of paraformaldehyde–fixative solution in a water bath
set at 37°C.
2. While observing each myofiber under the stereo dissecting microscope, use a
pipetman to gently, without agitating the culture or touching the myofiber,
add 500 µL of the warm paraformaldehyde–fixative solution to the culturing
medium of each of the 24 wells, for 10 min at room temperature.
3. Use a pipet to remove the paraformaldehyde–fixative-medium solution and rinse
each well three times with TBS.
4. Add 500 µL of TBS-TRX100 for 5 min at room temperature.
5. Add 500 µL of blocking solution (TBS-NGS) to each of the 24 wells to block
nonspecific antibody binding.
6. Follow steps 5–15 as described in Subheading 220.127.116.11.
1. Vitrogen collagen in solution is a sterile solution of purified, pepsin-solubilized
bovine dermal collagen type I dissolved in 0.012 N HCl and stored at 4°C until
used (Cohesion Technologies, Palo Alto, CA). In our studies, Vitrogen collagen
is made isotonic by mixing 6 vol of stock Vitrogen collagen with 1 vol of 7X
DMEM. The isotonic solution is prepared just prior to coating dishes because it
gels rapidly at room temperature. To obtain consistent coating, the culture dishes
should be precooled and coated on ice. When removed from the ice, these dishes
warm up rapidly and are ready for myofiber addition. Vitrogen collagen in solu-
tion is the recommended product and the use of collagen from other companies
would require prescreening to ensure compatibility.
2. Matrigel is a solubilized basement membrane preparation extracted from the
Engelbreth–Holm–Swarm mouse sarcoma, a tumor rich in extracellular matrix
proteins. Its major component is laminin, followed by collagen IV, entactin, and
heparan sulfate proteoglycan (20). To ensure Matrigel stability, we follow the
manufacturer’s handling instructions and aliquot 200 µL each into 2-mL cryo-
genic vials sealed with O-rings (Corning Inc., cat. no. 430488). These aliquots
are stored at –20°C.
3. For additional details about the FDB muscle anatomy, refer to http://www.
bartleby.com/107/illus443.html and http://www.bartleby.com/107/131.html.
For additional details about the EDL muscle anatomy, refer to http://www.
bartleby.com/107/illus437.html, http://www.bartleby.com/107/illus441.html, and
http://www.bartleby.com/107/129.html. We recommend these links as good
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300 Shefer and Yablonka-Reuveni
resources for anatomical description and schematic images of the muscles
although they refer to human muscles.
4. Falcon Primaria (Becton Dickinson Biosciences) 24-well multiwell dishes have
typically been used for single myofiber isolation; however, we find that the stan-
dard, less expensive Falcon 24-well multiwell dishes are as good.
5. Horse serum (HS) should be preselected by comparing sera from various suppli-
ers. We select HS based on its capacity to support proliferation and differentia-
tion of primary chicken myoblasts cultured at standard and clonal densities (21).
6. The Controlled Processed Serum Replacement 2 (CPSR-2; Sigma–Aldrich) that
had been routinely used in our earlier studies (4,9,10,15,19) has been discontin-
ued. The source of both the discontinued CPSR-2 and the currently available
CPSR-3 is dialyzed bovine plasma; the discontinued product was further pro-
cessed in a manner that also reduced lipids. The alternative serum replacement
product contains bovine serum albumin, insulin, and transferrin and its use for
mouse myofiber cultures has been previously described (18).
7. Fetal bovine serum (FBS) should be preselected by comparing sera from several
suppliers. We select FBS based on the capacity of the serum to support prolifera-
tion and differentiation of mouse primary myoblasts cultured at various cell den-
sities. Only sera able to support growth and differentiation over a wide range of
cell densities are employed in our studies. Primary myogenic cultures are pre-
pared as described in refs. 4 and 22.
8. We prepare chicken embryo extract (CEE) in our laboratory using 10-d-old White
Leghorn embryos (23). The procedure is similar to a previously described method
(24) but uses the entire embryo. We recommend this approach over purchasing
CEE if the investigator can obtain embryonated chicken eggs, as the quality is
higher and the cost lower than that of purchased CEE.
9. Preparation of chicken embryo extract. All steps are performed in a sterile manner.
a. Embryonated chicken eggs (8 dozen, White Leghorn; from Charles River) are
maintained in a standard egg incubator (incubation conditions: a dry tempera-
ture of 38°C, a wet temperature of 30°C, and relative humidity of 56%).
The following egg incubator is well suited for basic research use: Marsh
Automatic Incubator, Model PRO-FI, cat. no. 910-028, manufactured by Lyon
Electric Company Inc., Chula Vista, CA.
b. After 10 d, batches of 15–30 eggs are removed from the incubator and trans-
ferred into the tissue culture hood.
c. Place the eggs lengthwise in the rack and spray with 70% ethanol to sterilize.
Wait for several minutes until the ethanol evaporates.
d. Crack open one egg at a time into a 150-mm Petri dish.
e. Remove the embryo from surrounding membranes by holding it with fine
forceps. Rinse the embryo by transferring it through three 150-mm Petri
dishes containing minimal essential medium (MEM; Gibco–Invitrogen,
cat. no. 11095-080, MEM is supplemented with antibiotics as described for
DMEM in Subheading 2.7.1.). Swirl embryo a few times in each dish for a
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Mouse Myofiber Cultures301
f. Empty the egg remains from the initial 150-mm dish (described in step d)
into a waste beaker and repeat steps d–f until the final rinse dish contains
about 30 embryos.
g. The embryos are transferred with fine forceps into a 60-mL syringe, forced
through with the syringe plunger, and the suspension is collected into a
500-mL sterile glass bottle.
h. The extract is diluted with an equal volume of MEM and gently agitated for
2 h at room temperature. To ensure good agitation, keep maximum volume to
one-half bottle capacity.
i. The extract is frozen at –70°C for a minimum of 48 h. It is then thawed,
dispensed to sterile glass Corex tubes, and centrifuged at 15,000g for 10 min
to remove particulate material.
j. The supernatant is pooled, divided into 5-mL aliquots, and kept frozen until
k. Prior to use, the CEE should again be centrifuged at about 700g for 10 min to
remove aggregates, passed through a 0.45-µm filter, followed by a
0.22-µm filter (to clear remaining particles and to ensure sterility).
10. Methanol is a colorless, flammable liquid with an alcohol-like odor. Use nitrile
gloves, safety goggles, and a fume hood when handling. It is important to refer to
the MSDS instructions and institutional regulations for further information
regarding storage, handling, and first-aid.
11. Preparation of Tris-buffered saline (TBS). To make 1 L of 10X TBS:
a. Weigh 60.5 g of Tris base into a beaker.
b. Add 700 mL deionized water to the beaker.
c. Place the beaker on top of a magnetic stirrer.
d. When the powder has dissolved, adjust the pH to 7.4.
e. Add deionized water to bring the volume up to 1 L, mix well, and store at 4°C.
To make 1 L of TBS:
a. Weigh 8.766 g NaCl in a beaker
b. Add 100 mL of 10X TB to the beaker and mix vigorously.
c. When the powder has dissolved, add deionized water to bring the volume up
to 1 L; mix well and store at 4°C.
d. In a sterile environment: filter through a 0.45-µm disposable filter unit
(Nalgene, cat. no. 0001530020) into a bottle.
e. Store at 4°C.
12. Paraformaldehyde is a white powder with a formaldehyde-like odor. It is a rapid
fixative and a potential carcinogen. When handling paraformaldehyde, wear
gloves, mask, and goggles. It is important to refer to the MSDS instructions and
institutional regulations for further information regarding storage, handling and
13. Preparation of 100 mL of 4% paraformaldehyde with 0.03 M sucrose in a fume hood:
a. Mix 4 g of paraformaldehyde powder and 80 mL of deionized water in a glass
beaker; cover with parafilm.
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302 Shefer and Yablonka-Reuveni
b. Warm the solution to 60°C with continuous stirring to dissolve the powder.
c. Allow the solution to cool to room temperature.
d. Add about 1–4 drops of 1 N NaOH, until the opaque color of the solution
e. Add 10 mL of 1 M sodium phosphate.
f. Adjust the pH to 7.2–7.4 using color pH strips.
g. Add 1.026 g of sucrose.
h. Bring volume to 100 mL.
i. Filter through a 0.45-µm disposable filter unit (Nalgene, cat. no. 0001530020)
into a bottle.
j. Store at 4°C in an aluminum-foil-wrapped bottle for no more than 1 mo.
14. Collagenase concentration, as well as the optimal time for enzymatic digestion,
should be adjusted for younger or older mice and for different strains of mice.
15. FDB and EDL myofiber isolation protocols include gentle agitation during enzy-
matic digestion. However, if maintaining quiescence of satellite cells is an
important aspect of the study, we recommend to avoid continuous agitation.
Instead dishes should be gently swirled every 10–15 min.
16. The time required for the myofiber suspension to settle (at 1g) through 10 mL of
10% HS can vary between 5 and 15 min and the investigator should adjust this
time. A prolonged period results in a preparation with more debris and remaining
single cells released from the digested tissue. Depending on mouse age, the num-
ber of rounds of myofiber settling in the 15-mL glass Corex tubes, as well as the
amount of medium in the tube, might also need to be adjusted.
17. The tissue culture dishes are dry when the bottom appears opaque white.
18. For some antibodies the cultures may be blocked for just 2–4 h at room tempera-
ture if overnight blocking is not desired.
19. For even and continuous distribution of the antibodies (both primary and second-
ary), it is recommended to place the dishes on a three-dimensional rotator
(Labline Maxi Rotator; VWR Scientific Products, cat. no. 57018-500). It is espe-
cially important when staining myofibers in 24-well multiwell dishes because
the antibody aliquots tend to rapidly accumulate at the well periphery, leading to
uneven staining across the culture.
20. DAPI is potentially harmful. Avoid prolonged or repeated exposure; we typically
dissolve the entire powder in its original container and generate a concentrated
stock solution. A ready-made DAPI reagent is available from Molecular Probes.
It is important to refer to the MSDS instructions and institutional regulations for
further information regarding storage, handling, and first-aid.
We are grateful to Monika Wleklinski-Lee and Stefanie Kästner for helpful
comments on the manuscript. ZYR thanks Stefanie Kästner and Anthony
Rivera for their valuable contributions to the FDB myofiber studies. GS thanks
Dr. Terrance Partridge and members of his research team for advice on EDL
myofiber isolation during her former studies.
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Mouse Myofiber Cultures 303
The studies described in this chapter have been supported by grants to ZYR
from the National Institute of Health (AG13798 and AG21566), the Coopera-
tive State Research, Education and Extension Service/US Department of Agri-
culture (National Research Initiative Agreement no. 99-35206-7934), and the
Nathan Shock Center of Excellence in the Basic Biology of Aging, University
of Washington. Earlier support from the Muscular Dystrophy Association and
the USDA (NRI Agreements no. 93-37206-9301 and 95-37206-2356) has
facilitated our initial studies on the isolation and culture of rat myofibers.
1. Mauro, A. (1961) Satellite cells of skeletal muscle fibers. J. Biophys. Biochem.
Cytol. 9, 493–495.
2. Hawke, T. J. and Garry, D. J. (2001) Myogenic satellite cells: physiology to
molecular biology. J. Appl. Physiol. 91, 534–551.
3. Yablonka-Reuveni, Z., Quinn, L. S., and Nameroff, M. (1987) Isolation and clonal
analysis of satellite cells from chicken pectoralis muscle. Dev. Biol. 119, 252–259.
4. Kästner, S., Elias, M. C., Rivera, A. J., and Yablonka-Reuveni, Z. (2000) Gene
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