Proc. NatL Acad. Sci. USA
Vol. 78, No. 9, pp. 5623-5627, September 1981
Cell Biology
Isolation and characterization of human muscle cells
(differentiation/contractile protein synthesis)
HELEN M. BLAU AND CECELIA WEBSTER
Department of Pharmacology, Stanford University School of Medicine, Stanford, California 94305
Communicated by Robert T. Schimke, May 11, 1981
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ABSTRACT We have developed an in vitro system for the
study of postnatal human muscle under standardized conditions.
The technique utilizes cloning to isolate pure populations of muscle
cells. By manipulating culture conditions we can maximize either
proliferation or differentiation of individual clones or of clones
pooled to yield mass cultures of muscle cells. The muscle phenotype is stable; cells can be stored in liquid nitrogen for long-term
use without loss of proliferative or differentiative potential. We
have determined proliferative capacity of muscle cells from an
analysis of clonal growth kinetics; we determined differentiative
capacity from morphological evidence (cell fusion, striations, contractions, and the appearance of acetylcholine receptors) and biochemical analysis of muscle protein synthesis (creatine kinase, aactin, tropomyosin, and myosin light chains). Our approach eliminates the variability in cellular composition that has complicated
studies of primary muscle to date. We can now study in a controlled fashion the interactions and contributions of different cell
types to the development of normal and genetically dystrophic
human muscle.
Most biochemical studies of human muscle in vitro have used
either explants in organ culture or dissociated monolayers of
primary cells. With both techniques, muscle cells are inevitably
contaminated by diverse cell types including nerve, adipocytes,
and fibroblasts (see refs. 1 and 2 for reviews). It is well known
that the ratio of muscle to nonmuscle cells influences the behavior ofthe muscle cells present (3) and varies in disease states
(ref. 4; unpublished data). The proportion of muscle cells can
be increased to 90% ofthe total cell population by preincubation
on substrates not coated with collagen (5). Nonetheless, quantitative experiments using such mixed cultures remain extremely difficult. Unlike rat and mouse for which several cell
lines exist (3-6), there are no established human muscle lines.
As a result, previous studies of pure muscle cell populations
from humans have utilized individual clones derived from single
cells to analyze clonal growth kinetics and morphology (4, 7-9).
Because attempts to produce differentiation of these cloned
muscle cells in mass cultures have met with little success (7, 10),
biochemical studies of the differentiation process have been
limited.
To date, the majority of studies of human muscle in tissue
culture have used embryonic muscle, a source with distinct
disadvantages. First, because definitive diagnosis of genetically
determined muscular dystrophies is only possible postnatally,
cells from fetuses have the uncertainty of, at best, a 50% risk
for a disease. Second, cells isolated from the muscle of individuals of different ages have been shown to differ in the differentiative properties they express when cultured in vitro (11).
Consequently, muscle from embryonic sources may not display
the characteristic pathological abnormalities of the dystrophy
in culture. This influence of developmental stage on cellular
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differentiative potential in culture is an important consideration
in the analysis of normal and dystrophic myogenesis.
In this report, we describe the isolation, growth, and expansion of clones of human muscle cells of postnatal origin and the
conditions that maximize the proliferative or differentiative capacity of these muscle cells, either as individual clones or as
pooled clones or mass cultures. The pure populations of myogenic cells can be analyzed separately or in cell mixtures of
known composition in investigations ofinduction during muscle
development, of cell-cell interactions at the neuromuscular
junction, and of the etiology of human muscular dystrophies.
MATERIALS AND METHODS
Source of Human Muscle. Muscle samples were obtained
from 17 normal patients during surgical treatment for orthopedic nonmuscle problems in accordance with the guidelines of the Human Subjects Committee of Stanford University.
Cell Culture Conditions. Growth medium (GM) contained
Ham's nutrient mixture F-10 with 0.5% chicken embryo extract
and either 20% (vol/vol) fetal calf serum (GM-1) or horse serum
(GM-2). Fusion medium (FM) contained Dulbecco's modified
Eagle's medium and 2% (vol/vol) horse serum. Conditioned
medium (CM) was GM-2 exposed to confluent human musclederived fibroblast cultures for 24 hr. filtered through a 0.2-pym
Nalgene filter, and diluted 1:1 with fresh GM-2. CM can be
stored at 40C for 2 weeks or at -70'C for 6 months. All cultures
were grown in the presence of penicillin G (200 units/ml) and
streptomycin sulfate (200 pug/ml) on a collagen substrate (calf
skin collagen, Calbiochem; 1.4 mg/ml in distilled water, autoclaved). All cells were grown at 37°C in a humidified Forma
incubator containing 5% CO2 and 95% air. Cells were stored
frozen in 10% dimethyl sulfoxide (Mallinkrodt) in horse or fetal
calf serum.
F-10 and Dulbecco's modified Eagle's media, chicken embryo extract, and fetal calf serum were obtained from GIBCO,
and horse serum was from Kansas City Biologicals. Horse
serum lots were pretested for those that would best support
fusion. Tissue culture dishes, flasks, and multiwells were from
Falcon Plastics, and penicillin and streptomycin were from
GIBCO.
Assays for Muscle Gene Expression. Creatine kinase (CPK)
activity was assayed as described (12). Synthesis of a-actin was
determined by a modification of the method of Blau and Epstein
(12). Cells were labeled for 3 hr in methionine-free medium
supplemented with [3S]methionine (250 4Ci/ml; 1102 Ci/
mmol; Amersham) and then harvested; 0.60-2.7 X 10r cpm in
35 1Lg of protein was layered in 20 p.1 on each gel. y-, f3-, and
a-actin species were resolved by fractionating labeled proteins
on isoelectric focusing polyacrylamide gels, followed by autoradiography. Relative rates of actin synthesis were quantitated
Abbreviations: CPK, creatine kinase; mU, milliunits; GM, growth medium; FM, fusion medium; CM, conditioned medium.
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5624
Cell Biology: Blau and Webster
by scanning autoradiograms with a densitometer (E. C. Apparatus, St. Petersburg, FL) and cutting out and weighing the area
under the relevant peaks. Other major contractile proteins were
identified on two-dimensional gels (13) by comigration of labeled proteins with human contractile proteins purified directly
from adult human muscle (14).
The distribution of acetylcholine receptors was assayed by
autoradiographic analysis of the binding of '"I-labeled-a-bungarotoxin to intact myotubes in culture (15, 16). Cells were labeled for 45 min in FM with '"I-labeled-a-bungarotoxin (specific activity, 95-140 Ci/mmol; New England Nuclear) at 50
nM, a concentration determined to be sufficient to saturate the
receptors. Specificity of binding was controlled by preincubating replicate dishes of cells at each time point with unlabeled
a-bungarotoxin (5 juM) for 20 min prior to labeling. For assays
of total receptor, cells were solubilized in 1 M NaOH and assayed in a Micromedic 4/600 gamma counter with 75% efficiency. Total protein was determined by the method of Lowry
et al. (17).
Isolation and Selection of Muscle Cells from Adult Tissues.
A relatively small number of cells positioned between the basement membrane and the sarcoplasmic reticulum ofadult muscle
fibers are myoblasts capable of proliferation (18). After tissue
dissociation, it is these satellite cells that give rise to clones in
culture. Postnatal muscle samples can be stored at 40C in F-10
medium for up to 24 hr prior to dissociation without adverse
effects on the yield and viability of satellite cells. For dissociation, a 0.1- to 0.3-cm3 piece of skeletal muscle tissue in F-10
medium at 40C is carefully dissected to remove as much connective tissue as possible and minced to obtain fragments
smaller than 1 mm . To remove residual debris, the fragments
are washed with F-10 three times at 40C and once at 370C. The
tissue is then dissociated for a total of 40-60 min by two or three
successive treatments with- 25 ml of 0.05% trypsin-EDTA
(GIBCO) at 370C in a Wheaton graduated trypsinization flask
with constant stirring. The cells collected in the supernatant
after each trypsin treatment are pooled and cooled to 4°C on
ice. Horse serum is added to a final concentration of 10% (vol/
vol) to terminate further protease activity. The dissociated cells
are then centrifuged (2 min; 25°C); the cell pellet is resuspended
in CM and either plated in culture or frozen in liquid nitrogen
at a density of 0.1 cm3 of tissue per ml for future use.
Enrichment of the cell population for muscle is accomplished
by preplating the cells at 37C for 20 min on a non-collagencoated dish, a substrate to which fibroblasts preferentially adhere (5). Because cell counts and efficiency of plating cannot
be accurately determined for human postnatal muscle cells due
to debris resulting from myotube destruction during tissue dissociation, unattached cells are plated at a range of densities approximated from the expected satellite cell number, 107/cm3
of human muscle (19). Those yielding between 5 and 50 clones
per 60-mm collagen-coated plate are used. Cultures are maintained in 2-3 ml of CM; they are not fed for the first 3-6 days
and are fed only every 4 days thereafter. For future use, cloned
cells can be frozen at a density of 5 x 106/ml. The efficacy of
the isolation and selection procedure is demonstrated by the
fact that 96-100% of 828 clones from four different samples
were myogenic (see Table 1).
RESULTS
Clonal Analysis: Generation of Pure Muscle Cell Populations. A major problem in the study of postnatal human cells
in culture is their limited longevity-approximately 45 doublings compared to 60-70 for muscle cells from 80-day fetuses
(4). Our approach works within the time frame imposed by se-
Proc. Nad Acad. Sci. USA 78 (1981)
nescence. Clones containing 1000-2000 cells are harvested
prior to fusion; groups of three are pooled, grown to 60-80%
confluence in GM-1 in order to prevent initiation of myogenesis, and then frozen for long-term storage at a density of 5
X 106 cells per ml. Simultaneously, individual clones are tested
separately for their myogenic potential by plating a few cells of
each in 16-mm tissue culture wells and scoring these for fusion.
This approach ensures that the frozen cells are homogeneously
muscle, have not initiated differentiation, and spend a minimum of time in culture prior to use.
Potential Yield of Muscle Cells per Biopsy. The yield of
muscle cells from a small biopsy is sufficient for many kinds of
biochemical and morphological analyses. From a 0. 1-cm3 piece
of tissue, of which 50% is connective tissue and fat, 5 x 103
viable, proliferative satellite cells can be obtained. In our experience, each satellite cell is capable of giving rise to at least
1 X 107 cells, equivalent to one confluent T-75 flask or approximately 2 mg of protein.
Proliferative Capacity of Frozen Stored Muscle. The proliferative capacity of frozen cells was compared with that of
fresh cells. Clonal growth kinetics were determined by randomly selecting and circling clones and then visually counting
the number of cells in each on subsequent days by using an inverted microscope with phase-contrast optics. Although the
range in cell number per clone on day 4 suggests a high degree
of heterogeneity, the growth curves are remarkably similar (Fig.
1). The apparent heterogeneity simply reflects the time in culture required for individual cells to become established and
begin proliferation. For example, the clones indicated by A and
4 have lag times of 2 and 4 days, respectively, but similar growth
kinetics.
Table 1 shows data for the growth properties of clones of
freshly dissociated and frozen-thawed cells derived from four
muscle biopsy specimens. The ranges in doubling times (12-20
hr) and in the lag times prior to the initiation of proliferation
(2-4 days) suggest that there is variability among clones of the
same sample, of different anatomic muscles, and of different
individuals with respect to these growth values. However, of
importance is the observation that, on average, fresh and frozen
cells behave similarly. In addition, cells frozen at the same density (0.1 cm3 of dissociated tissue per ml) but in small volumes
(0.2 ml), are not adversely affected with respect to either their
growth or their differentiative properties and exhibit only a
slight decrease in total cell yield (data not shown). These results
demonstrate that dissociated cells can be frozen in numerous
aliquots for use at different times.
a
b
1000
~~~~~~500-
7
/
6
6 5
-
416 0
Oi
4
~~~50-
2
/
3
0
I~~~~~~~~1
1~ ~~~~~~~1
14]
A~~~~~~~~~
nm
0 24 6 81012 1416
0
8
6
4
2
Time in culture, days
Cells/clone
FIG. 1. Kinetics of growth of clones of fresh cells. The number of
cells in 20 individual clones was counted on day 4 (a) and daily thereafter (b); growth curves for six representatives are shown. The clones
with the least and most cells are indicatd by 4 and A, respectively.
Cell Biology: Blau and Webster
Proc. NatL Acad. Sci. USA 78 (1981)
5625
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Table 1. Clonal growth and differentiation of fresh and
frozen-thawed cells
Muscle
Aliquot
Doubling
Lag
Fusion,
sample* size, mlt
time, hrt
% off
time, hrt
Fresh:
1
20.7 ±0.8
42.2± 4.7
2
16.8 ± 0.5
89.2 ± 2.6
97 (151)
3
14.3 ± 0.6
55.0 ± 3.6
98(98)
4
13.5 ± 0.4
69.0 ± 4.4
100 (107)
Mean
16.3 ± 1.6
63.9 ± 15.8
98 ± 1
Frozen:
2
1.0
14.4 ± 0.8
62.9 ± 0.8
100 (50)
2
0.2
14.9 ± 0.7
70.9 ± 3.9
96 (53)
3
1.0
12.8 ± 0.4
72.4
3.8
100 (156)
3
0.5
13.1 ± 0.4
70.8 ± 4.6
99 (121)
3
0.2
12.6 ± 0.7
61.8 ± 3.8
99 (92)
Mean
13.6 ± 0.5
67.8 2.2
99 ± 1
* Each sample was from a separate muscle biopsy. Growth kinetics for
individual clones of samples 3 and 4, two biopsies from the same individual, are shown in Fig. 1. Sample 2 was from the biceps femoris;
all other samples were from the vastus lateralis.
t Cells were frozen at the same density (0.1 cm3 of dissociated tissue
per ml) but in different volumes (0.2, 0.5, 1.0 ml).
t Cell doubling and lag times prior to initiation of proliferation are
expressed as mean ± SEM for at least 10 clones in each case.
§ The percentage fusion is the proportion of total colonies that differentiated as muscle. The number of clones scored is in parentheses.
Differentiative Capacity of Frozen Stored Muscle. All assays of differentiation described below were performed with
mass cultures of frozen-thawed muscle cells. When plated at
greater than 1 x 105 cells per 35-mm dish, frozen cells, like
fresh cells, reached confluence and fused extensively until 61
± 5% of the cell nuclei were found within myotubes. The source
ofthe serum markedly affected the differentiation of these cells
(Fig. 2). We have exploited this finding to grow parallel cultures
of cells in low concentrations (2%, vol/vol) of horse serum to
test for differentiative capacity, while maintaining the majority
in a high concentration (20%, vol/vol) of fetal calf serum to
promote proliferation. Decreases in serum concentration have
similarly been used to promote differentiation of myogenic cells
of other species (ref. 20; S. D. Hauschka, personal communication). The appearance of striations (Fig. 3) and rhythmic contractions are further evidence of differentiation.
To assess differentiation biochemically, the synthesis of aactin, was compared to the synthesis of nonmuscle (& and yactins. Prior to differentiation, a-actin represented 20% of the
total actin synthesized but at 7 days after a change to FM, when
approximately 65% ofcells are found in myotubes, a-actin comprised 40% of the total actin synthesized (Fig. 4). We have observed similar increases in a-actin relative to total actin synthesis in primary cultures of rat and chicken [from 13% and 17%
to 22% and 32%, respectively (unpublished data)], in good
agreement with the findings of Garrels and Gibson (21) and
Rubenstein and Spudich (22). The ratio of a- to (3 and y-actins
in our differentiated cultures is not quite as high as that found
in extracts of biopsied adult human muscle (data not shown).
This finding is not surprising, given the persistence of unfused
myoblasts in these cultures, and agrees with observations in
pure populations of rat myogenic cells (21). Thus, in the course
of in vitro growth and differentiation of human muscle, the synthesis of a-actin increases 2-fold and this isoform becomes the
predominant actin species (Table 2).
The CPK activity increased 18-fold from 140 to 2515 milliunits (mU)/mg of protein after the cells had been in FM for
5 days (Table 2). This increase was largely due to de novo synthesis of the muscle-specific isozyme (data not shown). Com-
FIG. 2. Differentiation of frozen-thawed cells in mass culture.
(Upper) Cells grown in medium containing horse serum differentiate.
(Lower) Cells grown in medium containing fetal calf serum continue
to proliferate. (x70.)
parable CPK specific activities, 720 and 1000 mU/mg, have
been reported in well-differentiated L6 and rat primary cells,
respectively (23).
Total acetylcholine receptors increased 33-fold during the
course of myogenesis in vitro. This increase proved to be largely
due to the production of unclustered receptors, as shown by
autoradiography (Fig. 5). The rapid increase (day 3) and subsequent decrease (day 7) in receptors shown in Table 2 were
observed in three independent experiments and are in good
agreement with the findings of Prives et al. (24) for chicken
muscle which differentiates in the absence of nerve. To stabilize
receptors on the chicken myotube surface, a neural factor appears to be necessary. Peak amounts of receptor synthesized in
our human cell cultures are comparable to those observed in
.
.-i
0-1'I.
3.
Fri ,
..............................
.---*- ....
FIG. 3. Striations in frozen-thawed cells in mass culture. Four
days after growth in FM, cells were fixed in 2% glutaraldehyde in
Hanks' balanced salt solution for 20 min at 37TC and stained with 2%
orcein in 45% propionic acid. (x380.)
5626
B
Proc. Natl. Acad. Sci. USA 78 (1981)
Cell Biology: Blau and Webster
iL
A
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FIG. 4. Synthesis of actins by frozen-thawed cells in mass culture.
Myoblasts (A) and myotubes (B) were labeled with [55S]methionine on
day 0 and day 7 after change of medium from GM-1 to FM. Proteins
were separated by electrophoresis on isoelectric focusing gels and visualized by autoradiography (Upper) and densitometric scanning
(Lower).
muscle of other species grown in vitro (6, 24-26).
The synthesis of additional muscle-specific components was
determined by the fractionation of labeled proteins on two-dimensional gels (Fig. 6). Most dramatic is the initiation of synthesis of the tropomyosins and myosin light chains, similarly
associated with the differentiation of muscle of other species
(27-30). The isoforms observed upon differentiation of our cultures in vitro were the adult human forms; ['S]methionine-labeled proteins comigrated with accumulated myosin light
chains and tropomyosins purified from adult muscle and visualized by Coomassie brilliant blue staining (data not shown).
Numerous other changes in the pattern of protein synthesis
occurred but have not yet been characterized.
DISCUSSION
Our procedure for the growth of human postnatal muscle in
culture capitalizes on the presence of satellite cells, the small
percentage of cells in mature muscle fibers capable of proliferation and muscle regeneration (31, 32). We have optimized
conditions for growth, frozen storage, and differentiation of human cells within 45 doublings, the time frame imposed by senescence (4, 33). The total cell yield from a typical 0. 1-cm3 tissue
biopsy specimen results in a minimum of 10 g of protein, an
amount adequate for many kinds of biochemical and morphological analyses.
FIG. 5. Distribution of acetylcholine receptors on frozen-thawed
cells in mass culture. Cells were labeled with "2'-labeled-a-bungarotoxin and processed for autoradiography. (Upper) Myoblasts and myotubes as shown with phase-contrast optics. (Lbwer) The same field in
dark-field illumination reveals the pattern of silver grains and the
location of receptors. Arrows indicate undifferentiated myoblasts
which do not have receptor. (x 100.)
The proliferative and differentiative capacities of fresh and
frozen muscle cells have been ascertained. The observed range
in doubling times and lag times is likely to be due to minor
differences in cell density and growth conditions in culture (refs.
7, 8, and 34; unpublished data). Growth kinetics, on the other
hand, were more consistent among clones of a given human
sample than those observed for primary or secondary rat cells
(5). The ability to store small aliquots offrozen cells without loss
of proliferative or differentiative capacity provides flexibility,
permitting replication of experiments, easy exchange of mate-
Table 2. Expression of muscle functions in mass cultures
AcChoR
a-Actin
synthesized, %
fmol/mg
CPK mU/mg* protein*
Days in FM total actin
0
1
2
3
5
7
0.2
0.4
140
225
795
1730
2515
2450
3.1
15.4
48.2
89.0
103.5
75.6
33
18
2
Increase, -fold
Cells were plated at 1 x 105 per 35-mm dish and grown as mass cultures in GM-1 until nearly confluent. Days indicate elapsed time after
change to FM. All studies were performed in duplicate; mean values
are shown.
* An average 35-mm dish contained 0.2 mg of protein. AcChoR, acetylcholine receptor.
A
*Tm
?tTm
\Lc
Lc\Lc
ALc
\; C\ L c
FIG. 6. Patterns of protein synthesis in frozen-thawed cells in
mass culture. Proteins synthesized by myoblasts (Left) and myotubes
(Right). In each case, four pooled clones were labeled with
[(5S]methionine and the proteins were assayed by two-dimensional gel
electrophoresis and autoradiography. Arrows indicate muscle proteins: a-, 3-, and y-actin, tropomyosins (Tm), and four myosin light
chains (Lc), identified by comigration with purified human muscle contractile proteins.
Cell Biology: Blau and Webster
rial among laboratories, and comparative studies of properties
of normal and dystrophic muscle under identical conditions.
Although the specific reasons why our conditions are effective remain unknown, it is clear that our procedure results in
a degree of differentiation of mass cultures previously only observed in individual clones (7, 10). Our method differs from
others in that F-10 and CM are used only to promote proliferation and not for differentiation. For optimal differentiation,
we have defined a low nutrient medium with a high calcium
concentration which routinely results in the formation of
striated, contractile myotubes at incubator CO2 levels of 5%.
Nutrient deprivation appears to enhance fusion. When F-10 is
replaced by Dulbecco's modified Eagle's medium and the
serum concentration is decreased from 20% to 2%, extensive
differentiation occurs. In addition, the 6-fold higher calcium
concentration of the latter medium compared to F-10 may be
critical. pH also affects differentiation. CO2 levels >5% are inhibitory; again, the 3-fold higher bicarbonate concentration in
Dulbecco's modified Eagle's medium relative to F-10 may aid
in maintaining a stable pH.
Under our conditions, human muscle, like muscle of other
species (6, 20-29), exhibited marked increases in the synthesis
of a-actin, CPK, and acetylcholine receptor. In addition, synthesis ofthe myosin light chains and tropomyosin were initiated
in the course of myogenesis in vitro. It should be noted that
these proteins, which are characteristic of differentiated muscle, are the adult human forms because they comigrate on gels
with contractile proteins purified directly from adult human
muscle tissue. -In addition, the observed increase in CPK activity is due to the de novo appearance of the muscle-specific
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isozyme.
A distinct advantage of our approach is that it uses postnatal
muscle, permitting application of the methods to dystrophic
muscle obtained from individuals with diagnosed genetic muscle disease rather than from fetuses at risk for a disorder. Postnatal satellite cells are also more likely to have matured sufficiently in vivo to express the relevant muscle properties in vitro
(11).
In summary, we have determined the requirements for a
reproducible procedure for the isolation, growth, and differentiation of human muscle cell populations for in vitro study.
These isolated human muscle cells will be invaluable to studies
of cell-cell interactions, permitting identification of functions
intrinsic to muscle and those induced or contributed by nerve,
fibroblasts, or the extracellular matrix in the course of normal
and genetically dystrophic human muscle development.
We are grateful to Dr. D. Yaffe and Ms. B. Dieckmann for encouragement and advice early in this work, to Drs. E. Bleck and L. Rinsky
for muscle samples, to Drs. S. Guttman, S. Packman, and P. Byers for
critical-reading of the manuscript, and to Ms. C. Spain for expert secretarial assistance. This work was supported by grants to H. B. from the
Proc. NatL Acad. Sci. USA 78 (1981)
5627
Muscular Dystrophy Association of America, National Institutes of
Health Grant GM 26717, and Basil O'Connor Starter Grant 5-179 from
the March of Dimes-National Foundation.
1. Rowland, L. P. (1976) Arch. Neurol. 33, 315-321.
2. Rowland, .L. P. (1980) Muscle and Nerve 3, 3-20.
3. Yaffe, D. & Saxel, 0. (1977) Nature (London) 270, 725-727.
4. Hauschka, S. D., Linkhart, T. A., Clegg, C. & Merfill, G. (1979)
in Muscle Regeneration, ed. Mauro, A. (Raven, New York), pp.
311-322.
5. Richler, C. & Yaffe, D. (1970) Dev. BioL 23, 1-22.
6. Noble, M. D., Brown, T. H. & Peacock, J. H. (1978) Proc. Nati
Acad. Sci. USA 75, 3488-3492.
7. Hauschka, S. D. (1974) Dev. Biol 37, 329-344.
8. Hauschka, S. D. (1974) Dev. Biot 37, 345-368.
9. Hauschka, S. D., Hainey, C., Angello, J. C., Linkhart, T. A.,
Bonner, P. H. & White, N. K. (1977) in Pathogenesis of Human
Muscular Dystrophies, International Congress Series, ed. Rowland, L. P. (Excerpta Medica, Amsterdam), pp. 834-855.
10. Holtzer, H., Jones, K. W. & Yaffe, D. (1975)J. Neurot Sci. 26,
115-124.
11. Koenig, J. & Vigny, M. (1978) Nature (London) 271, 75-77.
12. Blau, H. M. & Epstein, C. J. (1979) Cell 17, 95-108.
13. O'Farrell, P. H. (1975) 1. BioL Chem. 250, 4007-4021.
14. Kielley, W. W. & Harrington, W. F. (1960) Biochem. Biophys.
Acta 41, 401-421.
15. Blau, H. M. & Kafatos, F. C. (1978)J. Cell Biol 78, 131-151.
16. Teng, N. N. H. & Fiszman, M. Y. (1976)J. Supramot Struct. 4,
381-387.
17. Lowry, 0. H., Rosebrough, N. J., Farr, A. L. & Randall, R. J.
(1951)J. Biot Chem. 193, 265-275.
18. Mauro, A. (1961)J. Biophys. Biochem. Cytot 9, 493-495.
19. Muir, A. R. (1970) in Regeneration of Striated Muscle and Myogenesis, International Congress Series, eds. Mauro, A., Shafiq,
S. A. & Milhorat, A. T. (Excerpta Medica, Amsterdam), pp.
91-100.
20. Yaffe, D. & Saxel, 0. (1977) Differentiation 7, 159-166.
21. Garrels, J. I. & Gibson, W. (1976) Cell 9, 793-805.
22. Rubenstein, P. A. & Spudich, J. A. (1977) Proc. NatL Acad. Sci.
USA 74, 120-123.
23. Shainberg, A., Yagil, G. & Yaffe, D. (1971) Dev. Biol 25, 1-29.
24. Prives, J., Silman, I. & Amsterdam, A. (1976) Cell 7, 543-550.
25. Betz, H. & Changeux, J.-P. (1979) Nature (London) 278,
749-752.
26. Libby, P., Bursztajn, S. & Goldberg, A. L. (1980) Cell 19,
481-491.
-27. Garrels, J. I. (1979) Dev. BioL 73, 134-152.
28. Devlin, R. B. & Emerson, C. P., Jr. (1978) Cell 13, 599-611.
29. Devlin, R. B. & Emerson, C. P., Jr. (1979) Dev. BioL 69,
202-216.
30. Stockdale, F. E., Raman, N. & Baden, H. (1981) Proc. Natl Acad.
Sci. USA 78, 931-935.
31. Konigsberg, U. R., Lipton, B. H. &,Konigsberg, I. R. (1975) Dev.
Biol 45, 260-275.
32. Bischoff, R. (1974) Anat. Rec. 180, 645-662.
33. Hayflick, L. (1965) Exp. Cell Res. 37, 614-636.
34. Konigsberg, I. R. & Hauschka, S. D. (1965) in Reproduction:
Molecular, Subcellular, and Cellular, ed. Locke, M. (Academic,
New York), pp. 243-290.