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Secondary myogenesis of normal muscle produces abnormal myotubes.

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THE ANATOMICAL RECORD 204:199-207 (1982)
Secondary Myogenesis of Normal Muscle Produces Abnormal
Myotu bes
MARCIA ONTELL, DONNA HUGHES, AND DIANNA BOURKE
Department of Anatomy and Cell Biology, University of Pittsburgh School of Medicine,
Pittsburgh, Pennsylvania 15261
ABSTRACT
The extensor digitorum longus muscles of 2-, 4-, and 12-weekold 129-ReJ mice were subjected to homotopic, whole-muscle transplantation.
Subsequent to myofiber necrosis and phagocytosis, a new population of myotubes
was produced. The three-dimensional cytoarchitecture of these newly formed myotubes was determined in spaced, serial, ultrathin sections. Myotubes, which for
long distances along their length appeared to be separate and discrete, were found
to branch and recombine, forming a complex syncytium.
It has been well established that mammalian striated muscle is capable of regeneration
in response to injury or disease. The regeneration may involve repair of existing fibers, or,
when the trauma to the muscle is sufficient to
cause whole-myofiber necrosis, de novo formation of myotubes may occur. This wave of
secondary myogenesis occurs, as does myogenesis during fetal development, by the fusion
of mononucleated cells (Carlson, 1973; Lipton
and Schultz, 1979).It is generally believed that
the cells responsible for this regenerative response are myosatellite cells, mononucleated
cells found sandwiched between the basal lamina and the sarcolemma of intact muscle fibers
(Mauro, 1961). However, the possibility that
other types of cells may also participate in secondary myogenesis has not been entirely excluded (Carlson, 1973; Snow, 1977a).
One of the most widely studied models of
secondary myogenesis is the whole-muscle
transplantation system, first described by Studitsky and Bosova (1960) and Bosova (1962)
and modified by Carlson and Gutmann (1974).
In this system the entire muscle, cut free of its
tendons, nerves, and vascular supply, is placed
into a suitably prepared bed. The tendons of
the transplanted muscle are attached to rigid
structures (bones or other tendons), restoring
the muscle to its original resting length. No
microsurgical procedures are performed to anastomose the blood vessels or nerves of the
grafted muscle to the surrounding vasculature
or nerves. With the exception of a few fibers a t
the periphery of the implant, all of the myofibers become ischemic and undergo total fiber
necrosis (Carlson and Gutmann, 1975;Lischka
et al., 1977).Macrophages remove the necrotic
0003-276W82/2043-0199$03.00
1982 ALAN R LISS, INC.
fibers (Snow, 1977a1, leaving only their relatively intact basal lamina (Carlson et al., 1979;
Hansen-Smith and Carlson, 1979). New myotubes are formed, within the old basal lamina1
tubes, by a secondary wave of myogenesis
(Hansen-Smith and Carlson, 1979; Carlson et
al., 1979). Ultimately, each new myotube becomes enclosed in its own basal lamina, and
the old basal lamina is lost.
Despite the large number of morphological
(cf. Carlson, 1978), physiological (cf. Faulkner
et al., 1980; Hakelius et al., 19751, and histochemical (Maxwell et al., 1978; Hakelius et al.,
1975; Carlson and Gutmann, 1975) studies of
whole-muscletransplants, little is known about
the three-dimensional cytoarchitecture of the
regenerating myotubes formed in this system.
In the present study, spaced serial ultrathin
sections have been used to define the myotubes’
cytoarchitecture.
MATERIALS AND METHODS
Orthotopic whole muscle transplants (Carlson and Gutmann, 1974) were performed on
the extensor digitorum longus muscles of Metafane (Pitmann-Moore)-anesthetized2-, 4-, and
12-week-old 129-ReJ mice (obtained from a
normal colony maintained at Jackson Laboratory). The muscle was removed by severing
its tendons, being careful not to leave any muscle attached to the tendon stumps. After soaking the muscle at room temperature for 15-20
minutes in 0.75% Marcaine (Breon Laboratories), a known myotoxic agent (Benoit and Belt,
1970;Libelius et al., 1970)which prevents surReceived March 18,1982;accepted July 9,1982
200
M. ONTELL, D. HUGHES, AND D. BOURKE
viva1 of the peripheral myofibers (Carlson,
19761, the muscle was replaced into its original
muscle bed and sutured to the tendon stumps.
Operated mice were allowed to move freely
about the cage and were given fresh tetracycline (400 mg/liter) in their water daily for up
to 5 days following surgery.
Fig. 1. Light micrograph of a transverse section through
the widest girth of a control extensor digitorurn longus muscle taken from a 2-week-old mouse. The polygonal-shaped
fibers are arranged into discrete fascicles. Toluidine blue.
x 90.
At various time periods after surgery (1, 3,
5, 7, and 20 days), the mice were killed by
cardiac puncture. Extensor digitorurn longus
muscles were exposed and bathed, in situ, for
30 minutes in 2.0% gluteraldehyde in 0.125 M
cacodylate buffer (pH 7.25). The transplant was
excised in toto, placed in fresh fixitive for 2
hours, postfixed in 2.0% osmium tetroxide in
cacodylate buffer, dehydrated in ethanol and
embedded in Epon 812. Epon blocks, containing entire muscles, were placed on a sliding
microtome and oriented for transverse sectioning. Sets were cut consisting of ten 15-pm-thick
sections and one 6-pm-thick section. The 6-pmthick sections were mounted on glass slides
and studied with a phase-contrast microscope.
All 15-pm-thick sections were cleared in Epon
between two layers of polystyrenefilm and cured
in an oven a t 60°C (Davidowitz et al., 1976).
Selected 15-pm-thick sections were adhered to
a preformed Epon block, and semithin and ultrathin sections were cut using an ultramicrotome. Semithin sections were stained with toluidine blue. Ultrathin sections were collected
on slot copper grids, stained with uranyl acetate and lead citrate (Reynolds, 1963), and observed using a Philips 300 electron microscope.
This permitted the study of chosen fascicles of
regenerating myotubes a t known intervals
along their length. Similar studies were performed on normal muscles taken from 2-, 4-,
and 12-week-old mice.
Fig. 2. Light micrograph of a transverse section through the widest girth of a 1-day-old,Marcaine-treated,
orthotopically transplanted extensor digitomm longus muscle, performed on a 2-week-old mouse. The muscle
fibers are rounded and swollen, and appear necrotic. Toluidine blue. x 90. Inset X 180.
201
SECONDARY MYOGENESIS
The cytoarchitecture of regenerating fibers
was initially examined in early transplants (7
days postoperative) of young (2- and 4-weekold) animals and was subsequently examined
in 20-day transplants. In order to determine
whether the cytoarchitecture of the regenerated fibers was related to the young age of the
animals at the time of transplantation, similar
studies were performed on 12-week-old mice.
RESULTS
The sequence of degeneration and regeneration of transplanted normal mouse muscle was
essentially similar to what has been described
Fig. 3. Light micrograph of a transverse section through
the widest girth of a T-day-old, Marcaine-treated, orthotopically grafted extensor digitorurn longus muscle, performed
on a 2-week-old mouse. The regenerating fibers are found
throughout the width of the graft. The myotubes in the graft
are arranged into discrete fascicles. They are smaller than
the myofibers in the control muscle (Fig. 1).Toluidine blue.
Fig. 5. Electron micrograph showing a necrotic fiber (N)
found in the central core of the 7-day-old transplanted muscle Seen in Figure 4. The necrotic fiber displays marked
coagulation necrosis, and both phagocytic cells (arrow) and
between the cOaWlurn and
myotubes (M) are found
the Persistent basal lamina (arrowhead). N, necrotic myofiher. Uranyl acetate and lead citrate.
59200.
x 90.
Fig. 4. Phase micrograph of a transverse section through the widest girth of a typical 7-day-old orthotopic,
whole-extensor digitorurn longus transplant performed on a 12-week-old mouse. A peripheral ring of regenerating myotubes surrounds the central core of necrotic fibers. x 90.
202
M. ONTELL, D. HUGHES, AND D. BOURKE
Figs. 615. A typical group of regenerating myotubes
found in the 7-day-old graft, seen in Figure 3,is followed in
spaced, serial, ultrathin sections. Figure 7 is 30 pm distal
to Figure 6 and 15 pm proximal to Figure 8.Figure 9 is 15
pm distal to Figure 8 and 15 pm proximal to Figure 10.
Figure 11 is 60 pm distal to Figure 10 and 80 km proximal
to Figure 12.Figure 13 is 140 km distal to Figure 12 and
15 pm proximal to Figure 14.Figure 15 is 15 pm distal to
Figure 14. Over a distances of 400 pm, three branching
points are seen, interconnecting what appear to be, in single
three independent myotubes
sections (Figs.6,10,11,12,13),
(1,2, 3). Close to the branching point, one diameter of the
myotube elongates (Figs. 8,15).Subsequently a constriction
(arrows)occurs perpendicular to the long diameter (Figs. 7,
9,14).Ultimately, two myotubes are formed, each in its own
basal lamina (Figs. 6,10,13). Uranyl acetate and lead citrate. x 1,900.
SECONDARY MYOGENESIS
in the rat (Carlson et al., 1979); however, in
the mouse both processes occurred more rapidly. Within 24 hours after transplantation,
rounded, swollen myofibers were seen throughout the muscle. (Compare control muscle in
Fig. 1 with 1-day transplant in Fig. 2.) All of
the fibers showed necrotic changes along their
entire lengths (Fig. 2). In young mice (2-4weeks
old), macrophages were found within necrotic
fibers located in the center of the graft by 3
days after transplantation, and regenerated
myotubes extended into the center of the graft
by 5 days postoperative. In older mice, with
larger muscles, the core remained necrotic for
longer periods, and regeneration in the center
of the graft was delayed. At low magnification,
in the light microscope, the 7-day grafts (Fig.
3) performed on young mice were virtually indistinguishable from control muscles (Fig. 1)
in that they were packed with myofibers arranged into discrete fascicles. However, a t
higher magnification, the nuclei of the regenerated fibers, unlike the nuclei in control fibers, were seen to be centrally located. Sevenday grafts performed on older animals contained regenerated myotubes only in their periphery (Fig. 4). Fine structural examination
of the center of the grafts, where necrotic fibers
persisted, revealed both macrophages and crescent-shaped immature myotubes sandwiched
between the basal lamina of the necrotic fiber
and the necrotic coagulum (Fig. 5).
During the initial phase of this study the
cytoarchitecture of the myotubes of 7-day-old
grafts performed on young mice was determined. Fascicles composedof mature myotubes
(i.e., myotubes packed with myofibrils), surrounded by their own, newly formed, basal
laminae, were followed in closely spaced ( s 15
pm), serial, ultrathin sections for distances
along their length (Figs. 6-15). Branching and
recombination of regenerating myofibers was
repeatedly observed (Figs. 7, 9,141 in multiple
203
randomly chosen fascicles across the width of
each graft. The branching example chosen for
illustration (Figs. 6-15) was taken from a segment of the graft that was approximately 3%
of the graft’s total length. It is entirely possible
that the branching pattern was even more
complex than indicated in the illustrated segment.
Approaching the branching region, one diameter of the regenerating fiber would elongate (Figs. 8, 15). Closer to the branch point,
a constriction of the myofiber would occur, perpendicular to the fiber’s long diameter (Figs.
7,9,14). The basal lamina followed the contour
of the constriction. Gradually, the constricted
area became increasingly attenuated, resulting in the fiber’s giving rise to two “daughter”
fibers (Figs. 6, 10, 13), each in its own basal
lamina. No more than two daughter fibers were
seen at each branch point. No accumulations
of specialized organelles characterized the region of the branching (Figs. 16, 17). A similar
branching and recombination of myofibers was
also found in 20-day-old transplants performed
on young animals (not shown). The branched
regenerating fibers showed no evidence of any
degenerative alterations.
In order to determine whether the branched
regenerating fibers were a function of the immaturity of the muscle at the time of transplantation (2 and 4 weeks postnatal), the cytoarchitecture of the regenerating myotubes
found in a 7-day transplant, performed on 12week-old mice, was studied. A similar branching pattern was noted.
In order to determine how early in the regenerative process the branching pattern was
established, fascicles of necrotic fibers that displayed immature, crescent-shaped, regenerating myotubes wedged between the muscle
coagulum and the old basal lamina (Figs. 5,
18)were followed in a similar series of spaced,
ultrathin sections. Even at this stage, branch-
204
M. ONTELL, D. HUGHES, AND D. BOURKE
DISCUSSION
Fig. 16. Higher magnificationofbranchingregion of myotube 1 + 2, seen in Figure 7.A constriction (arrow) occurs
in a plane perpendicular to the elongated diameter of the
myotube. No specialized organelles are seen at the branching point. Uranyl acetate and lead citrate. x 6,900.
Fig. 17. Higher magnificationof branching region of myotube 3 + 2, seen in Figure 14. A constriction (arrow) occurs
in a plane perpendicular to the elongated diameter of the
myotube. No specialized organelles are seen at the branching point. Uranyl acetate and lead citrate. x 6,900.
ing and recombination were observed between
the myotubes that shared a single, old, basal
lamina1 tube (Figs. 19, 20).
No branching myofibers were found in any
of the control muscles.
Original reports of successful regeneration
of muscle fibers following whole-muscle transplantation (Studitsky and Bosova, 1960; Bosova, 1962)have stimulated a renewed interest
in skeletal muscle regeneration (for review see
Carlson, 1978). Despite the widespread recognition that it is possible to replace necrotic
myofibers by new myotubes, produced by a second wave of myogenesis, there are few electron
microscopic studies of regenerating myotubes
in whole-muscle transplants (Carlson et al.,
1979; Hansen-Smith and Carlson, 1979;
Schmalbruch, 1977). While single, ultrathin
section studies have indicated that regenerating fibers appear to be identical to normal
developing muscle, in the present report the
application of a spaced, serial, ultrathin sectioning technique has clearly demonstrated that
a substantial percentage of the myotubes formed
as a result of secondary myogenesis undergo
complex branching and recombination, resulting in cytoplasmic continuity of myotubes that
for extensive regions along their length appear
to be independent myotubes. Comparison of the
cytoarchitecture of these branched myotubes
with similar studies of spaced, serial, ultrathin
sections of normal developing myotubes reveals that during no stage of fetal (Kozeka and
Ontell, unpublished results) or neonatal development (Ontell, 1977) are normal muscle
fibers branched, and no evidence of myofiber
branching has been found in the control muscles used in this study.
In the present study, the whole-muscle
transplants performed in younger mice have
regenerated at a much more rapid rate than
in older mice. It has not been determined
whether this difference is the result of the size
of the transplanted muscle and/or of an agerelated ability of the transplanted muscle to
undergo regeneration. Clearly, the smaller the
Fig. 18. Regenerating myotube (M)found closely apposed to a necrotic myofiber (N).
The myofilamenta are clearly
visible (arrow). Uranyl acetate and lead citrate.
X
28,350.
Figs. 19,20. Spaced, serial, ultrathin sectionsofa typical
group of necrotic myofibers found in the central region of a
7-day-old, orthotopically transplanted extensor digitorum
longus muscle, performed on a 12-week-old mouse. Figure
18 is
15 pm proximal to Figure 19. A single crescentshaped myotube (A) is found sandwiched between the basal
lamina and the necrotic coagulum of fiber 1(Fig. 18).Fifteen
microns distal to Figure 18,myotube A has branched giving
rise to two daughter myotubes (B,C) (Fig. 19). Uranyl acetate and lead citrate. X 1,900.
-
SECONDARY MYOGENESIS
205
graft, the more rapid its revascularization, and
it has been demonstrated that revascularization is necessary for secondary myogenesis
(Hansen-Smith et al., 1980). However, if satellite cells are the source of new myofibersfound
in transplanted muscle, then the younger muscle may have an advantage, because it has been
repeatedly demonstrated that the frequency of
myosatellite cells in normal muscle decreases
with age (Allbrook et al., 1971; Schultz, 1974;
Ontell, 1974;Snow, 1977b).Whatever the cause
of the differences in rates of regeneration between the younger and older murine muscles
used in this study, the branching phenomenon
occurs in both groups. Therefore, branching is
unrelated to the age of the muscle a t the time
of transplantation.
The ultimate fate of the branched striated
myotubes has not yet been determined. The
branched myotubes found in the longest-term
studies reported in the present paper (20 days
postoperative) show no evidence of degenerative changes. It should be noted that motor end
plates are regularly encounted by 14 days postoperative in these transplants (Ontell, unpublished results). The cytoarchitecture and pattern of motor innervation of the regenerating
fibers in longer-term transplants (100 days) are
currently under investigation.
The use of spaced, serial, ultrathin sections
to define the cytoarchitecture of myotubes
formed by a secondary wave of myogenesis is
particularly beneficial, because multiple myotubes are formed in a single, old, basal laminal tube. It is not possible at the light microscopic level of resolution to determine whether
the newly formed myotubes exhibit continuity
with each other or whether they are merely
closely apposed (Schmalbruch, 1976). Whereas
branching may be suggested in single ultrathin longitudinal sections, a section through a
myotube with an irregular contour may suggest branching when, in fact, the myotube is
only indented (cf. Ontell and Feng, 1981).
The present study is the first application of
a serial ultrathin sectioning technique to the
problem of defining the cytoarchitecture of normal regenerating myotubes. Jirmanova and
Thesleff (1972), studying regeneration after
methyl bupivacaine treatment, and Schmalbruch (1976), studying regeneration subsequent to injection with hot Ringer’s solution,
have reported what appears in single, longitudinal, ultrathin sections to be fusion between
adjacent myotubes.
In a previous study, using spaced, serial, ultrathin sections, it has been demonstrated that
206
M. ONTELL, D. HUGHES. AND D. BOURKE
regenerating myofibers that form spontaneously (i.e., without secondary trauma) in
murine dystrophic muscle display a similar
pattern of branching and recombination (Ontell and Feng, 1981).At that time, it could not
be determined whether branching was unique
to dystrophy or whether it was a characteristic
of any or all regenerating myofibers. The present study has established that the branched
regenerating fibers found in murine dystrophic
muscle do not reflect an alteration in the regenerative pattern produced by the disease,
since healthy muscle (obtained from mice derived from a normal colony) can produce regenerating myotubes with the same type of
branched cytoarchitecture when subjected to
whole-muscle transplantation. Interestingly,
cultures of normal muscle produce myotubes
with a branched pattern (Murray, 1960).
Since it is assumed that the myotubes formed
during secondary myogenesis are formed, as
are the myotubes during primary myogenesis
(fetal development), by the fusion of mononucleated cells into a multinucleated syncytium
(Snow, 1977a), i t is interesting to speculate
about those factors that may be responsible for
the cytoarchitectural differences between the
two types of myotubes. One factor may be the
age of the myoblast at the time of myotube
formation. For example, slight alterations in
the phenotypic expression of fusion-related factors ke., membrane-associated variations) may
occur with age. Thus, the satellite cell may be
released from, or subjected to, different fusion
limitations from those the embryonic myoblast
is. Second, there are different environmental
factors brought to bear on secondary myogenesis as compared to primary myogenesis. The
necrotic environment alone may be responsible
for alterations in myotube structure. One of
the most marked differences between secondary myogenesis and fetal development is the
presence of an intact basal lamina in regenerating muscle, which is used as a scaffold
within which regenerating myotubes are formed
(Vracko and Benditt, 1972; Burden et al., 1979).
It may be that mechanical forces, exerted by
the basal lamina and the necrotizing fiber a t
a critical stage of myogenesis, foster the production of abnormal myotubes. In the present
study, it has been demonstrated that branching is already present at a time when the immature, crescent-shaped myotubes are wedged
between the muscle coagulum and the old basal
lamina. An alternate explanation for the
branching phenomenon may be related to the
basal lamina’s segregation and partial isola-
tion of the satellite cells, which were originally
associated with a single myofiber. Although it
has not been documented whether additional
satellite cells or other types of cells migrate
into the necrotic fibers and participate, along
with the “resident” satellite cells, in de novo
myotube formation, it is likely that the “resident” satellite cells play the major role as the
source of myogenic cells during regeneration.
Could it be that some sort of “self‘ recognition
could allow these “resident” satellite cells to
fuse in a less regulated manner with all of the
myogenic cells in the old basal lamina?
The branching pattern observed in the regenerating myotubes in whole-muscle transplants, the branching myotubes observed in
the regenerating fibers formed spontaneously
in dystrophic muscle (Ontell and Feng, 19811,
and the suggestion that branching may also
be observed between myotubes formed subsequent t~ injections of myotoxic substance (Jirmanova and Thesleff, 1972;Schmalbruch, 1976)
appear to challenge the paradigm that secondary myogenesis produces myotubes that are
indistinguishable from the myotubes produced
by primary myogenesis.
ACKNOWLEDGMENTS
Supported by NIH grant NS 13688 and a
grant from the Muscular Dystrophy Association of America.
The competent technical assistance of Ms. F.
Shagas, Ms. G. Diluiso, and Ms. J. Lieberman
is gratefully acknowledged.
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