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Modification of the phenotypic expression of murine dystrophyA morphological study.

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THE ANATOMICAL RECORD 214:17-24 (1986)
Modification of the Phenotypic Expression of
Murine Dystrophy: A Morphological Study
Department of Anatomy and Cell Biology, University of Pittsburgh School of Medicine,
Pittsburgh, PA 15261
The extensor digitorum longus muscles of 4-6-week-old normal
mice (129 R e 4 and dystrophic mice (129 ReJ dy/dy) were orthotopically transplanted.
Grafted muscles were examined 1,3,7,14,20,50,and 100 days post-transplantation.
The myofibers of both types of grafts underwent a similar time course of necrosis
and regeneration. Other than during the initial necrotic response, no evidence of
necrotic myofibers was found in either type of grafted muscle. At 100 days posttransplantation, the grafted normal and dystrophic muscles were essentially similar,
except that the dystrophic graft was of smaller size. Based on a comparison of the
number of myofibers found a t the 100-day grafts’ widest girths [631 & 59 SEM, for
normal grafts (Bourke and Ontell, 1984);631 f 74 SEM, for dystrophic grafts], it is
suggested that the regenerative capability of traumatized 4-6-week-old dystrophic
muscle is similar to that of traumatized normal muscle. At 100 days post-transplantation, the grafted dystrophic muscle appeared “healthier” than untraumatized
muscle from age-matched dystrophic mice, having less variation in myofiber diameter, better fascicular organization, and less connective tissue. The transplantation
system demonstrates the possibility of modifying the expression of genetic programming of myopathic disorders using environmental manipulation.
In 1955, Michaelson et al. identified a myopathic
mouse strain (dy)which is caused by a n autosomal recessive mutation. The disorder is characterized by the progressive deterioration of striated muscle, and myofibers
are replaced by connective and/or fatty tissue. Despite
extensive morphological and physiological studies, the
etiology of the disease remains controversial. Originally
described at a primary myopathy (Michaelson et al.,
1955; Banker and Denny-Brown, 1959; Cosmos et al.,
1973), deficits are also found in the nervous system
(Biscoe et al., 1974; Bradley and Jaros, 1973; Bray et al.,
1983; Harris and Ribchester, 1979; McComas and Mrozek, 1967)of this dystrophic mouse.
The progressive deterioration of the dystrophic muscle
fibers occurs despite evidence that untraumatized dystrophic muscle possesses regenerative capability. Fascicles of small diameter regenerating myotubes are found
in these muscles (Michaelson et al., 1955). In explanation of this apparent paradox, it has been variously
hypothesized: that the regenerative capacity of dystrophic muscle is unable to keep pace with the degenerative changes; that the myotubes formed by the regenerative response are genetically programmed, as are
fetal dystrophic myotubes, to undergo progressive deterioration; and that the regenerating myotubes are unable to survive because of their dystrophic environment.
While it has been established that severely traumatized normal striated muscle displays extensive regenerative capability (Carlson, 1973), there are few studies
evaluating the ability of dystrophic muscle to regenerate subsequent to secondary trauma sufficient to induce
necrosis of all of its myofibers (Cosmos, 1974; Hironaka
and Miyata, 1973; Laird and Timmer, 1966; Neerunjun
and Dubowitz, 1975a,b; Neerunjun et al., 1976; Salafsky,
1971). These studies, using a variety of animal models
of dystrophy and various types of trauma, have provided
conflicting results. In the present study, normal and
dystrophic (dy) murine muscles of 4-6-week-old mice
have been subjected to the severe trauma of orthotopic
whole muscle transplantation. We report that the regenerative capacity of these grafted dystrophic muscles is
similar to that of grafted normal muscles. Moreover,
using morphological criteria, the regenerated myofibers
found in grafted dystrophic muscle appear to be refractory to degenerative changes even when the graft is
maintained for long periods in a dystrophic environment.
Orthotopic transplantation of whole extensor digitorum longus muscles (Carlson and Gutmann, 1974) was
performed on 30 normal and 30 dystrophic mice (129
ReJ and 129 ReJ dy/dy, respectively) obtained from colonies maintained a t the Jackson Laboratory. Muscles
were transplanted when the mice were 4-6 weeks old.
The muscles were removed by severing their tendons,
vascular supply, and nerve supply; care was taken not
to leave any muscle on the attached tendon stumps.
After soaking the muscles for -20 minutes in 0.75%
Marcaine, a known myotoxic agent (Benoit and Belt,
1970; Libelius et al., 1970) which prevents the survival
Received February 12,1985;accepted July 11,1985.
of the muscles’ peripheral fibers (Carlson, 19761, the
muscles were replaced in their respective beds in the
correct proximalidistal orientation and resutured to the
attached tendon stumps. No attempt was made to anastomose blood vessels or to align nerves. Grafted muscles
were exposed 1, 3, 7, 14, 20, 50, and 100 days posttransplantation and fixed in situ in 2.0% glutaraldehyde
in 0.125 M cacodylate buffer (pH 7.3) for 30 minutes.
Grafts were removed from the hindlimb and placed in
fresh fixative for 1.5 hours. The grafts were postfixed in
buffered 2.0% osmium tetroxide, dehydrated in ethanol,
and embedded in Epon 812. Untraumatized muscles obtained from age-matched normal and dystrophic mice
were similarly fixed and embedded. Epon blocks containing entire untraumatized muscles or grafted muscles were placed on a sliding microtorne (equipped with
a steel knife) and oriented for sectioning in a plane
perpendicular to the long axis of the muscle fibers. (The
extensor digitorum longus is a pennate muscle and a
plane perpendicular to the long axis of the myofibers is
not perpendicular to the long axis of the muscle.) The
muscles were serially sectioned, from origin to insertion,
into sets consisting of ten 15-pm-thick sections and one
8-pm-thick section. Fifteen-micron-thick sections were
cleared in Epon between two layers of polystyrene film
and cured in an oven a t 60°C (Davidowitz et al., 1976).
The 8-pm-thick sections were mounted on glass slides
and studied using a Leitz microscope equipped with
phase-contrast optics and a n eyepiece reticule which was
calibrated with a stage micrometer. This permitted us
to evaluate changes observed along the entire lengths of
the muscles. The eyepiece reticule was used to determine which of the 8-pm-thick sections contained the
maximal cross-sectional area of the muscle (hereafter
termed the muscle’s widest girth). The 15-pm-thick section adjacent to the muscle’s widest girth was cut out of
the polystyrene sandwich and attached to a preformed
Epon block. The reembedded sections were cut into
semithin (0.5 pm) or ultrathin (70-90 nm) sections with
a n ultramicrotome. Semithin sections were stained with
toluidine blue. Ultrathin sections were collected on slot
grids, stained with uranyl acetate and lead citrate (Reynolds, 1963), and observed using a Philips 300 electron
Morphometric analyses were carried out on grafts 100
days post-transplantation and on age-matched untraumatized muscles (N = 6 for grafted normal muscle, N =
6 for grafted dystrophic muscle, N = 4 for untraumatized muscle). The maximal cross-sectional area of the
grafted normal and dystrophic muscles was determined
from light micrographs ( x 145)of a toluidine blue section
taken from the region of the muscle’s widest girth using
a Bioquant-IBM morphometric analysis system. The
maximal cross-sectional area of grafted muscles was
compared with that of unoperated normal and dystrophic muscles (Ontell et al., 1984). The number of
myofiber profiles found at the widest girth of the dystrophic grafted muscles was determined using the electron microscope, and this was compared with the number
of myofiber profiles found a t the widest girths of normal
grafts (Bourke and Ontell, 1984) and with the number
of myofiber profiles found at the widest girth of untraumatized normal and dystrophic muscle (Ontell et al.,
1984). The frequency of finding central nuclei in the
grafted myofibers was determined by the examination
of 300 myofiber profiles at the widest girth of each
specimen using the electron microscope. Electron micrographs of the entire cross section of untraumatized and
grafted normal and dystrophic muscles, taken a t the
muscles’ widest girths, were printed at a magnification
of ~ 1 , 1 0 0(The
electron microscope and enlarger were
calibrated at each use.) Approximately 300 myofiber
profiles for each specimen were randomly chosen, and
the appropriate micrographs placed on a digitizing board
of a Bioquant-IBM morphometric analysis system. The
perimeter of each fiber was traced and the cross-sectional area was computed by the Bioquant system. A
calculations program determined the diameter of each
myofiber (assuming each myofiber to be circular in cross
section). The mean fiber diameters and histograms of
myofiber diameter distributions were calculated by the
Bioquant system for each muscle and for all muscles in
a given group.
In contrast to the untraumatized extensor digitorum
longus muscle of normal 4-6-week-old mice, which consisted of fascicles of myofibers with similar morphological characteristics, three types of myofibers, distinguished by their selective response to the disease
process, could be identified in the 4-6-week-old dystrophic muscle. There was a large population of relatively “healthy” myofibers, a few necrotic myofibers,
and some regenerating myofibers (Fig. 1). The “healthy”
myofibers were polygonally shaped (in transverse sections), contained subsarcolemmal myonuclei and regular myofibril arrays, and they were morphologically
indistinguishable from the myofibers found in untraumatized normal muscle. The necrotic myofibers were
swollen and appeared rounded in transverse section (Fig.
lb). Regenerating fibers (Fig. lc) resembled myofibers
found in embryonic muscle, being of small diameter and
containing centrally located nuclei, variable amounts of
myofilaments, and lipid and glycogen deposits. [A detailed ultrastructural description of necrotic and regenerating myofibers in untraumatized murine dystrophic
muscle is found in Ontell (1981) and Ontell and Feng
The time course of the degenerative and early regenerative changes in normal whole muscle transplants of the
mouse extensor digitorum longus muscle has been described (Ontell et al., 1982) and will only be briefly
mentioned in order to compare regeneration in the dystrophic grafts with that observed in the normal grafts.
Subsequent to transplantation, the myofibers of the normal and dystrophic grafts underwent a similar time
course of necrosis, phagocytosis, and replacement by
newly formed regenerating myotubes (Figs. 2-7). Within
24 hours after transplantation, all of the myofibers of
the grafts were swollen and exhibited necrotic changes
all along their lengths (Fig. 2). Macrophages were found
among and within the necrotic myofibers. Small-diameter regenerating myofibers began to appear in the periphery of the grafted muscles 3 days posttransplantation, and at 7 days post-transplantation, regenerated myofibers were found across the width of the
grafts (Fig. 3). By 2 weeks post-transplantation, myofibers organized into discrete fascicles were found
throughout the grafted muscles (Fig. 4). Myoneural junctions were first observed in grafted muscles at 14 days
Fig. 1. Light micrograph of untraumatized, 4-week-old dystrophic
extensor digitorum longus muscle. Region of necrotic fiber indicated
by crossed arrow in a is enlarged in b. Region of regenerating focus
indicated by arrow in a is enlarged in c. At this stage, relatively
“healthy” fibers predominate (b, asterisk). The necrotic fibers are
rounded and swollen (Fig. lh, crossed arrow). Regenerating myotubes
; X500.
(c, arrow) contain central nuclei. Toluidine blue. a, ~ 7 5b,c,
Fig. 2. Grafted dystrophic extensor digitorum longus muscle, 1 day
post-transplantation. Area surrounding arrow in a is enlarged in b.
All myofibers are necrotic, appearing swollen and rounded in cross
section. Macrophages (crossed arrow) are found between the myofibers.
Toluidine blue. a, ~ 5 0b,; x500.
Fig. 3. Grafted dystrophic extensor digitorum longus muscle, 1 week
post-transplantation. Area surrounding arrow in a is enlarged in b. A t
this stage small-diameter regenerating myotubes with centrally located myonuclei are found throughout the graft. Toluidine blue. a,
~ 7 5b,; ~ 5 0 0 .
Fig. 4. Grafted dystrophic extensor digitorum longus muscle, 2 weeks
post-transplantation. Area surrounding arrow in a is enlarged in h. At
this stage, the graft is significantly larger than at 1 week post-transplantation (Fig. 3a), and the myofihers have increased in diameter
(compare b with Fig. 3b). No necrotic fibers are found. Crossed arrow
in Figure 4a indicates adjacent, untraumatized, dystrophic muscle.
Toluidine blue. a, ~ 7 5b,; ~ 5 0 0 .
post-transplantation (Fig. 9). Subsequently, the girths of
the grafts increased, and the regenerating myotubes
increased in diameter (Figs. 5-71 by the accumulation of
additional myofibrils. Aside from the initial massive
necrotic reaction, a t no stage in the regenerative response were necrotic myofibers observed in either type
of graft.
At 100 days post-transplantation, the normal and dystrophic grafts were virtually indistinguishable from each
other, except for the larger size of the normal graft (Figs.
6, 7, Table 1).While some myonuclei had undergone
peripheral migration, 64% of the myonuclei of the normal grafts and 53%of the myonuclei of dystrophic grafts
remained centrally or eccentrically located. At low mag-
TABLE 1. Morphometric analyses of untraumatized and grafted normal and dystrophic murine EUL’
Maximal cross-sectionalarea of EDL (rnm’)
No. of myofibers
Mean myofiber diameter (pm)
0.75 5 0 . O l 2
922 k 28’
29.7 f 0.3
0.47 f 0.06
631 k 5g3
21.0 0.1
0.39 f 0.032
547 f 452
21.8 i- 0.3
631 f 74
15.8 -t 0.1
‘EDL, extensor digorum longus. Untraumatized muscles were from 17-week-old mice; grafted muscles were 100 post-transplantation
(transplants performed on 4-6-week-old mice). N = 6 for normal and dystrophic grafted muscles. All analysis performed at muscle’s widest
irth. Values expressed as mean k SEM.
‘Values taken from Ontell et al. (1984).
3Value taken from Bourke and Ontell (1984).
nification, the normal 100-day transplant (Fig. 7) appeared to be a smaller version of the 17-week-oldnormal
muscle (not shown); however, even at low magnification
considerable differences were observed between 100-day
dystrophic transplants and age-matched untraumatized
dystrophic muscle (Figs. 6, 8). The grafted dystrophic
muscle appeared “healthier” than did the untraumatized dystrophic muscle from age-matched mice, having
less variation in myofiber diameter, better fascicular
organization, and substantially less connective tissue.
The maximal cross-sectional area of the normal and
dystrophic grafts was less than that of their respective
untraumatized muscles, with the maximal cross-sectional area of the dystrophic graft being 59% of that of
the normal graft. However, the numbers of myofibers
present at the widest girths of both types of grafts were
similar (Table 1).While normal grafts contained only
68% of the myofibers found in age-matched untrauma-
Fig. 5. Grafted dystrophic extensor digitorum longus muscle. 50 days
post-transplantation. Area surrounding arrow in Figure 5a is enlarged
in b. At this stage, the polygonally shaped myofibers are organized
into discrete fascicles. No necrotic fibers are found in the grafted
muscles. Toluidine blue. a, x 75; b, x 500.
Fig. 6. Grafted dystrophic extensor digitorum longus muscle, 100
days post-transplantation. Area surrounding the arrow in Figure 6a is
enlarged in b. This graft is similar to 100-day grafted normal muscles
(Fig. 71, except that the maximal cross-sectional area of the dystrophic
graft is smaller. The dystrophic graft has the same number of myofibers as the normal graft; however, the myofibers of‘ the dystrophic
graft are smaller in diameter (compare h with Fig. 7b). Approximately
53% of myonuclei are centrally or eccentrically located (crossed arrow).
The grafted, dystrophic muscle appears “healthier” than age-matchcd
untraumatized dystrophic muscle (compare with Fig. 8), having less
connective tissue and a smaller range of myofiber diameters. No necrotic fibers are found in the grafted muscles. Toluidine blue. a, ~ 7 5 ;
b, x500.
Fig. 7. Grafted normal extensor digitorum longus muscle, 100 days
post-transplantation. Area surrounding arrow in a is enlarged in h.
The graft is similar to 100-day grafted dystrophic muscle. Approximately 64% of myonuclei are centrally located (crossed arrows). No
necrotic fibers are found in the grafted muscles. Toluidine blue. a, x 75;
b, x500.
Fig. 8 . Untraumatized, 1’7-week-old dystrophic extensor digitorum
longus muscle. Area surronding arrow in a is enlarged in b. At this
stage, muscle deterioration is advanced (compare with Fig. la), and
there is marked variation in myofiber diameter distributions. Regionally, the fascicular arrangement of the myofibers is disturbed. Large
amounts of connective tissue separate the myofibers (Fig. 8b, crossed
arrow). Based on morphological criteria, the grafted dystrophic muscle
F i g . 6) appears “healthier” than untraumatized dystrophic muscle.
Toluidine blue. a, ~ 7 5b,; x500.
tized normal muscle, the number of myofiber profiles
present in the dystrophic graft was similar to that found
in age-matched untraumatized dystrophic muscle (Table
1). The mean diameter of the myofibers of the dystrophic
graft was smaller than that of the normal graft (Table
1).Histograms of the diameter distributions of the myofiber profiles found at the widest girths of untraumatized and grafted normal and dystrophic muscles are
found in Figure 10. Myofibers in untraumatized dystrophic muscles had a wider diameter distribution than
myofibers in untraumatized normal muscle (Fig. 10a),
due to the presence in dystrophic muscle of small-diameter “regenerating” myofibers and a few large “hypertrophied” myofibers. When the normal extensor
digitorum longus muscle was transplanted, there was a
shift to the left in the diameter distribution of the myofibers, the myofibers of the grafted muscle failing to
achieve the normal myofiber diameter distribution (Fig.
lob). Transplantation of dystrophic muscle resulted in a
population of myofibers with much less diameter variation then that found in untraumatized dystrophic muscle (Fig. 1Oc). The extent of variation in the diameter
distribution of the myofibers in the two types of grafts
was similar; however, there was a shift to the left in the
diameter distribution of the myofibers found in the dystrophic grafts (Fig. 1Od).
The source of the regenerating myofibers in traumatized mammalian muscle is believed to be the myosatellite cell (cf. Carlson, 19731, a small mononucleated cell
found wedged between the sarcolemma and the basal
lamina of the original myofiber (Mauro, 1961). It is hypothesized that these cells are capable of surviving the
ischemia which characterizes the early stage of grafting
and that the trauma of transplantation exerts a mitogenic effect on the myosatellite cell population. Subsequent to multiple cell divisions, these cells fuse, forming
multinucleated myotubes (Carlson, 1973).Therefore, two
factors must be present to insure a n effective early regenerative response. First, there must be an adequate
population of mitotically competent myosatellite cells.
Second, the myosatellite cells of the grafted muscles
must be fusion competent. Recently, we have shown that
the size of the myosatellite cell population of untraumatized normal and dystrophic murine muscles are similar
and that the myosatellite cells of dystrophic muscle have
a higher labelling index than myosatellite cells of normal age-matched muscles (Ontell et al., 1984).This elevated labelling index may actually enhance the intial
Fig. 9. Grafted dystrophic extensor digitorurn longus muscle, 2 weeks post-transplantation. Myoneural
junctions begin to appear in t h e dystrophic grafts a t 2 weeks post-transplantation. The postsynaptic
membrane (arrows) is slightly thickened and the axon terminal is filled with synaptic vesicles (crossed
arrow). Uranyl acetate and lead citrate. ~ 9 , 5 5 3 .
[email protected]
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Fig. 10. Comparison of the diameter distributions of myofibers found a t the widest girth of untraumatized and grafted normal and dystrophic extensor digitorurn l o n y s muscles.
regenerative response of the dystrophic grafts. It also extensor digitorum longus muscle are innervated by
has been determined that the myosatellite cells of dys- approximately one-half of the motoneurons innervating
trophic muscle are fusion competent (Ontell et al., 1984). untraumatized normal muscles (Klueber et al., 1984).
Therefore, the ability of the grafted dystrophic muscles Innervation of normal grafts appears to be principally
to undergo a n effective, early regenerative response has the result of sprouting of the cut nerve which had innernot been entirely unexpected.
vated the muscle prior to transplantation (Yip and OnAn unexpected observation, however, is the persist- tell, unpublished data). If a similar reduction of
ence of a viable and apparently “healthy” graft 100 days motoneurons to grafted dystrophic muscle occurs and if
after the transplantation of the dystrophic muscle into dystrophic motoneurons are not equally affected by the
a dystrophic hindlimb. Previous studies in which minced dystrophic process, perhaps only the “healthier” motodystrophic muscles, rather than whole dystrophic mus- neurons are capable of innervating the dystrophic grafts.
cles, have been orthotopically transplanted have re- This could explain why the regenerated dystrophic mussulted in grafts which failed to survive or which cle, in the present study, fails to show evidence of dysexhibited dystrophic characteristics (Cosmos et al., 1979; trophic changes. Clearly the question of why regenerated
Neerunjan and Dubowitz, 1975a,b). Moreover, Hironaka dystrophic myofibers show modification of programmed,
and Miyata (1975) have reported degenerative changes dystrophic deterioration requires further investigation.
in whole grafted murine dystrophic muscle. However, in
Since the 4-week-old unoperated dystrophic muscle
their experiments, the muscles were grafted when the contains - 57% of the number of myofibers found in the
mice were older and free access of the nerve to the 4-week-old unoperated normal muscle I4-week-old norregenerating muscle belly was inhibited by wrapping mal muscle contains 1,235 f 29 (SEM) myofibers and 4the grafts in Saran Wrap. Their protocol also differed week-old dystrophic muscle contains 708 i 8 (SEMI
from that used in the present study in that Marcaine myofibers (Ontell et al., 198411 and since the normal and
was not used on their grafts prior to muscle implanta- dystrophic grafted muscles contain the same number of
tion. Therefore, some of the original peripheral myofi- myofibers, it is clear that relative to myofiber number
bers may have survived their transplantation procedure. the dystrophic muscle exhibits a regenerative response
These surviving fibers may be responsible for the popu- to trauma at least equivalent to that of normal muscle.
lation of degenerating myofibers found in their grafts. How then can we account for differences in the diameter
One might question why the regenerated myofibers of distributions of the myofibers found in normal and dysgrafted dystrophic muscles examined in this study ap- trophic grafts? One factor which should be considered is
pear to be refractory to dystrophic changes. If the exclu- that the total body weight of the dystrophic mouse is
sive cause of the degenerative response of untraumatized significantly less than that of the normal mouse (Rowe
dystrophic muscle is a genetically programmed primary and Goldspink, 1969a,b), and, therefore, a smaller musdefect in the myofiber (cf. Cosmos et al., 1973),one would cle could be anticipted in the smaller animal. Given that
expect that the fate of regenerated dystrophic myofibers both types of grafts have a similar number of myofibers,
[being formed by myosatellite cells, which are believed it might be expected that the myofibers in the dysto be the progeny of fetal myoblasts that have not fused trophic grafts would be of smaller diameter. Another
during fetal development (Mauro, 196111 would be simi- factor which could account for the differences in myofilar to that of the myofibers formed during fetal develop- ber diameter would be the fact that the dystrophic musment. Clearly, this is not the case. It is possible, however, cle has been placed into a limb which, because of
that environmental factors may have modified the phe- deterioration of surrounding muscles, is no longer a notypic expression of what might still be a primary tively used in locomotion. Myofiber diameter differences
myopathy. These factors may include differences in the between normal and dystrophic grafts may reflect the
time course of the establishment of neuromuscular re- reduced workload to which the dystrophic graft has been
lationships. Fetal myotubes of the normal murine exten- sujected. Coan and Tomanek (1981) have demonstrated
sor digitorum longus muscle become innervated within that an increased workload results in an increase in
a day or 2 of their formation (Ontell and Kozeka, 1984), myofiber diameter of the grafted rat soleus muscle. The
while regenerating myotubes in grafted murine muscles two hypotheses which we have proposed to explain difdo not become innervated until about 1 week after sec- ferences in diameter distributions of the myofibers found
ondary myogenesis has been completed. Since it has in normal and dystrophic grafts suggest a possible modbeen hypothesized that dystrophic myotubes exhibit de- ification of dystrophic myofiber size due t o environmenlayed maturation (Vrbova, 1983), the delayed innerva- tal influences. However, we cannot a t the present time
tion of the regenerating myotubes may be beneficial for exclude the possibility that differences in myofiber sizes
the long-term survival of the dystrophic muscle.
are genetically controlled. By transplanting a dysAn alternate hypothesis which may account for the trophic muscle into a normal host, i t may be possible t o
failure of the grafted dystrophic muscle to exhibit degen- determine whether differences in myofiber diameters
erative changes is that differences exist in the motoneu- are the result of genetic or environmental factors.
rons supplying transplanted and untraumatized
Using morphological criteria, the 100-day-oldgrafts of
dystrophic muscle. This factor may be of extreme impor- dystrophic muscles appear “healthier” than agetance if one adheres to the “sick motoneuron” theory of matched, untraumatized dystrophic muscles. However.,
dystrophy (McComas et al., 1971). This theory suggests we have not evaluated the condition of these grafts at
that the dystrophic myopathy is secondary to neuro- any subsequent time period. It may be that some degenpathological changes involving the motoneuron. While erative changes ultimately occur in dystrophic grafts.
no information is currently available regarding the This does not, however, lessen the importance of the
number of motoneurons innervating grafted dystrophic present observation. Myotubes first appear in the fetal
muscle, it is known that the grafts of the normal mouse extensor digitorum longus muscle a t the 12th day in
utero (Ontell and Kozeka, 1984). Within 22 days after
myotube formation (by 14 days postnatal), dystrophic
myofibers show evidence of histopathological changes
(Banker, 1967). In contrast, regenerated dystrophic myofibers in grafted muscles remain free of histopathological changes for a n extended time period.
The present study provides the first evidence that is
possible to experimentally alter the genetically programmed morphological deterioration of murine dystrophic muscle, in a dystrophic animal, by inducing the
muscle to undergo massive de nouo niyotube formation
subsequent to extensive trauma. We are currently evaluating the functional capacity of these muscles. While a
previous study has demonstrated reduced isometric
twitch and tetanic tension for orthotopically grafted
whole dystrophic muscles, as compared to age-matched
unoperated dystrophic muscles (Hironaka and Miyata,
1975),differences i n experimental procedures (see above)
have resulted in their grafts exhibiting degenerative
changes. The improved morphological appearance of the
regenerated dystrophic muscle in the present study will
be of greater significance if it can be shown to be accompanied by an improved physiological capacity.
This study was supported by the Muscular Dystrophy
Association, National Institutes of Health grant N S
13688, and the Health Research and Services Foundation. The excellent technical assistance of Ms. Gloria
Diluiso and Mrs. Barbara Hahn is gratefully
Banker, B.Q. (1967) A phase and electron microscopic study of dystrophic muscle I. The pathological chmges in the 2 week-old Bar
Harbor 129 dystrophic niouse. J . Neuropathol. Exp. Neurol..
Ranker, R.Q.. and U. D e u n - B r o ~ . (19591
A study of denervated m u s ~
cle in normal and dystrophic mice. J. Neuropathol. Exp. Ncurol.,
Benoit, P.W., and W.D. Belt 11970)Destruction and regeneration of
skeletal muscle after treatment with a local anesthetic, bupivacaine (Marcaine).J. Anat., 107:547-556.
Biscoe, T.J., K.W.T. Caddy, D.J. Pallot, U.M.M. Pehrson, and C.A.
Stirling (19741 The neurological lesion i n t h e dystrophic mouse.
Brain Res., 76r534-536.
Rourke, D.L., and M. Ontell (19841 Branched myolibers i n long-term
whole muscle transplants: A quantitative study. Anat. Rec.,
Bradley, W.G., and E. Jaros (1973) Axoplasmic flow i n axonal neuropathies. 11. Axoplasmic flow in mice with motor neuron disease and
muscular dystrophy. Brain, 96.247-258.
Bray, G.M., S. David, T. Carlstedt, and A. .4guayo (1983) Effects of
crush injury on the abnormalities in the spinal roots and peripheral nerves of dystrophic mice. Muscle Nerve, 6497-503.
Carlson, B.M. (1973) The regeneration of skeletal muscle-A review.
Am J. Anat., 137:119-150.
Carlson, B.M. (1976)A quantitative study of muscle fiber survival and
regeneration in normal, predenervated, and Marcaine-treated free
muscle grafts in the rat. Exp. Neurol., 52421-432.
Carlson, B.M., and E. Gutmann (1974) Transplantation and cross
transplantation of free musclc grafts in the rat. Experientia.
30: 1292-1294.
Coan, M.R., and R.J. Tomanek (1981) The growth of regenerating
soleus muscle transplants after abalation of the gastrocnemius
muscle. Exp. Neurol., 71:278-294.
Cosmos, E. (1974) Muscle transplants: Role in the etiology of hereditary muscular dystrophy. In: Exploratory Concepts in Muscular
Dystrophy, A.T. Milhorat, ed. Excerpta Medica, Amsterdam, pp.
Cosmos, E., J. Butler, J. Mazliah, and E.P. Allard (1979) Viability of
muscles grafted between normal animals and animals with hereditary muscular dystrophy. In: Muscle Regeneration. A. Mauro, ed.
Raven Press, New York, pp. 523-539.
Cosmos, E., J. Butler, and A. Milhorat (1973) Hereditary muscular
dystrophy: A possible myogenic defect i n the differentiation of
muscle. In: Basic Research in Myology. B.A. Kakulas, ed. Excerpta
Medica, Amsterdam, pp. 632-640.
Davidowitz, J., B.R. Pachter, and G.M. Breinin (1976) “Clearing” steel
knife epon sections in a polystyrene film sandwich. Stain Technol.,
52: 139-140.
Harris, J.B., and R.R. Ribchester (1979) Muscular dystrophy in the
mouse: Neuromuscular transmission and the concept of functional
denervation. Ann. N.Y. Acad. Sci., 317:152-170.
Hironaka, T., and Y. Miyata (1973) Muscle transplantation in t h e
aetiological elucidation of murine muscular dystrophy. Nature,
Hironaka, T., and Y. Miyata (1975) Transplantation of skeletal muscle
in normal and dystrophic mice. Exp. Neurol., 47:l-15.
Klueber, K., J.W. Yip, and M. Ontell (1984) Size and location of the
motoneuron pool supplying normal and orthotopically transplanted muscles. Brain Res., 305r192-195.
Laird, J., and R. Timmer (1966) Transplantation of skeletal muscle
into a host with muscular dystrophy. Tex. Rep. Biol. Med., 24.169179.
Libelius, R., B. Sonesson, B.A. Stamenovic, and S. Thesleff (1970)
Denervation-like changes in skeletal muscle after treatment with
a local anaesthetic (Marcaine). J. Anat., 206:297-309.
Mauro, A. (1961) Satellite cell of skeletal muscle fibers. J. Biophys.
Biochem. Cytol., 9~493-495.
McComas, A.J., and K . Mrozek (1967) Denervated muscle fibers in
hereditary mouse dystrophy. J. Neurol. Neurosurg. Psychiatr.,
McComas, A.J., R.E.P. Sica, and M.J. Campbell (1971) “Sick” motorneurones-A unifying concept of muscle disease. Lancet, 1:321325.
Michaelson, A.M., E.S. Russel, and P.J. Harman (1955) Dystrophia
muscularis: A hereditary primary myopathy in t h e house mouse.
Proc. Natl. Acad. Sci. USA, 41:1079-1084.
Neerunjun, J.S., and V. Dubowitz (1975aj Muscle transplantation between normal and dystrophic mice. 1. Histological studies. Neuropathol. Appl. Neurobiol., lt111-124.
Neerunjun, J.S., and V. Dubowitz (1975b) Muscle transplantation between normal and dystrophic mice. 2. Histochemical studies. Neuropathol. Appl. Neurobiol., 1:125-140.
Neerunjun, J.S., D.A. Jones, and V. Dubowitz (1976) Functional properties of muscles transplanted between normal and dystrophic mice.
Exp. Neurol., 52556-564.
Ontell, M (1981) Muscle fiber necrosis in murine dystrophy. Muscle
Nerve, 4.204-213,
Ontell, M., and K.C. Feng (1981) The three dimensional cytoarchitecture and pattern of motor innervation of branched striated myotubes. Anat. Rec., 200:ll-31.
Ontell, M., K.C. Feng, K. Klueber, R.F. Dunn, and F. Taylor (1984)
Myosatellite cells, growth, and regeneration in murine dystrophic
muscle: A quantitative study. Anat. Rec., 208.159-174.
Ontell, M, D. Hughes, and D.L. Bourke (1982) Secondary myogenesis
of normal muscle produces abnormal myotubes. Anat. Rec.,
Ontell, M., and K. Kozeka (1984) The organogenesis of murine striated
muscle: A cytoarchitectural study. Am. J. Anat., 171r133-148.
Reynolds, E.S. (1963) The use of lead citrate a t high pH a s a n electron
opaque stain in electron microscopy. J. Cell Biol., 17r.208-212.
Rowe, R.W.D., and G. Goldspink (1969a) Muscle fiber growth in five
different muscles in both sexes of mice. I. Normal mice. J. Anat.,
Rowe, R.W.D., and G. Goldspink (1969b) Muscle fiber growth in five
different muscles in both sexes of mice. 11. Dystrophic mice. J.
Anat., 104.531-538.
Salafsky, B. (1971) Functional studies of regenerated muscles from
normal and dystrophic mice. Nature, 229:270-272.
Vrbova, G. (1983) Duchenne dystrophy viewed a s a disturbance of
nerve-muscle interactions. Muscle Nerve. 6.671-675.
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expressions, murine, stud, phenotypic, modification, morphological, dystrophy
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