Download cross-sectional myofibre and myofibril growth in immobilized

J. Cell Sci. 29, 297-312 (1978)
Printed in Great Britain © Company of Biologists Limited igyS
297
CROSS-SECTIONAL MYOFIBRE AND
MYOFIBRIL GROWTH IN IMMOBILIZED
DEVELOPING SKELETAL MUSCLE
CHARLES R. SHEAR
Department of Anatomy, University of Maryland, School of Medicine,
655 W. Baltimore Street, Baltimore, Maryland 21201, U.S.A.
SUMMARY
The effect of immobilization on skeletal muscle fibre and myofibril growth in the developing
chick latissimus dorsi posterior muscle was investigated. Immobilization, immediately upon
hatching, results in a 25 % decrease in mean myofibre cross-sectional area within 24 h. Longer
periods of immobilization further retard normal myofibre growth until by day 27 after hatching
the mean myofibre cross-sectional area was 70 % less than normal. The variation in
individual myofibre cross-sectional areas (measured as the coefficient of variation, CV =
standard deviation/mean x 100) was greater in immobilized muscles. The larger immobilized
myofibres showed atypical myofibril and sarcotubular complex organization at the ultrastructural level of observation whereas the smaller immobilized myofibres appeared normal,
except for their size. In 1- to 27-day-old chicks, freeing the immobilized muscles for 24 h
resulted in myofibril reorganization and sarcotubular proliferation but little myofibre growth.
Freeing the immobilized muscles for 72 h resulted in a nearly complete recovery, in chicks up
to 18 days post-hatching, of both cross-sectional myofibre area and ultrastrucrural morphology.
Myofibres immobilized for longer than 18 days and subsequently freed for 72 h develop a
normal myofibrillar and sarcotubular morphology but remain 23 to 29 % smaller in crosssectional area than myofibres from control muscles.
Normal myofibril growth and organization are severely retarded in the immobilized
myofibres. The evidence presented here suggests that sarcomere shortening causes growing
myofibrils to rupture. The ruptured myofibrils are filled with proliferating elements of the
sarcotubular complex which, in turn, completely subdivide (split) the myofibril.
INTRODUCTION
It is generally agreed that the number of striated muscle fibres in vertebrate muscles
remains constant once differentiation is complete. The amount or duration of postnatal muscle differentiation varies among the different vertebrate classes and is
apparently dependent upon the relative maturity of the animal at birth (Morpurgo,
1897; Goldspink, 1962; Chiakulas & Pauly, 1965). However, differentiated muscles
are able to increase their mass during normal postnatal growth and in response to
altered functional demands (MacCallum, 1898; Adams, Denny-Brown & Pearson,
1962; Shear & Goldspink, 1971; Goldspink, 1972; Shear, 1975). In the developing
and fully differentiated muscle, the increase in total muscle mass is due to longitudinal and cross-sectional muscle fibre growth and to a lesser extent, to the growth of
non-contractile connective tissue elements within the muscle (Goldspink, 1968;
Bridge & Allbrook, 1970; Shear & Goldspink, 1971; Williams & Goldspink, 1971;
Shear, 1975).
298
C. R. Shear
During normal post-hatching development of the chick, the fast-phasic muscle
fibres of the latissimus dorsi posterior (PLD) and the slow-tonic muscle fibres
of the latissimus dorsi anterior (ALD) increase 100-fold in cross-sectional area.
This increase in fibre size is the result of an increase in myofibrillar (contractile)
and sarcoplasmic (non-contractile) components of the growing fibre. In young
fibres of both types, newly synthesized contractile proteins are assembled into
myofilaments (Morkin, 1970) and incorporated into the existing myofilament
lattice (Larson, Jenkison & Hudgson, 1970; Larson, Fulthorpe & Hudgson, 1973).
The growing myofibrils of the PLD muscle fibres incorporate new contractile
filaments, double their size, and then split or divide longitudinally into 2
daughter myofibrils (Shear & Goldspink, 1971). The fibres have a 'Fibrillenstruktur'
myofibril morphology with myofibrils of uniform size, each surrounded by a well
developed sarcotubular complex (sarcoplasmic reticulum and transverse tubular
system) and each I band traversed by a straight and narrow Z disk (Kriiger, 1950;
Kriiger & Giinther, 1955; Hess, 1961).
In a recent study of fish myotomal muscle fibres, peripheral myofibrils were found
to incorporate new contractile protein at a higher rate and to split more frequently
than central myofibrils (Patterson & Goldspink, 1976). In the present investigation,
the location of the dividing myofibrils within the growing chick muscle fibre has been
investigated. The role of the sarcotubular complex in subdividing the contractile
filament mass into discrete myofibrils has also been examined.
The homogenous fast-phasic, Fibrillenstruktur fibre composition of the newly
hatched, developing, and adult chick PLD muscle serves as an ideal model with which
to study myofibril growth and division. In the present study immobilization was
employed to limit whole muscle shortening and hence sarcomere shortening in newly
hatched chick myofibres because; (1) the load placed on the myofibres by their skeletal
attachments remains unchanged; (2) the myofibres may be fixed at a predetermined
resting length; and (3) the trauma caused by denervation and tenotomy is absent. In
order to determine whether immobilization had irreversibly limited growth in the
developing myofibres, immobilized muscles were freed and light-microscopic and
ultrastructural analysis made of whole fibre cross-sectional area and myofibril
morphology. A partial account of these results was reported before the Third International Congress on Muscle Diseases (Shear, 1975).
MATERIALS AND METHODS
Latissimus dorsi posterior muscles (PLD) from female White Leghorn chickens, ranging in
age from 1 to 27 days old (1, 3, 6, 9, 12, 18, 21 and 27 days after hatching), were used in this
study. The PLD muscle originates from the tips of the neural spines of the ninth to the eleventh
thoracic vertebrae and inserts by means of a short, flat tendon on the medial surface of the
proximal portion of the humerus. Whole muscle shortening was restricted byfixingthe position
of one wing at a predetermined body or rest length, i.e., that length at which maximum isometric tension is obtained. The unpublished observations of Shear & Goldspink show maximum
isometric tension for the PLD muscle, in chicks of all ages, to occur with the wings folded next
to the body. The position of the fixed wing and hence the length of the immobilized muscle is
most important. Studies on whole muscle weight and immobilization indicate the importance
of fixing the wing at rest or body length. Muscles immobilized at greater than rest length show
Myofibre and myofibril growth
299
an initial increase in total dry weight (Thomsen & Luco, 1944; Ferguson, Vaughan & Ward,
1
9S7)i while those tenotomized or fixed at a length shorter than rest length lose weight more
rapidly (Summers & Hines, 1951; Price, Howes & Blumberg, 1964). Apparently the prolonged
maintenance of long sarcomere length results in the serial addition of new sarcomeres to the
ends of the existing myofibrils, while shortened sarcomere lengths cause the loss of existing
sarcomeres (Williams & Goldspink, 1971). In order to avoid sarcomere degeneration which
might confuse an ultrastructural interpretation of myofibril growth, care was taken to immobilize PLD muscles at their proper resting lengths. Immediately upon hatching, the position of
one wing was fixed next to the body with a surgical adhesive cast. Thereafter the casts were
examined at daily intervals and adjustments made to allow for increased limb development.
Newly hatched chicks became adapted to single wing immobilization within the first 90 min
after hatching. Chicks with immobilized wings were maintained in battery brooders, separate
from non-immobilized birds, and were given food and water ad libitum. Experimental chickens
were assigned to 3 groups, according to the experimental designs described below. The chicks
were sacrificed by massive intraperitoneal or intravenous injections of tubocurarine chloride,
thus blocking myoneural transmission which might otherwise cause the muscle fibres to shorten.
Immobilized. One wing of the newly hatched chicks was immobilized and the animals
sacrificed 1, 3, 6, 9, 12, 18, 21 and 27 days post-hatching. Both the immobilized and nonimmobilized contralateral muscles were removed and prepared for electron microscopy (n — 3,
for each age) or used to determine whole muscle dry weight (n = 3, for each age).
Immobilized-freed. Animals with one wing immobilized for 3, 6, 9, 15, 18 and 24 days posthatching and then freed of the casts for 72 h were sacrificed and the PLD muscles from both
sides prepared for electron microscopy (n = 3, for each age) or dry weight measurement
(n = 3, for each age). Another group of chicks was immobilized for 2, 20 and 26 days posthatching, freed 24 h and the muscles prepared for electron microscopy (n = 3, for each age)
or dry weight measurement (n = 3, for each age). As in the preceding group, the chicks
resumed use of the immobilized wing within 2-5 h following removal of the cast.
Non-immobilised. A population of non-immobilized animals was maintained as described
above and sacrificed 1, 3, 6, 9, 12, 18, 21, and 27 days after hatching. Both the right and left
PLD muscles of each chick were removed and prepared for electron microscopy (n = 3, for
each age) or dry weight measurement (n = 3, for each age). The fibre cross-sectional area
values and whole muscle weights proved too variable, between different birds of the same age,
to permit comparisons between non-immobilized and immobilized animals. In this investigation the muscle contralateral to the immobilized side served as a control for each bird.
For electron microscopy, the muscles were exposed, dissected free of connective tissue, and
cold (4 °C) phosphate-buffered 2-5% glutaraldehyde fixative (pH 7-2) containing 0-54%
glucose was pipetted on to the muscles for 5 to 15 min. The muscles were then tied to an
applicator stick, removed, and placed in a larger volume of the same fixative at 4 °C for an
additional 2 h. After fixation the mid-belly region of the muscle was minced, washed briefly in
cold (4 °C) o-i M phosphate buffer and postfixed for 2 h in 1 % OsO, buffered to pH 7-2 with
o-i M phosphate buffer and containing 0-54 % glucose. The muscles were dehydrated in ethanol
and embedded in Araldite, CIBA CY 212 (Glauert, 1965). Ultrathin sections showing a silvergrey interference colour were collected on naked grids and stained with uranyl acetate and lead
citrate solutions (Reynolds, 1963). The sections were examined with a Philips 300 or Zeiss
EM 9S-2 electron microscope and the magnifications calibrated using a carbon replica of
2200 lines/mm diffraction grating. Semithin sections for light microscopy were cut at 1 fim
and stained with toluidine blue.
The cross-sectional area of 100fibresfrom each muscle were measured directly from electron
micrographs using an electronic rolling disk planimeter (Los Angeles Scientific Supply Co.,
Inc.). Only those fibres showing discrete, punctate thick and thin filaments were considered to
represent true transverse sections and to be suitable for fibre cross-sectional area determination.
All of the fibres within each randomly selected fasciculus were measured and the mean myofibre
cross-sectional area ± the standard error was calculated for each muscle. The coefficient of
variation (standard deviation/mean x 100) was calculated for each immobilized, immobilizedfreed and non-immobilized contralateral muscle. In this way the distribution and variability of
myofibre area, between chicks of different ages, could be compared (Snedecor & Cochran, 1967).
In order to determine whether a small change in mean myofibre size results in a measurable
C. R. Shear
3oo
change in whole muscle weight, the dry weights were calculated for each of the groups described
above. The muscles were removed, weighed, dried for 2 h at 55 °C, dessicated for 24 h and
reweighed.
RESULTS
Immobilization of the PLD muscle resulted in a reduction of mean myofibre
cross-sectional growth in chicks of all age groups. Representative histograms showing
the distribution of myofibre area in immobilized and non-immobilized muscles are
shown in Figs. 2 A, B; 3 A, C. In Fig. 1, the mean myofibre cross-sectional area for each
100
90
o
o
o
80
I
70
c
o
60
o
o
o
50
40
o
o
30
1
3
6
9
12
18 21
Age, days post-hatching
27
Fig. 1. Graph showing the relationship between the mean cross-sectional area of PLD
myofibres immobilized continuously ( • ) , immobilized PLD myofibres freed for 24 h
(O), and immobilized PLD myofibres freed for 72 h (A) at different post-hatching
ages. Each point represents the mean value of 100 myofibres measured from one
muscle and is expressed as a percentage of the mean cross-sectional value of 100
control myofibres.
immobilized muscle is shown as a per cent of the mean cross-sectional area of the nonimmobilized contralateral muscle. Owing to the small transverse size of many fibres
(less than 5 /tm diameter), and the difficulty in distinguishing muscle cells from
myoblastic, satellite and fibroblast cells, all of the cross-sectional area measurements
were taken directly from electron micrographs.
Even after only 1 day of immobilization, the mean myofibre area ranged from 19 to
22% less than corresponding contralateral muscles (Figs. 1, 2A). After 3 days the
Myofibre and myofibril growth
301
mean myofibre area was 27 to 3 1 % less for immobilized muscles (Figs. 1, 2B).
Between 3 and 12 days of post-hatching immobilization, there was a slow but steady
reduction in mean myofibre area until by day 12 the mean myofibre area was 34-40%
less in immobilized muscles (Figs. 1, 3 A). By day 21, the mean myofibre area declined
to 60-66% of the corresponding contralateral muscles (Figs. 1, 3 c), and by day 27
the immobilized fibres were 70-72% less than their controls.
30-
A
>o
*****
§20-
DC
^^^^
—L_
o
lative
a>
_r
i
15
i
1
V"
25
36
60
84
35
12
Myofibre area, /im2
myofibre cross-sectional area
Fig. 2. Representative histograms demonstrating myofibre
distributions in immobilized (shaded, narrow lines) and control (unshaded, heavy lines)
PLD muscles. A, 1 day post-hatching; B, 3 days post-hatching.
Considerably greater variation in myofibre cross-sectional area was seen in immobilized muscles between 1 and 12 days post-hatching than in corresponding control
myofibres (Figs. 2 A, B; 3 A). The coefficient of variation (CV) was 17-25% greater in
immobilized myofibres through day 12 post-hatching and 6-15 % greater in myofibres
immobilized for 18, 21 and 27 days than in myofibres of corresponding control
muscles. After 1 day of immobilization the muscles weigh 8-14% less than contralateral muscles and by 27 days the immobilized muscles weigh 27-35 % l e s s t n a n their
controls. In many of the immobilized muscles a pronounced proliferation of the perimysial connective tissue was observed, accounting in part for the less dramatic loss in
total dry weight.
Freeing immobilized muscles for 72 h resulted in a surprising recovery of myofibre
area in 6- to 18-day-old chicks (Figs, i, 3B). In these immobilized-freed, 72 b
muscles, the mean myofibre areas were only 1-6% less than in corresponding control
muscles. Freeing the muscles for 72 h reduced the variability in cross-sectional
myofibre area in the 6- to 18-day-old chicks, i.e., as compared to continuous immobilization. The immobilized-freed, 72-h myofibres show CVs 2-5% greater than their
controls. In 21- to 27-day-old chicks, the myofibres of muscles immobilized and then
freed for 72 h were 23-29 % smaller in cross-sectional area than their corresponding
control myofibres and their CVs ranged from 6 to 19% greater than CVs for the
contralateral muscles (Figs. 1, 3D). The immobilized-freed, 72-h muscles weigh
2-3% less than corresponding control muscles through day 18 post-hatching and
10-13% l e s s m the 21- and 27-day-old animals.
20
CEL 29
C. R. Shear
302
30-
30
i
I
I i
I
I
r
210
45
135
225
315
Myofibre area, ;im 2
Fig. 3. Representative histograms demonstrating myofibre cross-sectional area
distributions in immobilized (shaded, narrow lines) and control (unshaded, heavy
lines) PLD muscles: A, 12 days post-hatching; c, 21 days post-hatching. Representative
histograms demonstrating myofibre cross-sectional area distributions in immobilizedfreed 72-h (shaded, narrow lines) and control (unshaded, heavy lines) P L D muscles:
B, 12 days post-hatching; D, 21 days post-hatching.
Fig. 4. Transverse section of an untreated PLD myofibre from a 21-day-old chick.
The myofibrils are discrete and well separated from one another throughout the
fibre cross-section and at all levels of the sarcomere. x 6400.
Fig. 5. Transverse section of a small P L D myofibre immobilized for 21 days posthatching. Although the myofibrillar and sarcotubular organization (arrows) appear
normal, many of the fibres are very much smaller in cross-sectional area than controls,
x 21800.
Fig. 6. Transverse section of a large PLD myofibre immobilized for 21 days posthatching. The myofibrils are not well separated and the sarcotubular complex is poorly
developed, x 21 800.
Fig. 7. Transverse section of a large P L D myofibre immobilized for 21 days posthatching. The M line is clearly visible indicating that groups of adjacent thick filaments remain.in longitudinal register, x 116000.
Myofibre and myofibril growth
3°3
304
C. R. Shear
Muscles immobilized for 2, 20 and 26 days post-hatching and freed for 24 h had
mean myofibre cross-sectional areas of 20-65 % less than their corresponding controls
(Fig. 1). As in the continually immobilized muscles, the CVs ranged between 8 and
28% greater than corresponding control values. Whole muscle dry weights ranged
from 5 to 30% less for immobilized-freed, 24-h muscles than control muscles, the
greatest differences occurring in the 21- and 27-day-old chicks (20-25% and 23-30%
respectively).
Electron-microscopic examination shows 2 types of myofibrillar morphology in the
developing, immobilized myofibres (Figs. 4-6). At all ages the small myofibres show
distinct myofibrils and an extensive sarcotubular system (Fig. 5). The myofibrillar
morphology of the smaller immobilized myofibres is similar to that of the nonimmobilized myofibres (Fig. 4) with triads, i.e., one transverse tubule and 2 adjacent
terminal cisternae of the sarcoplasmic reticulum, located at the centre of the I band
between the Z disk and the A band. The larger fibres generally have ill-defined myofibrils and a poorly developed sarcotubular complex (Fig. 6) similar to the Felderstruktur morphology of the typical avian tonic muscle fibres (Shear & Goldspink,
1971). As in the tonic myofibres the triads are generally located at the Z disk in the
larger immobilized fibres but are difficult to distinguish since both the transverse
tubules and terminal cisternae are less well developed than in the small immobilized
or control myofibres. Unlike the tonic myofibres, the M lines are clearly visible even
in transverse sections (Fig. 7).
An extensive sarcotubular system is seen in large and small myofibres freed for 24 h
and the myofibrils are more clearly defined (Figs. 8, 9) than in immobilized myofibres
(Figs. 5, 6). A large number of the immobilized-freed, 24-h myofibrils show signs of
dividing or splitting apart (Figs. 8-10). Many of the larger immobilized-freed myofibres regain a nearly normal myofibrillar and sarcotubular organization, 24 h after
cast removal (Figs. 9, 10). In these fibres triads are generally found at the level of the
A-I band and occasionally at the Z disk. The location of the triads is more variable,
even along the length of a single myofibril, than in control myofibrils. The sarcotubular complex of the smaller immobilized-freed, 24-h myofibres is more extensive
than in the larger immobilized-freed, 24-h myofibres or in the control myofibres
(Fig. 8). Since both large and small immobilized-freed myofibres show notable proliferation of the sarcotubular complex within the first 24 h following removal of the
cast and only a small increase in mean myofibre cross-sectional area, transverse tubular
Fig. 8. Transverse section of a small PLD myofibre immobilized for 20 days and freed
for 24 h. Note the numerous myofibrillar separations and the extensive sarcotubular
complex, x 25 600.
Fig. 9. Transverse section of a large PLD myofibre immobilized for 20 days and freed
for 24 h. The sarcoplasmic reticulum is less extensive than in the small immobilized,
24-h fibres and triads can be identified at the A-I bands (arrows), x 28500.
Fig. 10. Longitudinal section of a dividing myofibril (arrows) within a large myofibre of a 27-day-old chick immobilized for 26 days and freed for 24 h. Myofilaments
associated with adjacent daughter myofibrils are occasionally encountered along
dividing myofibrils (triangles), x 24700.
Myofibre and myofibril growth
• • • • • ' "
•
•'Sh*-
;
•••
_
\
:
C. R. Shear
Myofibre and myofibril growth
307
and sarcoplasmic reticular development appear to precede renewed myofibrillar
growth. The myofibrillar morphology and organization of the sarcotubular complex
in fibres of muscles immobilized and freed for 72 h is similar to the controls.
The large number of centrally disrupted myofibrils in immobilized-freed, 24-h
fibres provides an ideal model for the study of longitudinal myofibril division. Ultrathin serial transverse sections were made through a single myofibril, from an immobilized-freed 24-h fibre of a 27-day-old chick, undergoing longitudinal division (Figs. 1114). The myofibril split is first seen as a disruption at the centre of the Z disk (Fig. 11).
Transverse serial sections through the I bands, A bands and Z disks of the 2 sarcomeres immediately preceding the disruption showed normal filament lattice arrangements with no disruptions. Serial sections through 4 successive sarcomeres adjacent
to the Z disk shown in Fig. 11, show the disruption to be continuous (Figs. 12-14).
The myofibrillar disruptions within the first 2 sarcomeres contain no visible elements
of the sarcotubular complex (Figs. 11, 12). Elements of the sarcotubular complex,
probably sarcoplasmic reticulum, have entered the disruptions of the last 2 sarcomeres and appear to be advancing through the myofibril along the previously existing
disruption (Figs. 13, 14). Transverse serial sections through a large number of disrupted myofibrils show similar patterns of myofilament disruption and sarcotubular
complex development. Within the small fibres, nearly 50% of the myofibrils freed
for 24 h show central disruptions or splits. In the large immobilized-freed, 24-h
fibres, 22% of the myofibrils show signs of splitting. The frequency of myofibril
splitting in the immobilized-freed, 72-h myofibres and in control myofibres is only
10-12% at all ages studied. In both the experimental and control myofibres, myofibrils in various stages of splitting occur at the periphery and centre of the cells with
equal frequency.
DISCUSSION
The results presented here indicate that sarcomeres of the fast-twitch myofibres of
chick PLD muscle must shorten in order to grow and develop normally. Even in the
very young chicks, myofibres showed retarded growth after only 24 h immobilization.
The decrease in immobilized muscle dry weight reflects a loss of insoluble contractile
protein. The greater percentage decrease in cross-sectional myofibre area than in comparable muscle dry weight suggests increased development of the vascular and/or connective tissue elements in the immobilized muscles. Although the vascular system
appeared normal in the immobilized muscles, the interfascicular connective tissue did
appear more extensive than in the controls. Since freeing the muscles for 72 h resulted
in nearly normal mean myofibre cross-sectional areas and whole muscle dry weights
Fig. 11. Transverse section of a PLD myofibril immobilized for 26 days and freed for
24 h. Note the irregular Z-disk lattice and the central disruption (arrow), x 90600.
Transverse serial sections through the A bands of 3 adjacent sarcomeres along the
same myofibril are shown in the following 3 figures (Figs. 12-14).
Fig. 12. The central A band disruption (arrow) is free of sarcotubular complex in the
sarcomere immediately adjacent to the Z disk preceding, x 90 600.
Myofibre and myofibril growth
309
(up to 18 days post-hatching), there appears to be some regulatory interaction between the myofibre and connective tissue elements comprising the PLD muscle. In
a recent study of immobilized adult cat skeletal muscles, whole muscle weight was
found to decrease by a greater percentage than did fibre size (Cooper, 1972). However,
in that study the percentage change in fibre size was determined from transverse
diameter measurements rather than cross-sectional area measurements. When crosssectional areas are calculated, using the reported mean myofibre diameter measurements of Cooper (1972), the changes in muscle weight and fibre size are similar to
those reported in the present investigation.
In light of the uniform physiological, ultrastructural, and histochemical make-up,
characteristic of newly hatched PLD myofibres (Gutmann, Hanslikova & HoleCkovd,
1969; Shear & Goldspink, 1971; Jirmanova & Zelena, 1973; Gordon, Perry, Spurway
& Vrbova, 1975), it is surprising that in the present study the myofibres are not equally
affected by immobilization. The cross-innervation studies of Hnik, Jirmanova,
Vyklicky & Zelend (1967) and Jirmanova & Zelend (1973), on young and newly hatched
chick PLD muscles, show that a change in fibre type can only be brought about in
newly hatched muscle fibres. These authors suggest that the ability to transform young
myofibres is largely dependent upon the maturity of a particular myofibre at the time
the foreign motor neuron establishes contact. Although the majority of the newly
hatched cross-innervated fibres are transformed physiologically, many myofibres
show only partial ultrastructural transformation (70% per muscle), i.e., as regards
the sarcotubular and myofibrillar organization. Their findings, together with those
reported in the present investigation indicate that the myofibres comprising the young
chick PLD muscle may not be equally differentiated on the day of hatching even though
they are 100% innervated (Hirano, 1967; Gutmann et al. 1969; Grim, 1971) and
show uniform histochemical properties (Gordon et al. 1975). In mammals the ultrastructural maturation of fibre type is a postnatal process related in some way to the
postnatal differentiation of distinct types of motor end-plates (Walker, Schrodt &
Bingham, 1968; Schiaffino & Margreth, 1969; Padykula & Gauthier, 1970). It has been
demonstrated (Chinoy & Hess, 1974) that avian motor end-plates undergo similar
post-hatching differentiation. Thus, depending on the state of such motor end-plate
differentiation, immobilization would affect the myofibres differently. Although the
immobilized myofibres do not show a measurable increase in cross-sectional area after
the first 24 h following removal of the cast, they do show considerable alteration at the
ultrastructural level as is evidenced by the proliferation of the sarcotubular complex
and subdivision of myofibrillar material.
The results of this investigation indicate that the sarcomeres of myofibrils in developing fast-twitch fibres must shorten in order to grow, split and divide longitudin-
Fig. 13. As in Figs. 11 and 12, showing the A band of the next adjacent sarcomere.
Note the presence of sarcotubular complex within the disruption (arrow), x 90600.
Fig. 14. As in Figs. 11—13, showing the A band of the next adjacent sarcomere. Note
that elements of the sarcotubular complex (arrows) are more abundant than in the
previous sarcomere and appear to proceed along a single axis, x 90600.
310
C.R. Shear
ally. Myofibrillar division is important in that it subdivides the myofibrillar mass,
allowing a well organized sarcoplasmic reticulum and transverse tubular system to
develop. The myofibres of the PLD muscle require a well developed and highly
ordered sarcotubular complex in order to release and sequester Ca2+ rapidly (Peachey
& Porter, 1959; Peachey, 1968, 1970). Evidently the development of the sarcotubular
complex is also dependent upon sarcomere shortening since immobilized fibres
showed a decreased development of that complex in fibres of the PLD normally
capable of extensive sarcotubular development (Shear & Goldspink, 1971). The
retarded development of the sarcotubular complex in the large immobilized fibres
is apparently reversible to the extent that muscles freed of the casts and allowed to
shorten already showed extensive proliferation of that complex by 24 h, and by 72 h
all of the fibres possessed a well developed highly organized sarcoplasmic reticulum
and transverse tubular system and a normal myofibril morphology.
Workers in several different laboratories have reported a lateral expansion of the
myofilament lattice (A and I band) during isotonic contraction (Carlsen & Knappeis,
1963; Brandt, Lopez, Reuben & Grundfest, 1967; Pringle, 1968; Goldspink, 1971;
April, 1975 a, b; and April & Wong, 1976). In vertebrate skeletal myofibres, Goldspink (1971, 1972) has found that during isotonic contraction the thick myosin
filament lattice expands more than the Z disk lattice, thus displacing the peripheral
thin filaments and causing them to pull in an oblique direction on their associated and
common Z disk. According to Goldspink the resulting force acting on one Z disk by
the thin filaments of 2 adjacent one-half sarcomeres might be sufficient to split the
myofibrillar mass longitudinally. Such a force might be expected to increase as the
total myofibrillar cross-sectional area increases, as indeed occurs when new contractile
proteins are synthesized and the filaments assembled and incorporated into the
growing sarcomere. The occurrence of a sarcotubular complex within the ruptured
myofibrillar mass suggests that proliferation of these membranous elements further
help to completely subdivide the growing myofibril.
In a recent study of myofibril growth in fish muscle, Patterson & Goldspink (1976)
found that the most peripheral myofibrils incorporated new contractile filaments and
divided while the more central myofibrils showed little growth or division. Also, these
authors suggest that new myofilaments are added to the daughter myofibrils while
they are splitting. In our observations of chick myofibril development, central and
peripheral myofibrils were seen to grow and divide with nearly equal frequency. As
in the fish myofibres, presumably newly assembled thick and thin filaments were
observed in various stages of incorporation at the periphery of the splitting myofibrils.
A possible explanation for the difference between fish and chick myofibrillar growth
is that in young, newly-hatched chick myofibres the ribosomes, upon which the contractile filaments are synthesized, are more uniformly distributed than in the young
adult fish myofibres. This, in turn would permit myofilament assembly and incorporation in a uniform fashion throughout the cross-section of the chick myofibre. We do
find abundant ribosomes and polysomes throughout the cross-section of the growing
myofibre. Similar observations, dealing with ribosomal distribution in developing
human and avian myofibres have been reported by Larson et al. (1970, 1973). In a
Myofibre and myofibril growth
311
recently completed study on regenerating mammalian skeletal muscle fibres (Shear &
O'Steen, in preparation), splitting myofibrils were observed with equal frequency
throughout the cell. Based upon our current understanding of the mechanisms of
protein synthesis in eukaryotic cells, the distribution of ribosomal material would
seem an important variable affecting myofibrillar growth. Further studies are required
to relate changes in both ribosomal and myonuclear distribution to myofilament
assembly in the growing myofibril.
The author wishes to acknowledge the skilful technical assistance rendered by Mrs Carol G.
Shear. This research was supported by Grant No. AM 20131, awarded by the National Institute of Arthritis, Metabolism and Digestive Diseases, PHS/DHEW.
REFERENCES
ADAMS, R. D., DENNY-BROWN, D. & PEARSON, C. M. (1962). Diseases of Muscle, A Study in
Pathology, pp. 135-264. New York: Hoeber Medical; Harper and Row.
APRIL, E. W. (1975a). The myofilament lattice: Studies on isolated fibres. IV. Lattice equilibria in striated muscle. J. Mechanochem. Cell Motil. 3, 111-121.
APRIL, E. W. (19756). Liquid-crystalline characteristics of the thick filament lattice of striated
muscle. Nature, Lond. 357, 139-141.
APRIL, E. W. & WONG, D. (1976). Non-isovolumic behavior of the unit cell of skinned striated
muscle fibres. J. molec. Biol. 101, 107—114.
BRIDGE, D. T . & ALLBROOK, D. (1970). Growth of striated muscle in the Australian marsupial
(Steonix brachyurus). J. Anat. 106, 285-295.
BRANDT, P., LOPEZ, E., REUBEN, J. & GRUNDFEST, H. (1967). T h e relationship between myo-
filament packing density and sarcomere length in frog striated muscle. J. Cell Biol. 33,255-263.
CARLSEN, F. & KNAPPEIS, G. G. (1963). Further investigations of the ultrastructure of the
Z disk in skeletal muscle. Acta physiol. scand. 59, 213-215.
CHIAKULAS, J. J. & PAULY, J. E. (1965). A study of postnatal growth of skeletal muscle in the
rat. Anat. Rec. 152, 55-62.
CHINOY, N. J. & HESS, A. (1974). An electron microscopic, cytochemical and biochemical
study on developing chicken latissimus dorsi muscles with special reference to motor endplate structure. Illrd int. Cong. Muscle Diseases, 334, pp. 8-9. Amsterdam: Excerpta Medica.
COOPER, R. R. (1972). Alterations during immobilization and regeneration of skeletal muscle in
cats. J. Bonejt Surg. 48-A, 1239-1271.
FERGUSON, A. B., VAUGHAN, L. &WARD, L. (1957). A study of disuse atrophy of skeletal muscle
in the rabbit. J. Bonejt Surg. 39-A, 583-586.
GLAUERT, A. M. (1965). The fixation and embedding of biological specimens. In Techniques for
Electron Microscopy (ed. D. H. Kay), pp. 166-212. Philadelphia: Davis.
GOLDSPINK, G. (1962). Studies of postembryonic growth and development of skeletal muscle.
I: Evidence of 2 phases in which striated muscle fibres are able to exist. Proc. R. Ir. Acad.
62B, 135-150.
GOLDSPINK, G. (1968). Sarcomere length during postnatal growth of mammalian muscle fibres.
J. Cell Sci. 3, 539-548.
GOLDSPINK, G. (1971). Changes in striated muscle fibres during contraction and growth with
particular reference to myofibril splitting. J. Cell Sci. 9, 123-138.
GOLDSPINK, G. (1972). Postembryonic growth and differentiation of striated muscle. In Tlie
Structure and Function of Muscle, vol. 1, 2nd edn (ed. G. H. Bourne), pp. 179-236. New
York: Academic Press.
GORDON, T., PERRY, R., SPURWAY, N. C. & VRBOVA, G. (1975). Development of slow and fast
muscles in chick embryos. Proc. Physiol. Soc. (Cambridge), 25 P.
GRIM, M. (1971). Development of the primordia of the latissimus dorsi muscle of chicken.
Folia morph., Praha 19, 252-258.
GUTMANN, E., HANZL^KOVA, V. & HOLECKOVA, E. (1969). Development of fast and slow muscles
of the chicken in vivo and their latent period in tissue culture. Expl Cell Res. 56, 33-38.
312
C.R. Shear
HESS, A. (1961). Structural differences of fast and slow extrafusal muscle fibres and their nerve
endings in chickens. J. Physiol., Land. 157, 221-231.
HIRANO, H. (1967). Ultrastructural study on the morphogenesis of the neuromuscular junction
in the skeletal muscle of the chick. Z. Zellforsch. mikrosk. Anat. 79, 198—208.
HNfK, P., JIRMANOVA, I., VYKLICKY, L. & ZELENA, J. (1967). Fast and slow muscles of the chick
after nerve cross-union. J. Physiol., Lond. 193, 309-325.
JIRMANOVA, I. & ZELENA, J. (1973). Ultrastructural transformation of fast chicken muscle
fibres induced by nerve cross-union. Z. Zellforsch. mikrosk. Anat. 146, 103-121.
KROGER, P. (1950). Untersuchungen am Vogelflugel. Zool. Anz. 145, 445-460.
KROCER, P. & GONTHER, P. G. (1955), Fasern m i t ' Fibrillenstruktur' und Fasern mit 'Felderstruktur' in der guergestreiften skelet Muskulatur der Saiiger und des Menschen. Z. ges.
Anat. 118, 313-323LARSON, P. F., FULTHORPE, J. J. & HUDGSON, P. (1973). The alignment of polysomes along
myosin filaments in developing myofibrils. J. Anat. 116, 327-334.
LARSON, P. F., JENKISON, M. & HUDGSON, P. (1970). The morphological development of chick
embryo skeletal muscle grown in tissue culture as studied by electron microscopy. J. neurol.
Set. 10, 385-405.
MACCULLUM, J. B. (1898). On the histogenesis of the striated muscle fibre and the growth of the
human sartorius muscle. Bull. Johns Hopkins Hosp. 9, 208-215.
MORKIN, E. (1970). Postnatal muscle fibre assembly: localization of newly synthesized myofibrillar proteins. Science, N.Y. 167, 1499-1501.
MORPUHGO, B. (1897). Ueber-Activitats Hypertrophie der willkurlichen Muskeln. Virchows
Arch. path. Anat. Physiol. 150, 522-554.
PADYKULA, H. A. & GAUTHIER, G. F. (1970). The ultrastructure of neuromuscular junctions
of mammalian red, white and intermediate skeletal muscle fibres. J. Cell Biol. 46, 27-41.
PATTERSON, S. & GOLDSPINK, G. (1976). Mechanism of myofibril growth and proliferation in
fish muscle. J. Cell Sci. 22, 607-616.
PEACHEY, L. D. (1968). Muscle. A. Rev. Physiol. 30, 401-429.
PEACHEY, L. D. (1970). Form of the sarcoplasmic reticulum and T system of striated muscle.
In Physiology and Biochemistry of Muscle as a Food, vol. 2 (ed. E. J. Briskey, R. G. Cassens &
B. B. Marsh), pp. 213-310. Madison: University of Wisconsin Press.
PEACHEY, L. D. & PORTER, K. R. (1959). Intercellular impulse conduction in muscle cells.
Science, N.Y. 129, 721-722.
PRICE, H. M., HOWES, E. L. & BLUMBERG, J. M. (1964). Ultrastructural alterations in skeletal
muscle fibers injured by cold. II. Cells of the sarcolemmal tube: Observations on 'discontinuous' regeneration and myofibril formation. Lab. Invest. 13, 1279-1302.
PRINGLE, J. W. S. (1968). Mechano-chemical transformation in striated muscle. In Aspects of
CellMotility, XXIInd Symp. Soc. exp. Biol., pp. 67-86. Cambridge & London: Cambridge
University Press.
REYNOLDS, E. S. (1963). The use of lead citrate at high pH as an electron-opaque stain in
electron microscopy, J. Cell Biol. 17, 208-213.
SCHIAFFINO, S. & MARGRETH, A. (1969). Coordinated development of the sarcoplasmic reticulum and T system during postnatal differentiation of rat skeletal muscle. J. Cell Biol. 41,
85S-875SHEAR, C. R. (1975). Myofibril proliferation in developing skeletal muscle. In Recent Advances
in Myology, 360 (ed. W. G. Bradley), pp. 364-373. Amsterdam: Excerpta Medica.
SHEAR, C. R. & GOLDSPINK, G. (1971). Structural and physiological changes associated with
the growth of avian fast and slow muscle. J. Morph. 135, 351-372.
SNEDECOR, G. W. & COCHRAN, W. G. (1967). Sampling from a normally distributed population.
In Statistical Methods, 6th edn, pp. 32-65. Ames: Iowa State University Press.
SUMMERS, T . B. & HINES, H. M. (195 I ) . Effect of immobilization in various positions upon the
weight and strength of skeletal muscle. Arch. Phys. Med. 32, 142-145.
THOMSEN, P. & Luco, J. V. (1944). Changes in weight and neuromuscular transmission in
muscles of immobilized joints. J. Neurophysiol. 7, 245-251.
WALKER, S. M., SCHRODT, G. R. & BINGHAM, M. (1968). Electron microscopic study of the
sarcoplasmic reticulum at the Z-line level in skeletal muscle fibres of foetal and newborn rats.
J. Cell Biol. 39, 469-475WILLIAMS, P. E. & GOLDSPINK, G. (1971). Longitudinal growth of striated muscle fibres. J.
Cell Sci. 9, 751-767.
[Received 30 May 1977)