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Department of Animal Sciences, The Hebrew University of Jerusalem, Rehovot, Israel
2To whom correspondence should be addressed.
 |
ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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KEY WORDS: • broiler chickens • starvation • growth • skeletal muscle • satellite cells
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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).
These cells become committed to their particular developmental program
during late embryonic development (reviewed in Grounds and Yablonka-Reuveni 1993
). Located beneath the basal lamina,
satellite cells are capable of proliferating, differentiating and
joining existing fibers or fusing with each other to form new fibers
(reviewed in Campion 1984
, Merly et al. 1998
). At birth or hatch,
skeletal muscle consists of a high percentage of proliferating
satellite cells; however, these decrease rapidly toward the end of the
growth period (Cardiasis and Cooper 1975
). Thereafter,
satellite cells become mitotically quiescent and are activated only
after stress signals such as muscle injury (Schultz et al. 1978
).
Several growth factors, including fibroblast growth factor
(FGF),3
insulin-like growth factor (IGF),
transforming growth factor beta (TGF-ß) (reviewed by Florini et al. 1991
), platelet-derived growth factor (PDGF) and hepatocyte growth
factor (HGF) influence proliferation and differentiation of cultured
satellite cells (Allen et al. 1995
, Allen and Rankin 1990
, Florini et al. 1991
,
Gal-Levi et al. 1998
, Yablonka-Reuveni et al. 1990
). We recently reported that myogenesis of chicken
satellite cells is affected by growth hormone (Halevy et al. 1996
, Hodik et al. 1997
). Growth hormone
receptor (GH-R) was regulated in satellite cells during their
differentiation in vitro and in skeletal muscle growth in vivo
(Halevy et al. 1996
).
During the postnatal period in birds, there is an adaptation period
during which the animal moves from embryonic metabolic dependence on
lipids derived from the yolk to carbohydrate-rich exogenous feed.
Immediately posthatch, some of the metabolic requirements are met by
yolk which is transferred to both the circulation and the small
intestine (Noy and Sklan 1998
), and dramatic changes in
intestinal size and function occur with intake of feed.
Under commercial hatchery conditions, eggs hatch over a 36–48-h period
after which birds are removed from the hatchery, and additional
processing and transport to the farm result in some birds being exposed
to food for the first time > 50 h after hatching. This
period of starving has been shown to depress growth, with both short-
and long-term effects (Noy and Sklan 1997
).
The activity of the satellite cells in very young animals is transient
but crucial for muscle growth. For instance, in broiler chicks selected
for rapid growth, the first week of life is the most important for
muscle production (Moss et al. 1964
). We therefore
examined the effect of early nutrition on this activity at this
critical time. We examined the chick model because these animals are
grown mainly for their yield of muscle and do not generally have access
to feed until they are 24 to 50 h old.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Male broiler chicks (Ross) were transported to the laboratory within
1 h of hatching from a commercial hatchery (Kvuzat Yavne, Yavne,
Israel). Upon arrival, chicks were weighed and randomly divided into
two treatment groups (n = 50): one group had free
access to water and to a commercial diet formulated to meet or exceed
NRC requirements (National Research Council 1994
,
Halevy et al. 1994a
) for the entire experimental period.
A second group was maintained without access to feed or water for
48 h (from d 0 to d 2), after which access to food and water was
as in the control group. In additional experiments, food and water were
taken from the chicks from 48 to 96 h after hatch (between d 2 to
d 4) or from 96 to 144 h after hatch (between d 4 to d 6). All
chicks were maintained in temperature-controlled brooders. Chicks
were monitored daily for body and breast muscle weight for 5 d
posthatch and then at weekly intervals until 41 d of age. Each
experiment was repeated three times. All experimental procedures were
approved by the Animal Welfare Committee of the Faculty of Agriculture,
The Hebrew University of Jerusalem, and the animals were maintained in
accordance with the guidelines for the care and use of laboratory
animals.
Cell cultures.
Chicken skeletal muscle satellite cells were cultured from the pectoral
muscle of chicks as described by Halevy and Lerman (1993)
. An enriched
population of myogenic cells was recovered, and < 5% of the
cells were nonmyogenic (Halevy et al. 1994b
). Under
these conditions, the coefficient of variation of cell preparations was
5.3%. Cells were counted using a hemocytometer, plated on 0.1%
gelatin-coated plates at 5 x 104
cells/cm2 in Dulbecco’s Modified Eagle’s Medium
(DMEM) supplemented with 10% horse serum (HS), and grown for 1 d.
Cells were maintained at 37°C in a humidified atmosphere, 95% air
and 5% CO2. Cell cultures were prepared under
exactly the same conditions
Thymidine incorporation.
DNA synthesis was assessed by [3H]thymidine
incorporation as described previously (Halevy and Lerman 1993
). Cells were incubated for 17 h, in 24-well plates,
and [3H]-thymidine (New England Nuclear,
Boston, MA) was added (at 74 kBq/well) for an additional 2 h of
incubation. The cells were then detached with 2.5 g/L of
trypsin-EDTA and precipitated with 0.61 mol/L of trichloroacetic
acid. Radioactivity in the dissolved precipitates was counted using a
Tri-Carb 1600CA scintillation counter (Packard Instruments, Downers
Grove, IL). Equal plating efficiency was verified by measuring cell
numbers in parallel wells.
RNA preparation and hybridization.
Total RNA was isolated from cell cultures using the
guanidinium-thiocyanate-phenol technique as described by Chomczynski and Sacchi (1987)
. Northern blots were performed by loading total RNA
on a formaldehyde gel and blotting as described previously
(Halevy et al. 1996
) using nylon membranes for transfer
(Schleicher and Schuell, Dassel, Germany). The membrane was hybridized
with a chicken GH-R specific probe (kindly provided by J. Burnside, University of Delaware, Newark, DE), labeled with a
random-primed labeling kit (Boehringer, Mannheim, Germany), in
hybridization buffer solution (100 g/L of dextran sulfate, 10 g/L of
SDS, 1 mol/L of NaCl and 200 mg/L of denatured herring sperm DNA) at
60°C. Filters were then washed and autoradiographed. Densitometry
anlaysis was performed on bands using NIH software.
5-Bromo-2'-deoxyuridine (BrdU) labeling.
Two-day-old chicks were injected intraperitoneally with 1 mL/100 g body weight of aqueous solution containing BrdU, a marker for the S phase, and 5-fluoro-2'-deoxyuridine (10:1, v/v) (Zymed Laboratories, San Francisco, CA). At 2, 12 and 24 h postinjection, the birds were killed and samples of the breast muscle were removed for further analysis.
Histological assessments.
Muscle samples were removed and immediately fixed in fresh 4% paraformaldehyde, dehydrated, cleared and embedded in paraffin. Sections were cut at 5 µm and placed on glass slides. For all the assays, sections were deparaffinized in xylene and rehydrated in a graded alcohol series, then processed by either hematoxylin-eosin staining or immunohistochemistry. To quench endogenous peroxidase, all sections were incubated in 0.15% (v/v) of hydrogen peroxide for 30 min followed by incubation with 1% of HS in PBS to block nonspecific binding of antibodies. Sections were then incubated overnight at 4°C in a monoclonal antibody to proliferating cell nuclear antigen (PCNA), a marker for dividing cells (1:1000 dilution in blocking buffer; Zymed). After rinsing for 1 h in PBS, sections were incubated for 2 h at room temperature in horseradish peroxidase-conjugated anti-mouse IgG, diluted 1:200 in blocking buffer. A solution of 1 g/L of diaminobenzidine hydrochloride (Sigma Chemicals, St. Louis, MO) was mixed (1:1) with 0.03% of hydrogen peroxide. Sections were incubated with the peroxidase substrate for 1 h and then rinsed with PBS. Incorporated BrdU was stained by monoclonal antibody to BrdU followed by the use of peroxidase-ABC (Zymed BrdU staining kit) according to the manufacturer’s directions. After immunostaining, sections were counterstained with methyl green or hematoxylin, dehydrated and mounted in Histmount (Zymed). Negative control slides without primary antibody were examined in all cases. Digitized maps of the sections were analyzed using Image Pro Plus 3.0 software. Four to six random fields were analyzed in each section, and the stained nuclei were calculated as percentage of total nuclei for each of the fields. Central vs. peripheral nuclei in the breast muscle were quantified in a total of 700–900 cells per section which were counted over eight to ten fields at a magnification of 400 times.
Statistical analysis.
Data were analyzed by one-way ANOVA using the General Linear Models
procedures of SAS for effects of diet and age. Differences between
means were tested using t tests, and differences were
considered significant when P < 0.05 unless otherwise
stated (SAS Institute 1986
).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Body weight (BW) was significantly higher in the fed group than in the
starved group from the 1st d posthatch and remained so during the
entire experimental period (Fig. 1
). Lack of access to feed caused a decrease in BW, which increased only
after feed was ingested. The difference between the treatments
expressed as percentage BW was maximal on d 2 with a difference between
starved and fed chicks of > 50% of the starved birds BW (Fig. 1A
). Once feed was ingested, growth rate in the starved
birds was not different from that of fed birds. However, the BW
difference between the treatments remained significant until the end of
the experiment at 41 d (Fig. 1B
). In starved chicks,
the lower BW was accompanied by a lower relative breast muscle weight
from d 2 until the end of the experiment (Fig. 2
). Breast muscle of control birds reached nearly 15.5 g/100 g BW,
whereas in the starved group it comprised < 14 g/100 g.
|
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The effects of food deprivation from d 2 to 4 and from d 4 to 6
posthatch were progressively lower relative to that of immediate
starving at hatch (Table 1
). On d 8, BW in all starved groups was lower than that in the fed
chicks; however the group that starved from d 4 to 6 caught up in BW
and both absolute and relative breast muscle weights by d 41.
|
Lack of access to feed for the first 48 h influenced breast muscle
morphology as shown in Figure 3
. On d 3, thin young multinucleated myofibers were observed in the
muscle of starved chicks where 28.3 ± 1.4% of total nuclei were
central (Fig. 3A
). In contrast, muscle derived from the fed
chicks was populated with more mature fibers containing nuclei located
mainly in the peripheral zone of the fiber (Fig. 3B
); only
9.2 ± 0.7% nuclei were found to be located in the central zone
of the fiber. On d 4, although the myofibers in the muscle derived from
the starved chicks were more developed than those from d 3 (Fig. 3C
), their development continued to lag behind that of
muscle fibers in the fed group (Fig. 3D
). Young myofibers
with centered nuclei were still evident in the starved birds (Fig. 3C
), whereas mostly large fibers were observed in the fed
birds (Fig. 3D
). The percentage of central nuclei was 14.2
± 1.3 in the starved group whereas in the fed chicks it was only
5.6 ± 3.7 (P < 0.05).
|
The results above indicated that starvation during the very first days
of life retards both body and skeletal muscle growth. This raised the
possibility that the delay in muscle growth could be due to
insufficient proliferation of satellite cells. Breast muscle was
removed from chicks that were either fed or starved for the first
2 d posthatch, or on various days after hatch and satellite cells
were prepared and counted. In the fed group there was a trend for an
increase in the number of satellite cells per gram of muscle until d 3
(P < 0.1), a significant decline between d 4 and d
5 (P < 0.05); then numbers remained constant
(Fig. 4A
). In contrast, the number of satellite cells was higher in
the starved group than in the fed group on d 2, decreased on d 3 and
then increased again on d 4. Thereafter the number of satellite cells
decreased slowly, reaching the same low level as the fed group on d 8.
|
). However, after 2 d of starvation,
labeled thymidine incorporation decreased significantly to very low
levels. In contrast, thymidine incorporation increased by more than
100% in cells prepared from the fed chicks. On d 3, satellite cells of
the starved chicks became active, and thymidine incorporation increased
significantly until d 4 then declined from d 5. In contrast, labeled
thymidine incorporation in satellite cells from the fed group declined
after d 2 and remained at low levels with a small increase on d 4.
For the first 2 d of the experiment, GH-R mRNA levels in the
cells from the starved chicks were significantly lower than those in
the fed group (Fig. 5
upper). Densitometry analysis revealed that the difference between the
starved and fed chicks was ~100% on d 1 and 200% on d 2 (Fig. 5
lower). On d 3, 24 h after starved chicks had begun to receive
food, GH-R gene expression was induced in satellite cells and was
significantly higher than the expression in the fed group (Fig. 5)
.
|
The studies on satellite cell number and activity in culture revealed
that 2 d of starvation posthatch caused a dramatic decline in cell
proliferation, resulting in a more than fivefold difference in
thymidine incorporation by cells from starved vs. fed chicks. In
contrast, the actual number of cells prepared from the starved chicks
on d 2 was higher than that of controls. To examine this apparent
discrepancy, in vivo BrdU labeling was performed. Samples taken 12 h postinjection exhibited the highest incorporation of BrdU (data not
shown), and the position of the stained cells suggested that most were
satellite cells. The number of cells incorporating BrdU was markedly
higher in muscle from the fed than from starved chicks where only a few
BrdU-incorporating cells were observed (Fig. 6
). Morphological analysis revealed that the percentages of total cells
that were labeled were 26.7 ± 1.5 and 7.5 ± 0.5 in the fed
and starved chicks, respectively (P < 0.05). In
contrast, the total number of nuclei in the muscle of starved chicks
was significantly higher than that in the muscle derived from the fed
chicks (1277 ± 56 and 867 ± 35 cells per field,
respectively, P < 0.05).
|
, E
and Table 2
). However, on d 2 there was a marked reduction in PCNA-expressing
cells in muscle from starved chicks compared to muscle of fed chicks
where their numbers were substantially increased (Fig. 7B
, F
and Table 2
). The percentage of these cells was ~83% less in the
muscle of starved than that of fed chicks (Table 2)
. After feed
ingestion, PCNA staining increased in the muscle from starved chicks (d
3) and was higher on d 4 (Fig. 7C
, D
, Table 2
). In the
muscle of fed birds, PCNA staining significantly decreased from d 2 to
d 3 and from d 3 to d 4 (Fig. 7G
, H
, Table 2
).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In this study we used several different techniques to follow the kinetics of the satellite cell population, including culturing the cells, thymidine incorporation and determining the numbers of satellite cells at specific stages in the cell cycle. These measurements consistently indicated, within treatments, different patterns of satellite cell activity when birds had no access to food close to hatch.
After 1 d of starving, satellite cell activity did not differ
between groups. However, on that day and after 48 h of starving,
overall cell counts were significantly higher in the starved group than
in the fed group (Figs. 4
, 6
and 7)
. This phenomenon could be due to
muscle atrophy which may result in alteration of myofiber size and in
intramuscular adipose and connective tissue. However, the
differences in breast muscle weight were less than those in cell
number. Therefore, we postulate that a 1-d starvation period activates
satellite cells and brings them into the cell cycle, thus producing
more cells. This phenomenon has been observed to be produced by
mechanical stress, injection of toxic agents and muscle injury (cold,
crush, mince) (Bischoff and Heintz 1994
, reviewed in
Carlson and Faulkner 1983
, Grounds 1998
).
Prolongation of the starvation period caused a dramatic reduction in
satellite cell activity on d 2, and the cells were no longer in the
cell cycle. This reduced the number of cells on the following day. A
reduction in satellite cell number and loss of activity have been
previously documented in malnourished children (Hansen-Smith et al. 1979
), indicating a general phenomenon occurring due to
starvation during the early phase of growth.
In light of these results, it appears that the length of starvation has
an important effect on satellite cell activity. Short-term
starvation may enhance satellite cell number. However, longer
starvation leads to a nearly complete cell-cycle arrest and the
number of satellite cells decrease. It is unlikely that a direct lack
of energy and protein resources caused the in vivo decrease in cell
proliferation in the starved birds (Figs. 6
, 7
, Table 2
) because when
cultured satellite cells prepared from these birds were exposed in
vitro to high energy and serum sources, they did not proliferate (Fig. 4B)
. We contend that these cells were under the influence of some
cell-cycle arrest signal which persisted in culture. Only after the
starved chicks began to eat was the signal turned off, and cells were
then able to proliferate in vivo as well as in vitro. It is tempting to
speculate that cells present in growing tissues in general, and in
skeletal muscle in particular, are sensitive to fluctuations in
nutritional sources above a certain threshold level.
Another variable used to examine cell activity was the expression of
GH-R. Previous studies have demonstrated that GH-R gene
expression peaks in proliferating chick satellite cells in culture and
in the skeletal muscle tissue of very young chicks (Halevy et al. 1996
), suggesting that GH-R expression in the muscle is
correlated with the active satellite cell population. Here, the
resultant pattern of GH-R expression was similar to that of
thymidine incorporation but occurred slightly earlier (see Fig. 4B
). It may well be that GH-R gene expression is more
rapidly affected by stress and therefore precedes the effect on
thymidine incorporation, implying an early post-transcriptional
response to starving conditions. Previously, we demonstrated that
GH-R gene expression in cultured satellite cells is modulated by GH
as well as by basic FGF (Hodik et al. 1997
). The latter
is known to be expressed and secreted by proliferating satellite cells
(Alterio et al. 1990
). Whether systemic and/or local
factors mediate feed-restriction effects on satellite cells and
whether those factors are specific only for those cells remain to be
elucidated.
It appears that the timing of starvation may influence its impact on
body and muscle growth. The closer the starvation period was to hatch,
the lower the birds’ ability to mount a compensatory growth response.
Thus starving for 2 d from d 2 or d 4 posthatch had less of an
effect than starving immediately (Table 1)
. In fact, chicks
starved from d 4 to 6 had full growth compensation by d 41. It has been
shown that feed restriction in the second week posthatch results in
growth retardation that is not only fully recoverable but also results
in compensatory growth (Plavnik and Hurwitz 1988
,
1990
). In this study, chicks starved during the first
days of life did not regain their BW or breast muscle weight by d 41.
This was despite the induction of satellite cell activity and satellite
cell numbers in the starved group in response to food administration to
values above those found in the control fed group. This could imply
full recovery of satellite cell growth in the starved chicks. However,
despite comparable numbers of proliferating satellite cells at
different ages, the maturation of muscle fibers in the starved group
was delayed, probably leading to less hypertrophy and muscle mass.
Since the process of fiber maturation is very rapid and programmed in
broilers, it appears that any delay, such as that caused by starving,
has an irreversible effect on fiber maturation.
In conclusion, these findings are important for achieving optimal muscle growth in all species and have economic implications in animals produced for meat. Sufficient feed in the immediate postnatal period may be critical in influencing later muscle development.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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3 Abbreviations used: BW, body weight; BrdU,
5-bromo-2'-deoxyuridine; DMEM, Dulbecco’s Modified Eagle’s Medium;
FGF, fibroblast growth factor; GH-R, growth hormone receptor; HGF,
hepatocyte growth factor; HS, horse serum; IGF, insulin-like growth
factor; PCNA, proliferating cell nuclear antigen; PDGF,
platelet-derived growth factor; TGF-ß, transforming growth factor beta.
Manuscript received July 23, 1999. Initial review completed August 30, 1999. Revision accepted November 12, 1999.
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REFERENCES |
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 O. Halevy, Y. Nadel, M. Barak, I. Rozenboim, and D. Sklan Early Posthatch Feeding Stimulates Satellite Cell Proliferation and Skeletal Muscle Growth in Turkey Poults J. Nutr., May 1, 2003; 133(5): 1376 - 1382. [Abstract] [Full Text] [PDF]
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L. Lefaucheur, P. Ecolan, Y.-M. Barzic, J. Marion, and J. Le Dividich Early Postnatal Food Intake Alters Myofiber Maturation in Pig Skeletal Muscle J. Nutr., January 1, 2003; 133(1): 140 - 147. [Abstract] [Full Text] [PDF]
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P. E. Mozdziak, J. J. Evans, and D. W. McCoy Early Posthatch Starvation Induces Myonuclear Apoptosis in Chickens J. Nutr., May 1, 2002; 132(5): 901 - 903. [Abstract] [Full Text] [PDF]
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O. Halevy, A. Krispin, Y. Leshem, J. P. McMurtry, and S. Yahav Early-age heat exposure affects skeletal muscle satellite cell proliferation and differentiation in chicks Am J Physiol Regulatory Integrative Comp Physiol, July 1, 2001; 281(1): R302 - R309. [Abstract] [Full Text] [PDF]
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