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The Babraham Institute, Cambridge CB2 4AT, UK
3To whom correspondence should be addressed.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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rhomboideus < cardiac < soleus.
There was a positive correlation with the proportion of oxidative
myofibers (P < 0.001) but not with type I
myofibers (P > 0.10). Compared with the high
intake pigs, hepatic GHR mRNA was downregulated in the low intake pigs
by 59% (P < 0.01), whereas in the four muscles
examined it was upregulated as follows: longissimus, 124%
(P < 0.05); rhomboideus, 19% (P
> 0.4); soleus, 65% (P < 0.05); cardiac,
51% (P < 0.05). Moreover, the proportion of
skeletal muscle fibers with high oxidative capacity was also greater in
the low intake group (P < 0.05). We conclude that
postnatal GHR gene expression and its regulation by mild undernutrition
are related to the metabolic, contractile and specific functional
properties of different muscles.
KEY WORDS: • growth hormone receptor • muscle function • postnatal development • pigs • skeletal and cardiac muscle • undernutrition
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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), whose abundance is regulated developmentally and
nutritionally in a tissue-specific manner. Thus, GHR gene
expression and GH-specific binding occur at an earlier stage of
fetal development in skeletal muscle than in liver (Duchamp et al. 1996
, Lee et al. 1993
,
Schnoebelen-Combes et al. 1996
). Moreover, postnatal
undernutrition induces a marked decrease in hepatic GHR gene expression
but an increase in its expression in longissimus dorsi muscle
(Dauncey et al. 1994
, Weller et al. 1994
).
The specific functions of GH range from key roles in growth via the
GH-insulin-like growth factor-I (IGF-I) somatotrophic axis to
direct metabolic actions (Ross and Buchanan 1990
). The
latter are especially important in peripheral tissues such as muscle,
in which GH has important antilipogenic, lipolytic and diabetogenic
actions. Normal GH secretion, GH binding to its receptor and GHR
expression in early life will therefore be critical factors in
determining optimal metabolic function in muscle. In muscle, GH can
influence not only metabolism but also fiber type, i.e., in fully
differentiated adult rat skeletal muscles it induces a transition from
type II fast oxidative-glycolytic to type I slow oxidative
myofibers (Ayling et al. 1989
, Tsang et al. 1996
). GH also increases the force of contraction in heart and
induces myosin phenoconversion toward the low ATPase activity V3
isoform (Sacca et al. 1994
). In addition, GHR expression
in longissimus dorsi and trapezius muscles is differentially affected
by moderate food restriction in adults (Combes et al. 1997
). From these observations, we speculate that GHR
expression is related to the metabolic and contractile properties of
muscle, which in turn are largely dependent on muscle fiber type.
Moreover, if this is the case, the increase in muscle GHR gene
expression in response to mild postnatal undernutrition (Dauncey et al. 1994
) is also likely to be related to muscle type.
The objectives of this study, therefore, were as follows: 1) to determine whether expression of the GHR gene in morphologically and functionally distinct skeletal and cardiac muscles is related to their metabolic and contractile properties, and 2) to test the hypothesis that the increase in muscle GHR gene expression in response to mild postnatal undernutrition is related to muscle type.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Studies were undertaken in the young pig because it is a good
metabolic, hormonal and developmental model for the human infant
(Tumbleson and Schook 1996
). Specifically, it provides
many similarities in relation to the regulation of muscle development
and function (Dauncey and Gilmour 1996
, Goldspink 1980
, Harrison et al. 1997
, Lefaucheur et al. 1995
). Eight pairs of littermate pigs of the Large White
breed were weaned at 3 wk and kept at thermal neutrality (26°C). They
were housed in pairs for the first 1–2 d and then kept in separate
pens to allow careful control of food intake and environmental
temperature. Food (UltraWean, Dalgety, Bristol, UK) was freely
available for the first 2 d and was then provided as 2 meals/d for
the next 3 wk, with one littermate fed an optimal food intake [60 g
food/(kg·d); 6%] and the other fed a low intake [20 g
food/(kg·d); 2%]. All pigs grew during this period but the rate of
growth was much slower in those with the low compared with the high
intake. The food contained 14 kJ gross energy/g wet food, and comprised
32% carbohydrate, 22.5% protein, 5.5% fat, 3.5% fiber and 6% ash,
with added vitamins and minerals. Water was freely available and
lighting was on from 0900–2100 h daily.
The aim of this study was to determine the long-term effects of nutritional status, rather than the acute effects of feeding. Therefore, because several hormonal and metabolic variables are influenced by food intake, tissue sampling was carried out 15–16 h after the last meal. At 6 wk, pigs were sedated with an intramuscular injection of ketamine hydrochloride (1.0 mL Vetalar, 100 g/L; Parke-Davis Veterinary, Pontypool, Gwent, UK) and killed with a 0.7 mL/kg body intracardiac injection of pentobarbitone sodium (Lethobarb, 200 g/L; Duphar Veterinary, Southampton, Hampshire, UK). Samples of liver and a range of morphologically and functionally distinct muscles were dissected rapidly; care was taken to sample from the same relative anatomical position. Muscles were longissimus dorsi (longissimus; dorsal lumbar), rhomboideus (dorsal interscapular), soleus (hind-limb), diaphragm and cardiac. All procedures were approved by the UK government under the Animals (Scientific Procedures) Act 1986, and by The Babraham Institute’s Animal Welfare, Experimentation and Ethics Committee.
For assessment of GHR gene expression, liver and muscle samples were divided into 5-g portions, frozen in liquid nitrogen and stored at -70°C. For assessment of metabolic and contractile properties of muscle, samples of 1 cm3 were mounted on cork blocks, surrounded by Cryo-m-bed (Bright Instrument, Cambridge, UK), and frozen immediately in isopentane cooled in liquid nitrogen before storage at -70°C. Serial 10-µm cross sections were later cut on a cryostat at -22°C and used for measurement of type I slow myosin heavy-chain (MyHC) protein level by immunocytochemistry, and oxidative enzyme activity by histochemistry.
In relation to endocrine systems of particular relevance to muscle
development, we previously undertook extensive investigations using
pigs of similar age and treatment to those in this study. Those studies
focused on the GH-IGF axis, the thyroid axis and glucocorticoids
(Dauncey et al. 1989
and 1994
, Dauncey and Buttle 1990
, Morovat and Dauncey 1998
) and demonstrated
that a complete understanding of these systems can be achieved only by
frequent sampling over a 24-h period. Such an approach would have been
impossible in this study; therefore the current findings are discussed
in relation to the earlier investigations.
Construction of GHR probes.
Using oligonucleotide primers taken from the published porcine GHR
sequence (Cioffi et al. 1990
), polymerase chain reaction
(PCR) was carried out on porcine liver cDNA to generate a DNA fragment
from the intracellular domain of the receptor; probes for either the
intracellular or extracellular domains can be used to detect mRNA for
the complete GHR (Dauncey et al. 1994
). The 5' primer
[5'-(TCTAGA)CAGCAAAGGATTAAGATG-3'], representing nucleotides 886–903
of the GHR sequence, contained an XbaI recognition site (in brackets),
and the 3' primer [5'-AACCCAAGAGTCATC-3'] was the complement of
nucleotides 1033–1048. The resulting PCR product was digested with
XbaI and EcoRI to give a 140-bp fragment by virtue of an internal EcoRI
site at position 1022. The fragment was cloned unidirectionally into
Bluescript (Stratagene, Cambridge, UK) and verified by DNA sequencing.
An antisense riboprobe was generated by first linearizing the plasmid
by digestion with XbaI and then transcribing with T7
polymerase in the presence of [
32P]UTP. The probe had
a full length of 200 nucleotides, of which 140 were protected in the
RNase protection assay.
Total RNA extraction and RNase protection analysis.
The methods used have been described in detail previously
(Dauncey et al. 1994
, Duchamp et al. 1996
, Weller et al. 1993
, White and Dauncey 1998a
). In brief, total RNA was isolated from 0.5-g
samples of frozen tissue using the guanidinium thiocyanate method and
quantified by absorbance at 260 nm, where 1 optical density (OD) unit
= 40 mg RNA/L solution. The integrity of total RNA extracted was
routinely checked by Northern gel electrophoresis. This demonstrated
that both the 28S and 18S subunits were intact, in the appropriate 2:1
ratio, indicating excellent integrity of the preparations. Each lane
also showed an identical level of loading as determined by ethidium
bromide staining and image analysis. In addition, the equivalence of
RNA samples from different tissues and treatments was checked by
analyzing total poly(A)+ content. This approach was used
because, although it is considerably more time-consuming than the
use of an RNase protection assay probe designed for the 28S or 18S
subunits, use of the latter is misleading. Titration analysis has shown
that because the 28S and 18S subunits are such highly abundant RNA, it
is not possible to produce enough radiolabeled probe to detect them
accurately. The 28S and 18S subunits represent 90–95% total RNA, and
a 50-µg RNA sample contains between 20 and 30 pmoles
of 18S RNA. To obtain the required excess of 18S control probe, at
least 125 pmoles of radiolabeled RNA need to be added per sample. The
maximum theoretical yield of RNA from a standard in-vitro transcription
reaction is less than 0.5 µg, which for a 150-bp probe is equivalent
to 10 pmoles. Thus, to achieve the required molar excess would require
the addition of at least 12 labelling reactions per sample. Since only
a diluted sample of one such labelling reaction can be used in the
RNase protection assay, the use of such a probe is erroneous and highly
misleading, i.e., all samples appear to have the same signal intensity
because the amount of target 18S RNA considerably exceeds the amount of
probe. Thus, the probe will be an unreliable control, and differences
will not be the result of true differences in the amount of total RNA.
Rather, they will represent poor experimental technique, such as loss
of sample during phenol/chloroform extraction, loss of pellet during
washing steps or poor resuspension of the final "protected" sample
before loading. Numerous problems are also associated with the use of
"housekeeping" genes as RNA controls (Ivell 1998
).
We therefore developed a highly optimized RNase protection assay
procedure that minimizes these common errors. To summarize, rather than
misinterpreting the data by using a 28S/18S or "housekeeping"
control probe, the following alternative approach was used, which our
experience suggests is more appropriate: 1) testing the
prepared RNA by denaturing formamide gel electrophoresis;
2) measuring total poly(A)+ content; and
3) using a rigorously optimized RNase protection assay
on a large number of animals in each treatment group. Additional
factors that further minimize the risk of artifacts include the use of
duplicate samples throughout and an experimental design that enables
the use of paired t tests or one-way randomized
block design ANOVA. Samples from littermate piglets subjected to each
of the treatments are loaded onto the same gel; hence, a series of
differences between two pigs can be compared, rather than simply the
average difference between the two groups.
RNase protection assays were carried out in duplicate with 50
µg total RNA extracted from the different tissues,
using the following procedure. Samples were hybridized with a small
molar excess of the radiolabeled GHR riboprobe to ensure linearity of
the assay with respect to RNA. After 16 h hybridization at 45°C,
excess nonprotected RNA was digested with RNase A (50 mg/L, 
1
U/sample) and RNase T1 (3 x105
U/L, 80 U/sample). The protected
hybridization products were purified by extraction in
phenol/chloroform/isoamyl alcohol (25:24:1) and separated on 6%
polyacrylamide sequencing gels. The dried gels were exposed to
X-ray film at -70°C, and relative intensities of the protected
bands were quantified by image analysis. The system was linear over the
range of optical density values measured.
Immunocytochemistry.
The proportion of type I slow-twitch/cardiac ß fibers in each muscle was determined using indirect immunoperoxidase staining with a slow MyHC (type I) specific monoclonal antibody (Clone No. MHCs, Biogenesis, Poole, Dorset, UK). The antibody had been raised in the mouse using native type I myosin from rabbit soleus as an immunogen. Frozen sections were placed on APTES-coated slides, fixed in 4% paraformaldehyde for 10 min, and washed in Tris-buffered saline (TBS) for 20 min. Sections were blocked with normal horse serum for 30 min and incubated with the primary MyHC (type I) antiserum diluted 1:50 in normal horse serum for 1 h at room temperature. After 3 x 10 min washes in TBS-Tween, sections were incubated with a peroxidase-labeled anti-mouse immunoglobulin G antibody (Vector Laboratories, Peterborough, UK) and diluted 1:200 in normal horse serum. Sections were washed, incubated with an avidin-biotin complex (ABC reagent; Vector), rinsed and incubated in 0.03% H2O2 in TBS + 1 g/L diaminobenzidine until the stain was seen to develop. After dehydration through ethanol and xylene, the sections were mounted in a xylene-based mounting medium.
Enzyme histochemistry.
The proportion of oxidative fibers in each muscle was determined using histochemical staining for succinate dehydrogenase (SDH). Air-dried frozen sections were incubated at 37°C in medium containing 0.125 mol/L sodium succinate, 1 g/L nitro blue tetrazolium, 0.005 mol/L MgCl2, 0.05 mol/L Tris-HCl, pH 7.4, for 1 h. Sections were fixed in formol saline for 10 min, washed in water and mounted in glycerin jelly. Myofibers were observed as having high (++), intermediate (+) or low (-) SDH activity; hence the proportion of total oxidative fibers (++ plus +), and those with high oxidative activity (+) could be assessed.
Fiber type distribution.
The relative proportions of type I slow-twitch and oxidative myofibers in each muscle sample were determined in four random fields of view within a standard field of 119,000 µm2. The numbers of specific fibers were counted with a Seescan A010 research-grade image analysis system (Cambridge, UK), and values were expressed as a percentage of the total fiber count.
Statistical analysis.
The results for GHR mRNA in different muscles were tested for significance by ANOVA for randomized block design using the statistical package Genstat (Lawes Agricultural Trust, Rothamsted, Hertfordshire, UK), in which muscles were main effects and gels were blocks because skeletal and cardiac muscle RNA samples from each pig were loaded onto the same gel. The relation between GHR mRNA and the proportion of oxidative fibers was assessed using a regression model with different intercepts for each pig but a common slope. A similar analysis was carried out for GHR mRNA vs. proportion of type I slow-twitch fibers. Student’s paired t test was used to test for significance between the two treatment groups. Results are presented as means ± SEM. Differences of < 0.05 were considered significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Pigs in the high energy intake group (6%) grew very rapidly during the treatment period, reaching a mean body weight that was almost twice that of their littermates in the low energy intake group (2%). At the start of investigation, at 3 wk of age, there was no difference in body weights, which were 5.6 ± 0.4 and 5.6 ± 0.3 kg for the 6 and 2% groups, respectively. By 6 wk, values had increased to 12.8 ± 0.6 and 7.2 ± 0.4 kg, respectively. During the last 2 wk of investigation, the average growth rate of pigs in the high intake group was 452 ± 30 g/d, whereas that of pigs in the low intake group was 122 ± 7 g/d (P < 0.001).
Muscle-specific differences in GHR gene expression.
An autoradiograph showing GHR mRNA abundance in liver and functionally
distinct muscles from a pig in the optimal 6% energy intake group is
illustrated in Figure 1
. Overall mean values and SEM for skeletal and cardiac
muscles in the 6% group are presented in Figure 2
. There were muscle-specific differences in GHR gene expression
(P < 0.001).
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Table 1
presents results for the metabolic (oxidative vs. nonoxidative) and
contractile (type I slow vs. type II fast) properties of the four
skeletal muscles examined in the 6% group. The proportion of oxidative
myofibers was greatest in soleus, intermediate in rhomboideus and
diaphragm, and lowest in longissimus. The greatest proportion of type I
fibers occurred in soleus and the least in longissimus. Cardiac muscle
comprised 100% oxidative and 100% type I/cardiac ß fibers. Further
details on MyHC isoform expression and metabolic enzyme activity in the
four skeletal muscles examined can be found elsewhere (White et al. 2000
).
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There were striking muscle-specific differences in GHR mRNA
expression (P < 0.001), and GHR expression was greater
in slow-oxidative than in fast-glycolytic muscles. The relation
between GHR mRNA abundance and metabolic/contractile properties of the
four skeletal muscles investigated, i.e., longissimus, rhomboideus,
soleus and diaphragm, is shown in Figure 3
, which indicates that the relation between GHR and oxidative capacity
was especially close. The correlation between GHR mRNA and total
oxidative fibers was significant (P < 0.001); GHR mRNA
increased by 0.056 ± 0.018 for each unit increase in proportion
of oxidative fibers. By contrast, there was no significant relationship
between GHR mRNA and total type I slow fibers (P > 0.10).
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A low food intake was found to result in a lower hepatic GHR mRNA level
(P < 0.01) and more GHR mRNA in all four muscles
examined (longissimus, rhomboideus, soleus and cardiac). Moreover, the
influence of nutrition was muscle specific; the upregulation of GHR was
most striking in longissimus (P < 0.05), intermediate
in soleus and cardiac (P < 0.05) and least in
rhomboideus muscle (P > 0.4). These findings are
illustrated in Figure 4
, which shows the percentage difference in GHR mRNA in pigs in the 2%
group compared with the 6% group. Thus, the greatest effect of
nutrition occurred in the muscle with the lowest abundance of GHR, the
least oxidative capacity and lowest proportion of type I
slow-twitch fibers. During food deprivation or severe
undernutrition, the total RNA content of many tissues can decrease
(Champigny and Ricquier, 1990
), and degradation of RNA
can result in the OD260 being less representative
of the total RNA in the sample. In all tissues, except soleus
(P < 0.05), there was no influence of nutrition on
total RNA (P > 0.05). Together with Northern analysis
revealing excellent integrity of RNA samples, the finding of no overall
reduction in tissue RNA content reinforces the point that the effects
of mild undernutrition were being investigated and that all pigs grew
throughout the study.
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. In all muscles, the proportion of myofibers with high oxidative
activity was greater in the 2% compared with the 6% group, i.e.,
longissimus, P < 0.05; rhomboideus, P
< 0.001; soleus, P < 0.05. By contrast, the
proportion of myofibers with a high level of type I slow MyHC protein
expression was affected only in rhomboideus, i.e., 2% group > 6% group (P < 0.05).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Muscle GHR gene expression is related to its functional properties.
Postnatal GHR gene expression was found to be greater in type I slow
oxidative than type II fast glycolytic/oxidative-glycolytic muscle, and
there was a particularly close relation between GHR mRNA and the
proportion of oxidative myofibers. GH exerts a variety of functions in
muscle, and the present findings (see Figs. 2
, 3
) strongly support the
close association of its metabolic action in muscle with oxidation of
energy substrates. Our results for muscle-specific differences in
GHR mRNA expression suggest that slow oxidative muscles will be more
responsive to GH than fast glycolytic muscles. The extent to which
endogenous changes in GH, induced by nutrition and thermal environment
(Dauncey and Buttle, 1990
, Ross and Buchanan, 1990
), can affect myofiber type remains to be established.
Nevertheless, exogenous GH can induce a transition from type II fast
oxidative-glycolytic to type I slow oxidative myofibers in adult
rats (Ayling et al. 1989
, Tsang et al. 1996
). Taken together, these findings suggest that GH and its
receptor are important factors in determining the metabolic and
contractile properties of muscle, and in defining the specific
functions of morphologically distinct muscles.
Postnatal undernutrition induces upregulation of GHR gene expression in skeletal and cardiac muscles.
We reported previously that mild postnatal undernutrition, which
nevertheless enables growth to continue but at a reduced rate,
increases GHR mRNA expression in longissimus muscle (Dauncey et al. 1994
). We have now extended this finding by demonstrating
that GHR mRNA in rhomboideus, soleus and cardiac muscles is upregulated
by postnatal undernutrition, although to varying extents. Because the
pigs grew throughout the study, the probability is that the effect on
muscle GHR mRNA will be even greater when nutritional restriction
prevents growth and induces the catabolic state. The upregulation of
muscle GHR in the low food intake group is probably a specific effect
and not related simply to an alteration in the developmental pattern of
GHR because previous studies have shown that there are no clear
age-related changes in muscle GHR in pigs
(Schnoebelen-Combes et al. 1996
).
These findings contrast with adult pigs in which GHR mRNA abundance in
longissimus was not affected by food restriction, although it was
upregulated in trapezius (Combes et al. 1997
). In this
earlier study, the restricted food intake was 70% that of the
controls. The difference between these results and the earlier study
may therefore be related to stage of development and/or magnitude of
undernutrition. Indeed, we have recently found that mild postnatal
dietary lysine deficiency, which nevertheless allowed pigs to
gain body weight, upregulates GHR mRNA expression in both longissimus
and rhomboideus (M. Katsumata, R. Takada and M. J. Dauncey,
unpublished results).
In general, the metabolic role of GH in muscle is anti-insulin and
includes antilipogenic, lipolytic and diabetogenic actions, which all
act to divert energy from muscle. The upregulation of muscle GHR gene
expression in response to postnatal undernutrition can thus be viewed
as an adaptation that spares neural and bone growth at the expense of
muscle growth. When energy supplies are limited, they will be conserved
predominantly for brain, nerve and/or bone rather than for muscle
growth. These results suggest that the nutritionally induced changes in
GHR mRNA expression are closely related to concomitant changes in
metabolic properties of the muscle (Fig. 5)
, and specifically the
greater proportion of myofibers with high oxidative capacity and the
transition from glycolytic/oxidative-glycolytic to oxidative
myofibers that occurs during undernutrition. GH induces a transition
from fast oxidative-glycolytic to slow oxidative myofibers in fully
differentiated adult rat skeletal muscles (Ayling et al. 1989
, Tsang et al. 1996
). This raises the
hypothesis that the nutritionally induced upregulation of GHR
expression, together with changes in other hormone systems
(Dauncey and Gilmour 1996
, White et al. 2000
), may induce the transition in myofiber type. Moreover,
the change in fiber type will itself enhance energetic efficiency
during undernutrition.
Mechanisms mediating the nutritionally induced changes in GHR
expression may involve thyroid hormones and glucocorticoids acting as
signals of energy status, together with tissue-specific changes in
the transcriptional regulator superfamily of nuclear hormone receptors
(Chatterjee and Tata 1992
, Dauncey 1997
,
Lazar 1993
). In cultured porcine hepatocytes, energy in
the form of glucose appears to control GHR mRNA expression and
interacts with the effects of thyroid hormones and glucocorticoids
(Brameld et al. 1999
). Both of these groups of hormones
in turn play key roles in regulating GHR expression during development
(Cabello and Wrutniak 1989
, Duchamp et al. 1996
, Li et al. 1996
). Consistent with these
findings, the response is tissue specific, i.e., prenatal
hypothyroidism induces downregulation of hepatic GHR mRNA but a marked
upregulation of muscle GHR mRNA expression (Duchamp et al. 1996
). Considerable evidence indicates that nutrition, and
especially energy status, induces marked changes in thyroid hormones
and glucocorticoids (Danforth and Burger 1989
,
Dauncey 1990
, Morovat and Dauncey 1995
,
Vance and Thorner 1989
). Postnatally, a reduction in
food intake that is not severe enough to prevent growth induces
hypothyroidism by reducing thyroid gland activity, plasma thyroid
hormone levels and nuclear thyroid hormone binding capacity of skeletal
muscle. Recent evidence also suggests that mild undernutrition may
induce muscle-specific differences in thyroid hormone receptor
isoform expression (White and Dauncey 1998b
and 1999
),
suggesting a mechanism by which nuclear hormone receptors may
selectively mediate upregulation of GHR gene expression in functionally
distinct muscles.
Upregulation of GHR gene expression by undernutrition is greatest in fast glycolytic/oxidative-glycolytic muscle.
This study has demonstrated that the magnitude of upregulation of GHR
gene expression in response to postnatal undernutrition differs among
morphologically distinct muscles. The greatest effect of a low energy
intake occurred in longissimus, which contains mainly fast
glycolytic/oxidative-glycolytic fibers and has the lowest abundance of
GHR mRNA. An insight into the metabolic significance of this
muscle-specific response may be gained from expression studies of
two recently cloned genes, uncoupling proteins (UCP) 2 and 3
(Samec et al. 1998a
and 1998b
). These proteins have been
implicated in regulation of lipids as fuel substrates in muscle, and
both genes are upregulated in rats during food deprivation. Moreover,
the magnitude of upregulation is markedly greater in gastrocnemius and
tibialis anterior, predominantly fast glycolytic/oxidative-glycolytic
muscles, than in the slow oxidative soleus muscle (Samec et al. 1998a
and 1998b
). These differences in UCP gene expression were
attributed to differences in responsiveness of myofiber types to food
deprivation. Gastrocnemius and tibialis anterior have a high capacity
to shift between glucose and lipids as energy sources, whereas soleus
has a higher dependency on lipids. During fasting, when glucose is
unlikely to be a major energy source and fat stores are mobilized,
upregulation of UCP homologues will enhance the oxidative capacity of
muscle, and especially those muscles that have a high capacity to shift
their energy source between glucose and lipids.
Our observations on GHR gene expression lend support to the central
role of fast-twitch glycolytic/oxidative-glycolytic muscles in
metabolic fuel mobilization during undernutrition. Although our
treatment was not as severe as food deprivation, it is probable that
pigs in the low food intake group were more highly dependent on lipids,
and especially fatty acids, as an energy source than were those in the
high intake group. Muscle likely adapted to the suboptimal energy
status by shifting its energy source from glucose to lipids via
upregulation of GHR gene expression. Upregulation was greatest in the
fast glycolytic/oxidative-glycolytic longissimus because of its
capacity to shift substrate utilization between glucose and lipids,
compared with slower oxidative muscles such as soleus and rhomboideus.
In relation to potential regulatory factors of muscle gene expression,
it has been suggested that circulating free fatty acids constitute an
interorgan signal that regulates expression of UCP2 and UCP3 in
oxidative muscles such as soleus, whereas other factors regulate
expression of these UCP homologues in fast
glycolytic/oxidative-glycolytic muscles such as gastrocnemius and
tibialis anterior (Samec et al. 1998a
). Thus, plasma
metabolites such as free fatty acids and glucose, together with thyroid
hormones, glucocorticoids, insulin or GH itself may function as
interorgan signals regulating muscle-specific GHR gene expression
in response to postnatal nutritional status.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Funded by the Biotechnology and Biological Sciences Research Council (The Babraham Institute, K.A.B. and M.J.D.); Japanese Science and Technology Agency (Postdoctoral Fellowship to M.K.); Italian Ministry of University and Scientific and Technological Research, Research Project 40% - Professor V. Dell’Orto (D.C.); and Medical Research Council (Postgraduate Studentship to P.W.).
4 Abbreviations used: GH, growth hormone; GHR, growth hormone receptor; IGF-I, insulin-like growth factor-I; MyHC, myosin heavy chain; OD, optical density; PCR, polymerase chain reaction; SDH, succinate dehydrogenase; TBS, Tris-buffered saline; UCP, uncoupling protein.
Manuscript received January 26, 2000. Initial review completed March 23, 2000. Revision accepted June 1, 2000.
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