1 Respiratory Division, McGill University Health Centre, Montreal, Quebec H3A 1A1; 2 Respiratory Muscle Biology Group, Meakins-Christie Laboratories, McGill University, Montreal, Quebec, Canada H2X 2P2; and 3 Universidade Federal do Rio de Janeiro, Instituto de Biofisica Carlos Chagas Filho, 21949-900 Rio de Janeiro, Brazil
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ABSTRACT |
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Although prolonged diaphragm denervation (DNV) produces myofiber atrophy and a loss of type I myosin heavy chain (MHC) expression, short-term DNV leads to significant diaphragm hypertrophy. The purpose of this study was to explore the regulation of MHC isoform expression and muscle remodeling during DNV hypertrophy of the diaphragm. Both unilateral and bilateral DNV led to similar changes, with a significant increase in total RNA content and muscle mass but no change in dry-to-wet weight ratio. Sarcomere number was also increased in diaphragm myofibers after DNV (~20%), suggesting an adaptive response to muscle stretch. There was hypertrophy of type I myofibers and increased coexpression of type I and type II MHCs within single myofibers by immunocytochemistry as well as increased type I MHC (25-46%) and decreased type IIb MHC (14-39%) by SDS-PAGE. Contractility parameters were also consistent with a type II-to-type I MHC phenotype transformation. Importantly, DNV-induced modulation of MHC isoform mRNA transcript levels did not correspond to changes in their cognate proteins, suggesting a major degree of posttranscriptional control. We conclude that DNV hypertrophy of the diaphragm is associated with reciprocal changes in type I and type II MHC isoforms that are directly opposed to the type I-to-type II MHC phenotype transformation reported in the diaphragm DNV atrophy model. Furthermore, in contradistinction to most hypertrophy models, control of MHC gene expression and myofibrillar remodeling after short-term DNV appears to entail major involvement of posttranscriptional regulatory mechanisms.
myosin heavy chain isoforms; denervation hypertrophy; muscle stretch; posttranscriptional regulation; unilateral and bilateral diaphragm paralysis
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INTRODUCTION |
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DENERVATION (DNV) of the diaphragm may result from a number of identifiable causes including trauma, malignancy, infection, and diffuse neuropathies involving the phrenic nerve. Inactivation of the diaphragm may also be found in patients in the Intensive Care Unit who are undergoing mechanical ventilation for acute respiratory failure due to administration of neuromuscular blocking agents. The diaphragm differs from traditionally studied limb skeletal muscles by its longer duty cycle (27) as well as its persistent rhythmic activation throughout both wakefulness and sleep without any period of sustained rest. In addition, differences in the intrinsic genetic programming of myoblast precursor cells derived from different skeletal muscles have been reported (10). Because of the unique physiological demands placed on the diaphragm as well as factors intrinsic to the biology and adaptive capacities of the muscle itself, regulation of diaphragm myofiber gene expression in response to various environmental cues may differ from that observed in limb muscles. In particular, the response to changes in activity level and altered neural input may not be identical to that observed in other muscles.
Myosin heavy chain (MHC) constitutes the predominant protein in
skeletal muscle as well as the molecular basis for the traditional histochemical classification into type I (slow-twitch) and type II
(fast-twitch) fiber types (9). Differences in myofibrillar ATPase
staining, maximum shortening velocity, and the economy of force
generation are directly correlated with the presence of distinct MHC
isoforms (3, 8, 9). In the mammalian diaphragm, these consist of type I
(also known as -cardiac), type IIa, type IIx, and type IIb MHC
isoforms (18). Immunocytochemical analysis has demonstrated alterations
in the pattern of MHC isoform expression along with muscle atrophy
after >4 wk of hemidiaphragm DNV (6, 7, 14). The changes consist of a
shift toward fast type II MHC isoform expression by the vast majority
of diaphragm myofibers, thereby producing an increasingly homogeneous
population of atrophic diaphragm myofibers lacking in characteristics
associated with the slow type I MHC phenotype (6, 7, 14).
In contrast, shorter periods of hemidiaphragm DNV lead to significant muscle hypertrophy on the inactivated side (12), which is accompanied by increased protein and DNA synthesis that is maximal during the first week after DNV (35, 37). This hypertrophic response to DNV has generally been attributed to rhythmic passive stretching of paralyzed hemidiaphragm myofibers by persistent contractions of the normally innervated contralateral hemidiaphragm (12). However, although passive stretch is a known stimulus for muscle protein synthesis and growth (16), this interpretation has recently been challenged by a report (38) that hypertrophy also occurs in regions of the diaphragm not subjected to intermittent stretch by contralateral hemidiaphragm activity. Therefore, the degree to which the myofiber hypertrophy observed after unilateral hemidiaphragm inactivation is due to mechanical stretch caused by contralateral hemidiaphragm contractions, as opposed to other consequences of removing innervation, is presently unclear.
The previous reports of ubiquitous expression within all diaphragm myofibers of type II MHC isoforms after longer term DNV (6, 7, 14) are consistent with the proposition that skeletal muscles express fast myosin isoforms by default in the absence of intact innervation (1, 4, 13, 17, 29). However, diaphragm inactivation over this time period (6, 7) is accompanied by a substantial loss of muscle mass. We were therefore interested in determining whether a similar pattern of increasingly homogeneous fibers with augmented type II MHC expression is also found during the earlier post-DNV period when diaphragm hypertrophy rather than atrophy is present.
Accordingly, in the present study, we examined the expression of MHC isoforms at the level of both mRNA and protein to ascertain the dynamics of MHC gene regulation during DNV hypertrophy of the diaphragm. Here we report that both type I and type II diaphragm myofibers respond in a qualitatively similar manner to short-term DNV, with a relative shift toward increased type I MHC rather than toward type II MHC isoform expression within both fiber-type populations. We also demonstrate evidence for a complex pattern of MHC isoform gene regulation in this setting, which appears to involve a substantial degree of posttranscriptional control. Finally, by directly comparing the effects of unilateral and bilateral diaphragm inactivation on the above parameters, we have attempted to gain insight into the potential role of muscle stretch produced by contralateral hemidiaphragm contractions in the induction of these changes.
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METHODS |
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Six-week-old male Sprague-Dawley rats were randomly divided into three groups: 1) control (CTL), 2) unilateral diaphragm DNV (UNI-DNV), and 3) bilateral diaphragm DNV (BIL-DNV). All measurements were based on a study of six rats from each experimental group. The rats were housed at room temperature with a 12:12-h light-dark cycle and provided with rat chow and water ad libitum. Animals were euthanized on the 8th post-DNV day; this time point was selected on the basis of the fact that a relatively stable elevation in the rate of protein synthesis (~50% above control value) is present between the 5th and 10th days after diaphragm DNV (35). The animals were euthanized by an overdose of pentobarbital sodium administered by intraperitoneal injection. All aspects of the study were approved by the Institutional Animal Care and Use Committee.
Surgical procedures. The animals were anesthetized by administration of pentobarbital sodium (45 mg/kg ip) in preparation for surgery. Under aseptic conditions, a midline incision was made in the lower neck, and the cervical portions of both phrenic nerves were visualized in all animals. The incision was closed without further manipulation in the CTL group, whereas the phrenic nerves were isolated and transected (removing at least 5 mm and separating the opposing ends to prevent reinnervation) in the UNI-DNV (left side only) and BIL-DNV groups. In the two DNV groups, diaphragm paralysis was apparent clinically in all animals by the observation of paradoxical inward motion of the abdomen during inspiration.
Diaphragm contractile properties. To ascertain the functional consequences of altered MHC isoform expression and to determine the physiologically optimal muscle length (Lo) for subsequent sarcomere measurements (see Determination of diaphragm mass and sarcomere number), in vitro contractility studies were performed. After euthanasia, the diaphragm was quickly removed (<60 s) en bloc with its intact rib cage as previously described (22). The freshly dissected diaphragm was immediately transferred to Ringer solution (119 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl2, 1.2 mM KH2PO4, 1.2 mM MgSO4, 20 mM NaHCO3 and 6 µM D-tubocurarine chloride) perfused with 95% O2-5% CO2 (pH 7.4) and chilled to ~15°C. In each rat, a diaphragm strip with intact attachments to both rib cage and central tendon was dissected free from the costal diaphragm. Measurements were performed on the left hemidiaphragm in all animals, and great care was taken to ensure that the strips (3-4 mm in width) were taken from the identical portion of the muscle in each animal by centering the dissection on the insertion point of the phrenic nerve. The diaphragm strip was then mounted vertically in a jacketed tissue bath chamber filled with continuously perfused Ringer solution. The temperature of the bath was maintained at 25°C, and a thermoequilibration period of 10 min was observed before contractile measurements were initiated. The rib cage was securely anchored to a platform near the base of the chamber while the opposite end consisting of central tendon was tied to the lever arm of a force transducer-length servomotor system (model 300B dual mode, Cambridge Technology, Watertown, MA) (22). The latter was mounted on a mobile micrometer stage (Newport Instruments, Toronto, Canada), which permitted fine adjustments of muscle strip length. Diaphragm strip length was incrementally adjusted to obtain Lo, defined as that length at which maximal force was achieved. Supramaximal stimulation was induced via two platinum plate electrodes placed into the bath on both sides of the strip. Stimuli with a monophasic pulse duration of 2 ms were delivered with a computer-controlled electrical stimulator (model S44, Grass Instruments, Quincy, MA) connected in series to a current amplifier (model 6824A, Hewlett-Packard). Muscle force was displayed on a storage oscilloscope (Tektronix, Beaverton, OR), and the data were simultaneously acquired to a computer at a sampling rate of 1,000 Hz for later analysis. The following parameters were measured: twitch contraction time (time to peak force), half-relaxation time (time for force to decrease to one-half of its peak value), maximal twitch force, and maximal tetanic force. After a rest period of 15 min, the fatiguability of the diaphragm was assessed by measuring the loss of force in response to repeated stimulations (30 Hz, 0.33 duty cycle, 90 trains/min). The fatigue index was calculated as the percentage of initial force remaining after 2 min of muscle stimulation (22). The muscle strip was then removed from the bath and its length from the rib to the central tendon was measured under a dissecting microscope with a microcaliper accurate to 0.1 mm. Diaphragm strip cross-sectional area was approximated by dividing muscle mass by its length and density (22); this allowed specific force (force/cross-sectional area) to be calculated, which is expressed as newtons per square centimeter.
Determination of diaphragm mass and sarcomere number. The remainder of the costal diaphragm was divided into two halves for determination of left (including the diaphragm strip used for the above contractility measurements) and right hemidiaphragm wet weights. The dry weight-to-wet weight ratio of the tissue samples was also measured. Given the previously proposed role of mechanical stretch in DNV hypertrophy of the diaphragm (12), we determined sarcomere number in isolated myofibers because this parameter is known to increase rapidly after muscle stretch (36). The diaphragm strips used to determine contractile properties were fixed at Lo in 4% glutaraldehyde-Millonig buffer (0.12 M NaH2PO4, 0.07 M NaOH, and 5 g/l of glucose, pH 6.8) for 4-6 h (25). Diaphragm strips were then removed from the fixing solution, rinsed, and mechanically dissociated into single fibers by a low-speed homogenizer (model PCU 11, Kinematica). The dissociated fibers were pelleted by centrifugation at 34,500 g for 30 min, plated on glass slides, and stained with 1% p-phenylenediamine. The dissociated fibers were viewed microscopically (×1,000), allowing images to be photographed with a video camera and stored on a Macintosh computer. Sarcomere length was determined from the calibrated computer image with the public domain program National Institutes of Health Image (version 1.49). For each animal, images from three different randomly selected fibers were used for the analysis, and sarcomere length measurements were made on 20 contiguous sarcomeres/fiber. Total sarcomere number was then calculated by dividing the Lo of the diaphragm strip by the mean sarcomere length (31).
Immunocytochemical analysis of MHC
isoforms. In each rat, a second costal diaphragm strip
(immediately anterior to that used for determination of contractile
properties and sarcomere characteristics) was removed and snap-frozen
in isopentane precooled with liquid N2. To control for possible
effects of muscle length on fiber cross-sectional area, diaphragm
strips were pinned at
Lo (as determined from the first strip) before they were frozen (22). Indirect immunofluorescence was then used to classify MHC fiber types as previously described (22, 23). Briefly, the diaphragm was embedded in
mounting medium, and serial sections (6 µm thick) midway between the
central tendon and rib cage were cut with a cryostat at
20°C. After the sections were air-dried, they were treated
with antibodies (22, 23) specific for slow type I (NOQ7.5.4D, 1:100)
and fast type II (MY-32, 1:200) MHC isoforms (the latter reacts with
all type II MHCs: IIa, IIx, and IIb). The primary antibodies (raised in
mouse) were diluted in PBS and incubated with tissue sections at
4°C overnight, rinsed, and then incubated for an additional hour
with fluorescein isothiocyanate-conjugated goat anti-mouse IgG (1:200)
secondary antibody at room temperature. By reacting serial sections
with the above antibodies, it was also possible to identify hybrid
fibers that were in transition from a type I to a type II MHC phenotype
or vice versa (i.e., fibers coexpressing both type I and II MHC
isoforms), which is not possible with traditional myofibrillar ATPase
histochemistry. Serially stained diaphragm sections were examined under
epifluorescence microscopy and categorized as type I, type II, or
hybrid fibers. The images were captured to a computer and the number of
fibers in each category was determined from randomly selected fields (minimum of 200 fibers/muscle); individual fiber cross-sectional areas
were then directly measured from the calibrated computer image. These
measurements allowed determination of the contribution of each fiber
type to the muscle tissue component of the diaphragm in terms of
relative proportion as well as relative area (i.e., the product of
fiber proportion and mean cross-sectional area for each fiber type
expressed as a percentage of the cumulative cross-sectional area of all
fiber types within the microscopic field examined).
MHC isoform separation by SDS-PAGE.
Myosin isolation and separation of MHC isoforms by SDS-PAGE were
performed with minor modifications of a previously published protocol
(18). Briefly, muscle samples were minced and extracted on ice in 4 volumes of buffer (300 mM NaCl, 100 mM
NaH2PO4,
50 mM
Na2HPO4,
1 mM MgCl2, 10 mM
Na4P2O7,
and 10 mM EDTA) at pH 6.5. Extracts were centrifuged at 13,000 g for 30 min at 4°C, and the
supernatants were recovered, diluted in 9 volumes of 1 mM EDTA-0.1%
-mercaptoethanol, and stored overnight at 4°C to allow
precipitation of myosin filaments. This solution was subsequently
centrifuged at 13,000 g for 30 min at
4°C to form a pellet, which was resuspended (1:1) in 0.5 M NaCl-10
mM
NaH2PO4
and then diluted 1:100 in SDS buffer composed of 62.5 mM
Tris · HCl, 2% (wt/vol) SDS, 25% (vol/vol)
glycerol, 350 mM dithiothreitol, and 0.01% (wt/vol) bromphenol
blue (pH 6.8). The samples were boiled for 2 min and stored at
80°C. Separation of MHC isoforms was accomplished with slab
gels (10.5 cm × 7.5 cm × 0.75 mm) consisting of a 2.5-cm
stacking gel and a 5-cm separating gel. Both gels were prepared from a
30% acrylamide stock solution containing 29% (wt/vol) acrylamide and
1% (wt/vol)
N,N'methylene-bis-acrylamide. Final concentrations of polyacrylamide were 4 (stacking) and 8% (separating) with the addition of glycerol (40% vol/vol).
Polymerization was activated by addition of 0.03% (wt/vol)
ammonium persulfate and 0.10% (vol/vol)
tetramethylethylenediamine. Approximately 1 µg of myosin was loaded
onto each lane, and electrophoresis was performed with Tris-glycine
running buffer (pH 8.3) in a vertical slab gel apparatus at 80 V for
22-24 h in a cold room. The gels were then stained with silver,
and individual bands were quantified with a scanning densitometer.
Northern blot analysis of MHC isoform
mRNA. Total cellular RNA was isolated from frozen
hemidiaphragm samples with the guanidinium thiocyanate-phenol-chloroform extraction method (23). Total RNA was
quantitated and purity was assessed by measuring spectrophotometric absorption at 260 and 280 nm. Aliquots (10 µg) of RNA were size fractionated by electrophoresis on a 1% agarose-6.6% formaldehyde gel, transferred to nylon membranes (GeneScreen, Biotechnology Systems,
NEN Research Products, Boston, MA), and immobilized by ultraviolet
cross-linking. The membranes were prehybridized at 56°C overnight
in a solution containing 50% formamide, 5× saline-sodium citrate
(SSC; 1× SSC is 0.15 M NaCl and 0.015 M sodium citrate, pH 7.0),
0.5% SDS, 2.5× Denhardt's solution, 50 µg/ml of tRNA, and 100 µg/ml of salmon sperm DNA. The membranes were then hybridized overnight at the same temperature to
32P-labeled cRNA probes
(riboprobes) transcribed from MHC isoform-specific cDNA sequences
corresponding to portions of the 3'-untranslated region of each
major MHC isoform found in rat diaphragm; the sequences of the probes
used in this study have been reported earlier (9). The membranes were
washed in 2× SSC-1% SDS at 65°C followed by 0.1×
SSC-0.1% SDS and then exposed on a storage phosphor screen (Fuji Photo
Film, Stamford, CT) as well as on X-ray film with 2 intensifying
screens at 80°C. The same blots were alternately stripped
(confirmed by phosphorimager) and reprobed at separate times. Relative
amounts of MHC isoform mRNA were determined by phosphorimager (Fujix
Bio-Imaging Analyzer System, Bas 1000, Fuji Photo Film); the MHC mRNA
phosphorimager scores were normalized to 18S rRNA with a random
primer-labeled cDNA probe as previously described (32). The slow-twitch
soleus muscle was used as a positive control for type I MHC expression,
whereas the fast-twitch extensor digitorum longus muscle served this
role for the type II MHCs. Liver tissue RNA was also examined to
provide a negative control for sarcomeric MHC expression.
Statistical analysis. All data are reported as means ± SE and were analyzed with a Statistix 3.5 statistical-analysis program (Analytical Software, St. Paul, MN). Differences among the three experimental groups were initially tested by one-way ANOVA. For variables identified as significant by ANOVA, pairwise comparisons between group mean values were made with the least significant difference test. For MHC isoform mRNA and protein quantitation by gel electrophoresis, each individual MHC isoform measurement was first normalized to the mean CTL group value. Statistical significance was defined as P < 0.05.
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RESULTS |
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Diaphragm mass and sarcomere number. There was no difference in initial body weight among the three groups, which amounted to 157 ± 5, 157 ± 5, and 157 ± 9 g in the CTL, UNI-DNV, and BIL-DNV groups, respectively. A small decrease in final body weight was observed in the BIL-DNV (194 ± 16 g) compared with UNI-DNV (215 ± 13 g) and CTL (216 ± 8 g) groups (P < 0.05). Table 1 shows the effects of unilateral and bilateral hemidiaphragm inactivation on muscle mass and sarcomere number. Absolute hemidiaphragm weight increased by ~20 and 13% in the UNI-DNV and BIL-DNV groups, respectively. [It should be noted that in rats the left hemidiaphragm normally weighs ~10% less than the right hemidiaphragm (37).] When hemidiaphragm weight was normalized to total body weight, significant increases amounting to ~20 (UNI-DNV group) and 26% (BIL-DNV group) were also observed. Importantly, the changes were confined to the site of DNV (i.e., for UNI-DNV group, left hemidiaphragm only; for BIL-DNV group, left and right hemidiaphragms). As shown in Table 1, the diaphragm dry weight-to-wet weight ratio for the three experimental groups did not differ, indicating that greater hemidiaphragm mass in the denervated animals was not due to increased muscle fluid content. In addition, there was a significant increase in total RNA content per milligram of diaphragm tissue after both UNI-DNV and BIL-DNV.
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Analysis of single myofibers isolated from the left midcostal diaphragm in all groups revealed that the mean number of sarcomeres present along the fiber length was ~20% higher in the UNI-DNV (6,949 ± 219) and BIL-DNV (6,733 ± 180) groups compared with the CTL group (5,632 ± 186; P < 0.05). Therefore, the increase in total diaphragm mass after DNV appeared to be related, at least in part, to the addition of new sarcomeres in series. Mean sarcomere length did not differ between the CTL (2.87 ± 0.07 µm) and BIL-DNV (2.83 ± 0.03 µm) groups, whereas it was slightly decreased in the UNI-DNV group (2.61 ± 0.04 µm; P < 0.05).
MHC isoform immunohistochemistry. Figure 1 shows serial diaphragm sections reacted with antibodies against type I and type II MHC isoforms. As can be seen from the micrographs, the UNI-DNV and BIL-DNV sections both exhibited a substantial proportion of hybrid fibers (indicated by numbered fibers) that coexpressed both type I and type II MHCs, whereas such hybrid fibers were rarely observed in CTL diaphragms. The relative proportions of type I, type II, and hybrid fibers for all animals are depicted in Fig. 2. After either unilateral or bilateral diaphragm inactivation, there was no change in the proportion of myofibers expressing type I MHC in an exclusive fashion. However, the relative proportion of hybrid fibers with type I-type II coexpression was markedly increased in the UNI-DNV and BIL-DNV groups compared with the CTL group. Therefore, this phenotypic transformation occurred at the expense of myofibers that had previously expressed type II MHCs exclusively. The data would thus appear to indicate that in the two DNV groups a fast-to-slow MHC isoform transformation occurred within individual type II diaphragm myofibers.
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As shown in Table 2, there was also significant hypertrophy of type I MHC fibers in the UNI-DNV group compared with the CTL group, with a similar tendency being present in the BIL-DNV group. No significant differences in mean fiber cross-sectional area for the type II or hybrid fiber populations were observed among the three groups of animals. By combining the data obtained for fiber-type proportions and mean fiber cross-sectional area, the relative contributions of type I, type II, and hybrid fibers to the total cross-sectional area of the diaphragm were also calculated. With respect to myofibers that solely expressed type I MHC, there was an increased contribution to overall diaphragm cross-sectional area in the UNI-DNV and BIL-DNV groups compared with the CTL group, although this did not achieve statistical significance. The relative contribution to total diaphragm cross-sectional area of the hybrid fibers was significantly elevated in both the UNI-DNV and BIL-DNV groups compared with the CTL group. Accordingly, the total area contribution of myofibers that solely expressed type II MHCs fell significantly in the two DNV groups compared with the CTL group.
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Electrophoretic quantitation of MHC isoforms. Figure 3 shows a representative example of electrophoretic separation of MHC isoforms in diaphragm samples from the three experimental groups as well as in control hindlimb muscles. Because the separation between type IIa and type IIx MHC bands was not distinct in all cases, these two isoforms with similar contractile characteristics (3) were considered together in the overall quantitative analysis. As can be seen, the order of migration for MHC isoforms was type I > type IIb > type IIa/IIx. Figure 3 indicates that in the two DNV samples there were reciprocal changes in the relative contributions of type I (increased) and type IIb (decreased) MHCs. The group mean data for MHC isoform quantitation are depicted in Fig. 4, with each MHC isoform expressed as a percentage of the mean CTL value. In keeping with the immunohistochemical findings, there was a significant increase in type I MHC expression after both UNI-DNV and BIL-DNV that occurred primarily at the expense of the type IIb MHC isoform.
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Diaphragm myofiber contractile function. To determine whether the above type II-to-type I switch in MHC isoform expression was of sufficient magnitude to induce changes in the physiological function of diaphragm myofibers, in vitro contractile properties were assessed and are shown in Table 3. In keeping with the increased type I MHC and reduced type IIb MHC expression noted by immunohistochemical and/or electrophoretic analyses, the UNI-DNV and BIL-DNV groups demonstrated significantly greater fatigue resistance than the CTL group. Furthermore, there was a prolongation of contraction time and half-relaxation time in the UNI-DNV and BIL-DNV groups compared with the CTL group, which is also consistent with the shift toward a slower MHC phenotype after DNV. Maximal twitch force as well as maximal tetanic force was reduced in the UNI-DNV and BIL-DNV groups. In addition, Lo was found to be significantly greater in the UNI-DNV (18.4 ± 0.6 mm) and BIL-DNV (18.6 ± 0.6 mm) groups compared with the CTL group (16.3 ± 0.3 mm; P < 0.05), a finding that is consistent with the previously mentioned increase in sarcomere number in the two DNV groups.
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MHC isoform mRNA levels. Figure 5 shows representative lanes from Northern blot analysis of RNA prepared from CTL, UNI-DNV, and BIL-DNV diaphragms. As expected, each of the sarcomeric MHC probes recognized a distinct 6-kb mRNA in muscle tissue, whereas no signal was detected in RNA extracted from liver (data not shown). Group mean data are depicted in Fig. 6, with MHC mRNA signals being normalized to 18S rRNA and expressed as a percentage of the mean CTL value obtained for each individual MHC isoform. Despite the evidence for increased type I MHC at the protein level, normalized type I MHC mRNA levels were significantly reduced in both DNV groups compared with the CTL group. In addition, type IIa MHC and type IIx MHC mRNA levels were also markedly decreased in the two DNV groups. For type IIa MHC, the DNV-induced reduction in mRNA levels was greater in the UNI-DNV group than in the BIL-DNV group, whereas no such difference between DNV groups was found for either type I MHC or type IIx MHC mRNA levels. Type IIb MHC mRNA expression in the diaphragm was not significantly altered by DNV.
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DISCUSSION |
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The response to diaphragm DNV is a dynamic process that leads to substantially increased rates of protein synthesis (35), with attendant muscle remodeling. Myosin represents the major component of skeletal muscle protein, and differences in the economy of force generation are largely determined by the relative contributions of specialized MHC isoforms (3, 8). Studies in rat hindlimb muscle have generally indicated that continuous innervation is required for the development as well as for the maintenance of a slow type I MHC phenotype (1, 4, 13, 17, 29). Prior studies of long-term (>4-wk) rat diaphragm DNV are also consistent with this proposition (6, 7, 14). In the present study, by contrast, we observed a change in the pattern of MHC isoform expression from a type II (fast) to type I (slow) profile within ~1 wk of diaphragm DNV. Our data indicate that this occurred not simply through the preferential hypertrophy of type I fibers previously described after diaphragm DNV (38, 40) but also by the apparent transformation of type II fibers toward the type I MHC phenotype. Therefore, the alterations in type I MHC gene expression within individual diaphragm myofibers were of both a quantitative (hypertrophy) and qualitative (coexpression with type II MHCs) nature.
The predominant view to date has been that switches in MHC isoform expression of this nature are regulated primarily at a pretranslational level in skeletal muscle (15, 33). For example, during chronic low-frequency electrical stimulation of rat tibialis anterior muscle, alterations in MHC isoform mRNA levels correlate well with changes in protein synthesis within 2 days after the onset of stimulation (33). However, there have been reports of discrepancies between alterations in MHC isoform mRNA and protein levels in situations associated with muscle atrophy such as hindlimb unweighting and spaceflight (5, 34), suggesting that posttranscriptional (i.e., translational and/or posttranslational) events also play an important role under these conditions. In contradistinction to the above, to our knowledge, there have been no previous demonstrations of antithetical changes in MHC isoform mRNA and protein in the setting of muscle hypertrophy.
We provide the first evidence that the differential expression of MHC isoforms in the setting of diaphragm hypertrophy induced by DNV involves posttranscriptional regulatory mechanisms. In this regard, an increased contribution of type I MHC protein was found despite apparent downregulation at the mRNA level. However, it should be noted that although type I MHC mRNA normalized to rRNA was significantly reduced after diaphragm inactivation, absolute levels of type I MHC mRNA could have been relatively well maintained due to the observed increase in total RNA concentration. This is provided, of course, that the increase in total RNA reflected events in muscle cells as opposed to those occurring primarily in other cell types such as fibroblasts and capillary endothelium. If one also assumes a constant ratio of rRNA to total RNA for all experimental groups, absolute levels of type I MHC mRNA would have represented ~85 and 125% of the CTL values in the UNI-DNV and BIL-DNV groups, respectively. On the other hand, a strong trend toward decreased type IIb MHC protein occurred in the absence of corresponding reductions in either normalized or absolute mRNA transcript levels. In fact, with the use of the same assumptions outlined above, the absolute levels of type IIb MHC mRNA would have moved in the opposite direction, amounting to ~120 and 160% of the CTL values for the UNI-DNV and BIL-DNV groups, respectively. This makes it unlikely that the disparities between MHC isoform mRNA and corresponding protein can be explained by a simple phase lag between cellular events occurring at these two levels. Rather, the data suggest that points of control other than MHC gene transcription and/or mRNA stability play a major role in the regulation of MHC isoform expression after short-term diaphragm inactivation.
In principle, posttranscriptional control of MHC isoform expression could be achieved at a number of different levels. First, it is conceivable that the cytosolic fraction (i.e., the "effective" mRNA utilized for translation into protein) could be altered through changes in posttranscriptional transport out of the nucleus. In this regard, increased nucleocytoplasmic transport of type I MHC mRNA out of normal cardiomyocyte myonuclei has been reported to occur in response to undefined cytosolic factors obtained from hypertrophic cardiomyocytes (24). A second possibility is that translational efficiency (i.e., the rate of translation per ribosome) and/or capacity (i.e., ribosomal content) might be altered after diaphragm DNV. Indeed, an increase in translational capacity is suggested by the increase in overall RNA concentration observed after DNV. However, although this explanation could partly clarify the relative upregulation of type I MHC protein seen after diaphragm inactivation, it cannot account for the directionally opposite pattern of changes in type IIb MHC mRNA and protein expression. Finally, because the rate of protein degradation is substantially elevated in the denervated diaphragm despite ongoing hypertrophy (35), a third and perhaps most likely explanation is that differential degradation rates could exist for each of the MHC isoforms, thereby exerting an important degree of posttranslational control over MHC transitions.
A particularly interesting aspect of the diaphragm remodeling associated with short-term DNV was the presence of sarcomere addition, which provides strong, albeit indirect, evidence for diaphragm stretch under these conditions (36). Previous work in rabbit limb muscle has demonstrated significant addition of new sarcomeres to myofibers after only 4 days of stretch (36). However, the fact that sarcomere addition was observed after bilateral as well as unilateral phrenicotomy suggests that the traditional idea of contralateral hemidiaphragm activation being the primary source of stretch in this setting is incorrect. Additional potential sources of diaphragm stretch include the actions of other intact accessory muscle groups, recruited as a means of aiding ventilation (20, 21), on the paralyzed and flaccid diaphragm. In this regard, persistent activity of other inspiratory muscles (e.g., intercostals) and consequent negative excursions in intrathoracic pressure, as well as expiratory recruitment of abdominal muscles, would tend to move the diaphragm cephalad and thereby lengthen the muscle (20, 21). Moreover, to the extent that functional residual capacity has also been reported to be decreased in the setting of diaphragm paralysis (30), passive changes in resting diaphragm length might provide a further stimulus for alterations in sarcomere number, as has been found to be the case in emphysema (31). The fact that diaphragm stretch was minimal to absent by sonomicrometry in acutely anesthetized rabbits with UNI-DNV (38) may relate to certain methodological issues. First, because anesthesia has been shown to inhibit the activation of both intercostal (11) and abdominal (19) muscles, sonomicrometry data obtained in the anesthetized state would tend to underestimate these factors. Second, sonomicrometry measurements in the above study (38) focused on phasic rather than potential tonic or resting changes in diaphragm muscle length that would be caused by an alteration in functional residual capacity.
Because of their inherent complexities, the diaphragm DNV models examined in this study do not allow precise dissection of the relative influences on MHC isoform expression of DNV per se versus stretch. It should also be noted that because the animals studied here were relatively young and in a period of rapid growth, effects of the above stimuli on the denervated diaphragm could differ in other age groups. DNV of rat hindlimb muscle shortly after birth interferes with development of the slow type I phenotype, whereas the fast type II program of development remains largely unaffected (4, 13). In adult rat hindlimb muscles, Jakubiec-Puka et al. (17) recently reported a shift toward greater relative expression of the more intermediate type II MHC isoforms (i.e., IIa and IIx), which was evident by 14 days post-DNV. This occurred at the expense of type I and type IIb fiber-type populations in the slow-twitch soleus and fast-twitch extensor digitorum longus, respectively (17). Therefore, these authors speculated that in mature rat muscle, maintenance of both the slowest (type I) and fastest (type IIb) MHC phenotypes is dependent on intact innervation (17). Similar findings have been reported by other investigators (1, 29). Muscle stretch, on the other hand, has been reported to cause a general repression of type II MHC gene expression as well as induction of the type I MHC gene both in vivo (15) and in vitro (26).
The pattern of reciprocal changes in type I-type IIb MHC protein observed after both UNI-DNV and BIL-DNV appears most consistent with previously described stretch effects (15, 26). Zhan et al. (39) recently used immunohistochemistry to examine shifts in MHC expression 2 wk after hemidiaphragm UNI-DNV and also found qualitatively similar changes. On the other hand, prior studies demonstrating type II MHC predominance after longer term (>4-wk) diaphragm DNV seem more in keeping with reports in denervated hindlimb muscles (6, 7, 14). One potential explanation for these findings is that the level of passive tension on the denervated diaphragm may be gradually reduced as new sarcomeres are added to its myofibers, thus creating a situation whereby the effects of DNV alone become increasingly dominant over time. Alternatively, it is also possible that alterations in MHC isoform expression after diaphragm DNV follow a biphasic pattern due to factors completely unrelated to muscle stretch. For example, mammalian skeletal muscles (including the diaphragm) demonstrate a form of spontaneous muscle activity after DNV referred to as "fibrillation," which is believed to be caused by enhanced sodium conductance (28). Because changes in the magnitude and/or pattern of external electrical stimulation of muscle are known to have differential effects on MHC isoform expression (1), time-dependent changes in such spontaneous muscle activity could also conceivably underlie the biphasic response of MHC isoforms to diaphragm DNV.
It is noteworthy that, in the clinical setting, neuromuscular blocking agents are often employed to induce paralysis of respiratory muscles in patients undergoing mechanical ventilation. This practice may essentially lead to a state of reversible, short-term "chemical" DNV of the diaphragm (2). However, in contrast to the presumed stretching of paralyzed diaphragm myofibers during spontaneous breathing, the diaphragm should be passively shortened with each breath delivered by the ventilator. Therefore, this circumstance should in theory allow the effects of diaphragm DNV and stretch to be dissociated from one another. The extent and nature of diaphragm remodeling that may result from the imposition of diaphragm paralysis during mechanical ventilation are currently unknown. If alterations in MHC isoform expression similar to those observed in this study do occur, this would strongly suggest that removal of normal innervation rather than stretch is the primary stimulus for these changes. Furthermore, this could have important clinical implications because a shift toward greater type I MHC expression, although potentially beneficial from the standpoint of enhancing fatigue resistance, would also reduce the range of shortening velocities and power output available to the diaphragm. In addition, the results of our study suggest that changes in sarcomere number caused by altered diaphragm length take place over a very short time period after the onset of diaphragm paralysis. These changes might actually be deleterious in the clinical setting because alterations in sarcomere number induced by diaphragm paralysis and mechanical ventilation could be inappropriate for the prevailing diaphragm length encountered during spontaneous breathing. Under these conditions, the changes in diaphragm sarcomere number would be maladaptive and further contribute to the already reduced capacity for power output, thus impeding the ability to discontinue ventilatory support. The results of the present investigation provide a conceptual framework for testing the above hypotheses and related issues in future studies.
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ACKNOWLEDGEMENTS |
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We thank Drs. Alan Kelly and Neal Rubinstein for the use of antibody NOQ7.5.4D and Dr. Stefano Schiaffino for the myosin heavy chain cDNAs employed in this study. We also thank Neola Matusiewicz for expert technical assistance.
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FOOTNOTES |
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This investigation was supported by a Clinician-Scientist Award from the Medical Research Council of Canada (to B. J. Petrof), a Medical Research Scholar Award from the Fonds de la Recherche en Santé du Québec (to S. B. Gottfried), a Postdoctoral Fellowship from the Canadian Lung Association (to L. Yang), and a grant from the Association Pulmonaire du Quebec.
Address for reprint requests: B. J. Petrof, Respiratory Division, Rm. L411, Royal Victoria Hospital, 687 Pine Ave. West, Montreal, Quebec, Canada H3A 1A1.
Received 21 March 1997; accepted in final form 13 March 1998.
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REFERENCES |
---|
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---|
1.
Ausoni, S.,
L. Gorza,
S. Schiaffino,
K. Gundersen,
and
T. Lomo.
Expression of myosin heavy chain isoforms in stimulated fast and slow rat muscles.
J. Neurosci.
10:
153-160,
1990[Abstract].
2.
Berg, D. K.,
and
Z. W. Hall.
Increased extrajunctional acetylcholine sensitivity produced by chronic acetylcholine sensitivity produced by chronic post-synaptic neuromuscular blockade.
J. Physiol. (Lond.)
244:
659-676,
1975[Abstract].
3.
Bottinelli, R.,
S. Schiaffino,
and
C. Reggiani.
Force-velocity relations and myosin heavy chain isoform compositions of skinned fibres from rat skeletal muscle.
J. Physiol. (Lond.)
437:
655-672,
1991[Abstract].
4.
Butler-Browne, G. S.,
L. B. Bugaisky,
S. Cuenoud,
K. Schwartz,
and
R. G. Whalen.
Denervation of newborn rat muscles does not block the appearance of adult fast myosin heavy chain.
Nature
299:
830-833,
1982[Medline].
5.
Caiozzo, V. J.,
F. Haddad,
M. J. Baker,
R. E. Herrick,
N. Prietto,
and
K. M. Baldwin.
Microgravity-induced transformation of myosin isoforms and contractile properties of skeletal muscle.
J. Appl. Physiol.
81:
123-132,
1996
6.
Carraro, U.,
L. Dalla Libera,
C. Catani,
and
D. Danieli-Betto.
Chronic denervation of rat diaphragm: selective maintenance of adult fast myosin heavy chains.
Muscle Nerve
5:
515-524,
1982[Medline].
7.
Carraro, U.,
D. Morale,
I. Mussini,
S. Lucke,
M. Cantini,
R. Betto,
C. Catani,
L. D. Libera,
D. D. Betto,
and
D. Noventa.
Chronic denervation of rat hemidiaphragm: maintenance of fiber heterogeneity with associated increasing uniformity of myosin isoforms.
J. Cell Biol.
100:
161-174,
1985[Abstract].
8.
Crow, M. T.,
and
M. J. Kushmerick.
Chemical energetics of slow- and fast-twitch muscles of the mouse.
J. Gen. Physiol.
79:
147-166,
1982[Abstract].
9.
DeNardi, C.,
S. Ausoni,
P. Moretti,
L. Gorza,
M. Velleca,
M. Buckingham,
and
S. Schiaffino.
Type 2X-myosin heavy chain is coded by a muscle fiber type-specific and developmentally regulated gene.
J. Cell Biol.
123:
823-835,
1993[Abstract].
10.
DiMario, J. X.,
S. E. Fernyak,
and
F. E. Stockdale.
Myoblasts transferred to the limbs of embryos are committed to specific fibre fates.
Nature
362:
165-167,
1993[Medline].
11.
Drummond, G. B.
Reduction of tonic ribcage muscle activity by anesthesia with thiopental.
Anesthesiology
67:
695-700,
1987[Medline].
12.
Feng, T. P.,
and
D. X. Lu.
New lights on the phenomenon of transient hypertrophy in the denervated hemidiaphragm of the rat.
Sci. Sin.
14:
1772-1784,
1965[Medline].
13.
Gambke, B.,
G. E. Lyons,
J. Haselgrove,
A. M. Kelly,
and
N. A. Rubinstein.
Thyroidal and neural control of myosin transitions during development of rat fast and slow muscles.
FEBS Lett.
156:
335-339,
1983[Medline].
14.
Gauthier, G. F.,
and
A. W. Hobbs.
Effects of denervation on the distribution of myosin isozymes in skeletal muscle fibers.
Biochem. Biophys. Res. Commun.
76:
331-347,
1982.
15.
Goldspink, G.,
A. Scutt,
P. T. Loughna,
D. J. Wells,
T. Jaenicke,
and
G. F. Gerlach.
Gene expression in skeletal muscle in response to stretch and force generation.
Am. J. Physiol.
262 (Regulatory Integrative Comp. Physiol. 31):
R356-R363,
1992
16.
Gutmann, E.,
S. Schiaffino,
and
V. Hanzlikova.
Mechanism of compensatory hypertrophy in skeletal muscle of the rat.
Exp. Neurol.
31:
451-464,
1971[Medline].
17.
Jakubiec-Puka, A.,
J. Kordowska,
C. Catani,
and
U. Carraro.
Myosin heavy chain isoform composition in striated muscle after denervation and self-reinervation.
Eur. J. Biochem.
193:
623-628,
1990[Abstract].
18.
Laframboise, W. A.,
J. F. Watchko,
B. S. Brozanski,
M. J. Daood,
and
R. D. Guthrie.
Myosin heavy chain expression in respiratory muscles of the rat.
Am. J. Respir. Cell Mol. Biol.
6:
335-339,
1992[Medline].
19.
Leevers, A. M.,
and
J. D. Road.
The effect of anesthesia on abdominal muscle resting length and shortening in awake dogs.
Lung
173:
105-115,
1995[Medline].
20.
Lisboa, C.,
C. D. Pare,
G. Pertuze,
R. Contreras,
R. Moreno,
S. Guillemi,
and
E. Cruz.
Inspiratory muscle function in unilateral diaphragmatic paralysis.
Am. Rev. Respir. Dis.
134:
488-492,
1986[Medline].
21.
Newsom Davis, J.,
M. Goldman,
L. Loh,
and
M. Casson.
Diaphragm function and alveolar hypoventilation.
Q. J. Med.
45:
87-100,
1976[Medline].
22.
Petrof, B. J.,
S. B. Gottfried,
J. Eby,
J. LaManca,
and
S. Levine.
Growth hormone does not prevent corticosteroid-induced changes in rat diaphragm structure and function.
J. Appl. Physiol.
79:
1571-1577,
1995
23.
Petrof, B. J.,
A. M. Kelly,
N. A. Rubinstein,
and
A. I. Pack.
Effect of hypothyroidism on myosin heavy chain expression in rat pharyngeal dilator muscles.
J. Appl. Physiol.
73:
179-187,
1992
24.
Rajamanickam, C.,
N. Selvamurugan,
S. Arun,
and
M. A. Q. Siddiqui.
Effect of cytosol on the regulation of expression of myosin heavy chain genes during cardiac hypertrophy.
Cell. Mol. Biol.
38:
81-89,
1992[Medline].
25.
Robbins, N.,
A. Olek,
S. S. Kelly,
P. Takach,
and
M. Christopher.
Quantitative study of motor endplates in muscle fibres dissociated by a simple procedure.
Proc. R. Soc. Lond. B Biol. Sci.
209:
555-562,
1980[Medline].
26.
Shyu, K. G.,
J. J. Chen,
N. L. Shih,
D. L. Wang,
H. Chang,
W. P. Lien,
and
C. C. Liew.
Regulation of human cardiac myosin heavy chain genes by cyclical mechanical stretch in cultured cardiocytes.
Biochem. Biophys. Res. Commun.
210:
567-573,
1995[Medline].
27.
Sieck, G. C.
Physiological effects of diaphragm muscle denervation and disuse.
Clin. Chest Med.
15:
641-659,
1994[Medline].
28.
Smith, J. W.,
and
S. Thesleff.
Spontaneous activity in denervated mouse diaphragm muscle.
J. Physiol. (Lond.)
257:
171-186,
1976[Abstract].
29.
Spector, S. A.
Effects of elimination of activity on contractile and histochemical properties of rat soleus muscle.
J. Neurosci.
5:
2177-2188,
1985[Abstract].
30.
Spitzer, S. A.,
A. D. Korczyn,
and
J. Kalaci.
Transient bilateral diaphragmatic paralysis.
Chest
64:
355-357,
1973[Medline].
31.
Supinski, G. S.,
and
S. G. Kelsen.
Effect of elastase-induced emphysema on the force-generating ability of the diaphragm.
J. Clin. Invest.
70:
978-988,
1982[Medline].
32.
Szyf, M.,
D. S. Milstone,
B. P. Schimmer,
K. L. Parker,
and
J. G. Seidman.
Cis modification of the steroid 21-hydroxylase gene prevents its expression in the Y1 mouse adrenocortical tumor cell line.
Mol. Endocrinol.
90:
1144-1152,
1990.
33.
Termin, A.,
and
D. Pette.
Changes in myosin heavy-chain isoform synthesis of chronically stimulated rat fast-twitch muscle.
Eur. J. Biochem.
204:
569-573,
1992[Abstract].
34.
Thomason, D. B.,
and
F. W. Booth.
Atrophy of the soleus muscle by hindlimb unweighting.
J. Appl. Physiol.
68:
1-12,
1990
35.
Turner, L. V.,
and
P. J. Garlick.
The effect of unilateral phrenicectomy on the rate of protein synthesis in rat diaphragm in vivo.
Biochim. Biophys. Acta
349:
109-113,
1974[Medline].
36.
Williams, P.,
P. Watt,
V. Bicik,
and
G. Goldspink.
Effect of stretch combined with electrical stimulation on the type of sarcomeres produced at the ends of muscle fibers.
Biochem. Biophys. Res. Commun.
93:
500-509,
1986.
37.
Zak, R.,
D. Grove,
and
M. Rabinowitz.
DNA synthesis in the rat diaphragm as an early response to denervation.
Am. J. Physiol.
216:
647-654,
1969[Medline].
38.
Zhan, W.-Z.,
G. A. Farkas,
M. A. Schroedeer,
L. E. Gosselin,
and
G. C. Sieck.
Regional adaptations of rabbit diaphragm muscle fibers to unilateral denervation.
J. Appl. Physiol.
79:
941-950,
1995
39.
Zhan, W.-Z.,
H. Miyata,
Y. S. Prakash,
and
G. C. Sieck.
Metabolic and phenotypic adaptations of diaphragm muscle fibers with inactivation.
J. Appl. Physiol.
82:
1145-1153,
1997
40.
Zhan, W. Z.,
and
G. C. Sieck.
Adaptations of diaphragm and medial gastrocnemius muscles to inactivity.
J. Appl. Physiol.
72:
1445-1453,
1992