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. 2003 Dec 12;554(Pt 3):891–903. doi: 10.1113/jphysiol.2003.051318

Respiratory muscle injury, fatigue and serum skeletal troponin I in rat

Jeremy A Simpson 1, Jennifer Van Eyk 1, Steve Iscoe 1
PMCID: PMC1664786  PMID: 14673191

Abstract

To evaluate injury to respiratory muscles of rats breathing against an inspiratory resistive load, we measured the release into blood of a myofilament protein, skeletal troponin I (sTnI), and related this release to the time course of changes in arterial blood gases, respiratory drive (phrenic activity), and pressure generation. After ∼1.5 h of loading, hypercapnic ventilatory failure occurred, coincident with a decrease in the ratio of transdiaphragmatic pressure to integrated phrenic activity (Pdi/∫Phr) during sighs. This was followed at ∼1.9 h by a decrease in the Pdi/∫Phr ratio during normal loaded breaths (diaphragmatic fatigue). Loading was terminated at pump failure (a decline of Pdi to half of steady-state loaded values), ∼2.4 h after load onset. During 30 s occlusions post loading, rats generated pressure profiles similar to those during occlusions before loading, with comparable blood gases, but at a higher neural drive. In a second series of rats, we tested for sTnI release using Western blot–direct serum analysis of blood samples taken before and during loading to pump failure. We detected only the fast isoform of sTnI, release beginning midway through loading. Differential detection with various monoclonal antibodies indicated the presence of modified forms of fast sTnI. The release of fast sTnI is consistent with load-induced injury of fast glycolytic fibres of inspiratory muscles, probably the diaphragm. Characterization of released fast sTnI may provide insights into the molecular basis of respiratory muscle dysfunction; fast sTnI may also prove useful as a marker of impending respiratory muscle fatigue.

Research on respiratory muscle (diaphragmatic) fatigue and failure has been devoted primarily to two main aspects: determination of their underlying molecular basis (or bases), and detection and quantification of fatigue. In animals, loading of the inspiratory muscles with, for example, an inspiratory resistance, reduces ventilation (hypercapnic ventilatory failure) and causes eventual pump failure (sometimes called task failure), defined as apnoea or an inability to generate a target pressure. Acute increases in arterial or alveolar PCO2 are a sign of ventilatory failure but do not indicate if the failure is of central (reduced drive) or peripheral (impaired contractility) origin, or some combination.

Experiments using either acute (Reid et al. 1994; Jiang et al. 1998a, 1998b, 2001) or chronic (Zhu et al. 1997; Reid & Belcastro,1999, 2000) loads have been supplemented only occasionally by studies of muscle injury (for reviews, see Reid & MacGowan, 1998; Road & Jiang, 1998). Because histological assessment of diaphragmatic injury in man is possible only under limited circumstances (MacGowan et al. 2001; Orozco-Levi et al. 2001), other techniques must be used to determine if contractile dysfunction, including fatigue, is present. This, however, cannot be done just by measuring the subject's maximal inspiratory pressure because such measurements can include components (decreased volitional output, inhibition of motoneurones, neurotransmission failure) unrelated to muscle contractility (for reviews, see Gandevia et al. 1998; Gandevia, 2001). Moreover, this measurement is restricted to co-operative subjects. To avoid this problem, the transdiaphragmatic twitch pressure during bilateral phrenic stimulation is now considered to be ‘the best method to detect diaphragmatic fatigue in humans’ (Mador et al. 2002). It nevertheless remains too complicated for general use, despite its introduction in 1981 by Aubier et al. (1981a) and recommendation in 1990 by a workshop group of the National Heart, Lung and Blood Institute. As Sliwinski and Macklem stated in 1997: ‘Although fatigue of the inspiratory muscles has been well documented, its prevalence in patients……[is] unknown, because of the lack of a simple, clinically available diagnostic test’ (Sliwinski & Macklem, 1997). This situation has not changed since. Finally, measurement of diaphragmatic twitch pressure detects fatigue only after it occurs and if a reference value is available; it therefore has limited prognostic value.

A paradigm for evaluating respiratory muscle status is available from the current practice of assessing patients with suspected myocardial infarction by measuring their serum levels of the cardiac isoforms of the myofilament proteins troponin I and T. There are three isoforms of troponin I, cardiac, fast skeletal and slow skeletal, found in cardiac, fast glycolytic and slow oxidative fibres, respectively. Cardiac troponin I (cTnI) is released from cardiac myocytes into the blood with injury and is the current gold standard for the diagnosis and management of patients with myocardial injury (Collinson, 2001; Ni, 2001; Apple et al. 2002). The few studies in which skeletal TnI (sTnI) has been evaluated as a marker of skeletal muscle damage used an indirect method, immunoassay, to measure sTnI; they indicate that it is more sensitive and specific than traditional markers (creatine kinase and myoglobin) of skeletal muscle injury (Sorichter et al. 1997; Onuoha et al. 2001). In this study, we used isoform-specific monoclonal antibodies for direct detection of fast and slow sTnI released into blood; their presence would serve as an indicator of load-induced injury of the respiratory muscles (primarily the diaphragm). Anaesthetized rats breathing against an inspiratory resistive load (IRL) consistently developed hypercapnic ventilatory failure followed by diaphragmatic fatigue, followed by pump failure. We detected only the fast isoform of sTnI in the blood, a response consistent with injury to fast fibres of respiratory (diaphragmatic) muscles. sTnI may prove useful in detecting and/or determining the role of injury in the development of respiratory muscle dysfunction.

Methods

Experiments, approved by the Animal Care Committee of Queen's University and in conformity with the guidelines of the Canadian Council on Animal Care, were conducted on pentobarbital-anaesthetized (65 mg kg−1i.p., supplemented as required to prevent a pedal reflex) Sprague Dawley rats (300–460 g). Atropine sulphate (0.05 mg kg−1, subcutaneous) was administered to reduce tracheal secretions. Once a surgical plane of anaesthesia was established, the rat was placed supine; body temperature was maintained at ∼37.5°C with a servo-controlled heating pad.

Surgical procedures

After making an incision in the midline of the neck, the sternohyoid muscles were excised. A tracheal cannula was inserted; one port was connected to a pressure transducer to measure tracheal pressure (Ptr). The right carotid artery (for measuring blood pressure and sampling arterial blood gases; Radiometer ABL-5, Copenhagen, Denmark) and jugular vein were cannulated. The left phrenic nerve (all 11 rats of series 1 and 1 of 5 rats of series 2) was isolated from underlying tissue with parafilm, placed over a bipolar silver hook electrode, and the nerve–electrode assembly then covered with low melting point paraffin wax. Phrenic nerve activity was amplified and filtered (100–10 000 Hz; Grass P-511, Quincy, MA, USA) and integrated (Paynter filter, time constant 50 ms). We recorded transdiaphragmatic pressure (Pdi) as the difference in pressures measured in the stomach (Pab) and oesophagus (Pes) (Millar SPR 524, Houston, TX, USA). All signals (blood pressure, integrated phrenic activity (∫Phr), and Ptr, Pes, Pab, and Pdi) were acquired (CED Spike2, Cambridge, UK) to computer.

Inspiratory resistive loading (IRL)

A two-way valve (Hans Rudolf 2300, Kansas City, MO, USA) was attached to the other port of the tracheal cannula; a short segment of rubber tubing on which a small clamp was placed was attached to the inspiratory side of this valve. IRL was applied by gradually tightening the clamp over 15 min until the rat generated a tidal Ptr∼60% (i.e. about –20 cmH2O) of the peak Ptr obtained during a previous 30 s occlusion; loading was considered to have started (normalized time = 0) at the end of this 15 min period. (In preliminary experiments, too rapid application of the ‘final’ IRL resulted in respiratory arrest within 10 min.) Blood samples were taken before loading and at 30 min intervals thereafter. IRL was discontinued when Ptr declined to ∼–10 cmH2O (pump failure; normalized time = 1), characterized by a rapid fall in pressure generation and respiratory frequency. After 15 min, a second 30 s occlusion was performed. Rats were then killed with an overdose of pentobarbital.

Serum analysis

Frozen (series 1) and fresh (series 2) samples were analysed for sTnI using Western blot–direct serum analysis (WB-DSA) as previously described (Labugger et al. 2000; Simpson et al. 2002). Anti-TnI monoclonal antibodies (mAb) of confirmed isoform specificity were chosen for WB-DSA: sTnI, FI-32 and FI-23 (fast only, Spectral Diagnostics, Toronto, ON, Canada); MYNT-S (preferential for slow in rat, courtesy of N. Matsumoto (Matsumoto et al. 1997)); and 3I-35 (fast, slow, and cardiac; Spectral Diagnostics). Specificity of all antibodies was confirmed by Western blot analysis of cardiac and skeletal tissue from human and rat as previously described (Simpson et al. 2000).

Statistics

Raw, not normalized, data were compared using paired t tests or repeated measures analysis of variance with post hoc analysis as indicated.

Results

Characterization of IRL model: assessment of hypercapnic ventilator failure, diaphragmatic fatigue, and pump failure

Figure 1 illustrates representative changes in Pdi, Pab, Pes, Ptr and ∫Phr during IRL in a rat of series 1. Extracts (a, control; b, load onset; c, midway; d, during pump failure) of the tracing are shown on an expanded time scale in the lower panels. Pdi in all rats was due almost entirely to changes in Pes. Pump failure in all rats occurred after an average of 2.4 h (range 1–4.7 h).

Figure 1. Effects of inspiratory resistive loading on (traces from top down): transdiaphragmatic (Pdi), abdominal (Pab), oesophageal (Pes) and tracheal (Ptr) pressures and integrated phrenic activity (∫Phr) in a representative rat.

Figure 1

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Arrows indicate 30 s occlusions before and after loading. Selected time points (a, control; b, load onset; c, midway during loading; d, during pump failure) are shown on expanded time scales to show individual breaths. Large ‘spikes’ throughout loading are sighs, best observed in expanded traces b and c.

In order to adjust for the variability in IRL duration, we normalized load duration: ‘c’ is preload control, 0 is load onset and 1 is termination of IRL, i.e. pump failure, for each rat. The absolute and normalized temporal profiles of changes in blood gases are shown in Fig. 2A and B. IRL onset was associated with an immediate decrease in PaO2 which fell throughout IRL; PaCO2 did not increase until ∼60% (time = 0.6) of IRL duration (hypercapnic ventilatory failure) (Fig. 2). PaO2 and PaCO2, just before pump failure, were 39.6 ± 7.9 (s.d.) and 59.9 ± 18.5 mmHg, respectively.

Figure 2. Arterial blood gases versus time (A) and normalized time (B) during inspiratory resistive loading.

Figure 2

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A, plots of arterial PO2 (PaO2) and PCO2 (PaCO2) as a function of time (C is preload control, R is 15 min post load) in all rats. Numbers above abscissa indicate number of rats breathing against load at that time. B, blood gases versus normalized time, 1 = termination of loading. Arrows indicate onset of hypercapnic failure and pump failure. * Significantly different from control (P < 0.001, repeated measures ANOVA). Data points and error bars are means ±s.d.

During IRL, respiratory frequency (Fig. 3A), inspiratory duration, and duty cycle (inspiratory duration/total breath duration, TI/TTOT, Fig. 3B) remained at control levels until abruptly declining at pump failure (time = 1). In contrast, diaphragmatic contractility, indicated by Pdi/∫Phr (pressure output/neural drive, during normal loaded breaths), fell at time = 0.8 below the value at time = 0.1 (Fig. 4A), preceding pump failure by an average 29.1 min (range 18–47 min). This fall in Pdi/∫Phr (indicating diaphragmatic fatigue) was due to contractile impairment (peripheral fatigue) and not decreased drive (i.e. central fatigue) because, on average, the rate of increase in phrenic activity during inspiration (∫Phr/TI) increased throughout loading (Fig. 4B).

Figure 3. Respiratory frequency and respiratory duty cycle (TI/TTOT) versus normalized time (1 = termination of load) during inspiratory resistive loading.

Figure 3

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Arrow indicates pump failure. * Significantly different from control (P < 0.001, repeated measures ANOVA). Data points and error bars are means ±s.e.m..

Figure 4. Diaphragmatic contractility (Pdi/∫Phr) and respiratory drive (∫Phr/TI) versus normalized time for all rats.

Figure 4

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Pressure output declined towards the end of loading despite increased drive. Arrows indicate onset of diaphragmatic fatigue and pump failure. * Significantly different from time = 0.1 (P < 0.001, repeated measures ANOVA). Data points and error bars are means ±s.e.m.

Sighs during IRL were evident as large ‘spikes’ of ∫Phr, Pdi, Pes and Ptr (see extracts b and c in Fig. 1) and represent a reflex attempt to improve gas exchange; they are mediated by pulmonary vagal and chemoreceptor inputs (Glowgowska et al. 1972). Sighs persisted during IRL because of the load-induced hypoventilation, the continuing hypoxaemia reducing the central (medullary) threshold needed to elicit them despite the load-induced reduction in tidal volume. Because fast twitch fibres can be recruited during sighs (Sieck & Fournier, 1989), we characterized drive (∫Phr, but not ∫Phr/TI because of the biphasic nature of the phrenic discharge during sighs) and pressure generation (Pdi) during sighs to determine when fatigue appeared during IRL (Fig. 5). Figure 5A shows that pressure generation during sighs increased during loading and remained at a plateau until pump failure (time = 1.0). A plot of the Pdi/∫Phr ratio (Fig. 5B) versus normalized load duration revealed an initial increase with loading (possibly reflecting recruitment of fast fatiguable motor units contributing disproportionately to Pdi) which was maintained until time = 0.6 at which time a greater drive was required to maintain pressure generation, indicating the recruitment of both damaged and ‘healthy’ motor units.

Figure 5. Plots of normalized Pdi(top) and normalized Pdi/∫Phr (bottom) during sighs versus normalized time in all rats.

Figure 5

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During IRL, Pdi maintained a plateau (no differences between values from time = 0.1 to time = 0.95) until pump failure (time = 1; P < 0.001, one-way repeated measures ANOVA on raw data). In contrast, Pdi/∫Phr increased during loading but fell at time = 0.6 below the value at time = 0.1 (P < 0.016, one-way repeated measures ANOVA on raw data). Arrow indicates onset of sigh fatigue; * significantly different from values at time = 0.1. Data points and error bars are means ±s.e.m.

We compared the temporal profiles of pressure generation (Pdi) during 30 s occlusions pre- and post-IRL (e.g. arrows in Fig. 1; expanded traces in Fig. 6A and B). Rats generated the same Pdi profiles on average (Fig. 6C, top) pre- and post-IRL; this is not surprising because chemical drives were similar (control PaO2 (mean ±s.d.) 73.0 ± 3.3 mmHg, recovery PaO2 66.8 ± 9.9 mmHg, P= 0.640; control PaCO2 36.5 ± 8.0 mmHg, recovery PaCO2 39.7 ± 8.9 mmHg, P= 0.995; control pH 7.42 ± 0.03, recovery pH 7.36 ± 0.07, P= 0.302 (all two-tailed paired t tests)). However, pressure generation post-IRL required significantly greater drive (∫Phr), as illustrated in the bottom pnael of Fig. 6C in which are plotted the averaged temporal profiles of Pdi/∫Phr for all rats. This difference was apparent even for unoccluded and unloaded inspirations (Fig. 6C, time = c). Thus, although pressure (Pdi) generation during 30 s occlusions had recovered by 15 min after removal of the load, when supplemented by knowledge of respiratory drive (∫Phr), contractility was still impaired.

Figure 6. Diaphragmatic contractility before and after inspiratory resistive loading.

Figure 6

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A and B, representative tracings from one rat (same as in Fig. 1) showing Pdi profiles during 30 s occlusions before and after inspiratory resistive loading. C, top: averaged Pdi data for all rats, grouped by 5 s bins; there were no differences between Pdi generated during unloaded breaths (‘c’) and occlusions pre-(▪) and post-IRL (○). C, bottom, averaged Pdi/Phr during unloaded breaths (‘c’) and during pre- and post-IRL occlusions for all rats. * Ratio significantly lower post IRL (P < 0.005, repeated measures ANOVA with post hoc Tukey's test for non-parametric data).

Detection of skeletal troponin I in serum as a marker of muscle injury

In rats of series 1, we did not detect sTnI by WB-DSA in samples of frozen serum; we did detect it in fresh serum samples from the rats of series 2. Figure 7A shows tracings of the Pdi, Ptr and ∫Phr in the one rat of series 2 in which we measured Pdi and phrenic activity and from which we obtained serum samples at the indicated times. The ratio of Pdi/∫Phr declined after approximately 1.5 h of loading. The Western blot (Fig. 7B) reveals the presence of fast sTnI before loading, a subsequent decrease in band intensity, and a pronounced reappearance (lane 3) even before fatigue onset (lane 4; decrease in Pdi/∫Phr ratio). Similar release patterns of sTnI (i.e. load-induced release occurring midway through loading) were observed in the four other rats of series 2. Only the fast isoform of sTnI was detected, never slow sTnI or cTnI. In some rats, loaded and sham, fast sTnI was detected, at low levels, in control samples obtained ∼1 h following completion of surgery and prior to loading (e.g. Fig. 7B, control (ct)). These levels decreased over the first few serum samples but subsequently increased only in loaded rats (e.g. Fig. 7B). The initial fall in levels of fast sTnI reflects either haemodilution (haematocrit progressively decreased from 47 ± 2 to 35 ± 1 (P < 0.001) during loading due to flushing of the catheters with saline after blood sampling and administration of anaesthetic) and/or rapid clearance from the blood.

Figure 7. Release of fast sTnI during inspiratory resistive loading.

Figure 7

Figure 7

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A, tracings of (from top down) Pdi, Ptr and ∫Phr during loading (onset indicated by *) in one rat. Times of blood samples indicated by arrows. B, Western blot probed for the presence of fast sTnI using mAb FI-32 at times corresponding to arrows in top panel.

Assessment of post-translational modifications

Antibody binding and subsequent detection can be influenced by post-translational modifications (PTM) of the analyte. Modifications to cTnI, before and after release, can affect detection (Katrukha et al. 1998; Morjana, 1998; Wu et al. 1998; Labugger et al. 2000). We used at least two different antibodies to positively identify fast sTnI. Different mAbs revealed different profiles of this protein in serial blood samples taken during IRL. mAbs FI-32 and FI-23 (specificity previously determined (Simpson et al. 2002); they have distinct epitopes but may have a minor degree of overlap according to surface plasmon resonance (Takahashi et al. 1996)) consistently revealed different temporal patterns of fast sTnI release in different rats (Fig. 8A). Both mAb FI-23 and FI-32 always revealed a progressive increase in fast sTnI starting about midway through IRL; mAb FI-23, however, always showed a pronounced increase in immunoreactivity just before pump failure. In contrast, mAb 3I-35 did not reveal fast sTnI (Fig. 8B). These results were unaffected by either prolonging exposures or altering mAb concentrations, suggesting the presence of PTM of fast sTnI.

Figure 8. Comparisons of Western blot analysis of serial samples of serum from two rats (A and B) probed for fast sTnI using mAbs FI-32 and FI-23 (A) and mAbs FI-32 and 3I-35 (B).

Figure 8

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mAbs FI-32 and FI-23 revealed different sTnI profiles during IRL; while both revealed an increase in fast sTnI immunoreactivity throughout loading, mAb FI-23 always revealed a pronounced increase just before pump failure. In contrast, mAb 3I-35 did not detect fast sTnI.

Discussion

Our major findings are that IRL caused (1) a defined time course of pathophysiological events leading to pump failure (Fig. 9); (2) persistent impaired contractility of the diaphragm (Fig. 6); (3) release of fast sTnI into the blood, the timing of release coinciding with the onset of fatigue during sighs and hypercapnic failure; and (4) post-translational modification of fast sTnI in blood, modifications which affected its detection.

Figure 9. Time course of events leading to IRL-induced pump failure in rats.

Figure 9

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Our results in anaesthetized rats are consistent with loading causing increased respiratory drive sufficient to have prevented a fall in alveolar ventilation (hypercapnic failure) until about 60% of loading duration, at which time fast sTnI was detected in serum. Both hypercapnic ventilatory failure and the appearance of fast sTnI in blood coincided with a drop in the sigh Pdi/∫Phr ratio, indicating that fast (fatiguable and/or fatigue resistant) fibres were injured and, for the first time, that information derived from sighs may help predict diaphragmatic fatigue (Fig. 9). Fatigue compromised pressure generation and led, eventually, to overt respiratory muscle fatigue (a drop in the Pdi/∫Phr ratio of non-sigh-loaded breaths), followed by pump failure. This end-stage failure was characterized by rapid decreases in both respiratory frequency and duty cycle (TI/TTOT), changes consistent with ‘fatigue’ of the respiratory central pattern generator (Yanos et al. 1990, 1994; De Vito & Roncoroni, 1993; Ferguson, 1995; Simpson et al. 2000).

Failure, fatigue, injury and recovery

Hypercapnic ventilatory failure is evident as an acute increase in arterial or end-tidal PCO2 (e.g. Sassoon et al. 1996; Sliwinski & Macklem, 1997; Reid & Belcastro, 2000; Lyall et al. 2001) and pump failure as an inability to sustain a target pressure (e.g. Morales et al. 1993; Laghi et al. 1998; Supinski et al. 1999), often, but not always (e.g. Supinski et al. 1999), in the absence of information about respiratory drive, or as apnoea (e.g. Yanos et al. 1990; Sassoon et al. 1996; Supinski et al. 1995, 1997; Ciufo et al. 2001). Neither the inability to sustain a desired pressure nor apnoea proves fatigue (and the authors did not suggest this) but one advantage of our model in which neural drive is measured is that one can differentiate fatigue from pump failure.

Determining if fatigue is present is complicated and usually involves detecting a deficit in respiratory muscle contractile function. In animals, particularly in models in which the diaphragm is excised and stimulated in vitro, fatigue is deemed to be present when less force is generated at a given stimulus frequency. In humans, evaluating diaphragmatic contractile function requires either the subject's co-operation (a sniff or maximal inspiratory pressure generation) or sophisticated techniques (power spectrum analysis of the diaphragm or measurement of Pdi during an elicited twitch). Early suggestions that changes in the power spectrum of its electromyogram can be used have not been confirmed (see Roussos & Zakynthinos, 1996). Instead, investigators now measure Pdi elicited by bilateral supramaximal stimulation of the phrenic nerves and this technique can be used to differentiate between peripheral and central fatigue (Bellemare &Bigland-Ritchie, 1984, 1987). Indeed, central fatigue is deemed to be present when phrenic stimulation elicits an increased Pdi during a maximal inspiratory effort.

In our rats, fatigue was indicated by the fall in the Pdi/∫Phr ratio and was present earlier when measured during sighs (Fig. 5) than during normal loaded breaths (Fig. 4). This probably reflects the increased contribution of fast motor units to pressure generation during sighs (Sieck & Fournier, 1989) and, since these units are more susceptible to fatigue, it is not surprising that this decrease occurred before diaphragmatic fatigue (analysis of normal loaded breaths). We do not know if the release of fast sTnI resulted from damage sustained during their recruitment during sighs, during ‘normal’ loaded breaths, or both.

The absence of changes in timing (TI/TTOT) and respiratory drive (∫Phr/Ti) indicate that central fatigue and neurotransmission failure at the level of the phrenic motoneurones, respectively, did not contribute to peripheral fatigue (a fall in Pdi/∫Phr), whether during sighs or normal loaded breaths. During loading, Pdi/∫Phr decreased (at ∼60% and at ∼80% of IRL duration for sighs and ‘normal’ breaths, respectively) despite progressive increases in respiratory drive (∫Phr/TI) throughout loading, a result consistent with peripheral fatigue, not central failure. We cannot, however, exclude transmission failure at the neuromuscular junction for which the evidence in rabbits (Aldrich, 1987; Osborne & Road, 1995; Sassoon et al. 1996), sheep (Bazzy & Donnelly, 1993) and dogs (Aubier et al. 1981b; Nava & Bellemare, 1989; De Vito & Roncoroni, 1993) is contradictory.

The fast recovery of pressure generation during occlusions following pump failure in our model is compatible with fatigue as defined by the study group of the National Heart Lung and Blood Institute (1990) but the persistence of a depressed Pdi/∫Phr ratio (i.e. increased drive necessary to maintain the same pressure generation) is not. Our results are similar to those of previous work in cats on IRL (Iscoe, 1996) and in rabbits subjected to brief loads after prolonged IRL (Road & Cairns, 1997). They are also consistent with an earlier report in man of rapid recovery of pressure-generating capacity following progressive inspiratory threshold loading to failure coexisting with reduced twitch amplitudes in response to phrenic nerve stimulation (Eastwood et al. 1994). The simplest explanation for the increased phrenic activity post IRL is that more phrenic drive was required to maintain normocapnia in rats in which injured inspiratory (diaphragmatic) motor units could not contribute to pressure generation. Thus, even though pressure generation had recovered, at least some fatigue-related mechanism(s) was still present because increased drive was necessary to produce this pressure (and therefore ventilation and PaCO2).

Anaesthetized rat model of diaphragmatic injury and fatigue

The rat, unlike the mouse, is large enough to permit the insertion of pressure tip catheters to measure Pdi. It also has a blood volume that permits multiple samples for blood gas analysis and scanning for such potential biomarkers as sTnI (Simpson et al. 2002), thereby minimizing the risk of circulatory compromise. In addition, its diaphragm, like that of most other (larger) mammals including man (Metzger et al. 1985), is mixed whereas that of the mouse is composed only of fast oxidative fibres (Green et al. 1984), allowing determination of the effects of stress on different fibre types. This is important because, under conditions of loading, we obtained evidence that fast motor units fatigued before slow ones (if the slow ones ever did) and this coincided with the release of fast sTnI. In contrast, in some models, especially in vitro diaphragm or diaphragmatic strips, fatigue is induced using supramaximal stimulation of the muscle or, less often, of the phrenic nerves, and this excites all motor units and at the same frequency, something that never happens normally (Iscoe et al. 1976; Sieck & Fournier, 1989). Imposition of the same contractile regimen on motor units with different contractile characteristics may therefore injure and fatigue fibre types, with consequent effects on sTnI release, in a manner very different from that under ‘normal’ circumstances. Indeed, under conditions of increased chemical drive, as elicited here by IRL, motor units are recruited in a stereotyped pattern, from slow oxidative to fast fatiguable (Sieck & Fournier, 1989), in which compensation for injured diaphragmatic motor units can be accomplished by recruiting other motor units in the diaphragm and, possibly, synergistic muscles, and by generating more force with doublets at the onset of discharge (Burke et al. 1970; Iscoe, 1996).

Type of load

We chose IRL because it is technically simple and elicited respiratory muscle dysfunction (hypercapnic failure followed by diaphragmatic fatigue and then pump failure) over a period (<5 h) relevant to acute respiratory failure, whether of peripheral or central (or some combination) origin. It differs from the many hours to days associated with such interventions as tracheal banding (e.g. Reid et al. 1994, 1992), induced emphysema (e.g. Lewis et al. 1992), or repetitive resistive loading (Zhu et al. 1997), all of which elicit varying degrees of adaptation and remodelling (e.g. fibre-type and isoform switching; Levine et al. 1997) that would complicate analysis of the underlying molecular cause(s) of acute dysfunction (Simpson et al. 2003).

Diaphragmatic injury and hypercapnic failure

The loss of skeletal myofilament proteins, including sTnI, and its appearance in blood suggests impairment or loss of contractile function of at least some muscle cells. We detected, within the limits to do so given the few mAbs specific for sTnI, release of fast sTnI about the time of hypercapnic failure, suggesting that any compensation was inadequate to maintain alveolar ventilation and prevent CO2 retention. Although we have previously shown that the anti-TnI mAb specific to the slow isoform can detect slow sTnI in serum using WB-DSA (Simpson et al. 2002), we do not yet know if the fast and slow anti-TnI mAbs have similar affinities for their respective antigens. Thus, low levels of slow sTnI may have been present but below the level of detection of the anti-slow TnI mAb used, as suggested by our inability to detect any surgery-induced release of slow sTnI.

Skeletal troponin I

The potential of the skeletal troponin isoforms, fast and slow sTnI, as markers of muscle injury has, in comparison to their cardiac counterpart, received little attention. sTnI may be useful in assessing the effects of intense exercise (Rama et al. 1996; Sorichter et al. 1997); for orthopedic and soft tissue injuries, it is more sensitive and specific than creatine kinase and myoglobin (Onuoha et al. 2001). We recently demonstrated that fast and slow sTnI were released into the blood of a patient with rhabdomyolysis (Simpson et al. 2002); both isoforms displayed proteolysis, fast sTnI also appearing as a doublet, a finding similar to that for cTnI in patients with acute myocardial infarction (Labugger et al. 2000). Here, we report that, in anaesthetized rats, IRL caused release of fast sTnI, presumably from the diaphragm. Unlike that patient, we detected only intact fast sTnI which was neither degraded (only 1 of the 5 rats had minor amounts of degradation) nor present as a doublet.

Fast sTnI was present in some samples taken before IRL and in sham (unloaded) rats; this was likely to be due to damage caused by surgery, even in rats in which we did not record phrenic activity but still had to cannulate the trachea and the carotid artery and jugular vein.

The presence of more fast sTnI in the blood of our loaded rats proves that load-induced injury was present and this could have contributed to the impaired ability of the diaphragm to translate increased respiratory drive (∫Phr) into pressure (Pdi). The extent of membrane injury (i.e. if necrosis is obligatory) needed for release of intracellular proteins is unknown, a controversy recently reviewed by Mair (1999) for markers of myocardial infarction.

sTnI as a biomarker

For serum levels of sTnI to be a useful diagnostic of respiratory muscle dysfunction, its release must be related to the timing and severity of the patient's injury. However, several issues must be addressed, issues identical to those that still, a decade after their introduction, plague commercial assays for cTnI. These assays vary in their lower limit of detection by up to 50 fold; some assays fail to detect cTnI even when present (Apple et al. 2002). This is caused, at least in part, by PTM, differences in access to the epitope because of the protein's ternary structure and/or complex formation (with serum proteins or with one or more of the other two troponin subunits, troponin T and C), all of which can influence immunoreactivity and therefore detectability. WB-DSA eliminates most of these confounding factors because it denatures, reduces and separates electrophoretically the proteins. Moreover, the use of multiple mAbs provides information about PTM of the protein. mAb 3I-35, unlike both FI-32 and FI-23 (Fig. 7), did not detect fast sTnI in serum because of either its lower affinity or, more likely, a PTM to the analyte. The differential binding of mAbs FI-32 and FI-23 (Fig. 7) confirms the presence of a PTM. Thus, reliance on one mAb for detection can be misleading in terms of both the levels of the protein present and the timing of its release. Last, unlike cTnI, which is specific to the heart, sTnI can be released from any skeletal muscle, not just respiratory muscles.

Time course of fast sTnI release

The time course of appearance of a muscle protein (biomarker) in blood will depend on factors affecting its release and clearance. Release of myofilament proteins is influenced, at least in part, by compartmentalization and local blood flow. Soluble cytosolic proteins may be released before insoluble structurally bound proteins (Adams, 1999; Mair, 1999). In skeletal muscle, there are cytosolic and structurally bound pools of sTnI (Sorichter et al. 1997) similar to those for cTnI (Mair et al. 1995). Although a change in level of cTnI is not detected within the first few hours after onset of symptoms of myocardial infarction (Wu et al. 1999), we could detect an increase in fast sTnI as early as 1 h after load onset. This faster ‘response’ is likely to be related to differences in perfusion. Because cTnI is most often used to detect acute myocardial infarction, a condition caused by low or no blood flow, its washout from the infarcted area will be slow. In experiments in which reperfusion is not limited, i.e. cultured cells (Van Nieuwenhoven et al. 1996) and perfused Langendorff rat hearts (Vorderwinkler et al. 1996), cTnI levels increase within 5 min, a result supported by measurements of cTnI levels after reperfusion of human myocardium following cardioplegic arrest (Bleier et al. 1998). During IRL, perfusion of the diaphragm and other respiratory muscles increases (Mayock et al. 1992; Pang et al. 1993; Janssens et al. 1995; Rochester & Bettini, 1976; Robertson et al. 1977); if anything, therefore, washout of skeletal cytosolic proteins should be enhanced.

Cause of sTnI release

Our data provide no information about the mechanism(s) underlying sTnI release. Candidates include mechanical stress associated with the high pressures generated against the load and the local metabolic conditions in the inspiratory muscles, particularly the diaphragm, the likely source of the sTnI because of its fatigue. There are no previous studies using a similar protocol to which we can compare our data. However, Jiang et al. (1998a) observed injury of the diaphragm 3 days after IRL. Because there was no damage to the gastrocnemius exposed to the same hypercapnic and acidotic arterial blood (hypoxaemia was prevented by administering supplemental oxygen), hypercapnia/acidosis was not responsible for the injury. The role of hypoxaemia in causing injury in our experiments is unresolved; inspiratory loading, at least in other animal models (Mayock et al. 1992; Rochester & Bettini, 1976; Robertson et al. 1977; Janssens et al. 1995), elicits marked increases in diaphragmatic perfusion that, by offsetting decreased arterial oxygen content, could maintain oxygen delivery. Indeed, Mayock et al. (1992) showed in piglets that 1 h of hypoxaemia did not compromise diaphragmatic pressure generation more than loading alone, suggesting an absence of injury.

Summary

In summary, in anaesthetized rats, IRL elicited hypercapnic ventilatory failure approximately coincident with injury and fatigue of fast motor units of the diaphragm (release of fast sTnI into the blood and a reduction in sigh Pdi/∫Phr). This precipitated diaphragmatic fatigue during non-sigh-loaded breaths (decreased pressure generation despite increased drive) and eventual pump failure, probably due to central fatigue (decrease in respiratory frequency and duty cycle). A model is shown in Fig. 9. Detection of load-induced injury of slow twitch, fatigue-resistant fibres, if present, may require development of more sensitive antibodies. Measurements of sTnI in blood may help in assessing respiratory muscle dysfunction and may be useful as a marker of impending contractile fatigue and respiratory failure.

Acknowledgments

We thank Sheila Gordon for superb technical assistance and Drs John Fisher, Peter Macklem and Jerry Dempsey for helpful comments. This work was supported by grants from the Canadian Institutes of Health Research, the Ontario Thoracic Society, a Block Term Grant to Queen's University from the Ontario Thoracic Society, the William M. Spear Endowment Fund, and funds from the Faculty of Health Sciences, Queen's University. J.A.S. also received an Ontario Graduate Student Science and Technology Award and support from both the School of Graduate Studies and Research, Queen's University and the Ontario Thoracic Society via grants to Queen's University.

References

  1. Adams JEI. Clinical application of markers of cardiac injury: basic concepts and new considerations. Clin Chim Acta. 1999;284:127–134. doi: 10.1016/s0009-8981(99)00074-1. [DOI] [PubMed] [Google Scholar]
  2. Aldrich TK. Transmission fatigue of the rabbit diaphragm. Respir Physiol. 1987;69:307–319. doi: 10.1016/0034-5687(87)90085-5. [DOI] [PubMed] [Google Scholar]
  3. Apple FS, Wu AH, Jaffe AS. European Society of Cardiology and American College of Cardiology guidelines for redefinition of myocardial infarction: how to use existing assays clinically and for clinical trials. Am Heart J. 2002;144:981–986. doi: 10.1067/mhj.2002.124048. [DOI] [PubMed] [Google Scholar]
  4. Aubier M, Farkas G, De Troyer A, Mozes R, Roussos C. Detection of diaphragmatic fatigue in man by phrenic stimulation. J Appl Physiol. 1981a;50:538–544. doi: 10.1152/jappl.1981.50.3.538. [DOI] [PubMed] [Google Scholar]
  5. Aubier M, Trippenbach T, Roussos C. Respiratory muscle fatigue during cardiogenic shock. J Appl Physiol. 1981b;51:499–508. doi: 10.1152/jappl.1981.51.2.499. [DOI] [PubMed] [Google Scholar]
  6. Bazzy AR, Donnelly DF. Diaphragmatic failure during loaded breathing: Role of neuromuscular transmission. J Appl Physiol. 1993;74:1679–1683. doi: 10.1152/jappl.1993.74.4.1679. [DOI] [PubMed] [Google Scholar]
  7. Bellemare F, Bigland-Ritchie B. Assessment of human diaphragm strength and activation using phrenic nerve stimulation. Respir Physiol. 1984;58:263–277. doi: 10.1016/0034-5687(84)90003-3. [DOI] [PubMed] [Google Scholar]
  8. Bellemare F, Bigland-Ritchie B. Central components of diaphragmatic fatigue assessed by phrenic nerve stimulation. J Appl Physiol. 1987;62:1307–1316. doi: 10.1152/jappl.1987.62.3.1307. [DOI] [PubMed] [Google Scholar]
  9. Bleier J, Vorderwinkler KP, Falkensammer J, Mair P, Dapunt O, Puschendorf B, Mair J. Different intracellular compartmentations of cardiac troponins and myosin heavy chains: a causal connection to their different early release after myocardial damage. Clin Chem. 1998;44:1912–1918. [PubMed] [Google Scholar]
  10. Burke RE, Rudomin P, Zajac FE. Catch property in single mammalian motor units. Science. 1970;168:122–124. doi: 10.1126/science.168.3927.122. [DOI] [PubMed] [Google Scholar]
  11. Ciufo R, Dimarco A, Stofan D, Nethery D, Supinski G. Dichloroacetate reduces diaphragmatic lactate formation but impairs respiratory performance. Am J Respir Crit Care Med. 2001;164:1669–1674. doi: 10.1164/ajrccm.164.9.9902054. [DOI] [PubMed] [Google Scholar]
  12. Collinson PO. Testing for cardiac markers at the point of care. Clin Laboratory Med. 2001;21:351–362. [PubMed] [Google Scholar]
  13. De Vito EL, Roncoroni AJ. Central and peripheral diaphragmatic fatigue in loaded normal and vagotomized dogs. J Appl Physiol. 1993;74:2820–2827. doi: 10.1152/jappl.1993.74.6.2820. [DOI] [PubMed] [Google Scholar]
  14. Eastwood PR, Hillman DR, Finucane KE. Ventilatory responses to inspiratory threshold loading and role of muscle fatigue in task failure. J Appl Physiol. 1994;76:185–195. doi: 10.1152/jappl.1994.76.1.185. [DOI] [PubMed] [Google Scholar]
  15. Ferguson GT. Respiratory failure due to altered central drive during inspiratory loading in rabbits. Respir Physiol. 1995;99:75–87. doi: 10.1016/0034-5687(94)00073-9. [DOI] [PubMed] [Google Scholar]
  16. Gandevia SC. Spinal and supraspinal factors in human muscle fatigue. Physiol Rev. 2001;81:1725–1789. doi: 10.1152/physrev.2001.81.4.1725. [DOI] [PubMed] [Google Scholar]
  17. Gandevia SC, Allen GM, Butler JE, Gorman RB, McKenzie DK. Human respiratory muscles: sensations, reflexes and fatiguability. Clin Exp Pharmacol Physiol. 1998;25:757–763. doi: 10.1111/j.1440-1681.1998.tb02150.x. [DOI] [PubMed] [Google Scholar]
  18. Glowgowska M, Richardson PS, Widdicombe JG, Winning AJ. The role of the vagus nerves, peripheral chemoreceptors and other afferent pathways in the genesis of augmented breaths in cats and rabbits. J Physiol. 1972;16:179–196. doi: 10.1016/0034-5687(72)90050-3. [DOI] [PubMed] [Google Scholar]
  19. Green HJ, Reichmann H, Pette D. Inter- and intraspecies comparisons of fibre type distribution and of succinate dehydrogenase activity in type I, IIA and IIB fibres of mammalian diaphragms. Histochemistry. 1984;81:67–73. doi: 10.1007/BF00495403. [DOI] [PubMed] [Google Scholar]
  20. Iscoe S. Phrenic motoneuron discharge during sustained inspiratory resistive loading. J Appl Physiol. 1996;81:2260–2266. doi: 10.1152/jappl.1996.81.5.2260. [DOI] [PubMed] [Google Scholar]
  21. Iscoe S, Dankoff J, Migicovsky R, Polosa C. Recruitment and discharge frequency of phrenic motoneurones during inspiration. Respir Physiol. 1976;26:113–128. doi: 10.1016/0034-5687(76)90056-6. [DOI] [PubMed] [Google Scholar]
  22. Janssens S, Derom E, Vanhaecke J, Decramer M. Theophylline increases oxygen consumption during inspiratory resistive loading. Am J Respir Crit Care Med. 1995;151:1000–1005. doi: 10.1164/ajrccm.151.4.7697222. [DOI] [PubMed] [Google Scholar]
  23. Jiang TX, Reid WD, Belcastro A, Road JD. Load dependence of secondary diaphragm inflammation and injury after acute inspiratory loading. Am J Respir Crit Care Med. 1998a;157:230–236. doi: 10.1164/ajrccm.157.1.9702051. [DOI] [PubMed] [Google Scholar]
  24. Jiang TX, Reid WD, Road JD. Delayed diaphragm injury and diaphragm force production. Am J Respir Crit Care Med. 1998b;157:736–742. doi: 10.1164/ajrccm.157.3.9707056. [DOI] [PubMed] [Google Scholar]
  25. Jiang TX, Reid WD, Road JD. Free radical scavengers and diaphragm injury following inspiratory resistive loading. Am J Respir Crit Care Med. 2001;164:1288–1294. doi: 10.1164/ajrccm.164.7.2005081. [DOI] [PubMed] [Google Scholar]
  26. Katrukha AG, Bereznikova AV, Filatov VL, Esakova TV, Kolosova OV, Pettersson K, Lovgren T, Bulgarina TV, Trifonov IR, Gratsiansky NA, Pulkki K, Voipio-Pulkki LM, Gusev NB. Degradation of cardiac troponin I: implication for reliable immunodetection. Clin Chem. 1998;44:2433–2440. [PubMed] [Google Scholar]
  27. Labugger R, Organ L, Collier C, Atar D, Van Eyk JE. Extensive troponin I and T modification detected in serum from patients with acute myocardial infarction. Circulation. 2000;102:1221–1226. doi: 10.1161/01.cir.102.11.1221. [DOI] [PubMed] [Google Scholar]
  28. Laghi F, Topeli A, Tobin MJ. Does resistive loading decrease diaphragmatic contractility before task failure. J Appl Physiol. 1998;85:1103–1112. doi: 10.1152/jappl.1998.85.3.1103. [DOI] [PubMed] [Google Scholar]
  29. Levine S, Kaiser L, Leferovich J, Tikunov B. Cellular adaptations in the diaphragm in chronic obstructive pulmonary disease. N Engl J Med. 1997;337:1799–1806. doi: 10.1056/NEJM199712183372503. [DOI] [PubMed] [Google Scholar]
  30. Lewis MI, Zhan WZ, Sieck GC. Adaptations of the diaphragm in emphysema. J Appl Physiol. 1992;72:934–943. doi: 10.1152/jappl.1992.72.3.934. [DOI] [PubMed] [Google Scholar]
  31. Lyall RA, Donaldson N, Polkey MI, Leigh PN, Moxham J. Respiratory muscle strength and ventilatory failure in amyotrophic lateral sclerosis. Brain. 2001;124:2000–2013. doi: 10.1093/brain/124.10.2000. [DOI] [PubMed] [Google Scholar]
  32. MacGowan NA, Evans KG, Road JD, Reid WD. Diaphragm injury in individuals with airflow obstruction. Am J Respir Crit Care Med. 2001;163:1654–1659. doi: 10.1164/ajrccm.163.7.2001042. [DOI] [PubMed] [Google Scholar]
  33. Mador MJ, Khan S, Kufel TJ. Bilateral anterolateral magnetic stimulation of the phrenic nerves can detect diaphragmatic fatigue. Chest. 2002;121:452–458. doi: 10.1378/chest.121.2.452. [DOI] [PubMed] [Google Scholar]
  34. Mair J. Tissue release of cardiac markers: from physiology to clinical applications. Clin Chem Laboratory Med. 1999;37:1077–1084. doi: 10.1515/CCLM.1999.157. [DOI] [PubMed] [Google Scholar]
  35. Mair J, Morandell D, Genser N, Lechleitner P, Dienstl F, Puschendorf B. Equivalent early sensitivities of myoglobin, creatine kinase MB mass, creatine kinase isoform ratios, and cardiac troponins I and T for acute myocardial infarction. Clin Chem. 1995;41:1266–1272. [PubMed] [Google Scholar]
  36. Matsumoto N, Nakamura T, Yasui Y, Torii J. Immunohistochemical differentiation of fiber types in human skeletal muscle using monoclonal antibodies to slow and fast isoforms of troponin I subunit. Biotech Histochem. 1997;72:191–197. doi: 10.3109/10520299709082237. [DOI] [PubMed] [Google Scholar]
  37. Mayock DE, Standaert TA, Woodrum DE. Effect of inspiratory resistive loaded breathing and hypoxemia on diaphragmatic function in the piglet. J Appl Physiol. 1992;73:1888–1893. doi: 10.1152/jappl.1992.73.5.1888. [DOI] [PubMed] [Google Scholar]
  38. Metzger JM, Scheidt KB, Fitts RH. Histochemical and physiological characteristics of the rat diaphragm. J Appl Physiol. 1985;58:1085–1091. doi: 10.1152/jappl.1985.58.4.1085. [DOI] [PubMed] [Google Scholar]
  39. Morales CF, Anzueto A, Andrade F, Levine SM, Maxwell LC, Lawrence RA, Jenkinson SG. Diethylmaleate produces diaphragmatic impairment after resistive breathing. J Appl Physiol. 1993;75:2406–2411. doi: 10.1152/jappl.1993.75.6.2406. [DOI] [PubMed] [Google Scholar]
  40. Morjana NA. Degradation of human cardiac troponin I after myocardial infarction. Biotechnol Appl Biochem. 1998;28:105–111. [PubMed] [Google Scholar]
  41. National Heart Lung and Blood Institute. NHLBI Workshop Summary: Respiratory muscle fatigue; report of the Respiratory Muscle Fatigue Workshop Group. Am Rev Respir Dis. 1990;142:474–480. doi: 10.1164/ajrccm/142.2.474. [DOI] [PubMed] [Google Scholar]
  42. Nava S, Bellemare F. Cardiovascular failure and apnea in shock. J Appl Physiol. 1989;66:184–189. doi: 10.1152/jappl.1989.66.1.184. [DOI] [PubMed] [Google Scholar]
  43. Ni CY. Cardiac troponin I: a biomarker for detection and risk stratification of minor myocardial damage. Clin Laboratory. 2001;47:483–492. [PubMed] [Google Scholar]
  44. Onuoha GN, Alpar EK, Dean B, Tidman J, Rama D, Laprade M, Pau B. Skeletal troponin-I release in orthopedic and soft tissue injuries. J Orthop Sci. 2001;6:11–15. doi: 10.1007/s007760170018. [DOI] [PubMed] [Google Scholar]
  45. Orozco-Levi M, Lloreta J, Minguella J, Serrano S, Broquetas JM, Gea J. Injury of the human diaphragm associated with exertion and chronic obstructive pulmonary disease. Am J Respir Crit Care Med. 2001;164:1734–1739. doi: 10.1164/ajrccm.164.9.2011150. [DOI] [PubMed] [Google Scholar]
  46. Osborne S, Road JD. Diaphragm and phrenic nerve activities during inspiratory loading in anesthetized rabbits. Respir Physiol. 1995;99:321–330. doi: 10.1016/0034-5687(94)00109-d. [DOI] [PubMed] [Google Scholar]
  47. Pang LM, Kim YJ, Bazzy AR. Blood flow to respiratory muscles and major organs during inspiratory flow resistive loads. J Appl Physiol. 1993;74:428–434. doi: 10.1152/jappl.1993.74.1.428. [DOI] [PubMed] [Google Scholar]
  48. Rama D, Margaritis I, Orsetti A, Marconneet P, Gros P, Larue C, Trinquier S, Pau B, Calzolari C. Troponin I immunoenzymometric assays for detection of muscle damage applied to monitoring a triathlon. Clin Chem. 1996;42:2033–2035. [PubMed] [Google Scholar]
  49. Reid WD, Belcastro AN. Chronic resistive loading induces diaphragm injury and ventilatory failure in the hamster. Respir Physiol. 1999;118:203–218. doi: 10.1016/s0034-5687(99)00089-4. [DOI] [PubMed] [Google Scholar]
  50. Reid WD, Belcastro AN. Time course of diaphragm injury and calpain activity during resistive loading. Am J Respir Crit Care Med. 2000;162:1801–1806. doi: 10.1164/ajrccm.162.5.9906033. [DOI] [PubMed] [Google Scholar]
  51. Reid WD, Huang J, Bryson S, Walker DC, Belcastro AN. Diaphragm injury and myofibrillar structure induced by resistive loading. J Appl Physiol. 1994;76:176–184. doi: 10.1152/jappl.1994.76.1.176. [DOI] [PubMed] [Google Scholar]
  52. Reid WD, MacGowan N. Respiratory muscle injury in animal models and humans. Mol Cell Biochem. 1998;179:63–80. doi: 10.1023/a:1006803703128. [DOI] [PubMed] [Google Scholar]
  53. Reid WD, Noonan J, Chung F, Tesler-Mabe C. Ventilatory failure induced by tracheal banding in the hamster. J Appl Physiol. 1992;73:1671–1675. doi: 10.1152/jappl.1992.73.4.1671. [DOI] [PubMed] [Google Scholar]
  54. Road JD, Cairns AM. Phrenic motoneuron firing rates before, during, and after prolonged inspiratory resistive loading. J Appl Physiol. 1997;83:776–783. doi: 10.1152/jappl.1997.83.3.776. [DOI] [PubMed] [Google Scholar]
  55. Road JD, Jiang TX. Determinants of diaphragmatic injury. Mol Cell Biochem. 1998;179:81–86. doi: 10.1023/a:1006855819966. [DOI] [PubMed] [Google Scholar]
  56. Robertson CH, Jr, Foster GH, Johnson RL., Jr The relationship of respiratory failure to the oxygen consumption of, lactate production by, and distribution of blood flow among respiratory muscles during increasing inspiratory resistance. J Clin Invest. 1977;59:31–42. doi: 10.1172/JCI108619. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Rochester DF, Bettini G. Diaphragmatic blood flow and energy expenditure in the dog. Effects of inspiratory airflow resistance and hypercapnia. J Clin Invest. 1976;57:661–672. doi: 10.1172/JCI108322. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Roussos C, Zakynthinos S. Fatigue of the respiratory muscles. Intensive Care Med. 1996;22:134–155. doi: 10.1007/BF01720721. [DOI] [PubMed] [Google Scholar]
  59. Sassoon CS, Gruer SE, Sieck GC. Temporal relationships of ventilatory failure, pump failure, and diaphragm fatigue. J Appl Physiol. 1996;81:238–245. doi: 10.1152/jappl.1996.81.1.238. [DOI] [PubMed] [Google Scholar]
  60. Sieck GC, Fournier M. Diaphragm motor unit recruitment during ventilatory and nonventilatory behaviors. J Appl Physiol. 1989;66:2539–2545. doi: 10.1152/jappl.1989.66.6.2539. [DOI] [PubMed] [Google Scholar]
  61. Simpson JA, Iscoe S, Van Eyk JE. Myofilament proteomics: understanding contractile dysfunction in cardiorespiratory disease. In: Van Eyk JE, Dunn M, editors. Proteomic and Genomic Analysis of Cardiovascular Disease. Weinheim: Wiley-VCH; 2003. pp. 317–337. [Google Scholar]
  62. Simpson JA, Labugger R, Hesketh GG, Arsigny C, O'Donnell DE, Matsumoto N, Collier CP, Iscoe S, Van Eyk JE. Differential detection of skeletal troponin I isoforms in serum of a patient with rhabdomyolysis: markers of muscle injury. Clin Chem. 2002;48:1112–1114. [PubMed] [Google Scholar]
  63. Simpson JA, Van Eyk JE, Iscoe S. Hypoxemia-induced modification of troponin I and T in canine diaphragm. J Appl Physiol. 2000;88:753–760. doi: 10.1152/jappl.2000.88.2.753. [DOI] [PubMed] [Google Scholar]
  64. Sliwinski P, Macklem PT. Inspiratory muscle dysfunction as a cause of death in COPD patients. Monaldi Arch Chest Dis. 1997;52:380–383. [PubMed] [Google Scholar]
  65. Sorichter S, Mair J, Koller A, Gebert W, Rama D, Calzolari C, Artner-Dworzak E, Puschendorf B. Skeletal troponin I as a marker of exercise-induced muscle damage. J Appl Physiol. 1997;83:1076–1082. doi: 10.1152/jappl.1997.83.4.1076. [DOI] [PubMed] [Google Scholar]
  66. Supinski G, Nethery D, Stofan D, Szweda L, Dimarco A. Oxypurinol administration fails to prevent free radical-mediated lipid peroxidation during loaded breathing. J Appl Physiol. 1999;87:1123–1131. doi: 10.1152/jappl.1999.87.3.1123. [DOI] [PubMed] [Google Scholar]
  67. Supinski GS, Stofan D, Ciufo R, Diamrco A. N-acetylcysteine administration and loaded breathing. J Appl Physiol. 1995;79:340–347. doi: 10.1152/jappl.1995.79.1.340. [DOI] [PubMed] [Google Scholar]
  68. Supinski GS, Stofan D, Ciufo R, Dimarco A. N-acetylcysteine administration alters the response to inspiratory loading in oxygen-supplemented rats. J Appl Physiol. 1997;82:1119–1125. doi: 10.1152/jappl.1997.82.4.1119. [DOI] [PubMed] [Google Scholar]
  69. Takahashi M, Lee L, Shi Q, Gawad Y, Jackowski G. Use of enzyme immunoassay for measurement of skeletal troponin-I utilizing isoform-specific monoclonal antibodies. Clin Biochem. 1996;29:301–308. doi: 10.1016/0009-9120(96)00016-1. [DOI] [PubMed] [Google Scholar]
  70. Van Nieuwenhoven FA, Musters RJ, Post JA, Verkleij AJ, van der Vusse GJ, Glatz JF. Release of proteins from isolated neonatal rat cardiomyocytes subjected to simulated ischemia or metabolic inhibition is independent of molecular mass. J Mol Cell Cardiol. 1996;28:1429–1434. doi: 10.1006/jmcc.1996.0133. [DOI] [PubMed] [Google Scholar]
  71. Vorderwinkler KP, Mair J, Puschendorf B, Hempel A, Schluter KD, Piper HM. Cardiac troponin I increases in parallel to cardiac troponin T, creatine kinase and lactate dehydrogenase in effluents from isolated perfused rat hearts after hypoxia-reoxygenation-induced myocardial injury. Clin Chim Acta. 1996;251:113–117. doi: 10.1016/0009-8981(96)06303-6. [DOI] [PubMed] [Google Scholar]
  72. Wu AH, Apple FS, Gibler WB, Jesse RL, Warshaw MM, Valdes RJ. National Academy of Clinical Biochemistry Standards of Laboratory Practice: recommendations for the use of cardiac markers in coronary artery diseases. Clin Chem. 1999;45:1104–1121. [PubMed] [Google Scholar]
  73. Wu AH, Feng YJ, Moore R, Apple FS, McPherson PH, Buechler KF, Bodor G. Characterization of cardiac troponin subunit release into serum after acute myocardial infarction and comparison of assays for troponin T and I. American Association for Clinical Chemistry Subcommittee on cTnI Standardization. Clin Chem. 1998;44:1198–1208. [PubMed] [Google Scholar]
  74. Yanos J, Keamy MF, III, Leisk L, Hall JB, Walley KR, Wood LDH. The mechanism of respiratory arrest in inspiratory loading and hypoxemia. Am Rev Respir Dis. 1990;141:933–937. doi: 10.1164/ajrccm/141.4_Pt_1.933. [DOI] [PubMed] [Google Scholar]
  75. Yanos J, Patti MJ, Banner AS. Mechanism of respiratory arrest in an animal model of acute fatal bronchoconstriction. J Appl Physiol. 1994;77:236–244. doi: 10.1152/jappl.1994.77.1.236. [DOI] [PubMed] [Google Scholar]
  76. Zhu E, Petrof BJ, Gea J, Comtois N, Grassino AE. Diaphragm muscle fiber injury after inspiratory resistive breathing. Am J Respir Crit Care Med. 1997;155:1110–1116. doi: 10.1164/ajrccm.155.3.9116995. [DOI] [PubMed] [Google Scholar]

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