Inhibition of the Ubiquitin–Proteasome Pathway Does Not Protect against Ventilator-induced Accelerated Proteolysis or Atrophy in the Diaphragm

MECHANICAL ventilation (MV) is used in critical care medicine to maintain adequate alveolar ventilation in patients. Common indications for MV include pulmonary disorders, heart failure, neuromuscular diseases, coma, and surgery. Although MV can be a life-saving intervention, prolonged MV results in the rapid development of ventilator-induced diaphragm dysfunction (VIDD) which occurs due to diaphragm atrophy and contractile dysfunction.^1–3  From a clinical perspective, VIDD is important because respiratory muscle weakness is predicted to contribute to difficulties in weaning patients from the ventilator.⁴  However, there is currently no drug therapy or clinical standard of care to address this problem. Therefore, improving our understanding of the processes that promote VIDD is essential to develop a therapeutic strategy to prevent MV-induced diaphragm weakness. In this regard, we determined whether targeting the ubiquitin–proteasome pathway (UPP) via epoxomicin administration is sufficient to protect against VIDD. The clinical importance of VIDD and the rationale supporting our approach are further highlighted in the next section.

The development of VIDD occurs rapidly and has been reported to occur within 12 to 18 h after the initiation of MV.^1,2  Reports indicate that approximately 33% of all adult patients admitted to an intensive care unit require MV, and weaning procedures account for 40 to 50% of the total time spent in the critical care unit.⁵  Although numerous factors can contribute to weaning difficulties, the most frequent cause of weaning failure in patients is inspiratory (i.e., diaphragm) muscle weakness. Therefore, developing strategies to prevent VIDD is clinically important. Previous work reveals that MV-induced diaphragm atrophy occurs due to both decreased protein synthesis and increased proteolysis.^3,6,7  However, because of the rapid development of VIDD, MV-induced increases in diaphragm protein breakdown seem to play the dominant role.³  In regard to MV-induced proteolysis, all four primary proteolytic systems are activated in the diaphragm during MV, including calpain, caspase-3, autophagy, and the UPP.^3,8,9  However, only calpain and caspase-3 have been shown to participate in the development of VIDD and the role that autophagy and the UPP play remains unknown.^9–11  The absence of studies investigating the role of the UPP in the development of VIDD is surprising, given that the UPP is considered to be an important proteolytic system responsible for the breakdown of myofibrillar proteins and the UPP plays a required role in inactivity-induced atrophy in limb skeletal muscles.¹²  In this regard, a recent study by Agten et al.¹³  administered the protease inhibitor bortezomib to animals during prolonged MV in an attempt to elucidate the role that the UPP plays in the development of VIDD. However, bortezomib inhibited caspase-3 activation but did not inhibit the MV-induced increase in 20S proteasome activity in the diaphragm. Thus, additional work is needed to determine the contribution of UPP to VIDD and thus forms the basis for the current experiments. Therefore, these experiments tested the hypothesis that pharmacological inhibition of the UPP will protect the diaphragm against VIDD.

Young adult female Sprague–Dawley rats were used in these experiments. Animals were assigned to one of three experimental groups (n = 8 per group): (1) acutely anesthetized control animals (CON); (2) 12-h mechanically ventilated animals, no treatment (MV); and (3) 12 h of MV, treatment with the proteasome inhibitor epoxomicin (MV + epoxomicin). The Institutional Animal Care and Use Committee of the University of Florida (Gainesville, Florida) approved these experiments. In an effort to comply with the U.S. Animal Welfare Act and Public Health Service Policy by using the absolute minimum number of animals required to achieve the science, measurements made using the control and MV groups were also used in a previously published article.¹¹  Investigators were blinded during the analysis of the data when possible (diaphragm contractile force and cross-sectional area).

Control animals were acutely anesthetized with an intraperitoneal injection of sodium pentobarbital (100 mg/kg body weight). After reaching a surgical plane of anesthesia, the diaphragm was quickly removed and the costal diaphragm was divided into several segments. One strip of the medial costal diaphragm was stored for histological measurements, another strip was immediately used for specific force measurements, and the remaining portion of the costal diaphragm was rapidly frozen in liquid nitrogen and stored at −80°C for subsequent biochemical analyses.

All surgical procedures were performed using aseptic techniques. Animals in the MV groups were anesthetized with an intraperitoneal injection of sodium pentobarbital (60 mg/kg body weight). Animals were then tracheostomized, and mechanically ventilated with a pressure-controlled ventilator (Servo Ventilator 300; Siemens AG, Munich, Germany) for 12 h with the following settings: upper airway pressure limit, 20 cm H₂O; respiratory rate, 80 breaths/min; and positive end-expiratory pressure, 1 cm H₂O. Previous work by our laboratory has demonstrated that the full support mode of MV used in this study results in the complete cessation of inspiratory electromyogram activity in the diaphragm.² 

The carotid artery was cannulated to permit the continuous measurement of blood pressure and the collection of blood during the protocol. Arterial blood samples (150 μl per sample) were removed periodically and analyzed for arterial partial pressure of oxygen, carbon dioxide, and pH using an electronic blood gas analyzer (GEM Premier 3000; Instrumentation Laboratory, Lexington, MA). Ventilator adjustments were made if arterial partial pressure of oxygen exceeded 40 to 45 mmHg. Moreover, arterial partial pressure of oxygen was maintained greater than 60 mmHg throughout the experiment by increasing the Fio₂ (22 to 27%). A venous catheter was inserted into the jugular vein for continuous infusion of sodium pentobarbital (10 mg/kg of body weight per hour).

Body temperature was maintained at 37°C by use of a recirculating water heating blanket, and heart rate was monitored via a lead II electrocardiograph. Continuous care during the MV protocol included lubricating the eyes, expressing the bladder, removing airway mucus, rotating the animal, and passively moving the limbs. Animals also received an intramuscular injection of glycopyrrolate (0.04 mg/kg of body weight) every 2 h during MV to reduce airway secretions. On completion of MV, the diaphragm was quickly removed and a section was stored for histochemical analyses, another strip was immediately used for contractile measurements, and the remaining portion was frozen in liquid nitrogen and stored at −80°C for subsequent biochemical analyses.

To prevent MV-induced proteasome activation in the diaphragm, we administered the highly selective proteasome inhibitor, namely epoxomicin (0.55 μg/kg body weight) (Boston Biochem, Boston, MA). This dose was chosen based on previous reports demonstrating the effectiveness of this dose¹⁴  and the efficacy of this dose was confirmed in our preliminary experiments. The inhibitor was dissolved in 60% dimethyl sulfoxide and given intravenously as a bolus immediately before the initiation of MV. Importantly, preliminary work demonstrated that the use of dimethyl sulfoxide as the vehicle had no additional effects on diaphragm contractile dysfunction after 12 h of MV (table 1).

On sacrifice or the completion of the MV period, the entire diaphragm was removed and placed in a dissecting chamber containing a Krebs–Hensleit solution equilibrated with 95% O₂–5% CO₂ gas. A muscle strip, including the tendinous attachments at the central tendon and rib cage was dissected from the midcostal region. The strip was suspended vertically between two lightweight Plexiglas clamps with one end connected to an isometric force transducer (model FT-03; Grass Instruments, Quincy, MA) within a jacketed tissue bath. The force output was recorded via a computerized data-acquisition system (Super Scope II; GW Instruments, Somerville, MA; Apple Computer, Cupertino, CA). Lo (i.e., the length at which peak twitch tension is achieved) was determined by systematically adjusting the length of the muscle using a micrometer while evoking single twitches. Thereafter, all contractile properties were measured isometrically at Lo. To measure maximal isometric twitch force, each strip was stimulated supramaximally with 120-V pulses at 1 Hz, and to measure the force–frequency response, each strip was stimulated supramaximally with 120-V pulses at 15 to 160 Hz. The duration of each train was 500 ms to achieve a force plateau. Contractions were separated by a 2-min recovery period. For comparative purposes, diaphragmatic (bundles of fibers) force production was normalized as specific Po. The total muscle cross-sectional area at right angles to the long axis was calculated by using the following algorithm¹⁵ : Total muscle cross-sectional area (mm²) = [muscle mass/(fiber length × 1.056)], where 1.056 is the density of muscle (in g/cm³). Fiber length was expressed in centimeters measured at Lo.¹⁶ 

Sections from frozen diaphragm samples were cut at 10 μm thickness using a cryotome (Shandon Inc., Pittsburgh, PA) and stained as described previously.⁹  Cross-sectional area was determined using Scion software (National Institutes of Health, Bethesda, MD).

Myofibrillar protein isolation and easily releasable myofilament protein assay were performed with a previously described protocol.^17,18  The final supernatant containing the released myofilaments was collected, and the pellet containing the residual myofibrillar protein fraction was resuspended. Proteins from both fractions were then assayed by the Bradford method (Sigma-Aldrich, St. Louis, MO). Myofibrillar protein (released myofibrillar protein + intact myofibrillar protein) was then corrected for muscle weight and expressed as myofibrillar protein concentration (microgram myofibrillar protein/milligram diaphragm muscle). Released myofilaments were expressed as a percentage of the combined amount of protein in the two fractions.

Protein abundance was determined in diaphragm samples via Western blot analysis. In brief, diaphragm tissue samples were homogenized 1:10 (wt/vol) in 5 mM Tris (pH 7.5) and 5 mM EDTA (pH 8.0) with a protease inhibitor cocktail (Sigma-Aldrich) and centrifuged at 1,500g for 10 min at 4°C. After collecting the resulting supernatant, diaphragm protein content was assessed by the method of Bradford (Sigma-Aldrich). Proteins from the supernatant fraction were separated via polyacrylamide gel electrophoresis. Proteins were transferred to nitrocellulose membranes, blocked in 5% nonfat milk, and incubated with primary antibodies directed against proteins of interest. 4-hydroxynonenal (4-HNE) (Abcam, Cambridge, MA), poly-ubiqutinated proteins (pUB) (Boston Biochem), active calpain (Cell Signaling, Beverly, MA), cleaved caspase-3 (Cell Signaling), spectrin (Santa Cruz, Santa Cruz, CA), muscle ring finger-1 (MuRF1) (ECM Biosciences, Versailles, KY), and atrogin-1 (ECM Biosciences) were assessed. Finally, tubulin (Santa Cruz) protein content was measured to normalize for equal protein loading and transfer. Membranes were developed using autoradiographic film and images of the film were captured and analyzed using the 440CF Kodak Imaging System (Kodak, New Haven, CT).

Total RNA was isolated from muscle tissue with TRIzol Reagent (Life Technologies, Carlsbad, CA) according to the manufacturer’s instructions. RNA content (microgram per milligram muscle weight) was evaluated by spectrophotometry. RNA (5 μg) was then reverse transcribed with the Superscript III First-Strand Synthesis System for reverse transcription-polymerase chain reaction (Life Technologies), using oligo(dT)20 primers and the protocol outlined by the manufacturer.

One microliter of complementary DNA was added to a 25 μl of polymerase chain reaction mixture for real-time polymerase chain reaction using Taqman chemistry and the ABI Prism 7000 Sequence Detection system (ABI, Foster City, CA). Relative quantification of gene expression was performed using the comparative computed tomography method (ABI, User Bulletin No. 2). β-Glucuronidase, a lysosomal glycoside hydrolase, was chosen as the reference gene based on previous work showing unchanged expression with our experimental interventions.^19,20  MuRF1 and atrogin-1/MaFbx microRNA transcripts were assayed using predesigned rat primer and probe sequences commercially available from ABI (Assays-on-Demand).

A section of the ventral costal diaphragm was homogenized 1:10 (vol: vol) in 5 mM Tris-HCL (pH 7.5) and 5 mM EDTA (pH 8.0) and centrifuged at 1,500g for 10 min at 4°C. The cytosolic fraction was centrifuged at 10,000g for 10 min at 4°C, followed by an additional spin of the supernatant at 100,000g for 1 h at 4°C. The in vitro chymotrypsin-like activity of the 20S proteasome was measured fluorometrically using the techniques described in the study by Stein et al.²¹ 

In vitro diaphragmatic specific force production was analyzed using a two-way mixed design group × frequency (3 × 5) ANOVA with repeated measures on frequency. Follow-up tests were conducted using the Bonferroni procedure. For the data on cross-sectional area, we compared the groups at each fiber type (i.e., I, IIa, and IIx/IIb) with one-way ANOVAs. Data for 4-HNE, easily releasable myofilaments, MuRF1, atrogin-1, pUB, 20S proteasome activity, calpain, spectrin breakdown product 145 kDa, caspase-3, and spectrin breakdown product 120 kDa were compared between groups by one-way ANOVA. When the one-way ANOVAs revealed statistically significant differences, a conservative Bonferroni adjustment was performed to identify group differences. Physiological variables (i.e., heart rate, systolic blood pressure, arterial partial pressure of oxygen and carbon dioxide, and arterial pH) were compared using the independent t test. Data points 2SDs different from the average were removed from the analyses. All statistical comparisons were performed using two-tailed tests. Overall significance was established at P value less than 0.05 and adjusted (i.e., P = 0.05/3 = 0.0167) with Bonferroni technique when examining the group main effect means. Data are presented as mean ± SD. All statistical analyses were performed using PASW Statistics 21 (SPSS, Inc., Chicago, IL).

Before the initiation of MV, no significant differences existed in body weight between the experimental groups. Importantly, 12 h of MV did not significantly alter body weight between groups (data not shown). In addition, heart rate and systolic blood pressure were maintained relatively constant during the 12 h of MV, and no significant differences existed between experimental groups in any of these measures at the completion of 12 h of MV (table 2). The arterial partial pressures of oxygen and carbon dioxide and pH were also maintained relatively constant during MV with no significant differences existing between groups (table 2). In addition, the colonic (body) temperature was homeostatic and relatively constant (36° to 37°C) during the MV period. At the completion of the MV protocol, there were no visual abnormalities of the lungs or peritoneal cavity, no visible indication of lung injury, and no evidence of infection, indicating that our aseptic surgical technique was successful.

To determine the role of the UPP in MV-induced diaphragm contractile dysfunction, we measured the in vitro specific force generation of diaphragm muscle strips. Similar to published reports, 12 h of MV resulted in a significant decrease in specific force generation.^2,22,23  Animals treated with epoxomicin were partially protected against MV-induced diaphragm contractile dysfunction. However, diaphragmatic specific force production in the MV + epoxomicin group remained significantly lower compared with that in control animals (fig. 1).

To determine whether the UPP is required for MV-induced diaphragm atrophy, we measured the cross-sectional area of diaphragm myofibers after 12 h of MV. Our results demonstrate that compared with control, MV promotes significant atrophy in fiber type I in diaphragm muscle. Importantly, our data also reveal that inhibition of the 20S proteasome does not prevent this MV-induced diaphragm atrophy (fig. 2).

Oxidative stress has been linked to accelerated rates of proteolysis and diaphragm atrophy during prolonged MV.^7,24  4-HNE is formed during the lipid peroxidation cascade and measurement of 4-HNE–modified protein adducts is an excellent biomarker of oxidative damage in muscle. In this regard, MV resulted in a significant increase in the accumulation of 4-HNE–modified proteins in the diaphragm. However, treatment of animals with epoxomicin failed to inhibit the increase in MV-induced oxidative damage in the diaphragm (fig. 3).

Easily releasable myofilaments were measured as an index of sarcomeric protein release in the diaphragm during MV. Our results reveal that MV produces a significant increase in the percentage of easily releasable myofibrillar proteins in the diaphragm and that inhibiting the 20S proteasome with epoxomicin does not attenuate this release of sarcomeric proteins from the myofibrillar lattice (fig. 4).

During periods of inactivity, the UPP relies on E3 ubiquitin ligases to tag damaged and misfolded muscle proteins, targeting them for degradation. To determine whether inhibition of the 20S proteasome affects muscle E3 ligase expression, we measured the microRNA expression and protein abundance of MuRF1 and atrogin-1/MaFbx. Compared with control, 12 h of MV significantly increased the expression of both MuRF1 and atrogin-1/MaFbx in the diaphragm. However, proteasome inhibition did not prevent the MV-induced increases in the protein expression of either MuRF1 or atrogin-1/MaFbx. However, epoxomicin treatment did result in a significant reduction in MuRF1 microRNA expression compared with MV (fig. 5, A–D). In addition to MuRF1 and atrogin-1/MaFbx, the presence of ubiquitinated proteins can serve as a marker of proteasome activity and changes in protein turnover. Therefore, we measured the levels of ubiquitinated proteins in the diaphragm to determine whether epoxomicin inhibited the formation of poly-ubiquitinated protein conjugates. Our results demonstrate that treatment of MV animals with epoxomicin results in a significant increase in ubiquitin protein concentration in the diaphragm (fig. 6A).

To verify that epoxomicin was effective in preventing proteasome activity, we measured the chymotrypsin-like activity of the 20S proteasome in the diaphragm. Importantly, our results verified that administration of epoxomicin prevents the MV-induced increase in 20S proteasome activity in the diaphragm (fig. 6B).

Given that previous work has shown both calpain and caspase-3 are required for VIDD,^9–11  we determined whether diaphragmatic calpain and/or caspase-3 activity was inhibited by epoxomicin. Similar to previous results, we report that the protein abundance of active calpain-1 and the abundance of the 145-kDa calpain-specific spectrin breakdown product were significantly increased after 12 h of MV in the diaphragm and treatment with epoxomicin did not attenuate this increase in calpain activity (fig. 7). In addition, we measured cleaved (active) caspase-3 and the 120-kDa caspase-specific spectrin breakdown product as markers of caspase-3 activity. MV resulted in a significant increase in caspase-3 activity in the diaphragm and this activity remained increased in MV animals treated with epoxomicin (fig. 8).

Our results reveal that inhibition of 20S proteasome activity is not sufficient to protect the diaphragm against MV-induced diaphragm fiber atrophy, but inhibition of 20S activity does provide partial protection against MV-induced diaphragmatic contractile dysfunction. A detailed discussion of these findings follows.

The ubiquitin–proteasome system is a key proteolytic system in muscle fibers that plays an important role during muscle wasting.^25–27  The total proteasome complex (26S) is comprised a core proteasome subunit (20S) coupled with two regulatory complexes (19S).^28,29  The primary function of this system is to degrade proteins that have been poly-ubiquitinated. In this regard, previous work from our laboratory suggests that proteolytic processing by the UPP is increased in the diaphragm during prolonged MV.³⁰  However, the contribution of this pathway to VIDD remains unknown. Historically, the UPP has been characterized as the dominant proteolytic pathway in disuse muscle atrophy.^26,31  Therefore, we investigated the role that the UPP plays in MV-induced diaphragm atrophy using the 20S proteasome inhibitor epoxomicin. Our results reveal that treatment of mechanically ventilated animals with epoxomicin partially attenuates the MV-induced decrease in diaphragm contractile force production, but does not protect against MV-induced diaphragmatic fiber atrophy. Our interpretation of these findings is that although the UPP is active during MV, this proteolytic system does not play a dominant role in the development of VIDD during the first 12 h of MV.

Protein oxidation can promote protein recognition and degradation by many proteolytic systems (i.e., calpain, caspase-3, and UPP). Indeed, previous work has demonstrated that protein oxidation can enhance proteins susceptibility to degradation by altering the proper folding of proteins resulting in increased recognition and proteolysis of damaged proteins.^29,32,33  Regulatory proteins that influence force production include actin, myosin, tropomyosin, and the troponin complex (reviewed in the study by Smith and Reid³⁴ ). Each of these proteins seems to be sensitive to oxidation and, therefore, to modifications that diminish force production, rendering them potential targets for proteolytic degradation.^29,32,34  In addition to oxidative modification of myofibrillar proteins, reports indicate that oxidative stress promotes the increased expression of proteins necessary for UPP function. Specifically, exposure of myotubes to hydrogen peroxide is sufficient to increase the expression of specific ubiquitin ligases.^35–37  Therefore, increased reactive oxygen species production seems to promote muscle protein degradation via the UPP. In this regard, we measured the abundance of 4-HNE–modified proteins to determine whether inhibition of 20S proteasome activity results in a reduction of MV-induced oxidative damage in the diaphragm. In agreement with numerous reports, our results confirm that prolonged MV increases diaphragmatic oxidative stress,^7,22,38  and treatment of animals with epoxomicin does not attenuate the MV-induced accumulation of 4-HNE in the diaphragm.

Mechanical ventilation–induced diaphragm atrophy results from an imbalance in the rate of muscle protein synthesis and degradation, with increased protein degradation playing the dominant role.^39,40  In this regard, myofibrillar proteins constitute 50 to 60% of muscle protein and are degraded at a faster rate than other classes of muscle proteins during disuse muscle atrophy.⁴¹  In addition, myofibrillar proteins have also been shown to be the principal protein target of the UPP.⁴²  The degradation of myofibrillar proteins is a process that requires the coordination of several proteolytic systems, including the calpain, caspase-3, and the UPP. Specifically, it is predicted that the UPP is unable to cleave intact sarcomeric proteins, and that release of these myofilaments from the sarcomeric lattice is required for UPP degradation of actin and myosin.^40,43 

As discussed previously, the breakdown of muscle proteins by the UPP requires ubiquitination of the targeted proteins. This poly-ubiquitination is a three-step process in which activation of specific protein ligases leads to the activation and conjugation of ubiquitin to protein substrates followed by the degradation of poly-ubiquitin-tagged proteins by the proteasome. Specifically, ubiquitin monomers are activated by E1 or ubiquitin-activating proteins and transferred to E2 ubiquitin-conjugating proteins. Finally, in coordination with the E3 ligases, ubiquitin is attached to the protein substrate.⁴⁴  In reference to substrate recognition and degradation, expression of E2 and E3 ligases determines the specificity of the UPP and transcriptional levels of these proteins strongly correlates with muscle catabolism.^40,43,44  Furthermore, expression of the muscle-specific E3 ligases MuRF1 and atrogin-1/MaFbx has been reported to be essential for skeletal muscle atrophy.^25–27  Similarly, Bodine et al.²⁶  demonstrated that these ligases are critical regulators of muscle atrophy and the deletion of either ligase during denervation is sufficient to protect muscle size and/or mass.

Although the importance of both MuRF1 and atrogin-1/MaFbx has been demonstrated in other models of disuse muscle atrophy, the role that these ligases play in VIDD remains unknown. In regard to E3 ligase expression, the current results demonstrate that 12 h of MV is sufficient to increase the transcription of both MuRF1 and atrogin-1/MaFbx in the diaphragm.^22,23  However, treatment of mechanically ventilated animals with epoxomicin did not reduce the expression of these E3 ligases in the diaphragm. Our data also demonstrate that 12 h of MV increases protein ubiquitination in the diaphragm and the levels of ubiquitinated proteins in the diaphragm tended to be higher in the MV animals treated with epoxomicin. This increased protein ubiquitination in MV animals is likely due to an accumulation of poly-ubiquitinated proteins resulting from increased MuRF1 and atrogin-1/MaFbx in the diaphragm. Furthermore, the increase in ubiquitinated proteins in the diaphragms of MV animals treated with epoxomicin is expected because these tagged proteins cannot be degraded by the UPP because of epoxomicin-induced inhibition of 20S proteasome activity.

Finally, although it has been proposed that the UPP is the primary proteolytic system essential for the degradation of myofibrillar proteins, it has also been demonstrated that the activity of other proteases play an obligatory role in diaphragm protein breakdown during prolonged MV.^7,9–11  Specifically, calpain and caspase-3 activation both seem to play a required role in MV-induced diaphragm muscle turnover.^9–11  In this regard, it has been hypothesized that actomyosin complexes and other myofilaments must first be released from the sarcomere, unfolded and ubiquitinated for UPP degradation.^45–49  This suggests that the rate-limiting step in muscle contractile protein degradation is the processing required for UPP proteolysis. Furthermore, evidence indicates that both calpain and caspase-3 can promote actomyosin disassociation.^45,49,50  Therefore, it has been speculated that activation of calpain and caspase-3 is required for UPP degradation of myofibrillar proteins.^32,50  A long-standing rationale for the prediction that the UPP is essential for disuse muscle atrophy is that the two major contractile proteins actin and myosin are reported to be weak substrates for calpain and caspase-3.⁴⁵  However, recent evidence indicates that oxidized myofibrillar proteins can be directly degraded by both calpain and caspase-3 which demonstrates that the UPP may not be required for the degradation of myofibrillar proteins in skeletal muscles exposed to oxidative stress.³²  Indeed, work from our laboratory demonstrates that prolonged MV results in oxidation of both actin and myosin in the diaphragm and that oxidation greatly increases the susceptibility of actin and myosin along with other skeletal muscle myofibrillar proteins to degradation by calpain and caspase-3.^32,51  Therefore, during prolonged MV where myofibrillar proteins in the diaphragm are exposed to oxidizing conditions, activation of the UPP system may not be required for the rapid atrophy that occurs within the first 12 h of prolonged MV.

This study provides the first evidence that the UPP plays a limited role in the development of VIDD during the first 12 h of MV. Indeed, although the UPP is activated in the diaphragm during MV, inhibition of 20S proteasome activity is not sufficient to protect the diaphragm against MV-induced contractile dysfunction and atrophy during the first 12 h of MV. Therefore, these results along with previous reports suggest that activation of calpain and caspase-3 plays a key role in the development of VIDD.^32,45,50  Hence, although the proteasome has been considered to be the primary protease responsible for myofibrillar protein degradation during locomotor muscle atrophy, its contribution to VIDD during the first 12 h of ventilator support seems to be limited. In regard to the impact of epoxomicin on diaphragm contractile function at high stimulation frequencies, it is feasible that the autophagy/lysosomal system can degrade poly-ubiquitinated proteins. Therefore, it is possible that activation of the autophagy/lysosomal system is upregulated in when UPP activity is inhibited. This is a testable hypothesis that is worthy of future experimentation.

This work was supported by the National Institutes of Health (Bethesda, Maryland) award (RO1 HL780839; to S.K.P.).

The authors declare no competing interests.