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FASEB J. Jan 2011; 25(1): 99–110.
PMCID: PMC3005430

Toll-like receptor 4 mediates lipopolysaccharide-induced muscle catabolism via coordinate activation of ubiquitin-proteasome and autophagy-lysosome pathways

Abstract

Cachectic muscle wasting is a frequent complication of many inflammatory conditions, due primarily to excessive muscle catabolism. However, the pathogenesis and intervention strategies against it remain to be established. Here, we tested the hypothesis that Toll-like receptor 4 (TLR4) is a master regulator of inflammatory muscle catabolism. We demonstrate that TLR4 activation by lipopolysaccharide (LPS) induces C2C12 myotube atrophy via up-regulating autophagosome formation and the expression of ubiquitin ligase atrogin-1/MAFbx and MuRF1. TLR4-mediated activation of p38 MAPK is necessary and sufficient for the up-regulation of atrogin1/MAFbx and autophagosomes, resulting in myotube atrophy. Similarly, LPS up-regulates muscle autophagosome formation and ubiquitin ligase expression in mice. Importantly, autophagy inhibitor 3-methyladenine completely abolishes LPS-induced muscle proteolysis, while proteasome inhibitor lactacystin partially blocks it. Furthermore, TLR4 knockout or p38 MAPK inhibition abolishes LPS-induced muscle proteolysis. Thus, TLR4 mediates LPS-induced muscle catabolism via coordinate activation of the ubiquitin-proteasome and the autophagy-lysosomal pathways.—Doyle, A., Zhang, G., Abdel Fattah, E. A., Eissa, N. T., Li, Y.-P. Toll-like receptor 4 mediates lipopolysaccharide-induced muscle catabolism via coordinate activation of ubiquitin-proteasome and autophagy-lysosome pathways.

Keywords: autophagosomes, atrogin-1/MAFbx, p38 MAPK, 3-methyladenine, lactacystin

Progressive loss of muscle mass and strength (muscle wasting) is the major component of cachexia, a frequent complication of many inflammatory conditions, including sepsis, AIDS, cancer, congestive heart failure (CHF), chronic obstructive pulmonary disease (COPD), renal failure, and diabetes, due largely to excessive muscle protein degradation. Cachectic muscle wasting is distinct from muscle atrophy caused by starvation, aging, primary depression, malabsorption, and hyperthyroidism and is associated with increased morbidity and mortality (1). Despite recent progress in research on this debilitating and often lethal condition, the understanding of the pathogenesis of cachectic muscle wasting is still limited. To a different extent, two major cellular proteolytic systems, the ubiquitin-proteasome pathway (UPP) and the autophagy-lysosomal pathway (ALP), are implicated in various forms of muscle atrophy. The UPP is thought to degrade myofibrillar proteins and most soluble proteins (2), while the ALP is capable of degrading most cytoplasmic constituents, including ubiquitinated protein aggregates and organelles (3, 4). The UPP mediates cachectic muscle wasting associated with aforementioned inflammatory diseases, as well as physiological muscle atrophy due to starvation, denervation, or disuse (5). Expression of muscle-enriched ubiquitin ligase atrogin-1/MAFbx and MuRF1 that are rate-limiting for the activity of the UPP (6, 7) are up-regulated by hypophosphorylated FoxO family of transcription factors resulting from suppressed activity of the PI3K/AKT signaling pathway in starvation, denervation, and disuse (810). Paradoxically, TNF-α, a major inflammatory cytokine that mediates cachectic muscle wasting (11) activates AKT in muscle cells (12), which would inactivate FoxOs leading to down-regulation of atrogin-1/MAFbx and MuRF1. The capability of TNF-α to up-regulate atrogin-1/MAFbx and MuRF1 has been attributed to its activation of p38 MAPK (13) and NF-κB (14), respectively. Yet, anti-TNF-α strategies have yielded mixed results in the intervention of cachectic muscle wasting (15, 16), perhaps due to the fact that other inflammatory cytokines that are elevated in inflammatory conditions such as IL-1 and IL-6 also activate the UPP via similar mechanisms (17, 18). Therefore, to effectively intervene in the up-regulation of atrogin-1/MAFbx and MuRF1 by inflammatory cytokines, the production of the relevant cytokines by the immune cells should be targeted. On the other hand, the ALP has been shown to also mediate physiological muscle atrophy (starvation and denervation), where activated FoxO3 (hypophosphorylated) up-regulates autophagosome formation (19, 20). Therefore, it has been proposed that the UPP and the ALP serve complementary roles in degrading distinct cellular constituents and might be activated coordinately by common signaling mechanisms (21). Notwithstanding, whether the ALP mediates cachectic muscle wasting, and if so, how inflammatory mediators activate the ALP is unknown.

Sepsis is an inflammatory condition that causes a severe and rapid loss of body protein, much of which originates from skeletal muscle (22). Many studies have provided evidence that muscle atrophy in sepsis is primarily the result of increased protein breakdown (23, 24) via the ubiquitin–proteasome pathway (25, 26). Indeed, both atrogin-1/MAFbx and MuRF1 are up-regulated in the muscle of septic models induced by either cecal ligation and puncture (27) or lipopolysaccharide (LPS) administration (28). LPS-mediated endotoxemia, which induces a number of catabolic factors in sepsis (e.g., TNF-α, IL-1, IL-6, glucocorticoids, and glucocorticoid receptors), also suppresses the signaling of anabolic factor IGF-I (29). LPS may also mediate muscle wasting in other inflammatory conditions since elevated serum LPS levels have been found in patients with such diseases as cancer (30), chronic heart failure (31), and type 2 diabetes (32), which frequently induce muscle wasting. However, the key event that coordinates the complex catabolic signaling of LPS leading to muscle loss has not been identified.

LPS is a potent agonist of Toll-like receptor 4 (TLR4) (33). We postulated that an upstream mediator of inflammatory response serves as a master regulator of inflammation-induced muscle wasting by regulating the activity of both the UPP and the ALP, and that TLR4 is likely the potential master regulator. Toll-like receptors are a family of pattern recognition receptors that have emerged as important mediators of innate immunity (34, 35). Among the TLR family, TLR4 is most likely to have important roles in regulating the events that lead to excessive muscle protein degradation for the following reasons. First, TLR4 plays a central role in macrophage-mediated innate immunity by activating such inflammatory signaling molecules as p38 MAPK and NF-κB, which, in turn, up-regulate macrophage production of proinflammatory cytokines, including TNF-α, IL-1, and IL-6 (36, 37). Systemically elevated proinflammatory cytokines are thought to mediate muscle wasting via up-regulating atrogin-1/MAFbx and MuRF1 (38). Second, expressed in muscle cells (39, 40), TLR4 could mediate the activation of p38 MAPK and NF-κB in muscle cells, and thus up-regulate atrogin-1/MAFbx and MuRF1 directly independent of humoral factors. Third, a newly found mechanism of TLR4 regulation of immunity involves its activation of autophagy in macrophages. TLR4 activation leads to increased autophagosome formation via a p38 MAPK-dependent mechanism (41). A TLR4-mediated signaling pathway that activates autophagy may also exist in muscle cells. Fourth, macrophage-released proinflammatory cytokines may enhance the activation of autophagy via activating p38 MAPK in muscle cells. Thus, TLR4 could mediate inflammatory muscle wasting via the activation of both the UPP and the ALP. In the present study, we tested the hypothesis that TLR4 is a master regulator of muscle wasting induced by endotoxemia via coordinate activation of the UPP and the ALP. The data that we obtained support our hypothesis and reveal a surprisingly critical role of autophagy in TLR4-mediated muscle wasting.

MATERIALS AND METHODS

Myogenic cell culture

Murine C2C12 myoblasts (American Type Culture Collection, Manassas, VA, USA) were cultured in growth medium (DMEM supplemented with 10% FBS) at 37°C under 5% CO2. At 85% confluence, myoblast differentiation was induced by incubation for 96 h in differentiation medium (DMEM supplemented with 4% heat-inactivated horse serum) to form myotubes, as described previously (13). LPS dissolved in saline was added to culture medium every 24 h for up to 48 h, and 10 μM SB202190 dissolved in DMSO (0.1% final concentration of DMSO, which did not alter the parameters we measured) was added 30 min prior to LPS when indicated. Chloroquine dissolved in saline was used in 50 μM final concentration when indicated.

Animal use

Experimental protocols were approved in advance by the University of Texas Health Science Center at Houston Animal Welfare Committee. Muscle catabolism was induced by intraperitoneal (i.p.) injection of LPS (1 mg/kg; Sigma-Aldrich, St. Louis, MO, USA) or an equal volume of vehicle (PBS) to male adult C57BL/6 mice (8 wk of age) purchased from Jackson Laboratory (Bar Harbor, ME, USA), GFP-LC3 transgenic mice (42) or TLR4−/− mice (43) in C57BL/6 background. When indicated, mice were preconditioned by daily i.p. injection of SB202190 (5 mg/kg; Sigma-Aldrich), 3-methyladenine (3-MA, 10 mg/kg; Sigma-Aldrich), or vehicle (PBS containing 0.1% DMSO) for 4 consecutive d prior to LPS injection. At indicated times, tibialis anterior (TA) and extensor digitorum longus (EDL) muscles were collected from the mice immediately after euthanasia.

Transfection

Plasmid encoding GFP-LC3 fusion protein (41) was transfected into C2C12 myoblasts at ~50% confluence by using deacylated polyethylenimine (PEI) 22000 (44), a gift from Dr. Guangwei Du (University of Texas Health Science Center, Houston, TX, USA). The on-target smart pool siRNA specific for TLR4 and control siRNA purchased from Dharmacon (Lafayette, CO, USA) and Ambion (Austin, TX, USA), respectively, were introduced into C2C12 myoblasts by electroporation (5 μg per reaction) using the Nucleofector system (Lonza, Basel, Switzerland), according to manufacturer's protocol. On the next day, differentiation was induced to form myotubes.

Adenovirus transduction

Ad5 cytomegalovirus encoding MKK6bE (45), a constitutively active mutant of mitogen-activated protein kinase kinase (MKK6), or green fluorescence protein (GFP; prepared by The Vector Development Core of Baylor College of Medicine) were used at 800 MOI to transduce C2C12 cells that had been incubated in differentiation medium for 48 h. Cells were further incubated for 48 h to allow recombinant protein expression and myotube formation before experimenting.

Western blot analysis

Muscle homogenate and myotube lysate were prepared in 4°C RIPA buffer containing 50 mM Tris-HCl (pH 7.5), 150 mM NaCl, 2 mM EDTA, 1% Nonidet P-40, 0.1% SDS, 2 mM phenylmethylsulfonylfluoride (PMSF), 0.1% sodium deoxycholate, 1 mM NaF; 1:100 protease inhibitor cocktail (Sigma-Aldrich) containing AEBSF, pepstatin A, E-64, bestatin, leupeptin and aprotinin; and 1:100 phosphatase inhibitor cocktail (Sigma-Aldrich) containing sodium vanadate, sodium molybdate, sodium tartrate, and imidazole. Muscle homogenate was then sonicated for 5 s. Insoluble muscle or cellular debris was removed by centrifugation at 16,000 g (4°C). The supernatant was collected, and protein concentration was determined using the Bio-Rad protein assay (Bio-Rad, Richmond, CA, USA) with BSA as a standard. Western blot analysis was performed as described previously (13). Densitometry analysis of detected protein bands was performed using Kodak 1D Image Analysis software (Eastman Kodak, Rochester, NY, USA). Antibodies to total and phosphorylated p38 (T181/Y182), AKT (S473), and FoxO1 (T24)/FoxO3a (T32) were from Cell Signaling (Beverly, MA, USA). Antibodies to TLR4 and MuRF1 were from Santa Cruz Biotechnology (Santa Cruz, CA, USA), antibody for atrogin-1/MAFbx was from ECM Biosciences (Versailles, KY, USA). Two antibodies against LC3B from Cell Signaling were used in different experiments, one detects both LC3-I and LC3-II (catalog no. 2775S), and the other detects LC3II only (3866S), as indicated in figure legends.

Real-time PCR

Total RNA was extracted from myotubes or muscle using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) and dissolved in diethylpyrocarbonate-treated water. Total RNA was reverse transcribed to cDNA using 50 ng of total RNA in a 20-μl reaction mixture containing 200 U of RevertAid M-MuLV reverse transcriptase (Fermentas, Burlington, ON, Canada). Quantitative PCR was performed using the SYBR green method in a total reaction mixture of 20 μl containing 0.2 U of JumpStart Taq polymerase (Sigma-Aldrich), 10 pmol of each forward and reverse primer, and 1 μl cDNA in a MyiQ real-time PCR detection system (Bio-Rad) in triplicate. Sequences of specific primers are Atrogin1/MAFbx, F 5′-CACATTCTCTCCTGGAAGGGC-3′, R 5′-TTGATAAAGTCTTGAGGGGAAAGTG-3′; MuRF1, F 5′-CACGAAGACGAGAAGATCAACATC-3′, R 5′-AGCCCCAAACACCTTGCA-3′; and GAPDH, F 5′-CATGGCCTTCCGTGTTCCTA-3′, R 5′-GCGGCACGTCAGATCCA-3′. Detected levels of target mRNAs were calculated using the ΔΔCt method and normalized to GAPDH in arbitrary units.

Fluorescence microscopy

Cryosections of TA muscle (10 μm) expressing GFP-LC3 and C2C12 myotubes transfected with GFP-LC3 were examined using a Zeiss Axioskop 40 microscope and a Zeiss Axiocam MRM camera system controlled by Axiovision Release 4.6 imaging software (Carl Zeiss, Oberkochen, Germany). Acquired images were edited using Photoshop software (Adobe Systems, San Jose, CA, USA). Anti-myosin heavy chain (MHC; MF20; Development Studies Hybridoma Bank, University of Iowa, Iowa City, IA, USA) and FITC-conjugated secondary antibody were used for staining myotubes.

Measurement of myotube diameter

Myotube diameter was measured using the method of Menconi et al. (46) with modifications. Briefly, myotube cultures were photographed under a phase-contrast microscope at ×40 after indicated treatment. The diameters were measured in a total of 100 myotubes from ≥10 random fields using computerized image analysis (Scion Image, Frederick, MD, USA). Myotubes were measured at 3 points along their length. The measurements were conducted in a masked fashion, and results were expressed as a percentage of the diameter in the control group.

Tyrosine release assay

Tyrosine release was measured using a protocol modified from Fulks et al. (47). Excised mouse EDL was preincubated for 30 min at 37°C in Krebs Henseleit buffer (120 mM NaCl, 4.8 mM KCl, 25 mM NaHCO3, 2.5 mM CaCl2, 1.2 mM KH2PO4, and 1.2 mM MgSO4, pH 7.4, supplemented with 5 mM glucose, 5 mM HEPES, 0.1% BSA, 0.17 mM leucine, 0.20 mM valine, 0.10 mM isoleucine, 0.1 U/ml insulin, and 0.5 mM cycloheximide; chemicals from Sigma-Aldrich). The buffer was saturated with 95% O2-5% CO2. EDL was then transferred into fresh buffer and incubated for 2 h. Tyrosine released into the buffer was determined by a fluorometric method established by Waalkes and Udenfriend (48). Briefly, 0.5 ml of 30% trichloroacetic acid was added to 2 ml of buffer collected from tyrosine release assay, after 10 min incubation at room temperature, the mixture was centrifuged. Two milliliters of the deproteinized supernatant was then mixed with 1 ml each of nitrosonaphthol reagent and nitric acid reagent, and incubated at 55°C for 30 min. After cooling to room temperature, 10 ml of ethylene dichloride was added and mixed vigorously to extract the unchanged nitrosonaphthol reagent. After centrifugation, the aqueous top layer was transferred to a cuvette and read in a TD-700 fluorometer (Turner Designs, Sunnyvale, CA, USA). Fluorescence of the tyrosine derivative resulting from its activation at 460 nm was measured at 570 nm. Concentration of tyrosine derivative was calculated using a standard curve. Rate of tyrosine release was normalized to the weight of the muscle as nanomoles of tyrosine per gram of muscle in 2 h.

Statistical analysis

Data were analyzed with Student's t test or 1-way ANOVA using SigmaStat software (Systat Software, San Jose, CA, USA). A value of P < 0.05 was considered to be statistically significant. All experiments were repeated ≥3 times, and data are presented as means ± se.

RESULTS

LPS induces myotube atrophy independent of the AKT/FoxO signaling pathway

To test our hypothesis, we carried out in vitro evaluation of the response to LPS by C2C12 myotubes that are known to express TLR4 (40). C2C12 myotubes that were incubated with LPS for 48 h underwent a dose-dependent loss of myofibrillar protein MHC, as detected by Western blot analysis (Fig. 1A). Immunofluorescence staining of the myotubes with an MHC-specific antibody revealed that exposure to 100 ng/ml LPS reduced the diameter of myotubes (Fig. 1B), while the number of myotubes remained unchanged. Thus, LPS can induce myotube atrophy via its direct action on muscle cells without the involvement of immune cells, the major source of inflammatory cytokines.

Figure 1.
LPS induces myotube atrophy. A) LPS induces myotube MHC loss. C2C12 myotubes were treated with saline or 10 to 1000 ng/ml LPS for 48 h. MHC content in cell lysate was evaluated by Western blot analysis. B) LPS reduces myotube size. C2C12 myotubes were ...

LPS has been shown to activate AKT (49) and p38 MAPK (41) in some cultured nonmuscle cells. AKT activation in muscle cells inhibits proteolysis via inactivating FoxO transcription factors, resulting in down-regulation of autophagosome formation (19, 20), as well as expression of ubiquitin ligase atrogin1/MAFbx and MuRF1 (8, 9). On the other hand, activation of p38 MAPK mediates atrogin1/MAFbx up-regulation by inflammatory cytokine TNF-α in muscle cells (13) and autophagosome formation in macrophages (41). To understand the role of AKT and p38 MAPK in mediating LPS-induced myotube atrophy, we evaluated activity of AKT and p38 MAPK in LPS-treated C2C12 myotubes. We observed that LPS activated both AKT and p38 MAPK within 30 to 60 min (Fig. 2A). As expected, the activation of AKT resulted in hyperphosphorylation of FoxO1 and FoxO3a (Fig. 2A), which would render these transcription factors inactive due to sequestration in the cytoplasm away from their target genes (50). The observed activation of AKT and inactivation of FoxO1/3a by LPS would down-regulate the activity of the UPP and ALP and thus could not explain the myotube atrophy observed above. To investigate whether p38 MAPK mediates LPS induction of myotube atrophy, we observed that pretreatment of myotubes with p38 MAPK inhibitor SB202190 blocked myotube atrophy induced by LPS (Fig. 2B) without altering the number of myotubes. Conversely, adenovirus-mediated overexpression of MKK6bE, which directly activates p38 MAPK (45), resulted in p38 MAPK activation, MHC loss (Fig. 2C) and myotube atrophy (Fig. 2D). Thus, p38 MAPK is a key mediator of LPS-induced myotube atrophy.

Figure 2.
LPS-induced myotube atrophy is mediated by p38 MAPK. A) LPS activates AKT and p38 MAPK in myotubes. C2C12 myotubes were treated with 100 ng/ml LPS for indicated periods. Cell lysate was analyzed for phosphorylation of p38 MAPK (T180/Y182), AKT (S473), ...

TLR4 mediates LPS-induced myotube atrophy via simultaneous activation of the UPP and the ALP

Next, we evaluated whether TLR4/p38 MAPK signaling mediates LPS up-regulation of ubiquitin ligase expression and, potentially, autophagosome formation in C2C12 myotubes. LPS treatment rapidly up-regulated atrogin1/MAFbx mRNA in 1 h, and the effect peaked in 2 h. On the other hand, MuRF1 mRNA that is known not to be influenced by p38 MAPK activity (51) was not up-regulated until 3 h of LPS treatment (Fig. 3A). Preincubation of myotubes with p38 MAPK inhibitor SB202190 reversed LPS up-regulation of atrogin-1/MAFbx mRNA. In addition, Western blot analysis confirmed a similar increase in atrogin1/MAFbx protein level in response to LPS treatment, and blockade of the LPS effect by SB202190 (Fig. 3B). To verify whether p38 MAPK activation up-regulates atrogin1/MAFbx, MKK6bE was overexpressed in myotubes to activate p38 MAPK. Overexpressed MKK6bE up-regulated atrogin-1/MAFbx, which was blocked by SB202190. In contrast, overexpressed MKK6bE did not alter MuRF1 expression (Fig. 3C). Thus, p38 MAPK activation is necessary and sufficient for the up-regulation of atrogin-1/MAFbx expression. To evaluate the effect of LPS on autophagosome formation in myotubes, the increased formation of LC3-II, a biochemical indication of induction of autophagosome formation (52), was evaluated by Western blot analysis. An increase in LC3-II level started in 2 h of LPS treatment and peaked at 8 h. As a positive control, lysosome inhibitor chloroquine was incubated with myotubes, which resulted in an increase in LC3-II accumulation (Fig. 3D). In addition, LPS-induced LC3-II increase was blocked by preincubation of myotubes with SB202190 (Fig. 3E). To verify LPS effect on the formation of autophagosomes, a plasmid encoding GFP-LC3 (41) was expressed in myotubes, and localization of GFP-LC3 was monitored with fluorescence microscopy. LPS treatment of myotubes stimulated autophagosome formation, as indicated by the appearance of punctate structures containing GFP-LC3, which was blocked by SB202190 pretreatment (Fig. 3F). Furthermore, overexpression of MKK6bE increased LC3-II levels (Fig. 3G). Thus, p38 MAPK mediates LPS up-regulation of autophagosome formation, and p38 MAPK activation is sufficient to up-regulate autophagosome formation. These data reveal that LPS coordinately activates the UPP and the ALP in myotubes via the activation of p38 MAPK.

Figure 3.
LPS up-regulates atrogin-1/MAFbx expression and autophagosome formation in myotubes via the activation of p38 MAPK. A) LPS up-regulates the expression of atrogin-1/MAFbx and MuRF1 in myotubes. C2C12 myotubes were treated with saline or 100 ng/ml LPS for ...

To verify whether TLR4 mediates LPS activation of the UPP and the ALP, TLR4-specific siRNA was employed to knock down TLR4 expression in C2C12 myotubes. TLR4 knockdown blocked LPS activation of p38 MAPK and AKT (Fig. 4A), LPS up-regulation of both ubiquitin ligases (Fig. 4B) and autophagosomes (Fig. 4C), and LPS-induced myotube atrophy (Fig. 4D). Therefore, TLR4 indeed mediates LPS-induced myotube atrophy via coordinate activation of the UPP and the ALP.

Figure 4.
TLR4 mediates LPS activation of p38 MAPK, UPP, ALP, and myotube atrophy. C2C12 myoblasts were transfected with TLR4-specific siRNA or control siRNA. After differentiation, myotubes were treated with 100 ng/ml of LPS or saline for various periods for the ...

TLR4 mediates muscle atrophy in mice via coordinate activation of the UPP and the ALP

The in vivo effects of LPS on muscle catabolism involve the activation of TLR4 in multiple systems and the ensuing interactions of a complex array of humoral factors that are not present in the myotube cultures. Thus, it is necessary to evaluate the significance of TLR4 in mediating muscle proteolysis through the two proteolytic pathways in vivo. We first evaluated whether LPS up-regulates muscle autophagosome formation in mouse muscle. We observed that in TA of transgenic GFP-LC3 mice (42), LPS administration up-regulated autophagosome formation, as indicated by the formation of punctate structures containing GFP-LC3 (Fig. 5A). Then, we evaluated the contribution of the UPP and the ALP to mouse muscle protein degradation induced by LPS using the tyrosine release assay (47). To evaluate the contribution of the UPP to LPS-induced muscle protein degradation, tyrosine release from excised EDL of mice that had been administered LPS was measured with or without the presence of the specific proteasome inhibitor lactacystin. Tyrosine release from EDL of LPS-treated mice increased by 40%, as compared with saline-treated control, and lactacystin reduced LPS-induced tyrosine release by 53% (Fig. 5B). To evaluate the contribution of the ALP to LPS-induced muscle protein degradation, mice were pretreated with 3-MA, which inhibits autophagosome formation (53) or saline (control) prior to LPS administration. Surprisingly, 3-MA completely abolished LPS-induced tyrosine release (Fig. 5C). LPS up-regulation of autophagosome formation and its inhibition by 3-MA in TA were verified by Western blot analysis of LC3-II levels (Fig. 5D). It was also verified that 3-MA did not alter LPS up-regulation of atrogin-1/MAFbx and MuRF1 mRNA (data not shown). These data indicate that autophagy may contribute more significantly than the UPP to LPS-induced muscle protein degradation in vivo.

Figure 5.
LPS induces muscle atrophy in mice via the activation of the ALP, as well as the UPP. A) LPS administration up-regulates autophagosome formation in mouse TA. LPS (1 mg/kg) or saline was injected (i.p.) to male GFP-LC3 transgenic mice (8 wk of age). In ...

Next, whether TLR4 mediates LPS-induced muscle catabolism in vivo was investigated by utilizing a line of TLR4−/− mice (43). Unlike in wild-type mice, LPS failed to stimulate tyrosine release in the EDL of TLR4−/− mice (Fig. 6A), while its up-regulation of the mRNA (Fig. 6B) and protein (Fig. 6C) of ubiquitin ligases and autophagy formation (Fig. 6D) in TA were abolished. Therefore, LPS induces muscle catabolism by activating TLR4, and TLR4 is a master regulator of the activity of the UPP and the ALP.

Figure 6.
TLR4 mediates LPS-induced muscle atrophy in mice by activating the UPP and the ALP. LPS (1 mg/kg) or saline was injected (i.p.) to TLR4−/− or wild-type (WT) male mice (8 wk of age). TA and EDL were collected at 18 h after injection for ...

We previously observed that unlike in myotubes, LPS administration to mice down-regulates AKT activity in muscle, although muscle catabolism is correlated to p38 MAPK activity, not AKT activity (51). Therefore, it is necessary to determine whether p38 MAPK has a predominant role in vivo in mediating TLR4 activation of the UPP and the ALP in muscle. In GFP-LC3 transgenic mice, LPS up-regulation of autophagosome formation was blocked by pretreatment with p38 MAPK inhibitor SB202190 (Fig. 7A). Examination of LC3-II levels confirmed that LPS-induced LC3-II increase is p38 MAPK dependent (Fig. 7B). In addition, LPS up-regulation of atrogin-1/MAFbx, but not of MuRF1, was attenuated by SB202190 pretreatment (Fig. 7C). Consequently, SB202190 effectively abolished LPS-induced tyrosine release from EDL (Fig. 7D). These data confirm that p38 MAPK is a key mediator of TLR4 activation of the UPP and the ALP in vivo. Therefore, p38 MAPK is a potential therapeutic target for TLR4-mediated muscle wasting.

Figure 7.
p38 MAPK mediates TLR4 activation of the UPP and the ALP in mouse muscle. Wild-type or GFP-LC3 transgenic male mice (8 wk of age) were pretreated with daily injection (i.p.) of SB202190 (5 mg/kg) or vehicle for 4 d, followed by LPS (1 mg/kg) or saline ...

DISCUSSION

The present study demonstrates for the first time that TLR4 acts as a master regulator of muscle proteolysis via coordinate activation of the UPP and ALP, and that p38 MAPK plays a key role in mediating the catabolic signaling of TLR4. The simultaneous activation of these two proteolytic pathways by TLR4 indicates that LPS-induced muscle atrophy results from loss of not only myofibrillar proteins but also organelles, including mitochondria. Myofibrillar proteins comprise 60–70% of cellular protein in adult muscle. Our observation that proteasome inhibitor lactacystin inhibits about half of LPS-induced muscle protein degradation appears consistent with the notion that the UPP mediates the degradation of myofibrillar proteins (2). On the other hand, our observation that autophagy inhibitor 3-MA completely prevents LPS-induced muscle protein degradation is surprising and suggests that the ALP mediates the degradation of myofibrillar, as well as nonmyofibrillar proteins. The capability of the ALP to degrade most cytoplasmic constituents, including soluble proteins and ubiquitinated protein aggregates (3, 4), might contribute to the degradation of myofibrillar proteins. In addition, autophagy-mediated loss of mitochondria would affect muscle energy balance, which may contribute to the muscle weakness seen in cachexia. Thus, the ALP appears to play a more significant role than the UPP in TLR4-mediated muscle wasting.

Using C2C12 myotubes, we showed that LPS directly stimulates muscle catabolism through activating the two cellular proteolytic systems independent of humoral factors. The rapid activation of p38 MAPK by LPS within 30 min suggests that the LPS actions result from direct activation of TLR4 in myotubes instead of an indirect effect through stimulating cytokine synthesis by myotubes. Although atrogin-1/MAFbx expression is up-regulated within 1 h of LPS treatment, MuRF1 expression responded differently, which suggests the two genes are up-regulated by LPS via a different mechanism. Given that MuRF1 up-regulation is dually mediated by the inactivation of AKT (9) and the activation of NF-κB (14), and that TLR4 mediates the activation of NF-κB in C2C12 myotubes (40), it is likely that LPS activation of NF-κB mediates the up-regulation of MuRF1 observed at a later time (3 h). Interestingly, up-regulation of myotube atrogin-1/MAFbx expression and autophagosome formation by LPS requires activation of p38 MAPK, while AKT is simultaneously activated. The latter is supposed to have the opposite effect on muscle protein degradation, thus, p38 MAPK has a predominant role in mediating muscle catabolism induced by LPS. Furthermore, the observation that direct activation of p38 MAPK by overexpressed MKK6bE up-regulates atrogin-1/MAFbx expression and autophagosome formation indicates that p38 MAPK activation is not only necessary but also sufficient for the activation of the UPP and the ALP. By the same token, other inflammatory activators of p38 MAPK, such as TNF-α, IL-1, and reactive oxygen species (ROS) (12, 13, 54) may also mediate muscle catabolism by activating the ALP, as well as the UPP.

The observation that LPS-induced tyrosine release is completely abolished in TLR4 knockout mice confirms that TLR4 is indeed the key mediator of LPS-induced muscle catabolism. On the other hand, because the TLR4-knockout mice used in the present study were not tissue-specific, we could not separate the direct action of muscle TLR4 activation from the indirect action of macrophage TLR4 activation on the UPP and the ALP activity. Our in vivo data indicate that unlike in physiological muscle atrophy (starvation and denervation), in which the AKT/FoxO signaling pathway mediates the coordinate activation of the UPP and the ALP (19, 20), in a pathological condition (endotoxemia), TLR4 activation simultaneously stimulates the activity of the two muscle proteolytic pathways via p38 MAPK. The observation that p38 MAPK inhibitor SB202190 blocks TLR4-mediated activation of the two muscle proteolytic pathways further confirms a predominant role of p38 MAPK in the up-regulation of muscle protein degradation in this catabolic model. It appears that one of the two independent signaling pathways involving AKT and p38 MAPK in muscle cells coordinately activate the UPP and the ALP, depending on whether the stimuli are physiological (such as starvation and denervation) or pathological (such as endotoxemia).

It was previously shown in macrophages that LPS-induced autophagy is regulated through a TRIF-dependent and MyD88-independent TLR4 signaling pathway. Receptor-interacting protein (RIP1) and p38 MAPK are downstream components of this pathway. This signaling pathway did not affect cell viability and was not mediated by c-Jun amino-terminal kinase (JNK), indicating that it is distinct from the autophagic death signaling pathway (41). The TLR4/TRIF/RIP1/p38 MAPK signaling pathway may also be responsible for LPS activation of both the ALP and the UPP in muscle demonstrated in the present study. There has been no information on how p38 MAPK regulates the autophagy activity and atrogin-1/MAFbx gene expression, which will be an interesting topic for our future projects.

Elevated serum LPS levels are also seen in such procatabolic diseases as cancer (30), chronic heart failure (31), and type 2 diabetes (32). In addition, TLR4 can be activated by endogenous agonists, including heat-shock proteins (55) and S100A8/S100A9 (calprotectin) (56) that are part of the response to cellular damage/stress. Thus, TLR4-mediated muscle catabolism may take place in a number of cachectic conditions in addition to endotoxemia.

Because of the importance of the UPP in degrading myofibrillar proteins, there has been interest in developing inhibitors of the UPP to combat cachectic muscle wasting. Our findings suggest that the intervention of cachectic muscle wasting should also include a strategy for inhibiting aberrant autophagy activity. Notwithstanding, autophagy was also found to be required for maintaining muscle mass (57). Thus, excessive inhibition of autophagy may have side effects. A desirable therapeutic strategy is to target the signaling pathway that up-regulates autophagosome formation, as well as ubiquitin ligase expression in response to pathological stimuli. Because of the important role of TLR4 in innate immunity, inhibition of TLR4 may not be a highly desirable therapeutic strategy for muscle wasting. Considering that p38 MAPK responds to a variety of inflammatory mediators, such as TNF-α, IL-1, and ROS (ROS), this downstream regulator of both the UPP and the ALP is a better therapeutic target.

Acknowledgments

This study was supported by a National Institute of Arthritis and Musculoskeletal and Skin Diseases R01 grant to Y.-P.L. (AR052511).

The authors thank S. Akira (Osaka University, Oskaka, Japan) for sharing the TLR4−/− mice, and J. Han (Scripps Research Institute, La Jolla, CA, USA) for sharing the adenovirus construct of MKK6bE.

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