Impaired Ribosomal Biogenesis by Noncanonical Degradation of β-Catenin during Hyperammonemia

Increased ribosomal biogenesis occurs during tissue hypertrophy, but whether ribosomal biogenesis is impaired during atrophy is not known. We show that hyperammonemia, which occurs in diverse chronic disorders, impairs protein synthesis as a result of decreased ribosomal content and translational capacity.

ABSTRACT

Increased ribosomal biogenesis occurs during tissue hypertrophy, but whether ribosomal biogenesis is impaired during atrophy is not known. We show that hyperammonemia, which occurs in diverse chronic disorders, impairs protein synthesis as a result of decreased ribosomal content and translational capacity. Transcriptome analyses, real-time PCR, and immunoblotting showed consistent reductions in the expression of the large and small ribosomal protein subunits (RPL and RPS, respectively) in hyperammonemic murine skeletal myotubes, HEK cells, and skeletal muscle from hyperammonemic rats and human cirrhotics. Decreased ribosomal content was accompanied by decreased expression of cMYC, a positive regulator of ribosomal biogenesis, as well as reduced expression and activity of β-catenin, a transcriptional activator of cMYC. However, unlike the canonical regulation of β-catenin via glycogen synthase kinase 3β (GSK3β)-dependent degradation, GSK3β expression and phosphorylation were unaltered during hyperammonemia, and depletion of GSK3β did not prevent ammonia-induced degradation of β-catenin. Overexpression of GSK3β-resistant variants, genetic depletion of IκB kinase β (IKKβ) (activated during hyperammonemia), protein interactions, and in vitro kinase assays showed that IKKβ phosphorylated β-catenin directly. Overexpressing β-catenin restored hyperammonemia-induced perturbations in signaling responses that regulate ribosomal biogenesis. Our data show that decreased protein synthesis during hyperammonemia is mediated via a novel GSK3β-independent, IKKβ-dependent impairment of the β-catenin–cMYC axis.

INTRODUCTION

Increased ribosomal biogenesis and ribosome abundance are critical for augmented protein synthesis during cellular proliferation and hypertrophy (1–3). In contrast, whether and how ribosomal biogenesis is perturbed during tissue atrophy, especially in skeletal muscle, the major protein store in the body, are unclear (4–6). The rate of cellular protein synthesis depends on both translational capacity and translational efficiency, which in turn are determined by ribosomal content and function, respectively (6). The ribosome is a complex organelle that comprises rRNA and ribosomal proteins (RPs). Ribosomal biogenesis occurs by the coordinated assembly of rRNA and RPs. RNA polymerase I (PolI) transcribes the 47S precursor rRNA from clusters of ribosomal DNA (rDNA) tandem repeat genes, and RNA PolII transcribes the mRNA for the large subunits (RPLs) and small subunits (RPSs) and most regulatory, noncoding RNAs, while RNA PolIII transcribes tRNA and 5S rRNA (7). Of these, transcription of 47S pre-rRNA is believed to be a rate-limiting step in ribosome biogenesis (8). Expressions of ribosomal proteins and rRNA have been used as measures of ribosomal biogenesis (6), even though ribosomal proteins and rRNA incorporated into ribosomes may be relatively stable, but there are few studies on their perturbations during cellular stress and atrophy.

The transcription factor cMYC increases ribosomal biogenesis by regulating the expression of a number of ribosomal components that are transcribed by RNA polymerase II (6, 9–13). cMYC binds to rDNA and increases PolI-mediated transcription of rRNA (11). Transcription of upstream binding factor 1 (UBF1), RPS, and RPL is also regulated by cMYC (6, 14). Additional mechanisms by which cMYC regulates rDNA include PolI cofactor recruitment as well as transcription of PolIII subunits and components (9, 15). Consistent with the regulatory function of cMYC on multiple ribosomal components, overexpression of cMYC results in cell growth and hypertrophy with increased ribosomal biogenesis; whether reduced cMYC mediates muscle atrophy and decreased ribosomal biogenesis is currently unknown (16, 17). cMYC expression is regulated by a number of mechanisms, including activation of β-catenin, a transcriptional activator of cMYC. β-Catenin is a critical component of the Wnt signaling pathway and has been best studied in the context of cell proliferation and organ regeneration, two processes dependent on extensive protein synthesis (18, 19). β-Catenin also regulates protein synthesis in the skeletal muscle (18, 19), and expression of the β-catenin target cMYC is increased during skeletal muscle hypertrophy (18). Stabilization of skeletal muscle β-catenin has also been reported in rat muscle following exercise (20). Consistently, conditional inactivation of β-catenin in mice by targeted depletion resulted in small myotube size in a cell-autonomous manner (19); whether impaired β-catenin signaling is involved in muscle atrophy is, however, not known. In the canonical pathway, in the absence of Wnt ligand, β-catenin is phosphorylated at Ser³³ by glycogen synthase kinase 3β (GSK3β), followed by ubiquitination and degradation in the proteasome (21–25). More recently, IκB kinase β (IKKβ) has been reported to cause the phosphorylation and degradation of β-catenin in mesenchymal stem cell differentiation (26), but whether this pathway is relevant in mature skeletal muscle protein synthesis is also not known.

While there is fairly robust evidence supporting a regulatory role of the β-catenin–cMYC axis in increased ribosomal biogenesis and translational changes during skeletal muscle hypertrophy (4–6, 27), there are limited data as to whether decreased ribosomal biogenesis occurs during muscle atrophy. Ribosomal abundance, measured by total RNA quantity, and molecular regulators of ribosomal biogenesis were decreased in type II fibers in aged rats with muscle loss despite an increase in cMYC expression (28). The reduction in ribosomal biogenesis in this model was shown to be due to an increase in proteasomal activity, suggesting translational deficits rather than global transcriptional mechanisms as mediators of aging-related muscle loss (28). In contrast, others have reported unaltered or even increased skeletal muscle ribosomal biogenesis with aging (29–31). These conflicting observations may be indicative of a compensatory increase in ribosome biogenesis during muscle atrophy or a context-dependent response. Despite muscle loss being frequent under conditions other than aging, such as chronic diseases, it is not known if skeletal muscle ribosomal biogenesis is altered under these conditions.

Despite a number of preclinical models of chronic disease, there have been limited mechanistic studies of muscle loss in chronic disease (32–36). Hyperammonemia, which occurs in chronic diseases with muscle loss (37), is now increasingly recognized to initiate a sequence of perturbations that culminate in impaired protein synthesis and muscle loss (38, 39). Ammonia is a cytotoxic compound generated during cellular metabolic processes (40), and ammonia metabolism is disordered in cirrhosis of the liver, heart failure, and chronic lung disease (37, 41, 42), with muscle loss being frequent under all these conditions. Reduced muscle mass and hyperammonemia have been reported by others and us in patients with cirrhosis of the liver (39, 43, 44). The phenotype of lower muscle mass has also been reported in the hyperammonemic portacaval anastomosis (PCA) rat, and a sarcopenic phenotype has been reported in myotubes during hyperammonemia (38, 45, 46). We have also previously reported a myostatin-mediated impairment of mTORC1 signaling in response to hyperammonemia (38, 47). These studies show that the level of mTORC1, a critical regulator of translational efficiency, specifically translation initiation, is decreased during muscle atrophy (48, 49). Hyperammonemia induces muscle atrophy, anabolic resistance, decreased protein synthesis, and impaired mTORC1 signaling responses (37, 47, 50). However, whether hyperammonemia perturbs ribosomal biogenesis and the β-catenin–cMYC axis has not been evaluated. In these studies, we show that active β-catenin and its downstream signaling responses are decreased in myotubes and HEK cells as well as in skeletal muscle from patients with cirrhosis and the PCA rat. We also show that hyperammonemia-induced loss of β-catenin is caused by proteasomal degradation via a noncanonical GSK3β-independent, IKKβ-dependent pathway.

RESULTS

Decreased skeletal muscle protein synthesis and ribosomal biogenesis during hyperammonemia.

Consistent with data reported previously by us and others (51, 52), hyperammonemia impaired protein synthesis in myotubes, as determined by puromycin incorporation (Fig. 1A). RNA sequencing (RNA-Seq) comparing untreated controls and myotubes exposed to hyperammonemia for 24 h identified a large number of differentially expressed genes. In particular, levels of mRNAs for ribosomal proteins were significantly decreased in ammonia-treated C2C12 myotubes only at 24 h (Fig. 1B and C). The lack of major differences between the transcriptome at 3 h and untreated controls, in contrast to the differences in those observed with 24-h hyperammonemic myotubes, may be due to an adaptive response in the early stages during hyperammonemia progressing to a maladaptive response with a longer duration of hyperammonemia. To determine the underlying molecular mechanism, the RNA sequence data were submitted for Ingenuity Pathway Analysis (IPA) core analysis, and differentially expressed genes were categorized based on the Ingenuity Pathway Knowledge Base (IPKB); the most enriched categories of the canonical pathway are listed in Fig. 1D, and the differentially expressed genes are provided in Table S1 in the supplemental material. In addition, networks built to connect key genes regulated by β-catenin showed downregulation of target genes, including those regulating ribosomal proteins (TCF-LEF, axin-2, and Frizzled), and that this pathway was perturbed during hyperammonemia (Fig. 1E). To further confirm that translation is indeed affected by hyperammonemia and to establish clinical relevance, we quantified total RNA as a measure of ribosomal abundance and translational capacity (28, 53). Hyperammonemic C2C12 myotubes, gastrocnemius muscle from PCA rat, and human skeletal muscle from patients with cirrhosis consistently had low expression levels of total RNA compared to controls (Fig. 1F). Also, expression levels of the 45S rRNA and the processed 28S and 18S rRNAs were reduced in myotubes and skeletal muscle from the hyperammonemic PCA rat and human patients with cirrhosis, compared to controls (Fig. 1G and H). We further used polysome fractionation to investigate the effects of hyperammonemia on ribosome density and assembly. Relative to control cells, hyperammonemia induced a reduction in absorbance in gradient fractions corresponding to polysomes, accompanied by a concomitant decrease in absorbance in the monosome fraction. The ratio of polysomes to the sum of polysomes and monosomes was 93.6% ± 2.1% of simultaneous untreated controls (n = 3; P < 0.05) (Fig. 1I).

FIG 1.

Hyperammonemia impairs protein synthesis in myotubes. (A) Representative immunoblotting and densitometry of puromycin incorporation in untreated C2C12 myotubes (UnT) or those treated with 10 mM ammonium acetate (AmAc) for 6 h and 24 h. (B) Heat map of transcriptome analysis comparing untreated C2C12 myotubes to those treated with 10 mM AmAc for 3 or 24 h. (C) Heat map of C2C12 myotubes after 24 h of treatment with AmAc compared with untreated cells. (D) Ingenuity Pathway Knowledge Base (IPKB) analysis of the most enriched canonical pathways present in differentiated C2C12 myotubes after 24 h of hyperammonemia. (E) Ingenuity Pathway Analysis of the β-catenin (CTNNB1) node from the entire transcriptome (from panel B). Note that the majority of connections are downregulated upon AmAc treatment (indicated in red). (F) Cellular RNA levels from untreated myotubes or those treated with 10 mM AmAc for 6 or 24 h; from the gastrocnemius muscle of portocaval anastomosis (PCA) and sham-operated, pair-fed control (Sham) rats (n = 5 in each group); and from the skeletal muscle of patients with cirrhosis (CIR) and healthy controls (CTL) (n = 5 in each group). (G) Fold changes of the expression levels of the 45S rRNA in C2C12 myotubes treated with 10 mM AmAc for 30 min or 3, 6, or 24 h in the gastrocnemius muscle of portocaval anastomosis (PCA) and sham-operated pair-fed control rats (n = 5 each) and the skeletal muscle of patients with cirrhosis and controls (n = 5 each). rRNA expression was normalized against β-actin. (H) RNA gel of 28S and 18S bands in untreated myotubes and those treated with 10 mM AmAc for different times. (I) Effect of AmAc treatment on the polysome/monosome ratio in C2C12 myotubes. Representative polysome profiles from a sucrose gradient (20 to 47%) were obtained by taking the optical density at 254 nm (OD₂₅₄) for untreated control and 10 mM AmAc-treated C2C12 cell extracts. Positions of 80S and polysome species are indicated. A.U., arbitrary units. All cellular experiments were performed in at least 3 biological replicates, and data are expressed as means ± SD. *, P < 0.05; **, P < 0.01; ***, P < 0.001 (versus the control).

Lower levels of rRNA were accompanied by reduced levels of the ribosomal component protein RPL5, -23, and -32 as well as RPS9, -19, and -26 mRNAs in hyperammonemic myotubes (Fig. 2A and B) and of RPL5, -23, and -32 proteins in hyperammonemic myotubes, gastrocnemius muscle from the PCA rat, and skeletal muscle from patients with cirrhosis (Fig. 2C to E). Interestingly, the ammonia-induced reduction in the expression of RPL5, -23, and -32 proteins was not limited only to skeletal muscle but was also observed in nonmuscle HEK293 cells (Fig. 2F). Ribosomal proteins may have relatively long half-lives after their incorporation into ribosomes; however, our studies on protein expression in response to cycloheximide (CHX) as a measure of protein degradation and MG132 as a measure of protein translation support the use of these inhibitors to study the stability of RPL5, -23, and -32 proteins and their expression in response to hyperammonemia (Fig. 2G). Consistently, during hyperammonemia, the expression of ribosomal proteins was significantly decreased, and MG132 treatment prevented ammonia-induced reduction of the RPLs studied (Fig. 2G). We also noted that the expression of a housekeeping gene, β-actin, was unaltered, but that of a nonhousekeeping, regulatable gene, total P70S6 kinase (P70S6K), was decreased by cycloheximide (which shows that it is regulated by translation) but did not change in response to hyperammonemia (Fig. 2G). An unbiased proteomics analysis of myotubes treated with ammonium acetate (AmAc) compared to untreated cells showed alterations in ribosomal proteins (Table 1). There was a reduction in the expression of 12 ribosomal proteins that are incorporated into the 40S subunit and 20 ribosomal proteins that are incorporated into the 60S subunit. Together, these data show that during hyperammonemia, rRNA and protein levels are decreased in skeletal muscle during hyperammonemia.

FIG 2.

Hyperammonemia impairs skeletal muscle ribosomal biogenesis. (A) Relative fold changes in expression of ribosomal RPL5, RPL23, and RPL32 mRNAs, comparing myotubes treated with 10 mM AmAc for 6 or 24 h with untreated (UnT) controls, measured by RT-quantitative PCR (qRT-PCR). (B) Relative fold changes in expression of ribosomal RPS9, RPS19, and RPS26 mRNAs, comparing myotubes treated with 10 mM AmAc for 6 or 24 h with untreated controls, measured by qRT-PCR. (C to E) Representative immunoblotting and densitometry of expression of ribosomal proteins RPL5, RPL23, and RPL32 from untreated C2C12 myotubes or those treated with 10 mM AmAc for 24 h (C); from the gastrocnemius muscle of portocaval anastomosis (PCA) and sham-operated, pair-fed control (Sham) rats (n = 5 each) (D); and from the skeletal muscle of patients with cirrhosis (CIR) and controls (CTL) (n = 5 each) (E). (F) Representative immunoblotting and densitometry of ribosomal biogenesis markers RPL5, RPL23, and RPL32 in untreated HEK cells and those treated with 10 mM AmAc for 6 or 24 h. (G) Representative immunoblotting and densitometry of expression of ribosomal proteins RPL5, -23, and -32 show that with 24 h of treatment with the translational inhibitor cycloheximide (CHX), there is reduced expression of ribosomal proteins, which is blocked by proteasome inhibition. However, the expression of a housekeeping gene, β-actin, did not change in response to these interventions, and the expression of a regulatable, nonhousekeeping gene (P70S6K) did not change, except with cycloheximide treatment, which resulted in lower expression levels than in untreated cells. All cellular experiments were performed in at least 3 biological replicates, and all data were normalized to β-actin and are expressed as means ± SD. *, P < 0.05; **, P < 0.01; ***, P < 0.001 (versus untreated cells or controls for all panels).

TABLE 1.

Unbiased proteomics analysis of myotubes treated with AmAc for 24 h compared to untreated cells^a

NCBI accession no.	Protein	Designation	Mol wt (kDa)	LFQ intensity at 24 h				LFQ intensity for Ct				Ratio of 24-h/Ct LFQ intensity	P value by t test	MS/MS count
				Expt 1	Expt 2	Expt 3	Avg	Expt 1	Expt 2	Expt 3	Avg
P60867	40S ribosomal protein S20	Rps20	13.373	1.81E+09	1.91E+09	1.87E+09	1.86E+09	2.07E+09	1.97E+09	2.04E+09	2.03E+09	0.92	0.018	152
P63325	40S ribosomal protein S10	Rps10	18.916	4.04E+08	4.42E+08	4.33E+08	4.26E+08	4.64E+08	5.07E+08	4.97E+08	4.89E+08	0.87	0.022	117
P62242	40S ribosomal protein S8	Rps8	24.205	2.17E+09	2.73E+09	2.46E+09	2.46E+09	2.85E+09	3.19E+09	3.01E+09	3.02E+09	0.81	0.040	300
P62754	40S ribosomal protein S6	Rps6	28.68	1.55E+09	1.90E+09	1.75E+09	1.74E+09	2.35E+09	2.29E+09	2.34E+09	2.32E+09	0.75	0.005	251
P62267	40S ribosomal protein S23	Rps23	15.807	5.13E+08	6.54E+08	5.60E+08	5.76E+08	7.20E+08	7.31E+08	9.23E+08	7.92E+08	0.73	0.050	133
P62281	40S ribosomal protein S11	Rps11	18.431	1.24E+09	1.37E+09	1.27E+09	1.29E+09	1.82E+09	1.49E+09	1.41E+09	1.57E+09	0.82	0.100	198
P62855	40S ribosomal protein S26	Rps26	13.015	6.72E+08	7.12E+08	6.49E+08	6.77E+08	6.86E+08	8.06E+08	7.41E+08	7.44E+08	0.91	0.164	78
P63276	40S ribosomal protein S17	Rps17	15.524	1.54E+09	1.58E+09	1.65E+09	1.59E+09	1.65E+09	1.63E+09	1.84E+09	1.71E+09	0.93	0.190	165
P25444	40S ribosomal protein S2	Rps2	31.231	1.58E+09	1.77E+09	1.69E+09	1.68E+09	1.90E+09	1.72E+09	1.75E+09	1.79E+09	0.94	0.219	232
P62264	40S ribosomal protein S14	Rps14	16.273	1.53E+09	1.65E+09	1.56E+09	1.58E+09	1.74E+09	1.53E+09	1.65E+09	1.64E+09	0.96	0.435	154
P62274	40S ribosomal protein S29	Rps29	6.6767	5.77E+08	6.43E+08	5.40E+08	5.87E+08	6.81E+08	6.02E+08	5.73E+08	6.19E+08	0.95	0.512	56
P62301	40S ribosomal protein S13	Rps13	17.222	5.37E+08	4.54E+08	4.94E+08	4.95E+08	6.56E+08	4.41E+08	4.94E+08	5.30E+08	0.93	0.634	120
P27659	60S ribosomal protein L3	Rpl3	46.109	2.41E+09	2.73E+09	2.56E+09	2.56E+09	3.05E+09	3.08E+09	3.09E+09	3.07E+09	0.83	0.005	377
Q9CPR4	60S ribosomal protein L17	Rpl17	21.423	1.37E+09	1.60E+09	1.36E+09	1.44E+09	1.88E+09	1.79E+09	1.70E+09	1.79E+09	0.81	0.023	225
P47963	60S ribosomal protein L13	Rpl13	24.305	1.84E+09	2.10E+09	1.96E+09	1.97E+09	2.44E+09	2.50E+09	2.69E+09	2.54E+09	0.77	0.006	229
Q9Z2Q5	39S ribosomal protein L40, mitb	Mrpl40	24.301	5.55E+07	4.90E+07	4.49E+07	4.98E+07	6.26E+07	7.00E+07	6.07E+07	6.45E+07	0.77	0.025	46
P41105	60S ribosomal protein L28	Rpl28	15.733	5.35E+08	6.39E+08	5.51E+08	5.75E+08	7.57E+08	7.45E+08	7.53E+08	7.52E+08	0.77	0.006	156
P61255	60S ribosomal protein L26	Rpl26	17.258	5.93E+08	7.08E+08	6.82E+08	6.61E+08	9.70E+08	9.00E+08	8.02E+08	8.91E+08	0.74	0.019	141
P62717	60S ribosomal protein L18a	Rpl18a	20.732	2.82E+08	3.72E+08	3.69E+08	3.41E+08	4.67E+08	4.90E+08	5.20E+08	4.92E+08	0.69	0.010	108
P12970	60S ribosomal protein L7a	Rpl7a	29.976	6.47E+08	6.23E+08	6.11E+08	6.27E+08	9.71E+08	9.43E+08	8.53E+08	9.22E+08	0.68	0.001	191
P47911	60S ribosomal protein L6	Rpl6	33.509	1.35E+09	1.65E+09	1.53E+09	1.51E+09	2.33E+09	2.27E+09	2.14E+09	2.25E+09	0.67	0.002	246
Q6ZWV7	60S ribosomal protein L35	Rpl35	14.552	3.94E+08	5.16E+08	4.34E+08	4.48E+08	6.72E+08	7.17E+08	6.46E+08	6.78E+08	0.66	0.005	73
P62918	60S ribosomal protein L8	Rpl8	28.024	9.82E+08	1.28E+09	1.12E+09	1.13E+09	1.82E+09	1.59E+09	1.78E+09	1.73E+09	0.65	0.006	205
Q8BP67	60S ribosomal protein L24	Rpl24	17.779	3.58E+08	4.28E+08	3.97E+08	3.94E+08	6.99E+08	6.28E+08	6.35E+08	6.54E+08	0.60	0.001	110
Q9D1R9	60S ribosomal protein L34	Rpl34	13.293	1.38E+08	1.62E+08	1.75E+08	1.58E+08	3.30E+08	2.45E+08	2.56E+08	2.77E+08	0.57	0.014	48
Q9CR57	60S ribosomal protein L14	Rpl14	23.564	1.84E+08	1.79E+08	1.99E+08	1.87E+08	3.77E+08	3.54E+08	2.95E+08	3.42E+08	0.55	0.004	75
P35980	60S ribosomal protein L18	Rpl18	21.644	1.65E+08	2.64E+08	1.81E+08	2.03E+08	3.06E+08	4.06E+08	5.40E+08	4.17E+08	0.49	0.045	92
P14148	60S ribosomal protein L7	Rpl7	31.419	3.59E+08	5.29E+08	3.65E+08	4.18E+08	7.76E+08	8.50E+08	9.51E+08	8.59E+08	0.49	0.004	161
P47964	60S ribosomal protein L36	Rpl36	12.215	3.93E+08	4.58E+08	4.83E+08	4.45E+08	5.49E+08	5.12E+08	6.25E+08	5.62E+08	0.79	0.052	93
P19253	60S ribosomal protein L13a	Rpl13a	23.464	5.08E+08	7.59E+08	5.99E+08	6.22E+08	7.35E+08	7.91E+08	7.84E+08	7.70E+08	0.81	0.121	157
P62892	60S ribosomal protein L39	Rpl39	6.4066	3.29E+08	3.83E+08	3.53E+08	3.55E+08	3.75E+08	4.06E+08	4.84E+08	4.22E+08	0.84	0.138	55
Q9D823	60S ribosomal protein L37	Rpl37	11.078	500,000	1,372,000	3,744,100	1.87E+06	2,985,600	2.45E+07	500,000	9.33E+06	0.20	0.387	7

^aLFQ, label-free quantitation; Ct, control group.

^bmit, mitochondrial.

Hyperammonemia impairs the β-catenin–cMYC axis.

Since the β-catenin–cMYC axis regulates ribosomal components and ribosomal biogenesis (6, 54), we determined whether hyperammonemia impaired ribosomal biogenesis and protein synthesis via perturbations in this axis. cMYC regulates protein synthesis by increasing ribosomal biogenesis and has also been suggested to mediate skeletal muscle hypertrophy (6, 10). We observed reductions in the levels of cMYC and UBF1, a ribosomal DNA transcription factor, during hyperammonemia, with lower levels of ribosomal biogenesis in all models evaluated (Fig. 3A to C). Expression of β-catenin, a transcriptional activator of cMYC, was also decreased during hyperammonemia in all 3 models studied (Fig. 3D to F). These effects of hyperammonemia extend beyond those in the skeletal muscle because similar findings were observed in HEK293 cells during hyperammonemia (Fig. 3G). β-Catenin-dependent signaling is activated by dephosphorylation, followed by nuclear translocation and upregulation of its target genes. Reporter assays using Topflash/Fopflash showed that decreased expression of active, dephosphorylated β-catenin was associated with reduced transcriptional activity during hyperammonemia (Fig. 3H). Importantly, the cells still retained their ability to activate β-catenin, as two chemical stabilizers of β-catenin, 6-bromoindirubin-3′-oxime (BIO) and lithium chloride (LiCl), were able to increase β-catenin levels, nuclear translocation, and transactivation (Fig. 3H and I). These data show that hyperammonemia decreased β-catenin stability and transcriptional activity.

FIG 3.

Reduced skeletal muscle β-catenin expression during hyperammonemia. (A to C) Representative immunoblotting and densitometry of cMYC and upstream binding factor 1 (UBF1) expression in untreated (UnT) murine C2C12 myotubes or those treated with 10 mM AmAc for 24 h (A); skeletal muscle from portocaval anastomosis (PCA) and sham-operated, pair-fed (Sham) control rats (n = 5 each) (B); and skeletal muscle of patients with cirrhosis (CIR) and controls (CTL) (n = 5 each) (C). (D to F) Representative immunoblotting and densitometry of total (T) and active, dephosphorylated (A) β-catenin expression from untreated C2C12 myotubes or those treated with 10 mM AmAc for 24 h (D); from the gastrocnemius muscle of PCA and sham-operated, pair-fed control rats (n = 5 each) (E); and from the skeletal muscle of patients with cirrhosis and control subjects (n = 5 each) (F). (G) Representative immunoblotting and densitometry of total and activated β-catenin as well as its transcriptional targets cMYC and UBF1 in untreated HEK cells or those treated with 10 mM ammonium acetate for 6 or 24 h. (H) Luciferase activity measuring transcriptional activity of β-catenin using Topflash reporter activity (normalized to Renilla) comparing untreated myotubes and those treated for 24 h with 10 mM AmAc, 5 mM lithium chloride (LiCl), or 2 μM BIO (6-bromoindirubin-3′-oxime) (small-molecule inhibitor of GSK3β with resultant activation of β-catenin), expressed as fold changes over untreated cells. (I) Representative photomicrographs of immunofluorescence of active, unphosphorylated β-catenin expression (green) in murine C2C12 myotubes treated with 10 mM AmAc or 2 μM BIO for 24 h. Nuclei are counterstained with 4′,6-diamidino-2-phenylindole (DAPI) (blue). Scale bars, 40 μm. **, P < 0.01; ***, P < 0.001 (versus untreated cells). All data are expressed as means ± SD. All cellular experiments were performed in at least 3 biological replicates, and all blots were normalized to β-actin. The antibody against active β-catenin recognizes the unphosphorylated, stable form of β-catenin.

Hyperammonemia results in β-catenin degradation that is independent of GSKβ.

Regulation of β-catenin is mediated primarily by stabilization through the Wnt signaling pathway, which phosphorylates and inactivates GSK3β, the canonical inhibitor of β-catenin (55). Even though GSK3β is the best-recognized regulator of β-catenin (56), during skeletal muscle hyperammonemia, phosphorylation of GSK3β was unaltered (Fig. 4A to C). Interestingly, we showed that muscle atrophy during hyperammonemia was not reversed by silencing of either GSKα or GSKβ in myotubes (Fig. 4D). Consistently, either chemical inhibitors of GSK3β, BIO and LiCl, or genetic depletion using short hairpin RNA (shRNA) targeting GSK3α (shGSK3α) and shGSK3β increased active β-catenin expression in untreated cells but did not restore active β-catenin expression during hyperammonemia (Fig. 4E and F). Expression of total P70S6K was used to show that the expression of a regulatable, nonhousekeeping gene did not change in response to hyperammonemia (Fig. 4F). Chemical inhibitors of GSK3β, BIO and LiCl, could not restore the β-catenin transcriptional activity, as shown by a Topflash assay (Fig. 5A). Finally, using the proteasome inhibitor MG132, we could prevent β-catenin loss during hyperammonemia, showing that canonical proteasome-dependent degradation of β-catenin was unaltered (Fig. 5B). To further determine if mTORC1 contributes to the effects of hyperammonemia on ribosome biogenesis, we treated myotubes with rapamycin and observed that expression levels of RPL5, -23, and -32 mRNAs were unaltered by blocking mTORC1, while phosphorylation of P70S6 kinase, a known target of mTORC1 signaling, was inhibited by rapamycin (Fig. 5C and D).

FIG 4.

Skeletal muscle hyperammonemia does not alter GSK3β expression or activity. (A to C) Representative immunoblots of phosphorylated and total GSK3β from untreated (UnT) C2C12 myotubes or those treated with 10 mM AmAc for 6 or 24 h (A), skeletal muscle from portocaval anastomosis (PCA) and sham-operated pair-fed control (Sham) rats (n = 5 each) (B), and skeletal muscle of patients with cirrhosis (CIR) and controls (CTL) (n = 5 each) (C). (D) Representative photomicrographs and myotube diameters of differentiated C2C12 myotubes depleted of GSK3β or GSK3α or transfected with a vector with a random construct (shRan), treated with and without 10 mM ammonium acetate (AmAc). Scale bars, 40 μm. (E) Representative immunoblots of dephosphorylated active (A) and total (T) β-catenin, as well as GSK3β, in untreated C2C12 myotubes or those treated with 10 mM AmAc for 24 h in the presence or absence of GSK3β inhibitors, 5 mM lithium chloride (LiCl) or 2 μM BIO (6-bromoindirubin-3′-oxime), that activate β-catenin. Densitometry of blots from active β-catenin were normalized to β-actin. (F) Representative immunoblots of GSK3α and GSK3β and immunoblots and densitometry of RPL5, RPL23, RPL32, P70S6K, and β-actin in myotubes with silencing of GSK3α and GSK3β with and without 10 mM AmAc for 24 h. Expression of total P70S6K was used to determine the response of a nonhousekeeping, regulatable protein with hyperammonemia. All cellular experiments were performed in at least 3 biological replicates, blots were normalized to β-actin, and data are expressed as means ± SD. **, P < 0.01; ***, P < 0.001 (versus untreated cells). The antibody against active β-catenin recognizes the unphosphorylated, stable form of β-catenin.

FIG 5.

Hyperammonemia effects on ribosomal proteins are mediated by GSK- and mTORC1-independent effects. (A) Luciferase activity measuring transcriptional activity of β-catenin using the Topflash reporter assay (normalized to Renilla) comparing untreated (UnT) myotubes and those treated for 24 h with 10 mM AmAc in the presence or absence of 5 mM LiCl or 2 μM BIO. Data are depicted as fold changes over Topflash activity. (B) Representative immunoblots for β-catenin phosphorylated at Ser^33/37 in untreated C2C12 myotubes or those treated with 10 mM AmAc for 6 or 24 h. Proteasome activity was blocked with 10 μM MG132 for the last 1 h to prevent ubiquitin-mediated degradation of phosphorylated β-catenin. (C) Fold changes in mRNAs for RPL5, -23, and -32 in myotubes treated for 24 h with and without 10 mM AmAc and/or rapamycin (Rap) (1 μg/ml). (D) Representative immunoblots of phosphorylated P70S6 kinase in response to 10 mM ammonium acetate for 24 h or 1 μg/ml rapamycin. All cellular experiments were performed in at least 3 biological replicates, blots were normalized to β-actin, and data are expressed as means ± SD. **, P < 0.01; ***, P < 0.001 (versus untreated cells). The antibody against active β-catenin recognizes the unphosphorylated, stable form of β-catenin.

Noncanonical degradation of β-catenin during hyperammonemia.

We then determined how increased β-catenin degradation occurred, independently of the canonical GSK3β pathway, during hyperammonemia. We previously reported increased IKK-dependent signaling during hyperammonemia (38). The IKK complex is comprised of both IKKα and IKKβ, in addition to other components (57–59). We genetically depleted IKK to determine whether β-catenin stability is regulated by IKKα or IKKβ. As shown in Fig. 6A, only knockdown of IKKβ restores β-catenin levels during hyperammonemia. Genetic depletion of IKKβ prevented not only hyperammonemia-induced reduction in active β-catenin but also expression of its targets cMYC and UBF1 (Fig. 6A and B). Silencing of IKKβ also partially reversed myotube atrophy during hyperammonemia, while random shRNA or shIKKα did not result in a reversal of the sarcopenic phenotype response to hyperammonemia (Fig. 6C). Consistently, depletion of only IKKβ resulted in nuclear translocation of active β-catenin by immunofluorescence studies (Fig. 6D). The Topflash reporter assay also showed that silencing of IKKβ restored the ammonia-induced reduction in the transcriptional activity of β-catenin (Fig. 6E) and partially reversed ammonia-induced impaired protein synthesis (Fig. 7A). These effects are likely via direct protein-protein interactions because IKKβ and β-catenin were associated with each other in both myotubes and HEK293 cells during hyperammonemia (Fig. 7B and C).

FIG 6.

Skeletal muscle IKKβ is a novel β-catenin kinase during hyperammonemia. (A) Representative immunoblotting and densitometry of active dephosphorylated and total β-catenin, cMYC, and UBF1 in C2C12 myotubes transfected with shRNAs targeting IKKβ or a scrambled shRNA (shRan) that were either untreated (UnT) or treated with 10 mM AmAc for 24 h. (B) Representative immunoblots of IKKβ and IKKα and immunoblots and densitometry of total and active β-catenin in untreated C2C12 myotubes or those treated with 10 mM AmAc for 24 h, stably expressing IKKβ or IKKα shRNA or a scrambled control (shRan). (C) Representative photomicrographs and myotube diameters of differentiated C2C12 myotubes depleted of IKKα or IKKβ or transfected with a vector with a random construct (shRan), treated with and without 10 mM AmAc. Scale bars, 40 μm. (D) Representative photomicrographs of immunofluorescence of active, dephosphorylated β-catenin expression (red) in myotubes stably expressing shRNAs targeting IKKβ or IKKα or a scrambled shRNA (shRan) that were either untreated or treated with 10 mM AmAc for 24 h. Nuclei were counterstained with DAPI (blue). Scale bars, 40 μm. (E) Luciferase activity for β-catenin transcriptional activity quantified by a Topflash assay (normalized to Renilla) in untreated C2C12 myotubes or those treated with 10 mM AmAc for 24 h, stably expressing either an shRNA targeting IKKβ shRNA or a scrambled control (shRan). All cellular experiments were performed in at least 3 biological replicates, all blots were normalized to β-actin, and data are expressed as means ± SD. *, P < 0.05; **, P < 0.01; ***, P < 0.001 (shIKKβ versus shRan). The antibody against active β-catenin recognizes the unphosphorylated, stable form of β-catenin. Bars, 40 μm.

FIG 7.

Hyperammonemia regulates β-catenin via IKKβ. (A) [³H]phenylalanine incorporation in untreated myotubes and those treated for 24 h with AmAc and depleted of IKKβ or IKKα showing a partial reversal of AmAc-induced protein synthesis with silencing of IKKβ. (B) Immunoprecipitate of β-catenin probed for IKKβ and immunoprecipitate of IKKβ probed for β-catenin in untreated (UnT) murine C2C12 myotubes or those treated with 10 mM AmAc for 24 h. (C) Immunoprecipitate of IKKβ probed for β-catenin and immunoprecipitate of β-catenin probed for IKKβ in untreated HEK cells or those treated with 10 mM AmAc for 24 h. (D) Representative immunoblotting and densitometry of total β-catenin levels in C2C12 myotubes stably transfected with β-catenin mutants that are resistant to GSK3β phosphorylation (S33Y, S45Y, and N-terminal deletion [ΔN-ter] constructs). The β-catenin N-terminal deletion construct has a lower molecular weight. (E) In vitro kinase assay with purified β-catenin and either IKKβ or GSK3β (positive control) and negative controls. The blots were probed with phospho-β-catenin (Ser^33/37/41) antibody. The purity of reagents was tested by immunoblotting. All cellular experiments and in vitro kinase assays were performed in at least 3 biological replicates, all blots were normalized to β-actin, and data are expressed as means ± SD. *, P < 0.05; **, P < 0.01; ***, P < 0.001; NS, not significant (versus mock-transfected untreated control myotubes).

Finally, we investigated whether the IKKβ-mediated phosphorylation of β-catenin that occurs during hyperammonemia is independent of GSK3β. C2C12 myotubes stably transfected with either the S33Y, S45K, or N-terminal deletion mutant of β-catenin had reduced expression of β-catenin during hyperammonemia (Fig. 7D). These data demonstrate that despite mutations in the phosphorylation sites of GSK3β, β-catenin was still degraded during hyperammonemia, suggesting that a GSK3β-independent, IKKβ-dependent pathway is active during hyperammonemia. Consistently, in vitro kinase assays showed that purified, active IKKβ phosphorylated β-catenin, as did GSK3β (Fig. 7E).

Overexpression of β-catenin reverses protein synthesis and ribosomal biogenesis.

To determine whether β-catenin degradation is not only necessary but also sufficient for decreased ribosomal biogenesis and protein synthesis during hyperammonemia, gains of β-catenin function were evaluated in myotubes. Overexpression of β-catenin in myotubes increased the expression of its targets, the cMYC and UBF1 genes (Fig. 8A) and ribosomal proteins (Fig. 8B) and reversed the reduction in protein synthesis (Fig. 8C), 45S ribosome RNA (Fig. 8D), and the muscle atrophy phenotype during hyperammonemia (Fig. 8E).

FIG 8.

Overexpression of β-catenin restores reduced cMYC and protein synthesis. (A) Representative immunoblotting and densitometry of total (T) and active (A) β-catenin, cMYC, and UBF1 in untreated (UnT) or 10 mM AmAc-treated murine C2C12 myotubes overexpressing wild-type β-catenin or a mock construct. (B) Representative immunoblotting and densitometry of ribosomal proteins RPL5, RPL23, and RPL32 in untreated C2C12 myotubes or those treated with 10 mM AmAc for 24 h, stably overexpressing either wild-type β-catenin or a mock construct. (C) Representative immunoblotting and densitometry of puromycin incorporation in untreated C2C12 myotubes or those treated with 10 mM AmAc for 24 h, overexpressing wild-type β-catenin or a mock construct. (D) Relative fold changes in 45S rRNA expression in untreated myotubes or those treated with 10 mM AmAc for 24 h, overexpressing wild-type β-catenin or a mock vector. (E) Representative photomicrographs and myotube diameters of differentiated C2C12 myotubes with β-catenin overexpression compared with mock vector-transfected myotubes treated with and without 10 mM AmAc. Scale bars, 40 μm. All cellular experiments were performed in at least 3 biological replicates, all blots were normalized to β-actin, and data are expressed as means ± SD. ***, P < 0.001 (mock versus β-catenin overexpression). The antibody against active β-catenin recognizes the unphosphorylated, stable form of β-catenin.

DISCUSSION

We provide novel evidence that during hyperammonemia, loss of β-catenin and an ensuing decrease in cMYC expression result in impaired ribosomal biogenesis and translational capacity and a consequent reduction in protein synthesis. Complementing reports that an increase in β-catenin signaling contributes to skeletal muscle hypertrophy, we show that impaired β-catenin–cMYC ribosomal biogenesis occurs with skeletal muscle atrophy.

Canonical regulation of β-catenin is mediated by GSK3β-dependent phosphorylation, and the activity of β-catenin is regulated by phosphorylation followed by proteasome-mediated degradation (21, 25). During hyperammonemia, we observed no alteration in the phosphorylation of GSK3β or reversal of β-catenin expression or function in response to classical GSK3 inhibitors. Instead, β-catenin is phosphorylated by a different Ser/Thr kinase. Loss- and gain-of-function studies show that IKKβ is the kinase primarily responsible for hyperammonemia-induced loss of skeletal muscle. Consistent with the critical role of β-catenin in regulating ribosomal biogenesis, hyperammonemia resulted in impaired ribosomal biogenesis and decreased protein synthesis, with a sarcopenic phenotype. Importantly, we also observed hyperammonemia-induced IKKβ/β-catenin-dependent decreases in ribosomal biogenesis and muscle loss in vivo in rats and in patients with hyperammonemia.

Our studies show that hyperammonemia impairs protein synthesis in both muscle and HEK cells. These data are consistent with reports by others that ammonia impairs protein synthesis in a variety of systems, including skeletal muscle (52, 60–62). Protein synthesis is regulated at multiple levels, including initiation, elongation, and termination. Most studies to date have focused on the signaling pathways regulating protein synthesis via kinase- and phosphatase-dependent mechanisms (63). Since skeletal muscle is the largest protein store in the body, ribosomal biogenesis is critical for protein homeostasis (1). Ribosome biogenesis is initiated by the synthesis and processing of rRNA and proteins and their assembly into the two major subunits, a large 60S subunit, which comprises 28S, 5S, and 5.8S rRNAs and 47 large ribosomal proteins (RPLs), and a 40S subunit, which comprises 18S rRNA and 33 small ribosomal proteins (RPSs) (1, 64). Ribosome biogenesis is increased during muscle hypertrophy and growth (6), but there are very limited data about the converse, that is, the decreased ribosomal biogenesis that occurs during muscle atrophy (6, 28, 65). Similarly, translational capacity, determined by total ribosome content, has been reported to be increased during muscle hypertrophy, but whether ribosome content is altered during atrophy has not been reported (6). We show that the levels of the 18S and 28S subunits of the rRNA, as well as the RPL proteins and mRNA, are decreased in multiple models of muscle atrophy during hyperammonemia. Our observations of reduced ribosome content and biogenesis during hyperammonemia-induced muscle loss are consistent with those reported for disuse-related muscle atrophy, where lower skeletal muscle RNA content was noted in rats during hindlimb immobilization and spinal cord injury (66, 67) or in humans during inactivity (68). Importantly, this phenomenon is different from aging-related sarcopenia and sciatic denervation-induced atrophy, where the skeletal muscle total RNA concentration is not decreased (31, 65). Also, our data show that ribosomal alterations in muscle atrophy are not necessarily the reverse of hypertrophy, because the changes in hypertrophy are believed to be due to translational rather than transcriptional perturbations (27). Interestingly, our observations using polysome profiling show that hyperammonemia does not seem to affect the proper maturation and assembly of the ribosomes. These data suggest that different catabolic stimuli regulate ribosome content and biogenesis in skeletal muscle, indicating context-dependent, upstream regulatory mechanisms.

The transcription factor cMYC modulates cellular functions by upregulating the transcription of rRNA and ribosomal proteins as well as rRNA processing, ribosome assembly, and nuclear-cytoplasmic export of mature ribosome subunits (6, 12, 15). In addition, cMYC also regulates upstream binding factor 1 (UBF1) and other rDNA regulatory factors to enhance RNA polymerase I-dependent transcription (11, 28, 69). We also show decreased UBF1 expression, suggesting that the entire cMYC-dependent regulatory axis is perturbed during hyperammonemia. Even though SL1, another transcription factor, forms a complex with UBF1 for transcriptional activation for RNA polymerase I (70, 71), there are no sequence-specific DNA binding sites for SL1, suggesting that reduced expression of UBF1 in hyperammonemia is sufficient to cause perturbations in ribosomal biogenesis. Our observations on ribosomal regulation in conjunction with polysome profiling cannot exclude the possibility that hyperammonemia can perturb mature ribosomal content. By blocking translation and proteasome degradation, we also noted that some of the ribosomal proteins are indeed susceptible to rapid regulation during cellular stress.

β-Catenin is a well-recognized upstream activator of cMYC expression both via direct transcriptional activation as well as through indirect activation of paired-like homeodomain transcription factor 2 (Pitx2), which activates cMYC (72, 73). In our studies, we noted loss of β-catenin and reduced expression of its transcriptional target cMYC. Interestingly, overexpression of β-catenin in C2C12 myotubes was able to restore cMYC expression as well as protein synthesis, showing for the first time a direct link between the loss of β-catenin–cMYC and decreased ribosomal content and protein synthesis. Thus, our studies on the mechanism by which hyperammonemia resulted in decreased active β-catenin point to a critical role of a heretofore little-recognized aspect of β-catenin degradation. Classical activation of β-catenin occurs during Wnt signaling, and inactivation is initiated by priming phosphorylation at Ser⁴⁵, followed by subsequent phosphorylation at other sites, including the GSK3β target Ser³³ (25, 55, 56). Phosphorylated β-catenin is then rapidly degraded via proteasome-mediated proteolysis. GSK3β is activated by dephosphorylation and inactivated by phosphorylation at the Ser⁹ site, with consequent changes in β-catenin stabilization and transcriptional activity (21, 25, 56, 74). In skeletal muscle, GSK3β is phosphorylated and inactivated by the phosphatidylinositol 3-kinase/Akt pathway (18, 19, 75). Interestingly, GSK3β expression and phosphorylation status were unaltered in skeletal muscle and myotubes during hyperammonemia. In addition to the classical GSK3β-dependent regulation of β-catenin, others have reported that the Ser/Thr kinase IKKβ phosphorylates and inactivates β-catenin (76). We previously reported IKKβ activation in the skeletal muscle during hyperammonemia (38). In the present studies, using association, loss- and gain-of-function, and in vitro kinase studies, we observed β-catenin phosphorylation and proteasome-mediated degradation via an IKKβ-dependent mechanism. These observations are similar to the recently reported regulation of β-catenin via IKKβ in mesenchymal stem cell differentiation (26), but our observations suggest that some of the effects of IKKβ can be mediated via β-catenin degradation. Interestingly, deletion of the priming phosphorylation sites on Ser^33,45 or the N-terminal domain of β-catenin, a target of GSK3β, did not affect β-catenin degradation during hyperammonemia. We also noted that despite mutation of the GSK3β target Ser³³ on β-catenin, β-catenin continues to be degraded via the proteasome. These observations suggest that during hyperammonemia, phosphorylation of β-catenin occurs via a noncanonical GSK3β-independent, IKKβ-dependent mechanism, following which the degradation of β-catenin occurs through the established proteasomal pathway. Whether IKKβ-mediated loss of β-catenin is relevant for other causes of muscle loss that are characterized by a decreased ribosomal content (e.g., immobilization and spinal cord injury) and whether it occurs in other organ systems need to be determined. These observations are therefore of high biological relevance, given that this regulatory mechanism of β-catenin degradation as a dominant pathway has not been described to date. Our observations also have therapeutic translational potential for dealing with muscle loss in chronic diseases, given the availability of small-molecule inhibitors of IKKβ (77).

In summary, our studies in myotubes and skeletal muscle in vivo show that during hyperammonemia, reduced protein synthesis and ribosomal content are accompanied by decreased ribosomal biogenesis. The classical regulation of ribosomal biogenesis by the β-catenin–cMYC axis is preserved. However, instead of GSK3β-dependent phosphorylation, IKKβ is the major upstream kinase implicated in β-catenin degradation (Fig. 9). Impaired ribosome biogenesis during muscle atrophy and noncanonical regulation of β-catenin during hyperammonemia have the potential for broad application, given that both perturbations of ammonia metabolism and muscle loss occur in a diverse group of chronic diseases.

FIG 9.

Schematic model of regulation of ribosomal biogenesis. Canonical regulation of β-catenin is mediated by Wnt ligand binding to its cognate receptor, Frizzled, leading to the phosphorylation and inactivation of GSK3β and the stabilization of β-catenin. Nuclear translocation of β-catenin transcriptionally activates target genes, including cMYC, which in turn induces ribosomal biogenesis and protein synthesis. During hyperammonemia, noncanonical regulation via GSK3β-independent, IKKβ-dependent phosphorylation and degradation of β-catenin results in impaired ribosomal biogenesis and protein synthesis.

MATERIALS AND METHODS

Materials.

All chemicals were of molecular biology grade and obtained from Sigma-Aldrich (St. Louis, MO). Anti-active β-catenin (catalog number 05-665) (this recognizes the unphosphorylated, stable form of β-catenin), anti-IKKβ (catalog number 05-535), and antipuromycin (catalog number MABE343) antibodies were obtained from EMD Millipore (Burlington, MA). Anti-phospho-β-catenin Ser^33/37, anti-phospho-GSK3β (catalog number 5558T), anti-GSK3β (catalog number 12456), and anti-RPL5 (catalog number 14568) antibodies were obtained from Cell Signaling (Danvers, MA). Anti-GSK3α (catalog number NB100-81943) and -GSK3β (catalog number NBP1-47470) were obtained from Novus Biologicals (Centennial, CO). Anti-total β-catenin (catalog number sc-7963) was obtained from Santa Cruz Biotechnology (Dallas, TX). Anti-cMYC (catalog number GTX103436), anti-RPL23 antibody (catalog number GTX130214), and anti-RPL32 antibody (catalog number GTX130214) were obtained from GeneTex, Inc. (Irvine, CA). Purified GSK3β, IKKβ, and β-catenin were obtained from SignalChem (Richmond, BC, Canada). shIKKβ, and shIKKα were from Joseph A. DiDonato at the Cleveland Clinic. shGSk3β and shGSK3α were obtained from Sigma-Aldrich (St. Louis, MO).

Human cirrhosis with hyperammonemia.

Vastus lateralis muscle specimens from patients with cirrhosis and healthy volunteer control subjects were obtained using a Bergstromm needle. The technique and clinical characteristics of the patients and controls were previously reported (43). Human studies were performed after obtaining written informed consent, and the studies were approved by the Institutional Review Board at the Cleveland Clinic and conformed to the Helsinki Declaration on human studies.

Hyperammonemic rat.

Gastrocnemius muscle from hyperammonemic PCA and sham-operated, pair-fed control rats were used as described previously (45). The PCA rat has been shown to have lower muscle mass than pair-fed controls, and the lower muscle mass is partially reversible by ammonia lowering or follistatin, a myostatin antagonist (45, 47, 50). Pair feeding was performed to ensure that reduced food intake by PCA rats did not contribute to the differences from the sham-operated rats. Animals were housed in individual cages in a 12-h-light/12-h-dark controlled environment. The details of animal maintenance were previously described (45). All animal studies were approved by the Institutional Animal Care and Use Committee at the Cleveland Clinic.

Cell culture.

In vitro cell culture studies were performed in differentiated C2C12 myotubes (ATCC CRL 1722; ATCC, Manassas, VA) as described by us previously (39). In brief, myoblasts were grown to confluence in proliferation medium (Dulbecco’s modified Eagle’s medium [DMEM] with 10% fetal calf serum), which was replaced by differentiation medium (DMEM plus 2% horse serum) for 48 h. Differentiation to myotubes has been reported by us to occur using this time course based on the fusion index and creatine kinase activity (78). Cells were treated with 10 mM ammonium acetate for the indicated times. This concentration has been reported by us to generate clinically relevant intracellular concentrations of ammonia (38, 79) and a muscle loss phenotype with reduced myotube diameter (45). To demonstrate that ammonia-induced β-catenin degradation occurs in other cell types as well, human embryonic kidney cells (ATCC CRL 1573; ATCC, Manassas, VA) were grown in Eagle’s minimum essential medium with 10% fetal bovine serum to confluence and treated with 10 mM ammonium acetate for 6 and 24 h, with untreated cells serving as controls. 6-Bromoindirubin-3′-oxime (BIO) at 2 μM and lithium chloride at 5 mM were used to inhibit GSK3β and thereby stabilize β-catenin expression (80).

RNA sequencing and bioinformatics analyses.

Total RNA was extracted from C2C12 myotubes that were untreated or treated with 10 mM ammonium acetate for 24 h using the RNeasy Plus minikit (Qiagen, Germantown, MD). Total RNA quality was evaluated using an Agilent 2100 bioanalyzer (Agilent Technologies, Santa Clara, CA), and the amount of the isolated RNA was determined using a NanoDrop ND-1000 spectrophotometer (Infinigen Biotechnology, Inc., City of Industry, CA). RNA-Seq libraries were generated using an Illumina TruSeq kit and by sequencing on an Illumina HiSEQ4000 instrument (all according to protocols provided by manufacturer) by the University of Chicago Genomics Facility. Sequence alignment and bioinformatics analyses were done by Novogene (Chula Vista, CA), using TopHat2 for mapping sequences, with a mismatch parameter set at 2 and other parameters at default settings. All the sequencing analyses (quality control, alignment, gene expression quantification, and differential expression analysis) were performed by Novogene using their standard pipeline for mouse RNA-Seq/transcriptome studies. TopHat2 has been widely used in RNA sequence analysis publications and has been used by Novogene as the mapping tool because TopHat2 can generate a database of splice junctions based on the gene model annotation file and thus provides a better mapping result than other nonsplice mapping tools. However, recent programs, including HiSat and STAR, are more robust in terms of resource utilization and mapping accuracy, but despite the limitations of TopHat2, the newer alignment programs generally yield similar results. For quantification of gene expression, we used HTSeq v0.6.1 to count the number of reads mapped to each gene. Read counts were normalized by conversion to fragments per kilobase of transcript per million mapped reads (FPKM). Differential expression analysis of two conditions/groups (two biological replicates under each condition) was performed using the DESeq2 R package (1.18.0). DESeq2 determines differential expression using a model based on the negative binomial distribution, and statistical tests were done using a Wald chi-square test. The resulting P values were adjusted using the Benjamini-Hochberg approach for controlling the false discovery rate (FDR) (81). Genes with an adjusted P value of <0.05 found by DESeq2 were assigned as differentially expressed. Heat maps were plotted by the mean of FPKM values in groups. Row and column were clustered by hclust, and the values were centered and scaled in the row direction.

The reference genome for mouse, GRCm38, was released by the Genome Reference Consortium in January 2012. It is based on the Mus musculus strain C57BL/6J. This assembly was used by UCSC to create their mm10 database that corresponds to GenBank assembly accession number GCA_000001635.8.

Outcomes from RNA sequencing analyses were uploaded into Qiagen’s IPA system and overlaid with the global molecular network in the Ingenuity Pathway Knowledge Base (IPKB) to determine the canonical pathways and gene networks to classify differentially expressed genes. Heat map and hierarchical cluster analyses were used to demonstrate the expression patterns of these differentially expressed genes.

Ribosomal abundance and components.

Total RNA (from an equal number of cells) was used as a measure of ribosome abundance and translational capacity, since the majority of total RNA is rRNA, and nearly two-thirds of the ribosome is RNA (28, 53, 82, 83). Eukaryotic 80S ribosomes comprise a large, 60S subunit and a small, 40S subunit (84). The 60S subunit is composed of rRNA and 47 ribosomal large proteins required for peptide bond formation, the critical step in protein synthesis. Expression levels of ribosomal proteins were quantified using real-time PCR and immunoblotting.

Polysome profiling.

For polysome profiling, differentiated C2C12 myotubes were treated with 10 mM ammonium acetate for 24 h, and equal numbers of untreated myotubes were used as a control. At the end of 24 h, untreated and AmAc-treated myotubes were treated with 100 μg/ml cycloheximide (CHX) for 10 min, followed by two washes with 1× phosphate-buffered saline (PBS) containing 100 μg/ml cycloheximide. Cells were then collected in 1× PBS containing 100 μg/ml cycloheximide and centrifuged for 3 min at 2,000 rpm, and the pellet was suspended in 200 μl of ice-cold lysis buffer (50 mM HEPES [pH 7.4], 250 mM KCl, 5 mM MgCl₂, 250 mM sucrose, 1% Triton X-100, 1.3% sodium deoxycholate, 1× protease cocktail inhibitor [catalog number P8340; Sigma], 1 mM 1,4-dithiothreitol [DTT], 200 U/ml RNase inhibitor [catalog number N261A; Promega]) and homogenized by aspiration and injection via a 26-gauge needle. The lysate was placed on ice for 10 min, followed by centrifugation at 13,000 × g for 10 min. Supernatants were transferred into a fresh Eppendorf tube, snap-frozen in liquid nitrogen, and stored at −80°C for subsequent analyses. Buffers containing 20 and 47% sucrose were prepared in a solution containing 10 mM HEPES (pH 7.4), 5 mM MgCl₂, 75 mM KCl, 1 mM DTT, 1× protease inhibitor cocktail, 4 U/ml RNase inhibitor, and 100 μg/ml cycloheximide and used for the gradient generation. Linear gradients were prepared using a Foxy Jr. gradient maker (Teledyne Isco, Lincoln, NE) and stored at 4°C for a minimum of 1 h. Two hundred microliters of the cell lysate was loaded on top of a 20 to 47% linear sucrose density gradient, followed by ultracentrifugation at 29,000 rpm for 4 h at 4°C. Gradients were eluted immediately after ultracentrifugation, fractions were collected using an Isco model fraction collector, and absorbance was measured using a UV absorbance monitor (Teledyne Isco, Lincoln, NE). The ratio of polysomes to polysomes and monosomes was quantified in ammonia-treated myotubes and normalized to that in untreated controls in 3 biological replicates. These experiments were performed using an equal number of myotubes. The use of an equal quantity of proteins or RNA may not be the best approach for these experiments because a lower number of ribosomes in a cell will result in lower total RNA (80 to 90% rRNA) and lower total protein concentrations due to decreased protein synthesis.

Ribosomal protein kinetics.

Since some of the ribosome proteins can be very stable after incorporation into ribosomes, the half-life of ribosome proteins was evaluated by a cycloheximide chase assay using previously reported methods, with some modifications (85). Since cycloheximide inhibits the translation of new protein synthesis, and MG132 prevents proteasome degradation, the expression of ribosomal proteins in response to cycloheximide is a measure of the rate of degradation, and the response to MG132 can be used as a measure of the rate of translation. The use of different combinations of cycloheximide, MG132, and hyperammonemia will allow for determination of protein stability. Briefly, C2C12 myotubes differentiated for 48 h were either untreated or treated with 20 μg/ml of CHX (Sigma-Aldrich) for 24 h to block translation as a measure of protein stability. To determine the proteasomal breakdown of the ribosomal proteins, myotubes were also treated with and without a potent proteasome inhibitor, MG132 (20 μM), for the last hour during the 24-h treatment with 10 mM ammonium acetate. Cells were harvested in ice-cold radioimmunoprecipitation assay (RIPA) buffer with 1× protease inhibitor cocktail at various time points. Total protein from the cell lysate was quantified using the bicinchoninic acid assay, and equal concentrations of samples were loaded onto 10% SDS-PAGE gels for immunoblotting for the expression of the ribosomal proteins RPL5, -23, and -32. Additionally, to dissect the role of mTORC1 inhibition during 24 h of hyperammonemia, myotubes were treated with and without rapamycin (1 μg/ml) for the last 12 h, expression levels of ribosomal protein mRNA were quantified by real-time PCR, and total and phosphorylated P70S6 kinase protein levels were determined by immunoblotting.

Proteomics studies in myotubes.

Untreated C2C12 myotubes and myotubes treated with 10 mM AmAc for 24 h were washed in ice-cold phosphate-buffered saline and centrifuged at 800 rpm at 4°C, and the cell pellets were transferred to 1.5-ml centrifuge tubes. A 200-μl aliquot of 8 M urea in 0.1 M Tris-HCl (pH 8) with a Complete Mini protease inhibitor cocktail (Roche) was added to each sample. The samples were placed on ice and lysed four times for 15 s each at 70% energy using an ultrasonic cell disruptor. Lysed samples were centrifuged, and the supernatant was transferred to a new Eppendorf tube. Five microliters of the supernatant was used for quantifying protein concentrations using the bicinchoninic acid assay. A 40-μg aliquot of each sample was removed for digestion. For protein digestion, the samples were reduced with DTT and alkylated with iodoacetamide prior to in-solution digestion. All samples were digested using trypsin (Promega, Madison, WI) by adding 1 μg trypsin in 50 mM ammonium bicarbonate and incubating the samples overnight at room temperature. The peptides were desalted using Sep-Pak C₁₈ columns (Waters Corporation, Milford, MA). These extracts were evaporated in a SpeedVac and then resuspended in 1% acetic acid to make up a final concentration of ∼0.2 μg/μl for liquid chromatography-mass spectrometry (LC-MS) analysis.

Unbiased proteomics were performed on a Thermo Scientific Fusion Lumos hybrid mass spectrometry system (Thermo Scientific, San Jose, CA). The high-performance liquid chromatography (HPLC) column used was a Dionex 15-cm- by 75-μm-internal-diameter Acclaim PepMap C₁₈, 2-μm, 100-Å, reversed-phase capillary chromatography column (Thermo Fisher, Waltham, MA). Five-microliter volumes of the extract were injected, and the peptides eluted from the column by an acetonitrile–0.1% formic acid gradient at a flow rate of 0.3 μl/min were introduced into the source of the mass spectrometer online. The microelectrospray ion source is operated at 1.9 kV. The acquisition method was a data-dependent-based MS/MS method that begins with a full MS scan in an Orbitrap analyzer (60,000 resolution), followed by selection of the parent ions in the MS scans, fragmentation by high-energy collision-induced dissociation (HCD), and detection with the Orbitrap analyzer. The data were analyzed using MaxQuant V1.5.2.8 with the search engine Andromeda, which is integrated in MaxQuant software, and the parameters used were default settings for an Orbitrap instrument. The database used to search the tandem MS (MS/MS) spectra was the UniProt mouse protein database containing 25,035 entries with an automatically generated decoy database (reversed sequences). Oxidation of methionine and acetylation of protein N termini were set as dynamic modifications, and carbamidomethylation of cysteine was set as a static modification. The false discovery rate (FDR) was set to 1% for both peptide and protein, with a minimum length of 7 amino acids, and two unique or razor peptides were required for positive identification. The “match between runs” feature of MaxQuant was used to transfer identifications to other LC-MS/MS runs based on their masses and retention times (maximum deviation of 0.7 min), and this was also used in quantification experiments. Quantifications were performed with the label-free quantitation method available in the MaxQuant program (86).

Quantitative real-time PCR.

Total RNA was extracted using the TRIzol reagent (Thermo Fisher Scientific, Waltham, MA) and quantified as previously described (47). Reverse transcription was performed using a Clontech Advantage RT-for-PCR kit (TaKaRa Bio USA, Mountain View, CA), followed by quantification using real-time PCR (RT-PCR) on a Stratagene RT-PCR thermocycler. The primer sequences are shown in Table 2.

TABLE 2.

Primers used in this study

Primer	Positions	Sequence	Product size (bp)
Hu 45S rRNA FW	851–868	5′-GAACGGTGGTGTGTCGTT-3′	130
Hu 45S rRNA Re	961–980	5′-GCGTCTCGTCTCGTCTCACT-3′
Mu 45S rRNA FW	434–455	5′-CTGAGAAACGGCTACCACATC-3′	107
Mu 45S rRNA Re	519–541	5′-GCCTCGAAAGAGTCCTGTATTG-3′
RT 45S rRNA FW	984–1004	5′-AAGACGAACCAGAGCGAAAG-3′	98
RT 45S rRNA Re	1062–1082	5′-TCGGAACTACGACGGTATCT-3′
Mu RPS26 FW	215–235	5′-GTGCTTCCCAAGCTCTATGT-3′	81
Mu RPS26 Re	276–296	5′-GCGGGATCGATTCCTAACAA-3′
Mu RPS21 FW	147–166	5′-CAAGACCTGCGTGGTAACT-3′	101
Mu RPS21 Re	229–245	5′-GTTGCTTGCGGAGCATTT-3′
Mu RPS19 FW	221–240	5′-AAGTCCGGGAAGCTGAAAG-3′	103
Mu RPS19 Re	308–328	5′-GAAGCAGCTCGTGTGTAGAA-3′
Mu RPL5 FW	485–507	5′-GGAGGTGAATGGAGGTGAATAC-3′	97
Mu RPL5 Re	562–582	5′-GTCTTGCCCGAACTACAACT-3′
Mu RPL23 FW	203–224	5-CTCTGTGAAGGGAATCAAGGG-3′	129
Mu RPL23 Re	310–332	5′-TACATCCAGCAGTGGTAATTCG-3′
Mu RPL32 FW	225–245	5′-CGGTTATGGGAGCAACAAGA-3′	128
Mu RPL32 Re	331–353	5′-TCTTACTGTGCTGAGATTGCTC-3′
Hu RPS26 FW	1043–1063	5′-TCCCAAAGTGCTGGGATTAC-3′	84
Hu RPS26 Re	1103–1127	5′-CCAACCATACGACCACAATTAAAG-3′
Hu RPS21 FW	174–195	5′-AGGTCACAGGCAGGTTTAATG-3′	98
Hu RPS21 Re	252–272	5′-ATTCCATTCTCCGATTGGCC-3′
Hu RPS19 FW	18–40	5′-GTATTCTCCACCACTGTTCCTT-3′	105
Hu RPS19 Re	101–123	5′-CTCTCTCTATTACACTCCGGGA-3′
Hu RPL5 FW	135–157	5′-TAGAAGACGACGAGAGGGTAAA-3′	115
Hu RPL5 Re	227–250	5′-TGATAGTTCGTGTGACAAACAGA-3′
Hu RPL23 FW	872–894	5′-ACTGCTCAATGACTGGCTATACA-3′	105
Hu RPL23 Re	956–977	5′-TAATCCCAACACTTTCGGAGG-3′
Hu RPL32 FW	708–730	5′-GCACAGACTAGCCTTAGTCATC-3′	92
Hu RPL32 Re	777–800	5′-GGTAGGTTCTGAGACACTTGAAGA-3′

Immunoblotting.

Protein extraction from skeletal muscle and myotubes followed by quantification was performed as previously described (51). In brief, cell lysates (30 μg protein) were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, electrotransferred to a polyvinylidene difluoride (PVDF) membrane, and incubated overnight with the appropriate primary antibody, followed by washes and incubation with horseradish peroxidase (HRP)-tagged secondary antibody. Signals were revealed by an enhanced chemiluminescence (ECL) reagent (GE Healthcare, Marlborough, MA) and visualized by autoradiography. Expression of β-actin was used to confirm equal loading of protein for all immunoblot assays. Blots were quantified using ImageJ software (87).

Immunoprecipitation assays.

Cells were harvested in lysis buffer containing 20 mM Tris (pH 7.4), 137 mM NaCl, 1% NP-40, 1 mM phenylmethylsulfonyl fluoride (PMSF), 20% glycerol, 10 mM sodium fluoride, 1 mM sodium orthovanadate, and 2 μg/ml leupeptin and aprotinin. The lysates were cleared by centrifugation at 12,000 × g for 15 min at 4°C. The supernatant solutions were used for immunoprecipitation. Proteins (β-catenin and IKKβ) were immunoprecipitated by incubating the cell lysates with protein A/G-agarose beads (Santa Cruz Biotechnology, Santa Cruz, CA) and the respective antibodies at 1:500 dilutions, overnight at 4°C. Associated proteins were probed with the appropriate antibodies at a 1:1,000 dilution overnight, followed by incubation with HRP-tagged secondary antibody. Signals were revealed with ECL reagent (GE Healthcare, Marlborough, MA) and visualized by autoradiography. Input protein loading and IgG for nonspecific responses were also blotted.

Transfection.

Mutant β-catenin constructs that were resistant to phosphorylation by GSK3β (S33Y) and casein kinase 1α (S45K) as well as N-terminally deleted β-catenin were generated as previously reported (88). Cassettes with shRNA to IKKβ and IKKα were made from Sigma Mission shRNA clones (3 clones each) as a multimer driven by the U6 promoter for each clone. Myoblasts were transfected using Mirus TransIT-2020 transfection reagent (Mirus, Madison, WI). To determine if the responses to chemical inhibition of GSK3β using either lithium chloride or BIO were specifically due to GSK3β, genetic depletion of GSK3α and GSK3β was achieved using shRNA, stable clones were selected, and protein synthesis in response to hyperammonemia was studied.

Protein synthesis in myotubes.

The rate of muscle protein synthesis was quantified in differentiated C2C12 myotubes in response to 10 mM ammonium acetate or the vehicle alone as previously described by us (51). Incorporation of either [³H]phenylalanine or puromycin was used for these assays. In brief, following incubation in 10 mM ammonium acetate or the vehicle, myotubes were washed, incubated with 1 μCi of [³H]phenylalanine for 3 h, and then washed and lysed, and the protein content and radioactivity were quantified. The rate of incorporation of [³H]phenylalanine was expressed as counts per minute per milligram of protein. Phenylalanine was the preferred amino acid because it is not synthesized (essential amino acid) or catabolized by the myotubes.

For puromycin incorporation as a readout for protein synthesis, a protocol described previously by us was used (51). In brief, ammonia-treated or untreated control myotubes were incubated with 1 μg/ml puromycin for 30 min, and cell lysates were subjected to immunoblotting using antipuromycin antibody (1:10,000 dilution). Densitometry of all the bands on each lane across the entire molecular weight range of puromycin-incorporated protein was quantified and normalized using β-actin protein. All experiments were done in at least 3 biological replicates.

Reporter assays.

β-Catenin signaling was assessed using Topflash and control Fopflash luciferase reporters. The Topflash reporter plasmid contains three copies of 2 sets of wild-type T-cell factor (TCF) binding sites driving luciferase gene expression; the control construct Fopflash contains mutated TCF binding sites. Topflash and Fopflash were obtained from Upstate Cell Signaling Solutions (Billerica, MA). A Renilla luciferase reporter vector was used as an internal control and was obtained from Promega (Madison, WI). These assays were performed as previously reported, and luciferase luminescence normalized to values for Renilla transfection (89).

In vitro kinase assays.

Kinase reactions were performed by mixing purified glutathione S-transferase (GST)-tagged β-catenin (500 ng) with either purified GSK3β (10 ng) or IKKβ (10 ng) in kinase assay buffer containing 5 mM morpholinepropanesulfonic acid (MOPS) (pH 7.2), 2.5 mM glycerol phosphate, 5 mM MgCl₂, 1 mM EGTA, 0.4 mM EDTA, and 0.05 mM DTT with 50 μM ATP. Reaction mixtures without β-catenin, GSK3β, or IKKβ served as negative controls. The reaction mixture was incubated at 30°C for 60 min, and the reaction was stopped by adding SDS-PAGE sample buffer, followed by incubation at 95°C for 5 min. Samples were separated on a 10% SDS-PAGE gel, transferred to a PVDF membrane, and incubated with phospho-β-catenin (Ser^33/37/Thr⁴¹) that detects only the phosphorylated form but not unphosphorylated β-catenin (Cell Signaling Technology, Danvers, MA). Purity of the proteins used for the kinase assay was tested by immunoblotting.

Statistical analyses.

All experiments were performed in at least 3 biological replicates in myotubes or in vitro experiments. All data are expressed as means ± standard deviations (SD) unless otherwise stated. Analysis of variance with a Bonferroni post hoc or Student t test was performed to compare quantitative variables, and a P value of <0.05 was considered significant.

Data availability.

All sequencing files have been uploaded to the NCBI Sequence Read Archive (SRA) (BioProject accession number PRJNA495054).