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. Author manuscript; available in PMC: 2021 Jul 1.
Published in final edited form as: Metabolism. 2020 May 1;108:154257. doi: 10.1016/j.metabol.2020.154257

Increased AMP deaminase activity decreases ATP content and slows protein degradation in cultured skeletal muscle

Patrick R Davis a,1, Spencer G Miller a,2, Nicolas A Verhoeven a, Joshua S Morgan b, David A Tulis b, Carol A Witczak a,b,c,3, Jeffrey J Brault a,b,c,2,*
PMCID: PMC7319876  NIHMSID: NIHMS1589986  PMID: 32370945

Abstract

Background:

Protein degradation is an energy-dependent process, requiring ATP at multiple steps. However, reports conflict as to the relationship between intracellular energetics and the rate of proteasome-mediated protein degradation.

Methods:

To determine whether the concentration of the adenine nucleotide pool (ATP + ADP + AMP) affects protein degradation in muscle cells, we overexpressed an AMP degrading enzyme, AMP deaminase 3 (AMPD3), via adenovirus in C2C12 myotubes.

Results:

Overexpression of AMPD3 resulted in a dose- and time-dependent reduction of total adenine nucleotides (ATP, ADP and AMP) without increasing the ADP/ATP or AMP/ATP ratios. In agreement, the reduction of total adenine nucleotide concentration did not result in increased Thr172 phosphorylation of AMP-activated protein kinase (AMPK), a common indicator of intracellular energetic state. Furthermore, LC3 protein accumulation and ULK1(Ser 555) phosphorylation were not induced. However, overall protein degradation and ubiquitin-dependent proteolysis was slowed by overexpression of AMPD3, despite unchanged content of several proteasome subunit proteins and proteasome activity in vitro under standard conditions.

Conclusions:

Altogether, these findings indicate that a physiologically relevant decrease in ATP content, without a concomitant increase in ADP or AMP, is sufficient to decrease the rate of protein degradation and activity of the ubiquitin-proteasome system in muscle cells. This suggests that adenine nucleotide degrading enzymes, such as AMPD3, may be a viable target to control muscle protein degradation and perhaps muscle mass.

Keywords: Energetics, adenine nucleotides, protein degradation, ubiquitin-proteasome system, C2C12 myoblasts

1. Introduction

Protein degradation in skeletal muscle is largely mediated by two proteolytic processes: the ubiquitin-proteasome system (UPS) and the autophagic-lysosomal pathway. Both of these processes require cellular energy as ATP. The UPS targets specific proteins by attachment of a polyubiquitin chain and then degrades that protein by the multi-subunit proteasome [1]. The UPS is highly energy-dependent [2, 3], requiring ATP at multiple steps including the ubiquitin-activating enzyme (E1) [4], assembly of the 26S proteasome [5, 6], 19S gate opening [7, 8], and protein unfolding [9]. The other major proteolytic process, the autophagic-lysosomal pathway, is not directly energy-dependent but does require ATP for proton pumping in order to create the acidic environment where lysosomal proteases are active [10]. Further, the autophagic-lysosomal pathway is activated by AMP-activated protein kinase, which itself is sensitive to the AMP/ATP ratio [11]. Therefore, ATP concentration or the ATP/ADP ratio, which partly determines the apparent Gibbs free energy of ATP hydrolysis (ΔGATP = ΔG°ATP + RTln([ADP] • [Pi]/[ATP]), may be an important regulator of protein degradation rate.

While the dependence of the UPS on ATP is well documented (for reviews see [12–14]), there are disparate reports on how intracellular ATP within the physiological range (3–5 mM in mammalian cells [15]) affect protein degradation. Several studies have demonstrated that the rate of proteasome activity in concentrated proteasome preparations from heart tissue is maximal between 50 and 100 μM ATP [16–18], indicating that low levels of ATP accelerate the rate of protein degradation [16]. Conversely, it has been reported that the rate of non-lysosomal (largely proteasome-mediated) protein degradation in fibroblasts [19], reticulocytes, and cell-free preparations [2] is maximal at physiological ATP concentrations and any reductions in intracellular ATP by metabolic inhibitors (e.g. dinitrophenol) slow the rate of protein degradation. Notably, the decrease in ATP with such inhibitors is coincident with increases in ADP and AMP concentrations, which decrease the apparent Gibbs free energy of ATP hydrolysis and lead to phosphorylation/activation AMP-activated protein kinase [20]. Therefore, by using metabolic inhibitors, it is not possible to test independently the effects of decreased ATP in live cells.

The purpose of this study was to determine whether decreased ATP content, without increased ADP and apparent decreased ΔGATP, was sufficient to decrease protein degradation in live cells. Therefore, we overexpressed the metabolic enzyme AMP deaminase isoform 3 (AMPD3; AMP + H2O ⇒ IMP + NH4+) in cultured skeletal muscle myotubes. AMPD3 is highly induced (mRNA increases of up to 100-fold) during skeletal muscle atrophy [21, 22] and been shown to decrease total adenine nucleotide (AdN: ATP + ADP + AMP) content in cardiac cells [23]. Given that UPS-mediated protein degradation depends on ATP and is a major system for degrading intracellular proteins, we hypothesized that decreased ATP content would reduce the overall rate of protein degradation in live cells.

2. Materials and methods

2.1. Cell Culture

Mouse skeletal muscle myoblasts (C2C12, ATCC) were grown on gelatin (Sigma, G9391) coated plates in DMEM supplemented with 10% heat-inactivated FBS (Life Technologies), 100 IU/ml penicillin, and 100 mg/ml streptomycin. When cells reached 75–90% confluence, the media was switched to low serum DMEM (2% horse serum, Hyclone) with penicillin/streptomycin to induce differentiation into multi-nucleated myotubes. Five to six days later, when fusion of myoblasts to myotubes appeared complete, cultures were transduced with serotype 5 (DE1/E3) adenoviruses encoding GFP alone as a control or encoding AMPD3-IRES-GFP (Vector Biolabs; Malvern, PA). Prior to use, viruses were purified by cesium chloride gradient, and viral titers were determined by the end-point dilution assay in HEK 293 cells. Transduction efficiency, as determined by visual inspection of GFP fluorescence, was near 100% in all experiments.

2.2. Nucleotide measurements

Myotubes were washed twice with ice-cold PBS. After PBS was removed completely, proteins were precipitated using ice-cold 0.5 M PCA plus 5 mM EDTA, which was prepared daily. Cells were scraped free from the wells and transferred to Eppendorf tubes. After a brief sonication, samples were centrifuged at 4°C to separate the precipitated protein pellets from the extracts. Protein pellets were re-suspended in 0.2 N NaOH for protein quantification by BCA Assay (Pierce). Extracts were neutralized by the addition of ice-cold 1 M KOH and centrifuged at 4°C to remove potassium perchlorate salt. Samples were stored at −80°C until analysis. Adenine nucleotide concentrations (ATP, ADP, AMP) and degradation products (IMP and inosine) were determined by ultra-performance liquid chromatography (UPLC) using a Waters Acquity UPLC H-Class system and an Acquity UPLC HSS T3 1.8 μm, 2.1 mm × 150 mm column (Waters), as done previously [24].

2.3. Protein analysis

Proteins were extracted using RIPA buffer including protease inhibitors (Complete, Roche) and quantified by BCA Assay (Pierce). Equal amounts of protein were separated by SDS-PAGE then transferred to polyvinylidene difluoride (PVDF) membranes. Equal total protein loading per lane and membrane transfer were confirmed by Ponceau S staining of the membrane [25]. Antibodies were purchased from Abcam (AMPD3, ab118230), Enzo (19S Rpt6, PW9265; 19S Rpt5, PW8770; 20S β5, PW8895), Thermo Fisher (GFP, MA5–15256) and Cell Signaling (AMPKα, #5831; P-AMPKα (Thr172), #2531; LC3B, #3868; ULK1, #8054; P-ULK1 (Ser555), #5869). Secondary antibodies conjugated to HRP (Cell Signaling #7074, ThermoFisher #31444) were used and visualized with Western Chemiluminescence HRP Substrate (EMD Millipore). Band intensities were captured using a Bio-Rad Chemi Doc XRS imager and analyzed using Image Lab Software (Version 5.2.1, Bio-Rad). Estimated molecular weights of specific proteins were calculated relative to PageRuler Plus protein ladder (ThermoFisher), which was included on every gel.

2.4. AMPD activity

Total AMPD activity was determined by the formation of IMP in cell homogenates as done previously [26]. Myotubes were collected by scraping cells in homogenization buffer (100 mM KCl, 50 mM imidazole, 1 mM DTT, pH 7.0) and then lysed with a Dounce Homogenizer. Homogenates were added to reaction buffer (150 mM KCl, 50 mM Imidazole, 10 mM AMP, pH 7.0, 30°C) to start the reaction. Fractions of the reaction mixture were removed at predetermined times, and cold 0.5N PCA was immediately added to stop enzymatic activity. PCA-treated fractions were neutralized with ice-cold 1.0 N KOH, and IMP was measured in supernatants by UPLC as described above. AMPD activity was calculated as the rate of IMP formation per total protein content per minute.

2.5. Proteasome activity

Activity of the chymotrypsin-like site of the proteasome was measured as described by Kisselev and Goldberg [27] using the fluorogenic peptide substrate Suc-LLVY-amc (100 μM final, Bachem). In brief, myotubes were lifted from plates with trypsin-EDTA and centrifuged in 1.7 mL Eppendorf tubes. Pellets were re-suspended and proteasomes were concentrated in a cytosolic extraction buffer (50 mM Tris-HCl, 250 mM sucrose, 5 mM MgCl2, 0.5 mM EDTA, 1 mM DTT, 2 mM ATP, 0.025% Digitonin). Cytosolic extracts were added to 10 volumes of proteasome activity buffer (50 mM Tris-HCl, 40 mM KCl, 5 mM MgCl2, 1 mM DTT, 0.5 mM ATP, 0.05 mg/mL BSA) in a black 96-well plate (Greiner). Samples were excited at 380 nm and absorbance was measured at 460 nm continually over 15 min. Results were normalized to total protein content (DC protein assay, Bio-Rad).

2.6. Autophagy flux

The flux through autophagy was determined by the accumulation of LC3 II in myotubes that had been treated with a lysosomal acidification inhibitor [28]. Cells were treated with 0.1 μM concanamycin A or DMSO (vehicle) for 6 hours after which proteins were collected in RIPA buffer. LC3 II was identified by immunoblot as described above.

2.7. Ubiquitin/proteasome-dependent proteolysis by Ub-GFP accumulation

To determine the effect of AMPD3 on ubiquitin-dependent proteasome activity, C2C12 myoblasts were transfected with plasmids encoding Ub-R-GFP (Addgene #11939) or UbG76V-GFP (Addgene #11941) [29] using FuGENE HD (Promega) per manufacturer’s recommended protocol. These GFP constructs represent different classes of substrates for the proteasome: Ub-R-GFP functions as an N-end rule substrate; and UbG76V-GFP, in which the ubiquitin is uncleavable, acts as a ubiquitin fusion degradation signal substrate [30]. Given that these constructs are the same size and use the same CVM promoter and thus are synthesized at the same rate, accumulation of GFP indicates a slowing of ubiquitin-dependent proteasomal degradation [29].

For studies of myotubes, myoblasts were transfected with the Ub-GFP plasmids in 6-well plates using Fugene HD. Twenty-four hours later, media was switched to differentiation media. Myotubes were allowed to grow for 4 or 5 days before transduction of adenovirus encoding AMDP3. Proteins were collected in RIPA buffer 24 or 48 hours later, all on the same day of differentiation. The relative expression of GFP and AMPD3 was determined by immunoblot.

For studies in myoblasts, C2C12 myoblasts were transfected in 24-well plates using Fugene HD with plasmids in 5 different combinations: (1) Ub-R-GFP + AMPD3, (2) UbG76V-GFP + AMPD3, (3) Ub-R-GFP + empty vector, (4) UbG76V-GFP + empty vector, and (5) empty vector alone. To demonstrate the responsiveness of this assay to proteasome inhibition, in a parallel experiment myoblasts were transfected similarly with the Ub-R-GFP, UbG76V-GFP, and empty vector but instead of co-transfection with AMPD3, myoblasts were treated with 1μM bortezomib (Biovision, cat # 1846) or vehicle (DMSO) 4 hours before cells were collected. The same total amount of plasmid (0.4 μg/well) was used in all conditions. Thirty hours after plasmids were given, myoblasts were lifted from the wells using 0.25% trypsin / 2.21 mM EDTA and transferred to 1.7 ml centrifuge tubes. After adding 10% FBS + DMEM to inhibit further activity of trypsin/EDTA, individual cells were then analyzed for GFP intensity by flow cytometry (BD Accuri C6 Flow Cytometer) using CFlow Plus software (Accuri).

2.8. Long-lived protein degradation

Degradation of long-lived proteins was determined from the release of radiolabeled tyrosine as previously described [31]. On day 4 of differentiation, C2C12 myotube proteins were radiolabeled with 5 μCi / ml L-[3,5-3H]-tyrosine (PerkinElmer, #NET127) for 24 hours. Cells were then washed and incubated in differentiation media supplemented with 2mM non-radioactive tyrosine for 2 hours to allow the breakdown of short-lived proteins. Cells were washed again and then transduced with either GFP or AMPD3 adenovirus in fresh media with 2mM tyrosine (time 0). Samples of the media were taken at 0, 4, 24, 28, and 48 hours. Proteins were precipitated by the addition of 10% TCA. After the last time point, media was removed completely, and the cells were solubilized in 0.2 N NaOH. Radioactivity was measured by scintillation counting of TCA soluble supernatants of the media samples and in the solubilized cell fraction. Degradation rates were calculated as the fraction of total radioactivity incorporated per hour.

2.9. Protein synthesis

Myotubes were pulsed with 5 μCi/ml L-[3,5-3H]-tyrosine for two hours and then rapidly washed twice with ice-cold phosphate buffered saline. Proteins were precipitated with 10% TCA and pelleted by centrifugation. Protein pellets were re-suspended in 0.1N NaOH for total protein measurement by BCA assay (Pierce) and total radioactivity measurement by scintillation counter. Protein synthesis rate was expressed as radioactivity incorporated into TCA precipitable proteins per microgram of protein [32].

2.10. Statistics

All data are expressed as mean ± standard deviation. To assess normality, quantile-quantile (Q-Q) plots were generated for all data sets. All plots of cumulative distribution appeared linear, which indicates that data were drawn from Gaussian (normal) distributions.Significant differences were assessed using two-way ANOVA (for comparisons of % degraded, protein synthesis, LC3 II, and proteasome subunits) with Sidak post-hoc analysis for multiple comparisons if significance was detected. One-way ANOVA was used for comparisons of nucleotides, mRNA, ULK1, and AMPK with a Bonferroni post-hoc analysis for multiple comparisons if significance was detected. Unpaired student t-tests were used for comparisons of degradation rate, AMPD activity, total protein, ubiquitin, and proteasome activity. All analyses were performed using GraphPad Prism, version 8.4.1. All experiments were repeated at least once using independent cell preparations to verify the reproducibility of our results.

3. Results

3.1. AMPD3 decreases adenine nucleotides in C2C12 myotubes

To determine appropriate conditions to decrease ATP content without increasing the ADP/ATP ratio, initial experiments tested different doses and durations of treatments with AMPD3 encoding adenovirus in myotubes. Increasing amounts of AMPD3 adenovirus from 1 to 20 × 106 pfu / ml media for 48 hours resulted in significant decreases in the concentrations of ATP, ADP (Figure 1A), and AMP (Figure 1B). Concomitantly, IMP levels increased from non-detectable with no AMPD3 virus to >1 μmol/g protein with all doses of AMPD3 adenovirus (Figure 1C). Accordingly, the sum of adenine nucleotides and IMP decreased with increasing does of the AMPD3 adenovirus (Figure 1D). Notably, the ADP/ATP ratio decreased with the lowest dose AMPD3 virus, and the AMP/ATP ratio was unchanged (Figure 1E). Further, no morphological differences were noted at these doses of AMPD3 adenovirus, but cytopathic effects (change in myotube shape and/or detachment) became evident by light microscopy when adenovirus was greater than 20 × 106 pfu / ml (data not shown). Therefore, subsequent experiments used 2 × 106 pfu / ml AMPD3 adenovirus because this low dose resulted in a significant reduction in ATP but avoided the extreme decrease in total adenine nucleotides as well as potential toxicity.

Figure 1. AMP deaminase 3 (AMPD3) expression decreases adenine nucleotides (AdN) and increases IMP in a dose- and time-dependent manner.

Figure 1.

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C2C12 myotubes were transduced with AMPD3 adenovirus, and nucleotides were extracted 48 hours later. (A) ATP, ADP, (B) AMP, and (C) IMP were measured by UPLC. (D) The sum of nucleotides (ATP + ADP + AMP + IMP) and (E) nucleotide ratios ADP/ATP and AMP/ATP were then calculated. C2C12 myotubes were transduced with 2 × 106 PFU/ml of GFP or AMPD3 adenovirus, and nucleotides were extracted 24, 48, or 72 hours later. (F) ATP, ADP, (G) AMP, and (H) IMP were measured by UPLC. (I) The sum of nucleotides (ATP + ADP + AMP + IMP) and (J) nucleotide ratios ADP/ATP and AMP/ATP were then calculated. Data are presented as mean ± SD. n = 3 per dose or time point.

To determine the time-course changes in adenine nucleotides, extracts were collected 24, 48, or 72 hours after myotubes were transduced with 2 × 106 pfu / ml AMPD3 or GFP (control) adenovirus. ATP and ADP concentrations were significantly reduced by 48 hours and remained reduced at 72 hours compared to GFP controls (Figure 1F). AMP decreased (Figure 1G) and IMP increased at 24, 48, and 72 hours post-transduction with AMPD3 compared to controls (Figure 1H). The total adenine nucleotide pool (ATP+ADP+AMP) plus IMP significantly decreased at 48 hours and remained so at 72 hours (Figure 1I). The ADP/ATP ratio, which is an indication of the ΔGATP, did not change over the time course (Figure 1J). The GFP adenovirus had no significant effect on any of the nucleotide measures as compared to non-transduced myotubes.

3.2. Increased AMPD3 activity does not induce AMPK phosphorylation

As expected, overexpression of AMPD3 by adenoviral delivery resulted in a robust, time-dependent increase in AMPD3 protein content, whereas AMPD1 was not reproducibly detectable (Figure 2A). Further, the rate of AMP deaminase activity, which includes the activity of all AMPD isoforms, doubled at 24 h and was 9-fold greater at 48 hours (Figure 2B). However, consistent with our measures of the adenine nucleotide ratios (Figure 1J), overexpression of AMPD3 did not result in significant changes to the amount of alpha subunit of the energy-sensing kinase, AMP-activated protein kinase (AMPK), nor the phosphorylation of AMPKα at Thr172 (Figure 2C, D). Therefore, despite the decreased measures of ATP (Figure 1F), AMPD3 overexpression does not result in an energy deficit that would be detectable by AMPK.

Figure 2. Increased AMPD3 activity does not induce phosphorylation of AMPK (Thr172).

Figure 2.

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(A) Protein content of AMPD3 and AMPD1 were measured by immunoblot following transduction with AMPD3 adenovirus. (B) Total AMPD activity was assessed as rate of IMP formation in homogenates of control (GFP) and AMPD3 transduced myotubes. (C) Representative blots and (D) quantification of total AMPK and P-AMPK (Thr172) content as measured by immunoblot. Data are presented as mean ± SD. n= 6 for AMPD activity assays. n = 5 for western immunoblotting. **p<0.01 vs GFP, ****p<0.0001 vs GFP

3.3. Autophagy flux is not affected by AMPD3

To determine whether the decrease in ATP content might alter autophagic flux, we measured the accumulation of LC3 I and II in the presence of concanamycin A [28], an inhibitor of the vacuolar H+-ATPase that leads to neutralization of lysosomal pH and inhibition of lysosomal acid-dependent proteases. Concanamycin A treatment led to a clear increase in LC3 I and LC3II (Figure 3A). However, there was no difference between AMPD3 and GFP groups in the accumulation of LC3 I, the accumulation of LC3 II, or the ratio of LC3 II /LC3 I (Figure 3B–D). We next tested whether ULK1, a critical kinase in the activation of autophagy that is activated by AMPK [11], was activated by AMPD3. The phosphorylation of ULK1 Ser555, but not the total amount of ULK1, decreased over time following AMPD3 overexpression (Figure 3E–G). Further, AMPD3 tended to reduce the P-ULK1/ULK1 ratio (Figure 3H). Thus, these findings suggest that AMPD3 overexpression and a decrease in ATP is not sufficient to stimulate lysosomal protein degradation.

Figure 3. Autophagy flux is not affected by AMPD3.

Figure 3.

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C2C12 myotubes were treated with concanamycin A (Concan A) to inhibit lysosomal acidification. (A) Representative blot and quantification of (B) LC3-I, (C) LC3-II and (D) ratio of LC3 II / LC3 I in GFP or AMPD3 transduced C2C12 myotubes. (E) Representative blot and quantification of (F) ULK1, (G) phospho-ULK1 (Ser555) and (H) ratio of P-ULK1 / ULK1 in AMPD3 transduced C2C12 myotubes for up to 48 hours. Data are presented as mean ± SD. n = 6 per group. #p<0.001 main effect vs vehicle, **p<0.01 vs 0 hr

3.4. Ubiquitin/proteasome-dependent proteolysis slows without changes in proteasome content

To examine ubiquitin/proteasome-specific protein degradation in myotubes, myoblasts were transfected with plasmids encoding mono-ubiquitinated GFP constructs [29]. The ubiquitin moiety increases the susceptibility for ubiquitin chain elongation and thus targets the GFP protein to the proteasome. Therefore, a greater accumulation of GFP is an indication of slower degradation by the proteasome. Transduction of myotubes with AMPD3 adenovirus for 48 h, but not at 24 h, resulted in a significant increase (p<0.001) in GFP protein of both proteasome substrates Ub-R-GFP (Figure 4A) and UbG76V-GFP groups compared to controls (Figure 4B), suggesting that flux through the proteasome is slower. Importantly, the amount of proteasome subunit proteins β5, Rpt5, and Rpt6 were not different with 48 hours of AMPD3 (Figure 4C). Further, proteasome activity, measured using concentrated proteasome preparations in standard in vitro conditions, was not different in myotubes overexpressing AMPD3 (Figure 4D).

Figure 4. Ubiquitin-proteasome mediated protein degradation slows without changes in proteasome content.

Figure 4.

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C2C12 myoblasts were transfected with plasmids encoding ubiquitinated GFP constructs, which are targeted to the proteasome, and then differentiated into myotubes. Myotubes were then transduced with AMPD3 adenovirus. GFP protein content was determined by immunoblot in (A) Ub-R-GFP and (B) UbG76V-GFP transfected cells after 0 (control), 24, or 48 hours of AMPD3 transduction. n = 4 per group. (C) Protein content of proteasome subunits after 48 hours of GFP or AMPD3 transduction in C2C12 myotubes was determined by immunoblot. n = 6 per group. (D) Proteasome activity was measured in-vitro under standard conditions in concentrated proteasome preparations. n = 9 per group. Data are presented as mean ± SD. ***p<0.001 vs control

Measures of GFP by immunoblot are unable to discern whether GFP intensity differs per cell or differs by the percent of cells expressing GFP. Therefore, myoblasts were transfected with the Ub-GFP constructs with or without AMPD3, and then 30 hours later GFP intensity was measured on a cell-by-cell basis using flow cytometry (Figure 5A). AMPD3 overexpression led to a significant increase in GFP intensity per GFP positive myoblast for both Ub-R-GFP and UbG76V-GFP (Figure 5B). To confirm our system detects changes in proteasomal activity, a parallel study used the specific proteasome inhibitor bortezomib. As expected, bortezomib treatment lead to increases in GFP intensity of both Ub-GFP constructs (Figure 5C).

Figure 5. AMPD3 slows ubiquitin/proteasome-mediated protein degradation.

Figure 5.

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C2C12 myoblasts were transfected with plasmids encoding Ub-R-GFP or UbG76V-GFP, and AMPD3 or empty vector, or just with empty vector alone. (A) Myoblasts were analyzed for GFP intensity by flow cytometry and (B) quantified. (C) In a parallel experiment, the proteasome inhibitor Bortezomib was used as a positive control for reduced proteasome-mediated protein degradation. n = 3 per group. Data are presented as mean ± SD. ***p<0.001 vs empty vector/vehicle

3.5. AMPD3 decreases overall protein degradation but does not alter protein synthesis

The observed accumulation of Ub-GFP in myotubes (Figure 4) and myoblasts (Figure 5) when AMPD3 is over-expressed suggests a slowing of protein degradation. However, because GFP accumulation is dependent on the rate of protein synthesis (which is rapidly driven by the CMV promoter) and duration of the experiment, the fold-changes in GFP accumulation do not necessarily indicate similar fold changes in rates of overall protein degradation. Therefore, we next tested the effects of AMPD3 on the rate of long-lived protein degradation, which include the myofibrillar proteins. The rate of proteolysis was measured by the release of [3H]-tyrosine in AMPD3 compared to GFP myotubes. No significant differences were detected in proteolysis rate during the first 24 h after adding the adenoviruses, likely related to the modest changes in adenine nucleotides at that time point (cf. Figure 1). However, the rate decreased significantly with AMPD3 when measured between 28 and 48 hours (Figure 6A).

Figure 6. AMPD3 decreases overall protein degradation without altering protein synthesis.

Figure 6.

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(A) Protein degradation was determined by the release of label to the media. Samples were collected at 0, 4, 24, 28, and 48 hours post GFP or AMPD3 transduction. Rate of overall protein degradation was calculated as % of total incorporated L-[3,5-3H]-tyrosine released into the media per hour. (B) Protein synthesis as measured by incorporation of 3H-tyrosine into TCA-precipitable protein. n = 4 per group. Data are presented as mean ± SD. **p<0.01 vs GFP

Protein synthesis rates were determined by giving a 2 hour pulse of [3H]-tyrosine and measuring the incorporation of radioactivity into protein extracts from myotubes. There were no differences in protein synthesis rates between AMPD3 or GFP myotubes at 24 and 48 hours (Figure 6B).

4. Discussion

Protein degradation, especially through the UPS, is an energy-dependent process, but the effects of decreased ATP levels (without an increase in ADP and AMP) on protein degradation have not been examined in live cells. In this study, we tested the hypothesis that a reduction in ATP would reduce the rate of protein degradation in muscle cells. In C2C12 myotubes, we found that overexpression of AMPD3 reduced adenine nucleotides (ATP, ADP, and AMP) in a dose- and time-dependent manner but did not increase the ADP/ATP ratio, AMP/ATP ratios or phosphorylation of AMPK Thr172, which are all markers for increased energetic stress [24, 33]. Furthermore, AMPD3 slowed the overall rate of protein degradation but had no effect on the rate of overall protein synthesis. Importantly, the activity of UPS, as measured by accumulation of Ub-GFP constructs, was decreased by AMPD3 overexpression, but the autophagic lysosomal pathway, the other major proteolytic pathway, was unaffected. Therefore, proteasomal-mediated protein degradation, but not autophagic-lysosomal degradation or protein synthesis, are sensitive to accelerated degradation of adenine nucleotides and decreases in ATP concentration in the physiological range in intact cells.

A major finding of the present study is that AMPD3 may be an important long-term regulator of adenine nucleotide content in muscle. The skeletal muscle adenine nucleotide pool is generally tightly regulated. During exercise, stimulated muscle contractions, or hypoxia, when ATP consumption outpaces ATP synthesis, ADP and AMP increase substantially. To preserve the free energy of ATP hydrolysis, AMP is deaminated to IMP [34]. For instance, skeletal muscle ATP has been shown to decrease up to 50% with roughly a roughly stoichiometric increase in IMP during intense electrically-stimulated contractions [35] or during ischemia [36]. Importantly, IMP remains largely intracellular and is resynthesized to ATP after cessation of contractions [35]. Therefore, the sum of ATP plus IMP remains constant. However, the current study demonstrates that AMP deamination can lead to a persistent reduction in ATP and other adenine nucleotides without an equivalent increase in IMP. In support of these findings, overexpression of 5’ nucleotidase (AMP → adenosine + Pi) also leads to reduced nucleotide triphosphate levels [37, 38]. On the contrary, overexpression of AMPD1 does not result in a decrease in ATP [23], likely due to differences in regulatory properties [39] or specific activity [40] of the different AMPD isoforms. Regardless, AMP degradation may be used as a general means to decrease intracellular ATP content and could be used to test the ATP dependence of other processes.

While overexpression of AMPD3 reduced ATP concentrations, it did not alter the AMP/ATP ratio or phosphorylation of AMPK Thr172, which is sensitive to the adenylate energy charge and is indicative of AMPK activation [33, 41]. AMPK is a known activator of lysosomal protein degradation through its phosphorylation of ULK1 Ser555 [42]. While we did not observe any change in P-AMPK, we did detect a significant reduction in the phosphorylation of ULK1 Ser555. However, this did not appear to influence the rate of autophagy flux as measured by the accumulation of LC3I and LC3II in concanamycin A treated cells. Therefore, enhanced degradation of the adenine nucleotides by AMPD3 can disassociate a decrease in ATP from markers of energetic stress.

The direct role of ATP concentration on proteasome activity in cell and tissue homogenates has been studied previously [16, 17]. While these outstanding in vitro studies allow precise manipulation of all reaction components, the cell studies as done herein have several advantages. First, intact cell studies retain the intracellular organization and proteasome-to-total protein concentrations, both of which may affect intracellular diffusion and interaction of proteins with proteases. This is especially important for skeletal muscle, where the lattice of the myofibrillar proteins specifically restricts movements of ions and proteins [43]. Second, protein-protein binding partners and post-translational modifications are preserved in intact cell studies. For example, preparation of tissue/cell homogenates may result in loss of proteasome post-translational modifications (e.g. phosphorylation) or binding partners, both of which can profoundly impact proteasome activity [12]. Third, because the studies were performed in live cells with no observable cytotoxic impact, the concentrations of adenine nucleotide and protein concentrations are necessarily in the physiological range. Fourth, rates of total protein degradation (i.e. both UPS and autophagic-lysosome) can be measured in intact cells, which is not possible in cell homogenates because the acidic environment of intact lysosomes is lost by homogenization.

Our findings demonstrate that a decrease in ATP is associated with a slowed rate of protein degradation. However, our cell studies were not designed to investigate which step of the UPS may be critically affected by lower ATP levels. Regardless, our data provide several clues as to possible mechanisms. First, our data suggest that the UPS activity is preferentially decreased; no changes in autophagy/lysosomal activation were observed. Second, the decrease in UPS activity was not due to a decrease in proteasome content or inherent activity (as measured in vitro at fixed ATP levels), suggesting that ubiquitination or access of proteins to the proteasome may be limited. Third, since the decrement in proteolysis was observed without a change in the apparent free energy of ATP hydrolysis, the impacted component is likely not highly energy dependent but, instead, dependent on ATP amount. For example, the role of ATP as a hydrotrope [44], to enhance protein solubility and perhaps delivery to the proteasome, is not dependent on free energy of ATP hydrolysis. Likewise, ATP binding and not strictly ATP hydrolysis may be rate limiting under some conditions in unfolding proteins at the proteasome [45, 46]. However, because we are using intact cells, we cannot discount the possibility that that factors other than ATP may be limiting protein degradation. Future studies will be required to identify which specific steps are slowed by increased degradation of AMP.

Interestingly, atrophic skeletal muscle, observed during cancer cachexia, diabetes, and chronic kidney disease, has an increased expression of AMPD3 and decreased levels of adenine nucleotides, especially ATP [47]. Since, protein degradation rates are typically accelerated during these atrophic conditions [48], our findings suggest that AMPD3 may function to temper protein degradation in these chronic diseases. Since even high levels of AMPD3 protein, as reported herein using overexpression viruses, do not result in a decrease of protein synthesis rates, further enhancement of AMPD activity may be an attractive target for treatment of diseases characterized by muscle protein degradation and subsequent muscle atrophy.

5. Conclusions

The overexpression of AMPD3 in skeletal muscle cells resulted in a reduction in ATP content and UPS-mediated protein degradation while maintaining ADP/ATP and AMP/ATP ratios. The altered energetic state of the cell did not influence the activation of AMPK, lysosomal protein degradation, or protein synthesis. These findings indicate that adenine nucleotide degradation and cellular energetics may play key roles in regulating the activity of the UPS in muscle cells under physiological conditions.

Highlights.

  • Increased AMP degradation leads to decreased ATP content in muscle cells

  • Decreased ATP content can be dissociated from decreased free energy of ATP hydrolysis

  • Decreased ATP content does not alter protein synthesis or autophagic/lysosomal flux

  • Decreased ATP content is associated with decreased protein degradation in live cells

Funding

This work was supported by the National Institute of Arthritis and Musculoskeletal and Skin Diseases (R01AR070200), National Institute of Diabetes and Digestive and Kidney Diseases (R01DK103562), National Heart, Lung, and Blood Institute (R15HL135699) of the National Institutes of Health, and the American College of Sports Medicine Foundation.

Nonstandard Abbreviations

AMPD3

AMP deaminase isoform 3

AMPK

AMP-activated protein kinase

DMEM

Dulbecco’s modified eagle medium

DTT

Dithiothreitol

FBS

Fetal bovine serum

GFP

Green fluorescent protein

HEK

Human embryonic kidney

IRES

Internal ribosomal entry site

PCA

Perchloric acid

RIPA

Radioimmunoprecipitation assay

TCA

Trichloroacetic acid

UPLC

Ultra-performance liquid chromatography

UPS

Ubiquitin-proteasome system

Footnotes

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Declarations of Interest: None

References

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