Chronology of UPR activation in skeletal muscle adaptations to chronic contractile activity

Abstract

The mitochondrial and endoplasmic reticulum unfolded protein responses (UPR^mt and UPR^ER) are important for cellular homeostasis during stimulus-induced increases in protein synthesis. Exercise triggers the synthesis of mitochondrial proteins, regulated in part by peroxisome proliferator activator receptor-γ coactivator 1α (PGC-1α). To investigate the role of the UPR in exercise-induced adaptations, we subjected rats to 3 h of chronic contractile activity (CCA) for 1, 2, 3, 5, or 7 days followed by 3 h of recovery. Mitochondrial biogenesis signaling, through PGC-1α mRNA, increased 14-fold after 1 day of CCA. This resulted in 10–32% increases in cytochrome c oxidase activity, indicative of mitochondrial content, between days 3 and 7, as well as increases in the autophagic degradation of p62 and microtubule-associated proteins 1A/1B light chain 3A (LC3)-II protein. Before these adaptations, the UPR^ER transcripts activating transcription factor-4, spliced X-box-binding protein 1, and binding immunoglobulin protein were elevated (1.3- to 3.8-fold) at days 1–3, while CCAAT/enhancer-binding protein homologous protein (CHOP) and chaperones binding immunoglobulin protein and heat shock protein (HSP) 70 were elevated at mRNA and protein levels (1.5- to 3.9-fold) at days 1–7 of CCA. The mitochondrial chaperones 10-kDa chaperonin, HSP60, and 75-kDa mitochondrial HSP, the protease ATP-dependent Clp protease proteolytic subunit, and the regulatory protein sirtuin-3 of the UPR^mt were concurrently induced 10–80% between days 1 and 7. To test the role of the UPR in CCA-induced remodeling, we treated animals with the endoplasmic reticulum stress suppressor tauroursodeoxycholic acid and subjected them to 2 or 7 days of CCA. Tauroursodeoxycholic acid attenuated CHOP and HSP70 protein induction; however, this failed to impact mitochondrial remodeling. Our data indicate that signaling to the UPR is rapidly activated following acute contractile activity, that this is attenuated with repeated bouts, and that the UPR is involved in chronic adaptations to CCA; however, this appears to be independent of CHOP signaling.

skeletal muscle is a highly adaptable tissue, responsive to multiple stressors leading to changes in whole body metabolism. It has been established that exercise induces many physiological adaptations that are beneficial for muscle performance capabilities, such as remodeling of the mitochondrial reticulum through mitochondrial biogenesis, as well as removal of damaged or dysfunctional organelles through a process termed autophagy (28, 39, 51). Mitochondria are the essential cellular organelles, with primary roles in energy production in the form of ATP (27). At the onset of contractile activity, early signaling events involved in mitochondrial biogenesis are initiated within skeletal muscle and converge largely on the activation of the transcriptional coactivator peroxisome proliferator activator receptor-γ coactivator 1α (PGC-1α) (20, 27, 52, 54). The vast majority of component proteins are transcribed in the nucleus and require transport to the mitochondria via chaperone and import machineries to facilitate protein shuttling and organelle assembly (11, 25, 30, 65). Less than 1% of total mitochondrial proteins are derived from mitochondrial DNA; the remainder are provided by the nucleus (3, 24, 53). Therefore, to produce a functional organelle capable of adequate ATP supply, contractile activity-induced mitochondrial biogenesis requires the integration, coordination, and timely expression of the nuclear and mitochondrial genomes.

Under conditions of cell stress, such as unaccustomed exercise, proteins may become misfolded and unable to reach their appropriate destination within the mitochondrion, leading to a proteotoxic cellular environment (21, 46). However, the unfolded protein response (UPR) is gaining attention as a protective mechanism that helps maintain homeostasis under cell stress. Two such UPR pathways have been observed. The first, an endoplasmic reticulum (ER)-mediated UPR (UPR^ER) senses misfolded proteins within the ER lumen through the resident chaperone binding immunoglobulin protein [BiP (also known as GRP78)] (50). BiP preferentially binds unfolded proteins, thus disassociating and, thereby, activating the ER membrane-bound kinases, inositol-requiring enzyme-1α (IRE1α) and protein kinase R-like ER kinase (PERK), as well as releasing a potent transcription factor, activating transcription factor (ATF) 6 (ATF6) to increase a host of chaperones and proteases while simultaneously reducing global translation (55). The second is a mitochondrial UPR (UPR^mt), which senses proteotoxic stress in the matrix and intermembrane space to independently activate a variety of mitochondria-specific chaperones and proteases and regain homeostasis (6, 46, 60). Together, the UPR^ER and UPR^mt initiate an acute response to reduce protein load and an adaptive response to increase protein-handling ability. Ultimately, if homeostasis is not reached, autophagy will be induced (29, 46).

The molecular mechanisms involved in the metabolic adaptations observed with exercise have yet to be fully elucidated. It has recently been found that exercise is capable of eliciting a UPR in untrained muscle, and in an intensity-dependent manner (32, 42, 57, 63). However, despite these recent findings, no work has focused on the role of the UPR^ER or UPR^mt with exercise in mitochondrial biogenesis and the induction of autophagy in skeletal muscle. Therefore, our objectives were to examine 1) the chronology of UPR^ER and UPR^mt activation with contractile activity in relation to mitochondrial biogenesis and autophagy and 2) whether the UPR is required for these exercise-associated adaptations by administration of the chemical chaperone mimetic tauroursodeoxycholic acid (TUDCA), a naturally occurring bile acid capable of blocking ER stress-induced UPR activation (44, 64). Based on these objectives, we hypothesize that the UPR will be rapidly induced in the acute phase of the contractile period and that UPR signaling is required for sufficient adaptation in response to the chronic contractile activity (CCA) stimulus.

METHODS

Animals and CCA.

The CCA model has been described previously as a useful model of chronic exercise (33, 58). Briefly, 72 male Sprague-Dawley rats (Charles River, St. Constant, QC, Canada) were given food and water ad libitum and kept on a 12:12-h light-dark schedule. At 12–18 wk of age, animals (350–550 g body wt) were anesthetized with isoflurane. Wire electrodes (Medwire, Leico Industries, New York, NY) were passed unilaterally and subcutaneously from the left hindlimb to the top of the back to connect to an external stimulation unit secured with surgical tape. Animals were allowed 5–7 days of recovery following surgery before tibialis anterior (TA) and extensor digitorum longus (EDL) muscles were subjected to stimulation (6 V, 10 Hz continuous, 0.1-ms pulse duration, 999 ms of rest per second). For all animals, TA and EDL contractions were induced by electrical stimulation of the common peroneal nerve for 3 h/day followed by 21 h of recovery. Animals in the time-course study were subjected to 1, 2, 3, 5, or 7 days of contractile activity, whereas vehicle/TUDCA-treated animals were subjected to 2 or 7 days of stimulation. At 3 h after the final bout of contractile activity, animals were euthanized and the TA and EDL of control and stimulated hindlimbs were extracted and flash-frozen in liquid nitrogen and then pulverized into fine powder for subsequent experimental analysis. Immediately following tissue removal, animals were euthanized via cardiac excision. Animal protocols were approved by the York University Animal Care Committee.

TUDCA and vehicle treatment.

Animals were randomly divided into TUDCA- or vehicle-treated groups. TUDCA (EMD Millipore, Billerica, MA) was dissolved in phosphate-buffered saline (Wisent Bio Products, Saint-Jean-Baptiste, QC, Canada) at 200 mg/ml and pH 7.5 as in previous studies using this dose (19, 48, 49). TUDCA (400 mg/kg) or an equal volume of sterile phosphate-buffered saline (vehicle) was administered daily via intraperitoneal injection beginning 3 days prior to surgery. Injections continued throughout the recovery and stimulation period and ended 1 day prior to tissue extraction to avoid the acute effects of drug treatment.

Cytochrome c oxidase activity.

Cytochrome c oxidase (COX) activity was used as a marker of mitochondrial content. Pulverized whole muscle homogenates were prepared and sonicated for 10 s on ice at a power output of 20–30%. A buffered test solution containing fully reduced horse heart cytochrome c (catalog no. C-2506, Sigma, Oakville, ON, Canada) was prepared. A multipipette was used to add 240 μl of test solution to 50 μl of whole muscle homogenate in a 96-well plate. COX enzyme activity was determined spectrophotometrically as the maximal rate of oxidation of fully reduced cytochrome c (catalog no. C-2506, Sigma), measured by the change in absorbance at 550 nm and 30°C in a microplate reader (Synergy HT, Bio-Tek Instruments, Winooski, VT). For each sample, COX activity was calculated as an average of three trials.

In vitro RNA isolation and reverse transcription.

Total RNA was isolated from frozen, whole muscle TA powders as described previously (43). Briefly, tissue powder (∼70 mg) was added to TRIzol reagent, homogenized, and mixed with chloroform. Samples were centrifuged at 4°C at 16,000 g for 15 min, and the upper aqueous phase of the sample was transferred to a new tube along with isopropanol and left overnight at −20°C to precipitate. Samples were once again centrifuged at 4°C at 16,000 g for 10 min. The resultant supernate was discarded, and the pellet was resuspended in 30 μl of molecular-grade sterile H₂O (Wisent Bio Products, Saint-Jean-Baptiste, QC, Canada). The concentration and purity of the RNA were measured using a spectrophotometer (NanoDrop 2000). SuperScript III reverse transcriptase (Invitrogen, Carlsbad, CA) was used to reverse-transcribe 1.5 μg of total RNA into cDNA.

Real-time PCR.

Sequences from GenBank were used to design primers with Primer 3 version 0.4.0 software (Massachusetts Institute of Technology, Cambridge, MA) for genes of interest (Table 1). Primer specificity was confirmed by OligoAnalyzer 3.1 (Integrated DNA Technologies, Toronto, ON, Canada). mRNA expression was measured with SYBR Green chemistry (PerfeCTa SYBR Green SuperMix, ROX, Quanta BioSciences, Gaithersburg, MD). Each well contained SYBR Green SuperMix, forward and reverse primers (20 μM), sterile H₂O, and 10 ng of cDNA. All real-time PCR amplification was detected in a 96-well plate using a StepOnePlus Real-Time PCR System (Applied Biosystems, Foster City, CA). The final reaction volume of each well was 25 μl. Samples were run in duplicates to ensure accuracy. The PCR program consisted of an initial holding stage (95°C for 10 min) followed by 40 amplification cycles (60°C for 1 min, 95°C for 15 s) and was completed with a final melting stage (95°C for 15 s, 60°C for 1 min, 95°C for 15 s). Analysis of melt curves generated by the instrument for SYBR Green analyses was used to control for nonspecific amplification and primer dimers. Negative control wells contained H₂O in place of cDNA.

Table 1.

List of primer oligonucleotide sequences used in real-time quantitative PCR analysis for Rattus norvegicus

		Primer
Gene, bp	Accession No.	Forward	Reverse
Ppargc1a (PGC-1α), 142	NM_031347.1	5′-CAT CGC AAT TCT CCC TTG TAT-3′	5′-CAG ACT CCC GCT TCT CAT ACT-3′
ATF4, 107	NM_024403.2	5′-CTC TCG CCA AAG AGA TTC AGT A-3′	5′-ACA AGC ACA AAG CAC CTG ACT A-3′
Hspa5 (BiP), 137	NM_013083.2	5′-TTG AAA CTG TGG GAG GTG T-3′	5′-GGG TCG TTC ACC TTC GTA GA-3′
Ddit3 (CHOP), 130	NM_001109986.1	5′-GAG CTG GAA GCC TGG TAT GA-3′	5′-GGG ATG CAG GGT CAA GAG TA-3′
ClpP, 145	XM_217313.8	5′-GAG CGA TAC GTG GGA GAC A-3′	5′-ACG TTG CTT CCT TAC TCA GCA-3′
Hspe1 (CPN10), 124	NM_012966.1	5′-CAC GGA GGC ACC AAA GTA GT-3′	5′-GGA ATG GGC AGC TTC ATG T-3′
Esr1 (Erα), 126	NM_012689.1	5′-CAT GAT GAA AGG CGG GAT A-3′	5′-AGG TTG GCA GCT CTC ATG T-3′
GAPDH, 122	NM_017008.3	5′-CTC TCT GCT CCT CCC TGT TCT-3′	5′-GGT AAC CAG GCG TCC GAT AC-3′
Hspd1 (HSP60), 138	NM_022229.2	5′-AGG CAG GTT CCT CAC CAA TAA-3′	5′-GCA TGG ACA ATG ACA GCA GTA-3′
Hspa4 (HSP70), 101	NM_153629.1	5′-CAT GGT GCT GAC CAA GAT GA-3′	5′-GCT GCG AGT CGT TGA AGT A-3′
Map1lc3 (LC3), 101	NM_199500.2	5′-GCA CAG CAT GGT GAG TGT AT-3′	5′-AGG TTT CTT GGG AGG CAT AGA-3′
Lonp1 (LonP), 117	NM_133404.1	5′-CTT GTG GTT CCC AAG CAT GT-3′	5′-CGT CAG CCA GTC CAG GTA GT-3′
Hspa9 (mtHSP70), 116	NM_001100658.2	5′-CCT TCT GTG GTT GCC TTT ACA-3′	5′-CGT CCA ATA AGA CGC TTT GTA-3′
Bnip3l (NIX), 133	NM_080888.1	5′-CCC TGC ACA ACA ACA ACA AC-3′	5′-CCA TTC TTC CCA TTT CCA TTA C-3′
Nqo1 (NQO1), 148	NM_017000.3	5′-TTC TGT GGC TTC CAG GTC TTA-3′	5′-GCT GCT TGG AGC AAA GTA GA-3′
Sqstm1 (p62), 117	NM_175843.4	5′-GGA ACT GAT GGA GTC GGA TAA C-3′	5′-TCC GAT TCT GGC ATC TGT AG-3′
Rps12 (S12), 127	NM_031709.3	5′-ATG GAC GTC AAC ACT GCT CT-3′	5′-ATG CAA GCA CGC AGA GAT-3′
SirT3, 118	NM_001106313.2	5′-GCC CAA TGT CGC TCA CTA CT-3′	5′-CAG CTT TGA GGC AGG GAT A-3′
Uqcrc1, 135	NM_001004250.2	5′-TCG AGA GGT TTG CTC CAA GTA-3′	5′-CGC AGA CTT CCT GCC TAG A-3′
XBP1s, 86	NM_001271731.1	5′-TGC TGA GTC CGC AGC AGG T-3′	5′-AAT CTG AAG AGG CAA CAG CGT-3′
XBP1t, 114	NM_001271731.1	5′-CCT TCT CCC TTC AGC GAC AT-3′	5′-CAG TGG TGG GTG GCT TTA GA-3′

Alternative names used in this paper are shown in parentheses. PGC-1α, peroxisome proliferator activator receptor-γ coactivator 1α; ATF, activating transcription factor; BiP, binding immunoglobulin protein; CHOP, CCAAT-enhancer binding protein (C/EBP) homologous protein; ClpP, ATP-dependent Clp protease proteolytic subunit; CPN10, 10-kDa chaperonin; Erα, estrogen receptor-α; HSP, heat shock protein; LC3, microtubule-associated proteins 1A/1B light chain 3A; LonP, lon peptidase 1; mtHSP70, 75-kDa mitochondrial HSP; Nqo1, NAD(P)H dehydrogenase, quinone 1; SirT3, sirtuin-3; Uqcrc1, cytochrome bc₁ complex III subunit 2; XBP1s and XBP1t, spliced and unspliced X-box binding protein 1.

Real-time PCR quantification.

First, the threshold cycle (C_T) value of the endogenous reference gene was subtracted from the C_T value of the target gene: ΔC_T = C_T(target) − C_T(reference). Next, the ΔC_T value of the control tissue was subtracted from the ΔC_T value of the experimental tissue: ΔΔC_T = ΔC_T(experimental) − ΔC_T(control). Results are reported as fold changes using the ΔΔC_T method, calculated as 2^−ΔΔC_T. Primers detecting ribosomal protein S12 along with glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were chosen as endogenous reference genes.

Immunoblotting.

Whole muscle protein extracts were separated by 10–15% SDS-PAGE and transferred to nitrocellulose membranes with a wet electrotransfer apparatus (Mini Trans-Blot electrophoretic transfer cell, Bio-Rad, Mississauga, ON, Canada). Nitrocellulose membranes were blocked in a 5% skim milk solution and subsequently incubated overnight at 4°C with the appropriate concentration of primary antibody (Table 2). Membranes were then washed and incubated with the suitable horseradish peroxidase-conjugated secondary antibody for 1 h at room temperature and visualized with enhanced chemiluminescence. Immunoblots were normalized using GAPDH or the cytoskeletal protein aciculin, as they do not alter expression levels with our exercise stimulus and, thus, provide a reliable loading control. Quantification was performed with ImageJ software (NIH, Bethesda, MD), and values were normalized to the appropriate loading control.

Table 2.

List of antibodies

Antibody	Manufacturer	Catalog No.	Lot No.
Aciculin	In house	*
ATF4	Santa Cruz Biotechnology	SC-200	K1314
ATF6	Abcam	ab203119	GR235316-1
BiP	Cell Signaling	3183S	3
CHOP	Cell Signaling	2895	10
COX IV	Abcam	ab140643	GR192963-3
CPN10	Enzo Life Sciences	ADI-SPA-110	09011010
Phosphorylated (Ser51) eIF2α	Invitrogen	44728G	1513929A
GAPDH	Abcam	ab8245	GR137268-5
HSP60	Enzo Life Sciences	ADI-SPA-806	03041452
HSP70	Enzo Life Sciences	ADI-SPA-810	12071118
LC3-II (A and B)	Cell Signaling	4108S	2
mtHSP70	Enzo Life Sciences	ADI-SPA-825	04011310
p62	Sigma	P0067	015M4877V
PGC-1α	Millipore	AB3242	2691399
SirT3	Cell Signaling	5490S	2
Uqcrc2	Abcam	ab14745	GR160043-12

BiP, binding immunoglobulin protein; COX IV, cytochrome c oxidase subunit IV; eIF2α; eukaryotic initiation factor 2.

^*See Ref. 7.

Statistical analysis.

To compare control with stimulated muscles at a given time point and within vehicle/TUDCA conditions for mRNA, protein, and COX activity experiments, a repeated-measures two-way analysis of variance was performed followed by Bonferroni's post hoc test when necessary. mRNA analysis was performed using the ΔC_T value. All statistical analyses were performed using GraphPad Prism 6 software. Values are means ± SE. Significance was set at P < 0.05.

RESULTS

CCA induces changes in mitochondrial content.

We sought to establish a time-course relationship between mitochondrial adaptations in response to the CCA protocol over 1–7 days. To observe the activation of mitochondrial biogenesis signaling, we measured the mRNA expression of the transcriptional coactivator PGC-1α, regarded as the master regulator of this process. A single 3-h bout of contractile activity was sufficient to elicit a 14-fold increase in the transcript level, which remained markedly elevated throughout 7 days of CCA (P < 0.05; Fig. 1A). To effectively quantify the changes in mitochondrial content, we also measured whole muscle COX enzyme activity. COX activity displayed an 11% increase as early as day 3 (P = 0.06), which progressed to 32% after 7 days of contractile activity (P < 0.05; Fig. 1B). We corroborated these data with protein expression of PGC-1α, as well as the integral nuclear-encoded electron transport chain component proteins COX subunit IV (COX IV) and the cytochrome bc₁ complex III subunit 2 (Uqcrc2), after 7 days of chronic stimulation (Fig. 1C). CCA significantly induced Uqcrc2 by 1.9-fold, with trends toward 1.3- to 1.4-fold increases in PGC-1α and COX IV, consistent with previous reports from our laboratory (17, 26).

Fig. 1.

Chronic contractile activity (CCA) results in enhanced mitochondrial biogenesis, leading to increased organelle content following 5 days of activity. A and B: fold change in mRNA expression of the mitochondrial biogenesis transcriptional coactivator peroxisome proliferator activator receptor-γ coactivator 1α (PGC-1α, n = 6 per day) and indicator of mitochondrial content cytochrome c oxidase (COX) enzyme activity throughout 1–7 days of CCA (n = 6 per day). STM, stimulated; CTR, control. C: spliced Western blot and associated graph of PGC-1α (n = 5 per day) and nuclear-encoded electron transport chain components cytochrome bc₁ complex subunit 2 (Uqcrc2, n = 6 per day) and COX IV (n = 6 per day) protein content, illustrating mitochondrial adaptation following 7 days of CCA. Numbers adjacent to blots represent molecular weight markers (in kDa). C, control; S, stimulated. Values are means ± SE. *P < 0.05, stimulated vs. control tibialis anterior (TA) muscle at a given time point. †P < 0.05, main effect of CCA relative to control across all conditions.

CCA elicits an autophagy response.

To focus on the autophagy response as it pertains to mitochondrial remodeling and removal of dysfunctional organelles (9), we selected the adapter protein p62 and the autophagosomal membrane protein microtubule-associated proteins 1A/1B light chain 3A (LC3), which together play important roles in facilitating subsequent organelle degradation. The mRNA expression of p62 increased 2.3-fold at day 2 of CCA (P < 0.05; Fig. 2A), while LC3 mRNA exhibited no changes over the course of 7 days (Fig. 2B). Corresponding Western blot analyses detected a twofold elevation in p62 protein levels following 3 days of CCA that returned to control levels by day 5 (P < 0.05; Fig. 2C). We also measured protein expression of LC3, as a ratio of its active form, LC3-II, to its inactive form, LC3-I. We found that the LC3-II-to-LC3-I ratio was significantly reduced by ∼55–65% between days 3 and 7 following repeated bouts of contractile activity (Fig. 2D). These changes in p62 and the LC3-II-to-LC3-I ratio between days 3 and 7 are suggestive of a CCA-induced increase in autophagy flux.

Fig. 2.

Autophagy is induced in a time-dependent manner with CCA. A and B: fold change in transcript levels of autophagy markers p62 (n = 6 per day) and microtubule-associated proteins 1A/1B light chain 3A (LC3, n = 7 per day) over 1–7 days of CCA. C and D: corresponding protein expression of p62 (n = 7 per time point) and ratio of lipidated LC3-II to inactive LC3-I (n = 6 per time point) throughout the exercise protocol. Values are means ± SE. *P < 0.05, stimulated vs. control TA muscle at a given time point.

CCA induces the UPR^ER as acute and adaptive responses.

Given that we found an increase in mitochondrial content and autophagy signaling, we also measured selected UPR^ER transcripts and proteins as they relate to these adaptive mechanisms over the time course of contractile activity. The activation of the PERK and IRE1α branches was assessed by the mRNA induction of their respective targets. In general, we detected a significant main effect of CCA (P < 0.05) on the mRNA expression of all UPR^ER factors measured as early as day 1 and throughout our 7-day time course. Specifically, ATF4, which is activated downstream of PERK, exhibited a 1.5- to 1.8-fold increase at days 2 and 3 (P < 0.05), with levels returning to control values by day 5 (Fig. 3A). To estimate IRE1α activation, we measured spliced X-box-binding protein 1 (XBP1s), which was similarly enhanced during days 1–3 of contractile activity, reaching a 3.3-fold increase at day 2 and, by day 7, was reduced by 45% in the stimulated, relative to the control, muscle (P < 0.05; Fig. 3B). We also extended our analysis to include downstream targets of the UPR^ER. The transcription factor CCAAT-enhancer binding protein (C/EBP) homologous protein (CHOP) and the ER chaperone BiP were significantly enriched at the transcript level by 2.2- to 3.9-fold throughout days 1–7 of chronic stimulation (Fig. 3, C and E). Furthermore, CHOP protein levels displayed a main effect of CCA at all time points, leading to a 1.8-fold increase by day 7 (P < 0.05; Fig. 3D), while BiP protein remained unchanged until day 7, when it trended toward an increase of 47% (P = 0.2; Fig. 3F). Finally, the cytosolic heat shock protein (HSP) HSP70 chaperone was markedly augmented throughout days 1–7 (P < 0.05) at the mRNA level, with a similar effect of CCA throughout our time course, specifically reaching 2.5-fold increases at days 2 and 5 (Fig. 3, G and H).

Fig. 3.

The endoplasmic reticulum unfolded protein response (UPR^ER) is rapidly activated in response to CCA. A–C, E, and G: fold change in mRNA levels of the UPR^ER-associated transcription factors activating transcription factor (ATF) 4 (ATF4, n = 6 per day), active X-box-binding protein 1 (XBP1s, n = 6 per day), and CCAAT/enhancer-binding protein homologous protein (CHOP, n = 6 per day), as well as chaperones binding immunoglobulin protein (BiP, n = 6 per day) and heat shock protein (HSP) 70 (HSP70, n = 6 per day) during 1–7 days of CCA. D, F, and H: corresponding protein expression of CHOP (n = 7 per day), BiP (n = 7 per day), and HSP70 [n = 6 (days 1–5) and n = 5 (day 7)]. Values are means ± SE. *P < 0.05, stimulated vs. control TA muscle at a given time point. †P < 0.05, main effect of CCA relative to control across all conditions.

CCA elicits the UPR^mt concurrent with the UPR^ER.

To investigate whether the UPR^mt plays a coordinated role in mitochondrial adaptations to exercise, we expanded our mRNA and protein analyses to include mitochondria-specific factors. First, mRNA levels of the mitochondrial protease ATP-dependent Clp protease proteolytic subunit (ClpP), which both senses and responds to unfolded matrix proteins (1, 6, 60), was significantly upregulated by CCA throughout the time course, particularly reaching a 70% increase by day 2 and maintaining a 45% elevation at days 5 and 7 (P < 0.05; Fig. 4A). Similarly, the mitochondrial regulatory protein sirtuin-3 (45) displayed enhanced protein expression as a result of CCA after days 2–7, reaching elevations of 46% by day 7 (P < 0.05; Fig. 4B). Finally, consistent 25–80% elevations in mRNA levels of the mitochondrial chaperones 10-kDa chaperonin (CPN10), HSP60, and 75-kDa mitochondrial HSP (mtHSP70) were observed throughout the 7-day contractile activity protocol (P < 0.05; Fig. 4, C, E, and G). This was further reinforced by changes in HSP60 and CPN10 protein expression, which exhibited a main effect of CCA across all time points, culminating in a 75% enhancement of CPN10 by day 7 specifically (P < 0.05; Fig. 4, D, F, and H).

Fig. 4.

The mitochondrial unfolded protein response (UPR^mt) is enhanced in response to acute and chronic contractile activity. A, C, E, and G: fold change in mRNA levels of the mitochondrial protease ATP-dependent Clp protease proteolytic subunit (ClpP, n = 8 per day) and the mitochondrial chaperones 10-kDa chaperonin (CPN10, n = 6 per day), HSP60 (n = 8 per day), and 75-kDa mitochondrial HSP (mtHSP70, n = 8 per day) throughout 7 days of muscle contractile activity. B, D, F, and H: coterminous expression of the UPR^mt regulatory protein sirtuin-3 (SirT3, n = 8 per day) and the chaperone proteins CPN10 (n = 7 per day), HSP60 (n = 7 per day), and mtHSP70 (n = 7 per day). Values are means ± SE. *P < 0.05, stimulated vs. control TA muscle at a given time point. †P < 0.05, main effect of CCA relative to control across all conditions.

TUDCA treatment does not impact CCA-induced activation of the UPR^ER or UPR^mt.

TUDCA has previously been shown to be capable of preventing ER stress-associated UPR activation (15, 35, 44). Therefore, we wanted to test the impact of the drug on mitochondrial adaptations, since our data illustrate that UPR activation seems to precede organelle remodeling in response to chronic muscle activity. Despite a ∼50% attenuation (P < 0.05) of the CCA-induced activation of CHOP protein (Fig. 5, A and B), a hallmark of TUDCA activity (44, 59), along with a 31% reduction in HSP70 adaptation to CCA (P < 0.05; Fig. 5, D and E), we were unable to detect any differences in the expression of other UPR^ER targets in vehicle- and TUDCA-treated animals. The principal mechanism of action of TUDCA is in facilitation of protein handling and translation (7, 14); therefore, it is no surprise that the mRNA expression of CHOP (Fig. 5C) and HSP70 (Fig. 5F) was significantly enriched following 2 days of muscle stimulation in either treatment group. The mRNA expression of ATF4 (Fig. 5G), XBP1s (Fig. 5H), and BiP (Fig. 5I) was markedly elevated on day 2 in both treatment groups, with levels dropping to equal to or below control following 7 days of CCA (P < 0.05). Consistent with our earlier findings, CCA had a significant main effect on mRNA expression of the mitochondrial chaperones CPN10 (Fig. 6A) and HSP60 (Fig. 6C) and the protease ClpP (Fig. 6D) following 2 and 7 days of activity, with no significant difference detected between treatments. Specifically, CPN10 mRNA displayed ∼60–80% increases (P < 0.05) across all conditions, while HSP60 followed the same trend and displayed a significant 30% increase in TUDCA-treated animals subjected to 7 days of stimulation (P < 0.05). Moreover, CPN10 protein increased ∼20% after 2 days of CCA and as much as 70% after 7 days, while TUDCA treatment did not have an effect on the level of induction relative to vehicle treatment (P < 0.05; Fig. 6B).

Fig. 5.

Tauroursodeoxycholic acid (TUDCA) treatment attenuates CHOP and HSP70 induction but has no effect on general UPR signaling associated with CCA. A and B: CHOP protein expression represented as fold change and as relative control vs. stimulated levels for a given day/treatment (n = 7 per day/condition). AU, arbitrary units. D and E: HSP70 protein expressed as fold change and as relative control vs. stimulated levels for a given day/treatment (n = 7 per day/condition). C and F–I: relative mRNA levels of the UPR^ER markers CHOP, HSP70, ATF4, XBP1s, and BiP in control and stimulated muscle of vehicle (VEH)- and TUDCA (TUD)-treated animals subjected to 2 or 7 days of muscle stimulation (n = 8, per day/treatment). C_T, threshold cycle. Values are means ± SE. *P < 0.05, stimulated vs. control TA muscle at a given time point and treatment. #P < 0.05, 2 days stimulated vs. 7 days stimulated for the same drug treatment. ¶P < 0.05, TUDCA vs. vehicle for a given time point.

Fig. 6.

TUDCA treatment has no effect on UPR^mt signaling in response to CCA. A, C, and D: relative transcript levels of the mitochondrial chaperones CPN10 (n = 8 per day/treatment) and HSP60 (n = 6 per day/treatment), as well as the protease ClpP (n = 8 per day/treatment) in control and stimulated muscle of vehicle- and TUDCA-treated animals following 2 or 7 days of contractile activity. B: corresponding protein expression of CPN10 (n = 8 per day/treatment). Values are means ± SE. *P < 0.05, stimulated vs. control TA muscle at a given time point and treatment. †P < 0.05, main effect of CCA relative to control across all conditions.

TUDCA treatment has no effect on CCA-induced mitochondrial biogenesis or autophagy.

We assessed changes in mitochondrial content through COX enzyme activity in animals treated with vehicle or TUDCA and subjected to 2 or 7 days of CCA. Consistent with our earlier results, we observed 22% and 24% increases in mitochondria with 7 days of CCA in vehicle- and TUDCA-treated animals, respectively (P < 0.05; Fig. 7A). Similarly, PGC-1α mRNA expression was significantly increased 6.6- to 8.4-fold in both treatment groups stimulated for 2 or 7 days, with no difference in mRNA induction as a result of drug administration (Fig. 7B). To reinforce these data, we measured the protein expression of Uqcrc2 and found similar 1.7- to 2-fold increases in the stimulated leg relative to control across all groups, as CCA exhibited a main effect in all conditions (P < 0.05; Fig. 7C). Akin to our earlier findings, p62 mRNA exhibited a significant ∼1.7-fold increase after 2 days of stimulation in TUDCA-treated animals (Fig. 8A). These trends in p62 were also evident at the protein level at day 2 in vehicle- and TUDCA-treated animals (P < 0.05; Fig. 8C) and were completely attenuated after 7 days of CCA. No change in LC3 mRNA and protein levels was observed in any condition/time point (Fig. 8, B and D).

Fig. 7.

TUDCA treatment did not impair mitochondrial biogenesis or attenuate the increase in mitochondrial content following CCA. A: relative changes in mitochondrial content observed via COX activity (n = 8 per day/treatment) following 2 and 7 days of CCA in vehicle- and TUDCA-treated rats. B and C: coactivator of mitochondrial biogenesis PGC-1α mRNA levels (n = 8 per day/condition) and protein expression of nuclear-encoded Uqcrc2 (n = 7 per day/condition). Values are means ± SE. *P < 0.05, stimulated vs. control TA muscle at a given time point and treatment. †P < 0.05, main effect of CCA relative to control across all conditions.

Fig. 8.

Autophagy induction was not affected by TUDCA administration with CCA. A and B: relative mRNA expression of p62 (n = 6 per day/treatment) and LC3 (n = 7 per day/treatment) in vehicle- and TUDCA-treated animals following 2 or 7 days of CCA. C and D: corresponding protein levels of p62 (n = 7) and ratio of lipidated LC3-II to inactive LC3-I (n = 8). Values are means ± SE. *P < 0.05, stimulated vs. control TA muscle at a given time point and treatment. #P < 0.05, 2 days stimulated vs. 7 days stimulated for the same drug treatment.

DISCUSSION

Skeletal muscle accounts for a majority of the body mass and, thus, is an important determinant of metabolic status and overall health (13, 27). Exercise has been studied for its ability to produce adaptations in skeletal muscle to achieve greater oxidative capacity as a by-product of enhanced mitochondrial volume. However, the mechanism by which muscle achieves this new phenotype has yet to be fully resolved. Recently, the UPR has been implicated in exercise, which suggests that it plays a role in sensing and initiating adaptive responses (31, 32, 41, 63). In this study we employed a CCA model of exercise to analyze the signaling events that facilitate skeletal muscle adaptations to exercise over a 7-day time course of repeated bouts of contractile activity (33, 58).

CCA over the course of 7 days established an enhanced mitochondrial pool, corroborating previous work from our laboratory using this protocol (12, 17, 42, 58). Our observed 30% increase in COX activity by day 7 is similar to the level of adaptation achieved with 6 wk of regular endurance training (23), indicating that our 7-day protocol of chronic stimulation provides a useful time course of the signaling responses as they contribute to a new muscle phenotype. Of particular note, mitochondrial biogenesis-specific signaling, indicated by increased PGC-1α mRNA, was markedly induced following a single bout of CCA. However, increases in mitochondrial content were not detectable until day 3. Therefore, this 3-day window of signaling preceding adaptation is of particular interest, since PGC-1α plays a fundamental role in regulating transcriptional activity of a host of nuclear-encoded mitochondrial proteins integral to organelle biosynthesis (34). While it is known that PGC-1α regulates mitochondrial content, only recently has it been suggested that it might also have a more functional role in balancing other adaptive responses such as autophagy and the UPR through interaction with ATF6 (61–63).

When considering the beneficial adaptations to exercise, it is important to acknowledge the balance between mitochondrial biogenesis and autophagy in effectively maintaining a healthy organelle pool (22, 62). Autophagy is gaining recognition for its role in maintaining cell quality and viability beyond simply degradation (9). Evidence suggests that lack of adequate autophagy results in muscle degeneration, wasting, and reduced strength, as well as accumulation of protein aggregates and dysfunctional organelles, contributing to excessive oxidative stress (9, 18, 36, 37). Our data reinforce the notion of autophagy as a quality-control mechanism in muscle adaptation, as we detected concomitant changes in autophagy markers from day 3 to day 7 following CCA, matching the timing of the increases we observed in mitochondrial content. Specifically, the enhanced mRNA expression of p62 at day 2 followed by elevated protein expression at day 3 suggests an increase in transcriptional drive to enhance the level of this adapter protein in readiness for autophagy. Consistent with this, we detected a reduction of the ratio of LC3-II to LC3-I from day 3 to day 7, as well as a return of p62 protein expression to control values by day 5, indicating the subsequent degradation of these autophagosomal components as they are decomposed by the lysosome. These data suggest that repeated exercise bouts in the form of chronic contractile activity induce an increase in autophagy flux within muscle.

Given these observations, we proposed that an intermediate response could modulate, and coordinate, the balance between synthesis and degradation. In particular, we focused on the UPR^ER and the UPR^mt, as they have been implicated in mitochondrial adaptations (8, 38, 47, 56) and autophagy (5, 40, 45) in the presence of cell stress. We hypothesized that the drive for nuclear-encoded mitochondrial proteins through enhanced PGC-1α signaling may perturb the protein environment within the cell and, therefore, that the UPR would be integral to exercise-induced mitochondrial remodeling. Our observations indicate that the UPR responds to the CCA stimulus acutely and persists to provide a level of adaptation to chronic stimulation. The UPR^ER transcription factors ATF4 and XBP1s were significantly induced from day 1 to day 3 of CCA, preceding changes in mitochondria and autophagy, thus providing an early signaling response. Similarly, the transcription factor CHOP, which regulates cell fate by monitoring cell stress (4), displayed marked elevations in both mRNA and protein throughout days 1–7 of CCA, implying a role for the protein in shifting from acute signaling to chronic adaptations. The mRNA and protein expression changes of the ER and cytosolic chaperones BiP and HSP70 substantiate CHOP data, as they were also induced from day 1 to day 7 in response to CCA. Together, our results provide the first time-course relationship of UPR signaling over a period of progressive adaptation in skeletal muscle mitochondria through exercise.

Another unique finding of our study was the parallel contractile activity-induced responses in the UPR^ER and the UPR^mt in skeletal muscle. To date, far less emphasis has been placed on the UPR^mt, particularly in mammalian cells. Nonetheless, we were interested in this response, as it pertains directly to the protein-folding capacity within mitochondria (10, 38). Our laboratory previously found that mitochondria-specific chaperones are induced as an adaptive response to 7 days of CCA (42). Our current data confirm these results and suggest that, similar to the time course of the UPR^ER, mitochondrial factors are induced immediately following a single bout of CCA and remain elevated throughout repeated contractile episodes. This further establishes the phasic nature of the UPR in providing rapid and adaptive protein-handling relief within multiple subcellular compartments upon alterations in exercise-induced changes in proteostasis.

Our results suggest that UPR signaling is initiated early in the CCA protocol and precedes mitochondrial adaptations and cellular remodeling and, therefore, may play a direct role in initiating and/or facilitating these events. As such, we utilized the chaperone mimetic drug TUDCA, a naturally occurring bile acid that is produced at low levels in the liver and used in the treatment of various ER stress-associated diseases (2, 16, 44). The effectiveness of TUDCA treatment is predominantly through augmentation of protein handling and suppression of CHOP- and caspase-12-mediated apoptosis during chronic ER stress (15, 35, 44, 59, 64). We found that TUDCA treatment sufficiently attenuated the increase in CHOP protein expression after 2 days of CCA and resulted in a similar reduction of HSP70 levels. Nevertheless, mRNA levels of both CHOP and HSP70 were unaffected by drug treatment, which corroborates the proposed mechanism of TUDCA action on protein handling and translation (7, 14). TUDCA also had no effect on mRNA expression of the UPR^ER transcription factors ATF4 and activated XBP1s, as well as the ER chaperone BiP, all of which displayed similar increases in response to CCA in TUDCA- and vehicle-treated animals. Since TUDCA facilitates protein handling, we next investigated its potential effect on activation of the UPR^mt. However, no effects on the signaling of the mitochondrial protein quality-control machinery ClpP, HSP60, and CPN10 were observed, indicating that the UPR^mt responds separately from the UPR^ER.

While various UPR targets were still induced in response to CCA in the presence of TUDCA treatment, we wanted to investigate whether the attenuation of CHOP protein could affect the outcome of mitochondrial remodeling. CHOP has been implicated as a regulator of cell fate during chronic stress (4), as well as in exercise-induced apoptosis (63), indicating that it is capable of monitoring the intensity and duration of cell stress and signaling toward remodeling (autophagy) or degradation (apoptosis). Despite effective attenuation of the CCA-induced CHOP expression by TUDCA, we observed no changes in mitochondrial content measured by COX enzyme activity and protein expression of the nuclear-encoded mitochondrial complex III subunit Uqcrc2. Similarly, mitochondrial biogenesis signaling was unaffected, as PGC-1α mRNA was considerably induced across all conditions. We also found that autophagy was unaffected by TUDCA treatment, despite the reduced activation of CHOP, as p62 and LC3 responded similarly to our initial time-course findings.

A limiting factor in the application of TUDCA treatment is the lack of specificity in inhibiting the three main branches/factors of the UPR: PERK, IRE1, and ATF6. Instead, TUDCA helps modulate protein handling within the cell to reduce the burden on the ER to prevent enhanced UPR activation and subsequent apoptosis. Therefore, specifically knocking out upstream UPR factors would be worthy of pursuit in the context of exercise adaptations. However, given the usefulness of TUDCA in diminishing CHOP protein increases, we were able to analyze the role of CHOP in CCA-induced adaptations in mitochondrial content and autophagy as a process of cellular quality control. Our findings suggest that the potential UPR^ER regulation of mitochondrial biogenesis acts independently of CHOP in our CCA model of exercise. However, this does not exclude the possibility that another UPR^ER-associated factor could influence this process. Specifically, ATF6 has been implicated in PGC-1α signaling (63); however, we were unable to observe any notable changes in ATF6 as a result of CCA in our study (data not shown), possibly as a result of the transient nature of UPR activation. Future work to examine a time course of UPR signaling following acute exercise would be useful to best elucidate which factors are the primary responders. The same study linking ATF6 and PGC-1α found that CHOP ablation partially rescued the exercise intolerance of muscle-specific PGC-1α knockout mice (63). Our data suggest that CHOP attenuation had no impact, positive or negative, on the mitochondrial or autophagic adaptations to contractile activity. It is important to note that the CCA stimulus is low-frequency (10-Hz) stimulation localized to the TA and EDL muscles, rather than whole body maximal exercise. In studies of whole body exercise, it has recently been observed that the UPR is sensitive to exercise intensity, as the higher the intensity, or novelty, of the stimulus, the greater the response (32, 41). We confirmed this trend, as UPR^ER transcription factor mRNAs returned to control levels by day 5, concomitant with enhanced protein-handling ability and mitochondrial content. Therefore, future avenues of research could include investigation of the effect of TUDCA on adaptations during an enhanced CCA stimulus, either in duration or intensity, to elicit a more stressful condition in the muscle and, possibly, induce an increased reliance on CHOP activation. Conversely, inducing ER stress to activate the UPR prior to an exercise protocol could further demonstrate its functional role in adaptation. As it is clear that the UPR is involved in some capacity in exercise-induced remodeling, elucidating the specific mechanism of action will be of particular interest as it relates to whole body metabolic health.

GRANTS

This work was supported by funding from the Natural Sciences and Engineering Research Council of Canada (NSERC) to D. A. Hood. D. A. Hood holds a Canada Research Chair in Cell Physiology. J. M. Memme was a recipient of a NSERC-Canada Graduate Scholarship.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

J.M.M. and D.A.H. developed the concept and designed the research; J.M.M. and A.N.O. performed the experiments; J.M.M., A.N.O., and D.A.H. analyzed the data; J.M.M. and D.A.H. interpreted the results of the experiments; J.M.M. prepared the figures; J.M.M. drafted the manuscript; J.M.M. and D.A.H. edited and revised the manuscript; J.M.M. and D.A.H. approved the final version of the manuscript.