Skeletal Muscle Mitochondrial Protein Synthesis and Respiration Increase With Low-Load Blood Flow Restricted as Well as High-Load Resistance Training

Thomas Groennebaek, Nichlas R Jespersen, Jesper Emil Jakobsgaard, Peter Sieljacks, Jakob Wang, Emil Rindom, Robert V Musci, Hans Erik Bøtker, Karyn L Hamilton, Benjamin F Miller, Frank V de Paoli, Kristian Vissing, Thomas Groennebaek, Nichlas R Jespersen, Jesper Emil Jakobsgaard, Peter Sieljacks, Jakob Wang, Emil Rindom, Robert V Musci, Hans Erik Bøtker, Karyn L Hamilton, Benjamin F Miller, Frank V de Paoli, Kristian Vissing

Abstract

Purpose: It is well established that high-load resistance exercise (HLRE) can stimulate myofibrillar accretion. Additionally, recent studies suggest that HLRE can also stimulate mitochondrial biogenesis and respiratory function. However, in several clinical situations, the use of resistance exercise with high loading may not constitute a viable approach. Low-load blood flow restricted resistance exercise (BFRRE) has emerged as a time-effective low-load alternative to stimulate myofibrillar accretion. It is unknown if BFRRE can also stimulate mitochondrial biogenesis and respiratory function. If so, BFRRE could provide a feasible strategy to stimulate muscle metabolic health. Methods: To study this, 34 healthy previously untrained individuals (24 ± 3 years) participated in BFRRE, HLRE, or non-exercise control intervention (CON) 3 times per week for 6 weeks. Skeletal muscle biopsies were collected; (1) before and after the 6-week intervention period to assess mitochondrial biogenesis and respiratory function and; (2) during recovery from single-bout exercise to assess myocellular signaling events involved in transcriptional regulation of mitochondrial biogenesis. During the 6-week intervention period, deuterium oxide (D2O) was continuously administered to the participants to label newly synthesized skeletal muscle mitochondrial proteins. Mitochondrial respiratory function was assessed in permeabilized muscle fibers with high-resolution respirometry. Mitochondrial content was assessed with a citrate synthase activity assay. Myocellular signaling was assessed with immunoblotting. Results: Mitochondrial protein synthesis rate was higher with BFRRE (1.19%/day) and HLRE (1.15%/day) compared to CON (0.92%/day) (P < 0.05) but similar between exercise groups. Mitochondrial respiratory function increased to similar degree with both exercise regimens and did not change with CON. For instance, coupled respiration supported by convergent electron flow from complex I and II increased 38% with BFRRE and 24% with HLRE (P < 0.01). Training did not alter citrate synthase activity compared to CON. BFRRE and HLRE elicited similar myocellular signaling responses. Conclusion: These results support recent findings that resistance exercise can stimulate mitochondrial biogenesis and respiratory function to support healthy skeletal muscle and whole-body metabolism. Intriquingly, BFRRE produces similar mitochondrial adaptations at a markedly lower load, which entail great clinical perspective for populations in whom exercise with high loading is untenable.

Keywords: bioenergetics; deuterium oxide; high-resolution respirometry; ischemic resistance training; mitochondrial biogenesis.

Figures

FIGURE 1
FIGURE 1
Study overview. The total study length was 9 weeks and comprised a single-trial study and a long-term study. For the single-trial study, subjects arrived in the early morning after overnight fasting. After 30 min supine rest, a whey protein supplement was administered. After consumption, participants completed a BFRRE, HLRE, or CON single-bout session. Biopsies were harvested immediately after BFRRE/HLRE/CON (45 min after protein administration) and again 3 h after BFRRE/HLRE/CON (3 h and 45 min after protein administration). During the subsequent 6-week intervention period, BFRRE and HLRE groups conducted training 3 times per week. Throughout the 6-week intervention period, D2O was orally administered to the participants with blood collected every second week to measure tracer enrichment. Biopsies and tests of exercise capacity were performed before and 4 days after the 6-week intervention period. BFRRE, blood flow restricted resistance exercise; CON, non-exercise control; HLRE, high-load resistance exercise; D2O, deuterium oxide; Ex, exercise (i.e., HLRE or BFRRE); C, non-exercise control (i.e., rest).
FIGURE 2
FIGURE 2
Muscle functional capacity for CON, BFRRE, and HLRE groups. (A) strength-endurance capacity. (B) Dynamic muscle strength. ∗∗ denotes difference from pre-values within groups (P < 0.01), C denotes difference from CON at corresponding time point (P < 0.05), B denotes difference from BFRRE at corresponding time point (P < 0.05). Data are presented as means ± SD. Overall effects are designated in the upper left corner of each graph.
FIGURE 3
FIGURE 3
Mitochondrial protein fractional synthesis rate (mito FSR) for CON, BFRRE, and HLRE groups. C denotes difference from CON (P < 0.05). Data are presented as means ± SD. Overall effect is designated in the upper left corner of the graph.
FIGURE 4
FIGURE 4
Citrate synthase activity normalized to total protein for CON, BFRRE, and HLRE groups. Data are presented as means ± SD. Overall effects are designated in the upper left corner of the graph.
FIGURE 5
FIGURE 5
Mitochondrial respiratory function in permeabilized muscle fibers for CON, BFRRE, and HLRE groups. (A) representative real-time oxygraph readout showing oxygen concentration in the chamber (blue line) and the calculated negative time derivative (oxygen flux) normalized to mg wet weight of muscle tissue (red line). Titrations are denoted with arrows and respiratory states are denoted with boxes. (B) state 2 respiration with glutamate and malate (GM). (C) state 3 respiration supported electron flow from complex I (GM3). (D) state 3 respiration supported by electron flow from complex I and II (GMS3). (E) state 4 respiration with oligomycin (4o). (F) maximal uncoupled respiration with FCCP (E). (G) respiratory control ratio (RCR) with complex I linked substrates. ∗ denotes difference from pre-values within groups (P < 0.05), ∗∗ denotes difference from pre-values within groups (P < 0.01), C denotes difference from CON at corresponding time point (P < 0.05), CC denotes difference from CON at corresponding time point (P < 0.01). Data are presented as means ± SD. Overall effects are designated in the upper left corner of each graph.
FIGURE 6
FIGURE 6
Phosphorylation levels of signaling proteins involved in regulation of mitochondrial adaptations for CON, BFRRE, and HLRE groups. (A) representative immunoblots for all proteins at all conditions. (B) phosphorylated 5′ AMP-activated protein kinase (p-AMPK). (C) phosphorylated acetyl-CoA carboxylase (p-ACC). (D) phosphorylated p38 mitogen-activated protein kinase (p-p38 MAPK). (E) phosphorylated calcium/calmodulin-dependent protein kinase II (p-CaMKII). (F) phosphorylated cAMP response-element binding protein (p-CREB). (G) phosphorylated p53 (p-p53). ∗ denotes difference from pre-values within groups (P < 0.05), ∗∗ denotes difference from pre-values within groups (P < 0.01), C denotes difference from CON at corresponding time point (P < 0.05), CC denotes difference from CON at corresponding time point (P < 0.01). Data are presented as mean fold changes ± SD. Overall effects are designated in the upper left corner of each graph.

References

    1. Abu-Elheiga L., Matzuk M. M., Abo-Hashema K. A., Wakil S. J. (2001). Continuous fatty acid oxidation and reduced fat storage in mice lacking acetyl-CoA carboxylase 2. Science 291 2613–2616. 10.1126/science.1056843
    1. American College of Sports Medicine [Acsm]. (2009). American College of Sports Medicine position stand. Progression models in resistance training for healthy adults. Med. Sci. Sports Exerc. 41 687–708. 10.1249/MSS.0b013e3181915670
    1. Anderson E. J., Lustig M. E., Boyle K. E., Woodlief T. L., Kane D. A., Lin C. T., et al. (2009). Mitochondrial H2O2 emission and cellular redox state link excess fat intake to insulin resistance in both rodents and humans. J. Clin. Invest. 119 573–581. 10.1172/jci37048
    1. Bartlett J. D., Close G. L., Drust B., Morton J. P. (2014). The emerging role of p53 in exercise metabolism. Sports Med. 44 303–309. 10.1007/s40279-013-0127-9
    1. Benziane B., Burton T. J., Scanlan B., Galuska D., Canny B. J., Chibalin A. V., et al. (2008). Divergent cell signaling after short-term intensified endurance training in human skeletal muscle. Am. J. Physiol. Endocrinol. Metab. 295 E1427–E1438. 10.1152/ajpendo.90428.2008
    1. Bergstrom J. (1975). Percutaneous needle biopsy of skeletal muscle in physiological and clinical research. Scand. J. Clin. Lab. Invest. 35 609–616. 10.3109/00365517509095787
    1. Brownsey R. W., Boone A. N., Elliott J. E., Kulpa J. E., Lee W. M. (2006). Regulation of acetyl-CoA carboxylase. Biochem. Soc. Trans. 34(Pt 2), 223–227. 10.1042/bst20060223
    1. Brzycki M. (1993). Strength testing—predicting a one-rep max from reps-to-fatigue. J. Phys. Educ. Rec. Dance 64 88–90. 10.1080/07303084.1993.10606684
    1. Busch R., Kim Y. K., Neese R. A., Schade-Serin V., Collins M., Awada M., et al. (2006). Measurement of protein turnover rates by heavy water labeling of nonessential amino acids. Biochim. Biophys. Acta 1760 730–744. 10.1016/j.bbagen.2005.12.023
    1. Camera D. M., Edge J., Short M. J., Hawley J. A., Coffey V. G. (2010). Early time course of Akt phosphorylation after endurance and resistance exercise. Med. Sci. Sports Exerc. 42 1843–1852. 10.1249/MSS.0b013e3181d964e4
    1. Chin E. R. (2005). Role of Ca2+/calmodulin-dependent kinases in skeletal muscle plasticity. J. Appl. Physiol. 99 414–423. 10.1152/japplphysiol.00015.2005
    1. Clark B. C. (2009). In vivo alterations in skeletal muscle form and function after disuse atrophy. Med. Sci. Sports Exerc. 41 1869–1875. 10.1249/MSS.0b013e3181a645a6
    1. Coffey V. G., Zhong Z., Shield A., Canny B. J., Chibalin A. V., Zierath J. R., et al. (2006). Early signaling responses to divergent exercise stimuli in skeletal muscle from well-trained humans. FASEB J. 20 190–192. 10.1096/fj.05-4809fje
    1. Counts B. R., Dankel S. J., Barnett B. E., Kim D., Mouser J. G., Allen K. M., et al. (2016). Influence of relative blood flow restriction pressure on muscle activation and muscle adaptation. Muscle Nerve 53 438–445. 10.1002/mus.24756
    1. Donges C. E., Burd N. A., Duffield R., Smith G. C., West D. W., Short M. J., et al. (2012). Concurrent resistance and aerobic exercise stimulates both myofibrillar and mitochondrial protein synthesis in sedentary middle-aged men. J. Appl. Physiol. 112 1992–2001. 10.1152/japplphysiol.00166.2012
    1. Drake J. C., Peelor F. F., III, Biela L. M., Watkins M. K., Miller R. A., Hamilton K. L., et al. (2013). Assessment of mitochondrial biogenesis and mTORC1 signaling during chronic rapamycin feeding in male and female mice. J. Gerontol. A Biol. Sci. Med. Sci. 68 1493–1501. 10.1093/gerona/glt047
    1. Drake J. C., Wilson R. J., Yan Z. (2016). Molecular mechanisms for mitochondrial adaptation to exercise training in skeletal muscle. FASEB J. 30 13–22. 10.1096/fj.15-276337
    1. Fahs C. A., Loenneke J. P., Thiebaud R. S., Rossow L. M., Kim D., Abe T., et al. (2015). Muscular adaptations to fatiguing exercise with and without blood flow restriction. Clin. Physiol. Funct. Imaging 35 167–176. 10.1111/cpf.12141
    1. Farup J., de Paoli F., Bjerg K., Riis S., Ringgard S., Vissing K. (2015). Blood flow restricted and traditional resistance training performed to fatigue produce equal muscle hypertrophy. Scand. J. Med. Sci. Sports 25 754–763. 10.1111/sms.12396
    1. Ganesan G., Cotter J. A., Reuland W., Cerussi A. E., Tromberg B. J., Galassetti P. (2015). Effect of blood flow restriction on tissue oxygenation during knee extension. Med. Sci. Sports Exerc. 47 185–193. 10.1249/mss.0000000000000393
    1. Garber C. E., Blissmer B., Deschenes M. R., Franklin B. A., Lamonte M. J., Lee I. M., et al. (2011). American College of Sports Medicine position stand. Quantity and quality of exercise for developing and maintaining cardiorespiratory, musculoskeletal, and neuromotor fitness in apparently healthy adults: guidance for prescribing exercise. Med. Sci. Sports Exerc. 43 1334–1359. 10.1249/MSS.0b013e318213fefb
    1. Gram M., Vigelso A., Yokota T., Hansen C. N., Helge J. W., Hey-Mogensen M., et al. (2014). Two weeks of one-leg immobilization decreases skeletal muscle respiratory capacity equally in young and elderly men. Exp. Gerontol. 58 269–278. 10.1016/j.exger.2014.08.013
    1. Granata C., Jamnick N. A., Bishop D. J. (2018). Training-induced changes in mitochondrial content and respiratory function in human skeletal muscle. Sports Med. 48 1809–1828. 10.1007/s40279-018-0936-y
    1. Groennebaek T., Vissing K. (2017). Impact of resistance training on skeletal muscle mitochondrial biogenesis, content, and function. Front. Physiol. 8:713 10.3389/fphys.2017.00713
    1. Gurtler A., Kunz N., Gomolka M., Hornhardt S., Friedl A. A., McDonald K., et al. (2013). Stain-Free technology as a normalization tool in Western blot analysis. Anal. Biochem. 433 105–111. 10.1016/j.ab.2012.10.010
    1. Henneman E. (1957). Relation between size of neurons and their susceptibility to discharge. Science 126 1345–1347. 10.1126/science.126.3287.1345
    1. Hesselink M. K., Schrauwen-Hinderling V., Schrauwen P. (2016). Skeletal muscle mitochondria as a target to prevent or treat type 2 diabetes mellitus. Nat. Rev. Endocrinol. 12 633–645. 10.1038/nrendo.2016.104
    1. Holloway G. P., Holwerda A. M., Miotto P. M., Dirks M. L., Verdijk L. B., van Loon L. J. C. (2018). Age-associated impairments in mitochondrial ADP sensitivity contribute to redox stress in senescent human skeletal muscle. Cell Rep. 22 2837–2848. 10.1016/j.celrep.2018.02.069
    1. Hood D. A., Tryon L. D., Carter H. N., Kim Y., Chen C. C. (2016). Unravelling the mechanisms regulating muscle mitochondrial biogenesis. Biochem. J. 473 2295–2314. 10.1042/bcj20160009
    1. Horman S., Vertommen D., Heath R., Neumann D., Mouton V., Woods A., et al. (2006). Insulin antagonizes ischemia-induced Thr172 phosphorylation of AMP-activated protein kinase alpha-subunits in heart via hierarchical phosphorylation of Ser485/491. J. Biol. Chem. 281 5335–5340. 10.1074/jbc.M506850200
    1. Hughes L., Paton B., Rosenblatt B., Gissane C., Patterson S. D. (2017). Blood flow restriction training in clinical musculoskeletal rehabilitation: a systematic review and meta-analysis. Br. J. Sports Med. 51 1003–1011. 10.1136/bjsports-2016-097071
    1. Irrcher I., Ljubicic V., Hood D. A. (2009). Interactions between ROS and AMP kinase activity in the regulation of PGC-1alpha transcription in skeletal muscle cells. Am. J. Physiol. Cell Physiol. 296 C116–C123. 10.1152/ajpcell.00267.2007
    1. Irving B. A., Lanza I. R., Henderson G. C., Rao R. R., Spiegelman B. M., Nair K. S. (2015). Combined training enhances skeletal muscle mitochondrial oxidative capacity independent of age. J. Clin. Endocrinol. Metab. 100 1654–1663. 10.1210/jc.2014-3081
    1. Jakobsgaard J. E., Christiansen M., Sieljacks P., Wang J., Groennebaek T., de Paoli F., et al. (2018). Impact of blood flow-restricted bodyweight exercise on skeletal muscle adaptations. Clin. Physiol. Funct. Imaging 10.1111/cpf.12509 [Epub ahead of print].
    1. Jespersen N. R., Yokota T., Stottrup N. B., Bergdahl A., Paelestik K. B., Povlsen J. A., et al. (2017). Pre-ischaemic mitochondrial substrate constraint by inhibition of malate-aspartate shuttle preserves mitochondrial function after ischaemia-reperfusion. J. Physiol. 595 3765–3780. 10.1113/JP273408
    1. Kelley D. E., He J., Menshikova E. V., Ritov V. B. (2002). Dysfunction of mitochondria in human skeletal muscle in type 2 diabetes. Diabetes Metab. Res. Rev. 51 2944–2950. 10.2337/diabetes.51.10.2944
    1. Kido K., Yokokawa T., Ato S., Sato K., Fujita S. (2017). Effect of resistance exercise under conditions of reduced blood insulin on AMPK alpha Ser485/491 inhibitory phosphorylation and AMPK pathway activation. Am. J. Physiol. Regul. Integr. Comp. Physiol. 313 R110–R119. 10.1152/ajpregu.00063.2017
    1. Kjobsted R., Hingst J. R., Fentz J., Foretz M., Sanz M. N., Pehmoller C., et al. (2018). AMPK in skeletal muscle function and metabolism. FASEB J. 32 1741–1777. 10.1096/fj.201700442R
    1. Konopka A. R., Laurin J. L., Musci R. V., Wolff C. A., Reid J. J., Biela L. M., et al. (2017). Influence of Nrf2 activators on subcellular skeletal muscle protein and DNA synthesis rates after 6 weeks of milk protein feeding in older adults. Geroscience 39 175–186. 10.1007/s11357-017-9968-8
    1. Korthuis R. J., Granger D. N., Townsley M. I., Taylor A. E. (1985). The role of oxygen-derived free radicals in ischemia-induced increases in canine skeletal muscle vascular permeability. Circ. Res. 57 599–609. 10.1161/01.RES.57.4.599
    1. Kudo N., Barr A. J., Barr R. L., Desai S., Lopaschuk G. D. (1995). High rates of fatty acid oxidation during reperfusion of ischemic hearts are associated with a decrease in malonyl-CoA levels due to an increase in 5’-AMP-activated protein kinase inhibition of acetyl-CoA carboxylase. J. Biol. Chem. 270 17513–17520. 10.1074/jbc.270.29.17513
    1. Larsen S., Nielsen J., Hansen C. N., Nielsen L. B., Wibrand F., Stride N., et al. (2012). Biomarkers of mitochondrial content in skeletal muscle of healthy young human subjects. J. Physiol. 590 3349–3360. 10.1113/jphysiol.2012.230185
    1. Lauver J. D., Cayot T. E., Rotarius T., Scheuermann B. W. (2017). The effect of eccentric exercise with blood flow restriction on neuromuscular activation, microvascular oxygenation, and the repeated bout effect. Eur. J. Appl. Physiol. 117 1005–1015. 10.1007/s00421-017-3589-x
    1. Loenneke J. P., Kim D., Fahs C. A., Thiebaud R. S., Abe T., Larson R. D., et al. (2015). The effects of resistance exercise with and without different degrees of blood-flow restriction on perceptual responses. J. Sports Sci. 33 1472–1479. 10.1080/02640414.2014.992036
    1. McGee S. L., Hargreaves M. (2010). AMPK-mediated regulation of transcription in skeletal muscle. Clin. Sci. 118 507–518. 10.1042/cs20090533
    1. Meinild Lundby A. K., Jacobs R. A., Gehrig S., de Leur J., Hauser M., Bonne T. C., et al. (2017). Exercise training increases skeletal muscle mitochondrial volume density by enlargement of existing mitochondria and not de novo biogenesis. Acta Physiol. 222:e12905. 10.1111/apha.12905
    1. Miller B. F., Hamilton K. L. (2012). A perspective on the determination of mitochondrial biogenesis. Am. J. Physiol. Endocrinol. Metab. 302 E496–E499. 10.1152/ajpendo.00578.2011
    1. Miller B. F., Wolff C. A., Peelor F. F., III, Shipman P. D., Hamilton K. L. (2015). Modeling the contribution of individual proteins to mixed skeletal muscle protein synthetic rates over increasing periods of label incorporation. J. Appl. Physiol. 118 655–661. 10.1152/japplphysiol.00987.2014
    1. Pesta D., Hoppel F., Macek C., Messner H., Faulhaber M., Kobel C., et al. (2011). Similar qualitative and quantitative changes of mitochondrial respiration following strength and endurance training in normoxia and hypoxia in sedentary humans. Am. J. Physiol. Regul. Integr. Comp. Physiol. 301R1078–R1087. 10.1152/ajpregu.00285.2011
    1. Picard M., Taivassalo T., Gouspillou G., Hepple R. T. (2011a). Mitochondria: isolation, structure and function. J. Physiol. 589(Pt 18), 4413–4421. 10.1113/jphysiol.2011.212712
    1. Picard M., Taivassalo T., Ritchie D., Wright K. J., Thomas M. M., Romestaing C., et al. (2011b). Mitochondrial structure and function are disrupted by standard isolation methods. PLoS One 6:e18317. 10.1371/journal.pone.0018317
    1. Porter C., Reidy P. T., Bhattarai N., Sidossis L. S., Rasmussen B. B. (2015). Resistance exercise training alters mitochondrial function in human skeletal muscle. Med. Sci. Sports Exerc. 47 1922–1931. 10.1249/MSS.0000000000000605
    1. Rahbek S. K., Farup J., de Paoli F., Vissing K. (2015). No differential effects of divergent isocaloric supplements on signaling for muscle protein turnover during recovery from muscle-damaging eccentric exercise. Amino Acids 47 767–778. 10.1007/s00726-014-1907-8
    1. Richter E. A., Ruderman N. B. (2009). AMPK and the biochemistry of exercise: implications for human health and disease. Biochem. J. 418 261–275. 10.1042/bj20082055
    1. Robbins J. L., Jones W. S., Duscha B. D., Allen J. D., Kraus W. E., Regensteiner J. G., et al. (2011). Relationship between leg muscle capillary density and peak hyperemic blood flow with endurance capacity in peripheral artery disease. J. Appl. Physiol. 111 81–86. 10.1152/japplphysiol.00141.2011
    1. Robinson M. M., Dasari S., Konopka A. R., Johnson M. L., Manjunatha S., Esponda R. R., et al. (2017). Enhanced protein translation underlies improved metabolic and physical adaptations to different exercise training modes in young and old humans. Cell Metab. 25 581–592. 10.1016/j.cmet.2017.02.009
    1. Robinson M. M., Turner S. M., Hellerstein M. K., Hamilton K. L., Miller B. F. (2011). Long-term synthesis rates of skeletal muscle DNA and protein are higher during aerobic training in older humans than in sedentary young subjects but are not altered by protein supplementation. FASEB J. 25 3240–3249. 10.1096/fj.11-186437
    1. Rontoyanni V. G., Nunez Lopez O., Fankhauser G. T., Cheema Z. F., Rasmussen B. B., Porter C. (2017). Mitochondrial bioenergetics in the metabolic myopathy accompanying peripheral artery disease. Front. Physiol. 8:141. 10.3389/fphys.2017.00141
    1. Ryu D., Mouchiroud L., Andreux P. A., Katsyuba E., Moullan N., Nicolet-Dit-Felix A. A., et al. (2016). Urolithin A induces mitophagy and prolongs lifespan in C. elegans and increases muscle function in rodents. Nat. Med. 22 879–888. 10.1038/nm.4132
    1. Salvadego D., Domenis R., Lazzer S., Porcelli S., Rittweger J., Rizzo G., et al. (2013). Skeletal muscle oxidative function in vivo and ex vivo in athletes with marked hypertrophy from resistance training. J. Appl. Physiol. 114 1527–1535. 10.1152/japplphysiol.00883.2012
    1. Scalzo R. L., Peltonen G. L., Binns S. E., Shankaran M., Giordano G. R., Hartley D. A., et al. (2014). Greater muscle protein synthesis and mitochondrial biogenesis in males compared with females during sprint interval training. FASEB J. 28 2705–2714. 10.1096/fj.13-246595
    1. Schrauwen-Hinderling V. B., Kooi M. E., Hesselink M. K., Jeneson J. A., Backes W. H., van Echteld C. J., et al. (2007). Impaired in vivo mitochondrial function but similar intramyocellular lipid content in patients with type 2 diabetes mellitus and BMI-matched control subjects. Diabetologia 50 113–120. 10.1007/s00125-006-0475-1
    1. Scott B. R., Loenneke J. P., Slattery K. M., Dascombe B. J. (2015). Exercise with blood flow restriction: an updated evidence-based approach for enhanced muscular development. Sports Med. 45 313–325. 10.1007/s40279-014-0288-1
    1. Sieljacks P., Knudsen L., Wernbom M., Vissing K. (2018). Body position influences arterial occlusion pressure: implications for the standardization of pressure during blood flow restricted exercise. Eur. J. Appl. Physiol. 118 303–312. 10.1007/s00421-017-3770-2
    1. Sieljacks P., Matzon A., Wernbom M., Ringgaard S., Vissing K., Overgaard K. (2016). Muscle damage and repeated bout effect following blood flow restricted exercise. Eur. J. Appl. Physiol. 116 513–525. 10.1007/s00421-015-3304-8
    1. Simmons E., Fluckey J. D., Riechman S. E. (2016). Cumulative muscle protein synthesis and protein intake requirements. Annu. Rev. Nutr. 36 17–43. 10.1146/annurev-nutr-071813-105549
    1. Smiles W. J., Conceicao M. S., Telles G. D., Chacon-Mikahil M. P., Cavaglieri C. R., Vechin F. C., et al. (2017). Acute low-intensity cycling with blood-flow restriction has no effect on metabolic signaling in human skeletal muscle compared to traditional exercise. Eur. J. Appl. Physiol. 117 345–358. 10.1007/s00421-016-3530-8
    1. Taylor C. W., Ingham S. A., Ferguson R. A. (2016). Acute and chronic effect of sprint interval training combined with postexercise blood-flow restriction in trained individuals. Exp. Physiol. 101 143–154. 10.1113/ep085293
    1. Valentine R. J., Coughlan K. A., Ruderman N. B., Saha A. K. (2014). Insulin inhibits AMPK activity and phosphorylates AMPK Ser(4)(8)(5)/(4)(9)(1) through Akt in hepatocytes, myotubes and incubated rat skeletal muscle. Arch. Biochem. Biophys. 562 62–69. 10.1016/j.abb.2014.08.013
    1. Valsecchi F., Ramos-Espiritu L. S., Buck J., Levin L. R., Manfredi G. (2013). cAMP and mitochondria. Physiology 28 199–209. 10.1152/physiol.00004.2013
    1. Vissing K., Andersen J. L., Schjerling P. (2005). Are exercise-induced genes induced by exercise? FASEB J. 19 94–96. 10.1096/fj.04-2084fje
    1. Vissing K., McGee S., Farup J., Kjolhede T., Vendelbo M., Jessen N. (2013). Differentiated mTOR but not AMPK signaling after strength vs endurance exercise in training-accustomed individuals. Scand. J. Med. Sci. Sports 23 355–366. 10.1111/j.1600-0838.2011.01395.x
    1. Wilkinson D. J., Brook M. S., Smith K., Atherton P. J. (2017). Stable isotope tracers and exercise physiology: past, present and future. J. Physiol. 595 2873–2882. 10.1113/jp272277
    1. Wilkinson S. B., Phillips S. M., Atherton P. J., Patel R., Yarasheski K. E., Tarnopolsky M. A., et al. (2008). Differential effects of resistance and endurance exercise in the fed state on signalling molecule phosphorylation and protein synthesis in human muscle. J. Physiol. 586 3701–3717. 10.1113/jphysiol.2008.153916
    1. Zhan M., Jin B., Chen S. E., Reecy J. M., Li Y. P. (2007). TACE release of TNF-alpha mediates mechanotransduction-induced activation of p38 MAPK and myogenesis. J. Cell Sci. 120(Pt 4), 692–701. 10.1242/jcs.03372
    1. Zhang Y., Uguccioni G., Ljubicic V., Irrcher I., Iqbal S., Singh K., et al. (2014). Multiple signaling pathways regulate contractile activity-mediated PGC-1alpha gene expression and activity in skeletal muscle cells. Physiol. Rep. 2:e12008. 10.14814/phy2.12008
    1. Zizola C., Schulze P. C. (2013). Metabolic and structural impairment of skeletal muscle in heart failure. Heart Fail. Rev. 18 623–630. 10.1007/s10741-012-9353-8

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