Skeletal muscle is the most abundant tissue in the human body and is capable of nearly instantaneously increasing its rate of energy turnover more than 100-fold at the onset of muscle contractions. Not surprisingly, mitochondrial deficits in muscle have been linked to the development of a large number of pathologies including diabetes and aging. Moreover, reactive oxygen species (ROS), a byproduct of mitochondrial energy conversion, are well known to be toxic in most tissues and have been implicated in the etiology of diseases such as Alzheimer’s, Parkinson’s, and cancer. Thus, a better understanding of the mechanisms regulating mitochondrial energy conversion is critical not only to improving skeletal muscle function but also to the development of potential treatments for a wide array of human diseases.
The focus of Dr. Glancy’s muscle energetics research program is to determine how mitochondria are optimized within muscle cells to help maintain energy homeostasis during the large change in energy demand caused by muscle contraction. Particularly, his work aims to answer four broad questions: 1) how is mitochondrial energy conversion acutely up-regulated to meet energetic demand during a bout of muscle contractions?, 2) what chronic mitochondrial adaptations are available to the muscle cell to improve the capacity for matching energy supply with demand?, 3) how does each chronic adaptation, or combinations thereof, alter the capacity for acute up-regulation of mitochondrial conversion?, and 4) how do acute mismatches between mitochondrial energy conversion and cellular energy demand signal chronic mitochondrial adaptations in skeletal muscle?
Recently, Dr. Glancy demonstrated that skeletal muscle mitochondria form a highly connected network resembling that of an electrical power grid, and, indeed, were capable of electrical conduction of the mitochondrial membrane potential throughout the cell. The discovery of this rapid energy distribution mechanism overturned longstanding ideas regarding diffusion as the primary energy distribution pathway in skeletal muscle. Current work in this area is focused on providing a better understanding of the regulation and development of these mitochondrial networks as well as their role in overall muscle function.
Despite much interest, control of mitochondrial function in vivo remains largely unclear as direct measures of mitochondrial enzymes in live animals have been limited. As a result, another major focus of the Glancy lab is to develop and utilize novel, direct measurements of in vivo mitochondrial function in skeletal muscle under different workloads. The goal of these studies is to unravel the signaling cascade involved in the upregulation of mitochondrial energy conversion during muscle contraction.
Muscle mitochondria form highly connected networks. a, 3D surface rendering of 25.53 × 24.06 × 4.23 μm FIB-SEM volume segmented to show spatial relationships between mitochondria (green) and other structures (nucleus (N), cyan; capillary (V), magenta; red blood cell, red; myofibrils, grey). b, Removing myofibrils highlights different morphologies within intra-fibrillar mitochondrial (IFM) network. c–e, Zooming in reveals projections from paravascular mitochondria (PVM) into I-band mitochondria (IBM) (c), and numerous interactions between IBM and cross-fiber connection mitochondria (CFCM) (d) and fiber parallel mitochondria (FPM) (d, e). Scale bars, 3 µm. Representative of eight separate volumes analyzed from four animals. From Nature 523, 617–620 (30 July 2015).
Capacity for membrane potential conduction. a, 3D rendering of muscle fibre immunostained for both complex IV and complex V. Confocal image colored according to complex IV/complex V ratio. Relatively higher complex IV, green pixels; relatively higher complex V, red pixels. Nuclei, blue. b, c, Separation of PVM around fibre periphery (b) and IFM (c) within fiber as used for calculations. d, PVM have relatively greater capacity for membrane potential generation while IFM have greater capacity for membrane potential utilization. Images are representative of data from 12 fibers, 5 mice. Error bars indicate standard error. Asterisk indicates significantly different from PVM (paired t-test, P < 0.05). Scale bars, 15 μm. From Nature 523, 617–620 (30 July 2015).
In Vivo, 3D Assessment of Mitochondrial and Muscular Structure. 2D images of a large field of view, 3D image stack collected from C57BL/6 mouse TA muscle in vivo. (A) Representative XY image plane provides longitudinal view of in vivo skeletal muscle fibers. (B) Representative YZ image plane shows cross-sectional view of in vivo skeletal muscle fibers. Green color represents endogenous mitochondrial NAD(P)H fluorescence. Red color is from di-8-ANEPPS fluorescent dye injected into the vasculature. Blue color is from fluorescent CFSE dye in the interstitial space. Because CFSE was also injected into the vasculature, the signal from the residual CFSE remaining in the blood vessels was subtracted out using the di-8-ANEPPS signal to yield just the CFSE signal from the interstitial space. X-, Y-, and Z-pixel sizes are 0.85 lm. Images sizes are 825 lm in X and Y and 280 lm in Z. From Microcirculation 21 (2), 131–147, February 2014.
The video above is a 360° rotation of a 3D rendering of a portion of the skeletal muscle mitochondrial network. Non-white colors indicate individual mitochondria. From Nature 523, 617–620 (30 July 2015).
The video above is a high resolution 3D image stack of a live mouse Tibialis anterior muscle, showing blood flowing in the capillaries (red) embedded in the sarcolemma of muscle fibers. Mitochondrial NAD(P)H – green. Extracellular space – blue. Time in the movie represents progression along the z-axis of the 3D volume.