The 3D nanostructure of the heart, its dynamic deformation during cycles of contraction and relaxation, and the effects of this deformation on cell function remain largely uncharted territory. Over the past decade, the first inroads have been made towards 3D reconstruction of heart cells, with a native resolution of around 1 nm3, and of individual molecules relevant to heart function at a near-atomic scale. These advances have provided access to a new generation of data and have driven the development of increasingly smart, artificial intelligence-based, deep-learning image-analysis algorithms. By high-pressure freezing of cardiomyocytes with millisecond accuracy after initiation of an action potential, pseudodynamic snapshots of contraction-induced deformation of intracellular organelles can now be captured. In combination with functional studies, such as fluorescence imaging, exciting insights into cardiac autoregulatory processes at nano-to-micro scales are starting to emerge. In this Review, we discuss the progress in this fascinating new field to highlight the fundamental scientific insight that has emerged, based on technological breakthroughs in biological sample preparation, 3D imaging and data analysis; to illustrate the potential clinical relevance of understanding 3D cardiac nanodynamics; and to predict further progress that we can reasonably expect to see over the next 10 years.
Electron microscopy (EM) methods are currently the only means of obtaining (sub-)nanometre-scale information on most biological structures.
After decades of dwindling interest, seminal developments in sample preparation, imaging and analysis have led to a renaissance of EM.
However, data acquisition and processing remain time-consuming and laborious, and the inability to observe dynamic events in live cells limits the uptake and utility of EM.
Recent developments promise the advent of temporally resolved, structure–function-correlative, large-volume, 3D EM.
Initial applications of these new developments show EM to be a powerful driver of modern fundamental and translational heart research.
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Palade, G. E. The fine structure of mitochondria. Anat. Rec. 114, 427–451 (1952).
Beams, H. W. & Evans, T. C. Electron microscope studies on the structure of cardiac muscle. Anat. Rec. 105, 59–81 (1949).
Kisch, B., Grey, C. E. & Kelsch, J. J. Electron histology of the heart. Exp. Med. Surg. 6, 346–365 (1948).
Hill, A. V. The abrupt transition from rest to activity in muscle. Proc. R. Soc. B 136, 399–420 (1949).
Lindner, E. Die submikroskopische Morphologie des Herzmuskels. Z. Zellforsch. Mikrosk. Anat. 45, 702–746 (1957).
Moore, D. H. & Ruska, H. Electron microscope study of mammalian cardiac muscle cells. J. Biophys. Biochem. Cytol. 3, 261–268 (1957).
Hanson, J. & Huxley, H. E. Structural basis of the cross-striations in muscle. Nature 172, 530–532 (1953).
Costantin, L. L., Franzini-Armstrong, C. & Podolsky, R. J. Localization of calcium-accumulating structures in striated muscle fibers. Science 147, 158–160 (1965).
Porter, K. R. & Palade, G. E. Studies on the endoplasmic reticulum. Its form and distribution in striated muscle cells. J. Biophys. Biochem. Cytol. 3, 269–300 (1957).
Fawcett, D. W. & McNutt, N. S. The ultrastructure of the cat myocardium. I. Ventricular papillary muscle. J. Cell Biol. 42, 1–45 (1969).
Franzini-Armstrong, C. Studies of the triad: structure of the junction in frog twitch fibers. J. Cell Biol. 47, 488–499 (1970).
Block, B. A., Imagawa, T., Campbell, K. P. & Franzini-Armstrong, C. Structural evidence for direct interaction between the molecular components of the transverse tubule/sarcoplasmic reticulum junction in skeletal muscle. J. Cell Biol. 107, 2587–2600 (1988).
Fleischer, S., Ogunbunmi, E. M., Dixon, M. C. & Fleer, E. A. Localization of Ca2+ release channels with ryanodine in junctional terminal cisternae of sarcoplasmic reticulum of fast skeletal muscle. Proc. Natl Acad. Sci. USA 82, 7256–7259 (1985).
Lai, F. A., Erickson, H. P., Rousseau, E., Liu, Q. Y. & Meissner, G. Purification and reconstitution of the calcium release channel from skeletal muscle. Nature 331, 315–319 (1988).
Franzini-Armstrong, C., Protasi, F. & Ramesh, V. Shape, size, and distribution of Ca2+ release units and couplons in skeletal and cardiac muscles. Biophys. J. 77, 1528–1539 (1999).
Revel, J. P. & Karnovsky, M. J. Hexagonal array of subunits in intercellular junctions of the mouse heart and liver. J. Cell Biol. 33, 7–12 (1967).
Legato, M. J. The correlation of ultrastructure and function in the mammalian myocardial cell. Prog. Cardiovasc. Dis. 11, 391–409 (1969).
Johnson, E. A. & Sommer, J. R. A strand of cardiac muscle. Its ultrastructure and the electrophysiological implications of its geometry. J. Cell Biol. 33, 103–129 (1967).
Levin, K. R. & Page, E. Quantitative studies on plasmalemmal folds and caveolae of rabbit ventricular myocardial cells. Circ. Res. 46, 244–255 (1980).
Schaper, J. & Schaper, W. Ultrastructural correlates of reduced cardiac function in human heart disease. Eur. Heart J. 4, 35–42 (1983).
Heggtveit, H. A. Contributions of electron microscopy to the study of myocardial ischaemia. Bull. World Health Organ. 41, 865–872 (1969).
Lichtig, C. & Brooks, H. Myocardial ultrastructure and function during progressive early ischemia in the intact heart. J. Thorac. Cardiovasc. Surg. 70, 309–315 (1975).
Klar, T. A., Jakobs, S., Dyba, M., Egner, A. & Hell, S. W. Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission. Proc. Natl Acad. Sci. USA 97, 8206–8210 (2000).
Kohl, T., Westphal, V., Hell, S. W. & Lehnart, S. E. Superresolution microscopy in heart – cardiac nanoscopy. J. Mol. Cell. Cardiol. 58, 13–21 (2013).
Gwosch, K. C. et al. MINFLUX nanoscopy delivers 3D multicolor nanometer resolution in cells. Nat. Methods 17, 217–224 (2020).
M’Saad, O. & Bewersdorf, J. Light microscopy of proteins in their ultrastructural context. Nat. Commun. 11, 3850 (2020).
De Mazière, A. M. G. L., van Ginneken, A. C. G., Wilders, R., Jongsma, H. J. & Bouman, L. N. Spatial and functional relationship between myocytes and fibroblasts in the rabbit sinoatrial node. J. Mol. Cell. Cardiol. 24, 567–578 (1992).
Pinali, C., Bennett, H., Davenport, J. B., Trafford, A. W. & Kitmitto, A. Three-dimensional reconstruction of cardiac sarcoplasmic reticulum reveals a continuous network linking transverse-tubules: this organization is perturbed in heart failure. Circ. Res. 113, 1219–1230 (2013).
Rog-Zielinska, E. A. et al. Electron tomography of rabbit cardiomyocyte three-dimensional ultrastructure. Prog. Biophys. Mol. Biol. 121, 77–84 (2016).
Dubochet, J. Cryo-EM–the first thirty years. J. Microsc. 245, 221–224 (2012).
Dubochet, J., Lepault, J., Freeman, R., Berriman, J. A. & Homo, J. C. Electron microscopy of frozen water and aqueous solutions. J. Microsc. 128, 219–237 (1982).
Bers, D. M. Cardiac excitation–contraction coupling. Nature 415, 198–205 (2002).
Maleckar, M. M., Edwards, A. G., Louch, W. E. & Lines, G. T. Studying dyadic structure-function relationships: a review of current modeling approaches and new insights into Ca2+ (mis)handling. Clin. Med. Insights Cardiol. 11, 1179546817698602 (2017).
Knollmann, B. C. et al. Casq2 deletion causes sarcoplasmic reticulum volume increase, premature Ca2+ release, and catecholaminergic polymorphic ventricular tachycardia. J. Clin. Invest. 116, 2510–2520 (2006).
Swift, F. et al. Extreme sarcoplasmic reticulum volume loss and compensatory T-tubule remodeling after SERCA knockout. Proc. Natl Acad. Sci. USA 109, 3997–4001 (2012).
Jones, P. P., MacQuaide, N. & Louch, W. E. Dyadic plasticity in cardiomyocytes. Front. Physiol. 9, 1773 (2018).
Lugo, C. A., Cantalapiedra, I. R., Peñaranda, A., Hove-Madsen, L. & Echebarria, B. Are SR Ca2+ content fluctuations or SR refractoriness the key to atrial cardiac alternans? Insights from a human atrial model. Am. J. Physiol. Heart Circ. Physiol. 306, 1540–1552 (2014).
Wu, H.-D. et al. Ultrastructural remodelling of Ca2+ signalling apparatus in failing heart cells. Cardiovasc. Res. 95, 430–438 (2012).
van Oort, R. J. et al. Disrupted junctional membrane complexes and hyperactive ryanodine receptors after acute junctophilin knockdown in mice. Circulation 123, 979–988 (2011).
Langer, G. A. & Peskoff, A. Calcium concentration and movement in the diadic cleft space of the cardiac ventricular cell. Biophys. J. 70, 1169–1182 (1996).
Frank, J. S. & Langer, G. A. The myocardial interstitium: its structure and its role in ionic exchange. J. Cell Biol. 60, 586–601 (1974).
Novotová, M. et al. Structural variability of dyads relates to calcium release in rat ventricular myocytes. Sci. Rep. 10, 8076 (2020).
Scriven, D. R. L., Asghari, P. & Moore, E. D. W. Microarchitecture of the dyad. Cardiovasc. Res. 98, 169–176 (2013).
Colman, M. A., Pinali, C., Trafford, A. W., Zhang, H. & Kitmitto, A. A computational model of spatio-temporal cardiac intracellular calcium handling with realistic structure and spatial flux distribution from sarcoplasmic reticulum and t-tubule reconstructions. PLoS Comp. Biol. 13, e1005714 (2017).
Sun, X. H. et al. Molecular architecture of membranes involved in excitation-contraction coupling of cardiac muscle. J. Cell Biol. 129, 659–671 (1995).
Hopwood, D. Cell and tissue fixation. Histochem. J. 17, 389–442 (1985).
Gerdes, A. M., Kriseman, J. & Bishop, S. P. Morphometric study of cardiac muscle: the problem of tissue shrinkage. Lab. Invest. 46, 271–274 (1982).
Dobro, M. J., Melanson, L. A., Jensen, G. J. & McDowall, A. W. Plunge freezing for electron cryomicroscopy. Meth. Enzymol. 481, 63–82 (2010).
Padrón, R., Alamo, L., Craig, R. & Caputo, C. A method for quick-freezing live muscles at known instants during contraction with simultaneous recording of mechanical tension. J. Microsc. 151, 81–102 (1988).
Moor, H. & Riehle, U. Snap freezing under high pressure: a new fixation technique for freeze-etching. In Proc. 4th European Regional Conference on Electron Microscopy (ed. Bocciarelli, S.) 2, 33–34 (Rome, 1968).
Gilkey, J. C. & Staehelin, L. A. Advances in ultrarapid freezing for the preservation of cellular ultrastructure. Microsc. Res. Tech. 3, 177–210 (1986).
Rog-Zielinska, E. A. et al. Nano-scale morphology of cardiomyocyte t-tubule/sarcoplasmic reticulum junctions revealed by ultra-rapid high-pressure freezing and electron tomography. J. Mol. Cell. Cardiol. 153, 86–92 (2021).
Cannell, M. B., Kong, C. H., Imtiaz, M. S. & Laver, D. R. Control of sarcoplasmic reticulum Ca2+ release by stochastic RyR gating within a 3D model of the cardiac dyad and importance of induction decay for CICR termination. Biophys. J. 104, 2149–2159 (2013).
Koh, X., Srinivasan, B., Ching, H. S. & Levchenko, A. A 3D Monte Carlo analysis of the role of dyadic space geometry in spark generation. Biophys. J. 90, 1999–2014 (2006).
Dulhunty, A. F. & Franzini-Armstrong, C. The relative contributions of the folds and caveolae to the surface membrane of frog skeletal muscle fibres at different sarcomere lengths. J. Physiol. 250, 513–539 (1975).
Ingber, D. E. Tensegrity-based mechanosensing from macro to micro. Prog. Biophys. Mol. Biol. 97, 163–179 (2008).
Irving, T. C., Konhilas, J., Perry, D., Fischetti, R. & de Tombe, P. P. Myofilament lattice spacing as a function of sarcomere length in isolated rat myocardium. Am. J. Physiol. Heart Circ. Physiol. 279, 2568–2573 (2000).
Talmon, Y., Burns, J. L., Chestnut, M. H. & Siegel, D. P. Time-resolved cryotransmission electron microscopy. J. Electron. Microsc. Tech. 14, 6–12 (1990).
Kohl, P., Cooper, P. J. & Holloway, H. Effects of acute ventricular volume manipulation on in situ cardiomyocyte cell membrane configuration. Prog. Biophys. 82, 221–227 (2003).
MacDonald, E. A. et al. Sinoatrial node structure, mechanics, electrophysiology and the chronotropic response to stretch in rabbit and mouse. Front. Physiol. 11, 809 (2020).
Oda, T. & Yanagisawa, H. Cryo-electron tomography of cardiac myofibrils reveals a 3D lattice spring within the Z-discs. Commun. Biol. 3, 585 (2020).
Risi, C. et al. Ca2+-induced movement of tropomyosin on native cardiac thin filaments revealed by cryoelectron microscopy. Proc. Natl Acad. Sci. USA 114, 6782–6787 (2017).
Sharma, M. R., Jeyakumar, L. H., Fleischer, S. & Wagenknecht, T. Three-dimensional structure of ryanodine receptor isoform three in two conformational states as visualized by cryo-electron microscopy. J. Biol. Chem. 275, 9485–9491 (2000).
Efremov, R. G., Leitner, A., Aebersold, R. & Raunser, S. Architecture and conformational switch mechanism of the ryanodine receptor. Nature 517, 39–43 (2015).
Dhindwal, S. et al. A cryo-EM-based model of phosphorylation- and FKBP12.6-mediated allosterism of the cardiac ryanodine receptor. Sci. Signal. 10, eaai8842 (2017).
des Georges, A. et al. Structural basis for gating and activation of RyR1. Cell 167, 145–157 (2016).
Chi, X. et al. Molecular basis for allosteric regulation of the type 2 ryanodine receptor channel gating by key modulators. Proc. Natl Acad. Sci. USA 116, 25575–25582 (2019).
McNary, T. G. et al. Mechanical modulation of the transverse tubular system of ventricular cardiomyocytes. Prog. Biophys. Mol. Biol. 110, 218–225 (2012).
Rog-Zielinska, E. A. et al. Beat-by-beat cardiomyocyte T-tubule deformation drives tubular content exchange. Circ. Res. 128, 203–215 (2021).
Rog-Zielinska, E. A., O’Toole, E. T., Hoenger, A. & Kohl, P. Mitochondrial deformation during the cardiac mechanical cycle. Anat. Rec. 302, 146–152 (2019).
Sinha, B. et al. Cells respond to mechanical stress by rapid disassembly of caveolae. Cell 144, 402–413 (2011).
Wendt-Gallitelli, M. F. & Isenberg, G. X-ray microanalysis of single cardiac myocytes frozen under voltage-clamp conditions. Am. J. Physiol. 256, 574–583 (1989).
Brette, F. & Orchard, C. T-tubule function in mammalian cardiac myocytes. Circ. Res. 92, 1182–1192 (2003).
Burton, R. A. B. et al. Caveolae in rabbit ventricular myocytes: distribution and dynamic diminution after cell isolation. Biophys. J. 113, 1047–1059 (2017).
Scriven, D. R., Klimek, A., Asghari, P., Bellve, K. & Moore, E. D. Caveolin-3 is adjacent to a group of extradyadic ryanodine receptors. Biophys. J. 89, 1893–1901 (2005).
Page, E. Quantitative ultrastructural analysis in cardiac membrane physiology. Am. J. Physiol. 235, 147–158 (1978).
Wong, J. et al. Nanoscale distribution of ryanodine receptors and caveolin-3 in mouse ventricular myocytes: dilation of t-tubules near junctions. Biophys. J. 104, 22–24 (2013).
Bang, B. H. & Bang, F. B. Graphic reconstruction of the third dimension from serial electron microphotographs. J. Ultrastruct. Res. 1, 138–139 (1957).
Pinali, C. & Kitmitto, A. Serial block face scanning electron microscopy for the study of cardiac muscle ultrastructure at nanoscale resolutions. J. Mol. Cell. Cardiol. 76, 1–11 (2014).
Hayashi, T. et al. Three-dimensional electron microscopy reveals new details of membrane systems for Ca2+ signaling in the heart. J. Cell Sci. 122, 1005–1013 (2009).
McIntosh, R., Nicastro, D. & Mastronarde, D. New views of cells in 3D: an introduction to electron tomography. Trends Cell. Biol. 15, 43–51 (2005).
Asghari, P. et al. Cardiac ryanodine receptor distribution is dynamic and changed by auxiliary proteins and post-translational modification. eLife 9, e51602 (2020).
Asghari, P. et al. Nonuniform and variable arrangements of ryanodine receptors within mammalian ventricular couplons. Circ. Res. 115, 252–262 (2014).
Kong, C. H. T., Rog-Zielinska, E. A., Kohl, P., Orchard, C. H. & Cannell, M. B. Solute movement in the t-tubule system of rabbit and mouse cardiomyocytes. Proc. Natl Acad. Sci. USA 115, 7073–7080 (2018).
Rog-Zielinska, E. A. et al. Species differences in the morphology of transverse tubule openings in cardiomyocytes. Europace 20, 120–124 (2018).
Tsushima, K. et al. Mitochondrial reactive oxygen species in lipotoxic hearts induce post-translational modifications of AKAP121, DRP1, and OPA1 that promote mitochondrial fission. Circ. Res. 122, 58–73 (2018).
Lavorato, M. et al. Increased mitochondrial nanotunneling activity, induced by calcium imbalance, affects intermitochondrial matrix exchanges. Proc. Natl Acad. Sci. USA 114, 849–858 (2017).
Huang, X. et al. Kissing and nanotunneling mediate intermitochondrial communication in the heart. Proc. Natl Acad. Sci. USA 110, 2846–2851 (2013).
Glancy, B. et al. Mitochondrial reticulum for cellular energy distribution in muscle. Nature 523, 617–620 (2015).
Aston, D. et al. High resolution structural evidence suggests the sarcoplasmic reticulum forms microdomains with acidic stores (lysosomes) in the heart. Sci. Rep. 7, 40620 (2017).
Iribe, G. et al. Axial stretch of rat single ventricular cardiomyocytes causes an acute and transient increase in Ca2+ spark rate. Circ. Res. 104, 787–795 (2009).
Peper, J. et al. Caveolin3 stabilizes McT1-mediated lactate/proton transport in cardiomyocytes. Circ. Res. 128, 102–120 (2021).
Gherghiceanu, M. & Popescu, L. M. Heterocellular communication in the heart: electron tomography of telocyte–myocyte junctions. J. Cell. Mol. Med. 15, 1005–1011 (2011).
Quinn, T. A. et al. Electrotonic coupling of excitable and nonexcitable cells in the heart revealed by optogenetics. Proc. Natl Acad. Sci. USA 113, 14852–14857 (2016).
Wang, Z. et al. The molecular basis for sarcomere organization in vertebrate skeletal muscle. Cell 184, 2135–2150 (2021).
Chen, W. & Kudryashev, M. Structure of RyR1 in native membranes. EMBO Rep. 21, 49891 (2020).
Wagenknecht, T., Hsieh, C. & Marko, M. Skeletal muscle triad junction ultrastructure by focused-ion-beam milling of muscle and cryo-electron tomography. Eur. J. Transl. Myol. 25, 49–56 (2015).
De Winter, D. A. M., Hsieh, C., Marko, M. & Hayles, M. F. Cryo-FIB preparation of whole cells and tissue for cryo-TEM: use of high-pressure frozen specimens in tubes and planchets. J. Microsc. 281, 125–137 (2021).
Weber, M. S., Wojtynek, M. & Medalia, O. Cellular and structural studies of eukaryotic cells by cryo-electron tomography. Cells 8, 57 (2019).
Kitmitto, A. Applications of electron cryo-microscopy to cardiovascular research. Methods Mol. Med. 129, 315–327 (2006).
Agip, A. N. A. et al. Cryo-EM structures of complex I from mouse heart mitochondria in two biochemically defined states. Nat. Struct. Mol. Biol. 25, 548–556 (2018).
Risi, C. et al. High-resolution cryo-EM structure of the cardiac actomyosin complex. Structure 29, 50–60 (2021).
von der Ecken, J., Heissler, S. M., Pathan-Chhatbar, S., Manstein, D. J. & Raunser, S. Cryo-EM structure of a human cytoplasmic actomyosin complex at near-atomic resolution. Nature 534, 724–728 (2016).
Elad, N. et al. The role of integrin-linked kinase in the molecular architecture of focal adhesions. J. Cell Sci. 126, 4099–4107 (2013).
Samsó, M. A guide to the 3D structure of the ryanodine receptor type 1 by cryo-EM. Protein Sci. 26, 52–68 (2017).
Dulhunty, A. F., Beard, N. A. & Casarotto, M. G. Recent advances in understanding the ryanodine receptor calcium release channels and their role in calcium signalling. F1000Res. 7, 1851 (2018).
Lee, C. H. & MacKinnon, R. Structures of the human HCN1 hyperpolarization-activated channel. Cell 168, 111–120 (2017).
Noreng, S., Li, T. & Payandeh, J. Structural pharmacology of voltage-gated sodium channels. J. Mol. Biol. 433, 166967 (2021).
Merino, F. et al. Structural transitions of F-actin upon ATP hydrolysis at near-atomic resolution revealed by cryo-EM. Nat. Struct. Mol. Biol. 25, 528–537 (2018).
Wu, M., Gu, J., Guo, R., Huang, Y. & Yang, M. Structure of mammalian respiratory supercomplex I1III2IV1. Cell 167, 1598–1609 (2016).
Zhu, J., Vinothkumar, K. R. & Hirst, J. Structure of mammalian respiratory complex I. Nature 536, 354–358 (2016).
Daghistani, H. M., Rajab, B. S. & Kitmitto, A. Three-dimensional electron microscopy techniques for unravelling mitochondrial dysfunction in heart failure and identification of new pharmacological targets. Br. J. Pharmacol. 176, 4340–4359 (2019).
Turk, M. & Baumeister, W. The promise and the challenges of cryo-electron tomography. FEBS Lett. 594, 3243–3261 (2020).
Renken, C. et al. Structure of frozen–hydrated triad junctions: a case study in motif searching inside tomograms. J. Struct. Biol. 165, 53–63 (2009).
Schorb, M., Haberbosch, I., Hagen, W. J. H., Schwab, Y. & Mastronarde, D. N. Software tools for automated transmission electron microscopy. Nat. Methods 16, 471–477 (2019).
Kremer, J. R., Mastronarde, D. N. & McIntosh, J. R. Computer visualization of three-dimensional image data using IMOD. J. Struct. Biol. 116, 71–76 (1996).
Buchholz, T. O. et al. Content-aware image restoration for electron microscopy. Methods Cell. Biol. 152, 277–289 (2019).
Fang, L. et al. Deep learning-based point-scanning super-resolution imaging. Nat. Methods 18, 406–416 (2021).
Bepler, T., Kelley, K., Noble, A. J. & Berger, B. Topaz-Denoise: general deep denoising models for cryoEM and cryoET. Nat. Commun. 11, 5208 (2020).
Heinrich, L. et al. Whole-cell organelle segmentation in volume electron microscopy. Nature 599, 141–146 (2021).
Berg, S. et al. ilastik: interactive machine learning for (bio)image analysis. Nat. Methods 16, 1226–1232 (2019).
Belevich, I., Joensuu, M., Kumar, D., Vihinen, H. & Jokitalo, E. Microscopy Image Browser: a platform for segmentation and analysis of multidimensional datasets. PLoS Biol. 14, 1002340 (2016).
Vergara, H. M. et al. Whole-body integration of gene expression and single-cell morphology. Cell 184, 4819–4837 (2021).
Perez, A. J. et al. A workflow for the automatic segmentation of organelles in electron microscopy image stacks. Front. Neuroanat. 8, 126 (2014).
Martinez-Sanchez, A., Garcia, I., Asano, S., Lucic, V. & Fernandez, J. J. Robust membrane detection based on tensor voting for electron tomography. J. Struct. Biol. 186, 49–61 (2014).
Hussain, A. et al. An automated workflow for segmenting single adult cardiac cells from large-volume serial block-face scanning electron microscopy data. J. Struct. Biol. 202, 275–285 (2018).
Khadangi, A., Boudier, T. & Rajagopal, V. EM-net: deep learning for electron microscopy image segmentation. bioRxiv https://doi.org/10.1101/2020.02.03.933127 (2020).
Hatano, A. et al. Isolation and reconstruction of cardiac mitochondria from SBEM images using a deep learning-based method. J. Struct. Biol. 214, 107806 (2021).
Xu, C. S. et al. An open-access volume electron microscopy atlas of whole cells and tissues. Nature 599, 147–151 (2021).
Lucchi, A., Smith, K., Achanta, R., Knott, G. & Fua, P. Supervoxel-based segmentation of mitochondria in EM image stacks with learned shape features. IEEE Trans. Med. Imaging 31, 474–486 (2012).
Wei, D. et al. MitoEM dataset: large-scale 3D mitochondria instance segmentation from EM images. Med. Image Comput. Comput. Assist. Interv. 12265, 66–76 (2020).
Maron, B. J., Ferrans, V. J. & Roberts, W. C. Ultrastructural features of degenerated cardiac muscle cells in patients with cardiac hypertrophy. Am. J. Pathol. 79, 387–434 (1975).
Maron, B. J., Ferrans, V. J. & Jones, M. The spectrum of degenerative changes in hypertrophied human cardiac muscle cells: an ultrastructural study. Recent. Adv. Stud. Card. Struct. Metab. 8, 447–466 (1975).
Burch, G. E. Ultrastructural myocardial changes produced by viruses. Recent. Adv. Stud. Card. Struct. Metab. 6, 501–523 (1975).
Brandenburg, S. et al. Junctophilin-2 expression rescues atrial dysfunction through polyadic junctional membrane complex biogenesis. JCI Insight 4, e127116 (2019).
Pinali, C. et al. Post-myocardial infarction T-tubules form enlarged branched structures with dysregulation of junctophilin-2 and bridging integrator 1 (BIN-1). J. Am. Heart Assoc. 6, e004834 (2017).
Collins, H. E. et al. Mitochondrial morphology and mitophagy in heart diseases: qualitative and quantitative analyses using transmission electron microscopy. Front. Aging 2, 670267 (2021).
Lavorato, M. et al. Dyad content is reduced in cardiac myocytes of mice with impaired calmodulin regulation of RyR2. J. Muscle Res. Cell Motil. 36, 205–214 (2015).
Takemura, G. et al. Electron microscopic findings are an important aid for diagnosing mitochondrial cardiomyopathy with mitochondrial DNA mutation 3243A>G. Circ. Heart Fail. 9, e003283 (2016).
Erlandson, R. A. Role of electron microscopy in modern diagnostic surgical pathology. Mod. Surg. Pathol. 1, 71–84 (2009).
Beikoghli Kalkhoran, S. et al. Assessing the effects of mitofusin 2 deficiency in the adult heart using 3D electron tomography. Physiol. Rep. 5, e13437 (2017).
Pinali, C. et al. Three-dimensional structure of the intercalated disc reveals plicate domain and gap junction remodeling in heart failure. Biophys. J. 108, 498–507 (2015).
Toomer, K. A. et al. Primary cilia defects causing mitral valve prolapse. Sci. Transl. Med. 11, eaax0290 (2019).
Ceska, T., Chung, C.-W., Cooke, R., Phillips, C. & Williams, P. A. Cryo-EM in drug discovery. Biochem. Soc. Trans. 47, 281–293 (2019).
Scapin, G., Potter, C. S. & Carragher, B. Cryo-EM for small molecules discovery, design, understanding, and application. Cell. Chem. Biol. 25, 1318–1325 (2018).
Jiang, D. et al. Structure of the cardiac sodium channel. Cell 180, 122–134 (2020).
Jiang, D. et al. Structural basis for voltage-sensor trapping of the cardiac sodium channel by a deathstalker scorpion toxin. Nat. Commun. 12, 128 (2021).
Wang, W. & MacKinnon, R. Cryo-EM structure of the open human ether-à-go-go-related K+ channel hERG. Cell 169, 422–430 (2017).
Wang, M.-C. et al. The three-dimensional structure of the cardiac L-type voltage-gated calcium channel: comparison with the skeletal muscle form reveals a common architectural motif. J. Biol. Chem. 279, 7159–7168 (2004).
Zhao, Y. et al. Molecular basis for ligand modulation of a mammalian voltage-gated Ca2+ channel. Cell 177, 1495–1506 (2019).
Iyer, K. A. et al. Structural mechanism of two gain-of-function cardiac and skeletal RyR mutations at an equivalent site by cryo-EM. Sci. Adv. 6, eabb2964 (2020).
Feng, X. et al. A fast and effective microfluidic spraying-plunging method for high-resolution single-particle cryo-EM. Structure 25, 663–670.e3 (2017).
Unwin, N. & Fujiyoshi, Y. Gating movement of acetylcholine receptor caught by plunge-freezing. J. Mol. Biol. 422, 617–634 (2012).
Lu, Z. et al. Gas-assisted annular microsprayer for sample preparation for time-resolved cryo-electron microscopy. J. Micromech. Microeng. 24, 115001 (2014).
Mäeots, M.-E. et al. Modular microfluidics enables kinetic insight from time-resolved cryo-EM. Nat. Commun. 11, 3465 (2020).
Jorgensen, A. O. & Campbell, K. P. Evidence for the presence of calsequestrin in two structurally different regions of myocardial sarcoplasmic reticulum. J. Cell Biol. 98, 1597–1602 (1984).
Thomas, M. J. et al. Localization and function of the Na+/Ca2+-exchanger in normal and detubulated rat cardiomyocytes. J. Mol. Cell. Cardiol. 35, 1325–1337 (2003).
Darkow, E. et al. The lectin LecA sensitizes the human stretch-activated channel TREK-1 but not Piezo1 and binds selectively to cardiac non-myocytes. Front. Physiol. 11, 457 (2020).
Sartori-Rupp, A. et al. Correlative cryo-electron microscopy reveals the structure of TNTs in neuronal cells. Nat. Commun. 10, 342 (2019).
Shu, X. et al. A genetically encoded tag for correlated light and electron microscopy of intact cells, tissues, and organisms. PLoS Biol. 9, e1001041 (2011).
Micheva, K. D. & Smith, S. J. Array tomography: a new tool for imaging the molecular architecture and ultrastructure of neural circuits. Neuron 55, 25–36 (2007).
The authors thank the staff at the Electron Microscopy Core Facility EMBL Heidelberg for many years of on-site support and advice, as well as A. Vlachos, J. Madl and J. O’Reilly, all at the University of Freiburg, for helpful comments on the manuscript. E.A.R.-Z. is a German Research Foundation Emmy Noether Fellow (DFG #396913060). The authors are members of the German Research Foundation Collaborative Research Centre SFB1425 (DFG #422681845).
The authors declare no competing interests.
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- Electron microscopy
(EM). A microscopy method in which the sample is exposed to a beam of accelerated electrons, while images are obtained either by trans-illumination of samples with spatially variable permeability for electrons, or reconstructed from electrons reflected by the sample surface in scanning mode; this allows in-plane linear resolutions of <1 nm (102–103 times higher than conventional photon-based microscopy).
- Electron tomography
(ET). A 3D EM method in which a sample (typically 200–300 nm thick) is tilted relative to the imaging plane of the electron beam, and individual transmitted images are captured (usually between −60o and +60o tilt levels, and often along two mutually perpendicular tilt axes); using these images, the sample volume can be reconstructed with a native voxel size <1 nm3.
A method of imaging samples while frozen and hydrated; by omitting fixatives, solvent substitution, resin embedding and heavy metal staining, researchers can visualize native nanostructural details down to the level of single molecules.
- Serial block-face or focused ion beam SEM
3D EM methods in which, between individual runs of SEM image acquisition, a thin layer of the sample surface is removed, either mechanically or using a focused ion beam; the sample volume is reconstructed from voxels whose resolution is limited by the surface removal technique, usually yielding voxels of ≥10 nm3.
The property of an object, here the unitary 3D imaging readouts (voxels), that does not have edge lengths of equal size; usually, the edge lengths in the imaging plane are identical, while that between imaging planes is larger (the voxel shape is a square cuboid).
The property of an object, here the unitary 3D imaging readouts (voxels), that has edge lengths of equal size (the voxel shape is a cube).
- Single-particle analysis
(SPA). A variant of cryo-EM that uses post-acquisition methods to three-dimensionally reconstruct individual molecules (such as proteins) by combining multiple images (usually thousands) of a population of molecules at random angular orientations; this approach allows researchers to achieve near-atomic-scale structural resolution.
- Correlative light and electron microscopy
(CLEM). An approach in which a single sample is first imaged using fluorescence microscopy (to visualize the presence or dynamics of suitable reporters in live or fixed cells) and then, aided by meticulous transfer of coordinates, re-imaged using EM; correlation of the resulting data sets allows researchers to interrelate nanoscale to mesoscale structural and functional information on cells and subcellular structures.
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Kohl, P., Greiner, J. & Rog-Zielinska, E.A. Electron microscopy of cardiac 3D nanodynamics: form, function, future. Nat Rev Cardiol 19, 607–619 (2022). https://doi.org/10.1038/s41569-022-00677-x