This work received support through Deutsche Forschungsgemeinschaft SFB 1002 (subprojects A05 and B05 to S.E.L.) and KFO 155 (subproject 4 to S.E.L.), a Halbach Foundation award to S.E.L. supporting E.W.; a grant from the German Cardiac Society to S.B.; and a DAAD exchange program supporting T.K. as visitor at the University of Maryland. The research leading to these results has received funding from the European Community&#8217;s Seventh Framework Program FP7/2007-2013 under grant agreement No. HEALTH-F2-2009-241526, EUTrigTreat (to S.E.L.). S.E.L. is a principal investigator of the German Center of Cardiovascular Research (DZHK).

<ol>
	<li>Prosser, B. L., Ward, C. W., Lederer, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subcellular+Ca2++signaling+in+the+heart+the+role+of+ryanodine+receptor+sensitivity.">Subcellular Ca2+ signaling in the heart the role of ryanodine receptor sensitivity.</a> J Gen Physiol. 136 (2), 135-142 (2010).</li><li>Wehrens, X. H., Lehnart, S. E., Marks, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+calcium+release+and+cardiac+disease.">Intracellular calcium release and cardiac disease.</a> Annu Rev Physiol. 67, 69-98 (2005).</li><li>Cheng, H., Cannell, M. B., Lederer, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Propagation+of+excitation-contraction+coupling+into+ventricular+myocytes.">Propagation of excitation-contraction coupling into ventricular myocytes.</a> Pflugers Arch. 428 (3-4), 415-417 (1994).</li><li>Williams, G. S., Chikando, A. C., Tuan, H. T., Sobie, E. A., Lederer, W. J., Jafri, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+calcium+sparks+and+calcium+leak+in+the+heart.">Dynamics of calcium sparks and calcium leak in the heart.</a> Biophys J. 101 (6), 1287-1296 (2011).</li><li>Sperelakis, N., Rubio, <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=R.+orderly+lattice+of+axial+tubules+which+interconnect+adjacent+transverse+tubules+in+guinea-pig+ventricular+myocardium.">R. orderly lattice of axial tubules which interconnect adjacent transverse tubules in guinea-pig ventricular myocardium.</a> J Mol Cell Cardiol. 2 (3), 211-220 (1971).</li><li>Soeller, C., Cannell, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=B.+of+the+transverse+tubular+system+in+living+cardiac+rat+myocytes+by+2-photon+microscopy+and+digital+image-processing+techniques.">B. of the transverse tubular system in living cardiac rat myocytes by 2-photon microscopy and digital image-processing techniques.</a> Circ Res. 84 (3), 266-275 (1999).</li><li>Song, L. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=et+al.+ryanodine+receptors+in+the+failing+heart.">et al. ryanodine receptors in the failing heart.</a> Proc Natl Acad Sci USA. 103 (11), 4305-4310 (2006).</li><li>Oort, R. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disrupted+junctional+membrane+complexes+and+hyperactive+ryanodine+receptors+after+acute+junctophilin+knockdown+in+mice.">Disrupted junctional membrane complexes and hyperactive ryanodine receptors after acute junctophilin knockdown in mice.</a> Circulation. 123 (9), 979-988 (2011).</li><li>Wagner, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stimulated+emission+depletion+live-cell+super-resolution+imaging+shows+proliferative+remodeling+of+T-tubule+membrane+structures+after+myocardial+infarction.">Stimulated emission depletion live-cell super-resolution imaging shows proliferative remodeling of T-tubule membrane structures after myocardial infarction.</a> Circ Res. 111 (4), 402-414 (2012).</li><li>Asghari, P., Schulson, M., Scriven, D. R., Martens, G., Moore, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axial+tubules+of+rat+ventricular+myocytes+form+multiple+junctions+with+the+sarcoplasmic+reticulum.">Axial tubules of rat ventricular myocytes form multiple junctions with the sarcoplasmic reticulum.</a> Biophys J. 96 (11), 4651-4660 (2009).</li><li>Lukyanenko, V., Ziman, A., Lukyanenko, A., Salnikov, V., Lederer, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+groups+of+ryanodine+receptors+in+rat+ventricular+cells.">Functional groups of ryanodine receptors in rat ventricular cells.</a> J Physiol. 583 (Pt 1), 251-269 (2007).</li><li>Shacklock, P. S., Wier, W. G., Balke, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Local+Ca2++transients+(Ca2++sparks)+originate+at+transverse+tubules+in+rat+heart+cells.">Local Ca2+ transients (Ca2+ sparks) originate at transverse tubules in rat heart cells.</a> J Physiol. 487 (Pt 3), 601-608 (1995).</li><li>Reynolds, J. O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Junctophilin-2+is+necessary+for+T-tubule+maturation+during+mouse+heart+development.">Junctophilin-2 is necessary for T-tubule maturation during mouse heart development.</a> Cardiovasc Res. 100 (1), 44-53 (2013).</li><li>Di Maio, A., Karko, K., Snopko, R. M., Mejia-Alvarez, R., Franzini-Armstrong, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T-tubule+formation+in+cardiac+myocytes+two+possible+mechanisms.">T-tubule formation in cardiac myocytes two possible mechanisms.</a> J Muscle Res Cell Motil. 28 (4-5), 231-241 (2007).</li><li>He, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reduction+in+density+of+transverse+tubules+and+L-type+Ca(2+)+channels+in+canine+tachycardia-induced+heart+failure.">Reduction in density of transverse tubules and L-type Ca(2+) channels in canine tachycardia-induced heart failure.</a> Cardiovasc Res. 49 (2), 298-307 (2001).</li><li>Heinzel, F. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Remodeling+of+T-tubules+and+reduced+synchrony+of+Ca2++release+in+myocytes+from+chronically+ischemic+myocardium.">Remodeling of T-tubules and reduced synchrony of Ca2+ release in myocytes from chronically ischemic myocardium.</a> Circ Res. 102 (3), 338-346 (2008).</li><li>Lyon, A. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Loss+of+T-tubules+and+other+changes+to+surface+topography+in+ventricular+myocytes+from+failing+human+and+rat+heart.">Loss of T-tubules and other changes to surface topography in ventricular myocytes from failing human and rat heart.</a> Proc Natl Acad Sci USA. 106 (16), 6854-6859 (2009).</li><li>Crossman, D. J., Ruygrok, P. N., Soeller, C., Cannell, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Changes+in+the+organization+of+excitation-contraction+coupling+structures+in+failing+human+heart.">Changes in the organization of excitation-contraction coupling structures in failing human heart.</a> PLoS One. 6 (3), e17901 (2011).</li><li>Kemi, O. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+exercise+training+on+transverse+tubules+in+normal,+remodeled,+and+reverse+remodeled+hearts.">The effect of exercise training on transverse tubules in normal, remodeled, and reverse remodeled hearts.</a> J Cell Physiol. 226 (9), 2235-2243 (2011).</li><li>Sachse, F. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subcellular+structures+and+function+of+myocytes+impaired+during+heart+failure+are+restored+by+cardiac+resynchronization+therapy.">Subcellular structures and function of myocytes impaired during heart failure are restored by cardiac resynchronization therapy.</a> Circ Res. 110 (4), 588-597 (2012).</li><li>Arakel, E. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tuning+the+electrical+properties+of+the+heart+by+differential+trafficking+of+KATP+ion+channel+complexes.">Tuning the electrical properties of the heart by differential trafficking of KATP ion channel complexes.</a> J Cell Sciences. 127 (Pt 9), 2106-2119 (2014).</li><li>Richards, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transverse+tubules+are+a+common+feature+in+large+mammalian+atrial+myocytes+including+human.">Transverse tubules are a common feature in large mammalian atrial myocytes including human.</a> Am J Physiol Heart Circ Physiol. 301 (5), H1996-H2005 (2011).</li><li>Trafford, A. W., Clarke, J. D., Richards, M. A., Eisner, D. A., Dibb, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+signalling+microdomains+and+the+t-tubular+system+in+atrial+mycoytes+potential+roles+in+cardiac+disease+and+arrhythmias.">Calcium signalling microdomains and the t-tubular system in atrial mycoytes potential roles in cardiac disease and arrhythmias.</a> Cardiovasc Res. 98 (2), 192-203 (2013).</li><li>Greiser, M., Schotten, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+remodeling+of+intracellular+Ca2++signaling+during+atrial+fibrillation.">Dynamic remodeling of intracellular Ca2+ signaling during atrial fibrillation.</a> J Mol Cell Cardiol. 58, 134-142 (2013).</li><li>Voigt, N., Zhou, X. B., Dobrev, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+human+atrial+myocytes+for+simultaneous+measurements+of+Ca2++transients+and+membrane+currents.">Isolation of human atrial myocytes for simultaneous measurements of Ca2+ transients and membrane currents.</a> J Vis Exp. (77), 10-3791 (2013).</li><li>Kaestner, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+and+genetic+manipulation+of+adult+cardiac+myocytes+for+confocal+imaging.">Isolation and genetic manipulation of adult cardiac myocytes for confocal imaging.</a> J Vis Exp. (31), (2009).</li><li>Louch, W. E., Sheehan, K. A., Wolska, B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+in+cardiomyocyte+isolation,+culture,+and+gene+transfer.">Methods in cardiomyocyte isolation, culture, and gene transfer.</a> J Mol Cell Cardiol. 51 (3), 288-298 (2011).</li><li>King, N. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+intact+cardiac+myocyte+mechanics+cross-bridge+and+titin-based+stress+in+unactivated+cells.">Mouse intact cardiac myocyte mechanics cross-bridge and titin-based stress in unactivated cells.</a> J Gen Physiol. 137 (1), 81-91 (2011).</li><li>Schindelin, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fiji,+an+open-source+platform+for+biological-image+analysis.">Fiji, an open-source platform for biological-image analysis.</a> Nat Methods. 9 (7), 676-682 (2012).</li><li>Arganda-Carreras, I., Fernandez-Gonzalez, R., Munoz-Barrutia, A., Ortiz-De-Solorzano, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+reconstruction+of+histological+sections+Application+to+mammary+gland+tissue.">3D reconstruction of histological sections Application to mammary gland tissue.</a> Microsc Res Tech. 73 (11), 1019-1029 (2010).</li><li>Liu, Z. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scale+space+approach+to+directional+analysis+of+images.">Scale space approach to directional analysis of images.</a> Appl Opt. 30 (11), 1369-1373 (1991).</li><li>Powell, T., Twist, V. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rapid+technique+for+the+isolation+and+purification+of+adult+cardiac+muscle+cells+having+respiratory+control+and+a+tolerance+to+calcium.">A rapid technique for the isolation and purification of adult cardiac muscle cells having respiratory control and a tolerance to calcium.</a> Biochem Biophys Res Commun. 72 (1), 327-333 (1976).</li><li>Watanabe, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhibitory+effect+of+2,3-butanedione+monoxime+(BDM)+on+Na(+)/Ca(2+)+exchange+current+in+guinea-pig+cardiac+ventricular+myocytes.">Inhibitory effect of 2,3-butanedione monoxime (BDM) on Na(+)/Ca(2+) exchange current in guinea-pig cardiac ventricular myocytes.</a> Br J Pharmacol. 132 (6), 1317-1325 (2001).</li><li>Kohl, T., Westphal, V., Hell, S. W., Lehnart, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Superresolution+microscopy+in+heart+-+cardiac+nanoscopy.">Superresolution microscopy in heart - cardiac nanoscopy.</a> J Mol Cell Cardiol. 58, 13-21 (2013).</li></ol>

The authors declare that no conflict of interest exists.

Although cardiac myocytes have been isolated and studied for decades32, a recent review concluded that consistent high-quality myocyte cell isolations remain challenging27. This reflects relatively complex protocols for isolation of primary cardiac myocytes vis-&#224;-vis a lack of common standard approaches, shared metadata, and transparent cell quality documentation. Cell isolation protocols are usually customized by individual groups, produce variable outcomes of cell isolates, depend on individual model settings (e.g., species, age, coexisting heart conditions), and are usually adjusted for particular experimental conditions. Within the context of the quantitative TATS membrane studies and protocols presented here, an essential level of quality assessment and documentation concerns the confocal or superresolution microscopy of individual cell membrane structures prone to metabolic and isolation protocol dependent changes, both in AM or VM. Importantly, even if high yields of cell isolates suggest healthy intact myocytes, investigators need to document and critically judge each individual cell carefully against morphological criteria of surface and TATS membrane integrity versus non-specific damage due to isolation procedures versus specific changes due to different types of interventions as compared to control conditions. An important variable during cardiac cell isolation is the specific activity of a given collagenase lot. To select a new lot of collagenase, the enzymatic activity of several collagenase samples should be tested against each other by evaluating cardiac myocyte yield and quality, and according to manufacturer instructions. Ideally, a new lot of collagenase is identified with collagenase activity similar to previous successfully used lots (for extended evaluation of possible enzyme activities refer to the &#8220;collagenase lot selection tool&#8221; in the materials and methods table).&#160;Taken together, quantitative approaches of TATS membrane visualization depend critically on cell isolation quality and, vice versa, cardiac cell isolations leading to unspecific membrane damage as documented by TATS microscopy should trigger critical review and correction of the isolation procedures. Since cell isolation quality and TATS membrane visualization and quantification are intrinsically linked, the protocols discussed in this article cover all major aspects as a continuous strategy.
An additional challenge and common issue of cardiac studies, cell damage and/or cell loss occur due to metabolically compromising interventions e.g., following myocardial infarction9, yet need to be judged against potential inadvertent damage e.g., following unnoticed air embolism during cell isolation. Isolation of cardiac myocytes from diseased hearts may lead to additional, significant cell loss and decreased cell yields. Therefore, comparison of the total number of isolated intact cells between control and diseased hearts can be meaningful if cell isolation and counting are consistently applied through standardized protocols. Consequently, it is critical to judge cell integrity through an appropriate control group, which reflects the best possible myocyte cell isolation quality. Importantly, individual cell quality and live cell microscopy of healthy versus diseased versus myocytes inadvertently damaged by the isolation procedure may significantly influence the analysis of TATS membrane networks. The protocols presented here therefore emphasize the integrity and stability of physiological membrane components during cell isolation and live cell microscopy of intact membranes. The entire workflow is designed as a continuous strategy to achieve and preserve intact TATS membrane components while excluding damaged cells, since these will exhibit isolation dependent membrane artifacts like disrupted membrane tubules, membrane blebs, and altered TATS networks erroneously under control conditions and compromise further quantitative analysis. Vice versa, the same strategies are crucial for intervention studies with the potential to disrupt TATS membranes, which depend critically on meaningful comparison controls between true healthy versus true diseased cell with TATS membrane changes.
In addition, we address procedures to achieve the technically much more challenging isolation of AM cells. Despite progress and improved protocols, it is important to emphasize that it is not trivial to reproduce high quality cell isolations of VMs and even less reliable for AMs. This is due to the overall lower yield of AM cells where even small errors or variations during cell isolation may lead to complete failure of AM cell isolation, whereas a mild degree of VM cell damage might be less apparent in cell suspension due to relatively high cell numbers compared to AM. Since AM cells might become curved after isolation, analysis through several ROIs can be advantageous as outlined under step 4.3. Following a detailed procedure of cell isolation steps, we provide a protocol for direct integral membrane staining and confocal or STED superresolution imaging of TATS networks both for VMs and AMs. These protocols enable both quantitative analysis and differentiation of select components of TATS membranes through previously established parameters. As compared to VMs, the 3D organization and functional behaviors of the atrial TATS network in AMs are currently less understood.
The procedures to image TATS membranes in living cells (steps 3.1 to 3.7) were developed with commercial confocal (Table of Materials/Equipment) and custom-made STED fluorescence microscopes9. To optimize the microscope settings for fluorescent image generation and quantitative TATS analysis, the following points are of general importance:
<ul>
	<li>Objective 
	In order to resolve small details of TATS membrane structures, empirically test which objective provides the highest image quality while focusing several micrometer deep in the cell. Certain confocal microscopes may perform better with water or glycerol objectives, in contrast to the 63X 1.4 NA oil objective used here. Objectives with 100X magnification are used for superresolution STED microscopy, trading off a smaller field of view for nanometric resolution34.</li>
	<li>Excitation and Gain 
	The optimal settings of the excitation power and detector gain depend on the microscope light path, laser performance, and sample properties. Ideally, laser power and gain are adjusted to exploit the full range of the detector, yet avoid image saturation. Commercial microscopy software packages usually provide lookup tables that visualize the lower and upper limit of the dynamic range. Furthermore, to minimize dye bleaching employ the lowest possible laser power that still provides sufficient structural TATS membrane details. Also, excitation power should be low enough to avoid cumulative photo damage leading to myocyte contractures and death.</li>
	<li>Pixel Size 
	Use a pixel size compatible with Nyquist sampling, approximately half the resolution achieved with the given settings. For confocal imaging a pixel size of 100 nm x 100 nm is compatible, which will also limit bleaching. For superresolution microscopy significantly smaller pixel sizes are used e.g., 20 nm x 20 nm for STED microscopy9.</li>
	<li>Dwell Time 
	Confocal microscopes provide an averaging function. In general, use the shortest possible pixel dwell time to avoid bleaching in combination with signal averaging e.g., line-averaging &#8805; 8 to improve the signal-to-noise-ratio.</li>
	<li>Document the Applied Microscopy Settings Through Meta-data 
	Once the settings how to image details of TATS membrane structures were optimized on a particular confocal microscope, safe and/or document the settings (protocol meta-data). Acquire all images within a (or between) group(s) of cells with the same objective, excitation power, gain, pixel size, pixel dwell time, and averaging function. Equal imaging conditions allow for direct comparison and quantitation within a (or between) group(s) of cells.</li>
	<li>For general guidance and further details about principles and applications of confocal microscopy refer to the Handbook of Biological Confocal Microscopy (Pawley J.B., 3rd Edition, 2006, Springer Science+Business Media, LLC).</li>
</ul>
In contrast to the direct analysis strategies presented here, previous publications describing TATS membranes and disease related changes, have used regional aggregate readouts of T-tubule density as quantitative strategy16,17, or indirect regional strategies based on Fourier transformation analysis of striated membrane signals in order to assess T-tubule component regularity7. In contrast, the quantitative approaches described here are directly related to individual TATS components and provide a number of additional parameters including membrane network properties and specific components like the percentage&#160;of A-tubules. Furthermore, the TATS network density can be quantified as the normalized length of the entire extracted skeleton per ROI area. The number of triple junctions of three individual, continuously connected tubule components can be used as a measure of the branching complexity of the TATS membrane network. We note that any analysis of smallest TATS components depends on the staining procedures. In our experience, 800 &#181;l of a 50 &#181;M di-8-ANEPPS solution are sufficient to stain complete TATS networks in a cell pellet containing 50,000 VM cells9. However, if the cell pellet contains a lower number of cardiac myocytes, if powerful fluorescence detectors are available, and if confocal imaging of the overall TATS network distribution rather than smallest membrane details and quantitative changes are of interest, lower dye concentrations may be used based on empirical testing. Finally, a software macro written for the described analysis can be used to automate the image processing steps to facilitate analysis of larger datasets, which is particularly useful for comparison between different treatment groups (e.g., drugs), cell types (e.g., AM versus VM), and pathophysiological interventions (e.g., sham versus myocardial infarction).
For image analysis of TATS networks, the following sequence of principle steps is applied: 1) rolling ball background&#8211;subtraction (4.5.1) to remove spatial variations in background intensity; 2) local contrast enhancement (4.5.2.); 3) image smoothing (4.5.3); 4) statistical region merging (4.5.4); 5) defining the threshold of image binarization (4.5.6); and 6) calculation of the skeleton data (4.5.8). A critical step during the skeletonization of fluorescent TATS images is the image binarization shown in Figure 6. The associated thresholding steps ultimately define which true membrane structures are detected to represent the underlying TATS components versus potentially false structures identified by error from background noise. Identification of the correct threshold for binary image analysis should correspond with the true TATS membrane structures, which depends on a sufficiently high signal-to-noise (SNR) ratio each for confocal and superresolution microscopy approaches. Therefore, a sufficient image quality should be established first and subsequently combined with critical judgment of individual cell quality including documentation by bright field images as outlined. Alternative options to adapt the image segmentation protocol for a given microscope data output and/or physiological questions include image deconvolution and other thresholding procedures like &#8220;Otsu&#8221; or &#8220;Iso-data&#8221; available as ImageJ plugins. Regardless of the final segmentation procedure, we consider the comparison between extracted and raw data by image overlay a mandatory quality control step. In summary, morphological and membrane integrity of individual isolated myocytes, sufficient staining of intracellular TATS membranes, parameter optimization for fluorescence imaging, and overlay control of extracted skeleton data will all contribute to the quality of fluorescent TATS images and quantitative results.
If larger species than mouse are used for cell isolation, the protocols can be readily adapted as appropriate. For the next larger species, rat hearts can be cannulated with a blunt 14&#160;G cannula (outer diameter 2.1 mm) and perfused at 8 ml/min. Significantly older or diseased hearts may require even larger cannula sizes. In general, cardiac perfusion may be conducted either by constant pressure e.g., using a 1 m high water column between reservoir and aorta or by constant flow using a peristaltic pump. For cell isolation from small rodent hearts like mice and rats constant flow may be advantageous since collagenase digestion will eventually disrupt coronary resistance vessels leading to excessive perfusion rates from leaking vessel beds which will be controlled to some degree by constant flow protocols. In contrast, constant pressure perfusion is advantageous if monitoring of the flow rate and correct cannulation are a priority, which is advantageous for intervention models with altered blood vessel resistance behaviors as well as for training of the cell isolation procedures.
As outlined above, sufficient cell quality is very important for quantitative studies of endogenous membrane systems. However, during heart perfusion and collagenase digestion numerous factors can critically affect the quality of the cell isolation, which should never be underestimated during protocol optimization or trouble shooting27. In particular, the activity of a given collagenase lot should be determined for the specific tissue of interest e.g., atria or ventricles prior to execution of the experimental bona fide studies to establish isolation conditions to be maintained throughout the remainder of the study. Furthermore, the water quality, pH, temperature, optimization and cleaning of the perfusion setup will minimize the risk of inadvertent damage from contaminants and emboli, and potentially additional factors have to be monitored to establish optimal homeostatic conditions during cell isolation. BDM (2,3-butanedione-monoxime) a reversible inhibitor of myosin ATPase cross-bridges is commonly used during tissue dissection and digestion to sustain cardiac muscle relaxation, which increases the yield of cell isolations. Nevertheless, investigators need to be aware that BDM may exert non-specific phosphatase activities leading to off-target effects e.g., inhibition of Na+/Ca2+ exchange currents under certain conditions33. For some experiments it might be advantageous to replace BDM by blebbistatin as cardioplegic solution, an inhibitor with a high affinity for myosin at micromolar concentrations which is, however, toxic and relatively expensive and may have other off-target effects. Resting healthy cardiomyocytes should not show any contractions in the absence of electrical stimulation and such cells should be excluded from further analysis. On the other hand, cardiac myocyte contraction and relaxation in response to electrical stimulation at physiological extracellular Ca2+ concentrations can be used to establish normal contractile behavior as an additional measure to assess functional cell quality and/or abnormal behavior in heart disease versus healthy control cells.
In summary, the protocols for single cell isolation and quantitative image analysis described here have been successfully applied for confocal and superresolution microscopy of the TATS membrane network in VM9 and AM cells21 as well as for quantitative analysis of microtubule networks in fixed cardiac myocytes (data not shown). These and future applications of the protocols may open avenues for a variety of experimental questions such as the characterization of TATS membranes at different developmental stages or the analysis of membrane associated protein or organelle structures that contact the TATS network to exert highly localized, domain specific signaling functions in AM and VM cells.

In healthy striated muscle cells, tubular membrane structures with &#8220;transverse&#8221; orientations (T-tubules) perpendicular to the main cell axis are abundant. Consequently, T-tubules have been characterized as continuous extensions of the muscle cell main &#8220;lateral&#8221; surface membrane (sarcolemma), which deeply penetrate the cytosol towards the cell center. The physiological role of T-tubules continuous with the outer surface membrane is the rapid electrical coupling of remote intracellular compartments formed by SER organelle contact domains throughout the relatively large cardiac muscle cell volume by nanometric proximity coupling of voltage-activated L-type Ca2+ channels (Cav1.2) inward current&#160;(ICa) activating adjacent ryanodine receptor (RyR2) SER Ca2+ releases. In ventricular myocytes (VMs), the non-continuous membrane contacts (&#8220;junctions&#8221;) between the junctional SER domains and T-tubules are thought to control thousands of individual intracellular Ca2+ release nanodomains in each cell1.
For any given contact domain, the juxtaposed membrane portions each of the T-tubule and the peripheral (junctional) SER are approximately 15 nm close to each other, hence defined as nanodomain. Thereby, very small individual cytoplasmic subspaces are segregated which enable quasi cell-autonomous compartment behaviors. When an incoming action potential activates Cav1.2 channels in the T-tubules of VMs, a relatively small Ca2+ inward current will rapidly increase the subspace Ca2+ concentration [Ca2+]S in the attoliter sized nanodomain1. Next, the [Ca2+]S increase activates Ca2+-gated ryanodine receptors (RyR2) within nanometer proximity in the juxtaposed SER membrane junction, and this coupling process occurs throughout all electrically coupled myocyte nanodomains. RyR2s occur as dense multi-channel clusters with an estimated stoichiometry of 1 Cav1.2 channel for 5-10 RyR2 channels2. Since the SER-to-cytosol [Ca2+] gradient is very steep (ratio 104:1) and RyR2s function as high-conductance Ca2+ release channels in functionally coupled clusters, RyR2 activation results in a quantitatively large Ca2+ release current from T-tubule coupled junctional SER domains increasing local subspace [Ca2+]S to 100 &#181;M or higher within 1-2 msec3,4. This cardiac signal amplification behavior is also referred to as Ca2+ induced Ca2+ release (CICR). Taken together, T-tubules are essential membrane structures that rapidly activate Ca2+ release signals through junctional nanodomain SER contacts and cell-wide CICR during excitation-contraction (EC) coupling.
In addition to T-tubules, axial tubules (A-tubules) with a significantly different orientation parallel to the main (longitudinal) cell axis have been documented by electron microscopy (EM), confocal and 2-photon microscopy studies. For instance, a cell-wide continuous lattice of A-tubules between myofibrils interconnected with T-tubules near sarcomere Z-lines was shown by extracellular tracers and EM imaging of fixed guinea pig VMs5. Using extracellular dextran-linked fluorescein staining and live 2-photon imaging of rat VMs, a complex reticular 3D tubule network was visualized consisting of ~60% T-tubules and ~40% A-tubules6. This study not only led to 3D visualization of abundant A-tubules, but also to the realization that sectioning for EM visualization is inherently limited for the analysis of complex and dynamic membrane networks like the transverse-axial tubule system (TATS). Consequently, confocal live cell imaging of TATS membranes directly stained with di-8-ANNEPS was developed. If live cell TATS networks are analyzed by Fourier transformation, the regular appearance of T-tubule components in space near sarcomere Z-lines is reflected by the ensemble power spectrum from a region of striated signals7. This indirect analysis strategy has been used to detect cell-wide regional changes in TATS component regularity in disease models7. For instance, shRNA mediated junctophilin-2 knock-down caused heart failure and the isoform specific protein deficiency resulted in T-tubule reorganization with nanodomain Ca2+ release dysfunction8. We have recently extended the analysis of TATS membrane networks through direct quantitative approaches and further by live cell superresolution microscopy of individual TATS components in mouse VMs using stimulated emission depletion (STED) nanoscopy9. Nanometric image resolution allowed for direct analysis of smaller individual TATS components, which approximate a 50:50 distribution of transverse versus axial tubule orientations, quantitatively confirming two abundant yet differentially oriented individual TATS components in healthy mouse hearts9. These strategies will be further outlined in the protocol section below.
While the physiological role of the abundant A-tubule components in the adult heart has remained enigmatic, EM studies have documented SER membrane structures associated with A-tubules suggesting endogenous Ca2+ release nanodomains in guinea pig and rat VMs5,10. Confocal analysis of Cav1.2 and RyR2 found a high degree of colocalization at A-tubule junctions10. Since ~20% of spontaneous Ca2+ sparks in rat VMs originated relatively far away from Z-line striations where T-tubules typically occur, one argument has been that A-tubule associated nanodomains may indeed exist and function as Ca2+ release sites11,12. Interestingly, T-tubule formation and maturation occurs only after birth and parallels the growth of cardiac cells, e.g., through sprouting of precursor sarcolemmal invaginations at P5 and immature branched TATS network assemblies at P10 in mice13. It appears that Junctophilin-2 is particularly important for postnatal TATS network maturation since shRNA knock-down prevented the anchoring of T-tubule membranes to SER junctions leading to delayed Ca2+ release and pathological TATS organization consistent with immature A-tubule dominated architectures in VMs13. These observations may ultimately lead to proof-of-concept that T-tubules form through membrane invagination processes whereas A-tubules may morph through additional or even alternative intracellular mechanisms14.
Characterization of TATS membrane changes in heart disease has become an important research area for pathophysiological questions. Initial reports in a dog model of pacing-induced heart failure showed a loss of T-tubules and Cav1.2 current (ICa)15. A pig model of ischemic cardiomyopathy showed reduced T-tubule densities and a reduced synchrony of intracellular Ca2+ release16. Using a spontaneously hypertensive rat (SHR) model of heart failure, a loss of T-tubules was associated with reduced nanodomain coupling of Cav1.2 and RyR2 by the proposed mechanism of &#8220;RyR2 orphaning&#8221;7. A loss of T-tubules has also been shown in human VMs from ischemic, dilated, and hypertrophic cardiomyopathy samples17. Furthermore, an increase in A-tubules was reported in tissue sections of human dilated cardiomyopathy18. Following myocardial infarction, we showed a differential mechanism of TATS reorganization in mouse VMs with significant decreases of T-tubules in contrast to increases of A-tubule components9. Importantly, improved local membrane contrast achieved through live cell superresolution STED microscopy enabled detailed quantitative component analysis through direct measurements, which showed significant proliferation of the A-tubules with overall increases of the TATS network length and branching complexity9. Moreover, it was shown that exercise training may reverse T-tubule remodeling in rats after myocardial infarction19 and that cardiac resynchronization therapy may lead to reverse remodeling of T-tubules in dogs with atrial tachypacing induced heart failure20. Taken together, studies both in diseased human and animal VMs as well as potential therapeutic interventions will arguably benefit from high quality cell isolation procedures and detailed quantitative analysis strategies as outlined in the protocol and results sections below.
Furthermore, as demonstrated recently by lateral surface versus TATS membrane trafficking of KATP channel isoforms 21, it is important to consider atrial myocytes (AMs) as biologically distinct as well as comparative cardiac cell model versus VMs. T-tubules were recently documented in sheep and human AMs22. Current evidence suggests that few T-tubules exist in AM cells and typically in larger mammals like sheep and humans, but not in small rodents23. In contrast to VMs, in AMs intracellular Ca2+ release appears to occur from the cell surface propagating by diffusion towards the cell center which results in marked spatial and temporal Ca2+ gradients23. Within this framework it seems important to elucidate mechanisms of intracellular Ca2+ signaling instability for common disease forms such as atrial fibrillation24. In summary, both AM and VM cell isolation and each for healthy and diseased hearts are commonly employed protocols. Only if cell isolation is done properly as judged by microscopic documentation of sufficient cell quality, AM and VM samples should be carried forward for quantitative TATS analysis. Accordingly, the following protocol sections are critically dependent on high quality cell isolates from mouse or other species followed by live cell microscopy to analyze intact TATS membranes. As pointed out previously, characterization of TATS membranes is a challenging research area with a propensity for fixation and preparation artifacts6, membrane changes due to osmotic alterations, and resolution limitations of conventional light microscopy9. We note that recent state-of-the-art protocols for isolation of human AMs for Ca2+ imaging and patch-clamp and of rat VMs for cell culture have been previously published in this journal25,26.

<table><tbody><tr><td colspan="4">&lt;strong&gt;Chemicals and Enzymes&lt;/strong&gt;</td></tr><tr><td>2,3-Butanedione monoxime</td><td>Sigma-Aldrich, Munich, Germany</td><td>B0753</td><td></td></tr><tr><td>Bovine calf serum</td><td>Thermo Scientific, Schwerte, Germany</td><td>SH30073</td><td>Triple 0.1 &amp;micro;m sterile filtered.</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich, Munich, Germany</td><td>21115</td><td>Diluted 1:10 in MQ water to obtain 100 mM CaCl&lt;sub&gt;2 &lt;/sub&gt;stock concentration.</td></tr><tr><td>Collagenase type II</td><td>Worthington via Cell Systems, Troisdorf, Germany</td><td>on request</td><td>Enzymatic activity depends on individual collagenase batches. Collagenase II and other enzyme activities (Caseinase, Clostripain, Tryptic) can be assessed in the &quot;collagenase lot selection tool&quot;. Determine cell yield and quality individually for each new lot of collagenase.</td></tr><tr><td>Glucose</td><td>Carl Roth, Karlsruhe, Germany</td><td>HN06.1</td><td></td></tr><tr><td>Heparin</td><td>Rotexmedica, Trittau, Germany</td><td>PZN-03862340</td><td>Diluted in 0.9% NaCl and injected subcutaneuosly in abdominal skin.</td></tr><tr><td>HEPES</td><td>Carl Roth, Karlsruhe, Germany</td><td>9105.4</td><td></td></tr><tr><td>Forene 100% (V/V)</td><td>Abbott, Libertyville, IL, USA</td><td>B506</td><td>Active agent: isoflurane, 250 ml. Use approximately 2 Vol% in air/oxygen dispenser instrument.</td></tr><tr><td>KCl</td><td>Carl Roth, Karlsruhe, Germany</td><td>6781.3</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Carl Roth, Karlsruhe, Germany</td><td>3904.2</td><td></td></tr><tr><td>Laminin (2 mg/ml)</td><td>BD Biosciences, Heidelberg, Germany</td><td>354232</td><td>Lamination is described under step 2.1.</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Carl Roth, Karlsruhe, Germany</td><td>2189.2</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Carl Roth, Karlsruhe, Germany</td><td>8283.2</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Carl Roth, Karlsruhe, Germany</td><td>4984.2</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Carl Roth, Karlsruhe, Germany</td><td>HN01.1</td><td></td></tr><tr><td>Taurin</td><td>Carl Roth, Karlsruhe, Germany</td><td>4721.2</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Dyes&lt;/strong&gt;</td></tr><tr><td>Di-8-ANEPPS</td><td>Molecular Probes, Life Technologies, Darmstadt, Germany</td><td>D-3167</td><td>Stock solution 2 mM in DMSO</td></tr><tr><td>Trypan blue</td><td>Sigma-Aldrich, Munich, Germany</td><td>T8154</td><td>Trypan blue is gently mixed 1:1 via tip-cut 1 ml plastic pipette with cell suspension prior to cell counting in Neubauer cytometer.</td></tr><tr><td colspan="4">&lt;strong&gt;Langendorff Perfusion Setup&lt;/strong&gt;</td></tr><tr><td>Circulation thermostat</td><td>Lauda, Lauda-K&amp;ouml;nigshofen, Germany</td><td></td><td>Please refer to Louch &lt;em&gt;et al.&lt;/em&gt; (JMCC 2011). Heat up thermostat und buffers in perfusion tubing to 37 &amp;deg;C 15 min prior to use.</td></tr><tr><td>Flexible silicone tubing Tygon for peristaltic pump</td><td>VWR, Darmstadt, Germany</td><td>224-2252</td><td>Tubing needs to be changed regularly.</td></tr><tr><td>Flexible silicone tubing Tygon for thermostat</td><td>VWR, Darmstadt, Germany</td><td>228-4340</td><td></td></tr><tr><td>Heating coil surroundung perfusion tubing</td><td>Rettberg, G&amp;ouml;ttingen, Germany</td><td>custom-made</td><td>Heating coil and tubing needs to be cleaned thoroughly via MQ water after using. Do not use detergents. Glass components should be bathed regularly&amp;nbsp; in 10 mM NaOH overnight.</td></tr><tr><td>Peristaltic pump</td><td>Ismatec, Wertheim, Germany</td><td>ISM830</td><td></td></tr><tr><td>Three way stop cock Discofix C Luer Lock 10 cm</td><td>Braun, Melsungen, Switzerland</td><td>16500C</td><td></td></tr><tr><td>Three way stop cock Discofix 3SC</td><td>Braun, Melsungen, Switzerland</td><td>4095146</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Instruments&lt;/strong&gt;</td></tr><tr><td>42 mm glass coverslips</td><td>Menzel Gl&amp;auml;ser via Thermo Scientific, Schwerte, Germany</td><td>on request</td><td>0.13-0.16 mm thickness</td></tr><tr><td>Cannula 21 G</td><td>Becton, Dickinson and Company, Franklin Lake, NY, USA</td><td>304432</td><td>Cut to a length of ~5 mm, roughened with sandpaper.</td></tr><tr><td>Coverslips for Neubauer cytometer 24 x 24 mm</td><td>Menzel Gl&amp;auml;ser via Thermo Scientific, Schwerte, Germany</td><td>on request</td><td>0.38-0.42 mm thickness</td></tr><tr><td>Graefe forceps, 0.5 mm tips, slight curve</td><td>Fine Science Tools, Heidelberg, Germany</td><td>11151-10</td><td></td></tr><tr><td>LSM 710 NLO</td><td>Carl Zeiss, Jena, Germany</td><td></td><td>63X 1.4 NA oil objective</td></tr><tr><td>Neubauer improved cytometer</td><td>Labor Optik, Friedrichsdorf, Germany</td><td>1100000</td><td>Counting procedure: Wipe cytometer and coverslip provided with the counting chamber with 70 % ethanol. Press coverslip gently on the counting chamber so that the two glass surfaces are in contact and Newton&#039;s rings can be observed. Subsequently, 10-20 &amp;micro;l cell suspension can be applied to the edge of the coverslip to be sucked into the void by capillary action. Count the intact vs. defect myocytes using the squares of the cytometer grid which reflects 100 nl. Repeat counting procedure on the second grid provided on the cytometer. Calculate the density of cells in your original cell suspension by taking account of any dilutions and counting shortcuts.</td></tr><tr><td>POC-R2 Imaging Chamber</td><td>Pecon, Erbach, Germany</td><td></td><td>Cell suspension volume: 800 &amp;micro;l; desired plating density: ~1,000 AM and ~10,000 VM</td></tr><tr><td>Spring scissors, 8 mm blades straight, blunt</td><td>Fine Science Tools, Heidelberg, Germany</td><td>15025-10</td><td></td></tr><tr><td>Student dumont #7 forceps, inox</td><td>Fine Science Tools, Heidelberg, Germany</td><td>91197-00</td><td></td></tr><tr><td>Student iris scissors, curved, 11.5 cm</td><td>Fine Science Tools, Heidelberg, Germany</td><td>91461-11</td><td></td></tr><tr><td>Student iris scissors, straight, 11.5 cm</td><td>Fine Science Tools, Heidelberg, Germany</td><td>91460-11</td><td></td></tr><tr><td>Student surcigal scissors, straight, sharp, 12 cm</td><td>Fine Science Tools, Heidelberg, Germany</td><td>91402-12</td><td></td></tr><tr><td>Tissue forceps, 1 x 2 teeth, slim, 10 cm</td><td>Fine Science Tools, Heidelberg, Germany</td><td>11023-10</td><td></td></tr></tbody></table>

analysis of tubular membrane networks in cardiac myocytes from atria and ventricles

In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. While the outer surface membrane and associated TATS membrane components appear to be continuous, there are substantial differences in lipid and protein content. In ventricular myocytes (VMs), certain TATS components are highly abundant contributing to rectilinear tubule networks and regular branching 3D architectures. It is thought that peripheral TATS components propagate action potentials from the cell surface to thousands of remote intracellular sarcoendoplasmic reticulum (SER) membrane contact domains, thereby activating intracellular Ca2+ release units (CRUs). In contrast to VMs, the organization and functional role of TATS membranes in atrial myocytes (AMs) is significantly different and much less understood. Taken together, quantitative structural characterization of TATS membrane networks in healthy and diseased myocytes is an essential prerequisite towards better understanding of functional plasticity and pathophysiological reorganization. Here, we present a strategic combination of protocols for direct quantitative analysis of TATS membrane networks in living VMs and AMs. For this, we accompany primary cell isolations of mouse VMs and/or AMs with critical quality control steps and direct membrane staining protocols for fluorescence imaging of TATS membranes. Using an optimized workflow for confocal or superresolution TATS image processing, binarized and skeletonized data are generated for quantitative analysis of the TATS network and its components. Unlike previously published indirect regional aggregate image analysis strategies, our protocols enable direct characterization of specific components and derive complex physiological properties of TATS membrane networks in living myocytes with high throughput and open access software tools. In summary, the combined protocol strategy can be readily applied for quantitative TATS network studies during physiological myocyte adaptation or disease changes, comparison of different cardiac or skeletal muscle cell types, phenotyping of transgenic models, and pharmacological or therapeutic interventions.

Watch this Scientific Journal Video about Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles at JoVE.com

Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles

In cardiac myocytes, tubular membrane structures form intracellular networks. We describe optimized protocols for i) isolation of myocytes from mouse heart including quality control, ii) live cell staining for state-of-the-art fluorescence microscopy, and iii) direct image analysis to quantify the component complexity and the plasticity of intracellular membrane networks.

In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. ...

analysis-tubular-membrane-networks-cardiac-myocytes-from-atria

Research

JoVE Journal

Bioengineering

20.5K Views.  Heart Research Center Goettingen. In cardiac myocytes, tubular membrane structures form intracellular networks. We describe optimized protocols for i) isolation of myocytes from mouse heart including quality control, ii) live cell staining for state-of-the-art fluorescence microscopy, and iii) direct image analysis to quantify the component complexity and the plasticity of intracellular membrane networks.

Video: Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles

Analysis of Cardiomyocyte Development using Immunofluorescence in Embryonic Mouse Heart

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated discs, and costameres requires the coordinated assembly of multiple components in time and space. Disruption in assembly of these components leads to developmental heart defects. Immunofluorescent staining techniques are used commonly in cultured cardiomyocytes to probe myofibril maturation, but this ex vivo approach is limited by the extent to which myocytes will fully differentiate in culture, lack of normal in vivo mechanical inputs, and absence of endocardial cues. Application of immunofluorescence techniques to the study of developing mouse heart is desirable but more technically challenging, and methods often lack sufficient sensitivity and resolution to visualize sarcomeres in the early stages of heart development. Here, we describe a robust and reproducible method to co-immunostain multiple proteins or to co-visualize a fluorescent protein with immunofluorescent staining in the embryonic mouse heart and use this method to analyze developing myofibrils, intercalated discs, and costameres. This method can be further applied to assess cardiomyocyte structural changes caused by mutations that lead to developmental heart defects.

Mutations that lead to congenital heart defects benefit from in vivo investigation of cardiac structure during development, but high-resolution structural studies in the mouse embryonic heart are technically challenging. Here we present a robust immunofluorescence and image analysis method to assess cardiomyocyte-specific structures in the developing mouse heart.

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated ...

Developmental Biology

Isolation and Functional Characterization of Human Ventricular Cardiomyocytes from Fresh Surgical Samples

Cardiomyocytes from diseased hearts are subjected to complex remodeling processes involving changes in cell structure, excitation contraction coupling and membrane ion currents. Those changes are likely&#160;to be responsible for the increased arrhythmogenic risk and the contractile alterations leading to systolic and diastolic dysfunction in cardiac patients. However, most information on the alterations of myocyte function in cardiac diseases has come from animal models.
Here we describe and validate a protocol to isolate viable myocytes from small surgical samples of ventricular myocardium from patients undergoing cardiac surgery operations. The protocol is described in detail. Electrophysiological and intracellular calcium measurements are reported to demonstrate the feasibility of a number of single cell measurements in human ventricular cardiomyocytes obtained with this method.
The protocol reported here can be useful for future investigations of the cellular and molecular basis of functional alterations of the human heart in the presence of different cardiac diseases. Further, this method can be used to identify novel therapeutic targets at cellular level and to test the effectiveness of new compounds on human cardiomyocytes, with direct translational value.

Current knowledge on the cellular basis of cardiac diseases mostly relies on studies on animal models. Here we describe and validate a novel method to obtain single viable cardiomyocytes from small surgical samples of human ventricular myocardium. Human ventricular myocytes can be used for electrophysiological studies and drug testing.

Cardiomyocytes from diseased hearts are subjected to complex remodeling processes involving changes in cell structure, excitation contraction coupling and ...

Medicine

Creating a Structurally Realistic Finite Element Geometric Model of a Cardiomyocyte to Study the Role of Cellular Architecture in Cardiomyocyte Systems Biology

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal microscopy, the scientific community has unprecedented access to large datasets at sub-micrometer resolution that characterize the architectural remodeling that accompanies changes in cardiomyocyte function in health and disease. However, these datasets have been under-utilized for&#160;investigating the role of cellular architecture remodeling in cardiomyocyte function. The purpose of this protocol is to outline how to create an accurate finite element model of a cardiomyocyte using high resolution electron microscopy and confocal microscopy images. A detailed and accurate model of cellular architecture has significant potential to provide new insights into cardiomyocyte biology, more than experiments alone can garner. The power of this method lies in its ability to computationally fuse information from two disparate imaging modalities of cardiomyocyte ultrastructure to develop one unified and detailed model of the cardiomyocyte. This protocol outlines steps to integrate electron tomography and confocal microscopy images of adult male Wistar (name for a specific breed of albino rat) rat cardiomyocytes to develop a half-sarcomere finite element model of the cardiomyocyte. The procedure generates a 3D finite element model that contains an accurate, high-resolution depiction (on the order of ~35 nm) of the distribution of mitochondria, myofibrils and ryanodine receptor clusters that release the necessary calcium for cardiomyocyte contraction from the sarcoplasmic reticular network (SR) into the myofibril and cytosolic compartment. The model generated here as an illustration does not incorporate details of the transverse-tubule architecture or the sarcoplasmic reticular network and is therefore a minimal model of the cardiomyocyte. Nevertheless, the model can already be applied in simulation-based investigations into the role of cell structure in calcium signaling and mitochondrial bioenergetics, which is illustrated and discussed using two case studies that are presented following the detailed protocol.

This protocol outlines a novel method to create a spatially detailed finite element model of the intracellular architecture of cardiomyocytes from electron microscopy and confocal microscopy images. The power of this spatially detailed model is demonstrated using case studies in calcium signaling and bioenergetics.

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal ...

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

The study of electrophysiological properties of cardiac ion channels with the patch-clamp technique and the exploration of cardiac cellular Ca2+ handling abnormalities requires isolated cardiomyocytes. In addition, the possibility to investigate myocytes from patients using these techniques is an invaluable requirement to elucidate the molecular basis of cardiac diseases such as atrial fibrillation (AF).1 Here we describe a method for isolation of human atrial myocytes which are suitable for both patch-clamp studies and simultaneous measurements of intracellular Ca2+ concentrations. First, right atrial appendages obtained from patients undergoing open heart surgery are chopped into small tissue chunks ("chunk method") and washed in Ca2+-free solution. Then the tissue chunks are digested in collagenase and protease containing solutions with 20 &mu;M Ca2+. Thereafter, the isolated myocytes are harvested by filtration and centrifugation of the tissue suspension. Finally, the Ca2+ concentration in the cell storage solution is adjusted stepwise to 0.2 mM. We briefly discuss the meaning of Ca2+ and Ca2+ buffering during the isolation process and also provide representative recordings of action potentials and membrane currents, both together with simultaneous Ca2+ transient measurements, performed in these isolated myocytes.

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

We describe the isolation of human atrial myocytes which can be used for intracellular Ca2+ measurements in combination with electrophysiological patch-clamp studies.

The study of  electrophysiological properties of cardiac ion channels with the patch-clamp  technique and the exploration of cardiac cellular Ca2+ ...

Biology

Isolation and Physiological Analysis of Mouse Cardiomyocytes

Cardiomyocytes, the workhorse cell of the heart, contain exquisitely organized cytoskeletal and contractile elements that generate the contractile force used to pump blood. Individual cardiomyocytes were first isolated over 40 years ago in order to better study the physiology and structure of heart muscle. Techniques have rapidly improved to include enzymatic digestion via coronary perfusion. More recently, analyzing the contractility and calcium flux of isolated myocytes has provided a vital tool in the cellular and sub-cellular analysis of heart failure. Echocardiography and EKGs provide information about the heart at an organ level only. Cardiomyocyte cell culture systems exist, but cells lack physiologically essential structures such as organized sarcomeres and t-tubules required for myocyte function within the heart. In the protocol presented here, cardiomyocytes are isolated via Langendorff perfusion. The heart is removed from the mouse, mounted via the aorta to a cannula, perfused with digestion enzymes, and cells are introduced to increasing calcium concentrations. Edge and sarcomere detection software is used to analyze contractility, and a calcium binding fluorescent dye is used to visualize calcium transients of electrically paced cardiomyocytes; increasing understanding of the role cellular changes play in heart dysfunction. Traditionally used to test drug effects on cardiomyocytes, we employ this system to compare myocytes from WT mice and mice with a mutation that causes dilated cardiomyopathy. This protocol is unique in its comparison of live cells from mice with known heart function and known genetics. Many experimental conditions are reliably compared, including genetic or environmental manipulation, infection, drug treatment, and more. Beyond physiologic data, isolated cardiomyocytes are easily fixed and stained for cytoskeletal elements. Isolating cardiomyocytes via perfusion is an extremely versatile method, useful in studying cellular changes that accompany or lead to heart failure in a variety of experimental conditions.

Individual cardiomyocytes from wild type and mutant mice can be isolated from the heart in order to study their contractility and calcium transients. This allows characterization of the contribution of cellular dysfunction to heart dysfunction from any cause.

Cardiomyocytes, the workhorse cell of the heart, contain exquisitely organized cytoskeletal and contractile elements that generate the contractile force ...

Isolation of Atrial Myocytes from Adult Mice

The electrophysiological properties of atrial myocytes importantly affect overall cardiac function. Alterations in the underlying ionic currents responsible for the action potential can cause pro-arrhythmic substrates that underlie arrhythmias, such as atrial fibrillation, which are highly prevalent in many conditions and disease states. Isolating adult mouse atrial cardiomyocytes for use in patch-clamp experiments has greatly advanced our knowledge and understanding of the cellular electrophysiology in the healthy atrial myocardium and in the setting of atrial pathophysiology. In addition, studies using genetic mouse models have elucidated the role of a vast array of proteins in regulating atrial electrophysiology. Here we provide a detailed protocol for the isolation of cardiomyocytes from the atrial appendages of adult mice using a combination of enzymatic digestion and mechanical dissociation of these tissues. This approach consistently and reliably yields isolated atrial cardiomyocytes that can then be used to characterize cellular electrophysiology by measuring action potentials and ionic currents in patch-clamp experiments under a number of experimental conditions.

This protocol is used to isolate single atrial cardiomyocytes from the adult mouse heart using a chunk digestion approach. This approach is used to isolate right or left atrial myocytes that can be used to characterize atrial myocyte electrophysiology in patch-clamp studies.

The electrophysiological properties of atrial myocytes importantly affect overall cardiac function. Alterations in the underlying ionic currents ...

Modifications of the Langendorff Method for Simultaneous Isolation of Atrial and Ventricular Myocytes from Adult Mice

A single cardiomyocyte is a vital tool in the cellular and subcellular level studies of cardiac biology and diseases as a fundamental unit of contraction and electrical activity. Hence, isolating viable, high-quality cardiomyocytes from the heart is the initial and most crucial experimental step. Comparing the various protocols for isolating the cardiomyocytes of adult mice, the Langendorff retrograde perfusion is the most successful and reproducible method reported in the literature, especially for isolating ventricular myocytes. However, isolating quality atrial myocytes from the perfused heart remains challenging, and few successful isolation reports are available. Solving this complicated problem is extremely important because apart from ventricular disease, atrial disease accounts for a large part of heart diseases. Therefore, further investigations on the cellular level to reveal the mechanisms are warranted. In this paper, a protocol based on the Langendorff retrograde perfusion method is introduced and some modifications in the depth of aorta cannulation and the steps that may affect the digestion process to isolate atrial and ventricular myocytes were simultaneously made. Moreover, the isolated cardiomyocytes are confirmed to be amenable to patch clamp investigation.

This protocol describes modifications of the Langendorff method including the depth of aorta cannulation for simultaneous Isolation of atrial and ventricular myocytes from adult mice.

A single cardiomyocyte is a vital tool in the cellular and subcellular level studies of cardiac biology and diseases as a fundamental unit of contraction ...

Analysis of Cardiac Contractile Dysfunction and Ca2+ Transients in Rodent Myocytes

Contractile dysfunction and Ca2+ transients are often analyzed at the cellular level as part of a comprehensive assessment of cardiac-induced injury and/or remodeling. One approach for assessing these functional alterations utilizes unloaded shortening and Ca2+ transient analyses in primary adult cardiac myocytes. For this approach, adult myocytes are isolated by collagenase digestion, made Ca2+ tolerant, and then adhered to laminin-coated coverslips, followed by electrical pacing in serum-free media. The general protocol utilizes adult rat cardiac myocytes but can be readily adjusted for primary myocytes from other species. Functional alterations in myocytes from injured hearts can be compared to sham myocytes and/or to in vitro therapeutic treatments. The methodology includes the essential elements needed for myocyte pacing, along with the cell chamber and platform components. The detailed protocol for this approach incorporates the steps for measuring unloaded shortening by sarcomere length detection and cellular Ca2+ transients measured with the ratiometric indicator Fura-2 AM, as well as for raw data analysis.

Analysis of Cardiac Contractile Dysfunction and Ca2+ Transients in Rodent Myocytes

A set of protocols are presented that describe the measurement of contractile function via sarcomere length detection along with calcium (Ca2+) transient measurement in isolated rat myocytes. The application of this approach for studies in animal models of heart failure is also included.

Contractile dysfunction and Ca2+ transients are often analyzed at the cellular level as part of a comprehensive assessment of cardiac-induced injury ...

Simultaneous Isolation and Culture of Atrial Myocytes, Ventricular Myocytes, and Non-Myocytes from an Adult Mouse Heart

The isolation and culturing of cardiac myocytes from mice has been essential for furthering the understanding of cardiac physiology and pathophysiology. While isolating myocytes from neonatal mouse hearts is relatively straightforward, myocytes from the adult murine heart are preferred. This is because compared to neonatal cells, adult myocytes more accurately recapitulate cell function as it occurs in the adult heart in vivo. However, it is technically difficult to isolate adult mouse cardiac myocytes in the necessary quantities and viability, which contributes to an experimental impasse. Furthermore, published procedures are specific for the isolation of either atrial or ventricular myocytes at the expense of atrial and ventricular non-myocyte cells. Described here is a detailed method for isolating both atrial and ventricular cardiac myocytes, along with atrial and ventricular non-myocytes, simultaneously from a single mouse heart. Also provided are the details for optimal cell-specific culturing methods, which enhance cell viability and function. This protocol aims not only to expedite the process of adult murine cardiac cell isolation, but also to increase the yield and viability of cells for investigations of atrial and ventricular cardiac cells.

A method is described for the simultaneous isolation of myocytes and non-myocytes from both the atria and ventricles of a single adult mouse heart. This protocol results in consistent yields of highly viable cardiac myocytes and non-myocytes and details optimal cell-specific culture conditions for phenotyping and in vitro analysis.

The isolation and culturing of cardiac myocytes from mice has been essential for furthering the understanding of cardiac physiology and pathophysiology. ...

Analyzing the &alpha;-Actinin Network in Human iPSC-Derived Cardiomyocytes Using Single Molecule Localization Microscopy

The maturation of iPSC-derived cardiomyocytes is a critical issue for their application in regenerative therapy, drug testing and disease modeling. Despite the development of multiple differentiation protocols, the generation of iPSC cardiomyocytes resembling an adult-like phenotype remains challenging. One major aspect of cardiomyocytes maturation involves the formation of a well-organized sarcomere network to ensure high contraction capacity. Here, we present a super resolution-based approach for semi-quantitative analysis of the &#945;-actinin network in cardiomyocytes. Using photoactivated localization microscopy a comparison of sarcomere length and z-disc thickness of iPSC-derived cardiomyocytes and cardiac cells isolated from neonatal tissue was performed. At the same time, we demonstrate the importance of proper imaging conditions to obtain reliable data. Our results show that this method is suitable to quantitatively monitor the structural maturity of cardiac cells with high spatial resolution, enabling the detection of even subtle changes of sarcomere organization.

Analyzing the &#945;-Actinin Network in Human iPSC-Derived Cardiomyocytes Using Single Molecule Localization Microscopy

The formation of a proper sarcomere network is important for the maturation of iPSC-derived cardiomyocytes. We present a super resolution-based approach that allows for the quantitative evaluation of the structural maturation of stem cell derived cardiomyocytes, to improve culture conditions promoting cardiac development.

The maturation of iPSC-derived cardiomyocytes is a critical issue for their application in regenerative therapy, drug testing and disease modeling. ...

Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles

Samenvatting

Meer video's verkennen

Hoofdstukken in deze video