This study was supported by the EU structural Fund (ESF/14-BM-A55-0024/18). In addition, H.L. is supported by the FORUN Program of Rostock University Medical Centre (889001 and 889003) and the Josef and K&#228;the Klinz Foundation (T319/29737/2017). C.I.L. is supported by the Clinician Scientist Program of the Rostock University Medical Center. R.D is supported by the DFG (DA1296/6-1), the DAMP foundation, the German Heart Foundation (F/01/12) and the BMBF (VIP+ 00240).
We thank Madeleine Bartsch for her technical support in iPSC cell culture and cardiac differentiation.

All steps in this protocol involving neonatal and adult mice were performed according to the ethical guidelines for animal care of the Rostock University Medical Centre.
1. Cultivation and dissociation of iPSC-derived cardiomyocytes
<ol>
	<li>Differentiate hiPSC-CMs for 25 days using a 2D monolayer method as described previously17.</li>
	<li>Prewarm dissociation medium to 37 &#176;C and support medium to room temperature.</li>
	<li>Wash cells twice with PBS.</li>
	<li>Add prewarmed dissociation medium to cells and incubate for 12 min at 37 &#176;C and 5% CO2.</li>
	<li>Add support medium to the cells. 
	NOTE: The volume of added support medium needs to be twice the volume of dissociation medium used in step 1.4.</li>
	<li>Dislodge the cells using a 5 mL serological pipette.</li>
	<li>Centrifuge cells at 200 x g for 5 min and resuspend the pellet in 3 mL of hiPSC-CM culture medium17.</li>
	<li>Seed cells in 8 well glass bottom chambers at a cell density of 75,000 cells/well and culture for 3 days.</li>
</ol>
2. Adult cardiomyocyte isolation
<ol>
	<li>Isolation and cultivation of adult cardiomyocytes from NMRI mice was performed as reported previously18.</li>
	<li>Seed cells in 8 well glass chamber slide and culture for one day.</li>
</ol>
3. Isolation and cultivation of neonatal cardiomyocytes
<ol>
	<li>Isolation procedure of neonatal cardiomyocytes, obtained from NMRI mice, was performed as described previously19.</li>
	<li>Seed isolated cell in 8 well glass chamber slide at a cell density of 75,000 cells/well and culture for 3 days in neonatal CM culture medium19.</li>
</ol>
4. Immunofluorescence labeling of the &#945;-actinin network
NOTE: For optimal results, cells are cultured in 8 well glass bottom chambers. Labeling should be performed one day before imaging.
<ol>
	<li>Prewarm 4% PFA at 37 &#176;C.</li>
	<li>Fix iPSC-CM by adding 4% paraformaldehyde directly into the culture media (1:1 dilution) and incubate at 37 &#176;C for 15 min. The final concentration of PFA for fixation is 2%.</li>
	<li>Incubate fixed cells in 0.2% Triton-X, diluted in PBS, for 5 min at room temperature.</li>
	<li>Wash cells twice with PBS, 5 min each.</li>
	<li>Add 1% BSA solution (diluted in PBS) and incubate for 60 min at room temperature.</li>
	<li>Prepare 150 &#181;L of primary antibody solution by diluting &#945;-actinin antibody 1:100 in 1% BSA, containing 0.05% Triton-X. Add to cells and incubate at room temperature for 60 min.</li>
	<li>Wash cells twice with 0.2% BSA solution, 5 min each.</li>
	<li>Prepare 150 &#181;L of secondary antibody solution by diluting goat-anti-mouse Alexa647 antibody 1:100 in 1% BSA, containing 0.05% Triton-X. Add to the cells and incubate at room temperature for 40 min.</li>
	<li>Wash cells twice with 0.2% BSA solution, 5 min each.</li>
	<li>Wash cells twice with PBS, 5 min each. Keep labeled cells in the dark at 4 &#176;C until PALM imaging.</li>
</ol>
5. Preparation of the PALM imaging buffer
NOTE: It is critical to freshly prepare the PALM imaging buffer for each experiment.
<ol>
	<li>Prepare 50% glucose solution by dissolving 25 g glucose in 50 mL of distilled water.</li>
	<li>Prepare basic buffer containing 50 mM Tris-HCl, 10% glucose and 10 mM sodium chloride.
	<ol>
		<li>Adjust the pH level to ~8.0 using hydrochloric acid.</li>
	</ol>
	</li>
	<li>Prepare pyranose oxidase solution by dissolving 0.6 mg pyranose oxidase in 316 &#181;L of basic buffer.</li>
	<li>Prepare catalase solution by dissolving 7 mg of catalase in 500 &#181;L of basic buffer. Mix thoroughly and centrifuge at 10,000 x g for 3 min. Keep the supernatant for further use. Catalase solution can be kept at 4 &#176;C for several days.</li>
	<li>Prepare 500 &#181;L of PALM imaging buffer by mixing 316 &#181;L of pyranose oxidase solution, 25 &#181;L of catalase solution, 100 &#181;L of 50% glucose solution, 50 &#181;L of cysteamine, 5 &#181;L of cyclooctatetraene and 3.5 &#181;L of &#946;-Mercaptoethanol. The final catalytic activity of pyranose oxidase and catalase need to be 7.5 U, 35,00U respectively. 
	NOTE: The PALM imaging buffer provides optimal imaging conditions for 3-5 h. If blinking capability of the fluorescent dye decreases, prepare a new buffer aliquot.</li>
</ol>
6. PALM image acquisition
<ol>
	<li>Switch on the microscope at least 3 h before use and bring the sample to room temperature before imaging to allow thermal equilibration. If the microscope is equipped with an incubation chamber, adjust temperature to 30 &#176;C.</li>
	<li>Clean the objective and bottom of the chamber slide using an appropriate cleaning solvent.</li>
	<li>Add 300 &#181;L of PALM imaging buffer into one well of labeled cells and insert the chamber slide into the stage holder of the microscope.</li>
	<li>Set the PALM image acquisition parameters.
	<ol>
		<li>Select 1.57 N.A. 100x oil objective for acquisition.</li>
		<li>Select PALM mode and activate the TIRF settings.</li>
		<li>Adjust the number of frames to be acquired. Usually 5, 000-10, 000 frames are sufficient to obtain optimal results. However, the number of acquired frames strongly depends on the labeling efficiency and the blinking capability of the fluorescent dye and may be adjusted by the user.</li>
		<li>Set UV laser power to 0.1% and 647 laser to 0.2%.</li>
		<li>Set the gain level to 50-100.</li>
		<li>Switch on the laser illumination and select a target cell. The gain level can be increased if signal intensity is to low (depending on fluorescence labeling).</li>
		<li>Switch off the laser illumination</li>
	</ol>
	</li>
	<li>PALM image acquisition
	<ol>
		<li>Reduce gain to 0 and increase the laser power (ex 647) to 100%.</li>
		<li>Bleach the target cell for ~5 s.</li>
		<li>Increase gain to 50 and start PALM image acquisition. 
		NOTE: Gain needs to be adjusted to get sufficient signal intensity while oversaturated pixels should be avoided. If signal intensity reduces, increase the gain level.</li>
	</ol>
	</li>
	<li>Optionally, steadily enhance the UV laser power (0.1%-10%) to increase signal intensity and to promote blinking of the fluorophore.</li>
</ol>
7. Reconstruction of PALM data
<ol>
	<li>Open the Image J software and import PALM data.</li>
	<li>Open Thunderstorm Plugin and &#8220;Run analysis&#8221;
	<ol>
		<li>In the &#8220;Camera setup&#8221; menu enter pixel size and EM gain. 
		NOTE: When using 100x objectives and 1.6x magnification lens, 100 nm pixel size is appropriate. However, as the pixel size depends on the hardware features of the microscope and camera used for PALM imaging, users need to carefully check and adapt this parameter. EM gain values can be obtained from the metadata.</li>
		<li>In the &#8220;Run analysis menu&#8221; set parameters as follows: B-spline order: 3, B-spline scale: 2.0, peak intensity threshold: stf(Wave.F1), fitting radius: 3, initial sigma: 1.6, magnification: 5.0, update frequency: 50, lateral shifts: 2. Confirm by clicking the &#8220;Ok&#8221; button.</li>
	</ol>
	</li>
	<li>Post processing of reconstructed PALM image
	<ol>
		<li>In the &#8220;Plot histogram&#8221; menu select the &#8220;Sigma&#8221; parameter.</li>
		<li>Use the &#8220;Rectangle&#8221; tool to select a ROI, excluding possible artefacts and apply ROI to the filter. ROI values will appear in the filter command box.</li>
		<li>Add &#8220;&#38; uncertainty &#60;25&#8221; to the ROI values. A possible filter command will look like this: &#8220;(sigma &#62; 48.6821 &#38; sigma &#60; 1117.40) &#38; uncertainty &#60;25&#8221;. Apply selected sigma values.</li>
		<li>In the &#8220;Remove duplicates&#8221; menu, enter a distance threshold of &#8220;10 nm&#8221; and apply.</li>
		<li>In the &#8220;Merging menu&#8221;, set maximum distance to &#8220;20&#8221;, maximum frames per molecule to &#8220;0&#8221; and maximum off frames to &#8220;1&#8221;. Apply settings.</li>
		<li>In the &#8220;Drift correction menu&#8221;, select cross correlation and set &#8220;Number of bins&#8221; to &#8220;5&#8221; and &#8220;Magnification&#8221; to &#8220;5.0&#8221;. Apply drift correction settings.</li>
		<li>Save the final PALM image and export post processed data if desired.</li>
	</ol>
	</li>
</ol>
8. Analysis of sarcomere filaments
<ol>
	<li>Analysis of sarcomere length
	<ol>
		<li>Open Image J software and import the reconstructed PALM image.</li>
		<li>Draw a line between selected sarcomere structures perpendicular to the z-disc to measure the shortest distance between actinin filaments.</li>
		<li>Select &#8220;Plot profile&#8221; in the &#8220;Analyze&#8221; menu and acquire the length between two peaks. As sarcomere length may vary within one cell, a minimum of 20 sarcomeres should be measured in different areas of the target cell.</li>
	</ol>
	</li>
	<li>Analysis of z-Disc thickness
	<ol>
		<li>Open Image J software and import the reconstructed PALM image.</li>
		<li>Convert the reconstructed PALM image into an 8-bit mode image.</li>
		<li>Open the ridge detection plugin and enter the following parameters: line width: 20, high contrast: 230, low contrast: 10, sigma: 0.79, lower threshold: 25.84, minimum line length: 20.</li>
		<li>Set &#8220;Estimate width&#8221;, &#8220;Extend line&#8221; and &#8220;Display results&#8221;.</li>
		<li>Click &#8220;Ok&#8221; and use &#8220;Mean line width&#8221; from results table for further analyses.</li>
	</ol>
	</li>
</ol>

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<li>Venkataramani, V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Enhanced+labeling+density+and+whole-cell+3D+dSTORM+imaging+by+repetitive+labeling+of+target+proteins%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Enhanced labeling density and whole-cell 3D dSTORM imaging by repetitive labeling of target proteins.</a> Scientific Reports. 8 (1), 1-7 (2018).</li>
<li>Erdélyi, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Origin+and+compensation+of+imaging+artefacts+in+localization-based+super-resolution+microscopy%5BTitle%5D)+AND+%22Methods%22%5BJournal%5D)">Origin and compensation of imaging artefacts in localization-based super-resolution microscopy.</a> Methods. 88, 122-132 (2015).</li>
<li>McGorty, R., Schnitzbauer, J., Zhang, W., Huang, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Correction+of+depth-dependent+aberrations+in+3D+single-molecule+localization+and+super-resolution+microscopy%5BTitle%5D)+AND+%22Optics+Letters%22%5BJournal%5D)">Correction of depth-dependent aberrations in 3D single-molecule localization and super-resolution microscopy.</a> Optics Letters. 39 (2), 275(2014).</li>
<li>Ribeiro, M. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+cardiomyocyte+show+of+force%3A+A+fluorescent+alpha-actinin+reporter+line+sheds+light+on+human+cardiomyocyte+contractility+versus+substrate+stiffness%5BTitle%5D)+AND+%22Journal+of+Molecular+and+Cellular+Cardiology%22%5BJournal%5D)">A cardiomyocyte show of force: A fluorescent alpha-actinin reporter line sheds light on human cardiomyocyte contractility versus substrate stiffness.</a> Journal of Molecular and Cellular Cardiology. 141, 54-64 (2020).</li>
<li>Pioner, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Isolation+and+mechanical+measurements+of+myofibrils+from+human+induced+pluripotent+stem+cell-derived+cardiomyocytes%5BTitle%5D)+AND+%22Stem+Cell+Reports%22%5BJournal%5D)">Isolation and mechanical measurements of myofibrils from human induced pluripotent stem cell-derived cardiomyocytes.</a> Stem Cell Reports. 6 (6), 885-896 (2016).</li>
</ol>

The authors declare no conflict of interest.

The generation of functional iPSC-derived CM in vitro is important for regenerative therapies, disease modeling and the development of drug-screening platforms. However, insufficient maturity of these CM is a major obstacle in cardiovascular research20. In this regard, high resolution imaging techniques are needed that enable monitoring of the structural maturation state of iPSC-derived CM. At the same time, super resolution microscopy can be a valuable tool to precisely analyze the function of sp...

Cardiovascular diseases (CVD) such as myocardial infarction or cardiomyopathy remain the major cause of death in the western world1. As the human heart possesses only poor regenerative capacity, there is a need for strategies to promote the recovery from CVDs. This includes cell replacement therapies to replenish lost cardiomyocytes (CM), as well as the development of new anti-arrhythmic drugs for efficient and safe pharmaceutical intervention. Induced pluripotent stem cell (iPSC) have been shown to be a promising cell source for the unlimited generation of human CM in vitro, suitable for regenerative therapies, disease modeling, and for the development of drug screening assays2,3,4.
Although many different cardiac differentiation protocols exist, iPSC-derived CM still lack certain phenotypical and functional aspects that impede the in vitro and in vivo application5,6. Beside electrophysiologic, metabolic, and molecular changes, the cardiac maturation process involves the structural organization of sarcomeres, which are the fundamental units required for force generation and cell contraction7. While adult CMs exhibit a well-organized contractile apparatus, iPSC-derived CMs commonly demonstrate disarranged sarcomere filaments, associated with a reduced contraction ability and altered contraction dynamics8,9. In contrast to mature CM that show uniaxial contraction pattern, the disoriented structures in immature CM results in a radial contraction of the whole cell or promote the appearance of contraction focal points9,10.
For improving cardiac maturation, multiple approaches have been applied, including 3D cell culture methods, electrical and mechanical stimulation, as well as the use of extracellular matrices mimicking in vivo conditions11,12,13. To evaluate the success and efficiency of these different culture conditions, techniques are needed to monitor and estimate the degree of the structural maturation of iPSC CM, e.g., by microscopic techniques. In contrast to conventional confocal imaging, the resolution in case of photoactivated localization microscopy (PALM) is approximately 10x higher. This technique in turn allows for a more accurate analysis, detecting even subtle alterations of cellular structures14. Considering the high resolution of PALM-based imaging, the overall goal of this method was the microscopic evaluation of sarcomere maturity in iPSC-derived CMs by precise determination of z-Disc thickness and sarcomere length. In previous studies, these structural features have been shown to be appropriate parameters to assess cardiac maturity15. For example, diseased iPSC-CM lacking full length dystrophin exhibit reduced sarcomere length and z-band width when compared to wild type cells16. Likewise, the length of individual sarcomeres was measured to investigate the impact of topographic cues on cardiac development16. Hence, we applied this approach to evaluate the structural maturation of the sarcomere network in iPSC-CM by quantitatively measuring the sarcomere length and z-disc thickness.

<table><tbody><tr><td>human iPSC cell line</td><td>Takara</td><td>Y00325</td><td></td></tr><tr><td>&amp;micro;-Slide 8 Well Glass Bottom</td><td>ibidi</td><td>80827</td><td></td></tr><tr><td>0.5ml eppendorf tube</td><td>Eppendorf</td><td>30121023</td><td></td></tr><tr><td>Bovine serum albumin</td><td>Sigma Aldrich</td><td>A906</td><td></td></tr><tr><td>Cardiomyocyte Dissociation Kit</td><td>Stem Cell Technologies</td><td>05025</td><td></td></tr><tr><td>Catalase</td><td>Sigma Aldrich</td><td>C40-1G</td><td></td></tr><tr><td>Cyclooctatetraene</td><td>Sigma Aldrich</td><td>138924-1G</td><td></td></tr><tr><td>Cysteamine</td><td>Sigma Aldrich</td><td>30070-10g</td><td></td></tr><tr><td>Dulbecco&#039;s phosphate-buffered saline without Ca2+ and Mg2+</td><td>Thermo Fisher</td><td>14190169</td><td></td></tr><tr><td>F(ab&#039;)2-Goat anti-Mouse IgG Alexa Fluor 647</td><td>Thermo Fisher</td><td>A-21237</td><td></td></tr><tr><td>Fiji image processing software (Image J)</td><td></td><td></td><td></td></tr><tr><td>Glucose</td><td>Carl Roth</td><td>X997.2</td><td></td></tr><tr><td>Hydrochloric acid</td><td>Sigma Aldrich</td><td>H1758</td><td></td></tr><tr><td>LSM 780 ELYRA PS.1 system</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Paraformaldehyde</td><td>Merck</td><td>8187150100</td><td></td></tr><tr><td>Pyranose oxidase</td><td>Sigma Aldrich</td><td>P4234-250UN</td><td></td></tr><tr><td>sarcomeric &amp;alpha;-actinin antibody [EA-53]</td><td>Abcam</td><td>ab9465</td><td></td></tr><tr><td>Sodium chloride</td><td>Sigma Aldrich</td><td>S7653</td><td></td></tr><tr><td>sterile water</td><td>Carl Roth</td><td>3255.1</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>X100</td><td></td></tr><tr><td>Trizma base</td><td>Sigma Aldrich</td><td>T1503</td><td></td></tr><tr><td>&amp;beta;-Mercaptoethanol</td><td>Sigma Aldrich</td><td>63689</td><td></td></tr></tbody></table>

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.

For estimating the degree of structural maturation of CM, neonatal, fully mature adult, and iPSC CM were initially labeled the CM with &#945;-actinin antibody to visualize the sarcomere network. Following PALM acquisition, images were reconstructed, and z-disc thickness was measured using plugin-based image processing software for the automatic detection of the width of individual filaments. Sarcomere length was calculated by measuring the distance between two adjacent intensity peaks, corresponding to neighboring filame...

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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.

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. ...

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Biology

6.7K Views.  Rostock University Medical Center. Deze methode kan worden gebruikt voor de kwantitatieve evaluatie van sarcomere rijping in iPSC-afgeleide cardiomyocyten. Met behulp van deze superresolutie-gebaseerde aanpak, is het mogelijk om zelfs subtiele veranderingen in sarcomere organisatie, die niet mogelijk is met conventionele confocale beeldvorming te detecteren. Schakel ten minste drie uur voor gebruik de microscoop in en breng het monster op kamertemperatuur.Wanneer de microscoop klaar is, voeg 300 microliter palm imaging ...

Video: Analyzing the α-Actinin Network in Human iPSC-Derived Cardiomyocytes Using Single Molecule Localization Microscopy

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

Visualization of Cell Cycle Variations and Determination of Nucleation in Postnatal Cardiomyocytes

Cardiomyocytes are prone to variations of the cell cycle, such as endoreduplication (continuing rounds of DNA synthesis without karyokinesis and cytokinesis) and acytokinetic mitosis (karyokinesis but no cytokinesis). Such atypical cell cycle variations result in polyploid and multinucleated cells rather than in cell division. Therefore, to determine cardiac turnover and regeneration, it is of crucial importance to correctly identify cardiomyocyte nuclei, the number of nuclei per cell, and their cell cycle status. This is especially true for the use of nuclear markers for identifying cell cycle activity, such as thymidine analogues Ki-67, PCNA, or pHH3. Here, we present methods for recognizing cardiomyocytes and their nuclearity and for determining their cell cycle activity. We use two published transgenic systems: the Myh6-H2B-mCh transgenic mouse line, for the unequivocal identification of cardiomyocyte nuclei, and the CAG-eGFP-anillin mouse line, for distinguishing cell division from cell cycle variations. Combined together, these two systems ease the study of cardiac regeneration and plasticity.

To distinguish cell division from cell cycle variations in cardiomyocytes, we present protocols using two transgenic mouse lines: Myh6-H2B-mCh transgenic mice, for the unequivocal identification of cardiomyocyte nuclei, and CAG-eGFP-anillin mice, for distinguishing cell division from cell cycle variations.

Cardiomyocytes are prone to variations of the cell cycle, such as endoreduplication (continuing rounds of DNA synthesis without karyokinesis and ...

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.

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. ...

Bioengineering

Fabrication of 3D Cardiac Microtissue Arrays using Human iPSC-Derived Cardiomyocytes, Cardiac Fibroblasts, and Endothelial Cells

Generation of human cardiomyocytes (CMs), cardiac fibroblasts (CFs), and endothelial cells (ECs) from induced pluripotent stem cells (iPSCs) has provided a unique opportunity to study the complex interplay among different cardiovascular cell types that drives tissue development and disease. In the area of cardiac tissue models, several sophisticated three-dimensional (3D) approaches use induced pluripotent stem cell-derived cardiomyocytes (iPSC-CMs) to mimic physiological relevance and native tissue environment with a combination of extracellular matrices and crosslinkers. However, these systems are complex to fabricate without microfabrication expertise and require several weeks to self-assemble. Most importantly, many of these systems lack vascular cells and cardiac fibroblasts that make up over 60% of the nonmyocytes in the human heart. Here we describe the derivation of all three cardiac cell types from iPSCs to fabricate cardiac microtissues. This facile replica molding technique allows cardiac microtissue culture in standard multi-well cell culture plates for several weeks. The platform allows user-defined control over microtissue sizes based on initial seeding density and requires less than 3 days for self-assembly to achieve observable cardiac microtissue contractions. Furthermore, the cardiac microtissues can be easily digested while maintaining high cell viability for single-cell interrogation with the use of flow cytometry and single-cell RNA sequencing (scRNA-seq). We envision that this in vitro model of cardiac microtissues will help accelerate validation studies in drug discovery and disease modeling.

Here, we describe an easy-to-use methodology to generate 3D self-assembled cardiac microtissue arrays composed of pre-differentiated human-induced pluripotent stem cell-derived cardiomyocytes, cardiac fibroblasts, and endothelial cells. This user-friendly and low cell requiring technique to generate cardiac microtissues can be implemented for disease modeling and early stages of drug development.

Generation of human cardiomyocytes (CMs), cardiac fibroblasts (CFs), and endothelial cells (ECs) from induced pluripotent stem cells (iPSCs) has provided ...

Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically carried out in low throughput. Rapid and ongoing expansion of induced pluripotent stem cell (iPSC) technology presents a new standard in cardiovascular research and alternate methods are now necessary to increase throughput of electrophysiological data at a single cell level. VF2.1Cl is a recently derived voltage sensitive dye which provides a rapid single channel, high magnitude response to fluctuations in membrane potential. It possesses kinetics superior to those of other existing voltage indicators and makes available functional data equivalent to that of traditional microelectrode techniques. Here, we demonstrate simplified, non-invasive action potential characterization in externally paced human iPSC derived cardiomyocytes using a modular and highly affordable photometry system.

Here we describe optical acquisition and characterization of action potentials from induced pluripotent stem cell derived cardiomyocytes using a high-speed modular photometry system.

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically ...

Hybrid Cell Analysis System to Assess Structural and Contractile Changes of Human iPSC-Derived Cardiomyocytes for Preclinical Cardiac Risk Evaluation

Cardiac contractility assessment is of immense importance for the development of new therapeutics and their safe transition into clinical stages. While human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) hold promise to serve as a human-relevant model in preclinical phases of drug discovery and safety pharmacology, their maturity is still controversial in the scientific community and under constant development. We present a hybrid contractility and impedance/extracellular field potential (EFP) technology, adding significant pro-maturation features to an industry-standard 96-well platform.
The impedance/EFP system monitors cellular functionality in real-time. Besides the beat rate of contractile cells, the electrical impedance spectroscopy readouts detect compound-induced morphological changes like cell density and integrity of the cellular monolayer. In the other component of the hybrid cell analysis system, the cells are cultured on bio-compliant membranes that mimic the mechanical environment of real heart tissue. This physiological environment supports the maturation of hiPSC-CMs in vitro, leading to more adult-like contractile responses including positive inotropic effects after treatment with isoproterenol, S-Bay K8644, or omecamtiv mecarbil. Parameters such as the amplitude of contraction force (mN/mm2) and beat duration also reveal downstream effects of compounds with influence on electrophysiological properties and calcium handling.
The hybrid system provides the ideal tool for holistic cell analysis, allowing preclinical cardiac risk assessment beyond the current perspectives of human-relevant cell-based assays.

The analysis of changes in contractile function and cellular integrity of human iPSC-derived cardiomyocytes is of immense importance for nonclinical drug development. A hybrid 96-well cell analysis system addresses both parameters in a real-time and physiological manner for reliable, human-relevant results, necessary for a safe transition into clinical stages.

Cardiac contractility assessment is of immense importance for the development of new therapeutics and their safe transition into clinical stages. While ...

Generation of Human Cardiomyocytes: A Differentiation Protocol from Feeder-free Human Induced Pluripotent Stem Cells

In order to investigate the events driving heart development and to determine the molecular mechanisms leading to myocardial diseases in humans, it is essential first to generate functional human cardiomyocytes (CMs). The use of these cells in drug discovery and toxicology studies would also be highly beneficial, allowing new pharmacological molecules for the treatment of cardiac disorders to be validated pre-clinically on cells of human origin. Of the possible sources of CMs, induced pluripotent stem (iPS) cells are among the most promising, as they can be derived directly from readily accessible patient tissue and possess an intrinsic capacity to give rise to all cell types of the body 1. Several methods have been proposed for differentiating iPS cells into CMs, ranging from the classical embryoid bodies (EBs) aggregation approach to chemically defined protocols 2,3. In this article we propose an EBs-based protocol and show how this method can be employed to efficiently generate functional CM-like cells from feeder-free iPS cells.

Pluripotent stem cells, either embryonic or induced pluripotent stem (iPS) cells, constitute a valuable source of human differentiated cells, including cardiomyocytes. Here, we will focus on cardiac induction of iPS cells, showing how to use them to obtain functional human cardiomyocytes through an embryoid bodies-based protocol.

In order to investigate the events driving heart development and to determine the molecular mechanisms leading to myocardial diseases in humans, it is ...

Model of Ischemic Heart Disease and Video-Based Comparison of Cardiomyocyte Contraction Using hiPSC-Derived Cardiomyocytes

Ischemic heart disease is a significant cause of death worldwide. It has therefore been the subject of a tremendous amount of research, often with small-animal models such as rodents. However, the physiology of the human heart differs significantly from that of the rodent heart, underscoring the need for clinically relevant models to study heart disease. Here, we present a protocol to model ischemic heart disease using cardiomyocytes differentiated from human induced pluripotent stem cells (hiPS-CMs) and to quantify the damage and functional impairment of the ischemic cardiomyocytes. Exposure to 2% oxygen without glucose and serum increases the percentage of injured cells, which is indicated by staining of the nucleus with propidium iodide, and decreases cellular viability. These conditions also decrease the contractility of hiPS-CMs as confirmed by displacement vector field analysis of microscopic video images. This protocol may furthermore provide a convenient method for personalized drug screening by facilitating the use of hiPS cells from individual patients. Therefore, this model of ischemic heart disease, based on iPS-CMs of human origin, can provide a useful platform for drug screening and further research on ischemic heart disease.

We present a model of ischemic heart disease using cardiomyocytes derived from human induced pluripotent stem cells, together with a method for quantitative evaluation of tissue damage caused by ischemia. This model can provide a useful platform for drug screening and further research on ischemic heart disease.

Ischemic heart disease is a significant cause of death worldwide. It has therefore been the subject of a tremendous amount of research, often with ...

Medicine

Sarcomere Shortening of Pluripotent Stem Cell-Derived Cardiomyocytes using Fluorescent-Tagged Sarcomere Proteins.

Pluripotent stem cell-derived cardiomyocytes (PSC-CMs) can be produced from both embryonic and induced pluripotent stem (ES/iPS) cells. These cells provide promising sources for cardiac disease modeling. For cardiomyopathies, sarcomere shortening is one of the standard physiological assessments that are used with adult cardiomyocytes to examine their disease phenotypes. However, the available methods are not appropriate to assess the contractility of PSC-CMs, as these cells have underdeveloped sarcomeres that are invisible under phase-contrast microscopy. To address this issue and to perform sarcomere shortening with PSC-CMs, fluorescent-tagged sarcomere proteins and fluorescent live-imaging were used. Thin Z-lines and an M-line reside at both ends and the center of a sarcomere, respectively. Z-line proteins &#8212;&#160;&#945;-Actinin (ACTN2), Telethonin (TCAP), and actin-associated LIM protein (PDLIM3) &#8212; and one M-line protein &#8212; Myomesin-2 (Myom2) &#8212;&#160;were tagged with fluorescent proteins. These tagged proteins can be expressed from endogenous alleles as knock-ins or from adeno-associated viruses (AAVs). Here, we introduce the methods to differentiate mouse and human pluripotent stem cells to cardiomyocytes, to produce AAVs, and to perform and analyze live-imaging. We also describe the methods for producing polydimethylsiloxane (PDMS) stamps for a patterned culture of PSC-CMs, which facilitates the analysis of sarcomere shortening with fluorescent-tagged proteins. To assess sarcomere shortening, time-lapse images of the beating cells were recorded at a high framerate (50-100 frames per second) under electrical stimulation (0.5-1 Hz). To analyze sarcomere length over the course of cell contraction, the recorded time-lapse images were subjected to SarcOptiM, a plug-in for ImageJ/Fiji. Our strategy provides a simple platform for investigating cardiac disease phenotypes in PSC-CMs.

This method can be used to examine sarcomere shortening using pluripotent stem cell-derived cardiomyocytes with fluorescent-tagged sarcomere proteins.

Pluripotent stem cell-derived cardiomyocytes (PSC-CMs) can be produced from both embryonic and induced pluripotent stem (ES/iPS) cells. These cells ...

Subtype-specific Optical Action Potential Recordings in Human Induced Pluripotent Stem Cell-derived Ventricular Cardiomyocytes

Cardiomyocytes generated from human induced pluripotent stem cells (iPSC-CMs) are an emerging tool in cardiovascular research. Rather than being a homogenous population of cells, the iPSC-CMs generated by current differentiation protocols represent a mixture of cells with ventricular-, atrial-, and nodal-like phenotypes, which complicates phenotypic analyses. Here, a method to optically record action potentials specifically from ventricular-like iPSC-CMs is presented. This is achieved by lentiviral transduction with a construct in which a genetically-encoded voltage indicator is under the control of a ventricular-specific promoter element. When iPSC-CMs are transduced with this construct, the voltage sensor is expressed exclusively in ventricular-like cells, enabling subtype-specific optical membrane potential recordings using time-lapse fluorescence microscopy.

Here we present a method to optically image action potentials, specifically in ventricular-like induced pluripotent stem cell-derived cardiomyocytes. The method is based on the promoter-driven expression of a voltage-sensitive fluorescent protein.

Analyzing the α-Actinin Network in Human iPSC-Derived Cardiomyocytes Using Single Molecule Localization Microscopy

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