The authors thank Portuguese Foundation for Science and Technology (FCT), European Union, Quadro de Refer&#234;ncia Estrat&#233;gico Nacional (QREN), Fundo Europeu de Desenvolvimento Regional (FEDER) and Programa Operacional Factores de Competitividade (COMPETE) for funding UnIC (UID/IC/00051/2013) research unit. This project is supported by FEDER through COMPETE 2020 - Programa Operacional Competitividade E Internacionaliza&#231;&#227;o (POCI), the project DOCNET (NORTE-01-0145-FEDER-000003), supported by Norte Portugal regional operational programme (NORTE 2020), under the Portugal 2020 partnership agreement, through the European Regional Development Fund (ERDF), the project NETDIAMOND (POCI-01-0145-FEDER-016385), supported by European Structural And Investment Funds, Lisbon&#39;s regional operational program 2020. Patr&#237;cia Rodrigues was funded by FCT (SFRH/BD/96026/2013) and Jo&#227;o Almeida-Coelho was by Universidade do Porto/FMUP and FSE-Fundo Social Europeu, NORTE 2020-Programa Operacional Regional do Norte, (NORTE-08-5369-FSE-000024-Programas Doutorais).

All animal experiments comply with the Guide for the Care and Use of Laboratory Animals (NIH Publication no. 85-23, revised 2011) and the Portuguese law on animal welfare (DL 129/92, DL 197/96; P 1131/97). The competent local authorities approved this experimental protocol (018833).
1. Stock solution preparation (Table 1)
<ol>
	<li>Prepare 1,000 mL of relaxing solution for cardiomyocytes&#39; isolation (RELAX-ISO) by following instructions in Table 2. Dissolve the reagent above in &#8776;500 mL and adjust the pH to 7.0 with KOH. Adjust the final volume to 1000 mL.
	<ol>
		<li>Distribute RELAX-ISO in 50 mL tubes. Store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare 250 mL of activating solution by following the instructions in Table 3. Dissolve the reagents above in &#8776;100 mL of ultra-pure water. Adjust the pH to 7.1 with 5 M KOH at 15 &#176;C. 
	NOTE: Usually, it is necessary to add a significant amount of KOH to reach the desired pH. Put the volumetric balloon in a box with ice to cool down the solution to 15 &#176;C.
	<ol>
		<li>Adjust the final volume to 250 mL. Agitate this solution continuously with a magnetic stirrer until the moment of mixing it with the relaxing solution.</li>
	</ol>
	</li>
	<li>Prepare 100 mL of relaxing solution by following the instructions in Table 4. Dissolve the reagents above in &#8776;50 mL of ultra-pure water. Adjust the pH to 7.1 with KOH 5 M at 15 &#176;C. 
	NOTE:&#160;Usually, it is necessary to add a significant amount of KOH to reach a&#160;pH of 7.1. Place the volumetric balloon in a box filled with ice to cool down the solution to 15 &#176;C. The ionic strength of the solutions used during the measurements amounted to 180 mM.
	<ol>
		<li>Adjust the final volume to 100 mL. Agitate this solution continuously with a magnetic stirrer until the moment of mixing it with activating solution.</li>
	</ol>
	</li>
	<li>Mix activating and relaxing solutions in the proportions presented in Table 5 to obtain pCa solutions between 5.0 and 6.0.
	<ol>
		<li>Always keep relaxing and activating solutions agitating while mixing both.</li>
		<li>Aliquot each mixture to 2 mL&#160;microtubes.&#160;Store all the microtubes at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare a different batch of pCa solution (4.5 to 6.0) for each protocol.</li>
</ol>
2. Calibration of the force transducer
NOTE: The calibration of the force transducer is a routine procedure that should be performed every couple of months or whenever it is suspected to be uncalibrated. The force transducer is highly sensitive and is easily broken. It should be gently handled in every step of its usage, including calibration, gluing of the cardiomyocyte and cleaning.
<ol>
	<li>Detach the force transducer from the rest of the apparatus.</li>
	<li>With the help of a clamp, place the force transducer horizontally in such a way that the needle points downward in the same orientation that the cardiomyocyte will develop force. This will facilitate hanging a series of masses with known weights (elastic band, suture or pin). 
	NOTE: Check the characteristics of the force transducer before proceeding to this step to check the scale factor [mg/volt] and to avoid excessive weight on the transducer. For the force transducer model, the scale factor is 50 (50 mg correspond to 1 volt) and we use 5 weights between 12.5 and 250 mg.</li>
	<li>Turn the force transducer on and let it warm up for 30 min.</li>
	<li>Start by hanging the lighter mass on the force transducer and registering the corresponding voltage measured at FORCE OUT.
	<ol>
		<li>Repeat this procedure for up to five weights.</li>
	</ol>
	</li>
	<li>Plot force applied to the force transducer (load) versus voltage and check for linearity.</li>
	<li>If there is no linearity, adjust the zero and gain potentiometers in the circuit board of the transducer. Check its specific instruction for further information.
	<ol>
		<li>Turn the zero potentiometer until the output voltage reads 0.0 V.</li>
		<li>Hand a medium weight on the transducer needle and adjust the gain potentiometer to read the corresponding voltage (for instance, 50 mg correspond to 1 V). Remove the weight and re-adjust the zero potentiometer to 0.0.</li>
		<li>Repeat step 2.6.2 until the output with and without the weight are correct.</li>
	</ol>
	</li>
	<li>Mount the force transducer back into the apparatus.</li>
</ol>
3. Setting the experimental apparatus
<ol>
	<li>Thaw one vial of each of the activating, 4.5, 5.0, 5.2, 5.4, 5.6, 5.8, 6.0 and relaxing solutions and maintain them on ice. 
	NOTE: ATP and PCr are labile compounds that should be maintained at cold temperatures.</li>
	<li>Prepare the microscope, testing apparatus and associated computer for use (Figure 1).</li>
	<li>Adjust the temperature so that the in-chamber thermometer reads 15 &#176;C. Perform all experiments at this temperature except for kinase and phosphatase incubations (20 &#176;C).</li>
	<li>Turn on the force-transducer and the motor.</li>
</ol>
4. Extraction and permeabilization of skinned cardiomyocytes
<ol>
	<li>Defrost 50 mL of RELAX-ISO solution.</li>
	<li>Turn on the centrifuge and fast cool it up to 4 &#176;C.</li>
	<li>Thaw 3-5 &#181;g of a myocardial sample in a Petri dish containing&#160;2.5 mL of RELAX-ISO solution.</li>
	<li>Cut the tissue in small pieces with a scalpel blade (Figure 2). Cut the sample in a precise way to avoid unnecessary cells damage.</li>
	<li>Transfer the 2.5 mL of RELAX-ISO solution with the tissue to a Potter-Elvehjem glass using&#160;a cut pipette tip.</li>
	<li>Mechanically disrupt the tissue with a grinder at a rotation speed of 30-40 rpm. Press the tissue 3 times for 2 s each to obtain a good cell suspension.</li>
	<li>Prepare 10% Triton in RELAX-ISO solution (250 &#181;L of Triton with 2.25 mL of RELAX-ISO) in a 15 mL tube and add this solution to the cell suspension.</li>
	<li>Gently mix by inverting the tube 3 times.</li>
	<li>Incubate at room temperature for 1 min and 4 min on ice.</li>
	<li>Wash out the Triton by adding RELAX-ISO up to the top of the 15 mL tube; gently mixing (inverting 3 times the tube) and finally spinning down the cells in an angled centrifuge (1 min at 348 x g). Remove the supernatant up to 3 mL above the cell pellet. 
	NOTE: Remove the supernatant gently to avoid disturbing the cell pellet. Still, some cells in supernatant will be inevitably lost.</li>
	<li>Repeat the step 4.10 at least 4 times or until no more bubbles produced by Triton residues are observed. 
	NOTE: The more wash-out steps are made, the more cells are lost with the discarded supernatant.</li>
	<li>In the last wash-out, remove the supernatant up to a volume of 5-10 mL of cell suspension.</li>
</ol>
5. Selecting and gluing the skinned cardiomyocyte
<ol>
	<li>Put a cell suspension drop on a coverslip on top of a glass slide in the microscope slide holder (Figure 1).</li>
	<li>Select a single rod-shaped cardiomyocyte with a good striation pattern and size (Figure 2).</li>
	<li>Find the needle tips of the force-transducer and the motor using the lowest magnification of the inverted microscope.</li>
	<li>Rotate the coverslip to position the selected cardiomyocyte horizontally so that its ends are aligned with the needle of the force transducer and the motor (Figure 2).</li>
	<li>Place a thin line of glue on the side of the coverslip with the help of a swab tip (Figure 2).</li>
	<li>Immerse the needle tips of the force transducer and the motor into the glue line to create a glue halo around both tips. 
	NOTE: Steps 5.6 - 5.10 are accomplished through careful use of the motorized micropositioners.</li>
	<li>Quickly move the needle tips close to the focal plane of the cardiomyocyte.</li>
	<li>Move the needle tip of the force transducer down so that it glues to one edge of the cardiomyocyte.</li>
	<li>Repeat this procedure with the tip of the motor and the other extremity of the cell. 
	NOTE: This procedure must take less than 2-3 min as the glue starts to cure very fast.</li>
	<li>After 5-8 min, lift the needles &#8776;15 &#181;m to avoid gluing the cell to the coverslip. This is done by moving up both micromanipulators simultaneously.</li>
	<li>Let the glue cure. This procedure can last from 15 to 45 min, depending on the type of glue. In our case, the cardiomyocyte is adequately glued after &#8776;15 min.</li>
</ol>
6. Recording force measurements of active, passive and Ca2+ sensitivity
<ol>
	<li>Fill the first experimental well with the relaxing solution (55-100 &#181;L in the experimental apparatus) and the second experimental well with activating solution.</li>
	<li>Using the camera software, place the region of interest (ROI) in an area of the cardiomyocyte with a clear pattern of striation. 
	NOTE: For cardiac myocytes, the operating sarcomere length varies between 1.8 and 2.2 &#181;m, and the optimal sarcomere length is around 2.15 &#181;m.</li>
	<li>Measure the distance between the two extremes of the cardiomyocyte (from the motor to the transducer glue halo, Figure 3) after the optimal sarcomere length has been set (2.2 &#181;m). Record the value as myocyte length in the software.</li>
	<li>Measure cardiomyocyte width and depth, the latter with the aid of a prism mirror placed perpendicular to the cell. 
	NOTE: A powerful, external light source will be required to visualize the cell through the prism. In case there is no prism, and assuming that cardiac cells have an elliptical shape, cardiomyocyte depth can be inferred as 70% of cardiomyocyte width.</li>
	<li>Calculate cross-sectional area (CSA, mm2) assuming an elliptical shape of the cardiomyocyte. 
	<img alt="Equation 1" src="/files/ftp_upload/60427/60427eq1.jpg" /></li>
	<li>Gently move the microscope stage so that the cell moves from the coverslip to the well containing relaxing solution on the back of the stage. 
	NOTE: This procedure can easily damage the cell. Before moving the cell, gently move the needles up a bit more. Avoid removing the cell out of the solution.</li>
	<li>Select the protocol in software that contains two cell shortening (80% of its initial length), that will occur when the cell is emerged in Ca2+ solution and in relaxing solution, respectively (Figure 1, Supplementary File). 
	NOTE: First &#34;Slack&#34; of the cell will be performed within activating solution and the second &#34;Slack&#34; within relaxing solution. By doing this, the user will be able to calculate the total force (Ftotal) of the cell from the 1st and the passive force (Fpassive) of the cell from the 2nd. Use the formula to calculate active force, Factive = Ftotal - Fpassive. The cell is shortened 80% in order to detach all cross-bridges record force.</li>
	<li>Elicit isometric contraction by moving the microscope stage so that the cardiomyocyte moves from the relaxing to the activating solution (pCa=4.5(1)). 
	NOTE: If the cell is functional, it will immediately contract.</li>
	<li>Upon reaching force plateau, start recording the force data. 
	NOTE: The tests can be done individually. Depending on software there is the possibility to create a sequence of tests that will correspond to the different Ca2+ solutions within a Ca2+-sensitivity protocol (Figure 1, Supplementary File).</li>
	<li>Wait ~10 s and then switch the cell immersed in activating solution. 
	NOTE: It is important to wait 10 s before immersing the cell in relaxing solution. If the cell is moved too early, important data to calculate the redevelopment force of the cell (ktr value) might be lost.</li>
	<li>Quickly move the stage so that the cardiomyocyte immerses in the relaxing solution.</li>
	<li>Wait until the test stops.</li>
	<li>Repeat steps 6.8 - 6.12 so that the cell is activated twice in activating solution (pCa=4.5(2)). 
	NOTE: Typically, after the first activation, cardiomyocyte ends can slightly detach&#160;from the needle tips, changing the cardiomyocyte length, CSA and/or the sarcomere length. Readjust to the desired sarcomere length and introduce the corrected dimensions in the software.
	<ol>
		<li>Continue to step 6.13.2 for Ca2+ sensitivity protocol or save the data, if values of passive and active force of the cell are the only parameters needed and detach the cell from needles and clean them with acetone to remove the glue.</li>
		<li>Adjust the sarcomere length of the cell to 2.2 &#181;m by slightly stretching it again, if necessary.</li>
		<li>Replace the activating solution by the next Ca2+ solution (55-100 &#181;L here). Repeat steps 6.8 - 6.12.</li>
		<li>Replace the activating solution by the next Ca2+ solution (55-100 &#181;L here) and repeat steps 6.8 - 6.12 until all solutions have been tested (5.0, 5.2, 5.4, 5.6, 5.8, 6.0).</li>
		<li>Lastly, re-activate the cell with activating solution (pCa4.5(3)). Repeat steps 6.8- 6.12.</li>
	</ol>
	</li>
</ol>
7. Incubation with kinases and phosphatases
<ol>
	<li>After performing the selected baseline protocol, dilute the kinase/phosphatase in Relaxing solution at the recommended concentration. 
	NOTE: It is recommended to carry out a dose-response curve prior to the experiment.</li>
	<li>Set the temperature of the experimental wells to 20 &#176;C.</li>
	<li>Fill the experimental wells with kinase/phosphatase, relaxing solution and activating solution (55-100 &#181;L).</li>
	<li>Gently move the microscope stage so that the cell becomes immersed in the well containing kinase/phosphatase.</li>
	<li>Incubate the cardiomyocyte with the kinase/phosphatase for at least 30 min or according to the manufacturers&#39; instructions.</li>
	<li>Repeat the selected baseline protocol.</li>
</ol>
8. Finalizing the experiment
<ol>
	<li>Unglue the cardiomyocyte from the tips of the force transducer and motor by stretching the cell.</li>
	<li>Carefully remove the glue halo from the needle tips using a cotton swab soaked in acetone.</li>
	<li>Shut down the equipment.</li>
</ol>
9. Analyzing the data
<ol>
	<li>Collect all files from each cardiomyocyte tested. 
	NOTE: Each test will correspond to one file. This means that for each Ca2+ solution or sarcomere length, there will be a corresponding file.</li>
	<li>Calculate active and passive forces of a single cardiomyocyte.
	<ol>
		<li>Open the file corresponding to first activation (pCa=4.5(1)) using a spreadsheet (Figure 3A, Appendix A in Supplementary File). 
		NOTE: We used a custom-made program to perform the analysis. Please see Appendix A in the Supplementary File.</li>
		<li>Average &#8776;60 values before and average &#8776;60 values after the 1st slack of the cell (when the cell is immersed in Ca2+ solution). These 2 values correspond to a and b, respectively.</li>
		<li>Repeat the same analysis for the 2nd slack of the cell (when the cell is immersed in relaxing solution). These 2 values correspond to c and d, respectively.</li>
		<li>Calculate the difference between a and b (total force, Ftotal).</li>
		<li>Calculate the difference between c and d (passive force, Fpassive).</li>
		<li>Calculate active force, Factive = Ftotal - Fpassive.</li>
		<li>Normalize all force values to CSA (see formula above) to obtain the total tension (Ttotal), passive tension (Tpassive) and active tension (Tactive).</li>
		<li>Repeat step 9.2.1 to 9.2.5 for the 2nd activation (pCa=4.5(2)).</li>
		<li>Consider these values as those representing Ttotal, Tactive and Tpassive of the cardiomyocyte under analysis. 
		NOTE: The first activation of the cell with pCa4.5(1) is usually associated with alterations in cell dimensions. For this reason, the 2nd activation with pCa4.5 is more accurate and is the one to be used.</li>
		<li>Repeat the steps 9.2.1 to 9.2.5 for each file/pCa tested solutions (5.0, 5.2, 5.4, 5.6, 5.8, 6.0, 4.5(3)).</li>
	</ol>
	</li>
	<li>Calculate pCa50 and nHill of a single cardiomyocyte. 
	NOTE: For this analysis, use the non-normalized Factive values from the files 4.5(2), 5.0, 5.2, 5.4, 5.6, 5.8, 6.0 and 4.5(3).
	<ol>
		<li>Place in a spreadsheet file all non-normalized values of Factive for each Ca2+ solution tested (4.5(2),5.0, 5.2, 5.4, 5.6, 5.8, 6.0, 4.5(3)).</li>
		<li>Calculate the correction factor = Factive [4.5(2)] - Factive [4.5(3)] / 7.</li>
		<li>Calculate the corrected values of Factive for each Ca2+ solutions (5.0, 5.2, 5.4, 5.6, 5.8, 6.0) by subtracting Factive - correction factor.</li>
		<li>Calculate the relative force (Frelative) for each Ca2+ solutions by normalizing each Factive values by the corresponding corrected value. 
		NOTE: The Frelative [4.5(3)] should equal 1. Each experimental protocol begins and ends with a control activation at saturating Ca2+ concentration (pCa 4.5(2) and 4.5(3)). This allows force normalization and assessment of the rundown of the preparations through the comparison of changes in maximal Ca2+-activated force (Fmax). If at the end of the experimental protocol, the cardiomyocyte produces less than at least 70% of the maximum force of the first contraction, that cell/measurement should be excluded from the analysis.</li>
		<li>Use the Frelative and the corresponding pCa values to fit to a sigmoidal curve with the following equation F(Ca) = CanHill/ (Ca50nHill + CanHill).</li>
		<li>Extrapolate pCa and nHill values from the equation above mentioned.</li>
	</ol>
	</li>
	<li>Calculate the rate of force redevelopment (ktr) of a single cardiomyocyte.
	<ol>
		<li>Perform a fit to the curve that corresponds to the values immediately after the 1st cell slack.</li>
		<li>Calculate the slope of the curve and this value will correspond to the rate of force redevelopment.</li>
		<li>Repeat step 9.4.1 and 9.4.2 for each Ca2+ solution. 
		NOTE: A poor curve fit will be obtained for the lowest Ca-solutions (R squared&#8804;0.90).</li>
	</ol>
	</li>
</ol>

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Y., Mosca, E., de Tombe, P. P., Cazorla, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+contribution+of+cardiac+sarcomeric+proteins+in+the+myofibrillar+force+response+to+stretch.">Differential contribution of cardiac sarcomeric proteins in the myofibrillar force response to stretch.</a> Pflugers Archiv. 457 (1), 25-36 (2008).</li><li>Falcao-Pires, I., 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=Diabetes+mellitus+worsens+diastolic+left+ventricular+dysfunction+in+aortic+stenosis+through+altered+myocardial+structure+and+cardiomyocyte+stiffness.">Diabetes mellitus worsens diastolic left ventricular dysfunction in aortic stenosis through altered myocardial structure and cardiomyocyte stiffness.</a> Circulation. 124 (10), 1151-1159 (2011).</li><li>Lim, C. 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=Anthracyclines+induce+calpain-dependent+titin+proteolysis+and+necrosis+in+cardiomyocytes.">Anthracyclines induce calpain-dependent titin proteolysis and necrosis in cardiomyocytes.</a> Journal Biology Chemistry. 279 (9), 8290-8299 (2004).</li><li>Woulfe, K. 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=A+Novel+Method+of+Isolating+Myofibrils+From+Primary+Cardiomyocyte+Culture+Suitable+for+Myofibril+Mechanical+Study.">A Novel Method of Isolating Myofibrils From Primary Cardiomyocyte Culture Suitable for Myofibril Mechanical Study.</a> Frontiers Cardiovascular Medicine. 6, 12 (2019).</li><li>Ait Mou, Y., Bollensdorff, C., Cazorla, O., Magdi, Y., de Tombe, P. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exploring+cardiac+biophysical+properties.">Exploring cardiac biophysical properties.</a> Global Cardiology Science Practice. 2015, 10 (2015).</li></ol>

The authors have no conflict of interest.

In vitro assessment of cardiac function using skinned cardiomyocytes represents an important technique to clarify the modifications occurring at cardiomyocyte level in physiological (e.g., stretch) and pathological context (e.g., ischemia). This methodology has several advantages such as requiring a minimal amount of myocardium to assess function in cardiomyocytes obtained from defrosted samples; using cardiomyocytes from a wide range of species (mice13, rat1,14,15, rabbit16, pig17, dog18, guinea pig19 and human20) and different cardiac locations, including the atria, left and right ventricles or a specific region of the infarcted heart. Moreover, this technique allows delivering specific concentrations of Ca2+ and energy (ATP) while measuring the function of regulatory and contractile structures in their native configuration.
Despite the simplicity of this technique, there are some critical steps. It is essential to guarantee the quality of each step from the beginning, including sample collection. Myofilament proteins are susceptible to proteases21. Thus it is mandatory to store samples in liquid nitrogen immediately after its collection. Fresh samples, which were not previously frozen, will develop significantly higher forces, so it is not advisable to mix measurement done in fresh and frozen samples in the same protocol. The second most critical step is the cardiomyocytes&#39; extraction. During this procedure, it is crucial to maintain the sample on ice most of the time. A protease inhibitor cocktail can be used to reduce the risk of protein degradation during the extraction/permeabilization22. Thirdly, samples should be cut in smaller pieces using precise scalpel movements since we noted reduced quality cardiomyocytes when this step was disregarded. Another critical step is washing the cardiomyocytes since it is difficult to have the right balance between washing out Triton (permeabilizes the cell but promotes its ungluing) and keeping as&#160;many cells in the supernatant as possible. It is important to first try the extraction and number of washouts for each sample, species or protocol. For instance, in our hands, we noted that ZSF1 obese rat tissue extractions have a &#34;fatty&#34; aspect, which made these cells more slippery during the gluing but not more difficult to measure. The way we circumvent this problem was by performing more experiments to have a reasonable number of cells per animal. Moreover, it is crucial to select a good cell to glue, namely with good striation and reasonable length. If the cardiomyocyte does not have these features, it will mostly detach from the needle tips or develop no/low force. It is also important to use the correct glue for cardiomyocyte attachment, considering the time of gluing and its efficacy to glue the cell to the needle. In our hands, the silicone glue (Table of Materials) cures fast (10-15 min) and strong enough. Finally, the last critical step is related with carefully lifting the cardiomyocyte 5 min after gluing the cell (to avoid gluing the cell to the coverslip) and before moving it to the wells (to avoid the cell to be dragged by the microscope stage). Table 7 summarizes the troubleshooting associated with this technique, its underlying causes and possible solutions to overcome frequent problems.
The major limitation of this method is that it cannot answer all the questions related to the myofilament contractility, such as how fast the myofilaments activate/deactivate. In the in vivo setting, membrane depolarization, intracellular Ca2+ increase and its diffusion to myofilaments need to occur for the myocytes to contract, whereas in skinned cardiomyocytes Ca2+ diffusion to myofilaments occurs immediately when the cell is submerged in the Ca2+ solution. This faster rate of Ca2+ diffusion will bias myofilaments activation/deactivation analysis23.
These experiments are influenced by different factors, including the temperature, solution pH, mechanical perturbation (slack-re-stretch vs. slack) and cell attachment procedures (pin tie vs. glue), all of these variables accounting for literature discrepancies in terms of ktr and the sarcomere length-dependent increase in force4,12.
Future progress of the technique includes performing functional studies in intact rather than permeabilized cardiomyocytes. This technique has the disadvantage of relying on cardiomyocytes freshly isolated (not previously frozen). Another important issue not directly related to this methodology but that may significantly impact it is related to the maximal period of sample frozen storage. Specifically, it is mandatory to establish the degree of myofilament degradation throughout storage time (i.e., for how long frozen samples can be stored in order to assure good quality functional data derived from the extracted cardiomyocytes).

Traditionally, assessment of myocardial mechanical properties has been attempted mostly in multicellular preparations, such as papillary muscles and trabeculae1,2. Multicellular cardiac muscles strips include a heterogeneous population of cells, including contractile cardiomyocytes with an unknown pattern of orientation and force generation, electrical activity and stress/strain distributions as well as a surrounding connective tissue matrix3,4. A preparation without collagen and containing a single cardiomyocyte would allow measurement of sarcomere length and cross-bridge contractile properties in a very precise and controlled manner5,6. Therefore, over the last four decades, several methodologies were developed allowing investigating the mechanical, contractile, and relaxation properties of a single cardiomyocyte6,7. The contractile function of these cells is strongly dependent on sarcomere length and cross-bridge cycling kinetics3. Thus, it is desirable to investigate muscle function directly in single isolated cardiac cells, considering that it allows assessing sarcomere length and performance as well as cross-bridge function and contractile properties. However, isolating and attaching functional cardiomyocytes with a reasonable optical sarcomere resolution while recording force measurement at the &#181;N level is still challenging and evolving3,6. Other challenges are the logistics that need to be installed to isolate cardiomyocytes from freshly collected biopsies. The unpredictability of human biopsies collection, for instance, may jeopardize the feasibility of the experiments.
Moreover, ethical concerns regarding the Replacement, Reduction and Refinement of animal experimentation for scientific procedures (principles of the 3Rs) have promoted study changes at the cellular and tissue level, preferably in human biopsies, or in smaller animal samples. Indeed a progressive refinement of methodologies to assess cardiac function in vitro on a smaller level of complexity allows proper integration of the results to the whole body and translate them to the clinical scenario7. Altogether, using samples stored at -80 &#176;C to extract cardiomyocytes may be an appealing alternative.
The myocardial tissue is cut into small pieces and homogenized with a mortar and a pestle. The result of this homogenization is a suspension of skinned bundled and isolated cells with varying degrees of sarcolemmal damage, wherein the myoplasm is exposed to the bathing medium and all the cellular components are washed out. Structures such as the myofibrils that are further away from the sarcolemma are preserved. Thus, sarcomere shortening and functional properties associated with the myofibrillar apparatus are kept intact and can be recorded8,9.
The cardiomyocyte force measurement system consists of an electromagnetic motor, used to adjust cardiomyocyte length, and a force transducer, that measures isometric cardiomyocyte contraction. A permeabilized, or skinned, cardiomyocyte is placed in an experimental chamber containing a relaxing solution ([Ca2+] &#60; 10 nM) and silicon-glued to 2 thin needles: one attached to the motor and the other to the force transducer. An optical system is used to determine cardiomyocyte morphology and sarcomere length. The experimental protocol often consists of a series of force recordings upon buffer solutions containing different Ca2+ concentrations, the determination of actin-myosin cross-bridge kinetics and the measurement of the passive tension of the mounted cardiomyocytes at pre-defined sarcomere lengths (Figure 1). Isolation of permeabilized cardiomyocytes from myocardial samples frozen in liquid nitrogen (and subsequently stored at -80 &#176;C) is a technique that utilizes cellular mechanics and protein biochemistry for measuring maximal Ca2+-activated (active) force per cross-sectional area (Tactive, kN&#8729;m-2), Ca2+-independent (passive) tension (Tpassive, kN&#8729;m-2), myofilaments Ca2+-sensitivity (pCa50), myofilaments cooperativity (nHill), the rate of force redevelopment (ktr) as well as sarcomere length dependencies of Tactive, Tpassive, pCa50, nHill and ktr.
The goal of this protocol is to illustrate and summarize the potential of the cardiomyocyte force measurement system as a reliable procedure to assess the functional mechanical properties of single skinned cardiomyocytes isolated from frozen samples from different species.

<table><tbody><tr><td>Acetone</td><td>Sigma</td><td>34580</td><td></td></tr><tr><td>Adenosine 5&amp;rsquo;-triphosphate disodium salt hydrate (Na2ATP)</td><td>Sigma</td><td>A2383</td><td></td></tr><tr><td>Calcium carbonate (CaCO3)</td><td>Merck</td><td>1.02067.0500</td><td></td></tr><tr><td>Imidazole</td><td>VWR</td><td>24720.157</td><td></td></tr><tr><td>Magnesium chloride hexahydrate (MgCl2.6H2O)</td><td>Merck</td><td>1.05833.0250</td><td></td></tr><tr><td>Magnesium chloride solution (MgCl2 1M)</td><td>Sigma</td><td>M1028</td><td></td></tr><tr><td>N,N-Bis(2-hydroxyethyl)taurine (BES)</td><td>Sigma</td><td>B9879</td><td></td></tr><tr><td>Phosphocreatine dissodium salt hydrate (Na2PCr)</td><td>Sigma</td><td>P7936</td><td></td></tr><tr><td>Potassium chloride (KCl)</td><td>Merck</td><td>1.04936.1000</td><td></td></tr><tr><td>Potassium hydroxide (KOH)</td><td>Merck</td><td>8.14353.1000</td><td></td></tr><tr><td>Propionic acid (C3H6O2)</td><td>Merck</td><td>8.00605.0500</td><td></td></tr><tr><td>Silicone Squeeze Tube</td><td>Marineland</td><td>31003</td><td></td></tr><tr><td>Tritiplex (EGTA)</td><td>Merck</td><td>1.08435.0025</td><td></td></tr><tr><td>Triton&amp;reg; X-100 10%</td><td>Merck</td><td>648463</td><td></td></tr><tr><td>Tissue homogeneizer (GKH GT Motor Control)</td><td>Terre Haute Glas-col</td><td></td><td></td></tr><tr><td>Length Controller (Model 315C-I)</td><td>Aurora Scientific</td><td></td><td></td></tr><tr><td>Force Transducer (Model 403 A)</td><td>Aurora Scientific</td><td></td><td></td></tr><tr><td>Software ASI 600A</td><td>Aurora Scientific</td><td></td><td></td></tr><tr><td>Sotware VSL (Model 900B)</td><td>Aurora Scientific</td><td></td><td></td></tr><tr><td>Inverted Microscope (IX51)</td><td>Olympus</td><td></td><td></td></tr></tbody></table>

in vitro assessment of cardiac function using skinned cardiomyocytes

In this article, we describe the steps required to isolate a single permeabilized (&#34;skinned&#34;) cardiomyocyte and attach it to a force-measuring apparatus and a motor to perform functional studies. These studies will allow measurement of cardiomyocyte stiffness (passive force) and its activation with different calcium (Ca2+)-containing solutions to determine, amongst others: maximum force development, myofilament Ca2+-sensitivity (pCa50), cooperativity (nHill) and the rate of force redevelopment (ktr). This method also enables determination of the effects of drugs acting directly on myofilaments and of the expression of exogenous recombinant proteins on both active and passive properties of cardiomyocytes. Clinically, skinned cardiomyocyte studies highlight the pathophysiology of many myocardial diseases and allow in vitro assessment of the impact of therapeutic interventions targeting the myofilaments. Altogether, this technique enables the clarification of cardiac pathophysiology by investigating correlations between in vitro and in vivo parameters in animal models and human tissue obtained during open heart or transplant surgery.

Watch this Scientific Journal Video about In Vitro Assessment of Cardiac Function Using Skinned Cardiomyocytes at JoVE.com

In Vitro Assessment of Cardiac Function Using Skinned Cardiomyocytes

This protocol aims to describe step-by-step the technique of extraction and assessment of cardiac function using skinned cardiomyocytes. This methodology allows measurement and acutemodulation of myofilament function using small frozen biopsies that can be collected from different cardiac locations, from mice to men.

In this article, we describe the steps required to isolate a single permeabilized (&#34;skinned&#34;) cardiomyocyte and attach it to a force-measuring ...

in-vitro-assessment-of-cardiac-function-using-skinned-cardiomyocytes

Research

JoVE Journal

Medicine

6.1K Views.  Universidade do Porto. Dit protocol is bedoeld om stapsgewijze de techniek van extractie en beoordeling van de hartfunctie te beschrijven met behulp van gevilde cardiomyocyten. Deze methode maakt het mogelijk om de myofilamentfunctie te meten en te acuut temodificeren met kleine bevroren biopsieën die kunnen worden verzameld vanaf verschillende hartlocaties, van muizen tot mannen.

Video: In Vitro Assessment of Cardiac Function Using Skinned Cardiomyocytes

Automated Contraction Analysis of Human Engineered Heart Tissue for Cardiac Drug Safety Screening

Cardiac tissue engineering describes techniques to constitute three dimensional force-generating engineered tissues. For the implementation of these procedures in basic research and preclinical drug development, it is important to develop protocols for automated generation and analysis under standardized conditions. Here, we present a technique to generate engineered heart tissue (EHT) from cardiomyocytes of different species (rat, mouse, human). The technique relies on the assembly of a fibrin-gel containing dissociated cardiomyocytes between elastic polydimethylsiloxane (PDMS) posts in a 24-well format. Three-dimensional, force-generating EHTs constitute within two weeks after casting. This procedure allows for the generation of several hundred EHTs per week and is technically limited only by the availability of cardiomyocytes (0.4-1.0 x 106/EHT). Evaluation of auxotonic muscle contractions is performed in a modified incubation chamber with a mechanical interlock for 24-well plates and a camera placed on top of this chamber. A software controls a camera moved on an XYZ axis system to each EHT. EHT contractions are detected by an automated figure recognition algorithm, and force is calculated based on shortening of the EHT and the elastic propensity and geometry of the PDMS posts. This procedure allows for automated analysis of high numbers of EHT under standardized and sterile conditions. The reliable detection of drug effects on cardiomyocyte contraction is crucial for cardiac drug development and safety pharmacology. We demonstrate, with the example of the hERG channel inhibitor E-4031, that the human EHT system replicates drug responses on contraction kinetics of the human heart, indicating it to be a promising tool for cardiac drug safety screening.

Here, we show the generation of human engineered heart tissue from induced pluripotent stem cells (hiPSC)-derived cardiomyocytes. We present a method to analyze contraction force and exemplary alteration of contraction pattern by the hERG channel inhibitor E-4031. This method shows high level of robustness and suitability for cardiac drug screening.

Cardiac tissue engineering describes techniques to constitute three dimensional force-generating engineered tissues. For the implementation of these ...

Bioengineering

Simultaneous Electrical and Mechanical Stimulation to Enhance Cells' Cardiomyogenic Potential

Cardiovascular diseases are the leading cause of death in developed countries. Consequently, the demand for effective cardiac cell therapies has motivated researchers in the stem cell and bioengineering fields to develop in vitro high-fidelity human myocardium for both basic research and clinical applications. However, the immature phenotype of cardiac cells is a limitation on obtaining tissues that functionally mimic the adult myocardium, which is mainly characterized by mechanical and electrical signals. Thus, the purpose of this protocol is to prepare and mature the target cell population through electromechanical stimulation, recapitulating physiological parameters. Cardiac tissue engineering is evolving toward more biological approaches, and strategies based on biophysical stimuli, thus, are gaining momentum. The device developed for this purpose is unique and allows individual or simultaneous electrical and mechanical stimulation, carefully characterized and validated. In addition, although the methodology has been optimized for this stimulator and a specific cell population, it can easily be adapted to other devices and cell lines. The results here offer evidence of the increased cardiac commitment of the cell population after electromechanical stimulation. Electromechanically stimulated cells show an increased expression of main cardiac markers, including early, structural, and calcium-regulating genes. This cell conditioning could be useful for further regenerative cell therapy, disease modeling, and high-throughput drug screening.

Here we present a protocol for training a cell population using electrical and mechanical stimuli emulating cardiac physiology. This electromechanical stimulation enhances the cardiomyogenic potential of the treated cells and is a promising strategy for further cell therapy, disease modeling, and drug screening.

Cardiovascular diseases are the leading cause of death in developed countries. Consequently, the demand for effective cardiac cell therapies has motivated ...

Preclinical Cardiac Electrophysiology Assessment by Dual Voltage and Calcium Optical Mapping of Human Organotypic Cardiac Slices

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between animal and clinical trials. Numerous animal and cell models have been used to examine the effects of drugs, yet these responses often differ in humans. Human cardiac slices offer an advantage for drug testing in that they are directly derived from viable human hearts. In addition to having preserved multicellular structures, cell-cell coupling, and extracellular matrix environments, human cardiac tissue slices can be used to directly test the effect of innumerable drugs on adult human cardiac physiology. What distinguishes this model from other heart preparations, such as whole hearts or wedges, is that slices can be subjected to longer-term culture. As such, cardiac slices allow for studying the acute as well as chronic effects of drugs. Furthermore, the ability to collect several hundred to a thousand slices from a single heart makes this a high-throughput model to test several drugs at varying concentrations and combinations with other drugs at the same time. Slices can be prepared from any given region of the heart. In this protocol, we describe the preparation of left ventricular slices by isolating tissue cubes from the left ventricular free wall and sectioning them into slices using a high precision vibrating microtome. These slices can then either be subjected to acute experiments to measure baseline cardiac electrophysiological function or cultured for chronic drug studies. This protocol also describes dual optical mapping of cardiac slices for simultaneous recordings of transmembrane potentials and intracellular calcium dynamics to determine the effects of the drugs being investigated.

This protocol describes the procedure for sectioning and culturing human cardiac slices for preclinical drug testing and details the use of optical mapping for recording transmembrane voltage and intracellular calcium signals simultaneously from these slices.

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between ...

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 ...

Cardiac Spheroids as in vitro Bioengineered Heart Tissues to Study Human Heart Pathophysiology

Despite several advances in cardiac tissue engineering, one of the major challenges to overcome remains the generation of a fully functional vascular network comprising several levels of complexity to provide oxygen and nutrients within bioengineered heart tissues. Our laboratory has developed a three-dimensional in vitro model of the human heart, known as the "cardiac spheroid" or "CS". This presents biochemical, physiological, and pharmacological features typical of the human heart and is generated by co-culturing its three major cell types, such as cardiac myocytes, endothelial cells, and fibroblasts. Human induced pluripotent stem cells-derived cardiomyocytes (hiPSC-CMs or iCMs) are co-cultured at ratios approximating the ones found in vivo with human cardiac fibroblasts (HCFs) and human coronary artery endothelial cells (HCAECs) in hanging drop culture plates for three to four days. The confocal analysis of CSs stained with antibodies against cardiac Troponin T, CD31 and vimentin (markers for cardiac myocytes, endothelial cells and fibroblasts, respectively) shows that CSs present a complex endothelial cell network, resembling the native one found in the human heart. This is confirmed by the 3D rendering analysis of these confocal images. CSs also present extracellular matrix (ECM) proteins typical of the human heart, such as collagen type IV, laminin and fibronectin. Finally, CSs present a contractile activity measured as syncytial contractility closer to the one typical of the human heart compared to CSs that contain iCMs only. When treated with a cardiotoxic anti-cancer agent, such as doxorubicin (DOX, used to treat leukemia, lymphoma and breast cancer), the viability of DOX-treated CSs is significantly reduced at 10 &#181;M genetic and chemical inhibition of endothelial nitric oxide synthase, a downstream target of DOX in HCFs and HCAECs, reduced its toxicity in CSs. Given these unique features, CSs are currently used as in vitro models to study heart biochemistry, pathophysiology, and pharmacology.

This protocol aims to fabricate 3D cardiac spheroids (CSs) by co-culturing cells in hanging drops.&#160;Collagen-embedded CSs are treated with doxorubicin (DOX, a cardiotoxic agent) at physiological concentrations to model heart failure. In vitro testing using DOX-treated CSs may be used to identify novel therapies for heart failure patients.

Despite several advances in cardiac tissue engineering, one of the major challenges to overcome remains the generation of a fully functional vascular ...

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 ...

Evaluation of Cardiac Contractility Modulation Therapy in 2D Human Stem Cell-Derived Cardiomyocytes

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are currently being explored for multiple in vitro applications and have been used in regulatory submissions. Here, we extend their use to cardiac medical device safety or performance assessments. We developed a novel method to evaluate cardiac medical device contractile properties in robustly contracting 2D hiPSC-CMs monolayers plated on a flexible extracellular matrix (ECM)-based hydrogel substrate. This tool enables the quantification of the effects of cardiac electrophysiology device signals on human cardiac function (e.g., contractile properties) with standard laboratory equipment. The 2D hiPSC-CM monolayers were cultured for 2-4 days on a flexible hydrogel substrate in a 48-well format.
The hiPSC-CMs were exposed to standard cardiac contractility modulation (CCM) medical device electrical signals and compared to control (i.e., pacing only) hiPSC-CMs. The baseline contractile properties of the 2D hiPSC-CMs were quantified by video-based detection analysis based on pixel displacement. The CCM-stimulated 2D hiPSC-CMs plated on the flexible hydrogel substrate displayed significantly enhanced contractile properties relative to baseline (i.e., before CCM stimulation), including an increased peak contraction amplitude and accelerated contraction and relaxation kinetics. Furthermore, the utilization of the flexible hydrogel substrate enables the multiplexing of the video-based cardiac-excitation contraction coupling readouts (i.e., electrophysiology, calcium handling, and contraction) in healthy and diseased hiPSC-CMs. The accurate detection and quantification of the effects of cardiac electrophysiological signals on human cardiac contraction is vital for cardiac medical device development, optimization, and de-risking. This method enables the robust visualization and quantification of the contractile properties of the cardiac syncytium, which should be valuable for nonclinical cardiac medical device safety or effectiveness testing. This paper describes, in detail, the methodology to generate 2D hiPSC-CM hydrogel substrate monolayers.

Here, we demonstrate a non-invasive cardiac medical device contractility evaluation method using 2D human induced pluripotent stem cell-derived cardiomyocyte (hiPSC-CM) monolayers, plated on a flexible substrate, coupled with video-based microscopy. This tool will be useful for the in vitro evaluation of the contractile properties of cardiac electrophysiology devices.

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are currently being explored for multiple in vitro applications and have been used ...

High-Throughput Cardiotoxicity Screening Using Mature Human Induced Pluripotent Stem Cell-Derived Cardiomyocyte Monolayers

Human induced stem cell-derived cardiomyocytes (hiPSC-CMs) are used to replace and reduce the dependence on animals and animal cells for preclinical cardiotoxicity testing. In two-dimensional monolayer formats, hiPSC-CMs recapitulate the structure and function of the adult human heart muscle cells when cultured on an optimal extracellular matrix (ECM). A human perinatal stem cell-derived ECM (maturation-inducing extracellular matrix-MECM) matures the hiPSC-CM structure, function, and metabolic state in 7 days after plating. 
Mature hiPSC-CM monolayers also respond as expected to clinically relevant medications, with a known risk of causing arrhythmias and cardiotoxicity. The maturation of hiPSC-CM monolayers was an obstacle to the widespread adoption of these valuable cells for regulatory science and safety screening, until now. This article presents validated methods for the plating, maturation, and high-throughput, functional phenotyping of hiPSC-CM electrophysiological and contractile function. These methods apply to commercially available purified cardiomyocytes, as well as stem cell-derived cardiomyocytes generated in-house using highly efficient, chamber-specific differentiation protocols. 
High-throughput electrophysiological function is measured using either voltage-sensitive dyes (VSDs; emission: 488 nm), calcium-sensitive fluorophores (CSFs), or genetically encoded calcium sensors (GCaMP6). A high-throughput optical mapping device is used for optical recordings of each functional parameter, and custom dedicated software is used for electrophysiological data analysis. MECM protocols are applied for medication screening using a positive inotrope (isoprenaline) and human Ether-a-go-go-related gene (hERG) channel-specific blockers. These resources will enable other investigators to successfully utilize mature hiPSC-CMs for high-throughput, preclinical cardiotoxicity screening, cardiac medication efficacy testing, and cardiovascular research.

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) offer an alternative to using animals for preclinical cardiotoxicity screening. A limitation to the widespread adoption of hiPSC-CMs in preclinical toxicity screening is the immature, fetal-like phenotype of the cells. Presented here are protocols for robust and rapid maturation of hiPSC-CMs.

Human induced stem cell-derived cardiomyocytes (hiPSC-CMs) are used to replace and reduce the dependence on animals and animal cells for preclinical ...

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 ...

Biology

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.

In Vitro Assessment of Cardiac Function Using Skinned Cardiomyocytes

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