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>

<ol>
	<li>Leite-Moreira, A. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stretch-induced+compliance:+a+novel+adaptive+biological+mechanism+following+acute+cardiac+load.">Stretch-induced compliance: a novel adaptive biological mechanism following acute cardiac load.</a> Cardiovascular Research. 114 (5), 656-667 (2018).</li><li>Falcao-Pires, I., Fontes-Sousa, A. P., Lopes-Conceicao, L., Bras-Silva, C., Leite-Moreira, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulation+of+myocardial+stiffness+by+&#946;-adrenergic+stimulation+-+its+role+in+normal+and+failing+heart.">Modulation of myocardial stiffness by &#946;-adrenergic stimulation - its role in normal and failing heart.</a> Physiological Research. 60 (4), 599-609 (2011).</li><li>Cokkinos, D. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Introduction to Translational Cardiovascular Research. , 371-387 (2015).</li><li>van der Velden, J., Stienen, G. J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+Disorders+and+Pathophysiology+of+Sarcomeric+Proteins.">Cardiac Disorders and Pathophysiology of Sarcomeric Proteins.</a> Physiological Reviews. 99 (1), 381-426 (2019).</li><li>Garnier, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Attachment+procedures+for+mechanical+manipulation+of+isolated+cardiac+myocytes:+a+challenge.">Attachment procedures for mechanical manipulation of isolated cardiac myocytes: a challenge.</a> Cardiovascular Research. 28 (12), 1758-1764 (1994).</li><li>Brady, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+properties+of+isolated+cardiac+myocytes.">Mechanical properties of isolated cardiac myocytes.</a> Physiological Reviews. 71 (2), 413-428 (1991).</li><li>Falcao-Pires, I., Leite-Moreira, A. F., Cokkinos, D. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+20.">Chapter 20.</a> Introduction to Translational Cardiovascular Research. 20, 371-387 (2015).</li><li>Liang, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Teaching+calcium-induced+calcium+release+in+cardiomyocytes+using+a+classic+paper+by+Fabiato.">Teaching calcium-induced calcium release in cardiomyocytes using a classic paper by Fabiato.</a> Advances Physiology Education. 32 (1), 1-10 (2008).</li><li>Roche, S. M., Gumucio, J. P., Brooks, S. V., Mendias, C. L., Claflin, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+Maximum+Isometric+Force+Generated+by+Permeabilized+Skeletal+Muscle+Fibers.">Measurement of Maximum Isometric Force Generated by Permeabilized Skeletal Muscle Fibers.</a> Journal of Visualized Experiments. (100), e52695 (2015).</li><li>Huxley, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscle+structure+and+theories+of+contraction.">Muscle structure and theories of contraction.</a> Progress Biophysics and Biophysical Chemistry. 7, 255-318 (1957).</li><li>Sequeira, V., 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=Synergistic+role+of+ADP+and+Ca(2+)+in+diastolic+myocardial+stiffness.">Synergistic role of ADP and Ca(2+) in diastolic myocardial stiffness.</a> Journal Physiology. 593 (17), 3899-3916 (2015).</li><li>Edes, I. F., 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=Rate+of+tension+redevelopment+is+not+modulated+by+sarcomere+length+in+permeabilized+human,+murine,+and+porcine+cardiomyocytes.">Rate of tension redevelopment is not modulated by sarcomere length in permeabilized human, murine, and porcine cardiomyocytes.</a> American Journal Physiology Regulatory Integrative Comparative Physiology. 293 (1), R20-R29 (2007).</li><li>King, N. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+intact+cardiac+myocyte+mechanics:+cross-bridge+and+titin-based+stress+in+unactivated+cells.">Mouse intact cardiac myocyte mechanics: cross-bridge and titin-based stress in unactivated cells.</a> Journal General Physiology. 137 (1), 81-91 (2011).</li><li>Hamdani, N., 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=Myocardial+titin+hypophosphorylation+importantly+contributes+to+heart+failure+with+preserved+ejection+fraction+in+a+rat+metabolic+risk+model.">Myocardial titin hypophosphorylation importantly contributes to heart failure with preserved ejection fraction in a rat metabolic risk model.</a> Circulation Heart Failure. 6 (6), 1239-1249 (2013).</li><li>Miranda-Silva, D., 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=Characterization+of+biventricular+alterations+in+myocardial+(reverse)+remodelling+in+aortic+banding-induced+chronic+pressure+overload.">Characterization of biventricular alterations in myocardial (reverse) remodelling in aortic banding-induced chronic pressure overload.</a> Scientific Reports. 9 (1), 2956 (2019).</li><li>Rodrigues, P. G., 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=Early+myocardial+changes+induced+by+doxorubicin+in+the+nonfailing+dilated+ventricle.">Early myocardial changes induced by doxorubicin in the nonfailing dilated ventricle.</a> American Journal Physiology Heart Circulatory Physiology. 316 (3), H459-H475 (2019).</li><li>van der Velden, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Alterations+in+myofilament+function+contribute+to+left+ventricular+dysfunction+in+pigs+early+after+myocardial+infarction.">Alterations in myofilament function contribute to left ventricular dysfunction in pigs early after myocardial infarction.</a> Circulation Research. 95 (11), e85-e95 (2004).</li><li>Wakili, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple+potential+molecular+contributors+to+atrial+hypocontractility+caused+by+atrial+tachycardia+remodeling+in+dogs.">Multiple potential molecular contributors to atrial hypocontractility caused by atrial tachycardia remodeling in dogs.</a> Circulation: Arrhythmia Electrophysiology. 3 (5), 530-541 (2010).</li><li>Ait Mou, Y., le Guennec, J. 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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Acetone</td>
			<td>Sigma</td>
			<td>34580</td>
			<td></td>
		</tr>
		<tr>
			<td>Adenosine 5&#8217;-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&#174; 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

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5.9K Views.  Universidade do Porto. 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.

Video: In Vitro Assessment of Cardiac Function Using Skinned Cardiomyocytes

Assessment of Cardiac Function and Energetics in Isolated Mouse Hearts Using 31P NMR Spectroscopy

Bioengineered mouse models have become powerful research tools in determining causal relationships between molecular alterations and models of cardiovascular disease. Although molecular biology is necessary in identifying key changes in the signaling pathway, it is not a surrogate for functional significance. While physiology can provide answers to the question of function, combining physiology with biochemical assessment of metabolites in the intact, beating heart allows for a complete picture of cardiac function and energetics. For years, our laboratory has utilized isolated heart perfusions combined with nuclear magnetic resonance (NMR) spectroscopy to accomplish this task. Left ventricular function is assessed by Langendorff-mode isolated heart perfusions while cardiac energetics is measured by performing 31P magnetic resonance spectroscopy of the perfused hearts. With these techniques, indices of cardiac function in combination with levels of phosphocreatine and ATP can be measured simultaneously in beating hearts. Furthermore, these parameters can be monitored while physiologic or pathologic stressors are instituted. For example, ischemia/reperfusion or high workload challenge protocols can be adopted. The use of aortic banding or other models of cardiac pathology are apt as well. Regardless of the variants within the protocol, the functional and energetic significance of molecular modifications of transgenic mouse models can be adequately described, leading to new insights into the associated enzymatic and metabolic pathways. Therefore, 31P NMR spectroscopy in the isolated perfused heart is a valuable research technique in animal models of cardiovascular disease.

Assessment of Cardiac Function and Energetics in Isolated Mouse Hearts Using 31P NMR Spectroscopy

Langendorff-mode isolated heart perfusion, in conjunction with 31P NMR spectroscopy, combines the fields of biochemistry and physiology into one experiment. The protocol allows for the dynamic measurement of high energy phosphate content and turnover in the heart while concurrently monitoring physiologic function. When performed correctly, this is a valuable technique in the assessment of cardiac energetics.

Bioengineered mouse models have become powerful research tools in determining causal relationships between molecular alterations and models of ...

Assessment of Cardiac Function and Myocardial Morphology Using Small Animal Look-locker Inversion Recovery (SALLI) MRI in Rats

Small animal magnetic resonance imaging is an important tool to study cardiac function and changes in myocardial tissue. The high heart rates of small animals (200 to 600 beats/min) have previously limited the role of CMR imaging. Small animal Look-Locker inversion recovery (SALLI) is a T1 mapping sequence for small animals to overcome this problem 1. T1 maps provide quantitative information about tissue alterations and contrast agent kinetics. It is also possible to detect diffuse myocardial processes such as interstitial fibrosis or edema 1-6. Furthermore, from a single set of image data, it is possible to examine heart function and myocardial scarring by generating cine and inversion recovery-prepared late gadolinium enhancement-type MR images 1.
	The presented video shows step-by-step the procedures to perform small animal CMR imaging. Here it is presented with a healthy Sprague-Dawley rat, however naturally it can be extended to different cardiac small animal models.

Assessment of Cardiac Function and Myocardial Morphology Using Small Animal Look-locker Inversion Recovery SALLI MRI in Rats

Contrast enhanced cardiac magnetic resonance (CMR) imaging allows comprehensive in vivo assessment of the heart in small animal models of cardiovascular disease. Here we detail the procedures to perform CMR imaging, reconstruction and analysis using Small Animal Lock-Locker Inversion Recovery (SALLI) in rats.

Small animal magnetic resonance  imaging is an important tool to study cardiac function and changes in myocardial  tissue. The high heart rates of small ...

Assessment of Right Ventricular Structure and Function in Mouse Model of Pulmonary Artery Constriction by Transthoracic Echocardiography

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, the RV is significantly affected in pulmonary diseases such as pulmonary artery hypertension (PAH). In addition, the RV is remarkably sensitive to cardiac pathologies, including left ventricular (LV) dysfunction, valvular disease or RV infarction4. To understand the role of RV in the pathogenesis of cardiac diseases, a reliable and noninvasive method to access the RV structurally and functionally is essential.
A noninvasive trans-thoracic echocardiography (TTE) based methodology was established and validated for monitoring dynamic changes in RV structure and function in adult mice. To impose RV stress, we employed a surgical model of pulmonary artery constriction (PAC) and measured the RV response over a 7-day period using a high-frequency ultrasound microimaging system. Sham operated mice were used as controls. Images were acquired in lightly anesthetized mice at baseline (before surgery), day 0 (immediately post-surgery), day 3, and day 7 (post-surgery). Data was analyzed offline using software.
Several acoustic windows (B, M, and Color Doppler modes), which can be consistently obtained in mice, allowed for reliable and reproducible measurement of RV structure (including RV wall thickness, end-diastolic&#160;and end-systolic dimensions), and function (fractional area change, fractional shortening, PA peak velocity, and peak pressure gradient) in normal mice and following PAC.
Using this method, the pressure-gradient resulting from PAC was accurately measured in real-time using Color Doppler mode and was comparable to direct pressure measurements performed with a Millar high-fidelity microtip catheter. Taken together, these data demonstrate that RV measurements obtained from various complimentary views using echocardiography are reliable, reproducible and can provide insights regarding RV structure and function. This method will enable a better understanding of the role of RV cardiac dysfunction.

Right ventricle (RV) dysfunction is critical to the pathogenesis of cardiovascular disease, yet limited methodologies are available for its evaluation. Recent advances in ultrasound imaging provide a noninvasive and accurate option for longitudinal RV study. Herein, we detail a step-by-step echocardiographic method using a murine model of RV pressure overload.

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, ...

Transcutaneous Assessment of Renal Function in Conscious Rodents

Glomerular filtration rate (GFR) is the gold standard to assess overall kidney function. However, traditional methods to evaluate GFR are cumbersome and time-consuming. In addition, serial blood or urine samples are required, with the associated stress for the experimental animals. A recent technique significantly reduces the investment in time and resources, minimizing the invasiveness and the animal stress, but being equally valid as the traditional approaches. The method measures transcutaneously renal function. Using an optical device and the exogenous renal marker fluorescein isothiocyanate (FITC)-sinistrin, this technique is capable of measuring the elimination kinetics of the marker through the skin. With neither blood nor urine samples nor the associated laboratory assays needed, the results of the transcutaneous measurement are almost instantaneously available. The method has been already validated in different species and successfully applied in several models of renal pathology. Moreover, due to its minimally invasive characteristics, it is suitable for sequential measurements within the same animal. Here is provided a detailed protocol to carry out the transcutaneous assessment of renal function in rodents.

Determination of glomerular filtration rate (GFR) is the gold standard to assess overall kidney function. However, traditional procedures to measure this parameter are cumbersome and require a large investment of time. Here we describe a faster and minimally invasive method to determinate GFR transcutaneously.

Glomerular filtration rate (GFR) is the gold standard to assess overall kidney function. However, traditional methods to evaluate GFR are cumbersome and ...

Preparation and In Vitro Characterization of Magnetized miR-modified Endothelial Cells

To date, the available surgical and pharmacological treatments for cardiovascular diseases (CVD) are limited and often palliative. At the same time, gene and cell therapies are highly promising alternative approaches for CVD treatment. However, the broad clinical application of gene therapy is greatly limited by the lack of suitable gene delivery systems. The development of appropriate gene delivery vectors can provide a solution to current challenges in cell therapy. In particular, existing drawbacks, such as limited efficiency and low cell retention in the injured organ, could be overcome by appropriate cell engineering (i.e., genetic) prior to transplantation. The presented protocol describes the efficient and safe transient modification of endothelial cells using a polyethyleneimine superparamagnetic magnetic nanoparticle (PEI/MNP)-based delivery vector. Also, the algorithm and methods for cell characterization are defined. The successful intracellular delivery of microRNA (miR) into human umbilical vein endothelial cells (HUVECs) has been achieved without affecting cell viability, functionality, or intercellular communication. Moreover, this approach was proven to cause a strong functional effect in introduced exogenous miR. Importantly, the application of this MNP-based vector ensures cell magnetization, with accompanying possibilities of magnetic targeting and non-invasive MRI tracing. This may provide a basis for magnetically guided, genetically engineered cell therapeutics that can be monitored non-invasively with MRI.

Preparation and In Vitro Characterization of Magnetized miR-modified Endothelial Cells

This manuscript describes the efficient, non-viral delivery of miR to endothelial cells by a PEI/MNP vector and their magnetization. Thus, in addition to genetic modification, this approach allows for magnetic cell guidance and MRI detectability. The technique can be used to improve the characteristics of therapeutic cell products.

To date, the available surgical and pharmacological treatments for cardiovascular diseases (CVD) are limited and often palliative. At the same time, gene ...

Assessment of Human Adipose Tissue Microvascular Function Using Videomicroscopy

While obesity is closely linked to the development of metabolic and cardiovascular disease, little is known about mechanisms that govern these processes. It is hypothesized that pro-atherogenic mediators released from fat tissues particularly in association with central/visceral adiposity may promote pathogenic vascular changes locally and systemically, and the notion that cardiovascular disease may be the consequence of adipose tissue dysfunction continues to evolve. Here, we describe a unique method of videomicroscopy that involves analysis of vasodilator and vasoconstrictor responses of intact small human arterioles removed from the adipose depot of living human subjects. Videomicroscopy is used to examine functional properties of isolated microvessels in response to pharmacological or physiological stimuli using a pressured system that mimics in vivo conditions. The technique is a useful approach to gain understanding of the pathophysiology and molecular mechanisms that contribute to vascular dysfunction locally within the adipose tissue milieu. Moreover, abnormalities in the adipose tissue microvasculature have also been linked with systemic diseases. We applied this technique to examine depot-specific vascular responses in obese subjects. We assessed endothelium-dependent vasodilation to both increased flow and acetylcholine in adipose arterioles (50 - 350 &#181;m internal diameter, 2 - 3 mm in length) isolated from two different adipose depots during bariatric surgery from the same individual. We demonstrated that arterioles from visceral fat exhibit impaired endothelium-dependent vasodilation compared to vessels isolated from the subcutaneous depot. The findings suggest that the visceral microenvironment is associated with vascular endothelial dysfunction which may be relevant to clinical observation linking increased visceral adiposity to systemic disease mechanisms. The videomicroscopy technique can be used to examine vascular phenotypes from different fat depots as well as compare findings across individuals with different degrees of obesity and metabolic dysfunction. The method can also be used to examine vascular responses longitudinally in response to clinical interventions.

Videomicroscopy systems are used to examine functional properties of isolated adipose tissue arterioles in response to physiological and pharmacological stimuli. This technique can be used to examine microvascular phenotypes in different adipose tissue domains in obese humans.

While obesity is closely linked to the development of metabolic and cardiovascular disease, little is known about mechanisms that govern these processes. ...

Normothermic Ex Situ Heart Perfusion in Working Mode: Assessment of Cardiac Function and Metabolism

The current standard method for organ preservation (cold storage, CS), exposes the heart to a period of cold ischemia that limits the safe preservation time and increases the risk of adverse post-transplantation outcomes. Moreover, the static nature of CS does not allow for organ evaluation or intervention during the preservation interval. Normothermic ex situ heart perfusion (ESHP) is a novel method for preservation of the donated heart that minimizes cold ischemia by providing oxygenated, nutrient-rich perfusate to the heart. ESHP has been shown to be non-inferior to CS in the preservation of standard-criteria donor hearts and has also facilitated the clinical transplantation of the hearts donated after the circulatory determination of death. Currently, the only available clinical ESHP device perfuses the heart in an unloaded, non-working state, limiting assessments of myocardial performance. Conversely, ESHP in working mode provides the opportunity for comprehensive evaluation of cardiac performance by assessment of functional and metabolic parameters under physiologic conditions. Moreover, earlier experimental studies have suggested that ESHP in working mode may result in improved functional preservation. Here, we describe the protocol for ex situ perfusion of the heart in a large mammal (porcine) model, which is reproducible for different animal models and heart sizes. The software program in this ESHP apparatus allows for real-time and automated control of the pump speed to maintain desired aortic and left atrial pressure and evaluates a variety of functional and electrophysiological parameters with minimal need for supervision/manipulation.

Normothermic ex situ heart perfusion (ESHP), preserves the heart in a beating, semi-physiologic state. When performed in a working mode, ESHP provides the opportunity to perform sophisticated assessments of donor heart function and organ viability. Here, we describe our method for myocardial performance evaluation during ESHP.

The current standard method for organ preservation (cold storage, CS), exposes the heart to a period of cold ischemia that limits the safe preservation ...

Echocardiographic Assessment of Cardiac Anatomy and Function in Adult Rats

The use of experimental animal models has become crucial in cardiovascular science. Most studies using rodent models are focused on two-dimensional imaging to study the cardiac anatomy of the left ventricle and M-mode echo to assess its dimensions. However, this could limit a comprehensive study. Herein, we describe a protocol that allows an assessment of the heart chamber size, left ventricular function (systolic and diastolic) and valvular function. A conventional medical ultrasound machine was used in this protocol and different echo views were obtained through left parasternal, apical and suprasternal windows. In the left parasternal window, the long and short axis were acquired to analyze left chamber dimensions, right ventricle and pulmonary artery dimensions, and mitral, pulmonary and aortic valve function. The apical window allows the measurement of heart chamber dimensions and evaluation of systolic and diastolic parameters. It also allows Doppler assessment with detection and quantification of heart valve disturbances (regurgitation or stenosis). Different segments and walls of the left ventricle are visualized throughout all views. Finally, the ascending aorta, aortic arch, and descending aorta can be imaged through the suprasternal window. A combination of ultrasound imaging, Doppler flow and tissue Doppler assessment have been obtained to study cardiac morphology and function. This represents an important contribution to improve the assessment of cardiac function in adult rats with impact for research using these animal models.

A non-invasive protocol for transthoracic echocardiography assessment of cardiac anatomy and function for adult rats is presented in the current study. The heart valves, all four cardiac chambers and the ascending aorta, aortic arch and descending aorta are studied in detail.

The use of experimental animal models has become crucial in cardiovascular science. Most studies using rodent models are focused on two-dimensional ...

Using a Chemical Biopsy for Graft Quality Assessment

Kidney transplantation is a life-saving treatment for a large number of people with end-stage renal dysfunction worldwide. The procedure is associated with an increased survival rate and greater quality of patient&#8217;s life when compared to conventional dialysis. Regrettably, transplantology suffers from a lack of reliable methods for organ quality assessment. Standard diagnostic techniques are limited to macroscopic appearance inspection or invasive tissue biopsy, which do not provide comprehensive information about the graft. The proposed protocol aims to introduce solid phase microextraction (SPME) as an ideal analytical method for comprehensive metabolomics and lipidomic analysis of all low molecular compounds present in kidneys allocated for transplantation. The small size of the SPME probe enables performance of a chemical biopsy, which enables extraction of metabolites directly from the organ without any tissue collection. The minimum invasiveness of the method permits execution of multiple analyses over time: directly after organ harvesting, during its preservation, and immediately after revascularization at the recipient&#8217;s body. It is hypothesized that the combination of this novel sampling method with a high-resolution mass spectrometer will allow for discrimination of a set of characteristic compounds that could serve as biological markers of graft quality and indicators of possible development of organ dysfunction.

The protocol presents utilization of the chemical biopsy approach followed by comprehensive metabolomic and lipidomic analysis for quality assessment of kidney grafts allocated for transplantation.

Kidney transplantation is a life-saving treatment for a large number of people with end-stage renal dysfunction worldwide. The procedure is associated ...

Semi-Minimal Invasive Method to Induce Myocardial Infarction in Rats and the Assessment of Cardiac Function by an Isolated Working Heart System

Myocardial infarction (MI) remains the main contributor to morbidity and mortality worldwide. Therefore, research on this topic is mandatory. An easily and highly reproducible MI induction procedure is required to obtain further insight and better understanding of the underlying pathological changes. This procedure can also be used to evaluate the effects or potency of new and promising treatments (as drugs or interventions) in acute MI, subsequent remodeling and heart failure (HF). After intubation and pre-operative preparation of the animal, an anesthetic protocol with isoflurane was performed, and the surgical procedure was conducted&#160;quickly. Using a minimally invasive approach, the left anterior descending artery (LAD) was located and occluded by a ligature. The occlusion can be performed acutely for subsequent reperfusion (ischemia/reperfusion injury). Alternatively, the vessel can be ligated permanently to investigate the development of chronic MI, remodeling or HF. Despite common pitfalls, the drop-out rates are minimal. Various treatments such as remote ischemic conditioning can be examined for their cardioprotective potential pre-, peri- and post-operatively. The post-operative recovery was quick as the anesthesia was precisely controlled and the duration of the operation was short. Post-operative analgesia was administered for three days. The minimally invasive procedure reduces the risk of infection and inflammation. Furthermore, it facilitates rapid recovery. The &#8220;working heart&#8221; measurements were performed ex vivo and enabled precise control of preload, afterload and flow. This procedure requires specific equipment and training for adequate performance. This manuscript provides a detailed step-by-step introduction for conducting these measurements.

This article presents an efficient method to perform myocardial ischemia and subsequent chronic reperfusion in rats using a minimally invasive approach. In addition, left ventricular hemodynamic function of rats is assessed by echocardiography and isolated working heart methods.&#160;

Myocardial infarction (MI) remains the main contributor to morbidity and mortality worldwide. Therefore, research on this topic is mandatory. An easily ...