I'm trying to isolate left ventricular myocytes from wildtype C57BL/6 mice. Currently the isolation step seems to be going okay - usually I get >50% viability of cells (although this is variable between isolations) however when I stimulate the cells they barely contract. The cells look viable - little spontaneous contraction, clear defined striations etc. They contract a tiny amount so I know the stimulation between the electrodes is working, but it's very little compared to previous single cell studies I did around a year ago with the same isolation protocol. Also recently I've used rat myocytes, prepared by another lab member, which showed far better contraction. I'm unsure what the problem could be - initially I thought perhaps the calcium concentration of the solutions - through the isolation they are perfused with Krebs-Henseleit (1mM CaCl2) , followed by low calcium (35 μM CaCl2) then enzyme solution (200μM). The cells are then spun down and re-suspended in enzyme solution (without collagenase & thermolysin) and DMEM is added step-wise to increase the calcium concentration to 0.84mM CaCl2. When the cells are under the microscope they are superfused with normal tyrode with 1mM CaCl2. Sometimes it looks like the cells initially contract a little, but the contraction quickly drops when stimulated at either 0.5Hz or 1Hz to negligible contraction (visibly very little and also zero Fluo-4 fluorescence).

Has anyone encountered this problem before or aware of something I could be doing wrong?

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