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Encyclopedia of Experiments: Biology

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Zebrafish Electrocardiography (ECG)


Zebrafish Electrocardiography (ECG): A Minimally Invasive Assay to Evaluate Cardiac Function



- To begin, place an anesthetized fish ventral side up into a slit in a wet sponge. Insert the positive electrode into the fish's midline at the level of the bulbous arteriosis. Next, insert the negative electrode below the ventricle and 0.5 to 1 millimeter to the left of the midline to create a bipolar lead across which differences in electrical potential can be measured. Finally, insert the reference electrode near the anal region to ground the electrical signal.

Begin recording, and after a short time, review the trace to ensure that four criteria are met. All wave forms must be visible and distinct from each other. The P wave, which represents atrial depolarization, must be positive. The net sum of the Q, R, and S waves, which represents ventricular depolarization, must be positive. And finally, the T wave, which represents atrial repolarization, must be positive. If the criteria are not met, adjust the electrodes until a proper trace is recorded.

After recording, wake the fish by submerging it in oxygenated water and squirting water over the gills until the gills or the fish begin to move regularly. In the example, we will perform surface electrocardiography on an adult zebrafish.

- On the day of the experiment, transport the zebrafish from the aquarium to the laboratory. To set up the in vivo ECG recording system, connect the essential pieces of equipment and insert the three color-coded stainless steel electrodes of the ECG lead into the three color-matched access portals of the amplifier. For induction of level 4 anesthesia, immerse an adult zebrafish in a dish containing a solution of anesthetics at the lowest predetermined concentrations approved by the Institutional Animal Care and Use Committee.

Once the zebrafish has maintained level for anesthesia for three seconds, use a pair of blunt forceps to immediately transfer the fish onto a damp sponge with a slit ventral surface up for placement of the three ECG electrodes. Gently insert the positive electrode in the ventral midline at the level of the bulbous arteriosis 1 to 2 millimeters above an imaginary line connecting the two lower edges of the operculums.

Position the negative electrode caudally and 0.5 to 1 millimeter left laterally to the positive electrode at a distance greater than the maximal apical-basal length of the adult zebrafish ventricle. Then, position the reference electrode caudally near the anal region. For ECG recording, start the system and open the ECG data acquisition program. Select the desired setting from the dropdown menus for range, low pass, and high pass.

Press Start to initiate continuous gap-free ECG recording at a sampling rate of 1 kilohertz. To optimize lead positioning for maximal signal to noise ratio, press stop to stop the recording and review the trace soon after the very first recording attempt for each heart. If the ECG is expected to be normal, confirm that all ECG wave forms are distinct and readily visible and that the P wave, net QRS complex, and T wave are all positive.

- The single most critical step is lead positioning to maximize the signal to noise ratio. Apply all four validating criteria after the very first ECG recording attempt for each fish to obtain correct feedback.

- If a normal ECG is expected, reposition the electrodes as necessary until all of these four validating criteria are satisfied. If a normal T wave is expected but the T wave is too small, reposition the electrodes to maximize the T wave amplitude. Resume the ECG recording after optimizing the lead positioning saving the ECG sweeps for subsequent analysis.

At the end of the ECG recording session, carefully remove the electrodes without injuring the fish. In survival studies, transfer the fish to fresh, oxygenated fish water free of tricaine. Please note that in this video to facilitate readers viewing of the fish recovery from anesthesia, oxygenation is discontinued.

To facilitate recovery from the anesthesia in survival studies, use a Pasteur pipette to vigorously squirt water over the gills until the fish resumes regular gill movement or swimming. Then monitor the fish for full recovery from anesthesia before returning the fish to the aquarium. The fish is considered fully recovered from anesthesia when it can swim upright for at least 5 seconds.

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