Detachable glass microelectrodes for recording action potentials in active moving organs

Abstract

Here, we describe new detachable floating glass micropipette electrode devices that provide targeted action potential recordings in active moving organs without requiring constant mechanical constraint or pharmacological inhibition of tissue motion. The technology is based on the concept of a glass micropipette electrode that is held firmly during cell targeting and intracellular insertion, after which a 100-µg glass microelectrode, a "microdevice," is gently released to remain within the moving organ. The microdevices provide long-term recordings of action potentials, even during millimeter-scale movement of tissue in which the device is embedded. We demonstrate two different glass micropipette electrode holding and detachment designs appropriate for the heart (sharp glass microdevices for cardiac myocytes in rats, guinea pigs, and humans) and the brain (patch glass microdevices for neurons in rats). We explain how microdevices enable measurements of multiple cells within a moving organ that are typically difficult with other technologies. Using sharp microdevices, action potential duration was monitored continuously for 15 min in unconstrained perfused hearts during global ischemia-reperfusion, providing beat-to-beat measurements of changes in action potential duration. Action potentials from neurons in the hippocampus of anesthetized rats were measured with patch microdevices, which provided stable base potentials during long-term recordings. Our results demonstrate that detachable microdevices are an elegant and robust tool to record electrical activity with high temporal resolution and cellular level localization without disturbing the physiological working conditions of the organ.NEW & NOTEWORTHY Cellular action potential measurements within tissue using glass micropipette electrodes usually require tissue immobilization, potentially influencing the physiological relevance of the measurement. Here, we addressed this limitation with novel 100-µg detachable glass microelectrodes that can be precisely positioned to provide long-term measurements of action potential duration during unconstrained tissue movement.

Keywords: action potential duration; electrophysiology; glass micropipette electrodes; transmembrane potential.

Fig. 1.

Action potential (AP) signals were recorded from excised perfused unconstrained beating hearts using detachable sharp microdevices. A: the fire-polished end of a glass capillary tube (the “holding micropipette”) holds the sharp microdevice via gentle suction. The microdevice is akin to a “bee stinger” and has a thin silver wire that attaches to the headstage of the preamplifier. B: the holding micropipette, sharp microdevice, and stabilization fork are advanced toward the epicardial surface of a beating guinea pig heart. C: The microdevice was deployed to float within the epicardial tissue. The stabilization fork and holding micropipette have been retracted. D: five diagrams (i−v) showing the steps for implanting and releasing a sharp microdevice into myocardial tissue to record APs from a cardiac myocyte. E: a representative signal from a sharp microdevice floating within the left ventricular (LV) epicardium of a beating guinea pig heart. This signal exhibited stable and consistent APs.

Fig. 2.

AP duration (APD) was stable even if the AP amplitude drifted. A: instantaneous heart rate [in beats/min (bpm)] during normal sinus rhythm was measured from the ECG and is plotted above the simultaneously acquired AP signal (gray) generated by a sharp microdevice embedded in a guinea pig LV. APD measured for each beat is plotted in black on top of the AP signal. This image illustrates an extreme example of AP amplitude drift. These data address a potential limitation of microdevice recordings, which is that consistent AP amplitudes within the setting of significant contractile motion were sometimes difficult to achieve. However, for this 160-s signal, APD at 90% repolarization (APD₉₀) fluctuated no more than 4%, with a SD of 2.6%. B: even though amplitudes drifted by as much as 42% (A), the level of APD fluctuation was small compared with APD changes observed during metabolic perturbations, such as those shown in Fig. 3. C: histogram of RR intervals for the heart rate signal shown in A. This indicates that APD fluctuations could be attributed to subtle changes in heart rate.

Fig. 3.

APD measured from sharp microdevice signals during sinus rhythm reproduced the expected electrophysiological effects of ischemia and reperfusion. Instantaneous heart rate (in bpm) measured from the ECG of a perfused contracting rat heart is plotted above a simultaneously acquired 1,000-s AP signal generated by a microdevice embedded in the LV epicardium. APD measured for each beat is plotted in black on top of the AP signal. Specific time periods (impalement, normal perfusion, ischemia, and reperfusion) are indicated as different colors. APD₉₀ histograms for the normal perfusion, ischemia, and reperfusion periods are shown to reveal the correlation between coronary flow and temporal changes in APD.

Fig. 4.

AP signals recorded using sharp microdevices embedded within the contracting LV of three species. APD measured from these signals reproduced the effect of ATP-sensitive K⁺ (K_ATP) channel activation, either due to hypoxia or a K_ATP channel agonist (pinacidil). Bar plots denote averages ± SE. A paired t-test compared the APD of APs with those of control perfusion. *Significant differences (P < 0.05). A, D, and G: images showing representative hearts from a rat and guinea pig as well as a human LV wedge preparation. The black circle denotes the location of a microdevice. B and E: representative APs from a rat and guinea pig heart during 5 min of hypoxia. C and F: APDs progressively decreased (n = 4 and n = 6 for rats and guinea pigs, respectively, P < 0.05) after 2 min of hypoxia. H: representative APs from a human LV wedge preparation during a 9-min recording after pharmacological activation of K_ATP channels using pinacidil. I: APDs progressively decreased (n = 4, P < 0.05) after 3 min of administration of pinacidil.

Fig. 5.

Simultaneous measurements from two sharp microdevices and the bath-conducted ECG. A: image showing an excised perfused contracting rat heart and the three ECG electrodes. A microdevice embedded in the LV and another microdevice embedded in the left atrium (LA) are also shown. B: the ECG signal (top), APs recorded from the LA (middle), and APs recorded from the LV (bottom). The activation sequence of AP signals follows the morphology of the ECG. The R-R interval was 275 ms, the average LA APD₉₀ was 43.9 ± 0.85 ms, and the average LV APD₉₀ was 64.1 ± 0.95 ms.

Fig. 6.

AP signals were recorded from the hippocampus of anesthetized rats using patch microdevices whereby the sealing wax was melted to deploy the microdevice. A: each patch microdevice was held by inserting it into the end of a fire-polished glass capillary tube (the “holding micropipette”) and sealing the junction with wax. This maintained a contiguous pathway between the microdevice and the holding micropipette so that pressure or suction could be applied to the inside of the microdevice. B: Sequential images showing the release of a patch microdevice using a small heater coil to melt the wax seal. C: three diagrams (i−iii) showing the steps for implanting and releasing a patch microdevice into the brain to record APs from a CA1 hippocampal neuron. D: patch microdevice floating within the brain craniotomy of an anesthetized rat.

Fig. 7.

Representative AP signals recorded from the pyramidal CA1 layer of the hippocampus of a rat using a patch microdevice. Eight different neurons were successfully patched with a microdevice. A and C: whole cell patch recordings from two different CA1 layer neural cells. B and D: zoomed in views of APs denoted by the red boxes shown within the signal on the left. E and F: representative current injection recordings measured using the same microdevice and neurons that were patched to acquire the signals shown in A and C.