Friday, October 7, 2011


Last month, I had my first experience with patch clamping neurons. In the neuro program here, the two weeks before the official quarter begins are spent in boot camp. The graduate students in this interdisciplinary field tend to come from various backgrounds, from electrical engineering to psychology to molecular biology. Boot camp, then, is meant to get everybody on the same page about certain important neuroscience concepts and techniques.

Neurons communicate with each other through the use of electrical signals. If you recall, a cell membrane is hydrophobic in the middle, which means that water and charged particles cannot pass through. A neuron uses this property to maintain a voltage across its membrane - it pumps certain ions in and certain ones out in such a way that there are more negatively-charged ions inside than outside, meaning the inside of the cell is more negative than the outside.

The electrochemical gradient of the sodium ion (Na+) depends on both the concentration and the electrical potential difference between the inside of the cell, and is set up in such a way that if a path across the membrane opens up, these cations will immediately flow inward. When a neuron is signaling, sodium-selective channels open and create that path through the membrane. This causes the inside of the neuron to become more positive than the outside, flipping the sign across  the membrane; this is the beginning of an action potential. There are certain channels that only open up when the voltage change passes a certain threshold, but these allow neighboring regions of the neuron to pass sodium as well, such that this flip of voltage is passed down the length of the axon. Potassium (K+) channels open with a slight delay and do the opposite, so the membrane voltage is flipped back slightly past normal in a wave following the original depolarization.

In patch clamping, you start with a tiny microelectrode that consists of a very thin pipette containing a conducting solution, whose tip is on the order of 2-3 microns (1/50th of a millimeter). Using a control box called a micro-manipulator, you watch through a high-power microscope and move the electrode in increments of less than a micron until you can rest it on the surface of the cell you're trying to record from. Cell membranes are flexible, and if you've done this right, the membrane should stick to the outside of your pipet and form a seal across the open tip.

Now, for a whole-cell recording, you have to break into the cell. You want to break open the patch of membrane across the open tip without tearing the membrane away from the outside of the pipet, so that you have the only hole into the cell. This requires a most advanced form of technology, on par with expensive microscopes and carefully pulled microelectrodes: your mouth.

No, seriously. Electrophysiology is still a developing technique, and there is not yet that much standardization between labs. In order to break the membrane, you want to apply suction in a small pulse (or multiple pulses, depending on the lab's preferred technique). Doing this by mouth is actually pretty accurate once you get the hang of it, though sometimes you end up sucking large portions of the membrane into the pipette before you ever break it open. Luckily, the thinness of the tip means that even if you apply a lot of suction, only a small portion of that will be evident to the cell, and it does take a surprisingly large amount of suction to break in.

So now you're in, and thanks to slightly more modern technology, you can use your single electrode to clamp the voltage or the current at a set amount while simultaneously recording the current or voltage, respectively. And then you hope that this cell is not leaky, not dying, and will actually have an interesting response to stimulation. That's how science goes.

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