Protocol for Electrical Recordings in DHBs
Requirements: (in addition to those needed for making DHBs)
· Ground and stabbing Ag/AgCl electrode (treated with sodium hypochlorite)
· A white light microscope (4 – 10 x objective lens) with faraday cage and micromanipulator.
· Patch clamp amplifier (Axopatch 200B) with headstage, connected to computer via MiniDigi 1A. The computer will need either WinEDR V3.2.6 or Axoscope 10.2 software.
· A torch
Before you begin, switch the Axopatch 200B on to allow it to let it warm up for at least 30 min.
Ensure the headstage is resting flat in the Faraday cage (or other suitable heatsink).
Device preparation:
Prepare device for making DHBs as usual, but with the following differences:
· Electrically insulate the stage insert by sticking down a sheet of Parafilm with a hole cut out in the centre where the wells are.
· Stick the coverslip to the bottom of the device with Scotch tape, covering the entire perimeter. This will minimize evaporation of water from the substrate agarose which could cause an offset across the bilayer.
· Immediately after filling the device with substrate agarose, insert the ground electrode into the hole directly opposite the filling hole. Poke it through a ToughTag with a slit cut in it and stick this to the device around the electrode to minimize evaporation from this hole. The seal won’t be perfect but it will make a difference. You may need to pipette a little agarose around the electrode to keep it securely in place.
· Carefully rest the device on the insulated stage insert, and secure in place using tape.
Make bilayers as usual. Once bilayers are stable, you are ready to stab.
Prepare electrode:
This can be done while droplets are brewing
· Secure the stabbing electrode in the micromanipulator such that the wire is vertical. Check that the wire is able to reach the bottom of the wells, and that the faraday cage lid will shut in this position.
· Make a lighting-bolt shaped kink in the stabbing electrode – this will make the wire more steady when moving it in the Z-direction with the micromanipulator.
· Clean the stabbing electrode with EtOH then MQ and dry well. Coat the tip in agarose by pipetting ~10 µl molten 0.7% agarose onto a cover slip and dipping the very tip of the electrode in the pool before it sets. This will act as an anchor inside the droplet to prevent the electrode from sliding out.
Prepare software:
Using Axoscope 10.2 (Bay6):
Before you do any recording, you will need to do the following:
· Sampling rate: 1000 Hz (box on left hand side of main display window)
· File -> Set data file names – specify where you want your files to be saved, otherwise it can be tedious searching through someone else’s folder trying to identify your data. When you hit record, the trace will automatically be saved there. If you don’t hit record there is no way of getting traces obtained in “play” mode.
The following settings shouldn’t change at all, but if the values on the software don’t match the values on the patch clamp amplifier then it’s a good thing to check:
· Acquire -> Edit Protocol -> Acquisition Mode -> set to “Gap Free”
-> Trial Length -> set to “Use available disk space”
-> Sample rate -> set to 1000 Hz (the digitiser won't go any higher)
· Configure -> Lab Bench -> In #0 (current) set units to pA, set scale factor to 0.001
-> In #1 (voltage) set units to mV, set scale factor to 0.01
Using WinEDR (Bay 3 + 4):
Before you do any recording, you will need to do the following:
Prepare patch clamp amplifier:
Most of the fixtures on the front of the patch clamp amplifier aren’t relevant to doing electrophysiology in DHBs. Below are the important ones. Best to check they’re all set correctly before you begin recording.
· Offset set to 5 mV (you may need to adjust this once you start recording to counterbalance electrical drift)
· Under “Meter” switch to VHOLD/IHOLD to see applied potential on the display. Set it to a low value to begin with, so you can ramp it up once you start recording.
o This can be switched to “I” to check if the currect matches what’s shown on the computer screen
o IRMS tells you the root mean squared value of the current (a good indication of noise)
· Under “commands” section
o Switch to 1x (not 5x)
o Switch below that one to “OFF”
o External command to “OFF”
· Under “mode” select V-Clamp
· Under “Config” select Whole Cell β=1
· Set “output gain” to x1
· Set “Lowpass Bessel filter” to 1 kHz (This won’t actually make a difference in bay 6 because the digitization rate is already 1 kHz, so applying a finer filter is pointless)
· Set “Leak subtraction” to ∞
Stabbing a droplet
This can be tricky at first, and takes some practice. Only do this when everything else is ready.
· Position the electrode over the well containing the droplet you wish to stab and lower it using the micromanipulator until the tip touches the surface of the oil that fills the well.
· Bay 6 only: Turn off the microscope light and switch off at the mains (this will drastically reduce noise, and must be done before droplet is stabbed otherwise the jolt is likely to rupture the bilayer). In one hand, point the torch down at the device so that you can easily see the bilayer and a dark shadowy blob where the electrode is. You may need to adjust the angle the torch is held at, and move the electrode in the X-Y direction until you can clearly see where it is.
· Position the electrode over the centre of the droplet and slowly lower it downwards (turning the micromanipulator Z-control in a clockwise direction brings it down). You may need to adjust X and Y as the tip of the electrode becomes clearer.
· When you touch the top of the droplet with the electrode, the droplet will wobble very slightly. When this happens, slowly move the electrode in X or Y beyond the edge of the droplet. If the circular droplet is deformed by doing this, or the droplet translocated, then you are in. If not, move back into the centre of the droplet and go down a little further.