Intracellular recording and staining in Lymnaea

Rationale


The most detailed information about neural activity comes from intracellular recording. Not only does this technique show the pattern of neural action potentials, but it also records the cellular events which give rise to the action potential, including variations in the resting potential, changes in cellular impedance and synaptic potentials. Using this technique with manipulation of the composition of the saline (changing ionic concentrations or adding agonists or antagonists), the transmitters and ions responsible for these potentials can be identified. The intracellular injection of dyes is regularly used to elucidate the shape of particular nerve cells.

The neurons of the pond snail Lymnaea stagnalis are particularly large and are intensely colored (yellow/ orange/ red). The neurons survive well in a simple saline and are relatively robust. All these features simplify the methods required for successful intracellular recording. These techniques are described on the schedules below. Read these before you begin. Everyone has his (or her) own technique and it is not easy to explain why some students are more successful than others.

A lot of patience and practice is needed!
 

Safety

Intracellular recording requires skills with electrical apparatus, salines, sharps, optical apparatus and may involve toxic chemicals and lasers. I don't expect beginners to master all the skills at once, but I do expect everything to be done carefully. These notes are here to help, as a reminder of my oral instructions. Safety hints are given in the right hand boxes with details behind their links.

 

Dissection Technique

Extract a snail by crushing its shell with stout forceps. Under the microscope, pin the snail foot-down on the plain sylgard dish and cut it open along the dorsal mid-line. Pin back the body wall and identify the reproductive and digestive organs. (see figure) Keep the preparation moist with standard snail saline throughout.
Care with pins
Dissect away the digestive gland, the albumen gland (this coats the eggs) and kidney. Cut off the penis and spermatheca to reveal the oesophagus. This arises from the brown buccal mass: the anterior part is yellow but the upper part is black. Identify the red CNS (running in a circumoesophageal ring. Just anterior to the yellow salivary glands) and the nerves which supply the lips, tentacles, body wall, foot, etc.. Cut through the oesophagus near its origin and remove it. Increase the magnification. Dissect out the CNS, leaving the nerves as long as possible.

Pin the CNS out in the perfusion dish filled with fresh saline, dorsal side up. (You can tell the dorsal surface by the brown "dorsal bodies" on the cerebral ganglia.) The nerves are quite elastic and you should use them to anchor the preparation, like guy ropes. Try to angle the pins so that the tension in the nerves keeps the CNS preparation taut against the sylgard surface; if it can float away when you bring up an electrode you will have endless difficulties getting a second electrode in. Examine the CNS, and sort out which ganglion is which (see figures). Cut through the cerebral comissure to reveal the dorsal surface of the pedal ganglion. Pull away the white-spotted connective tissue with a pair fine forceps in each hand. You may need to remove some of the aorta which runs over the surface of the pedal ganglion. Don't squeeze the CNS tissue. Can you locate R.Pe.D.1 - the largest neuron in Lymnaea? Digest the ganglionic sheath with protease: scatter it on the surface of the bath to give a solution of 1 mg/ml. Wash thoroughly after 5 minutes.

Intracellular recording technique

Break the glass tubing and place it centrally in the electrode puller. Ask for help to load the puller until you are very familiar. If the filament looks bent or your tubing touches it Get Help at Once. If the electrodes are the wrong shape you have no hope of success  
Fill an electrode from the syringe provided with saturated K2SO4 or use 4M K acetate. Dip the tip in the black ink (Rotring ink 595617)  CAUTION: THIS INK IS CORROSIVE AND INDELIBLE!
Check the reference electrode (a pellet of Silver/Silver Chloride) is in the bath. Place the electrode in the electrode holder with the silver wire about half-way down the barrel. Use the micromanipulator controls to move the tip of the electrode so it just touches the saline solution. Look at it under the microsope and see if the tip looks broken; replace if it looks damaged Don't touch the wire with wet hands

Put used electrodes in the sharps bin

Turn on the computer and equipment, and log in. In Dasylab, read the voltage on the digital meter and adjust the blue knob labelled "D-V" until the digital meter reads nearly zero (the color of the reading should change to green).  
Measure the resistance of the electrode, by pressing the "R-TEST" button. This gives 150mV for every 10MW . Reject the electrode if the voltage change is below 15mV (probably broken) or more than 500mV (probably blocked with gunge, or wire not connected to the back of the electrode).  
Now advance the electrode towards your chosen cell very slowly, using the fine control of the micromanipulator. When the electrode reaches the cell, the trace will gently drift upwards (sometimes downwards, depending on the place and shape of the electrode). This is an artifact of the recording technique. Tap the micromanipulator very, very gently. Do you see a sudden drop in potential of 40-80mV? This is the resting potential. You may also see a train of action potentials, which gradually slow down. This is caused by the hole made when tapping, which quickly seals over.

If you don't get into a cell by tapping gently then the electrode may jump in if you press the "-C" button briefly. (This overbalances the capacitance compensation and causes electrical oscillations. This in turn, vibrates the tip of the electrode.)

 
If you don't get in:
  • What is the electrode resistance - this will show if your electrode is blocked or if the tip has broken off. More electrodes can be made easily - so don't be afraid to change the electrode.
  • If you don't get into the cells quickly, you may need more protease. If the cell you are poking starts to move off as you come up, you have used too much protease.
  • If the whole ganglion tries to move away, check its pinned well down with taut nerves!
  • If these suggestions don't help, ask for advice. (Today's free offer!!)
 

When you're in:

Getting into a cell is a great achievement! Does your cell show synaptic and action potentials? How does it respond to current injection ? Record these using the chart recorder (press "plot"  on the oscilloscope) or click the Record button on DasyLab. DasyLab will automatically record the trace, using today's date and time to identify the filename.

Staining neurons with Lucifer Yellow

Fill a fresh electrode by standing its barrel in the Lucifer Yellow solution (3% in distilled water). Penetrate your chosen cell - a small one will stain quicker! Set the pulse generator to produce c.500 ms pulses once per second and the level control to between -1 and -2 nA. Periodically check that you are still in the cell by turning off the injection current. Depending on its size, it will take 5-10 minutes to fill sufficient to see the main features and after 30-40 minutes, the cell will be well filled with Lucifer yellow. You can check this under the blue filter in the fiber optic light source. Slowly remove the electrode from the cell by winding back the micromanipulator.

Staining neurons with 5-6 carboxyfluorescein

Dissolve the 5-6 CF (Kodak) in distilled water and then add KOH until the ph is 7.9 - wacth for a color change. Use the method given above for Lucifer yellow

Viewing filled neurons

Replace the saline with 50% glycerol in saline and leave for 10 minutes. Place in 100% glycerol for another 10 minutes, then place on a microscope cavity slide in fresh glycerol. Cover with a cover slip, avoiding bubbles. Check you have time on the confocal miscroscope and examine the preparation on it with the 60x objective


Schedule devised by Chris Elliott on September 2, 1997; revised 15 July 2001; glycerol mounting add 27 oct 2004