A sogar coated cover slip made by smearing A drop of sogar across the glass and curing overnight is placed in the circular cavity of the prefabricated recording chamber and covered with recording solution. A worm placed in the center is glued to the sill guard surface starting from the head or tail applying glue to the dorsal side of the worm using a glue pipette. Once gluing is complete, an incision adjacent to the vulva at the interface between the cuticle and glue is made with a glass needle and extended towards the head following extraction of viscera and eggs from the body cavity.
The cut edge of the incision is spot welded with the glue to the sogar, exposing the ventral nerve and body wall muscles. The ventral medial muscle anterior to the vulva is patch clamped and the stimulating electrode is placed upstream on the nerve cord. Hello, I'm Janet Richmond, an associate professor in the Department of Biological Sciences at the University of Illinois Chicago.
Today I'm gonna show you a dissection technique which allows us to expose the neuromuscular junctions of sea elgan. For electrophysiological recording. We use this prep to study synaptic transmission.
Okay, let's get started. Before working with the nematodes, some hardware must be crafted at least one day before the procedure. A dissection recording chamber, S guard coated glass cover slips, old pipettes, a glue applicator, and an extraction pipette.
Begin by constructing the chamber. Cut a sheet of one 16th inch thick plexiglass to match the microscope stage. Used to do the recordings at the center of the plexiglass.
Drill out a circular hole large enough to accommodate a 22 millimeter diameter circular cover glass glue. A 48 by 60 millimeter cover glass to the bottom side of the chamber below the hole using a pellet of low melt para plaque tissue embedding wax in each corner of the cover glass. The wax is quickly melted on a hot plate, leaving a thin layer of wax between the plexiglass and cover glass slide after cooling, trim any excess wax with a blade.
The cover slip that fits the chamber is a 22 millimeter S guard coated cover glass. For sard to work, it must be made fresh according to the manufacturer's instructions. Once prepared, a small drop of sard is placed on each cover glass and smeared using a blade to make an even surface.
The coated cover slips are then placed in covered containers and placed in a 65 degree Celsius oven overnight to cure. Tomorrow they'll be ready for the dissection throughout the protocol. One millimeter outer diameter for a silicate glass pipettes pulled to a tip that has about a four mega resistance are required, so it is good to keep a jar available.
See the Joe Publication making patch, pipettes and sharp electrodes with a programmable polar by Miriam Goodman. For technical details to perform electrophysiology, the nematode must be glued in place. And for this, a glue applicator is required.
The glue applicator is simply a two millimeter plastic tube with an inner diameter of one millimeter. One end of the tube will be held in the mouth. It can be outfitted with a mouthpiece, and the other end of the tube will hold a one millimeter pipette that is filled with glue.
A specialized extraction pipette is also used during the dissection. This is made of a tube and pipette just like the glue applicator, except that the pipette is slightly broken back to accommodate the eggs and viscera. Begin by drawing a wax line along the perimeter of the chamber.
Now press and place a S guard coated cover slip syl guard upwards. The wax line immobilizes the cover glass and prevents the recording solution from seeping under and dislodging the cover slip during dissection and recording The chamber is then filled with extracellular recording solution. Read the accompanying text for the formulation with the chamber prepared.
Using a nematode pick to place several nematodes into the solution. With practice, the swimming nematodes can be glued down without any additional procedures to prevent movement. Load a glue container fashioned from a PCR cap held in wax or modeling clay with a working stock of the Sano acrylic glue to work with the glue.
Finish building the glue applicator by attaching your recording pipette to the end of the glue applicator. Then under the dissection scope, fill the glue applicator by mouth suction. Adhering the glue to a nematode is challenging and requires practice because the glue polymerizes on contact with extracellular recording solution.
To remedy this, maintain a steady outward flow of glue or it will harden and plug the pipette. If the pipette becomes plugged, it can sometimes be unblocked by gently tapping the tip on the cover slip. Practice working with the glue.
And when you can write your name in glue on the sil guard, you are probably ready to glue the nematode cuticle to glue a nematode. First, choose one near the center of the chamber where there is room to operate. Second, attach a small amount of glue to either the head or tail of the nematode and rapidly draw the nematode down onto the sil guard surface.
Then apply a stream of glue along the dorsal edge of the nematodes cuticle. This forces the nematode into a C position with the vulva facing inward. Any gaps in this glue line should be filled in for maximal nematode to syl guard adhesion.
This is a tricky process and it can help to practice on hypo motil mutants. Like on 31, There's a bulb right there just to the right. Now I'm gonna move this prep round and then I'm gonna bring in the the glass needle and start making the incision To make the cuticle incision switch to a handheld dissection pipette.
This pipette needs to be sharper than the glue recording pipettes and have a short, sturdy shaft using the highest magnification. On the dissecting scope, align the pipette parallel to the longitudinal axis of the nematode and insert the tip near the vulva at the cuticle glue interface. This incision releases the nematodes hydrostatic pressure, forcing eggs and intestines out through the incision point.
Continue cutting the cuticle towards the head of the nematode with a slicing motion similar to opening a letter until you reach the pharynx. Now switch over to the extraction pipette to vacuum out the internal organs of the nematode. After clearing the incision, the opened cuticle will retain its cylindrical shape.
Next, get a fresh gluing pipette. Use it to spot weld the cuticle at the edge of the incision onto the syl guard surface. It is important to use minimal glue spots at this stage to avoid damaging or obscuring the njs.
The ventral nerve cord and body wall muscles are now exposed, but the basement membrane that covers the NMJ must still be removed. To do this, suck off the extracellular recording solution from the chamber and apply an extracellular recording solution with at 0.4 milligrams per milliliter. After 10 to 20 seconds, remove the collagenase and wash the preparation several times with fresh recording solution.
The nematode is now ready for electrophysiological recordings. At the NMJ position, the recording chamber on an upright microscope stage using a 10 times objective to center the nematode switch to a 40 times water immersion objective with DIC optics and check that the body wall muscles and nerve cord are intact. Position the nematode horizontally so electrodes can be introduced from both the left and right sides.
Typically, we patch the ventral medial body while muscles interior to the vulva, but muscles on both the dorsal and ventral side of the worm can be readily patched With a couple of neurons that look like half circle semicircle up against the muscle. Now patch the muscle using standard patch clamping techniques to obtain a gig seal. In our experience recording electrodes of about four megas resistance work well for the whole cell voltage clamp recording mode.
Apply more suction and zap the membrane until the gigo seal is ruptured. So we talked about having a four mega resistance. So if you look at that there, Yeah, That says 4.9 megas.
It's bigger, normal. It's in the ballpark that we use.Okay.Yeah. Now we're gonna try and go big on, so what?
What? We want that to come all the way down.Yeah. Got it.Yay.
In a good recording body wall muscles of wild type adult nematodes typically have a cell membrane capacitance of approximately 70 picofarad and can be stably recorded for at least 10 minutes at a holding potential of negative 60 millivolts with minimal holding currents, approximately 50 pico amps. Once a muscle cell has been patched, stimulation from a second electrode light activated channel, Rodin or addition of hyperosmotic solution will all generate measurable responses. To elicit an electrically evoked response, we typically place a stimulating electrode containing extracellular solution on the ventral nerve cord interior to the patched muscle.
This loose patch configuration requires one to two millisecond pulses of a current between 10 and 40 volts. Unfortunately, after a few evoked responses, the nerve cord is irreversibly damaged and for no known reason, only cholinergic evoked responses can be elicited using this configuration. Alternatively, nematodes expressing a light activated channel where dosin transgene in different neuronal populations can be used to evoke release.
In preparation for this method, be aware that nematodes used for this application must be grown on bacterial laced with retinol. In this preparation, a high powered 470 nanometer LED filtered through A GFP excitation filter four 50 to four 90 nanometers is used to repeatedly activate the channel opsin causing evoked release of different neurotransmitters depending on which cell types express the channel. The last technique to review can be used to measure the size of the readily releasable pool of synaptic vesicles from several neurons onto the recorded muscle.
This is accomplished by pressure pulsing a hyperosmotic extracellular solution onto the patched muscle for one to three seconds. The result will be a barrage of asynchronous post-synaptic events. Whole cell voltage clamped body wall muscle recordings are shown for each of the three stimulation protocols described above, electrically evoked light evoked and hyperosmotic responses.
In all cases, the ventral medial body wall muscle is clamped at negative 60 millivolts. Wild type muscle cells typically exhibit high rates of endogenous inward miniature synaptic currents. In our standard recording solutions, a two millisecond depolarization of the anterior ventral nerve cord causes a synchronous release from several synapses resulting in a large evoked response of around two to 2.5 nano amps.
In wild type worms, representative traces to the right of the wild type traces in A and B show synaptic release in a severely uncoordinated mutant on 18 panel C shows a light evoked response from the neuromuscular junction of a wild type worm expressing channel red dosin. In cholinergic neurons, a two millisecond light pulse typically produces a response of 1.5 to 1.8 nano amps. Pressure ejection of 850 Milli osmo hyper osmotic saline onto the muscle for one second produces an asynchronous barrage of miniature synaptic currents representing the readily releasable pool of vesicles.
I've just shown you the dissection technique we use to expose the neuromuscular junctions of sea elegance and perform electrophysiological recordings. It's important to remember that this technique is challenging and it will likely take you months to perfect. However, once acquired, it's like riding a bike.
The techniques will stay with you, and the more you do it, the faster and more reliable the preparation will become. So that's it. Thanks for watching and good luck with your experiments.
