Neuroscience
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Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons
Chapters
Summary April 18th, 2017
This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.
Transcript
The overall goal of this preparation is to allow the electrophysiological study of zebrafish spinal neurons in situ at the embryonic and larval stages. The electrophysiological analyses compliment the powerful genetic and imaging approaches that are also possible with the zebrafish model. While the method was originally developed to study rohon beard neurons, we have easily adapted the approach over time and now we can study various types of spinal neurons.
Demonstrating the procedure will be Dr.Rosa Moreno, a research associate, with whom I've had the pleasure of collaborating for many years. To begin this procedure remove the black bulb from one glass bore, and replace it with a white rubber adaptor from another glass bore. Next, cut a piece of flexible tubing to a length of about 38 centimeters.
Then, attach the flexible tubing via a small, straight polypropylene fitting. To prepare the silicone elastomer add the base and the curing agent to a plastic conical tube at a ratio of four to one. Afterward, mix the elastomer base and curing agent thoroughly.
Pour the mixture into two 100 millimeter Petri dishes up to about one millimeter deep in one Petri dish and about 2.5 millimeters deep in the other. Allow the silicone elastomer to cure by exposure to air for four to five days. If cured elastomer is needed sooner, incubate it at 60 degrees Celsius.
Afterward, cut out a 3.8 centimeter by 6.3 centimeter rectangle from the one millimeter thick cured elastomer to serve as the bottom of the chamber. From the 2.5 millimeter thick cured silicone cut a rectangle of 3.8 centimeters by 6.3 centimeters. Then from that, cut out an internal rectangle of 2.5 centimeters by five centimeters to generate a frame for the top of the chamber.
To make the dissection recording chamber place the thin silicone rectangle directly on top of the glass slide, making sure to remove any air bubbles between the silicone and the glass. Then place the silicone rectangle frame on top of the thin bottom layer of silicone. In this procedure place an embryo in a dissection chamber containing two to three milliliters of ringer's solution.
Immobilize the embryo by adding 100 microliters of 0.4%tricaine solution to the chamber. Then pull some injection micro pipettes with long thin tips from the thin wall glass with a micro pipette puller. Attach a pulled glass micro pipette to the glass floor of the glue dispenser.
Under the dissecting microscope use tweezers to break the tip of the glass micro pipette resulting in a tip of about 75 micrometers in diameter. Now, load the glass micro pipette with three to five microliters of glue by applying suction through the mouth piece. Bring the glue filled tip to the dissection chamber.
Then bring the tip of the glue loaded micro pipette near the head of the embryo while maintaining slight positive pressure to the micro pipette via the mouth tube. Once the tip is near the head of the embryo, apply sufficient positive pressure to expel a small drop of glue onto the bottom of the chamber. Use a dissecting pin to move the embryo towards the drop of glue so that the head makes contact with the glue.
Orient the embryo dorsal side up, and press on the head to ensure good contact with the glue. As the glue slowly hardens use the dissecting pin to reposition the embryo so that it lies ventral side down, dorsal side up. Dip the dissecting pin into the drop of glue and drag a few threads of the glue over and across the head of the embryo to further secure its position.
Once the head is securely attached to the silicone and the glue has solidified sacrifice the embryo by transection at the hind brain level with another glass micro pipette. Use a fresh glass micro pipette to superficially cut the skin several times at a position caudal the hind brain. Pierce the skin superficially by moving the pipette perpendicular to rostro caudal axis on each side of the trunk for dorsally mounted specimens, or on the exposed side of the trunk for laterally mounted specimens.
To create a flap of skin for the tweezers to grab, scrape the skin several times with the micro pipette, at the level of and perpendicularly to the initial cut. Using tweezers, gradually lift the skin flap, and pull the skin caudally. After that, deliver a small drop of glue near the tail of the embryo.
Use this glue to attach the tail to the bottom of the dissection chamber. As the glue hardens, adjust the position of the trunk with the dissecting pin to ensure that the trunk remains dorsally oriented, and that it is firmly attached to the chamber. Following this initial dissection, rinse the preparation extensively with ringer's solution to remove the tricaine and debris, and allow the preparation to rest for five minutes.
Next, replace the dissection solution with extra cellular recording solution. If needed, add an immobilizing agent to the preparation. Next, transfer the dissection chamber with the mounted embryo to the stage of an upright compound microscope, equipped with a 40 X water immersion long working distance objective.
Then mount an empty bore of silicate thick wall glass micro pipette onto the electrode holder of the head stage. Attach the tubing at one end to the air outlet of the electrode holder and at the other end to a three way stop cock. After that, place a mouth piece at one end of another piece of tubing, and attach it to the three way stop cock at the other end.
Subsequently, bring the micro pipette tip to the most dorsal portion of the spinal cord, and gently pierce the meninges. Follow with swift, short, sideways movements to loosen the meninges. After the micro pipette tip has traversed the meninges advance and raise the micro pipette to pull the meninges away from the spinal cord.
Subsequently move the micro pipette rostorally advancing over one to two hemi segments to expose the rohon beard cells. In our experience, rohon beard neurons are more fragile than other spinal neurons. To maintain them healthy we expose one or two rohon beards at a time.
Voltage clamp recordings of outward and inward currents were obtained from the rohon beard neurons in one, two, and seven days post fertilization embryos and larvae. The holding potential was 80 millivolts, and the currents were elicited by a depolarizing step to positive 20 millivolts. In current clamp mode, 0.35 nano amp current injection for one millisecond elicits a single action potential from the rohon beard neurons in both embryos and larvae.
In the absence of electrical stimulation, rohon beard neurons do not show spontaneous spiking. The resting potential of the rohon beard neurons becomes increasingly more negative as the development progresses. Once this method is mastered, embryos may be mounted and dissected to expose rohon beards within 10 minutes.
As mentioned in the protocol, it is important to understand the factors that determine how quickly the glue hardens. This video highlights the critical steps for recording from spinal neurons in zebrafish.
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