Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish

This protocol enables the researcher to monitor the activity and response of single sensory neurons to a controlled stimulation in an intact behaving vertebrae. This patch electrophysiology is a quick and direct way to record from the spiking activity of individual neurons in real time. And that provides better time resolution and sensitivity at a lower cost than most optical imaging techniques.

The lateral line hair cells are homologous to those of the human inner ear. So this technique is poised to reveal the properties of hair cell circuits that apply to human deafness and hearing disorders. Electrophysiology is a craft using fine motor skills which needs to be seen and physically imitated.

Text only instructions leave too much room for misinterpretation. To begin, dispense a thin layer of self-mixing silicone elastomer such as Sylgard into a cover glass bottomed tissue culture dish to make a silicone elastomer bottom recording dish. To make dissection pins, provide a negative charge of five volts to a 100 milliliter beaker of Etchant using a DC power supply and attach a tungsten wire to the positively charged output.

Repeatedly dip the tip of the wire into the Etchant bath until the tip narrows to a sharp point. Under a stereo microscope, cut the wire approximately one millimeter from the tip with a straight edge razor blade. Repeat this three times then insert pins into the cured recording dish using fine forceps.

To prepare recording electrodes, pull borosilicate glass capillary tubes using a micropipette puller with box filament. The electrode should have a 30 micrometer diameter tip with a taper that will be used to record afferent neurons from the posterior lateral line. Pull an additional borosilicate glass capillary tube into a pair of electrodes with smaller one to five micrometer tip diameters.

Holding one electrode in each hand, gently run the tips across one another to break the tips. Using a microforge, polish the beveled tip until smooth. The final tip diameter should be between 30 to 50 micrometers.

Immobilize zebrafish larvae and transfer them into a 35 millimeter petri dish using a large tip to transfer pipette. Remove as much of the surrounding solution as possible. Immerse larvae in 10 microliters of 0.1%alpha-Bungarotoxin for approximately five minutes.

Wash the paralyzed larva with extracellular solution for 10 minutes. Then, use a transfer pipette to move the larva from the extracellular solution bath to the silicone bottomed recording dish. Under a stereo microscope, gently position the larvae with fine tipped forceps above the center of this silicone mat, lateral side up with the body's anterior and posterior ends running left to right.

Using fine tipped forceps, insert the edge pin orthogonally to the silicone through the dorsal notochord of the larvae directly dorsal to the anus. Insert the second pin through the notochord near the end of the tail and insert the third pin through the notochord dorsal of the gas bladder. Insert the fourth pin through the otic vesicle while providing slight rotation as the pin inserts into the encapsulant.

As a slight rotation is applied, watch for the tissue between the cleithrum and otic vesicle to reveal the cluster of afferent somata. Place the pin larva under the 10 times objective on a fixed stage of the DIC microscope and orient the mioceptal clefts of the muscle blocks parallel to the left head stage vector. Place the ground wire into the bath solution and ensure that it is connected to the left head stage.

Fill the VR recording electrode with 30 microliters of extracellular solution using a flexible gel-loading pipette tip and insert it into the left head stage pipette holder. Increase the magnification to 40 times and lower the recording pipette into the dish solution while applying positive pressure produced by a pneumatic transducer. Bring the electrode tip over a myoseptum between the two myomeres ventral to the lateral line until the cleft is centered in the VR electrode tip aperture Lower the pipette until the lagging edge of the tip aperture gently contacts the epithelium.

After initial contact, maneuver the pipette diagonally to ensure the leading edge makes contact and can generate a seal. Apply negative pressure with the pneumatic transducer and hold it. In the patch clamp software, click on the Play button on the toolbar to monitor the VR signal.

Ensure that the VR recording is being achieved once motor neuron activity with Wells stereotyped burst signal dynamics are observed. Fill the afferent recording electrode with 30 microliters of extracellular solution and insert it into the right head stage pipette holder. Then lower it into the dish solution while applying positive pressure produced by a pneumatic transducer.

Locate the electrode and bring the electrode tip over the specimen. Using a micromanipulator, lower the afferent electrode tip until it is holding position above the cleithrum. Increase the magnification to 40 times immersion and locate the intersection of the posterior lateral line nerve and cleithrum.

Bring the electrode tip over the afferent ganglion and lower the pipette until the tip contacts the epithelium. Gently, maneuver the electrode so that the entire tip circumference contacts the afferent ganglion. Apply negative pressure with the pneumatic transducer and hold it.

After clicking Play in the patch clamp software, ensure that the whole cell loose patch recording of afferent neurons is achieved when spikes occur spontaneously roughly every 100 to 200 milliseconds. Once afferent neuron and motor neuron activity are both detected, click on the Record button on the toolbar in peak clamp 10 to capture simultaneous gap free recordings in both the channels. Perform data pre-processing as described in the text manuscript.

Using this gap-free recording protocol, the real-time activity of afferent and VR neurons can be measured simultaneously. Custom written, pre-processing scripts generate plots to assist in visualization of spike detection using parameters such as threshold, minimum duration and minimum interspike interval. Isolated signals were obtained by adjusting the lower bound and upper bound detection variables in the pre-processing script.

Ventral roots spike detection follows identical parameters with additional inputs. Bursts within a motor command are defined by a minimum of two spikes within 0.1 milliseconds lasting for five milliseconds at least and delineated by a minimum of three bursts with interburst intervals of less than 200 milliseconds Pre-processing scripts will overlay sections of afferent activity entered on a well-defined period of interest. In this case, mean spontaneous activity shows dramatic changes in response to the onset of motor activity depicted on the X-axis as time equal to zero.

Significant differences were detected in afferent spike rates between pre-swim spike rates and spike rates of both swim and post-swim periods. Afferent spike rates were negatively correlated with swim duration. There is no correlation detected between relative spike rate and swim frequency.

Animal health is the most important thing when trying to attempt this procedure. It is crucial to monitor fast blood flow to not only ensure animal welfare but also increase the likelihood of neural recording success. By leveraging genetic tools available in Zebrafish, This electrophysiology protocol can be complimented by transgenic lines to powerfully investigate anatomical and functional circuit connectivity in hair cell systems and beyond.

This simplified modification of the patch clamp technique allows us to monitor how sensory systems function in vivo.