Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish

All animal care and experiments were performed in accordance with protocols approved by the University of Florida's Institutional Animal Care and Use Committee.

1. Preparation of materials for electrophysiological recordings

1. Make a silicone elastomer-bottomed recording dish.
   1. Dispense a thin layer of self-mixing silicone elastomer components (e.g., Sylgard) into a cover glass bottomed tissue culture dish until it levels with the rim of shallow well. Approximately 0.5 mL is sufficient.
   2. Cover and cure the dish for a minimum of 48 h at room temperature.
2. Make dissection pins.
   1. Provide a negative charge (5 V) to a 100 mL beaker of etchant (3M KOH) using a DC power supply and attach a tungsten wire (0.002 inch; 50.8 µm diameter) to the positively charged output.
      ​CAUTION: Negative and positive wires should not contact one another during this procedure as you may run the risk of producing sparks, which could pose a potential fire hazard.
   2. Etch tungsten wire by quickly and repeatedly dipping the tip of the wire into the etchant bath until the tip narrows to a sharp point. Under a stereomicroscope, cut the wire approximately 1 mm from the tip with a straight edge razor blade. Repeat three more times and then insert pins into cured recording dish using fine forceps.
3. Prepare recording electrodes.
   1. Pull a borosilicate glass capillary tube (inner diameter: 0.86 mm, outer diameter: 1.50 mm) using a horizontal micropipette puller with box filament into electrodes with a 30 µm diameter tip with slight taper (Figure 1 Ai) that will be used to record afferent neurons from the posterior lateral line.
   2. Pull an additional borosilicate glass capillary tube into a pair of electrodes with smaller tip diameters (1-5 µm). Holding one electrode in each hand, gently run the tips across one another to break them to a ~30° angle. Using a microforge, polish the beveled tip until smooth. The final tip diameter should be between 30-50 µm and will be used as the ventral root (VR) recording electrode (Figure 1 Aii).
   3. Mark the side of the VR electrode with the leading edge with permanent ink to aid in orienting the tip aperture downward when inserting the electrode into the pipette holder of the headstage (step 3.2).
      ​NOTE: Mechanical modification of the VR recording electrode is imprecise and step 1.3.2 may require multiple attempts until a suitable tip morphology is attained before polishing. VR recording electrode is beveled to conform to the body curve of the larvae. Once fabricated, the VR recording electrode can be used repeatedly as long as the tip remains clear and clean between experiments.
4. Generate the protocol in electrophysiology recording programs.
   1. Ensure the right headstage is connected to the Channel 1 input in the back of the microelectrode current and voltage clamp amplifier for the afferent neuron recordings and the left headstage is connected to the Channel 2 input for the VR recordings.
      NOTE: Computer specifications for the current and voltage clamp amplifier minimally require 1 Ghz or a better processor, Windows XP Pro or Mac OS X 10.46.6, CD-ROM drive with 512 MB RAM, 500 MB hard drive space, and 2 USB ports.
   2. Open the computer-controlled amplifier software.
   3. Set Channel 1 and Channel 2 to current clamp mode by clicking the IC button for each channel.
   4. Input the following parameters under both I-Clamp 1 and I-Clamp 2 tabs . Primary Output: 100x AC Membrane Potential (100,000 mV/mV), Gain: 1,000, Bessel: 1 kHz , AC: 300 Hz , Scope: Bypass . Secondary Output: 100x AC Membrane Potential (100 mV/mV) , Gain: 1 , Lowpass Filter: 10 Hz.
   5. Save channel parameters as Ch1_Aff and Ch2_VR.
   6. Install and open the patch clamp electrophysiology software.
   7. Click on Configure and select Lab Bench to set up the analog signals by adding Ch1_Aff and Ch2_Vr a Digitizer Channels (e.g., Analog IN #0 and Analog IN #1) that are connected, by BNC coaxial cable, to the corresponding Channel 1 and Channel 2 Scaled Outputs of the amplifier. Click on OK.
      ​NOTE: The minimum computer requirements for the digitizer are: a 1 Ghz or better processor, Windows XP Pro or Mac OS X 10.46.6, CD-ROM drive 512 MB RAM, 500 MB hard drive space, and 2 USB ports.
   8. Click on Acquire and select New Protocol.
   9. In the Mode/Rate tab, select Gap Free; under Acquisition Mode, set Trial Length to either Use available disk space (i.e., record until stopped) or a desired set Duration (hh:mm:ss), and then set the Sampling rate per signal (Hz) a 20,000 to maximize the resolution.
   10. Under the Inputs tab, select the Analog IN Channels that were previously configured (in step 1.4.6) and select Ch1_Aff and Ch2_VR for the corresponding channel.
   11. Under the Outputs tab, select Cmd 0 and Cmd 1 for Channel #0 and Channel #1, respectively.
       1. Connect Channel 1 Command on the amplifier to Analog Ouput 0 on the digitizer via BNC coaxial cable and repeat for Channel 2 Command to Analog Output 1.
   12. The remaining tabs can remain under default settings. Click on OK and save the protocol.

2. Solution preparation

1. Prepare Hank's solution: 137 mM NaCL, 5.4 mM KCl, 0.25 mM Na₂HPO₄, 0.44 mM KH₂PO₄, 1.3 mM CaCl₂, 1.0 mM MgSO₄, 4.2 mM NaHCO₄; pH 7.3. Dilute to 10% Hank's solution by adding the appropriate volume of deionized water to the stock.
2. Prepare extracellular solution: 134 mM NaCl, 2.9 mM KCl, 1.2 mM MgCl₂, 2.1 mM CaCl₂, 10 mM glucose, 10 mM HEPES buffer; pH 7.8 adjusted with NaOH. Vacuum filter extracellular solution through the filter with 0.22 µm pore size.
3. Prepare α -bungarotoxin: Dissolve 1 mg of lyophilized α-bungarotoxin in 10 mL extracellular solution to produce 0.1% dilution.
4. Prepare Euthanasia solution: 50% (mg/L) buffered pharmaceutical-grade MS-222 in 10% Hank's solution.
   CAUTION: α-bungarotoxin is a potent neurotoxin that paralyzes muscles by blocking cholinergic receptors. Gloves are required and eye protection is recommended while handling the paralytic.

3. Preparation of larvae for electrophysiological recordings

1. Immobilize zebrafish larvae.
   1. Use larvae from the laboratory-bred population of zebrafish (Danio rerio; 4-7 days post fertilization) and house in embryo solution (10% Hank's solution) at 27 °C.
   2. Transfer larva from housing into a small Petri dish (35 mm) using a large-tipped transfer pipette and remove as much of the surrounding solution as possible.
      NOTE: Removal of the embryo solution prevents dilution of the paralytic and increases its efficacy. The corner of a task wipe can be used to wick away the remaining embryo solution, and it is critical not to contact the larvae or leave larvae exposed to air.
   3. Immerse larvae in 10 µL of 0.1% α-bungarotoxin for approximately 5 min.
      NOTE: The time necessary to immobilize larvae varies between preparations. A healthy preparation depends on closely monitoring sustained fast blood flow and decreased motor responses. Brief overexposure to the paralytic can lead to a slow decline in overall health of the preparation even after a thorough washout. It is best to apply a wash before the larva is completely immobilized while it still shows signs of subtle muscle vibrations.
   4. Wash paralyzed larva with extracellular solution and bathe for 10 min.
      NOTE: The washout allows for the larva to transition from subtle muscle vibrations to complete paralysis. Also, α-bungarotoxin is an antagonist of the nicotinic acetylcholine receptor (nAChR) α9-subunit, which is a critical component of the endogenous feedback circuit this protocol observes; however, this effect has been shown to be reversible in Xenopus and zebrafish hair cells after a 10 min washout^36,29.
2. Pin fish in the recording dish.
   NOTE: Anesthesia (e.g., MS-222, Tricaine) is not required while preparing larval zebrafish (4-7 dpf) as it interferes with animal health. In fact, larval zebrafish are exempt from certain vertebrate protocols.
   1. Using transfer pipette, move the larva from the extracellular solution bath to the silicone-bottomed recording dish. Fill the remainder of the dish with extracellular solution.
   2. Under a stereomicroscope, gently position the larvae with fine-tipped forceps above the center of the silicone mat, lateral side up, with the body's anterior and posterior ends running left to right, respectively. Then grasp an etched pin from the silicone mat using fine-tipped forceps and insert the 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 (Figure 1B).
      NOTE: While inserting pins, it is important to target the center of the notochord width to prevent impinging blood flow or damaging surrounding musculature. The notochord is dorsal to the posterior lateral line nerve, so with proper pinning, no damage is expected to the lateral line sensory neurons. Insert after first contact as to not disturb the surrounding tissue. Ideally, the diameter of the pin is less than half the width of the notochord to ensure clean insertion. If the pins exceed this width, repeat step 1.2 until the desired pin width is attained. If the blood flow slows after pinning, repeat from step 1.3 onward with a new specimen.
   3. Insert the fourth pin through the otic vesicle while providing slight rotation towards the anterior as the pin inserts into the encapsulant (Figure 1B). As a slight rotation is applied, watch for the tissue between the cleithrum and otic vesicle to reveal the cluster of afferent somata.
      ​NOTE: Angled insertion of the fourth pin is to ensure the exposure of the posterior lateral line afferent ganglion, which is otherwise obstructed by the large otic vesicle.

4. Ventral root recording

1. Place pinned larva under the 10x water immersion objective on a fixed stage differential interference contrast (DIC) upright microscope and orient the myseptal clefts of the muscle blocks parallel to the left headstage approach vector (Figure 1B).
   1. A fixed stage DIC microscope floated on an optical air table works best to prevent vibrations from interfering with the recordings. Using a motorized positioner, the microscope can then move freely around the fixed stage in which the preparation and headstages are mounted (Figure 1C).
   2. Place the ground wire into the bath solution and ensure that it is connected to the left headstage.
2. Fill the VR recording electrode with 30 µL of extracellular solution using a flexible gel-loading pipette tip and insert into the left headstage pipette holder.
3. Lower the recording pipette into the dish solution with a micromanipulator while applying positive pressure (1-2 mmHg) produced by a pneumatic transducer. Under 10x magnification, confirm the orientation of the tip aperture is facing downward.
   1. The pneumatic transducer should be connected to the pipette holder port via silicone tubing.
4. Place the VR electrode onto the myoseptum
   1. Using the micromanipulator again, lower the VR electrode tip until it is holding position above the larva. Increase the magnification to 40x immersion.
   2. Bring the electrode tip over a myoseptum between two myomeres ventral to the lateral line until the cleft is centered in the VR electrode tip aperture (Figure 1D).
   3. 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.
   4. Apply negative pressure (~100 mm Hg) with the pneumatic transducer and hold.
      NOTE: Proper orientation of the VR pipette relative to the myoseptum and continuous negative pressure optimizes detecting motor neuron activity with a high signal-to-noise ratio through the skin.
5. Detect motor neuron activity.
   1. The left headstage should be connected to the amplifier, which relays the amplified signal into a digitizer that outputs the said signal into the patch clamp electrophysiology software to be monitored on an adjacent computer (see section 1.4).
   2. In the patch clamp software, click on the Play button on the tool bar to monitor the VR signal (Figure 1E).
   3. Ensure that the VR recording is being achieved once motor neuron activity with well-stereotyped burst signal dynamics are observed^29,37 (Figure 1E).
      ​NOTE: Fictive swim bouts are the activity patterns of the VR motor neurons that continue to transmit despite the preparation being paralyzed. Therefore, fictive swims are an accessible means of determining the behavioral state of the animal and measuring locomotor parameters while simultaneously performing afferent neuron recordings that require an immobilized preparation. In our hands, once a VR recording is achieved, a healthy preparation will elicit voluntary fictive swim bouts every few seconds. Remember, a healthy preparation has sustained fast blood flow. Be advised, obtaining a sufficient VR signal may take several minutes and the signal-to-noise ratio may improve after initial detection. In the interest of time, it is acceptable to move on to section 5.
   4. If a VR recording is still not achieved after completing section 5, release negative pressure, raise the electrode, and repeat from step 4.4.2 onward on a different myoseptum if the larva health is still optimal.

5. Afferent neuron recording

1. Fill the afferent recording electrode with 30 µL of extracellular solution; insert into the right headstage pipette holder (Figure 1B,C), and lower into the dish solution while applying positive pressure (1-2 mm Hg) produced by a pneumatic transducer.
2. Locate and loosely attach to posterior lateral line afferent ganglion.
   1. Using a micromanipulator, lower the afferent electrode tip until it is holding position above the cleithrum.
   2. Increase the magnification to 40x immersion and locate the intersection of the posterior lateral line nerve and cleithrum. Follow the lateral line nerve anterior from the cleithrum to where the fibers innervate the posterior lateral line afferent ganglion, distinguishable by the discrete cluster of soma (Figure 1F).
   3. 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.
   4. Apply negative pressure (20-50 mm Hg) with the pneumatic transducer and hold.
      NOTE: The negative pressure applied to the afferent ganglion is gentler than the suction applied during the ventral root recording. Increasing negative pressure can improve signal-to-noise, but afferent neuron health declines under sustained aggressive suction, which decreases the probability of a successful recording.
3. Record afferent neuron activity.
   1. Ensure that the right headstage is connected in a similar sequence as described in step 4.5.1.
   2. In pClamp10, click on the Play button on the tool bar to monitor afferent neuron and VR signal simultaneously.
   3. Ensure that the whole cell, loose patch recording of afferent neurons is achieved once spikes occur spontaneously, roughly every 100-200 ms^1,29 (Figure 1E).
   4. Gradually, increase the afferent neuron recording electrode pressure back to atmospheric (0 mm Hg) and hold for the remainder of the recording.

6. Data acquisition

1. Simultaneous recording.
   1. Once afferent neuron and motor neuron activity are both detected, click on the Record button on the tool bar in pClamp10 to capture simultaneous gap free recordings in both the channels.
   2. Record for the desired duration (Figure 1E).
      ​NOTE: A recording in a healthy preparation can last for many hours and remain responsive to external stimuli.
   3. Save the recording as a supported file type (.abf, pClamp10) to preserve metadata such as acquisition parameters.

7. Euthanasia

1. Apply positive pressure (10 mm Hg) to both afferent and VR recording electrodes using pneumatic transducers and raise the electrodes from the recording dish using the micromanipulators.
2. Transfer the recording dish from the fixed stage DIC microscope to the dissection stereomicroscope.
3. Using fine tip forceps, remove tungsten pins from notochord and otic capsule, and transfer the larvae using a transfer pipette into a Petri dish (35 mm) containing 5 mL of euthanasia solution for a minimum of 5 min.

8. Pre-processing and data-analysis

NOTE: Data pre-processing and analysis will require a basic understanding of command line coding.

1. Convert the recording file for pre-processing.
   1. Install the Matlab software.
   2. Download the custom written script, abfload.m³⁸ and save the file into the same folder that stores the raw recording file.
   3. In the Matlab Editor window, open abfload.m and click on Run on the tool bar. Select Change Folder, if prompted.
   4. Execute the function in the Command Window with the raw recording file as the input,
      > [d,si,h] = abfload('[raw recording file name].abf')
      and save the output workspace with a file name that includes larva designation, age, and experimental date such as [specimen number]_[age in dpf]_[raw recording file name (includes experimental date by default)].
      ​NOTE: The converted file name should only include underscore delineated integers.
2. Perform data pre-processing.
   1. Download Matlab script AffVR_preprocess.m and associated functions, custom written by the authors.
   2. In the Editor window, open AffVR_preprocess.m.
   3. Under Variables, adjust lower bound spike detection thresholds for both afferent and VR recordings (spk_detect_lb and vr_detect_lb, lines 8 and 14, respectively) depending on the recording of the signal-to-noise ratio.
      NOTE: Spike detection relies on manual thresholding to sort and isolate signals from more than one afferent neuron by amplitude. Baseline noise and activity from other units are filtered out by thresholding to only include spike amplitudes within a percent (e.g., spk_detect_lb) of the maximum (e.g., spk_detect_ub). Thresholding also ensures accurate binning of motor activity spikes into bursts and collective fictive swim bouts. Generally, start with a threshold lower bound of 0.5, and gradually decrease until accurate detection is achieved. For example, setting spk_detect_lb as 0.5 and spk_detect_ub as 1.0 will detect all spikes equal to or greater than 50% of the spike amplitude maximum and exclude noise or additional neurons of lower spike amplitudes (Figure 2A). Alternatively, one could also lower the spk_detect_ub from 1.0 to exclude neurons of higher spike amplitudes in order to isolate the neurons of lower spike amplitudes. Pre-processing figure outputs (see step 7.2.6) will inform whether adjustments and repeating from step 7.2.2 is necessary.
   4. In the Editor window, click on Run to execute AffVR_preprocess.m.
   5. Navigate to the previously converted .mat data file (see step 7.1.3); select and click on Open to begin automated preprocessing.
      NOTE: While the custom script is running, outputs will appear in the Command Window informing its progress through processing steps such as "filtering data …", "detecting ventral root activity …", and "generating figures …".
   6. Figures generated by AffVR_preprocess.m will visualize afferent spike and VR detection and indicate whether any analysis variables should be adjusted (Figure 2).
   7. Enter "Y" on the keyboard to save pre-processed metadata into a preprocess_output folder.
   8. Pre-processed data will automatically output as data_out.xls.
3. Analyze the pre-processed data as desired.