We thank Ms. LourdesCarreon G Tan for her help maintaining the medaka facility and Anthony Peltier for the illustrative figures. This work was funded by NMBU and by the Research Council of Norway, grant numbers 244461 (Aquaculture program) and 248828 (Digital Life Norway program).

All animal handling was performed according to the recommendations for the care and welfare of research animals at the Norwegian University of Life Sciences, and under the supervision of authorized investigators.
1. Preparation of Instruments and Solutions
NOTE: All solutions should be sterile. Careful attention should be given to the pH and the osmolality (osmol/kg water) of all solutions, which should be carefully adapted to the extracellular environment of the studied species. pH and osmolality should be adjusted with precise electronic equipment such as pH meter and freezing point osmometer respectively.
<ol>
	<li>Make 500 mL of extra-cellular solution without Ca2+ (Ca2+ free EC): 150 mM NaCl, 5 mM KCl, 1.3 mM MgCl2, 4 mM glucose, 10 mM Hepes.
	<ol>
		<li>Dissolve 4.38 g of NaCl, 186.37 mg of KCl, 61.89 mg of MgCl2, 360.32 mg of Glucose and 1.19 g of HEPES in 450 mL of ultrapure (0.055 uS/cm at 25 &#176;C, the resistivity of 18.2 MOhm) H2O in a glass beaker using a magnetic agitator.</li>
		<li>Adjust the pH to 7.75 with NaOH.</li>
		<li>Transfer the solution into a 500 mL volumetric flask. Fill the volumetric flask with ultrapure H2O until 500 mL.</li>
		<li>Mix the solution well before measuring the osmolality. Adjust to 290&#8211;300 mOsm with mannitol. Filter sterilize the solution using water vacuum system and 0.2 &#181;m filter.</li>
	</ol>
	</li>
	<li>Make 500 mL of extra-cellular solution (EC): 150 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1.3 mM MgCl2, 4 mM glucose, 10 mM Hepes.
	<ol>
		<li>Dissolve 4.38 g of NaCl, 186.37 mg of KCl, 147.01 mg of CaCl2, 61.89 mg of MgCl2, 360.32 mg of Glucose and 1.19 g of HEPES in 450 mL of ultrapure H2O in a glass beaker using a magnetic agitator.</li>
		<li>Adjust the pH to 7.75 with NaOH.</li>
		<li>Transfer the solution into a 500 mL volumetric flask. Fill the volumetric flask with ultrapure H2O until 500 mL.</li>
		<li>Mix the solution well before measuring the osmolality. Adjust to 290&#8211;300 mOsm with mannitol. Filter sterilize the solution using water vacuum system and 0.2 &#181;m filter.</li>
	</ol>
	</li>
	<li>Make 500 mL of extra-cellular solution with 0.1% bovine serum albumin (EC with BSA): 150 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1.3 mM MgCl2, 4 mM glucose, 10 mM Hepes.
	<ol>
		<li>Dissolve 4.38 g of NaCl, 186.37 mg of KCl, 147.01 mg of CaCl2, 61.89 mg of MgCl2, 360.32 mg of Glucose and 1.19 g of HEPES in 450 mL of ultrapure H2O in a glass beaker using a magnetic agitator.</li>
		<li>Adjust the pH to 7.75 with NaOH.</li>
		<li>Transfer the solution into a 500 mL volumetric flask. Fill the volumetric flask with ultrapure H2O until 500 mL.</li>
		<li>Mix the solution well before measuring the osmolality. Adjust to 290&#8211;300 mOsm with mannitol. Filter sterilize the solution using water vacuum system and 0.2 &#181;m filter. Add 50 mg BSA.</li>
	</ol>
	</li>
	<li>Make 250 mL of intracellular solution (IC): 110 mM KOH, 20 mM KCl, 10 mM Hepes, 20 mM sucrose.
	<ol>
		<li>Dissolve 1.54 g KOH, 327.75 mg of KCl, 595.75 mg of HEPES and 1.712 g of sucrose in 450 mL of ultrapure H2O in a glass beaker using a magnetic agitator.</li>
		<li>Adjust the pH to 7.2 with (N-morpholino) ethane sulfonic (MES) acid.</li>
		<li>Transfer the solution into a 250 mL volumetric flask. Fill the volumetric flask with ultrapure H2O until 250 mL.</li>
		<li>Mix the solution well before measuring the osmolality. Adjust to 280&#8211;290 mOsm (10 mOsm lower than EC) with sucrose. Filter sterilize the solution using water vacuum system and 0.2 &#181;m filter.</li>
	</ol>
	</li>
	<li>Make 100 mL of 2% low melting agarose solution in Ca2+ and BSA free EC (2 g of low melting agarose in 100 mL of Ca2+ free EC). Dissolve using a microwave and place the melted agarose in a water bath at 40 &#176;C. Let it cool down until it reaches 40 &#176;C before use on the tissues. 
	NOTE: EC, Ca2+ free EC may be stored at 4 &#176;C for several weeks if sterile and IC solutions at -20 &#176;C, for several months. Agarose can be stored for a week at 4 &#176;C and can be reused 3&#8211;5 times by briefly microwaving. Excessive reuse will lead to evaporation and alteration of agarose and salt concentration and quality.</li>
	<li>Make agar bridge:
	<ol>
		<li>Heat 7.5 mm long borosilicate glass capillaries with outside diameter (O.D.) 2 mm and inside diameter (I.D.) 1.16 mm using a Bunsen burner at a focal point about 1/3 from one end until the glass starts to bend. Remove the glass from the flame and allow the glass to bend with an angle of about 120 degrees (Figure 2A).</li>
		<li>Melt 2% agar in normal EC without glucose using a microwave, and while the agarose is liquid fill the pre-made glass capillary using capillarity forces by putting one of the ends of the glass into the liquid and waiting for few minutes. Store bridges until use at 4 &#176;C in EC without glucose.</li>
	</ol>
	</li>
	<li>Prepare borosilicate glass with filament patch pipettes using a suitable pipette puller. Refer to the pipette puller manual to obtain the desired electrode resistance. 
	&#8203;NOTE: The taper of the pipette should be as short as possible. Short pipette taper is achieved by using 4&#8211;6 pulling steps. The electrode resistance should be between 2 and 6 M&#8486; depending on cell size.</li>
	<li>To avoid movement of tissue during the patch-clamp experiments coat the surface of the recording chamber with 0.1% of polyethylenimine (PEI) in borate buffer.
	<ol>
		<li>Prepare 500 mL of borate buffer (25 mM):
		<ol>
			<li>Dissolve 4.768 g of Na2B4O7/10H2O in 450 mL of ultrapure H2O in a glass beaker using a magnetic agitator.</li>
			<li>Adjust pH to 8.4 with HCl.</li>
			<li>Transfer the solution into a 500 mL volumetric flask. Fill the volumetric flask with ultrapure H2O until 500 mL and 0.2 &#181;m filter sterilize the solution using water vacuum system.</li>
		</ol>
		</li>
		<li>Prepare a 1% stock solution of PEI (1 mL 50% PEI to 49 mL 25 mM borate buffer) and a 0.1% PEI dilution by diluting 10 &#181;L of 1% stock solution in 100 &#181;L of borate buffer for final use.</li>
		<li>Add 2 mL of 0.1% PEI on the glass of the slice holder and let the coating incubate for 1 minute. Then briefly wash the glass twice with 5 mL of ultrapure H2O and let air dry until use.</li>
	</ol>
	</li>
	<li>Prepare amphotericin B solutions.
	<ol>
		<li>Make 60 mg/mL stock solution by dissolving 3 mg of amphotericin B powder in 50 &#181;L dimethyl sulfoxide (DMSO), in a 1 mL tube protected from light. Vortex at the maximum speed for 30 s and sonicate for 15 min to homogenize. Store aliquots into 3&#8211;5 tubes at -20 &#176;C and use one aliquot per day. 
		NOTE: If the amphotericin B stock solution is not clear and yellow color but milky (still yellow) after 15&#8211;20 min sonication make a new stock solution.</li>
		<li>Make IC solution with amphotericin B by pipetting 2 mL of the IC into a clean glass beaker with more than 10 mL capacity. Add 8 &#181;L of amphotericin B stock solution into the pipette solution and mix by pipetting. Wrap the beaker with aluminum foil and place on ice until use and renew it every 3 h.</li>
	</ol>
	</li>
</ol>
2. Dissection and Slicing with Vibratome
<ol>
	<li>Prepare the dissection tools before dissecting, including one sharp and one strong forceps with scissors. Clean the tools (tissue holder, brush, forceps) with ethanol and make sure that they are used only for dissection purposes and never in contact with fixed tissues. Keep them in a separate box to avoid contamination.</li>
	<li>Euthanize the fish in cold ice water for 1 min.</li>
	<li>Follow the subsequent steps to quickly dissect the medaka brain and pituitary with sharp forceps (use no more than 3 min).
	<ol>
		<li>While holding the fish with the strong forceps severe the two optic nerves behind the eyes with scissors or sharp forceps.</li>
		<li>Open the dorsal part of the skull from the posterior to the anterior side by breaking the skull bones step by step with strong forceps.</li>
		<li>Peel off the skull on one side with strong forceps.</li>
		<li>Severe the spinal cord with scissors or sharp forceps.</li>
		<li>Gently, grab the spinal cord with the forceps, flip the brain from posterior to anterior and place it into a dish with the ice-cold Ca2+ free EC.</li>
	</ol>
	</li>
	<li>Fill a 1.5&#8211;2 cm3 metal mold with liquid agarose and place it on the ice.</li>
	<li>Just before the agarose solidifies (at around 25&#8211;30 &#176;C), gently grab the brain by the spinal cord with the forceps, dry the forceps with a piece of fine paper and put the tissue into the agarose. Quickly mix the agarose to dilute the traces of EC left around the tissue and orientate the tissue and let the mold on ice for a few seconds until the agarose hardens. 
	NOTE: It is essential for this step to be fast. The agarose solidifies quickly as the metal mold is cooled down by the ice.</li>
	<li>Trim a square block of agarose containing the tissue with a scalpel blade and glue it on the vibratome specimen holder using surgery (non-toxic) glue.</li>
	<li>Fill the cuvette of the vibratome receiving the specimen (brain-pituitary) holder with ice-cold Ca2+ free EC.</li>
	<li>Place the specimen holder into the vibratome and cut the agarose block to make parasagittal sections of 150 &#181;m, using high frequency and the low speed for the sectioning. Collect the selected sections and place them into the patch-clamp recording chamber with 3 mL of ice-cold Ca2+ free EC.</li>
	<li>Place the grid harp (Figure 2B) on the tissue section. 
	NOTE: The harp will together with the coating stabilize the tissue and prevent it from moving during the patch-clamp recordings.</li>
	<li>After the harp is in place, change the Ca2+ free EC medium to the normal EC with Ca2+ and BSA. Let the tissue rest for 10 min.</li>
</ol>
3. Perforated Patch-clamp and Electrophysiological Recordings
<ol>
	<li>Before starting the experiment, chloride the silver wire electrodes by the following steps: First, use a fine sandpaper (p180) to clean the silver wire electrodes. Second, rinse with 2 mL, 70% ethanol. Third, dip the wires in 2 mL, 2&#8211;5% chlorine for 10 min. Finally, rinse with 2 mL, ultrapure H2O.</li>
	<li>Place the recording chamber with the brain-pituitary slice in the microscope stage and locate the target area (pituitary) using 10X objective and inspect the cells using 40X objective. 
	NOTE: If the preparation is successful, very few round cells should be visible. These round cells are usually damaged cells that are detaching from the tissue.</li>
	<li>Place the grounding electrode through the 2% agar bridge into the EC bath.</li>
	<li>Locate a healthy cell using the 40X objective. 
	NOTE: A healthy cell should be firmly attached to the tissue slice and not rounded up and detached from the slice.</li>
	<li>Set the amplifier in voltage-clamp (VC; Figure 4A point 1) mode. Open the seal test window and press the bath configuration (Figure 4B point 1 and 2. Use a 5-mV pulse to monitor pipette resistance (Figure 4B point 3).</li>
	<li>Backfill the tip of the patch pipette with the antifungal free IC solution before adding the IC solution with amphotericin B using a micro-filler syringe (see Figure 3).</li>
	<li>Add a slightly positive air pressure into the patch pipette using 1 mL syringe. 
	NOTE: This makes a small liquid flow from the patch pipette that avoid contamination of the tip.</li>
	<li>Once in the bath, assess the patch pipette resistance and make sure no particles are attached to the tip of the pipette (Figure 4C). 
	NOTE: A small piece of tape is attached onto the monitor displaying the video recording form the field of view. This makes the positioning of the patch pipette relative to cell easier when the objective is not focusing on the cell.</li>
	<li>Guide the patch pipette down to the cell using micromanipulators. 
	NOTE: Make sure that the pipette is clean by looking for small particles that may have attached to the tip of the pipette. Sudden changes in resistance may also be a result of particles stuck in the pipette tip.</li>
	<li>Readjust the pipette when a few microns above the cell so that the tip of the pipette will touch the cell 1/3 from the middle of the cell, as shown in Figure 5. When touching the cell, release the pressure and apply the gentle suction to make a seal. 
	NOTE: The time from the patch pipette enters the bath to a successful seal should be kept as short as possible. Not more than 1&#8211;1.5 min in order to avoid amphotericin B reaching the tip of the pipette and leak into the bath.</li>
	<li>Immediately switch to the patch window in the patch-clamp software (Figure 4D point 1) and have a holding potential between -50 and -60 mV (Figure 4D point 2). Zero out the fast capacitance made up by the glass pipette (Figure 4D point 3).</li>
	<li>Switch to the cell window in patch-clamp software (Figure 4E point 1 and 2) and start monitoring access resistance, Ra, displayed on the window. Zero out the membrane capacitance in the amplifier software (Figure 4E point 3) after sufficient access. 
	NOTE: Sufficient access might take 10 to 30 min (Figure 4E point 2). A good access for current clamp recordings should be below 20 M .</li>
	<li>Switch to current clamp, IC, in amplifier software window (Figure 4F point 1). Adjust the fast-capacitive currents in the I-clamp window of the amplifier software (Figure 4F point 2 and 3) to the same value as in 3.12. Check membrane potential (Figure 4F point 1). 
	NOTE: Importantly, if the membrane potential is shallow (typical above -35 mV) the cell might be damaged. Cancel the recording and find a new cell.</li>
	<li>After finding a healthy cell (firmly attached to the tissue with membrane potential below -40 mV), start the experimental recordings as detailed in the manufacturer&#39;s protocol 33,34.</li>
</ol>

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<li>Wittbrodt, J., Shima, A., Schartl, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Medaka--a+model+organism+from+the+far+East%5BTitle%5D)+AND+%22Nat+Rev+Genet%22%5BJournal%5D)">Medaka--a model organism from the far East.</a> Nat Rev Genet. 3 (1), 53-64 (2002).</li>
<li>Ball, J. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Hypothalamic+control+of+the+pars+distalis+in+fishes%2C+amphibians%2C+and+reptiles%5BTitle%5D)+AND+%22Gen+Comp+Endocrinol%22%5BJournal%5D)">Hypothalamic control of the pars distalis in fishes, amphibians, and reptiles.</a> Gen Comp Endocrinol. 44 (2), 135-170 (1981).</li>
<li>pCLAMP 10 Data Acquisition and Analysis For Comprehensive Electrophysiology User Guide. <a target="_blank" href="NULL">Molecular Devices Corporation</a>. , (2006).</li>
<li>MultiClamp 700B COMPUTER-CONTROLLED MICROELECTRODE AMPLIFIER Theory and Operation. <a target="_blank" href="NULL">Axon Instruments / Molecular Devices Corp</a>. , (2005).</li>
<li>Dominguez-Mancera, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Leptin+regulation+of+inward+membrane+currents%2C+electrical+activity+and+LH+release+in+isolated+bovine+gonadotropes%5BTitle%5D)+AND+%22Biochem+Biophys+Res+Commun%22%5BJournal%5D)">Leptin regulation of inward membrane currents, electrical activity and LH release in isolated bovine gonadotropes.</a> Biochem Biophys Res Commun. 491 (1), 53-58 (2017).</li>
<li>Schmidt-Nielsen, K. <a target="_blank" href="https://books.google.com/books/about/Animal_Physiology.html?id=Af7IwQWJoCMC">Animal Physiology : adptation and environment fifth edition</a>. , Cambridge university press. 613(1997).</li>
<li>Burton, R. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Evolutionary+determinants+of+normal+arterial+plasma+pH+in+ectothermic+vertebrates%5BTitle%5D)+AND+%22J+Exp+Biol%22%5BJournal%5D)">Evolutionary determinants of normal arterial plasma pH in ectothermic vertebrates.</a> J Exp Biol. 205, Pt 5 641-650 (2002).</li>
<li>Burton, R. F. <a target="_blank" href="https://www.sciencedirect.com/science/article/pii/0300962995021027">The dependence of normal arterial blood pH on sodium concentration in teleost fish.</a> Comparative Biochemistry and Physiology Part A: Physiology. 114 (2), 111-116 (1996).</li>
<li>Heming, T. A., Blumhagen, K. A. <a target="_blank" href="https://www.sciencedirect.com/science/article/pii/0166445X88900306">Plasma acid-base and electrolyte states of rainbow trout exposed to alum (aluminum sulphate) in acidic and alkaline environments.</a> Aquatic Toxicology. 12 (2), 125-139 (1988).</li>
<li>Reeves, R. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+interaction+of+body+temperature+and+acid-base+balance+in+ectothermic+vertebrates%5BTitle%5D)+AND+%22Annu+Rev+Physiol%22%5BJournal%5D)">The interaction of body temperature and acid-base balance in ectothermic vertebrates.</a> Annu Rev Physiol. 39, 559-586 (1977).</li>
<li>Baicu, S. C., Taylor, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Acid-base+buffering+in+organ+preservation+solutions+as+a+function+of+temperature%3A+new+parameters+for+comparing+buffer+capacity+and+efficiency%5BTitle%5D)+AND+%22Cryobiology%22%5BJournal%5D)">Acid-base buffering in organ preservation solutions as a function of temperature: new parameters for comparing buffer capacity and efficiency.</a> Cryobiology. 45 (1), 33-48 (2002).</li>
<li>Miyanishi, H., Inokuchi, M., Nobata, S., Kaneko, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Past+seawater+experience+enhances+seawater+adaptability+in+medaka%2C+Oryzias+latipes%5BTitle%5D)+AND+%22Zoological+Lett%22%5BJournal%5D)">Past seawater experience enhances seawater adaptability in medaka, Oryzias latipes.</a> Zoological Lett. 2, 12(2016).</li>
<li>Cass, A., Finkelstein, A., Krespi, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+ion+permeability+induced+in+thin+lipid+membranes+by+the+polyene+antibiotics+nystatin+and+amphotericin+B%5BTitle%5D)+AND+%22J+Gen+Physiol%22%5BJournal%5D)">The ion permeability induced in thin lipid membranes by the polyene antibiotics nystatin and amphotericin B.</a> J Gen Physiol. 56 (1), 100-124 (1970).</li>
<li>Holz, R., Finkelstein, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+water+and+nonelectrolyte+permeability+induced+in+thin+lipid+membranes+by+the+polyene+antibiotics+nystatin+and+amphotericin+B%5BTitle%5D)+AND+%22J+Gen+Physiol%22%5BJournal%5D)">The water and nonelectrolyte permeability induced in thin lipid membranes by the polyene antibiotics nystatin and amphotericin B.</a> J Gen Physiol. 56 (1), 125-145 (1970).</li>
</ol>

The authors have nothing to disclose

Electrophysiological recordings using the patch-clamp technique on brain-pituitary slices require careful optimization. Well-optimized protocols for conducting live-cell investigations specifically in teleosts are limited, with the majority of publications using protocols based on mammalian systems. In this regard, it is important to be aware of the fact that several physiological parameters like pH and osmolality are not only species dependent, but also very much dependent on whether the organism in question lives on la...

The pituitary is a key endocrine organ in vertebrates located below the hypothalamus and posterior to the optic chiasm. It produces and secretes six to eight hormones from the specific cell types. Pituitary hormones constitute an intermediate between the brain and peripheral organs and drive a wide range of essential physiological processes including growth, reproduction, and regulation of homeostasis. Similar to neurons, endocrine cells of the pituitary are electrically excitable with the ability to fire action potentials spontaneously 1. The role of these action potentials is cell dependent. In several cell types of the mammalian pituitary, action potentials can elevate the intracellular Ca2+ sufficiently for a sustained release of hormone 2. In addition, the pituitary receives both stimulatory and inhibitory information from the brain that affects the membrane potential of the cells 3,4,5,6. Typically, stimulatory input increases the excitability and often involves the release of Ca2+ from intracellular stores as well as increased firing frequency&#160;7. Understanding how the cell utilizes the ion channel composition and adapts to these input signals from the brain is key to understanding hormone synthesis and release.
The patch-clamp technique was developed in the late 1970s by Sakmann and Neher 8,9,10 and further improved by Hamill 11, and allows detailed investigations of electrophysiological properties of cells down to single ion channels. Moreover, the technique can be used for studying both current and voltage. Today, patch-clamping is the gold standard for measuring electrophysiological properties of the cell. Four major configurations of the tight seal patch-clamp technique have been developed 11; the cell-attached, the inside-out, the outside-out, and the whole-cell patch. The three first configurations are typically used for single ion channel investigations. For the fourth, following the cell-attached configuration, a hole in the cell membrane is made using sub-atmospheric pressure. This configuration also allows investigations of the ion channel composition of the whole cell 12. However, one limitation of this technique is that cytoplasmic molecules are diluted by the patch pipette solution 13 (Figure 1A), thus affecting the electrical and physiological responses of the studied cells. Indeed, some of those molecules may play important roles in the transduction of the signal or in the regulation of different ion channels. To avoid this, Lindau and Fernandez 14 developed a method where a pore-forming compound is added to the patch pipette. Following the cell-attached configuration, the compound will incorporate into the plasma membrane under the patch and slowly perforate the membrane creating electrical contact with the cytosol (Figure 1B). Several different antifungals such as nystatin 15 and amphotericin B 16, or surfactants such as the saponin beta-escin 17,18 can be used. These compounds create pores large enough to allow monovalent cation and Cl- diffusion between the cytosol and the patch pipette while preserving the cytosolic levels of macromolecules and larger ions like Ca2+ 15,16.
The challenge of using perforated patch is the potentially high series resistance. Series resistance (Rs) or access resistance is the combined resistance over the patch pipette relative to the ground. During patch-clamp recordings, the Rs will be in parallel with membrane resistance (Rm). Rm and Rs in parallel work as a voltage divider. With the high Rs, the voltage will fall over the Rs giving errors in the recordings. The error will become larger with larger currents recorded. In addition, the voltage divider is also frequency dependent creating a low-pass filter, thus affecting the temporal resolution. In effect, the perforated patch may not always allow recordings of large and fast currents like the voltage gated Na+ currents (for detailed readings see reference&#160;19). Also, Rs may vary during patch-clamp recordings, again leading to changes in the recorded current. Thus, false positives may occur in situations where Rs changes during drug application.
The electrophysiology on the sliced tissue was first introduced by the Andersen lab to study electrophysiological characteristics of the neurons in the brain 20. The technique paved the way for detailed investigations of single cells as well as cell-cell communications and cell circuits in a more intact environment. A similar technique for making pituitary slices was introduced in 1998 by Gu&#233;rineau et al.&#160;21. However, it was not before 2005, that brain-pituitary slice preparation was used successfully for patch-clamp studies in teleost 22. In this study, the authors also reported the use of perforated patch-clamp recordings. However, by far, most of the electrophysiological investigations of pituitary cells have been conducted in mammals, and only a handful of other vertebrates, including teleost fish 1,2,22,23. In teleosts, almost all studies were performed on primary dissociated cells 24,25,26,27,28,29,30.
In the present paper, we outline an optimized protocol for preparation of healthy brain-pituitary slices from the model fish medaka. The approach represents several advantages compared to primary dissociated cell cultures. First, the cells are recorded in a relatively preserved environment compared to dissociated cell culture conditions. Second, slice preparations allow us to study indirect pathways mediated by cell-cell communication 22, which is not possible in dissociated cell culture conditions. Furthermore, we demonstrate how to conduct electrophysiological recordings on the obtained tissue slices using the perforated whole-cell patch-clamp technique with amphotericin B as the pore-forming agent.
Medaka is a small freshwater fish native to Asia, primarily found in Japan. The physiology, embryology, and genetics of medaka have been extensively studied for over 100 years 31, and it is a commonly used research model in many laboratories. Of particular importance to this paper is the distinct morphological organization of the hypothalamus-pituitary complex in teleost fish: Whereas in mammals and birds the hypothalamic neurons release their neuro-hormones regulating pituitary endocrine cells into the portal system of the median eminence, there is a direct nervous projection of hypothalamic neurons onto the endocrine cells of the pituitary in teleost fish 32. Thus, carefully conducted brain-pituitary slicing is of particular importance in fish, allowing us to investigate electrophysiological characteristics of the pituitary cells in a well-preserved brain-pituitary network, and in particular how pituitary cells control their excitability and thereby Ca2+ homeostasis.

<table><tbody><tr><td>Vibratome</td><td>Leica</td><td>VT1000 S</td><td></td></tr><tr><td>Chirurgical glue</td><td>WPI</td><td>VETBOND</td><td>3M Vetbond Tissue Adhesive</td></tr><tr><td>Stainless steel blades</td><td>Campden Instruments</td><td>752-1-SS</td><td></td></tr><tr><td>metal molds</td><td>SAKURA</td><td>4122</td><td></td></tr><tr><td>steel harp</td><td>Warner instruments</td><td>64-1417</td><td></td></tr><tr><td>PBS</td><td>SIGMA</td><td>D8537</td><td></td></tr><tr><td>Ultrapure LMP agarose</td><td>invitrogen</td><td>166520-100</td><td></td></tr><tr><td>patch pipettes</td><td>Sutter Instrument</td><td>BF150-110-10HP</td><td>Borosilicate with filament O.D.:1.5 mm, I.D.:1.10 mm</td></tr><tr><td>Microscope Slicescope</td><td>Scientifica</td><td>pro6000</td><td></td></tr><tr><td>P-Clamp10</td><td>Molecular Devices</td><td>#1-2500-0180</td><td>sofware</td></tr><tr><td>Digitizer Digidata 1550A1</td><td>Molecular Devices</td><td>DD1550</td><td></td></tr><tr><td>Amplifier Multiclap 700B Headstage CV-7B</td><td>Molecular Devices</td><td>1-CV-7B</td><td></td></tr><tr><td>GnRH</td><td>Bachem</td><td>4108604</td><td>H-Glu-His-Trp-Ser-His-Gly-Leu-Ser-Pro-Gly-OH trifluoroacetate salt&amp;nbsp;</td></tr><tr><td>pipette puller</td><td>Sutter Instrument</td><td>P-1000</td><td></td></tr><tr><td>amphotericin B</td><td>SIGMA</td><td>A9528</td><td>pore-forming antibiotic</td></tr><tr><td>polyethylenimine&amp;nbsp;</td><td>SIGMA</td><td>P3143</td><td>50% PEI solution</td></tr><tr><td>microfiler syringe</td><td>WPI</td><td>MF28/g67-5</td><td></td></tr><tr><td>glass for the agar bridge</td><td>Sutter Instrument</td><td>BF200-116-15</td><td>Borosilicate with filament O.D.:2.0 mm, I.D.:1.16 mm Fire polished</td></tr><tr><td>Micro-Manager software</td><td></td><td></td><td>Open Source Microscopy Software</td></tr><tr><td>optiMOS sCMOS camera</td><td>Qimaging</td><td>&amp;nbsp;01-OPTIMOS-R-M-16-C</td><td></td></tr><tr><td>sonicator</td><td>Elma</td><td>D-7700 singen</td><td></td></tr><tr><td>NaCl</td><td>SiGMA</td><td>S3014</td><td></td></tr><tr><td>KCl</td><td>SiGMA</td><td>P9541</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>SiGMA</td><td>M8266</td><td></td></tr><tr><td>D-Glucose</td><td>SiGMA</td><td>G5400</td><td></td></tr><tr><td>Hepes</td><td>SiGMA</td><td>H4034</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>SiGMA</td><td>C8106</td><td></td></tr><tr><td>Sucrose</td><td>SiGMA</td><td>84097</td><td></td></tr><tr><td>D-mannitol</td><td>SiGMA</td><td>63565</td><td></td></tr><tr><td>MES-acid</td><td>SIGMA</td><td>M0895</td><td></td></tr><tr><td>BSA</td><td>SIGMA</td><td>A2153</td><td></td></tr></tbody></table>

healthy brain-pituitary slices for electrophysiological investigations of pituitary cells in teleost fish

Electrophysiological investigations of pituitary cells have been conducted in numerous vertebrate species, but very few in teleost fish. Among these, the clear majority have been performed on dissociated primary cells. To improve our understanding of how teleost pituitary cells, behave in a more biologically relevant environment, this protocol shows how to prepare viable brain-pituitary slices using the small freshwater fish medaka (Oryzias latipes). Making the brain-pituitary slices, pH and osmolality of all solutions were adjusted to values found in body fluids of freshwater fish living at 25 to 28 &#176;C. Following slice preparation, the protocol demonstrates how to conduct electrophysiological recordings using the perforated whole-cell patch-clamp technique. The patch-clamp technique is a powerful tool with unprecedented temporal resolution and sensitivity, allowing investigation of electrical properties from intact whole cells down to single ion channels. Perforated patch is unique in that it keeps the intracellular environment intact preventing regulatory elements in the cytosol from being diluted by the patch pipette electrode solution. In contrast, when performing traditional whole-cell recordings, it was observed that medaka pituitary cells quickly lose their ability to fire action potentials. Among the various perforation techniques available, this protocol demonstrates how to achieve perforation of the patched membrane using the fungicide Amphotericin B.

This protocol demonstrates a step by step protocol of how to achieve reliable electrophysiological recordings from pituitary (gonadotrope) cells, using a medaka transgenic line [Tg(lhb-hrGfpII)] where the target cells (Lh-producing gonadotropes) are labeled with green fluorescent protein (GFP).
Initially, the electrophysiological investigations were conducted using whole-cell configuration. However, spontaneous action p...

Watch this Scientific Journal Video about Healthy Brain-pituitary Slices for Electrophysiological Investigations of Pituitary Cells in Teleost Fish at JoVE.com

Healthy Brain-pituitary Slices for Electrophysiological Investigations of Pituitary Cells in Teleost Fish

The article describes an optimized protocol for making viable brain-pituitary tissue slices, using the teleost fish medaka (Oryzias latipes), followed by electrophysiological recordings of pituitary cells using the patch-clamp technique with the perforated patch configuration.

Electrophysiological investigations of pituitary cells have been conducted in numerous vertebrate species, but very few in teleost fish. Among these, the ...

healthy-brain-pituitary-slices-for-electrophysiological

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Neuroscience

7.7K Views.  Norwegian University of Life Sciences. The article describes an optimized protocol for making viable brain-pituitary tissue slices, using the teleost fish medaka (Oryzias latipes), followed by electrophysiological recordings of pituitary cells using the patch-clamp technique with the perforated patch configuration.

Video: Healthy Brain-pituitary Slices for Electrophysiological Investigations of Pituitary Cells in Teleost Fish

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate retina. Two classes of retinal neurons, photoreceptors and bipolar cells, accomplish this by using ribbon-type active zones, which enable sustained and high-throughput neurotransmitter release over long time periods. ON-type mixed bipolar cell (Mb) terminals in the goldfish retina, which depolarize to light stimuli and receive mixed rod and cone photoreceptor input, are suitable for the study of ribbon-type synapses both due to their large size (~10-12 &mu;m diameter) and to their numerous lateral and reciprocal synaptic connections with amacrine cell dendrites. Direct access to Mb bipolar cell terminals in goldfish retinal slices with the patch-clamp technique allows the measurement of presynaptic Ca2+ currents, membrane capacitance changes, and reciprocal synaptic feedback inhibition mediated by GABAA and GABAC receptors expressed on the terminals. Presynaptic membrane capacitance measurements of exocytosis allow one to study the short-term plasticity of excitatory neurotransmitter release 14,15. In addition, short-term and long-term plasticity of inhibitory neurotransmitter release from amacrine cells can also be investigated by recordings of reciprocal feedback inhibition arriving at the Mb terminal 21. Over short periods of time (e.g. ~10 s), GABAergic reciprocal feedback inhibition from amacrine cells undergoes paired-pulse depression via GABA vesicle pool depletion 11. The synaptic dynamics of retinal microcircuits in the inner plexiform layer of the retina can thus be directly studied.
The brain-slice technique was introduced more than 40 years ago but is still very useful for the investigation of the electrical properties of neurons, both at the single cell soma, single dendrite or axon, and microcircuit synaptic level 19. Tissues that are too small to be glued directly onto the slicing chamber are often first embedded in agar (or placed onto a filter paper) and then sliced 20, 23, 18, 9. In this video, we employ the pre-embedding agar technique using goldfish retina. Some of the giant bipolar cell terminals in our slices of goldfish retina are axotomized (axon-cut) during the slicing procedure. This allows us to isolate single presynaptic nerve terminal inputs, because recording from axotomized terminals excludes the signals from the soma-dendritic compartment. Alternatively, one can also record from intact Mb bipolar cells, by recording from terminals attached to axons that have not been cut during the slicing procedure. Overall, use of this experimental protocol will aid in studies of retinal synaptic physiology, microcircuit functional analysis, and synaptic transmission at ribbon synapses.

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Here we describe a protocol for the preparation of agar-embedded retinal slices that are suitable for electrophysiology and Ca2+ imaging. This method allows one to study ribbon-type synapses in retinal microcircuits using direct patch-clamp recordings of single presynaptic nerve terminals.

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be performed in an intact brain preparation where the neural circuitry of the CNS remains intact. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with green fluorescent protein for identification in the intact brain. Fish have multiple populations of GnRH neurons, and their functions are dependent on their location in the brain and the GnRH gene that they express1 . We have focused our demonstration on GnRH3 neurons located in the terminal nerves (TN) associated with the olfactory bulbs using the intact brain of transgenic medaka fish (Figure 1B and C). Studies suggest that medaka TN-GnRH3 neurons are neuromodulatory, acting as a transmitter of information from the external environment to the central nervous system; they do not play a direct role in regulating pituitary-gonadal functions, as do the well-known hypothalamic GnRH1 neurons2, 3 .The tonic pattern of spontaneous action potential firing of TN-GnRH3 neurons is an intrinsic property4-6, the frequency of which is modulated by visual cues from conspecifics2 and the neuropeptide kisspeptin 15. In this video, we use a stable line of transgenic medaka in which TN-GnRH3 neurons express a transgene containing the promoter region of Gnrh3 linked to enhanced green fluorescent protein7 to show you how to identify neurons and monitor their electrical activity in the whole brain preparation6.

In this video, we will demonstrate how to record electrical activity from identified single neurons in a whole brain preparation, which preserves complex neural circuits. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with a fluorescent protein for identification in the intact brain preparation.

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Labeling of Blood Vessels in the Teleost Brain and Pituitary Using Cardiac Perfusion with a DiI-fixative

Blood vessels innervate all tissues in vertebrates, enabling their survival by providing the necessary nutrients, oxygen, and hormonal signals. It is one of the first organs to start functioning during development. Mechanisms of blood vessel formation have become a subject of high scientific and clinical interest. In adults however, it is difficult to visualize the vasculature in most living animals due to their localization deep within other tissues. Nevertheless, visualization of blood vessels remains important for several studies such as endocrinology and neurobiology. While several transgenic lines have been developed in zebrafish, with blood vessels directly visualized through expression of fluorescent proteins, no such tools exist for other teleost species. Using medaka (Oryzias latipes) as a model, the current protocol presents a quick and direct technique to label blood vessels in brain and pituitary by perfusing through the heart with fixative containing DiI. This protocol allows improvement of our understanding on how brain and pituitary cells interact with blood vasculature in whole tissue or thick tissue slices.

The article describes a quick protocol for labeling blood vessels in a teleost fish by cardiac perfusion of DiI diluted in fixative, using medaka (Oryzias latipes) as a model and focusing on brain and pituitary tissue.

Blood vessels innervate all tissues in vertebrates, enabling their survival by providing the necessary nutrients, oxygen, and hormonal signals. It is one ...

Bioengineering

Preparation of a High-quality Primary Cell Culture from Fish Pituitaries

Primary cell culture is a powerful tool commonly used by scientists to study cellular properties and mechanisms of isolated cells in a controlled environment. Despite vast differences in the physiology between mammals and fish, primary cell culture protocols from fish are often based on mammalian culture conditions, often with only minor modifications. The environmental differences affect not only body temperature, but also blood serum parameters such as osmolality, pH, and pH buffer capacity. As cell culture media and similar working solutions are meant to mimic characteristics of the extracellular fluid and/or blood serum to which a cell is adapted, it is crucial that these parameters are adjusted specifically to the animal in question.
The current protocol describes optimized primary culture conditions for medaka (Oryzias latipes). The protocol provides detailed steps on how to isolate and maintain healthy dissociated pituitary cells for more than one week and includes the following steps: 1. the adjustment of the osmolality to the values found in medaka blood plasma, 2. the adjustment of the incubation temperature to normal medaka temperature (here in the aquarium facility), and 3. the adjustment of the pH and bicarbonate buffer to values comparable to other fish species living at similar temperatures. The results presented using the described protocol promote physiologically meaningful results for medaka and can be used as a reference guide by scientists making primary cell cultures from other non-mammalian species.

Here we describe a protocol to prepare and maintain primary pituitary cell cultures from medaka (Oryzias latipes). The optimized conditions in this protocol take important parameters such as temperature, osmolality, and pH into consideration by mimicking the physiological conditions of the fish, thereby enabling physiologically more meaningful results.

Primary cell culture is a powerful tool commonly used by scientists to study cellular properties and mechanisms of isolated cells in a controlled ...

Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ganglion cells and project it to the visual cortex. In addition, relay neurons receive top-down excitation from the cortex. The two main excitatory inputs to the relay neurons differ in several aspects. Each relay neuron receives input from only a few retinogeniculate synapses, which are large terminals with many release sites. This is reflected by the comparably strong excitation, the relay neurons receive, from retinal ganglion cells. Corticogeniculate synapses, in contrast, are simpler with few release sites and weaker synaptic strength. The two synapses also differ in their synaptic short-term plasticity. Retinogeniculate synapses have a high release probability and consequently display a short-term depression. In contrast, corticogeniculate synapses have a low release probability. Corticogeniculate fibers traverse the reticular thalamic nuclei before entering the lateral geniculate nucleus. The different locations of the reticular thalamic nucleus (rostrally from the lateral geniculate nucleus) and optic tract (ventro-laterally from the lateral geniculate nucleus) allow stimulating corticogeniculate or retinogeniculate synapses separately with extracellular stimulation electrodes. This makes the lateral geniculate nucleus an ideal brain area where two excitatory synapses with very different properties impinging onto the same cell type, can be studied simultaneously. Here, we describe a method to investigate the recording from relay neurons and to perform detailed analysis of the retinogeniculate and corticogeniculate synapse function in acute brain slices. The article contains a step-by-step protocol for the generation of acute brain slices of the lateral geniculate nucleus and steps for recording activity from relay neurons by stimulating the optic tract and corticogeniculate fibers separately.

Here, we present protocols for the preparation of acute brain slices containing the lateral geniculate nucleus and the electrophysiological investigation of retinogeniculate and corticogeniculate synapses function. This protocol provides an efficient way to study synapses with the high-release and low-release probability in the same acute brain slices.

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ...

Preparing Acute Brain Slices from the Dorsal Pole of the Hippocampus from Adult Rodents

Whole-cell patch-clamp recordings from acute rodent brain slices are a mainstay of modern neurophysiological research, allowing precise measurement of cellular and synaptic properties. Nevertheless, there is an ever increasing need to perform correlated analyses between different experimental modes in addition to slice electrophysiology, for example: immunohistochemistry, molecular biology, in vivo imaging or electrophysiological recording; to answer evermore complex questions of brain function. However, making meaningful conclusions from these various experimental approaches is not straightforward, as even within relatively well described brain structures, a high degree of sub-regional variation of cellular function exists. Nowhere is this better exemplified than in the CA1 of the hippocampus, which has well-defined dorso-ventral properties, based on cellular and molecular properties. Nevertheless, many published studies examine protein expression patterns or behaviorally correlated in vivo activity in the dorsal extent of the hippocampus; and explain findings mechanistically with cellular electrophysiology from the ventro-medial region. This is further confounded by the fact that many acute slice electrophysiological experiments are performed in juvenile animals, when other experimental modes are performed in more mature animals. To address these issues, this method incorporates transcardial perfusion of mature (&#62;60 day old rodents) with artificial cerebrospinal fluid followed by preparation of modified coronal slices including the septal pole of the dorsal hippocampus to record from CA1 pyramidal cells. This process leads to the generation of healthy acute slices of dorsal hippocampus allowing for slice-based cellular electrophysiological interrogation matched to other measures.

The purpose of this protocol is to describe a method to produce slices of the dorsal hippocampus for electrophysiological examination. This procedure employs perfusion with chilled ACSF prior to slice preparation with a near-coronal slicing angle which allows for preservation of healthy principal neurons.

Whole-cell patch-clamp recordings from acute rodent brain slices are a mainstay of modern neurophysiological research, allowing precise measurement of ...

Adult Zebrafish Electrophysiology: A Method to Measure the Extracellular Electrical Activity of Neurons In Vivo

Source: Johnston, L., et al. <a href="https://www.jove.com/v/51065/" target="_blank">Electrophysiological Recording in the Brain of Intact Adult Zebrafish</a>. &#160;J. Vis. Exp. (2013)
The protocol describes the procedure to perform electrophysiology in adult zebrafish. This method can be used for measuring neural activity in vivo.

Source: Johnston, L., et al. Electrophysiological Recording in the Brain of Intact Adult Zebrafish. &#160;J. Vis. Exp. (2013)
The protocol describes the ...