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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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Signal+transduction+involved+in+GnRH2-stimulation+of+identified+LH-producing+gonadotropes+from+lhb-GFP+transgenic+medaka+(Oryzias+latipes).">Signal transduction involved in GnRH2-stimulation of identified LH-producing gonadotropes from lhb-GFP transgenic medaka (Oryzias latipes).</a> Mol Cell Endocrinol. 372 (1-2), 128-139 (2013).</li><li>Hodne, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+differences+between+fshb-+and+lhb-expressing+gonadotropes+in+primary+culture.">Electrophysiological differences between fshb- and lhb-expressing gonadotropes in primary culture.</a> Endocrinology. 154 (9), 3319-3330 (2013).</li><li>Wittbrodt, J., Shima, A., Schartl, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Medaka--a+model+organism+from+the+far+East.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hypothalamic+control+of+the+pars+distalis+in+fishes,+amphibians,+and+reptiles.">Hypothalamic control of the pars distalis in fishes, amphibians, and reptiles.</a> Gen Comp Endocrinol. 44 (2), 135-170 (1981).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=pCLAMP+10+Data+Acquisition+and+Analysis+For+Comprehensive+Electrophysiology+User+Guide.">pCLAMP 10 Data Acquisition and Analysis For Comprehensive Electrophysiology User Guide.</a> Molecular Devices Corporation. , (2006).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MultiClamp+700B+COMPUTER-CONTROLLED+MICROELECTRODE+AMPLIFIER+Theory+and+Operation.">MultiClamp 700B COMPUTER-CONTROLLED MICROELECTRODE AMPLIFIER Theory and Operation.</a> Axon Instruments / Molecular Devices Corp. , (2005).</li><li>Dominguez-Mancera, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Leptin+regulation+of+inward+membrane+currents,+electrical+activity+and+LH+release+in+isolated+bovine+gonadotropes.">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="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Animal Physiology : adptation and environment fifth edition. , 613 (1997).</li><li>Burton, R. 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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.

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			<td>752-1-SS</td>
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			<td height="21" style="height:21px;width:309px;">metal molds</td>
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			<td height="21" style="height:21px;width:309px;">amphotericin B</td>
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			<td style="width:157px;">A9528</td>
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			<td height="21" style="height:21px;">polyethylenimine&#160;</td>
			<td style="width:171px;">SIGMA</td>
			<td style="width:157px;">P3143</td>
			<td style="width:428px;">50% PEI solution</td>
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			<td height="21" style="height:21px;width:309px;">microfiler syringe</td>
			<td style="width:171px;">WPI</td>
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		<tr height="42">
			<td height="42" style="height:42px;width:309px;">glass for the agar bridge</td>
			<td style="width:171px;">Sutter Instrument</td>
			<td style="width:157px;">BF200-116-15</td>
			<td style="width:428px;">Borosilicate with filament O.D.:2.0mm, I.D.:1.16mm Fire polished</td>
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			<td height="21" style="height:21px;">Micro-Manager software</td>
			<td style="width:171px;"></td>
			<td style="width:157px;"></td>
			<td style="width:428px;">Open Source Microscopy Software</td>
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			<td height="42" style="height:42px;width:309px;">optiMOS sCMOS camera</td>
			<td style="width:171px;">Qimaging</td>
			<td style="width:157px;">&#160;01-OPTIMOS-R-M-16-C</td>
			<td style="width:428px;"></td>
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			<td height="21" style="height:21px;width:309px;">sonicator</td>
			<td style="width:171px;">Elma</td>
			<td style="width:157px;">D-7700 singen</td>
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			<td style="width:157px;">S3014</td>
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			<td height="21" style="height:21px;width:309px;">KCl</td>
			<td style="width:171px;">SiGMA</td>
			<td style="width:157px;">P9541</td>
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			<td height="21" style="height:21px;width:309px;">MgCl2</td>
			<td style="width:171px;">SiGMA</td>
			<td style="width:157px;">M8266</td>
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			<td height="21" style="height:21px;width:309px;">D-Glucose</td>
			<td style="width:171px;">SiGMA</td>
			<td style="width:157px;">G5400</td>
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			<td height="21" style="height:21px;width:309px;">Hepes</td>
			<td style="width:171px;">SiGMA</td>
			<td style="width:157px;">H4034</td>
			<td></td>
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			<td height="21" style="height:21px;width:309px;">CaCl2</td>
			<td style="width:171px;">SiGMA</td>
			<td style="width:157px;">C8106</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:309px;">Sucrose</td>
			<td style="width:171px;">SiGMA</td>
			<td align="right" style="width:157px;">84097</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:309px;">D-mannitol</td>
			<td style="width:171px;">SiGMA</td>
			<td align="right" style="width:157px;">63565</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:309px;">MES-acid</td>
			<td style="width:171px;">SIGMA</td>
			<td style="width:157px;">M0895</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:309px;">BSA</td>
			<td style="width:171px;">SIGMA</td>
			<td style="width:157px;">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 ...

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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

Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

When startled adult D. melanogaster react by jumping into the air and flying away. In many invertebrate species, including D. melanogaster, the "escape" (or "startle") response during the adult stage is mediated by the multi-component neuronal circuit called the Giant Fiber System (GFS). The comparative large size of the neurons, their distinctive morphology and simple connectivity make the GFS an attractive model system for studying neuronal circuitry. The GFS pathway is composed of two bilaterally symmetrical Giant Fiber (GF) interneurons whose axons descend from the brain along the midline into the thoracic ganglion via the cervical connective. In the mesothoracic neuromere (T2) of the ventral ganglia the GFs form electro-chemical synapses with 1) the large medial dendrite of the ipsilateral motorneuron (TTMn) which drives the tergotrochanteral muscle (TTM), the main extensor for the mesothoracic femur/leg, and 2) the contralateral peripherally synapsing interneuron (PSI) which in turn forms chemical (cholinergic) synapses with the motorneurons (DLMns) of the dorsal longitudinal muscles (DLMs), the wing depressors. The neuronal pathway(s) to the dorsovental muscles (DVMs), the wing elevators, has not yet been worked out (the DLMs and DVMs are known jointly as indirect flight muscles - they are not attached directly to the wings, but rather move the wings indirectly by distorting the nearby thoracic cuticle) (King and Wyman, 1980; Allen et al., 2006). The di-synaptic activation of the DLMs (via PSI) causes a small but important delay in the timing of the contraction of these muscles relative to the monosynaptic activation of TTM (~0.5 ms) allowing the TTMs to first extend the femur and propel the fly off the ground. The TTMs simultaneously stretch-activate the DLMs which in turn mutually stretch-activate the DVMs for the duration of the flight. The GF pathway can be activated either indirectly by applying a sensory (e.g."air-puff" or "lights-off") stimulus, or directly by a supra-threshold electrical stimulus to the brain (described here). In both cases, an action potential reaches the TTMs and DLMs solely via the GFs, PSIs, and TTM/DLM motoneurons, although the TTMns and DLMns do have other, as yet unidentified, sensory inputs. Measuring "latency response" (the time between the stimulation and muscle depolarization) and the "following to high frequency stimulation" (the number of successful responses to a certain number of high frequency stimuli) provides a way to reproducibly and quantitatively assess the functional status of the GFS components, including both central synapses (GF-TTMn, GF-PSI, PSI-DLMn) and the chemical (glutamatergic) neuromuscular junctions (TTMn-TTM and DLMn-DLM). It has been used to identify genes involved in central synapse formation and to assess CNS function.

Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

The Giant Fiber System is a simple neuronal circuit of adult Drosophila melanogaster containing the largest neurons in the fly. We describe the protocol for monitoring synaptic transmission through this pathway by recording post synaptic potentials in dorsal longitudinal (DLM) and tergotrochanteral (TTM) muscles following direct stimulation of the Giant Fiber interneurons.

When startled adult D. melanogaster react by jumping into the air and flying away. In many invertebrate species, including D. melanogaster, the "escape" ...

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Electrophysiological and Morphological Characterization of Neuronal Microcircuits in Acute Brain Slices Using Paired Patch-Clamp Recordings

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with simultaneous intracellular biocytin filling allows a correlated analysis of their structural and functional properties. With this method it is possible to identify and characterize both pre- and postsynaptic neurons by their morphology and electrophysiological response pattern. Paired recordings allow studying the connectivity patterns between these neurons as well as the properties of both chemical and electrical synaptic transmission. Here, we give a step-by-step description of the procedures required to obtain reliable paired recordings together with an optimal recovery of the neuron morphology. We will describe how pairs of neurons connected via chemical synapses or gap junctions are identified in brain slice preparations. We will outline how neurons are reconstructed to obtain their 3D morphology of the dendritic and axonal domain and how synaptic contacts are identified and localized. We will also discuss the caveats and limitations of the paired recording technique, in particular those associated with dendritic and axonal truncations during the preparation of brain slices because these strongly affect connectivity estimates. However, because of the versatility of the paired recording approach it will remain a valuable tool in characterizing different aspects of synaptic transmission at identified neuronal microcircuits in the brain.

Patch-clamp recordings and simultaneous intracellular biocytin filling of synaptically coupled neurons in acute brain slices allow a correlated analysis of their structural and functional properties. The aim of this protocol is to describe the essential technical steps of electrophysiological recording from neuronal microcircuits and their subsequent morphological analysis.

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

Preparation of Horizontal Slices of Adult Mouse Retina for Electrophysiological Studies

Vertical slice preparations are well established to study circuitry and signal transmission in the adult mammalian retina. The plane of sectioning in these preparations is perpendicular to the retinal surface, making it ideal for the study of radially oriented neurons like photoreceptors and bipolar cells. However, the large dendritic arbors of horizontal cells, wide-field amacrine cells, and ganglion cells are mostly truncated, leaving markedly reduced synaptic activity in these cells. Whereas ganglion cells and displaced amacrine cells can be studied in a whole-mounted preparation of the retina, horizontal cells and amacrine cells located in the inner nuclear layer are only poorly accessible for electrodes in whole retina tissue.
To achieve maximum accessibility and synaptic integrity, we developed a horizontal slice preparation of the mouse retina, and studied signal transmission at the synapse between photoreceptors and horizontal cells. Horizontal sectioning allows&#160;(1) easy and unambiguous visual identification of horizontal cell bodies for electrode targeting, and (2) preservation of the extended horizontal cell dendritic fields, as a prerequisite for intact and functional cone synaptic input to horizontal cell dendrites.
Horizontal cells from horizontal slices exhibited tonic synaptic activity in the dark, and they responded to brief flashes of light with a reduction of inward current and diminished synaptic activity. Immunocytochemical evidence indicates that almost all cones within the dendritic field of a horizontal cell establish synapses with its peripheral dendrites. The horizontal slice preparation is therefore well suited to study the physiological properties of horizontally extended retinal neurons as well as sensory signal transmission and integration across selected synapses.

We developed a horizontal slice preparation of the adult mammalian retina with the plane of sectioning parallel to the retinal surface. Dendritic fields of nonradially oriented retinal neurons were not truncated, allowing the study of signal processing with patch-clamp and imaging techniques.

Vertical slice preparations are well established to study circuitry and signal transmission in the adult mammalian retina. The plane of sectioning in ...

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 ...

Assessment of Long-term Depression Induction in Adult Cerebellar Slices

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from parallel fibers (PF) to Purkinje cells (PC) is considered the basis for motor learning, and deficiencies of both LTD and motor learning are observed in various gene-manipulated animals. Common motor learning sets, such as adaptation of the optokinetic reflex (OKR), the vestibular-ocular reflex (VOR), and rotarod test were used for evaluation of motor learning ability. However, results obtained from the GluA2-carboxy terminus modified knock-in mice demonstrated normal adaptation of the VOR and the OKR, despite lacking PF-LTD. In that report, induction of LTD was only attempted using one type of stimulation protocol at room temperature. Thus, conditions to induce cerebellar LTD were explored in the same knock-in mutants using various protocols at near physiological temperature. Finally, we found stimulation protocols, by which LTD could be induced in these gene-manipulated mice. In this study, a set of protocols are proposed to evaluate LTD-induction, which will more accurately allow examination of the causal relationship between LTD and motor learning. In conclusion, experimental conditions are crucial when evaluating LTD in gene-manipulated mice.

In some gene-manipulated animals, using a single protocol may fail to induce LTD in cerebellar Purkinje cells, and there may be a discrepancy between LTD and motor learning. Multiple protocols are necessary to assess LTD-induction in gene-manipulated animals. Standard protocols are shown.

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from ...

Wireless Electrophysiological Recording of Neurons by Movable Tetrodes in Freely Swimming Fish

The neural mechanisms governing fish behavior remain mostly unknown, although fish constitute the majority of all vertebrates. The ability to record brain activity from freely moving fish would advance research on the neural basis of fish behavior considerably. Moreover, precise control of the recording location in the brain is critical to studying coordinated neural activity across regions in fish brain. Here, we present a technique that records wirelessly from the brain of freely swimming fish while controlling for the depth of the recording location. The system is based on a neural logger associated with a novel water-compatible implant that can adjust the recording location by microdrive-controlled tetrodes. The capabilities of the system are illustrated through recordings from the telencephalon of goldfish.

A novel wireless technique for recording extracellular neural signals from the brain of freely swimming goldfish is presented. The recording device is composed of two tetrodes, a microdrive, a neural data logger, and a waterproof case. All parts are custom-made except for the data logger and its connector.

The neural mechanisms governing fish behavior remain mostly unknown, although fish constitute the majority of all vertebrates. The ability to record brain ...