Authors would like to thank Peter Mombaerts for the generous gift of OR-GFP mice; Anne Lefranc and the CSGA animal facility for excellent animal care. Funding was provided by CNRS through an ATIP and ATIP Plus grants, by Conseil R&#233;gional de Bourgogne (FABER and PARI grants), by Universit&#233; de Bourgogne (BQR program).

This protocol follows the animal care guidelines of the Universit&#233; de Bourgogne and was approved by the Universit&#233; de Bourgogne ethics committee.
1. Animals
<ol>
	<li>Use genetically engineered OR-IRES-tauGFP mice available at the Jackson Laboratory. These mice were developed in Dr. Peter Mombaerts&#8217; laboratory in order to analyze axon targeting and development of the olfactory system19. For example, the MOR23-IRES-tauGFP line, stock number 006643, bears the official strain name B6;129P2-Olfr16tm2Mom/MomJ; similarly, the SR1-IRES-tauGFP line, stock number 6717 bears the official name B6;129P2-Olfr124tm1Mom/MomJ.</li>
	<li>Regarding the age of the animals: for a better outcome of the protocol, use animals between 2 and 4 weeks of age. In this age group, the dissection is easier (softer bones, firmer olfactory epithelium) and the dendritic knobs are bigger compare to older animals.</li>
</ol>
2. Preparation of Electrodes and Solutions
<ol>
	<li>For the stimulating pipettes: purchase prepulled stimulating pipettes. Otherwise, manually prepare them.
	<ol>
		<li>Using a flame, bend six 1 mm glass pipettes at about 1 cm from the tip. Insert these six pipettes plus a straight 7th in 1.5 cm heat-shrink tubing strengthen by an eyelet.</li>
		<li>Heat shrink the tubing to maintain the barrels attached together. Attach an additional heat-shrink tubing to the other extremity of the barrels. Pull this stimulating pipette with a multi-barrel puller.</li>
		<li>Add some white liquid glue around the eyelet to strengthen the bended tips. Let dry O/N.</li>
	</ol>
	</li>
	<li>Prepare 1 or 2 L of normal Ringer&#8217;s extracellular solution (in mM: NaCl 124, KCl 3, MgSO4 1.3, CaCl2 2, NaHCO3 26, NaH2PO4 1.25, glucose 15; pH 7.6 and 305 mOsm). Keep at 4 &#176;C until use.</li>
	<li>Prepare intracellular stock solution (in mM): KCl 70, KOH 53, methanesulfonic acid 30, EGTA 5, HEPES 10, sucrose 70; pH 7.2 (KOH) and 310 mOsm. Keep at 4 &#176;C until use. Good for several weeks.</li>
	<li>Prepare intracellular recording solution with nystatin extemporaneously (at the last minute) before experiment.
	<ol>
		<li>Weigh 3 mg of nystatin, add 50 &#181;l&#160;of DMSO; vortex 20 sec then sonicate 2-3 min until entirely diluted.</li>
		<li>Add 20 &#181;l of DMSO-nystatin solution in 5 ml&#160;of intracellular stock solution. Vortex 20 sec, then sonicate 3 min. Keep this solution at 4 &#176;C and protect from direct light, nystatin is light-sensitive. This solution can be used for a few hours. Replace every day.</li>
		<li>Once the olfactory epithelium preparation is under the microscope, take some of this solution in a 1 ml&#160;syringe with a flame-elongated yellow tip to fill the electrodes; protect from direct light. Once at RT, the nystatin solution is not stable; replace the solution in the 1 ml syringe every hour or keep it in ice.</li>
	</ol>
	</li>
	<li>Pull recording electrodes with a puller to obtain long neck and small tip (~2 &#181;m) with a resistance of 15-20 M&#8486; with the internal nystatin solution.</li>
	<li>Prepare odorant solution at 0.5 M in DMSO under fume hood; aliquot and keep at -20 &#176;C until use. Dilute odorant in Ringer&#8217;s solution until the desired concentration. Fill the stimulating pipette.</li>
</ol>
3. Preparation of Olfactory Epithelium
Note: OR-IRES-tauGFP mice express the tauGFP under the control of the OR promoter. In these mice, all neurons expressing the OR of interest are labeled with GFP. This protocol is adapted for ORs expressed in all zones. However, dissections and recordings are easier for ORs expressed in the dorsal zone.
<ol>
	<li>Anesthetize the animal by injecting a mix of ketamine and xylazine (150 mg/kg and 10 mg/kg body weight, respectively).&#160;Decapitation can be performed with sharp scissors for young mice or with a properly maintained rodent guillotine for older mice.
	<ol>
		<li>Using ring dissecting scissors make a longitudinal medial incision through the dorsal skin. Remove the skin by pulling it apart. Using the scissors, cut the lower jaws at the jaw joint. Remove the upper front teeth by a coronal cut parallel to the teeth.</li>
		<li>Make a coronal cut of the head behind the eyes and keep only the anterior part of the head. Dip it in ice-cold Ringer solution for the dissection under the scope.</li>
	</ol>
	</li>
	<li>Dissect under the scope: in the ventral side, make a longitudinal cut along the upper jaw/the teeth. Cut the dorsal bones longitudinally, following the dorso-lateral side of the nasal cavity. Remove most bones and palate; transfer the septum and the epithelium attached to it in oxygenated Ringer at RT.</li>
	<li>For the final dissection: Right before starting the recording session, peel away the epithelium from the underlying septum with forceps. Detach the epithelium with forceps and with two 4-5 mm scissor cuts (use microvannas scissors) at the anterior part of the septum where the adhesion is stronger.
	<ol>
		<li>Carefully remove the vomeronasal organ by cutting it out along its dorsal connection to the septal epithelium. Transfer the epithelium to a recording chamber with the mucus layer facing up; keep it flat in the chamber with a harp.</li>
	</ol>
	</li>
	<li>Install chamber under an upright microscope equipped with fluorescence optics and a sensitive camera. Visualize the preparation on the computer screen at high magnification through a 40X water-immersion objective (numerical aperture 0.8) and an extra 2X or 4X magnification achieved by a magnifying lens. Perfuse continuously with oxygenated Ringer at RT (1-2 ml/min).</li>
</ol>
4. Recording Session
<ol>
	<li>Search for fluorescent cell: excite the preparation at 480 nm for EGFP, which emits light in the 530-550 nm range; target one dendritic knob which is reliably visible in fluorescence and under bright field.</li>
	<li>Fill electrode with intracellular solution with nystatin; remove bubbles by gently tapping on the electrode.</li>
	<li>Insert electrode on electrode holder, apply positive pressure in the pipette; Resistance should be 15-20 M&#8486;.</li>
	<li>Bring electrode close to the cell; once resistance reaches ~40 M&#8486;, release positive pressure and apply slight negative pressure to reach a gigaseal.</li>
	<li>Once seal is reached, set the membrane potential at about -75 mV;</li>
	<li>Once the cell is opened, proceed with stimulation protocols, pharmacological treatments. To measure the response to a single odorant stimulation, record 200-500 msec of spontaneous activity, stimulate for 500 msec and measure the response of the cell for up to 10 sec&#160;13. For pharmacological treatments, perfuse the pharmacological agents at the desired concentration20.</li>
</ol>
5. Data Analysis
<ol>
	<li>Analyze currents elicited by odorant stimulation as followed: the maximum amplitude, the rise-time (time necessary to reach 90% of the maximum amplitude, in msec), the total current (area under the curve in pAs), the time at 50% (time between the onset and the offset of the response at 50% of the maximum amplitude, in msec).
	<ol>
		<li>Using the peak transduction currents (maximum amplitude reached) at different concentrations, draw and fit a dose response curve using the Hill equation: I &#160;= Imax/(1 + (K1/2/C)n), where I represents the peak current, Imax the maximum response at saturating concentrations, K1/2 the concentration at which half of the maximum response was reached, C the concentration of odorant and n the Hill coefficient.</li>
	</ol>
	</li>
	<li>Analyze recordings of membrane potential in current clamp for the maximum depolarization, the total potential elicited (area under the curve in mVsec); record spontaneous spiking activity over 30 sec to 1 min recordings; record excitability through spikes elicited by injection of currents (5-10 pA).</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Odorant+responsiveness+of+embryonic+mouse+olfactory+sensory+neurons+expressing+the+odorant+receptors+S1+or+MOR23.">Odorant responsiveness of embryonic mouse olfactory sensory neurons expressing the odorant receptors S1 or MOR23.</a> Eur J Neurosci. 38 (2), 2210-2217 (2013).</li><li>Zhang, J., Huang, G., Dewan, A., Feinstein, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Uncoupling+stimulus+specificity+and+glomerular+position+in+the+mouse+olfactory+system.">Uncoupling stimulus specificity and glomerular position in the mouse olfactory system.</a> Mol Cell Neurosci. 51 (3-4), 79-88 (2012).</li><li>Zhang, J., Pacifico, R., Cawley, D., Feinstein, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrasensitive+detection+of+amines+by+a+trace+amine-associated+receptor.">Ultrasensitive detection of amines by a trace amine-associated receptor.</a> J Neurosci. 33 (7), 3228-3239 (2013).</li><li>Lee, A. 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C., Tan, J., Luo, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dual+functions+of+mammalian+olfactory+sensory+neurons+as+odor+detectors+and+mechanical+sensors.">Dual functions of mammalian olfactory sensory neurons as odor detectors and mechanical sensors.</a> Nat Neurosci. 10 (3), 348-354 (2007).</li><li>Duchamp-Viret, P., Chaput, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Odor+response+properties+of+rat+olfactory+receptor+neurons.">Odor response properties of rat olfactory receptor neurons.</a> Science. 284 (5423), 2171-2174 (1999).</li><li>Heydel, J. 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The authors declare that they have no competing financial interest.

The ability of this protocol to correctly monitor the properties of healthy OSNs depends heavily on the quality of the preparation. Therefore, the dissection steps are critical. First it is critical to pay attention to the quality (pH, osmolarity), oxygenation and temperature (ice-cold but not frozen) of the dissection medium. Second, the manipulation of the epithelium with dissecting tools must be as limited as possible to avoid damages. Finally, it is critical to obtain a preparation as flat as possible in order to access the largest OSN population as possible.
The dissection quality is fundamental to obtain a healthy preparation. However, different approaches can be used for the dissection: here we present a ventral to dorsal dissection but some users can prefer to start the dissection dorsally to open the nasal cavity rapidly and have a rapid access to the septal recess of the olfactory epithelium. Each user can hence adapt the protocol for the most efficient strategy to reach a healthy preparation of the area of interest.
Once the preparation is under the microscope, permanent monitoring of the preparation&#8217;s health is required. Indeed, an unhealthy preparation leads to a low probability to reach a gigaseal. A low efficiency in obtaining gigaseals can be improved by changing the area of interest on the preparation in order to find healthier OSNs. A more drastic solution is to replace the preparation entirely by a new one. Once the seal is reached, a low probability to reach the &#8220;opened&#8221; state can take place. This may be due to the internal solution&#8217;s quality. The internal solution must be prepared shortly before the experiment. The use of vortex and sonication are critical since the solubility of nystatin is rather low. To improve the probability of opening, the internal solution should be again vortexed for 10 sec and then sonicated for 30 to 60 sec. This usually improves the efficiency of the cell opening.
To stimulate the recorded OSN&#160;with odorants, the protocol presented uses a 7 barrel stimulating pipette. This type of pipette implies a limitation in the number of odorants and/or concentration of odorants tested on each recorded cells to seven. This limitation is relatively not significant in the case of a dose response curve since it can cover up to 7 log units of concentration. However, in the case of screening experiments, in which different odorants are tested, a seven barrel pipette may show limitations. In screening experiments, the goal is to find a ligand for labeled OSNs expressing an orphan OR. Using this protocol will require the use of odorant mixtures. Each mixture eliciting a response will be then divided into individual odorants. This was done to screen amine odorants on TAAR expressing OSNs16. In these experiments, the fluorescent neurons expressed TAARs and the ligand was unknown. Using the protocol, authors could screen several dozens of amine odorants in mixtures. The mixtures eliciting a response were then broken down to single odorants in order to find the most efficient ligand.
The distance between the recording site and the stimulating pipette should be chosen wisely in order to minimize the mechanical stimulation of the cell20. If the pressure is too high (above 30 psi in our recording conditions) and/or the distance too small (below 20 &#181;m), the mechanical response in some OSNs can even mask the odorant response. Pressure and distance between the puffing pipette and the recording site are also critical to control solution exchange during stimulation. The concentration reaching the neuron was previously evaluated to be as close as possible to the concentration present in the pipette through a series of tests13. These tests used a solution colored with blue dye to measure the onset and offset of the stimulation. Using 20 psi pressure and 30 &#181;m distance, the maximum intensity of the stimulus was reached within 300 msec. The offset of the stimulus was also carefully evaluated: since the preparation is under continuous flow of perfused solution, the stimulus was washed out within 0.8 to 0.9 sec13.
The odor coding properties of OSNs were historically measured either through extracellular recordings in vivo or through experiments on isolated cells. In mammals, extracellular recordings were performed at first in vivo in rats21. In extracellular recordings, the OR expressed in the recorded cell is unknown thus limiting the success ratio to find a cell responding to the odorant tested. The protocol allows the recording and characterization of defined receptors in neurons in the cellular environment of the epithelium. The receptors&#8217; properties can therefore be analyzed at different levels: the specific ligand/OR interactions during the first step of the transduction pathway; consequently, the agonist/antagonist interactions of diverse ligands tested; the properties of mixtures integration of defined ORs and the role of the transduction pathway for OSNs&#39; specificity; the influence on olfactory coding of cell-to-cell interactions (such as GAP junction) within the epithelium; and finally the modulation of the transduction pathway using pharmacological agents.
In the dissociated cell recordings, as mentioned earlier, the dissociation process removes the mucus and the cell-to-cell interactions within the olfactory epithelium. This may induce changes in the coding properties of the OSNs. Calcium imaging experiments on isolated cells were frequently reported. For example, dose-response curves of M71 expressing OSNs in response to acetophenone were reported7. These dose-response curves are steeper than the dose-responses curves obtained using the protocol presented here4. These discrepancies may be due to the cell-to-cell interactions in the intact preparation and also to the removal of mucus in isolated cells. Mucus is known to contain many protein and enzymes involved in perireceptor events fundamental for the olfactory transduction22,23. In the protocol reported here, the preservation of the structure of the olfactory epithelium and the putative presence of the mucus contribute to preserve the native features of odorant coding in OSNs. The concentrations used in the protocol are indicated in mol/L and usually are in the 10-7-10-3 range. These concentrations are higher than concentrations used in air-phased recordings (either in local field potential or extracellular recordings). These discrepancies might be due to i) the liquid phase of the recordings (and therefore of the stimulation) and ii) the slow washout of the mucus by the perfusion during recordings. In fact, the washout of the mucus and the section of the OSNs&#39; axons during dissection may support that the preparation presented be potentially qualified as a &#34;pseudo-intact&#34;&#160;rather than a &#34;fully-intact&#34; preparation. This preparation represents, however, recording conditions as close to in vivo as possible and yet eliciting patch-clamp recordings. As any in vitro preparation from a mammalian organism, the survival remains time-limited. This limit might be due to the washout of the mucus, the section of OSNs&#39; &#160;axons as well as the absence of blood circulation within the preparation.
The protocol presented here allows one to record membrane properties of olfactory sensory neurons. This technique can have multiple applications such as analysis of membrane properties of OSNs, pharmacological investigations of OSNs, modulation of odorant coding properties of OSNs, and deorphanization of ORs. The events recorded during calcium imaging are slow and long lasting. Patch-clamp recordings in the present protocol lead to data about rapid events in olfactory transduction pathway at the membrane level.
The recordings presented here are performed in the patch-clamp configuration. In this configuration, the membrane properties of the recorded cell can be analyzed, whether it is the olfactory transduction pathway machinery or the voltage gated currents involved. Experiments using this protocol can provide data about the modulation of the odorant coding properties of OSNs. Pharmacological agents can be used to investigate the olfactory transduction in different conditions: for example, MDL12330A, a blocker of adenylate cyclase III (ACIII)24,25 can be applied in the perfusion solution to investigate the role of ACIII in an odorant response. Additionally, this protocol can be used to investigate how olfactory coding properties are modulated in different conditions such as the olfactory environment18 or under the influence of hormones involved in feeding behaviors26. Finally, this protocol can also lead to data about OR/ligand interaction and deciphering agonist/antagonist activities on defined ORs or even in randomly chosen receptors27.
Finally, with the limitations due to the 7 barrel stimulating pipette, this protocol can be also be used for deorphanizing ORs. As mentioned earlier, in this case, several odorant mixtures can be tested. Mixtures eliciting a response should then be divided into individual odorants tested on other cells.
By targeting OSNs expressing defined ORs, this protocol provides powerful data about the ORs, ligands/ORs interactions, transduction pathway properties and to compare properties of defined populations of OSNs in different conditions.

Olfactory sensory neurons (OSN) represent the first step of olfactory perception. Located in the olfactory epithelium lining the nasal cavity in rodents, they transform the chemical information of odorants into action potentials sent through their axon to the brain. To better understand the olfactory coding mechanisms, it is necessary to characterize the transduction and membrane properties of OSNs. Until recently, most of the techniques used to characterize the properties of mammalian OSNs were carried out on dissociated OSNs1-4. The dissociation process uses various mechanical and chemical (i.e., enzymes) processes to free the OSNs from their environment. These processes induce a low number of available cells for recordings. This low number can be even more critical in the case of GFP labelled cells. Dissociation also removes the local cell-to-cell interactions between OSNs and other cells of the olfactory epithelium that may enhance survival and modulation of OSNs&#39; properties. In order to bypass the dissociation procedure, an intact preparation was developed5.
Each OSN expresses one olfactory receptor (OR) selected from a large multigene family6. There are ~1,000 ORs expressed in the main olfactory epithelium in the mouse. Due to the large number of OR in wild type animals, the chances to record OSNs expressing the same OR are very low. To overcome these limitations, gene targeted mice are available&#160;in which all OSNs expressing an identified OR are labeled with a fluorescent protein7-9. These labeled OSNs were used to do functional analysis in dissociated preparations7,10,11 with the drawbacks mentioned earlier. An intact epithelium preparation5 from genetically labeled mice therefore circumvents these issues. It allows the monitoring of the activity of OSNs expressing precisely defined ORs in an environment as close to in vivo as possible. Besides, patch-clamp recordings of OSNs also allow precise analysis of membrane properties, transduction pathway pharmacology, ligand/OR interactions. All these topics can hardly be analyzed using extracellular recordings. We used this technique to monitor the responses of OSNs expressing the odorant receptors SR1 and MOR2312,13. The feasibility of the technique was confirmed by other groups on MOR23 expressing OSNs14 as well as on other ORs expressing neurons15,16. The monitoring of a defined population of OSNs can lead to the analysis of their properties in many different contexts such as development14, aging17, odorant induced plasticity18, and the role of variations in the odorant receptor&#8217;s sequence in odor coding15. This protocol thus provides a powerful tool to monitor the functional properties of defined OSNs at the membrane level.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td colspan="4">heavy equipment</td>
		</tr>
		<tr>
			<td>vibration table with Faraday cage</td>
			<td>TMC</td>
			<td>63-500 SERIES</td>
			<td>required : isolates the recording system from vibrations induced by the environment (movements of experimenter, vibrations of equipment such as fans for cooling computers, etc); can also be purchased with a Faraday cage, or equipped by a custom made Faraday cage; this cage is recommended to avoid electric noise from the environment</td>
		</tr>
		<tr>
			<td colspan="4">optics</td>
		</tr>
		<tr>
			<td>microscope</td>
			<td>Olympus</td>
			<td>BX51WI</td>
			<td>upright microcope equipped with epifluorescence; fixed or moving stage depending on the user&#39;s preference</td>
		</tr>
		<tr>
			<td>objectives</td>
			<td>Olympus</td>
			<td>LUMPLFL40XW</td>
			<td>at least 2 objectives required: a 4x or 10x for coarse approach to the cell; and a 40x immersion long distance example Olympus LUMPLFL40XW/IR/0,8/WD:3.3 MM</td>
		</tr>
		<tr>
			<td>magnifier</td>
			<td>Olympus</td>
			<td>U-TVCAC</td>
			<td>ABSOLUTELY REQUIRED: placed in the light path between the objective and the camera; allows to magnify the image on the screen in order to reach precisely the knob with the recording electrode</td>
		</tr>
		<tr>
			<td>camera</td>
			<td>Olympus</td>
			<td>DP72</td>
			<td>a good camera is required to see the neurons in fluorescence as well as in bright field; the controlling software is simple and allows to take pictures and do live camera image to monitor the approach of the electrode to the cell. An ultrasensitive camera is not necessary</td>
		</tr>
		<tr>
			<td>filters</td>
			<td>Olympus/Chroma</td>
			<td></td>
			<td>depending on the fluorescent protein used in the mice; example for GFP: excitation : BP460-490: emission: HQ530/50m</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">recording electrodes /system</td>
		</tr>
		<tr>
			<td>amplifier</td>
			<td>HEKA</td>
			<td>EPC10 USB</td>
			<td>monitors the currents flowing through the recording electrode and also controls the puffing by sending a TTL signal to the spritzer; the EPC10 setup is controled by computer</td>
		</tr>
		<tr>
			<td>software</td>
			<td>HEKA</td>
			<td>Patchmaster</td>
			<td>controls the amplifier during the experiment</td>
		</tr>
		<tr>
			<td>micromanipulator</td>
			<td>Sutter</td>
			<td>MP225</td>
			<td>precision micromanipulator, allows precise movements down to 1/1Oth of a micrometer; this model is very stable; avoid hydraulic manipulators that may drift</td>
		</tr>
		<tr>
			<td>electrode puller</td>
			<td>sutter</td>
			<td>P97</td>
			<td>with a FT345-B wide trough filament;&#160; to prepare recording pipets of about 2&#181;m diameter with a long tip to reach the cells; the resistance should be 15 to 20Mohm with perforated patch clamp solution</td>
		</tr>
		<tr>
			<td>glass</td>
			<td>sutter</td>
			<td>BF120-69-10</td>
			<td>in our recording conditions, this glass is ideal for recording pipets</td>
		</tr>
		<tr>
			<td>recording chamber</td>
			<td>warner instruments</td>
			<td>RC-26G</td>
			<td>a chamber is needed to set the preparation under the microscope. To maintain the preparation in the center of the chamber, a net/anchor should be used.</td>
		</tr>
		<tr>
			<td colspan="4">stimulation</td>
		</tr>
		<tr>
			<td>glass</td>
			<td>WPI</td>
			<td>TW100F-4</td>
			<td>attached in groups of 7, these pipettes are used to prepare prepulled stimulating pipettes</td>
		</tr>
		<tr>
			<td>multibarrel puller</td>
			<td>MDI</td>
			<td>PMP-107-Z</td>
			<td>by association of pull and twist, this puller allows us to prepare puffing electrodes with 7 barrels</td>
		</tr>
		<tr>
			<td>precision pressure injector&#160;</td>
			<td>Toohey Company</td>
			<td>P/N T25-1-900 Single Channel&#160;&#160;&#160;</td>
			<td>this precision pressure injector&#160; controls the pressure ejected in the multibarrel puller; it is controlled manually or by the amplifier by a 5V&#160; TTLs</td>
		</tr>
		<tr>
			<td>micromanipulator</td>
			<td>Narishige</td>
			<td>YOU-1</td>
			<td>a coarse manipulator is enough to bring the puffing electrode close to the recording site</td>
		</tr>
		<tr>
			<td>tubings</td>
			<td>N/A</td>
			<td></td>
			<td>tygon tubing to bring the pressure from the puffer to the puffing pipette</td>
		</tr>
		<tr>
			<td colspan="4">solutions/perfusion/chemicals</td>
		</tr>
		<tr>
			<td>vacuum pump</td>
			<td>gardner denver</td>
			<td>300 series</td>
			<td>a vibrating membrane pump is more quiet and efficient than other types of pumps</td>
		</tr>
		<tr>
			<td>perfusion system</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>gravity perfusion system with polyethlylen tubing to bring in and out the external solution from the recording chamber</td>
		</tr>
		<tr>
			<td>nystatin</td>
			<td>Sigma-Aldrich</td>
			<td>N3503</td>
			<td>mandatory to perpare internal solution for perforated patch clamp</td>
		</tr>
		<tr>
			<td>DIMETHYL SULFOXIDE</td>
			<td>Sigma-Aldrich</td>
			<td>D5879</td>
			<td>used to disolve nystatin for internal solution for perforated patch&#160;</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>S9625</td>
			<td>extracellular solution</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>P4504</td>
			<td>intracellular/extracellular solution</td>
		</tr>
		<tr>
			<td>Calcium chloride di hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>C7902</td>
			<td>extracellular solution</td>
		</tr>
		<tr>
			<td>Sodium phosphate monobasic monohydrate (NaH2PO4)</td>
			<td>Sigma-Aldrich</td>
			<td>S9638</td>
			<td>extracellular solution</td>
		</tr>
		<tr>
			<td>Magnesium sulfate heptahydrate (MgSO4 7H2O)</td>
			<td>Sigma-Aldrich</td>
			<td>63140</td>
			<td>extracellular solution</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G8270</td>
			<td>extracellular solution</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>S6297</td>
			<td>extracellular solution</td>
		</tr>
		<tr>
			<td>EGTA (Ethylene glycol-bis(2-aminoethylether)-N,N,N&#8242;,N&#8242;-tetraacetic acid)</td>
			<td>Sigma-Aldrich</td>
			<td>E3889</td>
			<td>internal solution</td>
		</tr>
		<tr>
			<td>Potassium hydroxyde</td>
			<td>Sigma-Aldrich</td>
			<td>P1767</td>
			<td>internal solution</td>
		</tr>
		<tr>
			<td>MethylSulfoxide</td>
			<td>Sigma-Aldrich</td>
			<td>47,135-6</td>
			<td>intracellular solution</td>
		</tr>
		<tr>
			<td>Hepes-Na</td>
			<td>Sigma-Aldrich</td>
			<td>H7006</td>
			<td>intracellular solution</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma-Aldrich</td>
			<td>S0389</td>
			<td>intracellular solution</td>
		</tr>
	</tbody>
</table>

perforated patch-clamp recording of mouse olfactory sensory neurons in intact neuroepithelium: functional analysis of neurons expressing an identified odorant receptor

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of olfactory-driven behaviors and their modulation. These coding properties depend heavily on the initial interaction between odor molecules and the olfactory receptor (OR) expressed in the OSNs. The identity, specificity and ligand spectrum of the expressed OR are critical. The probability to find the ligand of the OR expressed in an OSN chosen randomly within the epithelium is very low. To address this challenge, this protocol uses genetically tagged mice expressing the fluorescent protein GFP under the control of the promoter of defined ORs. OSNs are located in a tight and organized epithelium lining the nasal cavity, with neighboring cells influencing their maturation and function. Here we describe a method to isolate an intact olfactory epithelium and record through patch-clamp recordings the properties of OSNs expressing defined odorant receptors. The protocol allows one to characterize OSN membrane properties while keeping the influence of the neighboring tissue. Analysis of patch-clamp results yields&#160;a precise quantification of ligand/OR interactions, transduction pathways and pharmacology, OSNs&#39; coding properties and their modulation at the membrane level.&#160;

The outcome of this protocol depends on the quality of the dissection. This dissection steps must be short (less than 10 to 15 min) and precise (i.e., to avoid damages of the epithelium). The Figure 1 illustrates how an ideal preparation looks like at different magnification levels. At a low magnification under bright field the different cell types (such as knobs of OSNs, supporting cells) are distinguishable (Figure 1A). At the highest magnification level, typically 80X to 160X, in bright field, the dendritic knobs of the OSNs should be clearly distinguishable from the supporting cells (Figure 1B). Under fluorescent light, only the dendritic knobs and cilia of GFP labelled cells are visible (Figure 1C). By comparing the 2 images, the labelled cells can be approached with the recording pipette (Figure 1D).
The temperature of the dissection solution and timing of the dissection are critical. The first part of the dissection, the preparation of the septum (section 3.1.1 to 3.2) should take place within 5 to 10 min in ice-cold solution. The final dissection (3.3) should last less than 5 min at room temperature. In case the dissection lasted for too long, or was performed in dissection solution too warm, the preparation invariably looks rapidly damaged: dendritic knobs are floating above the surface of the epithelium, and resemble dead cells.
Once a seal is reached and the cell opens under the effects of nystatin, typical voltage gated currents can be observed (Figure 2A). The shape and characteristics of these currents can be used to monitor the health of the cell: in a dying cell or if the seal&#8217;s quality is decreasing, these currents&#8217; amplitude will decrease. Under the current clamp configuration, action potentials can be recorded either spontaneously (Figure 2B) or by injection of a depolarizing current (Figure 2C). The firing properties induced by current injection characterize the excitability of the recorded neuron. The excitability of different OSNs&#8217; population can be compared (using classical frequency and ISI calculations).
A multibarrel pipette loaded with different odorants and/or different concentrations of odorants can be used to stimulate the cells by pressure ejection. This allows to measure the odorant induced responses. The odorants and concentrations must be chosen depending on the OR expressed in the cell of interest, for example Lyral is the ligand for MOR23, acetophenone for M71, eugenol for mOREG. In the experiment illustrated on the Figure 3, the recorded MOR23-IRES-tauEGFP neurons responded to different concentrations of Lyral, a ligand for MOR23, under current clamp mode (Figure 3A) or under the voltage clamp mode (Figure 3B). In voltage clamp mode recordings, different characteristics can be monitored to quantify the response (maximum amplitude, rise-time, total current elicited, etc.) as performed classically in electrophysiology. Using the maximum amplitude of the odorant-induced response, dose-response can be plotted and fitted using the Hill equation. These results provide information about the encoding properties of each OSN: detection threshold, temporal dynamic, dynamic range and saturation level. Examples of dose-response curves are shown on Figure 3C. All these details can be compared between individual OSNs to measure potential heterogeneity within a population; they can also be compared between OSNs with or without application of a treatment to measure potential modulation and plasticity.
<img alt="Figure 1" src="/files/ftp_upload/52652/52652fig1.jpg" />
<img alt="Figure 1B" src="/files/ftp_upload/52652/52652fig1B.jpg" /> 
Figure 1: Representative images of a healthy preparation. (A) Intact olfactory epithelium extracted from the nasal cavity of a SR1-IRES-tauGFP transgenic mouse observed under bright field condition at the 40X magnification. OSN dendritic knobs (black arrow heads) are enclosed in a mesh of supporting cells (SC) and Bowman glands (BG). (B) Dendritic knobs of SR1 expressing OSNs observed under bright field condition (white and red arrow heads). (C) The same field as in (B) under fluorescent light showing dendritic knobs of SR1 expressing OSNs. (D) Recording pipette approaching a SR1-expressing OSN under bright field. The red arrowhead represents the same SR1 OSN in (B-D). Scale bar: 5 &#181;m.
<img alt="Figure 2" src="/files/ftp_upload/52652/52652fig2.JPG" /> 
Figure 2: Representative membrane properties results obtained with the protocol: patch-clamp recordings on the dendritic knob of a SR1-IRES-tauGFP OSN. (A) Voltage gated currents elicited by increasing depolarizing steps from the membrane potential -67 mV&#160;to +40 mV. (B) Spontaneous activity recorded in the current clamp configuration; action potentials can be observed during the 15 sec recording epoch. (C) Action potentials elicited by a +7 pA excitatory current; this protocol provides information about the excitability of the cell.
<img alt="Figure 3A" src="/files/ftp_upload/52652/52652fig3A.jpg" />
<img alt="Figure 3B" src="/files/ftp_upload/52652/52652fig3B.jpg" />
<img alt="Figure 3C" src="/files/ftp_upload/52652/52652fig3C.jpg" /> 
Figure 3: Representative examples of odorant induced responses in a MOR23-IRES-tauEGFP neuron. Increasing concentrations of Lyral, a ligand of the MOR23 receptor, induce increasing responses both in current clamp (A) and in voltage clamp (B). The membrane potential was clamped at -67 mV. (C) Examples of individual dose-response curves acquired on three M71 neurons in response to increasing concentrations of acetophenone and fitted with the Hill equation.

Watch this Scientific Journal Video about Perforated Patch-clamp Recording of Mouse Olfactory Sensory Neurons in Intact Neuroepithelium: Functional Analysis of Neurons Expressing an Identified Odorant

Perforated Patch-clamp Recording of Mouse Olfactory Sensory Neurons in Intact Neuroepithelium: Functional Analysis of Neurons Expressing an Identified Odorant Receptor

Analyzing the physiological properties of olfactory sensory neurons still faces technical limitations. Here we record them through perforated patch-clamp in an intact preparation of the olfactory epithelium in gene-targeted mice. This technique allows the characterization of membrane properties and responses to specific ligands of neurons expressing defined olfactory receptors.

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of ...

perforated-patch-clamp-recording-mouse-olfactory-sensory-neurons

Nanyang Technological University

Research

JoVE Journal

Neuroscience

26.6K Views.  Université de Bourgogne. Analyzing the physiological properties of olfactory sensory neurons still faces technical limitations. Here we record them through perforated patch-clamp in an intact preparation of the olfactory epithelium in gene-targeted mice. This technique allows the characterization of membrane properties and responses to specific ligands of neurons expressing defined olfactory receptors.

Video: Perforated Patch-clamp Recording of Mouse Olfactory Sensory Neurons in Intact Neuroepithelium: Functional Analysis of Neurons Expressing an Identified Odorant Receptor

Analyzing Responses of Mouse Olfactory Sensory Neurons Using the Air-phase Electroolfactogram Recording

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and other social interactions. The electroolfactogram (EOG) recording is an informative, easy to conduct, and reliable method to assay olfactory function at the level of the olfactory epithelium. Since the 1956 description of the EOG by Ottoson in frogs1, the EOG recording has been applied in many vertebrates including salamanders, rabbits, rats, mice, and humans (reviewed by Scott and Scott-Johnson, 2002, ref. 2). The recent advances in genetic modification in mice have rekindled interest in recording the EOG for physiological characterization of olfactory function in knock-out and knock-in mice. EOG recordings have been successfully applied to demonstrate the central role of olfactory signal transduction components3-8, and more recently to characterize the contribution of certain regulatory mechanisms to OSN responses9-12.
Odorant detection occurs at the surface of the olfactory epithelium on the cilia of OSNs, where a signal transduction cascade leads to opening of ion channels, generating a current that flows into the cilia and depolarizes the membrane13. The EOG is the negative potential recorded extracellularly at the surface of the olfactory epithelium upon odorant stimulation, resulting from a summation of the potential changes caused by individual responsive OSNs in the recording field2. Comparison of the amplitude and kinetics of the EOG thus provide valuable information about how genetic modification and other experimental manipulations influence the molecular signaling underlying the OSN response to odor. 
Here we describe an air-phase EOG recording on a preparation of mouse olfactory turbinates. Briefly, after sacrificing the mouse, the olfactory turbinates are exposed by bisecting the head along the midline and removing the septum. The turbinate preparation is then placed in the recording setup, and a recording electrode is placed at the surface of the olfactory epithelium on one of the medial turbinates. A reference electrode is electrically connected to the tissue through a buffer solution. A continuous stream of humidified air is blown over the surface of the epithelium to keep it moist. The vapor of odorant solutions is puffed into the stream of humidified air to stimulate the epithelium. Responses are recorded and digitized for further analysis.

The electroolfactogram (EOG) recording is an informative, easy-to-conduct, and reliable way of assessing olfactory function at the level of the olfactory epithelium. This protocol describes a recording setup, mouse tissue preparation, data collection, and basic data analysis.

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and ...

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

Lentivirus-mediated Genetic Manipulation and Visualization of Olfactory Sensory Neurons in vivo

Development of a precise olfactory circuit relies on accurate projection of olfactory sensory neuron (OSN) axons to their synaptic 
targets in the olfactory bulb (OB). The molecular mechanisms of OSN axon growth and targeting are not well understood. Manipulating gene expression and subsequent 
visualizing of single OSN axons and their terminal arbor morphology have thus far been challenging. To study gene function at the single cell level within a specified 
time frame, we developed a lentiviral based technique to manipulate gene expression in OSNs in vivo. Lentiviral particles are delivered to OSNs by microinjection 
into the olfactory epithelium (OE). Expression cassettes are then permanently integrated into the genome of transduced OSNs. Green fluorescent protein expression 
identifies infected OSNs and outlines their entire morphology, including the axon terminal arbor. Due to the short turnaround time between microinjection and 
reporter detection, gene function studies can be focused within a very narrow period of development. With this method, we have detected GFP expression within as 
few as three days and as long as three months following injection. We have achieved both over-expression and shRNA mediated knock-down by lentiviral microinjection. 
This method provides detailed morphologies of OSN cell bodies and axons at the single cell level in vivo, and thus allows characterization of candidate gene function 
during olfactory development.

Lentivirus-mediated Genetic Manipulation and Visualization of Olfactory Sensory Neurons in vivo

We present a lentiviral technique for genetic manipulation and visualization of single olfactory sensory neuron axon and its terminal arborization in vivo.

Development of a precise olfactory circuit relies on accurate projection of olfactory sensory neuron (OSN) axons to their synaptic 
targets in the ...

Flash Photolysis of Caged Compounds in the Cilia of Olfactory Sensory  Neurons

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds1. Caged compounds are molecules made physiologically inactive by a chemical cage that can be broken by a flash of ultraviolet light. Here, we show how to obtain patch-clamp recordings combined with photolysis of caged compounds for the study of olfactory transduction in dissociated mouse olfactory sensory neurons. The process of olfactory transduction (Figure 1) takes place in the cilia of olfactory sensory neurons, where odorant binding to receptors leads to the increase of cAMP that opens cyclic nucleotide-gated (CNG) channels2. Ca entry through CNG channels activates Ca-activated Cl channels. We show how to dissociate neurons from the mouse olfactory epithelium3 and how to activate CNG channels or Ca-activated Cl channels by photolysis of caged cAMP4 or caged Ca5. We use a flash lamp6,7 to apply ultraviolet flashes to the ciliary region to uncage cAMP or Ca while patch-clamp recordings are taken to measure the current in the whole-cell voltage-clamp configuration8-11.

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds. Here, we show how to obtain patch-clamp recordings combined with photolysis of caged cAMP or caged Ca for the study of olfactory transduction in dissociated mouse olfactory sensory neurons.

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds1. ...

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

Odorant-induced Responses Recorded from Olfactory Receptor Neurons using the Suction Pipette Technique

Animals sample the odorous environment around them through the chemosensory systems located in the nasal cavity. Chemosensory signals affect complex behaviors such as food choice, predator, conspecific and mate recognition and other socially relevant cues. Olfactory receptor neurons (ORNs) are located in the dorsal part of the nasal cavity embedded in the olfactory epithelium. These bipolar neurons send an axon to the olfactory bulb (see Fig. 1, Reisert &amp; Zhao1, originally published in the Journal of General Physiology) and extend a single dendrite to the epithelial border from where cilia radiate into the mucus that covers the olfactory epithelium. The cilia contain the signal transduction machinery that ultimately leads to excitatory current influx through the ciliary transduction channels, a cyclic nucleotide-gated (CNG) channel and a Ca2+-activated Cl- channel (Fig. 1). The ensuing depolarization triggers action potential generation at the cell body2-4.
In this video we describe the use of the "suction pipette technique" to record odorant-induced responses from ORNs. This method was originally developed to record from rod photoreceptors5 and a variant of this method can be found at jove.com modified to record from mouse cone photoreceptors6. The suction pipette technique was later adapted to also record from ORNs7,8. Briefly, following dissociation of the olfactory epithelium and cell isolation, the entire cell body of an ORN is sucked into the tip of a recording pipette. The dendrite and the cilia remain exposed to the bath solution and thus accessible to solution changes to enable e.g. odorant or pharmacological blocker application. In this configuration, no access to the intracellular environment is gained (no whole-cell voltage clamp) and the intracellular voltage remains free to vary. This allows the simultaneous recording of the slow receptor current that originates at the cilia and fast action potentials fired by the cell body9. The difference in kinetics between these two signals allows them to be separated using different filter settings. This technique can be used on any wild type or knockout mouse or to record selectively from ORNs that also express GFP to label specific subsets of ORNs, e.g. expressing a given odorant receptor or ion channel.

Olfactory receptor neurons (ORNs) convert odor signals first into a receptor current that in turn triggers action potentials that are conveyed to second order neurons in the olfactory bulb. Here we describe the suction pipette technique to record simultaneously the odorant-induced receptor current and action potentials from mouse ORNs.

Animals sample the odorous environment around them through the chemosensory systems located in the nasal cavity. Chemosensory signals affect complex ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

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 ...

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

The marine gastropod mollusk Aplysia californica has a venerable history as a model of nervous system function, with particular significance in studies of learning and memory. The typical preparations for such studies are ones in which the sensory and motoneurons are left intact in a minimally dissected animal, or a technically elaborate neuronal co-culture of individual sensory and motoneurons. Less common is the isolated neuronal preparation in which small clusters of nominally homogeneous neurons are dissociated into single cells in short term culture. Such isolated cells are useful for the biophysical characterization of ion currents using patch clamp techniques, and targeted modulation of these conductances. A protocol for preparing such cultures is described. The protocol takes advantage of the easily identifiable glutamatergic sensory neurons of the pleural and buccal ganglia, and describes their dissociation and minimal maintenance in culture for several days without serum.

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

We describe the dissection of the nervous system of the marine sea hare Aplysia after anesthesia, the isolation of neurons for short term-tissue culture, and recordings of single cell ion currents via the patch clamp technique.

The marine gastropod mollusk Aplysia californica has a venerable  history as a model of nervous system function, with particular significance in  studies ...

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

The freshwater leech, Hirudo medicinalis, is a versatile model organism that has been used to address scientific questions in the fields of neurophysiology, neuroethology, and developmental biology. The goal of this report is to consolidate experimental techniques from the leech system into a single article that will be of use to physiologists with expertise in other nervous system preparations, or to biology students with little or no electrophysiology experience. We demonstrate how to dissect the leech for recording intracellularly from identified neural circuits in the ganglion. Next we show how individual cells of known function can be removed from the ganglion to be cultured in a Petri dish, and how to record from those neurons in culture. Then we demonstrate how to prepare a patch of innervated skin to be used for mapping sensory or motor fields. These leech preparations are still widely used to address basic electrical properties of neural networks, behavior, synaptogenesis, and development. They are also an appropriate training module for neuroscience or physiology teaching laboratories.

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

This article describes three nervous system preparations using leeches: intracellular recording from neurons in ventral ganglia, culturing neurons from ventral ganglia, and recording from a patch of innervated skin to map sensory fields.