We thank Ivan Rodriguez and Benoit von der Weid for generating the FPR-rs3-i-venus mouse line, their constructive criticism and fruitful discussions. This work was funded by grants of the Volkswagen Foundation (I/83533), the Deutsche Forschungsgemeinschaft (SP724/6-1) and by the Excellence Initiative of the German federal and state governments. MS is a Lichtenberg Professor of the Volkswagen Foundation.

All animal procedures were in compliance with local and European Union legislation on the protection of animals used for experimental purposes (Directive 86/609/EEC) and with recommendations put forward by the Federation of European Laboratory Animal Science Associations (FELASA). Both C57BL/6 mice and Fpr-rs3-i-Venus mice were housed in groups of both sexes at room temperature on a 12 hr light/dark cycle with food and water available ad libitum. For experiments young adults (6-20 weeks) of either sex were used. No obvious gender-dependent differences were observed.
1. Solution Preparation
<ol>
	<li>Prepare extracellular solution S1: 4-(2-Hydroxy-ethyl)piperazine-1-ethanesulfonic acid (HEPES) buffered extracellular solution containing (in mM) 145 NaCl, 5 KCl, 1 CaCl2, 1 MgCl2, 10 HEPES; pH = 7.3 (adjusted with NaOH); osmolarity = 300 mOsm (adjusted with glucose).</li>
	<li>Prepare extracellular solution S2: Carbogen-oxygenated (95% O2, 5% CO2) extracellular solution containing (in mM) 125 NaCl, 25 NaHCO3, 5 KCl, 1 CaCl2, 1 MgSO4, 5 BES; pH = 7.3; osmolarity = 300 mOsm (adjusted with glucose).</li>
	<li>Prepare 4% low-gelling temperature agarose solution (S3) for tissue embedding: 4% low-gelling temperature agarose. Dissolve 2 g low-gelling agarose in 50 ml of S1 in a 100 ml glass laboratory bottle together with a magnetic stirrer. To melt the agarose, put the bottle in a water bath (with lid not tightly closed) at 70 &#176;C for approximately 20 min or until solution becomes transparent while stirring. Cool down and keep melted agarose in a second water bath at 42 &#176;C until further use.</li>
	<li>Prepare intracellular solution S4: Pipette solution containing (in mM) 143 KCl, 2 KOH, 1 EGTA, 0.3 CaCl2 (free Ca2+ = 110 nM), 10 HEPES, 2 MgATP, 1 NaGTP; pH = 7.1 (adjusted with KOH); osmolarity = 290 mOsm.</li>
	<li>Modify/adjust composition of S1, S2 and S4 according to individual experimental design (e.g., pharmacological blockers to isolate certain types of voltage-activated currents).</li>
</ol>
2. Workspace Preparation
<ol>
	<li>Fill oxygenating slice storing chamber with S2 at least 30 min prior to dissection, place on ice for temperature and pH adjustment and oxygenate solution continuously.</li>
	<li>Fill reservoir for slice superfusion at recording setup with S2 and oxygenate continuously.</li>
	<li>Prior to dissection, fill vibratome chamber with S2 and arrange crushed ice around the chamber. Alternatively, transfer spare ice cold oxygenated solution from oxygenating chamber into vibratome chamber and keep oxygenating continuously.</li>
	<li>Arrange surgical tools and consumables.</li>
	<li>Place ice gel pack under dissecting microscope, cover with a paper towel to prevent VNO tissue from freezing to bottom of the dish.</li>
	<li>Clean razor blade by briefly rinsing in 70% ethanol and distilled water and mount to vibratome. Replace for every slicing session.</li>
</ol>
3. VNO Dissection and Embedding
<ol>
	<li>Sacrifice animal by brief exposure to CO2 and decapitate using sharp surgical scissors. Note: As time from sacrificing the animal to putting the VNO capsule on ice is critical, minimize the time to less than 2 min. To maintain tissue viability, embed both fully dissected VNOs in agarose in less than 30 min.</li>
	<li>Remove the lower jaw with large surgical scissors. Enter through the mouth cavity and cut the mandible bones and muscle of each side separately.</li>
	<li>Place the remaining part of the head upside down in the large Petri dish.</li>
	<li>Pull away the skin of the upper jaw and around the tip of the nose with medium forceps to gain better access to the incisors.</li>
	<li>Use bone scissors to cut away the largest part of the incisors at a ~45&#176; angle in the rostral direction (Figure 1A). This will ease removal of the VNO capsule from the nasal cavity. 
	Note: Do not cut to the root of the tooth to prevent damage to the tip of the VNO capsule.</li>
	<li>Grab the rigid upper palate at its rostral part with medium forceps and carefully peel back in one piece at a flat angle (Figure 1B). 
	Note: Repeatedly rinse with ice-cold S2.</li>
	<li>Use micro spring scissors to cut the bony fusion between the tip of the vomer bone and the jaw. Insert the scissor tips with the curved part of the tip pointing outward away from the VNO and carefully cut the bone in small steps on both the left and right side lateral to the VNO capsule.</li>
	<li>To remove the VNO capsule, use micro spring scissors to cut through the vomer bone at the caudal part and carefully lift the vomer bone out of the nasal cavity using medium forceps. Immediately transfer the VNO to a small petri dish under a stereo microscope on an ice gel pack where the remaining steps of the dissection will be performed.</li>
	<li>Rinse the VNO in a small amount of ice-cold S2 to prevent the tissue from drying out.</li>
	<li>Separate the cartilaginous capsules that contain the VNO soft tissues by grabbing the back of the vomer bone with medium forceps. Position the capsule for a dorsal view so that a split between both VNOs becomes visible (Figure 1Di).</li>
	<li>Use the tip of fine forceps to separate both cartilage VNO capsules from the central bone while keeping the vomer bone pinned down at the rear part. 
	Note: Use forceps only at the rim of the cartilage capsule and be very careful not to pierce through the cartilage as the delicate sensory epithelium is easily damaged.</li>
	<li>Once the two VNOs are separated, start removing the cartilage encapsulating the first VNO.</li>
	<li>Grab the top rim of the capsule with one fine forceps and split away the cartilage wall that was previously attached to the vomer bone (medial side).</li>
	<li>To remove remaining cartilage turn the VNO with its curved lateral side to the bottom of the dish and securely pin down the cartilage on one side using forceps. Carefully move the second fine forceps from the back side at a very flat angle between cartilage and VNO to loosen the connection between tissue and cartilage.</li>
	<li>Slowly peel the VNO away from the cartilage by holding it at its caudal tip, to avoid damaging the sensory epithelium.</li>
	<li>Once the VNO is levered from the capsule, make sure to remove all remaining small cartilage parts as any remaining pieces of cartilage will detach the tissue from surrounding agarose during the slicing process.</li>
	<li>Place a small drop of ice-cold S2 on the first dissected VNO to prevent tissue damage. A large blood vessel on the lateral side will become visible indicating that the organ is still intact and was not grossly damaged during the dissection (Figure 2Aii). In case the blood vessel got damaged, it will still be worthwhile to carry on with slicing the VNO as long as the overall morphology was not clearly impaired.</li>
	<li>Immediately dissect the second VNO.</li>
	<li>To embed the VNOs, fill both small petri dishes to the rim with melted S3 (stored in water bath at 42 &#176;C; see 1.3).</li>
	<li>Hold the VNO on the broader caudal end with fine forceps to avoid damage to the sensory epithelium.</li>
	<li>Immerse the VNO in the agarose and move it horizontally back and forth several times to remove the film of extracellular solution as well as any air bubbles from its surface.</li>
	<li>Position the VNO vertically with the caudal tip facing the bottom of the dish. Instead of pinching the VNO with the forceps directly, adjust orientation by moving the forceps tip in close proximity to the VNO. 
	Note: Orientation during embedding is crucial as it determines slice plane and accessibility to sensory neurons during the experiment.</li>
	<li>Place dishes on gel ice pack and wait until agarose has solidified. 
	Note: Do not change VNO orientation once the agarose has started solidifying as this will detach the tissue from the surrounding agarose.</li>
</ol>
4. Coronal VNO Tissue Slicing
<ol>
	<li>Use small spatula to remove agarose block from the small dish into the lid of a large Petri dish, flip the agarose upside down leaving the caudal tip of the VNO facing upwards.</li>
	<li>Cut the block into a pyramidal shape using a surgical scalpel (3-4 mm at the tip, 8-10 mm at the bottom). Take care not to damage the embedded tissue.</li>
	<li>Use super glue to fix the pyramid-shaped block to the center of the vibratome specimen plate and wait ~1 min for the glue to dry completely.</li>
	<li>Transfer the specimen plate to the slicing chamber and prepare the second VNO, accordingly. Keep plate with the second specimen at 4 &#176;C until use.</li>
	<li>Use a vibrating blade microtome with the following settings: thickness: 150-200 &#181;m; speed: 3.5 a.u. = 0.15 mm/sec; frequency: 7.5 a.u. = 75 Hz. Transfer slices to oxygenating chamber until use after briefly inspecting slice morphology under the dissecting microscope. Slices can be kept for several hr.</li>
</ol>
5. Single-cell Electrophysiological Recordings
<ol>
	<li>For recordings, use an upright fixed-stage microscope equipped with water immersion objectives, Dodt or infrared differential interfering contrast (IR-DIC), and epi-fluorescence as well as a cooled CCD-camera. For data acquisition, use a patch-clamp amplifier, head stage, AD/DA interface board and a PC (including recording software).</li>
	<li>Prepare a stock of 10-20 patch pipettes (4-7 M&#8486;). Pull pipettes from borosilicate glass capillaries (1.50 mm OD/0.86 mm ID) using a micropipette puller and fire-polish with a microforge. 
	Note: Fire polishing the pipette tips is crucial when patching the rather small VSNs. This will help to prevent rupturing the cell membrane when applying negative pressure in order to obtain a high resistance seal. After polishing the pipette opening should be approximately 1 &#181;m.</li>
	<li>Keep pipettes in a pipette storage jar to prevent damage and dust accumulation until use.</li>
	<li>Prepare perfusion system by filling solution reservoirs and tubing according to experimental design. 
	Note: Remove air bubbles from tubing completely as they will strongly interfere with electrophysiological recordings.</li>
	<li>Adjust pressure to achieve ~3 ml/min flow. 
	Note: High pressures will cause movement of the tissue slice as well as termination of electrophysiological recordings.</li>
	<li>Transfer VNO slice to imaging chamber and fix the slice position using stainless steel anchor wired with 0.1 mm thick synthetic fibers (Figure 2B-C). 
	Note: Do not cover the tissue slice with one of the synthetic fiber threads but rather the agarose surrounding the slice.</li>
	<li>Transfer imaging chamber to recording setup and continuously superfuse slice with oxygenated S2 at room temperature via a bath application.</li>
	<li>Adjust the suction capillary to the surface of the solution to create a slow suction for constant exchange of bath solution. Adjust the 8-in-1 multi-barrel &#34;perfusion pencil&#34; above and close to the non-sensory part of the VNO slice that contains the blood vessel (Figure 2C, 3A)50. It will be beneficial to arrange perfusion pencil and recording pipette to be facing the slice from opposite directions.</li>
	<li>Connect reference electrode and bath solution using an L-shaped agar bridge (filled with 150 mM KCl).</li>
	<li>Fill patch pipette with pipette solution S4.</li>
	<li>Mount the pipette over the silver chloride-coated patch electrode connected to the head stage without scraping off the coating and attach firmly.</li>
	<li>Apply slight positive pressure (approximately 1 ml on a 10 ml plastic syringe) to the patch pipette before entering the bath.</li>
	<li>Lower the pipette into the bath using micro manipulators far enough to be able to submerge the objective without hitting it (Figure 2D).</li>
	<li>Monitor pipette resistance (Rpip, between 4-7 M&#8486;) using electrophysiology software connected to the head stage. 
	Note: If Rpip is &#60;4 M&#8486; the glass tip is broken. If R pip is &#62;10 M&#8486;, the tip is most likely clogged and the pipette must be replaced.</li>
	<li>Visualize the VNO slice with a CCD-camera using infrared-optimized differential interference contrast (DIC) and identify FPR-rs3-i-Venus expressing cells (or similarly labeled neurons) using fluorescence illumination and an appropriate filter cube.</li>
	<li>Focus and target fluorescent or non-fluorescent cells depending on experimental design.</li>
	<li>To approach the cell body, use hand wheels for maximum sensitivity. Due to positive pressure, a small dent in the cell soma membrane becomes visible once the pipette tip is in close proximity.</li>
	<li>Release positive pressure and apply slight negative pressure to suck in the cell membrane in order to gain a high resistance seal (1-20 G&#8486;). Apply short and gentle suction to disrupt the cell membrane and establish the whole-cell configuration.</li>
	<li>Monitor access resistance constantly during experiment. 
	Note: Only include neurons exhibiting small and stable access resistance (&#8804;3% of Rinput; change &#60;20% over the course of the experiment) into analysis.</li>
</ol>

The authors have nothing to disclose.

The VNO is a chemosensory structure that detects semiochemicals. To date, the majority of vomeronasal receptors remains to be deorphanized as only few receptor-ligand pairs have been identified. Among those, V1rb2 was described to be specifically activated by the male urinary pheromone 2-heptanone30, V2rp5 to be activated by the male specific pheromone ESP157 as well as V2r1b and V2rf2 to be activated by the MHC peptides SYFPEITHI48 and SEIDLILGY58, respectively. A prerequisite...

Most animals rely heavily on their chemical senses to interact with their surroundings. The sense of smell plays an essential role for finding and evaluating food, avoiding predators and locating suitable mating partners. In most mammals, the olfactory system consists of at least four anatomically and functionally distinct peripheral subsystems: the main olfactory epithelium1,2, the Grueneberg ganglion3,4, the septal organ of Masera5,6 and the vomeronasal organ. The VNO comprises the peripheral sensory structure of the accessory olfactory system (AOS), which plays a major role in detecting chemical cues that convey information about identity, gender, social rank and sexual state7-10. The VNO is located at the base of the nasal septum right above the palate. In mice, it is a bilateral blind-ending tube enclosed in a cartilaginous capsule11-13. The organ consists of both a crescent-shaped medial sensory epithelium that harbors the VSNs and of a non-sensory part on the lateral side. Between both epithelia lies a mucus-filled lumen which is connected to the nasal cavity via the narrow vomeronasal duct14. A large lateral blood vessel in the non-sensory tissue provides a vascular pumping mechanism to facilitate entry of relatively large, mostly non-volatile molecules such as peptides or small proteins into the VNO lumen through negative pressure15,16. The structural components of the VNO are present at birth and the organ reaches adult size shortly before puberty17. However, whether the rodent AOS is already functional in juveniles is still subject to debate18-20.
VSNs are distinguished by both their epithelial location and the type of receptor they express. VSNs show a bipolar morphology with an unmyelinated axon and a single apical dendrite that protrudes towards the lumen and ends in a microvillous dendritic knob. VSN axons fasciculate to form the vomeronasal nerve that leaves the cartilaginous capsule at the dorso-caudal end, ascends along the septum, passes the cribriform plate and projects to the accessory olfactory bulb (AOB)21,22. The vomeronasal sensory epithelium consists of two layers: the apical layer is located closer to the luminal side and harbors both V1R- and all but one type of FPR-rs-expressing neurons. These neurons coexpress the G-protein &#945;-subunit G&#945;i2 and project to the anterior part of the AOB23-25. Sensory neurons located in the more basal layer express V2Rs or FPR-rs1 alongside G&#945;o and send their axons to the posterior region of the AOB26-28.
Vomeronasal neurons are likely activated by rather small semiochemicals29-33 (V1Rs) or proteinaceous compounds34-38 (V2Rs) that are secreted into various bodily fluids such as urine, saliva and tear fluid37,39-41. In situ experiments have shown that VSNs are also activated by formylated peptides and various antimicrobial/inflammation-linked compounds25,42. Moreover, heterologously expressed FPR-rs proteins share agonist spectra with FPRs expressed in the immune system, indicating a potential role as detectors for sickness in conspecifics or spoiled food sources25 (see reference43).
Fundamental to understanding receptor-ligand relationships and downstream signaling cascades in specific VSN populations is a detailed evaluation of their basic biophysical characteristics in a native environment. In the past, the analysis of cellular signaling has greatly benefitted from genetically modified animals that mark a defined population of neurons by coexpressing a fluorescent marker protein30,44-49. In this protocol, a transgenic mouse line that expresses FPR-rs3 together with a fluorescent marker (Fpr-rs3-i-Venus) is used. This approach exemplifies how to employ such a genetically modified mouse strain to perform electrophysiological analysis of an optically identifiable cell population using single neuron patch-clamp recordings in acute coronal VNO tissue slices. An air pressure-driven multi-barrel perfusion system for sensory stimuli and pharmacological agents allows quick, reversible and focal neuronal stimulation or inhibition during recordings. Whole-cell recordings in slice preparations allow for a detailed analysis of intrinsic properties, voltage-activated conductances, as well as action potential discharge patterns in the cell&#39;s native environment.

<table border="0" cellpadding="0" cellspacing="0" width="846">
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			<td height="21" style="height:21px;width:323px;">Large petri dish, 90 mm</td>
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			<td style="width:275px;">removing skin and palate</td>
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			<td height="42" style="height:42px;width:323px;">Objective&#160;</td>
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			<td height="21" style="height:21px;width:323px;">Oscilloscope</td>
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			<td height="21" style="height:21px;width:323px;">Osmometer&#160;</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:323px;">Perfusion system 8-in-1</td>
			<td style="width:133px;">AutoMate Scientific</td>
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			<td style="width:275px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:323px;">pH Meter five easy</td>
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			<td style="width:116px;"></td>
			<td style="width:275px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:323px;">Pipette storage jar</td>
			<td style="width:133px;">World Precision Instruments</td>
			<td style="width:116px;">e212</td>
			<td style="width:275px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:323px;">Recording chamber&#160;</td>
			<td style="width:133px;">Luigs &#38; Neumann</td>
			<td style="width:116px;">Slice mini chamber</td>
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		<tr height="42">
			<td height="42" style="height:42px;width:323px;">Razor blades</td>
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		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:323px;">Oxygenating slice storage chamber; alternative commercial chambers are e.g. BSK1 Brain Slice Keeper (Digitimer) or the Pre-chamber (BSC-PC; Warner Instruments)</td>
			<td style="width:133px;">custom-made</td>
			<td style="width:116px;"></td>
			<td style="width:275px;"></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:323px;">Stereo microscope</td>
			<td style="width:133px;">Leica Microsystems</td>
			<td style="width:116px;">S4E</td>
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			<td height="21" style="height:21px;width:323px;">Trigger interface&#160;</td>
			<td style="width:133px;">HEKA Elektronik</td>
			<td style="width:116px;">TIB-14 S</td>
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			<td height="21" style="height:21px;width:323px;">Vibratome&#160;</td>
			<td style="width:133px;">Leica Microsystems</td>
			<td style="width:116px;">VT 1000 S</td>
			<td style="width:275px;"></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:323px;">Water bath&#160;</td>
			<td style="width:133px;">Memmert</td>
			<td style="width:116px;">WNB 45</td>
			<td style="width:275px;"></td>
		</tr>
	</tbody>
</table>

in-depth physiological analysis of defined cell populations in acute tissue slices of the mouse vomeronasal organ

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory neurons (VSNs) express a specific type of G protein-coupled receptor (GPCR) from at least three different chemoreceptor gene families allowing sensitive and specific detection of chemosensory cues. These families comprise the V1r and V2r gene families as well as the formyl peptide receptor (FPR)-related sequence (Fpr-rs) family of putative chemoreceptor genes. In order to understand the physiology of vomeronasal receptor-ligand interactions and downstream signaling, it is essential to identify the biophysical properties inherent to each specific class of VSNs.
The physiological approach described here allows identification and in-depth analysis of a defined population of sensory neurons using a transgenic mouse line (Fpr-rs3-i-Venus). The use of this protocol, however, is not restricted to this specific line and thus can easily be extended to other genetically modified lines or wild type animals.

To gain insight into the biophysical and physiological properties of defined cell populations, we perform acute coronal tissue slices of the mouse VNO (Figure 1-2). After dissection, slices can be kept in ice-cold oxygenated extracellular solution (S2) for several hr. At the recording setup, a constant exchange with fresh oxygenated solution (Figure 2D) ensures tissue viability throughout the experiment. We here employ a transg...

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In-depth Physiological Analysis of Defined Cell Populations in Acute Tissue Slices of the Mouse Vomeronasal Organ

Here, we describe a physiological approach that allows identification and in-depth analysis of a defined population of sensory neurons in acute coronal tissue slices of the mouse vomeronasal organ using whole-cell patch-clamp recordings.

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory ...

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JoVE Journal

Neuroscience

7.5K Views.  RWTH Aachen University. Here, we describe a physiological approach that allows identification and in-depth analysis of a defined population of sensory neurons in acute coronal tissue slices of the mouse vomeronasal organ using whole-cell patch-clamp recordings. 

Video: In-depth Physiological Analysis of Defined Cell Populations in Acute Tissue Slices of the Mouse Vomeronasal Organ

Imaging Pheromone Sensing in a Mouse Vomeronasal Acute Tissue Slice Preparation

Peter Karlson and Martin L&uuml;scher used the term pheromone for the first time in 19591 to describe chemicals used for intra-species communication. Pheromones are volatile or non-volatile short-lived molecules2 secreted and/or contained in biological fluids3,4, such as urine, a liquid known to be a main source of pheromones3. Pheromonal communication is implicated in a variety of key animal modalities such as kin interactions5,6, hierarchical organisations3 and sexual interactions7,8 and are consequently directly correlated with the survival of a given species9,10,11. In mice, the ability to detect pheromones is principally mediated by the vomeronasal organ (VNO)10,12, a paired structure located at the base of the nasal cavity, and enclosed in a cartilaginous capsule. Each VNO has a tubular shape with a lumen13,14 allowing the contact with the external chemical world. The sensory neuroepithelium is principally composed of vomeronasal bipolar sensory neurons (VSNs)15. Each VSN extends a single dendrite to the lumen ending in a large dendritic knob bearing up to 100 microvilli implicated in chemical detection16. Numerous subpopulations of VSNs are present. They are differentiated by the chemoreceptor they express and thus possibly by the ligand(s) they recognize17,18. Two main vomeronasal receptor families, V1Rs and V2Rs19,20,21,22, are composed respectively by 24023 and 12024 members and are expressed in separate layers of the neuroepithelium. Olfactory receptors (ORs)25 and formyl peptide receptors (FPRs)26,27 are also expressed in VSNs.
Whether or not these neuronal subpopulations use the same downstream signalling pathway for sensing pheromones is unknown. Despite a major role played by a calcium-permeable channel (TRPC2) present in the microvilli of mature neurons28 TRPC2 independent transduction channels have been suggested6,29. Due to the high number of neuronal subpopulations and the peculiar morphology of the organ, pharmacological and physiological investigations of the signalling elements present in the VNO are complex.
Here, we present an acute tissue slice preparation of the mouse VNO for performing calcium imaging investigations. This physiological approach allows observations, in the natural environment of a living tissue, of general or individual subpopulations of VSNs previously loaded with Fura-2AM, a calcium dye. This method is also convenient for studying any GFP-tagged pheromone receptor and is adaptable for the use of other fluorescent calcium probes. As an example, we use here a VG mouse line30, in which the translation of the pheromone V1rb2 receptor is linked to the expression of GFP by a polycistronic strategy.

In mice, the ability to detect pheromones is principally mediated by the vomeronasal organ (VNO). Here, an acute tissue slice preparation of VNO for performing calcium imaging is described. This physiological approach allows observations of subpopulations and/or individual neurons in a living tissue and is convenient for receptor-ligand identification.

Peter Karlson and Martin L&uuml;scher used the term pheromone for the first time in 19591 to describe chemicals used for intra-species communication. ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

Electrophysiological Characterization of GFP-Expressing Cell Populations in the Intact Retina

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack conspicuous morphological features or show a low population density. This applies particularly to retinal amacrine cells, an exceptionally multiform class of interneurons that comprise roughly 30 subtypes in mammals1. Though being a crucial part of the visual processing by shaping the retinal output2, most of these subtypes have not been studied up to now in a functional context because encountering these cells with a recording electrode is a rare event.
Recently, a multitude of transgenic mouse lines is available that express fluorescent markers like green fluorescent protein (GFP) under the control of promoters for membrane receptors or enzymes that are specific to only a subset of neurons in a given tissue3,4. These pre-labeled cells are therefore accessible to directed microelectrode targeting under microscopic control, permitting the systematic study of their physiological properties in situ. However, excitation of fluorescent markers is accompanied by the risk of phototoxicity for the living tissue. In the retina, this approach is additionally hampered by the problem that excitation light causes appropriate stimulation of the photoreceptors, thus inflicting photopigment bleaching and transferring the retinal circuits into a light-adapted condition. These drawbacks are overcome by using infrared excitation delivered by a mode-locked laser in short pulses of the femtosecond range. Two-photon excitation provides energy sufficient for fluorophore excitation and at the same time restricts the excitation to a small tissue volume minimizing the hazards of photodamage5. Also, it leaves the retina responsive to visual stimuli since infrared light (&gt;850 nm) is only poorly absorbed by photopigments6.
In this article we demonstrate the use of a transgenic mouse retina to attain electrophysiological in situ recordings from GFP-expressing cells that are visually targeted by two-photon excitation. The retina is prepared and maintained in darkness and can be subjected to optical stimuli which are projected through the condenser of the microscope (Figure 1). Patch-clamp recording of light responses can be combined with dye filling to reveal the morphology and to check for gap junction-mediated dye coupling to neighboring cells, so that the target cell can by studied on different experimental levels.

This article depicts the recording of individual cells from fluorescently tagged neuronal populations in the intact mouse retina. By using two-photon infrared excitation transgenetically labeled cells were targeted for patch-clamp recording to study their light responses, receptive field properties, and morphology.

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

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 Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to neuroblasts able to move along the rostral migratory stream (RMS) towards the olfactory bulb (OB). This long-distance migration is required for the subsequent maturation of newborn neurons in the OB, but the molecular mechanisms regulating this process are still unclear. Investigating the signaling pathways controlling neuroblast motility may not only help understand a fundamental step in neurogenesis, but also have therapeutic regenerative potential, given the ability of these neuroblasts to target brain sites affected by injury, stroke, or degeneration.
In this manuscript we describe a detailed protocol for in vivo postnatal electroporation and subsequent time-lapse imaging of neuroblast migration in the mouse RMS. Postnatal electroporation can efficiently transfect SVZ progenitor cells, which in turn generate neuroblasts migrating along the RMS. Using confocal spinning disk time-lapse microscopy on acute brain slice cultures, neuroblast migration can be monitored in an environment closely resembling the in vivo condition. Moreover, neuroblast motility can be tracked and quantitatively analyzed. As an example, we describe how to use in vivo postnatal electroporation of a GFP-expressing plasmid to label and visualize neuroblasts migrating along the RMS. Electroporation of shRNA or CRE recombinase-expressing plasmids in conditional knockout mice employing the LoxP system can also be used to target genes of interest. Pharmacological manipulation of acute brain slice cultures can be performed to investigate the role of different signaling molecules in neuroblast migration. By coupling in vivo electroporation with time-lapse imaging, we hope to understand the molecular mechanisms controlling neuroblast motility and contribute to the development of novel approaches to promote brain repair.

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

Neuroblast migration is a fundamental event in postnatal neurogenesis. We describe a protocol for efficient labeling of neuroblasts by in vivo postnatal electroporation and subsequent visualization of their migration using time-lapse imaging of acute brain slices. We include a description for the quantitative analysis of neuroblast dynamics by video tracking.

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

Slice It Hot: Acute Adult Brain Slicing in Physiological Temperature

Here we present a protocol for preparation of acute brain slices. This procedure is a critical element for electrophysiological patch-clamp experiments that largely determines the quality of results. It has been shown that omitting the cooling step during cutting procedure is beneficial in obtaining healthy slices and cells, especially when dealing with highly myelinated brain structures from mature animals. Even though the precise mechanism whereby elevated temperature supports neural health can only be speculated upon, it stands to reason that, whenever possible, the temperature in which the slicing is performed should be close to physiological conditions to prevent temperature related artifacts. Another important advantage of this method is the simplicity of the procedure and therefore the short preparation time. In the demonstrated method adult mice are used but the same procedure can be applied with younger mice as well as rats. Also, the following patch clamp experiment is performed on horizontal cerebellar slices, but the same procedure can also be used in other planes as well as other posterior areas of the brain.

In this paper we show a method for preparing acute brain slices in physiological temperature, using a conventional physiological solution without special modifications for the cutting (such as adding sucrose) and without intracardial perfusion of the animal before slice preparation.

Here we present a protocol for preparation of acute brain slices. This procedure is a critical element for electrophysiological patch-clamp experiments ...

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have validated the utility of this method for enhancing neuronal preservation and overall brain slice viability. The implementation of this technique by early adopters has facilitated detailed investigations into brain function using diverse experimental applications and spanning a wide range of animal ages, brain regions, and cell types. Steps are outlined for carrying out the protective recovery brain slice technique using an optimized NMDG artificial cerebrospinal fluid (aCSF) media formulation and enhanced procedure to reliably obtain healthy brain slices for patch clamp electrophysiology. With this updated approach, a substantial improvement is observed in the speed and reliability of gigaohm seal formation during targeted patch clamp recording experiments while maintaining excellent neuronal preservation, thereby facilitating challenging experimental applications. Representative results are provided from multi-neuron patch clamp recording experiments to assay synaptic connectivity in neocortical brain slices prepared from young adult transgenic mice and mature adult human neurosurgical specimens. Furthermore, the optimized NMDG protective recovery method of brain slicing is compatible with both juvenile and adult animals, thus resolving a limitation of the original methodology. In summary, a single media formulation and brain slicing procedure can be implemented across various species and ages to achieve excellent viability and tissue preservation.

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have ...