This research was supported by R00 DC011780 (JPM: NINDS, NIH), F30 DC011673 (GFH: NINDS, NIH) and UT Southwestern startup funds (JPM).

All experiments were carried out in accordance with protocols approved by the UT Southwestern Institutional Animal Care and Use Committee, and were chosen so as to minimize stress, discomfort, and pain experienced by the experimental animals.
1. Dissection Chamber
A custom dissection chamber and small, thin plastic plank are required for best results (Figure 1). Construct or obtain such a chamber in advance of attempting this protocol.
2. Dissection Solutions
<ol>
	<li>Prepare 1 L of standard artificial cerebrospinal fluid (aCSF) for use in Steps 3.22-5.4. Add NaCl 125 mM, KCl 2.5 mM, CaCl2 2 mM, MgCl2 1 mM, NaHCO3 25 mM, NaH2PO4 1.25 mM, myo-inositol 3 mM, sodium pyruvate 2 mM, sodium ascorbate 0.4 mM, and glucose 25 mM to 1 L of pyrogen free water.</li>
	<li>Prepare the initial dissection aCSF (200 ml) for use in steps 3.3-3.22. Take 200 ml of Standard aCSF and raise the MgCl2 concentration by 9 mM by adding 0.366 g of MgCl2 hexahydrate.</li>
	<li>Prepare standard Ringer&#8217;s solution to carry stimuli to the VNO through the cannula for use in steps 4.11-5.4 containing NaCl 115 mM, KCl 5 mM, CaCl2, 2 mM, MgCl2, 2 mM, NaHCO3 25 mM, HEPES 10 mM, glucose 10 mM.</li>
	<li>Bubble the 0.8 L of standard aCSF and 0.5 L of Ringer&#8217;s with 95% O2, 5% CO2 gas in a 37 &#176;C water bath for a minimum of 15 min until use.</li>
	<li>Bubble the 200 ml of initial dissection aCSF with 95% O2, 5% CO2 gas on ice for 15-20 min until it reaches 4 &#176;C.</li>
</ol>
3. Primary Dissection
<ol>
	<li>Prepare the dissection area. Place three 35 mm Petri dishes, oxygenated aCSF, decapitation scissors, curved dissecting scissors, straight dissecting scissors, Adson forceps, a #11 scalpel blade and handle and a razor blade on ice.</li>
	<li>After oxygenated aCSF has cooled to 4 &#176;C, deeply anesthetize the animal in a desiccation chamber using 2-5 ml Isothesia (99.9% isofluorane). Perform tail and foot pinch tests to ensure the animal is deeply anesthetized.</li>
	<li>Once deep anesthesia is confirmed, decapitate the animal and immediately place the head in a Petri dish filled with the chilled dissection aCSF.&#160;NOTE: Immerse the tissue in chilled dissection aCSF to keep it cool throughout the procedure when not in use.</li>
	<li>Remove the lower aspect (ventral pharynx, lower jaw, and tongue) with a curved scissors.</li>
	<li>Using Adson forceps deglove (peel) the superficial skin and muscle attachments from the skull. Peel the scalp from caudal to rostral until the only site of attachment is over the front teeth. Cut the tissue at the attachment point using a curved scissors. Remove the eyes by grasping it with forceps and pull it away from the preparation. Near the end of the degloving procedure, avoid excess force on the delicate nasal cartilage by using the Adson forceps to free muscle attachments from the upper jaw bones.&#160;NOTE: Detach the scalp from the caudal aspect of the skull with straight scissors.</li>
	<li>With the straight scissors, cut down the midline of the skull from caudal to rostral until the scissor tips are a few mm into the frontal bones (just prior to the rhinal sinus).</li>
	<li>Remove the parietal and frontal bones of the skull using Adson forceps.&#160;NOTE: Pieces of the frontal bone will often remain attached to the nasal bones caudal to the rhinal sinus. Remove these pieces with care taken not to contact the olfactory bulbs.</li>
	<li>Use the scalpel to make a coronal cut through the frontal lobe 2 mm caudal to the sinuses and then remove the brain caudal to the cut. Ensure the tip of the scalpel contacts the bony contour of the ventral skull to completely sever the posterior tissue.&#160;NOTE: This enables removal of majority of the brain without putting mechanical stress on the lateral olfactory tracts or olfactory bulbs.</li>
	<li>Remove the exposed caudal aspects of the ventral skull with straight scissors. Leave the sinuses and remaining nervous tissue.</li>
	<li>Remove the zygomatic arch and facial muscles on the anatomical right side of the skull using straight scissors</li>
	<li>Turn the snout over (ventral side up) to expose the roof of the mouth.</li>
	<li>Cut and remove the palate immediately caudal to the incisors. Do this by inserting the tip of the scalpel blade under the palate parallel to the roof of the mouth and then move the blade towards the incisors.</li>
	<li>Grasp the freed (rostral) edge of the palate with Adson forceps and remove by pulling caudally.</li>
	<li>On the anatomical left side of the skull, use the scalpel blade to extend the palatine foramen caudally, starting from the small foramen near the molars. Achieve this by inserting the tip of the scalpel blade into the void near the molars and rotating the wrist.&#160;NOTE: Use extreme caution to only use the tip of the blade &#8211; a deep cut can damage the septal tissue.</li>
	<li>Using the tip of the scalpel blade, cut from the palatine foramen through the incisors, starting rostral to the anatomical left vomeronasal organ. Use multiple scoring cuts. Do not damage the septal tissue and vomeronasal nerves by inserting the blade too deep.</li>
	<li>Turn the tissue so the dorsal skull is facing up and then orient the straight razor blade vertically (parallel to the midline) at the caudal edge of the preparation.</li>
	<li>At the ventral edge of the tissue, touch the cutting edge of the razor blade immediately to the left of the midline. Gently rock the blade and move along the left palatine foramen (cut region introduced in Step 3.14).</li>
	<li>At the dorsal edge of the tissue, place the cutting edge of the razor immediately to the left of the midline (less than 1 mm).</li>
	<li>Keeping the blade parallel to the midline, rock the razor blade slowly with pressure towards the anterior of the tissue. Notice resistance at the cribriform plate. Stop immediately after breaking through this resistance.&#160;NOTE: At this point, the right hemisphere is loosened from the left hemisphere. The only remaining connection is between the hemispheres along the dorsal snout anterior to the cribriform plate.</li>
	<li>Separate the hemispheres by grasping them near the caudal sides with gloved fingers and gently rotating them laterally and anteriorly.&#160;NOTE: This step is a source of experimental variability, especially in mice with deviated or brittle snouts (e.g., older mice). Also, while attempting to rotate the hemispheres away from each other, if strong resistance is encountered score the dorsal snout (anterior to the cribriform plate) to weaken the connective tissues. While doing so, take extreme caution not to insert the scalpel deeply, which can damage the septal tissue and vomeronasal nerves.</li>
	<li>Apply a small amount (50-100 &#181;l) of tissue glue to the plank and gently place the lateral edge of the right hemisphere onto the glue using the Adson forceps. Remove excess moisture from the lateral side of the preparation using a paper towel to prevent premature glue polymerization and loose adhesion to the plank.&#160;NOTE: Apply a few drops of the chilled dissection aCSF on the sample to accelerate the glue polymerization after tissue is placed onto the glue. This ensures that the tissue remains tightly held to the plank.</li>
	<li>Place a small amount (3-5 mm diameter, 2 mm deep) of vacuum grease in the middle of the secondary dissection chamber and place the side of the plank opposite the tissue firmly against the grease. Immediately fill the dissection chamber with chilled dissection aCSF, transfer to the fine dissection area and begin perfusion with oxygenated standard aCSF. Orient the plank so that the olfactory bulb is directly in the stream of freshly oxygenated standard aCSF.</li>
</ol>
4. Secondary Dissection
<ol>
	<li>Remove any visible contralateral (left) olfactory bulb or cortical tissue from the preparation using fine forceps. Pinch together the fine forceps (forming a semi blunt point), and then place it at the dorsal and caudal portion of the tissue near the midline. Gently run the pinched forceps along the midline, peeling the tissue away from the right hemisphere slowly in anterior motion. Remove all of the contralateral tissue, including the contralateral olfactory bulb tissue. Take extreme care not to stab through the tissue, which can cause damage to the AOB or vomeronasal nerve.</li>
	<li>Observe the white dura mater along the dorsal/medial surface of the olfactory bulb. Tear this dura mater along the dorsal/caudal edge of the olfactory bulb by grasping at the whitest portion of the dura with two pairs of fine forceps and pull them away from that point. The dura is tightly wrapped around the olfactory bulbs, and can easily itself slice through tissue while it is being removed. Avoid damage to the medial surface of the olfactory bulb and the AOB itself.&#160;NOTE: If the dura resists tearing easily at this point, introduce a small cut with fine spring tip dissection scissors to facilitate separation.</li>
	<li>Slowly and carefully peel away the frontal cortex with fine forceps without damaging the underlying olfactory bulbs. Observe a collection of blood vessels appearing as a semi-transparent net immediately caudal to the AOB. Remove this net with fine forceps to facilitate electrode penetration in electrophysiological experiments. Grasp the net with forceps as it separates from the AOB during frontal cortex retraction and remove it by pulling caudally and medially, being careful not to touch the AOB. Once the frontal cortex has been separated from the olfactory bulbs, remove it by cutting with forceps ventral and posterior to the AOB.&#160;NOTE: Take extreme care with net removal, for if it is done too roughly the glomerular layer can become damaged.</li>
	<li>In the exposed contralateral nasal cavity, remove any remaining contralateral septal tissue (flimsy yellowish sheet containing blood vessels) using the fine forceps. Grasp the tissue at the caudal/dorsal region (near the septal bone) and then pull/lift the tissue towards the anterior side.&#160;NOTE: The contralateral septal tissue can be inadvertently removed during the hemisection step (Step 3.20). In such case, observe a white, avascular septal cartilage and the slightly transparent, vascularized septal bone. If this is the case, Step 4.4 can be skipped.</li>
	<li>Carefully remove the contralateral VNO by running a single point of the fine forceps between the septal cartilage and the VNO &#8220;wing&#8221; (a thin piece of bony tissue that runs along the dorsal edge of both VNOs.
	<ol>
		<li>Following the wing detachment from the cartilage, move the tip of forceps ventrally into the contralateral VNO near the midline. Repeat this step at several locations along the rostral/caudal axis of the contralateral VNO to loosen it, enabling it to be removed by grasping with forceps and lifting away.</li>
	</ol>
	</li>
	<li>Prepare to remove the septal cartilage by making a cut at the ventral/caudal edge (where it narrows to a white avascular strip running along the ventral edge of the septal bone) with fine forceps.</li>
	<li>Remove the septal cartilage by pinching together the fine forceps to make a semi blunt point. Identify the cut made in Step 4.6, insert the pinched forceps behind (lateral) to that cut, and then slowly lift the cartilage away from the septal tissue and proceeding rostrally.&#160;NOTE: Do not disrupt the underlying septal tissue. As the cartilage is lifted, observe the underlying sheet of septal tissue stretch due to loose adhesions to the cartilage. A periodic, gentle, and stable movement of the pinched fine forceps along this site of loose adhesion can greatly facilitate successful separation of the two structures.</li>
	<li>Use a pair of #3 forceps with a bent tip to remove the septal bone. Approach the septal bone at an angle nearly parallel to the plane of the septal bone. Insert the curved tine between the septal tissue and septal bone. Grasp the bone with the forceps and gently move the bone back and forth until it cracks near the cribriform plate.&#160;NOTE: In some cases, the septal bone will resist breaking near the cribriform plate. In this case, use fine forceps to gently score the septal bone along the dorsal/ventral axis just anterior to the cribriform plate, then repeat Step 4.8.</li>
	<li>Remove the white cartilage just rostral to the VNO by pinching with one pair of fine forceps at the entrance and pulling rostrally with another pair of fine forceps.</li>
	<li>Attach the polyimide cannula to a fast perfusion device and start the flow of Ringer&#8217;s solution (0.1-0.3 ml/min) through the polyimide cannula.&#160;NOTE: Measure flow rate with an in-line, small volume flow meter. Use of computer controlled pneumatic stimulus switching device reduces flow rate changes during stimulation. Compare neural responses to test stimuli with responses to a mock stimulus (control Ringer&#8217;s solution) to ensure responses are stimulus specific.</li>
	<li>Orient the cannula parallel to the VNO entrance. When the cannula is satisfactorily parallel to the VNO entrance, watch as the outlet pressure causes the VNO to &#8220;inflate,&#8221; leading to evacuation of the blood vessel. Immediately insert the cannula into the VNO, keeping the angle of the cannula constant so that it remains parallel to the VNO opening.&#160;NOTE: It can be helpful to hold the cannula with one pair of fine forceps against the plastic plank and use another pair of fine forceps to angle the end of the cannula slightly upwards. If one inadvertently places the cannula behind the VNO, the outflow can also cause the VNO blood vessel to evacuate, leading to the impression a proper cannulation was performed. Proper location can be confirmed by incorporating a dye in the Ringer&#8217;s solution. When the cannula is properly placed, the dye should only exit via the VNO entrance. Additionally, one can confirm proper placement by ensuring that the cannula is easily visible through the semi-transparent medial aspect of the VNO.</li>
	<li>Move the plastic plank slowly until the AOB is in the direct flow of the standard aCSF, being careful not to dislodge the cannula from the VNO. The dissection is complete and assessment of physiological function can commence immediately. Step 5 briefly describes one approach for making such an assessment.</li>
</ol>
5. Evaluation
<ol>
	<li>Penetrate the surface of the AOB with a 2-4 M&#937; borosilicate glass microelectrode attached to an extracellular amplifier and oscilloscope.</li>
	<li>Slowly advance the microelectrode to a depth of 100-200 &#181;m from the surface at a rate of 50 &#181;m/min.&#160;NOTE: At this tissue depth, one will encounter occasional spontaneous action potential waveforms exemplified by sharp, brief (~1-2 msec) deflections in the microelectrode voltage.</li>
	<li>Upon encountering an action potential waveform, stop advancing the microelectrode.</li>
	<li>Observe and/or record the action potential rate while changing the stimulus delivery solution from control Ringer&#8217;s to a known activator of VSN activity (e.g., Ringer&#8217;s solution containing 50 mM KCl). If the dissection was successful, a stimulus-dependent change in action potential rate or local field potential can be observed.&#160;NOTE: Not all AOB neurons will respond to every stimulus with an increase in firing rate (including Ringer&#8217;s solution containing 50 mM KCl). A stimulus-dependent decrease in firing rate also indicates functional connectivity and a successful dissection. In the event that two or more neurons are encountered that are unresponsive to VNO stimulation, or if no action potentials are observed after multiple electrode penetrations, this suggests an unsuccessful dissection. If this is the case, a new dissection can be initiated at Step 3.</li>
</ol>

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The authors have no significant conflicts of interest to disclose.

The VNO-AOB ex vivo preparation described in this protocol is a useful alternative to anesthetized in vivo5-7 and acute live slice17 experiments of AOB function. Unlike acute AOB slice experiments, which also expose circuit elements for electrophysiological and optical recordings, this preparation retains all sensory afferents and intra-AOB connections. Although this can also be said of anesthetized in vivo approaches, the presence of anesthetics necessarily alter...

Sensory processing in the mammalian brain typically spans multiple reciprocally-connected neuronal circuits, each of which extracts particular features from sensory input. In sensory pathways, early information processing is vital for normal perception and behavior. In the accessory olfactory system (AOS), the accessory olfactory bulb (AOB) is the principal neural circuit linking the sensory periphery to downstream structures that dictate hormonal balance1,2, aggression3, and arousal4. As such, information processing within this circuit is strongly linked to changes in animal behavior.
The accessory olfactory bulb is located in mice and rats at the dorsal/caudal/posterior aspect of the main olfactory bulb (MOB) beneath the dense, vascularized rhinal sinus. The AOB receives afferent innervation from axons of peripheral vomeronasal sensory neurons (VSNs) that reside in the vomeronasal organ (VNO), a small blind-ended tube in the anterior snout just above the soft palate. These axons traverse the delicate sheet of septal tissue at the medial boundary of the nasal passages. Several studies have probed AOB neural responses to sources of AOS odors (such as mouse urine) in vivo using anesthetized mice5-7 or freely-exploring animals8. The heroic anesthetized in vivo studies involved (a) tracheotomies to ensure deep anesthesia and prevent the aspiration of liquid stimuli5-7, (b) stimulation of the sympathetic cervical ganglion6 or direct cannulation of the vomeronasal organ5,7 to introduce nonvolatile odors and (c) craniotomies with or without frontal lobe ablations to allow electrode advancement into the AOB6. Awake/behaving studies8-10 involved surgical implantation of a microdrive. In sum, these experimental paradigms are powerful, but extremely difficult and often require anesthesia.
Interestingly, several studies attempted to maintain sensory structures and downstream neural circuits alive outside the body (ex vivo) with some success11-15. Because the connections between the VNO and AOB remain ipsilateral, and because the midline septal tissue can be exposed to oxygenated superfusate in a single hemisphere, we sought to develop such a single-hemisphere ex vivo approach to isolate these structures while maintaining their functional connectivity. We recently succeeded in achieving this goal16. This preparation keeps both the VNO and AOB alive and functionally connected for at least 4 - 6 hr because both the axons (along the midline soft septal tissue) and AOB are relatively shallow &#60;600 &#181;m features that are accessible to superfused oxygenated artificial cerebrospinal fluid (aCSF). This VNO-AOB ex vivo preparation allows introduction of controlled stimuli into the VNO via a thin cannula, and direct visual access to the small AOB for targeted electrode placement and/or live fluorescence microscopy. This method is advantageous if one desires to study these circuits in the absence of anesthetics. Because this approach severs centrifugal connections, it is not well suited to inquiries into centrifugal modulation of AOB function. The VNO-AOB ex vivo preparation is difficult to learn, but once achieved produces a reliable platform upon which to investigate circuit organization, information processing, and neural plasticity in this powerful sensory circuit.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Straight Scissors</td>
			<td>Fine Science Tools</td>
			<td>14002-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Scissors-Straight</td>
			<td>Fine Science Tools</td>
			<td>14060-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Scissors-Curved</td>
			<td>Fine Science Tools</td>
			<td>14061-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Adson Forceps</td>
			<td>Fine Science Tools</td>
			<td>11006-12</td>
			<td></td>
		</tr>
		<tr>
			<td>#3 Scalpel Handle</td>
			<td>Fine Science Tools</td>
			<td>10003-12</td>
			<td></td>
		</tr>
		<tr>
			<td>#11 Scalpel Blades</td>
			<td>Fisher Scientific</td>
			<td>3120030</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Carbon Steel Razor Blades</td>
			<td>Fisher Scientific</td>
			<td>12-640</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm Petri Dish</td>
			<td>Fisher Scientific</td>
			<td>08-772-21</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection Chamber</td>
			<td>Custom&#160;</td>
			<td>N/A</td>
			<td>See Fig. 1</td>
		</tr>
		<tr>
			<td>Delrin plastic plank 0.6 cm x 1.5 cm x 0.1 cm</td>
			<td>Custom&#160;</td>
			<td>N/A</td>
			<td></td>
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			<td>1/16&#34; Male Luer</td>
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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.

Achieving success with this preparation takes extensive practice, and has several steps at which it can fail. One should expect to require many attempts before achieving success. The custom dissection chamber is required for successful completion of this protocol, and should be obtained prior to starting the later stages of the dissection. The chamber design presented in Figure 1 is sufficient for this purpose, and can be made of relatively inexpensive plastics with minimal machining demands. If one lack...

Watch this Scientific Journal Video about Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb at JoVE.com

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.

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 ...

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Research

JoVE Journal

Neuroscience

10.7K Views.  UT Southwestern Medical Center. 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.

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

Multielectrode Array Recordings of the Vomeronasal Epithelium

Understanding neural circuits requires methods to record from many neurons simultaneously. For in vitro studies, one currently available technology is planar multielectrode array (MEA) recording. Here we document the use of MEAs to study the mouse vomeronasal organ (VNO), which plays an essential role in the detection of pheromones and social cues via a diverse population of sensory neurons expressing hundreds of types of receptors. Combining MEA recording with a robotic liquid handler to deliver chemical stimuli, the sensory responses of a large and diverse population of neurons can be recorded. The preparation allows us to remove the intact neuroepithelium of the VNO from the mouse and stimulate with a battery of chemicals or potential ligands while monitoring the electrical activity of the neurons for several hours. Therefore, this technique serves as a useful method for assessing ligand activity as well as exploring the properties of receptor neurons. We present the techniques needed to prepare the vomeronasal epithelium, MEA recording, and chemical stimulation.

Multielectrode array (MEA) recordings provide a method for studying the electrical activity of large populations of neurons. Here, we present the details of a MEA preparation to record from the mouse vomeronasal epithelium while simultaneously stimulating the tissue.

Understanding neural circuits requires methods to record from many neurons simultaneously. For in vitro studies, one currently available technology is ...

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

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

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

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

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

Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines

In vivo electroporation is a powerful method for delivering DNA expression plasmids, RNAi reagents, and morpholino anti-sense oligonucleotides to specific regions of developing embryos, including those of C. elegans, chick, Xenopus, zebrafish, and mouse 1. In zebrafish, in vivo electroporation has been shown to have excellent spatial and temporal resolution for the delivery of these reagents 2-7. The temporal resolution of this method is important because it allows for incorporation of these reagents at specific stages in development. Furthermore, because expression from electroporated vectors occurs within 6 hours 7, this method is more timely than transgenic approaches. While the spatial resolution can be extremely precise when targeting a single cell 2, 6, it is often preferable to incorporate reagents into a specific cell population within a tissue or structure. When targeting multiple cells, in vivo electroporation is efficient for delivery to a specific region of the embryo; however, particularly within the developing nervous system, it is difficult to target specific cell types solely through spatially discrete electroporation. Alternatively, enhancer trap transgenic lines offer excellent cell type-specific expression of transgenes 8. Here we describe an approach that combines transgenic Gal4-based enhancer trap lines 8 with spatially discrete in vivo electroporation 7, 9 to specifically target developing neurons of the zebrafish olfactory bulb. The Et(zic4:Gal4TA4,UAS:mCherry)hzm5 (formerly GA80_9) enhancer trap line previously described 8, displays targeted transgenic expression of mCherry mediated by a zebrafish optimized Gal4 (KalTA4) transcriptional activator in multiple regions of the developing brain including hindbrain, cerebellum, forebrain, and the olfactory bulb. To target GFP expression specifically to the olfactory bulb, a plasmid with the coding sequence of GFP under control of multiple Gal4 binding sites (UAS) was electroporated into the anterior end of the forebrain at 24-28 hours post-fertilization (hpf). Although this method incorporates plasmid DNA into multiple regions of the forebrain, GFP expression is only induced in cells transgenically expressing the KalTA4 transcription factor. Thus, by using the GA080_9 transgenic line, this approach led to GFP expression exclusively in the developing olfactory bulb. GFP expressing cells targeted through this approach showed typical axonal projections, as previously described for mitral cells of the olfactory bulb 10. This method could also be used for targeted delivery of other reagents including short-hairpin RNA interference expression plasmids, which would provide a method for spatially and temporally discrete loss-of-function analysis.

Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines

The temporal and spatial resolution of genetic manipulations determines the spectrum of biological phenomena that they can perturb. Here we use temporally and spatially discrete in vivo electroporation, combined with transgenic lines of zebrafish, to induce expression of a GFP transgene specifically in neurons of the developing olfactory bulb.

In vivo electroporation is a powerful method for delivering DNA expression plasmids, RNAi reagents, and morpholino anti-sense oligonucleotides to specific ...

Imaging Odor-Evoked Activities in the Mouse Olfactory Bulb using Optical Reflectance and Autofluorescence Signals

In the brain, sensory stimulation activates distributed populations of neurons among functional modules which participate to the coding of the stimulus. Functional optical imaging techniques are advantageous to visualize the activation of these modules in sensory cortices with high spatial resolution. In this context, endogenous optical signals that arise from molecular mechanisms linked to neuroenergetics are valuable sources of contrast to record spatial maps of sensory stimuli over wide fields in the rodent brain.
Here, we present two techniques based on changes of endogenous optical properties of the brain tissue during activation. First the intrinsic optical signals (IOS) are produced by a local alteration in red light reflectance due to: (i) absorption by changes in blood oxygenation level and blood volume (ii) photon scattering. The use of in vivo IOS to record spatial maps started in the mid 1980's with the observation of optical maps of whisker barrels in the rat and the orientation columns in the cat visual cortex1. IOS imaging of the surface of the rodent main olfactory bulb (OB) in response to odorants was later demonstrated by Larry Katz's group2. The second approach relies on flavoprotein autofluorescence signals (FAS) due to changes in the redox state of these mitochondrial metabolic intermediates. More precisely, the technique is based on the green fluorescence due to oxidized state of flavoproteins when the tissue is excited with blue light. Although such signals were probably among the first fluorescent molecules recorded for the study of brain activity by the pioneer studies of Britton Chances and colleagues3, it was not until recently that they have been used for mapping of brain activation in vivo. FAS imaging was first applied to the somatosensory cortex in rodents in response to hindpaw stimulation by Katsuei Shibuki's group4.
The olfactory system is of central importance for the survival of the vast majority of living species because it allows efficient detection and identification of chemical substances in the environment (food, predators). The OB is the first relay of olfactory information processing in the brain. It receives afferent projections from the olfactory primary sensory neurons that detect volatile odorant molecules. Each sensory neuron expresses only one type of odorant receptor and neurons carrying the same type of receptor send their nerve processes to the same well-defined microregions of &tilde;100&mu;m3 constituted of discrete neuropil, the olfactory glomerulus (Fig. 1). In the last decade, IOS imaging has fostered the functional exploration of the OB5, 6, 7 which has become one of the most studied sensory structures. The mapping of OB activity with FAS imaging has not been performed yet.
Here, we show the successive steps of an efficient protocol for IOS and FAS imaging to map odor-evoked activities in the mouse OB.

This article presents the protocols of intrinsic optical signals and flavoproteins autofluorescence signals imaging to map odor-evoked activities at the surface of the olfactory bulb in mice.

In the brain, sensory stimulation activates distributed populations of neurons among functional modules which participate to the coding of the stimulus. ...

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 ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. Choroid plexus epithelial cells (CPECs) covering the surface of choroid plexus tissue harbor multiple unique cilia, but most of the functions of these cilia remain to be investigated. To uncover the function of CPEC cilia with particular reference to their motility, an ex vivo observation system was developed to monitor ciliary motility during embryonic, perinatal and postnatal periods. The choroid plexus was dissected out of the brain ventricle and observed under a video-enhanced contrast microscope equipped with differential interference contrast optics. Under this condition, a simple and quantitative method was developed to analyze the motile profiles of CPEC cilia for several hours ex vivo. Next, the morphological changes of cilia during development were observed by scanning electron microscopy to elucidate the relationship between the morphological maturity of cilia and motility. Interestingly, this method could delineate changes in the number and length of cilia, which peaked at postnatal day (P) 2, while the beating frequency reached a maximum at P10, followed by abrupt cessation at P14. These techniques will enable elucidation of the functions of cilia in various tissues. While related techniques have been published in a previous report1, the current study focuses on detailed techniques to observe the motility and morphology of CPEC cilia ex vivo.

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

In this study, a detailed light microscopic technique was optimized for real-time observation and analysis of the motion of CPEC cilia ex vivo together with an electron microscopic method for ultrastructural analysis.

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. ...

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. However, there is piling evidence that olfaction is not solely limited to chemical sensitivity, but also includes temperature sensitivity. Premetamorphic Xenopus laevis are translucent animals, with protruding nasal cavities deprived of the cribriform plate separating the nose and the olfactory bulb. These characteristics make them well suited for studying olfaction, and particularly thermosensitivity. The present article describes the complete procedure for measuring temperature responses in the olfactory bulb of X. laevis larvae. Firstly, the electroporation of olfactory receptor neurons (ORNs) is performed with spectrally distinct dyes loaded into the nasal cavities in order to stain their axon terminals in the bulbar neuropil. The differential staining between left and right receptor neurons serves to identify the &#947;-glomerulus as the only structure innervated by contralateral presynaptic afferents. Secondly, the electroporation is combined with focal bolus loading in the olfactory bulb in order to stain mitral cells and their dendrites. The 3D brain volume is then scanned under line-illumination microscopy for the acquisition of fast calcium imaging data while small temperature drops are induced at the olfactory epithelium. Lastly, the post-acquisition analysis allows the morphological reconstruction of the thermosensitive network comprising the &#947;-glomerulus and its innervating mitral cells, based on specific temperature-induced Ca2+ traces. Using chemical odorants as stimuli in addition to temperature jumps enables the comparison between thermosensitive and chemosensitive networks in the olfactory bulb.

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

Here we describe a protocol for measuring and analyzing temperature responses in the olfactory bulb of Xenopus laevis. Olfactory receptor neurons and mitral cells are differentially stained, after which calcium changes are recorded, reflecting a sensitivity of some neural networks in the bulb to temperature drops induced at the nose.

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. ...

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.

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 ...

Quadruple Immunostaining of the Olfactory Bulb for Visualization of Olfactory Sensory Axon Molecular Identity Codes

The mouse olfactory system is often used to study mechanisms of neural circuit formation&#12288;because of its simple anatomical structure. An Olfactory Sensory Neuron (OSN) is a bipolar cell with a single dendrite and a single unbranched axon. An OSN expresses only one Olfactory Receptor (OR) gene, OSNs expressing a given type of OR converge their axons to a few sets of invariant glomeruli in the Olfactory Bulb (OB). A remarkable feature of OSN projection is that the expressed ORs play instructive roles in axonal projection. ORs regulate the expression of multiple axon-sorting molecules and generate the combinatorial molecular code of axon-sorting molecules at the OSN axon termini. Thus, to understand the molecular mechanisms of OR-specific axon guidance mechanisms, it is vital to characterize their expression profiles at the OSN axon termini within the same glomerulus. The aim of this article was to introduce methods for collecting as many glomeruli as possible on a single OB section and for performing immunostaining using multiple antibodies. This would allow the comparison and analysis of the expression patterns of axon-sorting molecules without staining variation between OB sections.

Olfactory sensory neurons express a wide variety of axon-sorting molecules to establish proper neural circuitry. This protocol describes an immunohistochemical staining method to visualize combinatorial expressions of axon-sorting molecules at the axon termini of olfactory sensory neurons.

The mouse olfactory system is often used to study mechanisms of neural circuit formation&#12288;because of its simple anatomical structure. An Olfactory ...