This work was funded through an NIH R15 AREA grant to John Horne (NIMH; R15MH083221).

1. Transgenic embryos 
<ol>
	<li>Here we are using a transgenic line of zebrafish created through a transposon-mediated enhancer trap method 8. The transgenic insert includes two coding sequences: KalTA4, a zebrafish-optimized version of the Gal4 transcriptional activator, and the red fluorescent protein, mCherry. The enhancer trap method yields lines of transgenic zebrafish that express this transgenic cassette under control of an endogenous enhancer sequence. Thus, depending on the identity of the specific enhancer sequence, expression of both KalTA4 and mCherry is localized to specific regions of the developing brain. For the Et(zic4:Gal4TA4,UAS:mCherry)hzm5 line that we are using here, expression is localized to several sites in the hindbrain, forebrain, and, of particular importance here, the olfactory bulb 8 (see also Fig. 1). Given this pattern of expression, and the localization of the cassette within the genome, the transgenes are predicted to be under control of the zic4 promoter 8 (see also <a href="http://www.helmholtz-muenchen.de/en/idg/groups/neuroimaging/lines_distel/main">http://www.helmholtz-muenchen.de/en/idg/groups/neuroimaging/lines_distel/main</a>.).</li>
	<li>Adult Et(zic4:Gal4TA4,UAS:mCherry)hzm5 fish heterozygous for the transgenic insert are typically mated with WT (AB) fish, leading to approximately 50% transgenic offspring. (For a higher percentage of transgenic offspring, two heterozygotes can be mated.)</li> 
	<li>In order to block pigment formation and facilitate fluorescent imaging, treat transgenic embryos with 100 &mu;M N-phenylthiourea (PTU) beginning at 6-8 hpf.</li>
</ol>
2. Mounting embryos 
<ol>
	<li>At 24-28 hpf, transfer embryos to egg water without PTU. Identify transgenic embryos by expression of mCherry using a fluorescent microscope. Remove the chorionic membrane from transgenic embryos using fine forceps. Transfer embryos freed from their chorion to a Petri dish containing electroporation buffer plus 0.02% tricaine [Electroporation Buffer: 180 mM NaCl, 5 mM KCl, 1.8 mM CaCl2 , 5 mM HEPES, pH 7.2] 4. Allow 2 minutes for the anesthetic to take effect before proceeding to the mounting step.</li>
 	<li>Prepare a 0.5% solution of low-melting-point agarose in electroporation buffer plus tricaine by microwaving the solution, and then placing the solution into a 34-37&deg;C incubator. Once the temperature of the agarose solution has equilibrated, place a large drop (&tilde;500 &mu;l) of the agarose solution to the center of a 60 mm Petri dish. Add 6-8 anesthetized embryos to this drop of agarose. </li>
	<li> Using fine forceps (or the fine ends of cut microloader pipet tips), position the embryos such that they are all facing in the same direction and aligned in a vertical row. To solidify the agarose, and trap the embryos in position, place the dish onto a large Petri dish with 2-3 mm of ice.</li>
	<li> Once the agarose has solidified, flood the dish with electroporation buffer plus tricaine, ensuring that the solution completely covers the solidified agarose drop.</li>
</ol>
3. Micro-injection of DNA expression plasmid
<ol>
<li>Prepare the GFP expression plasmid using a midi- or maxi-prep plasmid isolation kit (e.g. Qiagen). Suspend the plasmid to a final concentration of 0.5 &mu;g/&mu;l in sterile water plus 0.03% phenol red. For the olfactory bulb targeting experiment described here, we are using an expression plasmid with the coding sequence of EGFP under the control of multiple Gal4 binding sites (14 tandem UAS sequences and the basal fish promoter, E1b). Using this plasmid, only cells transgenically expressing the KalTA4 transcription factor will be able to express GFP; thus, mediating targeted expression of GFP. 
Note: here we are using phenol red for visualization of DNA injections, which should be applicable for targeting other regions of the developing brain, as long as the region can be accessed from the ventricles. For targeting of other cell types that require fluorescent visualization, Bodipy or Quantam Dot dyes could potentially be used to visualize injections under fluorescent illumination.</li>
<li> Microinjection pipettes can be fabricated using either a horizontal or vertical pipette puller. Heat and pull-strength settings will vary depending on the type of puller, the type of heating element, and the type of glass capillary tubes. 
Using a Sutter P-30 vertical puller, use a heat setting of 980 and a pull-force of 960 to produce long, tapered, sharp pipettes from thin-walled borosilicate capillary glass with filaments [Sutter Instrument #BF100-78-10]. Microinjetion pipettes pulled with these settings are sealed and must be broken back before injection.</li>
<li>Use a microloader pipette tip to add 1-2 &mu;l of DNA plasmid solution to the injection pipette.</li>
<li>Mount and secure the loaded pipette into the pressure injection system pipette holder.</li>
<li>Break back the loaded pipette with fine forceps until pressure pulses produce puffed release of the red DNA solution. Alternatively, tips can be broken back by quickly penetrating a submerged, folded laboratory Kimwipe. 
Note: because the injection pipettes are initially sealed and must be manually broken back, the tip size and injection volume are variable. Ideal injection pipettes should require 3-5 injection pulses to completely fill the developing forebrain ventricle of 24 hpf embryos (100 ms injection pulses at 40 psi).</li>
<li>Inject the plasmid DNA solution into the ventricles of the developing brain such that DNA is forced adjacent to, and intercalated into, the region of brain destined to form the olfactory bulb. The olfactory bulb is derived from cells in the most anterior wall of the forebrain ventricle. Thus, by inserting the injection pipette into the ventricles such that the pipette is pointing toward the anterior end of the developing brain, pressure injection forces the DNA into and adjacent to the prospective olfactory bulb. Inject DNA until red dye is clearly visible at the anterior end of the forebrain ventricle. Because the DNA begins to diffuse immediately, diluting the DNA, it is important to proceed to the electroporation step as soon as possible after injection (e.g., within 10 sec).</li>
</ol>
4. Electroporation 
<ol>
	<li>Electroporation is carried out using self-constructed electrodes made from Grass platinum subdermal electrodes (Grass Technology #E2). The electrodes are attached to a hand-held probe, and the final distance between the platinum electrodes should be 1 mm (for details on construction see 9). Square-wave electroporation pulses are produced by connecting the electrodes to a Grass SD-9 Stimulator (Grass Technology, Model SD-9).</li> 
	<li>Set the SD-9 stimulator to produce single, 5 ms electroporation pulses at 70 volts. Position the electrodes such that the positive electrode is positioned adjacent to the anterior end of the embryo, while the negative electrode is posterior to the embryo head. Manually initiate &tilde;7-8 electroporation pulses using the single mode switch of the SD-9 stimulator, separating each pulse by &tilde;1 sec. Appropriate voltages will vary depending on the age of the embryos and the voltage source used. Younger embryos require lower voltages to avoid embryo damage, whereas older embryos require higher voltages to initiate electroporation 7.</li> 
<li>Allow the embryos to recover for at least 10 min after electroporation before freeing them from the agarose.</li>
<li>Using fine forceps, free the embryos from the agarose by gently tracing the outline of the embryos while avoiding actual contact with the embryos themselves.</li>
<li>Once freed, return the embryos to egg water (with PTU if needed). Keep them at 28&deg;C until they are to be imaged for GFP expression.</li>
</ol>
5. Mounting embryos for imaging
<ol>
<li>Transfer embryos from egg water plus PTU to egg water plus 0.02% tricaine. Allow several minutes for the anesthetic to take effect before proceeding to the mounting step.</li>
<li>Prepare a 0.5% solution of low-melting-point agarose in egg water plus tricaine by microwaving the solution, and then placing the solution into a 34-37&deg;C incubator. Once the temperature of the agarose solution has equilibrated, place a large drop (&tilde;300 &mu;l) of the agarose solution to the center of a 35 mm Petri dish. Add one anesthetized embryo to this drop of agarose. </li> 
<li> Using fine forceps (or the fine ends of cut microloader pipet tips), position the embryos such that they are upright, making it possible to visualize a dorsal view of the developing brain with an upright microscope (inverted scopes will require a different embryo geometry, and a different dish allowing visualization from below; see 12). Solidify the agarose by placing the dish onto a large Petri dish with 2-3 mm of ice. Re-orientation with fine forceps will likely be required during the solidification to ensure that the embryo does not fall over onto its side.</li>
<li>Once the agarose has solidified, flood the dish with egg water plus tricaine ensuring that the solution completely covers the solidified agarose drop.</li>
</ol>
6. Representative Results: 
Although GFP expression can be observed as early as 6 hours after electroporation, here we are showing images from an embryo 4 days after electroporation such that the neurite structure of the targeted cells has sufficiently developed. The Et(zic4:Gal4TA4,UAS:mCherry)hzm5 transgenic enhancer trap line displays constitutive expression of mCherry (and KalTA4) in the hindbrain, cerebellum, forebrain, and olfactory bulb in embryos at 5 days post-fertilization (Fig. 1A and 1C). Embryos that have had a UAS-GFP expression plasmid incorporated by targeted electroporation (as described above), show GFP expression that is restricted to cells of the developing olfactory bulb (Fig. 1B, 1D, and 1E). Only cells transgenically-expressing the KalTA4 transcription factor are able to activate expression from this UAS-GFP plasmid. Incorporation of this plasmid into non-transgenic embryos does not lead to any detectable GFP expression. Thus, it is the combined action of targeted electroporation of UAS-GFP and localized transgenic expression of the KalTA4 driver that leads to exclusive expression of GFP in cells of the developing olfactory bulb. The GFP-expressing cells show typical axonal projections of olfactory bulb mitral cells (Fig. 1B and 1D) 10. In order to visualize axon projections, images in figure 1B and 1D had to be exposed at high levels such that the region of cell bodies were well over-exposed. A lower level of exposure (Fig. 1E) shows that cell bodies expressing GFP are indeed localized to the olfactory bulb (compare Fig. 1C and 1E; gray line marks the border of the olfactory bulb).
<img src="/files/ftp_upload/2964/2964fig1.jpg" alt="Figure 1"/> 
Figure 1. Targeted GFP expression in the olfactory bulb of a 5 dpf zebrafish embryo. The Et(zic4:Gal4TA4,UAS:mCherry)hzm5 transgenic enhancer trap line, displays constitutive expression of mCherry in the hindbrain, cerebellum, forebrain, and olfactory bulb (A and C, mCherry fluorescence shown in red). This expression of mCherry is mediated by transgenic Gal4-expression. Embryos that have been electroporated with a UAS-GFP expression vector at 28 hpf, display targeted GFP expression in the olfactory bulb at 5 days post-fertilization (B, D, and E, GFP fluorescence shown in green. Same embryo as in A and C). A lower exposure image at bottom (E) shows that GFP expression is limited to cell bodies of the olfactory bulb. The gray line indicates the border of the olfactory bulb. Bright field (gray scale), green fluorescence (green), and red fluorescence (red) images were all take with an Olympus BX60 fluorescence microscope.

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No conflicts of interest declared.

Here we have described an in vivo electroporation method in zebrafish that utilizes an enhancer trap Gal4 transgenic line to target expression of the electroporated transgene to a specific population of cells in the developing olfactory bulb. This approach combines the excellent temporal resolution of in vivo electroporation 7 with the cell-type specific expression mediated by enhancer trap transgenic lines 8. Although here we have described the targeting of the olfactory bulb, in vivo electroporation can be used to target other regions of the developing nervous system 2-7, and enhancer trap lines are available with targeted expression of Gal4 in many different specific cell types or tissues 8, 11. Of course, this electroporation technique should also be suitable for other combinatioral genetic approaches such as the LexA or Tet systems 13, 14, 15. A major advantage of electroporation is that there is not need to make additional transgenic lines given that a suitable Gal4-line is available. This saves six months. 
In vivo electroporation allows for the incorporation of oligonucleotides into neurons and their precursors at whatever specific stage of nervous systems development is of interest 7. This temporal resolution is particularly advantageous for loss-of-function analysis because it can circumvent problems with targeting genes that have essential functions at earlier developmental stages. The method we describe here can also be used to incorporate loss-of-function reagents including RNAi oligonucleotides or constructs and morpholino anti-sense oligonucleotides 4, 7. Plasmid-driven reagents such as dominant-negative protein expression plasmids or short-hairpin RNAi plasmids can also be placed under control of a Gal4 UAS, allowing for the precise cell type-specific expression we have shown here for expression of GFP.

Reagents:
<ul>
	<li>Agarose, low-melting-point (Sigma, A9414)</li>
	<li>Plasmid midi- or maxi-prep kit (e.g. Qiagen, #12643]</li>
	<li>Phenol red (Sigma, P3532)</li>
</ul>
Equipment:
<ul>
	<li>Capillary glass with filament; ID = 0.78 mm; OD = 1.0 mm (Sutter Instruments, #BF100-78-10)</li>
	<li>Grass SD-9 Stimulator (Grass Technology, Model SD-9) </li>
	<li>Grass platinum subdermal electrodes, straight needle (Grass Technology, #E2)</li>
	<li>Micro-loader pipette tips (Eppendorf, #X22703R)</li>
	<li>Pipette puller (e.g. Sutter, Model P-30)</li>
	<li>Pipette puller (e.g. Sutter, Model P-30)</li>
</ul>

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.

Watch this Scientific Journal Video about Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines at JoVE.com

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.

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 ...

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

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

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

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

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

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

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

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. ...

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

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 ...

Olfactory Neurons Obtained through Nasal Biopsy Combined with Laser-Capture Microdissection: A Potential Approach to Study Treatment Response in Mental Disorders

Bipolar disorder (BD) is a severe neuropsychiatric disorder with poorly understood pathophysiology and typically treated with the mood stabilizer, lithium carbonate. Animal studies as well as human genetic studies indicate that lithium affects molecular targets that are involved in neuronal growth, survival and maturation, and notably molecules involved in Wnt signaling. Given the ethical challenge to obtaining brain biopsies for investigating dynamic molecular changes associated with lithium-response in the central nervous system (CNS), one may consider the use of neurons obtained from olfactory tissues to achieve this goal.The olfactory epithelium contains olfactory receptor neurons at different stages of development and glial-like supporting cells. This provides a unique opportunity to study dynamic changes in the CNS of patients with neuropsychiatric diseases, using olfactory tissue safely obtained from nasal biopsies. To overcome the drawback posed by substantial contamination of biopsied olfactory tissue with non-neuronal cells, a novel approach to obtain enriched neuronal cell populations was developed by combining nasal biopsies with laser-capture microdissection. In this study, a system for investigating treatment-associated dynamic molecular changes in neuronal tissue was developed and validated, using a small pilot sample of BD patients recruited for the study of the molecular mechanisms of lithium treatment response.

In this study, a novel platform to investigate intraneuronal molecular signatures of treatment response in bipolar disorder (BD) was developed and validated. Olfactory epithelium from BD patients was obtained through nasal biopsies. Then laser-capture microdissection was combined with Real Time RT PCR to investigate the molecular signature of lithium response in BD.

Bipolar disorder (BD) is a severe neuropsychiatric disorder with poorly understood pathophysiology and typically treated with the mood stabilizer, lithium ...

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. ...

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 ...