This study is supported by NIH DC052256 and DC006015, and NSF 0324769 to QG and T32-DC008072 to BS.

1. Preparation
This procedure is biosafety level 2, therefore all the following preparations are made in a biohazard hood where the microinjection procedure is performed.
<ol>
	<li>Prepare and preheat a 37&deg;C waterbed: Fill a plastic storage bag with water &#x2013; seal and set on an emptied heating block incubator. This will allow mice to recover following injection.</li>
	<li>Fill a biohazard waste container with 10% bleach to a suitable volume for immersion of all waste from this procedure.</li>
	<li>Prepare a small open dish of 70% ethanol and a second dish with sterilized ddH2O and place in the biological safety cabinet. </li>
	<li>Sterilize a 5&mu;L capacity Hamilton syringe with a 33-gauge needle by spraying down the body and needle with 70% ethanol with the plunger pulled out all the way. Repeatedly aspirate ethanol from the dish and eject it all the way out at least 5 times.</li>
	<li>Rinse the syringe cartridge by aspirating and ejecting sterile water at least 5 times. Place syringe safely aside in the culture hood and allow it to dry.</li>
	<li>Remove viral stocks from -80&deg;C freezer storage and place on ice to allow thawing. Microinjection requires 2&mu;L of virus stock per mouse at a titre of 109 infectious units/mL. Lentiviruses can be obtained from commercial sources or produced in the lab according to the established protocol (Lois et al., 2002).</li>
	<li>Prepare an injection stage that provides an area raised above the hood surface - can use a 10cm culture dish covered with a sheet of Kimwipe.</li>
	<li>Arrange alcohol wipes near the injection station.</li>
</ol>
2. Microinjection of lentiviral particles into the olfactory epithelium
All animal procedures performed in this method are done according to IACUC approved guidelines. The Mus musculus strain C57BL/6J is the animal model used in all experiments demonstrated here.
<ol>
	<li>Anaesthetize mouse pups by hypothermia by taking a mouse pup (P0-P3) out of the cage and putting the pup on a cut-open glove on ice for 5 minutes.</li>
	<li>While waiting for the pup to be fully anesthetized, aspirate 2&mu;L of viral stock into the sterilized Hamilton syringe.</li>
	<li>Place the mouse pup on top of the injection stage. Gently pull skin away from the top of the head at the ear toward the top of the neck. This should provide a taught skin surface over the nasal cavity and will ease injection.</li>
	<li>Insert needle through the top of the nasal cavity about 3mm below the skin surface to reach the olfactory epithelium. Microinject 0.5&mu;L of virus solution per shot, choosing two sites per side, both left and right (four injections total per mouse = 2&mu;L of virus needed). Injection sites should be staggered apart from one another. For targeting of proper injection sites, search the dorsal skull surface for the outline of the olfactory bulb. This line is barely visible at birth through the semi-translucent skin and skull. To make two injections per side, aim one injection just rostral to one bulb outline at a medial axis. Make the second insertion lateral to the bulb outline, yet make sure to angle the injection medially so that you may reach the olfactory epithelium that resides ventrally underneath the bulb. Avoid penetrating the olfactory bulb!</li>
	<li>Blot dry any residual virus solution or blood that may escape from the injection site or nostril openings using a Kimwipe. Immediately dispose of Kimwipes in the diluted liquid bleach biohazard container.</li>
	<li>Place the injected mouse onto the waterbed incubator.</li>
	<li>Repeat steps 1-4 for all mice to be injected, remembering you will need about 2&mu;L of virus stock per mouse.</li>
	<li>Let mice recover on the waterbed incubator for 30 minutes before returning them to their original nesting cage.</li>
	<li>Dispose of microinjection stage, sanitizing wipes and any extra cleaning paper products in the liquid bleach biohazard disposal.</li>
	<li>Clean the Hamilton syringe by repeated aspiration and ejection of ethanol and then water as done in the preparation steps. Allow the syringe to dry.</li>
	<li>For maximum transduction efficiency repeat this procedure again in 4 hours. Infected OSNs may be visualized throughout the length of the cell in as few as three days.</li>
</ol>
3. Immunohistochemistry of floating olfactory bulb sections
<ol>
	<li>Sacrifice the injected mouse at the desired time point and perfusion fix the tissue with 4% paraformaldehyde. Dissect out the OB. Incubate in paraformaldehyde overnight and then 30% sucrose in PBS overnight. Embed in O.C.T. embedding medium on dry ice. </li>
	<li>Coronal OB sections are obtained at a 60&mu;m in thickness using a cryostat. Sweep bulb sections into a 10cm culture dish with 5mL 1x PBS (pH 7.4). Sections should be kept floating in PBS to allow O.C.T. to dissolve. Transfer floating sections and PBS into a 50mL conical vial.</li>
	<li>Allow sections to settle to the bottom of the vial. Vacuum off PBS without sucking up any sections. Rinse tissue sections with 20mL PBS. Repeat settle and vacuuming steps. Resuspend tissue with 10mL PBS and transfer floating sections to a 15mL conical vial for immunostaining. Once tissue sections have settled, vacuum off PBS supernatant without disturbing tissue. Resuspend sections with 0.3% Triton X-100 in 1x PBS, pH 7.4 (PBS-Tx) allowing detergent to perforate cell membranes. Incubate for 10 minutes at room temperature. Vacuum off supernatant and repeat incubation with PBS-Tx twice.</li>
	<li>Incubate tissue sections in 10mL of blocking solution (5% horse serum in PBS-Tx) for two hours at room temperature.</li>
	<li>Rinse blocking solution at least twice with PBS-Tx for 15 minutes each at room temperature.</li>
	<li>Prepare 2mL of a primary antibody solution in PBS-Tx. See Reagents and Equipment section for antibodies used in this experiment.</li>
	<li>Replace second PBS-Tx blocking wash with primary antibody solution. Incubate on a slowly rocking platform (~30rpm) at 4&deg;C for 24-48 hours to ensure thorough penetration into the 60&mu;m floating sections.</li>
	<li>After incubation, the primary antibody solution can be retrieved carefully using a pipette and stored at 4&deg;C for reuse up to two more times within two weeks without any significant loss of staining intensity.</li>
	<li>Wash sections three times in PBS-Tx for 20 minutes each at room temperature with gentle rotation (~30rpm).</li>
	<li>Prepare 2mL of secondary antibody dilution in PBS-Tx. Vacuum off the last wash and add secondary antibody solution to tissue sections. Wrap the outside of the staining vial in aluminum foil to block light, which can cause fluorophore photobleaching. Place wrapped vial on a slowly rocking platform and incubate at 4&deg;C overnight.</li>
	<li>Discard secondary antibody solution by vacuuming away. Wash sections once with PBS-Tx for 20 minutes each at room temperature. Rinse two additional times with PBS (pH 7.4) for 20 minutes each at room temperature. Mount the stained tissue sections onto glass slides using Fluoromount G.</li>
	<li>Screen through stained sections for positively infected OSN axons and axon termini with a fluorescent microscope.</li>
	<li>To construct a three dimensional view of a labeled OSN axon and terminal arbor, make a series of optical sections through the 60&mu;m depth of the target OB tissue slice (along the z axis) using confocal microscopy.</li>
	<li>This stack of Z-section images can subsequently be projected together to reconstruct a 360&deg; rotating perspective of a single labeled OSN axon, including terminal arbor detail in its entirety. Any one single rotation angle can be captured and exported as an image file (i.e. JPG, TIFF) for 2D format presentation.</li>
</ol>
4. Representative Results
With the methods described here, we can transduce olfactory sensory neurons by lentivirus in vivo. We have thus far visualized infected OSNs with GFP and mCherry fluorescent reporters, as well with a myc-tagged fusion protein via immunocytochemistry. This technique provides an abundance of infected OSNs. Still, transduction occurs sporadically enough to provide individually labeled OSNs, thus allowing for analysis of single cell morphology. Axons and axon terminals are imaged within OB tissue sections amidst their native environment by confocal microscopy (Figure 1). The entire OSN axon terminal arbor can typically be captivated within a 40-50&mu;m optical section. High magnification of infected OSN axon terminals allows detailed analysis of axonal arbor morphology at the glomerular layer.
<img src="/files/ftp_upload/2951/2951fig1.jpg" /> 
Figure 1. Trajectory of single olfactory sensory neuron axon and its terminal arbor. A) A Coronal section of the olfactory bulb of a P7 mouse showing the olfactory nerve layer (ONL) immunostained for OMP (red) and the glomerular layer (GL) immunostained with both OMP and Vglut2 (grey). A single olfactory sensory neuron axon expressing GFP (green) travels within the ONL and penetrates into the glomerular structure. B) The axon terminal arbor morphology within the glomerulus clearly highlighted by GFP expression. Vglut2 co-staining in grey illuminates the total glomular area within which the GFP-positive axon elaborates a terminal abor. C) Lentiviral infected OSNs expressing GFP (green) in the OE. The cytostructure of the olfactory epithelium is shown with DAPI staining (blue). GFP expression allows visualization of OSN cell bodies and their cellular processes in the OE. D) An OSN axon and its terminal morphology with Rap1GAP2 expression knock-down by shRNA. Lentiviruses carrying a dual expression cassette with Rap1GAP2 shRNA under the mouse U6 promoter and GFP under a CMV promoter were applied to the OE. Infected olfactory axon terminals exhibited a more simplified terminal arbor when compared to that of the GFP only control. Bar = 20&mu;m.

<ol>
	<li>Chen, H., Kohno, K., Gong, Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conditional+ablation+of+mature+olfactory+sensory+neurons+mediated+by+diphtheria+toxin+receptor.">Conditional ablation of mature olfactory sensory neurons mediated by diphtheria toxin receptor.</a> J Neurocytol. 34, 1-2 (2005).</li><li>Doi, K., Nibu, K., Ishida, H., Okado, H., Terashima, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adenovirus-mediated+gene+transfer+in+olfactory+epithelium+and+olfactory+bulb:+a+long-term+study.">Adenovirus-mediated gene transfer in olfactory epithelium and olfactory bulb: a long-term study.</a> Ann Otol Rhinol Laryngol. 114, 629-633 (2005).</li><li>Lois, C., Hong, E. J., Pease, S., Brown, E. J., Baltimore, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Germline+transmission+and+tissue-specific+expression+of+transgenes+delivered+by+lentiviral+vectors.">Germline transmission and tissue-specific expression of transgenes delivered by lentiviral vectors.</a> Science. 295, 868-872 (2002).</li><li>Zhao, H., Otaki, J. M., Firestein, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adenovirus-mediated+gene+transfer+in+olfactory+neurons+in+vivo.">Adenovirus-mediated gene transfer in olfactory neurons in vivo.</a> J Neurobiol. 30, 521-530 (1996).</li></ol>

This protocol is bio-safety level 2 compliant. All appropriate workspace, personal protection, and viral waste containment measures have been established accordingly. All animal procedures were performed according to IACUC guidelines.

Microinjection of lentivirus results in permanent transfer of gene constructs into OSN genomic DNA. This approach allows us to perform short-term or long-term manipulations of candidate genes via overexpression or shRNA mediated knock-down. In addition, we can fluorescently label single OSNs in an existing transgenic mouse line to observe co-localization of odorant receptor populations. Microinjection is required for lentiviral transduction of OSNs. We have attempted flushing lentivirus suspension into the nasal cavity t...

<table><tbody><tr><td colspan="4">&lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Microinjection:&lt;/em&gt; Newborn mice were microinjected using a 5&amp;#x03BC;L Hamilton syringe (Hamilton #7647-01) fitted with a 33 gauge needle (Hamilton #7762-06).&lt;/p&gt;&lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Immunocytochemistry Reagents:&lt;/em&gt; Primary antibodies: rabbit polyclonal antibody GFP (Molecular Probes# A-6455), chicken polyclonal antibody against OMP (custom) (Chen et al., 2005), guinea pig polyclonal antibody against VGlut2 (Millipore# AB2251). Secondary antibodies: Cy2-conjugated donkey anti-rabbit (Jackson Immunolab# 711-225-152), Cy3-conjugated donkey-anti-chicken (Jackson Immunolab# 703-165-155) and Cy5-conjugated goat anti-guinea pig (Jackson Immunolab# 106-175-008). Sections were mounted on glass slides with Fluoromount G (Southern Biotech# 0010-01) with 50ng/mL DAPI chromatin stain solution.&lt;/p&gt;&lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Confocal Imaging:&lt;/em&gt; Z-stack images were taken using an Olympus Flouview FV1000 confocal microscope and images collected and processed into 3D projections using Olympus FV10-ASW 2.01 confocal acquisition and analysis software.&lt;/p&gt;</td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Lentivirus-mediated Genetic Manipulation and Visualization of Olfactory Sensory Neurons in vivo at JoVE.com

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.

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 ...

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15.6K Views.  University of California, Davis. We present a lentiviral technique for genetic manipulation and visualization of single olfactory sensory neuron axon and its terminal arborization in vivo.

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

Stable and Efficient Genetic Modification of Cells in the Adult Mouse V-SVZ for the Analysis of Neural Stem Cell Autonomous and Non-autonomous Effects

Relatively quiescent somatic stem cells support life-long cell renewal in most adult tissues. Neural stem cells in the adult mammalian brain are restricted to two specific neurogenic niches: the subgranular zone of the dentate gyrus in the hippocampus and the ventricular-subventricular zone (V-SVZ; also called subependymal zone or SEZ) in the walls of the lateral ventricles. The development of in vivo gene transfer strategies for adult stem cell populations (i.e. those of the mammalian brain) resulting in long-term expression of desired transgenes in the stem cells and their derived progeny is a crucial tool in current biomedical and biotechnological research. Here, a direct in vivo method is presented for the stable genetic modification of adult mouse V-SVZ cells that takes advantage of the cell cycle-independent infection by LVs and the highly specialized cytoarchitecture of the V-SVZ niche. Specifically, the current protocol involves the injection of empty LVs (control) or LVs encoding specific transgene expression cassettes into either the V-SVZ itself, for the in vivo targeting of all types of cells in the niche, or into the lateral ventricle lumen, for the targeting of ependymal cells only. Expression cassettes are then integrated into the genome of the transduced cells and fluorescent proteins, also encoded by the LVs, allow the detection of the transduced cells for the analysis of cell autonomous and non-autonomous, niche-dependent effects in the labeled cells and their progeny.

Here we describe a procedure based on the use of lentiviral particles for the long-term genetic modification of neural stem cells and/or their adjacent ependymal cells in the adult ventricular-subventricular neurogenic niche which allows the separate analysis of cell autonomous and non-autonomous, niche-dependent effects on neural stem cells.

Relatively quiescent somatic stem cells support life-long cell renewal in most adult tissues. Neural stem cells in the adult mammalian brain are ...

Developmental Biology

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 ...

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 ...

Viral Tracing of Genetically Defined Neural Circuitry

Classical methods for studying neuronal circuits are fairly low throughput. Transsynaptic viruses, particularly the pseudorabies (PRV) and rabies virus (RABV), and more recently vesicular stomatitis virus (VSV), for studying circuitry, is becoming increasingly popular. These higher throughput methods use viruses that transmit between neurons in either the anterograde or retrograde direction.
Recently, a modified RABV for monosynaptic retrograde tracing was developed. (Figure 1A). In this method, the glycoprotein (G) gene is deleted from the viral genome, and resupplied only in targeted neurons. Infection specificity is achieved by substituting a chimeric G, composed of the extracellular domain of the ASLV-A glycoprotein and the cytoplasmic domain of the RABV-G (A/RG), for the normal RABV-G1. This chimeric G specifically infects cells expressing the TVA receptor1. The gene encoding TVA can been delivered by various methods2-8. Following RABV-G infection of a TVA-expressing neuron, the RABV can transmit to other, synaptically connected neurons in a retrograde direction by nature of its own G which was co-delivered with the TVA receptor. This technique labels a relatively large number of inputs (5-10%)2 onto a defined cell type, providing a sampling of all of the inputs onto a defined starter cell type.
We recently modified this technique to use VSV as a transsynaptic tracer9. VSV has several advantages, including the rapidity of gene expression. Here we detail a new viral tracing system using VSV useful for probing microcircuitry with increased resolution. While the original published strategies by Wickersham et al.4 and Beier et al.9 permit labeling of any neurons that project onto initially-infected TVA-expressing-cells, here VSV was engineered to transmit only to TVA-expressing cells (Figure 1B). The virus is first pseudotyped with RABV-G to permit infection of neurons downstream of TVA-expressing neurons. After infecting this first population of cells, the virus released can only infect TVA-expressing cells. Because the transsynaptic viral spread is limited to TVA-expressing cells, presence of absence of connectivity from defined cell types can be explored with high resolution. An experimental flow chart of these experiments is shown in Figure 2. Here we show a model circuit, that of direction-selectivity in the mouse retina. We examine the connectivity of starburst amacrine cells (SACs) to retinal ganglion cells (RGCs).

A method of tracing synaptically connected neurons is described. We use TVA specificity of an upstream cell to probe whether a cell population of interest receives synaptic input from genetically defined cell types.

Classical methods for studying neuronal circuits are fairly low throughput. Transsynaptic viruses, particularly the pseudorabies (PRV) and rabies virus ...

Stereotaxic Injection of a Viral Vector for Conditional Gene Manipulation in the Mouse Spinal Cord

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central nervous system. We demonstrate a stereotaxic injection technique that allows targeted gene expression or silencing in the dorsal horn of the mouse spinal cord. The surgical procedure is brief. It requires laminectomy of a single vertebra, providing for quick recovery of the animal and unimpaired motility of the spine. Controlled injection of a small vector suspension volume at low speed and use of a microsyringe with beveled glass cannula minimize the tissue lesion. The local immune response to the vector depends on the intrinsic properties of the virus employed; in our experience, it is minor and short-lived when a recombinant adeno-associated virus is used. A reporter gene such as enhanced green fluorescent protein facilitates monitoring spatial distribution of the vector, and the efficacy and cellular specificity of the transfection.

Viral vectors allow for targeted gene manipulation. We demonstrate a method for conditional gene expression or ablation in the mouse spinal cord, using stereotaxic injection of a viral vector into the dorsal horn, a prominent site of synaptic contact between primary somatosensory afferents and neurons of the central nervous system.

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central ...

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 ...

Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. Few studies have examined gene transfer to the central auditory system. The dorsal cochlear nucleus (DCN) of the brainstem, which contains second order neurons of the auditory pathway, is a potential site for gene transfer. In this protocol, a technique for direct and maximal exposure of the murine DCN via a posterior fossa approach is demonstrated. This approach allows for either acute or survival surgery. Following direct visualization of the DCN, a host of experiments are possible, including injection of opsins into the cochlear nucleus and subsequent stimulation by an optical fiber coupled to a blue light laser.&#160;Other neurophysiology experiments, such as electrical stimulation and neural injector tracings are also feasible.&#160;The level of visualization and the duration of stimulation achievable make this approach applicable to a wide range of experiments.

The goal of this protocol is to outline a surgical approach to provide direct access to the dorsal cochlear nucleus in a murine model.

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. ...

Laser-guided Neuronal Tracing In Brain Explants

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake through electroporation in brain explants with targeted laser illumination and matching filter goggles during the procedure. The described technique of in vitro electroporation by itself yields relatively good visual control for targetting certain areas of the brain. By combining it with laser excitation of fluorescent genetic markers and their read-out through band-passing filter goggles, which can pick up the emissions of the genetically labeled cells/nuclei and the fluorescent tracing dye, a researcher can substantially increase the accuracy of injections by finding the area of interest and controlling for the dye-spread/uptake in the injection area much more efficiently. In addition, the laser illumination technique allows to study the functionality of a given neurocircuit by providing information about the type of neurons projecting to a certain area in cases where the GFP expression is linked to the type of transmitter expressed by a subpopulation of neurons.

We describe a technique to label neurons and their processes via anterograde or retrograde tracer injections into brain nuclei using an in vitro preparation. We modified an existing method of in vitro tracer electroporation by taking advantage of fluorescently labeled mouse mutants and basic optical equipment in order to increase labeling accuracy.

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake ...

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as accessibility to numerous imaging approaches, electrophysiological techniques and behavioral assays. The Xenopus tadpole olfactory system is particularly well suited to investigate the function of synapses established during normal development or reformed after injury. Here, we describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae. We outline a combination of in vivo measurements of presynaptic calcium responses in glomeruli of the olfactory bulb with olfactory-guided behavior assays. Methods can be combined with the transection of olfactory nerves to study the rewiring of synaptic connectivity. Experiments are presented using both wild-type and genetically modified animals expressing GFP reporters in central nervous system cells. Application of the approaches described to genetically modified tadpoles can be useful for unraveling the molecular bases that define vertebrate behavior.

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system in vivo. We describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae in normal rearing conditions or after injury.

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as ...

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. Traditional in vitro electrophysiological techniques use electrical stimulation to evoke synaptic transmission. A major drawback of this method is its nonspecific nature; all axons in the region of the stimulating electrode will be activated, making it difficult to attribute an effect to a particular afferent connection. This issue can be overcome by replacing electrical stimulation with optogenetic-based stimulation. We describe a method for combining optogenetics with in vitro patch-clamp recordings. This is a powerful tool for the study of both basal synaptic transmission and synaptic plasticity of precise anatomically defined synaptic connections and is applicable to almost any pathway in the brain. Here, we describe the preparation and handling of a viral vector encoding channelrhodopsin protein for surgical injection into a pre-synaptic region of interest (medial prefrontal cortex) in the rodent brain and making of acute slices of downstream target regions (lateral entorhinal cortex). A detailed procedure for combining patch-clamp recordings with synaptic activation by light stimulation to study short- and long-term synaptic plasticity is also presented. We discuss examples of experiments that achieve pathway- and cell-specificity by combining optogenetics and Cre-dependent cell labeling. Finally, histological confirmation of the pre-synaptic region of interest is described along with biocytin labeling of the post-synaptic cell, to allow further identification of the precise location and cell type.

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Here we present a protocol describing viral transduction of discrete brain regions with optogenetic constructs to permit synapse-specific electrophysiological characterization in acute rodent brain slices.

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. ...

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

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