This work was supported by a grant from the DFG to G.M.T.

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J., Sink, H., Kolodkin, A. L., Goodman, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Semaphorin+II+can+function+as+a+selective+inhibitor+of+specific+synaptic+arborizations.">Semaphorin II can function as a selective inhibitor of specific synaptic arborizations.</a> Cell. 81, 631-639 (1995).</li><li>Murray, M. J., Whitington, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+roundabout+on+growth+cone+dynamics,+filopodial+length,+and+growth+cone+morphology+at+the+midline+and+throughout+the+neuropile.">Effects of roundabout on growth cone dynamics, filopodial length, and growth cone morphology at the midline and throughout the neuropile.</a> Journal of Neuroscience. 19, 7901-7912 (1999).</li><li>Murray, M. J., Merritt, D. J., Brand, A. H., Whitington, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+dynamics+of+axon+pathfinding+in+the+Drosophilia+CNS:+a+time-lapse+study+of+an+identified+motorneuron.">In vivo dynamics of axon pathfinding in the Drosophilia CNS: a time-lapse study of an identified motorneuron.</a> J. Neurobiol. 37, 607-621 (1998).</li><li>Patel, N. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+neuronal+subsets+and+other+cell+types+in+whole-mount+Drosophila+embryos+and+larvae+using+antibody+probes.">Imaging neuronal subsets and other cell types in whole-mount Drosophila embryos and larvae using antibody probes.</a> Methods Cell Biol. 44, 445-487 (1994).</li><li>Rickert, C., Kunz, T., Harris, K. -. L., Whitington, P. M., Technau, G. 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M., Seifert, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axon+growth+from+limb+motorneurons+in+the+locust+embryo:+the+effect+of+target+limb+removal+on+the+pattern+of+axon+branching+in+the+periphery.">Axon growth from limb motorneurons in the locust embryo: the effect of target limb removal on the pattern of axon branching in the periphery.</a> Developmental Biology. 106, 438-449 (1984).</li><li>Whitington, P. M., Leach, D., Sandeman, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolutionary+change+in+neural+development+within+the+arthropods:+axonogenesis+in+the+embryos+of+two+crustaceans.">Evolutionary change in neural development within the arthropods: axonogenesis in the embryos of two crustaceans.</a> Development. 118, 449-461 (1993).</li><li>Williams, D. 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The authors declare that they have no competing financial interests.

One major advantage of Drosophila as a model system is that it allows analysis of development and function on the level of single cells. This is especially helpful regarding the nervous system, where the diversity of cell types is exceptionally high and the function and morphology of neighboring cells can be totally different.
The method we present here allows the labeling of individual neurons with a dye that can either be transformed into a permanent stain or examined directly by fluorescence microscopy. It reveals the morphology of neural processes in great detail (Figure 4). One of the major advantages of the method is that it enables the morphology of virtually any type of neuron in the CNS to be examined (for the embryonic ventral nerve cord, see Rickert et al., 2011; for the embryonic brain, see Kunz et al., 2012). In addition, it does not require complex combinations of genetic elements to be present in the embryo, as several of the genetic labeling techniques do. Dye injections may be performed into cells of animals that carry reporter constructs, thereby allowing individual cell morphologies to be visualized in the context of particular gene expression patterns (Figures 4D and 5) in either wild type or mutant backgrounds. We have also used the technique to monitor the dynamics of axon outgrowth in living embryonic neurons 11, 12.
The method can be readily adapted to label individual neurons in embryos of other organisms, provided the embryo is transparent enough to be examined under DIC illumination. We have used it to visualize neural morphology in a wide variety of arthropod embryos, including grasshoppers 18, centipedes 17, crustaceans 19 and silverfish 16.
The major disadvantage of the technique lies in its technical difficulty. We trust that the current detailed account of the method will assist interested researchers in mastering it.

Knowledge of neuronal morphology at the level of individual cells is a key prerequisite for understanding neuronal connectivity and CNS function. Thus, from the earliest days of neuroscience, researchers have sought to develop single cell labeling techniques (see 7 for a historical treatment of this issue). Classical methods such as Golgi staining provide excellent resolution of neuronal morphology but are not suitable if one seeks to label a particular type of neuron in a directed way, as staining occurs in a random fashion. The development of methods for staining single cells by intracellular or juxtacellular injection of dyes from a microelectrode addressed the requirement of specific labeling.
The application of single neuron dye injection to Drosophila presented a major challenge, because of the small size of both the organism and its neurons. Nonetheless, single neuron staining of embryonic Drosophila neurons was achieved in the mid-1980s in the laboratory of Corey Goodman 15. While the method has been eminently useful and has provided some key insights into mechanisms for neuronal development in Drosophila over the last 20-30 years (e.g. 2, 10), many workers have shied away from it, largely because of its technical demands.
The availability in more recent years of genetic techniques for neuronal labeling in Drosophila has also contributed to the unpopularity of single neuron dye injection. GAL4-directed expression of membrane-targeted GFP constructs can provide excellent resolution of neural morphology 1, 20. However, this method has certain limitations: the often-unavoidable expression of GFP in multiple cells can obscure the structure of individual neurons and a GAL4 driver line may not be available to drive expression in a particular neuron of interest. The MARCM (Mosaic Analysis with a Repressible Cell Marker) method 8 can provide labeling of essentially any neuron at the individual cell level, but cannot be successfully used in the embryo and early larva because of the slow turnover of the GAL80 protein.
Given these limitations of genetic labeling, we believe that single neuron dye injection in the Drosophila embryo remains a valuable technique and deserves broader application. To promote this goal, we provide here a detailed description of the method. An illustration of its power is provided by our recent account of the morphology of the complete set of interneurons in abdominal neuromeres of the late Drosophila embryo 14. This study, which revealed both the morphological variability of individual neuronal cell types and the principles of neuromere organization in the CNS of the embryo, would not have been possible with any other currently available labeling method.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>REAGENTS</td>
 </tr>
 <tr>
 <td>DAB</td>
 <td>Sigma-Aldrich</td>
 <td><a href="http://www.sigmaaldrich.com/catalog/product/sigma/d5905" target="_blank">D-5905</a></td>
 <td>2-3 mg/ml in 100 mM TRIS-Hcl, pH 7.4</td>
 </tr>
 <tr>
 <td>DiI</td>
 <td>Invitrogen/Molecular Probes</td>
 <td><a href="http://products.invitrogen.com/ivgn/product/D282" target="_blank">D-282</a></td>
 <td>1 mg/ml in ethanol</td>
 </tr>
 <tr>
 <td><a href="http://www.merckmillipore.com/germany/chemicals/formaldehydloesung-min-37-prozent/MDA_CHEM-104002/p_Wt6b.s1L6pAAAAEWpOEfVhTl" target="_blank">Formaldehyde</a></td>
 <td>Merck Millipore</td>
 <td>&nbsp;</td>
 <td>7,4 % in PBS</td>
 </tr>
 <tr>
 <td>Glycerol</td>
 <td><a href="http://www.carlroth.com/catalogue/" target="_blank">Roth</a></td>
 <td>3783</td>
 <td>70% in PBS</td>
 </tr>
 <tr>
 <td>Heptane Glue</td>
 <td>Beiersdorf AG</td>
 <td>Cello 31-39-30 *</td>
 <td>dilute ca. 1 to 1 with n-Heptane</td>
 </tr>
 <tr>
 <td>PBS</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>1x</td>
 </tr>
 <tr>
 <td>TRIS</td>
 <td><a href="http://www.carlroth.com/catalogue/" target="_blank">Roth</a></td>
 <td>4855</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Vectashield</td>
 <td>Vector Laboratories</td>
 <td><a href="http://www.vectorlabs.com/catalog.aspx?prodID=427" target="_blank">H1000</a></td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>EQUIPMENT</td>
 </tr>
 <tr>
 <td>Confocal Microscope</td>
 <td>Leica</td>
 <td>TCS SP2</td>
 <td>to view and document labelings in fluorescence</td>
 </tr>
 <tr>
 <td>DC - amplifier</td>
 <td>Dragan Corporation</td>
 <td>Cornerstone ION-100</td>
 <td>to perform the labelings</td>
 </tr>
 <tr>
 <td>Coverslips</td>
 <td><a href="http://www.menzel.de/" target="_blank">Menzel</a></td>
 <td>BB018018A1</td>
 <td>18 x 18 mm</td>
 </tr>
 <tr>
 <td>Coverslips</td>
 <td><a href="http://www.menzel.de/" target="_blank">Menzel</a></td>
 <td>BB024060A1</td>
 <td>24 x 60 mm</td>
 </tr>
 <tr>
 <td>Dissecting Microscope</td>
 <td>Leica</td>
 <td>MZ8</td>
 <td>to prepare and disect embryos</td>
 </tr>
 <tr>
 <td>Flat Capillaries</td>
 <td><a href="http://www.hilgenberg-gmbh.de/" target="_blank">Hilgenberg</a></td>
 <td>&nbsp;</td>
 <td>outer diameter 1 mm; glas thickness 0.1 mm</td>
 </tr>
 <tr>
 <td>Injection Capillaries</td>
 <td>Science Products</td>
 <td>GB 100 TF 8P</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micromanipulator R</td>
 <td>Leica</td>
 <td>&nbsp;</td>
 <td>to perform the labelings</td>
 </tr>
 <tr>
 <td>Model P-97</td>
 <td>Sutter Instruments</td>
 <td>&nbsp;</td>
 <td>to pull capillaries</td>
 </tr>
 <tr>
 <td>Object Slides</td>
 <td><a href="http://www.marienfeld-superior.com/" target="_blank">Marienfeld Superior</a></td>
 <td>1000000</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Scientific Microscope</td>
 <td>Zeiss</td>
 <td>Axioskop 2 mot</td>
 <td>to view labelings after photoconversion</td>
 </tr>
 <tr>
 <td>Scientific Microscope</td>
 <td>Olympus</td>
 <td>BX50</td>
 <td>to perform the labelings</td>
 </tr>
 <tr>
 <td>Sony MC3255 Video Camera</td>
 <td>Sony/AVT Horn</td>
 <td>Sony MC3255</td>
 <td>to record labelings after photoconversion</td>
 </tr>
 </tbody>
</table>
Table 1.

labeling of single cells in the central nervous system of drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Figure 4 illustrates typical results of the technique, we describe here. Figure 4A shows an example of a DiI filled single interneuron that was cleanly photoconverted. It nicely demonstrates the amount of detail these preparations offer. When viewed under DIC optics the spatial context of the labeled cell within the non-labeled surrounding tissue becomes visible, e.g. the position of the cell body within the cortex and of the fiber projection within the neuropile.
Figure 4B is a case where the dye drop was a little too big resulting in several neighboring cells becoming labeled simultaneously. This often makes it difficult to relate individual projections to distinct cell bodies. It also shows that when directly viewed under the fluorescent microscope (no photoconversion) background resolution is much lower.
The specimen in Figure 4C illustrates a case where multiple cells being were individually labeled in neighboring segments. Upon photoconversion the preparation was subsequently stained with an antibody against Fas2 5following standard protocols 13 to provide landmarks (fascicles) in the neuropile. Although in this preparation most cell bodies are well separated, and allowing reliable mapping of their projections, it also demonstrates what may occur when the photoconversion period is prolonged (in order to also convert the weakly labeled cells): the more intensely labeled cells tend to get overstained and start to swell (compare with A).
Figure 4D shows that observation by confocal microscopy reveals the morphology of labeled cells in detail. DiI labeling was performed in a strain that carried a GFP reporter construct. This may provide important information about the spatial context (and identity) of the dye filled cell within specific populations of neuronal or glial cell types (as defined by gene expression).
As a paradigm to evaluate morphological variability of an identified neuron we have chosen one of the Apterous positive neurons 14. This neuron lies in a dorsal and medial position close to the neuropile (dAP, Figure 5A, Lundgren et al., 1995) separate from the other Apterous positive neurons and is therefore easily identifiable in apGal4-UASGFP animals. Figure 5B shows a dAP cell that was filled with DiI and photoconverted. The cell body lies at the level of the anterior commissure and shows an axon that first grows towards the midline and then turns anterior in a medial connective region. It has growth cone like swellings at the tip and the turning point. 19 labels of this cell, drawn as maximum projections from image stacks, are summarized in Figure 5C. Cell morphology becomes visible to great detail. Only few cells show growth cone like structures; six of the 19 cells also send a branch posterior that is always shorter than the anterior branch and may well be a transient feature. In Figure 5D all 19 dAP cells labeled are stacked with each cell having only an opacity of 12%. This results in areas being darker the more cells share them. While the cell body varies more than one cell diameter around the center position, the mediolateral position of the axon within the neuropile varies much less - when the neuropile width is divided into nine parts it resides always in one of the medial two positions.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/50150/50150fig1highres.jpg" src="/files/ftp_upload/50150/50150fig1.jpg" /> 
 Figure 1. Schematic overview of the steps of the neuron labeling method and its two variants.
<img alt="Figure 2" fo:content-width="4in" fo:src="/files/ftp_upload/50150/50150fig2highres.jpg" src="/files/ftp_upload/50150/50150fig2.jpg" /> 
 Figure 2. Diagram illustrating the various steps in preparing Drosophila embryos for neuron dye injection, according to variant B. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50150/50150fig2large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 3" fo:content-width="5.5in" fo:src="/files/ftp_upload/50150/50150fig3highres.jpg" src="/files/ftp_upload/50150/50150fig3.jpg" /> 
 Figure 3. Schematic representation of the setup for dye injection. The upper right insert is an enlarged view of the arrangement of the specimen, the injection micropipette and the bath electrode. A: Micromanipulator, B: Fixed stage microscope. The micromanipulator is set at a shallow angle - approximately 10&deg;. C: Micropipette holder with micropipette, D: Specimen on slide, E: Bath electrode fixed with plasticine, F: Connection to DC amplifier.
<img alt="Figure 4" fo:content-width="5in" fo:src="/files/ftp_upload/50150/50150fig4highres.jpg" src="/files/ftp_upload/50150/50150fig4.jpg" /> 
 Figure 4. Examples of DiI-labeled interneurons in the ventral nerve cord of stage 17 embryos. Dorsal views, anterior is to left. Midline: dotted lines.
<ol class="ualpha">
 <li>An example of a perfectly photoconverted single cell, showing a prominent contralateral axonal projection and ipsilateral dendritic fibers (arrowhead). DIC optics.</li>
 <li>shows a preparation in which more than one cell was labeled - this can obscure the clear assignment of a single cell body to its projection.</li>
 <li>8 cells have been labeled in 3 consecutive segments. In addition, in this preparation the photoconversion was followed by a standard protocol antibody staining against Fas2 as a landmark.</li>
 <li>shows a single DiI-filled neuron documented with confocal microscopy in a fly strain carrying a GFP reporter construct.</li>
</ol>
<img alt="Figure 5" fo:content-width="5in" fo:src="/files/ftp_upload/50150/50150fig5highres.jpg" src="/files/ftp_upload/50150/50150fig5.jpg" /> 
 Figure 5. Targeted DiI labeling (marked by GFP expression) reveals the range of morphological variability of an identified neuron 14. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50150/50150fig5large.jpg" target="_blank"> Click here to view larger figure</a>.
<ol class="ualpha">
 <li>The dAP cell (artificially colored blue) can be unambiguously recognized within the apterousGal4-UASGFP pattern in the abdomen (dAP, Figure 5A, Lundgren et al. 1995). GFP expressing cells have been stained with an anti-GFP antibody. The midline is marked with dotted lines. ac = anterior commissure, pc = posterior commissure.</li>
 <li>shows an example of a photoconverted DiI-filled dAP cell. The cell was identified prior to dye filling in an apterousGal4-UASGFP embryo by its expression of GFP.</li>
 <li>A gallery of 19 fills of this cell, drawn as maximum projections from image stacks. Dorsal views. Anterior is up. The neuropile is in light grey, the cortex area is in darker grey.</li>
 <li>The 19 dAP cells are stacked with each cell having an opacity of 12%. The region occupied by most of the cells has the darkest shade of grey. The neuropile of all specimens (blue) was scaled to a similar width and the anterior commissures next to the cell were aligned so they overlapped. While the cell body varies more than one cell diameter around the central position, the mediolateral position of the axon within the neuropile shows less variation.</li>
</ol>

Watch this Scientific Journal Video about Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster at JoVE.com

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

labeling-single-cells-central-nervous-system-drosophila

Nanyang Technological University

Research

JoVE Journal

Neuroscience

12.7K Views.  University of Mainz. We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

Video: Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

Optic Nerve Transection: A Model of Adult Neuron Apoptosis in the Central Nervous System

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve 
transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous
 chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals 
through the vitreous fluid ensures that they act upon the entire RGC population. Moreover, RGCs can be selectively transfected by applying short interfering RNAs 
(siRNAs), plasmids, or viral vectors to the cut end of the optic nerve 5-7 or injecting vectors into their target, the superior colliculus 8. 
This allows researchers to study apoptotic mechanisms in the desired neuronal population without confounding effects on other bystander neurons or surrounding glia. 
An additional benefit is the ease and accuracy with which cell survival can be quantified after injury. The retina is a flat, layered tissue and RGCs are localized in 
the innermost layer, the ganglion cell layer. The survival of RGCs can be tracked over time by applying a fluorescent tracer (3% Fluorogold) to the cut end of the 
optic nerve at the time of axotomy, or by injecting the tracer into the superior colliculus (RGC target) one week prior to axotomy. The tracer is retrogradely transported, labeling 
the entire RGC population. Because the ganglion cell layer is a monolayer (one cell thick), RGC densities can be quantified in flat-mounted tissue, without the need 
for stereology. Optic nerve transection leads to the apoptotic death of 90% of injured RGCs within 14 days postaxotomy 9-11. RGC apoptosis has a 
characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a time window for 
experimental manipulations directed against pathways involved in apoptosis.

Optic Nerve transection is a widely used model of adult CNS injury. Ninety percent of retinal ganglion cells (RGCs) whose axons are completely transected (axotomy) die within 14 days after axotomy. This model is easily amenable to experimental manipulations and highly reproducible.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be ...

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, both in preclinical and clinical studies. However, to date results regarding functional outcome and/or tissue regeneration following stem cell transplantation are quite diverse. Generally, a clinical benefit is observed without profound understanding of the underlying mechanism(s)1. Therefore, multiple efforts have led to the development of different molecular imaging modalities to monitor stem cell grafting with the ultimate aim to accurately evaluate survival, fate and physiology of grafted stem cells and/or their micro-environment. Changes observed in one or more parameters determined by molecular imaging might be related to the observed clinical effect. In this context, our studies focus on the combined use of bioluminescence imaging (BLI), magnetic resonance imaging (MRI) and histological analysis to evaluate stem cell grafting.
BLI is commonly used to non-invasively perform cell tracking and monitor cell survival in time following transplantation2-7, based on a biochemical reaction where cells expressing the Luciferase-reporter gene are able to emit light following interaction with its substrate (e.g. D-luciferin)8, 9. MRI on the other hand is a non-invasive technique which is clinically applicable10 and can be used to precisely locate cellular grafts with very high resolution11-15, although its sensitivity highly depends on the contrast generated after cell labeling with an MRI contrast agent. Finally, post-mortem histological analysis is the method of choice to validate research results obtained with non-invasive techniques with highest resolution and sensitivity. Moreover end-point histological analysis allows us to perform detailed phenotypic analysis of grafted cells and/or the surrounding tissue, based on the use of fluorescent reporter proteins and/or direct cell labeling with specific antibodies.
In summary, we here visually demonstrate the complementarities of BLI, MRI and histology to unravel different stem cell- and/or environment-associated characteristics following stem cell grafting in the CNS of mice. As an example, bone marrow-derived stromal cells, genetically engineered to express the enhanced Green Fluorescent Protein (eGFP) and firefly Luciferase (fLuc), and labeled with blue fluorescent micron-sized iron oxide particles (MPIOs), will be grafted in the CNS of immune-competent mice and outcome will be monitored by BLI, MRI and histology (Figure 1).

This article describes an optimized sequence of events for multimodal imaging of cellular grafts in rodent brain using: (i) in vivo bioluminescence and magnetic resonance imaging, and (ii) post mortem histological analysis. Combining these imaging modalities on a single animal allows cellular graft evaluation with high resolution, sensitivity and specificity.

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, ...

Production of Lentiviral Vectors for Transducing Cells from the Central Nervous System

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing therapeutic approaches. Lentiviral vectors are attractive tools in transduction of neurons and other cell types in CNS as they transduce both dividing and non-dividing cells, support sustained expression of transgenes, and have relatively large packaging capacity and low toxicity 1-3. Lentiviral vectors have been successfully used in transducing many neural cell types in vitro 4-6 and in animals 7-10.
Great efforts have been made to develop lentiviral vectors with improved biosafety and efficiency for gene delivery. The current third generation replication-defective and self-inactivating (SIN) lentiviral vectors are depicted in Figure 1. The required elements for vector packaging are split into four plasmids. In the lentiviral transfer plasmid, the U3 region in the 5' long terminal repeat (LTR) is replaced with a strong promoter from another virus. This modification allows the transcription of the vector sequence independent of HIV-1 Tat protein that is normally required for HIV gene expression 11. The packaging signal (&Psi;) is essential for encapsidation and the Rev-responsive element (RRE) is required for producing high titer vectors. The central polypurine tract (cPPT) is important for nuclear import of the vector DNA, a feature required for transducing non-dividing cells 12. In the 3' LTR, the cis-regulatory sequences are completely removed from the U3 region. This deletion is copied to 5' LTR after reverse transcription, resulting in transcriptional inactivation of both LTRs. Plasmid pMDLg/pRRE contains HIV-1 gag/pol genes, which provide structural proteins and reverse transcriptase. pRSV-Rev encodes Rev which binds to the RRE for efficient RNA export from the nucleus. pCMV-G encodes the vesicular stomatitis virus glycoprotein (VSV-G) that replaces HIV-1 Env. VSV-G expands the tropism of the vectors and allows concentration via ultracentrifugation 13. All the genes encoding the accessory proteins, including Vif, Vpr, Vpu, and Nef are excluded in the packaging system. The production and manipulation of lentiviral vectors should be carried out according to NIH guidelines for research involving recombinant DNA (<a href="http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf" target="_blank">http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf</a>). An approval from individual Institutional Biological and Chemical Safety Committee may be required before using lentiviral vectors. Lentiviral vectors are commonly produced by cotransfection of 293T cells with lentiviral transfer plasmid and the helper plasmids encoding the proteins required for vector packaging. Many lentiviral transfer plasmids and helper plasmids can be obtained from Addgene, a non-profit plasmid repository (<a href="http://www.addgene.org/" target="_blank">http://www.addgene.org/</a>). Some stable packaging cell lines have been developed, but these systems provide less flexibility and their packaging efficiency generally declines over time 14, 15. Commercially available transfection kits may support high efficiency of transfection 16, but they can be very expensive for large scale vector preparations. Calcium phosphate precipitation methods provide highly efficient transfection of 293T cells and thus provide a reliable and cost effective approach for lentiviral vector production.
In this protocol, we produce lentiviral vectors by cotransfection of 293T cells with four plasmids based on the calcium phosphate precipitation principle, followed by purification and concentration with ultracentrifugation through a 20% sucrose cushion. The vector titers are determined by fluorescence- activated cell sorting (FACS) analysis or by real time qPCR. The production and titration of lentiviral vectors in this protocol can be finished with 9 days. We provide an example of transducing these vectors into murine neocortical cultures containing both neurons and astrocytes. We demonstrate that lentiviral vectors support high efficiency of transduction and cell type-specific gene expression in primary cultured cells from CNS.

In this protocol we describe production, purification and titration of lentiviral vectors. We provide an example of lentiviral vector-mediated gene delivery in primary cultured neurons and astrocytes. Our methods may also apply to other cell types in vitro and in vivo.

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing ...

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are tightly regulated, affecting the final dynamics of protein abundance. Various regulatory mechanisms exist at the translation step, rendering mRNA levels alone an unreliable indicator of gene expression. In addition, local regulation of mRNA translation has been particularly implicated in neuronal functions, shifting &#39;translatomics&#39; to the focus of attention in neurobiology. The presented method can be used to bridge transcriptomics and proteomics.
Here we describe essential modifications to the technique of polyribosome fractionation, which interrogates the translatome based on the association of actively translated mRNAs to multiple ribosomes and their differential sedimentation in sucrose gradients. Traditionally, working with in vivo samples, particularly of the central nervous system (CNS), has proven challenging due to the restricted amounts of material and the presence of fatty tissue components. In order to address this, the described protocol is specifically optimized for use with minimal amount of CNS material, as demonstrated by the use of single mouse spinal cord and brain. Briefly, CNS tissues are extracted and translating ribosomes are immobilized on mRNAs with cycloheximide. Myelin flotation is then performed to remove lipid rich components. Fractionation is performed on a sucrose gradient where mRNAs are separated according to their ribosomal loading. Isolated fractions are suitable for a range of downstream assays, including new genome wide assay technologies.

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

This protocol illustrates essential modifications of polyribosome fractionation in order to study the translatome of in vivo CNS samples. It allows global assessment of translation and transcription regulation through the isolation and comparison of total RNA to ribosome bound RNA fractions.

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...