This work was supported by the intramural program of the National Institutes of Neurological Disorders and Stroke at the National Institutes of Health. We would also like to thank Dr. George Mentis for his earlier contributions to the method and to the data in Figure 3.

Fluorescent dextrans have been used as anatomical tools and for imaging neuronal activity1-4. Fields et al., (2009)4 published a protocol for applying ion- and voltage-sensitive dyes to axonal tracts with a focus on the spinal cord as a model system. Here we describe a more detailed procedure for applying fluorescent and/or ion-sensitive dye to the cut ventral roots, dorsal roots or any neuronal tract of the spinal cord for in-vitro optophysiological and morphological studies.
1. Type-I and Type-II Pipettes
Start by pulling two short sections of polyethylene tubing (PE90, Clay Adams Brand) over the flame of an alcohol lamp7 to produce pipettes with tapered tips. One pipette (Type-I) should be short (3-7 mm) with a small tip that can tightly hold the target root or tract (Outer Diameter: 0.2-0.4 mm; Inner Diameter: 0.1-0.3 mm). Then pull a second pipette (Type-II) with a longer (8-12 cm) thinner, shaft and a very fine tip (Outer Diameter: 0.2-0.3 mm; inner Diameter: 0.1-0.2 mm) that can be inserted into the Type-I pipette as far as its tip. The second thinner pipette will be used to aspirate and introduce artificial cerebrospinal fluid (aCSF) and dye solution respectively from the Type-I pipette.
2. Positioning of Type-I Pipette
Dissect and isolate the spinal cord8,9 and place it in a bath, superfused with chilled aCSF (~16 &deg;C) throughout the loading process. Place the Type-I pipette on an electrode holder (H1/12 electrode holder, Narishige) so that its back opening can be easily connected to a syringe (1ml U-100 Insulin syringe, Becton Dickenson or comparable) via a flexible tubing (PharMed BPT, Cole-Parmer, #AY242002; 10 cm) (Fig.1A-B).
 
3. Dye Application
<ol>
	<li>Using the syringe, first draw aCSF into the Type-I pipette (Fig.1B; 2A) followed by the axonal tract or root to be filled (Fig. 2B). The flexible tubing should then be removed from the back opening. The Type-II pipette is attached to another syringe and inserted into the Type-I pipette holding the axonal tract or root. Using the syringe attached to the type-II pipette, aspirate aCSF so that the residual aCSF just covers the axonal tract (Fig.1C; 2C-D). Then withdraw the type-II pipette from the type-I pipette. Monitor the aCSF level for a few minutes to verify a good seal on the axonal tract or nerves (see Section 3 in Troubleshooting for "good seal").</li>
	<li>Dissolve the dye in aCSF or 0.2% triton X-100 in double distilled water and draw it into the Type-II pipette while paying attention not to include air bubbles. Insert the Type-II pipette containing the dye into the Type-I pipette and into the residual aCSF solution (Fig. 2E). Slowly release the dye into the aCSF using gentle positive pressure (Fig. 2F). Be careful not to introduce any air bubbles or to cause displacement of the target tissue from inside the tip. Withdraw the type-II pipette after sufficient amount of dye has been released (3-5 &mu;L of dye solution).</li>
</ol>
4. Incubation
Incubate the tissue in the dark between 6 to 20 hours depending on the type of experiment. Once sufficient time has been allowed for filling, gently pull away the electrode from the tissue leaving the tissue ready for imaging or histology.
Troubleshooting
<ol>
	<li>If the target tissue does not enter easily into the tip of the Type-I pipette or is loose once inside, then the tip diameter is the wrong size. If this occurs, pull and use a different pipette.</li>
	<li>Before adding the dye to the aCSF in the type-I pipette, make sure that enough aCSF remains so that it can be reached with the Type-II pipette tip (Fig. 2D). Otherwise, an air gap will exist once the dye is added. If this happens, do not add the dye and remove the axonal tract from the pipette and then draw it back in with sufficient aCSF to be reached by the Type-II pipette.</li>
	<li>If the aCSF level in the Type-I pipette increases after it was aspirated with the Type-II pipette, it means that the seal with the tissue is not good. Use a different electrode, otherwise the dye will be diluted during the labeling procedure.</li>
	<li>If an air bubble is introduced into the Type-I pipette while injecting the dye, drain out the bubble by advancing the Type-II pipette tip close to the bubble and gently applying a weak negative pressure using the attached syringe.</li>
</ol>
5. Representative Results
To illustrate an application of the method, we simultaneously loaded motoneurons, sensory afferents and spinal interneurons with three different dextran-conjugated fluorescent dyes (Fig. 3A). As illustrated in Fig. 3B, three classes of neuron are labeled; sensory afferents (red), interneurons projecting into the ventral funiculus (green) and motoneurons (blue).
<img alt="Figure 1" src="/files/ftp_upload/4008/4008fig1.jpg" /> 
Figure 1. Schematic showing Pipette Type-I and Type-II assembly. (A) A Type-I pipette (light blue) is placed on an electrode holder so that it's back opening can be easily connected to the flexible elastomer tubing (B) and into which a Type-II pipette can be inserted (C).
<img alt="Figure 2" src="/files/ftp_upload/4008/4008fig2.jpg" /> 
Figure 2. Schematic showing the loading process of motoneurons with fluorescent dyes. (A) A Type-I pipette is placed near the ventral root to be filled. (B) Suction is applied to the pipette to draw aCSF into the pipette followed by the root. (C-D) Disconnect the flexible elastomer tubing from the backend of Type-I pipette and reduce the amount of aCSF by applying suction to the Type-II pipette. (E) Insert the tip of the Type-II pipette containing the dissolved dye into the remaining aCSF in the Type-I pipette. (E-F) Slowly fill the Type-I pipette with the dye and then remove the Type-II pipette.
<img alt="Figure 3" src="/files/ftp_upload/4008/4008fig3.jpg" /> 
Figure 3. Application of the filling procedure to the isolated mouse spinal cord. (A) Schematic showing the three Type-I pipettes used to fill different neuronal classes in the spinal cord. Motoneurons (blue; Cascade Blue dextran) and sacral sensory afferents (red; Texas-Red dextran) were backfilled through the ventral and dorsal roots, respectively. Fluorescein dextran was used to load spinal interneurons (green) whose axons ascend through the ventral funiculus (VF). 1 mg of each dextran dye (10,000MW; Molecular Probes) was dissolved in 6 &mu;L distilled water containing 0.2% Triton X-100 (B). Confocal image of a section from the sacral cord of a spinal cord in which the neurons were back-labeled as described in A. Note that the labeled sensory afferents are primarily ipsilateral with a few contralateral projections. The labeled motoneurons are ipsilateral to the filled ventral root whereas the labeled interneurons are contralateral to the side of the fill (arrow head). Calibration bar 200 &mu;m.

<ol>
	<li>Nance, D. M., Burns, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+dextrans+as+sensitive+anterograde+neuroanatomical+tracers:+applications+and+pitfalls.">Fluorescent dextrans as sensitive anterograde neuroanatomical tracers: applications and pitfalls.</a> Brain Res. Bull. 25, 139-145 (1990).</li><li>Glover, J. C., Petursdottir, G., Jansen, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+dextran-amines+used+as+axonal+tracers+in+the+nervous+system+of+the+chicken+embryo.">Fluorescent dextran-amines used as axonal tracers in the nervous system of the chicken embryo.</a> J. Neurosci. Methods. 18, 243-254 (1986).</li><li>McPherson, D. R., McClellan, A. D., O'Donovan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+imaging+of+neuronal+activity+in+tissue+labeled+by+retrograde+transport+of+Calcium+Green+Dextran.">Optical imaging of neuronal activity in tissue labeled by retrograde transport of Calcium Green Dextran.</a> Brain Research Protocols. 1, 157-164 (1997).</li><li>Fields, D. R., Shneider, N., Mentis, G. Z., O'Donovan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+nervous+system+activity.">Imaging nervous system activity.</a> Curr. Protoc. Neurosci. Chapter 2, Unit 2.3 (2009).</li><li>O'Donovan, M. J., Ho, S., Sholomenko, G., Yee, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+imaging+of+neurons+retrogradely+and+anterogradely+labelled+with+calcium-sensitive+dyes.">Real-time imaging of neurons retrogradely and anterogradely labelled with calcium-sensitive dyes.</a> J. Neurosci. Methods. 46, 91-106 (1993).</li><li>Wenner, P., Tsau, Y., Cohen, L. B., O'Donovan, M. J., Dan, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Voltage-sensitive+dye+recording+using+retrogradely+transported+dye+in+the+chicken+spinal+cord:+staining+and+signal+characteristics.">Voltage-sensitive dye recording using retrogradely transported dye in the chicken spinal cord: staining and signal characteristics.</a> J. Neurosci. Methods. 70, 111-120 (1996).</li><li>Garudadri, S., Gallarda, B., Pfaff, S., Alaynick, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spinal+Cord+Electrophysiology+II:+Extracellular+Suction+Electrode+Fabrication.">Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication.</a> J. Vis. Exp. (48), e2580 (2011).</li><li>Smith, J. C., Feldman, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+brainstem-spinal+cord+preparations+for+study+of+motor+systems+for+mammalian+respiration+and+locomotion.">In vitro brainstem-spinal cord preparations for study of motor systems for mammalian respiration and locomotion.</a> J. Neurosci. 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Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+functional+impairment+of+sensory-motor+connectivity+in+a+mouse+model+of+spinal+muscular+atrophy.">Early functional impairment of sensory-motor connectivity in a mouse model of spinal muscular atrophy.</a> Neuron. 69, 453-467 (2011).</li><li>Blivis, D., Mentis, Z., O'Donovan, J. M. G., Lev-Tov, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studies+of+sacral+neurons+involved+in+activation+of+the+lumbar+central+pattern+generator+for+locomotion+in+the+neonatal+rodent+spinal+cord.">Studies of sacral neurons involved in activation of the lumbar central pattern generator for locomotion in the neonatal rodent spinal cord.</a> Soc. Neurosci. Abstr. 564.8, (2009).</li><li>O'Donovan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+the+spatiotemporal+organization+of+neural+activity+in+the+developing+spinal+cord.">Imaging the spatiotemporal organization of neural activity in the developing spinal cord.</a> Dev. Neurobiol. 68, 788-803 (2008).</li><li>Kasumacic, N., Glover, J. C., Perreault, M. -. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Segmental+patterns+of+vestibular-mediated+synaptic+inputs+to+axial+and+limb+motoneurons+in+the+neonatal+mouse+assessed+by+optical+recording.">Segmental patterns of vestibular-mediated synaptic inputs to axial and limb motoneurons in the neonatal mouse assessed by optical recording.</a> J. Physiol. (Lond). 588, 4905-4925 (2010).</li><li>Szokol, K., Glover, J. C., Perreault, M. -. 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We have nothing to disclose.

We describe here a simple and cost-effective protocol for introducing dyes into neurons, nerves and spinal tracts. This method involves exposing identified anatomical pathways to a highly concentrated dye solution for the duration of the loading process. This results in retrograde labeling of the target site with little background compared to microinjections, bath application and electroporation techniques. However, the method is limited to sites in the nervous system where a tract of axons can be isolated. Furthermore,...

<table border="1">
 <tbody>
 <tr>
 <td colspan="2">Material Name</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td rowspan="2">Tubing</td>
 <td>PE90 (IDxOD:0.034"x0.050"; Wall Thikness:0.008")</td>
 <td>Clay Adams Brand Intermedic</td>
 <td>427421</td>
 <td>For Type I &amp; Type II pipettes</td>
 </tr>
 <tr>
 <td>NSF-51(IDxOD:1/16x1/8; Wall Thikness:1/32)</td>
 <td>PharMed BPT, Cole-Parmer</td>
 <td>AY242002</td>
 <td>For Flexible tubing</td>
 </tr>
 <tr>
 <td>Syringe</td>
 <td>1ml insulin Syringe U-100</td>
 <td>Becton Dickenson</td>
 <td>329650</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Needle</td>
 <td>19G x1-1/2" Metal Hub Needle</td>
 <td>MONOJECT</td>
 <td>200136</td>
 <td>Connecting Type II pipette to syringe</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>Alcohol Lamp</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Stopcock</td>
 <td>Three-way Stopcock with Male Luer Slip Adapter</td>
 <td>Baxter Healthcare Corp.</td>
 <td>2C6241</td>
 <td>Use with syringe</td>
 </tr>
 <tr>
 <td rowspan="3">Holder</td>
 <td>H-1 electrode holder</td>
 <td>Narishige</td>
 <td>H-1/12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Magnet stand</td>
 <td>Narishige</td>
 <td>GJ-8</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micromanipulator</td>
 <td>Narishige</td>
 <td>M-3333</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tweezers</td>
 <td>S&amp;T DUMONT Swiss 00632-11</td>
 <td>DUMONT</td>
 <td>JF-5 TC</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
 </table>
*Those are suggested materials. Can be replaced with any compatible products

retrograde loading of nerves, tracts, and spinal roots with fluorescent dyes

Retrograde labeling of neurons is a standard anatomical method1,2 that has also been used to load calcium and voltage-sensitive dyes into neurons3-6. Generally, the dyes are applied as solid crystals or by local pressure injection using glass pipettes. However, this can result in dilution of the dye and reduced labeling intensity, particularly when several hours are required for dye diffusion. Here we demonstrate a simple and low-cost technique for introducing fluorescent and ion-sensitive dyes into neurons using a polyethylene suction pipette filled with the dye solution. This method offers a reliable way for maintaining a high concentration of the dye in contact with axons throughout the loading procedure.

Watch this Scientific Journal Video about Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes at JoVE.com

Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes

We describe a simple and low cost technique for introducing high concentration of fluorescent and calcium-sensitive dyes into neurons or any neuronal tract using a polyethylene suction pipette.

Retrograde labeling of neurons is a standard anatomical method1,2 that has also been used to load calcium and voltage-sensitive dyes into neurons3-6. ...

retrograde-loading-nerves-tracts-spinal-roots-with-fluorescent

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12.2K Views.  National Institute of Neurological Disorders and Stroke, National Institutes of Health. We describe a simple and low cost technique for introducing high concentration of fluorescent and calcium-sensitive dyes into neurons or any neuronal tract using a polyethylene suction pipette.

Video: Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Optical Imaging of Neurons in the Crab Stomatogastric Ganglion with Voltage-sensitive Dyes

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous activity of participating neurons leads to the emergence of the integral functionality of the network. Here we present the methodology of application of this technique to identified pattern generating neurons in the crab stomatogastric ganglion. We demonstrate the loading of these neurons with the fluorescent voltage-sensitive dye Di-8-ANEPPQ and we show how to image the activity of dye loaded neurons using the MiCAM02 high speed and high resolution CCD camera imaging system. We demonstrate the analysis of the recorded imaging data using the BVAna imaging software associated with the MiCAM02 imaging system. The simultaneous voltage-sensitive dye imaging of the detailed activity of multiple neurons in the crab stomatogastric ganglion applied together with traditional electrophysiology techniques (intracellular and extracellular recordings) opens radically new opportunities for the understanding of how central pattern generator neural networks work.

Here we present the methodology for fast and high resolution fluorescent voltage-sensitive dye imaging of detailed activity of neurons in the crab stomatogastric ganglion.

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Promotion of Survival and Differentiation of Neural Stem Cells with Fibrin and Growth Factor Cocktails after Severe Spinal Cord Injury

Neural stem cells (NSCs) can self-renew and differentiate into neurons and glia. Transplanted NSCs can replace lost neurons and glia after spinal cord injury (SCI), and can form functional relays to re-connect spinal cord segments above and below a lesion. Previous studies grafting neural stem cells have been limited by incomplete graft survival within the spinal cord lesion cavity. Further, tracking of graft cell survival, differentiation, and process extension had not been optimized. Finally, in previous studies, cultured rat NSCs were typically reported to differentiate into glia when grafted to the injured spinal cord, rather than neurons, unless fate was driven to a specific cell type. To address these issues, we developed new methods to improve the survival, integration and differentiation of NSCs to sites of even severe SCI. NSCs were freshly isolated from embryonic day 14 spinal cord (E14) from a stable transgenic Fischer 344 rat line expressing green fluorescent protein (GFP) and were embedded into a fibrin matrix containing growth factors; this formulation aimed to retain grafted cells in the lesion cavity and support cell survival. NSCs in the fibrin/growth factor cocktail were implanted two weeks after thoracic level-3 (T3) complete spinal cord transections, thereby avoiding peak periods of inflammation. Resulting grafts completely filled the lesion cavity and differentiated into both neurons, which extended axons into the host spinal cord over remarkably long distances, and glia. Grafts of cultured human NSCs expressing GFP resulted in similar findings. Thus, methods are defined for improving neural stem cell grafting, survival and analysis of in vivo findings.

Fibrin matrices containing growth factors were used to retain grafted neural stem cells into sites of complete spinal cord transection. Grafted cells completely filled the lesion cavity and differentiated into multiple neural cell types, including neurons that extended axons into host spinal cord over long distances.

Neural stem cells (NSCs) can self-renew and differentiate into neurons and glia. Transplanted NSCs can replace lost neurons and glia after spinal cord ...

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate goal is to image neuronal activity in vivo, one must be able to image activity of a single cell to ensure successful in vivo preparations. This procedure will describe how to image membrane potential in a single cell to provide a foundation to eventually image in vivo. Here we describe methods for imaging GEVIs consisting of a voltage-sensing domain fused to either a single fluorescent protein (FP) or two fluorescent proteins capable of F&#246;rster resonance energy transfer (FRET) in vitro. Using an image splitter enables the projection of images created by two different wavelengths onto the same charge-coupled device (CCD) camera simultaneously. The image splitter positions a second filter cube in the light path. This second filter cube consists of a dichroic and two emission filters to separate the donor and acceptor fluorescent wavelengths depending on the FPs of the GEVI. This setup enables the simultaneous recording of both the acceptor and donor fluorescent partners while the membrane potential is manipulated via whole cell patch clamp configuration. When using a GEVI consisting of a single FP, the second filter cube can be removed allowing the mirrors in the image splitter to project a single image onto the CCD camera.

A method for imaging changes in membrane potential using genetically encoded voltage indicators is described.

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate ...

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic nerves innervating the respiratory diaphragm. In spinal cord slices, phrenic motor neurons cannot be identified from other motor neurons on morphological or biochemical criteria. We provide the description of procedures for visualizing phrenic motor neuron cell bodies in mice, following intrapleural injections of cholera toxin subunit beta (CTB) conjugated to a fluorophore. This fluorescent neuroanatomical tracer has the ability to be caught up at the diaphragm neuromuscular junction, be carried retrogradely along the phrenic axons and reach the phrenic cell bodies. Two methodological approaches of intrapleural CTB delivery are compared: transdiaphragmatic versus transthoracic injections. Both approaches are successful and result in similar number of CTB-labeled phrenic motor neurons. In conclusion, these techniques can be applied to visualize or quantify the phrenic motor neurons in various experimental studies such as those focused on the diaphragm-phrenic circuitry.

Here, we describe a protocol for identifying phrenic motor neurons in mice after intrapleural delivery of fluorophore conjugated cholera toxin subunit beta. Two techniques are compared to inject the pleural cavity: transdiaphragmatic versus transthoracic approaches.

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic ...

Activity-based Training on a Treadmill with Spinal Cord Injured Wistar Rats

Spinal cord injury (SCI) results in lasting deficits that include both mobility and a multitude of autonomic-related dysfunctions. Locomotor training (LT) on a treadmill is widely used as a rehabilitation tool in the SCI population with many benefits and improvements to daily life. We utilize this method of activity-based task-specific training (ABT) in rodents after SCI to both elucidate the mechanisms behind such improvements and to enhance and improve upon existing clinical rehabilitation protocols. Our current goal is to determine the mechanisms underlying ABT-induced improvements in urinary, bowel, and sexual function in SCI rats after a moderate to severe level of contusion. After securing each individual animal in a custom-made adjustable vest, they are secured to a versatile body weight support mechanism, lowered to a modified three-lane treadmill and assisted in step-training for 58 minutes, once a day for 10 weeks. This setup allows for the training of both quadrupedal and forelimb-only animals, alongside two different non-trained groups. Quadrupedal-trained animals with body weight support are aided by a technician present to assist in stepping with proper hind limb placement as necessary, while forelimb-only trained animals are raised at the caudal end to ensure no hind limb contact with the treadmill and no weight-bearing. One non-trained SCI group of animals is placed in a harness and rests next to the treadmill, while the other control SCI group remains in its home cage in the training room nearby. This paradigm allows for the training of multiple SCI animals at once, thus making it more time-efficient in addition to ensuring that our pre-clinical animal model mimics the clinical representation as close as possible, particularly with respect to the body weight support with manual assistance.

This protocol demonstrates our model of activity-based locomotor treadmill training for rats with spinal cord injury (SCI). Included is both quadrupedal and forelimb-only groups, in addition to two distinct types of non-trained control groups. Investigators are able to assess training effects on SCI rats using this protocol.

Spinal cord injury (SCI) results in lasting deficits that include both mobility and a multitude of autonomic-related dysfunctions. Locomotor training (LT) ...

Dissection of Pelvic Autonomic Ganglia and Associated Nerves in Male and Female Rats

The bilateral major pelvic ganglia (MPG; synonym, pelvic ganglia) are the primary source of postganglionic sympathetic and parasympathetic neurons innervating pelvic organs of rodents; the functionally equivalent structure in humans is the inferior hypogastric plexus. The major pelvic ganglia also provide the route by which lumbar and sacral sensory axons reach the pelvic organs. These complex, mixed ganglia can prove challenging to identify and dissect for further experimental study of normal autonomic mechanisms or to establish preclinical models of disease, injury or visceral pain. Here we describe a protocol to access and visualize these ganglia and their associated nerve tracts. We provide this protocol with schematics for both male and female rats, as the ganglion size and landmarks for identification differ between sexes. The protocol describes removal of the ganglion for in vitro studies, but this method can be integrated into a surgical recovery protocol for experimental interventions (e.g., nerve crush, nerve resection) or for mapping neuronal circuits (e.g., by microinjection of neural tracers). We also demonstrate the primary structures of the ganglion and its associated nerves immediately following dissection and following immunohistochemical staining.

The major pelvic ganglia contain parasympathetic and sympathetic neurons that innervate pelvic organs. Here we describe a dissection method and provide schematics for identification of these ganglia and their associated nerves. These methods can be applied to experimental manipulation of these ganglia in vivo or removal post-mortem for further study.