The authors would like to thank the Kucenas Lab for valuable discussions and Quorum Technologies, Inc. for superb technical support. The work was supported by the UVa Fund for Excellence in Science and Technology (FEST) (S.K.).

1. Preparation and Mounting of Zebrafish Embryos for Ablation and Live Imaging
<ol>
	<li>Prepare a stock of 0.8% low melt agarose in egg water. Aliquot into 13X 100 mm disposable culture tubes and store at 4 &deg;C until needed.</li>
	<li>Cross adult zebrafish containing stably integrated transgenes to fluorescently label motor neurons and glial cell types of interest. Collect zebrafish embryos in egg water and place in 28.5 &deg;C incubator for correct staging later 3.</li>
	<li>At approximately 24 hr post fertilization (hpf), remove egg water and add 0.002% 1-phenyl 2-thiourea (PTU) in egg water. Return embryos to the incubator.</li>
	<li>Between 24 and 96 hpf, embryos should be screened for the presence of desired transgenes on a fluorescent dissecting scope. Place selected embryos in fresh PTU egg water and return to the incubator.</li>
	<li>When larvae have reached 6 days post fertilization (dpf) (or desired age), remove them from the incubator, select a few larvae for mounting, and transfer them to a smaller dish.</li>
	<li>Remove the water from the dish and immediately replace with approximately 0.02% Tricane in PTU egg water. Allow larvae to sit in anesthetic approximately 5 min.</li>
	<li>Place an aliquot of 0.8% low melt agarose in a beaker with tap water and microwave for 30 sec, or until the agarose is melted. Allow the beaker to cool until the agarose in the culture tube feels lukewarm to the touch (not hot).</li>
	<li>Select an anesthetized larva and transfer it to a single or multi-well 35 mm glass bottom dish. Remove any water that was transferred with the larva, then immediately cover the larva with enough warm agarose to fill the glass-bottomed portion of the dish but not create a large dome.</li>
	<li>As the agarose hardens, use a dissecting needle to position the larva on its side at the bottom of the dish, then tilt the larva just slightly on its back. It is imperative for injury and subsequent imaging, that the mounted larva be touching the glass on the bottom of the dish. Use the dissecting needle to maintain the larva in this position until the agarose is solidified and the larva is immobilized.</li>
	<li>Once the agarose is fully hardened, slowly pipette enough Tricane water (this can be the same Tricane used for anesthesia) into the dish to completely cover the agarose and larva.</li>
</ol>
2. Laser Calibration and Testing
<ol>
	<li>Turn on all confocal microscope instrumentation, appropriate diode lasers for exciting zebrafish transgenes, and the nitrogen-pumped dye laser. Be sure the "blank" beam splitter is in place, the laser attenuator is fully open, and the 435 nm Coumarin dye cell is in place. On the computer, open imaging software.</li>
	<li>Move a 63x 1.2NA water immersion objective into position and apply a small drop of immersion medium for water objectives onto the objective. A 63x objective will allow for good laser ablation accuracy, and water immersion allows for a larger working distance, which is necessary when working with 6-7 dpf zebrafish larvae.</li>
	<li>Obtain a glass slide with one mirrored side to use for calibration of the laser. Place the slide, mirror side down, onto the microscope stage.</li>
	<li>Using the eyepiece under brightfield illumination, find and focus on a scratch or etch in the mirror. The brightfield light will be shining down onto the glass slide, but will only pass through to the objective in places where the mirror has been scratched or etched away, causing the mirror to look black through the eyepiece, and the etchings to look like spots or lines of light.</li>
	<li>View and focus the etchings on the computer screen using imaging software. Open the window that controls the laser calibration and power settings. Set the number of pulses to "1" and the attenuation plate to "3" % transmission. The number of pulses represents the number of times the laser will fire within each region of interest (ROI) and the % transmission represents the laser power. Be sure the correct calibration setting is selected for the 63x 1.2NA water immersion objective.</li>
	<li>Select the ellipse tool from the main toolbar, and click the image on the computer screen to create a single circular ROI. Do this 3 more times until there are 4 circular ROIs spaced randomly over the image.</li>
	<li>Some systems may include a safety feature that will not allow the laser to fire if the light path to the oculars is open. If this is the case, manually close the light path to the oculars at this time.</li>
	<li>Fire the laser. This will create 4 small spot etchings, 1 within each circular ROI. If the laser does not fire, check to be sure the laser is turned on and connected properly, and that the light path to the oculars is closed. If the laser fires but no etching is seen, check to be sure the "blank" beam splitter is in place. If everything is in place, increase the laser power and "frap" until the etchings are visible. If a laser power setting greater than 10 is necessary to etch the glass, this may indicate the laser plasma cartridge needs to be replaced.</li>
	<li>If the etched spots appear centered within the selected ROIs, no calibration is necessary (proceed to 2.12). If the spots are not centered, the calibration setting needs to be updated.</li>
	<li>To calibrate, click the update the calibration setting. The laser will fire and an image will appear containing a single etched point. If the point does not appear, cancel the calibration, increase the laser power, and start the calibration again.</li>
	<li>Click in the center of the spot. The laser will fire again, and another point will appear. Click that point, and repeat this process until the calibration is complete and 9 spots appear in a grid fashion. Repeat steps 2.6-2.9 to check that the calibration was successful.</li>
	<li>Remove the glass slide from the microscope stage and clean the objective. Open the light path to the oculars, and replace the "blank" beam splitter with the "100%ILL" beam splitter.</li>
</ol>
3. Nerve Transection Using Laser Ablation and Time-lapse Confocal Imaging of Glial Cell Behaviors
<ol>
	<li>Remove the stage used to hold the glass slide and replace with a stage suitable for holding 35 mm glass bottomed dishes. Continue to use the 63x 1.2NA water immersion objective.</li>
	<li>Apply a small drop of water immersion medium to the objective, and place the dish with mounted larva on the stage. Stabilize the dish with clips.</li>
	<li>Using the eyepiece and widefield illumination, focus the larva and locate the motor nerves. Scan the nerves in hemisegments 10-20 and select a motor nerve for transection.</li>
	<li>Bring up a live-view of the nerve on the computer screen using imaging software. Select Z-planes and acquire an image of the axons and glial cells of the uninjured nerve.</li>
	<li>Prepare time-lapse imaging settings in the imaging software to capture Z-projections of all cell types in 5-30 min intervals, depending on the experiment. It is best to create all necessary settings for the time-lapse before performing the ablation, so there is no delay in starting the time-lapse once the injury is complete.</li>
	<li>Remove the "100%ILL" beam splitter and replace with the "blank". Close the light path to the oculars.</li>
	<li>Return to the live-view of the nerve. Use the appropriate fluorescent channel to view the axons that will be transected.</li>
	<li>Using the ellipse tool, create a thin elliptical ROI in the area to be ablated, then create smaller ROIs within the selected region.</li>
	<li>In the window that controls the laser settings, set the number of pulses to "2" and the attenuation plate to "18" % transmission. Fire the laser within the selected ROIs. If fluorescence remains within the ROIs, increase the laser power, wait approximately 10 sec, and fire the laser again. Do this until you reach a setting that causes fluorescence to disappear within the ROIs.</li>
	<li>Wait 10 or more seconds and check the ablated area again for fluorescence. Beware that an ROI may initially appear ablated, when it is actually photobleached. If fluorescence returns, increase the laser power and fire the laser again. Repeat this until florescence disappears within the ROIs and does not return within 10 sec. It is best to start with a lesser laser power and increase to the necessary power. The necessary laser power may vary based on individual microscopy systems, age of Coumarin dye, specimen mounting, age of larvae, and thickness of the tissue. Once an ideal laser power has been established, this power setting may be used again on subsequent nerves in the same experiment.</li>
	<li>Once the ablation is completed, begin time-lapse imaging.</li>
	<li>When the time-lapse is complete, use imaging software to compile data and create color composite Z-projections for each time point. Create a QuickTime movie to analyze the behavior of axons and glial cells simultaneously .</li>
</ol>

<ol>
	<li>Geuna, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+3:+Histology+of+the+peripheral+nerve+and+changes+occurring+during+nerve+regeneration.">Chapter 3: Histology of the peripheral nerve and changes occurring during nerve regeneration.</a> Int. Rev. Neurobiol. 87, 27-46 (2009).</li><li>Hirata, K., Kawabuchi, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myelin+phagocytosis+by+macrophages+and+nonmacrophages+during+Wallerian+degeneration.">Myelin phagocytosis by macrophages and nonmacrophages during Wallerian degeneration.</a> Microsc. Res. Tech. 57, 541-547 (2002).</li><li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+Embryonic+Development+of+the+Zebrafish.">Stages of Embryonic Development of the Zebrafish.</a> Developmental Dynamics. 203, 253-310 (1995).</li><li>Martin, S. M., O'Brien, G. S., Portera-Cailliau, C., Sagasti, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wallerian+degeneration+of+zebrafish+trigeminal+axons+in+the+skin+is+required+for+regeneration+and+developmental+pruning.">Wallerian degeneration of zebrafish trigeminal axons in the skin is required for regeneration and developmental pruning.</a> Development. 137, 3985-3994 (2010).</li><li>O'Brien, G. S., Rieger, S., Martin, S. M., Cavanaugh, A. M., Portera-Cailliau, C., Sagasti, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+axotomy+and+time-lapse+confocal+imaging+in+live+zebrafish+embryos.">Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos.</a> J. Vis. Exp. (24), e1129 (2009).</li><li>Parrinello, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EphB+signaling+directs+peripheral+nerve+regeneration+through+Sox2-dependent+Schwann+cell+sorting.">EphB signaling directs peripheral nerve regeneration through Sox2-dependent Schwann cell sorting.</a> Cell. 143, 145-155 (2010).</li><li>Rosenberg, A. F., Wolman, M. A., Franzini-Armstrong, C., Granato, 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+nerve-macrophage+interactions+following+peripheral+nerve+injury.">In vivo nerve-macrophage interactions following peripheral nerve injury.</a> J. Neurosci. 32, 3898-3909 (2012).</li><li>Stoll, G., Muller, H. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nerve+injury,+axonal+degeneration+and+neural+regeneration:+basic+insights.">Nerve injury, axonal degeneration and neural regeneration: basic insights.</a> Brain Pathol. 9, 313-325 (1999).</li><li>Villegas, R., Martin, S. M., O'Donnell, K., Carrillo, S., Sagasti, A., Allende, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+degeneration+and+regeneration+in+developing+zebrafish+peripheral+axons+reveals+a+requirement+for+extrinsic+cell+types.">Dynamics of degeneration and regeneration in developing zebrafish peripheral axons reveals a requirement for extrinsic cell types.</a> Neural Development. 7, 19 (2012).</li><li>Waller, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experiments+on+the+section+of+the+glossopharyngeal+and+hypoglossal+nerves+of+the+frog,+and+observations+of+the+alterations+pro-+duced+thereby+in+the+structure+of+their+primitive+fibres.">Experiments on the section of the glossopharyngeal and hypoglossal nerves of the frog, and observations of the alterations pro- duced thereby in the structure of their primitive fibres.</a> Philos. Trans. R. Soc. Lond. , 423-429 (1849).</li></ol>

The authors have nothing to disclose.

The most critical steps of this experimental design are: 1) properly mounting larvae for injury and subsequent in vivo imaging and 2) calibrating the laser and selecting the correct power settings in order to create a clean nerve transection that results in minimal extra-tissue damage. To help ensure a successful axotomy for in vivo imaging and subsequent analysis, mount multiple larvae in either individual glass bottom dishes or in a glass bottom dish with dividers. After calibrating the laser, ...

Zebrafish have been used extensively to study development of the nervous system because of their optical transparence and ease of transgenesis, which when coupled, allow for spectacular imaging of dynamic cell behaviors in a living embryo. Additionally, because zebrafish and mammals share nearly all of the genes required for nervous system formation, cellular and molecular information collected in this model organism is directly relatable to other vertebrate species. Although incredibly powerful for neural developmental studies, the zebrafish and its unique attributes have the potential to also elucidate the mechanisms that maintain and rebuild the nervous system after injury. Zebrafish larvae maintain their translucence into late larval stages and pigmentation can be effectively blocked with either the use of pharmacological inhibitors of melanin production or genetic mutants that lack pigment cells. Thus, using this model organism to study injury and regeneration in older animals is possible and offers the unique opportunity to directly investigate the cellular and molecular mechanisms that rebuild the nervous system. In this manuscript, we describe how to efficiently and reproducibly injure nerves in the PNS of zebrafish larvae. This injury paradigm lends itself to studying not only degeneration, but also the responses of peripheral glia and immune cells as well as the interactions between these populations during regeneration.
The PNS is a complex network of motor and sensory nerves that is necessary to pass information between the central nervous system (CNS) and the skin, organs and muscle of the body, allowing an organism to interact with its environment and survive. Along these nerves, peripheral glia, including myelinating and non-myelinating Schwann cells and perineurial glia, as well as connective tissue, encase the axons and ultimately form the mature nerve. Injury of these nerves initiates a process known as Wallerian degeneration 10. This mechanism of axonal fragmentation, immune recruitment, debris clearance and regeneration is very stereotyped and genetically regulated 1. Previous studies in mammalian systems have described the roles of Schwann cells during nerve degeneration and regeneration 1, 2, 6, 8. In these studies of fixed tissue or cell culture, Schwann cells not only recruited macrophages to the injury site to aid in debris clearance, but also aided in myelin phagocytosis themselves. While these studies have been incredibly informative, we have never before visualized glial responses to peripheral axon injury in vivo in real time, and no other studies have investigated the relationship between the different classes of peripheral glia during these events.
Recently, several labs have investigated Wallerian degeneration using zebrafish and laser-mediated axon injury similar to what we describe here 4, 5, 7, 9. In some of these studies, superficial sensory axons were axotomized in young larvae using a custom built, two-photon confocal microscope 4, 5, 9. In another study, which is very similar to our own, deeper axons within the ventral motor nerve were transected in 5 day old larvae using a commercially available laser ablation system 7. In both of these experimental set-ups, the focus was on Wallerian degeneration and both axons and immune cells were imaged. To expand on these studies, we describe injuring motor axons in older larvae with more mature, myelinated nerves and assay the response of all nerve-associated peripheral glia during degeneration and regeneration.
To do this, we transect motor nerves in 6 and 7 day post fertilization (dpf) larvae and visualize the responses of individual glial populations as well as investigate the interactions between these populations along injured axons. Using double and triple transgenic lines that label peripheral glia, including Schwann cells and perineurial glia, as well as a marker for axons, we use a commercially available laser ablation system consisting of a nitrogen-pumped dye laser (wavelength 435 nm) attached to a spinning disc confocal system to create axon transections. This experimental set-up allows us to visualize live, larval zebrafish, injure specific peripheral motor axon tracts and time-lapse image the responses of distinct glial populations to axon injury and their relationship to one another. This protocol can be further adapted to create nerve injuries in zebrafish of different ages, with different transgenic lines or genetic mutants to address different scientific questions.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Phenylthiourea</td>
 <td>Sigma</td>
 <td>P7629-100G</td>
 </tr>
 <tr>
 <td>Finquel Tricaine Methanesulfonate MS-222</td>
 <td>Argent Chemical</td>
 <td>C-FINQ-UE-100G</td>
 </tr>
 <tr>
 <td>Low melting point agarose</td>
 <td>Sigma</td>
 <td>A9414-10G</td>
 </tr>
 <tr>
 <td>Quad CELLview Cell Culture Dishes, Glass Bottom, Sterile, Greiner Bio One</td>
 <td>VWR/Greiner</td>
 <td>89125-444</td>
 </tr>
 <tr>
 <td>Single well glass bottom Petri dishes 35 x 10 mm, 12 mm thick</td>
 <td>Willco Wells</td>
 <td><a href="http://www.willcowells.com/en/products/willco-dish-glass-bottom-dishes/gwst-3512/" target="_blank">GWSt-3512</a></td>
 </tr>
 <tr>
 <td>MicroPoint Laser System with all components</td>
 <td>Andor Technology - purchased through Quorum Technologies, Inc.</td>
 <td>2203-SYS</td>
 </tr>
 <tr>
 <td>MicroPoint Laser Courmarin dye (435 nm)</td>
 <td>Andor Technology </td>
 <td>MP-27-435-DYE</td>
 </tr>
 <tr> </tr>
 </tbody>
 </table>

motor nerve transection and time-lapse imaging of glial cell behaviors in live zebrafish

The nervous system is often described as a hard-wired component of the body even though it is a considerably fluid organ system that reacts to external stimuli in a consistent, stereotyped manner, while maintaining incredible flexibility and plasticity. Unlike the central nervous system (CNS), the peripheral nervous system (PNS) is capable of significant repair, but we have only just begun to understand the cellular and molecular mechanisms that govern this phenomenon. Using zebrafish as a model system, we have the unprecedented opportunity to couple regenerative studies with in vivo imaging and genetic manipulation. Peripheral nerves are composed of axons surrounded by layers of glia and connective tissue. Axons are ensheathed by myelinating or non-myelinating Schwann cells, which are in turn wrapped into a fascicle by a cellular sheath called the perineurium. Following an injury, adult peripheral nerves have the remarkable capacity to remove damaged axonal debris and re-innervate targets. To investigate the roles of all peripheral glia in PNS regeneration, we describe here an axon transection assay that uses a commercially available nitrogen-pumped dye laser to axotomize motor nerves in live transgenic zebrafish. We further describe the methods to couple these experiments to time-lapse imaging of injured and control nerves. This experimental paradigm can be used to not only assess the role that glia play in nerve regeneration, but can also be the platform for elucidating the molecular mechanisms that govern nervous system repair.

The assay described here can be used to assess the response of glial cells and other nerve-associated cell populations to axonal injury in vivo. Movie 1 shows an example of a nerve injury created using this method and the response of surrounding glial cells. This experiment was performed in Tg(nkx2.2a:megfp);Tg(olig2:dsred) zebrafish, in which perineurial glia express a membrane targeted EGFP and motor neurons express cytosolic DSRed. The injury was made along the rostral projection of ...

Watch this Scientific Journal Video about Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish at JoVE.com

Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

Although the peripheral nervous system (PNS) is capable of significant repair after injury, little is known about the cellular and molecular mechanisms that govern this phenomenon. Using live, transgenic zebrafish and a reproducible nerve transection assay, we can study dynamic glial cell behaviors during nerve degeneration and regeneration.

The  nervous system is often described as a hard-wired component of the body even  though it is a considerably fluid organ system that reacts to external ...

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11.8K Views.  University of Virginia. Although the peripheral nervous system (PNS) is capable of significant repair after injury, little is known about the cellular and molecular mechanisms that govern this phenomenon. Using live, transgenic zebrafish and a reproducible nerve transection assay, we can study dynamic glial cell behaviors during nerve degeneration and regeneration.

Video: Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

Methods for Experimental Manipulations after Optic Nerve Transection in the Mammalian CNS

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. Viral vectors, plasmids or short interfering RNAs (siRNAs) can also be delivered to the vitreous chamber in order to infect or transfect retinal cells 5-12. The high tropism of Adeno-Associated Virus (AAV) vectors is beneficial to target RGCs, with an infection rate approaching 90% of cells near the injection site 6, 7, 13-15. Moreover, RGCs can be selectively transfected by applying siRNAs, plasmids, or viral vectors to the cut end of the optic nerve 16-19 or injecting vectors into their target the superior colliculus 10. This allows researchers to study apoptotic mechanisms in the injured neuronal population without confounding effects on other bystander neurons or surrounding glia. 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 window for experimental manipulations directed against pathways involved in apoptosis. Manipulations that directly target RGCs from the transected optic nerve stump are performed at the time of axotomy, immediately after cutting the nerve. In contrast, when substances are delivered via an intraocular route, they can be injected prior to surgery or within the first 3 days after surgery, preceding the initiation of apoptosis in axotomized RGCs. In the present article, we demonstrate several methods for experimental manipulations after optic nerve transection.

Optic Nerve transection is a widely used model of adult CNS injury. This model is ideal for performing a number of experimental manipulations that target the retina globally or directly target the injured neuronal population of retinal ganglion cells.

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 ...

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to neuroblasts able to move along the rostral migratory stream (RMS) towards the olfactory bulb (OB). This long-distance migration is required for the subsequent maturation of newborn neurons in the OB, but the molecular mechanisms regulating this process are still unclear. Investigating the signaling pathways controlling neuroblast motility may not only help understand a fundamental step in neurogenesis, but also have therapeutic regenerative potential, given the ability of these neuroblasts to target brain sites affected by injury, stroke, or degeneration.
In this manuscript we describe a detailed protocol for in vivo postnatal electroporation and subsequent time-lapse imaging of neuroblast migration in the mouse RMS. Postnatal electroporation can efficiently transfect SVZ progenitor cells, which in turn generate neuroblasts migrating along the RMS. Using confocal spinning disk time-lapse microscopy on acute brain slice cultures, neuroblast migration can be monitored in an environment closely resembling the in vivo condition. Moreover, neuroblast motility can be tracked and quantitatively analyzed. As an example, we describe how to use in vivo postnatal electroporation of a GFP-expressing plasmid to label and visualize neuroblasts migrating along the RMS. Electroporation of shRNA or CRE recombinase-expressing plasmids in conditional knockout mice employing the LoxP system can also be used to target genes of interest. Pharmacological manipulation of acute brain slice cultures can be performed to investigate the role of different signaling molecules in neuroblast migration. By coupling in vivo electroporation with time-lapse imaging, we hope to understand the molecular mechanisms controlling neuroblast motility and contribute to the development of novel approaches to promote brain repair.

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

Neuroblast migration is a fundamental event in postnatal neurogenesis. We describe a protocol for efficient labeling of neuroblasts by in vivo postnatal electroporation and subsequent visualization of their migration using time-lapse imaging of acute brain slices. We include a description for the quantitative analysis of neuroblast dynamics by video tracking.

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

Ex Vivo Oculomotor Slice Culture from Embryonic GFP-Expressing Mice for Time-Lapse Imaging of Oculomotor Nerve Outgrowth

Accurate eye movements are crucial for vision, but the development of the ocular motor system, especially the molecular pathways controlling axon guidance, has not been fully elucidated. This is partly due to technical limitations of traditional axon guidance assays. To identify additional axon guidance cues influencing the oculomotor nerve, an ex vivo slice assay to image the oculomotor nerve in real-time as it grows towards the eye was developed. E10.5 IslMN-GFP embryos are used to generate ex vivo slices by embedding them in agarose, slicing on a vibratome, then growing them in a microscope stage-top incubator with time-lapse photomicroscopy for 24-72 h. Control slices recapitulate the in vivo timing of outgrowth of axons from the nucleus to the orbit. Small molecule inhibitors or recombinant proteins can be added to the culture media to assess the role of different axon guidance pathways. This method has the advantages of maintaining more of the local microenvironment through which axons traverse, not axotomizing the growing axons, and assessing the axons at multiple points along their trajectory. It can also identify effects on specific subsets of axons. For example, inhibition of CXCR4 causes axons still within the midbrain to grow dorsally rather than ventrally, but axons that have already exited ventrally are not affected.

An ex vivo slice assay allows oculomotor nerve outgrowth to be imaged in real time. Slices are generated by embedding E10.5 IslMN:GFP embryos in agarose, slicing on a vibratome, and growing in a stage-top incubator. The role of axon guidance pathways is assessed by adding inhibitors to the culture media.

Accurate eye movements are crucial for vision, but the development of the ocular motor system, especially the molecular pathways controlling axon ...

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of sciatic nerve injury rodent models. In this protocol, we describe a novel kinematic analysis method that uses a three-dimensional (3D) motion capture apparatus for functional evaluations using a rat sciatic nerve crush injury model. First, the rat is familiarized with treadmill walking. Markers are then attached to the designated bone landmarks and the rat is made to walk on the treadmill at the desired speed. Meanwhile, the posterior limb movements of the rat are recorded using four cameras. Depending on the software used, marker tracings are created using both automatic and manual modes and the desired data are produced after subtle adjustments. This method of kinematic analysis, which uses a 3D motion capture apparatus, offers numerous advantages, including superior precision and accuracy. Many more parameters can be investigated during the comprehensive functional evaluations. This method has several shortcomings that require consideration: The system is expensive, can be complicated to operate, and may produce data deviations due to skin shifting. Nevertheless, kinematic analysis using a 3D motion capture apparatus is useful for performing functional anterior and posterior limb evaluations. In the future, this method may become increasingly useful for generating accurate assessments of various traumas and diseases.

We introduce a kinematic analysis method that uses a three-dimensional motion capture apparatus containing four cameras and data processing software for performing functional evaluations during fundamental research involving rodent models.

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of ...

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding environment and initiate contacts with potential synaptic partners. Although the connection between dendritic branch dynamics and synaptogenesis is well established, how developmental and activity-dependent processes regulate dendritic branch dynamics is not well understood. This is partly due to the technical difficulties associated with the live imaging and quantitative analyses of these fine structures using an in vivo system. We established a method to study dendrite dynamics using Drosophila larval ventral lateral neurons (LNvs), which can be individually labeled using genetic approaches and are accessible for live imaging. Taking advantage of this system, we developed protocols to capture branch dynamics of the whole dendritic arbor of a single labeled LNv through time-lapse live imaging. We then performed post-processing to improve image quality through drift correction and deconvolution, followed by analyzing branch dynamics at the single-branch level by annotating spatial positions of all branch terminals. Lastly, we developed R scripts (Supplementary File) and specific parameters to quantify branch dynamics using the coordinate information generated by the terminal tracing. Collectively, this protocol allows us to achieve a detailed quantitative description of branch dynamics of the neuronal dendritic arbor with high temporal and spatial resolution. The methods we developed are generally applicable to sparsely labeled neurons in both in vitro and in vivo conditions.

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Here, we describe the method we employed to image highly motile dendritic filopodia in a live preparation of the Drosophila larval brain, and the protocol we developed to quantify time-lapse 3D imaging datasets for quantitative assessments of dendrite dynamics in developing neurons.

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding ...

Purification of Fibroblasts and Schwann Cells from Sensory and Motor Nerves in Vitro

The principal cells in the peripheral nervous system are the Schwann cells (SCs) and the fibroblasts. Both these cells distinctly express the sensory and motor phenotypes involved in different patterns of neurotrophic factor gene expression and other biological processes, affecting nerve regeneration. The present study has established a protocol to obtain highly purified rat sensory and motor SCs and fibroblasts more rapidly. The ventral root (motor nerve) and the dorsal root (sensory nerve) of neonatal rats (7-days-old) were dissociated and the cells were cultured for 4-5 days, followed by isolation of sensory and motor fibroblasts and SCs by combining differential digestion and differential adherence methods sequentially. The results of immunocytochemistry and flow cytometry analyses showed that the purity of the sensory and motor SCs and fibroblasts were &#62;90%. This protocol can be used to obtain a large number of sensory and motor fibroblasts/SCs more rapidly, contributing to the exploration of sensory and motor nerve regeneration.

Here, we present a method to purify fibroblasts and Schwann cells from sensory and motor nerves in vitro.

The principal cells in the peripheral nervous system are the Schwann cells (SCs) and the fibroblasts. Both these cells distinctly express the sensory and ...